Skidmore Faculty Development Grant funded the preparation and publication of this manuscript.

1. Mounting Embryos&#160;(Upon Completion of Selected Visualization Technique, such as Immunocytochemistry, in situ&#160;Hybridization,&#160;etc.)
<ol>
	<li>Place embryos in a 35 mm Petri dish filled with buffer of choice (Figure 1A). The choice of buffer is based on the labeling procedure. Typically PBS-based buffers are suitable.</li>
	<li>Under a dissecting microscope, and using forceps, secure the head with one pair of forceps while pulling the yolk away with the other pair (Figure 1B). Alternatively, use an insect pin in an insect pin holder to gently pull away the yolk.</li>
	<li>Using the embryo poker (fishing line (0.41 mm diameter) glued to either a capillary tube or a Pasteur pipette) move the embryos to a part of the Petri dish that does not have a lot of yolk debris (Figures 1C-D).</li>
	<li>Using a glass Pasteur pipette, aspirate the embryos in as little liquid as possible, and gently pipette them onto a slide (Figure 1E).</li>
	<li>Gently wick away excess liquid using a laboratory wipe. Avoid contact with embryos.</li>
	<li>Add one drop of mounting medium to embryos. For fluorescent labels, use an antifade reagent of choice. For colorimetric signals, 70% glycerol may be used as a mounting medium.</li>
	<li>Using the embryo poker, orient the embryos on their sides in rows (Figure 1F).</li>
	<li>Add one &#34;dab&#34; of petroleum jelly or high vacuum grease to each corner of a coverslip (Figure 1G). This prevents damage of embryo from excessive compression.</li>
	<li>Place coverslip (petroleum jelly side down) on top of embryos (Figure 1H).</li>
	<li>Gently tap the coverslip on each corner until the mounting media (70% glycerol or anti-fade mounting reagent) is touching the coverslip (Figure 1H).</li>
	<li>If necessary, add more mounting media by pipetting right next to the coverslip, using a pipettor in the range of 20-200 ml. It will wick under.</li>
	<li>Using a laboratory wipe, completely clean around the coverslip. Be certain to not apply pressure to the coverslip, as this will damage embryos. This area must be dry and have no petroleum jelly or mounting media on it.</li>
	<li>Use nail polish to seal around the edge of the coverslip (Figure 1I).</li>
</ol>

