We thank the AECOM Zebrafish Core Facility and Cincinnati Children&#39;s Veterinary Services for fish maintenance, the Cincinnati Children&#39;s Imaging Core for technical assistance, Didar Saparov for assistance with video production and Hannah Seawall for editing the manuscript. Research reported in this publication was supported by the National Institute&#160;of General Medical Sciences of the National Institutes of Health under Award Number R35GM140805 to E.M.&#214;. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

This protocol involves use of live vertebrate embryos younger than 1 day post-fertilization. All the animal experiments were performed under the ethical guidelines of Cincinnati Children&#39;s Hospital Medical Center; animal protocols were reviewed and approved by the Institutional Animal Care and Use Committee (Protocol # 2017-0048).
1. Embryo collection
<ol>
	<li>Set up pairs of zebrafish in crossing tanks the night before the embryo collection day. For precise staging control of embryo development, use barriers between mating pairs.</li>
	<li>Raise barriers before preferred spawning time and collect eggs within 15 minutes in a 100 mm Petri dish.
	<ol>
		<li>Clean debris from the Petri dish. If more than 50 embryos are collected from a single clutch, split the clutch into multiple Petri dishes accordingly.</li>
	</ol>
	</li>
	<li>Incubate embryos in fish system water at 28 &#176;C until they reach 50% epiboly stage (5 hours post-fertilization). A standardized embryo growth medium such as E3 can also be used instead of the aquarium system water until step 3.2.
	<ol>
		<li>Remove unfertilized eggs under a stereoscope and move the embryos to a 23.5 &#176;C incubator overnight (O/N). Embryos should be at 8-10 somites stage the morning following collection day.</li>
	</ol>
	</li>
</ol>
2. Tool preparation
<ol>
	<li>Sterilize the microsurgery knife blade, needle tips (used for dissection of the tissue), and glass Pasteur pipette by soaking in 100% ethanol (EtOH) and fire glazing.</li>
	<li>Use two layers of transparent tape (~100-120 &#181;m thickness) on 25 mm x 75 mm microscope slides. Cut ~18 mm x 18 mm square wells in the center of each slide&#39;s tape covering with a scalpel.
	<ol>
		<li>Wipe the prepared slide chambers with 70% EtOH. These wells will hold ~40 &#181;L of medium.</li>
	</ol>
	</li>
</ol>
3. Sample preparation
<ol>
	<li>Dechorionate embryos using the tip of two needle syringes under a stereoscope. Transfer embryos into a separate Petri dish with fish system water to rinse.</li>
	<li>Using a fire-glazed sterile glass Pasteur pipette, transfer embryos in a 6 cm Petri dish containing dissection medium (Leibovitz-15 cell culture medium with L-Glutamine without Phenol Red, 0.8 mM CaCl2 and 1&#215; antibiotic-antimycotic solution). 
	NOTE: Continue to use a sterilized glass pipette for all transfers following this step.
	<ol>
		<li>Use a glass Petri dish for explanting procedure to avoid polystyrene chips during dissection.</li>
	</ol>
	</li>
	<li>Put 50 &#181;L of tissue growth medium (dissection medium, and 10% FBS) in the slide chamber.</li>
	<li>Stabilize an embryo for dissection under the stereoscope with a needle tip at the yolk-tissue intersection near the hindbrain.</li>
	<li>Keeping the embryonic tissue stable with a needle, use the microsurgical knife with the blade held at 45&#176; to cut the tissue anterior to the hindbrain apart and peel the yolk off the embryonic tissue starting from the anterior and moving towards the tail bud (Figure 1A). 
	NOTE: Be careful to not lose the skin tissue while cleaning the yolk. The skin would easily peel off as a flanking single layer elastic tissue around the embryo during the dissection, so it is easy to recognize.</li>
	<li>Once the yolk is fully removed from the embryonic body, cut the flanking skin tissue from the tailbud. Keeping the last formed 3-4 somites intact, cut the more anterior tissue out (full-axis explant).
	<ol>
		<li>The yolk should come off mainly intact from this procedure. In case of a ruptured yolk, significant granules of yolk can stay attached to the ventral surface of the tissue. If so, use a lash tool to clean off remaining yolk granules gently. 
		NOTE: An imbalance of skin tissues on the lateral sides of an explant would not allow the tissue to maintain a straight growth direction. The explant instead will bend towards the side of more stretched skin. This imbalance can be corrected under the stereoscope by rupturing the skin layer with the aid of microsurgical knife.</li>
		<li>For skinless explants, press on a tip of the skin layer with needle and peel the tissue explant off with the microsurgical knife. These explants will not elongate their body axis in culture.</li>
