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 border="0" cellpadding="0" cellspacing="0" width="686">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:433px;">Methyl cellulose</td>
			<td style="width:87px;">Sigma</td>
			<td style="width:167px;">M0387</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Individual components for E3/HCR:</td>
			<td></td>
			<td style="width:167px;">For 1L: 14,61g NaCl, 0,63g KCl, 2,43g CaCl2-2H2O, 1,99g MgSO4</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">NaCl (Sodium chloride)</td>
			<td style="width:87px;">Sigma-Aldrich</td>
			<td style="width:167px;">S9888</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">KCl (Potassium chloride)</td>
			<td>Sigma-Aldrich</td>
			<td style="width:167px;">P9541</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">CaCl2 (Calcium chloride dihydrate)</td>
			<td>Sigma-Aldrich</td>
			<td style="width:167px;">223506</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">MgSO4 (Magnessium sulfate heptahydrate)</td>
			<td>Sigma-Aldrich</td>
			<td style="width:167px;">230391</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Hepes (1M ) buffer solution</td>
			<td style="width:87px;">ThermoFisher</td>
			<td>15630-080</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:433px;">Name</td>
			<td style="width:87px;">Company</td>
			<td style="width:167px;">Catalog Number</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Antibiotics:</td>
			<td style="width:87px;"></td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Pen/Strep</td>
			<td style="width:87px;">gibco by Life technologies</td>
			<td>15140-120</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Ampicilin sodium salt</td>
			<td style="width:87px;">Sigma Life Science</td>
			<td style="width:167px;">A0166</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Kanamycin sulfate from Streptomyces kanamyceticus</td>
			<td style="width:87px;">Sigma Life Science</td>
			<td style="width:167px;">K1377</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:433px;">Name of Reagent/ Equipment</td>
			<td style="width:87px;">Company</td>
			<td style="width:167px;">Catalog Number</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">50 mg/ml Pronase from Streptomyces griseus</td>
			<td style="width:87px;">Roche</td>
			<td style="width:167px;">11459643001</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">capillary glass used for needles (Capillary Glass &#38; Filaments)</td>
			<td style="width:87px;">Sutter Instrument</td>
			<td style="width:167px;">ITEM#: BF 100-50-10</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Teasing needles with wooden handles</td>
			<td style="width:87px;">Fisher Scientific</td>
			<td>S07894</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Glass Pasteur pipettes</td>
			<td style="width:87px;">Fisherbrand</td>
			<td style="width:167px;">13-678-20A</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">10 mL pipette pump (green) (Pipette Pump Pipettor)</td>
			<td style="width:87px;">Novatech International</td>
			<td style="width:167px;">F37898-0000</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">100 mm diameter/ 20 mm deep plastic petri dishes (PETRI DISH, 100/20 MM, PS, CLEAR, WITH VENTS, 
			HEAVY DESIGN, 15 PCS./BAG )</td>
			<td>Greiner Bio-one</td>
			<td>664102</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Dextran, Cascade Blue, 10,000 MW, Anionic, Lysine Fixable</td>
			<td>ThermoFisher</td>
			<td>D-1976</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">PTU. working stock is 0.003% (50X is 0.15%). for 500ml, 0.75 g N-Phenylthiourea</td>
			<td style="width:87px;">Sigma-Aldrich</td>
			<td style="width:167px;">P7629</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Tricaine (powder) (Tricaine Methanesulfonato, Tricaine-S)</td>
			<td style="width:87px;">Western Chemical Inc</td>
			<td style="width:167px;">MS 222</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">LMP agarose (Ultrapure LMP agarose)</td>
			<td style="width:87px;">Invitrogen</td>
			<td style="width:167px;">16520100</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">plastic transfer pipette (just the wide ended one I think)</td>
			<td style="width:87px;">Fisherbrand</td>
			<td style="width:167px;">137115AM</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Glass-bottom 6-well plates used for imaging</td>
			<td style="width:87px;">MatTek</td>
			<td>P06G1.5-20-F</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">plastic western gel loading tip fixed on the end of a wood-handled dissecting needle (GELoader tips)</td>
			<td style="width:87px;">Eppendorf</td>
			<td style="width:167px;">22351656</td>
		</tr>
		<tr height="21">
			<td colspan="2" height="21" style="height:21px;">glass cover slips, slides and vacuum grease if mounting for an upright microscope:</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:433px;">Vaccum grease ( Dow Corning&#174; high-vacuum silicone grease 
			colorless, weight 5.3 oz (tube) )</td>
			<td style="width:87px;">Dow Corning</td>
			<td>Z273554</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:433px;">Glass cover slips</td>
			<td style="width:87px;">Corning Life Sciences</td>
			<td style="width:167px;">2960-244</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

