Identification of Quiescent Cells in a Zebrafish T-Cell Acute Lymphoblastic Leukemia Model Using Cell Proliferation Staining

Abstract

Cellular quiescence is a state of growth arrest or slowed proliferation that is described in normal and cancer stem cells (CSCs). Quiescence may protect CSCs from antiproliferative chemotherapy drugs. In T-cell acute lymphoblastic leukemia (T-ALL) patient-derived xenograft (PDX) mouse models, quiescent cells are associated with treatment resistance and stemness. Cell proliferation dyes are popular tools for the tracking of cell division. The fluorescent dye is covalently anchored into amine groups on the membrane and macromolecules inside the cell. This allows for the tracking of labeled cells for up to 10 divisions, which can be resolved by flow cytometry.

Ultimately, cells with the highest proliferation rates will have low dye retention, as it will be diluted with each cell division, while dormant, slower-dividing cells will have the highest retention. The use of cell proliferation dyes to isolate dormant cells has been optimized and described in T-ALL mouse models. Complementary to the existing mouse models, the rag2:Myc-derived zebrafish T-ALL model provides an excellent venue to interrogate self-renewal in T-ALL due to the high frequency of leukemic stem cells (LSCs) and the convenience of zebrafish for large-scale transplant experiments.

Here, we describe the workflow for the staining of zebrafish T-ALL cells with a cell proliferation dye, optimizing the concentration of the dye for zebrafish cells, passaging successfully stained cells in vivo, and the collection of cells with varying levels of dye retention by live cell sorting from transplanted animals. Given the absence of well-established cell surface makers for LSCs in T-ALL, this approach provides a functional means to interrogate quiescent cells in vivo. For representative results, we describe the engraftment efficiency and the LSC frequency of high and low dye-retaining cells. This method can help investigate additional properties of quiescent cells, including drug response, transcriptional profiles, and morphology.

Introduction

Adult stem cells are responsible for the regeneration of differentiated cell types in a given organ and are predominantly present in a dormant, non-dividing state^1,2. For example, hematopoietic stem cells (HSCs), which maintain the blood, largely remain quiescent, and only a small fraction enters the cell cycle to self-renew or differentiate to generate mature blood components³. Similarly, in cancers, a rare subpopulation of cells called cancer stem cells (CSCs) possess the self-renewal ability and are responsible for the long-term maintenance of the malignancy⁴. Cancer stem cells exist in vivo in a state of quiescence or slow growth, which may enable them to escape the anti-proliferative cancer treatments⁵, evade clearance by the immune system⁶, reduce oxidative stress, and enhance their DNA repair pathways⁷. Even a low number of CSCs left behind after treatment can potentially repopulate the tumor, resulting in a patient's relapse⁸. Accordingly, understanding cellular quiescence holds great promise for the identification of potential vulnerabilities of CSCs and the development of new ways to target them.

Cell proliferation dyes, such as the carboxyfluorescein succinimidyl ester (CFSE) stain and its derivatives, are commonly used to track the frequency of cell divisions⁹. The dye permeates across the cell membrane and, once inside the cell, undergoes activation by intracellular esterases into a fluorescent product. The resultant fluorescent compound is retained inside the cell through the covalent amide bonds formed between the succinimidyl moiety and the amine functional groups of intracellular proteins¹⁰. With each cellular division, the fluorescent compound is divided equally between the two resulting cells, causing a two-fold signal dilution. This dye enables the detection of up to 10 cell divisions through flow cytometry analysis¹¹.

This approach has been previously utilized to enrich CSC populations in vitro by identifying slow-cycling populations of cells with high retention of the dye^11,12. In T-ALL, CFSE has been used to track tumor growth in vivo in patient-derived xenografts in mice. Following cell labeling and three weeks of transplant, flow cytometry analysis showed a rare population of cells that still retained CFSE fluorescence. This population was associated with stemness, treatment resistance, and high similarity to relapse-causing cells in patients¹³. Accordingly, this dye provides a useful tool for the study of leukemia stem cell (LSC) phenotypes in T-ALL.

