We thank Cecilia Moens for training in zebrafish transplantation; Marc Tye for excellent fish care; and Emma Carlson for feedback on the manuscript. This work was supported by NIH grant NS121595 to A.J.I.

All aspects of this procedure that pertain to work with live zebrafish have been approved by the University of Minnesota Institutional Animal Care and Use Committee (IACUC) and are performed in compliance with IACUC guidelines.
1. One-time initial setup of transplant apparatus (Figure 1)
<ol>
	<li>Assemble the transplant microscope per manufacturer&#39;s instructions. 
	NOTE: This protocol uses an upright fluorescence microscope with a 40x water dipping objective.</li>
	<li>Assemble the coarse and fine micromanipulators and mount them on the right side of the microscope according to the manufacturer&#39;s instructions. 
	NOTE: Because it is important that the stage not move in the Z dimension relative to the needle, we use a fixed stage microscope with the micromanipulators mounted to the microscope base. If a fixed stage microscope is not available, the micromanipulator can be mounted to the stage.</li>
	<li>Attach the microelectrode holder to the electrode handle, and mount on the micromanipulator.</li>
	<li>Connect one side of the three-way stopcock to the microsyringe pump. Connect the other side of the stopcock to the polyethylene tubing using the chromatography adapter.</li>
	<li>Connect the opposite side of the polyethylene tubing to the microelectrode holder using the chromatography adapter.</li>
	<li>Fill the 10 mL reservoir syringe with light mineral oil and mount it on the top side of the stopcock.</li>
	<li>Fill the microsyringe pump and tubing (the hydraulic line) with mineral oil from the reservoir, making sure to remove all air bubbles. 
	NOTE: Once the apparatus is set up, it will require only occasional maintenance of the hydraulic line. The reservoir syringe can be removed and refilled with mineral oil as necessary.</li>
</ol>
2. Prepare embryo pushers.
<ol>
	<li>Cut a 2 cm piece of fishing line and insert it partially into the narrow end of a P1000 pipette tip, leaving ~1.5 cm exposed. Secure the fishing line with a small drop of superglue and let dry.</li>
</ol>
3. Prepare solutions.
<ol>
	<li>To prepare 1% low melting point agarose dissolved in embryo media (LMA), dissolve agarose in boiling embryo media34. Make 1-2 mL aliquots in round bottom test tubes and store at 4 &#176;C.</li>
	<li>Prepare Ringer&#39;s solution with penicillin-streptomycin and tricaine (RPT) by combining the following ingredients to final concentrations of 116 mM NaCl, 2.9 mM KCl, 1.8 mM CaCl2, 5 mM HEPES pH 7.2, 50 units/mL penicillin, and 50ug/mL streptomycin, stirring until dissolved, and passing through a vacuum filter. Store at room temperature. Add tricaine immediately before use to a concentration of 0.02%.</li>
</ol>
4. Prepare donor and host animals for transplantation.
<ol>
	<li>Raise host and donor animals of the appropriate genotypes, and with cells of interest fluorescently labeled, to the desired age. 
	NOTE: For these transplants, we used Tg(isl1:EGFPCAAX)fh474 donor animals32 and Tg(isl1:mRFP)fh1 host animals35 and transplanted neurons from donor to host vagus motor nuclei at 3dpf.</li>
	<li>Prepare transplant slides by applying a rectangular outline of clear nail polish to each glass slide (Figure 2A). The nail polish creates a hydrophobic barrier to hold in the RPT added in step 4.8. Allow to dry fully before using. 
	NOTE: Slides can be prepared ahead of time and stored indefinitely.</li>
	<li>Melt one LMA aliquot by placing a tube of LMA in a 50mL beaker containing 40mL of water and microwaving for 1 min at 50% power or until melted. Maintain LMA at 40 &#176;C in a dry bath heater.</li>
	<li>Dechorionate host and donor animals with forceps (if applicable) and anesthetize them by placing them in small Petri dishes filled with room-temperature RPT. Wait 5 min for animals to be fully anesthetized before proceeding.</li>
	<li>Mount individual donor and host animals: Using Pasteur pipette and pipette pump, transfer animals from RPT to LMA, then transfer animals in small drops of LMA onto slides within the nail polish outline. Minimize the amount of RPT transferred into the LMA so that the LMA does not become diluted. Before the LMA solidifies, use an embryo pusher to orient the animals, ensuring that each animal is mounted with the intended needle insertion site to the right, and all animals are aligned vertically (Figure 2B); allow the agarose to fully set (~5 min) before proceeding. 
	NOTE: Because many cells can be taken from each donor animal, it is more efficient to mount 2-4 host animals with one donor on each slide.</li>
	<li>Using a scalpel, cut a straight vertical slice through all agarose drops just to the right of the mounted animals (Figure 2C). Remove loose agarose from slide with laboratory wipes.</li>
	<li>Using a scalpel, cut wedges out of the agarose to expose the needle insertion site (Figure 2C). Remove loose agarose from slide with laboratory wipes.</li>
	<li>Apply RPT to the slide until the agarose drops are fully submerged (Figure 2D).</li>
	<li>Mount the prepared slide on the transplantation microscope. Bring the donor animal into focus under the 40x objective. To do this, lower the objective until it breaks the surface of the media, then raise the objective to the desired focal plane. 
	NOTE: Liquid contact by the objective must be maintained until transplantation is complete (Figure 3C).</li>
	<li>Bring the fluorescently labeled donor cell(s) of interest into focus; then, move the stage to the left in preparation to locate the transplant needle. 
	NOTE: At this point, do not adjust microscope on the Y- or Z-axes, to ensure that you can easily re-find the donor animal in section 5.</li>
</ol>
5. Prepare the transplant needle.
<ol>
	<li>Insert a micropipette into the micropipette puller and pull a microinjection needle. 
