This work was supported in part by grants from the NIH (HL118532, HL120142, CA187692), the David &#38; Lucile Packard Foundation, the Camille &#38; Henry Dreyfus Foundation, and the Burroughs Welcome Fund. A.S.P. was supported in part by a Charlotte Elizabeth Procter Honorific Fellowship.

1. Preparation of Solutions
<ol>
	<li>To prepare a 5 mg/ml solution of insulin, dilute the powdered insulin stock with 5 mM hydrochloric acid (HCl) in dH2O (500 mg insulin in 100 ml solvent). Prepare 100 ml solvent by adding 50 &#956;l of concentrated HCl to 100 ml distilled water (dH2O).</li>
	<li>To make a 1x solution of PBS, dilute the 10x phosphate-buffered saline (PBS) stock solution to 1x with dH2O under sterile conditions.</li>
	<li>Prepare the polydimethylsiloxane (PDMS) elastomer solution as follows: mix the PDMS prepolymer together with the curing agent in a 10:1 (w:w) ratio.</li>
	<li>Prepare the cell culture medium as follows: to 500 ml of stock Dulbecco&#39;s Modified Eagle Medium: Nutrient Mixture F-12 (DMEM/F-12), add 10 ml of sterile fetal bovine serum (FBS), 500 &#956;l of gentamicin reagent, and 500 &#956;l of 5 mg/ml insulin.</li>
	<li>To prepare the 1% bovine serum albumin (BSA) in 1x PBS solution, add 1% (w:v) solution of BSA powder to 1x PBS and mix thoroughly.</li>
	<li>To prepare a 4 mg/ml solution of neutralized bovine type I collagen, add 50 &#181;l 10x Hank&#39;s balanced salt solution (HBSS) buffer, 30 &#181;l 0.1 N sodium hydroxide (NaOH), 30 &#181;l cell culture medium, and 400 &#181;l of stock collagen to a chilled 1.5 ml microcentrifuge tube. Mix slowly by pipetting up and down; avoid introducing bubbles. To remove any bubbles, briefly centrifuge the mixture at 4 &#176;C.</li>
	<li>Prepare a fixative solution by diluting 16% paraformaldehyde 1:4 (v:v) in 1x PBS.</li>
	<li>Prepare a nuclei labeling solution by diluting the stock nuclei labeling solution 1:1,000 (v:v) in 1x PBS.</li>
	<li>Prepare a 0.3% phosphate-buffered saline and detergent (PBST) solution by mixing 1x PBS with 0.3% (v:v) detergent.</li>
	<li>Prepare blocking buffer by diluting goat serum 1:10 (v:v) in 0.3% PBST.</li>
	<li>Prepare a primary antibody solution for a particular protein of interest by diluting primary antibody (for example, rabbit anti-focal adhesion kinase (FAK)) 1:200 (v:v) in blocking buffer.</li>
	<li>Prepare a secondary antibody solution by diluting a fluorescent probe-conjugated antibody (for example, goat anti-rabbit) 1:1,000 (v:v) in blocking buffer.</li>
</ol>
2. Preparation of Elastomeric Stamps for 3D Micropatterning
Note: Elastomeric stamps are made with PDMS.
<ol>
	<li>Make a PDMS solution at the same ratio as described in step 1.3 and mix thoroughly. Place the mixture in a vacuum chamber for 15-30 min to remove any air bubbles that were introduced during the mixing process.</li>
	<li>Pour the degassed solution into a plastic weigh boat or Petri dish containing a silicon master that is lithographically patterned with features of desired geometry. For best results, cure the PDMS solution at 60 &#176;C for ~12 hr. 
	Note: The silicon masters are highly customizable; this protocol uses rectangular structures with dimensions of 50 &#956;m x 200 &#956;m x 50 &#956;m spaced 200 &#956;m apart. The silicon masters can be made using standard photolithography techniques. Briefly, an epoxy-based photoresist is spin-coated on top of a silicon wafer. A mask detailing the desired stamp features is placed on top of the photoresist-coated wafer, which is then exposed to UV light. The desired features not blocked by the mask are exposed to the light and the photoresist at these locations becomes crosslinked, while the remainder of the photoresist coating is soluble and can be washed away.</li>
	<li>After the PDMS has cured around the desired features on the silicon master, remove the PDMS and master from the plastic container. Carefully separate the PDMS from the silicon wafer and remove excess PDMS from around the imprinted features using a clean razor blade.</li>
	<li>Now cut the PDMS patterned with imprinted features into individual rectangular stamps (~8-mm x 5-mm) with a razor blade. Store these feature-side-up in a clean 100-mm-diameter Petri dish.</li>
	<li>Additionally, use the PDMS solution described in step 1.3 to make supports for the rectangular stamps.</li>
	<li>Using a spin coater, spread ~2-3 g of the PDMS solution evenly on a 100-mm-diameter Petri dish and cure the PDMS for ~12 hr at 60 &#176;C as in step 2.1. Then, using a razor blade, cut the thin layer of PDMS into rectangles (~5 mm x 2 mm). 
	Note: Two supports are needed for each PDMS stamp made in step 2.3. The supports can be stored in the 100-mm-diameter Petri dish that contains the PDMS stamps.</li>
	<li>Before the PDMS stamps and supports can be used for cell culture, sterilize by immersion in 70% ethanol and dry using an aspirator in a biosafety cabinet (cell culture hood).</li>
</ol>
3. Preparation of 3D Epithelial Tissues
<ol>
	<li>In a biosafety cabinet, add a drop of approximately 50&#160;&#181;l of 1% BSA in PBS to the top of each stamp. Place the droplet covered stamps at 4 &#176;C for a minimum of 4 hr to ensure that the BSA adsorbs to the surface of the stamp.</li>
	<li>Aspirate BSA solution from the PDMS stamps.</li>
	<li>Wash the surfaces of the stamps twice with cell culture medium (50 &#181;l per wash per stamp should be sufficient), aspirating after each wash.</li>
	<li>Handle the sterilized PDMS stamps and supports using curved stainless steel tweezers. In a biosafety cabinet, lay out one 35-mm-diameter tissue culture dish for each PDMS stamp. In each dish, lay down two PDMS supports separated by a distance slightly less than the length of the PDMS stamps.</li>
	<li>Using the tweezers, sterilize as many circular glass coverslips (15 mm in diameter, #1) as there are PDMS stamps using 70% ethanol. Aspirate the excess liquid from the coverslips while holding them with the tweezers. Store the washed coverslips in a separate 100-mm-diameter Petri dish.</li>
	<li>Dispense ~50 &#181;l of the collagen mixture to evenly coat the surface of each PDMS stamp.</li>
	<li>Pick up each collagen-coated PDMS stamp with the tweezers and gently invert them. Lower the inverted stamps on top of the PDMS supports laid out in 35-mm-diameter tissue culture dished such that the collagen is between the stamp and the bottom of the tissue culture dish. Incubate the dishes at 37 &#176;C for 30 min.</li>
	<li>Dispense ~50 &#181;l of the remaining collagen mixture onto each of the circular coverslips (later, these will be placed on top of the engineered tissues to completely encapsulate them in collagen) and incubate them at 37 &#176;C for 30 min.</li>
	<li>Obtain a 100-mm-diameter tissue culture dish containing epithelial cells (ex. EpH4 mouse mammary epithelial cells) at ~40% confluency. Aspirate the cell culture medium and wash once with 10 ml 1x PBS. Add 2 ml of trypsin to the cells and incubate at 37 &#176;C for 5-10 min.</li>
	<li>Add 8 ml of fresh culture medium to the tissue culture dish containing trypsinized cells, detaching any remaining adherent cells from the dish. Mix gently by pipetting and move the cell mixture to a 15 ml conical tube and centrifuge at 100 &#215; g for 5 min.</li>
	<li>Aspirate the supernatant from the conical tube and resuspend the cell pellet in cell culture medium. Using a hemocytometer, count the cells to determine the concentration of the suspension. Adjust the suspension volume to obtain a final concentration of ~106-107cells/ml.</li>
	<li>Remove the gelled collagen samples from the incubator. Using the tweezers, gently lift the PDMS stamps straight upwards to detach them from the molded collagen and discard them.</li>
	<li>Dispense ~30 &#181;l of the concentrated cell suspension onto the surface of each collagen gel containing molded cavities of desired geometry. Observe the cells under a brightfield microscope with a 10X/0.25 NA objective while gently shaking the dishes side to side to promote cell settling within the cavities. The cavities should be filled within ~5 min.</li>
	<li>To remove excess cells from around the cavities, tilt each tissue culture dish on its side and gently dispense ~400 &#956;l cell culture medium over the surface of the collagen gel. Aspirate the liquid and repeat the wash 1-2 more times, checking the collagen gels under the microscope in between each wash.</li>
