Real-Time Monitoring of Energy Metabolism in Human NK Cells Using an Extracellular Flux Analyzer

All procedures involving sample collection have been performed in accordance with the institute's IRB guidelines.

1. Reagent preparation

1. Reagents for the isolation of natural killer (NK) cells
   NOTE: Prepare these reagents in a cell culture hood.
   1. Prepare NK separation buffer: Supplement phosphate-buffered saline, PBS (pH 7.4) with 1 mM ethylenediaminetetraacetic acid (EDTA) and 2% fetal calf serum (FCS) that has previously been heat-inactivated (at 56 °C for 30 min).
   2. Prepare NK culture medium: Supplement Iscove's Modified Dulbecco's Media (IMDM) with 10% human serum (HS). Sterile filter and store at 2-8 °C. Warm the medium to 37 °C before adding to the cells.
   3. Resuspend Human interleukin, IL-15 in PBS at 200 µg/mL.
2. Reagents for extracellular flux assay
   1. Prepare assay media: For mitochondrial stress test medium, add 1 mM sodium pyruvate, 2 mM L-glutamine, and 10 mM glucose to the base medium; for glycolysis stress test medium, add 2 mM L-glutamine to the base medium.      
      NOTE: The concentrations of glucose, pyruvate, and glutamine are provided as recommended by the extracellular flux analyzer manufacturer. However, media composition can be altered by researchers to perfectly match that of the culture medium, if desired.
   2. Adjust the pH of both media to 7.4 with 0.1 N sodium hydroxide (NaOH) using a benchtop pH meter, sterile filter through a 0.2 µM pore size, and store at 2-8 °C. Warm media to 37 °C, check pH, and readjust to 7.4 before use if required.   
      NOTE: Only ambient carbon dioxide (CO₂) is contained in the atmosphere of the extracellular flux analyzer, the use of media without bicarbonate is critical.
   3. Prepare stock solutions for reagents: Oligomycin (ATP synthase inhibitor), 10 mM stock solution in Dimethyl sulfoxide (DMSO); 2,4-dinitrophenol (DNP, uncoupler), 1 M stock solution in DMSO; antimycin A (complex III inhibitor), 10 mM stock solution in DMSO; rotenone (complex I inhibitor), 10 mM stock solution in DMSO. Make 30 µL aliquots of all reagents and store them at -20 °C.         
      NOTE: These guidelines are intended for reagents purchased individually and prepared by the researcher. If reagents are purchased from the analyzer manufacturer instead, follow their guidelines for reagent preparation.

