DNL was supported by the University of North Carolina at Chapel Hill (UNC) School of Medicine as a Simmons Scholar. TWR was supported by UNC PREP Grant R25 GM089569. Work using the UNC Neuroscience Center Microscopy Core Facility was supported, in part, by funding from the NIH-NINDS Neuroscience Center Support Grant P30 NS045892 and the NIH-NICHD Intellectual and Developmental Disabilities Research Center Support Grant U54 HD079124.

All animal procedures were performed with the approval of the University of North Carolina at Chapel Hill Animal Care and Use Committee (IACUC).
1. Brain dissection and culture of primary mouse astrocytes
NOTE: The following protocol was adapted from published methods, which follows the original procedure developed by McCarthy and deVellis13,14,15. Mixed cell cultures of McCarthy/deVellis (MD) astrocytes are prepared from postnatal day 2&#8722;4 (P2&#8722;P4) mouse cortices, treated with an anti-mitotic factor, and purified to yield the final enriched astrocyte culture. Four cortices from mouse pups of either sex are sufficient to generate one T75 tissue culture flask culture.
<ol>
	<li>Coat the bottom of T75 flasks (100% angled neck access, 0.22 &#181;m hydrophobic vented filter cap) with poly-D-lysine (1 mg/mL in 0.1 M Tris buffer, pH 8.5) and incubate at 37 &#176;C for 24 h prior to start of dissection.</li>
	<li>Before beginning the dissection procedure, warm 30 mL of astrocyte culture media (Dulbecco&#8217;s modified Eagle&#8217;s medium [DMEM] high glucose, 10% heat-inactivated fetal bovine serum, 1% penicillin/streptomycin) at 37 &#176;C and thaw a 5 mL aliquot of 2.5% trypsin on ice. Prepare two dissecting dishes (60 mm diameter) on ice filled with 6 mL of Hank&#8217;s balanced salt solution (HBSS) without Ca2+ or Mg2+.</li>
	<li>Disinfect all surgical tools (fine tip tweezers, dissecting scissors, Vannas scissors straight, Graefe forceps with curved tips) in 70% ethanol prior to and in between each dissection.</li>
	<li>Spray the head and neck of the P2&#8722;P4 mouse pups with 70% ethanol before quickly euthanizing them by decapitation with surgical scissors. Perform a posterior to anterior midline incision along the scalp to expose the skull. Carefully cut the skull from the neck to the nose.</li>
	<li>Perform two additional lateral incisions superior to the eyes. Using fine tip tweezers gently flip the skull flaps to the side. Remove the brain and place it in the first dissecting dish filled with ice-cold HBSS without Ca2+ or Mg2+. Keep the dish on ice and continue harvesting the rest of the brains. 
	NOTE: Perform the remainder of the dissection procedure under a dissecting scope.</li>
	<li>Remove the olfactory bulbs using Vannas scissors or fine tweezers (Figure 1Ai). With the curved tip forceps at the intersection of the transverse and interhemispheric fissures, gently insert a second pair of curved tip forceps between the cortex and diencephalon, slowly opening the forceps to separate the cerebral hemispheres from the rest of the brain (Figure 1Aii,Aiii). Remove the unwanted tissue (Figure 1Aiv).</li>
	<li>For each cortical hemisphere, carefully remove the meninges with fine tip tweezers. Optionally, remove the hippocampus from the cortices. Transfer the dissected mouse cortices into a second dish filled with ice-cold HBSS without Ca2+ or Mg2+ and keep it on ice while dissecting the remaining brains. 
	NOTE: Conduct the next steps under aseptic conditions in a laminar flow hood.</li>
	<li>Cut the dissected cortices into small pieces (approximately 2&#8722;4 mm) using sterile sharp surgical scissors or a scalpel. Transfer the dissected tissue to a 50 mL conical tube containing enzyme solution (22.5 mL HBSS without Ca2+ or Mg2+, 2.5 mL 2.5% trypsin). Incubate at 37 &#176;C for 30 min with gentle inversion every 10 min.</li>
	<li>Collect the digested tissue by centrifugation at 300 x g for 5 min and remove the enzyme solution by decantation (do not use a vacuum aspirator). Add 10 mL of astrocyte culture media and dissociate the tissue into single cells by pipetting the solution 20&#8722;30x using a 10 mL serological pipette.</li>
	<li>Filter the cell suspension through a 70 &#181;m cell strainer to minimize cell clumps and undigested tissue. Collect the filtrate in a clean 50 mL conical tube and adjust the total volume to 20 mL with astrocyte culture media. At this point, evaluate the efficiency of the cortical dissociation by mixing 10 &#181;L of the cell suspension with 10 &#181;L of trypan blue and counting cells with a hemocytometer. 
	NOTE: One preparation from four P2&#8722;P4 mouse cortices yields 10&#8722;15 x 106 single cells.</li>
	<li>Aspirate the poly-D-lysine coating solution from the T75 culture flasks and wash twice with sterile water. Plate the 20 mL cell suspension and incubate at 37 &#176;C and 5% CO2.</li>
</ol>
2. Purification and maintenance of astrocyte cultures
NOTE: Perform all media changes under aseptic conditions in a laminar flow hood using sterile filtered (0.22 &#181;m filter membrane) astrocyte media warmed to 37 &#176;C.
<ol>
	<li>Replace the cell media 48 h after plating (day in vitro 2, DIV2).</li>
	<li>At DIV5, replace the media with fresh astrocyte culture media supplemented with 10 &#181;M cytosine arabinoside (AraC). 
	NOTE: This step facilitates the removal of contaminant cells, including fibroblasts and other glial cells. It is important to treat the cultures with AraC no longer than 48 h as longer incubation times will reduce astrocyte viability.</li>
	<li>At DIV7, change media to astrocyte culture media without AraC and start to check the culture confluency daily. Once cells have reached 80% confluence, coat two-T75 flasks for each unpurified astrocyte culture flask with poly-D-lysine and incubate at 37 &#176;C for at least 8 h to overnight.</li>
	<li>The following day, begin the astrocyte purification by shaking the culture flasks at 180 rpm for 30 min in a 37 &#176;C orbital shaking incubator. Remove the media and replace it with fresh astrocyte culture media. Shake cells at 240 rpm for 6 h in a 37 &#176;C orbital shaking incubator. Prewarm 1x phosphate-buffered saline (PBS), astrocyte culture media, and 0.25% trypsin-EDTA to 37 &#176;C 20&#8722;30 min before the end of the shaking period. 
	NOTE: CO2 incubation is not required during the shaking steps. If desired, the media collected after the 30-min and 6-h shaking steps may be used to culture microglia and oligodendrocytes, respectively. AraC treatment of the mixed culture will, however, reduce the abundance of the other glial cells.</li>
	<li>Remove the culture flasks from the shaker and immediately replace the media with 10 mL of warm 1x PBS per flask. Remove PBS and add 3 mL of 0.25% trypsin-EDTA per flask. Incubate at 37 &#176;C and 5% CO2, checking every 5 min for detachment.</li>
	<li>Once cells have detached from the flask, stop trypsinization by adding 10 mL of astrocyte culture media. Transfer detached astrocytes to a 50 mL conical tube and pellet cells by centrifugation at 300 x g for 5 min. Aspirate supernatant and re-suspend the cells in 40 mL of astrocyte culture media.</li>
	<li>Wash the poly-D-lysine-coated T75 flasks twice with sterile water and plate 20 mL of the purified astrocyte suspension. Return to the 37 &#176;C and 5% CO2 incubator. Change the astrocyte culture media every two days for an additional 10 days until astrocytes mature. 
	NOTE: Each T75 flask of purified astrocytes should be sufficient to produce 2 new T75 flasks with 3 x 105 cells each at plating.</li>
	<li>Repeat steps 2.5&#8722;2.7 for subsequent passes or to plate astrocytes for microscopy. 
	NOTE: The purity of the astrocyte cultures after the addition of AraC and following purification can be assessed qualitatively through brightfield microscopy, or more precisely by quantifying the percentage of glial fibrillary acid protein (GFAP)-positive cells per field of view in fluorescent images (Figure 1B,C).</li>
</ol>
3. Transfection of fluorescently tagged plasmids
<ol>
