We gratefully acknowledge Dr. Erica Stults (Academic Research &#38; Application Scientist, WPI Information Technology Services) for her assistance with 3D printing, Amanda Zo&#235; Reidinger, Ph.D., Chris Nycz, and Karen Levi, M.E., for their input on mold design, Kathy Suqui and Jennifer Mann for their assistance testing mold designs, and Michael O&#8217;Keefe for his assistance with filming.&#160; This work was supported by NSF IGERT DGE 1144804 (MWR, HAS), NIH R15 HL097332 (MWR, TAH), NSF REU EEC0754996 (BA), NIH 1R01 EB023907 (MWR, HAS) and NIH R15 HL137197 (MWR, HAS).
<div></div>

1. Preparing the 3D-printed Mold
<ol>
	<li>Prepare a CAD drawing of the mold in the desired dimensions.</li>
	<li>Send the CAD file to a high-resolution 3D printer. Select the appropriate plastic printing material, with glossy finish.</li>
	<li>After printing, thoroughly wash the mold with detergent and water.</li>
</ol>
2. Casting PDMS Negatives
<ol>
	<li>Measure the desired amount of PDMS base on a balance. For a mold with a 60 mm diameter and 2 mm well posts, use 25 g.</li>
	<li>Add curing agent at a 1:10 (w/w) ratio to the PDMS base.</li>
	<li>Stir vigorously until the two components are thoroughly combined. Insufficient mixing may result in incomplete curing of PDMS, resulting in a &#34;tacky&#34; surface.</li>
	<li>Place a piece of laboratory tape around the border of 3D-printed molds, to enable the formation of a PDMS base layer above the printed wells.</li>
	<li>Pour the PDMS mixture into the mold and place the mold into a vacuum chamber to de-gas until all bubbles are released.</li>
	<li>Cure the PDMS at 50 &#176;C for 2&#8211;4 h, or until solidified enough to remove.</li>
	<li>Remove the lab tape and carefully extract the PDMS negative.</li>
	<li>Incubate the PDMS at 60 &#176;C for an additional 1 h (after removal from the 3D-printed mold) to ensure that mold is fully cured.</li>
	<li>Wash the PDMS thoroughly with detergent and water to remove any residue. Insufficient washing may result in poor ring formation for the first uses of the PDMS negative.</li>
</ol>
3. Fabricating Agarose Wells
<ol>
	<li>Prepare agarose wells 1 day prior to seeding cells.</li>
	<li>Make a 2% (w/v) agarose solution in DMEM and autoclave.</li>
	<li>Pipet 4 mL molten agarose into an autoclaved PDMS negative. Then pipet agarose directly into the posts of PDMS negatives. Remove any air bubbles with a pipet tip, as bubbles may cause inhomogeneities in the well dimensions.</li>
	<li>NOTE: Do not overfill molds; the top surface of the agarose must be flat, so that agarose wells will be level upon removal from PDMS and placement into 6-well culture plates. Leftover agarose may be re-autoclaved and used again, although not more than once.</li>
	<li>After the agarose cools (approximately 10 min for 4 mL agarose; larger molds may need longer cooling times), carefully separate the agarose wells from the PDMS negatives using blunt forceps and transfer into a well of a 6-well plate.</li>
	<li>Submerge the agarose wells in complete culture medium, and equilibrate overnight in a 37 &#176;C incubator prior to use.</li>
</ol>
4. Seeding Tissue Rings
<ol>
	<li>Culture rat aortic smooth muscle cells21 (RaSMCs) in DMEM containing 10% fetal bovine serum (FBS), 1% L-glutamine, 1% non-essential amino acids, 1% sodium pyruvate, and 1% penicillin-streptomycin until they reach 70% confluence. 
	NOTE: Cells should be cultured at 5% CO2 and 37 &#176;C in 15 cm tissue culture plates.</li>
	<li>After molds are equilibrated (section 3), prepare RaSMCs for seeding.</li>
	<li>Rinse plates twice with 5 mL per plate of phosphate buffered saline (PBS).</li>
	<li>Add 3 mL 0.25% trypsin per 15 cm Petri dish. Move the plates to a 37 &#176;C incubator for 2&#8211;3 min, or until cells have lifted off the plate.</li>
	<li>Neutralize trypsin with an equal volume (3 mL per plate) of complete culture medium. Resuspend the cells thoroughly to break up clumps.</li>
	<li>Dilute an aliquot of cell suspension 1:1 with trypan blue dye, and count the cells with a hemocytometer.</li>
	<li>Centrifuge the total volume of cell suspension for 5 min at 200 x g to pellet cells.</li>
	<li>Resuspend cells at a concentration of 10 million cells per mL; this will result in 500,000 cells per 50 &#181;L. 
	NOTE: Different cell types may require different cell concentrations.</li>
	<li>Aspirate all medium from the agarose mold. Be careful to remove all medium from individual wells, but to not puncture the bottoms of the wells.</li>
	<li>Pipet 50 &#181;L of suspension into each well.</li>
	<li>Carefully add 2 mL of fresh medium around the outside of the agarose mold. Be careful not to let medium overflow into the wells of the agarose. Place plates in the incubator overnight (approximately 16 h).</li>
	<li>After overnight incubation, aspirate the medium from outside of the molds, and add 4.5 mL fresh medium to each well&#160;of the 6-well plate so that molds and rings are completely submerged. Change medium daily.</li>
</ol>

