Isolation of Primary Cancer-Associated Fibroblasts from a Syngeneic Murine Model of Breast Cancer for the Study of Targeted Nanoparticles

1. Establishing a syngeneic 4T1 model of breast cancer

NOTE: The present protocol describes the isolation of primary CAFs from a mouse 4T1 breast tumor. The animal study described here has been approved by the Italian Ministry of Health (aut. number 110/2018-PR).

1. Tumor cell culture and implantation
   1. Thaw 1 × 10⁶ 4T1-luc cells in a T75 flask with 10 mL of Roswell Park Memorial Institute (RPMI) 1640 medium supplemented with 10% fetal bovine serum (FBS), 1% penicillin/streptomycin (P/S), and 1% L-glutamine. Add Mycoplasma removal agent (1:100) in the culture medium.
      NOTE: 4T1-luc cells stably express luciferase and can be visualized by bioluminescence (BLI) upon proper stimulation with D-luciferin. This allows in vivo monitoring of the viability and proliferation of tumor cells upon implantation in mice (Figure 1).
   2. Maintain the cells at 37 °C and 5% CO₂ in a humidified atmosphere until ~80% confluence, by changing the medium every 2 days.
      NOTE: Continue treatment with Mycoplasma removal agent for 1 week (~2/3 passages) before injection in mice to ensure that injected cells are mycoplasma-free.
   3. On the day of the procedure, detach cells using 1 mL of trypsin-ethylenediaminetetraacetic acid (EDTA) solution (trypsin 1:250) for 5 min at 37 °C. Stop trypsin activity by adding culture medium, count the cells with trypan blue (1:1), and calculate the number of cells/mL.
   4. Pipette the volume corresponding to 1 × 10⁶ cells and centrifuge for 5 min at 400 × g. Pour off the supernatant, and resuspend the pellet in 1 mL of RPMI 1640 base medium. Keep the cells on ice until ready for injection.
      NOTE: For each mouse, 1 × 10⁵ cells are needed; however, always calculate for an excess of 2 mice (corresponding to 2 × 10⁵ cells) to ease syringe loading.
   5. One day before the operation, shave the fur of 8 BALB/c mice (female, seven-week-old) to expose the area of the abdominal mammary glands on the right side. Use an electric shaver and apply a thin layer of depilatory cream for 4 min; then, wash away the cream with water and a piece of paper.
   6. Induce anesthesia by continuous inhalation of 2% isoflurane gas for ~5 min, and perform a subcutaneous injection in the region of the abdominal mammary glands with a 27 G tuberculin syringe. Turn the needle upwards to enter the skin subcutaneously; then, hold the skin upwards and slowly inject 100 μL of cell suspension (1 × 10⁵ cells). Slightly turn the needle around before slowly retracting the syringe.
      NOTE: Remember to bring the cell suspension to room temperature 15 min before injection.
2. Tumor growth and dissection
   1. At 5 days post-cell-injection, inject 150 mg/kg D-luciferin intraperitoneally (100 μL) 5 min before imaging. Capture BLI images using an in vivo imaging system setting 10 s exposure, medium binning, and f/stop at 4. Define the luminescent area of the tumor as the region of interest (ROI), and quantify the total signal in the ROI (photon/sec/m²) using imaging software.
   2. Repeat the procedure of imaging (1.2.1) every 5 days from the injection to monitor tumor growth in terms of an increase in BLI.
   3. To establish the tumor volume, hold the mouse and expose its belly. Use calipers to measure the tumor length (L) and width (W) once a week. Calculate the tumor volume (V) using equation 1.
      ​[V = (L x W²)/2]     (1)
   4. At 20 days post-cell-injection, sacrifice the animals by cervical dislocation, dissociate the tumors from the skin with scissors, and collect them in a tissue storage solution (see the Table of Materials).
      NOTE: Tumor samples may be used immediately for dissociation into single cells (section 2) or stored at 4 °C for up to 48 h.

2. Tumor dissociation into single cells

NOTE: For the following steps, use sterile reagents and disposables in a laminar flow hood. Work with 4 tumors at a time.