The aim of the work is to extend the application of the cell proliferation dye to study quiescence in vivo using a zebrafish T-ALL model. In particular, the rag2:Myc-driven zebrafish T-ALL model¹⁴ provides an excellent venue for the study of self-renewal due to the high frequency of LSCs compared to mouse models and human disease¹⁵. In addition, the use of zebrafish allows for large-scale transplantation studies, which can be done at a much lower cost of care and maintenance compared to their mouse counterparts¹⁶. Zebrafish are also excellent for live imaging applications, as fluorescently labeled tumor cells can be readily viewed using a simple fluorescence microscope to estimate the rate of tumor development¹⁶.

In this protocol, we describe the workflow for staining zebrafish T-ALL cells with cell proliferation dye followed by in vivo propagation of stained cells in syngeneic CG1 zebrafish. Upon leukemia development, we describe the sorting of cells that retained the dye and their use for a subsequent limiting dilution transplantation experiment to quantify rates of LSC self-renewal. This protocol can be extended for additional applications, including in vivo drug screening of potential compounds for the targeting of quiescent LSCs. In addition, collected cells can be used for different downstream analyses, such as transcriptomic profiling, proteomics, and metabolomics, offering unique insights into the behavior of quiescent LSCs in T-ALL.

Protocol

In this protocol, we are using GFP-labeled zebrafish T-ALL cells that were generated previously in the CG1 strain and thus can be directly injected into recipient syngeneic CG1 zebrafish¹⁵. Briefly, leukemia was generated by DNA microinjection of rag2:Myc and rag2:GFP into single-cell CG1 zebrafish embryos. Animals were monitored for leukemia development starting at 3 weeks post injection, using fluorescence microscopy. GFP-positive leukemia cells were FACS isolated and serially transplanted in recipient CG1 zebrafish to generate clones with a high frequency of LSC. Details of the entire protocol are described by Blackburn et al.¹⁵.

Alternatively, primary rag2:Myc-derived T-ALL can be generated by DNA microinjection of zebrafish embryos¹⁷. The DNA microinjection of rag2:Myc with a rag2-driven fluorescent reporter such as GFP can result in the development of B-cell, T-cell, and mixed ALL¹⁸. The leukemia clones used in this protocol were previously verified to be T-ALL¹⁵. All experimental procedures involving zebrafish were reviewed and approved by the University of Kentucky's Institutional Animal Care and Use Committee, protocol 2019-3399.

1. Propagating zebrafish T-ALL cells in vivo

1. Thaw one vial containing 1 mL of frozen GFP-labeled zebrafish T-ALL cells in a 37 °C water bath with gentle shaking, until thawed. Once thawed, transfer to ice.
2. Slowly pipette the contents of the tube into a 15 mL conical tube with 4 mL of fish media (0.9x PBS + 5% FBS) to dilute the freezing media.
3. Spin down for 5 min at 2,500 × g at 4 °C. Remove media from the pellet. Resuspend in 0.5 mL of fish media and keep the cells on ice.
4. Count the cells and dilute them if needed to 100,000 cells/5 µL.
5. Anesthetize the adult CG1 zebrafish by adding 200 µL of 4 mg/mL anesthetic (MS-222) to 25 mL of fish system water in a Petri dish. Hold the CG1 fish, ventral side up and inject 5-6 µL into the intraperitoneal (IP) space using a Hamilton microliter syringe. Move the animals back to their tanks and monitor for a return to normal swimming before putting them back on the system.
   NOTE: Adult zebrafish, 60 days or older were used for the transplantation studies described in this protocol.
6. Monitor leukemia growth once per week using a fluorescence microscope. See Section 2 below for details of the screening procedure.

