Name: sensitive measurement of mitophagy by flow cytometry using the ph-dependent fluorescent reporter mt-keima
Uploaded: 2018-08-12T14:00:00
Description: Watch this Scientific Journal Video about Sensitive Measurement of Mitophagy by Flow Cytometry Using the pH-dependent Fluorescent Reporter mt-Keima at JoVE.com

This work was supported by a grant from National Research Foundation of Korea (2016R1D1A1B03931949) (to J. U.), and by the National Research foundation of Korea (NRF) grant funded by the Korea government(MSIT) (No. 2016R1A2B2008887, No. 2016R1A5A2007009) (to J.Y.)

1. Generation of HeLa cells expressing mito-Keima (mt-Keima)
<ol>
	<li>Preparation of mt-Keima lentivirus
	<ol>
		<li>Coat a 100-mm culture dish by adding 2 mL of 0.001% poly-L-lysine/phosphate-buffered saline (PBS) and allow it to stand for 5 min at room temperature.</li>
		<li>Remove the poly-L-lysine solution using a glass pipette connected to a vacuum and wash the culture dish by adding 2 mL of 1x PBS.</li>
		<li>Plate 1.5 x 106 HEK293T cells on the coated culture dish with 10 mL of DMEM...

<ol>
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Here, we present a rapid and sensitive method for using flow cytometry to measure cellular mitophagy activity in cells expressing mt-Keima. Cells undergoing a high level of mitophagy exhibit an increased ratio of PE-CF594 (561 nm)/BV605 (405 nm) excitation. Thus, mitophagy activity can be expressed as the percentage of cells exhibiting a high 561/405 ratio. We calculated the percentage of cells in the &#34;high&#34; gate region on a dot plot of PE-CF594 (561 nm) versus BV605 (405 nm), and the results showed that treatmen...

