Intracellular Staining and Flow Cytometry to Identify Lymphocyte Subsets within Murine Aorta, Kidney and Lymph Nodes in a Model of Hypertension

Vanderbilt University's Institutional Animal Care and Use Committee has approved the procedures described herein. Mice are housed and cared for in accordance with the Guide for the Care and Use of Laboratory Animals (National Academies Press. Revised 2010).

1. Isolation of the Aortic Draining Lymph Nodes, Kidney and Aorta from Mice

1. Euthanize the mice by CO₂ inhalation. Spray the chest with 70% ethanol and carefully open the skin and the chest wall with scissors to expose the heart.
2. To perfuse the vasculature, perform a small incision in the right atrium and steadily inject at least 10 ml of cold PBS (approximately 1 ml/sec) into the apex of the left ventricle using a 21 or 23 G needle. Perfuse until all organs have blanched. The heart blanches first. Blanching of the liver indicates that the perfusion has been well performed.
3. Using small forceps and fine scissors, gently cut away and pull out the intestines, stomach, spleen, pancreas and liver to better visualize the aorta.
   NOTE: This step has to be done very precisely as damage to the gastrointestinal tract can induce contamination.
4. Cut out and remove each lung with scissors. Rinse the chest cavity with phosphate buffered saline (PBS) using a needleless syringe. Remove excess blood and fluid using sterile gauze.
5. Remove the abdominal aortic draining lymph nodes using fine dissecting forceps. Make sure not to rupture the capsule. Remove the remaining fat on the surface of each lymph node with the dissecting forceps. Cut out the two kidneys with fine scissors.
6. Dissect the entire aorta (thoracic and abdominal aorta) with fine, curved scissors. Start at the level of the heart and carefully dissect away from the esophagus and the vertebrae down to the level of the iliac bifurcation making sure to keep the surrounding perivascular fat attached to the aorta.
   NOTE: Further dissection of the aorta to remove surrounding structures can be done under a microscope. Specifically, remove arterial branches (carotid arteries, subclavian arteries, celiac, mesenteric, and renal arteries) and lymph nodes. Do keep a consistent layer of perivascular fat around each aorta since this is a site where many inflammatory cells reside in the setting of hypertension. Place each tissue in separate tubes containing cold PBS.

2. Generation of Single Cell Suspensions from Each Tissue

1. The kidneys
   NOTE: The kidney can be dissociated into a single cell suspension by combining mechanical dissociation plus enzymatic digestion.
   1. Prepare the kidney digestion solution by adding collagenase D (2 mg/ml) and DNase I (100 µg/ml) to RPMI 1640 medium (supplemented with 5% fetal bovine serum (FBS)). Prepare 10 ml per kidney sample.
   2. Use forceps to transfer the two kidneys into a dissociation tube containing 10 ml of kidney digestion solution.
   3. Perform the mechanical dissociation using a semi-automated dissociator device according to the manufacturer's instructions.
   4. After mechanical dissociation, detach the tube from the device and perform the enzymatic digestion. Incubate the samples for 20 min at 37 ºC under continuous rotation. Immediately stop the enzymatic digestion by adding cold RPMI 1640 medium supplemented with 5% FBS.
   5. Transfer the solution by pipetting into a 50 ml conical tube after filtering through a 40 µm strainer. Tease the remaining tissue apart by pressing with the plunger of a 1 ml syringe. Pellet the cells (500 x g, 8 min, 4 ºC).
   6. Resuspend the pellet into 3 ml of 36% density gradient centrifugation media. Transfer the suspension into a 15 ml conical tube. Gently add 3 ml of 72% density gradient centrifugation media beneath the 3 ml 36% cell suspension without any disruption. Centrifuge (1,000 x g without brake, 20 min, 4 °C).
   7. The immune cells will be located at the interface. Remove the topmost yellowish layer containing cellular debris and then bring the volume up to 15 ml using cold PBS. Shake well to breakdown the gradient. Spin the cells (300 x g, 8 min, 4 °C) and resuspend the pellet in 3 ml of RPMI with 5% FBS. Count the number of live cells on a hemocytometer using Trypan blue exclusion.
2. The aorta
   1. Prepare the aorta digestion solution by adding collagenase A (1 mg/ml), collagenase B (1 mg/ml), and DNase I (100 µg/ml) to RPMI 1640 medium (supplemented with 5% FBS).
   2. Using fine forceps, transfer the aorta into a 2 ml tube containing 1 ml of aorta digestion solution. Mince the whole aorta into very small pieces with fine scissors in the 1 ml digestion solution. Keep on ice. Incubate the samples for 30 min at 37 °C under continuous rotation.
   3. Transfer the solution to a tissue culture dish and stop the enzymatic digestion by adding 5 times the volume of cold RPMI with 5% FBS.
   4. Transfer the solution by pipetting into a 50 ml conical tube after filtering through a 40 µm strainer. Tease the remaining tissue apart into a single cell suspension by pressing with the plunger of a 1 ml syringe. Flush the strainer and pellet the cells (300 x g, 8 min, 4 ºC).
   5. Wash the pellet by adding 10 ml of RPMI with 5% FBS. Spin the cells (300 x g, 8 min, 4 °C) and resuspend the pellet in 3 ml of RPMI with 5% FBS. Count the number of live cells on a hemocytometer using Trypan blue exclusion.
3. The aortic draining lymph nodes
   1. Place a 40 µm strainer in a tissue culture dish (60 mm x 15 mm). Place the lymph nodes on the strainer and tease them apart into a single cell suspension by pressing with the plunger of a 1 ml syringe. Flush the strainer with 2 ml of RPMI with 5% FBS. Collect the cells in the dish.
   2. Transfer the cells into a 15 ml falcon tube and pellet the cells (300 x g, 8 min, 4 ºC). Resuspend the pellet in 500 µl of RPMI with 5% FBS. Count the number of live cells on a hemocytometer using Trypan blue exclusion.

