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

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 deVellis^13,14,15. Mixed cell cultures of McCarthy/deVellis (MD) astrocytes are prepared from postnatal day 2−4 (P2−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.

1. Coat the bottom of T75 flasks (100% angled neck access, 0.22 µm hydrophobic vented filter cap) with poly-D-lysine (1 mg/mL in 0.1 M Tris buffer, pH 8.5) and incubate at 37 °C for 24 h prior to start of dissection.
2. Before beginning the dissection procedure, warm 30 mL of astrocyte culture media (Dulbecco’s modified Eagle’s medium [DMEM] high glucose, 10% heat-inactivated fetal bovine serum, 1% penicillin/streptomycin) at 37 °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’s balanced salt solution (HBSS) without Ca²⁺ or Mg²⁺.
3. 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.
4. Spray the head and neck of the P2−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.
5. 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.
6. 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).
7. 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.
8. Cut the dissected cortices into small pieces (approximately 2−4 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 °C for 30 min with gentle inversion every 10 min.
9. 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 20−30x using a 10 mL serological pipette.
10. Filter the cell suspension through a 70 µm 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 µL of the cell suspension with 10 µL of trypan blue and counting cells with a hemocytometer.
    NOTE: One preparation from four P2−P4 mouse cortices yields 10−15 x 10⁶ single cells.
11. 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 °C and 5% CO₂.

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 µm filter membrane) astrocyte media warmed to 37 °C.

1. Replace the cell media 48 h after plating (day in vitro 2, DIV2).
2. At DIV5, replace the media with fresh astrocyte culture media supplemented with 10 µM 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.
3. 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 °C for at least 8 h to overnight.
4. The following day, begin the astrocyte purification by shaking the culture flasks at 180 rpm for 30 min in a 37 °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 °C orbital shaking incubator. Prewarm 1x phosphate-buffered saline (PBS), astrocyte culture media, and 0.25% trypsin-EDTA to 37 °C 20−30 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.
5. 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 °C and 5% CO₂, checking every 5 min for detachment.
6. 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.
7. 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 °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.
8. Repeat steps 2.5−2.7 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).

3. Transfection of fluorescently tagged plasmids

1. The day prior to transfection or labeling, seed astrocytes at desired density for imaging 24−48 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.
2. 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 μL of the lipofection reagent in 50 μL of reduced-serum media.
3. Dilute 5 μg of high purity DNA in 250 μL of reduced-serum media. Add 5 μL 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.
4. Remove astrocyte culture media and add transfection mix (step 3.3) to cells dropwise. After a 6-h incubation at 37 °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 24−72 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.

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.

1. Dilute the lysosomal-labeling probe (Table of Materials) in 200 µL of astrocyte culture media to a working concentration of 1 μM and apply to astrocytes from step 3.1 at a density of 2 x 10⁴ cells/well. Incubate for 30 min at 37 °C.
2. Wash the cells once with warm astrocyte culture media and replace with imaging media (Table of Materials). Proceed to live imaging immediately.

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.

1. 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.
2. 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.
3. Save time-lapse images and export them as AVI or TIFF stack files.

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.

1. 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.
2. 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.
3. 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.
4. 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.
   1. Run the Analyze Kymo macro of the KymoToolBox plugin. A window will open asking to define the "outward" 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 "from left to right".
   2. 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 µm/second is an appropriate choice. Modify this value to adjust the software's sensitivity to each cargo of interest. Adjust the line width to match the thickness of each particle trajectory.
   3. 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’s position on the original time-lapse video and kymograph.