High-resolution Cell Transplantation in Embryonic and Larval Zebrafish

Lindsey S. Qian; Rabab Ibrahim; Adam  J. Isabella

doi:10.3791/67218

JoVE Journal > Developmental Biology

Biologia dello sviluppo

High-resolution Cell Transplantation in Embryonic and Larval Zebrafish

Published: July 05, 2024

doi:

10.3791/67218

Lindsey S. Qian¹, Rabab Ibrahim^1,2, Adam J. Isabella^1,2

¹Department of Genetics, Cell Biology, and Development,University of Minnesota, ²Graduate Program in Molecular, Cellular, Developmental Biology and Genetics,University of Minnesota

Summary

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.

Abstract

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.

Introduction

Cell transplantation has a long and storied history as a foundational technique in developmental biology. Around the turn of the 20^th 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'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.

Protocol

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)

Assemble the transplant microscope per manufacturer's instructions.
NOTE: This protocol uses an upright fluorescence microscope with a 40x water dipping objective.
Assemble the coarse and fine micromanipulators and mount them on the right side of the microscope according to the manufacturer'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.
Attach the microelectrode holder to the electrode handle, and mount on the micromanipulator.
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.
Connect the opposite side of the polyethylene tubing to the microelectrode holder using the chromatography adapter.
Fill the 10 mL reservoir syringe with light mineral oil and mount it on the top side of the stopcock.
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.

2. Prepare embryo pushers.

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.

3. Prepare solutions.

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 °C.
Prepare Ringer'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%.

4. Prepare donor and host animals for transplantation.

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.
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.
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 °C in a dry bath heater.
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.
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.
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.
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.
Apply RPT to the slide until the agarose drops are fully submerged (Figure 2D).
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).
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.

5. Prepare the transplant needle.

Insert a micropipette into the micropipette puller and pull a microinjection needle.
NOTE: The needle'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.
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 µm diameter), needles should be broken to an inner diameter of 10 µm, which corresponds to an outer diameter of 20 µm. If needed, the size, shape, and/or angle of the needle tip can be refined with a microforge or microgrinder³⁶.
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.
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.
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° from horizontal (Figure 3C).
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.
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'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.

6. Transplantation

NOTE: All needle movements should be done using the fine micromanipulator for this section.

Move the stage to the right until the donor animal is brought back into view, being cautious to avoid accidental penetration with the needle.
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.
Insert the needle into the donor animal.
NOTE: Repeated in-and-out motions with the needle can aid in penetrating the skin.
Immediately re-stabilize the oil meniscus in the needle tip with the microsyringe pump, as described in step 5.7.
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.
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.
Being careful to avoid contacting animals or agarose with the needle, reposition the stage to bring the first host animal into view.
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).
Insert the needle into the host animal and immediately re-stabilize the oil meniscus in the needle tip with the microsyringe pump.
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.
Remove the needle from the host and immediately re-stabilize the oil meniscus in the needle tip with the microsyringe pump.
Repeat steps 6.7-6.11 for all remaining hosts.

7. Host animal recovery

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.
Remove the slide from the transplant microscope and place it under a dissecting microscope.
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'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.

Representative Results

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.

Figure 1: Overview of the transplant apparatus. (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. Please click here to view a larger version of this figure.

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's solution with penicillin-streptomycin and tricaine. Scale bars = 5 mm. Please click here to view a larger version of this figure.

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 µm Please click here to view a larger version of this figure.

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 µm Please click here to view a larger version of this figure.

Figure 5: Representative results. (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'). The donor neuron has extended several neurites (arrows in A'). (B) The vagus motor nucleus (B-B') and pharyngeal axon branches (B''-B''', axon branches labeled) of a 2 dpt host (magenta) containing a donor neuron (green, black in B', B'''). 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'), likely representing a fragment of a dead donor neuron (arrowhead), is present. Scale bars = 20 µm. Abbreviations: dpt = days post transplantation; hpt = hours post transplantation. Please click here to view a larger version of this figure.

Discussion

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.

Divulgazioni

The authors have nothing to disclose.

Acknowledgements

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.

Materials

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

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