	<li>The day prior to transfection or labeling, seed astrocytes at desired density for imaging 24&#8722;48 h post-transfection. A recommended density for 24-well plates or 14 mm glass bottom wells is 2 x 104 cells/well. 
	NOTE: This protocol is optimized for transfection of astrocytes plated on 12 mm glass coverslips on 24-well plates or 14 mm glass bottom wells using a lipofection-based method. Reagents should be scaled depending on the size of the culture dish in which the astrocytes are growing.</li>
	<li>Dilute the lipofection reagent (Table of Materials) in reduced-serum media (Table of Materials). To optimize the ratio of lipofection reagent to DNA, test a range of adequate dilutions. For example, if using the reagent employed in this protocol (Table of Materials), dilute 2, 3, 4, and 5 &#956;L of the lipofection reagent in 50 &#956;L of reduced-serum media.</li>
	<li>Dilute 5 &#956;g of high purity DNA in 250 &#956;L of reduced-serum media. Add 5 &#956;L of lipofection enhancer reagent (Table of Materials). Combine the tube containing the lipofection reagent with an equal volume of the DNA-lipofection enhancer mix. Mix by pipetting and incubate at room temperature (RT) for 15 min.</li>
	<li>Remove astrocyte culture media and add transfection mix (step 3.3) to cells dropwise. After a 6-h incubation at 37 &#176;C and 5% CO2, replace the transfection complex with an appropriate volume of astrocyte culture media (2 mL for 14 mm glass bottom dishes). Incubate for an additional 24&#8722;72 h before proceeding to the image acquisition. Closely monitor the duration of the incubation step to achieve the best balance between protein expression and cell viability.</li>
</ol>
4. Labeling of late endosomes/lysosomes using fluorescent probes
NOTE: Some cargos can be labeled using fluorescent dyes with high affinity for cargo-specific proteins. The following example permits the labelling of late endosomes/lysosomes with a fluorescent acidotropic probe.
<ol>
	<li>Dilute the lysosomal-labeling probe (Table of Materials) in 200 &#181;L of astrocyte culture media to a working concentration of 1 &#956;M and apply to astrocytes from step 3.1 at a density of 2 x 104 cells/well. Incubate for 30 min at 37 &#176;C.</li>
	<li>Wash the cells once with warm astrocyte culture media and replace with imaging media (Table of Materials). Proceed to live imaging immediately.</li>
</ol>
5. Image acquisition using a time-lapse imaging system
NOTE: Time-lapse live imaging should be done using a fluorescence microscope equipped with a high-speed camera, definite focus, incubation chamber, and a 40x oil objective with a high numeric aperture (e.g., Plan-Apochromat 1.4NA). A variety of acquisition software is available for time-lapse imaging. Selection of the microscopy system and acquisition software should be based on their availability and suitability for the goals of the particular study. Some general guidelines are provided below.
<ol>
	<li>Place the culture chamber or dish in the proper adaptor on the microscope stage. Using epifluorescence light, select the cell(s) that express fluorescent proteins or probe to record. Adjust the fluorescent sample illumination to visualize the selected cell(s) using the digital camera. Avoid using high illumination that might cause photobleaching and phototoxicity. Adjust focus and zoom.</li>
	<li>Acquire single Z-stack time-lapse series at a frequency of 1 frame every 2 s for time intervals ranging between 300 s and 500 s using the zoom and definite focus functions. 
	NOTE: Trafficking dynamics (velocity, frequency of motion, etc.) will vary depending on the protein of interest, thus, the time course of acquisition may require adjustment. For instance, mitochondria are less motile than lysosomes and exhibit frequent pauses. In this instance, it is more appropriate to adjust the acquisition parameters to 1 frame every 5 s for 800 s.</li>
	<li>Save time-lapse images and export them as AVI or TIFF stack files.</li>
</ol>
6. Image analysis
NOTE: Image analysis was performed using the KymoToolBox plugin for ImageJ/Fiji16. Detailed step-by-step written instructions are available online17. The following abbreviated steps should be sufficient to create the kymographs and extract the particle transport parameters.
<ol>
	<li>Open time-lapse image sequence in ImageJ/Fiji software. Import images from the File menu as AVI or TIFF stack files. Convert the images to 8-bit or 16-bit by selecting one of these image types using the Type tool from the Image menu. If working with RGB files, split channels using the Split Channel tool available under the Color feature in the Image menu.</li>
	<li>To generate a kymograph (representation in position and time of particle trajectories), draw each track in which particles are moving by tracing a line along the trajectories of the particles using the segmented line tool. To correctly assign directionality of the movement, draw the polyline using the same desired directional convention for all movies so that the polarity is consistent within and across all cells being studied. For instance, draw the polyline from the center of the cell towards the periphery or vice versa. Adjust the line width to match the thickness of the track by double clicking the Line tool.</li>
	<li>Run the Draw Kymo macro of the KymoToolBox plugin using a line width of 10. A prompt asking to calibrate the image in time (number of frames, frame rate) and space (x,y resolution) will appear. Once calibrated, the kymographs are generated and can be saved as TIFF files for data presentation or further particle analysis.</li>
	<li>Assign particle trajectories by manually tracking each particle in the kymograph using the segmented line tool for the full duration of the acquisition. Record each particle trajectory as a region of interest (ROI) using the ROI manager tool found in the Analyze/Tools menu. Save all ROIs per each time-lapse video for further analysis. 
	NOTE: It is critical to assign the correct trajectory for individual particles on the kymograph to achieve the most accurate calculation of transport parameters.
	<ol>
		<li>Run the Analyze Kymo macro of the KymoToolBox plugin. A window will open asking to define the &#34;outward&#34;&#160;direction of particle motion (from left to right or vice versa) from the drop-down menu. This selection depends on the position of the fluorescent cargos imaged relative to the nucleus or the cell periphery, as established in step 6.2. For instance, if the nucleus is positioned to the left of the cargos, and the polyline in step 6.2 was drawn away from the nucleus, define the outward particle motion as &#34;from left to right&#34;.</li>
		<li>Define the limit speed (minimum speed at which a cargo is considered motile) in the Analyze Kymo window. For cargos that move at fast intracellular transport speeds, 0.1 &#181;m/second is an appropriate choice. Modify this value to adjust the software&#39;s sensitivity to each cargo of interest. Adjust the line width to match the thickness of each particle trajectory.</li>
		<li>Select the Log all data and Log extrapolated coordinates options in the Analyze kymo window. This will calculate various cargo transport parameters, including mean speed, inward speed, outward speed, cumulative distance traveled, distance traveled in either the inward or outward direction, number of switches for each particle, percentage of time moving in each direction or paused, etc. Data calculated for individual tracks are then pooled per kymograph and saved in text files specific to each image. 
		NOTE: The Show Colored Kymo option of Analyze Kymo macro generates a color-coded overlay of the path over the original kymograph. Particles traveling outward are colored in green, inward in red, and stationary particles in blue. The Report Coordinates on Original Stack option of the Analyze Kymo macro generates a color-coded overlay of the extrapolated object&#8217;s position on the original time-lapse video and kymograph.</li>
	</ol>
	</li>
</ol>

