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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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&#039;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&#039;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 &amp;micro;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

Measuring Near Plasma Membrane and Global Intracellular Calcium Dynamics in Astrocytes

The brain contains glial cells.  Astrocytes, a type of glial cell, have long been known to provide a passive supportive role to neurons. However, increasing evidence suggests that astrocytes may also actively participate in brain function through functional interactions with neurons.  However, many fundamental aspects of astrocyte biology remain controversial, unclear and/or experimentally unexplored. One important issue is the dynamics of intracellular calcium transients in astrocytes. This is relevant because calcium is well established as an important second messenger and because it has been proposed that astrocyte calcium elevations can trigger the release of transmitters from astrocytes. However, there has not been any detailed or satisfying description of near plasma membrane calcium signaling in astrocytes. Total internal reflection fluorescence (TIRF) microscopy is a powerful tool to analyze physiologically relevant signaling events within about 100 nm of the plasma membrane of live cells. Here, we use TIRF microscopy and describe how to monitor near plasma membrane and global intracellular calcium dynamics almost simultaneously. The further refinement and systematic application of this approach has the potential to inform about the precise details of astrocyte calcium signaling. A detailed understanding of astrocyte calcium dynamics may provide a basis to understand if, how, when and why astrocytes and neurons undergo calcium-dependent functional interactions.

We describe how to measure near membrane and global intracellular calcium dynamics in cultured astrocytes using total internal reflection and epifluorescence microscopy.

The brain contains glial cells.  Astrocytes, a type of glial cell, have long been known to provide a passive supportive role to neurons. However, ...

Biology

Deriving the Time Course of Glutamate Clearance with a Deconvolution Analysis of Astrocytic Transporter Currents

The highest density of glutamate transporters in the brain is found in astrocytes. Glutamate transporters couple the movement of glutamate across the membrane with the co-transport of 3 Na+ and 1 H+ and the counter-transport of 1 K+. The stoichiometric current generated by the transport process can be monitored with whole-cell patch-clamp recordings from astrocytes. The time course of the recorded current is shaped by the time course of the glutamate concentration profile to which astrocytes are exposed, the kinetics of glutamate transporters, and the passive electrotonic properties of astrocytic membranes. Here we describe the experimental and analytical methods that can be used to record glutamate transporter currents in astrocytes and isolate the time course of glutamate clearance from all other factors that shape the waveform of astrocytic transporter currents. The methods described here can be used to estimate the lifetime of flash-uncaged and synaptically-released glutamate at astrocytic membranes in any region of the central nervous system during health and disease.

We describe an analytical method to estimate the lifetime of glutamate at astrocytic membranes from electrophysiological recordings of glutamate transporter currents in astrocytes.

The highest density of glutamate transporters in the brain is found in astrocytes. Glutamate transporters couple the movement of glutamate across the ...

Inducing Plasticity of Astrocytic Receptors by Manipulation of Neuronal Firing Rates

Close to two decades of research has established that astrocytes in situ and in vivo express numerous G protein-coupled receptors (GPCRs) that can be stimulated by neuronally-released transmitter. However, the ability of astrocytic receptors to exhibit plasticity in response to changes in neuronal activity has received little attention. Here we describe a model system that can be used to globally scale up or down astrocytic group I metabotropic glutamate receptors (mGluRs) in acute brain slices. Included are methods on how to prepare parasagittal hippocampal slices, construct chambers suitable for long-term slice incubation, bidirectionally manipulate neuronal action potential frequency, load astrocytes and astrocyte processes with fluorescent Ca2+ indicator, and measure changes in astrocytic Gq GPCR activity by recording spontaneous and evoked astrocyte Ca2+ events using confocal microscopy. In essence, a &#8220;calcium roadmap&#8221; is provided for how to measure plasticity of astrocytic Gq GPCRs. Applications of the technique for study of astrocytes are discussed. Having an understanding of how astrocytic receptor signaling is affected by changes in neuronal activity has important implications for both normal synaptic function as well as processes underlying neurological disorders and neurodegenerative disease.

Here we describe an adaptation of protocols used to induce homeostatic plasticity in neurons for the study of plasticity of astrocytic G protein-coupled receptors. Recently used to examine changes in astrocytic group I mGluRs in juvenile mice, the method can be applied to measure scaling of various astrocytic GPCRs, in tissue from adult mice in situ and in vivo, and to gain a better appreciation of the sensitivity of astrocytic receptors to changes in neuronal activity.

Close to two decades of research has established that astrocytes in situ and in vivo express numerous G protein-coupled receptors (GPCRs) that can be ...

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 ...

Three-dimensional Tissue Engineered Aligned Astrocyte Networks to Recapitulate Developmental Mechanisms and Facilitate Nervous System Regeneration

