Using Live-Cell Imaging to Measure the Effects of Pathological Proteins on Axonal Transport in Primary Hippocampal Neurons

These protocols were approved by the Michigan State University Institutional Animal Care and Use Committee. This protocol has been successfully applied to Tau Knockout mice in the C57BL/6J background and wild-type Sprague Dawley rats. Other strains should be acceptable as well.

1. Primary hippocampal neuron harvest

1. Coat a 4-well glass bottom chamber slide with filtered 0.5 mg/mL poly-d-lysine (PDL) in borate buffer (12.5 mM sodium borate decahydrate and 50 mM boric acid). Add 750 µL/well and incubate at room temperature overnight.
2. Wash the chamber slide 4x with sterile deionized water. Air dry after the last wash and keep at 4 °C for short-term storage (<1 week).
   NOTE: Do not allow the PDL to dry on the slide prior to and between washes as this is toxic to the cultures. We find that longer-term storage of PDL-coated slides may affect neuronal health and that maintaining consistent intervals between coating and plating reduces inter-run variability in culture viability.
3. Obtain a timed-pregnant mouse or rat.
   NOTE: Timed-pregnant animals can be ordered from various companies or bred in-house. Embryonic day zero refers to the day the vaginal plug is found, and we have had success culturing E16-18 mouse primary hippocampal and cortical neurons with this approach. Rat neurons are also harvested on embryonic day 18.
4. On E18, euthanize the pregnant female by an approved method. Cut through the abdominal wall and remove the two uterine horns. Transfer them to a Petri dish containing ice-cold 0.9% saline. Rinse until the saline is clean.
   NOTE: We administer an overdose of sodium pentobarbital (at least 100 mg/kg) via intraperitoneal injection.
5. Decapitate the pups. Beginning at the foramen magnum (the opening in the back of the skull), use microdissection scissors to cut the skin and skull along the midline, angling the lower tip of the scissors up against the internal part of the skull to avoid damage to the brain tissue.
6. Remove the overlying skin and carefully remove the skull to expose the brain.
7. Use forceps to hold the mouth/nose area and turn the skull so the brain is facing downward over a Petri dish filled with 0.9% saline. Flip the brain out of the skull and cut the nerves and brain stem to release the brain into the saline.
8. Place the Petri dish under a dissecting microscope for dissection of the hippocampus. Insert scissors into the brain's midline, angle the scissors so that the top blade is pushing one of the cerebral hemispheres outward, and cut to separate the hemisphere away from the subcortical regions.
9. Make a vertical cut through the cortex so that the frontal cortex is anterior to, and the hippocampus is posterior to, the cut site. Insert the lower tip of the scissors into the resulting hole and carefully cut along the apex of the cortex stopping at the posterior end.
10. Using the forceps, carefully swing out the cortex and finish the posterior cut. The hippocampus should resemble a crescent shape. Carefully trim the remaining cortex so the hippocampus maintains its rounded, crescent shape.
11. Using forceps and scissors, gently remove the meninges from the hippocampus.
    NOTE: It is important to remove all the meninges from the brain to reduce the number of non-neuronal cells that are collected while refraining from damaging or tearing the hippocampal tissue.
12. Cut the hippocampus into four equal pieces and place the tissue into a 15 mL conical tube filled with ice-cold calcium- and magnesium-free buffer (CMF; Dulbecco's PBS with 0.1% glucose, 2.5 µg/mL Amphotericin B, and 50 µg/mL Gentamicin).
    ​NOTE: For best results, collect the hippocampi from 5-10 pups per dissociation. Each hippocampus should provide approximately 300,000 cells for mice and 500,000 cells for rats.