		<li>In addition to the full axis explants, alternative explants can be made at this step. For instance, dissect out already segmented somites using the microsurgical knife (full-PSM explants) or dissect the PSM into its half anteroposterior (half-PSM explants). Please see section 5.1 for an application of such alternative explants.</li>
	</ol>
	</li>
	<li>Immediately transfer the dissected explant to a 22 mm x 22 mm coverslip on which the imaging will be performed.
	<ol>
		<li>Arrange the explant flat on the dorsoventral axis, ventral side touching the coverslip (Figure 1B). Gently remove the excess media around the tissue explant using a 20 &#181;L filtered tip pipette. 
		NOTE: Delayed transfer of dissected explants to the coverslip results in deformations of the tissue, as it is relieved from the geometric constraints of the yolk.</li>
	</ol>
	</li>
	<li>Swiftly and carefully flip the coverslip with the explant over the growth medium filled slide chamber.
	<ol>
		<li>To prevent bubble formation, place a side of the square coverslip on the tape chamber and release the other side gently. Take care to not move/deform the explant in this step.</li>
		<li>Gently remove excess media bleeding out of the chamber by pressing the slide chamber on a laboratory tissue. The coverslip will sit stably on the slide chamber for live imaging, due to the surface tension of the liquid medium without any sealing.</li>
		<li>For long term culturing (&#62;6 hours), use a bigger chamber. 22 mm x 50 mm rectangular coverslips together with two parallel lanes of tape layers on slides can be utilized in such cases. A ~1 mm wide gap can be left in between two tape lanes to facilitate air access into the growth medium.</li>
	</ol>
	</li>
	<li>Repeat steps 3.3-3.8 to prepare more explants. Prepared explants will elongate their A-P body axis with an average of ~30 &#181;m/h speed and segment their somites with ~40 min intervals at 25 &#176;C (Figure 2A, Video 1).
	<ol>
		<li>For non-elongating explants, apply gentle pressure to the sides of slide holding the sample in step 3.8.2 while sucking the excess media out on a laboratory tissue. Alternatively, explants can be cultured in single tape layer slide chambers. Also, chemically activating the slide chamber surface with Type I Collagen will result in non-elongating explants (Figure 2B, Video 2).
		<ol>
			<li>Perform coating of the chamber with Type I Collagen in advance by fully covering the slide chambers with 15-20 mL of prediluted collagen solution at room temperature for 1 h. Use a laminar flow hood for this protocol to maintain sterility. Carefully rinse chambers with dissection medium at the end.</li>
		</ol>
		</li>
		<li>For embryos older than 15 somite stage, mount the tail explant tissue laterally instead of a flat (dorsoventral) mount (Video 3). To prevent muscular twitching, include 0.004% tricaine solution in the culture media as an anesthetic agent3.</li>
	</ol>
	</li>
</ol>
4. Live image acquisition
<ol>
	<li>Image samples either on a dissection scope for widefield transmitted light imaging of somite segmentation sizes and periods, or with structured illumination/confocal/light sheet microscopy using transgenic reporter fish lines.</li>
	<li>Equilibrate the temperature of tissue explants with the imaging room temperature for at least 15 min.
	<ol>
		<li>For more precise temperature control, use a commercial temperature control system mounted on an inverted microscope.</li>
	</ol>
	</li>
	<li>Set the image acquisition frame intervals to 2 - 10 min depending on the biological process of interest. 
	NOTE: Zebrafish somite segmentation is a fast process, ranging between 20 - 55 min for viable temperatures of 30 &#176;C to 21.5 &#176;C in whole embryos. Explants will elongate and segment ~30% slower than the whole embryos.
	<ol>
		<li>Pay attention to leave enough delay between sets of channel acquisitions to avoid possible phototoxicity to live tissue. Do not expose the tissue to the excitation beam for more than half of the imaging duration and lower the beam intensity as much as possible. 
		NOTE: Accumulation of reactive oxygen species (ROS) is generally the major cause of phototoxicity in live samples4. Ascorbic acid as a ROS scavenger can be supplemented to growth medium at 4 mM concentration to buffer ROS activity and alleviate phototoxicity. Adverse effects of phototoxicity might be hard to notice during live imaging. Tail explants are advantageous in this aspect since some visual markers of phototoxicity such as mitotic arrest, impeded tissue development (i.e., formation of somites, tail elongation), and disintegrating tissue are easier to notice. Please refer to the provided reference4 for a detailed discussion.</li>
	</ol>
	</li>
	<li>Use single cell-stage RNA injected embryos to acquire 4-D images intended to be segmented and analyzed in cellular resolution level.
	<ol>
