Our research aims to understand the fundamental regulatory mechanisms controlling gene expression in the brain. And we're particularly interested in understanding how disruption of these mechanisms contributes to neurodevelopmental and neuropsychiatric conditions. We focus on understanding how genetic and environmental risk factors combine to alter cellular function and produce behavioral impairments.
This protocol includes methods for high-throughput screening of histone modifications in isolated microglia, allowing us to screen through dozens of different epigenetic modifications before committing to high-throughput sequencing and functional follow-up studies. Our protocol provides measures of global histone modifications, similar to western blot. However, our technique is both faster, requires fewer input cells, is more quantitative and can be multiplexed.
Begin by placing the isolated mouse brain tissue and one milliliter of digestion buffer per brain in separate Petri dishes on ice. Thoroughly chop the brain tissues into small pieces using a clean scalpel blade. Using a plastic transfer pipette with its tip cut off, carefully transfer minced brains into individual wells inside a 24 well plate on ice.
Cover the plate with a large piece of transparent, flexible film, close the lid and incubate on ice for 30 minutes. Next, transfer the digested brain solution into individual seven milliliter iced glass Dounce homogenizers on ice containing five milliliters of cold FACS buffer. Gently dounce each brain with the loose pestle until a single cell suspension is obtained.
Softly dounce with the tight pestle three to four times to ensure a single cell suspension before transferring to 15 milliliter polypropylene tubes. Begin by adding 2.125 milliliters of isotonic density gradient to a 15 milliliter polypropylene tube containing murine brain homogenate. Top up each tube with FACS buffer to a final volume of 8.5 milliliters.
Gently invert the tubes 20 times to mix thoroughly. Using a narrow graduated transfer pipette, gently underlay four milliliters of pink 37%density gradient, establishing two clean layers. Switch transfer pipettes and underlay two milliliters of the blue 70%density gradient below the 37%layer.
Transfer the tubes to a centrifuge cooled to four degrees Celsius and spin them at 500G for 20 minutes with the braking ramp set to the lowest setting. After centrifugation, discard the myelin from the top of the 15 milliliter tube using a clean transfer pipette. Carefully collect the top fragment of the density gradient into a clean 15 milliliter polypropylene tube using another transfer pipette.
Next, gather all cells in the sample by slowly circling the pipette along the sides of the tube, while collecting the immune-enriched fragment. Transfer the immune-enriched fragment into a new 15 milliliter polypropylene tube. Wash the sample by adding 10 milliliters of FACS to the tube.
Gently invert the tube 20 times to mix thoroughly. Spin the tubes in a cooled centrifuge with the braking ramp set to the lowest setting to pellet the isolated immune cells. Transfer a mouse brain enriched immune pellet from a 15 milliliter tube to a round bottom 96 well plate.
After centrifuging the plate at 500G for five minutes, remove the supernatant by flicking. Resuspend the cells in 100 microliters of FACS buffer. Then transfer 10 microliters from each well into the control wells.
Centrifuge at 500G for five minutes. After removing the supernatant from the centrifuged cells add 50 microliters of double concentration FC block. Incubate for 10 minutes on ice.
Now, add 50 microliters of the antibody master mix at double the final desired concentration and incubate it on ice in the dark. Then add 200 microliters of the FACS buffer to each well and centrifuge at 500G for five minutes. Discard the supernatant by flicking.
Then wash the cells in 300 microliters of FACS buffer. Centrifuge in resuspend the cells in 200 microliters of FACS buffer. Finally, transfer the suspension to a flow sort tube containing 300 microliters of FACS buffer.
To analyze the antibody panel, confirm that the cytometer has a minimum of four lasers including red, yellow, blue and violet. Establish the compensation matrix with either compensation beads or single stain cell controls. To calibrate and standardize, run rainbow fluorescent beads at the beginning of each experiment by adjusting the photomultiplier tube voltage until the bead peaks align with the target values from the previous experiments.
Set the photomultiplier tube voltage and gain for the experiment. Then use antibody captured compensation beads to establish a compensation matrix. Next, plot the side scatter area on log versus the forward scatter height on linear in a dot plot.
Gate out debris and select for cell size using the S1 gate. Choose singlet cells in a dot plot of forward scatter width versus forward scatter height with S2 gate. For establishing fluorophore gates, use the relevant FMO for each fluorophore channel.
With single parameter histograms, determine the positive signal in each channel and establish the gates accordingly. Carefully record the samples using the established gating strategy. Identify microglia using P2 RY 12 plus signal and determine protein expression in the respective channel for microglia only.
Establish analysis gates on the cytometer analysis software user interface by replicating the same gating strategy used during recording. Use the add statistics function to select the median for the population of interest on the compensated channel height. Export the MFI values for the respective channels to a spreadsheet using the table editor for further statistical analysis.
Lipopolysaccharide treatment induced an increase in histone H3 lysine 27 acetylation when the MFI is normalized within sets. Histogram analysis of the stained cells showed normally distributed populations with cell shifts to increased fluorescence. A similar increase in histone H3 lysine 27 acetylation was seen in the stained cells.
