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. Author manuscript; available in PMC: 2014 Dec 15.
Published in final edited form as: J Immunol. 2013 Nov 4;191(12):6022–6029. doi: 10.4049/jimmunol.1301821

Staphylococcus aureus LukGH promotes formation of neutrophil extracellular traps

Natalia Malachowa *, Scott D Kobayashi *, Brett Freedman *, David W Dorward †, Frank R DeLeo *,1
PMCID: PMC3903389  NIHMSID: NIHMS529976  PMID: 24190656

Abstract

Staphylococcus aureus secretes numerous virulence factors that facilitate evasion of the host immune system. Among these molecules are pore-forming cytolytic toxins, including Panton-Valentine leukocidin (PVL), leukotoxins GH (LukGH; also known as LukAB) and DE (LukDE), and gamma-hemolysin (HlgABC). PVL and LukGH have potent cytolytic activity in vitro, and both toxins are proinflammatory in vivo. Although progress has been made towards elucidating the role of these toxins in S. aureus virulence, our understanding of the mechanisms that underly the proinflammatory capacity of these toxins, and the associated host response towards them, is incomplete. To address this deficiency in knowledge, we assessed the ability of LukGH to prime human PMNs for enhanced bactericidal activity and further investigated the impact of the toxin on neutrophil function. We found that unlike PVL, LukGH did not prime human neutrophils for increased production of reactive oxygen species nor did it enhance binding and/or uptake of S. aureus. Unexpectedly, LukGH promoted release of neutrophil extracellular traps (NETs), which in turn, ensnared but did not kill S. aureus. Furthermore, we found that electropermeabilization of human neutrophils—used as a separate means to create pores in the neutrophil plasma membrane—similarly induced formation of NETs, a finding consistent with the notion that NETs can form during non-specific cytolysis. We propose that the ability of LukGH to promote formation of NETs contributes to the inflammatory response and host defense against S. aureus infection.

Keywords: Staphylococcus aureus, neutrophil, leukotoxin, priming, extracellular trap

INTRODUCTION

Staphylococcus aureus is a leading cause of bacterial infections worldwide and renowned for causing a diverse clinical spectrum of disease. The success of S. aureus as a pathogen is facilitated by production of a litany of virulence factors that promote evasion of the host innate immune system. S. aureus secretes a wide range of toxins including several leukotoxins that have potent cytolytic activity towards polymorphonuclear leukocytes (PMNs) (1). S. aureus expresses at least four different two-component leukotoxins: Panton-Valentine leukocidin (PVL), LukGH, LukDE, and γ-hemolysin. These leukotoxins are comprised of two polypeptide subunits that sequentially assemble and form a β-barrel pore across the plasma membrane. Although the ability of the bicomponent leukotoxins to lyse PMNs in vitro is well characterized (2–6), the impact of sublytic concentrations of leukocidins on neutrophil function and/or the overall host immune response is less clear. Indeed, previous studies have shown that leukotoxins such as PVL may not achieve cytolytic concentrations in vivo and suggest an additional role that extends beyond cell lysis (7, 8).

PVL is a potent proinflammatory toxin and elicits a pronounced inflammatory response following injection of purified toxin into the skin in experimental animal models. In addition, sublytic levels of PVL induce several myeloid cellular responses, including release of myeloperoxidase and chemotactic factors such as IL-8 and LTB4 (9–12), and primes PMNs for enhanced bactericidal activity (13). Recently we demonstrated that although LukGH is cytolytic towards rabbit neutrophils in vitro and promotes inflammation in the skin, isogenic USA300 strains devoid of the toxin promoted more severe infection than the parent strain. Although the finding suggests that LukGH may contribute to host defense against S. aureus, the mechanism underlying the ability of the leukotoxin to enhance the innate immune response is unknown. To address this deficiency in knowledge, we investigated the ability of sublytic levels of LukGH to alter neutrophil function.

MATERIALS AND METHODS

Human PMN isolation

Human PMNs were purified from heparinized blood of healthy donors as previously described (14), in accordance with a protocol approved by the NIAID Institutional Review Board for human subjects. Donors were informed of the procedure risks and provided written consent prior to enrollment. Purity and viability of purified PMNs was at least 99%.

