We thank Julie R. Perlin for helpful comments on the manuscript. D.I.S. is supported by grants from the American Society of Hematology, the Cooley's Anemia Foundation, and the NIH (K01DK085217 and R03DK100672). E.J.H. is a Howard Hughes Medical Institute Fellow of the Helen Hay Whitney Foundation. B.L. is a Howard Hughes Medical Institute Medical Research Fellow. B.W.B. is supported by an Irvington Fellowship from the Cancer Research Institute and a Young Investigator Award from the Conquer Cancer Foundation of ASCO. L.I.Z. is supported by grants from the NIH (R01CA103846, P01HL032262, and R01HL04880), Taub Foundation for MDS Research, Harvard Stem Cell Institute, and is an investigator of the Howard Hughes Medical Institute.

This protocol was approved by Boston Children&#39;s Hospital Animal Care and Use Committee. This protocol is modified from a previously published method 15.
1. Preparation of Reagents (Days or Weeks in Advance)
<ol>
	<li>Prepare 1.5% agarose-coated dishes. Add 1.5 g agarose to 100 ml Zebrafish E3 medium in an Erlenmeyer flask. Heat, dissolve the agarose, and then pour approximately 5 ml into 100 mm diameter x 20 mm deep petri dishes. 
	Note: These are used for dechorionation of embryos (Step 3.1) and for the parabiotic surgery (Step 3.3). Store agarose-coated dishes at 4 &#176;C for a few weeks; take them out of the fridge and warm to RT the morning of the day of parabiotic surgery.</li>
	<li>Prepare 4% methyl cellulose. Dissolve 4 g of methyl cellulose powder in 100 ml of E3 in an Erlenmeyer flask with a stir bar. Place the flask on a stir plate at 4 &#176;C, set to the lowest speed, and let it mix for up to 2 days.
	<ol>
		<li>Intermittently use a spatula to break up white clumps of methyl cellulose. The methyl cellulose crystals do not fully dissolve at this concentration. When the solution becomes clear and appears homogenous with no white clumps remaining, aliquot the solution into 1.5 ml microfuge tubes and freeze at -20 &#176;C for long term storage. 
		Note: Lower concentrations of methyl cellulose can be used; however, this higher percentage, more viscous methyl cellulose has yielded the best results.</li>
	</ol>
	</li>
	<li>Prepare High Calcium Ringer (HCR) solution. To make 400 ml of HCR solution, mix 8 ml of 5M NaCl (116 mM final), 400 &#181;l of 3M KCl (2.9 mM final), 800 &#181;l of 5M CaCl2 (10 mM final), and 2ml of 1M HEPES (5 mM final) with 390 ml of E3 media. 
	Note: Store HCR solution at 4 &#176;C for a few weeks; warm to RT the morning of the day of parabiotic surgery.</li>
	<li>Prepare 5 - 6 modified Pasteur pipettes for transferring blastula pairs. Briefly heat the end of the pipette over a Bunsen burner. Then, using large forceps, bend the end of the pipette to approximately 45&#160;&#176;C while it is still hot.
	<ol>
		<li>To ensure there are no sharp edges that might damage embryos, hold the tip of the pipette briefly in the flame for around 3 sec. This will smooth/polish the end of the pipette. This modified glass Pasteur pipette is used in conjunction with a 10 ml pipette pump (Figure 1) in Step 3.5 below.</li>
	</ol>
	</li>
	<li>Prepare glass needle tools for surgical stitching of blastula pairs. Pull 2 - 3 glass needles as done for microinjection of zebrafish embryos. Use lab-film to fix each glass needle to the end of a wood- or plastic-handled teasing needle (Figure 1A). Using forceps, break off the end of the needle at a diameter of approximately 20 - 30 &#181;m. Use a stage micrometer to guide this step (Figure 1B). 
	Note: The size of the needle tip is important. If the tip is too large it becomes difficult to control precisely and can cause excessive damage. If the tip is too small it is difficult to sufficiently wound the embryos.</li>
</ol>
2. Setting up Breeding Pairs of Zebrafish and Collection of Embryos (Day -1 to Day 0)
<ol>
	<li>Set up breeding pairs of adult zebrafish. Set up fish the night before the parabiosis surgery is to be conducted. Use dividers to separate males and females. To increase the odds that the desired embryos will be obtained, set up several pairs of each line. We find that typically 20 - 50% of the pairs will spawn.</li>