11.4K 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

Mechanical Vessel Injury in Zebrafish Embryos

Zebrafish (Danio rerio) embryos have proven to be a powerful model for studying a variety of developmental and disease processes. External development and optical transparency make these embryos especially amenable to microscopy, and numerous transgenic lines that label specific cell types with fluorescent proteins are available, making the zebrafish embryo an ideal system for visualizing the interaction of vascular, hematopoietic, and other cell types during injury and repair in vivo. Forward and reverse genetics in zebrafish are well developed, and pharmacological manipulation is possible. We describe a mechanical vascular injury model using micromanipulation techniques that exploits several of these features to study responses to vascular injury including hemostasis and blood vessel repair. Using a combination of video and timelapse microscopy, we demonstrate that this method of vascular injury results in measurable and reproducible responses during hemostasis and wound repair. This method provides a system for studying vascular injury and repair in detail in a whole animal model.

This article describes a method for creating a mechanical vessel injury in zebrafish embryos. This injury model provides a platform for studying hemostasis, injury-related inflammation, and wound healing in an organism ideally suited for real-time microscopy.

Zebrafish (Danio rerio) embryos have proven to be a powerful model for studying a variety of developmental and disease processes.  External development ...

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 ...

Affinity Labeling Detection of Endogenous Receptors from Zebrafish Embryos

By combining the powers of Affinity Labeling and Immunoprecipitation (AFLIP), a technique for the detection of low abundance receptors in zebrafish embryos has been implemented. This technique takes advantage of the selectivity and sensitivity conferred by affinity labeling of a given receptor by its ligand with the specificity of the immunoprecipitation. We have used AFLIP to detect the type III TGF-&#946; receptor (TGFBR3), also know as betaglycan, during early zebrafish development. AFLIP was instrumental in validating the efficacy of a TGFBR3 morphant zebrafish phenotype. In the first step, embryo protein extracts are prepared and used to generate 125I-TGF-&#946;2-TGFBR3 complexes that are purified by immunoprecipitation. Later, these complexes are covalently cross-linked and revealed using SDS-PAGE separation and autoradiography detection. This technique requires the availability of a labeled ligand for, and a specific antibody against, the receptor to be detected, and shall be easily adapted to identify any growth factor or cytokine receptor that meets these requirements.

A novel technique for the detection of low abundance endogenous receptors present in zebrafish embryos is described. We have named it AFLIP because it consists of affinity labeling of the receptor by its ligand linked to immunoprecipitation.

By combining the powers of Affinity Labeling and Immunoprecipitation (AFLIP), a technique for the detection of low abundance receptors in zebrafish ...

Ploidy Manipulation of Zebrafish Embryos with Heat Shock 2 Treatment

Manipulation of ploidy allows for useful transformations, such as diploids to tetraploids, or haploids to diploids. In the zebrafish Danio rerio, specifically the generation of homozygous gynogenetic diploids is useful in genetic analysis because it allows the direct production of homozygotes from a single heterozygous mother. This article describes a modified protocol for ploidy duplication based on a heat pulse during the first cell cycle, Heat Shock 2 (HS2). Through inhibition of centriole duplication, this method results in a precise cell division stall during the second cell cycle. The precise one-cycle division stall, coupled to unaffected DNA duplication, results in whole genome duplication. Protocols associated with this method include egg and sperm collection, UV treatment of sperm, in vitro fertilization and heat pulse to cause a one-cell cycle division delay and ploidy duplication. A modified version of this protocol could be applied to induce ploidy changes in other animal species.

A modified protocol for ploidy manipulation uses a heat shock to induce a one-cycle stall in cytokinesis in the early embryo. This protocol is demonstrated in the zebrafish but may be applicable to other species.

Manipulation of ploidy allows for useful transformations, such as diploids to tetraploids, or haploids to diploids. In the zebrafish Danio rerio, ...

