This work was funded by National Institutes of Health Grant RO1DC002756 and the Purdue Research Foundation. We thank Doris Wu and Wiese Chang for advice with experiments and Rodney McPhail for help with figures.

Recipes
Chick Ringers
<table border="1">
 <tbody>
 <tr>
 <td>7.2 g</td>
 <td>NaCl</td>
 </tr>
 <tr>
 <td>0.23 g</td>
 <td>CaCl2 + 2H2O</td>
 </tr>
 <tr>
 <td>0.37 g</td>
 <td>KCl</td>
 </tr>
 <tr>
 <td>0.115 g</td>
 <td>Na2HPO4 </td>
 </tr>
 <tr>
 <td>900 ml</td>
 <td>Water (Tissue culture grade)</td>
 </tr>
 </tbody>
</table>
Note: Adjust pH to 7.4. Bring final volume to 1000 ml with water.
10X PBS
<table border="1">
 <tbody>
 <tr>
 <td>40 g</td>
 <td>NaCl</td>
 </tr>
 <tr>
 <td>1 g</td>
 <td>KCl</td>
 </tr>
 <tr>
 <td>7 g </td>
 <td>Na2HPO4</td>
 </tr>
 <tr>
 <td>1.2 g </td>
 <td>KH2PO4</td>
 </tr>
 <tr>
 <td>450 ml</td>
 <td>DEPC water</td>
 </tr>
 </tbody>
</table>
Note: Adjust pH to 7.4. Bring final volume to 500 ml with water. Working concentration (1X) is made by diluting 1 part 10X stock with 9 parts of tissue-culture grade water, such as that obtained with a Millipore UV irradiation system to generate water with 18 M&Omega;-cm conductivity.
SAG explant holding medium
<ul>
 <li>DMEM/F12 with L-glutamine, 10 mM Hepes</li>
 <li>10% Insulin, Transferring, Selenium (ITS+1)</li>
 <li>1% pen-strep</li>
</ul>
Explant culture medium
<ul>
 <li>SAG explant holding medium</li>
 <li>10 ng/ml Neurotrophin-3 (NT-3)</li>
 <li>10 ng/ml ciliary neurotrophic factor (CNTF)</li>
</ul>
Spinal cord dissection and holding medium
<ul>
 <li>L-15</li>
 <li>10% fetal calf serum</li>
 <li>1% pen-strep</li>
</ul>
Incubate eggs at 37-38&deg;C. Sterilize dissecting tools and Sylgard dissecting dish in 70% ethanol. Dissecting dishes and tools can also be UV sterilized overnight.
1. Bead preparation
<ol>
 <li>Place beads in a microtube with 1 ml sterile PBS, mix, and wait for the beads to settle to the bottom of the tube.</li>
 <li>Wash the beads several times by removing the supernatant after the beads have settled and resuspend in PBS.</li>
 <li>Incubate the beads in purified protein (diluted in PBS) or PBS alone (Control) for 1 hour at room temperature.</li>
 <li>Rinse the beads in PBS and store in a 24-well plate with 1ml PBS until placed in collagen.</li>
</ol>
2. Statoacoustic ganglion dissection
<ol>
 <li>On E4 remove the embryo from the egg and place it in a dish with chick Ringer's solution to remove embryonic membranes.</li>
 <li>Place the embryo in a Sylgard&copy; lined petri dish filled with cold HBSS and position the embryo on its side with the otic vesicle facing up. Pin the embryo to the dish using dissecting pins.</li>
 <li>Using two dissecting pins, make a horizontal cut through the skin immediately ventral to the canal pouches, approximately where it meets the cochlear duct. Make a vertical cut anterior and posterior to the SAG. Use forceps or a dissecting pin to lift and remove the flap of skin bordered by the three cuts.
 Note: The SAG is located at the anteroventral edge of the otocyst, about midway between the dorsal pouch which is readily visible with brightfield base illumination, and the ventral cochlear duct which projects ventromedially and is obscured by the pharyngeal arches. The cochlear duct elongates so its dorsoventral dimension will progressively increase on E4 between Hamburger-Hamilton stages 23-25 9.</li>
 <li>Pull the otocyst in a posterior direction to separate it from the SAG. Displace tissue surrounding the SAG with dissection pins and carefully remove the SAG from the embryo using #55 forceps. Remove large chunks of mesenchymal tissue and protruding nerve bundles from the SAG.</li>
 <li>Using a wide-mouth pipet tip, transfer the explant to a 24-well plate with 0.5 ml SAG explant holding medium. Keep explants on ice or at room temperature for up to 4 hours.</li>
</ol>
3. Spinal cord dissection
<ol>
 <li>Remove the E6 embryo from the egg and place it in a dish containing chick Ringer's solution. Remove the head and embryonic membranes.</li>
 <li>Place the body in a Sylgard&copy; dissecting dish containing cold spinal cord dissection medium. Position and pin the embryo "ventral down" and caudal facing the experimenter. Place dissecting pins through each limb and through the anterior end.</li>
 <li>Using forceps and/or dissecting pins, carefully remove skin and tissue, moving in a ventral direction, until the dorsal surface of the spinal cord is visible.</li>
 <li>Cut the dorsal midline of the spinal cord with a dissecting pin, in a posterior to anterior direction, along the entire length of the spinal cord. This creates an "open book" as the left and right sides of the spinal cord separate.</li>
 <li>Remove surrounding tissue, laterally, to isolate the spinal cord and expose the dorsal root ganglia (DRG). Remove the DRG and meninges by rubbing a dissecting pin or forceps between the spinal cord and DRG.</li>
 <li>Remove the spinal cord from the body by holding the caudal-most end of the spinal cord with forceps and lifting up and away from the experimenter. Remove remaining DRG and meninges.</li>
