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. Author manuscript; available in PMC: 2022 May 15.
Published in final edited form as: Methods Mol Biol. 2021;2294:3–16. doi: 10.1007/978-1-0716-1350-4_1

Cancer cell invasion and metastasis in zebrafish models (Danio rerio).

Sarah Martinez Roth 1, Eric B Berens 1, Ghada M Sharif 1, Eric Glasgow 1, Anton Wellstein 1
PMCID: PMC9107928  NIHMSID: NIHMS1803786  PMID: 33742390

Abstract

Cancer cell vascular invasion and extravasation at metastatic sites are hallmarks of malignant progression of cancer and associated with poor disease outcome. Here we describe an in vivo approach to study the invasive ability of cancer cells into the vasculature and their hematogenous metastatic seeding in zebrafish (Danio rerio). In one approach, extravasation of fluorescently labeled cancer cells is monitored in zebrafish embryos whose vasculature is marked by a contrasting fluorescent reporter. After injection into the precardiac sinus of 2-day old embryos cancer cells can extravasate from the vasculature into tissues over the next few days. Extravasated cancer cells are identified and counted in live embryos via fluorescence microscopy. In a second approach intravasation of cancer cells can be evaluated by changing their injection site to the yolk sac of zebrafish embryos. In addition to monitoring the impact of drivers of malignant progression, candidate inhibitors can be studied in this in vivo model system for their efficacy as well as their toxicity for the host.

Keywords: Cancer, Extravasation, Zebrafish, Embryo, Fluorescence, Vascular Invasion, Metastasis

1. Introduction

Metastatic progression accounts for the majority of cancer related deaths, however the mechanisms underlying this complex process differ between cancer types and remains poorly understood [1]. The migration of cancer cells from primary tumors to distant tissues involves a multi-step process named the invasion-metastasis cascade. The process begins with the local invasion of tumor cells into the surrounding tissues. These cells then intravasate into the circulatory system and survive hematogenous transport. Cancer cells arrest in the microvessels of distant tissues and extravasate through the vascular lining into the parenchyma to form metastatic seeds. Proliferation of these colonies lead to clinically detectable lesions [2]. Crucial rate-limiting steps in the metastatic cascade include intravasation and extravasation of cancer cells. Cancer cells disrupt and migrate through endothelial junctions whilst also interacting with other circulating cells such as leukocytes and platelets that contribute to this process [3]. The tumor microenvironment (TME) also continually changes over the course of cancer progression, emphasizing the need to consider the TME’s role in metastasis [4]. Developing an animal model that allows the tracking of single cells or cell subpopulations is essential to understanding the key components of the invasion-metastasis cascade.

Currently, there are various in-vitro model systems that are employed to study cancer cell vascular invasion. The most common of in-vitro assays include transwell assays to observe cancer migration through endothelial barriers [5], and electric cell-substrate impendance sensing (ECIS) assay to assess the disruption of an intact monolayer by cancer cells in real-time over a time lapse [6, 7]. However, there are limitations in these existing in-vitro assays, in that they lack the fluid dynamics and stromal factors that physiologically impact cancer cell attachment to the endothelial wall. Microfluidic devices are at the forefront of studying metastasis in-vitro. These devices create a perfusable vascular networks that arise from 3D cultures of endothelia cells with supporting stroma [8, 9]. Still, these techniques lack the microenvironment of a physiologically intact circulatory system.

Mouse models have been the most widely used animal model in cancer research because of the gene homology with humans and similar body organization, organ function and immune system. Some limitations include the high number of animals needed to reach statistical significance. Still, mice remain the most widely used in vivo model to study vascular invasion because they closely mimic physiologic metastasis [10]. The assays in mice involve injection of cancer cells into the circulation to monitor extravasation and colonization of distant organs. Cancer cells injected into the tail vein will result mostly in seeding of the lungs, as a distant organ of metastasis [11, 12]. Intracardiac injection will result in metastatic seeding of the bone marrow [13, 14, 15] and brain [16]. Injection into the spleen or portal vein results in seeding of the liver [17] whilst carotid artery injection directs cancer cells to the brain [18, 19]. Monitoring and quantitation of organ colonization is generally determined via luminescence and can detect manifest metastases. Real-time imaging of extravasation can be very instructive though it requires surgical implantation and complex microscopy that has to adapt to the breathing of an anesthetized animal [20, 21, 22]. This complexity of the setup limits the extravasation analysis.