2. NK cells isolation from peripheral blood

1. Peripheral blood mononuclear cells (PBMCs) preparation from human blood
   NOTE: Perform these steps in a cell culture hood. Decontaminate all residues and material in contact with blood with bleach and discard them into the appropriate container to be incinerated.
   1. Pipette 20 mL of lymphocyte separation medium (LSM) into a 50 mL conical tube.
   2. Carefully, while keeping the tube at a 30˚ angle, pipette 20 mL of blood over the LSM, very gently and touch the wall of the tube. Avoid the mixing of blood with the LSM and create a visible and well-defined interphase between the two fluids.    
      NOTE: Peripheral blood or enriched leukapheresis products can be used.
   3. Centrifuge the tubes for 25 min, 1000 x g at room temperature. Do not use a brake, as it could make both phases in the tube (LSM and blood) mix.
   4. Carefully take out the tubes from the centrifuge and place them in a rack. Check for the presence of a conspicuous layer of cells (mononuclear cells) that will form at the interphase between LSM (clear) and plasma (yellow), while red blood cells pellet at the bottom of the tube.
   5. Gently aspirate the mononuclear cell layer with a 10 mL plastic pipette (around 5-8 mL) and place it in a new 50 mL conical tube. The lymphocyte interphase of up to 2 different tubes can be pooled together.
   6. Wash the mononuclear cells 2x by resuspending in 45 mL PBS and centrifuging at 800 x g for 5 min, at room temperature.
      NOTE: After this step, a pellet of peripheral blood mononuclear cells (PBMCs) is obtained and the researcher can proceed to the NK isolation step.
2. NK isolation from PBMCs
   1. Count the cells from 2.1.6 and resuspend them in NK Separation Buffer (1x 10⁸ PBMCs/mL).
   2. Take 10 mL of the cell resuspension (10⁹ PBMCs ) and place them into a 50 mL tube.
   3. Add 500 µL (50 µL/mL of buffer) of NK cell isolation antibody mix and 10 µL (1 µL/ mL of buffer) of anti-CD3 positive isolation antibody mix to the PBMCs and incubate at room temperature for 10 min.
   4. Vortex the magnetic beads and add 1 mL to the mix of PBMCs with antibodies (100 µL beads/ml PBMCs). Incubate for 10 min at room temperature with occasional stirring.
   5. Add 35 mL of NK isolation buffer (3.5 ml buffer/ml PBMCs), mix, and place on the magnet for 15 min (2.2 x 10⁷ PBMCs/mL). After that time, the beads and cells positively selected (all but NK cells) will have adhered to the walls of the tube.
   6. Carefully collect the supernatant (containing NK cells) with a 50 mL plastic pipette without touching the sides or the bottom of the tube.
   7. Count the cells using a cell counter and centrifuge at 800 x g for 5 min.
   8. Resuspend isolated NK cells at 5 x 10⁶ cells/mL in IMDM containing 10% HS and place them in an incubator at 37 ˚C with 5% CO₂ until the experiment is performed.     
      NOTE: This NK isolation protocol states cell numbers and reagent volumes adapted for the use of a 50 mL tube and a big magnet. This protocol can be scaled up (in case more PBMCs are obtained) by repeating the isolation steps in different tubes or scaled down by reducing the volume in the tube. For final volumes of 14-45 mL a 50 mL polystyrene tube is used, for volumes 4-14, a 15 mL polystyrene tube is used and for volumes 1-4 mL, a 5 mL polystyrene tube is used. The magnet is different and fit the appropriate tube in each case. The amount of antibody mix and magnetics beads can also be scaled according to the number of cells and the final volume.
3. NK cell population staining for flow cytometry
   1. Take 0.25 x 10⁶ cells per sample from step 2.2.8, remove the medium by centrifuging at 800 x g for 5 min at room temperature and resuspend the cell pellet in 500 µL of PBS.
   2. Add viability dye according to the manufacturer's recommendation (1 µL of DMSO-reconstituted dye to 1 mL of cell resuspension) and incubate at room temperature for 30 min.
   3. Wash 2x by resuspending in 5 mL PBS and centrifuging at 800 x g for 5 min, at room temperature.
   4. Stain with the corresponding anti-human antibodies in 100 µL of IMDM containing 10% HS for 30 min on ice protected from light (see Table 1). All the antibodies can be combined and used in a single staining step.
   5. Analyze the samples by flow cytometry to evaluate the purity of the NK cell population obtained. Use the cell gating and analysis method that has been previously described.
4. NK cells stimulation with soluble IL-15
   1. Resuspend 0.75 x 10⁶ cells from steps 2.2.8 or 2.4.3 in 100 µL of IMDM containing 10% HS in a well of a 96 well-plate (round bottom).
   2. Dilute human IL-15 to 1 µg/mL in IMDM containing 10% HS. Add 100 µL of the diluted human IL-15 to the cells to reach a final concentration of 0.5 µg/mL.
      NOTE: 0.5 µg/mL is a saturating concentration of IL-15. Lower concentrations of IL-15 or other cytokines such as IL-2, IL-12 or IL-18 may be tested by researchers if desired.
   3. Place the cells in the incubator at 37 °C and stimulate for 48 h before performing the extracellular flux assay. Resuspend unstimulated (control) cells at the same concentration and volume in IMDM containing 10% HS without IL-15, and place them in the same incubator for 48 h.

3. Hydration of sensor cartridge

NOTE: The 96 probe tips of the sensor cartridge contain individual solid-state fluorophores for O₂ and H⁺ that need to be hydrated in order to detect O₂ and pH changes.

1. Turn on the analyzer and let it warm up to 37 °C.
2. Open the sensor cartridge package and separate the sensor cartridge from the utility plate. Add 200 µL of the calibrant solution in each well of the utility plate and put back the sensor cartridge onto the plate, validating that the sensors are completely submerged in the solution. For optimum results incubate the cartridge overnight at 37 °C in a CO₂-free incubator that is properly humidified to prevent evaporation. Prevent bubble formation under the sensors during hydration.
   NOTE: The minimum cartridge hydration time is 4 h at 37 °C in a CO₂-free incubator. Alternatively, overnight hydration of the sensor cartridge with 200 µL of sterile water at 37 °C in a CO₂-free incubator, followed by an incubation of the sensor cartridge with 200 µL of pre-warmed calibrant solution 45 – 60 min prior to the start of the run, can be used.

