We give special thanks to Lino Kim, Steve Guzman and Ujit Satyarthi for technical assistance during the development of this method. Myron Hawthorne, Richard Spengel, and Roberto Rojas machined the chambers used here and provided much-needed technical assistance. Notably, Roberto Rojas produced Figure 4. We are also thankful for Scott Fraser&#x2019;s invaluable advice prior to the development of the above chemotaxis assay. This work was partly supported by an NIH-MBRS SCORE-5S06GM048680-13 to MEdB and by an award from the CSU, Northridge Graduate Thesis Support Program to CW.

1. Day 1: Isolation of trunk neural tubes for overnight culture on coverslips
<ol>
	<li>Incubate chick eggs for 56 h at 38&deg; C. Remove the eggs from incubation, mildly spray them with 70% Ethanol, and then allow them to dry. Break the eggs open into a UV-sterilized glass tray.</li>
	<li>Extract each embryo from its yolk and place it in chick Ringer's. Do this by first cutting around its blood islands with curved scissors; then, with blunt forceps, pick the embryo up by its extraembryonic membrane and place it in a sterile plastic Petri dish containing chick Ringer&prime;s solution.</li>
	<li>Isolate the trunk of each embryo by trimming off excess extraembryonic membranes as well as cranial, vagal, and sacral axial levels using a tungsten needle (Fig. 1). First, select about 9 embryos that are between stages HH15-17 11. For stages HH15 and up, the forebrain and hindbrain axes form an acute angle and therefore the head appears to tilt caudally. At stage HH17, the tail bud is present and tilts ventrally but does not yet contain somites. With a tungsten needle, trim off extraembryonic membranes to about 2 mm from the embryo and cut off any embryonic tissues anterior to somite 10. Also remove all caudal embryonic tissues starting from around the fifth most newly formed somite.</li>
	<li>Place the isolated embryonic trunks in Dispase (0.24 U/ml DMEM) and incubate them for 1 h 15 min at 37&deg; C and 5% CO2. Once the trunks begin incubation, start preparing 6 coverslips (CS) for culturing NT explants (steps 1.5-1.8).</li>
	<li>Rinse 6 CS in 70% ethanol (diluted in sterile, ultrapure water), and then allow them to dry. Using a lab marker, draw a circle in the center of each CS that is about 1 cm in diameter (this circle will later help you identify where a fibronectin coat has been applied). On the same face of each CS, write the word "be" (or some other asymmetrical word or shape) outside the drawn circle (this will help you identify whether the marked side of the CS is facing up or down).</li>
	<li>Place each CS in a separate 40 x 10 mm sterile dish with the marked surface facing down and allow the dish to sit open underneath a germicidal UV lamp for 10 min.</li>
	<li>Apply 60 &mu;l fibronectin (FN; 10 &mu;g/ml DMEM) to the unmarked surface of the CS while making sure the entire area within the 1 cm circle is coated. Place the dishes to incubate at 37&deg; C for 30 min and then carefully aspirate the fibronectin from each CS.</li>
	<li>Add 250 &mu;l "culture" medium [DMEM with L-Glutamine (2 mM), Penicillin (100U/ml), Streptomycin (100 mcg/ml), and 8% fetal bovine serum (FBS)] to the FN-coated area of the CS. Place the dishes containing each CS at 37&deg; C and 5% CO2 again until the NTs have been isolated.</li>
	<li>Transfer all the incubated embryo trunks to one 5 cm glass Petri dish containing L15 medium and start dissecting out each NT using fine forceps and a tungsten needle (Fig. 1). Carefully slice along the border of the NT and somites with a sharp tungsten needle while being cautious not to damage the NT. It is often easier to begin isolating each NT from the caudal-most end of the trunk.</li>
	<li>Select 6 of the straightest and longest NTs to culture overnight (NTs between roughly 8 and 15 somites long are recommended). Using a micropipette tip primed with culture medium, transfer each of the 6 NTs to its own previously prepared CS (from steps 1.5 through 1.8). Be sure that the NT does not remain floating at the surface. If the NT is floating, drip medium onto it until it sinks using a micropipette.</li>
	<li>Place each dish at 37&deg; C and 5% CO2 overnight. Be careful to ensure each NT is within the FN-coated area of its respective CS immediately prior to placing the dish in the incubator (by using the circle drawn in step 1.5 as a reference). A micropipette can be used to better adjust the position of each NT if needed.</li>
	<li>Place at least 2 ml of culture medium (without serum) into a sterile 15 ml centrifuge tube and incubate overnight at 37&deg; C and 5% CO2. Leave the cap slightly unscrewed to allow the pH of the medium to adjust overnight. Pre-incubating the medium is important to help prevent bubble formation in your chamber, which may disrupt the establishment of a molecular gradient. Such "preincubated " medium should be used in all future steps. When not being used, this medium should be incubating at 37&deg; C.</li>
</ol>
2. Day 2: Loading the modified Zigmond chamber and time-lapse analysis of cell migration
<ol>
	<li>Out of the 6 NTs cultured, select the 3 cultures best suited for analysis. Generally, NCC cultures that have at least one long, straight edge should be selected (Fig. 2A). The 3 best cultures will be used to load and film 3 modified Zigmond chambers throughout the day, each with a different treatment. Out of the 3 cultures, choose one for loading the first chamber and return the others to the incubator for later use.</li>
	<li>Using a cotton swab, apply a thin, even layer of petroleum jelly surrounding the reservoirs and bridge of one modified Zigmond chamber.</li>
	<li>With a tungsten needle, gently remove the NT from the CS, while leaving the surrounding NCCs attached to the surface of the CS. Mark the dish with a pen to remember the orientation of the straightest edge of the NCC culture.</li>
	<li>Place a few drops of preincubated medium onto the bridge. Pick up the CS with fine forceps, dab the edge of the CS against a Kimwipe to remove most of the old culture medium, then immediately place the CS on the modified Zigmond chamber so that the straight edge of the culture to be filmed is centered over the length of the bridge and roughly perpendicular to the bridge-reservoir border (Fig. 2A,B).</li>
	<li>Using an inverted microscope, move the straight NCC border to be on the side of the bridge closest to the reservoir that will contain the suspected chemotactant (Fig. 2B; for controls this will correspond to whichever reservoir is loaded second). Also, more finely align the straight edge of the culture to be perpendicular to the bridge-reservoir border.</li>
	<li>Carefully, but securely press the CS into the petroleum jelly present on the Zigmond chamber, making sure it is completely sealed on to the chamber, then place additional petroleum jelly along the edge of the CS to further ensure it will be airtight. Fine-adjust the angle of the NCC border again to correct for any movement during the sealing process.</li>
	<li>Load the reservoir that will not contain the suspected chemotactant first (Fig. 2B). Do so by loading a 1 ml syringe (25 G x 1.5 in. attached needle) with roughly 300 &mu;l preincubated medium and inject the medium into the reservoir until full (while being careful not to generate any bubbles in the reservoir). Plug the reservoir on both sides with a sufficient amount of petroleum jelly prior to loading the next reservoir.</li>
	<li>Repeat step 2.7, except this time using preincubated medium containing the candidate chemotactant. It is critical when generating a molecular gradient across the culture to always load the reservoir containing the molecule to be tested after loading the reservoir lacking the tested molecule.</li>
	<li>Place the loaded Zigmond chamber at 37&deg; C to incubate for 1 h prior to filming. Image the straightest border of the NCC culture for 3 h at 90 s intervals while incubating at roughly 37&deg; C (Fig. 2A,B). Prior to creating any movie, be sure to align the camera so that the edge of the images to be obtained are aligned with and touching the edge of the bridge that borders the last reservoir loaded (Fig. 2B, upper panel; dashed box represents ideal position for imaging). This will facilitate later software analysis by standardizing the directionality of the molecular gradient applied and the distance from the reservoir filmed in each movie produced.</li>