	<li>After the excess cells have been cleared from around the cell-filled cavities in the collagen mold, place the tissue culture dishes into a cell culture incubator at 37 &#176;C for 15 min. Then, using the tweezers, gently invert the collagen-coated class coverslips and place them on top of the cell-filled collagen molds such that the collagen from the coverslips forms a cap over the cell-filled cavities. Incubate the samples at 37 &#176;C for 15 min.</li>
	<li>Once the collagen caps have adhered to the cell-filled collagen mold, dispense ~2-2.5 ml cell culture medium slowly over the glass coverslip on top of the gels. Culture the samples at 37 &#176;C for 1-3 days. 
	Note: 24 hr after initial seeding, the epithelial tissues can be treated with growth factors such as epidermal growth factor (EGF) or hepatocyte growth factor (HGF) to induce branching.</li>
</ol>
4. Immunofluorescence and Image Analysis&#160;
<ol>
	<li>Aspirate the cell culture medium from the tissue culture dishes and add enough fixative solution to cover the cell-containing gels. Incubate the dishes at room temperature for 15 min on a shaker at 200 rpm.</li>
	<li>Aspirate the fixative, fill the tissue culture dishes with 1x PBS, and incubate at room temperature for 15 min on a shaker at 200 rpm. Repeat twice for three total washes.</li>
	<li>To label nuclei: Aspirate the PBS from the tissue culture dish and replace with Hoechst solution. Incubate at room temperature for 15-20 min. To stain for FAK or another marker that is detectable with antibodies, skip to Step 4.5.</li>
	<li>Aspirate the nuclear labeling solution and wash the cell-containing gels three times with 1x PBS as in Step 4.2. Stained samples can be stored in 1x PBS at 4 &#176;C until further use.</li>
	<li>To stain for a protein of interest: Aspirate the PBS from the tissue culture dishes, add 300 &#956;l of 0.3% PBST, and incubate the sample at room temperature for 15 min.</li>
	<li>Aspirate the PBS, cover the gels with blocking buffer, and incubate on a shaker (200 rpm) at room temperature for ~4 hr.</li>
	<li>Aspirate the blocking buffer, cover the gels with primary antibody solution, and incubate on a shaker at 200 rpm overnight at 4 &#176;C.</li>
	<li>Aspirate the primary antibody solution, add 0.3% PBST solution, and incubate on a shaker at 200 rpm at room temperature for 30 min. Aspirate the PBST and repeat every 30 min for 3-4 hr.</li>
	<li>Repeat steps 4.7 and 4.8, this time incubating with the secondary antibody solution. Wrap the tissue culture dishes with aluminum foil to prevent photobleaching of the secondary antibody. After the final wash, stained samples can be stored in 1x PBS at 4 &#176;C until further use.</li>
	<li>To visualize samples, use a 10X/0.30 NA objective focused on the midplane of the epithelial tissues.</li>
	<li>To visualize cell nuclei, image fixed samples labeled with a nuclear marker using a 10X/0.30 NA objective under UV illumination.</li>
	<li>To visualize samples stained for a protein of interest, image using a 10X/0.30 NA objective on an inverted fluorescence microscope.</li>
	<li>To create frequency maps of protein staining of multiple tissues (typically 50 or more) of identical initial geometry, first import each of the images using standard image analysis software (see Materials) and convert them to 8-bit grayscale.</li>
	<li>To threshold these images, convert them to binary (i.e., halftone/black and white) by defining a grayscale cutoff point; grayscale values below the threshold become black, and those above the threshold become white. Then, combine each of the individual binary images into a single image stack.</li>
	<li>Next, register the stacked images (i.e., align or match them) in the analysis software. Many image analysis software packages come with a freely available registration plugin.
	<ol>
		<li>Use each image in a stack as a template for the alignment of the next image, such that the images in the entire stack are aligned by propagation. Finally, overlay (project) the images in the aligned image stack based on average intensity to form a single image with a pixel frequency map. This image can then be color-coded using image editing software of choice10,11.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Affolter, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tube+or+not+tube:+remodeling+epithelial+tissues+by+branching+morphogenesis.">Tube or not tube: remodeling epithelial tissues by branching morphogenesis.</a> Dev Cell. 4 (1), 11-18 (2003).</li><li>Zhu, W., Nelson, C. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=PI3K+signaling+in+the+regulation+of+branching+morphogenesis.">PI3K signaling in the regulation of branching morphogenesis.</a> Biosystems. 109 (3), 403-411 (2012).</li><li>Sternlicht, M. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Key+stages+in+mammary+gland+development:+the+cues+that+regulate+ductal+branching+morphogenesis.">Key stages in mammary gland development: the cues that regulate ductal branching morphogenesis.</a> Breast Cancer Res. 8 (1), 201 (2006).</li><li>Fata, J. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+MAPK(ERK-1,2)+pathway+integrates+distinct+and+antagonistic+signals+from+TGFalpha+and+FGF7+in+morphogenesis+of+mouse+mammary+epithelium.">The MAPK(ERK-1,2) pathway integrates distinct and antagonistic signals from TGFalpha and FGF7 in morphogenesis of mouse mammary epithelium.</a> Dev Biol. 306 (1), 193-207 (2007).</li><li>Ip, M. M., Darcy, K. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Three-dimensional+mammary+primary+culture+model+systems.">Three-dimensional mammary primary culture model systems.</a> J Mammary Gland Biol Neoplasia. 1 (1), 91-110 (1996).</li><li>Lo, A. T., Mori, H., Mott, J., Bissell, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Constructing+three-dimensional+models+to+study+mammary+gland+branching+morphogenesis+and+functional+differentiation.">Constructing three-dimensional models to study mammary gland branching morphogenesis and functional differentiation.</a> J Mammary Gland Biol Neoplasia. 17 (2), 103-110 (2012).</li><li>Nelson, C. M., Vanduijn, M. M., Inman, J. L., Fletcher, D. A., Bissell, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tissue+geometry+determines+sites+of+mammary+branching+morphogenesis+in+organotypic+cultures.">Tissue geometry determines sites of mammary branching morphogenesis in organotypic cultures.</a> Science. 314 (5797), 298-300 (2006).</li><li>Gjorevski, N., Nelson, C. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Endogenous+patterns+of+mechanical+stress+are+required+for+branching+morphogenesis.">Endogenous patterns of mechanical stress are required for branching morphogenesis.</a> Integr Biol (Camb). 2 (9), 424-434 (2010).</li><li>Gjorevski, N., Nelson, C. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mapping+of+mechanical+strains+and+stresses+around+quiescent+engineered+three-dimensional+epithelial+tissues.">Mapping of mechanical strains and stresses around quiescent engineered three-dimensional epithelial tissues.</a> Biophys J. 103 (1), 152-162 (2012).</li><li>Gjorevski, N., Piotrowski, A. S., Varner, V. D., Nelson, C. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamic+tensile+forces+drive+collective+cell+migration+through+three-dimensional+extracellular+matrices.">Dynamic tensile forces drive collective cell migration through three-dimensional extracellular matrices.</a> Sci Rep. 5, 11458 (2015).</li><li>Zhu, W., Nelson, C. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=PI3K+regulates+branch+initiation+and+extension+of+cultured+mammary+epithelia+via+Akt+and+Rac1+respectively.">PI3K regulates branch initiation and extension of cultured mammary epithelia via Akt and Rac1 respectively.</a> Dev Biol. 379 (2), 235-245 (2013).</li><li>Barcellos-Hoff, M. H., Aggeler, J., Ram, T. G., Bissell, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+differentiation+and+alveolar+morphogenesis+of+primary+mammary+cultures+on+reconstituted+basement+membrane.">Functional differentiation and alveolar morphogenesis of primary mammary cultures on reconstituted basement membrane.</a> Development. 105 (2), 223-235 (1989).</li><li>Hirai, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Epimorphin+functions+as+a+key+morphoregulator+for+mammary+epithelial+cells.">Epimorphin functions as a key morphoregulator for mammary epithelial cells.</a> J Cell Biol. 140 (1), 159-169 (1998).</li><li>Pavlovich, A. L., Manivannan, S., Nelson, C. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adipose+stroma+induces+branching+morphogenesis+of+engineered+epithelial+tubules.">Adipose stroma induces branching morphogenesis of engineered epithelial tubules.</a> Tissue Eng Part A. 16 (12), 3719-3726 (2010).</li></ol>

