The authors would like to acknowledge grant numbers UO1CA143069, R01CA142833, R01CA114462, RO1CA179556, T32-AG000213-24, and T32-GM008692-18 for funding this work. We also acknowledge Jeremy Bredfelt and Yuming Liu of LOCI for the development of and assistance with the CT-FIRE analysis.

1. Neutralization, Dilution and Polymerization of Collagen Solutions for 3D Investigation and Cellular Contraction Assays
<ol>
	<li>On ice in sterile tissue culture hood, neutralize collagen (1:1) with sterile, ice-cold 100 mM HEPES in 2x PBS, pH 7.4, in a 15 ml conical tube. Mix thoroughly with plastic pipette until solution is homogenous and mixing swirls are no longer visible. Be careful not to introduce air bubbles during the mixing process. Store briefly on ice.</li>
	<li>Dilute neutralized collagen to the appropriate concentration with cell media (such as RPMI, DMEM, etc.).
	<ol>
		<li>To calculate the amount of neutralized collagen required to make up desired collagen concentration, use the equation N = (D*V)/(S/2), where N is amount of neutralized collagen required, D is the desired collagen concentration, V is the final volume of the desired collagen concentration, and S is the starting stock collagen concentration.</li>
		<li>Bring up the amount of neutralized collagen to volume by the addition of a cell and cell media solution. Mix thoroughly and store on ice until ready to cast.
		<ol>
			<li>Example: For a 2 mg/ml gel in a 1 ml gel volume (typical volume for a gel cast in 6 well plate or 50 mm glass bottom dish), first multiply the desired concentration (2 mg/ml) by the desired final volume (1 ml). Take this number (2 mg) and divide it by half of the stock concentration listed on bottle (9.49 mg/ml). In this case, 0.42 ml of the neutralized collagen are diluted with 0.58 ml of media/cell mixture.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Pipet the ice-cold collagen/cell/media mixture into either a non-tissue culture treated 6-well plate or a 50 mm glass bottom dish. Use pipet tip to evenly spread out the solution. 
	NOTE: It is important to use a non-tissue culture treated plate to minimize the attachment or growth of the cells outside of the collagen gel.</li>
	<li>Allow to polymerize at room temperature for approximately 10 - 15 min. The gels should turn opaque upon polymerization.</li>
	<li>After it has turned opaque, move plate or dish to 37 &#730;C for additional 45 - 60 min to finish polymerization.</li>
	<li>After 45 - 60 min, add 2 - 3 ml of media and release gels from sides of the well by running a p200 pipet tip around perimeter of well or dish. Swirl dish gently to release the gel. The collagen gel should be floating in media.</li>
</ol>
2. Western Blotting, Cell Morphogenesis and Gel Gontractility Assay
<ol>
	<li>To assess protein levels, morphology or cellular contractility in relation to ECM stiffness, begin by pouring collagen gels, according to 1.1 - 1.6, seeded with cells for a 7 - 10 day assay. Determine each cell line seeding rate empirically depending on growth rate and confluency. For a 10 day assay, the seeding density can range from 20,000 - 100,000 cells/gel.</li>
	<li>To measure cell contractility, measure the gel diameter by using a ruler or a camera every 24 hr or at the appropriate time interval. 
	Note: Additionally, corresponding images collected from a microscope can be examined for morphological features characteristic of the cell line seeded in the gel, such as acini-like structures, epithelial tubules, cellular protrusions, and lameliapodia.</li>
	<li>To assess protein levels, lyse the gels in RIPA buffer and process for Western blot analysis of proteins of interest as described in Wozniak et al. 17 and Gallagher et al. 18.
