The research work reported in this publication was supported by the National Institute of General Medical Sciences of the National Institutes of Health under the Award Number R35GM124913, Purdue University PI4D incentive program, and PVM Internal Competitive Basic Research Funds Program. The content is solely the responsibility of the authors and does not necessarily represent the official views of the funding agents. We thank Koichi Kawakami for the Tol2 construct, Michael Lin for the ASAP1 construct, and Leonard Zon for the ubi promoter construct through Addgene.

The zebrafish are housed in an AAALAC-approved animal facility, and all experiments were carried out according to the protocols approved by the Purdue Animal Care and Use Committee (PACUC).
1. Tol2 Transposon Plasmid Construct Preparation
NOTE: Tol2, a transposon that was discovered in medaka fish, has widely been used in the zebrafish research community8,9. It has been successfully adopted to the Gateway site-specific recombination-based cloning system and known as the Tol2 kit10. The Tol2 kit allows for a more convenient way of creating customized expression constructs, while also increasing the efficiency of transgenesis. Thus, it was an easy decision to take advantage of this system, and create a ubiquitous ASAP1 expression zebrafish line using a validated ubiquitin promoter to drive ASAP111.
<ol>
	<li>Creating a middle entry ASAP1 construct: pDONR221-ASAP1
	<ol>
		<li>Acquire the genetically encoded voltage sensor ASAP1 construct,&#160;pcDNA3.1/Puro-CAG-ASAP1 (Plasmid#52519), from Addgene. To amplify the ASAP1 coding region, set up a PCR using the customized primers (attB1-ASAP1F and attB2-ASAP1R) flanked with attB sequences at the 5&#39; end of the primers (Figure 1). Phusion DNA polymerase was chosen for its high efficiency, and PCR conditions were optimized based on the previously published protocol12.</li>
		<li>Load the 50 &#181;L PCR products into a 1% TAE gel using a regular pipette with 200 &#181;L tips, and perform electrophoresis at 160 V in a horizontal gel tank for about 30 min.</li>
		<li>Check the gel under a UV transilluminator (353 nm), excise out the desired band using a blade/scalpel under a UV transilluminator as previously published13,14, and put the DNA containing gel sample into a clean 1.5 mL microcentrifuge tube.</li>
		<li>Perform gel purification for the ASAP1 PCR products. Recover the DNA in the excised gel using a commercial DNA gel purification kit following manufacturer&#39;s instruction, and elute the DNA into 20 &#181;L of water. Take 1 &#181;L as a sample, and measure the DNA concentration using a spectrophotometer. Flow the software instructions using water as a blank control15.</li>
		<li>Take 100 ng of purified PCR product and mix it with 150 ng of pDONR221 plasmid in 10 &#181;L of TE buffer (10 mM Tris, 1 mM EDTA pH 8.0)16. Add 2 &#181;L of BP Clonase II into the reaction and incubate the reaction at room temperature overnight.</li>
		<li>On the second day, add 1 &#181;L of proteinase K into the reaction and incubate the reaction at 37 &#176;C for 30 min.</li>
		<li>Perform transformation. Transfer the reaction and mix it with 50 &#181;L of Top10 competent E. coli cells and incubate the cells on ice for 30 min. Then, transfer the reaction tube into a 42 &#176;C water bath for 1 min. Immediately remove the tube and incubate on ice for 2 min. Next, put the tube into a 37 &#176;C shaker and incubate it for 1 h.</li>
		<li>Take the tube out and plate the cells onto a kanamycin LB plate. Next, incubate the plate overnight (16-18 h) at 37 &#176;C.</li>
		<li>Pick single and well-separated colonies and inoculate them into 14 mL cell culture tubes with 3 mL of LB medium. Culture them overnight at 37 &#176;C in a shaker with a rotation speed of 250 rpm (rotation per minute).</li>
		<li>Perform miniprep using a commercial miniprep kit following its instruction manual17.</li>
		<li>Sequence 3-4 plasmids with Sanger sequencing to identify positive pDONR221-ASAP1 clones using M13F and M13R sequencing primers.</li>
	</ol>
	</li>
	<li>Creating the Tol2 construct for microinjection: pDestTol2-ubi-ASAP1
	<ol>
		<li>Choose the sequencing verified pDONR221-ASAP1 clone and measure its DNA concentration using a spectrophotometer following&#160;the software instruction using water as a blank control15.</li>
		<li>Take 100 &#181;g of pDONR221-ASAP1 and mix it with 100 &#181;g of Tol2 kit 5-end plasmid (pENTR5&#39;_ubi, Addgene #27320), 100 &#181;g of p3E-polyA (Tol2 kit #302) and 150 &#181;g of pDestTol2pA2 (Tol2 kit #394). Adjust total volume to 8 &#181;L using TE buffer (10 mM Tris, 1 mM EDTA, pH 8.0) in a 1.5 mL microcentrifuge tube and mix well by a 2-5 s brief vortex. Then, add 2 &#181;L of LR Clonase II plus, and incubate the reaction at room temperature overnight.</li>
		<li>On the second day, add 1 &#181;L of proteinase K into the reaction using a 10 &#181;L pipette, and incubate the reaction at 37 &#176;C for 30 min.</li>
		<li>Perform transformation and identify positive clones as described above (steps 1.1.7-11).</li>
		<li>Measure the DNA concentration of sequencing verified pDestTol2-ubi-ASAP1 clone (Figure 1B) using a spectrophotometer15. Usually the concentration is around 200ng/&#181;L.</li>
