The overall goal of this cell culture protocol is to generate highly pure cultures of microglia from developing or adult rodents that can be maintained without exposure to serum. This procedure can help answer key questions in neuroscience related to microglia morphology, fibrous ketosis, and microglia's interaction with other CNS cell types. The main advantage to this protocol is that microglia can be rapidly isolated from rodent tissue of any age and cultured without exposure to serum.
To remove the brain, cut the occipital condyle from the spinal cord of an animal on both sides with dissection scissors, being careful not to damage the brain. Then, cut skin along the top of the skull, and cut up one side along the parietal and frontal bone towards the nasal bone. Carefully pull back the top of the skull with forceps, quickly remove the brain, and place it into prechilled DPBS on ice.
Repeat the procedure for all remaining animals. After the brains have been collected in cold DPBS, chop each brain into one cube millimeter chunks in a petri dish on ice with a cold scalpel blade and transfer to an ice cold dounce homogenizer with five to seven milliliters of ice cold douncing buffer. Next, use a loose fitting dounce homogenizer to dissociate the tissue with 10 to 20 gentle and incomplete strokes.
Take care not to directly crush the tissue at the bottom of the homogenizer. Instead, impel the tissue through the space between the sides of the piston and the homogenizer. It is important to slowly homogenize the tissue with multiple rounds of douncing to preserve microglia viability.
Afterward, carefully remove the piston to prevent introducing air bubbles. Allow poorly dissociated tissue chunks to settle to the bottom of the homogenizer and transfer the supernatant to a new, chilled, 50 milliliter conical tube. Subsequently, replace the removed volume with fresh douncing buffer and repeat the dissociation procedure for a total of three to four rounds, or until all tissue has been dissociated.
In this procedure, add ice cold douncing buffer to the 50 milliliter conical tube containing the cell suspension and adjust the total volume to 33.5 milliliters. Next, add 10 milliliters of MSB to the cell suspension and mix thoroughly by inverting the tube several times. This will result in a 23%final concentration of MSB in a volume of 43.5 milliliters.
After that, centrifuge the cells for 15 minutes at 500 x g at 4 degrees Celsius with slow braking. Then, remove the top layer of myelin and debris and the supernatant with a pipette. Take care when removing the top layer to ensure as much of it is removed as possible.
Following that, resuspend the cell pellet in 12 milliliters of panning buffer. Gently triturate the cell suspension to break up any remaining clumps of cells. In this step, pass the cell suspension through a 70 micrometer cell strainer to remove large debris or cell clumps.
Rinse the OX42 coated panning dish three times with DPBS. Don't allow the plate to completely dry between washes. Pour off the last DPBS wash and apply the filtered cell suspension to the panning dish.
Gently swirl the plate to distribute the cells. Then, incubate the plate on a flat surface at room temperature for 20 minutes to allow cells to adhere. Do not incubate longer than 20 minutes or cells will become very difficult to recover from the dish.
Afterward, rinse the panning dish with DPBS 10 times to remove nonadherent cells. Microglia will be firmly attached to the plate, so swirl the plate with each rinse to ensure the removal of other nonadherent cells. Pour off the last DPBS wash and replace with 15 milliliters of DPBS and 200 microliters of trypsin.
To trypsinize the cells, incubate the dish for less than or equal to 10 minutes at 37 degrees Celsius. After 10 minutes of trypsinization, microglia will still be stuck to the plate. Pour off the mixture and gently wash the plate two times with DPBS to remove the trypsin.
Then, replace with 12 milliliters of ice cold MGM. Subsequently, place the panning dish on ice for two minutes to help weaken cell and substrate interaction and make sure that the dish is flat to prevent areas of the panning dish from drying out. After that, pipette vigorously with a 10 milliliter pipette and pipette controller on high speed to recover cells from the panning dish.
Draw a 16 by 16 grid with the stream of liquid from the pipette to try and remove all of the cells. It is important to find a balance between overall cell recovery and the health of your culture. Repeated pipetting of the cell supernatant against the panning dish will result in reduced viability of your culture.
Check the cells under a microscope at 20x magnification to make sure that they have detached from the plate. Mark the spots on top of the dish where cells are stuck and repeat pipetting in those areas. Collect the cell suspension and allocate three to four milliliters of supernatant in each 15 milliliter conical tube.
Spin it for 15 minutes at 500 x g at four degrees Celsius with slow braking. After 15 minutes, aspirate the supernatant, leaving 0.5 milliliters of MGM with the cell pellet. Then, resuspend each pellet in remaining MGM and pull the cells from all the tubes.
Following that, count the cells with a hemocytometer. Now, plate 15 microliters of collagen IV coating directly in the center of a 24 well anionic cationic coated tissue culture plate and incubate for 15 minutes at 37 degrees Celsius. After counting the cells with a hemocytometer, dilute them to 2.3 x 10 to the five per milliliter in MGM.
Aspirate collagen IV spot and immediately plate 15 microliters of cell suspension to this spot to yield 3.5 x 10 to the three cells per spot. Next, incubate for five to ten minutes at 37 degrees Celsius to allow the cells to adhere. Then, gently add 500 microliters of CO2 equilibrated TIC to the well.
If plating on cover slips, first coat the sterile glass cover slips with 10 micrograms per milliliter PDL in water for one hour at room temperature. Then, wash them three times with water and let them dry in a hood under UV light before proceeding with spot plating on the cover slips. In order to demonstrate successful isolation and culture of highly pure microglia, we isolated microglia from P21 rats and cultured these cells in TIC or TIC supplemented with 10%FCS for eight days.
Cells were then fixed and stained with microglial markers and proliferation markers. Cells cultured in TIC show a ramified morphology, low levels of proliferation, and expression of microglia markers. In comparison, cells cultured in TIC supplemented with 10%FCS also stained for microglial markers, but show high levels of proliferation and amoeboid morphology, properties often seen in activated microglia.
In addition to staining for cell markers, RNA-Seq on freshly isolated cells can provide a powerful and sensitive tool for evaluating the RNA profile of the cells and can help inform the starting purity of cultures. CD11b micropanning typically results in a small percentage of CD11b positive neutrophils and monocyte carryover, in addition to CD11b positive microglia. These cells will die in TIC after a few days, but can complicate analysis of earlier timepoints.
Here is a time lapse movie of P21 rat microglia after 14 DIV in TIC. After two weeks, serum free cultures show dynamic surveillance behavior with rapid process extensions and retractions throughout the dish. Here is a time lapse movie of P21 rat microglia after five DIV in TIC before and after exposure to fetal calf serum.
A rapid morphological change is evident after serum exposure. Once mastered, this protocol can be done in four to five hours, if performed properly. Following this procedure, other assays to monitor phagocytosis, chemotaxis, survival, cytokine release, transcriptional responses to stimuli, can be performed in order to answer additional questions about microglia function.
After watching this video, you should have a good understanding on how to rapidly isolate microglia from adult rodent tissue and culture them under serum free conditions.
