A Patient-Derived Xenograft Model for Venous Malformation

Patient tissue samples were obtained from participants after informed consent from the Collection and Repository of Tissue Samples and Data from Patients with Tumors and Vascular Anomalies under an approved Institutional Review Board (IRB) per institutional policies at Cincinnati Children’s Hospital Medical Center (CCHMC), Cancer and Blood Disease Institute and with approval of the Committee on Clinical Investigation. All animal procedures described below have been reviewed and approved by the CCHMC Institutional Animal Care and Use Committee.

1. Preparation of materials and stock solutions

1. Preparation of complete endothelial cell growth medium (EGM)
   1. Supplement endothelial basal medium (EBM) with the following growth factors present in the kit (see Table of Materials): human Fibroblast Growth Factor-Beta (hFGF-β), vascular endothelial growth factor (VEGF), Long Arg3 Insulin-Like Growth Factor- I (R³-IGF-I), ascorbic acid, epidermal growth factor (EGF), gentamycin sulphate and amphotericin-B, and heparin. Add 1% Penicillin/Streptomycin/L-Glutamine (PSG) solution and 20% fetal bovine serum (FBS).
      NOTE: We do not recommend adding hydrocortisone.
   2. Sterile filter the solution under a laminar flow hood through a 0.2 µm bottle top filter into an autoclaved glass bottle and aliquot media into 50 mL conical tubes and store at 4 °C up to a week or at -20 °C for up to one year.
2. Prepare collagenase A stock solution
   1. Prepare 50 mg/mL collagenase A stock solution in 1x phosphate buffer saline (PBS).
   2. Sterile filter the solution under a laminar flow hood with 0.2 µm filter and syringe, and store 100 µL aliquots at -20 °C.
3. Prepare tissue collection medium (Buffer A)
   1. Prepare a Ca²⁺/Mg²⁺ stock solution by adding 0.927 g of calcium chloride dihydrate (CaCl₂.2H₂O) and 1 g of magnesium sulfate heptahydrate (MgSO₄.7H₂O) to 500 mL of distilled water. Sterile filter the solution through a 0.2 µm bottle top filter into an autoclaved glass bottle and store at room temperature (RT).
   2. Supplement Dulbecco's Modified Eagle Medium (DMEM) with 10% Ca²⁺/Mg²⁺ solution and 2% FBS.
   3. Sterile filter the solution under a laminar flow hood with 0.2 µm filter and syringe and store aliquots of 5 mL at -20 °C.
4. Fibronectin coating of tissue culture plates
   1. Prepare coating buffer (Buffer B) by dissolving 5.3 g of sodium carbonate (Na₂CO₃; 0.1 M) in 500 mL of deionized water. Adjust the pH of the buffer to 9.4 using 1 M hydrochloric acid (HCl). Filter under a laminar flow hood through a 0.2 µm bottle top filter into an autoclaved glass bottle and store at RT.
   2. For coating, pipette 2 mL/5 mL/10 mL of Buffer B per 60 mm/100 mm/145 mm tissue culture plate, respectively. Add 1 µg/cm² of human plasma fibronectin purified protein solution and gently distribute the liquid onto the plate.
   3. Incubate plate at 37 °C, 5% CO₂ for 20 min.
   4. Aspirate Buffer B and wash the plate with PBS prior to culturing cells.

