This work was supported by the Deutsche Forschungsgemeinschaft (BA 3679/5-1; FOR 2289: BA 3679/4-2). A.K. is supported by an ERASMUS+ fellowship. We thank Iris B&#252;nzli-Ehret, Rita Rosner, and Andrea Schlicksupp for the preparation of primary neurons. We thank Dr. Tobias Dick for providing pLPCX-mito-Grx1-roGFP2. Experiments shown in Figure 4 were performed at the Nikon Imaging Center, University of Heidelberg. Figure 2 was prepared with BioRender.com.

<ol>
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The authors declare that they have no conflict of interest.

Quantitative and dynamic measurements of the mitochondrial redox state provide important information about mitochondrial and cellular physiology. Several fluorogenic chemical probes are available that detect reactive oxygen species, &#34;redox stress,&#34; or &#34;oxidative stress.&#34; However, the latter terms are not well-defined and often lack specificity9,17,18. Compared to chemical dyes, Grx1-roGFP2 offers several advantages9,19: (i) as an excitation-ratiometric sensor, it provides inherent normalization for sensor expression level, absolute fluorescence intensity, and cell or organelle density; (ii) as a genetically encoded probe, it can be targeted to organelles or specific subcellular compartments such as presynaptic terminals or postsynaptic dendritic spines; (iii) genetic expression of the sensor obviates the need for dye loading, which in the case of chemical dyes can be stressful for primary neurons; (iv) in contrast to general redox-sensitive chemical dyes, Grx1-roGFP2 is highly specific for the glutathione redox potential; (v) oxidation and reduction of the sensor are reversible, enabling the measurement of transient redox changes; (vi) in contrast to chemical dyes that allow for the detection of relative changes only, Grx1-roGPF2 fluorescence ratios can be calibrated to determine absolute redox potentials in mV9.
This protocol describes dynamic measurements of the mitochondrial glutathione redox potential in primary neurons. Analysis of mitochondria within intact cells has several advantages15. Compared to the study of isolated mitochondria, mitochondria within cells retain physiologically relevant contacts with other organelles (e.g., endoplasmic reticulum-mitochondria contacts, mitochondria-lysosome contacts) and are exposed to cellular concentrations of signaling molecules and metabolites. Compared to the study of intact animals, primary neurons provide better accessibility for genetic and pharmacological manipulations and microscopy. However, for certain questions, analysis of isolated mitochondria will be better suited as this allows precise control over delivered substrates and inhibitors. Notably, mito-Grx1-roGFP2 can also be used in isolated mitochondria20. To study neuronal mitochondria within intact tissues or animals, transgenic fly and mouse lines are available that express mito-Grx1-roGFP2 in the nervous system21,22,23. Notably, the excitation spectrum of roGFP2 is amenable to two-photon excitation, enabling ex vivo and in vivo two-photon imaging in tissue explants and intact animals, respectively23,24. Spectral variants of thiol redox-sensitive fluorescent proteins are also available such as yellow rxYFP-Grx1p25 and red Grx1-roCherry26.These provide additional flexibility for multiplexing and, in principle, enable the simultaneous measurement of redox-responses in different cell types or cellular compartments.
As described in the protocol, the 405:488 nm ratio can be calibrated by measuring maximal and minimal fluorescence ratios after the application of DA and DTT, respectively. This normalization is not essential because the sensor provides a certain degree of inherent normalization due to its ratiometric nature. However, we recommend performing max/min calibration after each imaging run, mostly for two reasons. First, it ensures that sensor responses were not saturated during the experiment. Second, the absolute 405:488 nm ratio is an arbitrary number that depends on the microscope settings of both channels. Even when keeping the same settings, one cannot be sure that laser power and detector performance will stay the same between experiments, especially when these are several weeks apart. One workaround would be to set the average baseline ratio of control cells to 1.0 within each experiment. This allows for relative quantification of treatment-induced redox perturbations in a given experiment but limits the comparability of different experiments. Especially when comparing the basal redox state of cells from different experiments or cell types, it is important to obtain max/min calibrated absolute quantification. Occasionally, the 405:488 nm ratio does not fall below the baseline ratio after DTT treatment, especially if the mitochondria were already in a relatively reduced state at baseline. This typically is a sign of sensor bleaching during prolonged time-lapse experiments. In this case, repeat the experiment with microscope settings that cause less light exposure.
It is paramount to use microscope settings that do not cause bleaching of the sensor or phototoxicity-induced oxidation of mitochondria. Make sure to take sufficient time for protocol optimization before starting actual experiments. Once acceptable settings have been defined, they can be used in subsequent experiments with minimal adjustments.&#160;mito-Grx1-roGFP2 imaging can be performed at room temperature or 35-37 &#176;C on a heated stage. The sensor will work in both cases; however, the kinetics of signaling cascades and enzyme reactions inside the cells will naturally differ based on temperature.&#160;Therefore, kinetics and amplitudes of sensor responses may vary depending on the chosen temperature. The choice of temperature is up to the user, depending on the specific needs of the experiment.
Finally, we would like to point out two limitations of the sensor. First, the intensity of emitted light is considerably lower with excitation at 405 nm than with excitation at 488 nm, often resulting in rather noisy 405 nm images. Higher laser power is needed to improve the signal to noise in the 405 nm channel. However, excitation at 405 nm is typically more phototoxic and generates more autofluorescence than excitation at 488 nm, limiting the permissible 405 nm laser intensity. For example, in this study, weak roGFP fluorescence combined with considerable 405 nm-excited tissue autofluorescence prevented the acquisition of robust data from mito-Grx1-roGFP2-expressing ganglion cells in live retina flatmounts (Depp and Bas-Orth, unpublished observation). Second, although the 405:488 nm ratio is insensitive to pH changes within a physiological range around pH 5.5 to pH 8.5 (Figure 4)9, pH-dependent quenching of roGFP2 emission can cause the typically weak 405 nm signal to drop below the detection limit of the microscope, preventing the acquisition of quantifiable ratio images. For example, this was the case during NMDA-induced acidosis in the neuronal cytosol13.

