This work was supported by the Clinician Scientist Program In Vascular Medicine (PRIME), funded by the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation), project number MA 2186/14-1.

1. Preparatory work
<ol>
	<li>3D printing 
	NOTE: The following steps require the download of the &#34;Standard Triangle Language&#34; (STL) files provided as Supplemental File 1, Supplemental File 2, Supplemental File 3, Supplemental File 4, and Supplemental File 5 with this manuscript. Collector parts are provided as STL-files. The connecting flange is provided as &#34;STandard for the Exchange of Product model data&#34; (STEP) file to allow adjustment of the collector to fit individual setups. Furthermore, a technical drawing for the central metal rods is provided for conventional manufacturing as Supplemental File 6.
	<ol>
		<li>Open the slicing software (see the Table of Materials) and configure the active printhead for nonconductive polylactic acid (PLA) and a 0.4 mm nozzle. 
		NOTE: Slicing software, filament, and nozzle diameter might vary depending on the available 3D-printer.</li>
		<li>Upload the STL-files Specimen_mount_A (Supplemental File 3) and Speciment_mount_B (Supplemental File 4) into the slicing software.</li>
		<li>Rotate the models, so the triangular surfaces are placed on the build plate.</li>
		<li>Mark all parts, right-click, and select Multiply Selected Models. Enter 1 in the prompt Number of Copies and click OK. Place a total of four models on the build plate.</li>
		<li>Set slice thickness to 0.1 mm, wall thickness to 1 mm, infill density to 40%, and uncheck the Generate Support box.</li>
		<li>Click the Slice button | Save to Removable to save the printing file to a USB drive.</li>
		<li>Load nonconductive PLA into the printer and start the print file.</li>
		<li>After the print is completed, remove the models from the build plate and check for warping at the bottom corners.</li>
		<li>In the slicing software, keep the material parameters and replace the models with Collector_Flange (Supplemental File 1 and Supplemental File 5) and Leaflet_Template&#160;(Supplemental File 2).</li>
		<li>Rotate the flange, so the flat circular surface is against the build plate. Additionally, rotate the leaflet template, so the square surface is placed directly on the build plate.</li>
		<li>Mark the flange and multiply the model as in step 1.1.4. Type 1 to receive 1 copy and 1 original of the flange model on the build plate.</li>
		<li>Mark the leaflet model and multiply by 8 to receive a total of nine leaflet models, following the steps described in 1.1.4.</li>
		<li>Set slice thickness to 0.1 mm, wall thickness to 1 mm, infill density to 80%, and uncheck the Generate Support box.</li>
		<li>Click the Slice button | Save to Removable to save the new printing file to a USB drive.</li>
		<li>Load conductive PLA into the printer and start the printing process.</li>
		<li>After completion of the print, remove the models from the build plate. Remove individual filament fibers at the bottom of the leaflet negative carefully with a wire cutter if these are present in the leaflet models (as no support structures were used).</li>
	</ol>
	</li>
	<li>Spinning solution 
	CAUTION: Tetrahydrofuran (THF) and dimethylformamide (DMF) are harmful solvents that should not be inhaled or contact the skin. It is strongly recommended to wear solvent-resistant gloves and protective goggles while handling them. When handling them, work under an exhaust hood as they are extremely volatile.
	<ol>
		<li>Place a scale under the exhaust hood and position a 200 mL screw-cap glass bottle on it. Tare the scale.</li>
		<li>Pour 50 mL of DMF and 50 mL of THF into the glass bottle. Note the weight of the solvents.</li>
		<li>Place a magnetic bar inside the bottle, place the bottle on a magnetic stirrer, and switch it on.</li>
		<li>Multiply the noted weight by 0.15 (= 15% w/v) and transfer the corresponding amount of polyurethane (PU) slowly into the glass bottle containing the solvent mixture (DIN 1310).</li>
		<li>Close the bottle and stir for at least 12 h at room temperature to obtain a homogeneous solution.</li>
	</ol>
	</li>
</ol>
2. Electrospinning setup
<ol>
	<li>Assembly 
	NOTE: As the leaflet scaffolds created with the presented collector are relatively small, the optional use of a large diameter drum mandrel (D: 110 mm) is recommended. This allows the creation of larger, multilayered scaffolds, which will be beneficial for microscopic, biocompatibility, and biomechanical assessment.
	<ol>
		<li>Assemble the collector using the 3D printed parts and six M3 x 15 screws.</li>
		<li>Use three screws to secure the metal rods to one of the flanges.</li>
		<li>Slide one Specimen_mount_B between the metal bars. Make sure the spaces for templates point in the opposite direction of the flange.</li>
		<li>Fill the three slots of the Specimen_mount_B with heart valve leaflet templates.</li>
		<li>Place Specimen_mount_A on top and fill the spaces with templates.</li>
		<li>Slide another Specimen_mount_A in and fill the spaces with templates.</li>
		<li>Fixate the templates by putting the second Specimen_mount_B on top.</li>
		<li>Put the second flange on top and use the M3 screws to secure it. 
		NOTE: Ensure that the leaflet templates are all oriented in the same direction (straight edge of the leaflet parallel to the metal rods).</li>
		<li>Place the assembled leaflet collector in the electrospinning setup and tightly secure the flanges to the motor axis (i.e., M6 screws and wing nuts) (Figure 1). 
		CAUTION: As conductive PLA is more brittle than regular PLA, use a torque wrench at 1.4 Nm when fastening bolts that put pressure on the material to avoid snapping.</li>
		<li>Place a needle holder 30 cm from the collector.</li>
		<li>Fix a 14 gauge (G)&#160;needle with a flat tip in the needle holder and fixate it at the height of the collector&#39;s axis.</li>
		<li>Connect a flexible, solvent-resistant (e.g., polytetrafluoroethylene (PTFE)) tube to the Luer-lock port of the needle. 
		NOTE: DMF and THF dissolve many plastics. It is necessary to use solvent-resistant materials when working with these solvents, e.g., metal and glass tools. When plastic tools are required (i.e., syringe or tubing), make sure to use solvent-resistant materials.</li>
		<li>Guide the tube to the syringe pump for later connection of the polymer-filled syringe.</li>
	</ol>
	</li>
	<li>Connection of power supply unit (PSU) 
	CAUTION: During the setup, ensure that the power supply is disconnected from the main power source.
	<ol>
		<li>Connect two shielded, high-voltage cables to the power supply&#39;s anode and cathode.</li>
		<li>Using a crocodile clip, connect the cable connected to the cathode (- pole) to the 14 G needle. Check the connection between clip and needle. Next, guide the high-voltage cable, so it runs outside the spinning area to avoid interference.</li>
		<li>Connect the collector to the anode (+ pole) using a crocodile clip and the second high voltage cable. Use a slip ring or a sliding contact using a stripped cable to create contact at the collector&#39;s flange.</li>
	</ol>
	</li>
	<li>Preparation of the syringe 
	&#8203;NOTE: This step should be performed immediately before the spinning process starts.
	<ol>
		<li>Fill a 20 mL Luer-lock syringe with the spinning solution prepared in step 1.2.</li>
		<li>Connect the syringe to the solvent-resistant tube and manually push the solution into the tubing system until a droplet is visible at the tip of the needle.</li>
		<li>Place the syringe in the syringe pump. After turning the pump on, enter the following parameters: diameter: 19.129 mm; volume: 5 mL; speed 3 mL/h.</li>
	</ol>
	</li>
</ol>
3. Electrospinning process
<ol>
	<li>Motor test run 
	NOTE: Manufacturing the collector using 3D-printing might lead to off-center motion of the collector. Therefore, a test run with lower revolution speeds but without high voltage is highly recommended.
	<ol>
		<li>Open the motor control software by double-clicking the icon on the computer.</li>
		<li>Connect to the motor control by clicking the Connect button.</li>
		<li>After connecting, select the Profile velocity operation mode and click on the Operation tab located in the upper left corner of the screen.</li>
		<li>Select the Profile velocity tab below the Quick stop button framed by a red line. Type in the following settings: Target velocity: 200 rpm; profile acceleration: 100; profile deceleration: 200; quick stop: 5000. 
		NOTE: The direction of rotation should be upwards on the needle side, which can be adjusted by changing the sign in the &#34;target velocity&#34; field from &#34;+&#34; to &#34;-.&#34;</li>
		<li>Start the test run and check the collector for any unbalance. If the collector runs smoothly, proceed with the protocol. Otherwise, stop the motor and readjust the collector as described in step 2.1.9.</li>
		<li>Stop the motor by clicking the Switch on enabled button and change Target velocity to 2,000 rpm.</li>
	</ol>
	</li>
	<li>Manufacturing process 
	NOTE: Electrospinning is a process with a high dependency on environmental parameters. Optimal electrospinning results were obtained between 15-20% relative humidity at a temperature between 21 and 24 &#176;C.
	<ol>
		<li>First Layer 
		NOTE: During the setup phase, a dried droplet of PU might have formed at the tip of the needle. Remove the droplet using a long, nonconductive tool, if necessary.
		<ol>
			<li>In the motor control software, click the Enable Operation button to switch on the motor.</li>
			<li>Switch on the high-voltage power supply and adjust the voltage for both anode and cathode: minus pole (needle): 18 kV; plus pole (collector): 1.5 kV.</li>
			<li>Start the syringe pump at a flow rate of 3 mL/h.</li>
			<li>Set a timer to 20 min.</li>
			<li>Observe the needle tip for the formation of a tailor cone. Depending on the shape of the cone at the needle tip, adjust the voltage at the cathode in increments of &#177;100 V until a stable tailor cone is established. 
			NOTE: If the drop is hanging, the voltage is too low. However, unsteady flow can indicate that voltage is set too high.</li>
			<li>Wait for 20 min for the cusp templates to be covered adequately with fibers.</li>
			<li>Turn off the syringe pump.</li>
			<li>Turn off the PSU by flipping the power switch.</li>
			<li>Stop the motor by clicking the Switch on enabled button in the motor control software. 
			CAUTION: To prevent injury from moving parts in the system, wait until the collector has completely stopped to open the test chamber.</li>
		</ol>
		</li>
		<li>Second Layer
		<ol>
			<li>In the motor control software, alter the input field Target velocity to 10 rpm.</li>
			<li>Repeat steps 3.2.1.1-3.2.1.9.</li>
		</ol>
		</li>
		<li>Third Layer 
		NOTE: Before the scaffolds are fully dry, they are extremely sensitive to mechanical stress. Be very careful when performing steps 3.2.3.2-3.2.3.6. Avoid touching the scaffolds/fibers during these steps, as the scaffold might be rendered useless.
		<ol>
			<li>Carefully open the screws connecting the collector flanges to the motor axis and remove the leaflet collector (Figure 2B) from the electrospinning device.</li>
			<li>Using a scalpel, cut the electrospun fibers along the outer contour of each leaflet template (Figure 2C).</li>
			<li>Remove the flange on one side of the collector.</li>
			<li>Pull out the 3D-printed inserts and separate the leaflet templates from the nonconductive triangular holders.</li>
			<li>Rotate all leaflet templates by 90&#176; and reassemble the collector.</li>
			<li>Insert the collector into the electrospinning setup and tightly secure it.</li>
			<li>Again, check for any unbalance before continuing the spinning process.</li>
			<li>In the motor control software, alter the input field Target velocity to 2,000 rpm.</li>
			<li>Repeat steps 3.2.1.1-3.2.1.9. 
			NOTE: After finishing the electrospinning process, it is highly recommended to flush the tubing and needle with pure DMF to prevent clogging of the tubing.</li>
		</ol>
		</li>
		<li>Fluorescent dyed scaffolds (optional) 
		NOTE: Fluorescent dyes are used to make the fibers visible under a conventional fluorescence microscope. This is only necessary while implementing the method and for quality control after new settings have been applied. The use of fluorescent dyes is not recommended when manufacturing scaffolds using established settings.
		<ol>
			<li>Divide the spinning solution prepared in step 1.2 into three equal portions in separate bottles.</li>
			<li>Using a scale, measure 1 mg of fluorescent dye for every gram (0.1 wt%) of polymer solution. Repeat for all three fluorescent dyes (i.e., Fluorescein, Texas Red, 4&#39;,6-diamidino-2-phenylindole [DAPI]).</li>
			<li>Add the dye to the spinning solution, close the bottle lid, and stir for 2-3 h or until homogenization. 
			NOTE: To prevent the fluorescent dyes from fading, protect the spinning solution from light as much as possible, i.e., by placing an opaque cover over the magnetic stirrer. The process for fluorescent dyed scaffolds is very similar to the standard process described in steps 3.2.1-3.2.3.</li>
			<li>In step 3.2.1, replace the standard syringe with a syringe filled with the spinning solution containing the first fluorescent dye.</li>
			<li>In step 3.2.2, replace the currently used tubing and needle with new or cleaned ones. Subsequently, place a syringe with the spinning solution containing the second fluorescent dye in the syringe pump.</li>
			<li>In step 3.2.3 again, replace the tubing and needle with new or cleaned ones and replace the syringe with one that is filled with spinning solution containing the third fluorescent dye. 
			&#8203;NOTE: To avoid delay during the manufacturing process, it is beneficial to use three sets of tubing and needles. Alternatively, the tube and needle can be flushed thoroughly with THF and DMF in-between the production of layers until no spinning solution containing fluorescent dye is left in the system.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
4. Postprocessing and sample acquisition
<ol>
	<li>Postprocessing scaffolds
	<ol>
		<li>Remove the collector from the electrospinning device.</li>
		<li>Using a scalpel, cut each template free at its base as described in step 3.2.3.2.</li>
		<li>Open the collector, as described above, and place the templates, base down, on a tray.</li>
		<li>Place the tray in a drying cabinet overnight at 40 &#176;C.</li>
		<li>After the samples are completely dried, use a scalpel to carefully cut along the edges of the leaflet template to remove surplus fibers.</li>
		<li>Afterward, carefully peel the leaflet scaffold of the template and place it on a tray for further processing.</li>
	</ol>
	</li>
</ol>