2. Primary hippocampal neuron dissociation and plating

1. Remove the CMF with a Pasteur pipette and rinse the tissue with fresh cold CMF 4x, swirling gently in between rinses.
2. Remove the CMF and add 3 mL of filtered 0.125% trypsin solution (dilute 2.5% trypsin in prewarmed [37 °C] CMF). Incubate the tissue for exactly 15 min at 37 °C; gently swirl every 5 min.
   NOTE: Make the trypsin solution immediately before use.
3. Prepare 5 mL of trypsin inactivation solution (TIS; Hanks' Balanced Salt Solution, 20% newborn calf serum and 1x DNase I solution [10x stock: 5 mM sodium acetate, 1 µM calcium chloride, 0.5 mg/mL DNase I]).
4. Remove the trypsin solution and wash 2x with cold CMF. Add 3 mL of cold TIS.
5. Dissociate cells through trituration using a 3 mL syringe with progressively smaller diameter needles: triturate 30x using a 14 G needle, 30x using a 15 G needle, 20x using a 16 G needle, 20x using an 18 G needle, and 15x using a 21 G needle. Use 2 s draws with the first four needles and 3 s draws with the 21 G needle.
   NOTE: Other methods of trituration (e.g., fire-polished glass pipets^14,15) may also be used. We find that using the standardized metal syringe needles substantively reduces variability in cell clumping and viability.
6. Using the 21 G needle, place a droplet of the cell suspension on a microscope slide and check for cell clumps. If several clumps are observed, triturate three more times through a 22 G needle and re-check. Repeat this process until a homogeneous single-cell suspension is obtained, being careful to triturate gently throughout the process.
7. Filter 5 mL of fetal bovine serum (FBS) using a 0.22 µm syringe filter. Gently layer the cell suspension onto the FBS in a 15 mL conical tube. Centrifuge the cells (200 × g at 4 °C) for 5 min.
8. Remove as much FBS as possible without disrupting the pellet. Resuspend in 1 mL of warm (37 °C) antibiotic-free neurobasal media (NBM; supplemented with 1x glutamine substitute and 1x B-27).
9. Dilute the cell suspension in Trypan blue (0.4%) and obtain a cell count.
   NOTE: A 1:1 dilution works well for an automatic cell counter and use a 1:60 dilution for manual counting with a hemacytometer. To continue onto cell plating, the neuron viability should be higher than 90% at this point. This significantly enhances the likelihood of a healthy neuron culture after dissociation.
10. Plate the cells at a density of 175,000 cells/well (70,000 cells/mm²) in a prewarmed PDL-coated four-well chamber slide in 750 µL of antibiotic-free NBM. Additionally, plate 200,000 cells/well in four wells of a PDL-coated 24-well plate in 1.5 mL of antibiotic-free NBM for use as conditioned medium in the transfection described in section 3. Incubate the cells in a humidity-controlled, 37 °C, and 5% CO₂ chamber.
    ​NOTE: The density of cells can be adjusted based on the results. High densities can make it more difficult to image the neurons while low densities can negatively affect the health of the culture and neuron survival. In our experience, rat neurons can be plated at a lower density compared to mouse neurons.

3. Neuron transfection

NOTE: Neurons can be transfected on DIV 7 or DIV 8 without notable changes in transfection efficiency or transport results. These volumes are for four wells of a four-well glass-bottom chamber slide in 750 µL volume/well. This protocol describes transfection using lipid transfection reagents. Allow media and transfection reagents to warm to room temperature prior to use.

