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.&#160;We thank Dr. Jennifer Rhodes for maintaining the zebrafish and microinjection facility at FCCC.

Zebrafish maintenance, feeding, and husbandry occurred under standard aquaculture conditions at 28.5 &#176;C, as described31. All zebrafish-related experiments were done at this temperature; however, following xenotransplantation, the animals were cultured at 34 &#176;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)
<ol>
	<li>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.</li>
	<li>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).</li>
</ol>
2. Embryo collection (2 days before injection)
<ol>
	<li>The next morning, remove the dividers, allowing the fish to breed.</li>
	<li>Visualize the embryos in the tanks 20 minutes (min) after removing the dividers.</li>
	<li>Collect the embryos using a sieve in a 90 mm Petri dish containing embryo water made as described in The Zebrafish Book31.
	<ol>
		<li>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&#8722; (31.86 mg/L), Na+ (17.77 mg/L), SO4&#8722; (4.47 mg/L), Mg2+ (2.14 mg/L), and Ca2+ (0.68 mg/L).</li>
	</ol>
	</li>
	<li>Allow the zebrafish to breed for an extra hour and collect the resulting embryos.</li>
	<li>Pool the embryos for the experiment.</li>
	<li>In the evening, remove all unfertilized or dead embryos, which are recognizable by their abnormal morphology, and provide fresh embryo water.</li>
</ol>
3. Embryo maintenance and tool preparation for injections (1 day before injection)
<ol>
	<li>The following morning, remove any additional dead embryos and provide fresh embryo water.</li>
	<li>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.</li>
	<li>Pull non-filament needles from glass capillaries (Borosilicate) using the needle puller. Needles are pulled (under heat pressure)&#160;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, &#160;section 6.3). 
	NOTE: Glass capillaries can be purchased with or without central filaments. Capillaries lacking central filaments are preferred for cell injections.</li>
	<li>Place the needles in a covered 90 mm Petri plate in the grooves made using clay (kids modeling clay) until use (Figure 2A).</li>
</ol>
4. Preparation and labeling of leukemia cells with CM-Dil (day of injection)
<ol>
	<li>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.</li>
	<li>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.</li>
	<li>Perform CM-Dil staining 
	NOTE: CM-Dil staining enables monitoring of the injection bolus.
	<ol>
		<li>Make a stock solution of CM-Dil by resuspending a 50 &#181;g vial of CM-Dil in 50 &#181;L of dimethyl sulfoxide (DMSO; 1 mg/mL or ~1 mM final).</li>
		<li>Produce a working solution by diluting the stock (4.8 &#181;L of stain/mL) in 1% fetal bovine serum (FBS)/Hank&#39;s balanced salt solution (HBSS) containing any supportive supplements needed by the cells to be used.</li>
	</ol>
	</li>
	<li>Resuspend cells at 1 x 106/100 &#181;L in the working solution of the stain.</li>
	<li>Incubate at 37 &#176;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.</li>
	<li>Wash with 10 mL of 1x HBSS at room temperature (RT).</li>
	<li>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.</li>
	<li>Resuspend the stained tumor cells at 40,000 cells/&#181;L in 1% FBS/PBS and any supportive supplements needed and maintain the cell suspension at 34 &#176;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.</li>
</ol>
5. Dechorionation
<ol>
	<li>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).</li>
	<li>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.</li>
	<li>Remove the stripped chorions by changing the embryo water.</li>
</ol>
6. Setting up the microinjector and needle
<ol>
	<li>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.</li>
	<li>Load the tumor cell line suspension (~5 &#181;L) into the microneedle carefully in a single pass, avoiding formation of&#160;air bubbles, which will disrupt the cell stream inside the needle.</li>
	<li>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&#160;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)</li>
</ol>
7. Embryo preparation for injection
<ol>
	<li>Select healthy embryos under the microscope, culling any with developmental anomalies such as heart edema or a short or curved trunk.</li>
	<li>Anesthetize the embryos using Tricaine methanesulfonate (MS-222; 0.16 g/L of embryo water) for 1 min in a 90 mm Petri plate.</li>
	<li>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).</li>
	<li>Remove excess water using the Pasteur pipette, leaving the minimum amount of embryo water needed to keep the embryos&#160;alive.</li>
</ol>
8. Injection procedure