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<li>Zamani, M., Shakhssalim, N., Ramakrishna, S., Naji, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Electrospinning%3A+application+and+prospects+for+urologic+tissue+engineering%5BTitle%5D)+AND+%22Frontiers+in+Bioengineering+and+Biotechnology%22%5BJournal%5D)">Electrospinning: application and prospects for urologic tissue engineering.</a> Frontiers in Bioengineering and Biotechnology. 8, 579925(2020).</li>
<li>Heilingoetter, A., Smith, S., Malhotra, P., Johnson, J., Chiang, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Applications+of+Electrospinning+for+Tissue+Engineering+in+Otolaryngology%5BTitle%5D)+AND+%22Annals+of+Otology%2C+Rhinology+%26+Laryngology%22%5BJournal%5D)">Applications of Electrospinning for Tissue Engineering in Otolaryngology.</a> Annals of Otology, Rhinology & Laryngology. 130 (4), 395-404 (2020).</li>
<li>Xue, J., Xie, J., Liu, W., Xia, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Electrospun+nanofibers%3A+new+concepts%2C+materials%2C+and+applications%5BTitle%5D)+AND+%22Accounts+of+Chemical+Research%22%5BJournal%5D)">Electrospun nanofibers: new concepts, materials, and applications.</a> Accounts of Chemical Research. 50 (8), 1976-1987 (2017).</li>
</ol>