	<li>For controls, repeat steps 2.2-2.9 for each of the two other NCC cultures selected (in step 2.1), but fill each reservoir with an appropriate medium. For one type of control fill both reservoirs with preincubated medium not containing the molecule to be tested. For a second control treatment, prime the bridge with a few drops of preincubated medium containing the suspected chemotactant prior to mounting the CS. Then, load both reservoirs with the same medium containing the suspected chemotactant.</li>
	<li>Use ImageJ (NIH) Manual Tracking <a href="http://rsb.info.nih.gov/ij/plugins/track/track.html">(rsb.info.nih.gov/ij/plugins/track/track.html)</a> and Chemotaxis and Migration Tool v1.01 <a href="http://www.ibidi.de/applications/ap_chemo.html">(www.ibidi.de/applications/ap_chemo.html)</a> plugins to track the migration of peripheral NCCs along the straight border of the culture just imaged and to analyze various parameters of the migratory trajectories obtained (Fig. 2B-C).</li>
</ol>
3. Representative Results:
A sample of cell trajectories from a movie where many trunk NCCs were responsive to a candidate chemoattractant using the above technique is shown (Fig. 2D). Most cells in this example of a positive response displayed a net movement up the chemoattractant gradient (as shown in red). Trajectory data can be used to analyze other properties of cell migration as well.
In order to visually evaluate an applied gradient in a modified Zigmond chamber, an Alexa Fluor 488 IgM conjugate (MW ~ 900 kDa) was loaded into the second reservoir of a modified Zigmond chamber (at roughly 40 &mu;g/ml H2O). A gradient was established by 1 h and still somewhat present after 26 h, but greatly diminished by 50 h (Fig. 3). If the molecule to be tested is smaller, then the applied gradient will degrade faster than what is shown.
<img alt="Figure 1" src="/files/ftp_upload/3330/3330fig1.jpg" /> 
Figure 1. Explantation of trunk-level NTs for overnight culturing on fibronectin-coated coverslips. Because trunk NCCs delaminate from the dorsal NT located adjacent to somites 8-28, this segment of the NT is isolated by microdissection and cultured overnight on a fibronectin-coated CS to allow for emigration of NCCs from the NT explant. Isolated NTs that are between 8-15 somites long and relatively straight are best suited for overnight culturing as they tend to yield NCC cultures with longer straight borders. Regions of the neural tube that give rise to other neural crest axial levels are shown in a smaller font. s, somite.
<img alt="Figure 2" src="/files/ftp_upload/3330/3330fig2.jpg" /> 
Figure 2. Method for evaluating the migration of explanted trunk NCCs using a modified Zigmond chamber. (A) Elongated trunk NCC cultures are prepared by overnight culturing of NTs and resultant NCC cultures with at least one long, straight border are selected for experimentation. The longest straight border of a selected culture is then positioned perpendicular to the bridge-reservoir border, and therefore parallel to the vector of the future applied gradient. (B) After fine-adjusting the position of the NCC culture on the Zigmond chamber and sealing the coverslip to the chamber, the chamber is loaded. When testing chemotaxis, the reservoir that will not contain the suspected chemotactant (-) is loaded first and sealed. Then, the other reservoir is loaded with the suspected chemotactant (+) and sealed. Peripheral NCCs along the previously selected border can then be imaged and tracked using the Manual Tracking plugin for ImageJ (bottom panel). (C) Numerous migratory characteristics in response to the applied gradient can be assessed based on tracking data. For instance, a chemotaxis index can be derived by dividing a cell's displacement along the x-axis by the total distance it has migrated. (D) An example of an attractive response is shown by a cell trajectory plot initially generated by the Chemotaxis and Migration Tool plugin for ImageJ. The starting point of each trajectory is set to the origin (0,0). Note how many more cells migrate toward the chemoattractant source.The center of mass of all cells at their final position (blue cross; all cells equally weighted) is also closer to the chemoattractant source. NCCs, neural crest cells; Red tracks, cells that migrated toward the reservoir loaded with a suspected chemoattractant; Black tracks, cells that migrated away; (+), higher chemotactant concentration; (-), lower chemotactant concentration.
<img alt="Figure 3" src="/files/ftp_upload/3330/3330fig3.jpg" /> 
Figure 3. Intensity profiles across the bridge of a modified Zigmond chamber at different times following the addition of an Alexa Fluor 488 IgM conjugate. The chamber was loaded in a similar manner to that described in the protocol with the main exceptions that preincubated water (instead of preincubated medium) was used to dilute the antibody to 40 &mu;g/ml, and small air pockets were present at the ends of the bridge (away from the slice where the above intensity profiles where taken). Initially, no gradient was present across most of the bridge. By 1 h a gradient was established and remained present through to 26 h. By 50 h the presence of the gradient was inconsistent at different areas of the bridge, and when present, the steepness of the gradient was greatly diminished. All profiles were generated from an identical slice across the bridge (from one bridge-reservoir border to the other) using AxioVision 4.6 software. Note that even while air pockets were present, the gradient was not disrupted. High, high intensity; Low, low intensity; x-axis, distance across entire width of bridge (2 mm); (+), reservoir loaded with the Alexa Fluor 488 IgM conjugate; (-), reservoir not loaded with the conjugate.
<img alt="Figure 4" src="/files/ftp_upload/3330/3330fig4.jpg" /> 
Figure 4. Modified Zigmond chamber specifications. Shown is a diagram of the modified Zigmond chamber used here along with its dimensional specifications (&plusmn;0.2 mm). Measurements can be moderately adjusted in order to match individual preferences.
Supplemental Protocol: Fabrication of a Modified Zigmond Chamber
Please refer to Figure 4 as a reference for the protocol below:
<ol>
	<li>Purchase a sheet of 3/16&Prime; thick polished acrylic (4.45 mm actual thickness).</li>
	<li>Using a table saw, cut chamber blanks oversized to the rough dimensions of 33.25 mm x 64.57 mm. This allows 3.175 mm extra material for machining.</li>
	<li>Set the chamber blank on a vise. With a milling machine and a 6.35 mm (1/4&Prime;) end mill bit, finish machining the sides of the chamber to their exact dimensions: 30.07 mm x 61.39 mm.</li>
	<li>Position the chamber blank on the milling machine and locate the center of the blank along both the x and y axes with an edge finder; then zero the center location.</li>
	<li>Acquire the chamber height (z-axis) by touching the end mill bit to the top surface and zero the height.</li>
	<li>Using a 3.91 mm (0.154&Prime;) end mill bit, offset the bit 3.03 mm along the x-axis (positive direction) for the first reservoir. Begin machining into the chamber to a depth of 2.84 mm while moving along the y-axis (positive direction) to 7.62 mm (0.300&Prime;) and then traverse to 7.62 mm (0.300&Prime;) in the opposite (negative) direction to a complete reservoir length of 15.24 mm (0.600&Prime;). Offset the bit to 3.03 mm (0.119&Prime;) along the x-axis (negative direction) and repeat the same process for the second reservoir.</li>
	<li>Position the chamber on its edge and drill a hole using a 1.09 mm (0.043 in.) drill bit on the end of each reservoir (4 total) that connects the end of the reservoir to the side of the chamber for loading medium during experimentation.</li>
	<li>Soak the chamber well in warm soapy water to help remove any chemical contaminants.</li>
	<li>Soak and rinse the chamber well in double-distilled water to remove any soap. The chambers are now ready to use as described above.</li>
</ol>

