The author thanks Dr. Jin Gao for PGS synthesis, Dr. Peter Crapo for insightful discussion for bioreactor setup, Drs. Mohamed Ezzelarab and Wei Wu for explanting baboon carotid arteries. This study was supported by a grant from the National Institutes of Health (R01 HL089658).

1. Tubular Scaffold Fabrication
<ol>
<li>Prepare glass tubing (length = 70 mm, outer diameter = 9.0 mm, wall thickness = 1.0 mm; Small Parts, Miramar, FL) as a mold for scaffold.</li>
<li>Prepare 1.0 wt /vol % hyaluronic acid (HA) (Sigma-Aldrich, St. Louis, MO) by dissolving 100 mg of HA into 10 ml of deionized water. Mix using a rotator overnight until there are no air bubbles in solution.</li>
<li>Pour 1.0 % HA solution into the top of glass tubing. Let it flow slowly down along the inner wall of the mold. Flip over the tubing when the solution reached the bottom of the mold using the rotator and repeat this step until the inner wall of the mold is evenly coated by the solution.</li>
<li>After coating, dry all glass tubings into the vacuum oven at 37&deg;C for 24 h.</li>
<li>Grind and sieve salt particles (size = 25-32 &mu;m) used as porogens.</li>
<li>Assemble a glass tubing (coated with 1.0 % HA solution inside), mandrel (stainless steel encased by PTFE tubing), heat-shrinkable (HS) sleeve, and PTFE ring as described previously8.</li>
<li>Add salt particles of the desired size to the glass mold using a spatula through the silicon rubber funnel. Tap the mold gently to ensure even distribution of salt particles. Scrape off the excess salt particles across the top of the mold using the side of spatula.</li>
<li>Turn on the hybridization incubator for 30 min before transferring the salt-packed mold. Turn off the hybridization incubator and Put the salt-packed mold into the hybridization incubator for salt fusion.</li> 
<li>Remove the molds from the hybridization incubator and dry them into the vacuum oven at 37&deg;C for 24 h.</li>
<li>Remove the salt-packed molds from the vacuum oven and allow them to cool at room temperature. Remove the stainless steel mandrel by pushing it down with holding the PTFE ring. If the mandrel is too tight to the mold, use the needle pliers to pull it out. Remove the PTFE ring from the bottom of the mold.</li>
<li>Put the molds into the oven at 120 &deg;C for 5 min to shrink HS sleeve. Check if the HS sleeve is shrunk. Remove the HS sleeve from the salt-packed molds and allow them to cool at room temperature. Keep the molds into the dessicator until they are used.</li>
<li>Prepare the PGS solution by dissolving 3 g of PGS into 15 ml of tetrahydrofuran (THF) (Sigma-Aldrich, St. Louis, MO) to form a 20 wt/vol % solution.</li>
<li>Add the PGS solution to the lumen of salt-packed molds by dropping it using a spoid in the hume hood. Lean the glass mold about 45&deg; and drop the PGS solution into inner lumen with rotating the mold slowly. Check if the PGS solution flows down along the wall of mold. If dry spot is observed, add more solution.</li> 
<li>After adding the PGS solution, place the molds in the hume hood for at least 30 min for evaporating THF.</li> 
<li>Put the molds into the vacuum oven for pre-determined curing time (ex. 24/ 36/ 48 h) at 100 mTorr and 150&deg;C for curing PGS.</li>
<li>After curing, take out the molds and place them at room temperature to cool down. Dip each mold into the deionized water with vertical position slowly. Don't dip them into the water fast because air bubbles might cause tear the scaffold.</li>
<li>Transfer the molds to the water bath carefully and place them at an angle for 1 h using silicon tube to dissolve HA easily. Check if HA is dissolved from the mold. If no HA is released from the mold, push the one end of the scaffold slowly using spatula to release it from the mold. Repeat this step at the other end of the scaffold and shake the mold back and forth slowly in the water bath. Check if the scaffold is moved inside the mold.</li>
<li>Transfer the scaffold to another water bath with stirrer carefully. Don&#x2019;t grab the scaffold firmly, which might crack the scaffold. Place the scaffold in the water bath for at least 3 days with changing water to leach out salt particles.</li>
<li>After leaching out salt particles, transfer each scaffold to the 15 ml centrifuge tube filled with the deionized water and place them in the dry-ice box for 1 h to freeze. Put centrifuge tubes into the lyophilizer (all tube cabs must be opened) and lyophilize them for 2 days.</li>
<li>Put all scaffolds into the dessicator before until they are used.</li>
</ol>
2. Scaffold Preparation for Cell Seeding
<ol>
<li>Take out scaffolds from the dessicator and cut them as 25-30mm of length.</li> 
