JAL was supported by NIH grant K08EY030923; CMC was supported by a K01 grant (5K01AR060169) from the NIH National Institute of Arthritis and Musculoskeletal Diseases and a Novel Research Grant (637405) from the Lupus Research Alliance.

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
<ol>
	<li>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.</li>
	<li>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.</li>
	<li>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&#8217;s Buffered Saline Solution (HBSS) with calcium and magnesium in 1.7 mL microcentrifuge tube.</li>
</ol>
2. Digestion of ocular tissue
<ol>
	<li>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.</li>
	<li>Under a dissecting microscope at 10x total magnification, remove the remaining conjunctiva and optic nerve using fine forceps and spring scissors in cold HBSS.</li>
	<li>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&#730; away from this first wound for digestion buffer egress.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Place the dissociation tube inverted onto electronic dissociator and run cycle &#8220;m_brain_03_01&#8221;, 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.</li>
	<li>Place the dissociation tube in shaking incubator for 30 min at 200 rpm at 37 &#730;C.</li>
	<li>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.</li>
	<li>Stop the reaction by adding 10 mL of cold flow buffer and place tubes on ice.</li>
</ol>
3. Preparation of cell suspension
<ol>
	<li>To create single-cell suspension, pour the halted eye digestion reaction through a 40 &#956;m 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.</li>
	<li>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.</li>
	<li>Repeat step 3.2 using 5 mL of Flow buffer.</li>
	<li>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.</li>
	<li>Centrifuge the conical tube at 400 x g for 10 min at 4 &#730;C. Decant the supernatant without disturbing the cell pellet.</li>
	<li>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).</li>
	<li>Stop the reaction with 20 mL of Flow buffer.</li>
	<li>Centrifuge the conical tube at 400 x g for 10 min at 4 &#730;C. Decant the supernatant without disturbing the cell pellet.</li>
	<li>Dissociate the pellet by flicking tube against another tube. Add 5 mL of cold 1x HBSS per tube.</li>
	<li>Centrifuge tubes at 400 x g for 10 min at 4 &#730;C. Aspirate supernatant.</li>
</ol>
4. Staining of cell suspension for mononuclear phagocytes
<ol>
	<li>Prepare 0.5 mL of Live/Dead stain per sample by diluting Live/Dead dye 1:5,000 in cold HBSS.</li>
	<li>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.</li>
	<li>Take a small aliquot (10 &#956;L) 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.</li>
	<li>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.</li>
	<li>Wash samples by adding 400 &#956;L of cold Flow buffer. Centrifuge tubes in a micro titer tube rack at 400 x g for 10 min at 4 &#730;C. Aspirate the supernatant without disturbing the cell pellet.</li>
	<li>Resuspend cells completely in 500 &#956;L of cold Flow buffer. Centrifuge tubes in a micro titer tube rack at 400 x g for 10 min at 4 &#730;C. Aspirate the supernatant without disturbing the cell pellet.</li>
	<li>Block up to 5 x 106 living cells in 50 &#181;L of Fc 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.</li>
	<li>Add Fc block to the pellet making sure to completely dissociate the sample. Incubate for 20 min at 4 &#730;C.</li>
	<li>Add 50 &#956;L 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 Fc block and mix completely. Incubate for 30 min at 4 &#730;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&#8217;s recommendation, and calculate the staining index. Perform a test run of varying antibody concentrations (both above and below the manufacturer&#8217;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.</li>
	<li>Wash samples by adding 700 &#956;L of cold Flow buffer. Centrifuge tubes in a micro titer tube rack at 400 x g for 10 min at 4 &#730;C. Aspirate the supernatant without disturbing the cell pellet.</li>
	<li>Optionally, after antibody staining, the samples can be fixed with 500 &#956;L 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 &#730;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.</li>
	<li>Wash samples again using 500 &#956;L of cold Flow buffer. Centrifuge tubes in a micro titer tube rack at 400 x g for 10 min at 4 &#730;C. Aspirate half of the supernatant without disturbing the cell pellet.</li>
	<li>Resuspend pellets and transfer to a 96 well, U-bottom assay plate. Centrifuge plate at 400 x g for 5 min at 4 &#730;C. To decant the supernatant, turn plate over and in one strong shake dislodge the liquid into a sink basin, without disturbing pellet.</li>
	<li>In a new 1.2 mL micro titer tube, place 50 &#956;L of well vortexed count beads.</li>
	<li>Resuspend pellets in 96 well plate in 200 &#956;L of cold Flow buffer. Add cell mixture on the top of beads from prior step. Run on flow cytometer in total volume of 250 &#956;L / micro titer tube.</li>
</ol>
5. Collection of flow cytometry data
<ol>
	<li>Turn on and prepare flow cytometer. Instructions listed here are for a modified four laser cytometer (Table 2).</li>
	<li>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.</li>
	<li>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.
	<ol>
		<li>Create the SCC for BV421 by adding 2 drops of compensation beads to a 1.2 mL titer tube along with 1 &#956;L 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 &#730;C. Add 0.2 mL of flow buffer.</li>
		<li>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 &#956;L 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.</li>
		<li>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 &#730;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.</li>
		<li>Vortex all bead mixtures completely. Store SCCs at 4 &#730;C for up to 3 days. 
		NOTE: The SCC for fluorescent reporter mice is unstained cells.</li>
		<li>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).</li>
	</ol>
	</li>
	<li>Run samples on &#8220;Low&#8221; 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).</li>
	<li>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.</li>
	<li>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.</li>
	<li>Clean and shut down flow cytometer according to manufacturer or facility instructions.</li>
</ol>
6. Annotation of flow cytometry data
<ol>
	<li>Open a new worksheet using analysis software. Import FCS files from cytometer for all samples and single-color controls.</li>
	<li>Use SCCs to create a compensation matrix.</li>
	<li>Apply the compensation matrix to your stained samples and FMOs.</li>
	<li>Use the FMOs to determine positive staining to draw gates.</li>
</ol>