The authors declare that they have no competing financial interests.

The protocol described above outlines a method to produce identical epithelial tissues of pre-defined shape, enabling spatial control of the mechanical stress experienced by cells in the tissue. An elastomeric mold is used to create cavities in type I collagen that are then filled with epithelial cells and covered with an additional collagen layer such that cells are completely encapsulated in a 3D collagen matrix environment. Further culture of these tissues and treatment with growth factors to induce branching from the initial architecture make this system amenable for the study of branching morphogenesis of epithelial cells. There are several critical steps in the protocol. The first is lifting the PDMS molds straight upwards from the gelled collagen prior to cell seeding. Any horizontal movement during this step will distort the cavities, and they will no longer retain the desired geometry. Then, after the cavities have been seeded with cells, it is important to carefully wash off any excess cells that remain on the surface, as these may interfere with the behavior of the patterned tissues, particularly if they are very close to the tissues. Washing too rigorously, however, may result in cells being washed out of the cavities. Lastly, in the final micropatterning step when the cell-containing collagen gels are being immersed in culture medium, it is important to add the medium slowly and directly on top of the coverslip. Adding medium to the side of the dish away from the coverslip may result in detachment of the coverslip and the collagen lid. Each of these steps needs to be performed with care to achieve the best results.
Over the past few decades, several culture models have emerged for the study of epithelial morphogenesis, many of which involve the ex vivo culture of intact organs or explants of organs. Major advantages of culturing tissue taken directly from an animal are that tissue architecture remains intact and that cell-cell interactions are maintained. However, these models are not readily amenable to studies investigating the specific effects of a particular cell type or physical microenvironment. Additionally, organs and organ explants can be fragile and difficult to culture ex vivo. Others used primary mammary epithelial cells12, in which gene expression can be more readily manipulated, though this method does not conserve in vivo tissue architecture. Moreover, like intact organs, these cells can also be difficult to culture for long periods of time5. There is a need, therefore, for more robust culture models of epithelial morphogenesis that can be more precisely controlled and readily perturbed to gather reproducible data. 
A widely used 3D cell culture technique is the formation of epithelial cell clusters embedded in collagen or other ECM proteins11,13. Disadvantages of this technique, however, include the inability to precisely control or predict the location of clusters within the gel, cluster size and shape, and branching sites. In addition, it is difficult to manipulate or measure the mechanical cues experienced by cells in different regions of the tissue clusters. Engineered tissues enable better control of these parameters.
The engineered tissue protocol described here is an easy to use, versatile, and reproducible 3D cell culture method that eliminates the heterogeneity commonly found in ex vivo culture systems and 3D clusters. Such engineered cell culture models provide a platform for more precise control of physical and biochemical signals, and results can be easily combined with computational models to predict the effects of perturbations or measure cellular forces using techniques such as traction force microscopy (TFM)9,10. A caveat of the model system, however, is that it does not completely replicate the cellular, chemical, and physical microenvironment within the mammary gland in vivo. Nonetheless, essentially all parts of the system are modifiable: including the cells used (in which gene expression can also be altered by transfection), tissue geometry, ECM composition, and other biochemical factors (ex. growth factors). Primary mammary epithelial cells can also be cultured in this system; they form a lumen and undergo branching morphogenesis from the same locations as immortalized cell lines7. Moreover, the model can be modified to incorporate other cell types to more appropriately represent the native mammary gland microenvironment14, or to investigate the underlying mechanisms that give rise to any branched organ.

