Fluorescence-Based Quantification of Mitochondrial Membrane Potential and Superoxide Levels Using Live Imaging in HeLa Cells (Video) | JoVE

Mohammad Fazli; Chantell  S. Evans

doi:10.3791/65304

Biology

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

Published: May 12, 2023 doi: 10.3791/65304

Mohammad Fazli¹, Chantell S. Evans¹

¹Department of Cell Biology, Duke University Medical Center

Summary

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.

Abstract

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 (Parkin^T240R) 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.

Introduction

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'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 (Parkin^WT), a Parkin mutant (Parkin^T240R), or an empty control vector. Parkin^T240R 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 µM) or severe (20 µM) 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.

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Protocol

1. Preparation of biological samples

NOTE: Perform the following steps using sterile technique in a biosafety cabinet. Spray the surface of the cabinet and all materials with 70% ethanol.

HeLa cell culturing and transfection
1. Culture 30,000 HeLa cells in Dulbecco's modified eagle medium (DMEM) containing 4.5 g/L glucose supplemented with 10% fetal bovine serum and 1% L-glutamine solution (HeLa media; see Table of Materials). Plate the cells on 35 mm glass-bottom imaging dishes (see Table of Materials) containing 2 mL of prewarmed HeLa media. Maintain the HeLa cultures at 5% CO₂ and 37 °C²⁶^,²⁷.
2. The day after plating, inspect the 35 mm imaging dishes using a light microscope to ensure the cells are ~50%-60% confluent. Once confluent, transfect the HeLa cells with empty yellow fluorescent protein (YFP) vector, YFP-Parkin^WT, or YFP-Parkin^T240R (Figure 1A).
3. Prepare two separate mixtures in sterile microcentrifuge tubes for each transfection. Mix by tapping the tube or gently pipetting up and down.
  1. In tube 1, mix 200 µL of reduced serum media (see Table of Materials) with 2 µg of the plasmid DNA.
  2. In tube 2, mix 200 µL of reduced serum media with 6 µL of transfection reagent (see Table of Materials)²⁸.
4. Incubate the tubes for 5 min at room temperature (RT). Add tube 2 to tube 1, mix by pipetting up and down, and incubate for 20 min at RT.
5. Add the transfection complexes dropwise to the existing imaging dishes containing the HeLa cell cultures, ensuring equal distribution across the entire dish. Place the imaging dishes in the 5% CO₂ and 37 °C incubator overnight.
  NOTE: Label each dish with the appropriate transfected plasmid. For each experiment, label the three YFP-Parkin^WT dishes (dimethyl sulfoxide [DMSO; see Table of Materials], 5 µM CCCP, and 20 µM CCCP). Repeat the labeling with the three YFP-Parkin^T240R dishes and the three empty YFP vector dishes.
Preparation of CCCP and fluorescent dyes
CAUTION: Fluorescent dyes are often light-sensitive. Store the dyes in the dark using aluminum foil or brown centrifuge tubes to reduce exposure to light.
1. Make a 5 mM working stock of CCCP (see Table of Materials) by dissolving 5 mg of CCCP in 4.89 mL of DMSO. Make a 20 mM working stock of CCCP by dissolving 5 mg of CCCP in 1.22 mL of DMSO.
2. Dilute 10 µL of a 1 mM MitoTracker Deep Red (see Table of Materials) stock solution in 390 µL of DMSO to make a 25 µM working stock.
3. Dissolve 50 µg of MitoSOX Red²⁵ (see Table of Materials) in 13 µL of DMSO to create a 5 mM working stock.
4. Dissolve 10 mg of TMRE²⁴ (see Table of Materials) in 19.4 mL of DMSO to make a 1 mM stock solution. Dilute the TMRE by adding 10 µL of 1 mM TMRE into 990 µL of DMSO to make a 10 µM working stock.

2. CCCP treatment and mitochondrial labeling with fluorescent probes in HeLa cells

CAUTION: Perform the following steps quickly to ensure the HeLa cells are not out of the incubator for an extended period.

