Name: zebrafish in situ spinal cord preparation for electrophysiological recordings from spinal sensory and motor neurons
Uploaded: 2017-04-18T14:00:00
Description: Watch this Scientific Journal Video about Zebrafish In Situ Spinal Cord Preparation for Electrophysiological Recordings from Spinal Sensory and Motor Neurons at JoVE.com

This work was supported by grants from the NIH (F32 NS059120 to RLM and R01NS25217 and P30NS048154 to ABR).

All animal procedures were approved by the Institutional Animal Care and Use Committee (IACUC; Office of Laboratory Animal Resources, University of Colorado Anschutz Medical Campus).
1. Zebrafish Husbandry
<ol>
	<li>Raise and maintain adult zebrafish (Danio rerio) at 28.5 &#176;C on a 10 h dark/14 h light cycle and with appropriate water treatment and exchange52.</li>
	<li>Raise zebrafish embryos/larvae at 28.5 &#176;C in embryo medium until they reach the de...

The methods described here allow for the electrical and morphological characterization of sensory and motor neurons of zebrafish embryos after minimal dissection of the spinal cord. Neurons remain healthy for at least 1 h, the time limit imposed on these recordings. Neurons have been recorded using the standard whole-cell configuration, as well as from nucleated patches; the latter method minimizes space-clamp issues that can preclude a detailed biophysical study of ion currents9.
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George Streisinger pioneered the use of Danio rerio, commonly known as zebrafish, as a model system for the genetic analysis of vertebrate development10. The model offers several advantages including: (1) relatively simple and inexpensive animal husbandry; (2) external fertilization, allowing easy access to embryos from the earliest developmental stages; and (3) a transparent embryo, permitting direct and repeated observations of cells, tissues, and organs as they form.
Over the ensuing decades, several advances further increased the power of the zebrafish model. In particular, forward genetic screens and whole-genome sequencing efforts played key roles in the identification of mutations and genes critical to many developmental processes11,12,13,14,15,16. Gateway cloning methods have allowed the routine application of transgenic approaches17,18. Recent advances in genome editing, exemplified by transcription activator-like (TALENs) and clustered regularly interspaced short palindromic repeats (CRISPR)-Cas9 nucleases, allow for the targeted introduction of mutations, as well as knock-out and knock-in approaches19,20,21,22. Combined, these methods make zebrafish a powerful model for the study of the genetic mechanisms underlying specific behaviors and several human diseases23,24,25,26,27.
This work focuses on developmental regulation and the role of electrical activity in neuronal development. The focus is on the spinal cord, for which the zebrafish model provides several advantages. First, it is relatively easy to access zebrafish at embryonic and larval stages; therefore, one can study spinal cord function during developmental stages that have fewer neurons and simpler circuitry28,29. Moreover, the zebrafish spinal cord has a diverse set of neurons, similar to other vertebrates, as demonstrated by characteristic and distinguishing patterns of transcription factors30,31,32,33,34,35.
The majority of studies in zebrafish that aim to uncover the mechanisms that underlie the function of spinal cord circuits, especially ones that support locomotion, are understandably focused on larval stages36,37,38,39,40,41,42,43. However, many of the neurons that form the spinal locomotive networks initiate their differentiation at early embryonic stages, ~9-10 h post-fertilization (hpf)44,45,46,47,48,49,50,51. In view of this, understanding how the morphological and electrical properties of spinal neurons arise and change between the embryonic and larval stages is important for an overall understanding of locomotor circuit formation and function.
