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Method Article

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

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DOI:

10.3791/66164

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July 12th, 2024

 ,  ,  , 
1Nuclear Dynamics and Cancer Program, Fox Chase Cancer Center

In This Article

Summary

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.

Abstract

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.

Introduction

Analysis of the behavior of tumors in response to genetic alteration or drug treatment in vivo is an essential element of cancer research1,2,3,4. Such studies most often involve the use of immunocompromised mouse (Mus musculus) models5; however, xenotransplantation studies in mice are limited in many respects, including limited capacity, extended duration, significant expense, and the requirement for sophisticated imaging equipment to monitor the progression of internal tumors6,7. By contrast, the zebrafish model (Danio rerio) enables greater capacity, shorter duration, lower expense, and, due to their transparency, simple monitoring of disease progression8,9.

Zebrafish is a well-developed vertebrate model system with ex-utero development and high fecundity, with individual females producing more than 100 embryos10. Moreover, zebrafish embryos are transparent, enabling easy visualization of developmental processes using fluorescence-related techniques such as reporters. Finally, the conservation of critical developmental processes makes them an ideal model for many types of studies, including the behavior of transplanted malignant cells11,12. Wild-type zebrafish embryos develop melanocytes, which render them optically opaque by 2 weeks of age, but this has been overcome by the generation of casper embryos (roya9; mitfaw2), which remain transparent throughout life13. Because of their optical properties, casper zebrafish are ideal recipients of transplanted tumor cells14,15,16. Xenotransplantation of tumor cells into zebrafish has gained importance in the past 2 decades17,18,19,20,21. Zebrafish embryos have innate immunity; however, they lack adaptive immunity during their larval stage, rendering them functionally immunocompromised, which enables them to serve as effective hosts for transplanted tumor xenografts22.

Protocols have been developed for tumor engraftment in zebrafish embryos as well as adults that have considered a number of different variables23,24,25,26,27. These have explored numerous sites of tumor deposition in zebrafish, including injections in yolk, peri-vitelline space, and heart and at different developmental stages16,28. The ambient temperature of aquaculture for zebrafish xenografts is also important as zebrafish rearing typically occurs at 28 °C, while mammalian cells grow at 37 °C. Consequently, a compromise temperature must be employed that is tolerated by the fish yet supports tumor growth, and 34 °C appears to achieve both goals29. Analysis of the behavior and progression of tumors following xenotransplantation is another major area of focus, and this involves the use of a variety of imaging modalities as well as survival analysis30. One of the major advantages of the zebrafish model is the availability of large numbers of study animals to provide immense statistical power to in vivo studies of tumor behavior; however, previous approaches have severely limited this potential because of the requirement of tedious mounting procedures for injections.

Here, we address this limitation through the development of a simple, rapid method with which to stage embryos that enables high throughput and monitoring of injection quality using the transparent casper zebrafish line. This entails the injection of xenografts into the yolk sac of the casper zebrafish embryos at 2 days post fertilization (dpf). We observe the survival of embryos following xenotransplantation as part of tumor behavior analysis. We further show the assessment of disease burden after xenotransplantation by making single cell suspensions and analyzing by flow cytometry (Figure 1).

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Protocol

Zebrafish maintenance, feeding, and husbandry occurred under standard aquaculture conditions at 28.5 °C, as described31. All zebrafish-related experiments were done at this temperature; however, following xenotransplantation, the animals were cultured at 34 °C for the duration of the experiment, in accordance with procedures approved by the Institutional Animal Care and Use Committee (IACUC).

1. Breeding (3 days before injection)

  1. Provide dry feed (extra feed; 5-6 granules per fish) to fish pairs a week prior to breeding to maximize animal health and increase the number of embryos produced by breeding pairs.
  2. On the evening prior to embryo harvest, set up breeding animals in breeding tanks with a divider separating male and female fish, using harem matings of 2 females for each male.
    NOTE: For experiments with 4 arms of 100 animals per arm, employ 20 breeding pairs per experiment. To determine the number of breeding pairs needed, a good estimate is 50 embryos per casper breeding pair. Another option is to employ a more robust, pigmented zebrafish strain and treat with 1-phenyl 2-thiourea (PTU) to prevent pigmentation31. In practice, the experiment should be scaled such that one has enough embryos to inject twice the number desired at 1 day post injection (dpi).