		<li>Use 300 pg of RNA from in vitro transcribed membranes and nuclear fluorescent reporter marker plasmids such as pCS-membrane-ceruleanFP (Addgene plasmid #53749) or pCS-memb-mCherry (Addgene plasmid #53750) in combination with pCS2+ H2B-mTagBFP2 (Addgene plasmid #99267) or pCS2+ H2B-TagRFP-T (Addgene plasmid #99271) in injections. For a sample movie with cell membrane and nuclei markers please see Video 4. 
		&#8203;NOTE: Average cell size of PSM tissue is about ~5 &#181;m in diameter, of which nuclei comprises ~3 - 4 &#181;m. A pixel size of ~0.5 &#181;m and a z-sectioning of ~1 &#181;m should be recorded for proper cell segmentation.</li>
	</ol>
	</li>
</ol>
5. Immunostaining of tail explants
NOTE: Tissues grown after various dissection scenarios (elongating, non-elongating, tail bud dissected, half PSM etc.) as flat-mounted tail explants5 can be recovered from slide chambers for further immunostaining quantifications of proteins of interest. Here, we present the protocol used for di-phosphorylated extracellular signal regulated kinase (ppERK) staining of explants as FGF signaling gradient readout.
<ol>
	<li>After formation of somites until the desired stage, cautiously shift the coverslip halfway to the corner of the slide chamber without lifting.</li>
	<li>With the help of ~100 &#181;L of supplementary dissection medium in a glass Pasteur pipette, recover the explants from the slide and transfer into a 64-well cell culture plate. 
	NOTE: Beginning with this step, all solution replacements can be performed under a dissection scope with a separate glass pipette for fixed samples. This will ensure not losing explant tissues in wells or transferring them in between.</li>
	<li>After transferring all explants, suck the excessive medium out of the wells one-by-one and put 100 &#181;L of 4% paraformaldehyde in PBS (PFA) into each well. 
	CAUTION: PFA is a toxic solution with carcinogenic effects. Proper PPE should be used while handling.</li>
	<li>Fix explants in a 64-well plate at room temperature for 1 h on a shaker.
	<ol>
		<li>Tissue explants are more sensitive to deformations than whole embryos. Adjust the shaker speed accordingly.</li>
	</ol>
	</li>
	<li>Wash the fixative out with 150 &#181;L of PBS-Tw (0.1% Tween20 in PBS) three times. Collect the first wash in a specific &#34;PFA Waste&#34; container.</li>
	<li>Dehydrate explants in 4&#215;5 min steps by replacing ~40 &#181;L of solution each time with 100% methanol (MeOH). 
	CAUTION: MeOH is a toxic chemical which is volatile and flammable. Work in a well-ventilated space and use proper PPE for handling.</li>
	<li>As the last step of dehydration, remove all solution from wells and replace with 100 &#181;L of MeOH. Incubate at -20 &#176;C for 15 min. 
	NOTE: Use a specific &#34;MeOH Waste&#34; container to collect the solutions until step 5.11.</li>
	<li>Add 50 &#181;L of MeOH and shake at room temperature for 5 min.</li>
	<li>Rehydrate explants in 4&#215;5 min steps by replacing ~40 &#181;L of solution each time with PBS-T (0.1% Triton-X 100 in PBS). Use a specific &#34;MeOH Waste&#34; container to collect the solutions.</li>
	<li>As the last step of rehydration, remove all solution from wells and replace with 100 &#181;L of PBS-T.</li>
	<li>For tissue permeabilization treat explants with 1.5% Triton-X 100 in PBS for 20 min at room temperature on the shaker.</li>
	<li>Wash samples with MAB-D-T (0.1% Triton-X 100 detergent and 1% dimethyl sulfoxide (DMSO) in 150 mM NaCl 100 mM maleic acid buffer pH 7.5) 3&#215;5 min. 
	CAUTION: DMSO is flammable and a toxic mutagen. Proper PPE should be used while handling.</li>
	<li>Incubate explants in 100 &#181;L/well serum blocking solution (2% fetal bovine serum in MAB-D-T) for 2 hours at room temperature.</li>
	<li>Replace all the blocking solution with 50-100 &#181;L/well primary antibody solution (1:1000 dilution of monoclonal mouse antibody against ppERK in serum block). Incubate samples O/N (&#62;16 h) at 4 &#176;C on shaker.</li>
	<li>Wash the primary antibody solution with MAB-D-T 5&#215;5 min.</li>
	<li>Incubate samples in secondary antibody solution (Alexa Fluor 597 goat anti-mouse IgG2b (1:200) and Hoechst 33342 (1:5000) in MAB-D-T) O/N at 4 &#176;C on a shaker or for 3 h at room temperature. 
	NOTE: Beginning at this step, cover the 64-well plate with aluminum foil to avoid light exposure of secondary antibody treated samples. 
	CAUTION: Hoechst 33342 is a potential carcinogen. Proper PPE should be used while handling.</li>
	<li>Wash the secondary antibody solution with PBS-Tw 3&#215;5 min.</li>
	<li>Fix samples with PFA for 15 minutes at room temperature.</li>
	<li>Wash the fixative with PBS-Tw and equilibrate samples within 60% glycerol. Mount explants on microscope slides with nail polish and 60% glycerol for imaging. For representative immunostaining results please see Figure 3.</li>
</ol>

