We thank Peter Johnson of the Georgetown University Microscopy Core for assistance with imaging the zebrafish embryos. The Microscopy &amp; Imaging Shared Resource and the Zebrafish Shared Resource are partially supported by NIH/NCI grant P30-CA051008. This work was also supported by NIH/NCI CA71508 (AW) and CA177466 (AW).

Ethics Statement: Zebrafish embryos were generated according to an approved IACUC protocol. These experiments were carried out in compliance with recommendations by the Georgetown University Animal Care and Use Committee.
1. Organize Embryos for Injection and Create Stock Solutions
<ol>
	<li>Generate requisite zebrafish larvae to assess cancer cell vascular invasion.
	<ol>
		<li>Set up pair-wise or group in-cross mating with Tg(kdrl:grcfp)zn1;mitfab692;ednrb1b140 fish. 
		NOTE: We generated Tg(kdrl:grcfp)zn1;mitfab692;ednrb1b140 zebrafish, by crossing Tg(kdrl:grcfp)zn125, which express green reef coral fluorescent protein in endothelial cells, with a line that lacks pigment cells, mitfab692;ednrb1b140, developed at the Zebrafish International Resource Center.</li>
		<li>Collect eggs, clean, and remove unfertilized or deformed embryos the next day22.</li>
		<li>Incubate embryos at 28.5 &#176;C until ready for injection with cancer cells, to occur when the zebrafish embryos are 2-days post-fertilization (2 dpf).</li>
	</ol>
	</li>
	<li>Make injection plates.
	<ol>
		<li>Melt 25 ml of 1.5% agarose in dH2O for each plate.</li>
		<li>Pour 12 ml of the agarose into a 100 mm x 15 mm petri dish and let it harden.</li>
		<li>Re-melt then pour the remaining agarose in the plate.</li>
		<li>Immediately position a cut glass mold (3 mm x 7.2 cm wide x 7.5 cm long) so that it is at a 30 degree angle to the agarose and positioned in the center of the plate. 
		NOTE: This will create a steep 60&#176; wall and a 30&#176; sloped ramp.</li>
		<li>Tape the glass mold in place and let the agarose harden.</li>
		<li>Gently remove the glass mold and be careful not to tear the agarose. 
		NOTE: Molds can be stored in dH2O at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Equilibrate injection plate with fish water (0.3 g/L sea salt).
	<ol>
		<li>Rinse plate twice with distilled water.</li>
		<li>Equilibrate plate by adding 10 ml of fish water to the plate and place on shaker for 10 min.</li>
		<li>Equilibrate plate a second time with fish water.</li>
	</ol>
	</li>
	<li>Split 2-day post-fertilization (dpf) embryos into injection groups by transferring them into dishes containing fish water22.</li>
	<li>Prepare recovery dishes for each group to utilize after injection. Ensure that the recovery dish contains 10 ml fish water, plus penicillin (25 &#181;g/ml) and streptomycin (50 &#181;g/ml).</li>
	<li>Prepare 2x tricaine solution by adding 4 ml of buffered tricaine stock (4 mg/ml,10 mM Tris, pH 7) to 50 ml of fish water plus penicillin and streptomycin.</li>
	<li>Dissolve 15 mg low melting-point agarose in 10 ml of 2x tricaine solution to generate a mounting anesthetic medium that will immobilize live embryos for imaging. 
	NOTE: Mounting medium consists of 1.5 % agarose.</li>
	<li>Pull microinjection needles.
	<ol>
		<li>Place glass capillary tubing in a vertical pipette puller. Pull long tapered pipettes using 20 mAmp current and a 2-coil heating element.</li>
	</ol>
	</li>
</ol>
2. Labeling Cancer Cells with Lipophilic Fluorescent Dye
<ol>
	<li>Maintain cancer cells in their recommended culture conditions. 
	NOTE: These lines were maintained in DMEM + 10% FBS: BT-474, MCF-7, MDA-MB-231, MDA-MB-468, and SK-BR-3. These lines were maintained in RPMI + 10% FBS: HCC 1806 and T-47D. All cell lines were maintained at 37 oC and 5% CO2 during incubation.</li>
	<li>Generate a single-cell suspension by dissociating an adherent culture of cancer cells.
	<ol>
		<li>Wash cells first with PBS and then treat with 0.05% trypsin-EDTA solution. 
		NOTE: Trypsin exposure time will depend on the cell line.</li>
		<li>Neutralize the trypsin solution with serum-containing cell culture media after the cells detach.</li>
	</ol>
	</li>
	<li>Centrifuge the trypsin-neutralized cell suspension for 5 min at 200 x g, then resuspend the cell pellet in fresh culture media for cell counting.</li>
	<li>Count the cell suspension using an automated counter and prepare 1 million cells in 200 &#181;l of cell culture media.