	<li>The next morning, pull the dividers and allow mating. Collect any embryos using a fine-mesh tea strainer, then transfer them to a Petri dish containing E3 embryo medium18.</li>
	<li>Micro-inject embryos as desired (e.g., with 25 pg plasmid DNA, 200 pg mRNA, or with 1 - 6 ng morpholino) within 1 hr of collection. Allow them to grow until 256-cell stage. If the DNA expression construct is placed under the control of a heat shock promoter (e.g., hsp70, Figure 4B), control the timing of gene expression by heat shocking the parabionts at 37 &#176;C at a later stage (e.g., at 36 hpf). 
	Note: As an alternative to a fluorescent transgene, fluorescent dextran can be added to the DNA, RNA, or morpholino injection mix (e.g.,&#160;dextran, cascade blue at 2 mg/ml). The dextran dye will not interfere with normal development and will allow for the injected embryo to be identified under the appropriate fluorescent light after parabiotic fusion. Alternatively, the parabiosis of a pigmented strain (e.g.,&#160;AB) with a non-pigmented strain (e.g.,&#160;Casper) could be used to identify the injected embryo at later stages.</li>
	<li>Incubate the embryos at 28.5 &#176;C.Use a stereoscope to monitor their development periodically as they near the 256-cell developmental stage (approximately 2.5 hpf). 
	Note: If needed, shift embryos to different temperatures to ensure stage matching between parabiotic partners (e.g., morpholino-injected embryos often develop slower than their un-injected counterparts. Shifting the un-injected embryos to RT while the morpholino-injected embryos are placed at 28.5 &#176;C will result in their stages aligning after a couple hours of development).</li>
</ol>
3. Generation of Parabiotic Zebrafish Embryos by Surgical Fusion of Developing Blastulae
<ol>
	<li>As the embryos approach the 256-cell stage (approximately 2.5 hpf), transfer the embryos from each genetic background (i.e., genetic mutant, transgenic lines and/or genetic manipulation) into separate agarose-coated dishes (e.g., one dish for morpholino-injected embryos and a second dish for control embryos). Alternatively, transfer the embryos to clean, scratch-free glass beakers or glass Petri dishes. 
	Note: Avoid using uncoated plastic dishes as the embryos will stick to the plastic and be damaged once out of their chorions.</li>
	<li>Decant the E3 media until just enough liquid remains to cover the embryos (approximately 7 - 10 ml). Add 200 &#181;l of 50 mg/ml mixture of proteases isolated from the extracellular fluid of Streptomyces griseus (proteases). Swirl the embryos gently every couple of minutes.
	<ol>
		<li>When 50% of the embryos have come out of their chorions, which typically takes 5 - 10 min, rinse them three times with a generous volume (approximately 15 - 20 ml) of fresh E3 embryo medium.
		<ol>
			<li>Dechorionate any embryos remaining in their chorions by gently drawing them up into the modified glass Pasteur pipette and then gently dispelling them. Once the embryos have been dechorionated and washed, replace the E3 media with HCR solution. As an alternative to proteases, remove the chorions manually using a fine forceps. Aim to have 60 - 100 dechorionated embryos available for each experimental condition. 
			Note: This is several fold more than will ultimately be used, but many embryos are likely to be damaged during handling or the surgery for parabiosis. Once the embryos are dechorionated, keep them submerged and do not allow them to touch the surface of the water, as the surface tension will cause them to be destroyed. Keep in mind that using a higher dose of proteases allows the embryos to come out of their chorions sooner and in a more synchronized fashion. When using a lower dose of proteases, the embryos take longer to emerge from their chorions and do so in a less synchronous fashion, which can result in a portion of the embryos being damaged by prolonged exposure to proteases.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Thaw the methylcellulose (made in Step 1.2 above) and centrifuge in a tabletop centrifuge on max speed for 10 min&#160;to pellet undissolved crystals which will damage the embryos or rupture their yolks.
	<ol>
