This work was supported in part by a MOST 111-2311-B-001-019-MY3 research grant from the National Science and Technology Council in Taiwan.

1. Preparation of cells expressing diKillerRed or self-labeling proteins (SLPs) in the peroxisome lumen
<ol>
	<li>Seed the desired cells on glass-bottomed cell culture dishes. For the experiment here, seed 2 x 105 human SHSY5Y cells in 840 &#181;L of culture medium (DMEM/F-12 supplemented with 10% fetal bovine serum and 1% penicillin/streptomycin) or 6 x 104 mouse NIH3T3 cells in 840 &#181;L of culture medium (DMEM supplemented with 10% bovine serum and 1% penicillin/streptomycin) on a 35 mm culture dish with a 20 mm diameter glass microwell. 
	NOTE: The number of cells seeded should be adjusted proportionally according to the glass microwell area when using glass bottom dishes of other specifications.</li>
	<li>Grow the cells under 5% CO2/37 &#176;C for 24 h.</li>
	<li>Transfect the cells with desired plasmids using a transfection reagent.
	<ol>
		<li>Use PMP34-TagBFP to label the peroxisomes in live cells. Use either peroxisome-targeting diKillerRed-VKSKL or SLP-VKSKL (+ dye-labeled SLP ligands) for ROS generation in the peroxisome lumen (through light-activated ROS production). In this protocol, we use the self-labeling protein HaloTag-VKSKL21. To monitor the translocation of Stub1/Hsp70, the accumulation of ubiquitinated proteins, and the formation of autophagosomes on ROS-stressed peroxisomes, transfect EGFP-Stub1, EGFP-Hsp70, EGFP-Ub, or EGFP-LC3B, respectively. To check ROS generation in the peroxisomal lumen, co-transfect diKillerRed-VKSKL with the peroxisome-localized fluorescent redox probe roGFP2-VKSKL22.</li>
		<li>For SHSY5Y cell transfection, dilute plasmids in 55 &#181;L of serum-free DMEM/F-12, and dilute 1 &#181;L of transfection reagent in 55 &#181;L of serum-free DMEM/F12. Combine the diluted DNA with the diluted reagent. Mix gently, and incubate for 30 min at room temperature.</li>
		<li>Add the complex to the 20 mm microwell previously plated with 100 &#181;L of culture medium for SHSY5Y without antibiotics (DMEM/F-12 with 10% fetal bovine serum). Gently shake the dish back and forth.</li>
		<li>For NIH3T3 cell transfection, substitute reduced serum medium for the serum-free DMEM/F-12 for diluting the plasmids and transfection reagent. Replace the plating culture medium with SHSY5Y with culture medium for NIH3T3 cells without antibiotics (DMEM with 10% bovine serum).</li>
	</ol>
	</li>
	<li>After 2 h of transfection, remove the transfection reagent-containing medium, and add 1 mL of fresh culture medium. Incubate the NIH3T3 or SHSY5Y cells at 37&#176;C and 5% CO2 for 24 h. 
	NOTE: Other cell lines can also be used to study Stub1-mediated pexophagy (e.g., HeLa cells).</li>
</ol>
2. Staining peroxisomes with dye-labeled SLP ligands (for light-activated ROS production)
<ol>
	<li>At 24 h post-transfection, remove the culture medium, and add 100 &#181;L of 200 nM HaloTag TMR ligand or 200 nM Janelia Fluor 646 HaloTag ligand onto the 20 mm glass microwell. 
	NOTE: The dye-labeled SLP ligands should be diluted with the respective culture medium for each cell line. Choosing Janelia Fluor 646-labeled ligands will free up the blue, green, and red channels for downstream fluorescence imaging.</li>
	<li>Transfer the dish to a 37&#176;C, 5% CO2 incubator. Stain for 1 h. After 1 h, remove the staining medium thoroughly, and add 1 mL of fresh culture medium.</li>
</ol>
3. ROS-stressing of peroxisomes on a laser-scanning confocal microscope
<ol>
	<li>Place a glass-bottomed dish containing the desired cells on a laser-scanning confocal microscope equipped with a stage-top incubator.</li>
	<li>Click on Tool Window, choose Ocular, and click on the DIA button (Figure 2A) to turn on the transmitted light. View the dish through the microscope eyepiece, and bring the cells into focus by turning the focus knob.</li>
	<li>Choose LSM, and click on the Dye &#38; Detector Select button (Figure 2B). Double-click on the desired dyes in the pop-up window to select the appropriate laser and filter settings for imaging the cells (Figure 2C). Click on Livex4 in the Live panel to initiate fast laser scanning to start screening for the fluorescence signals of the cells (Figure 2D).</li>
	<li>Move the stage around using the joystick controller, and locate the medium diKillerRed-VKSKL- or SLP-VKSKL-expressing cells to apply ROS stress (on the peroxisomes). Acquire an image of the desired cell.</li>
	<li>Click on Tool Window, and choose LSM Stimulation (Figure 2E). Click on the ellipse button, and in the live image window, use the mouse to draw a circular ROI of 15 &#181;m in diameter (Figure 2F) containing the desired peroxisomes acquired in step 3.4 to which ROS stress will be applied (see Figure 3 for an example).</li>
	<li>To apply ROS stress, use 561 nm for cells with the diKillerRed-PTS1- or TMR-labeled SLP ligand, and use 640 nm for cells stained with the Janelia Fluor 646-labeled SLP ligand.</li>
