The authors wish to thank the Veterinary Pathology Centre at the University of Surrey School of Veterinary Medicine for histology services. We also thank Drs L. Dixon, A. Reis, and M. Henstock from The Pirbright Institute (Pirbright, UK) for their support in procuring the animal tissues, and the Department of Biochemical Sciences at the University of Surrey, especially the technical team, for their continuing support. RSM was supported by the Doctoral College studentship award (University of Surrey), DM and PC were supported by the National Centre for the Replacement, Refinement &#38; Reduction of Animals in Research (grant numbers: NC/R001006/1 and NC/T001216/1).

This protocol describes the assembly and use of a system composed of two EasyFlow (bioreactor) inserts: one representing the reaction chamber (C), containing the perfused artery sample, and one functioning as a medium reservoir (R) (Figure 1 and Figure 2A). Carotid arteries were obtained from 4-6-week-old male and female piglets (6-12 kg) at The Pirbright Institute, UK. Animal procedures were carried out under the Home Office Animals (Scientific Procedures) Act (1986) (ASPA) and approved by the Animal Welfare and Ethical Review Board (AWERB) at The Pirbright Institute. The animals were housed in accordance with the Code of Practice for the Housing and Care of Animals Bred. All procedures were conducted by Personal License holders who were trained and competent under Project License PPL70/8852. Piglets were culled according to the schedule one method under the ASPA.
1. Insert manufacturing
<ol>
	<li>Fabricate the insert by 3D printing, using the provided 3D model (Supplementary File 1). 
	NOTE: 3D modeling enables easy changes to the design to adapt to new applications. Alternative materials and alternative manufacturing techniques can also be used to produce the insert. Due to the complex inner structure, selective laser sintering and stereolithography are appropriate alternatives15. Polyamide 12 (PA12; see Table of Materials) is a good material candidate due to its superior performance in terms of liquid retention and resistance to repeated heat sterilization cycles16.</li>
	<li>Fabricate the silicon gaskets and polycarbonate washers by laser cutting17, using the designs provided in Supplementary File 2. 
	&#8203;NOTE: Laser cutting is easily outsourced and a cheap manufacturing method. Washers can be manufactured from stainless steel, providing a more resistant component for repeated use. All other components are commercially available items. A complete list of the required materials is provided in Table 1. The commercial details of the items are included in the Table of Materials.</li>
</ol>
2. Device sterilization, assembly, and priming
<ol>
	<li>Sterilize all components following the instructions in Table 1.</li>
	<li>Under laminar flow conditions, assemble two fabricated inserts (step 1), as shown in Figure 1A.</li>
	<li>Assemble the perfusion system as in Figure 2, following the steps below:
	<ol>
		<li>Attach two connected three-way valves to the reservoir&#39;s R1 port (media exchange port).</li>
		<li>Connect the resulting outlet to the peristaltic pump&#39;s head using a tube equipped with a one-way valve (one-way valve tube).</li>
		<li>Connect the pump head to the reaction chamber&#39;s C4 port using system tubing equipped with a one-way valve. 
		NOTE: This branch can optionally be equipped with a pressure sensor enabling constant monitoring.</li>
		<li>Connect the reaction chamber port C1 to soft wall tubing, and this to the resistance channel (small inner diameter). 
		NOTE: The length of the resistance tubing will greatly influence the pressure present in the system. This should be further investigated to ensure adequate perfusion conditions.</li>
		<li>Extend the resistance channel with the system tubing, creating a return channel and closing the luminal circulation loop by connecting to the reservoir at R5 (Figure 2C).</li>
		<li>Create an overflow channel by connecting the reaction chamber C3 to the reservoir R3 with system tubing.</li>
		<li>Attach a ventilation filter through R2.</li>
		<li>Create a pressure dampener by connecting a syringe containing air and media to the reaction chamber C6. 
		NOTE: The correct air-to-media ratio will depend on the required pressure dampening.</li>
	</ol>
	</li>
	<li>Prime the system with perfusion medium (Dulbecco&#39;s modified eagle medium [DMEM] + 10% [v/v] fetal bovine serum [FBS] + 1% [v/v] penicillin-streptomycin + 1% [v/v] amphotericin B + 30% [v/v] dextran; see Table of Materials) through the media exchange ports and the reservoir. 
	&#8203;NOTE: Priming the system reduces the risk of trapped bubbles in the system and identifies any potential leaks. The recommended volume of media is about 100-120 mL. The volume used will depend on the dead volume of the tubing used for experimentation.</li>
</ol>
3. Sample harvesting and preparation
<ol>
	<li>Collect the left and right common carotid arteries, minimizing direct handling of the arterial tissue13.</li>
	<li>Place the tissue in cold transport media (DMEM+ 20% [v/v] FBS + 2% [v/v] penicillin-streptomycin + 1% [v/v] amphotericin B, see Table of Materials) for transfer.</li>
	<li>In a laminar flow cabinet, remove the excess connective tissue and trim the ends of the tissue using a scalpel blade. Wash the tissue twice in cold transport media.</li>
	<li>Place tissue on an orbital shaker in transport media for at least 30 min for thorough washing.</li>
	<li>Connect a non-branching segment of the artery to the bioreactor system using two barbed luer connectors and fix it in place using a vessel bond (vascular silicone ties; Table 1). 
	NOTE: The vessel bond provides the proper tension and retraction to secure the vessel. The oval section prevents tissue damage.</li>
	<li>Gently flow media through the artery to verify patency.</li>
	<li>After securing the artery to the fabricated insert, fill the reaction space with perfusion media (step 2.4). Finally, gently fill the luminal circulation loop with media to eliminate any remaining air from the system.</li>
	<li>Connect the reaction space with the previously assembled perfusion system (section 2), completing the circulation. 
	&#8203;NOTE: It is recommended to perform a quality check by immunohistochemistry of processed tissue. This identifies any damage due to excessive handling during preparation.</li>
</ol>
4. Perfusion culture and media change
<ol>
	<li>Place the perfusion system in a 37 &#176;C incubator with 5% CO2 and then connect it to a peristaltic pump (see Table of Materials)</li>
	<li>Connect any additional (facultative) acquisition systems, such as pressure sensors (see Table of Materials).</li>
	<li>Leave the system to equilibrate overnight with a low media flow rate of ~10-15 mL/min. 
	NOTE: Initial experiments to determine pump settings, media flow, system pressure, and optimal pressure dampeners/clamps settings are required to ensure appropriate conditions are met.</li>
	<li>The next day, increase the flow incrementally (+1 rpm every hour, equivalent to a ~2.5 mL/min increase every hour, in the current system) until the final flow rate (35 mL/min) is achieved. Monitor the system periodically for leaks. 
	NOTE: To calculate the accurate flowrate based on the peristaltic pump speed, users must first perform adequate pump calibration18. Using the formula (1), volumetric flow rate (Q) can be calculated based on the volume dispensed (V) within a given time (t). To calculate the estimated flow velocity (<img alt="Equation 3" src="/files/ftp_upload/65465/65465eq03.jpg" style="margin: auto;" />) we can use the previously calculated flow rate (Q) and the area of vessel cross-section (A), described in equation (2). 
	<img alt="Equation 1" src="/files/ftp_upload/65465/65465eq01.jpg" style="margin: auto;" />&#160; &#160; (1) 
	<img alt="Equation 2" src="/files/ftp_upload/65465/65465eq02.jpg" style="margin: auto;" />&#160; &#160; (2)</li>
	<li>Every 3 days, exchange 50% of the media (~50 mL) by connecting one syringe filled with fresh media to the media exchange port closer to the pump and an empty syringe to the port closer to the reservoir to collect the spent medium (Figure 2). 
	NOTE: Using two three-way valves as media exchange ports facilitates the continuous operation of the perfusion during the medium exchange.</li>
	<li>At the end of the experiment, harvest the tissue from the reaction chamber by trimming the ends connected to the bioreactor using sterile surgical scissors.</li>
	<li>Dismantle, clean, and sterilize the system for further use.</li>
</ol>
5. Sample analysis
<ol>
	<li>Fix the harvested sample in 4% paraformaldehyde (PFA) overnight at 4 &#176;C.</li>
	<li>Embed the tissue in optimal cutting temperature (OCT; see Table of Materials)19 and freeze by immersion in iso-pentane cooled in liquid nitrogen.</li>
	<li>Obtain cryo-sections of 3-5 mm thickness using a cryostat.</li>
	<li>Perform immunohistochemistry (haematoxylin and eosin [H&#38;E]) and/or immunofluorescence. Follow the typical immunofluorescence protocol:
	<ol>
		<li>Block the slices for 1 h at room temperature with 20% [v/v] goat serum in phosphate-buffered saline (PBS; see Table of Materials).</li>
		<li>Incubate the slices overnight at 4 &#176;C with primary rabbit antibody against CD31 diluted 1:50 in PBS (see Table of Materials).</li>
		<li>Wash three times with PBS for 5 min.</li>
		<li>Incubate for 1 h at 37 &#176;C with fluorescently labeled goat anti-rabbit secondary antibody diluted 1:200 in PBS and &#945;-smooth muscle actin (SMA) directly conjugated fluorescent antibody diluted 1:200 in PBS (see Table of Materials).</li>
		<li>Wash three times with PBS for 5 min.</li>
		<li>Stain the nuclei with DAPI diluted 1:1,000 in PBS for 10 min at room temperature.</li>
		<li>Incubate with 0.1% (w/v) Sudan Black (see Table of Materials) in 70% (v/v) ethanol for 10 min at room temperature to reduce tissue autofluorescence.</li>
		<li>Wash the tissue with abundant deionized water. Mount the slides in the mounting medium.</li>
		<li>Image the samples with a confocal laser scanning microscope.</li>
	</ol>
	</li>
</ol>

