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>GGGGACAAGTTTGTACAAAAAA&lt;br /&gt;GCAGGCTTCACCATGGAGACGA&lt;br /&gt;CTGTGAGGTATGAACA</td><td>ASAP1 coding region amplification for subcloning</td></tr><tr><td>Attb2-ASAP1-R primer</td><td>IDT DNA</td><td>GGGGACCACTTTGTACAAGAAA&lt;br /&gt;GCTGGGTCTTAGGTTACCACTTC&lt;br /&gt;AAGTTGTTTCTTCTGTGAAGCCA</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 MM3301R</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 &amp;amp; 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 ...

visualization-cellular-electrical-activity-zebrafish-early-embryos

Universidad de Cartagena UDEC

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

Time-lapse Live Imaging of Clonally Related Neural Progenitor Cells in the Developing Zebrafish Forebrain

Precise patterns of division, migration and differentiation of neural progenitor cells are crucial for proper brain development and function1,2. To understand the behavior of neural progenitor cells in the complex in vivo environment, time-lapse live imaging of neural progenitor cells in an intact brain is critically required. In this video, we exploit the unique features of zebrafish embryos to visualize the development of forebrain neural progenitor cells in vivo. We use electroporation to genetically and sparsely label individual neural progenitor cells. Briefly, DNA constructs coding for fluorescent markers were injected into the forebrain ventricle of 22 hours post fertilization (hpf) zebrafish embryos and electric pulses were delivered immediately. Six hours later, the electroporated zebrafish embryos were mounted with low melting point agarose in glass bottom culture dishes. Fluorescently labeled neural progenitor cells were then imaged for 36hours with fixed intervals under a confocal microscope using water dipping objective lens. The present method provides a way to gain insights into the in vivo development of forebrain neural progenitor cells and can be applied to other parts of the central nervous system of the zebrafish embryo.

The present video demonstrates a method which takes advantage of the combination of electroporation and confocal microscopy to perform live imaging on individual neural progenitor cells in the developing zebrafish forebrain. In vivo analysis of the development of forebrain neural progenitor cells at a clonal level can be achieved in this way.

Precise patterns of division, migration and differentiation of neural progenitor cells are crucial for proper brain development and function1,2. To ...

Neuroscience

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 ...

Live Imaging of the Zebrafish Embryonic Brain by Confocal Microscopy

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the developing zebrafish brain (Gutzman et al, 2008). Such analysis affords a new understanding of the underlying cell biology required for development of the 3D structure of the vertebrate brain, and significantly increases our ability to study neural tube morphogenesis. The embryonic zebrafish brain is shaped beginning at 18 hours post fertilization (hpf) as the ventricles within the neuroepithelium inflate. By 24 hpf, the initial steps of neural tube morphogenesis are complete. Using the method described here, embryos at the one cell stage are injected with mRNA encoding membrane-targeted green fluorescent protein (memGFP). After injection and incubation, the embryo, now between 18 and 24 hpf, is mounted, inverted, in agarose and imaged by confocal microscopy. Notably, the zebrafish embryo is transparent making it an ideal system for fluorescent imaging. While our analyses have focused on the midbrain-hindbrain boundary and the hindbrain, this method could be extended for analysis of any region in the zebrafish to a depth of 
80-100 &mu;m.

In this video, we demonstrate a method by which to analyze the developing vertebrate brain in live zebrafish embryos at single cell resolution by confocal microscopy. This includes the method by which we inject the single-cell zebrafish embryo and subsequently mount and image the developing brain.

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the ...

Biology

Forebrain Electrophysiological Recording in Larval Zebrafish

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

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

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

Live Imaging of Cell Motility and Actin Cytoskeleton of Individual Neurons and Neural Crest Cells in Zebrafish Embryos

The zebrafish is an ideal model for imaging cell behaviors during development in vivo. Zebrafish embryos are externally fertilized and thus easily accessible at all stages of development. Moreover, their optical clarity allows high resolution imaging of cell and molecular dynamics in the natural environment of the intact embryo. We are using a live imaging approach to analyze cell behaviors during neural crest cell migration and the outgrowth and guidance of neuronal axons.
Live imaging is particularly useful for understanding mechanisms that regulate cell motility processes. To visualize details of cell motility, such as protrusive activity and molecular dynamics, it is advantageous to label individual cells. In zebrafish, plasmid DNA injection yields a transient mosaic expression pattern and offers distinct benefits over other cell labeling methods. For example, transgenic lines often label entire cell populations and thus may obscure visualization of the fine protrusions (or changes in molecular distribution) in a single cell. In addition, injection of DNA at the one-cell stage is less invasive and more precise than dye injections at later stages.
Here we describe a method for labeling individual developing neurons or neural crest cells and imaging their behavior in vivo. We inject plasmid DNA into 1-cell stage embryos, which results in mosaic transgene expression. The vectors contain cell-specific promoters that drive expression of a gene of interest in a subset of sensory neurons or neural crest cells. We provide examples of cells labeled with membrane targeted GFP or with a biosensor probe that allows visualization of F-actin in living cells1.
Erica Andersen, Namrata Asuri, and Matthew Clay contributed equally to this work.