	</ol>
	</li>
</ol>
2. Prepare Tol2 Transposase mRNA and Injection Solution
<ol>
	<li>Streak E. coli glycerol stock of pCS2FA-transposase plasmid (Tol2 kit #396) onto a LB plate (with 100 &#181;g/mL ampicillin) using a sterilized inoculation loop. Incubate the plate at 37 &#176;C in an incubator overnight. The next day, pick a single colony and inoculate it into 3 mL of LB (100 &#181;g/mL ampicillin) using a sterilized 10 &#181;L pipette tip. Culture it overnight at 37 &#176;C in a shaker with a rotation speed of 250 rpm.</li>
	<li>Perform miniprep on the E. coli culture using a commercial miniprep kit following its instruction manual. Elute plasmid DNA into 30-50 &#181;L of TE buffer by 1-minute centrifuge at 14,000 rpm, and measure its DNA concentration with&#160;a spectrophotometer.</li>
	<li>Linearize 1-2 &#181;g of plasmid with Not I endonuclease and purify the DNA with a DNA cleaning kit following its instruction manual after Not I digestion. Elute the DNA into 5 &#181;L of water by centrifuging at 14,000 rpm for 1 minute. The expected concentration is about 200-300 ng/&#181;L.</li>
	<li>Perform in vitro transcription with Not I linearized pCS2FA-transposase as a DNA template using a commercial SP6 transcription kit.</li>
	<li>Once the reaction is finished, purify Tol2 transposase mRNA using a commercial RNA cleaning kit following the manufacturer&#39;s instructions. Finally, elute mRNA into 20 &#181;L RNAase-free water and measure the RNA concentration in a spectrophotometer. The expected concentration is about 1-3 &#181;g/&#181;L. Samples can be stored in a -80 &#176;C freezer if needed.</li>
	<li>Prepare the microinjection solution by mixing 20 ng/&#181;L pDestTol2-ubi-ASAP1 and Tol2 transposase mRNA (100 ng/&#181;L) in a microcentrifuge tube by pipetting. To prevent nucleic acid degradation caused by repeated thawing and refreezing, aliquot 6 &#181;L per tube and store it in a -80&#160;&#176;C freezer for future use.</li>
</ol>
3. Microinjection
<ol>
	<li>Set up 4-6 breeding tanks with at least 2 males and 2 females the afternoon before injection. These fish must not be fed in the afternoon. This step will reduce the amount of fish waste and save time to clean them out the next morning, while also helping to induce a breeding response.</li>
	<li>The following morning, remove the prepared injection solution (pDestTol2-ubi-ASAP1 construct and Tol2 mRNA) from the -80 &#176;C freezer, and place it on ice.</li>
	<li>Pull the dividers in the fish breeding tanks and allow the fish to mate. In general, fish lay eggs within 20-30 minutes after pulling out the divider. If not, wait 1-2 hours longer. Some fish may not lay eggs at all. In this case, repeat this experiment for fish embryo collection.</li>
	<li>While waiting, make sure there are needles prepared with the tip broken at an angle creating a beveled edge with forceps, or by breaking on a delicate task wiper.
	<ol>
		<li>Pull needles from capillary glass on a micropipette puller using the following parameters: heat 545; pull 60; velocity 80; time 250; pressure 500. Break the needle with forceps underneath a dissection scope (with eye-piece ruler for diameter estimation) by holding the forceps at an angle approximately 45&#176;. Desired needle diameters can be variable depending on the microinjector settings, but smaller diameter is preferred for decreasing embryo mortality.</li>
	</ol>
	</li>
	<li>Once the fish have laid eggs, collect them in a 10-cm diameter Petri dish and bring them to the dissection scope. Remove all abnormal embryos and fish waste.</li>
	<li>Pipet the fertilized embryos into the prepared 3% agarose injection mold. Remove excess water to help keep the embryos in place.</li>
	<li>Once all of the rows are filled with viable embryos, arrange them so that the single cells all face the same direction toward the needle, which is about a 45&#176; angle horizontally. This will make injection much easier later.</li>
	<li>While wearing gloves, use a 20 &#181;L loading pipet tip and remove 5 &#181;L of the prepared construct from the tube on ice.</li>
	<li>Carefully insert the tip into the back end of the broken capillary tube all the way to where it begins to tapper, as to get the reagent as close to the tip. If there are still air bubbles, shake the needle, making sure to not break the tip.</li>
	<li>Insert the needle straight into the microinjection needle holder and carefully tighten until the needle stays in place. Adjust the angle to about 45&#176;.</li>
	<li>Once the needle is prepared and attached, turn on the microscope and gas pressure tank. Commercial CO2 tanks are generally good for this purpose. The injection volume is adjusted by the holding and ejection pressure: approximate 0.5 psi for holding and 30 psi for ejection. Be sure to check that the solution comes out when pressing the pedal.</li>
	<li>Using a stage micrometer with a drop of mineral oil, adjust the volume and flow of the solution to ~150 &#181;m in diameter (about 2 nL). Ensure that the back pressure will let a small amount drip out of the needle. If there is not enough back pressure, capillary action will cause liquid to enter the needle and destroy the mRNA.</li>