1. Place the tumor sample in a Petri dish, and carefully remove any fragment of skin, fat, and necrotic areas with the help of tweezers and scalpel. Then, reduce the tumor into small pieces of approximately 1–2 mm and transfer them to a tube (Figure 2A).
   NOTE: 4T1 tumors become highly necrotic while growing, with a tendency to ulceration. It is important to carefully remove the necrotic areas to avoid any interference of debris with the next steps.
2. Prepare a digestion mix for tumor dissociation: mix 2.35 mL of RPMI 1640 medium, 100 μL of enzyme D, 50 μL of enzyme R, and 12.5 μL of enzyme A. Soak the tumor fragments in the solution and tightly close the tube.
   NOTE: The indicated volumes of digestion mix are valid for tumors up to 1 g and may be adjusted for larger tumors according to their weight. 4T1 tumors grown for 20 days normally are in a range of 0.5–0.8 g.
3. Turn the tube upside down, checking that all the pieces of tumor stand in the bottom of the tube toward the cap. Attach the tube to a mechanical dissociator in the proper housing, and run a specific dissociation program designed for tough tumors (see the manufacturer’s instructions).
   NOTE: Special tubes may be required to fit into the dissociator. C-tubes bear a rotor inside the cap to promote mechanical dissociation of the tissue.
4. Detach the tube, maintain it upside down, and incubate the sample at 37 °C for 40 min with gentle shaking. Then, attach the tube to the dissociator in the proper housing, and run the following specific dissociation program designed for tough tumors two times. Ensure there are no large pieces of tissue at the end of the procedure (Figure 2B).
   NOTE: To dissociate the tumor, other enzyme cocktails could be used to degrade the extracellular matrix. However, in this protocol, a commercially available Tumor Dissociation kit containing an optimized enzyme mix (enzymes D, R, A) and a semi-automated dissociator that mechanically dissociates the tissue were used. This combination ensured a proper degradation of the extracellular matrix together with maintenance of cell integrity and cell surface epitopes.
5. Filter the sample through a 40 μm cell strainer in a 50 mL tube, wash the filter with 10 mL of RPMI 1640 medium, and centrifuge the tube for 7 min at 300 × g. If the cell pellet appears red, lyse erythrocytes by adding 1 mL of Ammonium-Chloride-Potassium (ACK) lysing buffer for 5 min at room temperature, wash with 10 mL of RPMI 1640, transfer the cell suspension in a 15 mL tube, and centrifuge again.
6. Resuspend the pellet in 1 mL of PBE buffer composed of phosphate-buffered saline (PBS), 0.5% bovine serum albumin (BSA), and 2 mM EDTA, and count the cells with trypan blue (1:1). In case of cell clumps, filter the cell suspension through a 70 μm cell strainer, previously soaked with PBS.
   NOTE: Especially when tumors are big, it is important to check the cell count and eventually to split each tumor sample in different tubes with no more than 10⁷ cells in each tube before proceeding to step 2.7.
7. Prepare the dead cell removal binding buffer by diluting the 20x binding buffer stock solution (see the Table of Materials) with sterile double-distilled water. Wash cells with 5 mL of 1× binding buffer (Figure 2C).
   NOTE: Keep the buffer at 4 °C.
8. Centrifuge for 7 min at 300 × g, and resuspend the cell pellet with 0.1 mL of dead cell removal microbeads (see the Table of Materials). Mix well and incubate at room temperature for 15 min.
9. During the incubation with beads, prepare a magnetic stand under the hood and hang ferromagnetic separation columns (one column for up to 10⁷ cells) on it with the tips pointing down. Equilibrate the columns with 0.5 mL of cold 1× binding buffer. Wait until the solution has flowed through under gravity.
   NOTE: It is important to not exceed the number of cells per column to avoid any risk of clotting the column bed and reducing the recovery yield. When working with more than 10⁷ cells, either use more columns according to the total number of cells or use bigger columns. In such cases, adjust the reagent volumes described in the following steps to the ones indicated by the manufacturer.
10. At the end of incubation, add 400 μL of cold 1× binding buffer to the bead/cell suspension, load the entire volume onto the column, and collect the effluent (corresponding to unlabeled cells) in a 15 mL tube.
11. Wash the column four times with 0.5 mL of cold 1× binding buffer, and collect the total effluent (corresponding to the live cell fraction) in the same tube.
12. Count the cells with trypan blue (1:1). Place 1 × 10⁵ cells in two tubes for Fluorescence Activated Cell Sorting (FACS), and keep them at 4 °C until process validation (see step 4.1): use the first tube to set the analysis parameters and the regions of positivity (control tube) and the other one for biomarker analysis (sample tube). Use the remaining cells for CAFs extraction (see section 3).
    NOTE: Removal of dead cells is important to reduce non-specific reactions with the microbeads in section 3. However, to improve the yield, avoid the removal of dead cells if the proportion of dead cells is <20%.