	<li>Select the laser wavelength for applying the ROS stress. Enter the desired laser percentage and the duration (Figure 2G).</li>
	<li>Click on Acquire, and click on the Stimulation button (Figure 2H) to initiate the scanning with 561/640 nm light through the ROIs for 30 s each. This corresponds to a ~5% laser power setting for both 561 nm and 640 nm on the confocal microscope used here. This operation will lead to ROS production in the peroxisomes. 
	NOTE: One can select ROIs of different sizes. One should proportionally adjust the illumination time for each ROI according to its area.</li>
	<li>After 30 s of laser illumination, peroxisomes within the ROIs will have lost their diKillerRed-PTS1/TMR/Janelia Fluor 646 signals. This signal loss allows one to distinguish the illuminated from the non-illuminated peroxisomes (damaged vs. healthy) within a cell.</li>
</ol>
4. Monitoring pexophagy in live cells by time-lapse imaging
<ol>
	<li>Follow the translocation of EGFP-tagged Stub1, Hsp70, Ub, p62, or LC3B onto the illuminated/ROS-stressed peroxisomes by time-lapse imaging using 10 min intervals between frames (for examples, see Figures 3-7). 
	NOTE: EGFP-Stub1 or EGFP-Hsp70 signals start to appear 10-20 min post illumination and stay on all the damaged peroxisomes until ~40 min post illumination. The EGFP-Ub signals appear 30 min post illumination and last for 2-3 h. EGFP-p62 translocation appears 60 min post illumination, and EGFP-LC3B signals become visible ~3 h after illumination.</li>
	<li>Quantify the EGFP signals on the damaged peroxisomes (from Stub1, Ub, p62, or LC3B) using ImageJ. Perform the following steps for using ImageJ to quantify a three-channel image (PMP34-TagBFP/EGFP-LC3B/HaloTag-VKSKL).
	<ol>
		<li>Open the image, and apply Elliptical selections or Freehand selections to enclose the region containing all the illuminated peroxisomes (PMP34-TagBFP signal positive and HaloTag ligand signal bleached; Figure 8A).</li>
		<li>Click on Duplicate to pick out the PMP34-TagBFP channel (Figure 8A). Set a threshold using the drop-down menu Image &#62; Adjust &#62; Threshold to mark the peroxisomes (Figure 8B).</li>
		<li>Select Analyze &#62; Analyze particles to determine the exact regions of the illuminated peroxisomes. ImageJ records these regions of interest (ROIs) in the ROI manager (Figure 8C).</li>
		<li>Click on Duplicate to pick out the EGFP channel for analyzing the fluorescence signal from EGFP-conjugated Ub, p62, or LC3B (Figure 8D). Select all the ROIs in the ROI Manager (Figure 8E).</li>
		<li>Apply Measure in the ROI Manager to measure the EGFP signals on the peroxisomes (Figure 8E). Find the value of the area and integrated density (the sum of all the pixel intensities) for each ROI in the Results table (Figure 8E).</li>
		<li>To obtain the average EGFP fluorescence intensities at different time points after illumination, divide the integrated EGFP intensity by the total area of illuminated peroxisomes (PMP34-TagBFP signal positive and HaloTag ligand signal negative areas). To obtain the accumulated signal on the peroxisomes, subtract the averaged intensity at a time point by the averaged intensity at time = 0, and then normalize by the averaged intensity at time = 0.</li>
	</ol>
	</li>
</ol>
5. Globally damaging all the peroxisomes (in every cell) on a culture dish
<ol>
	<li>Perform step 1 (preparation of cells expressing peroxisome-targeted diKillerRed [diKillerRed-VKSKL] or SLP [SLP-VKSKL]) and step 2 (staining peroxisomes with the SLP ligand [for light-activated ROS production]).</li>
	<li>After 1 h of TMR-labeled ligand staining, remove the staining medium, and add 1 mL of culture medium containing 30 mM 3-AT (3-amino-1,2,4-triazole; a catalase inhibitor). 
	NOTE: Catalase is a ROS scavenger expressed in the peroxisome lumen, so 3-AT treatment helps to augment the light-elicited oxidative damage on the peroxisomes.</li>
	<li>Place the culture dish above an LED (555-570 nm). Illuminate all the cells with a power density of 1.8 W/cm2 for 9 h in an incubator. 
	NOTE: To perform this step outside an incubator, place the culture dish on a 37&#176;C heating plate. Add 20 mM HEPES to the 3-AT-containing medium, and cover the culture dish with a transparent film to minimize gas exchange. HEPES is a buffering agent that maintains physiological pH in cell culture outside a CO2 incubator during LED illumination.</li>
	<li>Validate the success of LED-elicited damage by imaging the EGFP-Ub, EGFP-p62, or EGFP-LC3B signals on the peroxisomes. Using the illumination condition outlined above, 82% of SHSY5Y cells stably expressing HaloTag-VKSKL displayed Stub1-mediated pexophagy.</li>
	<li>Harvest the cells for downstream biochemical analysis.</li>
</ol>

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The authors declare no competing financial interests.

