The authors wish to thank Dr. Alistair Garratt (Max Delbr&uuml;ck Center, Berlin) for helpful comments. This work was supported by the collaborative research center (SFB665) of the German Research Council (DFG).

1. Dissection procedure
Note: Experimental use of mice should follow officially approved guidelines for the care and use of laboratory animals.
<ol>
	<li>Before preparation, set up your dissecting microscope and lay out the surgical instruments needed for dissection including large and small scissors, large toothed forceps, curved forceps and four sets of Dumont No.5 dissecting forceps (two of which have inside-polished tips) (for details see the table of specific reagents and equipment). Place a sheet of filter paper in a 100-mm Sylgard-coated Petri dish. Pour cold PBS in the wells of a 12-well plate, a 100-mm petri dish and a 12-ml tube and leave on ice. Pipette 2ml of fixation buffer (4% paraformaldehyde in PBS, pH 7.4) in each well of a second 12-well plate and place on ice.</li>
	<li>After anesthetization, sacrifice the timed pregnant dam at E12.5 (detection of vaginal plug was designated as E0.5), place the mouse with its ventral side up on three sheets of paper towels and soak the abdominal area with 70% ethanol.</li>
	<li>Open the abdominopelvic cavity, isolate the bilateral uterine horns and transfer them into one of the 100-mm dishes with ice-cold PBS.</li>
	<li>Under microscopic control, make a longitudinal incision of the uterine wall with small, straight scissors and cut away each amniotic sac from the placenta.</li>
	<li>Peel away the amniotic sac from each embryo and cut the umbilical cord.</li>
	<li>Using small scissors, decapitate the embryos and put the heads or a part of them aside for the isolation of genomic DNA if genotyping should be required. Subsequently, transfer the torsi to the respective wells of the 12-well plate with cold PBS.</li>
	<li>Wet the filter paper in the Sylgard dish with a few drops of PBS and position an embryonic torso with its dorsal side up on the paper. For better stability, straighten its tail and extremities away from the body.</li>
	<li>Working under a dissecting microscope with a magnification of approximately 16x, carefully pinch the skin of the embryo above the spinal cord with two pairs of fine tipped forceps and gently tear it apart. Beginning in the middle, proceed first towards the tail and then resuming from the middle towards the anterior side.</li>
	<li>Wet the embryo from time to time with two or three drops of PBS to prevent it from drying.</li>
	<li>To prevent that DRG are teared off from the spinal cord when it is removed from the embryo detach the DRG and the spinal cord from the surrounding cartilaginous vertebral column by horizontal sliding movements with the blade of a fine forceps with inside-polished tips. Starting in the middle of the embryo's right side, work your way towards the tail and then to the anterior end. Afterwards, repeat the procedure on the other side. Be careful not to rip off the associated DRG.</li>
	<li>To detach completely the loosened spinal cord from the embryo, pick up its cervical part with fine forceps and pull it out in one piece towards its caudal end.</li>
	<li>Submerge the isolated spinal cord with attached DRG in a well of the 12-well plate filled with fixation buffer and proceed with the preparation of spinal cords from the remaining embryos.</li>
	<li>Fix spinal cords at least two hours on ice.</li>
</ol>
2. DiI-labeling of DRG neurons
<ol>
	<li>For each spinal cord to be labeled, prepare two glass needles with a micropipette puller. We use a Sutter model P-97 electrode puller (programme setting: 1. HEAT=625, VEL= 35, TIME=175; 2. HEAT=600, VEL= 25, TIME=175; 3. HEAT=630, VEL= 30, TIME=150).</li>
	<li>Under microscopic control dip the tip of each glass needle three to five times in a 5 % (w/v) solution of DiI in ethanol. Evaporation of the ethanol creates a thin layer of DiI crystals on the needle.</li>
	<li>Working under a dissecting microscope with a magnification of approximately 40x, place a spinal cord with its ventral side up on a slide. Using spring scissors, open the cord on the ventral side at its whole length by cutting through the floorplate to allow the later mounting of the preparation in an inverted open-book mode (see section C).</li>