The day after transfection, add 2 µL of either 5 or 20 mM CCCP (final concentration: 5 µM and 20 µM, respectively) to each experimental plate. For control plates, add 2 µL of DMSO. Return the plates to the 5% CO₂ and 37 °C incubator for 1.5 h (Figure 1A).
NOTE: The CCCP treatment is for a total of 2 h. However, the mitochondria are labeled with fluorescent probes during the final 30 min of the CCCP treatment.
After 1.5 h, remove the plates from the incubator and add fluorescent probes to the imaging dishes. Return the plates to the 5% CO₂ and 37 °C incubator for 30 min (Figure 1A).
1. TMRE experiment: Add 2 µL of 25 µM MitoTracker (final concentration: 25 nM) and 2 µL of 10 µM TMRE (final concentration: 10 nM).
2. MitoSOX experiment: Add 2 µL of 25 µM MitoTracker (final concentration: 25 nM) and 1 µL of 5 mM MitoSOX (final concentration: 2.5 µM).
After the 2 h CCCP treatment, prepare the HeLa cells for imaging (Figure 1A,B).
1. TMRE experiment: HeLa cells are immediately ready to image; ensure that the TMRE remains in the HeLa cell culture media.
2. MitoSOX experiment: Wash the cells 3x with 2 mL of prewarmed HeLa media to remove free MitoSOX dye. Add 2 mL of prewarmed media. The HeLa cells are now ready to image.
  NOTE: Wash the HeLa cells with fresh, prewarmed HeLa media that contains the same DMSO or CCCP concentration as the experimental condition; do not include MitoSOX or MitoTracker.

3. Confocal microscope image acquisition setup

NOTE: Image the HeLa cells using a confocal microscope (see Table of Materials) equipped with a 63x/1.40 numerical aperture (NA) oil immersion objective and an environmental chamber (see Table of Materials).

At 1 h before imaging, turn on the CO₂ by opening the tank valve. Press the On button to turn on the environmental controller for the microscope. Use the Up and Down Arrows on the touchpad to adjust the temperature to 37 °C and the CO₂to 5%. Press Set when complete.
NOTE: Ensure the doors to the environmental chamber are closed and wait for the conditions to stabilize.
Laser settings
1. Turn on the White Light Laser (WLL) and set the laser power to 85% and the excitation control to maximum power. Click the Acquire tab, select Add Laser, and in the dialog box that appears, toggle the WLL to On. Click the Laser Power button and enter 85%. Click the Excitation Control button and select Maximum Power from the dropdown menu (Figure 2A).
2. TMRE experiment: Set the excitation and emission spectra for YFP, MitoTracker, and TMRE.
  1. For YFP, set the excitation laser to 514 nm and the emission spectra window to 524-545 nm. In the Acquire tab, click the Add New Setting button; then, click Add Laser and drag it into Setting 1. Double-click on the Excitation Line and enter 514 as the wavelength in the dialog box. Double-click on the corresponding detector and enter 524 for the beginning and 545 for the end wavelength.
  2. For MitoTracker Deep Red, set the excitation laser to 641 and the emission spectra window to 650-750 nm. In the Acquire tab, click the Add Laser button and drag it into Setting 1. Double-click on the Excitation Line and enter 641 as the wavelength in the dialog box. Double-click on the corresponding detector and enter 650 for the beginning and 750 for the end wavelength.
  3. For TMRE, set the excitation laser to 555 nm and the emission spectra window to 557-643 nm. In the Acquire tab, click the Add New Setting button; then, click the Add Laser button and drag it into Setting 2. Double-click on the Excitation Line and enter 555 as the wavelength in the dialog box. Double-click on the corresponding detector and enter 557 for the beginning and 643 for the end wavelength.
3. MitoSOX experiment: Set the excitation and emission spectra for YFP, MitoTracker, and MitoSOX (Figure 2B).
  1. For YFP, set the excitation laser to 507 nm and the emission spectra window to 517-540 nm. In the Acquire tab, click the Add New Setting button; then, click the Add Laser button and drag it into Setting 1. Double-click on the Excitation Line and enter 507 as the wavelength in the dialog box. Double-click on the corresponding detector and enter 517 for the beginning and 540 for the end wavelength.
  2. For MitoSOX, set the excitation laser to 547 nm and the emission spectra window to 564-636 nm. In the Acquire tab, click the Add New Setting button; then, click the Add Laser button and drag it into Setting 2. Double-click on the Excitation Line and enter 547 as the wavelength in the dialog box. Double-click on the corresponding detector and enter 564 for the beginning and 636 for the end wavelength.
  3. For MitoTracker Deep Red, set the excitation laser to 641 nm and the emission spectra window to 652-742 nm. In the Acquire tab, click the Add New Setting button; click the Add Laser button and drag it into Setting 3. Double-click on the Excitation Line and enter 641 as the wavelength in the dialog box. Double-click on the corresponding detector and enter 652 for the beginning and 742 for the end wavelength.
Image acquisition settings (Figure 2C)
1. Set the format to 1,024 x 1,024, the scanning speed to 600 Hz, and the line average to 3.
  1. Select the Acquisition tab and click the Format button. From the dropdown menu, select 1,024 x 1,024. Click the Speed button and select 600 from the dropdown menu. Then, click the Line Average button, and from the dropdown menu, select 3.
  2. Turn Bidirectional Scanning on and set the phase and zoom factor to 22.61 and 1.50, respectively.
    1. In the Acquisition tab, toggle the Bidirectional button to On. Click on the Phase X setting and set it to 22.61. Click on the Zoom Factor setting and set it to 1.50.

