We thank members of the Gross laboratory for the critical reading of the manuscript and their comments and advice. We thank the Caenorhabditis Genetics Center (CGC), which is funded by the National Institutes of Health Office of Research Infrastructure Programs (P40 OD010440), for providing some of the strains. This research was supported by a grant from Vitalunga Ltd and the Israel Science Foundation (grant No. 989/19). The graphical abstract figure (Figure 1) was generated with BioRender.com.

NOTE: For the convenience of the readers, we have divided the protocol into two parts: one focuses on the protocol for measuring mitophagy in C. elegans, and the other focuses on the protocol for measuring mitophagy in liver cells. The list of materials can be found in the Table of Materials provided.
1. The C. elegans protocol
<ol>
	<li>Preparing the nematode growth medium (NGM) plates and Escherichia coli OP50 bacterial stock 
	NOTE: As a point of clarification, while we followed standard protocols for the preparation of the NGM plates and OP50 bacterial stock25,26, we recognize that there may be variations in these protocols between different laboratories. Therefore, we have included the complete protocols to ensure accurate replication of the experiment.
	<ol>
		<li>Make a 1 M potassium phosphate buffer, pH 6, by adding ~150 mL of 1 M K2HPO4 to 500 mL of 1 M KH2PO4 solution until pH 6 is reached. Sterilize the buffer by passing it through a 0.22 &#181;m vacuum filter/storage system.</li>
		<li>Make 0.1 M of calcium chloride (CaCl2) and magnesium sulfate (MgSO4), and sterilize them with a 0.2 &#181;m syringe filter.</li>
		<li>Prepare 5 mg/mL cholesterol in absolute ethanol. 
		NOTE: Since the cholesterol is prepared in ethanol, do not filter it.</li>
		<li>Prepare NGM-agar by dissolving 1.5 g of sodium chloride (NaCl), 1.25 g of peptone, and 8.5 g of agar in 500 mL of double-distilled water (DDW). Autoclave and let it cool to ~55 &#176;C.</li>
		<li>Under sterile conditions, add 12.5 mL of potassium phosphate buffer (pH 6), 0.5 mL of 0.1 M CaCl2, 0.5 mL of 0.1 M MgSO4, and 1 mL of 5 mg/mL cholesterol. Mix well after every addition.</li>
		<li>Add 4 mL of the melted NGM-agar to each 35 mm plate. Leave the dishes overnight to solidify at room temperature (RT, ~21 &#176;C).</li>
		<li>To make Luria-Bertani (LB) agar plates, dissolve 5 g of NaCl, 5 g of tryptone, 2.5 g of yeast extract, and 7.5 g of agar in 400 mL of distilled, deionized water (DDW), adjust the pH of the solution to 7.0, bring the volume up to 500 mL with DDW, and autoclave. Once the solution has cooled to 55 &#176;C, pour 25 mL of the mixture into each 90 mm Petri dish, and allow the plates to dry for 2 days at room temperature. Next, streak out OP50 bacteria on the dried LB plates from the glycerol stock, and incubate at 37 &#176;C overnight to obtain single colonies.</li>
		<li>Prepare 2x yeast tryptone (YT) by dissolving 8 g of tryptone, 5 g of yeast extract, and 2.5 g of sodium chloride (NaCl) in 0.5 L of DDW. Adjust the pH to 7, and autoclave.</li>
		<li>Upon cooling, inoculate an OP50 bacteria colony from the freshly streaked LB plate into 50 mL of 2x YT medium in a 250 mL Erlenmeyer flask. Shake at 37 &#176;C and 250 rpm to an optical density (OD600) of approximately 0.6.</li>
	</ol>
	</li>
	<li>Preparing the vehicle and experimental plates
	<ol>
		<li>Add 100 &#181;L of OP50 bacteria to the center of each 35 mm NGM-agar plate. Dry overnight at RT (room temperature, 21 &#176;C).</li>
		<li>Prepare 0.5 M VL-850 in DMSO, and dilute to 10 mM VL-850 using M9 buffer (22 mM KH2PO4, 42 mM Na2HPO4, 86 mM NaCl, pH 7, and 1 mM MgSO4)26. Confirm that the pH is 7.0 (if not, titrate with 0.1 M HCl), and filter-sterilize the solution with a 0.22 &#181;m syringe filter. Prepare the vehicle as described above, but without the drug (in this case, VL-850). Make 50 mM FCCP in DMSO, dilute to 1 mM FCCP with M9 buffer, and filter-sterilize the solution with a 0.22 &#181;m syringe filter.</li>
		<li>Add 25 &#181;L of vehicle (negative control), FCCP (positive control; 5 &#181;M), or VL-850 (experimental treatment; 62.5 &#181;M) to separately seeded NGM plates on the bacterial lawn.</li>
		<li>Cover the plates with aluminum foil, and leave them to dry at RT (room temperature, 21 &#176;C). Use the plates after ~16 h.</li>
	</ol>
	</li>
	<li>Obtaining synchronized young adult C. elegans hermaphrodites
	<ol>
		<li>Make 1 L of M9 buffer with 22 mM KH2PO4, 42 mM Na2HPO4, and 86 mM NaCl. Sterilize by autoclaving, and let it cool. Once cooled, add 1 mL of 1 mM MgSO4 (0.22 &#181;m filter-sterilized).</li>
		<li>Mix 0.8 mL of 2.5 N sodium hydroxide and 1 mL of a 5% solution of sodium hypochlorite with 2.2 mL of DDW to create a 4 mL alkaline hypochlorite solution (final concentration of 0.5 N for sodium hydroxide and 1.25% for sodium hypochlorite).</li>
		<li>Collect the worms (gravid hermaphrodites) into a 15 mL conical tube by washing the NGM plates with 1 mL of M9 buffer 3x to ensure all the mothers have been collected into the tube.</li>
		<li>Sediment the worms by centrifugation at 500 &#215; g for 1 min, and discard the supernatant until a volume of 1 mL remains.</li>
		<li>Add 1 mL of alkaline hypochlorite solution, and mix by inverting the tube 5x. Gently vortex the tube for 3 min to assist the release of eggs, and observe the state of worms under a dissecting stereoscope.</li>
		<li>When approximately 50% of the worms are broken, add 5 mL of M9 buffer&#160;and immediately sediment the eggs by centrifuging for 1 min at 500 &#215; g.</li>
		<li>Remove the supernatant carefully without disturbing the pellet. Add 5 mL of M9 buffer, and repeat the washing procedure 2x.</li>
