Simplified Human Neutrophil Extracellular Traps (NETs) Isolation and Handling

NOTE: All experiments were conducted in accordance with local institutional ethical guidelines.

1. Blood Drawing

1. Through antecubital venipucture, collect 14 tubes of blood from a healthy volunteer into green top (heparinized) tubes and invert each tube before settling on ice. Collect blood within 10 – 15 min of experiment to ensure optimal neutrophil yield.

2. Neutrophil Isolation from Whole Blood

1. Wash each green top tube of blood with 5 ml of PBS without Ca and Mg to dilute the blood. In each of 4 x 50 ml conical tubes, add 15 ml of Lymphocyte Separation Media (LSM) at RT. Using a 18 G needle mounted on a 60 ml syringe, carefully layer the diluted blood onto the LSM, creating a sharp LSM-blood interface.
   NOTE: Avoid mixing of the layers as much as possible.
2. Centrifuge at 800 x g for 30 min at 21 °C without break.
   NOTE: Sudden breaks can cause mixing of the different layers.
3. Observe the erythrocytes and neutrophils sediment at the bottom. Aspirate and discard the top 2 layers (plasma and LSM) making sure to get rid of the interface between these layers which contains lymphocytes and mononuclear cells, leaving only the bottom red layer.
4. To each tube, add 20 ml of Phosphate Buffer Saline (PBS) and 20 ml of 6% Dextran solution. Gently invert each tube to mix with the layer of erythrocytes and neutrophils and allow to sit at RT for 30 min.
   NOTE: This will allow red blood cells (RBCs) to sediment at the bottom.
5. After 30 min, transfer the neutrophil rich supernatant into fresh tubes and discard the RBC pellets. Centrifuge supernatants at 450 x g at 4 °C for 5 min. After centrifugation, discard supernatant. The pellet will contain mostly neutrophils and few RBCs.
6. Prepare a lysing solution by adding 0.5 ml of lysing buffer to 4.5 ml of sterile water. Lyse remaining RBC with the prepared 5 ml of lysing solution, transferring all resuspended pellets into one tube. Allow lysis at RT in the dark for 10 min.
7. Centrifuge at 450 x g at 4 °C for 5 min. Discard supernatant and wash pellet with 5 ml PBS without Ca and Mg and centrifuge again at 450 x g at 4 °C to get rid of any remaining lysing solution. The pellet obtained at this point contains the neutrophils. Resuspend the pellet in 30 ml of cold 3% RPMI and put on ice.
   NOTE: 3% RPMI is made by supplementing RPMI media with 3% Fetal Bovine Serum (FBS)
8. Verify purity of the sample using Methylene blue staining. >95% of cells should be granulocytes with multi-lobar nuclei. Use Trypan blue staining to verify >95% viability of cells and count the final neutrophil yield. Dilute neutrophils in ice cold 3% RPMI to obtain a final concentration of 5 x 10⁶ neutrophils/ml.

3. NETs Generation

1. Stimulate neutrophils with 500 nM of PMA (per 30 ml of neutrophil solution) and incubate on a 150 x 25mm flat tissue culture dish with 20 mm grid for 4 hr at 37 °C 5% CO₂. This will allow NETosis.
2. After 4 hr of stimulation, gently aspirate and discard the media, leaving the layer of NETs and neutrophils adhered at the bottom. Do not disrupt this layer.
3. Using a total of 15 ml of cold PBS without Ca and Mg per dish, wash the bottom of each dish by pipetting 15 ml of PBS on the bottom of the dish in order to lift off all adherent material from the bottom.
4. Collect solution obtained from washing each dish (step 3.3) in a 15 ml conical tube and centrifuge for 10 min at 450 x g at 4 °C. Neutrophils and any remaining cells will pellet at the bottom, leaving a cell-free NET-rich supernatant.
5. Divide supernatant into 1.5 ml micro-centrifuge tubes and spin for 10 min at 18,000 x g at 4 °C. This will allow all DNA to pellet.
   NOTE: Centrifugation in larger tubes is easier and less time consuming if such high speeds are attainable on the regular centrifuge, otherwise will need to divide supernatants in smaller tubes in order to use high speed micro-centrifuge.
6. Discard supernatant and resuspend all pellets obtained together in ice cold PBS to a concentration corresponding to 2 x 10⁷ neutrophils per 100 μl of PBS. This will yield the cell-free NET stock that can be used for subsequent experiments.
7. Measure DNA concentration in the sample obtained using spectrophotometry or alternate DNA quantification tool. An adequate concentration in the sample should range between 140 – 180 ng/μl.

4. NET-Cancer Cell Static Adhesion Assay

1. Add 100 μl of the previously obtained NET stock per well in a 96 well flat bottom plate and allow to coat wells O/N at 4 °C in the dark.
2. Between 12 and 20 hr later, verify formation of a uniform monolayer of cell free NETs at the bottom of the wells under the microscope (Figure 1). At this point, gently aspirate all non-adherent material out of the wells, making sure not to disrupt the NET monolayer at the bottom.
3. Add 100 μl of 1% Bovine Serum Albumin (BSA) blocking solution to each well and leave for 1 h at RT.
4. Detach A549 cancer cells from a T-75 flask of using 2 ml of 0.25% Trypsin solution. Once detached, add 10 ml of A549 media to trypsinized cells and centrifuge at 450 x g at 4 °C for 5 min. Discard supernatant and resuspend cells in media to obtain a concentration of 2 x10⁴ cancer cells per 100 μl media.
   NOTE: A549 cells were grown separately. Briefly, cells were cultured and maintained in DMEM F12 media containing 10% FBS and 1% Penicillin Streptomycin and incubated at 37 °C 5% CO₂. Once 70 – 80% cell confluence was reached, they were detached using 0.25% Trypsin-EDTA and resuspended in the same media described above.
5. Stain cancer cells using CFSE by adding 1 μl of CFSE per ml of media and leave to stain at RT for 10 min. After staining, centrifuge down at 450 x g at 4 °C for 5 min then discard supernatant and resuspend cells in initial volume of media to obtain a concentration of 2 x10⁴ cancer cells per 100 μl media.
6. After 1 hr of NETs blocking, gently aspirate blocking solution and add 2 x10⁴ cancer cells in media, which is equivalent to 100 μl, per well over the NET monolayer and allow to adhere for 90 min at 37 °C 5% CO₂. Gently aspirate the cells and add 100 μl of PBS to each well to wash any non-adherent cells.
7. In some wells, add 1000U of DNAse1 per well for 10 min before washing to degrade NETs. This will serve as negative control. In other wells, add 100 μl of sterile water per well for 10 min, which will serve as the vehicle control (VC).
   NOTE: 3 replicates per condition are usually performed to increase sample size.
8. Aspirate and discard all solution in the wells leaving only NETs and adherent cancer cells at the bottom. Add 100 μl of 4% formaldehyde solution per well to fix adherent cancer cells to NETS and read assay under the fluorescence microscope (Figure 1). Plot and analyze the results (Figure 3).