We thank Chelsea Tiernan and Kyle Christensen for their efforts in developing and optimizing aspects of these protocols. This work was supported by National Institutes of Health (NIH) Grants R01 NS082730 (N.M.K.), R01 AG044372 (N.M.K.), R01 AG067762 (N.M.K.), and F31 AG074521 (R.L.M.); NIH/National Institute on Aging, Michigan Alzheimer&#39;s Disease Research Center Grant 5P30AG053760 (N.M.K. and B.C.); Office of the Assistant Secretary of Defense for Health Affairs through the Peer Reviewed Alzheimer&#39;s Research Program Award W81XWH-20-1-0174 (B.C.); Alzheimer&#39;s Association Research Grants 20-682085 (B.C.); and the Secchia Family Foundation (N.M.K.).

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
<ol>
	<li>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 &#181;L/well and incubate at room temperature overnight.</li>
	<li>Wash the chamber slide 4x with sterile deionized water. Air dry after the last wash and keep at 4 &#176;C for short-term storage (&#60;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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Remove the overlying skin and carefully remove the skull to expose the brain.</li>
	<li>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.</li>
	<li>Place the Petri dish under a dissecting microscope for dissection of the hippocampus. Insert scissors into the brain&#39;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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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&#39;s PBS with 0.1% glucose, 2.5 &#181;g/mL Amphotericin B, and 50 &#181;g/mL Gentamicin). 
	&#8203;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.</li>
</ol>
2. Primary hippocampal neuron dissociation and plating
<ol>
	<li>Remove the CMF with a Pasteur pipette and rinse the tissue with fresh cold CMF 4x, swirling gently in between rinses.</li>
	<li>Remove the CMF and add 3 mL of filtered 0.125% trypsin solution (dilute 2.5% trypsin in prewarmed [37 &#176;C] CMF). Incubate the tissue for exactly 15 min at 37 &#176;C; gently swirl every 5 min. 
	NOTE: Make the trypsin solution immediately before use.</li>
	<li>Prepare 5 mL of trypsin inactivation solution (TIS; Hanks&#39; Balanced Salt Solution, 20% newborn calf serum and 1x DNase I solution [10x stock: 5 mM sodium acetate, 1 &#181;M calcium chloride, 0.5 mg/mL DNase I]).</li>
	<li>Remove the trypsin solution and wash 2x with cold CMF. Add 3 mL of cold TIS.</li>
	<li>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 pipets14,15) may also be used. We find that using the standardized metal syringe needles substantively reduces variability in cell clumping and viability.</li>
	<li>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.</li>
	<li>Filter 5 mL of fetal bovine serum (FBS) using a 0.22 &#181;m syringe filter. Gently layer the cell suspension onto the FBS in a 15 mL conical tube. Centrifuge the cells (200 &#215; g at 4 &#176;C) for 5 min.</li>
	<li>Remove as much FBS as possible without disrupting the pellet. Resuspend in 1 mL of warm (37 &#176;C) antibiotic-free neurobasal media (NBM; supplemented with 1x glutamine substitute and 1x B-27).</li>
	<li>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.</li>
	<li>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 &#181;L 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 &#176;C, and 5% CO2 chamber. 
	&#8203;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.</li>
</ol>
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 &#181;L volume/well. This protocol describes transfection using lipid transfection reagents. Allow media and transfection reagents to warm to room temperature prior to use.
<ol>
	<li>Prepare transfection stock reagent by adding 4.4 &#181;L of transfection reagent to 127.6 &#181;L of reduced-serum medium. Rock gently by hand to mix and incubate at room temperature for 30 min.</li>
	<li>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 &#181;L12. Add 0.8 &#181;L of transfection enhancer reagent (2 &#181;L per &#181;g 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.</li>
	<li>Add 32 &#181;L of lipid transfection mix to each tube of DNA. Rock gently by hand to mix. Incubate at room temperature for 30 min.</li>
	<li>Slowly add 64 &#181;L 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 &#176;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.</li>
	<li>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 &#181;L 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.</li>
	<li>Perform a full medium change 18 h after transfection with&#160;conditioned medium containing the secondary antibody. Add 1 &#181;L 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.</li>
</ol>
4. Neuron imaging
<ol>
	<li>Prepare for imaging 2 days after transfection. Warm the live cell chamber attachment for the confocal microscope to 37 &#176;C (including the 60x objective and lens oil) and equilibrate the chamber to 5% CO2.</li>
	<li>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.</li>
	<li>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).</li>
	<li>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 &#181;m 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.</li>
	<li>Set the 561 laser power to 1.40, the pinhole to 7.8, the Size to 128 px2, and the Speed to 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.</li>
	<li>Set the ROI by selecting Draw Rectangular ROI&#8230; 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.</li>
	<li>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.
	<ol>
		<li>Acquire Image to collect the prestimulation reference image.</li>
		<li>Stimulation; ROI: S1; Interval: NoDelay; Duration 1.64 s, Loops: 7. This step bleaches background fluorescence with several pulses of the laser.</li>
		<li>Acquire Image. This step collects a poststimulation reference image.</li>
		<li>Waiting; No Action; 2 min. This step provides a recovery period for fluorescent cargo to repopulate the bleached ROI.</li>
		<li>Acquisition; Interval: NoDelay; Duration 5 min, Loops: 8,615. This step collects images at the maximum rate for the 5 min imaging period.</li>
	</ol>
	</li>
	<li>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., &#62;25 frames/s) to better decipher cargo movements.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Fix the cells by removing the medium and incubating with prewarmed 4% paraformaldehyde&#160;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.</li>
	<li>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 &#946;-III tubulin (Tuj1) to verify that the neurons remained healthy throughout the imaging process. 
	&#8203;NOTE: We observe co-expression of both DNA constructs in nearly all transfected cells using these methods.</li>
</ol>
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.
<ol>
	<li>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).</li>
	<li>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&#39;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&#39;s trace on the kymograph. After the polyline is laid over the entire track, click OK. Repeat until all tracks are assigned.</li>
	<li>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 &#181;m 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.</li>
	<li>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.</li>
</ol>