		<li>Remove the supernatant until 2 mL remains, and rotate the tube (360&#176; rotation) at 20 rpm for ~16 h (RT, 21 &#176;C) to obtain synchronized L1 larvae. From this tube, take a 5 &#181;L drop on a glass slide, count the number of larvae under a stereoscope, repeat this step 3x, take an average of the three counts, and estimate the number of worms per microliter (&#181;L). Based on these calculations, add ~200 larvae per NGM plate seeded with OP50 bacteria.</li>
		<li>Grow the L1 larvae at 21 &#176;C for ~48 h until the young adult stage.</li>
	</ol>
	</li>
	<li>Drug treatment and microscopy procedure
	<ol>
		<li>Put 100 worms on each of the experimental or control plates. Ensure that the negative, positive, and experimental plates contain the vehicle, 5 &#181;M FCCP, and 62.5 &#181;M VL-850, respectively. Incubate at 21 &#176;C for 6 h.</li>
		<li>Use 1 mL of M9 buffer to wash the worms from each plate into a 1.7 mL microcentrifuge tube. Spin the tube briefly (~3 s) in a minicentrifuge. Next, discard the supernatant by pipetting the M9 buffer out gently without disturbing the worm pellet. Repeat this wash step 2x, and then gently remove the supernatant without disturbing the worm pellet.</li>
		<li>Add 200 &#181;L of M9 buffer, which contains 0.1% v/v Poloxamer 188, 0.1% v/v Pluronic F127, and 2 &#181;L of the staining kit reagent, to the worm pellet. Let the mixture rotate at 20 rpm (360&#176; rotation) for 1 h at RT (room temperature, 21 &#176;C). To protect the dyes from light, cover the tubes with aluminum foil.</li>
		<li>Spin the worms gently, as described in step 1.3.4. Then, remove the staining solution without disturbing the worm pellet. Next, wash the worms as described in step 1.3.2, and transfer them into a seeded NGM-agar plate containing the appropriate treatment-for example, worms treated with FCCP are transferred into a plate containing FCCP. Cover the plates with aluminum foil because the staining reagent is light-sensitive. 
		&#8203;NOTE: We transferred the worms to culture plates containing bacteria and the corresponding treatment agent to minimize the background noise due to excess dye. For instance, worms treated with FCCP were transferred to FCCP-supplemented plates, and so forth.</li>
		<li>Wash the worms off the plates into fresh microcentrifuge tubes using 1 mL of M9 buffer; wash the worms in the same manner 2x. Next, fix the worms with 1% formaldehyde on ice for 30 min, and wash the worms with 1 mL of M9 buffer 3x to remove residual formaldehyde. After the washes, spin the worms down to a pellet, and aspirate the maximum amount of supernatant, keeping the worm pellet intact in 10 &#181;L of M9.</li>
		<li>To prepare 2.5% agarose, weigh 0.125 g of agarose into a 10 mL borosilicate glass test tube, add 5 mL of M9 buffer, and dissolve the agarose by gently heating the tube with a Bunsen burner. Transfer the melted agarose to a dry bath set at 75 &#176;C, and using a 1 mL tip, put 100 &#181;L of the melted agarose onto a Deckgl&#228;ser microscope cover glass (24 mm x 60 mm). Immediately, put another slide perpendicularly on the agarose drop, forming a cross shape. Wait ~2 min, and gently separate the slides by (gently) pushing the upper cover glass, thus leaving the agarose pad on the bottom cover slide. 
		NOTE: Be careful while heating the agarose in the tube, and ensure the tube is held away from the body. Cut the edge of the 1 mL tip to minimize coagulation of the agarose.</li>
		<li>Transfer the worms onto the agarose pad with a Pasteur glass pipette (i.e., the entire amount in the tube, ~10 &#181;L). Remove the excess liquid with a wick made of a laboratory wipe, and then cover the worms with a smaller cover slide (24 mm x 40 mm). Apply transparent nail polish to the periphery of the smaller cover slide to prevent evaporation. Put the slide in a dark box to protect it from light.</li>
		<li>Use a confocal microscope to image the worms within 24 h at the appropriate wavelengths (see below) using a 60x magnification lens.
		<ol>
			<li>Place the slide onto the microscope stage.</li>
			<li>Open the imaging software, and right-click on the grey area of the software. Open Acquisition | Ti2 Full Pad | ND Acquisition | LUTs by clicking on the options on the pop-up that emerges as a result of the right click.</li>
			<li>Under Ti2 Full Pad, select 60x.</li>
			<li>Under Acquisition, select Eyepiece DIA, and bring the worms into focus using the microscope fine-focus knob. Under Acquisition, select Spinning disk, and choose the 16-bit - No binning option. For each fluorescence filter, set the Exposure Time to 500&#160;ms and 20 ms for Brightfield. Once these parameters are set, select Run Now, and wait for the output image to be generated as an ND2 file. 
			NOTE: The exposure time needs to be determined experimentally, as different imaging setups have different characteristics. Use lookup tables (LUTs) to examine the fluorescence intensity for each wavelength.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Image analysis
	<ol>
		<li>Open the confocal images (here, Nikon ND2 files) in ImageJ27 with the colocalization plugin. Each ND file contains image planes taken at three wavelengths (using DAPI, GFP/FITC [green], and Texas Red [red] filters) and visible light. To access these images, open the ND file in the ImageJ server, and select Split Images in the dialog box. Work with brightfield (BF), green channel, and red channel images.</li>
		<li>Generate duplicates of these images to keep the original image untouched by clicking on Image | Duplicate or using the keyboard shortcut Shift + D.</li>
