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

The authors declare no competing financial interests.

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.
An important challenge is to achieve the firm attachment of the embryo or larva to the chamber in order to remove the skin and to perform the limited dissection required to provide access to the neurons of interest. The preparation also needs to be properly secured to the recording chamber for the whole-cell patch-clamp methods. A method that meets this challenge via the use of veterinarian suture glue to attach the embryo or larva to the dissection/recording chamber is described here, an approach that has been used for the dissection of other model organisms (e.g., Drosophila)63. From our experience training others, we find that the most critical step to master is the controlled and precise delivery of small amounts of glue. Here, a glue dispenser device that allows a user to apply negative and positive pressure to load glue into or to expel it from the tip of a micropipette is discussed. Using the suture glue, embryos and larvae can be firmly attached to the chamber and oriented either dorsal-side up or laterally. In this way, different access options for a variety of neurons are available. Also, the layer of silicone elastomer on the chamber bottom can be even thinner than the 1 mm specified here, providing potential optical advantages. Another method, more commonly used to attach the preparation to a recording chamber, involves use of fine tungsten pins1,2,3,4,5. While that methods differ, both allow electrophysiological access to zebrafish spinal neurons, affording researchers with options that can be selected based on the goals and challenges of the experiment.
The zebrafish preparation described here allows for the electrical and morphological study of spinal neurons in situ during their earliest stages of differentiation. By recording from spinal neurons using these methods, we have gained insight into the cellular effects of several mutations, even prior to the identification of the lesioned gene6,64,65.

