This work was supported by grants from the Canadian Institutes of Health Research (CIHR), the Peter Lougheed Medical Research Foundation, the McGill Program in Neuroengineering, the Fonds de Recherche en Sant&eacute; du Qu&eacute;bec (FRSQ), and the Canada Foundation for Innovation (CFI). S&eacute;bastien D. Langlois was supported by a Master's Training Award from the Fonds de la recherch&eacute; en sant&eacute; du Qu&eacute;bec (FRSQ) and by a Frederick Banting and Charles Best Canada Graduate Scholarships Master's Award from the Canadian Institutes of Health Research (CIHR). We are thankful to Jessica M.T. Pham for assistance with figures.

1. Dissection of embryonic rat dorsal spinal cord
General recommendations
Keep L-15 medium on ice and frequently change the medium in the dissection dish to keep the embryos cool. This helps preserve tissue integrity. All steps are performed with two pairs of Dumont #5 forceps unless stated. To avoid contamination, spray all tools and working surfaces with 70% ethanol and keep dissection medium bottle closed. To transfer embryos between dishes, use a cut plastic pipette or a perforated spoon. It is critical not to damage the spinal cord (nicking, stretching) to successfully complete the dissection.
Preparation
<ul>
	<li>Cold L-15 medium</li>
	<li>50 mL of L-15 + 10% heat-inactivated horse serum (HiHS). Keep on ice.</li>
</ul>
Spinal cord dissection
<ol>
	<li>Euthanize an E13 pregnancy-staged rat (E0 = first day following mating day) with a CO2 chamber according to institutional guidelines.</li>
	<li>Spray 70% ethanol on the abdomen. Pinch and pull up the skin of the lower abdominal region with forceps and cut with surgical scissors. Repeat to cut through the muscle and peritoneal layers to reach the abdominal cavity. Create a V-shape incision by cutting tissue along the sides of the abdomen, up to the thorax. Lift and pull back the tissue to expose the abdominal cavity.</li>
	<li>The uterus is attached at three locations: at the lower center abdomen, and at both upper lateral corners. Lift the uterus by grabbing on tissue between embryonic sacs. Cut the connective tissue to remove the uterus and place in a Petri dish filled with L-15 on ice.</li>
	<li>The next steps are done under a dissection microscope. To separate an embryo from extraembryonic tissues and embryonic membranes, grab the tissue between uterine sacs with one pair of forceps,. With the other pair, pinch the more transparent side of the sac to cut through the superficial membranes (the darker side is the placenta). Squeeze out the embryo by gently pressing the placenta side of the sac. Remove all embryos and place in a Petri dish filled with L-15 on ice.</li>
	<li>Place one embryo in a 10 cm Petri dish filled with ice-cold L-15. Using microscissors, cut away the head and posterior parts at angles shown in figure 1a. Cutting at these angles will help position the embryo when placed "ventral face down". </li>
	<li>Position the embryo "ventral face down" (anterior pointing away, posterior pointing towards the experimenter) (fig. 1b).</li>
	<li>Firmly hold the embryo from one side, using the forceps to "pin down" the embryo (fig. 1c). Do not move the forceps, as it will tear the tissue. With the other pair of forceps, peel off the skin covering the back of the embryo starting from the region in between the "holding" forceps (fig 1d-e). This will expose the spinal cord which, at this point, is wrapped by membranes (the meninges). Grab the skin from the sides rather than from above the spinal cord to avoid nicking the spinal cord. </li>
	<li>Partially detach the tissue to the right side of the spinal cord (fig. 1f). Starting at the level of the holding forceps, poke the tissue with closed forceps, as close as possible to the spinal cord. Then, slowly open the forceps to tear the tissue. This should detach the dorsal root ganglia (DRGs) from the spinal cord, and should also disrupt ventral organs. Leave some tissue attached at the anterior and posterior ends. Leaving tissue attached adds weight to the embryo, thus preventing it from being pulled upward during the next step. In addition, that tissue is used to hold on to the embryo in later steps.</li>
 <li>Starting from the anterior end, use the hook-shaped tungsten needle to cut the meninges and open the spinal cord along the roofplate (fig. 1g).</li>
	<li>Rotate the embryo 180 degrees (posterior pointing away, anterior pointing towards the experimenter) (fig. 1h). This allows the experimenter to use the same hand to detach the tissue from the other side of the embryo. </li>
	<li>Hold the embryo by grabbing onto the previously detached side, and disrupt the tissue from the remaining side using the same method (see 1.8.). When done, completely detach the tissue on both sides of the spinal cord. Alternative: Completely detach the tissue remaining on the left side, but leave some tissue attached at the anterior and posterior end on the right side. This can sometimes help maintain the spinal cord in position in step 1.12.</li>
	<li>Place the spinal cord on its side and remove most of the remaining mesenchymal tissue and DRGs (fig. 1i).</li>
	<li>At this step, the meninges and the spinal cord are two "sheets" of tissue apposed onto each other. Pin down the larger, anterior-most portion of the spinal cord, and peel off a short segment of meninges from the spinal cord (fig. 1j, left). Now, hold the two separated segments by grabbing across with forceps, and peel off the meninges with a smooth, constant movement (fig. 1j). Uneven "peeling" will lead to breakage of the spinal cord and/or the meninges layer. It is usually not possible to recover the portion of the spinal cord still attached to meninges.</li>