2. Isolation of endothelial cells from VM patient tissue

1. Isolation of EC from solid VM tissue
   NOTE: VM tissue is resected by debulking surgery^12,13 under an IRB approved protocol.
   1. Wash VM tissue (tissue sample weight typically ranges between 0.5 g and 1.5 g) in 5% PSG in PBS.
   2. Transfer tissue to a 100 mm cell culture dish, mince tissue sample into small pieces using sterile surgical dissection tools, and transfer into a 50 mL conical tube.
   3. Add 100 µL of collagenase A stock solution to 5 mL of Buffer A for a final concentration of 1 mg/mL, then add this to the tissue and digest the minced tissue at 37 °C for 30 min while shaking contents every 5 min.
   4. Carefully grind the digested tissue at RT using a 6 mm, smooth-surface pestle grinder within the 50 mL conical tube.
   5. Continue to carefully grind digested tissue using a pestle while adding 5 mL of cold PBS supplemented with 0.5% bovine serum albumin (BSA) and 2% PSG. Repeat this step four times.
   6. Filter the solution through a 100 μm cell strainer into a 50 mL conical tube to remove tissue fragments.
   7. Centrifuge cell suspension for 5 min at 400 x g at RT. Proceed to step 2.3.
2. Isolation of VM EC from lesional blood obtained from patient sclerotherapy
   1. Obtain human VM lesional blood from sclerotherapy under an IRB approved protocol.
   2. Dilute lesional blood (sample volume typically ranges between 0.5 mL and 5 mL) in PBS to a final volume of 40 mL.
   3. Centrifuge cell suspension for 5 min at 200 x g at RT.
3. Initial cell plating
   1. Discard supernatant and resuspend the single-cell pellet in 1 mL of EGM.
   2. Add 9 mL of complete EGM onto fibronectin-coated (1 µg/cm²) 100 mm plates and seed 1 mL of cell suspension.
   3. Incubate cells at 37 °C in a humidified 5% CO₂ atmosphere.
   4. Every other day remove 2 mL of media and add 2 mL of sterile filtered FBS.
   5. Once cell cultures reach a confluency of 40‒50% change the medium to complete EGM. It typically takes between 2‒3 weeks for the cells to reach this confluency.
   6. Observe daily for the appearance of EC colonies. They can be recognized by their typical “cobblestone-like” morphology (Figure 1A). Between 3‒5 EC-colonies appear in each sample.
   7. Change the medium every other day for another 5‒7 days until individual EC colonies start to touch one another.
4. Manual isolation of individual EC colonies
   1. To harvest EC colonies, wash the plate with 5 mL of PBS and manually aspirate with a serological pipette.
   2. Take the plate to a microscope and circle the locations of multiple EC colonies using a marking pen both on lid and bottom before returning it to laminar flow hood.
   3. Detach EC colonies by pipetting 50 µL of 0.05% trypsin-EDTA solution on the marked areas.
   4. Using a small cell scraper or a pipette tip, gently scrape cells from plate.
   5. Tilt the plate to the nearest edge, and rinse with 1 mL of EGM per marked area and collect cells.
   6. Count the number of cells using a hemocytometer or automated cell counter.
   7. Plate the collected EC colonies at a density of 1 x 10⁴ cells/cm² onto fibronectin-coated (1 µg/cm²) cell culture dishes containing fresh EGM. The next day, change medium to complete EGM.
   8. Change the medium to complete EGM every other day for 2‒3 weeks until cells reach 80% confluency.
   9. Trypsinize EC with 2 mL of pre-warmed 0.05% trypsin-EDTA per 100 mm dish at 37 °C for 2 min and neutralize trypsin by adding 4 mL of EGM.
   10. Collect the cell suspension into one 15 mL conical tube and centrifuge at 400 x g at RT for 5 min. Aspirate the supernatant and resuspend cells in 2 mL of EGM.
   11. Count the number of cells using a hemocytometer or automated cell counter. Typical cell numbers obtained are 1 x 10⁶ cells per 60 mm cell culture plate or 2 x 10⁶ cells per 100 mm tissue culture plate.
   12. Pellet the cells by centrifugation at 400 x g at RT for 5 min and aspirate the supernatant. Proceed to step 3.2.1.