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><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&amp;nbsp;</td></tr><tr><td>&amp;alpha;-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&amp;nbsp;</td><td>Cole-Palmer</td><td>6365-90</td><td></td></tr><tr><td>Vetbond tissue adhesive&amp;nbsp;</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>&amp;nbsp;Model P-97&amp;nbsp;</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&amp;nbsp;&amp;nbsp;</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&amp;nbsp;</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&amp;nbsp;</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&amp;nbsp;</td><td>Axon Instruments</td><td></td><td>PClamp 10 - Clampex and Clampfit&amp;nbsp;</td></tr><tr><td>Micropipette puller</td><td>Sutter Instruments</td><td>Model P-97</td><td></td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td colspan="4">&lt;strong&gt;Dissection and Recording Solutions (in mM)&lt;/strong&gt;</td></tr><tr><td>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><td></td><td></td><td></td></tr><tr><td>The intracellular solution is filtered (0.2 mm syringe filters) and stored frozen (-20 &amp;deg;C) in small aliquots that are individually thawed on the day of use.&amp;nbsp;</td><td></td><td></td><td></td></tr><tr><td>Dissection/Ringer&amp;rsquo;s solution</td><td></td><td></td><td>145 NaCl, 3 KCl, 1.8 CaCl&lt;sub&gt;2&lt;/sub&gt;.2H&lt;sub&gt;2&lt;/sub&gt;O, 10 HEPES; pH 7.4 (with NaOH)</td></tr><tr><td>Pipette (intracellular) recording solution</td><td></td><td></td><td>135 KCl, 10 EGTA-acid, 10 HEPES; pH 7.4 (with KOH).</td></tr><tr><td>Bath (extracellular) recording solution/voltage and current-clamp</td><td></td><td></td><td>125 NaCl, 2 KCl, 10 CaCl&lt;sub&gt;2&lt;/sub&gt;.2H&lt;sub&gt;2&lt;/sub&gt;O, 5 HEPES; pH 7.4 (with NaOH).</td></tr><tr><td>Alexa-594 hydrazine salt stock solution.&amp;nbsp;</td><td></td><td></td><td>Prepare a 13.2 mM stock in ddH&lt;sub&gt;2&lt;/sub&gt;O, aliquot (~100 &amp;micro;l) and store at -20 &amp;deg;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&amp;nbsp; with a centrifuge tube filter.</td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Immobilizing agents&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>0.4% ethyl 3-aminobenzoate methanesulfonate salt (Tricaine)</td><td></td><td></td><td>Prepare a 0.4% stock solution in 0.2 M Tris, pH 9 (0.4 g Tricaine/100 mL 0.2 M Tris</td></tr><tr><td colspan="4">Adjust pH to 7 with NaOH and store at -20 &amp;deg;C.</td></tr><tr><td colspan="4">For use, dilute the stock solution ~25 fold in embryo media</td></tr><tr><td>250 mM &amp;alpha;-bungarotoxin</td><td></td><td></td><td>Prepare a 250 mM stock in ddH&lt;sub&gt;2&lt;/sub&gt;O (1 mg/500 mL), prepare 100 &amp;micro;L aliquots, and sotre at -20 &amp;deg;C.</td></tr><tr><td colspan="4">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></td><td>Prepare a 1 mM stock in ddH2O (1 mg/3 mL), prepare 100 &amp;micro;L aliquots, and sotre at -20 &amp;deg;C.</td></tr><tr><td colspan="4">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 neurons (Figure 5, inset table)6,54. Moreover, as primary sensory neurons, Rohon-Beard neurons lack synaptic input. Therefore, in the absence of electrical stimulation, no changes in membrane potential should occur while recording in current-clamp mode (Figure 5B&#39;). Since the initial recordings from Rohon-Beard cells in zebrafish were performed6, various transgenic lines (e.g., Tg(islet2b:gfp), Tg(ngn:gfp), and Tg(isletss:gfp)) have been generated that express fluorescent reporters in these neurons, further facilitating their identification55,56,57.
<img alt="Figure 5" src="/files/ftp_upload/55507/55507fig5.jpg" /> 
Figure 5: Whole-cell voltage- and current-clamp recordings from Rohon-Beard neurons in 1 and 2 dpf embryos and 7 dpf larvae. (A) Voltage-clamp recordings of outward and inward currents were obtained from Rohon-Beard neurons in 1- (thin black line), 2- (thick black line), and 7-dpf (gray line) embryos/larvae. The holding potential was -80 mV and currents were elicited by a depolarizing step to +20 mV. (B) Single action potentials are elicited by brief (1 ms) current injections (~0.35 nA) to Rohon-Beard neurons of 1- (thin black line), 2- (thick black line), and 7-dpf (gray line) embryos/larvae. (B&#39;) In the absence of electrical stimulation, no changes in membrane potential, such as spontaneous postsynaptic depolarizations, occur in Rohon-Beard neurons. The inset table summarizes the values of resting membrane potentials recorded from Rohon-Beard neurons of 1- (n = 21) and 2- (n = 9) dpf embryos and 7- (n = 7) dpf larvae. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55507/55507fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Transgenic lines that allow unequivocal identification of other spinal neuron subtypes are also available. Among these, the mnx1 transgenic line Tg(mnx1:gfp) expresses green fluorescent protein (gfp) in a subset of spinal motor neurons soon after their specification (~14-16 hpf)58,59. Due to the stereotypical positioning of the primary motor neurons within each hemisegment (Figure 6A), together with the expression of gfp in the mnx1 transgenic, it is possible to identify the various primary motor neuron subtypes (Figure 6B and 6C). Including a fluorescent dye in the recording electrode solution allows for the visualization of axonal trajectories, providing additional confirmation of motor neuron identity, as some interneurons also express gfp in the Tg(mnx1:gfp) line. Alternatively, another transgenic line that allows the identification of motor neurons is the ET2 line60.
