This work was supported by grants from the National Institutes of Health, including a microphysiological systems consortium grant from the National Center for Advancing Translational Sciences (UH2TR000503), a Ruth L. Kirschstein National Research Service Award Postdoctoral Fellowship (F32DK098905 for WJM) and pathway to independence award (DK095984 for AB) from the National Institute of Diabetes and Digestive and Kidney Diseases.

1. Preparation of the Native Soluble Collagen Solution
<ol>
	<li>Prepare or purchase 200 mg of acidified, soluble, type I collagen from rat tails at 1&#8211;3 mg/ml using standard isolation protocols, such as reported by Piez et al.15</li>
	<li>Scale the amount of starting material based on the desired end volume of modified collagen solutions. Approximately make 25&#8211;30 ml of methylated and 25&#8211;30 ml of succinylated collagen solutions, each at 3 mg/ml, from 200 mg of soluble native collagen.</li>
</ol>
2. Collagen Methylation
<ol>
	<li>Dilute 100 mg of the native, acidified (pH 2&#8211;3) collagen solution to a concentration of 0.5 mg/ml with ice cold sterile water and keep the solution on ice to prevent gelation.</li>
	<li>Adjust the pH of the collagen solution to 9&#8211;10 with a few drops of 1 N NaOH and stir at RT for 30 min. Observe the collagen precipitate, causing the solution to become cloudy.</li>
	<li>Spin down the precipitated collagen solution at 3,000 &#215; g for 25 min. A clear, gel-like precipitate should be visible in the bottom of the tubes. Aspirate and properly dispose of the supernatant fluid.</li>
	<li>Resuspend the precipitated collagen in 200 ml of methanol with 0.1 N HCl and allow the methylation reaction to occur with stirring at RT for 4 days. The collagen will not dissolve, but should break apart into very small visible pieces that will make the solution cloudy.</li>
	<li>After the methylation, centrifuge the solution at 3,000 &#215; g for 25 min to pellet the methylated collagen. Aspirate and dispose of the acidified methanol supernatant.</li>
	<li>Dissolve the methylated collagen in 25 ml of sterile PBS, giving a concentration of approximately 3 mg/ml, with repeated pipetting, and filter the solution through a 60 &#181;m cell strainer. Adjust the pH of the solution to 7.3&#8211;7.4 using 20 &#181;l increments of 1 N NaOH.</li>
	<li>Assess the concentration of the solution using a commercial rat collagen ELISA kit or hydroxyproline assay kit. Dilute the solution to 3 mg/ml with sterile PBS.</li>
	<li>Sterilize the methylated collagen solution by transferring it to a glass bottle with a screw cap, carefully layering 3 ml of chloroform at the bottom of the bottle, allow the bottle to set O/N&#160;at 4 &#176;C, and then aseptically remove and store the top layer, which is the methylated collagen.</li>
	<li>Store the solution at 4 &#176;C for use within 1 month.</li>
</ol>
3. Collagen Succinylation
<ol>
	<li>Dilute the other 100 mg of the native, acidified (pH 2&#8211;3) collagen solution to a concentration of 0.5 mg/ml with ice cold sterile water and keep the solution on ice to prevent gelation.</li>
	<li>Adjust the pH of the collagen solution to 9&#8211;10 with a few drops of 1 N NaOH and stir at RT for 30 min. The collagen should precipitate, causing the solution to become cloudy.</li>
	<li>Dissolve 40 mg of succinic anhydride (0.4 mg per mg of collagen) in 10 ml of acetone (1/20th the volume of the collagen solution). Slowly (in approximately 0.5 ml increments) add this mixture to the collagen solution with stirring while continuously monitoring the pH. Maintain the pH above 9.0 by adding 1 or 2 drops of 1 N NaOH as the pH approaches 9.0.</li>
	<li>Continue stirring at RT for 120 min after adding all of the succinic anhydride in acetone. Observe the solution to become clear as the succinylated collagen dissolves. Periodically check the pH to ensure it remains above 9.0.</li>
	<li>Adjust the pH of the solution to 4.0 with 20 &#181;l increments of 1 N HCl. Observe the solution cloudy again, as the succinylated collagen precipitates.</li>
	<li>After the succinylation, centrifuge the solution at 3,000 &#215; g for 25 min to pellet the succinylated collagen. Aspirate and dispose of the acidified supernatant with unreacted succinic anhydride.</li>
	<li>Dissolve the succinylated collagen in 25 ml of sterile PBS, giving a concentration of approximately 3 mg/ml, with repeated pipetting, and filter the solution through a 60 &#181;m cell strainer. Adjust the pH of the solution to 7.3&#8211;7.4 using 20 &#181;l increments of 1 N NaOH.</li>
	<li>Assess the concentration of the solution using a commercial collagen rat ELISA kit or hydroxyproline assay kit. Dilute the solution to 3 mg/ml with sterile PBS.</li>
	<li>Sterilize the succinylated collagen solution by transferring it to a glass bottle with a screw cap, carefully layering 3 ml of chloroform at the bottom of the bottle, allow the bottle to set O/N at 4 &#176;C, and then aseptically remove and store the top layer, which is the succinylated collagen.</li>
	<li>Store the solution at 4 &#176;C for use within 1 month.</li>
</ol>
4. Verification of the Collagen Modifications
<ol>
	<li>Prepare 1 ml samples from each of the native, methylated, and succinylated collagen solutions and dilute to a concentration of 0.1 mg/ml with sterile pure water for hydrogen ion titration. Further dilute the buffer at least 1,000-fold by dialysis against water through a 10 kDa cutoff membraneusing 3 solution changes for at least 4 hr&#160;each.</li>
