Human Neutrophil Flow Chamber Adhesion Assay

All donors recruited for this study gave written informed consent and the study was approved by the University of Alabama at Birmingham Institutional Review Board.

1. HUVEC Initial Culture and Subculture

1. Culture human umbilical vein endothelial cells (HUVEC) in vitro in complete growth medium that is comprised of endothelial cell basal medium (see List of Materials) supplemented with endothelial cell growth factors.
   Note: The growth factors used in this study include: 5 ng/ml human recombinant Epidermal Growth Factor (hEGF), 1.0 mg/ml hydrocortisone, 50 mg/ml Gentamicin and 50 ng/ml amphotericin-B (GA-1000), 2% Fetal Bovine Serum (FBS), 0.5 ng/ml Vascular Endothelial Growth Factor (VEGF), 10 ng/ml human Fibroblast Growth Factor-Basic with heparin (hFGF-B), 20 ng/ml human recombinant Insulin-like Growth Factor (R³-IGF-1), 1 µg/ml ascorbic acid, and 22.5 µg/ml heparin. The complete medium is prepared initially at 37 °C and can then be stored at 4 °C for use within 1 month of preparation.
2. To prepare a flask for cells, complete growth medium is added to a 75 cm² tissue culture flask (1 ml/5 cm²) and then the flask is allowed to equilibrate to 37 °C/5% CO₂ in a humidified incubator for at least 30 min. Human HUVEC will be seeded directly into this equilibrated culture flask at a density of 2,500-5,000 cells/cm².
3. While the media is equilibrating, quickly thaw the stock HUVEC cryovial in a 37 °C water bath. Disperse the cells in the original storage vial by vortexing and then add them directly to the culture flask containing the pre-equilibrated HUVEC complete growth medium to achieve a density of 2,500-5,000 cells/cm². Gently rock the flask to evenly distribute the cells and then return the flask to the incubator. These cells now represent the 1^st passage.
4. Medium should be changed every two days until the cells are 70-80% confluent.
5. The HUVECs can now be harvested from the culture flask with Trypsin/EDTA. The culture media is aspirated from the culture flasks followed by a PBS rinse to remove any residual protein and calcium from the cells. A 0.25% Trypsin-EDTA solution is added and within 2-6 min cell detachment should be evident as assessed by light microscopy.
6. When 90% of the cells are rounded up off the plate, stop the trypsinization by adding an equal volume of 2x trypsin inhibitor. To facilitate the harvest of cells, add another equal amount of complete growth medium. Transfer the detached cells to a sterile 15 ml centrifuge tube.
7. Centrifuge the detached cells at 225 x g for 5 min at room temperature. Aspirate the supernatant, and then resuspend the cells in 2-3 ml of complete growth medium. Determine the cell concentration and viability using a hemacytometer and Trypan Blue.
8. To utilize the harvested cells for study, re-seed additional 75 cm² flasks with cells at a density of 10,000 cells/cm² and proceed to step 2.2. Alternatively, cells can be frozen at this point for future studies. For cell freezing, pellet the cells by centrifugation at 225 x g for 5 min at room temperature. Aspirate off the supernatant and resuspend the cell pellet in FBS containing 10% DMSO at a concentration of 1 x 10⁶ cells/ml. The cell suspension is then transferred to cryovials and stored at -80 °C after freezing of the cells in a cell-freezing container.

2. HUVEC Layer Preparation

1. Use of frozen 2^nd passage HUVECs: revival and culture. If using actively growing cells, proceed to step 2.2.
   1. Thaw the cryovial containing 2^nd passage HUVECs from step 1.8 quickly in a 37 °C water bath. Transfer cells from the cryovial to a 15 ml sterile centrifuge tube and add 10 ml growth medium.
   2. Centrifuge the cells at 200 x g for 5 min at room temperature and aspirate the supernatant to remove the residual DMSO.
   3. Resuspend the cell pellet with complete growth medium and transfer the cells into a 75 cm² flask. Add growth medium to a total volume about 20-25 ml and incubate at 37 °C/5% CO₂.
2. Visually examine the cell culture for confluence each day with media changes every two days. Usually within 2-4 days, cells will reach 80-90% confluence at which time they will be ready for use in the flow chamber assay.
3. To prepare the culture dish that will be used in the flow chamber (see Section 5), add 1 ml of 10 µg/ml fibronectin and 0.05% (w/v) gelatin to a 35 mm tissue culture dish and pipette several times to make sure the whole plate surface is coated. Remove the excessive fibronectin and gelatin solution and air dry the plate for at least 30 min to optimize the protein matrix formation. The fibronectin and gelatin solution may be reused up to 10x.
4. Harvest the HUVEC cells from step 2.2 using trypsin-EDTA as described in steps 1.5-1.7. Seed 500,000 cells into each coated tissue culture dish. Add 2 ml growth medium in each dish and incubate at 37 °C/5% CO₂.
5. Allow cells to grow to confluence as in step 2.2. Cells should be visually inspected daily with medium changes every two days. Prior to the flow chamber assay, prime the HUVEC with 20 ng/ml human TNF-α for 4-6 hr to upregulate and stimulate adhesion molecule expression.