2. Embryo collection (2 days before injection)

  1. The next morning, remove the dividers, allowing the fish to breed.
  2. Visualize the embryos in the tanks 20 minutes (min) after removing the dividers.
  3. Collect the embryos using a sieve in a 90 mm Petri dish containing embryo water made as described in The Zebrafish Book31.
    1. To make embryo water, add 1.5 mL of stock salts to 1 L of distilled water and methylene blue to 0.1% final. Make the stock salt solution by dissolving 40 g of sea salts (Table of Materials) in 1 L of distilled water. The ionic composition of embryo water is K+ (0.68 mg/L), Cl− (31.86 mg/L), Na+ (17.77 mg/L), SO4− (4.47 mg/L), Mg2+ (2.14 mg/L), and Ca2+ (0.68 mg/L).
  4. Allow the zebrafish to breed for an extra hour and collect the resulting embryos.
  5. Pool the embryos for the experiment.
  6. In the evening, remove all unfertilized or dead embryos, which are recognizable by their abnormal morphology, and provide fresh embryo water.

3. Embryo maintenance and tool preparation for injections (1 day before injection)

  1. The following morning, remove any additional dead embryos and provide fresh embryo water.
  2. Prepare an agarose plate by heating 1.5% agarose in embryo water and pour the heated mixture into a 90 mm Petri plate. One 90 mm dish requires 30-35 mL of the mixture.
  3. Pull non-filament needles from glass capillaries (Borosilicate) using the needle puller. Needles are pulled (under heat pressure) producing a closed end; clip them with forceps to generate an optimal orifice. Assess the suitability of the needle orifice by determining the volume of fluid displaced per unit time (see below,  section 6.3).
    NOTE: Glass capillaries can be purchased with or without central filaments. Capillaries lacking central filaments are preferred for cell injections.
  4. Place the needles in a covered 90 mm Petri plate in the grooves made using clay (kids modeling clay) until use (Figure 2A).

4. Preparation and labeling of leukemia cells with CM-Dil (day of injection)

  1. Maintain the cells to be transplanted under conditions optimal for their growth. The murine leukemia cells employed here were either sufficient (M82; Rpl22+/+) or deficient (M109; Rpl22-/-) for the ribosomal protein Rpl22, which functions as a tumor suppressor32.
  2. Pellet cells in a 50 mL conical tube. Count, then centrifuge at 300 x g at room temperature (RT) for 5 min. Discard the supernatant.
    NOTE: The number of cells needed will be dictated by the experimental scope and conditions, but 1 x 106 cells is a good starting point.
  3. Perform CM-Dil staining
    NOTE: CM-Dil staining enables monitoring of the injection bolus.
    1. Make a stock solution of CM-Dil by resuspending a 50 µg vial of CM-Dil in 50 µL of dimethyl sulfoxide (DMSO; 1 mg/mL or ~1 mM final).
    2. Produce a working solution by diluting the stock (4.8 µL of stain/mL) in 1% fetal bovine serum (FBS)/Hank's balanced salt solution (HBSS) containing any supportive supplements needed by the cells to be used.
  4. Resuspend cells at 1 x 106/100 µL in the working solution of the stain.
  5. Incubate at 37 °C for 10 min.
    NOTE: The staining conditions must be optimized for the cells employed (time, etc). The cells used here required two distinct 10 min incubations at different temperatures to achieve optimal staining.
  6. Wash with 10 mL of 1x HBSS at room temperature (RT).
  7. Pellet cells (centrifugation at 300 x g for 5 min at RT), decant the supernatant, then resuspend with 10 mL of HBSS and repeat the same for a second wash.
  8. Resuspend the stained tumor cells at 40,000 cells/µL in 1% FBS/PBS and any supportive supplements needed and maintain the cell suspension at 34 °C until injection.
    NOTE: Supportive supplements were required for maintenance of the cell lines used in this study (e.g., 1% FBS and cytokines). The supplements and xenotransplantation may need to be adapted to the particular cell lines under study. PBS was selected here instead of media to avoid any potential toxicity in the yolk.

