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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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&#34;&#160;</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&#160;</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&#34; Long 8 x 0.15 mm Angle Sharp Scissors</td>
			<td>Roboz</td>
			<td>RS-5658</td>
			<td>autoclave</td>
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		<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&#39;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&#160;</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&#39; 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&#34; 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 &#181;m</td>
			<td>VWR</td>
			<td>514-1263</td>
			<td></td>
		</tr>
		<tr>
			<td>Thumb dressing forceps, serrated, 4.5&#34;</td>
			<td>Roboz</td>
			<td>RS-8100</td>
			<td>autoclave</td>
		</tr>
		<tr>
			<td>&#181;-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 ...

using-live-cell-imaging-to-measure-effects-pathological-proteins-on

Research

JoVE Journal

Neuroscience

360 Views.  Michigan State University. 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.

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

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

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

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

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

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

Neuromodulation and Mitochondrial Transport: Live Imaging in Hippocampal Neurons over Long Durations

To understand the relationship between mitochondrial transport and neuronal function, it is critical to observe mitochondrial behavior in live cultured neurons for extended durations1-3. This is now possible through the use of vital dyes and fluorescent proteins with which cytoskeletal components, organelles, and other structures in living cells can be labeled and then visualized via dynamic fluorescence microscopy. For example, in embryonic chicken sympathetic neurons, mitochondrial movement was characterized using the vital dye rhodamine 1234. In another study, mitochondria were visualized in rat forebrain neurons by transfection of mitochondrially targeted eYFP5. However, imaging of primary neurons over minutes, hours, or even days presents a number of issues. Foremost among these are: 1) maintenance of culture conditions such as temperature, humidity, and pH during long imaging sessions; 2) a strong, stable fluorescent signal to assure both the quality of acquired images and accurate measurement of signal intensity during image analysis; and 3) limiting exposure times during image acquisition to minimize photobleaching and avoid phototoxicity.
Here, we describe a protocol that permits the observation, visualization, and analysis of mitochondrial movement in cultured hippocampal neurons with high temporal resolution and under optimal life support conditions. We have constructed an affordable stage-top incubator that provides good temperature regulation and atmospheric gas flow, and also limits the degree of media evaporation, assuring stable pH and osmolarity. This incubator is connected, via inlet and outlet hoses, to a standard tissue culture incubator, which provides constant humidity levels and an atmosphere of 5-10% CO2/air. This design offers a cost-effective alternative to significantly more expensive microscope incubators that don't necessarily assure the viability of cells over many hours or even days. To visualize mitochondria, we infect cells with a lentivirus encoding a red fluorescent protein that is targeted to the mitochondrion. This assures a strong and persistent signal, which, in conjunction with the use of a stable xenon light source, allows us to limit exposure times during image acquisition and all but precludes photobleaching and phototoxicity. Two injection ports on the top of the stage-top incubator allow the acute administration of neurotransmitters and other reagents intended to modulate mitochondrial movement. In sum, lentivirus-mediated expression of an organelle-targeted red fluorescent protein and the combination of our stage-top incubator, a conventional inverted fluorescence microscope, CCD camera, and xenon light source allow us to acquire time-lapse images of mitochondrial transport in living neurons over longer durations than those possible in studies deploying conventional vital dyes and off-the-shelf life support systems.

We describe a protocol that allows imaging of mitochondria in living neurons via fluorescence microscopy over long durations. Imaging over extended periods is accomplished through lentivirus-mediated expression of a mitochondrially targeted fluorescent protein and use of an inexpensive stage-top incubator that was designed and built in our laboratory.

To understand the relationship between mitochondrial transport and neuronal function, it is critical to observe mitochondrial behavior in live cultured ...