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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Glia+in+mammalian+development+and+disease.">Glia in mammalian development and disease.</a> Development. 142 (22), 3805-3809 (2015).</li><li>Allaman, I., B&#233;langer, M., Magistretti, P. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Astrocyte-neuron+metabolic+relationships:+for+better+and+for+worse.">Astrocyte-neuron metabolic relationships: for better and for worse.</a> Trends in Neurosciences. 34 (2), 76-87 (2011).</li><li>Chung, W. S., Barres, B. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+role+of+glial+cells+in+synapse+elimination.">The role of glial cells in synapse elimination.</a> Current Opinion in Neurobiology. 22 (3), 438-445 (2012).</li><li>Chung, W. S., Allen, N. J., Eroglu, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Astrocytes+Control+Synapse+Formation,+Function,+and+Elimination.">Astrocytes Control Synapse Formation, Function, and Elimination.</a> Cold Spring Harbor Perspectives in Biology. 7 (9), 020370 (2015).</li><li>Shin, J. W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Distribution+of+glutamate+transporter+GLAST+in+membranes+of+cultured+astrocytes+in+the+presence+of+glutamate+transport+substrates+and+ATP.">Distribution of glutamate transporter GLAST in membranes of cultured astrocytes in the presence of glutamate transport substrates and ATP.</a> Neurochemistry Research. 34 (10), 1758-1766 (2009).</li><li>Potokar, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cytoskeleton+and+vesicle+mobility+in+astrocytes.">Cytoskeleton and vesicle mobility in astrocytes.</a> Traffic. 8 (1), 12-20 (2007).</li><li>Potokar, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Regulation+of+AQP4+surface+expression+via+vesicle+mobility+in+astrocytes.">Regulation of AQP4 surface expression via vesicle mobility in astrocytes.</a> Glia. 61 (6), 917-928 (2013).</li><li>Potokar, M., Lacovich, V., Chowdhury, H. 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The authors have nothing to disclose.

