This study was inspired by the work of Levine et al., which described this experimental method and provided a comprehensive description of vascular development in Xenopus laevis. We thank the members of our laboratory for their input. This study was supported by the Yonsei University Future-leading Research Initiative of 2015 (2015-22- 0095) and the Bio &#38; Medical Technology Development Program of the National Research Foundation (NRF) funded by the ministry of Science, ICT &#38; Future Planning (NRF-2013M3A9D5072551)

All experiments complied with protocols approved by the Yonsei University College of Medicine Institutional Animal Care and Use Committees.
1. Preparation of Xenopus tropicalis Embryos
NOTE: Xenopus tropicalis embryos were produced as previously described10, with slight modification. Xenopus tropicalis embryos were staged according to the tables of Nieuwkoop and Faber11 .
<ol>
	<li>Induction of ovulation
	<ol>
		<li>To pre-prime, inject human chorionic gonadotropin (hCG) into the dorsal lymph sac of a pair of frogs (one male and one female) on the day prior to the day of mating. Inject 20 units per female frog and 10 units per male. House a male and a female in the same tank overnight (for more than 18 h).</li>
		<li>To prime, on the following day, inject 200 units of hCG to the pre-primed female and 100 units of hCG to the pre-primed male; the primed female will start to lay eggs in approximately 3 h and will continue to do so throughout the day. Keep the male and female pair together for natural mating or in separate tanks for in vitro fertilization (step 1.2).</li>
	</ol>
	</li>
	<li>Fertilization 
	NOTE: Eggs can be fertilized by natural mating or in vitro fertilization. In natural mating, eggs are fertilized as they are laid, and therefore, the timing of fertilization cannot be controlled. Alternatively, a primed female frog can be housed in a separate tank and unfertilized eggs can be collected for in vitro fertilization, which generates hundreds of embryos fertilized at the same time. The latter approach is useful when blastomere-targeted microinjection will be performed prior to vessel labeling. In this approach, however, a male is sacrificed.
	<ol>
		<li>Natural mating
		<ol>
			<li>Leave primed female and male pair in the same tank. The male will clasp the female, leading to immediate fertilization of the laid eggs. Collect the eggs with a glass or plastic pipette and transfer them to a Petri dish. To prevent the eggs from sticking to the pipette, coat the inner surface of the pipette with 0.5% bovine serum albumin (BSA).</li>
		</ol>
		</li>
		<li>In vitro fertilization (IVF)
		<ol>
			<li>Separate the primed female from the male after priming. The female will start to lay eggs spontaneously in approximately 3 h. Transfer a few laid eggs using a BSA-coated pipette to a Petri dish and check their quality under a microscope. If healthy eggs (clear pigment, elastic ball-shaped) were laid, prepare to dissect out the testes from the primed male.</li>
			<li>Make 0.4% Tricaine methanesulfonate (MS222) for euthanasia and inject 300 &#181;L into the dorsal lymph sac of the primed male. In approximately 10 min, the male will lose his sense of balance and will stay afloat; confirm that the male is euthanized by pinching a hind leg with a pair of blunt forceps and observing no response.</li>
			<li>Dissect out the two testes, clean them by quickly tapping them on lint-free paper, and transfer them into a microtube containing 1 mL of 60% Leibovitz&#39;s L-15 Medium (60%) supplemented with 10% fetal bovine serum (FBS); a detailed procedure is described elsewhere10.</li>
			<li>Gently mince the testes with a pestle to extract the sperm. At this concentration (i.e., 2 testes in 1 mL of 10% FBS in 60% L-15), a few drops of the testis solution (0.1 mL) can fertilize approximately three hundred eggs. The remaining sperm can be stored at 4 &#176;C in a tightly sealed tube and used for IVF the next day.</li>
			<li>Squeeze the eggs from the female into a small Petri dish. Hold the upper part of the female upside down on the palm and put the forefinger between its hind legs. Spread its hind legs with the forefinger and the other hand to expose the cloaca. Touch the cloaca to the Petri dish. Coat the Petri dish with 0.5% BSA to evenly distribute the eggs as a monolayer during IVF.</li>
			<li>Add a few drops (0.1 mL) of sperm-containing L-15 medium to the eggs. For synchronous fertilization, gently stir the sperm solution across the dish. After 4 min at room temperature, fill the Petri dish with 0.01x Modified Barth&#39;s Saline (MBS), incubate for 10 min, and replace the solution with 0.1x MBS. The fertilized eggs will undergo the first cleavage in approximately 1 h at room temperature. 1x MBS is 88 mM NaCl, 1 mM KCl, 2.5 mM NaHCO3, 5 mM HEPES, 1 mM MgSO4, and 0.7 mM CaCl2, pH 7.5.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>(Optional) Injection of antisense morpholino oligonucleotides (MOs) and/or mRNAs
	<ol>
		<li>To investigate the effect of a gene of interest on vessel formation, perform the microinjection of antisense MO and/or mRNAs.</li>
		<li>For such experiments, de-jelly the eggs with 2% L-cysteine in 1x MBS (pH 7.5-7.8). Replace 0.1x MBS in the dish containing the fertilized embryos with 2% L-cysteine in 1x MBS. Gently swirl the dish to mix the solution. 
		NOTE: It usually takes about 5 min for the embryos to be de-jellied. During this step, monitor the eggs frequently to ensure that they are not over-exposed to the solution. If the eggs are over-exposed, they will lose elasticity and become flattened, which will cause aberrant development and lethality.</li>