 <li>Hold one end of the the spinal cord with forceps (or pin it to the dish) and use Vannas scissors to cut along the ventral midline, to bisect the spinal cord.</li>
 <li>Hold one end of the explant with forceps to immobilize the tissue and use Vannas scissors to cut small explants (100-500 um in length) along the length of the tissue. The floor plate can be recognized as a clear thickening of tissue along one explant edge.</li>
 <li>Use a wide-mouth pipet tip to transfer explants to a 24 well plate with 0.5 ml cold dissection medium. Keep explants on ice to maintain tissue viability, for up to 4 hours.</li>
</ol>
4. Explant culture with purified proteins to test neurite responsiveness
<ol>
 <li>Prepare a 1.5 mg/ml collagen solution in a 15 ml conical tube, on ice, according to the manufacturer's instructions.</li>
 <li>Verify the collagen solution has a pH of 7-7.4 using pH indicator paper. Adjust the pH by adding NaOH in 1 &mu;l volumes.</li>
 <li>Transfer explants to a 24-well plate using a wide mouth pipet tip and aspirate excess liquid. Multiple explants can be cultured in one well, but should be placed at least 500 &mu;m apart from each other.</li>
 <li>Add 0.5 ml of collagen to the well containing the explant. Position the explant on the bottom surface of the well. Be sure to set each culture completely before proceeding with the next well because the collagen will begin to polymerize at room temperature.</li>
 <li>Place the culture plate on a 37&deg;C slide warmer for 30-45 min to polymerize the collagen. When the collagen has polymerized it will resemble a gel.</li>
 <li>Add 0.5 ml warm explant culture medium supplemented with either purified proteins (diluted in PBS) or PBS (Control) and incubate at 37&deg;C, 5% CO2. Effects on neurite outgrowth should be visible by 24 hours. NOTE: We have used the same protocol to culture SAG explants up to 42 hours.</li>
</ol>
5. Explant co-culture with protein-coated beads to test directional neurite outgrowth
<ol>
 <li>Begin with explant culture steps 4.1- 4.4.</li>
 <li>Using a pipet, transfer 1-5 beads from PBS to the well containing the collagen solution and explant.</li>
 <li>Use forceps to position beads 50-500 &mu;m from the edge of the explant.</li>
 <li>Place the plate on a 37&deg;C slide warmer for 30-45 min to polymerize the collagen. The tissue or bead may become displaced as the collagen polymerizes. Check the cultures and re-position the tissue and beads during the first 5 min.</li>
 <li>Add 0.5 ml SAG media to each well and incubate for 24 hours.</li>
</ol>
6. Visualization of neurite outgrowth by immunohistochemistry
<ol>
 <li>Rinse cultures in PBS and fix for 1 hour at room temperature in 4% paraformaldehyde in PBS.</li>
 <li>Release the gels from the walls of the wells by tracing around the edge of the gels with the rounded end of a teflon micro spatula. Perform the immunostain with the gels in the wells. Rinse several times in PBS.</li>
 <li>Incubate in 0.5 ml blocking solution (10% calf serum, 0.1% Triton X-100, 0.1% sodium azide in PBS) overnight at 4&deg;C.</li>
 <li>Incubate in 0.5 ml primary antibody (anti-&beta;-Tubulin antibody diluted 1:500 in blocking solution) overnight at 4&deg;C.</li>
 <li>Rinse several times in PBS. Incubate in 0.5 ml secondary antibody (Alexa fluor 488 goat anti-mouse IgG2b antibody diluted 1:500) overnight at 4&deg;C. From this step forward, keep plates in dark or wrap in foil, due to light sensitivity of secondary antibodies.</li>
 <li>Rinse several times in PBS and store at 4&deg;C in PBS for up to one week.</li>
</ol>
7. Representative Results
<img alt="Figure 1" src="/files/ftp_upload/3600/3600fig1.jpg" /> 
Figure 1. Representative results of the neurite-promoting effects of purified NT-3 and NT-3-coated beads on E4 chick SAG explants. SAG neuron cell bodies and neurites were immunostained with &beta;-Tubulin and imaged with a 10x objective on a confocal microscope. After 24 hours in vitro, SAG explants displayed greater neurite outgrowth in the presence of 100 ng/ml purified human NT-3 added to the cell culture medium (1B), compared to controls (1A). SAG neurons were co-cultured with beads soaked in100 ng/ml NT-3 to demonstrate that a point source of NT-3 can locally promote neurite outgrowth. Explants displayed longer and denser neurite outgrowth on the side of the explant facing the bead compared to the opposite side (1D) and compared to control cultures with beads soaked in PBS (1C). Scale bar = 200 &mu;m.
<img alt="Figure 2" src="/files/ftp_upload/3600/3600fig2.jpg" /> 
Figure 2. Representative results of the neurite-promoting effects of purified Wnt5a and Wnt5a-coated beads on E6 chick spinal cord. Spinal cord explants were immunostained with &beta;-Tubulin and imaged with a confocal microscope. After 24 hours of growth in the presence of purified mouse Wnt5a (400ng/ml), neurite outgrowth from chick spinal cord explants was increased (2B) compared to controls cultured without Wnt5a (2A). Beads soaked in 500 ng/ml Wnt5a, placed 300-500 &mu;m from the edge of spinal cord explants, also promoted neurite outgrowth (2D) compared to control cultures (2C). Similar co-culture results were previously published with Wnt5a-expressing cells8. Scale bar = 200 &mu;m.