<ol>
	<li>Gribble, S. L., Nikolaus, O. B., Dorsky, R. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Regulation+and+function+of+Dbx+genes+in+the+zebrafish+spinal.">Regulation and function of Dbx genes in the zebrafish spinal.</a> Dev. Dyn. 236 (12), 3472-3483 (2007).</li><li>Bonner, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Midline+crossing+is+not+required+for+subsequent+pathfinding+decisions+in+commissural+neurons.">Midline crossing is not required for subsequent pathfinding decisions in commissural neurons.</a> Neural Dev. 7 (1), 18 (2012).</li><li>Ross, A. B. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Activation+of+Wnt+signaling+using+Lithium+Chloride:+Inquiry-Based+Undergraduate+Laboratory+Exercises.">Activation of Wnt signaling using Lithium Chloride: Inquiry-Based Undergraduate Laboratory Exercises.</a> Zebrafish. , (2012).</li><li>Bonner, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Proliferation+and+patterning+are+mediated+independently+in+the+dorsal+spinal+cord+downstream+of+canonical+Wnt+signaling.">Proliferation and patterning are mediated independently in the dorsal spinal cord downstream of canonical Wnt signaling.</a> Dev. Biol. 313 (1), 398-407 (2008).</li><li>Gribble, S. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tcf3+inhibits+spinal+cord+neurogenesis+by+regulating+sox4a+expression.">Tcf3 inhibits spinal cord neurogenesis by regulating sox4a expression.</a> Development. 136 (5), 781-789 (2009).</li><li>England, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Roles+of+Hedgehog+pathway+components+and+retinoic+acid+signalling+in+specifying+zebrafish+ventral+spinal+cord+neurons.">Roles of Hedgehog pathway components and retinoic acid signalling in specifying zebrafish ventral spinal cord neurons.</a> Development. 138 (23), 5121-5134 (2011).</li><li>Challa, A. K., Beattie, C. E., Seeger, 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=Identification+and+characterization+of+roundabout+orthologs+in+zebrafish.">Identification and characterization of roundabout orthologs in zebrafish.</a> Mech Dev. (1-2), 101-101 (2001).</li><li>Lee, J. S., Ray, R., Chien, C. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cloning+and+expression+of+three+zebrafish+roundabout+homologs+suggest+roles+in+axon+guidance+and+cell.">Cloning and expression of three zebrafish roundabout homologs suggest roles in axon guidance and cell.</a> 221 (2), 216-230 (2001).</li><li>Downes, G. B., Waterbury, J. A., Granato, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rapid+in+vivo+labeling+of+identified+zebrafish+neurons.">Rapid in vivo labeling of identified zebrafish neurons.</a> Genesis. 34 (3), 196-202 (2002).</li><li>Kim, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Notch-regulated+oligodendrocyte+specification+from+radial+glia+in+the+spinal+cord+of+zebrafish+embryos.">Notch-regulated oligodendrocyte specification from radial glia in the spinal cord of zebrafish embryos.</a> Dev. Dyn. 237 (8), 2081-2089 (2008).</li><li>Sylvain, N. J., Brewster, D. L., Ali, D. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Zebrafish+embryos+exposed+to+alcohol+undergo+abnormal+development+of+motor+neurons+and+muscle+fibers.">Zebrafish embryos exposed to alcohol undergo abnormal development of motor neurons and muscle fibers.</a> Neurotoxicol. Teratol. 32 (4), 472-480 (2010).</li><li>de Soysa, T. Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Macondo+crude+oil+from+the+Deepwater+Horizon+oil+spill+disrupts+specific+developmental+processes+during+zebrafish+embryogenesis.">Macondo crude oil from the Deepwater Horizon oil spill disrupts specific developmental processes during zebrafish embryogenesis.</a> BMC Biol. 10 (40), (2012).</li><li>Langenberg, T., Brand, M., Cooper, M. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+brain+development+and+organogenesis+in+zebrafish+using+immobilized+embryonic+explants.">Imaging brain development and organogenesis in zebrafish using immobilized embryonic explants.</a> Dev Dyn. 228 (3), 464-474 (2003).</li><li>Deschene, E. R., Barresi, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tissue+Targeted+Embryonic+Chimeras:+Zebrafish+Gastrula+Cell+Transplantation.">Tissue Targeted Embryonic Chimeras: Zebrafish Gastrula Cell Transplantation.</a> J. Vis. Exp. (31), (2009).</li></ol>

No competing financial interests exist for either author.

