The authors gratefully acknowledge Neil Whitehouse (WPI, Higgins Machine Shop) for his assistance with CNC machining. Additionally, we would like to thank Adriana Hera (WPI Computing and Communications Center) for her help with MATLAB programming, as well as Kate Beverage and Joseph Cotnoir (WPI Academic Technology Center) for assistance with Camtasia. Sophie Burke and Jacleen Becker (WPI Academic Technology Center) provided supplemental video footage. This work was funded by the National Institutes of Health (R15 HL097332), the UMass Medical School-WPI Pilot Research Initiative, the American Heart Association (undergraduate research fellowship to J.Z.H), and Worcester Polytechnic Institute (Summer Undergraduate Research Fellowship to J.Z.H. and institutional start-up funds to M.W.R.).

1. Cell seeding mold fabrication
Begin by milling a 1/2" thick piece of polycarbonate to create 15, round-bottomed, annular wells with a center post diameter of 2 mm. The milled channels are 6 mm deep and 3.75 mm wide. Clean and dry the polycarbonate mold to remove any plastic debris from the milling process.
Mix polydimethylsiloxane (PDMS) at a 10:1 ratio (w/w) of base to curing agent, degas to remove all air bubbles, and pour onto the polycarbonate mold. Degas again to remove any remaining bubbles, and cure in the oven at 60&deg;C for 4 hours.
Once cured, carefully remove the PDMS template by slowly peeling it away from the polycarbonate, wash with soap and water, and autoclave. Also autoclave a solution of two percent agarose (w/v) dissolved in Dulbecco's modified Eagle medium (DMEM).
Place the PDMS template on a level surface and fill with molten agarose by first pipetting agarose into each of the center post molds, and then pipetting into the space around it. Allow the agarose to solidify (approximately 15 minutes), then invert the mold to release the agarose from the PDMS. Cut excess agarose from around each of the wells, and place the agarose wells into 12-well plates.
2. Cell culture and ring seeding
Add cell culture media (DMEM with 10% FBS and 1% penicillin-streptomycin) around the outside of the agarose mold, without covering the top of the molds. Place the plates in the incubator and allow them to equilibrate for approximately 15 minutes while you prepare the cells.
At 90% confluence, trypsinize rat aortic smooth muscle cells (rSMCs) and resuspend at a concentration of 5x106 cells/ml. Pipette 100 &mu;L of cell suspension into each of the annular agarose wells, using a circular motion to apply cells to each well.
Make sure incubator shelves are level, then place the plates in the incubator and allow them to sit undisturbed for 24 hours. Exchange medium every two days thereafter by aspirating the media from around the well, and re-filling each well until the agarose well is completely submerged.
3. Tissue ring harvesting and thickness measurements
At the conclusion of culture, remove the ring from the agarose mold by sliding it over the top of the center post.
Place the ring in a small Petri dish with phosphate buffered saline (PBS), center the sample, and acquire an image using a digital imaging system.
Use an image analysis program for edge detection and measure the thickness of the ring in 4 positions (top, bottom, left and right) around its circumference. Calculate the mean thickness value (t) for each individual ring, and use this value to calculate the cross-sectional area (2&pi;(t/2)2).
4. Mechanical testing of rings
Set up the tensile testing machine (Instron EPS 1000) in the horizontal position. Attach a 1N &plusmn;1mN load cell and custom thin wire grips (created by bending stainless steel wire).
Mount the ring sample on the two thin wire grips. Extend the grips until a 5 mN tare load is applied to the sample. Enter the cross-sectional area (calculated from thickness measurements) and record the gauge length.
Pre-cycle the rings 8 times between the tare load and 50 kPa stress at a rate of 10 mm/min. After the eighth pre-cycle, pull to failure at 10 mm/min. Failure is noted as a decrease in force by 40% from one measurement to the next. For each ring sample, measure the ultimate tensile stress (UTS) and failure strain and calculate the maximum tangent modulus (MTM) from the acquired data.
5. Tissue ring assembly to fabricate tissue tubes
