We thank the members of the Evans lab for their thoughtful feedback on this manuscript. This work is supported by Duke Whitehead Scholars, Duke Science and Technology Scholars, and Howard Hughes Medical Institute (HHMI) Hanna Gray Fellowship. Figure 1A was made using BioRender.com.

Quantification of Mitochondrial Membrane Potential and Superoxide Levels

The authors have no competing interests to declare.

The workflow outlined here can be used to quantify mitochondrial membrane potential and superoxide levels robustly and reproducibly using fluorescence-based imaging30. There are important technical limitations to consider when designing these experiments. HeLa cells were transfected with an empty YFP vector, YFP-ParkinWT, or YFP-ParkinT240R. The empty YFP vector was used as a control to confirm that the experimental findings were specific to Parkin. For the transient transfection, a mass ratio of 1:3 for DNA to transfection reagent was experimentally determined to yield the highest transfection rate, where 2 &#181;g of plasmid DNA was used for each construct28. Keeping the YFP-tag consistent for all constructs was important, as fluorescent proteins differ in innate brightness31,32. If multiple fluorescent protein tags must be used, then fluorescent proteins with similar brightness and photostability should be selected33.
To measure mitochondrial membrane potential and superoxide levels, two well-documented dyes were selected. The fluorescence signal of TMRE is based on the mitochondrial membrane potential, while MitoSOX fluorescence intensity is a function of superoxide levels. For fluorescence-based imaging, there should be no spectral overlap between fluorescent probes. However, the initial protocol was performed using mCherry-ParkinWT and mCherry-ParkinT240R, which overlapped with the red-shifted spectra of TMRE and MitoSOX. To avoid spectral overlap and minimize potential crosstalk, YFP-tagged Parkin constructs were selected. This adjustment necessitated altering the MitoTracker dye from green to far-red. In addition, the HeLa cell plating density was optimized; initially, HeLa cells were plated at 45,000 cells per imaging dish, but this resulted in over-confluent dishes. High cell confluency can reduce transfection efficiency, promote cell death, and alter cellular metabolism34,35,36. To avoid these potential impacts, the cells were plated at a density of 30,000 cells per imaging dish.
The main limitation of this protocol is the potential for user bias, specifically when selecting cells and ROI locations. The protocol uses changes in the TMRE and MitoSOX fluorescence intensities as a functional readout for membrane potential and superoxide levels. Therefore, cell selection using these probes could potentially bias the experimental outcomes. To combat this bias, we chose cells based solely on YFP expression. Nonoverlapping ROIs were randomly selected across the mitochondrial network within the cell. Though this method was previously used to measure fluorescence intensity27, additional bias-reducing measures could be taken. First, the study could be blinded using the Blind Analysis Tools in ImageJ to prevent ROI selection that supports expected outcomes. Second, fluorescence intensities could be measured using advanced image analysis software that eliminates the need for ROIs.
While this protocol uses confocal microscopy for fluorescence-based quantification, additional methods, such as flow cytometry, could be used to quantify fluorescence changes in TMRE and MitoSOX37. Here, we used confocal microscopy rather than flow cytometry to visualize the mitochondrial network morphology. The outlined protocol could easily be adapted to measure TMRE and MitoSOX fluorescence with flow cytometry yielding similar experimental results. In addition, oxygen consumption rate (OCR) assays could be used to detect changes in the function of the mitochondrial electron transport chain without cationic dyes. While OCR provides a sensitive measure of mitochondrial dysfunction, it is not specific to membrane potential or superoxide concentration, but instead provides a global measure of mitochondrial function38. These assays could be performed in conjunction with TMRE and MitoSOX experiments to assess mitochondrial health39.
Though this protocol specifically focuses on the effect of the mitochondrial uncoupler CCCP40, damaging reagents with alternative mechanisms could be utilized. For example, antimycin A and oligomycin A are electron transport chain inhibitors commonly used to induce mitochondrial damage via reactive oxygen species (ROS) production38. While we specifically measured mitochondrial superoxide levels using MitoSOX, intracellular ROS could be measured using CellROX. In HeLa cells, it was necessary to overexpress Parkin due to the low endogenous expression. Future studies could observe the effects of Parkin expression on mitochondrial network health using cell culture systems that express endogenous Parkin41. Since mitochondria are critical regulators of energy metabolism and cellular homeostasis, mitochondrial dysfunction is associated with numerous diseases, including diabetes42, Alzheimer&#39;s disease43,44,45, cancer46, and liver disease47. Therefore, this workflow could be adapted to study mitochondrial health and dysregulation in relevant cell lines and primary cultures.