A Novel 3D Printed Bioreactor for Ex Vivo Culture of Blood Vessels in Perfusion

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The authors do not have any conflicts of interest to disclose.

Ex vivo vascular perfusion systems constitute a unique platform to study the function and behavior of vascular cells within their native tissues under controlled conditions, which enables the dissection of complex processes such as post-injury vascular remodeling22. However, most reported bioreactors are in-house made systems based on custom-made components and are often difficult to replicate by others23. Alternative commercial solutions exist, but lack flexibility in the design and can be relatively costly24.
We have developed an alternative system that provides an easy, cheap, and reproducible platform that can be manufactured using open-source 3D printing techniques13. The present article describes the system&#39;s setup to enable reproducible applications by end-users. This setup enables the application of physiological and pathological conditions of pressure (40-180 mmHg), flow rate (6-30 mL/min), and in combination with media mimicking the blood viscosity, with varying degrees of shear stress.
Reproducibility is an essential aspect of the scientific process, as it allows researchers to validate the findings of others and build upon them to progress our understanding of vascular diseases. Furthermore, tools that enable and promote collaborations between groups are essential for advancing scientific knowledge. EasyFlow represents an example of such open-source, accessible solutions that can be easily produced and adopted by labs working on a wide range of projects within the field of vascular sciences and beyond.
We report that this device culture maintains arterial tissue viability for at least 7 days, and can be used to model specific steps of vascular disease. Using this, one could model physiological flow rates and pressure conditions13. Importantly, this perfusion culture is cost-effective due to the low production costs and the low volume of medium necessary to operate the system.
The 3D design can also be adapted to new applications, and new materials for printing can be tested. Even in its current format, the sample lodging space can be easily adapted to samples of different sizes by changing the length of the fittings or the bore of the luer connectors. It is important to note that, given the modular nature of the device and its small dimensions, this bioreactor can be used in several setups (Figure 5) and can be applied to multiplex cultures, where several samples can be exposed to different conditions at the same time in separate bioreactors.
It is envisioned that the use of the system be expanded in the future to support the culture of blood vessels from different origins (e.g., different species) and of different natures (e.g., veins, lymphatic), and perhaps applied to the culture of other hollow tissues (e.g., trachea, intestine). In particular, research shows that culturing tissue-engineered scaffolds in constant perfusion helps the homogenous distribution of cells within the construct and maturation of the resulting tissue25,26. In addition, seeding vascular grafts in perfusion contributes to achieving a more uniformly cellularized vascular lumen, as compared to static methods27. For this reason, we envision the system being applied to tissue engineering to help address the current challenges, allowing for the future development of reproducible synthetic blood vessel substitutes28.
The protocol described here presents some important steps critical to the success of the flow culture. Establishing and monitoring appropriate flow conditions is not trivial and needs to be performed on each system when it is set up for the first time, to ensure that the culture conditions are physiological. The flow and pressure were monitored using pressure sensors and ultrasound imaging. Another critical point is ensuring that the tissue is viable and intact at the beginning of the culture. This requires a fresh source, careful handling and can be verified by histological analysis. Additionally, troubleshooting must be performed at the start of every experiment to identify any potential bacterial contamination or media leakage source.
It is important to highlight that the perfusion system described, while providing physiological pressure and flow conditions, is unable to entirely mimic the complex pressure wave patterns recorded in vivo. This limitation is ascribable to using a peristaltic pump and can be resolved using more specialized equipment to reproduce advanced hemodynamic conditions. The culture of blood vessels in a bioreactor is also unable to address studies where the immune system or the interaction with other organs is critical.
In conclusion, a simple 3D printed perfusion system that can mimic the physiological hemodynamic environment is presented, which is expected to contribute to the standardization of ex vivo blood vessel cultures. Its potential for customization and application to long-term culture makes it an essential tool for advancing the understanding of these complex biological systems in physiology and pathological conditions.

