Generation of Large Numbers of Myeloid Progenitors and Dendritic Cell Precursors from Murine Bone Marrow Using a Novel Cell Sorting Strategy

This method can enable researchers to acquire sufficient numbers of cell types to answer key questions in the field of developmental immunology. The main advantage of this technique is that it relies on a few select markers to isolate large numbers of cells that are traditionally found in low numbers ex vivo. The implication of this technique extends towards the therapeutic bone marrow transplantation, because it allows for the isolation of large numbers of progenitors.

To begin the protocol, saturate the hind legs and torso of a previously prepared mouse with 75%ethanol, and make shallow cuts through the skin around the hip joint with curved tissue scissors. Then, remove and strip the hind legs. Using forceps, firmly pull the skin from the hip down towards the ankle, revealing the muscle, and use scissors to remove the skin flap.

Cut just above the femur and hip joint, and remove the whole hind leg by cutting through the bone. Working in a sterile biosafety cabinet, transfer the legs to one of the previously-prepared petri dishes. Use scissors to cut just below the ankle, and carefully remove as much of the muscle and elastic connective tissue as possible.

Transfer the cleaned bone to the second prepared petri dish. Next, separate the femur, knee, and tibia. Using forceps, hold the leg at the knee and locate the marrow, a faint red line inside the bone cavity at the top of the femur and toward the end of the tibia.

With scissors, cut the tibia just above where the marrow appears to end. Cut just below the knee joint, cut just above the knee joint. Then, flush bone marrow from the femur and tibia.

Fill a 10 milliliter syringe with complete media from the 50 milliliter conical tube, and cap the syringe with a 23 gauge needle. Holding the bone with forceps above the third prepared petri dish, insert the needle into the bone canal, and push the media through, flushing out the cells. Repeat this step until no more color can be seen through the bone.

Continue the procedure by crushing the epiphysis. While still in the second petri dish, hold the kneecap firmly with forceps, and mash the knees with the tip of the syringe. Continue until the epiphysis are no longer red.

Using the syringe, transfer the cells from the second and third petri dishes to the 50 milliliter tube. Break up clumps by gently pipetting up and down, and try to avoid forming bubbles. Centrifuge the cells.

Then remove the supernatant with the serological pipette, dislodge the pellet by flicking, and lyse red blood cells by incubating them in one milliliter of ammonium chloride potassium or ACK lysis buffer, for one minute at room temperature. Using a serological pipette, add 40 milliliters of HBSS buffer. Using a serological pipette, filter the cells through a 70 micrometer cell strainer into a new 50 milliliter conical tube, and centrifuge the cells.

Using a serological pipette, remove the supernatant and culture the bone marrow cells in complete media with 10 nanograms per milliliter of Recombinant Mouse GM-CSF at a density of one times 10 to the 6th cells per milliliter. Using a serological pipette, transfer the cells to tissue culture plates, and incubate them at 37 degrees Celsius in 5%carbon dioxide, until ready to proceed with staining. Gently, but thoroughly, pipette the cells up and down to dislodge loosely adherent cells.

Using a serological pipette, transfer cells to a 50 milliliter conical tube, and centrifuge the cells. Gently pour off the supernatant, and wash the pelleted cells by adding 30 milliliters of FACS wash buffer, or SFWB, with the serological pipette. Then centrifuge the cells and repeat the wash.

Next, suspend and stain cells per the antibody manufacturer's instructions. Resuspend five times 10 to the 7th cells in one milliliter of FWB, and add two micrograms each of anti Ly-6C, and anti CD115, labeled with fluorophores. To further distinguish CMP from MODC, add two micrograms of anti CD11C antibodies.

Incubate the samples for 30 minutes on ice. After incubation, use a serological pipette to add 10 milliliters of FWB, and centrifuge the cells. Gently pour off the supernatant, and wash the pelleted cells by adding 30 milliliters of FWB with a serological pipette.

Centrifuge the cells, and repeat the wash. Before suspending the cells, flick the tube thoroughly to dislodge the pellet. Use a serological pipette to resuspend cells at one times 10 to the 7th cells per milliliter of FWB, and filter them through a 35 micrometer cell filter.

Use a serological pipette to transfer the filtered cells into a polypropelyne tube, and place the tube on ice until they are ready to be sorted. Run the unstained control through the cell sorter, and apply a gate to exclude small debris and highly granular particles. Run the single fluorescent control samples through the cell sorter, and adjust the compensation as needed.

Run a sample of the multi-labeled sample, and observe four distinct populations. Apply a gate to isolate each of the four major populations. Prepare collection tubes by adding enough fetal calf serum, or FCS, to achieve at least 20%final concentration when full.

For example, if using five milliliter tubes, add one milliliter of FCS before sorting, and remove the tube when it reaches five milliliters total volume. To prevent membrane turnover and antibody uptake, keep all samples at four degrees Celsius throughout the sort. After the desired number of cells have been collected, use a serological pipette to transfer the cells to a new conical tube, and centrifuge the cells.

Carefully remove the supernatant and resuspend the cells in 10 milliliters of FWB, and centrifuge the cells again. Repeat the FWB suspension for a total of two washes. Finally, remove the supernatant after the second wash, and proceed based on the experiment's design.

To keep as many channels available for analysis as possible, viable cells were routinely selected, based on forward and side scatter, excluding very small and very granular events. To determine if this gating strategy reliably excluded dead cells, samples were stained with 7-Aminoactinomycin D.Cells analyzed immediately post harvest had approximately 10%of 7-AAD positive cells, when a typical FSC, SSC gate was applied to freshly isolated cells from the bone marrow. A similar proportion of dead cells was also present at stay 1 and stay 3 of culture.

By day five, the number of dead cells within the gate was reduced to 5%thus, using such a viability gate is generally appropriate for sorting on day five and after. Flow cytometry revealed that within the Ly6C negative, CD115 population, CD3, CD45R positive cells persisted strongly through day zero through three. On day four, only a few CD3, CD45R positive cells remained, and by day five and six, there were no CD3, CD45R expressing cells present;thus, within four days of culture in GM-CSF, lineage positive cells were essentially absent, and were not detected at all at days five and six of culture.

While attempting this procedure, it's important to remember cellular composition is dependent on the length of culture. Sorting three days post-harvest yields high numbers of early stages and few late stage, and vice-versa for cultures sorted after five days.