Cellular migration is a critical wound healing process that can drastically alter the outcome of a healing wound. This protocol allows researchers to better understand the effect of various compounds on cellular migration during wound healing. Which can often times lead to targeted treatment strategies for clinical medicine.
The scratch assay is a cost-effective and high-throughput approach for generating meaningful wound healing data prior to costly and time-intensive in vivo testing. The scratch assay as a method is particularly relevant in wound healing applications. Most specifically, the skin.
However, the assay is really a migration assay, and could be applied to any cell type where migration of cells is of interest to measure or to capture. Similar to classic cell and tissue culture, standard operating procedures, training, technique, and practice, practice, practice. Those are key to success with the scratch assay.
Demonstrating the procedure will be Nathan Cruz, a master's student, and Oscar Lujan, an undergraduate researcher. To begin this procedure, prepare the biosafety II cabinet using a 70%isopropyl aqueous solution to perform a sanitation wipe, beginning at the back and side walls, and working down to the baseplate. Also wipe down all materials that will be used in the assay before placing them in the cabinet.
Next, obtain vials containing the desired cryopreserved cell line, and place one in a one-centimeter-deep water bath at 37 degrees Celsius to thaw. Add 10 milliliters of medium supplemented with 10%FBS into a T75 tissue culture flask. Open the vial containing the cell stock, and carefully aspirate the solution.
Then transfer the cells into the T75 flask. Remove one milliliter of media from the flask, and transfer it back into the vial, and then transfer it back into the flask to maximize the cell yield from the vial. Tighten the flask's cap and gently rock it to uniformly disperse the cells in suspension across the entire surface.
After this, label the flask with the information shown here, and place it in a humidified incubator at 37 degrees Celsius and 5%carbon dioxide for 72 hours. First, retrieve the tissue culture flask from the incubator, and observe the adhered cells as outlined in the text protocol. Aspirate the entire volume of media from the flask and transfer it into a waste container.
Add five milliliters of balanced salt solution to the flask, and rock it gently to rinse out any excess media, dead cells, or waste product. Then remove the entire volume of salt solution from the flask and discard it into a waste container. Next, add two milliliters of trypsin to the flask, and gently rock it to disperse the enzyme over the adhered cells.
Ensure that the adherent surface is completely saturated, and place the tissue flask in the incubator for two to three minutes. After this, gently tap the side of the flask three to five times to help facilitate cell detachment. Wipe down the flask with 70%alcohol, and place it in the biosafety cabinet.
Add five milliliters of stock media to the tissue flask to stop the enzyme substrate reaction. Aspirate the entire volume of fluid from the flask, and transfer it into a sterile 15-milliliter conical tube. Centrifuge at 200 times g and at 25 degrees Celsius for five minutes.
Then use 70%alcohol to wipe down the tube containing the pelleted cells, and place it inside the biosafety cabinet. Pipette off the media, leaving behind approximately 250 microliters of fluid with the cell pellet. Run the bottom of the conical tube across the vial slots of a microcentrifuge tube rack to create a swift vibration and disturb and resuspend the cell pellet.
When completed correctly, the new cell solution will appear as a cloudy mixture with no remaining pellet. Add two milliliters of media to the cell resuspension. Pipette the cells gently up and down the side of the tube 20 times to break up any clumps.
And record the total cell suspension volume. Next, use a sterile pipette to add 20 microliters of trypan blue stock to an empty well of a 96-well plate. Using a sterile pipette tip, transfer 20 microliters of cell stock from the 15-milliliter tube to the well containing the trypan blue solution.
Pipette gently to mix the contents and transfer 10 microliters of the mixture between the cover slip and the counting chamber on a hemocytometer. Count only the cells in the center and four corner squares, separately tallying both the total cell count and the non-viable cell count for the five identified squares. After calculating the viable cell count, mark a horizontal line across the bottom of a 12-well plate to serve as a reference mark during imaging.
Dilute the cell stock with media to a density of 19, 000 cells per milliliter. Then seed each well of the 12-well plate with one milliliter of the diluted cell stock, mixing the cell stock periodically to ensure even cell seeding across all 12 wells. Label each plate with the information shown here.
And place it in an incubator at 37 degrees Celsius and 5%carbon dioxide until the cells grow to 100%confluence. Retrieve a 12-well plate with cells from the incubator. Using a microscope with a 10x objective, observe the cells and determine the confluence within each well.
Next, aspirate and dispose of the growth media from each well. Assemble a P200 pipetter with a 1-200 microliter sterile tip. In each well, glide the pipette tip across the cell surface from the 12:00 location to the 6:00 location to produce a scratch mock wound.
Rinse each well with one milliliter of balanced salt solution to clear away excess debris and cell clumps. Rock the plate from side to side to facilitate washing. Then remove and dispose of the balanced salt solution from each well.
Using a P1000 pipetter with a 1, 000-microliter tip, add 1, 000 microliters from the prepared arsenic stocks into wells that have been randomly assigned with a treatment group. Use a new pipette tip for each well, and record the time when the treatments are added. Incubate the plates at 37 degrees Celsius with 5%carbon dioxide.
To image the plates, capture images of each well at 10x objective magnification every four hours over a 24-hour period. Reference the line previously drawn on the bottom of the plate to image the same location along the scratch within each well at each time point. In this study, an in vivo scratch test assay is used to observe the effects of arsenic on cell migration.
Uniform scratches are observed across all treatment groups, with a mean scratch width of approximately 0.8 millimeters. The control cells show a completely healed scratch after 20 hours. The two arsenic groups do not show this level of healing.
With the 10-micromolar treatment inhibiting closure altogether for 24 hours. These results demonstrate that the presence of arsenic slows cellular migration. The raw images captured during the assay are analyzed using automated software to measure the wound width, then calculate the percent closure, and finally calculate the area under the curve.
The 10-micromolar treatment group shows statistically-slowed cellular migration of human neonatal dermal fibroblasts compared to all other treatment groups. It is important to thoroughly practice this technique to increase consistency when scratching the cells. Unintentional variability between assays can potentially skew data and alter conclusions.
Further molecular techniques such as quantitative polymerase chain reaction and enzyme-linked immunosorbent assays can be employed post-scratch assay to identify messenger signals and proteins that may be the target for changes to cell migration observed in the assay. This assay has contributed to researchers'understanding of cancer metastasis, new emerging cancer therapies, stem cells and regenerative medicine, and many other scientific milestones.