S. aureus culture conditions

S. aureus USA300 strain LAC was cultured in trypticase soy broth (TSB, Difco, Detroit, MI) at 37°C with shaking at 225 rpm. Overnight cultures were diluted 1:200 into fresh TSB media and bacteria were cultured to mid-logarithmic phase of growth (OD600 = 0.75).

Purification of LukGH from USA300 culture supernatant

S. aureus strain SF8300ΔhlgABC/ΔlukDE/Δpvl containing the plasmid pTX-15-lukGH was used to produce and purify LukGH as described (5). The quality of purified protein was assessed by SDS-PAGE, and protein concentration was determined by use of a BCA Protein Assay kit in accordance with the manufacturer’s protocol (Pierce Protein Research Products/Thermo Fisher Scientific, Rockford, IL). Purified LukGH was stored in 0.2 M NaCl 30 mM sodium phosphate buffer, pH 6.5 at −80°C.

PMN plasma membrane permeability and cytolysis assays

LukGH-mediated PMN plasma membrane permeability (pore formation) of human PMNs was evaluated by ethidium bromide (EtBr) uptake (7, 15). To exclude cell debris, intact neutrophils were gated (during analysis) based upon typical FSC (forward scatter) and SSC (side scatter) characteristics. Briefly, PMNs were incubated with 0.1–5 nM LukGH for 30 min. Heat inactivation of LukGH was accomplished by incubating the protein for 10 min at 95°C. To verify that PMN membrane permeability and lysis were caused by LukGH, we inhibited cytolysis with rabbit polyclonal antibody specific for the LukH-specific peptide sequence KDKRNVTNKDKNSC (GenScript, Piscataway, NJ). To inhibit PMN cytolysis, LukGH was incubated for 15 min at room temperature with 100-, 40-, or 20 µg/ml of anti-LukH (αLukH) prior to combining with PMNs.

Human neutrophil lysis was determined by lactate dehydrogenase (LDH) release using the Cytotoxicity Detection kit (Roche Applied Sciences, Indianapolis, IN) as described previously (16). Briefly, neutrophils (1×106) were combined with 0.1–5 nM of LukGH alone or a mixture of LukGH containing 100-, 40-, or 20 µg/ml of αLukH in 96-well tissue culture plates. After 3 h of incubation at 37°C, the plate was centrifuged at 1600 rpm for 7 min at 4°C and cell supernatants were diluted 1:1 with RPMI-1640 medium (Invitrogen/Life Technologies, Grand Island, NY) buffered with 10 mM HEPES (RPMI/H) for detection of extracellular LDH as per manufacturer instructions.

Electropermeabilization of human neutrophils

Freshly isolated neutrophils (400 µl; 107 PMNs/ml) were pre-chilled on ice in 0.2-cm gap Gene Pulser® cuvettes (Bio-Rad Laboratories, Hercules, CA). Cells were electropermeabilized in a GenePulser Xcell (Bio-Rad Laboratories) using the following settings (determined empirically): (i) 600V and 10 µF or (ii) 800 V and 25 µF. The efficiency of electroporation was confirmed by PMN permeability assays as described above, and cells were examined visually under the microscope to monitor gross cell integrity.

PMN bactericidal activity assays

Bacteria were opsonized with 50% autologous normal human serum for 30 min at 37°C. Opsonized bacteria were washed with sterile Dulbecco’s phosphate-buffered saline (DPBS, Sigma-Aldrich, St. Louis, MO) and re-suspended in RPMI/H at a final concentration of 107 cells/ml. To determine whether LukGH primes neutrophils for enhanced bactericidal activity, PMNs (2 × 106 cells/ml; 500 µl in a1.5-ml tube) were incubated with 0.1 or 1 nM LukGH for 30 min at 37°C. Bacteria were added at a MOI of 1:1 to LukGH pretreated neutrophils and samples were incubated at 37°C with rotation for up to 2 h. In addition, we evaluated several permutations of our neutrophil bactericidal activity and LDH release assays, such as varied bacteria:neutrophil ratios (0.1 and 10:1), survival of USA300 wild-type and isogenic ΔlukGH mutant strains, impact of calcium and magnesium, and differential effects of anticoagulants used in blood collection (EDTA versus heparin). None of these permutations had a significant impact on bacterial survival or LDH release. For assays that contained DNase I (StemCell Technologies Inc., Vancouver, BC), the enzyme was added to the samples at a final concentration of 50 µg/ml prior to adding bacteria or 15 min before the 2 h time point. At 2 h, 25-µl aliquots of the assay contents were evaluated by light microscopy (cytospin followed by Wright-Giemsa stain) and the remaining sample was mixed with saponin on ice for 15 min and subsequently diluted and plated on trypticase soy agar. CFUs were enumerated on the following day and were used to determine percent S. aureus survival relative to a time-matched control assay that did not contain PMNs.