The zebrafish model has emerged as a system to study cancer cell metastasis that is easier to implement, accommodates higher throughput and allows for state-of-the-art imaging [23, 24]. This model allows for the assessment of cancer cell invasion and extravasation in an intact circulatory system over a shorter timescale when compared with murine model [25, 26, 27, 28]. The zebrafish genome overlaps with the human genome by 70% and most organs and cellular function are conserved. An overview of zebrafish in biomedical research is provided in an award-winning short movie for an educated lay audience [29] ( www.zebrafishfilm.org ). One major benefit of the model is the potential for cost efficient, relatively high throughput using dozens of easy to visualize transparent embryos that can grow and develop in small volumes accommodated by 96-well format dishes in a single experiment [30]. The model also allows the observation of very few to single-cell behaviors that help elucidate the impact of subpopulations present in heterogeneous tumor cell populations (see e.g. Fig. 3) [31]. Therefore, transplantation of cells in the optically clear, immune-permissive zebrafish embryos can provide a unique perspective to understand cancer metastasis and to visualize single-cell cancer progression in real-time [32]. The zebrafish model of cancer invasion and metastasis can also be used to assess drug effects or screen for candidate inhibitors in an in vivo model. Drugs to be analyzed can be added to the growth media of the fish embryos at an appropriate time to read out the effect on the host embryo as well as the cancer cells transplanted for evaluation of invasion and metastasis. Several examples from different areas of study were published during the past years [33, 34, 35, 36] including the application to individualized analysis of human cancer biopsies [37].

Figure 3. Intravasation of two distinct cancer cell populations after yolk sac injection.

Figure 3.

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(A) Brightfield image of a wild-type zebrafish embryo 4 days after fertilization. The boxed areas indicates the yolk sac. (B) Yolk sac with green (invasive = low density growth) and red (non-invasive = high density growth) Q-dot labeled cancer cells. (C) The caudal region with cancer cells that intravasated into the vasculature and reached the tail region. (D) The percentage of embryos (n = 75) with intravasated cells in the caudal region two days after injection (Original data in Ref. [39]).

In the approach described, first we utilize a transparent zebrafish strain that has its endothelia tagged with a green reef coral fluorescent protein reporter driven by the kdrl promoter, the receptor for vascular endothelial growth factor [38]. The vascular invasive ability of a commonly used cancer cell line is shown as a representative example and the general steps are outlined in Figure 1. In this example, MDA-MB-231, a human breast cancer cell line, was labeled with a red fluorescent lipophilic dye and injected into the precardiac sinus of 2-day old embryos. Between 48 and 96 hours after the injection, cancer cells that have extravasated out of the vasculature and invaded into tissues in the caudal region of embryos can be scored efficiently on a fluorescent microscope (Fig 2). In the experiment shown here, the breast cancer cell line was cultured at different cell densities for a few days resulting in inhibition of the Hippo pathway and in vitro invasion of an endothelial monolayer by cells grown at low density [39]. In the zebrafish embryos cells grown at low density extravasated from the vasculature and invaded tissues at a significantly higher rate confirming the biologic significance of the in vitro findings (Fig 2B,C).

Figure 1: Overview of the cancer cell extravasation assay in zebrafish embryos.

Figure 1:

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Cancer cells are incubated with Q-dots (Thermofisher) or a fluorescent dye that is taken up by the cells as a label (red). Cells in suspension are injected into the precardiac sinus of the embryos (2 days old). During the next 2 to 4 days, cancer cells traffic in the vasculature, can invade tissues in the caudal region of the embryo and are imaged after mounting in an anesthetic agarose medium. The vascular endothelia in the embryos express GFP (Green Fluorescent Protein) to identify the location of cancer cells either inside the vasculature or inside the tissues by confocal microscopy. (Reproduced with permission; Ref. [41]).

Figure 2. Extravasation of cancer cells (red) in zebrafish embryos with GFP-labeled vasculature (green).

Figure 2.

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(A) Schematic of injection of cancer cells into the precardiac sinus. Note that some of the cancer cells have entered tissues 2 days after injection. (B) The embryo's caudal region is magnified to show extravasated cancer cells. Punctate fluorescent labeling is seen upon magnification. (C) Quantitation of extravasated cancer cells in tissues. Twelve and five Zebrafish were scored for extravasated cancer cells grown at low and high density respectively (Original data in Ref. [39]).

