Fabrication of Engineered Vascular Flaps Using 3D Printing Technologies

All animal procedures were approved and conducted under the supervision of the Technion Pre-Clinical Research Authority (PCRA Technion, ethics approval no. 058-05-20). Male Sprague-Dawley rats (275-350 g) were used for these studies. See the Table of Materials for details related to all materials, equipment, and software used in this protocol.

1. Vascular scaffold fabrication

1. Create a 3D computer model of the desired vascular scaffold shape using computer-aided design (CAD) software or download a 3D file of a desired vascular structure from online repositories.
   1. Create a cylinder with an inner diameter of 0.9 mm, an outer diameter of 2 mm, and a height of 18 mm. Add radial fenestrations with a diameter of 0.22 mm alongside the cylinder wall.
2. Create a mold for the scaffold using the 3D design software and export it as an .STL file. Next, add a funnel to the design to ease the filling of the mold later.
3. Import the .STL file into the slicing software for the fused deposition modeling (FDM) 3D printer. Slice the model with a layer height of 0.1 mm, and export it as .gcode or send it directly to the 3D printer.
4. 3D print the mold on an FDM 3D printer equipped with a 0.25 mm nozzle, using water-soluble butene-diol vinyl alcohol copolymer (BVOH) filament. Store the printed molds in a vacuum chamber until use.
   CAUTION: Avoid handling the BVOH filament with bare hands as it is moisture-sensitive. In addition, it is recommended to store the filament in a vacuum chamber to avoid exposure to moisture.
5. Prepare a polymer solution of a 1:1 mixture of poly-L-lactic-acid (PLLA) and polylactic-co-glycolic-acid (PLGA) in 1,4-dioxane.
   1. Weigh 350 mg of PLLA and 350 mg of PLGA and transfer to a 20 mL glass vial.
   2. Measure 10 mL of 1,4-dioxane and transfer to the glass vial using a glass pipette. Add a small magnetic stir bar to the glass vial.
      CAUTION: 1,4-Dioxane is a hazardous organic solvent. Use appropriate personal safety equipment and work inside a fume hood.
   3. Place the glass vial inside a hot water bath heated to 70 °C and mix the contents overnight until all the polymers dissolve fully. Store the solution in an airtight glass container until use.
6. Fill the BVOH molds with the PLLA:PLGA solution.
   1. Using a positive displacement pipette, measure 30 µL of the polymer solution and fill in the mold.
      NOTE: The volume needed to fill the mold will change according to the volume of the designed vessel. Keep adding the polymer solution until the funnel of the mold is entirely filled.
   2. Centrifuge the mold at 100 × g for 2 min. Top up the mold with a further 20 µL of the polymer solution to ensure complete filling.
7. Freeze the filled molds at -80 °C for at least 30 min to ensure all the polymer solution freezes. Remove the solvent from the molds by lyophilizing them overnight.
   NOTE: The color of the polymer in the mold should turn from clear to white after the freeze-drying process.
8. To remove the sacrificial mold material, transfer the molds after the freeze-drying process to a 5 L deionized water bath under gentle stirring. Replace the water in the bath when it gets cloudy. When all the BVOH dissolves, air-dry the scaffolds and store them in a vacuum chamber until use.

2. Coating the vascular scaffold with fibronectin

1. Disinfect the PLLA:PLGA vascular scaffolds by immersing them in 70% ethanol for 30 min.
2. Wash the scaffolds 3x 5 min with PBS.
3. Prepare a dilution of 50 µg/mL human fibronectin in PBS and coat the scaffolds.
   1. Mix 500 µL of stock fibronectin solution (1 mg/mL) with 9.5 mL of PBS.
   2. Immerse the disinfected vascular scaffolds in this fibronectin solution and incubate them at 37 °C for 60 min to allow for protein adsorption.
4. After incubation, rinse the scaffolds with PBS to remove the unbound fibronectin. Store these coated scaffolds at 4 °C for up to 1 week.

