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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Reagent or resource</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>Chemicals</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&#39;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>Glass/Plastic Disposables</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>0.22 &#956;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&#160;mL vacuum filter/storage bottle system, 0.22&#160;&#956;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&#228;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 &#956;-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>Instruments</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

Research

JoVE Journal

Biology

2.1K Views.  The Hebrew University of Jerusalem. 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.

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

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 ...

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 ...

Basic Caenorhabditis elegans Methods: Synchronization and Observation

Research into the molecular and developmental biology of the nematode Caenorhabditis elegans was begun in the early seventies by Sydney Brenner and it has since been used extensively as a model organism 1. C. elegans possesses key attributes such as simplicity, transparency and short life cycle that have made it a suitable experimental system for fundamental biological studies for many years 2. Discoveries in this nematode have broad implications because many cellular and molecular processes that control animal development are evolutionary conserved 3.
C. elegans life cycle goes through an embryonic stage and four larval stages before animals reach adulthood. Development can take 2 to 4 days depending on the temperature. In each of the stages several characteristic traits can be observed. The knowledge of its complete cell lineage 4,5 together with the deep annotation of its genome turn this nematode into a great model in fields as diverse as the neurobiology 6, aging 7,8, stem cell biology 9 and germ line biology 10. 
An additional feature that makes C. elegans an attractive model to work with is the possibility of obtaining populations of worms synchronized at a specific stage through a relatively easy protocol. The ease of maintaining and propagating this nematode added to the possibility of synchronization provide a powerful tool to obtain large amounts of worms, which can be used for a wide variety of small or high-throughput experiments such as RNAi screens, microarrays, massive sequencing, immunoblot or in situ hybridization, among others. 
Because of its transparency, C. elegans structures can be distinguished under the microscope using Differential Interference Contrast microscopy, also known as Nomarski microscopy. The use of a fluorescent DNA binder, DAPI (4',6-diamidino-2-phenylindole), for instance, can lead to the specific identification and localization of individual cells, as well as subcellular structures/defects associated to them.

Basic Caenorhabditis elegans Methods: Synchronization and Observation

The easiness of maintaining and propagating the nematode C. elegans make it a nice model organism to work with. The possibility of synchronizing worms allows the work with a significant amount of subjects at the same developmental stage, what facilitates the study of one particular process in many animals.

Research into the molecular and developmental biology of the nematode Caenorhabditis elegans was begun in the early seventies by Sydney Brenner and it has ...

Prostaglandin Extraction and Analysis in Caenorhabditis elegans

Caenorhabditis elegans is emerging as a powerful animal model to study the biology of lipids1-9. Prostaglandins are an important class of eicosanoids, which are lipid signals derived from polyunsaturated fatty acids (PUFAs)10-14. These signalling molecules are difficult to study because of their low abundance and reactive nature. The characteristic feature of prostaglandins is a cyclopentane ring structure located within the fatty acid backbone. In mammals, prostaglandins can be formed through cyclooxygenase enzyme-dependent and -independent pathways10,15. C. elegans synthesizes a wide array of prostaglandins independent of cyclooxygenases6,16,17. A large class of F-series prostaglandins has been identified, but the study of eicosanoids is at an early stage with ample room for new discoveries. Here we describe a procedure for extracting and analyzing prostaglandins and other eicosanoids. Charged lipids are extracted from mass worm cultures using a liquid-liquid extraction technique and analyzed by liquid chromatography coupled to electrospray ionization tandem mass spectrometry (LC-ESI-MS/MS). The inclusion of deuterated analogs of prostaglandins, such as PGF2 &alpha;-d4 as an internal standard is recommended for quantitative analysis. Multiple reaction monitoring or MRM can be used to quantify and compare specific prostaglandin types between wild-type and mutant animals. Collision-induced decomposition or MS/MS can be used to obtain information on important structural features. Liquid chromatography mass spectrometry (LC-MS) survey scans of a selected mass range, such as m/z 315-360 can be used to evaluate global changes in prostaglandin levels. We provide examples of all three analyses. These methods will provide researchers with a toolset for discovering novel eicosanoids and delineating their metabolic pathways.