	<li>Adjust the pH of the solutions to 7.3 with small amounts of NaOH and HCl. Using pH 7.3 as an arbitrary reference, perform hydrogen ion titration on each of the samples as described by Tanford16 to create titration curves for the native, methylated, and succinylated collagen solutions.</li>
	<li>Plot the change in pH per volume of acid added versus the number of bound H+ ions per molecule. The high pH &#8220;amino&#8221; range should show a shift in the succinylated collagen towards the neutral point (a loss of amine groups), and the low pH &#8220;carboxyl&#8221; range should show a leftward shift in the methylated collagen (a loss of carboxyl groups) and a rightward shift in the succinylated collagen (a gain in carboxyl groups), compared to the native collagen.</li>
	<li>Assess the efficacy of the succinylation reaction by determining the % of amino groups in the native collagen replaced by succinylation using the 2,4,6-trinitrobenzenesulfonic acid (TNBA) colorimetric method, following standard protocols17,18.</li>
</ol>
5. Fabrication of Microfluidic Devices and Cell Seeding
<ol>
	<li>Fabricate microfluidic devices using standard methods9. Use replica molding of PDMS from SU-8 masters on silicon defined using photolithography to create microfluidic cell culture chambers with 100 &#181;m tall, 0.4&#8211;1.5 mm wide, and 1&#8211;10 mm long channels for cell growth.</li>
	<li>Use a plasma cleaner to oxidize the surfaces of the device and a glass slide, and then press together to bond. After sterilizing the device by exposure to UV light for at least 30 min, fill the chamber with 50 &#181;g/ml fibronectin in sterile PBS and incubate at 37 &#176;C for 45 min.</li>
	<li>Seed the device with cells, such as primary rat or human hepatocytes, which require a collagen gel for stabilization of phenotype or differentiation state. We use 20 &#181;l of freshly isolated primary rat hepatocytes7,19 or commercially available cryopreserved primary human hepatocytes seeded at 14&#215;106 cells/ml per device.</li>
	<li>Allow the cells to attach for 4&#8211;6 hr, and then wash out unattached cells and replace the plating media with growth media. Incubate O/N to ensure full cell spreading and create a confluent monolayer of cells.</li>
</ol>
6. Layer-by-layer Collagen Deposition
<ol>
	<li>In a laminar flow tissue culture hood, prepare sufficient volumes of methylated and succinylated collagen solutions for 10 applications of each solution per device, as well as a few mls of media. Keep the solutions on ice. We use 20 &#181;l (10&#8211;15 times the total device volume) of collagen per layer per device.</li>
	<li>Beginning with the methylated (polycationic) solution, alternate flushing the devices with 20 &#181;l of methylated and then succinylated collagen solutions, waiting 1 min between each application. Flush the device a total of 10 times per solution, which should take approximately 20 min. Work quickly to minimize the amount of time the cells are without media.</li>
	<li>Observe collagen slowly accumulate at the inlet/outlet depending on its size. If the resistance to fluid flow increases, flush the device once or twice with media, and then continue layering.</li>
	<li>After applying all of the layers, rinse the device twice with fresh media and return to the incubator. Depending on the cell type, the extracellular matrix-induced morphological changes, such as enhanced polarization in hepatocytes, should be visible within a few hours.</li>
	<li>Prepare representative devices for transmission electron microscopy using standard methods9 to verify the presence of a collagen matrix assembly on top of the cultured cells.</li>
</ol>
7. Stabilization of Cell Phenotype and Function
<ol>
	<li>Image the cell morphology, viability, and polarity using standard methods for the cell type. For hepatocytes, collect images over 14 days using phase microscopy, LIVE/DEAD staining for viability, and CMFDA dye for apical polarization to demonstrate the effects of the collagen deposition.</li>
	<li>For cell morphology, flush the device through the inlet by pipette with 20 &#181;l of PBS to rinse, inject 20 &#181;l of PBS, and image the device using phase contrast microscopy.</li>
	<li>For LIVE/DEAD staining, flush the device through the inlet by pipette with 20 &#181;l of PBS to rinse, inject 20 &#181;l of LIVE/DEAD staining solution with DAPI prepared following the manufacturer&#39;s instructions, incubate for 30 min at 37 &#176;C, rinse the device again with 20 &#181;l of PBS, and image the device using fluorescence microscopy.</li>
	<li>For bile canaliculi staining, flush the device through the inlet by pipette with 20 &#181;l of PBS to rinse, inject 20 &#181;l of 2 &#181;M CMFDA staining solution with DAPI prepared following the manufacturer&#39;s instructions, incubate for 30 min at 37 &#176;C, rinse the device again with 20 &#181;l of PBS, and image the device using fluorescence microscopy.</li>
	<li>Collect the spent media and measure cellular metabolic products at appropriate time points over the culture duration to determine the cell function. For hepatocytes, measure the amount of albumin and urea in the spent media.</li>
	<li>Perform endpoint functional analyses on the cells themselves, such as assessing enzyme activity levels, antibody staining, or RNA analysis. For hepatocytes, induce and measure cytochrome P450 enzyme activities or phase II conjugation enzyme glutathione S-transferase.</li>
</ol>