Here, we describe an experimental approach to express, visualize, and track fluorescently tagged organelles and membrane proteins of interest using time-lapse video microscopy in high-purity primary mouse cortical MD astrocytes. We also outline a methodology for measuring particle dynamics. Direct visualization of protein and organelle dynamics in primary astrocytes provides a powerful tool to study the regulation of intracellular transport in these cells in vitro.
The methodology for establis...

Astrocytes are the most abundant cells in the adult central nervous system, where they perform unique developmental and homeostatic functions1. Astrocytes modulate synaptic development through direct contact with pre- and postsynaptic terminals as part of the tri-partite synapse, which contains neurotransmitter receptors, transporters, and cell adhesion molecules that facilitate synapse formation and neuron-astrocyte communication2. In addition, astrocytes actively control synaptic transmission and prevent neuronal excitotoxicity by rapidly removing excitatory neurotransmitters from the synaptic cleft, recycling neurotransmitters, and participating in synaptic pruning3,4,5,6. To enable these highly interactive functions, astrocytes communicate among themselves, with other glial cells, and with neurons through specialized membrane proteins, including cell adhesion molecules, aquaporins, ion channels, neurotransmitter transporters, and gap junction molecules. Astrocytes actively change the surface levels of these proteins in response to fluctuations in their intra- and extracellular environment7. Furthermore, changes in the levels and distribution of mitochondria, lipid droplets, and degradative and recycling organelles modulate energy supply, metabolite availability, and cellular clearing processes that are essential for astrocyte function and survival.
The dynamic changes in membrane protein and organelle trafficking and positioning in astrocytes are facilitated by the concerted function of motor proteins and adaptors that promote cargo motility8,9. Similarly, surface levels of membrane proteins are modulated through internalization and recycling events10. These cargos are transported via an intricate network of actin, microtubules, and possibly intermediate filaments tracks8. Studies based on immunofluorescence staining of end-binding protein 1 (EB1), which accumulates at the growing microtubule plus ends, suggest that in astrocytes bundles of microtubules radiate out from the perinucleus and extend their plus end towards the periphery11. However, a comprehensive examination of the organization and polarity of microtubules and other cytoskeletal elements using live-cell imaging is still lacking. While many of the mechanisms underlying the dynamics of organelles and membrane proteins have been extensively studied in neurons and other cell types, cargo motility in astrocytes is less well understood. Most of our current knowledge about changes in protein and organelle distribution in astrocytes is based on traditional antibody-based labeling of fixed preparation, which precludes precise spatial and temporal examination of cargo dynamics7,12.
Here, we describe a method to label membrane proteins and organelles for live imaging in high purity primary mouse astrocyte cultures. Using this protocol, we provide examples in which we track the dynamic localization of green fluorescent protein (GFP)-tagged membrane proteins in transfected astrocytes, including the gap junction protein connexin 43 (Cx43-GFP) and the excitatory amino acid transporter 1 (EAAT1-GFP). We also describe the use of a fluorescent acidotropic probe to visualize acidic organelles and follow their trafficking dynamics in live astrocytes. Finally, we demonstrate how to analyze the time-lapse data to extract and evaluate transport parameters for individual cargos.