Custom tube holders are machined from round 5 cm polycarbonate disks with rectangular cutouts (2 cm x 3 cm). Drill threaded holes through these disks to allow two holders to be screwed together. Autoclave the custom tube holders, and then transfer to the biosafety cabinet and place in an empty Petri dish.
Use surgical scissors to cut the ends of 1.9 mm diameter silicone tubes at an angle to create beveled edges, and then autoclave the silicone tubes. While the tubes are being autoclaved, fill a Petri dish with media.
Remove the rings from the agarose wells and place them in the Petri dish with media. Place autoclaved silicone tubes in the same Petri dish to wet them before use.
Use forceps to place a beveled end of a silicone tube into the center of a ring and gently slide the ring onto the silicone tubing. Repeat with the desired number of rings. Slide the rings into contact with one another by gently pushing them successively in both directions along the tube. This method can be used for rings harvested after only one day in culture.
Once rings are mounted, align silicone tubes within the polycarbonate holders and screw the two parts of the holder together. Place the holder in a 100 mm Petri dish and add 55 mL media. Exchange media every 3 days for the duration of culture.
To remove the tissue tubes, release the silicone tubes from the polycarbonate holder and use forceps to slide the tissue tubes off the silicone tube and into a Petri dish filled with PBS.
6. Representative Results:
When the protocol is performed correctly, cells aggregate to form tissue rings with an inner diameter equal to the diameter of the corresponding mold post within 24 hours. The ring edges are usually smooth in appearance and (if cultured on a level surface), are uniform in thickness around the entire circumference. The tissue rings are easy to handle and can be removed from their wells for subsequent mechanical and histological analysis (see in Gwyther et al., 20118). Tissue ring morphology, matrix composition, and mechanical properties vary depending on the number and type of cells seeded.
Representative results from tissue rings made from varying cell types, seeding conditions, and culture length are shown in Table 1. Two mm ID rings created from 5x105 rSMCs exhibited a higher UTS than rings created from our previously published 2 mm rings (made from 6.6x105 rSMCs).8 Similar to rat cells, human smooth muscle cells (hSMC) readily aggregated to form tissue rings, which contained a large amount of collagen after only 14 days in culture (data not shown). Human mesenchymal stem cells (hMSC) also aggregated and formed cohesive rings, but lacked mechanical strength and broke during the initial precycling phases of uniaxial tensile testing.
<table border="1">
	<tbody>
		<tr>
			<td>&nbsp;</td>
			<td>n</td>
			<td>Cell number</td>
			<td>Thickness (mm)</td>
			<td>Length of culture (days)</td>
			<td>UTS (kPa)</td>
			<td>MTM (kPa)</td>
			<td>Failure strain (mm/mm)</td>
		</tr>
		<tr>
			<td>rSMC rings</td>
			<td>6</td>
			<td>660,000</td>
			<td>0.94&plusmn;0.12</td>
			<td>14</td>
			<td>97&plusmn;30</td>
			<td>497&plusmn;91</td>
			<td>0.50&plusmn;0.08</td>
		</tr>
		<tr>
			<td>rSMC rings</td>
			<td>4</td>
			<td>500,000</td>
			<td>0.53&plusmn;0.02</td>
			<td>7</td>
			<td>113&plusmn;8</td>
			<td>189&plusmn;15</td>
			<td>0.88&plusmn;0.05</td>
		</tr>
		<tr>
			<td>hSMC rings</td>
			<td>3</td>
			<td>750,000</td>
			<td>0.51&plusmn;0.05</td>
			<td>14</td>
			<td>160&plusmn;30</td>
			<td>270&plusmn;20</td>
			<td>0.92&plusmn;0.08</td>
		</tr>
		<tr>
			<td>hMSC rings</td>
			<td>3</td>
			<td>750,000</td>
			<td>0.40&plusmn;0.07</td>
			<td>14</td>
			<td>N/A</td>
			<td>N/A</td>
			<td>N/A</td>
		</tr>
	</tbody>
</table>
Table 1. Table showing the culture parameters and mechanical properties of 2 mm rings generated from different cell types, seeding concentrations, and culture durations.
<img alt="Figure 1" src="/files/ftp_upload/3366/3366fig1.jpg" /> 
Figure 1. (A) Schematic of the tissue ring generation process. (B) Custom polycarbonate mold with milled annular wells. Central post diameters are 2 mm. (C) PDMS template after it was peeled away from the polycarbonate mold. (D) Aggregated tissue ring cultured in an agarose mold with a 2 mm post. (E) Two mm diameter tissue ring in PBS. Scale bars = 6 mm (B, C) and 2 mm (D, E).
<img alt="Figure 2" src="/files/ftp_upload/3366/3366fig2.jpg" /> 
Figure 2. Representative plot of stress-strain curve produced from uniaxial tensile testing.