The authors declare no conflicts of interest.

The described protocol presents two innovations in the field of (cardiovascular) tissue engineering: low-cost manufacturing of completely 3D-printed phantoms for electrospinning and the usage of a versatile collector to produce adaptable, multilayered heart valve leaflets.
Recently, 3D-printing has become a valuable tool for the production of laboratory equipment, e.g., bioreactors or manufacturing and testing setups11,12. Therefore, it was possible to manufacture the electrospinning setup presented in this study in a short amount of time and for an affordable budget (Table 1). This stays in line with previous findings for the low-cost production of electrospinning setups by using 3D-printing13. 
Moreover, to the best of the authors' knowledge, this is the first time that a conductive 3D-printing material was used to create an electrospinning collector for heart valve leaflets. So far, 3D-printed collectors were either fabricated by metal laser sintering14 or using nonconductive polymer printing and subsequent postprocessing with a conductive coating15. In contrast to this novel approach, those procedures are at a significant disadvantage as they are more expensive, take much longer, or require more manual labor.
Electrospinning depends on a multitude of variables that impact the morphology of the created fibers. Although different commercial electrospinning setups are available on the market, many research groups use highly individualized setups to match their specific needs16. Taking this into account, the described values in this protocol (voltage, distance, and rotation speed) might need to be adapted for individual setups and should be seen as a starting point rather than fixed values. Furthermore, it is known that environmental parameters can have a significant influence on electrospinning results17,18. Therefore, it is highly recommended to control at least temperature and humidity within the electrospinning rig. Optimal electrospinning results were obtained between 15-20% relative humidity at a temperature between 21 and 24 &#176;C. To follow this protocol, the following equipment is essential: a motor capable of accelerating a collector weighing approximately 300 g to a revolution speed of 2,000 rpm, a syringe pump suited for small volume flow rates of 1-3 mL/h, and a dual-pole power supply unit capable of &plusmn;20 kV direct current (DC).
In line with previous studies, it was possible to visualize the fibrous structure of the electrospun scaffolds by fluorescence microscopy19. It was possible to successfully demonstrate the multilayered structure of the scaffold, including the varying fiber orientations. Especially when working with multiple layers or multiple materials, the introduction of fluorescent dyes should be considered as a standard procedure for stringent quality control. It could improve the visual assessment of results after changes in the parameters or workflow protocol. The application of dye in scaffolds to be used for in vivo or in vitro assessment cannot be recommended. This is important to avoid interference with established analytical methods.
Mimicking natural heart valve morphology is of great importance to produce a tissue-engineered replicate to be used as a heart valve prosthesis (Figure 4B). It has been shown that the specific valve geometry has a high impact on in vivo remodeling20. In this context, 3D-printing of the leaflet geometry for electrospinning is of advantage, as iterations are easy and quick to implement. Even the production of personalized valve geometries is conceivable and subsequent development of individual and personalized 3D models of heart valve abnormalities, for example, for teaching purposes, is possible.
Further improvement of tissue-engineered heart valve properties is at the center of current research efforts, as several research groups have worked on developing multilayered scaffolds with defined fiber orientations. Masoumi et al. fabricated composite scaffolds from a molded polyglycerol sebacate layer and electrospun polycaprolactone (PCL) fiber mats21. Thus, a triple layer could be created out of two orientated electrospun layers separated by a sheet of microfabricated polyglycerol sebacate. However, in contrast to the scaffolds on hand, they were neither in a 3D shape nor did they adequately mimic the middle layer (spongiosa). Another approach to producing a bioinspired tissue-engineered heart valve was pursued by Jana et al.22,23. They successfully produced triple-layered scaffolds with orientated fibers using aluminum collectors for PCL-based electrospinning. Again, these scaffolds also presented morphological imperfections, as they only have a 2D appearance, and the final scaffold is pervaded by spokes. 
Even though the protocol gives detailed information on how 3D, triple-layered heart valve leaflets are produced, there are several more steps needed to create an actual heart valve prosthesis. A stent of 24 mm diameter is recommended for the leaflets described here. Complementary to the stent used, the leaflets can be provided with additional support structures for stitching. To allow maximal flexibility, the leaflets shown here are not individualized to a specific stent design. This can be done by simply altering the template using CAD software.
Although used for heart valve tissue engineering, the presented method will be readily applicable for electrospinning setups in orthopedics24, urology25, otolaryngology26, and others. The production of sophisticated and/or individualized 3D constructs is feasible by the implementation of other 3D-printed collectors. Although the material of the collector has changed, the principle of electrospinning stays intact27. Therefore, the usage of different polymers is theoretically possible, although adjustment of the electrospinning parameters may be necessary.
Overall, the presented protocol describes an easy and cost-effective way to manufacture multilayered heart valve leaflets. The application of 3D-printing allows for fast adaptation and modifications of the collector and the inserts. This allows the production of patient-specific prostheses without a complicated manufacturing process of, for example, metal collectors. Multiple samples can be created in one run under identical conditions. Therefore, material destructive tests can be performed on the samples with the benefit of having (nearly) identical ones remaining to build the actual valve. The inclusion of the printing files as Supplemental Files in this study is meant to support the advancement of multilayered heart valve scaffolds. This new electrospinning technique also has a high potential for other fields of regenerative medicine, as modified collectors and other 3D-printed, spinning templates are easy to implement.