The development of branched epithelial tissues, known as branching morphogenesis, is regulated by cell-derived, physical, and environmental factors. In the mammary gland, branching morphogenesis is an iterative process through which guided collective cell migration creates a tree-like architecture. The first step is primary bud formation from the milk ducts, followed by branch initiation and elongation1,2. Invasion of branches into the surrounding stroma is induced by the systemic release of steroid hormones at puberty. New primary buds then initiate from the ends of existing branches, and this process continues to create an epithelial tree3. Although many important biochemical signals have been identified, a comprehensive understanding of the cell biological mechanisms that guide this complex developmental process is currently lacking. Moreover, mechanistic studies on the influences of specific cues are difficult to deconstruct from experiments in vivo, as precise spatiotemporal perturbations and measurements are often not possible.
Three-dimensional (3D) culture techniques, such as whole organ culture, primary organoids, and cell culture models, are useful tools for systematically investigating the mechanisms underlying tissue morphogenesis4-6. These can be particularly useful for determining the influences of specific factors individually, such as mechanical forces and biochemical signals, on a variety of cell behaviors, including migration, proliferation, and differentiation.6 Engineered cell culture models, in particular, readily enable the perturbation of individual cells and their microenvironment.
One such culture model uses a microfabrication-based approach to engineer model mammary epithelial tissues with controlled 3D structure that consistently and reproducibly form branches that migrate collectively when induced with the appropriate growth factors. The major advantage of the model is the ability to precisely manipulate and measure the effects of physical and biochemical factors, such as patterns of mechanical stress, with high statistical confidence. This technique, together with computational modeling, has already been used to determine the relative contributions of physical and biochemical signals in the guidance of the normal development of mammary epithelial tissues and other branched epithelia7-11. Presented here is a detailed protocol for building these model tissues, which can be readily extended to other types of cells and extracellular matrix (ECM) gels, and which serves as a potential tool for the testing of therapeutics.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Polydimethylsiloxane (PDMS)</td>
			<td>Ellsworth Adhesives</td>
			<td>Sylgard 184</td>
			<td></td>
		</tr>
		<tr>
			<td>PDMS curing agent</td>
			<td>Ellsworth Adhesives</td>
			<td>Sylgard 184</td>
			<td></td>
		</tr>
		<tr>
			<td>Lithographically patterned silicon master</td>
			<td>self-made</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Plastic weigh boat</td>
			<td>Fisher Scientific</td>
			<td>08-732-115</td>
			<td></td>
		</tr>
		<tr>
			<td>100-mm-diameter Petri dishes</td>
			<td>BioExpress</td>
			<td>D-2550-2</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethyl Alcohol 200 Proof</td>
			<td>Pharmco-Aaper</td>
			<td>111000200</td>
			<td>Make a 70% EtOH (v:v) solution by mixing with dH2O</td>
		</tr>
		<tr>
			<td>Razor blade</td>
			<td>American Safety Razor</td>
			<td>620179</td>
			<td></td>
		</tr>
		<tr>
			<td>1:1 Dulbecco&#8217;s Modified Eagle&#8217;s Medium : Ham&#8217;s F12 Nutrient Mixture (DMEM/F12) (1:1)</td>
			<td>Hyclone</td>
			<td>SH30023FS</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum (FBS)</td>
			<td>Atlanta Biologicals</td>
			<td>S11150H</td>
			<td></td>
		</tr>
		<tr>
			<td>10x Hank&#8217;s balanced salt solution (HBSS)</td>
			<td>Life Technologies</td>
			<td>14185-052</td>
			<td></td>
		</tr>
		<tr>
			<td>Insulin</td>
			<td>Sigma Aldrich</td>
			<td>I6634-500MG</td>
			<td></td>
		</tr>
		<tr>
			<td>Gentamicin</td>
			<td>Life Technologies</td>
			<td>15750-060</td>
			<td></td>
		</tr>
		<tr>
			<td>10X Phosphate-buffered saline (PBS)</td>
			<td>Fisher Scientific</td>
			<td>BP399-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium hydroxide (NaOH)</td>
			<td>Sigma Aldrich</td>
			<td>221465-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine type I collagen (non-pepsinized)</td>
			<td>Koken</td>
			<td>IAC-50</td>
			<td></td>
		</tr>
		<tr>
			<td>Albumin from bovine serum (BSA)</td>
			<td>Sigma Aldrich</td>
			<td>A-7906</td>
			<td></td>
		</tr>
		<tr>
			<td>Curved stainless steel tweezers</td>
			<td>Dumont</td>
			<td>7</td>
			<td></td>
		</tr>
		<tr>
			<td>35-mm-diameter tissue culture dishes</td>
			<td>BioExpress</td>
			<td>T-2881-6</td>
			<td></td>
		</tr>
		<tr>
			<td>15 mL conical tubes</td>
			<td>BioExpress</td>
			<td>C-3394-2</td>
			<td></td>
		</tr>
		<tr>
			<td>1.5 mL Eppendorf Safe-Lock Tube</td>
			<td>USA Scientific</td>
			<td>1615-5500</td>
			<td></td>
		</tr>
		<tr>
			<td>Circular #1 glass coverslips, 15-mm in diameter</td>
			<td>Bellco Glass Inc.</td>
			<td>Special order</td>
			<td></td>
		</tr>
		<tr>
			<td>0.05% 1X Trypsin-EDTA</td>
			<td>Life Technologies</td>
			<td>25300-054</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>VWR</td>
			<td>100503-916</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Perkin Elmer</td>
			<td>N9300260</td>
			<td>Detergent</td>
		</tr>
		<tr>
			<td>HGF</td>
			<td>Sigma Aldrich</td>
			<td>H 9661</td>
			<td>Resuspended in dH2O at 50 mg/mL</td>
		</tr>
		<tr>
			<td>Rabbit anti-mouse FAK antibody</td>
			<td>Life Technologies</td>
			<td>AMO0672</td>
			<td></td>
		</tr>
		<tr>
			<td>Goat anti-rabbit Alexa 488 antibody</td>
			<td>Life Technologies</td>
			<td>A-11034</td>
			<td></td>
		</tr>
		<tr>
			<td>Adobe Photoshop</td>
			<td>Adobe</td>
			<td>N/A</td>
			<td>Used for color-coding pixel frequency maps.</td>
		</tr>
		<tr>
			<td>FIJI (ImageJ)</td>
			<td>NIH</td>
			<td>N/A</td>
			<td>Free image analysis software used for thresholding, registering, and overlaying images to create a pixel frequency map. The StackReg plugin was used for registering binary images.</td>
		</tr>
	</tbody>
</table>

