The study protocol for using human blood samples was approved by the ethics committee of The Hospital for Sick Children, Toronto. All the procedures including healthy human volunteer recruitment for blood donation were performed in accordance with the ethics committee guidelines. All the volunteers participating in this study gave their signed consent prior to the blood donation.

Peripheral blood from healthy male donors were drawn in K2 EDTA blood collection tubes (Becton, Dickinson and Co.) in the hospital at nursing station. The neutrophils isolation was performed using PolymorphPrep (Axis-Shield), according to the company protocol (as previously reported). Briefly, equal volume of blood was laid over PolymorphPrep solution and centrifuged for 35 min at room temperature without applying breaks. After centrifugation, the polymorphonuclear neutrophil layer was collected and washed with washing solution [0.425% (w/v) NaCl with 10 mM HEPES] to eliminate all the residues of PolymorphPrep. Then, red blood cells were lysed twice with 0.2% (w/v) NaCl hypotonic solution for 30 s followed by adding an equal volume of 1.6% (w/v) NaCl solution with 20 mM HEPES buffer to obtain the isotonic condition. After lysis, two more washes were done to eliminate red blood cells debris and soluble components. Neutrophils were resuspended in RPMI medium (Invitrogen) containing 10 mM HEPES (pH 7.2) and counted using hemocytometer. Viability of the purified neutrophils were checked by Trypan blue exclusion assay. Neutrophils purity was determined by imaging Cytospin preparations. Only neutrophil preparations having more than 95–98% viability and purity were used in the experiments. Cells were kept at 37°C and 5% (v/v) CO₂ incubator for the entire experimental period.

The volume of HCl (5 M) or NaOH (5 M) necessary to modify the initial media pH (7.2) to 6.6, 6.8, 7.0, 7.2, 7.4, 7.6, and 7.8 was predetermined using a pH meter. The isotonic RPMI media with different pHs were added to the neutrophil suspension for further experiments. Each experiment performed in this study had a technical duplicate, and the assay was repeated with different donors to obtain biological replicates (n = 3–5; specific details are given in the figure legends).

The NETotic index was measured by Sytox Green, a cell-impermeable DNA dye (Life Technologies), fluorescence. Briefly, neutrophils were resuspended at 1 × 10⁶ cells per mL in RPMI media (pH 7.2), containing 5 µM of Sytox Green dye. Later, 50,000 neutrophils were seeded into 96-well plates containing 50 µL of media with predetermined pH to adjust the pH of corresponding wells. The cells were stimulated with NOX-independent agonists calcium ionophores A23187 (4 µM) or ionomycin (5 µM) and kept at 37°C and 5% (v/v) CO₂ incubator. Sytox Green fluorescence intensities were detected by a POLARstar OMEGA fluorescence microplate reader (BMG Labtech) every 30-min intervals for 240 min. The plates were briefly (~2–5 min) taken out of the incubator for the readings. The NETotic index was calculated based on the value of 100% NET formation obtained by lyzing the cells with 0.5% (v/v) Triton X-100 (representing total DNA presenting in the sample). The baseline green fluorescence at time 0-min was subtracted from the fluorescence at each time point and was then divided by the fluorescence values of cell lysed with Triton X-100. To determine whether pH affects SYTOX Green fluorescence, the same number of cells were lyzed with Triton in various pH buffers, and the green fluorescence of 5 µM SYTOX Green was recorded every 30 min for 240 min.