	<ol>
		<li>Verify cell viability with trypan blue dye exclusion before injection into zebrafish embryos. 
		NOTE: Only viable cell populations should be injected into zebrafish embryos, as injection of dead cells will not reflect true vascular invasion.</li>
	</ol>
	</li>
	<li>Add 2 &#181;l of red lipophilic dye to the cancer cell suspension for a 1:100 dilution, mix well, and then incubate the mixture at 37 &#176;C for 20 min. 
	NOTE: Concentration of the dye and labeling time may need to be optimized for each cell line.</li>
	<li>Following the incubation, add 1 ml of fresh media to the tube and then centrifuge for 5 min at 200 x g.</li>
	<li>Wash away residual fluorescent dye from the cancer cells.
	<ol>
		<li>Aspirate the supernatant from the cell pellet, resuspend the pellet in 1 ml of fresh culture media, and centrifuge for 5 min at 200 x g.</li>
		<li>Repeat the washing step a second time: aspirate the supernatant, resuspend the cell pellet in 1 ml of fresh media and then centrifuge again for 5 min at 200 x g.</li>
		<li>Repeat the washing step third time: aspirate the supernatant, resuspend the cell pellet in 1 ml of fresh media and then centrifuge again for 5 min at 200 x g.</li>
	</ol>
	</li>
	<li>Aspirate the supernatant and resuspend the cell pellet containing 1 million labeled cancer cells in 500 &#181;l of fresh media. 
	NOTE: 0.5 mM EDTA can be added to the media to prevent cell clumping.</li>
</ol>
3. Injecting Cancer Cells into the Pre-cardiac Sinus of Zebrafish Embryos
<ol>
	<li>Attach microinjection dispense system to a pressurized air source and turn on the microinjection dispense system power source.
	<ol>
		<li>Test pressure by depressing the foot pedal. A brief pulse of air should emit from the needle holder.</li>
	</ol>
	</li>
	<li>Equilibrate the injection plates twice with the 2x tricaine solution.
	<ol>
		<li>For each equilibration step, add 20 ml of 2x tricaine solution to the injection plate and place on shaker for 10 min.</li>
	</ol>
	</li>
	<li>Use plastic pipette to transfer a group of embryos to a small dish containing the 2x tricaine solution.</li>
	<li>Backfill the microinjection injection needle with cancer cells using a gel-loading pipet tip.
	<ol>
		<li>Place the needle in an electrode storage jar with the pointed end facing down so cells settle near the tip.</li>
	</ol>
	</li>
	<li>Transfer 20 - 30 anesthetized embryos to an injection plate by collecting the embryos with plenty of 2x tricaine in a plastic pipette.
	<ol style="margin-left: 40px;">
		<li>Allow embryos to settle in the tip of the pipette.</li>
		<li>Gently expel embryos into the trough of the injection plate, spreading the embryos along the length of the trough.</li>
		<li>Align embryos with heads facing up and bellies facing the steep wall of the trough. 
		NOTE: Tricaine solution should cover both the trough length and the flat agarose surface, with embryos only residing in the trough. Embryos are now ready for injection.</li>
	</ol>
	</li>
	<li>Inject 50 - 100 cancer cells (2 - 5 nl) into the precardiac sinus of the zebrafish embryos using the microinjection dispense system.
	<ol>
		<li>Attach the needle to the needle holder of a micromanipulator.</li>
		<li>Position the injection plate under the stereoscope with the 60 degree wall to the left and focus on the top embryo at 25x magnification.</li>
		<li>Position the micromanipulator so that, when extended, the needle will pierce the embryo.</li>
		<li>Extend the needle by eye until it is nearly touching the embryo.</li>
		<li>Looking under the microscope, align the needle so that it will pierce the embryo upon further extension.</li>
		<li>Pierce the embryo through the yolk sac placing the tip just at, but not in, the pre-cardiac sinus.</li>
		<li>Inject cells by depressing the foot pedal. The force of the injection expels the cells into the cardiac sinus. Retract the needle.</li>
		<li>Using the right hand, extend and retract the injection needle. With the left hand, make fine adjustments to position next embryo.</li>
		<li>Return the needle to the electrode storage jar while setting up to inject another plate.</li>
	</ol>
	</li>
	<li>Transfer the embryos to the recovery dish once the entire plate is injected.
	<ol>