	<ol>
		<li>IMPORTANT: In comparisons of contractility between cell lines, normalize contractility to total DNA, which can be extracted from the gels as outlined by Lui, et al.19, or to an unchanging, housekeeping protein (histones, GAPDH, etc.) by Western blot analysis, as described in Wozniak et al.17 and Gallagher et al.18. If counting cells within the gel using a hemocytometer, make sure to normalize cell number to total gel area as the contracting gel will concentrate cells.</li>
	</ol>
	</li>
	<li>Feed gels every 3 - 4 days by removing 1 ml of media and replacing it with 1 ml of fresh media. Make sure to feed gels after the measurement is taken as the addition of fresh media/serum will cause a spike in contractility.</li>
</ol>
3. Generation of PDMS Microchannels for Collagen Fiber Alignment
Note: To generate aligned collagen matrices, a mold for PDMS microchannels (Figure 2A) requires a SU-8 silicon master made via soft-lithography15.
<ol>
	<li>To make PDMS channels, mix PDMS thoroughly in a disposable cup with a craft stick. For a 6 inch silicone master, mix 20 g of elastomer base with 2 g of curing agent.</li>
	<li>De-gas the PDMS mixture by placing the disposable cup in a vacuum chamber under a vacuum pressure of 550 mm Hg. De-gas for 1 - 1.5 hr.</li>
	<li>While de-gassing the PDMS, prepare the silicone master for pouring. Prepare by placing a clean sheet of transparency film onto a hotplate followed by the silicone master. Ensure that the microchannel mold faces up. 
	NOTE: During PDMS casting and curing, the silicone master will be sandwiched between two sheets of transparency film.</li>
	<li>After 1 - 1.5 hr, remove the de-gassed PDMS from the vacuum chamber, and slowly pour over the silicone master. 
	IMPORTANT: Avoid air bubbles. Continue to pour in center of master, and allow gravity to spread it evenly. 
	NOTE: The PDMS drop does not need to extend all the way to the edge of the master.</li>
	<li>After PDMS has been poured onto the master, apply a second sheet of transparency film on top of the silicone master and PDMS. Carefully, roll the second transparency sheet down on top of the PDMS/silicone master to avoid air bubbles. Do not rush. PDMS should be now evenly spread over master.</li>
	<li>Gently place a rubber sheet on top of the transparency, followed by a 1/8&#34; acrylic sheet.</li>
	<li>Add three 10 lb weights on top of the acrylic sheet. Initially, the weights will &#34;float&#34;. Allow them to settle and stabilize before proceeding further.</li>
	<li>Set hotplate temperature to 70 &#730;C and cure PDMS for 4 hr. Allow to cool for minimum of 1 additional hr before disturbing.</li>
	<li>Carefully peel top sheet of transparency film from the wafer, and remove the channels with a forceps.</li>
	<li>Store in dust-free dish until ready to use.</li>
</ol>
4. Prepping PDMS Microchannels for Use
<ol>
	<li>Place channels upside down on new, clean transparency film. Clean all ports (Figure 2b) using circular motion with a sharp forceps. Remove any fragments of PDMS.</li>
	<li>Clean channels by using a piece of packing tape as a tack cloth. Apply tape to surface of bench (sticky side up), and then set channel on top. Press down on channels with round end of forceps to ensure good contact. Remove and repeat on both sides until visible debris is removed.</li>
	<li>Transfer cleaned, prepped PDMS channels into a 50 ml conical with 70% EtOH. Vortex at maximum speed for 30 sec. Discard EtOH and replace with fresh 70% EtOH. Store in 70% EtOH until ready to use.</li>
	<li>In tissue culture hood and using aseptic techniques, transfer the PDMS channels to a clean and sterile cover glass or glass bottom dishes. Put the channel side up and UV treat until EtOH has evaporated.</li>