		<li>Wash the de-jellied eggs five times with 0.1x MBS. Perform the microinjection in 4% Ficoll dissolved in 0.1x MBS. Typically, inject 4 ng of MO and 600 pg of mRNA per blastomere at the two-cell stage. Inject into one cell to manipulate gene expression on only one side of the embryo. Use the uninjected side of the same embryo as a control.</li>
		<li>As another control for the specificity of gene manipulation, inject the same amount of a control MO (CoMO) or control mRNA (e.g., beta-galactosidase mRNA). CoMO must have the same molecular weight as the gene-specific MO and must not target any gene in the Xenopus genome. For easy visualization of the injected side, use fluorescein-tagged MO or co-inject a tracer (e.g., EGFP mRNA).</li>
		<li>Leave the injected embryos in 4% Ficoll for 2 h and then transfer them to a new 60 mm Petri dish filled with 0.1x MBS.</li>
	</ol>
	</li>
	<li>Raising
	<ol>
		<li>Raise the embryos at 23 &#176;C. Although the speed of development can be slowed down or accelerated by using temperatures below or above 23 &#176;C, respectively (range: 16-26 &#176;C), it is recommended to raise them at 23 &#176;C.</li>
	</ol>
	</li>
	<li>(Optional) Treatment with pharmacological reagents
	<ol>
		<li>For pharmacological manipulations, administer drugs to the developing embryos by bath application.</li>
	</ol>
	</li>
</ol>
2. Preparation of DiI-AcLDL Injection
<ol>
	<li>Make a tapering glass pipette using a standard micropipette puller. Place a borosilicate glass tube in the micropipette puller and use the following settings: pressure, 200; heat, 30; pull, 30; velocity, 120; time, 200.</li>
	<li>Store DiI-AcLDL stock solution at 4 &#176;C. Take the upper clear part of the solution for injection, but not the precipitated particles on the bottom of tube. Any debris in the glass pipette will disturb the proper ejection of the solution.</li>
	<li>Fill the prepared glass pipette with a suitable amount of DiI-AcLDL solution (~5 &#181;L) using a microloader.</li>
	<li>Clip off the tip of the glass pipette little by little using a fine pair of forceps (number 55). Set up the pressure-controlled injection system so that approximately 5 nL of solution is ejected at a time; 50-60 nL of DiI-AcLDL solution is enough to stain the vessels of one embryo.</li>
	<li>Do not leave the DiI-AcLDL-loaded pipette in air for an extended period of time in order to avoid drying. Keep the tip of the pipette submerged in 0.04% MS222 in&#160;0.1X MBS solution. Just before injecting it into the embryo (step 4.2.1), check to see whether the same amount of dye ejects.</li>
</ol>
3. Injection Setup
<ol>
	<li>Sylgard mold
	<ol>
		<li>To prepare a small Sylgard mold, mix Sylgard base and curing agent with a weight ratio of 10 to 1. Fill approximately half of a small Petri dish (35 mm or 60 mm dish) with this mixed solution and then leave it to solidify for approximately 48 h at room temperature.</li>
	</ol>
	</li>
	<li>Anesthesia
	<ol>
		<li>Use 0.04% MS222 in 1x MBS (pH 7.5-8.0) for anesthesia. When long anesthesia is needed (e.g., during live imaging), use 0.04% MS222 in 0.1x MBS to reduce the osmotic imbalance. To adjust the pH, add a few drops of NaOH (~20 &#181;L) and check the pH using pH paper.</li>
		<li>Wait until the transferred embryos stop moving. Touch the dorsal fins of the embryos and inspect whether they respond to confirm complete anesthesia.</li>
	</ol>
	</li>
	<li>Positioning embryos for injection
	<ol>
		<li>Indent the surface of the hardened Sylgard mold with a thin and sharp blade. Make a V-shaped concave indentation in which to place the embryos (see Figure 2D). Place an anesthetized embryo, ventral-side up, in a slightly oblique fashion (approximately 30&#176;) (Figure 2D).</li>
		<li>Fill the mold with MS222 solution and place the embryos in the mold.</li>
	</ol>
	</li>
</ol>
4. DiI-AcLDL Injection
<ol>
	<li>Incision of skin overlying the heart
	<ol>
		<li>Prepare two pins (Minutiens, 0.10 mm) to cut the skin overlying the heart. Tightly fix each pin to a pin-holder; the heart should be readily identifiable as it pulsates (Figure 2A-2C, pink).</li>
		<li>Puncture the skin of an embryo with one pin. Insert the pin through the hole in the space between the heart and the skin (Figure 2C, solid arrow). Do not puncture the heart. Position the inserted part of this pin in the region where the incision will be made.</li>
		<li>Make a linear incision by gently rubbing the other pin held outside the skin against the inserted pin. Gently widen this incision with two pins,exposing the heart (Figure 2B&#39; and 2D&#39;, arrow). The pipette loaded with DiI-AcLDL can now directly approach the heart (Figure 2D&#39;).</li>
	</ol>
	</li>
	<li>Injection
	<ol>
		<li>Insert the tip of the loaded glass pipette into the heart and inject 50-60 nL of DiI-AcLDL solution in approximately 10 ejection pulses. Do not inject an excessive amount of solution, as it causes the heart to rupture. Check whether the heart continues to beat after the injection. After every injection, check whether the same amount of DiI-AcLDL is ejected; otherwise, the tip of the pipette may be blocked (see step 2.5).</li>
	</ol>
	</li>
	<li>Recovery and visual screening
	<ol>
		<li>Transfer the injected embryos to a new Petri dish filled with 0.1x MBS for recovery. After 5-10 min, the tadpoles should regain consciousness and start to move. At this time, the labeled vessels can be imaged. It is recommended to visually screen successfully labeled embryos under a fluorescence microscope (Figure 3A and 3B). If an insufficient amount of DiI-AcLDL is injected into an embryo, it can be injected again (Figure 3C).</li>