	NOTE: The needle&#39;s shape should be similar to those used for 1-cell-stage zebrafish embryo injections or patch clamping. We use a micropipette puller with the following program: PRESSURE = 500, HEAT = ramp + 80, PULL = 90, VELOCITY = 70, TIME = 250 (see Table of Materials), although correct settings for each puller will likely need to be empirically determined.</li>
	<li>Align the needle tip with the divisions of the stage micrometer under a dissecting scope. Using the micrometer as a measurement guide, use a sharpened pair of forceps to break the needle to a bore size that is slightly larger than the diameter of the cells of interest (Figure 3A). Make sure that the break is as clean as possible, as jagged edges can contribute to needle clogging. NOTE: For vagus motor neurons (5-7 &#181;m diameter), needles should be broken to an inner diameter of 10 &#181;m, which corresponds to an outer diameter of 20 &#181;m. If needed, the size, shape, and/or angle of the needle tip can be refined with a microforge or microgrinder36.</li>
	<li>Using a 50 mL syringe filled with light mineral oil and equipped with a pipette filler, completely fill the needle with mineral oil (Figure 3B). Ensure that there are no air bubbles. Set the filled needle aside until ready for mounting. 
	NOTE: To ensure there are no air bubbles, insert the pipette filler all the way into the needle and release oil while slowly pulling the filler out of the needle.</li>
	<li>On the transplant apparatus, turn the three-way stopcock so that its long arm faces the microsyringe pump and depress the reservoir syringe plunger to flush all air bubbles from the hydraulic line. Maintain light pressure on the plunger (to prevent backflow of air into line) and turn the three-way stopcock so that its long arm faces the reservoir syringe.</li>
	<li>Insert the needle into the holder, taking care not to introduce any air bubbles. Adjust the angle of the needle so that it faces directly to the left at an angle of 10-15&#176; from horizontal (Figure 3C).</li>
	<li>Use the coarse and fine micromanipulators to maneuver the tip of the needle under the microscope objective and bring it into focus (Figure 3D). 
	NOTE: Coarse positioning of the needle in the X and Y planes can be done by directly observing the area beneath the objective while you maneuver the needle, as the needle will reflect the transmitted light beam when it is positioned under the objective. The microscope eyepieces can then be used for fine positioning. Do not adjust the microscope stage position or focus during this step. Use the micromanipulators to bring the needle tip into the existing focal position.</li>
	<li>If liquid is flowing into or out of the needle tip, adjust the pressure using the syringe pump until a stable meniscus between the mineral oil and Ringer&#39;s solution is observed (Figure 3D). 
	NOTE: If an oil bubble remains at the needle tip after stabilization, it can be removed by using the X plane adjustment on the coarse micromanipulator to move the needle to the right until its tip has been withdrawn from the RPT, then back to the left until it is repositioned under the objective.</li>
</ol>
6. Transplantation
NOTE: All needle movements should be done using the fine micromanipulator for this section.
<ol>
	<li>Move the stage to the right until the donor animal is brought back into view, being cautious to avoid accidental penetration with the needle.</li>
	<li>Re-center and focus on the cells to be transplanted and align the needle with the cells just outside the animal (Figure 4A). 
	NOTE: You may need to switch between brightfield and fluorescence often to ensure proper alignment. The use of a neutral density filter can help in visualizing both simultaneously.</li>
	<li>Insert the needle into the donor animal. 
	NOTE: Repeated in-and-out motions with the needle can aid in penetrating the skin.</li>
	<li>Immediately re-stabilize the oil meniscus in the needle tip with the microsyringe pump, as described in step 5.7.</li>
	<li>Position the needle tip against the cells of interest and apply gentle suction with the microsyringe pump (Figure 4B). 
	NOTE: Gentle in-and-out movements during suction can help loosen cells.</li>
	<li>When an adequate number of cells have been taken up, remove the needle from the donor and immediately re-stabilize the oil meniscus in the needle tip with the microsyringe pump.</li>
	<li>Being careful to avoid contacting animals or agarose with the needle, reposition the stage to bring the first host animal into view.</li>
	<li>Center and focus on the area in which the cells are to be placed and align the needle with this region just outside the animal (Figure 4C).</li>
	<li>Insert the needle into the host animal and immediately re-stabilize the oil meniscus in the needle tip with the microsyringe pump.</li>
	<li>Position the needle tip in the deposition site and apply gentle pressure with the microsyringe pump until the correct number of donor cells are released from the needle (Figure 4D). 
	NOTE: If donor and host cells are labeled with different fluorophores, a multi-band filter to allow simultaneous visualization can be very helpful at this stage.</li>
	<li>Remove the needle from the host and immediately re-stabilize the oil meniscus in the needle tip with the microsyringe pump.</li>
	<li>Repeat steps 6.7-6.11 for all remaining hosts.</li>
</ol>
7. Host animal recovery
<ol>
	<li>Raise the 40x objective and use the coarse micromanipulator to maneuver the needle back to the loading position. 
	NOTE: The needle may be reused for multiple slides.</li>
	<li>Remove the slide from the transplant microscope and place it under a dissecting microscope.</li>
	<li>Unmount the host animals by carefully removing them from the LMA drop with forceps. Using a glass Pasteur pipette, transfer the host animals into a dish with fresh Ringer&#39;s solution with penicillin/streptomycin. Ensure that the animals recover from anesthesia and maintain them in an embryo incubator until ready to image. Euthanize the donor animals according to approved protocols. 
	NOTE: Our method for euthanasia is to immerse animals in an ice bath for at least 1 hour followed by immersion in 500 mg/L sodium hypochlorite.</li>
</ol>