		<li>After centrifugation, use the clear upper fraction (approximately 75% of the volume). Transfer 1 - 1.5 cm diameter drops of methyl cellulose into a 1.5% agarose-coated Petri dish. Take care to avoid drawing up any of the pelleted crystals from the bottom of the tube.</li>
		<li>Use a marker to predetermine the position of each droplet on the bottom side of the dish. Place 12 - 15 drops (2 - 3 aliquots&#39; worth) of methyl cellulose per 100 mm diameter petri dish. Prepare as many plates as needed per experiment. 
		Note: Use each droplet for the fusion of a single blastula pair.&#160;The mark will designate the location of each droplet once the media is added-at which point the drops become nearly invisible.</li>
	</ol>
	</li>
	<li>Prepare one 40 ml aliquot of antibiotic-supplemented HCR solution per agarose-coated dish prepared above in Step 3.3. To each 40 ml aliquot of HCR add penicillin-streptomycin at 50 U/ml, ampicillin at 50 U/ml, and kanamycin at 0.5 &#181;g/ml.
	<ol>
		<li>When ready to perform blastula fusions, carefully fill each of the dishes containing methyl cellulose droplets with 40 ml of the antibiotic-supplemented HCR solution. If the HCR solution is added too quickly, it can disrupt the droplets.</li>
	</ol>
	</li>
	<li>Attach the modified glass Pasteur pipette (made in Step 1.4 above) to a 10 ml pipette pump. Under a stereoscope collect one dechorionated embryo from each background (e.g.,&#160;one morpholino-injected embryo and one wild-type embryo).
	<ol>
		<li>Before dispensing the two embryos, use the Pasteur pipette to make a small depression in the middle of the methyl cellulose drop,&#160;then gently deposit both embryos into the depression. 
		Note: During the transfer process, turn the Pasteur pipette slightly upwards so that the embryos settle into the bend of the Pasteur pipette so as to avoid their making contact with the surface of the liquid at the top or bottom of the pipette (which can lead to their rupture).</li>
	</ol>
	</li>
	<li>Immediately after depositing the embryos into the methyl cellulose, use a wood- or plastic-handled teasing needle with a gel-loading pipet tip on the end (Figure 1A) to carefully reposition the embryos within the methyl cellulose such that their animal poles are directly touching one another.
	<ol>
		<li>Utilize gentle sweeping motions through the methyl cellulose close to the embryos to reposition them without directly touching them. Let the embryos sit for 5 min so the methyl cellulose will settle in around them. During this time, load the next pair into another drop. 
		Note: Avoid making direct contact with the embryos and take special care not to touch any part of the yolk sac, which will easily rupture.</li>
	</ol>
	</li>
	<li>Using the glass needle tool, carefully wound each embryo at their point of contact. With a gentle sewing motion, cells from the first embryo can be pulled into the second embryo and then back again. At the end of this motion, hold the needle in place for 2 - 3 sec and then draw it away very slowly. 
	Note: There is a period of about 2 - 3 sec after the embryos are wounded where the two tissues seem to be more adhesive and more likely to attach to one another. Thus, we believe that holding the needle in place briefly after wounding helps to keep the wounded tissues of each embryo in contact for the initial period of time immediately after wounding.</li>
	<li>Allow these embryos to sit for 10 - 15 min at RT. In the meantime, continue to stitch new pairs in other drops of methyl cellulose. After 10 - 15 min, assess whether the first pair has remained attached (Figure 2A). 
	Note: If the embryos are longer attached, repeat the stitching process for as long as the embryos are younger than around the 30% epiboly stage. Typically this allows for up to three attempts to stitch the embryos together. If the embryos appear to have maintained an attachment but the connection is not substantial, perform additional stitching at the edge of the connection to reinforce it. Avoid shaking or moving dishes during the parabiotic surgery and incubation periods. 
	Note: Despite their early developmental stage and perceived fragility, the cell masses of the embryos can tolerate a substantial amount of manipulation (i.e., wounding and stitching), and still develop normally. The yolk, however, can rupture with only the slightest touch, killing the embryo. When stitching the two blastulae with the glass needle, do not to make contact with the interface between the cell mass and yolk of each embryo.</li>