<ol>
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No conflicts of interest declared.

Recently, there has been an increased interest in cell-based or "scaffold-less" tissue engineering methods to address some of the limitations of scaffold-based tissue engineering approaches. Given that cell-derived tissues are created from cells and the matrix they produce, they inherently contain much higher cell densities, contain no exogenous materials, and can be made entirely from human cells and proteins. Vascular grafts made from human cells can attain substantial mechanical strength in the absence of exogenous scaffolds (e.g., burst pressures 3400 mmHg compared to 1600 mmHg for human saphenous veins)12. Although cell-based vascular tissues exhibit improved cell density and mechanical strength, most current fabrication methods (such as "sheet-based engineering"3,4,12 or "bioprinting"5,13) require extensive culture periods (&gt; 3 months) or specialized equipment for 3D tissue construction. The cell-derived tissue ring method described here enables rapid cellular self-assembly to form robust 3D tissue constructs within a short period of time and without the use of specialized equipment.
This protocol details the procedure we developed to create 2 mm inner diameter rat smooth muscle cell-derived tissue rings. In the present example, tissue rings were cultured for 7 days (then cultured for an additional 7 days for ring fusion and tube formation). However, 2 mm rat (and human) smooth muscle cell rings can be removed from the wells and are cohesive enough for handling (e.g., transfer onto silicone tubes) as early as one day after cell seeding. In addition, robust tissue rings with varying inner diameters (2, 4, and 6 mm) can be created with this method by simply changing the post diameter of the original polycarbonate mold.8 We also recently modified the polycarbonate mold design to enable five 2 mm seeding wells to be cast in a single multi-well agarose chamber, which uses less PDMS and agarose, and fits in one well of a 6-well plate (data not shown). Changes in the post diameter, the width of the seeding well, the radius of curvature of the rounded bottom, the number of seeding wells, or the depth of the seeding wells can all be modified simply by changing the specifications in the CAD file for CNC machining of the polycarbonate mold. Finally, a single polycarbonate mold can be used to fabricate an unlimited number of PDMS templates, and each PDMS template can be cleaned, autoclaved and reused dozens of times.
In addition to changing the size of the tissue rings, we have made rings from many different cell types, including: primary rat SMCs (Cell Applications, R354-05), primary human coronary artery SMCs (Lonza, CC-2583), primary human dermal fibroblasts11 (generous gift of Dr. George Pins, WPI Dept. of Biomedical Engineering), rat lung fibroblasts (RFL-6, ATCC CCL-192), and mesenchymal stem cells (Lonza, PT-2501). Each of these cell types aggregates and contracts around the center posts to form tissue rings; although the cellular organization, ECM composition, and mechanical properties of the constructs vary for each cell type. The seeding parameters for each cell type must be empirically determined based on the size of the cells and their ability to aggregate. Therefore, while this system of creating cell-derived tissue rings is extremely versatile, the protocol may need slight adjustments for optimal tissue formation with different cell types.
Tissue ring geometry facilitates easy loading and assessment of tissue material properties by uniaxial tensile testing, as described. There is also substantial precedent for using blood vessel ring segments to measure vascular contraction and physiological function. Preliminary studies indicate that cell-derived tissue rings can be mounted on a wire myograph device for measurement of pharmacological responsiveness and contractile force generation (data not shown). Altogether, the ability to rapidly fabricate self-assembled cell rings for histological, mechanical, physiological, and biochemical analysis suggests a powerful new tool that may be useful for modeling vascular tissue structure and function in health and disease.