		<li>Discard the tadpoles that have other organs labeled (Figure 3D). Continue until the required number of successfully labeled embryos is obtained.</li>
	</ol>
	</li>
	<li>(Optional) Fixation of the embryos
	<ol>
		<li>For more thorough imaging of blood vessels, use confocal microscopy; it might be easier to fix the labeled embryos for confocal microscopy. First anesthetize the labeled embryos in 0.04% MS222 in 1x MBS (pH 7.5-8.0) and then fix them in 4% paraformaldehyde in 1X&#160;phosphate-buffered saline (PBS) for 1 h at room temperature. Fixed embryos can be stored for several days at 4 &#176;C; protect the injected samples from light to avoid photobleaching of the DiI.</li>
	</ol>
	</li>
</ol>
5. Imaging of DiI-AcLDL
<ol>
	<li>Stereoscopic imaging
	<ol>
		<li>Take photographs under a fluorescence stereoscopic microscope (Figure 3) to assess the overall morphology of the vasculature.</li>
		<li>Melt 1% agarose in 1x PBS&#160;for fixed embryos or in 1x MBS for live imaging. Pour melted agarose into a Petri dish and wait until it solidifies. Make a shallow indentation on the surface of agarose gel that will fit the embryo to be imaged when lying on its lateral side. Fill the dish with buffer (MS222 solution for anesthetized embryos or 1x PBS for fixed embryos).</li>
	</ol>
	</li>
	<li>Fluorescence microscopy (Rhodamine filter set)
	<ol>
		<li>As DiI is a green-excited and red-emitting dye, image using a standard filter set for rhodamine or spectrally related fluorophores (maximum excitation at 554 nm and emission at 571 nm). Place the injected embryos in the indentations on the agarose mold and take photographs (Figure 3).</li>
	</ol>
	</li>
	<li>Confocal microscopy 
	NOTE: For a closer analysis of vascular architecture, use confocal microscopy.
	<ol>
		<li>If using an inverted microscope, position the embryos on a glass-bottom dish. Add a few drops of 1x PBS and immobilize them by placing a cover slip on top. Remove excess PBS. If live embryos are to be imaged, use anesthetized embryos and use 0.015% MS222 in 0.1x MBS instead of PBS.</li>
		<li>For low-magnification imaging, use a 4X to 10X objective to find the area of interest; the rostral part of the posterior cardinal vein (PCV) is a useful region to investigate developmental angiogenesis (Figure 4, dashed boxes).
		<ol>
			<li>Use an acquisition setting for a red emission fluorophore (such as Rhodamine).</li>
			<li>Make z-stacked images by imaging the lateral half of an embryo. First, focus on the PCV on the lateral side near the objective and then set the lower limit of the focus at the position where the vessels nearest to the objective are in focus. Set the upper limit at the position where the medial part of the PCV being imaged can be focused. Image this entire lateral half of the embryo. Use software that stores metadata on the acquisition settings, such as x, y, and z scales, as they are required to calculate the vessel lengths.</li>
			<li>If needed, use the tile scan mode to image the entire vasculature of an embryo. Empirically determine the number of horizontal and vertical tiles.</li>
		</ol>
		</li>
		<li>For high-magnification imaging, follow the procedures above to image the rostral part of the PCV and 4 rostral-most intersomitic veins (ISVs) (Figure 4, dashed box). Adjust the number of stacks (z-stack) and tiles (tile scan) appropriately.</li>
	</ol>
	</li>
</ol>
6. Quantification of DiI-labeled Vessels
NOTE: DiI-labeled vessels can be traced manually or using software. We use &#34;Simple neurite tracer,&#34; a free ImageJ (NIH) plugin, which allows semi-automatic tracing of tube-like structures such as blood vessels and neurites12. Using this free software, the following parameters can be calculated. A detailed procedure for the use of this software is described elsewhere (https://imagej.net/Simple_Neurite_Tracer)12. Below, we use examples of the vasculature of embryos in which Tie2 signaling is inhibited or enhanced (Figure 4A-4C)5. Antisense Tie2MO-injected embryos exhibit reduces angiogenesis and shortens ISVs (Figure 4B), whereas the co-injection of constitutively active Tie2 mutant mRNA (caTie2 mRNA) over-rescues this phenotype, resulting in exuberant ISV branches (Figure 4C).
<ol>
	<li>ISV lengths
	<ol>
		<li>Using Simple neurite tracer, calculate the length of each traced vessel. Calculate the lengths of the 4 rostral-most ISVs.</li>
	</ol>
	</li>
	<li>ISV branches 
	NOTE: In wildtype embryos, only a few short branches sprout from the rostral-most ISVs at stage 42.
	<ol>
		<li>In Simple neurite tracer, trace these small vessels after making the ISV from which they originate the primary branch. Calculate the following parameters of these branches originating from the ISVs.</li>
		<li>Total branch number
		<ol>
			<li>Once all of the ISVs and the associated branches are traced, calculate the total number of branches (Figure 5B-5D).</li>
		</ol>
		</li>
		<li>Branch order
		<ol>
			<li>As Simple neurite tracer records the order of each branch (Figure 5A), calculate the number of branches for each order. The majority of vessels in wildtype embryos should be primary branches (i.e., ISVs).</li>
		</ol>
		</li>
		<li>Vein complexity index (VCI)
		<ol>
			<li>As VCI is a useful parameter for vessel complexity, which gives more weight to higher-order branches, calculate the VCI for each embryo using the formula in Figure 5A. For example, Tie2MO-injected embryos have a lower VCI compared to control (Figure 5B and 5C), and caTie2 mRNA-injected embryos have a higher VCI (Figure 5D).</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