<ol>
	<li>Check under the light microscope to ensure that the cells have accumulated in the tip of the needle.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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 (&#34;good bolus&#34;) from those with inferior staining (&#34;inferior bolus&#34;; Figure 3, yellow arrow).</li>
	<li>Discard the embryos with an inferior bolus or use them to assess the impact of a different cell dose on disease progression.</li>
	<li>Remove any dead embryos by the end of the day since their death is related to injection trauma rather than tumor growth. Remove&#160;from analysis embryos that do not retain cells&#160;since the cells likely leaked out of the injection site.</li>
	<li>Maintain the injected embryos at 34 &#176;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.</li>
</ol>
9. Survival analysis
<ol>
	<li>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.</li>
	<li>Check embryo health and score death for the duration of the analysis. 
	NOTE: The experimental duration was 7 days for this&#160;experiment&#160;but may be shorter or longer depending on the aggressiveness of the xenotransplanted tumor.</li>
	<li>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.</li>
</ol>
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.
<ol>
	<li>Isolate embryos at the dpi stage of choice. 5 dpi is displayed here (Figure 5).</li>
	<li>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.&#160;Anesthetize the embryos as described above. 
	NOTE: Embryos may be subdivided as replicates to provide statistical significance, as shown here (Figure 5B).</li>
	<li>Transfer the embryos to 1.5 mL centrifuge tubes.</li>
	<li>Use 100 &#956;L of calcium-free Ringer&#39;s solution (recipe31) per sample to dissolve the yolk since low calcium softens the embryonic tissues, enabling more effective tissue dissociation.</li>
	<li>Pipette up and down intermittently for 5 min to remove the yolk using a 200 &#956;L pipette tip.</li>
	<li>Pre-heat 0.05% trypsin/PBS (without phenol red indicator) to 29 &#176;C and supplement it with 27 &#956;L 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.</li>
	<li>Add 1 mL of the trypsin/collagenase solution to each sample of deyolked embryos and incubate at 29 &#176;C for 30-35 min.</li>
	<li>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.</li>
	<li>Stop the reaction using 200 &#956;L of FBS.</li>
	<li>Mix well and incubate at 29 &#176;C for an additional 5 min to ensure complete inactivation of the trypsin. 
	NOTE: A temperature of 29 &#176;C is employed for the tissue dissociation protocol to prevent heat shock-induced death of zebrafish cells, which occurs at 37 &#176;C; however, if preservation of zebrafish cells is not required, digestion can be performed at 37 &#176;C.</li>
	<li>Pellet the cell suspension at 300 x g for 5 min at 4 &#176;C and discard the supernatant.</li>
	<li>Resuspend the cell pellet in 4 &#176;C PBS and pellet as above.</li>
	<li>Repeat the wash, then strain the cells through a 70 &#181;m cell strainer.</li>
	<li>Pellet and proceed with staining for flow cytometry analysis. 
	NOTE: If culture of primary zebrafish cells is required, wash twice more with 4 &#176;C PBS and resuspend in L15 media (with antibiotics and 10% FBS).</li>
</ol>
11. Fluorescence-activated cell sorting (FACS): Staining and sorting of xenotransplanted cells
<ol>
	<li>Resuspend the cell suspension in a staining medium (HBSS with 1% FBS) and pellet at 300 x g for 5 min.</li>
	<li>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 &#181;L of anti-mouse CD45 (APC-CD45) per sample was employed at a 1:50 dilution (Figure 5).</li>
	<li>Incubate for 20 min at 4 &#176;C before washing as above with 1 mL of the staining medium to remove the unbound antibody.</li>
	<li>After discarding the supernatant, resuspend the cell pellet in 200 &#181;L of staining medium containing the vital dye Helix NP Blue (1 &#181;M), which will enable live/dead discrimination.</li>
	<li>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.</li>
</ol>
12. Flow cytometry
<ol>
	<li>Turn on the flow cytometer that is available using a low flow rate (500 events per second or less) to set the parameters.</li>
	<li>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.</li>
	<li>Use the uninjected embryo sample to establish settings that accommodate both transplanted and zebrafish cells.</li>
	<li>Increase the flow rate to 8000 events per second and record 1 million events for each sample.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>If needed, sort the tumor cells in the embryos by flow cytometry for downstream molecular analysis.</li>
</ol>

