The overall goal of this protocol is to assess bone marrow-derived macrophage phagocytosis of brain-derived myelin debris. This protocol is designed to answer key questions in the field of neurotrauma. The advantage of this method is that allows for efficient and relatively cost-effective analysis of macrophage responses in vitro, specifically the phagocytic capacity of bone marrow-derived macrophages for myelin debris.
This method can easily be adapted to investigate the effects of external modulators. Having saturated the mouse with 70%ethanol, make an opening in the abdominal skin. Pull the skin back to reveal the legs for dissection.
Take care not to puncture the peritoneal cavity during this process. Once exposed, remove the legs by cutting at the ankle and the hip. Place the legs in ice-cold PBS supplemented with antibiotics.
Repeat for the other leg. Wash the tissue in fresh ice-cold PBS supplemented with antibiotics to dislodge any particulates that may have adhered to the tissue during dissection. A gentle scraping motion applied along at the muscle will dislodge a majority of the unwanted tissue.
With a scalpel and tweezers, free the tibia from the muscle and connective tissue. Once freed, remove both ends to expose the marrow cavity. Repeat this process for the femur.
Affix a 25-gauge needle to a 20-millimeter syringe filled with complete bone marrow-derived macrophage media. Then flush the bone marrow cavity. The bone will appear white when sufficiently flushed.
Agitate the collected marrow aspirates with an 18-gauge needle for 30 to 90 seconds in order to obtain a single cell suspension. Next pass the suspension through a sterile 70-micrometer pore size cell strainer. Evenly divide the cell suspensions in 145-centimeter cell culture dishes.
Each mouse will provide enough for a total of three dishes. After seven days of culture, these dishes will typically be between 70 to 80%confluent with mature macrophages. To isolate crude myelin debris from brains, a series of sucrose gradient ultracentrifugation steps are carried out.
To begin, 10 to 12 brains are dissected from mice eight to 10 weeks of age. The brains are then homogenized in 0.32 molar sucrose solution, and then layered onto a 0.383 molar sucrose solution cushion to create a discontinuous gradient. Following centrifugation, the crude myelin debris is found at the interface between the layers.
This is collected and dispensed into a Tris buffer solution and centrifuged again to pellet the myelin debris. The pellet is resuspended again in Tris buffer solution and split between two tubes for additional cenetrifugation. To begin, remove the entire brain and place it in a dish with ice-cold 0.32 molar sucrose solution.
Using sterile surgical scissors, cut the collected brains into pieces approximately five millimeters in size. This step aids in the subsequent homogenization process. Collect the tissue into a 50-milliliter tube.
Using a rotary homogenizer, homogenize the tissue into a smooth slurry. Make sure that all large visible brain solids have been thoroughly homogenized. Into an ultracentrifuge tube containing 20 milliliters of 0.83 molar sucrose solution, generate and homogenate, taking care to maintain the separation.
Carefully balance each tube using the 0.32 molar sucrose solution. Once finished, transfer the tubes to an appropriate centrifuge rotor and spin for 45 minutes at 100, 000 times gravity, four degrees Celsius. Ensure that acceleration and deceleration are set to their minimum values.
Once centrifugation is complete, the myelin debris will be visible between the sucrose density interface. Gently collect the myelin debris. Once collected, add Tris buffer to the myelin debris.
Homogenize again with a rotary homogenizer. Divide the homogenate between six clean ultracentrifuge tubes and balance the tubes as needed with additional Tris buffer. Centrifuge the collected myelin debris again at 100, 000 times gravity, four degrees Celsius for 45 minutes, with acceleration and deceleration set to their maximum values.
After centrifugation, the myelin will have been pelleted. Using the Tris buffer, resuspend the pellet. Combine the resuspensions and ultracentrifuge again.
After centrifugation, a tightly-packed pellet is formed. Decant the supernate. Resuspend the pellet in sterile PBS.