<table><tbody><tr><td>Polydimethylsiloxane (PDMS)</td><td>Dow Corning</td><td>Sylgard 184</td><td></td></tr><tr><td>Agarose</td><td>Lonza Inc.</td><td>50000</td><td></td></tr><tr><td>DMEM</td><td>Mediatech, Inc.</td><td>15-017-CV</td><td></td></tr><tr><td>Fetal Bovine Serum (FBS)</td><td>PAA Laboratories</td><td>A05-201</td><td></td></tr><tr><td>Penicillin/Streptomycin</td><td>Mediatech, Inc.</td><td>30-002-CI</td><td></td></tr><tr><td>Digital imaging system</td><td>DVT Corporation</td><td>Model 630</td><td></td></tr><tr><td>Uniaxial testing machine</td><td>Instron</td><td>ElectroPuls E1000</td><td></td></tr><tr><td>Edge Detection Software</td><td>DVT Corporation</td><td>Framework 2.4.6</td><td></td></tr></tbody></table>

directed cellular self-assembly to fabricate cell-derived tissue rings for biomechanical analysis and tissue engineering

Each year, hundreds of thousands of patients undergo coronary artery bypass surgery in the United States.1 Approximately one third of these patients do not have suitable autologous donor vessels due to disease progression or previous harvest. The aim of vascular tissue engineering is to develop a suitable alternative source for these bypass grafts. In addition, engineered vascular tissue may prove valuable as living vascular models to study cardiovascular diseases. Several promising approaches to engineering blood vessels have been explored, with many recent studies focusing on development and analysis of cell-based methods.2-5 Herein, we present a method to rapidly self-assemble cells into 3D tissue rings that can be used in vitro to model vascular tissues.
To do this, suspensions of smooth muscle cells are seeded into round-bottomed annular agarose wells. The non-adhesive properties of the agarose allow the cells to settle, aggregate and contract around a post at the center of the well to form a cohesive tissue ring.6,7 These rings can be cultured for several days prior to harvesting for mechanical, physiological, biochemical, or histological analysis. We have shown that these cell-derived tissue rings yield at 100-500 kPa ultimate tensile strength8 which exceeds the value reported for other tissue engineered vascular constructs cultured for similar durations (&lt;30 kPa).9,10 Our results demonstrate that robust cell-derived vascular tissue ring generation can be achieved within a short time period, and offers the opportunity for direct and quantitative assessment of the contributions of cells and cell-derived matrix (CDM) to vascular tissue structure and function.

Watch this Scientific Journal Video about Directed Cellular Self-Assembly to Fabricate Cell-Derived Tissue Rings for Biomechanical Analysis and Tissue Engineering at JoVE.com

Directed Cellular Self-Assembly to Fabricate Cell-Derived Tissue Rings for Biomechanical Analysis and Tissue Engineering

This article outlines a versatile method to create cell-derived tissue rings by cellular self-assembly. Smooth muscle cells seeded into ring-shaped agarose wells aggregate and contract to form robust three-dimensional (3D) tissues within 7 days. Millimeter-scale tissue rings are conducive to mechanical testing and serve as building blocks for tissue assembly.

Each year, hundreds of thousands of patients undergo coronary artery bypass surgery in the United States.1 Approximately one third of these patients do ...

directed-cellular-self-assembly-to-fabricate-cell-derived-tissue

Research

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Bioengineering

18.9K Views.  Worcester Polytechnic Institute. This article outlines a versatile method to create cell-derived tissue rings by cellular self-assembly. Smooth muscle cells seeded into ring-shaped agarose wells aggregate and contract to form robust three-dimensional (3D) tissues within 7 days. Millimeter-scale tissue rings are conducive to mechanical testing and serve as building blocks for tissue assembly.