		<li>To reduce the background, generate another duplicate of the image, as mentioned above. Subtract the background with a Rolling Radius of 100, and select the Create Background (don&#39;t subtract) option to generate an image with the background of the given image. Next, go to Process | Image Calculator, and subtract the first duplicated image from the second duplicated image. Use the resulting images for the colocalization analysis.</li>
		<li>To use the colocalization plugin, convert the green channel and red channel images to 8-bit. To do this, click on Image | Type | 8 bit.</li>
		<li>Click on Plugins | Colocalization. To measure the colocalization of mitochondria and lysosome signals using the colocalization plugin (see above), use the following parameters: Ratio = 75%, Threshold red channel = 80.0, Threshold green channel = 50.0. The output is an 8-bit binary image containing colocalized puncta and a combination of the three 8-bit images (green, red, and the colocalized image) in an RGB image.</li>
		<li>Focus on the puncta in the head body-wall muscle of the worms by manually selecting this area and creating a mask by clicking on Edit | Selection | Create Mask, which selects the region of interest (Figure 2A, B). Selectively remove other stained entities (e.g., the pharyngeal muscles) to analyze the head body-wall muscle region.</li>
		<li>To select the particles in the region of interest (ROI), select the colocalized 8-bit and mask images using the image calculator. Then, use the operation AND (Figure 3A) to select the puncta in the ROI. This generates an image with puncta in the ROI (Figure 3B).</li>
		<li>To analyze the area of the colocalized mitochondria and lysosomes, select Analyze | Analyze Particles, and measure the summation of the puncta between 0.1625 &#181;m2 and 4 &#181;m2.</li>
	</ol>
	</li>
</ol>
2. The Hep-3B cancer cell protocol
<ol>
	<li>Preparing the drug stock solution
	<ol>
		<li>Prepare 100 mM VL-850 in DMSO. Dilute it to 5 mM with 0.5 M HEPES buffer, pH 7.3, and sterilize the solution using a 0.22 &#181;m syringe filter. Then, prepare the vehicle as described before, but without the drug (in this case, VL-850).</li>
	</ol>
	</li>
	<li>Culturing Hep-3B cells for the experiment, drug treatment, and microscopy procedure
	<ol>
		<li>Grow Hep-3B cells in 10 cm tissue culture plates containing Dulbecco&#39;s Modified Eagle Medium (DMEM) supplemented with 10% heat-inactivated fetal bovine serum (FBS), 2% L-glutamine, and 1% tetracycline (hereafter referred to as complete DMEM). Incubate the cells at 37 &#176;C and 5% CO2.</li>
		<li>Choose a plate of Hep-3B cells displaying 70%-80% cell confluency (logarithmic growth phase), remove the medium, and wash the plate with 5 mL of prewarmed phosphate-buffered saline (PBS). Remove the PBS, and incubate the cells with 1 mL of prewarmed 0.25% trypsin/0.02% EDTA for ~3 min at 37 &#176;C; observe the cells under a tissue culture microscope (10x). Stop the trypsin digestion when the cells become round and start dissociating from the plate by adding 5 mL of complete DMEM. Centrifuge the cells at 1,000 &#215; g for 5 min, remove the supernatant, and resuspend the cells in 5 mL of complete DMEM.</li>
		<li>Determine the cell concentration. Mix 50 &#181;L of the cell suspension with 50 &#181;L of trypan blue. Count the cells using an automated cell counter or by using a hemocytometer 5x, and take an average of these counts to ensure the accuracy of the counts.</li>
		<li>Seed 42,000 Hep3G cells in each 8-well &#181;-slide in 400 &#181;L of complete DMEM (as described above). Incubate the cells for 24 h at 37 &#176;C and 5% CO2.</li>
		<li>After 24 h at ~80%-85% confluency, remove 250 &#181;L of medium from each of the wells, and add 50 &#181;L of medium with the appropriate treatment or vehicle. To follow this protocol, treat the cells with 100 &#181;M VL-850, 5 &#181;M FCCP, and a vehicle as a control.</li>
		<li>After the 6 h incubation with the compounds, add 50 &#181;L of medium to each well containing the staining reagent (0.5 &#181;L of dye for 250 &#181;L in each well). Incubate the cells with the dye for 30 min at 37 &#176;C and 5% CO2. 
		NOTE: As the staining reagent is light-sensitive, minimize the exposure to light by covering the samples with aluminum foil and working in a dim-light environment (if possible).</li>
		<li>Using a 200 &#181;L pipette, gently remove all the medium (250 &#181;L) from each well, and then wash the cells with 200 &#181;L of prewarmed PBS.</li>
		<li>Fix the cells with 200 &#181;L of fixing solution containing 4% formaldehyde and 2.5% glutaraldehyde prepared in PBS for 15 min at RT.</li>
		<li>Decant the fixative solution, and wash briefly with 200 &#181;L of PBS.</li>
		<li>Add 200 &#181;L of PBS, keep the cells covered and protected from light at 4 &#176;C, and image within 24 h. 
		NOTE: We used the spinning disk confocal microscope in four channels, including DIC, TRITC, FITC, and DAPI, as in step 1.4.8. We imaged ~300 cells per treatment.</li>
	</ol>
	</li>
	<li>Image analysis
	<ol>
		<li>Perform steps 1.5.1-1.5.5. To obtain the ROI of the cells, generate an image that highlights the area of cells. For this, choose a colocalized points (RGB) image (Figure 4A). To select the entire cell area, select Process | Binary | Create Mask to obtain a binary image (Figure 4B). To analyze the area of the cell, select Analyze | Analyze Particles, and measure all the particles in the image from 0 to infinity, which is the default setting for Analyze Particles.</li>
		<li>To analyze the area of the colocalized mitochondria and lysosomes, select a colocalized 8-bit image, convert it to binary by selecting select Process | Binary | Make Binary, select Analyze | Analyze Particles, and measure the summation of puncta between 0.1625 &#181;m2 and 4 &#181;m2. To measure the colocalized puncta, divide the area of the colocalized mitochondria and lysosomes by the total cell area.</li>
	</ol>
	</li>
</ol>