The protocol presented here was first developed by Ali H. Brivanlou and colleagues to investigate developmental events during vascular formation in Xenopus laevis4, but, as shown in this manuscript, it can be applied to other small animals. Dye injection into the heart is simple to perform, and the entire vascular network can be imaged under a fluorescence dissection microscope, as well as a confocal microscope. If the dye is injected into the heart during vessel development, the dynamics...

Vasculogenesis, the formation of new blood vessels from newly born endothelial cells, and angiogenesis, the formation of new vessels from pre-existing vessels, are two distinct processes that shape embryonic vasculature1. Any dysregulation in these processes results in various heart diseases and structural abnormalities of vessels. Furthermore, tumor growth is associated with uncontrolled vessel growth. As such, molecular mechanisms underlying vasculogenesis and angiogenesis are the subject of intense investigation2.
Xenopus and zebrafish are attractive vertebrate models for vasculogenesis and angiogenesis studies, for several reasons. First, their embryos are small; therefore, it is relatively easy to image the entire vasculature. Second, embryonic development is rapid; it only takes a couple of days for the entire vasculature to develop, during which time the developing vasculature can be imaged. Third, genetic and pharmacological interventions before and during vessel formation are easy to perform, such as through the microinjection of antisense morpholino nucleotides (MOs) into the developing embryo or through the bath application of drugs3,4,5.
The unique advantage of Xenopus over zebrafish is that embryological manipulations can be performed because Xenopus follows stereotypical holoblastic cleavages and the embryonic fate map is well defined6. For example, it is possible to generate an embryo in which only one lateral side is genetically manipulated by injecting an antisense MO to one cell at the two-cell stage. It is also possible to transplant the heart primordium from one embryo to another to determine whether the gene exerts its function by a cell-intrinsic or -extrinsic mechanism7. Although these techniques have mostly been developed in Xenopus laevis, which is allotetraploid and is therefore not ideal for genetic studies, they can be directly applied to Xenopus tropicalis, a closely related diploid species8.
One way to visualize the vasculature in a live Xenopus embryo is to inject a fluorescent dye to label the blood vessels. Acetylated low density lipoprotein (AcLDL) labeled with a fluorescent molecule such as DiI is a very useful probe. Unlike unacetylated LDL, AcLDL does not bind to the LDL receptor9 but is endocytosed by macrophages and endothelial cells. The injection of DiI-AcLDL into the heart of a live animal results in the specific fluorescent labeling of endothelial cells, and the entire vasculature can be imaged by fluorescence microscopy in live or fixed embryos4.
Here, we present detailed protocols for the visualization and quantification of blood vessels using DiI-AcLDL in Xenopus tropicalis (Figure 1). We provide key practical points, with examples of successful and unsuccessful experiments. In addition, we provide a straightforward method for the quantitative analysis of vascular complexity, which might be useful in assessing the effects of genetic and environmental factors on the shaping of the vascular network.