The vascular wall exists in a reactive steady state, which ensures both responsivities to external stimuli (i.e., change of pressure, vasoconstrictors) and a consistent non-activating surface preventing blood coagulation and inflammatory cell infiltration1. In response to aging- and lifestyle-dependent stimuli and upon direct damage, the vascular wall activates remodeling processes such as restenosis and atherosclerosis, which are known contributors to common cardiovascular diseases (CVDs), such as ischemic stroke and myocardial infarction2. While interventional approaches such as percutaneous revascularisation and stenting are available to tackle advanced manifestations of vascular disease, these are known to provoke further vascular damage, often leading to recurrence. In addition, only limited preventative and early-stage solutions are available. Understanding the mechanisms maintaining vascular wall homeostasis and driving its dysfunction is at the heart of developing new cures3.
Despite the constant development and advances in molecular biology and tissue engineering, animal studies remain a crucial component of vascular biology studies. In vivo animal studies have provided enormous insight into the mechanisms of vascular homeostasis and pathology; however, these procedures are costly, have relatively low throughput, and pose substantial ethical issues. In addition, small animals are poorly representative of human vascular physiology, and larger animal experiments are vastly more expensive and create further ethical considerations4,5. With the increasing demand for pharmaceutical and medical solutions for a rapidly aging population, the downsides of animal use are magnified, impacting the reproducibility, reliability, and transferability of results to patient care6.
In vitro systems offer a simplified platform to study basic mechanisms but fail to recapitulate the complexity of the whole tissue, the interactions between cells and the extracellular matrix, and the mechanical forces, which are critical determinants in the development of vascular diseases7.
Ex vivo studies performed on whole tissues maintained in artificially controlled environments mimic the in vivo complexity while enabling relatively high-throughput investigations8. Given the ability to closely control the culture conditions and environment, ex vivo models allow a broad range of complex studies and provide a suitable alternative to reduce the use of animal procedures in vascular biology. Static vascular ring cultures offered interesting insights but failed to incorporate the crucial hemodynamic element9. Indeed, the study of the vascular system ex vivo poses specific challenges related to the many dynamic forces which apply to the cells within the blood vessel wall. Stimuli such as luminal flow, turbulence, shear stress, pressure, and wall deformation significantly impact tissue pathophysiology10,11,12.
Perfusion bioreactors are essential for studying vascular homeostasis and remodeling in response to injury or hemodynamic changes13. Furthermore, perfusion culture can be used to improve the maturation and durability of tissue-engineered blood vessels (TEBVs), providing suitable alternatives for vascular grafts14.
Commercially available perfusion bioreactors are limited in flexibility and adaptability and are costly. Many of the existing in-house developed bioreactors are instead difficult to replicate in other labs, due to the limited descriptions and unavailability of specially made components7,8,9,10,11,12. To overcome these limitations, we have recently developed a new bioreactor (EasyFlow), which is economical to produce, accommodates a range of tissues, and enables relatively simple modifications to adapt to different research demands13. The insert is 3D printed and fits as in a lid of a standard 50 mL centrifuge tube. Its modular design and 3D printing manufacture make it accessible and reproducible across different labs, as well as easily modifiable to adapt to different scientific needs. This protocol describes the assembly and basic operation of the bioreactor system in an arterial perfusion setting.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>EasyFlow</td>
			<td>-</td>
			<td>-</td>
			<td>3D printed by MultiJet Fusion by Protolabs</td>
		</tr>
		<tr>
			<td>PA12 - 3D printing</td>
			<td>Protolabs</td>
			<td>-</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Peristaltic pump</td>
			<td>Heidolph</td>
			<td>&#160;PD5201</td>
			<td></td>
		</tr>
		<tr>
			<td>Culture media components:</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Amphotericin B solution, 250 mug/mL in deionized water</td>
			<td>Sigma-Aldrich</td>
			<td>A2942-20ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Dextran&#160; from Leuconostoc spp.</td>
			<td>Sigma-Aldrich</td>
			<td>D8802-25ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s Modified Eagle&#39;s Medium - high glucose, w/ 4500 mg/L glucose, L-glutamine, sodium pyruvate, and sodium bicarbonate</td>
			<td>Sigma-Aldrich</td>
			<td>D6429-6X500ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum</td>
			<td>Sigma-Aldrich</td>
			<td>F9665</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin</td>
			<td>Sigma-Aldrich</td>
			<td>P4333-100ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Immunostaining materials:</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Cryostat</td>
			<td>LEICA</td>
			<td>CM3050 S</td>