5. Dechorionation

  1. Dechorionate the casper zebrafish embryos manually at 2 dpf using insulin injection syringes under 2x magnification in a light microscope. Pierce the chorion with one needle while using the other needle to immobilize the chorion.
    NOTE: Using pronase for dechorionation is not recommended because it sometimes results in reduced embryo health. Dechorionating the embryos at 2 dpf is preferred because it is easier, and the embryos are less fragile than at earlier times (1 dpf).
  2. While dechorionating, be careful not to touch the embryos with the needles. Touching or damaging the yolk of embryos by inadvertent contact with the needles might cause death.
  3. Remove the stripped chorions by changing the embryo water.

6. Setting up the microinjector and needle

  1. Turn the microinjector and the pump on and set up the conditions suitable for microinjections of cells. An injection pressure of 9-11 psi and a time of injection of 0.5 seconds (s) are a good starting point for clipping the needle and setting the orifice.
  2. Load the tumor cell line suspension (~5 µL) into the microneedle carefully in a single pass, avoiding formation of air bubbles, which will disrupt the cell stream inside the needle.
  3. Cut the end of the needle with forceps (Dumont number 5) to produce an orifice that will support the ejection of 10-15 nanoliter (nL) of cell suspension per 0.2-0.3 s.
    NOTE: The above calculation of nL volume is done using calibration capillaries, where 1 mm = 30 nL. In brief, set the time to 0.5 s, and after every clip of the needle, press inject and collect the volume in the calibration capillary. Then, the length of the collected volume is measured using the scale under a microscope, and the needle clipping is stopped when 30 nL is injected in 0.5 s. Then, set the time of injections as 0.2-0.3 s. (injecting ~10-15 nL of cells)

7. Embryo preparation for injection

  1. Select healthy embryos under the microscope, culling any with developmental anomalies such as heart edema or a short or curved trunk.
  2. Anesthetize the embryos using Tricaine methanesulfonate (MS-222; 0.16 g/L of embryo water) for 1 min in a 90 mm Petri plate.
  3. Use a glass Pasteur pipette to pick up the embryos. Arrange 10-15 embryos in a lateral position on the 1.5% agarose plate (Figure 2B-D).
  4. Remove excess water using the Pasteur pipette, leaving the minimum amount of embryo water needed to keep the embryos alive.

8. Injection procedure

  1. Check under the light microscope to ensure that the cells have accumulated in the tip of the needle.
  2. Inject the embryos using the calibrated needle for 0.2-0.3 s (with 10-15 nL corresponding to 400-600 cells) in the yolk of the embryos.
  3. Repeat the injection for all the embryos, then collect them in fresh embryo water.
    NOTE: Because cells tend to accumulate on the needle tip, the tip will need to be reclipped slightly every 15-20 injections. This will also require resetting the pressure and time with each reclipping to ensure that a similar volume is injected.
  4. To ensure that the comparison of the behavior of two distinct sets of transferred cells is valid, monitor the bolus of cells transferred. Do this by sorting the embryos based on CM-Dil staining (in the RFP channel) at 1 hour post injection (hpi), separating those with optimal staining ("good bolus") from those with inferior staining ("inferior bolus"; Figure 3, yellow arrow).
  5. Discard the embryos with an inferior bolus or use them to assess the impact of a different cell dose on disease progression.
  6. Remove any dead embryos by the end of the day since their death is related to injection trauma rather than tumor growth. Remove from analysis embryos that do not retain cells since the cells likely leaked out of the injection site.
  7. Maintain the injected embryos at 34 °C for the duration of the experiment in a 90 mm dish with ~60 embryos per plate.
    NOTE: After 5 dpf, the yolk will have been consumed by the growing embryos, so embryos must be provided with paramecium food for the duration of the experiment. To ensure proper nutrition, paramecia should be given to the embryos daily from 6 dpf (4 dpi) to 9 dpf (7 dpi). Paramecia are propagated by culturing in flasks under optimum nutrition and temperature conditions, as described31.

