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 containing 10% FBS and 1% penicillin/streptomycin, and culture the cells at 37 &#176;C in a CO2 tissue culture incubator for one day.</li>
		<li>Transfect 2 &#181;g of mt-Keima lentiviral plasmid29&#160;DNA together with packaging DNAs (psPAX 2 &#181;g and pVSVG 0.25 &#181;g) using a transfection reagent for 8 h according to the manufacturer&#39;s instructions.</li>
		<li>Remove the transfection media and add 8 mL of fresh media.</li>
		<li>Collect the media containing viral particles 48 h later. Remove cellular debris from the collected media by centrifugation at 500 x g for 5 min and filter them with a 0.45-&#181;m syringe filter. 
		Note: For long-term use, make aliquots of lentiviral particles and store the samples at -80 &#176;C. Avoid subjecting the viral samples to freeze-thaw cycles for efficient viral infection.</li>
	</ol>
	</li>
	<li>Infecting HeLa-Parkin cells with mt-Keima lentivirus
	<ol>
		<li>Plate 5 x 104 HeLa cells expressing Parkin (HeLa-Parkin) in a 60-mm culture dish with 4 mL of DMEM containing 10% FBS and 1% penicillin/streptomycin at 37 &#176;C one day before harvesting the mt-Keima lentivirus. 
		Note: Because of the absence of endogenous Parkin expression in HeLa cells30, the HeLa-Parkin cells used here to more clearly show CCCP-induced mitophagy. This procedure can also be applied to other primary or immortalized cell lines.</li>
		<li>Remove the growth medium the next day and add 1 mL of mt-Keima lentiviral media. Add an additional 2 mL of growth medium containing 3 &#181;L of polybrene stock solution (8 mg/mL in distilled water). Incubate for one day in a CO2 tissue culture incubator.</li>
		<li>Remove the mt-Keima lentiviral mixture the next day and wash the cells twice with 1x PBS. Add 4 mL of growth medium and treat the cells with 2 &#181;g/mL puromycin for two days.</li>
		<li>After two days of puromycin selection, remove the growth medium, and wash the cells twice with 1x PBS. Add growth medium, and culture the cells in a CO2 incubator until use. 
		Note: This method can be applied to generate cells that stably express mt-Keima, including other cell lines and primary cells.</li>
	</ol>
	</li>
</ol>
2. Inducing mitophagy using CCCP treatment and preparing FACS samples
<ol style="margin-left: 40px;">
	<li>Plate 5 x 104 HeLa-Parkin cells expressing mt-Keima in a 60-mm culture dish with 4 mL of DMEM containing 10% FBS and 1% penicillin/streptomycin and culture them for one day at 37 &#176;C in a CO2 tissue culture incubator.</li>
	<li>Remove the growth medium the next day and add 4 mL of fresh DMEM containing 10 &#181;M carbonyl cyanide m-chlorophenylhydrazone (CCCP). Incubate HeLa-Parkin-mt-Keima cells in a CO2 tissue incubator for 6 h.</li>
	<li>Remove the growth medium 6 h later and wash the cells with 2 mL of 1x PBS. Detach the cells by treating them with trypsin/EDTA solution and inactivate the trypsin by adding growth medium containing 10% FBS. Transfer the cells into a 15-mL conical tube.</li>
	<li>Centrifuge the cells at 500 x g for 5 min.</li>
	<li>Carefully remove the growth medium by aspiration and resuspend the cells with 1 mL of cold 1x PBS.</li>
	<li>Transfer cells into the appropriate FACS tube and place it on ice. 
	Note: Prepare uninfected HeLa cells as a negative control. 
	Note: Cells are viable on ice for several hours, and mt-Keima fluorescence signals also remain stable.</li>
</ol>