<img alt="Figure 6" src="/files/ftp_upload/55507/55507fig6.jpg" /> 
Figure 6: Whole-cell voltage and current-clamp recordings from motor neurons of 1 dpf zebrafish embryos. (A) A cartoon depicts the specific morphological features of the primary motor neuron subtypes present in the zebrafish spinal cord. Primary motor neurons are identified by the position of their soma within a segment (i.e., rostral [RoP], medial [MiP], or caudal [CaP])45. In addition, each subtype extends an axon to the periphery via a distinct path. The combined use of the Tg(mnx1:gfp) line and dye labeling reveals the stereotypical axonal arbor and the identity of the motor neuron subtype during a recording. Using the methods presented here, it is possible to sequentially record from three different primary motor neuron subtypes within the same hemisegment. (B) Voltage-clamp recordings are shown that were obtained from RoP, MiP, and CaP, all in a single hemisegment. A voltage step to +20 mV was used to elicit currents from a holding potential of -80 mV. (C) During current-clamp recordings from RoP, MiP, and CaP, brief (1 ms, ~0.4 nA) current injections were applied to trigger an action potential. The membrane potential was held at ~-65 mV. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55507/55507fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
A principal difference between primary and secondary motor neurons is the larger somata of the earlier-born neurons. However, secondary motor neuron subtypes are not identifiable by soma size or position. For recordings from specific secondary motor neurons, two transgenic lines, Tg(gata2:gfp) and Tg(islet1:gfp), have been used for the identification of secondary motor neurons with ventrally and dorsally projecting axons, respectively55,61. However, a third secondary motor neuron subtype is present in the zebrafish spinal cord, with axons that project dorsally and ventrally62. Accordingly, dye can be used to fill secondary motor neurons during recordings to identify subtypes on the basis of morphology (Figure 7A)8,9. Often, during voltage-clamp (Figure 7B, asterisks) or current-clamp recordings (Figure 7C and 7C&#39;, arrowheads) from secondary motor neurons, spontaneous or synaptic events are recorded.
<img alt="Figure 7" src="/files/ftp_upload/55507/55507fig7.jpg" /> 
Figure 7: Whole-cell voltage and current-clamp recordings from secondary motor neurons of 2 dpf embryos. (A) In the Tg(gata2:gfp) line, two different secondary motor neuron subtypes express gfp62. In the left hemisegment, a ventral secondary motor neuron (asterisk and arrow indicate soma and axon, respectively). In the neighboring hemisegment, on the right (caudal), there is a ventral/dorsal secondary motor neuron (asterisk indicates soma; arrows indicate the two axons, one projecting ventrally [bottom arrow] and the other dorsally [top arrow]). These neurons were labeled with a red fluorescent dye during the recordings. To identify ventral/dorsal secondary motor neurons, it is critical to ensure that the dissection does not remove muscle in the adjacent caudal hemisegment, thereby damaging or removing the dorsal axon. After the recording, the neuron soma remains attached to the micropipette as it is pulled away from the preparation (right top asterisk). When using dyes to fill neurons during recordings, the dye often leaks while the electrode is in the bath, resulting in a red fluorescent background visible in the spinal cord and notochord (bottom asterisk in rostral [left] hemisegment). (B) Voltage-clamp recordings were obtained from ventral and ventral/dorsal secondary motor neurons. Voltage steps (to -30, -10, +10, +30, +50, +70, +90, and +110 mV) elicited outward and inward currents. Unclamped action potentials/depolarizations may be present in the recordings (asterisks). (C) During current-clamp recordings from secondary motor neurons, brief (1 ms) current injections of increasing amplitude were applied to the neurons to trigger an action potential (asterisks). (C&#39;) Examples of single action potentials triggered in secondary motor neurons by ~0.4-nA current injections are shown. At this stage, spontaneous action potentials are also observed (C and C&#39;, arrowheads). (D) Prolonged (100 ms) current injections (~0.35 nA) trigger the repetitive firing of action potentials. The membrane potential was held at ~-65 mV. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55507/55507fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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 ...