	<li>Once the needle is calibrated, begin injecting the construct into the single cell of the fertilized embryos. 
	NOTE: This takes a large amount of practice, patience, and finesse, due to the cell membrane being hard to pierce. It is important to inject the solution into the cell, not the yolk, for generating transgenic zebrafish. This is different from morpholino injection. It does not matter which side the needle enters the cell as long as the construct goes in the cell. The transgenesis will have a very low rate of success if injected into the yolk instead of the cell. Single-cell stage injection is also important, or somatic chimera fish will be created. This will reduce the chance of the transgene going into to the germ cells.</li>
	<li>Use the edge of the gel notch to provide a backing that keeps the embryo in place and allows the needle to apply pressure without moving the embryo. Once the tip of the needle is in the single cell, press the pedal to release the desired amount of solution. Repeat this process for all of the embryos.</li>
	<li>When completed, transfer the injected embryos into a labeled dish by rinsing them out of the agarose notch with fish system water and a disposable 3.4 mL transfer pipette. Store the embryos in a 28.5 &#176;C incubator to let them develop. Check back throughout the day removing dead fish embryos and replace water with 0.1% methylene blue in fish water.</li>
	<li>Around 6-8 hours after injection, take 10 individual injected fish embryos and prepare genomic DNA from them using the Hotshot method18.</li>
	<li>The following morning, use a dissection microscope with a fluorescence light source to sort out the embryos showing GFP in the non-yolk tissues. These fish embryos should contain the injected construct.</li>
	<li>Perform Tol2 excise assay to check the transposon activity as described previously19. If excised plasmid can be detected, keep the injected fish embryos and raise them. If no excised plasmid can be detected, repeat the Tol2 mRNA synthesis and microinjection process until achieving the positive results of the Tol2 excise assay.</li>
</ol>
4. Establish Transgenic ASAP1 fish, Tg(ubi:ASAP1)
<ol>
	<li>Raise the injected fish (F0 generation) to adulthood as described previously in the zebrafish book20. This usually takes about 4 months.</li>
	<li>Take a single adult F0 fish and cross it with a single opposite gender wildtype fish. Collect fish embryos after breeding later in the day. Keep the collected fish embryos in the 28.5 &#176;C incubator in fish water with 0.1% methylene blue.</li>
	<li>On the third day, check the fish embryos underneath a fluorescent dissection microscope with a GFP filter. Sort out green fish embryos, if there are any, and raise them to adulthood as F1&#160;generation transgenic fish, Tg(ubi:ASAP1). 
	NOTE: Mendelian ratio is not expected since most of the parental F0 fish are germ-line genetic chimeras.</li>
	<li>Cross single F1 adult fish with wildtype fish and collect fish embryos. Sort out green fish embryos and raise them to adulthood as F2 generation fish. 
	NOTE: Green and non-green fish embryos should be close to 1:1 if there is a single transgene.</li>
	<li>To view electric potential changes in tumor like malignant peripheral nerve sheath tumors (MPNST), cross the F2 generation Tg(ubi:ASAP1) fish with rpl35hi258/wt fish. It is known that visible tumors start to be found in the 6-8 month old adults21,22.</li>
</ol>
5. Imaging
<ol>
	<li>To image zebrafish embryos, take multiple F2 generation founder fish and cross them with wildtype fish in individual pairs. Collect fish embryos at different desired developmental stages according to the zebrafish staging guide23.</li>
	<li>For the early stages of fish embryos, peel and remove chorions of the embryos carefully using a pair of forceps under a dissection scope in a 10-cm diameter Petri dish with fish system water.</li>
	<li>Transfer a few fish embryos onto a concaved glass slide with 3% methylcellulose using a 3.4 mL disposable transfer pipette. Adjust the embryos to the desired positions to view the cellular GFP activity using a needle underneath a fluorescent dissection scope.</li>
	<li>For the moving stages of fish embryos (older than 12 somite stage), use 0.05% tricaine mesylate to anesthetize the fish embryos before transferring them to slides. Briefly, fish embryos were emerged into 0.05% tricaine mesylate in fish water until they stopped swimming and loss body balance. Also, add a drop of 0.05% tricaine mesylate with the methylcellulose on the slide.</li>
	<li>For less than 12 somite stage fish embryos, use an epi-fluorescence compound microscope with a compatible camera and software for imaging. For older than 12 somite stage fish embryos, use a fluorescence dissection microscope.</li>
	<li>To image tumor cell voltage, first identify the fish with MPNST tumors. Then, anesthetize the fish with 0.05% tricaine mesylate. For whole mount imaging, put the fish into a 10-cm diameter Petri dish. To view the tumor cell electrical activity, fish tumors may be dissected out after whole mount imaging.</li>
</ol>