We used the SNARF^®-4F 5-(and-6)-carboxylic acid (Thermo Fisher Scientific) dye to determine the changes in intracellular pH after adjusting extracellular pH. First, cells were preloaded with 10 µM dye for 30 min at 37°C and 5% (v/v) CO₂, washed and resuspended in fresh RPMI (pH 7.2) at a concentration of 1 × 10⁶ cells/mL. Neutrophils (0.5 × 10⁵ cells in 50 µL) were seeded into a 96-well plate containing the same volume of media to adjust the pH to different values (6.6, 6.8, 7.0, 7.2, 7.4, 7.6, and 7.8). These neutrophils were stimulated either with only media (negative control), A23187 (4 µM; Sigma-Aldrich) or ionomycin (4 µM; Sigma-Aldrich) and dual emission spectra of SNARF were measured. pH changes were recorded every 10 min up to 60 min. The emission spectrum of SNARF undergoes a pH-dependent wave length shift; therefore, the ratio (580/640 nm) of the fluorescence intensities from the dye at two emission wavelengths were used for intracellular pH determinations. Carboxy SNARF-4F is typically used by exciting the dye at one wave length (between 488 and 530 nm), while monitoring the fluorescence emission at two wave lengths (580 and 640 nm). The ratio of the fluorescence intensities of two emission wavelengths 640/580 nm were used as a proxy for intracellular pH. Under this format, increasing SNARF ratio reflects higher pH.

Intracellular calcium concentrations were assessed using Fluo-4-AM calcium indicator dye by a plate reader assay. Briefly, 1 × 10⁶ cells per mL were incubated in HBSS-Mg²⁺ (calcium free) media with 4 µM Fluo-4-AM for 15 min at 37°C and 5% (v/v) CO₂. After washing, the cells were resuspended in RPMI (pH 7.2) and 50,000 neutrophils (50 µL) were seeded in a well of 96-well plates containing 50 µL of media to adjust the pH of corresponding wells. Directly after seeding the cells, Fluo-4-AM fluorescence was measured using the SpectraMax Gemini EM fluorescence microplate reader (0-min; Molecular Devices). Cells were incubated for 10 min at 37°C to allow intracellular pH changes. After incubation, Fluo-4-AM fluoresce was measured. Cells were then stimulated with either calcium ionophore A23187 or ionomycin. After adding calcium ionophores or media controls, the Fluo-4-AM fluorescence intensities were recorded every 30 s up to 1,200 s (at 37°C). The plates were not taken out from the microplate reader until the end of experiment. The ratio comparing each time point with the 0-min reading was calculated and used for determining the calcium influx. To determine whether pH affects Fluo-4 AM fluorescence, the experiment was repeated with media containing Fluo-4 with no cells.

To detect the mROS production, a plate reader assay was performed using the MitoSOX probe. Briefly, 1 × 10⁶ cells per mL were incubated with 4 µM of MitoSOX Red for 15 min at 37°C and 5% CO₂ and seeded in a black clear bottom 96-well plate containing 50 µL of specific media to adjust the pH of corresponding wells. The cells were stimulated with 4 µM of calcium ionophore A23187 or ionomycin, and the fluorescence was measured every 4 min up to 120 min using the SpectraMax Gemini EM fluorescence microplate reader (Molecular Devices). To determine whether pH affects 4 µM MitoSox fluorescence, the experiment was repeated with media containing MitoSox with no cells.

The same method described above was used for obtaining the pH 6.6, 7.2, or 7.8 used in the immunofluorescence. A total of 100,000 neutrophils were seeded into 12-well chamber slides at 37°C (Ibidi, cat #81201) and the cells were activated with either media only (negative control), 4 µM of calcium ionophore A23187 or ionomycin for 30 min (PAD4 and Citrullinated Histone 3 staining) or 120 min (MPO and Citrullinated Histone 3 staining). The cells and NETs were fixed with paraformaldehyde [4% (w/v) for 15 min], permeabilized for 15 min with 0.1% Triton-X 100, and blocked with 5% (w/v) BSA for 1 h. The following antibodies were used for immunostaining: mouse anti-PAD4 antibody (ab128086, Abcam; 1:100 dilution) or mouse anti-myeloperoxidase antibody (ab25989, Abcam; 1:400 dilution) was used for staining PAD4 and MPO, respectively (with secondary antibody conjugated with a green fluorescence Alexa fluor 488 dye; 1:5,000 dilution; Thermo Fisher Scientific), while rabbit anti-citrullinated histone 3 antibody (ab5103, Abcam; 1:400 dilution) was used for detecting the presence of CitH3 (with secondary antibody conjugated with a far red fluorescence dye Alexa fluor 647; 1:5,000 dilution; Thermo Fisher Scientific). DAPI was used to stain the DNA (1:100 dilution). As isotype controls, we used Mouse IgG (eBioscience 14-4714-81; 1:100 dilution) and Rabbit IgG (Invitrogen; 31235; 1:100 dilution). After treating with the secondary antibody, slides were washed and mounted by glass cover slips (Fisher Scientific) with anti-fade fluorescent mounting medium (Dako). The images were then taken using an Olympus IX81 inverted fluorescence microscope with a Hamamatsu C9100-13 back-thinned EM-CCD camera and Yokogawa CSU × 1 spinning disk confocal scan head with Spectral Aurora Borealis upgrade, four separate diode-pumped solid-state laser lines (Spectral Applied Research, 405, 491, 561, and 642 nm). The images were taken at 40×/0.95 magnification and processed by Velocity software (version 6.3, Cell Imaging Perkin-Elmer).