Utilizing Larval Zebrafish for Efficient In Vivo Tumor Studies

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(174), e62767 (2021).</li><li>Martinez-Lopez, M., P&#243;voa, V., Fior, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Generation+of+zebrafish+larval+xenografts+and+tumor+behavior+analysis.">Generation of zebrafish larval xenografts and tumor behavior analysis.</a> J Vis Exp. (172), e62373 (2021).</li><li>Ren, J., Liu, S., Cui, C., Ten Dijke, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Invasive+behavior+of+human+breast+cancer+cells+in+embryonic+zebrafish.">Invasive behavior of human breast cancer cells in embryonic zebrafish.</a> J Vis Exp. (122), e55459 (2017).</li><li>Zhao, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+xenograft+model+in+zebrafish+for+high-resolution+investigating+dynamics+of+neovascularization+in+tumors.">A novel xenograft model in zebrafish for high-resolution investigating dynamics of neovascularization in tumors.</a> PloS One. 6 (7), e21768 (2011).</li><li>Cabezas-S&#225;inz, P., Pensado-L&#243;pez, A., S&#225;inz Jr, B., S&#225;nchez, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modeling+cancer+using+zebrafish+xenografts:+drawbacks+for+mimicking+the+human+microenvironment.">Modeling cancer using zebrafish xenografts: drawbacks for mimicking the human microenvironment.</a> Cells. 9 (9), 1978 (2020).</li><li>Haraoka, Y., Akieda, Y., Ishitani, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live-imaging+analyses+using+small+fish+models+reveal+new+mechanisms+that+regulate+primary+tumorigenesis.">Live-imaging analyses using small fish models reveal new mechanisms that regulate primary tumorigenesis.</a> Yakugaku Zasshi. 139 (5), 733-741 (2019).</li><li>Westerfield, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The Zebrafish Book. A Guide for the Laboratory Use of Zebrafish (Danio rerio). , (2000).</li><li>Rao, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Inactivation+of+ribosomal+protein+L22+promotes+transformation+by+induction+of+the+stemness+factor,+Lin28B.">Inactivation of ribosomal protein L22 promotes transformation by induction of the stemness factor, Lin28B.</a> Blood. 120 (18), 3764-3773 (2012).</li><li>Goel, M. K., Khanna, P., Kishore, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Understanding+survival+analysis:+Kaplan-Meier+estimate.">Understanding survival analysis: Kaplan-Meier estimate.</a> Int J Ayurveda Res. 1 (4), 274-278 (2010).</li><li>Usai, A., Di Franco, G., Gabellini, C., Morelli, L., Raffa, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Establishment+of+zebrafish+patient-derived+xenografts+from+pancreatic+cancer+for+chemosensitivity+testing.">Establishment of zebrafish patient-derived xenografts from pancreatic cancer for chemosensitivity testing.</a> J Vis Exp. (195), e63744 (2023).</li><li>Murali Shankar, N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preclinical+assessment+of+CAR-NK+cell-mediated+killing+efficacy+and+pharmacokinetics+in+a+rapid+zebrafish+xenograft+model+of+metastatic+breast+cancer.">Preclinical assessment of CAR-NK cell-mediated killing efficacy and pharmacokinetics in a rapid zebrafish xenograft model of metastatic breast cancer.</a> Front Immunol. 14, 1254821 (2023).</li><li>Takahi, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Xenograft+of+human+pluripotent+stem+cell-derived+cardiac+lineage+cells+on+zebrafish+embryo+heart.">Xenograft of human pluripotent stem cell-derived cardiac lineage cells on zebrafish embryo heart.</a> Biochem Biophys Res Commun. 674, 190-198 (2023).</li><li>Rudner, L. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Shared+acquired+genomic+changes+in+zebrafish+and+human+T-ALL.">Shared acquired genomic changes in zebrafish and human T-ALL.</a> Oncogene. 30 (41), 4289-4296 (2011).</li><li>Regan, J. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=RNA+sequencing+of+long-term+label-retaining+colon+cancer+stem+cells+identifies+novel+regulators+of+quiescence.">RNA sequencing of long-term label-retaining colon cancer stem cells identifies novel regulators of quiescence.</a> iScience. 24 (6), 102618 (2021).</li></ol>

The authors declare no competing interests.