	<li>Using a plastic pipette, transfer the isolated spinal cord to a Petri dish containing L-15 + 10% HiHS and leave on ice. Typically, spinal cords of all embryos are collected before dissecting the dorsal portions.</li>
</ol>
Dorsal spinal cord dissection
<ol start="15">
	<li>Place one spinal cord in a Petri dish containing L-15+10% HiHS, lying flat in an "open-book" configuration. Cut away the wider, anterior portion (which includes part of the hindbrain) (fig. 1k).</li>
	<li>The dorsal tissue is located in the lateral-most parts of the spinal cord (fig 1l, left). While pinning the cord with a straight tungsten needle, use the L-shaped tungsten needle to cut out a strip that is 1/5th the width of half the spinal cord. (fig. 1l, right). Place the dorsal strips in a 15 mL plastic tube containing L-15 + 10% HiHS on ice. Dorsal neural tube sections of ~12 E13 embryos will yield ~3.5-4 million cells for plating after dissociation. More cells can be obtained by performing a coarse dissection of the dorsal spinal cord (cutting wider dorsal strips), but commissural neuron purity will be reduced.</li>
</ol>
2. Commissural neuron culture
General recommendations
All steps should be performed under sterile conditions in a tissue culture hood unless otherwise stated. Use fresh medium and freshly-thawed supplements and reagents. The dissociation and trituration steps are performed in Ca2+/Mg2+-free HBSS to minimize Ca2+/Mg2+-dependent adhesion.
Preparation
<ul>
	<li>coated coverslips (use German Desag glass) or tissue culture plates (see coating procedure below).</li>
	<li>warm Neurobasal Plating Media (see below), in tissue culture dishes and CO2 equilibrated in the tissue culture incubator for at least 1.5 h before plating.</li>
	<li>2.5% trypsin in the 37 &deg;C waterbath</li>
	<li>2 bottles of HBSS Ca2+/Mg2+-free, one at 4 &deg;C, one at 37 &deg;C.</li>
	<li>two fire-polished glass Pasteur pipettes one with a diameter half the usual size, and one with a diameter slightly smaller than half. Use sterilized Pasteur pipettes. To fire-polish the pipettes, use a Bunsen burner (the top of the light blue flame) to melt the tip to slightly decrease the diameter. Because this step is performed outside the tissue culture hood, spray the pipettes with 70% ethanol before placing under the tissue culture hood. For more efficient recovery of neurons, coat the Pasteur pipettes with media containing serum before the beginning of the dissociation or just before the trituration step (fill pipettes with media and keep for 30 seconds). This will prevent cells from sticking to the inside of the Pasteur pipette during the trituration.</li>
	<li>sterile milliQ water for washing PLL-coated dishes or coverslips</li>
	<li>12.5% MgSO4 solution in HBSS</li>
</ul>
Poly-L-lysine coating
If using glass coverslips, acid-wash for 24 hours and sterilize prior to plating (see Kaech and Banker, 2006). Use German Desag glass coverslips.
To coat poly-L-lysine on glass coverslips or plastic tissue culture dishes:
<ul>
	<li>under a tissue culture hood, cover surfaces with a small dome of 100 ug/ml PLL solution for 1.75-2 hours.</li>
	<li>wash twice with milliQ water, at least 5 minutes per wash (can be performed during the cell dissociation step below).</li>
	<li>store in water until use. Do not let the PLL-coated surface dry. To reduce PLL wastage for coverslips coating, place the coverslips in sterile bacterial Petri dishes. Coat and wash the coverslips by placing a dome of liquid on the coverslips. The bacterial Petri dishes are hydrophobic, so the liquid should remain on the glass coverslips. Transfer the coverslips to tissue culture dishes just before plating the cells. For neurons plated on plastic tissue culture dishes, adhesion is normally higher, thus neurite elongation might be diminished.</li>
</ul>
Dissociation and plating
<ol>
	<li>Verify that the dorsal strips have settled down to the bottom of the tube. Remove most of the L-15+10% HiHS with a Pasteur pipette. Quickly wash the dorsal neural tube strips once by adding 3 ml of cold (4 &deg;C) HBSS.</li>
	<li>Let the dorsal neural tube strips settle for 2 minutes, then remove the HBSS with a Pasteur pipette.</li>
	<li>Add warm (37 &deg;C) HBSS to a volume of 4.7 ml. Then add 0.3 ml of 2.5 % trypsin to give a final concentration of 0.15% trypsin.</li>
	<li>Incubate at 37 &deg;C in the waterbath for 7 min. Mix once gently half-way through the incubation.</li>
 <li>Add 30 &mu;l DNAse (25 000 U/mL) for a final concentration of 150 U/mL. Add 60 &mu;l of MgSO4 and mix briefly, for a final concentration of 0.15%. For tissue from coarse dissections, incubate for an extra 1 minute at 37 &deg;C in the waterbath. At this stage, the dorsal neural tube sections should be fragmented. If the dorsal neural tube sections have not begun to fragment, it usually means that the 2.5% trypsin stock is old, and new stock should be thawed, or the wash with cold HBSS did not effectively remove the HiHS from the sample.</li>
	<li>Centrifuge the tissue fragments at 200 g for 4 min.</li>
	<li>Remove the supernatant with a Pasteur pipette, leaving ~50-100 &mu;l of liquid at the bottom of the tube.</li>
	<li>Flick the tube gently to loosen the pellet, then wash the cells by adding 5 ml of warm HBSS. Let settle at room temperature for 2 min.</li>
	<li>Centrifuge at 200 g for 5 min.</li>