Biosensing Motor Neuron Membrane Potential in Live Zebrafish Embryos

The protocols described here are designed to allow researchers to study cell communication without altering the integrity of the environment in which the cells are located. Specifically, they have been developed to analyze the electrical activity of excitable cells, such as spinal neurons. In such a scenario, it is crucial to preserve the integrity of the spinal cell, but it is also important to preserve the anatomy and physiological shape of the systems involved. Indeed, the comprehension of the manner in which the nervous system-and other complex systems-works must be based on a systemic approach. For this reason, the live zebrafish embryo was chosen as a model system, and the spinal neuron membrane voltage changes were evaluated without interfering with the physiological conditions of the embryos. 
Here, an approach combining the employment of zebrafish embryos with a FRET-based biosensor is described. Zebrafish embryos are characterized by a very simplified nervous system and are particularly suited for imaging applications thanks to their transparency, allowing for the employment of fluorescence-based voltage indicators at the plasma membrane during zebrafish development. The synergy between these two components makes it possible to analyze the electrical activity of the cells in intact living organisms, without perturbing the physiological state. Finally, this non-invasive approach can co-exist with other analyses (e.g., spontaneous movement recordings, as shown here).

Protocols described here allow for the study of the electrical properties of excitable cells in the most non-invasive physiological conditions by employing zebrafish embryos in an in vivo system together with a fluorescence resonance energy transfer (FRET)-based genetically encoded voltage indicator (GEVI) selectively expressed in the cell type of interest.

The protocols described here are designed to allow researchers to study cell communication without altering the integrity of the environment in which the ...

Induction of Hypoxia in Living Frog and Zebrafish Embryos

Here, we introduce a novel system for hypoxia induction, which we developed to study the effects of hypoxia in aquatic organisms such as frog and zebrafish embryos. Our system comprises a chamber featuring a simple setup that is nevertheless robust to induce and maintain a specific oxygen concentration and temperature in any experimental solution of choice. The presented system is very cost-effective but highly functional, it allows induction and sustainment of hypoxia for direct experiments in vivo and for various time periods up to 48 h.
To monitor and study the effects of hypoxia, we have employed two methods - measurement of levels of hypoxia-inducible factor 1alpha (HIF-1&#945;) in whole embryos or specific tissues and determination of retinal stem cell proliferation by 5-ethynyl-2&#8242;-deoxyuridine (EdU) incorporation into the DNA. HIF-1&#945; levels can serve as a general hypoxia marker in the whole embryo or tissue of choice, here embryonic retina. EdU incorporation into the proliferating cells of embryonic retina is a specific output of hypoxia induction. Thus, we have shown that hypoxic embryonic retinal progenitors decrease proliferation within 1 h of incubation under 5% oxygen of both frog and zebrafish embryos.
Once mastered, our setup can be employed for use with small aquatic model organisms, for direct in vivo experiments, any given time period and under normal, hypoxic or hyperoxic oxygen concentration or under any other given gas mixture.

We introduce a novel hypoxic chamber system for use with aquatic organisms such as frog and zebrafish embryos. Our system is simple, robust, cost-effective and allows the induction and sustainment of hypoxia in vivo and for up to 48 h. We present 2 reproducible methods to monitor the effectiveness of hypoxia.

Here, we introduce a novel system for hypoxia induction, which we developed to study the effects of hypoxia in aquatic organisms such as frog and ...

Development of an In Vitro Assay to Quantitate Hematopoietic Stem and Progenitor Cells (HSPCs) in Developing Zebrafish Embryos

Hematopoiesis is an essential cellular process in which hematopoietic stem and progenitor cells (HSPCs) differentiate into the multitude of different cell lineages that comprise mature blood. Isolation and identification of these HSPCs is difficult because they are defined ex post facto; they can only be defined after their differentiation into specific cell lineages. Over the past few decades, the zebrafish (Danio rerio) has become a model organism to study hematopoiesis. Zebrafish embryos develop ex utero, and by 48 h post-fertilization (hpf) have generated definitive HSPCs. Assays to assess HSPC differentiation and proliferation capabilities have been developed, utilizing transplantation and subsequent reconstitution of the hematopoietic system in addition to visualizing specialized transgenic lines with confocal microscopy. However, these assays are cost prohibitive, technically difficult, and time consuming for many laboratories. Development of an in vitro model to assess HSPCs would be cost effective, quicker, and present fewer difficulties compared to previously described methods, allowing laboratories to quickly assess mutagenesis and drug screens that affect HSPC biology. This novel in vitro assay to assess HSPCs is performed by plating dissociated whole zebrafish embryos and adding exogenous factors that promote only HSPC differentiation and proliferation. Embryos are dissociated into single cells and plated with HSPC-supportive colony stimulating factors that cause them to generate colony forming units (CFUs) that arise from a single progenitor cell. These assays should allow more careful examination of the molecular pathways responsible for HSPC proliferation, differentiation, and regulation, which will allow researchers to understand the underpinnings of vertebrate hematopoiesis and its dysregulation during disease.