Several important mitochondrial enzymes and signaling molecules are subject to thiol redox regulation1. Moreover, mitochondria are a major cellular source of reactive oxygen species and are selectively vulnerable to oxidative damage2. Accordingly, the mitochondrial redox potential directly affects bioenergetics, cell signaling, mitochondrial function, and ultimately cell viability3,4. The mitochondrial matrix contains high amounts (1-15 mM) of the thiol-disulfide redox buffer glutathione (GSH) to maintain redox homeostasis and mount antioxidant defenses5,6. GSH can be covalently attached to target proteins (S-glutathionylation) to control their redox status and activity and is used by a range of detoxifying enzymes that reduce oxidized proteins. Therefore, the mitochondrial glutathione redox potential is a highly informative parameter when studying mitochondrial function and pathophysiology.
roGFP2 is a variant of GFP that has been made redox-sensitive by the addition of two surface-exposed cysteines that form an artificial dithiol-disulfide pair7,8. It has a single emission peak at ~510 nm and two excitation peaks at ~400 nm and 490 nm. Importantly, the relative amplitudes of the two excitation peaks depend on the redox state of roGFP2 (Figure 1), making this protein a ratiometric sensor. In the Grx1-roGFP2 sensor, human glutaredoxin-1 (Grx1) has been fused to the N-terminus of roGFP29,10. Covalent attachment of the Grx1 enzyme to roGFP2 affords two major improvements of the sensor: it makes the sensor response specific for the GSH/GSSG glutathione redox pair (Figure 1), and it speeds up equilibration between GSSG and roGFP2 by a factor of at least 100,0009. Therefore, Grx1-roGFP2 enables specific and dynamic imaging of the cellular glutathione redox potential.
Grx1-roGFP2 imaging can be performed on a wide range of microscopes, including widefield fluorescence microscopes, spinning disc confocal microscopes, and laser scanning confocal microscopes. Expression of the sensor in primary neurons can be achieved by various methods that include lipofection11, DNA/calcium-phosphate coprecipitation12, virus-mediated gene transfer, or use of transgenic animals as the cell source (Figure 2). Pseudotyped recombinant adeno-associated viruses (rAAV) containing a 1:1 ratio of AAV1 and AAV2 capsid proteins 13,14 were used for the experiments in this article. With this vector, maximal sensor expression is typically reached 4-5 days after infection and stays stable for at least two weeks. We have successfully used Grx1-roGFP2 in primary hippocampal and cortical neurons from mice and rats.
In this article, rAAV-mediated expression of mitochondria-targeted Grx1-roGFP2 in primary rat hippocampal and cortical neurons is used to assess basal mitochondrial glutathione redox state and its acute perturbation. A protocol is provided for confocal live imaging with detailed instructions on how to (i) optimize laser scanning confocal microscope settings, (ii) run a live imaging experiment, and (iii) analyze data with FIJI.

