This work was supported in part by the National Institute of Health under award number R21GM143658 and by the National Science Foundation under grant number 2150798. The content is solely the responsibility of the authors and does not necessarily represent the official views of the funding agencies.

1. Fabrication of the two-piece channel and modular platform base
NOTE: The microfluidic channel is constructed using two parts &#8212; the microfluidic channel &#34;cutout&#34;,&#160;which is razor cut from a poly dimethyl siloxane (PDMS) sheet of defined thickness, and the channel cover, which reversibly bonds to the cutout and forms&#160;the channel. The channel is surrounded by a poly(methyl methacrylate) (PMMA) frame that will acts as a media reservoir (Figure 1). The PMMA frame can also be used to magnetically latch specialized modules for added functionality.
<ol>
	<li>Razor cutting channels from PDMS sheets: 
	NOTE: The microfluidic channel design for this step is provided in Supplementary Figure 1. The channel consists of five segments of length 5 mm each and widths of 10 mm, 5 mm, 2.5 mm, 1.25 mm, and 0.75 mm. The velocity of the collagen solution, injected at a constant flow rate (50-400 &#181;L&#183;min&#8722;1), increases locally along the channel as the channel width is decreased to generate extensional flow.
	<ol>
		<li>Mount a 250 &#181;m thick PDMS sheet onto a plastic carrier sheet and razor-cut the microfluidic design using a craft cutter at a blade depth of 0.5 mm, 1 cm&#183;s&#8722;1 speed, and high force.</li>
		<li>Using an ultrasonic bath, clean the microfluidic channel cutouts for 5 min. Rinse the sonicated channels in deionized (DI) water and dry them on a clean hotplate at 100 &#730;C for 5 min.</li>
		<li>Store the channels in a clean, covered Petri dish until use.</li>
	</ol>
	</li>
	<li>Fabricating the PDMS cover and surface passivation: 
	NOTE: The channel cutout from section 1.1 requires a cover that can be placed on top of it to create an enclosed channel that a COL1 solution can be injected into (Supplementary Figure 1). The cover can be removed after the COL1 has self-assembled. To minimize the chances of the COL1 in the channel binding to the cover, the cover is passivated using bovine serum albumin (BSA). The mold design for the PDMS cover is provided. The mold must be at least 2 mm in thickness and should have a footprint that is equal to the footprint of the microfluidic channel cutout. In this protocol, the mold footprint was 35 mm x 15 mm.
	<ol>
		<li>To prepare the mold, affix a sheet of pressure-sensitive adhesive (PSA) to a 2.5 mm thick sheet of PMMA and laser cut the desired mold shape.</li>
		<li>Clean the laser-cut PMMA mold with a damp, lint-free wipe and remove the backing from the PSA film. Attach the mold to a 100 mm diameter silicon wafer bt pressing down firmly.</li>
		<li>Prepare PDMS in a ratio of 10:1 (base:crosslinker). Mix vigorously for 1 min to ensure proper mixing and degas the mixture in a vacuum chamber to remove air bubbles.</li>
		<li>Pour the degassed PDMS solution into the PMMA/silicon mold and cure on a hotplate at 100 &#730;C for 20 min. Allow the mold to cool, remove the cured PDMS, and trim any excess with a razor blade. 
		NOTE: Make sure the silicon wafer facing sides (flat sides) of the PDMS covers are facing up throughout all the following steps.</li>
		<li>Use a 1 mm diameter biopsy punch to create inlet and outlet holes that correspond to the ends of the microfluidic channel. The channel cutout from step 1.1 may be used as a template to guide the position of the holes.</li>
		<li>Sonicate the cover in isopropanol (IPA) for 5 min, rinse with DI water, dry with a compressed air source, and then store in a clean, covered Petri dish. Place the Petri dish in a UV sterilization chamber (uncovered) for 1 min to sterilize.</li>
		<li>Cover and transfer to a biosafety cabinet (BSC).</li>
		<li>Pipette 300 &#181;L of 40 mg&#8226;mL-1 bovine serum albumin (BSA) in 1x phosphate buffered saline (PBS) onto the PDMS covers and spread the solution evenly using a pipette tip. Place in a refrigerator at 4 &#730;C for at least 4 h before use.</li>
		<li>Move the covers from the refrigerator into a BSC, wash 5x with 1x PBS, and let them dry in air for 10 min. 
		NOTE: The PDMS covers can be stored in the fridge for up to 1 week after coating them with BSA.</li>
	</ol>
	</li>
	<li>Glutaraldehyde treatment of coverslips 
	NOTE: Functionalizing the coverslips with glutaraldehyde covalently bonds COL1 to the coverslip and prevents the hydrogel from detaching.
	<ol>
		<li>Prepare a 2% (v/v) aminopropyl triethoxysilane (APTES) solution in a glass beaker by adding 1 mL of APTES to 49 mL of acetone.</li>
		<li>Dilute a 25% glutaraldehyde solution to 5% in DI water. Make 2 mL of solution for each 24 mm x 50 mm coverslip. For 24 mm x 24 mm or 22 mm x 22 mm coverslips, make 1 mL of solution for each.</li>
		<li>Clean the coverslips in a bath sonicator for 5 min using IPA. Rinse off the IPA from the coverslips using DI water. The IPA is completely rinsed when a smooth film of water can be seen on the coverslip.</li>