<table>
	<tbody>
		<tr>
			<td>2.5% Trypsin (10x)</td>
			<td>Gibco</td>
			<td>15090-046</td>
			<td></td>
		</tr>
		<tr>
			<td>Benchtop Centrifuge</td>
			<td>Thermo Scientific</td>
			<td>75-203-637</td>
			<td>Sorvall ST8 Centrifuge</td>
		</tr>
		<tr>
			<td>Cell Culture Grade Water</td>
			<td>Gen Clone</td>
			<td>25-511</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell Culture Microscope</td>
			<td>Zeiss</td>
			<td>WSN-AXIOVERT A1</td>
			<td>Vert.A1 inverted scope, Inverted tissue culture microscope with fluorescent capabilities</td>
		</tr>
		<tr>
			<td>Cytosine Arabinoside</td>
			<td>Sigma</td>
			<td>C1768-100MG</td>
			<td>(AraC) Dissolve lyophilized powder in sterile cell culture grade water to make a 10mM stock, aliquot and freeze for long term storage</td>
		</tr>
		<tr>
			<td>DAPI</td>
			<td>Sigma</td>
			<td>D9542-5MG</td>
			<td>Dissolve lyophilized powder in deionized water to a maximum concentration of 20 mg/ml, heat or sonicate as necessary. Use at 300 nM for counterstaining.</td>
		</tr>
		<tr>
			<td>Dissecting Microscope</td>
			<td>Zeiss</td>
			<td></td>
			<td>Stemi 305</td>
		</tr>
		<tr>
			<td>Dissecting Scissors</td>
			<td>F.S.T</td>
			<td>14558-09</td>
			<td></td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s Modified Eagle Medium</td>
			<td>Gen Clone</td>
			<td>25-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum</td>
			<td>Gemini</td>
			<td>100-106</td>
			<td>Heat-Inactivated</td>
		</tr>
		<tr>
			<td>FIJI (Fiji is Just Image J)</td>
			<td>NIH</td>
			<td>Version 1.52i</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine Tip Tweezers</td>
			<td>F.S.T</td>
			<td>11254-20</td>
			<td>Style #5</td>
		</tr>
		<tr>
			<td>Fluorescence light source</td>
			<td>Excelitas</td>
			<td>012-63000</td>
			<td>X-Cite 120Q</td>
		</tr>
		<tr>
			<td>GFAP antibody</td>
			<td>Cell Signaling</td>
			<td>3670S</td>
			<td>GFAP (GA5) mouse monoclonal antibody</td>
		</tr>
		<tr>
			<td>Glass Bottom Dishes</td>
			<td>Mattek corporation</td>
			<td>P35G-1.5-14-C</td>
			<td>35 mm Dish | No. 1.5 Coverslip | 14 mm Glass Diameter | Uncoated</td>
		</tr>
		<tr>
			<td>Graefe Forceps</td>
			<td>F.S.T</td>
			<td>11054-10</td>
			<td>Graefe Iris Forceps with curved tips</td>
		</tr>
		<tr>
			<td>Green fluorescent dye that stains acidic compartments (late endosomes and lysosomes)</td>
			<td>Life Technologies</td>
			<td>L7526</td>
			<td>LysoTracker Green DND 26. Pre-dissolved in DMSOS to a 1mM stock solution. Dilute to the final working concentration in the growth medium or buffer of choice.</td>
		</tr>
		<tr>
			<td>Hank&#39;s Balanced Salt Solution (10x)</td>
			<td>Gibco</td>
			<td>14065-056</td>
			<td>Magnesium and calcium free</td>
		</tr>
		<tr>
			<td>Imaging Media</td>
			<td>Life Technologies</td>
			<td>A14291DJ</td>
			<td>Live Cell Imaging Solution</td>
		</tr>
		<tr>
			<td>Inverted Confocal Microscope</td>
			<td>Zeiss</td>
			<td></td>
			<td>LSM 780</td>
		</tr>
		<tr>
			<td>KymoToolBox</td>
			<td></td>
			<td></td>
			<td>https://github.com/fabricecordelieres/IJ_KymoToolBox</td>
		</tr>
		<tr>
			<td>Lipofection Enhancer Reagent</td>
			<td>Life Technologies</td>
			<td>11514015</td>
			<td>Plus Reagent</td>
		</tr>
		<tr>
			<td>Lipofection Reagent</td>
			<td>Life Technologies</td>
			<td>15338100</td>
			<td>Lipofectamine LTX reagent</td>
		</tr>
		<tr>
			<td>Orbital shaking incubator</td>
			<td>New Brunswick Scientific</td>
			<td>8261-30-1008</td>
			<td>Innova 4230 , orbital shaking incubator with temperature and speed control</td>
		</tr>
		<tr>
			<td>Penicillin/Strepomycin solution (100x)</td>
			<td>Gen Clone</td>
			<td>25-512</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate Buffered Saline (10x)</td>
			<td>Gen Clone</td>
			<td>25-507x</td>
			<td></td>
		</tr>
		<tr>
			<td>Poly-D-Lysine Hydrobromide</td>
			<td>Sigma</td>
			<td>P7405</td>
			<td>Dissolve in Tris buffer, pH 8.5, at 1mg/mL. Freeze for long term storage. Avoid cycles of freezing and thawing</td>
		</tr>
		<tr>
			<td>Reduced serum medium</td>
			<td>Gibco</td>
			<td>31985-062</td>
			<td>OPTI-MEM</td>
		</tr>
		<tr>
			<td>Tissue Culture Flasks</td>
			<td>Olympus Plastics</td>
			<td>25-209</td>
			<td>75 cm^2. 100% angled neck access, 0.22um hydrophobic vented filter cap</td>
		</tr>
		<tr>
			<td>Tissue culture incubator</td>
			<td>Thermo Scientific</td>
			<td>51030285</td>
			<td>HERAcell VIOS 160i, tissue culture incubator with temperature, humidity, and CO2 control</td>
		</tr>
		<tr>
			<td>Tris-Base</td>
			<td>Sigma</td>
			<td>T1503</td>
			<td>8.402 g dissolved to one liter in water with 4.829 g Tris HCl to make 0.1 M Tris buffer, pH 8.5</td>
		</tr>
		<tr>
			<td>Tris-HCl</td>
			<td>Sigma</td>
			<td>T3253</td>
			<td>4.829 g dissolved to one liter in water with 8.402 g Tris Base to make 0.1 M Tris buffer, pH 8.5</td>
		</tr>
		<tr>
			<td>Trypsin-EDTA (0.25%), phenol red</td>
			<td>Gibco</td>
			<td>25200072</td>
			<td></td>
		</tr>
		<tr>
			<td>Vacuum-Driven Filter Systems</td>
			<td>Olympus Plastics</td>
			<td>25-227</td>
			<td>500 ml, PES membrane, 0.22 &#181;m</td>
		</tr>
		<tr>
			<td>Vannas scissors straight</td>
			<td>Roboz</td>
			<td>RS-5620</td>
			<td></td>
		</tr>
	</tbody>
</table>

visualizing and analyzing intracellular transport of organelles and other cargos in astrocytes

Astrocytes are among the most abundant cell types in the adult brain, where they play key roles in a multiplicity of functions. As a central player in brain homeostasis, astrocytes supply neurons with vital metabolites and buffer extracellular water, ions, and glutamate. An integral component of the &#8220;tri-partite&#8221; synapse, astrocytes are also critical in the formation, pruning, maintenance, and modulation of synapses. To enable these highly interactive functions, astrocytes communicate among themselves and with other glial cells, neurons, the brain vasculature, and the extracellular environment through a multitude of specialized membrane proteins that include cell adhesion molecules, aquaporins, ion channels, neurotransmitter transporters, and gap junction molecules. To support this dynamic flux, astrocytes, like neurons, rely on tightly coordinated and efficient intracellular transport. Unlike neurons,&#160;where intracellular trafficking has been extensively delineated, microtubule-based transport in astrocytes has been less studied. Nonetheless, exo- and endocytic trafficking of cell membrane proteins and intracellular organelle transport orchestrates astrocytes&#8217; normal biology, and these processes are often affected in disease or in response to injury. Here we present a straightforward protocol to culture high quality murine astrocytes, to fluorescently label astrocytic proteins and organelles of interest, and to record their intracellular transport dynamics using time-lapse confocal microscopy. We also demonstrate how to extract and quantify relevant transport parameters from the acquired movies using available image analysis software (i.e., ImageJ/FIJI) plugins.