This protocol describes imaging of individual neurons or neural crest cells in living zebrafish embryos. This method is used to examine cellular behaviors and actin localization using fluorescence confocal time-lapse microscopy.

The zebrafish is an ideal model for imaging cell behaviors during development in vivo. Zebrafish embryos are externally fertilized and thus easily ...

Visualizing the Developing Brain in Living Zebrafish using Brainbow and Time-lapse Confocal Imaging

Development of the vertebrate nervous system requires a precise coordination of complex cellular behaviors and interactions. The use of high resolution in vivo imaging techniques can provide a clear window into these processes in the living organism. For example, dividing cells and their progeny can be followed in real time as the nervous system forms. In recent years, technical advances in multicolor techniques have expanded the types of questions that can be investigated. The multicolor Brainbow approach can be used to not only distinguish among like cells, but also to color-code multiple different clones of related cells that each derive from one progenitor cell. This allows for a multiplex lineage analysis of many different clones and their behaviors simultaneously during development. Here we describe a technique for using time-lapse confocal microscopy to visualize large numbers of multicolor Brainbow-labeled cells over real time within the developing zebrafish nervous system. This is particularly useful for following cellular interactions among like cells, which are difficult to label differentially using traditional promoter-driven colors. Our approach can be used for tracking lineage relationships among multiple different clones simultaneously. The large datasets generated using this technique provide rich information that can be compared quantitatively across genetic or pharmacological manipulations. Ultimately the results generated can help to answer systematic questions about how the nervous system develops.

In vivo imaging is a powerful tool that can be used to investigate the cellular mechanisms underlying nervous system development. Here we describe a technique for using time-lapse confocal microscopy to visualize large numbers of multicolor Brainbow-labeled cells in real time within the developing zebrafish nervous system.

Development of the vertebrate nervous system requires a precise coordination of complex cellular behaviors and interactions. The use of high resolution in ...

Multicolor Time-lapse Imaging of Transgenic Zebrafish: Visualizing Retinal Stem Cells Activated by Targeted Neuronal Cell Ablation

High-resolution time-lapse imaging of living zebrafish larvae can be utilized to visualize how biological processes unfold (for review see 1). Compound transgenic fish which express different fluorescent reporters in neighboring cell types provide a means of following cellular interactions 2 and/or tissue-level responses to experimental manipulations over time. In this video, we demonstrate methods that can be used for imaging multiple transgenically labeled cell types serially in individual fish over time courses that can span from minutes to several days. The techniques described are applicable to any study seeking to correlate the "behavior" of neighboring cells types over time, including: 1) serial 'catch and release' methods for imaging a large number of fish over successive days, 2) simplified approaches for separating fluorophores with overlapping excitation/emission profiles (e.g., GFP and YFP), 3) use of hypopigmented mutant lines to extend the time window available for high-resolution imaging into late larval stages of development, 4) use of membrane targeted fluorescent reporters to reveal fine morphological detail of individual cells as well as cellular details in larger populations of cells, and 5) a previously described method for chemically-induced ablation of transgenically targeted cell types; i.e., nitroreductase (NTR) mediated conversion of prodrug substrates, such as metronidazole (MTZ), to cytotoxic derivatives 3,5.
As an example of these approaches, we will visualize the ablation and regeneration of a subtype of retinal bipolar neuron within individual fish over several days. Simultaneously we will monitor several other retinal cell types, including neighboring non-targeted bipolar cells and potential degeneration-stimulated retinal stem cells (i.e., M&#971;ller glia). This strategy is being applied in our lab to characterize cell- and tissue-level (e.g., stem cell niche) responses to the selective loss and regeneration of targeted neuronal cell types.

In this video, techniques for multicolor confocal time-lapse imaging and targeted cell ablation are provided. Time-lapse imaging is used to monitor the behavior of multiple cell types of interest in vivo. Targeted cell ablation facilitates the study neural circuit function and cell-specific neuronal regeneration paradigms.

High-resolution time-lapse imaging of living zebrafish larvae can be utilized to visualize how biological processes unfold (for review see 1). Compound ...