In a second approach in wildtype zebrafish embryos without fluorescent vasculature, the injection site of cancer cells is changed to the yolk sac (Fig 3A,B). The approach allows for the study of heterogeneous cancer cell populations that are labeled with differently colored Q-dots (Fig 3). Cancer cells that have intravasated in the yolk sac can traffic in the vasculature and are scored in the caudal region (Fig 3C) 24 to 48 hours after injection [39, 40]. In the example shown here, breast cancer cells grown at low or high cell density were labeled with green or red Q-dots respectively and co-inoculated at the same time into the yolk sacs of embryos for a direct comparison (Fig 3B). Cells grown at low density intravasated at a significantly higher rate than the high-density cells as evidenced by their appearance in the tail vasculature (Fig 3C, D). The extravasation and intravasation assays from Figures 2 and 3 provide complementary insights. Both approaches can also be used to assess the contribution of pathways by knockdown of candidate genes as well as treatment with small molecule or antibody-based inhibitors of potential driver molecules. Examples of such interventions can be found in [39, 40].

2. Materials

Injection station

  1. Needles: Capillary Glass, Standard, 1.2MM x 0.68MM, 4", A-M Systems, Inc, pulled using David Kopf 700C Vertical Pipette Puller, Hofstra Group. Pull long tapered pipettes using 20 mAmp current and a 2-coil heating element.

  2. Machinery needed for injection station set up: Electrode Storage Jar, 1.0MM, World Precision Instruments, Inc, latex rubber bulbs, 2mL, Pack of 72, Heathrow Scientific and micromanipulator, Narishige, picospritzer II, General Valve Corporation.

Manipulating fish

  1. Tools for manipulating embryos: Eyelash Brush, Ted Pella, Inc, transfer pipettes, and 5 3/4" Disposable Pasteur Pipets, borosilicate Glass.

  2. Fish water: 0.3 g/L, Instant Ocean Salt, Sea Salt, Pentair. Store at room temperature for up to 3 months.

  3. Tricane stock solution: Ethyl 3-aminobenzoate methanesulfonate salt (Tricaine, MS-222), Fluka: (4 mg/mL,10 mM Tris, pH 7) to 50 mL of fish water plus penicillin-G potassium and streptomycin sulfate.

Visualization

  1. Microscopy method used: for visualizing cells use Leica SP8 Confocal Microscope.

  2. Staining cells:
    1. For staining cancer cells for pre-cardiac sinus injection cancer cells were labeled with a lipophilic red fluorescent dye: vybrant diI cell-labeling solution (Thermofisher).
    2. For staining cancer cells for yolk sac injection cells were labelled with 565 nm or 655 nm Q-dots using the Qtracker Cell Labeling kit (Thermofisher).

3. Methods

Ethics Statement: Experiments were carried out in compliance with recommendations by the Georgetown University Animal Care and Use Committee. Zebrafish embryos were generated under an approved IACUC protocol.

3.1. Set up embryos for injection and make stock solutions

  1. Separate males and females the night before experiment in a breeding cage. The goal is to develop enough zebrafish larvae to assess cancer cell vascular invasion.

  2. Pull the gate on the breeding cage of pair-wise or group in-cross mating with Tg(kdrl:grcfp)zn1;mitfab692;ednrb1b140 fish (see Note 1).

  3. After about 30 mins, collect eggs, and remove unfertilized, deformed embryos and debris the next day under the microscope.

  4. Incubate embryos at 28.5 °C until ready for injection, 2-days post-fertilization (2 dpf), make sure all embryos are dechorionated.

  5. Make injection plates: melt 25 mL of 1.5% agarose in dH2O for each plate.

  6. Pour 12 mL of the agarose into a 100 mm x 15 mm petri dish and let it solidify.

  7. Re-melt then pour the remaining agarose in the plate.

  8. 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. (see Note 2).

  9. Tape the glass mold in place and let the agarose set.

  10. Slowly remove the glass mold. (see Note 3).

  11. Equilibrate injection plate with fish water (0.3 g/L sea salt) and to room temperature.

  12. Wash plate twice with distilled water.

  13. Equilibrate plate by adding 10 mL of fish water to the plate and place on shaker for 10 min. Do this two times as well.

  14. Split 2-day post-fertilization (dpf) embryos into injection groups by transferring them into petri dishes containing fish water.

  15. Set up recovery dishes for each group that will be used after injection. (see Note 4).

  16. 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.

  17. 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. (see Note 5)

  18. Pull the microinjection needles.

  19. Place glass capillary tubing in a vertical pipette puller. Pull long tapered pipettes using 20 mAmp current and a 2-coil heating element.