<ol>
	<li>Fekete, D. M., Campero, A. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Axon+guidance+in+the+inner+ear.">Axon guidance in the inner ear.</a> Int. J. Dev. Biol. 51 (6-7), 549-549 (2007).</li><li>Fritzsch, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+inner+ear+afferent+connections:+forming+primary+neurons+and+connecting+them+to+the+developing+sensory+epithelia.">Development of inner ear afferent connections: forming primary neurons and connecting them to the developing sensory epithelia.</a> Brain Res. Bull. 60 (5-6), 423-423 (2003).</li><li>Gu, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neuropilin-1+conveys+semaphorin+and+VEGF+signaling+during+neural+and+cardiovascular+development.">Neuropilin-1 conveys semaphorin and VEGF signaling during neural and cardiovascular development.</a> Dev. Cell. 5 (1), 45-45 (2003).</li><li>Tessarollo, L., Coppola, V., Fritzsch, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=NT-3+replacement+with+brain-derived+neurotrophic+factor+redirects+vestibular+nerve+fibers+to+the+cochlea.">NT-3 replacement with brain-derived neurotrophic factor redirects vestibular nerve fibers to the cochlea.</a> J. Neurosci. 24 (10), 2575-2575 (2004).</li><li>Avila, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Brain-derived+neurotrophic+factor+and+neurotrophin-3+support+the+survival+and+neuritogenesis+response+of+developing+cochleovestibular+ganglion+neurons.">Brain-derived neurotrophic factor and neurotrophin-3 support the survival and neuritogenesis response of developing cochleovestibular ganglion neurons.</a> Dev. Biol. 159 (1), 266-266 (1993).</li><li>Bianchi, L. M., Cohan, C. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+the+neurotrophins+and+CNTF+on+developing+statoacoustic+neurons:+comparison+with+an+otocyst-derived+factor.">Effects of the neurotrophins and CNTF on developing statoacoustic neurons: comparison with an otocyst-derived factor.</a> Dev. Biol. 159 (1), 353-353 (1993).</li><li>Juurlink, B. e. r. n. h. a. r. d. H. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chick+Spinal+Somatic+Motoneurons+in+Culture.">Chick Spinal Somatic Motoneurons in Culture.</a> Protocols for Neural Cell Culture.. 59, (2001).</li><li>Fantetti, K. N., Zou, Y., Fekete, D. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Wnts+and+Wnt+inhibitors+do+not+influence+axon+outgrowth+from+chicken+statoacoustic+ganglion+neurons.">Wnts and Wnt inhibitors do not influence axon outgrowth from chicken statoacoustic ganglion neurons.</a> Hear. Res. , (2011).</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+chick+embryo.">A series of normal stages in the development of the chick embryo.</a> Journal of Morphology. 88, 49-49 (1951).</li><li>Bourikas, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sonic+hedgehog+guides+commissural+axons+along+the+longitudinal+axis+of+the+spinal+cord.">Sonic hedgehog guides commissural axons along the longitudinal axis of the spinal cord.</a> Nat. Neurosci. 8 (3), 297-297 (2005).</li><li>Charron, F., Tessier-Lavigne, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Novel+brain+wiring+functions+for+classical+morphogens:+a+role+as+graded+positional+cues+in+axon+guidance.">Novel brain wiring functions for classical morphogens: a role as graded positional cues in axon guidance.</a> Development. 132 (10), 2251-2251 (2005).</li><li>Augsburger, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=BMPs+as+mediators+of+roof+plate+repulsion+of+commissural+neurons.">BMPs as mediators of roof plate repulsion of commissural neurons.</a> Neuron. 24 (1), 127-127 (1999).</li><li>Lyuksyutova, A. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Anterior-posterior+guidance+of+commissural+axons+by+Wnt-frizzled+signaling.">Anterior-posterior guidance of commissural axons by Wnt-frizzled signaling.</a> Science. 302 (5652), 1984-1984 (2003).</li></ol>

No conflicts of interest declared.