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...

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 &#176;C, while mammalian cells grow at 37 &#176;C. Consequently, a compromise temperature must be employed that is tolerated by the fish yet supports tumor growth, and 34 &#176;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).

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1-phenyl 2-thiourea (PTU)</td>
			<td>Sigma</td>
			<td>P7629</td>
			<td></td>
		</tr>
		<tr>
			<td>70 micron cell strainer</td>
			<td>Corning&#160;</td>
			<td>CLS431751-50EA</td>
			<td></td>
		</tr>
		<tr>
			<td>90 mm Petri dish</td>
			<td>Thermo Fisher Scientific</td>
			<td>S43565</td>
			<td></td>
		</tr>
		<tr>
			<td>Agarose</td>
			<td>Apex bioresearch</td>
			<td>20-102GP</td>
			<td></td>
		</tr>
		<tr>
			<td>APC APC anti-mouse CD45.2 Antibody</td>
			<td>Biolegend</td>
			<td>109814</td>
			<td></td>
		</tr>
		<tr>
			<td>BD FACSymphony A5 Cell Analyzer</td>
			<td>BD Biosciences</td>
			<td>BD FACSymphony A5</td>
			<td></td>
		</tr>
		<tr>
			<td>calibration capillaries</td>
			<td>Sigma&#160;</td>
			<td>P1424-1PAK</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell tracker CM-dil dye</td>
			<td>Invitrogen</td>
			<td>C7001</td>
			<td></td>
		</tr>
		<tr>
			<td>Collageanse IV</td>
			<td>Gibco</td>
			<td>17104019</td>
			<td></td>
		</tr>
		<tr>
			<td>Dumont forceps number 55</td>
			<td>Fine science tools</td>
			<td>11255-20</td>
			<td></td>
		</tr>
		<tr>
			<td>FBS</td>
			<td>Corning&#160;</td>
			<td>35-015-CV</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorescence microscope</td>
			<td>Nikon</td>
			<td>model SMZ1500</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass capillaries (Borosilicate)</td>
			<td>World precision instruments</td>
			<td>1B100-4</td>
			<td></td>
		</tr>
		<tr>
			<td>HBSS</td>
			<td>Corning&#160;</td>
			<td>21-023-CV</td>
			<td></td>
		</tr>
		<tr>
			<td>Helix NP Blue</td>
			<td>Biolegend</td>
			<td>425305</td>
			<td></td>
		</tr>
		<tr>
			<td>Instant Ocean Sea Salt</td>
			<td>Instant ocean</td>
			<td>SS15-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Light microscope</td>
			<td>Nikon</td>
			<td>model SMZ1000</td>
			<td></td>
		</tr>
		<tr>
			<td>Methylene blue</td>
			<td>Sigma</td>
			<td>M9140-100G</td>
			<td></td>
		</tr>
		<tr>
			<td>Microloader (long tips for laoding cells)</td>
			<td>eppendorf</td>
			<td>930001007</td>
			<td></td>
		</tr>
		<tr>
			<td>P1000 micropipette puller</td>
			<td>Sutter instruments</td>
			<td>model P-97</td>
			<td></td>
		</tr>
		<tr>
			<td>PM 1000 cell microinjector</td>
			<td>MicroData Instruments, Inc. (MDI)</td>
			<td>PM1000</td>
			<td></td>
		</tr>
		<tr>
			<td>Tricaine methanesulphate (Ethyl 3- aminobenzoate methanesulphate)</td>
			<td>Sigma</td>
			<td>E10521-10G</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypsin-EDTA (0.5%), no phenol red</td>
			<td>Gibco</td>
			<td>15400054</td>
			<td></td>
		</tr>
		<tr>
			<td>Zebrafish adult irradiated diet (dry feed)</td>
			<td>Zeigler</td>
			<td>388763</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

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 h...

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.

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 ...

a-simple-rapid-effective-method-for-tumor-xenotransplantation

Research

JoVE Journal

Cancer Research

1.1K Views.  Fox Chase Cancer Center. 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.