<ol>
	<li>Assheton, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Growth in length: Embryological Essays. , (1916).</li><li>Gomez, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Control+of+segment+number+in+vertebrate+embryos.">Control of segment number in vertebrate embryos.</a> Nature. 454 (7202), 335-339 (2008).</li><li>Westerfield, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The Zebrafish Book: a guide for the laboratory use of zebrafish (Danio rerio), 3rd edition. , (1995).</li><li>Icha, J., Weber, M., Waters, J. C., Norden, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Phototoxicity+in+live+fluorescence+microscopy,+and+how+to+avoid+it.">Phototoxicity in live fluorescence microscopy, and how to avoid it.</a> BioEssays. 39 (1700003), (2017).</li><li>Simsek, M. F., Ozbudak, E. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spatial+fold+change+of+Fgf+signaling+encodes+positional+information+for+segmental+determination+in+zebrafish.">Spatial fold change of Fgf signaling encodes positional information for segmental determination in zebrafish.</a> Cell Reports. 24 (1), 66-78 (2018).</li><li>Dubrulle, J., Pourqui&#233;, O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=fgf8+mRNA+decay+establishes+a+gradient+that+couples+axial+elongation+to+patterning+in+the+vertebrate+embryo.">fgf8 mRNA decay establishes a gradient that couples axial elongation to patterning in the vertebrate embryo.</a> Nature. 427 (6973), 419-422 (2004).</li><li>Diez del Corral, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Opposing+FGF+and+Retinoid+Pathways+Control+Ventral+Neural+Pattern,+Neuronal+Differentiation,+and+Segmentation+during+Body+Axis+Extension.">Opposing FGF and Retinoid Pathways Control Ventral Neural Pattern, Neuronal Differentiation, and Segmentation during Body Axis Extension.</a> Neuron. 40 (1), 65-79 (2003).</li><li>Stern, H. M., Hauschka, S. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neural+tube+and+notochord+promote+in+vitro+myogenesis+in+single+somite+explants.">Neural tube and notochord promote in vitro myogenesis in single somite explants.</a> Developmental Biology. 167 (1), 87-103 (1995).</li><li>Langenberg, T., Brand, M., Cooper, M. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+brain+development+and+organogenesis+in+zebrafish+using+immobilized+embryonic+explants.">Imaging brain development and organogenesis in zebrafish using immobilized embryonic explants.</a> Developmental Dynamics. 228 (3), 464-474 (2003).</li><li>Picker, A., Roellig, D., Pourqui&#233;, O., Oates, A. C., Brand, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tissue+micromanipulation+in+zebrafish+embryos.">Tissue micromanipulation in zebrafish embryos.</a> Methods in molecular biology. 546 (11), 153-172 (2009).</li><li>Manning, A. J., Kimelman, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tbx16+and+Msgn1+are+required+to+establish+directional+cell+migration+of+zebrafish+mesodermal+progenitors.">Tbx16 and Msgn1 are required to establish directional cell migration of zebrafish mesodermal progenitors.</a> Developmental Biology. 406 (2), 172-185 (2015).</li><li>Kimmel, C. B., Ballard, W. W., Kimmel, S. R., Ullmann, B., Schilling, T. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stages+of+embryonic+development+of+the+zebrafish.">Stages of embryonic development of the zebrafish.</a> Developmental Dynamics. 203 (3), 253-310 (1995).</li><li>Kaufmann, A., Mickoleit, M., Weber, M., Huisken, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multilayer+mounting+enables+long-term+imaging+of+zebrafish+development+in+a+light+sheet+microscope.">Multilayer mounting enables long-term imaging of zebrafish development in a light sheet microscope.</a> Development. 139, 3242-3247 (2012).</li></ol>