3. Endothelial cell selection and expansion

1. Preparation of anti-CD31-conjugated magnetic beads
   1. Vortex the vial containing the anti-CD31-conjugated magnetic beads for 30 s. The number of beads required is 8 x 10⁶ beads/2 x 10⁶ cells.
   2. Wash the desired amount of anti-CD31-conjugated magnetic beads with 1 mL of wash solution containing 0.1% BSA in PBS in a 1.5 mL microcentrifuge tube.
   3. Place the microcentrifuge tube in a cell isolation magnet for 1 min.
   4. Aspirate the supernatant and repeat the washing step.
2. Endothelial cell purification and plating
   1. Resuspend cell pellet obtained in 2.4.12 in 500 µL of wash solution containing 0.1% BSA in PBS.
   2. Add cell solution to microcentrifuge tube containing magnetic beads and resuspend thoroughly.
   3. Incubate for 20 min at 4 °C with gentle tilting.
   4. Add 500 µL of 0.1% BSA in PBS and mix well.
   5. Place the tube on a cell isolation magnet for 1 min.
   6. Gently aspirate all of the liquid, which contains the CD31-negative fraction of cells without touching the beads.
   7. Wash the bead pellet containing the CD31-positive cell fraction with 1 mL of 0.1% BSA in PBS and repeat magnetic separation.
   8. Repeat washing and magnetic separation step for a minimum of 3 times to purify CD31-positive cell fraction.
   9. Resuspend purified endothelial cells into a 15 mL conical tube and spin down for 5 min at 400 x g at RT.
   10. Remove supernatant and resuspend cell pellet in 1 mL of EGM.
   11. Add 9 mL of complete EGM onto fibronectin-coated (1 µg/cm²) 100 mm plates and seed 1 mL of cell suspension. Note that some magnetic beads are still attached to the cells in this initial seeding step. Most beads wash away during cell expansion, but small number of beads may persist in early passages.
   12. Incubate cells at 37 °C in a humidified 5% CO₂ atmosphere.
   13. Change the medium every other day until cells reach 80 % confluency (Figure 1B).
3. Endothelial cell expansion
   1. Once cells reach 80 % confluency, detach with 2 mL of 0.05% trypsin-EDTA per 100 mm dish at 37 °C for 2 min.
   2. Neutralize trypsin by adding 4 mL of EGM and collect cells into one 15 mL conical tube.
   3. Centrifuge at 400 x g at RT for 5 min.
   4. Aspirate the supernatant, resuspend cells in 2 mL of EGM, and count the number of cells with a hemocytometer or by automated cell counting.
   5. Seed cells at a density of 1 x 10⁴ cells/cm² into 145 mm fibronectin-coated (1 µg/cm²) tissue culture plates.
   6. Continue to passage cells until the desired cell number has been met. Consider that a confluent 145 mm tissue culture plate contains about 8‒9 x 10⁶ cells and the number of cells needed is 2.5 x 10⁶ cells per injection. Only cells between passage 3 and 8 should be considered for the xenograft.

4. VM patient-derived xenograft protocol

NOTE: In this protocol we use 5‒6 week old, male immunodeficient, athymic nude Foxn1^nu mice.

All animal procedures must be approved by the Institutional Animal Care and Use Committee (IACUC).

