We thank Ge Yang, Gaudenz Danuser, Khuloud Jaqaman, and Daniel Whisler for assistance with the adaptation and development of the software to quantitate cargo mapping analyses, and Emily Niederst for help in making the microfluidic devices. This work was supported in part by NIH-NIA grant AG032180 to L.S.B.G., and the Howard Hughes Medical Institute. L.S. was supported in part by a NIH Bioinformatics Training Grant T32 GM008806, S.E.E. was supported by a Damon Runyon Cancer Research Foundation Fellowship, NIH Neuroplasticity Training Grant AG000216, and by a grant from The Ellison Medical Foundation New Scholar in Aging Award, G.E.C was supported by an NIH/NCATS&#160;1 TL1 award TR001114 and by the Achievement Rewards for College Scientists foundation.

All experiments were conducted following approved protocols and according to institutional guidelines for the humane care of research animals. Neonate mice were euthanized by decapitation.
1. Preparation of Microfluidic Devices for Cell Culture
<ol>
	<li>Prepare polydimethyl siloxane (PDMS) microfluidic devices for growth of hippocampal neurons as described by Harris and colleagues17-19. Below are some modifications that were adapted to the cargo mapping protocol. 
	NOTE: Microfluidic devices are also commercially available (Materials List), thus access to a fabrication facility is not necessary.</li>
	<li>Prepare No. 1&#189; cover glasses (24 x 40 mm) by washing them three times with acetone, followed by three washes with 100% ethanol and three washes of water. Store coverslips in water at 4 &#176;C until use. These treatments help ensure that coverslips are clean from debris that might interfere with plating of cells, contamination, and survival. 
	NOTE: Acetone and ethanol are flammable and hazardous in case of skin contact (irritant), of eye contact (irritant), of ingestion, of inhalation. Dispose according to official regulations.</li>
	<li>When ready to use, place one cover glass in a 60 mm cell culture dish per microfluidic device, and let it air dry uncovered for 45 min inside a biosafety cabinet to avoid contamination.</li>
	<li>After devices are cut out from the masters and punches have been cut to create the reservoirs (Figure 1A, B), place them with the microfluidic channels side-up inside a biosafety cabinet equipped with an UV lamp for 45 min for sterilization. 
	NOTE: Leaving devices under UV treatment for longer than 2 hr might affect the integrity of the PDMS.</li>
	<li>Assemble the devices by placing one device onto each cover glass inside the 60 mm plastic cell culture dish so that the microfluidic channels face down. Apply gentle pressure to the top of the device (avoid touching the microchannels) to provide a non-permanent seal of the device to the cover glass. Cover each 60 mm cell culture dish with a lid.</li>
	<li>Using a micropipette, hydrate the devices by adding cell culture grade water to the top two microfluidic reservoirs (1 and 2 in Figure 1A), allowing water to flow through. Remove water with a micropipette and add 1 mg/ml poly-L-lysine into microfluidic reservoir 1 (200 &#181;l) and 2 (100 &#181;l). The volume differential will allow for the poly-L-lysine to flow through the microchannels (Figure 1A, B).</li>
	<li>To ensure that the devices inside the 60 mm cell culture dishes do not topple over inside a 37 &#176;C incubator, place three 60 mm cell culture dishes containing the devices into a 150 mm plastic cell culture dish and cover the dish with a lid. Incubate O/N in a 37 &#176;C incubator with 5% CO2. Typically more than 90% of the channels are usable per device and do not contain any air bubbles or physical blockage.</li>
	<li>On the next day, replace poly-L-lysine with water and leave device in incubator for at least 1 hr. Repeat this washing three times. Remove water and add Neurobasal-A growth medium (containing 2% B-27 supplement and 500 &#181;M GlutaMAX) to each device. Leave O/N in a 37 &#176;C incubator with 5% CO2.</li>
	<li>On the next day, plate hippocampal cells as described below.</li>
</ol>
2. Plating of Mouse Hippocampal Primary Neurons in Microfluidic Devices
NOTE: The preparation of cultured hippocampal neurons from neonate rats has been previously published in JoVE25. Below are some modifications that were adapted to plate mouse hippocampal neurons in microfluidic devices, and that were optimal for transfections.
<ol>
	<li>Before mouse brain dissection, pre-warm Dulbecco&#8217;s Modified Eagle Medium (DMEM) containing 10% fetal bovine serum (FBS; without antibiotics), Mixture A (125 mg DL-cysteine, 125 mg bovine serum albumin (BSA) and 3.125 g of D-glucose in 500 ml of phosphate buffered saline (PBS); filter sterilize through a 0.22 &#181;m filter), and deoxyribonuclease I solution (DNase I: 50 mg of DNase I and 0.5 ml of a 1.2 M MgSO4 solution in 100 ml Hank&#8217;s Balanced Salt Solution (HBSS) filter sterilize through a 0.22 &#181;m filter) in a 37 &#176;C water bath.
	<ol>
		<li>Pre-warm Neurobasal-A growth medium inside a 37 &#176;C incubator with 5% CO2 to equilibrate the pH.</li>
	</ol>
	</li>
	<li>Dissect hippocampi from 1-3 day old neonate mice according to established protocols26. Keep hippocampi in a 15 ml conical tube containing 10 ml dissection buffer (HBSS containing 10 mM HEPES pH 7.4, 50 mM glucose, 100 U/ml penicillin and 100 &#181;g/ml streptomycin), on ice. Put up to 4 hippocampi per 15 ml conical tube.</li>
	<li>Perform the remaining protocol is under sterile conditions inside a biosafety cabinet.</li>
	<li>Dilute 45 units of papain in 5 ml of Mixture A, vortex, and incubate for 5 min at 37 &#176;C until papain powder is dissolved. Add 1 ml of DNase I solution. Filter the mix through a 0.22 &#181;m syringe filter unit.</li>
	<li>Carefully aspirate HBSS&#8217; from 15 ml conical tube containing hippocampi. Add 10 ml cold (4 &#176;C) HBSS to hippocampi and let them settle to the bottom of the tube. Carefully aspirate HBSS from 15 ml conical tube.</li>