engineering three-dimensional epithelial tissues embedded within extracellular matrix

The architecture of branched organs such as the lungs, kidneys, and mammary glands arises through the developmental process of branching morphogenesis, which is regulated by a variety of soluble and physical signals in the microenvironment. Described here is a method created to study the process of branching morphogenesis by forming engineered three-dimensional (3D) epithelial tissues of defined shape and size that are completely embedded within an extracellular matrix (ECM). This method enables the formation of arrays of identical tissues and enables the control of a variety of environmental factors, including tissue geometry, spacing, and ECM composition. This method can also be combined with widely used techniques such as traction force microscopy (TFM) to gain more information about the interactions between cells and their surrounding ECM. The protocol can be used to investigate a variety of cell and tissue processes beyond branching morphogenesis, including cancer invasion.

General schematic of mammary epithelial tissue microfabrication
A general schematic of the microfabrication procedure outlining the experimental work flow is shown in Figure 1. The end result is an array of epithelial tissues of identical geometry and spacing that are completely embedded within an ECM gel. A representative experiment uses EpH4 mouse mammary epithelial cells cultured in a gel of bovine type I collagen at a concentration of 4 mg/ml. To ensure the highest quality of engineered tissues, the techniques outlined in the protocol should be followed closely. Figures 2A and 2B show low and high magnification views of arrays of rectangular wells that have been molded into a type I collagen gel prior to cell seeding. The shape of the wells is determined by the shape of the features on the silicon master. It is important to lift the PDMS mold straight up from the collagen so as not to distort the cavity geometry. Figure 2C shows rectangular wells in a type I collagen gel that have been filled with mammary epithelial cells (excess cells have been washed off the surface of the collagen). In this example, each 200 &#956;m x 50 &#956;m rectangular well contains approximately 80-100 cells.
Addition of growth factors induces morphogenesis 
24 hours after seeding, the tissues may be treated with growth factors such as HGF or EGF and cultured over several days to model branching morphogenesis. Typically, branches begin to form as early as 4 hours after growth factor stimulation. Figure 3A shows representative results from rectangular mammary epithelial tissues within a type I collagen gel 24 hours after the microfabrication procedure, after which cells have adhered to the collagen and to each other. No branches are observed prior to growth factor addition. Figure 3B shows a representative rectangular tissue that has undergone branching 24 hours after the addition of HGF at 10 ng/ml. In this case, branches occur at the ends of the tissues (as opposed to the middle), where the cells experience the highest mechanical stress9. Multiple tissues of identical initial geometry in the same gel can then be imaged to determine population averages of branch location and branch length, enabling high-throughput analysis.
Immunofluorescence staining to visualize protein localization
Immunofluorescence staining of the tissue arrays in the culture model allows us to determine protein localization within a tissue with high statistical confidence. Figure 4A shows representative results from a branching rectangular mammary epithelial tissue stained for focal adhesion kinase (FAK). Creating frequency maps of tissues of identical geometry can be used to visualize the average spatial localization of proteins of interest within the tissues, which can be compared to the localization of other proteins as well as branching activity. Figure 4B shows a frequency map of average FAK staining for 50 tissues showing FAK enrichment at the short ends of rectangular tissues, where branching typically occurs.
<img alt="Figure 1" src="/files/ftp_upload/54283/54283fig1.jpg" /> 
Figure 1. Schematic outlining the microfabrication procedure. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54283/54283fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/54283/54283fig2.jpg" /> 
Figure 2. Images taken during the microfabrication process. (A) Low and (B) and high magnification phase-contrast images of rectangular cavities in type I collagen created using an elastomeric PDMS mold. (C) Cavities from (A) and (B) are filled with mammary epithelial cells. Scale bars, 100 &#956;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54283/54283fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/54283/54283fig3.jpg" /> 
Figure 3. Microfabricated tissues undergo branching morphogenesis. (A) Phase-contrast image of a representative rectangular tissue 24 hr after microfabrication. (B) Phase-contrast image of a representative rectangular tissue that has started to undergo branching 24 hr after the addition of HGF at 10 ng/ml. White arrows indicate newly formed branches. Scale bars, 100 &#956;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54283/54283fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/54283/54283fig4.jpg" /> 
Figure 4. Immunofluorescence staining of microfabricated tissues. (A) Immunofluorescence staining for FAK in a mammary epithelial tissue after branch initiation. (B) A frequency map of average FAK staining in 50 tissues. Scale bars, 100 &#956;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54283/54283fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Engineering Three-dimensional Epithelial Tissues Embedded within Extracellular Matrix at JoVE.com

Engineering Three-dimensional Epithelial Tissues Embedded within Extracellular Matrix

This manuscript describes a soft lithography-based technique to engineer uniform arrays of three-dimensional (3D) epithelial tissues of defined geometry surrounded by extracellular matrix. This method is amenable to a wide variety of cell types and experimental contexts and allows for high-throughput screening of identical replicates.