	<li>Incubate the embryos O/N&#160;at 28.5 &#176;C in the same dish and media (do not change the media). By the following morning, the embryos will have moved out of the mostly dissolved methyl cellulose droplets. Remove any embryos that were not successfully fused. Decant the media and replace with a fresh 25 ml E3. 
	Note: If working with a pigmented strain, add 500 &#181;l 0.15% (50x) phenylthiourea (PTU) to a final concentration of 0.003% to block pigmentation. Embryos will need to remain continuously in PTU to maintain suppression of pigmentation.</li>
</ol>
4. Microscope Setup, Image Acquisition, Processing and Analysis
<ol>
	<li>Prepare 0.8% low melting point (LMP) agarose: Add 0.8 g LMP agarose to 100 ml E3 and heat until fully dissolved. While hot, aliquot 1 ml of LMP agarose into 1.5 ml microfuge tubes in a 37 &#176;C heating block. The LMP agarose will remain molten at 37 &#176;C. Add 40 &#181;l of 4 mg/ml (25x), pH 7.0 tricaine to each 1 ml aliquot of LMP agarose at 37 &#176;C. Note: If working with a pigmented strain, add 20 &#181;l of 0.15% (50x) PTU to each 1ml aliquot.
	<ol>
		<li>Store the remaining LMP agarose for several months in sealed 50 ml tubes at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Anesthetize parabiotic embryos to be imaged by adding 1 ml of 4 mg/ml (25x), pH 7.0 tricaine to the 25 ml of E3 that the embryos are already in.</li>
	<li>Gently swirl the dish so that the embryos will pool in the middle. Then, using a wide-tipped plastic transfer pipette draw up the embryos in as little liquid as possible. Turn the pipette upright and gently bounce the embryos so that they settle to the very bottom of the pipette.
	<ol>
		<li>With the embryos at the bottom of the pipette, transfer them to a 1 ml aliquot of LMP agarose by lightly touching the pipette tip to the surface of the agarose. Avoid transferring excess liquid as this will dilute the agarose.</li>
	</ol>
	</li>
	<li>After expelling the excess liquid from the pipette, use the pipette to gently mix the embryos within the agarose, then use the pipette to transfer all of the agarose and embryos to the well of a glass-bottom (No. 1.5 coverslip), 6-well plate. 
	Note: Be sure to discard the pipette after transferring the embryos to the imaging plate. Residual agarose will set inside the pipette, rendering it ineffective for further use. Rather, use a new pipette each time.</li>
	<li>Under a stereoscope, use a gel-loading pipette tip fixed on the end of a wood-handled teasing needle to position the embryos down towards the cover glass (to ensure they are within the working distance of the microscope objective) and in the desired orientation for imaging. 
	Note: Reposition continuously until the agarose has fully set. Take care to mount the parabiotic embryos according to which embryo of the pair is to be imaged. Given the random orientation of the conjoined embryos, for some pairs it may be difficult to image both embryos. In this scenario, recover the embryos from the agarose using a forceps and a wide-tipped plastic transfer pipette and then remount for imaging from a different perspective.</li>
	<li>Once the agarose has set, cover with 2 - 3 ml of E3 supplemented with tricaine (and PTU if necessary).</li>
	<li>Image the embryos using an inverted wide-field epi-fluorescence, laser-scanning confocal, or spinning disk confocal microscope. Select the objective most appropriate for the desired imaging. For a whole-embryo field of view, use a 4x objective. Select a higher magnification, such as a 20x objective, to image a specific tissue16, 19. 
	Note: If using an upright microscope, mount in LMP agarose (similar to above) on a glass cover slip. Invert the glass coverslip onto a glass microscope slide that has a ring of vacuum grease filled with the same E3-tricaine-PTU16, 19.</li>
	<li>Process acquired images using free image analysis software packages such as NIH Image J/FIJI16, 19. 
	Note: Count cell numbers and quantify dynamics such as cell migration and cell division using segmentation and tracking algorithms16, 19.</li>
</ol>