	<li>Remove the supernatant with a Pasteur pipette, leaving ~50-100 &mu;l of liquid at the bottom of the tube.</li>
	<li>Flick the tube gently to loosen the pellet and partially resuspend the cells. Then add 2 ml of warm HBSS.</li>
	<li>Use the small (half-diameter) fire-polished glass Pasteur pipette to dissociate the cells by slowly pipetting up-and-down 4-6 times. Avoid making bubbles, and pipette the liquid against the side of the tube. Do not over-triturate.</li>
	<li>Use the smallest fire-polished glass Pasteur pipette to further dissociate the cells by slowly pipetting up-and-down 3-4 times. Avoid making bubbles, and pipette the liquid against the side of the tube. Do not over-triturate. For tissue from coarse dissections, add an extra 1 ml of warm HBSS to the tube at the end of the dissociation. When dissociating the cells, it is not necessary to dissociate all the cell clumps and aggregates. Stop pipetting up-and-down or change to a Pasteur pipette with a smaller diameter if you see no further decrease in the size of the cell aggregates with further pipetting.</li>
	<li>Let any remaining tissue fragments settle in the tube for 1 min. It is not necessary to transfer the cells to a new tube.</li>
	<li>Take 20 &mu;l of cell suspension and add 5 &mu;l of trypan blue. Count cells on a haemocytometer. Neurons should be &ge;95% viable by trypan blue exclusion.</li>
	<li>Plate the neurons in Neurobasal Plating Media (see Recipes below).
<ul>
	<li>Suggested plating densities for obtaining isolated neurons (low-density cultures to avoid neurons clumping or overlapping each other):</li>
	<li>120 000 - 180 000 cells/well for a 6 well plate</li>
	<li>60 000 - 75 000 cells/well for a 12 well plate</li>
</ul>
</li>
	<li>16-18 hours later, change the media to Neurobasal Growth Media (see Recipes below) Do not use a vacuum pump to aspirate the media from the culture dish when changing media; gently use a pipette. This avoids dislodging the neurons.</li>
</ol>
Representative Results:
Four hours after plating, neurons should have adhered to the poly-L-lysine (PLL)-coated surface. Under phase contrast illumination, adhered cell bodies are typically relatively flat and oval-shaped (fig. 2a). Cells that have not well adhered appear as spheres that move slightly when the dish is very gently tapped on the side. Many factors can potentially impede the adhesion of cells (see discussion). 
After 30 hours in vitro, most neurons have extended an axon with a visible growth cone (fig 2c,d). If poor axonal growth is observed, verify that the Neurobasal Growth Media has been made with fresh medium and supplements. Neurons remain healthy for at least 6 days in these conditions. This procedure has proven to reliably yield preparations highly enriched in commissural neurons, with ~90% of neurons expressing DCC (Yam et al. 2009). The width of the dorsal spinal cord strip that is used for preparing the cell suspension will affect the purity of the culture, with greater purity when thinner strips are used. An example application is shown in figure 3 (immunofluorescence). See the article by Yam et al. (2009) for more examples.
<img src="/files/ftp_upload/1773/1773fig1tmb.jpg" border="0" alt="Figure 1" /> Figure 1. Schematics of spinal cord dissection steps. D = Dorsal, V =Ventral. <a href="https://www.jove.com/files/ftp_upload/1773/1773fig1.jpg">Click here to see a larger figure.</a>
<img src="/files/ftp_upload/1773/1773fig2.jpg" alt="Figure 2" /> Figure 2.Representative result of isolated commissural neurons plated on a PLL-coated glass coverslip. a,b) 4 hours after plating, neurons have adhered to the surface. Bar = 20 &mu;m. c,d) 30 hours after plating, most neurons have extended an axon with a visible growth cone. Bar = 20 &mu;m.
<img src="/files/ftp_upload/1773/1773fig3.jpg" alt="Figure 3" /> Figure 3. A commissural neuron immunostained for gamma-tubulin (green), with F-actin labeled by phalloidin (F-actin, red) and the nucleus by DAPI (blue).
Recipes and comments
Neurobasal Plating Media
<ul>
	<li>Neurobasal</li>
	<li>10% Heat-inactivated FBS (HiFBS)</li>
	<li>2 mM L-glutamine (from 200 mM stock solution) Optional: Penicillin/Streptomycin antibiotics (use half the normal concentration)</li>
</ul>
Neurobasal Growth Media
<ul>
	<li>Neurobasal</li>
	<li>B27 (1/50 dilution from stock)</li>
	<li>2 mM L-glutamine (from 200 mM stock solution)</li>
	<li>Optional: Penicillin/Streptomycin antibiotics (use half the normal concentration) Once the media is made, it can be kept at 4 &deg;C for two weeks. To equilibrate the temperature and the pH of media before plating the cells, place media in tissue culture Petri dishes and place in a tissue culture incubator for at least 1.5 hours.</li>
</ul>
Neurobasal
After a bottle of Neurobasal medium has been opened, it can be kept for one month at 4 &deg;C in the dark. Dispose of Neurobasal which has been opened for more than one month otherwise cell survival will be lower.
Heat-inactivated fetal bovine serum (HiFBS) or horse serum (HiHS)
To heat-inactivate FBS or HS, heat at 56 &deg;C in a water bath for 30 minutes. Swirl the bottle approximately every 10 minutes or so. (For accuracy use a bottle of similar size filled with water. Place a thermometer in the water bottle to see when 56 &deg;C is reached. Begin timing at this point.) Heat-inactivated FBS may need to be centrifuged to clear precipitates, and can be aliquoted and re-frozen at -20 &deg;C.
L-Glutamine
Always thaw a fresh aliquot of L-glutamine for each experiment.
B27
Aliquots of B27 can be kept at 20 &deg;C for long term storage, or at 4 &deg;C for up to one month.