The authors have nothing to disclose and declare no conflicts of interest.

This article presents a detailed protocol of a tissue culture explant technique we developed and used recently5 for zebrafish embryos. Our technique builds on the previous explant methods in chick8 and zebrafish9,10,11 model organisms. Tail explants prepared with this protocol can survive as long as &gt;12 h in a simple slide chamber, continuing to elongate its major body axis and ...

Metameric segmentation of organisms is widely used in nature. Repeated structures are essential for functionality of lateral organs such as vertebrae, muscles, nerves, vessels, limbs, or leaves in a body plan1. As a result of such physiological and geometric constraints of the axial symmetry, most phyla of Bilateria-such as annelids, arthropods, and chordates-exhibit segmentation of their embryonic tissues (e.g., ectoderm, mesoderm) antero-posteriorly.
Vertebrate embryos sequentially segment their paraxial mesoderm along the major body axis into somites with species-specific intervals, counts, and size distributions. Despite such robustness among individual embryos within a species, somite segmentation is versatile in between vertebrate species. Segmentation happens in a vast regime of time intervals (from 25 min in zebrafish to 5 h in humans), sizes (from ~20 &#181;m in tail somites of zebrafish to ~200 &#181;m in trunk somites of mice) and counts (from 32 in zebrafish to ~300 in corn snakes)2. More interestingly, fish embryos can develop in a wide range of temperatures (from ~20.5 &#176;C up to 34 &#176;C for zebrafish) while keeping their somites intact with proper size distributions by compensating for both segmentation intervals and axial elongation speeds. Beyond such interesting features, zebrafish stays as a useful model organism to study segmentation in vertebrates due to the external, synchronous and transparent development of a plenitude of sibling embryos as well as their accessible genetic tools. Adversely from a microscopy perspective, teleost embryos develop on a bulky spherical yolk, stretching and rounding the gastrulating tissue around it (Figure 1A). In this article, we present a flattened 3-D tissue explant culture for zebrafish tails. This explant system circumvents the spherical constraints of yolk mass, allowing access to high resolution live imaging of fish embryos for somite patterning.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61981/61981fig01.jpg" /> 
Figure 1: Slide Chamber Explant System for Zebrafish Embryos. (A) Zebrafish embryos have advantages for live imaging, such as the transparency of gastrulating embryonic tissue (blue), but the tissue forms around a bulky spherical yolk mass (yellow) which prevents near-objective, high-resolution imaging in intact embryos. Tail explants can be dissected starting with a microsurgical knife (brown) cut from the tissue anterior of somites (red) and continuing at the border with the yolk posteriorly. (B) Dissected tail explants can be placed on a coverslip (light blue) dorsoventrally; keeping neural tissue (light gray) on top and notochord (dark gray) at the bottom. <a href="https://www.jove.com/files/ftp_upload/61981/61981fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table><tbody><tr><td>1 mL Sub-Q Syringe with PrecisionGlide Needle</td><td>Becton, Dickinson and Co.</td><td>REF 309597</td><td>for dechorionating embryos and manipulations</td></tr><tr><td>200 Proof Ethanol, Anhydrous</td><td>Decon Labs</td><td>2701</td><td>for immunostaining</td></tr><tr><td>Antibiotic Antimycotic Solution (100&amp;times;)</td><td>Sigma-Aldrich</td><td>A5955</td><td>for tissue dissection media</td></tr><tr><td>Calcium Chloride Anhydrous, Powder</td><td>Sigma-Aldrich</td><td>499609</td><td>for tissue dissection media</td></tr><tr><td>Dimethylsulfoxide</td><td>Sigma-Aldrich</td><td>D5879</td><td>for immunostaining</td></tr><tr><td>Disposable Scalpel, #10 Stainless Steel</td><td>Integra-Miltex</td><td>MIL4-411</td><td>for preparing tape slide wells</td></tr><tr><td>Ethyl 3-aminobenzoate methanesulfonate salt (Tricaine)</td><td>Sigma-Aldrich</td><td>886-86-2</td><td>(optional) for anesthesizing tissues older than 20 somites stage</td></tr><tr><td>Fetal Bovine Serum (FBS)</td><td>ThermoFisher</td><td>A3160601</td><td>additional for tissue culture media</td></tr><tr><td>Goat anti-Mouse IgG2b, Alexa Fluor 594</td><td>Invitrogen</td><td>Cat#A-21145; RRID: AB_2535781</td><td>secondary antibody for immunostaining</td></tr><tr><td>L-15 Medium with L-Glutamine w/o Phenol Red</td><td>GIBCO</td><td>21083-027</td><td>for tissue dissection media</td></tr><tr><td>Methanol</td><td>Sigma-Aldrich</td><td>179337</td><td>for immunostaining</td></tr><tr><td>Microsurgical Corneal Knife 2.85 mm Angled Tip Double Bevel Blade</td><td>Surgical Specialties</td><td>72-2863</td><td>for tissue dissection</td></tr><tr><td>Mouse monoclonal anti-ppERK</td><td>Sigma-Aldrich</td><td>Cat#M8159; RRID:AB_477245</td><td>for ppERK immunostaining</td></tr><tr><td>NucRed Live 647 ReadyProbes Reagent</td><td>Invitrogen</td><td>R37106</td><td>(optional) for live staining of cell nuclei</td></tr><tr><td>Paraformaldehyde Powder, 95%</td><td>Sigma-Aldrich</td><td>158127</td><td>for fixation of samples for immunostaining</td></tr><tr><td>Rat Tail Collagen Coating Solution</td><td>Sigma-Aldrich</td><td>122-20</td><td>(optional) for chemically activating slide chambers</td></tr><tr><td>Stage Top Incubator</td><td>Tokai Hit</td><td>tokai-hit-stxg</td><td>(optional) for temperature control during live imaging</td></tr><tr><td>Transparent Tape 3/4&#039;&#039;</td><td>Scotch</td><td>S-9782</td><td>for preparing tape slide wells</td></tr><tr><td>Triton X-100</td><td>Sigma-Aldrich</td><td>X100</td><td>for immunostaining</td></tr><tr><td>Tween 20</td><td>Sigma-Aldrich</td><td>P1379</td><td>for immunostaining</td></tr><tr><td>Zebrafish: Tg(actb2:2xMCP-NLS-EGFP)</td><td>Campbell et&amp;nbsp;al., 2015</td><td>ZFIN: ZDB-TGCONSTRCT-150624-4</td><td>transgenic fish with nuclear localized EGFP</td></tr><tr><td>Zebrafish: Tg(Ola.Actb:Hsa.HRAS-EGFP)</td><td>Cooper et&amp;nbsp;al., 2005</td><td>ZFIN: ZDB-TGCONSTRCT-070117-75</td><td>transgenic fish with cell membrane localized EGFP</td></tr></tbody></table>

