This work was partially funded by the National Key Research and Development Program of China (2017YFA0105601, 2018YFA0107102), the National Natural Science Foundation of China (81970333,31901044,), and the Program for Professor of Special Appointment at Shanghai Institutions of Higher Learning (GZ2020008).

1. Cell culture and passaging
NOTE: The protocol is described using routinely cultured mouse embryonic fibroblasts (MEFs) as an example.
<ol>
	<li>Culture MEF cells in 10 cm cell culture dishes with 10 mL of Dulbecco&#39;s Modified Eagle Medium (DMEM). Incubate at 37 &#176;C and 5% CO2 and monitor the cells under a microscope at 100x magnification.</li>
	<li>Perform routine cell passaging.
	<ol>
		<li>When the cells reach 80%-90% confluency (every 3 days), wash the cells with 2 mL of Dulbecco&#39;s phosphate buffered saline (DPBS). Then add 2 mL of 0.05% trypsin-EDTA for 1 min to dissociate the cells, followed by 2 mL of DMEM to stop the action of trypsin-EDTA. Centrifuge the cell suspension at 100 x g for 3 min and resuspend the cell pellet in 1 mL of DMEM.</li>
		<li>Count the cells using an automated cell counter and cell counting chamber slides (see Table of Materials), and then inoculate 1.5 x 106 cells into a new 10 cm cell culture dish containing 10 mL of DMEM.</li>
	</ol>
	</li>
	<li>For the mitophagy assay, prepare a cell suspension as in step 1.2.1. Dilute the cell suspension to 1 x 105 cells/mL in fresh DMEM.</li>
	<li>Add 2 mL of the diluted cell suspension to a 20 mm confocal dish (see Table of Materials) and shake the culture dish in a &#34;cross&#34;. Incubate the cell culture dish in a 37 &#176;C, 5% CO2 incubator for 24 h.</li>
</ol>
2. Mitochondrial staining
<ol>
	<li>Remove the stock solution aliquots of the green-fluorescent mitochondrial dye and red-fluorescent lysosome dye (see Table of Materials) from the -20 &#176;C freezer.</li>
	<li>Prepare working solutions of the dyes by diluting the stock solutions 1:1,000 in DMEM and mix well. For example, add 2 &#181;L each of 1 mM mitochondrial dye and lysosome dye to 2 mL of DMEM to obtain a working concentration of 1 &#181;M for both dyes.</li>
	<li>Remove the medium from the confocal culture dish (step 1.4). Add 1 mL of the staining solution (prepared in step 2.2) to cover the cells. Place the cell culture dish in an incubator at 37 &#176;C, 5% CO2 for 20-30 min.</li>
</ol>
3. Confocal imaging
<ol>
	<li>Prepare 1 L of Krebs-Henseleit (KH) buffer (138.2 mM NaCl, 3.7 mM KCl, 0.25 mM CaCl2, 1.2 mM KH2PO4, 1.2 mM MgSO4.7H2O, 15 mM glucose, and 21.85 mM HEPES; final pH 7.4) and store at 4 &#176;C (for up to 1 month).</li>
	<li>On the day of confocal imaging, remove the KH buffer from the refrigerator in advance and pre-warm it to room temperature (20 to 25 &#176;C).</li>
	<li>Set the parameters of the confocal microscopy imaging software (see Table of Materials): For dual excitation images, use sequential excitation at 488 nm and 543 nm, and collect emission at 505-545 nm and &#62;560 nm, respectively. 
	NOTE: Set the imaging settings as follows. Scan Mode: frame; Speed: 9; Average: number, 1; Gain: 450 to 600; Pinhole: 30 to 200; laser: &#60;10%. It is best to start the imaging software first and then completely turn on the 488 nm laser. The 543 nm laser needs to be turned on and stabilized for 3-5 min before use (Figure 1A).</li>
	<li>Remove the culture medium containing the dye from the incubator (step 2.3) and add 1 mL of KH buffer to the dish.</li>
	<li>To induce mitophagy, treat the cells with carbonyl cyanide-4-(trifluoromethoxy) phenylhydrazone (FCCP) at 1 &#181;M (final concentration) in KH buffer for 10 min at room temperature, and immediately proceed to image the cells using the confocal microscope.</li>
	<li>Apply an appropriate amount of oil to the top of the 63x oil lens (see Table of Materials). Place the cell sample on the sample stage of the confocal microscope and move it directly above the objective lens.</li>
	<li>Use the imaging software to find the sample by clicking on the Locate tab in the top left corner of the software interface (Figure 1B). Select a green filter set for the experiment.</li>
	<li>Use the coarse adjustment knob to quickly focus by moving the objective lens up and down. After the cell sample is clearly visible through the eyepiece, search and focus the area of single cells and move it to the center of the field of view.</li>
	<li>Click on the Acquisition tab in the top left corner in the software interface to acquire images. Select only the 488 nm channel and the frame resolution 1024 x 1024 for preview.</li>
	<li>Click on the Live tab in the top left corner to start a live scan. Adjust the field of view to the sharpest and adjust the laser power by moving the slider left or right (Figure 1A). Keep the gain setting below 600 to avoid overexposure.</li>
	<li>Adjust the pinhole value to 156, gain value to 545, and digital offset value to 0.</li>
	<li>Select the best field of view, check the two channels (488 nm and 543 nm), and choose the frame resolution 1024 x 1024. Click Snap to acquire 2D images. Save the acquired images. 
	&#8203;NOTE: The green mitochondria dye has an excitation peak at 490 nm and an emission peak at 516 nm; it can be excited using a 488 nm laser. The red lysosome dye has an excitation peak at 576 nm and an emission peak at 590 nm; it can be excited using a 543 nm laser.</li>
</ol>
4. Image analysis
<ol>
	<li>Open the saved image with ImageJ and import the merged image into it.</li>
	<li>Manually count the number of yellow dots in each cell, which indicate that the lysosome is engulfing mitochondria.</li>
</ol>