1. Prepare transfection stock reagent by adding 4.4 µL of transfection reagent to 127.6 µL of reduced-serum medium. Rock gently by hand to mix and incubate at room temperature for 30 min.
2. Prepare DNA stocks during the lipid transfection incubation. Add 200 ng of fluorescent-cargo DNA (pFIN mApple-synaptophysin; 12,112 base pairs) and 200 ng of DNA to express the protein of interest (tau plasmids used were pCMV tau-Flag constructs; 5,036 base pairs) into reduced-serum or serum-free medium to a final volume of 32 µL¹². Add 0.8 µL of transfection enhancer reagent (2 µL per µg of DNA). Mix well by hand.
   NOTE: We find that a derivative of RFP works best (mApple or mCherry), but GFP can be used as well. Additionally, a DNA construct to express a gene-silencing small hairpin RNA (shRNA) can be included to test mechanisms of phenotypes observed with the expression of the protein of interest. To do this, include 150 ng of each of the three DNA constructs in the DNA mix and adjust the amount of transfection enhancer reagent accordingly. Changing the parameters of DNA or lipofection reagent can affect the transfection efficiency and can be adjusted based on the need for more or less efficient transfections.
3. Add 32 µL of lipid transfection mix to each tube of DNA. Rock gently by hand to mix. Incubate at room temperature for 30 min.
4. Slowly add 64 µL of DNA/transfection reagent mix to the cells by placing the tip of the pipette just below the surface of the medium and moving in a serpentine motion throughout the well. Gently rock the chamber slide by hand 3x after addition to all wells to mix further and place the chamber slide in the incubator. Incubate at 37 °C and 5% CO₂ for 2 h.
   NOTE: Minimize the amount of time neurons are outside of the incubator as they are very sensitive to changes in temperature and pH, which can occur quickly.
5. Perform a half medium change 2 h after transfection with conditioned media + Neurofascin antibody. Immediately prior to the medium change, pool 1.5 mL of conditioned antibiotic-free NBM from the cells cultured on the 24-well plate. Add 0.75 µL of Neurofascin 186 (NF-186) antibody (500 ng/mL) to the conditioned medium.
   NOTE: The NF-186 antibody detects an extracellular domain of neurofascin that is enriched in the axon initial segment (AIS) to definitively identify the axon for imaging.
6. Perform a full medium change 18 h after transfection with conditioned medium containing the secondary antibody. Add 1 µL of goat-anti-rabbit secondary antibody conjugated to AlexaFluor 647 to 3 mL of conditioned NBM (antibiotic-free) pooled from the 24-well plate. Mix well.

