DNA Is an Antimicrobial Component of Neutrophil Extracellular Traps

Summary

Comprising the first line of the innate immune response, neutrophils combat infectious microorganisms through the release of toxic molecules, phagocytosis of invaders and the production of the recently characterized neutrophil extracellular traps (NETs). The antimicrobial activity of NETs has been attributed to proteins bound to the DNA backbone. Our results demonstrate that the DNA lattice of each NET is potently antibacterial and elicits upregulation of protective surface modifications by the opportunistic bacterial pathogen Pseudomonas aeruginosa. These modifications, previously shown to protect bacteria from antimicrobial peptides, confer greater bacterial tolerance to DNA and NET-mediated antibacterial activity. Treatments that quench the cation chelating capacity of DNA restore bacterial viability and suppress the expression of surface modifications even in the presence of intact NETs. These observations highlight the dual function of DNA as an antibacterial component of NETs, but also a signal perceived by bacteria to induce broad host resistance strategies. Therefore, the ability of P. aeruginosa to sense and defend against the antibacterial activity of neutrophil extracellular traps may contribute to long-term survival in chronic infection sites including the Cystic Fibrosis lung.

Introduction

Neutrophils are central mediators of the innate immune defense system and perform their role by killing invading microbes through phagocytosis, degranulation, and the release of neutrophil extracellular traps (NETs) [1, 2]. The scaffold of NETs is composed of genomic DNA, which is enmeshed with antimicrobial proteins normally found in the nucleus, granules, or cytoplasm of neutrophils [1, 2]. Although largely characterized ex vivo using purified human neutrophils and the chemical inducer phorbol-12-myristate 13-acetate (PMA), the process of NETosis resulting in the generation of NETs has been observed in vivo during infection where these structures function to trap bacteria, fungi, protozoa and viruses [1–5]. The mechanism by which NETs kill microbial invaders remains controversial [5–7]. Given the detection of known antimicrobial proteins that decorate the genomic lattice structure [1, 2, 8, 9], current models describing the antimicrobial function of NETs focus on the role of NET-bound proteins. However, most of these proteins are present in low abundance and evidence of their antimicrobial function while bound to the NET structure is limited to a few proteins [1, 8–11]. NET-bound calprotectin is a zinc-chelating protein with antimicrobial activity against Candida and Klebsiella that can be neutralized with excess zinc [3, 9, 10]. Histones, the most abundant NET-bound proteins [10], possess direct membrane-acting antibacterial activity [12, 13] and were shown to contribute to killing of Staphylococcus and Shigella [1]. Cathepsin G, a granular serine protease, is required for the clearance of Neisseria by NETs [8]. To demonstrate the antibacterial contribution of the latter two NET-bound proteins, antibodies raised to histones or cathepsin G were shown to limit the bactericidal capacity of NETs towards these pathogens [1, 8].

Immunocompromised individuals, including those with Cystic Fibrosis (CF), are particularly susceptible to P. aeruginosa infection, which is a major cause of morbidity and mortality in these patients. Chronic P. aeruginosa infection of the CF lung leads to an intense inflammatory immune response, resulting in the recruitment of large numbers of neutrophils to the site of infection [14, 15]. CF sputum is highly enriched in neutrophil-derived DNA, including that of NET origin, indicating that neutrophils deploy NETs in an effort to combat infection of the lung [16, 17]. However, persistence of P. aeruginosa in the midst of sustained neutrophil presence and NETosis suggests that the pathogen is capable of evading this host immune response [18].

An important virulence strategy adopted by successful microbial pathogens is the tolerance of NETs. In a number of cases described so far, microbial invaders accomplish this goal by either avoiding or disarming neutrophil extracellular traps. Modification of the bacterial capsule and surface-localized lipoteichoic acid reduces the trapping of Streptococcus pneumonia in NETs [19]. The secretion of extracellular nucleases by Staphylococcus aureus, Streptococcus pneumonia, group A Streptococcus and Vibrio cholerae highlight a shared virulence strategy that functions to degrade the DNA-backbone of NETs, enabling evasion or liberation of the bacteria from entrapment [2, 20–22].

Although microbial or exogenous DNase is proposed to dissolve NET structures to avoid capture, it has not been considered that the DNA backbone itself may be antimicrobial. We have previously demonstrated that extracellular DNA is an efficient chelator of divalent metal cations [23]. This cation chelation has pleiotropic effects on P. aeruginosa depending on the concentration of extracellular DNA. At subinhibitory levels, sequestration of cations by DNA leads to the expression of genes controlled by the two component systems PhoPQ and PmrAB that sense Mg²⁺ limitation [23–25]. However, at higher concentrations, extracellular DNA causes dramatic disruptions to the bacterial envelope integrity, which leads to lysis and rapid cell death [23]. It is predicted that the phosphodiester backbone is required for cation sequestration of metal cations on the bacterial surface and the membrane-destabilizing antimicrobial activity of extracellular DNA [23]. Therefore, we hypothesize that the DNA backbone of NETs contributes to their antibacterial activity. Here we show that neutralizing the membrane destabilizing activity of extracellular DNA by quenching the capacity of the phosphodiester backbone to chelate cations protected bacteria from NETs. P. aeruginosa is capable of detecting the DNA lattice of NETs, and in response, upregulates genes required to modify the bacterial outer membrane surface to tolerate the toxic effects of DNA-mediated NET killing.