The dissection methods described here allow patch clamp recordings from spinal neurons and have been successfully applied at embryonic stages (~17-48 hpf) and larval stages (~3-7 days post fertilization [dpf]). This approach limits the amount of dissection required to provide access to the neurons of interest. The protocol differs from the majority of the other published methods for recording from zebrafish spinal neurons in that veterinarian suture glue is used, rather than a fine tungsten pin, to attach the embryo or larva to the recording chamber. The availability of two different approaches (i.e., suture glue versus the tungsten pin) for mounting the zebrafish embryos or larvae for electrophysiological analysis provides researchers with alternative options to achieve their specific experimental goals.
First, procedures for accessing and recording from a population of primary sensory neurons, Rohon-Beard cells, are described. The cell bodies of these neurons lie within the dorsal spinal cord. Rohon-Beard cells exist in numerous vertebrate species, differentiate early in development, and underlie the embryonic touch response6,44,47,48.
Second, procedures for accessing and recording from spinal motor neurons are detailed. Zebrafish spinal motor neurons arise during two waves of neurogenesis. The earlier-born primary motor neurons arise at the end of gastrulation (~9-16 hpf), with only 3-4 primary motor neurons present per hemisegment45,46,49. In contrast, the later-born population of secondary motor neurons is more numerous and arises during a prolonged period, starting at ~14 hpf45,50. Secondary motor neuron genesis in mid-trunk segments is mostly completed by 51 hpf50. Secondary motor neurons are considered to be the counterpart of motor neurons in amniotes46. Interestingly, supraspinal neurons, via dopamine, regulate locomotion in the larva and secondary motor neuron genesis in the embryo and young larva50,51. Primary and secondary motor neurons each comprise several different subtypes. Each primary motor neuron subtype projects a peripheral axon that innervates a characteristic muscle group, resulting in a stereotypical, identifying axonal trajectory. Generally, secondary motor neurons follow the axonal pathways previously established by primary motor neurons. Thus, with respect to axonal trajectories, primary and secondary motor neurons are similar, with the exception that axonal thickness and somata size are greater for primary motor neurons45.
Third, methods for recording from a few types of interneurons are discussed. However, in these cases, a limited amount of removal of other spinal cord cells is required, and thus the spinal cord is less intact than for recordings from Rohon-Beard cells or motor neurons.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Vacuum filter/Storage bottle, 0.22 mm pore</td>
			<td>Corning</td>
			<td>431096</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe filter 0.2 mm</td>
			<td>Whatman</td>
			<td>6780-2502</td>
			<td></td>
		</tr>
		<tr>
			<td>Tricaine</td>
			<td>Sigma</td>
			<td>A-5040</td>
			<td>Ethyl 3-aminobenzoate methanesulfonate salt&#160;</td>
		</tr>
		<tr>
			<td>a-bugarotoxin</td>
			<td>Tocris</td>
			<td>11032-79-4</td>
			<td></td>
		</tr>
		<tr>
			<td>Tetrodotoxin</td>
			<td>Tocris</td>
			<td>4368-28-9</td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa-549 hydrazine salt</td>
			<td>Molecular Probes</td>
			<td>A-10438</td>
			<td>fluorescent dye</td>
		</tr>
		<tr>
			<td>Spin-X centrifuge tube filter</td>
			<td>Corning</td>
			<td>8161</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass microscope slide</td>
			<td>Fisher</td>
			<td>12-550C</td>
			<td></td>
		</tr>
		<tr>
			<td>Sylgard silicone elastomer kit</td>
			<td>Dow Corning</td>
			<td>184</td>
			<td>silicone elastomer</td>
		</tr>
		<tr>
			<td>Petri dishes</td>
			<td>Falcon</td>
			<td>351029</td>
			<td></td>
		</tr>
		<tr>
			<td>Borosilicate glass capillaries</td>
			<td>Harvard Apparatus</td>
			<td>30-0038</td>
			<td>inner and outer diameters of 0.78 and 1.0 mm (thin walled glass capillaries)</td>
		</tr>
		<tr>
			<td>Borosilicate glass capillaries</td>
			<td>Drummond Scientific</td>
			<td>1-000-1000-100</td>
			<td>inner and outer diameters of 1.13 and 1.55 mm (thick walled glass capillaries)</td>
		</tr>
		<tr>
			<td>Miniature barbed polypropylene fitting&#160;</td>
			<td>Cole-Palmer</td>
			<td>6365-90</td>
			<td></td>
		</tr>
		<tr>
			<td>Vetbond tissue adhesive&#160;</td>
			<td>3M</td>
			<td>1469SB</td>
			<td></td>
		</tr>
		<tr>
			<td>Collagenase XI</td>