Cardiovascular disease is the leading cause of death in western countries 1. Although extensive research is done in this field, it is estimated that the burden of degenerative heart valve disease will increase even further during the next years2. Surgical or interventional heart valve replacement is possible as a therapeutic option. At this point, mechanical and bioprosthetic heart valves are available, both with individual drawbacks. Mechanical valves are thrombogenic and require lifelong anticoagulation. Although biological valves do not require anticoagulation, they show a lack of remodeling, a high rate of calcification, and concomitant degradation3.
Tissue-engineered heart valves might be able to address these drawbacks by introducing a microfibrous scaffold into the body that allows in vivo remodeling. Various methods, e.g., electrospinning (ESP), decellularization, micromolding, spray, dip-coat, and 3D-bioprinting, are available. These methods can be chosen for creating specific properties, being cheaper and faster, or just due to a lack of alternatives. Methods and materials can even be combined to create more complex structures4. For example, ESP has been a standard technique for creating scaffolds in tissue engineering, allowing for the combination of different materials and the adjustment of fiber diameters, fiber orientations, and porosities4. Furthermore, a variety of postprocessing techniques allow for optimized tissue remodeling, improved hemocompatibility, and adjustable biodegradation of electrospun scaffolds5,6,7.
Basic ESP uses either static or rotating collectors, which have a direct influence on the degree of fiber alignment and the obtained fiber diameters8. Due to manufacturing restrictions, classic ESP rotating collectors consist of rotating drums, discs, wires, or metal rods. The introduction of 3D-printing allows for the creation of more individualized collector geometries that are not limited by traditional manufacturing techniques. This individualization is especially useful for the creation of 3D constructs such as heart valve leaflets.
The natural three-layered (fibrosa, spongiosa, ventricularis) architecture of human heart valve leaflets is the tissues&#39; response to the mechanical forces and shear stress they are exposed to during the cardiac cycle9,10. The fibers of the lamina fibrosa are oriented circumferentially, whereas the fibers of the lamina spongiosa are randomly aligned and those of the lamina ventricularis radially. A triple-layer with the corresponding fiber orientations is thus proposed to mimic the properties of these valves in a tissue-engineered scaffold.
The workflow protocol describes an innovative method to produce three-layered, 3D heart valve leaflets using 3D-printing and electrospinning. Additionally, a quality control step is presented to ensure accurate fiber orientation in every layer.

<table><tbody><tr><td>BTC-FR2.5TN.D09</td><td>ZwickRoell GmbH &amp;amp; Co. KG</td><td></td><td>Traction engine (Tensile tests)</td></tr><tr><td>C5-E Motor Controller</td><td>Nanotec Electronic GmbH &amp;amp; Co. KG</td><td></td><td>Motor controll unit</td></tr><tr><td>CH1: CPN 30 kV | 0.3 mA</td><td>iseg Spezialelectronik GmbH</td><td></td><td>Power Supply Unit Anode</td></tr><tr><td>CH1: CPN 30 kV | 0.3 mA</td><td>iseg Spezialelektronik GmbH</td><td></td><td>Power Supply Unit Kathode</td></tr><tr><td>Conductive Composite PLA</td><td>ProtoPasta</td><td></td><td>Conductive PLA</td></tr><tr><td>Cura 4.7.1</td><td>Ultimaker BV</td><td></td><td>Slicing Software Ultimaker, step 1.1.2</td></tr><tr><td>DAPI Stock Solution c = 0.1 mg/mL</td><td>Sigma-Aldrich Chemie GmbH</td><td></td><td>DAPI</td></tr><tr><td>Disposable Scalpel No. 23</td><td>FEATHER</td><td></td><td>Scalpel</td></tr><tr><td>Fluorescein (C.I. 45350) M 376.28 g/mol</td><td>Carl Roth GmbH + Co. KG</td><td></td><td>Fluorescein</td></tr><tr><td>Fume Hood as per DIN 12924 Class 2</td><td>K&amp;ouml;ttermann GmbH</td><td></td><td>Fume Hood</td></tr><tr><td>Leica Applicatin Suite X 3.5.5.19976</td><td>Leica Microsystems GmbH</td><td></td><td>Software for Confocal Laser Scanning Microscope</td></tr><tr><td>Luerlock Syringe 20 mL</td><td>BD Plastipak</td><td></td><td>Luerlock Syringe</td></tr><tr><td>Metal needle plane 2.50/2.00 x 20 mm</td><td>Unimed S.A.</td><td></td><td>Needle with plane tip</td></tr><tr><td>Montage-complet-tubes; inner diameter x outer diameter: 1/16&quot; x 1/8&quot;, length 1.000 mm</td><td>Bohlender GmbH</td><td>F740-28</td><td>Solvent resistant tubes</td></tr><tr><td>N,N-Dimethylformamide &amp;ge;99.8%</td><td>Sigma-Aldrich Chemie GmbH</td><td></td><td>Dimethylformamide</td></tr><tr><td>Pellethane 2363 80AE</td><td>Velox GmbH Hamburg</td><td></td><td>Polyurethane</td></tr><tr><td>PLA</td><td>Ultimaker BV</td><td></td><td>PLA</td></tr><tr><td>Plug&amp;amp;Drive Studio (1.0.4)</td><td>Nanotec Electronic GmbH &amp;amp; Co. KG</td><td></td><td>Motor operation software</td></tr><tr><td>SEM Evo LS 10</td><td>Zeiss MicroImaging GmbH</td><td></td><td>Scanning Electron Microscope</td></tr><tr><td>SHT 31-D</td><td>Adafruit Industries</td><td></td><td>Temperature and Humidity Sensor</td></tr><tr><td>SolidWorks 2020 CAD Software</td><td>Dassault Syst&amp;egrave;mes</td><td></td><td>Commercial CAD Software</td></tr><tr><td>Sulforhodamine 101 50 mg</td><td>Sigma - Aldrich&amp;nbsp;</td><td>S 7635</td><td>Texas Red</td></tr><tr><td>Syringe Pump Model: Fusion 100</td><td>Chemyx Inc.</td><td></td><td>Syringe Pump</td></tr><tr><td>TCS SP8 inverted CEL BMi8</td><td>Leica Microsystems GmbH</td><td></td><td>Confocal Laser Scanning Microscope</td></tr><tr><td>testXpert V11.02</td><td>ZwickRoell GmbH &amp;amp; Co. KG</td><td></td><td>Software Tensile Test</td></tr><tr><td>Tetrahydrofuran &amp;ge;99.9%</td><td>Sigma-Aldrich Chemie GmbH</td><td></td><td>Tetrahydrofuran</td></tr><tr><td>Type 1511530000202 #980361</td><td>Binder Labortechnik GmbH</td><td></td><td>Heating Cabinet</td></tr><tr><td>Ultimaker 3 Extended</td><td>Ultimaker BV</td><td></td><td>3D Printer</td></tr></tbody></table>

combining 3d-printing and electrospinning to manufacture biomimetic heart valve leaflets

Electrospinning has become a widely used technique in cardiovascular tissue engineering as it offers the possibility to create (micro-)fibrous scaffolds with adjustable properties. The aim of this study was to create multilayered scaffolds mimicking the architectural fiber characteristics of human heart valve leaflets using conductive 3D-printed collectors.
Models of aortic valve cusps were created using commercial computer-aided design (CAD) software. Conductive polylactic acid was used to fabricate 3D-printed leaflet templates. These cusp negatives were integrated into a specifically designed, rotating electrospinning mandrel. Three layers of polyurethane were spun onto the collector, mimicking the fiber orientation of human heart valves. Surface and fiber structure was assessed with a scanning electron microscope (SEM). The application of fluorescent dye additionally permitted the microscopic visualization of the multilayered fiber structure. Tensile testing was performed to assess the biomechanical properties of the scaffolds.
3D-printing of essential parts for the electrospinning rig was possible in a short time for a low budget. The aortic valve cusps created following this protocol were three-layered, with a fiber diameter of 4.1 &#177; 1.6 &#181;m. SEM imaging revealed an even distribution of fibers. Fluorescence microscopy revealed individual layers with differently aligned fibers, with each layer precisely reaching the desired fiber configuration. The produced scaffolds showed high tensile strength, especially along the direction of alignment. The printing files for the different collectors are available as Supplemental File 1, Supplemental File 2, Supplemental File 3, Supplemental File 4, and Supplemental File 5.
With a highly specialized setup and workflow protocol, it is possible to mimic tissues with complex fiber structures over multiple layers. Spinning directly on 3D-printed collectors creates considerable flexibility in manufacturing 3D shapes at low production costs.

