Digestion of Whole Mouse Eyes for Multi-Parameter Flow Cytometric Analysis of Mononuclear Phagocytes

All procedures were approved by the Northwestern University Institutional Animal Care and Use Committee. C57BL/6 mice were group housed at a barrier facility at the Center for Comparative Medicine at Northwestern University (Chicago, IL). All animals were housed in 12/12 h light/dark cycle with free access to food and water.

1. Collection of ocular tissue

1. Sacrifice 10-12-week-old C57BL/6J mice of either sex using regulated administration of carbon dioxide until mouse is unresponsive and no longer inhaling. Perform cervical dislocation or other approved secondary method to ensure euthanasia.
   NOTE: Transcardial perfusion can be performed to remove circulating leukocytes that are not in the ocular tissues. In this experiment, systemic perfusion does not reduce the number of microglia, monocytes, macrophages, or dendritic cells detected in untreated or laser treated whole eyes. Therefore, the majority of these cell types are located in the ophthalmic tissues and this step is optional.
2. To enucleate eyes, push down near the eye socket while placing a pair of curved forceps gently under the eye as it stands out of the socket. Pinch forceps closed under the eye without squeezing the actual eye. Dislodge eye from the surrounding conjunctiva (please see prior publication)²³.
3. Pull eye laterally until the optic nerve is the only remaining connection to detach the eye. The optic nerve often severs with tension, but a small scissor is sometimes necessary to cut the optic nerve. Place the eye in cold 1x Hank’s Buffered Saline Solution (HBSS) with calcium and magnesium in 1.7 mL microcentrifuge tube.

2. Digestion of ocular tissue

1. Prepare the digestion buffer containing 0.1 mg/mL digestion enzyme and 0.2 mg/mL DNase in cold HBSS. Reconstitute 5 mg digestion enzyme in 2 mL of sterile water initially. Prepare an initial dilution of 10 mg/mL DNase in HBSS. For every 10 mL of the digestion buffer (sufficient for 3 animals) mix 9.4 mL of cold HBSS with 0.4 mL of reconstituted digestion enzyme and 0.2 mL of diluted DNase I.
   NOTE: These concentrations were empirically determined over multiple experiments to provide optimal levels of antibody staining of single cells, while reducing cell death. Collagenase D was tried as an alternative to digestion enzyme with reduced cell viability and decreased CD45⁺ cells. Please see Discussion for more details regarding the iterative process to optimize digestion.
2. Under a dissecting microscope at 10x total magnification, remove the remaining conjunctiva and optic nerve using fine forceps and spring scissors in cold HBSS.
3. Move clean eyes to a dry dissecting dish or small weigh boat. Inject 0.15-0.20 mL/eye of digestion buffer via syringe fitted with 30 G needle into the vitreous cavity after two perforating wounds are made in the eye through the visual axis and 90˚ away from this first wound for digestion buffer egress.
4. Mince whole eyes using spring scissors and fine forceps with all pieces smaller than 0.5 mm x 0.5 mm. Dissociate lens tissue with blunt mechanical disruption via forceps to prevent clogging pipette tips in subsequent steps. For each sample, rinse forceps and scissors each with 0.75 mL of digestion buffer into small dish. Use a new dish for each sample.
5. Transfer contents of the dish to a dissociation tube on ice using P1000 pipette taking care to not lose any material in the pipette transfer. Rinse the dish with additional 1.5 mL of digestion buffer bringing the total volume in dissociation tube to 3.25-3.50 mL.
6. Place the dissociation tube inverted onto electronic dissociator and run cycle “m_brain_03_01”, consisting of 1 min unidirectional rotation at 200 rpm at room temperature, which is a pre-loaded program. Ensure all tissue is at the bottom of the dissociation tube in contact with digestion buffer and the rotor for this and all remaining steps.
7. Place the dissociation tube in shaking incubator for 30 min at 200 rpm at 37 ˚C.
8. Repeat steps 2.6 and 2.7 one additional time. Following second 30 min incubation perform one final mechanical digestion using electronic dissociator as in step 2.6.
9. Stop the reaction by adding 10 mL of cold flow buffer and place tubes on ice.

