Isolation and Whole-Cell Patch-Clamp Recording of Hippocampal Microglia from Adult Mice

Our research focuses on acutely isolating microglia for whole-cell recordings, including potassium currents to understand the ion-channel activity. The protocol is also adaptable for isolating microglia for various brain regions, providing insights into zero across different neurology conditions.

The most recent developments in our research field include the use of traditional pancreatitis enzyme digestion, combined with microglia purification through fluorescence active cell sorting. These advancements have enhanced the accuracy and efficiency of isolating specific microglia populations for further study.

Our protocol investigates the electro-physiological properties of microglia directly from the adult mouse hippocampus, rather than relying on primary cultured CS. This approach allows for a more accurate analysis of microglia functions in their native environment.

Our results have paved the way for new scientific questions about the biochemical and the functional property of microglia under both physiological and disease conditions, offering researchers valuable guidance in exploring these aspects further.

[Presenter] To begin, place the anesthetized animal supine on a dissecting table and pin their limbs. Expose the chest and perfuse the heart with ice-cold, sterile PBS to remove intravascular circulating blood cells. Then hold the head and cut the skin along the brain midline with scissors to expose the skull. Next, cut the posterior of the skull bilaterally and between the eye sockets with small scissors. Cut the skull along the sagittal suture and remove the skull with tweezers. After that, extract the brain tissue quickly. Now, place the brain tissue in 10-centimeter culture dishes containing ice-cold PBS. Dissect the cerebral cortex bilaterally with tweezers. Isolate the hippocampal tissue and cut it into smaller pieces. Transfer the pieces into a 15-milliliter centrifuge tube. Then re-suspend the small pieces of hippocampal tissue in two milliliters of ice-cold Hanks' Balanced Salt Solution or HBSS. And centrifuge at 300 G for three minutes at room temperature. After discarding the supernatant, add 1,950 microliters of enzyme mixture one to the 15-milliliter centrifuge tube containing the hippocampal tissue pieces. Shake the centrifuge tube continuously in a 37-degree Celsius water bath for five minutes. Next, add 30 microliters of freshly prepared enzyme mixture two to the 15-milliliter centrifuge tube. Pipette the tissue fragments up and down 20 times with a one-milliliter pipette tip and incubate in a water bath for five minutes with continuous shaking at 37 degrees Celsius. After removing the centrifuge tube, pipette the suspension up and down multiple times with a one-milliliter pipette until no large tissue pieces remain. Then filter the cell suspension through a 70-micrometer cell screening filter and collect the filtrate in a clean 50-milliliter centrifuge tube. Transfer the filtrate to a new 15-milliliter centrifuge tube and centrifuge at 300 G for 10 minutes at four degrees Celsius. Then discard the supernatant with a one-milliliter pipette. Now, pipette the cell suspension up and down 20 times with 15 milliliters of HBSS to wash it and centrifuge at 300 G for 10 minutes at four degrees Celsius. After discarding the supernatant, re-suspend the cell pellet in one milliliter of magnetic-activated cell sorting buffer. To begin, transfer one milliliter of the prepared single-cell suspension into a two-milliliter microcentrifuge tube and centrifuge at 300 G for three minutes at four degrees celsius. Once the supernatant is removed, re-suspend the single-cell pellet in 90 microliters of magnetic-activated cell sorting or max buffer. Add 10 microliters of CD11b or c microbeads and incubate the suspension at four degrees Celsius for 15 minutes on a horizontal shaker. Then add one milliliter of max buffer directly to the tube to wash the single-cell suspension. After centrifugation, re-suspend the cell pellet in 500 microliters of max buffer. Using a one-milliliter pipette, transfer the re-suspended cell suspension to the sorting column in the magnetic field. While it remains in the magnetic field, wash the column three times with 500 microliters of max buffer. Then remove the sorting column from the magnetic field. Add one milliliter of max buffer to the column and slowly push it through with the piston. Collect the flow through. Now, seed the selected cells into a 24-well plate containing a cover slip coated with Poly-L-lysine. Add 200 microliters of culture medium and incubate for 20 minutes in a 37-degree Celsius incubator with 95% oxygen, 5% carbon dioxide, and 60 to 80% humidity. To begin, take the cultured microglia isolated from mice brain tissue. Locate the spherical microglia under a water microscope using the 40x objective. Next, fill the electrode with intracellular solution and attach it to the amplifier head stage of the electrophysiology setup. Apply positive pressure. Switch to the 4x objective and locate the electrode. Then lower the electrode into the bath solution. Set the amplifier to voltage clamp mode to perform the membrane test in bath mode. After that, switch back to the 40x objective and move the electrode tip closer to the microglia. Next, apply a tiny negative pressure to hold the cell. When the resistance reaches 100 megaohms, switch from bath to patch mode in the membrane test. When the resistance reaches 300 megaohms, release the negative pressure. When the resistance indicates the formation of a giga-ohm seal between the electrode and the microglial membrane. Switch from patch to cell mode in the membrane test. Then apply a small amount of suction and initiate an electric shock to rupture the membrane. After that switch to voltage clamp mode, open the recording program and record potassium currents for 200 milliseconds.