<ol>
	<li>Charron, F., Stein, E., Jeong, J., McMahon, A. P., 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=The+morphogen+sonic+hedgehog+is+an+axonal+chemoattractant+that+collaborates+with+netrin-1+in+midline+axon+guidance.">The morphogen sonic hedgehog is an axonal chemoattractant that collaborates with netrin-1 in midline axon guidance.</a> Cell. , 113-1111 (2003).</li><li>Fabre, P., Shimogori, T., Charron, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Segregation+of+ipsilateral+retinal+ganglion+cell+axons+at+the+optic+chiasm+requires+the+Shh+Receptor+Boc.">Segregation of ipsilateral retinal ganglion cell axons at the optic chiasm requires the Shh Receptor Boc.</a> Journal of Neuroscience. 30, 266-275 (2010).</li><li>Helms, A. W., Johnson, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Progenitors+of+dorsal+commissural+interneurons+are+defined+by+MATH1+expression.">Progenitors of dorsal commissural interneurons are defined by MATH1 expression.</a> Development. 125, 919-928 (1998).</li><li>Kaech, S., Banker, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Culturing+hippocampal+neurons.">Culturing hippocampal neurons.</a> Nature Protocols. 1, 2406-2415 (2006).</li><li>Okada, A., Charron, F., Morin, S., Shin, D. S., Wong, K., Fabre, P. J., Tessier-Lavigne, M., McConnell, S. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Boc+is+a+receptor+for+sonic+hedgehog+in+the+guidance+of+commissural+axons.">Boc is a receptor for sonic hedgehog in the guidance of commissural axons.</a> Nature. 444, 369-373 (2006).</li><li>Yam, P. T., Langlois, S. D., Morin, S., Charron, F. <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+axons+through+a+noncanonical,+Src-family-kinase-dependent+signaling+pathway.">Sonic hedgehog guides axons through a noncanonical, Src-family-kinase-dependent signaling pathway.</a> Neuron. 62, 349-362 (2009).</li></ol>