Robust Zebrafish Cell Transplantation for Regeneration and Development Research

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I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+use+of+zebrafish+to+understand+immunity.">The use of zebrafish to understand immunity.</a> Immunity. 20 (4), 367-379 (2004).</li><li>Lam, S. H., Chua, H. L., Gong, Z., Lam, T. J., Sin, Y. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+and+maturation+of+the+immune+system+in+zebrafish,+Danio+rerio:+a+gene+expression+profiling,+in+situ+hybridization+and+immunological+study.">Development and maturation of the immune system in zebrafish, Danio rerio: a gene expression profiling, in situ hybridization and immunological study.</a> Dev Comp Immunol. 28 (1), 9-28 (2004).</li><li>Wienholds, E., Schulte-Merker, S., Walderich, B., Plasterk, R. H. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Target-selected+inactivation+of+the+zebrafish+rag1+gene.">Target-selected inactivation of the zebrafish rag1 gene.</a> Science. 297 (5578), 99-102 (2002).</li><li>Petrie-Hanson, L., Hohn, C., Hanson, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+of+rag1+mutant+zebrafish+leukocytes.">Characterization of rag1 mutant zebrafish leukocytes.</a> BMC Immunol. 10, 8 (2009).</li><li>Roh-Johnson, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Macrophage-dependent+cytoplasmic+transfer+during+melanoma+invasion+in+vivo.">Macrophage-dependent cytoplasmic transfer during melanoma invasion in vivo.</a> Dev Cell. 43 (5), 549-562 (2017).</li><li>Bukrinsky, A., Griffin, K. J. P., Zhao, Y., Lin, S., Banerjee, U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Essential+role+of+spi-1-like+(spi-1l)+in+zebrafish+myeloid+cell+differentiation.">Essential role of spi-1-like (spi-1l) in zebrafish myeloid cell differentiation.</a> Blood. 113 (9), 2038-2046 (2009).</li></ol>

The authors have no conflicts of interest to disclose.

Developmental and regenerative biology has for over a century relied on transplantation experiments to examine principles of cell signaling and cell fate determination. The zebrafish model already represents a powerful fusion of genetic and transplantation approaches. Transplantation at blastula and gastrula stages to generate mosaic animals is common but limited in what types of questions it can address. Later-stage transplantation is rare, although methods to transplant embryonic spinal motor neurons and retinal gangli...

