Measuring Microglial Uptake of Intravitreally-Delivered Fluorescent Particles Using Flow Cytometry

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.