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

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D., Wells, G., Campanella, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+pharmacological+regulation+of+cellular+mitophagy.">The pharmacological regulation of cellular mitophagy.</a> Nature Chemical Biology. 13 (2), 136-146 (2017).</li><li>Porta-de-la-Riva, M., Fontrodona, L., Villanueva, A., Cer&#243;n, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Basic+Caenorhabditis+elegans+methods:+Synchronization+and+observation.">Basic Caenorhabditis elegans methods: Synchronization and observation.</a> Journal of Visualized Experiments. (64), e4019 (2012).</li><li>Wood, W. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The Nematode Caenorhabditis Elegans. , (1988).</li><li>Schneider, C. A., Rasband, W. S., Eliceiri, K. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=NIH+Image+to+ImageJ:+25+years+of+image+analysis.">NIH Image to ImageJ: 25 years of image analysis.</a> Nature Methods. 9 (7), 671-675 (2012).</li><li>Knowles, B. B., Howe, C. C., Aden, D. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Human+hepatocellular+carcinoma+cell+lines+secrete+the+major+plasma+proteins+and+hepatitis+B+surface+antigen.">Human hepatocellular carcinoma cell lines secrete the major plasma proteins and hepatitis B surface antigen.</a> Science. 209 (4455), 497-499 (1980).</li><li>Ant&#243;n, Z., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Human+Atg8-cardiolipin+interactions+in+mitophagy:+Specific+properties+of+LC3B,+GABARAPL2+and+GABARAP.">Human Atg8-cardiolipin interactions in mitophagy: Specific properties of LC3B, GABARAPL2 and GABARAP.</a> Autophagy. 12 (12), 2386-2403 (2016).</li></ol>

The authors have no conflicts of interest to declare.

Multiple mitophagy pathways involve various proteins and biomolecules (e.g., cardiolipin29). However, the endpoint of these pathways is similar-the degradation of mitochondria by lysosomal enzymes12,13. Indeed, several methods use this endpoint to quantify mitophagy. However, some methods, such as electron microscopy, demand access to costly equipment, trained experts, and an extended preparation time for the specimens and analysis. Furthe...

