Imaging the Mitochondrial Redox State in Primary Neurons Using a Ratiometric Indicator

Abstract

Source: Katsalifis, A., et al. Live Imaging of the Mitochondrial Glutathione Redox State in Primary Neurons using a Ratiometric Indicator. J. Vis. Exp. (2021)

This video demonstrates the imaging of mitochondrial redox state in primary neurons using a ratiometric fluorescent indicator. By applying two excitation wavelengths under a fluorescence microscope, the fluorescence intensity ratio is measured to assess mitochondrial oxidation. The protocol includes N-methyl-D-aspartate (NMDA) stimulation to induce calcium influx and reactive oxygen species (ROS) production, followed by calibration using oxidizing and reducing agents to quantify NMDA-induced mitochondrial oxidation.

Protocol

1. Loading cells with Tetramethylrhodamine ethyl ester perchlorate (TMRE)

NOTE: In this protocol, TMRE is used in non-quench mode at a final concentration of 20 nM. In general, the lowest possible concentration of TMRE that still provides sufficient signal intensity on the microscope of choice should be used. Due to uneven evaporation, the volume of medium in different wells can differ in long-term primary cultures. To ensure consistent TMRE concentration in all wells, do not add TMRE directly to the wells. Instead, replace the medium in each well with the same amount of TMRE-containing medium. The protocol below is designed for primary neurons in 24-well plates containing ~1 mL of medium per well.

1. Working in a tissue culture laminar flow hood, collect 500 µL of medium from each well into a single conical tube.
2. Per well, add 0.5 µL of 20 µM TMRE stock into the conical tube (e.g., 12 µL for 24 wells).
3. Carefully aspirate the remaining medium from the first well and replace it with 500 µL of TMRE-containing medium. Continue, well-by-well, with the remaining wells.
   NOTE: Take care not to let the cells dry out and not to disturb the cells.
4. Return the cells to the incubator and wait for at least 60 min for dye equilibration.
   NOTE: Loading time can be extended to several hours without adverse effects.
5. To ensure consistent TMRE concentration and equilibration throughout the imaging experiment, make sure to include a final concentration of 20 nM TMRE in the imaging buffer and all stimulation solutions.

2. Optimization of scanning confocal microscope settings

NOTE: This step aims to find the best compromise between image quality and cell viability during live imaging. This section describes the optimization of settings for reduction-oxidation-sensitive green fluorescent protein (roGFP) imaging. If multiparametric imaging is performed, similar optimization, including checking for a stable baseline without signs of bleaching or phototoxicity, needs to be performed for the additional indicators.

1. Start the confocal microscope and load standard settings for GFP imaging (488 nm excitation, 505 - 550 nm emission).
2. Set the detector to 12 bits or 16 bits.NOTE: Usually, 8 bits are not sufficient for quantitative imaging.
3. Activate the sequential scan mode and add second sequence/track (405 nm excitation, 505 - 550 nm emission).
4. For both channels, select a pseudocolor lookup table that indicates over- and under-exposed pixels (e.g., GLOW OU).
5. Select an objective that is suitable for the object of interest.
   NOTE: 10x-40x are suitable for single-cell analysis, 63x-100x are suitable for single-mitochondrion analysis.
6. Mount a coverslip with cells into the imaging chamber, add 1 mL of imaging buffer, and place the chamber on the microscope.
7. Use the eyepiece and transmitted light to focus the cells.
   NOTE: Do not use epifluorescence light to locate and focus cells. Even at low power, this will adversely affect the cells.
8. Record images with different pixel formats. Based on these images, select the lowest pixel number that gives an acceptable resolution of the structure of interest.
   NOTE: Typically, 512 x 512 pixels work well for single-cell imaging with 20x and 40x objectives, and 1024 x 1024 or 2048 x2048 pixels typically work well for single-mitochondrion imaging with a 63x objective.
9. Record images with different pinhole sizes. Based on these images, select the largest pinhole size that gives an acceptable resolution of the structure of interest.
   NOTE: Typically, 3-7 airy units work well.
10. Record images with different laser intensities.
    1. Adjust the detector gain and threshold accordingly. Based on these images, select the lowest laser intensity that gives acceptable signal intensity and signal-to-background ratio.
       1. To determine the signal-to-background ratio, measure the signal intensity in a region of interest (ROI) that contains cells or mitochondria (ROI1) and in an ROI without cells or mitochondria (ROI2). Then, divide the intensity of ROI1 by the intensity of ROI2.
          NOTE: Aim for a signal-to-background ratio of >3 and signal intensities of individual ROIs of 200-1,000 for 405 nm excitation with 1-3% laser power and intensities of individual ROIs of 300-1,500 for 488 nm excitation with 1% laser power.
11. Record images with different scan speeds and number of frame averages. Record 4-5 images for each combination of settings. Based on these image series, select the highest speed and lowest average settings that give acceptable image noise and image-to-image variability.
    NOTE: A scan speed of 600 Hz and 1-2 frames for averaging work well in most cases.
12. Using a new coverslip, record a time-lapse series with the optimized settings.
    NOTE: The duration and image interval of the series should resemble those of the planned experiments.
13. At the end of the time-lapse series, add 1 mL of 2x Diamide (DA) solution to the recording chamber. Image for an additional 2 min.
14. Aspirate the imaging buffer using a peristaltic pump or handheld pipette. Add 1 mL of 1x Dithiothreitol (DTT) solution. Image for an additional 5 min.
15. Analyze the time-lapse experiment.
    1. Verify that none of the two channels gets over- or under-exposed during DA- and DTT-treatment with the optimized settings.
    2. Ensure that none of the two channels shows considerable bleaching during the time-lapse recording; aim for <2% loss of intensity between the first and last images.
    3. Verify that the 405:488 ratio does not change considerably during imaging.
16. Repeat the whole procedure in an iterative manner, using several coverslips, until settings that consistently provide acceptable results have been defined.