Video: Author Spotlight: Advancing Cancer Therapeutics Using Tumor Xenotransplantation in Zebrafish

A Mimic of the Tumor Microenvironment: A Simple Method for Generating Enriched Cell Populations and Investigating Intercellular Communication

Understanding the early heterotypic interactions between cancer cells and the surrounding non-cancerous stroma is important in elucidating the events leading to stromal activation and establishment of the tumor microenvironment (TME). Several in vitro and in vivo models of the TME have been developed; however, in general these models do not readily permit isolation of individual cell populations, under non-perturbing conditions, for further study. To circumvent this difficulty, we have employed an in vitro TME model using a cell growth substrate consisting of a permeable microporous membrane insert that permits simple generation of highly enriched cell populations grown intimately, yet separately, on either side of the insert&#39;s membrane for extended co-culture times. Through use of this model, we are capable of generating greatly enriched cancer-associated fibroblast (CAF) populations from normal diploid human fibroblasts following co-culture (120 hr) with highly metastatic human breast carcinoma cells, without the use of fluorescent tagging and/or cell sorting. Additionally, by modulating the pore-size of the insert, we can control for the mode of intercellular communication (e.g., gap-junction communication, secreted factors) between the two heterotypic cell populations, which permits investigation of the mechanisms underlying the development of the TME, including the role of gap-junction permeability. This model serves as a valuable tool in enhancing our understanding of the initial events leading to cancer-stroma initiation, the early evolution of the TME, and the modulating effect of the stroma on the responses of cancer cells to therapeutic agents.

We adapted a permeable microporous membrane insert to mimic the tumor microenvironment (TME). The model consists of a mixed cell culture, allows simplified generation of highly enriched individual cell populations without using fluorescent tagging or cell sorting, and permits studying intercellular communication within the TME under normal or stress conditions.

Understanding the early heterotypic interactions between cancer cells and the surrounding non-cancerous stroma is important in elucidating the events ...

The Drosophila Imaginal Disc Tumor Model: Visualization and Quantification of Gene Expression and Tumor Invasiveness Using Genetic Mosaics

Drosophila melanogaster has emerged as a powerful experimental system for functional and mechanistic studies of tumor development and progression in the context of a whole organism. Sophisticated techniques to generate genetic mosaics facilitate induction of visually marked, genetically defined clones surrounded by normal tissue. The clones can be analyzed through diverse molecular, cellular and omics approaches. This study describes how to generate fluorescently labeled clonal tumors of varying malignancy in the eye/antennal imaginal discs (EAD) of Drosophila larvae using the Mosaic Analysis with a Repressible Cell Marker (MARCM) technique. It describes procedures how to recover the mosaic EAD and brain from the larvae and how to process them for simultaneous imaging of fluorescent transgenic reporters and antibody staining. To facilitate molecular characterization of the mosaic tissue, we describe a protocol for isolation of total RNA from the EAD. The dissection procedure is suitable to recover EAD and brains from any larval stage. The fixation and staining protocol for imaginal discs works with a number of transgenic reporters and antibodies that recognize Drosophila proteins. The protocol for RNA isolation can be applied to various larval organs, whole larvae, and adult flies. Total RNA can be used for profiling of gene expression changes using candidate or genome-wide approaches. Finally, we detail a method for quantifying invasiveness of the clonal tumors. Although this method has limited use, its underlying concept is broadly applicable to other quantitative studies where cognitive bias must be avoided.

The Drosophila Imaginal Disc Tumor Model: Visualization and Quantification of Gene Expression and Tumor Invasiveness Using Genetic Mosaics

This protocol demonstrates how to generate fluorescently marked, genetically defined clonal tumors in the Drosophila eye/antennal imaginal discs (EAD). It describes how to dissect the EAD and brain from the third instar larvae and how to process them to visualize and quantify gene expression changes and tumor invasiveness.

Drosophila melanogaster has emerged as a powerful experimental system for functional and mechanistic studies of tumor development and progression in the ...