The mitochondrial network is a series of interconnected organelles that play a crucial role in energy production1, innate immunity2,3, and cell signalling4,5. Mitochondrial dysregulation has been associated with neurodegenerative diseases such as Parkinson&#39;s disease (PD)6,7. PD is a progressive neurodegenerative disorder affecting dopaminergic neurons of the substantia nigra that impacts nearly 10 million people worldwide8. PD has been genetically linked to mitophagy, a mitochondrial quality control pathway necessary for maintaining cellular homeostasis that selectively removes damaged mitochondria9,10. Studies have identified multiple independent mitophagy pathways, including FUN14 domain containing 1 (FUNDC1)-mediated mitophagy, Bcl-2 interacting protein 3 (BNIP3)-facilitated mitophagy, NIX-dependent mitophagy, and the well-characterized PTEN-induced kinase 1 (PINK1)/Parkin-regulated mitophagy10,11. PINK1 (a putative kinase) and Parkin (an E3 ubiquitin ligase) work in tandem to phospho-ubiquitinate damaged mitochondria, which drives the formation of autophagosomes that engulf the damaged organelle and fuse with lysosomes to initiate degradation12,13,14,15,16. Mutations in Parkin have been associated with PD-linked phenotypes such as neurodegeneration via the loss of dopaminergic neurons17,18.
Here, a protocol is described in which HeLa cells, routinely used immortalized cells derived from cervical cancer, are used to investigate the role of Parkin in maintaining mitochondrial network health. HeLa cells express negligible levels of endogenous Parkin and therefore require exogenous Parkin expression19. To study the role of Parkin in mitochondrial network health, HeLa cells are transfected with either wild-type Parkin (ParkinWT), a Parkin mutant (ParkinT240R), or an empty control vector. ParkinT240R is an autosomal recessive juvenile parkinsonism mutation that affects Parkin E3 ligase activity, significantly reducing the efficiency of the mitophagy pathway20. HeLa cells are subject to mild (5 &#181;M) or severe (20 &#181;M) concentrations of carbonyl cyanide m-chlorophenyl hydrazone (CCCP), a mitochondrial uncoupling agent. Treatment with severe concentrations of CCCP is routinely used to induce Parkin-mediated mitophagy in various cell lines, such as HeLa and COS-7 cells21,22,23.
Following treatment, the protocol employs live imaging of the mitochondrial network using two currently available mitochondrial-targeted fluorescent dyes. Tetramethylrhodamine, ethyl ester, perchlorate (TMRE) is a cationic dye that fluoresces based on mitochondrial membrane potential24, while MitoSOX is a mitochondrial superoxide indicator where the fluorescence intensity is a function of superoxide concentration25. Finally, the outlined protocol uses a fluorescence-based quantification and simple workflow to effectively quantify the mitochondrial membrane potential and superoxide levels with minimal scope for user bias. Although this protocol was designed to study mitochondrial function in HeLa cells, it can be adapted for additional cell lines and primary cell types to quantitively characterize mitochondrial network health.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Chemicals, Peptides, and Recombinant Proteins</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>CCCP (carbonyl cyanide m-chlorophenyl hydrazone)&#160;</td>
			<td>Sigma-Aldrich</td>
			<td>C2759</td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM (1x) with 4.5 g/L glucose</td>
			<td>Gibco</td>
			<td>11-965-084</td>
			<td></td>
		</tr>
		<tr>
			<td>DMSO, Anhydrous</td>
			<td>ThermoFisher Scientific</td>
			<td>D12345</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum</td>
			<td>Hyclone</td>
			<td>SH3007103</td>
			<td></td>
		</tr>
		<tr>
			<td>FuGENE 6 (Tranfection Reagent)</td>
			<td>Promega</td>
			<td>E2691</td>
			<td></td>
		</tr>
		<tr>
			<td>GlutaMAX 100x (L-Glutamine Solution)&#160;</td>
			<td>Gibco&#160;</td>
			<td>35-050-061</td>
			<td></td>
		</tr>
		<tr>
			<td>Hoescht 33342</td>
			<td>ThermoFisher Scientific</td>
			<td>62249</td>
			<td></td>
		</tr>
		<tr>
			<td>MitoSOX&#160; Red&#160;</td>
			<td>ThermoFisher Scientific</td>
			<td>M36008</td>
			<td></td>
		</tr>
		<tr>
			<td>MitoTracker Deep Red</td>
			<td>ThermoFisher Scientific</td>
			<td>M7514</td>
			<td></td>
		</tr>
		<tr>
			<td>Opti-MEM (Redued Serum media)</td>
			<td>ThermoFisher scientific</td>
			<td>31985070</td>
			<td></td>
		</tr>
		<tr>
			<td>Tetramethylrhodamine, Ethyl Ester, Perchlorate (TMRE)&#160;</td>
			<td>ThermoFisher Scientific</td>
			<td>T669</td>
			<td></td>
		</tr>
		<tr>
			<td>Experimental models: Organisms/Strains</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>HeLa-M (Homo sapiens)</td>
			<td>A. Peden (Cambridge Institute for Medical Research)</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Recombinant DNA</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>EYFP Empty Vector</td>
			<td>N/A</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>YFP-Parkin T240R</td>
			<td>This Paper</td>
			<td>Generated by site-directed mutagenesis from YFP-Parkin</td>
			<td></td>
		</tr>
		<tr>
			<td>YFP-Parkin WT</td>
			<td>Addgene; PMID:19029340</td>
			<td>RRID:Addgene_23955</td>
			<td></td>
		</tr>
		<tr>
			<td>Software and Algorithms</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Adobe Illustrator</td>
			<td>Adobe Inc.</td>
			<td>https://www.adobe.com/products/illustrator</td>
			<td>(Schindelin, 2012)</td>
		</tr>
		<tr>
			<td>Excel (Spreadsheet Software)</td>
			<td>Microsoft Office&#160;</td>
			<td>https://www.microsoft.com/en-us/microsoft-365/excel</td>
			<td></td>
		</tr>
		<tr>
			<td>ImageJ</td>
			<td></td>
			<td>https://imagej.net/software/fiji/</td>
			<td></td>
		</tr>
		<tr>
			<td>Leica Application Suite (LAS X)</td>
			<td>Leica</td>
			<td>https://www.leica-microsystems.com/products/microscope-software/p/leica-las-x-ls/</td>
			<td></td>
		</tr>
		<tr>
			<td>Microsoft Excel</td>
			<td>Microsoft Office</td>
			<td>https://www.microsoft.com/excel</td>
			<td></td>
		</tr>
		<tr>
			<td>Prism9 (Statistical Analysis Software)</td>
			<td>GraphPad Software</td>
			<td>https://www.graphpad.com</td>
			<td></td>
		</tr>
		<tr>
			<td>Other</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>35 mm Dish, No. 1.5 Coverslip, 20 mm Glass Diameter, Uncoated</td>
			<td>MatTek</td>
			<td>P35G-1.5-20-C</td>
			<td></td>
		</tr>
		<tr>
			<td>Cage Incubator (Environmental Chamber)</td>
			<td>Okolab</td>
			<td>https://www.oko-lab.com/cage-incubator</td>
			<td></td>
		</tr>
		<tr>
			<td>DMiL Inverted Microscope</td>
			<td>Leica</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>LIGHTNING Deconvolution Software</td>
			<td>Leica</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>STELLARIS 8 confocal microscope</td>
			<td>Leica</td>
			<td>N/A</td>
			<td></td>
		</tr>
	</tbody>
</table>

fluorescence-based quantification of mitochondrial membrane potential and superoxide levels using live imaging in hela cells

