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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S., 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=Multilineage+differentiation+of+rhesus+monkey+embryonic+stem+cells+in+three&#8208;dimensional+culture+systems.">Multilineage differentiation of rhesus monkey embryonic stem cells in three&#8208;dimensional culture systems.</a> Stem Cells. 21 (3), 281-295 (2003).</li><li>Whelan, M. C., Senger, D. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Collagen+I+initiates+endothelial+cell+morphogenesis+by+inducing+actin+polymerization+through+suppression+of+cyclic+AMP+and+protein+kinase+A.">Collagen I initiates endothelial cell morphogenesis by inducing actin polymerization through suppression of cyclic AMP and protein kinase A.</a> J Biol Chem. 278 (1), 327-334 (2003).</li><li>Dunn, J. C., Tompkins, R. G., Yarmush, M. 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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 border="0" cellpadding="0" cellspacing="0">
	<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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9.3K 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

Separating Beads and Cells in Multi-channel Microfluidic Devices Using Dielectrophoresis and Laminar Flow

Microfluidic devices have advanced cell studies by providing a dynamic fluidic environment on the scale of the cell for studying, manipulating, sorting and counting cells. However, manipulating the cell within the fluidic domain remains a challenge and requires complicated fabrication protocols for forming valves and electrodes, or demands specialty equipment like optical tweezers. Here, we demonstrate that conventional printed circuit boards (PCB) can be used for the non-contact manipulation of cells by employing dielectrophoresis (DEP) for bead and cell manipulation in laminar flow fields for bioactuation, and for cell and bead separation in multichannel microfluidic devices. First, we present the protocol for assembling the DEP electrodes and microfluidic devices, and preparing the cells for DEP. Then, we characterize the DEP operation with polystyrene beads. Lastly, we show representative results of bead and cell separation in a multichannel microfluidic device. In summary, DEP is an effective method for manipulating particles (beads or cells) within microfluidic devices.

Dielectrophoresis (DEP) is an effective method to manipulate cells. Printed circuit boards (PCB) can provide inexpensive, reusable and effective electrodes for contact-free cell manipulation within microfluidic devices. By combining PDMS-based microfluidic channels with coverslips on PCBs, we demonstrate bead and cell manipulation and separation within multichannel microfluidic devices.

Microfluidic devices have advanced cell studies by providing a dynamic fluidic environment on the scale of the cell for studying, manipulating, sorting ...

Simple Microfluidic Devices for in vivo Imaging of C. elegans, Drosophila and Zebrafish

Micro fabricated fluidic devices provide an accessible micro-environment for in vivo studies on small organisms. Simple fabrication processes are available for microfluidic devices using soft lithography techniques 1-3. Microfluidic devices have been used for sub-cellular imaging 4,5, in vivo laser microsurgery 2,6 and cellular imaging 4,7. In vivo imaging requires immobilization of organisms. This has been achieved using suction 5,8, tapered channels 6,7,9, deformable membranes 2-4,10, suction with additional cooling 5, anesthetic gas 11, temperature sensitive gels 12, cyanoacrylate glue 13 and anesthetics such as levamisole 14,15. Commonly used anesthetics influence synaptic transmission 16,17 and are known to have detrimental effects on sub-cellular neuronal transport 4. In this study we demonstrate a membrane based poly-dimethyl-siloxane (PDMS) device that allows anesthetic free immobilization of intact genetic model organisms such as Caenorhabditis elegans (C. elegans), Drosophila larvae and zebrafish larvae. These model organisms are suitable for in vivo studies in microfluidic devices because of their small diameters and optically transparent or translucent bodies. Body diameters range from ~10 &mu;m to ~800 &mu;m for early larval stages of C. elegans and zebrafish larvae and require microfluidic devices of different sizes to achieve complete immobilization for high resolution time-lapse imaging. These organisms are immobilized using pressure applied by compressed nitrogen gas through a liquid column and imaged using an inverted microscope. Animals released from the trap return to normal locomotion within 10 min.
We demonstrate four applications of time-lapse imaging in C. elegans namely, imaging mitochondrial transport in neurons, pre-synaptic vesicle transport in a transport-defective mutant, glutamate receptor transport and Q neuroblast cell division. Data obtained from such movies show that microfluidic immobilization is a useful and accurate means of acquiring in vivo data of cellular and sub-cellular events when compared to anesthetized animals (Figure 1J and 3C-F 4).
Device dimensions were altered to allow time-lapse imaging of different stages of C. elegans, first instar Drosophila larvae and zebrafish larvae. Transport of vesicles marked with synaptotagmin tagged with GFP (syt.eGFP) in sensory neurons shows directed motion of synaptic vesicle markers expressed in cholinergic sensory neurons in intact first instar Drosophila larvae. A similar device has been used to carry out time-lapse imaging of heartbeat in ~30 hr post fertilization (hpf) zebrafish larvae. These data show that the simple devices we have developed can be applied to a variety of model systems to study several cell biological and developmental phenomena in vivo.