			<td></td>
		</tr>
		<tr>
			<td>DAPI</td>
			<td>Sigma-Aldrich</td>
			<td>D9542-10MG</td>
			<td></td>
		</tr>
		<tr>
			<td>Goat serum</td>
			<td>Sigma-Aldrich</td>
			<td>G9023-10ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Goat &#945;-Rabbit Alexa Fluor 488</td>
			<td>Thermo Fisher Scientific</td>
			<td>A11008</td>
			<td></td>
		</tr>
		<tr>
			<td>Invitrogen&#160;eBioscience Fluoromount G</td>
			<td>Thermo Fisher Scientific</td>
			<td>50-187-88</td>
			<td></td>
		</tr>
		<tr>
			<td>MX35 Premier + Microtome Blade</td>
			<td>Thermo Scientific</td>
			<td>3052835</td>
			<td></td>
		</tr>
		<tr>
			<td>Optimal Cooling Tempearure Compound - OCT</td>
			<td>Agar Scientific</td>
			<td>AGR1180</td>
			<td></td>
		</tr>
		<tr>
			<td>Rabbit &#945;-CD31 antibody</td>
			<td>Abcam</td>
			<td>ab28364</td>
			<td></td>
		</tr>
		<tr>
			<td>Sudan Black B</td>
			<td>Santa Cruz Biotechnology</td>
			<td>SC-203760</td>
			<td></td>
		</tr>
		<tr>
			<td>X72 SuperFrost Plus Adhesion slide, 25x75x1mm, White, 90&#176; Ground Edges, Frosted Area 20mm, 72/box</td>
			<td>Fisher Scientific</td>
			<td>J1800AMNZ</td>
			<td></td>
		</tr>
		<tr>
			<td>&#945;-Smooth Muscle Actin (SMA) Alexa Fluor&#174; 647-conjugated antibody</td>
			<td>R&#38;D Systems</td>
			<td>IC1420R</td>
			<td></td>
		</tr>
		<tr>
			<td>Material for laser cutting of components:</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Clear Plastic Sheet, 1250 mm x 610 mm x 1 mm (for laser cutting of&#160; washers)</td>
			<td>RS Components</td>
			<td>258-6590</td>
			<td></td>
		</tr>
		<tr>
			<td>RS PRO Translucent Rubber Sponge Sheet, 600 mm x 600 mm x 1.5 mm (for laser cutting of&#160; silicone seals)</td>
			<td>RS Components</td>
			<td>840-5541</td>
			<td></td>
		</tr>
		<tr>
			<td>Optional pressure monitors:</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Pressure sensor</td>
			<td>Parker Hannifin</td>
			<td>080-699PSX-3P-5</td>
			<td></td>
		</tr>
		<tr>
			<td>SciPres Pressure Monitor</td>
			<td>Parker Hannifin</td>
			<td>206-200-M</td>
			<td></td>
		</tr>
		<tr>
			<td>Pre-sterilized single use plasticware:</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>0.2 um filter</td>
			<td>Sarstedt</td>
			<td>70.1114.210</td>
			<td></td>
		</tr>
		<tr>
			<td>20 mL Sterile syringe</td>
			<td>IMS Euro</td>
			<td>40004</td>
			<td></td>
		</tr>
		<tr>
			<td>50 mL Centrifuge Tube</td>
			<td>Thermo Fisher Scientific</td>
			<td>Sarstedt - 62.547.254</td>
			<td></td>
		</tr>
		<tr>
			<td>Small components:</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Cable ties</td>
			<td>-</td>
			<td>-</td>
			<td></td>
		</tr>
		<tr>
			<td>Masterflex Adapter Fittings, Female Luer to Hose Barb</td>
			<td>Cole-Parmer</td>
			<td>WZ-30800-10</td>
			<td>Barb Adaptor</td>
		</tr>
		<tr>
			<td>Masterflex&#160;Polycarbonate Luer Fittings</td>
			<td>Cole-Parmer</td>
			<td>AU-45504-84</td>
			<td></td>
		</tr>
		<tr>
			<td>Nylon Miniature Check Valve</td>
			<td>Cole-Parmer</td>
			<td>98553-00</td>
			<td></td>
		</tr>
		<tr>
			<td>RS PRO Translucent Rubber Sponge Sheet, 600 mm x 600 mm x 1.5 mm (for laser cutting of&#160; silicone seals)</td>
			<td>RS Components</td>
			<td>840-5541</td>
			<td></td>
		</tr>
		<tr>
			<td>Stainless Steel M2 Hex Nuts</td>
			<td>RS Components</td>
			<td>527-218</td>
			<td></td>
		</tr>
		<tr>
			<td>Stainless Steel M2 x 6 mm Screws</td>
			<td>RS Components</td>
			<td>418-7426</td>
			<td></td>
		</tr>
		<tr>
			<td>Stainless Steel M5 Hex Nuts</td>
			<td>RS Components</td>
			<td>189-585</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical vessel loop</td>
			<td>Vascular Silicone Ties,International Medical Supplies</td>
			<td>&#160;10-1003</td>
			<td></td>
		</tr>
		<tr>
			<td>Three-way valves</td>
			<td>IMS Euro</td>
			<td>&#160;91000</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical Equipment</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Anatomical Forceps, GRAEFE, Curved, 10 cm SKU: BD-07</td>
			<td>International Medical Supplies</td>
			<td>SKU: BD-07</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro Forceps, Angled, 0.3 mm, 11 cm</td>
			<td>International Medical Supplies</td>
			<td>SKU: BD-361</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro Scissors Noyes, Curved, 12 cm</td>
			<td>International Medical Supplies</td>
			<td>SKU: FD-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Troge Surgical Scalpels - Size 23 - Box of 100</td>
			<td>International Medical Supplies</td>
			<td>63114</td>
			<td></td>
		</tr>
		<tr>
			<td>Tubing:</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Eppendorf silicone tubing (I.D.1.6 mm, O.D.4.7 mm)</td>
			<td>Eppendorf</td>
			<td>M0740-2396</td>
			<td>System tubing</td>
		</tr>
		<tr>
			<td>Masterflex&#160;PharMed BPT 3-Stop Tubing</td>
			<td>ISMATEC</td>
			<td>95714-48</td>
			<td>Soft wall tubing (for clamp)</td>
		</tr>
		<tr>
			<td>RS PRO Transparent Hose Pipe, 0.8 mm ID, Silicone</td>
			<td>RS Components</td>
			<td>667-8432</td>
			<td>Resistance tubing&#160;(small inner diameter)</td>
		</tr>
		<tr>
			<td>Tygon for food (I.D. 4.8 mm, W.T. 1.6 mm)</td>
			<td>Heidolph</td>
			<td>525-30027-00-0</td>
			<td>One way valve tube</td>
		</tr>
		<tr>
			<td>Verderflex Yellow Hose Pipe, 6.4 mm ID, Verderprene</td>
			<td>RS Components</td>
			<td>125-4042</td>
			<td>Pump Tubing</td>
		</tr>
	</tbody>
</table>