Results

Pseudomonas aeruginosa induces and tolerates NETosis

P. aeruginosa has been shown to induce the formation of neutrophil extracellular traps in purified human neutrophils [18] and NETs have been observed in CF sputum [16, 17], where P. aeruginosa is a predominant pathogen. PMA-induced NETs contained known neutrophil proteins embedded in the DNA lattice, including myeloperoxidase (MPO) and histones [2] (Fig. 1A). We also show that Gfp-tagged P. aeruginosa is efficiently trapped and aggregated in NETs (Fig. 1A). However, in vivo NETosis during P. aeruginosa infection has not been reported. Therefore intravital confocal microscopy was used to determine whether P. aeruginosa elicited NETosis in the mouse skin infection model [26]. Infection with P. aeruginosa led to the production of large NET-like structures that stained with the DNA-binding dye Sytox green and entrapped ChFP-labeled P. aeruginosa (Fig. 1B and C). In addition to the presence of NETs, neutrophils remained chemotactic and phagocytosed bacteria, suggesting that multiple neutrophil clearance mechanisms are employed in vivo to combat P. aeruginosa (Fig. 1D and S1 Movie).

Given that P. aeruginosa induces the production of NETs in vitro and in vivo (Fig. 1), we sought to compare the relative abilities of P. aeruginosa, S. aureus, E. coli and the chemical inducer PMA to elicit NETosis. In the presence of purified human neutrophils, PMA, E. coli and S. aureus induced significantly more NET formation, relative to P. aeruginosa within 1 hour of coincubation (Fig. 2A). However, at later time points (3h), P. aeruginosa elicited the formation of similar amounts of NET structures. Furthermore, quantification of the number of NETs and NET-area using the skin infection model confirmed that P. aeruginosa weakly induces NETosis relative to S. aureus in vivo (Fig. 2B and C).

One possible explanation for the reduced NET formation in the presence of P. aeruginosa is the production of a microbial secreted DNase that degrades NETs more efficiently than other organisms. Therefore, we measured the DNase activity in overnight culture supernatants, as well as in supernatants isolated from coincubating bacteria with PMA-treated neutrophils. We demonstrate that there was little, if any, DNase activity produced by P. aeruginosa, S. aureus or E. coli under these conditions (S1 Fig.). Since cations are a requirement for DNase activity, we were able to restore DNase activity in S. aureus supernatants after the addition of excess cations (S1 Fig.). However, under the cation-free conditions used to quantitate antibacterial NET function, even in the presence of PMA-stimulated neutrophils, DNase activity was not detected and thus P. aeruginosa appears to limit NETosis through an uncharacterized mechanism.

In order to characterize the bactericidal capacity of NETs, neutrophils were treated with PMA to stimulate maximal NETosis and cytochalasin D to block phagocytosis, thus restricting bacterial killing to extracellular NET function [1, 8, 21, 22]. Importantly, the addition of cytochalasin D had no effect on NETosis induced in PMA-treated neutrophils (S2 Fig.). We used the conventional method of direct bacterial counts to enumerate the number of bacteria before and after challenge with PMA-induced NETs. Direct counts of NET-exposed bacteria revealed that P. aeruginosa was most tolerant to NET killing, whereas S. aureus and E. coli were significantly more sensitive (Fig. 3A). The addition of deoxyribonuclease (DNase) restored bacterial survival of the NET-sensitive organisms E. coli and S. aureus, confirming that killing was mediated by extracellular NET function (Fig. 3A). Further, the kinetics of bacterial killing by PMA-generated NETs was determined by measuring the loss of luminescence from chromosomally-tagged luminescent P. aeruginosa strain, PAO1::p16Slux [23], and plasmid-borne luminescent E. coli / pσ70-lux [27]. This approach confirmed that P. aeruginosa was more tolerant to NET killing than E. coli, where luminescence rapidly decreased upon neutrophil challenge (Fig. 3B).

Direct contact and the phosphate backbone are required for the antimicrobial, cation chelating activity of extracellular DNA

The addition of exogenous DNase I and the production of secreted DNases by bacteria are the most effective means to disable NET killing [1, 2, 20, 21]. It is thought that DNase treatment dissolves NET structures, thereby releasing and diluting the antimicrobial proteins bound to NETs. We have previously shown that extracellular DNA has a potent antibacterial activity, as purified salmon DNA (2% w/v) causes several log orders of bacterial killing within minutes and breaks the integrity of both the inner and outer membranes, leading to lysis [23]. DNA is a very efficient cation chelator and the antimicrobial activity of DNA can be blocked with addition of excess divalent metal cations [23]. It is predicted that the cation chelation is mediated by the phosphodiester backbone.

To confirm the mechanism by which extracellular DNA kills bacteria, we monitored the loss of P. aeruginosa viability in the presence of dilute extracellular DNA (0.125% w/v; Fig. 4A). Bacterial survival was restored if DNA was pretreated with DNase or excess 5mM Mg²⁺, which degrades DNA or saturates its cation chelating ability, respectively, thus neutralizing the antibacterial activity (Fig. 4A). Pretreatment of extracellular DNA with calf intestinal alkaline phosphatase (PTase), which cleaves 5’-phosphates, also blocked the observed antibacterial activity (Fig. 4A). The addition of decreasing amounts of DNase, PTase or Mg²⁺, resulted in marked, dose-dependent, decreases in bacterial survival when challenged with 0.15% DNA (Fig. 4B). The DNA-mediated damage to the outer membrane leads to the formation of ChFP-enriched outer membrane vesicle-like structures (OMVs; Fig. 4C) [23]. However, incubation of P. aeruginosa with extracellular DNA pretreated with DNase, PTase and Mg²⁺ greatly diminished the number of ChFP-labeled OMVs relative to control conditions (Fig. 4D), confirming that the bactericidal mechanism of extracellular DNA is through disruption of the bacterial membrane.