	<li>Once dry, flip the PDMS so the channels are facing down toward coverglass. Press the PDMS channel down on the glass to make a good seal. Add a patch of sterile PDMS to cover/close the center port (port B, Figure 2b). Allow to thoroughly dry before proceeding.</li>
	<li>Pre-coat the inside of the channel with 10 &#181;g/ml collagen in sterile water. To coat, place a 100 &#181;l droplet on the top of a channel, and draw through with vacuum. After 1 hr at 37 &#730;C, transfer channels filled with collagen coating solution to a refrigerator. Chill for approximately 15 - 30 min.</li>
	<li>Remove collagen coating solution with aspirator or pipet, and begin collagen preparation.</li>
</ol>
5. Collagen Preparation for Use in Microchannels
<ol>
	<li>On ice, neutralize collagen (1:1) with ice-cold 100 mM HEPES in 2x PBS, pH 7.4. Mix thoroughly until homogenous (For more details, see section 1.1).</li>
	<li>Dilute neutralized collagen to the appropriate concentrations with cell media (For more details, see section 1.2). Incubate for 15 min on ice.</li>
	<li>At the same time, chill mounted channels on ice. 
	NOTE: The goal is to have all the components for the channel process at 4 &#730;C or below. Collagen nucleation temperature and time are key parameters to the polymerization process, and may be the starting point for further optimization, if needed.</li>
	<li>Count cells with a hemacytometer and resuspend at the appropriate seeding density at this time. For ease of calculations, resuspend to 1 - 3 million cells/ml. (For more details, also see section 2.1).</li>
	<li>Once cells have been counted and 15 min have passed, proceed to collagen pouring.</li>
</ol>
6. Pouring Aligned and Random Collagen Microchannels
<ol>
	<li>Before drawing collagen through channels, adjust and set vacuum pressure with an inline vacuum regulator. Vacuum pressure provides the force to induce and control the rate of collagen flow, which determines the degree of alignment.
	<ol>
		<li>For random or unaligned matrices, use a wide channel (3mm wide x 200 &#181;m tall) with a vacuum pressure of 10 mm Hg or less.</li>
		<li>For aligned matrices, use narrow channel (1mm wide x 200 &#181;m tall) with a vacuum pressure of 60 mm Hg or greater.</li>
	</ol>
	</li>
	<li>Remove mounted channel from ice and place on clean, sanitized surface of laminar flow hood.</li>
	<li>Work quickly and load 120 - 150 &#181;l of neutralized collagen into port A (Figure 2b).</li>
	<li>Draw collagen through the channel by placing a 25 ml pipet attached to the vacuum line over port c (Figure 2b). Draw collagen through in a single, uniform motion. IMPORTANT: For random or unaligned matrices, draw collagen slowly across channel (approximately 0.5 - 1 mm per second), and stop once it reaches the end. For aligned matrices, draw the collagen across more rapidly, but take care to avoid air bubbles.</li>
	<li>Carefully remove excess collagen from the port region with p200 pipetman or aspirator.</li>
	<li>Place sterile PDMS patches over both ports A and C. All ports should now be covered.</li>
	<li>After 2 - 3 mins, remove center PDMS patch (port B) and add 2 - 3 &#956;l of cells (5 - 10 thousand cells) into the center port. Allow to partially polymerize (turn opaque) at room temperature for another 10 - 15 min.</li>
	<li>After 10 - 15 min, transfer to 37 &#730;C for additional 15 - 30 min to finish polymerization. Remove PDMS covers and add media to completely cover the channel and culture cells as needed. Cells can be fed by removing a ml of old media, and replacing it with a ml of new media.</li>
</ol>