3. Extraction of primary CAFs from breast tumor

NOTE: For section 3, use the mouse Tumor-Associated Fibroblast Isolation kit containing Non-Tumor-Associated Fibroblast Depletion Cocktail and Tumor-Associated Fibroblast Microbeads suited for magnetic labeling of the cells (see the Table of Materials).

1. Depletion of non-cancer-associated fibroblasts
   1. Moisten a 70 μm cell strainer with PBS, and filter the cell suspension to remove any clumps. Then, centrifuge the cells for 10 min at 300 × g and aspirate the supernatant.
      NOTE: If a single tumor was divided into two samples for step 2.7, pool the cell suspension two by two before proceeding with step 3.1.2.
   2. Resuspend the pellet in 80 μL of cold PBE buffer, and add 20 μL of Non-Tumor-Associated Fibroblast Depletion Cocktail. Mix well and incubate at 4 °C for 15 min in the dark.
   3. During the incubation with beads, prepare ferromagnetic depletion columns (1 per sample) on a magnetic stand, and equilibrate the columns with 2 mL of cold PBE buffer. Wait until the solution has flowed through.
   4. At the end of incubation, add 400 µL of cold PBE buffer to the bead/cell suspension, load the entire volume onto the column, and collect the effluent (corresponding to unlabeled cells) in a 15 mL tube.
   5. Wash the column twice with 2 mL of cold PBE buffer, and collect the total effluent in the same tube. Centrifuge for 10 min at 300 × g and resuspend in 0.1 mL of cold PBE buffer.
      NOTE: Depletion of non-cancer-associated fibroblasts drastically reduces the total amount of cells recovered. It is generally necessary and recommended to pool cell suspensions from four different samples to obtain a visible pellet and enough cells for step 3.2. This will ensure an optimal yield.
   6. Place 20 µL of the cell suspension in a FACS tube and keep it at 4 °C for process validation (see section 4). Use the remaining cells for CAF selection (see step 3.2).
2. Positive selection of cancer-associated fibroblasts
   1. Add 20 μL of Tumor-Associated Fibroblast Microbeads to 80 μL of cell suspension. Mix well and incubate at 4 °C for 15 min in the dark.
   2. During incubation with beads, prepare ferromagnetic separation columns (1 per sample) on a magnetic stand, and equilibrate the columns with 0.5 mL of cold PBE buffer. Wait until the solution has flowed through.
   3. At the end of the incubation, add 400 μL of cold PBE buffer to the bead/cell suspension, load the entire volume onto the column, and let the unlabeled cells flow through into a 15 mL tube.
   4. Wash the column three times with 0.5 mL of cold PBE buffer, and collect the total effluent in the same tube.
      NOTE: The flow-through contains unlabeled CD90.2-negative cells, which may be discarded as non-cancer-associated fibroblasts.
   5. Remove the column from the magnet and place it in a 1.5 mL tube. Add 1 mL of cold PBE onto the column, and immediately push the plunger into the column to flush the cells out.
   6. Centrifuge for 10 min at 400 × g and resuspend in 0.1 mL of cold PBE buffer. Place 20 μL of cell suspension in a FACS tube and keep it at 4 °C for process validation (see section 4).
      NOTE: After centrifuging, the pellet may be hardly visible, especially when the original tumors are small. Use conical bottom centrifuge tubes and be careful not to lose any cells when aspiring the supernatant.
   7. Dilute the remaining cells in an appropriate volume of Dulbecco’s Modified Eagle’s Medium (DMEM)/Ham’s F-12 supplemented with 15% FBS, 2 mM L-glutamine, 1% P/S, and 1% non-essential amino acids, and seed the cells in a tissue-culture plate.
   8. Check the cell density under a microscope, and place the plate in an incubator at 37 °C and 5% CO₂ to let the cells adhere and grow.
      ​NOTE: If the cells are too dense, immediately split the cell suspension into two wells of the tissue culture plate before placing the plate in the incubator.