Video: Directed Cellular Self-Assembly to Fabricate Cell-Derived Tissue Rings for Biomechanical Analysis and Tissue Engineering

Manipulating Living Cells to Construct Stable 3D Cellular Assembly Without Artificial Scaffold

Regenerative medicine and tissue engineering offer several advantages for the treatment of intractable diseases, and several studies have demonstrated the importance of 3-dimensional (3D) cellular assemblies in these fields. Artificial scaffolds have often been used to construct 3D cellular assemblies. However, the scaffolds used to construct cellular assemblies are sometimes toxic and may change the properties of the cells. Thus, it would be beneficial to establish a non-toxic method for facilitating cell-cell contact. In this paper, we introduce a novel method for constructing stable cellular assemblies by using optical tweezers with dextran. One of the advantages of this method is that it establishes stable cell-to-cell contact within a few minutes. This new method allows the construction of 3D cellular assemblies in a natural hydrophilic polymer and is expected to be useful for constructing next-generation 3D single-cell assemblies in the fields of regenerative medicine and tissue engineering.

We demonstrate a novel method for constructing a single-cell-based 3-dimensional (3D) assembly without an artificial scaffold.

Regenerative medicine and tissue engineering offer several advantages for the treatment of intractable diseases, and several studies have demonstrated the ...

Biochemistry

Tissue Engineering: Construction of a Multicellular 3D Scaffold for the Delivery of Layered Cell Sheets

Many tissues, such as the adult human hearts, are unable to adequately regenerate after damage.2,3 Strategies in tissue engineering propose innovations to assist the body in recovery and repair. For example, TE approaches may be able to attenuate heart remodeling after myocardial infarction (MI) and possibly increase total heart function to a near normal pre-MI level.4 As with any functional tissue, successful regeneration of cardiac tissue involves the proper delivery of multiple cell types with environmental cues favoring integration and survival of the implanted cell/tissue graft. Engineered tissues should address multiple parameters including: soluble signals, cell-to-cell interactions, and matrix materials evaluated as delivery vehicles, their effects on cell survival, material strength, and facilitation of cell-to-tissue organization. Studies employing the direct injection of graft cells only ignore these essential elements.2,5,6 A tissue design combining these ingredients has yet to be developed. Here, we present an example of integrated designs using layering of patterned cell sheets with two distinct types of biological-derived materials containing the target organ cell type and endothelial cells for enhancing new vessels formation in the &#8220;tissue&#8221;. Although these studies focus on the generation of heart-like tissue, this tissue design can be applied to many organs other than heart with minimal design and material changes, and is meant to be an off-the-shelf product for regenerative therapies. The protocol contains five detailed steps. A temperature sensitive Poly(N-isopropylacrylamide) (pNIPAAM) is used to coat tissue culture dishes. Then, tissue specific cells are cultured on the surface of the coated plates/micropattern surfaces to form cell sheets with strong lateral adhesions. Thirdly, a base matrix is created for the tissue by combining porous matrix with neovascular permissive hydrogels and endothelial cells. Finally, the cell sheets are lifted from the pNIPAAM coated dishes and transferred to the base element, making the complete construct.

For creation of highly organized structures of complex tissue, one must assemble multiple material and cell types into an integrated composite. This combinatorial design incorporates organ-specific layered cell sheets with two distinct biologically-derived materials containing a strong fibrous matrix base, and endothelial cells for enhancing new vessels formation.

Many tissues, such as the adult human hearts, are unable to adequately regenerate after damage.2,3 Strategies in tissue engineering propose innovations to ...

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. ...

