Generation of Zebrafish Larval Xenografts and Tumor Behavior Analysis

The zebrafish (Danio rerio) model was handled and maintained according to the standard protocols of the European Animal Welfare Legislation, Directive 2010/63/EU (European Commission, 2016) and Champalimaud Fish Platform. All protocols were approved by the Champalimaud Animal Ethical Committee and Portuguese institutional organizations-ORBEA (Órgão de Bem-Estar e Ética Animal/Animal Welfare and Ethics Body) and DGAV (Direção Geral de Alimentação e Veterinária/Directorate General for Food and Veterinary).

NOTE: Before starting the main experiment, practice with the human colorectal cancer (CRC) cell line HCT116. This cell line is easy to prepare (highly proliferative), easy to inject and engrafts very efficiently (around 95-100%). Start with cells in excess (~12×10⁶ cells, T-75 flask) and excess fish (400 fish) until becoming proficient in the technique, since many cells and fish will be lost during training. Experimenters are ready once the engraftment of ~95% is achieved in HCT116 xenografts. See Figure 1 for a schematic of the complete protocol.

1. Setting up for injection

1. Two weeks before injection, expand cells in culture (see Table 1 for a detailed guide of the optimal in vitro confluence for injection of several cell lines).
2. Three days before the injection, cross the zebrafish of the desired background.

2. 24 h before injection

1. Clean the zebrafish embryo plates (discard all dead and non-developed embryos) and refresh E3 medium.
2. In the cell culture room, discard the cell culture medium from the flasks planned for injection, wash once with 1x phosphate-buffered saline (PBS) to remove the dead cells and add fresh medium.
3. Prepare the tools for the injection procedure, such as: microinjection needles, agarose plates and hairpins to align embryos for injection (Figure 2A-D, detailed below).
   1. Microinjection needles (Figure 2A)
      1. Use borosilicate glass capillaries (4 inches, OD 1.0 mm, No Filament in a micropipette puller (heat: ≈500; fil: 10; vel: 50; dec: 60; pull: 100).
   2. Plate preparation (Figure 2B)
      1. Prepare 2% agarose in H₂O, heat it up and pour one layer of dissolved agarose in the lid of a clean Petri dish. Let it polymerize and with the help of a ruler, make three to four straight agarose lines for the alignment of the embryos.
   3. Hairpin assembly (Figure 2C-D)
      1. Place 1 hair inside a glass capillary tube leaving approximately 1 centimeter of hair outside the tube.
      2. Curl the outside tip of the hair with the help of forceps into the glass capillary tube forming a loop of ~0.5 mm length.
      3. Seal the edge of the capillary tube with a drop of nail polish. This will also help fix the loop in place. Let it dry. Repeat the procedure on the other edge of the tube.
      4. Cut a piece of electrical tape (more resistant, impermeable, and rigid than regular adhesive tape).
      5. Seal the tape around the capillary to protect it from breaking.

3. Injection day

1. Separate the hatched embryos from unhatched eggs. Adding 1x pronase (0.6 mg/mL, Table 3) to the embryo medium at this stage can boost hatching. Place the embryos into the incubator (at 28 °C) until injection.
   NOTE: Do not leave the embryos in the pronase solution for longer than 1 hour, since the enzyme will act on the hatched embryos increasing their risk of mortality. Ensure that the developmental stage of the embryos is the one corresponding to 48 hpf (Figure 3A, A') to avoid the risk of edema and mortality.
2. Prepare 1x Tricaine (from a 25x stock).
   ​NOTE: A detailed recipe can be found in Table 3 and at the Zebrafish Information Network – ZFIN webpage¹⁷.

4. Cell labeling for injection

NOTE: Labelling of cells can be performed either directly in a flask or in a 1.5 mL microcentrifuge tube after enzymatic detachment. See Discussion for more information.

1. Remove the cell culture medium and wash the flask twice (2x) with 1X PBS.
2. Label the cells with a lipophilic dye of choice either directly in the flask (2 mL solution/T75 flask) or in a 1.5 mL microcentrifuge tube after enzymatic detachment. Avoid exposure to light and incubate cells at 37 °C (see Table 1 and Table 2 for conditions/solutions).
3. If labelling in the flask
   1. Remove the dye, wash with 1x PBS and detach the cells with EDTA and a cell scraper.
   2. Transfer cells to 1.5 mL microcentrifuge tubes. Centrifuge for 5 minutes at 300 x g then go to step 4.5.
4. If labelling in the 1.5 mL microcentrifuge tube:
   1. Centrifuge 5 minutes at 300 x g to remove the dye and discard the supernatant. Resuspend in 1x PBS to wash.
   2. Centrifuge 5 minutes at 300 x g and then go to step 4.5.
5. Discard the supernatant and resuspend the pellet with cell culture medium (for a 50 µL pellet add ~ 150-200 µL of medium).
6. Quantify cell viability using a Neubauer chamber with Trypan blue exclusion or other method of choice.
7. Centrifuge for 4 minutes at 300 x g and discard the supernatant. Resuspend the cells in the injection medium.
   NOTE: The recommended cell concentration (in general from 0.25-0.5×10⁶ cells/µL) and medium can be found in Table 1.
8. From this point onwards, keep the cells on ice.