Visualizing Proteins in Primary Hippocampal Neurons by Live&#45;Cell Imaging

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The authors have no conflicts of interest to disclose.

There is growing evidence that multiple pathological proteins associated with a variety of neurodegenerative disorders disrupt fast axonal transport in neurons. This represents a potential common mechanism of neurotoxicity across these diseases. To better understand the process by which these proteins disrupt transport, we need tools and models that allow us to address specific questions. The method described here allows the examination of mechanisms engaged by pathological proteins to negatively affect cargo transport i...

Neurons depend on the bidirectional transport of cargo along the axon to maintain functional synapses and neural connectivity. Axonal transport deficits are thought to be critical contributors to the pathogenesis of several neurodegenerative diseases, including Alzheimer&#39;s disease (AD) and other tauopathies, Parkinson&#39;s disease, amyotrophic lateral sclerosis, and Huntington&#39;s disease1,2,3. Indeed, pathological modifications to several disease-related proteins negatively affect transport (reviewed in 4). Developing methods to investigate the mechanisms by which pathological proteins exert toxicity in disease is necessary to understand neurodegenerative disorders and identify potential targets for therapeutic intervention.
Several important insights into axon transport, including the discovery of conventional kinesin, the kinase- and phosphatase-dependent pathways regulating motor proteins, and mechanisms by which pathological proteins disrupt the regulation of axon transport, were made using the isolated squid axoplasm model4,5. Perfusion of squid axoplasm with pathological forms of tau protein inhibits anterograde fast axonal transport (FAT), an effect dependent on the exposure of the phosphatase-activating domain of tau, which activates protein phosphatase 1 (PP1)6,7,8. PP1 activates glycogen synthase kinase 3 (GSK3), which in turn phosphorylates kinesin light chains causing the release of cargo. Another pathological protein in AD is amyloid-&#946;. Oligomeric forms of amyloid-&#946; inhibit bidirectional FAT through casein kinase 2, which phosphorylates kinesin light chains9. Furthermore, pathological huntingtin protein harboring a polyglutamine expansion and a familial ALS-linked SOD1 mutant each disrupt axonal transport in the squid axoplasm through c-Jun N-terminal kinase and p38 mitogen-activated protein kinase activity, respectively10,11.
While the squid axoplasm model continues to be a valuable tool in understanding the effects of pathological proteins on axonal transport, limited access to the equipment and specimens prevents it from being more widely used. We developed a transport assay using live-cell confocal microscopy imaging of rodent (mouse and rat) primary neurons. This model represents an easily adaptable and manipulatable mammalian neuron-based approach using widely available cell sources and microscopy systems. For example, a variety of pathological proteins (e.g., harboring disease-related modifications) are expressed to identify how specific modifications of these proteins affect transport in axons. Similarly, a variety of fluorescently-tagged cargo proteins can be used to examine cargo-specific changes. Moreover, the underlying molecular mechanisms are studied relatively easily by targeting expression (i.e., knockdown or overexpression) of selected proteins that may mediate these effects. This method also is easily adapted for primary neurons derived from a wide variety of animal models.
We present a detailed protocol describing the live-cell axon transport assay that was previously used in primary hippocampal neurons to show that mutant tau proteins associated with frontotemporal lobar dementias (FTLD; P301L or R5L tau) increase the pausing frequency of fluorescently-labeled cargo proteins&#160;bidirectionally12. Furthermore, the knockdown of the PP1&#947; isoform rescued the pausing effects12. This provides support for the model of pathological tau-induced disruption that is mediated through aberrant activation of a signaling pathway initiated by PP1 as described above6,7,12. In a separate study, we showed that pseudophosphorylation of tau at S199/S202/T205 (the pathogenic AT8 phosphoepitope relevant to tauopathies) increased cargo pause frequency and anterograde segment velocity. These effects were dependent on the N-terminal phosphatase activating domain of tau13. These examples highlight the utility of this model for identifying the mechanisms of how pathological proteins disrupt axon transport in mammalian neurons.
This paper provides a detailed description of the method beginning with the harvest, dissociation, and culturing of primary mouse hippocampal neurons, followed by the transfection of the neurons with cargo proteins fused with a fluorescent protein, and finally, the live-cell imaging and image analysis approach. We demonstrate how this method is used to study the effects of modified tau on the bidirectional transport of the vesicle-associated protein, synaptophysin, as an example. However, there is flexibility in the pathogenic protein and transport cargo protein of interest, which makes this a versatile approach to study axonal transport.