Extracellular DNA-mediated damage to membrane integrity was confirmed by using flow cytometry (Fig. 4E). Treatment of P. aeruginosa with DNA resulted in a new population of cells that were dual positive for SYTO9 and PI, indicative of membrane damage and increased PI uptake, and possibly dead cells. The increased staining of DNA-exposed bacteria by membrane-impermeable propidium iodide (PI) was not observed in DNase and Mg²⁺ pretreatments and was reduced in PTase pretreated DNA samples (Fig. 4E and F). To ensure cation chelation was responsible for the observed bacterial membrane destabilization, we assessed whether the known cation chelator EDTA could cause membrane disruption. Like DNA, EDTA caused major outer membrane disruptions and the release of OMVs, as well as a dramatic increase in PI-staining of EDTA-treated cells monitored by flow cytometry (S3 Fig.) [28]. To determine whether the antibacterial capacity of DNA requires direct bacterial contact or can be mediated through passive cation sequestration, we exposed P. aeruginosa to high concentrations of DNA spatially separated by an ion-permeable barrier. Blocking direct interaction between P. aeruginosa and DNA resulted in bacterial survival after a prolonged exposure, compared to the rapid antibacterial activity of DNA in direct contact with the bacteria (S4 Fig.). Together, these results demonstrate that the antibacterial activity of extracellular DNA requires direct contact, and the phosphate backbone for cation chelation, leading to membrane disruption and bacterial cell death.

Bactericidal activity of NETs is blocked by neutralizing the cation chelation capacity of DNA

The bactericidal activity of neutrophil extracellular traps is attributed to direct contact and exposure of bacteria to the antimicrobial proteins embedded in the DNA scaffold of NETs [1, 2]. Given the antimicrobial activity of DNA, we propose that the DNA backbone of the NET itself is antibacterial. Therefore, if DNA contributes to bacterial killing, treatments that quench the cation chelation potential of the DNA backbone will block bactericidal activity of NETs. To address this possibility, PMA-activated neutrophils were treated with the addition of DNase, PTase or excess Mg²⁺ and bacterial viability was monitored. The DNA-targeted treatments completely protected P. aeruginosa and E. coli from killing by neutrophil extracellular traps (Fig. 5A). To confirm these results, we monitored the luminescence of P. aeruginosa PAO1::p16Slux co-incubated with PMA-activated neutrophils and observed that the antibacterial effects of NETs were neutralized by treatment with exogenous DNase, Mg²⁺ cations and PTase (Fig. 5B).

To determine whether the restored bacterial viability in the presence of NETs was due to preventing damage to the bacterial envelope, we performed flow cytometry to assess membrane integrity. Increased PI staining of NET-exposed bacteria is an indicator of membrane damage, which was completely blocked by addition of DNase (Fig. 5C). The addition of excess Mg²⁺ or treatment with exogenous PTase also limited membrane damage (Fig. 5C). Importantly, both Mg²⁺ and PTase treatments neutralized the antimicrobial activity of NETs (Fig. 5A and B) did not disrupt overall NET architecture, as MPO and histones were still present within the treated NET structures (Fig. 6). To assess the function of a NET-bound protein, we measured elastase activity in PMA-treated neutrophils and showed no difference in elastase activity when NETs were treated with exogenous PTase or Mg²⁺ (S5 Fig.). NET structures remain intact and contain functional proteins (elastase) after treatment with PTase or excess Mg²⁺, but are no longer antibacterial for E. coli and P. aeruginosa (Fig. 5). Together these results suggest an antibacterial mechanism wherein the DNA backbone of NETs target and destabilize the bacterial membrane and promotes cell death.

Extracellular DNA elicits induction of surface modifications that protect P. aeruginosa from NETs

Subinhibitory concentrations of extracellular DNA sequester Mg²⁺ and trigger the expression of multiple surface modifications that are known to protect the bacterial outer membrane from antimicrobial peptide (AP) damage and killing [23–25]. The arn operon (PA3552-PA3559) is required for the covalent addition of aminoarabinose to the phosphates of lipid A and the spermidine synthesis genes (PA4773-PA4774; speDE homologs) are required for production of the polycation spermidine on the outer surface [23, 24]. Both modifications substitute for divalent metal cations, mask the negative charges of the outer surface and thus contribute to AP resistance [23, 24, 29–31]. Given that these modifications stabilize the bacterial envelope, we sought to determine whether these surface modification pathways provided a more general mechanism to resist bacterial membrane damage. We noted that P. aeruginosa strains with mutations in the arn or spermidine biosynthetic pathways were significantly less capable of surviving exposure to DNA (Fig. 7A).

