Funding was provided by the National Institutes of Health (EB 001013), Natural Sciences and Engineering Research Council of Canada and Canadian Institutes of Health Research.  We thank Dr. A. P. McGuigan for her expertise and help in development of modules.

1) Preparation of Tubing
<ol>
<li>	Cut 2 to 3 m lengths of polyethylene tubing (0.76 mm I.D. x 1.22 mm O.D.).</li>
<li>	Thread a 20 G1 needle into one end of the tubing.</li>
<li>	Coil the tubing and place in a convertors self-seal pouch (7 1/2" x 13"). The two ends of the tubing should be placed at the unsealed end of the bag and taped in place with gas sterilization tape.</li>
<li>	Seal bag and gas sterilize.</li>
</ol>
2) Neutralization of Collagen
Note: 1 ml of collagen fills about 2 m of polyethylene tubing.
<ol>
<li> Place the necessary amount of collagen needed for the desired length of tubing into a 15 ml conical tube and keep on ice. </li>
<li> Add 128 uL of 10x &alpha;-MEM per mL of collagen to the tube and mix by repeatedly pipetting the solution. Note: (1) The solution should turn yellow as the &alpha;-MEM is mixed into the acidified collagen and (2) try to minimize bubbles when mixing.</li>
<li> Neutralize the collagen by adding 0.8 M sodium bicarbonate until the collagen has a pH of 7.4. For a general estimate of the pH add sodium bicarbonate until the yellow collagen solution turns a light pink color. Note: It typically takes 30 to 60 &mu;L per each mL of collagen.</li>
<li> Keep neutralized collagen on ice until ready to use as the neutralized collagen will gel if not kept at 4&deg;C.</li>
</ol>
3) Embedding of Cells (Optional)
Note: Depending on the cell type the cells can be embedded at a concentration of between 1 million cells per mL to 20 million cells per mL. Use the medium required for the cells that you want to embed.
<ol>
<li> Remove media from cells and wash with 5 mL PBS.</li>
<li> Remove PBS and add 3 mL trypsin-EDTA.</li>
<li> Incubate the cells in trypsin-EDTA for 5 min at 37&deg;C.</li>
<li> Resuspend cells in 7 mL media.</li>
<li> Count cells.</li>
<li> Transfer the required number of cells for embedding to a 15 mL conical tube.</li>
<li> Pellet the cells at 300 x g for 5 min.</li>
<li> Aspirate media off the cell pellet.</li>
<li> Resuspend the cells in the neutralized collagen and keep on ice.</li>
</ol>
4) Gelling of Collagen
<ol>
<li> In a biological safety cabinet cut off the sealed end of the bag containing the polyethylene tubing.</li>
<li> Attach a 3 ml syringe to the 20G needle in the one end of the tubing using sterilized tweezers. Keep the tubing in the bag.</li>
<li> Insert the other end of the tubing into the neutralized collagen until the tip is at the bottom of the conical tube using sterilized tweezers. Keep the conical tube on ice.</li>
<li> Pull the collagen into the polyethylene tubing by retracting the plunger of the syringe slowly. 
Note: If the collagen is pulled into the tubing to quickly then bubbles will form in the collagen.</li>
<li> After pulling the collagen through the tubing, pull the 20G needle out of tubing and put both ends back into the bag. Tape the bag shut.</li>
<li> Incubate the bag at 37&deg;C for 60 min. If the collagen contains cells then flip the bag over every 15 min. so that the cells do not settle to one side of the module.</li>
</ol>
5) Cutting of Tubing Containing the Gelled Collagen
<ol>
<li> Autoclave headers and blade for the cutting device and tray to hold tubing.</li>
<li> Assemble the cutting device.</li>
<li> Place tubing in a sterile tray in front of cutting device and load the tubing into the 
cutting device.</li>
<li> Set up a 50 mL conical tube containing 25 mL of media at the outlet of the cutting 
device to collect the cut modules. 
Note: Use media containing FBS even for collagen-only modules as the modules do not separate from the tubing without FBS in the media. If the modules contain cells use the media required for the embedded cells. If there are no embedded cells in the modules use the media for the endothelial cells.</li>