<table><tbody><tr><td>35 mm Petri dish</td><td>SPL</td><td>10035</td><td>Sylgard mold frame</td></tr><tr><td>60 mm Petri dish</td><td>SPL</td><td>10060</td><td>Embryo raising tray</td></tr><tr><td>Borosilicate Glass</td><td>Sutter instrument</td><td>B100-50-10</td><td>Needle for injection</td></tr><tr><td>BSA</td><td>Sigma</td><td>A3059-10G</td><td>Coating reagent</td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>D.S.P.GR Reagent</td><td></td><td>0.1x MBS component</td></tr><tr><td>Coverslip</td><td>Superior</td><td>HSU-0111520</td><td>For confocal imaging</td></tr><tr><td>DiI-AcLDL</td><td>Thermo Fisher Scientific</td><td>L3484</td><td>Vessel staining solution</td></tr><tr><td>FBS</td><td>Hyclone</td><td>SH.30919.02</td><td>For storage of testis</td></tr><tr><td>Fiber Optical Illuminator</td><td>World Precision Instruments</td><td>Z-LITE-Z</td><td>Light</td></tr><tr><td>Ficoll</td><td>Sigma</td><td>F4375</td><td>Injection buffer</td></tr><tr><td>Flaming/Brown Micropipette Puller</td><td>Sutter instrument</td><td>P-97</td><td>Injection needle puller</td></tr><tr><td>Forcep</td><td>Fine Science Tool</td><td>11255-20</td><td>For embryo hatching and&lt;br/&gt; needle tip cutting</td></tr><tr><td>Glass Bottom dish</td><td>SPL</td><td>100350</td><td>For confocal imaging</td></tr><tr><td>hCG</td><td>MNS Korea</td><td></td><td>For priming of frogs</td></tr><tr><td>HEPES</td><td>Sigma</td><td>H3375</td><td>Buffering agent</td></tr><tr><td>Incubator</td><td>Lab. Companion</td><td>ILP-02</td><td>For raising embryos</td></tr><tr><td>KCl</td><td>DAEJUNG</td><td>6566-4400</td><td>MBS component</td></tr><tr><td>L15 medium</td><td>Gibco</td><td>11415-114</td><td>For storage of testis</td></tr><tr><td>L-cysteine</td><td>Sigma</td><td>168149-100G</td><td>De-jellying reagent</td></tr><tr><td>MgSO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma</td><td>M7506</td><td>MBS component</td></tr><tr><td>Microtube</td><td>Axygen</td><td>MCT-175-C-S</td><td>For storage of testis</td></tr><tr><td>MS222</td><td>Sigma</td><td>E10521</td><td>Anesthetic powder</td></tr><tr><td>NaCl</td><td>DAEJUNG</td><td>7647-14-5</td><td>MBS component</td></tr><tr><td>NaOH</td><td>Sigma</td><td>S-0899</td><td>pH adjusting reagent</td></tr><tr><td>Paraformaldehyde</td><td>Sigma</td><td>P6148</td><td>Fixatives</td></tr><tr><td>PBS</td><td>BIOSESANG</td><td>P2007</td><td>Buffer for imaging</td></tr><tr><td>pH paper</td><td>Sigma</td><td>P4536-100EA</td><td>For confirming pH</td></tr><tr><td>PICO-LITER INJECTOR</td><td>Waner instruments</td><td>PLI-100A</td><td>For injection</td></tr><tr><td>Pin</td><td>Pinservice</td><td>26002-10</td><td>For incision</td></tr><tr><td>Pinholder</td><td>Scitech Korea</td><td>26016-12</td><td>For incision</td></tr><tr><td>Precision Stereo Zoom Binocular Microscope</td><td>World Precision Instruments</td><td>PZMIII</td><td>For visual screening</td></tr><tr><td>Standard Manual Control Micromanipulator&amp;nbsp;</td><td>Waner instruments</td><td>W4 64-0056</td><td>For microinjection</td></tr><tr><td>SYLGARD 184 Kit</td><td>Dow Corning</td><td></td><td>For DiI injection</td></tr><tr><td>Transfer pipette</td><td>Korea Ace Scientific Co.</td><td>YM.B78-400</td><td>For eggs and&lt;br/&gt; embryo collection</td></tr></tbody></table>

visualization and quantitative analysis of embryonic angiogenesis in xenopus tropicalis

Blood vessels supply oxygen and nutrients throughout the body, and the formation of the vascular network is under tight developmental control. The efficient in vivo visualization of blood vessels and the reliable quantification of their complexity are key to understanding the biology and disease of the vascular network. Here, we provide a detailed method to visualize blood vessels with a commercially available fluorescent dye, human plasma acetylated low density lipoprotein DiI complex (DiI-AcLDL), and to quantify their complexity in Xenopus tropicalis. Blood vessels can be labeled by a simple injection of DiI-AcLDL into the beating heart of an embryo, and blood vessels in the entire embryo can be imaged in live or fixed embryos. Combined with gene perturbation by the targeted microinjection of nucleic acids and/or the bath application of pharmacological reagents, the roles of a gene or of a signaling pathway on vascular development can be investigated within one week without resorting to sophisticated genetically engineered animals. Because of the well-defined venous system of Xenopus and its stereotypic angiogenesis, the sprouting of pre-existing vessels, vessel complexity can be quantified efficiently after perturbation experiments. This relatively simple protocol should serve as an easily accessible tool in diverse fields of cardiovascular research.