3. rhCollMA bioink preparation

1. Prepare a phosphate buffer for neutralizing the acidic rhCollMA bioink stock.
   1. Prepare a 10x stock of phosphate buffer by dissolving 5.495 g of Na₂HPO₄, 1.55 g of NaH₂PO₄, and 30 mg of NaCl in 50 mL of deionized water.
   2. Prepare a 10-fold dilution of the 10x stock phosphate buffer by combining 1 part of 10x phosphate buffer with 9 parts of deionized water.
2. Combine 900 µL of stock rhCollMA solution with 100 µL of 10x phosphate buffer to neutralize the acidic rhCollMA solution.
3. Dilute the neutralized rhCollMA solution to the desired concentration of 10 mg/mL by mixing it with the required amount of 1x phosphate buffer.
   NOTE: The stock concentration of rhCollMA varies between lots. Therefore, the volume of 1x phosphate buffer needed to reach the desired concentration will vary.
4. Prepare the porogen-photoinitiator solution (PEO-LAP) to be combined with the diluted neutral rhCollMA.
   1. Prepare a 1.6% (w/v) solution of poly(ethylene oxide) (PEO) in phenol red-free DMEM by dissolving 160 mg of PEO in 10 mL of DMEM.
   2. Add 20 mg of lithium phenyl-2,4,6-trimethylbenzoylphosphinate (LAP) to this solution to obtain 0.2% (w/v) LAP in the solution. From this point, cover the solution with aluminum foil or keep it in a dark place to protect it from light.
   3. Sterilize the PEO-LAP solution by passing it through a 0.22 µm filter. Store this solution at 4 °C for up to 1 week.
5. Prior to bioprinting, mix the diluted neutral rhCollMA solution with the PEO-LAP solution in a 1:1 ratio to obtain the final bioink solution. Use pipetting, rather than a vortex, to mix the solutions while avoiding air bubbles.
6. If a lot of bubbles get introduced into the solution, centrifuge it at 2,000 × g for 30 s. After centrifugation, mix the bioink solution again to ensure homogeneity.

4. Granular support bath preparation

1. Prepare gelatin granular support material.
   1. In a biological safety cabinet, divide the contents of one 2 g tube of lyophilized support material into two sterile 50 mL conical tubes, each containing approximately 1 g.
   2. Add 40 mL of cold (4 °C) phenol-red free DMEM to each tube, and vortex vigorously until all the lyophilized support material dissolves.
   3. Let the resulting slurry sit at 4 °C to allow the support material to rehydrate.
   4. Degas the support material in a vacuum chamber for 30 min to reduce bubbles.
   5. Centrifuge the support material at 2,000 × g for 5 min. Ensure that the material is at the bottom of the tube and the supernatant is clear.
   6. Aspirate the supernatant, while being careful not to aspirate the support material at the bottom.
      NOTE: The support material should not flow while slanting the tube after removing the supernatant. If the material flows, centrifuge it again using the same settings but without adding new supernatant.
2. Transfer approximately 4 mL of the prepared, compacted support material into each well of a 12-well plate using a positive displacement pipette. Tap the well plate on a hard surface to force the support material to spread evenly in the well.
   NOTE: The minimal recommended volume of support material is 3x the volume of the construct that will be printed in it.
3. Place the well plate with the support material on a cooled bioprinter stage, or at 4 °C, until its use to prevent the gelatin particles from melting.

5. Incorporation of endothelial cells and support cells with the bioink

1. Prepare endothelial cell medium according to the manufacturer's instructions by mixing the basal medium with its corresponding medium kit component, including an antibiotic solution, fetal bovine serum, and endothelial cell growth supplements.
2. Prepare dental pulp stem cell medium by combining 500 mL of low-glucose DMEM, 58 mL of fetal bovine serum, 5.8 mL of non-essential amino acids (NEAA), 5.8 mL of glutamine substitute, 5.8 mL of 1 M HEPES, and 5.8 mL of penicillin-streptomycin-nystatin solution.
3. Prepare a suspension containing 2 × 10⁶ ZsGreen1-expressing human adipose microvascular endothelial cells (HAMEC-ZsGreen1) and 6 × 10⁶ dental pulp stem cells (DPSCs) in 10 mL of endothelial cell medium.
4. Centrifuge the cell suspension at 200 × g for 4 min to obtain a cell pellet. Aspirate the supernatant medium.
5. Resuspend the cell pellet in 1 mL of rhCollMA bioink prepared previously (containing PEO-LAP) to obtain a bioink with a total cell concentration of 8 × 10⁶ cells per 1 mL of bioink.
   CAUTION: After combining the bioink with the cells, proceed immediately to the next step. If cells are left in the suspension for a long time, the cells will sink to the bottom of the tube, and their viability will deteriorate. For experiments not involving cells, either skip Steps 5.1-5.5, or incorporate fluorescent beads with the bioink instead of cells for enhanced microscopic visualization of the printed constructs.
6. Transfer the cells-bioink mixture into printing cartridges.
   1. Fit a 0.22 mm internal diameter needle to a 3 mL amber printing cartridge and place the cartridge in a 50 mL conical tube.
   2. Transfer 1 mL of the cells-bioink mixture to the printing cartridge by filling it from the top, using a positive displacement pipette to reduce bubbles.
7. Install the printing cartridge in the appropriate tool at the bioprinter.