<table><tbody><tr><td>0.4% Trypan blue</td><td>Gibco</td><td>15250-061</td><td></td></tr><tr><td>1.7 mL microcentrifuge tubes</td><td>DOT</td><td>RN1700-GMT</td><td></td></tr><tr><td>2.5% trypsin</td><td>Gibco</td><td>15090-046</td><td></td></tr><tr><td>3 mL syringe with 21 G needle</td><td>Fisher</td><td>14-826-84</td><td></td></tr><tr><td>10 mL plastic syringe</td><td>Fisher</td><td>14-823-2A</td><td></td></tr><tr><td>14 G needle</td><td>Fisher</td><td>14-817-203</td><td></td></tr><tr><td>15 G needle</td><td>Medline</td><td>SWD200029Z</td><td></td></tr><tr><td>16 G needle</td><td>Fisher</td><td>14-817-104</td><td></td></tr><tr><td>18 G needle</td><td>Fisher</td><td>14-840-97</td><td></td></tr><tr><td>22 G needle</td><td>Fisher</td><td>14-840-90</td><td></td></tr><tr><td>32% paraformaldehyde</td><td>Fisher</td><td>50-980-495</td><td></td></tr><tr><td>AlexaFluor 647 goat anti-rabbit IgG (H+L)</td><td>Invitrogen</td><td>A21244</td><td>RRID:AB_2535813</td></tr><tr><td>Amphotericin B</td><td>Gibco</td><td>15290-026</td><td></td></tr><tr><td>Arruga Micro Embryonic Capsule Forceps, Curved; 4&quot;&amp;nbsp;</td><td>Roboz</td><td>RS-5163</td><td>autoclave</td></tr><tr><td>B-27 Supplement (50x), serum free</td><td>Gibco</td><td>A3582801</td><td></td></tr><tr><td>BioCoat 24-well Poly D lysine plates&amp;nbsp;</td><td>Fisher</td><td>08-774-124</td><td></td></tr><tr><td>boric acid</td><td>Sigma</td><td>B6768-1KG</td><td></td></tr><tr><td>Calcium chloride</td><td>Sigma</td><td>C7902</td><td></td></tr><tr><td>Castroviejo 3 1/2&quot; Long 8 x 0.15 mm Angle Sharp Scissors</td><td>Roboz</td><td>RS-5658</td><td>autoclave</td></tr><tr><td>Cell counting device</td><td></td><td></td><td>automatic or manual</td></tr><tr><td>Confocal microscope with live cell chamber attachment</td><td></td><td></td><td></td></tr><tr><td>Confocal imaging software</td><td></td><td></td><td></td></tr><tr><td>D-(+)-glucose</td><td>Sigma</td><td>G7528</td><td></td></tr><tr><td>DNase I (Worthington)</td><td>Fisher</td><td>NC9185812</td><td></td></tr><tr><td>Dulbecco&#039;s Phosphate Buffered Saline</td><td>Gibco</td><td>14200-075</td><td></td></tr><tr><td>EGTA</td><td>Fisher</td><td>O2783-100</td><td></td></tr><tr><td>Fatal-Plus Solution</td><td>Vortech Pharmaceuticals, LTD</td><td>NDC 0298-9373-68</td><td>sodium pentobarbital; other approved methods of euthanasia may be used&amp;nbsp;</td></tr><tr><td>Fetal bovine serum</td><td>Invitrogen</td><td>16000044</td><td></td></tr><tr><td>Gentamicin Reagent Solution</td><td>Gibco</td><td>15710-072</td><td></td></tr><tr><td>GlutaMAX</td><td>Gibco</td><td>35050-061</td><td>glutamine substitute</td></tr><tr><td>Hanks&#039; Balanced Salt Solution</td><td>Gibco</td><td>24020-117</td><td></td></tr><tr><td>ImageJ version 1.51n</td><td>ImageJ</td><td></td><td>Life-Line version 2017 May 30: https://imagej.net/software/fiji/downloads</td></tr><tr><td>KymoAnalyzer (version 1.01)</td><td>Encalada Lab</td><td></td><td>Package includes all 6 macros: https://www.encalada.scripps.edu/kymoanalyzer</td></tr><tr><td>Lipofectamine 3000</td><td>Invitrogen</td><td>100022050</td><td>Use with P3000 transfection enhancer reagent</td></tr><tr><td>Magnesium chloride</td><td>Fisher</td><td>AC223211000</td><td></td></tr><tr><td>MES hydrate</td><td>Sigma</td><td>M8250</td><td></td></tr><tr><td>Micro Dissecting Scissors 3.5&quot; Straight Sharp/Sharp</td><td>Roboz</td><td>RS-5910</td><td>autoclave</td></tr><tr><td>Neurobasal Plus medium</td><td>Gibco</td><td>A3582901</td><td></td></tr><tr><td>Neurofascin (A12/18) Mouse IgG2a</td><td>UC Davis/NIH NeuroMab</td><td>75-172</td><td>RRID:AB_2282826; 250 ng/mL; Works in rat neurons, NOT in mouse neurons</td></tr><tr><td>Neurofascin 186 (D6G60) Rabbit IgG</td><td>Cell Signaling</td><td>15034</td><td>RRID:AB_2773024; 500 ng/mL; Works in mouse neurons, we have not tested in rat neurons</td></tr><tr><td>newborn calf serum</td><td>Gibco</td><td>16010-167</td><td></td></tr><tr><td>Opti-MEM</td><td>Gibco</td><td>31985-062</td><td></td></tr><tr><td>P3000</td><td>Invitrogen</td><td>100022057</td><td></td></tr><tr><td>Petri dish, 100 x 10 mm glass</td><td>Fisher</td><td>08-748B</td><td>For dissection; autoclave</td></tr><tr><td>Petri dish, 100 x 20 mm glass</td><td>Fisher</td><td>08-748D</td><td>To place uterine horns in; autoclave</td></tr><tr><td>Poly-D-lysine</td><td>Sigma</td><td>P7886-100MG</td><td></td></tr><tr><td>Polypropylene conical centrifuge tubes (15 mL)</td><td>Fisher</td><td>14-955-238</td><td></td></tr><tr><td>Polypropylene conical centrifuge tubes (50 mL)</td><td>Fisher</td><td>14-955-238</td><td></td></tr><tr><td>Potassium chloride</td><td>Fisher</td><td>P217-500</td><td></td></tr><tr><td>Sodium acetate</td><td>Sigma</td><td>S5636</td><td></td></tr><tr><td>sodium borate decahydrate</td><td>VWR</td><td>MK745706</td><td></td></tr><tr><td>Straight-Blade Operating Scissors Blunt/Sharp</td><td>Fisher</td><td>13-810-2</td><td>autoclave</td></tr><tr><td>Syringe Filters, 0.22 &amp;micro;m</td><td>VWR</td><td>514-1263</td><td></td></tr><tr><td>Thumb dressing forceps, serrated, 4.5&quot;</td><td>Roboz</td><td>RS-8100</td><td>autoclave</td></tr><tr><td>&amp;micro;-Slide 4 Well Glass Bottom</td><td>Ibidi</td><td>80427</td><td></td></tr></tbody></table>