	<li>Add 1 ml papain/DNase I mix to up to 4 hippocampi and incubate for 20-30 min at 37 &#176;C. Tilt conical tube every 5 min to bathe hippocampi with mix. Alternatively place hippocampi in a 37 &#176;C water bath shaker with gentle shaking (~100 rpm).</li>
	<li>Aspirate papain/DNase I mix and wash hippocampi twice with pre-warmed DMEM (37 &#176;C) containing 10% FBS.</li>
	<li>After second wash, add 2 ml of pre-warmed DMEM containing 10% FBS to hippocampi and manually triturate hippocampi by pipetting up and down ~10-12 times using a 1 ml pipette tip to dissociate neurons.</li>
	<li>Let triturate settle for ~1 min as this will allow non-dissociated neurons to settle to the bottom of the conical tube. Transfer supernatant containing dissociated neurons to a new 15 ml conical tube avoiding transfer of larger tissue that has settled at the bottom of the tube.</li>
	<li>Spin cells at 1,000 x g for 2 min and carefully remove supernatant without disturbing the cell pellet. Resuspend cells in 80 &#956;l Neurobasal-A growth medium by pipetting gently.</li>
	<li>Remove medium from microfluidic reservoir 1 and 1&#8217;. Apply 20 &#181;l of cells (~275,000) to microfluidic reservoir 1, and allow them to flow through. Place device in 37 &#176;C incubator with 5% CO2 for 20 min.</li>
	<li>Look under microscope to ensure that cells have adhered to the cover glass. Add Neurobasal-A growth medium to top off all reservoirs. Return device with cells to a 37 &#176;C incubator with 5% CO2.</li>
	<li>Check devices every ~2-3 days and top off reservoirs with Neurobasal-A containing 2% B-27 supplement (without GlutaMAX) to avoid evaporation. Neurons start to extend their axons through microfluidic channels ~2-3 days after plating. Subject neurons to transfection typically 7-10 days after plating.</li>
</ol>
3. Transfection of Hippocampal Neurons Grown in Microfluidic Device
<ol>
	<li>Per device, prepare a mix of 1.2 &#181;l Lipofectamine 2000 in 30 &#181;l Neurobasal-A, and a mix of 0.5 &#181;g fluorescent cargo fusion plasmid in 30 &#181;l Neurobasal-A and incubate 5 min at RT. Add Lipofectamine mix to DNA mix and incubate for 20 min at RT. Add 60 &#181;l of Neurobasal-A containing 4% B-27 supplement to the DNA/Lipofectamine mix and mix by flicking the tube.</li>
	<li>Remove all media from microfluidic reservoir 2 and 2&#8217; and carefully fill up wells with Neurobasal-A containing 2% B-27 supplement without spilling over the medium into the other compartments.</li>
	<li>Remove media from microfluidic reservoir 1 and 1&#8217;. Add 120 &#181;l of DNA/Lipofectamine mix to microfluidic reservoir 1 and let it flow. Return device to 37 &#176;C incubator with 5% CO2 for 3-4 hr.</li>
	<li>Remove all media from microfluidic reservoir 1 and 1&#8217; and replace with Neurobasal-A containing 2% B-27 supplement for 24 hr (or until ready to image). Typically, 5-7 axons that grow through microchannels are transfected under those conditions, which represent approximately 1-3% transfection efficiency consistent with efficiencies published for hippocampal cultured cells 26.</li>
</ol>
4. &#8220;Cargo Mapping&#8221; Analyses
<ol>
	<li>Live Imaging of cargo movement
	<ol>
		<li>Perform the imaging on an inverted epifluorescence light microscope equipped with a 37 &#176;C incubator and a CO2 control chamber.</li>
		<li>Mount the coverslip with hippocampal neurons grown in a microfluidic device onto the microscope stage. To avoid neuronal death, work expeditiously to maintain the samples at the microscope for no longer than 1 hr.</li>
		<li>Using a high-resolution objective (100X), find axons of transfected cells that extend through channels of microfluidic chambers and record the number of the channels in which the transfected axon was found. Count the number of channels using a hand tally counter.
		<ol>
			<li>For optimal recording of live movement and subsequent colocalization analysis, choose axons that grow relatively flat through the channels so that imaging of most vesicles can be performed in focus.</li>
		</ol>
		</li>
		<li>Remove most of the medium from reservoirs 2 and 2&#8217; with a 1 ml plastic transfer pipette. To facilitate subsequent alignment of fixed images with movement trajectories on kymographs, align the right edge of the microchannels with the field of view during imaging (Figure 1B, C).</li>
		<li>To facilitate fixing of the samples during imaging, perform live imaging with two people. Alternatively, if live imaging is performed by a single person, use a microscope system equipped with drift correction.</li>
		<li>Start imaging live cargo movement with time-lapse specifications specific to the transport dynamics of the cargo being analyzed (microscope specifications and settings for imaging YFP-PrPC are specified in the legend of Figure 2 and Materials List).</li>
		<li>After sufficient live movement data have been collected, have one person fill up reservoirs 2 and 2&#8217; with pre-warmed 4% paraformaldehyde in PBS containing 0.04 g/ml sucrose to fix the cells. Avoid touching the microfluidic device, as this will disrupt the focus of the live imaging. Have the other person adjust the focus while the fixative is added. Fixation is successful when immediate stop of movement is observed. Add fixative to reservoir 1 and 1&#39; afterwards. 
		NOTE: Temporary photobleaching of the cargo fluorescence might also be observed, but the fluorescence persists after IF staining for motor proteins is completed (next step). 
		NOTE: Paraformaldehyde is harmful by inhalation, in contact with skin, and if swallowed. Irritating to eyes, respiratory system and skin. Dispose according to official regulations.</li>
	</ol>
	</li>
	<li>IF staining of motor proteins for colocalization analysis 