<ol>
	<li>Le Douarin, N. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+avian+embryo+as+a+model+to+study+the+development+of+the+neural+crest:+a+long+and+still+ongoing+story.">The avian embryo as a model to study the development of the neural crest: a long and still ongoing story.</a> Mechanisms of Development. 121, 1089-1102 (2004).</li><li>Baker, C. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Neural Crest and Cranial Ectodermal Placodes. , (2005).</li><li>Gammill, L. S., Roffers-Agarwal, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Division+of+labor+during+trunk+neural+crest+development.">Division of labor during trunk neural crest development.</a> Dev. Biol. 344, 555-565 (2010).</li><li>Kulesa, P. M., Gammill, L. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neural+crest+migration:+patterns,+phases+and+signals.">Neural crest migration: patterns, phases and signals.</a> Dev. Biol. 344, 566-568 (2010).</li><li>Wang, H. U., Anderson, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Eph+family+transmembrane+ligands+can+mediate+repulsive+guidance+of+trunk+neural+crest+migration+and+motor+axon+outgrowth.">Eph family transmembrane ligands can mediate repulsive guidance of trunk neural crest migration and motor axon outgrowth.</a> Neuron. 18, 383-396 (1997).</li><li>Krull, C. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Interactions+of+Eph-related+receptors+and+ligands+confer+rostrocaudal+pattern+to+trunk+neural+crest+migration.">Interactions of Eph-related receptors and ligands confer rostrocaudal pattern to trunk neural crest migration.</a> Curr. Biol. 7, 571-580 (1997).</li><li>Gammill, L. S., Gonzalez, C., Gu, C., Bronner-Fraser, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Guidance+of+trunk+neural+crest+migration+requires+neuropilin+2/semaphorin+3F+signaling.">Guidance of trunk neural crest migration requires neuropilin 2/semaphorin 3F signaling.</a> Development, Cambridge, England. , 133-199 (2006).</li><li>De Bellard, M. E., Rao, Y., Bronner-Fraser, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dual+function+of+Slit2+in+repulsion+and+enhanced+migration+of+trunk,+but+not+vagal,+neural+crest+cells.">Dual function of Slit2 in repulsion and enhanced migration of trunk, but not vagal, neural crest cells.</a> The Journal of cell biology. 162, 269-279 (2003).</li><li>Kasemeier-Kulesa, J. C., McLennan, R., Romine, M. H., Kulesa, P. M., Lefcort, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CXCR4+controls+ventral+migration+of+sympathetic+precursor+cells.">CXCR4 controls ventral migration of sympathetic precursor cells.</a> J. Neurosci. 30, 13078-13088 (2010).</li><li>Zigmond, S. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ability+of+polymorphonuclear+leukocytes+to+orient+in+gradients+of+chemotactic+factors.">Ability of polymorphonuclear leukocytes to orient in gradients of chemotactic factors.</a> The Journal of Cell Biology. 75, 606-616 (1977).</li><li>Hamburger, V., Hamilton, H. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+series+of+normal+stages+in+the+development+of+the+chicken+embryo.">A series of normal stages in the development of the chicken embryo.</a> J. Morph. 88, 49-52 (1951).</li><li>Boyden, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+chemotactic+effect+of+mixtures+of+antibody+and+antigen+on+polymorphonuclear+leucocytes.">The chemotactic effect of mixtures of antibody and antigen on polymorphonuclear leucocytes.</a> The Journal of Experimental Medicine. 115, 453-466 (1962).</li><li>Davis, E. M., Trinkaus, J. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Significance+of+cell-to+cell+contacts+for+the+directional+movement+of+neural+crest+cells+within+a+hydrated+collagen+lattice.">Significance of cell-to cell contacts for the directional movement of neural crest cells within a hydrated collagen lattice.</a> Journal of Embryology and Experimental Morphology. 63, 29-51 (1981).</li><li>Zicha, D., Dunn, G. A., Brown, A. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+new+direct-viewing+chemotaxis+chamber.">A new direct-viewing chemotaxis chamber.</a> Journal of Cell Science. 99, 769-775 (1991).</li></ol>

No conflicts of interest declared.