6. Bioprinting microvascular networks using rhCollMA bioink

1. Create a 3D CAD design for the bioprinted microvascular network.
   1. Sketch a 2D pattern of a 4 mm square containing a 2 mm diameter circular channel in its center. Extrude the sketch by 4 mm to obtain a 4 mm x 4 mm x 4 mm cube with a central channel. Export this object as an .STL file.
2. Import a 12-well plate .STL template into the solid modeling tab in the bioprinter slicing software and place it in the designated area of the virtual print bed.
3. Import the .STL file of the cube shape into the solid modeling tab in the bioprinter slicing software. Click on the use external slicer checkbox | configure slicer under the object properties section. In the popup window, choose a rectilinear pattern for the fill pattern, and type 30% in the fill density. Click accept.
4. Place a copy of the cube shape in each desired virtual well by clicking on the cube and moving it using the mouse.
5. Create new material settings for rhCollMA bioink in the materials tab of the bioprinter slicing software by clicking the add material button. Choose the material's settings for printing. For pressure, type 2 psi in the corresponding box. For speed, type 20 mm/s in the corresponding box.
   NOTE: Each bioink material has its own different printing settings that can be saved in the materials list in the application.
6. Assign the line width and line height values to 0.24 mm and the acceleration value to 400 mm/s², by typing these values in the corresponding boxes.
7. In the solid modeling tab, click on the cube object, and then click on the rhCollMA material from the materials section to assign it to the cube shape. In the bioassembly tab, click on the send print job button to send the print job to the bioprinter.
8. Load the printing cartridge loaded with the cellular bioink into the 3 mL ambient pneumatic dispense tool of the 3D extrusion-based bioprinter.
9. Set the print bed temperature to 4 °C by clicking on the cool button under the Control-Heating/Cooling tab in the bioprinter interface screen. Load the plate with the support bath on the printer bed.
   NOTE: The material's settings for print pressure and speed might change due to differences in ambient temperature or bioink batch variability. For instance, on colder days, a higher pressure or slower speed is needed to extrude the same amount of material.
10. In the print tab on the bioprinter interface screen, click on the print job sent in Step 6.7. and click start | go to begin the print job.
11. After printing, expose the well-plate to a 405 nm light source with an intensity of 3 mW/cm² for 30 s to initiate the crosslinking of the rhCollMA bioink.
12. After crosslinking, incubate the well plate at 37 °C and 5% CO₂ for at least 20 min until all the support bath melts.
13. Gently aspirate the liquified support bath and replace it with endothelial cell medium. Incubate the constructs at 37 °C and 5% CO₂ until use in further steps.
    ​NOTE: Due to the contraction of the construct following printing, it is recommended to perform the assembly Step 7 on the same day as Step 6.

7. Assembling the bioprinted microvascular network with the vascular scaffold to obtain the engineered vascularized flap

1. Immediately after support material removal of the bioprinted microvascular network, place a fibronectin-coated PLLA:PLGA vascular scaffold in the main channel of the bioprinted structure.
2. Incubate the assembled constructs at 37 °C and 5% CO₂ for 2 days.
3. Line the lumen of the vascular scaffold with endothelial cells.
   1. Prepare a cell suspension of tdTomato-expressing human adipose microvascular endothelial cells (HAMEC-tdTomato) at a concentration of 1 × 10⁷ cells/mL_.
   2. Place a 20 µL droplet of this cell suspension on a hydrophobic surface (i.e., a non-tissue culture [nonTC] 10 cm dish).
   3. Gently place the vascular scaffolds on top of the droplet, such that the lumen of the scaffold on one end makes contact with the cell droplet.
   4. Aspirate the droplet from the opposing end of the scaffold (i.e., the end that is not contacting the droplet), filling its lumen with the cell suspension.
   5. Place the seeded scaffold in a microcentrifuge tube and put it in a rotator inside in a humidified incubator for 60 min. Next, transfer the scaffold into a 12-well plate and add 2 mL of endothelial cell medium.
4. Culture the engineered flaps for 7 days, while replacing the medium every 2 days with fresh endothelial cell medium.