We have described a method to dissect and culture commissural neurons from embryonic rat spinal cord. This procedure has been routinely used in our lab to reliably prepare neurons to study the cellular and molecular mechanisms of axon guidance. For cell biology and immunochemistry experiments, dissection of one litter produces sufficient neurons. When more cells are needed, such as in many biochemistry experiments, dissection of two litters may be required. For a trained person, dissection and dissociation of ~20 embryos...

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		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Neurobasal medium, liquid</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>21103-049</td>
<td>See Recipes and Comments</td>
</tr>
<tr>
<td>B27 supplement 50X</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>17504</td>
<td>See Recipes and Comments</td>
</tr>
<tr>
<td>Poly-L-lysine 0.01% solution</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>P4707</td>
<td>&nbsp;</td>
<tr>
<td>L-15 medium, powder</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>41300-070</td>
<td>&nbsp;</td>
<tr>
<td>Trypsin 2.5% (10X)</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>15090-046</td>
<td>&nbsp;</td>
<tr>
<td>DNAse I, 25000 U/mL</td>
<td>&nbsp;</td>
<td>Worthington</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>MgSO4</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>M2643</td>
<td>&nbsp;</td>
<tr>
<td>HBSS, Ca2+/Mg2+-free</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>14170-112</td>
<td>&nbsp;</td>
<tr>
<td>L-glutamine 200mM, liquid</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>25030-081</td>
<td>See Recipes and Comments</td>
</tr>		</tbody>
		</table>
		</div>
			<div>
			Dissection of embryonic rat dorsal spinal cord (see also Table I)
<ul>
	<li>E13 pregnancy-staged rat</li>
	<li>Ethanol 70%</li>
	<li>Surgical scissors, Fine Science Tools</li>
	<li>Forceps, Dumont #5, Fine Science Tools</li>
	<li>Petri dishes</li>
	<li>L-15 medium</li>
	<li>Dissection microscope, Leica</li>
	<li>Plastic transfer pipettes</li>
	<li>Microscissors, Fine Science Tools</li>
	<li>Tungsten needles and pin holders (one hook-shaped needle, one L-shapes needle, one straight needle), Fine Science Tools</li>
	<li>Heat-inactivated Horse Serum (HiHS)</li>
	<li>15 ml plastic tubes</li>
</ul>
Commissural neuron culture (see also Table I)
<ul>
	<li>Tissue culture incubators (37&deg;C, 5% CO2, humidity controlled)</li>
	<li>German Desag glass coverslips and/or plastic tissue culture dishes</li>
	<li>Poly-L-lysine, Sigma</li>
	<li>Sterile milliQ H2O</li>
	<li>Neurobasal, Invitrogen</li>
	<li>Heat-inactivated Fetal Bovine Serum (HiFBS)</li>
	<li>L-Glutamine</li>
	<li>B27, Invitrogen</li>
	<li>Penicillin/Streptomycin antibiotics</li>
	<li>37&deg;C waterbath</li>
	<li>Sterile Pasteur Pipettes</li>
	<li>Bunsen gas burner</li>
	<li>HBSS, Ca2+/Mg2+-free, Invitrogen</li>
	<li>DNAse, Worthington</li>
	<li>MgSO4</li>
	<li>Centrifuge</li>
	<li>Hemacytometer</li>
	<li>Trypan Blue Solution</li>
</ul>		</div>