5. Injection procedure

1. Anesthetize the embryos in 1x Tricaine for 5 minutes.
2. With a plastic Pasteur pipette, transfer a small amount (~50) of anesthetized embryos to the agarose plate and carefully align them with the help of a hairpin loop. Make sure to maintain distance between the embryos, especially between the yolk of one and the head of the next one (Figure 4A).
   NOTE: The number of embryos to align will vary according to the level of expertise of the researcher performing the injections. Start with a few (~10-20). For a schematic of the correct positioning of the embryos in the agar/agarose plate see Figure 4A.
3. Ensure the aligned embryos do not dry out in the agarose plate to prevent mortality, by carefully adding 1-3 drops of 1x Tricaine solution to the plate.
4. Lightly tap the microcentrifuge tube to resuspend the cells. Backload the injection needle with the cell suspension using a microloader tip avoiding air bubbles, as they can compromise the integrity of the embryos.
5. Open the air pressure valve (40 psi), set up the microinjector and carefully place the microinjection needle into the holder.
   NOTE: Use the following recommended microinjector settings: Hold pressure – vent (3 psi); Eject pressure – vent; Range – 100 ms.
6. Cut the microinjection needle close to the tip with Dumont forceps #5 or similar under the stereomicroscope.
   NOTE: The tip must be blunt and thin enough to allow the cells to pass without clogging as well as to avoid damaging the embryos and losing cells. A thick microinjection needle tip will injure the embryo and promote the formation of edema or zebrafish death. Graticule is not used for needle calibration. See Discussion for a detailed explanation.
7. Before injecting the embryos, test the microinjector pressure starting by the lowest eject pressure until in ~1-3 pulses a volume similar to the size of the zebrafish embryo's eye is achieved.
   NOTE: The use of a fluorescence stereomicroscope whenever possible at the beginning of the training is recommended. This will allow an easier identification of fluorescently labelled cells.
8. Carefully pierce in the middle of the embryo's yolk, lowering the angle of the needle and cautiously push until the tip of the needle reaches the perivitelline space (PVS) (Figure 4B-D).
9. Press the microinjector pedal and inject the cells into the PVS. Use the eye of the embryo as a guide. Try to inject a volume of cells similar to the size of the embryo's eye and as far as possible from the heart to prevent cardiac edema.
10. Carefully remove the needle and move onto the next embryo.
11. Adjust the microinjector pressure if needed.
    NOTE: Cells tend to start clogging, so pressure may be increased. If needed, it is possible to cut the capillary (to increase the diameter) while reducing the pressure.
12. Transfer the injected embryos to a clean Petri dish (Table 4) with 1x Tricaine solution and leave them to rest for 5-10 minutes. This will give time for the wound to close.
    NOTE: To transfer the embryos from the agar plate to the Petri dish add a few drops of 1x Tricaine solution on top of the embryos and carefully collect them with a plastic Pasteur pipette. Drop the collected embryos in a new Petri dish with 1x Tricaine solution.
13. Remove the 1x Tricaine solution and add fresh E3 medium.
14. Incubate the xenografts at 34 °C (a compromised temperature between human cell lines survival and zebrafish development⁸).

6. Metastatic assay

1. At approximately 1-hour post-injection (hpi), screen the injected embryos on a fluorescent stereomicroscope and sort the xenografts into 2 groups, according to the absence (Figure 5A) or presence (Figure 5B) of cells in circulation.
   ​NOTE: Even if only one cancer cell is detected in the heart or circulation, include these xenografts in the group of xenografts with cells in circulation. Alternatively, cells can be directly injected into circulation to increase the numbers of xenografts in this group.