3. Preparation of cell suspension

1. To create single-cell suspension, pour the halted eye digestion reaction through a 40 μm filter placed on the top of a 50 mL conical centrifuge tube. Using the plunger from a 2.5 mL syringe, push any remaining pieces of undigested eye tissue through the filter.
2. Wash the dissociation tube with 10 mL of Flow buffer and rinse filter before using the same plunger to again pass undigested eye tissue pieces through the filter.
3. Repeat step 3.2 using 5 mL of Flow buffer.
4. Wash the dissociation tube and filter with a final 10 mL of Flow buffer.
   NOTE: Total volume of digestion and flow buffer is about 38-40 mL for each sample.
5. Centrifuge the conical tube at 400 x g for 10 min at 4 ˚C. Decant the supernatant without disturbing the cell pellet.
6. Prepare 1x lysing solution by adding 10x lysing solution to sterile water. Break up the pellet by flicking tube against another tube. To lyse red blood cells, add 1 mL of 1x lysing solution for 30 s with swirling at room temperature (RT).
7. Stop the reaction with 20 mL of Flow buffer.
8. Centrifuge the conical tube at 400 x g for 10 min at 4 ˚C. Decant the supernatant without disturbing the cell pellet.
9. Dissociate the pellet by flicking tube against another tube. Add 5 mL of cold 1x HBSS per tube.
10. Centrifuge tubes at 400 x g for 10 min at 4 ˚C. Aspirate supernatant.

4. Staining of cell suspension for mononuclear phagocytes

1. Prepare 0.5 mL of Live/Dead stain per sample by diluting Live/Dead dye 1:5,000 in cold HBSS.
2. Add Live/Dead stain to each sample using a P1000 pipette making sure to dissociate the pellet completely. Transfer samples to 1.2 mL micro titer tubes.
3. Take a small aliquot (10 μL) to count cells using a hemocytometer or automatic cell counter (see 4.4). Incubate samples with Live/Dead stain for 15 min at room temperature in the dark.
4. Count cells using Trypan blue to determine the number of live cells per sample.
   NOTE: Typically, 2 eyes from a 10-12-week-old C57BL/6J mouse contain less than 2 x 10⁶ living cells.
5. Wash samples by adding 400 μL of cold Flow buffer. Centrifuge tubes in a micro titer tube rack at 400 x g for 10 min at 4 ˚C. Aspirate the supernatant without disturbing the cell pellet.
6. Resuspend cells completely in 500 μL of cold Flow buffer. Centrifuge tubes in a micro titer tube rack at 400 x g for 10 min at 4 ˚C. Aspirate the supernatant without disturbing the cell pellet.
7. Block up to 5 x 10⁶ living cells in 50 µL of F_c block (1:50 dilution in cold Flow buffer of purified rat anti-mouse CD16/CD32). If more than 5 x 10⁶ living cells, increase volumes appropriately.
8. Add F_c block to the pellet making sure to completely dissociate the sample. Incubate for 20 min at 4 ˚C.
9. Add 50 μL of antibody staining solution (see Table 1 for choice and amount of each antibody that is included in the solution) per 5 x 10⁶ living cells directly to cells in F_c block and mix completely. Incubate for 30 min at 4 ˚C.
   NOTE: Antibody quantities are dependent upon the flow cytometer configuration and should be titrated independently for every lab and instrument. See Table 2 for instrument configuration of filters and mirrors. To titrate antibodies, start with manufacturer’s recommendation, and calculate the staining index. Perform a test run of varying antibody concentrations (both above and below the manufacturer’s recommendation) and choose the lowest concentration of antibody where the staining index is maintained. Also, ensure that positive staining that is less bright than the single-color control. Use unstained cells to ensure that negatively stained cells are on scale and have a peak at or less than 1 x 10². Use a fluorescence minus one control to define the positively stained cells.
10. Wash samples by adding 700 μL of cold Flow buffer. Centrifuge tubes in a micro titer tube rack at 400 x g for 10 min at 4 ˚C. Aspirate the supernatant without disturbing the cell pellet.
11. Optionally, after antibody staining, the samples can be fixed with 500 μL of 2% paraformaldehyde in Flow buffer. Fix samples for 15 min at room temperature. Then, perform one wash as described in 4.10. Store fixed samples at 4 ˚C. Count beads should be added on the day of flow cytometry data collection. Fixed samples should be run on flow cytometer in 1-2 days.
    CAUTION: Paraformaldehyde is a corrosive and a health hazard.
12. Wash samples again using 500 μL of cold Flow buffer. Centrifuge tubes in a micro titer tube rack at 400 x g for 10 min at 4 ˚C. Aspirate half of the supernatant without disturbing the cell pellet.
13. Resuspend pellets and transfer to a 96 well, U-bottom assay plate. Centrifuge plate at 400 x g for 5 min at 4 ˚C. To decant the supernatant, turn plate over and in one strong shake dislodge the liquid into a sink basin, without disturbing pellet.
14. In a new 1.2 mL micro titer tube, place 50 μL of well vortexed count beads.
15. Resuspend pellets in 96 well plate in 200 μL of cold Flow buffer. Add cell mixture on the top of beads from prior step. Run on flow cytometer in total volume of 250 μL / micro titer tube.