<li>Prepare two silicone rubber stoppers (diameter = 20 mm, wall thickness = 6.35 mm, middle hole diameter = 3 mm) and connect two PTFE tubings (outer diameter = 3.97 mm, inner diameter = 2.38 mm, wall thickness = 0.79 mm, length = 60 mm and 40 mm) through the middle holes of each stopper.</li>
<li>Cut a HS ring as a 1.5 mm of length and put it on top of one end of the scaffold. Push one PTFE tubing (connected silicone rubber stopper) into the one end of scaffold as small overlapping portion as possible.</li>
<li>Put the scaffold and PTFE tubing into the oven at 120 &deg;C for 10 min to shrink HS ring. Check if HS ring is shrunk and scaffolds are connected to PTFE tubing firmly.</li>
<li>Take out the scaffold and PTFE tubing and place them at room temperature to cool down. Put the scaffold-PTFE tubing assembly into the polycarbonate tube (outer diameter = 15.5 mm, inner diameter = 9 mm, length = 50 mm) as a bioreactor chamber.</li> 
<li>Connect another end of PTFE tubing to the scaffold as steps 2.3) and 2.4).</li> 
<li>Adjust two silicone rubber stoppers to attach their inner surfaces to both ends of the polycarbonate tube. Attach both outer surfaces of silicone rubber stoppers to two aluminum alloy plates (top and bottom; width = 20 mm, length = 70 mm, thickness = 6.35 mm). Put two threaded rods into the side hole of plate and fix the polycarbonate tube (included the scaffold inside) to plates by tightening thumb nut screw on the plate.</li>
<li>Measure the seedable length of each scaffold (a length between two HS rings) and calculate inner surface area (&pi; x inner diameter x seedable length) of each scaffold for cell seeding.</li> 
<li>Wrap one bioreactor chamber unit with aluminum foil and sterilize it by autoclaving at 120 &deg;C for 30 min. Sterilize each part (medium reservoir, pump tubing, gas exchanger, manifolds, and needle valve) of the bioreactor by autoclaving at 120 &deg;C for 30 min and assemble them inside the cell culture hood.</li>
<li>Pre-treat and rinse scaffolds with a series of perfusions (70, 50, and 25 % ethanol for 1 h, phosphate-buffered saline (PBS; Mediatech, Herndon, VA) for 2 h, and SMC culture medium for 24 h by using a peristaltic pump at 1.0 ml/min in the flow circuit.</li>
</ol>
3. Cell Seeding and Culture 
<ol>
<li>Isolate primary arterial smooth muscle cells (SMCs) from the carotid arteries of juvenile male baboons (Papio Anubis) and characterize them in two-dimensional (2D) culture before passaging and seeding as described previously9.</li>
<li>Expand SMCs using the MCDB 131 medium (Mediatech, Herndon, VA) with 10% fetal bovine serum (Lonza, Walkersville, MD), 1 % L-glutamine (Mediatech), 50 &mu;g/ml ascorbic acid (Fisher Scientific, Fair Lawn, NJ), and an antibiotic-antimycotic solution (10,000 IU/ml penicillin, 10,000 &mu;g/ml streptomycin, and 25 &mu;g/ml amphotericin B; Mediatech).</li>
<li>Detach each bioreactor chamber from the flow circuit in the bioreactor. Seed SMCs (passage 4-6) to the scaffold with a density of 2x106 cells/cm2 by injecting suspension with 5 ml syringe into the lumen of scaffolds after cutting off both abluminal (inlet) and luminal (outlet) flows.</li> 
<li>Put all bioreactor chambers into the hybridization incubator at 37 &deg;C and rotate them at 2 rpm for 4 h to allow the cells to uniformly spread into the scaffold.</li>
<li>Take out bioreactor chambers from the hybridization incubator reattach them to flow circuit of the bioreactor in the hood. Transfer all bioreactor system into cell culture incubator.</li>
<li>Connect the pump tubing (inner diameter = 1.6 mm) to the cartilage of peristaltic pump and load fresh culture medium in the reservoir. Start the through-wall perfusion (luminal to abluminal) at 1.0 ml/min for 15 min followed by luminal flow only.</li> 
<li>Increase the flow rate and pressure gradually from 1.0 ml/min (luminal shear stress: 1.1 dynes/cm2) and 20 mmHg on day 1 to 14.2 ml/min (15.3 dynes/cm2) and 120 mmHg on day 21, respectively. Change the pump tubing (inner diameter = 3.2 mm) at day 4 and the medium once every week.</li>
</ol>
4. Tissue Harvesting and Sample Preparation for Analysis
<ol>
<li>After 21-day culture, stop the pump and clamp tubings (inlet and outlet) of the medium reservoir. Detach the pump tubing from the cartilage of peristaltic pump and transfer bioreactor system into the hood.</li>
<li>Clamp inlet and outlet tubings of each bioreactor chamber and detach it from the flow loop. Open the inlet tubing and prepare a petri-dish at the outlet tubing to collect culture medium inside the polycarbonate tube. Open the outlet tubing slowly and remove the medium from the chamber.</li>