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

Nanyang Technological University

Research

JoVE Journal

Neuroscience

11.4K Views.  University of Colorado Anschutz Medical Campus (UCAMC).  Dit manuscript beschrijft werkwijzen voor elektrofysiologische opnames van spinale neuronen van zebravis embryo's en larven. Het preparaat handhaaft neuronen in situ en vaak gaat om minimale dissectie. Deze werkwijzen maken het elektrofysiologisch onderzoek van verschillende spinale neuronen van de initiële elektrische prikkelbaarheid overname door de vroege larvale stadia. 

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

Paired Patch Clamp Recordings from Motor-neuron and Target Skeletal Muscle in Zebrafish

Larval zebrafish represent the first vertebrate model system to allow simultaneous patch clamp recording from a spinal motor-neuron and target muscle. This is a direct consequence of the accessibility to both cell types and ability to visually distinguish the single segmental CaP motor-neuron on the basis of morphology and location. This video demonstrates the microscopic methods used to identify a CaP motor-neuron and target muscle cells as well as the methodologies for recording from each cell type. Identification of the CaP motor-neuron type is confirmed by either dye filling or by the biophysical features such as action potential waveform and cell input resistance. Motor-neuron recordings routinely last for one hour permitting long-term recordings from multiple different target muscle cells. Control over the motor-neuron firing pattern enables measurements of the frequency-dependence of synaptic transmission at the neuromuscular junction. Owing to a large quantal size and the low noise provided by whole cell voltage clamp, all of the unitary events can be resolved in muscle. This feature permits study of basic synaptic properties such as release properties, vesicle recycling, as well as synaptic depression and facilitation. The advantages offered by this in vivo preparation eclipse previous neuromuscular model systems studied wherein the motor-neurons are usually stimulated by extracellular electrodes and the muscles are too large for whole cell patch clamp. The zebrafish preparation is amenable to combining electrophysiological analysis with a wide range of approaches including transgenic lines, morpholino knockdown, pharmacological intervention and in vivo imaging. These approaches, coupled with the growing number of neuromuscular disease models provided by mutant lines of zebrafish, open the door for new understanding of human neuromuscular disorders.

Larval zebrafish represent the first vertebrate model system to allow simultaneous patch clamp recording from a spinal motor-neuron and target skeletal muscle. This video demonstrates the microscopic methods used to identify a segmental CaP motor-neuron and target muscle cells as well as the methodologies for recording from each cell type.

Larval zebrafish represent the first vertebrate model system to allow simultaneous patch clamp recording from a spinal motor-neuron and target muscle.  ...

Biosensing Motor Neuron Membrane Potential in Live Zebrafish Embryos

The protocols described here are designed to allow researchers to study cell communication without altering the integrity of the environment in which the cells are located. Specifically, they have been developed to analyze the electrical activity of excitable cells, such as spinal neurons. In such a scenario, it is crucial to preserve the integrity of the spinal cell, but it is also important to preserve the anatomy and physiological shape of the systems involved. Indeed, the comprehension of the manner in which the nervous system-and other complex systems-works must be based on a systemic approach. For this reason, the live zebrafish embryo was chosen as a model system, and the spinal neuron membrane voltage changes were evaluated without interfering with the physiological conditions of the embryos. 
Here, an approach combining the employment of zebrafish embryos with a FRET-based biosensor is described. Zebrafish embryos are characterized by a very simplified nervous system and are particularly suited for imaging applications thanks to their transparency, allowing for the employment of fluorescence-based voltage indicators at the plasma membrane during zebrafish development. The synergy between these two components makes it possible to analyze the electrical activity of the cells in intact living organisms, without perturbing the physiological state. Finally, this non-invasive approach can co-exist with other analyses (e.g., spontaneous movement recordings, as shown here).

Protocols described here allow for the study of the electrical properties of excitable cells in the most non-invasive physiological conditions by employing zebrafish embryos in an in vivo system together with a fluorescence resonance energy transfer (FRET)-based genetically encoded voltage indicator (GEVI) selectively expressed in the cell type of interest.

The protocols described here are designed to allow researchers to study cell communication without altering the integrity of the environment in which the ...

Developmental Biology

Forebrain Electrophysiological Recording in Larval Zebrafish

Epilepsy affects nearly 3 million people in the United States and up to 50 million people worldwide. Defined as the occurrence of spontaneous unprovoked seizures, epilepsy can be acquired as a result of an insult to the brain or a genetic mutation. Efforts to model seizures in animals have primarily utilized acquired insults (convulsant drugs, stimulation or brain injury) and genetic manipulations (antisense knockdown, homologous recombination or transgenesis) in rodents. Zebrafish are a vertebrate model system1-3 that could provide a valuable alternative to rodent-based epilepsy research. Zebrafish are used extensively in the study of vertebrate genetics or development, exhibit a high degree of genetic similarity to mammals and express homologs for ~85% of known human single-gene epilepsy mutations. Because of their small size (4-6 mm in length), zebrafish larvae can be maintained in fluid volumes as low as 100 &mu;l during early development and arrayed in multi-well plates. Reagents can be added directly to the solution in which embryos develop, simplifying drug administration and enabling rapid in vivo screening of test compounds4. Synthetic oligonucleotides (morpholinos), mutagenesis, zinc finger nuclease and transgenic approaches can be used to rapidly generate gene knockdown or mutation in zebrafish5-7. These properties afford zebrafish studies an unprecedented statistical power analysis advantage over rodents in the study of neurological disorders such as epilepsy. Because the "gold standard" for epilepsy research is to monitor and analyze the abnormal electrical discharges that originate in a central brain structure (i.e., seizures), a method to efficiently record brain activity in larval zebrafish is described here. This method is an adaptation of conventional extracellular recording techniques and allows for stable long-term monitoring of brain activity in intact zebrafish larvae. Sample recordings are shown for acute seizures induced by bath application of convulsant drugs and spontaneous seizures recorded in a genetically modified fish.

