This article describes a protocol for isolating and analyzing mouse retinal myeloid cells, including microglia, monocytes, and neutrophils. The method employs flow cytometry to identify and characterize these immune cells based on specific surface markers.
Begin with a multi-well plate containing a single-cell suspension of mouse retinal myeloid cells, including microglia, monocytes, and neutrophils.
Microglia — or retinal phagocytic immune cells, exhibit fluorescence due to engulfed fluorophore-labeled bioparticles of fungal origin, preinjected intravitreally.
Centrifuge and add Fc-receptor antibodies, blocking non-specific interaction sites.`
Introduce a cocktail of fluorophore-labeled antibodies targeting CD11b, Ly6C, and Ly6G surface proteins, interacting with myeloid cells expressing them.
Remove unbound antibodies. Treat with propidium iodide, which permeates through damaged membranes, exclusively staining dead cells.
In flow cytometry, excite propidium iodide to exclude dead cells and identify the non-fluorescent live cells.
Illuminate the cells with a violet laser; excited live cells signify the CD11b-expressing myeloid cells.
Next, illuminate with a red laser, exciting the near-far-red fluorophore within CD11b-positive cells, indicating Ly6C-expressing monocytes and Ly6G-expressing neutrophils.
Excite the cells with a blue laser; the emission of green fluorescence from the remaining CD11b-positive cells confirms the microglial uptake of fluorescent particles.
Three hours after intravitreal injection, use 45-degree angle forceps to gently press against the eyelid to proptose the eyeball. Position the forceps behind the eyeball, and pull. Then, transfer the eyeball to the dry area of a Petri dish containing a small volume of PBS with calcium and magnesium under a dissecting microscope, and use one tip of the super-fine forceps to perforate the eye in the corneal limbus. Next, holding the eyeball with fine 45-degree angled forceps, use spring scissors to cut around the corneal limbus until roughly half of the circumference is cut.
Transfer the eyeball into PBS and use a second pair of fine 45-degree angled forceps to tear the cornea and sclera apart. The lens and retina will come out intact. Ensure the lens and retina are separated and transfer the retina to a 5.4-milliliter polystyrene test tube containing 2 milliliters of PBS with calcium and magnesium.
To obtain a single-cell suspension of the retinal cells, use a neuronal tissue dissociation kit according to the manufacturer's instructions and resuspend the cells in 200 microliters of staining buffer. Then, transfer the sample to one well of a 96-well U-bottom plate. After centrifugation, invert the plate to discard the supernatant, and block the FC receptors with 25 microliters of stain buffer containing CD16/CD32 antibody per well for 5 minutes at room temperature. Next, label the cells with the antibodies of interest for 15 minutes at room temperature in the dark.
Then, pellet the cells and follow this by a wash in 200 microliters of fresh staining buffer per well. Now, resuspend the pellets in 200 microliters of stain buffer and viability dye and transfer the samples to 1.2-milliliter microtiter tubes. Then, wash the wells with an additional 100 microliters of staining buffer and viability dye and pool the washes with the corresponding samples for analysis by flow cytometry.
繁體中文