Simple Microfluidic Devices for in vivo Imaging of C. elegans, Drosophila and Zebrafish

A simple microfluidic device has been developed to perform anesthetic free in vivo imaging of C. elegans, intact Drosophila larvae and zebrafish larvae. The device utilizes a deformable PDMS membrane to immobilize these model organisms in order to perform time lapse imaging of numerous processes such as heart beat, cell division and sub-cellular neuronal transport. We demonstrate the use of this device and show examples of different types of data collected from different model systems.

Micro fabricated fluidic devices provide an accessible micro-environment for in vivo studies on small organisms. Simple fabrication processes are ...

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 ...

Using Adhesive Patterning to Construct 3D Paper Microfluidic Devices

We demonstrate the use of patterned aerosol adhesives to construct both planar and nonplanar 3D paper microfluidic devices. By spraying an aerosol adhesive through a metal stencil, the overall amount of adhesive used in assembling paper microfluidic devices can be significantly reduced. We show on a simple 4-layer planar paper microfluidic device that the optimal adhesive application technique and device construction style depends heavily on desired performance characteristics. By moderately increasing the overall area of a device, it is possible to dramatically decrease the wicking time and increase device success rates while also reducing the amount of adhesive required to keep the device together. Such adhesive application also causes the adhesive to form semi-permanent bonds instead of permanent bonds between paper layers, enabling single-use devices to be non-destructively disassembled after use. Nonplanar 3D origami devices also benefit from the semi-permanent bonds during folding, as it reduces the likelihood that unrelated faces may accidently stick together. Like planar devices, nonplanar structures see reduced wicking times with patterned adhesive application vs uniformly applied adhesive.

We demonstrate the use of patterned aerosol adhesives to construct 3D paper microfluidic devices. This method of adhesive application forms semi-permanent bonds between layers, enabling single-use devices to be non-destructively disassembled after use and to ease folding complex nonplanar structures.

We demonstrate the use of patterned aerosol adhesives to construct both planar and nonplanar 3D paper microfluidic devices. By spraying an aerosol ...

Designing Microfluidic Devices for Studying Cellular Responses Under Single or Coexisting Chemical/Electrical/Shear Stress Stimuli

Microfluidic devices are capable of creating a precise and controllable cellular micro-environment of pH, temperature, salt concentration, and other physical or chemical stimuli. They have been commonly used for in vitro cell studies by providing in vivo like surroundings. Especially, how cells response to chemical gradients, electrical fields, and shear stresses has drawn many interests since these phenomena are important in understanding cellular properties and functions. These microfluidic chips can be made of glass substrates, silicon wafers, polydimethylsiloxane (PDMS) polymers, polymethylmethacrylate (PMMA) substrates, or polyethyleneterephthalate (PET) substrates. Out of these materials, PMMA substrates are cheap and can be easily processed using laser ablation and writing. Although a few microfluidic devices have been designed and fabricated for generating multiple, coexisting chemical and electrical stimuli, none of them was considered efficient enough in reducing experimental repeats, particular for screening purposes. In this report, we describe our design and fabrication of two PMMA-based microfluidic chips for investigating cellular responses, in the production of reactive oxygen species and the migration, under single or coexisting chemical/electrical/shear stress stimuli. The first chip generates five relative concentrations of 0, 1/8, 1/2, 7/8, and 1 in the culture regions, together with a shear stress gradient produced inside each of these areas. The second chip generates the same relative concentrations, but with five different electric field strengths created within each culture area. These devices not only provide cells with a precise, controllable micro-environment but also greatly increase the experimental throughput.