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The authors declare that they have no conflict of interest.

Parabiotic fusion has been a powerful tool to investigate cellular functions of candidate genes in adult murine models and chick embryos10-14. More recently, a blastula fusion method has been described for generating conjoined zebrafish embryos15. In the present protocol, video-based tutorials are used to demonstrate and better describe the methodology for creating parabiotic zebrafish embryos of different genetic backgrounds in order to study temporal, cell-intrinsic, and cell-extrinsic roles of ca...

Creation of genetic mosaics (chimeras) between wild-type and genetically modified animals&#160;is a well-established and classical strategy for investigating cell-intrinsic versus cell-extrinsic functions of candidate genes1-6. Blastula transplantation in zebrafish has been widely utilized to generate chimeric embryos for studies of cell-autonomy7-9. Depending on the tissue of interest, however, it can be challenging to predictably target donor cells to the desired tissue (e.g., blood) 1-3, 7-9. Mouse geneticists have long utilized parabiotic surgical methods to generate conjoined organisms with a shared circulation 10-14. Because the parabiotic animals share a common bloodstream, interactions between cells that originated from one animal with cells of the other animal of a different genetic background can be studied 10-16. Recently, Karima Kissa&#39;s group elegantly demonstrated the ability to create conjoined zebrafish embryos and then use this system to study hematopoietic stem and progenitor cell (HSPC) migration15. In addition, zebrafish parabiosis was recently used to investigate the role of endothelial cadherin 5 in hematopoietic stem cell (HSC) emergence and migration,16 and to study the role of stromal cells in the HSPC niche during zebrafish development 17.&#160;
Unlike mice, zebrafish embryos are transparent and allow for direct visualization of cells during parabiotic development, making the system uniquely powerful. The utility of parabiosis in zebrafish, however, is limited by a steep learning curve, and parabiotic surgery on the delicate embryos can be technically challenging without detailed instruction and visual demonstration. The goal of this protocol is to provide a set of clear, step-by-step instructions to accompany video-based tutorials on how to generate parabiotic zebrafish embryos for studying temporal, cell-intrinsic, or cell-extrinsic functions of a candidate gene(s) by surgical fusion of developing blastulae. Key modifications and additional recommendations for increasing parabiont survival and new experimental applications are included.

<table><tbody><tr><td>Methyl cellulose</td><td>Sigma</td><td>M0387</td><td></td></tr><tr><td>Individual components for E3/HCR:</td><td></td><td>For 1L: 14,61g NaCl, 0,63g KCl, 2,43g CaCl&lt;sub&gt;2&lt;/sub&gt;-2H&lt;sub&gt;2&lt;/sub&gt;O, 1,99g MgSO&lt;sub&gt;4&lt;/sub&gt;</td><td></td></tr><tr><td>NaCl (Sodium chloride)</td><td>Sigma-Aldrich</td><td>S9888</td><td></td></tr><tr><td>KCl (Potassium chloride)</td><td>Sigma-Aldrich</td><td>P9541</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt; (Calcium chloride dihydrate)</td><td>Sigma-Aldrich</td><td>223506</td><td></td></tr><tr><td>MgSO&lt;sub&gt;4&lt;/sub&gt; (Magnessium sulfate heptahydrate)</td><td>Sigma-Aldrich</td><td>230391</td><td></td></tr><tr><td>Hepes (1M ) buffer solution</td><td>ThermoFisher</td><td>15630-080</td><td></td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td></td></tr><tr><td>&lt;strong&gt;Antibiotics:&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Pen/Strep</td><td>gibco by Life technologies</td><td>15140-120</td><td></td></tr><tr><td>Ampicilin sodium salt</td><td>Sigma Life Science</td><td>A0166</td><td></td></tr><tr><td>Kanamycin sulfate from Streptomyces kanamyceticus</td><td>Sigma Life Science</td><td>K1377</td><td></td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td></td></tr><tr><td>50 mg/ml Pronase from Streptomyces griseus</td><td>Roche</td><td>11459643001</td><td></td></tr><tr><td>capillary glass used for needles (Capillary Glass &amp;amp; Filaments)</td><td>Sutter Instrument</td><td>ITEM#: BF 100-50-10</td><td></td></tr><tr><td>Teasing needles with wooden handles</td><td>Fisher Scientific</td><td>S07894</td><td></td></tr><tr><td>Glass Pasteur pipettes</td><td>Fisherbrand</td><td>13-678-20A</td><td></td></tr><tr><td>10 ml pipette pump (green) (Pipette Pump Pipettor)</td><td>Novatech International</td><td>F37898-0000</td><td></td></tr><tr><td>100 mm diameter/ 20 mm deep plastic petri dishes (PETRI DISH, 100/20 MM, PS, CLEAR, WITH VENTS,&lt;br/&gt; HEAVY DESIGN, 15 PCS./BAG )</td><td>Greiner Bio-one</td><td>664102</td><td></td></tr><tr><td>Dextran, Cascade Blue, 10,000 MW, Anionic, Lysine Fixable</td><td>ThermoFisher</td><td>D-1976</td><td></td></tr><tr><td>PTU. working stock is 0.003% (50x is 0.15%). for 500 ml, 0.75 g N-Phenylthiourea</td><td>Sigma-Aldrich</td><td>P7629</td><td></td></tr><tr><td>Tricaine (powder) (Tricaine Methanesulfonato, Tricaine-S)</td><td>Western Chemical Inc</td><td>MS 222</td><td></td></tr><tr><td>LMP agarose (Ultrapure LMP agarose)</td><td>Invitrogen</td><td>16520100</td><td></td></tr><tr><td>plastic transfer pipette (just the wide ended one I think)</td><td>Fisherbrand</td><td>137115AM</td><td></td></tr><tr><td>Glass-bottom 6-well plates used for imaging</td><td>MatTek</td><td>P06G1.5-20-F</td><td></td></tr><tr><td>plastic western gel loading tip fixed on the end of a wood-handled dissecting needle (GELoader tips)</td><td>Eppendorf</td><td>22351656</td><td></td></tr><tr><td>glass cover slips, slides and vacuum grease if mounting for an upright microscope:</td><td></td><td></td><td></td></tr><tr><td>Vaccum grease ( Dow Corning&amp;reg; high-vacuum silicone grease&lt;br/&gt; colorless, weight 5.3 oz (tube) )</td><td>Dow Corning</td><td>Z273554</td><td></td></tr><tr><td>Glass cover slips</td><td>Corning Life Sciences</td><td>2960-244</td><td></td></tr></tbody></table>