9. Survival analysis

  1. Monitor the embryos for the next 7 dpi, changing the embryo water daily. Water changes may be reduced to alternate days for convenience if the study involves drug treatment.
  2. Check embryo health and score death for the duration of the analysis.
    NOTE: The experimental duration was 7 days for this experiment but may be shorter or longer depending on the aggressiveness of the xenotransplanted tumor.
  3. Use CM-Dil fluorescence to assess disease burden (Figure 4A) and determine the impact of genetic alterations or drug treatments on survival using Kaplan Meier analysis and depict graphically (e.g., with GraphPad Prism; Figure 4B)33.

10. Single-cell suspension of embryos for flow cytometry analysis

NOTE: Disease burden can be assessed by flow cytometry analysis after xenotransplantation; however, doing so requires indelible marking of the tumor cells. Retrovirally or lentivirally-delivered red fluorescent protein (RFP) or mCherry is effective as it provides a good signal over the autofluorescence of zebrafish cells, which obscures signal from green fluorescent protein.

  1. Isolate embryos at the dpi stage of choice. 5 dpi is displayed here (Figure 5).
  2. Gather 30-40 embryos per condition as a starting point, but the embryo number may differ depending on the stage and aggressiveness of the transplanted cells. Anesthetize the embryos as described above.
    NOTE: Embryos may be subdivided as replicates to provide statistical significance, as shown here (Figure 5B).
  3. Transfer the embryos to 1.5 mL centrifuge tubes.
  4. Use 100 μL of calcium-free Ringer's solution (recipe31) per sample to dissolve the yolk since low calcium softens the embryonic tissues, enabling more effective tissue dissociation.
  5. Pipette up and down intermittently for 5 min to remove the yolk using a 200 μL pipette tip.
  6. Pre-heat 0.05% trypsin/PBS (without phenol red indicator) to 29 °C and supplement it with 27 μL of collagenase IV (100 mg/mL) per mL of Trypsin solution. A volume of 1 mL of solution will be needed for each sample of embryos.
  7. Add 1 mL of the trypsin/collagenase solution to each sample of deyolked embryos and incubate at 29 °C for 30-35 min.
  8. Pipette the embryos up and down in this solution using a 1 mL pipette tip every 5 min until the structure of the embryos (backbone) is no longer visible.
  9. Stop the reaction using 200 μL of FBS.
  10. Mix well and incubate at 29 °C for an additional 5 min to ensure complete inactivation of the trypsin.
    NOTE: A temperature of 29 °C is employed for the tissue dissociation protocol to prevent heat shock-induced death of zebrafish cells, which occurs at 37 °C; however, if preservation of zebrafish cells is not required, digestion can be performed at 37 °C.
  11. Pellet the cell suspension at 300 x g for 5 min at 4 °C and discard the supernatant.
  12. Resuspend the cell pellet in 4 °C PBS and pellet as above.
  13. Repeat the wash, then strain the cells through a 70 µm cell strainer.
  14. Pellet and proceed with staining for flow cytometry analysis.
    NOTE: If culture of primary zebrafish cells is required, wash twice more with 4 °C PBS and resuspend in L15 media (with antibiotics and 10% FBS).

11. Fluorescence-activated cell sorting (FACS): Staining and sorting of xenotransplanted cells

  1. Resuspend the cell suspension in a staining medium (HBSS with 1% FBS) and pellet at 300 x g for 5 min.
  2. Resuspend the cell pellet in the staining medium with an antibody reactive with the transplanted cells to provide a second marker (in addition to RFP or mCherry) with which to distinguish transplanted cells from zebrafish cells. Here, 50 µL of anti-mouse CD45 (APC-CD45) per sample was employed at a 1:50 dilution (Figure 5).
  3. Incubate for 20 min at 4 °C before washing as above with 1 mL of the staining medium to remove the unbound antibody.
  4. After discarding the supernatant, resuspend the cell pellet in 200 µL of staining medium containing the vital dye Helix NP Blue (1 µM), which will enable live/dead discrimination.
  5. Transfer the cell mix to 5 mL round bottom polycarbonate tubes for flow cytometry analysis.
    NOTE: Staining a tumor cell control in parallel provides clarity for drawing gates during flow cytometry analysis.