Conducting chemotaxis research on trunk NCCs has proven challenging for a series of reasons. Trunk NCCs constitute a heterogeneous stem cell population that will differentiate if cultured long-term; therefore, trunk NCCs must be obtained from primary explantation of the trunk-level NT. Conventional methods to study the chemotactic response of homogenously distributed cell populations in vitro are difficult to test on trunk NCCs since they first require that cells are isolated and homogenously replated in ...

<table border="1">
	<tbody>
		<tr>
			<td>Name of the reagent</td>
			<td>Company</td>
			<td>Catalogue number</td>
			<td>Comments (optional)</td>
		</tr>
		<tr>
			<td>DMEM</td>
			<td>Omega Scientific</td>
			<td>DM-22</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Penicillin Streptomycin Solution</td>
			<td>Omega Scientific</td>
			<td>PS-20</td>
			<td>100X Stock Concentration</td>
		</tr>
		<tr>
			<td>L-Glutamine</td>
			<td>Omega Scientific</td>
			<td>GS-60</td>
			<td>100X Stock Concentration</td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum</td>
			<td>Omega Scientific</td>
			<td>FB-11</td>
			<td>Lot# 105247 (or another that is comparable)</td>
		</tr>
		<tr>
			<td>Modified Zigmond chamber</td>
			<td>Home made</td>
			<td>N/A</td>
			<td>Reservoir volume: ~ 160 &mu;l ea; for additional specifications, see Fig. 4 and the supplemental fabrication protocol</td>
		</tr>
		<tr>
			<td>Cell culture dish</td>
			<td>Denville</td>
			<td>T6040</td>
			<td>40 x 10 mm</td>
		</tr>
		<tr>
			<td>Fibronectin</td>
			<td>BD</td>
			<td>354008</td>
			<td>10X Stock prepped by diluting 1 mg FN in 1 ml H2O and 9 ml DMEM</td>
		</tr>
		<tr>
			<td>Coverslips</td>
			<td>Fisher</td>
			<td>12-548-B</td>
			<td>Precleaned; 22 x 22 mm</td>
		</tr>
		<tr>
			<td>L15 medium</td>
			<td>Thermo Scientific</td>
			<td>SH30525.02</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Petroleum Jelly</td>
			<td>Comforts</td>
			<td>011110794642</td>
			<td>100%</td>
		</tr>
		<tr>
			<td>Centrifuge tube</td>
			<td>Biologix</td>
			<td>10-9152</td>
			<td>15 ml</td>
		</tr>
		<tr>
			<td>Dispase</td>
			<td>Cell Systems</td>
			<td>4Z0-850</td>
			<td>10X Stock Concentration</td>
		</tr>
		<tr>
			<td>Syringe</td>
			<td>BD</td>
			<td>309602</td>
			<td>1 ml</td>
		</tr>
		<tr>
			<td>Needle</td>
			<td>BD</td>
			<td>305127</td>
			<td>25 G x 1.5 in.</td>
		</tr>
		<tr>
			<td>Alexa Fluor 488-IgM</td>
			<td>Invitrogen</td>
			<td>A21042</td>
			<td>Stock is 2 mg/ml; 7 moles dye/mole IgM</td>
		</tr>
		<tr>
			<td>Dissecting Forceps</td>
			<td>FST</td>
			<td>Misc.</td>
			<td>Dumont #5 or 55; straight tipped; stainless steel or titanium</td>
		</tr>
		<tr>
			<td>Tungsten Needle</td>
			<td>N/A</td>
			<td>N/A</td>
			<td>Home made; placed in a pin holder</td>
		</tr>
		<tr>
			<td>Blunt Forceps</td>
			<td>Tiemann</td>
			<td>160-18</td>
			<td>Used for transferring embryos to Ringer's from egg yolk</td>
		</tr>
	</tbody>
</table>
Supplemental Protocol: Fabrication of a Modified Zigmond Chamber
Please refer to Figure 4 as a reference for the protocol below:
<ol>
	<li>Purchase a sheet of 3/16" thick polished acrylic (4.45 mm actual thickness).</li>
	<li>Using a table saw, cut chamber blanks oversized to the rough dimensions of 33.25 mm x 64.57 mm. This allows 3.175 mm extra material for machining.</li>
	<li>Set the chamber blank on a vise. With a milling machine and a 6.35 mm (1/4") end mill bit, finish machining the sides of the chamber to their exact dimensions: 30.07 mm x 61.39 mm.</li>
	<li>Position the chamber blank on the milling machine and locate the center of the blank along both the x and y axes with an edge finder; then zero the center location.</li>
	<li>Acquire the chamber height (z-axis) by touching the end mill bit to the top surface and zero the height.</li>
	<li>Using a 3.91 mm (0.154") end mill bit, offset the bit 3.03 mm along the x-axis (positive direction) for the first reservoir. Begin machining into the chamber to a depth of 2.84 mm while moving along the y-axis (positive direction) to 7.62 mm (0.300") and then traverse to 7.62 mm (0.300") in the opposite (negative) direction to a complete reservoir length of 15.24 mm (0.600"). Offset the bit to 3.03 mm (0.119") along the x-axis (negative direction) and repeat the same process for the second reservoir.</li>
	<li>Position the chamber on its edge and drill a hole using a 1.09 mm (0.043 in.) drill bit on the end of each reservoir (4 total) that connects the end of the reservoir to the side of the chamber for loading medium during experimentation.</li>
	<li>Soak the chamber well in warm soapy water to help remove any chemical contaminants.</li>
	<li>Soak and rinse the chamber well in double-distilled water to remove any soap. The chambers are now ready to use as described above.</li>
</ol>

analysis of trunk neural crest cell migration using a modified zigmond chamber assay