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The authors have nothing to disclose.

Multi-parameter flow cytometry allows for the quantitative analysis of multiple cell types in a complex tissue. In this report, we describe the flow cytometric detection of mononuclear phagocyte populations, including monocytes, dendritic cells, and macrophage subsets, after laser injury in the mouse eye. Liyanage et al recently reported a similar gating strategy to detect neutrophils, eosinophils, lymphocytes, DCs, macrophages, infiltrating macrophages, and monocytes27. Our gating strategy primar...

The innate immune system includes multiple cell types that stimulate complement activation and inflammation. Innate immune cells include natural killer (NK) cells, mast cells, basophils, eosinophils, neutrophils, and mononuclear phagocytes. Mononuclear phagocytes, which are composed of monocytes, macrophages, and dendritic cells, have been implicated in the pathophysiology of multiple ophthalmic conditions including uveitis, diabetic retinopathy, and age-related macular degeneration (AMD)1. In this protocol, we will focus on the identification of mononuclear phagocytes using multi-parameter flow cytometric analysis in a mouse model of neovascular AMD2. This protocol is adaptable for mouse models of diabetic retinopathy and/or uveitis, but more extensive ocular dissection is recommended because of the systemic nature of these diseases.
Mononuclear phagocytes express overlapping cell surface markers. Long-lasting tissue resident macrophages and microglia originate from the yolk sac-derived erythromyeloid progenitor3, while recycling macrophages and dendritic cells differentiate from the bone marrow-derived macrophage dendritic cell progenitor4. Mouse cell surface markers common to monocytes, macrophages, and dendritic cells include CD45, CD11b5, F4/806, Cx3cr17, and the intracellular marker Iba18. In order to overcome this challenge, transcriptomic analysis of macrophages, monocytes, and dendritic cells from multiple tissues defines CD64 as a macrophage-specific cell surface marker6. Macrophages have been described in the iris, choroid, ciliary body, and optic nerve in healthy eyes3. Alternatively, dendritic cell identification is more difficult; the most specific method of dendritic cell identification requires fate mapping using the Zbtb46-GFP reporter mouse9. Independent of this reporter line, expression of CD11c and MHCII in conjunction with the absence of CD64 can identify potential dendritic cells6,10. Dendritic cells have been identified in cornea, conjunctiva, iris, and choroid in normal eyes11. Microglia are specialized macrophages located in the retina, protected by the blood-retinal barrier, and derived from yolk sac progenitor cells12. As a result, retinal microglia can be differentiated from monocyte-derived macrophages by their dim levels of CD45 expression13 and high levels of Tmem119, which is available as a flow cytometry antibody14. Upon microglia activation, however, CD45 can be up-regulated15 and Tmem119 may be down-regulated3, demonstrating the complexity of microglia biology and this is likely relevant in both AMD and its mouse model. Finally, monocytes can be divided into at least two subtypes, including classical and non-classical. Classical monocytes display CCR2+Ly6ChighCX3CR1low expression, and non-classical monocytes demonstrate CCR2-Ly6ClowCX3CR1high markers5.
Due to the necessity of the quantitative analysis of marker expression, i.e., high versus low/dim levels, multi-parameter flow cytometry is the ideal method for discrimination between monocytes, macrophages, microglia, and dendritic cells in the eye and other tissues. Additional advantages include the identification of sub-populations, the ability to use fluorescence-activated cell sorting (FACS) to sort cell populations for transcriptomic or proteomic analysis, and fate mapping. The major disadvantage of multi-parameter flow cytometry is the lack of tissue architecture. This can be overcome by the ophthalmic dissection into the various ocular subcompartments: cornea, conjunctiva, iris, lens, retina, and choroid-sclera complex. Additionally, confirmatory immunofluorescence imaging can be performed, but is limited by the number of markers and lack of robust quantitation.
Genome wide association studies have linked multiple complement genes with AMD16. Complement activation leads to anaphylatoxin production, leukocyte recruitment, and resultant inflammation. In complement receptor deficient mice, laser injury reduces mononuclear phagocyte recruitment and laser-induced choroidal neovascularization (CNV) area17. Similarly, the C-C motif chemokine receptor 2 (CCR2) knockout mouse, which is deficient in monocyte recruitment to the tissue, demonstrates both decreased mononuclear phagocyte recruitment and laser-induced CNV area18. These data link complement and mononuclear phagocytes with experimental CNV and possibly neovascular AMD. In support of this association, complement receptors are dysregulated on peripheral blood monocytes in patients with neovascular AMD19,20. These data demonstrate a strong association between AMD and mononuclear phagocytes.
In this manuscript, we will use the experimental laser induced CNV model to characterize the mononuclear phagocyte populations in the mouse eye using multi-parameter flow cytometry. Laser-induced CNV is the standard mouse model of neovascular AMD, which demonstrated the efficacy of current first line neovascular AMD therapy21. This protocol will describe enucleation of mouse eyes, ocular dissection, digestion into a single cell suspension, antibody staining, determination of laser voltages using single color controls, and gating strategy using fluorescence minus one (FMO) controls. For a detailed description of the laser-induced CNV model, please see a previous publication22. Using this protocol, we will define microglia, monocytes, dendritic cells, and macrophage populations. Furthermore, we will use MHCII and CD11c to further define macrophage subsets within the laser induced CNV model.