Lateral mounting of fixed zebrafish embryos aids visualization of spinal cord development. Through removal of the yolk ball, the working distance required for exceptional microscopic imaging is reduced. Our representative results were limited to the 24 hpf stage, however, this technique can be used as early as 18 hpf, though earlier embryos are more difficult to dissect due to the size of the embryo and the fragility of the tissue. This technique is also applicable to embryos older than 24 hpf. Aided by the optical clarity of the embryonic zebrafish, the ability to perform forward genetic screens, reverse genetics, and transgenic approaches, several developmental processes may be investigated. This includes mitotic indices of neuronal progenitors with markers like BrdU and anti-phospho-histone 3 4-6&#160;and patterning through expression of neuronal and glial progenitor markers in the pax, nkx, dbx, and olig families1,4-6. Reagents that report glial and neuronal determination (such as glial fibrillary acid protein (GFAP) and HuC/D)1,4,5,10&#160;can be used. Neuronal subtype differentiation can be analyzed through expression of islet, vsx, engrailed, and gata genes1,4-6. And finally, analysis of axon guidance is possible through antibodies such as anti-acetylated tubulin, 3A10, and znp-1 2,11,12. Though other vertebrate model systems (mouse and chick) are used to study spinal cord development, only the zebrafish permits simultaneous viewing of all developmental axes in the intact embryo.
The obvious limitation to this approach is that the embryos are fixed, which precludes live imaging. In fact, removal of (or damage to) the yolk results in rapid embryonic death, thus it is not possible to reduce the working distance in live embryos using this technique. However, high magnification spinal cord imaging in live embryos is possible. In order to preserve the yolk during live imaging, embryos can be placed in depression slides, which provide a larger space between the specimen and the coverslip. Alternatively, several 22 mm x&#160;22 mm coverslips can be glued on either slide of a 3 in x&#160;1 in&#160;microscopic slide. The coverslips serve as a &#34;bridge&#34; for the overlying coverslip. If embryos are older than 18 hpf, tricaine is used to anesthetize embryos to prevent movement. While a working distance of 0.288 mm is required for deyolked embryos at 24 hpf, when working with live embryos embedded in agarose, with intact yolks, a microscopic lens with a working distance of 3.3 mm is sufficient. Alternatively, explant cultures derived from deyolked embryos may be visualized using a plasma clot immobilization technique13. Various populations of cells can be observed through the use of genetically encoded fluorescent proteins such as GFP, mCherry, or the photoconvertible dye Kaede (to name a few). Further, fluorescently labeled cells within chimeric embryos can also be viewed with this approach14.
A consistent mounting technique that is stable for months is a critical tool for developmental and cell biologists. This straightforward technique is reproducible, allowing easy comparison between different experimental trials. Furthermore, the alignment of embryos in rows readily permits identification of specific embryos for later analysis.

Visualization of the spinal cord in zebrafish is inhibited by a number of factors. Due to the thickness of the overlying somites&#160;and the internal location of the spinal cord, a considerably long working distance is required for high cellular resolution. The yolk ball (which is still present during early stages of neurogenesis) further increases the working distance required, and is easily damaged by pressure from a coverslip. In addition, debris from damaged yolk prohibits clear visualization of tissues. Though cross sections in the dorso-ventral (DV) axis are possible, they do not readily permit simultaneous visualization in the anterior-posterior (AP) axis1.
To overcome these obstacles, embryos are dissected and mounted on slides. This procedure provides several benefits. First, zebrafish embryos readily lie with the lateral side facing upwards, which facilitates AP axis visualization. Second, removal of the yolk ball decreases the required working distance, and limits debris. Third, this mounting procedure allows for both fluorescent and brightfield microscopy. Fourth, mounted embryos are stable for months at 4 &#176;C, allowing prolonged specimen visualization. Finally, progression of development occurs in that anterior segments are more mature than posterior segments. In order to ensure that staged matched spinal hemisegments are compared between embryos, overlying somite tissue is used as a guide. For example, the tenth most posterior somite overlies the tenth spinal hemisegment, which is developmentally equivalent in stage-matched embryos. This mounting procedure of intact embryos allows easy identification of somites.
We use genetics and gene knock-down technologies to study the mechanisms of axon guidance in the developing spinal cord. In particular, we have determined that robo2, robo3, and dcc are required for Commissural Primary Ascending (CoPA) axon pathfinding. Using the mounting procedure described here, we were able to examine ventral growth, midline crossing, commissure width, dorsal and anterior growth2,3. This mounting procedure may also be applied to DV or AP patterning of the spinal cord. We have used this procedure to determine disparate roles of downstream Wnt effectors in DV patterning of the spinal cord. Using this mounting, we were able to obtain resolution of DV markers at the single cell level, and determine minute patterning shifts as a result of altered Wnt signal reception4,5. This mounting procedure also allows for mitotic index calculation through anti-phosphohistone 3 or BrdU labeling (progenitor proliferation) differentiation5.