2. Harvesting of fluorescently labeled leukemia cells

1. Approximately 21-28 days post-transplant, anesthetize the adult CG1 zebrafish by adding 200 µL (4-5 drops) of 4 mg/mL anesthetic to 25 mL of fish system water in a Petri dish.
2. Examine the CG1 zebrafish for leukemia burden by assessing the percentage of the GFP-labeled leukemia cells in the animal with an epifluorescent microscope, using the appropriate filter to detect the GFP fluorescence. The GFP-positive leukemia is ready to be harvested if it has spread across >70% of the animal.
3. Euthanize the animal by adding 1 mL of 4 mg/mL anesthetic in a Petri dish containing 9 mL of fish system water. Allow all signs of life to cease; for instance, observe the fish for the cessation of opercular movement (~5 min).
4. Move the zebrafish into a new Petri dish and add 1 mL of fish media to serve as a buffer for the cells. Using a new razor blade, first decapitate the zebrafish, then macerate the tissue using the same razor blade. Pipet up and down to dislodge large clumps of cells.
5. Pass the cells through a 40 µm cell strainer into a 50 mL conical tube to dissociate into a single-cell suspension. Keep the cells on ice.

3. Optimization of the cell proliferation dye (CellTrace Far Red) concentration for staining and zebrafish cell viability

NOTE: For this protocol, and since the tumor cells are GFP-labeled, we used a far-red cell proliferation dye (CellTrace Far Red) to avoid spectral overlap, which will be referred to as CT-FR.

1. Prepare a 1 mM solution of the fluorescent dye immediately before use by adding the appropriate volume of DMSO as per the manufacturer's recommendations. Have working solutions of 1 mM, 0.5 mM, 0.1 mM, and 0.01 mM; dilute as needed in fish media to test cell viability following staining.
2. Count the cells and collect 1 × 10⁶ cells in 1.5 mL microcentrifuge tubes.
3. Centrifuge at 2,500 × g at 4 °C for 5 min. Remove the liquid and resuspend the cells in 1 mL of 0.9x PBS.
4. Split the cell suspension between four tubes of 250 µL each for treatment with CT-FR.
5. Add 250 µL of 0.9x PBS to bring the volume up to 500 µL each with approximately 250,000 cells/tube.
6. Add the required amounts of the working solutions described above, to each of the four tubes to achieve the following final concentrations 1 µM, 0.5 µM, 0.1 µM, and 0.05 µM. Incubate for 20 min at 37 °C, protected from light.
7. Pellet the cells by spinning down at 2,500 × g for 5 min at room temperature. Remove the liquid and wash the cells with 500 µL of fish media to remove the excess dye.
8. Pellet again, remove the liquid, and resuspend in 25 µL of fish media. Use 1 µL to count cells in a 1:10 dilution in 50% Trypan Blue.
9. Examine the cells under the microscope for cell viability; live cells will not retain the Trypan blue and appear bright under the microscope, while dead cells will take up the dye and appear dark. Examine the cells under the appropriate fluorescence channel first to identify tumor cells and then use the brightfield channel to assess cell viability.
10. Select the concentration that results in successful staining while maintaining cell viability (>90% trypan blue-negative cells).
    NOTE: For the zebrafish T-ALL cells tested in this experiment, 1 µM concentration was used for successful staining, and >90% of the cell viability.

4. Propagating CT-FR-stained T-ALL zebrafish cells in vivo

1. Using the optimized CT-FR concentration from Section 3, stain the zebrafish cells. Leave some tumor cells unstained for the generation of the GFP-labeled T-ALL tumor, as a GFP-positive control during FACS sorting. Dilute both stained and unstained cells to 50,000 cells/5 µL for transplant.
2. Anesthetize the adult CG1 zebrafish by adding 200 µL of 4 mg/mL anesthetic to 25 mL of fish system water in a Petri dish.
3. Inject 5 µL of the cell suspension into the IP cavity of the zebrafish as described in Section 1 for both the stained and the unstained cells. Inject 5-6 zebrafish per group to account for the potential mortality post injection.
4. Examine the animals by fluorescent microscopy under sedation 2x weekly for leukemia development.
   NOTE: After ~10-14 days, the leukemia should occupy about 30-40% of this body. Plan for leukemia harvest and subsequent sort within this time frame. Waiting for the tumor to expand to a large size might result in a greater dilution of the fluorescent cell dye and difficulty in detecting its signal. Consider optimization of the timing of tumor harvest for the tumor model chosen. In addition, more frequent tumor monitoring is recommended here to ensure the capture of the desired tumor size.