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.

Watch this Scientific Journal Video about Dissection and Culture of Commissural Neurons from Embryonic Spinal Cord at JoVE.com

Dissection and Culture of Commissural Neurons from Embryonic Spinal Cord

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 ...

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Neuroscience

16.8K Aufrufe.  Institut de Recherches Cliniques de Montréal. Dieses Video zeigt eine Methode zu sezieren und Kultur kommissuralen Neuronen aus E13 Ratte Rückenmark. Dissoziiert kommissuralen Neuronen sind nützlich, um die zellulären und molekularen Mechanismen der Axon-Wachstum und Anleitung zu studieren.

Serie: Dissection and Culture of Commissural Neurons from Embryonic Spinal Cord

Nucleofection and Primary Culture of Embryonic Mouse Hippocampal and Cortical Neurons

Hippocampal and cortical neurons have been used extensively to study central nervous system (CNS) neuronal polarization, axon/dendrite outgrowth, and synapse formation and function. An advantage of culturing these neurons is that they readily polarize, forming distinctive axons and dendrites, on a two dimensional substrate at very low densities. This property has made them extremely useful for determining many aspects of neuronal development. Furthermore, by providing glial conditioning for these neurons they will continue to develop, forming functional synaptic connections and surviving for several months in culture. In this protocol we outline a technique to dissect, culture and transfect embryonic mouse hippocampal and cortical neurons. Transfection is accomplished by electroporating DNA into the neurons before plating via nucleofection. This protocol has the advantage of expressing fluorescently-tagged fusion proteins early in development (~4-8hrs after plating) to study the dynamics and function of proteins during polarization, axon outgrowth and branching. We have also discovered that this single transfection before plating maintains fluorescently-tagged fusion protein expression at levels appropriate for imaging throughout the lifetime of the neuron (&gt; 2 months in culture). Thus, this methodology is useful for studying protein localization and function throughout CNS development with little or no disruption of neuronal function.

This protocol outlines the steps required to dissect, transfect via electroporation and culture mouse hippocampal and cortical neurons. Short-term cultures may be used for studies of axon outgrowth and guidance, while long-term cultures can be used for studies of synaptogenesis and dendritic spine analysis.

Hippocampal and cortical neurons have been used extensively to study central nervous system (CNS) neuronal polarization, axon/dendrite outgrowth, and ...

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.

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 ...