<ol>
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E., Forgacs, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Scaffold-free+vascular+tissue+engineering+using+bioprinting.">Scaffold-free vascular tissue engineering using bioprinting.</a> Biomaterials. 30 (30), 5910-5917 (2009).</li><li>Tan, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=3D+printing+facilitated+scaffold-free+tissue+unit+fabrication.">3D printing facilitated scaffold-free tissue unit fabrication.</a> Biofabrication. 6 (2), 1-11 (2014).</li><li>Dean, D. M., Napolitano, A. P., Youssef, J., Morgan, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rods,+tori,+and+honeycombs:+the+directed+self-assembly+of+microtissues+with+prescribed+microscale+geometries.">Rods, tori, and honeycombs: the directed self-assembly of microtissues with prescribed microscale geometries.</a> The FASEB Journal. 21 (14), 4005-4012 (2007).</li><li>Gwyther, T. A., Hu, J. Z., Billiar, K. L., Rolle, M. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Directed+cellular+self-assembly+to+fabricate+cell-derived+tissue+rings+for+biomechanical+analysis+and+tissue+engineering.">Directed cellular self-assembly to fabricate cell-derived tissue rings for biomechanical analysis and tissue engineering.</a> J Vis Exp. (57), e3366 (2011).</li><li>Zhu, W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=3D+printing+of+functional+biomaterials+for+tissue+engineering.">3D printing of functional biomaterials for tissue engineering.</a> Curr Opin Biotechnol. 40, 103-112 (2016).</li><li>Jakab, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tissue+engineering+by+self-assembly+and+bio-printing+of+living+cells.">Tissue engineering by self-assembly and bio-printing of living cells.</a> Biofabrication. 2 (2), 022001 (2010).</li><li>Lemire, J. M., Potter-Perigo, S., Hall, K. L., Wight, T. N., Schwartz, S. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Distinct+Rat+Aortic+Smooth+Muscle+Cells+Differ+in+Versican/PG-M+Expression.">Distinct Rat Aortic Smooth Muscle Cells Differ in Versican/PG-M Expression.</a> Arteriosclerosis, Thrombosis, and Vascular Biology. 16 (6), 821-829 (1996).</li><li>Strobel, H. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cellular+self-assembly+with+microsphere+incorporation+for+growth+factor+delivery+within+engineered+vascular+tissue+rings.">Cellular self-assembly with microsphere incorporation for growth factor delivery within engineered vascular tissue rings.</a> Tissue Eng Part A. 23 (3-4), 143-155 (2017).</li><li>Strobel, H. A., Calamari, E. L., Beliveau, A., Jain, A., Rolle, M. 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The authors have nothing to disclose.