<table><tbody><tr><td>&lt;strong&gt;reagents&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Calcium chloride (CaCl&lt;sub&gt;2&lt;/sub&gt;&amp;middot;2H&lt;sub&gt;2&lt;/sub&gt;O)</td><td>Sigma-Aldrich</td><td>C3306</td><td></td></tr><tr><td>Diamide (DA)</td><td>Sigma-Aldrich</td><td>D3648</td><td></td></tr><tr><td>Dithiothreitol (DTT)</td><td>Carl Roth GmbH</td><td>6908.1</td><td></td></tr><tr><td>Glucose (2.5 M stock solution)</td><td>Sigma-Aldrich</td><td>G8769</td><td></td></tr><tr><td>Glucose</td><td>Sigma-Aldrich</td><td>G7528</td><td></td></tr><tr><td>Glycine</td><td>neoFroxx GmbH</td><td>LC-4522.2</td><td></td></tr><tr><td>HEPES (1 M stock solution)</td><td>Sigma-Aldrich</td><td>15630-080</td><td></td></tr><tr><td>HEPES</td><td>Sigma-Aldrich</td><td>H4034</td><td></td></tr><tr><td>Magnesium chloride (MgCl&lt;sub&gt;2&lt;/sub&gt;&amp;middot;6H&lt;sub&gt;2&lt;/sub&gt;O)</td><td>Sigma-Aldrich</td><td>442611-M</td><td></td></tr><tr><td>N-methyl-D-aspartate (NMDA)</td><td>Sigma-Aldrich</td><td>M3262</td><td></td></tr><tr><td>Potassium chloride (KCl)</td><td>Sigma-Aldrich</td><td>P3911</td><td></td></tr><tr><td>Sodium chloride (NaCl)</td><td>neoFroxx GmbH</td><td>LC-5932.1</td><td></td></tr><tr><td>Sodium pyruvate (0.1 M stock solution)</td><td>Sigma-Aldrich</td><td>S8636</td><td></td></tr><tr><td>Sodium pyruvate</td><td>Sigma-Aldrich</td><td>P8574</td><td></td></tr><tr><td>Sucrose</td><td>Carl Roth GmbH</td><td>4621.1</td><td></td></tr><tr><td>Tetramethylrhodamine ethyl ester perchlorate (TMRE)</td><td>Sigma-Aldrich</td><td>87917</td><td></td></tr><tr><td>&lt;strong&gt;equipment&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>imaging chamber</td><td>Life Imaging Services (Basel, Switzerland)</td><td>10920</td><td>Ludin Chamber Type 3 for &amp;Oslash;12mm coverslips</td></tr><tr><td>laser scanning confocal microscope, microscope</td><td>Leica</td><td>DMI6000</td><td></td></tr><tr><td>laser scanning confocal microscope, scanning unit</td><td>Leica</td><td>SP8</td><td></td></tr><tr><td>peristaltic pump</td><td>VWR</td><td>PP1080 181-4001</td><td></td></tr><tr><td>spinning disc confocal microscope, camera</td><td>Hamamatsu</td><td>C9100-02 EMCCD</td><td></td></tr><tr><td>spinning disc confocal microscope, incubationsystem</td><td>TokaiHit</td><td>INU-ZILCF-F1</td><td></td></tr><tr><td>spinning disc confocal microscope, microscope</td><td>Nikon</td><td>Ti microscope</td><td></td></tr><tr><td>spinning disc confocal microscope, scanning unit</td><td>Yokagawa</td><td>CSU-X1</td><td></td></tr><tr><td>&lt;strong&gt;software&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>FIJI</td><td></td><td>https://fiji.sc</td><td></td></tr><tr><td>StackReg plugin</td><td></td><td>https://github.com/fiji-BIG/StackReg/blob/master/src/main/java/StackReg_.java</td><td></td></tr><tr><td>TurboReg plugin</td><td></td><td>https://github.com/fiji-BIG/TurboReg/blob/master/src/main/java/TurboReg_.java</td><td></td></tr></tbody></table>

live imaging of the mitochondrial glutathione redox state in primary neurons using a ratiometric indicator

Mitochondrial redox homeostasis is important for neuronal viability and function. Although mitochondria contain several redox systems, the highly abundant thiol-disulfide redox buffer glutathione is considered a central player in antioxidant defenses. Therefore, measuring the mitochondrial glutathione redox potential provides useful information about mitochondrial redox status and oxidative stress. Glutaredoxin1-roGFP2 (Grx1-roGFP2) is a genetically encoded, green fluorescent protein (GFP)-based ratiometric indicator of the glutathione redox potential that has two redox-state-sensitive excitation peaks at 400 nm and 490 nm with a single emission peak at 510 nm. This article describes how to perform confocal live microscopy of mitochondria-targeted Grx1-roGFP2 in primary hippocampal and cortical neurons. It describes how to assess steady-state mitochondrial glutathione redox potential (e.g., to compare disease states or long-term treatments) and how to measure redox changes upon acute treatments (using the excitotoxic drug N-methyl-D-aspartate (NMDA) as an example). In addition, the article presents co-imaging of Grx1-roGFP2 and the mitochondrial membrane potential indicator, tetramethylrhodamine, ethyl ester (TMRE), to demonstrate how Grx1-roGPF2 can be multiplexed with additional indicators for multiparametric analyses. This protocol provides a detailed description of how to (i) optimize confocal laser scanning microscope settings, (ii) apply drugs for stimulation followed by sensor calibration with diamide and dithiothreitol, and (iii) analyze data with ImageJ/FIJI.

