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 ganglion cells at 16-18 hpf and 30-33 hpf, respectively, have been reported26,27,28. Drawing inspiration from these previous efforts, this work further enhances the power of zebrafish transplantation by describing an approach that is applicable to late embryonic and larval stages, opening the door for its use in regeneration studies, and that integrates modern fluorescent transgenic approaches to enhance labeling, transfer, and long-term tracking of cells.
This protocol presents new opportunities to transplant cells of many different differentiation stages and ages, including in the regenerative context. For example, transplantation of differentiated neurons has been used to examine how signaling dynamics affect neuronal identity by changing the spatial and temporal contexts in which neurons develop32; to examine the timing of neuronal determination26; as a means to injure axons and track their regrowth with single-cell resolution33; to examine the cell-autonomous role of a receptor during axon regeneration via mutant-to-wild type transplants33; and to examine the role of target memory in regenerative axon guidance33. While we have thus far focused on neuronal transplantation, we are not aware of any characteristic of neurons that make them uniquely suited to this approach; rather, we believe that this approach is applicable to many cell types. We hope and expect that researchers will find creative ways to examine the dynamics of development and regeneration in many contexts using this technique.
Critical considerations in this protocol include selection of cell labels, suitability of cells for transplantation, and tissue accessibility. A bright, stable, permanent fluorescent label for donor cells is important to facilitate both transplantation and subsequent tracking. Both injected dyes and transgenes may be used, although transgenes can be more permanent and more specific to the cells of interest. Resistance to photobleaching also prolongs the time one can take in performing transplantation. The donor cells must be sufficiently loosely adherent and of appropriate shape to be drawn into the needle from the surrounding tissue/extracellular matrix without excessive damage. Modifications for different cell types include adjusting the needle bore size, which should be slightly larger than the diameter of the cell of interest; properly orienting the animal during mounting to facilitate access to the cells of interest; and optimizing the level of suction or physical disruption required to loosen and pull up cells. As the animal ages, certain tissues may become visually and/or physically inaccessible, although we are not aware of any specific age limits. For example, the skull will likely eventually limit access to neurons. Modifications may include the use of nonpigmented animals to more easily visualize internal tissues, or the making of preliminary small incisions, adjustments to microcapillary tube thickness, or pulled needle shape to facilitate needle entry.
Limitations of this protocol include cells of certain types and ages not being amenable to transplantation, as described above, and the difficulty of maintaining tissue-level structures during transplantation. Because this procedure requires that individual cells be dissociated from their surroundings during removal, cell-cell interactions and higher-order structures will be lost. The innate immune system, which becomes functional during embryogenesis, may respond to tissue damage or the presence of dead cells and debris at the transplantation site, which could affect donor cell survival37,38. At later stages, donor cells may also be rejected by the adaptive immune system, which becomes functional at 3-6 weeks post fertilization39,40. This issue may be avoided by transplanting into hosts lacking adaptive or innate immunity41,42,43,44.
Potential difficulties with the protocol that may require troubleshooting include the following: First, material within the needle should move in a smooth and responsive manner during suction and pressure. Jumpy and inconsistent movement is likely caused by the presence of air bubbles in the hydraulic line or a partial clog in the needle; therefore, caution should be taken to remove all air bubbles from the needle and hydraulic line during setup. If air bubbles are found, return the needle to loading position using the coarse micromanipulator, remove the needle from the holder, and flush the needle and line with mineral oil before reassembly. Needle clogging can be mitigated by avoiding jagged needle breaks. Clogs can sometimes be resolved by expelling mineral oil out of the needle tip to push out the clog or by fully retracting and then reinserting the needle into the RPT; otherwise, a new needle should be used. Second, there may be difficulty loosening cells during removal. Breaking the needle tip at an angle for a beveled shape, moving the needle back and forth during suction, and adjusting suction strength can help loosen the cells for detachment from tissue. A microgrinder can also be used to prepare beveled needles with a specific opening size and angle36. Lastly, there may be damage to cells during removal. Severely damaged cells may lose their defined shape and appear as small fragments or amorphous fluorescent blobs in the injection needle. Failure of cells to survive after transplant may also be an indication of damage and can be easily assayed by an absence of fluorescently marked cells in the host. Troubleshooting for repeated cell damage could include increasing the bore size of the needle and limiting the amount of suction applied to reduce shear force. A partially clogged needle may also increase shear forces, leading to damage.