<li>Detach abluminal gauge needle and luminal silicone tubing from the chamber and detach aluminum plates by releasing thumb screws. Remove the polycarbonate tube and detach one end of the tissue construct from the PTFE tubing.</li>
<li>Prepare a petri-dish filled with 10 ml of PBS and detach other end of the construct from the PTFE tubing. Put construct into the PBS and rinse three times.</li>
<li>Cut the construct with a razor blade as 5-mm of length. Freeze it at -80 &deg;C freezer for biochemical assay or transfer it to 15 ml centrifuge tube filled with 5 ml of fresh medium for mechanical testing or snap freeze into the Tissue-Tek optimal cutting temperature compound (Sakura Finetek Inc., Torrance, CA) for histology/immunofluorescence staining.</li> 
</ol>
5. Representative Results:
The tubular PGS scaffolds were fabricated using biodegradable elastomer by salt fusion method (Fig. 1A). Each bioreactor chamber provided scaffolds with both luminal and abluminal flows and could be detached as a separate unit from a main flow loop (Fig. 1B). Bioreactor system was designed for culturing four scaffolds at a time by controlling and monitoring flow as well as pressure (Fig. 1C &amp; D). 
Schematic of bioreactor culture was shown in Fig. 2. After seeding SMCs, each bioreactor chamber was rotated at 37 &deg;C for 4 h to distribute cells uniformly in the lumen of scaffold. And then, pulsatile flow was applied to scaffolds until day 14 with gradually increasing flow rate (Fig. 2A) and pressure (Fig. 2B). After day 14, flow and pressure kept constant until the end of culture (day 21).
Surface morphology of the PGS scaffold was examined by scanning electron microscopy (SEM). Scanning electron micrographs showed that scaffold had consistent wall thicknesses (539 &plusmn; 18 &mu;m) (Fig. 3A) and randomly distributed macro- and micro-pores on the luminal surface (Fig. 3B). Morphometric parameters of the scaffold was measured from microcomputed tomography (micro-CT) and imaging analysis. Average pore size is 23.3 &plusmn; 3.9 &#x03BC;m and pore interconnectivity is 99.4 &plusmn; 0.62 %, which means that all pores are fully interconnected in scaffold. Porosity measured by ethanol displacement is 75.6 &plusmn; 2.7 %.
Cellular morphology of the PGS construct was examined by SEM (Fig. 4). Multilayered SMCs completely covered the luminal surface and they were perpendicularly oriented to flow direction. These results show that mechanical conditioning from pulsatile flow bioreactor supports SMC orientation in the scaffold. 
The presence of extracellular matrix (ECM) and elastic fibers were examined by H&amp;E staining and elastin autofluorescence (Fig. 5). H&amp;E staining demonstrated that cells and ECM proteins completely covered the lumen of the PGS construct. Elastin autofluorescence also showed circumferentially organized elastic fibers at the luminal surface of the construct. Production of ECM proteins in PGS constructs were measured from biochemical assays. The insoluble elastin and collagen contents were 20.2 &plusmn; 9.1 &mu;g/mg of tissue and 6.3 &plusmn; 1.9 &mu;g/mg of tissue, respectively.
<img src="/files/ftp_upload/2691/2691fig1.jpg" alt="Figure 1"/> 
Figure 1. Scaffold fabrication and bioreactor system. (A) Schematic of tubular scaffold fabrication. (B) Bioreactor chamber. Scaffolds were connected to PTFE tubing, placed into the polycarbonate tube, and fixed by silicone rubber stoppers and aluminum alloy plates. Each chamber has two flow paths: luminal (by silicone tubing) and abluminal (by gauge needle) flows. (C) Bioreactor system placed inside the incubator. It includes medium reservoir, peristaltic pump module, gas exchanger, pressure transducers, two manifolds (top and bottom), and needle valve. (D) Bioreactor system placed outside the incubator. It includes pressure monitor, flow control unit, data acquisition system, and computer.
<img src="/files/ftp_upload/2691/2691fig2.jpg" alt="Figure 2"/> 
Figure 2. Schematic of the bioreactor culture. (A) Culture protocol. (B) Applied pressure profiles at each time point (Day 1, 4, 7, and 14). 
<img src="/files/ftp_upload/2691/2691fig3.jpg" alt="Figure 3"/> 
Figure 3. Surface morphology of the PGS scaffold. (A) Cross-section. (B) Lumen. 
<img src="/files/ftp_upload/2691/2691fig4.jpg" alt="Figure 4"/> 
Figure 4. Cellular morphology of the PGS construct. (A) Lumen. (B) Cross-section of 45&deg; cut. Arrows in both figures represent the flow direction.
<img src="/files/ftp_upload/2691/2691fig5.jpg" alt="Figure 5"/> 
Figure 5. Histology and elastin autofluorescence of the PGS construct. (A) H&amp;E staining. (B) Corresponding elastin autofluorescence. L: lumen. Magnification: 40X. Scale bar: 50 &mu;m.