Development of an In Vitro Assay to Quantitate Hematopoietic Stem and Progenitor Cells HSPCs in Developing Zebrafish Embryos

Here, we present a simple method to quantitate hematopoietic stem and progenitor cells (HSPCs) in embryonic zebrafish. HSPCs from dissociated zebrafish are plated in methylcellulose with supportive factors, differentiating into mature blood. This allows the detection of blood defects and allows drug screening to be easily conducted.

Hematopoiesis is an essential cellular process in which hematopoietic stem and progenitor cells (HSPCs) differentiate into the multitude of different cell ...

Visualization of Cellular Electrical Activity in Zebrafish Early Embryos and Tumors

Bioelectricity, endogenous electrical signaling mediated by ion channels and pumps located on the cell membrane, plays important roles in signaling processes of excitable neuronal and muscular cells and many other biological processes, such as embryonic developmental patterning. However, there is a need for in vivo electrical activity monitoring in vertebrate embryogenesis. The advances of genetically encoded fluorescent voltage indicators (GEVIs) have made it possible to provide a solution for this challenge. Here, we describe how to create a transgenic voltage indicator zebrafish using the established voltage indicator, ASAP1 (Accelerated Sensor of Action Potentials 1), as an example. The Tol2 kit and a ubiquitous zebrafish promoter, ubi, were chosen in this study. We also explain&#160;the processes of Gateway site-specific cloning, Tol2 transposon-based zebrafish transgenesis, and the imaging process for early-stage fish embryos and fish tumors using regular epifluorescent microscopes. Using this fish line, we found that there are cellular electric voltage changes during zebrafish embryogenesis, and fish larval movement. Furthermore, it was observed that in a few zebrafish malignant peripheral nerve sheath tumors, the tumor cells were generally polarized compared to the surrounding normal tissues.

Here, we show the process of creating a cellular electric voltage reporter zebrafish line to visualize embryonic development, movement, and fish tumor cells in vivo.

Bioelectricity, endogenous electrical signaling mediated by ion channels and pumps located on the cell membrane, plays important roles in signaling ...

Surgical Size Reduction of Zebrafish for the Study of Embryonic Pattern Scaling

In the developmental process, embryos exhibit a remarkable ability to match their body pattern to their body size; their body proportion is maintained even in embryos that are larger or smaller, within certain limits. Although this phenomenon of scaling has attracted attention for over a century, understanding the underlying mechanisms has been limited, owing in part to a lack of quantitative description of developmental dynamics in embryos of varied sizes. To overcome this limitation, we developed a new technique to surgically reduce the size of zebrafish embryos, which have great advantages for in vivo live imaging. We demonstrate that after balanced removal of cells and yolk at the blastula stage in separate steps, embryos can quickly recover under the right conditions and develop into smaller but otherwise normal embryos. Since this technique does not require special equipment, it is easily adaptable, and can be used to study a wide range of scaling problems, including robustness of morphogen mediated patterning.&#160;

Here, we describe a method for reducing the size of zebrafish embryos without disrupting normal developmental processes. This technique enables the study of pattern scaling and developmental robustness against size change.

In the developmental process, embryos exhibit a remarkable ability to match their body pattern to their body size; their body proportion is maintained ...

Whole Mount Immunohistochemistry in Zebrafish Embryos and Larvae

Immunohistochemistry is a widely used technique to explore protein expression and localization during both normal developmental and disease states. Although many immunohistochemistry protocols have been optimized for mammalian tissue and tissue sections, these protocols often require modification and optimization for non-mammalian model organisms. Zebrafish are increasingly used as a model system in basic, biomedical, and translational research to investigate the molecular, genetic, and cell biological mechanisms of developmental processes. Zebrafish offer many advantages as a model system but also require modified techniques for optimal protein detection. Here, we provide our protocol for whole-mount fluorescence immunohistochemistry in zebrafish embryos and larvae. This protocol additionally describes several different mounting strategies that can be employed and an overview of the advantages and disadvantages each strategy provides. We also describe modifications to this protocol to allow detection of chromogenic substrates in whole mount tissue and fluorescence detection in sectioned larval tissue. This protocol is broadly applicable to the study of many developmental stages and embryonic structures.

Here, we present a protocol for fluorescent antibody-mediated detection of proteins in whole preparations of zebrafish embryos and larvae.

Immunohistochemistry is a widely used technique to explore protein expression and localization during both normal developmental and disease states. ...