generation of parabiotic zebrafish embryos by surgical fusion of developing blastulae

Surgical parabiosis of two animals of different genetic backgrounds creates a unique scenario to study cell-intrinsic versus cell-extrinsic roles for candidate genes of interest, migratory behaviors of cells, and secreted signals in distinct genetic settings. Because parabiotic animals share a common circulation, any blood or blood-borne factor from one animal will be exchanged with its partner and vice versa. Thus, cells and molecular factors derived from one genetic background can be studied in the context of a second genetic background. Parabiosis of adult mice has been &#160;used extensively to research aging, cancer, diabetes, obesity, and brain development. More recently, parabiosis of zebrafish embryos has been used to study the developmental biology of hematopoiesis. In contrast to mice, the transparent nature of zebrafish embryos permits the direct visualization of cells in the parabiotic context, making it a uniquely powerful method for investigating fundamental cellular and molecular mechanisms. The utility of this technique, however, is limited by a steep learning curve for generating the parabiotic zebrafish embryos. This protocol provides a step-by-step method on how to surgically fuse the blastulae of two zebrafish embryos of different genetic backgrounds to investigate the role of candidate genes of interest. In addition, the parabiotic zebrafish embryos are tolerant to heat shock, making temporal control of gene expression possible. This method does not require a sophisticated set-up and has broad applications for studying cell migration, fate specification, and differentiation in vivo during embryonic development.

Consistent with previously published studies15, successful parabiotic fusion of zebrafish embryos depends on the staging and orientation of the two embryos&#160;and the concentration of methylcellulose. With just a few simple, inexpensive tools, surgically fused developing blastulae were generated that grew into parabiotic embryos with shared circulation. These tools included a modified Pasteur pipette, a 10 ml pipette pump, and wood handled teasing needles which were used eith...

Watch this Scientific Journal Video about Generation of Parabiotic Zebrafish Embryos by Surgical Fusion of Developing Blastulae at JoVE.com

Generation of Parabiotic Zebrafish Embryos by Surgical Fusion of Developing Blastulae

This protocol provides step-by-step instruction on how to generate parabiotic zebrafish embryos of different genetic backgrounds. When combined with the unparalleled imaging capabilities of the zebrafish embryo, this method provides a uniquely powerful means to investigate cell-autonomous versus non-cell-autonomous functions for candidate genes of interest.

Surgical parabiosis of two animals of different genetic backgrounds creates a unique scenario to study cell-intrinsic versus cell-extrinsic roles for ...

generation-parabiotic-zebrafish-embryos-surgical-fusion-developing

Research

JoVE Journal

Developmental Biology

11.5K Views.  Boston Children’s Hospital. This protocol provides step-by-step instruction on how to generate parabiotic zebrafish embryos of different genetic backgrounds. When combined with the unparalleled imaging capabilities of the zebrafish embryo, this method provides a uniquely powerful means to investigate cell-autonomous versus non-cell-autonomous functions for candidate genes of interest.