Here we have presented a versatile method for the rapid fabrication of self-assembled tissue rings with easily customized dimensions using 3D printing. Our method is similar to that reported in Svoronos et al.6, where 3D-printed honeycomb and dog-bone shaped wax molds were used to cast PDMS negatives. However, the molds have been modified to contain several unique design features. A notch (Figure 4A(1)) provides orientation of the mold to allow each ring to be labeled and monitored individually. The central hole (Figure 4A(2)) helps improve diffusion of medium into the wells. CAD file numbers are printed directly on the mold; therefore, PDMS negatives are each labeled with version number and post diameter (Figure 4A3). The tapered outer walls (Figure 3(1), 5 &#176;), at the top of the well troughs (Figure 3(2), 45 &#176;), and central hole (Figure 3(3), 5 &#176;) make it easier to remove PDMS negatives from the 3D-printed molds, and agarose wells are easier to remove from the PDMS negatives (Figure 4A(2), A(4)).
We have demonstrated the versatility of this system by fabricating self-assembled ring-shaped tissues of a variety of diameters and cell types, including primary human smooth muscle cells (SMCs)18,22, rat aortic SMCs7,23, human mesenchymal stem cells (hMSCs)13, and SMCs derived from induced pluripotent stem cells (iPSCs)11 (Table 1). In ongoing work, we are evaluating the formation of rings from additional cell types such as endothelial cells, and fusing cartilage rings of varying sizes for potential applications in tracheal replacement. In addition to completely cell-derived constructs, we have also used this system to fabricate rings with incorporated cross-linked gelatin microspheres13,22. Microspheres can be incorporated within tissue rings during self-assembly to provide additional mechanical strength, or for localized delivery of growth factors13,22.
When fabricating tissue rings, optimization of cell number may be required for different cell types. Minimum cell numbers may vary based on the size and type of cells. For example, hSMCs derived from iPSCs are seeded at 600,000 cells/ring11, hMSCs and primary hSMCs are seeded at 400,000 cells/ring13,22, and rat aortic SMCs are seeded at 500,000 cells/ring18. Trough dimensions may also affect ring formation and the minimum number of cells required for ring formation24. For studies with human cells and 3D-printed molds, a trough width of 2 mm was used. The original polycarbonate molds had a trough width of 3.75 mm, which required 750,000 hSMCs to form a 2 mm cell ring18. With the reduced trough width, we were able to reduce the number of cells necessary for ring formation by 46%, to 400,000 cells per ring25. Quantities of cells seeded per ring are summarized in Table 1.
When choosing a 3D-printed material, many factors need to be considered. Because PDMS is typically cured at 60 &#176;C, the 3D-printed material must have a high enough melting temperature to avoid damage during PDMS curing. The melting temperature of the material used in this study (a proprietary material, see Table of Materials) is not available. However, when baked at 60 &#176;C for 1 h, we observed that the material began to produce an odor. Thus, we decided to lower the curing temperature to 50 &#176;C and increase the curing time in order to bake the PDMS without damaging the 3D-printed material. Adjustments in curing time may be necessary if molds are modified to form larger PDMS negatives. An additional curing period at 60 &#176;C after PDMS removal from the 3D-printed molds prevents the final PDMS negative from remaining tacky, while limiting the temperature the 3D-printed mold is exposed to. Note that some materials inhibit curing of PDMS, so ensure that the selected material is compatible with PDMS. Finally, the mold material toxicity must also be considered. While the 3D-printed mold will not be in direct contact with cells, it is possible that some residue from the mold may be transferred to the PDMS negative during the curing procedure. We found that very thorough washing with detergent was sufficient to remove any residue from the PDMS negative. However, we previously observed that inadequate washing led to poor ring formation in agarose wells for the first few uses of the PDMS negative. The use of PDMS cast from other 3D-printed materials may require additional investigation to verify that detergent is sufficient to remove mold residues, including any potential leachates. Periodic testing may also be necessary, as it possible that repeated heating cycles (even to 50 &#176;C) may damage the mold over time, and cause increases in residue after repeated use. To date, we have used a single 3D-printed mold to produce more than 30 PDMS negatives that have been used to successfully generate tissue rings.
Overall, 3D printing allows greater versatility for the fabrication of agarose molds than machining of polycarbonate. It provides a higher resolution than is possible with tooling, and mold design is not limited by the dimensions of the tools available. This allows for greater customization, and the addition of features such as tapering that may not be possible with machining. This system may be applied to fabricating self-assembled tissues in other shapes as well, in addition to rings6,17. Using the ring fabrication method, we have developed tissue rings from a variety of cell types and sizes for potential applications in tracheal tissue engineering13, engineered blood vessels7, and modeling vascular diseases11.