Mitochondria are essential for all aerobic animals, including humans. They convert the chemical energy of biomolecules to adenosine triphosphate (ATP) via oxidative phosphorylation1, synthesize heme2, degrade fatty acids through &#946; oxidation3, regulate calcium4 and iron5 homeostasis, control cell death by apoptosis6, and generate reactive oxygen species (ROS), which play a vital role in redox homeostasis7. Two complementary and opposite processes maintain the integrity and proper function of the mitochondria: the synthesis of new mitochondrial components (biogenesis) and the selective removal of damaged ones through mitochondrial autophagy (i.e., mitophagy)8.
Several mitophagy pathways are mediated by enzymes, such as PINK1/Parkin, and receptors, including FUNDC1, FKBP8, and BNIP/NIX9,10. Notably, the selective degradation of mitochondrial components can occur independently of the autophagosome machinery (i.e., through mitochondrial-derived vesicles)11. However, the endpoints of the different selective mitophagy pathways are similar&#160;(i.e.,mitochondrial degradation by lysosomal enzymes)12,13. For this reason, various methods for identifying and measuring mitophagy rely on the colocalization of mitochondrial and lysosomal markers14,15,16,17 and decreased levels of mitochondrial proteins/mitochondrial DNA18.
Below is a concise description of the existing experimental methodologies for measuring mitophagy in animal cells using fluorescence microscopy, emphasizing the mitophagy endpoint phase.
Mitophagy biosensors 
Mitochondrial degradation occurs within the acidic environment of the lysosome19. Therefore, mitochondrial components, including proteins, experience a shift from a neutral to an acidic pH at the endpoint of the mitophagy process. This pattern underpins the mechanism of action of several mitophagy biosensors, including mito-Rosella18 and tandem mCherry-GFP-FIS114. These sensors contain a pH-sensitive green fluorescence protein (GFP) and a pH-insensitive red fluorescence protein (RFP). Therefore, at the endpoint of mitophagy, the green-to-red fluorescence ratio drops significantly due to the quenching of the GFP fluorophore. The major limitations of these sensors are (1) possible F&#246;rster resonance energy transfer (FRET) between the fluorophores; (2) the differential maturation rate of GFP and RFP; (3) dissociation between the GFP and RFP due to proteolytic cleavage of the polypeptide that connects them; (4) fluorescence-emission overlap; and (5) differential fluorophore brightness and quenching15,16.
A sensor that overcomes some of these limitations is the Keima mitochondrial sensor17. The mt-Keima sensor (derived from the coral protein Keima) displays a single emission peak (620 nm). However, its excitation peaks are pH-sensitive. As a result, there is a transition from a green excitation (440 nm) to a red one (586 nm) when shifting from a high pH to an acidic pH16,17. A more recent mitophagy sensor, Mito-SRAI, has advanced the field by allowing for measurements in fixed biological samples20. However, despite the many advantages of genetic sensors, such as the ability to express them in specific tissues/cells and target them to distinct mitochondrial compartments, they also have limitations. One limitation is that the genetic sensors need to be expressed in cells or animals, which can be time-consuming and resource-intensive.
Additionally, the expression of the sensors within mitochondria themselves may influence the mitochondrial function. For example, expressing mitochondrial GFP (mtGFP) in the C. elegans worm body wall muscles expands the mitochondrial network21. This phenotype depends on the function of the stress-activated transcription factor ATFS-1, which plays an essential role in the activation of unfolded protein response in mitochondria (UPRmt)21. Therefore, although genetically encoded mitochondria/mitophagy biosensors are extremely useful for monitoring mitochondria homeostasis in vivo, they may affect the very process they are designed to measure.
Mitochondria/lysosome-specific antibodies and dyes 
Another strategy for testing mitochondrial/lysosome colocalization is to use antibodies against mitochondrial/lysosomal proteins, such as the mitochondrial outer membrane protein TOM20 and lysosomal-associated membrane protein 1 (LAMP1)22. In most cases, secondary antibodies that are conjugated to a fluorophore are used to detect the fluorescence signal via microscopy. Another strategy is to combine genetic constructs with mitochondrial/lysosomal dyes, such as expressing a LAMP1::GFP fusion construct in cells while staining them with a red mitochondrial dye (e.g., Mitotracker Red)16. These methodologies, while effective, require specific antibodies and often involve working with fixed specimens or generating cells/transgenic animals expressing fluorescently labeled mitochondria/lysosomes.
Here, we outline the utilization of a commercial lysosome/mitochondria/nuclear staining kit for assessing the mitophagy-activating properties of synthetic diamine O,O (octane-1,8-diyl)bis(hydroxylamine), hereafter referred to as VL-85023, in C. elegans worms and the human cancer cell line Hep-3B (Figure 1). The staining kit contains a mixture of lysosomal/mitochondrial/nuclear-targeted dyes that specifically stain these organelles23. We previously used this kit to demonstrate the mitophagy activity of 1,8 diaminooctane (hereafter referred to as VL-004) in C. elegans23. Importantly, we validated the staining kit results with the mito-Rosella biosensor and qPCR measurements of the mitochondrial:nuclear DNA content23. This staining kit offers the following advantages. First, there is no need to generate transgenic animals or cells expressing a mitochondrial biosensor. Therefore, we can study unmodified wild-type animals or cells and, thus, save much time, money, and labor. Moreover, as mentioned, expressing mitochondrial biosensors can change the mitochondrial function. Second, the kit is cost-effective, easy to use, and fast. Third, although we demonstrate the method in C. elegans and human cells, it could be modified for other cell types and organisms.
With that said, like any method, the staining kit protocol has drawbacks. For example, the incubation of the worms with the reagent is carried out in the absence of food (we have seen that even dead bacteria significantly decrease the staining efficiency). Although the incubation time is relatively short, it is possible that even in this time frame, homeostatic responses may be altered, including mitophagy. In addition, the binding of the dyes to the ER/mitochondrial/nuclear proteins and other biomolecules may affect the activities of these organelles. Moreover, unlike mitophagy measurement with genetic sensors, we work with worms and cells that have undergone chemical fixation. Therefore, it is impossible to continue monitoring the same worms/cells at different times. Hence, we recommend combining different methodologies to validate the function of mitophagy in a particular physiological process. Below, we present new data demonstrating that VL-850 induces robust mitophagy in C. elegans worms and Hep-3B cells. Therefore, these data further support the hypothesis that VL-850 extends the lifespan of C. elegans and protects C. elegans from oxidative damage through the induction of healthy mitophagy. We have used the proton ionophore carbonyl cyanide 4-(trifluoromethoxy)phenylhydrazone (FCCP), which is a potent mitophagy inducer24, as a positive control.