Automated Robotic Dispensing Technique for Surface Guidance and Bioprinting of Cells

This manuscript describes the introduction of cell guidance features followed by the direct delivery of cells to these features in a hydrogel bioink using an automated robotic dispensing system. The particular bioink was selected as it allows cells to sediment towards and sense the features. The dispensing system bioprints viable cells in hydrogel bioinks using a backpressure assisted print head. However, by replacing the print head with a sharpened stylus or scalpel, the dispensing system can also be employed to create topographical cues through surface etching. The stylus movement can be programmed in steps of 10 &#181;m in the X, Y and Z directions. The patterned grooves were able to orientate mesenchymal stem cells, influencing them to adopt an elongated morphology in alignment with the grooves&#39; direction. The patterning could be designed using plotting software in straight lines, concentric circles, and sinusoidal waves. In a subsequent procedure, fibroblasts and mesenchymal stem cells were suspended in a 2% gelatin bioink, for bioprinting in a backpressure driven extrusion printhead. The cell bearing bioink was then printed using the same programmed coordinates used for the etching. The bioprinted cells were able to sense and react to the etched features as demonstrated by their elongated orientation along the direction of the etched grooves.

This protocol describes a bioprinting methodology using an automated robotic depositing system that incorporates etched topographical guidance cues with the precision deposition of a cell bearing hydrogel bioink. The printed cells are directly delivered to the etched features and are able to sense and orientate with them.

This manuscript describes the introduction of cell guidance features followed by the direct delivery of cells to these features in a hydrogel bioink using ...

Scaling of Engineered Vascular Grafts Using 3D Printed Guides and the Ring Stacking Method

Coronary artery disease remains a leading cause of death, affecting millions of Americans. With the lack of autologous vascular grafts available, engineered grafts offer great potential for patient treatment. However, engineered vascular grafts are generally not easily scalable, requiring manufacture of custom molds or polymer tubes in order to customize to different sizes, constituting a time-consuming and costly practice. Human arteries range in lumen diameter from about 2.0-38 mm and in wall thickness from about 0.5-2.5 mm. We have created a method, termed the "Ring Stacking Method," in which variable size rings of tissue of the desired cell type, demonstrated here with vascular smooth muscle cells (SMCs), can be created using guides of center posts to control lumen diameter and outer shells to dictate vessel wall thickness. These tissue rings are then stacked to create a tubular construct, mimicking the natural form of a blood vessel. The vessel length can be tailored by simply stacking the number of rings required to constitute the length needed. With our technique, tissues of tubular forms, similar to a blood vessel, can be readily manufactured in a variety of dimensions and lengths to meet the needs of the clinic and patient.

Scalable engineered blood vessels would improve clinical applicability. Using easily sizable 3D-printed guides, rings of vascular smooth muscle were created and stacked into a tubular form, forming a vascular graft. Grafts can be sized to meet the range of human coronary artery dimensions by simply changing the 3D-printed guide size.

Coronary artery disease remains a leading cause of death, affecting millions of Americans. With the lack of autologous vascular grafts available, ...

Microfluidic Bioprinting for Engineering Vascularized Tissues and Organoids

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

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

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

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 ...

Controlling Electrospun Polymer Morphology for Tissue Engineering Demonstrated Using hepG2 Cell Line

Electrospinning affords researchers the opportunity to fabricate reproducible micro to nanoscale polymer fibers. The 3D fibrous architecture of electrospun polymers is regarded as a structural imitation of the extracellular matrix (ECM). Hence, electrospun fibers fabricated from biocompatible polymers have been widely investigated by tissue engineering researchers for their potential role as an artificial ECM for guiding tissue growth both in vitro and in vivo. All cells are acutely sensitive to their mechanical environment. This has been demonstrated by the discovery of multiple mechanotransduction pathways intrinsically linked to the cytoskeletal actin filaments. The cytoskeleton acts as a mechanical sensor that can direct the functionality and differentiation of the host cell depending on the stiffness and morphology of its substrate. Electrospun fibers can be tuned both in terms of fiber size and morphology to easily modulate the mechanical environment within a fibrous polymer scaffold. Here, methods for electrospinning polycaprolactone (PCL) for three distinct morphologies at two different fiber diameters are described. The morphological fiber categories consist of randomly oriented fibers, aligned fibers, and porous cryogenically spun fibers, with 1 &#181;m and 5 &#181;m diameters. The methods detailed within this study are proposed as a platform for investigating the effect of electrospun fiber architecture on tissue generation. Understanding these effects will allow researchers to optimize the mechanical properties of electrospun fibers and demonstrate the potential of this technology more thoroughly.