Neurotrauma and neurodegenerative disease often result in lasting neurological deficits due to the limited capacity of the central nervous system (CNS) to replace lost neurons and regenerate axonal pathways. However, during nervous system development, neuronal migration and axonal extension often occur along pathways formed by other cells, referred to as &#34;living scaffolds&#34;. Seeking to emulate these mechanisms and to design a strategy that circumvents the inhibitory environment of the CNS, this manuscript presents a protocol to fabricate tissue engineered astrocyte-based &#34;living scaffolds&#34;. To create these constructs, we employed a novel biomaterial encasement scheme to induce astrocytes to self-assemble into dense three-dimensional bundles of bipolar longitudinally-aligned somata and processes. First, hollow hydrogel micro-columns were assembled, and the inner lumen was coated with collagen extracellular-matrix. Dissociated cerebral cortical astrocytes were then delivered into the lumen of the cylindrical micro-column and, at a critical inner diameter of &#60;350 &#181;m, spontaneously self-aligned and contracted to produce long fiber-like cables consisting of dense bundles of astrocyte processes and collagen fibrils measuring &#60;150 &#181;m in diameter yet extending several cm in length. These engineered living scaffolds exhibited &#62;97% cell viability and were virtually exclusively comprised of astrocytes expressing a combination of the intermediate filament proteins glial-fibrillary acidic protein (GFAP), vimentin, and nestin. These aligned astrocyte networks were found to provide a permissive substrate for neuronal attachment and aligned neurite extension. Moreover, these constructs maintain integrity and alignment when extracted from the hydrogel encasement, making them suitable for CNS implantation. These preformed constructs structurally emulate key cytoarchitectural elements of naturally occurring glial-based &#34;living scaffolds&#34; in vivo. As such, these engineered living scaffolds may serve as test-beds to study neurodevelopmental mechanisms in vitro or facilitate neuroregeneration by directing neuronal migration and/or axonal pathfinding following CNS degeneration in vivo.

We showcase the development of self-assembled, three-dimensional bundles of longitudinally aligned astrocytic somata and processes within a novel biomaterial encasement. These engineered &#34;living scaffolds&#34;, exhibiting micron-scale diameter yet extending centimeters in length, may serve as test-beds to study neurodevelopmental mechanisms or facilitate neuroregeneration by directing neuronal migration and/or axonal pathfinding.

Neurotrauma and neurodegenerative disease often result in lasting neurological deficits due to the limited capacity of the central nervous system (CNS) to ...

Bioengineering

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 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 ...

A Method for 3D Reconstruction and Virtual Reality Analysis of Glial and Neuronal Cells

Serial sectioning and subsequent high-resolution imaging of biological tissue using electron microscopy (EM) allow for the segmentation and reconstruction of high-resolution imaged stacks to reveal ultrastructural patterns that could not be resolved using 2D images. Indeed, the latter might lead to a misinterpretation of morphologies, like in the case of mitochondria; the use of 3D models is, therefore, more and more common and applied to the formulation of morphology-based functional hypotheses. To date, the use of 3D models generated from light or electron image stacks makes qualitative, visual assessments, as well as quantification, more convenient to be performed directly in 3D. As these models are often extremely complex, a virtual reality environment is also important to be set up to overcome occlusion and to take full advantage of the 3D structure. Here, a step-by-step guide from image segmentation to reconstruction and analysis is described in detail.

The described pipeline is designed for the segmentation of electron microscopy datasets larger than gigabytes, to extract whole-cell morphologies. Once the cells are reconstructed in 3D, customized software designed around individual needs can be used to perform a qualitative and quantitative analysis directly in 3D, also using virtual reality to overcome view occlusion.

Serial sectioning and subsequent high-resolution imaging of biological tissue using electron microscopy (EM) allow for the segmentation and reconstruction ...

Visualizing Astrocyte Morphology Using Lucifer Yellow Iontophoresis

Astrocytes are essential components of neural circuits. They tile the entire central nervous system (CNS) and are involved in a variety of functions, which include neurotransmitter clearance, ion regulation, synaptic modulation, metabolic support to neurons, and blood flow regulation. Astrocytes are complex cells that have a soma, several major branches, and numerous fine processes that contact diverse cellular elements within the neuropil. In order to assess the morphology of astrocytes, it is necessary to have a reliable and reproducible method to visualize their structure. We report a reliable&#160;protocol to perform intracellular iontophoresis of astrocytes using fluorescent Lucifer yellow (LY) dye in lightly fixed brain tissue from adult mice. This method has several features that are useful to characterize astrocyte morphology. It allows for three-dimensional reconstruction of individual astrocytes, which is useful to perform morphological analyses on different aspects of their structure. Immunohistochemistry together with LY iontophoresis can also be utilized to understand the interaction of astrocytes with different components of nervous system and to evaluate the expression of proteins within the labelled astrocytes. This protocol can be implemented in a variety of mouse models of CNS disorders to rigorously examine astrocyte morphology with light microscopy. LY iontophoresis provides an experimental approach to evaluate astrocyte structure, especially in the context of injury or disease where these cells are proposed to undergo significant morphological changes.

Astrocytes are morphologically complex cells, exemplified by their multiple processes and bushy territories. To analyze their elaborate morphology, we present a reliable protocol to perform intracellular Lucifer yellow iontophoresis in lightly fixed tissue.

Astrocytes are essential components of neural circuits. They tile the entire central nervous system (CNS) and are involved in a variety of functions, ...

Investigation of Spatial Interaction Between Astrocytes and Neurons in Cleared Brains

Combining viral vector transduction and tissue clearing using the CLARITY method makes it possible to simultaneously investigate several types of brain cells and their interactions. Viral vector transduction enables the marking of diverse cell types in different fluorescence colors within the same tissue. Cells can be identified genetically by activity or projection. Using a modified CLARITY protocol, the potential sample size of astrocytes and neurons has grown by 2-3 orders of magnitude. The use of CLARITY allows the imaging of complete astrocytes, which are too large to fit in their entirety in slices, and the examination of the somata with all their processes. In addition, it provides the opportunity to investigate the spatial interaction between astrocytes and different neuronal cell types, namely, the number of pyramidal neurons in each astrocytic domain or the proximity between astrocytes and specific inhibitory neuron populations. This paper describes, in detail, how these methods are to be applied.

Combining viral vector transduction and brain clearing using the CLARITY method allows the investigation of a large number of neurons and astrocytes simultaneously.

Combining viral vector transduction and tissue clearing using the CLARITY method makes it possible to simultaneously investigate several types of brain ...