We thank M. Rousseau and S. Kroll for help with photography and videography and M. Ingram, J. Nguyen, D. Shea, and N. Wen for contributions to initial fabrication protocol development. This research was sponsored by the Wyss Institute for Biologically Inspired Engineering at Harvard University and the Defense Advanced Research Projects Agency under Cooperative Agreements #W911NF-12-2-0036 and #W911NF-16-C-0050, and FDA grant #HHSF223201310079C, NIH grants #R01-EB020004 and #UG3-HL141797-01, and Bill and Melinda Gates Foundation grants #OPP1163237 and #OPP1173198 to DEI. The views and conclusions contained in this document are those of the authors and should not be interpreted as representing the official policies, either expressed or implied, of the Defense Advanced Research Projects Agency, Food and Drug Administration, the National Institutes of Health, or the U.S. Government.

1. General Preparation
<ol>
	<li>To avoid debris, clean work area using packing tape and wipe down area with a cleanroom wipe and isopropyl alcohol.</li>
	<li>For all steps requiring PDMS, mix PDMS at a 10:1 ratio (10 g of cross linking agent, 100 g of elastomer base). Mix by hand or with a commercially available mixer. Use a planetary centrifugal mixer here: mixing for 2 minutes at 2000 rpm, then degassing the PDMS for 2 minutes at 2200 rpm.</li>
	<li>Clean all molds with air gun to blow out debris prior to use. 
	CAUTION: Do not use metal forceps to remove debris as it will damage the surface of the molds.</li>
</ol>
2. Top Channel Preparation
<ol>
	<li>Wipe down the glossy side of each polyurethane piece with ethanol and cleanroom wipes. Make sure all residual ethanol is dried from the polyurethane surface.</li>
	<li>Place the glossy side of the polyurethane over the open side of the MiP mold to create a seal on the open side of the mold, leaving only a well-like opening at the top of the mold for pouring PDMS. 
	NOTE: Check that every mold is covered securely by polyurethane piece or the PDMS will leak from molds during pouring.</li>
	<li>Place the mold and polyurethane assemblies into a MiP jig, with the textured side against the end of the MiP jig. Continue to do this until all molds have been placed in the jig.</li>
	<li>Tighten the MiP jig by turning the handle using a wrench until the jig spacing is 25 mm in width.</li>
	<li>Make a boat of aluminum foil surrounding the MiP jig to prevent excess PDMS from leaking onto surfaces.</li>
	<li>Pour PDMS into each of the molds well until full. 
	NOTE: Each chip top component requires about 3 mL of PDMS.</li>
	<li>Once the entire jig is filled, place the jig into the vacuum desiccator. Pull vacuum at -80 kPa for 1 h to degas PDMS.</li>
	<li>After 1 h, remove the MiP jig from the desiccator and place in 60 &#176;C oven for at least 4 h to cure PDMS.</li>
	<li>Disassemble the MiP jig using a wrench, loosen the jig by turning the handle counter clockwise. Once molds are free from compression, remove molds from jig.</li>
	<li>Remove the polyurethane strips from each mold and discard.</li>
	<li>Carefully de-mold PDMS parts from their molds and lay them feature-side up.</li>
	<li>Line up the blade of tile scraper at end tab notch and cut away each end to singulate the top components.</li>
	<li>Check parts for any of the following failure modes and discard any unsatisfactory parts: scratches in the main channel, large debris above the channel area, large bubbles, deformed vacuum channels.</li>
	<li>Store finished parts in square Petri dishes within pressure positive cabinets at room temperature.</li>
</ol>
3. Bottom Channel Preparation
<ol>
	<li>Pour approximately 10.5 g of PDMS into molds until the PDMS reaches the top of the cavity.
	<ol>
		<li>Inspect the bottom channel mold for PDMS cured to the bottom of mold.</li>
		<li>If dirty, scrape old PDMS from the bottom of the mold since an uneven surface on the bottom of the mold can cause uneven thickness of the final parts. 
		NOTE: For small &#60;2 cm2 areas that are uncovered, the air gun can be used very gently to move PDMS over the space.</li>
	</ol>
	</li>
	<li>Place molds into vacuum desiccator for 1 h.</li>
	<li>After 1 h, move the molds into a level 60 &#176;C oven for &#62;4 h.</li>
	<li>Place mold on table in laminar flow hood. Loosen PDMS from one edge of the mold.</li>
	<li>Grip one corner and gently peel back the PDMS from the mold surface.</li>
	<li>When fully removed, lay on work surface, so that channel features are face up.</li>
	<li>Cut parts along outside edges with tile cutters, placing tile cutter blade in notched PDMS as in step 2.12.</li>
	<li>Lay parts feature side up on tape to remove any debris.</li>
	<li>Remove part from packing tape. Drag the loose end of the part across the slide. The loose end will laminate with the glass. 
	NOTE: It is critical to avoid stretching the part while laying it down. If bubble is trapped between part and glass, gently lift the part with the forceps and re-lay.</li>
	<li>Perform quality control inspection of parts. Check parts for any failure modes and discard any unsatisfactory parts, including ones that contain scratches in the main channel, large debris, large bubbles, or deformed vacuum channels.</li>
	<li>Cover features with tape.</li>
	<li>Store parts in a positive pressure cabinet at room temperature.</li>
</ol>
4. PDMS Membrane Preparation
<ol>
	<li>Check that the wafers are free of PDMS on the back.</li>
	<li>Place each membrane wafer in the designated slots in the AMF trays.</li>
	<li>Use the 1 mL syringe to place 0.09 mL of PDMS onto the center of each membrane wafer post array. Let PDMS sit for a minimum of 5 min to allow PDMS to spread throughout the posts of the membrane wafer. 
	NOTE: Do not proceed to next step until at least 75% of the post array is covered in PDMS. The quality of membranes improves the longer the PDMS is allowed to wick into post region so longer wait times in this step are preferred.</li>
	<li>Plasma treat the polycarbonate strip at 20 W for 45 s, O2 gas at 0.80 mbar in a plasma machine.</li>
	<li>Remove the polycarbonate sheet from the plasma machine and use scissors to cut the polycarbonate sheets into 45 mm x 45 mm squares. 
	NOTE: Minimize contact with the plasma treated surface to prevent dust from sticking to the polycarbonate.</li>
	<li>Gently lay the plasma treated side of the polycarbonate squares onto the liquid PDMS centered on the membrane wafer. Ensure that the polycarbonate and the PDMS are in contact. 
