This study was supported by a Synapsis Foundation fellowship awarded to K.R. and the Lausanne University Hospital (CHUV). The authors thank Nikon for their help, in particular J. Gannevat.

The present protocol has been performed with the approval of an ethical committee (agreement VD3602, Lausanne, Switzerland) and follows European guidelines for the use of animals.
1. Rat hippocampal astrocyte primary culture
<ol>
	<li>Sacrifice five rat pups (Wistar IGS Rat) by decapitation at postnatal day 1-2.</li>
	<li>Remove the brain and keep it in a Petri dish containing 5 mL of fresh astrocyte medium (DMEM with GlutaMAX supplemented with 1% Penicillin/Streptomycin and 10% fresh Horse Serum).</li>
	<li>Isolate the hippocampus. Dissociate in 5 mL of the astrocytic medium by three passages through a 21 G needle and three passages through a 25 G needle.</li>
	<li>Transfer the dissociated cells to a 15 mL centrifuge tube and count in a hemocytometer.</li>
	<li>Plate 20,000-25,000 cells/cm2 in multiwell dishes (9.4 mm x 10.7 mm x 9.3 mm) and store them at 37 &#176;C under an atmosphere containing 5% CO2 for the rest of the experiment.</li>
	<li>Replace the medium entirely at 3 days in vitro (DIV3).</li>
	<li>At DIV8, add 0.6 pg of p24 antigen per cell of lentiviral vector coding for mitochondrial biosensor MitoTimer (LV-G1-MitoTimer22) diluted in phosphate-buffered saline (PBS + 0.01% BSA).</li>
	<li>At DIV9, wash with pre-warmed 1x sterile PBS (37 &#176;C) and add fresh astrocyte medium.</li>
	<li>At DIV11, perform 2 washes with pre-warmed 1x sterile PBS (37 &#176;C) and add fresh astrocyte medium without phenol red. 
	&#8203;NOTE: Choice of the coating depends on the intended assay. As a standard coating for astrocyte primary culture, it is recommended to use 0.2 mg/mL poly-D-lysine or 8.7 &#181;g/cm2 of basement membrane matrix (e.g., Matrigel). To visualize the mitochondrial system of astrocytes, it is essential to work on flattened and stretched cells. In this context, the combination of basement membrane matrix on IBIDI u-slides is the most suitable. It is also crucial not to work with phenol red, which is toxic for the cell under repeated exposure to light.</li>
</ol>
2. Long-term monitoring of the mitochondrial system.
<ol>
	<li>Assess the astrocytic mitochondrial system a minimum of 3-5 days after the lentiviral infections with LV-G1-MitoTimer (to allow a sufficient level of fluorescence in the mitochondria).</li>
	<li>Image the cells with an inverted microscope controlled by the acquisition and analysis software and a module for full automation of acquisition. 
	NOTE: Refer to the Table of Materials for the details of the software and module used in this study.</li>
	<li>Ensure that the microscope is equipped with a cage incubator (see Table of Materials) to maintain astrocyte culture at 5% CO2 and 37 &#176;C throughout the experiment.</li>
	<li>Capture fluorescence images using sequential excitation at 490 nm (for green channel) and 550 nm (for red channel) with detection of green (500-540 nm for green channel) and red (550-600 nm for red channel) fluorescence signals.</li>
	<li>Select 5 astrocytes per well with a mitochondrial network expressing sufficient levels of LV-G1-MitoTimer using a magnification of 40x. Take care to select astrocytes as flat and large as possible and not located in clusters of cells (to work on individual cells).</li>
	<li>Save the coordinates of the 5 selected cells in an xlm file (map.xlm). This map.xlm allows the user to return to the same cells over time.</li>
	<li>Acquire image sequences (1 image/s for 60 s) using a magnification of 150x (100x oil immersion objective, 1.5x intermediate magnification) for each coordinates.</li>
	<li>Repeat image acquisition (1 image/s for 60 s) for the same astrocytes 6 h, 12 h, and 24 h after treatments. 
	&#8203;NOTE: Use a microscope equipped with a fast and efficient autofocus system. In this context, the hardware solution called Perfect Focus System (PFS) was used. PFS uses near-infrared 870-nm LED and CCD line sensors to combat axial focus fluctuations in real-time during long-term imaging investigations.</li>
</ol>
3. Analysis of individual mitochondrial morphology and LV-G1-MitoTimer ratio
NOTE: NIS General Analysis 3 (GA3) from Nikon was used to automatize morphometric analysis in this study.
<ol>
	<li>For each image sequence (baseline, 6 h, 12 h, 24 h), select the first frame for red and green channels by clicking on ND Processing &#62; Select Frame.</li>
	<li>Merge the red and the green channels by selecting Conversions &#62; Merge Channel.</li>
	<li>To correct image shading, select Preprocessing &#62; Auto Shading Correction.</li>
	<li>Apply the Rolling Ball algorithm by selecting Preprocessing &#62; Rolling Ball.</li>
	<li>To generate binary masks for each mitochondrion, select Segmentation &#62; Threshold.</li>
	<li>Remove any objects truncated by the border by selecting Binary processing &#62; Touching Border.</li>
	<li>To measure the surface area, select Measurement &#62; Object Area.</li>
	<li>To measure the diameter, select Measurement &#62; Eq Diameter.</li>
	<li>To measure the length, select Measurement &#62; Length.</li>
	<li>To measure the width, select Measurement &#62; Width.</li>
	<li>To measure roughness, select Measurement &#62; Roughness.</li>
	<li>To measure circularity, select Measurement &#62; Circularity.</li>
	<li>To measure elongation, select Measurement &#62; Elongation.</li>
	<li>Compose a group (right-click) with the above measurement and rename it as MorphoData.</li>
	<li>Measure mean green intensity by selecting Measurement &#62; Mean Intensity.</li>
	<li>Measure mean red intensity by selecting Measurement &#62; Mean Intensity.</li>
	<li>To measure the Red/Green ratio, select Measurement &#62; Ratio.</li>
	<li>Compose a group (right-click) with the above measurement and rename it as RatioData.</li>
	<li>Export the table to a CSV file by selecting Reference &#62; Table to CSV.</li>
	<li>Save the GA3 script of analysis by selecting Save As 
	NOTE: A non-exhaustive list of the criteria used in this procedure is summarized in Figure 1, Table 1, and Table 2. The analysis GA3 script file is available in supplemental (Supplemental Coding File 1 and Figure 2). The thresholds used were adapted to individualize as many mitochondria as possible (excluding large mitochondrial networks where possible). Wherever possible, mitochondrial networks are either individualized manually or excluded from the analyses. Due to intercellular variability, perform these analyses on at least 20-25 cells per condition (with a minimum of 50 mitochondria by cell).</li>
</ol>
4. Analysis of mitochondrial motility
NOTE: Due to the high complexity of mitochondrial movements, manual motility analysis is preferred. Here, Nikon&#39;s NIS Element system was used to manually track mitochondria.
<ol>
	<li>Open the tracking module in NIS, select View &#62; Analysis &#62; Tracking.</li>
	<li>Click on Define New ROI.</li>
	<li>With the Automatic Detection Tool, select 25-50 mitochondria on the first image of the image sequence.</li>
	<li>Click on Track Autodetected ROIs Analyze.</li>
	<li>If necessary, delete the incorrect ROI tracks.</li>
	<li>Export the table to a CSV file. 
	&#8203;NOTE: A non-exhaustive list of the criteria used in this procedure is summarized in Figure 1 and Table 3. A tracking analysis generally gives a path depicting the movements of the center of every tracked object. The different tracking options must be set in the tracking option. Favor the selection of isolated mitochondria that are sufficiently distant from each other to facilitate tracking. Keep only the objects that could consistently be tracked over the whole sequence. Proceed the same way to remove outliers due to bad quality tracking. Consequently, the set of objects analyzed in this section is different from the one analyzed for static morphological analyses and will generally be smaller.</li>
</ol>
5. Data transformation, normalization, and statistical analysis
NOTE: Mainly due to the high heterogeneity of the mitochondria, data generated often have a non-normal distribution.
<ol>
	<li>Manually log-transform the measurements prior to processing.</li>
	<li>For each analysis and time frame, manually normalize the resulting data by the mean of those obtained in the reference acquisition.</li>
	<li>In addition, consider a second/double normalization to a control condition, for example, to evaluate the effect of a treatment.</li>
	<li>Then, perform statistical analysis using a two-way matched ANOVA. Here GraphPad Prism V8 was used.</li>
</ol>