Given the role of these pathways for tolerance of exogenous DNA, we then investigated whether the DNA component of NETs induced expression of the arn or spermidine synthesis genes in P. aeruginosa. Expression of both pathways was strongly induced 2–6 fold following co-incubation with NETs produced by PMA-activated neutrophils (Fig. 7B). To confirm that DNA was the component of NETs that led to induction of the bacterial gene expression response, the addition of excess Mg²⁺ cations and enzymatic treatment with DNase (Fig. 7C) and PTase (S6 Fig.) all blocked the induction of the arn and spermidine operons response to NETs. While these treatments specifically neutralize DNA, we also considered the possibility that NET-bound antimicrobial proteins including histones or LL-37 may elicit these protective responses. We have previously shown that sub-MIC concentrations of antimicrobial peptides induce both outer surface modifications [29]. Therefore, to assess the relative capacity of each NET component to act as a bacterial signal, we compared the ability of purified histones, the well-characterized APs polymyxin B and colistin, and DNA to induce the expression of the spermidine synthesis pathway. Although all NET components induced expression of the PA4773-PA4774 spermidine synthesis pathway (S7 Fig.), DNA was the most potent inducer of this bacterial response (S7 Fig.).

The upregulation of protective outer membrane modifications (aminoarabinose-modified LPS and surface spermidine production) by NETs, and by the individual NET components of DNA and histones, suggests that these modifications are required to defend the membrane against assault from multiple innate immune components that target the bacterial membrane. Both modifications result in stable substitutions for divalent metal cations in the outer membrane, and protect P. aeruginosa from antimicrobial peptides [23, 24, 29–31] and DNA killing (Fig. 7A). P. aeruginosa mutants in the arn and spermidine synthesis genes also exhibited increased susceptibility to the disruptive effects of NETs (Fig. 7D). We next compared the susceptibilities of P. aeruginosa, E. coli and S. aureus to histone and DNA killing. Surprisingly, P. aeruginosa was the most susceptible to DNA killing, while S. aureus was the most tolerant, the opposite pattern of NET susceptibility (Figs. 3A and 8A). The bactericidal capacity of purified histones was modest, where after 2 hours exposure P. aeruginosa was the more histone tolerant (Fig. 8B). Taken together, these results highlight the ability of P. aeruginosa to quickly respond and defend against the DNA and histone mediated-antibacterial effects of NETs by stabilizing the outer membrane.

Discussion

Given the observations that exogenous or secreted microbial DNases protect bacteria against NET killing and that extracellular DNA has rapid, membrane-damaging antibacterial activity, we sought to test the hypothesis that the DNA backbone of NETs contributes to their bactericidal function. Extracellular DNA possessed contact-mediated antibacterial activity that could be neutralized by enzymatic and cationic treatments that degrade or quench the capacity of DNA to chelate cations (Fig. 4 and S3 Fig.). NETs exposed to the same treatments that target the DNA scaffold were unable to cause bacterial membrane damage or to cause bacterial killing of P. aeruginosa and E. coli (Fig. 5). Therefore, we propose a novel bactericidal mechanism of NETs whereby the removal of surface-stabilizing cations by the DNA phosphodiester backbone results in bacterial lysis (Fig. 4B and C). To address the controversy surrounding the bacterial killing ability of NETs [5–7], we used multiple viability assays to measure NET killing and membrane damage, which included direct bacterial counts, luminescence viability assays and flow cytometry of PI-stained cells [32]. Taken together, these data support the general notion that NETs are directly antimicrobial.

Deciphering the specific antimicrobial mechanisms of NETs has been limited to a few candidate proteins [1, 8–11]. Most studies have focused on the NET-bound proteins as the antimicrobial components, given their important role during phagocytosis and degranulation. Although granular, cytoplasmic and nuclear proteins derived from neutrophils can be detected in NETs by immunofluorescence, the abundance of most NET-bound proteins is low (<1–6%), relative to histones, which comprise 65% of the total protein content [10]. Although classically characterized as chromatin structural proteins, NET-bound histones possess antimicrobial activity that can be neutralized through the addition of anti-histone antibodies [1, 12, 13]. However, other neutrophil proteins exhibit altered or reduced enzymatic activity when enmeshed in the NET backbone raising questions as to their antimicrobial capacity. For example, S. aureus tolerates myeloperoxidase in NETs unless supplemented with the addition of exogenous H₂O₂ [11]. Neutrophil elastase activity increases in DNase treated sputum from Cystic Fibrosis patients, suggesting that DNA may inhibit elastase activity, thus limiting the role of elastase as a potential NET-bound factor [17].

We observed that both the antibacterial activity of NETs and the ability to induce protective bacterial responses were blocked by treatments that target extracellular DNA, suggesting that the NET scaffold is not simply a passive structure (Figs. 5 and 7) [23–25, 31]. The most potent inducing triggers of the P. aeruginosa surface modifications are purified eDNA, followed by APs and purified histones (S7 Fig.). Since being widely induced by these components, it is not surprising that the aminoarabinose-modified LPS and spermidine synthesis pathways protect the outer membrane from DNA, NETs (Fig. 7) and antimicrobial proteins [23, 24, 29–31]. The low potency of the tested histones may be explained by the fact that our assays were performed with a mixture of full-length histones, which had modest antibacterial activity (Fig. 8). Recent evidence highlights that histones are proteolytically processed by proteases such as elastase during the process of nucleus decondensation, prior to NET release [33] It is therefore likely that potent bactericidal histone-derived peptides are present in NETs as an important antibacterial component of NETs.