		<li>Dry the coverslips on a hot plate for 5 min at 100 &#176;C. Place the dried coverslips into clean Petri dishes, ensuring they do not overlap.</li>
		<li>Using a corona discharge wand, expose the coverslips to a corona discharge for 1 min each. 
		NOTE: This step should be performed in a well-ventilated area or a chemical hood.</li>
		<li>Remove the coverslips from the Petri dish and then immerse into the APTES solution for 10 s within 5 min of corona exposure, ensuring the coverslip is submerged. 
		NOTE: Make sure to keep track of which side was treated with plasma and keep it facing upward.</li>
		<li>Then, immerse the coverslip into acetone for 10 s and dry with compressed air. Place the dry coverslip back into the Petri dish, treated side facing up. 
		NOTE: Repeat steps 1.3.5-1.3.7 for all the coverslips.</li>
		<li>Pipette 1 mL of glutaraldehyde solution onto the surface of each coverslip. Cover as much surface as possible without allowing the solution to spill over the edge of the coverslip. More glutaraldehyde solution can be added if necessary. Make sure not to scratch the surface with the pipette tip.</li>
		<li>Let the coverslips sit in contact with the solution for 30 min and then rinse with DI water for 20 s. Dry the coverslips using compressed air and place them back in the Petri dishes, plasma treated side up. 
		NOTE: The coverslips can be stored for up to 1 week at room temperature (RT).</li>
	</ol>
	</li>
	<li>Laser cutting the modular magnetic base 
	NOTE: The design of the modular magnetic base is provided in Supplementary Figure 1. The modular base serves as a well to hold the media and can also be used to magnetically attach specialized modules, as described in previously published works22,37,38,39.
	<ol>
		<li>Cut out the designs from the PMMA layer using appropriate laser settings such as the number of passes and power. 
		NOTE: The laser settings should be adjusted such that the magnets can be press fitted in the PMMA layer. Each laser is different, and the cut parameters must be optimized experimentally. For a 45 W laser, 100% speed, 100% power, and 3 passes are recommended for cutting 2 mm through 2 mm thick PMMA.</li>
		<li>Wash the laser-cut part using soap and water to remove debris from the laser-cutting process. 
		NOTE: Do not use solvents to clean the laser-cut parts. Solvents may result in the propagation of microcracks in the laser-cut edges.</li>
	</ol>
	</li>
	<li>Assembling the platform
	<ol>
		<li>Push magnets (3/16 in diameter, 1/16 in thick) by hand into the laser-cut base. A soft hammer or screw-driver end may be used to help push in the magnet. The thickness of the magnets must be less than the thickness of the PMMA base to ensure the magnets are flush with the surface of the base. 
		NOTE: The magnets allow the user to add functionality to the platform by attaching additional modules.</li>
		<li>Peel off the backing from the PSA sheet and attach the base to a glutaraldehyde-treated coverslip with the functionalized side facing up.</li>
		<li>Gently place the PDMS channel cutout into the cavity defined by the frame. Press down with wide-tip tweezers to remove air bubbles and ensure conformal contact.</li>
		<li>Place the BSA-treated channel cover on top of the channel cutout, with the BSA side facing down. Ensure the fluid inlet and outlet ports are aligned with the channel.</li>
		<li>The device is ready for the COL1 injection.</li>
	</ol>
	</li>
</ol>
2. Injecting the COL1 solution into the microchannel and removing the cover for cell culture applications
<ol>
	<li>Preparing the COL1 solution
	<ol>
		<li>Place all the required reagents (COL1 stock solution [6 mg&#183;mL&#8722;1], ultrapure water, 10x PBS, 0.1 M NaOH) on ice in the biosafety cabinet.</li>
		<li>Calculate the volume of reagents as follows 
		<img alt="Equation 1" src="/files/ftp_upload/64457/64457eq01v2.jpg" style="margin: auto;" /> 
		<img alt="Equation 2" src="/files/ftp_upload/64457/64457eq02.jpg" style="margin: auto;" /> 
		<img alt="Equation 3" src="/files/ftp_upload/64457/64457eq03.jpg" style="margin: auto;" /> 
		<img alt="Equation 4" src="/files/ftp_upload/64457/64457eq04.jpg" style="margin: auto;" /> 
		NOTE: Therefore, to make 1 mL of 2.5 mg&#183;mL&#8722;1 neutralized collagen from a 6 mg&#183;mL&#8722;1 stock, add 416.67 &#181;L of collagen, 429.16 &#181;L of DI water, 100 &#181;L of 10x PBS, and 54.16 &#181;L of 0.1 M NaOH for a final pH of 7. Add 20 &#181;L of 0.1 M NaOH to achieve a pH of 9.0.</li>
		<li>Add the reagents into an empty 5 mL vial in the following order: i) COL1 stock, ii) DI water, iii) 10x PBS, iv) 0.1 M NaOH. 
		NOTE: Use a displacement pipette to handle the COL1 solution. Significant sample loss may occur if a regular pipette is used, leading to fluctuations in final solution concentration.</li>
		<li>Raise the pH of the solution to the desired pH (typically between 7-9).</li>
	</ol>
	</li>
	<li>Injecting the COL1 solution
	<ol>
		<li>Place a syringe pump, a chilled sterile syringe, chilled neutralized COL1 solution, and a sterile 20 G 90&#730; angle tip Luer lock needle into a biosafety cabinet. Load the COL1 solution into the syringe, taking care to avoid introducing bubbles.</li>
