CAM-Delam Assay to Score Metastatic Properties by Quantifying Delamination and Invasion Capacity of Cancer Cells

In brief, Figure 1 summarizes the overall steps in the CAM-Delam assay. The below protocol is based on 30 cultured fertilized chicken eggs and the use of two different cancer cell lines seeded separately in three rings/egg and analyzed at four time points.

1. Egg incubation

1. Use fertilized chick eggs from a local hatchery, which guarantee over 90% fertilization
   NOTE: The eggs should not have been stored for more than 4 days at room temperature (RT). In Sweden, ethical permission for the experimental use of embryonic chickens is only required from embryonic day (E) 14 and beyond.
2. If needed, carefully mechanically wipe off any dirt or feathers on the eggs using a dry paper towel or wet in water.
3. Clean and sterilize the egg incubator with 70% ethanol before use (every time).
4. Place the desired number of eggs in egg trays and incubate the chick eggs horizontally in an egg incubator at 37.5-38 °C with 70% humidity. Consider this as incubation day 0 (Figure 1A).

2. Preparation of weighing boat, plastic boxes, and dissection instruments

1. Use 70% ethanol to sterilize 30 weighing boats and 30 small plastic boxes (~0.4 L) with transparent caps.
2. Dry the weighing boats and boxes in a laminar hood overnight (ON) and store in a closed plastic box until further use on incubation day 3.
3. Sterilize 2 L of distilled or deionized H₂O per 30 incubated eggs.
4. Sterilize two pairs of scissors and three pairs of forceps by spraying 70% ethanol. Air-dry the instruments and then store them in a sterilized box until incubation day 3.
   ​NOTE: These actions (Steps 2.1-2.4) can be done any day prior to incubation day 3. Use gloves to avoid contamination.

3. Opening the eggs and transfer to the internal humidified chamber

NOTE: Use gloves and a face mask to avoid contamination.

1. Add approximately 50 mL of sterilized H₂O to the sterilized plastic boxes. Keep the water-filled boxes with closed lids, to avoid contamination, at RT until use.
2. On incubation day 3, crack the eggshell using the sharp part of scissors and cut a straight opening in the shell.
3. Manually break open the eggshell over a weighing boat and collect the egg white, the yolk, and its attached healthy embryo in the weighing boat (Figure 1B). Look for an intact embryo with a beating heart, intact yolk, and developed blood vessels for the experiment (>95% of the incubated eggs). Discard eggs with a malformed embryo, an embryo without a beating heart, a broken egg yolk, or damaged yolk blood vessels.
4. Gently transfer the weighing boat into an internal humidified chamber.
   NOTE: For Steps 3.2.-3.4., work as fast as possible to avoid contamination, and if possible, use a laminar hood.
5. Incubate the internal humidified chamber in the egg incubator (Figure 1C).
6. Culture the shell-less eggs from day 3 to day 10 in the egg incubator before being used (see Step 3.6.), and when needed, add water to the incubator to maintain 70% humidity.
7. Check the internal humidified chambers through the transparent plastic lid for dead embryos or contaminated shell-less eggs every day, and remove them from the incubator.

4. Preparation of silicone rings

1. To prepare silicone rings, cut a silicone tube with an inner and outer diameter of 4 mm and 5 mm, respectively, in ~1 mm thickness, preferably using a paper cutter (Figure 1D).
2. Transfer the silicone rings to small glass bottles (Figure 1D), cover with metal foil, and sterilize them using an autoclave or similar.
   NOTE: Avoid repeated autoclaving of unused silicone rings that might be contaminated, since such treatment decreases the capacity of the silicone rings to attach to the membrane, with subsequent leakage of the cancer cell/collagen/RPMI solution outside of the rings.
3. Store the sterile silicone rings at RT.
   ​NOTE: This step can be done any day prior to incubation day 10.

5. Preparation of cancer cells

NOTE: Solutions such as cell culture medium, trypsin, and 1x PBS are stored at 4 °C and should be heated to 37 °C in a water bath before adding to the cells. After heating, rinse the bottles in 70% ethanol and dry before use.