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The authors have nothing to disclose

3D collagen gels are a valuable addition to our toolbox to understand how cells interpret and respond to their local microenvironment. This manuscript has provided a very basic protocol for embedding cells within a 3D collagen matrix, and to reproducibly generate matrices with random or aligned collagen fibers. Both protocols work as adaptable platforms where different collagen isoforms, crosslinkers, or other matrix proteins could potentially be added at the time of polymerization. It is also easy to modify the platform...

Many cellular processes have been extensively studied in 2 dimensions, thereby forming a collective knowledge of basic cell signaling mechanisms. These studies, however, neglect important cellular cues received from the structural and mechanical properties of the local cellular microenvironment and extracellular matrix (ECM). To better understand how cells interact within a physiological context, it is important to study them in 3-dimensional (3D) assays. The ECM for these 3D assays can either be cell-derived or reconstituted from purified proteins. Regardless of the source of the ECM, 3D matrix assays have proven to be invaluable for understanding how cells navigate and interact within the physiological world. For example, cells grown in 3D matrices display distinct modes of locomotion that depend on the mechanical nature of their surrounding ECM which are not observed in 2D experiments4-6. Moreover, cells cultured in 3D also have fewer and less pronounced stress fibers and focal adhesions than their counterparts grown on hard surfaces such as glass or plastic7. 
The importance of contextual 3D assays is not limited to cell migration, however. Some other cell signaling events can only be investigated through the use of 3D assays. During tissue and cell differentiation, the stiffness of the extracellular environment and ECM provides signals that can influence morphogenic events. For example, mammary epithelial tubulogenesis only occurs in low stiffness 3D matrices, but not in stiff matrices nor on 2D substrata8,9. When cultured within stiff 3D matrices, these same epithelial cells take on an aberrant phenotype with increased proliferation and cell membrane protrusions driven through altered FAK and ERK signaling10. Many other signaling pathways and cellular processes are known to be similarly affected by the stiffness of the local cellular environment, and these signaling cascades highlight the importance of investigating signaling events and cellular phenotype in the context of appropriate local mechanical properties of a 3D ECM. 
Collagen I is a particularly relevant protein to use for in vitro studies as it is the most abundant component of the ECM and is responsible for many of the mechanical properties of the cellular microenvironment. While it was originally thought of as merely a structural protein, its role is now known to be much more complex. Collagen fiber composition, architecture, orientation, density, and stiffness all provide a concentrated milieu of signaling information5. During the progression of certain diseases, such as chronic inflammation and tumorigenesis, the collagen network is dramatically remodeled2,11. More specifically in breast cancers, increased collagen deposition and tissue stiffness accompany and likely contribute to tumor progression. In these early tumors, the stiffened collagen network appears strained and more aligned, such that most of the fibers encapsulate the growing tumor2. As the tumor progresses, the collagen continues to reorganize, and regions of the fibrillar network become orientated perpendicular to the tumor boundary2,12. Perpendicular alignment serves as a prognostic biomarker where these patients have a poorer disease free progression and overall survival1. One explanation for this correlation is that the poor outcomes are a consequence of increased dissemination of tumor cells via persistent cell migration in aligned collagen networks3. 
To understand how cells specifically respond to alignment and organization that is observed in tumor progression, it is necessary to generate both random and aligned 3D collagen matrices for experimentation. There are three basic methodologies to induce alignment within fibrillar networks. The first technique utilizes a strain-inducing device where the collagen between two points is contracted or stretched to generate alignment. Fibers parallel to the axis of force are pulled taut while fibers perpendicular to the axis are compressed and buckled. While strain-induced techniques typically offer superb alignment, this approach requires bulky equipment that is not easily adaptable to many platforms3,13. Alternatively, cell-induced strain can be created by placing localized plugs of cells that subsequently contract and align the collagen13. This method has the problem of being variable, as many parameters may be subject to change. The second method utilizes magnetic beads and a magnetic field during polymerization to induce collagen alignment13,14. Good results can be obtained from this method with unsophisticated equipment, but it does require the use of antibodies or some other method to magnetize the polymer. Therefore, it can be somewhat expensive to use, and the stiffness of the collagen gel is potentially modified by the increased connections in the network. Moreover, the magnetic beads used in this process are often autofluorescent, which is problematic for imaging experiments. Lastly, alignment can be generated by PDMS microfluidic channels3,15,16. In this method, collagen alignment is achieved by flowing polymerizing collagen through small microfluidic channels. These microfluidic channels can be made in a multitude of designs, and are easily adaptable to many platforms. Moreover, they are very economical as very small quantities of collagen and other reagents are used due to their diminutive sizes.
Provided here is a simple protocol for embedding cells within a 3-dimensional, fibrillar collagen hydrogel. In addition, a more advanced technique, wherein PDMS microfluidic channels are used to control the organization and alignment of the collagen matrix is also provided. 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 3-dimensional, physiological microenvironment.