5. Preparing for sorting of CT-FR-stained zebrafish T-ALL cells

1. Prepare the following controls for the cell sorting: No color control (unstained, non-transgenic, single-cell suspension, from a wild type zebrafish), GFP-Labeled T-ALL cells, and newly stained CT-FR WT zebrafish cells (for setting a maximum threshold of Far-Red signal).
2. Preparing unstained and CT-FR controls
   1. Harvest wild-type (WT) fish cells by euthanizing a non-transplanted CG1 zebrafish by adding 1 mL of 4 mg/mL anesthetic in a Petri dish containing 9 mL of fish system water.
   2. Move the zebrafish into a new Petri dish and add 1 mL of fish media to serve as a buffer for the cells. Use a new razor blade to first decapitate and then macerate the zebrafish using the same razor blade; pipet up and down to dislodge large cell clumps.
   3. Pass the cells through a 40 µm cell strainer into a 50 mL conical tube to dissociate into a single-cell suspension. Keep the cells on ice.
   4. For the CT-FR control, transfer 250,000 WT cells into a microcentrifuge tube and pellet the cells by centrifugation at 2,500 × g for 5 min at room temperature. Remove the fish media and add 500 µL of 0.9x PBS+ 1% FBS.
   5. Add 0.5 µL of the 1 mM CT-FR dye stock solution for a final concentration of 1 µM. Gently pipet with 1,000 µL pipet to mix. Incubate at 37 °C for 20 min, protected from light.
   6. Remove the liquid and wash the cells with 500 µL of fish media to remove the excess dye.
   7. Pellet the cells by centrifugation again, remove the liquid, and resuspend the cells in 500 µL of 0.9x PBS+ 1% FBS and pass through a 35 µm filter cap into a FACS tube. Keep on ice.
3. Preparing the donor plate
   NOTE: The donor plate is a 96-well v-bottom plate into which the CT-FR-stained T-ALL cells from transplanted zebrafish will be sorted. Each well of the plate contains 5,000 unstained, WT CG1 cells in 50 µL of fish media to serve as a carrier to help pellet the sorted cells in subsequent steps.
   1. Count 500,000 WT CG1 cells and transfer them into a 15 mL conical tube.
   2. Dilute this cell stock to 100 cells/µL by adding 5 mL of fish media.
   3. Dispense 50 µL of the cell suspension into each well of the 96-well plate. Mix the cell suspension stock tube after each row to maintain cell suspension homogeneity. Keep the plate at 4 °C until sorting.
4. Preparing the transplanted fish samples
   1. Select transplanted fish at 30-40% T-ALL burden. Collect fish transplanted with both the CT-FR-stained cells and unstained GFP tumor cells.
   2. Harvest cells as described in Section 2.
      NOTE: Since the dye-retaining cells are expected to be present at a low abundance, prepare to sort a large number of cells from the CT-FR GFP sample.
   3. Dilute into appropriate concentration for sorting ~3-5 × 10⁶ cells/mL in 0.9x PBS + 1% FBS and pass through a 35 µm filter cap into a FACS tube. Add DAPI (1 mg/mL) at 1:1,000 dilution as a vital dye to exclude dead cells. Keep on ice.
   4. For the unstained GFP tumor cells, transfer 250,000 cells into a microcentrifuge tube and pellet at 2,500 × g for 5 min at room temperature. Remove the fish media, add 500 µL of PBS, and pass through a 35 µm filter cap into a FACS tube. Keep on ice.

6. Sorting

NOTE: Keep all tubes and the donor plate on ice the entire time except when used for sorting.