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The authors have nothing to disclose.

Although the cellular and tissue level electrical activities during embryonic development and human disease were discovered a long time ago, the in vivo dynamic electrical changes and their biological roles still remain largely unknown. One of the major challenges is to visualize and quantify the electrical changes. Patch clamp technology is a break-through for tracking single cells, but its application to vertebrate embryos is limited because they are composed of many cells. The current chemical voltage dyes ar...

Bioelectricity refers to endogenous electrical signaling mediated by ion channels and pumps located on the cell membrane1. Ionic exchanges across the cellular membrane, and the coupled electrical potential and current changes, are essential for signaling processes of excitable neuronal and muscular cells. In addition, bioelectricity and ion gradients have a variety of other important biological functions including energy storage, biosynthesis, and metabolite transportation. Bioelectrical signaling was also discovered as a regulator of embryonic pattern formation, such as body axes, the cell cycle, and cell differentiation1. Thus, it is critical for understanding many human congenital diseases that result from the mis-regulation of this type of signaling. Although patch clamp has been&#160;widely used for recording single cells, it is still far from ideal for the simultaneous monitoring of multiple cells during embryonic development in vivo. Furthermore, voltage sensitive small molecules are also not ideal for in vivo applications due to their specificities, sensitivities, and toxicities.
The creation of a variety of genetically encoded fluorescent voltage indicators (GEVIs) offers a new mechanism to overcome this issue, and allows for easy application to study embryonic development, even though they were originally intended for monitoring neural cells2,3. One of the currently available GEVIs is the Accelerated Sensor of Action Potentials 1 (ASAP1)4. It is composed of an extracellular loop of a voltage-sensing domain of voltage sensitive phosphatase and a circularly permuted green fluorescent protein. Therefore, ASAP1 allows visualization of cellular electric potential changes (polarization: bright green; depolarization: dark green). ASAP1 has 2 ms on-and-off kinetics, and can track subthreshold potential change4. Thus, this genetic tool allows for a new level of efficacy in real-time bioelectric monitoring in live cells. Further understanding of the roles of bioelectricity in embryonic development and many human diseases, such as cancer, will shed new light on the underlying mechanisms, which is critical for disease treatment and prevention.
Zebrafish have been proven a powerful animal model to study developmental biology and human diseases including cancer5,6. They share 70% orthologous genes with humans, and they have similar vertebrate biology7. Zebrafish provide relatively easy care, a large clutch size of eggs, tractable genetics, easy transgenesis, and transparent external embryonic development, which make them a superior system for in vivo imaging5,6. With a large source of mutant fish lines already present and a fully sequenced genome, zebrafish will provide a relatively&#160;unlimited range of scientific discovery.
To investigate the in vivo real-time electrical activity of cells, we take advantage of the zebrafish model system and ASAP1. In this paper, we describe how to incorporate the fluorescent voltage biosensor ASAP1 into the zebrafish genome using Tol2 transposon transgenesis, and visualize cellular electrical activity during embryonic development, fish larval movement, and in live tumor.