P. aeruginosa appears to mount a multifunctional, outer membrane defense strategy to combat multiple antimicrobial components enmeshed in NETs. Consistent with this model, we noticed unexpected susceptibility patterns that also suggest that NET killing may be the result of the combinatorial effect of DNA and NET-bound proteins. The observation that S. aureus is susceptible to NET killing but tolerant to DNA (Figs. 3 and 8), suggests that NET killing of S. aureus is likely dependent on other anti-staphylococcal proteins enmeshed in the DNA lattice. The DNA susceptibility phenotype of P. aeruginosa may explain the potency of DNA as the strongest inducer of the protective outer surface modifications that contribute to the observed NET tolerance. It is intriguing to speculate that the NET-bound, antimicrobial proteins act in concert with the antibacterial activity of DNA to provide broad-spectrum protection against a wide range of microbial pathogens.

Modification of the bacterial cell surface and the production of secreted DNases are virulence strategies utilized by microbial pathogens to evade NET killing [19–22]. Here we report that the spermidine and the arn surface modification pathways are required to tolerate the antibacterial action of both DNA and NETs (Fig. 7). The covalent addition of aminoarabinose to the lipid A component of LPS masks the negative charges of core LPS phosphates, and the polycationic nature of spermidine (+3 charge) substitutes for surface divalent metal cations, and may also bind and neutralize DNA. In addition, spermidine possesses an antioxidant activity that protects bacterial membrane lipids from oxidative damage [24] and therefore may protect P. aeruginosa from NET-induced oxidative damage [11]. Combined, these results suggest that the spermidine and arn surface modifications possess multiple protective roles that may contribute to resisting a broad range of antimicrobial components present within NETs.

We propose that bacterial surface-bound, divalent metal cations are displaced by direct contact with extracellular DNA, and that DNA-induced surface modifications prevent outer membrane disruption and bacterial killing by NETs. Therefore, the antibacterial mechanism of cation chelation exerted by DNA is distinct from that of other previously characterized antimicrobial cation chelating proteins such as calprotectin. We have previously demonstrated that DNA chelates diverse metal cations (Mg²⁺, Ca²⁺, Zn²⁺, Mn²⁺) [23] while calprotectin chelates zinc and manganese [34]. Additionally, the antimicrobial function of calprotectin is contact-independent whereas the bactericidal function of DNA requires contact (S4 Fig.). Further, sequestration of zinc and manganese by calprotectin does not target the microbial membrane but rather sequesters cation cofactors required by bacterial enzymes such as superoxide dismutase, which protects bacteria from superoxide [34].

In summary, we have identified that the DNA backbone is a bona fide antibacterial component of neutrophil extracellular traps. The DNA scaffold structure also acts as a warning signal perceived by P. aeruginosa. Overall, these results support a model where the membrane-destabilizing activity of the DNA scaffold contributes to the bactericidal capacity of NETs, while the cation chelating activity acts as a signal perceived by P. aeruginosa that leads to upregulation of protective surface modifications. These results highlight a dynamic bacterial-host interaction between an opportunistic pathogen that causes chronic infections in the lungs of individuals with Cystic Fibrosis, an infection site known to be rich in neutrophil DNA and neutrophil extracellular traps [16, 17]. This ability to sense and defend against NETosis may help explain the long-term persistence of P. aeruginosa in CF lung infections.

Materials and Methods

Bacterial strains and growth conditions

All strains and plasmids used in this study are shown in S1 Table. Bacterial cultures were routinely grown at 37°C in LB or BM2 defined minimal media with 0.5 mM MgSO₄, unless otherwise stated. S. aureus was grown overnight in BHI media. When necessary, the following antibiotics were used: 50 µg/mL tetracycline for P. aeruginosa mini-Tn 5-lux mutants, and 50 µg/mL kanamycin for E. coli DH5α/ pσ70-lux. Mid-log cultures were used for co-incubation experiments with neutrophils or extracellular DNA.

Human neutrophil isolation

Neutrophils were isolated from healthy donors as previously described [35]. Whole blood was collected and mixed 5:1 in acid citrate dextrose, followed by removal of red blood cells using dextran sedimentation and hypotonic lysis with KCl. After all red blood cells were lysed, the cell pellet was subjected to Ficol-Histopaque density centrifugation. The subsequent pellet was resuspended in 2 mL of HBSS (Hank’s balanced salt solution, no cations; Invitrogen 14175–095). The viable cell concentration was determined using a haemocytometer and Trypan blue staining.

NET imaging

Glass cover slips were HAS-coated and placed in 6-well tissue culture plates. Neutrophils were added at 2.0×10⁶ cells/mL per well, adhered (30 min, 5% CO₂, 37°C) and treated with cytochalasin D (10 µg/well) and PMA (25 nM) to activate NETosis [35]. Mid-log bacterial cultures were diluted in HBSS (no cations) (5.0x10⁷ CFU/mL) for an MOI of 25:1, centrifuged to the neutrophils, and coincubated for 1–4 hours (5% CO₂, 37°C). Cells on cover slips were fixed with 4% paraformaldehyde, washed with 250 µL of 10% FBS (Invitrogen) in PBS and stained with either DNA dyes and/or various primary and secondary antibodies (described below).