	NOTE: For quantitation of relative levels of motor proteins (as determined by antibody staining), associated to individual moving cargoes, validate and titrate antibodies to ensure that the antibody-antigen binding is saturated. This is done by determining the specificity of the antibody using neurons in which the protein of interest has been either knocked-out or knocked-down, and by performing IF analysis with increasing antibody concentrations. For negative controls, neurons are stained with secondary antibodies in the absence of primary antibodies. More detailed description of antibody validation can be found in Encalada et al.16, and Szpankowski et al.20.
	<ol>
		<li>Remove the coverslip with hippocampal neurons grown in microfluidic device from the microscope stage. Do not detach the microfluidic device from the coverslip, as the microchannels need to remain intact in order to superimpose the live and fixed images. Perform the following steps in a humidity chamber.</li>
		<li>To ensure the flow of solutions into the microchannels, maintain a buffer volume differential between reservoirs 1, 1&#8217;, and 2, 2&#8217; of at least 30 &#181;l. in all subsequent steps. For example, add 100 &#181;l media volume to microfluidic reservoir 2, 2&#8217;, and 70 &#181;l media volume to microfluidic reservoir 1, 1&#8217;.</li>
		<li>Remove fixative from chambers of the microfluidic device and wash cells three times with PBS waiting 10 min between washes.</li>
		<li>Permeabilize cells by adding 0.1% Triton X-100 diluted in PBS for 5 min to all reservoirs.</li>
		<li>Remove solution from all reservoirs and wash with PBS. Repeat washing three times waiting 10 min between washes.</li>
		<li>Remove solution from all reservoirs and apply blocking buffer containing 10% Normal Donkey Serum and 3% protease free and immunoglobulin G (IgG)-free BSA in PBS and incubate for at least 30 min at RT.</li>
		<li>Spin primary antibody stock solution at 20,000 x g for 5 min to remove precipitates. Dilute antibody to an appropriate concentration in blocking buffer. Replace the blocking buffer from microfluidic device with antibody solution. Incubate samples for 2 hr at RT or O/N at 4 &#176;C.</li>
		<li>Remove solution from all reservoirs and wash three times with PBS, waiting 10 min between washes.</li>
		<li>Spin secondary antibody stock solution at 20,000 x g for 5 min to remove precipitates. Dilute antibody to an appropriate concentration in blocking buffer. Replace PBS with secondary antibody solution. Incubate samples for 1 hr at RT.</li>
		<li>Remove solution from all reservoirs and wash three times with PBS, waiting 10 min between washes.</li>
		<li>Replace PBS with ~10-20 &#181;l mounting medium by directly pipetting mounting medium into the opening of the channels connecting the reservoirs. Let mounting media dry for ~20 min &#8211; 1 hr at RT.</li>
		<li>Store devices at 4 &#176;C protected from light to avoid photobleaching, and image axons within two days (maximum one week) of staining.</li>
		<li>After staining is completed, mount coverslip with hippocampal neurons grown in microfluidic device onto the microscope stage. Locate the region of the transfected axon imaged in step 4.1.1-4.1.7.</li>
		<li>Acquire Z-stack images (300 nm steps) for each of the three channels: cargo, motor 1, motor 2, using a high-resolution objective (e.g., a high numerical aperture oil or water objective). 
		NOTE: Acquisition of Z-stacks is important as axons often do not grow perfectly flat in microchannels and therefore not all fluorescent puncta are in the same focal plane.</li>
	</ol>
	</li>
	<li>Cargo Mapping
	<ol>
		<li>Generate a kymograph of cargo movement using image analysis software. The ImageJ plugin developed by Rietdorf and Seitz (EMBL) is recommended for generating kymographs (for download see Materials List). 
		NOTE: ImageJ is a public domain, Java-based image processing program developed at the National Institutes of Health27,28 (for download see Materials List).</li>
		<li>Align the fixed images of the fluorescent cargo (generated in step 4.2.14) manually by superimposing them onto the position of the trajectories in the kymograph at the point of fixation (Figure 2A, B), using a commercially available graphing software program (Materials List). Manually align by first superimposing the fixed cargo image onto the kymograph and manually choosing the cargo puncta that corresponds to a specific trajectory on the kymograph. For best alignment results, use the Z slice that is in best focus for the cargo mapped. Several Z slices may be used.</li>
		<li>For each image in the Z-stack (cargo, motor 1 and motor 2; generated in step 4.2.14) determine the X-Y coordinates and intensity amplitudes for each fluorescent puncta using an established algorithm to fit 2D Gaussians to the point spread function that represents each point source in each of the three channels16,20,21 (Figure 2D). 
		NOTE: To obtain the X-Y coordinates, a custom built MATLAB software package called &#8220;Motor Colocalization&#8221; was developed16,20, which incorporates a Gaussian fitting algorithm developed by Jaqaman, Danuser and colleagues21-23. The software package and detailed instructions for its use can be obtained upon request.</li>
		<li>For each of the individually cargo puncta that were mapped onto the mobile or stationary trajectories on the kymograph, select the X-Y coordinates and intensity amplitudes from the results of the &#8220;Motor Colocalization&#8221; program (these cargoes are individually labeled in Figure 2B). Use several Z-stack images acquired in step 4.2.14) to map more cargoes that are found on different focal planes.</li>
		<li>Compare the individual X-Y coordinates for the mapped cargo to those obtained for each of the motor channels in the corresponding Z slice (Figure 2D). Determine and select the X-Y motor coordinates that are within a 300 nm radius of the cargo. For cargoes and motors that have signal within the same focal plane, use the &#8220;Motor Colocalization&#8221; software package to determine the puncta that are located within a 300 nm radius.</li>
	</ol>
	</li>
</ol>

