The overall goal of this procedure is to provide a simple way to study the effects of chemotactic and chemorepulsive agents on cell migration. This method can help answer key questions in the cell interaction fields such as the effects of different growth factors on wound healing. The main advantage of this technique is that it is easily customizable for different applications and simple to use for researchers of all skill levels.
Demonstrating the procedure will be Aniqa Chowdhury, an undergraduate student, and Tyler Harvey, a graduate student from my laboratory. To begin, create a CAD file with the desired shape of the micro-channel. Adjust the channel's width and length to achieve the required gradient.
Here, 2.254 centimeter squares are connected by a 0.127 centimeter wide channel. Next, obtain a piece of acrylic that is the desired thickness in order to create the proper depth of the microchannel. An acrylic sheet width of about one sixteenth of an inch is ideal as thicker sheets are difficult to cut and very thin sheets can break or warp during the process.
Import the CAD file into a laser cutting apparatus and place the acrylic piece on the cutting surface. Then cut out the chamber, according to the manufacturer's protocol. Place the freshly cut barbell shaped acrylic cutout in a small Petri dish and cover it until it is needed.
In a weigh boat, add ten parts of an elastomer base, with one part of the curing agent and stir using a micro pipette for five minutes. Transfer the mixture into a vacuum chamber, and pull a vacuum for 30 minutes to remove any trapped air bubbles from the PDMS. When finished, turn off the vacuum and remove the weigh boat.
Pour the degassed PDMS on top of a barbell shaped acrylic cutout in a small Petri dish. Make sure that the PDMS completely covers the insert. Allow the PDMS to cure for at least 18 hours at room temperature, then carefully cut the PDMS around the acrylic piece with a scalpel and remove the insert.
In a standard cell culture cabinet, place the microchannel chamber in a Petri dish, feature side up and completely cover it with 70%ethanol. Then shut the hood and turn on the UV light. Expose the chamber to UV light for one to two hours.
Following exposure, open the laminar flow hood and wait 15 minutes for the flow hood to re-establish flow. Next, remove the 70%ethanol and wash the chambers twice with sterile PBS. Use one milliliter PBS for each wash.
Following the final wash, remove the PBS and allow the chambers to dry overnight with the lids off. Harvest the cell type to be tested and add 100 microliters of the cell suspension to 400 microliters of the 0.4%trypan blue and mix gently. Apply 100 microliters of this solution to the hemocytometer by gently filling both chambers under the glass cover slip.
Use a microscope with a 10x objective to focus on the grid on the hemocytometer. Then use a hand tally counter to count the live, unsustained cells within all four sets of 16 squares on the hemocytometer. Divide the total cell number by four and multiply this number by 50, 000 to get the number of cells per milliliter of cell suspension.
Based on this calculation, add 2500 cells and 100 microliters of media to one well in each chamber. Allow the cells to sit in the chamber for 60 minutes then add additional media. To begin, add agarose to water and heat the solution in a microwave for one minute, or until the agarose has completely dissolved and the solution is clear.
Allow the solution to cool for 15 to 30 seconds before pouring into a Petri dish. To remove any bubbles, place the Petri dish into a vacuum chamber and allow the agarose to solidify while under a vacuum for 30 minutes. Once solidified, cut a two millimeter by two millimeter by two millimeter cube of agarose with a scalpel from the Petri dish.
Completely submerge the cube in a solution containing the desired concentration of the chemo tactic factor, and soak the agarose block overnight at room temperature. Next place the agarose block containing the chemo tactic factor at the far end of the microchannel chamber. Utilize a marker, such as polystyrene microbeads or a microscopic defect in the chamber.
To identify the relative starting location of the cells. Use the imaging software to take time lapsed images of the moving cell front every hour for 12 hours. The series of images shown here documents the movement of a cell front across the channel in response to a gradient of fetal bovine serum placed at the opposite end of the channel.
The migration distance was measured based on the distance of the growth front from a microscopic defect in the device. From these measurements, the average cell front velocity was found to be 13.4 micrometers per hour. Once mastered, this technique can be completed in four hours, with an additional 18 hours of wait time for PDMS to cure, and an additional 12 hours for the time lapse imaging.
After it's development, this technique paved the way for researchers of different skill levels, including undergraduate students with little lab experience to easily explore wound healing and cell migration in vitro. While attempting this procedure, it's important to remember to carefully remove the acrylic insert from the PDMS mold in order to ensure successful production of a microchannel chamber. After watching this video, you should have a good understanding of how to produce a simple microchannel chamber for studying cell migration in response to different chemical cues.
Following this procedure, other cues can be studied such as multiple soluble chemical gradients, substrates, fluid sheer and electrical fields in order to study the effect they have on cell migration. Don't forget that working with laser cutters can be extremely hazardous and precautions such as obtaining proper training, and using personal protective equipment should always be taken while performing this procedure.