Mitochondria are dynamic organelles critical for metabolic homeostasis by controlling energy production via ATP synthesis. To support cellular metabolism, various mitochondrial quality control mechanisms cooperate to maintain a healthy mitochondrial network. One such pathway is mitophagy, where PTEN-induced kinase 1 (PINK1) and Parkin phospho-ubiquitination of damaged mitochondria facilitate autophagosome sequestration and subsequent removal from the cell via lysosome fusion. Mitophagy is important for cellular homeostasis, and mutations in Parkin are linked to Parkinson's disease (PD). Due to these findings, there has been a significant emphasis on investigating mitochondrial damage and turnover to understand the molecular mechanisms and dynamics of mitochondrial quality control. Here, live-cell imaging was used to visualize the mitochondrial network of HeLa cells, to quantify the mitochondrial membrane potential and superoxide levels following treatment with carbonyl cyanide m-chlorophenyl hydrazone (CCCP), a mitochondrial uncoupling agent. In addition, a PD-linked mutation of Parkin (ParkinT240R) that inhibits Parkin-dependent mitophagy was expressed to determine how mutant expression impacts the mitochondrial network compared to cells expressing wild-type Parkin. The protocol outlined here describes a simple workflow using fluorescence-based approaches to quantify mitochondrial membrane potential and superoxide levels effectively.