3. Ex Vivo Stimulation for Cytokine Detection by Flow Cytometry

1. Prepare the cell culture media: RPMI with 5% FBS, 0.1 mM of non-essential amino acids, 50 µM β-mercaptoethanol and 1% penicillin/streptomycin.
2. Centrifuge each cell suspension obtained from the kidney, aorta and lymph nodes (300 x g, 8 min, 4 ºC). Resuspend each pellet with the cell culture media at a concentration of 1 x 10⁶ cells per ml.
3. Stimulate cells by adding 2 µl of the cell activation cocktail (containing Phorbol 12-Myristate 13-Acetate (PMA), the calcium ionophore ionomycin, and the Golgi inhibitor Brefeldin A) for every 1 ml of cell culture.
   NOTE: The final concentrations of each component are: 81 nM of PMA, 1.33 µM of ionomycin and 5 µg/ml of Brefeldin A. Treatment with PMA and ionomycin activates cells to produce cytokines. Cytokines are secreted proteins that must be trapped in the cells to be detected. Brefeldin A causes the accumulation of normally secreted proteins in the endoplasmic reticulum and Golgi apparatus. Thus, the technique described here enhances the detectability of cytokine-producing lymphocytes by flow cytometry.
4. Plate 1 to 2 million cells in a 12-well (aorta or kidney) or 24-well (lymph node) low-adherence plates. Place the plate in a 37 °C humidified CO₂ incubator for 3 hr.
   NOTE: The number of plated cells may be lower in lymph nodes from control mice. Also, the time of stimulation has to be empirically determined for each cell population and cytokine studied. In the case of T cells isolated from aorta, kidney or lymph nodes, we recommend stimulating for 3 to 4 hr. A longer stimulation will not increase the cytokine secretion and will induce a further downregulation of several T cell surface markers like CD3, CD4 and CD8. (Note that even a 3-4 hr stimulation will induce some downregulation of these markers.)

4. Surface Staining

1. Transfer the cells into a polystyrene FACS tube and centrifuge (300 x g, 8 min, 4 °C). Wash the cells twice with FACS buffer (Ca²⁺ and Mg⁺ free PBS containing 4% FBS and 2mM EDTA) and divide the cell suspension equally into different FACS tubes based on the number of antibody panels desired and the number of control tubes needed (see below).
2. To perform compensation for each panel, prepare unstained control tubes and single stained tubes for each tissue type and antibody panel. In addition, prepare fluorescence minus one (FMO) control tubes for each antibody panel by adding all but one of the antibodies.
3. Adjust the volume of each tube to 2 ml with FACS Buffer. Centrifuge the cell suspension (300 x g, 8 min, 4 °C), remove the supernatant and block Fc receptors by incubating the cells with 0.5 µg of anti-CD16/CD32 antibodies per million cells for 10 min on ice.
   NOTE: CD16 is low affinity IgG Fc receptor III (FcR III) and CD32 is FcR II. CD16/CD32 are expressed on granulocytes, mast cells, monocytes/macrophages, natural killer cells, B cells, dendritic cells and some activated mature T cells.
4. Perform viability staining: Wash the cells with 1x PBS, centrifuge (300 x g, 8 min, 4 °C) and resuspend in 1,000 µl of 1x PBS. Add 1 µl of a cell-impermeant amine-reactive dye (viability stain). Incubate for 15 min at 4 °C protected from light.
5. During the incubation, prepare the cocktail of antibodies (for surface staining) in an appropriate volume of FACS buffer. Wash the cells twice with 1-2 ml of FACS buffer, centrifuge (300 x g, 8 min, 4 °C) and resuspend each pellet with 100 µl of the antibody mixture. Incubate for 30 min at 4 °C protected from light.
   NOTE: For each flow cytometry antibody, it is useful to determine the optimal amount of antibody needed by performing a dose titration curve.
6. Wash the cells twice with 2 ml of FACS buffer and pellet the cells (300 x g, 8 min, 4 °C).