This protocol details how to trigger Stub1-mediated pexophagy within cell cultures by elevating peroxisomal ROS levels with light. As the protocol relies on dye-assisted ROS generation, one needs to ensure sufficient expression of diKillerRed-VKSKL or dye-labeled SLP ligand staining within the cells of interest. Given that different cell types or cells of different genetic backgrounds can harbor peroxisomes with slightly different properties, one may need to tune the exact illumination conditions to ensure the induction ...

Peroxisomes are single-membrane-bound organelles present in most eukaryotic cells. Peroxisomes are a metabolic compartment essential for carrying out the beta-oxidation of very long-chain fatty acids, purine catabolism, and ether phospholipid and bile acid synthesis1. Peroxisome-derived acetyl-CoA controls lipid homeostasis by regulating the central signaling in metabolism2. Therefore, it is no surprise that compromised peroxisomal functions are implied in various diseases, including neurodegenerative disorders, aging, cancers, obesity, and diabetes3,4,5. An essential process in the maintenance of peroxisomal operation is pexophagy. Pexophagy is a catabolic process for the selective turnover of peroxisomes by autophagy. Cells use pexophagy to help control the quantity and quality of peroxisomes, thereby ensuring proper peroxisomal function. A recent study demonstrated that peroxisome loss caused by mutations in PEX1 peroxisomal biogenesis factors 1 and 6 results from uncontrolled pexophagy6. Notably, 65% of all peroxisome biogenesis disorder (PBD) patients harbor deficiencies in the peroxisomal AAA ATPase complex, composed of PEX1, PEX6, and PEX26 in mammalian cells7.
A number of methods can be used to initiate and study pexophagy. In yeast, pexophagy is triggered when the supplied nutrients are switched from peroxisome-dependent carbon sources to peroxisome-independent carbon sources (to lower cellular peroxisome numbers)8. For example, the transfer of methanol-grown Pichia pastoris cells from methanol medium to glucose medium and ethanol medium induces micropexophagy and macropexophagy, respectively8,9,10. Micropexophagy sequesters clustered peroxisomes for degradation by remodeling the vacuole to form cup-like vacuolar sequestration membranes and a lid-like structure termed the micropexophagy-specific membrane apparatus (MIPA). In macropexophagy, individual peroxisomes are engulfed by double-membrane structures known as pexophagosomes, followed by fusion with the vacuole for degradation8,9,10. The phosphorylation of pexophagy receptors, such as Atg36p in Saccharomyces cerevisiae and Atg30p in Pichia pastoris, is critical for the receptors to recruit core autophagy machinery and to facilitate peroxisomal targeting to autophagosomes8,11.
In mammalian cells, pexophagy can be induced by ubiquitination. Tagging the peroxisomal membrane proteins PMP34 or PEX3 with ubiquitin on the cytosolic side induces pexophagy12. The overexpression of PEX3 induces peroxisome ubiquitination and peroxisome elimination by lysosomes13. In addition, the fusion of PEX5 with a C-terminal EGFP impairs the export of monoubiquitinated PEX5 and results in pexophagy14. On the other hand, pexophagy can also be triggered by H2O2 treatment. Peroxisomes produce reactive oxygen species (ROS); specifically, the peroxisomal enzyme Acox1, which catalyzes the initial step of beta-oxidation of very long-chain fatty acids (&#62; 22 carbon), produces not only acetyl-CoA but also peroxisomal ROS. In response to the heightened ROS levels under H2O2 treatment, mammalian cells activate pexophagy to lower the ROS production and alleviate stress. It has been reported that H2O2 treatment drives the recruitment of ataxia-telangiectasia mutated (ATM) to peroxisomes. ATM then phosphorylates PEX5 to promote peroxisomal turnover by pexophagy15.
Since peroxisomes are ROS-generating centers, they are also prone to ROS damage. ROS-elicited peroxisomal injuries force cells to activate pexophagy to initiate peroxisome quality control pathways (removal of the damaged peroxisome by autophagy). Here, we outline an approach for the on-demand triggering of ROS-elicited peroxisomal injury. The protocol takes advantage of light-activated ROS production within organelles16,17,18,19,20 (Figure 1). Dye-labeled peroxisomes are illuminated, leading to ROS production within the peroxisomal lumen, which specifically triggers peroxisomal injury. Using this protocol, it is shown that ROS-stressed peroxisomes are removed through a ubiquitin-dependent degradation pathway. ROS-stressed peroxisomes recruit the ubiquitin E3 ligase Stub1 to allow their engulfment into autophagosomes for individual removal by pexophagy16. One can use this protocol to compare the fate of injured and healthy peroxisomes within the same cell by time-lapse microscopy. The method can also be used to globally damage all the peroxisomes (in all cells) on a culture dish, allowing for the biochemical analysis of the pexophagy pathway.