<table>
	<tbody>
		<tr>
			<td>0.6 ml microcentrifuge tubes</td>
			<td>Fisher Scientific</td>
			<td>05-408-120</td>
			<td></td>
		</tr>
		<tr>
			<td>1.7 ml microcentrifuge tubes</td>
			<td>Costar</td>
			<td>3620</td>
			<td></td>
		</tr>
		<tr>
			<td>10 ml pipettes</td>
			<td>Fisher Scientific</td>
			<td>431031</td>
			<td></td>
		</tr>
		<tr>
			<td>15 ml conicals</td>
			<td>ThermoFisher</td>
			<td>339650</td>
			<td></td>
		</tr>
		<tr>
			<td>1x HBSS, 500 ml</td>
			<td>Gibco</td>
			<td>14025-092</td>
			<td></td>
		</tr>
		<tr>
			<td>2.5 ml syringe</td>
			<td>Henke Sass Wolf</td>
			<td>7886-1</td>
			<td></td>
		</tr>
		<tr>
			<td>20% paraformaldehyde solution</td>
			<td>Electron Microscopy Sciences</td>
			<td>15713-S</td>
			<td></td>
		</tr>
		<tr>
			<td>25 ml pipettes</td>
			<td>Fisher Scientific</td>
			<td>431032</td>
			<td></td>
		</tr>
		<tr>
			<td>30 G needle</td>
			<td>Exel</td>
			<td>26437</td>
			<td></td>
		</tr>
		<tr>
			<td>3ml luer lock syringe</td>
			<td>Fisher Scientific</td>
			<td>14955457</td>
			<td></td>
		</tr>
		<tr>
			<td>41 x 41 x 8 mm polystyrene weigh dish</td>
			<td>Fisher Scientific</td>
			<td>08-732-112</td>
			<td></td>
		</tr>
		<tr>
			<td>50 ml conicals</td>
			<td>Falcon</td>
			<td>352070</td>
			<td></td>
		</tr>
		<tr>
			<td>96-well, U-bottom assay plate without lid</td>
			<td>Falcon</td>
			<td>353910</td>
			<td></td>
		</tr>
		<tr>
			<td>Amine reactive beads</td>
			<td>ThermoFisher</td>
			<td>A10628</td>
			<td></td>
		</tr>
		<tr>
			<td>Analysis software</td>
			<td>FlowJo</td>
			<td>v 10</td>
			<td></td>
		</tr>
		<tr>
			<td>Anti-rat and anti-hamster Ig kappa bead set</td>
			<td>BD Biosciences</td>
			<td>552845</td>
			<td></td>
		</tr>
		<tr>
			<td>C57BL/6J mice</td>
			<td>Jackson Laboratory</td>
			<td>664</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell counter</td>
			<td>Invitrogen</td>
			<td>AMQAX1000</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell counter slides</td>
			<td>Invitrogen</td>
			<td>C10283</td>
			<td></td>
		</tr>
		<tr>
			<td>Compensation beads</td>
			<td>ThermoFisher</td>
			<td>01-1111-42</td>
			<td></td>
		</tr>
		<tr>
			<td>Count beads</td>
			<td>ThermoFisher</td>
			<td>01-1234-42</td>
			<td></td>
		</tr>
		<tr>
			<td>Curved forceps</td>
			<td>Fisher Scientific</td>
			<td>16-100-123</td>
			<td></td>
		</tr>
		<tr>
			<td>Digestion enzyme</td>
			<td>Sigma Aldrich</td>
			<td>5401020001</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissecting microscope</td>
			<td>National Optical and Scientific Instruments, Inc.</td>
			<td>DC3-420T</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissociation tubes</td>
			<td>Miltenyi Biotec</td>
			<td>130-096-334</td>
			<td></td>
		</tr>
		<tr>
			<td>DNase</td>
			<td>Roche</td>
			<td>10104159001</td>
			<td></td>
		</tr>
		<tr>
			<td>Electronic dissociator</td>
			<td>Miltenyi Biotec</td>
			<td>130-095-937</td>
			<td></td>
		</tr>
		<tr>
			<td>FACS Diva software</td>
			<td>BD Biosciences</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Fc block, rat anti-mouse CD16/CD32</td>
			<td>BD Biosciences</td>
			<td>553142</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine forceps</td>
			<td>Integra Miltex</td>
			<td>17035X</td>
			<td></td>
		</tr>
		<tr>
			<td>Flow buffer</td>
			<td>Miltenyi Biotec</td>
			<td>130-091-221</td>
			<td></td>
		</tr>
		<tr>
			<td>Flow cytometer</td>
			<td>BD Biosciences</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Flow tubes</td>
			<td>Falcon</td>
			<td>352008</td>
			<td></td>
		</tr>
		<tr>
			<td>Hamster anti-mouse CD11c BV421</td>
			<td>BD Biosciences</td>
			<td>562782</td>