Prostaglandin Extraction and Analysis in Caenorhabditis elegans

In this paper, we describe an optimized procedure for extracting and analyzing prostaglandins and other eicosanoids from C. elegans using LC-MS/MS.

Caenorhabditis elegans is emerging as a powerful animal model to study the biology of lipids1-9. Prostaglandins are an important class of eicosanoids, ...

Using Caenorhabditis elegans as a Model System to Study Protein Homeostasis in a Multicellular Organism

The folding and assembly of proteins is essential for protein function, the long-term health of the cell, and longevity of the organism. Historically, the function and regulation of protein folding was studied in vitro, in isolated tissue culture cells and in unicellular organisms. Recent studies have uncovered links between protein homeostasis (proteostasis), metabolism, development, aging, and temperature-sensing. These findings have led to the development of new tools for monitoring protein folding in the model metazoan organism Caenorhabditis elegans. In our laboratory, we combine behavioral assays, imaging and biochemical approaches using temperature-sensitive or naturally occurring metastable proteins as sensors of the folding environment to monitor protein misfolding. Behavioral assays that are associated with the misfolding of a specific protein provide a simple and powerful readout for protein folding, allowing for the fast screening of genes and conditions that modulate folding. Likewise, such misfolding can be associated with protein mislocalization in the cell. Monitoring protein localization can, therefore, highlight changes in cellular folding capacity occurring in different tissues, at various stages of development and in the face of changing conditions. Finally, using biochemical tools ex vivo, we can directly monitor protein stability and conformation. Thus, by combining behavioral assays, imaging and biochemical techniques, we are able to monitor protein misfolding at the resolution of the organism, the cell, and the protein, respectively.

Using Caenorhabditis elegans as a Model System to Study Protein Homeostasis in a Multicellular Organism

To study the relationship between protein homeostasis, stress and aging, we monitored changes in protein folding by following protein dysfunction, protein localization in the cell and protein stability at the organismal, cellular and protein levels, using the genetically tractable metazoan Caenorhabditis elegans as a model system.

The folding and assembly of proteins is essential for protein function, the long-term health of the cell, and longevity of the organism. Historically, the ...

Cultivation of Caenorhabditis elegans in Three Dimensions in the Laboratory

The use of genetic model organisms such as Caenorhabditis elegans has led to seminal discoveries in biology over the last five decades. Most of what we know about C. elegans is limited to laboratory cultivation of the nematodes that may not necessarily reflect the environments they normally inhabit in nature. Cultivation of C. elegans in a 3D habitat that is more similar to the 3D matrix that worms encounter in rotten fruits and vegetative compost in nature could reveal novel phenotypes and behaviors not observed in 2D. In addition, experiments in 3D can address how phenotypes we observe in 2D are relevant for the worm in nature. Here, a new method in which C. elegans grows and reproduces normally in three dimensions is presented. Cultivation of C. elegans in Nematode Growth Tube-3D (NGT-3D) can allow us to measure the reproductive fitness of C. elegans strains or different conditions in a 3D environment. We also present a novel method, termed Nematode Growth Bottle-3D (NGB-3D), to cultivate C. elegans in 3D for microscopic analysis. These methods allow scientists to study C. elegans biology in conditions that are more reflective of the environments they encounter in nature. These can help us to understand the overlying evolutionary relevance of the physiology and behavior of C. elegans we observe in the laboratory.

Cultivation of Caenorhabditis elegans in Three Dimensions in the Laboratory

We present a simple method to construct 3D nematode cultivation systems called NGT-3D and NGB-3D. These can be used to study nematode fitness and behaviors in habitats that are more similar to natural Caenorhabditis elegans habitats than the standard 2D laboratory C. elegans culture plates.

