The authors would like to thank Simona Rodighiero for her priceless support with the FRET imaging analysis.

1. pHuC_Mermaid Plasmid Generation
NOTE: Mermaid is a biosensor developed by pairing the voltage-sensing domain (VSD) of the Ciona intestinalis (now Ciona robusta)5 voltage sensor containing phosphatase (Ci-VSP) with the FRET partner fluorophores Umi-Kinoko Green (mUKG: donor) and a monomeric version of the orange-emitting fluorescent protein Kusabira Orange (mKOk: acceptor). For this biosensor, conformational changes of the VSD domain, induced by membrane depolarization, increase the proximity of the donor and acceptor fluorescent proteins, thus increasing the energy transfer between them (increasing the FRET Ratio)3. The VSD assures the efficient localization of the biosensor at the plasma membrane. The neuronal expression of the biosensor (in the spinal cord, the signal is detectable in both interneurons and motor neurons) is achieved by cloning the Mermaid open reading frame (ORF) under the zebrafish pan-neural promoter HuC6.
<ol>
	<li>Amplify by polymerase chain reaction (PCR) the Mermaid ORF from the pCS4+ Mermaid plasmid3 with Pfu (proofreading) DNA polymerase using the T3 Universal primer and the Mermaid SmaI primer (5&#8242;TATCCCGGGATTCGACGGTTCAGATTTTA) in order to insert a SmaI restriction site upstream of the Mermaid ORF.
	<ol>
		<li>Use the following PCR mixture: 0.5 &#181;L of Pfu DNA polymerase, 7.5 &#181;L of the specific 10x buffer, 1 &#181;L of 10 mM dNTPs, 2.5 &#181;L of dimethyl sulfoxide (DMSO), 0.5 &#181;L (50 ng) of the plasmid preparation, and 1 &#181;L of 20-&#181;M stock solutions of each primer in a total volume of 50 &#181;L.</li>
		<li>Heat the PCR mixture for 15 s at 95 &#176;C, prime it for 15 s at 50 &#176;C, and elongate it for 4 min at 72 &#176;C for a total of 35 cycles.</li>
	</ol>
	</li>
	<li>Gel-purify the specific blunt PCR product using a commercial gel purification kit, following the manufacturer&#39;s protocol.</li>
	<li>Clone the DNA fragment into the pCMV-SC blunt vector following the manufacturer&#39;s protocol.</li>
	<li>Linearize the Mermaid-positive plasmid (named pCMV-SC_Mermaid) with the SmaI restriction enzyme for the following insertion of the HuC promoter.
	<ol>
		<li>Set up the restriction reaction as follow: 0.5 &#181;L (10 U) of SmaI restriction enzyme, 2 &#181;L of the kit 10x buffer, and 5 &#181;g of plasmid DNA in a total volume of 20 &#181;L. Incubate the reaction at 25 &#176;C for 1 h.</li>
	</ol>
	</li>
	<li>Gel-purify the digested plasmid using a commercial gel purification kit following the manufacturer&#39;s protocol.</li>
	<li>PCR-amplify the HuC promoter (pHuC), with Pfu DNA polymerase and zebrafish genomic DNA as template, using a pair of HuC-specific primers (HuCprom-forw1_SalI: 5&#8242;-GTAGTCGACCAGACTTGTCAAAAGGGTCCA and HuCprom-rev1: 5&#8242;-TCCATTCTTGACGTACAAAGATG) and spanning a 3,150-bp region upstream of the ATG.
	<ol>
		<li>Set up the PCR mixture using the following scheme: 0.5 &#181;L of Pfu DNA polymerase, 7.5 &#181;L of the specific 10x buffer, 1 &#181;L of 10 mM dNTPs, 2.5 &#181;L of DMSO, 200 ng of genomic DNA, and 1 &#181;L of 20-&#181;M stock solutions of each primer in a total volume of 50 &#181;L.</li>
		<li>After an initial step of 2 min at 95 &#176;C, heat the PCR mixture for 15 s at 95 &#176;C, prime it for 15 s at 50&#176;C, and elongate it for 4 min at 72 &#176;C for a total of 35 cycles.</li>
	</ol>
	</li>
	<li>Gel-purify the specific blunt PCR product using a commercial gel purification kit following the manufacturer&#39;s protocol.</li>
	<li>Ligate an equimolar amount of the purified pCMV-SC_Mermaid (step 1.5) and the pHuC DNA (step 1.7) using 1 &#181;L of T4 DNA ligase and 1 &#181;L of the specific 10X buffer in a total volume of 10 &#181;L. Incubate the reaction for 16 h at 4 &#176;C.</li>
	<li>Transform an aliquot of competent cells with 5 &#181;L of the ligation reaction (step 1.8) following the manufacturer&#39;s instructions.</li>
	<li>Select the pHuC-positive clones (pHuC_Mermaid) with the promoter inserted in the proper orientation by a SalI-EcoRV double digestion (the SalI restriction site has been inserted upstream of the promoter fragment in step 1.6, while the EcoRV restriction site is positioned downstream of the polyadenylation region, PolyA, of the pCMV-SC plasmid).
	<ol>
		<li>Set up the restriction reaction as follow: 0.5 &#181;L (10 U) of both SalI and EcoRV restriction enzymes, 2 &#181;L of the kit10X buffer, and 1 &#181;g of pHuC_Mermaid DNA in a total volume of 20 &#181;L. Incubate the reaction at 37 &#176;C for 1 h. Run the reactions onto an agarose gel.</li>
	</ol>
	</li>
</ol>
2. Embryo Microinjection
<ol>
	<li>Transfer the fertilized eggs obtained from wild-type (AB strain) or Sod1-G93R adult zebrafish4 to a 10 mm Petri dish using a plastic pipette.</li>
	<li>Rinse the embryos in cold (4 &#176;C) fish water and immediately microinject them into the yolk with 200 pg of pHuC_Mermaid plasmid using a microinjector (for an overall and detailed description of the microinjection procedure, see References 7 and 8).</li>