Neural crest cells (NCCs) are a transient population of cells present in vertebrate development that emigrate from the dorsal neural tube (NT) after undergoing an epithelial-mesenchymal transition 1,2. Following EMT, NCCs migrate large distances along stereotypic pathways until they reach their targets. NCCs differentiate into a vast array of cell types including neurons, glia, melanocytes, and chromaffin cells 1-3. The ability of NCCs to reach and recognize their proper target locations is foundational for the appropriate formation of all structures containing trunk NCC-derived components 3. Elucidating the mechanisms of guidance for trunk NCC migration has therefore been a matter of great significance. Numerous molecules have been demonstrated to guide NCC migration 4. For instance, trunk NCCs are known to be repelled by negative guidance cues such as Semaphorin, Ephrin, and Slit ligands 5-8. However, not until recently have any chemoattractants of trunk NCCs been identified 9. 
Conventional in vitro approaches to studying the chemotactic behavior of adherent cells work best with immortalized, homogenously distributed cells, but are more challenging to apply to certain primary stem cell cultures that initially lack a homogenous distribution and rapidly differentiate (such as NCCs). One approach to homogenize the distribution of trunk NCCs for chemotaxis studies is to isolate trunk NCCs from primary NT explant cultures, then lift and replate them to be almost 100% confluent. However, this plating approach requires substantial amounts of time and effort to explant enough cells, is harsh, and distributes trunk NCCs in a dissimilar manner to that found in in vivo conditions. 
Here, we report an in vitro approach that is able to evaluate chemotaxis and other migratory responses of trunk NCCs without requiring a homogenous cell distribution. This technique utilizes time-lapse imaging of primary, unperturbed trunk NCCs inside a modified Zigmond chamber (a standard Zigmond chamber is described elsewhere10). By exposing trunk NCCs at the periphery of the culture to a chemotactant gradient that is perpendicular to their predicted natural directionality, alterations in migratory polarity induced by the applied chemotactant gradient can be detected. This technique is inexpensive, requires the culturing of only two NT explants per replicate treatment, avoids harsh cell lifting (such as trypsinization), leaves trunk NCCs in a more similar distribution to in vivo conditions, cuts down the amount of time between explantation and experimentation (which likely reduces the risk of differentiation), and allows time-lapse evaluation of numerous migratory characteristics.

Watch this Scientific Journal Video about Analysis of Trunk Neural Crest Cell Migration using a Modified Zigmond Chamber Assay at JoVE.com

Analysis of Trunk Neural Crest Cell Migration using a Modified Zigmond Chamber Assay

An approach to analyze the migration of explanted cells (trunk neural crest cells) is described. This method is inexpensive, gentle, and capable of distinguishing chemotaxis from both chemokinesis and other influences on migratory polarity such as those derived from cell-cell interactions within the primary trunk neural crest cell culture.

Neural crest cells (NCCs) are a transient population of cells present in vertebrate development that emigrate from the dorsal neural tube (NT) after ...

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12.4K Views.  California State University, Northridge. An approach to analyze the migration of explanted cells (trunk neural crest cells) is described. This method is inexpensive, gentle, and capable of distinguishing chemotaxis from both chemokinesis and other influences on migratory polarity such as those derived from cell-cell interactions within the primary trunk neural crest cell culture.

Video: Analysis of Trunk Neural Crest Cell Migration using a Modified Zigmond Chamber Assay

Isolation and Expansion of the Adult Mouse Neural Stem Cells Using the Neurosphere Assay

Isolation and expansion of the putative neural stem cells (NSCs) from the adult murine brain was first described by Reynolds and Weiss in 1992 employing a chemically defined serum-free culture system known as the neurosphere assay (NSA). In this assay, the majority of differentiated cell types die within a few days of culture but a small population of growth factor responsive precursor cells undergo active proliferation in the presence of epidermal growth factor (EGF) and/ basic fibroblastic growth factor (bFGF). These cells form colonies of undifferentiated cells called neurospheres, which in turn can be subcultured to expand the pool of neural stem cells. Moreover, the cells can be induced to differentiate, generating the three major cell types of the CNS i.e. neurons, astrocytes, and oligodendrocytes. This assay provides an invaluable tool to supply a consistent, renewable source of undifferentiated CNS precursors, which could be used for in vitro studies and also for therapeutic purposes.
This video demonstrates the NSA method to generate and expand NSCs from the adult mouse periventricular region, and provides technical insights to ensure one can achieve reproducible neurosphere cultures. The procedure includes harvesting the brain from the adult mouse, micro-dissection of the periventricular region, tissue preparation and culture in the NSA. The harvested tissue is first chemically digested using trypsin-EDTA and then mechanically dissociated in NSC medium to achieve a single cell suspension and finally plated in the NSA. After 7-10 days in culture, the resulting primary neurospheres are ready for subculture to reach the amount of cells required for future experiments.

This video protocol demonstrates the neurosphere assay method to generate and expand neural stem cells from the adult mouse periventricular region, and provides technical insights to ensure one can achieve reproducible neurosphere cultures.

Isolation and expansion of the putative neural stem cells (NSCs) from the adult murine brain was first described by Reynolds and Weiss in 1992 employing a ...

Establishing Embryonic Mouse Neural Stem Cell Culture Using the Neurosphere Assay

In mammalians, stem cells acts as a source of undifferentiated cells to maintain cell genesis and renewal in different tissues and organs during the life span of the animal. They can potentially replace cells that are lost in the aging process or in the process of injury and disease. The existence of neural stem cells (NSCs) was first described by Reynolds and Weiss (1992) in the adult mammalian central nervous system (CNS) using a novel serum&#8208;free culture system, the neurosphere assay (NSA). Using this assay, it is also feasible to isolate and expand NSCs from different regions of the embryonic CNS. These in vitro expanded NSCs are multipotent and can give rise to the three major cell types of the CNS. While the NSA seems relatively simple to perform, attention to the procedures demonstrated here is required in order to achieve reliable and consistent results. This video practically demonstrates NSA to generate and expand NSCs from embryonic day 14-mouse brain tissue and provides technical details so one can achieve reproducible neurosphere cultures. The procedure includes harvesting E14 mouse embryos, brain microdissection to harvest the ganglionic eminences, dissociation of the harvested tissue in NSC medium to gain a single cell suspension, and finally plating cells in NSA culture. After 5-7 days in culture, the resulting primary neurospheres are passaged to further expand the number of the NSCs for future experiments.