<table>
	<tbody>
		<tr>
			<td>14mL cell culture tubes</td>
			<td>VWR</td>
			<td>60818-725</td>
			<td>E.Coli culture</td>
		</tr>
		<tr>
			<td>Agarose electrophoresis tank</td>
			<td>Thermo Scientific</td>
			<td>Owl B2</td>
			<td>DNA eletrophoresis</td>
		</tr>
		<tr>
			<td>Agarose RA</td>
			<td>Amresco</td>
			<td>N605-500G</td>
			<td>For making the injection gels</td>
		</tr>
		<tr>
			<td>Attb1-ASAP1-F primer</td>
			<td>IDT DNA</td>
			<td>GGGGACAAGTTTGTACAAAAAAGCAGGCTTCACCATGGAGACGACTGTGAGGTATGAACA</td>
			<td>ASAP1 coding region amplification for subcloning</td>
		</tr>
		<tr>
			<td>Attb2-ASAP1-R primer</td>
			<td>IDT DNA</td>
			<td>GGGGACCACTTTGTACAAGAAAGCTGGGTCTTAGGTTACCACTTCAAGTTGTTTCTTCTGTGAAGCCA</td>
			<td>ASAP1 coding region amplification for subcloning</td>
		</tr>
		<tr>
			<td>Bright field dissection scope</td>
			<td>Nikon</td>
			<td>SMZ 745</td>
			<td>Dechorionation, microinjection, mounting</td>
		</tr>
		<tr>
			<td>Color camera</td>
			<td>Zeiss</td>
			<td>AxioCam MRc</td>
			<td>Fish embryo image recording</td>
		</tr>
		<tr>
			<td>Concave slide</td>
			<td>VWR</td>
			<td>48336-001</td>
			<td>For holding fish embryos during imaging process</td>
		</tr>
		<tr>
			<td>Disposable transfer pipette 3.4 ml</td>
			<td>Thermo Scientific</td>
			<td>13-711-9AM</td>
			<td>Fish embryos and water transfer</td>
		</tr>
		<tr>
			<td>Endonuclease enzyme, Not I</td>
			<td>NEB</td>
			<td>R0189L</td>
			<td>For linearizing plasmid DNA</td>
		</tr>
		<tr>
			<td>Epifuorescent compound scope</td>
			<td>Zeiss</td>
			<td>Axio Imager.A2</td>
			<td>Fish embryo imaging</td>
		</tr>
		<tr>
			<td>Epifuorescent stereo dissection scope</td>
			<td>Zeiss</td>
			<td>Stereo Discovery.V12</td>
			<td>Fish embryo imaging</td>
		</tr>
		<tr>
			<td>Fluorescent light source</td>
			<td>Lumen dynamics</td>
			<td>X-cite seris 120</td>
			<td>Light source for fluorescence microscopes</td>
		</tr>
		<tr>
			<td>Forceps #5</td>
			<td>WPI</td>
			<td>500342</td>
			<td>Dechorionation and needle breaking</td>
		</tr>
		<tr>
			<td>Gateway BP Clonase II Enzyme mix</td>
			<td>Thermo Scientific</td>
			<td>11789020</td>
			<td>Gateway BP recombination cloning</td>
		</tr>
		<tr>
			<td>Gateway LR Clonase II Plus enzyme</td>
			<td>Thermo Scientific</td>
			<td>12538120</td>
			<td>Gateway LR recombination cloning</td>
		</tr>
		<tr>
			<td>Gel DNA Recovery Kit</td>
			<td>Zymo Research</td>
			<td>D4002</td>
			<td>DNA gel purification</td>
		</tr>
		<tr>
			<td>Loading tip</td>
			<td>Eppendorf</td>
			<td>930001007</td>
			<td>For loading injection solution into capilary needles</td>
		</tr>
		<tr>
			<td>Methylcellulose (1600cPs)</td>
			<td>Alfa Aesar</td>
			<td>43146</td>
			<td>Fish embryo mounting</td>
		</tr>
		<tr>
			<td>Methylene blue</td>
			<td>Sigma-Aldrich</td>
			<td>M9140</td>