Cell transplantation has a long and storied history as a foundational technique in developmental biology. Around the turn of the 20th century, approaches using physical manipulations to perturb the developmental process, including transplantation, transformed embryology from an observational science into an experimental one1,2. In one landmark experiment, Hans Spemann and Hilde Mangold ectopically transplanted the dorsal blastopore lip of a salamander embryo onto the opposite side of a host embryo, inducing the nearby tissue to form a secondary body axis3. This experiment showed that cells could induce other cells to adopt certain fates, and subsequently, transplantation developed as a powerful method for interrogating critical questions in developmental biology regarding competence and cell fate determination, cell lineage, inductive ability, plasticity, and stem cell potency1,4,5.
More recent scientific advances have expanded the power of the transplantation approach. In 1969, Nicole Le Douarin&#39;s discovery that nucleolar staining could distinguish species of origin in quail-chick chimeras allowed for the tracking of transplanted cells and their progeny6. This concept was later supercharged by the advent of transgenic fluorescent markers and advanced imaging techniques5, and has been leveraged to track cell fate6,7, identify stem cells and their potency8,9, and track cell movements during brain development10. Additionally, the rise of molecular genetics facilitated transplantation between hosts and donors of distinct genotypes, supporting precise dissection of autonomous and non-autonomous functions of developmental factors11.
Transplantation has also made important contributions to the study of regeneration, particularly in organisms with strong regenerative abilities such as planarians and axolotl, by elucidating the cellular identities and interactions that regulate the growth and patterning of regenerating tissues. Transplant studies have revealed principles of potency12, spatial patterning13,14, contributions of specific tissues15,16, and roles for cellular memory12,17 in regeneration.
Zebrafish are a leading vertebrate model for the study of development and regeneration, including in the nervous system, due to their conserved genetic programs, high genetic tractability, external fertilization, large clutch size, and optical clarity18,19,20. Zebrafish are also highly amenable to transplantation at early developmental stages. The most prominent approach is the transplantation of cells from a labeled donor embryo to a host embryo at the blastula or gastrula stage to generate mosaic animals. Cells transplanted during the blastula stage will scatter and disperse as epiboly begins, producing a mosaic of labeled cells and tissues across the embryo21. Gastrula transplants allow for some targeting of transplanted cells according to a rough fate map as the shield forms and the A-P and D-V axes can be determined21. The resulting mosaics have been valuable in determining whether genes act cell autonomously, testing cell commitment, and mapping tissue movement and cell migration throughout development5,11. Mosaic zebrafish can be generated in several ways, including electroporation22, recombination23, and F0 transgenesis24 and mutagenesis25, but transplantation provides the greatest manipulability and precision in space, time, and number and types of cells. The current state of zebrafish transplantation is largely constrained to progenitor cells at early stages, with a few exceptions including transplantation of spinal motor neurons26,27, retinal ganglion cells28,29, and neural crest cells in the first 10-30 h post fertilization (hpf)30, and of hematopoietic and tumor cells in adult zebrafish5,31. Expanding transplantation methods to a broad range of ages, differentiation stages, and cell types would greatly enhance the power of this approach to provide insights into developmental and regenerative processes.
Here, we demonstrate a flexible and robust technique for high resolution cell transplantation effective in zebrafish embryos and larvae up to at least 7 days post fertilization. Transgenic host and donor fish expressing fluorescent proteins in target tissues can be used to extract single cells and transplant them with near-perfect spatial and temporal resolution. The optical clarity of zebrafish embryos and larvae allows for the transplanted cells to be imaged live as the host animal develops or regenerates. This approach has previously been used to examine how spatiotemporal signaling dynamics influence neuronal identity and axon guidance in the embryo32, and to examine the logic by which intrinsic and extrinsic factors promote axon guidance during regeneration in larval fish33. While we focus here on the transplantation of differentiated neurons, our method is widely applicable to both undifferentiated and differentiated cell types across many stages and tissues to address questions in development and regeneration.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>10 mL &#34;reservoir syringe&#34;</td>
			<td>Fisher Scientific</td>
			<td>14-955-459</td>
			<td></td>
		</tr>
		<tr>
			<td>150 mL disposable vacuum filter, .2 &#181;m, PES</td>
			<td>Corning</td>
			<td>431153</td>
			<td></td>
		</tr>
		<tr>
			<td>20 x 12 mm heating block</td>
			<td>Corning</td>
			<td>480122</td>
			<td></td>
		</tr>
		<tr>
			<td>3-way stopcock</td>
			<td>Braun Medical Inc.</td>
			<td>455991</td>
			<td></td>
		</tr>
		<tr>
			<td>3 x 1 Frosted glass slide</td>
			<td>VWR</td>
			<td>48312-004</td>
			<td></td>
		</tr>
		<tr>
			<td>40x water dipping objective</td>
			<td>Nikon</td>
			<td>MRD07420</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium chloride dihydrate</td>
			<td>Sigma-Aldrich</td>
			<td>C3306</td>
			<td></td>
		</tr>
		<tr>
			<td>Coarse Manipulator</td>
			<td>Narishige</td>
			<td>MN-4</td>
			<td></td>
		</tr>
		<tr>
			<td>Custom microsyringe pump</td>
			<td>University of Oregon</td>
			<td>N/A</td>
			<td>Manufactured by University of Oregon machine shop (tsa.uoregon@gmail.com). A commercially available alternative is listed below.</td>
		</tr>
		<tr>
			<td>Dumont #5 Forceps</td>
			<td>Fine Science Tools</td>
			<td>1129500</td>
			<td></td>
		</tr>
		<tr>
			<td>Eclipse FN1 &#34;Transplant Microscope&#34;</td>
			<td>Nikon</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>electrode handle</td>
			<td>World Precision Instruments</td>
			<td>5444</td>
			<td></td>
		</tr>
		<tr>
			<td>Feather Sterile Surgical Blade, #11</td>
			<td>VWR</td>
			<td>21899-530</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine micromanipulator, Three-axis Oil hydraulic&#160;</td>
			<td>Narishige</td>
			<td>MMO-203</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES pH 7.2</td>
			<td>Sigma-Aldrich</td>
			<td>H3375-100G</td>
			<td></td>
		</tr>
		<tr>
			<td>High Precision #3 Style Scalpel Handle</td>
			<td>Fisher Scientific</td>
			<td>12-000-163</td>
			<td></td>
		</tr>
		<tr>
			<td>Kimble Disposable Borosilicate Pasteur Pipette, Wide Tip, 5.75 in</td>
			<td>DWK Life Sciences</td>
			<td>63A53WT</td>
			<td></td>
		</tr>
		<tr>
			<td>KIMBLE Chromatography Adapter&#160;</td>
			<td>DWK Life Sciences</td>
			<td>420408-0000</td>
			<td></td>
		</tr>
		<tr>
			<td>Kimwipes</td>
			<td>Kimberly-Clark Professional</td>
			<td>34120</td>
			<td></td>
		</tr>
		<tr>
			<td>Light Mineral Oil</td>
			<td>Sigma-Aldrich</td>
			<td>M3516-1L</td>
			<td></td>
		</tr>
		<tr>
			<td>LSE digital dry bath heater, 1 block, 120 V</td>
			<td>Corning</td>
			<td>6875SB</td>
			<td></td>
		</tr>
		<tr>
			<td>Manual microsyringe pump</td>
			<td>World Precision Instruments</td>
			<td>MMP</td>
			<td>Commercial alternative to custom microsyringe pump</td>
		</tr>
		<tr>
			<td>Microelectrode Holder</td>
			<td>World Precision Instruments</td>
			<td>MPH310</td>
			<td></td>
		</tr>
		<tr>
			<td>MicroFil Pipette Filler</td>
			<td>World Precision Instruments</td>
			<td>MF28G67-5</td>
			<td></td>
		</tr>
		<tr>
			<td>Nail Polish</td>
			<td>Electron MIcroscopy Sciences</td>
			<td>72180</td>
			<td></td>
		</tr>
		<tr>
			<td>Nuclease-free water</td>
			<td>VWR</td>
			<td>82007-334</td>
			<td></td>
		</tr>
		<tr>
			<td>P-97 Flaming/Brown Type Micropipette Puller</td>
			<td>Sutter Instruments</td>
			<td>P-97</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin-streptomycin</td>
			<td>Sigma-Aldrich</td>
			<td>p4458-100ML</td>
			<td>5,000 units penicillin and 5 mg streptomycin/mL</td>
		</tr>
		<tr>
			<td>pipette pump 10 mL</td>
			<td>Bel-Art</td>
			<td>37898-0000</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium chloride</td>
			<td>Sigma-Aldrich</td>
			<td>P3911</td>
			<td></td>
		</tr>
		<tr>
			<td>Professional Super Glue</td>
			<td>Loctite</td>
			<td>LOC1365882</td>
			<td></td>
		</tr>
		<tr>
			<td>Round-Bottom Polystyrene Test Tubes</td>
			<td>Falcon</td>
			<td>352054</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium chloride</td>
			<td>Sigma-Aldrich</td>
			<td>S9888</td>
			<td></td>
		</tr>
		<tr>
			<td>Stage micrometer</td>
			<td>Meiji Techno America</td>
			<td>MA285</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringes without Needle, 50 mL</td>
			<td>BD Medical</td>
			<td>309635</td>
			<td></td>
		</tr>
		<tr>
			<td>Tricaine Methanosulfonate</td>
			<td>Syndel USA</td>
			<td>SYNCMGAUS03</td>
			<td></td>
		</tr>
		<tr>
			<td>Trilene XL smooth casting Fishing line</td>
			<td>Berkley</td>
			<td>XLFS6-15</td>
			<td></td>
		</tr>
		<tr>
			<td>Tubing, polyethylene No. 205</td>
			<td>BD Medical</td>
			<td>427445</td>
			<td></td>
		</tr>
		<tr>
			<td>UltraPure Low Melting Point Agarose</td>
			<td>Invitrogen</td>
			<td>16520050</td>
			<td></td>
		</tr>
		<tr>
			<td>Wiretrol II calibrated micropipettes</td>
			<td>Drummond</td>
			<td>50002010</td>
			<td></td>
		</tr>
	</tbody>
</table>