This video protocol demonstrates the application of the neurosphere assay for the isolation and expansion of neural stem cells from the ganglionic eminences of embryonic day 14-mouse brain.

In mammalians, stem cells acts as a source of undifferentiated cells to maintain cell genesis and renewal in different tissues and organs during the life ...

Neural-Colony Forming Cell Assay: An Assay To Discriminate Bona Fide Neural Stem Cells from Neural Progenitor Cells

The neurosphere assay (NSA) is one of the most frequently used methods to isolate, expand and also calculate the frequency of neural stem cells (NSCs). Furthermore, this serum-free culture system has also been employed to expand stem cells and determine their frequency from a variety of tumors and normal tissues. It has been shown recently that a one-to-one relationship does not exist between neurosphere formation and NSCs. This suggests that the NSA as currently applied, overestimates the frequency of NSCs in a mixed population of neural precursor cells isolated from both the embryonic and adult mammalian brain. This video practically demonstrates a novel collagen based semi- solid assay, the neural-colony forming cell assay (N-CFCA), which has the ability to discriminate stem from progenitor cells based on their long-term proliferative potential, and thus provides a method to enumerate NSC frequency. In the N-CFCA, colonies &ge;2 mm in diameter are derived from cells that meet all the functional criteria of a NSC, while colonies &lt; 2mm are derived from progenitors. The N-CFCA procedure can be used for cells prepared from different sources including primary and cultured adult or embryonic mouse CNS cells. Here we use cells prepared from passage one neurospheres generated from embryonic day 14 mice brain to perform N-CFCA. The cultures are replenished with proliferation medium every seven days for three weeks to allow the plated cells to exhibit their full proliferative potential and then the frequency of neural progenitor and bona fide neural stem cells is calculated respectively by counting the number of colonies that are &lt; 2mm and the ones that are &ge;2mm in reference to the number of cells that were initially plated.

This video protocol demonstrates how to discriminate and enumerate bona fide neural stem cells in a mixed population of neural precursor cells using the neural colony-forming cell assay.

The neurosphere assay (NSA) is one of the most frequently used methods to isolate, expand and also calculate the frequency of neural stem cells (NSCs). ...

Isolation and Culture of Neural Crest Stem Cells from Human Hair Follicles

Hair follicles undergo lifelong growth and hair cycle is a well-controlled process involving stem cell proliferation and quiescence. Hair bulge is a well-characterized niche for adult stem cells1. This segment of the outer root sheath contains a number of different types of stem cells, including epithelial stem cells2, melanocyte stem cells3 and neural crest like stem cells4-7. Hair follicles represent an accessible and rich source for different types of human stem cells. We and others have isolated neural crest stem cells (NCSCs) from human fetal and adult hair follicles4,5. These human stem cells are label-retaining cells and are capable of self-renewal through asymmetric cell division in vitro. They express immature neural crest cell markers but not differentiation markers. Our expression profiling study showed that they share a similar gene expression pattern with murine skin immature neural crest cells. They exhibit clonal multipotency that can give rise to myogenic, melanocytic, and neuronal cell lineages after in vitro clonal single cell culture. Differentiated cells not only acquire lineage-specific markers but also demonstrate appropriate functions in ex vivo conditions. In addition, these NCSCs show differentiation potential toward mesenchymal lineages. Differentiated neuronal cells can persist in mouse brain and retain neuronal differentiation markers. It has been shown that hair follicle derived NCSCs can help nerve regrowth, and they improve motor function in mice transplanted with these stem cells following transecting spinal cord injury8. Furthermore, peripheral nerves have been repaired with stem cell grafts9, and implantation of skin-derived precursor cells adjacent to crushed sciatic nerves has resulted in remyelination10. Therefore, the hair follicle/skin derived NCSCs have already shown promising results for regenerative therapy in preclinical models.
Somatic cell reprogramming to induced pluripotent stem (iPS) cells has shown enormous potential for regenerative medicine. However, there are still many issues with iPS cells, particularly the long term effect of oncogene/virus integration and potential tumorigenicity of pluripotent stem cells have not been adequately addressed. There are still many hurdles to be overcome before iPS cells can be used for regenerative medicine. Whereas the adult stem cells are known to be safe and they have been used clinically for many years, such as bone marrow transplant. Many patients have already benefited from the treatment. Autologous adult stem cells are still preferred cells for transplantation. Therefore, the readily accessible and expandable adult stem cells in human skin/hair follicles are a valuable source for regenerative medicine.

This article presents a robust protocol for isolation and culture of neural crest stem cells from human hair follicles.

Hair follicles undergo lifelong growth and hair cycle is a well-controlled process involving stem cell proliferation and quiescence. Hair bulge is a ...

Reproducible Mouse Sciatic Nerve Crush and Subsequent Assessment of Regeneration by Whole Mount Muscle Analysis

Regeneration in the peripheral nervous system (PNS) is widely studied both for its relevance to human disease and to understand the robust regenerative response mounted by PNS neurons thereby possibly illuminating the failures of CNS regeneration1. Sciatic nerve crush (axonotmesis) is one of the most common models of peripheral nerve injury in rodents2. Crushing interrupts all axons but Schwann cell basal laminae are preserved so that regeneration is optimal3,4. This allows the investigator to study precisely the ability of a growing axon to interact with both the Schwann cell and basal laminae4. Rats have generally been the preferred animal models for experimental nerve crush. They are widely available and their lesioned sciatic nerve provides a reasonable approximation of human nerve lesions5,4. Though smaller in size than rat nerve, the mouse nerve has many similar qualities. Most importantly though, mouse models are increasingly valuable because of the wide availability of transgenic lines now allows for a detailed dissection of the individual molecules critical for nerve regeneration6, 7. Prior investigators have used multiple methods to produce a nerve crush or injury including simple angled forceps, chilled forceps, hemostatic forceps, vascular clamps, and investigator-designed clamps8,9,10,11,12. Investigators have also used various methods of marking the injury site including suture, carbon particles and fluorescent beads13,14,1. We describe our method to obtain a reproducibly complete sciatic nerve crush with accurate and persistent marking of the crush-site using a fine hemostatic forceps and subsequent carbon crush-site marking. As part of our description of the sciatic nerve crush procedure we have also included a relatively simple method of muscle whole mount we use to subsequently quantify regeneration.