<table><tbody><tr><td>High Concentration Rat Tail Collagen</td><td>Corning</td><td>354249</td><td></td></tr><tr><td>SylGard184 elastomer kit</td><td>Corning</td><td>NC9285739</td><td>Elastomer for PDMS channels</td></tr><tr><td>HEPES</td><td>Fisher</td><td>BP310</td><td>For HEPES neutralization buffer</td></tr><tr><td>KCl&amp;nbsp;</td><td>Fisher</td><td>BP366</td><td>For HEPES neutralization buffer</td></tr><tr><td>KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Fisher</td><td>BP362</td><td>For HEPES neutralization buffer</td></tr><tr><td>Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;</td><td>Fisher</td><td>S374</td><td>For HEPES neutralization buffer</td></tr><tr><td>NaCl</td><td>Fisher</td><td>BP358</td><td>For HEPES neutralization buffer</td></tr><tr><td>Levy Improved Neubauer Hemacytometer</td><td>Fisher</td><td>15170-208</td><td>cell counting</td></tr><tr><td>6-well non-tissue culture plate&amp;nbsp;</td><td>Corning</td><td>351146</td><td></td></tr><tr><td>50 mm glass bottom dish</td><td>MatTek</td><td>P50g-1.5-30-f</td><td></td></tr><tr><td>Bel-Art Plastic Vacuum Desiccator</td><td>Bel-Art</td><td>F4200-2021</td><td>Degassing chamber for PDMS</td></tr><tr><td>transparency film&amp;nbsp;</td><td>3M</td><td>pp2950</td><td>Plastic film for pouring pdms channels</td></tr><tr><td>ThermoScientific CimaRec</td><td>ThermoScientific&amp;nbsp;</td><td>HP141925</td><td>Hot plate for curing PDMS microchannels</td></tr><tr><td>Vacuum regulator</td><td>Precision Medical</td><td>PM3100</td><td>Vacuum regulator for collagen microchannels</td></tr><tr><td>8&quot; x 8&quot; rubber sheet</td><td>Amazon - Rubber-Cal</td><td>Silicone - 60A</td><td>rubber sheet for pouring PDMS microchannel</td></tr><tr><td>8&quot; x 8&quot; x 0.125&quot; acrylic sheet</td><td>Amazon</td><td>Plexiglass sheets</td><td>for pouring PDMS microchannels</td></tr><tr><td>10 lb weights</td><td>Amazon</td><td>CAP Barbell</td><td>for pouring PDMS microchannels</td></tr><tr><td>15 ml Conical tubes</td><td>Fisher</td><td>352097</td><td></td></tr><tr><td>50 ml Conical tubes</td><td>Fisher</td><td>352098</td><td></td></tr><tr><td>Plastic pipets</td><td>Dot Scientific</td><td>229202B, 229206B, and 667225B</td><td>2&amp;nbsp;ml, 5&amp;nbsp;ml, and 25&amp;nbsp;ml</td></tr><tr><td>70% EtOH</td><td>Fisher</td><td>NC9663244</td><td></td></tr></tbody></table>

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.

While 3D assays can be done within the same stiffness of collagen gel, varying the gel stiffness can be used to determine how the cells will respond to mechanical changes in their cellular microenvironment. A stiff collagen hydrogel is defined as a gel where the embedded cells are unable to locally contract the surrounding collagen. The intrinsic contractility of different cell types is unique, and thus it is best to begin with a simple contractility curve to establish the collagen concen...

Watch this Scientific Journal Video about Preparation of 3D Collagen Gels and Microchannels for the Study of 3D Interactions In Vivo at JoVE.com

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.

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 ...

preparation-3d-collagen-gels-microchannels-for-study-3d-interactions

Universidad de Cartagena UDEC

Research

JoVE Journal

Bioengineering

16.9K Views.  University of Wisconsin-Madison. 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.

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

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

This protocol describes the fabrication of a type of micro-tissues called modules.  The module approach generates uniform, scalable and vascularized tissues.  The modules can be made of collagen as well as other gelable or crosslinkable materials.  They are approximately 2 mm in length and 0.7 mm in diameter upon fabrication but shrink in size with embedded cells or when the modules are coated with endothelial cells.  The modules individually are small enough that the embedded cells are within the diffusion limit of oxygen and other nutrients but modules can be packed together to form larger tissues that are perfusable.  These tissues are modular in construction because different cell types can be embedded in or coated on the modules before they are packed together to form complex tissues.  There are three main steps to making the modules: (1) neutralizing the collagen and embedding cells in it, (2) gelling the collagen in the tube and cutting the modules and (3) coating the modules with endothelial cells.

Creation of micro-tissues using cylindrical collagen gels, called modules, that contain embedded cells and which surface is coated with endothelial cells.

This protocol describes the fabrication of a type of micro-tissues called modules.  The module approach generates uniform, scalable and vascularized ...