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J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ImageJ+for+microscopy.">ImageJ for microscopy.</a> Biotechniques. 43, 25-30 (2007).</li><li>Schneider, C. A., Rasband, W. S., Eliceiri, K. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=NIH+Image+to+ImageJ:+25+years+of+image+analysis.">NIH Image to ImageJ: 25 years of image analysis.</a> Nat. Methods. 9, 671-675 (2012).</li><li>Moore, D. S., McCabe, G. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Introduction to the Practice of Statistics. , (2005).</li></ol>

The authors have nothing to disclose.

The protocol presented here enables the correlation of directionality of movement of individual fluorescent microtubule-based moving cargo particles with the relative type and amount of associated motor proteins in live neurons. Previously, the total motor composition of axonal vesicular cargoes was assayed on heterogeneous populations of biochemically purified vesicles and organelles9,15. However, characterizing motor composition for a single type of cargo inside cells has been chal...

Intracellular transport is critical in all cell types for the delivery of proteins, membranes, organelles, and signaling molecules to various cellular domains1. Neurons are highly specialized cells with long, polarized projections that critically depend on intracellular transport of essential cargoes for their long-distance delivery to various axonal microdomains. This transport is mediated by kinesins and dyneins &#8212; two large families of molecular motor proteins &#8212; that bind to cargoes and track along polarized microtubules in anterograde and retrograde directions, respectively. While retrograde movement is mainly mediated by dynein, movement in the anterograde direction is facilitated by a large, functionally diverse family of kinesin motors. Consequently, anterograde transport of axonal cargoes could be mediated by various family members of the kinesin superfamily1-5. Though some cargoes move persistently in either direction, most cargoes move bidirectionally and reverse frequently on their way to their final destinations1,5-13. Furthermore, it has been shown that motors of opposing directionality associate simultaneously to cargoes, raising the question as to how regulated movement of cargoes is coordinated by opposite-polarity motors5-7. Together, transport of axonal cargoes is a concerted process that is regulated by the composition of motors and their specific biochemical activities, which in turn are dependent on various adaptors and regulatory binding partners14.
To faithfully describe the mechanism of axonal transport for a specific cargo and to uncover the underlying regulation of that transport, it is paramount to determine the composition of motor proteins and their regulatory binding partners associated with individual cargoes during their live transport. Other methods, for example biochemical approaches, provide estimates of average motor compositions on purified heterogeneous vesicle populations, but these estimates do not reveal the type or amounts of motors associated to single moving vesicles. Also, reconstitution of vesicle transport along preassembled microtubules in vitro enabled measuring the amount of one type of motor on a single vesicle level15. However, these experiments did not directly correlate the amount of motors with the transport characteristics of those vesicles, and measured transport in the absence of cellular regulatory factors.
A protocol is presented here, which determines the motor composition (type and relative amount of motors) of individual moving vesicles from immunofluorescence (IF) data measuring endogenously expressed motor proteins, and correlates these parameters to the live transport of the exact same vesicles in neurons16. This method entails precise mapping of IF-to-live cargo movement data. This is accomplished by growing hippocampal mouse neurons in microfluidic devices following established protocols17-19. These devices allow for the identification and correlation (&#8220;mapping&#8221;) of axons and single moving cargoes in fixed and live light microscopy modalities (Figure 1). Cultured neurons are transfected with fluorescently labeled cargo proteins whose transport is imaged at high spatial and temporal resolution to obtain detailed movement information that is plotted in kymographs. During the course of imaging, neurons are fixed with paraformaldehyde, and subsequently stained with antibodies against endogenous motor proteins. Fixed cargo and motor images are superimposed onto live movement kymographs to &#8220;map&#8221; (colocalize) them to the live cargo movement trajectories16. To correlate the live movement of cargoes with the association of motor proteins, colocalization is analyzed using a custom made MATLAB software package called &#8220;Motor Colocalization&#8221;16,20. Fluorescently labeled cargoes and motors generate diffraction-limited punctate features that can partially overlap. To resolve the position of overlapping puncta, the software first automatically fits Gaussian functions to each point spread function, representing individual fluorescent puncta, to determine their precise X-Y sub-pixel position coordinates and intensity amplitudes21-23. The positions of motors and cargoes are subsequently compared to each other to determine colocalization16,20. Therefore, this method more precisely assigns colocalization between fluorescent puncta as compared to other methods24.
The strength of this method is the ability to assess the colocalization of motors with individual cargo in fixed cells, for which live movement trajectories (e.g., the direction in which they were moving at the time of fixation) have been recorded. With this method, kinesins and dyneins were found to associate simultaneously to vesicles that carry the normal prion protein (PrPC-cellular), a neuronally enriched cargo that moves bidirectionally or remains stationary in axons16. This analysis allowed the formulation of a working model for the regulation of PrPC vesicle movement in which anterograde (kinesin) and retrograde (dynein) motors coordinate their activities in order to move the vesicles in either direction or to remain stationary while associated to the cargo. Another strength of this method is its potential broad applicability for characterizing colocalization/association of many fluorescently labeled cargoes that move in virtually any cell type, with any other protein(s) of interest. Thus, live/fixed correlation could potentially allow for the detection of transient protein-cargo interactions, as many individual fluorescently labeled moving particles can be analyzed over a desired period of time. Given the broad applicability and the type of questions that this method can address, this protocol will be of interest to a wide audience of cell biologists including those studying trafficking and transport in neurons or in other cell types.