	<li>Using a plastic pipette, transfer the embryos to a Petri dish and incubate them in fish water at 28 &#176;C until they reach the desired developmental stage (20-24 h post-fertilization, hpf) for the following analyses.</li>
</ol>
3. Spontaneous Tail Coiling Analysis
NOTE: Evaluate the spontaneous tail coiling behavior in 20-24 hpf embryos with or without the drug riluzole.
<ol>
	<li>Transfer an embryo to a 90-mm round Petri dish filled with fish water containing 0.2% DMSO (riluzole vehicle) and manually dechorionate it using two jeweler&#39;s forceps with sharp tips. Incubate the embryo for 5 min.</li>
	<li>Detect the tail coiling at RT during a 1 min video recording using a digital camera mounted on a stereomicroscope. Acquire time series at a time resolution of 30 frames/s.</li>
	<li>Calculate the frequency of spontaneous tail coilings by counting the number of bends (both contralateral and ipsilateral) per time unit.</li>
	<li>To evaluate the effect of the drug riluzole, gently use a plastic Pasteur pipette to transfer the embryo to a new 90 mm Petri dish filled with fish water containing 5 &#181;M riluzole.
	<ol>
		<li>Incubate the embryo for 5 min before recording a 1-min video and performing the behavioral analysis as above.</li>
	</ol>
	</li>
</ol>
4. Imaging Setup for Mermaid Biosensor Visualization in Living Embryos: Simultaneous Detection of Donor and Acceptor Signals
<ol>
	<li>Mount the 20 - 24 hpf embryos in 1% low-melting-point agarose in fish water at 37 &#176;C inside a 35 mm glass-bottomed imaging dish. Orient the embryos on their sides. Wait until the agarose solidifies at room temperature.</li>
	<li>Transfer the imaging dish to the stage of a inverted confocal mounted on an inverted microscope. Identify motor neurons expressing the biosensor with a 20X objective (0.7 numerical aperture, NA) by exciting the mUKG with the 488 nm argon laser line and recording its emission between 495 and 525 nm.</li>
	<li>For a FRET measurement, excite mUKG, the donor of the FRET pair, with the 488 nm laser line. Simultaneously detect, with a resonant scanner operating at 8,000 Hz in the bidirectional mode, the fluorescence emitted by the donor (between 495 and 525 nm) and the fluorescence emitted by the mKOk acceptor (between 550 and 650 nm, FRET channel). If available, use the 473 nm laser. 
	NOTE: The excitation efficiency of the donor will be slightly reduced (85% instead of 93% with 488 nm), but the cross-excitation of the acceptor will be reduced as well (from 17% with the 488 nm to 9% with 473 nm laser line).</li>
	<li>To reduce phototoxicity and fluorophore bleaching, minimize the illumination of the sample by reducing the power of the laser line (on the beam path window of the acquisition software).</li>
	<li>Optimize the excitation to match gain and offset parameters that are set at the beginning of the experiment and kept constant throughout the session. To set the offset, change the color of the image to intensity values (by using the Q look-up table) and, while scanning with the laser off, turn the offset knob (smart offset) so that the background pixels have an intensity slightly higher than zero. With the same look-up table, by switching the laser on while scanning, turn the gain knob (smart gain) to maximize the signal-to-noise ratio, being careful to avoid saturated pixels.</li>
	<li>Using an opened pinhole (2 airy units), acquire 16-bit images to provide a sufficient dynamic range for quantitative analyses. Avoid averaging to increase the acquisition speed and to minimize photobleaching.</li>
	<li>In the software acquisition window, select an image field size of 512 x 64 pixels (pixel size: 605 nm) from the drop-down menu.</li>
	<li>From the acquisition mode window, select xyt (time lapse on a single xy plane)from the drop-down menu and record the changes in embryo spinal neuron voltage by acquiring a single xy plane, setting the acquisition parameters to record one image every 30 ms for 1 min.</li>
	<li>To evaluate the effect of riluzole administration on membrane depolarization in the same neuron, acquire a new dataset 5 min after the addition of fish water containing 5 &#181;M riluzole.</li>
	<li>For FRET analysis, use the ImageJ macro Biosensor_FRET (expressing single-chain FRET biosensors.</li>
	<li>Evaluate the basal membrane FRET ratio of each neuron at t1 as ((FRET mean - FRET background)/(Donor mean - Donor background)), where FRET and Donor mean intensity is the mean fluorescence intensity calculated in the same region of interest (ROI) drawn around the cell for each channel acquired and FRET and Donor background is the mean fluorescence intensity calculated in an ROI of the field of view without the fluorescent sample. 
	NOTE: A detailed step-by-step description of the use of the plugin can be found at the <a href="http://www.med.unc.edu/microscopy/resources/imagej-plugins-and-macros/biosensor-fret">www.med.unc.edu/microscopy/resources/imagej-plugins-and-macros/biosensor-fret</a> website.</li>
	<li>Use a graphing software to compare the frequency, amplitude, and duration of depolarization between different experimental paradigms. Compare two groups using an unpaired Student&#39;s t-test and consider mean values as statistically different when P &#60;0.05.</li>
</ol>

