Super-resolution Imaging of the Natural Killer Cell Immunological Synapse on a Glass-supported Planar Lipid Bilayer

1. Preparation of Liposomes

1. Calculate the amount of chloroform-suspended stock solutions of 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC) and 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-cap biotinyl (Biotin-PE) to make diluted stocks at the desired final concentration. To make final concentrations of 400 μM DOPC and 80 μM Biotin-PE phospholipids at 10 ml each, start by placing 629 μl of 10 mg/ml DOPC and 88 μl 10 mg/ml Biotin-PE into separate glass chromatography tubes.
   NOTE: it is important to clean glass Hamilton syringes and glass chromatography tubes by cleaning solution (1 L 95% ethanol to 120 ml water containing 60 g potassium hydroxide,KOH), while transferring chloroform-suspended stock solution of DOPC and Biotin-PE.
2. Dry the chloroform with a stream of argon in the chemical hood. Seal the chromatography tube by parafilm.
3. Subject the newly dried liposomes to a high vacuum in a lyophilizer O/N to remove any residual chloroform. For same-day completion, dry for 60-90 min.
4. While the lyophilizer runs, prepare some dilution buffer. For this protocol, prepare 25 ml consisting of 25 mM Tris, pH 8.0; 150 mM NaCl; and 2% (by weight) n-octyl-β-D-glucopyranoside (OG) detergent. Mix the first two ingredients together first, then displace oxygen with argon before adding the dry OG powder. After preparation, filter the OG solution with 0.2 micron cellulose acetate membrane, and store at 4°C.
5. In addition, prepare two screw-top bottles of 1 L of Tris-saline buffer at the same concentrations, but without OG. Place a distilled water-cleaned magnetic stir bar in the bottom of each. Prepare 6 additional liters of the Tris-saline buffer. Remove oxygen from all the bottles with argon and place them at 4° C as well.
6. After lyophilization, dissolve the dried lipids in the Tris-saline OG buffer to make a 4 mM solution of each. Following the example volumes, add 2 ml to the DOPC tube, and 0.2 ml to the Biotin-PE tube.
7. Mix together the biotin-PE lipids with the DOPC lipids. This improves the mobility of the SLB, as the coupled biotin can impair the fluidity of the phosphate head groups. To make a final concentration of 80 μM Biotin-PE, mix 0.2 ml of 4 mM Biotin-PE and 1 ml of 4 mM DOPC. Then add 8.8 ml of the Tris-saline OG buffer.
8. For a final concentration of 400 μM DOPC, simply mix 1ml of 4 mM DOPC with 9 ml of Tris-saline OG.
9. Fill the sonicator with ice water. Put the glass tube containing the diluted phospholipid in the center of the sonicator by using a utility clamp. Sonicate the diluted phospholipid for 10 min until the solution becomes clear.
   NOTE: Add ice into the sonicator water bath to keep the temperature low since sonication will generate heat.
10. Fill in the tubes with argon to displace the oxygen in the air above the liquid, and seal them with parafilm.

2. Dialysis of Liposomes

1. Cut two sections of dry dialysis tubing (Molecular Weight cut-off: 12-14,000, diameter: 6.4 mm) of appropriate length (in this example, 40 cm), one for each phospholipid dilution, from the roll.
2. Rehydrate the tubing sections by allowing them to soak in a 200 ml of distilled water in a glass beaker for 2 min.
3. Microwave this for 5 min at a high setting, or at least until the water comes to a boil.
4. Tie a knot at one end of each tube and rinse out the interior with a few milliliters of Tris-saline-OG buffer. Thereafter, meticulously squeeze out as much of this wash buffer as possible to minimize the amount of buffer remaining inside.
5. In a laminar flow hood, add the diluted phospholipids into each tube and clamp the open ends with a small dialysis tube closure so as to exclude all air. Complete air exclusion will require the sacrifice of a small volume of the sample by clamping below the “water line”.
6. Immerse the samples in the prepared previously bottles of Tris-saline buffer without OG. Displace oxygen in the bottle with argon before re-sealing and place to stir O/N at 4°.
7. Transfer the tubing into a new bottle of Tris-saline buffer without OG every 12 h for at least 3 times.
8. Shortly before removal of the dialyzed lipids, prepare a number of small tubes into which to aliquot the lipids by filling each with argon to displace the oxygen.
9. After 36 h, take the dialysis bottles into the laminar flow hood and remove the dialysis tubes from the bottles. Have a bench diaper or beaker on hand to collect the wet runoff.
10. Cut the dialysis tubing above the clip, then remove the clip and carefully transfer the dialyzed lipid solution via pipet into 1 ml aliquots in pre-prepared tubes filled with argon gas on ice.
11. Aliquot the aqueous liposome solution, use the argon stream to displace oxygen again in each tube.
12. Store the liposomes at 4°. Do not freeze.