			<td>Suppresses fungal outbreaks in Petri dishes</td>
		</tr>
		<tr>
			<td>Microinjection mold</td>
			<td>Adaptive Science Tools</td>
			<td>TU-1</td>
			<td>To prepare agaorse mold tray for holding fish embryos during injection</td>
		</tr>
		<tr>
			<td>Microinjector</td>
			<td>WPI</td>
			<td>Pneumatic Picopump PV820</td>
			<td>Microinjection injector</td>
		</tr>
		<tr>
			<td>Micro-manipulator</td>
			<td>WPI</td>
			<td>Microinjector mm33 rechts</td>
			<td>Microinjection operation</td>
		</tr>
		<tr>
			<td>Micropipette puller</td>
			<td>Sutter instrument</td>
			<td>P-1000</td>
			<td>For preparing capillary needle</td>
		</tr>
		<tr>
			<td>Mineral oil</td>
			<td>Amresco</td>
			<td>J217-500ml</td>
			<td>For calibrating injection volume</td>
		</tr>
		<tr>
			<td>mMESSAGE mMACHINE SP6 Transcription Kit</td>
			<td>Thermo Scientific</td>
			<td>AM1340</td>
			<td>mRNA in vitro transcription</td>
		</tr>
		<tr>
			<td>Monocolor camera</td>
			<td>Zeiss</td>
			<td>AxioCam MRm</td>
			<td>Fish embryo image recording</td>
		</tr>
		<tr>
			<td>Plasmid Miniprep Kit</td>
			<td>Zymo Research</td>
			<td>D4020</td>
			<td>Prepare small amount of plasmid DNA</td>
		</tr>
		<tr>
			<td>Plastic Petri dishes</td>
			<td>VWR</td>
			<td>25384-088</td>
			<td>For holding fish or fish embryos during imaging process</td>
		</tr>
		<tr>
			<td>RNA Clean &#38; Concentrator-5</td>
			<td>Zymo Research</td>
			<td>R1015</td>
			<td>mRNA cleaning after in vitro transcription</td>
		</tr>
		<tr>
			<td>Spectrophotometer</td>
			<td>Thermo Scientific</td>
			<td>NanoDrop 2000</td>
			<td>For measuring DNA and RNA concentrations</td>
		</tr>
		<tr>
			<td>Stage Micrometer</td>
			<td>Am Scope</td>
			<td>MR100</td>
			<td>Microinjection volume calibration</td>
		</tr>
		<tr>
			<td>Thermocycler</td>
			<td>Bio-Rad</td>
			<td>T100</td>
			<td>DNA amplification for gene cloning</td>
		</tr>
		<tr>
			<td>Thin wall glass capillaries</td>
			<td>WPI</td>
			<td>TW100F-4</td>
			<td>Raw glass for making cappilary needle</td>
		</tr>
		<tr>
			<td>Tol2-exL1 primer</td>
			<td>IDT DNA</td>
			<td>GCACAACACCAGAAATGCCCTC</td>
			<td>Tol2 excise assay</td>
		</tr>
		<tr>
			<td>Tol2-exR primer</td>
			<td>IDT DNA</td>
			<td>ACCCTCACTAAAGGGAACAAAAG</td>
			<td>Tol2 excise assay</td>
		</tr>
		<tr>
			<td>TOP10 Chemically Competent E. coli</td>
			<td>Thermo Scientific</td>
			<td>C404006</td>
			<td>Used for transformation during gene cloning</td>
		</tr>
		<tr>
			<td>Tricaine mesylate</td>
			<td>Sigma-Aldrich</td>
			<td>A5040</td>
			<td>For anesthetizing fish or fish embryos</td>
		</tr>
		<tr>
			<td>UV trans-illuminator 302nm</td>
			<td>UVP</td>
			<td>M-20V</td>
			<td>DNA visualization</td>
		</tr>
		<tr>
			<td>Water bath</td>
			<td>Thermo Scientific</td>
			<td>2853</td>
			<td>For transformation process of gene cloning</td>
		</tr>
	</tbody>
</table>