ex vivo perfusion culture of large blood vessels in a 3d printed bioreactor

Vascular disease forms the basis of most cardiovascular diseases (CVDs), which remain the primary cause of mortality and morbidity worldwide. Efficacious surgical and pharmacological interventions to prevent and treat vascular disease are urgently needed. In part, the shortage of translational models limits the understanding of the cellular and molecular processes involved in vascular disease. Ex vivo perfusion culture bioreactors provide an ideal platform for the study of large animal vessels (including humans) in a controlled dynamic environment, combining the ease of in vitro culture and the complexity of the live tissue. Most bioreactors are, however, custom manufactured and therefore difficult to adopt, limiting the reproducibility of the results. This paper presents a 3D printed system that can be easily produced and applied in any biological lab, and provides a detailed protocol for its setup, enabling users&#39; operation. This innovative and reproducible ex vivo perfusion culture system enables the culture of blood vessels for up to 7 days in physiological conditions. We expect that adopting a standardized perfusion bioreactor will support a better understanding of physiological and pathological processes in large animal blood vessels and accelerate the discovery of new therapeutics.

This study has established a versatile and affordable perfusion system (EasyFlow)13. The 3D printed design of the system facilitates the adoption of the system by other labs and therefore encourages reproducibility.
The fabricated perfusion insert is housed in a 50 mL centrifuge tube, creating an isolated environment. Using two perfusion inserts, a perfusion loop can be established containing a reservoir and a reaction chamber, where the biological sample is incubated. The perfusion system is then connected to a peristaltic pump and optional acquisition systems, such as pressure sensors and ultrasound devices, to monitor culture conditions (Figure 2).
Porcine carotid artery samples were collected and processed before being cultured in perfusion for 7 days. To ensure the quality of the tissue before culture, preliminary experiments were performed where the samples were fixed at the time of excision, after tissue preparation, and after perfusion. Fluorescent staining for endothelial (CD31) and smooth muscle (&#945;SMA) markers was used to assess the maintenance of tissue integrity. Examples of well-preserved and damaged tissues are presented in Figure 3 for comparison. The images show the importance of gentle handling of the tissue at the time of excision, as incorrect processing (excessive stretch, crushing, etc.) may result in loss of the endothelium ahead of the culture. Results also show the importance of the gradual establishment of perfusion to avoid luminal damage.
H&#38;E staining was performed alongside immunofluorescence (IF) staining to show maintenance of the morphology and overall distribution of the cells in the vessel wall after 7 days of culture (Figure 4). The application of physiological culture conditions in the device ensures maintenance of the endothelial coverage of the lumen, alignment of the smooth muscle cells in the media, and preservation of the vasa vasorum in the adventitia.
The compact and modular design of the bioreactor system also allows for a wide range of system setups. The smaller setup comprises a single bioreactor and is ideal for pharmacological and low-volume studies (recirculating perfusion20, total volume of 50-70 mL). To increase the length of culture and reduce the number of media changes, a single reservoir circulation system, like the one described in this protocol, or a constant feeding system is more ideal, as they have a larger volume of medium in circulation. A double circulation21 setup can be established to explore experimental settings where the localization of the stimuli is critical. The current device can also be combined with more sophisticated control systems to enable precise feedback control of parameters such as pH and dissolved oxygen (Figure 5).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65465/65465fig01.jpg" /> 
Figure 1: EasyFlow assembly schematics.&#160;(A) 3D rendered schematics aiding the assembly of the perfusion insert. (B) Detailed schematics are provided to facilitate assembly of the connection sites. (C) A cross-section view of the reaction space highlights the essential components of the EasyFlow insert and the connection of the tissue to the insert. <a href="https://www.jove.com/files/ftp_upload/65465/65465fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/65465/65465fig02.jpg" /> 
Figure 2: Schematic representation of the perfusion system assembly.&#160;(A) Assembled perfusion system containing all important components, highlighting their relative position in a 3D-rendered environment. Not all the components are to scale. (B) Individual isometric views of components are also presented. (C) A top view of the assembled perfusion system is shown to aid the assembly and connection of the different components. The ports have been labeled and numbered counterclockwise to navigate the various connection sites across the perfusion system. This principle has been applied to the reservoir (R) and reaction chamber (C). The various channels connecting the two chambers have also been assigned names for reference. <a href="https://www.jove.com/files/ftp_upload/65465/65465fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/65465/65465fig03.jpg" /> 
Figure 3: Tissue maintenance during processing. (A) Confocal images show the normal structure of an artery at the time of harvest, (B) after cleaning and processing, and (C) after perfusion culture, demonstrating the maintenance of a well-preserved morphology. (D) Examples of damaged tissue due to excessive or incorrect handling during processing or (E) due to the application of non-physiological culture conditions (i.e., abrupt initiation of high flow) show depletion of the luminal coverage and disruption of the media. (F) Negative control indicates the specificity of the staining. CD31: green; &#945;SMA: red; DAPI: blue. Scale bars: 100 &#181;m. <a href="https://www.jove.com/files/ftp_upload/65465/65465fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/65465/65465fig04.jpg" /> 
Figure 4: Histological evaluation of tissue before and after perfusion culture.&#160;(A and B) Arterial tissue was evaluated by histology at the time of harvest and (C and D) after 7 days of culture with the bioreactor system. (A and C) H&#38;E staining reveals preservation of the arterial wall structure and organization. (B and D) Immunofluorescence staining of the same tissue evidence endothelial coverage, smooth muscle cell alignment, and vasa vasorum in the adventitia. CD31: green; SMA: red; DAPI: blue. Scale bars: 100 &#181;m. <a href="https://www.jove.com/files/ftp_upload/65465/65465fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/65465/65465fig05.jpg" /> 
Figure 5: Schematic representation of possible perfusion setups. Various alternative setups can be accommodated with the perfusion insert to enable different experimental studies. (A) The recirculating perfusion setup minimizes the volume of medium required. (B) The constant feeding setup provides a steady supply of medium to the tissue. (C) Single reservoir circulation (as described in this paper) provides a larger volume of media for long-term incubations and includes a buffer zone for air exchange and pressure equilibration. (D) The double circulation setup provides two distinct loops feeding the inner (inside the artery) and outer (reaction space) circulations independently. (E) The dynamic perfusion setup includes continuous gasses and pH control by a proportional-integral-derivative (PID) controller. <a href="https://www.jove.com/files/ftp_upload/65465/65465fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Table 1: Components for fabricating the inserts. <a href="https://www.jove.com/files/ftp_upload/65465/65465_Table 1.xlsx" target="_blank">Please click here to download this Table.</a>
Supplementary File 1:&#160;3D model of the EasyFlow insert for 3D printing manufacture. <a href="https://www.jove.com/files/ftp_upload/65465/Supplementary File 1.stl" target="_blank">Please click here to download this File.</a>
Supplementary File 2: Design schematics of the constructed device, section view, printout, and sealing components. <a href="https://www.jove.com/files/ftp_upload/65465/Supplementary File 2.pdf" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about EasyFlow  An Economical and Adaptable Perfusion Bioreactor for Large Blood Vessel Culture at JoVE.com

Author Spotlight: EasyFlow - An Economical and Adaptable Perfusion Bioreactor for Large Blood Vessel Culture 

This protocol presents the setup and operation of a newly developed, 3D printed bioreactor for the ex vivo culture of blood vessels in perfusion. The system is designed to be easily adopted by other users, practical, affordable, and adaptable to different experimental applications, such as basic biology and pharmacological studies.