Quantification of differences in steady-state mitochondrial redox state after growth factor withdrawal 
To demonstrate the quantification of steady-state differences in mitochondrial redox state, primary neurons grown in standard medium were compared to neurons cultured without growth factors for 48 h before imaging. Growth factor withdrawal results in apoptotic neuronal cell death after 72 h16. Cells were imaged after 48 h to test if this is preceded by changes in mitochondrial redox state. Primary rat cortical neurons grown on poly-L-ornithine-coated coverslips were infected with rAAV-mito-Grx1-roGFP2 on days in vitro 6 (DIV6) and were imaged on DIV12. Live imaging was performed at room temperature according to section 4 of this protocol on an inverted laser scanning confocal microscope equipped with a 40x/1.10 water immersion objective. Confocal settings were pinhole 7 airy units, pixel size 568.7 nm (512 x 512 pixels), scan speed 600 Hz, laser power 405 nm 3%, laser power 488 nm 1%, emission bandwidth 505-550 nm, and frame average 4. There was no major difference between the raw 405:488 nm ratios of the two conditions (Figure 3B). After data normalization, a subset of cells with an increased 405:488 nm ratio was detectable in the growth factor withdrawal group (Figure 3C). This indicates that mitochondrial redox changes might precede neuronal cell death, and it underscores the relevance of max/min data normalization for the comparison of basal redox states between groups.
Dynamic changes in mitochondrial redox state upon treatment of neurons with NMDA 
The NMDA-type glutamate receptor (NMDAR) plays a central role in neuronal plasticity but can also mediate neuronal damage and cell death. Pathological activation of the NMDAR leads to several adverse effects on mitochondria that include matrix calcium overload, mitochondrial oxidation and fragmentation, and mitochondrial permeability transition. In a previous study, the above-described protocol was used to investigate a causal relationship between NMDA-induced mitochondrial calcium overload and mitochondrial oxidation13. Primary rat hippocampal neurons grown on poly-D-lysine/laminin-coated coverslips were infected with rAAV-mito-Grx1-roGFP2 on DIV4 and imaged on DIV12. Live imaging was performed at 37 &#176;C on an inverted spinning disc confocal microscope equipped with a 20x/0.75 multi immersion objective (water immersion was used) and an on-stage incubation system. Mito-Grx1-roGFP2 was sequentially excited every 20 s using the 405 nm and 488 nm laser lines, and emission was collected with a 527/55 nm emission filter for both excitation wavelengths. Treatment of neurons with 30 &#181;M NMDA caused oxidation of mitochondria within a few minutes (Figure 4A,B). Notably, NMDA-induced mitochondrial acidosis caused significant quenching of roGFP2 fluorescence, in line with its well-known pH-sensitivity8. To confirm that this pH-dependent quenching did not affect the 405:488 ratio9, mitochondria were fully oxidized by DA before the addition of NMDA in a control experiment. Pretreatment with DA precludes any further oxidation of mitochondria by NMDA and, accordingly, the 405:488 ratio did not change in this experiment despite a considerable quenching of roGFP2 fluorescence intensity (Figure 4C).
Multiparametric analysis of NMDA-induced changes of dendritic mitochondria 
To assess the temporal sequence of NMDA-induced changes in mitochondrial morphology, membrane potential, and redox state, parallel imaging of TMRE- and mito-Grx1-roGFP2 fluorescence was performed at high spatial and temporal resolution. Primary rat cortical neurons grown on poly-L-ornithine-coated coverslips were infected with rAAV-mito-Grx1-roGFP2 on DIV6 and imaged on DIV12. Live imaging was performed at room temperature on an inverted confocal laser scanning microscope using a 63x/1.40 oil immersion objective, a scan speed of 600 Hz, a pixel size of 90.2 nm (1024 x 1024 pixels at 2x scan zoom), a pinhole size of 3 airy units, and a frame average of 2. Every 30 s, three images were recorded in sequential mode: 405 nm excitation/505-550 nm emission; 488 nm excitation/505-550 nm emission; 552 nm excitation/560-600 nm emission. Treatment of neurons with 60 &#181;M NMDA resulted in a loss of TMRE signal and an increase in the 405:488 nm roGFP ratio, followed by some delayed rounding up of mitochondria (Figure 5).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63073/63073fig01.jpg" /> 
Figure 1: Schematic representation of Grx1-roGFP2 function and roGFP2 excitation spectra. (A) Oxidative stress and the action of antioxidant defense systems oxidize the cellular glutathione pool. Grx1 in the Grx1-roGPF2 fusion protein promotes the rapid equilibration of the roGFP2 redox state with the redox state of the glutathione pool. The redox status of the roGFP2 pool can be assessed by monitoring the ratio of GFP-fluorescence emission at 510 nm after excitation at 405 nm and 488 nm. Reduced species are shown in blue; oxidized species are shown in red. (B) Excitation spectra of fully reduced (blue) and oxidized (red) roGFP2. Upon oxidation of roGFP2, fluorescence emission at 400 nm excitation increases, whereas emission at 490 nm excitation decreases. This figure has been modified from a previous publication13. Excitation spectra in B were drawn based on Figure 1B from 8. Dotted lines in B indicate the wavelengths of commonly used 405 nm and 488 nm laser lines. Abbreviations: GSH = glutathione; GSSG = oxidized glutathione; Grx = glutaredoxin; roGFP = redox-sensitive green fluorescent protein variant. <a href="https://www.jove.com/files/ftp_upload/63073/63073fig01large.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/63073/63073fig02.jpg" /> 