DiI-Labeling of DRG Neurons to Study Axonal Branching in a Whole Mount Preparation of Mouse Embryonic Spinal Cord

Here we present a technique to label the trajectories of small groups of DRG neurons into the embryonic spinal cord by diffusive staining using the lipophilic tracer 1,1'-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine perchlorate (DiI)1. The comparison of axonal pathways of wild-type with those of mouse lines in which genes are mutated allows testing for a functional role of candidate proteins in the control of axonal branching which is an essential mechanism in the wiring of the nervous system. Axonal branching enables an individual neuron to connect with multiple targets, thereby providing the physical basis for the parallel processing of information. Ramifications at intermediate target regions of axonal growth may be distinguished from terminal arborization. Furthermore, different modes of axonal branch formation may be classified depending on whether branching results from the activities of the growth cone (splitting or delayed branching) or from the budding of collaterals from the axon shaft in a process called interstitial branching2 (Fig. 1).
The central projections of neurons from the DRG offer a useful experimental system to study both types of axonal branching: when their afferent axons reach the dorsal root entry zone (DREZ) of the spinal cord between embryonic days 10 to 13 (E10 - E13) they display a stereotyped pattern of T- or Y-shaped bifurcation. The two resulting daughter axons then proceed in rostral or caudal directions, respectively, at the dorsolateral margin of the cord and only after a waiting period collaterals sprout from these stem axons to penetrate the gray matter (interstitial branching) and project to relay neurons in specific laminae of the spinal cord where they further arborize (terminal branching)3. DiI tracings have revealed growth cones at the dorsal root entry zone of the spinal cord that appeared to be in the process of splitting suggesting that bifurcation is caused by splitting of the growth cone itself4 (Fig. 2), however, other options have been discussed as well5.
This video demonstrates first how to dissect the spinal cord of E12.5 mice leaving the DRG attached. Following fixation of the specimen tiny amounts of DiI are applied to DRG using glass needles pulled from capillary tubes. After an incubation step, the labeled spinal cord is mounted as an inverted open-book preparation to analyze individual axons using fluorescence microscopy.

The stereotyped projections of sensory afferents into the rodent spinal cord offer an easily accessible experimental system to study axonal branching through the tracing of single axons.

Here we  present a technique to label the trajectories of small groups of DRG neurons  into the embryonic spinal cord by diffusive staining using the ...

Dissection, Culture, and Analysis of Xenopus laevis Embryonic Retinal Tissue

The process by which the anterior region of the neural plate gives rise to the vertebrate retina continues to be a major focus of both clinical and basic research. In addition to the obvious medical relevance for understanding and treating retinal disease, the development of the vertebrate retina continues to serve as an important and elegant model system for understanding neuronal cell type determination and differentiation1-16. The neural retina consists of six discrete cell types (ganglion, amacrine, horizontal, photoreceptors, bipolar cells, and M&uuml;ller glial cells) arranged in stereotypical layers, a pattern that is largely conserved among all vertebrates 12,14-18.
While studying the retina in the intact developing embryo is clearly required for understanding how this complex organ develops from a protrusion of the forebrain into a layered structure, there are many questions that benefit from employing approaches using primary cell culture of presumptive retinal cells 7,19-23. For example, analyzing cells from tissues removed and dissociated at different stages allows one to discern the state of specification of individual cells at different developmental stages, that is, the fate of the cells in the absence of interactions with neighboring tissues 8,19-22,24-33. Primary cell culture also allows the investigator to treat the culture with specific reagents and analyze the results on a single cell level 5,8,21,24,27-30,33-39. Xenopus laevis, a classic model system for the study of early neural development 19,27,29,31-32,40-42, serves as a particularly suitable system for retinal primary cell culture 10,38,43-45.
Presumptive retinal tissue is accessible from the earliest stages of development, immediately following neural induction 25,38,43. In addition, given that each cell in the embryo contains a supply of yolk, retinal cells can be cultured in a very simple defined media consisting of a buffered salt solution, thus removing the confounding effects of incubation or other sera-based products 10,24,44-45.
However, the isolation of the retinal tissue from surrounding tissues and the subsequent processing is challenging. Here, we present a method for the dissection and dissociation of retinal cells in Xenopus laevis that will be used to prepare primary cell cultures that will, in turn, be analyzed for calcium activity and gene expression at the resolution of single cells. While the topic presented in this paper is the analysis of spontaneous calcium transients, the technique is broadly applicable to a wide array of research questions and approaches (Figure 1).