			<td></td>
		</tr>
		<tr>
			<td>Ice bucket</td>
			<td>Fisher Scientific</td>
			<td>07-210-123</td>
			<td></td>
		</tr>
		<tr>
			<td>Live/Dead dye</td>
			<td>ThermoFisher</td>
			<td>65-0866-14</td>
			<td></td>
		</tr>
		<tr>
			<td>Lysing solution, 10x solution</td>
			<td>BD Biosciences</td>
			<td>555899</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro titer tube and rack</td>
			<td>Fisher Scientific</td>
			<td>02-681-380</td>
			<td></td>
		</tr>
		<tr>
			<td>Mouse anti-mouse CD64 PE</td>
			<td>BioLegend</td>
			<td>139304</td>
			<td></td>
		</tr>
		<tr>
			<td>Mouse anti-mouse NK1.1 PECF594</td>
			<td>BD Biosciences</td>
			<td>562864</td>
			<td></td>
		</tr>
		<tr>
			<td>P1000 pipet tips</td>
			<td>Denville Scientific</td>
			<td>P2404</td>
			<td></td>
		</tr>
		<tr>
			<td>P1000 pipettor</td>
			<td>Gilson</td>
			<td>FA10006M</td>
			<td></td>
		</tr>
		<tr>
			<td>P2 pipet tips</td>
			<td>eppendorf</td>
			<td>22491806</td>
			<td></td>
		</tr>
		<tr>
			<td>P2 pipettor</td>
			<td>Gilson</td>
			<td>FA10001M</td>
			<td></td>
		</tr>
		<tr>
			<td>P20 pipettor</td>
			<td>Gilson</td>
			<td>FA10003M</td>
			<td></td>
		</tr>
		<tr>
			<td>P200 pipet tips</td>
			<td>Denville Scientific</td>
			<td>P2401</td>
			<td></td>
		</tr>
		<tr>
			<td>P200 pipettor</td>
			<td>Gilson</td>
			<td>FA10005M</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipet man</td>
			<td>Fisher Scientific</td>
			<td>FB14955202</td>
			<td></td>
		</tr>
		<tr>
			<td>Rat anti-mouse B220 PECF594</td>
			<td>BD Biosciences</td>
			<td>562313</td>
			<td></td>
		</tr>
		<tr>
			<td>Rat anti-mouse CD11b APC-Cy7</td>
			<td>BD Biosciences</td>
			<td>557657</td>
			<td></td>
		</tr>
		<tr>
			<td>Rat anti-mouse CD19 AlexaFluor 700</td>
			<td>BD Biosciences</td>
			<td>557958</td>
			<td></td>
		</tr>
		<tr>
			<td>Rat anti-mouse CD19 PE</td>
			<td>BD Biosciences</td>
			<td>553786</td>
			<td></td>
		</tr>
		<tr>
			<td>Rat anti-mouse CD4 PECF594</td>
			<td>BD Biosciences</td>
			<td>562314</td>
			<td></td>
		</tr>
		<tr>
			<td>Rat anti-mouse CD45 FITC</td>
			<td>ThermoFisher</td>
			<td>11-0451-82</td>
			<td></td>
		</tr>
		<tr>
			<td>Rat anti-mouse CD45 PE-Cy7</td>
			<td>BD Biosciences</td>
			<td>552848</td>
			<td></td>
		</tr>
		<tr>
			<td>Rat anti-mouse CD8 PECF594</td>
			<td>BD Biosciences</td>
			<td>562315</td>
			<td></td>
		</tr>
		<tr>
			<td>Rat anti-mouse Ly6C</td>
			<td>BD Biosciences</td>
			<td>561085</td>
			<td></td>
		</tr>
		<tr>
			<td>Rat anti-mouse Ly6G PECF594</td>
			<td>BD Biosciences</td>
			<td>562700</td>
			<td></td>
		</tr>
		<tr>
			<td>Rat anti-mouse MHC II AlexaFluor 700</td>
			<td>BioLegend</td>
			<td>107622</td>
			<td></td>
		</tr>
		<tr>
			<td>Rat anti-mouse Siglec F PECF594</td>
			<td>BD Biosciences</td>
			<td>562757</td>
			<td></td>
		</tr>
		<tr>
			<td>Rat anti-mouse Tim4 AlexaFluor647</td>
			<td>BD Biosciences</td>
			<td>564178</td>
			<td></td>
		</tr>
		<tr>
			<td>Shaking incubator</td>
			<td>Labnet</td>
			<td>311DS</td>
			<td></td>
		</tr>
		<tr>
			<td>Spring scissors</td>
			<td>Fine Science Tools</td>
			<td>15024-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile cell strainer, 40 mm nylon mesh</td>
			<td>Fisher Scientific</td>
			<td>22363547</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile water, 500 ml</td>
			<td>Gibco</td>
			<td>A12873-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Swinging bucket centrifuge</td>
			<td>eppendorf</td>
			<td>5910 R</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypan blue, 0.4% solution</td>
			<td>Gibco</td>
			<td>15250061</td>
			<td></td>
		</tr>
	</tbody>
</table>