<table border="1" cellpadding="0" cellspacing="0">
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		<tr>
			<td style="width:132px;">
			Petri dishes 35mm X 10mm
			</td>
			<td style="width:115px;">
			VWR
			</td>
			<td style="width:116px;">
			25373-041
			</td>
		</tr>
		<tr>
			<td style="width:132px;">
			Dumont Forceps #3
			</td>
			<td style="width:115px;">
			Fisher Scientific
			</td>
			<td style="width:116px;">
			NC9839169
			</td>
		</tr>
		<tr>
			<td style="width:132px;">
			Cover glass
			</td>
			<td style="width:115px;">
			Fisher Scientific
			</td>
			<td style="width:116px;">
			12-541-B22X22-1.5
			</td>
		</tr>
		<tr>
			<td style="width:132px;">
			Slides
			</td>
			<td style="width:115px;">
			Fisher Scientific
			</td>
			<td style="width:116px;">
			12-550-343
			</td>
		</tr>
		<tr>
			<td style="width:132px;">
			SlowFade Gold
			</td>
			<td style="width:115px;">
			Fisher Scientific
			</td>
			<td style="width:116px;">
			S36936
			</td>
		</tr>
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			<td style="width:132px;">
			ProLong Gold
			</td>
			<td style="width:115px;">
			Life Technologies
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			<td style="width:116px;">
			P36934
			</td>
		</tr>
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			<td style="width:132px;">
			Petroleum Jelly
			</td>
			<td style="width:115px;">
			Any grocery store
			</td>
			<td style="width:116px;">
			 
			</td>
		</tr>
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			<td style="width:132px;">
			Loop Holders
			</td>
			<td style="width:115px;">
			VWR
			</td>
			<td style="width:116px;">
			80094-482
			</td>
		</tr>
		<tr>
			<td style="width:132px;">
			Insect Pins
			</td>
			<td style="width:115px;">
			Fine Science Tools
			</td>
			<td style="width:116px;">
			26002-10
			</td>
		</tr>
		<tr>
			<td style="width:132px;">
			Nickel Plated Pin Holders
			</td>
			<td style="width:115px;">
			Fine Science Tools
			</td>
			<td style="width:116px;">
			26016-12
			 
			</td>
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	</tbody>
</table>
 
 
 
 
 
 
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			Name of Equipment
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			Company
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			Catalog number
			</td>
		</tr>
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			<td style="width:118px;">
			Olympus Stereo microscope
			</td>
			<td style="width:118px;">
			Olympus
			</td>
			<td style="width:118px;">
			SZ61
			</td>
		</tr>
	</tbody>
</table>

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.