Mitochondria are organelles that are essential for cell proliferation and physiology. Mitochondria are responsible for generating more than 80% of the ATP supply via oxidative phosphorylation, and they also provide various metabolic intermediates for biosynthesis and metabolism1,2. In addition to their roles in energy supply and metabolism, mitochondria play central roles in many other important processes, including reactive oxygen species (ROS) generation, the regulation of cell death, and intracellular Ca2+ dynamics3. Alterations in mitochondrial function have been associated with human diseases, including cancer, diabetes mellitus and various neurodegenerative diseases4,5. Increased mitochondrial dysfunction and mitochondrial DNA mutations are also thought to contribute to normal aging processes6,7,8. Therefore, quality control is a crucial issue in mitochondria. Mitochondria are highly dynamic organelles that can continuously change their shape between elongated networks and short, fragmented form. Mitochondria dynamics plays an important role in maintaining the function of mitochondria as well as their degradation through mitophagy9.
Mitophagy is a mechanism that involves the selective degradation of whole mitochondria using the autophagy machinery. Mitophagy is the principle mechanism underlying mitochondrial turnover and the removal of damaged or dysfunctional mitochondria. In this process, mitochondria are first surrounded by a membrane, resulting in the formation of autophagosomes, which then fuse with lysosomes for hydrolytic degradation9. Previous genetic studies in Drosophila have suggested that two Parkinson&#39;s disease-related genes, PTEN-induced putative kinase 1 (PINK1) (PARK6) and Parkin (PARK2), function in the same pathway10,11. Subsequence studies have shown that the PINK1-Parkin pathway is responsible for eliminating damaged mitochondria and defects in this pathway result in dysfunctional mitophagy and may contribute to human diseases12,13. Defects in mitophagy processes have recently been found in various human diseases, including cancer, heart disease, liver diseases and neurodegenerative disease 14. In addition, recent studies have also shown that mitophagy is critical to many physiological processes, such as differentiation, development, and the immune response15,16,17,18,19, suggesting that mitophagy may play a more active role in controlling cell functions.
Despite recent confirmation that mitophagy plays an important role in both quality control in mitochondria and human disease, the molecular mechanisms underlying mitophagy remain poorly understood. Although mitophagy-related mechanisms have been systematically studied in yeast, studies aimed at exploring mitophagy-related mechanisms in mammalian cells have mainly focused on the PINK1-Parkin pathway. Previous studies have established that the PINK1-Parkin pathway is primarily responsible for the selective removal of damaged mitochondria via mitophagy12,20,21. However, recent studies have reported that in some models, mitophagy can be activated even in the absence of functional PINK122,23,24. These results suggest that in addition to the PINK1-Parkin pathway, there an additional unidentified pathway that can activate mitophagy.
The absence of a convenient method to assess mitophagy activity is a major obstacle to exploring the pathways that regulate mitophagy and its role in pathophysiological events. Electron microscopy is a powerful tool to directly visualize autophagosome-engulfed mitochondria. However, electron microscopy has limitations in quantifying mitophagy activity. Although strategies that use microtubule-associated protein-1 light chain 3 (LC3)-conjugated fluorescent probes, such as GFP-LC3, are currently the most widely used approaches25, the transient nature of the LC3-based signal and its high rate of false-positive signals limit its sensitivity for detecting mitophagy in cells26. A combination of western blotting to measure mitochondrial protein level, the quantification of mitochondrial DNA, and fluorescence microscopy analysis to colocalize mitochondria with either autophagosomes or lysosomes would be a good approach for assessing mitophagy. However, quantification limitations and a lack of convenience of existing methodologies call for new approaches. The introduction of a new pH-dependent fluorescent protein, the mitochondrial target Keima (mt-Keima), greatly improved the ability to detect mitophagy27. By fusing the mitochondrial-targeting sequence of cytochrome c oxidase subunit VIII (COX VIII) into Keima, mt-Keima is directed to the mitochondrial matrix. The large shift in the excitation peak of Keima from 440 nm to 586 nm (corresponding to pH 7 and pH 4, respectively) can be utilized to assess mitophagy with good sensitively in both in vitro and in vivo experiments28,29. More importantly, the resistance of mt-Keima to lysosomal degradation causes the integration of the mt-Keima signal in lysosomes, allowing for the easy quantitative measurement of mitophagy activity. The fluorescence change that occurs in mt-Keima can be analyzed using either confocal microscopy or flow cytometry28,29. However, a flow cytometry-based method for measuring mitophagy activity using mt-Keima has not been provided in detail to date.
Here, we describe a detailed protocol for a flow cytometry-based technique for measuring mitophagy activity in cells using mt-Keima. Although we have shown here that flow cytometry analysis successfully detected CCCP-induced mitophagy in HeLa cells expressing Parkin, this technique can be applied to a wide variety of cell types.