<li> Set the cutting device to cut 2 mm lengths of tubing and cut the tubing.</li>
<li> Incubate cut modules at 37&deg;C for 60 min in the 50 mL conical tube.</li>
</ol>
6) Removing Collagen Modules from Tubing
<ol>
<li> Vortex 50 mL conical tube containing modules in the tubing for 10 sec. 
	Note: Be gentle when separating modules embedded with high cell density to avoid breakage. </li>
<li> Let modules settle to the bottom of the tube. Takes about 5 min.</li>
<li> Using a wide mouth 10 mL serological pipette transfer the settled modules to a 15 mL 
conical tube.</li>
<li> Repeat steps 6.1 to 6.3 twice.</li>
<li> Go to step 7 to coat modules with endothelial cells or transfer modules to a non-tissue 
culture treated plate.</li>
</ol>
7) Coating the modules with endothelial cells
<ol>
<li> Settle modules into the bottom of a 15 mL conical tube with 5 mL of media for the embedded 
cells.</li>
<li> Remove media from the endothelial cells and wash with 5 mL PBS.</li>
<li> Remove PBS and add 3 mL trypsin-EDTA.</li>
<li> Incubate the cells in trypsin-EDTA for 5 min at 37&deg;C.</li>
<li> Resuspend cells in 7 mL media.</li>
<li> Count cells. Need 5x106 endothelial cells per mL of packed modules at the bottom of 
the conical tube.</li>
<li> Pellet the cells at 300 x g for 5 min and resuspend in 5 mL endothelial cell media.</li>
<li> Add the 5 mL of the media containing endothelial cells to the modules. 
Note: This gives a 50:50 ratio of the two media types which we have found to work for the survival and function of both cell types. Test the use a co-culture medium formulated to ensure proper maintenance of phenotypes of the two cells types before using it. </li>
<li> Incubate the modules with the endothelial cells both statically and dynamically.</li>
<li> For static incubation mix modules with endothelial cells by inversion and set the tube upright at 37&deg;C for 15 min. Repeat.</li>
<li> Dynamically seed the endothelial cells by gently rocking the tube at 37&deg;C for 60 min.</li>
<li> Repeat static incubation twice.</li>
<li> Place the modules in a non-tissue culture treated plate and incubate overnight at 37&deg;C.</li>
<li> The next day place the modules in a new non-tissue culture treated plate with fresh media (50:50 mix if using co-culture system) to remove unattached endothelial cells.</li>
<li> Incubate modules that are coated with endothelial cells until they cover 100% of the surface of the modules. This usually takes about 7 days.</li>
</ol>
8) Representative Results:
The modules will look cylindrical when they first are removed from the tube. If the modules contain embedded cells and/or are coated with endothelial cells they can contract up to 50% in volume and develop an oval shape (Figure 1). The modules will also become denser and more opaque when viewed by light microscopy (Figure 1). Also when the endothelial cells are confluent on the surface of the modules there is a formation of tight junctions which can be seen by immunofluorescence staining of VE-Cadherin (Figure 2). Mass transfer analysis has demonstrated that modules are capable of supporting high cell densities (8x107 cells/cm3) without developing a necrotic core due to inadequate oxygen transport, which is often problematic in larger tissues (e.g. modules of large diameter, D = 1.4mm) (Figure 3).
<img src="/files/ftp_upload/2177/2177fig1.jpg" alt="Figure 1" /> 
Figure 1: Module fabrication and contraction. Module contraction occurred during the three days following HUVEC-C (endothelial cell line) seeding results significantly smaller module diameter and length (p &lt; 0.001) (A). Embedded HepG2 cells were uniformly distributed within modules at time of fabrication and retained high viability (B). [Live cells green; dead cells red] [adapted from Corstorphine 2010]