4. Neuron imaging

1. Prepare for imaging 2 days after transfection. Warm the live cell chamber attachment for the confocal microscope to 37 °C (including the 60x objective and lens oil) and equilibrate the chamber to 5% CO₂.
2. Use a high magnification objective and live-cell chamber to image the cells.
   NOTE: We use a 60x 1.40 NA objective on a laser scanning confocal microscope but other confocal (e.g., spinning disc) or widefield fluorescence microscopy systems can also be used if they are able to achieve high imaging frame rates.
3. Visually identify transfected neurons in the channel containing the cargo protein using the eyepieces. Next, image the neurons to identify the AIS in the Cy5 channel (Figure 1).
4. Set the field of view to a rectangular box of 32 pixels by 128 pixels and move it to image a region of the axon ~50-150 µm distal to the AIS. Note and record the directionality of the cargo transport and the orientation of the ROI. The dots on the box will appear as the top and right, respectively, of any subsequent images and movies. Record which side of the image is closest to the cell body and which is closest to the axon terminus so the kymograph polyline in step 5.1 can be drawn with the correct orientation.
   NOTE: When selecting the region of interest, the imaged area of an isolated axon should be relatively straight and intersect the field of view at each end. The entire segment should be within the same focal plane. The area should clearly display active transport of cargo. Excessively bright neurons typically express the cargo protein and proteins of interest at levels that are likely to be toxic to the neuron and should be avoided.
5. Set the 561 laser power a 1.40, the pinhole a 7.8, the Size a 128 px², and the Speed a 32 frames/s. Adjust the gain to visualize the transport of individual cargo vesicles without it being obscured by background signal (typically between 10 and 50 results in high-quality kymographs).
   NOTE: The extremely low laser power is achievable because of highly sensitive GaAsP detectors and prevents potential phototoxic effects and photobleaching of moving cargo. The pinhole size is set as wide as possible to increase the focal depth to ensure that the entire ROI is imaged as the axon may not lay perfectly flat on the surface of the slide.
6. Set the ROI by selecting Draw Rectangular ROI… from the ROI dropdown menu. Draw the ROI so that it covers the entire field of view. Right-click on the ROI and select Use as Stimulation ROI: S1.
7. Prepare the ND Setup by including the following steps in the ND Setup tab.
   NOTE: This is the five-step acquisition protocol that is set up once and then will be performed by the software each time the movies are collected.
   1. Acquire Image to collect the prestimulation reference image.
   2. Stimulation; ROI: S1; Interval: NoDelay; Duration 1.64 s, Loops: 7. This step bleaches background fluorescence with several pulses of the laser.
   3. Acquire Image. This step collects a poststimulation reference image.
   4. Waiting; No Action; 2 min. This step provides a recovery period for fluorescent cargo to repopulate the bleached ROI.
   5. Acquisition; Interval: NoDelay; Duration 5 min, Loops: 8,615. This step collects images at the maximum rate for the 5 min imaging period.
8. After the ND Setup protocol is set up, click the Apply Stimulation Settings button on the ND Setup tab. Click Run Now when ready to image cargo transport. This will initiate the ND Acquisition protocol from step 4.7.
   NOTE: Although the frame rate is set to 32 frames/s, the practical frame rate may be slower (~28.7 frames/s). This value should be recorded for accurate kymograph analysis in section 5. Ideally, live-cell transport imaging is done with the fastest frame rate possible (e.g., >25 frames/s) to better decipher cargo movements.
9. Reset the field of view to the entire image; then, collect and save an image of the full neuron in the 561 and 647 channels with the ROI box included. Record the XY coordinates where the image was taken for postfixation confirmation of the expression of the protein of interest.
10. Collect transport videos from at least two neurons for analysis. Ensure that each independent replicate comes from the means of variables measured from an individual primary neuron harvest (i.e., not individual neurons).
    NOTE: For a given primary neuron harvest, we collect videos from 2-4 neurons per condition.
11. Fix the cells by removing the medium and incubating with prewarmed 4% paraformaldehyde for 20 min at room temperature. Remove the buffer and wash cells for 3 x 5 min each in TBS.
    NOTE: For paraformaldehyde fixation, we use cytoskeleton buffer, a common buffer that maintains cytoskeletal structure (10 mM 2-(N-morpholino)ethanesulfonic acid [MES], 138 mM KCl, 3 mM MgCl₂, and 4 mM EGTA, pH 6.1). Other buffers (e.g., TBS, PBS) can also be used for fixation.
12. Confirm that the analyzed neurons express both the protein of interest and the fluorescently tagged cargo protein through immunocytofluorescence staining. Stain for human tau (Tau12) to confirm exogenous tau expression and β-III tubulin (Tuj1) to verify that the neurons remained healthy throughout the imaging process.
    ​NOTE: We observe co-expression of both DNA constructs in nearly all transfected cells using these methods.

5. Generate and analyze Kymographs

NOTE: Kymographs can be generated and analyzed using a variety of different programs. We briefly describe the freely-available KymoAnalyzer (v. 1.01) software using six ImageJ (v. 1.51n) plugins¹⁶. These plugins can be downloaded from the developers at the Encalada lab website (https://www.encalada.scripps.edu/kymoanalyzer). More detailed instructions on the use of this software can be found at this site.

1. Group the movies generated in step 4.9 based on experimental conditions and replicate. Run the BatchKymographGeneration macro and select the appropriate folder. Draw a kymograph polyline to trace the axon beginning with the proximal end in that particular image. The resulting kymographs are stored in the selected folder (Figure 2).
2. Assign tracks manually by running the Tracks macro, selecting an individual movie folder, selecting yes to add a track, and then clicking to track each particle's trajectory. Perform the initial click at the top of the track and click again at every point the particle changes velocity or direction so that the polyline overlays the particle's trace on the kymograph. After the polyline is laid over the entire track, click OK. Repeat until all tracks are assigned.
3. Run the remaining macros, assigning frame rate and pixel size based on imaging parameters. First, run CargoPopulation, then NetCargoPopulation, and then Segments. Enter the pixel size and frame rate of the movies when prompted (0.22 µm and 28.7 frames/s, respectively, with these settings). Finally, run the poolData macro to calculate and pool transport parameter data from all kymographs in the selected folder.
4. Plot the results from the data files and compare the various conditions with the independent replicates represented by the mean values of all analyzed neurons within an experimental condition for an individual neuron harvest.