		<li>Tilt the injection plate to pool the embryos at the bottom, washing any remaining embryos out of the trough, and collecting them with the plastic pipette.</li>
		<li>Allow the embryos to settle in the bottom of the pipette.</li>
		<li>Transfer the embryos to the recovery dish in a minimal volume of tricaine.</li>
	</ol>
	</li>
	<li>Incubate recovery dish at 28 &#176;C for 1 hr.
	<ol>
		<li>Separate viable zebrafish embryos from dead embryos and other debris.</li>
	</ol>
	</li>
	<li>Incubate dish at 33 &#176;C until ready for scoring, typically 24 - 96 hr. 
	NOTE: This temperature is determined as a compromise between 37 &#176;C, the ideal temperature for cancer cells, and 28.5 &#176;C, the ideal temperature for zebrafish.</li>
</ol>
4. Scoring Extravasation
<ol>
	<li>Anesthetize the batch of embryos to be scored by placing them in a dish with tricaine solution.</li>
	<li>Place an anesthetized larvae on a depression microscopy slide in a drop of tricaine.
	<ol>
		<li>Orient larvae laterally for optimal imaging of the caudal region.</li>
	</ol>
	</li>
	<li>Count the number of cancer cells that have successfully invaded out of the vasculature by focusing up and down through the tail region to clearly discern intact cells. 
	NOTE: It is best to have at least two individuals involved in this process, where the individual scoring the fish is blind to the experimental condition being assessed.
	<ol>
		<li>Score larvae on a compound fluorescence microscope with the 10x objective lens. Use the 20x objective for any difficult calls.</li>
	</ol>
	</li>
</ol>
5. Mounting Embryos onto Slides and Subsequent Fluorescence Imaging
<ol>
	<li>Melt 1.5% agarose/tricaine solution and bring to 37 &#176;C.</li>
	<li>Anesthetize the embryo to be imaged by placing it in tricaine solution.</li>
	<li>Transfer the embryo in a drop of tricaine solution to the imaging surface. Optionally use a glass-bottom dish or microscope slide.</li>
	<li>Use a glass pipette to remove the excess tricaine solution, retaining the embryo on the imaging surface.</li>
	<li>Overlay one drop of melted agarose solution over the embryo.</li>
	<li>Quickly, before the agarose polymerizes, use a delicate tool, like an eyelash brush, to orient the embryo laterally for imaging, giving extra care to ensure the embryo is flattened along the imaging surface.</li>
	<li>Submerge the now polymerized agarose drop under tricaine solution.</li>
	<li>Subject the live zebrafish embryo to microscopic imaging.</li>
</ol>
6. Modification: Injecting Cancer Cells Into the Yolk Sac of Zebrafish Embryos
<ol>
	<li>Prepare the microinjection dispense system and injection plates as previously described in section (3) of this protocol.</li>
	<li>Label two cell populations with contrasting fluorescent dyes as previously described in section (2) of this protocol.</li>
	<li>Inject 5 - 10 nl of 2 x 10^7 cells/ml into the yolk sac. Keep the injection volume constant to inject identical cell numbers (100 - 200 cancer cells) from each cell population 
	NOTE: One may use transparent zebrafish embryos lacking fluorescent vasculature for this assay. Passive entry of particles into the vasculature can be controlled for by injecting fluorescent beads (&#60; 10 &#181;m) or, alternatively, a cell line that does not intravsate.</li>
	<li>Recover the injected embryos as previously described in section (3) of this protocol and then screen for successful injections.
	<ol>
		<li>Use a stereoscope to screen and transfer viable embryos that were successfully injected to a new dish. 
		NOTE: All embryos should have a consistently sized mass of cells located in the yolk. Embryos are discarded if the mass size differs or if any cells are located outside of the yolk.</li>
		<li>Transfer the viable embryos to a new dish if cancer cells are clearly seen in the yolk sac.</li>
	</ol>
	</li>
	<li>Incubate dish at 33 &#176;C until ready for scoring, typically 24 - 48 hr. 
	NOTE: This temperature is determined as a compromise between 37 &#176;C, the ideal temperature for cancer cells, and 28.5 &#176;C, the ideal temperature for zebrafish.</li>
	<li>To score intravasation, follow the guidelines described in section (4) of this protocol, but instead count the number of cancer cells that have successfully invaded into the vasculature of the caudal region.</li>
</ol>