<img src="/files/ftp_upload/2177/2177fig2.jpg" alt="Figure 2" /> 
Figure 2: VE-Cadherin immunofluorescent staining of EC tight cell junctions 10 days after RAEC where coated on the surface of a module. Scale bar = 50 &mu;m.

<img src="/files/ftp_upload/2177/2177fig3.jpg" alt="Figure 3" /> 
Figure 3: Masson's trichrome staining of typical modules (0.76 mm initial diameter) and large modules (1.4 mm initial diameter) demonstrate the effect of oxygen diffusion limitations. Seven days following fabrication, a large number of dead cells had formed within the core of the large modules (lower right panel), leaving only a thin rim (~200&mu;m thick) of viable cells. Conversely, the small modules retained a uniform and high distribution of live cells. [Modules embedded with HepG2 cells and coated with endothelial cells.] [Taken from Corstorphine 2010]

<img src="/files/ftp_upload/2177/2177fig4.jpg" alt="Figure 4" /> 
Figure 4: (A) HMEC-1 seeded on poloxamine - collagen modules. After Day 1 of seeding the poloxamine-collagen modules retain their cylindrical shape and there is limited cell attachment properties compared to collagen only modules. Scale bar = 200 &mu;m. (B) Light microscope image of a collagen module containing PLGA-based biodegradable microspheres. Scale bar = 500 &mu;m.

<img src="/files/ftp_upload/2177/2177fig5.jpg" alt="Figure 5" /> 
Figure 5: Examples of in vitro assays. (A) Angiogenesis assay: capillary-like formations on the module seeded with endothelial cells can be easily detected and quantified after 5 days of incubation with adipose-derived stem cells. (B) Confocal microscopy images of a live/dead assay on modules where the staining for live cells is green and for dead cells is red.

<img src="/files/ftp_upload/2177/2177fig6.jpg" alt="Figure 6" /> 
Figure 6: Collagen modules containing embedded mesenchymal stem cells and a surface layer of endothelial cells. Modules were exposed to shear stress (~0.64 dyn/cm2) for 7 days in a microfluidic chamber and begin to show fusion at points of contact. Endothelial cells are stained with vWF (brown) and mesenchymal stem cell nuclei appear blue (H&amp;E). Scale bar = 100&mu;m.

<img src="/files/ftp_upload/2177/2177fig7.jpg" alt="Figure 7" /> 
Figure 7: Hundreds of collagen modules containing Adipose-Derived Stem cells (ASC) and coated with Human Microvascular Endothelial Cells (HMEC) are implanted under mouse skin to study ASC/HMEC interaction in vivo for fat regeneration applications.

<ol>
	<li>Chamberlain, M. D., Gupta, R., Sefton, M. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chimeric+vessel+tissue+engineering+driven+by+endothelialized+modules.">Chimeric vessel tissue engineering driven by endothelialized modules.</a> , (2010).</li><li>Corstorphine, L. E., Sefton, M. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effectiveness+factor+and+diffusion+limitations+in+collagen+gel+modules+containing+HepG2+cells.">Effectiveness factor and diffusion limitations in collagen gel modules containing HepG2 cells.</a> J Tissue Eng Regen Med. , (2010).</li><li>Gupta, R., Van Rooijen, N., Sefton, M. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fate+of+endothelialized+modular+constructs+implanted+in+an+omental+pouch+in+nude+rats.">Fate of endothelialized modular constructs implanted in an omental pouch in nude rats.</a> Tissue Eng. Part A 15, 2875-2887 (2009).</li><li>Leung, B. M., Sefton, M. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+modular+tissue+engineering+construct+containing+smooth+muscle+cells+and+endothelial+cells.">A modular tissue engineering construct containing smooth muscle cells and endothelial cells.</a> Ann Biomed Eng. 35, 2039-2049 (2007).</li><li>McGuigan, A. P., Leung, B., Sefton, M. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fabrication+of+cell-containing+gel+modules+to+assemble+modular+tissue-engineered+constructs+[corrected].">Fabrication of cell-containing gel modules to assemble modular tissue-engineered constructs [corrected].</a> Nat Protoc. 1, 2963-2969 (2006).</li><li>McGuigan, A. P., Sefton, M. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Vascularized+organoid+engineered+by+modular+assembly+enables+blood+perfusion.">Vascularized organoid engineered by modular assembly enables blood perfusion.</a> Proc Natl Acad Sci U S A. 103, 11461-11466 (2006).</li><li>McGuigan, A. P., Sefton, M. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modular+tissue+engineering:+fabrication+of+a+gelatin-based+construct.">Modular tissue engineering: fabrication of a gelatin-based construct.</a> J Tissue Eng Regen Med. 1, 136-145 (2007).</li><li>McGuigan, A. P., Sefton, M. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Design+and+fabrication+of+sub-mm-sized+modules+containing+encapsulated+cells+for+modular+tissue+engineering.">Design and fabrication of sub-mm-sized modules containing encapsulated cells for modular tissue engineering.</a> Tissue Eng. 13, 1069-1078 (2007).</li><li>McGuigan, A. P., Sefton, M. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Design+criteria+for+a+modular+tissue-engineered+construct.">Design criteria for a modular tissue-engineered construct.</a> Tissue Eng. 13, 1079-1089 (2007).</li><li>McGuigan, A. P., Sefton, M. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+thrombogenicity+of+human+umbilical+vein+endothelial+cell+seeded+collagen+modules.">The thrombogenicity of human umbilical vein endothelial cell seeded collagen modules.</a> Biomaterials. 29, 2453-2463 (2008).</li><li>Sosnik, A., Leung, B., McGuigan, A. P., Sefton, M. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Collagen/poloxamine+hydrogels:+cytocompatibility+of+embedded+HepG2+cells+and+surface-attached+endothelial+cells.">Collagen/poloxamine hydrogels: cytocompatibility of embedded HepG2 cells and surface-attached endothelial cells.</a> Tissue Eng. 11, 1807-1816 (2005).</li></ol>