	NOTE: Avoid air pockets between the polycarbonate and the PDMS.</li>
	<li>Place the pre-cut PDMS spacer on the center of the polycarbonate square.</li>
	<li>Place the pre-cut textured polycarbonate sheet on the PDMS block to keep the assembly from bonding to the compression plate.</li>
	<li>Insert tray so that Tray 3 is in the back, Tray 2 is in the middle, and Tray 1 is in the front. Tray 1 has a notch for alignment.</li>
	<li>Open the output pressure valve and very slowly open the input pressure valve. Only then close the output pressure valve. 
	NOTE: This is so that the output 4 kg of force is gradually applied to each membrane wafer as opposed to instantly, which may break the wafers.</li>
	<li>Flip the AMF switch to ON to begin the curing cycle. Cure wafer under 4 kg (16 kPa) of compression and a ramping temperature cycle listed in Table 1.</li>
</ol>
<table border="0" cellpadding="0" cellspacing="0" style="width:323px;" width="322">
	<colgroup>
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="39">
			<td height="39" style="height:39px;width:91px;">Step</td>
			<td style="width:131px;">Temperature (&#176;C)</td>
			<td style="width:101px;">Duration (min)</td>
		</tr>
		<tr height="20">
			<td align="right" height="20" style="height:20px;">1</td>
			<td align="right">20</td>
			<td align="right">20</td>
		</tr>
		<tr height="20">
			<td align="right" height="20" style="height:20px;">2</td>
			<td align="right">35</td>
			<td align="right">10</td>
		</tr>
		<tr height="20">
			<td align="right" height="20" style="height:20px;">3</td>
			<td align="right">45</td>
			<td align="right">10</td>
		</tr>
		<tr height="20">
			<td align="right" height="20" style="height:20px;">4</td>
			<td align="right">50</td>
			<td align="right">60</td>
		</tr>
		<tr height="20">
			<td align="right" height="20" style="height:20px;">5</td>
			<td align="right">60</td>
			<td align="right">120</td>
		</tr>
		<tr height="18">
			<td align="right" height="18" style="height:19px;">6</td>
			<td align="right">20</td>
			<td>Hold</td>
		</tr>
	</tbody>
</table>
Table 1 - Membrane curing conditions
<ol start="12">
	<li>Close the input pressure valve and open the output pressure valve to release pressure from the air cylinders.</li>
	<li>Remove the trays and bring them to the laminar flow hood.</li>
	<li>Carefully peel off the textured polycarbonate and carefully remove the PDMS spacer. 
	NOTE: Watch the PDMS spacer to ensure it does not also peel off the polycarbonate carrier, if this occurs start peeling from a different corner.
	<ol>
		<li>Inspect the PDMS membrane through the polycarbonate carrier for areas with through-holes and use a marker to trace the outline of the through-hole area and mark any holes or defects in the membranes.</li>
		<li>Using wafer handling forceps, loosen wafers from the tray.</li>
		<li>Remove each membrane from the wafer and place in Petri dish. 
		NOTE: The PDMS membrane will de-mold from the membrane wafer and will be adhered to the polycarbonate backing. If PDMS starts detaching from the polycarbonate carrier, peel from a different region.</li>
		<li>Store membranes and wafers in Petri dishes in a positive pressure cabinet at room temperature.</li>
	</ol>
	</li>
</ol>
5. Top Assembly and Preparation
<ol>
	<li>Using matte tape, clean the PDMS membranes as well as the insides of the Petri dish to remove debris.</li>
	<li>Thoroughly tape the feature side of each top component to remove debris.</li>
	<li>Place top channel part (&#34;top&#34;) feature side up in Petri dish with PDMS membrane. 
	NOTE: Be aware that some membranes may be used for one or two top parts depending on size of the usable area. The main channels of each top part should fit within the marked area of the membrane.</li>
	<li>Load the Petri dishes into the plasma machine.</li>
	<li>Plasma treat membrane and top at 20 W for 45 s, O2 gas at 0.80 mbar.</li>
	<li>Once the bonding cycles has finished, remove the dishes and lay the activated parts feature side down on top of the membrane and ensure part is fully laminated with membrane with no bubbles.</li>
	<li>Place parts into 60 &#176;C oven for at least 2 h to anneal.</li>
	<li>Using a scalpel, trace around the perimeter of the bonded top to separate top-membrane assembly from the polycarbonate carrier. 
	NOTE: Do not cut the carrier.</li>
	<li>Once the part is traced, peel the assembly from the polycarbonate. The PDMS membrane that is bonded to the top should peel from the carrier.</li>
	<li>Using sharp tipped forceps, remove the membrane from the ports that access the bottom channel, and remove any debris or dust with forceps under a stereoscope. 
	NOTE: Do not leave any part of the membrane covering the access port.</li>
</ol>
6. Chip Assembly
<ol>
	<li>Feature side up, plasma treat assemblies with bottom components using the conditions in step 5.5.</li>
	<li>Under an inverted microscope, align the top assembly with microscope slide to the bottom half.</li>
	<li>Place in 60 &#176;C oven for at least 2 h.</li>
	<li>Chip Quality Control Inspection 
	NOTE: Pay close attention to the main ports and channel of chip. Check for failure modes by eye and also under the microscope.
	<ol>
		<li>To check that the chip is bonded fully, tug lightly on each corner of the chip to check for delaminating parts.</li>
		<li>Look at the channel of the chip to check for a wrinkled or sagging membrane, which will appear as a wavy pattern or a light deflection in the channel.</li>
		<li>Perform a microscope inspection to inspect for debris in the main channel. 
		NOTE: Debris in non-critical areas, such as the vacuum channels is acceptable.</li>
		<li>With the chip still on the inverted microscope, inspect main channel and vacuum channels for delamination. 
		NOTE: Delamination in non-critical areas (e.g., the edge of the chip) is acceptable.</li>
		<li>Check that the main channels are aligned to within 50-60 &#181;m (1-2 membrane pores). 
		NOTE: It is crucial that the channels are not overlapping with vacuum channels.</li>
		<li>Check that the membrane between the main channels and the inlet and outlet channels is intact without any apparent holes. 
		NOTE: Any hole in the membrane can lead to a leaky chip or cell growth outside the channels.</li>
	</ol>
	</li>
	<li>Store chips in Petri dishes in a positive pressure cabinet at room temperature.</li>
</ol>