using live&#45;cell imaging to measure the effects of pathological proteins on axonal transport in primary hippocampal neurons

Bidirectional transport of cargos along the axon is critical for maintaining functional synapses, neural connectivity, and healthy neurons. Axonal transport is disrupted in multiple neurodegenerative diseases, and projection neurons are particularly vulnerable because of the need to transport cellular materials over long distances and sustain substantial axonal mass. Pathological modifications of several disease-related proteins negatively affect transport, including tau, amyloid-&#946;, &#945;-synuclein, superoxide dismutase, and huntingtin, providing a potential common mechanism by which pathological proteins exert toxicity in disease. Methods to study these toxic mechanisms are necessary to understand neurodegenerative disorders and identify potential therapeutic interventions.
Here, cultured primary rodent hippocampal neurons are co-transfected with multiple plasmids to study the effects of pathological proteins on fast axonal transport using live-cell confocal imaging of fluorescently-tagged cargo proteins. We begin with the harvest, dissociation, and culturing of primary hippocampal neurons from rodents. Then, we co-transfect the neurons with plasmid DNA constructs to express fluorescent-tagged cargo protein and wild-type or mutant tau (used as an exemplar of pathological proteins). Axons are identified in live cells using an antibody that binds an extracellular&#160;domain of neurofascin, an axon initial segment protein, and an axonal region of interest is imaged to measure fluorescent cargo transport.
Using KymoAnalyzer, a freely available ImageJ macro, we extensively characterize the velocity, pause frequency, and directional cargo density of axonal transport, all of which may be affected by the presence of pathological proteins. Through this method, we identify a phenotype of increased cargo pause frequency associated with the expression of pathological tau protein. Additionally, gene-silencing shRNA constructs can be added to the transfection mix to test the role of other proteins in mediating transport disruption. This protocol is easily adaptable for use with other neurodegenerative disease-related proteins and is a reproducible method to study the mechanisms of how those proteins disrupt axonal transport.

Using these methods, we characterized axonal transport in the presence of wild-type or disease-related forms of tau protein to examine potential mechanisms of pathological tau-induced neurotoxicity in disease12,13. The KymoAnalyzer software calculates and pools a variety of different parameters from all kymographs within a given folder. Transport rates are calculated only when cargo is in motion (segmental velocity) as well as the overall rate, including pauses (...

Read this Scientific Article Protocol about Author Spotlight: Understanding the Impact of Pathological Proteins on Axonal Transport in Neurodegenerative Diseases at JoVE.com

Author Spotlight: Understanding the Impact of Pathological Proteins on Axonal Transport in Neurodegenerative Diseases

Here, we demonstrate how to combine transfection of primary hippocampal rodent neurons with live-cell confocal imaging to analyze pathological protein-induced effects on axonal transport and identify mechanistic pathways mediating these effects.