No conflicts of interest declared.

We have fabricated several different micro-tissues using modules embedded with different cell types1-11. We have successfully embedded primary cardiomyocytes, islets, adipose stem cells and mesenchymal stromal cells as well as several cell lines including HepG2, NIH 3T3 and clone 9 liver cells. We have coated the modules with several types of endothelial cells including rat aortic endothelial cells, human umbilical vein endothelial cells and human microvascular endothelial cells. Modules have also been prod...

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		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Intramedic tubing PE60</td>
<td>&nbsp;</td>
<td>Becton Dickson</td>
<td>427416</td>
<td>Different diameter of tubing can be used to change the diameter of the modules</td>
</tr>
<tr>
<td>Phosphate-buffered saline (PBS)</td>
<td>&nbsp;</td>
<td>Gibco</td>
<td>20012-027</td>
<td>&nbsp;</td>
<tr>
<td>Trypsin-EDTA</td>
<td>&nbsp;</td>
<td>Gibco</td>
<td>25200-072</td>
<td>&nbsp;</td>
<tr>
<td>Purcol acidificed collagen, 3 mg/mL</td>
<td>&nbsp;</td>
<td>Cedarlane</td>
<td>5005-B</td>
<td>&nbsp;</td>
<tr>
<td>Sodium bicarbonate</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>S5761</td>
<td>&nbsp;</td>
<tr>
<td>20G needle</td>
<td>&nbsp;</td>
<td>Becton Dickson</td>
<td>305175</td>
<td>Diameter of the needle needs to be similar to the diameter of the tubing</td>
</tr>
<tr>
<td>3 mL syringe</td>
<td>&nbsp;</td>
<td>BD Biosciences</td>
<td>309585</td>
<td>&nbsp;</td>
<tr>
<td>15 mL tube</td>
<td>&nbsp;</td>
<td>BD Biosciences</td>
<td>352096</td>
<td>&nbsp;</td>
<tr>
<td>50 mL tube</td>
<td>&nbsp;</td>
<td>BD Biosciences</td>
<td>352070</td>
<td>&nbsp;</td>
<tr>
<td>Convertors Self-Seal Pouch 7 1/2&#x201D; x 13&#x201D;</td>
<td>&nbsp;</td>
<td>Cardinal Health</td>
<td>92713</td>
<td>&nbsp;</td>
<tr>
<td>10 ml Wide Tip serological pipet</td>
<td>&nbsp;</td>
<td>BD Falcon</td>
<td>357504</td>
<td>&nbsp;</td>
<tr>
<td>10 ml serological pipet</td>
<td>&nbsp;</td>
<td>BD Falcon</td>
<td>357551</td>
<td>&nbsp;</td>
<tr>
<td>5 ml serological pipet</td>
<td>&nbsp;</td>
<td>BD Falcon</td>
<td>357543</td>
<td>&nbsp;</td>		</tbody>
		</table>
		</div>

fabrication of micro-tissues using modules of collagen gel containing cells

This protocol describes the fabrication of a type of micro-tissues called modules.  The module approach generates uniform, scalable and vascularized tissues.  The modules can be made of collagen as well as other gelable or crosslinkable materials.  They are approximately 2 mm in length and 0.7 mm in diameter upon fabrication but shrink in size with embedded cells or when the modules are coated with endothelial cells.  The modules individually are small enough that the embedded cells are within the diffusion limit of oxygen and other nutrients but modules can be packed together to form larger tissues that are perfusable.  These tissues are modular in construction because different cell types can be embedded in or coated on the modules before they are packed together to form complex tissues.  There are three main steps to making the modules: (1) neutralizing the collagen and embedding cells in it, (2) gelling the collagen in the tube and cutting the modules and (3) coating the modules with endothelial cells.