Cellular self-assembly approaches to fabricating tissue engineered blood vessels are an alternative to scaffold-based approaches. Self-assembled, scaffold-free tissues may have greater cell density, enhanced matrix deposition and strength, and improved biological function compared to scaffold-based tissues1,2,3,4. However, forming 3D tissues without the use of exogenous scaffold support with specific sizes and shapes remains a challenge. Some methods fuse together layers of cell sheets to form thicker constructs, although this process can be time consuming and labor intensive5. Alternatively, cells can be seeded into non-adhesive molds and allowed to aggregate into spheroids, rings, and other tissue shapes6,7,8.
Self-assembled tissue rings require fewer cells, shorter culture times, and less reagents than larger tubular engineered tissues, but can still be mechanically tested, examined histologically, or used for contractility and other functional testing7,9,10,11. Because they can be rapidly fabricated and easily tested, tissue rings are ideal for screening large numbers of culture parameters, and have potential for use as disease models11 or tools for drug screening12. Additionally, rings can be fused into more complex tissue structures such as blood vessels or trachea7,13, and rings may fuse more completely than other shapes such as spheroids14,15.
Agarose is widely used as a mold material for fabricating self-assembled tissues due to its biocompatibility, permeability, and non-cell adhesive properties. For example, Norotte et al. fabricated agarose molds from extruded rods, which enabled limited control over mold shape and required specialized equipment15. Tan et al. deposited alginate droplets as building units to fabricate custom hydrogel molds in various shapes (pyramid, square)16. However, the large diameter of the alginate spheroids (300 &#181;m) resulted in features with low resolution. Such a low resolution may result in uneven mold surfaces that can adversely affect cell aggregation consistency. Alternatively, agarose can be cast into polymer negatives to create non-adhesive molds with smooth features and specific dimensions6,7,17.
We previously reported a system for fabricating custom annular agarose cell-seeding wells from a PDMS negative cast in a milled polycarbonate mold7,18. Agarose was poured into the PDMS negative and allowed to set7,18. Cells were then seeded into agarose wells, where they aggregated to form self-assembled, scaffold-free tissue rings in less than 24 h7,18. PDMS negatives are autoclavable, can be reused many times, and are soft and flexible, making it easy to remove the solidified agarose wells. When this system was initially reported in Gwyther et al.7, PDMS negatives were cast from milled polycarbonate molds (Figure 1A). After agarose casting, the cell seeding wells were individually cut out and placed into wells of a 12-well plate7,18. The design was more recently modified such that a single agarose mold produces 5 rings and fits in a well of a 6-well plate, eliminating the need to cut out individual wells and reducing the amount of PDMS and agarose required to produce each ring (Figure 1B). A smaller cell seeding trough width was used to reduce the number of seeded cells required to achieve ring formation. Despite these changes, the resolution and customization of molds were restricted to available standard endmill dimensions, and micromilling can be prohibitively expensive. Additionally, computer numerical control (CNC) machining can be time consuming and cumbersome due to the need to reserve time on heavily utilized custom equipment, additional computer-aided manufacturing (CAM) software to convert the computer-aided design (CAD) file to a programmable tool path, and reliable fixturing of the polycarbonate part during machining.
In the present study, we examined the use of 3D printing as an alternative to CNC machining. 3D printing is widely used for engineering custom implants, fabricating scaffold materials, and for direct printing of cells and tissue spheroids15,19,20. We used a high-resolution 3D printer, and specialized 3D printing material that enabled us to print a rigid mold with a smooth, glossy surface finish (see Table of Materials). Our technique allows for fabrication of highly customizable, high-resolution plastic molds that can be used for casting PDMS negatives and agarose wells. Design iterations are summarized in Figure 1. The mold design was further modified in the 3D-printed mold version to include tapered outer walls and a center hole in order to ease removal of both the PDMS negatives from 3D-printed molds and the agarose wells from PDMS negatives. These tapered features cannot be achieved with standard machining processes. The distance from the bottom of the wells to the bottom of the mold was increased in this iteration, resulting in a thicker agarose base below the posts to reduce the risk of posts breaking during agarose well removal. The mold and ring fabrication procedure is shown schematically in Figure 2.

<table>
	<tbody>
		<tr>
			<td>SeaKem LE Agarose</td>
			<td>Lonza</td>
			<td>50040</td>
			<td></td>
		</tr>
		<tr>
			<td>PDMS</td>
			<td>Dow Corning</td>
			<td>Sylgard 184</td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM</td>
			<td>Corning Cellgro</td>
			<td>15-017-CV</td>
			<td></td>
		</tr>
		<tr>
			<td>VeroWhite</td>
			<td>StrataSys</td>
			<td>RGD835</td>
			<td></td>
		</tr>
		<tr>
			<td>3D printer</td>
			<td>StrataSys</td>
			<td>Objet 260 Connex</td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM</td>
			<td>Corning Cellgro</td>
			<td>15-017-CV</td>
			<td></td>
		</tr>
		<tr>
			<td>FBS</td>
			<td>Thermo Fisher</td>
			<td>16000069</td>
			<td></td>
		</tr>
		<tr>
			<td>L-glutamine</td>
			<td>Corning Cellgro</td>
			<td>25-015-CI</td>
			<td></td>
		</tr>
		<tr>
			<td>Non-essential amino acids</td>
			<td>Corning Cellgro</td>
			<td>25-025-CI</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium pyruvate</td>
			<td>Corning Cellgro</td>
			<td>25-000-CI</td>
			<td></td>
		</tr>
		<tr>
			<td>Pen-strep</td>
			<td>Corning Cellgro</td>
			<td>30-002-CI</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypsin</td>
			<td>Corning Cellgro</td>
			<td>25-053-CI</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypan blue</td>
			<td>Corning Cellgro</td>
			<td>25-900-CI</td>
			<td></td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>Lonza</td>
			<td>17-516F</td>
			<td></td>
		</tr>
		<tr>
			<td>6-well plate</td>
			<td>Corning</td>
			<td>353046</td>
			<td></td>
		</tr>
		<tr>
			<td>WKY 3M-22 rat aortic smooth muscle cells</td>
			<td>Provided by T. Wight [ref 21]</td>
			<td>N/A</td>
			<td></td>
		</tr>
	</tbody>
</table>