3.2. Labeling cells with red lipophilic fluorescent dye

  1. Grow cells of interest in their recommended culture conditions (see Note 6).

  2. Generate a single-cell suspension by dissociating an adherent culture of cancer cells.

  3. Wash cells with 1X PBS and add 0.05% trypsin-EDTA solution (see Note 7).

  4. Neutralize the trypsin solution with serum-containing cell culture media after the cells detach.

  5. Centrifuge the cell suspension for 5 min at 200 x g, then resuspend the cell pellet in media for cell counting.

  6. Count the cell suspension and prepare 1 million cells per 200 μl of cell culture media (see Note 8).

  7. Check cell viability with trypan blue dye before injection into zebrafish embryos (see Note 9).

  8. Add 2 μl of red lipophilic dye to the cell suspension for a 1:100 dilution, mix well, and then incubate the mixture at 37 °C for 20 min (see Note 10).

  9. Following the incubation, add 1 mL of fresh media to the tube and then centrifuge again for 5 min at 200 x g.

  10. Wash the residual fluorescent dye from the cells.

  11. Aspirate the supernatant and resuspend the pellet in 1 mL of fresh culture media, and centrifuge for 5 min at 200 x g.

  12. 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.

  13. Repeat the washing step: aspirate the supernatant, resuspend the cell pellet in 1 mL of fresh media and then centrifuge for the third time for 5 min at 200 x g.

  14. Aspirate the supernatant and resuspend the cell pellet containing 1 million labeled cancer cells in 500 μL of fresh media (see Note 11).

3.3. Injecting cells into the precardiac sinus of zebrafish embryos

  1. Attach microinjection dispense system to a pressurized air source and turn on the microinjection dispense system power.

  2. Test pressure by depressing the foot pedal. A brief pulse of air should come out from the needle holder.

  3. Equilibrate the injection plates twice with 2X tricaine solution.

  4. For each equilibration step, add 20 mL of 2X tricaine solution to the injection plate and place on shaker for 10 min.

  5. Use a transfer pipette to move a group of embryos to a small dish containing the 2X tricaine solution.

  6. Fill the microinjection injection needle from the back with cancer cells using a gel-loading pipet tip.

  7. Place the needle in an electrode storage jar with the pointed end facing down so cells settle near the tip, make sure to cut your tip so the appropriate number of cells are injected per embryo.

  8. Transfer 20-30 anesthetized embryos to an injection plate with a transfer pipette with 2X tricaine.

  9. Allow embryos to settle in the tip of the pipette.

  10. Gently move embryos into the trough of the injection plate, spreading the fish along the length of the trough.

  11. Align embryos with heads facing up and bellies facing the steep wall of the trough (see Note 12).

  12. Inject 50-100 cancer cells (2-5 nL) into the precardiac sinus of the zebrafish embryos using the microinjection dispense system.

  13. Attach the needle to the needle holder of a micromanipulator.

  14. Position the injection plate under the stereoscope with the 60° wall to the left and focus on the top embryo at 25x magnification (Fig 1).

  15. Position the micromanipulator so that, the needle will pierce the embryo.

  16. Extend the needle until it is nearly touching the embryo.

  17. Looking under the microscope, align the needle so that it will pierce the embryo upon further extension.

  18. Pierce the embryo through the yolk sac placing the tip just at the pre-cardiac sinus.

  19. Inject cells by depressing the foot pedal. The force of the injection expels the cells into the cardiac sinus. Retract the needle (Fig 2B).

  20. Extend and retract the injection needle with one hand. Make fine adjustments to position next embryo with your other hand.

  21. Retract the needle as high as it goes while setting up to inject another plate.

  22. Transfer the embryos to the recovery dish once the entire plate is injected.

  23. Tilt the injection plate to pool the embryos at the bottom of the petri dish and wash embryos off the plate with fish water, and collecting them with the transfer pipette.

  24. Allow the embryos to settle in the bottom of the transfer pipette.

  25. Transfer the embryos to the recovery dish.

  26. Incubate recovery dish at 28 °C for 1 hr. After the incubation, separate viable zebrafish embryos from dead embryos.

  27. Incubate dish at 33 °C until ready for scoring, typically 24-96 hrs (see Note 13).

3.4. Scoring extravasation of cancer cells in zebrafish embryos

  1. Anesthetize one batch of embryos at a time to be scored by placing them in a dish with 2X tricaine solution.

  2. Place an anesthetized larva on a depression slide in a drop of 2X tricaine.

  3. Orient larvae laterally for imaging of the caudal region (Fig 2B).

  4. Count the number of cancer cells that have successfully invaded out of the vasculature by focusing on the cell location to clearly discern an intact cell (see Note 14).