<table><tbody><tr><td>35 mm culture dish with a 20 mm diameter glass microwell&amp;nbsp;</td><td>MatTek</td><td>P35G-1.5-20-C</td><td>20 mm glass bottomed</td></tr><tr><td>3-amino-1,2,4-triazole (3-AT)</td><td>Sigma Aldrich</td><td>A8056</td><td></td></tr><tr><td>bovine serum</td><td>ThermoFisher Scientific</td><td>16170060</td><td></td></tr><tr><td>Cell culture incubator</td><td>Nuaire</td><td>NU-4750</td><td></td></tr><tr><td>diKillerRed-PTS1</td><td>Academia Sinica</td><td></td><td>made by appending the KillerRed tandem dimer with PTS1(VKSKL)</td></tr><tr><td>Dulbecco&#039;s Modified Eagle Medium (DMEM)</td><td>ThermoFisher Scientific</td><td>11965092</td><td></td></tr><tr><td>Dulbecco&#039;s Modified Eagle Medium/Nutrient Mixture F-12 (DMEM/F-12)</td><td>ThermoFisher Scientific</td><td>11330032</td><td></td></tr><tr><td>EGFP-C1</td><td>Clontech</td><td>pEGFP-C1</td><td>The backbone of EGFP-C1 was used for cloning EGFP-Stub1, EGFP-Hsp70, EGFP-p62</td></tr><tr><td>EGFP-Hsp70</td><td>Academia Sinica</td><td></td><td>Hsp70 gene (HSPA1A) PCR amplified from HeLa cDNA and cloned into EGFP-C1</td></tr><tr><td>EGFP-LC3B</td><td>Addgene</td><td>11546</td><td></td></tr><tr><td>EGFP-p62</td><td>Academia Sinica</td><td></td><td>generated by inserting the human SQSTM1 gene (through PCR amplification of the HeLa cell cDNA) into EGFP-C1</td></tr><tr><td>EGFP-Stub1</td><td>Academia Sinica</td><td></td><td>generated by inserting the mouse Stub1 gene (through PCR amplification of the total mouse kidney cDNA) into EGFP-C1</td></tr><tr><td>EGFP-Ub</td><td>Addgene</td><td>11928</td><td></td></tr><tr><td>fetal bovine serum</td><td>ThermoFisher Scientific</td><td>10437028</td><td></td></tr><tr><td>HaloTag TMR ligand&amp;nbsp;</td><td>Promega</td><td>G8252</td><td></td></tr><tr><td>HaloTag-PTS1</td><td>Academia Sinica</td><td></td><td>PTS1 appended and cloned into EGFP-C1 backbone</td></tr><tr><td>HEPES</td><td>ThermoFisher Scientific</td><td>15630080</td><td></td></tr><tr><td>Inverted Confocal Microscope&amp;nbsp;</td><td>Olympus</td><td>FV3000RS</td><td>405 nm Ex, 488 nm Ex, 561 nm Ex,&amp;nbsp; microscope with a TOKAI HIT chamber incubator and the UNIV2-D35 dish attachment</td></tr><tr><td>Janelia Fluor 646 HaloTag Ligand</td><td>Promega</td><td>GA1120</td><td></td></tr><tr><td>LED</td><td>VitaStar</td><td>PAR64</td><td>80 W, 555-570 nm</td></tr><tr><td>lipofectamine 2000&amp;nbsp;</td><td>ThermoFisher Scientific</td><td>11668</td><td>transfection reagent</td></tr><tr><td>NIH3T3 cell</td><td>ATCC</td><td>CRL-1658</td><td>adherent</td></tr><tr><td>Opti-MEM</td><td>ThermoFisher Scientific</td><td>319850</td><td>reduced serum media</td></tr><tr><td>penicillin/streptomycin</td><td>ThermoFisher Scientific</td><td>15140</td><td></td></tr><tr><td>PMP34-TagBFP</td><td>Academia Sinica</td><td></td><td>PMP34 PCR amplified from HeLa cDNA and cloned intoTagBFP-C (Evrogen FP171)</td></tr><tr><td>roGFP2-PTS1</td><td>Academia Sinica</td><td></td><td>generated by appending eroGFP (taken from Addgene plasmid 20131) with the amino acid sequence VKSKL, and cloned into the EGFP-C1</td></tr><tr><td>SHSY5Y cell</td><td>ATCC</td><td>CRL-2266</td><td>adherent</td></tr></tbody></table>

monitoring stub1-mediated pexophagy

Mammalian cells can turn over peroxisomes through Stub1-mediated pexophagy. The pathway potentially permits cellular control of the quantity and quality of peroxisomes. During this process, heat shock protein 70 and the ubiquitin E3 ligase, Stub1, translocate onto peroxisomes to be turned over to initiate pexophagy. The Stub1 ligase activity allows the accumulation of ubiquitin and other autophagy-related modules on targeted peroxisomes. Elevating reactive oxygen species (ROS) levels within the peroxisomal lumen can activate Stub1-mediated pexophagy. One can, therefore, use dye-assisted ROS generation to trigger and monitor this pathway. This article outlines the procedures for using two classes of dyes, fluorescent proteins and synthetic fluorophores, to initiate pexophagy within mammalian cell cultures. These dye-assisted ROS generation-based protocols can not only be used to target all the peroxisomes within a cell population globally but can also permit the manipulation of individual peroxisomes within single cells. We also describe how Stub1-mediated pexophagy can be followed using live-cell microscopy.