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The authors have no conflicts of interest to disclose.

This technique utilizes the zebrafish model to efficiently test the vascular invasive ability of cancer cells (see Figure 1). Here we applied the technique to a panel of breast cancer cell lines in order to provide a baseline onto which other investigators can then build their own studies (see Table 1; Figures 2 - 3). The observation that MDA-MB-231 cells readily invaded into the caudal region of zebrafish embryos would make this cell line ideal for testing agents that m...

Metastatic disease is a major cause of cancer mortality and many mechanisms that enable cancer cell dissemination remain to be discovered1. In order for a cancer cell to successfully metastasize, it must first invade through the stroma that surrounds a primary tumor, enter (intravasate) into the circulatory system, survive in transit, exit (extravasate) from the circulation, and lastly establish a viable colony at the distant organ site2. Intravasation and extravasation are thus crucial steps in the metastatic cascade, yet every cancer cell is not inherently adept at disrupting and migrating through endothelial junctions3. In fact, there are a series of unique selection pressures that surround cancer cell vascular invasion and the process can be further influenced by a variety of endogenous and exogenous factors4. For these reasons, techniques that probe the aggressive behavior of advanced stage cancer often focus on vascular invasive ability as a means to predict metastatic spread.
Various model systems exist to facilitate the study of cancer cell vascular invasion in vitro. The most used in vitro assays involve either transwell systems to assess cancer cell migration through an endothelial barrier5 or Electric Cell-Substrate Impedance Sensing (ECIS) technology to monitor the real-time disruption of an intact endothelial monolayer by cancer cells6. These assays typically lack the fluid dynamics and stromal factors that would otherwise impact cancer cell attachment to an endothelial wall. This issue is somewhat circumvented by perfusable vascular networks that arise from the 3D culture of endothelia with supporting stromal cells, and these 3D microfluidic systems now represent the forefront of current in vitro options7,8. Still, these approaches omit the robust microenvironment of a functional circulatory system and therefore only in part substitute for in vivo models.
The most widely used in vivo model of vascular invasion is the mouse, in which experimental metastasis assays are commonly performed because they occur on relatively short timescales and are generally indicative of metastatic ability9. These assays involve direct injection of cancer cells into circulation and therefore model the end stages of metastasis, namely extravasation and cancer cell colonization of organs. The experimental metastasis assays differ based on the site of cancer cell injection and the organs ultimately analyzed. In the first assay type, cancer cells are injected into the tail vein of mice and cancer cell seeding in the lungs is monitored10,11. The second assay involves performing intracardiac injections to direct metastatic seeding toward the bone microenvironment12-14, but also the brain15. In the third assay, cancer cells are injected into the spleen in order to permit colonization of the liver16 whereas the fourth delivery route into the carotid artery carries cancer cells to the brain17,18. Irrespective of the cancer cell delivery method, organ colonization is the accepted experimental endpoint and is generally determined via luminescence, histology, or PCR-based techniques. Despite the physiologic advantages of conducting experimental metastasis assays within a murine host, these experiments still require weeks to months to complete and analyze.
The zebrafish (Danio rerio) model has recently emerged as a new system to study cancer progression19,20, and allows for the assessment of cancer cell vascular invasion within a functional circulatory system over a much shorter timescale when compared with mice21-24. The method utilizes a transparent zebrafish strain that has its endothelia tagged with a green reef coral fluorescent protein reporter driven by the kdrl promoter, the zebrafish receptor for vascular endothelial growth factor25. In the assay, cancer cells are labeled with a red fluorescent marker and injected into the precardiac sinus of 2-day old embryos. Anywhere between 48 to 96 hr after the injection, cancer cells that have invaded out of the vasculature and into the caudal region of embryos can be scored efficiently on a fluorescent microscope. Here we apply the technique to a panel of commonly used human breast cancer cell lines to demonstrate stark differences in their vascular invasive ability. Furthermore, we demonstrate that changing the injection site to the embryo yolk sac allows for the study of heterogeneous cell interactions, as cancer cell populations can be differentially labeled with fluorescent dyes and injected into zebrafish embryos lacking fluorescent vasculature. In this latter assay, cancer cells that have invaded the yolk and intravasated into the vasculature are scored in the caudal region 24 to 48 hr after injection. Due to the effectiveness and convenience of this model, zebrafish are increasingly employed to rapidly test the vascular invasive ability of cancer cells under a physiologic setting.