In this protocol, fluorescence-based quantification was used to measure the membrane potential and superoxide levels of the mitochondrial network following CCCP treatment (Figure 1). This workflow used HeLa cells, an immortalized cell line derived from cervical cancer. HeLa cells are routinely used to study mitochondrial biology and are relatively flat, making it easy to visualize the mitochondrial network using microscopy. To investigate the role of Parkin in maintaining mitochondrial network health, HeLa cells were transiently transfected with an empty control vector, ParkinWT, or ParkinT240R (a mutant that disrupts mitophagy)21. HeLa cells were plated in 35 mm glass-bottom imaging dishes and were transfected once ~50%-60% confluency was reached (Figure 1A). Since HeLa cells rapidly divide, this is typically the day after plating the cells into the imaging dishes. A light microscope was used to observe the YFP fluorescence signal the following day to assess the transfection efficiency. Experiments were only performed following a successful transfection.
After inspection, the HeLa cells were treated with mild (5 &#181;M) or severe (20 &#181;M) concentrations of CCCP to depolarize the mitochondrial network (step&#160;1.2 shows detailed instructions for preparing CCCP and fluorescent probes). The CCCP treatment was performed for a total of 2 h. During the final 30 min of the CCCP treatment, the mitochondria were labeled with MitoTracker and TMRE/MitoSOX (Figure 1A). It should be noted that TMRE and MitoSOX have overlapping fluorescence spectra and cannot be used simultaneously. Instead, we used separate imaging dishes for the TMRE and MitoSOX experiments. For TMRE experiments, the HeLa cells were immediately ready to image at the end of the 2 h CCCP treatment. The TMRE concentration must remain constant; therefore, TMRE remained in the HeLa media. However, free MitoSOX must be removed before imaging. For the MitoSOX experiments, the cells were washed three times with HeLa media to remove the free dye. For this step, it is essential to use HeLa media containing the same DMSO or CCCP concentration as the experimental conditions. Hoechst 33342, a nuclei marker, was initially used to assess the HeLa cells following the transient transfection and CCCP treatment (Figure 1B).
Subsequently, confocal microscopy was performed to visualize TMRE and MitoSOX fluorescence intensities, to measure the mitochondrial membrane potential and superoxide levels, respectively. The images were acquired using a 63x (1.4 NA) oil immersion objective with parameters of bidirectional scanning, a spatial resolution of 1,024 x 1,024 pixels, a scanning speed of 600 Hz, line averaging of 3, a phase of 22.61, and a zoom factor of 1.5 (Figure 2C). The DMSO control plate was imaged first for all experiments to set the gain and intensity values for TMRE and MitoSOX. To make quantitative comparisons across conditions, these values must be set in the DMSO condition and remain constant throughout the experiment. CCCP induces mitochondrial depolarization and a loss of membrane potential, resulting in a reduced TMRE fluorescence intensity. Therefore, the control experiments had the highest TMRE intensity, and the gain and intensity values were set close to saturation. In contrast, higher superoxide levels increase the MitoSOX intensity. Therefore, the control had the lowest MitoSOX fluorescence intensity, and the gain and intensity values were set low where a dim signal was present. Since MitoTracker, TMRE, and MitoSOX are vital dyes and label all cells, they should not be used when selecting cells to image. Instead, the cells were selected based on the YFP signal to ensure they were transfected. Single-plane images were acquired, focusing on the bottom of the HeLa cell, where a large population of mitochondria was located.
Quantification of TMRE and MitoSOX fluorescence intensity 
Quantification of the TMRE and MitoSOX fluorescence intensities was analyzed using ImageJ (Figure 3). A 6 micron x 6 micron ROI was created and stored in the ROI manager. The ROI was placed in five random nonoverlapping regions of each cell to measure the TMRE or MitoSOX intensity. The fluorescence intensity corresponds to the mean gray value in ImageJ. The five intensity values were averaged for each cell to calculate the mean fluorescence intensity per cell. These values were plotted and analyzed for statistical significance across treatment conditions.
The results for the TMRE and MitoSOX fluorescence intensities are shown in Figure 4 and Figure 5, respectively. As expected, treatment with the known uncoupling agent CCCP decreased the TMRE fluorescence intensity compared to the control conditions (Figure 4A,B). In addition, severe (20 &#181;M) CCCP treatment induced superoxide production and increased the MitoSOX fluorescence intensity (Figure 5A,B). In mild (5 &#181;M) CCCP stress conditions, the expression of ParkinWT and ParkinT240R resulted in higher TMRE intensity compared to the empty YFP control vector. Similarly, the MitoSOX intensity was lower in cells expressing ParkinWT and ParkinT240R compared to cells expressing the YFP control vector (Figure 5A,B). These results suggest that Parkin expression helps maintain the mitochondrial network health by preserving higher mitochondrial membrane potentials and low superoxide levels. Thus, the protocol outlined here can be used to accurately compare fluorescence intensity to analyze the role of Parkin in controlling mitochondrial membrane potential and superoxide formation.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65304/65304fig01.jpg" /> 
Figure 1: Experimental workflow. (A) Schematic of the experimental workflow used to transfect, treat with CCCP, and label the mitochondria network, and image TMRE and MitoSOX fluorescence intensity in HeLa cells.&#160;(B)&#160;Representative images of cells expressing an empty YFP vector (top), YFP-ParkinWT (middle), and YFP-ParkinT240R (bottom; magenta). Hoechst 33342 (white)&#160;is used to label the DNA. Scale bar = 10 &#181;m. Abbreviations: CCCP = carbonyl cyanide m-chlorophenyl hydrazone; TMRE = tetramethylrhodamine-ethyl ester-perchlorate; YFP = yellow fluorescent protein. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65304/65304fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/65304/65304fig02.jpg" /> 
Figure 2: The user interface to set up confocal acquisition settings. (A) Laser settings dialog box. The red rectangle highlights the white light laser, power state, laser power, and wavelengths. (B) Experimental channel set up for a MitoSOX experiment. The settings, excitation lines, and emission spectra windows are shown for YFP, MitoSOX, and MitoTracker. (C) Acquisition settings show the format, speed, bidirectionality, phase X, zoom factor, and line average.(D) A representative acquired image. HeLa cells are expressing YFP (magenta), and the mitochondria are labeled with MitoSOX (green) and MitoTracker (cyan). Scale bar = 10 &#181;m. Abbreviations: WLL = white light laser; YFP = yellow fluorescent protein. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65304/65304fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/65304/65304fig03.jpg" /> 
Figure 3: ImageJ workflow to quantify TMRE and MitoSOX fluorescence intensities. (A)&#160;The ROI manager panel with an example ROI in ImageJ. (B) Bio-formats import options panel. The red rectangle highlights the Split Channels option that should be selected. (C) Brightness and contrast settings parameters.&#160;(D)&#160;Set measurements parameter. The red rectangles highlight the Area and Mean gray value options that should be selected.&#160;(E) Representative image of a HeLa cell labeled with MitoSOX. The red square illustrates the ROI from panel A. Scale bar = 10 &#181;m.&#160;(F) Results panel showing the experimental area and mean gray values from the ROIs. The mean gray value (orange) represents the fluorescence intensity values. Abbreviations: TMRE = tetramethylrhodamine-ethyl ester-perchlorate; ROI = region of interest. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65304/65304fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/65304/65304fig04.jpg" /> 
Figure 4: TMRE fluorescence intensity after mitochondrial damage. (A) Representative images of HeLa cells following a 2 h treatment with DMSO, 5 &#181;M CCCP, or 20 &#181;M CCCP to induce mitochondrial damage. Cells are exogenously expressing an empty YFP vector, YFP-ParkinWT, or YFP-ParkinT240R (magenta), and labeled with MitoTracker (cyan) and TMRE (green). Scale bar = 30 &#181;m. (B) Quantification of TMRE fluorescence intensity for cells expressing empty YFP vector (blue), YFP-ParkinWT (orange), and YFP-ParkinT240R (purple) in DMSO- and CCCP-treated conditions. ****p &#60; 0.0001 by two-way ANOVA with a multiple comparison test. (C-E) The quantification of the TMRE fluorescence intensities from panel B is separated to highlight differences in DMSO (C), 5 &#181;M CCCP (D), and 20 &#181;M CCCP (E). * p &#60; 0.05; ***p &#60; 0.001; ****p &#60; 0.0001 by Kruskal-Wallis ANOVA with Dunn&#39;s multiple comparison test. Ns = not significant. Mean &#177; SEM; n= 79-104 from three independent biological replicates. Abbreviations: CCCP = carbonyl cyanide m-chlorophenyl hydrazone; TMRE = tetramethylrhodamine-ethyl ester-perchlorate; YFP = yellow fluorescent protein; DMSO = dimethyl sulfoxide. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65304/65304fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/65304/65304fig05.jpg" /> 
Figure 5: MitoSOX fluorescence intensity following damage to the mitochondrial network. (A) Representative images of HeLa cells treated for 2 h with DMSO, 5 &#181;M CCCP, or 20 &#181;M CCCP to uncouple the mitochondria membrane potential. Cells were transfected with empty YFP vector, YFP-ParkinWT, or YFP-ParkinT240R (magenta), and labeled with MitoTracker (cyan) and MitoSOX (green). Scale bar = 30 &#181;m. (B) Quantification of MitoSOX fluorescence intensity for cells expressing empty YFP vector (blue), YFP-ParkinWT (orange), and YFP-ParkinT240R (purple) for control and treated conditions. ****p &#60; 0.0001 by two-way ANOVA with Dunn&#39;s multiple comparison test. (C-E) The quantification of the MitoSOX fluorescence intensities from panel B is separated to highlight differences in DMSO (C), 5 &#181;M CCCP (D), and 20 &#181;M CCCP (E). *p &#60; 0.05; ****p &#60; 0.0001 by Kruskal-Wallis ANOVA with Dunn&#39;s multiple comparison test. Ns = not significant. Mean &#177; SEM; n = 87-107 from three independent biological replicates. Abbreviations: CCCP = carbonyl cyanide m-chlorophenyl hydrazone; TMRE = tetramethylrhodamine-ethyl ester-perchlorate; YFP = yellow fluorescent protein; DMSO = dimethyl sulfoxide. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65304/65304fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Fluorescence-Based Quantification of Mitochondrial Membrane Potential and Superoxide Levels Using Live Imaging in HeLa Cells at JoVE.com