Micro-fabricated devices integrated with fluidic components provide an in vitro platform for cell studies mimicking the in vivo micro-environment. We developed polymethylmethacrylate-based microfluidic chips for studying cellular responses under single or coexisting chemical/electrical/shear stress stimuli.

Microfluidic devices are capable of creating a precise and controllable cellular micro-environment of pH, temperature, salt concentration, and other ...

Multi-step Variable Height Photolithography for Valved Multilayer Microfluidic Devices

Microfluidic systems have enabled powerful new approaches to high-throughput biochemical and biological analysis. However, there remains a barrier to entry for non-specialists who would benefit greatly from the ability to develop their own microfluidic devices to address research questions. Particularly lacking has been the open dissemination of protocols related to photolithography, a key step in the development of a replica mold for the manufacture of polydimethylsiloxane (PDMS) devices. While the fabrication of single height silicon masters has been explored extensively in literature, fabrication steps for more complicated photolithography features necessary for many interesting device functionalities (such as feature rounding to make valve structures, multi-height single-mold patterning, or high aspect ratio definition) are often not explicitly outlined. 
Here, we provide a complete protocol for making multilayer microfluidic devices with valves and complex multi-height geometries, tunable for any application. These fabrication procedures are presented in the context of a microfluidic hydrogel bead synthesizer and demonstrate the production of droplets containing polyethylene glycol (PEG diacrylate) and a photoinitiator that can be polymerized into solid beads. This protocol and accompanying discussion provide a foundation of design principles and fabrication methods that enables development of a wide variety of microfluidic devices. The details included here should allow non-specialists to design and fabricate novel devices, thereby bringing a host of recently developed technologies to their most exciting applications in biological laboratories.

Multilayer microfluidic devices often involve the fabrication of master molds with complex geometries for functionality. This article presents a complete protocol for multi-step photolithography with valves and variable height features tunable to any application. As a demonstration, we fabricate a microfluidic droplet generator capable of producing hydrogel beads.

Microfluidic systems have enabled powerful new approaches to high-throughput biochemical and biological analysis. However, there remains a barrier to ...

Fabrication of Three-dimensional Paper-based Microfluidic Devices for Immunoassays

Paper wicks fluids autonomously due to capillary action. By patterning paper with hydrophobic barriers, the transport of fluids can be controlled and directed within a layer of paper. Moreover, stacking multiple layers of patterned paper creates sophisticated three-dimensional microfluidic networks that can support the development of analytical and bioanalytical assays. Paper-based microfluidic devices are inexpensive, portable, easy to use, and require no external equipment to operate. As a result, they hold great promise as a platform for point-of-care diagnostics. In order to properly evaluate the utility and analytical performance of paper-based devices, suitable methods must be developed to ensure their manufacture is reproducible and at a scale that is appropriate for laboratory settings. In this manuscript, a method to fabricate a general device architecture that can be used for paper-based immunoassays is described. We use a form of additive manufacturing (multi-layer lamination) to prepare devices that comprise multiple layers of patterned paper and patterned adhesive. In addition to demonstrating the proper use of these three-dimensional paper-based microfluidic devices with an immunoassay for human chorionic gonadotropin (hCG), errors in the manufacturing process that may result in device failures are discussed. We expect this approach to manufacturing paper-based devices will find broad utility in the development of analytical applications designed specifically for limited-resource settings.

We detail a method to fabricate three-dimensional paper-based microfluidic devices for use in the development of immunoassays. Our approach to device assembly is a type of multilayer, additive manufacturing. We demonstrate a sandwich immunoassay to provide representative results for these types of paper-based devices.

Paper wicks fluids autonomously due to capillary action. By patterning paper with hydrophobic barriers, the transport of fluids can be controlled and ...