Phagocytosis assays were performed as described by Voyich et al. (17). To quantitate the percent PMNs with bound/ingested S. aureus, 50 PMNs (per blood donor) from different fields of view were scored for associated bacteria (includes bound and ingested).

PMN priming assays

Lucigenin–enhanced chemiluminescence was used to measure neutrophil superoxide production (18). Assays were performed in white 96-well cell culture plates (Nunclon, Nunc™, Roskilde, Denmark) coated with 20% human serum for 30 min at 37°C. A 100-µL aliquot of PMNs in RPMI/H (107 PMNs/ml) was added to each well, followed by addition of LukGH (1 nM final concentration) or a mixture of LPS/LBP (final concentration 100 ng/ml each; R&D Systems) and the assay was incubated at 37°C for 30 min. Plates were subsequently transferred to a SynergyMX plate reader (BioTek, Winooski, VT) and lucigenin (bis- N-methylacridinium nitrate; Sigma-Aldrich) or a mixture of lucigenin and 1 µM fMLF (formyl-methionyl-leucyl-phenylalanine; Sigma-Aldrich) was injected into the wells using an automated injector. Luminescence in each sample was recorded for 10 min with 24-s minimal intervals.

Alternatively, superoxide production was measured by reduction of ferricytochrome c as described previously (13), but with a few modifications. Freshly purified human PMNs (107 cells/ml) were incubated for 15 min at 37°C with 0.1 or 1 nM LukGH or a mixture of LPS/LBP (positive control) as described above. Subsequently, cells were added to a 96-well cell culture plate containing 100 µM ferricytochrome c, ± 1 µM fMLF, and ± 40 µg/ml superoxide dismutase (SOD; Sigma-Aldrich). Release of superoxide was monitored for 20 min using a SpectraMax 384 Plus microplate reader (Molecular Devices LLC., Sunnyvale, CA).

Surface expression of CD11b was assessed by flow cytometry (FACSCalibur, BD Bioscience, San Jose, CA) as described elsewhere (13).

Myeloperoxidase (MPO) release/activity assays

MPO assays were performed in 96-well cell culture plates (Costar, Corning, NY) coated with 20% pooled normal human sera. 100 µl of RPMI/H or media containing LukGH (final concentration 0.1 or 1 nM) was added to the wells, followed by 100 µl of PMNs (107 cells/ml) or electropermeabilized PMNs (107 cells/ml). Plates were centrifuged at 450g for 7 min at 4°C, and then incubated for 2 h at 37°C. 10 µl of DNase I (50 µg/ml final concentration) was added to the appropriate wells during the last 15 min of incubation. At time points indicated, plates were centrifuged at 450g and 4°C for 7 min. To quantify extracellular MPO, culture supernatants were diluted 1:200 with RPMI/H buffer and MPO concentration was determined using an EnzCheck MPO activity kit (Invitrogen/Life Technologies).