We present a method to dissect and culture E4 SAG and E6 spinal cord explants, from chick, under serum-free conditions. This procedure is currently used in our lab to study the effects of various secreted ligands on SAG neuron survival and neurite outgrowth. Novel aspects of this protocol include the use of a serum-free system for culturing explants and the use of beads soaked in growth factors to study effects on SAG neurite outgrowth. A bead method similar to ours has been described for the chick spinal cord10</su...

<table border="1">
<tbody>
<tr>
<td>Reagent</td>
<td>Company</td>
<td>Catalogue number</td>
<td>Comment</td>
</tr>
<tr>
<td>Equipment</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Dissection microscope</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>We use a Leica M80 stereomicroscope with brightfield base illumination</td>
</tr>
<tr>
<td>Slide warmer</td>
<td>C.S. &amp; E.</td>
<td>&nbsp;</td>
<td>Collagen polymerization</td>
</tr>
<tr>
<td>Chicken egg incubator</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>37&deg;C/5&#37; CO2 humidified cell culture incubator</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Dissection Materials</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Sylgard&copy; coated petri dish</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>24-well cell culture plate</td>
<td>Corning</td>
<td>3526</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>&#35;5 Dumont forceps</td>
<td>Fine Science Tools</td>
<td>11252-30</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>&#35;55 Dumont forceps</td>
<td>Fine Science Tools</td>
<td>11295-51</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Stainless steel dissecting pins </td>
<td>Fine Science Tools</td>
<td>26002-10</td>
<td>Embryo pinning, fine dissection</td>
</tr>
<tr>
<td>Moria perforated spoon</td>
<td>Fine Science Tools</td>
<td>10370-17</td>
<td>Embryo harvest/transfer</td>
</tr>
<tr>
<td>200 &micro;l wide mouth pipet tips</td>
<td>Dot Scientific Inc</td>
<td>118-96R</td>
<td>Explant transfer</td>
</tr>
<tr>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>E4 and E6 chicken eggs</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Chick Ringer&rsquo;s solution</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>Embryo harvest</td>
</tr>
<tr>
<td>HBSS</td>
<td>Sigma</td>
<td>H8264</td>
<td>SAG dissection</td>
</tr>
<tr>
<td>L15 medium </td>
<td>Sigma</td>
<td>L5520</td>
<td>Spinal cord dissection medium</td>
</tr>
<tr>
<td>Fetal calf serum</td>
<td>Atlanta Biologicals</td>
<td>S11150</td>
<td>Spinal cord dissection medium</td>
</tr>
<tr>
<td>Vannas Scissors</td>
<td>World Precision Instruments</td>
<td>501778</td>
<td>Spinal cord dissection</td>
</tr>
<tr>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Collagen preparation</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Rat-tail collagen Type I</td>
<td>BD Biosciences</td>
<td>354249</td>
<td>Explant culture</td>
</tr>
<tr>
<td>10X PBS (Sterile)</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>To neutralize collagen</td>
</tr>
<tr>
<td>1N NaOH (Sterile)</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>To neutralize collagen</td>
</tr>
<tr>
<td>Distilled water (Sterile)</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>To neutralize collagen</td>
</tr>
<tr>
<td>pH indicator papers (6.0-8.1)</td>
<td>Whatman</td>
<td>2629-990</td>
<td>To check collagen pH</td>
</tr>
<tr>
<td>15 ml conical tubes</td>
<td>Dot Scientific Inc</td>
<td>818-PG</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>500 &micro;l wide mouth pipet tips</td>
<td>Dot Scientific Inc</td>
<td>119-R100</td>
<td>To pipet collagen</td>
</tr>
<tr>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Explant culture</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>DMEM/F12 medium</td>
<td>Sigma</td>
<td>D8437</td>
<td>Explant culture medium</td>
</tr>
<tr>
<td>ITS+1</td>
<td>Sigma</td>
<td>I2521</td>
<td>Medium supplement</td>
</tr>
<tr>
<td>Penicillin-Streptomycin</td>
<td>Invitrogen</td>
<td>15140-122</td>
<td>Medium supplement</td>
</tr>
<tr>
<td>CNTF (rat)</td>
<td>Sigma</td>
<td>C3835</td>
<td>Medium supplement</td>
</tr>
<tr>
<td>NT-3 (human)</td>
<td>Sigma</td>
<td>N1905</td>
<td>Medium supplement</td>
</tr>
<tr>
<td>Purified mouse Wnt5a </td>
<td>R&amp;D Systems</td>
<td>645-WN-010</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Affi-gel Blue gel beads</td>
<td>Bio-Rad</td>
<td>153-7302</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Immunohistochemistry</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Paraformaldehyde </td>
<td>Sigma</td>
<td>P6148</td>
<td>Fixation</td>
</tr>
<tr>
<td>PBS</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>Rinses</td>
</tr>
<tr>
<td>Triton X-100</td>
<td>Sigma</td>
<td>T9284</td>
<td>Blocking solution </td>
</tr>
<tr>
<td>Sodium azide </td>
<td>Sigma</td>
<td>S8032</td>
<td>Blocking solution</td>
</tr>
<tr>
<td>Calf serum</td>
<td>Invitrogen</td>
<td>16170-078</td>
<td>Blocking solution</td>
</tr>
<tr>
<td>Mouse monoclonal IgG2b anti-&beta; Tubulin 1+II antibody</td>
<td>Sigma</td>
<td>T8535</td>
<td>Labels cell bodies and neurites</td>
</tr>
<tr>
<td>Alexa fluor 488 goat anti &ndash;mouse IgG2b antibody</td>
<td>Invitrogen</td>
<td>A21141</td>
<td>Detects &beta; Tubulin antibody</td>
</tr>
<tr>
<td>Teflon micro spatula</td>
<td>VWR</td>
<td>57949-033</td>
<td>Release collagen gels from well</td>
</tr>
</tbody>
</table>

dissection and culture of chick statoacoustic ganglion and spinal cord explants in collagen gels for neurite outgrowth assays