Imaging and 3D Reconstruction of Cerebrovascular Structures in Embryonic Zebrafish

Zebrafish are a powerful tool to study developmental biology and pathology in vivo. The small size and relative transparency of zebrafish embryos make them particularly useful for the visual examination of processes such as heart and vascular development. In several recent studies transgenic zebrafish that express EGFP in vascular endothelial cells were used to image and analyze complex vascular networks in the brain and retina, using confocal microscopy. Descriptions are provided to prepare, treat and image zebrafish embryos that express enhanced green fluorescent protein (EGFP), and then generate comprehensive 3D renderings of the cerebrovascular system. Protocols include the treatment of embryos, confocal imaging, and fixation protocols that preserve EGFP fluorescence. Further, useful tips on obtaining high-quality images of cerebrovascular structures, such as removal the eye without damaging nearby neural tissue are provided. Potential pitfalls with confocal imaging are discussed, along with the steps necessary to generate 3D reconstructions from confocal image stacks using freely available open source software.

Imaging of cerebrovascular development in larval zebrafish is described. Techniques to facilitate 3D imaging and modify cerebrovascular development using chemical treatments are also provided.

Zebrafish are a powerful tool to study developmental biology and pathology in vivo.  The small size and relative transparency of zebrafish embryos make ...

Semi-automated Imaging of Tissue-specific Fluorescence in Zebrafish Embryos

Zebrafish embryos are a powerful tool for large-scale screening of small molecules. Transgenic zebrafish that express fluorescent reporter proteins are frequently used to identify chemicals that modulate gene expression. Chemical screens that assay fluorescence in live zebrafish often rely on expensive, specialized equipment for high content screening. We describe a procedure using a standard epifluorescence microscope with a motorized stage to automatically image zebrafish embryos and detect tissue-specific fluorescence. Using transgenic zebrafish that report estrogen receptor activity via expression of GFP, we developed a semi-automated procedure to screen for estrogen receptor ligands that activate the reporter in a tissue-specific manner. In this video we describe procedures for arraying zebrafish embryos at 24-48 hours post fertilization (hpf) in a 96-well plate and adding small molecules that bind estrogen receptors. At 72-96 hpf, images of each well from the entire plate are automatically collected and manually inspected for tissue-specific fluorescence. This protocol demonstrates the ability to detect estrogens that activate receptors in heart valves but not in liver.

Described here is a protocol for semi-automated imaging of tissue-specific fluorescence in zebrafish embryos.

Zebrafish embryos are a powerful tool for large-scale screening of small molecules. Transgenic zebrafish that express fluorescent reporter proteins are ...

In vivo Imaging of Fully Active Brain Tissue in Awake Zebrafish Larvae and Juveniles by Skull and Skin Removal

Understanding the ephemeral changes that occur during brain development and maturation requires detailed high-resolution imaging in space and time at cellular and subcellular resolution. Advances in molecular and imaging technologies have allowed us to gain numerous detailed insights into cellular and molecular mechanisms of brain development in the transparent zebrafish embryo. Recently, processes of refinement of neuronal connectivity that occur at later larval stages several weeks after fertilization, which are for example control of social behavior, decision making or motivation-driven behavior, have moved into focus of research. At these stages, pigmentation of the zebrafish skin interferes with light penetration into brain tissue, and solutions for embryonic stages, e.g., pharmacological inhibition of pigmentation, are not feasible anymore.
Therefore, a minimally invasive surgical solution for microscopy access to the brain of awake zebrafish is provided that is derived from electrophysiological&#160;approaches. In teleosts, skin and soft skull cartilage can be carefully removed by micro-peeling these layers, exposing underlying neurons and axonal tracts without damage. This allows for recording neuronal morphology, including synaptic structures and their molecular contents, and the observation of physiological changes such as Ca2+ transients or intracellular transport events. In addition, interrogation of these processes by means of pharmacological inhibition or optogenetic manipulation is feasible. This brain exposure approach provides information about structural and physiological changes in neurons as well as the correlation and interdependence of these events in live brain tissue in the range of minutes or hours. The technique is suitable for in vivo brain imaging of zebrafish larvae up to 30 days post fertilization, the latest developmental stage tested so far. It, thus, provides access to such important questions as synaptic refinement and scaling, axonal and dendritic transport, synaptic targeting of cytoskeletal cargo or local activity-dependent expression. Therefore, a broad use for this mounting and imaging approach can be anticipated.

Here we present a method to image the zebrafish embryonic brain in vivo upto larval and juvenile stages. This microinvasive procedure, adapted from electrophysiological approaches, provides access to cellular and subcellular details of mature neuron and can be combined with optogenetics and neuropharmacological studies for characterizing brain function and drug intervention.

Understanding the ephemeral changes that occur during brain development and maturation requires detailed high-resolution imaging in space and time at ...