a 3-d tail explant culture to study vertebrate segmentation in zebrafish

Vertebrate embryos pattern their major body axis as repetitive somites, the precursors of vertebrae, muscle, and skin. Somites progressively segment from the presomitic mesoderm (PSM) as the tail end of the embryo elongates posteriorly. Somites form with regular periodicity and scale in size. Zebrafish is a popular model organism as it is genetically tractable and has transparent embryos that allow for live imaging. Nevertheless, during somitogenesis, fish embryos are wrapped around a large, rounding yolk. This geometry limits live imaging of PSM tissue in zebrafish embryos, particularly at higher resolutions that require a close objective working distance. Here, we present a flattened 3-D tissue culture method for live imaging of zebrafish tail explants. Tail explants mimic intact embryos by displaying a proportional slowdown of axis elongation and shortening of rostrocaudal somite lengths. We are further able to stall axis elongation speed through explant culture. This, for the first time, enables us to untangle the chemical input of signaling gradients from the mechanistic input of axial elongation. In future studies, this method can be combined with a microfluidic setup to allow time-controlled pharmaceutical perturbations or screening of vertebrate segmentation without any drug penetration concerns.

This protocol enables flat geometric culturing of live zebrafish tail explants. Tissue culture presents three major advantages over whole embryos: 1) control of axis elongation speed, 2) control over various signaling (morphogen) sources by simple dissection, and 3) near-objective, high magnification and high NA live imaging.
Chemically untreated slide chambers allow the tail explant to elongate its major axis (Figure 2A) by the skin ectoderm wrapping around the t...

Watch this Scientific Journal Video about A 3-D Tail Explant Culture to Study Vertebrate Segmentation in Zebrafish at JoVE.com

A 3-D Tail Explant Culture to Study Vertebrate Segmentation in Zebrafish

Here, we present the protocol for 3-D tissue culture of the zebrafish posterior body axis, enabling live study of vertebrate segmentation. This explant model provides control over axis elongation, alteration of morphogen sources, and subcellular resolution tissue-level live imaging.

Vertebrate embryos pattern their major body axis as repetitive somites, the precursors of vertebrae, muscle, and skin. Somites progressively segment from ...

a-3-d-tail-explant-culture-to-study-vertebrate-segmentation

Research

JoVE Journal

Developmental Biology

4.2K Views.  Cincinnati Children’s Hospital Medical Center. Here, we present the protocol for 3-D tissue culture of the zebrafish posterior body axis, enabling live study of vertebrate segmentation. This explant model provides control over axis elongation, alteration of morphogen sources, and subcellular resolution tissue-level live imaging.

Video: A 3-D Tail Explant Culture to Study Vertebrate Segmentation in Zebrafish

Capturing Tissue Repair in Zebrafish Larvae with Time-lapse Brightfield Stereomicroscopy

The zebrafish larval tail fin is ideal for studying tissue regeneration due to the simple architecture of the larval fin-fold, which comprises of two layers of skin that enclose undifferentiated mesenchyme, and because the larval tail fin regenerates rapidly within 2-3 days. Using this system, we demonstrate a method for capturing the repair dynamics of the amputated tail fin with time-lapse video brightfield stereomicroscopy. We demonstrate that fin amputation triggers a contraction of the amputation wound and extrusion of cells around the wound margin, leading to their subsequent clearance. Fin regeneration proceeds from proximal to distal direction after a short delay. In addition, developmental growth of the larva can be observed during all stages. The presented method provides an opportunity for observing and analyzing whole tissue-scale behaviors such as fin development and growth in a simple microscope setting, which is easily adaptable to any stereomicroscope with time-lapse capabilities.

We present a protocol for capturing the dynamics of zebrafish larval tail fin regeneration on a whole-tissue scale using brightfield-based stereomicroscopy. This technique enables capturing the regeneration dynamics with single cell resolution. This methodology can be adapted to any stereomicroscope equipped with a CCD camera and time-lapse software.

The zebrafish larval tail fin is ideal for studying tissue regeneration due to the simple architecture of the larval fin-fold, which comprises of two ...

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 ...

A Versatile Mounting Method for Long Term Imaging of Zebrafish Development

Zebrafish embryos offer an ideal experimental system to study complex morphogenetic processes due to their ease of accessibility and optical transparency. In particular, posterior body elongation is an essential process in embryonic development by which multiple tissue deformations act together to direct the formation of a large part of the body axis. In order to observe this process by long-term time-lapse imaging it is necessary to utilize a mounting technique that allows sufficient support to maintain samples in the correct orientation during transfer to the microscope and acquisition. In addition, the mounting must also provide sufficient freedom of movement for the outgrowth of the posterior body region without affecting its normal development. Finally, there must be a certain degree in versatility of the mounting method to allow imaging on diverse imaging set-ups. Here, we present a mounting technique for imaging the development of posterior body elongation in the zebrafish D. rerio. This technique involves mounting embryos such that the head and yolk sac regions are almost entirely included in agarose, while leaving out the posterior body region to elongate and develop normally. We will show how this can be adapted for upright, inverted and vertical light-sheet microscopy set-ups. While this protocol focuses on mounting embryos for imaging for the posterior body, it could easily be adapted for the live imaging of multiple aspects of zebrafish development.

