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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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&#215;)</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&#39;&#39;</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&#160;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&#160;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...

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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 ...

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Research

JoVE Journal

Developmental Biology

4.1K 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

Zebrafish Keratocyte Explants to Study Collective Cell Migration and Reepithelialization in Cutaneous Wound Healing

Due to their unique motile properties, fish keratocytes dissociated from explant cultures have long been used to study the mechanisms of single cell migration. However, when explants are established, these cells also move collectively, maintaining many of the features which make individual keratocytes an attractive model to study migration: rapid rates of motility, extensive actin-rich lamellae with a perpendicular actin cable, and relatively constant speed and direction of migration. In early explants, the rapid interconversion of cells migrating individually with those migrating collectively allows the study of the role of cell-cell adhesions in determining the mode of migration, and emphasizes the molecular links between the two modes of migration. Cells in later explants lose their ability to migrate rapidly and collectively as an epithelial to mesenchymal transition occurs and genes associated with wound healing and inflammation are differentially expressed. Thus, keratocyte explants can serve as an in vitro model for the reepithelialization that occurs during cutaneous wound healing and can represent a unique system to study mechanisms of collective cell migration in the context of a defined program of gene expression changes. A variety of mutant and transgenic zebrafish lines are available, which allows explants to be established from fish with different genetic backgrounds. This allows the role of different proteins within these processes to be uniquely addressed. The protocols outlined here describe an easy and effective method for establishing these explant cultures for use in a variety of assays related to collective cell migration.

Zebrafish keratocytes migrate in cell sheets from explants and provide an in vitro model for the study of the mechanisms of collective cell migration in the context of epithelial wound healing. These protocols detail an effective way to establish primary explant cultures for use in collective cell migration assays.

Due to their unique motile properties, fish keratocytes dissociated from explant cultures have long been used to study the mechanisms of single cell ...

Use of the TetON System to Study Molecular Mechanisms of Zebrafish Regeneration

The zebrafish has become a very important model organism for studying vertebrate development, physiology, disease, and tissue regeneration. A thorough understanding of the molecular and cellular mechanisms involved requires experimental tools that allow for inducible, tissue-specific manipulation of gene expression or signaling pathways. Therefore, we and others have recently adapted the TetON system for use in zebrafish. The TetON system facilitates temporally and spatially-controlled gene expression and we have recently used this tool to probe for tissue-specific functions of Wnt/beta&#8211;catenin signaling during zebrafish tail fin regeneration. Here we describe the workflow for using the TetON system to achieve inducible, tissue-specific gene expression in the adult regenerating zebrafish tail fin. This includes the generation of stable transgenic TetActivator and TetResponder lines, transgene induction and techniques for verification of tissue-specific gene expression in the fin regenerate. Thus, this protocol serves as blueprint for setting up a functional TetON system in zebrafish and its subsequent use, in particular for studying fin regeneration.

Here we outline the workflow for using the TetON system to achieve tissue-specific gene expression in the adult regenerating zebrafish tail fin.

The zebrafish has become a very important model organism for studying vertebrate development, physiology, disease, and tissue regeneration. A thorough ...

Hepatocyte-specific Ablation in Zebrafish to Study Biliary-driven Liver Regeneration

The liver has a great capacity to regenerate. Hepatocytes, the parenchymal cells of the liver, can regenerate in one of two ways: hepatocyte- or biliary-driven liver regeneration. In hepatocyte-driven liver regeneration, regenerating hepatocytes are derived from preexisting hepatocytes, whereas, in biliary-driven regeneration, regenerating hepatocytes are derived from biliary epithelial cells (BECs). For hepatocyte-driven liver regeneration, there are excellent rodent models that have significantly contributed to the current understanding of liver regeneration. However, no such rodent model exists for biliary-driven liver regeneration. We recently reported on a zebrafish liver injury model in which BECs extensively give rise to hepatocytes upon severe hepatocyte loss. In this model, hepatocytes are specifically ablated by a pharmacogenetic means. Here we present in detail the methods to ablate hepatocytes and to analyze the BEC-driven liver regeneration process. This hepatocyte-specific ablation model can be further used to discover the underlying molecular and cellular mechanisms of biliary-driven liver regeneration. Moreover, these methods can be applied to chemical screens to identify small molecules that augment or suppress liver regeneration.

To help assess the molecular mechanisms underlying zebrafish biliary-driven liver regeneration, we established a liver injury model in which the nitroreductase-expressing hepatocytes are genetically ablated upon metronidazole treatment. In this protocol, we describe how to adeptly manipulate, monitor and analyze hepatocyte ablation and biliary-driven liver regeneration.

The liver has a great capacity to regenerate. Hepatocytes, the parenchymal cells of the liver, can regenerate in one of two ways: hepatocyte- or ...