	<li>Position the spinal cord with the dorsal side up on a slide. Then manually approach the cord with the DiI-covered tip of the glass needle under microscopic visualization and carefully pierce every second DRG on one side of the spinal cord. Almost no traces of DiI should be visible in the pierced DRG to ensure the labeling of only a small number of neurons. Take a second needle for the other side of the cord. Aiming only every second DRG avoids an overlap of labeled axonal projections from neighboring DRG which might complicate the differentiation of individual axons.</li>
	<li>Return the spinal cord to the fixation buffer and incubate in the dark for at least six hours at ambient temperature or overnight at 4 &deg;C to give time for the dye to diffuse along the axon within the plasma membrane. Allow for a longer incubation period (up to two days at 4 &deg;C) if the analysis of collateral growth into the spinal cord is desired.</li>
</ol>
3. Mounting and microscopic analysis
<ol>
	<li>Following dye diffusion, position a single spinal cord with its dorsal side down on a coverslip in a drop of PBS. Take care that the attached DRG are correctly oriented laterally and do not intermingle. Aspirate excess fluid and flatten out the cord in an inverted open-book mode using forceps.</li>
	<li>Mount the preparation on a microscope slide using PBS.</li>
	<li>To prevent the increase of unwanted background staining fluorescence microscopic analysis of labeled axonal projections should be carried out the day of mounting. Until then keep the slides in the dark at 4 &deg;C.</li>
</ol>
4. Representative Results:
The spinal cord of the mouse receives afferent projections from 8 pairs of cervical, 13 pairs of thoracic, 5 pairs of lumbar and 4 pairs of sacral DRG totaling to 60 spinal ganglia. After some training a spinal cord with most DRG still attached can be isolated from an embryo in under five minutes. This procedure is suited for the isolation of spinal cords with attached DRG from E11.5 to E13.5 mouse embryos. However, best results are achieved from E12.5. Exemplary outcomes of the labeling procedure described here are shown in Figure 3. The labeling of DRG over the entire length of the spinal cord may then be used to quantify axonal branching behaviour at different vertebral levels (Fig. 4).
<img alt="Figure 1" src="/files/ftp_upload/3667/3667fig1.jpg" /> 
Figure 1. Scheme depicting the two principal modes of axonal branching. (A) Branching by the activities of the growth cone that could lead to a bifurcation - as indicated here - as well as complex terminal arbors or (B) collateral formation at the axon shaft (interstitial branching). The latter mode is the dominating branching type of projecting cortical and thalamocortical axons6,7.
<img alt="Figure 2" src="/files/ftp_upload/3667/3667fig2.jpg" /> 
Figure 2. Afferent projections of DRG neurons into the embryonic spinal cord display both types of axonal branching: axons first branch at the DREZ by bifurcation (1) and from the resulting daughter branches collaterals form after a waiting period by interstitial branching (2).
<img alt="Figure 3" src="/files/ftp_upload/3667/3667fig3.jpg" /> 
Figure 3. Visualization of single axonal trajectories of embryonic mouse DRG neurons. (A) Spinal cord with attached DRG prepared from an E12.5 mouse embryo. (Scale bar, 1 mm.) (B-D) Dorsal views of DiI-labeled DRG of wild-type mice at increasing magnification are shown. In B every second DRG is labeled by DiI. Fluorescence images are inverted, caudal is at the left and in C and D, lateral is at the bottom. In C a small number of axons is labeled and at higher magnification the presence of T-like branches can be identified in the DREZ of the spinal cord. (Scale bars, 250 &mu;m (B), 100 &mu;m (C) and 50 &mu;m (D).)
<img alt="Figure 4" src="/files/ftp_upload/3667/3667fig4.jpg" /> 
Figure 4. Quantification of T-shaped branches, single rostral or caudal turns in wild-type and C-type natriuretic peptide (CNP)-deficient mice at E13.5. The numbers of single axons counted are given in brackets for different trunk levels for each genotype. C- or R-turns - growth only in caudal or rostral direction, respectively.
<img alt="Figure 5" src="/files/ftp_upload/3667/3667fig5.jpg" /> 
Figure 5. A cGMP signaling pathway triggers sensory axon bifurcation at the DREZ of the spinal cord. (A) Scheme of the cGMP signaling pathway composed of the ligand CNP, the receptor guanylyl cyclase Npr2 and the serine/threonine kinase cGKI&alpha; in embryonic DRG neurons. Npr2 generates cGMP from GTP upon stimulation by CNP. (B) DiI tracing of single axons of DRG neurons in wild-type and CNP-deficient mice. (Scale bar, 25 &mu;m.)