The sensory organs of the chicken inner ear are innervated by the peripheral processes of statoacoustic ganglion (SAG) neurons. Sensory organ innervation depends on a combination of axon guidance cues1 and survival factors2 located along the trajectory of growing axons and/or within their sensory organ targets. For example, functional interference with a classic axon guidance signaling pathway, semaphorin-neuropilin, generated misrouting of otic axons3. Also, several growth factors expressed in the sensory targets of the inner ear, including Neurotrophin-3 (NT-3) and Brain Derived Neurotrophic Factor (BDNF), have been manipulated in transgenic animals, again leading to misrouting of SAG axons4. These same molecules promote both survival and neurite outgrowth of chick SAG neurons in vitro5,6.
Here, we describe and demonstrate the in vitro method we are currently using to test the responsiveness of chick SAG neurites to soluble proteins, including known morphogens such as the Wnts, as well as growth factors that are important for promoting SAG neurite outgrowth and neuron survival. Using this model system, we hope to draw conclusions about the effects that secreted ligands can exert on SAG neuron survival and neurite outgrowth. 
SAG explants are dissected on embryonic day 4 (E4) and cultured in three-dimensional collagen gels under serum-free conditions for 24 hours. First, neurite responsiveness is tested by culturing explants with protein-supplemented medium. Then, to ask whether point sources of secreted ligands can have directional effects on neurite outgrowth, explants are co-cultured with protein-coated beads and assayed for the ability of the bead to locally promote or inhibit outgrowth. We also include a demonstration of the dissection (modified protocol7) and culture of E6 spinal cord explants. We routinely use spinal cord explants to confirm bioactivity of the proteins and protein-soaked beads, and to verify species cross-reactivity with chick tissue, under the same culture conditions as SAG explants. These in vitro assays are convenient for quickly screening for molecules that exert trophic (survival) or tropic (directional) effects on SAG neurons, especially before performing studies in vivo. Moreover, this method permits the testing of individual molecules under serum-free conditions, with high neuron survival8.

Watch this Scientific Journal Video about Dissection and Culture of Chick Statoacoustic Ganglion and Spinal Cord Explants in Collagen Gels for Neurite Outgrowth Assays at JoVE.com

Dissection and Culture of Chick Statoacoustic Ganglion and Spinal Cord Explants in Collagen Gels for Neurite Outgrowth Assays

We demonstrate how to dissect and culture chick E4 statoacoustic ganglion and E6 spinal cord explants. Explants are cultured under serum-free conditions in 3D collagen gels for 24 hours. Neurite responsiveness is tested with growth factor-supplemented medium and with protein-coated beads.

The sensory organs of the chicken inner ear are innervated by the peripheral processes of statoacoustic ganglion (SAG) neurons. Sensory organ innervation ...

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Research

JoVE Journal

Neuroscience

12.0K Views.  Purdue University. We demonstrate how to dissect and culture chick E4 statoacoustic ganglion and E6 spinal cord explants. Explants are cultured under serum-free conditions in 3D collagen gels for 24 hours. Neurite responsiveness is tested with growth factor-supplemented medium and with protein-coated beads.

Video: Dissection and Culture of Chick Statoacoustic Ganglion and Spinal Cord Explants in Collagen Gels for Neurite Outgrowth Assays

Dissection and Culture of Commissural Neurons from Embryonic Spinal Cord

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies of these neurons are located in the dorsal spinal cord and their axons follow stereotyped trajectories during embryonic development. Commissural axons initially project ventrally towards the floorplate. After crossing the midline, these axons turn anteriorly and project towards the brain. Each of these steps is regulated by the action of several guidance cues. Cultures highly enriched in commissural neurons are ideally suited for many experiments addressing the mechanisms of axon pathfinding, including turning assays, immunochemistry and biochemistry. Here, we describe a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. First, the spinal cord is isolated and dorsal strips are dissected out. The dorsal tissue is then dissociated into a cell suspension by trypsinization and mechanical disruption. Neurons are plated onto poly-L-lysine-coated glass coverslips or tissue-culture dishes. After 30 hours in vitro, most neurons have extended an axon. The purity of the culture (Yam et al. 2009), typically over 90%, can be assessed by immunolabeling with the commissural neuron markers DCC, LH2 and TAG1 (Helms and Johnson, 1998). This neuronal preparation is a useful tool for in vitro studies of the cellular and molecular mechanisms of commissural axon growth and guidance during spinal cord development.