3. FACS analysis of mitophagy
Note: In this study, cells were analyzed with a flow cytometer equipped with a 405-nm and 561-nm laser. Cells were excited with a violet laser (405 nm) with emission detected at 610 &#177; 10 nm with a BV605 detector and with a yellow-green laser (561 nm) with emission detected at 610 &#177; 10 nm by a PE-CF594 detector simultaneously (Figure 1).
<ol>
	<li>Gating live cells
	<ol>
		<li>Turn on the flow cytometer and computer and log into the analysis software.</li>
		<li>Generate a new experiment or duplicate an existing experiment. To excite cells with the violet (405 nm) and yellow-green (561 nm) lasers and detect emission at 610 &#177; 10 nm detector, select BV605 and PE-CF594 in addition to forward scatter (FSC) and side scatter (SSC) in the parameter window as required fluorophores. 
		Note: All fluorophores should be added in a linear mode.</li>
		<li>Vortex each cell sample briefly to disperse cell aggregates.</li>
		<li>Place the tube containing a control HeLa-Parkin cell sample that is not infected with mt-Keima lentivirus on the sample injection port and press the &#34;run&#34; button on the cytometer.</li>
		<li>In the dot plot of FSC versus SSC, adjust the FSC and SSC voltage to place cells in the center of the dot plot.</li>
		<li>Draw a P1 gate using the Polygon icon around live cells and eliminate dead cells and cell debris (Figure 2A).</li>
	</ol>
	</li>
	<li>Adjust voltages for violet and green laser
	<ol>
		<li>Place the tube containing HeLa-Parkin cells infected with mt-Keima lentivirus on the sample injection port and press the &#34;run&#34; button on the cytometer.</li>
		<li>In the dot plot of BV605 (405 nm) versus SSC, adjust the BV605 voltage so that HeLa-Parkin cells expressing mt-Keima are clearly distinguished from control cells that do not express mt-Keima (Figure 2B).</li>
		<li>In the dot plot of PE-CF594 (561 nm) versus SSC, adjust the PE-CF594 voltage so that HeLa-Parkin cells expressing mt-Keima are clearly distinguished from control cells (Figure 2B).</li>
	</ol>
	</li>
	<li>Assess the percentage of cells in mitophagy
	<ol>
		<li>Acquire the BV605 versus PE-CF594 dot plot using a linear scale. Draw a gate using the Polygon icon around cells expressing mt-Keima, and eliminate cells not expressing mt-Keima (Figure 3A). The mt-Keima fluorescence-positive cell population is referred to as the &#34;mt-Keima&#34; gate.</li>
		<li>Create another dot plot of BV605 versus PE-CF594 showing only &#34;mt-Keima&#34;-gated cells. In this dot plot, draw a gate around untreated mt-Keima-positive cells. The basal mitophagy activity of control HeLa cells is low, and the ratio of emission at PE-CF594/BV605 is less than 1. Thus, this gate is referred to as the &#34;low&#34; gate.</li>
		<li>Draw another gate containing the above region of the &#34;low&#34; gate. This gate is referred to as the &#34;high&#34; gate because it contains cells with high mitophagy activity, that is, cells with a high ratio of emission at PE-CF594/BV605 (Figure 3B).</li>
		<li>To calculate the percentage of cells in the &#34;high&#34; gate, select the &#34;Show Population Hierarchy&#34; menu. The &#34;Percent of Parent&#34; (%Parent) value represents the percentage of cells in the &#34;high&#34; gate among the mt-Keima-positive population.</li>
		<li>Set at least 10,000 events to record in the &#34;mt-Keima&#34; stopping gate and run each sample.</li>
	</ol>
	</li>
</ol>