5. Collection of flow cytometry data

1. Turn on and prepare flow cytometer. Instructions listed here are for a modified four laser cytometer (Table 2).
2. Run a test sample, which includes a small aliquot from each sample, to ensure that all cells are on scale for each detector. Adjust voltages so that all cells are on scale.
3. Run single color controls (SCC) for every fluorophore used in staining cocktail (Table 3). Place micro titer tube into a larger 5 mL polystyrene tube for placement onto the flow cytometer stage.
   NOTE: Root fluorophores like BV421 (Pacific Blue), FITC, PE, Alexa647 (APC), and Alexa700 do not require the same antibody as the cocktail (i.e., CD19 PE for CD64 PE). However, tandem fluorophores like PE-CF594, PE-Cy7, or APC-Cy7 require the same antibody for SCC as cocktail due to potential dissociation of the conjugated fluorophores.
   1. Create the SCC for BV421 by adding 2 drops of compensation beads to a 1.2 mL titer tube along with 1 μL of hamster anti-mouse CD11c BV421. Compensation beads are used because CD11c BV421 is an IgG lambda subtype rather than kappa as described in 5.3.3. Incubate for 15 mins at 4 ˚C. Add 0.2 mL of flow buffer.
   2. Create the SCC for the Live/Dead dye by adding one drop of positive staining beads (green cap) from the amine reactive beads, followed by 1 μL of Live/Dead dye. Incubate for 15 min at room temperature in the dark. Add one drop of the negative beads (white cap) from the amine reactive beads. Add 0.2 mL of flow buffer.
   3. Create the remaining SCCs by adding one drop of positive staining beads (green cap) from the anti-rat and anti-hamster Ig kappa bead set, followed by the antibody quantity listed (Table 3) in a 1.2 mL titer tube. Incubate for 15 min at 4 ˚C. Add one drop of the negative beads (white cap) from the anti-rat and anti-hamster Ig kappa bead set. Add 0.2 mL of flow buffer.
   4. Vortex all bead mixtures completely. Store SCCs at 4 ˚C for up to 3 days.
      NOTE: The SCC for fluorescent reporter mice is unstained cells.
   5. When running SCCs, set voltages such that the SCC sample has a peak greater than any fluorescence for a sample of cells in the same detector channel. In addition, voltages should be set so that any SCC has at least a 0.5 Log difference between the histogram peak in the channel of interest versus any other channel (Figure 1).
4. Run samples on “Low” keeping events/second below 4000. If flow rate cannot be lowered appropriately, dilute samples with additional cold flow buffer. Record the volume added as this will be necessary for the count calculation (see representative results).
5. Collect at least 3 x 10⁶ events per sample. This may be higher than the cell number due to pigment granules and debris counting as events on your cytometer.
6. Record a Fluorescence Minus One (FMO) for each fluorophore other than Live/Dead for proper gating strategy in section 6.4. An FMO is a sample that has been stained with all fluorophores except one.
7. Clean and shut down flow cytometer according to manufacturer or facility instructions.

6. Annotation of flow cytometry data

1. Open a new worksheet using analysis software. Import FCS files from cytometer for all samples and single-color controls.
2. Use SCCs to create a compensation matrix.
3. Apply the compensation matrix to your stained samples and FMOs.
4. Use the FMOs to determine positive staining to draw gates.