1. For sorting parameters and gates, start by applying the forward and side scatter gating to identify the cells of interest based on their size and eliminate cell debris. Next, use forward scatter height versus forward scatter area to exclude duplicate cells and keep only singlets. Finally, use the no-color and single-color controls to draw the quadrants to identify GFP-positive and CT-FR-positive cell populations.
   NOTE: The CT-FR control is expected to be much brighter than the samples for sorting since the staining for the control was done fresh with no cell divisions; cells from the CT-FR GFP samples are expected to appear at the lower end of the CT-FR positive gate.
2. Sort the number of cells to be transplanted into each fish into one well of the donor plate.
   NOTE: For this experiment, we used a limiting dilution assay to estimate the frequency of leukemia stem cells in CT-FR High and CT-FR Low. This assay used a dose of 25 cells, transplanted into 3 animals, and a dose of 10 cells, transplanted into 10 animals. A typical limiting dilution assay using the zebrafish rag2:Myc-derived T-ALL would involve a group with a higher number of cells (500 or 1,000) since the LSC frequency in this model is estimated to be 1 LSC in hundreds of cells. However, in this assay, we are using one of the T-ALL clones previously described by Blackburn et al.¹⁵. The LSC frequency in the sample used here is ~1 LSC in tens of cells; we predict leukemia to be detected in animals transplanted in the 25 cells group. When optimizing this assay for your disease model, care should be taken while selecting the higher cell number group to ensure the engraftment of the tumor.
3. After sorting, keep the plate on ice until ready for transplant.
   NOTE: Sorting procedures can be stressful and affect cell viability. Before running the actual limiting dilution experiment, a pilot sorting experiment was done. Sorted cells were transplanted and leukemia development was monitored to determine if the sorting was detrimental to cell viability. Alternatively, cell viability can be assessed by staining the sorted cells with Trypan blue and examining them under the microscope before transplant.

7. Transplanting sorted cells at limiting dilution and tracking tumor onset

1. Spin down the plate using a large benchtop centrifuge at 2,500 × g at 4 °C.
2. Carefully remove 45 µL of the supernatant from each well, leaving 5 µL liquid behind and making sure to avoid touching the cell pellet at the bottom of the well.
3. Resuspend the cells with a 20 µL pipet. Resuspend 2-3 wells with the pipet and then using the Hamilton microliter syringe, inject the cell suspensions from each well into the desired number of CG1 zebrafish via IP transplant. Then, resuspend the cells in the next group of wells to avoid resettling of the cells.
4. Examine the zebrafish by fluorescence microscopy under sedation once every week for T-ALL development for at least 6 weeks.

8. Determination of the leukemia stem cell frequency

1. Record the number of positive animals per cell dose and treatment group at 6 weeks post transplant, considering animals with any amount of GFP signal to be positive. Consider animals to have significant leukemia burden when the GFP signal occupies >50% of the body.
2. Use the web-based ELDA (Extreme Limiting Dilution Analysis) statistical software (http://bioinf.wehi.edu.au/software/elda/)¹⁹ to calculate the frequency of LSCs within each cell population.
   NOTE: A schematic illustration of the workflow is provided in Figure 1.

Figure 1: Workflow for the use of the cell tracking stain to isolate quiescent cells in the zebrafish T-ALL model. Schematic illustration of staining zebrafish T-ALL cells with the cell proliferation stain, propagation into a CG1 zebrafish (top panel), and sorting of cells based on the retention of the cell proliferation stain (middle panel). Collected cells were used for a limiting dilution assay to determine the frequency of LSCs (bottom panel). Abbreviations: T-ALL = T-cell acute lymphoblastic leukemia; LSCs = leukemic stem cells; CT-FR = CellTrace Far Red; FITC = fluorescein isothiocyanate; PE = phycoerythrin. Please click here to view a larger version of this figure.

Representative Results

We followed the above-described protocol to sort cells that have retained the cell proliferation dye, CT-FR, and used them for a Limiting dilution assay (LDA) to estimate LSC frequency in the CT-FR High and CT-FR Low populations. To set the gating for the flow cytometry experiment, we used a no fluorophore (no color) control in addition to single-color controls (Figure 2A). Those quadrants were used to identify FITC- and PE-positive and -negative populations for the setup of the sorting gates. The CT-FR single-color positive control was prepared on the same day of the sorting by staining WT cells from an un-transplanted zebrafish, and thus, the signal intensity is expected to be much higher than the signal from the propagated cells in the sample.