This protocol is aimed at the development of a triple-layered leaflet scaffold destined for use in cardiovascular tissue engineering of heart valves. It mimics the collagen configuration of the three layers in the native human heart valve. Each layer consists of fibers with an overall diameter of 4.1 &#177; 1.6 &#181;m (Figure 1).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63604/63604fig01.jpg" /> 
Figure 1: Fiber characteristics. Analysis of fibers: Total fiber count; Diameter in &#181;m: mean, mode, standard deviation, minimal diameter, maximal diameter. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63604/63604fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The leaflet templates are designed to fit a &#216; 24 mm aortic valve prosthesis (Figure 2C). After drying, the leaflet scaffolds kept their shape of a 3D heart valve cusp (Figure 3A).
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/63604/63604fig02.jpg" /> 
Figure 2: Electrospinning setup. (A) Assembled 3D-printed collector in the rotary setup; (B) CAD rendering of the 3D-printable collector; (C) CAD rendering of the heart valve leaflet negative shown in B; triangle indicates zoomed-in part. Abbreviation: CAD = computer-aided design. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63604/63604fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
SEM imaging was used to assess the aligned and unaligned layers (TEMP F3512-21). Photographs were taken at 100x, 500x, and 2,000x magnification in three different locations on a scaffold. Aligned fiber scaffolds appear with a smooth surface and strict orientation in the circumferential direction (Figure 3B). Visual analysis of the 2,000x image with respect to the fiber orientation confirms the primary alignment of the fibers (Figure 3C). Unaligned fiber scaffolds show a similarly smooth surface compared to the aligned fibers. Fiber orientation is disordered, with many prominent intersections between fibers (Figure 3D). Subsequent visual analysis confirms the unalignment of fibers with no primary orientation visible (Figure 3E).
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/63604/63604fig03.jpg" /> 
Figure 3: Electrospun leaflet and SEM imaging. (A) Electrospun multilayered leaflet and 3D-printed leaflet collector; (B) SEM image of unaligned fibers (magnification 1,000x); (C) Fiber orientation analysis of unaligned fibers; (D) SEM image of aligned fibers (magnification 1,000x); (E) Fiber orientation analysis of aligned fibers. Scale bars = 10 mm (A), 100 &#181;m (B, D). Abbreviation: SEM = scanning electron microscopy. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63604/63604fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Imaging of fluorescent dyed multilayered scaffolds revealed three individual layers with distinct fiber orientations (Figure 4D). The bottom layer (Figure 4A; blue) shows aligned fibers in horizontal orientation with very little intersection between the fibers. The middle layer (Figure 4B; green) shows unaligned fibers with no primary fiber orientation. The top layer (Figure 4C; red) shows aligned fibers in a perpendicular orientation. Visual analysis of the top and bottom layers reveals an average angle between the two layers of 89&#176;, which is in accordance with the 90&#176; rotation of the collector during the spinning process (Figure 4E).
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/63604/63604fig04.jpg" /> 
Figure 4: Fluorescence microscopy of multilayered scaffold. (A) Fluorescence image of the first layer with primary orientation from bottom left to top right; (B) Fluorescence image of the second layer with unaligned fiber orientation; (C) Fluorescence image of the third layer with primary orientation from bottom right to top left; (D) Fluorescence image of all three layers combined in one scaffold; (E) Fiber orientation analysis for all three layers (Layer 1: blue; Layer 2: green; Layer 3: red); magnification = 400x (A-D); scale bars = 100 &#181;m (A-D). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63604/63604fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Thickness measurement was done on 21 samples (Figure 5A) (TEMP F3510-21). All samples were created applying the same parameters. Temperature and humidity could differ between 20.3 &#176;C and 26.1 &#176;C and 35% and 55% humidity, respectively. The results showed a relatively linear increase in thickness of ~2.65 &#181;m per min.
Another experiment showed the consistency of the results after 60 min of spinning under matching parameters (Figure 5B). Humidity and temperature could differ between 35% and 50% humidity and 20.3 &#176;C to 26.1 &#176;C, respectively. The results were scaffolds between 126 and 181 &#181;m in thickness. The average thickness was 151.11 &#177; 13.17 &#181;m. The increase in thickness was ~2.52 &#181;m per min, on average.
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/63604/63604fig05.jpg" /> 
Figure 5: Thickness measurement. (A) Thickness of scaffolds per time spun; n = 21; Correlation coefficient (r) = 0.653; p** = 0.00132; (B) Thickness of samples after 60 min; n = 13; red line: mean. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63604/63604fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Tensile tests for aligned and unaligned fiber scaffolds were performed in two directions, along the circumferential direction and perpendicular to it. Each grout consisted of 15 specimens. Samples were taken out of plane scaffolds according to DIN 53504:2017-03. The thickness was measured at three different spots on each sample and used to calculate the maximal force values per square mm.
The thickness values lay between 0.03 and 0.2 mm. The comparison of ultimate tensile strength revealed a significant difference (p &#60; 0.001) between orientations for the aligned fiber scaffolds (Figure 6A). The scaffolds reached a maximum strength of 12.26 &#177; 2.59 N/mm2 along the circumferential orientation. The tensile strength was reduced to 3.86 &#177; 1.08 N/mm2 in the perpendicular direction.
Unaligned fiber scaffolds show no difference in the ultimate tensile strength for the different orientations (F1: 7.19 &#177; 1.75 N/mm2, F2: 7.54 &#177; 1.59 N/mm2; p = 0.60). The comparative analysis of the elongation at break for the aligned fiber scaffolds revealed significant differences (p &#60; 0.001) in distensibility between the directions (Figure 6B). The extensibility reached 187.01 &#177; 39.37% in the circumferential direction compared to 107.16 &#177; 30.04% in the perpendicular direction.
In contrast, the elongation at break for the unaligned fiber mats revealed uniform extensibility in both directions (F1: 269.74 &#177; 24.78 % ; F2: 285.01 &#177; 25.58 %; p = 0.69). Representative stress-strain curves show huge differences in the behavior of the material, depending on the direction in which the tensile force is applied. Unaligned fiber mats showed linear elastic behavior, while aligned fiber mats showed nonlinearity in the axial direction.
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/63604/63604fig06.jpg" /> 
Figure 6: Tensile tests of aligned and unaligned fibers. (A) Ultimate tensile strength for aligned and unaligned fiber mats in circumferential and axial directions; n = 15; (B) Elongation at break for aligned and unaligned fiber mats in circumferential and axial directions; n = 15; (C) Representative stress-strain curves of aligned and unaligned scaffolds, pulled in axial and circumferential directions, respectively. (***p &#60; 0.001). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63604/63604fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td style="min-width: 40px;"></td>
			<td colspan="2" style="min-width: 60px;"></td>
			<td colspan="2" style="min-width: 60px;"></td>
			<td colspan="2" style="min-width: 60px;">Manufacturing Metrics</td>
			<td style="min-width: 60px;"></td>
			<td style="min-width: 60px;"></td>
			<td style="min-width: 60px;"></td>
		</tr>
		<tr>
			<td style="min-width: 40px;"></td>
			<td colspan="2" style="min-width: 60px;">Name</td>
			<td colspan="2" style="min-width: 60px;">Material</td>
			<td style="min-width: 60px;">Amount</td>
			<td style="min-width: 60px;">Total Time</td>
			<td style="min-width: 60px;">Total Weight [g]</td>
			<td style="min-width: 60px;">Cost [&#8364; per kg]</td>
			<td style="min-width: 60px;">Total Cost</td>
		</tr>
		<tr>
			<td style="min-width: 40px;">1</td>
			<td colspan="2" style="min-width: 60px;">Specimen_Mount_A</td>
			<td colspan="2" style="min-width: 60px;">Regular PLA</td>
			<td style="min-width: 60px;">2</td>
			<td style="min-width: 60px;">18:19</td>
			<td style="min-width: 60px;">159</td>
			<td style="min-width: 60px;">51.33 &#8364;</td>
			<td style="min-width: 60px;">8.16 &#8364;</td>
		</tr>
		<tr>
			<td style="min-width: 40px;">2</td>
			<td colspan="2" style="min-width: 60px;">Specimen_Mount_B</td>
			<td colspan="2" style="min-width: 60px;">Regular PLA</td>
			<td style="min-width: 60px;">2</td>
			<td style="min-width: 60px;">19:42</td>
			<td style="min-width: 60px;">161</td>
			<td style="min-width: 60px;">51.33 &#8364;</td>
			<td style="min-width: 60px;">8.26 &#8364;</td>
		</tr>
		<tr>
			<td style="min-width: 40px;">3</td>
			<td colspan="2" style="min-width: 60px;">Collector Flange</td>
			<td colspan="2" style="min-width: 60px;">Conductive PLA</td>
			<td style="min-width: 60px;">2</td>
			<td style="min-width: 60px;">10:40</td>
			<td style="min-width: 60px;">95</td>
			<td style="min-width: 60px;">99.98 &#8364;</td>
			<td style="min-width: 60px;">9.50 &#8364;</td>
		</tr>
		<tr>
			<td style="min-width: 40px;">4</td>
			<td colspan="2" style="min-width: 60px;">Leaflet_Inlet</td>
			<td colspan="2" style="min-width: 60px;">Conductive PLA</td>
			<td style="min-width: 60px;">9</td>
			<td style="min-width: 60px;">05:32</td>
			<td style="min-width: 60px;">31</td>
			<td style="min-width: 60px;">99.98 &#8364;</td>
			<td style="min-width: 60px;">3.10 &#8364;</td>
		</tr>
		<tr>
			<td style="min-width: 40px;"></td>
			<td colspan="2" style="min-width: 60px;">Total</td>
			<td colspan="2" style="min-width: 60px;"></td>
			<td style="min-width: 60px;"></td>
			<td style="min-width: 60px;"></td>
			<td style="min-width: 60px;"></td>
			<td style="min-width: 60px;"></td>
			<td style="min-width: 60px;">29.02 &#8364;</td>
		</tr>
	</tbody>
</table>
Table 1: Manufacturing metrics. Table specifying quantity, manufacturing time, amount of material needed, and costs for 3D-printed parts. Abbreviation: PLA = polylactic acid.
Supplemental File 1: Adaptable collector flange. Step-file to adapt and print collector flange. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63604/Collector_Flange_adaptable.STEP" target="_blank">Please click here to download this File.</a>
Supplemental File 2: Leaflet template. STL-file to print leaflet template. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63604/Leaflet_Template.STL" target="_blank">Please click here to download this File.</a>
Supplemental File 3: Specimen mount A. STL-file to print specimen mount A. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63604/Specimen_Mount_A.STL" target="_blank">Please click here to download this File.</a>
Supplemental File 4: Specimen mount B. STL-file to print specimen mount B. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63604/Specimen_Mount_B.STL" target="_blank">Please click here to download this File.</a>
Supplemental File 5: Collector flange. STL-file to print collector flange. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63604/Collector_Flange.STL" target="_blank">Please click here to download this File.</a>
Supplemental File 6: Connecting metal rod. Technical drawing to construct connecting metal rods. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63604/Connecting_Rod.zip" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Combining 3D-Printing and Electrospinning to Manufacture Biomimetic Heart Valve Leaflets at JoVE.com

