This protocol is designed to digest whole mouse eyes into a single cell suspension for multi-parameter flow cytometry of mononuclear phagocytes which include macrophages, monocytes, microglia, and dendritic cells. The main advantage of this technique is the ability to use quantitative analysis with fluorophore intensity to differentiate closely-related cell types using cell surface marker expression. Our protocol is optimized for the analysis of macrophage heterogeneity, including both resident and infiltrating populations in the laser-induced choroidal neovascularization model.
However, these concepts and techniques can be equally applied to other models like diabetic retinopathy, experimental autoimmune uveitis, as well as manifestations of lupus. When attempting this protocol, understanding your cytometer's configuration is a critical starting point as all cytometers are not configured the exact same way. Thus, volumes of fluorescently conjugated antibodies may need to be titrated and/or fluorophores may need to be adjusted to optimize the protocol based on your cytometer's configuration.
Begin by preparing the digestion buffer according to manuscript directions. Using a dissection microscope and 10X total magnification, remove the remaining conjunctiva and optic nerve from the eyes. Move clean eyes to a dry dissecting dish or a small weigh boat, then make two perforating wounds with a 90 degree angle between them through the visual axis.
Use a syringe with a 30 gauge needle to inject 0.15 to 0.2 milliliters of digestion buffer into the vitreous cavity of each eye. Mince the eyes with spring scissors and fine forceps, then dissociate the lens tissue with blunt mechanical disruption to prevent clogging pipette tips in subsequent steps. Rinse the forceps and scissors with 0.75 milliliters of digestion buffer and replace the dish for each sample.
Use a P1000 pipette to transfer the contents of the dish to a dissociation tube on ice, taking care to not lose any material. Rinse the dish with an additional 1.5 milliliters of digestion buffer and add it to the dissociation tube, bringing the total volume to between 3.25 and 3.5 milliliters. Dissociate the tissue on an electronic dissociator as described in the text manuscript, making sure that all tissue is at the bottom of the inverted dissociation tube and in contact with the digestion buffer.
Then incubate the dissociation tube at 37 degrees Celsius and 200 RPM for 30 minutes. Repeat the tissue dissociation two more times, incubating the sample at 37 degrees Celsius and 200 RPM for 30 minutes in between the dissociations. To stop the reaction, add 10 milliliters of cold flow buffer and place the sample tubes on ice.
To create a single cell suspension, strain the halted eye digestion reaction through a 40 micrometer filter placed on the top of a 50 milliliter conical centrifuge tube. Use the plunger from a 2.5 milliliter syringe to push any remaining pieces of undigested eye tissue through the filter. Wash the dissociation tube with 10 milliliters of flow buffer and rinse the filter.
Then use the same plunger to again pass the undigested eye tissue pieces through the filter. Repeat this process with five milliliters of flow buffer. Then wash the dissociation tube and filter with a final 10 milliliters of flow buffer.
Centrifuge the conical tube at 400 times G for 10 minutes at four degrees Celsius and decant the supernatant without disturbing the cell pellet. Break up the pellet by flicking the tube against another tube. Then add one milliliter of lysing solution and swirl the tube at room temperature for 30 seconds to lyse the red blood cells.
Stop the reaction with 20 milliliters of flow buffer and centrifuge of the conical tube at 400 times G for 10 minutes at four degrees Celsius. Decant the supernatant without disturbing the cell pellet. Flick the tube against another tube.
Then add five milliliters of cold HBSS. Repeat the centrifugation and remove the supernatant. Add 0.5 milliliters of live dead stain to each sample using a P1000 pipette, making sure to dissociate the pellet completely.
Transfer the samples to 1.2 milliliter microtiter tubes and incubate them for 15 minutes in the dark. Meanwhile, count a 10 microliter aliquot of the sample with trypan blue and a hemocytometer. After the incubation, wash each sample by adding 400 microliters of cold flow buffer.
Centrifuge the tubes in a microtiter tube rack at 400 times G for 10 minutes at four degrees Celsius and aspirate the supernatant without disturbing the cell pellet. Resuspend the cells completely in 500 microliters of cold flow buffer. Then repeat the centrifugation and remove the supernatant.
Block up to five million living cells in 50 microliters of FC block. Completely dissociate the cell pellet and incubate it at four degrees Celsius for 20 minutes. Then add 50 microliters of antibody staining solution to each sample.
Mix it completely and incubate it for another 30 minutes at four degrees Celsius. Forward scatter area versus side scatter area properties for all analyzed events from two eyes of a single mouse are shown here. Bead counts were cleaned and confirmed by plotting PE over APC-Cy7, creating a tight cluster of PE positive and APC-Cy7 negative events.
Next, singlets in live cells were identified from all events. Singlets were positively correlated in forward scatter height versus forward scatter area while doublet and triplet cells had greater forward scatter area than forward scatter height. Live cells are forward scatter area positive and live dead stain negative.
The initial gating strategy for the delineation of mononuclear phagocytes from live singlet cells is shown here. Live singlets were visualized using a CD45 versus CD11b plot. The absence of CD45 positive cells in the CD45 FMO confirmed the gate selection.
Next, neutrophils, eosinophils, B cells, and K cells, and T cells were excluded by plotting CD45 positive live singlets with lineage gate versus CD11b. CD45-dim cells were differentiated from CD45-high cells. Macrophage subsets were also identified.
Microglia were delineated in the CD45-dim cells with CD64 positive MHC2 low staining. MHC2 negative macrophages were identified as CD64 positive MHC2 negative. CD11c negative and CD11c positive macrophages were demonstrated as CD64 positive CD11c negative.
and CD64 positive CD11c positive in the MHC2 positive cells. It was found that laser treatment increased the amount of MHC2 negative, CD11c negative, and CD11c positive macrophages. Dendritic cell counts were also upregulated by laser treatment while microglia and monocyte counts were not affected.
Systemic perfusion had no effect on macrophage numbers. Following this procedure, florescence activated cell sorting rather than analysis alone allows for characterization of macrophage heterogeneity and its impact upon function via transcriptomic or proteomic studies.