<table border="0" cellpadding="0" cellspacing="0" style="width:635px;" width="635">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">REAGENTS</td>
			<td style="width:133px;"></td>
			<td style="width:119px;"></td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">poly-L-lysine</td>
			<td style="width:133px;">Sigma-Aldrich</td>
			<td style="width:119px;">P2636</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">FBS</td>
			<td style="width:133px;">GIBCO</td>
			<td style="width:119px;">16000-044</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">penicillin/streptomycin</td>
			<td style="width:133px;">wellgene</td>
			<td style="width:119px;">LS202-02</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">PBS</td>
			<td style="width:133px;">Hyclone</td>
			<td style="width:119px;">SH30013.02</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">HEK293T</td>
			<td style="width:133px;">ATCC</td>
			<td style="width:119px;">CRL-3216</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">DMEM</td>
			<td style="width:133px;">GIBCO</td>
			<td style="width:119px;">12800-082</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">OPTI-MEM&#160;</td>
			<td style="width:133px;">GIBCO</td>
			<td style="width:119px;">31985-070</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Turbofect</td>
			<td style="width:133px;">Thermos scientific</td>
			<td style="width:119px;">R0531</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">0.45 &#956;m syringe filter</td>
			<td style="width:133px;">sartorius</td>
			<td align="right" style="width:119px;">16555</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">HeLa</td>
			<td style="width:133px;">ATCC</td>
			<td style="width:119px;">CCL-2</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">polybrene</td>
			<td style="width:133px;">Sigma-Aldrich</td>
			<td style="width:119px;">H9268</td>
			<td>8 mg/ml</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">puromycin</td>
			<td style="width:133px;">Sigma-Aldrich</td>
			<td style="width:119px;">P8833</td>
			<td>2 mg/ml&#160;</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Carbonyl cyanide m-chlorophenyl hydrazine (CCCP)</td>
			<td style="width:133px;">Sigma-Aldrich</td>
			<td style="width:119px;">C2759</td>
			<td>10 mM</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">trypsin-EDTA</td>
			<td style="width:133px;">wellgene</td>
			<td style="width:119px;">LS015-01</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">EQUIPMENTS</td>
			<td style="width:133px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">BD LSRFortessa</td>
			<td style="width:133px;">BD Bioscience</td>
			<td style="width:119px;">LSRFortessa</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">FACSDIVA</td>
			<td style="width:133px;">BD Bioscience</td>
			<td style="width:119px;">FACSDIVA (v8.0.1)</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

An example of the flow cytometry analysis of CCCP-induced mitophagy in HeLa-Parkin cells is shown in Figure 4. Using the flow cytometry analysis method described above, we can detect a dramatic increase in mitophagic cells in the &#34;high&#34; gate. The percentage of cells in the &#34;high&#34; gate was increased more than 10-fold compared with untreated control cells (Figure 4A). This increase in mitophagy activity was complete...

Watch this Scientific Journal Video about Sensitive Measurement of Mitophagy by Flow Cytometry Using the pH-dependent Fluorescent Reporter mt-Keima at JoVE.com

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

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 ...