Using Live&#45;Cell Imaging to Measure the Effects of Pathological Proteins on Axonal Transport in Primary Hippocampal Neurons

Bidirectional transport of cargos along the axon is critical for maintaining functional synapses, neural connectivity, and healthy neurons. Axonal ...

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Research

JoVE Journal

Neuroscience

413 Views.  Michigan State University. Neuronen vertrouwen op bidirectioneel transport van vracht langs het axon om functionele synapsis en neurale connectiviteit te behouden. Tekorten in het transport worden verondersteld een cruciale bijdrage te leveren aan de pathogenese van verschillende neurodegeneratieve ziekten. We willen mechanismen identificeren waarmee ziekte-geassocieerde modificaties van tau-eiwit het axonale transport verstoren.We hebben vastgesteld dat meerdere pathologische v...

Video: Auteur in de schijnwerpers: inzicht in de impact van pathologische eiwitten op axonaal transport bij neurodegeneratieve ziekten

Live Imaging of Dense-core Vesicles in Primary Cultured Hippocampal Neurons

Observing and characterizing dynamic cellular processes can yield important information about cellular activity that cannot be gained from static images.  Vital fluorescent probes, particularly green fluorescent protein (GFP) have revolutionized cell biology stemming from the ability to label specific intracellular compartments and cellular structures. For example, the live imaging of GFP (and its spectral variants) chimeras have allowed for a dynamic 
analysis of the cytoskeleton, organelle transport, and membrane dynamics in a multitude of organisms and cell types [1-3].  Although live imaging has become prevalent, this approach still poses many technical challenges, particularly in primary cultured neurons.  One challenge is the expression of GFP-tagged proteins in post-mitotic neurons; the other is the ability to capture fluorescent images while minimizing phototoxicity, photobleaching, 
and maintaining general cell health.  Here we provide a protocol that describes a lipid-based transfection method that yields a relatively low transfection rate (~0.5%), however is ideal for the imaging of fully polarized neurons.  A low transfection rate is essential so that single axons and dendrites can be characterized as to their orientation to the cell body to confirm directionality of transport, i.e., anterograde v. retrograde.  Our approach to imaging 
GFP expressing neurons relies on a standard wide-field fluorescent microscope outfitted with a CCD camera, image capture software, and a heated imaging chamber. We have imaged a wide variety of organelles or structures, for example, dense-core vesicles, mitochondria, growth cones, and actin without any special optics or excitation requirements other than a fluorescent light source. Additionally, spectrally-distinct, fluorescently 
labeled proteins, e.g., GFP and dsRed-tagged proteins, can be visualized near simultaneously to characterize co-transport or other coordinated cellular events.  The imaging approach described here is flexible for a variety of imaging applications and can be adopted by a laboratory for relatively little cost provided a microscope is available.

Live cell imaging is of particular utility when studying the dynamics of organelle trafficking. Here we describe a protocol for live imaging of dense-core vesicles in cultured neurons using wide-field fluorescence microscopy. This protocol is flexible and can be adapted to image other organelles such as mitochondria, endosomes, and peroxisomes.

Observing and characterizing dynamic cellular processes can yield important information about cellular activity that cannot be gained from static images.  ...

Biology

Isolation of CA1 Nuclear Enriched Fractions from Hippocampal Slices to Study Activity-dependent Nuclear Import of Synapto-nuclear Messenger Proteins

Studying activity dependent protein expression, subcellular translocation, or phosphorylation is essential to understand the underlying cellular mechanisms of synaptic plasticity. Long-term potentiation (LTP) and long-term depression (LTD) induced in acute hippocampal slices are widely accepted as cellular models of learning and memory. There are numerous studies that use live cell imaging or immunohistochemistry approaches to visualize activity dependent protein dynamics. However these methods rely on the suitability of antibodies for immunocytochemistry or overexpression of fluorescence-tagged proteins in single neurons. Immunoblotting of proteins is an alternative method providing independent confirmation of the findings.&#160;The first limiting factor in preparation of subcellular fractions from individual tetanized hippocampal slices is the low amount of material. Second, the handling procedure is crucial because even very short and minor manipulations of living slices might induce activation of certain signaling cascades. Here we describe an optimized workflow in order to obtain sufficient quantity of nuclear enriched fraction of sufficient purity from the CA1 region of acute hippocampal slices from rat brain. As a representative example we show that the ERK1/2 phosphorylated form of the synapto-nuclear protein messenger Jacob actively translocates to the nucleus upon induction of LTP and can be detected in a nuclear enriched fraction from CA1 neurons.

We provide a detailed protocol for induction of long-term potentiation in the CA1 region of the hippocampus and the subsequent isolation of nuclear enriched fractions from the tetanized area of the slice. This approach can be used to determine activity dependent nuclear protein import in cellular models of learning and memory.

Studying activity dependent protein expression, subcellular translocation, or phosphorylation is essential to understand the underlying cellular ...