Figure 2: Workflow of the method. Expression of the excitation-ratiometric redox-sensitive fluorescent protein roGFP2 in neurons can be achieved through several methods that include transfection, lipofection, viral gene transfer, and transgenic animals. The sensor can be used to study the neuronal redox state in cultured primary neurons, ex vivo tissue explants, and intact animals. roGFP2 imaging can be performed on a variety of microscopes that include widefield fluorescent microscopes, confocal microscopes, and 2-photon microscopes. Analysis of roGFP2 imaging data can be performed with the freely available software ImageJ/FIJI. Abbreviation: roGFP = redox-sensitive green fluorescent protein variant. <a href="https://www.jove.com/files/ftp_upload/63073/63073fig02large.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/63073/63073fig03.jpg" /> 
Figure 3: Growth factor withdrawal causes oxidation of neuronal mitochondria. (A)&#160;Representative 405:488 nm ratio images of neurons cultured in the presence (+ GF) or absence (- GF) of growth factors for 48 h before imaging. Color-coded scales represent non-normalized 405:488 nm ratios (lower ratios correspond to a reduced state; higher ratios correspond to an oxidized state). Scale bars = 50 &#181;m. (B) Quantification of the 405:488 nm ratio in individual neurons. (C) Max/min calibrated 405:488 nm ratio of individual neurons. Round symbols represent single cells; bar represents mean. N = 40-44 cells from 3 coverslips from one preparation. Abbreviation: GF = growth factor. <a href="https://www.jove.com/files/ftp_upload/63073/63073fig03large.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/63073/63073fig04.jpg" /> 
Figure 4: NMDA-induced oxidation of neuronal mitochondria. (A) Representative 405:488 nm ratio images before and after treatment with 30 &#181;M NMDA and after max/min calibration with DA and DTT. At this magnification, roGFP signal is mostly detected in the soma and proximal dendrites. The color-coded scale represents non-normalized 405:488 nm ratios (lower ratios correspond to a reduced state; higher ratios correspond to an oxidized state). Scale bars = 50 &#181;m. (B) Quantification of the imaging run shown in A. NMDA induces a rapid and sustained mitochondrial oxidation that can be calibrated using DA and DTT. (C) NMDA-induced mitochondrial acidosis causes a drop of GFP fluorescence upon both 405 nm and 488 nm excitation (upper panel). To isolate the pH-driven effect and confirm that the 405:488 nm ratio is pH-insensitive, neurons were first maximally oxidized using DA and subsequently challenged with NMDA in the presence of DA (lower panel). Under these conditions, NMDA still causes mitochondrial acidosis but no further mitochondrial oxidation. Accordingly, although both the 405 nm and 488 nm trace show a pH-driven drop in fluorescence intensity, the 405:488 nm ratio remains stable. This figure is modified from 13. Abbreviations: GFP = green fluorescent protein; roGFP = redox-sensitive green fluorescent protein variant; NMDA = N-methyl-D-aspartate; DA = diamide; DTT = dithiothreitol. <a href="https://www.jove.com/files/ftp_upload/63073/63073fig04large.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/63073/63073fig05.jpg" /> 
Figure 5: NMDA-induced changes in membrane potential, redox state, and morphology of dendritic mitochondria. (A) Three high-magnification images from a time-lapse experiment, acquired at t = 1, 4, and 9 min, showing dendritic and axonal mitochondria. Colorization represents TMRE intensity (see calibration bar). (B) 405:488 nm roGFP ratio images from the same time-lapse experiment as in (A) at t = 1, 4, and 9 min. Note that due to limited AAV infection efficiency, only a subset of neurons expresses mito-Grx1-roGFP2, and therefore, not all TMRE-positive mitochondria are roGFP-positive. The color-coded calibration bar represents non-normalized 405:488 nm ratios (lower ratios correspond to a reduced state; higher ratios correspond to an oxidized state). (C) Quantification of TMRE intensity, roGFP 405:488 nm ratio, and roundness of a single mitochondrion (indicated by arrows in A, B). After 1 min of baseline recording, 60 &#181;M NMDA was added to the bath solution. Scale bars = 5 &#181;m. The y-axis in C depicts the 405:488 nm roGFP ratio relative to baseline T0 (green dashed line), the TMRE fluorescence intensity relative to baseline T0 (red dotted line), and the FIJI/ImageJ shape descriptor &#34;roundness&#34; (black solid line). Abbreviations: GFP = roGFP = redox-sensitive green fluorescent protein variant; mito-Grx1-roGFP2 = Glutaredoxin-1 fused to N-terminus of roGFP; AAV = adeno-associated virus; TMRE = tetramethylrhodamine, ethyl ester; NMDA = N-methyl-D-aspartate. <a href="https://www.jove.com/files/ftp_upload/63073/63073fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Live Imaging of the Mitochondrial Glutathione Redox State in Primary Neurons using a Ratiometric Indicator at JoVE.com