Video: Generation of Parabiotic Zebrafish Embryos by Surgical Fusion of Developing Blastulae

Microbead Implantation in the Zebrafish Embryo

The zebrafish has emerged as a valuable genetic model system for the study of developmental biology and disease. Zebrafish share a high degree of genomic conservation, as well as similarities in cellular, molecular, and physiological processes, with other vertebrates including humans. During early ontogeny, zebrafish embryos are optically transparent, allowing researchers to visualize the dynamics of organogenesis using a simple stereomicroscope. Microbead implantation is a method that enables tissue manipulation through the alteration of factors in local environments. This allows researchers to assay the effects of any number of signaling molecules of interest, such as secreted peptides, at specific spatial and temporal points within the developing embryo. Here, we detail a protocol for how to manipulate and implant beads during early zebrafish development.

The zebrafish is an excellent model system for genetic and developmental studies. Bead implantation is a valuable tissue manipulation technique that can be used to interrogate developmental mechanisms by introducing alterations in local cellular environments. This protocol describes how to perform microbead implantation in the zebrafish embryo.

The zebrafish has emerged as a valuable genetic model system for the study of developmental biology and disease. Zebrafish share a high degree of genomic ...

Generating Chimeric Zebrafish Embryos by Transplantation

One of the most powerful tools used to gain insight into complex developmental processes is the analysis of chimeric embryos. A chimera is defined as an organism that contains cells from more than one animal; mosaics are one type of chimera in which cells from more than one genotype are mixed, usually wild-type and mutant. In the zebrafish, chimeras can be readily made by transplantation of cells from a donor embryo into a host embryo at the appropriate embryonic stage. Labeled donor cells are generated by injection of a lineage marker, such as a fluorescent dye, into the one-cell stage embryo. Labeled donor cells are removed from donor embryos and introduced into unlabeled host embryos using an oil-controlled glass pipette mounted on either a compound or dissecting microscope. Donor cells can in some cases be targeted to a specific region or tissue of the developing blastula or gastrula stage host embryo by choosing a transplantation site in the host embryo based on well-established fate maps.

A step-by-step guide to generating targeted chimeric zebrafish embryos by transplantation at the blastula or gastrula stage.

One of the most powerful tools used to gain insight into complex developmental processes is the analysis of chimeric embryos. A chimera is defined as an ...

Biology

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 ...

Spinal Cord Transection in the Larval Zebrafish

Mammals fail in sensory and motor recovery following spinal cord injury due to lack of axonal regrowth below the level of injury as well as an inability to reinitiate spinal neurogenesis. However, some anamniotes including the zebrafish Danio rerio exhibit both sensory and functional recovery even after complete transection of the spinal cord. The adult zebrafish is an established model organism for studying regeneration following spinal cord injury, with sensory and motor recovery by 6&#160;weeks post-injury. To take advantage of in vivo analysis of the regenerative process available in the transparent larval zebrafish as well as genetic tools not accessible in the adult, we use the larval zebrafish to study regeneration after spinal cord transection. Here we demonstrate a method for reproducibly and verifiably transecting the larval spinal cord. After transection, our data shows sensory recovery beginning at 2&#160;days post-injury (dpi), with the C-bend movement detectable by 3 dpi and resumption of free swimming by 5 dpi. Thus we propose the larval zebrafish as a companion tool to the adult zebrafish for the study of recovery after spinal cord injury.

After spinal transection, adult zebrafish have functional recovery by six weeks post-injury. To take advantage of larval transparency and faster recovery, we present a method for transecting the larval spinal cord. After transection, we observe sensory recovery beginning at 2&#160;days post-injury, and C-bend movement by 3 days post-injury.

Mammals fail in sensory and motor recovery following spinal cord injury due to lack of axonal regrowth below the level of injury as well as an inability ...

Neuroscience

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 ...

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 ...