			<td>Sigma</td>
			<td>C7657</td>
			<td></td>
		</tr>
		<tr>
			<td>Microelectrode puller</td>
			<td>Sutter Instruments</td>
			<td>&#160;Model P-97&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>Amplifier</td>
			<td>Molecular Devices</td>
			<td>Axopatch 200B</td>
			<td></td>
		</tr>
		<tr>
			<td>Head stage</td>
			<td>Molecular Devices</td>
			<td>CV203BU</td>
			<td></td>
		</tr>
		<tr>
			<td>Motorized micromanipulator&#160;&#160;</td>
			<td>Sutter Instruments</td>
			<td>MP-285</td>
			<td></td>
		</tr>
		<tr>
			<td>Tygon tubing</td>
			<td>Fisher</td>
			<td>14-169-1B</td>
			<td>ID 1/16 IN, OD 1/8 IN and WALL 1/32 IN (flexible laboratory tubing)</td>
		</tr>
		<tr>
			<td>Electrode holder</td>
			<td>Molecular Devices</td>
			<td>1-HC-U</td>
			<td></td>
		</tr>
		<tr>
			<td>Pharmaseal Three-Way Stopcocks&#160;</td>
			<td>Baxter</td>
			<td>K75</td>
			<td></td>
		</tr>
		<tr>
			<td>Digitizer</td>
			<td>Axon Instruments</td>
			<td>Digidata 1440A</td>
			<td></td>
		</tr>
		<tr>
			<td>Inverted microscope&#160;</td>
			<td>Zeiss</td>
			<td>Axioskop2 FS plus</td>
			<td></td>
		</tr>
		<tr>
			<td>40x/0.80w Achroplan objective</td>
			<td>Zeiss</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Data acquisition and analysis software&#160;</td>
			<td>Axon Instruments</td>
			<td colspan="2">PClamp 10 - Clampex and Clampfit&#160;</td>
		</tr>
		<tr>
			<td>Micropipette puller</td>
			<td>Sutter Instruments</td>
			<td>Model P-97</td>
			<td></td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td colspan="2">Dissection and Recording Solutions(in mM)</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td colspan="4">All solutions, except the intracellular, are stable for ~ 2 - 3 months when filtered (0.22 mm filter cups) and stored at room temperature (RT).</td>
		</tr>
		<tr>
			<td colspan="4">The intracellular solution is filtered (0.2 mm syringe filters) and stored frozen (-20&#176;C) in small aliquots that are individually thawed on the day of use.&#160;</td>
		</tr>
		<tr>
			<td colspan="2" rowspan="1">Dissection/Ringer&#8217;s solution</td>
			<td></td>
			<td colspan="2">145 NaCl, 3 KCl, 1.8 CaCl2.2H2O, 10 HEPES; pH 7.4 (with NaOH)</td>
		</tr>
		<tr>
			<td colspan="2">Pipette (intracellular) recording solution</td>
			<td></td>
			<td colspan="2">135 KCl, 10 EGTA-acid, 10 HEPES; pH 7.4 (with KOH).</td>
		</tr>
		<tr>
			<td colspan="2">Bath (extracellular) recording solution/voltage and current-clamp</td>
			<td></td>
			<td colspan="2">125 NaCl, 2 KCl, 10 CaCl2.2H2O, 5 HEPES; pH 7.4 (with NaOH).</td>
		</tr>
		<tr>
			<td colspan="2">Alexa-594 hydrazine salt stock solution.&#160;</td>
			<td></td>
			<td colspan="2">Prepare a 13.2 mM stock in ddH2O, aliquot (~ 100 &#181;l) and store at -20&#176;C. For use, dilute the stock solutiond 132 fold with pipette solution to a final concentration of 100 mM. After dilution, filter the Alexa-594 containing pipette solution&#160; with a centrifuge tube filter.</td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Immobilizing agents</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td colspan="2">0.4 % ethyl 3-aminobenzoate methanesulfonate salt (Tricaine)</td>
			<td></td>
			<td colspan="2">Prepare a 0.4% stock solution in 0.2M Tris, pH9 (0.4 g Tricaine/100 ml 0.2 M Tris</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td colspan="2">Adjust pH to 7 with NaOH and store at -20&#176;C.</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td colspan="2">For use, dilute the stock solution ~ 25 fold in embryo media</td>
		</tr>
		<tr>
			<td>250 &#956;M &#945;-bungarotoxin</td>
			<td></td>
			<td colspan="2">Prepare a 250 &#956;M stock in ddH2O (1 mg/500 &#956;l), prepare 100 &#181;l aliquots, and sotre at -20&#176;C.</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td colspan="2">For use, dilute 2,500-fold with extracellular solution to a final concentration of 100 nM.</td>
		</tr>
		<tr>
			<td>1 mM Tetrodotoxin</td>
			<td></td>
			<td colspan="2">Prepare a 1 mM stock in ddH2O (1 mg/3 ml), prepare 100 &#181;l aliquots, and sotre at -20&#176;C.</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td colspan="2">For use, dilute 2,000-fold with extracellular solution to a final concentration of 500 nM.</td>
		</tr>
	</tbody>
</table>