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D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Initial+trajectories+of+sensory+axons+toward+laminar+targets+in+the+developing+mouse+spinal+cord.">Initial trajectories of sensory axons toward laminar targets in the developing mouse spinal cord.</a> J. Comp. Neurol. 380 (2), 215-215 (1997).</li><li>Schmidt, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+receptor+guanylyl+cyclase+Npr2+is+essential+for+sensory+axon+bifurcation+within+the+spinal+cord.">The receptor guanylyl cyclase Npr2 is essential for sensory axon bifurcation within the spinal cord.</a> J. Cell Biol. 179 (2), 331-331 (2007).</li><li>Gibson, D. 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The authors have nothing to disclose.

Stereotyped projection patterns comprising both types of axonal branch formation together with ease of preparation in combination with the use of fixed tissue for DiI labeling makes the embryonic spinal cord with attached DRG a favourable model to study axonal branching. The application of minute amounts of DiI using coated glass needles allows - in contrast to the bulk labeling of DRG - the visualization of small groups of DRG neurons and thereby the analysis of individual axons and their branching patterns. The describ...

<table border="1">
<tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments (optional)</td>
 </tr>
 <tr>
 <td>Stereomicroscope Stemi DRC</td>
 <td>Zeiss</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Phosphate-buffered solution (PBS)</td>
 <td>Biochrom AG</td>
 <td>L182-50</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Paraformaldehyde</td>
 <td>Merck</td>
 <td>8.18715.1000</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Standard surgical scissors</td>
 <td>Fine Science Tools</td>
 <td>14001-13</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Toothed standard forceps</td>
 <td>Fine Science Tools</td>
 <td>11021-14</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Extra fine iris scissors</td>
 <td>Fine Science Tools</td>
 <td>14088-10</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Curved forceps</td>
 <td>Fine Science Tools</td>
 <td>11003-13</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Dumont No.5 fine tips forceps</td>
 <td>Fine Science Tools</td>
 <td>11254-20</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Dumont No.5 mirror finish forceps</td>
 <td>Fine Science Tools</td>
 <td>11252-23</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Vannas-T&uuml;bingen spring scissors</td>
 <td>Fine Science Tools</td>
 <td>15008-08</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Filter paper</td>
 <td>Fisher Scientific</td>
 <td>FB59041</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Sylgard 184</td>
 <td>World Precission Instruments</td>
 <td>SYLG184</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>100-mm Petri dishes</td>
 <td>Greiner</td>
 <td>663102</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>12-ml polypropylene tube</td>
 <td>Carl Roth GmbH</td>
 <td>ECO3.1</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>12-well culture plate</td>
 <td>Becton Dickinson</td>
 <td>35-3043</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Ethanol </td>
 <td>Merck</td>
 <td>1.00983.2500</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Flaming/Brown micropipette puller P-97</td>
 <td>Sutter Instrument Co.</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Borosilicate glass capillaries</td>
 <td>Harvard Apparatus</td>
 <td>30-0066</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>DiI (1,1&prime;-Dioctadecyl-3,3,3&prime;,3&prime;-tetramethyl-indocarbocyanine perchlorate)</td>
 <td>Sigma-Aldrich</td>
 <td>468495</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Microscope slides SuperFrost Plus</td>
 <td>Carl Roth GmbH</td>
 <td>H867.1</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Glass cover slips</td>
 <td>Carl Roth GmbH</td>
 <td>1870.2</td>
 <td>&nbsp;</td>
 </tr>
</tbody>
</table>

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.

Watch this Scientific Journal Video about DiI-Labeling of DRG Neurons to Study Axonal Branching in a Whole Mount Preparation of Mouse Embryonic Spinal Cord at JoVE.com

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

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 ...

dii-labeling-drg-neurons-to-study-axonal-branching-whole-mount

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24.8K Views.  Max Delbrück Center for Molecular Medicine. 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.

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

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 ...

Visualization of the Embryonic Nervous System in Whole-mount Drosophila Embryos

The Drosophila embryo is an attractive model system for investigating the cellular and molecular basis of neuronal development. Here we describe the procedure for the visualization of Drosophila embryonic nervous system using antibodies to neuronal proteins. Since the entire embryonic peripheral nervous and central nervous systems are well characterized at the level of individual cells (Dambly-Chaudi&egrave;re et al., 1986; Bodmer et al., 1987; Bodmer et al., 1989), any aberrations to these systems can be easily identified using antibodies to different neuronal proteins. The developing embryos are collected at certain times to ensure that the embryos are in the proper developmental stages for visualization. After collection, the outer layers of the embryo, the chorion membrane and the vitelline envelope that surrounds the embryo, are removed before fixation. Embryos are then incubated with neuronal antibodies and visualized using fluorescently labeled secondary antibodies. Embryos at stages 12-17 are visualized to access the embryonic nervous system. At stage 12 the CNS germ band starts shortening and by stage 15 the definitive pattern of the commissure has been achieved. By stage 17 the CNS contracts and the PNS is fully developed (Campos-Ortega et al. 1985). Thus changes in the pattern of the PNS and CNS can be easily observed during these developmental stages.