Equally divide the resuspended myelin debris into preweighed microcentrifuge tubes. After centrifugation for 10 minutes at 22, 000 times gravity, remove the PBS supernate. After weighing the pellet, resuspend to a concentration of 100 milligrams per milliliter with PBS.
The resulting crude myelin debris is now suitable for study of phagocytosis. CFSE labeling myelin debris can be used to track early internalization as well as intracellular trafficking. Unlabeled myelin debris is compatible with Oil Red-O lipid staining to assess the storage and metabolism of the internalized lipids.
If you are using myelin debris that has been stored at negative 80 degrees Celsius, you'll first want to resuspend with a 29-gauge needle. Resuspend 10 milligrams of pelleted myelin debris in 200 microliters of PBS. Then add two microliters of five-millimolar CFSE solution.
Pipette to mix. After a 30 minute incubation, protected from light and subsequent washing, resuspend the myelin debris to 100 milligrams per milliliter with PBS. Following culture and fixation with 4%paraformaldehyde, add 100%propylene glycol to each well.
After incubating for five minutes at room temperature, add Oil Red-O staining solution to each well. Incubate for eight minutes at 60 degrees Celsius with gentle agitation. Gently tilt the plate to allow for complete aspiration of the Oil Red-O stain.
To each well add 85%propylene glycol. After five minutes of incubation at room temperature, wash wells with PBS and stain cells with your choice of nuclear dye. Representative light microscopy images of bone marrow cells during culture.
24 hours after initial seeding, few adherent cells are present. However, in the presence of macrophage colony-stimulating factor, M-CSF, leukocyte precursor cells begin to undergo differentiation. After seven days of culture in the presence of M-CSF, precursor cells are fully differentiated into mature bone marrow-derived macrophages capable of phagocytosis.
Using this method, a single eight-to 10-week-old C57 black 6J mouse will typically yield 18 to 24 million bone marrow macrophages. To visualize internalization, CFSE-labeled myelin debris is added to bone marrow-derived macrophage cultures. Cells were treated with one milligram per milliliter CFSE-labeled myelin debris for one hour prior to washing and fixation with 4%paraformaldehyde.
Internalized myelin debris can be visualized using standard GFP filter sets on an epifluorescent-capable microscope. To demonstrate how myelin lipid accumulation changes over time, macrophages were treated with unlabeled myelin debris for 90 minutes, then washed and fixed at various time points. Immediately following washing, few lipid droplets are visible.
However, as time progresses, more lipid droplets become visible, reaching a maximum approximately 24 hours after washing. Macrophages will begin to metabolize for efflux to lipid stores, reducing the number of stainable droplets. To quantify Oil Red-O staining, five randomly-obtained images from each time point sample were obtained using an epifluorescent-capable microscope.
The Oil Red-O-stained area was determined for each well using images captured in the DsRed channel. The total number of cells is determined by counting the number of nuclei present in each field. This graph shows the resulting quantification.
A steady increase in Oil Red-O-positive signal is typical, as the bone marrow-derived macrophages convert internalized myelin lipids into neutral lipids. A decrease in Oil Red-O-positive signal follows the peak at 24 hours as macrophages begin to metabolize or efflux the lipids. Having watched this video, you should have a good idea of how to isolate and culture primary bone marrow-derived macrophages, as well as how to study their interaction with freshly isolated brain-derived myelin debris.
The most important part of these procedures is the proper maintenance of aseptic technique at all times. Because macrophages can robustly respond to small quantities of pathogen-associated molecules, it is critical to minimize any potential interaction that can bias results. The main advantage of these techniques is that they make us of biologically relevant primary cells and fresh myelin debris, while eliminating the amount of specialized equipment needed.
Additionally, with some user optimization, these methods can be easily adapted to address a wide variety of questions regarding the response of bone marrow-derived macrophages to myelin debris. By making use of both florescently-labeled myelin debris and Oil Red-O staining of myelinated macrophage, it is possible to investigate potential therapeutic interventions for patients suffering from a variety of neurotrauma. The overall goal of such work is for remote macrophage-mediated cellular debris clearance to promote inflammation resolution.