digestion of whole mouse eyes for multi-parameter flow cytometric analysis of mononuclear phagocytes

The innate immune system plays important roles in ocular pathophysiology including uveitis, diabetic retinopathy, and age-related macular degeneration. Innate immune cells, specifically mononuclear phagocytes, express overlapping cell surface markers, which makes identifying these populations a challenge. Multi-parameter flow cytometry allows for the simultaneous, quantitative analysis of multiple cell surface markers in order to differentiate monocytes, macrophages, microglia, and dendritic cells in mouse eyes. This protocol describes the enucleation of whole mouse eyes, ocular dissection, digestion into a single cell suspension, and staining of the single cell suspension for myeloid cell markers. Additionally, we explain the proper methods for determining voltages using single color controls and for delineating positive gates using fluorescence minus one controls. The major limitation of multi-parameter flow cytometry is the absence of tissue architecture. This limitation can be overcome by multi-parameter flow cytometry of individual ocular compartments or complimentary immunofluorescence staining. However, immunofluorescence is limited by its lack of quantitative analysis and reduced number of fluorophores on most microscopes. We describe the use of multi-parametric flow cytometry to provide highly quantitative analysis of mononuclear phagocytes in laser-induced choroidal neovascularization. Additionally, multi-parameter flow cytometry can be used for the identification of macrophage subsets, fate mapping, and cell sorting for transcriptomic or proteomic studies.

Figure 1 showed uncompensated frequency histograms for the SCCs and cells for the red laser: Alexa647, Alexa700, and APC-Cy7. In Figure 1A, the magenta line depicted the peak of the SCC for Alexa647, while the cyan line showed&#160;the intended 0.5 Log difference. Notice that the peak in Alexa700 and APC-Cy7 was to the left (less bright) of the cyan line. Also note that the cells were all to the left of the magenta line. Any cells to the right of the SCC peak we...

Watch this Scientific Journal Video about Digestion of Whole Mouse Eyes for Multi-Parameter Flow Cytometric Analysis of Mononuclear Phagocytes at JoVE.com

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

This protocol provides a method to digest whole eyes into a single cell suspension for the purpose of multi-parameter flow cytometric analysis in order to identify specific ocular mononuclear phagocytic populations, including monocytes, microglia, macrophages, and dendritic cells.

The innate immune system plays important roles in ocular pathophysiology including uveitis, diabetic retinopathy, and age-related macular degeneration. ...

digestion-whole-mouse-eyes-for-multi-parameter-flow-cytometric

Research

JoVE Journal

Immunology and Infection

4.8K Views.  Northwestern University. This protocol provides a method to digest whole eyes into a single cell suspension for the purpose of multi-parameter flow cytometric analysis in order to identify specific ocular mononuclear phagocytic populations, including monocytes, microglia, macrophages, and dendritic cells.

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

Whole-animal Imaging and Flow Cytometric Techniques for Analysis of Antigen-specific CD8+ T Cell Responses after Nanoparticle Vaccination

Traditional vaccine adjuvants, such as alum, elicit suboptimal CD8+ T cell responses. To address this major challenge in vaccine development, various nanoparticle systems have been engineered to mimic features of pathogens to improve antigen delivery to draining lymph nodes and increase antigen uptake by antigen-presenting cells, leading to new vaccine formulations optimized for induction of antigen-specific CD8+ T cell responses. In this article, we describe the synthesis of a &#8220;pathogen-mimicking&#8221; nanoparticle system, termed interbilayer-crosslinked multilamellar vesicles (ICMVs) that can serve as an effective vaccine carrier for co-delivery of subunit antigens and immunostimulatory agents and elicitation of potent cytotoxic CD8+ T lymphocyte (CTL) responses. We describe methods for characterizing hydrodynamic size and surface charge of vaccine nanoparticles with dynamic light scattering and zeta potential analyzer and present a confocal microscopy-based procedure to analyze nanoparticle-mediated antigen delivery to draining lymph nodes. Furthermore, we show a new bioluminescence whole-animal imaging technique utilizing adoptive transfer of luciferase-expressing, antigen-specific CD8+ T cells into recipient mice, followed by nanoparticle vaccination, which permits non-invasive interrogation of expansion and trafficking patterns of CTLs in real time. We also describe tetramer staining and flow cytometric analysis of peripheral blood mononuclear cells for longitudinal quantification of endogenous T cell responses in mice vaccinated with nanoparticles.