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The authors have nothing to disclose.

Ultrathin pure collagen assemblies can be deposited on charged cells or material surfaces using layer-by-layer deposition of modified collagens. The results of this study demonstrate that methylation and succinylation of native collagen create polycationic and polyanionic collagen solutions (Figure 1) that can be used with the layer-by-layer technique to deposit ultrathin collagen matrix assemblies on cells (Figure 2) or other charged material surfaces. Such ultrathin matrix layers can s...

Although microfluidics allows for the exquisite control of the cellular microenvironment, culturing cells, especially primary cells, within microfluidic devices can be challenging. Many traditional cell culture techniques have been developed to sustain and stabilize cell function when cultured in tissue culture plates, but translating those techniques to microfluidic devices is often difficult. 
One such technique is the culture of cells on or sandwiched between collagen gels as a model of the physiological 3D cell environment.1 Type I collagen is one of the most frequently used proteins for biomaterials applications because of its ubiquity in extracellular matrix, natural abundance, robust cell attachment sites, and biocompatibility.2 Many cells benefit from 3D culture with collagen, including cancer cells3,45, microvascular endothelial cells6, and hepatocytes7, among others. While the use of collagen gels is easy in open formats, such as tissue culture plates, the limited channel dimensions and enclosed nature of microfluidic devices makes the use of liquids that gel impractical without blocking the entire channel.
To overcome this problem, we combined the layer-by-layer deposition technique8 with chemical modifications of native collagen solutions to create ultrathin collagen assemblies on top of cells cultured in microfluidic devices. These layers can stabilize cell morphology and function similar to collagen gels and can be deposited on cells in microfluidic devices without blocking the channels with polymerized matrix. The goal of this method is to modify native collagen to create polycationic and polyanionic collagen solutions and to stabilize cells in microfluidic culture by depositing thin collagen matrix assemblies onto the cells. This technique has been used to stabilize the morphology and function of primary hepatocytes in microfluidic devices.9
Although layer-by-layer deposition has previously been reported with natural and synthetic polyelectrolytes10 to cover hepatocytes in plate culture11,12 and as a seeding layer for hepatocytes in microfluidic devices13,14, this method describes the deposition of a pure collagen layer on top of hepatocytes, mimicking the 3D collagen culture techniques. In this protocol, we use hepatocytes as example cells that can be maintained using 3D collagen layers. The many other types of cells that benefit from 3D culture in collagen may similarly benefit from culture after layer-by-layer deposition of an ultrathin collagen matrix assembly.