The protocol for establishing primary mouse MD astrocytes outlined above should yield reproducible, high quality cultures. Although cultures initially contain a mix of astrocytes, fibroblasts, and other glial cells, including microglia and oligodendrocytes (Figure 1Bi,Biv; red arrowheads), the addition of AraC to the mixed culture between DIV5-DIV7 minimizes the proliferation of these contaminant cells. The combined AraC treatment and shaking-based purification strategy enri...

Watch this Scientific Journal Video about Visualizing and Analyzing Intracellular Transport of Organelles and Other Cargos in Astrocytes at JoVE.com

Visualizing and Analyzing Intracellular Transport of Organelles and Other Cargos in Astrocytes

Here we describe an in vitro live-imaging method to visualize intracellular transport of organelles and trafficking of plasma membrane proteins in murine astrocytes. This protocol also presents an image analysis methodology to determine cargo transport itineraries and kinetics.

Astrocytes are among the most abundant cell types in the adult brain, where they play key roles in a multiplicity of functions. As a central player in ...

visualizing-analyzing-intracellular-transport-organelles-other-cargos

Universidad de Cartagena UDEC

Research

JoVE Journal

Neuroscience

7.7K Views.  University of North Carolina at Chapel Hill. Here we describe an in vitro live-imaging method to visualize intracellular transport of organelles and trafficking of plasma membrane proteins in murine astrocytes. This protocol also presents an image analysis methodology to determine cargo transport itineraries and kinetics.

Video: Visualizing and Analyzing Intracellular Transport of Organelles and Other Cargos in Astrocytes

Fluorescence Recovery After Photobleaching (FRAP) of Fluorescence Tagged Proteins in Dendritic Spines of Cultured Hippocampal Neurons

FRAP has been used to quantify the mobility of GFP-tagged proteins. Using a strong excitation laser, the fluorescence of a GFP-tagged protein is bleached in the region of interest. The fluorescence of the region recovers when the unbleached GFP-tagged protein from outside of the region diffuses into the region of interest. The mobility of the protein is then analyzed by measuring the fluorescence recovery rate. This technique could be used to characterize protein mobility and turnover rate.
In this study, we express the (enhanced green fluorescent protein) EGFP vector in cultured hippocampal neurons. Using the Zeiss 710 confocal microscope, we photobleach the fluorescence signal of the GFP protein in a single spine, and then take time lapse images to record the fluorescence recovery after photobleaching. Finally, we estimate the percentage of mobile and immobile fractions of the GFP in spines, by analyzing the imaging data using ImageJ and Graphpad softwares.
This FRAP protocol shows how to perform a basic FRAP experiment as well as how to analyze the data.

Fluorescence Recovery After Photobleaching FRAP of Fluorescence Tagged Proteins in Dendritic Spines of Cultured Hippocampal Neurons

FRAP has been used to quantify the mobility of Green Fluorescence Protein (GFP)-tagged proteins in cultured cells. We examined the mobile/immobile fractions of the GFP by analyzing the fluorescence recovery percentage after photobleaching. In this study, FRAP was performed at spines of hippocampal neurons.

FRAP has been used to quantify the mobility of GFP-tagged proteins. Using a strong excitation laser, the fluorescence of a GFP-tagged protein is bleached ...

Isolation and Culture of Mouse Cortical Astrocytes

Astrocytes are an abundant cell type in the mammalian brain, yet much remains to be learned about their molecular and functional characteristics. In vitro astrocyte cell culture systems can be used to study the biological functions of these glial cells in detail. This video protocol shows how to obtain pure astrocytes by isolation and culture of mixed cortical cells of mouse pups. The method is based on the absence of viable neurons and the separation of astrocytes, oligodendrocytes and microglia, the three main glial cell populations of the central nervous system, in culture. Representative images during the first days of culture demonstrate the presence of a mixed cell population and indicate the timepoint, when astrocytes become confluent and should be separated from microglia and oligodendrocytes. Moreover, we demonstrate purity and astrocytic morphology of cultured astrocytes using immunocytochemical stainings for well established and newly described astrocyte markers. This culture system can be easily used to obtain pure mouse astrocytes and astrocyte-conditioned medium for studying various aspects of astrocyte biology.

Astrocytes have been recognized to be versatile cells participating in fundamental biological processes that are essential for normal brain development and function, and central nervous system repair. Here we present a rapid procedure to obtain pure mouse astrocyte cultures to study the biology of this major class of central nervous system cells.

Astrocytes  are an abundant cell type in the mammalian brain, yet much remains to be  learned about their molecular and functional characteristics. In ...

Imaging Intracellular Ca2+ Signals in Striatal Astrocytes from Adult Mice Using Genetically-encoded Calcium Indicators

Astrocytes display spontaneous intracellular Ca2+ concentration fluctuations ([Ca2+]i) and in several settings respond to neuronal excitation with enhanced [Ca2+]i signals. It has been proposed that astrocytes in turn regulate neurons and blood vessels through calcium-dependent mechanisms, such as the release of signaling molecules. However, [Ca2+]i imaging in entire astrocytes has only recently become feasible with genetically encoded calcium indicators (GECIs) such as the GCaMP series. The use of GECIs in astrocytes now provides opportunities to study astrocyte [Ca2+]i signals in detail within model microcircuits such as the striatum, which is the largest nucleus of the basal ganglia. In the present report, detailed surgical methods to express GECIs in astrocytes in vivo, and confocal imaging approaches to record [Ca2+]i signals in striatal astrocytes in situ, are described. We highlight precautions, necessary controls and tests to determine if GECI expression is selective for astrocytes and to evaluate signs of overt astrocyte reactivity. We also describe brain slice and imaging conditions in detail that permit reliable [Ca2+]i imaging in striatal astrocytes in situ. The use of these approaches revealed the entire territories of single striatal astrocytes and spontaneous [Ca2+]i signals within their somata, branches and branchlets. The further use and expansion of these approaches in the striatum will allow for the detailed study of astrocyte [Ca2+]i signals in the striatal microcircuitry.