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D.E.I. is a founder and holds equity in Emulate, Inc., and chairs its scientific advisory board. J.P. is presently an employee of Emulate, Inc. R.N., Y.C., J.P., and D.E.I. are inventors on intellectual property that has been licensed to Emulate, Inc.

The fabrication process relies on high resolution 3D printed molds to pattern the PDMS top and bottom Organ Chip body components coupled with micromolded porous PDMS membranes. This critical approach was selected due to ease of prototyping combined with rapid transition into scaled up fabrication and replacement of tooling. The top component molds are designed to pattern ports in precise locations with defined vertical profiles during the casting step. This not only avoids the labor involved in manually punching access ports but also reduces debris in the workplace, enables reproducible port alignment to interface manifolds or instrumentation, and produces parts with control over the fit and sealing of inserted tubing or pins for fluidic and pneumatic connections. The molds are stacked on top of each other in a compression jig, separated by compliant polyurethane sheets to facilitate through-hole casting of ports. By stacking multiple parts in a single jig, a single user can cast large quantities of components complete with ports in a single step. Material selection and manufacturing method for the molds are critical to provide the necessary feature resolution, low surface roughness, and high degree of flatness for device assembly and subsequent imaging applications. Stereolithography can meet these requirements, although materials with high deflection temperatures (&#62; 80 &#176;C) and compatibility with PDMS curing reduce the available polymer range. Various commercially available resins, including glass-filled resins, meet these criteria.
The elastic porous PDMS membrane is arguably the most unique and critical component of an Organ Chip while being the most complex to fabricate. A deep reactive ion etches (DRIE) process outsourced to a vendor is used to microfabricate 50 x 50 mm hexagonal arrays of pillars (7 &#181;m diameter, 40 &#181;m apart, 50 &#181;m tall, C4F8 coated) that are used to pattern pores in the PDMS membrane. The quality of the pillar arrays is critical to achieving robust membrane casting. In particular, pillars must be etched to tight tolerances with smooth vertical profiles to avoid undercuts or excessive sidewall roughness that can lead to mold failure. Care should be taken to avoid &#34;grassing&#34; at the bottom of the etched region, which can affect membrane demolding and cell attachment. Membrane fabrication with successful through-hole patterning and device integration is the single most complex section of the protocol. Critically, placing 0.09 mL of PDMS on each wafer and allowing adequate time for it to spread is essential to avoid incomplete through-hole molding. Properly plasma treating the polycarbonate backing is required to achieving robust backing of the membrane for demolding and bonding steps without wrinkling or stretching. The backing provides a robust means of demolding the cast membrane from the fragile silicon wafer.
The compressive load applied to each wafer is also essential for uniform through-hole fabrication. Earlier efforts using weights hindered membrane production and resulted in poor yields due to non-uniform force distribution. To overcome the production bottleneck, we optimized the previously published membrane fabrication protocol18 and built an Automated Membrane Fabricator (AMF) to parallelize the process. The AMF consists of 24 pneumatic pistons supported over a programmable hot plate to provide controlled compressive force throughout a programmed PDMS curing process. A polycarbonate backing film is placed on the uncured PDMS and then uniformly compressed using pneumatic pistons of the AMF while being heated to polymerize the PDMS. Critically, the gradual curing process described in the protocol results in higher quality membranes than a single step to the maximum temperature, where feathering patterns resulting from bubble development during the curing process were observed. While optional, the AMF increases throughput significantly beyond what is possible using weights in an oven.
Troubleshooting the resulting Organ Chips takes place at two levels: during the fabrication process and during Organ Chip culture. We have developed a visual method for quality assurance (QA) of through-hole formation in the cast membranes that greatly accelerates the production process while improving the quality and reliability of assembled Organ Chips. This QA method allows for process troubleshooting, and we recommend keeping a record of process conditions to enable tracking fabrication problems that may occur during cell culture. During Organ Chip culture, inert tracer dyes are the simplest method of measuring barrier function to troubleshoot the fabrication process and cell culture steps. Lucifer Yellow has been used historically due to its small molecular mass and innate fluorescence, but Cascade Blue offers similar properties with a narrower emission spectrum that is less likely to interfere with downstream assays. Larger molecules, such as poly-ethyleneglycol (PEG)- or dextran-conjugated fluorophores are larger and consequently result in lower permeability overall and lower sensitivity. The apparent permeability (Papp, cm/s) of tracer dyes can be used to determine barrier function properties of organs or tissues (Figure 5). The following equation derived by Tran, et al.19 can be used to calculate Papp between the dosing channel and receiving channel, which partially corrects for tracer dye loss caused by absorption into PDMS by averaging the two output flows and not relying on mass balance assumptions at the outflow.
<img alt="Equation 1" src="/files/ftp_upload/58151/58151eq1v2.jpg" />
Vr is the volume in mL of receiving channel effluent after time t; Vd is the volume in mL of the dosing channel effluent after time t; A is the area of membrane through-hole region in cm2 (0.167 cm2 for this device); t is the time of effluent collection in seconds; Cr is the measured change in concentration of the tracer dye in the receiving channel effluent; Cd is the measured concentration of the tracer dye in the dosing channel effluent. Key assumptions for this equation to be valid include: 1) steady tracer dye dosing concentration over time t, 2) the concentration of Cr is small compared to Cd, and 3) the permeability of the system is uniformly distributed across the culture region. Although this equation can be used for static systems, care must be taken to check that the assumptions hold true. Electrical methods, including trans-epithelial electrical resistance (TEER) are commonly implemented in Transwell studies and recently have been incorporated into PDMS Organ Chips for instant and continuous barrier function measurements as well20,21.

Limitations of this protocol include the elasticity of PDMS as well as the manual casting and assembly process that limits production rates. PDMS is a versatile polymer that is well-suited for Organ Chips requiring mechanical strain actuation, but its elasticity can hinder production. Parts can be difficult to handle without deformation and membranes require backing films for manipulation. As a result, automation of Organ Chip production can be limited. The casting process, unlike hot embossing or injection molding used for thermoplastic polymers, is batch-based and therefore also limits throughput.
Organ Chips enable in vitro studies of human organ- and body-level functions in vivo by perfusing a common medium through the vascular channels. By reconstituting physiological tissue-tissue interfaces, flux of molecules between the vascular and parenchymal compartments, mechanical cues, and fluidic shear and transport, these devices promote histodifferentiation and are capable of recapitulating in vivo-like functions of both normal and diseased organs. The compartmentalization of tissues and fluids in two compartments mimics their in vivo functions, and Organ Chip studies are amenable to time-resolved pharmacokinetic experimentation and modeling as well as in vitro-in vivo extrapolation9,10 that is difficult or impossible in single channel MPS14,15,16. The microchannel structures can be leveraged for other applications, including investigating the impact of dynamic tobacco smoke exposure with bidirectional breathing in human small airway epithelium to develop novel biomarkers of lung damage22. The defined positions of the planar membranes and high optical clarity of the devices make them uniquely suited for image-based analyses and integration of embedded sensors. The mechanical stimulation enabled by integrated vacuum channels and elastomeric materials provides functionalities not possible in Transwell systems. We have demonstrated that mechanical strain is essential for recapitulation of certain in vivo physiological functions, including nanoparticle absorption in the lung4, pulmonary edema3 and differentiation of mature iPS-derived glomerular podocytes8.
Future applications of this protocol may include integration of various sensing modalities that can be used to provide real-time readouts of Organ Chip response to stimuli such as drugs, toxins, or radiation. The protocol presented here could be extended to non-PDMS materials with different optical, mechanical, and chemical properties, including biodegradable materials. The Organ Chip protocol presented here should enable researchers to fabricate devices that offer a high degree of control over the microenvironment of healthy and pathophysiologic tissues and organs, which can be leveraged for therapeutic development, including target discovery, toxicity and pharmacokinetic assessments, as well as for personalized medicine.