1. Preparation of materials on the day before injection
   1. Pre-chill syringes, needles, and pipet tips in -20 °C freezer overnight.
   2. Slowly thaw the basement membrane extracellular matrix (BMEM) overnight on ice bucket placed at 4 °C to avoid increased viscosity of the gel.
2. Preparation of cell suspension for injection
   1. Trypsinize EC with 5 mL of pre-warmed 0.05% trypsin-EDTA per 145 mm dish at 37 °C for 2 min.
   2. Neutralize trypsin by adding 5 mL of EGM and collect cells into one 15 mL conical tube.
   3. Centrifuge at 400 x g at RT for 5 min.
   4. Aspirate the supernatant, resuspend cells in 3 mL of EGM, and count the number of cells using a hemocytometer or automated cell counter.
   5. Determine the total number of cells that are needed for all planned injections.
      NOTE: The recommended number of cells is 2.5 x 10⁶ cells per injection. A total of 2 injections are typically performed in each mouse for technical duplicates. It is necessary to calculate a 10% excess of cell number to account for loss during transfer into the syringe.
   6. Transfer the volume containing the calculated cell number into a new 50 mL conical tube and pellet cells by centrifugation at 400 x g at RT for 5 min.
   7. Aspirate the supernatant, leaving a small volume (about 50‒70 µL) to loosen the pellet.
3. Syringe preparation
   1. Calculate an excess volume of 20 µL/injection to account for loss during transfer to the syringe. Resuspend the cell pellet with 220 µL of BMEM per injection on ice. The injected volume of cell suspension will be 200 μL per lesion.
   2. Mix the cell suspension thoroughly on ice to obtain a homogenous cell suspension and avoid creating bubbles.
   3. Using a 1 mL pipet and 1 mL syringe, simultaneously pipet BMEM-cell mixture into the syringe opening by suction force while pulling plunger of syringe.
   4. Luer lock a 26G x 5/8 inch sterile needle to the syringe and keep prepared syringes flat on ice prior to injection.
4. Subcutaneous injection into mouse
   1. Anesthetize the mice with 5% isoflurane/oxygen mixture at a flow rate of 1 L/min using an isoflurane vaporizer. Ensure proper sedation of animals (e.g., unresponsiveness to toe pinches). Maintain anesthesia via continuous administration of 1.5% isoflurane/oxygen delivered via nose cone.
   2. Place mice on their stomach, exposing the back region where grafting will occur and disinfect the injection region with 70% ethanol.
   3. Gently roll the prepared syringe to resuspend any settled cells. Flick bubbles to the needle end of the syringe and expel a small volume of the cell suspension to ensure the removal of all bubbles.
      NOTE: Two injections can be performed for each mouse – on the left and the right side of mouse back.
   4. Pinch and create a ‘tent-like’ structure using your thumb and index finger and insert the needle subcutaneously right under the skin. Ensure that the needle is only skin deep by releasing pinched skin to prevent injection into muscle tissue.
   5. Holding needle at 45° angle carefully inject 200 µL of the cell-suspension to create a small spherical mass (Figure 1C).
   6. Record mouse weight with a scale, ear tag the mouse, and return to cage.
   7. Monitor mice following sedation to ensure they return to normal activity.
5. Lesion growth monitoring
   1. Using a caliper, measure the length and width of each plug (Figure 1D).
   2. Document measurements every other day up until lesion collection.

5. Tissue collection and processing

1. Euthanize mice 9 days post-implantation in a CO₂ chamber and check vital signs to confirm death. Perform cervical dislocation on the mice as a secondary method to euthanize the mouse.
2. Harvest the xenograft lesion/plug from the flank of the mouse by dissection using surgical forceps and scissors.
   NOTE: In order to prevent rupturing the blood-filled vessels within the plug, it is important to avoid touching the lesion plug with dissection tools and leave excessive surrounding tissue such as skin attached to the plug.
3. Immerse the resected lesion in PBS to wash.
4. Set up a camera stage with a camera. Align plugs onto a cutting board with a ruler. Take an image of all plugs to record gross vascularity of lesions (Figure 1E).
5. Fix plugs by submerging in 10% formalin overnight at RT.
6. Wash plugs in PBS the following day and move them into 70% ethanol.
7. Process lesion plugs for paraffin embedding (pathology core).

6. Lesion sectioning

1. Use a microtome to cut 5 μm sections from the collected murine lesions onto positively charged slides.
   NOTE: For the subsequent analysis, sections in the center of the plug (about 50‒70 µm into the tissue) are of importance.
2. Melt paraffin at 60 °C for 1 h prior to staining.
3. De-paraffinize and re-hydrate tissue sequentially under a chemical fume flow hood. Therefore, incubate slide in xylene for 10 min, 100% ethanol (EtOH) for 5 min, 90% EtOH for 3 min, and 80% EtOH for 3 min.
4. Rinse slide in deionized water for 5 min.

7. Hematoxylin and Eosin (H&E)

1. Incubate sections in Hematoxylin for 2 min.
2. Place slides in a staining jar and rinse in a sink by a steady stream of tap water until water is clear.
3. De-hydrate slides by incubating tissue sequentially in 70% EtOH for 1 min, 80% EtOH for 1 min, 90% EtOH for 1 min, 100% EtOH for 1 min, and fresh 100% EtOH for 1 min.
4. Stain sections in Eosin Y for 30 s.
5. Rinse in fresh 100% EtOH until solution is clear.
6. Incubate slide in xylenes for 2 min. Let slides dry for 5‒10 min under the fume hood.
7. Dispense a drop of permanent, non-aqueous mounting medium over xenografts sections and place coverslip on top.
8. Allow slides to dry overnight before imaging.