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</ol>

The authors have nothing to disclose.

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><tbody><tr><td>&lt;strong&gt;REAGENTS&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>poly-L-lysine</td><td>Sigma-Aldrich</td><td>P2636</td><td></td></tr><tr><td>FBS</td><td>GIBCO</td><td>16000-044</td><td></td></tr><tr><td>penicillin/streptomycin</td><td>wellgene</td><td>LS202-02</td><td></td></tr><tr><td>PBS</td><td>Hyclone</td><td>SH30013.02</td><td></td></tr><tr><td>HEK293T</td><td>ATCC</td><td>CRL-3216</td><td></td></tr><tr><td>DMEM</td><td>GIBCO</td><td>12800-082</td><td></td></tr><tr><td>OPTI-MEM&amp;nbsp;</td><td>GIBCO</td><td>31985-070</td><td></td></tr><tr><td>Turbofect</td><td>Thermos scientific</td><td>R0531</td><td></td></tr><tr><td>0.45 &amp;mu;m syringe filter</td><td>sartorius</td><td>16555</td><td></td></tr><tr><td>HeLa</td><td>ATCC</td><td>CCL-2</td><td></td></tr><tr><td>polybrene</td><td>Sigma-Aldrich</td><td>H9268</td><td>8 mg/ml</td></tr><tr><td>puromycin</td><td>Sigma-Aldrich</td><td>P8833</td><td>2 mg/ml&amp;nbsp;</td></tr><tr><td>Carbonyl cyanide m-chlorophenyl hydrazine (CCCP)</td><td>Sigma-Aldrich</td><td>C2759</td><td>10 mM</td></tr><tr><td>trypsin-EDTA</td><td>wellgene</td><td>LS015-01</td><td></td></tr><tr><td>&lt;strong&gt;EQUIPMENTS&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>BD LSRFortessa</td><td>BD Bioscience</td><td>LSRFortessa</td><td></td></tr><tr><td>FACSDIVA</td><td>BD Bioscience</td><td>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 ...

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JoVE Journal

Biology

14.8K Views.  Dong-A University. 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.

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

A Fluorescence Microscopy Assay for Monitoring Mitophagy in the Yeast Saccharomyces cerevisiae

Autophagy is important for turnover of cellular components under a range of different conditions. It serves an essential homeostatic function as well as a quality control mechanism that can target and selectively degrade cellular material including organelles1-4. For example, damaged or redundant mitochondria (Fig. 1), not disposed of by autophagy, can represent a threat to cellular homeostasis and cell survival. In the yeast, Saccharomyces cerevisiae, nutrient deprivation (e.g., nitrogen starvation) or damage can promote selective turnover of mitochondria by autophagy in a process termed mitophagy 5-9.
We describe a simple fluorescence microscopy approach to assess autophagy. For clarity we restrict our description here to show how the approach can be used to monitor mitophagy in yeast cells. The assay makes use of a fluorescent reporter, Rosella, which is a dual-emission biosensor comprising a relatively pH-stable red fluorescent protein linked to a pH-sensitive green fluorescent protein. The operation of this reporter relies on differences in pH between the vacuole (pH ~ 5.0-5.5) and mitochondria (pH ~ 8.2) in living cells. Under growing conditions, wild type cells exhibit both red and green fluorescence distributed in a manner characteristic of the mitochondria. Fluorescence emission is not associated with the vacuole. When subjected to nitrogen starvation, a condition which induces mitophagy, in addition to red and green fluorescence labeling the mitochondria, cells exhibit the accumulation of red, but not green fluorescence, in the acidic vacuolar lumen representing the delivery of mitochondria to the vacuole. Scoring cells with red, but not green fluorescent vacuoles can be used as a measure of mitophagic activity in cells5,10-12.

A Fluorescence Microscopy Assay for Monitoring Mitophagy in the Yeast Saccharomyces cerevisiae

A robust approach to monitor the delivery of organelles to the acidic lumen of the yeast vacuole for degradation and recycling is described. The method relies on the specific labeling of target organelles with a genetically encoded dual-emission fluorescence pH-biosensor, and visualization of individual cells using fluorescence microscopy.

Autophagy is important for turnover of cellular components under a range of different conditions. It serves an essential homeostatic function as well as a ...