The overall goal of this procedure is to provide a sensitive technique for measuring cellular mitophagy activity using flow cytometry. Here we demonstrate that this method was successfully used to assess the dramatic increase in HeLa cell mitophagy, after treatment with CCCP, a mitochondrial uncoupler. This technique allows mitophage activity to be measured very quickly and easily.The main advantage of this technique is that you can sensitively measure mitophage activity in living cells without any additional staining. Therefore, this technique will ultimately be able to help in understanding the mechanism and physiological role of mitophage. Demonstrating the procedure will be Jee-Hyun Um and Young Yeon Kim from my laboratory.Begin this procedure by preparing the mt-Keima, a mitochondria-targeting Keima lentivirus. First seed HEK293T cells in a poly-L-lysine coated culture dish and culture them at 37 degrees Celsius in a tissue culture incubator for one day. On the next day transfect HEK293T cells with mt-Keima lentiviral DNA and packaging DNase using a transfection reagent according to the manufacturer's instructions.After eight hours of transfection remove the transfection media and add eight milliliters of fresh media. Collect the media containing viral particles 48 hours later and remove cellular debris by centrifugation. Next filter the virus-containing media with a 0.45 micrometer syringe filter.Plate HeLa-Parkin cells in a 60 millimeter culture dish one day before harvesting the mt-Keima lentivirus and culture the cells with four milliliters of growth media for one day at 37 degrees Celsius in an incubator. On the next day remove the growth medium add one millimeter of mt-Keima virus media add an additional two milliliters of growth medium containing 3 microliters of polybrene stock solution and incubate the cells for one day in a CO2 incubator. At 24 hours after infection remove the virus media wash the cells twice with PBS and add four milliliters of growth media.To remove uninfected cells treat cells for two days with two micrograms per microliter puromycin. Prepare fresh media containing 10 micromolar CCCP. Remove the cultured media and add four milliliters of fresh growth media containing CCCP.Incubate the cells at 37 degrees Celsius in a CO2 incubator for six hours. Remove the growth medium and wash the cells once with PBS. Detach the cells by treatment with trypsin EDTA solution.Collect the cells by centrifugation and re-suspend them in one milliliter of PBS. Transfer the cells in a FACS tube and place them on ice. For flow cytometry turn on the flow cytometer and start the analyzing software.Generate a new experiment and select BV605 for the violet laser and PE-CF594 for the yellow-green laser in the parameter window. First run a control sample of cells not infected with the mt-Keima lentivirus. In the plot of forward scatter versus side scatter draw a gate, using the polygon icon, around the live cells and eliminate dead cells and cell debris.Then run HeLa-Parkin cells infected with the mt-Keima lentivirus. In the dot plot of BV605 versus side scatter adjust the voltage so that HeLa-Parkin cells expressing mt-Keima are clearly distinguished from control cells that do not express mt-Keima. Next in the dot plot of PE-CF594 versus side scatter adjust the voltage so that HeLa-Parkin cells expressing mt-Keima are clearly distinguished from control cells.Then in the dot plot of BV605 versus PE-CF594 select the mt-Keima-positive cell population by drawing a gate using the polygon icon. Next create another dot plot of BV605 versus PE-CF594 showing only mt-Keima gated cells. In this dot plot, draw a gate around untreated mt-Keima-positive cells.This gate is referred to as the low gate. Then draw another gate containing the region above the low gate. This gate is referred to as the high gate because it contains cells with high mitophagy activity.Then select the Show Population Hierarchy menu. The Percent of Parent value represents the percentage of cells in the high gate among the mt-Keima-positive population. Now run each sample and record at least 10 000 events in the mt-Keima stopping gate.Flow cytometry analysis of untreated HeLa cells expressing mt-Keima showed a low mitophagy level. CCCP treatment induced a dramatic increase in mitophagy cells in the high gate. The percentage of mitophagy cells increased by more than 10 fold at six hours after CCCP treatment.This increase in mitophagy by CCCP treatment was completely abolished by co-treatment with bafilomycin A.Once the cells expressing mito-Keima are ready mitophage measurement using flow cytometry can be conducted within one hour if prepared properly. For accurate mitophage analysis by flow cytometry it is important to set up the correct high and low gates. When analyzing new types of cells it is necessary to use well-known cells such as HeLa cells as a control.In addition it is important to remember that mitophage should be fully confirmed by additional characteristics such as a decrease in mitochondrial protein and a reduction in mitochondrial DNA. This technique can be applied to a wide variety of cell types. After watching this video you should have a good understanding of how to analyze mitophage by flow cytometry.

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Research

JoVE Journal

Biology

IP-FCM:  Immunoprecipitation Detected by Flow Cytometry

Immunoprecipitation detected by flow cytometry (IP-FCM) is an efficient method for detecting and quantifying protein-protein interactions. The basic principle extends that of sandwich ELISA, wherein the captured primary analyte can be detected together with other molecules physically associated within multiprotein complexes. The procedure involves covalent coupling of polystyrene latex microbeads with immunoprecipitating monoclonal antibodies (mAb) specific for a protein of interest, incubating these beads with cell lysates, probing captured protein complexes with fluorochrome-conjugated probes, and analyzing bead-associated fluorescence by flow cytometry. IP-FCM is extremely sensitive, allows analysis of proteins in their native (non-denatured) state, and is amenable to either semi-quantitative or quantitative analysis. As additional advantages, IP-FCM requires no genetic engineering or specialized equipment, other than a flow cytometer, and it can be readily adapted for high-throughput applications.

The IP-FCM method is presented, which allows a sensitive, robust, biochemical assessment of native protein-protein interactions, without requiring genetic engineering or large sample sizes.

Immunoprecipitation detected by flow cytometry (IP-FCM) is an efficient method for detecting and quantifying protein-protein interactions. The basic ...