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.

We have successfully recorded from Rohon-Beard neurons in 17 hpf embryos through 7 dpf larvae (Figure 5A and 5B). When Rohon-Beard cells were recorded, the preparation was mounted dorsal-side up. Such mounting allows for the unambiguous identification of Rohon-Beard cells based on their superficial dorsal positions and large soma sizes. The identification is additionally confirmed by the stereotypical hyperpolarized resting membrane potential of these neu...

Watch this Scientific Journal Video about Zebrafish In Situ Spinal Cord Preparation for Electrophysiological Recordings from Spinal Sensory and Motor Neurons at JoVE.com

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 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 ...

The overall goal of this preparation is to allow the electrophysiological study of zebrafish spinal neurons in situ at the embryonic and larval stages. The electrophysiological analyses compliment the powerful genetic and imaging approaches that are also possible with the zebrafish model. While the method was originally developed to study rohon beard neurons, we have easily adapted the approach over time and now we can study various types of spinal neurons.Demonstrating the procedure will be Dr.Rosa Moreno, a research associate, with whom I've had the pleasure of collaborating for many years. To begin this procedure remove the black bulb from one glass bore, and replace it with a white rubber adaptor from another glass bore. Next, cut a piece of flexible tubing to a length of about 38 centimeters.Then, attach the flexible tubing via a small, straight polypropylene fitting. To prepare the silicone elastomer add the base and the curing agent to a plastic conical tube at a ratio of four to one. Afterward, mix the elastomer base and curing agent thoroughly.Pour the mixture into two 100 millimeter Petri dishes up to about one millimeter deep in one Petri dish and about 2.5 millimeters deep in the other. Allow the silicone elastomer to cure by exposure to air for four to five days. If cured elastomer is needed sooner, incubate it at 60 degrees Celsius.Afterward, cut out a 3.8 centimeter by 6.3 centimeter rectangle from the one millimeter thick cured elastomer to serve as the bottom of the chamber. From the 2.5 millimeter thick cured silicone cut a rectangle of 3.8 centimeters by 6.3 centimeters. Then from that, cut out an internal rectangle of 2.5 centimeters by five centimeters to generate a frame for the top of the chamber.To make the dissection recording chamber place the thin silicone rectangle directly on top of the glass slide, making sure to remove any air bubbles between the silicone and the glass. Then place the silicone rectangle frame on top of the thin bottom layer of silicone. In this procedure place an embryo in a dissection chamber containing two to three milliliters of ringer's solution.Immobilize the embryo by adding 100 microliters of 0.4%tricaine solution to the chamber. Then pull some injection micro pipettes with long thin tips from the thin wall glass with a micro pipette puller. Attach a pulled glass micro pipette to the glass floor of the glue dispenser.Under the dissecting microscope use tweezers to break the tip of the glass micro pipette resulting in a tip of about 75 micrometers in diameter. Now, load the glass micro pipette with three to five microliters of glue by applying suction through the mouth piece. Bring the glue filled tip to the dissection chamber.Then bring the tip of the glue loaded micro pipette near the head of the embryo while maintaining slight positive pressure to the micro pipette via the mouth tube. Once the tip is near the head of the embryo, apply sufficient positive pressure to expel a small drop of glue onto the bottom of the chamber. Use a dissecting pin to move the embryo towards the drop of glue so that the head makes contact with the glue.Orient the embryo dorsal side up, and press on the head to ensure good contact with the glue. As the glue slowly hardens use the dissecting pin to reposition the embryo so that it lies ventral side down, dorsal side up. Dip the dissecting pin into the drop of glue and drag a few threads of the glue over and across the head of the embryo to further secure its position.Once the head is securely attached to the silicone and the glue has solidified sacrifice the embryo by transection at the hind brain level with another glass micro pipette. Use a fresh glass micro pipette to superficially cut the skin several times at a position caudal the hind brain. Pierce the skin superficially by moving the pipette perpendicular to rostro caudal axis on each side of the trunk for dorsally mounted specimens, or on the exposed side of the trunk for laterally mounted specimens.To create a flap of skin for the tweezers to grab, scrape the skin several times with the micro pipette, at the level of and perpendicularly to the initial cut. Using tweezers, gradually lift the skin flap, and pull the skin caudally. After that, deliver a small drop of glue near the tail of the embryo.Use this glue to attach the tail to the bottom of the dissection chamber. As the glue hardens, adjust the position of the trunk with the dissecting pin to ensure that the trunk remains dorsally oriented, and that it is firmly attached to the chamber. Following this initial dissection, rinse the preparation extensively with ringer's solution to remove the tricaine and debris, and allow the preparation to rest for five minutes.Next, replace the dissection solution with extra cellular recording solution. If needed, add an immobilizing agent to the preparation. Next, transfer the dissection chamber with the mounted embryo to the stage of an upright compound microscope, equipped with a 40 X water immersion long working distance objective.Then mount an empty bore of silicate thick wall glass micro pipette onto the electrode holder of the head stage. Attach the tubing at one end to the air outlet of the electrode holder and at the other end to a three way stop cock. After that, place a mouth piece at one end of another piece of tubing, and attach it to the three way stop cock at the other end.Subsequently, bring the micro pipette tip to the most dorsal portion of the spinal cord, and gently pierce the meninges. Follow with swift, short, sideways movements to loosen the meninges. After the micro pipette tip has traversed the meninges advance and raise the micro pipette to pull the meninges away from the spinal cord.Subsequently move the micro pipette rostorally advancing over one to two hemi segments to expose the rohon beard cells. In our experience, rohon beard neurons are more fragile than other spinal neurons. To maintain them healthy we expose one or two rohon beards at a time.Voltage clamp recordings of outward and inward currents were obtained from the rohon beard neurons in one, two, and seven days post fertilization embryos and larvae. The holding potential was 80 millivolts, and the currents were elicited by a depolarizing step to positive 20 millivolts. In current clamp mode, 0.35 nano amp current injection for one millisecond elicits a single action potential from the rohon beard neurons in both embryos and larvae.In the absence of electrical stimulation, rohon beard neurons do not show spontaneous spiking. The resting potential of the rohon beard neurons becomes increasingly more negative as the development progresses. Once this method is mastered, embryos may be mounted and dissected to expose rohon beards within 10 minutes.As mentioned in the protocol, it is important to understand the factors that determine how quickly the glue hardens. This video highlights the critical steps for recording from spinal neurons in zebrafish.