Imaging pHluorin-tagged Receptor Insertion to the Plasma Membrane in Primary Cultured Mouse Neurons

A better understanding of the mechanisms governing receptor trafficking between the plasma membrane (PM) and intracellular compartments requires an experimental approach with excellent spatial and temporal resolutions. Moreover, such an approach must also have the ability to distinguish receptors localized on the PM from those in intracellular compartments. Most importantly, detecting receptors in a single vesicle requires outstanding detection sensitivity, since each vesicle carries only a small number of receptors. Standard approaches for examining receptor trafficking include surface biotinylation followed by biochemical detection, which lacks both the necessary spatial and temporal resolutions; and fluorescence microscopy examination of immunolabeled surface receptors, which requires chemical fixation of cells and therefore lacks sufficient temporal resolution1-6 . To overcome these limitations, we and others have developed and employed a new strategy that enables visualization of the dynamic insertion of receptors into the PM with excellent spatial and temporal resolutions 7-17 . The approach includes tagging of a pH-sensitive GFP, the superecliptic pHluorin 18, to the N-terminal extracellular domain of the receptors. Superecliptic pHluorin has the unique property of being fluorescent at neutral pH and non-fluorescent at acidic pH (pH &lt; 6.0). Therefore, the tagged receptors are non-fluorescent when within the acidic lumen of intracellular trafficking vesicles or endosomal compartments, and they become readily visualized only when exposed to the extracellular neutral pH environment, on the outer surface of the PM. Our strategy consequently allows us to distinguish PM surface receptors from those within intracellular trafficking vesicles. To attain sufficient spatial and temporal resolutions, as well as the sensitivity required to study dynamic trafficking of receptors, we employed total internal reflection fluorescent microscopy (TIRFM), which enabled us to achieve the optimal spatial resolution of optical imaging (~170 nm), the temporal resolution of video-rate microscopy (30 frames/sec), and the sensitivity to detect fluorescence of a single GFP molecule. By imaging pHluorin-tagged receptors under TIRFM, we were able to directly visualize individual receptor insertion events into the PM in cultured neurons. This imaging approach can potentially be applied to any membrane protein with an extracellular domain that could be labeled with superecliptic pHluorin, and will allow dissection of the key detailed mechanisms governing insertion of different membrane proteins (receptors, ion channels, transporters, etc.) to the PM.

By tagging the extracellular domains of membrane receptors with superecliptic pHluorin, and by imaging these fusion receptors in cultured mouse neurons, we can directly visualize individual vesicular insertion events of the receptors to the plasma membrane. This technique will be instrumental in elucidating the molecular mechanisms governing receptor insertion to the plasma membrane.

A better understanding of the mechanisms governing receptor trafficking between the plasma membrane (PM) and intracellular compartments requires an ...

Assaying DNA Damage in Hippocampal Neurons Using the Comet Assay

A number of drugs target the DNA repair pathways and induce cell kill by creating DNA damage. Thus, processes to directly measure DNA damage have been extensively evaluated. Traditional methods are time consuming, expensive, resource intensive and require replicating cells. In contrast, the comet assay, a single cell gel electrophoresis assay, is a faster, non-invasive, inexpensive, direct and sensitive measure of DNA damage and repair. All forms of DNA damage as well as DNA repair can be visualized at the single cell level using this powerful technique.
	The principle underlying the comet assay is that intact DNA is highly ordered whereas DNA damage disrupts this organization. The damaged DNA seeps into the agarose matrix and when subjected to an electric field, the negatively charged DNA migrates towards the cathode which is positively charged. The large undamaged DNA strands are not able to migrate far from the nucleus. DNA damage creates smaller DNA fragments which travel farther than the intact DNA. Comet Assay, an image analysis software, measures and compares the overall fluorescent intensity of the DNA in the nucleus with DNA that has migrated out of the nucleus. Fluorescent signal from the migrated DNA is proportional to DNA damage. Longer brighter DNA tail signifies increased DNA damage. Some of the parameters that are measured are tail moment which is a measure of both the amount of DNA and distribution of DNA in the tail, tail length and percentage of DNA in the tail. This assay allows to measure DNA repair as well since resolution of DNA damage signifies repair has taken place. The limit of sensitivity is approximately 50 strand breaks per diploid mammalian cell 1,2. Cells treated with any DNA damaging agents, such as etoposide, may be used as a positive control. Thus the comet assay is a quick and effective procedure to measure DNA damage.