A simple method to record extracellular field potentials in the larval zebrafish forebrain is described. The method provides a robust in vivo read-out of seizure-like activity. This technique can be used with genetically modified zebrafish larvae carrying epilepsy-related genes or seizures evoked by administration of convulsant drugs.

Epilepsy affects nearly 3 million people in the United States and up to 50 million people worldwide. Defined as the occurrence of spontaneous unprovoked ...

Patch Clamp Recordings from Embryonic Zebrafish Mauthner Cells

Mauthner cells (M-cells) are large reticulospinal neurons located in the hindbrain of teleost fish. They are key neurons involved in a characteristic behavior known as the C-start or escape response that occurs when the organism perceives a threat. The M-cell has been extensively studied in adult goldfish where it has been shown to receive a wide range of excitatory, inhibitory and neuromodulatory signals1. We have been examining M-cell activity in embryonic zebrafish in order to study aspects of synaptic development in a vertebrate preparation. In the late 1990s Ali and colleagues developed a preparation for patch clamp recording from M-cells in zebrafish embryos, in which the CNS was largely intact2,3,4. The objective at that time was to record synaptic activity from hindbrain neurons, spinal cord neurons and trunk skeletal muscle while maintaining functional synaptic connections within an intact brain-spinal cord preparation. This preparation is still used in our laboratory today. To examine the mechanisms underlying developmental synaptic plasticity, we record excitatory (AMPA and NMDA-mediated)5,6 and inhibitory (GABA and glycine) synaptic currents from developing M-cells. Importantly, this unique preparation allows us to return to the same cell (M-cell) from preparation to preparation to carefully examine synaptic plasticity and neuro-development in an embryonic organism. The benefits provided by this preparation include 1) intact, functional synaptic connections onto the M-cell, 2) relatively inexpensive preparations, 3) a large supply of readily available embryos 4) the ability to return to the same cell type (i.e. M-cell) in every preparation, so that synaptic development at the level of an individual cell can be examined from fish to fish, and 5) imaging of whole preparations due to the transparent nature of the embryos.

We have developed an intact brain-spinal cord preparation to record and monitor electrical activity via patch clamp recording from the Mauthner neurons and other reticulospinal cells in zebrafish embryos. Thus, we are able to record excitatory and inhibitory synaptic currents, voltage-gated channel activity and action potentials from key neurons in a developing embryo.

Mauthner cells (M-cells) are large  reticulospinal neurons located in the hindbrain of teleost fish. They are key  neurons involved in a characteristic ...

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

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

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

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

Electrophysiological Recording in the Brain of Intact Adult Zebrafish

Previously, electrophysiological studies in adult zebrafish have been limited to slice preparations or to eye cup preparations and electrorentinogram recordings. This paper describes how an adult zebrafish can be immobilized, intubated, and used for in vivo electrophysiological experiments, allowing recording of neural activity. Immobilization of the adult requires a mechanism to deliver dissolved oxygen to the gills in lieu of buccal and opercular movement. With our technique, animals are immobilized and perfused with habitat water to fulfill this requirement. A craniotomy is performed under tricaine methanesulfonate (MS-222; tricaine) anesthesia to provide access to the brain. The primary electrode is then positioned within the craniotomy window to record extracellular brain activity. Through the use of a multitube perfusion system, a variety of pharmacological compounds can be administered to the adult fish and any alterations in the neural activity can be observed. The methodology not only allows for observations to be made regarding changes in neurological activity, but it also allows for comparisons to be made between larval and adult zebrafish. This gives researchers the ability to identify the alterations in neurological activity due to the introduction of various compounds at different life stages.