The Stub1-mediated pexophagy induction scheme shown here takes advantage of dye-assisted ROS generation within the peroxisome lumen. This operation requires minimal light intensities. Peroxisomes containing fluorescent proteins or dyes can, therefore, be illuminated using standard laser-scanning confocal microscopes. Focal illumination leads to instantaneous and localized ROS production within individual peroxisomes, as indicated by the fluorescent reporter roGFP2-VKSKL (Figure 9).&#160;We d...

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Monitoring Stub1-Mediated Pexophagy

This protocol provides instructions for triggering and monitoring Stub1-mediated pexophagy in live cells.

Mammalian cells can turn over peroxisomes through Stub1-mediated pexophagy. The pathway potentially permits cellular control of the quantity and quality ...

Research

JoVE Journal

Biology

1.5K Views.  Academia Sinica. Peroxisomen voeren bèta-oxidatie uit en zijn gevoelig voor ROS-schade. Deze methode maakt het mogelijk om te onderzoeken hoe cellen omgaan met ROS-gestresste peroxisomen. Dit protocol wordt niet alleen gebruikt om zich wereldwijd te richten op alle peroxisomen binnen een celpopulatie, maar kan ook de manipulatie van de individuele peroxisomen binnen afzonderlijke cellen mogelijk maken.Kleurstof-geassisteerde ROS-generatie kan ook worden gebruikt om organellen buiten peroxisomen te ric...

Video: Monitoring Stub1-Mediated Pexophagy

Exploring the Regulation of Lipid Droplet Catabolism through Lipophagy

Macroautophagy, commonly referred to as autophagy, is a highly conserved cellular process responsible for the degradation of cellular components. This process is particularly prominent under conditions such as fasting, cellular stress, organelle damage, cellular damage, or aging of cellular components. During autophagy, a segment of the cytoplasm is enclosed within double-membrane vesicles known as autophagosomes, which then fuse with lysosomes. Following this fusion, the contents of autophagosomes undergo non-selective bulk degradation facilitated by lysosomes. However, autophagy also exhibits selective functionality, targeting specific organelles, including mitochondria, peroxisomes, lysosomes, nuclei, and lipid droplets (LDs). Lipid droplets are enclosed by a phospholipid monolayer that isolates neutral lipids from the cytoplasm, protecting cells from the harmful effects of excess sterols and free fatty acids (FFAs). Autophagy is implicated in various conditions, including neurodegenerative diseases, metabolic disorders, and cancer. Specifically, lipophagy -- the autophagy-dependent degradation of lipid droplets -- plays a crucial role in regulating intracellular FFA levels across different metabolic states. This regulation supports essential processes such as membrane synthesis, signaling molecule formation, and energy balance. Consequently, impaired lipophagy increases cellular vulnerability to death stimuli and contributes to the development of diseases such as cancer. Despite its significance, the precise mechanisms governing lipid droplet metabolism regulated by lipophagy in cancer cells remain poorly understood. This article aims to describe confocal imaging acquisition and quantitative imaging analysis protocols that enable the investigation of lipophagy associated with metabolic changes in cancer cells. The results obtained through these protocols may shed light on the intricate interplay between autophagy, lipid metabolism, and cancer progression. By elucidating these mechanisms, novel therapeutic targets may emerge for combating cancer and other metabolic-related diseases.

Lipophagy is a selective form of autophagy that involves the degradation of lipid droplets. Dysfunctions in this process are associated with cancer development. However, the precise mechanisms are not yet fully understood. This protocol describes quantitative imaging approaches to better understand the interplay between autophagy, lipid metabolism, and cancer progression.

Macroautophagy, commonly referred to as autophagy, is a highly conserved cellular process responsible for the degradation of cellular components. This ...

Cancer Research

Assessing LC3-II-Positive EV Release in Autophagy with Impaired Lysosome Fusion

Evaluation of LC3-II Release via Extracellular Vesicles in Relation to the Accumulation of Intracellular LC3-positive Vesicles