The use of genetic model organisms such as Caenorhabditis elegans has led to seminal discoveries in biology over the last five decades. Most of what we ...

Combined Nucleotide and Protein Extractions in Caenorhabditis elegans

A single biological sample holds a plethora of information, and it is now common practice to simultaneously investigate several macromolecules to capture a full picture of the multiple levels of molecular processing and changes between different conditions. This protocol presents the method of isolating DNA, RNA, and protein from the same sample of the nematode Caenorhabditis elegans to remove the variation introduced when these biomolecules are isolated from replicates of similarly treated but ultimately different samples. Nucleic acids and protein are extracted from the nematode using the acid guanidinium thiocyanate-phenol-chloroform extraction method, with subsequent precipitation, washing, and solubilization of each. We show the successful isolation of RNA, DNA, and protein from a single sample from three strains of nematode and HeLa cells, with better protein isolation results in adult animals. Additionally, guanidinium thiocyanate-phenol-chloroform-extracted protein from nematodes improves the resolution of larger proteins, with enhanced detectable levels as observed by immunoblotting, compared to the traditional RIPA extraction of protein.
The method presented here is useful when investigating samples using a multiomic approach, specifically for the exploration of the proteome and transcriptome. Techniques that simultaneously assess multiomics are appealing because molecular signaling underlying complex biological phenomena is thought to occur at complementary levels; however, it has become increasingly common to see that changes in mRNA levels do not always reflect the same change in protein levels and that the time of collection is relevant in the context of circadian regulations. This method removes any intersample variation when assaying different contents within the same sample (intrasample.)

Combined Nucleotide and Protein Extractions in Caenorhabditis elegans

Here, we present a protocol for the isolation of RNA, DNA, and protein from the same sample, in an effort to reduce variation, improve reproducibility, and facilitate interpretations.

A single biological sample holds a plethora of information, and it is now common practice to simultaneously investigate several macromolecules to capture ...

Measurement of Oxygen Consumption Rates in Intact Caenorhabditis elegans

Optimal mitochondrial function is critical for healthy cellular activity, particularly in cells that have high energy demands like those in the nervous system and muscle. Consistent with this, mitochondrial dysfunction has been associated with a myriad of neurodegenerative diseases and aging in general. Caenorhabditis elegans have been a powerful model system for elucidating the many intricacies of mitochondrial function. Mitochondrial respiration is a strong indicator of mitochondrial function and recently developed respirometers offer a state-of-the-art platform to measure respiration in cells. In this protocol, we provide a technique to analyze live, intact C. elegans. This protocol spans a period of ~7 days and includes steps for (1) growing and synchronization of C. elegans, (2) preparation&#160;of compounds to be injected and hydration of probes, (3) drug loading and cartridge equilibration, (4) preparation of worm assay plate and assay run, and (5) post-experiment data analysis.

Measurement of Oxygen Consumption Rates in Intact Caenorhabditis elegans

Mitochondrial respiration is critical for organismal survival; therefore, oxygen consumption rate is an excellent indicator of mitochondrial health. In this protocol, we describe the use of a commercially available respirometer to measure basal and maximal oxygen consumption rates in live, intact, and freely-motile Caenorhabditis elegans.

Optimal mitochondrial function is critical for healthy cellular activity, particularly in cells that have high energy demands like those in the nervous ...

