The Use of Mouse Mammary Tumor Cells in an In Vitro Invasion Assay as a Measure of Oncogenic Cell Behavior

All experiments and methods were performed as authorized by Villanova University Institutional Animal Care and Use Committee (IACUC).

1. Gene Expression in Cultured Mouse Mammary Tumor Cells

1. First, prepare the cell lines to be tested.
2. Use a breeding colony of BALB/cV mice. These mice carry the BALB/cV strain of mouse mammary tumor virus, transmitted to pups in milk^6,7. Fifty percent of breeding females develop mammary tumors by 10 months of age.
   1. To establish a tumor cell line, sacrifice a tumor-bearing dam using an IACUC-approved protocol. Soak the abdominal skin in 70% EtOH and remove the tumor under sterile conditions in a laminar flow hood. Tumors are subcutaneous, so take care to avoid puncturing the peritoneum.
   2. Place tumor fragments from non-necrotic areas in a petri dish with a small amount of sterile Hanks Balanced Saline Solution (HBSS) and mince very finely with a sterile razor blade.
   3. Transfer the dispersed cell clumps to a T25 cell culture flask containing 5 mL Dulbecco's Modified Eagle Medium or DMEM (4.5 g/L glucose, phenol red, and L-glutamine) supplemented with 50% Fetal Bovine Serum or FBS and 1% antibiotic/antimycotic solution. Grow cells in treated cell culture flasks in an incubator with 5% CO₂ and 100% humidity at 37 °C.
   4. Wash off non-adherent cells after 24-48 h and replace the DMEM with 50% FBS and 1% antibiotic/antimycotic solution. Return the cells to the incubator.
   5. Replace liquid culture media every 3 days.
   6. Split or passage the cells when they are 90% confluent. Remove culture media and wash adherent cells with HBSS (without calcium or magnesium). Incubate cells with 0.25% Trypsin-EDTA solution for 1-5 min at 37 °C until cells are rounded and detached. Dilute cells 1:10 in fresh culture media and add to a new flask. A T-25 cell culture flask requires 5-8 mL culture media, 5 mL of HBSS to wash, and 1 mL of trypsin-EDTA to passage. Return the flask to the incubator.
   7. Reduce the FBS concentration to 40% after the first passage, then 30% after the second passage, then 20% after the third passage, and finally 10% for all subsequent passages. The process of establishing growth in standard DMEM medium with 10% FBS requires four passages and 2 – 8 months.
3. Once the cell line(s) are established, alter the expression of suspected oncogenes by transfection of shRNA targeting mRNA or CRISPR for non-essential genes.
   1. Establish antibiotic-resistant stable cell lines with individual clones that are verified by western blot and/or RT-qPCR for consistent results, however, transient transfections can work as well since the assay only requires 22 h. Control cell lines with non-specific target sequences are also required.