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 system allows for the simple, customized fabrication of agarose cell seeding wells that enables cell self-assembly of ring-shaped 3D engineered tissues. 3D printing allows better resolution and greater flexibility in mold design than machining polycarbonate, where dimensions are constrained by the available tool sizes. With a high-resolution 3D printer, walls as thin as 0.254 mm can be printed, and trough dimensions are limited only by the resolution of the printer (15.2 &#181;m resolution for these studies). Typically, CNC endmills less than 0.3 mm are not commercially available, so it is not possible to create troughs of smaller widths. Micro-milling can produce smaller features, although the equipment required may be prohibitively expensive or inaccessible to many biomedical research labs. Trough dimensions and curvature are also limited by the endmill tip shape. Additionally, features such as angled or tapered walls are not possible, and small features may break during the machining process.
CAD drawings can be easily modified to produce 3D-printed molds and PDMS negatives of customizable dimensions. Figure 3 describes the dimensions of our current 2 mm post 3D-printed molds, compared to previous design iterations. The process is inexpensive; the 2 mm, 5-ring 3D-printed mold cost 44.67 USD to 3D print, and each part can be used to create multiple PDMS negatives. To date, we have created more than 30 PDMS negatives from a single 3D-printed mold. Each negative requires 25 g of PDMS at 0.11 USD per g (2.75 USD per PDMS negative). Each PDMS negative can be cleaned with detergent, autoclaved, and re-used for up to several years, depending on frequency of use. Compared to the original design18, the compact 3D-printed mold uses considerably less PDMS and agarose per 2 mm tissue ring (Figure 1E).
Cells seeded in the equilibrated agarose wells aggregate to form tissue rings in less than 24 h. Ring dimensions depend on the dimensions of the agarose wells. Here, we demonstrated that self-assembled rat smooth muscle cell rings can be fabricated in wells with 2, 4, or 12 mm diameter posts (Figure 4). While rings in this study were only cultured for 3 days, we have previously cultured hSMC rings for up to 2 weeks22, and hMSC-cartilage rings for up to 3 weeks13. As reported previously, rings can function as 3D in vitro models of human tissues for quantitative assessment of tissue function11 and mechanical strength13,22, and can also serve as modular building units to generate tube-shaped tissue constructs7,13. Cell seeding conditions and functional attributes of tissue rings engineered from these cell types are summarized in Table 1.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56618/56618fig1.jpg" /> 
Figure 1: Polycarbonate mold designs. Initially, 15-well molds were machined in polycarbonate (A). Later versions featured a more compact design (B), with 5 wells designed to fit in a single polycarbonate mold, and within one well of a 6-well plate. Here, we modified this design to use 3D-printed plastic as a more customizable alternative to machined polycarbonate (C). Shown in (D) are agarose wells fabricated from the initial design18 (left) compared to the current compact design (right), which requires significantly less agarose, and does not require manual separation of wells. Quantities of PDMS and agarose required for each design iteration are shown in (E). Center posts are 2 mm in diameter. Mold in (A) is 6 cm x 12 cm, (B) has a 5 cm diameter, and (C) has a 6 cm diameter. Scale bar in (D) = 3 cm. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56618/56618fig1large.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/56618/56618fig2.jpg" /> 
Figure 2: Fabrication of self-assembled tissue rings. A 3D-printed mold is used to cast a PDMS negative, which is then used to cast the agarose wells (A). Cells are then seeded directly into the agarose wells, where they aggregate in less than 24 h to form tissue rings (B). Dashed lines in (B) show the well outline. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56618/56618fig2large.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/56618/56618fig3.jpg" /> 
Figure 3: Cross-sectional view of 3D-printed mold CAD drawing. Dimensions for trough width (A), trough height (B), center hole (C), total diameter (D), outer lip (E), and outer wall height (F) are shown. The outer walls (1), upper portion of the wells (2), and center hole (3) are tapered (at a 5 &#176; angle for 1 and 3, 45 &#176; for 2) to improve ease of PDMS negative removal. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56618/56618fig3large.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/56618/56618fig4.jpg" /> 
Figure 4: CAD drawings with corresponding PDMS negatives and self-assembled SMC rings with 2 (A), 4 (B), and 12 (C) mm post diameters. In (A), (1) indicates a notch for providing orientation, (2) indicates a central hole to improve perfusion, (3) is a file number, which imprints directly onto the PDMS negatives, and (4) shows the outer wall, which is tapered to ease PDMS negative removal. Scale bars in agarose wells = 1 cm. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56618/56618fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Cell type</td>
			<td>Cells seeded per ring</td>
			<td>Ring diameter</td>
			<td>Culture Duration</td>
			<td>Functional analysis performed</td>
		</tr>
		<tr>
			<td>Rat SMC</td>
			<td>0.5, 1, or 3 x 106</td>
			<td>2, 4, or 12 mm</td>
			<td>3 days</td>
			<td>Current study; N/A</td>
		</tr>
		<tr>
			<td>iPSC derived SMC11</td>
			<td>0.6 x 106</td>
			<td>2 mm</td>
			<td>17 days</td>
			<td>Contractility testing, uniaxial tensile test, histological analysis</td>
		</tr>
		<tr>
			<td>Human SMC22</td>
			<td>0.4 x 106</td>
			<td>2 mm</td>
			<td>2, 7, or 14 days</td>
			<td>Uniaxial tensile test, histological analysis</td>
		</tr>
		<tr>
			<td>Human MSC14</td>
			<td>0.4 x 106</td>
			<td>2 mm</td>
			<td>21 days</td>
			<td>Uniaxial tensile test, histological analysis, ring fusion and tube compression testing, differentiation into cartilage tissue</td>
		</tr>
	</tbody>
</table>
Table 1: Cell numbers required for ring fabrication from different cell types.