high-resolution cell transplantation in embryonic and larval zebrafish

Development and regeneration occur by a process of genetically encoded spatiotemporally dynamic cellular interactions. The use of cell transplantation between animals to track cell fate and to induce mismatches in the genetic, spatial, or temporal properties of donor and host cells is a powerful means of examining the nature of these interactions. Organisms such as chick and amphibians have made crucial contributions to our understanding of development and regeneration, respectively, in large part because of their amenability to transplantation. The power of these models, however, has been limited by low genetic tractability. Likewise, the major genetic model organisms have lower amenability to transplantation. 
The zebrafish is a major genetic model for development and regeneration, and while cell transplantation is common in zebrafish, it is generally limited to the transfer of undifferentiated cells at the early blastula and gastrula stages of development. In this article, we present a simple and robust method that extends the zebrafish transplantation window to any embryonic or larval stage between at least 1 and 7 days post fertilization. The precision of this approach allows for the transplantation of as little as one cell with near-perfect spatial and temporal resolution in both donor and host animals. While we highlight here the transplantation of embryonic and larval neurons for the study of nerve development and regeneration, respectively, this approach is applicable to a wide range of progenitor and differentiated cell types and research questions.

The outcomes of transplantation experiments are directly observed by visualizing fluorescently labeled donor cells in host animals at appropriate timepoints post transplantation using a fluorescence microscope. Here, we transplanted individual anterior vagus neurons at 3 dpf. Host animals were then incubated for 12 or 48 h, anesthetized, mounted in LMA on a glass coverslip, and imaged with a confocal microscope (Figure 5). At 12 h post transplantation (hpt), we observe a successfully transpl...

Author Spotlight: Mapping Cellular Connectivity in the Zebrafish Nervous System During Development and Regeneration

Here we present a protocol to transplant cells with high spatial and temporal resolution in zebrafish embryos and larvae at any stage between at least 1 and 7 days post fertilization.

