Isolation and Culture of Bone Marrow Stromal Cells from a Mouse Tibia

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.

1. Harvest of Bones from Mice

1. Euthanize the animals using CO₂ followed by cervical dislocation and spray them with 70% ethanol.
   NOTE: Ensure that animals of the same sex and preferably the same age are used. We routinely use animals aged 7 to 8 weeks. We recommend pooling cells from at least 3 animals per experimental group to yield a better representative and reproducible cell population.
2. Place the euthanized mouse on its back on a dissecting board. Use forceps to create a tent of skin on the abdomen. Make a small incision (around 1 cm) using sterile dissecting scissors. Peel back the skin towards the limbs and the feet from the cut in order to expose the lower abdomen and the legs.
   1. Cut below the ankle joint to remove the foot. Cut along the iliac crest at the hip to separate the femur head from the hipbone. While the mouse is still on its back, make an incision in the middle of the hipbone to separate the two iliac bones.
3. Using lint-free wipes, carefully remove the muscle from the femurs, tibias, and iliac bones.
   NOTE: Cutting as much of the tendons and muscle from the bone as possible makes the cleaning faster and easier with the lint-free wipes (e.g., Kim wipes). Use care when manipulating the bones as they tend to break easily and their fragility may be increased with certain genotypes.
4. Once all the samples have been dissected, place them in PBS on ice and move to a sterile culture hood for the remaining steps of isolation.
   NOTE: Working aseptically as much as possible will reduce the potential for contamination in the cultures.
5. Make small cuts (approximately 1 to 2 mm) at both the proximal and distal ends of the bones before placing them in the sectioned tips within the centrifuge tubes.
   NOTE: Ensure to cut a sufficient amount so that marrow can be centrifuged out; however, care must be taken to avoid cutting too much as cell populations tend to vary based on proximity within the marrow space.
6. Centrifuge at 10,000 x g for 15 s at room temperature. Ensure that all the marrow is flushed and pelleted at the bottom of the 1.5 mL tube while the bones are inside the inserts (either 200 µL pipet tip or 0.75 mL microcentrifuge tube). Once all the marrow is collected, remove the inserts with the bones from the collection tubes.
   NOTE: If all the marrow is flushed, the bones will appear white. However, if marrow remains in certain bones, try cutting ends again and centrifuge.

2. Cell Suspension

1. Using a 25 G needle, pull the cell pellets up and down slowly to break up clumps and combine all the samples into a single 15 mL conical tube. Add 10 mL of BMSC Culture Media per 500 µL of samples.
2. Place a 70 µm filter on top of a 50 mL conical tube and pass the cell suspension through this filter in order to remove possible bone fragments.

3. Plating and Culture of Mixed Populations of Bone Marrow Cells (Total BMCs)

NOTE: Cells are cultured at 37 °C in an incubator with 5% CO₂.

1. Calculate the number of cells and plate them at 1.0 x 10⁶ cells/cm² in culture media. Allow 72 h of mixed culture for cells to attach.
   NOTE: We recommend diluting the cell suspension 20x in culture media and diluting again in trypan blue before counting the cells using a hemocytometer. For 7-week-old male C57Bl6/J mice, we routinely yield a cell number of around 50 x 10⁶ cells but age, sex, and strain can affect cell number greatly.
   NOTE: A mixed population of cells should attach at the bottom of the culture well.
2. After 72 h, remove the media and replace it with fresh media. Continue changing the media every other day and check for confluency. When the cells reach 80-100% confluency, they are ready for differentiation.
   NOTE: Because this culture is mixed with both BMSCs and HSCs, it can be directly used for osteoblastogenesis, adipogenesis as well as osteoclastogenesis.

4. Plating and Culture of the Split Population of BMSCs (Adherent BMSCs)

NOTE: Cells are cultured at 37 °C in an incubator with 5% CO₂.

1. Plate all the cells collected from the femurs, tibias, and iliac bones of one mouse into a 10 cm culture dish (55 cm² of culture area).
   NOTE: When pooling 2-3 C57BL/6J mice, we typically plate this 'total' bone marrow population in a 150 cm² flask.
2. After 48 h of mixed culture, remove the culture media (containing the non-adherent HSCs). If osteoclastogenesis experiments are to be performed, set this population of non-adherent cells aside (see section 7), if not, aspirate and discard these cells.
3. Wash the plate/flask with PBS carefully so as to not disturb the attached cells.
4. Remove PBS and lift the cells by adding 0.25% trypsin and incubating at 37 °C for 1-3 min. Quench the trypsin by adding cell media to the cell suspension. At this point, BMSCs, which are now lifted, can be counted (as previously done in step 4.1) and plated for future experiments.
5. Calculate the number of cells needed for plating. Centrifuge the required volume of cell suspension at 1,000 x g for 5 min at room temperature.
   Note: Cells are typically plated based on end-point experiments. For example, if protein/RNA is to be isolated we typically plate at a higher density (~1.0-2.0 x 10⁶ cells/well in a 6-well plate).
6. Remove the media and resuspend the cell pellet in the volume of culture media needed for plating. Plate the cells in culture media according to the experiments and allow them to attach for a few days. When the BMSCs reach confluency, proceed to perform differentiation.