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><tbody><tr><td>10 mL &quot;reservoir syringe&quot;</td><td>Fisher Scientific</td><td>14-955-459</td><td></td></tr><tr><td>150 mL disposable vacuum filter, .2 &amp;micro;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 &quot;Transplant Microscope&quot;</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&amp;nbsp;</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&amp;nbsp;</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 transplanted donor neuron (Figure 5A). We can confirm correct positioning of the cell, as it is situated among the host vagus neurons in the anterior region of the host vagus motor nucleus; the cell also appears intact and healthy, as indicated by the extension of several short neurite protrusions (Figure 5A). At 2 days post transplantation (dpt), we can again confirm that a single neuron has been successfully transplanted into the correct position of the host nucleus (Figure 5B). At this point, we observe that the neuron has extended a new axon to the 4th pharyngeal arch (Figure 5B). Direct observation also reveals when the procedure has failed. In this example, we observe a 2 dpt host in which no donor neuron is present; rather, a small green speck, likely a fragment of a donor neuron that died after transplantation, is apparent (Figure 5C). Although transplant success rates will vary based on user experience and the number and type of transplanted cells, we experience a rate of success (defined as the presence of a surviving neuron that has extended an axon in the host at 2-3 dpt) of 66% (n = 453 transplants) for vagus motor neuron transplants performed at 3-4 dpf33.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/67218/67218fig01.jpg" /> 
Figure 1: Overview of the transplant apparatus.&#160;(A) Upright fluorescence microscope; (B) 40x water dipping objective; (C) coarse micromanipulator; (D) fine micromanipulator; (E) microelectrode holder; (F) electrode handle; (G) three-way stopcock; (H) microsyringe pump; (I) polyethylene tubing; (J) chromatography adapters; (K) 10 mL reservoir syringe. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67218/67218fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/67218/67218fig02.jpg" /> 
Figure 2: Mounting animals for transplantation. (A) A 3 inch x 1 inch frosted glass slide with clear nail polish applied in a rectangular outline (dashed line). (B) Donor (arrow) and host (arrowhead) animals mounted in drops of agarose, vertically aligned with the transplantation target site (hindbrain) oriented toward the right. (C) Agarose with right edge and wedges cut out to expose transplantation target site. Inset: zoom of boxed region. (D) Transplant slide flooded with Ringer&#39;s solution with penicillin-streptomycin and tricaine. Scale bars = 5 mm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67218/67218fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/67218/67218fig03.jpg" /> 
Figure 3: Transplant needle preparation. (A) Broken transplant needle atop a stage micrometer. (B) Pipette filler (1) inserted into transplant needle (2) to fill needle with mineral oil. (C) Prepared transplant slide and needle in place on the transplant microscope. 1: Water meniscus between the 40x dipping objective and the flooded slide 2: Transplant needle 3: Microelectrode holder. (D) Tip of a broken transplant needle with stable oil meniscus (arrowhead) viewed through transplant microscope at 40x magnification. Scale bars = 100 &#181;m <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67218/67218fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/67218/67218fig04.jpg" /> 
Figure 4: The transplantation process under the microscope. (A) Needle positioned outside the donor animal, aimed at vagus motor neurons (green, arrowhead). (B) Needle positioned inside the donor animal after taking up fluorescently marked donor neurons (arrowheads). (C) Needle containing several donor neurons (arrowheads) positioned outside host animal, aimed at anterior host vagus motor neurons (red, asterisk). (D) Host animal post transplantation with needle withdrawn. A cluster of transplanted donor neurons expressing GFP (arrowhead) have been expelled into the host. Scale bars = 30 &#181;m <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67218/67218fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/67218/67218fig05.jpg" /> 
Figure 5: Representative results.&#160;(A,B) Successful transplants. (A) The anterior region of a 12 hpt host vagus motor nucleus (magenta) containing a donor neuron (green, black in A&#39;). The donor neuron has extended several neurites (arrows in A&#39;). (B) The vagus motor nucleus (B-B&#39;) and pharyngeal axon branches (B&#39;&#39;-B&#39;&#39;&#39;, axon branches labeled) of a 2 dpt host (magenta) containing a donor neuron (green, black in B&#39;, B&#39;&#39;&#39;). The donor neuron has extended a new axon to branch 4 (arrowheads). Figure 5A,B represent different animals. (C) Unsuccessful transplant. The vagus motor nucleus of a 2 dpt host (magenta). No donor neuron is present; rather, a small speck (green, black in C&#39;), likely representing a fragment of a dead donor neuron (arrowhead), is present. Scale bars = 20 &#181;m. Abbreviations: dpt = days post transplantation; hpt = hours post transplantation. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67218/67218fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