3. Determination of Antibody Density on the Lipid Bilayer

1. Prepare a dilution series of biotinylated, fluorescently-labeled antibody, 50 μl in volume for each dilution, at the following concentrations: 0 nM (blank), 10 nM, 50 nM, 100 nM, and 500 nM. (Hereafter referred to as “sample series”).
2. Add 1 μl of silica beads into 6 wells of a 96-well v-bottom plate. Be sure to shake the beads well before pipetting, as they tend to settle.
   NOTE: If having ordered dry beads rather than suspended, follow the manufacturer’s instructions to dilute.
3. To these beads, add 2 μl of mixed DOPC:Biotinyl phospholipids at a 1:1 ratio. Do this for each well.
4. Pulse the plate on a vortexer at medium strength 3 times for 10 sec each to encourage interaction with the beads and the phospholipids.
5. Add 150 μl of 5% casein to each well. Mix well by pipetting up and down three times.
6. Let the plate incubate in the casein solution for 10 min, then wash it out. To wash, fill each well to a total volume of 250 μl with HEPES buffered saline (HBS) with 1% human serum albumin (HSA). Centrifuge at 1,000 x g for 2 min. Pipette out and discard the top 200 μl of supernatant, and repeat twice for a total of three wash cycles.
7. Add 50 μl of streptavidin at a 333 ng/ml concentration. Pulse the plate again 3x 10 sec, and let it sit on a shaker for 15 min. Wash 3x as in step 6 to remove unbound streptavidin.
8. Add 50 μl of the fluorescently labeled biotinylated antibody from the previously prepared dilution series (see step 3.1) to each well, and replace the plate on the shaker for 20-30 min. Wash 3x as in step 6 to remove unbound antibody.
9. After the final wash, resuspend the beads in 100 μl of HBS/1% HSA, then transfer to a FACS tube. Repeat this twice to ensure the efficient removal of all beads from the well, for a total of 300 μl, and do this for each well.
10. Bring the resulting tubes to the flow cytometer. It is time to read them.
11. Add 1 drop from the bottle marked “B” from the fluorescence intensity calibration (FIC) bead kit (hereafter referred to as “standard series”) to a FACS tube, and dilute with 300 μl of HBS/1% HSA.
12. Read this tube, but do not record the data just yet. For now, simply ensure that the blank beads are properly zeroed. Make a histogram showing the fluorescence of the beads measured in the appropriate channel, then shift the voltage of the excitation laser down until the peak is at the far left side of the histogram.
13. Remove the tube and add 1 drop of each of the other 4 tubes in the series (labeled 1 through 4) to the same tube. Now, place the tube in the machine again, and record the resulting data, which should appear as 5 distinct peaks.
14. Read each individual tube from the sample series.
15. Using FACS analysis software, draw a gate spanning the entire width of each peak in the standard series histogram at the half-maximal point. That's one gate for each peak. Do the same for each sample. Note the MFI (mean fluorescence intensity) for each gate.
16. Using a spreadsheet program input the measured MFI values in the appropriate place. Also input the MESF (Molecules of Equivalent Soluble Fluorochrome) values (average number of fluorescent molecules coating each bead) for each bottle in the standard series. This information can be found by following the directions on the plastic jar the tubes came in.
17. Use the spreadsheet will plot the MFI against the MESF values, creating a linear correlation between the number of fluorophores and the measured intensity.
18. Use the module of ‘proteins & labels’ in the micro-volume spectrophotometer to determine the ratio of dye to protein in the labeled antibody.
19. Input the labeling efficiency of the sample protein, the average diameter of the beads coated with lipids, and the MFI value for each entry. The spreadsheet will automatically use the formula from the line graph generated in step 3.16 to compute the MESF values for each protein dilution and the seeding density of the sample protein at each concentration.