3. Purified Ligand Coating

1. Draw a circle of 0.5 cm diameter with a marker or pen at the center of a 35 mm tissue culture dish.
2. Plate 20 µl of 20 µg/ml protein A in the marked area. Use the pipette tip to spread the protein A solution to cover the whole area within the 0.5 cm diameter circle. It is important to not touch or scratch the surface of the dish. Incubate at 37 °C for 1 hr.
3. Wash the protein-A coated plate 3x with 1 ml of PBS (pH 8.0).
4. Plate 50 µl 1% BSA in the marked area to block non-specific binding on the plate. Incubate for 2 hr at 4 °C.
5. Wash the blocked plate 3x with 1 ml of PBS as in step 3.3.
6. Prepare the Fc-adhesion receptor ligand chimeric protein solutions for coating. In this experiment, an ICAM-1/Fc chimera solution at 25 µg/ml and a P-Selectin/Fc chimera at 0.5 µg/ml in PBS (pH 8.0) was used.
7. Coat the marked area with 50 µl of substrate. Incubate overnight at 4 °C. The dish should be used within two days and the coated area should not be allowed to dry out. Add PBS if necessary to maintain the 50 µl solution on the plate.

4. Neutrophil Separation (All Steps Performed at Room Temperature)

1. After obtaining informed consent, collect participant blood by phlebotomy into an anticoagulant blood collection tube or vacutainer (EDTA or heparin). After blood collection, dilute the blood 1:1 with PBS before separation.
2. Prepare a two layer Ficoll for separating PBMC and neutrophils in 50 ml centrifuge tubes. First add 15 ml heavy Ficoll (see List of Materials, ρ=1.118-1.120), and then carefully layer 10 ml light Ficoll (see List of Materials, ρ=1.077-1.080) on top of the heavy Ficoll. There should be a sharp border between the light Ficoll and heavy Ficoll layers. Finally, carefully layer 25 ml of the diluted blood sample on top of the light Ficoll without disturbing the Ficoll layer.
3. Centrifuge the tube at 250 x g for 30 min at room temperature. Note: The centrifuge rotor brake should be off for these spins to minimize potential disruption of the cell separation during rotor deceleration at the end of the centrifugation. After centrifugation, the following layers (from top to bottom) are present: the top layer is the plasma followed by the PBMC layer on top of the light Ficoll layer, the neutrophil layer with few red blood cells (RBC) is between the light and heavy Ficoll followed by the heavy Ficoll layer and RBCs at the bottom of the tube.
4. Harvest and transfer the neutrophil layer into a new 50 ml tube with a transfer pipette, add PBS to a final volume of 50 ml and centrifuge at 225 x g for 10 min at room temperature.
   1. After centrifugation, there may still be RBCs mixed with the neutrophils. Aspirate the supernatant down to the 10 ml mark. Resuspend the neutrophil-RBC pellet by gentle agitation of the tube (or a brief low speed vortexing) and then wash again with 50 ml of PBS.
5. After the second wash, aspirate the supernatant. To remove the contaminating RBCs, resuspend the pelleted cells in the residual PBS by agitation (or short vortexing), add 25 ml H₂O and gently vortex for 10 sec to lyse the RBC.
   1. Add 25 ml of 1.8% NaCl and immediately mix by centrifugation at 225 x g for 10 min at room temperature. The contaminating RBC should now be lysed leaving a white neutrophil cell pellet.
6. Wash the neutrophil cell pellet with PBS.
7. Resuspend the isolated neutrophils in RPMI medium with 10% FBS and determine the cell concentration under light microscope with a hemocytometer.
8. Adjust the cell density to 500,000 cells/ml with complete RPMI-10% FBS medium.

5. Flow Chamber Adhesion Assay

1. Assemble the flow chamber. Place the 35 mm dish containing the confluent HUVECs or purified adhesion receptor ligands on the microscope table. Connect the parallel plate flow chamber with syringe pump, vacuum system and leave one line open for the neutrophil input. Insert the flow chamber on top of the plate and fasten the flow chamber assembly¹³. (See Figure 1)
2. Start the video recording program on the computer connected to the microscope. Adjust the field and focus of the microscope until a clear field with fully grown HUVEC cells or a field within the ligand coating region is visible.
3. Using the syringe pump, rinse the flow chamber with RPMI medium. Make sure there are no air bubbles within the chamber or the neutrophil input line.
4. If desired, the neutrophils may be primed with low dose (10^-8 M) fMLP for 10 min. This will allow for a matching of the basal level of neutrophil activation between different donors¹⁴.
5. Use the syringe pump to inject the neutrophils into the flow chamber at defined speeds (a speed of 350 µl/min, which equals a shear stress of 1.5 dynes/cm² is used in this study). Record the video. Because neutrophil adhesion can occur rapidly, a video with a length of 4-5 min is usually sufficient to quantitate adhesion events for analysis.
6. An adherent cell is defined as a cell that moves less than one cell diameter within 5 sec on the HUVEC or ligand coated surface^15,16. Count the total number of adherent cells in the field present in the recorded video using this rule. By recording similar length videos with different donors’ neutrophil, one can calculate the adherent cells/min to compare the adhesion properties between different donors.