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The authors declare no competing interests.

Here, a novel method to longitudinally follow the dynamics and turnover of the mitochondrial system in a cultured astrocyte is proposed. Unlike a time-lapse approach on a fixed group of cells or one individual cell at a time (most often used in the literature)24,25, researchers can follow the evolution of the mitochondrial system during several days on the same individual cells. In contrast to single well live imaging where high levels of light exposure are required, and the selection of many individual cells is less feasible, the proposed method takes advantage of this microscope&#39;s ability to image several different cells in different areas of a well and to come back to those same cells at various time points to re-image them. Thanks to normalization to a baseline carried out for each measured criterion on each cell of interest, it takes the mitochondrial system&#39;s complexity into account and investigates the effect of treatment on each cell relative to its own baseline image. The microscope&#39;s ability to autonomously carry out this type of imaging on up to 16 wells at a time (imaging 5 cells per well) allows for the heterogeneity of the mitochondrial system to be properly taken into account during analysis without the experimental variability that comes with imaging various conditions on different days.
The quality of the cultures, the levels of viral infections expressing the LV-G1-MitoTimer biosensor, the type of microscope and objectives, and the selection of suitable cells are critical variables that must remain as consistent as possible in this protocol. The cell densities, the type of vector, and the viral titers can be adapted according to the question. Although previous work shows that LV-G1-MitoTimer expression has no deleterious consequences for mitochondrial function and dynamics21,22,26,27, it is essential to verify that the concentration is not toxic for the cells (for example, checking the total number of cells in control well). As a single focal plane is used, astrocytes should be: (1) as flat as possible, (2) isolated from other labeled cells (to simplify the analysis in case of displacement in the dish), and (3) possessing high fluorescence levels. As cells in culture can be highly variable in morphology, the mitochondrial system can be highly heterogeneous. In this context, analyzing ROIs (and not the whole cell) compensates for some problematic regions, such as the perinuclear regions, and decreases variability. It is essential to do the baseline on relatively similar cells and sample as many cells as possible. Consequently, high content acquisition and analysis microscopes are ideally suited. During this longitudinal monitoring, it is also important not to overexpose the cells to light to avoid biosensor bleaching.
This imaging method is not without its complexities, and throughout the protocol, several notes are included, which take into account troubleshooting done during previous tests with the microscope. For example, the choice of plate coating used depends on the intended assay, but recommendations for the most suitable choices for astrocyte primary cultures have been included. Additionally, image acquisition should be performed on at least 5 cells per condition due to intercellular variability. More specifically, some cells selected at baseline imaging will die, some will move out of the frame of the assigned image acquisition area, and some will change their morphology, making the mitochondria very difficult to individualize in analysis. Imaging many cells from the beginning increase the likelihood of a large enough sample size of cells to analyze at the end of the experiment. In addition to the more complex aspects of this imaging technique, there are some outright limitations as far as who can benefit from this type of imaging and analysis. In order to take full advantage of the automatization of image acquisition, the microscope used must have an autofocus system that can handle the speed of time intervals between images (i.e., every 3 s in this protocol) and can consistently focus on the cell in question before each image is taken. Additionally, without the JOBS software, which automates the entire image acquisition process, this method becomes arduous and potentially impossible depending on the number of cells being imaged as it would require manually finding and imaging each cell again at the appropriate time point. Finally, this imaging method is not immune to the issue of photobleaching. For this reason, as with any long-term acquisition method, it is important to choose fluorescent markers that are less susceptible to photobleaching and to tailor image acquisition to avoid this issue as much as possible.
This technique differs from others currently used in a crucial way. Unlike other time-lapse studies, this technique does not require imaging on the same position in the well the entire time, nor does it require manual movement of the plate to image other areas. This allows researchers the ability to image many cells in many conditions in one 24 h timeframe. Consequently, the ability to carry out this imaging and analysis on many cells in each well gives the same population information one would obtain from broadly studying a large group of cells while additionally providing specific measures from each cell imaged. While some specificities to this method may not apply to other image acquisition methods (outlined above), the benefits outweigh the complications with the type of analysis possible after acquisition. This technique allows researchers to see the exact ramifications of various treatments on the mitochondrial system, and consequently, on the cultured astrocytes.
Additionally, this method is highly customizable to many different scientific questions regarding mitochondrial behavior and roles in specific contexts. Here the outlined protocol deals specifically with cultured astrocytes. However, many other cell types can be used, and the treatments that can be tested are limited only by the questions being investigated. This type of imaging has the potential to advance the collective knowledge and understanding of mitochondrial behavior, the underlying mechanisms that lead to mitochondrial dysfunction, and the effects of many pathologies on the innate dynamics present in different types of cells.