7. 1 day post injection

1. On a fluorescent stereomicroscope carefully analyze each embryo and select those with properly injected tumors (Figure 6).
   NOTE: If needed, anesthetize the injected embryos with Tricaine 1X solution before screening.
2. Discard the following embryos/xenografts (Figure 6A-A''):
   •  without tumor/abnormal morphology/dead,
   •  with cardiac and/or yolk edema,
   •  with tumor cells only in the yolk,
   •  with very few cancer cells.
3. Sort selected xenografts according to their tumor size. Use the size of the eye for comparison (Figure 6B-B'', 6C).
   •  Tumors smaller than the size of the eye (+),
   •  Tumors the same size as the eye (++),
   •  Tumors larger than the size of the eye (+++).
4. Distribute the xenografts according to the desired experimental layout and start the drug assay (control vs drug, etc.). Replace the drugs and E3 medium daily (Table 4).
5. Incubate the xenografts maintaining the temperature of 34°C until the end of the assay.

8. 4 days post injection

1. On the final day of the assay, anesthetize the xenografts with 1x Tricaine solution and carefully align them on the agar plate.
   NOTE: Discard any dead or swollen xenografts. Drug treatments and some tumor cells can induce toxicity and eventually cause xenograft mortality. These xenografts are not considered for engraftment quantification.
2. To determine percentage of engraftment: on a fluorescent stereomicroscope analyze each live xenograft and assess the absence (Figure 6D) / presence (Figure 6E) of tumors in the PVS.
   
3. To determine percentage of metastasis: on a fluorescent stereomicroscope analyze each xenograft and assess the presence/absence of micrometastasis in the caudal hematopoietic tissue (CHT) of the 2 previously defined groups (CIRC and NO CIRC, Figure 6F)
   
4. According to the experimental set up, select the xenografts of interest and euthanize them with 25x Tricaine (Table 3).
5. Fix them in 4% formaldehyde (FA) for at least 4 hours at room temperature (RT) or overnight at 4 °C.
   NOTE: Use methanol-free formaldehyde (16% FA) diluted at 4% in PBS/0.1% Triton. Fill the tubes to the top with fixative. Position the tubes horizontally to ensure a homogeneous fixation of all xenografts, increasing permeability and preventing the zebrafish from aggregating at the bottom.
6. Alternatively fix them with PIPES (Per 1 mL: 100 µL of 1 M PIPES sodium salt (4 °C); 1 µL of 1 M MgSO₄ (RT); 4 µL of 0.5 M EGTA (RT); 93.7 µL of 16% FA (RT); 801.3 µL of ddH₂O).
   NOTE: PIPES preserves the fluorescence of RFP and mCherry transgenic lines better than 4% FA.
7. If the immunostaining will be performed on a different day, replace the FA with 100% methanol (MetOH). Xenografts fixed in 100% methanol can be stored at -20 °C indefinitely.
   ​NOTE: MetOH can impair the efficiency of some stainings (i.e., phalloidin) and quench some fluorescent labelling. Confirm the efficiency of the antibodies in MetOH fixed samples beforehand.

9. Whole mount immunostaining for confocal imaging

NOTE: The whole mount immunofluorescence technique takes 3 days divided as follows: The first day is for permeabilization of the xenografts and primary antibody incubation. The second day for washing and secondary antibody incubation and the third day for washing, fixation of xenografts and storage in the mounting media.

1. Day 1
   1. If the xenografts were stored in 100% MetOH, rehydrate them by a series of decreasing MetOH concentrations (75%, 50%, 25% MetOH diluted in PBS/0.1% Triton). If fixed in FA, replace it by PBS/0.1% Triton.
   2. Wash 4x for 5 minutes in PBS/0.1% Triton.
   3. Wash 1x for 5 minutes in H₂O.
      NOTE: The tubes must be positioned horizontally always in fixation, permeabilization and washing steps unless stated otherwise.
   4. Replace the H₂O for ice cold acetone and incubate at -20 °C for 7 minutes.
      NOTE: Place a 50 mL tube with acetone at -20 °C so it is ready to use. Microcentrifuge tubes must be positioned vertically on a rack so that the acetone does not leak out.
   5. Wash 2x for 10 minutes in PBS/0.1%Triton.
   6. Incubate with PBDX_GS blocking solution for 1 h at RT (PBDX_GS blocking buffer: 50 mL of 1x PBS; 0.5 g of bovine serum albumin – BSA; 0.5 mL of DMSO; 250 µL of 10% Triton; 750 µL of goat serum – GS (15 µL/1 mL)).
   7. Remove PBDX_GS and add ~40 µL of primary antibody dilution (generally 1:100).
      NOTE: The volume of the primary antibody dilution varies depending on the number of xenografts present in the microcentrifuge tube. Ensure that all the xenografts are submerged.
   8. Incubate for 1 hour at RT and then at 4 °C overnight. Position tubes vertically.
2. Day 2
   1. Remove primary antibody and wash 2x for 10 minutes in PBS/0.1% Triton.
   2. Wash 4x for 30 min in PBS/0.05% Tween.
      NOTE: The following steps must be performed in the dark (use aluminum foil to protect tubes from light).
   3. Remove PBS/0.05% Tween and add ~50-100 µL of secondary antibody dilution (generally 1:200 – 1:400) + DAPI (50 µg/mL) diluted in PBDX_GS.
   4. Incubate for 1 hour at RT and then at 4 °C overnight. Position tubes vertically and protect from light.
3. Day 3 (use aluminum foil to protect tubes from light)
   1. Remove secondary antibody dilution and wash 4x for 15 minutes in PBS/0.05% Tween.
   2. Fix at room temperature for 20 minutes in 4% FA.
   3. Wash 1x for 5 minutes in PBS/0.05% Tween.
   4. Remove PBS/0.05% Tween and add 1 drop of aqueous mounting medium to each microcentrifuge tube. Position the tubes vertically.
   5. Mount or store at 4 °C protected from light until mounting. Position the tubes vertically.