(Macro)autophagy represents a fundamental cellular degradation pathway. In this process, double-membraned vesicles known as autophagosomes engulf cytoplasmic contents, subsequently fusing with lysosomes for degradation. Beyond the canonical role, autophagy-related genes also modulate a secretory pathway involving the release of inflammatory molecules, tissue repair factors, and extracellular vesicles (EVs). Notably, the process of disseminating pathological proteins between cells, particularly in neurodegenerative diseases affecting the brain and spinal cord, underscores the significance of understanding this phenomenon. Recent research suggests that the transactive response DNA-binding protein 43 kDa (TDP-43), a key player in amyotrophic lateral sclerosis and frontotemporal lobar degeneration, is released in an autophagy-dependent manner via EVs enriched with the autophagosome marker microtubule-associated proteins 1A/1B light chain 3B-II (LC3-II), especially when autophagosome-lysosome fusion is inhibited.
To elucidate the mechanism underlying the formation and release of LC3-II-positive EVs, it is imperative to establish an accessible and reproducible method for evaluating both intracellular and extracellular LC3-II-positive vesicles. This study presents a detailed protocol for assessing LC3-II levels via immunoblotting in cellular and EV fractions obtained through differential centrifugation. Bafilomycin A1 (Baf), an inhibitor of autophagosome-lysosome fusion, serves as a positive control to enhance the levels of intracellular and extracellular LC3-II-positive vesicles. Tumor susceptibility gene 101 (TSG101) is used as a marker for multivesicular bodies. Applying this protocol, it is demonstrated that siRNA-mediated knockdown of syntaxin-6 (STX6), a genetic risk factor for sporadic Creutzfeldt-Jakob disease, augments LC3-II levels in the EV fraction of cells treated with Baf while showing no significant effect on TSG101 levels. These findings suggest that STX6 may negatively regulate the extracellular release of LC3-II via EVs, particularly under conditions where autophagosome-lysosome fusion is impaired. Combined with established methods for evaluating autophagy, this protocol provides valuable insights into the role of specific molecules in the formation and release of LC3-II-positive EVs.

Here, we present the methodology for concisely assessing autophagosome marker LC3-II levels in extracellular vesicles (EVs) by immunoblotting. Analysis for LC3-II levels in EVs, autolysosome formation, and omegasome formation suggests the new role of STX6 in the release of LC3-II-positive EVs when autophagosome-lysosome fusion is inhibited.

(Macro)autophagy represents a fundamental cellular degradation pathway. In this process, double-membraned vesicles known as autophagosomes engulf ...

Biochemistry

Monitoring of Ubiquitin-proteasome Activity in Living Cells Using a Degron (dgn)-destabilized Green Fluorescent Protein (GFP)-based Reporter Protein

Proteasome is the main intracellular organelle involved in the proteolytic degradation of abnormal, misfolded, damaged or oxidized proteins 1, 2. Maintenance of proteasome activity was implicated in many key cellular processes, like cell's stress response 3, cell cycle regulation and cellular differentiation 4 or in immune system response 5. The dysfunction of the ubiquitin-proteasome system has been related to the development of tumors and neurodegenerative diseases 4, 6. Additionally, a decrease in proteasome activity was found as a feature of cellular senescence and organismal aging 7, 8, 9, 10. Here, we present a method to measure ubiquitin-proteasome activity in living cells using a GFP-dgn fusion protein. To be able to monitor ubiquitin-proteasome activity in living primary cells, complementary DNA constructs coding for a green fluorescent protein (GFP)&#x2013;dgn fusion protein (GFP&#x2013;dgn, unstable) and a variant carrying a frameshift mutation (GFP&#x2013;dgnFS, stable 11) are inserted in lentiviral expression vectors. We prefer this technique over traditional transfection techniques because it guarantees a very high transfection efficiency independent of the cell type or the age of the donor. The difference between fluorescence displayed by the GFP&#x2013;dgnFS (stable) protein and the destabilized protein (GFP-dgn) in the absence or presence of proteasome inhibitor can be used to estimate ubiquitin-proteasome activity in each particular cell strain. These differences can be monitored by epifluorescence microscopy or can be measured by flow cytometry.

Monitoring of Ubiquitin-proteasome Activity in Living Cells Using a Degron dgn-destabilized Green Fluorescent Protein GFP-based Reporter Protein

A method to monitor ubiquitin-proteasome activity in living cells is described. A degron-destabilized GFP- (GFP-dgn) and a stable GFP-dgnFS fusion protein are generated and transduced into the cell using a lentiviral expression vector. This technique allows to generate a stable GFP-dgn/GFP-dgnFS expressing cell line in which ubiquitin-proteasome activity can be easily assessed using epifluorescence or flow cytometry.

Proteasome is the main intracellular organelle involved in the proteolytic degradation of abnormal, misfolded, damaged or oxidized proteins 1, 2. ...

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

Mitochondria are the powerhouses of cells and produce cellular energy in the form of ATP. Mitochondrial dysfunction contributes to biological aging and a wide variety of disorders including metabolic diseases, premature aging syndromes, and neurodegenerative diseases such as Alzheimer's disease (AD) and Parkinson's disease (PD). Maintenance of mitochondrial health depends on mitochondrial biogenesis and the efficient clearance of dysfunctional mitochondria through mitophagy. Experimental methods to accurately detect autophagy/mitophagy, especially in animal models, have been challenging to develop. Recent progress towards the understanding of the molecular mechanisms of mitophagy has enabled the development of novel mitophagy detection techniques. Here, we introduce several versatile techniques to monitor mitophagy in human cells, Caenorhabditis elegans (e.g., Rosella and DCT-1/ LGG-1 strains), and mice (mt-Keima). A combination of these mitophagy detection techniques, including cross-species evaluation, will improve the accuracy of mitophagy measurements and lead to a better understanding of the role of mitophagy in health and disease.

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

Mitophagy, the process of clearing damaged mitochondria, is necessary for mitochondrial homeostasis and health maintenance. This article presents some of the latest mitophagy detection methods in human cells, Caenorhabditis elegans, and mice.