Author Spotlight: Fluorescence-Based Quantification of Mitochondrial Membrane Potential and Superoxide Levels Using Live Imaging in HeLa Cells  

This technique describes an effective workflow to visualize and quantitatively measure mitochondrial membrane potential and superoxide levels within HeLa cells using fluorescence-based live imaging.

Fluorescence-Based Quantification of Mitochondrial Membrane Potential and Superoxide Levels Using Live Imaging in HeLa Cells

Mitochondria are dynamic organelles critical for metabolic homeostasis by controlling energy production via ATP synthesis. To support cellular metabolism, ...

fluorescence-based-quantification-mitochondrial-membrane-potential

Nanyang Technological University

Research

JoVE Journal

Biology

7.1K Views.  Duke University Medical Center. This technique describes an effective workflow to visualize and quantitatively measure mitochondrial membrane potential and superoxide levels within HeLa cells using fluorescence-based live imaging. 

Video: Author Spotlight: Fluorescence-Based Quantification of Mitochondrial Membrane Potential and Superoxide Levels Using Live Imaging in HeLa Cells  

Live-cell Imaging and Quantitative Analysis of Embryonic Epithelial Cells in Xenopus laevis

Embryonic epithelial cells serve as an ideal model to study morphogenesis where multi-cellular tissues undergo changes in their geometry, such as changes in cell surface area and cell height, and where cells undergo mitosis and migrate. Furthermore, epithelial cells can also regulate morphogenetic movements in adjacent tissues1. A traditional method to study epithelial cells and tissues involve chemical fixation and histological methods to determine cell morphology or localization of particular proteins of interest. These approaches continue to be useful and provide "snapshots" of cell shapes and tissue architecture, however, much remains to be understood about how cells acquire specific shapes, how various proteins move or localize to specific positions, and what paths cells follow toward their final differentiated fate. High resolution live imaging complements traditional methods and also allows more direct investigation into the dynamic cellular processes involved in the formation, maintenance, and morphogenesis of multicellular epithelial sheets. Here we demonstrate experimental methods from the isolation of animal cap tissues from Xenopus laevis embryos to confocal imaging of epithelial cells and simple measurement approaches that together can augment molecular and cellular studies of epithelial morphogenesis.

Live-cell Imaging and Quantitative Analysis of Embryonic Epithelial Cells in Xenopus laevis

Xenopus embryonic epithelia are an ideal model system to study cell behaviors such as polarity development and shape change during epithelial morphogenesis. Traditional histology of fixed samples is increasingly being complemented by live-cell confocal imaging. Here we demonstrate methods to isolate frog tissues and visualize live epithelial cells and their cytoskeleton using live-cell confocal microscopy.

Embryonic epithelial cells serve as an ideal model to study morphogenesis where multi-cellular tissues undergo changes in their geometry, such as changes ...

Monitoring Dynamic Changes In Mitochondrial Calcium Levels During Apoptosis Using A Genetically Encoded Calcium Sensor

Dynamic changes in intracellular calcium concentration in response to various stimuli regulates many cellular processes such as proliferation, differentiation, and apoptosis1. During apoptosis, calcium accumulation in mitochondria promotes the release of pro-apoptotic factors from the mitochondria into the cytosol2. It is therefore of interest to directly measure mitochondrial calcium in living cells in situ during apoptosis. High-resolution fluorescent imaging of cells loaded with dual-excitation ratiometric and non-ratiometric synthetic calcium indicator dyes has been proven to be a reliable and versatile tool to study various aspects of intracellular calcium signaling. Measuring cytosolic calcium fluxes using these techniques is relatively straightforward. However, measuring intramitochondrial calcium levels in intact cells using synthetic calcium indicators such as rhod-2 and rhod-FF is more challenging. Synthetic indicators targeted to mitochondria have blunted responses to repetitive increases in mitochondrial calcium, and disrupt mitochondrial morphology3. Additionally, synthetic indicators tend to leak out of mitochondria over several hours which makes them unsuitable for long-term experiments. Thus, genetically encoded calcium indicators based upon green fluorescent protein (GFP)4 or aequorin5 targeted to mitochondria have greatly facilitated measurement of mitochondrial calcium dynamics. Here, we describe a simple method for real-time measurement of mitochondrial calcium fluxes in response to different stimuli. The method is based on fluorescence microscopy of 'ratiometric-pericam' which is selectively targeted to mitochondria. Ratiometric pericam is a calcium indicator based on a fusion of circularly permuted yellow fluorescent protein and calmodulin4. Binding of calcium to ratiometric pericam causes a shift of its excitation peak from 415 nm to 494 nm, while the emission spectrum, which peaks around 515 nm, remains unchanged. Ratiometric pericam binds a single calcium ion with a dissociation constant in vitro of ~1.7 &mu;M4. These properties of ratiometric pericam allow the quantification of rapid and long-term changes in mitochondrial calcium concentration. Furthermore, we describe adaptation of this methodology to a standard wide-field calcium imaging microscope with commonly available filter sets. Using two distinct agonists, the purinergic agonist ATP and apoptosis-inducing drug staurosporine, we demonstrate that this method is appropriate for monitoring changes in mitochondrial calcium concentration with a temporal resolution of seconds to hours. Furthermore, we also demonstrate that ratiometric pericam is also useful for measuring mitochondrial fission/fragmentation during apoptosis. Thus, ratiometric pericam is particularly well suited for continuous long-term measurement of mitochondrial calcium dynamics during apoptosis.