Here, we present a versatile mounting method that allows for the long-term time-lapse imaging of the posterior body development of live zebrafish embryos without perturbing normal development.

Zebrafish embryos offer an ideal experimental system to study complex morphogenetic processes due to their ease of accessibility and optical transparency. ...

A 3D Culture Method of Spheroids of Embryonic and Liver Zebrafish Cell Lines

Fish cell lines are promising in vitro models for ecotoxicity assessment; however, conventional monolayer culture systems (2D culture) have well-known limitations (e.g., culture longevity and maintenance of some in vivo cellular functions). Thus, 3D cultures, such as spheroids, have been proposed, since these models can reproduce tissue-like structures, better recapturing the in vivo conditions. This article describes an effective, easy, and fast 3D culture protocol for the formation of spheroids with two zebrafish (Danio rerio) cell lines: ZEM2S (embryo) and ZFL (normal hepatocyte). The protocol consists of plating the cells in a round-bottom, ultra-low attachment, 96-well plate. After 5 days under orbital shaking (70 rpm), a single spheroid per well is formed. The formed spheroids present stable size and shape, and this method avoids the formation of multiple spheroids in a well; thus, it is not necessary to handpick spheroids of similar sizes. The ease, speed, and reproducibility of this spheroid method make it useful for high-throughput in vitro tests.

Here, we present an effective, easy, and fast 3D culture protocol for the formation of spheroids of two zebrafish (Danio rerio) cell lines: ZEM2S (embryo) and ZFL (normal hepatocyte).

Fish cell lines are promising in vitro models for ecotoxicity assessment; however, conventional monolayer culture systems (2D culture) have well-known ...

Environment

Dissection and Lateral Mounting of Zebrafish Embryos: Analysis of Spinal Cord Development

The zebrafish spinal cord is an effective investigative model for nervous system research for several reasons. First, genetic, transgenic and gene knockdown approaches can be utilized to examine the molecular mechanisms underlying nervous system development. Second, large clutches of developmentally synchronized embryos provide large experimental sample sizes. Third, the optical clarity of the zebrafish embryo permits researchers to visualize progenitor, glial, and neuronal populations. Although zebrafish embryos are transparent, specimen thickness can impede effective microscopic visualization. One reason for this is the tandem development of the spinal cord and overlying somite tissue. Another reason is the large yolk ball, which is still present during periods of early neurogenesis. In this article, we demonstrate microdissection and removal of the yolk in fixed embryos, which allows microscopic visualization while preserving surrounding somite tissue. We also demonstrate semipermanent mounting of zebrafish embryos. This permits observation of neurodevelopment in the dorso-ventral and anterior-posterior axes, as it preserves the three-dimensionality of the tissue.

Developmental processes such as proliferation, patterning, differentiation, and axon guidance can be readily modeled in the zebrafish spinal cord. In this article, we describe a mounting procedure for zebrafish embryos, which optimizes visualization of these events.

The zebrafish spinal cord is an effective investigative model for nervous system research for several reasons. First, genetic, transgenic and gene ...

Neuroscience

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 ...

Three and Four-Dimensional Visualization and Analysis Approaches to Study Vertebrate Axial Elongation and Segmentation

Somitogenesis is a hallmark of vertebrate embryonic development. For years, researchers have been studying this process in a variety of organisms using a wide range of techniques encompassing ex vivo and in vitro approaches. However, most studies still rely on the analysis of two-dimensional (2D) imaging data, which limits proper evaluation of a developmental process like axial extension and somitogenesis involving highly dynamic interactions in a complex 3D space. Here we describe techniques that allow mouse live imaging acquisition, dataset processing, visualization and analysis in 3D and 4D to study the cells (e.g., neuromesodermal progenitors) involved in these developmental processes. We also provide a step-by-step protocol for optical projection tomography and whole-mount immunofluorescence microscopy in mouse embryos (from sample preparation to image acquisition) and show a pipeline that we developed to process and visualize 3D image data. We extend the use of some of these techniques and highlight specific features of different available software (e.g., Fiji/ImageJ, Drishti, Amira and Imaris) that can be used to improve our current understanding of axial extension and somite formation (e.g., 3D reconstructions). Altogether, the techniques here described emphasize the importance of 3D data visualization and analysis in developmental biology, and might help other researchers to better address 3D and 4D image data in the context of vertebrate axial extension and segmentation. Finally, the work also employs novel tools to facilitate teaching vertebrate embryonic development.

Here, we describe computational tools and methods that allow visualization and analysis of three and four-dimensional image data of mouse embryos in the context of axial elongation and segmentation, obtained by in toto optical projection tomography, and by live imaging and whole-mount immunofluorescence staining using multiphoton microscopy.

Somitogenesis is a hallmark of vertebrate embryonic development. For years, researchers have been studying this process in a variety of organisms using a ...

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. ...