784 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

Isolation and Characterization of Single Cells from Zebrafish Embryos

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

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

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

In Vivo Imaging of Transgenic Gene Expression in Individual Retinal Progenitors in Chimeric Zebrafish Embryos to Study Cell Nonautonomous Influences

The genetic and technical strengths have made the zebrafish vertebrate a key model organism in which the consequences of gene manipulations can be traced in vivo throughout the rapid developmental period. Multiple processes can be studied including cell proliferation, gene expression, cell migration and morphogenesis. Importantly, the generation of chimeras through transplantations can be easily performed, allowing mosaic labeling and tracking of individual cells under the influence of the host environment. For example, by combining functional gene manipulations of the host embryo (e.g., through morpholino microinjection) and live imaging, the effects of extrinsic, cell nonautonomous signals (provided by the genetically modified environment) on individual transplanted donor cells can be assessed. Here we demonstrate how this approach is used to compare the onset of fluorescent transgene expression as a proxy for the timing of cell fate determination in different genetic host environments.
In this article, we provide the protocol for microinjecting zebrafish embryos to mark donor cells and to cause gene knockdown in host embryos, a description of the transplantation technique used to generate chimeric embryos, and the protocol for preparing and running in vivo time-lapse confocal imaging of multiple embryos. In particular, performing multiposition imaging is crucial when comparing timing of events such as the onset of gene expression. This requires data collection from multiple control and experimental embryos processed simultaneously. Such an approach can easily be extended for studies of extrinsic influences in any organ or tissue of choice accessible to live imaging, provided that transplantations can be targeted easily according to established embryonic fate maps.

In Vivo Imaging of Transgenic Gene Expression in Individual Retinal Progenitors in Chimeric Zebrafish Embryos to Study Cell Nonautonomous Influences

Live tracking of individual WT retinal progenitors in distinct genetic backgrounds allows for the assessment of the contribution of cell non-autonomous signaling during neurogenesis. Here, a combination of gene knockdown, chimera generation via embryo transplantation and in vivo time-lapse confocal imaging was utilized for this purpose.

The genetic and technical strengths have made the zebrafish vertebrate a key model organism in which the consequences of gene manipulations can be traced ...

Ortho- and Ectopic Zebrafish Xeno-Engraftment of Ocular Melanoma to Recapitulate Primary Tumor and Experimental Metastasis Development