Polydimethylsiloxane-polycarbonate Microfluidic Devices for Cell Migration Studies Under Perpendicular Chemical and Oxygen Gradients

This paper reports a microfluidic device made of polydimethylsiloxane (PDMS) with an embedded polycarbonate (PC) thin film to study cell migration under combinations of chemical and oxygen gradients. Both chemical and oxygen gradients can greatly affect cell migration in vivo; however, due to technical limitations, very little research has been performed to investigate their effects in vitro. The device developed in this research takes advantage of a series of serpentine-shaped channels to generate the desired chemical gradients and exploits a spatially confined chemical reaction method for oxygen gradient generation. The directions of the chemical and oxygen gradients are perpendicular to each other to enable straightforward migration result interpretation. In order to efficiently generate the oxygen gradients with minimal chemical consumption, the embedded PC thin film is utilized as a gas diffusion barrier. The developed microfluidic device can be actuated by syringe pumps and placed into a conventional cell incubator during cell migration experiments to allow for setup simplification and optimized cell culture conditions. In cell experiments, we used the device to study migrations of adenocarcinomic human alveolar basal epithelial cells, A549, under combinations of chemokine (stromal cell-derived factor, SDF-1&#945;) and oxygen gradients. The experimental results show that the device can stably generate perpendicular chemokine and oxygen gradients and is compatible with cells. The migration study results indicate that oxygen gradients may play an essential role in guiding cell migration, and cellular behavior under combinations of gradients cannot be predicted from those under single gradients. The device provides a powerful and practical tool for researchers to study interactions between chemical and oxygen gradients in cell culture, which can promote better cell migration studies in more in vivo-like microenvironments.

The control of chemical and oxygen gradients is essential for cell cultures. This paper reports a polydimethylsiloxane-polycarbonate (PDMS-PC) microfluidic device capable of reliably generating combinations of chemical and oxygen gradients for cell migration studies, which can be practically utilized in biological labs without sophisticated instrumentation.

This paper reports a microfluidic device made of polydimethylsiloxane (PDMS) with an embedded polycarbonate (PC) thin film to study cell migration under ...

A Microfluidic Flow Chamber Model for Platelet Transfusion and Hemostasis Measures Platelet Deposition and Fibrin Formation in Real-time

Microfluidic models of hemostasis assess platelet function under conditions of hydrodynamic shear, but in the presence of anticoagulants, this analysis is restricted to platelet deposition only. The intricate relationship between Ca2+-dependent coagulation and platelet function requires careful and controlled recalcification of blood prior to analysis. Our setup uses a Y-shaped mixing channel, which supplies concentrated Ca2+/Mg2+ buffer to flowing blood just prior to perfusion, enabling rapid recalcification without sample stasis. A ten-fold difference in flow velocity between both reservoirs minimizes dilution. The recalcified blood is then perfused in a collagen-coated analysis chamber, and differential labeling permits real-time imaging of both platelet and fibrin deposition using fluorescence video microscopy. The system uses only commercially available tools, increasing the chances of standardization. Reconstitution of thrombocytopenic blood with platelets from banked concentrates furthermore models platelet transfusion, proving its use in this research domain. Exemplary data demonstrated that coagulation onset and fibrin deposition were linearly dependent on the platelet concentration, confirming the relationship between primary and secondary hemostasis in our model. In a timeframe of 16 perfusion min, contact activation did not take place, despite recalcification to normal Ca2+ and Mg2+ levels. When coagulation factor XIIa was inhibited by corn trypsin inhibitor, this time frame was even longer, indicating a considerable dynamic range in which the changes in the procoagulant nature of the platelets can be assessed. Co-immobilization of tissue factor with collagen significantly reduced the time to onset of coagulation, but not its rate. The option to study the tissue factor and/or the contact pathway increases the versatility and utility of the assay.

This paper describes an experimental model of hemostasis that simultaneously measures platelet function and coagulation. Platelet and fibrin fluorescence is measured in real-time, and platelet adhesion rate, coagulation rate, and onset of coagulation are determined. The model is used to determine platelet procoagulant properties under flow in concentrates for transfusion.

Microfluidic models of hemostasis assess platelet function under conditions of hydrodynamic shear, but in the presence of anticoagulants, this analysis is ...