Immunofluorescence and confocal laser-scanning microscopy

Samples for immunofluorescence microscopy were prepared as described (19), but with the following modifications. Acid-washed cover slips were placed in wells of a 24-well tissue culture plate, coated with 100% pooled normal human serum and incubated for at least 1 h at 37°C. Coverslips were washed with DPBS, and 300 µl of 106 PMNs/ml was transferred to each well. PMNs or freshly electropermeabilized PMNs were incubated at room temperature for 15 min and then transferred on ice. PMNs were combined with RPMI/H or RPMI/H containing LukGH or PMA as indicated, the cells were centrifuged at 450g at 4°C for 8 min, followed by incubation for 2 h at 37°C. Plates were placed on ice and cells were washed with ice cold DPBS. PMNs were fixed in 4% paraformaldehyde for 30 min on ice followed by overnight blocking at 4°C in Stain Buffer with 2% FBS (BD Pharmingen™, BD Bioscience). Cells were washed once and incubated with 2 µg/ml rabbit anti-histone 2A (αH2A, Abcam, Cambridge, MA) and/or 1 µg/ml mouse anti-human myeloperoxidase (αMPO, BD Pharmingen) or 2 µg/ml rabbit anti-neutrophil elastase antibody (Abcam) in Stain Buffer (BD Pharmingen) at room temperature with gentle agitation. After a 90-min incubation, samples were washed 3 times with Stain Buffer and incubated for 1 h with secondary antibody (2 µg/ml goat anti-rabbit antibody conjugated with AlexaFluor594 or AlexaFluor488, Jackson, ImmunoReasearch Laboratories, Inc. West Grove, PA; and 2 µg/ml goat anti-mouse antibody conjugated with AlexaFluor 488, Molecular Probes/Life Technologies). Coverslips were washed 3× with DPBS and mounted on microscopy slides with DAPI Fluoromount G medium (Electron Microscopy Sciences, Hatfield, PA). Images were evaluated using a Zeiss 510 Meta scanning laser confocal microscope, LSM510 software version 4.2. The brightness and contrast of images were adjusted in Adobe Photoshop CS5 (Adobe Systems, San Jose, CA).

Scanning and transmission electron microscopy

Samples for scanning electron microscopy (SEM) were prepared as described elsewhere (17) with the following modifications. PMNs (3 × 105) were seeded on 100% serum coated silicon chips (Ted Pella, Inc., Redding, CA) contained in 24-well tissue culture plates. Cells were allowed to attach for 15 min at room temperature followed by addition of LukGH or PMA in RPMI/H medium as indicated. RPMI/H medium only was added to the PMN control and electropermeabilized PMNs. Samples were incubated at 37°C with 5% CO2 for 30 min. At time points indicated, cells were fixed and processed with microwave excitation. Sample chips were covered with approximately 0.5 ml of Karnovsky’s fixative and irradiated at 80 W for two cycles of 2 min on, 2 min off, and 2 min on in a Biowave model microwave processor, equipped with a vacuum chamber and Cold-spot® radiation attenuation system (Ted Pella, Inc.). Following a 45 sec wash in phosphate buffer at 80 W, the samples were post-fixed in 1% osmium tetroxide and phosphate buffer with 2 cycles of 2-2-2 as above. Samples were washed twice in water for 45 sec at 80 W, and dehydrated for 1 min each at 250 W in 70%, 100 %, and 100% ethanol. The samples were critical-point-dried through carbon dioxide, sputter coated with iridium in a model IBSe ion beam sputterer (South Bay Technologies, San Clemente, CA), and examined using a model S-8000 field emission scanning electron microscope (Hitachi High Technologies, America, Schaumburg, IL). For transmission electron microscopy (TEM), cells were seeded on 13 mm serum-coated Thermanox cover slips (VWR, West Chester, PA) and treated as described above (except neutrophils were incubated for up to 4 h). Samples were processed further using low-power microwave irradiation as described previously (20). Images were adjusted in Photoshop CS 5 (Adobe Systems) for contrast and brightness.

Statistical analyses

Data were compared using a one-way ANOVA and Tukey’s or Bonferroni’s post-test to correct for multiple comparisons (GraphPad Prism 5, GraphPad Software, San Diego CA) as indicated in the figure legend.

RESULTS

Impact of LukGH on PMN function

We first assessed the effect of varied concentrations of LukGH purified from S. aureus culture supernatants on formation of PMN plasma membrane pores (Fig. 1A) and cytolytic capacity (Fig. 1B). LukGH-mediated PMN pore formation and cytolysis were concentration-dependent (Fig. 1A-B). In addition, αLukH inhibited the ability of the purified LukGH to form pores in the PMN plasma membrane and cause cytolysis, thus demonstrating that activity was specific to the leukocidin.

Figure 1. LukGH-mediated PMN plasma membrane pore formation and cytolysis.

Figure 1

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(A) Pore formation. PMNs (1×106) were incubated with 0.1–5.0 nM LukGH or 5.0 nM heat-inactivated (h.i.) LukGH for 30 min, and plasma membrane permeability was evaluated using an EtBr uptake assay. Alternatively, LukGH was incubated with αLukH or IgG isotype control antibody at the indicated concentrations for 15 min prior to combining with PMNs. (B) PMN lysis. PMNs (1×106) were incubated with LukGH, h.i. LukGH, and/or antibody as described in panel A and lysis was determined by release of LDH as described in Methods. *P < 0.05 for the indicated comparisons using a one-way ANOVA and Tukey’s post-test.