Live Imaging of the Mitochondrial Glutathione Redox State in Primary Neurons using a Ratiometric Indicator

This article describes a protocol to determine differences in basal redox state and redox responses to acute perturbations in primary hippocampal and cortical neurons using confocal live microscopy. The protocol can be applied to other cell types and microscopes with minimal modifications.

Mitochondrial redox homeostasis is important for neuronal viability and function. Although mitochondria contain several redox systems, the highly abundant ...

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Nanyang Technological University

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JoVE Journal

Neuroscience

2.7K Views.  Heidelberg University. This article describes a protocol to determine differences in basal redox state and redox responses to acute perturbations in primary hippocampal and cortical neurons using confocal live microscopy. The protocol can be applied to other cell types and microscopes with minimal modifications.

Video: Live Imaging of the Mitochondrial Glutathione Redox State in Primary Neurons using a Ratiometric Indicator

Neuromodulation and Mitochondrial Transport: Live Imaging in Hippocampal Neurons over Long Durations

To understand the relationship between mitochondrial transport and neuronal function, it is critical to observe mitochondrial behavior in live cultured neurons for extended durations1-3. This is now possible through the use of vital dyes and fluorescent proteins with which cytoskeletal components, organelles, and other structures in living cells can be labeled and then visualized via dynamic fluorescence microscopy. For example, in embryonic chicken sympathetic neurons, mitochondrial movement was characterized using the vital dye rhodamine 1234. In another study, mitochondria were visualized in rat forebrain neurons by transfection of mitochondrially targeted eYFP5. However, imaging of primary neurons over minutes, hours, or even days presents a number of issues. Foremost among these are: 1) maintenance of culture conditions such as temperature, humidity, and pH during long imaging sessions; 2) a strong, stable fluorescent signal to assure both the quality of acquired images and accurate measurement of signal intensity during image analysis; and 3) limiting exposure times during image acquisition to minimize photobleaching and avoid phototoxicity.
Here, we describe a protocol that permits the observation, visualization, and analysis of mitochondrial movement in cultured hippocampal neurons with high temporal resolution and under optimal life support conditions. We have constructed an affordable stage-top incubator that provides good temperature regulation and atmospheric gas flow, and also limits the degree of media evaporation, assuring stable pH and osmolarity. This incubator is connected, via inlet and outlet hoses, to a standard tissue culture incubator, which provides constant humidity levels and an atmosphere of 5-10% CO2/air. This design offers a cost-effective alternative to significantly more expensive microscope incubators that don't necessarily assure the viability of cells over many hours or even days. To visualize mitochondria, we infect cells with a lentivirus encoding a red fluorescent protein that is targeted to the mitochondrion. This assures a strong and persistent signal, which, in conjunction with the use of a stable xenon light source, allows us to limit exposure times during image acquisition and all but precludes photobleaching and phototoxicity. Two injection ports on the top of the stage-top incubator allow the acute administration of neurotransmitters and other reagents intended to modulate mitochondrial movement. In sum, lentivirus-mediated expression of an organelle-targeted red fluorescent protein and the combination of our stage-top incubator, a conventional inverted fluorescence microscope, CCD camera, and xenon light source allow us to acquire time-lapse images of mitochondrial transport in living neurons over longer durations than those possible in studies deploying conventional vital dyes and off-the-shelf life support systems.

We describe a protocol that allows imaging of mitochondria in living neurons via fluorescence microscopy over long durations. Imaging over extended periods is accomplished through lentivirus-mediated expression of a mitochondrially targeted fluorescent protein and use of an inexpensive stage-top incubator that was designed and built in our laboratory.

To understand the relationship between mitochondrial transport and neuronal function, it is critical to observe mitochondrial behavior in live cultured ...

Determination of Mitochondrial Membrane Potential and Reactive Oxygen Species in Live Rat Cortical Neurons

Mitochondrial membrane potential (&Delta;&Psi;m) is critical for maintaining the physiological function of the respiratory chain to generate ATP. A significant loss of &Delta;&Psi;m renders cells depleted of energy with subsequent death. Reactive oxygen species (ROS) are important signaling molecules, but their accumulation in pathological conditions leads to oxidative stress. The two major sources of ROS in cells are environmental toxins and the process of oxidative phosphorylation. Mitochondrial dysfunction and oxidative stress have been implicated in the pathophysiology of many diseases; therefore, the ability to determine &Delta;&Psi;m and ROS can provide important clues about the physiological status of the cell and the function of the mitochondria. 
Several fluorescent probes (Rhodamine 123, TMRM, TMRE, JC-1) can be used to determine &Delta;&psi;m in a variety of cell types, and many fluorescence indicators (Dihydroethidium, Dihydrorhodamine 123, H2DCF-DA) can be used to determine ROS. Nearly all of the available fluorescence probes used to assess &Delta;&Psi;m or ROS are single-wavelength indicators, which increase or decrease their fluorescence intensity proportional to a stimulus that increases or decreases the levels of &Delta;&Psi;m or ROS. Thus, it is imperative to measure the fluorescence intensity of these probes at the baseline level and after the application of a specific stimulus. This allows one to determine the percentage of change in fluorescence intensity between the baseline level and a stimulus. This change in fluorescence intensity reflects the change in relative levels of &Delta;&Psi;m or ROS. In this video, we demonstrate how to apply the fluorescence indicator, TMRM, in rat cortical neurons to determine the percentage change in TMRM fluorescence intensity between the baseline level and after applying FCCP, a mitochondrial uncoupler. The lower levels of TMRM fluorescence resulting from FCCP treatment reflect the depolarization of mitochondrial membrane potential. We also show how to apply the fluorescence probe H2DCF-DA to assess the level of ROS in cortical neurons, first at baseline and then after application of H2O2. This protocol (with minor modifications) can be also used to determine changes in &#x2206;&#x03A8;m and ROS in different cell types and in neurons isolated from other brain regions.