There are currently no animal models for metastatic ocular melanoma. The lack of metastatic disease models has greatly hampered the research and development of novel strategies for the treatment of metastatic ocular melanoma. In this protocol we delineate a quick and efficient way to generate embryonic zebrafish models for both the primary and disseminated stage of ocular melanoma, using retro-orbital orthotopic and intravascular ectopic cell engraftment, respectively. Combining these two different engraftment strategies we can recapitulate the etiology of cancer in its totality, progressing from primary, localized tumor growth under the eye to a peri-vascular metastasis formation in the tail. These models allow us to quickly and easily modify the cancer cells prior to implantation with specific labeling, genetic or chemical interference; and to treat the engrafted hosts with (small molecular) inhibitors to attenuate tumor development. 
Here, we describe the generation and quantification of both orthotopic and ectopic engraftment of ocular melanomas (conjunctival and uveal melanoma) using fluorescently labelled stable cell lines. This protocol is also applicable for engraftment of primary cells derived from patient biopsy and patient/PDX derived material (manuscript in preparation). Within hours post engraftment cell migration and proliferation can be visualized and quantified. Both tumor foci are readily available for imaging with both epifluorescence microscopy and confocal microscopy. Using these models, we can confirm or refute the activity of either chemical or genetic inhibition strategies within as little as 8 days after the onset of the experiment, allowing not only highly efficient screening on stable cell lines, but also enables patient directed screening for precision medicine approaches.

Here, we present a protocol to establish versatile orthotopic and ectopic zebrafish xenograft models for ocular melanoma to assess the growth kinetics of the primary tumor, dissemination, extravasation and distant, peri-vascular metastasis formation and the effect of chemical inhibition thereon.

There are currently no animal models for metastatic ocular melanoma. The lack of metastatic disease models has greatly hampered the research and ...

Cancer Research

Culture and Transfection of Zebrafish Primary Cells

Zebrafish embryos are transparent and develop rapidly outside the mother, thus allowing for excellent in vivo imaging of dynamic biological processes in an intact and developing vertebrate. However, the detailed imaging of the morphologies of distinct cell types and subcellular structures is limited in whole mounts. Therefore, we established an efficient and easy-to-use protocol to culture live primary cells from zebrafish embryos and adult tissue.
In brief, 2 dpf zebrafish embryos are dechorionated, deyolked, sterilized, and dissociated to single cells with collagenase. After a filtration step, primary cells are plated onto glass bottom dishes and cultivated for several days. Fresh cultures, as much as long term differenciated ones, can be used for high resolution confocal imaging studies. The culture contains different cell types, with striated myocytes and neurons being prominent on poly-L-lysine coating. To specifically label subcellular structures by fluorescent marker proteins, we also established an electroporation protocol which allows the transfection of plasmid DNA into different cell types, including neurons. Thus, in the presence of operator defined stimuli, complex cell behavior, and intracellular dynamics of primary zebrafish cells can be assessed with high spatial and temporal resolution. In addition, by using adult zebrafish brain, we demonstrate that the described dissociation technique, as well as the basic culturing conditions, also work for adult zebrafish tissue.

We present an efficient and easy-to-use protocol for preparing primary cell cultures of zebrafish embryos for transfection and live cell imaging as well as a protocol to prepare primary cells from adult zebrafish brain.

Zebrafish embryos are transparent and develop rapidly outside the mother, thus allowing for excellent in vivo imaging of dynamic biological processes in ...

Tissue Targeted Embryonic Chimeras: Zebrafish Gastrula Cell Transplantation

Certain fundamental questions in the field of developmental biology can only be answered when cells are placed in novel environments or when small groups of cells in a larger context are altered.  Watching how one cell interacts with and behaves in a unique environment is essential to characterizing cell functions.  Determining how the localized misexpression of a specific protein influences surrounding cells provides insightful information on the roles that protein plays in a variety of developmental processes.  Our lab uses the zebrafish model system to uniquely combine genetic approaches with classical transplantation techniques to generate genotypic or phenotypic chimeras.  We study neuron-glial cell interactions during the formation of forebrain commissures in zebrafish. This video describes a method that allows our lab to investigate the role of astroglial populations in the diencephalon and the roles of specific guidance cues that influence projecting axons as they cross the midline. Due to their transparency zebrafish embryos are ideal models for this type of ectopic cell placement or localized gene misexpression. Tracking transplanted cells can be accomplished using a vital dye or a transgenic fish line expressing a fluorescent protein. We demonstrate here how to prepare donor embryos with a vital dye tracer for transplantation, as well as how to extract and transplant cells from one gastrula staged embryo to another.  We present data showing ectopic GFP+ transgenic cells within the forebrain of zebrafish embryos and characterize the location of these cells with respect to forebrain commissures.  In addition, we show laser scanning confocal timelapse microscopy of Alexa 594 labeled cells transplanted into a GFP+ transgenic host embryo.  These data provide evidence that gastrula staged transplantation enables the targeted positioning of ectopic cells to address a variety of questions in Developmental Biology.