Real-time Imaging of Axonal Transport of Quantum Dot-labeled BDNF in Primary Neurons

BDNF plays an important role in several facets of neuronal survival, differentiation, and function. Structural and functional deficits in axons are increasingly viewed as an early feature of neurodegenerative diseases, including Alzheimer&#8217;s disease (AD) and Huntington&#8217;s disease (HD). As yet unclear is the mechanism(s) by which axonal injury is induced. We reported the development of a novel technique to produce biologically active, monobiotinylated BDNF (mBtBDNF) that can be used to trace axonal transport of BDNF. Quantum dot-labeled BDNF (QD-BDNF) was produced by conjugating quantum dot 655 to mBtBDNF. A microfluidic device was used to isolate axons from neuron cell bodies. Addition of QD-BDNF to the axonal compartment allowed live imaging of BDNF transport in axons. We demonstrated that QD-BDNF moved essentially exclusively retrogradely, with very few pauses, at a moving velocity of around 1.06 &#956;m/sec. This system can be used to investigate mechanisms of disrupted axonal function in AD or HD, as well as other degenerative disorders.

Axonal transport of BDNF, a neurotrophic factor, is critical for the survival and function of several neuronal populations. Some degenerative disorders are marked by disruption of axonal structure and function. We demonstrated the techniques used to examine live trafficking of QD-BDNF in microfluidic chambers using primary neurons.

BDNF plays an important role in several facets of neuronal survival, differentiation, and function. Structural and functional deficits in axons are ...

Fluorescence and Bioluminescence Imaging of Subcellular Ca2+ in Aged Hippocampal Neurons

Susceptibility to neuron cell death associated to neurodegeneration and ischemia are exceedingly increased in the aged brain but mechanisms responsible are badly known. Excitotoxicity, a process believed to contribute to neuron damage induced by both insults, is mediated by activation of glutamate receptors that promotes Ca2+ influx and mitochondrial Ca2+ overload. A substantial change in intracellular Ca2+ homeostasis or remodeling of intracellular Ca2+ homeostasis may favor neuron damage in old neurons. For investigating Ca2+ remodeling in aging we have used live cell imaging in long-term cultures of rat hippocampal neurons that resemble in some aspects aged neurons in vivo. For this end, hippocampal cells are, in first place, freshly dispersed from new born rat hippocampi and plated on poli-D-lysine coated, glass coverslips. Then cultures are kept in controlled media for several days or several weeks for investigating young and old neurons, respectively. Second, cultured neurons are loaded with fura2 and subjected to measurements of cytosolic Ca2+ concentration using digital fluorescence ratio imaging. Third, cultured neurons are transfected with plasmids expressing a tandem of low-affinity aequorin and GFP targeted to mitochondria. After 24 hr, aequorin inside cells is reconstituted with coelenterazine and neurons are subjected to bioluminescence imaging for monitoring of mitochondrial Ca2+ concentration. This three-step procedure allows the monitoring of cytosolic and mitochondrial Ca2+ responses to relevant stimuli as for example the glutamate receptor agonist NMDA and compare whether these and other responses are influenced by aging. This procedure may yield new insights as to how aging influence cytosolic and mitochondrial Ca2+ responses to selected stimuli as well as the testing of selected drugs aimed at preventing neuron cell death in age-related diseases.

Fluorescence and Bioluminescence Imaging of Subcellular Ca2+ in Aged Hippocampal Neurons

Intracellular Ca2+ remodeling in aging may contribute to excitotoxicity and neuron damage, processes mediated by Ca2+ overload. We aimed at investigating Ca2+ remodeling in the aging brain using fluorescence and bioluminescence imaging of cytosolic and mitochondrial Ca2+ in long-term cultures of rat hippocampal neurons, a model of neuronal aging.

Susceptibility to neuron cell death associated to neurodegeneration and ischemia are exceedingly increased in the aged brain but mechanisms responsible ...

Live Images of GLUT4 Protein Trafficking in Mouse Primary Hypothalamic Neurons Using Deconvolution Microscopy

Type 2 diabetes mellitus (T2DM) is a global health crisis which is characterized by insulin signaling impairment and chronic inflammation in peripheral tissues. The hypothalamus in the central nervous system (CNS) is the control center for energy and insulin signal response regulation. Chronic inflammation in peripheral tissues and imbalances of certain chemokines (such as CCL5, TNF&#945;, and IL-6) contribute to diabetes and obesity. However, the functional mechanism(s) connecting chemokines and hypothalamic insulin signal regulation still remain unclear.
In vitro primary neuron culture models are convenient and simple models which can be used to investigate insulin signal regulation in hypothalamic neurons. In this study, we introduced exogeneous GLUT4 protein conjugated with GFP (GFP-GLUT4) into primary hypothalamic neurons to track GLUT4 membrane translocation upon insulin stimulation. Time-lapse images of GFP-GLUT4 protein trafficking were recorded by deconvolution microscopy, which allowed users to generate high-speed, high-resolution images without damaging the neurons significantly while conducting the experiment. The contribution of CCR5 in insulin regulated GLUT4 translocation was observed in CCR5 deficient hypothalamic neurons, which were isolated and cultured from CCR5 knockout mice. Our results demonstrated that the GLUT4 membrane translocation efficiency was reduced in CCR5 deficient hypothalamic neurons after insulin stimulation.