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			<td height="21" style="height:21px;">poly-L-lysine</td>
			<td>Sigma</td>
			<td>P5899-20mg</td>
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			<td height="21" style="height:21px;">D-MEM (Dulbecco&#8217;s Modified Eagle Medium) High Glucose, w/ L-Glutamine, w/o Sodium Pyruvate (1X)</td>
			<td>Life Technologies</td>
			<td>11965092</td>
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			<td height="21" style="height:21px;">FBS (Fetal Bovine Serum)</td>
			<td>Life Technologies</td>
			<td>10082147</td>
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			<td height="21" style="height:21px;">Neurobasal-A Medium (1X)&#160;</td>
			<td>Life Technologies</td>
			<td>10888022</td>
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			<td height="21" style="height:21px;">B-27 Serum Free Supplement</td>
			<td>Life Technologies</td>
			<td>10888022</td>
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			<td height="21" style="height:21px;">GlutaMAX I Supplement (100X)</td>
			<td>Life Technologies</td>
			<td>35050061</td>
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			<td height="21" style="height:21px;">HBSS (1X) (Hank&#8217;s Balanced Salt Solution)</td>
			<td>Life Technologies</td>
			<td>24020117</td>
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			<td height="21" style="height:21px;">DPBS (Dulbecco&#8217;s Phosphate Buffered Saline, no Magnesium, no Calcium (1X)</td>
			<td>Life Technologies</td>
			<td>14190250</td>
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			<td height="21" style="height:21px;">Penicillin/Streptomycin (100X)</td>
			<td>Life Technologies</td>
			<td>15140122</td>
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			<td height="21" style="height:21px;">Corning cellgro Water for Cell Culture&#160;</td>
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			<td>USB Corporation</td>
			<td>19925</td>
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			<td height="21" style="height:21px;">DL-cysteine HCl</td>
			<td>Sigma-Aldrich</td>
			<td>C9768</td>
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			<td height="21" style="height:21px;">BSA (bovine serum albumin)</td>
			<td>Sigma-Aldrich</td>
			<td>A7906</td>
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			<td>Sigma-Aldrich</td>
			<td>G6152</td>
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			<td height="21" style="height:21px;">DNAse I grade II</td>
			<td>Roche Applied Sciences</td>
			<td>10104159001</td>
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			<td height="21" style="height:21px;">Lipofectamine 2000</td>
			<td>Life Technologies</td>
			<td>&#160;11668027</td>
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			<td height="21" style="height:21px;">Formaldehyde Solution 16% EM Grade</td>
			<td>Fisher Scientific</td>
			<td>50980487</td>
			<td>Caution: Harmful by inhalation, in contact with skin and if swallowed. Irritating to eyes, respiratory system and skin. Dispose according to official regulations.</td>
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			<td>Fisher Scientific</td>
			<td>S5-500</td>
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			<td>Sigma</td>
			<td>H-3375</td>
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			<td>Jackson Immuno Research</td>
			<td>017-000-0121</td>
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			<td>Jackson Immuno Research</td>
			<td>001-000-162</td>
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			<td height="21" style="height:21px;">Acetone</td>
			<td>Fisher Scientific</td>
			<td>BP2403-4</td>
			<td></td>
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			<td height="21" style="height:21px;">Ethanol</td>
			<td>Fisher Scientific</td>
			<td>BP2818-100</td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;">ProLong Gold antifade reagent</td>
			<td>Life Technologies</td>
			<td>P36934</td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;">Cover Glass 1 1/2. 24X40mm</td>
			<td>Corning</td>
			<td>2980-244</td>
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			<td height="21" style="height:21px;">Axis microfluidic device, 450 &#181;m</td>
			<td>Millipore</td>
			<td>AX450</td>
			<td></td>
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			<td height="21" style="height:21px;">Adobe Photoshop</td>
			<td>Adobe</td>
			<td>N/A</td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;">Nikon Eclipse TE2000-U&#160;</td>
			<td>Nikon</td>
			<td>N/A</td>
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			<td height="21" style="height:21px;">Coolsnap HQ camera&#160;</td>
			<td>Roper Scientific</td>
			<td>N/A</td>
			<td></td>
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			<td height="21" style="height:21px;">60 mm cell culture dish</td>
			<td>Fisher Scientific</td>
			<td>12-565-95</td>
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			<td height="21" style="height:21px;">150 mm cell culture dish</td>
			<td>Fisher Scientific</td>
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			<td height="21" style="height:21px;">Anti-Kinesin light chain, V-17</td>
			<td>Santa Cruz&#160;</td>
			<td>sc-13362</td>
			<td>specificity verified using KLC1-/- neurons (Ref. 16 ), recommended dilution 1:100.</td>
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		<tr height="21">
			<td height="21" style="height:21px;">Anti Dynein Heavy Chain 1, R-325</td>
			<td>Santa Cruz&#160;</td>
			<td>sc-9115</td>
			<td>specificity verified using shRNA against DYN1HC1 (Ref. 16), recommended dilution 1:100</td>
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			<td height="21" style="height:21px;">Alexa Fluor 568 Donkey Anti-Rabbit IgG Antibody</td>
			<td>Life Technologies</td>
			<td>A10042</td>
			<td>&#160;recommended dilution 1:200.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Alexa Fluor 647 Donkey Anti-Rabbit IgG (H+L) Antibody</td>
			<td>Life Technologies</td>
			<td>A31573</td>
			<td>&#160;recommended dilution 1:200.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Alexa Fluor 568 Donkey Anti-Goat IgG (H+L) Antibody IgG Antibody</td>
			<td>Life Technologies</td>
			<td>A11057</td>
			<td>&#160;recommended dilution 1:200.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Alexa Fluor 647 Donkey Anti-Goat IgG (H+L) Antibody</td>
			<td>Life Technologies</td>
			<td>A21447</td>
			<td>&#160;recommended dilution 1:200.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;"></td>
			<td></td>
			<td></td>
			<td></td>
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			<td></td>
			<td></td>
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			<td height="21" style="height:21px;">Plugins and Macros</td>
			<td></td>
			<td></td>
			<td></td>
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			<td height="21" style="height:21px;">ImageJ</td>
			<td></td>
			<td></td>
			<td><a href="http://imagej.nih.gov/ij/index.html">&#160;http://imagej.nih.gov/ij/index.html.&#160;</a></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">ImageJ Kymoraph Plugin</td>
			<td></td>
			<td></td>
			<td><a href="http://www.embl.de/eamnet/html/body_kymograph.html">http://www.embl.de/eamnet/html/body_kymograph.html.</a></td>
		</tr>
	</tbody>
</table>