5. Fixation, Permeabilization and Intracellular Staining

1. Perform the intracellular staining with a fixation and permeabilization kit containing appropriate buffers by following the manufacturer's instructions. Fix and permeabilize the cells by resuspending them in the appropriate buffer. Incubate samples for 40 min at 4 °C protected from light.
2. Add 1 ml of 1x permeabilization and wash buffer directly to the fixed and permeabilized cells. Centrifuge (500 x g, 8 min, 4 °C) and remove the supernatant. Resuspend the pellet with 2 ml of 1x wash buffer.
3. Prepare the mixtures of antibodies (for intracellular staining only) in an appropriate volume of 1x wash buffer (usually 100 µl per tube).
   NOTE: As described above, the optimal antibody concentration needs to be determined for each antibody in the panel. For example, for antibodies to IL-17A or IL-17F, we use a dilution factor of 1:30, but for antibodies to IFNγ and T-bet, we use a dilution factor of 1:100.
4. Centrifuge the cells (500 x g, 8 min, 4 °C) and resuspend each pellet with 100 µl of the antibody mixture. Incubate for 40 min at 4 °C protected from light.
5. Wash the cells twice with 2 ml of 1x wash buffer and pellet the cells (500 x g, 8 min, 4 °C). Resuspend the pellet with 300 µl of 1x wash buffer and analyze on a flow cytometer with appropriate filters.
6. For quantification of the total number of cells per tissue sample, use counting beads. Prior to acquisition, add the recommended volume/number of counting beads to each tube.

6. Compensation, Gating, Normalization, and Tips

1. Compensation and gating
   1. Run unstained and single stained control tubes for compensation to adjust for spectral overlap.
      NOTE: The use of compensation beads rather than cell-based compensation controls is necessary in cases where the antigen of interest or cell population in a sample is present at such low levels that compensation using cells will be difficult. Also of note, when using tandem dyes, it is advisable to prepare compensations for each experiment since tandem dyes vary lot-to-lot and with each day.
   2. Run fluorescence minus one controls (FMOs) for all the fluorophores in the panel when starting a new multicolor experiment.
      NOTE: The FMO controls are samples that contain all the antibodies in the panel except for one. These controls permit accurate discrimination of positive versus negative signals for the omitted antibody to properly adjust the gates. These controls are particularly important when the expression levels are low or when the separation of the target population is poor (for example, when the target is a cytokine or transcription factor). Although not always necessary, in some cases, isotype controls may be preferred in place of FMO controls as they provide proper compensation for the antibody isotype and fluorophore conjugate without the specificity of the antigen.
   3. Set up the primary gate: adjust Forward Scatter Area (FSC-A) and Side Scatter Area (SSC-A) voltage in order to detect the leukocyte population.
   4. Exclude the dead cells.
      NOTE: The viability stain reacts with free amines present on both the surface and the interior of cells. In contrast to live cells, dead cells (with compromised membranes) allow the dye to enter into the cytoplasm, increasing the amount of protein labeling. Thus, dead cells will be brighter than live cells allowing easy discrimination. Gate on the live cells.
      NOTE: The viability of the cell suspensions from the aorta and the kidney may be lower than the viability of the cell suspensions from the lymph nodes due to the additional digestion step.
   5. Plot Forward Scatter Area (FSC-A) [y-axis] and Forward Scatter Height (FSC-H) [x-axis]. Singlets appear as a diagonal on this dot plot. Gate on this diagonal to exclude doublets. Then plot SSC-A [y-axis] and CD45 [x-axis].
   6. The CD45⁺ leukocyte population can be easily identified. Gate on this population. Subsequent populations can be identified from this population.
2. Normalization of cell counts
   1. Counting beads have characteristics that result in low FSC and high SSC. For example, if 50,000 beads are added into 300 µl of the single cell suspension, the equation for the calculation of the absolute number of any give cell type per tube is as follows:
           Number of cells per tube = (50,000 / number of beads counted by the flow cytometer) x number of cells counted.
      Using this calculation, the absolute number of any given cell type per tube can be determined. Based on how many tubes were generated from each organ, the number of cells per organ can be calculated.
      NOTE: Keep all cells and reagents at 4 °C during the protocol. Avoid vigorous vortexing because it can damage cells. Use minimal forces to pellet the cells. Avoid making bubbles in the FACS tube since it can precipitate cell death. Don't let the pellets dry out at any time.