High-resolution Cell Transplantation in Embryonic and Larval Zebrafish

Development and regeneration occur by a process of genetically encoded spatiotemporally dynamic cellular interactions. The use of cell transplantation ...

high-resolution-cell-transplantation-in-embryonic-and-larval-zebrafish

Research

JoVE Journal

Developmental Biology

721 Views.  University of Minnesota. Here we present a protocol to transplant cells with high spatial and temporal resolution in zebrafish embryos and larvae at any stage between at least 1 and 7 days post fertilization.

Video: Author Spotlight: Mapping Cellular Connectivity in the Zebrafish Nervous System During Development and Regeneration

Large-scale Zebrafish Embryonic Heart Dissection for Transcriptional Analysis

The zebrafish embryonic heart is composed of only a few hundred cells, representing only a small fraction of the entire embryo. Therefore, to prevent the cardiac transcriptome from being masked by the global embryonic transcriptome, it is necessary to collect sufficient numbers of hearts for further analyses. Furthermore, as zebrafish cardiac development proceeds rapidly, heart collection and RNA extraction methods need to be quick in order to ensure homogeneity of the samples. Here, we present a rapid manual dissection protocol for collecting functional/beating hearts from zebrafish embryos. This is an essential prerequisite for subsequent cardiac-specific RNA extraction to determine cardiac-specific gene expression levels by transcriptome analyses, such as quantitative real-time polymerase chain reaction (RT-qPCR). The method is based on differential adhesive properties of the zebrafish embryonic heart compared with other tissues; this allows for the rapid physical separation of cardiac from extracardiac tissue by a combination of fluidic shear force disruption, stepwise filtration and manual collection of transgenic fluorescently labeled hearts.

To analyse cardiac gene expression profiles during zebrafish heart development, total RNA has to be extracted from isolated hearts. Here, we present a protocol for collecting functional/beating hearts by rapid manual dissection from zebrafish embryos to obtain cardiac-specific mRNA.

The zebrafish embryonic heart is composed of only a few hundred cells, representing only a small fraction of the entire embryo. Therefore, to prevent the ...

Live Imaging of Innate Immune and Preneoplastic Cell Interactions Using an Inducible Gal4/UAS Expression System in Larval Zebrafish Skin

Here we describe a method to conditionally induce epithelial cell transformation by the use of the 4-Hydroxytamoxifen (4-OHT) inducible KalTA4-ERT2/UAS expression system1 in zebrafish larvae, and the subsequent live imaging of innate immune cell interaction with HRASG12V expressing skin cells. The KalTA4-ERT2/UAS system is both inducible and reversible which allows us to induce cell transformation with precise temporal/spatial resolution in vivo. This provides us with a unique opportunity to live image how individual preneoplastic cells interact with host tissues as soon as they emerge, then follow their progression as well as regression. Recent studies in zebrafish larvae have shown a trophic function of innate immunity in the earliest stages of tumorigenesis2,3. Our inducible system would allow us to live image the onset of cellular transformation and the subsequent host response, which may lead to important insights on the underlying mechanisms for the regulation of oncogenic trophic inflammatory responses. We also discuss how one might adapt our protocol to achieve temporal and spatial control of ectopic gene expression in any tissue of interest.

Studying the earliest events of preneoplastic cell progression and innate immune cell interaction is pivotal to understand and treat cancer. Here we describe a method to conditionally induce epithelial cell transformations and the subsequent live imaging of innate immune cell interaction with HRASG12V expressing skin cells in zebrafish larvae.

Here we describe a method to conditionally induce epithelial cell transformation by the use of the 4-Hydroxytamoxifen (4-OHT) inducible KalTA4-ERT2/UAS ...

An Efficient Method to Obtain Dedifferentiated Fat Cells

Tissue engineering and cell therapy hold great promise clinically. In this regard, multipotent cells, such as mesenchymal stem cells (MSCs), may be used therapeutically, in the near future, to restore function to damaged organs. Nevertheless, several technical issues, including the highly invasive procedure of isolating MSCs and the inefficiency surrounding their amplification, currently hamper the potential clinical use of these therapeutic modalities. Herein, we introduce a highly efficient method for the generation of dedifferentiated fat cells (DFAT), MSC-like cells. Interestingly, DFAT cells can be differentiated into several cell types including adipogenic, osteogenic, and chondrogenic cells. Although other groups have previously presented various methods for generating DFAT cells from mature adipose tissue, our method allows us to produce DFAT cells more efficiently. In this regard, we demonstrate that DFAT culture medium (DCM), supplemented with 20% FBS, is more effective in generating DFAT cells than DMEM, supplemented with 20% FBS. Additionally, the DFAT cells produced by our cell culture method can be redifferentiated into several tissue types. As such, a very interesting and useful model for the study of tissue dedifferentiation is presented.

We have modified the conditions for DFAT cell generation and provide herein information regarding the use of an improved growth medium for the production of these cells.

Tissue engineering and cell therapy hold great promise clinically. In this regard, multipotent cells, such as mesenchymal stem cells (MSCs), may be used ...

