強化された純度と汎用性を持つ初代ミクログリアの迅速かつ洗練されたCD11b磁気分離

The overall goal of CD11b magnetic microglia isolation is to achieve a high purity end yield of postnatal microglia in a relatively short time period for the purposes of versatile in vitro experimentation. This method can help answer key questions in the field of neuroscience, such as the signaling mechanisms pertinent to neural inflammation. The main advantage of this technique over currently available methods is that the isolation takes approximately 30 minutes, with no sacrifices in purity, yield, or viability.

Begin by transferring freshly decapitated heads from one-to two-day-old mouse pups to a laminar airflow hood. Then, use 4.5-inch straight micro-dissecting scissors to make a small incision in the skull and meninges by inserting the tips in the opening formed by the decapitation and cutting caudal to rostral. After making the incision, peel one of the hemisphere to the side.

Then use a pair of curved or hooked tweezers to remove the entire brain. Transfer the brain to a 50-milliliter tube containing two milliliters of 0.25%trypsin EDTA, and incubate for 15 minutes in a 37-degree Celsius water bath. Wash the brain with fresh growth medium by adding and removing the medium.

After completion of the washes, add two milliliters of growth medium per brain, and homogenize by triturating the brain. When the brain tissue does not get smaller, transition to a pipette with a smaller aperture. At the end of trituration, when the suspension is clear with no visible chunks, pass the suspension through a 70-micron cell strainer to create a single cell culture.

Then pipette eight to nine milliliters of growth medium into T75 flasks, two flasks per brain, and add on milliliter of brain homogenate to each flask. Grow the cells until isolation on the sixteenth day. After sixteen days, transfer the growth medium from the flask to a fresh 50-milliliter tube.

Add three milliliters of 0.25%trypsin EDTA to each T75 flask. Then shake the flasks for five minutes at room temperature on an orbital shaker. Centrifuge the growth medium at 0.4 times g for five minutes.

After shaking for five minutes, add a minimum of four milliliters of the centrifuged culture medium to stop the trypsin EDTA reaction, and triturate to ensure that all the cells have been detached. Then pass the cells through a 70-micromolar cell strainer to create a single cell culture. Perform a cell count, and then spin down at 0.4 times g for five minutes.

Next, take a five-milliliter polystyrene tube, add one milliliter of recommended medium, and mark the meniscus. Add recommended medium up to 2.5 milliliters, and mark this meniscus as well. Then, remove the medium, and re-suspend the pellet in 500 microliters of recommended medium.

Transfer this mix into a five-milliliter polystyrene tube, and then dilute to the one milliliter mark. Then add 50 microliters of rat serum for every one milliliter of suspended cells, and incubate for five minutes at room temperature. During the incubation, prepare the selection cocktail by mixing 25 microliters of component A and 25 microliters of component B.Once the five minutes have elapsed, add 50 microliters of the selection cocktail to the cells, and incubate for a further five minutes at room temperature.

Vortex the microspheres for 45 seconds. Then add 80 microliters of microspheres per one milliliter of sample, and incubate for three minutes at room temperature. Next, add recommended medium up to the 2.5 milliliter mark on the polystyrene tube.

Put the tube in the magnetic holder for three minutes at room temperature. After the incubation, slowly pour out the medium into a 15-milliliter polystyrene tube still in the magnet. Repeat the magnetic separation three more times, each time adding recommended medium up to the 2.5 milliliter mark.

After the last magnetic separation, add three milliliters of growth medium, and count the number of cells using a cell counter. Plate the cells in poly-D-lysine-coated plates for treatments. Plate the negative fraction from the 15-milliliter tube, which mostly contains astrocytes, in T75 flasks.

After at least six hours of incubation in a 37-degree Celsius incubator, replace all the growth medium in the flasks containing the negative fraction before returning to the incubator. This image shows immunocytochemistry of an isolated microglial culture probed for IBA, a marker of microglia in the green channel, and GFAP, a marker of astrocytes, in the red channel. Hoechst staining reveals the presence of cell nuclei.

The absence of GFAP-positive cells demonstrates that microglia isolated using the CD11b positive selection kit, too, have high purity. These immunoblotted proteins from an isolated microglial culture were probed for GFAP, which appears as an approximately 51-kilodalton band, and IBA1, which appears as an approximately 15-kilodalton band. Beta-actin was used as a loading control and appears at approximately 42-kilodaltons.

A problem with the old isolation kit was that PE fluorescence could be observed in the red channel as seen here. The modified procedure does not produce any auto-fluorescence from the magnetic beads, as seen in the red channel here. The microglia isolated using this procedure can be used for signaling studies.

Western blot analysis shows that Fyn and phospho-src tyrosine Y416 can be detected from microglia isolated with our newly-refined method. The negative fraction from the microglial separation contains astrocytes that can be used for signaling studies. Western blotting shows that the negative fraction contains GFAP-positive cells.

Western blot analysis shows that Fyn and phospho-src tyrosine Y416 can be detected from the GFAP-positive cells. Following this procedure, other methods, like multi-channel fluorescence microscopy, western blotting, and quantitative PCR, can be performed to answer questions regarding gene expression and protein signaling. After watching this video, you should have a good understanding of how to utilize this rapid isolation technique for the purposes of microglial experimentation.