Ex Vivo Perfusion Culture of Large Blood Vessels in a 3D Printed Bioreactor

Vascular disease forms the basis of most cardiovascular diseases (CVDs), which remain the primary cause of mortality and morbidity worldwide. Efficacious ...

ex-vivo-perfusion-culture-large-blood-vessels-3d-printed

Research

JoVE Journal

Bioengineering

3.1K Views.  University of Surrey. This protocol presents the setup and operation of a newly developed, 3D printed bioreactor for the ex vivo culture of blood vessels in perfusion. The system is designed to be easily adopted by other users, practical, affordable, and adaptable to different experimental applications, such as basic biology and pharmacological studies. 

Video: Author Spotlight: EasyFlow - An Economical and Adaptable Perfusion Bioreactor for Large Blood Vessel Culture 

High Resolution 3D Imaging of Ex-Vivo Biological Samples by Micro CT

Non-destructive volume visualization can be achieved only by tomographic techniques, of which the most efficient is the x-ray micro computerized tomography (&mu;CT).
High resolution &mu;CT is a very versatile yet accurate (1-2 microns of resolution) technique for 3D examination of ex-vivo biological samples1, 2. As opposed to electron tomography, the &mu;CT allows the examination of up to 4 cm thick samples. This technique requires only few hours of measurement as compared to weeks in histology. In addition, &mu;CT does not rely on 2D stereologic models, thus it may complement and in some cases can even replace histological methods3, 4, which are both time consuming and destructive. Sample conditioning and positioning in &mu;CT is straightforward and does not require high vacuum or low temperatures, which may adversely affect the structure. The sample is positioned and rotated 180&deg; or 360&deg;between a microfocused x-ray source and a detector, which includes a scintillator and an accurate CCD camera, For each angle a 2D image is taken, and then the entire volume is reconstructed using one of the different available algorithms5-7. The 3D resolution increases with the decrease of the rotation step. The present video protocol shows the main steps in preparation, immobilization and positioning of the sample followed by imaging at high resolution.

High Resolution 3D Imaging of Ex-Vivo Biological Samples by Micro CT

Non-destructive volume visualization can be achieved only by tomographic techniques, of which the most efficient is the x-ray micro computerized tomography ( CT).

Non-destructive volume visualization can be achieved only by tomographic techniques, of which the most efficient is the x-ray micro computerized ...

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic valve leaflets are composed of interstitial cells suspended within an extracellular matrix (ECM) and are lined with an endothelial cell monolayer. The valve withstands a harsh, dynamic environment and is constantly exposed to shear, flexion, tension, and compression. Research has shown calcific lesions in diseased valves occur in areas of high mechanical stress as a result of endothelial disruption or interstitial matrix damage1-3. Hence, it is not surprising that epidemiological studies have shown high blood pressure to be a leading risk factor in the onset of aortic valve disease4. 
The only treatment option currently available for valve disease is surgical replacement of the diseased valve with a bioprosthetic or mechanical valve5. Improved understanding of valve biology in response to physical stresses would help elucidate the mechanisms of valve pathogenesis. In turn, this could help in the development of non-invasive therapies such as pharmaceutical intervention or prevention. Several bioreactors have been previously developed to study the mechanobiology of native or engineered heart valves6-9. Pulsatile bioreactors have also been developed to study a range of tissues including cartilage10, bone11 and bladder12. The aim of this work was to develop a cyclic pressure system that could be used to elucidate the biological response of aortic valve leaflets to increased pressure loads. 
The system consisted of an acrylic chamber in which to place samples and produce cyclic pressure, viton diaphragm solenoid valves to control the timing of the pressure cycle, and a computer to control electrical devices. The pressure was monitored using a pressure transducer, and the signal was conditioned using a load cell conditioner. A LabVIEW program regulated the pressure using an analog device to pump compressed air into the system at the appropriate rate. The system mimicked the dynamic transvalvular pressure levels associated with the aortic valve; a saw tooth wave produced a gradual increase in pressure, typical of the transvalvular pressure gradient that is present across the valve during diastole, followed by a sharp pressure drop depicting valve opening in systole. The LabVIEW program allowed users to control the magnitude and frequency of cyclic pressure. The system was able to subject tissue samples to physiological and pathological pressure conditions. This device can be used to increase our understanding of how heart valves respond to changes in the local mechanical environment.

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

A cyclic pressure bioreactor capable of subjecting heart valve tissue to physiological and pathological pressure conditions has been designed. A LabVIEW program allows users to control pressure magnitude, amplitude and frequency. This device can be used to study the mechanobiology of heart valve tissue or isolated cells.

The aortic valve, located between the left ventricle and the aorta, allows for unidirectional blood flow, preventing backflow into the ventricle. Aortic ...

Nonhuman Primate Lung Decellularization and Recellularization Using a Specialized Large-organ Bioreactor

There are an insufficient number of lungs available to meet current and future organ transplantation needs. Bioartificial tissue regeneration is an attractive alternative to classic organ transplantation. This technology utilizes an organ&#39;s natural biological extracellular matrix (ECM) as a scaffold onto which autologous or stem/progenitor cells may be seeded and cultured in such a way that facilitates regeneration of the original tissue. The natural ECM is isolated by a process called decellularization. Decellularization is accomplished by treating tissues with a series of detergents, salts, and enzymes to achieve effective removal of cellular material while leaving the ECM intact. Studies conducted utilizing decellularization and subsequent recellularization of rodent lungs demonstrated marginal success in generating pulmonary-like tissue which is capable of gas exchange in vivo. While offering essential proof-of-concept, rodent models are not directly translatable to human use. Nonhuman primates (NHP) offer a more suitable model in which to investigate the use of bioartificial organ production for eventual clinical use.
The protocols for achieving complete decellularization of lungs acquired from the NHP rhesus macaque are presented. The resulting acellular lungs can be seeded with a variety of cells including mesenchymal stem cells and endothelial cells. The manuscript also describes the development of a bioreactor system in which cell-seeded macaque lungs can be cultured under conditions of mechanical stretch and strain provided by negative pressure ventilation as well as pulsatile perfusion through the vasculature; these forces are known to direct differentiation along pulmonary and endothelial lineages, respectively. Representative results of decellularization and cell seeding are provided.