Astrocytes are critical players in the maintenance of brain homeostasis. They are perhaps most well known to have significant structural roles in the brain, as part of the blood-brain barrier1 and by supporting neurons and synapses throughout the brain2. Astrocyte support of neurons is both structural3 and metabolic4,5, with astrocytes promoting neurogenesis and synaptogenesis while also providing key metabolites like lactate to active neurons4,6,7. Beyond the role of structural support, astrocytes are active cells that take part in Ca2+ signaling and buffering (including spontaneous mitochondrial Ca2+ influxes)8,9, K+ buffering10, and can adapt and react to the needs of the brain in times of injury11,12. Being such dynamic cells, astrocytes have robust energy requirements, which necessitate an efficient mitochondrial network. These mitochondria also have a crucial role in buffering excessive reactive oxygen species (ROS)13. In addition to their individual or local roles of energy generation and ROS buffering, mitochondria function as a network14. In this sense, they maintain equilibrium between fissioning and fusioning mitochondria, representing new/reduced mitochondria and older/oxidized mitochondria, respectively15. The overall redox state of a cell can be gauged by the redox state of the mitochondrial network. In pathology, this is a critical piece of information that can shed light on which cells may not be functioning optimally.
In recent years, many sensors have been developed to study the dynamics and functions of mitochondria in cells. For example, sensors measuring energy exchange (ATP), redox state (NADH/NAD+, ROS), and enzymatic functionality (cAMP, Ca2+, Zn2+) are currently used in the study of mitochondrial function16. Among them, MitoTimer permits to follow the changes in mitochondrial morphology (size, shape, surface area), mobility (speed, displacement), and dynamics (fusioning and fissioning events), as well as the overall mitochondrial turnover rate and redox state. MitoTimer is a mutant red fluorescent protein, drFP58317, with a mitochondrial signal from subunit VIII of human cytochrome c oxidase18,19 to visualize newly synthesized mitochondria in green (488 nm) and oxidized mitochondria in red (555 nm). Using the green (488 nm) and red (555 nm) fluorescence ratio permits simultaneous evaluation of individual mitochondria, their morphology analysis, fusion/fission events, and redox state history20,21. This unique property can be used to investigate many scientific questions regarding mitochondria&#39;s physiological and pathological roles and is therefore very promising for unveiling the underlying mechanisms of mitochondrial dynamics within many different cell types.
We recently developed a new lentiviral vector (LV-G1-MitoTimer-MiR124T, hereafter called LV-G1-MitoTimer) to study mitochondria&#39;s dynamic and functions specifically in astrocytes in vitro and in vivo22. LV-G1-MitoTimer uses a truncated version of the glial fibrillary acidic protein (GFAP) promoter gfaABC1D, with a B3 enhancer (gfaABC1D(B3), hereafter called G1) combined with the previously described miR124T neuronal detargeting system23. It allows exclusive expression of the mitochondrial biosensor in astrocytes in vitro and in vivo22. Presented here are the different steps to perform a culture of rat hippocampal astrocytes and equip them with the LV-G1-MitoTimer biosensor, as well as the different microscopy steps to follow the behavior of astrocyte mitochondria during several consecutive hours/days.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>&#181;-Slide 8 Well</td>
			<td>IBIDI</td>
			<td>80807</td>
			<td></td>
		</tr>
		<tr>
			<td>19 G needle</td>
			<td>Plexus SANTE</td>
			<td>PL001213</td>
			<td></td>
		</tr>
		<tr>
			<td>21 G needle</td>
			<td>Plexus SANTE</td>
			<td>PL000142</td>
			<td></td>
		</tr>
		<tr>
			<td>25 G needle</td>
			<td>Plexus SANTE</td>
			<td>PL000133</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovin Serum Albumin</td>
			<td>LIFE TECH</td>
			<td>15260037</td>
			<td></td>
		</tr>
		<tr>
			<td>Camera</td>
			<td>HAMAMATSU</td>
			<td>ORCA-flash4.0 V3 - C13440-20CU</td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM, high glucose, GlutaMAX(TM)</td>
			<td>THERMOFISHER</td>
			<td>61965059</td>
			<td></td>
		</tr>
		<tr>
			<td>Glutamax Supplement</td>
			<td>THERMOFISHER</td>
			<td>35050061</td>
			<td></td>
		</tr>
		<tr>
			<td>Horse Serum</td>
			<td>SIGMA</td>
			<td>16050122</td>
			<td></td>
		</tr>
		<tr>
			<td>Lens</td>
			<td>Nikon Instruments</td>
			<td>CFI Plan Fluor 100x Oil</td>
			<td></td>
		</tr>
		<tr>