2. In Vitro Invasion Assay

1. Grow adherent mouse mammary tumor cells in a T25 flask until 70-90% confluent for Day 1 of the experiment.
   NOTE: Other cell lines or experimental conditions may require different cell culture media. For example, if hormone-responsive breast cancer cells are being tested, charcoal-filtered serum may be needed to remove compounds that activate estrogen or other hormone receptors.
2. Prepare the Boyden chamber inserts by warming them to room temperature (from -20 °C storage) for about 20 min in a cell culture hood. Avoid freeze-thaw cycles for unused inserts. Use three inserts (replicates) to generate statistically valid data for each cell line for each experiment.
   1. Add pre-warmed 37 °C serum-free DMEM media to the well and then the insert. For inserts designed for 24-well dishes, add 500 µL of serum-free DMEM to the well and then 500 µL of serum-free DMEM to the insert so the porous membrane and gel are hydrated on both sides.
      NOTE: For larger inserts designed for a 6-well dish, use 2 mL of serum-free DMEM on both sides.
3. Place the dish with inserts into a 37 °C cell culture incubator with 5% CO₂ and 100% humidity for at least 2 h to thoroughly hydrate and acclimate.
4. Prepare cells by removing the growth media and rinsing the cells with 5 mL HBSS.
   1. Remove the HBSS and add 1 mL 0.25% trypsin-EDTA solution for 1 – 5 min at 37 °C or until cells appear rounded or show signs of detachment from the flask.
   2. Gently tap the flask to detach all the cells. Resuspend the cells in 5 mL of DMEM + 10% FBS and transfer to a sterile 15 mL centrifuge tube.
   3. Centrifuge the cells at 1,000 x g for 5 min to gently pellet the cells. Remove the media and replace with 5 mL serum-free DMEM to resuspend the cells.
   4. Repeat the centrifugation and suspension in serum-free media twice more for a total of 3 washes.
   5. Thoroughly resuspend the cells in the final 5 mL of serum-free DMEM and ensure there are no clumps of cells.
5. Determine the cell concentration using a hemocytometer. Count only viable cells. Dilute the cell suspension to 5 x 10⁴ cells/mL in 5 mL of serum-free DMEM.
   NOTE: This concentration yields 2,500 cells per insert in a 24-well dish. This number of cells is enough for aggressive cancer cells with high rates of migration and invasion. Less aggressive cell lines may require more cells, up to 10,000 cells per insert for observable invading cells. If reduced cell invasion is expected, consider maximizing invading cells by increasing cell number. If increased cell invasion is expected, begin with fewer cells.
6. Remove the cell culture dish from the incubator and gently aspirate the media from the insert. Lift the insert and aspirate the media from the well. Working quickly, add the chemoattractant to the lower chamber. For a 24-well dish, add 750 µL of DMEM with 5% FBS.
   1. Place the insert in the well and add the cells to the insert. For a 24-well dish, use 500 µL of cell suspension. For a 6-well dish insert, use 2.5 mL of chemoattract and 2 mL of cells. Ensure that no bubbles of air are present on either side of the membrane.
   2. Return the dish to the cell culture incubator for 22 h.
7. After 22 h, fix and stain the cells. Prepare a solution of 1% paraformaldehyde in 1x phosphate buffered saline (PBS) for fixation.
   1. In a clean 24-well cell culture dish, add 1 mL of fixative to individual wells so there is one well for each insert.
   2. Prepare the staining solution (freshly made) of 0.1% crystal violet (w/v) in a solution of PBS with 10% ethanol (v/v). Similarly, add 1 mL of staining solution to a clean well in a 24-well culture dish for each insert.
   3. Remove each insert one at a time with forceps and place a sterile cotton swab inside the insert and swab the upper side of the membrane to remove unmigrated cells.
   4. Repeat with a second cotton swab. The membrane is rather strong, so gentle pressure while swabbing does not compromise the integrity of the membrane.
   5. Remove any remaining media from the inside of the insert and add 750 µL of PBS to wash away detached cells.
   6. Remove the PBS and repeat the wash. Place the insert into a well containing fixative to fix the migrated cells on the underside of the membrane. Repeat for all inserts.
   7. Fix the inserts for 15 min at room temperature.
   8. Following fixation remove the insert. Wash the insert again with 750 µL PBS.
   9. Place the insert into the well with the staining solution to stain all migrated and fixed cells. Stain the cells for 15 min at room temperature.
      NOTE: For 6-well dish inserts, use 2 mL of fixative solution, 2 mL staining solution, and 2 mL of PBS wash.
8. Use a beaker with distilled water to destain the inserts. Remove the inserts and dip into the distilled water until the water running off the insert is clear.
   1. Remove any excess water droplets and place the inserts onto a filter paper sidewise to air dry (usually overnight).
9. Prepare the membrane for imaging by labeling a clean glass microscope slide for each insert and place a small drop of microscope immersion oil in the center of the slide.
   1. Detach the membrane from the insert using a scalpel to cut around the perimeter of the membrane on the inside of the plastic insert.
   2. Remove the membrane using forceps and place on the top of the oil drop on the slide to hold it in place.
      NOTE: The membranes are very thin and very light, so they can easily be lost by gentle air flow.

3. Imaging and Analysis

1. Since the porous membranes are clear and staining cells with crystal violet give them contrast, use a compound light microscope to view the cells. Use a camera and/or software to quantify for many samples but is not necessary. View cells at 5x, 10x, or 20x magnification. For quantification, use multiple, non-overlapping images at 10x magnification. Alternatively, count all migrated cells on the membrane at 10x magnification.
2. Determine the total number of invading cells or cells per area for all samples. For each experiment, perform each condition in the assay with three replicate inserts and repeat multiple times for statistically useful results.
3. When comparing different cell lines with different growth rates and migration rates, do a parallel experiment to compare cells that invade through a protein matrix and compare to a Boyden Chamber assembly without protein matrix (migrated cells only). Calculate percent invasion for each cell line (Number of invaded cells/number of migrated cells x 100%).
   NOTE: This approach can help compare invasion rate to migration rates. If comparing different conditions applied to the same cell line, calculating invasion alone is more relevant calculations. The outer edge of the membrane sometimes has a higher number of cells than the central areas and is, therefore, less accurate. If this occurs, exclude those cells from the quantification and ensure that all cells are removed by a cotton swab in a repeat experiment.