We demonstrate application of the fluorescence indicator, TMRM, in cortical neurons to determine the relative changes in TMRM fluorescence intensity before and after application of a specific stimulus. We also show application of the fluorescence probe H2DCF-DA to assess the relative level of reactive oxygen species in cortical neurons.

Mitochondrial membrane potential (&Delta;&Psi;m) is critical for maintaining the physiological function of the respiratory chain to generate ATP.  A ...

Ratiometric Biosensors that Measure Mitochondrial Redox State and ATP in Living Yeast Cells

Mitochondria have roles in many cellular processes, from energy metabolism and calcium homeostasis to control of cellular lifespan and programmed cell death. These processes affect and are affected by the redox status of and ATP production by mitochondria. Here, we describe the use of two ratiometric, genetically encoded biosensors that can detect mitochondrial redox state and ATP levels at subcellular resolution in living yeast cells. Mitochondrial redox state is measured using redox-sensitive Green Fluorescent Protein (roGFP) that is targeted to the mitochondrial matrix. Mito-roGFP contains cysteines at positions 147 and 204 of GFP, which undergo reversible and environment-dependent oxidation and reduction, which in turn alter the excitation spectrum of the protein. MitGO-ATeam is a F&ouml;rster resonance energy transfer (FRET) probe in which the &epsilon; subunit of the FoF1-ATP synthase is sandwiched between FRET donor and acceptor fluorescent proteins. Binding of ATP to the &epsilon; subunit results in conformation changes in the protein that bring the FRET donor and acceptor in close proximity and allow for fluorescence resonance energy transfer from the donor to acceptor.

We describe the use of two ratiometric, genetically encoded biosensors, which are based on GFP, to monitor mitochondrial redox state and ATP levels at subcellular resolution in living yeast cells.

Mitochondria have roles in many cellular processes, from energy metabolism and calcium homeostasis to control of cellular lifespan and programmed cell ...

Bioengineering

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 ...

Biology

Real-Time Fluorescent Measurement of Synaptic Functions in Models of Amyotrophic Lateral Sclerosis

Before neuronal degeneration, the cause of motor and cognitive deficits in patients with amyotrophic lateral sclerosis (ALS) and/or frontotemporal lobe dementia (FTLD) is dysfunction of communication between neurons and motor neurons and muscle. The underlying process of synaptic transmission involves membrane depolarization-dependent synaptic vesicle fusion and the release of neurotransmitters into the synapse. This process occurs through localized calcium influx into the presynaptic terminals where synaptic vesicles reside. Here, the protocol describes fluorescence-based live-imaging methodologies that reliably report depolarization-mediated synaptic vesicle exocytosis and presynaptic terminal calcium influx dynamics in cultured neurons.
Using a styryl dye that is incorporated into synaptic vesicle membranes, the synaptic vesicle release is elucidated. On the other hand, to study calcium entry, Gcamp6m is used, a genetically encoded fluorescent reporter. We employ high potassium chloride-mediated depolarization to mimic neuronal activity. To quantify synaptic vesicle exocytosis unambiguously, we measure the loss of normalized styryl dye fluorescence as a function of time. Under similar stimulation conditions, in the case of calcium influx, Gcamp6m fluorescence increases. Normalization and quantification of this fluorescence change are performed in a similar manner to the styryl dye protocol. These methods can be multiplexed with transfection-based overexpression of fluorescently tagged mutant proteins. These protocols have been extensively used to study synaptic dysfunction in models of FUS-ALS and C9ORF72-ALS, utilizing primary rodent cortical and motor neurons. These protocols easily allow for rapid screening of compounds that may improve neuronal communication. As such, these methods are valuable not only for the study of ALS but for all areas of neurodegenerative and developmental neuroscience research.

Two related methods are described to visualize subcellular events required for synaptic transmission. These protocols enable the real-time monitoring of the dynamics of presynaptic calcium influx and synaptic vesicle membrane fusion using live-cell imaging of in vitro cultured neurons.

Before neuronal degeneration, the cause of motor and cognitive deficits in patients with amyotrophic lateral sclerosis (ALS) and/or frontotemporal lobe ...

Imaging Mitochondrial Ca2+ Uptake in Astrocytes and Neurons using Genetically Encoded Ca2+ Indicators (GECIs)

Mitochondrial Ca2+ plays a critical role in controlling cytosolic Ca2+ buffering, energy metabolism, and cellular signal transduction. Overloading of mitochondrial Ca2+ contributes to various pathological conditions, including neurodegeneration and apoptotic cell death in neurological diseases. Here we present a cell-type specific and mitochondria targeting molecular approach for mitochondrial Ca2+ imaging in astrocytes and neurons in vitro and in vivo. We constructed DNA plasmids encoding mitochondria-targeting genetically encoded Ca2+ indicators (GECIs) GCaMP5G or GCaMP6s (GCaMP5G/6s) with astrocyte- and neuron-specific promoters gfaABC1D and CaMKII and mitochondria-targeting sequence (mito-). For in vitro mitochondrial Ca2+ imaging, the plasmids were transfected in cultured astrocytes and neurons to express GCaMP5G/6s. For in vivo mitochondrial Ca2+ imaging, adeno-associated viral vectors (AAVs) were prepared and injected into the mouse brains to express GCaMP5G/6s in mitochondria in astrocytes and neurons. Our approach provides a useful means to image mitochondrial Ca2+ dynamics in astrocytes and neurons to study the relationship between cytosolic and mitochondrial Ca2+ signaling, as well as astrocyte-neuron interactions.