Immunofluorescence microscopy of NETs

For NET visualization with antibodies, the primary anti-human MPO antibody (DakoCytomation- A0398) was diluted into 10% FBS in PBS (1/500). 30 µL was added to adhered neutrophils, incubated (30 min, 37°C) and washed twice with sterile PBS. 40 µL of the secondary anti-rabbit Cy 5 antibody (Jackson ImmunoResearch 60354) (1/500 dilution) was added. After 15 min incubation in the dark, cover slips were washed twice with PBS, and prepared with mounting media. Anti-DNA and anti-histone antibodies where obtained from Dr. Marvin Fritzler. Either the anti-DNA (1:10) or anti-histone (1:500) antibodies [36, 37] were added as described above. The anti-human secondary antibodies with Alexa Flour 647 (Invitrogen A21445, 1/500) were added to cover slips and mounted as described above. Images of human NETs were acquired using the Leica DMI 4000B inverted microscope equipped with ORCA R2 digital camera and Metamorph software for image acquisition using the 63X or 100X objectives. The following excitation and emission filters were used for blue fluorescence (Ex 390/40; Em 455/50), red fluorescence (Ex 555/25; Em 605/52), far red fluorescence (Ex 645/30; Em 705/72) and green fluorescence (Ex 490/20; Em 525/36). Images were formatted and analyzed using the Imaris 7.0.0 imaging software. All images shown are representative of at least three experiments.

Mouse skin infection model

Mice were anaesthetized (10 mg/kg xylazine hydrochloride and 200 mg/kg ketamine hydrochloride) and body temperature was maintained using a rectal probe and heating pad. The mice were pretreated with intradermal MIP-2 (0.2µg/injection) diluted in sterile normal saline 30 minutes prior to imaging. The right jugular vein was cannulated to administer additional anesthetic and fluorescent dyes. The microcirculation of the dorsal skin was prepared for microscopy as previously described [26]. Briefly, after shaving the mouse’s back, a midline dorsal incision was made extending from the tail region up to the level of the occiput. The skin was separated from the underlying tissue, remaining attached laterally to ensure the blood supply remained intact. The area of skin was then extended over a viewing pedestal and secured along the edges using 5.0 sutures. The loose connective tissue lying on top of the dermal microvasculature was carefully removed by dissection under an operating microscope. The exposed dermal microvasculature was immersed in isotonic saline and covered with a coverslip held in place with vacuum grease. Alexa Fluor 649 conjugated anti-mouse GR-1 antibody (10µl per mouse i.v.; eBioscience) was used visualization of neutrophils. To visualize NETs in vivo the membrane impermeable dyes SYTOX-green or SYTOX-orange were administered (diluted 1:1000 with sterile saline, 100µl per mouse i.v.). MIP-2 injection (0.2µg/injection) was initiated 30 min prior to Pseudomonas aeruginosa or S. aureus administration. Following baseline visualization, all bacteria were directly administered into the field of view using a tuberculin needle (1×10⁸ CFU/100µl of sterile saline, i.d.).

Intravital microscopy

Spinning disk confocal intravital microscopy was performed using an Olympus BX51WI (Olympus, Center Valley, PA) upright microscope equipped with a 20×/0.95 XLUM Plan Fl water immersion objective. The microscope was equipped with a confocal light path (WaveFx, Quorum, Guelph, ON) based on a modified Yokogawa CSU-10 head (Yokogawa Electric Corporation, Tokyo, Japan). Laser excitation at 488, 561 and 649nm (Cobalt, Stockholm, Sweden), was used in rapid succession and fluorescence in green, red and blue channels was visualized with the appropriate long pass filters (Semrock, Rochester, NY). Exposure time for all wavelengths was between 500 and 600ms. Sensitivity settings were maintained at the same level for all experiments. A 512×512 pixels back-thinned EMCCD camera (C9100–13, Hamamatsu, Bridgewater, NJ) was used for fluorescence detection. Volocity Acquisition software (Improvision Inc., Lexington, MA) was used to drive the confocal microscope. Images captured using the spinning disk were processed and analyzed in Volocity 6.0.1. NET area and NET number were quantified using the Volocity software.

Quantification of NET area and number

NET area was determined using Volocity imaging software. Briefly, in each field of view (FOV) the threshold of the corresponding fluorescent channel in which NET structures were stained was set to eliminate the background staining of the skin. The area and number of NET positive structures in the FOV was calculated and counted via the Volocity 6.0.1 software. Structures that showed no characteristic NET-like shape and resembled the staining of a nucleus of an obvious dead cell in the FOV were excluded from the quantification manually. NET image analysis was performed in at least two infected or uninfected animals and from 5 fields of view.

NET quantification from purified human neutrophils

NETs were quantitated by measuring the amount of extracellular DNA that stains with the cell impermeant dye Styox Green [35]. Cell culture media (CCM) consisted of 48.5 mL of RPMI 1640 (Invitrogen), 0.5 mL of 1.0 M HEPES and 1.0 mL of human serum albumin (HAS; Innovative Research). Neutrophils were diluted into CCM (2.0×10⁵ cell/well) and added to an HSA-coated, 96-well black, clear-bottom plate (Thermo Scientific). As a positive control, PMA (25 nM/well) was added to activate the neutrophils [35]. For bacterial activation of NETs, mid-log bacterial cultures were diluted in CCM (2.0×10⁶ CFU/well) for a multiplicity of infection (MOI) of 10:1 (bacteria to neutrophils). DNase (430 kU/well; VWR 31149) was added to degrade extracellular DNA in NETs. Bacteria were gently centrifuged onto the adhered neutrophils (800xg, 10 min). Sytox green (2.5 µM; Invitrogen) was added to each well and green fluorescence (Ex 490/8; Em 535/25) was measured with Perkin Elmer 1420 Multilabel Counter Victor³ between 1 and 4 hours. All values shown are the mean from at least three individual replicates and each experiment was performed at least 3 times. We noticed variation in the background level of NET staining between individual donors, but there was a reproducible and robust 2.5 to 10-fold increase in NETosis after PMA treatment of neutrophils from various donors (S2 Fig.).