This video demonstrates a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. Dissociated commissural neurons are useful to study the cellular and molecular mechanisms of axon growth and guidance.

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies ...

A Functional Motor Unit in the Culture Dish: Co-culture of Spinal Cord Explants and Muscle Cells

Human primary muscle cells cultured aneurally in monolayer rarely contract spontaneously because, in the absence of a nerve component, cell differentiation is limited and motor neuron stimulation is missing1. These limitations hamper the in vitro study of many neuromuscular diseases in cultured muscle cells. Importantly, the experimental constraints of monolayered, cultured muscle cells can be overcome by functional innervation of myofibers with spinal cord explants in co-cultures.
Here, we show the different steps required to achieve an efficient, proper innervation of human primary muscle cells, leading to complete differentiation and fiber contraction according to the method developed by Askanas2. To do so, muscle cells are co-cultured with spinal cord explants of rat embryos at ED 13.5, with the dorsal root ganglia still attached to the spinal cord slices. After a few days, the muscle fibers start to contract and eventually become cross-striated through innervation by functional neurites projecting from the spinal cord explants that connecting to the muscle cells. This structure can be maintained for many months, simply by regular exchange of the culture medium.
The applications of this invaluable tool are numerous, as it represents a functional model for multidisciplinary analyses of human muscle development and innervation. In fact, a complete de novo neuromuscular junction installation occurs in a culture dish, allowing an easy measurement of many parameters at each step, in a fundamental and physiological context.
Just to cite a few examples, genomic and/or proteomic studies can be performed directly on the co-cultures. Furthermore, pre- and post-synaptic effects can be specifically and separately assessed at the neuromuscular junction, because both components come from different species, rat and human, respectively. The nerve-muscle co-culture can also be performed with human muscle cells isolated from patients suffering from muscle or neuromuscular diseases3, and thus can be used as a screening tool for candidate drugs. Finally, no special equipment but a regular BSL2 facility is needed to reproduce a functional motor unit in a culture dish. This method thus is valuable for both the muscle as well as the neuromuscular research communities for physiological and mechanistic studies of neuromuscular function, in a normal and disease context.

Cultured muscle cells are an inadequate model to recapitulate innervated muscle in vivo. A functional motor unit can be reproduced in vitro by innervation of differentiated human primary muscle cells using rat embryo spinal cord explants. This article describes how co-cultures of spinal cord explants and muscle cells are established.

Human primary muscle cells cultured  aneurally in monolayer rarely contract spontaneously because, in the absence of  a nerve component, cell ...

Dissection and Culture of Mouse Dopaminergic and Striatal Explants in Three-Dimensional Collagen Matrix Assays

Midbrain dopamine (mdDA) neurons project via the medial forebrain bundle towards several areas in the telencephalon, including the striatum1. Reciprocally, medium spiny neurons in the striatum that give rise to the striatonigral (direct) pathway innervate the substantia nigra2. The development of these axon tracts is dependent upon the combinatorial actions of a plethora of axon growth and guidance cues including molecules that are released by neurites or by (intermediate) target regions3,4. These soluble factors can be studied in vitro by culturing mdDA and/or striatal explants in a collagen matrix which provides a three-dimensional substrate for the axons mimicking the extracellular environment. In addition, the collagen matrix allows for the formation of relatively stable gradients of proteins released by other explants or cells placed in the vicinity (e.g. see references 5 and 6). Here we describe methods for the purification of rat tail collagen, microdissection of dopaminergic and striatal explants, their culture in collagen gels and subsequent immunohistochemical and quantitative analysis. First, the brains of E14.5 mouse embryos are isolated and dopaminergic and striatal explants are microdissected. These explants are then (co)cultured in collagen gels on coverslips for 48 to 72 hours in vitro. Subsequently, axonal projections are visualized using neuronal markers (e.g. tyrosine hydroxylase, DARPP32, or &beta;III tubulin) and axon growth and attractive or repulsive axon responses are quantified. This neuronal preparation is a useful tool for in vitro studies of the cellular and molecular mechanisms of mesostriatal and striatonigral axon growth and guidance during development. Using this assay, it is also possible to assess other (intermediate) targets for dopaminergic and striatal axons or to test specific molecular cues.

Explants from the midbrain dopamine system and striatum are used in a collagen matrix assay for the in vitro analysis of mesostriatal and striatonigral pathway development. In this assay axonal outgrowth and guidance can be manipulated and quantified. It can also be modified for assessing other regions or molecular cues.

Midbrain dopamine (mdDA) neurons project via the medial forebrain bundle towards several areas in the telencephalon, including the striatum1. ...