An erratum was issued for: <a href="https://www-jove-com.remotexs.ntu.edu.sg/video/58151/scalable-fabrication-stretchable-dual-channel-microfluidic-organ">Scalable Fabrication of Stretchable, Dual Channel, Microfluidic Organ Chips</a>.&#160; The Representative Results, Discussion, and References sections have been updated.
In the Representative Results section, the legend for Figure 5 has been updated&#160;from:
Figure 5: Permeability of inert tracer Cascade Blue through the microporous PDMS membrane. Cascade Blue hydrazide dye in medium was loaded into the top channel of the Organ Chip and perfused at 60 &#181;L/h to measure the flux of the dye across the membrane into the bottom channel containing medium. Empty chips were compared to Gut Chips with Caco2-BBe1 cells in the apical channel and human vascular endothelial cells (HUVEC) in the basal channel cultured for 6 days. Error bars indicate standard error of the mean.
to:
Figure 5: Permeability of inert tracer Cascade Blue through the microporous PDMS membrane. Cascade Blue hydrazide dye in medium was loaded into the top channel of the Organ Chip and perfused at 60 &#181;l/h to measure the flux of the dye across the membrane into the bottom channel containing medium. Empty chips were compared to Gut Chips with Caco2-BBe1 cells in the apical channel and human vascular endothelial cells (HUVEC) in the basal channel cultured for 6 days. The apparent permeability (Papp, cm/s) of the microporous PDMS membrane was determined using the dye concentration in the outlet channels. The gut chip cell layers provide a significantly increased barrier to permeability. Error bars indicate standard error of the mean.
In the Discussion section, the fourth paragraph has been updated from:
Troubleshooting the resulting Organ Chips takes place at two levels: during the fabrication process and during Organ Chip culture. We have developed a visual method for quality assurance (QA) of through-hole formation in the cast membranes that greatly accelerates the production process while improving the quality and reliability of assembled Organ Chips. This QA method allows for process troubleshooting, and we recommend keeping a record of process conditions to enable tracking fabrication problems that may occur during cell culture. During Organ Chip culture, inert tracer dyes are the simplest method of measuring barrier function to troubleshoot the fabrication process and cell culture steps. Lucifer Yellow has been used historically due to its small molecular mass and innate fluorescence, but Cascade Blue offers similar properties with a narrower emission spectrum that is less likely to interfere with downstream assays. Larger molecules, such as poly-ethyleneglycol (PEG)- or dextran-conjugated fluorophores are larger and consequently result in lower permeability overall and lower sensitivity. The apparent permeability (Papp, cm/s) of tracer dyes can be used to determine barrier function properties of organs or tissues (Figure 4). The following equation can be used to calculate Papp between the dosing channel and receiving channel and is derived from equations used primarily for Transwell studies19,20 and corrects for tracer dye loss caused by absorption into PDMS by comparing the two output flows and not relying on mass balance assumptions at the outflow.
<img alt="Equation 1" src="/files/ftp_upload/58151/58151eq1.jpg" />
to:
Troubleshooting the resulting Organ Chips takes place at two levels: during the fabrication process and during Organ Chip culture. We have developed a visual method for quality assurance (QA) of through-hole formation in the cast membranes that greatly accelerates the production process while improving the quality and reliability of assembled Organ Chips. This QA method allows for process troubleshooting, and we recommend keeping a record of process conditions to enable tracking fabrication problems that may occur during cell culture. During Organ Chip culture, inert tracer dyes are the simplest method of measuring barrier function to troubleshoot the fabrication process and cell culture steps. Lucifer Yellow has been used historically due to its small molecular mass and innate fluorescence, but Cascade Blue offers similar properties with a narrower emission spectrum that is less likely to interfere with downstream assays. Larger molecules, such as poly-ethyleneglycol (PEG)- or dextran-conjugated fluorophores are larger and consequently result in lower permeability overall and lower sensitivity. The apparent permeability (Papp, cm/s) of tracer dyes can be used to determine barrier function properties of organs or tissues (Figure 5). The following equation derived by Tran, et al.19 can be used to calculate Papp between the dosing channel and receiving channel, which partially corrects for tracer dye loss caused by absorption into PDMS by averaging the two output flows and not relying on mass balance assumptions at the outflow.
<img alt="Equation 1" src="/files/ftp_upload/58151/58151eq1v2.jpg" />
The References section has been updated&#160;from:
<ol start="19">
	<li>Blaser, D.W. Determination of drug absorption parameters in Caco-2 cell monolayers with a mathematical model encompassing passive diffusion, carrier-mediated efflux, non-specific binding and phase II metabolism. at &#60;http://edoc.unibas.ch/655/1/DissB_7998.pdf&#62; (2007).</li>
	<li>Hubatsch, I., Ragnarsson, E.G.E., Artursson, P. Determination of drug permeability and prediction of drug absorption in Caco-2 monolayers. Nature Protocols. 2 (9), 2111-2119 (2007).</li>
	<li>Henry, O.Y.F. et al. Organs-on-chips with integrated electrodes for trans-epithelial electrical resistance (TEER) measurements of human epithelial barrier function. Lab on a Chip. 17 (13), 2264-2271 (2017).</li>
	<li>Maoz, B.M. et al. Organs-on-Chips with combined multi-electrode array and transepithelial electrical resistance measurement capabilities. Lab on a Chip. 17 (13), 2294-2302 (2017).</li>
	<li>Benam, K.H. et al. Matched-Comparative Modeling of Normal and Diseased Human Airway Responses Using a Microengineered Breathing Lung Chip. Cell Systems. 3 (5), 456-466.e4 (2016).</li>
</ol>
to:
<ol start="19">
	<li>Tran, T.T. et al. Exact kinetic analysis of passive transport across a polarized confluent MDCK cell monolayer modeled as a single barrier. Journal of Pharmaceutical Sciences. 93 (8), 2108&#8211;2123 (2004).</li>
	<li>Henry, O.Y.F. et al. Organs-on-chips with integrated electrodes for trans-epithelial electrical resistance (TEER) measurements of human epithelial barrier function. Lab on a Chip. 17 (13), 2264-2271 (2017).</li>
	<li>Maoz, B.M. et al. Organs-on-Chips with combined multi-electrode array and transepithelial electrical resistance measurement capabilities. Lab on a Chip. 17 (13), 2294-2302 (2017).</li>
	<li>Benam, K.H. et al. Matched-Comparative Modeling of Normal and Diseased Human Airway Responses Using a Microengineered Breathing Lung Chip. Cell Systems. 3 (5), 456-466.e4 (2016).</li>
</ol>

An erratum was issued for: <a href="https://www-jove-com.remotexs.ntu.edu.sg/video/58151/scalable-fabrication-stretchable-dual-channel-microfluidic-organ">Scalable Fabrication of Stretchable, Dual Channel, Microfluidic Organ Chips</a>.&#160; The Representative Results, Discussion, and References sections have been updated.