In CT-FR stained GPF T-ALL (Figure 2B), cells appearing in the upper right quadrant at a low frequency of 0.13% were considered CT-FR High, and a sorting gate labeled as "CT-FR high" was placed in this quadrant. Another gate was created in the lower right quadrant to collect leukemia cells that are CT-FR Low. Since the CT-FR High cells existed at low abundance in the tumor sample, a large number of cells (~2.5 × 10⁶) were sorted to collect the needed number of cells for the limiting dilution experiment.

After transplanting the CG1 zebrafish with the sorted cells for the LDA, zebrafish were monitored for 6 weeks. Cells from the CT-FR High had a four-fold higher LSC frequency compared to the CT-FR Low cell population (p = 0.0134) (Figure 3A). Representative images of animals at 28 days post-transplant are shown in (Figure 3B). In the CT-FR High, 10 cell dose group, 8 out of 9 animals (89%) of the animals developed leukemia, compared to 2 out of 10 (20%) in the CT-FR Low group (Figure 3A). Animals from the CT-FR High group had a significantly shorter latency to leukemia (p = 0.0014) (Figure 3C). Finally, animals from the CT-FR High had a significantly lower overall survival (p = 0.0408) (Figure 3D); at week 6, 70% of the CT-FR Low animals were alive compared to only 10% of the CT-FR High animals. These results demonstrate the enrichment of self-renewing LSCs within the slow-dividing, quiescent cell population in the rag2:Myc-driven zebrafish T-ALL model and the utilization of the cell proliferation stain to isolate this population.

Figure 2: Cell sorting using flow cytometry. (A) FACS plots from freshly CT-FR-stained WT cells and GFP-labeled T-ALL cells were used to create the gates for sorting CT-FR-stained. GFP⁺ T-ALL cells. (B) Representative example of the propagated CT-FR stained GFP⁺ T-ALL cells. Cells from the upper right quadrant were considered CT-FR High and cells from the lower right quadrant were considered CT-FR Low. Abbreviations: T-ALL = T-cell acute lymphoblastic leukemia; CT-FR = CellTrace Far Red; GFP = green fluorescent protein; FITC = fluorescein isothiocyanate; PE = phycoerythrin; FACS = fluorescence-activated cell sorting; WT = wild type. Please click here to view a larger version of this figure.

Figure 3: Significantly higher LSC frequency in cells with high retention of the cell proliferation stain. (A) Data from the limiting dilution assay stating the number of leukemia-positive animals post transplant with two increasing doses of cells (10 and 25 cells). The calculated LSC frequency and 95% confidence interval are stated (p = 0.0134). (B) Representative images of animals after 28 days of transplant; scale bar = 10 mm. (C) Latency to significant leukemia considered as the time needed for >50% of the animal to become GFP-positive, representing significant leukemia burden. (D) Survival curve of animal transplanted with 10 cells. Abbreviations: LSCs = leukemic stem cells; CT-FR = CellTrace Far Red. Please click here to view a larger version of this figure.

Discussion

LSCs are known to be resistant to conventional, anti-proliferative chemotherapy treatments, and finding targeted therapies against these cells holds great promise in reducing the occurrence of relapse and improving patient prognosis²⁰. Previous research described the use of fluorescent cell proliferation stains to identify a small population of quiescent cells associated with drug resistance and stemness in T-ALL PDX models¹³. In this work, we describe the use of a similar cell proliferation stain to resolve slow-dividing or quiescent cells in a zebrafish model of T-ALL. We used this tool to describe the enrichment of LSCs in the cells with the highest retention of the cell proliferation stain. The extension of these investigations to zebrafish offers numerous advantages. Zebrafish are highly fecund; each mating pair can generate hundreds of offspring every week, powering transplantation and limiting dilution studies that require a large number of animals^15,17,21. In addition, zebrafish are very convenient for in vivo imaging, enabling the rapid screening of positive animals by simple fluorescence microscopy¹⁶. Accordingly, this protocol can serve as the basis for additional investigation of quiescent cells in the zebrafish T-ALL model.