This protocol describes a method for real-time measurement of mitochondrial calcium fluxes by fluorescent imaging. The method takes advantage of a circularly permutated YFP-based dual-excitation ratiometric calcium sensor (ratiometric pericam-mt) selectively expressed in mitochondria.

Dynamic changes in intracellular calcium concentration in response to various stimuli regulates many cellular processes such as proliferation, ...

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to replenish depleted endoplasmic reticulum (ER) or sarcoplasmic reticulum (SR) Ca2+ stores1,2. Since Ca2+ is a ubiquitous second messenger, it is not surprising to see that SOCE plays important roles in a variety of cellular processes, including proliferation, apoptosis, gene transcription and motility. Due to its wide occurrence in nearly all cell types, including epithelial cells and skeletal muscles, this pathway has received great interest3,4. However, the heterogeneity of SOCE characteristics in different cell types and the physiological function are still not clear5-7.
The functional channel properties of SOCE can be revealed by patch-clamp studies, whereas a large body of knowledge about this pathway has been gained by fluorescence-based intracellular Ca2+ measurements because of its convenience and feasibility for high-throughput screening. The objective of this report is to summarize a few fluorescence-based methods to measure the activation of SOCE in monolayer cells, suspended cells and muscle fibers5,8-10. The most commonly used of these fluorescence methods is to directly monitor the dynamics of intracellular Ca2+ using the ratio of F340nm and F380nm (510 nm for emission wavelength) of the ratiometric Ca2+ indicator Fura-2. To isolate the activity of unidirectional SOCE from intracellular Ca2+ release and Ca2+ extrusion, a Mn2+ quenching assay is frequently used. Mn2+ is known to be able to permeate into cells via SOCE while it is impervious to the surface membrane extrusion processes or to ER uptake by Ca2+ pumps due to its very high affinity with Fura-2. As a result, the quenching of Fura-2 fluorescence induced by the entry of extracellular Mn2+ into the cells represents a measurement of activity of SOCE9. Ratiometric measurement and the Mn+2 quenching assays can be performed on a cuvette-based spectrofluorometer in a cell population mode or in a microscope-based system to visualize single cells. The advantage of single cell measurements is that individual cells subjected to gene manipulations can be selected using GFP or RFP reporters, allowing studies in genetically modified or mutated cells. The spatiotemporal characteristics of SOCE in structurally specialized skeletal muscle can be achieved in skinned muscle fibers by simultaneously monitoring the fluorescence of two low affinity Ca2+ indicators targeted to specific compartments of the muscle fiber, such as Fluo-5N in the SR and Rhod-5N in the transverse tubules9,11,12.

The extent of store-operated Ca2+ entry (SOCE) can be monitored using fluorescent Ca2+ indicators. Mn2+ quenching of such indicators assays SOCE in cultured cells and skeletal muscle fibers. A technique allowing spatial and temporal resolution of SOCE by confocal imaging of mechanically skinned muscle fibers is also described.

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to ...

4D Imaging of Protein Aggregation in Live Cells

One of the key tasks of any living cell is maintaining the proper folding of newly synthesized proteins in the face of ever-changing environmental conditions and an intracellular environment that is tightly packed, sticky, and hazardous to protein stability1. The ability to dynamically balance protein production, folding and degradation demands highly-specialized quality control machinery, whose absolute necessity is observed best when it malfunctions. Diseases such as ALS, Alzheimer's, Parkinson's, and certain forms of Cystic Fibrosis have a direct link to protein folding quality control components2, and therefore future therapeutic development requires a basic understanding of underlying processes. Our experimental challenge is to understand how cells integrate damage signals and mount responses that are tailored to diverse circumstances.
The primary reason why protein misfolding represents an existential threat to the cell is the propensity of incorrectly folded proteins to aggregate, thus causing a global perturbation of the crowded and delicate intracellular folding environment1. The folding health, or "proteostasis," of the cellular proteome is maintained, even under the duress of aging, stress and oxidative damage, by the coordinated action of different mechanistic units in an elaborate quality control system3,4. A specialized machinery of molecular chaperones can bind non-native polypeptides and promote their folding into the native state1, target them for degradation by the ubiquitin-proteasome system5, or direct them to protective aggregation inclusions6-9.
In eukaryotes, the cytosolic aggregation quality control load is partitioned between two compartments8-10: the juxtanuclear quality control compartment (JUNQ) and the insoluble protein deposit (IPOD) (Figure 1 - model). Proteins that are ubiquitinated by the protein folding quality control machinery are delivered to the JUNQ, where they are processed for degradation by the proteasome. Misfolded proteins that are not ubiquitinated are diverted to the IPOD, where they are actively aggregated in a protective compartment.
Up until this point, the methodological paradigm of live-cell fluorescence microscopy has largely been to label proteins and track their locations in the cell at specific time-points and usually in two dimensions. As new technologies have begun to grant experimenters unprecedented access to the submicron scale in living cells, the dynamic architecture of the cytosol has come into view as a challenging new frontier for experimental characterization. We present a method for rapidly monitoring the 3D spatial distributions of multiple fluorescently labeled proteins in the yeast cytosol over time. 3D timelapse (4D imaging) is not merely a technical challenge; rather, it also facilitates a dramatic shift in the conceptual framework used to analyze cellular structure.
We utilize a cytosolic folding sensor protein in live yeast to visualize distinct fates for misfolded proteins in cellular aggregation quality control, using rapid 4D fluorescent imaging. The temperature sensitive mutant of the Ubc9 protein10-12 (Ubc9ts) is extremely effective both as a sensor of cellular proteostasis, and a physiological model for tracking aggregation quality control. As with most ts proteins, Ubc9ts is fully folded and functional at permissive temperatures due to active cellular chaperones. Above 30 &deg;C, or when the cell faces misfolding stress, Ubc9ts misfolds and follows the fate of a native globular protein that has been misfolded due to mutation, heat denaturation, or oxidative damage. By fusing it to GFP or other fluorophores, it can be tracked in 3D as it forms Stress Foci, or is directed to JUNQ or IPOD.