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No conflicts of interest declared.

The fabrication technique using a biodegradable elastomer described here has several features. (1) We used hyaluronic acid (HA) as a mold release. Since HA is water soluble, scaffold was easily released from the glass mold after soaking it into water. In this report, we used 1.0 wt/vol % of HA solution because low concentration (&lt; 0.5 wt/vol %) of solution is not viscous and flows down so fast when we pouring it on top of glass tubing. To coat HA solution uniformly, we flipped over the glass tubing when the solution flew down the bottom of the tubing and repeated this step. This HA coating is a critical to our fabrication procedure for releasing final scaffolds. (2) We used heat-shrinkable (HS) sleeve for retaining salts in the glass tubing. Since salts were densely packed in the space between the inner wall of glass tubing and the HS sleeve, HS sleeve retained salts after removing mandrel and PTFE ring in the bottom of the tubing. We could remove HS sleeve easily by putting the mold into an oven at 120 &deg;C for 5 min, and then get tubular salt templates. (3) We used the salt fusion method. It is well known that salt fusion method can improve pore interconnectivity and mechanical properties by varying fusion time10. Furthermore, since we used PGS, macro-pores were produced by the salt particles during leaching process, while the micro-pores were likely generated by glycerol vapor formed during PGS curing as we described previously11. Thus, this method has a potential to fabricate porous tubular scaffolds with different macro- and micro-structures by varying salt particles as well as PGS curing condition.
The mechanical conditioning from the bioreactor has provided pulsatile flow perfusion (maximum mean flow = 14 ml/min, maximum shear stress = 15.3 dyne/cm2, frequency = 0.5 - 1.7 Hz) and physiologically relevant pressure with the PGS scaffold, which led to SMC growth and orientation (Fig. 4). These results are consistent with previous studies reporting that cyclic stretch at this frequency and shear stress increases SMC proliferation12, and ECM protein production13,14. In addition to SMC growth and orientation, PGS construct supported ECM protein production, especially circumferentially organized elastic fibers (Fig. 5) within 3-week culture in the bioreactor. Some studies using an elastomeric scaffold as a small-diameter arterial construct have demonstrated mechanical strength and burst pressure comparable to native arteries15, and rapid SMC integration in compliant scaffolds using spinner flask16,17, while no elastic fibers were found in these constructs. Our results suggest that cyclic radial distension from the bioreactor improved mechanical signal transduction more effectively to SMCs in PGS scaffold, which likely contributed to elastin synthesis and organization.
Since vascular SMCs were the only cells that produced ECM proteins in our approach, quiescent endothelium and improve mechanical strength are necessary to develop a clinically successful small-diameter arterial constructs. We have reported that endothelial cells co-cultured with SMCs generated a confluent monolayer and supported phenotype protein expression under our culture conditions and mechanical conditioning9. Therefore, based on our approach described here, modification of co-culture experiment conditions would be a next step to improve the functions of resultant constructs and generate nonthrombogenic, robust, and compliant arterial construct similar to native arteries.

<table border="1">
	<tbody>
 <tr>
 	<th>Name of the reagent</th>
 <th>Company</th>
 <th>Catalogue number</th>
 </tr>
 <tr>
 	<td>Hyaluronic acid sodium salt</td>
 <td>Sigma-Aldrich</td>
 <td>H7630</td>
 </tr>
 <tr>
 	<td>Tetrahydrofuran</td>
 <td>Sigma-Aldrich</td>
 <td>401757</td>
 </tr>
 <tr>
 	<td>MCDB 131</td>
 <td>Mediatech</td>
 <td>15-100-CV</td>
 </tr>
 <tr>
 	<td>Fetal bovine serum</td>
 <td>Lonza</td>
 <td>BW14-502F</td>
 </tr>
 <tr>
 	<td>L-glutamine</td>
 <td>Mediatech</td>
 <td>25-005-CV</td>
 </tr>
 <tr>
 	<td>Ascorbic acid</td>
 <td>Fisher Scientific</td>
 <td>A62-500</td>
 </tr>
 <tr>
 	<td>Antibiotic-antimycotic solution</td>
 <td>Mediatech</td>
 <td>30-004-CI</td>
 </tr>
 <tr>
 	<td>Phosphate Buffered Saline (PBS)</td>
 <td>Mediatech</td>
 <td>21-031-CV</td>
 </tr>
 <tr>
 	<td>Tissue-Tek optimal cutting temperature compound, 4583</td>
 <td>Sakura Finetek</td>
 <td>25608-930</td>
 </tr> 
 </tbody>
</table>

elastomeric pgs scaffolds in arterial tissue engineering

Cardiovascular disease is one of the leading cause of mortality in the US and especially, coronary artery disease increases with an aging population and increasing obesity1. Currently, bypass surgery using autologous vessels, allografts, and synthetic grafts are known as a commonly used for arterial substitutes2. However, these grafts have limited applications when an inner diameter of arteries is less than 6 mm due to low availability, thrombotic complications, compliance mismatch, and late intimal hyperplasia3,4. To overcome these limitations, tissue engineering has been successfully applied as a promising alternative to develop small-diameter arterial constructs that are nonthrombogenic, robust, and compliant. Several previous studies have developed small-diameter arterial constructs with tri-lamellar structure, excellent mechanical properties and burst pressure comparable to native arteries5,6. While high tensile strength and burst pressure by increasing collagen production from a rigid material or cell sheet scaffold, these constructs still had low elastin production and compliance, which is a major problem to cause graft failure after implantation. Considering these issues, we hypothesized that an elastometric biomaterial combined with mechanical conditioning would provide elasticity and conduct mechanical signals more efficiently to vascular cells, which increase extracellular matrix production and support cellular orientation.
The objective of this report is to introduce a fabrication technique of porous tubular scaffolds and a dynamic mechanical conditioning for applying them to arterial tissue engineering. We used a biodegradable elastomer, poly (glycerol sebacate) (PGS)7 for fabricating porous tubular scaffolds from the salt fusion method. Adult primary baboon smooth muscle cells (SMCs) were seeded on the lumen of scaffolds, which cultured in our designed pulsatile flow bioreactor for 3 weeks. PGS scaffolds had consistent thickness and randomly distributed macro- and micro-pores. Mechanical conditioning from pulsatile flow bioreactor supported SMC orientation and enhanced ECM production in scaffolds. These results suggest that elastomeric scaffolds and mechanical conditioning of bioreactor culture may be a promising method for arterial tissue engineering.