As a first step towards understanding the mechanism underlying the proinflammatory capacity of LukGH, we assessed the ability of the toxin to function as a neutrophil priming agent. Priming of PMNs elicits granule exocytosis and thereby promotes enrichment of surface receptors such as CD11b/CD18 at the plasma membrane (21). Thus, we tested the ability of 0.1 nM and 1 nM LukGH to upregulate expression of CD11b at the neutrophil surface (Fig. 2A). PMNs exposed to 0.1 or 1 nM LukGH had increased surface expression of CD11b (e.g., the mean FL2 for PMNs exposed to 0.1 nM LukGH at 15 min was 359.4 ± 56.4 and 610.4 ± 103.3 for control and LukGH-treated PMNs; P < 0.05). These results indicate that LukGH promotes fusion of secretory vesicles and/or specific granules with the plasma membrane, and is consistent with the ability of known PMN priming agents−including PVL−to up-regulate PMN surface expression of CD11b (13, 22, 23). Inasmuch as the hallmark of PMN priming is enhanced release of O2−, we assayed LukGH-treated PMNs for increased superoxide anion production following secondary stimulation with fMLF (Figs. 2B-C). As evidenced by lucigenin-enhanced chemiluminescence assays, neutrophils incubated with 1 nM LukGH did not produce superoxide when challenged with fMLF as a secondary stimulus (Fig. 2B). Consistent with these observations, PMNs exposed to either 0.1 nM LukGH or 1.0 nM LukGH did not demonstrate fMLF-mediated release of O2− in ferricytochrome c reduction assays (Fig. 2C). By contrast, PMNs primed with LPS (positive control for priming) showed markedly enhanced production of O2− in response to fMLF (Figs 2B-C). Although LukGH failed to prime neutrophils for enhanced O2− production, the finding that LukGH promoted increased CD11b surface expression lends to the possibility that PMN signal transduction pathways unlinked to NADPH oxidase activity are altered.

Figure 2. LukGH alters CD11b surface expression but fails to prime human PMNs for enhanced superoxide anion production.

Figure 2

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(A) Neutrophil surface expression of CD11b. PMNs were exposed to 0.1 nM LukGH, 1nM LukGH or 1 nM h.i. LukGH for 30 min and surface expression of CD11b was assessed by flow cytometry. Results are the mean ± SE of three to four PMN donors. *P < 0.05 for the indicated comparisons using a one-way ANOVA and Bonferroni’s posttest. (B-C) Priming for PMN superoxide. (B) PMN superoxide production was evaluated with lucigenin–enhanced chemiluminescence using the indicated stimuli. PMNs were incubated with 1 nM LukGH or 100 ng/ml LPS + LBP for 30 min and then activated with 1 µM fMLF for 10 min as indicated. (C) Superoxide production as assessed by reduction of ferricytochrome c. PMNs were incubated with 0.1 nM LukGH, 1 nM LukGH, and/or 100 ng/ml LPS + LBP for 15 min and then activated with 1 µM fMLF for 20 min as indicated. Results are the mean ± SE of three to seven experiments.

We next determined whether LukGH promotes enhanced PMN bactericidal activity against S. aureus (Fig. 3A). Compared with control assays, the number of S. aureus CFUs recovered from assays in which PMNs were exposed to 0.1 nM or 1 nM LukGH were significantly reduced at 60 and 120 min (e.g., survival of USA300 at 60 and 120 min was 110.2 ± 11.6 and 44.6 ± 10.1 for assays without LukGH compared to 63.1±9.4 and 11.0 ± 2.3 for those containing 0.1 nM LukGH; P < 0.05) (Fig. 3A). Unexpectedly, and in contrast to previous findings with PVL-primed PMNs, the reduction in recovered S. aureus CFUs from assays with LukGH-treated PMNs was not due to increased ingestion and/or binding of S. aureus (Figs. 3B-C). Collectively, these observations suggest that LukGH-enhanced PMN bactericidal activity is independent of NADPH-oxidase-derived superoxide anion or there is aggregation of bacteria (and thus reduced CFUs) by PMNs following treatment with LukGH.

Figure 3. Impact of LukGH on PMN phagocytosis and bactericidal activity.