8. Confocal microscopy and immunofluorescence staining of the engineered flaps

1. After 4 days and 7 days of culture, perform live-cell imaging of the assembled scaffolds using a confocal laser-scanning microscope.
   1. Define the fluorescence channels for ZsGreen1 and tdTomato fluorescent proteins on the microscope image acquisition software.
   2. Choose a 5x/0.16 objective lens with 0.5x zoom (pixel size 5 µm), and define a Z-stack with 22 slices with a thickness of 41 µm each.
   3. Define a 3 x 2 tile scan to image and capture the entire construct.
2. Adjust the laser intensity and gain values to obtain a clear fluorescent signal with no saturation and minimal background and acquire the images.
3. Perform a maximum-intensity projection of the acquired Z-stack to obtain a single 2D image using the microscope image acquisition software or similar image analysis software.
4. Perform higher magnification imaging of a region of interest.
   1. Switch to a 10x/0.3 objective lens with 0.5 zoom (pixel size 1.25 µm) and define a Z-stack with 9 slices of thicknesses of 41 µm.
   2. Perform Steps 8.2.-8.3.
5. After 7 days of culture, fix the engineered flaps by submerging them in 4% paraformaldehyde for 20 min.
6. Wash the constructs 3x 5 min with PBS.
7. Add 0.3% (v/v) Triton-X in PBS to the constructs and incubate for 15 min at room temperature to permeabilize the cells.
8. Wash the constructs 3x 5 min with PBS.
9. Prepare a blocking solution by dissolving 5% (w/v) bovine serum albumin (BSA) in PBS. Add the blocking solution to the engineered flaps and incubate for 1 h at room temperature.
10. Prepare the primary antibody solution by diluting mouse anti-smooth muscle actin (anti-SMA) 1:50 antibody in the 5% BSA solution prepared previously to stain the SMA-expressing support cells.
11. Incubate the scaffolds in the primary antibody solution overnight at 4 °C.
12. Wash 3x 5 min with PBS.
13. Prepare the secondary antibody solution by diluting Cy3-conjugated goat anti-mouse IgG 1:400 antibody and 4',6-diamidino-2-phenylindole (DAPI, 2.5 µg/mL) in PBS.
14. Apply the secondary antibody solution to the scaffolds and incubate for 3 h at room temperature.
15. Wash 3x 5 min with PBS.
16. Store the constructs at 4 °C in PBS for up to 1 month.
17. Image the stained scaffolds using a confocal laser-scanning microscope.
    1. Define the three fluorescence channels (DAPI, ZsGreen, and Cy3) on the microscope image acquisition software.
    2. Define a Z-stack with a section thickness of 2 µm spanning a total thickness of 50 µm. Next, define a tile scan to image larger areas of the construct.
    3. Adjust the acquisition parameters to obtain a clear fluorescent signal with no saturation and minimal background.
    4. Acquire the images using 20x/0.8 objective with 1.0x zoom (pixel size 0.31 µm).
18. Perform a maximum-intensity projection of the acquired Z-stack to obtain a single 2D image using the microscope image acquisition software or similar image analysis software.