The architecture of branched organs such as the lungs, kidneys, and mammary glands arises through the developmental process of branching morphogenesis, ...

engineering-three-dimensional-epithelial-tissues-embedded-within

Nanyang Technological University

Research

JoVE Journal

Bioengineering

7.5K Views.  Princeton University. This manuscript describes a soft lithography-based technique to engineer uniform arrays of three-dimensional (3D) epithelial tissues of defined geometry surrounded by extracellular matrix. This method is amenable to a wide variety of cell types and experimental contexts and allows for high-throughput screening of identical replicates.

Video: Engineering Three-dimensional Epithelial Tissues Embedded within Extracellular Matrix

Three-dimensional Optical-resolution Photoacoustic Microscopy

Optical microscopy, providing valuable insights at the cellular and organelle levels, has been widely recognized as an enabling biomedical technology. As the mainstays of in vivo three-dimensional (3-D) optical microscopy, single-/multi-photon fluorescence microscopy and optical coherence tomography (OCT) have demonstrated their extraordinary sensitivities to fluorescence and optical scattering contrasts, respectively. However, the optical absorption contrast of biological tissues, which encodes essential physiological/pathological information, has not yet been assessable. 
The emergence of biomedical photoacoustics has led to a new branch of optical microscopy optical-resolution photoacoustic microscopy (OR-PAM)1, where the optical irradiation is focused to the diffraction limit to achieve cellular1 or even subcellular2 level lateral resolution. As a valuable complement to existing optical microscopy technologies, OR-PAM brings in at least two novelties. First and most importantly, OR-PAM detects optical absorption contrasts with extraordinary sensitivity (i.e., 100%). Combining OR-PAM with fluorescence microscopy3 or with optical-scattering-based OCT4 (or with both) provides comprehensive optical properties of biological tissues. Second, OR-PAM encodes optical absorption into acoustic waves, in contrast to the pure optical processes in fluorescence microscopy and OCT, and provides background-free detection. The acoustic detection in OR-PAM mitigates the impacts of optical scattering on signal degradation and naturally eliminates possible interferences (i.e., crosstalks) between excitation and detection, which is a common problem in fluorescence microscopy due to the overlap between the excitation and fluorescence spectra. 
Unique for optical absorption imaging, OR-PAM has demonstrated broad biomedical applications since its invention, including, but not limited to, neurology5, 6, ophthalmology7, 8, vascular biology9, and dermatology10. In this video, we teach the system configuration and alignment of OR-PAM as well as the experimental procedures for in vivo functional microvascular imaging.

Optical-resolution photoacoustic microscopy (OR-PAM) is an emerging technology capable of imaging optical absorption contrasts in vivo with cellular resolution and sensitivity. Here, we provide a visualized instruction on the experimental protocols of OR-PAM, including system configuration, system alignment, typical in vivo experimental procedures, and functional imaging schemes.

Optical microscopy, providing valuable insights at the cellular and organelle levels, has been widely recognized as an enabling biomedical technology. As ...

Live-cell Imaging of Migrating Cells Expressing Fluorescently-tagged Proteins in a Three-dimensional Matrix

Traditionally, cell migration has been studied on two-dimensional, stiff plastic surfaces. However, during important biological processes such as wound healing, tissue regeneration, and cancer metastasis, cells must navigate through complex, three-dimensional extracellular tissue. To better understand the mechanisms behind these biological processes, it is important to examine the roles of the proteins responsible for driving cell migration. Here, we outline a protocol to study the mechanisms of cell migration using the epithelial cell line (MDCK), and a three-dimensional, fibrous, self-polymerizing matrix as a model system. This optically clear extracellular matrix is easily amenable to live-cell imaging studies and better mimics the physiological, soft tissue environment. This report demonstrates a technique for directly visualizing protein localization and dynamics, and deformation of the surrounding three-dimensional matrix. 
Examination of protein localization and dynamics during cellular processes provides key insight into protein functions. Genetically encoded fluorescent tags provide a unique method for observing protein localization and dynamics. Using this technique, we can analyze the subcellular accumulation of key, force-generating cytoskeletal components in real-time as the cell maneuvers through the matrix. In addition, using multiple fluorescent tags with different wavelengths, we can examine the localization of multiple proteins simultaneously, thus allowing us to test, for example, whether different proteins have similar or divergent roles. Furthermore, the dynamics of fluorescently tagged proteins can be quantified using Fluorescent Recovery After Photobleaching (FRAP) analysis. This measurement assays the protein mobility and how stably bound the proteins are to the cytoskeletal network.
By combining live-cell imaging with the treatment of protein function inhibitors, we can examine in real-time the changes in the distribution of proteins and morphology of migrating cells. Furthermore, we also combine live-cell imaging with the use of fluorescent tracer particles embedded within the matrix to visualize the matrix deformation during cell migration. Thus, we can visualize how a migrating cell distributes force-generating proteins, and where the traction forces are exerted to the surrounding matrix. Through these techniques, we can gain valuable insight into the roles of specific proteins and their contributions to the mechanisms of cell migration.

Cellular processes such as cell migration have traditionally been studied on two-dimensional, stiff plastic surfaces. This report describes a technique for directly visualizing protein localization and analyzing protein dynamics in cells migrating in a more physiologically relevant, three-dimensional matrix.

Traditionally, cell migration has been studied on two-dimensional, stiff plastic surfaces. However, during important biological processes such as wound ...