<table><tbody><tr><td>&lt;strong&gt;Reagent or resource&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Analytical balance</td><td>Mettler-Toledo</td><td></td><td></td></tr><tr><td>Bacto Agar</td><td>BD-Difco</td><td>214010</td><td></td></tr><tr><td>Bacto Peptone</td><td>BD-Difco</td><td>211677</td><td></td></tr><tr><td>Bacto Tryptone</td><td>BD-Difco</td><td>211705</td><td></td></tr><tr><td>Bacto Yeast extract</td><td>BD-Difco</td><td>212750</td><td></td></tr><tr><td>Calcium chloride</td><td>Sigma</td><td>C1016</td><td></td></tr><tr><td>Carbonyl cyanide 4-(trifluoromethoxy)phenylhydrazone (FCCP)</td><td>Sigma</td><td>C2920</td><td></td></tr><tr><td>&lt;strong&gt;Chemicals&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Cholestrol</td><td>Thermo Fisher</td><td>C/5360/48</td><td></td></tr><tr><td>DMEM high glucose</td><td>Biological Industries</td><td>01-055-1A</td><td></td></tr><tr><td>Double distilled water (DDW)</td><td></td><td></td><td></td></tr><tr><td>Dulbecco&#039;s Phosphate Buffered Saline (PBS)</td><td>Biological Industries</td><td>02-023-1A</td><td></td></tr><tr><td>FBS heat inactivated</td><td>Invitrogen</td><td>M7514</td><td></td></tr><tr><td>Gluteradehyde (25%)</td><td>Sigma</td><td>G5882</td><td></td></tr><tr><td>HEPES Buffer 1 M</td><td>Biological Industries</td><td>03-025-1B</td><td></td></tr><tr><td>L-gluatamine</td><td>Biological Industries</td><td>03-020-1B</td><td></td></tr><tr><td>Lysosome/Mitochondria/Nuclear Staining Cytopainter Reagent</td><td>Abcam</td><td>ab139487</td><td></td></tr><tr><td>Magnesium Sulfate</td><td>Sigma</td><td>M7506</td><td></td></tr><tr><td>Nonidet P 40</td><td>Sigma</td><td>74385</td><td></td></tr><tr><td>Paraformalydehyde (16%)</td><td>Electron Microscopy Sciences</td><td>15720</td><td></td></tr><tr><td>Poloxamer 188 Solution</td><td>Sigma</td><td>P5556</td><td></td></tr><tr><td>Potassium dihydrogen phosphate</td><td>Millipore</td><td>1.04873.1000</td><td></td></tr><tr><td>Potassium phosphate dibasic</td><td>Sigma</td><td>P3786</td><td></td></tr><tr><td>SeaKem LE Agarose</td><td>Lonza</td><td>50004</td><td></td></tr><tr><td>Sodium Chloride</td><td>Bio-Lab</td><td>1903059100</td><td></td></tr><tr><td>Sodium Hydroxide</td><td>Gadot</td><td>1310732</td><td></td></tr><tr><td>Sodium phosphate dibasic dodecahydrate</td><td>Sigma</td><td>4273</td><td></td></tr><tr><td>Tetracycline hydrochloride</td><td>Sigma</td><td>87128-25G</td><td></td></tr><tr><td>Trypsin-EDTA</td><td>Biological Industries</td><td>03-052-1A</td><td></td></tr><tr><td>VL-850: 1,8-diaminooxy-octane</td><td>Patented</td><td></td><td></td></tr><tr><td>&lt;strong&gt;Glass/Plastic Disposables&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>0.22 &amp;mu;m syringe filter</td><td>Millex GV</td><td>SLGV033RS</td><td></td></tr><tr><td>1.7 mL Micro Centrifuge Tubes</td><td>Lifegene</td><td>LMCT1.7B-500</td><td></td></tr><tr><td>10 cm Petri plates</td><td>Corning</td><td>430167</td><td></td></tr><tr><td>1,000 mL Erlenmeyer Flask</td><td>IsoLab, Germany</td><td></td><td></td></tr><tr><td>15 mL Sterile Polypropylene tube</td><td>Lifegene</td><td>LTB15-500</td><td></td></tr><tr><td>35 mm Petri dishes</td><td>Bar Naor</td><td>BN9015810</td><td></td></tr><tr><td>500&amp;nbsp;mL vacuum filter/storage bottle system, 0.22&amp;nbsp;&amp;mu;m</td><td>Lifegene</td><td>LG-FPE205500S</td><td></td></tr><tr><td>50 mL Sterile Polypropylene tube</td><td>Lifegene</td><td>LTB50-500</td><td></td></tr><tr><td>Deckgl&amp;auml;ser Microscope cover glass 24 x 60 mm</td><td>Marienfeld</td><td>101152</td><td></td></tr><tr><td>Glass test tubes (10 mL- 13 x 100 mm) Borosilicate glass</td><td>Pyrex</td><td>99445-13</td><td></td></tr><tr><td>iBiDi 8 well &amp;mu;-slides</td><td>iBiDi</td><td>80826</td><td></td></tr><tr><td>Microscope cover glass 24 x 40 mm</td><td>Bar Naor</td><td>BN1052421ECALN</td><td></td></tr><tr><td>Platinum iridium 0.25 mM wire</td><td>World Precision Instruments</td><td>PT1002</td><td></td></tr><tr><td>&lt;strong&gt;Instruments&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Cell counter CellDrop BF</td><td>DeNovix</td><td>CellDrop BF-UNLTD</td><td></td></tr><tr><td>Microspin FV-2400</td><td>Biosan</td><td>BS-010201-AAA</td><td></td></tr><tr><td>Nikon Yokogawa W1 Spinning Disk confocal microscope with DAPI, FITC, and TRITC filters and bright-field, with a 60x CFI Plan-Apochromat Lambda type lens (air lens) and NIS-Elements software</td><td>Nikon</td><td>CSU-W1</td><td></td></tr><tr><td>Olympus SZ61 stereo microscope</td><td>Olympus</td><td>SZ61</td><td></td></tr><tr><td>pH meter</td><td>Mettler-Toledo</td><td>MT30019032</td><td></td></tr><tr><td>Revolver Adjustable Lab Rotator</td><td>Labnet</td><td>H5600</td><td></td></tr></tbody></table>

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.