4. Isolating and Culturing Human NK Cells

1. Aliquot 15 ml peripheral blood or buffy coat into a 50 ml conical tube. Dilute this blood with PBS containing 1% FBS at the ratio of 1:1.
2. Add 13 ml of Ficoll gently to the bottom of the tube with a 10 ml serological pipette.
3. Centrifuge this tube for 20 min at 1,200 x g with the accelerator and the break off or at their lowest settings.
4. After centrifugation, use a serological pipette to collect the floating cloudy white middle layer of peripheral blood mononuclear cells (PBMCs), which should sit and the intersection between a clear yellow upper layer and a more cloudy pale-colored lower layer, both of which sit above a lowermost layer of red blood cells (RBCs). NOTE Do not to collect any RBCs in collecting the PBMCs.
5. Place the collected PBMCs in a new 50 ml conical tube, and dilute to capacity with PBS containing 1% FBS. Centrifuge again, this time with the brake and accelerator on maximum, for 5 min at 300 x g.
6. Discard the supernatant and resuspend the cells in 10 ml of PBS containing 1% FBS.
7. Count the cells while centrifuging once more at the same settings as in step 4.6.
8. Discard the supernatant once more, and resuspend the cells in R10 medium at a density of 10 million cells/ml.
9. Take 30 million cells in a 5 ml polystyrene tube, and isolate NK cells using a magnetic separation kit, following the manufacturer’s instructions. Following isolation, count the cells one more and resuspend at a density of 500,000 cells/ml in R10 complete medium (88% RPMI; 10% FBS; 1% HEPES; 1% sodium pyruvate) supplemented with IL-2 (100 U/ml). Culture at 37° in a CO₂ incubator, and replace medium 2-3 times weekly.