Quantitative Measurement of GLUT4 Translocation to the Plasma Membrane by Flow Cytometry

Glucose is the main source of energy for the body, requiring constant regulation of its blood concentration. Insulin release by the pancreas induces glucose uptake by insulin-sensitive tissues, most notably the brain, skeletal muscle, and adipocytes. Patients suffering from type-2 diabetes and/or obesity often develop insulin resistance and are unable to control their glucose homeostasis. New insights into the mechanisms of insulin resistance may provide new treatment strategies for type-2 diabetes.
The GLUT family of glucose transporters consists of thirteen members distributed on different tissues throughout the body1. Glucose transporter type 4 (GLUT4) is the major transporter that mediates glucose uptake by insulin sensitive tissues, such as the skeletal muscle. Upon binding of insulin to its receptor, vesicles containing GLUT4 translocate from the cytoplasm to the plasma membrane, inducing glucose uptake. Reduced GLUT4 translocation is one of the causes of insulin resistance in type-2 diabetes2,3.
The translocation of GLUT4 from the cytoplasm to the plasma membrane can be visualized by immunocytochemistry, using fluorophore-conjugated GLUT4-specific antibodies.
Here, we describe a technique to quantify total amounts of GLUT4 translocation to the plasma membrane of cells during a chosen duration, using flow cytometry. This protocol is rapid (less than 4 hours, including incubation with insulin) and allows the analysis of as few as 3,000 cells or as many as 1 million cells per condition in a single experiment. It relies on anti-GLUT4 antibodies directed to an external epitope of the transporter that bind to it as soon as it is exposed to the extracellular medium after translocation to the plasma membrane.

This protocol describes a rapid technique to quantify the translocation of GLUT4 from the cytoplasm to the plasma membrane of cells by flow cytometry.

Glucose is the main source of energy for the body, requiring constant regulation of its blood concentration. Insulin release by the pancreas induces ...

Flow Cytometry Purification of Mouse Meiotic Cells

The heterogeneous nature of cell types in the testis and the absence of meiotic cell culture models have been significant hurdles to the study of the unique differentiation programs that are manifest during meiosis. Two principal methods have been developed to purify, to varying degrees, various meiotic fractions from both adult and immature animals: elutriation or Staput (sedimentation) using BSA and/or percoll gradients. Both of these methods rely on cell size and density to separate meiotic cells1-5. Overall, except for few cell populations6, these protocols fail to yield sufficient purity of the numerous meiotic cell populations that are necessary for detailed molecular analyses. Moreover, with such methods usually one type of meiotic cells can be purified at a given time, which adds an extra level of complexity regarding the reproducibility and homogeneity when comparing meiotic cell samples.
Here, we describe a refined method that allows one to easily visualize, identify, and purify meiotic cells, from germ cells to round spermatids, using FACS combined with Hoechst 33342 staining7,8. This method provides an overall snapshot of the entire meiotic process and allows one to highly purify viable cells from most stage of meiosis. These purified cells can then be analyzed in detail for molecular changes that accompany progression through meiosis, for example changes in gene expression9,10and the dynamics of nucleosome occupancy at hotspots of meiotic recombination11.

An efficient method to obtain highly purified viable meiotic fractions from mouse testis is described, which combines a refined cell dissociation protocol with fluorescent activated cell sorting (FACS). This method takes advantage of differences in the DNA content and nuclear density of discrete meiotic fractions.

The heterogeneous nature of cell types in the testis and the absence of meiotic cell culture models have been significant hurdles to the study of the ...