<table><tbody><tr><td>0.05% Trypsin-EDTA</td><td>Life Technologies</td><td>25300-054</td><td></td></tr><tr><td>100 mm Dishes</td><td>Corning Incorporated</td><td>3160-100</td><td></td></tr><tr><td>5 3/4&quot; Disposable Pastur Pipets, borosilicate Glass</td><td>Fisher Brand</td><td>13-678-20B</td><td></td></tr><tr><td>60 mm Dish</td><td>Corning Incorporated</td><td>3160-60</td><td></td></tr><tr><td>Agarose, Low Melting</td><td>Fisher</td><td>BP165-25</td><td></td></tr><tr><td>Agarose, Molecular Grade</td><td>Bioline</td><td>BIO-41026</td><td></td></tr><tr><td>Capillary Glass, Standard, 1.2 mm x 0.68 mm, 4&quot;</td><td>A-M Systems, Inc</td><td>627000</td><td></td></tr><tr><td>David Kopf 700C Vertical Pipette Puller</td><td>Hofstra Group</td><td>3600</td><td></td></tr><tr><td>DMEM</td><td>Life Technologies</td><td>11995-065</td><td></td></tr><tr><td>Electrode Storage Jar, 1.0 mm</td><td>World Precision Instruments, Inc</td><td>E210</td><td></td></tr><tr><td>Ethyl 3-aminobenzoate methanesulfonate salt (Tricaine, MS-222)</td><td>Fluka</td><td>A5040</td><td></td></tr><tr><td>Eyelash Brush</td><td>Ted Pella, Inc</td><td>113</td><td></td></tr><tr><td>Fetal Bovine Serum, Heat Inactivated</td><td>Omega Scientific</td><td>FB-12</td><td></td></tr><tr><td>Fisherbrand Transfer Pipettes</td><td>ThermoFisher Scientific</td><td>13-711-7M</td><td></td></tr><tr><td>Gel Loading Pipet Tips</td><td>Fisher Brand</td><td>02-707-181</td><td></td></tr><tr><td>Glass Bottom Dishes (12.0 mm)</td><td>ThermoFisher Scientific</td><td>150680</td><td></td></tr><tr><td>Glass Depression Slide</td><td>VWR</td><td>470005-634</td><td></td></tr><tr><td>Instant Ocean Salt, Sea Salt</td><td>Pentair</td><td>IS50</td><td></td></tr><tr><td>Latex Rubber Bulbs, 2 ml, Pack of 72</td><td>Heathrow Scientific</td><td>HS20622B</td><td></td></tr><tr><td>SP8 Confocal Microscope</td><td>Leica</td><td></td><td></td></tr><tr><td>Micromanipulator</td><td>Narishige</td><td></td><td></td></tr><tr><td>Eclipse E600</td><td>Nikon</td><td></td><td></td></tr><tr><td>PBS</td><td>Life Technologies</td><td>10010-023</td><td></td></tr><tr><td>Penicillin-G Potassium</td><td>Fisher Biotech</td><td>BP914-100</td><td></td></tr><tr><td>Petri Plates, 100 mm x 15 mm</td><td>Fisher Brand</td><td>&amp;nbsp;FB0875713</td><td></td></tr><tr><td>Picospritzer II</td><td>General Valve Corporation</td><td></td><td></td></tr><tr><td>RPMI 1640 Medium&amp;nbsp;</td><td>Life Technologies</td><td>11875-093</td><td></td></tr><tr><td>Streptomycin Sulfate</td><td>Fisher Biotech</td><td>BP910-50</td><td></td></tr><tr><td>Vybrant DiI</td><td>ThermoFisher Scientific</td><td>V22885</td><td></td></tr><tr><td>Vybrant DiO</td><td>ThermoFisher Scientific</td><td>V22886</td><td></td></tr><tr><td>Zebrafish&amp;nbsp;</td><td>Georgetown Zebrafish Shared Resources</td><td></td><td></td></tr><tr><td>Cell lines were maintained in DMEM + 10% FBS, with the expection of BT-474 and HCC18-6 cells, which were mantained in RPMI + 10% FBS.</td><td></td><td></td><td></td></tr></tbody></table>

testing the vascular invasive ability of cancer cells in zebrafish (danio rerio)

Cancer cell vascular invasion and extravasation is a hallmark of metastatic progression. Traditional in vitro models of cancer cell invasion of endothelia typically lack the fluid dynamics that invading cells are otherwise exposed to in vivo. However, in vivo systems such as mouse models, though more physiologically relevant, require longer experimental timescales and present unique challenges associated with monitoring and data analysis. Here we describe a zebrafish assay that seeks to bridge this technical gap by allowing for the rapid assessment of cancer cell vascular invasion and extravasation. The approach involves injecting fluorescent cancer cells into the precardiac sinus of transparent 2-day old zebrafish embryos whose vasculature is marked by a contrasting fluorescent reporter. Following injection, the cancer cells must survive in circulation and subsequently extravasate from vessels into tissues in the caudal region of the embryo. Extravasated cancer cells are efficiently identified and scored in live embryos via fluorescence imaging at a fixed timepoint. This technique can be modified to study intravasation and/or competition amongst a heterogeneous mixture of cancer cells by changing the injection site to the yolk sac. Together, these methods can evaluate a hallmark behavior of cancer cells and help uncover mechanisms indicative of malignant progression to the metastatic phenotype.

Here we tested the vascular invasive ability of commonly used breast cancer cell lines in a zebrafish embryo model (Figure 1). Rigorous criteria were employed in scoring extravasation for these different cell lines, where positive events were only counted if the cancer cells had clearly extravasated, this being done chiefly to limit any false-positives that could arise from scoring cellular debris.
Our analysis ...