4. Image acquisition

CAUTION: The experimenter must make a visual judgment to select the cells based on the YFP fluorescence signal. Avoid oversaturated pixels, as they can significantly affect the fluorescence intensity quantification. Use an over/under look-up table that indicates pixel saturation to avoid acquiring saturated images.

Click on the Setting of interest and press Fast Live to provide a live preview of the fluorescence image.
Adjust the gain and intensity by double-clicking on the corresponding detector. In the dialog box that appears, adjust the Gain using the slider. To alter the intensity, double-click on the Excitation Line and use the Up and Down Arrows in the popup window to change the intensity. Click Stop to end the preview.
1. TMRE experiment: Image the DMSO control plate first. Adjust the gain and intensity of the TMRE signal (Setting 2) so that the mitochondrial network intensity is just below saturation; keep the gain and intensity constant for the experiment. Adjust the gain and intensity of the MitoTracker and YFP (Setting 1) so that the mitochondrial network is visible but dim.
2. MitoSOX experiment: Image the DMSO control plate first. Adjust the gain and intensity of the MitoSOX (Setting 2) signal so that the fluorescence is visible but dim; keep the gain and intensity constant for the experiment. Adjust the gain and intensity of the YFP (Setting 1) and MitoTracker (Setting 3) so that the mitochondrial network is visible but dim.
  NOTE: Record the gain and intensity of TMRE and MitoSOX, as these values must remain constant throughout the experiment. The fluorescence signals of YFP and MitoTracker are not quantified in this protocol. Therefore, the gain and intensity can be adjusted for each image. It is most effective to have the gain and intensity settings where the cells are easy to see but still dim, to ensure the cells are not exposed to excessive laser intensities that cause cell damage or photobleaching.
Once the gain and intensity settings are complete, click Start to acquire an image. Acquire images of 20 cells per experimental condition (e.g., five images with four cells per image; Figure 2D).

5. Fluorescence intensity quantification using ImageJ

NOTE: Imaging files are saved as ".lif" and are compatible with ImageJ²⁹. Compatible file types with ImageJ are specified on their website. It may be necessary to convert the file type if it is incompatible.

Create and save a region of interest (ROI).
1. In ImageJ, click File | New | Image. In the dialog box that appears, click OK.
2. Click the Rectangle Tool button and draw a 6 micron x 6 micron box. Open the ROI manager by clicking Analyze | Tools | ROI Manager and wait for a dialog box with the ROI manager to appear (Figure 3A). Add the rectangular ROI to the manager by clicking Add on the manager dialog box. Save the ROI by selecting More, then click the Save button and OK.
Measure the fluorescence intensity.
1. In Image J, click File and select Open. In the dialog box, select the imaging files for the experiment and click Open.
2. Wait for the Bio-formats Import Options window to appear (Figure 3B). Select Split Channels and click Open.
  NOTE: The split channels feature allows each channel in the image to be opened as a separate window. The channel order corresponds to the acquisition order.
  1. TMRE experiments: YFP (Setting 1) is the first channel (c = 0); MitoTracker (Setting 1) is the second channel (c = 1); and TMRE (Setting 2) is the third channel (c = 2).
  2. MitoSOX experiments: YFP (Setting 1) is the first channel (c = 0); MitoSOX (Setting 2) is the second channel (c = 1); and MitoTracker (Setting 3) is the third channel (c = 2).
3. Adjust the brightness by selecting Image | Adjust | Brightness (Figure 3C).
  NOTE: Do not press Set or Apply to ensure the image brightness is not permanently altered. Occasionally, the fluorescence intensity is not visible in the TMRE or MitoSOX channels. Adjust the brightness to allow for easier visualization. Altering the brightness does not affect the raw fluorescence intensity values.
4. Click Analyze and select Set Measurements. Check the Area and Mean gray value boxes and click OK (Figure 3D).
  NOTE: The mean gray value is the fluorescence intensity.
5. Open the ROI Manager and load the ROI saved in step 5.1 by clicking Analyze | Tools | ROI Manager. In the ROI Manager, click More, then Open from the list that appears, and select the saved ROI.
6. Measure the fluorescence intensity of five random regions in a single cell by selecting the saved ROI from the ROI manager and move the ROI to a random location within a cell (Figure 3E). Measure the fluorescence intensity by pressing M. Repeat this step with four additional nonoverlapping regions. When a dialog box with the area and mean gray values appears (Figure 3F), copy and paste the values into a spreadsheet for analysis.
  1. TMRE experiment: Select image (c = 2).
  2. MitoSOX experiment: Select image (c = 2).
7. Obtain the mean fluorescence intensity values. Using spreadsheet software (see Table of Materials), average the five mean gray values. Repeat this process for each cell.
8. Analyze and compare the TMRE or MitoSOX mean gray values across experimental conditions using a statistical software program (see Table of Materials; Figure 4 and Figure 5). Present data as bar graphs or violin plots.