<table><tbody><tr><td>collagen type I, rat tail</td><td>Life Technologies</td><td>A1048301</td><td>option for concentrated rat tail collagen</td></tr><tr><td>collagen type I, rat tail</td><td>Sigma-Aldrich</td><td>C3867-1VL</td><td>option for concentrated rat tail collagen</td></tr><tr><td>collagen type I, rat tail</td><td>EMD Millipore</td><td>08-115</td><td>option for concentrated rat tail collagen</td></tr><tr><td>collagen type I, rat tail</td><td>R%D Systems</td><td>3440-100-01</td><td>option for concentrated rat tail collagen</td></tr><tr><td>succinic anhydride</td><td>Sigma-Aldrich</td><td>239690-50G</td><td>succinylation reagent</td></tr><tr><td>anhydrous methanol</td><td>Sigma-Aldrich</td><td>322415-100ML</td><td>methylation reagent</td></tr><tr><td>sodium hydroxide</td><td>Sigma-Aldrich</td><td>S5881-500G</td><td>pH precipitation reagent</td></tr><tr><td>hydrochloric acid</td><td>Sigma-Aldrich</td><td>320331-500ML</td><td>pH precipitation reagent</td></tr><tr><td>rat collagen type I ELISA</td><td>Chondrex</td><td>6013</td><td>option for detecting collagen content</td></tr><tr><td>hydroxyproline assay kit</td><td>Sigma-Aldrich</td><td>MAK008-1KT</td><td>option for detecting collagen content</td></tr><tr><td>hydroxyproline assay kit</td><td>Quickzyme Biosciences</td><td>QZBtotcol1</td><td>option for detecting collagen content</td></tr></tbody></table>

layer-by-layer collagen deposition in microfluidic devices for microtissue stabilization

Although microfluidics provides exquisite control of the cellular microenvironment, culturing cells within microfluidic devices can be challenging. 3D culture of cells in collagen type I gels helps to stabilize cell morphology and function, which is necessary for creating microfluidic tissue models in microdevices. Translating traditional 3D culture techniques for tissue culture plates to microfluidic devices is often difficult because of the limited channel dimensions. In this method, we describe a technique for modifying native type I collagen to generate polycationic and polyanionic collagen solutions that can be used with layer-by-layer deposition to create ultrathin collagen assemblies on top of cells cultured in microfluidic devices. These thin collagen layers stabilize cell morphology and function, as shown using primary hepatocytes as an example cell, allowing for the long term culture of microtissues in microfluidic devices.

Native collagen can be modified using methylation and succinylation to create polycationic and polyanionic collagen solutions for use in layer-by-layer deposition. Succinylation modifies the &#949;-amino groups of native collagen with succinyl groups, and methylation modifies the carboxyl groups of native collagen with a methyl group (Figure 1A). These modifications to the collagen protein amino acid side chains alter the pH titration curves for the solutions. Succinylation reduces the number of amino gr...

Watch this Scientific Journal Video about Layer-by-layer Collagen Deposition in Microfluidic Devices for Microtissue Stabilization at JoVE.com

Layer-by-layer Collagen Deposition in Microfluidic Devices for Microtissue Stabilization

The creation of functional microtissues within microfluidic devices requires the stabilization of cell phenotypes by adapting traditional cell culture techniques to the limited spatial dimensions in microdevices. Modification of collagen allows the layer-by-layer deposition of ultrathin collagen assemblies that can stabilize primary cells, such as hepatocytes, as microfluidic tissue models.

Although microfluidics provides exquisite control of the cellular microenvironment, culturing cells within microfluidic devices can be challenging. 3D ...

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9.4K Views.  Massachusetts General Hospital, Harvard Medical School. The creation of functional microtissues within microfluidic devices requires the stabilization of cell phenotypes by adapting traditional cell culture techniques to the limited spatial dimensions in microdevices. Modification of collagen allows the layer-by-layer deposition of ultrathin collagen assemblies that can stabilize primary cells, such as hepatocytes, as microfluidic tissue models.

Video: Layer-by-layer Collagen Deposition in Microfluidic Devices for Microtissue Stabilization

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.

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 ...