characterizing the composition of molecular motors on moving axonal cargo using "cargo mapping" analysis

Understanding the mechanisms by which molecular motors coordinate their activities to transport vesicular cargoes within neurons requires the quantitative analysis of motor/cargo associations at the single vesicle level. The goal of this protocol is to use quantitative fluorescence microscopy to correlate (&#8220;map&#8221;) the position and directionality of movement of live cargo to the composition and relative amounts of motors associated with the same cargo. &#8220;Cargo mapping&#8221; consists of live imaging of fluorescently labeled cargoes moving in axons cultured on microfluidic devices, followed by chemical fixation during recording of live movement, and subsequent immunofluorescence (IF) staining of the exact same axonal regions with antibodies against motors. Colocalization between cargoes and their associated motors is assessed by assigning sub-pixel position coordinates to motor and cargo channels, by fitting Gaussian functions to the diffraction-limited point spread functions representing individual fluorescent point sources. Fixed cargo and motor images are subsequently superimposed to plots of cargo movement, to &#8220;map&#8221; them to their tracked trajectories. The strength of this protocol is the combination of live and IF data to record both the transport of vesicular cargoes in live cells and to determine the motors associated to these exact same vesicles. This technique overcomes previous challenges that use biochemical methods to determine the average motor composition of purified heterogeneous bulk vesicle populations, as these methods do not reveal compositions on single moving cargoes. Furthermore, this protocol can be adapted for the analysis of other transport and/or trafficking pathways in other cell types to correlate the movement of individual intracellular structures with their protein composition. Limitations of this protocol are the relatively low throughput due to low transfection efficiencies of cultured primary neurons and a limited field of view available for high-resolution imaging. Future applications could include methods to increase the number of neurons expressing fluorescently labeled cargoes.

Figure 1 shows an overview of the microfluidic device used to grow hippocampal neurons (Figure 1A, B). Neurons are plated in reservoir 1. The size of the microchannels prevents the diffusion of cell bodies (soma) into the axonal compartment while the length of the channels prevents dendritic projections from crossing all the way to the axonal compartment. After ~2-3 days in culture, neurons start extending their axons across microchannels into the axonal compartment (Figure 1B, C...

Watch this Scientific Journal Video about Characterizing the Composition of Molecular Motors on Moving Axonal Cargo Using "Cargo Mapping" Analysis at JoVE.com

Characterizing the Composition of Molecular Motors on Moving Axonal Cargo Using &quot;Cargo Mapping&quot; Analysis

Intracellular transport of cargoes, such as vesicles or organelles, is carried out by molecular motor proteins that track on polarized microtubules. This protocol describes the correlation of the directionality of transport of individual cargo particles moving inside neurons, to the relative amount and type of associated motor proteins.

Characterizing the Composition of Molecular Motors on Moving Axonal Cargo Using "Cargo Mapping" Analysis

Understanding the mechanisms by which molecular motors coordinate their activities to transport vesicular cargoes within neurons requires the quantitative ...

characterizing-composition-molecular-motors-on-moving-axonal-cargo

Research

JoVE Journal

Neuroscience

9.3K vistas.  The Scripps Research Institute. Intracellular transport of cargoes, such as vesicles or organelles, is carried out by molecular motor proteins that track on polarized microtubules. This protocol describes the correlation of the directionality of transport of individual cargo particles moving inside neurons, to the relative amount and type of associated motor proteins.

Video: Characterizing the Composition of Molecular Motors on Moving Axonal Cargo Using "Cargo Mapping" Analysis

High-density EEG Recordings of the Freely Moving Mice using Polyimide-based Microelectrode

Electroencephalogram (EEG) indicates the averaged electrical activity of the neuronal populations on a large-scale level. It is widely utilized as a noninvasive brain monitoring tool in cognitive neuroscience as well as a diagnostic tool for epilepsy and sleep disorders in neurology. However, the underlying mechanism of EEG rhythm generation is still under the veil. Recently introduced polyimide-based microelectrode (PBM-array) for high resolution mouse EEG1 is one of the trials to answer the neurophysiological questions on EEG signals based on a rich genetic resource that the mouse model contains for the analysis of complex EEG generation process. This application of nanofabricated PBM-array to mouse skull is an efficient tool for collecting large-scale brain activity of transgenic mice and accommodates to identify the neural correlates to certain EEG rhythms in conjunction with behavior. However its ultra-thin thickness and bifurcated structure cause a trouble in handling and implantation of PBM-array. In the presented video, the preparation and surgery steps for the implantation of PBM-array on a mouse skull are described step by step. Handling and surgery tips to help researchers succeed in implantation are also provided.

In this article, we described the surgery procedure and handling tips for implantation of ultra-thin polyimide-based microelectrode array (PBM-array) on the mouse skull for acquisition of high-density encephalography (EEG) in a mouse model.

Electroencephalogram (EEG) indicates the averaged electrical activity of the neuronal populations on a large-scale level. It is widely utilized as a ...