Chicken Embryo Spinal Cord Slice Culture Protocol

Slice cultures can facilitate the manipulation of embryo development both pharmacologically and through gene manipulations. In this reduced system, potential lethal side effects due to systemic drug applications can be overcome. However, culture conditions must ensure that normal development proceeds within the reduced environment of the slice. We have focused on the development of the spinal cord, particularly that of spinal motor neurons. We systematically varied culture conditions of chicken embryo slices from the point at which most spinal motor neurons had been born. We assayed the number and type of motor neurons that survived during the culture period and the position of those motor neurons compared to that in vivo. We found that serum type and neurotrophic factors were required during the culture period and were able to keep motor neurons alive for at least 24 hr and allow those motor neurons to migrate to appropriate positions in the spinal cord. We present these culture conditions and the methodology of preparing the embryo slice cultures using eviscerated chicken embryos embedded in agarose and sliced using a vibratome.

Slice cultures facilitate the manipulation of embryo development by gene and pharmacological perturbations. However, culture conditions must ensure that normal development can proceed within the reduced environment of the slice. We illustrate a protocol that facilitates normal spinal cord development to proceed for at least 24 hr.

Slice cultures can facilitate the manipulation of  embryo development both pharmacologically and through gene manipulations. In  this reduced system, ...

Dissection and Lateral Mounting of Zebrafish Embryos: Analysis of Spinal Cord Development

The zebrafish spinal cord is an effective investigative model for nervous system research for several reasons. First, genetic, transgenic and gene knockdown approaches can be utilized to examine the molecular mechanisms underlying nervous system development. Second, large clutches of developmentally synchronized embryos provide large experimental sample sizes. Third, the optical clarity of the zebrafish embryo permits researchers to visualize progenitor, glial, and neuronal populations. Although zebrafish embryos are transparent, specimen thickness can impede effective microscopic visualization. One reason for this is the tandem development of the spinal cord and overlying somite tissue. Another reason is the large yolk ball, which is still present during periods of early neurogenesis. In this article, we demonstrate microdissection and removal of the yolk in fixed embryos, which allows microscopic visualization while preserving surrounding somite tissue. We also demonstrate semipermanent mounting of zebrafish embryos. This permits observation of neurodevelopment in the dorso-ventral and anterior-posterior axes, as it preserves the three-dimensionality of the tissue.

Developmental processes such as proliferation, patterning, differentiation, and axon guidance can be readily modeled in the zebrafish spinal cord. In this article, we describe a mounting procedure for zebrafish embryos, which optimizes visualization of these events.

The zebrafish spinal cord is an effective investigative model for nervous system research for several reasons. First, genetic, transgenic and gene ...

Complete Spinal Cord Injury and Brain Dissection Protocol for Subsequent Wholemount In Situ Hybridization in Larval Sea Lamprey

After a complete spinal cord injury, sea lampreys at first are paralyzed below the level of transection. However, they recover locomotion after several weeks, and this is accompanied by short distance regeneration (a few mm) of propriospinal axons and spinal-projecting axons from the brainstem. Among the 36 large identifiable spinal-projecting neurons, some are good regenerators and others are bad regenerators. These neurons can most easily be identified in wholemount CNS preparations. In order to understand the neuron-intrinsic mechanisms that favor or inhibit axon regeneration after injury in the vertebrates CNS, we determine differences in gene expression between the good and bad regenerators, and how expression is influenced by spinal cord transection. This paper illustrates the techniques for housing larval and recently transformed adult sea lampreys in fresh water tanks, producing complete spinal cord transections under microscopic vision, and preparing brain and spinal cord wholemounts for in situ hybridization. Briefly, animals are kept at 16&#160;&#176;C and anesthetized in 1% Benzocaine in lamprey Ringer. The spinal cord is transected with iridectomy scissors via a dorsal approach and the animal is allowed to recover in fresh water tanks at 23 &#176;C. For in situ hybridization, animals are reanesthetized and the brain and cord removed via a dorsal approach.

Complete Spinal Cord Injury and Brain Dissection Protocol for Subsequent Wholemount In Situ Hybridization in Larval Sea Lamprey

Lampreys recover locomotion after a complete spinal cord injury. However, some spinal-projecting neurons are good regenerators and others are not. This paper illustrates the techniques for housing sea lamprey larvae (and recently transformed adults), producing complete spinal cord transections and preparing wholemount brains and spinal cords for in situ hybridization.

After a complete spinal cord injury, sea lampreys at first are paralyzed below the level of transection. However, they recover locomotion after several ...

RNA-Seq Analysis of Differential Gene Expression in Electroporated Chick Embryonic Spinal Cord

In ovo electroporation of the chick neural tube is a fast and inexpensive method for identification of gene function during neural development. Genome wide analysis of differentially expressed transcripts after such an experimental manipulation has the potential to uncover an almost complete picture of the downstream effects caused by the transfected construct. This work describes a simple method for comparing transcriptomes from samples of transfected embryonic spinal cords comprising all steps between electroporation and identification of differentially expressed transcripts. The first stage consists of guidelines for electroporation and instructions for dissection of transfected spinal cord halves from HH23 embryos in ribonuclease-free environment and extraction of high-quality RNA samples suitable for transcriptome sequencing. The next stage is that of bioinformatic analysis with general guidelines for filtering and comparison of RNA-Seq datasets in the Galaxy public server, which eliminates the need of a local computational structure for small to medium scale experiments. The representative results show that the dissection methods generate high quality RNA samples and that the transcriptomes obtained from two control samples are essentially the same, an important requirement for detection of differential expression genes in experimental samples. Furthermore, one example is provided where experimental overexpression of a DNA construct can be visually verified after comparison with control samples. The application of this method may be a powerful tool to facilitate new discoveries on the function of neural factors involved in spinal cord early development.