Mitochondria are the powerhouses of cells and produce cellular energy in the form of ATP. Mitochondrial dysfunction contributes to biological aging and a ...

Medicine

Live Cell Imaging of Early Autophagy Events: Omegasomes and Beyond

Autophagy is a cellular response triggered by the lack of nutrients, especially the absence of amino acids. Autophagy is defined by the formation of double membrane structures, called autophagosomes, that sequester cytoplasm, long-lived proteins and protein aggregates, defective organelles, and even viruses or bacteria. Autophagosomes eventually fuse with lysosomes leading to bulk degradation of their content, with the produced nutrients being recycled back to the cytoplasm. Therefore, autophagy is crucial for cell homeostasis, and dysregulation of autophagy can lead to disease, most notably neurodegeneration, ageing and cancer.	
	Autophagosome formation is a very elaborate process, for which cells have allocated a specific group of proteins, called the core autophagy machinery. The core autophagy machinery is functionally complemented by additional proteins involved in diverse cellular processes, e.g. in membrane trafficking, in mitochondrial and lysosomal biology. Coordination of these proteins for the formation and degradation of autophagosomes constitutes the highly dynamic and sophisticated response of autophagy. Live cell imaging allows one to follow the molecular contribution of each autophagy-related protein down to the level of a single autophagosome formation event and in real time, therefore this technique offers a high temporal and spatial resolution.	
	Here we use a cell line stably expressing GFP-DFCP1, to establish a spatial and temporal context for our analysis. DFCP1 marks omegasomes, which are precursor structures leading to autophagosomes formation. A protein of interest (POI) can be marked with either a red or cyan fluorescent tag. Different organelles, like the ER, mitochondria and lysosomes, are all involved in different steps of autophagosome formation, and can be marked using a specific tracker dye. Time-lapse microscopy of autophagy in this experimental set up, allows information to be extracted about the fourth dimension, i.e. time. Hence we can follow the contribution of the POI to autophagy in space and time.

Time-lapse microscopy of fluorescently labeled autophagy markers allows monitoring of the dynamic autophagy response with high temporal resolution. Using specific autophagy and organelle markers in a combination of 3 different colors, we can follow the contribution of a protein to autophagosome formation in a robust spatial and temporal context.

Autophagy is a  cellular response triggered by the lack of nutrients, especially the absence of  amino acids. Autophagy is defined by the formation of ...

Visualization of Endogenous Mitophagy Complexes In Situ in Human Pancreatic Beta Cells Utilizing Proximity Ligation Assay

Mitophagy is an essential mitochondrial quality control pathway, which is crucial for pancreatic islet beta cell bioenergetics to fuel glucose-stimulated insulin release. Assessment of mitophagy is challenging and often requires genetic reporters or multiple complementary techniques not easily utilized in tissue samples, such as primary human pancreatic islets. Here we demonstrate a robust approach to visualize and quantify formation of key endogenous mitophagy complexes in primary human pancreatic islets. Utilizing the sensitive proximity ligation assay technique to detect interaction of the mitophagy regulators NRDP1 and USP8, we are able to specifically quantify formation of essential mitophagy complexes in situ. By coupling this approach to counterstaining for the transcription factor PDX1, we can quantify mitophagy complexes, and the factors that can impair mitophagy, specifically within beta cells. The methodology we describe overcomes the need for large quantities of cellular extracts required for other protein-protein interaction studies, such as immunoprecipitation (IP) or mass spectrometry, and is ideal for precious human islet&#160;samples generally not available in sufficient quantities for these approaches. Further, this methodology obviates the need for flow sorting techniques to purify beta cells from a heterogeneous islet population for downstream protein applications. Thus, we describe a valuable protocol for visualization of mitophagy highly compatible for use in heterogeneous and limited cell populations.

This protocol outlines a method for quantitative analysis of mitophagy protein complex formation specifically in beta cells from primary human islet samples. This technique thus allows analysis of mitophagy from limited biological material, which are crucial in precious human pancreatic beta cell samples.

Mitophagy is an essential mitochondrial quality control pathway, which is crucial for pancreatic islet beta cell bioenergetics to fuel glucose-stimulated ...

Analysis of Autophagy in Penicillium chrysogenum by Using Starvation Pads in Combination With Fluorescence Microscopy

The study of cellular quality control systems has emerged as a highly dynamic and relevant field of contemporary research. It has become clear that cells possess several lines of defense against damage to biologically relevant molecules like nucleic acids, lipids and proteins. In addition to organelle dynamics (fusion/fission/motility/inheritance) and tightly controlled protease activity, the degradation of surplus, damaged or compromised organelles by autophagy (cellular &#8216;self-eating&#8217;) has received much attention from the scientific community. The regulation of autophagy is quite complex and depends on genetic and environmental factors, many of which have so far not been elucidated. Here a novel method is presented that allows the convenient study of autophagy in the filamentous fungus Penicillium chrysogenum. It is based on growth of the fungus on so-called &#8216;starvation pads&#8217; for stimulation of autophagy in a reproducible manner. Samples are directly assayed by microscopy and evaluated for autophagy induction / progress. The protocol presented here is not limited for use with P. chrysogenum and can be easily adapted for use in other filamentous fungi.