		<li>Attach the 90&#730; 20 G needle tip to the syringe, load the syringe into the syringe pump, needle facing down, and prime the needle with the COL1 solution.</li>
		<li>Set the syringe pump to the required flow rate, between 50-2,000 &#181;L/min.</li>
		<li>Place prepared PDMS channels (section 1) on a lab jack and level with the needle.</li>
		<li>Insert the needle into the inlet port of the PDMS channel (wide side). Inject the channel until a ~30 &#181;L drop of COL1 collects on the outlet side.</li>
		<li>Lower the lab jack and gently separate the needle from the newly filled channel.</li>
		<li>Repeat steps 2.2.5-2.2.9 until all channels have been filled with COL1 solution.</li>
		<li>Load the filled channels into the Petri dishes alongside a clean, lint-free wipe saturated with DI water to prevent dehydration of the newly formed COL1 gel.</li>
		<li>Cover the Petri dish and place the loaded channels in the incubator (37 &#730;C, 95% humidity) for 2 h before the peel-off step.</li>
	</ol>
	</li>
	<li>Peel off and media equilibration
	<ol>
		<li>Expose the polymerized COL1 gel by lifting off the PDMS cover using tweezers. The BSA treatment prevents COL1 from attaching to the cover.</li>
		<li>Add 650 &#181;L of EGM to the well.</li>
		<li>Leave the devices in the incubator (37 &#730;C, 95% humidity) for a minimum of 4 h to equilibrate the gel and media. Replace the media before seeding with cells</li>
	</ol>
	</li>
	<li>Seeding cells
	<ol>
		<li>Place warm 0.25% trypsin and culture media along with the required number of 5 mL and 10 mL pipettes in the biosafety cabinet.</li>
		<li>Culture human umbilical vein endothelial cells (HUVECs) to 80% confluency in endothelial growth media (EGM) at 37 &#730;C, in 95% humidity. Place the tissue culture flask containing HUVECs in the BSC after checking confluency under a microscope.</li>
		<li>Discard the media in the T25 flask and wash the cells 2x with 1x PBS. Add 1 mL of Trypsin to the flask and place it in the incubator (37 &#730;C, 95% humidity) for 3 min.</li>
		<li>Check the flask under the microscope after 3 min to ensure that the cells are completely detached from the surface.</li>
		<li>Add 3 mL of EGM (with serum) to the flask to neutralize the trypsin. Next, transfer the cell solution using a 5 mL pipette into a 15 mL conical tube. Centrifuge the conical tube containing cells at 150 x g for 5 min.</li>
		<li>Place the conical tube in the biosafety cabinet and discard the supernatant without disturbing the cell pellet. Resuspend the cells in 1 mL of fresh culture media.</li>
		<li>Add and mix 15 &#181;L of trypan blue and 15 &#181;L of the resuspended cell solution in a 1 mL conical tube.</li>
		<li>Inject trypan blue and cell solution on both sides of a cell counting slide and insert the slide into a cell counting device.</li>
		<li>Dilute the cell solution in endothelial growth media to the required concentration based on the cell concentration obtained from the cell counting device. 
		NOTE: A T25 flask of HUVECs at 80% confluency will yield ~750,000 cells&#183;ml&#8722;1 if diluted in 1 mL of media. The volume of the cell suspension to be seeded is calculated as area of the culture surface &#215; cell density/concentration of cell suspension. Example: To seed 20,000 cells&#183;cm&#8722;2 in the 35 mm x 15 mm well, one will need ~140 &#181;L of the cell suspension.</li>
		<li>Aspirate the media on the engineered COL1 matrix.</li>
		<li>Add the required volume of the cell solution onto the COL1 substrate and let the cells settle for a minimum of 4 h before imaging.</li>
	</ol>
	</li>
	<li>Labeling the cell nucleus and cytoskeleton
	<ol>
		<li>Aspirate the media from the cells and wash 3x with 1x PBS, 500 &#181;L in each wash. Cover the cell layer with 4% paraformaldehyde for 15 min at RT.</li>
		<li>Aspirate the paraformaldehyde and wash with 1x PBS Tween-20 for 5 min.</li>
		<li>Permeabilize the cell membrane using 500 &#181;L of 0.1% Triton-X solution in PBS for 15 min. Wash with 1x PBS Tween-20 for 5 min.</li>
		<li>Block the non-specific binding sites with 500 &#181;L of 4% BSA in PBS for 30 min at RT.</li>
		<li>Dilute the phalloidin-actin fluorescent label solution in 4% BSA (1 &#181;L of stock in 400 &#181;L of BSA).</li>
		<li>Aspirate the BSA solution, add the phalloidin solution to the cells, and wait for 30 min at RT.</li>
		<li>Dilute the nuclear stain in 4% BSA (1 &#181;L in 500 &#181;L), aspirate the phalloidin, add the working nuclear stain solution, and wait for 15 min at RT.</li>
		<li>Aspirate the nuclear stain working solution, wash with PBS Tween-20 3x for 5 min each, and replace with 1x PBS before imaging.</li>
		<li>Image using an epifluorescent microscope using the FITC channel (ex 491 nm/em 516 nm) and DAPI channel (ex 360 nm/em 460 nm) with a 40x lens. Image the collagen using a laser scanning confocal in the reflectance mode using the 488 nm laser line (15% power) and 40x water-immersion objective.</li>
	</ol>
	</li>
</ol>