Cellular viability depends on timely and efficient management of protein misfolding. Here we describe a method for visualizing the different potential fates of a misfolded protein: refolding, degradation, or sequestration in inclusions. We demonstrate the use of a folding sensor, Ubc9ts, for monitoring proteostasis and aggregation quality control in live cells using 4D microscopy.

One of the key tasks of any living cell is maintaining the proper folding of newly synthesized proteins in the face of ever-changing environmental ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...

Intravital Microscopy for Imaging Subcellular Structures in Live Mice Expressing Fluorescent Proteins

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, we use the salivary glands of live mice since they provide several advantages. First, they can be easily exposed to enable access to the optics, and stabilized to facilitate the reduction of the motion artifacts due to heartbeat and respiration. This significantly facilitates imaging and tracking small subcellular structures. Second, most of the cell populations of the salivary glands are accessible from the surface of the organ. This permits the use of confocal microscopy that has a higher spatial resolution than other techniques that have been used for in vivo imaging, such as two-photon microscopy. Finally, salivary glands can be easily manipulated pharmacologically and genetically, thus providing a robust system to investigate biological processes at a molecular level.
In this study we focus on a protocol designed to follow the kinetics of the exocytosis of secretory granules in acinar cells and the dynamics of the apical plasma membrane where the secretory granules fuse upon stimulation of the beta-adrenergic receptors. Specifically, we used a transgenic mouse that co-expresses cytosolic GFP and a membrane-targeted peptide fused with the fluorescent protein tandem-Tomato. However, the procedures that we used to stabilize and image the salivary glands can be extended to other mouse models and coupled to other approaches to label in vivo cellular components, enabling the visualization of various subcellular structures, such as endosomes, lysosomes, mitochondria, and the actin cytoskeleton.

Intravital microscopy is a powerful tool that enables imaging various biological processes in live animals. In this article, we present a detailed method for imaging the dynamics of subcellular structures, such as the secretory granules, in the salivary glands of live mice.

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, ...

Confocal Imaging of Single Mitochondrial Superoxide Flashes in Intact Heart or In Vivo

Mitochondrion is a critical intracellular organelle responsible for energy production and intracellular signaling in eukaryotic systems. Mitochondrial dysfunction often accompanies and contributes to human disease. Majority of the approaches that have been developed to evaluate mitochondrial function and dysfunction are based on in vitro or ex vivo measurements. Results from these experiments have limited ability in determining mitochondrial function in vivo. Here, we describe a novel approach that utilizes confocal scanning microscopy for the imaging of intact tissues in live aminals, which allows the evaluation of single mitochondrial function in a real-time manner in vivo. First, we generate transgenic mice expressing the mitochondrial targeted superoxide indicator, circularly permuted yellow fluorescent protein (mt-cpYFP). Anesthetized mt-cpYFP mouse is fixed on a custom-made stage adaptor and time-lapse images are taken from the exposed skeletal muscles of the hindlimb. The mouse is subsequently sacrificed and the heart is set up for Langendorff perfusion with physiological solutions at 37 &deg;C. The perfused heart is positioned in a special chamber on the confocal microscope stage and gentle pressure is applied to immobilize the heart and suppress heart beat induced motion artifact. Superoxide flashes are detected by real-time 2D confocal imaging at a frequency of one frame per second. The perfusion solution can be modified to contain different respiration substrates or other fluorescent indicators. The perfusion can also be adjusted to produce disease models such as ischemia and reperfusion. This technique is a unique approach for determining the function of single mitochondrion in intact tissues and in vivo.

Confocal Imaging of Single Mitochondrial Superoxide Flashes in Intact Heart or In Vivo

Confocal scanning microscopy is applied for imaging single mitochondrial events in perfused heart or skeletal muscles in live animal. Real-time monitoring of single mitochondrial processes such as superoxide flashes and membrane potential fluctuations enables the evaluation of mitochondrial function in a physiologically relevant context and during pathological perturbations.

Mitochondrion is a critical intracellular organelle responsible for energy production and intracellular signaling in eukaryotic systems. Mitochondrial ...