Isolation and Culture of Adult Zebrafish Brain-derived Neurospheres

The zebrafish is a highly relevant model organism for understanding the cellular and molecular mechanisms involved in neurogenesis and brain regeneration in vertebrates. However, an in-depth analysis of the molecular mechanisms underlying zebrafish adult neurogenesis has been limited due to the lack of a reliable protocol for isolating and culturing neural adult stem/progenitor cells. Here we provide a reproducible method to examine adult neurogenesis using a neurosphere assay derived from zebrafish whole brain or from the telencephalon, tectum and cerebellum regions of the adult zebrafish brain. The protocol involves, first the microdissection of zebrafish adult brain, then single cell dissociation and isolation of self-renewing multipotent neural stem/progenitor cells. The entire procedure takes eight days. Additionally, we describe how to manipulate gene expression in zebrafish neurospheres, which will be particularly useful to test the role of specific signaling pathways during adult neural stem/progenitor cell proliferation and differentiation in zebrafish.

Here we provide a reproducible method to examine adult neurogenesis using a neurosphere assay derived from the whole brain or from either the telencephalic, tectal or cerebellar regions of the adult zebrafish brain. Additionally, we describe the procedure to manipulate gene expression in zebrafish neurospheres.

The zebrafish is a highly relevant model organism for understanding the cellular and molecular mechanisms involved in neurogenesis and brain regeneration ...

Production and Administration of Therapeutic Mesenchymal Stem/Stromal Cell (MSC) Spheroids Primed in 3-D Cultures Under Xeno-free Conditions

Mesenchymal stem/stromal cells (MSCs) hold great promise in bioengineering and regenerative medicine. MSCs can be isolated from multiple adult tissues via their strong adherence to tissue culture plastic and then further expanded in vitro, most commonly using fetal bovine serum (FBS). Since FBS can cause MSCs to become immunogenic, its presence in MSC cultures limits both clinical and experimental applications of the cells. Therefore, studies employing chemically defined xeno-free (XF) media for MSC cultures are extremely valuable. Many beneficial effects of MSCs have been attributed to their ability to regulate inflammation and immunity, mainly through secretion of immunomodulatory factors such as tumor necrosis factor-stimulated gene 6 (TSG6) and prostaglandin E2 (PGE2). However, MSCs require activation to produce these factors and since the effect of MSCs is often transient, great interest has emerged to discover ways of pre-activating the cells prior to their use, thus eliminating the lag time for activation in vivo. Here we present protocols to efficiently activate or prime MSCs in three-dimensional (3D) cultures under chemically defined XF conditions and to administer these pre-activated MSCs in vivo. Specifically, we first describe methods to generate spherical MSC micro-tissues or &#39;spheroids&#39; in hanging drops using XF medium and demonstrate how the spheres and conditioned medium (CM) can be harvested for various applications. Second, we describe gene expression screens and in vitro functional assays to rapidly assess the level of MSC activation in spheroids, emphasizing the anti-inflammatory and anti-cancer potential of the cells. Third, we describe a novel method to inject intact MSC spheroids into the mouse peritoneal cavity for in vivo efficacy testing. Overall, the protocols herein overcome major challenges of obtaining pre-activated MSCs under chemically defined XF conditions and provide a flexible system to administer MSC spheroids for therapies.

Production and Administration of Therapeutic Mesenchymal Stem/Stromal Cell MSC Spheroids Primed in 3-D Cultures Under Xeno-free Conditions

The therapeutic potential of mesenchymal stem/stromal cells (MSCs) is well-documented, however the best method of preparing the cells for patients remains controversial. Herein, we communicate protocols to efficiently generate and administer therapeutic spherical aggregates or 'spheroids' of MSCs primed under xeno-free conditions for experimental and clinical applications.

Mesenchymal stem/stromal cells (MSCs) hold great promise in bioengineering and regenerative medicine. MSCs can be isolated from multiple adult tissues via ...

Application of Aorta-gonad-mesonephros Explant Culture System in Developmental Hematopoiesis

The limitation of using mouse embryos for hematopoiesis studies is the added inconvenience in operations, which is largely due to the intrauterine development of the embryo. Although genetic data from knockout (KO) mice are convincing, it is not realistic to generate KO mice for all genes as needed. In addition, performing in vivo rescue experiments to consolidate the data obtained from KO mice is not convenient. To overcome these limitations, the Aorta-Gonad-Mesonephros (AGM) explant culture was developed as an appropriate system to study hematopoietic stem cell (HSC) development. Especially for rescue experiments, it can be used to recover the impaired hematopoiesis in KO mice. By adding the appropriate chemicals into the medium, the impaired signaling can be reactivated or up-regulated pathways can be inhibited. With the use of this method, many experiments can be performed to identify the critical regulators of HSC development, including HSC related gene expression at mRNA and protein levels, colony formation ability, and reconstitution capacity. This series of experiments would be helpful in defining the underlying mechanisms essential for HSC development in mammals.