Tubes containing 0.5 × 10⁶ neutrophils in pH adjusted (6.6, 7.2, or 7.8) media were activated either by negative control (only media) or by calcium ionophore A23187 or ionomycin for 60 min (at 37°C and 5% CO₂ incubator). After incubation, cells were lysed using the lysis buffer containing 1% (w/v) Triton X-100, 25 mM NaF, 50 mM Tris, 10 mM KCl, 10 µg/mL Aprotinin, 2 mM PMSF, 1 mM Levanisole, 1 mM NaVO₃, 0.5 µM EDTA, 25 µM Leupeptin, 25 µM Pepstatin, 1 protease inhibitor cocktail tablets per 5 mL (Roche), and 1 phosphatase inhibitor cocktail tablet per 10 mL (Roche). A quarter volume of 5 × loading dye [125 mM Tris.HCl at pH 6.8, 6% (w/v) SDS, 8% (v/v) β-mercaptoethanol, 18% (v/v) glycerol, and 5 mM EDTA, 5 mM EGTA] was added followed by 10 min of heating at 95°C with 350 rcm shaking. The samples were separated in a 5% (w/v) stacking and 10% (w/v) resolving gel at 100 V and transferred on a nitrocellulose membrane for 70 min at 350 mA. After transfer, the membranes were blocked with 5% (w/v) BSA in 0.05% PBST for 2 h at room temperature. The membranes were incubated in the primary antibody at 4°C overnight followed by three washes with PBS with 0.1% Tween (PBST) for 30 min. The antibodies used were: anti-Histone H4 (ab16483, Abcam; rabbit pAb; 1:1,000) and anti-GAPDH (FL-335, Santa Cruz; rabbit pAb; 1:5,000). The membranes were then incubated with the secondary antibody [donkey anti-rabbit IgG-HRP (31458, Thermo Fisher; 1:10,000)] for 1 h and then washed three times with 0.1% PBST for 30 min. The densitometry analysis of the blots was done using the Image Studio software (LI-COR Biotechnology) and normalized to GAPDH.

All statistical analyses were performed on GraphPad Prism 7. One-way ANOVA with Dunnett and Tukey’s post-tests, two-way ANOVA with Bonferroni posttest or t-test was done as appropriate. A p-value of less than 0.05 was considered to represent significant differences between conditions. All the data are presented as mean ± SEM.

To determine the effect of pH on NOX-independent NETosis, we incubated purified peripheral blood neutrophils in media with seven different pHs (6.6–7.8) in the absence (negative control) or presence of calcium ionophores A23187 or ionomycin. The percentage DNA release in each condition was calculated by comparing the green fluorescent intensity of the cells at specific conditions with the cells treated with Triton (100% DNA release). The percentage DNA release (or NETotic index) calculated by SYTOX Green fluorescence showed that alkaline and acidic pHs increased and decreased NET formation, respectively (Figures 1A–C; Figures S1A–C in Supplementary Material; pH itself does not affect SYTOX Green fluorescence signal, Figure S2 in Supplementary Material). A regression line at 120 and 240-min time points of the Sytox Green-based kinetics showed a clear increase in NET formation with increasing pH; the changes were small and slow in the negative control, whereas the increase was much higher for A2387 and ionomycin (Figures S1D–F in Supplementary Material). For example, the slope of spontaneous NET formation after 120 min was ~10-fold less steep (3.754) than that of A23187 (35.92) and ionomycin (32.92)-mediated NET formation, indicating that the effect of pH on calcium ionophore-mediated NET formation was much higher than the spontaneous NET formation. After 4 h, the cells were fixed, and fluorescence images were captured to compare the differences in the overall structure of the nuclei and NETs between the lowest and the highest pH conditions (6.6 and 7.8; Figure 1D). These images confirmed the SYTOX Green plate reader data set. Overall, pH affected both spontaneous and calcium ionophore-mediated NET formation; pH above the normal blood pH of ~7.4 promoted higher levels of NET formation, whereas a more acidic pH suppressed NET formation. The effect was very dramatic in calcium ionophore-mediated NET formation compared to spontaneous NET formation.