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.

In a successful injection, more than 50% injected fish embryos will display some degree of green fluorescence&#160;in the somatic cells, and most of them will be positive by Tol2 transposon excise assay (Figure 2). After 2-4 generations of out-cross with wildtype fish (until the fluorescent fish reach 50%, the expected Mendelian ratio), the transgenic fish were used for the imaging experiment to track cell membrane potentials during embryonic development. Fir...

Watch this Scientific Journal Video about Visualization of Cellular Electrical Activity in Zebrafish Early Embryos and Tumors at JoVE.com

Visualization of Cellular Electrical Activity in Zebrafish Early Embryos and Tumors

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 ...

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Research

JoVE Journal

Developmental Biology

8.7K Views.  Purdue University. 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.

Video: Visualization of Cellular Electrical Activity in Zebrafish Early Embryos and Tumors

Measuring Protein Stability in Living Zebrafish Embryos Using Fluorescence Decay After Photoconversion (FDAP)

Protein stability influences many aspects of biology, and measuring the clearance kinetics of proteins can provide important insights into biological systems. In FDAP experiments, the clearance of proteins within living organisms can be measured. A protein of interest is tagged with a photoconvertible fluorescent protein, expressed in vivo and photoconverted, and the decrease in the photoconverted signal over time is monitored. The data is then fitted with an appropriate clearance model to determine the protein half-life. Importantly, the clearance kinetics of protein populations in different compartments of the organism can be examined separately by applying compartmental masks. This approach has been used to determine the intra- and extracellular half-lives of secreted signaling proteins during zebrafish development. Here, we describe a protocol for FDAP experiments in zebrafish embryos. It should be possible to use FDAP to determine the clearance kinetics of any taggable protein in any optically accessible organism.

Measuring Protein Stability in Living Zebrafish Embryos Using Fluorescence Decay After Photoconversion FDAP

Protein levels in cells and tissues are often tightly regulated by the balance of protein production and clearance. Using Fluorescence Decay After Photoconversion (FDAP), the clearance kinetics of proteins can be experimentally measured in vivo.

Protein stability influences many aspects of biology, and measuring the clearance kinetics of proteins can provide important insights into biological ...

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 ...

Transcriptome Profiling of In-Vivo Produced Bovine Pre-implantation Embryos Using Two-color Microarray Platform

Early embryonic loss is a large contributor to infertility in cattle. Moreover, bovine becomes an interesting model to study human preimplantation embryo development due to their similar developmental process. Although genetic factors are known to affect early embryonic development, the discovery of such factors has been a serious challenge. Microarray technology allows quantitative measurement and gene expression profiling of transcript levels on a genome-wide basis. One of the main decisions that have to be made when planning a microarray experiment is whether to use a one- or two-color approach. Two-color design increases technical replication, minimizes variability, improves sensitivity and accuracy as well as allows having loop designs, defining the common reference samples. Although microarray is a powerful biological tool, there are potential pitfalls that can attenuate its power. Hence, in this technical paper we demonstrate an optimized protocol for RNA extraction, amplification, labeling, hybridization of the labeled amplified RNA to the array, array scanning and data analysis using the two-color analysis strategy.

Transcriptome Profiling of In-Vivo Produced Bovine Pre-implantation Embryos Using Two-color Microarray Platform

Microarray technology allows quantitative measurement and gene expression profiling of transcripts on a genome-wide basis. Therefore, this protocol provides an optimized technical procedure in a two-color custom made bovine array using Day 7 bovine embryos to demonstrate the feasibility of using low amount of total RNA.

Early embryonic loss is a large contributor to infertility in cattle. Moreover, bovine becomes an interesting model to study human preimplantation embryo ...

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 ...

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 ...

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 ...