Quantification of Bacterial Viability and Gene Expression Using Plate Counts and Luminescence NET killing was examined using direct plate counting methods where a reduction in cell number indicated bacterial killing. All NET killing experiments were performed in HBSS solution lacking divalent cations. Isolated human neutrophils were mixed with mid-log bacteria in HBSS (no cations) in black, clear-bottom 96-well plates with of 2.0 × 10⁷ CFU bacteria and 2.0 × 10⁶ neutrophils (MOI 10:1). After a 1–4 hr incubation, 50 µL of DNase I solution (430 kU/mL) was added to every well, mixed, and incubated for 30 min at 37°C, in order to release bacteria trapped in NETs for accurate plate counts. 15 µL of suspension was serially diluted (1/10) in 0.9% NaCl solution in a sterile 96-well plate and 5 µL from each well was stamped onto LB agar plates to obtain bacterial plate count data for time zero (T₀) and after 4h (T₄). CFU/ml values from T₄ and T₀ time points were used to calculate the percentage survival by subtracting the T₄—T₀ plate counts and dividing the ‘bacteria and neutrophil’ conditions by the ‘bacteria alone’ conditions and multiplying by 100. For lux viability and gene expression assays [23], bacteria were centrifuged onto the adhered neutrophils and placed in the Victor³ plate reader for luminescence (CPS) measurements every 20 minutes for 3–4 hours. All values shown are the mean from at least six individual replicates and each experiment was performed at least 3 times.

Flow cytometry analysis of SYTO9/propidium iodide stained bacteria

SYTO9 stains the DNA in all cells and propidium iodide (PI) stains the DNA in dead cells and cells with damaged membranes [28, 32]. The sample of bacteria and neutrophils (~200 µL) was placed in 5 mL polystyrene round-bottom sample tubes and stained with SYTO9 and propidium iodide at final concentrations of 0.02 mM and 0.2 mM, respectively. The tubes were centrifuged at 300x gravity and incubated (RT, 15 min). Bacterial cells were analyzed using the BD LSRII flow cytometer (BD Bioscience, San Jose, USA) equipped with a blue laser (488nm) and a green laser (532nm). Unstained, mid-log bacterial cells were used to gate the forward scatter (FSC) and size scatter (SSC) parameters. For green and red fluorescence profiles, SYTO9 was excited by blue (488nm) laser with emission filters 525/50BP and 505LP and PI was excited using the green (532 nm) laser with emission filters 610/20BP and 600LP. All detectors were set to the logarithmic amplification with the following voltages, 500, 240, 596, and 489 and threshold was set at 200 for both FSC and SSC. For each sample, 50 000 events were acquired using the BD FACSDiva software 6.1.3. The Hierarchical gating strategy was used to determine double positive population of bacterial cells (stained with both SYTO9 and PI) where gate P1 is the total population of FSC and SSC gated events, as determined from bacteria alone control and then applied to all other samples. P2 is the population of events stained by SYTO9 and P3 is the population stained with both SYTO9 and PI. Neutrophils and mid-log bacteria controls do not contribute any autofluorescence or PI-stained events when stained with either or both of the SYTO9/PI dyes. Values displayed in each density plot represent the percentage of 50 000 cells (N value) in each quadrant gate and each experiment was performed with at least 5 times.

DNase, PTase or Mg²⁺ treatment of NETs

During the coincubation experiments of bacteria and PMA-activated neutrophils, exogenous Mg²⁺ was added at a final concentration of 5 mM MgSO₄. For the enzyme treatments, deoxyribonuclease (DNase I, VWR) was added at a final concentration of 430 kU/well and calf intestinal alkaline phosphatase (PTase, Invitrogen) at a final concentration of 16.6 U/well. Maximum enzyme amounts were added to bacterial-neutrophil mixtures that had no effect on bacterial viability and without the addition of enzyme buffers. The killing experiments were incubated for up to 4 hours in the 5% CO₂ incubator at 37°C. All % survival values shown are the mean from at least three individual replicates and each experiment was performed at least 3 times.