1. Culture the cancer cell lines of interest in the relevant culture medium in a cell culture incubator at 37 °C in the presence of 5% (v/v) CO₂.
   NOTE: Here, U251-GFP glioblastoma and PC-3U-GFP prostate cancer cells were cultured in complete RPMI medium-RPMI medium supplemented with 10% (v/v) fetal bovine serum (FBS) and 1% (v/v) penicillin-streptomycin (PS).
2. Plan the culture of cancer cells so that approximately 50 × 10⁶ cancer cells are ready for harvesting on incubation day 10 of the CAM-Delam assay.
3. Change the cell culture medium every 2-3 days, or when the medium color changes from pink to orange/yellow.
4. Optional: Induce hypoxia by treating the cancer cells with CoCl₂ to investigate the molecular mechanistic effects on delamination 1 day before cell harvesting.
   CAUTION: CoCl₂ has moderate toxicity. Handle with care in a fume hood, and follow the manufacturer's instructions.
   1. Prepare fresh 20 mM CoCl₂ stock solution by dissolving 0.0258 g in 10 mL of sterilized distilled H₂0 in a 15 mL conical tube wrapped with aluminum foil to protect from light.
   2. In a 50 mL conical tube, mix 250 µL of the 20 mM CoCl₂ stock solution in 25 mL of RPMI medium supplemented with 1% (v/v) PS (but without FBS) per 15 cm diameter cell culture dish. Vortex gently.
   3. Remove the complete RPMI medium from the cell culture dishes.
   4. Wash the cells with sterilized 1x PBS 2x.
   5. Add 25 mL of RPMI medium with 200 µM CoCl₂ cell trypsinization (see Step 5.6).
5. On egg incubation day 10, prepare 1 mL of a collagen/RPMI mix (ratio 1:3), containing 250 µL of type I collagen (5 mg/mL) and 750 µL of cell culture RPMI medium supplemented with 10% FBS and 1% (v/v) PS. Keep the collagen/RPMI-mix on ice.
6. Trypsinize the cancer cells to isolate the cells by first removing the cell culture medium and wash 2x with 1x PBS.
7. Add 3 mL of trypsin solution (0.05%) per 15 cm diameter cell culture dish and incubate for 2-3 min in a cell culture incubator until the cells detach. Use a table-top, inverted microscope to see the detached cells. If needed, gently tap the side of the flask to dislodge remaining clustered or attached cells.
8. Inactivate trypsin by adding 5 mL of complete RPMI medium to each cell culture dish and collect the cell suspension from all cell culture dishes into a 50 mL tube.
9. Count the cancer cells by the Trypan Blue exclusion method to distinguish live from dead cells using a cell counter.Add 10 µL of the cell suspension to 10 µL of 0.4% trypan blue stain. Mix the sample by pipetting up and down a few times, and then load 10 µL of the cell mixture per chamber into the sample slide in the cell counter. Repeat cell counting 3x to verify the cell number/mL.
10. Calculate and centrifuge the correct volume cell culture suspension containing 50 x 10⁶ live cancer cells in a 50 mL tube at 500 × g for 5 min.
11. Discard the supernatant and mix the cell pellet with 1 mL of the collagen/RPMI mix. As collagen is a gelatinous material, mix the cells very slowly and carefully with a 1 mL pipette to avoid losing cells in the formation of bubbles.
12. Keep the prepared cell suspension on ice and bring it to the egg working area.
    ​NOTE: Avoid keeping the prepared cancer cells on ice for too long. The prepared cancer cells must be placed on the CAM within 15-25 min.

6. Seeding the cancer cells on the CAM

NOTE: Use gloves and a face mask to avoid contamination.

1. At incubation day 10, take out the internal humidified chambers with the incubated shell-less eggs from the incubator.
   NOTE: Approximately 90% of the initial number of the incubated eggs can be used; see Figure 2.
2. Open the internal humidified chamber and place up to six silicone rings on the CAM using sterilized forceps.
   NOTE: Avoid placing the silicone rings near each other or close to bigger blood vessels.
3. Mix the cancer cell suspension by pipetting to get an even distribution of cancer cells, and add 20 µL (1 × 10⁶ cells) of the prepared cancer cell suspension inside a silicone ring (Figure 1E). When adding the cells, keep the pipette tip above the CAM to make sure not to damage the membrane.
   NOTE: According to previous findings⁷, different cancer cells can be seeded in separate rings on the same CAM.
4. Close the internal humidified chamber and put it in the egg incubator.