Imaging Intracellular Ca2+ Signals in Striatal Astrocytes from Adult Mice Using Genetically-encoded Calcium Indicators

The properties and functions of astrocyte intracellular Ca2+ signals in the striatum remain incompletely explored. We describe methods to express genetically encoded calcium indicators in striatal astrocytes using adeno-associated viruses of serotype 2/5 (AAV2/5), as well as procedures to reliably image Ca2+ signals within striatal astrocytes in situ.

Astrocytes display spontaneous intracellular Ca2+ concentration fluctuations ([Ca2+]i) and in several settings respond to neuronal excitation with ...

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 ...

Analyzing Dendritic Morphology in Columns and Layers

In many regions of the central nervous systems, such as the fly optic lobes and the vertebrate cortex, synaptic circuits are organized in layers and columns to facilitate brain wiring during development and information processing in developed animals. Postsynaptic neurons elaborate dendrites in type-specific patterns in specific layers to synapse with appropriate presynaptic terminals. The fly medulla neuropil is composed of 10 layers and about 750 columns; each column is innervated by dendrites of over 38 types of medulla neurons, which match with the axonal terminals of some 7 types of afferents in a type-specific fashion. This report details the procedures to image and analyze dendrites of medulla neurons. The workflow includes three sections: (i) the dual-view imaging section combines two confocal image stacks collected at orthogonal orientations into a high-resolution 3D image of dendrites; (ii) the dendrite tracing and registration section traces dendritic arbors in 3D and registers dendritic traces to the reference column array; (iii) the dendritic analysis section analyzes dendritic patterns with respect to columns and layers, including layer-specific termination and planar projection direction of dendritic arbors, and derives estimates of dendritic branching and termination frequencies. The protocols utilize custom plugins built on the open-source MIPAV (Medical Imaging Processing, Analysis, and Visualization) platform and custom toolboxes in the matrix laboratory language. Together, these protocols provide a complete workflow to analyze the dendritic routing of Drosophila medulla neurons in layers and columns, to identify cell types, and to determine defects in mutants.

Here, we show how to analyze dendritic routing of Drosophila medulla neurons in columns and layers. The workflow includes a dual-view imaging technique to improve the image quality and computational tools for tracing, registering dendritic arbors to the reference column array and for analyzing the dendritic structures in 3D space.

In many regions of the central nervous systems, such as the fly optic lobes and the vertebrate cortex, synaptic circuits are organized in layers and ...

Visualization and Live Imaging of Oligodendrocyte Organelles in Organotypic Brain Slices Using Adeno-associated Virus and Confocal Microscopy

Neurons rely on the electric insulation and trophic support of myelinating oligodendrocytes. Despite the importance of oligodendrocytes, the advanced tools currently used to study neurons, have only partly been taken on by oligodendrocyte researchers. Cell type-specific staining by viral transduction is a useful approach to study live organelle dynamics. This paper describes a protocol for visualizing oligodendrocyte mitochondria in organotypic brain slices by transduction with adeno-associated virus (AAV) carrying genes for mitochondrial targeted fluorescent proteins under the transcriptional control of the myelin basic protein promoter. It includes the protocol for making organotypic coronal mouse brain slices. A procedure for time-lapse imaging of mitochondria then follows. These methods can be transferred to other organelles and may be particularly useful for studying organelles in the myelin sheath. Finally, we describe a readily available technique for visualization of unstained myelin in living slices by Confocal Reflectance microscopy (CoRe). CoRe requires no extra equipment and can be useful to identify the myelin sheath during live imaging.

Myelinating oligodendrocytes promote rapid action potential propagation and neuronal survival. Described here is a protocol for oligodendrocyte-specific expression of fluorescent proteins in organotypic brain slices with subsequent time-lapse imaging. Further, a simple procedure for visualizing unstained myelin is presented.

Neurons rely on the electric insulation and trophic support of myelinating oligodendrocytes. Despite the importance of oligodendrocytes, the advanced ...

Analyzing the Size, Shape, and Directionality of Networks of Coupled Astrocytes

It has become increasingly clear that astrocytes modulate neuronal function not only at the synaptic and single-cell levels, but also at the network level. Astrocytes are strongly connected to each other through gap junctions and coupling through these junctions is dynamic and highly regulated. An emerging concept is that astrocytic functions are specialized and adapted to the functions of the neuronal circuit with which they are associated. Therefore, methods to measure various parameters of astrocytic networks are needed to better describe the rules governing their communication and coupling and to further understand their functions.
Here, using the image analysis software (e.g., ImageJFIJI), we describe a method to analyze confocal images of astrocytic networks revealed by dye-coupling. These methods allow for 1) an automated and unbiased detection of labeled cells, 2) calculation of the size of the network, 3) computation of the preferential orientation of dye spread within the network, and 4) repositioning of the network within the area of interest.
This analysis can be used to characterize astrocytic networks of a particular area, compare networks of different areas associated to different functions, or compare networks obtained under different conditions that have different effects on coupling. These observations may lead to important functional considerations. For instance, we analyze the astrocytic networks of a trigeminal nucleus, where we have previously shown that astrocytic coupling is essential for the ability of neurons to switch their firing patterns from tonic to rhythmic bursting1. By measuring the size, confinement, and preferential orientation of astrocytic networks in this nucleus, we can build hypotheses about functional domains that they circumscribe. Several studies suggest that several other brain areas, including the barrel cortex, lateral superior olive, olfactory glomeruli, and sensory nuclei in the thalamus and visual cortex, to name a few, may benefit from a similar analysis.