  5. Score larvae on a Nikon compound fluorescence microscope with the 10x objective lens. Use the 20x objective for any difficult calls (Fig 2C).

3.5. Mounting zebrafish onto slides and subsequent fluorescence imaging

  1. Melt 1.5% agarose/2X tricaine solution, and bring to 37 °C.

  2. Anesthetize the embryo by placing it in 2X tricaine solution.

  3. Transfer the embryo in a drop of 2X tricaine solution to the imaging station (see Note 15).

  4. Use a glass pipette to remove the excess 2X tricaine solution, retaining the embryo on the imaging surface.

  5. Overlay one drop of melted agarose solution over the embryo.

  6. Before the agarose polymerizes, use an eyelash brush, to orient the embryo laterally for imaging, making sure the embryo is flattened along the imaging surface.

  7. Submerge the now polymerized agarose drop under 2X tricaine solution.

  8. Image the zebrafish embryo using microscopy.

3.6. Alternative application: Injecting cancer cells into the yolk sac

  1. Prepare the microinjection dispense system and injection plates as previously described in section (3.3) of this protocol.

  2. Label two cell populations with contrasting fluorescent Q-dots from the Qtracker Cell Labeling kit (Thermofisher) or with flourescent dyes as described in section (3.2) above.

  3. Inject 5-10 nL of 2 x10^7 cells/mL into the yolk sac (Fig 3B). Keep the injection volume constant to inject identical cell numbers (100-200 cancer cells) from each cell population (see Note 16).

  4. Collect the injected embryos as previously described in section (3.3) of this protocol and then screen for successful injections.

  5. Screen and transfer viable embryos that were successfully injected to a new dish (see Note 17).

  6. Transfer the viable embryos to a new petri dish if cancer cells are clearly seen in the yolk sac (Fig 3B).

  7. Incubate dish at 33 °C until ready for scoring, typically 24-48 hrs after injection (see Note 18).

  8. To score intravasation in zebrafish embryos, follow the guidelines in section (3.4) of this protocol, but instead count the number of cancer cells that have successfully invaded into the vasculature of the caudal region (Fig 3C).

4. Notes

  1. 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.

  2. This will create a steep 60° wall with a 30° sloped ramp.

  3. Molds can be stored in dH2O at 4 °C, edges wrapped with parafilm.

  4. Recovery dish contains 10 mL fish water: penicillin (25 μg/mL) and streptomycin (50 μg/mL).

  5. Mounting medium consists of 1.5% agarose.

  6. This cell line was grown in DMEM + 10% FBS: MDA-MB-231.

  7. Time of trypsin exposure will depend on the cell line.

  8. Cell number is determined with an automated counter, feel free to use whatever you have.

  9. Only live cell populations should be injected into zebrafish embryos, as injection of dead cells will not reflect true vascular invasion.

  10. We have found that Dil tends to produce the best labeling, yet its fluorescence level can vary between cell lines. Concentration and labeling time may need to be optimized for each cell line. It is critical that the excess dye is washed away from the cells after labeling. This is achieved by the centrifugation steps indicated in the labeling section 3.2. Excess fluorescent dye can be toxic to cells or leak into the bloodstream, producing a faux fluorescent background.

  11. 0.5 mM EDTA can be added to the media to prevent cell clumping.

  12. Tricaine solution should cover both the trough length and the flat agarose surface, with embryos only residing in the trough.

  13. This temperature is determined as a compromise between 37 °C, the ideal temperature for cancer cells, and 28.5 °C, the temperature for zebrafish.

  14. It is best to have at least two individuals involved in this process, where one individual scoring the fish is blind to the experimental condition being assessed. It is imperative to generate consistent criteria that will be applied to all conditions.

  15. A glass-bottom dish or microscope slide can be used.

  16. A transparent zebrafish embryos lacking fluorescent vasculature can be used for this assay. Passive entry of particles into the vasculature can be controlled for by injecting fluorescent beads (<10 μm).

  17. All embryos should have a consistently sized mass of cells located in the yolk (Fig 3B). Embryos are discarded if the mass size differs or if any cells are located outside of the yolk.

  18. This temperature is considered a compromise between 37°C, the ideal temperature for cancer cells, and 28.5°C, the ideal temperature for zebrafish.

ACKNOWLEDGMENTS:

We thank Peter Johnson of the Georgetown University Microscopy Core for assistance with imaging the zebrafish embryos. The Microscopy & 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 F31-CA235970-01 (SMR), R01CA231291-01(AW).

Footnotes

DISCLOSURES:

The authors have no conflicts of interest to disclose.

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