This video shows the basic steps for performing whole transcriptome analysis on dissected chick embryonic spinal cord samples after transfection with in ovo electroporation.

In ovo electroporation of the chick neural tube is a fast and inexpensive method for identification of gene function during neural development. Genome ...

Spinal Cord Neurons Isolation and Culture from Neonatal Mice

We present a protocol for the isolation and culture of spinal cord neurons. The neurons are obtained from neonatal C57BL/6 mice and are isolated on postnatal day 1-3. A mouse litter, usually 4-10 pups born from one breeding pair, is gathered for one experiment, and spinal cords are collected individually from each mouse after euthanasia with isoflurane. The spinal column is dissected out and then the spinal cord is released from the column. The spinal cords are then minced to increase the surface area of delivery for an enzymatic protease that allows for the neurons and other cells to be released from the tissue. Trituration is then used to release the cells into solution. This solution is subsequently fractionated in a density gradient to separate the various cells in solution, allowing for neurons to be isolated. Approximately 1-2.5 x 106 neurons can be isolated from one litter group. The neurons are then seeded onto wells coated with adhesive factors that allow for proper growth and maturation. The neurons take approximately 7 days to reach maturity in the growth and culture medium and can be used thereafter for treatment and analysis.

This study presents a technique for the isolation of neurons from WT neonatal mice. It requires the careful dissection of the spinal cord from the neonatal mouse, followed by the separation of neurons from the spinal cord tissue through mechanical and enzymatic cleavage.

We present a protocol for the isolation and culture of spinal cord neurons. The neurons are obtained from neonatal C57BL/6 mice and are isolated on ...

Intra-Arterial Delivery of Neural Stem Cells to the Rat and Mouse Brain: Application to Cerebral Ischemia

Neural stem cell (NSC) therapy is an emerging innovative treatment for stroke, traumatic brain injury and neurodegenerative disorders. As compared to intracranial delivery, intra-arterial administration of NSCs is less invasive and produces a more diffuse distribution of NSCs within the brain parenchyma. Further, intra-arterial delivery allows the first-pass effect in the brain circulation, lessening the potential for trapping of cells in peripheral organs, such as liver and spleen, a complication associated with peripheral injections. Here, we detail the methodology, in both mice and rats, for delivery of NSCs through the common carotid artery (mouse) or external carotid artery (rat) to the ipsilateral hemisphere after an ischemic stroke. Using GFP-labeled NSCs, we illustrate the widespread distribution achieved throughout the rodent ipsilateral hemisphere at 1 d, 1 week and 4 weeks after postischemic delivery, with a higher density in or near the ischemic injury site. In addition to long-term survival, we show evidence of differentiation of GFP-labeled cells at 4 weeks. The intra-arterial delivery approach described here for NSCs can also be used for administration of therapeutic compounds, and thus has broad applicability to varied CNS injury and disease models across multiple species.

A method for delivering neural stem cells, adaptable for injecting solutions or suspensions, through the common carotid artery (mouse) or external carotid artery (rat) after ischemic stroke is reported. Injected cells are distributed broadly throughout the brain parenchyma and can be detected up to 30 d after delivery.

Neural stem cell (NSC) therapy is an emerging innovative treatment for stroke, traumatic brain injury and neurodegenerative disorders. As compared to ...

Induction of Complete Transection-Type Spinal Cord Injury in Mice

Spinal cord injury (SCI) largely leads to irreversible and permanent loss of function, most commonly as a result of trauma. Several treatment options, such as cell transplantation methods, are being researched to overcome the debilitating disabilities arising from SCI. Most pre-clinical animal trials are conducted in rodent models of SCI. While rat models of SCI have been widely used, mouse models have received less attention, even though mouse models can have significant advantages over rat models. The small size of mice equates to lower animal maintenance costs than for rats, and the availability of numerous transgenic mouse models is advantageous for many types of studies. Inducing repeatable and precise injury in the animals is the primary challenge for SCI research, which in small rodents requires high-precision surgery. The transection-type injury model has been a commonly used injury model over the last decade for transplantation-based therapeutic research, however a standardized method for inducing a complete transection-type injury in mice does not exist. We have developed a surgical protocol for inducing a complete transection type injury in C57BL/6 mice at thoracic vertebral level 10 (T10). The procedure uses a small tip drill instead of rongeurs to precisely remove the lamina, after which a thin blade with rounded cutting edge is used to induce the spinal cord transection. This method leads to reproducible transection-type injury in small rodents with minimal collateral muscle and bone damage and therefore minimizes confounding factors, specifically where behavioral functional outcomes are analyzed.

This protocol describes how to create a precise laminectomy for induction of stable transection-type spinal cord injury in the mouse model, with minimal collateral damage for spinal cord injury research.

Spinal cord injury (SCI) largely leads to irreversible and permanent loss of function, most commonly as a result of trauma. Several treatment options, ...