Organotypic Collagen I Assay: A Malleable Platform to Assess Cell Behaviour in a 3-Dimensional Context

Cell migration is fundamental to many aspects of biology, including development, wound healing, the cellular responses of the immune system, and metastasis of tumor cells. Migration has been studied on glass coverslips in order to make cellular dynamics amenable to investigation by light microscopy. However, it has become clear that many aspects of cell migration depend on features of the local environment including its elasticity, protein composition, and pore size, which are not faithfully represented by rigid two dimensional substrates such as glass and plastic 1. Furthermore, interaction with other cell types, including stromal fibroblasts 2 and immune cells 3, has been shown to play a critical role in promoting the invasion of cancer cells. Investigation at the molecular level has increasingly shown that molecular dynamics, including response to drug treatment, of identical cells are significantly different when compared in vitro and in vivo 4. 
Ideally, it would be best to study cell migration in its naturally occurring context in living organisms, however this is not always possible. Intermediate tissue culture systems, such as cell derived matrix, matrigel, organotypic culture (described here) tissue explants, organoids, and xenografts, are therefore important experimental intermediates. These systems approximate certain aspects of an in vivo environment but are more amenable to experimental manipulation such as use of stably transfected cell lines, drug treatment regimes, long term and high-resolution imaging. Such intermediate systems are especially useful as proving grounds to validate probes and establish parameters required to image the dynamic response of cells and fluorescent reporters prior to undertaking imaging in vivo 5. As such, they can serve an important role in reducing the need for experiments on living animals.

A method is described for the preparation of a 3-dimensional matrix consisting of collagen type I and primary human fibroblasts. This organotypic gel serves as a useful substrate to assess invasive cell migration because it mimics basic features of tissue stroma and is amenable to many forms of microscopy.

Cell migration is fundamental to many aspects of biology, including development, wound healing, the cellular responses of the immune system, and ...

Engineering Fibrin-based Tissue Constructs from Myofibroblasts and Application of Constraints and Strain to Induce Cell and Collagen Reorganization

Collagen content and organization in developing collagenous tissues can be influenced by local tissue strains and tissue constraint. Tissue engineers aim to use these principles to create tissues with predefined collagen architectures. A full understanding of the exact underlying processes of collagen remodeling to control the final tissue architecture, however, is lacking. In particular, little is known about the (re)orientation of collagen fibers in response to changes in tissue mechanical loading conditions. We developed an in&#160;vitro model system, consisting of biaxially-constrained myofibroblast-seeded fibrin constructs, to further elucidate collagen (re)orientation in response to i) reverting biaxial to uniaxial static loading conditions and ii) cyclic uniaxial loading of the biaxially-constrained constructs before and after a change in loading direction, with use of the Flexcell FX4000T loading device. Time-lapse confocal imaging is used to visualize collagen (re)orientation in a nondestructive manner.
Cell and collagen organization in the constructs can be visualized in real-time, and an internal reference system allows us to relocate cells and collagen structures for time-lapse analysis. Various aspects of the model system can be adjusted, like cell source or use of healthy and diseased cells. Additives can be used to further elucidate mechanisms underlying collagen remodeling, by for example adding MMPs or blocking integrins. Shape and size of the construct can be easily adapted to specific needs, resulting in a highly tunable model system to study cell and collagen (re)organization.

This model system starts from a myofibroblast-populated fibrin gel that can be used to study endogenous collagen (re)organization real-time in a nondestructive manner. The model system is very tunable, as it can be used with different cell sources, medium additives, and can be adapted easily to specific needs.

Collagen content and organization in developing collagenous tissues can be influenced by local tissue strains and tissue constraint. Tissue engineers aim ...

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. ...

Microfluidic Genipin Deposition Technique for Extended Culture of Micropatterned Vascular Muscular Thin Films

The chronic nature of vascular disease progression requires the development of experimental techniques that simulate physiologic and pathologic vascular behaviors on disease-relevant time scales. Previously, microcontact printing has been used to fabricate two-dimensional functional arterial mimics through patterning of extracellular matrix protein as guidance cues for tissue organization. Vascular muscular thin films utilized these mimics to assess functional contractility. However, the microcontact printing fabrication technique used typically incorporates hydrophobic PDMS substrates. As the tissue turns over the underlying extracellular matrix, new proteins must undergo a conformational change or denaturing in order to expose hydrophobic amino acid residues to the hydrophobic PDMS surfaces for attachment, resulting in altered matrix protein bioactivity, delamination, and death of the tissues.
Here, we present a microfluidic deposition technique for patterning of the crosslinker compound genipin. Genipin serves as an intermediary between patterned tissues and PDMS substrates, allowing cells to deposit newly-synthesized extracellular matrix protein onto a more hydrophilic surface and remain attached to the PDMS substrates. We also show that extracellular matrix proteins can be patterned directly onto deposited genipin, allowing dictation of engineered tissue structure. Tissues fabricated with this technique show high fidelity in both structural alignment and contractile function of vascular smooth muscle tissue in a vascular muscular thin film model. This technique can be extended using other cell types and provides the framework for future study of chronic tissue- and organ-level functionality.