Figure 3

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(A) Bactericidal activity. PMNs were treated with 0.1 or 1 nM LukGH for 30 min, incubated with S. aureus strain USA300, and bactericidal activity was determined as described in Methods. (B) PMN phagocytosis. PMN assays were performed as described in panel A and phagocytosis was determined at the indicated time points. (C) Association (bound + ingested) of bacteria with PMNs. Results are the mean ± SE of four to five experiments. *P < 0.05 for the indicated comparisons using a one-way ANOVA and Tukey’s post-test.

LukGH promotes formation of neutrophil extracellular traps (NETs)

To further investigate the mechanism underlying the ability of LukGH to promote a reduction of the number of S. aureus CFUs recovered from assays with human PMNs, we examined bacteria and PMNs from the bactericidal activity assays by standard light microscopy (Fig. 4). At 2 h following incubation, PMNs exposed to LukGH and challenged with S. aureus consistently showed evidence of concentration dependent necrosis and/or clumping, whereas this effect was limited with control, untreated cells (Fig. 4, panels A-C). Notably, cells exposed to LukGH often formed filamentous structures that resembled extracellular traps (Fig. 4, panels D and E). By comparison, neutrophils failed to exhibit either phenotype in the absence of exogenous LukGH (Fig. 4, panel A) or following addition of DNase I to PMNs treated with LukGH (Fig. 4, panels G and H). The finding that LukGH promoted the release of neutrophil DNA is consistent with the ability of the related S. aureus pore-forming toxin, PVL, to induce formation of NETs (24). Inasmuch as LukGH enhanced neutrophil aggregation and induced formation of putative extracellular traps, we next modified the PMN bactericidal activity assays to include DNase I digestion of extracellular DNA, and assessed recovery of S. aureus CFUs (Fig. 5). Compared with assays lacking DNase I, the number of recovered S. aureus CFUs was significantly greater in assays containing LukGH and DNase I (e.g., survival of USA300 was 27.34 ± 8.4% and 58.66 ± 2.8% for cells exposed to 0.1 nM LukGH with and without DNase I; P < 0.05; Fig 5). By comparison, DNase I had no significant impact on the number of recovered S. aureus CFUs from assays with PMNs alone. Moreover, the number of CFUs recovered from PMN assays that contained LukGH and DNase I was not significantly different than those containing DNase I only (e.g., survival of USA300 was 58.66 ± 2.8% and 66.71 ± 7.2% with and without 0.1 nM LukGH; Fig. 5). These data provide support to the idea that extracellular DNA—released as a result of exposure to LukGH—promotes aggregation of PMNs and bacteria, and this phenomenon results in gross underestimation of CFUs by the conventional bactericidal activity assay (lacking DNase I).

Figure 4. LukGH promotes extracellular release of neutrophil DNA.

Figure 4

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PMNs were incubated without (A, F) or with 0.1 nM (B, D, G) or 1 nM (C, E, H) LukGH for 30 min, and combined with S. aureus strain USA300 at a ratio of 1:1 bacteria per PMN (A-H). Alternatively, DNase I was added to prior to addition of bacteria (F, G, H). PMNs and bacteria were cultured for 2 h at 37°C with gentle rotation. Samples were stained with modified a Wright-Giemsa stain and evaluated by light microscopy (original magnification ×400). Black arrows indicate neutrophils, red arrows indicate S. aureus strain USA300, and green arrows indicate extracellular DNA. Images D and E (extracellular DNA), and B and C (clumping/necrosis) represent two distinct features of samples treated with LukGH.

Figure 5. LukGH-induced NETs do not promote killing of S. aureus.

Figure 5

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PMN bactericidal activity. PMNs were pre-treated with buffer, 0.1 nM LukGH, or 1 nM LukGH for 30 min, and bactericidal activity was assessed by determination of S. aureus CFUs after 2 h of culture. DNase I was added to samples as indicated 15 min prior to termination of the assay. Results are the mean ± SE of three to five experiments. *P < 0.05 for the indicated comparisons using a one-way ANOVA and Tukey’s post-test.