Zebrafish cell transplantation enables the combination of genetics and embryology to generate tissue specific chimeras. This video demonstrates gastrula staged cell transplantations that have allowed our lab to investigate the roles of astroglial populations and specific guidance cues during commissure formation in the forebrain.

Certain fundamental questions in the field of developmental biology can only be answered when cells are placed in novel environments or when small groups ...

Biology

Normal and Malignant Muscle Cell Transplantation into Immune Compromised Adult Zebrafish

Zebrafish have become a powerful tool for assessing development, regeneration, and cancer. More recently, allograft cell transplantation protocols have been developed that permit engraftment of normal and malignant cells into irradiated, syngeneic, and immune compromised adult zebrafish. These models when coupled with optimized cell transplantation protocols allow for the rapid assessment of stem cell function, regeneration following injury, and cancer. Here, we present a method for cell transplantation of zebrafish adult skeletal muscle and embryonal rhabdomyosarcoma (ERMS), a pediatric sarcoma that shares features with embryonic muscle, into immune compromised adult rag2E450fs homozygous mutant zebrafish. Importantly, these animals lack T cells and have reduced B cell function, facilitating engraftment of a wide range of tissues from unrelated donor animals. Our optimized protocols show that fluorescently labeled muscle cell preparations from &#945;-actin-RFP transgenic zebrafish engraft robustly when implanted into the dorsal musculature of rag2 homozygous mutant fish. We also demonstrate engraftment of fluorescent-transgenic ERMS where fluorescence is confined to cells based on differentiation status. Specifically, ERMS were created in AB-strain myf5-GFP; mylpfa-mCherry double transgenic animals and tumors injected into the peritoneum of adult immune compromised fish. The utility of these protocols extends to engraftment of a wide range of normal and malignant donor cells that can be implanted into dorsal musculature or peritoneum of adult zebrafish.

Here, we present a protocol for cell transplantation of zebrafish skeletal muscle and embryonal rhabdomyosarcoma (ERMS) into adult immune compromised rag2E450fs homozygous mutant zebrafish. This protocol allows for the efficient analysis of regeneration and malignant transformation of muscle cells.

Zebrafish have become a powerful tool for assessing development, regeneration, and cancer. More recently, allograft cell transplantation protocols have ...

Immunology and Infection

Transplantation of GFP-expressing Blastomeres for Live Imaging of Retinal and Brain Development in Chimeric Zebrafish Embryos

Cells change extensively in their locations and property during embryogenesis. These changes are regulated by the interactions between the cells and their environment. Chimeric embryos, which are composed of cells of different genetic background, are great tools to study the cell-cell interactions mediated by genes of interest. The embryonic transparency of zebrafish at early developmental stages permits direct visualization of the morphogenesis of tissues and organs at the cellular level. Here, we demonstrate a protocol to generate chimeric retinas and brains in zebrafish embryos and to perform live imaging of the donor cells. The protocol covers the preparation of transplantation needles, the transplantation of GFP-expressing donor blastomeres to GFP-negative hosts, and the examination of donor cell behavior under live confocal microscopy. With slight modifications, this protocol can also be used to study the embryonic development of other tissues and organs in zebrafish. The advantages of using GFP to label donor cells are also discussed. 

We demonstrate a protocol to generate chimeric zebrafish embryos for live imaging cellular behavior during embryogenesis.

Cells change extensively in their locations and property during embryogenesis. These changes are regulated by the interactions between the cells and their ...