Watch this Scientific Journal Video about Fabrication of Custom Agarose Wells for Cell Seeding and Tissue Ring Self-assembly Using 3D-Printed Molds at JoVE.com

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

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 ...

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Research

JoVE Journal

Bioengineering

11.7K Views.  Worcester Polytechnic Institute. 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.

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

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.

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 ...

Non-contact, Label-free Monitoring of Cells and Extracellular Matrix using Raman Spectroscopy

Non-destructive, non-contact and label-free technologies to monitor cell and tissue cultures are needed in the field of biomedical research.1-5 However, currently available routine methods require processing steps and alter sample integrity. Raman spectroscopy is a fast method that enables the measurement of biological samples without the need for further processing steps. This laser-based technology detects the inelastic scattering of monochromatic light.6 As every chemical vibration is assigned to a specific Raman band (wavenumber in cm-1), each biological sample features a typical spectral pattern due to their inherent biochemical composition.7-9 Within Raman spectra, the peak intensities correlate with the amount of the present molecular bonds.1 Similarities and differences of the spectral data sets can be detected by employing a multivariate analysis (e.g. principal component analysis (PCA)).10
Here, we perform Raman spectroscopy of living cells and native tissues. Cells are either seeded on glass bottom dishes or kept in suspension under normal cell culture conditions (37 &deg;C, 5% CO2) before measurement. Native tissues are dissected and stored in phosphate buffered saline (PBS) at 4 &deg;C prior measurements. Depending on our experimental set up, we then either focused on the cell nucleus or extracellular matrix (ECM) proteins such as elastin and collagen. For all studies, a minimum of 30 cells or 30 random points of interest within the ECM are measured. Data processing steps included background subtraction and normalization.

Raman spectroscopy is a suitable technique for the non-contact, label-free analysis of living cells, tissue-engineered constructs and native tissues. Source-specific spectral fingerprints can be generated and analyzed using multivariate analysis.

Non-destructive, non-contact and label-free technologies to monitor cell and tissue cultures are needed in the field of biomedical research.1-5 However, ...

Human Cartilage Tissue Fabrication Using Three-dimensional Inkjet Printing Technology

Bioprinting, which is based on thermal inkjet printing, is one of the most attractive enabling technologies in the field of tissue engineering and regenerative medicine. With digital control&#160;cells, scaffolds, and growth factors can be precisely deposited to the desired two-dimensional (2D) and three-dimensional (3D) locations rapidly. Therefore, this technology is an ideal approach to fabricate tissues mimicking their native anatomic structures. In order to engineer cartilage with native zonal organization, extracellular matrix composition (ECM), and mechanical properties, we developed a bioprinting platform using a commercial inkjet printer with simultaneous photopolymerization capable for 3D cartilage tissue engineering. Human chondrocytes suspended in poly(ethylene glycol) diacrylate (PEGDA) were printed for 3D neocartilage construction via layer-by-layer assembly. The printed cells were fixed at their original deposited positions, supported by the surrounding scaffold in simultaneous photopolymerization. The mechanical properties of the printed tissue were similar to the native cartilage. Compared to conventional tissue fabrication, which requires longer UV exposure, the viability of the printed cells with simultaneous photopolymerization was significantly higher. Printed neocartilage demonstrated excellent glycosaminoglycan (GAG) and collagen type II production, which was consistent with gene expression. Therefore, this platform is ideal for accurate cell distribution and arrangement for anatomic tissue engineering.

The methods described in this paper show how to convert a commercial inkjet printer into a bioprinter with simultaneous UV polymerization. The printer is capable of constructing 3D tissue structure with cells and biomaterials. The study demonstrated here constructed a 3D neocartilage.

