A Simple Fluorescence Assay for Quantification of Canine Neutrophil Extracellular Trap Release

All experiments were performed with ethical permission from the Iowa State University Institutional Animal Care and Use Committee.

1. Blood Collection

1. Draw 9 ml of blood from a saphenous, cephalic or jugular vein directly into anticoagulant (e.g., ethylenediaminetetraacetic acid (EDTA), 1.8 mg K₂EDTA/ml blood) vacutainers, or into a syringe with immediate transfer to EDTA-containing blood tubes. Gently roll or invert the blood tubes to ensure adequate mixing of the blood and anti-coagulant.

2. Neutrophil Isolation

1. In a 50 ml conical tube, mix blood with an equal volume of sterile room temperature 3% dextran-500 (made up in 0.9% sterile saline) by gentle inversion. Leave standing upright for 18-20 min at room temperature. Transfer the straw colored upper layer to a fresh tube until it can no longer be collected without contamination by the lower red layer. The erythrocytes in the original tube can be discarded.
2. Centrifuge the supernatant at 500 x g for 10 min at 4 °C. Draw off and discard the supernatant. Manually re-suspend the cell pellet in 10 ml room temperature sterile phosphate buffered saline (PBS). Note that vortexing should not be used to re-suspend neutrophils at any stage of this protocol. Manually drawing off, rather than pouring off supernatants is also recommended throughout the protocol to maximize cell yield.
3. Add 10 ml of a room temperature polysucrose and sodium diatrizoate cell separation solution (e.g., Histopaque 1077) to a 50 ml conical tube. Carefully layer the cell-containing solution over the cell separation solution. Note that using cell separation solution which has not equilibrated to room temperature may reduce cell yield.
4. Centrifuge at 300 x g for 30 min at room temperature with the brake off.
5. As neutrophils are in the lowest layer of the gradient, draw off and discard the upper 3 layers, composed of an upper layer of plasma and platelets, a narrow white band of mononuclear cells and a clear layer of the density gradient medium.
6. To lyse any remaining erythrocytes, re-suspend the neutrophil pellet in 10 ml of room temperature water.
7. After 30 sec, restore tonicity by adding 10 ml of sterile room temperature 1.8% sodium chloride and mix by gentle inversion.
8. Centrifuge at 500 x g for 5 min at 4 °C. Draw off the supernatant and discard.
9. If the pellet is still red, repeat the lysis step. If the pellet is white (or if lysis has already been performed twice), gently re-suspend cells in 10 ml of sterile PBS.
10. Count cells using an automated counter or a hemocytometer. Typical yields are at least 10⁶ cells per ml of blood.
11. Centrifuge at 500 x g for 5 min at 4 °C. Draw off the supernatant and discard.
12. Re-suspend cells to the desired concentration in sterile room temperature PBS. For the DNA release assay described in section 2, resuspend at 5 x 10⁶ cells per ml.
13. If desired, transfer a small volume of cells directly or using a cytocentrifuge to a glass slide. Allow cells to dry and stain using a rapid fixation and staining kit according to the manufactures' instructions. If the neutrophil isolation has been successful, when examined under the microscope at least 95% of nucleated cells should be granulocytes (neutrophils, basophils, eosinophils) with minimal erythrocyte contamination.

3. DNA Release Assay

1. Set up the DNA release assay immediately after neutrophil isolation.
2. If cell clumping is present on examination of the stock solution by light microscopy, pass the cell suspension through a sterile 70 µm sterile filter immediately before setting up the assay.
3. To each test well of a sterile 96 well plate, add 5 x 10⁴ cells/well; agonist and Roswell Park Memorial Institute-1640 (RPMI) medium without phenol red and supplemented with 0.5% heat-inactivated fetal calf serum to a final volume of 100-200 µl. Include at least two blank wells per agonist in which the neutrophil stock solution is replaced by an equal volume of PBS. Note that setting up blank wells for each agonist is important in ruling out DNA contamination of the agonist or interference by the agonist with the fluorescence assay.
4. Incubate at 39 °C for 30 min in a carbon dioxide (4%) incubator.
5. Add agonists at desired concentrations and a cell impermeable nucleic acid dye. Note the optimal concentration of cell impermeable dye will vary between different nucleic acid dyes. Non-stimulated controls in which agonists are replaced by an equal volume of media should be included in each experiment.
   NOTE: It is desirable to dilute agonists in similar volumes of culture medium to maintain consistent conditions between wells. Final well volumes of 200 µl have been used successfully and if this is altered, well volumes should remain identical for all conditions. Phorbol-12-myristate-13-acetate (PMA) (final concentration ≥0.1 µM) or platelet activating factor (PAF) (final concentration ≥31 µM) can be used as positive controls. Agonists should be measured at least in duplicate.
6. Incubate at 39 °C. Measure fluorescence using a microplate reader after 2 hours or at desired intervals. Note that optimal time intervals will vary with agonists used.
   NOTE: Wavelength will depend on dye employed.
7. Subtract blank fluorescence before calculating mean fluorescence.
8. To allow comparison between individuals and between plates, calculate a fold change in fluorescence by dividing the mean fluorescence of stimulated wells by the mean fluorescence of non-stimulated wells.