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The authors have no conflicts of interest to disclose.

The protocol described here provides a method for evaluating and monitoring the dynamic process of mitophagy in living cells, involving autophagosomes, lysosomes, and mitochondrial fission, through co-staining with cell-permeant mitochondria and lysosome dyes. The method can also be used to identify mitochondria and assess mitochondrial morphology. Both dyes used in this study should be protected from light, multiple freeze-thaw cycles should be avoided, and the dyes should be stored in single-use aliquots as much as pos...

Mitochondria are the powerhouses of nearly all eukaryotic cells1,2. In addition to ATP production through oxidative phosphorylation, mitochondria play a vital role in other processes such as bioenergetics, calcium homeostasis, free radical generation, apoptosis, and cellular homeostasis3,4,5. As mitochondria generate reactive oxygen species (ROS) from multiple complexes in the electron transport chain, they are constantly stimulated by potential oxidative stress, which can eventually lead to structural damage and dysfunction when the antioxidant defense system collapses6,7. Mitochondrial dysfunction has been found to contribute to many diseases, including metabolic disorders, neurodegeneration, and cardiovascular disease8. Therefore, it is crucial to maintain healthy mitochondrial populations and their proper function. Mitochondria are highly plastic and dynamic organelles; their morphology and function are controlled by mitochondrial quality control mechanisms, including post-translational modifications (PTM) of mitochondrial proteins, mitochondrial biogenesis, fusion, fission, and mitophagy9,10. Mitochondrial fission mediated by dynamin-related protein 1 (DRP1), a GTPase of the dynamin superfamily of proteins, results in small and round mitochondria and isolates the dysfunctional mitochondria, which can be cleared and degraded by mitophagy11,12.
Mitophagy is a cellular process that selectively degrades mitochondria by autophagy, usually occurring in damaged mitochondria following injury, aging, or stress. Subsequently, these mitochondria are delivered to lysosomes for degradation10. Thus, mitophagy is a catabolic process that helps maintain the quantity and quality of mitochondria in a healthy state in a wide range of cell types. It plays a crucial role in the restoration of cellular homeostasis under normal physiological and stress conditions13,14. Cells are characterized by a complex mitophagy mechanism, which is induced by different signals of cellular stress and developmental changes. Mitophagy regulatory pathways are classified as ubiquitin-dependent or receptor-dependent15,16; the ubiquitin-dependent autophagy is mediated by the kinase PINK1 and the recruitment of ubiquitin ligase Parkin E3 to the mitochondria17,18, while receptor-dependent autophagy involves the binding of autophagy receptors to the microtubule-associated protein light chain LC3 that mediates mitophagy in response to mitochondrial damage19.
Transmission electron microscopy (TEM) is the most commonly used method, and still one of the best methods, to observe and detect mitophagy20. The morphological features of mitophagy are autophagosomes or autolysosomes formed by the fusion of autophagosomes with lysosomes, which can be observed from electron microscopy images21. The weakness of electron microscopy (EM), however, is the inability to monitor the dynamic processes of mitophagy, such as mitochondrial depolarization, mitochondrial fission, and fusion of autophagosomes and lysosomes, in the living cell20. Thus, evaluating mitophagy through imaging living organelles is an attractive alternative method for mitochondrial research. The live cell imaging technique described here uses two fluorescent dyes to stain mitochondria and lysosomes. When mitophagy occurs, damaged or superfluous mitochondria engulfed by autophagosomes are stained green by the mitochondrial dye, while the red dye stains the lysosomes. The fusion of these autophagosomes and lysosomes, referred to as autolysosomes, causes the green and red fluorescence to overlap and manifest as yellow dots, thus indicating the occurrence of mitophagy22. The cell-permeant mitochondria dye (MitoTracker Green) contains a mildly thiol-reactive chloromethyl moiety to label mitochondria23. To label mitochondria, cells are simply incubated with the dye, which diffuses passively across the plasma membrane and accumulates in active mitochondria. This mitochondria dye can easily stain live cells, and is less effective in staining aldehyde-fixed or dead cells. The lysosome dye (LysoTracker Red) is a fluorescent acidotropic probe used for labeling and tracking acidic organelles in live cells. This dye exhibits a high selectivity for acidic organelles and can effectively label live cells at nanomolar concentrations24.
The procedures for using these fluorescent dyes in living cells, including loading the dyes and the visualization of mitochondria and lysosomes, are presented here. This method can help researchers observe mitophagy using live-cell fluorescent microscopy. It can also be used to quantify mitochondria and lysosomes, and assess mitochondrial morphology.