Bioprinting, which is based on thermal inkjet printing, is one of the most attractive enabling technologies in the field of tissue engineering and ...

Rapid and Low-cost Prototyping of Medical Devices Using 3D Printed Molds for Liquid Injection Molding

Biologically inert elastomers such as silicone are favorable materials for medical device fabrication, but forming and curing these elastomers using traditional liquid injection molding processes can be an expensive process due to tooling and equipment costs. As a result, it has traditionally been impractical to use liquid injection molding for low-cost, rapid prototyping applications. We have devised a method for rapid and low-cost production of liquid elastomer injection molded devices that utilizes fused deposition modeling 3D printers for mold design and a modified desiccator as an injection system. Low costs and rapid turnaround time in this technique lower the barrier to iteratively designing and prototyping complex elastomer devices. Furthermore, CAD models developed in this process can be later adapted for metal mold tooling design, enabling an easy transition to a traditional injection molding process. We have used this technique to manufacture intravaginal probes involving complex geometries, as well as overmolding over metal parts, using tools commonly available within an academic research laboratory. However, this technique can be easily adapted to create liquid injection molded devices for many other applications.

We have devised a method for low-cost and rapid prototyping of liquid elastomer rubber injection molded devices by using fused deposition modeling 3D printers for mold design and a modified desiccator as a liquid injection system.

Biologically inert elastomers such as silicone are favorable materials for medical device fabrication, but forming and curing these elastomers using ...

Synthesis of Cationized Magnetoferritin for Ultra-fast Magnetization of Cells

Many important biomedical applications, such as cell imaging and remote manipulation, can be achieved by labeling cells with superparamagnetic iron oxide nanoparticles (SPIONs). Achieving sufficient cellular uptake of SPIONs is a challenge that has traditionally been met by exposing cells to elevated concentrations of SPIONs or by prolonging exposure times (up to 72 hr). However, these strategies are likely to mediate toxicity. Here, we present the synthesis of the protein-based SPION magnetoferritin as well as a facile surface functionalization protocol that enables rapid cell magnetization using low exposure concentrations. The SPION core of magnetoferritin consists of cobalt-doped iron oxide with an average particle diameter of 8.2 nm mineralized inside the cavity of horse spleen apo-ferritin. Chemical cationization of magnetoferritin produced a novel, highly membrane-active SPION that magnetized human mesenchymal stem cells (hMSCs) using incubation times as short as one minute and iron concentrations as lows as 0.2 mM.

A protocol for the synthesis and cationization of cobalt-doped magnetoferritin is presented, as well as a method to rapidly magnetize stem cells with cationized magnetoferritin.

Many important biomedical applications, such as cell imaging and remote manipulation, can be achieved by labeling cells with superparamagnetic iron oxide ...

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, ...

Fabrication of Extracellular Matrix-derived Foams and Microcarriers as Tissue-specific Cell Culture and Delivery Platforms

Cell function is mediated by interactions with the extracellular matrix (ECM), which has complex tissue-specific composition and architecture. The focus of this article is on the methods for fabricating ECM-derived porous foams and microcarriers for use as biologically-relevant substrates in advanced 3D in vitro cell culture models or as pro-regenerative scaffolds and cell delivery systems for tissue engineering and regenerative medicine. Using decellularized tissues or purified insoluble collagen as a starting material, the techniques can be applied to synthesize a broad array of tissue-specific bioscaffolds with customizable geometries. The approach involves mechanical processing and mild enzymatic digestion to yield an ECM suspension that is used to fabricate the three-dimensional foams or microcarriers through controlled freezing and lyophilization procedures. These pure ECM-derived scaffolds are highly porous, yet stable without the need for chemical crosslinking agents or other additives that may negatively impact cell function. The scaffold properties can be tuned to some extent by varying factors such as the ECM suspension concentration, mechanical processing methods, or synthesis conditions. In general, the scaffolds are robust and easy to handle, and can be processed as tissues for most standard biological assays, providing a versatile and user-friendly 3D cell culture platform that mimics the native ECM composition. Overall, these straightforward methods for fabricating customized ECM-derived foams and microcarriers may be of interest to both biologists and biomedical engineers as tissue-specific cell-instructive platforms for in vitro and in vivo applications.

The tissue-specific extracellular matrix (ECM) is a key mediator of cell function. This article describes methods for synthesizing pure ECM-derived foams and microcarriers that are stable in culture without the need for chemical crosslinking for applications in advanced 3D in vitro cell culture models or as pro-regenerative bioscaffolds.

Cell function is mediated by interactions with the extracellular matrix (ECM), which has complex tissue-specific composition and architecture. The focus ...