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

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

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

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

Revealing the Cytoskeletal Organization of Invasive Cancer Cells in 3D

Cell migration has traditionally been studied in 2D substrates. However, it has become increasingly evident that there is a need to study cell migration in more appropriate 3D environments, which better resemble the dimensionality of the physiological processes in question. Migratory cells can substantially differ in their morphology and mode of migration depending on whether they are moving on 2D or 3D substrates. Due to technical difficulties and incompatibilities with most standard protocols, structural and functional analysis of cells embedded within 3D matrices still remains uncommon. This article describes methods for preparation and imaging of 3D cancer cell cultures, either as single cells or spheroids. As an appropriate ECM substrate for cancer cell migration, we use nonpepsinized rat tail collagen I polymerized at room-temperature and fluorescently labeled to facilitate visualization using standard confocal microscopes. This work also includes a protocol for 3D immunofluorescent labeling of endogenous cell cytoskeleton. Using these protocols we hope to contribute to a better description of the molecular composition, localization, and functions of cellular structures in 3D.

This article presents a method to fluorescently label collagen that can be further used for both fix and live imaging of 3D cell cultures. We also provide an optimized protocol to visualize endogenous cytoskeletal proteins of cells cultured in 3D environments.

Cell migration has traditionally been studied in 2D substrates. However, it has become increasingly evident that there is a need to study cell migration ...

Medicine

Concentric Gel System to Study the Biophysical Role of Matrix Microenvironment on 3D Cell Migration

The ability of cells to migrate is crucial in a wide variety of cell functions throughout life&#160;from embryonic development and wound healing&#160;to tumor and cancer metastasis. Despite intense research efforts, the basic biochemical and biophysical principles of cell migration are still not fully understood, especially in the physiologically relevant three-dimensional (3D) microenvironments. Here, we describe an in vitro assay designed to allow quantitative examination of 3D cell migration behaviors. The method exploits the cell&#8217;s mechanosensing ability and propensity to migrate into previously unoccupied extracellular matrix (ECM). We use the invasion of highly invasive breast cancer cells, MDA-MB-231, in collagen gels as a model system. The spread of cell population and the migration dynamics of individual cells over weeks of culture can be monitored using live-cell imaging and analyzed to extract spatiotemporally-resolved data. Furthermore, the method is easily adaptable for diverse extracellular matrices, thus offering a simple yet powerful way to investigate the role of biophysical factors in the microenvironment on cell migration.

The mechanical properties and microstructure of the extracellular matrix strongly affect 3D migration of cells. An in vitro method to study the spatiotemporal cell migration behavior in biophysically variable environments, at both population and individual cell levels, is described.

The ability of cells to migrate is crucial in a wide variety of cell functions throughout life&#160;from embryonic development and wound healing&#160;to ...

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 ...

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

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

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

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

Micropatterning and Assembly of 3D Microvessels

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

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

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

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.

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 ...

Generation of 3-D Collagen-based Hydrogels to Analyze Axonal Growth and Behavior During Nervous System Development

This protocol uses natural type I collagen to generate three-dimensional (3-D) hydrogel for monitoring and analyzing the axonal growth. The protocol is centered on culturing small pieces of embryonic or early postnatal rodent brains inside a 3-D hydrogel formed by the rat tail tendon-derived type I collagen with specific porosity. Tissue pieces are cultured inside the hydrogel and confronted to specific brain fragments or genetically-modified cell aggregates to produce and secrete molecules suitable for creating a gradient inside the porous matrix. The steps of this protocol are simple and reproducible but include critical steps to be considered carefully during its development. Moreover, the behavior of growing axons can be monitored and analyzed directly using a phase-contrast microscope or mono/multiphoton fluorescence microscope after fixation by immunocytochemical methods.

Here, we provide a method for analyzing the behavior of growing axons in 3D matrices, mimicking their natural development.

This protocol uses natural type I collagen to generate three-dimensional (3-D) hydrogel for monitoring and analyzing the axonal growth. The protocol is ...

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

Summary

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Chapters in this video

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

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

Revealing the Cytoskeletal Organization of Invasive Cancer Cells in 3D

Concentric Gel System to Study the Biophysical Role of Matrix Microenvironment on 3D Cell Migration

Engineering 3D Cellularized Collagen Gels for Vascular Tissue Regeneration

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

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

Micropatterning and Assembly of 3D Microvessels

Microengineering 3D Collagen Hydrogels with Long-Range Fiber Alignment

Generation of 3-D Collagen-based Hydrogels to Analyze Axonal Growth and Behavior During Nervous System Development