To determine the effect of calcium ionophores on intracellular pH (pH_i) of neutrophils in the presence of different extracellular pH (pH_e), we used a pH sensitive dual-wavelength Seminaphtharhodafluor (SNARF) dye. SNARF-loaded neutrophils were resuspended in media with seven different pH_e, from 6.6 to 7.8, with or without A23187 or ionomycin and SNARF ratios were calculated at 0, 10, and 20 min time points. The 20-min time points were chosen to analyze the intracellular pH change because the majority (87–98%) of the cells were alive at that time point, excluding any bias related to extracellular buffers entering the dying cells without any cellular regulation (Table S1 in Supplementary Material). In this format, increasing SNARF ratios (ratio 640/580 nm) reflect higher intracellular pH values (Figures 2A–C). In non-stimulated neutrophils, increasing pH_e increased the pH_i and reached a slope of 7.4x + 0.05x³ SNARF ratios per pH unit at 20 min (Figure 2A). By contrast, at 20 min stimulation with A23187 (a slope of 95x + 0.64x³; Figure 2B) or ionomycin (a slope of 59.3x + 0.4x³; Figure 2C), much higher increase in pH_i values were detected compared to their respective controls (Figures 2D–F; see the full non-linear equations on the graphs). For calcium ionophore conditions, a drastic increase in pH_i was apparent beyond pH_e of 7.6. These results indicate that both of the calcium ionophores drastically increase intracellular pH at the beginning of NOX-independent NET formation process.

Increase in intracellular calcium (Ca²⁺) is important for increasing NET formation-related cellular functions (5). However, the effect of pH on calcium influx in neutrophils is not clearly established. Therefore, we asked whether pH levels could affect the intracellular Ca²⁺ levels in resting and activated neutrophils. First, neutrophils were preloaded with the Ca²⁺ dye Fluo-4 AM and activated with media (negative control), A23187, or ionomycin in three different pH conditions (6.6, 7.2, and 7.8). Cytosolic Ca²⁺ concentration was measured every 30 s for 20 min. Plate reader assays showed that increased pH facilitated Ca²⁺ influx in resting and stimulated neutrophils (Figures 3A–D; increasing pH does not increase the fluorescence of Fluo-4 AM dye, Figure S3 in Supplementary Material). The magnitude of calcium increase is about the same (~0.2-fold) between two pHs; however, the total calcium levels are much higher in the presence of calcium ionophores. To confirm the effect of pH in Ca²⁺ influx, we performed fluorescence imaging, in resting (negative media) or calcium ionophore-activated neutrophils (A23187 or ionomycin), in three different pH conditions (6.6, 7.2, and 7.8; Figure 3E). Images were taken directly after cells stimulation. This data set confirmed the plate reader data showing an increase of the fluorescence in higher pH conditions. Therefore, increasing pH helps to increase intracellular calcium levels in neutrophils, particularly in the presence of calcium ionophores.