Dissection of Larval Zebrafish Gonadal Tissue

Although wild zebrafish possess a ZZ/ZW sex-determination system, domesticated zebrafish have lost the sex chromosome. They utilize a polygenic sex determination system, where several genes distributed throughout the genome collectively determine the sex identities of individual fish. Currently, the genes involved in regulating gonad development and how they work remain elusive. Normally, isolating gonadal tissue is the first step to examine the sex developmental processes. Here, we present a procedure to isolate gonadal tissue from 17 dpf (days post fertilization) and 25 dpf zebrafish larvae. The isolated gonadal tissue may be subsequently examined by morphology and gene expression profiling.

Here, we present a protocol for isolating gonadal tissue of larval zebrafish, which will facilitate investigations of zebrafish sex differentiation and maintenance.

Although wild zebrafish possess a ZZ/ZW sex-determination system, domesticated zebrafish have lost the sex chromosome. They utilize a polygenic sex ...

A Novel Mammary Fat Pad Transplantation Technique to Visualize the Vessel Generation of Vascular Endothelial Stem Cells

Endothelial cells (ECs) are the fundamental building blocks of the vascular architecture and mediate vascular growth and remodeling to ensure proper vessel development and homeostasis. However, studies on endothelial lineage hierarchy remain elusive due to the lack of tools to gain access as well as to directly evaluate their behavior in vivo. To address this shortcoming, a new tissue model to study angiogenesis using the mammary fat pad has been developed. The mammary gland develops mostly in the postnatal stages, including puberty and pregnancy, during which robust epithelium proliferation is accompanied by extensive vascular remodeling. Mammary fat pads provide space, matrix, and rich angiogenic stimuli from the growing mammary epithelium. Furthermore, mammary fat pads are located outside the peritoneal cavity, making them an easily accessible grafting site for assessing the angiogenic potential of exogenous cells. This work also describes an efficient tracing approach using fluorescent reporter mice to specifically label the targeted population of vascular endothelial stem cells (VESCs) in vivo. This lineage tracing method, coupled with subsequent tissue whole-mount microscopy, enable the direct visualization of targeted cells and their descendants, through which the proliferation capability can be quantified and the differentiation commitment can be fate-mapped. Using these methods, a population of bipotent protein C receptor (Procr) expressing VESCs has recently been identified in multiple vascular systems. Procr+ VESCs, giving rise to both new ECs and pericytes, actively contribute to angiogenesis during development, homeostasis, and injury repair. Overall, this manuscript describes a new mammary fat pad transplantation and in vivo lineage tracing techniques that can be used to evaluate the stem cell properties of VESCs.

This work demonstrates a novel approach to assess the proliferation, differentiation, and vessel-forming potential of vascular endothelial stem cells (VESCs) through mammary fat pad transplantation followed by whole-mount tissue preparation for microscopic observation. A lineage tracing strategy to investigate the behavior of VESCs in vivo is also presented.

Endothelial cells (ECs) are the fundamental building blocks of the vascular architecture and mediate vascular growth and remodeling to ensure proper ...

High-resolution Episcopic Microscopy (HREM) - Simple and Robust Protocols for Processing and Visualizing Organic Materials

We provide simple protocols for generating digital volume data with the high-resolution episcopic microscopy (HREM) method. HREM is capable of imaging organic materials with volumes up to 5 x 5 x 7 mm3 in typical numeric resolutions between 1 x 1 x 1 and 5 x 5 x 5 &#181;m3. Specimens are embedded in methacrylate resin and sectioned on a microtome. After each section an image of the block surface is captured with a digital video camera that sits on the phototube connected to the compound microscope head. The optical axis passes through a green fluorescent protein (GFP) filter cube and is aligned with a position, at which the bock holder arm comes to rest after each section. In this way, a series of inherently aligned digital images, displaying subsequent block surfaces are produced. Loading such an image series in three-dimensional (3D) visualization software facilitates the immediate conversion to digital volume data, which permit virtual sectioning in various orthogonal and oblique planes and the creation of volume and surface rendered computer models. We present three simple, tissue specific protocols for processing various groups of organic specimens, including mouse, chick, quail, frog and zebra fish embryos, human biopsy material, uncoated paper and skin replacement material.

High-resolution Episcopic Microscopy HREM - Simple and Robust Protocols for Processing and Visualizing Organic Materials

We provide simple and robust protocols for processing biopsy material of various species, embryos of biomedical model organisms and samples of other organic tissues in order to permit digital volume data generation with the high-resolution episcopic microscopy method.

We provide simple protocols for generating digital volume data with the high-resolution episcopic microscopy (HREM) method. HREM is capable of imaging ...

AFM and Microrheology in the Zebrafish Embryo Yolk Cell

Elucidating the factors that direct the spatio-temporal organization of evolving tissues is one of the primary purposes in the study of development. Various propositions claim to have been important contributions to the understanding of the mechanical properties of cells and tissues in their spatiotemporal organization in different developmental and morphogenetic processes. However, due to the lack of reliable and accessible tools to measure material properties and tensional parameters in vivo, validating these hypotheses has been difficult. Here we present methods employing atomic force microscopy (AFM) and particle tracking with the aim of quantifying the mechanical properties of the intact zebrafish embryo yolk cell during epiboly. Epiboly is an early conserved developmental process whose study is facilitated by the transparency of the embryo. These methods are simple to implement, reliable, and widely applicable since they overcome intrusive interventions that could affect tissue mechanics. A simple strategy was applied for the mounting of specimens, AFM recording, and nanoparticle injections and tracking. This approach makes these methods easily adaptable to other developmental times or organisms.