When elucidating the mechanisms that underlie various developmental events such as cellular patterning, differentiation, and axon guidance, it is important to be able to visualize cells within the context of their tissue. Dorsoventral patterning defects typically present as a ventral or dorsal shift in the expression domain of transcription factors in the pax, nkx, or dbx families4. At times, changes may be subtle, comprising only a few cell diameters4. Cross sectional analysis permits this type of analysis. However, reagents with weaker signals, or cross sections that are not perpendicular to the AP axis can limit interpretation. Further, cross sectional analysis does not readily take into account whether changes persist in the AP axis, and relies on the availability of a cryostat. This limitation is circumvented by lateral views of the spinal cord. As seen in Figure 2B, nkx6.1 mRNA is distributed in roughly the ventral half of the spinal cord at 24 hr post-fertilization (hpf), within the progenitor domain. Progenitor cells are distinguishable from post mitotic cells due to presence of the floor plate, which is found in the medial part of the spinal cord. Viewed with DIC optics, individual cells are clearly distinguishable, and a defined boundary between expressing and nonexpressing cells is evident.
Progenitor cell cycle exit is accompanied by changes in gene expression. For example, all post-mitotic neurons express HuC/D which is detectable with immunocytochemistry1,5. mRNA probes for islet, gata, and vsx families label specific post-mitotic neurons of consistent position and number within each spinal cord hemisegment6. In addition, axon guidance receptors such as those in the robo family are expressed in restricted populations of postmitotic neurons (Figure 2C)2,7,8. Lateral mounting of embryos allows accurate counts of post-mitotic neurons. Similarly, progenitors that continue to divide can be quantified with anti-phospho-histone 3 immunocytochemistry as well as BrdU labeling. Furthermore, cell death can be assessed with TUNEL labeling4-6.
At 24 hpf, axons of the following postmitotic neurons can be visualized with various methods, and are distinguishable at the single cell level: Dorsal Lateral Ascending (DoLA), Commissural Primary Ascending (CoPA), Commissural Secondary Ascending (CoSA), Ventral Longitudinal Descending (VeLD), Kolmer-Agdur (KA), Commissural Bifurcating/Longitudinal (CoB/L), Circumferential Ascending (CiA), Circumferential Descending (CiD), and Unipolar Commissural Descending (UCoD), Rohon-Beard (R-B), motoneurons (M)9. Axons extend from these neurons in the dorsal, ventral, anterior, and posterior directions. They branch, cross the midline, and exit both the dorsal and ventral spinal cord. When coupled with confocal laser scanning microscopy, these varied cellular behaviors are evident in laterally mounted embryos, using immunocytochemistry and genetically encoded fluorescent proteins such as GFP2. In Figure 2D, znp-1 immunocytochemistry was used to label developing motoneurons. Motoneurons exit the spinal cord ventrally to innervate the surrounding developing musculature.
<img alt="Figure 1" fo:content-width="6in" fo:src="/files/ftp_upload/50703/50703fig1highres.jpg" src="/files/ftp_upload/50703/50703fig1.jpg" width="500px" /> 
Figure 1.&#160;Dissection and lateral mount of zebrafish embryos.&#160;(A) After processing for immunofluorescence, in situ hybridization, etc., embryos are placed in a 35 mm Petri dish filled with PBT (PBS with 0.5% Triton X-100) or PTw (PBS with 0.1% Tween-20). The yolk ball (YB) is evident. (B) The yolk is removed by securing the head of the embryo followed by careful dissection. (C) An embryo poker (fishing line glued to a Pasteur pipette) is used to separate embryos and yolk debris (D). Cleaned embryos are pipetted onto a microscope slide (E) and aligned (F). (G) Petroleum jelly is applied to the corners of the coverslip. (H) The coverslip is placed gently on top of the embryos in mounting medium. (I) Nail polish is used to seal the edges of the coverslip.&#160;<a href="https://www.jove.com/files/ftp_upload/50703/50703fig1highres.jpg" target="_blank">Click here to view larger image</a>.
<img alt="Figure 2" fo:content-width="3.5in" fo:src="/files/ftp_upload/50703/50703fig2highres.jpg" src="/files/ftp_upload/50703/50703fig2.jpg" width="500px" /> 
Figure 2.&#160;Analysis of gene expression patterns and axon pathfinding in laterally mounted zebrafish embryos. (A) Drawing of a zebrafish embryo at 24 hpf. In B-D, roughly four hemisegments above the yolk sac extension are visualized (boxed area). (B) nkx6.1 is expressed in the ventral spinal cord, including the floor plate (FP). (C) robo3 is expressed in postmitotic neurons (PMN). The floor plate and progenitor zone is not evident in this more lateral focal plane. In B,C, the spinal cord is bounded by a bracket on the left hand side of the image. (D) Confocal microscopy was used to image znp-1 immunofluorescence, which labels motor axons exiting the spinal cord ventrally. In all images, anterior is to the left and dorsal is up. A 40X long working distance, water immersion (N.A. 0.8) lens was used (3.3 mm). <a href="https://www.jove.com/files/ftp_upload/50703/50703fig2highres.jpg" target="_blank">Click here to view larger image</a>.

Watch this Scientific Journal Video about Dissection and Lateral Mounting of Zebrafish Embryos: Analysis of Spinal Cord Development at JoVE.com

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

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 ...