All authors declare no competing interests.

Protocols to generate COL1 matrices with aligned fibers have been described using magnetic methods, the direct application of mechanical strain, and microfluidic techniques47. Microfluidic approaches are commonly used to create microphysiological systems because of their well-defined flow and transport characteristics, which enable precise control over the biochemical microenvironment. Since aligned COL1 fibers provide key instructive cues during pathophysiological processes such as wound healing,...

Cells reside in a complex 3D fibrous network called the extracellular matrix (ECM), the bulk of which is composed of the structural protein collagen type I (COL1)1,2. The biophysical properties of the ECM provide guidance cues to cells, and in response, cells remodel the ECM microarchitecture3,4,5. These reciprocal cell-matrix interactions can give rise to aligned COL1 fiber domains6 that promote angiogenesis and cell invasion in the tumor environment7,8,9 and influence cell morphology10,11,12, polarization13, and differentiation14. Aligned collagen fibers also promote wound healing15, play a key role in tissue development16, and contribute to long-range cell communication17,18. Therefore, replicating the native COL1 fiber microarchitecture in vitro is an important step toward developing structured models to study cell responses to aligned microenvironments.
Microfluidic cell culture systems have been established as a preferred technology to develop microphysiological systems (MPS)19,20,21,22,23. Leveraging favorable microscale scaling effects, these systems provide precise control over fluid flows, support the controlled introduction of mechanical forces, and define the biochemical microenvironment within a microchannel21,24,25,26,27. MPS platforms have been used to model tissue-specific microenvironments and study multi-organ interactions28. Simultaneously, hydrogels have been widely explored to recapitulate the 3D mechanics and biological influence of the ECM that are observed in vivo29,30. With a growing emphasis on integrating 3D culture with microfluidic platforms, numerous approaches can combine COL1 hydrogels in microfluidic devices31,32,33. However, the methods to align COL1 hydrogels in microfluidic channels have been limited to thin 2D &#34;mats&#34; (&#60;40 &#181;m in thickness) in channels &#60;1 mm wide, offering limited potential to model cell responses in aligned 3D microenvironments31,34,35,36.
To achieve aligned 3D COL1 hydrogels in a microfluidic system, it has been shown that, when a self-assembling COL1 solution is exposed to local extensional flows (velocity change along the streamwise direction), the resulting COL1 hydrogels display a degree of fiber alignment that is directly proportional to the magnitude of the extensional strain rate they experience37,38. The microchannel design in this protocol is unique in two ways; first, the segmented design introduces local extensional strain to the COL1 solution, and second, its &#34;two-piece&#34; construction allows the user to align COL1 fibers and then disassemble the channel to directly access the aligned fibers in an open format. This approach can further be adopted to develop modular microfluidic platforms that develop microphysiological systems with ordered COL1 matrices. The following protocol describes the process of fabricating segmented microchannels and details the use of the channels to align bovine atelo COL1. This protocol also provides instructions for culturing cells on COL1 in an open well format and discusses adding functionality to the platform using a modular, magnetic base layer.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>(3-Aminopropyl)triethoxysilane, 99% (APTES)</td>
			<td>Sigma Aldrich</td>
			<td>440140-100ML</td>
			<td></td>
		</tr>
		<tr>
			<td>20 Gauge IT Series Angled Dispensing Tip</td>
			<td>Jensen Global</td>
			<td>JG-20-1.0-90</td>
			<td></td>
		</tr>
		<tr>
			<td>3/16&#34; dia. x 1/16&#34; thick Nickel Plated Magnet</td>
			<td>KJ Magnetics</td>
			<td>D31</td>
			<td></td>