This method provides the means to test different polycaprolactone fiber morphologies and topographies for the purpose of tissue engineering. Small and large fibers are fabricated with random orientations, aligned orientations, and also porous cryogenically electrospun structures and used as platforms for cell culture.

Electrospinning affords researchers the opportunity to fabricate reproducible micro to nanoscale polymer fibers. The 3D fibrous architecture of ...

Simple, Affordable, and Modular Patterning of Cells using DNA

The relative positioning of cells is a key feature of the microenvironment that organizes cell-cell interactions. To study the interactions between cells of the same or different type, micropatterning techniques have proved useful. DNA Programmed Assembly of Cells (DPAC) is a micropatterning technique that targets the adhesion of cells to a substrate or other cells using DNA hybridization. The most basic operations in DPAC begin with decorating cell membranes with lipid-modified oligonucleotides, then flowing them over a substrate that has been patterned with complementary DNA sequences. Cells adhere selectively to the substrate only where they find a complementary DNA sequence. Non-adherent cells are washed away, revealing a pattern of adherent cells. Additional operations include further rounds of cell-substrate or cell-cell adhesion, as well as transferring the patterns formed by DPAC to an embedding hydrogel for long-term culture. Previously, methods for patterning oligonucleotides on surfaces and decorating cells with DNA sequences required specialized equipment and custom DNA synthesis, respectively. We report an updated version of the protocol, utilizing an inexpensive benchtop photolithography setup and commercially available cholesterol modified oligonucleotides (CMOs) deployed using a modular format. CMO-labeled cells adhere with high efficiency to DNA-patterned substrates. This approach can be used to pattern multiple cell types at once with high precision and to create arrays of microtissues embedded within an extracellular matrix. Advantages of this method include its high resolution, ability to embed cells into a three-dimensional microenvironment without disrupting the micropattern, and flexibility in patterning any cell type.

Here we present a protocol to micropattern cells at single-cell resolution using DNA-programmed adhesion. This protocol uses a benchtop photolithography platform to create patterns of DNA oligonucleotides on a glass slide and then labels cell membranes with commercially available complementary oligonucleotides. Hybridization of the oligos results in programmed cell adhesion.

The relative positioning of cells is a key feature of the microenvironment that organizes cell-cell interactions. To study the interactions between cells ...

Fibroblast Derived Human Engineered Connective Tissue for Screening Applications

Fibroblasts are phenotypically highly dynamic cells, which quickly transdifferentiate into myofibroblasts in response to biochemical and biomechanical stimuli. The current understanding of fibrotic processes, including cardiac fibrosis, remains poor, which hampers the development of new anti-fibrotic therapies. Controllable and reliable human model systems are crucial for a better understanding of fibrosis pathology. This is a highly reproducible and scalable protocol to generate engineered connective tissues (ECT) in a 48-well casting plate to facilitate studies of fibroblasts and the pathophysiology of fibrotic tissue in a 3-dimensional (3D) environment. ECT are generated around the poles with tunable stiffness, allowing for studies under a defined biomechanical load. Under the defined loading conditions, phenotypic adaptations controlled by cell-matrix interactions can be studied. Parallel testing is feasible in the 48-well format with the opportunity for the time-course analysis of multiple parameters, such as tissue compaction and contraction against the load. From these parameters, biomechanical properties such as tissue stiffness and elasticity can be studied.

Presented here is a protocol to generate engineered connective tissues for a parallel culture of 48 tissues in a multi-well plate with double poles, suitable for mechanistic studies, disease modeling, and screening applications. The protocol is compatible with fibroblasts from different organs and species and is exemplified here with human primary cardiac fibroblasts.

Fibroblasts are phenotypically highly dynamic cells, which quickly transdifferentiate into myofibroblasts in response to biochemical and biomechanical ...