4. Process validation

1. Flow cytometry
   NOTE: Section 4.1 does not require sterile conditions and can be performed outside the laminar hood. Perform section 4.1 in parallel for samples collected in step 2.11 (dissociated tumor), step 3.1.6 (after depletion of non-cancer-associated fibroblasts), and step 3.2.6 (after enrichment of cancer-associated fibroblasts).
   1. Centrifuge the FACS tubes prepared in steps 2.11, 3.1.6, and 3.2.6 for 10 min at 300 × g. Discard the supernatant.
      NOTE: Prepare at least one additional tube, which contains unstained cells acting as a control to set the parameters for analysis.
   2. In sample tubes, resuspend cells in 88 µL of PBE, and add 2 µL of anti-CD45 antibody conjugated with fluorescein isothiocyanate (FITC, 1:50 dilution, according to the manufacturer’s instructions) and 10 µL of anti-CD90.2 antibody conjugated with phycoerythrin (PE, 1:10 dilution, according to the manufacturer’s instructions). Mix thoroughly and incubate for 10 min at 4 °C in the dark. In the control tube (unstained), resuspend cells in 100 µL of PBE. Incubate for 10 min at 4 °C in the dark to replicate the procedure followed for antibody-stained cells.
      NOTE: Temperature and time of incubation must be checked carefully, as a variation in such variables might result in poor or non-specific staining, thus altering the results.
   3. After completing the incubation step, perform a wash by adding 1 mL of PBE. Centrifuge for 10 min at 300 x g. Discard the supernatant.
   4. Resuspend cells (all tubes) in 500 µL of PBS.
   5. Mix well and proceed with flow cytometry analysis. Set the channels to measure the fluorescence of the antibodies (i.e., FITC and PE).
   6. From the control tube, select all viable cells by drawing a first gate (P1) on the Forward versus Side scatter (FSC vs SSC) plot. Within P1, set a second gate (P2) comprising single cells only.
   7. Using the specific fluorescence channels of the antibodies (i.e., FITC and PE), set proper gates to discern positively stained cells.
   8. Start the analysis and record at least 10,000 events in P2. Read signals from both channels simultaneously.
      NOTE: If the overall number of cells is less than expected, it could be preferable to read the whole tube.
   9. (OPTIONAL) If further staining with additional antibodies/fluorophores is programmed, set an appropriate compensation matrix before initiating the analysis.
2. Monitoring of cell morphology and characteristics
   NOTE: Once seeded, the cells must be handled under sterile conditions.
   1. On the day after seeding, visualize the cells under an optical microscope to check cell adhesion and morphology. Aspirate the supernatant and replace with fresh medium.
      NOTE: CAFs are large spindle-shaped cells, which can be easily distinguished from the epithelial-like structure of 4T1 tumor cells.
   2. Maintain the cells at 37 °C and 5% CO₂ in a humidified atmosphere, changing the medium every 2 days.
   3. When cells reach ~80% confluence (approximately 4-6 days after seeding), detach the cells using TrypLE Select solution (see the Table of Materials) for 5 min at room temperature. Add fresh medium (4:1), centrifuge the cell suspension at 400 × g for 5 min, and resuspend the pellet in fresh medium.
   4. Split the cells 1:2, and expand the culture until the number of cells needed for the experiment is obtained.
   5. At each passage, check cell morphology and growth under an optical microscope. If cell morphology is not homogeneous (e.g., clones of small round-shape cells appearing in culture), analyze a sample of cells by flow cytometry as described in section 4.1. If homogeneous, proceed to step 4.2.7.
      NOTE: Changes in cell morphology may indicate contamination by non-cancer-associated fibroblasts, which need to be removed to ensure culture purity.
   6. Analyze the flow cytometry results and proceed according to one of the following scenarios: (i) if contamination of CD90.2-/CD45- cells occurs, collect all the cells, and repeat the positive selection by following step 3.2; (ii) if contamination of CD45+ cells also occurs, collect all the cells, and repeat the depletion in step 3.1; (iii) if no contamination occurs (only CD90.2+/CD45- cells are present), keep the cells in culture and proceed directly to 4.2.7.
      NOTE: As repeating the depletion process with microbeads will reduce cell recovery, it is recommended only when cells do not need to be used immediately.
   7. Count the cells with trypan blue (1:1), place 5 × 10⁵ cells in two FACS tubes, and centrifuge for 10 min at 300 × g. Discard the supernatant, and resuspend the pellet in 0.5 mL of blocking buffer (PBS supplemented with 2% BSA, 2% goat serum) for 15 min at room temperature.
      NOTE: One tube acts as a control consisting of unstained cells only, while cells in the second tube will be stained for flow cytometry analysis.
   8. Centrifuge for 10 min at 300 × g, discard the supernatant and resuspend in 99 µL of blocking buffer. Add 1 µg of anti-FAP antibody to the sample tube and 1 mL of blocking buffer to the control tube. Mix thoroughly and incubate for 15 min at room temperature.
      NOTE: FAP is a surface marker of reactive tumor stroma and may be used to identify and characterize CAFs.
   9. Wash thrice with PBS, centrifuge, and incubate with the appropriate secondary antibody conjugated with a fluorescent dye (1 μg) in blocking buffer for 15 min at room temperature.
   10. Wash thrice with PBS, and analyze by flow cytometry (follow steps 4.1.5 to 4.1.7 by setting the appropriate channel to detect the fluorophore used). Then, use the cells for experiments (step 5.2).
   11. ​(OPTIONAL) Freeze a sample of cells in 1 mL of 90% FBS and 10% dimethyl sulfoxide (DMSO); store at -80 °C for further use.