Optimized Staining and Proliferation Modeling Methods for Cell Division Monitoring using Cell Tracking Dyes

Fluorescent cell tracking dyes, in combination with flow and image cytometry, are powerful tools with which to study the interactions and fates of different cell types in vitro and in vivo.1-5 Although there are literally thousands of publications using such dyes, some of the most commonly encountered cell tracking applications include monitoring of:
<ol>
	<li>stem and progenitor cell quiescence, proliferation and/or differentiation6-8</li>
	<li>antigen-driven membrane transfer9 and/or precursor cell proliferation3,4,10-18 and</li>
	<li>immune regulatory and effector cell function1,18-21.</li>
</ol>
Commercially available cell tracking dyes vary widely in their chemistries and fluorescence properties but the great majority fall into one of two classes based on their mechanism of cell labeling. "Membrane dyes", typified by PKH26, are highly lipophilic dyes that partition stably but non-covalently into cell membranes1,2,11. "Protein dyes", typified by CFSE, are amino-reactive dyes that form stable covalent bonds with cell proteins4,16,18. Each class has its own advantages and limitations. The key to their successful use, particularly in multicolor studies where multiple dyes are used to track different cell types, is therefore to understand the critical issues enabling optimal use of each class2-4,16,18,24.
The protocols included here highlight three common causes of poor or variable results when using cell-tracking dyes. These are:
<ol>
	<li>Failure to achieve bright, uniform, reproducible labeling. This is a necessary starting point for any cell tracking study but requires attention to different variables when using membrane dyes than when using protein dyes or equilibrium binding reagents such as antibodies.</li>
	<li>Suboptimal fluorochrome combinations and/or failure to include critical compensation controls. Tracking dye fluorescence is typically 102 - 103 times brighter than antibody fluorescence. It is therefore essential to verify that the presence of tracking dye does not compromise the ability to detect other probes being used.</li>
	<li>Failure to obtain a good fit with peak modeling software. Such software allows quantitative comparison of proliferative responses across different populations or stimuli based on precursor frequency or other metrics. Obtaining a good fit, however, requires exclusion of dead/dying cells that can distort dye dilution profiles and matching of the assumptions underlying the model with characteristics of the observed dye dilution profile.</li>
</ol>
Examples given here illustrate how these variables can affect results when using membrane and/or protein dyes to monitor cell proliferation.

Successful use of cell tracking dyes to monitor immune cell function and proliferation involves several critical steps. We describe methods for: 1) obtaining bright, uniform, reproducible label-ing with membrane dyes; 2) selecting fluorochromes and data acquisition conditions; and 3) choosing a model to quantify cell proliferation based on dye dilution.

Fluorescent cell tracking dyes, in combination with flow and image cytometry, are powerful tools with which to study the interactions and fates of ...

Techniques for the Analysis of Extracellular Vesicles Using Flow Cytometry

Extracellular Vesicles (EVs) are small, membrane-derived vesicles found in bodily fluids that are highly involved in cell-cell communication and help regulate a diverse range of biological processes. Analysis of EVs using flow cytometry (FCM) has been notoriously difficult due to their small size and lack of discrete populations positive for markers of interest. Methods for EV analysis, while considerably improved over the last decade, are still a work in progress. Unfortunately, there is no one-size-fits-all protocol, and several aspects must be considered when determining the most appropriate method to use. Presented here are several different techniques for processing EVs and two protocols for analyzing EVs using either individual detection or a bead-based approach. The methods described here will assist with eliminating the antibody aggregates commonly found in commercial preparations, increasing signal&#8211;to-noise ratio, and setting gates in a rational fashion that minimizes detection of background fluorescence. The first protocol uses an individual detection method that is especially well suited for analyzing a high volume of clinical samples, while the second protocol uses a bead-based approach to capture and detect smaller EVs and exosomes.

Many different methods exist for the measurement of extracellular vesicles (EVs) using flow cytometry (FCM). Several aspects should be considered when determining the most appropriate method to use. Two protocols for measuring EVs are presented, using either individual detection or a bead-based approach.

Extracellular Vesicles (EVs) are small, membrane-derived vesicles found in bodily fluids that are highly involved in cell-cell communication and help ...

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+ ...