			<td>Light Engines</td>
			<td>LUMENCOR</td>
			<td>SPECTRA X</td>
			<td></td>
		</tr>
		<tr>
			<td>Linear-encoded motorized platine</td>
			<td>Nikon Instruments</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope</td>
			<td>Nikon Instruments</td>
			<td>ECLIPSE Ti2-E MICROSCOPE INVERSE</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope Stage Incubator with 3-channel manual gas mixer and gas bubbler/ humidity module</td>
			<td>OKOLAB</td>
			<td>H201-NIKON-TI-S-ER</td>
			<td></td>
		</tr>
		<tr>
			<td>PBS 1x liquid</td>
			<td>THERMOFISHER</td>
			<td>20012068</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin</td>
			<td>SIGMA</td>
			<td>15140122</td>
			<td></td>
		</tr>
		<tr>
			<td>Petri dishes 100 mm</td>
			<td>SIGMA</td>
			<td>P5731</td>
			<td></td>
		</tr>
		<tr>
			<td>Petri dishes 35 mm</td>
			<td>SIGMA</td>
			<td>CLS430165</td>
			<td></td>
		</tr>
		<tr>
			<td>Pregnant Rats</td>
			<td>CHARLES RIVERS</td>
			<td>3</td>
			<td></td>
		</tr>
		<tr>
			<td>Software Nikon NIS-HC</td>
			<td>Nikon Instruments</td>
			<td>NIS-Elements HC</td>
			<td></td>
		</tr>
		<tr>
			<td>Sofware Prism</td>
			<td>GraphPad</td>
			<td>V8.02</td>
			<td></td>
		</tr>
		<tr>
			<td>Stericup 500 mL</td>
			<td>MERCK MILLIPORE</td>
			<td>10412701</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Primary culture of astrocytes infected with LV-G1-MitoTimer exhibited typical mitochondrial networks. Before treatment, astrocytes expressing LV-G1-MitoTimer showed the heterogeneous mitochondrial size and various green/red fluorescence intensities (Figure 3, Figure 4, and Video 1). The mitochondrial system of astrocyte cultures expressing LV-G1-MitoTimer was monitored before and after incubation with H202 (10 &#181;M). The different mitochondrial features described above were calculated over 12 h (every 3 h) and normalized (cell by cell) to their initial state. At the morphological level (Figure 3B), the effects of H2O2 start to be visible at about 6 h. Indeed, the mitochondria were fragmented (decrease of length, surface area, and elongation factor). This fragmentation is even more obvious 12 h after the treatment. Note that the diameters, widths, and sphericity were not reduced. Concerning redox state and turnover (Figure 3C), 3 h after H2O2 treatment, the proportion of green mitochondria increased in astrocytes (the consequence of a rapid increase in mitochondrial biogenesis). At 6 h, the green/red ratio returned to baseline levels, but the number of purely red mitochondria increased significantly from basal levels. After 12 h, the consequences of the oxidative treatment of H2O2 were visible and resulted in a substantial increase in the ratio and number of red puncta. Concerning the dynamics and mobility (Figure 3D), 3 h after the treatment, all the criteria were transiently increased. In the longer term (12 h), the mitochondria moved more slowly and over shorter distances.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62957/62957fig01.jpg" /> 
Figure 1: Astrocyte culture expressing LV-G1-MitoTimer biosensor. (A) Confocal photographs of astrocytes expressing LV-G1-MitoTimer. (B) Selection of confocal photographs of reduced (green) balanced (orange) and oxidized (red) mitochondria with different levels of fragmentation. (C) Summary diagram of the different criteria available for analysis in an astrocyte expressing LV-G1-MitoTimer. Scale bar: (A) Upper panel: 50 &#181;m, lower panel: 10 &#181;m, (B) 1 &#181;m. <a href="https://www.jove.com/files/ftp_upload/62957/62957fig01large.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/62957/62957fig02.jpg" /> 
Figure 2: Mitochondrial morphology and ratio analysis. (A) GA3 script overview for the analysis of individual mitochondrial morphology and ratio. (B) Initial photographs of an astrocyte expressing LV-G1-MitoTimer analyzed with the GA3 script. (C) Example of binary masks generated for the mitochondrial system of astrocytes. Scale bar: 10 &#181;m (B-C). <a href="https://www.jove.com/files/ftp_upload/62957/62957fig02large.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/62957/62957fig03.jpg" /> 
Figure 3: The effects of H2O2 on the mitochondrial system of astrocytes. (A) Photographs of astrocytes expressing LV-G1-MitoTimer 1 h before and 6 h, 24 h after treatment with PBS (CTRL) and 10 &#181;M of H2O2. (B) Radar charts of mitochondrial morphology, (C) redox state and turnover, and (D) mobility criteria evaluated on astrocytes during baseline and 3 h, 6 h, and 12 h after H2O2 treatment. SA: Surface area; D: Diameter; L: Length; W: Width; R: Roundness; S: Sphericity; EF: Elongation factor (=L/W); G/R: Individual red/green ratio; Gpuncta: Percentage of green puncta mitochondria; Rpuncta: Percentage of red puncta mitochondria; Dis: Displacement; Tr: Track Length; Sp and SpV: Speed and speed variance; Str: Straightness. Scale bar: 20 &#181;m (A) and 2.5 &#181;m (inset). <a href="https://www.jove.com/files/ftp_upload/62957/62957fig03large.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/62957/62957fig04.jpg" /> 