Combining 3D-Printing and Electrospinning to Manufacture Biomimetic Heart Valve Leaflets

The presented method offers an innovative way for engineering biomimetic fiber structures in three-dimensional (3D) scaffolds (e.g., heart valve leaflets). 3D-printed, conductive geometries were used to determine shape and dimensions. Fiber orientation and characteristics were individually adjustable for each layer. Multiple samples could be manufactured in one setup.

Electrospinning has become a widely used technique in cardiovascular tissue engineering as it offers the possibility to create (micro-)fibrous scaffolds ...

combining-3d-printing-electrospinning-to-manufacture-biomimetic-heart

Nanyang Technological University

Research

JoVE Journal

Medicine

4.2K Views.  Ludwig Maximilians University Munich. Drielaagse structuren bestaande uit georiënteerde vezels zijn overal in het menselijk lichaam te vinden. Door deze methode toe te passen, kan men niet alleen hartklepblaadjes maken die natuurlijke omstandigheden nabootsen, maar ook een verscheidenheid aan ander weefsel. Dit is de eerste keer dat 3D-geprinte collectoren uit geleidend materiaal zijn gebruikt bij elektrospinning, een feit dat dit proces zeer flexibel en kosteneffectief maakt.Begin om te beginnen met 3D-printen door het S...

Video: Combining 3D-Printing and Electrospinning to Manufacture Biomimetic Heart Valve Leaflets

Tri-layered Electrospinning to Mimic Native Arterial Architecture using Polycaprolactone, Elastin, and Collagen: A Preliminary Study

Throughout native artery, collagen and elastin play an important role, providing a mechanical backbone, preventing vessel rupture, and promoting recovery under pulsatile deformations. The goal of this study was to mimic the structure of native artery by fabricating a multi-layered electrospun conduit composed of poly(caprolactone) (PCL) with the addition of elastin and collagen with blends of 45-45-10, 55-35-10, and 65-25-10 PCL-ELAS-COL to demonstrate mechanical properties indicative of native arterial tissue, while remaining conducive to tissue regeneration. Whole grafts and individual layers were analyzed using uniaxial tensile testing, dynamic compliance, suture retention, and burst strength. Compliance results revealed that changes to the middle/medial layer changed overall graft behavior with whole graft compliance values ranging from 0.8 - 2.8 % / 100 mmHg, while uniaxial results demonstrated an average modulus range of 2.0 - 11.8 MPa. Both modulus and compliance data displayed values within the range of native artery. Mathematical modeling was implemented to show how changes in layer stiffness affect the overall circumferential wall stress, and as a design aid to achieve the best mechanical combination of materials. Overall, the results indicated that a graft can be designed to mimic a tri-layered structure by altering layer properties.