This protocol describes using cultured Aorta-Gonad-Mesonephros for expression analyses, colony-forming units in the culture and spleen, and long-term reconstitution to determine the effect of regulatory factors and signaling pathways on hematopoietic stem cell development. This has been demonstrated as an effective system for studying hematopoietic stem cell biology and function.

The limitation of using mouse embryos for hematopoiesis studies is the added inconvenience in operations, which is largely due to the intrauterine ...

Organotypic Culture Method to Study the Development Of Embryonic Chicken Tissues

The embryonic chicken is commonly used as a reliable model organism for vertebrate development. Its accessibility and short incubation period makes it ideal for experimentation. Currently, the study of these developmental pathways in the chicken embryo is conducted by applying inhibitors and drugs at localized sites and at low concentrations using a variety of methods. In vitro tissue&#160;culturing is a technique that enables the study of tissues separated from the host organism, while simultaneously bypassing many of the physical limitations present when working with whole embryos, such as the susceptibility of embryos to high doses of potentially lethal chemicals. Here, we present an organotypic culturing protocol for culturing the embryonic chicken half head in vitro, which presents new opportunities for the examination of developmental processes beyond the currently established methods.

Here, we present an organotypic culturing protocol to grow embryonic chicken organs in vitro. Using this method, the development of embryonic chicken tissue can be studied, while maintaining a high degree of control over the culture environment.

The embryonic chicken is commonly used as a reliable model organism for vertebrate development. Its accessibility and short incubation period makes it ...

Culture and Transfection of Zebrafish Primary Cells

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

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

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

Adult Zebrafish Injury Models to Study the Effects of Prednisolone in Regenerating Bone Tissue

Zebrafish are able to regenerate various organs, including appendages (fins) after amputation. This involves the regeneration of bone, which regrows within roughly two weeks after injury. Furthermore, zebrafish are able to heal bone rapidly after trepanation of the skull, and repair fractures that can be easily introduced into zebrafish bony fin rays. These injury assays represent feasible experimental paradigms to test the effect of administered drugs on rapidly forming bone. Here, we describe the use of these 3 injury models and their combined use with systemic glucocorticoid treatment, which exerts bone inhibitory and immunosuppressive effects. We provide a workflow on how to prepare for immunosuppressive treatment in adult zebrafish, illustrate how to perform fin amputation, trepanation of calvarial bones, and fin fractures, and describe how the use of glucocorticoids affects both bone forming osteoblasts and cells of the monocyte/macrophage lineage as part of innate immunity in bone tissue.

Here, we describe 3 adult zebrafish injury models and their combined use with immunosuppressive drug treatment. We provide guidance on imaging of regenerating tissues and on detecting bone mineralization therein.

Zebrafish are able to regenerate various organs, including appendages (fins) after amputation. This involves the regeneration of bone, which regrows ...

Whole-Mount In Situ Hybridization in Zebrafish Embryos and Tube Formation Assay in iPSC-ECs to Study the Role of Endoglin in Vascular Development

Vascular development is determined by the sequential expression of specific genes, which can be studied by performing in situ hybridization assays in zebrafish during different developmental stages. To investigate the role of endoglin(eng) in vessel formation during the development of hereditary hemorrhagic telangiectasia&#160;(HHT), morpholino-mediated targeted knockdown of eng in zebrafish are used to study its temporal expression and associated functions. Here, whole-mount in situ RNA hybridization (WISH) is employed for the analysis of eng and its downstream genes in zebrafish embryos. Also, tube formation assays are performed in HHT patient-derived induced pluripotent stem cell-differentiated&#160;endothelial cells (iPSC-ECs; with eng mutations). A specific signal amplifying system using the whole amount In Situ Hybridization &#8211; WISH provides higher resolution and lower background results compared to traditional methods. To obtain a better signal, the post-fixation time is adjusted to 30 min after probe hybridization. Because fluorescence staining is not sensitive in zebrafish embryos, it is replaced with diaminobezidine (DAB) staining here. In this protocol, HHT&#160;patient-derived iPSC lines containing an eng mutation are differentiated into endothelial cells. After coating a plate with basement membrane matrix for 30 min at 37 &#176;C, iPSC-ECs are seeded as a monolayer into wells and kept at 37 &#176;C for 3 h. Then, the tube length and number of branches are calculated using microscopic images. Thus, with this improved WISH protocol, it is shown that reduced eng expression affects endothelial progenitor formation in zebrafish embryos. This is further confirmed by tube formation assays using iPSC-ECs derived from a patient with HHT. These assays confirm the role for eng in early vascular development.

Presented here is a protocol for whole-mount in situ RNA hybridization analysis in zebrafish and tube formation assay in patient-derived induced pluripotent stem cell-derived endothelial cells to study the role of endoglin in vascular formation.

Vascular development is determined by the sequential expression of specific genes, which can be studied by performing in situ hybridization assays in ...