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Research

JoVE Journal

Neuroscience

13.8K Views.  Skidmore College.  Processi di sviluppo quali la proliferazione, il modello, la differenziazione, e la guida degli assoni possono essere facilmente modellati nel midollo spinale zebrafish. In questo articolo, si descrive una procedura di montaggio per gli embrioni di zebrafish, che ottimizza la visualizzazione di questi eventi. 

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

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 ...

Lateral Fluid Percussion: Model of Traumatic Brain Injury in Mice

Traumatic brain injury (TBI) research has attained renewed momentum due to the increasing awareness of head injuries, which result in morbidity and mortality. Based on the nature of primary injury following TBI, complex and heterogeneous secondary consequences result, which are followed by regenerative processes 1,2. Primary injury can be induced by a direct contusion to the brain from skull fracture or from shearing and stretching of tissue causing displacement of brain due to movement 3,4. The resulting hematomas and lacerations cause a vascular response 3,5, and the morphological and functional damage of the white matter leads to diffuse axonal injury 6-8. Additional secondary changes commonly seen in the brain are edema and increased intracranial pressure 9. Following TBI there are microscopic alterations in biochemical and physiological pathways involving the release of excitotoxic neurotransmitters, immune mediators and oxygen radicals 10-12, which ultimately result in long-term neurological disabilities 13,14. Thus choosing appropriate animal models of TBI that present similar cellular and molecular events in human and rodent TBI is critical for studying the mechanisms underlying injury and repair.
Various experimental models of TBI have been developed to reproduce aspects of TBI observed in humans, among them three specific models are widely adapted for rodents: fluid percussion, cortical impact and weight drop/impact acceleration 1. The fluid percussion device produces an injury through a craniectomy by applying a brief fluid pressure pulse on to the intact dura. The pulse is created by a pendulum striking the piston of a reservoir of fluid. The percussion produces brief displacement and deformation of neural tissue 1,15. Conversely, cortical impact injury delivers mechanical energy to the intact dura via a rigid impactor under pneumatic pressure 16,17. The weight drop/impact model is characterized by the fall of a rod with a specific mass on the closed skull 18. Among the TBI models, LFP is the most established and commonly used model to evaluate mixed focal and diffuse brain injury 19. It is reproducible and is standardized to allow for the manipulation of injury parameters. LFP recapitulates injuries observed in humans, thus rendering it clinically relevant, and allows for exploration of novel therapeutics for clinical translation 20.
We describe the detailed protocol to perform LFP procedure in mice. The injury inflicted is mild to moderate, with brain regions such as cortex, hippocampus and corpus callosum being most vulnerable. Hippocampal and motor learning tasks are explored following LFP.

Lateral fluid percussion (LFP), an established model of traumatic brain injury in mice, is demonstrated. LFP fulfills three major criteria for animal models: validity, reliability and clinical relevance. The procedure, consisting of surgical craniotomy, fixation of hub followed by induction of injury, resulting in focal and diffuse injuries, is described.

Traumatic brain injury (TBI) research has attained renewed momentum due to the increasing awareness of head injuries, which result in morbidity 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 ...

Spinal Cord Transection in the Larval Zebrafish

Mammals fail in sensory and motor recovery following spinal cord injury due to lack of axonal regrowth below the level of injury as well as an inability to reinitiate spinal neurogenesis. However, some anamniotes including the zebrafish Danio rerio exhibit both sensory and functional recovery even after complete transection of the spinal cord. The adult zebrafish is an established model organism for studying regeneration following spinal cord injury, with sensory and motor recovery by 6&#160;weeks post-injury. To take advantage of in vivo analysis of the regenerative process available in the transparent larval zebrafish as well as genetic tools not accessible in the adult, we use the larval zebrafish to study regeneration after spinal cord transection. Here we demonstrate a method for reproducibly and verifiably transecting the larval spinal cord. After transection, our data shows sensory recovery beginning at 2&#160;days post-injury (dpi), with the C-bend movement detectable by 3 dpi and resumption of free swimming by 5 dpi. Thus we propose the larval zebrafish as a companion tool to the adult zebrafish for the study of recovery after spinal cord injury.