Analysis of Cell Migration within a Three-dimensional Collagen Matrix

The ability to migrate is a hallmark of various cell types and plays a crucial role in several physiological processes, including&#160;embryonic development, wound healing, and immune responses. However, cell migration is also a key mechanism in cancer enabling these cancer cells to detach from the primary tumor to start metastatic spreading. Within the past years various cell migration assays have been developed to analyze the migratory behavior of different cell types. Because the locomotory behavior of cells markedly differs between a two-dimensional (2D) and three-dimensional (3D) environment it can be assumed that the analysis of the migration of cells that are embedded within a 3D environment would yield in more significant cell migration data. The advantage of the described 3D collagen matrix migration assay is that cells are embedded within a physiological 3D network of collagen fibers representing the major component of the extracellular matrix. Due to time-lapse video microscopy real cell migration is measured allowing the determination of several migration parameters as well as their alterations in response to pro-migratory factors or inhibitors. Various cell types could be analyzed using this technique, including lymphocytes/leukocytes, stem cells, and tumor cells. Likewise, also cell clusters or spheroids could be embedded within the collagen matrix concomitant with analysis of the emigration of single cells from the cell cluster/ spheroid into the collagen lattice. We conclude that the 3D collagen matrix migration assay is a versatile method to analyze the migration of cells within a physiological-like 3D environment.

Cell migration is a biological phenomenon that is involved in a plethora of physiological, such as wound healing and immune responses, and pathophysiological processes, like cancer. The 3D-collagen matrix migration assay is a versatile tool to analyze the migratory properties of different cell types within in a 3D physiological-like environment.

The ability to migrate is a hallmark of various cell types and plays a crucial role in several physiological processes, including&#160;embryonic ...

Design and Construction of Artificial Extracellular Matrix (aECM) Proteins from Escherichia coli for Skin Tissue Engineering

Recombinant technology is a versatile platform to create novel artificial proteins with tunable properties. For the last decade, many artificial proteins that have incorporated functional domains derived from nature (or created de novo) have been reported. In particular, artificial extracellular matrix (aECM) proteins have been developed; these aECM proteins consist of biological domains taken from fibronectin, laminins and collagens and are combined with structural domains including elastin-like repeats, silk and collagen repeats. To date, aECM proteins have been widely investigated for applications in tissue engineering and wound repair. Recently, Tjin and coworkers developed integrin-specific aECM proteins designed for promoting human skin keratinocyte attachment and propagation. In their work, the aECM proteins incorporate cell binding domains taken from fibronectin, laminin-5 and collagen IV, as well as flanking elastin-like repeats. They demonstrated that the aECM proteins developed in their work were promising candidates for use as substrates in artificial skin. Here, we outline the design and construction of such aECM proteins as well as their purification process using the thermo-responsive characteristics of elastin.

Design and Construction of Artificial Extracellular Matrix aECM Proteins from Escherichia coli for Skin Tissue Engineering

Recombinant technologies have enabled material designers to create novel artificial proteins with customized functionalities for tissue engineering applications. For example, artificial extracellular matrix proteins can be designed to incorporate structural and biological domains derived from native ECMs. Here, we describe the construction and purification of aECM proteins containing elastin-like repeats.

Recombinant technology is a versatile platform to create novel artificial proteins with tunable properties. For the last decade, many artificial proteins ...

Epithelial Cell Repopulation and Preparation of Rodent Extracellular Matrix Scaffolds for Renal Tissue Development

This protocol details the generation of acellular, yet biofunctional, renal extracellular matrix (ECM) scaffolds that are useful as small-scale model substrates for organ-scale tissue development. Sprague Dawley rat kidneys are cannulated by inserting a catheter into the renal artery and perfused with a series of low-concentration detergents (Triton X-100 and sodium dodecyl sulfate (SDS)) over 26 hr to derive intact, whole-kidney scaffolds with intact perfusable vasculature, glomeruli, and renal tubules. Following decellularization, the renal scaffold is placed inside a custom-designed perfusion bioreactor vessel, and the catheterized renal artery is connected to a perfusion circuit consisting of: a peristaltic pump; tubing; and optional probes for pH, dissolved oxygen, and pressure. After sterilizing the scaffold with peracetic acid and ethanol, and balancing the pH (7.4), the kidney scaffold is prepared for seeding via perfusion of culture medium within a large-capacity incubator maintained at 37 &#176;C and 5% CO2. Forty million renal cortical tubular epithelial (RCTE) cells are injected through the renal artery, and rapidly perfused through the scaffold under high flow (25 ml/min) and pressure (~230 mmHg) for 15 min before reducing the flow to a physiological rate (4 ml/min). RCTE cells primarily populate the tubular ECM niche within the renal cortex, proliferate, and form tubular epithelial structures over seven days of perfusion culture. A 44 &#181;M resazurin solution in culture medium is perfused through the kidney for 1 hr during medium exchanges to provide a fluorometric, redox-based metabolic assessment of cell viability and proliferation during tubulogenesis. The kidney perfusion bioreactor permits non-invasive sampling of medium for biochemical assessment, and multiple inlet ports allow alternative retrograde seeding through the renal vein or ureter. These protocols can be used to recellularize kidney scaffolds with a variety of cell types, including vascular endothelial, tubular epithelial, and stromal fibroblasts, for rapid evaluation within this system.

This protocol describes decellularization of Sprague Dawley rat kidneys by antegrade perfusion of detergents through the vasculature, producing acellular renal extracellular matrices that serve as templates for repopulation with human renal epithelial cells. Recellularization and use of the resazurin perfusion assay to monitor growth is performed within specially-designed perfusion bioreactors.

This protocol details the generation of acellular, yet biofunctional, renal extracellular matrix (ECM) scaffolds that are useful as small-scale model ...

Characterizing Cell Migration Within Three-dimensional In Vitro Wound Environments

Currently, most in vitro models of wound healing, such as well-established scratch assays, involve studying cell migration and wound closure on two-dimensional surfaces. However, the physiological environment in which in vivo wound healing takes place is three-dimensional rather than two-dimensional. It is becoming increasingly clear that cell behavior differs greatly in two-dimensional vs. three-dimensional environments; therefore, there is a need for more physiologically relevant in vitro models for studying cell migration behaviors in wound closure. The method described herein allows for the study of cell migration in a three-dimensional model that better reflects physiological conditions than previously established two-dimensional scratch assays. The purpose of this model is to evaluate cell outgrowth via the examination of cell migration away from a spheroid body embedded within a fibrin matrix in the presence of pro- or anti-migratory factors. Using this method, cell outgrowth from the spheroid body in a three-dimensional matrix can be observed and is easily quantifiable over time via brightfield microscopy and analysis of spheroid body area. The effect of pro-migratory and/or inhibitory factors on cell migration can also be evaluated in this system. This method provides researchers with a simple method of analyzing cell migration in three-dimensional wound associated matrices in vitro, thus increasing the relevance of in vitro cell studies prior to the use of in vivo animal models.