<ol>
	<li>Kn&#246;pfel, T., Gallero-Salas, Y., Song, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genetically+encoded+voltage+indicators+for+large+scale+cortical+imaging+come+of+age.">Genetically encoded voltage indicators for large scale cortical imaging come of age.</a> Curr Opin Chem Biol. 27, 75-83 (2015).</li><li>Chemla, S., Chavane, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Voltage-sensitive+dye+imaging:+Technique+review+and+models.">Voltage-sensitive dye imaging: Technique review and models.</a> J Physiol Paris. 104 (1-2), 40-50 (2010).</li><li>Tsutsui, H., Karasawa, S., Okamura, Y., Miyawaki, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Improving+membrane+voltage+measurements+using+FRET+with+new+fluorescent+proteins.">Improving membrane voltage measurements using FRET with new fluorescent proteins.</a> Nat Methods. 5 (8), 683-685 (2008).</li><li>Benedetti, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=INaP+selective+inhibition+reverts+precocious+inter-+and+motorneurons+hyperexcitability+in+the+Sod1-G93R+zebrafish+ALS+model.">INaP selective inhibition reverts precocious inter- and motorneurons hyperexcitability in the Sod1-G93R zebrafish ALS model.</a> Sci Rep. 6, 24515 (2016).</li><li>Pennati, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Morphological+differences+between+larvae+of+the+Ciona+intestinalis+species+complex:+hints+for+a+valid+taxonomic+definition+of+distinct+species.">Morphological differences between larvae of the Ciona intestinalis species complex: hints for a valid taxonomic definition of distinct species.</a> PLoS ONE. 10 (5), 0122879 (2015).</li><li>Park, H. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+upstream+elements+in+the+HuC+promoter+leads+to+the+establishment+of+transgenic+zebrafish+with+fluorescent+neurons.">Analysis of upstream elements in the HuC promoter leads to the establishment of transgenic zebrafish with fluorescent neurons.</a> Dev Biol. 227 (2), 279-293 (2000).</li><li>Yuan, S., Sun, Z. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microinjection+of+mRNA+and+morpholino+antisense+oligonucleotides+in+zebrafish+embryos.">Microinjection of mRNA and morpholino antisense oligonucleotides in zebrafish embryos.</a> J Vis Exp. (27), (2009).</li><li>Rosen, J. N., Sweeney, M. F., Mably, J. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microinjection+of+zebrafish+embryos+to+analyze+gene+function.">Microinjection of zebrafish embryos to analyze gene function.</a> J Vis Exp. (25), (2009).</li><li>Drapeau, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+the+locomotor+network+in+zebrafish.">Development of the locomotor network in zebrafish.</a> Prog Neurobiol. 68 (2), 85-111 (2002).</li><li>Brustein, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Steps+during+the+development+of+the+zebrafish+locomotor+network.">Steps during the development of the zebrafish locomotor network.</a> J Physiol Paris. 97 (1), 77-86 (2003).</li><li>Drapeau, P., Ali, D. W., Buss, R. R., Saint-Amant, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+recording+from+identifiable+neurons+of+the+locomotor+network+in+the+developing+zebrafish.">In vivo recording from identifiable neurons of the locomotor network in the developing zebrafish.</a> J Neurosci Methods. 88, 1-13 (1999).</li><li>Kawakami, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tol2:+a+versatile+gene+transfer+vector+in+vertebrates.">Tol2: a versatile gene transfer vector in vertebrates.</a> Genome Biol. 8, (2007).</li><li>Sungmoo, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+Membrane+Potential+with+Two+Types+of+Genetically+Encoded+Fluorescent+Voltage+Sensors.">Imaging Membrane Potential with Two Types of Genetically Encoded Fluorescent Voltage Sensors.</a> J Vis Exp. (108), e53566 (2016).</li></ol>

The authors have nothing to disclose.