Watch this Scientific Journal Video about Fabrication of Micro-tissues using Modules of Collagen Gel Containing Cells at JoVE.com

Fabrication of Micro-tissues using Modules of Collagen Gel Containing Cells

Creation of micro-tissues using cylindrical collagen gels, called modules, that contain embedded cells and which surface is coated with endothelial cells.

This protocol describes the fabrication of a type of micro-tissues called modules.  The module approach generates uniform, scalable and vascularized ...

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12.6K Views.  University of Toronto. Creation of micro-tissues using cylindrical collagen gels, called modules, that contain embedded cells and which surface is coated with endothelial cells.

Video: Fabrication of Micro-tissues using Modules of Collagen Gel Containing Cells

Fabrication of Micropatterned Hydrogels for Neural Culture Systems using Dynamic Mask Projection Photolithography

Increasingly, patterned cell culture environments are becoming a relevant technique to study cellular characteristics, and many researchers believe in the need for 3D environments to represent in vitro experiments which better mimic in vivo qualities 1-3. Studies in fields such as cancer research 4, neural engineering 5, cardiac physiology 6, and cell-matrix interaction7,8have shown cell behavior differs substantially between traditional monolayer cultures and 3D constructs.
Hydrogels are used as 3D environments because of their variety, versatility and ability to tailor molecular composition through functionalization 9-12. Numerous techniques exist for creation of constructs as cell-supportive matrices, including electrospinning13, elastomer stamps14, inkjet printing15, additive photopatterning16, static photomask projection-lithography17, and dynamic mask microstereolithography18. Unfortunately, these methods involve multiple production steps and/or equipment not readily adaptable to conventional cell and tissue culture methods. The technique employed in this protocol adapts the latter two methods, using a digital micromirror device (DMD) to create dynamic photomasks for crosslinking geometrically specific poly-(ethylene glycol) (PEG) hydrogels, induced through UV initiated free radical polymerization. The resulting "2.5D" structures provide a constrained 3D environment for neural growth. We employ a dual-hydrogel approach, where PEG serves as a cell-restrictive region supplying structure to an otherwise shapeless but cell-permissive self-assembling gel made from either Puramatrix or agarose. The process is a quick simple one step fabrication which is highly reproducible and easily adapted for use with conventional cell culture methods and substrates.
Whole tissue explants, such as embryonic dorsal root ganglia (DRG), can be incorporated into the dual hydrogel constructs for experimental assays such as neurite outgrowth. Additionally, dissociated cells can be encapsulated in the photocrosslinkable or self polymerizing hydrogel, or selectively adhered to the permeable support membrane using cell-restrictive photopatterning. Using the DMD, we created hydrogel constructs up to ~1mm thick, but thin film (&lt;200 &mu;m) PEG structures were limited by oxygen quenching of the free radical polymerization reaction. We subsequently developed a technique utilizing a layer of oil above the polymerization liquid which allowed thin PEG structure polymerization.
In this protocol, we describe the expeditious creation of 3D hydrogel systems for production of microfabricated neural cell and tissue cultures. The dual hydrogel constructs demonstrated herein represent versatile in vitro models that may prove useful for studies in neuroscience involving cell survival, migration, and/or neurite growth and guidance. Moreover, as the protocol can work for many types of hydrogels and cells, the potential applications are both varied and vast.

Simple techniques are described for the rapid production of microfabricated neural culture systems using a digital micromirror device for dynamic mask projection lithography on regular cell culture substrates. These culture systems may be more representative of natural biological architecture, and the techniques described could be adapted for numerous applications.

Increasingly, patterned cell culture environments are becoming a relevant technique to study cellular characteristics, and many researchers believe in the ...