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

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

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

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

Immunology and Infection

In Vitro and In Vivo Detection of Mitophagy in Human Cells, C. Elegans, and Mice

Mitochondria are the powerhouses of cells and produce cellular energy in the form of ATP. Mitochondrial dysfunction contributes to biological aging and a wide variety of disorders including metabolic diseases, premature aging syndromes, and neurodegenerative diseases such as Alzheimer's disease (AD) and Parkinson's disease (PD). Maintenance of mitochondrial health depends on mitochondrial biogenesis and the efficient clearance of dysfunctional mitochondria through mitophagy. Experimental methods to accurately detect autophagy/mitophagy, especially in animal models, have been challenging to develop. Recent progress towards the understanding of the molecular mechanisms of mitophagy has enabled the development of novel mitophagy detection techniques. Here, we introduce several versatile techniques to monitor mitophagy in human cells, Caenorhabditis elegans (e.g., Rosella and DCT-1/ LGG-1 strains), and mice (mt-Keima). A combination of these mitophagy detection techniques, including cross-species evaluation, will improve the accuracy of mitophagy measurements and lead to a better understanding of the role of mitophagy in health and disease.

In Vitro and In Vivo Detection of Mitophagy in Human Cells, C. Elegans, and Mice

Mitophagy, the process of clearing damaged mitochondria, is necessary for mitochondrial homeostasis and health maintenance. This article presents some of the latest mitophagy detection methods in human cells, Caenorhabditis elegans, and mice.

Mitochondria are the powerhouses of cells and produce cellular energy in the form of ATP. Mitochondrial dysfunction contributes to biological aging and a ...

Medicine

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 ...

Time-Lapse Video Microscopy for Assessment of EYFP-Parkin Aggregation as a Marker for Cellular Mitophagy

Time-lapse video microscopy can be defined as the real time imaging of living cells. This technique relies on the collection of images at different time points. Time intervals can be set through a computer interface that controls the microscope-integrated camera. This kind of microscopy requires both the ability to acquire very rapid events and the signal generated by the observed cellular structure during these events. After the images have been collected, a movie of the entire experiment is assembled to show the dynamic of the molecular events of interest. Time-lapse video microscopy has a broad range of applications in the biomedical research field and is a powerful and unique tool for following the dynamics of the cellular events in real time. Through this technique, we can assess cellular events such as migration, division, signal transduction, growth, and death. Moreover, using fluorescent molecular probes we are able to mark specific molecules, such as DNA, RNA or proteins and follow them through their molecular pathways and functions. Time-lapse video microscopy has multiple advantages, the major one being the ability to collect data at the single-cell level, that make it a unique technology for investigation in the field of cell biology. However, time-lapse video microscopy has limitations that can interfere with the acquisition of high quality images. Images can be compromised by both external factors; temperature fluctuations, vibrations, humidity and internal factors; pH, cell motility. Herein, we describe a protocol for the dynamic acquisition of a specific protein, Parkin, fused with the enhanced yellow fluorescent protein (EYFP) in order to track the selective removal of damaged mitochondria, using a time-lapse video microscopy approach.

Herein, we describe in detail a time-lapse video microscopy approach to measuring the temporal recruitment of EYFP-Parkin during the selective removal of damaged mitochondria. This dynamic process of EYFP-Parkin-dependent removal of damaged mitochondria can be used as an indicator of cellular health under different experimental conditions.

Time-lapse video microscopy can be defined as the real time imaging of living cells. This technique relies on the collection of images at different time ...

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 ...

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 ...