The protocol presented here allowed us to explore the association between the electrical properties of zebrafish embryo spinal motor neurons and the spontaneous coiling behavior, the earliest stereotypic motor activity, which appears around 17 hpf of embryonic development and lasts until 24 hpf10.
Our approach provides researchers with a tool to study the neural system of intact embryos, fully preserving the complexity of the interactions between cells in a developing functional network. The in vivo imaging of zebrafish embryos has been performed by simply immobilizing them through immersion in low-melting agarose. The study of changes in cell membrane potential through the use of a biosensor represents one of the main advantages of this method. Canonical electrophysiological approaches, where neurons must be physically accessed by removing the enveloping tissues11, are procedures that might alter the electrical properties of the cells under examination. Moreover, the FRET-based biosensor approach allows for the study of cell electrical properties without the use of anesthetics (i.e., Tricaine, a sodium channel blocker), avoiding any potential disturbance associated with the administration of chemicals (i.e., riluzole here), which might be crucial in the experimental plan. This method allowed us to focus on the modulation of both the electrical activity and the frequency of the spontaneous tail coilings, a behavioral response that prevents, in most cases, electrophysiological recordings without the administration of anesthetics. Finally, the employment of GEVIs offers the highest temporal experimental control because it allows work at a specific embryonic developmental stage. Our imaging/recording sessions spanned a very limited time window, allowing us to precisely compare the developmental stages of the different embryos in real time. All the conventional electrophysiological recordings, on the contrary, are usually conducted in larger and less precise time spans.
Measuring voltage changes by imaging techniques is, however, intrinsically difficult because of the nature of the signal itself, which can vary in speed, in frequency, and in the size of the changes in membrane potential. An ideal voltage sensor should combine fast responsiveness, high sensitivity, and high photon emission. For most FRET-based GEVIs, ratio changes vary as a function of membrane voltage variations, but the amplitude of the fluorescence ratio changes is tightly associated with the duration of the depolarization events. Recently, new GEVIs were developed and characterized, thus making it possible to choose the best probe in relation to the type of experimental issue, model, and imaging device.
Voltage biosensors are integral membrane proteins. When transfected into cultured cells or expressed in living organisms, the fluorescence of the plasma membrane-localized sensor will be, to a certain extent, associated with the fluorescence of the sensor in other intracellular compartments-where the protein can be mislocalized or aggregated-thus generating a background signal. The level of expression and the proper membrane localization of each construct should be monitored in simplified experimental models (i.e., transfecting cell lines or primary cultures) before using the biosensors in living organisms.
In the present experimental setup, another potential critical step of the protocol might be represented by the efficiency of the transgenesis procedure. If necessary, the percentage of plasmid-positive cells can be drastically increased by employing the Tol2 transposon system12. An additional crucial aspect that needs to be considered is the loss of fluorescence emission in the donor fluorophore and the loss in FRET signal occurring during the imaging session because of the intense illumination. This should be avoided by minimizing the illumination of the sample, reducing the power of the laser line used. Similarly, the gain and offset of the photomultipliers must be carefully tuned at the beginning of every recording session to achieve the best signal-to-noise ratio in the acquired images as well as to avoid saturated pixels. All these parameters can vary depending on the type of cell examined, the characteristics of the biosensor, and the instruments used for image acquisition (wide field fluorescence microscopes can be also used, as recently described13). The researcher should always consider, however, that during in vivo imaging, the noise level might interfere with the specific signal of the sensor more than in in vitro recordings. Finally, if available, control measurements using mutant probes, which do not respond to voltage, would be a convenient way to evaluate the level of system-specific noise and/or artifacts.
The employment of zebrafish embryos represents a key feature of this approach thanks to their responsiveness to genetic manipulation and microscopic investigation. In fact, these characteristics make transient transgenesis and the FRET-based analysis of neuronal electrical properties feasible. In our opinion, with the best-suited promoter and biosensor combinations, it would potentially be possible to selectively analyze the electrical activity of any cell type of interest and to parallel such activity with the embryonic phenotype.

In vivo systemic component analysis allows scientists to investigate cellular behavior in the most reliable way. This is particularly true when the activity under scrutiny is heavily influenced by cell-cell interactions (both contact- and non-contact-dependent), as in the nervous system, where membrane voltage changes drive the communication among excitable cells. The comprehension of the information encoded by these electrical signals is the key to understanding the way the nervous system works in both physiological and disease states.
In order to study cell electrical properties in the most non-invasive physiological conditions, several genetically encoded voltage indicators have been recently developed1. As opposed to the previous generations of optical voltage sensors (mainly voltage-sensitive dyes)2, GEVIs allow for in vivo analyses of the intact neural system, and their expression can be limited to specific cell types or populations.
The zebrafish embryo is the in vivo &#34;substrate&#34; of choice to take advantage of the great potential attributed to GEVIs. In fact, thanks to its optical clarity and its simplified yet evolutionarily conserved nervous system, the zebrafish model allows for the straightforward identification and manipulation of every cellular component in a network. Indeed, the employment of the FRET-based GEVI Mermaid3 led to the identification of pre-symptomatic alterations in spinal motor neuron behavior in a zebrafish model of amyotrophic lateral sclerosis (ALS)4.
The following in vivo protocol describes how to monitor the electrical properties of spinal motor neurons in intact zebrafish embryos expressing Mermaid in a neuronal-specific manner. Moreover, it demonstrates how pharmacologically induced changes in such electrical properties can be associated with alterations in the frequency of embryonic spontaneous coilings, the stereotypic motor activity that characterizes the movement behavior of the zebrafish at very early stages of development.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Low Melting Point Agarose&#160;</td>
			<td>Sigma-Aldrich</td>
			<td>A9414</td>
		</tr>
		<tr>
			<td>DMSO&#160;</td>
			<td>Sigma-Aldrich</td>
			<td>W387520</td>
		</tr>
		<tr>
			<td>Riluzole</td>
			<td>Sigma-Aldrich</td>
			<td>R116</td>
		</tr>
		<tr>
			<td>Pfu Ultra HQ DNA polymerase&#160;</td>
			<td>Agilent Technologies - Stratagene Products Division</td>
			<td>600389</td>
		</tr>
		<tr>
			<td>T3 Universal primer&#160;</td>
			<td>Sigma-Aldrich</td>
			<td></td>
		</tr>
		<tr>
			<td>Wizard SV Gel and PCR Clean-Up system</td>
			<td>Promega</td>
			<td>A9280</td>
		</tr>
		<tr>
			<td>Universal SmaI primer&#160;</td>
			<td>Eurofins</td>
			<td></td>
		</tr>
		<tr>
			<td>StrataClone Mammalian Expression Vector System / pCMV-SC blunt vector&#160;</td>
			<td>Agilent Technologies - Stratagene Products Division&#160;</td>
			<td>240228</td>
		</tr>
		<tr>
			<td>SmaI&#160;</td>
			<td>New England Biolabs</td>
			<td>R0141S</td>
		</tr>
		<tr>
			<td>T4 DNA ligase</td>
			<td>Promega</td>
			<td>M1801</td>
		</tr>
		<tr>
			<td>SalI</td>
			<td>New England Biolabs</td>
			<td>R0138S</td>
		</tr>
		<tr>
			<td>EcoRV</td>
			<td>New England Biolabs</td>
			<td>R0195S</td>
		</tr>
		<tr>
			<td>35 mm, glass-bottomed imaging dish&#160;</td>
			<td>Ibidi</td>
			<td>81151</td>
		</tr>
		<tr>
			<td>forceps</td>
			<td>Sigma-Aldrich</td>
			<td>F6521</td>
		</tr>
		<tr>
			<td>Stereomicroscope</td>
			<td>Leica Microsystems</td>
			<td>M10 F</td>
		</tr>
		<tr>
			<td>Digital camera</td>
			<td>Leica Microsystems</td>
			<td>DFC 310 FX</td>
		</tr>
		<tr>
			<td>Leica Application Suite 4.7.1 software&#160;</td>
			<td>Leica Microsystems</td>
			<td></td>
		</tr>
		<tr>
			<td>QuickTime Player, v10.4</td>
			<td>Apple</td>
			<td></td>
		</tr>
		<tr>
			<td>Confocal microscope (inverted)</td>
			<td>Leica Microsystems</td>
			<td>TCS SP5</td>
		</tr>
		<tr>
			<td>Microinjector&#160;</td>
			<td>Eppendorf&#160;</td>
			<td>Femtojet</td>
		</tr>
		<tr>
			<td>ImageJ macro Biosensor_FRET&#160;</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>GraphPad Prism 6.0c&#160;</td>
			<td>GraphPad Software, Inc</td>
			<td></td>
		</tr>
	</tbody>
</table>