Analysis of Autophagy in Penicillium chrysogenum by Using Starvation Pads in Combination With Fluorescence Microscopy

A convenient and powerful method for studying autophagy in Penicillium chrysogenum by using starvation pads (mixtures of agarose and tap water in a microscope slide containing a central cavity) is presented here.

The study of cellular quality control systems has emerged as a highly dynamic and relevant field of contemporary research. It has become clear that cells ...

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

Time-lapse video microscopy can be defined as the real time imaging of living cells. This technique relies on the collection of images at different time points. Time intervals can be set through a computer interface that controls the microscope-integrated camera. This kind of microscopy requires both the ability to acquire very rapid events and the signal generated by the observed cellular structure during these events. After the images have been collected, a movie of the entire experiment is assembled to show the dynamic of the molecular events of interest. Time-lapse video microscopy has a broad range of applications in the biomedical research field and is a powerful and unique tool for following the dynamics of the cellular events in real time. Through this technique, we can assess cellular events such as migration, division, signal transduction, growth, and death. Moreover, using fluorescent molecular probes we are able to mark specific molecules, such as DNA, RNA or proteins and follow them through their molecular pathways and functions. Time-lapse video microscopy has multiple advantages, the major one being the ability to collect data at the single-cell level, that make it a unique technology for investigation in the field of cell biology. However, time-lapse video microscopy has limitations that can interfere with the acquisition of high quality images. Images can be compromised by both external factors; temperature fluctuations, vibrations, humidity and internal factors; pH, cell motility. Herein, we describe a protocol for the dynamic acquisition of a specific protein, Parkin, fused with the enhanced yellow fluorescent protein (EYFP) in order to track the selective removal of damaged mitochondria, using a time-lapse video microscopy approach.

Herein, we describe in detail a time-lapse video microscopy approach to measuring the temporal recruitment of EYFP-Parkin during the selective removal of damaged mitochondria. This dynamic process of EYFP-Parkin-dependent removal of damaged mitochondria can be used as an indicator of cellular health under different experimental conditions.

Time-lapse video microscopy can be defined as the real time imaging of living cells. This technique relies on the collection of images at different time ...

The Lactate Dehydrogenase Sequestration Assay &#8212; A Simple and Reliable Method to Determine Bulk Autophagic Sequestration Activity in Mammalian Cells

Bulk autophagy is characterized by the sequestration of large portions of cytoplasm into double/multi-membrane structures termed autophagosomes. Here a simple protocol to monitor this process is described. Moreover, typical results and experimental validation of the method under autophagy-inducing conditions in various types of cultured mammalian cells are provided. During bulk autophagy, autophagosomes sequester cytosol, and thereby also soluble cytosolic proteins, alongside other autophagic cargo. LDH is a stable and highly abundant, soluble cytosolic enzyme that is non-selectively sequestered into autophagosomes. The amount of LDH sequestration therefore reflects the amount of bulk autophagic sequestration. To efficiently and accurately determine LDH sequestration in cells, we employ an electrodisruption-based fractionation protocol that effectively separates sedimentable from cytosolic LDH, followed by measurement of enzymatic activity in sedimentable fractions versus whole-cell samples. Autophagic sequestration is determined by subtracting the proportion of sedimentable LDH in untreated cells from that in treated cells. The advantage of the LDH sequestration assay is that it gives a quantitative measure of the autophagic sequestration of endogenous cargo, as opposed to other methods that either involve ectopic expression of sequestration probes or semi-quantitative protease protection analyses of autophagy markers or receptors.

Here a simple and well-validated protocol for measuring bulk autophagic sequestration activity in mammalian cells is described. The method is based on quantifying the proportion of lactate dehydrogenase (LDH) in sedimentable cell fractions compared to total cellular LDH levels.

Bulk autophagy is characterized by the sequestration of large portions of cytoplasm into double/multi-membrane structures termed autophagosomes. Here a ...

The Microscopy-Based Assay to Study and Analyze the Recycling Endosomes using SNARE Trafficking

Recycling endosomes (REs) are tubular-vesicular organelles generated from early/sorting endosomes in all cell types. These organelles play a key role in the biogenesis of melanosomes, a lysosome-related organelle produced by melanocytes. REs deliver the melanocyte-specific cargo to premature melanosomes during their formation. Blockage in the generation of REs, observed in several mutants of Hermansky-Pudlak syndrome, results in hypopigmentation of skin, hair, and eye. Therefore, studying the dynamics (refer to number and length) of REs is useful to understand the function of these organelles in normal and disease conditions. In this study, we aim to measure the RE dynamics using a resident SNARE STX13.

Recycling endosomes are part of the endosomal tubular network. Here we present a method to quantify the dynamics of recycling endosomes using GFP-STX13 as an organelle marker.

Recycling endosomes (REs) are tubular-vesicular organelles generated from early/sorting endosomes in all cell types. These organelles play a key role in ...

Monitoring Stub1-Mediated Pexophagy

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