Imaging Denatured Collagen Strands In vivo and Ex vivo via Photo-triggered Hybridization of Caged Collagen Mimetic Peptides

Collagen is a major structural component of the extracellular matrix that supports tissue formation and maintenance. Although collagen remodeling is an integral part of normal tissue renewal, excessive amount of remodeling activity is involved in tumors, arthritis, and many other pathological conditions. During collagen remodeling, the triple helical structure of collagen molecules is disrupted by proteases in the extracellular environment. In addition, collagens present in many histological tissue samples are partially denatured by the fixation and preservation processes. Therefore, these denatured collagen strands can serve as effective targets for biological imaging. We previously developed a caged collagen mimetic peptide (CMP) that can be photo-triggered to hybridize with denatured collagen strands by forming triple helical structure, which is unique to collagens. The overall goals of this procedure are i)&#160;to image denatured collagen strands resulting from normal remodeling activities in vivo, and ii) to visualize collagens in ex vivo tissue sections using the photo-triggered caged CMPs. To achieve effective hybridization and successful in vivo and ex vivo imaging, fluorescently labeled caged CMPs are either photo-activated immediately before intravenous injection, or are directly activated on tissue sections. Normal skeletal collagen remolding in nude mice and collagens in prefixed mouse cornea tissue sections are imaged in this procedure. The imaging method based on the CMP-collagen hybridization technology presented here could lead to deeper understanding of the tissue remodeling process, as well as allow development of new diagnostics for diseases associated with high collagen remodeling activity.

Imaging Denatured Collagen Strands In vivo and Ex vivo via Photo-triggered Hybridization of Caged Collagen Mimetic Peptides

This procedure demonstrates in vivo near IR fluorescence imaging of collagen remodeling activities in mice as well as ex vivo staining of collagens in tissue sections using caged collagen mimetic peptides that can be photo-triggered to hybridize with denatured collagen strands.

Collagen is a major structural component of the extracellular matrix that supports tissue formation and maintenance. Although collagen remodeling is an ...

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 ...

Minced Tissue in Compressed Collagen: A Cell-containing Biotransplant for Single-staged Reconstructive Repair

Conventional techniques for cell expansion and transplantation of autologous cells for tissue engineering purposes can take place in specially equipped human cell culture facilities. These methods include isolation of cells in single cell suspension and several laborious and time-consuming events before transplantation back to the patient. Previous studies suggest that the body itself could be used as a bioreactor for cell expansion and regeneration of tissue in order to minimize ex vivo manipulations of tissues and cells before transplanting to the patient. The aim of this study was to demonstrate a method for tissue harvesting, isolation of continuous epithelium, mincing of the epithelium into small pieces and incorporating them into a three-layered biomaterial. The three-layered biomaterial then served as a delivery vehicle, to allow surgical handling, exchange of nutrition across the transplant, and a controlled degradation. The biomaterial consisted of two outer layers of collagen and a core of a mechanically stable and slowly degradable polymer. The minced epithelium was incorporated into one of the collagen layers before transplantation. By mincing the epithelial tissue into small pieces, the pieces could be spread and thereby the propagation of cells was stimulated. After the initial take of the transplants, cell expansion and reorganization would take place and extracellular matrix mature to allow ingrowth of capillaries and nerves and further maturation of the extracellular matrix. The technique minimizes ex vivo manipulations and allow cell harvesting, preparation of autograft, and transplantation to the patient as a simple one-stage intervention. In the future, tissue expansion could be initiated around a 3D mold inside the body itself, according to the specific needs of the patient. Additionally, the technique could be performed in an ordinary surgical setting without the need for sophisticated cell culturing facilities.

Tissue engineering often includes in vitro expansion in order to create autografts for tissue regeneration. In this study a method for tissue expansion, regeneration, and reconstruction in vivo was developed in order to minimize the processing of cells and biological materials outside the body.

Conventional techniques for cell expansion and transplantation of autologous cells for tissue engineering purposes can take place in specially equipped ...

Preparation and Photoacoustic Analysis of Cellular Vehicles Containing Gold Nanorods

Gold nanorods are attractive for a range of biomedical applications, such as the photothermal ablation and the photoacoustic imaging of cancer, thanks to their intense optical absorbance in the near-infrared window, low cytotoxicity and potential to home into tumors. However, their delivery to tumors still remains an issue. An innovative approach consists of the exploitation of the tropism of tumor-associated macrophages that may be loaded with gold nanorods in vitro. Here, we describe the preparation and the photoacoustic inspection of cellular vehicles containing gold nanorods. PEGylated gold nanorods are modified with quaternary ammonium compounds, in order to achieve a cationic profile. On contact with murine macrophages in ordinary Petri dishes, these particles are found to undergo massive uptake into endocytic vesicles. Then these cells are embedded in biopolymeric hydrogels, which are used to verify that the stability of photoacoustic conversion of the particles is retained in their inclusion into cellular vehicles. We are confident that these results may provide new inspiration for the development of novel strategies to deliver plasmonic particles to tumors.