		</tr>
		<tr>
			<td>3M (TC) 12X12-6-467MP</td>
			<td>DigiKey</td>
			<td>3M9726-ND</td>
			<td></td>
		</tr>
		<tr>
			<td>ACETONE ACS REAGENT &#8805;99.5%</td>
			<td>Signa Aldrich</td>
			<td>179124-4L</td>
			<td></td>
		</tr>
		<tr>
			<td>BD-20AC LABORATORY CORONA TREATER</td>
			<td>Electro-Technic Products</td>
			<td>12051A</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin (BSA), Fraction V, 98%, Reagent Grade, Alfa Aesar</td>
			<td>VWR</td>
			<td>AAJ64100-09</td>
			<td></td>
		</tr>
		<tr>
			<td>Clear cast acrylic sheet</td>
			<td>McMaster-Carr</td>
			<td>8560K181</td>
			<td></td>
		</tr>
		<tr>
			<td>Corning 100 mL Trypsin 10x, 2.5% Trypsin in HBSS [-] calcium, magnesium, phenol red, Porcine Parvovirus Tested</td>
			<td>VWR</td>
			<td>45000-666</td>
			<td></td>
		</tr>
		<tr>
			<td>Countess II Automated Cell Counter</td>
			<td>Thermo Fisher Scientific</td>
			<td>AMQAX1000</td>
			<td></td>
		</tr>
		<tr>
			<td>CT-FIRE software</td>
			<td>LOCI - University of Wisconsin</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>EGM-2 Endothelial Cell Growth Medium-2 BulletKit, (CC-3156 &#38; CC-4176), Lonza CC-3162, 500 mL</td>
			<td>Lonza</td>
			<td>CC-3162</td>
			<td></td>
		</tr>
		<tr>
			<td>Glutaraldehyde 50% in aqueous solution, Reagent Grade, Packaging=HDPE Bottle, Size=100 mL</td>
			<td>VWR</td>
			<td>VWRV0875-100ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Graphtec CELITE-50</td>
			<td>Graphtec</td>
			<td>CE LITE-50</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES (1 M)</td>
			<td>Thermo Fisher Scientific</td>
			<td>15-630-080</td>
			<td></td>
		</tr>
		<tr>
			<td>High-Purity Silicone Rubber .010&#34; Thick, 6&#34; X 8&#34; Sheet, 55A Durometer</td>
			<td>McMaster-Carr</td>
			<td>87315K62</td>
			<td></td>
		</tr>
		<tr>
			<td>Human Umbilical Vein Endothelial cells</td>
			<td>Thermo Fisher Scientific</td>
			<td>C0035C</td>
			<td></td>
		</tr>
		<tr>
			<td>Invitrogen&#160;Trypan Blue Stain (0.4%)</td>
			<td>Thermo Fisher Scientific</td>
			<td>T10282</td>
			<td></td>
		</tr>
		<tr>
			<td>Isopropanol</td>
			<td>Fisher Scientific</td>
			<td>A4154</td>
			<td></td>
		</tr>
		<tr>
			<td>Laser cutter</td>
			<td>Full Spectrum</td>
			<td>20x12 H-series</td>
			<td></td>
		</tr>
		<tr>
			<td>Microfluidics Syringe pump</td>
			<td>New Era Syringe Pumps</td>
			<td>NE-1002X</td>
			<td></td>
		</tr>
		<tr>
			<td>Microman E Single Channel Pipettor, Gilson, Model M1000E</td>
			<td>Gilson</td>
			<td>FD10006</td>
			<td></td>
		</tr>
		<tr>
			<td>Molecular Probes Alexa Fluor 488 Phalloidin</td>
			<td>Thermo Fisher Scientific</td>
			<td>A12379</td>
			<td></td>
		</tr>
		<tr>
			<td>Molecular Probes Hoechst 33342, Trihydrochloride, Trihydrate</td>
			<td>Thermo Fisher Scientific</td>
			<td>H3570</td>
			<td></td>
		</tr>
		<tr>
			<td>Nutragen Bovine Atelo Collagen</td>
			<td>Advanced BioMatrix</td>
			<td>5010-50ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Pbs (10x), pH 7.4</td>
			<td>VWR</td>
			<td>70011044.00</td>
			<td></td>
		</tr>
		<tr>
			<td>PBS pH 7.4</td>
			<td>Thermo Fisher Scientific</td>
			<td>10010049.00</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate-buffered saline (PBS, 10x), with Triton X-100</td>
			<td>Alfa Aesar</td>
			<td>J63521</td>
			<td></td>
		</tr>
		<tr>
			<td>Replacement carrier sheet for graphtec craft ROBO CC330L-20</td>
			<td>USCUTTER</td>
			<td>GRPCARSHTN</td>
			<td></td>
		</tr>
		<tr>
			<td>Restek Norm-Ject Plastic Syringe 1 mL Luer Slip</td>
			<td>Restek</td>
			<td>22766.00</td>
			<td></td>
		</tr>
		<tr>
			<td>Silicon wafer</td>
			<td>University wafer</td>
			<td>452</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Hydroxide, ACS, Packaging=Poly Bottle, Size=500 g</td>
			<td>VWR</td>
			<td>BDH9292-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>Sylgard 184</td>
			<td>VWR</td>
			<td>102092-312</td>
			<td></td>
		</tr>
		<tr>
			<td>Thermo Scientific&#160;Pierce 20x PBS Tween 20</td>
			<td>Thermo Fisher Scientific</td>
			<td>28352.00</td>
			<td></td>
		</tr>
	</tbody>
</table>