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).

An expression vector carrying the FRET-based Mermaid biosensor coding sequence under the control of the pHuC pan-neuronal promoter, which drives the synthesis of the protein exclusively in the nervous system, was delivered into single-cell fertilized eggs by means of a microinjection in order to obtain transient transgenic embryos (Figure 1, left panel). After mastering the microinjection technique, the percentage of Mermaid-positive embryos was close to 100%. Among these, only the embryos expressing the proper amount of Mermaid biosensor at the plasma membrane were considered for FRET analysis/image acquisition.
Spontaneous coilings, motor responses that consist exclusively of a full body contraction bringing the tip of the tail to the head9,10, were recorded at the stage of 20-24 hpf in regular fish medium (Figure 1, center panel and Video 1) and following incubation in 5 &#181;M riluzole (Video 2). The statistical analyses of the changes in the frequency of the spontaneous tail coilings triggered by the drug riluzole are reported elsewhere4.
To test whether alterations in the electrical activity of the spinal motor neurons were at the basis of the riluzole-induced changes in the spontaneous coiling frequency, the membrane potential in embryo spinal motor neurons were studied in a non-invasive fashion, measuring the fluorescence intensity ratio of the donor/acceptor FRET pair of the Mermaid biosensor (FRET ratio: Figure 1, right panel). Primary spinal motor neurons expressing the biosensor were identified (Figure 2A) and their basal spontaneous depolarization activities were recorded (Figure 2B and Videos 3 and 4). Together with the reduction in the frequency of tail coiling movements, riluzole administration reduced the frequency of spontaneous depolarization events (Figure 2C).
<img alt="Figure 1" src="/files/ftp_upload/55297/55297fig1.jpg" /> 
Figure 1: Flow Chart of the Experimental Procedure. Single-cell-stage embryos are collected and immediately microinjected with the pHuC_Mermaid plasmid to express the Mermaid FRET-voltage biosensor in a pan-neuronal fashion. At 20-24 hpf with incubation at 28 &#176;C, single embryos are transferred under a stereomicroscope and their spontaneous tail coiling activity, with and without the drug riluzole present in the fish water, is video recorded and analyzed. The same embryos undergo in vivo FRET analysis in order to measure changes in motor neuron membrane potential. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55297/55297fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/55297/55297fig2.jpg" /> 
Figure 2: In Vivo FRET Analysis to Measure Changes in Motor Neuron Membrane Potential. (A) Wild-type zebrafish embryos with the mosaic expression of the FRET-based voltage biosensor Mermaid in the spinal neuron. (a) The bright field image shows the area of the zebrafish trunk identified by the black box (sc: spinal cord; ms: myoseptum; m: myotome). By superimposing on (a) the fluorescence signal detected (b), the efficient expression of the biosensor in a primary spinal motor neuron (PM) can be clearly visualized. The donor (c) and the FRET (d) channel detected are shown with the respective green and magenta look-up tables (LUT) on the right side. All the embryos are mounted in a cranial-to-caudal orientation. Scale bar = 10 &#181;m. (B) The FRET ratio map displays the FRET ratio calculated every 0.03 s in the same motor neuron. It shows the basal spontaneous depolarization activity that the cell undergoes. For the Mermaid biosensor, the increase in the FRET ratio occurs when the membrane potential increases. Scale bar = 10 &#181;m.&#160;(C) Representative example of spontaneous mean FRET ratio changes occurring in a spinal motor neuron of a Sod1-G93R zebrafish embryo during a 1 min recording. Riluzole administration reduces the frequency of the spontaneous depolarizations. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55297/55297fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Movie 1" src="/files/ftp_upload/55297/55297video1.jpg" /> 
Movie 1: Spontaneous Tail Coiling in 24-hpf Wild-typecontrol Embryos. Representative recording showing spontaneous tail coiling in a 24 hpf wild-type control embryo incubated in fish water. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55297/Movie_1.mp4" target="_blank">Please click here to view this video. (Right-click to download.)</a>
<img alt="Movie 2" src="/files/ftp_upload/55297/55297video2.jpg" /> 
Movie 2: Spontaneous Tail Coiling in 24-hpf Wild-typeriluzole-treated Embryos. Representative recording showing spontaneous tail coiling in a 24 hpf wild-type control embryo incubated in 5 &#181;M riluzole (+R). Compared to controls, +R embryos showed a significant decrease in the frequency of spontaneous tail coiling behavior at 24 hpf. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55297/Movie_2.mp4" target="_blank">Please click here to view this video. (Right-click to download.)</a>
<img alt="Movie 3" src="/files/ftp_upload/55297/55297video3.jpg" /> 
Movie 3: Basal Spontaneous Depolarization Activity of a Primary Motor Neuron in the Spinal Cord of a Wild-type Zebrafish Embryo at 20 hpf. The Mermaid biosensor is so sensitive that it can detect the rapid changes in motor neuron membrane potential in the spinal cords of intact embryos. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55297/Movie_3.mov" target="_blank">Please click here to view this video. (Right-click to download.)</a>
<img alt="Movie 4" src="/files/ftp_upload/55297/55297video4.jpg" /> 
Movie 4: Spontaneous Depolarization Activity is Coupled to Muscular Contraction. A secondary motor neuron in the spinal cord of a wild-type zebrafish embryo at 20 hpf undergoes spontaneous depolarization. In this case, each depolarization is associated to a contraction of the zebrafish muscles, organized in a functionally synchronized syncytium at this developmental stage. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55297/Movie_4.mov" target="_blank">Please click here to view this video. (Right-click to download.)</a>