Visualization and Genetic Manipulation of Dendrites and Spines in the Mouse Cerebral Cortex and Hippocampus using In utero Electroporation

In utero electroporation (IUE) has become a powerful technique to study the development of different regions of the embryonic nervous system 1-5. To date this tool has been widely used to study the regulation of cellular proliferation, differentiation and neuronal migration especially in the developing cerebral cortex 6-8. Here we detail our protocol to electroporate in utero the cerebral cortex and the hippocampus and provide evidence that this approach can be used to study dendrites and spines in these two cerebral regions.
Visualization and manipulation of neurons in primary cultures have contributed to a better understanding of the processes involved in dendrite, spine and synapse development. However neurons growing in vitro are not exposed to all the physiological cues that can affect dendrite and/or spine formation and maintenance during normal development. Our knowledge of dendrite and spine structures in vivo in wild-type or mutant mice comes mostly from observations using the Golgi-Cox method 9. However, Golgi staining is considered to be unpredictable. Indeed, groups of nerve cells and fiber tracts are labeled randomly, with particular areas often appearing completely stained while adjacent areas are devoid of staining. Recent studies have shown that IUE of fluorescent constructs represents an attractive alternative method to study dendrites, spines as well as synapses in mutant / wild-type mice 10-11 (Figure 1A). Moreover in comparison to the generation of mouse knockouts, IUE represents a rapid approach to perform gain and loss of function studies in specific population of cells during a specific time window. In addition, IUE has been successfully used with inducible gene expression or inducible RNAi approaches to refine the temporal control over the expression of a gene or shRNA 12. These advantages of IUE have thus opened new dimensions to study the effect of gene expression/suppression on dendrites and spines not only in specific cerebral structures (Figure 1B) but also at a specific time point of development (Figure 1C).
Finally, IUE provides a useful tool to identify functional interactions between genes involved in dendrite, spine and/or synapse development. Indeed, in contrast to other gene transfer methods such as virus, it is straightforward to combine multiple RNAi or transgenes in the same population of cells. 
In summary, IUE is a powerful method that has already contributed to the characterization of molecular mechanisms underlying brain function and disease and it should also be useful in the study of dendrites and spines.

Visualization and Genetic Manipulation of Dendrites and Spines in the Mouse Cerebral Cortex and Hippocampus using In utero Electroporation

This article describes in detail a protocol to electroporate in utero the cerebral cortex and the hippocampus at E14.5 in mice. We also show that this is a valuable method to study dendrites and spines in these two cerebral regions.

In utero electroporation (IUE) has become a powerful technique to study the development of different regions of the embryonic nervous system 1-5. To date ...

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 ...

The Olfactory System as a Model to Study Axonal Growth Patterns and Morphology In Vivo

The olfactory system has the unusual capacity to generate new neurons throughout the lifetime of an organism. Olfactory stem cells in the basal portion of the olfactory epithelium continuously give rise to new sensory neurons that extend their axons into the olfactory bulb, where they face the challenge to integrate into existing circuitry. Because of this particular feature, the olfactory system represents a unique opportunity to monitor axonal wiring and guidance, and to investigate synapse formation. Here we describe a procedure for in vivo labeling of sensory neurons and subsequent visualization of axons in the olfactory system of larvae of the amphibian Xenopus laevis. To stain sensory neurons in the olfactory organ we adopt the electroporation technique. In vivo electroporation is an established technique for delivering fluorophore-coupled dextrans or other macromolecules into living cells. Stained sensory neurons and their axonal processes can then be monitored in the living animal either using confocal laser-scanning or multiphoton microscopy. By reducing the number of labeled cells to few or single cells per animal, single axons can be tracked into the olfactory bulb and their morphological changes can be monitored over weeks by conducting series of in vivo time lapse imaging experiments. While the described protocol exemplifies the labeling and monitoring of olfactory sensory neurons, it can also be adopted to other cell types within the olfactory and other systems.

The Olfactory System as a Model to Study Axonal Growth Patterns and Morphology In Vivo

We describe a protocol for in vivo labeling of olfactory sensory neurons by electroporation and subsequent confocal laser-scanning or multiphoton microscopy to visualize neuronal morphology and its development over time.

The olfactory system has the unusual capacity to generate new neurons throughout the lifetime of an organism. Olfactory stem cells in the basal portion of ...

Visualization of the Axonal Projection Pattern of Embryonic Motor Neurons in Drosophila

The establishment of functional neuromuscular circuits relies on precise connections between developing motor axons and target muscles. Motor neurons extend growth cones to navigate along specific pathways by responding to a large number of axon guidance cues that emanate from the surrounding extracellular environment. Growth cone target recognition also plays a critical role in neuromuscular specificity. This work presents a standard immunohistochemistry protocol to visualize motor neuron projections of late stage-16 Drosophila melanogaster embryos. This protocol includes a few key steps, including a genotyping procedure, to sort the desired mutant embryos; an immunostaining procedure, to tag embryos with fasciclin II (FasII) antibody; and a dissection procedure, to generate filleted preparations from fixed embryos. Motor axon projections and muscle patterns in the periphery are much better visualized in flat preparations of filleted embryos than in whole-mount embryos. Therefore, the filleted preparation of fixed embryos stained with FasII antibody provides a powerful tool to characterize the genes required for motor axon pathfinding and target recognition, and it can also be applied to both loss-of-function and gain-of-function genetic screens.

Visualization of the Axonal Projection Pattern of Embryonic Motor Neurons in Drosophila

This work details a standard immunohistochemistry method to visualize motor neuron projections of late stage-16 Drosophila melanogaster embryos. The filleted preparation of fixed embryos stained with FasII antibody provides a powerful tool to characterize the genes required for motor axon pathfinding and target recognition during neural development.

The establishment of functional neuromuscular circuits relies on precise connections between developing motor axons and target muscles. Motor neurons ...