4. Extracellular flux assay

1. Preparation of adhesive-coated plates         
   NOTE: Since the measurement of metabolic parameters takes place in a microchamber formed at the bottom of the 96-well assay plate, suspension cells must first adhere to the bottom of the well. A cell adhesive extracted from the mussel Mytilus edulis is employed. The manufacturer of the cell adhesive recommends a coating concentration of 1 to 5 µg/cm². The well of the analyzer cell culture microplate has a surface of approximately 0.110 cm². Thus, for a 5 µg/cm² concentration, around 0.55 µg of adhesive is required. As 25 µL of the adhesive solution will be used to coat each well, the optimal adhesive solution concentration for the analyzer cell culture microplates is around 22.4 µg/mL (22.4 µg/mL x 0.025 mL = 0.56 µg).
   1. Prepare 2.5 mL of cell adhesive solution (22.4 µg/mL) in 0.1 M sodium bicarbonate, pH 8.0. Bicarbonate provides the optimum pH for cell adherent performance which, according to the manufacturer, is between 6.5 and 8.0.
   2. Pipette 25 µL of the cell adhesive solution to each well of the assay plate and incubate at room temperature for 20 min. After that, remove the solution and wash 2x with 200 µL of sterile water/well. Let the wells dry by keeping the plate open for 15 minutes inside a cell culture hood.
      NOTE: Coated plates may be stored for up to 1 week at 4 °C.
2. Cell seeding in plates coated with adhesive
   1. Centrifuge cells from step 2.4.3 at 200 x g for 5 min at room temperature. Remove supernatants and wash cells in a warmed mitochondrial stress test medium (if a mitochondrial stress test is being performed) or glycolysis stress test medium (if a glycolysis stress test is being performed). Pellet cells again and resuspend to the preferred cell concentration in the same medium (resuspension volume will depend on the cell concentration chosen; since each well will contain 180 µL of the cell suspension, prepare 0.26 x 10⁶, 0.52 x 10⁶, 1.04 x 10⁶, 2.08 x 10⁶, 4.17 x 10⁶ and 8.33 x 10⁶ cells/mL cell suspensions for 0.047 x 10⁶, 0.094 x 10⁶, 0.187 x 10⁶, 0.375 x 10⁶, 0.75 x 10⁶ and 1. 5 x 10⁶ cells per well respectively).
   2. Plate 180 µL of cell suspension per well along the side of each well. A multichannel pipette is recommended. Use wells A1, A12, H1, and H12 of the analyzer culture plate as control wells for background correction. Add 180 µL of the assay medium in these wells (no cells). Additional control wells can be used if desired and if there is enough space in the plate.
      NOTE: The presence of serum may cause poor cell attachment.
   3. Incubate the plate for 30 min at 37 °C in a CO₂-free incubator. Prepare 10x compounds in the meantime (see step 4.3 below).
   4. Change the centrifuge settings to zero braking. Centrifuge the plate at 200 x g for 5 min. Observe the cells under the microscope to check that they form a monolayer at the bottom of the well. Transfer the plates back to the CO₂-free incubator for 25 min. For best results, the total time after plating should be no greater than around 1 h.
3. Preparation of 10x working solutions to load into sensor cartridge
   NOTE: Each of the 96 probe tips of the sensor cartridge harbors 4 ports (A, B, C, and D) that can be used to inject compounds sequentially into individual wells.
   1. To perform the mitochondrial stress test, prepare 2.5 mL each of 10 µM oligomycin, 1 mM DNP, and a mixture of 10 µM rotenone and 10 µM antimycin A, in mitochondrial stress assay medium (use the stock solutions from step 1.2.3). Final concentrations in the well after injection will be 1 µM oligomycin, 0.1 mM DNP, and 1 µM antimycin A/rotenone.
   2. To perform a glycolysis stress test, prepare 2.5 mL of a mixture of 10 µM rotenone and 10 µM antimycin A in a glycolysis stress assay medium (use the stock solutions from step 1.2.3). Dissolve glucose in a glycolysis stress test medium for a 100 mM solution and 2-deoxy-glucose (2-DG) in a glycolysis stress test medium for a 500 mM solution. Final concentrations in the well after injection will be 10 mM glucose, 1 µM antimycin A/rotenone, and 50 mM 2-DG.
   3. Warm solutions to 37 °C, check pH, and readjust to 7.4 if required. Load compounds prepared in step 4.3.1. (for a mitochondrial stress test) or 4.3.2 (for a glycolysis stress test) into ports A, B, and C of the hydrated sensor cartridge (from step 3.2) using a multichannel micropipette and the port-loading guides provided with the cartridge, as shown in Table 2.
      NOTE: To ensure proper injection in all wells during the assay, each series of ports that are used (e.g., all ports A) must contain the same injection volume across the entire sensor cartridge. This applies to background correction wells and even to those wells not used in the experiment.
   4. Incubate the loaded sensor cartridge at 37 °C in a CO₂-free incubator while setting up the program.
4. Setting up extracellular flux assay protocols
   1. Open the extracellular flux analyzer software, and using the Group Definitions and Plate Map tabs indicate groups of wells that have similar conditions (e.g., wells with the same number of cells, or wells with either resting cells or IL-15-stimulated cells). Also, indicate background correction wells (by default A1, A12, H1, and H12 will be set, but additional wells can be used) and empty wells.
   2. Set up the program described in Table 3 in the software using the Protocol tab.
   3. Begin the program using the Run Assay tab. Place the sensor cartridge (hydrated and loaded with 10x compounds) and utility plate onto the tray. After the calibration step (15 – 20 min), when prompted, replace the calibrant plate for the assay plate (without lid) with attached cells. After this, the run is fully automated (the machine will perform all measurements and injections).
      NOTE: It is possible to perform a mitochondrial stress test and a glycolytic stress test in the same plate, as long as the specific compounds are loaded into the proper ports (oligomycin, DNP, and antimycin/rotenone in ports A, B, and C respectively of the wells where a mitochondrial stress test is performed; glucose, antimycin/rotenone and 2-DG in ports A, B and C respectively of the wells where a glycolysis stress test is performed), the same volumes for injections are used in each series of ports and wells for each test are properly identified using the Group Definitions and Plate Map tabs of the software.
   4. After the completion of the run, retrieve the data and analyze them using the software.

Table 1: Reagents and antibodies used for flow cytometry.

Table 2: Compound loading.

Table 3: Program layout.