We show the preparation and address the feasibility of cellular vehicles containing gold nanorods for the photoacoustic imaging of cancer.

Gold nanorods are attractive for a range of biomedical applications, such as the photothermal ablation and the photoacoustic imaging of cancer, thanks to ...

Forming Giant-sized Polymersomes Using Gel-assisted Rehydration

Polymer vesicles, or polymersomes, are being widely explored as synthetic analogs of lipid vesicles based on their stability, robustness, barrier properties, chemical versatility and tunable physical characteristics. Typical methods used to prepare giant-sized (&gt; 4 &#181;m) vesicles, however, are both time and labor intensive, yielding low numbers of intact polymersomes. Here, we present for the first time the use of gel-assisted rehydration for the rapid and high-yielding formation of giant (&gt;4 &#181;m) polymer vesicles (polymersomes). Using this method, polymersomes can be formed from a wide array of rehydration solutions including several different physiologically-compatible buffers and full cell culture media, making them readily useful for biomimicry studies. This technique is also capable of reliably producing polymersomes from different polymer compositions with far better yields and much less difficulty than traditional methods. Polymersome size is readily tunable by altering temperature during rehydration or adding membrane fluidizers to the polymer membrane, generating giant-sized polymersomes (&gt;100 &#181;m).

We present a protocol to rapidly form giant polymer vesicles (pGVs). Briefly, polymer solutions are dehydrated on dried agarose films adhered to coverslips. Rehydration of the polymer films results in rapid formation of pGVs. This method greatly advances the preparation of synthetic giant vesicles for direct applications in biomimetic studies.

Polymer vesicles, or polymersomes, are being widely explored as synthetic analogs of lipid vesicles based on their stability, robustness, barrier ...

Fabrication and Characterization of Optical Tissue Phantoms Containing Macrostructure

The rapid development of new optical imaging techniques is dependent on the availability of low-cost, customizable, and easily reproducible standards. By replicating the imaging environment, costly animal experiments to validate a technique may be circumvented. Predicting and optimizing the performance of in vivo and ex vivo imaging techniques requires testing on samples that are optically similar to tissues of interest. Tissue-mimicking optical phantoms provide a standard for evaluation, characterization, or calibration of an optical system. Homogenous polymer optical tissue phantoms are widely used to mimic the optical properties of a specific tissue type within a narrow spectral range. Layered tissues, such as the epidermis and dermis, can be mimicked by simply stacking these homogenous slab phantoms. However, many in vivo imaging techniques are applied to more spatially complex tissue where three dimensional structures, such as blood vessels, airways, or tissue defects, can affect the performance of the imaging system. 
This protocol describes the fabrication of a tissue-mimicking phantom that incorporates three-dimensional structural complexity using material with optical properties of tissue. Look-up tables provide India ink and titanium dioxide recipes for optical absorption and scattering targets. Methods to characterize and tune the material optical properties are described. The phantom fabrication detailed in this article has an internal branching mock airway void; however, the technique can be broadly applied to other tissue or organ structures.

Optical tissue phantoms are essential tools for calibration and characterization of optical imaging systems and validation of theoretical models. This article details a method for phantom fabrication that includes replication of tissue optical properties and three-dimensional tissue structure.

The rapid development of new optical imaging techniques is dependent on the availability of low-cost, customizable, and easily reproducible standards. By ...

Preparation, Purification, and Use of Fatty Acid-containing Liposomes

Liposomes containing single-chain amphiphiles, particularly fatty acids, exhibit distinct properties compared to those containing diacylphospholipids due to the unique chemical properties of these amphiphiles. In particular, fatty acid liposomes enhance dynamic character, due to the relatively high solubility of single-chain amphiphiles. Similarly, liposomes containing free fatty acids are more sensitive to salt and divalent cations, due to the strong interactions between the carboxylic acid head groups and metal ions. Here we illustrate techniques for preparation, purification, and use of liposomes comprised in part or whole of single chain amphiphiles (e.g., oleic acids).