Induction of a robust mitophagy response in both C. elegans worms and Hep-3B cells with VL-850 
VL-850 protects C. elegans worms and human keratinocytes (HaCaT cells) from oxidative stress23. To further explore its mechanism of action, we examined whether VL-850 induces mitophagy in C. elegans and other human cells. To test this, we exposed C. elegans worms (young adults, 3 days post-L1) to 62.5 &#181;M VL-850, 5 &#181;M FCCP (positive cont...

Watch this Scientific Journal Video about Detection of Mitophagy in Caenorhabditis elegans and Mammalian Cells Using Organelle-Specific Dyes at JoVE.com

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.

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, ...

detection-mitophagy-caenorhabditis-elegans-mammalian-cells-using

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JoVE Journal

Biology

2.1K Views.  The Hebrew University of Jerusalem. Afbraak van beschadigde mitochondriën door mitofagie is essentieel voor de functie van het mitochondriale netwerk. Helaas neemt dit proces af met de leeftijd. We ontwikkelen en gebruiken geneesmiddelen die mitofagie activeren, waardoor gezond ouder worden wordt bevorderd om de functie van mitofagie bij veroudering en leeftijdsgerelateerde ziekten, waaronder de ziekte van Alzheimer, te begrijpen.Een van de grootste uitdagingen is om het mitofagieproces te meten zonder het te beïnvloed...

Video: Auteur in de schijnwerpers: detectie van mitofagie in Caenorhabditis elegans en zoogdiercellen met behulp van organelspecifieke kleurstoffen  

Isolation and In vitro Activation of Caenorhabditis elegans Sperm

Males and hermaphrodites are the two naturally found sexual forms in the nematode C. elegans. The amoeboid sperm are produced by both males and hermaphrodites. In the earlier phase of gametogenesis, the germ cells of hermaphrodites differentiate into limited number of sperm - around 300 - and are stored in a small 'bag' called the spermatheca. Later on, hermaphrodites continually produce oocytes1. In contrast, males produce exclusively sperm throughout their adulthood. The males produce so much sperm that it accounts for &gt;50% of the total cells in a typical adult worm2. Therefore, isolating sperm from males is easier than from that of hermaphrodites.
Only a small proportion of males are naturally generated due to spontaneous non-disjunction of X chromosome3. Crossing hermaphrodites with males or more conveniently, the introduction of mutations to give rise to Him (High Incidence of Males) phenotype are some of strategies through which one can enrich the male population3.
Males can be easily distinguished from hermaphrodites by observing the tail morphology4. Hermaphrodite's tail is pointed, whereas male tail is rounded with mating structures.
Cutting the tail releases vast number of spermatids stored inside the male reproductive tract. Dissection is performed under a stereo microscope using 27 gauge needles. Since spermatids are not physically connected with any other cells, hydraulic pressure expels internal contents of male body, including spermatids2.
Males are directly dissected on a small drop of 'Sperm Medium'. Spermatids are sensitive to alteration in the pH. Hence, HEPES, a compound with good buffering capacity is used in sperm media. Glucose and other salts present in sperm media help maintain osmotic pressure to maintain the integrity of sperm.
Post-meiotic differentiation of spermatids into spermatozoa is termed spermiogenesis or sperm activation. Shakes5, and Nelson6 previously showed that round spermatids can be induced to differentiate into spermatozoa by adding various activating compounds including Pronase E. Here we demonstrate in vitro spermiogenesis of C. elegans spermatids using Pronase E.
Successful spermiogenesis is pre-requisite for fertility and hence the mutants defective in spermiogenesis are sterile. Hitherto several mutants have been shown to be defective specifically in spermiogenesis process7. Abnormality found during in vitro activation of novel Spe (Spermatogenesis defective) mutants would help us discover additional players participating in this event.

Isolation and In vitro Activation of Caenorhabditis elegans Sperm

A protocol for isolating and activating spermatids from male C. elegans is described here. Cutting the posterior end of male releases spermatids. The spermatids can be activated by addition of protease.

Males and hermaphrodites are the two naturally found sexual forms in the nematode C. elegans. The amoeboid sperm are produced by both males and ...

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 ...

Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

Caenorhabditis elegans has often been used as a model system in studies of early developmental processes. The transparency of the 
embryos, the genetic resources, and the relative ease of transformation are qualities that make C. elegans an excellent model for early embryogenesis. Laser-based 
confocal microscopy and fluorescently labeled tags allow researchers to follow specific cellular structures and proteins in the developing embryo. For example, 
one can follow specific organelles, such as lysosomes or mitochondria, using fluorescently labeled dyes. These dyes can be delivered to the early embryo by means 
of microinjection into the adult gonad. Also, the localization of specific proteins can be followed using fluorescent protein tags. Examples are presented here 
demonstrating the use of a fluorescent lysosomal dye as well as fluorescently tagged histone and ubiquitin proteins. The labeled histone is used to visualize 
the DNA and thus identify the stage of the cell cycle. GFP-tagged ubiquitin reveals the dynamics of ubiquitinated vesicles in the early embryo. Observations 
of labeled lysosomes and GFP:: ubiquitin can be used to determine if there is colocalization between ubiquitinated vesicles and lysosomes. A technique for 
the microinjection of the lysosomal dye is presented. Techniques for generating transgenenic strains are presented elsewhere (1, 2). For imaging, embryos 
are cut out of adult hermaphrodite nematodes and mounted onto 2% agarose pads followed by time-lapse microscopy on a standard laser scanning confocal 
microscope or a spinning disk confocal microscope. This methodology provides for the high resolution visualization of early embryogenesis.

Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

This article describes a technique for the visualization of the early events of embryogenesis in the nematode Caenorhabditis elegans.

Caenorhabditis elegans has often been used as a model system in studies of early developmental processes.  The transparency of the 
embryos, the genetic ...

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

Methods to Assess Subcellular Compartments of Muscle in C. elegans

Muscle is a dynamic tissue that responds to changes in nutrition, exercise, and disease state. The loss of muscle mass and function with disease and age are significant public health burdens. We currently understand little about the genetic regulation of muscle health with disease or age. The nematode C. elegans is an established model for understanding the genomic regulation of biological processes of interest. This worm&#8217;s body wall muscles display a large degree of homology with the muscles of higher metazoan species. Since C. elegans is a transparent organism, the localization of GFP to mitochondria and sarcomeres allows visualization of these structures in vivo. Similarly, feeding animals cationic dyes, which accumulate based on the existence of a mitochondrial membrane potential, allows the assessment of mitochondrial function in vivo. These methods, as well as assessment of muscle protein homeostasis, are combined with assessment of whole animal muscle function, in the form of movement assays, to allow correlation of sub-cellular defects with functional measures of muscle performance. Thus, C. elegans provides a powerful platform with which to assess the impact of mutations, gene knockdown, and/or chemical compounds upon muscle structure and function. Lastly, as GFP, cationic dyes, and movement assays are assessed non-invasively, prospective studies of muscle structure and function can be conducted across the whole life course and this at present cannot be easily investigated in vivo in any other organism.

Methods to Assess Subcellular Compartments of Muscle in C. elegans

Skeletal muscle is essential for locomotion and is the bodies&#8217; main protein store. Muscle health measurements within C. elegans are described. Prospective changes to muscle structure and function are assessed using localized GFP and cationic dyes.

Muscle is a dynamic tissue that responds to changes in nutrition, exercise, and disease state. The loss of muscle mass and function with disease and age ...

Detecting Protein Subcellular Localization by Green Fluorescence Protein Tagging and 4',6-Diamidino-2-phenylindole Staining in Caenorhabditis elegans

In this protocol, a green fluorescence protein (GFP) fusion protein and 4',6-diamidino-2-phenylindole (DAPI) staining are used to track protein subcellular localization changes; in particular, a nuclear translocation under a heat stress condition. Proteins react correspondingly to external and internal signals. A common mechanism is to change its subcellular localization. This article describes a protocol to track protein localization that does not require an antibody, radioactive labeling, or a confocal microscope. In this article, GFP is used to tag the target protein EXL-1 in C. elegans, a member of the chloride intracellular channel proteins (CLICs) family, including mammalian CLIC4. An integrated translational exl-1::gfp transgenic line (with a promoter and a full gene sequence) was created by transformation and &#947;-radiation, and stably expresses the gene and gfp. Recent research showed that upon heat stress, not oxidative stress, EXL-1::GFP accumulates in the nucleus. Overlapping the GFP signal with both the nuclei structure and the DAPI signals confirms the EXL-1 subcellular localization changes under stress. This protocol presents two different fixation methods for DAPI staining: ethanol fixation and acetone fixation. The DAPI staining protocol presented in this article is fast and efficient and preserves both the GFP signal and the protein subcellular localization changes. This method only requires a fluorescence microscope with Nomarski, a FITC filter, and a DAPI filter. It is suitable for a small laboratory setting, undergraduate student research, high school student research, and biotechnology classrooms.

Detecting Protein Subcellular Localization by Green Fluorescence Protein Tagging and 4',6-Diamidino-2-phenylindole Staining in Caenorhabditis elegans

This protocol demonstrates how to track a protein nuclear translocation under heat stress by using a green fluorescence protein (GFP) fusion protein as a marker and 4',6-diamidino-2-phenylindole (DAPI) staining. The DAPI staining protocol is fast and preserves the GFP and protein subcellular localization signals.

In this protocol, a green fluorescence protein (GFP) fusion protein and 4',6-diamidino-2-phenylindole (DAPI) staining are used to track protein ...

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 ...

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. ...

Analyzing Mitochondrial Morphology in C. elegans

Imaging and Quantifying Mitochondrial Morphology in C. elegans During Aging

Mitochondria, important cellular organelles found in most eukaryotic cells, are major sites of energy production through aerobic respiration. Beyond this well-known role as the 'cellular powerhouse,' mitochondria are also involved in many other essential cellular processes, including the regulation of cellular metabolism, proliferation, immune signaling, and hormonal signaling. Deterioration in mitochondrial function during aging or under mitochondrial stress is often characterized by distinct changes in mitochondrial morphology and volume. The nematode C. elegans is an ideal model for studying these changes due to its transparent body and short lifespan, which facilitate live microscopy throughout its lifetime. However, even within the C. elegans field, numerous transgenic constructs and methods for mitochondrial imaging are available, each with its own limitations. Here, single-copy, matrix-localized GFP constructs are presented as a robust and reliable method for imaging mitochondrial morphology in C. elegans. This study specifically focuses on experimentally controllable factors to minimize errors and reduce variability between replicates and across studies when performing mitochondrial imaging during the aging process. Additionally, mitoMAPR is recommended as a robust method to quantify changes in mitochondrial morphology across tissue types during aging.

This protocol provides a standardized approach for imaging mitochondrial morphology in multiple tissues of C. elegans during aging.

Mitochondria, important cellular organelles found in most eukaryotic cells, are major sites of energy production through aerobic respiration. Beyond this ...

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.

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 ...

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

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