10. Mounting of xenografts

NOTE: Protect microcentrifuge tubes from the light throughout the process. Zebrafish xenografts are mounted between 2 coverslips (24 x 60 mm # 1.5). This allows to flip the mounted xenografts during confocal imaging so that both sides of the tumor (top and bottom) are accessible. Do not use plastic pipettes with mounting medium – the xenografts may get caught in the pipette. See Figure 7 for the schematic representation of the following steps.

1. Label the coverslip Y and seal the edges of the coverslip X with petroleum jelly or silicone grease to avoid the leakage of the mounting media.
2. Transfer the xenografts with a glass Pasteur pipette to the coverslip X.
3. Carefully align them with a hairpin and remove the excess aqueous mounting media.
4. Add aqueous mounting media to coverslip Y.
5. Carefully place coverslip Y on top of coverslip X. Do not press the coverslips as this can potentially disrupt the xenografts.
6. Place the assembled coverslips on top of a microscope slide and secure them with transparent adhesive tape. This allows an easier manipulation for confocal imaging and storage.

11. Confocal imaging

NOTE: An Apochromatic 25x immersion objective lens with water correction is optimal for imaging PVS tumors with single cell resolution (see Figure 8C-C" and Figure 9A for examples).

1. Acquire samples using the z-stack function with an interval of 5µm between each slice. For images aimed at 3D reconstruction, in particularly vessels, use an interval of 1-3µm between slices (Figure 8A).

12. Analysis and quantification

1. Use FIJI/ImageJ software or similar for confocal image processing and analysis.
2. Open the raw data (.czi, .lsm, etc) in FIJI software.
3. To select all or just a single channel in composite mode, click: Image > Colors > Channels Tool.
4. To adjust brightness and contrast levels, click: Image > Adjust > Color Balance.
5. To quantify tumor size
   1. Select three representative slices of the tumor, from the top, middle and bottom, per z-stack per xenograft (Figure 8 A).
      NOTE: Confocal resolution achieves ~60-70 µm of tumor depth. If the tumor is large, it may not be possible to image its total volume.
   2. Open a spreadsheet to annotate the data.
   3. Count every DAPI nuclei that corresponds to the tumor cells in the 3 selected slices (Figure 8 A, B). To do so:
      1. Open the cell counter plugin from FIJI/ImageJ by clicking Plugins > Analyze > Cell counter.
      2. In the cell counter tool, click Initialize, select a counter type, and click on the image to start the counting mode manually. For every click, the counter adds how many cells are counted (number of clicks).
      3. After counting one full slice, save the cell number in the corresponding excel document.
      4. Back to Fiji, click Reset from the Cell Counter window to delete the information if the same counter will be used. Otherwise, the information can be kept, and other counters can be used with other slices (or cells).
      5. Move to the second representative slice and repeat the previous steps. Gather all data.
         NOTE: DAPI counterstaining is commonly used to count cell numbers due to the clear definition of individual cells, however, other cell specific staining can be used.
   4. To obtain the total number of cells in the tumor use the following formula:
      
      NOTE: The 1.5 correction number was estimated for cells that have average nucleus diameter of ~10-12 µm. This correction may need adjustment if the cells are larger/smaller. See Dicussion for more details about this method.
6. To quantify other markers (immune cells, mitotic figures, PPH3, activated Caspase3, Ki67, etc.), quantify all slices using the same plugin (see Figure 8C-C'' for mitotic figures visualization). Divide the total number of counted cells by the corresponding tumor size and multiply by 100 to get the percentage.
   NOTE: Beware that some cells will be located between two slices, go back and forth in the z-stack to make sure one cell is not being counted twice.