Visualizing Mitophagy with Fluorescent Dyes for Mitochondria and Lysosome

Mitochondria, being the powerhouses of the cell, play important roles in bioenergetics, free radical generation, calcium homeostasis, and apoptosis. Mitophagy is the primary mechanism of mitochondrial quality control and is generally studied using microscopic observation, however in vivo mitophagy assays are difficult to perform. Evaluating mitophagy by imaging live organelles is an alternative and necessary method for mitochondrial research. This protocol describes the procedures for using the cell-permeant green-fluorescent mitochondria dye MitoTracker Green and the red-fluorescent lysosome dye LysoTracker Red in live cells, including the loading of the dyes, visualization of the mitochondria and the lysosome, and expected outcomes. Detailed steps for the evaluation of mitophagy in live cells, as well as technical notes about microscope software settings, are also provided. This method can help researchers observe mitophagy using live-cell fluorescent microscopy. In addition, it can be used to quantify mitochondria and lysosomes and assess mitochondrial morphology.

Mitophagy is the primary mechanism of mitochondrial quality control. However, the evaluation of mitophagy in vivo is hindered by the lack of reliable quantitative assays. Presented here is a protocol for the observation of mitophagy in living cells using a cell-permeant green-fluorescent mitochondria dye and a red-fluorescent lysosome dye.

Mitochondria, being the powerhouses of the cell, play important roles in bioenergetics, free radical generation, calcium homeostasis, and apoptosis. ...

Quantifying Mitophagy in C. elegans and a Liver Cancer Cell Line

Detection of Mitophagy in Caenorhabditis elegans and Mammalian Cells Using Organelle-Specific Dyes

Mitochondria are essential for various biological functions, including energy production, lipid metabolism, calcium homeostasis, heme biosynthesis, regulated cell death, and the generation of reactive oxygen species (ROS). ROS are vital for key biological processes. However, when uncontrolled, they can lead to oxidative injury, including mitochondrial damage. Damaged mitochondria release more ROS, thereby intensifying cellular injury and the disease state. A homeostatic process named mitochondrial autophagy (mitophagy) selectively removes damaged mitochondria, which are then replaced by new ones. There are multiple mitophagy pathways, with the common endpoint being the breakdown of the damaged mitochondria in lysosomes. 
Several methodologies, including genetic sensors, antibody immunofluorescence, and electron microscopy, use this endpoint to quantify mitophagy. Each method for examining mitophagy has its advantages, such as specific tissue/cell targeting (with genetic sensors) and great detail (with electron microscopy). However, these methods often require expensive resources, trained personnel, and a lengthy preparation time before the actual experiment, such as for creating transgenic animals. Here, we present a cost-effective alternative for measuring mitophagy using commercially available fluorescent dyes targeting mitochondria and lysosomes. This method effectively measures mitophagy in the nematode Caenorhabditis elegans and human liver cells, which indicates its potential efficiency in other model systems.

Author Spotlight: Detection of Mitophagy in Caenorhabditis elegans and Mammalian Cells Using Organelle-Specific Dyes  

Exploring mitophagy through electron microscopy, genetic sensors, and immunofluorescence requires costly equipment, skilled personnel, and a significant time investment. Here, we demonstrate the efficacy of a commercial fluorescence dye kit in quantifying the mitophagy process in both Caenorhabditis elegans and a liver cancer cell line.

Mitochondria are essential for various biological functions, including energy production, lipid metabolism, calcium homeostasis, heme biosynthesis, ...

Quantitative Analysis of Mitophagy in Murine Pancreatic &#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 &#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 ...

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

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Przeglądaj więcej filmów

A Fluorescence Microscopy Assay for Monitoring Mitophagy in the Yeast Saccharomyces cerevisiae

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

In Vitro and In Vivo Detection of Mitophagy in Human Cells, C. Elegans, and Mice

Immunodetection of Outer Membrane Proteins by Flow Cytometry of Isolated Mitochondria

Time-Lapse Video Microscopy for Assessment of EYFP-Parkin Aggregation as a Marker for Cellular Mitophagy

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

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

Visualizing Mitophagy with Fluorescent Dyes for Mitochondria and Lysosome

Detection of Mitophagy in Caenorhabditis elegans and Mammalian Cells Using Organelle-Specific Dyes

Complementary Approaches to Interrogate Mitophagy Flux in Pancreatic β-Cells