We describe whole-animal imaging and flow cytometry-based techniques for monitoring expansion of antigen-specific CD8+ T cells in response to immunization with nanoparticles in a murine model of vaccination.

Traditional vaccine adjuvants, such as alum, elicit suboptimal CD8+ T cell responses. To address this major challenge in vaccine development, various ...

Isolation and Flow Cytometric Analysis of Immune Cells from the Ischemic Mouse Brain

Ischemic stroke initiates a robust inflammatory response that starts in the intravascular compartment and involves rapid activation of brain resident cells. A key mechanism of this inflammatory response is the migration of circulating immune cells to the ischemic brain facilitated by chemokine release and increased endothelial adhesion molecule expression. Brain-invading leukocytes are well-known contributing to early-stage secondary ischemic injury, but their significance for the termination of inflammation and later brain repair has only recently been noticed.
Here, a simple protocol for the efficient isolation of immune cells from the ischemic mouse brain is provided. After transcardial perfusion, brain hemispheres are dissected and mechanically dissociated. Enzymatic digestion with Liberase&#160;is followed by density gradient (such as Percoll) centrifugation to remove myelin and cell debris. One major advantage of this protocol is the single-layer density gradient procedure which does not require time-consuming preparation of gradients and can be reliably performed. The approach yields highly reproducible cell counts per brain hemisphere and allows for measuring several flow cytometry panels in one biological replicate. Phenotypic characterization and quantification of brain-invading leukocytes after experimental stroke may contribute to a better understanding of their multifaceted roles in ischemic injury and repair.

Inflammation plays a central role in the pathogenesis of ischemic stroke. Increasing evidence suggests that it acts as a double-edged sword which exacerbates early brain injury, but also contributes to later repair. This protocol describes the isolation of immune cells from the ischemic brain and their subsequent flow cytometric phenotyping.

Ischemic stroke initiates a robust inflammatory response that starts in the intravascular compartment and involves rapid activation of brain resident ...

Isolation and Flow Cytometric Analysis of Glioma-infiltrating Peripheral Blood Mononuclear Cells

Our laboratory has recently demonstrated that natural killer (NK) cells are capable of eradicating orthotopically implanted mouse GL26 and rat CNS-1 malignant gliomas soon after intracranial engraftment if the cancer cells are rendered deficient in their expression of the &#946;-galactoside-binding lectin galectin-1 (gal-1). More recent work now shows that a population of Gr-1+/CD11b+ myeloid cells is critical to this effect. To better understand the mechanisms by which NK and myeloid cells cooperate to confer gal-1-deficient tumor rejection we have developed a comprehensive protocol for the isolation and analysis of glioma-infiltrating peripheral blood mononuclear cells (PBMC). The method is demonstrated here by comparing PBMC infiltration into the tumor microenvironment of gal-1-expressing GL26 gliomas with those rendered gal-1-deficient via shRNA knockdown. The protocol begins with a description of&#160;how to culture and prepare GL26 cells for inoculation into the syngeneic C57BL/6J mouse brain. It then explains the steps involved in the isolation and flow cytometric analysis of glioma-infiltrating PBMCs from the early brain tumor microenvironment. The method is adaptable to a number of in vivo experimental designs in which temporal data on immune infiltration into the brain is required. The method is sensitive and highly reproducible, as glioma-infiltrating PBMCs can be isolated from intracranial tumors as soon as 24 hr post-tumor engraftment with similar cell counts observed from time point matched tumors throughout independent experiments. A single experimentalist can perform the method from brain harvesting to flow cytometric analysis of glioma-infiltrating PBMCs in roughly 4-6 hr depending on the number of samples to be analyzed. Alternative glioma models and/or cell-specific detection antibodies may also be used at the experimentalists&#8217; discretion to assess the infiltration of several other immune cell types of interest without the need for alterations to the overall procedure.

Presented here is a straightforward method for the isolation and flow cytometric analysis of glioma-infiltrating peripheral blood mononuclear cells that yields time-dependent quantitative data on the number and activation status of immune cells entering the early brain tumor microenvironment.

Our laboratory has recently demonstrated that natural killer (NK) cells are capable of eradicating orthotopically implanted mouse GL26 and rat CNS-1 ...