Watch this Scientific Journal Video about Elastomeric PGS Scaffolds in Arterial Tissue Engineering at JoVE.com

Elastomeric PGS Scaffolds in Arterial Tissue Engineering

Elastomeric PGS scaffolds with vascular smooth muscle cells cultured in a pulsatile flow bioreactor may lead to promising small-diameter arterial constructs with native ECM production in a relatively short culture period.

Cardiovascular disease is one of the leading cause of mortality in the US and especially, coronary artery disease increases with an aging population and ...

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15.5K Görüntüleme.  University of Pittsburgh. Pulsatil akış biyoreaktör kültürü damar düz kas hücreleri ile Elastomerik PGS iskeleler, görece kısa bir kültür dönemde, yerli ECM üretim ile gelecek vaat eden küçük çaplı arter yapıları neden olabilir.

Video: Elastomeric PGS Scaffolds in Arterial Tissue Engineering

Tissue Engineering of the Intestine in a Murine Model

Tissue-engineered small intestine (TESI) has successfully been used to rescue Lewis rats after massive small bowel resection, resulting in return to preoperative weights within 40 days.1 In humans, massive small bowel resection can result in short bowel syndrome, a functional malabsorptive state that confers significant morbidity, mortality, and healthcare costs including parenteral nutrition dependence, liver failure and cirrhosis, and the need for multivisceral organ transplantation.2 In this paper, we describe and document our protocol for creating tissue-engineered intestine in a mouse model with a multicellular organoid units-on-scaffold approach. Organoid units are multicellular aggregates derived from the intestine that contain both mucosal and mesenchymal elements,3 the relationship between which preserves the intestinal stem cell niche.4 In ongoing and future research, the transition of our technique into the mouse will allow for investigation of the processes involved during TESI formation by utilizing the transgenic tools available in this species.5The availability of immunocompromised mouse strains will also permit us to apply the technique to human intestinal tissue and optimize the formation of human TESI as a mouse xenograft before its transition into humans. Our method employs good manufacturing practice (GMP) reagents and materials that have already been approved for use in human patients, and therefore offers a significant advantage over approaches that rely upon decellularized animal tissues. The ultimate goal of this method is its translation to humans as a regenerative medicine therapeutic strategy for short bowel syndrome.

This article and the accompanying video present our protocol for generating tissue-engineered intestine in the mouse, using an organoid units-on-scaffold approach.

Tissue-engineered small intestine (TESI) has successfully been used to rescue Lewis rats after massive small bowel resection, resulting in return to ...

Density Gradient Multilayered Polymerization (DGMP): A Novel Technique for Creating Multi-compartment, Customizable Scaffolds for Tissue Engineering

Complex tissue culture matrices, in which types and concentrations of biological stimuli (e.g. growth factors, inhibitors, or small molecules) or matrix structure (e.g. composition, concentration, or stiffness of the matrix) vary over space, would enable a wide range of investigations concerning how these variables affect cell differentiation, migration, and other phenomena. The major challenge in creating layered matrices is maintaining the structural integrity of layer interfaces without diffusion of individual components from each layer1. Current methodologies to achieve this include photopatterning2-3, lithography4, sequential functionalization5, freeze drying6, microfluidics7, or centrifugation8, many of which require sophisticated instrumentation and technical skills. Others rely on sequential attachment of individual layers, which may lead to delamination of layers9. 
DGMP overcomes these issues by using an inert density modifier such as iodixanol to create layers of varying densities10. Since the density modifier can be mixed with any prepolymer or bioactive molecule, DGMP allows each scaffold layer to be customized. Simply varying the concentration of the density modifier prevents mixing of adjacent layers while they remain aqueous. Subsequent single step polymerization gives rise to a structurally continuous multilayered scaffold, in which each layer has distinct chemical and mechanical properties. The density modifier can be easily removed with sufficient rinsing without perturbation of the individual layers or their components. This technique is therefore well suited for creating hydrogels of various sizes, shapes, and materials.
A protocol for fabricating a 2D-polyethylene glycol (PEG) gel, in which alternating layers incorporate RGDS-350, is outlined below. We use PEG because it is biocompatible and inert. RGDS, a cell adhesion peptide11, is used to demonstrate spatial restriction of a biological cue, and the conjugation of a fluorophore (Alexa Fluor 350) enables us to visually distinguish various layers. This procedure can be adapted for other materials (e.g. collagen, hyaluronan, etc.) and can be extended to fabricate 3D gels with some modifications10.

Density Gradient Multilayered Polymerization DGMP: A Novel Technique for Creating Multi-compartment, Customizable Scaffolds for Tissue Engineering

Here we describe a unique strategy for creating biocompatible, layered matrices with continuous interfaces between distinct layers for tissue engineering. Such a scaffold could provide an ideal customizable environment to modulate cell behavior by various biological, chemical or mechanical cues

Complex tissue culture matrices, in which types and concentrations of biological stimuli (e.g. growth factors, inhibitors, or small molecules) or matrix ...