Preparation of Single-Somite Explants from Zebrafish Embryos

The body axis of vertebrate embryos is periodically subdivided into 3D multicellular units called somites. While genetic oscillations and molecular prepatterns determine the initial length-scale of somites, mechanical processes have been implicated in setting their final size and shape. To better understand the intrinsic material properties of somites, a method is developed to culture single-somite explant from zebrafish embryos. Single somites are isolated by first removing the skin of embryos, followed by yolk removal and sequential excision of neighboring tissues. Using transgenic embryos, the distribution of various sub-cellular structures can be observed by fluorescent time-lapse microscopy. Dynamics of explanted somites can be followed for several hours, thus providing an experimental framework for studying tissue-scale shape changes at single-cell resolution. This approach enables direct mechanical manipulation of somites, allowing for dissection of the material properties of the tissue. Finally, the technique outlined here can be readily extended for explanting other tissues such as the notochord, neural plate, and lateral plate mesoderm.

We present a protocol for isolating single somites from zebrafish embryos, the dynamics of which can be followed in culture for several hours by fluorescence time-lapse microscopy, thus providing a methodology to quantify tissue-scale shape changes at single-cell resolution.

The body axis of vertebrate embryos is periodically subdivided into 3D multicellular units called somites. While genetic oscillations and molecular ...

In vivo Electroporation of Morpholinos into the Regenerating Adult Zebrafish Tail Fin

Certain species of urodeles and teleost fish can regenerate their tissues. Zebrafish have become a widely used model to study the spontaneous regeneration of adult tissues, such as the heart1, retina2, spinal cord3, optic nerve4, sensory hair cells5, and fins6.
The zebrafish fin is a relatively simple appendage that is easily manipulated to study multiple stages in epimorphic regeneration. Classically, fin regeneration was characterized by three distinct stages: wound healing, blastema formation, and fin outgrowth. After amputating part of the fin, the surrounding epithelium proliferates and migrates over the wound. At 33 &deg;C, this process occurs within six hours post-amputation (hpa, Figure 1B)6,7. Next, underlying cells from different lineages (ex. bone, blood, glia, fibroblast) re-enter the cell cycle to form a proliferative blastema, while the overlying epidermis continues to proliferate (Figure 1D)8. Outgrowth occurs as cells proximal to the blastema re-differentiate into their respective lineages to form new tissue (Figure 1E)8. Depending on the level of the amputation, full regeneration is completed in a week to a month.
The expression of a large number of gene families, including wnt, hox, fgf, msx, retinoic acid, shh, notch, bmp, and activin-betaA genes, is up-regulated during specific stages of fin regeneration9-16. However, the roles of these genes and their encoded proteins during regeneration have been difficult to assess, unless a specific inhibitor for the protein exists13, a temperature-sensitive mutant exists or a transgenic animal (either overexpressing the wild-type protein or a dominant-negative protein) was generated7,12. We developed a reverse genetic technique to quickly and easily test the function of any gene during fin regeneration.
Morpholino oligonucleotides are widely used to study loss of specific proteins during zebrafish, Xenopus, chick, and mouse development17-19. Morpholinos basepair with a complementary RNA sequence to either block pre-mRNA splicing or mRNA translation. We describe a method to efficiently introduce fluorescein-tagged antisense morpholinos into regenerating zebrafish fins to knockdown expression of the target protein. The morpholino is micro-injected into each blastema of the regenerating zebrafish tail fin and electroporated into the surrounding cells. Fluorescein provides the charge to electroporate the morpholino and to visualize the morpholino in the fin tissue.
This protocol permits conditional protein knockdown to examine the role of specific proteins during regenerative fin outgrowth. In the Discussion, we describe how this approach can be adapted to study the role of specific proteins during wound healing or blastema formation, as well as a potential marker of cell migration during blastema formation.

In vivo Electroporation of Morpholinos into the Regenerating Adult Zebrafish Tail Fin

We describe a method to conditionally knockdown the expression of a target protein during adult zebrafish fin regeneration. This technique involves micro-injecting and electroporating antisense oligonucleotide morpholinos into fin tissue, which allows testing the protein&#x2019;s role in various stages of fin regeneration, including wound healing, blastema formation, and regenerative outgrowth.

Certain species of urodeles and teleost fish can regenerate their tissues. Zebrafish have become a widely used model to study the spontaneous regeneration ...

A 3-D Tail Explant Culture to Study Vertebrate Segmentation in Zebrafish

Podsumowanie

Przeglądaj więcej filmów

Rozdziały w tym wideo

Capturing Tissue Repair in Zebrafish Larvae with Time-lapse Brightfield Stereomicroscopy

Microbead Implantation in the Zebrafish Embryo

A Versatile Mounting Method for Long Term Imaging of Zebrafish Development

A 3D Culture Method of Spheroids of Embryonic and Liver Zebrafish Cell Lines

Dissection and Lateral Mounting of Zebrafish Embryos: Analysis of Spinal Cord Development

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

Three and Four-Dimensional Visualization and Analysis Approaches to Study Vertebrate Axial Elongation and Segmentation

Generation of Naïve Blastoderm Explants from Zebrafish Embryos

Preparation of Single-Somite Explants from Zebrafish Embryos

In vivo Electroporation of Morpholinos into the Regenerating Adult Zebrafish Tail Fin