We present a method for microfluidic deposition of patterned genipin and fibronectin on PDMS substrates, allowing extended viability of vascular smooth muscle cell-dense tissues. This tissue fabrication method is combined with previous vascular muscular thin film technology to measure vascular contractility over disease-relevant time courses.

The chronic nature of vascular disease progression requires the development of experimental techniques that simulate physiologic and pathologic vascular ...

Preparation of 3D Collagen Gels and Microchannels for the Study of 3D Interactions In Vivo

Historically, most cellular processes have been studied in only 2 dimensions. While these studies have been informative about general cell signaling mechanisms, they neglect important cellular cues received from the structural and mechanical properties of the local microenvironment and extracellular matrix (ECM). To understand how cells interact within a physiological ECM, it is important to study them in the context of 3 dimensional assays. Cell migration, cell differentiation, and cell proliferation are only a few processes that have been shown to be impacted by local changes in the mechanical properties of a 3-dimensional ECM. Collagen I, a core fibrillar component of the ECM, is more than a simple structural element of a tissue. Under normal conditions, mechanical cues from the collagen network direct morphogenesis and maintain cellular structures. In diseased microenvironments, such as the tumor microenvironment, the collagen network is often dramatically remodeled, demonstrating altered composition, enhanced deposition and altered fiber organization. In breast cancer, the degree of fiber alignment is important, as an increase in aligned fibers perpendicular to the tumor boundary has been correlated to poorer patient prognosis1. Aligned collagen matrices result in increased dissemination of tumor cells via persistent migration2,3. The following is a simple protocol for embedding cells within a 3-dimensional, fibrillar collagen hydrogel. This protocol is readily adaptable to many platforms, and can reproducibly generate both aligned and random collagen matrices for investigation of cell migration, cell division, and other cellular processes in a tunable, 3-dimensional, physiological microenvironment.

Preparation of 3D Collagen Gels and Microchannels for the Study of 3D Interactions In Vivo

Collagen is a core component of the ECM, and provides essential cues for several cellular processes ranging from migration to differentiation and proliferation. Provided here is a protocol for embedding cells within 3D collagen hydrogels, and a more advanced technique for generating randomized or aligned collagen matrices using PDMS microchannels.

Historically, most cellular processes have been studied in only 2 dimensions.  While these studies have been informative about general cell signaling ...

Micropatterning and Assembly of 3D Microvessels

In vitro platforms to study endothelial cells and vascular biology are largely limited to 2D endothelial cell culture, flow chambers with polymer or glass based substrates, and hydrogel-based tube formation assays. These assays, while informative, do not recapitulate lumen geometry, proper extracellular matrix, and multi-cellular proximity, which play key roles in modulating vascular function. This manuscript describes an injection molding method to generate engineered vessels with diameters on the order of 100 &#181;m. Microvessels are fabricated by seeding endothelial cells in a microfluidic channel embedded within a native type I collagen hydrogel. By incorporating parenchymal cells within the collagen matrix prior to channel formation, specific tissue microenvironments can be modeled and studied. Additional modulations of hydrodynamic properties and media composition allow for control of complex vascular function within the desired microenvironment. This platform allows for the study of perivascular cell recruitment, blood-endothelium interactions, flow response, and tissue-microvascular interactions. Engineered microvessels offer the ability to isolate the influence from individual components of a vascular niche and precisely control its chemical, mechanical, and biological properties to study vascular biology in both health and disease.

This manuscript presents an injection molding method to engineer microvessels that recapitulate physiological properties of endothelium. The microfluidic-based process creates patent 3D vascular networks with tailorable conditions, such as flow, cellular composition, geometry, and biochemical gradients. The fabrication process and examples of potential applications are described.

In vitro platforms to study endothelial cells and vascular biology are largely limited to 2D endothelial cell culture, flow chambers with polymer or glass ...