Dissection, Culture, and Analysis of Xenopus laevis Embryonic Retinal Tissue

Xenopus laevis provides an ideal model system for studying cell fate specification and physiological function of individual retinal cells in primary cell culture. Here we present a technique for dissecting retinal tissues and generating primary cell cultures that are imaged for calcium activity and analyzed by in situ hybridization.

The process by which  the anterior region of the neural plate gives rise to the vertebrate retina  continues to be a major focus of both clinical and ...

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 ...

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 ...

Isolation, Culture and Long-Term Maintenance of Primary Mesencephalic Dopaminergic Neurons From Embryonic Rodent Brains

Degeneration of mesencephalic dopaminergic (mesDA) neurons is the pathological hallmark of Parkinson&#8217;s diseae. Study of the biological processes involved in physiological functions and vulnerability and death of these neurons is imparative to understanding the underlying causes and unraveling the cure for this common neurodegenerative disorder. Primary cultures of mesDA neurons provide a tool for investigation of the molecular, biochemical and electrophysiological properties, in order to understand the development, long-term survival and degeneration of these neurons during the course of disease. Here we present a detailed method for the isolation, culturing and maintenance of midbrain dopaminergic neurons from E12.5 mouse (or E14.5 rat) embryos. Optimized cell culture conditions in this protocol result in presence of axonal and dendritic projections, synaptic connections and other neuronal morphological properties, which make the cultures suitable for study of the physiological, cell biological and molecular characteristics of this neuronal population.

The causes of degeneration of midbrain dopaminergic neurons during Parkinson&#8217;s disease are not fully understood. Cellular culture systems provide an essential tool for study of the neurophysiological properties of these neurons. Here we present an optimized protocol, which can be utilized for in vitro modeling of neurodegeneration.

Degeneration of mesencephalic dopaminergic (mesDA) neurons is the pathological hallmark of Parkinson&#8217;s diseae. Study of the biological processes ...

Zebrafish In Situ Spinal Cord Preparation for Electrophysiological Recordings from Spinal Sensory and Motor Neurons

Zebrafish, first introduced as a developmental model, have gained popularity in many other fields. The ease of rearing large numbers of rapidly developing organisms, combined with the embryonic optical clarity, served as initial compelling attributes of this model. Over the past two decades, the success of this model has been further propelled by its amenability to large-scale mutagenesis screens and by the ease of transgenesis. More recently, gene-editing approaches have extended the power of the model.
For neurodevelopmental studies, the zebrafish embryo and larva provide a model to which multiple methods can be applied. Here, we focus on methods that allow the study of an essential property of neurons, electrical excitability. Our preparation for the electrophysiological study of zebrafish spinal neurons involves the use of veterinarian suture glue to secure the preparation to a recording chamber. Alternative methods for recording from zebrafish embryos and larvae involve the attachment of the preparation to the chamber using a fine tungsten pin1,2,3,4,5. A tungsten pin is most often used to mount the preparation in a lateral orientation, although it has been used to mount larvae dorsal-side up4. The suture glue has been used to mount embryos and larvae in both orientations. Using the glue, a minimal dissection can be performed, allowing access to spinal neurons without the use of an enzymatic treatment, thereby avoiding any resultant damage. However, for larvae, it is necessary to apply a brief enzyme treatment to remove the muscle tissue surrounding the spinal cord. The methods described here have been used to study the intrinsic electrical properties of motor neurons, interneurons, and sensory neurons at several developmental stages6,7,8,9.

Zebrafish In Situ Spinal Cord Preparation for Electrophysiological Recordings from Spinal Sensory and Motor Neurons

This manuscript describes methods for electrophysiological recordings from spinal neurons of zebrafish embryos and larvae. The preparation maintains neurons in situ and often involves minimum dissection. These methods allow for the electrophysiological study of a variety of spinal neurons, from the initial electrical excitability acquisition through the early larval stages.

Zebrafish, first introduced as a developmental model, have gained popularity in many other fields. The ease of rearing large numbers of rapidly developing ...

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 ...