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 goal of this procedure is to engineer a raise of identical three-dimensional tissues that are embedded in a collagen gel for use in many experimental contexts, such as determining the influence of mechanical stress on branching morphogenesis. This method can help answer key questions in tissue morphogenesis, such as determining the underlying mechanisms of collective cell behavior. The main advantage of this technique is that the geometry of the engineered tissues is controlled and reproducible, enabling precise manipulation and measurement of physical and biochemical factors.
Thus, sample size is large enough that conclusions could be made with high statistical confidence. Begin by preparing a silicon master photopatterned with your desired array using standard photolithography techniques. The pattern used in this video consists of rectangular structures that are spaced 200 microns apart.
Next, mix 50 grams of PDMS by combining the prepolymer and a curing agent in a disposable dish at a 10:1 ratio. Then, place the mixture under vacuum for 15 to 30 minutes to remove any air bubbles that were introduced during the mixing process. Place the silicon master textured side up into plastic weigh boat, and pour the degassed PDMS solution on top of the mold.
Then, place the dish in an oven and cure the PDMS at 60 degrees Celsius for 12 hours. Once cured, remove the PDMS and master from the plastic container. Carefully separate the PDMS from the silicon wafer.
Then, use a clean razor blade to remove any excess PDMS from around the imprinted features. Now, use a razor blade to cut the patterned PDMS into individual rectangular stamps. Store these stamps feature side up in a clean 100 millimeter diameter petri dish.
Next, prepare a fresh batch of degassed PDMS solution using the same 10:1 ratio of prepolymer to curing agent. Spin coat two to three grams of this solution onto a 100 millimeter diameter petri dish, and then cure the PDMS for 12 hours at 60 degrees Celsius. Then, use a razor blade to cut the thin layer of PDMS into rectangles to be used as supports for the stamps.
Two supports are needed for each stamp. Once all the supports are cut, sterilize the PDMS stamps and supports by immersing them in 70%ethanol and drying them using an aspirator in a biosafety cabinet. In a biosafety cabinet, add 50 microliters of 1%BSA in PBS to the top of each stamp.
Place the droplet covered stamps at four degrees Celsius for a minimum of four hours to ensure that the BSA absorbs to the surface of the stamp. Then, return the stamps to the biosafety cabinet and aspirate the BSA solution from the PDMS stamps. Next, wash the surfaces of the stamps twice with 50 microliters of cell culture medium.
Aspirate the medium after each wash. Lay out one 35 millimeter 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.
Then, use a pair of tweezers to pick up 15 millimeter diameter glass cover slips and sterilize them using 70%ethanol. Aspirate the excess liquid from the cover slips while holding them with the tweezers and then store them in a separate petri dish. Next, dispense 50 microliters of a four milligram per milliliter collagen mixture directly on to each stamp to evenly coat the surface of the PDMS stamps.
Using the tweezers, pick up each of the collagen coated PDMS stamps and gently invert them. Lower the inverted stamps collagen side down on top of the PDMS supports in the tissue culture dishes. Then, incubate the dishes at 37 degrees Celsius for 30 minutes.
Next, dispense 50 microliters of the remaining collagen mixture onto each of the circular cover slips and incubate them at 37 degrees Celsius for 30 minutes. At this point, harvest your cell type of interest. Here, we have EpH4 mouse mammory epithelial cells we suspended at a concentration of between one and 10 million cells per milliliter.
Now, 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. It is important to lift the stamp straight upwards so as not to distort the features in the collagen gel.
Dispense 30 microliters of the concentrated cell suspension onto the surface of each collagen gel containing molded cavities of the desired geometry. Observe the cells under a bright-field microscope while gently shaking the dishes side to side to promote cell settling within the cavities. The cavities should be filled within five minutes.
To remove excess cells from around the cavities, tilt each tissue culture dish on its side and gently dispense 400 microliters of cell culture medium over the surface of the collagen gel. It is important to wash gently by slowly dispensing culture medium over the gel while tilting the dish so that any excess cells on the gel surface are removed and the cells in the cavities of the gel are not displaced. Aspirate the liquid and repeat the wash twice more.
Check the collagen gels under the microscope in between each wash in order to observe when the excess cells have been rinsed away. After the excess cells have been cleared from around the cell filled cavities, place the tissue culture dishes into an incubator at 37 degrees Celsius for 15 minutes. Then, using the tweezers, gently invert the collagen coated glass cover slips and place them on top of the cell filled collagen molds so that the collagen from the cover slip forms a cap over the cell filled cavities.
Incubate the samples at 37 degrees Celsius for 15 minutes. Once the collagen caps have adhered to the cell filled collagen molds, dispense two to 2 1/2 milliliters of cell culture medium slowly over the glass cover slips on top of the gels and culture the samples at 37 degrees celsius for one to three days. These images are from low and high magnification views of the arrays 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. Here, the rectangular wells have been filled with mammory epithelial cells. In this example, each 200 by 50 micron rectangular well contains approximately 80 to 100 cells.
24 hours after cell seeding, cells were observed adhering to one another within the wells to form a tissue. 24 hours after the addition of a growth factor HGF to the culture medium, the tissues are seen undergoing branching morphogenesis. One of the proteins involved with cell migration is focal adhesion kinase, which as you can see from this composite image, increases at the ends of the wells where branching typically occurs.
Once mastered, this technique can be completed within two hours if performed properly. Following this procedure, other methods like traction force microscopy, real time RT-PCR, and Western Blotting, can be performed in order to determine the forces exerted by cells as they migrate and changes in transcript and protein levels of genes of interest. After watching this video, you should have a good understanding of how to set up this 3D cell culture model in which tissues of defined initial geometry are embedded within a collagen gel.
Thanks for watching and good luck with your experiments.