Watch this Scientific Journal Video about Biosensing Motor Neuron Membrane Potential in Live Zebrafish Embryos at JoVE.com

Biosensing Motor Neuron Membrane Potential in Live Zebrafish Embryos

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 ...

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6.6K Views.  Università degli Studi di Milano. 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.

Video: Biosensing Motor Neuron Membrane Potential in Live Zebrafish Embryos

Mechanical Vessel Injury in Zebrafish Embryos

Zebrafish (Danio rerio) embryos have proven to be a powerful model for studying a variety of developmental and disease processes. External development and optical transparency make these embryos especially amenable to microscopy, and numerous transgenic lines that label specific cell types with fluorescent proteins are available, making the zebrafish embryo an ideal system for visualizing the interaction of vascular, hematopoietic, and other cell types during injury and repair in vivo. Forward and reverse genetics in zebrafish are well developed, and pharmacological manipulation is possible. We describe a mechanical vascular injury model using micromanipulation techniques that exploits several of these features to study responses to vascular injury including hemostasis and blood vessel repair. Using a combination of video and timelapse microscopy, we demonstrate that this method of vascular injury results in measurable and reproducible responses during hemostasis and wound repair. This method provides a system for studying vascular injury and repair in detail in a whole animal model.

This article describes a method for creating a mechanical vessel injury in zebrafish embryos. This injury model provides a platform for studying hemostasis, injury-related inflammation, and wound healing in an organism ideally suited for real-time microscopy.

Zebrafish (Danio rerio) embryos have proven to be a powerful model for studying a variety of developmental and disease processes.  External development ...

Isolation and Characterization of Single Cells from Zebrafish Embryos

The zebrafish (Danio rerio) is a powerful model organism to study vertebrate development. Though many aspects of zebrafish embryonic development have been described at the morphological level, little is known about the molecular basis of cellular changes that occur as the organism develops. With recent advancements in microfluidics and multiplexing technologies, it is now possible to characterize gene expression in single cells. This allows for investigation of heterogeneity between individual cells of specific cell populations to identify and classify cell subtypes, characterize intermediate states that occur during cell differentiation, and explore differential cellular responses to stimuli. This study describes a protocol to isolate viable, single cells from zebrafish embryos for high throughput multiplexing assays. This method may be rapidly applied to any zebrafish embryonic cell type with fluorescent markers. An extension of this method may also be used in combination with high throughput sequencing technologies to fully characterize the transcriptome of single cells. As proof of principle, the relative abundance of cardiac differentiation markers was assessed in isolated, single cells derived from nkx2.5 positive cardiac progenitors. By evaluation of gene expression at the single cell level and at a single time point, the data support a model in which cardiac progenitors coexist with differentiating progeny. The method and work flow described here is broadly applicable to the zebrafish research community, requiring only a labeled transgenic fish line and access to microfluidics technologies.

This protocol describes a method for isolating single cells from zebrafish embryos, enriching for cells of interest, capturing zebrafish cells in microfluidic based single cell multiplex systems, and assessing gene expression from single cells.

The zebrafish (Danio rerio) is a powerful model organism to study vertebrate development. Though many aspects of zebrafish embryonic development have been ...

Observing Mitotic Division and Dynamics in a Live Zebrafish Embryo

Mitosis is critical for organismal growth and differentiation. The process is highly dynamic and requires ordered events to accomplish proper chromatin condensation, microtubule-kinetochore attachment, chromosome segregation, and cytokinesis in a small time frame. Errors in the delicate process can result in human disease, including birth defects and cancer. Traditional approaches investigating human mitotic disease states often rely on cell culture systems, which lack the natural physiology and developmental/tissue-specific context advantageous when studying human disease. This protocol overcomes many obstacles by providing a way to visualize, with high resolution, chromosome dynamics in a vertebrate system, the zebrafish. This protocol will detail an approach that can be used to obtain dynamic images of dividing cells, which include: in vitro transcription, zebrafish breeding/collecting, embryo embedding, and time-lapse imaging. Optimization and modifications of this protocol are also explored. Using H2A.F/Z-EGFP (labels chromatin) and mCherry-CAAX (labels cell membrane) mRNA-injected embryos, mitosis in AB wild-type, auroraBhi1045, and esco2hi2865 mutant zebrafish is visualized. High resolution live imaging in zebrafish allows one to observe multiple mitoses to statistically quantify mitotic defects and timing of mitotic progression. In addition, observation of qualitative aspects that define improper mitotic processes (i.e., congression defects, missegregation of chromosomes, etc.) and improper chromosomal outcomes (i.e., aneuploidy, polyploidy, micronuclei, etc.) are observed. This assay can be applied to the observation of tissue differentiation/development and is amenable to the use of mutant zebrafish and pharmacological agents. Visualization of how defects in mitosis lead to cancer and developmental disorders will greatly enhance understanding of the pathogenesis of disease.