Erratum: Scalable Fabrication of Stretchable, Dual Channel, Microfluidic Organ Chips

Here, we describe the fabrication of dual channel, vascularized Organ-on-a-Chip (Organ Chip) microfluidic culture devices using a scalable protocol amenable for use by research groups lacking access to cleanrooms and traditional soft lithography tools. These devices have been developed to recapitulate human organ-level functions for understanding normal and disease physiology, as well as drug responses in vitro1,2. Critical to engineering this functionality are two perfused microfluidic channels separated by a semi-permeable membrane (Figure 1). This design enables recreation of tissue-tissue interfaces between at least two types of tissues, typically organ parenchymal cells on one side of the porous membrane and vascular endothelium on the other, as well as their exposure to fluid flow. In addition, because the elastomeric polymer, poly (dimethyl siloxane) (PDMS), is used to fabricate the Organ Chip body and membrane components, cyclic mechanical strain can be applied to the entire engineered tissue-tissue interface via the elastic membrane to mimic the natural physical microenvironment of living organs, such as breathing motions in the lung and peristalsis in the intestine.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58151/58151fig1.jpg" /> 
Figure 1: Organ Chip cross section. Organ Chips consist of two channels separated by a porous, elastic membrane that can be seeded with cells on both sides. Top channel cross sections are 1 mm wide x 1 mm high, bottom channel cross sections are 1 mm wide x 0.2 mm high, and vacuum channels in both and bottom parts are 0.3 mm wide, 0.5 mm high, and spaced 0.3 mm from the fluidic channels.&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58151/58151fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
These stretchable, dual channel, Organ Chips have been used for demonstrating the impact of breathing motion on nanoparticle absorption in the lung and drug-induced pulmonary edema3,4; effects of peristaltic motion on differentiation5 and bacterial overgrowth in the intestine5,6,7; and influence of cyclic deformations due to the pulsation of the heart on differentiation and maturation of glomerular podocytes in the kidney8. Additionally, these two-lumen devices that contain an endothelium-lined vascular channel separated by an extracellular matrix (ECM)-coated membrane from parenchymal cells within a separately accessible channel are well suited for characterization of drug PK parameters and new target discovery, which has been limited in single perfusion channel systems. Moreover, multiple Organ Chips may be linked together via their vascular channels to effectively create a human body-on-chips, which could offer an attractive human in vitro platform for therapeutics development9,10. Unlike most micro-physiological systems (MPS)11,12,13, the Organ Chips contain two microfluidic channels separated by a porous membrane that facilitates vascular-parenchymal interactions to recapitulate in vivo organ function. This not only simplifies linking of different organs together by perfusing a common medium through the vascular channels, but the compartmentalization of tissues and fluids mimics in vivo functions and supports pharmacokinetic experimentation and modeling as well as in vitro-in vivo extrapolation9,10 that is difficult or impossible in single channel MPS14,15,16. The popularity of PDMS in microfluidic devices has led to the development of tools to overcome the material&#39;s inherent ability to absorb small molecules10,17. However, the large numbers of chips required to support biological studies where the use of microbial agents and PDMS-absorbing compounds make reuse of Organ Chips difficult necessitates a scalable manufacturing process even for small research groups. The protocol described here presents a method for the device fabrication suitable for the use in academic laboratories, including those lacking access to cleanrooms and soft lithography. This protocol aims to broaden access to Organ Chips by a broad range of researchers seeking to use the stretchable, dual-channel devices for exploring basic biological processes as well as translational therapeutic development.
Leveraging best practices from micromanufacturing fields coupled with design for manufacturing, a robust approach was developed for fabricating Organ Chip devices in large quantities with high reproducibility and yield. The fabrication protocol described here provides a scalable method for Organ Chip production. We describe the use of an optional Mold-in-Place Jig (MiP; design details in Supplemental Materials) coupled with polyurethane gasket strips to enable scaling up of casting PDMS components. The glossy side of polyurethane strips produce optically smooth PDMS parts while the textured side facilitates demolding. We also describe the use of an optional Automated Membrane Fabricator (AMF) that provides uniform compression of membrane wafer molds during curing for fabricating up to 24 membranes per batch. The design is broadly applicable for studies of organs that are composed of tissues that experience mechanical strain and perfusion, and these chips can be produced with low chip-to-chip variability in quantities required to meet the needs of small and large research groups alike. The workflow is amenable to a batch or assembly line format, and readily compatible with quality assessment protocols for control of production processes, personnel training, and responsive troubleshooting. We hope that this protocol will expand access to the capabilities of dual channel, stretchable, Organ Chips for basic and translational research.

<table border="0" cellpadding="0" cellspacing="0" style="width:1212px;" width="1211">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:501px;">Personal Protective Equipment</td>
			<td style="width:179px;"></td>
			<td style="width:133px;"></td>
			<td style="width:399px;"></td>
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		<tr height="20">
			<td height="20" style="height:20px;">Hairnet</td>
			<td>VWR</td>
			<td>89107-770</td>
			<td></td>
		</tr>
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			<td height="20" style="height:20px;">Tyvek lab coat</td>
			<td>VWR</td>
			<td>13450-506</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Extended cuff gloves</td>
			<td>VWR</td>
			<td>89521-898</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Equipment</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Cutting mat</td>
			<td>VWR</td>
			<td>102096-430</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Tile cutter</td>
			<td>McMaster-Carr</td>
			<td>26765A31</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Mold-in-place (MIP) top molds</td>
			<td>Protolabs, Inc.</td>
			<td>custom</td>
			<td>printed in Prototherm 12120</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Mold-in-place (MIP) bottom molds</td>
			<td>Protolabs, Inc.</td>
			<td>custom</td>
			<td>printed in Prototherm 12121</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Duckbill curved forceps</td>
			<td>VWR</td>
			<td>63041-864</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Sharp tipped forceps</td>
			<td>Electron Microscopy Sciences</td>
			<td>72700-D</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Metal spatula</td>
			<td>VWR</td>
			<td>&#160;82027-528</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Deep reactive ion etch (DRIE)&#160; pillar array wafers</td>
			<td>Sensera, Inc.</td>
			<td>custom</td>
			<td>Four 50 x 50 mm pillar arrays per wafer; pillars 7 um wide, 50 um tall, spaced hexagonally 40 um apart</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Textured polycarbonate .01&#8221; thick</td>
			<td>McMaster-Carr</td>
			<td>85585K33</td>
			<td>cut to 45 mm square</td>
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		<tr height="20">
			<td height="20" style="height:20px;">PDMS blocks (40 x 40 x 5 mm)</td>
			<td>n/a</td>
			<td>custom</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Laminar flow hood</td>
			<td>Germfree</td>
			<td>BVBI</td>
			<td>cast in-house</td>
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		<tr height="20">
			<td height="20" style="height:20px;">Air gun</td>
			<td></td>
			<td></td>
			<td></td>
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		<tr height="20">
			<td height="20" style="height:20px;">60&#176;C level oven</td>
			<td></td>
			<td></td>
			<td></td>
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		<tr height="20">
			<td height="20" style="height:20px;">Vacuum desiccator</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Mass balance</td>
			<td></td>
			<td></td>
			<td>accuracy to 0.1 g</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Plasma machine</td>
			<td>Diener</td>
			<td>Nano</td>
			<td>oxygen plasma capability is critical</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Supplies</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Sylgard 184 poly (dimethylsiloxane) (PDMS) base/curing agent kit</td>
			<td>Ellsworth Adhesives&#160;</td>
			<td>4019862</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Mixing cup</td>
			<td></td>
			<td></td>
			<td>Ensure adequate ventilation when handling prepolymer due to low levels of ethylbenzene</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">1 mL syringe</td>
			<td>VWR</td>
			<td>10099-395</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Cleanroom wipes</td>
			<td>VWR</td>
			<td>TWTX1080</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">25 x 75 mm glass microscope slides</td>
			<td>VWR</td>
			<td>48311-703</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Packing tape</td>
			<td>VWR</td>
			<td>500043-724</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Scotch tape</td>
			<td>VWR</td>
			<td>500026-873</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Die-cut Polyurethane (PU) strips</td>
			<td>Atlantic Gasket, Inc.</td>
			<td>custom: AGWI2X3</td>
			<td>&#160;1/8&#8221; thick; 60 Durometer Black Polyurethane; 2&#8221; x 3&#8221;</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Polycarbonate film .005&#8221; thick</td>
			<td>McMaster-Carr</td>
			<td>85585K102</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">100 x 100 x 15 mm square gridded petri dishes</td>
			<td>VWR</td>
			<td>60872-480</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">&#160;Aluminum foil</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Optional Equipment</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Thinky PDMS Mixer</td>
			<td>Thinky</td>
			<td>ARE-310</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Mold-in place (MIP) jig</td>
			<td>in-house</td>
			<td></td>
			<td>screw clamp compression jig</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Automated membrane fabricator (AMF)</td>
			<td>in-house</td>
			<td></td>
			<td>pneumatic compression piston array with programmable heater</td>
		</tr>
	</tbody>
</table>