This paper describes how an adult zebrafish can be immobilized, intubated, and used for in vivo electrophysiological experiments to allow recordings and manipulation of neural activity in an intact animal.

Previously, electrophysiological studies in adult zebrafish have been limited to slice preparations or to eye cup preparations and electrorentinogram ...

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 ...

Activity of Posterior Lateral Line Afferent Neurons during Swimming in Zebrafish

Sensory systems gather cues essential for directing behavior, but animals must decipher what information is biologically relevant. Locomotion generates reafferent cues that animals must disentangle from relevant sensory cues of the surrounding environment. For example, when a fish swims, flow generated from body undulations is detected by the mechanoreceptive neuromasts, comprising hair cells, that compose the lateral line system. The hair cells then transmit fluid motion information from the sensor to the brain via the sensory afferent neurons. Concurrently, corollary discharge of the motor command is relayed to hair cells to prevent sensory overload. Accounting for the inhibitory effect of predictive motor signals during locomotion is, therefore, critical when evaluating the sensitivity of the lateral line system. We have developed an in vivo electrophysiological approach to simultaneously monitor posterior lateral line afferent neuron and ventral motor root activity in zebrafish larvae (4-7 days post fertilization) that can last for several hours. Extracellular recordings of afferent neurons are achieved using the loose patch clamp technique, which can detect activity from single or multiple neurons. Ventral root recordings are performed through the skin with glass electrodes to detect motor neuron activity. Our experimental protocol provides the potential to monitor endogenous or evoked changes in sensory input across motor behaviors in an intact, behaving vertebrate.

We describe a protocol to monitor changes in the afferent neuron activity during motor commands in a model vertebrate hair cell system.

Sensory systems gather cues essential for directing behavior, but animals must decipher what information is biologically relevant. Locomotion generates ...

Controlled Semi-Automated Laser-Induced Injuries for Studying Spinal Cord Regeneration in Zebrafish Larvae

Zebrafish larvae possess a fully functional central nervous system (CNS) with a high regenerative capacity only a few days after fertilization. This makes this animal model very useful for studying spinal cord injury and regeneration. The standard protocol for inducing such lesions is to transect the dorsal part of the trunk manually. However, this technique requires extensive training and damages additional tissues. A protocol was developed for laser-induced lesions to circumvent these limitations, allowing for high reproducibility and completeness of spinal cord transection over many animals and between different sessions, even for an untrained operator. Furthermore, tissue damage is mainly limited to the spinal cord itself, reducing confounding effects from injuring different tissues, e.g., skin, muscle, and CNS. Moreover, hemi-lesions of the spinal cord are possible. Improved preservation of tissue integrity after laser injury facilitates further dissections needed for additional analyses, such as electrophysiology. Hence, this method offers precise control of the injury extent that is unachievable manually. This allows for new experimental paradigms in this powerful model in the future.

The present protocol describes a method to induce tissue-specific and highly reproducible injuries in zebrafish larvae using a laser lesion system combined with an automated microfluidic platform for larvae handling.

Zebrafish larvae possess a fully functional central nervous system (CNS) with a high regenerative capacity only a few days after fertilization. This makes ...

Medicine

A Non-invasive Approach to Study the Electrophysiology of Motor Neurons in Zebrafish Embryos

Source: Benedetti, L., et al. <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/55297">Biosensing Motor Neuron Membrane Potential in Live Zebrafish Embryos.</a> J. Vis. Exp. (2017).
This video demonstrates a non-invasive assay to study electrophysiology of motor neurons in zebrafish embryos. Upon measuring tail coiling in the embryos as a result of spontaneous depolarization of the spinal cord motor neurons, a sodium channel blocker is applied to reduce the coiling frequency. Next, the embryos are immobilized in agarose and placed under a fluorescence microscope. The genetically engineered motor neurons express a voltage-sensing biosensor with a donor and an acceptor fluorophore. A decrease in acceptor fluorophore emission indicates a reduction in the spontaneous depolarization of motor neurons due to the sodium channel blocker.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
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Zebrafish In Situ Spinal Cord Preparation for Electrophysiological Recordings from Spinal Sensory and Motor Neurons

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