Capillary Force Lithography for Cardiac Tissue Engineering

Cardiovascular disease remains the leading cause of death worldwide1. Cardiac tissue engineering holds much promise to deliver groundbreaking medical discoveries with the aims of developing functional tissues for cardiac regeneration as well as in vitro screening assays. However, the ability to create high-fidelity models of heart tissue has proven difficult. The heart&#8217;s extracellular matrix (ECM) is a complex structure consisting of both biochemical and biomechanical signals ranging from the micro- to the nanometer scale2. Local mechanical loading conditions and cell-ECM interactions have recently been recognized as vital components in cardiac tissue engineering3-5.
A large portion of the cardiac ECM is composed of aligned collagen fibers with nano-scale diameters that significantly influences tissue architecture and electromechanical coupling2. Unfortunately, few methods have been able to mimic the organization of ECM fibers down to the nanometer scale. Recent advancements in nanofabrication techniques, however, have enabled the design and fabrication of scalable scaffolds that mimic the in vivo structural and substrate stiffness cues of the ECM in the heart6-9.
Here we present the development of two reproducible, cost-effective, and scalable nanopatterning processes for the functional alignment of cardiac cells using the biocompatible polymer poly(lactide-co-glycolide) (PLGA)8 and a polyurethane (PU) based polymer. These anisotropically nanofabricated substrata (ANFS) mimic the underlying ECM of well-organized, aligned tissues and can be used to investigate the role of nanotopography on cell morphology and function10-14.
Using a nanopatterned (NP) silicon master as a template, a polyurethane acrylate (PUA) mold is fabricated. This PUA mold is then used to pattern the PU or PLGA hydrogel via UV-assisted or solvent-mediated capillary force lithography (CFL), respectively15,16. Briefly, PU or PLGA pre-polymer is drop dispensed onto a glass coverslip and the PUA mold is placed on top. For UV-assisted CFL, the PU is then exposed to UV radiation (&#955; = 250-400 nm) for curing. For solvent-mediated CFL, the PLGA is embossed using heat (120 &#176;C) and pressure (100 kPa). After curing, the PUA mold is peeled off, leaving behind an ANFS for cell culture. Primary cells, such as neonatal rat ventricular myocytes, as well as human pluripotent stem cell-derived cardiomyocytes, can be maintained on the ANFS2.

In this protocol, we demonstrate the fabrication of biomimetic cardiac cell culture substrata made from two distinct polymeric materials using capillary force lithography. The described methods provide a scalable, cost-effective technique to engineer the structure and function of macroscopic cardiac tissues for in vitro and in vivo applications.

Cardiovascular disease remains the leading cause of death worldwide1. Cardiac tissue engineering holds much promise to deliver groundbreaking medical ...

Tissue Engineering of a Human 3D in vitro Tumor Test System

Cancer is one of the leading causes of death worldwide. Current therapeutic strategies are predominantly developed in 2D culture systems, which inadequately reflect physiological conditions in vivo. Biological 3D matrices provide cells an environment in which cells can self-organize, allowing the study of tissue organization and cell differentiation. Such scaffolds can be seeded with a mixture of different cell types to study direct 3D cell-cell-interactions. To mimic the 3D complexity of cancer tumors, our group has developed a 3D in vitro tumor test system.	
	Our 3D tissue test system models the in vivo situation of malignant peripheral nerve sheath tumors (MPNSTs), which we established with our decellularized porcine jejunal segment derived biological vascularized scaffold (BioVaSc). In our model, we reseeded a modified BioVaSc matrix with primary fibroblasts, microvascular endothelial cells (mvECs) and the S462 tumor cell line. For static culture, the vascular structure of the BioVaSc is removed and the remaining scaffold is cut open on one side (Small Intestinal Submucosa SIS-Muc). The resulting matrix is then fixed between two metal rings (cell crowns).	
	Another option is to culture the cell-seeded SIS-Muc in a flow bioreactor system that exposes the cells to shear stress. Here, the bioreactor is connected to a peristaltic pump in a self-constructed incubator. A computer regulates the arterial oxygen and nutrient supply via parameters such as blood pressure, temperature, and flow rate. This setup allows for a dynamic culture with either pressure-regulated pulsatile or constant flow.	
	In this study, we could successfully establish both a static and dynamic 3D culture system for MPNSTs. The ability to model cancer tumors in a more natural 3D environment will enable the discovery, testing, and validation of future pharmaceuticals in a human-like model.

Tissue Engineering of a Human 3D in vitro Tumor Test System

Methods to create human 3D tumor tissues as test systems are described. These technologies are based on a decellularized Biological Vascularized Scaffold (BioVaSc), primary human cells and a tumor cell line, which can be cultured under static as well as under dynamic conditions in a flow bioreactor.

Cancer is one of the leading causes of death worldwide.  Current therapeutic strategies are predominantly developed in 2D culture  systems, which ...