scalable fabrication of stretchable, dual channel, microfluidic organ chips

A significant number of lead compounds fail in the pharmaceutical pipeline because animal studies often fail to predict clinical responses in human patients. Human Organ-on-a-Chip (Organ Chip) microfluidic cell culture devices, which provide an experimental in vitro platform to assess efficacy, toxicity, and pharmacokinetic (PK) profiles in humans, may be better predictors of therapeutic efficacy and safety in the clinic compared to animal studies. These devices may be used to model the function of virtually any organ type and can be fluidically linked through common endothelium-lined microchannels to perform in vitro studies on human organ-level and whole body-level physiology without having to conduct experiments on people. These Organ Chips consist of two perfused microfluidic channels separated by a permeable elastomeric membrane with organ-specific parenchymal cells on one side and microvascular endothelium on the other, which can be cyclically stretched to provide organ-specific mechanical cues (e.g., breathing motions in lung). This protocol details the fabrication of flexible, dual channel, Organ Chips through casting of parts using 3D printed molds, enabling combination of multiple casting and post-processing steps. Porous poly (dimethyl siloxane) (PDMS) membranes are cast with micrometer sized through-holes using silicon pillar arrays under compression. Fabrication and assembly of Organ Chips involves equipment and steps that can be implemented outside of a traditional cleanroom. This protocol provides researchers with access to Organ Chip technology for in vitro organ- and body-level studies in drug discovery, safety and efficacy testing, as well as mechanistic studies of fundamental biological processes.

The protocol presented here describes the scalable fabrication of PDMS Organ Chips. These devices enable culture of two distinct perfused tissue types on an elastic porous membrane (Figure 1). The PDMS channels are cast using 3D printed molds, which accelerates prototyping of new designs (Figure&#160;2A and 2B). Top channels are cast in molds under compression against a compliant polyurethane gasket to produce components with molded ports (Figure 2C) while bottom channel components are cast in trays and handled on microscope slide backing (Figure 2D). This fabrication approach combines multi-scale patterning of the parts into a single step, which saves time, improves reproducibility and traceability, and reduces debris generated by port punching and multiple cutting steps. The porous membranes are critical to the function of the Organ Chip, and the fabrication approach based on casting against patterned silicon wafers results in membranes of consistent thickness and surface finish (Figure 3). Handling via polycarbonate carriers allows for larger batch production and storage.
The assembled Organ Chip (Figure 4) consists of two perfusion channels in an optically transparent package. In the overlapping region, a porous PDMS membrane enables tissue-tissue interaction of metabolites, proteins, therapeutics, pathogens, and cells to recapitulate organ chip function while two parallel channels on either side are used to provide mechanical strain using cyclic vacuum actuation. The porosity of the PDMS membrane biomimetically supports the flux of metabolites, growth factors, and even cells between the vasculature and organ parenchyma (Figure 5). The apparent permeability (Papp, cm/s) of the membrane was determined using the dye concentration in the outlet channels with and without Caco2 gut cells. The gut chip cell layers provide a significantly increased barrier to permeability. The Organ Chip can be actuated using the parallel vacuum channels to quantitatively and reproducibly apply cyclic strain loading to the membrane and therefore the cultured tissues (Figure 6). This cyclic strain combined with media perfusion supports cellular differentiation to better mimic in vivo organ physiology, such as formation of villi in the Gut Chip.
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/58151/58151fig2.jpg" /> 
Figure 2: Channel fabrication with 3D printed molds. Organ chip parts are cast against high resolution 3D printed molds (A and B), which allows for greater design versatility and prototyping than traditional soft lithography. Top channel parts (C) are cured under compression eliminating the need for punching ports in the finished parts. Each triplicate casting is singulated with a single cut. Bottom channel parts (D) are placed on glass slides to facilitate ease of use and imaging. Scale bars are approximately 1 cm in all images. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58151/58151fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/58151/58151fig3.jpg" /> 
Figure 3: The porous PDMS membrane is cast using DRIE patterned silicon wafers. (A) Rendering of&#160;7 &#181;m diameter, 50 &#181;m tall micropillars etched using DRIE into a silicon wafer. (B) PDMS is cured on this array under 4 kg of compression (16 kPa) to create a 50 &#181;m thick membrane with an array of 7 &#181;m diameter though holes spaced hexagonally 40 &#181;m apart. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58151/58151fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/58151/58151fig4.jpg" /> 
Figure 4: Photograph of an assembled PDMS Organ Chip. Red dye fills the larger apical channel used for parenchymal cells while the blue dye highlights the basal channel typically used for vascular endothelium. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58151/58151fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/58151/58151fig5.jpg" /> 
Figure 5: Permeability of inert tracer Cascade Blue through the microporous PDMS membrane.&#160;Cascade Blue hydrazide dye in medium was loaded into the top channel of the Organ Chip and perfused at 60 &#181;l/h to measure the flux of the dye across the membrane into the bottom channel containing medium. Empty chips were compared to Gut Chips with Caco2-BBe1 cells in the apical channel and human vascular endothelial cells (HUVEC) in the basal channel cultured for 6 days. The apparent permeability (Papp, cm/s) of the microporous PDMS membrane was determined using the dye concentration in the outlet channels. The gut chip cell layers provide a significantly increased barrier to permeability. Error bars indicate standard error of the mean. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58151/58151fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/58151/58151fig6.jpg" /> 
Figure 6: Application of membrane strain using vacuum side channels. Plot indicates linear strain modulation of membrane in response to an applied vacuum pressure. Cyclic uniaxial strain is applied uniformly to the culture region of the Organ Chip using applied vacuum to parallel side channels. The Strain correlates linearly with decreasing vacuum pressure at approximately 1% strain for every -10 kPa change in vacuum pressure (R2 = 0.992). Error bars indicate standard deviation of the mean.&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58151/58151fig5large.jpg" style="font-size: 14px;" target="_blank">Please click here to view a larger version of this figure.</a>
Supplemental Materials:&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58151/MIP_Jig.7z" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Scalable Fabrication of Stretchable, Dual Channel, Microfluidic Organ Chips at JoVE.com

Scalable Fabrication of Stretchable, Dual Channel, Microfluidic Organ Chips

Here, we present a protocol that describes the fabrication of stretchable, dual channel, organ chip microfluidic cell culture devices for recapitulating organ-level functionality in vitro.