In this report we describe a method to crush mouse sciatic nerve. This method uses readily available hemostatic forceps and easily and reproducibly produces complete sciatic nerve crush. In addition, we describe a method to prepare muscle whole mounts suitable for analysis of nerve regeneration after sciatic nerve crush.

Regeneration in the peripheral nervous system (PNS) is widely studied both for its relevance to human disease and to understand the robust regenerative ...

Analysis of Neural Crest Migration and Differentiation by Cross-species Transplantation

Avian embryos provide a unique platform for studying many vertebrate developmental processes, due to the easy access of the embryos within the egg. Chimeric avian embryos, in which quail donor tissue is transplanted into a chick embryo in ovo, combine the power of indelible genetic labeling of cell populations with the ease of manipulation presented by the avian embryo.
Quail-chick chimeras are a classical tool for tracing migratory neural crest cells (NCCs)1-3. NCCs are a transient migratory population of cells in the embryo, which originate in the dorsal region of the developing neural tube4. They undergo an epithelial to mesenchymal transition and subsequently migrate to other regions of the embryo, where they differentiate into various cell types including cartilage5-13, melanocytes11,14-20, neurons and glia21-32. NCCs are multipotent, and their ultimate fate is influenced by 1) the region of the neural tube in which they originate along the rostro-caudal axis of the embryo11,33-37, 2) signals from neighboring cells as they migrate38-44, and 3) the microenvironment of their ultimate destination within the embryo45,46. Tracing these cells from their point of origin at the neural tube, to their final position and fate within the embryo, provides important insight into the developmental processes that regulate patterning and organogenesis.
Transplantation of complementary regions of donor neural tube (homotopic grafting) or different regions of donor neural tube (heterotopic grafting) can reveal differences in pre-specification of NCCs along the rostro-caudal axis2,47. This technique can be further adapted to transplant a unilateral compartment of the neural tube, such that one side is derived from donor tissue, and the contralateral side remains unperturbed in the host embryo, yielding an internal control within the same sample2,47. It can also be adapted for transplantation of brain segments in later embryos, after HH10, when the anterior neural tube has closed47.
Here we report techniques for generating quail-chick chimeras via neural tube transplantation, which allow for tracing of migratory NCCs derived from a discrete segment of the neural tube. Species-specific labeling of the donor-derived cells with the quail-specific QCPN antibody48-56 allows the researcher to distinguish donor and host cells at the experimental end point. This technique is straightforward, inexpensive, and has many applications, including fate-mapping, cell lineage tracing, and identifying pre-patterning events along the rostro-caudal axis45. Because of the ease of access to the avian embryo, the quail-chick graft technique may be combined with other manipulations, including but not limited to lens ablation40, injection of inhibitory molecules57,58, or genetic manipulation via electroporation of expression plasmids59-61, to identify the response of particular migratory streams of NCCs to perturbations in the embryo's developmental program. Furthermore, this grafting technique may also be used to generate other interspecific chimeric embryos such as quail-duck chimeras to study NCC contribution to craniofacial morphogenesis, or mouse-chick chimeras to combine the power of mouse genetics with the ease of manipulation of the avian embryo.62

An approach for analyzing migration and eventual fate of avian neural crest cells in quail-chick chimeric embryos is described. This method is a simple and straightforward technique for tracing neural crest cells during migration and differentiation that are otherwise difficult to distinguish within an unmanipulated chick embryo.

Avian embryos provide a unique platform for studying many vertebrate developmental processes, due to the easy access of the embryos within the egg. ...

The Neuroblast Assay: An Assay for the Generation and Enrichment of Neuronal Progenitor Cells from Differentiating Neural Stem Cell Progeny Using Flow Cytometry

Neural stem cells (NSCs) can be isolated and expanded in large-scale, using the neurosphere assay and differentiated into the three major cell types of the central nervous system (CNS); namely, astrocytes, oligodendrocytes and neurons. These characteristics make neural stem and progenitor cells an invaluable renewable source of cells for in vitro studies such as drug screening, neurotoxicology and electrophysiology and also for cell replacement therapy in many neurological diseases. In practice, however, heterogeneity of NSC progeny, low production of neurons and oligodendrocytes, and predominance of astrocytes following differentiation limit their clinical applications. Here, we describe a novel methodology for the generation and subsequent purification of immature neurons from murine NSC progeny using fluorescence activated cell sorting (FACS) technology. Using this methodology, a highly enriched neuronal progenitor cell population can be achieved without any noticeable astrocyte and bona fide NSC contamination. The procedure includes differentiation of NSC progeny isolated and expanded from E14 mouse ganglionic eminences using the neurosphere assay, followed by isolation and enrichment of immature neuronal cells based on their physical (size and internal complexity) and fluorescent properties using flow cytometry technology. Overall, it takes 5-7 days to generate neurospheres and 6-8 days to differentiate NSC progeny and isolate highly purified immature neuronal cells.

This video protocol demonstrates a novel method for the generation and subsequent purification of neuronal progenitor cells from a renewable source of neural stem cells (NSCs) based on their physical (size and internal granularity) and fluorescent properties using flow cytometry technology.

Neural stem cells (NSCs) can be isolated and expanded in large-scale, using the neurosphere assay and differentiated into the three major cell types of ...

