protocol for measuring mitophagy in caenorhabditis elegans

Quantifying Mitophagy in &lt;em&gt;C. elegans&lt;/em&gt; and a Liver Cancer Cell Line

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.

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

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

Degradation of damaged mitochondria by mitophagy is essential for the function of the mitochondrial network. Unfortunately, this process declines with age. We develop and use drugs that activate mitophagy, thereby promoting healthy aging to understand the function of mitophagy in aging and age-related diseases, including Alzheimer's.One of the biggest challenges is to measure the mitophagy process without affecting it. Recent studies have shown that GFP expression in the mitochondria of muscle cells induces stress that is MT UPR. Therefore, using dyes in non-transgenic animals or cells is essential and complements genetic technologies.Our protocol has three main advantages. First, there is no need to generate transgenic animals or cells expressing mitophagy biosensors. Second, no need for expensive infrastructure and expertise such as in case of electron microscopy.And third, the organelle dye cocktail is commercially available and the protocol is simple and fast. The new mitophagy-activating compound VL-850 induces robust mitophagy, and in this way protects from oxidative stress and promote lifespan. Now the big question is to determine the underlying mechanism of action.We explored the mechanism at multiple levels, from the cellular to the organismal using C.elegans and mammalian cells.

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Protocol for Measuring Mitophagy in Caenorhabditis elegans  

Protocol for Measuring Mitophagy in Caenorhabditis elegans

To begin, take the synchronized young adult C.Elegans hermaphrodites in a micro centrifuge tube. Add 200 microliters of M9 buffer, containing Poloxamer 188, Pluronic F127, and two microliters of the staining kit reagent to the worm pellet. Cover the tubes with aluminum foil to protect the dyes from light.Let the mixture rotate at 20 RPM for one hour at room temperature. Then spin the worms at 500 G for one minute. After that, remove the staining solution without disturbing the worm pellet.Wash the worms with M9 buffer thrice and transfer them into a seeded NGM agar plate containing the appropriate treatment. Wash the worms off the plates into fresh micro centrifuge tubes with one milliliter of M9 buffer. Afterward, fix the worms with 1%formaldehyde on ice for 30 minutes.And wash the worms with one milliliter of M9 buffer thrice to remove any residual formaldehyde. After pelleting the worms, aspirate the maximum amount of supernatant, keeping the pellet intact in 10 microliters of M9.Next, to prepare 2.5%agarose, weigh 0.125 grams of agarose into a 10 milliliter borosilicate glass test tube. Add five milliliters 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 degrees Celsius. Add 100 microliters of the melted agarose onto a deck glosser microscope cover glass using a one milliliter tip. Immediately, put another slide perpendicularly on the agarose drop forming a cross shape.After two minutes, gently separate the slides by pushing the upper cover glass leaving the agarose pad on the bottom cover slide. Transfer the worms onto the agarose pad with a Pasteur glass pipette. Use a wick made of a laboratory wipe to remove excess liquid.Then cover the worms with a smaller cover slip and apply transparent nail polish to the periphery to prevent evaporation. Place the slide in a dark box to protect it from light. Use a confocal microscope to image the worms within 24 hours at the appropriate wavelengths using a 60 times magnification lens.Next, open the imaging software and right click on the gray area. From the popup that appears, select acquisition. Ti2 full pad.ND acquisition, and lookup tables. Under Ti2 full pad, select 60 times and adjust the microscope fine focus knob to bring the worms into focus. Then select spinning disc and choose the 16 bit-no binning option.For each fluorescence filter, set the exposure time to 500 milliseconds and 20 milliseconds for bright field. Once these parameters are set, select run now and wait for the output image to be generated as an ND2 file.

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142 ビュー.  The Hebrew University of Jerusalem. はじめに、同期した若い成人のC.Elegans雌雄同体をマイクロ遠心管に入れます。ポロクサマー188、プルロニックF127、および2マイクロリットルの染色キット試薬を含む200マイクロリットルのM9バッファーをワームペレットに追加します。染料を光から保護するために、チューブをアルミホイルで覆います。混合物を室温で20 RPMで1時間回転させます。次に、ワームを500 G...