Visualization of the Embryonic Nervous System in Whole-mount Drosophila Embryos

We describe the procedure to prepare staged Drosophila embryos for the visualization of the embryonic nervous system during embryogenesis.

The Drosophila embryo is an attractive model system for investigating the cellular and molecular basis of neuronal development. Here we describe the ...

Whole Mount Preparation of the Adult Drosophila Ventral Nerve Cord for Giant Fiber Dye Injection

To analyze the axonal and dendritic morphology of neurons, it is essential to obtain accurate labeling of neuronal structures. Preparing well labeled samples with little to no tissue damage enables us to analyze cell morphology and to compare individual samples to each other, hence allowing the identification of mutant anomalies.
In the demonstrated dissection method the nervous system remains mostly inside the adult fly. Through a dorsal incision, the abdomen and thorax are opened and most of the internal organs are removed. Only the dorsal side of the ventral nerve cord (VNC) and the cervical connective 
(CvC) containing the big axons of the giant fibers (GFs)1 are exposed, while the brain containing the GF cell body and dendrites remains2 in the 
intact head. In this preparation most nerves of the VNC should remain attached to their muscles. 
Following the dissection, the intracellular filling of the giant fiber (GF) with a fluorescent dye is demonstrated. In the CvC the GF axons are located at the dorsal surface and thus can be easily visualized under a microscope with differential interference contrast (DIC) optics. This allows the injection of the GF axons with dye at this site to label the entire GF including the axons and their terminals in the VNC. This method results in reliable and strong staining of the GFs allowing the neurons to be imaged immediately after filling with an epifluorescent microscope. Alternatively, the fluorescent signal can be enhanced using standard immunohistochemistry procedures3 suitable for high resolution confocal microscopy.

Whole Mount Preparation of the Adult Drosophila Ventral Nerve Cord for Giant Fiber Dye Injection

An in vivo dissection of the adult Drosophila ventral nerve cord (VNC) is demonstrated. This particular dissection method causes little damage to the VNC allowing the subsequent labeling of the giant fiber neurons with fluorescent dye for high resolution imaging.

To analyze the axonal and dendritic morphology of neurons, it is essential to obtain accurate labeling of neuronal structures. Preparing well labeled ...

An Ex Vivo Laser-induced Spinal Cord Injury Model to Assess Mechanisms of Axonal Degeneration in Real-time

Injured CNS axons fail to regenerate and often retract away from the injury site. Axons spared from the initial injury may later undergo secondary axonal degeneration. Lack of growth cone formation, regeneration, and loss of additional myelinated axonal projections within the spinal cord greatly limits neurological recovery following injury. To assess how central myelinated axons of the spinal cord respond to injury, we developed an ex vivo living spinal cord model utilizing transgenic mice that express yellow fluorescent protein in axons and a focal and highly reproducible laser-induced spinal cord injury to document the fate of axons and myelin (lipophilic fluorescent dye Nile Red) over time using two-photon excitation time-lapse microscopy. Dynamic processes such as acute axonal injury, axonal retraction, and myelin degeneration are best studied in real-time. However, the non-focal nature of contusion-based injuries and movement artifacts encountered during in vivo spinal cord imaging make differentiating primary and secondary axonal injury responses using high resolution microscopy challenging. The ex vivo spinal cord model described here mimics several aspects of clinically relevant contusion/compression-induced axonal pathologies including axonal swelling, spheroid formation, axonal transection, and peri-axonal swelling providing a useful model to study these dynamic processes in real-time. Major advantages of this model are excellent spatiotemporal resolution that allows differentiation between the primary insult that directly injures axons and secondary injury mechanisms; controlled infusion of reagents directly to the perfusate bathing the cord; precise alterations of the environmental milieu (e.g., calcium, sodium ions, known contributors to axonal injury, but near impossible to manipulate in vivo); and murine models also offer an advantage as they provide an opportunity to visualize and manipulate genetically identified cell populations and subcellular structures. Here, we describe how to isolate and image the living spinal cord from mice to capture dynamics of acute axonal injury.