Watch this Scientific Journal Video about Testing the Vascular Invasive Ability of Cancer Cells in Zebrafish Danio Rerio at JoVE.com

Testing the Vascular Invasive Ability of Cancer Cells in Zebrafish Danio Rerio

This method utilizes zebrafish embryos to efficiently test the vascular invasive ability of cancer cells. Fluorescent cancer cells are injected into the precardiac sinus or yolk sac of developing embryos. Cancer cell vascular invasion and extravasation is assessed via fluorescence microscopy of the tail region 24 to 96 hr later.

Testing the Vascular Invasive Ability of Cancer Cells in Zebrafish (Danio Rerio)

Cancer cell vascular invasion and extravasation is a hallmark of metastatic progression.  Traditional in vitro models of cancer cell invasion of ...

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Research

JoVE Journal

Cancer Research

11.9K Views.  Georgetown University. This method utilizes zebrafish embryos to efficiently test the vascular invasive ability of cancer cells. Fluorescent cancer cells are injected into the precardiac sinus or yolk sac of developing embryos. Cancer cell vascular invasion and extravasation is assessed via fluorescence microscopy of the tail region 24 to 96 hr later.

Video: Testing the Vascular Invasive Ability of Cancer Cells in Zebrafish Danio Rerio

Mosaic Zebrafish Transgenesis for Functional Genomic Analysis of Candidate Cooperative Genes in Tumor Pathogenesis

Comprehensive genomic analysis has uncovered surprisingly large numbers of genetic alterations in various types of cancers. To robustly and efficiently identify oncogenic &#8220;drivers&#8221; among these tumors and define their complex relationships with concurrent genetic alterations during tumor pathogenesis remains a daunting task. Recently, zebrafish have emerged as an important animal model for studying human diseases, largely because of their ease of maintenance, high fecundity, obvious advantages for in vivo imaging, high conservation of oncogenes and their molecular pathways, susceptibility to tumorigenesis and, most importantly, the availability of transgenic techniques suitable for use in the fish. Transgenic zebrafish models of cancer have been widely used to dissect oncogenic pathways in diverse tumor types. However, developing a stable transgenic fish model is both tedious and time-consuming, and it is even more difficult and more time-consuming to dissect the cooperation of multiple genes in disease pathogenesis using this approach, which requires the generation of multiple transgenic lines with overexpression of the individual genes of interest followed by complicated breeding of these stable transgenic lines. Hence, use of a mosaic transient transgenic approach in zebrafish offers unique advantages for functional genomic analysis in vivo. Briefly, candidate transgenes can be coinjected into one-cell-stage wild-type or transgenic zebrafish embryos and allowed to integrate together into each somatic cell in a mosaic pattern that leads to mixed genotypes in the same primarily injected animal. This permits one to investigate in a faster and less expensive manner whether and how the candidate genes can collaborate with each other to drive tumorigenesis. By transient overexpression of activated ALK in the transgenic fish overexpressing MYCN, we demonstrate here the cooperation of these two oncogenes in the pathogenesis of a pediatric cancer, neuroblastoma that has resisted most forms of contemporary treatment.

The goal of this study is to demonstrate how the mosaic transgenesis strategy can be used in zebrafish to rapidly and efficiently assess the relative contributions of multiple oncogenes in tumor initiation and progression in vivo.

Comprehensive genomic analysis has uncovered surprisingly large numbers of genetic alterations in various types of cancers.  To robustly and efficiently ...

Developmental Biology

Invasive Behavior of Human Breast Cancer Cells in Embryonic Zebrafish

In many cases, cancer patients do not die of a primary tumor, but rather because of metastasis. Although numerous rodent models are available for studying cancer metastasis in vivo, other efficient, reliable, low-cost models are needed to quickly access the potential effects of (epi)genetic changes or pharmacological compounds. As such, we illustrate and explain the feasibility of xenograft models using human breast cancer cells injected into zebrafish embryos to support this goal. Under the microscope, fluorescent proteins or chemically labeled human breast cancer cells are transplanted into transgenic zebrafish embryos, Tg (fli:EGFP), at the perivitelline space or duct of Cuvier (Doc) 48 h after fertilization. Shortly afterwards, the temporal-spatial process of cancer cell invasion, dissemination, and metastasis in the living fish body is visualized under a fluorescent microscope. The models using different injection sites, i.e., perivitelline space or Doc are complementary to one another, reflecting the early stage (intravasation step) and late stage (extravasation step) of the multistep metastatic cascade of events. Moreover, peritumoral and intratumoral angiogenesis can be observed with the injection into the perivitelline space. The entire experimental period is no more than 8 days. These two models combine cell labeling, micro-transplantation, and fluorescence imaging techniques, enabling the rapid evaluation of cancer metastasis in response to genetic and pharmacological manipulations.