The aim of this study was to mimic the native three layered architecture of the arterial wall. To accomplish this, electrospinning was employed with the use of a 3-1 (input-output) nozzle and blends of polycaprolactone, elastin, and collagen.

Throughout native artery, collagen and elastin play an important role, providing a mechanical backbone, preventing vessel rupture, and promoting recovery ...

Bioengineering

Combination of Microstereolithography and Electrospinning to Produce Membranes Equipped with Niches for Corneal Regeneration

Corneal problems affect millions of people worldwide reducing their quality of life significantly. Corneal disease can be caused by illnesses such as Aniridia or Steven Johnson Syndrome as well as by external factors such as chemical burns or radiation. Current treatments are (i) the use of corneal grafts and (ii) the use of stem cell expanded in the laboratory and delivered on carriers (e.g., amniotic membrane); these treatments are relatively successful but unfortunately they can fail after 3-5 years. There is a need to design and manufacture new corneal biomaterial devices able to mimic in detail the physiological environment where stem cells reside in the cornea. Limbal stem cells are located in the limbus (circular area between cornea and sclera) in specific niches known as the Palisades of Vogt. In this work we have developed a new platform technology which combines two cutting-edge manufacturing techniques (microstereolithography and electrospinning) for the fabrication of corneal membranes that mimic to a certain extent the limbus. Our membranes contain artificial micropockets which aim to provide cells with protection as the Palisades of Vogt do in the eye.

We report a technique for the fabrication of micropockets within electrospun membranes in which to study cell behavior. Specifically, we describe a combination of microstereolithography and electrospinning for the production of PLGA (Poly(lactide-co-glycolide)) corneal biomaterial devices equipped with microfeatures.

Corneal problems affect millions of people worldwide reducing their quality of life significantly. Corneal disease can be caused by illnesses such as ...

Adapting the Electrospinning Process to Provide Three Unique Environments for a Tri-layered In Vitro Model of the Airway Wall

Electrospinning is a highly adaptable method producing porous 3D fibrous scaffolds that can be exploited in in vitro cell culture. Alterations to intrinsic parameters within the process allow a high degree of control over scaffold characteristics including fiber diameter, alignment and porosity. By developing scaffolds with similar dimensions and topographies to organ- or tissue-specific extracellular matrices (ECM), micro-environments representative to those that cells are exposed to in situ can be created. 
The airway bronchiole wall, comprised of three main micro-environments, was selected as a model tissue. Using decellularized airway ECM as a guide, we electrospun the non-degradable polymer, polyethylene terephthalate (PET), by three different protocols to produce three individual electrospun scaffolds optimized for epithelial, fibroblast or smooth muscle cell-culture. Using a commercially available bioreactor system, we stably co-cultured the three cell-types to provide an in vitro model of the airway wall over an extended time period.
This model highlights the potential for such methods being employed in in vitro diagnostic studies investigating important inter-cellular cross-talk mechanisms or assessing novel pharmaceutical targets, by providing a relevant platform to allow the culture of fully differentiated adult cells within 3D, tissue-specific environments.

Adapting the Electrospinning Process to Provide Three Unique Environments for a Tri-layered In Vitro Model of the Airway Wall

Advancements in biomaterial technologies enable the development of three-dimensional multi-cell-type constructs. We have developed electrospinning protocols to produce three individual scaffolds to culture the main structural cells of the airway to provide a 3D in vitro model of the airway bronchiole wall.

Electrospinning is a highly adaptable method producing porous 3D fibrous scaffolds that can be exploited in in vitro cell culture. Alterations to ...

Creation of Cardiac Tissue Exhibiting Mechanical Integration of Spheroids Using 3D Bioprinting

This protocol describes 3D bioprinting of cardiac tissue without the use of biomaterials, using only cells. Cardiomyocytes, endothelial cells and fibroblasts are first isolated, counted and mixed at desired cell ratios. They are co-cultured in individual wells in ultra-low attachment 96-well plates. Within 3 days, beating spheroids form. These spheroids are then picked up by a nozzle using vacuum suction and assembled on a needle array using a 3D bioprinter. The spheroids are then allowed to fuse on the needle array. Three days after 3D bioprinting, the spheroids are removed as an intact patch, which is already spontaneously beating. 3D bioprinted cardiac patches exhibit mechanical integration of component spheroids and are highly promising in cardiac tissue regeneration and as 3D models of heart disease.

This protocol describes 3D bioprinting of cardiac tissue without the use of biomaterials. 3D bioprinted cardiac patches exhibit mechanical integration of component spheroids and are highly promising in cardiac tissue regeneration and as 3D models of heart disease.

This protocol describes 3D bioprinting of cardiac tissue without the use of biomaterials, using only cells. Cardiomyocytes, endothelial cells and ...

Microfluidic Bioprinting for Engineering Vascularized Tissues and Organoids

Engineering vascularized tissue constructs and organoids has been historically challenging. Here we describe a novel method based on microfluidic bioprinting to generate a scaffold with multilayer interlacing hydrogel microfibers. To achieve smooth bioprinting, a core-sheath microfluidic printhead containing a composite bioink formulation extruded from the core flow and the crosslinking solution carried by the sheath flow, was designed and fitted onto the bioprinter. By blending gelatin methacryloyl (GelMA) with alginate, a polysaccharide that undergoes instantaneous ionic crosslinking in the presence of select divalent ions, followed by a secondary photocrosslinking of the GelMA component to achieve permanent stabilization, a microfibrous scaffold could be obtained using this bioprinting strategy. Importantly, the endothelial cells encapsulated inside the bioprinted microfibers can form the lumen-like structures resembling the vasculature over the course of culture for 16 days. The endothelialized microfibrous scaffold may be further used as a vascular bed to construct a vascularized tissue through subsequent seeding of the secondary cell type into the interstitial space of the microfibers. Microfluidic bioprinting provides a generalized strategy in convenient engineering of vascularized tissues at high fidelity.

We provide a generalized protocol based on a microfluidic bioprinting strategy for engineering a microfibrous vascular bed, where a secondary cell type could be further seeded into the interstitial space of this microfibrous structure to generate vascularized tissues and organoids.

Engineering vascularized tissue constructs and organoids has been historically challenging. Here we describe a novel method based on microfluidic ...

Melt Electrospinning Writing of Three-dimensional Poly(&#949;-caprolactone) Scaffolds with Controllable Morphologies for Tissue Engineering Applications

This tutorial reflects on the fundamental principles and guidelines for electrospinning writing with polymer melts, an additive manufacturing technology with great potential for biomedical applications. The technique facilitates the direct deposition of biocompatible polymer fibers to fabricate well-ordered scaffolds in the sub-micron to micro scale range. The establishment of a stable, viscoelastic, polymer jet between a spinneret and a collector is achieved using an applied voltage and can be direct-written. A significant benefit of a typical porous scaffold is a high surface-to-volume ratio which provides increased effective adhesion sites for cell attachment and growth. Controlling the printing process by fine-tuning the system parameters enables high reproducibility in the quality of the printed scaffolds. It also provides a flexible manufacturing platform for users to tailor the morphological structures of the scaffolds to their specific requirements. For this purpose, we present a protocol to obtain different fiber diameters using melt electrospinning writing (MEW) with a guided amendment of the parameters, including flow rate, voltage and collection speed. Furthermore, we demonstrate how to optimize the jet, discuss often experienced technical challenges, explain troubleshooting techniques and showcase a wide range of printable scaffold architectures.