<table><tbody><tr><td>Automated cell counter</td><td>Countstar</td><td>IC1000</td><td></td></tr><tr><td>Cell counting chamber slides</td><td>Countstar</td><td>12-0005-50</td><td></td></tr><tr><td>Dulbecco&#039;s modified Eagle medium (DMEM)</td><td>Corning</td><td>10-013-CV</td><td></td></tr><tr><td>Dulbecco&#039;s phosphate-buffered saline (DPBS)</td><td>Corning</td><td>21-031-CVC</td><td></td></tr><tr><td>Glass bottom cell culture dish (confocal dish)</td><td>NEST</td><td>801002</td><td></td></tr><tr><td>Image J (Rasband, NIH)</td><td>NIH</td><td></td><td>https://imagej.nih.gov/ij/download.html</td></tr><tr><td>Krebs&amp;ndash;Henseleit(KHB) buffer</td><td></td><td></td><td>Self-prepared</td></tr><tr><td>LysoTracker Red</td><td>Invitrogen</td><td>1818430</td><td>100 &amp;micro;mol/L, red-fluorescent lysosome dye</td></tr><tr><td>MitoTracker Green</td><td>Invitrogen</td><td>1842298</td><td>200 &amp;micro;mol/L stock, green-fluorescent mitochondria dye</td></tr><tr><td>Mouse Embryonic Fibroblasts</td><td></td><td></td><td>Self-prepared</td></tr><tr><td>Objective (63x oil lens)</td><td>ZEISS</td><td>ZEISS LSM 880</td><td></td></tr><tr><td>Trypsin-EDTA 0.25%</td><td>Gibico</td><td>Cat# 25200056</td><td></td></tr><tr><td>ZEISS LSM 880 Confocal Laser Scanning Microscope</td><td>ZEISS</td><td>ZEISS LSM 880</td><td></td></tr><tr><td>ZEN Microscopy Software 2.1 (confocal microscope imaging software)</td><td>ZEISS</td><td>ZEN 2.1</td><td></td></tr></tbody></table>

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.

MitoTracker Green is a green-fluorescent mitochondrial stain that is able to accurately localize to mitochondria. The dye can easily stain live cells and is less effective in staining aldehyde-fixed or dead cells (Figure 2). The red-fluorescent lysosome dye LysoTracker Red is capable of labeling and tracking acidic lysosomal organelles and can only stain live cells (Figure 2). Confocal microscope imaging allows the visualization of mitochondria and lysosomes sta...

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Visualizing Mitophagy with Fluorescent Dyes for Mitochondria and Lysosome

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. ...

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4.2K Views.  Tongji University. 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. 

Video: Visualizing Mitophagy with Fluorescent Dyes for Mitochondria and Lysosome

Labeling Stem Cells with Fluorescent Dyes for non-invasive Detection with Optical Imaging

Optical imaging (OI) is an easy, fast and inexpensive tool for in vivo monitoring of new stem cell based therapies. The technique is based on ex vivo labeling of stem cells with a fluorescent dye, subsequent intravenous injection of the labeled cells and visualization of their accumulation in specific target organs or pathologies. The presented technique demonstrates how we label human mesenchymal stem cells (hMSC) by simple incubation with the lipophilic fluorescent dye DiD (C67H103CIN2O3S) and how we label human embryonic stem cells (hESC) with the FDA approved fluorescent dye Indocyanine Green (ICG). The uptake mechanism is via adherence and diffusion of the lypophilic dye across the phospholipid cell membrane bilayer. The labeling efficiency is usually improved if the cells are incubated with the dye in serum-free media as opposed to incubation in serum-containing media. Furthermore, the addition of the transfection agent Protamine Sulfate significantly improves contrast agent uptake.