Self-reporting Scaffolds for 3-Dimensional Cell Culture

Culturing cells in 3D on appropriate scaffolds is thought to better mimic the in vivo microenvironment and increase cell-cell interactions. The resulting 3D cellular construct can often be more relevant to studying the molecular events and cell-cell interactions than similar experiments studied in 2D. To create effective 3D cultures with high cell viability throughout the scaffold the culture conditions such as oxygen and pH need to be carefully controlled as gradients in analyte concentration can exist throughout the 3D construct. Here we describe the methods of preparing biocompatible pH responsive sol-gel nanosensors and their incorporation into poly(lactic-co-glycolic acid) (PLGA) electrospun scaffolds along with their subsequent preparation for the culture of mammalian cells. The pH responsive scaffolds can be used as tools to determine microenvironmental pH within a 3D cellular construct. Furthermore, we detail the delivery of pH responsive nanosensors to the intracellular environment of mammalian cells whose growth was supported by electrospun PLGA scaffolds. The cytoplasmic location of the pH responsive nanosensors can be utilized to monitor intracellular pH (pHi) during ongoing experimentation.

Biocompatible pH responsive sol-gel nanosensors can be incorporated into poly(lactic-co-glycolic acid) (PLGA) electrospun scaffolds. The produced self-reporting scaffolds can be used for in&#160;situ monitoring of microenvironmental conditions whilst culturing cells upon the scaffold. This is beneficial as the 3D cellular construct can be monitored in real-time without disturbing the experiment.

Culturing cells in 3D on appropriate scaffolds is thought to better mimic the in vivo microenvironment and increase cell-cell interactions. The resulting ...

Electrospun Fibrous Scaffolds of Poly(glycerol-dodecanedioate) for Engineering Neural Tissues From Mouse Embryonic Stem Cells

For tissue engineering applications, the preparation of biodegradable and biocompatible scaffolds is the most desirable but challenging task. &#160;Among the various fabrication methods, electrospinning is the most attractive one due to its simplicity and versatility. Additionally, electrospun nanofibers mimic the size of natural extracellular matrix ensuring additional support for cell survival and growth. This study showed the viability of the fabrication of long fibers spanning a larger deposit area for a novel biodegradable and biocompatible polymer named&#160;poly(glycerol-dodecanoate) (PGD)1 by using a newly designed collector for electrospinning. PGD exhibits unique elastic properties with similar mechanical properties to nerve tissues, thus it is suitable for neural tissue engineering applications. The synthesis and fabrication set-up for making fibrous scaffolding materials was simple, highly reproducible, and inexpensive. In biocompatibility testing, cells derived from mouse embryonic stem cells could adhere to and grow on the electrospun PGD fibers. In summary, this protocol provided a versatile fabrication method for making PGD electrospun fibers to support the growth of mouse embryonic stem cell derived neural lineage cells.

Electrospun Fibrous Scaffolds of Polyglycerol-dodecanedioate for Engineering Neural Tissues From Mouse Embryonic Stem Cells

Synthesis and fabrication of electrospun long fibers spanning a larger deposit area via a newly designed collector from a novel biodegradable polymer named poly(glycerol-dodecanoate) (PGD) was reported. The fibers were able to support the growth of cells derived from mouse pluripotent stem cells.

For tissue engineering applications, the preparation of biodegradable and biocompatible scaffolds is the most desirable but challenging task. &#160;Among ...

Engineering 3D Cellularized Collagen Gels for Vascular Tissue Regeneration

Synthetic materials are known to initiate clinical complications such as inflammation, stenosis, and infections when implanted as vascular substitutes. Collagen has been extensively used for a wide range of biomedical applications and is considered a valid alternative to synthetic materials due to its inherent biocompatibility (i.e., low antigenicity, inflammation, and cytotoxic responses). However, the limited mechanical properties and the related low hand-ability of collagen gels have hampered their use as scaffold materials for vascular tissue engineering. Therefore, the rationale behind this work was first to engineer cellularized collagen gels into a tubular-shaped geometry and second to enhance smooth muscle cells driven reorganization of collagen matrix to obtain tissues stiff enough to be handled.
The strategy described here is based on the direct assembling of collagen and smooth muscle cells (construct) in a 3D cylindrical geometry with the use of a molding technique. This process requires a maturation period, during which the constructs are cultured in a bioreactor under static conditions (without applied external dynamic mechanical constraints) for 1 or 2 weeks. The &#8220;static bioreactor&#8221; provides a monitored and controlled sterile environment (pH, temperature, gas exchange, nutrient supply and waste removal) to the constructs. During culture period, thickness measurements were performed to evaluate the cells-driven remodeling of the collagen matrix, and glucose consumption and lactate production rates were measured to monitor the cells metabolic activity. Finally, mechanical and viscoelastic properties were assessed for the resulting tubular constructs. To this end, specific protocols and a focused know-how (manipulation, gripping, working in hydrated environment, and so on) were developed to characterize the engineered tissues.

In this work, we present a technique for the rapid fabrication of living vascular tissues by direct culturing of collagen, smooth muscle cells and endothelial cells. In addition, a new protocol for the mechanical characterization of engineered vascular tissues is described.

Synthetic materials are known to initiate clinical complications such as inflammation, stenosis, and infections when implanted as vascular substitutes. ...