7. Isolation of the CAM with associated cancer cells

1. After 14 h, 1.5 days, 2.5 days, and 3.5 days of incubation, take out approximately seven internal humidified chambers and open the lids one at a time.
2. With a pair of scissors, dissect the cultured cancer cells attached to the CAM (so-called CAM-Delam samples) by cutting outside the silicone ring (Figure 1F).
3. Immediately transfer the isolated CAM-Delam sample using forceps to 4% paraformaldehyde (PFA) in 0.1 M phosphate buffer (PB) in a Petri dish for fixation of the tissue. Keep on ice or at 4 °C for 1 h.
   NOTE: Store 4% PFA solution at 4 °C for a maximum of 5 days. CAUTION: PFA is toxic. When handling PFA powder and PFA solutions, use a fume hood and wear a face mask and gloves. Avoid inhaling powder or solution vapors. Follow the manufacturer's instructions.
4. Decapitate the chicken embryos and discard as biological waste in specific bins.
5. Remove the 4% PFA solution and add 30% sucrose in 0.1 M PB to the CAM-Delam samples and equilibrate at 4 °C for 1 h.
6. Under a dissection microscope, carefully remove the silicone ring using forceps. With a pair of scissors, cut the CAM-Delam sample in a rectangular shape with the cancer cells in the middle (Figure 1G).
7. With a pair of forceps, transfer the CAM-Delam samples to frozen section medium in a Petri dish to remove excess sucrose and then to embedding molds in frozen section medium.
8. Under a dissection microscope, position the CAM-Delam sample in a U-shape in a vertical direction in the embedding molds using any needle-like instrument (Figure 1G).
9. Freeze and store the CAM-Delam samples at −80 °C.

8. Sectioning CAM-Delam samples

1. Section the frozen CAM-Delam samples at 10 µm on 5-6 consecutive slides using cryosectioning.
2. Store the slides with sections at −80 °C or use directly for immunohistochemistry staining.

9. Immunohistochemistry staining

1. Bring the slides from −80 °C to RT for 5 min before starting the immunoprotocol.
2. Make a line with a hydrophobic marker on the slides where the sections end. Let them dry for a few minutes.
3. Place the slides in a humidified (H₂O) chamber and cover the sections with ~200-500 µL of blocking solution (10% fetal calf serum and 0.1% sodium azide in Tris-buffered saline with 0.1% Triton X-100 [TBST]) and incubate for 15-30 min.
   NOTE: From this step onwards, do not let the slides dry at any time.
4. Pour off the blocking solution and replace it with 100-150 µL of the primary antibody of interest diluted in the blocking solution and incubate ON at 4 °C. Preferably use rabbit anti-laminin-111 antibody to define the basal lamina together with a cancer cell marker, if not using GFP or other fluorescent-tagged cancer cell lines.
   NOTE: Here, a rabbit anti-von Willebrand Factor was also used to detect blood vessels in the CAM.
5. Prepare three glass cuvettes filled with TBST buffer.
6. Pour off the primary antibody solution, transfer the slides to the glass cuvettes, and wash at least 3x for 5 min each in TBST.
7. Remove excess TBST from the slide and from the hydrophobic barrier area with a soft paper tissue.
8. Cover the slide with 100-150 µL of a suitable secondary fluorescent antibody diluted in blocking solution combined with 4',6-diamidino-2-phenylindole, dihydrochloride (DAPI) and incubate in the dark at RT for 1 h.
9. Pour off the secondary antibody solution, transfer the slides to the glass cuvettes, and wash at least 3x for 5 min each in TBST.
10. Remove excess TBST from the slide and from the hydrophobic barrier area with a soft paper tissue.
11. Mount the slides by putting 1-2 drops of fluorescent mounting medium on the slide, and gently place a glass coverslip, avoiding air bubbles.
12. Let the slides dry for at least 1 h at 4 °C before analyzing, and store the slides 4 °C.

10. Microscopy imaging and delamination scoring

1. Photograph the sections using an epifluorescence microscope equipped with a digital camera, preferably at 10x magnification (Figure 1H).
2. Analyze the sections using the following CAM-Delam scoring categories (Figure 3):
   1. Look for intact basal lamina without visible alterations.
   2. Look for altered but undamaged basal lamina.
   3. Look for damaged basal lamina without cell invasion.
   4. Look for damaged basal lamina with cell invasion.