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

The overall goal of this procedure is to generate aligned and random 3D collagen matrices, for the purpose of observing cell migration or other cellular processes in the context of a 3D environment. This method is designed to capture collagen features that we find in real tissues, which can help answer the key questions such as the mechanisms that cells use to recognize an aligned or random extracellular matrix. The main advantage of this technique is that you can manipulate the organization of the 3D environment, image cells within and you can do this cost effectively with equipment that is commonly available.

Generally, individuals new to this method will struggle because they don't mix the collagen solution and the neutralization buffer well enough and they draw the collagen through the channels inconsistently. Demonstrating this procedure will be Brian Burkel, who is a senior scientist in my laboratory. To begin, thoroughly mix the collagen and HEPES buffer prior to adding cells and media to ensure proper neutralization of the collagen.

Then, add the cells and media to the neutralized collagen mixture and place it on ice as described in the accompanying text protocol. Pipette the ice cold mixture into a sterile non-tissue culture treated surface to minimize the attachment and growth of cells outside of the collagen gel. Use a pipette to evenly spread out the solution.

Allow the gel to polymerize at room temperature for approximately 10 to 15 minutes. Next, cover the gels, and move them to a 37 degree Celsius incubator for an additional 45 to 60 minutes to finish polymerization. Upon polymerization, the gel will turn opaque.

Once polymerized, add two to three milliliters of media. Then, release the gels from the sides of the well by running a 200 pipette around the perimeter of the well. Swirl the dish gently to release the gel from the bottom surface.

In order to create PDMS Microchannels for aligning collagen matrices. First prepare an SuA silicon master, using standard soft lithography techniques described elsewhere. Next, use a craft stick to thoroughly mix 20 grams of elastomer base with 2 grams of curing agent in a disposable cup.

Once well-mixed, degas the PDMS mixture by placing the disposable cup in a vacuum chamber under a vacuum pressure of 550 millimeters of mercury. Degas the mixture for approximately one to one and a half hours. While degassing the PDMS, prepare the silicon master by placing a clean sheet of transparency film onto a hot plate followed by the silicon master.

Ensure that the micro channel mold faces up. After one to one and a half hours, remove the degassed PDMS from the vacuum chamber. Slowly pour the PDMS into the center of the master and allow gravity to spread it evenly.

Next, carefully roll a second sheet of transparency film on top of the silicon master and PDMS to avoid air bubbles. Gently place a rubber sheet on top of the transparency, followed by a 1/8 inch sheet of acrylic, then, add the first of three ten pound weights on top of the acrylic sheet. Initially, the first weight will float, allow it to settle and stabilize before proceeding with the two remaining weight plates.

When ready to continue, set a hot plate to 70 degrees Celsius and cure the PDMS for four hours. Once cured, allow the PDMS to cool for a minimum of one additional hour and then, carefully peel the top transparency film for the wafer. Finally, remove the channels with the forceps.

Place the channels upside down on a new clean transparency film and clean all of the ports with using a circular motion with sharp forceps. Remove any fragments of PDMS that come loose. Next, place a piece of packing tape to the bench, sticky side up and set the PDMS channel side-down on top of the tape.

Press down on the channels with the round end of a pair of forceps to ensure good contact. Remove the channels and repeat this process on both sides until all of the visible debris has been removed. Then, transfer the clean, prepped PDMS channels into a 50 milliliter conical tube filled with 70%ethanol.

Vortex the channels at maximum speed for 30 seconds, and then replace the solution with fresh, 70%ethanol. Using aseptic techniques, transfer the PDMS channel side up, to a clean and sterile dish. Then, UV treat the channels until the ethanol has evaporated.

Once dry, flip the channels over onto a cover slip or glass bottom dish and press them down onto the glass to make a good seal. Add a patch of sterile PDMS to cover and close the center part of the channels and allow the setup to dry thoroughly before proceeding. Next, place a 100 microliter droplet of a 10 micro grams per milliliter collagen solution on the top of a channel and draw it through the channel using a vacuum.

Incubate the channels at 37 degrees Celsius for one hour and then transfer the channels, still filled with collagen coating solution into a refrigerator. Chill the set up for approximately for 15 to 30 minutes and then use an aspirator or pipette to remove the collagen coating solution from the channels. Begin by neutralizing the collagen solution, diluting it with cell media and incubating it on ice for 15 minutes.

Then, place 120 to 150 microliters of the collagen solution into port A of a pre chilled channel. Next, attach a 25 milliliter pipette to a vacuum line and place it over port C.Use it to draw in the mixture through a single uniform motion, stopping once the collagen reaches the end. The single most important step to generating random matrices is to slowly draw the collagen through the channel.

If it is drawn through too rapidly, it will result in increased alignment in the portion of the channel or throughout the entirety of the channel. Carefully remove any excess solution from the port regions and then place sterile PDMS patches over both ports, A and C.After two to three minutes, remove the center PDMS patch covering port B and add two to three micro litters of cells at a concentration of one to three million cells per milliliter into the center port. Allow the gel time to partially polymerize at room temperature for another 10 to 15 minutes and then transfer the constract to a 37 degree celsius incubator for an additional 15 to 30 minutes.

Once polymerized, remove the PDMS port covers and add media to completely cover the channel. Here 50, 000 four T one cells were enbedded in either a 2 mg/ml or a 3.5 mg/ml collagen gel for 5 days. The four T cell one line is a highly contractile Med Aesthetic cancer line but it's only able to contract the high density gel by approximately 10%Comporable low density gels are contracted 57%during the same time frame.

The degree of collagen alignment is modulated by the rate of collagen flow such that higher collagen flow rate yield better alignment. The use of narrower channels coupled with higher vacuum pressures enhances the alignment of the collagen network, while a wider channel used in conjunction with low vacuum pressures generates random matrices. Once mastered, pouring collagen gels or channels can be done in two and a half hours if performed properly.

While attempting this procedure, it's important to remember to set up the components prior to starting. Once you begin working with the collagen, you need to work quickly. After it's development, this technique paved the way for researchers in the field of 3D cell biology, to explore cell adhesion, migration and invasion into three dimensional extracellular environments.

After watching this video, you should have a good understanding of how to pour collagen gels of varying densities and how to generate, both random and aligned collagen matrices.