Timeline of experiments (Figures 1 and 2)
Shortly after fertilization, targeted microinjection can be performed to modulate gene expression. For example, an antisense MO that specifically binds to the initiation codon of the endogenous Tie2 mRNA can be injected, inhibiting the translation of Tie2 target mRNA by steric hindrance. A MO can be conjugated to fluorescein for the easy visual screening of successfully injected embryos. Al...

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Visualization and Quantitative Analysis of Embryonic Angiogenesis in Xenopus tropicalis

This protocol demonstrates a fluorescence-based method to visualize the vasculature and to quantify its complexity in Xenopus tropicalis. Blood vessels can be imaged minutes after the injection of a fluorescent dye into the beating heart of an embryo after genetic and/or pharmacological manipulations to study cardiovascular development in vivo.

Visualization and Quantitative Analysis of Embryonic Angiogenesis in Xenopus tropicalis

Blood vessels supply oxygen and nutrients throughout the body, and the formation of the vascular network is under tight developmental control. The ...

visualization-quantitative-analysis-embryonic-angiogenesis-xenopus

Research

JoVE Journal

Developmental Biology

8.4K Views.  Yonsei University College of Medicine.  Dit protocol toont een fluorescentie gebaseerde methode om de vasculatuur te visualiseren en de complexiteit ervan te kwantificeren in Xenopus tropicalis . Bloedvaten kunnen na de injectie van een fluorescerende kleurstof in het kloppende hart van een embryo worden gedetecteerd na genetische en / of farmacologische manipulaties om in vivo cardiovasculaire ontwikkeling te bestuderen. 

Video: Visualization and Quantitative Analysis of Embryonic Angiogenesis in Xenopus tropicalis

Understanding Early Organogenesis Using a Simplified In Situ Hybridization Protocol in Xenopus

Organogenesis is the study of how organs are specified and then acquire their specific shape and functions during development. The Xenopuslaevis embryo is very useful for studying organogenesis because their large size makes them very suitable for identifying organs at the earliest steps in organogenesis. At this time, the primary method used for identifying a specific organ or primordium is whole mount in situ hybridization with labeled antisense RNA probes specific to a gene that is expressed in the organ of interest. In addition, it is relatively easy to manipulate genes or signaling pathways in Xenopus and in situ hybridization allows one to then assay for changes in the presence or morphology of a target organ. Whole mount in situ hybridization is a multi-day protocol with many steps involved. Here we provide a simplified protocol with reduced numbers of steps and reagents used that works well for routine assays. In situ hybridization robots have greatly facilitated the process and we detail how and when we utilize that technology in the process. Once an in situ hybridization is complete, capturing the best image of the result can be frustrating. We provide advice on how to optimize imaging of in situ hybridization results. Although the protocol describes assessing organogenesis in Xenopus laevis, the same basic protocol can almost certainly be adapted to Xenopus tropicalis and other model systems.

Understanding Early Organogenesis Using a Simplified In Situ Hybridization Protocol in Xenopus

The Xenopus laevis embryo continues to be exceptionally useful in the study of early development due to its large size and ease of manipulation. A simplified protocol for whole mount in situ hybridization protocol is provided that can be used in the identification of specific organs in this model system.

Organogenesis is the study of how organs are specified and then acquire their specific shape and functions during development.  The Xenopuslaevis embryo ...

Using Confocal Analysis of Xenopus laevis to Investigate Modulators of Wnt and Shh Morphogen Gradients

This protocol describes a method to visualise ligands distributed across a field of cells. The ease of expressing exogenous proteins, together with the large size of their cells in early embryos, make Xenopus laevis a useful model for visualising GFP-tagged ligands. Synthetic mRNAs are efficiently translated after injection into early stage Xenopus embryos, and injections can be targeted to a single cell. When combined with a lineage tracer such as membrane tethered RFP, the injected cell (and its descendants) that are producing the overexpressed protein can easily be followed. This protocol describes a method for the production of fluorescently tagged Wnt and Shh ligands from injected mRNA. The methods involve the micro dissection of ectodermal explants (animal caps) and the analysis of ligand diffusion in multiple samples. By using confocal imaging, information about ligand secretion and diffusion over a field of cells can be obtained. Statistical analyses of confocal images provide quantitative data on the shape of ligand gradients. These methods may be useful to researchers who want to test the effects of factors that may regulate the shape of morphogen gradients.

Using Confocal Analysis of Xenopus laevis to Investigate Modulators of Wnt and Shh Morphogen Gradients

The manuscript here provides a simple set of methods for analysing the secretion and diffusion of fluorescently tagged ligands in Xenopus. This provides a context for testing the ability of other proteins to modify ligand distribution and allowing experiments that may give insight into mechanisms regulating morphogen gradients.

This protocol describes a method to visualise ligands distributed across a field of cells. The ease of expressing exogenous proteins, together with the ...