Liposomes containing single-chain amphiphiles, particularly fatty acids, exhibit distinct properties compared to those containing diacylphospholipids due to the unique chemical properties of single chain amphiphiles. Here we describe techniques for the preparation, purification, and use of liposomes comprised in part or whole of these amphiphiles.

Liposomes containing single-chain amphiphiles, particularly fatty acids, exhibit distinct properties compared to those containing diacylphospholipids due ...

Uptake of New Lipid-coated Nanoparticles Containing Falcarindiol by Human Mesenchymal Stem Cells

Nanoparticles are the focus of an increased interest in drug delivery systems for cancer therapy. Lipid-coated nanoparticles are inspired in structure and size by low-density lipoproteins (LDLs) because cancer cells have an increased need for cholesterol to proliferate, and this has been exploited as a mechanism for delivering anticancer drugs to cancer cells. Moreover, depending on drug chemistry, encapsulating the drug can be advantageous to avoid degradation of the drug during circulation in vivo. Therefore, in this&#160;study, this design is used to fabricate lipid-coated nanoparticles of the anticancer drug falcarindiol, providing a potential new delivery system of falcarindiol in order to stabilize its chemical structure against degradation and improve its uptake by tumors. Falcarindiol nanoparticles, with a phospholipid and cholesterol monolayer encapsulating the purified drug core of the particle, were designed. The lipid monolayer coating consists of 1,2-distearoyl-sn-glycero-3-phosphocholine (DSPC), cholesterol (Chol), and 1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N-[methoxy(polyethylene glycol)-2000] (DSPE PEG 2000) along with the fluorescent label DiI (molar ratios of 43:50:5:2). The nanoparticles are fabricated using the rapid injection method, which is a fast and simple technique to precipitate nanoparticles by good-solvent for anti-solvent exchange. It consists of a rapid injection of an ethanol solution containing the nanoparticle components into an aqueous phase. The size of the fluorescent nanoparticles is measured using dynamic light scattering (DLS) at 74.1 &#177; 6.7 nm. The uptake of the nanoparticles is tested in human mesenchymal stem cells (hMSCs) and imaged using fluorescence and confocal microscopy. The uptake of the nanoparticles is observed in hMSCs, suggesting the potential for such a stable drug delivery system for falcarindiol.

This article describes the encapsulation of falcarindiol in lipid-coated 74 nm nanoparticles. The cellular uptake of the nanoparticles&#160;by human stem cells into lipid droplets is monitored by fluorescent and confocal imaging. Nanoparticles are fabricated by the rapid injection method of solvent shifting, and their size is measured with the dynamic light scattering technique.

Nanoparticles are the focus of an increased interest in drug delivery systems for cancer therapy. Lipid-coated nanoparticles are inspired in structure and ...

Microfluidic Production of Lysolipid-Containing Temperature-Sensitive Liposomes

The presented protocol enables a high-throughput continuous preparation of low temperature-sensitive liposomes (LTSLs), which are capable of loading chemotherapeutic drugs, such as doxorubicin (DOX). To achieve this, an ethanolic lipid mixture and ammonium sulfate solution are injected into a staggered herringbone micromixer (SHM) microfluidic device. The solutions are rapidly mixed by the SHM, providing a homogeneous solvent environment for liposomes self-assembly. Collected liposomes are first annealed, then dialyzed to remove residual ethanol. An ammonium sulfate pH-gradient is established through buffer exchange of the external solution by using size exclusion chromatography. DOX is then remotely loaded into the liposomes with high encapsulation efficiency (&#62; 80%). The liposomes obtained are homogenous in size with Z-average diameter of 100 nm. They are capable of temperature-triggered burst release of encapsulated DOX in the presence of mild hyperthermia (42 &#176;C). Indocyanine green (ICG) can also be co-loaded into the liposomes for near-infrared laser-triggered DOX release. The microfluidic approach ensures high-throughput, reproducible and scalable preparation of LTSLs.

The protocol presents the optimized parameters for preparing thermosensitive liposomes using the staggered herringbone micromixer microfluidics device. This also allows co-encapsulation of doxorubicin and indocyanine green into the liposomes and the photothermal-triggered release of doxorubicin for controlled/triggered drug release.

The presented protocol enables a high-throughput continuous preparation of low temperature-sensitive liposomes (LTSLs), which are capable of loading ...