This video shows techniques for labeling of human embryonic stem cells and mesenchymal stem cells with fluorescent dyes. This technique can be used for an in vivo tracking of transplanted stem cells with optical imaging and for histopathological correlations with fluorescence microscopy.

Optical imaging (OI) is an easy, fast and inexpensive tool for in vivo monitoring of new stem cell based therapies. The technique is based on ex vivo ...

Visualizing the Live Drosophila Glial-neuromuscular Junction with Fluorescent Dyes

Our project identified GFP labeled glial structures at the developing larval fly neuromuscular synapse. To look at development of live glial-nerve-muscle synapses, we developed a larval tissue preparation that had features of live intact larvae, but also had good optical properties.  This new preparation also allowed for access of perfusates to the synapse.  We used fly larvae, immersed them in artificial hemolymph, and relaxed their normal rhythmic body contractions by chilling them. Next we dissected off the posterior segments of each animal and with a blunt insect pin pushed the mouth parts backward through the body cavity.  This everted the larval body wall, like turning a sock inside-out.  We completed the dissection with ultra-fine dissection scissors and thus exposed the visceral side of the body wall muscles. The glial structures at the NMJ expressed membrane targeted GFP under the control of glial specific promoters. The post-synaptic membrane, the SSR (Subsynaptic Reticula) in muscle expressed synaptically targeted dsRed. We needed to acutely label the motor neuron terminals, the third part of the synapse. To do this we applied primary antibodies to HRP, conjugated to a far-red emitting flurophore.  To test for dye diffusion properties into the perisynaptic space between the motor neuron terminals and the SSR, we applied a solution of large Dextran molecules conjugated to far-red emitting flurophore and collected images.

Visualizing the Live Drosophila Glial-neuromuscular Junction with Fluorescent Dyes

We described structural features of the Glia-neuromuscular synapses in a novel Inside-out tissue preparation of live fly larvae using fluorescent dyes with confocal microscopy. We labeled live neuron terminals with fluorescent primary antibodies to HRP, and also visualized the perisynaptic space with fluorescent Dextrans.

Our project identified GFP labeled glial structures at the developing larval fly neuromuscular synapse. To look at development of live glial-nerve-muscle ...

Visualizing Single-molecule DNA Replication with Fluorescence Microscopy

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA template is attached to a functionalized glass coverslip and replicated extensively after introduction of replication proteins and nucleotides (Figure 1). The growing product double-strand DNA (dsDNA) is extended with laminar flow and visualized by using an intercalating dye. Measuring the position of the growing DNA end in real time allows precise determination of replication rate (Figure 2). Furthermore, the length of completed DNA products reports on the processivity of replication. This experiment can be performed very easily and rapidly and requires only a fluorescence microscope with a reasonably sensitive camera.


This protocol demonstrates a simple single-molecule fluorescence microscopy technique for visualizing DNA replication by individual replisomes in real time.

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA ...

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 ...

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

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 ...

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 ...

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 ...

An In Vivo Assay to Detect Mitophagy in Transgenic Nematodes

Source: Fang, E. F., et al. <a href="https://app.jove.com/v/56301">In Vitro and In Vivo Detection of Mitophagy in Human Cells, C. Elegans, and Mice.</a> J. Vis. Exp. (2017).
This video demonstrates an in vivo assay to study mitophagy in Caenorhabditis elegans. Upon exposing the nematodes to a mitophagy-inducing drug, the occurrence of mitophagy is assessed by co-localization between DCT-1, a mitophagy receptor on the damaged mitochondria, and the autophagosomal membrane protein LGG-1.

Test in vivo do wykrywania mitofagii u nicieni transgenicznych

1. Detection of Mitophagy in C. elegans
NOTE: The nematode C. elegans provides a platform to assay mitophagy at the organismal level. Two strains can be ...

Visualizing Mitophagy with Fluorescent Dyes for Mitochondria and Lysosome

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Labeling Stem Cells with Fluorescent Dyes for non-invasive Detection with Optical Imaging

Visualizing the Live Drosophila Glial-neuromuscular Junction with Fluorescent Dyes

Visualizing Single-molecule DNA Replication with Fluorescence Microscopy

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

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

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

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

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

Complementary Approaches to Interrogate Mitophagy Flux in Pancreatic β-Cells

Test in vivo do wykrywania mitofagii u nicieni transgenicznych