A significant number of lead compounds fail in the pharmaceutical pipeline because animal studies often fail to predict clinical responses in human ...

scalable-fabrication-stretchable-dual-channel-microfluidic-organ

Nanyang Technological University

Research

JoVE Journal

Bioengineering

26.5K Views.  Harvard University. Here, we present a protocol that describes the fabrication of stretchable, dual channel, organ chip microfluidic cell culture devices for recapitulating organ-level functionality in vitro.

Video: Scalable Fabrication of Stretchable, Dual Channel, Microfluidic Organ Chips

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 ...

Microfluidic Chip Fabrication and Method to Detect Influenza

Fast and effective diagnostics play an important role in controlling infectious disease by enabling effective patient management and treatment. Here, we present an integrated microfluidic thermoplastic chip with the ability to amplify influenza A virus in patient nasopharyngeal (NP) swabs and aspirates. Upon loading the patient sample, the microfluidic device sequentially carries out on-chip cell lysis, RNA purification and concentration steps within the solid phase extraction (SPE), reverse transcription (RT) and polymerase chain reaction (PCR) in RT-PCR chambers, respectively. End-point detection is performed using an off-chip Bioanalyzer (Agilent Technologies, Santa Clara, CA). For peripherals, we used a single syringe pump to drive reagent and samples, while two thin film heaters were used as the heat sources for the RT and PCR chambers. The chip is designed to be single layer and suitable for high throughput manufacturing to reduce the fabrication time and cost. The microfluidic chip provides a platform to analyze a wide variety of virus and bacteria, limited only by changes in reagent design needed to detect new pathogens of interest.

An integrated microfluidic thermoplastic chip has been developed for use as a molecular diagnostic. The chip performs nucleic acid extraction, reverse transcriptase, and PCR. Methods for fabricating and running the chip are described.

Fast  and effective diagnostics play an important role in controlling infectious  disease by enabling effective patient management and treatment. Here, we ...

Microfluidic Picoliter Bioreactor for Microbial Single-cell Analysis: Fabrication, System Setup, and Operation

In this protocol the fabrication, experimental setup and basic operation of the recently introduced microfluidic picoliter bioreactor (PLBR) is described in detail. The PLBR can be utilized for the analysis of single bacteria and microcolonies to investigate biotechnological and microbiological related questions concerning, e.g.&#160;cell growth, morphology, stress response, and metabolite or protein production on single-cell level. The device features continuous media flow enabling constant environmental conditions for perturbation studies, but in addition allows fast medium changes as well as oscillating conditions to mimic any desired environmental situation. To fabricate the single use devices, a silicon wafer containing sub micrometer sized SU-8 structures served as the replication mold for rapid polydimethylsiloxane casting. Chips were cut, assembled, connected, and set up onto a high resolution and fully automated microscope suited for time-lapse imaging, a powerful tool for spatio-temporal cell analysis. Here, the biotechnological platform organism Corynebacterium glutamicum&#160;was seeded into the PLBR and cell growth and intracellular fluorescence were followed over several hours unraveling time dependent population heterogeneity on single-cell level, not possible with conventional analysis methods such as flow cytometry. Besides insights into device fabrication, furthermore, the preparation of the preculture, loading, trapping of bacteria, and the PLBR cultivation of single cells and colonies is demonstrated. These devices will add a new dimension in microbiological research to analyze time dependent phenomena of single bacteria under tight environmental control. Due to the simple and relatively short fabrication process the technology can be easily adapted at any microfluidics lab and simply tailored towards specific needs.

In this protocol the fabrication, setup and basic operation of a microfluidic picoliter bioreactor (PLBR) for single-cell analysis of prokaryotic microorganisms is introduced. Industrially relevant microorganisms were analyzed as proof of principle allowing insights into growth rate, morphology, and phenotypic heterogeneity over certain time periods, hardly possible with conventional methods.

In this protocol the fabrication, experimental setup and basic operation of the recently introduced microfluidic picoliter bioreactor (PLBR) is described ...

Microfluidic Fabrication of Polymeric and Biohybrid Fibers with Predesigned Size and Shape

A &#8220;sheath&#8221; fluid passing through a microfluidic channel at low Reynolds number can be directed around another &#8220;core&#8221; stream and used to dictate the shape as well as the diameter of a core stream.&#160;Grooves in the top and bottom of a microfluidic channel were designed to direct the sheath fluid and shape the core fluid.&#160;By matching the viscosity and hydrophilicity of the sheath and core fluids, the interfacial effects are minimized and complex fluid shapes can be formed.&#160;Controlling the relative flow rates of the sheath and core fluids determines the cross-sectional area of the core fluid.&#160;Fibers have been produced with sizes ranging from 300 nm to ~1 mm, and fiber cross-sections can be round, flat, square, or complex as in the case with double anchor fibers. Polymerization of the core fluid downstream from the shaping region solidifies the fibers.&#160;Photoinitiated click chemistries are well suited for rapid polymerization of the core fluid by irradiation with ultraviolet light.&#160;Fibers with a wide variety of shapes have been produced from a list of polymers including liquid crystals, poly(methylmethacrylate), thiol-ene and thiol-yne resins, polyethylene glycol, and hydrogel derivatives. Minimal shear during the shaping process and mild polymerization conditions also makes the fabrication process well suited for encapsulation of cells and other biological components.

Two adjacent fluids passing through a grooved microfluidic channel can be directed to form a sheath around a prepolymer core; thereby&#160;determining both shape and cross-section. Photoinitiated polymerization, such as thiol click chemistry, is well suited for rapidly solidifying the core fluid into a microfiber with predetermined size and shape.

A &#8220;sheath&#8221; fluid passing through a microfluidic channel at low Reynolds number can be directed around another &#8220;core&#8221; stream and ...

Fabrication and Implantation of Miniature Dual-element Strain Gages for Measuring In Vivo Gastrointestinal Contractions in Rodents.