Here, we describe xenograft zebrafish models using two different injection sites, i.e., perivitelline space and duct of Cuvier, to investigate the invasive behavior and to assess the intravasation and extravasation potential of human breast cancer cells, respectively.

In many cases, cancer patients do not die of a primary tumor, but rather because of metastasis. Although numerous rodent models are available for studying ...

Transplantation of Zebrafish Pediatric Brain Tumors into Immune-competent Hosts for Long-term Study of Tumor Cell Behavior and Drug Response

Tumor cell transplantation is an important technique to define the mechanisms controlling cancer cell growth, migration, and host response, as well as to assess potential patient response to therapy. Current methods largely depend on using syngeneic or immune-compromised animals to avoid rejection of the tumor graft. Such methods require the use of specific genetic strains that often prevent the analysis of immune-tumor cell interactions and/or are limited to specific genetic backgrounds. An alternative method in zebrafish takes advantage of an incompletely developed immune system in the embryonic brain before 3 days, where tumor cells are transplanted for use in short-term assays (i.e., 3 to 10 days). However, these methods cause host lethality, which prevents the long-term study of tumor cell behavior and drug response. This protocol describes a simple and efficient method for the long-term orthotopic transplantation of zebrafish brain tumor tissue into the fourth ventricle of a 2-day-old immune-competent zebrafish. This method allows: 1) long-term study of tumor cell behaviors, such as invasion and dissemination; 2) durable tumor response to drugs; and 3) re-transplantation of tumors for the study of tumor evolution and/or the impact of different host genetic backgrounds. In summary, this technique allows cancer researchers to assess engraftment, invasion, and growth at distant sites, as well as to perform chemical screens and cell competition assays over many months. This protocol can be extended to studies of other tumor types and can be used to elucidate mechanisms of chemoresistance and metastasis.

The transplantation of cancer cells is an important tool for the identification of cancer mechanisms and therapeutic responses. Current techniques depend on immune-incompetent animals. Here, we describe a method to transplant zebrafish tumor cells into immune-competent embryos for the long-term analysis of tumor cell behavior and in vivo drug responses.

Tumor cell transplantation is an important technique to define the mechanisms controlling cancer cell growth, migration, and host response, as well as to ...

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 ...

Drug Screening of Primary Patient Derived Tumor Xenografts in Zebrafish

Patient derived xenograft models are critical in defining how different cancers respond to drug treatment in an in vivo system. Mouse models are the standard in the field, but zebrafish have emerged as an alternative model with several advantages, including the ability for high-throughput and low-cost drug screening. Zebrafish also allow for in vivo drug screening with large replicate numbers that were previously only obtainable with in vitro systems. The ability to rapidly perform large scale drug screens may open up the possibility for personalized medicine with rapid translation of results back to clinic. Zebrafish xenograft models could also be used to rapidly screen for actionable mutations based on tumor response to targeted therapies or to identify new anti-cancer compounds from large libraries. The current major limitation in the field has been quantifying and automating the process so that drug screens can be done on a larger scale and be less labor-intensive. We have developed a workflow for xenografting primary patient samples into zebrafish larvae and performing large scale drug screens using a fluorescence microscope equipped imaging unit and automated sampler unit. This method allows for standardization and quantification of engrafted tumor area and response to drug treatment across large numbers of zebrafish larvae. Overall, this method is advantageous over traditional cell culture drug screening as it allows for growth of tumor cells in an in vivo environment throughout drug treatment, and is more practical and cost-effective than mice for large scale in vivo drug screens.

Zebrafish xenograft models allow for high-throughput drug screening and fluorescent imaging of human cancer cells in an in vivo microenvironment. We developed a workflow for large scale, automated drug screening on patient-derived leukemia samples in zebrafish using an automated fluorescence microscope equipped imaging unit.

Patient derived xenograft models are critical in defining how different cancers respond to drug treatment in an in vivo system. Mouse models are the ...

In Vivo Targeting of Xenografted Human Cancer Cells with Functionalized Fluorescent Silica Nanoparticles in Zebrafish

Developing nanoparticles capable of detecting, targeting, and destroying cancer cells is of great interest in the field of nanomedicine. In vivo animal models are required for bridging the nanotechnology to its biomedical application. The mouse represents the traditional animal model for preclinical testing; however, mice are relatively expensive to keep and have long experimental cycles due to the limited progeny from each mother. The zebrafish has emerged as a powerful model system for developmental and biomedical research, including cancer research. In particular, due to its optical transparency and rapid development, zebrafish embryos are well suited for real-time in vivo monitoring of the behavior of cancer cells and their interactions with their microenvironment. This method was developed to sequentially introduce human cancer cells and functionalized nanoparticles in transparent Casper zebrafish embryos and monitor in vivo recognition and targeting of the cancer cells by nanoparticles in real time. This optimized protocol shows that fluorescently labeled nanoparticles, which are functionalized with folate groups, can specifically recognize and target metastatic human cervical epithelial cancer cells labeled with a different fluorochrome. The recognition and targeting process can occur as early as 30 min postinjection of the nanoparticles tested. The whole experiment only requires the breeding of a few pairs of adult fish and takes less than 4 days to complete. Moreover, zebrafish embryos lack a functional adaptive immune system, allowing the engraftment of a wide range of human cancer cells. Hence, the utility of the protocol described here enables the testing of nanoparticles on various types of human cancer cells, facilitating the selection of optimal nanoparticles in each specific cancer context for future testing in mammals and the clinic.