5. CAFs targeting by engineered Ferritin nanoparticles

NOTE: A recombinant variant of human ferritin heavy chain (HFn) was used as bare nanoparticle or was conjugated with targeting moieties. Here, HFn nanoparticles functionalized with the variable portion of an anti-FAP antibody (Fab@FAP) were prepared by the NanoBioLab at University of Milano-Bicocca at two HFn:Fab@FAP molar ratios, 1:1 and 1:5, according to a previously described protocol³².

1. Fluorescent labeling of HFn nanoparticles
   NOTE: Both bare and functionalized HFn nanocages are fluorescently labelled with FITC.
   1. Dissolve the FITC powder in 99% ethanol to obtain a concentration of 2 mg/mL.
      NOTE: This solution should be prepared freshly.
   2. Incubate 50 µL of the prepared solution (200 µg of FITC) for every 1 mg of protein (i.e., 100 µL of HFn at 10 mg/mL). Add 50 µL of 1 M sodium bicarbonate (NaHCO₃), and adjust the volume to 500 µL with 0.1 M NaHCO_3.
      NOTE: Scale up the volumes according to the experimental needs.
   3. Incubate the reaction mixture at room temperature, in the dark, and with continuous stirring for 1 h.
   4. Remove the excess of FITC from the conjugate by gel filtration using a spin desalting column (7 kDa MWCO).
   5. Assess the concentration of the recovered labelled protein by using a spectrophotometer. Estimate amounts of protein and dye by measuring the absorbance at 280 nm and at 488 nm, respectively.
2. Binding of HFn nanoparticles to CAFs
   NOTE: Use CAFs from no later than passages 4–5 in culture.
   1. Place 5 × 10⁵ cells in each FACS tube, centrifuge for 10 min at 300 × g, and resuspend in 0.5 mL of PBS, 0.3% BSA supplemented with 0.1 mg/mL of the different preparations of nanoparticles (bare HFn, HFn-Fab@FAP 1:1, HFn-Fab@FAP 1:5) previously labeled with FITC.
      ​NOTE: Run each condition in triplicate. Prepare an additional tube as a control unlabeled sample to which no nanoparticles are added.
   2. Incubate for 2 h at 4 °C in the dark, centrifuge, and wash three times in PBS.
   3. Resuspend cells with 0.5 mL PBS, mix well, and analyze by flow cytometry. Set the channels to measure the fluorescence of FITC.
   4. After gating on live single cells, acquire 20,000 events. Use the control unlabeled tube to set the gate of FITC positivity and obtain the percentage of FITC⁺ events (corresponding to nanoparticle binding).

6. Statistical analysis and experimental replicates

1. Animals
   1. Perform three independent experiments of tumor growth and CAFs isolation, using 8 animals per single experiment, as described in 1.1.5. To optimize CAFs isolation yield, divide the excised tissues into subgroups (n=4) and process them, following steps described in section 2.
2. HFn interaction with cells
   1. Evaluate the interaction of functionalized and non-functionalized HFn with target and non-target cells (CAFs and 4T1, respectively) in terms of the percentage of positively stained cells by fluorescently labelled nanocages. Report the results as average ± standard deviation of three independent experiments.
3. Statistical analysis
   1. To calculate statistical significance in cell binding experiments with HFn and HFn-FAP, use the ordinary one-way ANOVA test.