Stem cell-like Xenopus Embryonic Explants to Study Early Neural Developmental Features In Vitro and In Vivo

Understanding the genetic programs underlying neural development is an important goal of developmental and stem cell biology. In the amphibian blastula, cells from the roof of the blastocoel are pluripotent. These cells can be isolated, and programmed to generate various tissues through manipulation of genes expression or induction by morphogens. In this manuscript protocols are described for the use of Xenopus laevis blastocoel roof explants as an assay system to investigate key in vivo and in vitro features of early neural development. These protocols allow the investigation of fate acquisition, cell migration behaviors, and cell autonomous and non-autonomous properties. The blastocoel roof explants can be cultured in a serum-free defined medium and grafted into host embryos. This transplantation into an embryo allows the investigation of the long-term lineage commitment, the inductive properties, and the behavior of transplanted cells in vivo. These assays can be exploited to investigate molecular mechanisms, cellular processes and gene regulatory networks underlying neural development. In the context of regenerative medicine, these assays provide a means to generate neural-derived cell types in vitro that could be used in drug screening.

Stem cell-like Xenopus Embryonic Explants to Study Early Neural Developmental Features In Vitro and In Vivo

In Xenopus embryos, cells from the roof of the blastocoel are pluripotent and can be programmed to generate various tissues. Here, we describe protocols to use amphibian blastocoel roof explants as an assay system to investigate key in vivo and in vitro features of early neural development.

Understanding the genetic programs underlying neural development is an important goal of developmental and stem cell biology. In the amphibian blastula, ...

Technique to Target Microinjection to the Developing Xenopus Kidney

The embryonic kidney of Xenopus&#160;laevis (frog), the pronephros, consists of a single nephron, and can be used as a model for kidney disease. Xenopus embryos are large, develop externally, and can be easily manipulated by microinjection or surgical procedures. In addition, fate maps have been established for early Xenopus embryos. Targeted microinjection into the individual blastomere that will eventually give rise to an organ or tissue of interest can be used to selectively overexpress or knock down gene expression within this restricted region, decreasing secondary effects in the rest of the developing embryo. In this protocol, we describe how to utilize established Xenopus fate maps to target the developing Xenopus kidney (the pronephros), through microinjection into specific blastomere of 4- and 8-cell embryos. Injection of lineage tracers allows verification of the specific targeting of the injection. After embryos have developed to stage 38 - 40, whole-mount immunostaining is used to visualize pronephric development, and the contribution by targeted cells to the pronephros can be assessed. The same technique can be adapted to target other tissue types in addition to the pronephros.

Technique to Target Microinjection to the Developing Xenopus Kidney

Here, we present a protocol to use fate maps and lineage tracers to target injections into individual blastomeres that give rise to the kidney of Xenopus laevis embryos.

The embryonic kidney of Xenopus&#160;laevis (frog), the pronephros, consists of a single nephron, and can be used as a model for kidney disease. Xenopus ...

In Vivo Imaging and Quantitation of the Host Angiogenic Response in Zebrafish Tumor Xenografts

Tumor angiogenesis is a key target of anti-cancer therapy and this method has been developed to provide a new model to study this process in vivo. A zebrafish xenograft is created by implanting mammalian tumor cells into the perivitelline space of two days-post-fertilization zebrafish embryos, followed by measuring the extent of the angiogenic response observed at an experimental endpoint up to two days post-implantation. The key advantage to this method is the ability to accurately quantitate the zebrafish host angiogenic response to the graft. This enables detailed examination of the molecular mechanisms as well as the host vs tumor contribution to the angiogenic response. The xenografted embryos can be subjected to a variety of treatments, such as incubation with potential anti-angiogenesis drugs, in order to investigate strategies to inhibit tumor angiogenesis. The angiogenic response can also be live-imaged in order to examine more dynamic cellular processes. The relatively undemanding experimental technique, cheap maintenance costs of zebrafish and short experimental timeline make this model especially useful for the development of strategies to manipulate tumor angiogenesis.

The aim of this method is to generate an in vivo model of tumor angiogenesis by xenografting mammalian tumor cells into a zebrafish embryo that has fluorescently-labelled blood vessels. By imaging the xenograft and associated vessels, a quantitative measurement of the angiogenic response can be obtained.

Tumor angiogenesis is a key target of anti-cancer therapy and this method has been developed to provide a new model to study this process in vivo. A ...

Cancer Research

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 ...

Live-cell Imaging and Quantitative Analysis of Embryonic Epithelial Cells in Xenopus laevis

Embryonic epithelial cells serve as an ideal model to study morphogenesis where multi-cellular tissues undergo changes in their geometry, such as changes in cell surface area and cell height, and where cells undergo mitosis and migrate. Furthermore, epithelial cells can also regulate morphogenetic movements in adjacent tissues1. A traditional method to study epithelial cells and tissues involve chemical fixation and histological methods to determine cell morphology or localization of particular proteins of interest. These approaches continue to be useful and provide "snapshots" of cell shapes and tissue architecture, however, much remains to be understood about how cells acquire specific shapes, how various proteins move or localize to specific positions, and what paths cells follow toward their final differentiated fate. High resolution live imaging complements traditional methods and also allows more direct investigation into the dynamic cellular processes involved in the formation, maintenance, and morphogenesis of multicellular epithelial sheets. Here we demonstrate experimental methods from the isolation of animal cap tissues from Xenopus laevis embryos to confocal imaging of epithelial cells and simple measurement approaches that together can augment molecular and cellular studies of epithelial morphogenesis.