A Simple Flow Cytometric Method to Measure Glucose Uptake and Glucose Transporter Expression for Monocyte Subpopulations in Whole Blood

Monocytes are innate immune cells that can be activated by pathogens and inflammation associated with certain chronic inflammatory diseases. Activation of monocytes induces effector functions and a concomitant shift from oxidative to glycolytic metabolism that is accompanied by increased glucose transporter expression. This increased glycolytic metabolism is also observed for trained immunity of monocytes, a form of innate immunological memory. Although in vitro protocols examining glucose transporter expression and glucose uptake by monocytes have been described, none have been examined by multi-parametric flow cytometry in whole blood. We describe a multi-parametric flow cytometric protocol for the measurement of fluorescent glucose analog 2-NBDG uptake in whole blood by total monocytes and the classical (CD14++CD16-), intermediate (CD14++CD16+) and non-classical (CD14+CD16++) monocyte subpopulations. This method can be used to examine glucose transporter expression and glucose uptake for total monocytes and monocyte subpopulations during homeostasis and inflammatory disease, and can be easily modified to examine glucose uptake for other leukocytes and leukocyte subpopulations within blood.

Monocytes are integral components of the human innate immune system that rely on glycolytic metabolism when activated. We describe a flow cytometry protocol to measure glucose transporter expression and glucose uptake by total monocytes and monocyte subpopulations in fresh whole blood.

Monocytes are innate immune cells that can be activated by pathogens and inflammation associated with certain chronic inflammatory diseases. Activation of ...

Flow Cytometric Analysis of Particle-bound Bet v 1 Allergen in PM10

Flow cytometry is a method widely used to quantify suspended solids such as cells or bacteria in a size range from 0.5 to several tens of micrometers in diameter. In addition to a characterization of forward and sideward scatter properties, it enables the use of fluorescent labeled markers like antibodies to detect respective structures. Using indirect antibody staining, flow cytometry is employed here to quantify birch pollen allergen (precisely Bet v 1)-loaded particles of 0.5 to 10 &#181;m in diameter in inhalable particulate matter (PM10, particle size &#8804;10 &#181;m in diameter). PM10 particles may act as carriers of adsorbed allergens possibly transporting them to the lower respiratory tract, where they could trigger allergic reactions.
So far the allergen content of PM10 has been studied by means of enzyme linked immunosorbent assays (ELISAs) and scanning electron microscopy. ELISA measures the dissolved and not the particle-bound allergen. Compared to scanning electron microscopy, which can visualize allergen-loaded particles, flow cytometry may additionally quantify them. As allergen content of ambient air can deviate from birch pollen count, allergic symptoms might perhaps correlate better with allergen exposure than with pollen count. In conjunction with clinical data, the presented method offers the opportunity to test in future experiments whether allergic reactions to birch pollen antigens are associated with the Bet v 1 allergen content of PM10 particles &#62;0.5 &#181;m.

Here, we present a protocol to quantify allergen-loaded particles by flow cytometry. Ambient particulate matter particles may act as carriers of adsorbed allergens. We show here that flow cytometry, a method widely used to characterize suspended solids &#62;0.5 &#181;m in diameter, can be used to measure these allergen-loaded particles.

Flow cytometry is a method widely used to quantify suspended solids such as cells or bacteria in a size range from 0.5 to several tens of micrometers in ...

Flow Cytometric Analysis of Natural Killer Cell Lytic Activity in Human Whole Blood

NK cell cytotoxicity is a widely used measure to determine the effect of outside intervention on NK cell function. However, the accuracy and reproducibility of this assay can be considered unstable, either because of user&#39;s errors or because of the sensitivity of NK cells to experimental manipulation. To eliminate these issues, a workflow that reduces them to a minimum was established and is presented here. To illustrate, we obtained blood samples, at various time points, from runners (n = 4) that were submitted to an intense bout of exercise. First, NK cells are simultaneously identified and isolated through CD56 tagging and magnetic-based sorting, directly from whole blood and from as little as one milliliter. The sorted NK cells are removed of any reagent or capping antibodies. They can be counted in order to establish an accurate NK cell count per milliliter of blood. Secondly, the sorted NK cells (effectors cells or E) can be mixed with 3,3&#39;-Diotadecyloxacarbocyanine Perchlorate (DiO) tagged K562 cells (target cells or T) at an assay-optimal 1:5 T:E ratio, and analyzed using an imaging flow-cytometer that allows for the visualization of each event and the elimination of any false positive or false negatives (such as doublets or effector cells). This workflow can be completed in about 4 h, and allows for very stable results even when working with human samples. When available, research teams can test several experimental interventions in human subjects, and compare measurements across several time points without compromising the data&#39;s integrity.

This work describes an advanced workflow for the accurate and fast determination of NK (Natural Killer) cell count and NK cell cytotoxicity in human blood samples.

NK cell cytotoxicity is a widely used measure to determine the effect of outside intervention on NK cell function. However, the accuracy and ...