Figure 4: Photographs of astrocytes expressing LV-G1-MitoTimer and showing a homogeneous and balanced mitochondrial network during baseline. Scale bar: 20 &#181;m <a href="https://www.jove.com/files/ftp_upload/62957/62957fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Video 1: Effect of H2O2 treatment on the mitochondrial system of cultured astrocytes. Astrocytic mitochondria before H2O2 treatment (baseline), as well as 6 h and 24 h after H2O2 treatment compared to non-treated control cell. <a href="https://www.jove.com/files/ftp_upload/62957/movie.mp4" target="_blank">Please click here to download this File.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Morphology criteria</td>
			<td>Range</td>
			<td colspan="3">Remarks</td>
		</tr>
		<tr>
			<td>Surface Area (SA)</td>
			<td>0.5&#8211;4 &#181;m2</td>
			<td colspan="3" rowspan="7">These criteria inform on the fragmented-elongated features of mitochondria. They generally evolve in the same direction. Fragmented mitochondria will have decreased surface area, diameter, length, and elongation factor while roundness, sphericity, and width may be unchanged or increased.</td>
		</tr>
		<tr>
			<td>Diameter (D)</td>
			<td>0.5&#8211;1.5 &#181;m</td>
		</tr>
		<tr>
			<td>Length (L)</td>
			<td>0.5&#8211;5 &#181;m</td>
		</tr>
		<tr>
			<td>Width (W)</td>
			<td>0.5&#8211;2 &#181;m</td>
		</tr>
		<tr>
			<td>Roundness (R)</td>
			<td>0&#8211;1</td>
		</tr>
		<tr>
			<td>Sphericity (S)</td>
			<td>0&#8211;1</td>
		</tr>
		<tr>
			<td>Elongation factor (EF = L/W)</td>
			<td>1&#8211;10</td>
		</tr>
	</tbody>
</table>
Table 1: Summary of selected parameters for mitochondrial morphology.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Redox State criteria</td>
			<td>Range</td>
			<td colspan="3">Remarks</td>
		</tr>
		<tr>
			<td>Individual ratio (G/R)</td>
			<td>0&#8211;10</td>
			<td colspan="3" rowspan="3">The ratio indicates the result of the redox state. It informs about the general state and age of the mitochondria in the cell. It is essential to consider that this ratio is the balance of biogenesis and degradation of mitochondria and the fission/fusion of oxidized mitochondria with reduced mitochondria. Therefore the evaluation of the number of green and red puncta can powerfully help interpret the results.&#160; Green puncta mitochondria are determined when the intensity of green is 10 times that of red. Red puncta mitochondria are determined when the intensity of red is 10 times greater than that of green. The redox state of an astrocyte is the average of all the mitochondria ratios of that cell.</td>
		</tr>
		<tr>
			<td>Percentage of green puncta mitochondria (Gpuncta)</td>
			<td>0%&#8211;100%</td>
		</tr>
		<tr>
			<td>Percentage of red puncta mitochondria (Rpuncta)</td>
			<td>0%&#8211;100%</td>
		</tr>
	</tbody>
</table>
Table 2: Summary of selected parameters for mitochondrial redox state.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Mobility criteria</td>
			<td>Range</td>
			<td colspan="3">Remarks</td>
		</tr>
		<tr>
			<td>Displacement (Dis)</td>
			<td>0&#8211;10 &#181;m</td>
			<td colspan="3" rowspan="5">Together these features inform the general motility dynamics of the network. Stationary mitochondria display short displacement &#38; track length with a low speed. On the other hand, oscillatory particles can be differentiated with a difference between the track length and displacement (resulting in low straightness) and an increased speed compared to static.</td>
		</tr>
		<tr>
			<td>Track Length (Tr)</td>
			<td>0&#8211;10 &#181;m</td>
		</tr>
		<tr>
			<td>Speed and speed variance (Sp and SpV)</td>
			<td>0&#8211;1.5&#160; &#181;m/s &#177; 0.2 &#181;m/s</td>
		</tr>
		<tr>
			<td>Straightness&#160;</td>
			<td rowspan="2">0&#8211;1</td>
		</tr>
		<tr>
			<td>(Str = displacement/track length)</td>
		</tr>
	</tbody>
</table>
Table 3: Summary of selected parameters for mitochondrial mobility.
Supplemental Coding File 1: GA3 script file for analysis of individual mitochondrial morphology. <a href="https://www.jove.com/files/ftp_upload/62957/Mito_Morpho.ga3" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Live-imaging of Mitochondrial System in Cultured Astrocytes at JoVE.com