Utilizing Larval Zebrafish for Efficient In Vivo Tumor Studies

A Simple, Rapid, and Effective Method for Tumor Xenotransplantation Analysis in Transparent Zebrafish Embryos

In vivo studies of tumor behavior are a staple of cancer research; however, the use of mice presents significant challenges in cost and time. Here, we present larval zebrafish as a transplant model that has numerous advantages over murine models, including ease of handling, low expense, and short experimental duration. Moreover, the absence of an adaptive immune system during larval stages obviates the need to generate and use immunodeficient strains. While established protocols for xenotransplantation in zebrafish embryos exist, we present here an improved method involving embryo staging for faster transfer, survival analysis, and the use of flow cytometry to assess disease burden. Embryos are staged to facilitate rapid cell injection into the yolk of the larvae and cell marking to monitor the consistency of the injected cell bolus. After injection, embryo survival analysis is assessed up to 7 days post injection (dpi). Finally, disease burden is also assessed by marking transferred cells with a fluorescent protein and analysis by flow cytometry. Flow cytometry is enabled by a standardized method of preparing cell suspensions from zebrafish embryos, which could also be used in establishing the primary culture of zebrafish cells. In summary, the procedure described here allows a more rapid assessment of the behavior of tumor cells in vivo with larger numbers of animals per study arm and in a more cost-effective manner.

Author Spotlight: Advancing Cancer Therapeutics Using Tumor Xenotransplantation in Zebrafish

We describe a protocol for xenotransplantation into the yolk of transparent zebrafish embryos that is optimized by a simple, rapid staging method. Post-injection analyses include survival and assessing the disease burden of xenotransplanted cells by flow cytometry.

In vivo studies of tumor behavior are a staple of cancer research; however, the use of mice presents significant challenges in cost and time. Here, we ...

A Simple and Effective Transplantation Device for Zebrafish Embryos

Classical embryological manipulations, such as removing cells and transplanting cells within or between embryos, are powerful techniques to study complex developmental processes. Zebrafish embryos are ideally suited for these manipulations since they are easily accessible, relatively large in size, and transparent. However, previously developed devices for cell removal and transplantation are cumbersome to use or expensive to purchase. In contrast, the transplantation device presented here is economical, easy to assemble, and simple to use. In this protocol, we first introduce the handling of the transplantation device as well as its assembly from commercially and widely available parts. We then present three applications for its use: generation of ectopic clones to study signal dispersal from localized sources, extirpation of cells to produce size-reduced embryos, and germline transplantation to generate maternal-zygotic mutants. Finally, we show that the tool can also be used for embryological manipulations in other species such as the Japanese rice fish medaka.

Embryological manipulations such as extirpation and transplantation of cells are important tools to study early development. This protocol describes a simple and effective transplantation device to perform these manipulations in zebrafish embryos.

Classical embryological manipulations, such as removing cells and transplanting cells within or between embryos, are powerful techniques to study complex ...

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 ...

High-resolution Cell Transplantation in Embryonic and Larval Zebrafish

Summary

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Chapters in this video

Isolation and Characterization of Single Cells from Zebrafish Embryos

In Vivo Imaging of Transgenic Gene Expression in Individual Retinal Progenitors in Chimeric Zebrafish Embryos to Study Cell Nonautonomous Influences

Ortho- and Ectopic Zebrafish Xeno-Engraftment of Ocular Melanoma to Recapitulate Primary Tumor and Experimental Metastasis Development

Culture and Transfection of Zebrafish Primary Cells

Tissue Targeted Embryonic Chimeras: Zebrafish Gastrula Cell Transplantation

Normal and Malignant Muscle Cell Transplantation into Immune Compromised Adult Zebrafish

Transplantation of GFP-expressing Blastomeres for Live Imaging of Retinal and Brain Development in Chimeric Zebrafish Embryos

A Simple, Rapid, and Effective Method for Tumor Xenotransplantation Analysis in Transparent Zebrafish Embryos

A Simple and Effective Transplantation Device for Zebrafish Embryos

Imaging and 3D Reconstruction of Cerebrovascular Structures in Embryonic Zebrafish