Screening of Axonal Degeneration in Carpal Tunnel Syndrome Using Ultrasonography and Nerve Conduction Studies

Axonal degeneration, indicative of surgical decompression, may coexist in carpal tunnel syndrome (CTS) as the disease progresses. However, the current diagnostic and severity gradation system cannot clearly indicate its coexistence, resulting in&#160;confusion of appropriate treatment prescription. There are also constraints in conventional methods for&#160;differentiation as well. This study aims at introducing an innovative, efficient, and quick screening protocol to differentiate axonal degeneration associated with CTS, using ultrasound and nerve conduction studies (NCS). It starts by using NCS to perform orthodromic stimulation at the wrist, to obtain the sensory conduction of the median and the ulnar nerves respectively. Meanwhile, the motor conduction&#160;of the median nerve is collected by stimulating the palm, wrist, and elbow, followed by the stimulation of the ulnar nerve at the wrist,&#160;below and above the elbow. Then, an ultrasound assessment is performed, using a linear array transducer, with cross-sectional area (CSA) and perimeter (P) at the wrist and at the one-third distal forearm calipered. Ratios (R-CSA, R-P) and changes from wrist to one-third distal forearm (&#916;CSA and &#916;P) are calculated according to a standard format. Potential axonal degeneration coexisting in CTS will be screened according to the criteria of NCS and cut-off values of ultrasound measurements established in a previous study. In terms of its noninvasiveness, low cost, convenience, and efficiency, it is easy to apply ultrasound complimentarily in clinical practice to prescreen patients with potential coexisting axonal degeneration. Nevertheless, the ultrasonographic imaging cannot directly reflect axonal degeneration. It still relies on conventional but invasive methods&#160;such as electromyography (EMG) and biopsy&#160;for confirmation if needed.

Here we present a protocol using nerve conduction studies and ultrasound to screen potential axonal degeneration associated with carpal tunnel syndrome. The criteria for differentiation are established. Compared to conventional approaches, this method is noninvasive, convenient, and efficient, with an overall satisfactory accuracy, sensitivity, and specificity.

Axonal degeneration, indicative of surgical decompression, may coexist in carpal tunnel syndrome (CTS) as the disease progresses. However, the current ...

Generation of 3-D Collagen-based Hydrogels to Analyze Axonal Growth and Behavior During Nervous System Development

This protocol uses natural type I collagen to generate three-dimensional (3-D) hydrogel for monitoring and analyzing the axonal growth. The protocol is centered on culturing small pieces of embryonic or early postnatal rodent brains inside a 3-D hydrogel formed by the rat tail tendon-derived type I collagen with specific porosity. Tissue pieces are cultured inside the hydrogel and confronted to specific brain fragments or genetically-modified cell aggregates to produce and secrete molecules suitable for creating a gradient inside the porous matrix. The steps of this protocol are simple and reproducible but include critical steps to be considered carefully during its development. Moreover, the behavior of growing axons can be monitored and analyzed directly using a phase-contrast microscope or mono/multiphoton fluorescence microscope after fixation by immunocytochemical methods.

Here, we provide a method for analyzing the behavior of growing axons in 3D matrices, mimicking their natural development.

This protocol uses natural type I collagen to generate three-dimensional (3-D) hydrogel for monitoring and analyzing the axonal growth. The protocol is ...

Analysis of Spliceosomal snRNA Localization in Human Hela Cells Using Microinjection

Biogenesis of spliceosomal snRNAs is a complex process involving both nuclear and cytoplasmic phases and the last step occurs in a nuclear compartment called the Cajal body. However, sequences that direct snRNA localization into this subnuclear structure have not been known until recently. To determine sequences important for accumulation of snRNAs in Cajal bodies, we employed microinjection of fluorescently labelled snRNAs followed by their localization inside cells. First, we prepared snRNA deletion mutants, synthesized DNA templates for in vitro transcription and transcribed snRNAs in the presence of UTP coupled with Alexa488. Labelled snRNAs were mixed with 70 kDa-Dextran conjugated with TRITC, and microinjected to the nucleus or the cytoplasm of human HeLa cells. Cells were incubated for 1 h and fixed and the Cajal body marker coilin was visualized by indirect immunofluorescence, while snRNAs and dextran, which serves as a marker of nuclear or cytoplasmic injection, were observed directly using a fluorescence microscope. This method allows for efficient and rapid testing of how various sequences influence RNA localization inside cells. Here, we show the importance of the Sm-binding sequence for efficient localization of snRNAs into the Cajal body.

Biogenesis of spliceosomal snRNAs is a complex process involving various cellular compartments. Here, we employed microinjection of fluorescently labelled snRNAs in order to monitor their transport inside the cell.

Biogenesis of spliceosomal snRNAs is a complex process involving both nuclear and cytoplasmic phases and the last step occurs in a nuclear compartment ...

Induction of Diffuse Axonal Brain Injury in Rats Based on Rotational Acceleration

Traumatic brain injury (TBI) is a major cause of death and disability. Diffuse axonal injury (DAI) is the predominant mechanism of injury in a large percentage of TBI patients requiring hospitalization. DAI involves widespread axonal damage from shaking, rotation or blast injury, leading to rapid axonal stretch injury and secondary axonal changes that are associated with a long-lasting impact on functional recovery. Historically, experimental models of DAI without focal injury have been difficult to design. Here we validate a simple, reproducible and reliable rodent model of DAI that causes widespread white matter damage without skull fractures or contusions.

This protocol validates a reliable, easy-to-perform and reproducible rodent model of brain diffuse axonal injury (DAI) that induces widespread white matter damage without skull fractures or contusions.

Traumatic brain injury (TBI) is a major cause of death and disability. Diffuse axonal injury (DAI) is the predominant mechanism of injury in a large ...

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

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.

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.