Live-imaging of Mitochondrial System in Cultured Astrocytes

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 ...

live-imaging-of-mitochondrial-system-in-cultured-astrocytes

Research

JoVE Journal

Neuroscience

3.9K Views.  Lausanne University Hospital (CHUV) and University of Lausanne. 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.

Video: Live-imaging of Mitochondrial System in Cultured Astrocytes

Live-imaging of PKC Translocation in Sf9 Cells and in Aplysia Sensory Neurons

Protein kinase Cs (PKCs) are serine threonine kinases that play a central role in regulating a wide variety of cellular processes such as cell growth and learning and memory. There are four known families of PKC isoforms in vertebrates: classical PKCs (&alpha;, &beta;I, &beta;II and &gamma;), novel type I PKCs (&#949; and &eta;), novel type II PKCs (&delta; and &theta;), and atypical PKCs (&zeta; and &iota;). The classical PKCs are activated by Ca2+ and diacylclycerol (DAG), while the novel PKCs are activated by DAG, but are Ca2+-independent. The atypical PKCs are activated by neither Ca2+ nor DAG. In Aplysia californica, our model system to study memory formation, there are three nervous system specific PKC isoforms one from each major class, namely the conventional PKC Apl I, the novel type I PKC Apl II and the atypical PKC Apl III. PKCs are lipid-activated kinases and thus activation of classical and novel PKCs in response to extracellular signals has been frequently correlated with PKC translocation from the cytoplasm to the plasma membrane. Therefore, visualizing PKC translocation in real time in live cells has become an invaluable tool for elucidating the signal transduction pathways that lead to PKC activation. For instance, this technique has allowed for us to establish that different isoforms of PKC translocate under different conditions to mediate distinct types of synaptic plasticity and that serotonin (5HT) activation of PKC Apl II requires production of both DAG and phosphatidic acid (PA) for translocation 1-2. Importantly, the ability to visualize the same neuron repeatedly has allowed us, for example, to measure desensitization of the PKC response in exquisite detail 3. In this video, we demonstrate each step of preparing Sf9 cell cultures, cultures of Aplysia sensory neurons have been described in another video article 4, expressing fluorescently tagged PKCs in Sf9 cells and in Aplysia sensory neurons and live-imaging of PKC translocation in response to different activators using laser-scanning microscopy.

In this video, we demonstrate visualization of PKC translocation in living cells using fluorescently tagged PKCs.

Protein kinase Cs (PKCs) are serine threonine kinases that play a central role in regulating a wide variety of cellular processes such as cell growth and ...

Time-lapse Live Imaging of Clonally Related Neural Progenitor Cells in the Developing Zebrafish Forebrain

Precise patterns of division, migration and differentiation of neural progenitor cells are crucial for proper brain development and function1,2. To understand the behavior of neural progenitor cells in the complex in vivo environment, time-lapse live imaging of neural progenitor cells in an intact brain is critically required. In this video, we exploit the unique features of zebrafish embryos to visualize the development of forebrain neural progenitor cells in vivo. We use electroporation to genetically and sparsely label individual neural progenitor cells. Briefly, DNA constructs coding for fluorescent markers were injected into the forebrain ventricle of 22 hours post fertilization (hpf) zebrafish embryos and electric pulses were delivered immediately. Six hours later, the electroporated zebrafish embryos were mounted with low melting point agarose in glass bottom culture dishes. Fluorescently labeled neural progenitor cells were then imaged for 36hours with fixed intervals under a confocal microscope using water dipping objective lens. The present method provides a way to gain insights into the in vivo development of forebrain neural progenitor cells and can be applied to other parts of the central nervous system of the zebrafish embryo.

The present video demonstrates a method which takes advantage of the combination of electroporation and confocal microscopy to perform live imaging on individual neural progenitor cells in the developing zebrafish forebrain. In vivo analysis of the development of forebrain neural progenitor cells at a clonal level can be achieved in this way.

Precise patterns of division, migration and differentiation of neural progenitor cells are crucial for proper brain development and function1,2. To ...

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 ...