Isolation and Flow Cytometric Analysis of Human Endocervical Gamma Delta T Cells

The female reproductive tract (FRT) mucosal immune system serves as the first line of defense. Better knowledge of the genital mucosa is therefore essential for understanding pathogenicity of different pathogens including HIV. Gamma delta (GD) T cells are the prototype of &#39;unconventional&#39; T cells and represent a relatively small subset of T cells defined by their expression of heterodimeric T-cell receptors (TCRs) composed of gamma and delta chains. This sets them apart from the classical and much better known CD4+ helper T cells and CD8+ cytotoxic T cells that are defined by alpha-beta TCRs. GD T cells often show tissue-specific localization and are enriched in epithelium. GD T cells orchestrate immune responses in inflammation, tumor surveillance, infectious disease, and autoimmunity.
Here, we present a method to reproducibly isolate and analyze human endocervical intraepithelial GD T lymphocytes. We have used endocervical cytobrush samples from women participating in the Women&#39;s Interagency HIV Infection Study (WIHS). Knowledge about GD T cells interactions during conditions in which there is an insult to the vaginal mucosal could be applied to any clinical study in which mucosal vulnerability is addressed, including the development of vaginal microbicides.In addition, knowledge about mucosal GD T cell responses has potential for application of GD T cell-based immune therapy in treating infectious diseases.

We describe here an isolation method to obtain human endocervical intraepithelial lymphocytes for the analysis of intraepithelial gamma delta T cells. This protocol can be extended for the purification of endocervical gamma delta T cells by magnetic beads or by cell sorting.

The female reproductive tract (FRT) mucosal immune system serves as the first line of defense. Better knowledge of the genital mucosa is therefore ...

Characterization of Human Monocyte Subsets by Whole Blood Flow Cytometry Analysis

Monocytes are key contributors in various inflammatory disorders and alterations to these cells, including their subset proportions and functions, can have pathological significance. An ideal method for examining alterations to monocytes is whole blood flow cytometry as the minimal handling of samples by this method limits artifactual cell activation. However, many different approaches are taken to gate the monocyte subsets leading to inconsistent identification of the subsets between studies. Here we demonstrate a method using whole blood flow cytometry to identify and characterize human monocyte subsets (classical, intermediate, and non-classical). We outline how to prepare the blood samples for flow cytometry, gate the subsets (ensure contaminating cells have been removed), and determine monocyte subset expression of surface markers &#8212; in this example M1 and M2 markers. This protocol can be extended to other studies that require a standard gating method for assessing monocyte subset proportions and monocyte subset expression of other functional markers.

Here we present a protocol for characterizing monocyte subsets by whole blood flow cytometry. This includes outlining how to gate the subsets and assess their expression of surface markers and giving an example of the assessment of the expression of M1 (inflammatory) and M2 markers (anti-inflammatory).

Monocytes are key contributors in various inflammatory disorders and alterations to these cells, including their subset proportions and functions, can ...

Digestion of the Murine Liver for a Flow Cytometric Analysis of Lymphatic Endothelial Cells

Within the liver, lymphatic vessels are found within the portal triad, and their described function is to remove interstitial fluid from the liver to the lymph nodes where cellular debris and antigens can be surveyed. We are very interested in understanding how the lymphatic vasculature might be involved in inflammation and immune cell function within the liver. However, very little has been published establishing digestion protocols for the isolation of lymphatic endothelial cells (LECs) from the liver or specific markers that can be used to evaluate liver LECs on a per cell basis. Therefore, we optimized a method for the digestion and staining of the liver in order to evaluate the LEC population in the liver. We are confident that the method outlined here will be useful for the identification and isolation of LECs from the liver and will strengthen our understanding of how LECs respond to the liver microenvironment.

The goal of this protocol is to identify lymphatic endothelial cell populations within the liver using described markers. We utilize collagenase IV and DNase and a gentle mincing of tissue, combined with flow cytometry, to identify a distinct population of lymphatic endothelial cells.

Within the liver, lymphatic vessels are found within the portal triad, and their described function is to remove interstitial fluid from the liver to the ...

Preparation of Whole Bone Marrow for Mass Cytometry Analysis of Neutrophil-lineage Cells

In this article, we present a protocol that is optimized to preserve neutrophil-lineage cells in fresh BM for whole BM CyTOF analysis. We utilized a myeloid-biased 39-antibody CyTOF panel to evaluate the hematopoietic system with a focus on the neutrophil-lineage cells by using this protocol. The CyTOF result was analyzed with an open-resource dimensional reduction algorithm, viSNE, and the data was presented to demonstrate the outcome of this protocol. We have discovered new neutrophil-lineage cell populations based on this protocol. This protocol of fresh whole BM preparation may be used for 1), CyTOF analysis to discover unidentified cell populations from whole BM, 2), investigating whole BM defects for patients with blood disorders such as leukemia, 3), assisting optimization of fluorescence-activated flow cytometry protocols that utilize fresh whole BM.

Here, we present a protocol to process fresh bone marrow (BM) isolated from mouse or human for high-dimensional mass cytometry (Cytometry by Time-Of-Flight, CyTOF) analysis of neutrophil-lineage cells.

In this article, we present a protocol that is optimized to preserve neutrophil-lineage cells in fresh BM for whole BM CyTOF analysis. We utilized a ...