High-throughput Measurement of Dictyostelium discoideum Macropinocytosis by Flow Cytometry

Large-scale non-specific fluid uptake by macropinocytosis is important for the proliferation of certain cancer cells, antigen sampling, host cell invasion and the spread of neurodegenerative diseases. The commonly used laboratory strains of the amoeba Dictyostelium discoideum have extremely high fluid uptake rates when grown in nutrient medium, over 90% of which is due to macropinocytosis. In addition, many of the known core components of mammalian macropinocytosis are also present, making it an excellent model system for studying macropinocytosis. Here, the standard technique to measure internalized fluid using fluorescent dextran as a label is adapted to a 96-well plate format, with the samples analyzed by flow cytometry using a high-throughput sampling (HTS) attachment.
Cells are fed non-quenchable fluorescent dextran for a pre-determined length of time, washed by immersion in ice-cold buffer and detached using 5 mM sodium azide, which also stops exocytosis. Cells in each well are then analyzed by flow cytometry. The method can also be adapted to measure membrane uptake and phagocytosis of fluorescent beads or bacteria.
This method was designed to allow measurement of fluid uptake by Dictyostelium in a high-throughput, labor and resource efficient manner. It allows simultaneous comparison of multiple strains (e.g. knockout mutants of a gene) and conditions (e.g. cells in different media or treated with different concentrations of inhibitor) in parallel and simplifies time-courses.

High-throughput Measurement of Dictyostelium discoideum Macropinocytosis by Flow Cytometry

Macropinocytosis, large-scale non-specific fluid uptake, is important in many areas of clinical biology including immunology, infection, cancer, and neurodegenerative diseases. Here, existing techniques have been adapted to allow high-throughput, single-cell resolution measurement of macropinocytosis in the macropinocytosis model organism Dictyostelium discoideum using flow cytometry.

Large-scale non-specific fluid uptake by macropinocytosis is important for the proliferation of certain cancer cells, antigen sampling, host cell invasion ...

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

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

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

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

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 ...

Quantitative Analysis of Mitophagy in Murine Pancreatic &amp;#946;-Cells

Complementary Approaches to Interrogate Mitophagy Flux in Pancreatic &#946;-Cells

Mitophagy is a quality control mechanism necessary to maintain optimal mitochondrial function. Dysfunctional &#946;-cell mitophagy results in insufficient insulin release. Advanced quantitative assessments of mitophagy often require the use of genetic reporters. The mt-Keima mouse model, which expresses a mitochondria-targeted pH-sensitive dual-excitation ratiometric probe for quantifying mitophagy via flow cytometry, has been optimized in &#946;-cells. The ratio of acidic-to-neutral mt-Keima wavelength emissions can be used to robustly quantify mitophagy. However, using genetic mitophagy reporters can be challenging when working with complex genetic mouse models or difficult-to-transfect cells, such as primary human islets. This protocol describes a novel complementary dye-based method to quantify &#946;-cell mitophagy in primary islets using MtPhagy. MtPhagy is a pH-sensitive, cell-permeable dye that accumulates in the mitochondria and increases its fluorescence intensity when mitochondria are in low pH environments, such as lysosomes during mitophagy. By combining the MtPhagy dye with Fluozin-3-AM, a Zn2+ indicator that selects for &#946;-cells, and Tetramethylrhodamine, ethyl ester (TMRE) to assess mitochondrial membrane potential, mitophagy flux can be quantified specifically in &#946;-cells via flow cytometry. These two approaches are highly complementary, allowing for flexibility and precision in assessing mitochondrial quality control in numerous &#946;-cell models.

Author Spotlight: Assessment of Mitophagy Flux in Pancreatic &amp;#946;-Cells Using Effective and Robust Complementary Approaches 

This protocol outlines two methods for the quantitative analysis of mitophagy in pancreatic &#946;-cells: first, a combination of cell-permeable mitochondria-specific dyes, and second, a genetically encoded mitophagy reporter. These two techniques are complementary and can be deployed based on specific needs, allowing for flexibility and precision in quantitatively addressing mitochondrial quality control.

Mitophagy is a quality control mechanism necessary to maintain optimal mitochondrial function. Dysfunctional &#946;-cell mitophagy results in insufficient ...