Live Imaging of Dorsal Root Axons after Rhizotomy

The primary sensory axons injured by spinal root injuries fail to regenerate into the spinal cord, leading to chronic pain and permanent sensory loss. Regeneration of dorsal root (DR) axons into spinal cord is prevented at the dorsal root entry zone (DREZ), the interface between the CNS and PNS. Our understanding of the molecular and cellular events that prevent regeneration at DREZ is incomplete, in part because complex changes associated with nerve injury have been deduced from postmortem analyses. Dynamic cellular processes, such as axon regeneration, are best studied with techniques that capture real-time events with multiple observations of each living animal. Our ability to monitor neurons serially in vivo has increased dramatically owing to revolutionary innovations in optics and mouse transgenics. Several lines of thy1-GFP transgenic mice, in which subsets of neurons are genetically labeled in distinct fluorescent colors, permit individual neurons to be imaged in vivo1. These mice have been used extensively for in vivo imaging of muscle2-4 and brain5-7, and have provided novel insights into physiological mechanisms that static analyses could not have resolved. Imaging studies of neurons in living spinal cord have only recently begun. Lichtman and his colleagues first demonstrated their feasibility by tracking injured dorsal column (DC) axons with wide-field microscopy8,9. Multi-photon in vivo imaging of deeply positioned DC axons, microglia and blood vessels has also been accomplished10. Over the last few years, we have pioneered in applying in vivo imaging to monitor regeneration of DR axons using wide-field microscopy and H line of thy1-YFP mice. These studies have led us to a novel hypothesis about why DR axons are prevented from regenerating within the spinal cord11.
In H line of thy1-YFP mice, distinct YFP+ axons are superficially positioned, which allows several axons to be monitored simultaneously. We have learned that DR axons arriving at DREZ are better imaged in lumbar than in cervical spinal cord. In the present report we describe several strategies that we have found useful to assure successful long-term and repeated imaging of regenerating DR axons. These include methods that eliminate repeated intubation and respiratory interruption, minimize surgery-associated stress and scar formation, and acquire stable images at high resolution without phototoxicity.

An in vivo imaging protocol to monitor primary sensory axons following dorsal root crush is described. The procedures utilize wide-field fluorescence microscopy and thy1-YFP transgenic mice, and permit repeated imaging of axon regeneration over 4 cm in the PNS and axon interactions with the interface of the CNS.

The primary sensory axons injured by spinal root injuries fail to regenerate into the spinal cord, leading to chronic pain and permanent sensory loss. ...

High-resolution Live Imaging of Cell Behavior in the Developing Neuroepithelium

The embryonic spinal cord consists of cycling neural progenitor cells that give rise to a large percentage of the neuronal and glial cells of the central nervous system (CNS). Although much is known about the molecular mechanisms that pattern the spinal cord and elicit neuronal differentiation1, 2, we lack a deep understanding of these early events at the level of cell behavior. It is thus critical to study the behavior of neural progenitors in real time as they undergo neurogenesis.
In the past, real-time imaging of early embryonic tissue has been limited by cell/tissue viability in culture as well as the phototoxic effects of fluorescent imaging. Here we present a novel assay for imaging such tissue for long periods of time, utilizing a novel ex vivo slice culture protocol and wide-field fluorescence microscopy (Fig. 1). This approach achieves long-term time-lapse monitoring of chick embryonic spinal cord progenitor cells with high spatial and temporal resolution.
This assay may be modified to image a range of embryonic tissues3, 4 In addition to the observation of cellular and sub-cellular behaviors, the development of novel and highly sensitive reporters for gene activity (for example, Notch signaling5) makes this assay a powerful tool with which to understand how signaling regulates cell behavior during embryonic development.

Imaging embryonic tissue in real-time is challenging over long periods of time. Here we present an assay for monitoring cellular and sub-cellular changes in chick spinal cord for long periods with high spatial and temporal resolution. This technique can be adapted for other regions of the nervous system and developing embryo.

The embryonic spinal cord consists of cycling neural progenitor cells that give rise to a large percentage of the neuronal and glial cells of the central ...

Ex vivo Live Imaging of Single Cell Divisions in Mouse Neuroepithelium

We developed a system that integrates live imaging of fluorescent markers and culturing slices of embryonic mouse neuroepithelium. We took advantage of existing mouse lines for genetic cell lineage tracing: a tamoxifen-inducible Cre line and a Cre reporter line expressing dsRed upon Cre-mediated recombination. By using a relatively low level of tamoxifen, we were able to induce recombination in a small number of cells, permitting us to follow individual cell divisions. Additionally, we observed the transcriptional response to Sonic Hedgehog (Shh) signaling using an Olig2-eGFP transgenic line 1-3 and we monitored formation of cilia by infecting the cultured slice with virus expressing the cilia marker, Sstr3-GFP 4. In order to image the neuroepithelium, we harvested embryos at E8.5, isolated the neural tube, mounted the neural slice in proper culturing conditions into the imaging chamber and performed time-lapse confocal imaging. Our ex vivo live imaging method enables us to trace single cell divisions to assess the relative timing of primary cilia formation and Shh response in a physiologically relevant manner. This method can be easily adapted using distinct fluorescent markers and provides the field the tools with which to monitor cell behavior in situ and in real time.

Ex vivo Live Imaging of Single Cell Divisions in Mouse Neuroepithelium

Here we develop the tools necessary for ex vivo live imaging to trace single cell divisions in the mouse E8.5 neuroepithelium

We developed a system that integrates live imaging of fluorescent markers and culturing slices of embryonic mouse neuroepithelium. We took advantage of ...

Live Imaging of Mitosis in the Developing Mouse Embryonic Cortex

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the developing cerebral cortex, abnormal mitosis of neural progenitors can cause defects in brain size and function. Hence, there is a critical need for tools to understand the mechanisms of neural progenitor mitosis. Cortical development in rodents is an outstanding model for studying this process. Neural progenitor mitosis is commonly examined in fixed brain sections. This protocol will describe in detail an approach for live imaging of mitosis in ex vivo embryonic brain slices. We will describe the critical steps for this procedure, which include: brain extraction, brain embedding, vibratome sectioning of brain slices, staining and culturing of slices, and time-lapse imaging. We will then demonstrate and describe in detail how to perform post-acquisition analysis of mitosis. We include representative results from this assay using the vital dye Syto11, transgenic mice (histone H2B-EGFP and centrin-EGFP), and in utero electroporation (mCherry-&#945;-tubulin). We will discuss how this procedure can be best optimized and how it can be modified for study of genetic regulation of mitosis. Live imaging of mitosis in brain slices is a flexible approach to assess the impact of age, anatomy, and genetic perturbation in a controlled environment, and to generate a large amount of data with high temporal and spatial resolution. Hence this protocol will complement existing tools for analysis of neural progenitor mitosis.