This protocol serves as a comprehensive guideline to fabricate scaffolds via electrospinning with polymer melts in a direct writing mode. We systematically outline the process and define the appropriate parameter settings for achieving targeted scaffold architectures.

This tutorial reflects on the fundamental principles and guidelines for electrospinning writing with polymer melts, an additive manufacturing technology ...

Fabrication of Engineered Vascular Flaps Using 3D Printing Technologies

Engineering implantable, functional, thick tissues requires designing a hierarchical vascular network. 3D bioprinting is a technology used to create tissues by adding layer upon layer of printable biomaterials, termed bioinks, and cells in an orderly and automatic manner, which allows for creating highly intricate structures that traditional tissue engineering techniques cannot achieve. Thus, 3D bioprinting is an appealing in vitro approach to mimic the native vasculature complex structure, ranging from millimetric vessels to microvascular networks. 
Advances in 3D bioprinting in granular hydrogels enabled the high-resolution extrusion of low-viscosity extracellular matrix-based bioinks. This work presents a combined 3D bioprinting and sacrificial mold-based 3D printing approach for fabricating engineered vascularized tissue flaps. 3D bioprinting of endothelial and support cells using recombinant collagen-methacrylate bioink within a gelatin support bath is utilized for the fabrication of a self-assembled capillary network. This printed microvasculature is assembled around a mesoscale vessel-like porous scaffold, fabricated using a sacrificial 3D printed mold, and is seeded with endothelial cells. 
This assembly induces the endothelium of the mesoscale vessel to anastomose with the surrounding capillary network, establishing a hierarchical vascular network within an engineered tissue flap. The engineered flap is then directly implanted by surgical anastomosis to a rat femoral artery using a cuff technique. The described methods can be expanded for the fabrication of various vascularized tissue flaps for use in reconstruction surgery and vascularization studies.

Engineered flaps require an incorporated functional vascular network. In this protocol, we present a method of fabricating a 3D printed tissue flap containing a hierarchical vascular network and its direct microsurgical anastomoses to rat femoral artery.

Engineering implantable, functional, thick tissues requires designing a hierarchical vascular network. 3D bioprinting is a technology used to create ...

Three-Dimensional Printing of a Complex Aortic Anomaly

Complex congenital aortic anomalies include diverse types of malformations that may be clinically asymptomatic or present with respiratory or esophageal symptoms. These anomalies may be associated with other congenital heart diseases. It is hard to identify the accurate anatomic vessel location from two-dimensional imaging data, such as computed tomography. As an additive manufacturing method, three-dimensional (3-D) printing can covert the acquired imaging data into 3-D physical models. This protocol describes the procedure for modeling the volumetric DICOM imaging into 3-D data and printing it as an anatomically realistic 3-D model. Using this model, surgeons can identify the vessel location of complex aortic anomalies, which is helpful for pre-operative planning and intra-operative guidance.

Here, we present a protocol to use three dimensional printed models for pre-operative planning and intra-operative reorganization of complicated vascular locations when handling a congenital aortic anomaly.

Complex congenital aortic anomalies include diverse types of malformations that may be clinically asymptomatic or present with respiratory or esophageal ...

Phototunable Bioprinted Hydrogels to Probe Cellular Responses to ECM Stiffening

3D Bioprinting Phototunable Hydrogels to Study Fibroblast Activation

Phototunable hydrogels can transform spatially and temporally in response to light exposure. Incorporating these types of biomaterials in cell-culture platforms and dynamically triggering changes, such as increasing microenvironmental stiffness, enables researchers to model changes in the extracellular matrix (ECM) that occur during fibrotic disease progression. Herein, a method is presented for 3D bioprinting a phototunable hydrogel biomaterial capable of two sequential polymerization reactions within a gelatin support bath. The technique of Freeform Reversible Embedding of Suspended Hydrogels (FRESH) bioprinting was adapted by adjusting the pH of the support bath to facilitate a Michael addition reaction. First, the bioink containing poly(ethylene glycol)-alpha methacrylate (PEG&#945;MA) was reacted off-stoichiometry with a cell-degradable crosslinker to form soft hydrogels. These soft hydrogels were later exposed to photoinitator and light to induce the homopolymerization of unreacted groups and stiffen the hydrogel. This protocol covers hydrogel synthesis, 3D bioprinting, photostiffening, and endpoint characterizations to assess fibroblast activation within 3D structures. The method presented here enables researchers to 3D bioprint a variety of materials that undergo pH-catalyzed polymerization reactions and could be implemented to engineer various models of tissue homeostasis, disease, and repair.

Author Spotlight: Understanding Chronic Lung Diseases Using 3D Printed Phototunable Hydrogels 

This article describes how to 3D bioprint phototunable hydrogels to study extracellular matrix stiffening and fibroblast activation.

Phototunable hydrogels can transform spatially and temporally in response to light exposure. Incorporating these types of biomaterials in cell-culture ...

3D Bioprinting for Rapid Vascular Network Fabrication Using Vessels-on-a-Plate

High-Throughput Bioprinting Method for Modeling Vascular Permeability in Standard Six-well Plates with Size and Pattern Flexibility

Vascular permeability is a key factor in developing therapies for disorders associated with compromised endothelium, such as endothelial dysfunction in coronary arteries and impaired function of the blood-brain barrier. Existing fabrication techniques do not adequately replicate the geometrical variation in vascular networks in the human body, which substantially influences disease progression; moreover, these techniques often involve multi-step fabrication procedures that hinder the high-throughput production necessary for pharmacological testing. This paper presents a bioprinting protocol for creating multiple vascular tissues with desired patterns and sizes directly on standard six-well plates, overcoming existing resolution and productivity challenges in bioprinting technology. A simplified fabrication approach was established to construct six hollow, perfusable channels within a hydrogel, which were subsequently lined with human umbilical vein endothelial cells to form a functional and mature endothelium. The computer-controlled nature of 3D bioprinting ensures high reproducibility and requires fewer manual fabrication steps than traditional methods. This highlights VOP's potential as an efficient high-throughput platform for modeling vascular permeability and advancing drug discovery.

Author Spotlight: Automated Bioprinting for High-Throughput Vascular Model Fabrication

We present a protocol for high-throughput production of vascular channels with flexible sizes and desired patterns on a standard six-well plate using 3D bioprinting technology, referred to as vessels-on-a-plate (VOP). This platform has the potential to advance the development of therapeutics for the disorders associated with compromised endothelium.

Vascular permeability is a key factor in developing therapies for disorders associated with compromised endothelium, such as endothelial dysfunction in ...

Combining 3D-Printing and Electrospinning to Manufacture Biomimetic Heart Valve Leaflets

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