Isolation of Macrophages from Mouse Dorsal Root Ganglion Using Mechanical Dissociation

Before starting the experiment, prepare the working solution of the density gradient medium by mixing nine volumes of the medium with one volume of calcium and magnesium-free 10X HBSS, and keep it on ice. Confirm that the mouse is fully anesthetized by the lack of response to the hind paw pinch.

Begin by placing an anesthetized mouse inside a chemical fume hood in the supine position with four paws secured with tape. Use forceps to lift the skin below the rib cage, and then with surgical scissors, make a small incision to expose the liver and diaphragm. Using the same scissors, cut the diaphragm and the rib cage, and then open the pleural cavity to expose the beating heart.

Next, with iris scissors, quickly cut the right atrial appendage. Once the bleeding is noted, insert a 30-gauge needle into the posterior end of the left ventricle, and slowly inject 10 milliliters of pre-chilled 1X PBS to perfuse the animal.

To perform dorsal laminectomy, place the mouse in the prone position. Use a size 11 scalpel to make two longitudinal lateral deep incisions starting from the thoracic region down to the sacral region. Then, remove the skin exposing the dorsal muscle layer. Then, with Friedman-Pearson Rongeur, peel off the connective tissues and muscles until the lumbosacral spine processes and bilateral transverse processes are exposed.

First, use a Friedman-Pearson Rongeur to carefully open the dorsal spinal column. Then, switch between a Friedman-Pearson Rongeur and Noyes Spring Scissors to remove the vertebral bones, exposing the spinal cord with intact spinal nerves attached. Carefully dissect ipsilateral and contralateral lumbar DRG.

Place it into 1 milliliter of ice-cold calcium and magnesium-free 1X HBSS in a Dounce tissue homogenizer. Homogenize the DRG tissue in the Dounce homogenizer with a loose pestle for 20 to 25 times. Place a sterile 70-micrometer nylon cell strainer in a sterile 50-milliliter conical tube. Use 800 microliters of ice-cold 1X HBSS to wet the cell strainer.

Use a pipette to collect the homogenized tissue suspension from the homogenizer onto the wet cell strainer, and collect it into the 50-milliliter conical tube. Rinse the homogenizer twice with 800 microliters of ice-cold 1X HBSS, collecting the liquid into the same 50-milliliter conical tube to increase the yield.

Add 1.5 milliliters of equilibrated ice-cold isotonic density gradient medium into a sterile 5-milliliter polystyrene FACS tube. Transfer the cell homogenate from the 50-milliliter conical tube into this FACS tube, and pipette up and down to mix well.

Add an additional 500 microliters of 1X HBSS to seal the top. Centrifuge the cells at 800 times g for 20 minutes at 4 degrees Celsius. Carefully aspirate the supernatant containing myelin in the medium, without disturbing the cell pellet at the bottom of the tube.

Add 100 microliters of PBS containing 5% fetal bovine serum to the pellet, and resuspend the DRG cells. Then, add alpha-mouse CX3CR1-APC antibody to the cells, and incubate in the dark at 4 degrees Celsius for 1 hour.

Wash the cells once with 5 milliliters of PBS. Centrifuge the cells at 360 times g for 8 minutes at 4 degrees Celsius. Aspirate the supernatant, and then, resuspend the cell pellet in 300 microliters of PBS for FACS analysis.