Here we present a protocol to assess the organization of astrocytic networks. The described method minimizes bias to provide descriptive measures of these networks such as cell count, size, area, and position within a nucleus. Anisotropy is assessed with a vectorial analysis.

It has become increasingly clear that astrocytes modulate neuronal function not only at the synaptic and single-cell levels, but also at the network ...

Analyzing Mitochondrial Transport and Morphology in Human Induced Pluripotent Stem Cell-Derived Neurons in Hereditary Spastic Paraplegia

Neurons have intense demands for high energy in order to support their functions. Impaired mitochondrial transport along axons has been observed in human neurons, which may contribute to neurodegeneration in various disease states. Although it is challenging to examine mitochondrial dynamics in live human nerves, such paradigms are critical for studying the role of mitochondria in neurodegeneration. Described here is a protocol for analyzing mitochondrial transport and mitochondrial morphology in forebrain neuron axons derived from human induced pluripotent stem cells (iPSCs). The iPSCs are differentiated into telencephalic glutamatergic neurons using well-established methods. Mitochondria of the neurons are stained with MitoTracker CMXRos, and mitochondrial movement within the axons are captured using a live-cell imaging microscope equipped with an incubator for cell culture. Time-lapse images are analyzed using software with &#34;MultiKymograph&#34;, &#34;Bioformat importer&#34;, and &#34;Macros&#34; plugins. Kymographs of mitochondrial transport are generated, and average mitochondrial velocity in the anterograde and retrograde directions is read from the kymograph. Regarding mitochondrial morphology analysis, mitochondrial length, area, and aspect ratio are obtained using the ImageJ. In summary, this protocol allows characterization of mitochondrial trafficking along axons and analysis of their morphology to facilitate studies of neurodegenerative diseases.

Impaired mitochondrial transport and morphology are involved in various neurodegenerative diseases. The presented protocol uses induced pluripotent stem cell-derived forebrain neurons to assess mitochondrial transport and morphology in hereditary spastic paraplegia. This protocol allows characterization of mitochondrial trafficking along axons and analysis of their morphology, which will facilitate the study of neurodegenerative disease.

Neurons have intense demands for high energy in order to support their functions. Impaired mitochondrial transport along axons has been observed in human ...

Axonal Transport of Organelles in Motor Neuron Cultures using Microfluidic Chambers System

Motor neurons (MNs) are highly polarized cells with very long axons. Axonal transport is a crucial mechanism for MN health, contributing to neuronal growth, development, and survival. We describe a detailed method for the use of microfluidic chambers (MFCs) for tracking axonal transport of fluorescently labeled organelles in MN axons. This method is rapid, relatively inexpensive, and allows for the monitoring of intracellular cues in space and time. We describe a step by step protocol for: 1) Fabrication of polydimethylsiloxane (PDMS) MFCs; 2) Plating of ventral spinal cord explants and MN dissociated culture in MFCs; 3) Labeling of mitochondria and acidic compartments followed by live confocal imagining; 4) Manual and semiautomated axonal transport analysis. Lastly, we demonstrate a difference in the transport of mitochondria and acidic compartments of HB9::GFP ventral spinal cord explant axons as a proof of the system validity. Altogether, this protocol provides an efficient tool for studying the axonal transport of various axonal components, as well as a simplified manual for MFC usage to help discover spatial experimental possibilities.

Axonal transport is a crucial mechanism for motor neuron health. In this protocol we provide a detailed method for tracking the axonal transport of acidic compartments and mitochondria in motor neuron axons using microfluidic chambers.

Motor neurons (MNs) are highly polarized cells with very long axons. Axonal transport is a crucial mechanism for MN health, contributing to neuronal ...

In Vitro Wedge Slice Preparation for Mimicking In Vivo Neuronal Circuit Connectivity

In vitro slice electrophysiology techniques measure single-cell activity with precise electrical and temporal resolution. Brain slices must be relatively thin to properly visualize and access neurons for patch-clamping or imaging, and in vitro examination of brain circuitry is limited to only what is physically present in the acute slice. To maintain the benefits of in vitro slice experimentation while preserving a larger portion of presynaptic nuclei, we developed a novel slice preparation. This &#8220;wedge slice&#8221; was designed for patch-clamp electrophysiology recordings to characterize the diverse monaural, sound-driven inputs to medial olivocochlear (MOC) neurons in the brainstem. These neurons receive their primary afferent excitatory and inhibitory inputs from neurons activated by stimuli in the contralateral ear and corresponding cochlear nucleus (CN). An asymmetrical brain slice was designed which is thickest in the rostro-caudal domain at the lateral edge of one hemisphere and then thins towards the lateral edge of the opposite hemisphere. This slice contains, on the thick side, the auditory nerve root conveying information about auditory stimuli to the brain, the intrinsic CN circuitry, and both the disynaptic excitatory and trisynaptic inhibitory afferent pathways that converge on contralateral MOC neurons. Recording is performed from MOC neurons on the thin side of the slice, where they are visualized using DIC optics for typical patch-clamp experiments. Direct stimulation of the auditory nerve is performed as it enters the auditory brainstem, allowing for intrinsic CN circuit activity and synaptic plasticity to occur at synapses upstream of MOC neurons. With this technique, one can mimic in vivo circuit activation as closely as possible within the slice. This wedge slice preparation is applicable to other brain circuits where circuit analyses would benefit from preservation of upstream connectivity and long-range inputs, in combination with the technical advantages of in vitro slice physiology.

Integration of diverse synaptic inputs to neurons is best measured in a preparation that preserves all pre-synaptic nuclei for natural timing and circuit plasticity, but brain slices typically sever many connections. We developed a modified brain slice to mimic in vivo circuit activity while maintaining in vitro experimentation capability.

In vitro slice electrophysiology techniques measure single-cell activity with precise electrical and temporal resolution. Brain slices must be relatively ...