Whole-organ decellularization produces natural biological scaffolds that may be used for regenerative medicine. The description of a nonhuman primate model of lung regeneration in which whole lungs are decellularized and then seeded with adult stem cells and endothelial cells in a bioreactor that facilitates vascular circulation and liquid media ventilation is presented.

There are an insufficient number of lungs available to meet current and future organ transplantation needs. Bioartificial tissue regeneration is an ...

Fabrication of Custom Agarose Wells for Cell Seeding and Tissue Ring Self-assembly Using 3D-Printed Molds

Engineered tissues are being used clinically for tissue repair and replacement, and are being developed as tools for drug screening and human disease modeling. Self-assembled tissues offer advantages over scaffold-based tissue engineering, such as enhanced matrix deposition, strength, and function. However, there are few available methods for fabricating 3D tissues without seeding cells on or within a supporting scaffold. Previously, we developed a system for fabricating self-assembled tissue rings by seeding cells into non-adhesive agarose wells. A polydimethylsiloxane (PDMS) negative was first cast in a machined polycarbonate mold, and then agarose was gelled in the PDMS negative to create ring-shaped cell seeding wells. However, the versatility of this approach was limited by the resolution of the tools available for machining the polycarbonate mold. Here, we demonstrate that 3D-printed plastic can be used as an alternative to machined polycarbonate for fabricating PDMS negatives. The 3D-printed mold and revised mold design is simpler to use, inexpensive to produce, and requires significantly less agarose and PDMS per cell seeding well. We have demonstrated that the resulting agarose wells can be used to create self-assembled tissue rings with customized diameters from a variety of different cell types. Rings can then be used for mechanical, functional, and histological analysis, or for fabricating larger and more complex tubular tissues.

This protocol describes a platform for fabricating self-assembled tissue rings in variable sizes using a customized 3D-printed plastic mold. PDMS negatives are cured in the 3D-printed mold; then agarose is cast in the cured PDMS negatives. Cells are seeded into the resulting agarose wells where they aggregate into tissue rings.

Engineered tissues are being used clinically for tissue repair and replacement, and are being developed as tools for drug screening and human disease ...

Rigid Embedding of Fixed and Stained, Whole, Millimeter-Scale Specimens for Section-free 3D Histology by Micro-Computed Tomography

For over a hundred years, the histological study of tissues has been the gold standard for medical diagnosis because histology allows all cell types in every tissue to be identified and characterized. Our laboratory is actively working to make technological advances in X-ray micro-computed tomography (micro-CT) that will bring the diagnostic power of histology to the study of full tissue volumes at cellular resolution (i.e., an X-ray Histo-tomography modality). Toward this end, we have made targeted improvements to the sample preparation pipeline. One key optimization, and the focus of the present work, is a straightforward method for rigid embedding of fixed and stained millimeter-scale samples. Many of the published methods for sample immobilization and correlative micro-CT imaging rely on placing the samples in paraffin wax, agarose, or liquids such as alcohol. Our approach extends this work with custom procedures and the design of a 3-dimensional printable apparatus to embed the samples in an acrylic resin directly into polyimide tubing, which is relatively transparent to X-rays. Herein, sample preparation procedures are described for the samples from 0.5 to 10 mm in diameter, which would be suitable for whole zebrafish larvae and juveniles, or other animals and tissue samples of similar dimensions. As proof of concept, we have embedded the specimens from Danio, Drosophila, Daphnia, and a mouse embryo; representative images from 3-dimensional scans for three of these samples are shown. Importantly, our methodology leads to multiple benefits including rigid immobilization, long-term preservation of laboriously-created resources, and the ability to re-interrogate samples.

We developed protocols and designed a custom apparatus to enable embedding of millimeter-scale specimens. We present sample preparation procedures with an emphasis on embedding in acrylic resin and polyimide tubing to achieve rigid immobilization and long-term storage of specimens for the interrogation of tissue architecture and cell morphology by micro-CT.

For over a hundred years, the histological study of tissues has been the gold standard for medical diagnosis because histology allows all cell types in ...

3D Printed Porous Cellulose Nanocomposite Hydrogel Scaffolds

This work demonstrates the use of three-dimensional (3D) printing to produce porous cubic scaffolds using cellulose nanocomposite hydrogel ink, with controlled pore structure and mechanical properties. Cellulose nanocrystals (CNCs, 69.62 wt%) based hydrogel ink with matrix (sodium alginate and gelatin) was developed and 3D printed into scaffolds with uniform and gradient pore structure (110-1,100 &#181;m). The scaffolds showed compression modulus in the range of 0.20-0.45 MPa when tested in simulated in vivo conditions (in distilled water at 37 &#176;C). The pore sizes and the compression modulus of the 3D scaffolds matched with the requirements needed for cartilage regeneration applications. This work demonstrates that the consistency of the ink can be controlled by the concentration of the precursors and porosity can be controlled by the 3D printing process and both of these factors in return defines the mechanical properties of the 3D printed porous hydrogel scaffold. This process method can therefore be used to fabricate structurally and compositionally customized scaffolds according to the specific needs of patients.&#160;

The three critical steps of this protocol are i) developing the right composition and consistency of the cellulose hydrogel ink, ii) 3D printing of scaffolds into various pore structures with good shape fidelity and dimensions and iii) demonstration of the mechanical properties in simulated body conditions for cartilage regeneration.

This work demonstrates the use of three-dimensional (3D) printing to produce porous cubic scaffolds using cellulose nanocomposite hydrogel ink, with ...

Labeling of Blood Vessels in the Teleost Brain and Pituitary Using Cardiac Perfusion with a DiI-fixative

Blood vessels innervate all tissues in vertebrates, enabling their survival by providing the necessary nutrients, oxygen, and hormonal signals. It is one of the first organs to start functioning during development. Mechanisms of blood vessel formation have become a subject of high scientific and clinical interest. In adults however, it is difficult to visualize the vasculature in most living animals due to their localization deep within other tissues. Nevertheless, visualization of blood vessels remains important for several studies such as endocrinology and neurobiology. While several transgenic lines have been developed in zebrafish, with blood vessels directly visualized through expression of fluorescent proteins, no such tools exist for other teleost species. Using medaka (Oryzias latipes) as a model, the current protocol presents a quick and direct technique to label blood vessels in brain and pituitary by perfusing through the heart with fixative containing DiI. This protocol allows improvement of our understanding on how brain and pituitary cells interact with blood vasculature in whole tissue or thick tissue slices.