Described here is a method for utilizing zebrafish embryos to study the ability of functionalized nanoparticles to target human cancer cells in vivo. This method allows for the evaluation and selection of optimal nanoparticles for future testing in large animals and in clinical trials.

Developing nanoparticles capable of detecting, targeting, and destroying cancer cells is of great interest in the field of nanomedicine. In vivo animal ...

Generation of Zebrafish Larval Xenografts and Tumor Behavior Analysis

Zebrafish larval xenografts are being widely used for cancer research to perform in vivo and real-time studies of human cancer. The possibility of rapidly visualizing the response to anti-cancer therapies (chemo, radiotherapy, and biologicals), angiogenesis and metastasis with single cell resolution, places the zebrafish xenograft model as a top choice to develop preclinical studies.

The zebrafish larval xenograft assay presents several experimental advantages compared to other models, but probably the most striking is the reduction of size scale and consequently time. This reduction of scale allows single cell imaging, the use of a relatively low number of human cells (compatible with biopsies), medium-high-throughput drug screenings, but most importantly enables a significant reduction of the time of the assay. All these advantages make the zebrafish xenograft assay extremely attractive for future personalized medicine applications.

Many zebrafish xenograft protocols have been developed with a wide diversity of human tumors; however, a general and standardized protocol to efficiently generate zebrafish larval xenografts is still lacking. Here we provide a step-by-step protocol, with tips to generate xenografts and guidelines for tumor behavior analysis, whole-mount immunofluorescence, and confocal imaging quantification.

Here, we provide a step-by-step protocol, with tips to generate xenografts and guidelines for tumor behavior analysis, whole-mount immunofluorescence, and confocal imaging quantification.

Zebrafish larval xenografts are being widely used for cancer research to perform in vivo and real-time studies of human cancer. The possibility of rapidly ...

Zebrafish Model of Neuroblastoma Metastasis

Zebrafish has emerged as an important animal model to study human diseases, especially cancer. Along with the robust transgenic and genome editing technologies applied in zebrafish modeling, the ease of maintenance, high-yield productivity, and powerful live imaging altogether make the zebrafish a valuable model system to study metastasis and cellular and molecular bases underlying this process in vivo. The first zebrafish neuroblastoma (NB) model of metastasis was developed by overexpressing two oncogenes, MYCN and LMO1, under control of the dopamine-beta-hydroxylase (d&#946;h) promoter. Co-overexpressed MYCN and LMO1 led to the reduced latency and increased penetrance of neuroblastomagenesis, as well as accelerated distant metastasis of tumor cells. This new model reliably reiterates many key features of human metastatic NB, including involvement of clinically relevant and metastasis-associated genetic alterations; natural and spontaneous development of metastasis in vivo; and conserved sites of metastases. Therefore, the zebrafish model possesses unique advantages to dissect the complex process of tumor metastasis in vivo.

This paper introduces the method of developing, characterizing, and tracking in real-time the tumor metastasis in zebrafish model of neuroblastoma, specifically in the transgenic zebrafish line with overexpression of MYCN and LMO1, which develops metastasis spontaneously.

Zebrafish has emerged as an important animal model to study human diseases, especially cancer. Along with the robust transgenic and genome editing ...

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 ...

Zebrafish Xenograft Assay: A Method to Assess the Anti-cancer Efficacy of Therapeutics on Tumorigenicity in Zebrafish In Vivo

Source: Lee, J. Y et al.,<a href="https://www.jove.com/v/57490/preclinical-assessment-bioactivity-anticancer-coumarin-ot48-spheroids">Preclinical Assessment of the Bioactivity of the Anticancer Coumarin OT48 by Spheroids, Colony Formation Assays, and Zebrafish Xenografts.</a>&#160;J. Vis. Exp. (2018)
This video describes the zebrafish xenograft assay to assess in vivo tumor formation and study the effect of therapeutic agents.

Source: Lee, J. Y et al.,Preclinical Assessment of the Bioactivity of the Anticancer Coumarin OT48 by Spheroids, Colony Formation Assays, and Zebrafish ...