Mitosis is critical to every living organism and defects often lead to cancer and developmental disorders. Using this imaging protocol and zebrafish as a model system, researchers can visualize mitosis in a live vertebrate organism and the multitude of defects that arise when mitotic processes are defective.

Mitosis is critical for organismal growth and differentiation. The process is highly dynamic and requires ordered events to accomplish proper chromatin ...

Affinity Labeling Detection of Endogenous Receptors from Zebrafish Embryos

By combining the powers of Affinity Labeling and Immunoprecipitation (AFLIP), a technique for the detection of low abundance receptors in zebrafish embryos has been implemented. This technique takes advantage of the selectivity and sensitivity conferred by affinity labeling of a given receptor by its ligand with the specificity of the immunoprecipitation. We have used AFLIP to detect the type III TGF-&#946; receptor (TGFBR3), also know as betaglycan, during early zebrafish development. AFLIP was instrumental in validating the efficacy of a TGFBR3 morphant zebrafish phenotype. In the first step, embryo protein extracts are prepared and used to generate 125I-TGF-&#946;2-TGFBR3 complexes that are purified by immunoprecipitation. Later, these complexes are covalently cross-linked and revealed using SDS-PAGE separation and autoradiography detection. This technique requires the availability of a labeled ligand for, and a specific antibody against, the receptor to be detected, and shall be easily adapted to identify any growth factor or cytokine receptor that meets these requirements.

A novel technique for the detection of low abundance endogenous receptors present in zebrafish embryos is described. We have named it AFLIP because it consists of affinity labeling of the receptor by its ligand linked to immunoprecipitation.

By combining the powers of Affinity Labeling and Immunoprecipitation (AFLIP), a technique for the detection of low abundance receptors in zebrafish ...

Ploidy Manipulation of Zebrafish Embryos with Heat Shock 2 Treatment

Manipulation of ploidy allows for useful transformations, such as diploids to tetraploids, or haploids to diploids. In the zebrafish Danio rerio, specifically the generation of homozygous gynogenetic diploids is useful in genetic analysis because it allows the direct production of homozygotes from a single heterozygous mother. This article describes a modified protocol for ploidy duplication based on a heat pulse during the first cell cycle, Heat Shock 2 (HS2). Through inhibition of centriole duplication, this method results in a precise cell division stall during the second cell cycle. The precise one-cycle division stall, coupled to unaffected DNA duplication, results in whole genome duplication. Protocols associated with this method include egg and sperm collection, UV treatment of sperm, in vitro fertilization and heat pulse to cause a one-cell cycle division delay and ploidy duplication. A modified version of this protocol could be applied to induce ploidy changes in other animal species.

A modified protocol for ploidy manipulation uses a heat shock to induce a one-cycle stall in cytokinesis in the early embryo. This protocol is demonstrated in the zebrafish but may be applicable to other species.

Manipulation of ploidy allows for useful transformations, such as diploids to tetraploids, or haploids to diploids. In the zebrafish Danio rerio, ...

Induction of Hypoxia in Living Frog and Zebrafish Embryos

Here, we introduce a novel system for hypoxia induction, which we developed to study the effects of hypoxia in aquatic organisms such as frog and zebrafish embryos. Our system comprises a chamber featuring a simple setup that is nevertheless robust to induce and maintain a specific oxygen concentration and temperature in any experimental solution of choice. The presented system is very cost-effective but highly functional, it allows induction and sustainment of hypoxia for direct experiments in vivo and for various time periods up to 48 h.
To monitor and study the effects of hypoxia, we have employed two methods - measurement of levels of hypoxia-inducible factor 1alpha (HIF-1&#945;) in whole embryos or specific tissues and determination of retinal stem cell proliferation by 5-ethynyl-2&#8242;-deoxyuridine (EdU) incorporation into the DNA. HIF-1&#945; levels can serve as a general hypoxia marker in the whole embryo or tissue of choice, here embryonic retina. EdU incorporation into the proliferating cells of embryonic retina is a specific output of hypoxia induction. Thus, we have shown that hypoxic embryonic retinal progenitors decrease proliferation within 1 h of incubation under 5% oxygen of both frog and zebrafish embryos.
Once mastered, our setup can be employed for use with small aquatic model organisms, for direct in vivo experiments, any given time period and under normal, hypoxic or hyperoxic oxygen concentration or under any other given gas mixture.

We introduce a novel hypoxic chamber system for use with aquatic organisms such as frog and zebrafish embryos. Our system is simple, robust, cost-effective and allows the induction and sustainment of hypoxia in vivo and for up to 48 h. We present 2 reproducible methods to monitor the effectiveness of hypoxia.

Here, we introduce a novel system for hypoxia induction, which we developed to study the effects of hypoxia in aquatic organisms such as frog and ...