9. Anastomosing the engineered flap directly to a rat's femoral artery using a cuff technique

1. Prepare and sterilize (autoclave) the following surgical tools: No. 15 scalpel, a pair of fine-toothed forceps, and a small pair of ring-handled scissors. In addition, prepare and sterilize the following microsurgical tools: a straight fine-pointed No. 3 jeweler's forceps, an angled jeweler's forceps, a round-handled needle holder with curved jaws, a pair of vessel dilators, dissecting scissors, adventitia scissors, as well as vessel clamps with their applicator.
2. Prepare a solution of 100 IU/mL heparinized saline by diluting 5 mL of stock heparin solution with 195 mL of saline.
3. Cut and sterilize polyimide cuffs.
   1. Cut 2.5 mm sections of a polyimide tube under a dissecting microscope.
   2. At one end of each section, make a 1.25 mm lengthwise incision in the wall of the tube to obtain a cuff handle. Make an angled cut at the other end of the tube to ease vessel eversion.
   3. Submerge the cuffs in 70% ethanol followed by two washes with heparinized saline.
4. Prepare a male Sprague-Dawley rat (275-350 g) for surgery.
   1. Seven days prior to surgery, begin administering a subcutaneous daily dose of cyclosporine (10 mg kg^-1) to achieve immune suppression.
   2. Prior to surgery, anesthetize the animal using 3% isoflurane inhalation according to institutional SOP using an induction chamber. Verify deep anesthesia by pinching both feet and checking for reflexes.
   3. Administer a subcutaneous dose of analgesic buprenorphine (0.03 mg kg^-1) and anticoagulative heparin (200 IU kg^-1).
   4. Apply a lubricating eye ointment to the animal's eyes to prevent dehydration.
   5. Shave the surgical area of the rat and disinfect the site with iodine and then 70% ethanol. Secure the animal's limbs to the table using adhesive tape.
5. Expose and isolate the common femoral artery.
   1. Make a 2 cm long, oblique incision through the skin, along the concavity between the leg and the abdomen.
   2. Using forceps and blunt-ended scissors, release the skin from the underlying tissue to visualize the inguinal fat pad.
   3. Using scissors and forceps, cut right through the fat pad around the upper, medial, and lower margins. Pay attention to not cut large vessels underneath the fat pad.
   4. Reflect the fat pad laterally, leaving the epigastric vessels intact. Place a wet gauze on the reflected fat pad to prevent drying and fix it in place. Ensure that the common femoral vessels are visible.
      NOTE: The fat pad will be later used to apply soft pressure to the anastomoses.
   5. Expose the femoral artery from the inguinal ligament proximally to the epigastric artery branching point distally by extracting it from its sheath.
      CAUTION: There is usually a branch of the femoral artery running underneath it, commonly called Murphy's branch. Pay attention not to damage this difficult-to-see branch.
   6. Divide the Murphy's branch by ligating it, and then ligate the femoral artery with two ligatures at the middle of the artery spaced 1 mm apart. Keep one end of the ligature suture long to use later to pull the vessel end through the cuff.
6. Cut the artery between the two ligatures using adventitia scissors.
   CAUTION: There should not be any bleeding when performing this cut. If the arterial end bleeds, clamp the artery and reapply a ligature.
7. Using the long end of the ligature suture, insert each vessel end into a polyimide cuff, as prepared in the previous step, such that the two cuffs' handles are pointing away from each other.
8. After insertion, apply a clamp simultaneously on the cuff handle and the vessel, securing the cuff in place.
9. Prepare a heparinized saline solution in a syringe equipped with a 27 G needle.
10. Pull on the ligated vessel end and cut it close to the ligature. Immediately wash the vessel end thoroughly with the heparinized saline until no blood is visible in the lumen.
    CAUTION: Ensure that the vessel end is washed well since any blood left inside will form a clot, which will be introduced to the bloodstream upon completing the procedure. This might lead to occlusion of the vessel and loss of perfusion to the leg.
11. Expand the lumen of the vessel by holding it by its sheath with one hand and dilating its lumen with the vessel dilator with the other hand.
12. Place a loose 6-0 polypropylene circumferential suture around the cuff body. Evert the vessel end using two vessel dilators over the cuff body and secure it in place by tightening the circumferential suture.
    NOTE: If any blood leaks through the clamped vessel during the manipulations, wash the lumen thoroughly with heparinized saline.
13. Rinse the engineered flap with heparinized saline, and then insert the vessel-lined cuff into the lumen of the vascular scaffold. Fix the scaffold in place by placing a circumferential 6-0 polypropylene suture around the scaffold and the cuff body. Do this step for both sides of the vascular scaffold.
14. Wrap a 0.2 µm polystyrene membrane filter around the anastomoses site to isolate the engineered flap from the surrounding tissue.
15. Release the distal clamp first; then, release the proximal clamp to reestablish blood flow through the vessel.
16. To stop any bleeding through the anastomoses, reflect the fat pad back over the femoral vessels and apply gentle pressure using a sterile gauze for 3 min.
17. Suture the fat pad back in place using 6-0 polypropylene; then, suture the skin using 5-0 PGA absorbable sutures.
18. Clean the wound area with saline and apply iodine solution.
19. For postoperative pain and animal management, prepare a 0.1% (v/v) of tramadol in drinking water. Moreover, administer a daily dose of heparin (200 IU kg^-1) and cyclosporine (10 mg kg^-1).
20. Place the animal in a clean cage by itself placed under a heating lamp. Continue to monitor the animal until it regains sufficient consciousness to maintain sternal recumbency.