The lack of tools to measure material properties and tensional parameters in vivo prevents validating their roles during development. We employed atomic force microscopy (AFM) and nanoparticle-tracking to quantify mechanical features on the intact zebrafish embryo yolk cell during epiboly. These methods are reliable and widely applicable avoiding intrusive interventions.

Elucidating the factors that direct the spatio-temporal organization of evolving tissues is one of the primary purposes in the study of development. ...

Establishment of Larval Zebrafish as an Animal Model to Investigate Trypanosoma cruzi Motility In Vivo

Chagas disease is a parasitic infection caused by Trypanosoma cruzi, whose motility is not only important for localization, but also for cellular binding and invasion. Current animal models for the study of T. cruzi allow limited observation of parasites in vivo, representing a challenge for understanding parasite behavior during the initial stages of infection in humans. This protozoan has a flagellar stage in both vector and mammalian hosts, but there are no studies describing its motility in vivo.The objective of this project was to establish a live vertebrate zebrafish model to evaluate T. cruzi motility in the vascular system. Transparent zebrafish larvae were injected with fluorescently labeled trypomastigotes and observed using light sheet fluorescence microscopy (LSFM), a noninvasive method to visualize live organisms with high optical resolution. The parasites could be visualized for extended periods of time due to this technique&#39;s relatively low risk of photodamage compared to confocal or epifluorescence microscopy. T. cruzi parasites were observed traveling in the circulatory system of live zebrafish in different-sized blood vessels and the yolk. They could also be seen attached to the yolk sac wall and to the atrioventricular valve despite the strong forces associated with heart contractions. LSFM of T. cruzi-inoculated zebrafish larvae is a valuable method that can be used to visualize circulating parasites and evaluate their tropism, migration patterns, and motility in the dynamic environment of the cardiovascular system of a live animal.

Establishment of Larval Zebrafish as an Animal Model to Investigate Trypanosoma cruzi Motility In Vivo

In this protocol, fluorescently labeled T. cruzi&#160;were injected into transparent zebrafish larvae, and parasite motility was observed in vivo using light sheet fluorescence microscopy.

Chagas disease is a parasitic infection caused by Trypanosoma cruzi, whose motility is not only important for localization, but also for cellular binding ...

Single-cell Photoconversion in Living Intact Zebrafish

Animal and plant tissue is composed of distinct populations of cells. These cells interact over time to build and maintain the tissue and can cause disease when disrupted. Scientists have developed clever techniques to investigate characteristics and natural dynamics of these cells within intact tissue by expressing fluorescent proteins in subsets of cells. However, at times, experiments require more selected visualization of cells within the tissue, sometimes at the single-cell or population-of-cells manner. To achieve this and visualize single cells within a population of cells, scientists have utilized single-cell photoconversion of fluorescent proteins. To demonstrate this technique, we show here how to direct UV light to an Eos-expressing cell of interest in an intact, living zebrafish. We then image those photoconverted Eos+ cells 24 h later to determine how they changed in the tissue. We describe two techniques: single cell photoconversion and photoconversions of populations of cell. These techniques can be used to visualize cell-cell interactions, cell-fate and differentiation, and cell migrations, making it a technique that is applicable in numerous biological questions.

Here, we present a protocol to show how cell photoconversion is achieved through UV exposure to specific areas expressing the fluorescent protein, Eos, in living animals.

Animal and plant tissue is composed of distinct populations of cells. These cells interact over time to build and maintain the tissue and can cause ...

Electroretinogram Recording in Larval Zebrafish using A Novel Cone-Shaped Sponge-tip Electrode

The zebrafish (Danio rerio) is commonly used as a vertebrate model in developmental studies and is particularly suitable for visual neuroscience. For functional measurements of visual performance, electroretinography (ERG) is an ideal non-invasive method, which has been well established in higher vertebrate species. This approach is increasingly being used for examining the visual function in zebrafish, including during the early developmental larval stages. However, the most commonly used recording electrode for larval zebrafish ERG to date is the glass micropipette electrode, which requires specialized equipment for its manufacture, presenting a challenge for laboratories with limited resources. Here, we present a larval zebrafish ERG protocol using a cone-shaped sponge-tip electrode. The novel electrode is easier to manufacture and handle, more economical, and less likely to damage the larval eye than the glass micropipette. Like previously published ERG methods, the current protocol can assess outer retinal function through photoreceptor and bipolar cell responses, the a- and b-wave, respectively. The protocol can clearly illustrate the refinement of visual function throughout the early development of zebrafish larvae, supporting the utility, sensitivity, and reliability of the novel electrode. The simplified electrode is particularly useful when establishing a new ERG system or modifying existing small-animal ERG apparatus for zebrafish measurement, aiding researchers in the visual neurosciences to use the zebrafish model organism.

Here, we present a protocol that simplifies the measurement of light evoked electroretinogram responses from larval zebrafish. A novel cone-shaped sponge-tip electrode can help to make the study of visual development in larval zebrafish using the electroretinogram ERG easier to achieve with reliable outcomes and lower cost.

The zebrafish (Danio rerio) is commonly used as a vertebrate model in developmental studies and is particularly suitable for visual neuroscience. For ...