Simultaneous Measurement of Mitochondrial Calcium and Mitochondrial Membrane Potential in Live Cells by Fluorescent Microscopy

Apart from their essential role in generating ATP, mitochondria also act as local calcium (Ca2+) buffers to tightly regulate intracellular Ca2+ concentration. To do this, mitochondria utilize the electrochemical potential across their inner membrane (&#916;&#936;m) to sequester Ca2+. The influx of Ca2+ into the mitochondria stimulates three rate-limiting dehydrogenases of the citric acid cycle, increasing electron transfer through the oxidative phosphorylation (OXPHOS) complexes. This stimulation maintains &#916;&#936;m, which is temporarily dissipated as the positive calcium ions cross the mitochondrial inner membrane into the mitochondrial matrix.
We describe here a method for simultaneously measuring mitochondria Ca2+ uptake and &#916;&#936;m in live cells using confocal microscopy. By permeabilizing the cells, mitochondrial Ca2+ can be measured using the fluorescent Ca2+ indicator Fluo-4, AM, with measurement of &#916;&#936;m using the fluorescent dye tetramethylrhodamine, methyl ester, perchlorate (TMRM). The benefit of this system is that there is very little spectral overlap between the fluorescent dyes, allowing accurate measurement of mitochondrial Ca2+ and &#916;&#936;m simultaneously. Using the sequential addition of Ca2+ aliquots, mitochondrial Ca2+ uptake can be monitored, and the concentration at which Ca2+ induces mitochondrial membrane permeability transition and the loss of &#916;&#936;m determined.

Mitochondria can utilize the electrochemical potential across their inner membrane (&#916;&#936;m) to sequester calcium (Ca2+), allowing them to shape cytosolic Ca2+ signaling within the cell. We describe a method for simultaneously measuring mitochondria Ca2+ uptake and &#916;&#936;m in live cells using fluorescent dyes and confocal microscopy.

Apart from their essential role in generating ATP, mitochondria also act as local calcium (Ca2+) buffers to tightly regulate intracellular Ca2+ ...

Ground State Depletion Super-resolution Imaging in Mammalian Cells

Advances in fluorescent microscopy and cell biology are intimately correlated, with the enhanced ability to visualize cellular events often leading to dramatic leaps in our understanding of how cells function. The development and availability of super-resolution microscopy has considerably extended the limits of optical resolution from ~250-20 nm. Biologists are no longer limited to describing molecular interactions in terms of colocalization within a diffraction limited area, rather it is now possible to visualize the dynamic interactions of individual molecules. Here, we outline a protocol for the visualization and quantification of cellular proteins by ground-state depletion microscopy&#160;for fixed cell imaging. We provide examples from two different membrane proteins, an element of the endoplasmic reticulum translocon, sec61&#946;, and a plasma membrane-localized voltage-gated L-type Ca2+ channel (CaV1.2). Discussed are the specific microscope parameters, fixation methods, photo-switching buffer formulation, and pitfalls and challenges of image processing.

This article outlines a protocol for the detection of one or more plasma and/or intracellular membrane proteins using Ground State Depletion (GSD) super-resolution microscopy in mammalian cells. Here, we discuss the benefits and considerations of using such approaches for the visualization and quantification of cellular proteins.

Advances in fluorescent microscopy and cell biology are intimately correlated, with the enhanced ability to visualize cellular events often leading to ...

Improving the Accuracy of Flow Cytometric Assessment of Mitochondrial Membrane Potential in Hematopoietic Stem and Progenitor Cells Through the Inhibition of Efflux Pumps

As cellular metabolism is a key regulator of hematopoietic stem cell (HSC) self-renewal, the various roles played by the mitochondria in hematopoietic homeostasis have been extensively studied by HSC researchers. Mitochondrial activity levels are reflected in their membrane potentials (&#916;&#936;m), which can be measured by cell-permeant cationic dyes such as TMRM (tetramethylrhodamine, methyl ester). The ability of efflux pumps to extrude these dyes from cells can limit their usefulness, however. The resulting measurement bias is particularly critical when assessing HSCs, as xenobiotic transporters exhibit higher levels of expression and activity in HSCs than in differentiated cells. Here, we describe a protocol utilizing Verapamil, an efflux pump inhibitor, to accurately measure &#916;&#936;m across multiple bone marrow populations. The resulting inhibition of pump activity is shown to increase TMRM intensity in hematopoietic stem and progenitor cells (HSPCs), while leaving it relatively unchanged in mature fractions. This highlights the close attention to dye-efflux activity that is required when &#916;&#936;m-dependent dyes are used, and as written and visualized, this protocol can be used to accurately compare either different populations within the bone marrow, or the same population across different experimental models.

Xenobiotic efflux pumps are highly active in hematopoietic stem and progenitor cells (HSPCs) and cause extrusion of TMRM, a mitochondrial membrane potential fluorescent dye. Here, we present a protocol to accurately measure mitochondrial membrane potential in HSPCs by TMRM in the presence of Verapamil, an efflux pump inhibitor.

As cellular metabolism is a key regulator of hematopoietic stem cell (HSC) self-renewal, the various roles played by the mitochondria in hematopoietic ...

Fluorescence-Based Quantification of Mitochondrial Membrane Potential and Superoxide Levels Using Live Imaging in HeLa Cells

Podsumowanie

Przeglądaj więcej filmów

Rozdziały w tym wideo

Live-cell Imaging and Quantitative Analysis of Embryonic Epithelial Cells in Xenopus laevis

Monitoring Dynamic Changes In Mitochondrial Calcium Levels During Apoptosis Using A Genetically Encoded Calcium Sensor

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

4D Imaging of Protein Aggregation in Live Cells

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Intravital Microscopy for Imaging Subcellular Structures in Live Mice Expressing Fluorescent Proteins

Confocal Imaging of Single Mitochondrial Superoxide Flashes in Intact Heart or In Vivo

Simultaneous Measurement of Mitochondrial Calcium and Mitochondrial Membrane Potential in Live Cells by Fluorescent Microscopy

Ground State Depletion Super-resolution Imaging in Mammalian Cells

Improving the Accuracy of Flow Cytometric Assessment of Mitochondrial Membrane Potential in Hematopoietic Stem and Progenitor Cells Through the Inhibition of Efflux Pumps