Extracellular DNA and histone killing assays

P. aeruginosa was grown to mid-log in LB medium (OD₆₀₀ = 0.2–0.4), washed and resuspended in 10 mM Tris buffer (pH 7.4; 1.0 × 10⁷ CFU/well). Cells were incubated with fish sperm DNA (0.125%, w/v; USB) or with DNA that had been pretreated with exogenous DNase I (150 kU/well), PTase (50 U/well) or 5 mM MgSO₄. DNA was pretreated for up to 3 hrs at 37°C in order to neutralize the antimicrobial activity. To determine if DNA killing required direct cell contact, 2% w/v fish sperm DNA (USB) was resuspended in 10 mM Tris pH 7.4 was placed in sealed dialysis membranes (MW cutoff 3500 Da) and allowed to dialyze into 10 mM Tris pH 7.4 for 4 hours, exchanging the buffer every hour. Cells from mid-log P. aeruginosa PAO1 cultures (1 × 10⁷ CFU) were washed into 10 mM Tris pH 7.4 and coincubated directly with 1% or 0.125% dialyzed DNA (final concentration), with 1% eDNA maintained inside dialysis tubing, or 10 mM Tris pH 7.4 alone as a negative control. For histone killing experiments, 1 × 10⁷ CFU mid-log growth phase P. aeruginosa PAO1, E. coli and S. aureus were washed into 10 mM Tris pH 7.4 and subsequently coincubated directly with 1.5 µg/mL calf thymus histones (Roche). Killing experiments were performed at RT in 96-well microplates and bacterial survival was assessed by colony counts (CFU/ml) every hour. All survival values shown are the mean from 4–8 individual replicates and each experiment was performed at least 3 times. Differences in bacterial survival were statistically analyzed by two-tailed student t-test.

Outer membrane damage visualization

PAO1::OM-lipoChFP was used as an indicator for outer membrane damage. This strain of PAO1 expresses a synthetic Cherry fluorescent lipoprotein (CSFP^OmlA-ChFP) anchored to the outer membrane encoded on plasmid pCHAP6656 [38]. PAO1::OM-lipoChFP was exposed to a lethal concentration of 2% w/v (20 mg/ml) extracellular sperm DNA (USB) or 2 mM EDTA and red fluorescence of untreated and DNA killed cells were monitored as described above. Fluorescent outer membrane vesicles (OMVs) were counted in 6 fields of view by ImageJ quantification using a manually controlled threshold cutoff.

DNA, histone, and CAP-mediated induction of PA4774::lux

Overnight cultures were grown in LB medium, diluted 1/100 (approximately 1 × 10⁷ CFU) into 100 µl of HBSS medium lacking cations (Life Technologies) in 96-well black plates with a transparent bottom (Thermo Scientific) and overlaid with 75 µl of mineral oil (Sigma Aldrich) to prevent evaporation. Microplate planktonic cultures were incubated at 37°C in a Wallac Victor³ luminescence plate reader (Perkin-Elmer) and optical density (growth, OD₆₀₀) and luminescence (gene expression, CPS) readings were taken every 20 minutes in the presence of 0.2% salmon sperm DNA, 0.125 µg/mL polymyxin B and colistin, and 0.1 µg/mL calf thymus histones (Roche). Mean gene expression was derived from triplicate samples at 180 minutes after initial dilution and error bars represent the standard deviation from 4 individual replicates. Differences were statistically assessed by two-tailed student t-test.

Deoxyribonuclease assay

Overnight cultures of S. aureus and E. coli were grown in BHI medium and P. aeruginosa in BM2 medium, normalized to an OD₆₀₀ = 1 and supernatants were collected by centrifugation at 8000 rpm for 3 minutes. 15 µL of supernatant was incubated with 5 µg of P. aeruginosa genomic DNA for 1 h at 37°C. Pseudomonas aeruginosa genomic DNA was purified using the Wizard Genomic DNA purification kit (Promega). DNA degradation was visualized on red safe (FroggaBio) stained 1% agarose gels. To test whether exposure to NETs induced DNase production, supernatants from S. aureus, E. coli and P. aeruginosa incubated in HBSS lacking cations with 10⁶ PMA-stimulated human neutrophils (MOI 10:1, same method as described in the NET killing experiments section) were collected by centrifugation at 8000 rpm for 3 minutes. 100 µL of the supernatants were then coincubated at 37°C with 5 µg salmon sperm DNA stained with 2.5 µM Sytox green. 90 kU of DNase I was included as a positive control. Reactions were placed in 96-well black plates with a transparent bottom and Sytox green fluorescence quantified after 1 hour in a Wallac Victor³ luminescence plate reader.

Neutrophil elastase assay

To determine whether phosphatase treatment or the presence of excess Mg²⁺ altered NET-bound protein function, 2 × 10⁵ human neutrophils were seeded in 96-well black plates with a transparent bottom and induced with 100 nM PMA. Immediately after PMA addition, 50 units of phosphatase (CIAP) and 5 mM MgSO₄ were added to wells. The plate was then placed in cell culture conditions for 2 hours (37°C, 5% CO₂). 300 µM elastase substrate I was added to all wells (Calbiochem), which were subsequently overlaid with 75 µL of mineral oil. The plate was then placed in a Wallac Victor³ luminescence plate reader at 37°C. Neutrophil elastase activity was monitored by measuring absorbance at OD₄₁₀ nm every 20 minutes for 8 hours.

Statistical analysis

Statistical analysis was performed using GraphPad Prism v4.0 software. One-way ANOVA with Bonferroni posts tests and two-tailed students t-tests were used to calculate significant differences for plate counts, luminescence and flow cytometry analyses. Significant differences refer to P< 0.05 or less, or as otherwise denoted.

Ethics statement

Human neutrophils were isolated from human blood samples with ethical approval by the University of Calgary Research Ethics Committee (Ethics ID# 23187), where all subjects provided written informed consent. All animal protocols were approved by the animal care committee of the University of Calgary under the protocol number AC12–0222. All protocols used were in accordance with the guidelines drafted by the University of Calgary Animal Care Committee and the Canadian Council on the Use of Laboratory Animals.

Supporting Information

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