The comet assay is an efficient way of detecting single- and double-strand breaks, including alkali-labile sites and DNA-DNA/DNA-protein cross-links on the DNA in all cells including hippocampal neurons. The method takes advantage of the differential migration of DNA in an electric field due to differences in amount of DNA damage.

A number of drugs target the DNA repair pathways and induce cell kill by creating DNA damage. Thus, processes to directly measure DNA damage have been ...

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety of brain disorders1,2. To better understand the mechanisms by which brain circuits react to an animal's experience requires the ability to monitor the experience-dependent molecular changes in a given set of neurons, over a prolonged period of time, in the live animal. While experience and associated neural activity is known to trigger gene expression changes in neurons1,2, most of the methods to detect such changes do not allow repeated observation of the same neurons over multiple days or do not have sufficient resolution to observe individual neurons3,4. Here, we describe a method that combines in vivo two-photon microscopy with a genetically encoded fluorescent reporter to track experience-dependent gene expression changes in individual cortical neurons over the course of day-to-day experience. 
One of the well-established experience-dependent genes is Activity-regulated cytoskeletal associated protein (Arc)5,6. The transcription of Arc is rapidly and highly induced by intensified neuronal activity3, and its protein product regulates the endocytosis of glutamate receptors and long-term synaptic plasticity7. The expression of Arc has been widely used as a molecular marker to map neuronal circuits involved in specific behaviors3. In most of those studies, Arc expression was detected by in situ hybridization or immunohistochemistry in fixed brain sections. Although those methods revealed that the expression of Arc was localized to a subset of excitatory neurons after behavioral experience, how the cellular patterns of Arc expression might change with multiple episodes of repeated or distinctive experiences over days was not investigated. 
In vivo two-photon microscopy offers a powerful way to examine experience-dependent cellular changes in the living brain8,9. To enable the examination of Arc expression in live neurons by two-photon microscopy, we previously generated a knock-in mouse line in which a GFP reporter is placed under the control of the endogenous Arc promoter10. This protocol describes the surgical preparations and imaging procedures for tracking experience-dependent Arc-GFP expression patterns in neuronal ensembles in the live animal. In this method, chronic cranial windows were first implanted in Arc-GFP mice over the cortical regions of interest. Those animals were then repeatedly imaged by two-photon microscopy after desired behavioral paradigms over the course of several days. This method may be generally applicable to animals carrying other fluorescent reporters of experience-dependent molecular changes4.

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

Experience-dependent molecular changes in neurons are essential for the brain's ability to adapt in response to behavioral challenges. An in vivo two-photon imaging method is described here that allows the tracking of such molecular changes in individual cortical neurons through genetically encoded reporters.

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety ...

Calcium Phosphate Transfection of Primary Hippocampal Neurons

Calcium phosphate precipitation is a convenient and economical method for transfection of cultured cells. With optimization, it is possible to use this method on hard-to-transfect cells like primary neurons. Here we describe our detailed protocol for calcium phosphate transfection of hippocampal neurons cocultured with astroglial cells.

Calcium phosphate precipitation is a convenient and economical method for transfection of cultured cells. With optimization, it is possible to use this ...

