Injection of Syngeneic Murine Melanoma Cells to Determine Their Metastatic Potential in the Lungs

Ethics Statement: The use of mice in research is only acceptable in instances where it might advance human understanding of fundamental biology or towards the eventual treatment of disease.

1. Ordering

1. Purchase female C57BL/6 mice (age of 6 – 8 weeks), 5 mice per group. Allow several days for the mice to adjust.

2. Preparation of B16-BL6 Cells

1. Remove cell culture plates from 37-degree incubator and place in a sterile hood. Plates should be approximately 70% confluent with B16-BL6 murine melanoma cells. A greater confluence may alter the cells metastatic potential.
2. Add 1 ml of 0.05% Trypsin-EDTA per plate, let sit for 1 min, and then aspirate the trypsin-media solution.
3. Add 1 ml of trypsin per plate and return plates to the incubator for 10 min.
4. After moving the plates from the incubator to the sterile hood, add 4 ml of serum-free Rosewell Park Memorial Institute Medium (RPMI) to each plate.
5. Collect the cells with a sterile, motorized pipette. Eject cells against the bottom of the plate, with the tip pressed at a very slight angle, to break up the cells. Repeat several times in order to ensure adequate separation of cell clumps, which might otherwise clog the ejection needle. Recollect and transfer to a 50 ml conical tube.
6. Use a hemocytometer to determine the cell density. Keeping the total number of B16-BL6 cells constant, determine the volume of serum-free RPMI needed to reach final concentrations of 0, 0.065, 0.015, 0.25, and 0.5 million cells/ml.
7. Equally aliquot the media in the 50 ml tube to 15 ml conical tubes. Centrifuge the conical tubes at 2,250 x g for 5 min. Decant the media. Add an appropriate volume of serum-free RPMI and confirm the desired cell density via hemacytometer. Adjust densities- by adding additional RPMI or spinning and resuspending in a smaller volume- accordingly.
8. Aliquot 500 µl of each cell suspension into labeled 1.8 ml tubes on ice.

3. Tail Vein Injection

1. Take a mouse, held by the tail, and guide it rear-first into the mouse restrainer. With the torso in the main chamber of the restrainer, with the tail outside it, pull the mouse towards the end of the restrainer. Insert the restrainer-plunger, continuing to push until the mouse is secure.
2. Rotate the mouse 90 degrees such that one of the lateral tail veins is facing upwards. Using an alcohol pad, vigorously clean the side of the mouse's tail, particularly where the tail vein is most visible.
   Note: Good lighting is essential for effective tail vein visualization on B57BL/6J mice. Heat lamps or heated surgery pads, while not necessary, will also help by increasing the volume of the tail vein.
3. In a sterile culture hood, collect 300 µl of cell culture media from the 2 million cells/ml tube. Invert the needle, flicking the side and pushing the plunger, to eject bubbles from the syringe. Eject culture-RPMI to result in a final volume of 250 µl.
4. Extend the mouse-tail with non-dominant hand. Hold the tail with the proximal end of the tail elevated with the forefinger of the non-dominant hand, and the distal portion of the tail depressed with the thumb.
   Note: In this way, the tail vein is held in a lateral position, parallel to the restrainer surface, with a congruent entry possible in the more distal part of the tail.
5. Insert the needle into the tail vein, towards the distal end, at a minute downward angle to begin with, and then adjust the needle to match the angle of the tail vein upon further insertion (lowering plunger end of the syringe relative to the needle).
6. Insert the needle approximately 1 cm into the tail vein. After insertion, begin to push the plunger, holding the needle steady.
   Note: If the needle is in the tail vein and the end of the needle is not obstructed the media should flow unimpeded into the vein. Effective injection is characterized by blood clearance from the vein. If there is resistance, or a bubble begins to form at the site in injection, remove the needle and try again at a more proximal location along the tail vein. 
7. Remove the needle from the vein and discard it. Hold gauze against the entry site to stop any bleeding.
8. Remove the plunger from the restrainer and place the mouse in the recipient cage.
9. Repeat for all other concentrations.
   Note: Lung foci establishment takes approximately 2 – 3 weeks, with variability depending on cell line and the desired size of the foci.⁹ In the interim, mice should be monitored for symptoms that could warrant early euthanasia, like excessive panting. 

4. Counting the Lung Foci

1. Euthanize the mouse by placing in a CO₂ chamber and exposing to 100% CO₂ introduced at 10-30% of chamber volume per minute for 5 min. Follow with a cervical dislocation. 
2. Splay the mouse on Styrofoam. Using surgical scissors, create a surgical incision along the sternum.
3. Make two additional incisions, both from the top of the sternum incision and branching towards the shoulders of the mouse, exposing the rib cage.
4. Make two cuts, each along the lateral sides of sternum, to remove the rib cage from the torso, exposing the lungs and heart.
5. Holding the heart with surgical tweezers, cut the esophagus and windpipe, freeing the heart and lungs from the mouse. Place beneath a dissecting microscope, at 10X magnification, for better visualization.
6. Dissect the heart from the lungs and remove the thymus, leaving only the two lungs.
7. With the lungs beneath a dissecting scope, begin to count the lung foci. Each foci should appear as a circular, brown obtrusion on the surface of the lungs.
   1. For consistency between mice, count the number of foci on the surface of a lobe, and then invert that lobe. Doing each of the four lobes individually allows for easier counting.
8. Save and fix the lungs if performing IHC, otherwise, discard the remains.