Live-cell Imaging and Quantitative Analysis of Embryonic Epithelial Cells in Xenopus laevis

Xenopus embryonic epithelia are an ideal model system to study cell behaviors such as polarity development and shape change during epithelial morphogenesis. Traditional histology of fixed samples is increasingly being complemented by live-cell confocal imaging. Here we demonstrate methods to isolate frog tissues and visualize live epithelial cells and their cytoskeleton using live-cell confocal microscopy.

Embryonic epithelial cells serve as an ideal model to study morphogenesis where multi-cellular tissues undergo changes in their geometry, such as changes ...

Biology

Utilizing Larval Zebrafish for Efficient In Vivo Tumor Studies

A Simple, Rapid, and Effective Method for Tumor Xenotransplantation Analysis in Transparent Zebrafish Embryos

In vivo studies of tumor behavior are a staple of cancer research; however, the use of mice presents significant challenges in cost and time. Here, we present larval zebrafish as a transplant model that has numerous advantages over murine models, including ease of handling, low expense, and short experimental duration. Moreover, the absence of an adaptive immune system during larval stages obviates the need to generate and use immunodeficient strains. While established protocols for xenotransplantation in zebrafish embryos exist, we present here an improved method involving embryo staging for faster transfer, survival analysis, and the use of flow cytometry to assess disease burden. Embryos are staged to facilitate rapid cell injection into the yolk of the larvae and cell marking to monitor the consistency of the injected cell bolus. After injection, embryo survival analysis is assessed up to 7 days post injection (dpi). Finally, disease burden is also assessed by marking transferred cells with a fluorescent protein and analysis by flow cytometry. Flow cytometry is enabled by a standardized method of preparing cell suspensions from zebrafish embryos, which could also be used in establishing the primary culture of zebrafish cells. In summary, the procedure described here allows a more rapid assessment of the behavior of tumor cells in vivo with larger numbers of animals per study arm and in a more cost-effective manner.

Author Spotlight: Advancing Cancer Therapeutics Using Tumor Xenotransplantation in Zebrafish

We describe a protocol for xenotransplantation into the yolk of transparent zebrafish embryos that is optimized by a simple, rapid staging method. Post-injection analyses include survival and assessing the disease burden of xenotransplanted cells by flow cytometry.

In vivo studies of tumor behavior are a staple of cancer research; however, the use of mice presents significant challenges in cost and time. Here, we ...

Whole-mount Immunohistochemical Analysis for Embryonic Limb Skin Vasculature: a Model System to Study Vascular Branching Morphogenesis in Embryo

Whole-mount immunohistochemical analysis for imaging the entire vasculature is pivotal for understanding the cellular mechanisms of branching morphogenesis. We have developed the limb skin vasculature model to study vascular development in which a pre-existing primitive capillary plexus is reorganized into a hierarchically branched vascular network. Whole-mount confocal microscopy with multiple labelling allows for robust imaging of intact blood vessels as well as their cellular components including endothelial cells, pericytes and smooth muscle cells, using specific fluorescent markers. Advances in this limb skin vasculature model with genetic studies have improved understanding molecular mechanisms of vascular development and patterning. The limb skin vasculature model has been used to study how peripheral nerves provide a spatial template for the differentiation and patterning of arteries. This video article describes a simple and robust protocol to stain intact blood vessels with vascular specific antibodies and fluorescent secondary antibodies, which is applicable for vascularized embryonic organs where we are able to follow the process of vascular development.

We introduce a whole-mount immunohistochemistry and laser scanning confocal microscopy with multiple labelling for analyzing intricate vascular network formation in mouse embryonic limb skin.

Whole-mount immunohistochemical analysis for imaging the entire vasculature is pivotal for understanding the cellular mechanisms of branching ...

Assessing Primary Neurogenesis in Xenopus Embryos Using Immunostaining

Primary neurogenesis is a dynamic and complex process during embryonic development that sets up the initial layout of the central nervous system. During this process, a portion of neural stem cells undergo differentiation and give rise to the first populations of differentiated primary neurons within the nascent central nervous system. Several vertebrate model organisms have been used to explore the mechanisms of neural cell fate specification, patterning, and differentiation. Among these is the African clawed frog, Xenopus, which provides a powerful system for investigating the molecular and cellular mechanisms responsible for primary neurogenesis due to its rapid and accessible development and ease of embryological and molecular manipulations. Here, we present a convenient and rapid method to observe the different populations of neuronal cells within Xenopus central nervous system. Using antibody staining and immunofluorescence on sections of Xenopus embryos, we are able to observe the locations of neural stem cells and differentiated primary neurons during primary neurogenesis.

Assessing Primary Neurogenesis in Xenopus Embryos Using Immunostaining

This article presents a convenient and rapid method for visualizing different neuronal cell populations in the central nervous system of Xenopus embryos using immunofluorescent staining on sections.

Primary neurogenesis is a dynamic and complex process during embryonic development that sets up the initial layout of the central nervous system. During ...

Visualization and Quantitative Analysis of Embryonic Angiogenesis in Xenopus tropicalis

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