microengineering 3d collagen hydrogels with long-range fiber alignment

Aligned collagen I (COL1) fibers guide tumor cell motility, influence endothelial cell morphology, control stem cell differentiation, and are a hallmark of cardiac and musculoskeletal tissues. To study cell response to aligned microenvironments in vitro, several protocols have been developed to generate COL1 matrices with defined fiber alignment, including magnetic, mechanical, cell-based, and microfluidic methods. Of these, microfluidic approaches offer advanced capabilities such as accurate control over fluid flows and the cellular microenvironment. However, the microfluidic approaches to generate aligned COL1 matrices for advanced in vitro culture platforms have been limited to thin "mats" (&lt;40 &#181;m in thickness) of COL1 fibers that extend over distances less than 500 &#181;m and are not conducive to 3D cell culture applications. Here, we present a protocol to fabricate 3D COL1 matrices (130-250 &#181;m in thickness) with millimeter-scale regions of defined fiber alignment in a microfluidic device. This platform provides advanced cell culture capabilities to model structured tissue microenvironments by providing direct access to the micro-engineered matrix for cell culture.

When a self-assembling COL1 solution flows through a channel with decreasing cross-sectional area, the streamwise velocity (vx) of the COL1 solution increases locally by a magnitude, &#8706;vx, along the length of the constriction between the two segments (&#8706;x), resulting in an extensional strain rate (&#949;&#775;) where &#949;&#775; = &#8706;vx/&#8706;x. The extensional strain rate can be calculated from the fluid velocity, which is measured using particle image velocimetry (PIV), ...

Watch this Scientific Journal Video about Microengineering 3D Collagen Hydrogels with Long-Range Fiber Alignment at JoVE.com

Microengineering 3D Collagen Hydrogels with Long-Range Fiber Alignment

This protocol demonstrates the use of a microfluidic channel with changing geometry along the fluid flow direction to generate extensional strain (stretching) to align fibers in a 3D collagen hydrogel (&lt;250 &#181;m in thickness). The resulting alignment extends across several millimeters and is influenced by the extensional strain rate.

Aligned collagen I (COL1) fibers guide tumor cell motility, influence endothelial cell morphology, control stem cell differentiation, and are a hallmark ...

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Research

JoVE Journal

Bioengineering

2.1K Views.  Rochester Institute of Technology. This protocol demonstrates the use of a microfluidic channel with changing geometry along the fluid flow direction to generate extensional strain (stretching) to align fibers in a 3D collagen hydrogel (

Video: Microengineering 3D Collagen Hydrogels with Long-Range Fiber Alignment

Electrospun Nanofiber Scaffolds with Gradations in Fiber Organization

The goal of this protocol is to report a simple method for generating nanofiber scaffolds with gradations in fiber organization and test their possible applications in controlling cell morphology/orientation. Nanofiber organization is controlled with a new fabrication apparatus that enables the gradual decrease of fiber organization in a scaffold. Changing the alignment of fibers is achieved through decreasing deposition time of random electrospun fibers on a uniaxially aligned fiber mat. By covering the collector with a moving barrier/mask, along the same axis as fiber deposition, the organizational structure is easily controlled. For tissue engineering purposes, adipose-derived stem cells can be seeded to these scaffolds. Stem cells undergo morphological changes as a result of their position on the varied organizational structure, and can potentially differentiate into different cell types depending on their locations. Additionally, the graded organization of fibers enhances the biomimicry of nanofiber scaffolds so they more closely resemble the natural orientations of collagen nanofibers at tendon-to-bone insertion site compared to traditional scaffolds. Through nanoencapsulation, the gradated fibers also afford the possibility to construct chemical gradients in fiber scaffolds, and thereby further strengthen their potential applications in fast screening of cell-materials interaction and interfacial tissue regeneration. This technique enables the production of continuous gradient scaffolds, but it also can potentially produce fibers in discrete steps by controlling the movement of the moving barrier/mask in a discrete fashion.

Here, we present a protocol to fabricate electrospun nanofiber scaffolds with gradated organization of fibers and explore their applications in regulating cell morphology/orientation. Gradients with regard to physical and chemical properties of the nanofiber scaffolds offer a wide variety of applications in the biomedical field.

The goal of this protocol is to report a simple method for generating nanofiber scaffolds with gradations in fiber organization and test their possible ...

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. ...

Planar Gradient Diffusion System to Investigate Chemotaxis in a 3D Collagen Matrix

The importance of cell migration can be seen through the development of human life. When cells migrate, they generate forces and transfer these forces to their surrounding area, leading to cell movement and migration. In order to understand the mechanisms that can alter and/or affect cell migration, one can study these forces. In theory, understanding the fundamental mechanisms and forces underlying cell migration holds the promise of effective approaches for treating diseases and promoting cellular transplantation. Unfortunately, modern chemotaxis chambers that have been developed are usually restricted to two dimensions (2D) and have complex diffusion gradients that make the experiment difficult to interpret. To this end, we have developed, and describe in this paper, a direct-viewing chamber for chemotaxis studies, which allows one to overcome modern chemotaxis chamber obstacles able to measure cell forces and specific concentration within the chamber in a 3D environment to study cell 3D migration. More compelling, this approach allows one to successfully model diffusion through 3D collagen matrices and calculate the coefficient of diffusion of a chemoattractant through multiple different concentrations of collagen, while keeping the system simple and user friendly for traction force microscopy (TFM) and digital volume correlation (DVC) analysis.