8. Immunohistochemistry

1. Prepare antigen retrieval buffer (Tris-EDTA) by weighing 0.6 g of Tris-base and 1 mL of 0.5 M EDTA to 500 mL of deionized water. Adjust pH to 9.0 using 1 M HCl. Add 250 µL of Tween-20.
2. Incubate de-paraffinized tissue slides (as obtained in step 6.2‒6.3) in a beaker with antigen retrieval buffer, stirring on a heating block, for 20 min at 95 °C.
3. Remove the beaker from heating block, allow solution to cool to 35 °C then wash in PBS for 3 min.
4. Block tissue sections in 5% normal horse serum in PBS for 30 min at RT.
5. Prepare a biotinylated Ulex europaeus agglutinin-I (UEA-I) working solution by diluting 20 µg/mL of biotinylated UEA-I in 5% normal horse serum in PBS.
6. Pipet 50‒100 µL of UEA-I working solution per section and incubate for 1 h at RT in a humidifying chamber.
7. Wash slides two times in PBS for 3 min.
8. Quench slide sections in 3% hydrogen peroxide for 5 min at RT.
9. Wash slides two times in PBS for 3 min.
10. Prepare 5 µg/mL of Streptavidin horseradish peroxidase-conjugated in 5% normal horse serum in PBS.
11. Pipet 50‒100 µL on each tissue slide and incubate for 1 h at RT in a humidifying chamber.
12. Wash slides two times in PBS for 3 min.
13. Prepare 3,3'Diaminobenzidine (DAB) solution according to manufacturer’s instructions and add 50‒100 µL per section.
14. Incubate sections for 10‒15 min, checking and monitoring for development of stain every 2‒5 min.
15. Wash slides three times in PBS for 3 min.
16. Add a drop of Hematoxylin and incubate for 3 min.
17. Place slides in a staining jar and rinse in a sink by a steady stream of tap water until water is clear.
18. Sequentially incubate slide in 80% EtOH for 1 min, 90% EtOH for 1 min, 100% EtOH for 1 min, and xylene for 2 min.
19. Let slides dry for 5‒10 min under the fume hood.
20. Dispense a drop of permanent, non-aqueous mounting medium over xenografts sections and place coverslip on top.
21. Allow slides to dry overnight before imaging.

9. Analysis of human-derived Vascular Channels

NOTE: Vascularity of VM lesions is quantified by measuring vascular area and vascular density. Only UEA-I positive, human-derived vascular channels are considered for quantification.

1. Take four to five images per lesion section with a bright field microscope at a 20x magnification (high power fields [HPF]). Take HPF images in an x-plane pattern within the lesion section to avoid overlap (Figure 1F-H). Include a scale bar on the images taken.
2. Open the HPF images in Image J (File > Open). Calibrate the pixels of the scale bar as follows. Use the straight line tool and go over the scale bar. To convert the measured pixels into mm click on Analyze > Set scale.
3. Click on Analyze > Set Measurements and select Area and Add to overlay.
4. Measure the total field area in a HPF using Analyze > Measure. Save this measurement for quantification in step 9.8.
5. Using the freehand selections tool, manually outline UEA-I⁺ vascular channels.
   NOTE: A vascular channel is defined as any area that is lined with UEA-I⁺ – EC that may contain blood cells.
6. Click on Analyze > Measure to quantify the outlined UEA-I⁺ vascular area (mm²/HPF).
7. Repeat this measurement for all five HPF taken within one plug.
8. Average the total vascular area of all five HPF. The obtained vascular area per HPF is subsequently divided by the HPF field area (in mm², step 9.5) and expressed as a percent (%).
9. For quantification of vascular density, count the number of UEA-I⁺ vascular channels of each HPF taken. The vascular density is the average number of UEA-I⁺ vascular channels counted per HPF area (vessels/mm²).