Modeling Age-Associated Neurodegenerative Diseases in Caenorhabditis elegans

Battling human neurodegenerative pathologies and managing their pervasive socioeconomic impact is becoming a global priority. Notwithstanding their detrimental effects on the human life quality and the healthcare system, the majority of human neurodegenerative disorders still remain incurable and non-preventable. Therefore, the development of novel therapeutic interventions against such maladies is becoming a pressing urgency. Age-associated deterioration of neuronal circuits and function is evolutionarily conserved in organisms as diverse as the lowly worm Caenorhabditis elegans and humans, signifying similarities in the underlying cellular and molecular mechanisms. C. elegans is a highly malleable genetic model, which offers a well-characterized nervous system, body transparency and a diverse repertoire of genetic and imaging techniques to assess neuronal activity and quality control during ageing. Here, we introduce and describe methodologies utilizing some versatile nematode models, including hyperactivated ion channel-induced necrosis (e.g., deg-3(d) and mec-4(d)) and protein aggregate (e.g., &#945;-syunclein and poly-glutamate)-induced neurotoxicity, to monitor and dissect the cellular and molecular underpinnings of age-related neuronal breakdown. A combination of these animal neurodegeneration models, together with genetic and pharmacological screens for cell death modulators will lead to an unprecedented understanding of age-related breakdown of neuronal function and will provide critical insights with broad relevance to human health and quality of life.

Modeling Age-Associated Neurodegenerative Diseases in Caenorhabditis elegans

Here, we introduce and describe widely accessible methodologies utilizing some versatile nematode models, including hyperactivated ion channel-induced necrosis and protein aggregate-induced neurotoxicity, to monitor and dissect the cellular and molecular underpinnings of age-associated neurodegenerative diseases.

Battling human neurodegenerative pathologies and managing their pervasive socioeconomic impact is becoming a global priority. Notwithstanding their ...

Automating Aggregate Quantification in Caenorhabditis elegans

A rise in the prevalence of neurodegenerative protein conformational diseases (PCDs) has fostered a great interest in this subject over the years. This increased attention has called for the diversification and improvement of animal models capable of reproducing disease phenotypes observed in humans with PCDs. Though murine models have proven invaluable, they are expensive and are associated with laborious, low-throughput methods. Use of the Caenorhabditis elegans nematode model to study PCDs has been justified by its relative ease of maintenance, low cost, and rapid generation time, which allow for high-throughput applications. Additionally, high conservation between the C. elegans and human genomes makes this model an invaluable discovery tool. Nematodes that express fluorescently tagged tissue-specific polyglutamine (polyQ) tracts exhibit age- and polyQ length-dependent aggregation characterized by fluorescent foci. Such reporters are often employed as proxies to monitor changes in proteostasis across tissues. Manual aggregate quantification is time-consuming, limiting experimental throughput. Furthermore, manual foci quantification can introduce bias, as aggregate identification can be highly subjective. Herein, a protocol consisting of worm culturing, image acquisition, and data processing was standardized to support high-throughput aggregate quantification using C. elegans that express intestine-specific polyQ. By implementing a C. elegans-based image processing pipeline using CellProfiler, an image analysis software, this method has been optimized to separate and identify individual worms and enumerate their respective aggregates. Though the concept of automation is not entirely unique, the need to standardize such procedures for reproducibility, elimination of bias from manual counting, and increase throughput is high. It is anticipated that these methods can drastically simplify the screening process of large bacterial, genomic, or drug libraries using the C. elegans model.

Automating Aggregate Quantification in Caenorhabditis elegans

The following protocol describes the development and optimization of a high-throughput workflow for worm culturing, fluorescence imaging, and automated image processing to quantify polyglutamine aggregates as an assessment of changes in proteostasis.

A rise in the prevalence of neurodegenerative protein conformational diseases (PCDs) has fostered a great interest in this subject over the years. This ...

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

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Isolation and In vitro Activation of Caenorhabditis elegans Sperm

Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

Basic Caenorhabditis elegans Methods: Synchronization and Observation

Prostaglandin Extraction and Analysis in Caenorhabditis elegans

Using Caenorhabditis elegans as a Model System to Study Protein Homeostasis in a Multicellular Organism

Cultivation of Caenorhabditis elegans in Three Dimensions in the Laboratory

Combined Nucleotide and Protein Extractions in Caenorhabditis elegans

Measurement of Oxygen Consumption Rates in Intact Caenorhabditis elegans

Modeling Age-Associated Neurodegenerative Diseases in Caenorhabditis elegans

Automating Aggregate Quantification in Caenorhabditis elegans