An Ex Vivo Laser-induced Spinal Cord Injury Model to Assess Mechanisms of Axonal Degeneration in Real-time

We present a protocol utilizing two-photon excitation time-lapse microscopy to simultaneously visualize the dynamics of axon and myelin injuries in real time. This proposed protocol permits studies of both intrinsic and extrinsic factors which can influence central myelinated axon fate after injury and contribute to permanent clinical disability.

Injured CNS axons fail to regenerate and often retract away from the injury site. Axons spared from the initial injury may later undergo secondary axonal ...

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 ...

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

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

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

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

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

Laminectomy and Spinal Cord Window Implantation in the Mouse

This protocol describes a method for spinal cord laminectomy and glass window implantation for in vivo imaging of the mouse spinal cord. An integrated digital vaporizer is utilized to achieve a stable plane of anesthesia at a low-flow rate of isoflurane. A single vertebral spine is removed, and a commercially available cover-glass is overlaid on a thin agarose bed. A 3D-printed plastic backplate is then affixed to the adjacent vertebral spines using tissue adhesive and dental cement. A stabilization platform is used to reduce motion artifact from respiration and heartbeat. This rapid and clamp-free method is well-suited for acute multi-photon fluorescence microscopy. Representative data are included for an application of this technique to two-photon microscopy of the spinal cord vasculature in transgenic mice expressing eGFP:Claudin-5 &#8212; a tight junction protein.

This protocol describes implantation of a glass window onto the spinal cord of a mouse to facilitate visualization by intravital microscopy.

This protocol describes a method for spinal cord laminectomy and glass window implantation for in vivo imaging of the mouse spinal cord. An integrated ...

Preparation of Acute Spinal Cord Slices for Whole-cell Patch-clamp Recording in Substantia Gelatinosa Neurons

Recent whole-cell patch-clamp studies from substantia gelatinosa (SG) neurons have provided a large body of information about the spinal mechanisms underlying sensory transmission, nociceptive regulation, and chronic pain or itch development. Implementations of electrophysiological recordings together with morphological studies based on the utility of acute spinal cord slices have further improved our understanding of neuronal properties and the composition of local circuitry in SG. Here, we present a detailed and practical guide for the preparation of spinal cord slices and show representative whole-cell recording and morphological results. This protocol permits ideal neuronal preservation and can mimic in vivo conditions to a certain extent. In summary, the ability to obtain an in vitro preparation of spinal cord slices enables stable current- and voltage-clamp recordings and could thus facilitate detailed investigations into the intrinsic membrane properties, local circuitry and neuronal structure using diverse experimental approaches.

Here, we describe the essential steps for whole-cell patch-clamp recordings made from substantia gelatinosa (SG) neurons in the in vitro spinal cord slice. This method allows the intrinsic membrane properties, synaptic transmission and morphological characterization of SG neurons to be studied.

Recent whole-cell patch-clamp studies from substantia gelatinosa (SG) neurons have provided a large body of information about the spinal mechanisms ...

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 ...

Understanding Motor Neuron Responses to Spinal Cord Stimulation

The Ex vivo Preparation of Spinal Cord Slice for the Whole-Cell Patch-Clamp Recording in Motor Neurons During Spinal Cord Stimulation

Spinal cord stimulation (SCS) can effectively restore locomotor function after spinal cord injury (SCI). Because the motor neurons are the final unit to execute sensorimotor behaviors, directly studying the electrical responses of motor neurons with SCS can help us understand the underlying logic of spinal motor modulation. To simultaneously record diverse stimulus characteristics and cellular responses, a patch-clamp is a good method to study the electrophysiological characteristics at a single-cell scale. However, there are still some complex difficulties in achieving this goal, including maintaining cell viability, quickly separating the spinal cord from the bony structure, and using the SCS to successfully induce action potentials. Here, we present a detailed protocol using patch-clamp to study the electrical responses of motor neurons to SCS with high spatiotemporal resolution, which can help researcher improve their skills in separating the spinal cord and maintaining the cell viability at the same time to smoothly study the electrical mechanism of SCS on motor neuron and avoid unnecessary trial and mistake.

Author Spotlight: Advancing Spinal Cord Stimulation - Exploring the Cellular Responses of Motor Neurons Through Patch-Clamp Electrophysiology 

This protocol describes a method using a patch-clamp to study the electrical responses of motor neurons to spinal cord stimulation (SCS) with high spatiotemporal resolution, which can help researchers improve their skills in separating the spinal cord and maintaining cell viability simultaneously.

Spinal cord stimulation (SCS) can effectively restore locomotor function after spinal cord injury (SCI). Because the motor neurons are the final unit to ...