Custom Engineered Tissue Culture Molds from Laser-etched Masters

As the field of tissue engineering has continued to mature, there has been increased interest in a wide range of tissue parameters, including tissue shape. Manipulating tissue shape on the micrometer to centimeter scale can direct cell alignment, alter effective mechanical properties, and address limitations related to nutrient diffusion. In addition, the vessel in which a tissue is prepared can impart mechanical constraints on the tissue, resulting in stress fields that can further influence both the cell and matrix structure. Shaped tissues with highly reproducible dimensions also have utility for in vitro assays in which sample dimensions are critical, such as whole tissue mechanical analysis.
This manuscript describes an alternative fabrication method utilizing negative master molds prepared from laser etched acrylic: these molds perform well with polydimethylsiloxane (PDMS), permit designs with dimensions on the centimeter scale and feature sizes smaller than 25 &#181;m, and can be rapidly designed and fabricated at a low cost and with minimal expertise. The minimal time and cost requirements allow for laser etched molds to be rapidly iterated upon until an optimal design is determined, and to be easily adapted to suit any assay of interest, including those beyond the field of tissue engineering.

Herein we present a rapid, facile, and low-cost method for fabricating custom polydimethylsiloxane molds that can be used for producing hydrogel-based engineered tissues with complex geometries. We additionally describe results from mechanical and histological assessments conducted on engineered cardiac tissues produced using this technique.

As the field of tissue engineering has continued to mature, there has been increased interest in a wide range of tissue parameters, including tissue ...

Agarose-based Tissue Mimicking Optical Phantoms for Diffuse Reflectance Spectroscopy

This protocol describes how to make agarose-based tissue-mimicking phantoms and demonstrates how to determine their optical properties using a conventional optical system with an integrating sphere. Measuring systems for the acquisition of the diffuse reflectance and total transmittance spectra are constructed with a broadband white light source, a light guide, an achromatic lens, an integrating sphere, a sample holder, an optical fiber probe, and a multi-channel spectrometer. An acrylic mold consisting of two rectangular acrylic pieces and a U-shaped acrylic piece is constructed to create an epidermal phantom and a dermal phantom with whole blood. The application of a sodium dithionite (Na2S2O4) solution to the dermal phantom enables the researcher to deoxygenate hemoglobin in red blood cells distributed in the dermal phantom. The inverse Monte Carlo simulation with the diffuse reflectance and total transmittance spectra measured by a spectrometer with an integrating sphere is performed to determine the absorption coefficient spectrum &#181;a(&#955;) and the reduced scattering coefficient spectrum &#181;s&#39;(&#955;) of each layer phantom. A two-layered phantom mimicking the diffuse reflectance of human skin tissue is also demonstrated by piling up the epidermal phantom on the dermal phantom.

Here, we demonstrate how agarose-based tissue-mimicking optical phantoms are made and how their optical properties are determined using a conventional optical system with an integrating sphere.

This protocol describes how to make agarose-based tissue-mimicking phantoms and demonstrates how to determine their optical properties using a ...

Seeding and Implantation of a Biosynthetic Tissue-engineered Tracheal Graft in a Mouse Model

Treatment options for congenital or secondary long segment tracheal defects have historically been limited due to an inability to replace functional tissue. Tissue engineering holds great promise as a potential solution with its ability to integrate cells and signaling molecules into a 3-dimensional scaffold. Recent work with tissue engineered tracheal grafts (TETGs) has seen some success but their translation has been limited by graft stenosis, graft collapse, and delayed epithelialization. In order to investigate the mechanisms driving these issues, we have developed a mouse model for tissue engineered tracheal graft implantation. TETGs were constructed using electrospun polymers polyethylene terephthalate (PET) and polyurethane (PU) in a mixture of PET and PU (20:80 percent weight). Scaffolds were then seeded using bone marrow mononuclear cells isolated from 6-8 week-old C57BL/6 mice by gradient centrifugation. Ten million cells per graft were seeded onto the lumen of the scaffold and allowed to incubate overnight before implantation between the third and seventh tracheal rings. These grafts were able to recapitulate the findings of stenosis and delayed epithelialization as demonstrated by histological analysis and lack of Keratin 5 and Keratin 14 basal epithelial cells on immunofluorescence. This model will serve as a tool for investigating cellular and molecular mechanisms involved in host remodeling.

Graft stenosis poses a critical obstacle in tissue engineered airway replacement. To investigate cellular mechanisms underlying stenosis, we utilize a murine model of tissue engineered tracheal replacement with seeded bone marrow mononuclear cells (BM-MNC). Here, we detail our protocol, including scaffold manufacturing, BM-MNC isolation, graft seeding, and implantation.

Treatment options for congenital or secondary long segment tracheal defects have historically been limited due to an inability to replace functional ...