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 ...

A Microfluidic Platform for Stimulating Chondrocytes with Dynamic Compression

Mechanical stimuli are known to modulate biological functions of cells and tissues. Recent studies have suggested that compressive stress alters growth plate cartilage architecture and results in growth modulation of long bones of children. To determine the role of compressive stress in bone growth, we created a microfluidic device actuated by pneumatic pressure, to dynamically (or statically) compress growth plate chondrocytes embedded in alginate hydrogel cylinders. In this article, we describe detailed methods for fabricating and characterizing this device. The advantages of our protocol are: 1) Five different magnitudes of compressive stress can be generated on five technical replicates in a single platform, 2) It is easy to visualize cell morphology via a conventional light microscope, 3) Cells can be rapidly isolated from the device after compression to facilitate downstream assays, and 4) The platform can be applied to study mechanobiology of any cell type that can grow in hydrogels.

This article provides detailed methods for fabricating and characterizing a pneumatically actuating microfluidic device for chondrocyte compression.

Mechanical stimuli are known to modulate biological functions of cells and tissues. Recent studies have suggested that compressive stress alters growth ...

Developing 3D Organized Human Cardiac Tissue within a Microfluidic Platform

The leading cause of death worldwide persists as cardiovascular disease (CVD). However, modeling the physiological and biological complexity of the heart muscle, the myocardium, is notoriously difficult to accomplish in vitro. Mainly, obstacles lie in the need for human cardiomyocytes (CMs) that are either adult or exhibit adult-like phenotypes and can successfully replicate the myocardium&#39;s cellular complexity and intricate 3D architecture. Unfortunately, due to ethical concerns and lack of available primary patient-derived human cardiac tissue, combined with the minimal proliferation of CMs, the sourcing of viable human CMs has been a limiting step for cardiac tissue engineering. To this end, most research has transitioned toward cardiac differentiation of human induced pluripotent stem cells (hiPSCs) as the primary source of human CMs, resulting in the wide incorporation of hiPSC-CMs within in vitro assays for cardiac tissue modeling.
Here in this work, we demonstrate a protocol for developing a 3D mature stem cell-derived human cardiac tissue within a microfluidic device. We specifically explain and visually demonstrate the production of a 3D in vitro anisotropic cardiac tissue-on-a-chip model from hiPSC-derived CMs. We primarily describe a purification protocol to select for CMs, the co-culture of cells with a defined ratio via mixing CMs with human CFs (hCFs), and suspension of this co-culture within the collagen-based hydrogel. We further demonstrate the injection of the cell-laden hydrogel within our well-defined microfluidic device, embedded with staggered elliptical microposts that serve as surface topography to induce a high degree of alignment of the surrounding cells and the hydrogel matrix, mimicking the architecture of the native myocardium. We envision that the proposed 3D anisotropic cardiac tissue-on-chip model is suitable for fundamental biology studies, disease modeling, and, through its use as a screening tool, pharmaceutical testing.

The goal of this protocol is to explain and demonstrate the development of a three-dimensional (3D) microfluidic model of highly aligned human cardiac tissue, composed of stem cell-derived cardiomyocytes co-cultured with cardiac fibroblasts (CFs) within a biomimetic, collagen-based hydrogel, for applications in cardiac tissue engineering, drug screening, and disease modeling.

The leading cause of death worldwide persists as cardiovascular disease (CVD). However, modeling the physiological and biological complexity of the heart ...

Layer-by-layer Collagen Deposition in Microfluidic Devices for Microtissue Stabilization

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Chapters in this video

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

Organotypic Collagen I Assay: A Malleable Platform to Assess Cell Behaviour in a 3-Dimensional Context

Engineering Fibrin-based Tissue Constructs from Myofibroblasts and Application of Constraints and Strain to Induce Cell and Collagen Reorganization

Engineering 3D Cellularized Collagen Gels for Vascular Tissue Regeneration

Microfluidic Genipin Deposition Technique for Extended Culture of Micropatterned Vascular Muscular Thin Films

Preparation of 3D Collagen Gels and Microchannels for the Study of 3D Interactions In Vivo

Micropatterning and Assembly of 3D Microvessels

Microfluidic Bioprinting for Engineering Vascularized Tissues and Organoids

A Microfluidic Platform for Stimulating Chondrocytes with Dynamic Compression

Developing 3D Organized Human Cardiac Tissue within a Microfluidic Platform