zebrafish-situ-spinal-cord-preparation-for-electrophysiological

Research

JoVE Journal

Neuroscience

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 ...

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 ...

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 ...

Preparation of Primary Mixed Glial Cultures from Adult Mouse Spinal Cord Tissue

It has been well-accepted that spinal cord glial responses contribute significantly to the development of neuropathic pain. Tremendous information regarding glial activities at the cellular and molecular levels has been obtained through in vitro cell culture systems. The in vitro systems utilized, mainly include primary glia derived from neonatal brain cortical tissue and immortalized cell lines. However, these systems may not reflect the characteristics of spinal cord glial cells in vivo. In order to further investigate the roles of spinal cord glial cells in the development of peripheral nerve injury-induced neuropathic pain using a culture system that better reflects the in vivo condition, our laboratory has developed a method to establish primary spinal cord mixed glial cultures from adult mice. Briefly, spinal cords are collected from adult mice and processed through papain digestion followed by myelin removal with a density-gradient medium. Single cell suspensions are cultured in complete Dulbecco&#39;s modified Eagle media (cDMEM) supplemented with 2-mercaptoethanol (2-ME) at 35.9 oC. These culture conditions were optimized specifically for the growth of mixed glial cells. Under these conditions, cells are ready to be used for experimentation between 12 - 14 d&#160;(cells are usually in log phase during this time) after the establishment of the culture (D 0) and can be kept in culture conditions up to D 21. This culture system can be used to investigate the responses of spinal cord glial cells upon stimulation with various substances and agents. Besides neuropathic pain, this system can be used to study glial responses in other diseases that involve pathological changes of spinal cord glial cells.

The development of neuropathic pain involves pathological changes of spinal cord glial cells. A reliable glial culture system derived from adult spinal cord tissue and designed for studying these cells in vitro is lacking. Therefore, we show here how&#160;to establish primary mixed glial cultures from adult mouse spinal cord tissue.

It has been well-accepted that spinal cord glial responses contribute significantly to the development of neuropathic pain. Tremendous information ...

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 ...

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 ...