Sandwich-like Microenvironments to Harness Cell/Material Interactions

Cell culture has been traditionally carried out on bi-dimensional (2D) substrates where cells adhere using ventral receptors to the biomaterial surface. However in vivo, most of the cells are completely surrounded by the extracellular matrix (ECM), resulting in a three-dimensional (3D) distribution of receptors. This may trigger differences in the outside-in signaling pathways and thus in cell behavior.
This article shows that stimulating the dorsal receptors of cells already adhered to a 2D substrate by overlaying a film of a new material (a sandwich-like culture) triggers important changes with respect to standard 2D cultures. Furthermore, the simultaneous excitation of ventral and dorsal receptors shifts cell behavior closer to that found in 3D environments. Additionally, due to the nature of the system, a sandwich-like culture is a versatile tool that allows the study of different parameters in cell/material interactions, e.g., topography, stiffness and different protein coatings at both the ventral and dorsal sides. Finally, since sandwich-like cultures are based on 2D substrates, several analysis procedures already developed for standard 2D cultures can be used normally, overcoming more complex procedures needed for 3D systems.

The following protocol describes the procedure to assemble sandwich-like cultures to be used as an intermediate stage between bi-dimensional (2D) and three-dimensional (3D) cellular environments. The engineered systems can have applications in microscopy, biomechanics, biochemistry and cell biology assays.

Cell culture has been traditionally carried out on bi-dimensional (2D) substrates where cells adhere using ventral receptors to the biomaterial surface. ...

Epithelial Cell Repopulation and Preparation of Rodent Extracellular Matrix Scaffolds for Renal Tissue Development

This protocol details the generation of acellular, yet biofunctional, renal extracellular matrix (ECM) scaffolds that are useful as small-scale model substrates for organ-scale tissue development. Sprague Dawley rat kidneys are cannulated by inserting a catheter into the renal artery and perfused with a series of low-concentration detergents (Triton X-100 and sodium dodecyl sulfate (SDS)) over 26 hr to derive intact, whole-kidney scaffolds with intact perfusable vasculature, glomeruli, and renal tubules. Following decellularization, the renal scaffold is placed inside a custom-designed perfusion bioreactor vessel, and the catheterized renal artery is connected to a perfusion circuit consisting of: a peristaltic pump; tubing; and optional probes for pH, dissolved oxygen, and pressure. After sterilizing the scaffold with peracetic acid and ethanol, and balancing the pH (7.4), the kidney scaffold is prepared for seeding via perfusion of culture medium within a large-capacity incubator maintained at 37 &#176;C and 5% CO2. Forty million renal cortical tubular epithelial (RCTE) cells are injected through the renal artery, and rapidly perfused through the scaffold under high flow (25 ml/min) and pressure (~230 mmHg) for 15 min before reducing the flow to a physiological rate (4 ml/min). RCTE cells primarily populate the tubular ECM niche within the renal cortex, proliferate, and form tubular epithelial structures over seven days of perfusion culture. A 44 &#181;M resazurin solution in culture medium is perfused through the kidney for 1 hr during medium exchanges to provide a fluorometric, redox-based metabolic assessment of cell viability and proliferation during tubulogenesis. The kidney perfusion bioreactor permits non-invasive sampling of medium for biochemical assessment, and multiple inlet ports allow alternative retrograde seeding through the renal vein or ureter. These protocols can be used to recellularize kidney scaffolds with a variety of cell types, including vascular endothelial, tubular epithelial, and stromal fibroblasts, for rapid evaluation within this system.

This protocol describes decellularization of Sprague Dawley rat kidneys by antegrade perfusion of detergents through the vasculature, producing acellular renal extracellular matrices that serve as templates for repopulation with human renal epithelial cells. Recellularization and use of the resazurin perfusion assay to monitor growth is performed within specially-designed perfusion bioreactors.

This protocol details the generation of acellular, yet biofunctional, renal extracellular matrix (ECM) scaffolds that are useful as small-scale model ...

Tissue Engineering by Intrinsic Vascularization in an In Vivo Tissue Engineering Chamber

In reconstructive surgery, there is a clinical need for an alternative to the current methods of autologous reconstruction which are complex, costly and trade one defect for another. Tissue engineering holds the promise to address this increasing demand. However, most tissue engineering strategies fail to generate stable and functional tissue substitutes because of poor vascularization. This paper focuses on an in vivo tissue engineering chamber model of intrinsic vascularization where a perfused artery and a vein either as an arteriovenous loop or a flow-through pedicle configuration is directed inside a protected hollow chamber. In this chamber-based system angiogenic sprouting occurs from the arteriovenous vessels and this system attracts ischemic and inflammatory driven endogenous cell migration which gradually fills the chamber space with fibro-vascular tissue. Exogenous cell/matrix implantation at the time of chamber construction enhances cell survival and determines specificity of the engineered tissues which develop. Our studies have shown that this chamber model can successfully generate different tissues such as fat, cardiac muscle, liver and others. However, modifications and refinements are required to ensure target tissue formation is consistent and reproducible. This article describes a standardized protocol for the fabrication of two different vascularized tissue engineering chamber models in vivo.

Tissue Engineering by Intrinsic Vascularization in an In Vivo Tissue Engineering Chamber

This is a guideline for constructing in vivo vascularized tissue using a microsurgical arteriovenous loop or a flow-through pedicle configuration inside a tissue engineering chamber. The vascularized tissues generated can be employed for organ regeneration and replacement of tissue defects, as well as for drug testing and disease modeling.

In reconstructive surgery, there is a clinical need for an alternative to the current methods of autologous reconstruction which are complex, costly and ...