Cell migration is an important part of human development and life. In order to understand the mechanisms that can alter cell migration, we present a planar gradient diffusion system to investigate chemotaxis in a 3D collagen matrix, which allows one to overcome modern diffusion chamber limitations of existing assays.

The importance of cell migration can be seen through the development of human life. When cells migrate, they generate forces and transfer these forces to ...

Engineered 3D Silk-collagen-based Model of Polarized Neural Tissue

Despite huge efforts to decipher the anatomy, composition and function of the brain, it remains the least understood organ of the human body. To gain a deeper comprehension of the neural system scientists aim to simplistically reconstruct the tissue by assembling it in vitro from basic building blocks using a tissue engineering approach. Our group developed a tissue-engineered silk and collagen-based 3D brain-like model resembling the white and gray matter of the cortex. The model consists of silk porous sponge, which is pre-seeded with rat brain-derived neurons, immersed in soft collagen matrix. Polarized neuronal outgrowth and network formation is observed with separate axonal and cell body localization. This compartmental architecture allows for the unique development of niches mimicking native neural tissue, thus enabling research on neuronal network assembly, axonal guidance, cell-cell and cell-matrix interactions and electrical functions.

Insight into the complex actions of the brain requires advanced research tools. Here we demonstrate a novel silk-collagen-based 3D engineered model of neural tissue resembling brain-like architecture. The model can be used to study neuronal network assembly, axonal guidance, cell-cell interactions and electrical activity.

Despite huge efforts to decipher the anatomy, composition and function of the brain, it remains the least understood organ of the human body. To gain a ...

Three-dimensional Biomimetic Technology: Novel Biorubber Creates Defined Micro- and Macro-scale Architectures in Collagen Hydrogels

Tissue scaffolds play a crucial role in the tissue regeneration process. The ideal scaffold must fulfill several requirements such as having proper composition, targeted modulus, and well-defined architectural features. Biomaterials that recapitulate the intrinsic architecture of in vivo tissue are vital for studying diseases as well as to facilitate the regeneration of lost and malformed soft tissue. A novel biofabrication technique was developed which combines state of the art imaging, three-dimensional (3D) printing, and selective enzymatic activity to create a new generation of biomaterials for research and clinical application. The developed material, Bovine Serum Albumin rubber, is reaction injected into a mold that upholds specific geometrical features. This sacrificial material allows the adequate transfer of architectural features to a natural scaffold material. The prototype consists of a 3D collagen scaffold with 4 and 3 mm channels that represent a branched architecture. This paper emphasizes the use of this biofabrication technique for the generation of natural constructs. This protocol utilizes a computer-aided software (CAD) to manufacture a solid mold which will be reaction injected with BSA rubber followed by the enzymatic digestion of the rubber, leaving its architectural features within the scaffold material.

An innovative biofabrication technique was developed to engineer three-dimensional constructs that resemble the architectural features, components, and mechanical properties of in vivo tissue. This technique features a newly developed sacrificial material, BSA rubber, which transfers detailed spatial features, reproducing the in vivo architectures of a wide variety of tissues.

Tissue scaffolds play a crucial role in the tissue regeneration process. The ideal scaffold must fulfill several requirements such as having proper ...

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

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

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

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

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

Tunable Hydrogels from Pulmonary Extracellular Matrix for 3D Cell Culture

Here we present a method for establishing multiple component cell culture hydrogels for in vitro lung cell culture. Beginning with healthy en bloc lung tissue from porcine, rat, or mouse, the tissue is perfused and submerged in subsequent chemical detergents to remove the cellular debris. Histological comparison of the tissue before and after processing confirms removal of over 95% of double stranded DNA and alpha galactosidase staining suggests the majority of cellular debris is removed. After decellularization, the tissue is lyophilized and then cryomilled into a powder. The matrix powder is digested for 48 hr in an acidic pepsin digestion solution and then neutralized to form the pregel solution. Gelation of the pregel solution can be induced by incubation at 37 &#176;C and can be used immediately following neutralization or stored at 4 &#176;C for up to two weeks. Coatings can be formed using the pregel solution on a non-treated plate for cell attachment. Cells can be suspended in the pregel prior to self-assembly to achieve a 3D culture, plated on the surface of a formed gel from which the cells can migrate through the scaffold, or plated on the coatings. Alterations to the strategy presented can impact gelation temperature, strength, or protein fragment sizes. Beyond hydrogel formation, the hydrogel stiffness may be increased using genipin.

This is a method to create a 3-dimensional cell culture scaffold from pulmonary extracellular matrix. Intact lung is processed into hydrogels that can support the growth of cells in three-dimensions.

Here we present a method for establishing multiple component cell culture hydrogels for in vitro lung cell culture. Beginning with healthy en bloc lung ...

