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

Summary

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.

Abstract

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.

Introduction

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 β 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/NIX^9,10. 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 (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 markers^14,15,16,17 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 Förster 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 quenching^15,16.

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 pH^16,17. 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 (UPR^mt)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.

Protocol

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

1. 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 stock^25,26, 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.
   1. 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 µm vacuum filter/storage system.
   2. Make 0.1 M of calcium chloride (CaCl₂) and magnesium sulfate (MgSO₄), and sterilize them with a 0.2 µm syringe filter.
   3. Prepare 5 mg/mL cholesterol in absolute ethanol.
      NOTE: Since the cholesterol is prepared in ethanol, do not filter it.
   4. 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 °C.
   5. 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 MgSO_4, and 1 mL of 5 mg/mL cholesterol. Mix well after every addition.
   6. Add 4 mL of the melted NGM-agar to each 35 mm plate. Leave the dishes overnight to solidify at room temperature (RT, ~21 °C).
   7. 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 °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 °C overnight to obtain single colonies.
   8. 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.
   9. 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 °C and 250 rpm to an optical density (OD₆₀₀) of approximately 0.6.
2. Preparing the vehicle and experimental plates
   1. Add 100 µL of OP50 bacteria to the center of each 35 mm NGM-agar plate. Dry overnight at RT (room temperature, 21 °C).
   2. 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 µm 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 µm syringe filter.
   3. Add 25 µL of vehicle (negative control), FCCP (positive control; 5 µM), or VL-850 (experimental treatment; 62.5 µM) to separately seeded NGM plates on the bacterial lawn.
   4. Cover the plates with aluminum foil, and leave them to dry at RT (room temperature, 21 °C). Use the plates after ~16 h.
3. Obtaining synchronized young adult C. elegans hermaphrodites
   1. Make 1 L of M9 buffer with 22 mM KH₂PO₄, 42 mM Na₂HPO_4, and 86 mM NaCl. Sterilize by autoclaving, and let it cool. Once cooled, add 1 mL of 1 mM MgSO₄ (0.22 µm filter-sterilized).
   2. 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).
   3. 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.
   4. Sediment the worms by centrifugation at 500 × g for 1 min, and discard the supernatant until a volume of 1 mL remains.
   5. 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.
   6. When approximately 50% of the worms are broken, add 5 mL of M9 buffer and immediately sediment the eggs by centrifuging for 1 min at 500 × g.
   7. Remove the supernatant carefully without disturbing the pellet. Add 5 mL of M9 buffer, and repeat the washing procedure 2x.
   8. Remove the supernatant until 2 mL remains, and rotate the tube (360° rotation) at 20 rpm for ~16 h (RT, 21 °C) to obtain synchronized L1 larvae. From this tube, take a 5 µL 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 (µL). Based on these calculations, add ~200 larvae per NGM plate seeded with OP50 bacteria.
   9. Grow the L1 larvae at 21 °C for ~48 h until the young adult stage.
4. Drug treatment and microscopy procedure
   1. Put 100 worms on each of the experimental or control plates. Ensure that the negative, positive, and experimental plates contain the vehicle, 5 µM FCCP, and 62.5 µM VL-850, respectively. Incubate at 21 °C for 6 h.
   2. 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.
   3. Add 200 µL of M9 buffer, which contains 0.1% v/v Poloxamer 188, 0.1% v/v Pluronic F127, and 2 µL of the staining kit reagent, to the worm pellet. Let the mixture rotate at 20 rpm (360° rotation) for 1 h at RT (room temperature, 21 °C). To protect the dyes from light, cover the tubes with aluminum foil.
   4. 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.
      ​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.
   5. 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 µL of M9.
   6. 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 °C, and using a 1 mL tip, put 100 µL of the melted agarose onto a Deckgläser 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.
   7. Transfer the worms onto the agarose pad with a Pasteur glass pipette (i.e., the entire amount in the tube, ~10 µL). 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.
   8. Use a confocal microscope to image the worms within 24 h at the appropriate wavelengths (see below) using a 60x magnification lens.
      1. Place the slide onto the microscope stage.
      2. 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.
      3. Under Ti2 Full Pad, select 60x.
      4. 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 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.
5. Image analysis
   1. 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.
   2. Generate duplicates of these images to keep the original image untouched by clicking on Image | Duplicate or using the keyboard shortcut Shift + D.
   3. 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'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.
   4. 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.
   5. 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.
   6. 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.
   7. 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).
   8. To analyze the area of the colocalized mitochondria and lysosomes, select Analyze | Analyze Particles, and measure the summation of the puncta between 0.1625 µm² and 4 µm².

2. The Hep-3B cancer cell protocol

1. Preparing the drug stock solution
   1. 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 µm syringe filter. Then, prepare the vehicle as described before, but without the drug (in this case, VL-850).
2. Culturing Hep-3B cells for the experiment, drug treatment, and microscopy procedure
   1. Grow Hep-3B cells in 10 cm tissue culture plates containing Dulbecco'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 °C and 5% CO₂.
   2. 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 °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 × g for 5 min, remove the supernatant, and resuspend the cells in 5 mL of complete DMEM.
   3. Determine the cell concentration. Mix 50 µL of the cell suspension with 50 µL 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.
   4. Seed 42,000 Hep3G cells in each 8-well µ-slide in 400 µL of complete DMEM (as described above). Incubate the cells for 24 h at 37 °C and 5% CO₂.
   5. After 24 h at ~80%-85% confluency, remove 250 µL of medium from each of the wells, and add 50 µL of medium with the appropriate treatment or vehicle. To follow this protocol, treat the cells with 100 µM VL-850, 5 µM FCCP, and a vehicle as a control.
   6. After the 6 h incubation with the compounds, add 50 µL of medium to each well containing the staining reagent (0.5 µL of dye for 250 µL in each well). Incubate the cells with the dye for 30 min at 37 °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).
   7. Using a 200 µL pipette, gently remove all the medium (250 µL) from each well, and then wash the cells with 200 µL of prewarmed PBS.
   8. Fix the cells with 200 µL of fixing solution containing 4% formaldehyde and 2.5% glutaraldehyde prepared in PBS for 15 min at RT.
   9. Decant the fixative solution, and wash briefly with 200 µL of PBS.
   10. Add 200 µL of PBS, keep the cells covered and protected from light at 4 °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.
3. Image analysis
   1. 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.
   2. 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 µm² and 4 µm². To measure the colocalized puncta, divide the area of the colocalized mitochondria and lysosomes by the total cell area.

Results

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 µM VL-850, 5 µM FCCP (positive control), and ve...

Discussion

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 enzymes^12,13. 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...

Materials

List of materials used in this article

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