Neural progenitor mitosis is a critical parameter of neurogenesis. Much of our understanding of neural progenitor mitosis is based on analysis of fixed tissue. Live imaging in embryonic brain slices is a versatile technique to assess mitosis with high temporal and spatial resolution in a controlled environment.

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the ...

Live Imaging of Drosophila Larval Neuroblasts

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given their relevance to development and disease, understanding the mechanisms that govern asymmetric stem cell division has been a robust area of study. Because they are genetically tractable and undergo successive rounds of cell division about once every hour, the stem cells of the Drosophila central nervous system, or neuroblasts, are indispensable models for the study of stem cell division. About 100 neural stem cells are located near the surface of each of the two larval brain lobes, making this model system particularly useful for live imaging microscopy studies. In this work, we review several approaches widely used to visualize stem cell divisions, and we address the relative advantages and disadvantages of those techniques that employ dissociated versus intact brain tissues. We also detail our simplified protocol used to explant whole brains from third instar larvae for live cell imaging and fixed analysis applications.

Live Imaging of Drosophila Larval Neuroblasts

This protocol details a streamlined method used to conduct live cell imaging in the context of an intact larval brain. Live cell imaging approaches are invaluable for the study of asymmetric neural stem cell divisions as well as other neurogenic and developmental processes, consistently uncovering mechanisms that were previously overlooked.

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given ...

Live Imaging of the Ependymal Cilia in the Lateral Ventricles of the Mouse Brain

Multiciliated ependymal cells line the ventricles in the adult brain. Abnormal function or structure of ependymal cilia is associated with various neurological deficits. The current ex vivo live imaging of motile ependymal cilia technique allows for a detailed study of ciliary dynamics following several steps. These steps include: mice euthanasia with carbon dioxide according to protocols of The University of Toledo&#8217;s Institutional Animal Care and Use Committee (IACUC); craniectomy followed by brain removal and sagittal brain dissection with a vibratome or sharp blade to obtain very thin sections through the brain lateral ventricles, where the ependymal cilia can be visualized. Incubation of the brain&#8217;s slices in a customized glass-bottom plate containing Dulbecco&#8217;s Modified Eagle&#8217;s Medium (DMEM)/High-Glucose at 37 &#176;C in the presence of 95%/5% O2/CO2 mixture is essential to keep the tissue alive during the experiment. A video of the cilia beating is then recorded using a high-resolution differential interference contrast microscope. The video is then analyzed frame by frame to calculate the ciliary beating frequency. This allows distinct classification of the ependymal cells into three categories or types based on their ciliary beating frequency and angle. Furthermore, this technique allows the use of high-speed fluorescence imaging analysis to characterize the unique intracellular calcium oscillation properties of ependymal cells as well as the effect of pharmacological agents on the calcium oscillations and the ciliary beating frequency. In addition, this technique is suitable for immunofluorescence imaging for ciliary structure and ciliary protein localization studies. This is particularly important in disease diagnosis and phenotype studies. The main limitation of the technique is attributed to the decrease in live motile cilia movement as the brain tissue starts to die.

Using high-resolution differential interference contrast (DIC) microscopy, an ex vivo observation of the beating of motile ependymal cilia located within the mouse brain ventricles is demonstrated by live-imaging. The technique allows a recording of the unique ciliary beating frequency and beating angle as well as their intracellular calcium oscillation pacing properties.

Multiciliated ependymal cells line the ventricles in the adult brain. Abnormal function or structure of ependymal cilia is associated with various ...

Protocol for Three-dimensional Confocal Morphometric Analysis of Astrocytes

As glial cells in the brain, astrocytes have diverse functional roles in the central nervous system. In the presence of harmful stimuli, astrocytes modify their functional and structural properties, a condition called reactive astrogliosis. Here, a protocol for assessment of the morphological properties of astrocytes is presented. This protocol includes quantification of 12 different parameters i.e. the surface area and volume of the tissue covered by an astrocyte (astrocyte territory), the entire astrocyte including branches, cell body, and nucleus, as well as total length and number of branches, the intensity of fluorescence immunoreactivity of antibodies used for astrocyte detection, and astrocyte density (number/1,000 &#181;m2). For this purpose three-dimensional (3D) confocal microscopic images were created, and 3D image analysis software such as Volocity 6.3 was used for measurements. Rat brain tissue exposed to amyloid beta1-40 (A&#946;1-40) with or without a therapeutic&#160;intervention was used to present the method. This protocol can also be used for 3D morphometric analysis of other cells from either in vivo or in vitro conditions.

Astrocytes in the CNS change their functional and structural properties in response to harmful stimuli. This report presents a protocol for assessment of three-dimensional astrocyte morphology in diseased conditions or after therapeutic interventions.

As glial cells in the brain, astrocytes have diverse functional roles in the central nervous system. In the presence of harmful stimuli, astrocytes modify ...