Production of Elastin-like Protein Hydrogels for Encapsulation and Immunostaining of Cells in 3D

Two-dimensional (2D) tissue culture techniques have been essential for our understanding of fundamental cell biology. However, traditional 2D tissue culture systems lack a three-dimensional (3D) matrix, resulting in a significant disconnect between results collected in vitro and in vivo. To address this limitation, researchers have engineered 3D hydrogel tissue culture platforms that can mimic the biochemical and biophysical properties of the in vivo cell microenvironment. This research has motivated the need to develop material platforms that support 3D cell encapsulation and downstream biochemical assays. Recombinant protein engineering offers a unique toolset for 3D hydrogel material design and development by allowing for the specific control of protein sequence and therefore, by extension, the potential mechanical and biochemical properties of the resultant matrix. Here, we present a protocol for the expression of recombinantly-derived elastin-like protein (ELP), which can be used to form hydrogels with independently tunable mechanical properties and cell-adhesive ligand concentration. We further present a methodology for cell encapsulation within ELP hydrogels and subsequent immunofluorescent staining of embedded cells for downstream analysis and quantification.

Recombinant protein-engineered hydrogels are advantageous for 3D cell culture as they allow for complete tunability of the polymer backbone and therefore, the cell microenvironment. Here, we describe the process of recombinant elastin-like protein purification and its application in 3D hydrogel cell encapsulation.

Two-dimensional (2D) tissue culture techniques have been essential for our understanding of fundamental cell biology. However, traditional 2D tissue ...

Measuring Global Cellular Matrix Metalloproteinase and Metabolic Activity in 3D Hydrogels

Three-dimensional (3D) cell culture systems often more closely recapitulate in vivo cellular responses and functions than traditional two-dimensional (2D) culture systems. However, measurement of cell function in 3D culture is often more challenging. Many biological assays require retrieval of cellular material which can be difficult in 3D cultures. One way to address this challenge is to develop new materials that enable measurement of cell function within the material. Here, a method is presented for measurement of cellular matrix metalloproteinase (MMP) activity in 3D hydrogels in a 96-well format. In this system, a poly(ethylene glycol) (PEG) hydrogel is functionalized with a fluorogenic MMP cleavable sensor. Cellular MMP activity is proportional to fluorescence intensity and can be measured with a standard microplate reader. Miniaturization of this assay to a 96-well format reduced the time required for experimental set up by 50% and reagent usage by 80% per condition as compared to the previous 24-well version of the assay. This assay is also compatible with other measurements of cellular function. For example, a metabolic activity assay is demonstrated here, which can be conducted simultaneously with MMP activity measurements within the same hydrogel. The assay is demonstrated with human melanoma cells encapsulated across a range of cell seeding densities to determine the appropriate encapsulation density for the working range of the assay. After 24 h of cell encapsulation, MMP and metabolic activity readouts were proportional to cell seeding density. While the assay is demonstrated here with one fluorogenic degradable substrate, the assay and methodology could be adapted for a wide variety of hydrogel systems and other fluorescent sensors. Such an assay provides a practical, efficient and easily accessible 3D culturing platform for a wide variety of applications.

Here, a protocol is presented for encapsulating and culturing cells in poly(ethylene glycol) (PEG) hydrogels functionalized with a fluorogenic matrix metalloproteinase (MMP)-degradable peptide. Cellular MMP and metabolic activity are measured directly from the hydrogel cultures using a standard microplate reader.

Three-dimensional (3D) cell culture systems often more closely recapitulate in vivo cellular responses and functions than traditional two-dimensional (2D) ...

Controlled Strain of 3D Hydrogels under Live Microscopy Imaging

External forces are an important factor in tissue formation, development, and maintenance. The effects of these forces are often studied using specialized in vitro stretching methods. Various available systems use 2D substrate-based stretchers, while the accessibility of 3D techniques to strain soft hydrogels, is more restricted. Here, we describe a method that allows external stretching of soft hydrogels from their circumference, using an elastic silicone strip as the sample carrier. The stretching system utilized in this protocol is constructed from 3D-printed parts and low-cost electronics, making it simple and easy to replicate in other labs. The experimental process begins with polymerizing thick (&#62;100 &#956;m) soft fibrin hydrogels (Elastic Modulus of ~100 Pa) in a cut-out at the center of a silicone strip. Silicone-gel constructs are then attached to the printed-stretching device and placed on the confocal microscope stage. Under live microscopy the stretching device is activated, and the gels are imaged at various stretch magnitudes. Image processing is then used to quantify the resulting gel deformations, demonstrating relatively homogenous strains and fiber alignment throughout the gel&#8217;s 3D thickness (Z-axis). Advantages of this method include the ability to strain extremely soft hydrogels in 3D while executing in situ microscopy, and the freedom to manipulate the geometry and size of the sample according to the user&#8217;s needs. Additionally, with proper adaptation, this method can be used to stretch other types of hydrogels (e.g., collagen, polyacrylamide or polyethylene glycol) and can allow for analysis of cells and tissue response to external forces under more biomimetic 3D conditions.

The presented method involves uniaxial stretching of 3D soft hydrogels embedded in silicone rubber while allowing live confocal microscopy. Characterization of the external and internal hydrogel strains as well as fiber alignment are demonstrated. The device and protocol developed can assess the response of cells to various strain regimes.

External forces are an important factor in tissue formation, development, and maintenance. The effects of these forces are often studied using specialized ...