3. Assessment of basal redox status

1. Start the microscope and load the optimized settings from section 2.
2. Set frame average to 3-5.
3. Mount a coverslip with cells into the imaging chamber, add 1 mL of imaging buffer, and place the chamber on the microscope.
4. Use the eyepiece and transmitted light to focus the cells.
   NOTE: Do not use epifluorescence light to locate and focus cells. Even at low power, this will adversely affect the cells.
5. Switch to scanning mode and use the 488 nm channel in live view to focus and locate cells for imaging.
6. Use the multipoint function to select 3-5 fields of view on the coverslip.
7. Record a baseline image.
8. Add 1 mL of 2x DA solution to the chamber.
9. After 1, 2, and 3 min, use live view to confirm/adjust the focus and then record an image.
   NOTE: Cells are typically fully oxidized after 2 min.
10. Replace the buffer in the imaging chamber with 1 mL of 1x DTT solution.
11. After 3 and 5 min, use live view to confirm/adjust the focus and then record an image.​
    NOTE: Cells are typically fully reduced after 4-5 min.

4. Live imaging of acute treatments

NOTE: The protocol below describes imaging of the mitochondrial redox response to NMDA treatment. Image intervals and duration of the experiment might need to be adjusted for other treatments.

1. Start the microscope and load the optimized settings from section 3.
2. Set the time-lapse interval to 30 s and duration to 25 min.
3. Mount a coverslip with cells into the imaging chamber, add 1 mL of imaging buffer, and place the chamber on the microscope.
   NOTE: To avoid thermal focus drift, leave the cells on the microscope stage for 10-15 min before starting time-lapse imaging.
4. Use the eyepiece and transmitted light to focus the cells.
   NOTE: Do not use epifluorescence light to locate and focus cells. Even at low power, this will adversely affect the cells.
5. Switch to the scanning mode and use the 488 nm channel in live view to focus and locate cells for imaging.
6. Optional: To increase the number of recorded cells per run, use the multipoint function to image 2-3 fields of view per coverslip.
7. Start the time-lapse acquisition and record 5 images as 2 min baseline recording.
8. Add 500 µL of 3x NMDA solution to the chamber (final concentration 30 µM) and record additional 20 images as a 10 min NMDA response.​
   NOTE: Neurons are very sensitive to changes in osmolarity. Therefore, make sure to minimize evaporation of the imaging buffer. For longer treatments, the imaging chamber should be covered with a lid.
9. Add 500 µL of 4x DA solution to the chamber and record 6 more images (3 min maximum calibration).
10. Aspirate the buffer from the imaging chamber and replace it with 1 mL of 1x DTT solution. Record 10 more images (5 min minimum calibration).
11. End the recording and save the image series.

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Reagents
Diamide (DA)	Sigma-Aldrich	D3648
Dithiothreitol (DTT)	Carl Roth GmbH	6908.1
Sucrose	Carl Roth GmbH	4621.1
Tetramethylrhodamine ethyl ester perchlorate (TMRE)	Sigma-Aldrich	87917
Equipment
Imaging chamber	Life Imaging Services (Basel, Switzerland)	10920	Ludin Chamber Type 3 for Ø12mm coverslips
Laser scanning confocal microscope, microscope	Leica	DMI6000
Laser scanning confocal microscope, scanning unit	Leica	SP8
Peristaltic pump	VWR	PP1080 181-4001
Spinning disc confocal microscope, camera	Hamamatsu	C9100-02 EMCCD
Spinning disc confocal microscope, incubationsystem	TokaiHit	INU-ZILCF-F1
Spinning disc confocal microscope, microscope	Nikon	Ti microscope
Spinning disc confocal microscope, scanning unit	Yokagawa	CSU-X1