Here we describe the use of T-ALL clones with high frequency of LSCs that were previously generated in CG1 zebrafish¹⁵, thus, frozen cells were simply thawed and propagated for use. Alternatively, rag2:Myc-driven T-ALL can be generated by DNA microinjection of zebrafish embryos, screening for animals that develop T-ALL and harvesting the leukemia from positive zebrafish as previously described¹⁷. The use of the syngeneic zebrafish also offers numerous advantages for this type of research. Since the animals are genetically identical, transplanted cells can robustly engraft within the host without needing to adapt to a new micro-environment, enabling accurate estimation of the LSC frequency^15,17,22.

As an alternative to using syngeneic animals, this protocol can be followed using a wild-type zebrafish strain that has undergone chemical or radiation-mediated immune ablation. If using immune-ablated animals, care should be taken to optimize the limiting dilution assay since more cells may be needed for successful engraftment in allogenic transplantation. In addition, due to immune system recovery after 21 days, a shorter duration of observation will likely be needed¹⁷. This protocol may also be extended to the xenotransplantation of human cells. Zebrafish have emerged as an efficient and cost-effective model for cancer xenotransplants complementary to their mouse counterparts¹⁶. The adult immune-deficient prkdc^−/−, il2rga^−/− zebrafish have been used to successfully engraft human cells at 37 °C for up to 60 days²³. With this model, instead of propagating and having the stained cells divided in vivo as described here, human cells can be stained and allowed to divide in vitro then sorted and transplanted into a prkdc^−/−, il2rga^−/− zebrafish for in vivo studies. Several limitations need to be considered when using zebrafish as a xenograft model. For instance, zebrafish must be adapted to survive at 37 °C. In addition, orthotopic transplants might not be possible for certain cancers due to the absence of certain organs in the zebrafish, such as lung and breast¹⁶.

In conclusion, the use of the cell proliferation stains in the zebrafish T-ALL model provides a promising avenue for the investigation of quiescent LSCs and their roles in disease development and treatment resistance. Leveraging the advantages of the CG1 zebrafish, including their cost-effectiveness and genetic homogeneity, this protocol offers a robust platform for the understanding of the LSC biology in T-ALL. The use of the zebrafish model can complement mouse and PDX models to enable the development of optimized and targeted therapies that will ultimately improve the prognosis of T-ALL patients.

Materials

Name	Company	Catalog Number	Comments
26 G/2” micro-syringe	Hamilton	87930	NA
35 µm filter cap FACS tubes	Falcon	352235	NA
40 µm cell strainer	CELLTREAT	229482	NA
96-well skirted PCR plate	Thermo Fisher Scientific	AB0800	NA
Cell sorter	Sony Biotechnology	SY3200	NA
CellTrace Far Red	Thermo Fisher Scientific	C34564	NA
Conical tubes	VWR	10026-078	NA
DAPI	Thermo Fisher Scientific	62248	NA
DMSO	Sigma-Aldrich	D4818	NA
Dulbecco'sPhosphate-buffered saline (PBS)	Caisson Labs	22110001	NA
Epifluorescence stereo microscope	Nikon	SMZ25	NA
Fetal Bovine Serum (FBS)	Sigma-Aldrich	12306C	NA
Fish system water	N/A	N/A	0.03-0.05% salinity, pH 6.5-8, buffered with sodium bicarbonate
Microcentrifuge tubes	Thermo Fisher Scientific	C2171	NA
MS-222	Pentaire	TRS-1	tricaine mesylate, an anesthetic
Petri dishes	Corning	07-202-011	NA
Razor blades	American Line	66-0089	NA
Trypan Blue	Thermo Fisher Scientific	T10282	NA