Gastrointestinal dysfunction remains a major cause of morbidity and mortality. Indeed, gastrointestinal (GI) motility in health and disease remains an area of productive research with over 1,400 published animal studies in just the last 5 years. Numerous techniques have been developed for quantifying smooth muscle activity of the stomach, small intestine, and colon. In vitro and ex vivo techniques offer powerful tools for mechanistic studies of GI function, but outside the context of the integrated systems inherent to an intact organism. Typically, measuring in vivo smooth muscle contractions of the stomach has involved an anesthetized preparation coupled with the introduction of a surgically placed pressure sensor, a static pressure load such as a mildly inflated balloon or by distending the stomach with fluid under barostatically-controlled feedback. Yet many of these approaches present unique disadvantages regarding both the interpretation of results as well as applicability for in vivo use in conscious experimental animal models. The use of dual element strain gages that have been affixed to the serosal surface of the GI tract has offered numerous experimental advantages, which may continue to outweigh the disadvantages. Since these gages are not commercially available, this video presentation provides a detailed, step-by-step guide to the fabrication of the current design of these gages. The strain gage described in this protocol is a design for recording gastric motility in rats. This design has been modified for recording smooth muscle activity along the entire GI tract and requires only subtle variation in the overall fabrication. Representative data from the entire GI tract are included as well as discussion of analysis methods, data interpretation and presentation.

Fabrication and Implantation of Miniature Dual-element Strain Gages for Measuring In Vivo Gastrointestinal Contractions in Rodents.

The in vivo measurement of smooth muscle contractions along the gastrointestinal tract of laboratory animals remains a powerful, though underutilized, technique. Flexible, dual element strain gages are not commercially available and require fabrication. This protocol describes the construction of reliable, inexpensive strain gages for acute or chronic implantation in rodents.

Gastrointestinal dysfunction remains a major cause of morbidity and mortality. Indeed, gastrointestinal (GI) motility in health and disease remains an ...

The Multi-organ Chip - A Microfluidic Platform for Long-term Multi-tissue Coculture

The ever growing amount of new substances released onto the market and the limited predictability of current in vitro test systems has led to a high need for new solutions for substance testing. Many drugs that have been removed from the market due to drug-induced liver injury released their toxic potential only after several doses of chronic testing in humans. However, a controlled microenvironment is pivotal for long-term multiple dosing experiments,&#160;as even minor alterations in extracellular conditions may greatly influence the cell physiology. We focused within our research program on the generation of a microengineered bioreactor, which can be dynamically perfused by an on-chip pump and combines at least two culture spaces for multi-organ applications. This circulatory system mimics the in vivo conditions of primary cell cultures better and assures a steadier,&#160;more quantifiable extracellular relay of signals to the cells. 
For demonstration purposes, human liver equivalents, generated by aggregating differentiated HepaRG cells with human hepatic stellate cells in hanging drop plates, were cocultured with human skin punch biopsies for up to 28 days inside the microbioreactor. The use of cell culture inserts enables the skin to be cultured at an air-liquid interface, allowing topical substance exposure. The microbioreactor system is capable of supporting these cocultures at near physiologic fluid flow and volume-to-liquid ratios, ensuring stable and organotypic culture conditions. The possibility of long-term cultures enables the repeated exposure to substances. Furthermore, a vascularization of the microfluidic channel circuit using human dermal microvascular endothelial cells yields a physiologically more relevant vascular model.

Here, we present a protocol to coculture primary cells, tissue models and punch biopsies in a microfluidic multi-organ chip for up to 28 days. Human dermal microvascular endothelial cells, liver aggregates and skin biopsies were successfully combined in a common media circulation.

The ever growing amount of new substances released onto the market and the limited predictability of current in vitro test systems has led to a high need ...

Electrotaxis Studies of Lung Cancer Cells using a Multichannel Dual-electric-field Microfluidic Chip

The behavior of directional cell migration under a direct current electric-field (dcEF) is referred to as electrotaxis. The significant role of physiological dcEF in guiding cell movement during embryo development, cell differentiation, and wound healing has been demonstrated in many studies. By applying microfluidic chips to an electrotaxis assay, the investigation process is shortened and experimental errors are minimized. In recent years, microfluidic devices made of polymeric substances (e.g., polymethylmethacrylate, PMMA, or acrylic) or polydimethylsiloxane (PDMS) have been widely used in studying the responses of cells to electrical stimulation. However, unlike the numerous steps required to fabricate a PDMS device, the simple and rapid construction of the acrylic micro&#64258;uidic chip makes it suitable for both device prototyping and production. Yet none of the reported devices facilitate the efficient study of the simultaneous chemical and dcEF effects on cells. In this report, we describe our design and fabrication of an acrylic-based multichannel dual-electric-field (MDF) chip to investigate the concurrent effect of chemical and electrical stimulation on lung cancer cells. The MDF chip provides eight combinations of electrical/chemical stimulations in a single test. The chip not only greatly shortens the required experimental time but also increases accuracy in electrotaxis studies.

Many microfluidic devices have been developed for use in the study of electrotaxis. Yet, none of these chips allows the efficient study of the simultaneous chemical and electric-field (EF) effects on cells. We developed a polymethylmethacrylate-based device that offers better-controlled coexisting EF and chemical stimulation for use in electrotaxis research.

The behavior of directional cell migration under a direct current electric-field (dcEF) is referred to as electrotaxis. The significant role of ...

Image-guided, Laser-based Fabrication of Vascular-derived Microfluidic Networks

This detailed protocol outlines the implementation of image-guided, laser-based hydrogel degradation for the fabrication of vascular-derived microfluidic networks embedded in PEGDA hydrogels. Here, we describe the creation of virtual masks that allow for image-guided laser control; the photopolymerization of a micromolded PEGDA hydrogel, suitable for microfluidic network fabrication and pressure head-driven flow; the setup and use of a commercially available laser scanning confocal microscope paired with a femtosecond pulsed Ti:S laser to induce hydrogel degradation; and the imaging of fabricated microfluidic networks using fluorescent species and confocal microscopy. Much of the protocol is focused on the proper setup and implementation of the microscope software and microscope macro, as these are crucial steps in using a commercial microscope for microfluidic fabrication purposes that contain a number of intricacies. The image-guided component of this technique allows for the implementation of 3D image stacks or user-generated 3D models, thereby allowing for creative microfluidic design and for the fabrication of complex microfluidic systems of virtually any configuration. With an expected impact in tissue engineering, the methods outlined in this protocol could aid in the fabrication of advanced biomimetic microtissue constructs for organ- and human-on-a-chip devices. By mimicking the complex architecture, tortuosity, size, and density of in vivo vasculature, essential biological transport processes can be replicated in these constructs, leading to more accurate in vitro modeling of drug pharmacokinetics and disease.

This protocol outlines the implementation of image-guided, laser-based hydrogel degradation to fabricate vascular-derived, biomimetic microfluidic networks embedded in poly(ethylene glycol) diacrylate (PEGDA) hydrogels. These biomimetic microfluidic systems may be useful for tissue engineering applications, generation of in vitro disease models, and fabrication of advanced "on-a-chip" devices.

This detailed protocol outlines the implementation of image-guided, laser-based hydrogel degradation for the fabrication of vascular-derived microfluidic ...

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 ...