Isolation and Culture of Oculomotor, Trochlear, and Spinal Motor Neurons from Prenatal Islmn:GFP Transgenic Mice

Oculomotor neurons (CN3s) and trochlear neurons (CN4s) exhibit remarkable resistance to degenerative motor neuron diseases such as amyotrophic lateral sclerosis (ALS) when compared to spinal motor neurons (SMNs). The ability to isolate and culture primary mouse CN3s, CN4s, and SMNs would provide an approach to study mechanisms underlying this selective vulnerability. To date, most protocols use heterogeneous cell cultures, which can confound the interpretation of experimental outcomes. To minimize the problems associated with mixed-cell populations, pure cultures are indispensable. Here, the first protocol describes in detail how to efficiently purify and cultivate CN3s/CN4s alongside SMNs counterparts from the same embryos using embryonic day 11.5 (E11.5) IslMN:GFP transgenic mouse embryos. The protocol provides details on the tissue dissection and dissociation, FACS-based cell isolation, and in vitro cultivation of cells from CN3/CN4 and SMN nuclei. This protocol adds a novel in vitro CN3/CN4 culture system to existing protocols and simultaneously provides a pure species- and age-matched SMN culture for comparison. Analyses focusing on the morphological, cellular, molecular, and electrophysiological characteristics of motor neurons are feasible in this culture system. This protocol will enable research into the mechanisms that define motor neuron development, selective vulnerability, and disease.

Isolation and Culture of Oculomotor, Trochlear, and Spinal Motor Neurons from Prenatal Islmn:GFP Transgenic Mice

This work presents a protocol to yield homogeneous cell cultures of primary oculomotor, trochlear, and spinal motor neurons. These cultures can be used for comparative analyses of the morphological, cellular, molecular, and electrophysiological characteristics of ocular and spinal motor neurons.

Oculomotor neurons (CN3s) and trochlear neurons (CN4s) exhibit remarkable resistance to degenerative motor neuron diseases such as amyotrophic lateral ...

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 ...

Optogenetic Phase Transition of TDP-43 in Spinal Motor Neurons of Zebrafish Larvae

Abnormal protein aggregation and selective neuronal vulnerability are two major hallmarks of neurodegenerative diseases. Causal relationships between these features may be interrogated by controlling the phase transition of a disease-associated protein in a vulnerable cell type, although this experimental approach has been limited so far. Here, we describe a protocol to induce phase transition of the RNA/DNA-binding protein TDP-43 in spinal motor neurons of zebrafish larvae for modeling cytoplasmic aggregation of TDP-43 occurring in degenerating motor neurons in amyotrophic lateral sclerosis (ALS). We describe a bacterial artificial chromosome (BAC)-based genetic method to deliver an optogenetic TDP-43 variant selectively to spinal motor neurons of zebrafish. The high translucency of zebrafish larvae allows for the phase transition of the optogenetic TDP-43 in the spinal motor neurons by a simple external illumination using a light-emitting diode (LED) against unrestrained fish. We also present a basic workflow of live imaging of the zebrafish spinal motor neurons and image analysis with freely available Fiji/ImageJ software to characterize responses of the optogenetic TDP-43 to the light illumination. This protocol enables the characterization of TDP-43 phase transition and aggregate formation in an ALS-vulnerable cellular environment, which should facilitate an investigation of its cellular and behavioral consequences.

We describe a protocol to induce phase transition of TAR DNA-binding protein 43 (TDP-43) by light in the spinal motor neurons using zebrafish as a model.

Abnormal protein aggregation and selective neuronal vulnerability are two major hallmarks of neurodegenerative diseases. Causal relationships between ...

A Minimally Invasive, Fast Spinal Cord Lateral Hemisection Technique for Modeling Open Spinal Cord Injuries in Rats

Open spinal cord injury techniques modeling laceration-like injuries are time-consuming and invasive because they involve laminectomy. This new technique eliminates laminectomy by removing two spinous processes and lifting, then tilting the caudal vertebral arch. The surgical area opens up without the need for laminectomy. Lateral hemisection is then performed with direct visible control under a microscope. The trauma is minimized, requiring only a small bone wound. 
This technique has several advantages: it is faster and, therefore, less of a burden for the animal, and the bone wound is smaller. Because the laminectomy is eliminated, there is less chance for unwanted injury to the spinal cord, and there are no bone splinters that can cause problems (bone splinters embedded in the spinal cord can cause swelling and secondary damage). The vertebral canal remains intact. The main limitation is that the hemisection can only be performed in the intervertebral spaces. 
The results show that this technique can be performed much faster than the traditional surgical approach, using laminectomy (11 min vs. 35 min). This technique can be useful for researchers working with animal models of open spinal cord injury as it is widely adaptable and does not require any additional specialized instrumentation.

Here, we describe a new, fast technique modeling open spinal cord injury in rats that eliminates laminectomy. Lateral hemisection is performed while viewing through a microscope. The technique is versatile and can also be used in the cervical, thoracic, and lumbar regions of the spinal cord of other animals.

Open spinal cord injury techniques modeling laceration-like injuries are time-consuming and invasive because they involve laminectomy. This new technique ...