Imaging Dendritic Spines of Rat Primary Hippocampal Neurons using Structured Illumination Microscopy

Dendritic spines are protrusions emerging from the dendrite of a neuron and represent the primary postsynaptic targets of excitatory inputs in the brain. Technological advances have identified these structures as key elements in neuron connectivity and synaptic plasticity. The quantitative analysis of spine morphology using light microscopy remains an essential problem due to technical limitations associated with light's intrinsic refraction limit. Dendritic spines can be readily identified by confocal laser-scanning fluorescence microscopy. However, measuring subtle changes in the shape and size of spines is difficult because spine dimensions other than length are usually smaller than conventional optical resolution fixed by light microscopy's theoretical resolution limit of 200 nm.
Several recently developed super resolution techniques have been used to image cellular structures smaller than the 200 nm, including dendritic spines. These techniques are based on classical far-field operations and therefore allow the use of existing sample preparation methods and to image beyond the surface of a specimen. Described here is a working protocol to apply super resolution structured illumination microscopy (SIM) to the imaging of dendritic spines in primary hippocampal neuron cultures. Possible applications of SIM overlap with those of confocal microscopy. However, the two techniques present different applicability. SIM offers higher effective lateral resolution, while confocal microscopy, due to the usage of a physical pinhole, achieves resolution improvement at the expense of removal of out of focus light. In this protocol, primary neurons are cultured on glass coverslips using a standard protocol, transfected with DNA plasmids encoding fluorescent proteins and imaged using SIM. The whole protocol described herein takes approximately 2 weeks, because dendritic spines are imaged after 16-17 days in vitro, when dendritic development is optimal. After completion of the protocol, dendritic spines can be reconstructed in 3D from series of SIM image stacks using specialized software.

This article describes a working protocol to image dendritic spines from hippocampal neurons in vitro using Structured Illumination Microscopy (SIM). Super-resolution microscopy using SIM provides image resolution significantly beyond the light diffraction limit in all three spatial dimensions, allowing the imaging of individual dendritic spines with improved detail.

Dendritic spines are protrusions emerging from the dendrite of a neuron and represent the primary postsynaptic targets of excitatory inputs in the brain. ...

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 ...

Assay to Measure Nucleocytoplasmic Transport in Real Time within Motor Neuron-like NSC-34 Cells

Nucleocytoplasmic transport refers to the import and export of large molecules from the cell nucleus. Recently, a number of studies have shown a connection between amyotrophic lateral sclerosis (ALS) and impairments in the nucleocytoplasmic pathway. ALS is a neurodegenerative disease affecting the motor neurons and resulting in paralysis and ultimately in death, within 2-5 years on average. Most cases of ALS are sporadic, lacking any apparent genetic linkage, but 10% are inherited in a dominant manner. Recently, hexanucleotide repeat expansions (HREs) in the chromosome 9 open reading frame 72 (C9orf72) gene were identified as a genetic cause of ALS and frontotemporal dementia (FTD). Importantly, different groups have recently proposed that these mutants affect nucleocytoplasmic transport. These studies have mostly shown the final outcome and manifestations caused by HREs on nucleocytoplasmic transport, but they do not demonstrate nuclear transport dysfunction in real time. As a result, only severe nucleocytoplasmic transport deficiency can be determined, mostly due to high overexpression or exogenous protein insertion. 
This protocol describes a new and very sensitive assay to evaluate and quantify nucleocytoplasmic transport dysfunction in real time. The rate of import of a NLS-NES-GFP protein (shuttle-GFP) can be quantified in real time using fluorescent microscopy. This is performed by using an exportin inhibitor, thus allowing the shuttle GFP only to enter the nucleus. To validate the assay, the C9orf72 HRE translated dipeptide repeats, poly(GR) and poly(PR), which have been previously shown to disrupt nucleocytoplasmic transport, were used. Using the described assay, a 50% decrease in the nuclear import rate was observed compared to the control. Using this system, minute changes in nucleocytoplasmic transport can be examined and the ability of different factors to rescue (even partially) a nucleocytoplasmic transport defect can be determined.

This protocol describes a method to sensitively measure the nucleocytoplasmic transport rate within motor neuron-like NSC-34 cells by quantifying the real-time change in the nuclear import of a NLS-NES-GFP protein.

Nucleocytoplasmic transport refers to the import and export of large molecules from the cell nucleus. Recently, a number of studies have shown a ...

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

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

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

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