This protocol describes a technique for observation of real-time Green Fluorescence Protein (GFP) tagged Glucose Transporter 4 (GLUT4) protein trafficking upon insulin stimulation and characterization of the biological role of CCR5 in the insulin&#8211;GLUT4 signaling pathway with Deconvolution Microscopy.

Type 2 diabetes mellitus (T2DM) is a global health crisis which is characterized by insulin signaling impairment and chronic inflammation in peripheral ...

Visualizing Axonal Growth Cone Collapse and Early Amyloid &#946; Effects in Cultured Mouse Neurons

Amyloid-&#946; (A&#946;) causes memory impairments in Alzheimer's disease (AD). Although therapeutics have been shown to reduce A&#946; levels in the brains of AD patients, these do not improve memory functions. Since A&#946; aggregates in the brain before the appearance of memory impairments, targeting A&#946; may be inefficient for treating AD patients who already exhibit memory deficits. Therefore, downstream signaling due to A&#946; deposition should be blocked before AD development. A&#946; induces axonal degeneration, leading to the disruption of neuronal networks and memory impairments. Although there are many studies on the mechanisms of A&#946; toxicity, the source of A&#946; toxicity remains unknown. To help identify the source, we propose a novel protocol that uses microscopy, gene transfection, and live cell imaging to investigate early changes caused by A&#946; in axonal growth cones of cultured neurons. This protocol revealed that A&#946; induced clathrin-mediated endocytosis in axonal growth cones followed by growth cone collapse, demonstrating that inhibition of endocytosis prevents A&#946; toxicity. This protocol will be useful in studying the early effects of A&#946; and may lead to more efficient and preventative AD treatment.

Here a protocol to investigate the early effects of amyloid-&#946; (A&#946;) in the brain is presented. This shows that A&#946; induces clathrin-mediated endocytosis and collapse of axonal growth cones. The protocol is useful in studying early effects of A&#946; on axonal growth cones and may facilitate prevention of Alzheimer's disease.

Amyloid-&#946; (A&#946;) causes memory impairments in Alzheimer's disease (AD). Although therapeutics have been shown to reduce A&#946; levels in the ...

A Scalable Method to Study Neuronal Survival in Primary Neuronal Culture with Single-cell and Real-Time Resolution

Neuronal loss is at the core of many neuropathologies, including stroke, Alzheimer's disease, and Parkinson's disease. Different methods were developed to study the process of neuronal survival upon cytotoxic stress. Most methods are based on biochemical approaches that do not allow single-cell resolution or involve complex and costly methodologies. Presented here is a versatile, inexpensive, and effective experimental paradigm to study neuronal survival. This method takes advantage of sparse fluorescent labeling of the neurons followed by live imaging and automated quantification. To this aim, the neurons are electroporated to express fluorescent markers and co-cultured with non-electroporated neurons to easily regulate cell density and increase survival. 
Sparse labeling by electroporation allows a simple and robust automated quantification. In addition, fluorescent labeling can be combined with the co-expression of a gene of interest to study specific molecular pathways. Here, we present a model of stroke as a neurotoxic model, namely, the oxygen-glucose deprivation (OGD) assay, which was performed in an affordable and robust homemade hypoxic chamber. Finally, two different workflows are described using IN Cell Analyzer 2200 or the open-source ImageJ for image analysis for semi-automatic data processing. This workflow can be easily adapted to different experimental models of toxicity and scaled up for high-throughput screening. In conclusion, the described protocol provides an approachable, affordable, and effective in vitro model of neurotoxicity, which can be suitable for testing the roles of specific genes and pathways in live imaging and for high-throughput drug screening.

Here, we present a simple, potent, and versatile methodology to investigate neuronal survival upon cytotoxic stress in primary cortical neurons with cellular resolution in real time or in fixed material.

Neuronal loss is at the core of many neuropathologies, including stroke, Alzheimer's disease, and Parkinson's disease. Different methods were developed to ...

Real-Time Fluorescent Measurement of Synaptic Functions in Models of Amyotrophic Lateral Sclerosis