The article describes a quick protocol for labeling blood vessels in a teleost fish by cardiac perfusion of DiI diluted in fixative, using medaka (Oryzias latipes) as a model and focusing on brain and pituitary tissue.

Blood vessels innervate all tissues in vertebrates, enabling their survival by providing the necessary nutrients, oxygen, and hormonal signals. It is one ...

A Human 3D Extracellular Matrix-Adipocyte Culture Model for Studying Matrix-Cell Metabolic Crosstalk

The extracellular matrix (ECM) plays a central role in regulating tissue homeostasis, engaging in crosstalk with cells and regulating multiple aspects of cellular function. The ECM plays a particularly important role in adipose tissue function in obesity, and alterations in adipose tissue ECM deposition and composition are associated with metabolic disease in mice and humans. Tractable in vitro models that permit dissection of the roles of the ECM and cells in contributing to global tissue phenotype are sparse. We describe a novel 3D in vitro model of human ECM-adipocyte culture that permits study of the specific roles of the ECM and adipocytes in regulating adipose tissue metabolic phenotype. Human adipose tissue is decellularized to isolate ECM, which is subsequently repopulated with preadipocytes that are then differentiated within the ECM into mature adipocytes. This method creates ECM-adipocyte constructs that are metabolically active and retain characteristics of the tissues and patients from which they are derived. We have used this system to demonstrate disease-specific ECM-adipocyte crosstalk in human adipose tissue. This culture model provides a tool for dissecting the roles of the ECM and adipocytes in contributing to global adipose tissue metabolic phenotype and permits study of the role of the ECM in regulating adipose tissue homeostasis.

We describe a 3D human extracellular matrix-adipocyte in vitro culture system that permits dissection of the roles of the matrix and adipocytes in contributing to adipose tissue metabolic phenotype.

The extracellular matrix (ECM) plays a central role in regulating tissue homeostasis, engaging in crosstalk with cells and regulating multiple aspects of ...

Evaluating Regional Pulmonary Deposition using Patient-Specific 3D Printed Lung Models

Development of targeted therapies for pulmonary diseases is limited by the availability of preclinical testing methods with the ability to predict regional aerosol delivery. Leveraging 3D printing to generate patient-specific lung models, we outline the design of a high-throughput, in vitro experimental setup for quantifying lobular pulmonary deposition. This system is made with a combination of commercially available and 3D printed components and allows the flow rate through each lobe of the lung to be independently controlled. Delivery of fluorescent aerosols to each lobe is measured using fluorescence microscopy. This protocol has the potential to promote the growth of personalized medicine for respiratory diseases through its ability to model a wide range of patient demographics and disease states. Both the geometry of the 3D printed lung model and the air flow profile setting can be easily modulated to reflect clinical data for patients with varying age, race, and gender. Clinically relevant drug delivery devices, such as the endotracheal tube shown here, can be incorporated into the testing setup to more accurately predict a device&#8217;s capacity to target therapeutic delivery to a diseased region of the lung. The versatility of this experimental setup allows it to be customized to reflect a multitude of inhalation conditions, enhancing the rigor of preclinical therapeutic testing.

We present a high-throughput, in vitro method for quantifying regional pulmonary deposition at the lobe level using CT scan-derived, 3D printed lung models with tunable air flow profiles.

Development of targeted therapies for pulmonary diseases is limited by the availability of preclinical testing methods with the ability to predict ...

Stepwise Cell Seeding on Tessellated Scaffolds to Study Sprouting Blood Vessels

The cardiovascular system is a key player in human physiology, providing nourishment to most tissues in the body; vessels are present in different sizes, structures, phenotypes, and performance depending on each specific perfused tissue. The field of tissue engineering, which aims to repair or replace damaged or missing body tissues, relies on controlled angiogenesis to create a proper vascularization within the engineered tissues. Without a vascular system, thick engineered constructs cannot be sufficiently nourished, which may result in cell death, poor engraftment, and ultimately failure. Thus, understanding and controlling the behavior of engineered blood vessels is an outstanding challenge in the field. This work presents a high-throughput system that allows for the creation of organized and repeatable vessel networks for studying vessel behavior in a 3D scaffold environment. This two-step seeding protocol shows that vessels within the system react to the scaffold topography, presenting distinctive sprouting behaviors depending on the compartment geometry in which the vessels reside. The obtained results and understanding from this high throughput system can be applied in order to inform better 3D bioprinted scaffold construct designs, wherein fabrication of various 3D geometries cannot be rapidly assessed when using 3D printing as the basis for cellularized biological environments. Furthermore, the understanding from this high throughput system may be utilized for the improvement of rapid drug screening, the rapid development of co-cultures models, and the investigation of mechanical stimuli on blood vessel formation to deepen the knowledge of the vascular system.

Engineered tissues heavily rely on proper vascular networks to provide vital nutrients and gases and remove metabolic waste. In this work, a stepwise seeding protocol of endothelial cells and support cells creates highly organized vascular networks in a high-throughput platform for studying developing vessel behavior in a controlled 3D environment.

The cardiovascular system is a key player in human physiology, providing nourishment to most tissues in the body; vessels are present in different sizes, ...

Ex Vivo Perfusion Culture of Large Blood Vessels in a 3D Printed Bioreactor

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High Resolution 3D Imaging of Ex-Vivo Biological Samples by Micro CT

Design of a Cyclic Pressure Bioreactor for the Ex Vivo Study of Aortic Heart Valves

Nonhuman Primate Lung Decellularization and Recellularization Using a Specialized Large-organ Bioreactor

Fabrication of Custom Agarose Wells for Cell Seeding and Tissue Ring Self-assembly Using 3D-Printed Molds

Rigid Embedding of Fixed and Stained, Whole, Millimeter-Scale Specimens for Section-free 3D Histology by Micro-Computed Tomography

3D Printed Porous Cellulose Nanocomposite Hydrogel Scaffolds

Labeling of Blood Vessels in the Teleost Brain and Pituitary Using Cardiac Perfusion with a DiI-fixative

A Human 3D Extracellular Matrix-Adipocyte Culture Model for Studying Matrix-Cell Metabolic Crosstalk

Evaluating Regional Pulmonary Deposition using Patient-Specific 3D Printed Lung Models

Stepwise Cell Seeding on Tessellated Scaffolds to Study Sprouting Blood Vessels