Mitochondrial reactive oxygen species production is related to NOX-independent NET formation (5, 14, 15). Therefore, first we verified the importance of mROS for NOX-independent NET formation by performing MitoSOX (a specific fluorescent probe for mROS detection) and SYTOX Green assays in the presence of an mROS scavenger (MitoTEMPO). Purified neutrophils were incubated with 200 µM MitoTEMPO for 15 min under physiologic pH conditions (7.4), and stimulated with media (negative control), A23187, or ionomycin. MitoTEMPO decreased mROS generation induced by both stimuli (Figures 4A,B). SYTOX Green assays showed that NET formation was also significantly decreased in the presence of mROS scavenger, confirming that mROS production is a key factor for NOX-independent NET formation (Figures 4C,D). Using PMA (a prototypical agonist for the NOX-dependent NET formation), we confirmed that mROS does not play a major role in NOX-dependent NET formation (Figure S4 in Supplementary Material).

Since calcium influx plays an important role in the mROS production (16–19), we asked whether increasing pH could affect the mROS production. To measure mROS, neutrophils were treated with 4 µM MitoSOX, and stimulated either with media control (negative control), A23187 or ionomycin in three different pH conditions (pH 6.6, 7.2, and 7.8). mROS fluorescence in these cells was measured every 4 min up to 120 min. Within 30 min, we found that neutrophils resuspended in higher pH media generated higher amount of mROS after stimulation with either A23187 or ionomycin (Figures 5A–F). Analysis of the MitoSox data after 120 min of stimulation indicates that resting neutrophils also generate mROS at higher pH, albeit lower levels than measured in calcium ionophore-stimulated neutrophils (Figure S5 in Supplementary Material). We also measured the MitoTEMPO-mediated suppression of mROS production measured by MitoSox and NET formation measured by three other pH conditions. These data suggest that increasing pH increases mROS generation and scavenging mROS suppresses NOX-independent NET formation (Figures S6 and S7 in Supplementary Material for the pHs 6.6, 7.2, and 7.8). Collectively, increasing pH increases mROS generation during the activation of neutrophils with calcium ionophores, and pH-dependent increase in mROS generation is an important component for promoting NOX-independent NET formation.

PAD4 activation is a key step in NOX-independent NET formation. Once bound to cytosolic-calcium, PAD4 is activated and translocated to nucleus (6, 20, 21). The translocated PAD4 deiminates positively charged arginine present on histones 3 into non-charged citrulline, helping chromatin decondensation (22, 23). This step is especially relevant to NOX-independent NET formation (5); NET images clearly showed a strong histone 3 citrullination. To determine the influence of the pH in NET formation, we performed a confocal immunofluorescence microscopy of PAD4 and citrullinated histone 3. Purified neutrophils were seeded in a chamber slide, stimulated with buffer, A23187, or ionomycin up to 30 min, fixed and stained the cells with monoclonal antibodies to PAD4, citrullinated histone 3, and DNA dye. Neutrophils activated with A23187 or ionomycin, showed increased immunostaining for PAD4 and citrullinated histone 3 in higher pH conditions (Figures 6A,B; Figure S8 in Supplementary Material for negative controls and Figure S9 in Supplementary Material for isotype controls). Furthermore, we performed confocal microscopy of the immunostained neutrophils to determine citrullination of histone 3 in different pH conditions. Images showed a strong co-localization of MPO, DNA, and citrullination of histone 3, in neutrophils incubated in higher pH, confirming that A23187 or ionomycin-stimulated neutrophils have a greater amount of citrullination of histone 3 and formed a greater amount of NETs with increasing pH (Figure 7; Figures S10–S12 in Supplementary Material for single channels and Figure S13 in Supplementary Material for isotype control). Therefore, increasing pH increases CitH3 formation that facilitates NOX-independent NET formation.

Histone 4 cleavage is another hallmark event of NET formation; therefore, we sought to determine whether the pH levels affect this process. The cells were seeded in three different media conditions (pH 6.6, 7.2, or 7.8) with media (negative control) or calcium ionophores (A23187 or ionomycin), and after 1 h the H4 cleavage was determined by Western blot analysis. Unstimulated controls had little or no histone cleavage, whereas both calcium ionophores promoted substantial histone cleavage. The pH-dependent effect was much clear for ionomycin condition (Figure 8). Therefore, increased cleavage of histone is another factor that could promote NET formation at higher pHs, particularly during the activation of neutrophils by calcium ionophores.