Co-analysis of Brain Structure and Function using fMRI and Diffusion-weighted Imaging

The study of complex computational systems is facilitated by network maps, such as circuit diagrams. Such mapping is particularly informative when studying the brain, as the functional role that a brain area fulfills may be largely defined by its connections to other brain areas. In this report, we describe a novel, non-invasive approach for relating brain structure and function using magnetic resonance imaging (MRI). This approach, a combination of structural imaging of long-range fiber connections and functional imaging data, is illustrated in two distinct cognitive domains, visual attention and face perception. Structural imaging is performed with diffusion-weighted imaging (DWI) and fiber tractography, which track the diffusion of water molecules along white-matter fiber tracts in the brain (Figure 1). By visualizing these fiber tracts, we are able to investigate the long-range connective architecture of the brain. The results compare favorably with one of the most widely-used techniques in DWI, diffusion tensor imaging (DTI). DTI is unable to resolve complex configurations of fiber tracts, limiting its utility for constructing detailed, anatomically-informed models of brain function. In contrast, our analyses reproduce known neuroanatomy with precision and accuracy. This advantage is partly due to data acquisition procedures: while many DTI protocols measure diffusion in a small number of directions (e.g., 6 or 12), we employ a diffusion spectrum imaging (DSI)1, 2 protocol which assesses diffusion in 257 directions and at a range of magnetic gradient strengths. Moreover, DSI data allow us to use more sophisticated methods for reconstructing acquired data. In two experiments (visual attention and face perception), tractography reveals that co-active areas of the human brain are anatomically connected, supporting extant hypotheses that they form functional networks. DWI allows us to create a "circuit diagram" and reproduce it on an individual-subject basis, for the purpose of monitoring task-relevant brain activity in networks of interest.

We describe a novel approach for simultaneous analysis of brain function and structure using magnetic resonance imaging (MRI). We assess brain structure with high-resolution diffusion-weighted imaging and white-matter fiber tractography. Unlike standard structural MRI, these techniques allow us to directly relate anatomical connectivity to functional properties of brain networks.

The study of complex computational systems is facilitated by network maps, such as circuit diagrams. Such mapping is particularly informative when ...

Isolation and Culture of Neural Crest Cells from Embryonic Murine Neural Tube

The embryonic neural crest (NC) is a multipotent progenitor population that originates at the dorsal aspect of the neural tube, undergoes an epithelial to mesenchymal transition (EMT) and migrates throughout the embryo, giving rise to diverse cell types 1-3. NC also has the unique ability to influence the differentiation and maturation of target organs4-6. When explanted in vitro, NC progenitors undergo self-renewal, migrate and differentiate into a variety of tissue types including neurons, glia, smooth muscle cells, cartilage and bone. 
NC multipotency was first described from explants of the avian neural tube7-9. In vitro isolation of NC cells facilitates the study of NC dynamics including proliferation, migration, and multipotency. Further work in the avian and rat systems demonstrated that explanted NC cells retain their NC potential when transplanted back into the embryo10-13. Because these inherent cellular properties are preserved in explanted NC progenitors, the neural tube explant assay provides an attractive option for studying the NC in vitro. 
To attain a better understanding of the mammalian NC, many methods have been employed to isolate NC populations. NC-derived progenitors can be cultured from post-migratory locations in both the embryo and adult to study the dynamics of post-migratory NC progenitors11,14-20, however isolation of NC progenitors as they emigrate from the neural tube provides optimal preservation of NC cell potential and migratory properties13,21,22. Some protocols employ fluorescence activated cell sorting (FACS) to isolate a NC population enriched for particular progenitors11,13,14,17. However, when starting with early stage embryos, cell numbers adequate for analyses are difficult to obtain with FACS, complicating the isolation of early NC populations from individual embryos. Here, we describe an approach that does not rely on FACS and results in an approximately 96% pure NC population based on a Wnt1-Cre activated lineage reporter23. 
The method presented here is adapted from protocols optimized for the culture of rat NC11,13. The advantages of this protocol compared to previous methods are that 1) the cells are not grown on a feeder layer, 2) FACS is not required to obtain a relatively pure NC population, 3) premigratory NC cells are isolated and 4) results are easily quantified. Furthermore, this protocol can be used for isolation of NC from any mutant mouse model, facilitating the study of NC characteristics with different genetic manipulations. The limitation of this approach is that the NC is removed from the context of the embryo, which is known to influence the survival, migration and differentiation of the NC2,24-28.

Isolation of embryonic neural crest from the neural tube facilitates the use of in vitro methods for studying migration, self-renewal, and multipotency of neural crest.

The embryonic neural crest (NC) is a multipotent progenitor population that originates at the dorsal aspect of the neural tube, undergoes an epithelial to ...

A Galvanotaxis Assay for Analysis of Neural Precursor Cell Migration Kinetics in an Externally Applied Direct Current Electric Field

The discovery of neural stem and progenitor cells (collectively termed neural precursor cells) (NPCs) in the adult mammalian brain has led to a body of research aimed at utilizing the multipotent and proliferative properties of these cells for the development of neuroregenerative strategies. A critical step for the success of such strategies is the mobilization of NPCs toward a lesion site following exogenous transplantation or to enhance the response of the endogenous precursors that are found in the periventricular region of the CNS. Accordingly, it is essential to understand the mechanisms that promote, guide, and enhance NPC migration. Our work focuses on the utilization of direct current electric fields (dcEFs) to promote and direct NPC migration - a phenomenon known as galvanotaxis. Endogenous physiological electric fields function as critical cues for cell migration during normal development and wound repair. Pharmacological disruption of the trans-neural tube potential in axolotl embryos causes severe developmental malformations1. In the context of wound healing, the rate of repair of wounded cornea is directly correlated with the magnitude of the epithelial wound potential that arises after injury, as shown by pharmacological enhancement or disruption of this dcEF2-3. We have demonstrated that adult subependymal NPCs undergo rapid and directed cathodal migration in vitro when exposed to an externally applied dcEF. In this protocol we describe our lab's techniques for creating a simple and effective galvanotaxis assay for high-resolution, long-term observation of directed cell body translocation (migration) on a single-cell level. This assay would be suitable for investigating the mechanisms that regulate dcEF transduction into cellular motility through the use of transgenic or knockout mice, short interfering RNA, or specific receptor agonists/antagonists.

In this protocol we demonstrate how to construct custom chambers that permit the application of a direct current electric field to enable time-lapse imaging of adult brain derived neural precursor cell translocation during galvanotaxis.

The discovery of neural stem and progenitor  cells (collectively termed neural precursor cells) (NPCs) in the adult  mammalian brain has led to a body of ...