A Protocol for Rapid Post-mortem Cell Culture of Diffuse Intrinsic Pontine Glioma (DIPG)

Diffuse Intrinsic Pontine Glioma (DIPG) is a childhood brainstem tumor that carries a universally fatal prognosis. Because surgical resection is not a viable treatment strategy and biopsy is not routinely performed, the availability of patient samples for research is limited. Consequently, efforts to study this disease have been challenged by a paucity of faithful disease models. To address this need, we describe here a protocol for the rapid processing of post-mortem autopsy tissue samples in order to generate durable patient-derived cell culture models that can be used in in vitro assays or in vivo orthotopic xenograft experiments. These models can be used to screen for potential drug targets and to study fundamental pathobiological processes within DIPG. This protocol can further be extended to analyze and isolate tumor and microenvironmental cells using Fluorescence-activated Cell Sorting (FACS), which enables subsequent analysis of gene expression, protein expression, or epigenetic modifications of DNA at the bulk cell or single cell level. Finally, this protocol can also be adapted to generate patient-derived cultures for other central nervous system tumors.

A Protocol for Rapid Post-mortem Cell Culture of Diffuse Intrinsic Pontine Glioma DIPG

This protocol describes a method for the rapid processing of post-mortem diffuse intrinsic pontine glioma samples for the establishment of patient-derived cell culture models or direct characterization of tumor and microenvironmental cells.

Diffuse Intrinsic Pontine Glioma (DIPG) is a childhood brainstem tumor that carries a universally fatal prognosis. Because surgical resection is not a ...

Moving Upwards: A Simple and Flexible In Vitro Three-dimensional Invasion Assay Protocol

Although 3D invasion assays have been developed, the challenge remains to study cells without affecting the integrity of their microenvironment. Traditional 3D assays such as the Boyden Chamber require that cells are displaced from the original culture location and moved to a new environment. Not only does this disrupt the cellular processes that are intrinsic to the microenvironment, but it often results in a loss of cells. These problems are especially challenging when dealing with cells that are either rare, or extremely sensitive to their microenvironment. Here, we describe the development of a 3D invasion assay that avoids both concerns. In this assay, cells are plated within a small well and an ECM matrix containing a chemoattractant is laid atop the cells. This requires no cell displacement, and allows the cells to invade upwards into the matrix. In this assay, cell invasion as well as cell morphology can be assessed within the collagen gel. Using this assay, we characterize the invasive capacity of rare and sensitive cells; the hybrid cells resulting from fusion between breast cancer cells MCF7 and mesenchymal/multipotent stem/stroma cells (MSCs).

Moving Upwards: A Simple and Flexible In Vitro Three-dimensional Invasion Assay Protocol

This protocol describes an inverted vertical invasion assay that could be used to quantify the migration and invasion capabilities of cells in a three-dimensional setting while preserving the cell microenvironment. This assay is suitable for rare and/or sensitive cells.

Although 3D invasion assays have been developed, the challenge remains to study cells without affecting the integrity of their microenvironment. ...

Simple and Rapid Method to Obtain High-quality Tumor DNA from Clinical-pathological Specimens Using Touch Imprint Cytology

It is critical to determine the mutational status in cancer before administration and treatment of specific molecular targeted drugs for cancer patients. In the clinical setting, formalin-fixed paraffin-embedded (FFPE) tissues are widely used for genetic testing. However, FFPE DNA is generally damaged and fragmented during the fixation process with formalin. Therefore, FFPE DNA is sometimes not adequate for genetic testing because of low quality and quantity of DNA. Here we present a method of touch imprint cytology (TIC) to obtain genomic DNA from cancer cells, which can be observed under a microscope. Cell morphology and cancer cell numbers can be evaluated using TIC specimens. Furthermore, the extraction of genomic DNA from TIC samples can be completed within two days. The total amount and quality of TIC DNA obtained using this method was higher than that of FFPE DNA. This rapid and simple method allows researchers to obtain high-quality DNA for genetic testing (e.g., next generation sequencing analysis, digital PCR, and quantitative real time PCR) and to shorten the turnaround time for reporting results.

Obtaining high-quality genomic DNA from tumor tissues is an essential first step for analyzing genetic alterations using next generation sequencing. In this article, we present a simple and rapid method to enrich tumor cells and obtain intact DNA from touch imprint cytology specimens.

It is critical to determine the mutational status in cancer before administration and treatment of specific molecular targeted drugs for cancer patients. ...