After spinal transection, adult zebrafish have functional recovery by six weeks post-injury. To take advantage of larval transparency and faster recovery, we present a method for transecting the larval spinal cord. After transection, we observe sensory recovery beginning at 2&#160;days post-injury, and C-bend movement by 3 days post-injury.

Mammals fail in sensory and motor recovery following spinal cord injury due to lack of axonal regrowth below the level of injury as well as an inability ...

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 ...

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 ...

Isolation of Adult Spinal Cord Nuclei for Massively Parallel Single-nucleus RNA Sequencing

Probing an individual cell&#39;s gene expression enables the identification of cell type and cell state. Single-cell RNA sequencing has emerged as a powerful tool for studying transcriptional profiles of cells, particularly in heterogeneous tissues such as the central nervous system. However, dissociation methods required for single cell sequencing can lead to experimental changes in the gene expression and cell death. Furthermore, these methods are generally restricted to fresh tissue, thus limiting studies on archival and bio-bank material. Single nucleus RNA sequencing (snRNA-Seq) is an appealing alternative for transcriptional studies, given that it accurately identifies cell types, permits the study of tissue that is frozen or difficult to dissociate, and reduces dissociation-induced transcription. Here, we present a high-throughput protocol for rapid isolation of nuclei for downstream snRNA-Seq. This method enables isolation of nuclei from fresh or frozen spinal cord samples and can be combined with two massively parallel droplet encapsulation platforms.

Here, we present a protocol to rapidly isolate high-quality nuclei from the fresh or frozen tissue for downstream massively parallel RNA sequencing. We include detergent-mechanical and hypotonic-mechanical tissue disruption and cell lysis options, both of which can be used for isolation of nuclei.

Probing an individual cell&#39;s gene expression enables the identification of cell type and cell state. Single-cell RNA sequencing has emerged as a ...

In Vivo Calcium Imaging of Lateral-line Hair Cells in Larval Zebrafish

Sensory hair cells are mechanoreceptors found in the inner ear that are required for hearing and balance. Hair cells are activated in response to sensory stimuli that mechanically deflect apical protrusions called hair bundles. Deflection opens mechanotransduction (MET) channels in hair bundles, leading to an influx of cations, including calcium. This cation influx depolarizes the cell and opens voltage-gated calcium channels located basally at the hair-cell presynapse. In mammals, hair cells are encased in bone, and it is challenging to functionally assess these activities in vivo. In contrast, larval zebrafish are transparent and possess an externally located lateral-line organ that contains hair cells. These hair cells are functionally and structurally similar to mammalian hair cells and can be functionally assessed in vivo. This article outlines a technique that utilizes a genetically encoded calcium indicator (GECI), GCaMP6s, to measure stimulus-evoked calcium signals in zebrafish lateral-line hair cells. GCaMP6s can be used, along with confocal imaging, to measure in vivo calcium signals at the apex and base of lateral-line hair cells. These signals provide a real-time, quantifiable readout of both mechanosensation- and presynapse-dependent calcium activities within these hair cells. These calcium signals also provide important functional information regarding how hair cells detect and transmit sensory stimuli. Overall, this technique generates useful data about relative changes in calcium activity in vivo. It is less well-suited for quantification of the absolute magnitude of calcium changes. This in vivo technique is sensitive to motion artifacts. A reasonable amount of practice and skill are required for proper positioning, immobilization, and stimulation of larvae. Ultimately, when properly executed, the protocol outlined in this article provides a powerful way to collect valuable information about the activity of hair-cells in their natural, fully integrated states within a live animal.

The zebrafish is a model system that has many valuable features including optical clarity, rapid external development, and, of particular importance to the field of hearing and balance, externally located sensory hair cells. This article outlines how transgenic zebrafish can be used to assay both hair-cell mechanosensation and presynaptic function in toto.

Sensory hair cells are mechanoreceptors found in the inner ear that are required for hearing and balance. Hair cells are activated in response to sensory ...

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 ...