5. Assembling the Glass-supported Planar Lipid Bilayer

1. Prepare 100 ml piranha solution by mixing 30% hydrogen peroxide with sulfuric acid at a ratio of 1:3 in a beaker.
   NOTE: Always carry out work with noxious agents like sulfuric acid in a properly designated chemical fume hood.
2. Into this solution, immerse 2 rectangular #1.5 coverslips in piranha solution for 20-30 min.
   NOTE: It is essential to clean the coverslips by piranha solution.
3. While the coverslips are being cleaned, take 1 tube of previously prepared 400 μM DOPC lipids and 1 tube of previously prepared 80 μM Biotin-PE lipids. Transport them on ice to the argon tank.
4. Displace oxygen in a new microcentrifuge tube with argon, then add together the DOPC and Biotin-PE at a 1:1 ratio. Specific volume will vary based on experimental needs, but should be at minimum 2 μl each. Displace oxygen in the mixture tube once more with argon gas, and the individual reagent tubes as well, before returning the latter to the refrigerator.
5. After they are finished cleaning, thoroughly rinse the coverslips with distilled water. Set the coverslips out to air dry for a few minutes.
6. Withdraw 1.5 μl of the liposome mixture prepared in step 5.4 and aliquot it in a single drop into one of the lane chambers of the chamber slide. The use of 2 drops per lane is typical, but not necessary.
7. Quickly and efficiently place the dry coverslip over the droplets. Ensure that the drops are sufficiently spaced so that they do not merge once the coverslip is placed. Furthermore, make sure that the drops remain circular and well-defined, without touching the edges of the chamber walls. Press down firmly in between and around each lane to ensure a watertight seal between the coverslip and slide.
8. Mark the positions of the drops using a marker pen
9. Pass 100 μl of aqueous 5% casein through the chamber to block the bilayer. Try to make sure that there are no bubbles in the flow chamber.
10. Inject 100 μl of streptavidin at a concentration of 333 ng/ml into each lane. Incubate for 10-15 min at RT. Thereafter, wash by running 3 ml of HBS/1% HSA through each lane to remove the excess streptavidin.
11. Add 100 μl of biotinylated fluorescently labeled anti-CD16 such as Alexa Fluor 568 at the protein concentration previously determined to be most effective in section 3. Incubate in the dark for 20-30 min. Wash again by running 3 ml of HBS/1% HSA through each lane.
12. Flow 100 μl of D-biotin at a concentration 25 nM through the chamber in order to bind any excess streptavidin and thus eliminate the chance of non-specific binding of streptavidin to the cells.
13. Count NK cells and resuspend at a concentration of 500,000/ml in HBS/1% HSA.
14. While spinning down the cells, wash the D-biotin out of the chamber with another 3 ml of HBS/1% HSA per lane.
15. Check the mobility of ligands on the SLB by fluorescence recovery after photobleaching (FRAP) on a total internal reflection fluorescence (TIRF) or confocal microscopy prior to adding NK cells.
16. Once the cells have finished spinning and have been resuspended at the desired concentration, add 100 μl to each lane.
17. Place the chamber in a 37° 5% CO₂ incubator for 30-60 min.
18. After this incubation period, fix the cells with 4% paraformaldehyde at RT for 10-20 min. Wash by running 3 ml of PBS through each lane to remove the paraformaldehyde.
19. Add 400 μl blocking buffer (5% normal donkey serum and 0.2% Tritron X-100 in PBS). Incubate at RT for 30 min.
20. Stain F-actin and perforin by adding 200 μl of diluted fluorescently-labeled phalloidin (1 unit/ml labeled phalloidin) and fluorescently-labeled anti-perforin mAb (500 ng/ml anti-perforin mAb). Incubate at RT for 1 hr.
21. Wash by running 3 ml of PBS. The chamber is ready for the imaging.

6. Imaging of the NK Synapse on Lipid Bilayer using STED

1. Turn on all necessary hardware modules.
2. Start up the image analysis software. Enable both resonant scanning and STED modules. After making these selections, wait for about 3-5 min for the software to initiate.
3. Click on the “Configuration” tab at the top of the screen.
4. Select “Laser Config” then turn on the white light and STED 592 nm lasers.
5. Choose the 100x objective, and align the excitation laser beam with the 592 nm depletion laser.
6. Select the “Laser Config” module, turn off the 592 depletion laser, and turn on the 660 nm depletion laser.
7. Place the slide upon the stage, over the lens. Bring the cells bound on the demarcated bilayer region into focus using the white light lamp and the oculars.
8. Return to the “Acquisition” tab, directly to the right of the “Configuration” tab.
9. Click the “Switch to Whitelight” tab, then turn that module on and drag the excitation laser line to the appropriate wavelength.
10. Select the desired detector from the list of those available, then set the detection range to encompass the appropriate range of wavelengths.
    NOTE: NEVER put the detection range directly beneath the excitation beam.
11. Click on the “Seqential” button in the left-hand “Acquire” toolbar to bring up the Sequential Scanning dialogue in the bottom of the left-hand toolbar. This allows the user to add multiple sequences, each with a different excitation beam for a different color. Click “Between Frames”, and then set the excitation frequency, detector, and detection range for each additional color as in steps 6.9 and 6.10.
12. Once all the settings are optimized, hit “Start” to begin the acquisition process.
13. Apply the free deconvolution software (Huygens) using a setting designed for STED deconvolution, as described previously¹³.