Digital Polymerase Chain Reaction Assay for the Genetic Variation in a Sporadic Familial Adenomatous Polyposis Patient Using the Chip-in-a-tube Format

The quantitative analysis of human genetic variation is crucial for understanding the molecular characteristics of serious medical conditions, such as tumors. Because digital polymerase chain reactions (PCR) enable the precise quantification of DNA copy number variants, they are becoming an essential tool for detecting rare genetic variations, such as drug-resistant mutations. It is expected that molecular diagnoses using digital PCR (dPCR) will be available in clinical practice in the near future; thus, how to efficiently conduct dPCR with human genetic material is a hot topic. Here, we introduce a method to detect Adenomatous polyposis coli (APC) somatic mosaicism using dPCR with the chip-in-a-tube format, which allows eight dPCR reactions to be simultaneously conducted. Care should be taken when filling and sealing the reaction mixture on the chips. This article demonstrates how to avoid the over- and underestimation of positive partitions. Furthermore, we present a simple procedure for collecting the dPCR product from the partitions on the chips, which can then be used to confirm the specific amplification. We hope that this methods report will help promote the dPCR with the chip-in-a-tube method in genetic research.

Digital polymerase chain reaction (PCR) is a useful tool for the high-sensitivity detection of single nucleotide variants and DNA copy number variants. Here, we demonstrate key considerations for measuring rare variants in the human genome using digital PCR with the chip-in-a-tube format.

The quantitative analysis of human genetic variation is crucial for understanding the molecular characteristics of serious medical conditions, such as ...

Analysis of Side Population in Solid Tumor Cell Lines

Cancer stem cells (CSCs) are an important cause of tumor growth, metastasis, and recurrence. Isolation and identification of CSCs are of great significance for tumor research. Currently, several techniques are used for the identification and purification of CSCs from tumor tissues and tumor cell lines. Separation and analysis of side population (SP) cells are two of the commonly used methods. The methods rely on the ability of CSCs to rapidly expel fluorescent dyes, such as Hoechst 33342. The efflux of the dye is associated with the ATP-binding cassette (ABC) transporters and can be inhibited by ABC transporter inhibitors. Methods for staining cultured tumor cells with Hoechst 33342 and analyzing the proportion of their SP cells by flow cytometry are described. This assay is convenient, fast, and cost-effective. Data generated in this assay can contribute to a better understanding of the effect of genes or other extracellular and intracellular signals on the stemness properties of tumor cells.

A convenient, fast, and cost-effective method to measure the proportion of side population cells in solid tumor cell lines is presented.

Cancer stem cells (CSCs) are an important cause of tumor growth, metastasis, and recurrence. Isolation and identification of CSCs are of great ...

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 ...

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 ...

Automated Dissection Protocol for Tumor Enrichment in Low Tumor Content Tissues

Tumor enrichment in low tumor content tissues, those below 20% tumor content depending on the method, is required to generate quality data reproducibly with many downstream assays such as next generation sequencing. Automated tissue dissection is a new methodology that automates and improves tumor enrichment in these common, low tumor content tissues by decreasing the user-dependent imprecision of traditional macro-dissection and time, cost, and expertise limitations of laser capture microdissection by using digital image annotation overlay onto unstained slides. Here, digital hematoxylin and eosin (H&#38;E) annotations are used to target small tumor areas using a blade that is 250 &#181;m2 in diameter in unstained formalin fixed paraffin embedded (FFPE) or fresh frozen sections up to 20 &#181;m in thickness for automated tumor enrichment prior to nucleic acid extraction and whole exome sequencing (WES). Automated dissection can harvest annotated regions in low tumor content tissues from single or multiple sections for nucleic acid extraction. It also allows for capture of extensive pre- and post-harvest collection metrics while improving accuracy, reproducibility, and increasing throughput with utilization of fewer slides. The described protocol enables digital annotation with automated dissection on animal and/or human FFPE or fresh frozen tissues with low tumor content and could also be used for any region of interest enrichment to boost adequacy for downstream sequencing applications in clinical or research workflows.

Digital annotation with automated tissue dissection provides an innovative approach to enriching tumor in low tumor content cases and is adaptable to both paraffin and frozen tissue types. The described workflow improves accuracy, reproducibility and throughput and could be applied to both research and clinical settings.

Tumor enrichment in low tumor content tissues, those below 20% tumor content depending on the method, is required to generate quality data reproducibly ...