Characterizing Cell Migration Within Three-dimensional In Vitro Wound Environments

The goal of this protocol is to evaluate the effect of pro- and anti-migratory factors on cell migration within a three-dimensional fibrin matrix.

Currently, most in vitro models of wound healing, such as well-established scratch assays, involve studying cell migration and wound closure on ...

Culturing Mammalian Cells in Three-dimensional Peptide Scaffolds

A useful technique for culturing cells in a self-assembling nanofiber three-dimensional (3D) scaffold is described. This culture system recreates an environment that closely mimics the structural features of non-polarized tissue. Furthermore, the particular intrinsic nanofiber structure of the scaffold makes it transparent to visual light, which allows for easy visualization of the sample under microscopy. This advantage was largely used to study cell migration, organization, proliferation, and differentiation and thus any development of their particular cellular function by staining with specific dyes or probes. Furthermore, in this work, we describe the good performance of this system to easily study the redifferentiation of expanded human articular chondrocytes into cartilaginous tissue. Cells were encapsulated into self-assembling peptide scaffolds and cultured under specific conditions to promote chondrogenesis. Three-dimensional cultures showed good viability during the 4 weeks of the experiment. As expected, samples cultured with chondrogenic inducers (compared to non-induced controls) stained strongly positive for toluidine blue (which stains glycosaminoglycans (GAGs) that are highly present in cartilage extracellular matrix) and expressed specific molecular markers, including collagen type I, II and X, according to Western Blot analysis. This protocol is easy to perform and can be used at research laboratories, industries and for educational purposes in laboratory courses.

Here, we present a protocol to obtain 3D-culture systems in self-assembling peptide scaffolds to promote the differentiation of dedifferentiated human articular chondrocytes into cartilage-like tissue.

A useful technique for culturing cells in a self-assembling nanofiber three-dimensional (3D) scaffold is described. This culture system recreates an ...

Production of Extracellular Matrix Fibers via Sacrificial Hollow Fiber Membrane Cell Culture

Engineered scaffolds derived from extracellular matrix (ECM) have driven significant interest in medicine for their potential in expediting wound closure and healing. Extraction of extracellular matrix from fibrogenic cell cultures in vitro has potential for generation of ECM from human- and potentially patient-specific cell lines, minimizing the presence of xenogeneic epitopes which has hindered the clinical success of some existing ECM products. A significant challenge in in vitro production of ECM suitable for implantation is that ECM production by cell culture is typically of relatively low yield. In this work, protocols are described for the production of ECM by cells cultured within sacrificial hollow fiber membrane scaffolds. Hollow fiber membranes are cultured with fibroblast cell lines in a conventional cell medium and dissolved after cell culture to yield continuous threads of ECM. The resulting ECM fibers produced by this method can be decellularized and lyophilized, rendering it suitable for storage and implantation.

The goal of this protocol is production of whole extracellular matrix fibers targeted for wound repair which are suitable for preclinical evaluation as part of a regenerative scaffold implant. These fibers are produced by the culture of fibroblasts on hollow fiber membranes and extracted by dissolution of the membranes.

Engineered scaffolds derived from extracellular matrix (ECM) have driven significant interest in medicine for their potential in expediting wound closure ...

Preparation of Tunable Extracellular Matrix Microenvironments to Evaluate Schwann Cell Phenotype Specification

Traumatic peripheral nervous system (PNS) injuries currently lack suitable treatments to regain full functional recovery. Schwann cells (SCs), as the major glial cells of the PNS, play a vital role in promoting PNS regeneration by dedifferentiating into a regenerative cell phenotype following injury. However, the dedifferentiated state of SCs is challenging to maintain through the time-period needed for regeneration and is impacted by changes in the surrounding extracellular matrix (ECM). Therefore, determining the complex interplay between SCs and differing ECM to provide cues of regenerative potential of SCs is essential. To address this, a strategy was created where different ECM proteins were adsorbed onto a tunable polydimethylsiloxane (PDMS) substrate which provided a platform where stiffness and protein composition can be modulated. SCs were seeded onto the tunable substrates and critical cellular functions representing the dynamics of SC phenotype were measured. To illustrate the interplay between SC protein expression and cellular morphology, differing seeding densities of SCs in addition to individual microcontact printed cellular patterns were utilized and characterized by immunofluorescence staining and western blot. Results showed that cells with a smaller spreading area and higher extent of cellular elongation promoted higher levels of SC regenerative phenotypic markers. This methodology not only begins to unravel the significant relationship between the ECM and cellular function of SCs, but also provides guidelines for the future optimization of biomaterials in peripheral nerve repair.

This methodology aims to illustrate the mechanisms by which extracellular matrix cues such as substrate stiffness, protein composition and cell morphology regulate Schwann cell (SC) phenotype.

Traumatic peripheral nervous system (PNS) injuries currently lack suitable treatments to regain full functional recovery. Schwann cells (SCs), as the ...

Generation of a Simplified Three-Dimensional Skin-on-a-chip Model in a Micromachined Microfluidic Platform

This work presents a new, cost-effective, and reliable microfluidic platform with the potential to generate complex multilayered tissues. As a proof of concept, a simplified and undifferentiated human skin containing a dermal (stromal) and an epidermal (epithelial) compartment has been modelled. To accomplish this, a versatile and robust, vinyl-based device divided into two chambers has been developed, overcoming some of the drawbacks present in microfluidic devices based on polydimethylsiloxane (PDMS) for biomedical applications, such as the use of expensive and specialized equipment or the absorption of small, hydrophobic molecules and proteins. Moreover, a new method based on parallel flow was developed, enabling the in situ deposition of both the dermal and epidermal compartments. The skin construct consists of a fibrin matrix containing human primary fibroblasts and a monolayer of immortalized keratinocytes seeded on top, which is subsequently maintained under dynamic culture conditions. This new microfluidic platform opens the possibility to model human skin diseases and extrapolate the method to generate other complex tissues.

Here, we present a protocol to generate a three-dimensional simplified and undifferentiated skin model using a micromachined microfluidic platform. A parallel flow approach allows the in situ deposition of a dermal compartment for the seeding of epithelial cells on top, all controlled by syringe pumps.

This work presents a new, cost-effective, and reliable microfluidic platform with the potential to generate complex multilayered tissues. As a proof of ...