Generation of Na&#239;ve Blastoderm Explants from Zebrafish Embryos

Due to their optical clarity and rapid development, zebrafish embryos are an excellent system for examining cell behaviors and developmental processes. However, because of the complexity and redundancy of embryonic signals, it can be challenging to discern the complete role of any single signal during early embryogenesis. By explanting the animal region of the zebrafish blastoderm, relatively na&#239;ve clusters of embryonic cells are generated that can be easily cultured and manipulated ex vivo. By introducing a gene of interest by RNA injection before explantation, one can assess the effect of this molecule on gene expression, cell behaviors, and other developmental processes in relative isolation. Furthermore, cells from embryos of different genotypes or conditions can be combined in a single chimeric explant to examine cell/tissue interactions and tissue-specific gene functions. This article provides instructions for generating zebrafish blastoderm explants and demonstrates that a single signaling molecule - a Nodal ligand - is sufficient to induce germ layer formation and extension morphogenesis in otherwise na&#239;ve embryonic tissues. Due to their ability to recapitulate embryonic cell behaviors, morphogen gradients, and gene expression patterns in a simplified ex vivo system, these explants are anticipated to be of great utility to many zebrafish researchers.

Zebrafish blastoderm explants are generated by isolating embryonic cells from endogenous signaling centers within the early embryo, producing relatively na&#239;ve cell clusters easily manipulated and cultured ex vivo. This article provides instructions for making such explants and demonstrates their utility by interrogating roles for Nodal signaling during gastrulation.

Due to their optical clarity and rapid development, zebrafish embryos are an excellent system for examining cell behaviors and developmental processes. ...

Robust Zebrafish Cell Transplantation for Regeneration and Development Research

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.

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.

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

Production of Haploid Zebrafish Embryos by In Vitro Fertilization

The zebrafish has become a mainstream vertebrate model that is relevant for many disciplines of scientific study. Zebrafish are especially well suited for forward genetic analysis of developmental processes due to their external fertilization, embryonic size, rapid ontogeny, and optical clarity &#8211; a constellation of traits that enable the direct observation of events ranging from gastrulation to organogenesis with a basic stereomicroscope. Further, zebrafish embryos can survive for several days in the haploid state. The production of haploid embryos in vitro is a powerful tool for mutational analysis, as it enables the identification of recessive mutant alleles present in first generation (F1) female carriers following mutagenesis in the parental (P) generation. This approach eliminates the necessity to raise multiple generations (F2, F3, etc.) which involves breeding of mutant families, thus saving the researcher time along with reducing the needs for zebrafish colony space, labor, and the husbandry costs. Although zebrafish have been used to conduct forward screens for the past several decades, there has been a steady expansion of transgenic and genome editing tools. These tools now offer a plethora of ways to create nuanced assays for next generation screens that can be used to further dissect the gene regulatory networks that drive vertebrate ontogeny. Here, we describe how to prepare haploid zebrafish embryos. This protocol can be implemented for novel future haploid screens, such as in enhancer and suppressor screens, to address the mechanisms of development for a broad number of processes and tissues that form during early embryonic stages.

Production of Haploid Zebrafish Embryos by In Vitro Fertilization

The zebrafish is a powerful model system for developmental biology and human disease research due to their genetic similarity with higher vertebrates. This protocol describes a methodology to create haploid zebrafish embryos that can be utilized for forward screen strategies to identify recessive mutations in genes essential for early embryogenesis.

The zebrafish has become a mainstream vertebrate model that is relevant for many disciplines of scientific study. Zebrafish are especially well suited for ...

Zebrafish Breeding and Embryo Handling

Zebrafish (Danio rerio) are an important model organism that is particularly valuable for research in developmental biology. Zebrafish are extremely fertile and can produce hundreds of progeny per week, so it is relatively easy to collect a large number of embryos for high sample numbers. Furthermore, zebrafish undergo rapid development and embryos are transparent, allowing for easy visualization of developmental processes.
This video covers the steps required for the collection of newly fertilized zebrafish embryos. A brief overview of zebrafish mating behavior is presented, followed by instructions for setting up crosses in specialized laboratory breeding tanks that allow for controlled mating. Also covered are the conditions required to initiate the release of eggs (known as spawning) the morning after tanks are set. Next, essential techniques for working with embryos are presented, including the inhibition of pigment development with the chemical PTU, and dechorionation:&#160;a procedure in which the shell-like membrane surrounding the embryo (the chorion) is removed. Finally, the video concludes with some practical applications of these techniques in developmental research.

Zebrafish (Danio rerio) are an important model organism that is particularly valuable for research in developmental biology. Zebrafish are extremely ...

Generation of Parabiotic Zebrafish Embryos by Surgical Fusion of Developing Blastulae

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