Imaging Mitochondrial Ca2+ Uptake in Astrocytes and Neurons using Genetically Encoded Ca2+ Indicators GECIs

This proctocol aims to provide a method for in vitro and in vivo mitochondrial Ca2+ imaging in astrocytes and neurons.

Mitochondrial Ca2+ plays a critical role in controlling cytosolic Ca2+ buffering, energy metabolism, and cellular signal transduction. Overloading of ...

Live-imaging of Mitochondrial System in Cultured Astrocytes

While much attention has been given to mitochondrial alterations at the neuronal level, recent evidence demonstrates that mitochondrial dynamics and function in astrocytes are implicated in cognition. This article describes the method for time-lapse imaging of astrocyte cultures equipped with a mitochondrial biosensor: MitoTimer. MitoTimer is a powerful and unique tool to assess mitochondrial dynamics, mobility, morphology, biogenesis, and redox state. Here, the different procedures for culture, image acquisitions, and subsequent mitochondrial analysis are presented.

This article describes the method for mitochondrial time-lapse imaging of astrocyte cultures equipped with MitoTimer biosensor and the resulting multiparametric analysis of mitochondrial dynamics, mobility, morphology, biogenesis, redox state, and turnover.

While much attention has been given to mitochondrial alterations at the neuronal level, recent evidence demonstrates that mitochondrial dynamics and ...

Measurement of Mitochondrial Membrane Potential In Vivo using a Genetically Encoded Voltage Indicator

Mitochondrial membrane potential (MMP, &#916;&#936;m) is critical for mitochondrial functions, including ATP synthesis, ion transport, reactive oxygen species (ROS) generation, and the import of proteins encoded by the nucleus. Existing methods for measuring &#916;&#936;m typically use lipophilic cation dyes, such as Rhodamine 800 and tetramethylrhodamine methyl ester (TMRM), but these are limited by low specificity and are not well-suited for in vivo applications. To address these limitations, we have developed a novel protocol utilizing genetically encoded voltage indicators (GEVIs). Genetically encoded voltage indicators (GEVIs), which generate fluorescent signals in response to membrane potential changes, have demonstrated significant potential for monitoring plasma membrane and neuronal potentials. However, their application to mitochondrial membranes remains unexplored. Here, we developed protein-based mitochondrial-targeted GEVIs capable of detecting &#916;&#936;m fluctuations in cells and the motor cortex of living animals. The mitochondrial potential indicator (MPI)offers a non-invasive approach to study &#916;&#936;m dynamics in real-time, providing a method to investigate mitochondrial function under both normal and pathological conditions.

This protocol describes the application of mitochondria-targeted genetically encoded voltage indicators (GEVIs). These GEVIs offer a significant advantage over traditional mitochondrial membrane potential dyes by enabling specific, in vivo, and real-time monitoring of mitochondrial membrane potential.

Mitochondrial membrane potential (MMP, &#916;&#936;m) is critical for mitochondrial functions, including ATP synthesis, ion transport, reactive oxygen ...

Quantification of Mitochondrial Membrane Potential and Superoxide Levels

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.

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.

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

Fluorescence-Based Monitoring of Mitochondrial Stress in Astrocytes

Source: Espourteille, J., et al. <a href="https://app.jove.com/v/62957">Live-imaging of Mitochondrial System in Cultured Astrocytes.</a> J. Vis. Exp. (2021)
This video demonstrates the evaluation of mitochondrial health in genetically modified astrocytes, which express a fluorescent protein within their mitochondria. The protein helps to track changes in mitochondrial health as its fluorescence changes from green to red, indicating stress or damage. Images are captured and analyzed using various parameters.

1. Rat hippocampal astrocyte primary culture

	Sacrifice five rat pups (Wistar IGS Rat) by decapitation at postnatal days 1-2.
	Remove the brain and keep ...

Live Imaging of the Mitochondrial Glutathione Redox State in Primary Neurons using a Ratiometric Indicator

Summary

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Chapters in this video

Neuromodulation and Mitochondrial Transport: Live Imaging in Hippocampal Neurons over Long Durations

Determination of Mitochondrial Membrane Potential and Reactive Oxygen Species in Live Rat Cortical Neurons

Ratiometric Biosensors that Measure Mitochondrial Redox State and ATP in Living Yeast Cells

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

Real-Time Fluorescent Measurement of Synaptic Functions in Models of Amyotrophic Lateral Sclerosis

Imaging Mitochondrial Ca²⁺ Uptake in Astrocytes and Neurons using Genetically Encoded Ca²⁺ Indicators (GECIs)

Live-imaging of Mitochondrial System in Cultured Astrocytes

Measurement of Mitochondrial Membrane Potential In Vivo using a Genetically Encoded Voltage Indicator

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

Fluorescence-Based Monitoring of Mitochondrial Stress in Astrocytes