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Representative Results

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, Parkin^WT, or Parkin^T240R (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 µM) or severe (20 µM) concentrations of CCCP to depolarize the mitochondrial network (step 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 µM) CCCP treatment induced superoxide production and increased the MitoSOX fluorescence intensity (Figure 5A,B). In mild (5 µM) CCCP stress conditions, the expression of Parkin^WT and Parkin^T240R resulted in higher TMRE intensity compared to the empty YFP control vector. Similarly, the MitoSOX intensity was lower in cells expressing Parkin^WT and Parkin^T240R 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.

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. (B) Representative images of cells expressing an empty YFP vector (top), YFP-Parkin^WT (middle), and YFP-Parkin^T240R (bottom; magenta). Hoechst 33342 (white) is used to label the DNA. Scale bar = 10 µm. Abbreviations: CCCP = carbonyl cyanide m-chlorophenyl hydrazone; TMRE = tetramethylrhodamine-ethyl ester-perchlorate; YFP = yellow fluorescent protein. Please click here to view a larger version of this figure.

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 µm. Abbreviations: WLL = white light laser; YFP = yellow fluorescent protein. Please click here to view a larger version of this figure.

Figure 3: ImageJ workflow to quantify TMRE and MitoSOX fluorescence intensities. (A) 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. (D) Set measurements parameter. The red rectangles highlight the Area and Mean gray value options that should be selected. (E) Representative image of a HeLa cell labeled with MitoSOX. The red square illustrates the ROI from panel A. Scale bar = 10 µm. (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. Please click here to view a larger version of this figure.

Figure 4: TMRE fluorescence intensity after mitochondrial damage. (A) Representative images of HeLa cells following a 2 h treatment with DMSO, 5 µM CCCP, or 20 µM CCCP to induce mitochondrial damage. Cells are exogenously expressing an empty YFP vector, YFP-Parkin^WT, or YFP-Parkin^T240R (magenta), and labeled with MitoTracker (cyan) and TMRE (green). Scale bar = 30 µm. (B) Quantification of TMRE fluorescence intensity for cells expressing empty YFP vector (blue), YFP-Parkin^WT(orange), and YFP-Parkin^T240R(purple) in DMSO- and CCCP-treated conditions. ****p < 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 µM CCCP (D), and 20 µM CCCP (E). * p < 0.05; ***p < 0.001; ****p < 0.0001 by Kruskal-Wallis ANOVA with Dunn's multiple comparison test. Ns = not significant. Mean ± 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. Please click here to view a larger version of this figure.

Figure 5: MitoSOX fluorescence intensity following damage to the mitochondrial network. (A) Representative images of HeLa cells treated for 2 h with DMSO, 5 µM CCCP, or 20 µM CCCP to uncouple the mitochondria membrane potential. Cells were transfected with empty YFP vector, YFP-Parkin^WT, or YFP-Parkin^T240R (magenta), and labeled with MitoTracker (cyan) and MitoSOX (green). Scale bar = 30 µm. (B) Quantification of MitoSOX fluorescence intensity for cells expressing empty YFP vector (blue), YFP-Parkin^WT(orange), and YFP-Parkin^T240R(purple) for control and treated conditions. ****p < 0.0001 by two-way ANOVA with Dunn'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 µM CCCP (D), and 20 µM CCCP (E). *p < 0.05; ****p < 0.0001 by Kruskal-Wallis ANOVA with Dunn's multiple comparison test. Ns = not significant. Mean ± 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. Please click here to view a larger version of this figure.

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Discussion

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-Parkin^WT, or YFP-Parkin^T240R. 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 µg 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-Parkin^WT and mCherry-Parkin^T240R, 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'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.

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Disclosures

The authors have no competing interests to declare.

Acknowledgments

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.

Materials

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

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References

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