Multi-Photon Time Lapse Imaging to Visualize Development in Real-time: Visualization of Migrating Neural Crest Cells in Zebrafish Embryos

Congenital eye and craniofacial anomalies reflect disruptions in the neural crest, a transient population of migratory stem cells that give rise to numerous cell types throughout the body. Understanding the biology of the neural crest has been limited, reflecting a lack of genetically tractable models that can be studied in vivo and in real-time. Zebrafish is a particularly important developmental model for studying migratory cell populations, such as the neural crest. To examine neural crest migration into the developing eye, a combination of the advanced optical techniques of laser scanning microscopy with long wavelength multi-photon fluorescence excitation was implemented to capture high-resolution, three-dimensional, real-time videos of the developing eye in transgenic zebrafish embryos, namely Tg(sox10:EGFP) and Tg(foxd3:GFP), as sox10 and foxd3 have been shown in numerous animal models to regulate early neural crest differentiation and likely represent markers for neural crest cells. Multi-photon time-lapse imaging was used to discern the behavior and migratory patterns of two neural crest cell populations contributing to early eye development. This protocol provides information for generating time-lapse videos during zebrafish neural crest migration, as an example, and can be further applied to visualize the early development of many structures in the zebrafish and other model organisms.

A combination of the advanced optical techniques of laser scanning microscopy with long wavelength multi-photon fluorescence excitation was implemented to capture high-resolution, three-dimensional, real-time imaging of neural crest migration in Tg(sox10:EGFP) and Tg(foxd3:GFP) zebrafish embryos.

Congenital eye and craniofacial anomalies reflect disruptions in the neural crest, a transient population of migratory stem cells that give rise to ...

Visualizing the Node and Notochordal Plate In Gastrulating Mouse Embryos Using Scanning Electron Microscopy and Whole Mount Immunofluorescence

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is the formation of the transient organizers, the node and notochordal plate, from cells that have passed through the primitive streak. The proper formation of these signaling centers is essential for the development of the body plan and techniques to visualize them are of high interest to mouse developmental biologists. The node and notochordal plate lie on the ventral surface of gastrulating mouse embryos around embryonic day (E) 7.5 of development. The node is a cup-shaped structure whose cells possess a single slender cilium each. The proper subcellular localization and rotation of the cilia in the node pit determines left-right asymmetry. The notochordal plate cells also possess single cilia albeit shorter than those of the node cells. The notochordal plate forms the notochord which acts as an important signaling organizer for somitogenesis and neural patterning. Because the cells of the node and notochordal plate are transiently present on the surface and possess cilia, they can be visualized using scanning electron microscopy (SEM). Among other techniques used to visualize these structures at the cellular level is whole mount immunofluorescence (WMIF) using the antibodies against the proteins that are highly expressed in the node and notochordal plate. In this report, we describe our optimized protocols to perform SEM and WMIF of the node and notochordal plate in developing mouse embryos to help in the assessment of tissue shape and cellular organization in wild-type and gastrulation mutant embryos.

The node and notochordal plate are transient signaling organizers in developing mouse embryos that can be visualized using several techniques. Here, we describe in detail how to perform two of the techniques to study their structure and morphogenesis: 1) scanning electron microscopy (SEM); and 2) whole mount immunofluorescence (WMIF).

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is ...

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. ...

Preparation of Small RNA Libraries for Sequencing from Early Mouse Embryos

MicroRNAs (miRNAs) are important for the complex regulation of cell fate decisions and developmental timing. In vivo studies of the contribution of miRNAs during early development are technically challenging due to the limiting cell number. Moreover, many approaches require a miRNA of interest to be defined in assays such as northern blotting, microarray, and qPCR. Therefore, the expression of many miRNAs and their isoforms have not been studied during early development. Here, we demonstrate a protocol for small RNA sequencing of sorted cells from early mouse embryos to enable relatively unbiased profiling of miRNAs in early populations of neural crest cells. We overcome the challenges of low cell input and size selection during library preparation using amplification and gel-based purification. We identify embryonic age as a variable accounting for variation between replicates and stage-matched mouse embryos must be used to accurately profile miRNAs in biological replicates. Our results suggest that this method can be broadly applied to profile the expression of miRNAs from other lineages of cells. In summary, this protocol can be used to study how miRNAs regulate developmental programs in different cell lineages of the early mouse embryo.

We describe a technique for profiling microRNAs in early mouse embryos. This protocol overcomes the challenge of low cell input and small RNA enrichment. This assay can be used to analyze changes in miRNA expression over time in different cell lineages of the early mouse embryo.

MicroRNAs (miRNAs) are important for the complex regulation of cell fate decisions and developmental timing. In vivo studies of the contribution of miRNAs ...