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

Summary

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.

Abstract

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 migration^2,3. 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.

Introduction

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 experiments^4-6. 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 substrata^8,9. 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 remodeled^2,11. 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 boundary^2,12. 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 platforms^3,13. 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 alignment^13,14. 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 channels^3,15,16. 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.

Protocol

1. Neutralization, Dilution and Polymerization of Collagen Solutions for 3D Investigation and Cellular Contraction Assays

1. 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.
2. Dilute neutralized collagen to the appropriate concentration with cell media (such as RPMI, DMEM, etc.).
   1. 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.
   2. 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.
      1. 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.
3. 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.
4. Allow to polymerize at room temperature for approximately 10 - 15 min. The gels should turn opaque upon polymerization.
5. After it has turned opaque, move plate or dish to 37 ˚C for additional 45 - 60 min to finish polymerization.
6. 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.

2. Western Blotting, Cell Morphogenesis and Gel Gontractility Assay

1. 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.
2. 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.
3. 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.
   1. 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.
4. 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.

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.

1. 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.
2. 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.
3. 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.
4. 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.
5. 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.
6. Gently place a rubber sheet on top of the transparency, followed by a 1/8" acrylic sheet.
7. Add three 10 lb weights on top of the acrylic sheet. Initially, the weights will "float". Allow them to settle and stabilize before proceeding further.
8. Set hotplate temperature to 70 ˚C and cure PDMS for 4 hr. Allow to cool for minimum of 1 additional hr before disturbing.
9. Carefully peel top sheet of transparency film from the wafer, and remove the channels with a forceps.
10. Store in dust-free dish until ready to use.

4. Prepping PDMS Microchannels for Use

1. 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.
2. 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.
3. 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.
4. 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.
5. 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.
6. Pre-coat the inside of the channel with 10 µg/ml collagen in sterile water. To coat, place a 100 µl droplet on the top of a channel, and draw through with vacuum. After 1 hr at 37 ˚C, transfer channels filled with collagen coating solution to a refrigerator. Chill for approximately 15 - 30 min.
7. Remove collagen coating solution with aspirator or pipet, and begin collagen preparation.

5. Collagen Preparation for Use in Microchannels

1. 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).
2. Dilute neutralized collagen to the appropriate concentrations with cell media (For more details, see section 1.2). Incubate for 15 min on ice.
3. At the same time, chill mounted channels on ice.
   NOTE: The goal is to have all the components for the channel process at 4 ˚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.
4. 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).
5. Once cells have been counted and 15 min have passed, proceed to collagen pouring.

6. Pouring Aligned and Random Collagen Microchannels

1. 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.
   1. For random or unaligned matrices, use a wide channel (3mm wide x 200 µm tall) with a vacuum pressure of 10 mm Hg or less.
   2. For aligned matrices, use narrow channel (1mm wide x 200 µm tall) with a vacuum pressure of 60 mm Hg or greater.
2. Remove mounted channel from ice and place on clean, sanitized surface of laminar flow hood.
3. Work quickly and load 120 - 150 µl of neutralized collagen into port A (Figure 2b).
4. 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.
5. Carefully remove excess collagen from the port region with p200 pipetman or aspirator.
6. Place sterile PDMS patches over both ports A and C. All ports should now be covered.
7. After 2 - 3 mins, remove center PDMS patch (port B) and add 2 - 3 μl of cells (5 - 10 thousand cells) into the center port. Allow to partially polymerize (turn opaque) at room temperature for another 10 - 15 min.
8. After 10 - 15 min, transfer to 37 ˚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.

Results

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...

Discussion

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...

Materials

Name	Company	Catalog Number	Comments
High Concentration Rat Tail Collagen	Corning	354249
SylGard184 elastomer kit	Corning	NC9285739	Elastomer for PDMS channels
HEPES	Fisher	BP310	For HEPES neutralization buffer
KCl	Fisher	BP366	For HEPES neutralization buffer
KH2PO4	Fisher	BP362	For HEPES neutralization buffer
Na2HPO4	Fisher	S374	For HEPES neutralization buffer
NaCl	Fisher	BP358	For HEPES neutralization buffer
Levy Improved Neubauer Hemacytometer	Fisher	15170-208	cell counting
6-well non-tissue culture plate	Corning	351146
50 mm glass bottom dish	MatTek	P50g-1.5-30-f
Bel-Art Plastic Vacuum Desiccator	Bel-Art	F4200-2021	Degassing chamber for PDMS
transparency film	3M	pp2950	Plastic film for pouring pdms channels
ThermoScientific CimaRec	ThermoScientific	HP141925	Hot plate for curing PDMS microchannels
Vacuum regulator	Precision Medical	PM3100	Vacuum regulator for collagen microchannels
8" x 8" rubber sheet	Amazon - Rubber-Cal	Silicone - 60A	rubber sheet for pouring PDMS microchannel
8" x 8" x 0.125" acrylic sheet	Amazon	Plexiglass sheets	for pouring PDMS microchannels
10 lb weights	Amazon	CAP Barbell	for pouring PDMS microchannels
15 ml Conical tubes	Fisher	352097
50 ml Conical tubes	Fisher	352098
Plastic pipets	Dot Scientific	229202B, 229206B, and 667225B	2 ml, 5 ml, and 25 ml
70% EtOH	Fisher	NC9663244