Before neuronal degeneration, the cause of motor and cognitive deficits in patients with amyotrophic lateral sclerosis (ALS) and/or frontotemporal lobe dementia (FTLD) is dysfunction of communication between neurons and motor neurons and muscle. The underlying process of synaptic transmission involves membrane depolarization-dependent synaptic vesicle fusion and the release of neurotransmitters into the synapse. This process occurs through localized calcium influx into the presynaptic terminals where synaptic vesicles reside. Here, the protocol describes fluorescence-based live-imaging methodologies that reliably report depolarization-mediated synaptic vesicle exocytosis and presynaptic terminal calcium influx dynamics in cultured neurons.
Using a styryl dye that is incorporated into synaptic vesicle membranes, the synaptic vesicle release is elucidated. On the other hand, to study calcium entry, Gcamp6m is used, a genetically encoded fluorescent reporter. We employ high potassium chloride-mediated depolarization to mimic neuronal activity. To quantify synaptic vesicle exocytosis unambiguously, we measure the loss of normalized styryl dye fluorescence as a function of time. Under similar stimulation conditions, in the case of calcium influx, Gcamp6m fluorescence increases. Normalization and quantification of this fluorescence change are performed in a similar manner to the styryl dye protocol. These methods can be multiplexed with transfection-based overexpression of fluorescently tagged mutant proteins. These protocols have been extensively used to study synaptic dysfunction in models of FUS-ALS and C9ORF72-ALS, utilizing primary rodent cortical and motor neurons. These protocols easily allow for rapid screening of compounds that may improve neuronal communication. As such, these methods are valuable not only for the study of ALS but for all areas of neurodegenerative and developmental neuroscience research.

Two related methods are described to visualize subcellular events required for synaptic transmission. These protocols enable the real-time monitoring of the dynamics of presynaptic calcium influx and synaptic vesicle membrane fusion using live-cell imaging of in vitro cultured neurons.

Before neuronal degeneration, the cause of motor and cognitive deficits in patients with amyotrophic lateral sclerosis (ALS) and/or frontotemporal lobe ...

Expanding the Toolkit for In Vivo Imaging of Axonal Transport

Axonal transport maintains neuronal homeostasis by enabling the bidirectional trafficking of diverse organelles and cargoes. Disruptions in axonal transport have devastating consequences for individual neurons and their networks, and contribute to a plethora of neurological disorders. As many of these conditions involve both cell autonomous and non-autonomous mechanisms, and often display a spectrum of pathology across neuronal subtypes, methods to accurately identify and analyze neuronal subsets are imperative.
This paper details protocols to assess in vivo axonal transport of signaling endosomes and mitochondria in sciatic nerves of anesthetized mice. Stepwise instructions are provided to 1) distinguish motor from sensory neurons in vivo, in situ, and ex vivo by using mice that selectively express fluorescent proteins within cholinergic motor neurons; and 2) separately or concurrently assess in vivo axonal transport of signaling endosomes and mitochondria. These complementary intravital approaches facilitate the simultaneous imaging of different cargoes in distinct peripheral nerve axons to quantitatively monitor axonal transport in health and disease.

Expanding the Toolkit for In Vivo Imaging of Axonal Transport

Using transgenic fluorescent mice, detailed protocols are described to assess in vivo axonal transport of signaling endosomes and mitochondria within motor and sensory axons of the intact sciatic nerve in live animals.

Axonal transport maintains neuronal homeostasis by enabling the bidirectional trafficking of diverse organelles and cargoes. Disruptions in axonal ...

Primary Cultures of Rat Astrocytes and Microglia and Their Use in the Study of Amyotrophic Lateral Sclerosis

This protocol demonstrates how to prepare primary cultures of glial cells, astrocytes, and microglia from the cortices of Sprague Dawley rats and how to use these cells for the purpose of studying the pathophysiology of amyotrophic lateral sclerosis (ALS) in the rat hSOD1G93A model. First, the protocol shows how to isolate and culture astrocytes and microglia from postnatal rat cortices, and then how to characterize and test these cultures for purity by immunocytochemistry using the glial fibrillary acidic protein (GFAP) marker of astrocytes and the ionized calcium-binding adaptor molecule 1 (Iba1) microglial marker. In the next stage, methods are described for dye-loading (calcium-sensitive Fluo 4-AM) of cultured cells and the recordings of Ca2+ changes in video imaging experiments on live cells.
The examples of video recordings consist of: (1) cases of Ca2+ imaging of cultured astrocytes acutely exposed to immunoglobulin G (IgG) isolated from ALS patients, showing a characteristic and specific response compared to the response to ATP as demonstrated in the same experiment. Examples also show a more pronounced transient rise in intracellular calcium concentration evoked by ALS IgG in hSOD1G93A astrocytes compared to non-transgenic controls; (2) Ca2+ imaging of cultured astrocytes during a depletion of calcium stores by thapsigargin (Thg), a non-competitive inhibitor of the endoplasmic reticulum Ca2+ ATPase, followed by store-operated calcium entry elicited by the addition of calcium in the recording solution, which demonstrates the difference between Ca2+ store operation in hSOD1G93A and in non-transgenic astrocytes; (3) Ca2+ imaging of the cultured microglia showing predominantly a lack of response to ALS IgG, whereas ATP application elicited a Ca2+ change. This paper also emphasizes possible caveats and cautions regarding critical cell density and purity of cultures, choosing the correct concentration of the Ca2+ dye and dye-loading techniques.

We present here a protocol on how to prepare primary cultures of glial cells, astrocytes, and microglia from rat cortices for time-lapse video imaging of intracellular Ca2+ for research on pathophysiology of amyotrophic lateral sclerosis in the hSOD1G93A rat model.

This protocol demonstrates how to prepare primary cultures of glial cells, astrocytes, and microglia from the cortices of Sprague Dawley rats and how to ...

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

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