In Vitro Ovule Cultivation for Live-cell Imaging of Zygote Polarization and Embryo Patterning in Arabidopsis thaliana

In most flowering plants, the zygote and embryo are hidden deep in the mother tissue, and thus it has long been a mystery of how they develop dynamically; for example, how the zygote polarizes to establish the body axis and how the embryo specifies various cell fates during organ formation. This manuscript describes an in vitro ovule culture method to perform live-cell imaging of developing zygotes and embryos of Arabidopsis thaliana. The optimized cultivation medium allows zygotes or early embryos to grow into fertile plants. By combining it with a poly(dimethylsiloxane) (PDMS) micropillar array device, the ovule is held in the liquid medium in the same position. This fixation is crucial to observe the same ovule under a microscope for several days from the zygotic division to the late embryo stage. The resulting live-cell imaging can be used to monitor the real-time dynamics of zygote polarization, such as nuclear migration and cytoskeleton rearrangement, and also the cell division timing and cell fate specification during embryo patterning. Furthermore, this ovule cultivation system can be combined with inhibitor treatments to analyze the effects of various factors on embryo development, and with optical manipulations such as laser disruption to examine the role of cell-cell communication.

In Vitro Ovule Cultivation for Live-cell Imaging of Zygote Polarization and Embryo Patterning in Arabidopsis thaliana

This manuscript describes an in vitro ovule cultivation method that enables live-cell imaging of Arabidopsis zygotes and embryos. This method is utilized to visualize the intracellular dynamics during zygote polarization and the cell fate specification in developing embryos.

In most flowering plants, the zygote and embryo are hidden deep in the mother tissue, and thus it has long been a mystery of how they develop dynamically; ...

Visualization of Cellular Electrical Activity in Zebrafish Early Embryos and Tumors

Bioelectricity, endogenous electrical signaling mediated by ion channels and pumps located on the cell membrane, plays important roles in signaling processes of excitable neuronal and muscular cells and many other biological processes, such as embryonic developmental patterning. However, there is a need for in vivo electrical activity monitoring in vertebrate embryogenesis. The advances of genetically encoded fluorescent voltage indicators (GEVIs) have made it possible to provide a solution for this challenge. Here, we describe how to create a transgenic voltage indicator zebrafish using the established voltage indicator, ASAP1 (Accelerated Sensor of Action Potentials 1), as an example. The Tol2 kit and a ubiquitous zebrafish promoter, ubi, were chosen in this study. We also explain&#160;the processes of Gateway site-specific cloning, Tol2 transposon-based zebrafish transgenesis, and the imaging process for early-stage fish embryos and fish tumors using regular epifluorescent microscopes. Using this fish line, we found that there are cellular electric voltage changes during zebrafish embryogenesis, and fish larval movement. Furthermore, it was observed that in a few zebrafish malignant peripheral nerve sheath tumors, the tumor cells were generally polarized compared to the surrounding normal tissues.

Here, we show the process of creating a cellular electric voltage reporter zebrafish line to visualize embryonic development, movement, and fish tumor cells in vivo.

Bioelectricity, endogenous electrical signaling mediated by ion channels and pumps located on the cell membrane, plays important roles in signaling ...

Visualize Drosophila Leg Motor Neuron Axons Through the Adult Cuticle

The majority of work on the neuronal specification has been carried out in genetically and physiologically tractable models such as C. elegans, Drosophila larvae, and fish, which all engage in undulatory movements (like crawling or swimming) as their primary mode of locomotion. However, a more sophisticated understanding of the individual motor neuron (MN) specification&#8212;at least in terms of informing disease therapies&#8212;demands an equally tractable system that better models the complex appendage-based locomotion schemes of vertebrates. The adult Drosophila locomotor system in charge of walking meets all of these criteria with ease, since in this model it is possible to study the specification of a small number of easily distinguished leg MNs (approximately 50 MNs per leg) both using a vast array of powerful genetic tools, and in the physiological context of an appendage-based locomotion scheme. Here we describe a protocol to visualize the leg muscle innervation in an adult fly.

Visualize Drosophila Leg Motor Neuron Axons Through the Adult Cuticle

Here we describe a protocol to visualize the axonal targeting with a florescent protein in adult legs of Drosophila by fixation, mounting, imaging, and post-imaging steps.

The majority of work on the neuronal specification has been carried out in genetically and physiologically tractable models such as C. elegans, Drosophila ...

Whole Mount Immunohistochemistry in Zebrafish Embryos and Larvae

Immunohistochemistry is a widely used technique to explore protein expression and localization during both normal developmental and disease states. Although many immunohistochemistry protocols have been optimized for mammalian tissue and tissue sections, these protocols often require modification and optimization for non-mammalian model organisms. Zebrafish are increasingly used as a model system in basic, biomedical, and translational research to investigate the molecular, genetic, and cell biological mechanisms of developmental processes. Zebrafish offer many advantages as a model system but also require modified techniques for optimal protein detection. Here, we provide our protocol for whole-mount fluorescence immunohistochemistry in zebrafish embryos and larvae. This protocol additionally describes several different mounting strategies that can be employed and an overview of the advantages and disadvantages each strategy provides. We also describe modifications to this protocol to allow detection of chromogenic substrates in whole mount tissue and fluorescence detection in sectioned larval tissue. This protocol is broadly applicable to the study of many developmental stages and embryonic structures.

Here, we present a protocol for fluorescent antibody-mediated detection of proteins in whole preparations of zebrafish embryos and larvae.

Immunohistochemistry is a widely used technique to explore protein expression and localization during both normal developmental and disease states. ...