Activation of Mouse Bone Marrow-Derived Macrophages in Response to Antibodies

Secure a euthanized mouse supine on a foam board with its limbs outstretched, then, clean the legs with 70% ethanol. Next, make a shallow cut about 2-millimeter deep to remove the skin and fur from the surface of each hind leg. Then, dissect the tissues to access the tibias and femurs.

To remove the tibias and femurs, cut the bone and muscles just below the hip joint and above the ankles. Then, remove as much of the muscle tissue from the bones as possible. Next, clean the bones with 70% ethanol and allow it to evaporate for 1 minute, then, place the bones in a 6-well plate filled with bone flush medium.

In the plate, trim 1 millimeter from either end of the femurs so that their inner cavities are exposed. Make certain a 26-gauge needle can access the cavities. Next, separate the tibia and femur from the knee joint. Then, insert the needle attached to a 10-milliliter syringe into a bone cavity and collect the bone marrow.

Pool the bone marrow from all 4 bones into a 50-milliliter conical tube with 5 milliliters of bone flush medium. Pipette the medium and marrow up and down several times, or vortex at a slow speed to gently disperse clumps in the suspension. Strings of marrow should be broken up into small flecks of marrow less than half a millimeter long.

Next, top off the tube to 50 milliliters with bone flush medium and transfer it into a 75-square-centimeter tissue culture flask. Incubate the bone marrow for an hour. During the incubation, the matured cells will adhere to the flask and the desired hematopoietic progenitors will remain in suspension. Transfer the suspension to a 50-milliliter conical tube, and spin down the cells into a pellet.

Next, discard the supernatant, and resuspend the cell pellet in 5 milliliters of MCSF culture medium. Then, adjust the cell density to 500,000 per milliliter and load 30 milliliters into a new 75-square-centimeter flask. Continue culturing the cells and on days 4, 7, and 10, verify that the cells are adherent, and slightly branched using an inverted phase contrast brightfield microscope.

When checking the cells, discard the medium containing the non-adherent cells. Then, wash the adherent cells with IMDM one time, and add 30 milliliters of fresh MSCF culture medium back into the flask. On day 10, replace the medium with 8 milliliters of enzyme-free EDTA-based cell dissociation buffer to collect the cells. Incubate the cells briefly, then, scrape the cells gently and verify that they've released using a microscope.

Once the cells are released, transfer the buffer with the cells to a 50-milliliter conical tube, then rinse the flask three times with 5 milliliters of IMDM to collect more cells. Pool all the collections, centrifuge them, and resuspend the pellet in 3 milliliters of MCSF. Now, count the cells and adjust their concentration to 1 million per milliliter.

Then, plate 100 microliters of cell suspension per well in a 96-well flat-bottom plate. The entire plate should be fillable. Culture the cells for about an hour until they are adherent. Then, stimulate them with different concentrations of LPS or intravenous immunoglobulin or both, prepared in IMDM. Perform all the manipulations in duplicate or triplicate and be certain to keep untreated controls, then, incubate the cells for 24 hours and analyze the supernatants.