12. Flow cytometry

  1. Turn on the flow cytometer that is available using a low flow rate (500 events per second or less) to set the parameters.
  2. Use the tumor cell control (the same cells used for xenotransplantation) to set the voltage for FSC (forward scatter), SSC (side scatter), BV421/CasB (viability), CE-594 (mCherry), and APC (CD45.2) channels.
    NOTE: Singly stained samples will be required to establish the compensation settings that eliminate fluorochrome overlap between distinct stains.
  3. Use the uninjected embryo sample to establish settings that accommodate both transplanted and zebrafish cells.
  4. Increase the flow rate to 8000 events per second and record 1 million events for each sample.
  5. Analyze the resulting data using appropriate analysis software, first selecting singlets by plotting FSC-H v/s FSC-A (height v/s area), followed by a plot for selecting viable cells. FlowJo software is widely used for such analysis.
  6. Using the tumor cell control, draw a gate around the transplanted cells by FSC-A v/s SSC-A, then use the indicator stains, in this case, CD45 and mCherry (Figure 5B).
    NOTE: The size gate selected for the tumor will also contain zebrafish cells, which provides the basis for normalization and determination of disease burden.
  7. Analyze the embryo samples using the same gate settings as above. The final plot of tumor stain and fluorescent protein indicator will provide a measure of disease burden (Figure 5C).
    NOTE: Here, the difference in disease burden was plotted for embryos receiving a good bolus of injected cells versus those receiving an inferior bolus.
  8. If needed, sort the tumor cells in the embryos by flow cytometry for downstream molecular analysis.

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Results

Xenotransplantation
A comprehensive view of the entire experiment and analysis is depicted in Figure 1, spanning from embryo production to the assessment of disease progression by both survival and disease burden analysis by flow cytometry. This approach brings several improvements that enhance the reproducibility and scalability of xenotransplantation, as well as adding a new way to assess disease burden. The success of these experiments is highly dependent upon the hea...

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Discussion

Zebrafish xenotransplantation has emerged as a rapid, robust, and cost-effective alternative to mouse studies12. Though several approaches to zebrafish xenotransplantation have been reported, our adaptation has resulted in significant improvements. In addition to standardizing parameters around the procedure, these improvements specifically focus on accelerating the rate at which tumor injections can be performed, thus enabling an increase in the number of animals per study arm and using tumor lab...

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Disclosures

The authors declare no competing interests.

Acknowledgements

This work was supported by NIH grants R37AI110985 and P30CA006927, an appropriation from the Commonwealth of Pennsylvania, the Leukemia and Lymphoma Society, and the Bishop Fund. This study was also supported by the core facilities at Fox Chase, including Cell Culture, Flow Cytometry, and Laboratory Animal facility. We thank Dr. Jennifer Rhodes for maintaining the zebrafish and microinjection facility at FCCC.

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
1-phenyl 2-thiourea (PTU)SigmaP7629
70 micron cell strainerCorning CLS431751-50EA
90 mm Petri dishThermo Fisher ScientificS43565
AgaroseApex bioresearch20-102GP
APC APC anti-mouse CD45.2 AntibodyBiolegend109814
BD FACSymphony A5 Cell AnalyzerBD BiosciencesBD FACSymphony A5
calibration capillariesSigma P1424-1PAK
Cell tracker CM-dil dyeInvitrogenC7001
Collageanse IVGibco17104019
Dumont forceps number 55Fine science tools11255-20
FBSCorning 35-015-CV
Fluorescence microscopeNikonmodel SMZ1500
Glass capillaries (Borosilicate)World precision instruments1B100-4
HBSSCorning 21-023-CV
Helix NP BlueBiolegend425305
Instant Ocean Sea SaltInstant oceanSS15-10
Light microscopeNikonmodel SMZ1000
Methylene blueSigmaM9140-100G
Microloader (long tips for laoding cells)eppendorf930001007
P1000 micropipette pullerSutter instrumentsmodel P-97
PM 1000 cell microinjectorMicroData Instruments, Inc. (MDI)PM1000
Tricaine methanesulphate (Ethyl 3- aminobenzoate methanesulphate)SigmaE10521-10G
Trypsin-EDTA (0.5%), no phenol redGibco15400054
Zebrafish adult irradiated diet (dry feed)Zeigler388763

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