Neutrophils have been reported to release extracellular traps in response to S. aureus (25, 26) and the pore-forming toxin PVL (24). We performed immunofluorescence staining of PMNs following LukGH treatment to verify the presence of extracellular DNA, histone 2A (H2A), and myeloperoxidase (MPO)−known features of NETs (27). Indeed, we found that histones and myeloperoxidase were associated with LukGH-induced NETs at 2 h, and the staining pattern was similar to PMNs exposed to phorbol myristate acetate (PMA, used as a positive control for the formation of NETs) (Fig. 6). Moreover, the nucleus of the cells treated with LukGH underwent morphological changes characteristic of those reported for NETosis (Supplemental Fig. 1). Given that PVL (24) and LukGH each induce NETs, we tested whether non-specific pore formation per se could induce NETs. Formation of PMN plasma membrane pores by electropermeabilization caused discrete pores in the plasma membrane without grossly disrupting cell architecture, as revealed by scanning electron microscopy (Supplemental Fig. 2). For example, exposure of neutrophils to 800 V at 25 µF promoted extensive NET formation, which was similar to that after exposure to PMA or LukGH (Fig. 6). We confirmed that PMNs treated with LukGH or those that were electropermeabilized maintained extracellular MPO enzymatic activity, which increased significantly following treatment with DNase I (Fig. 7A-B), thus suggesting release of NET-bound MPO. These findings are consistent with a previous report that demonstrated MPO activity is associated with NETs (28). Collectively, these data strongly suggest that although LukGH promotes formation of NETs−here shown to be a characteristic common to non-specific PMN damage−this process ensnares bacteria but does not enhance neutrophil microbicidal activity against S. aureus.

Figure 6. Electropermeabilization of human PMNs promotes formation of NETs.

Figure 6

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PMNs were treated with 0.1 nM LukGH or 1 nM LukGH, or were electropermeabilized using a pulse of 600 V at 10 µF or 800 V at 25 µF, and were cultured for 2 h at 37°C. Alternatively, PMNs were treated with 40 nM PMA (positive control) to induce formation of NETs. Extracellular DNA was confirmed by immunofluorescence microscopy to be NETs by staining for histones (H2A; red), myeloperoxidase (MPO; green), and DNA (DAPI; blue).

Figure 7. LukGH-induced NETs sequester MPO.

Figure 7

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PMN MPO activity assays. PMNs were treated with 0.1 nM LukGH or 1 nM LukGH, or cells were electropermeabilized and cultured for 2 h at 37°C. DNase I was added to samples as indicated 15 min prior to termination of the assay. NET-associated MPO was determined by chlorination (A) and peroxidase activity (B) assays. *P < 0.05 for the indicated comparisons using a one-way ANOVA and Tukey’s post-test.

DISCUSSION

Staphylococcus aureus two-component pore-forming toxins are a related group of secreted molecules that have potent cytolytic activity against host cells−most notably leukocytes. There are at least four S. aureus leukotoxins characterized with activity against PMNs: PVL, LukGH, LukDE, and γ-hemolysin. Inasmuch as neutrophils are the primary host defense against invading bacterial pathogens, significant emphasis has been placed on understanding the role of leukotoxins in S. aureus virulence and pathogenesis. Although progress has been made, the role of two-component cytolytic toxins in S. aureus pathogenesis is unclear—if not controversial (29–32). Furthermore, it has been proposed recently that the presence of PVL can be beneficial to the host during infection, since sublytic concentrations of this leukocidin have the ability to prime human neutrophils for enhanced function and greater bactericidal activity (13, 32). Based on previous findings with LukGH, which includes the ability of the molecule to induce an inflammatory response in animal models (5, 33), we evaluated the ability of LukGH to prime human PMNs for enhanced function. Although LukGH promoted increased surface expression of CD11b, a phenomenon characteristic of priming, the toxin failed to prime PMNs for enhanced superoxide production in response to a secondary stimulus.

PMNs exposed to LukGH had a significantly altered surface morphology that was not necessarily a reflection of—or caused by—permeabilized plasma membrane (Supplemental Fig. 2). This change in PMN surface morphology was particularly evident at earlier time points after exposure to LukGH and apparently not directly related to cell lysis (Supplemental Fig. 2). There was also noted donor-to-donor variability in the sensitivity of PMNs to LukGH-mediated membrane permeabilization and lysis. Such variability is perhaps explained by varied levels in surface receptor expression, since DuMont et al. (34) recently demonstrated that CD11b is a probable LukGH receptor—and there can be considerable variance in CD11b surface expression among human PMN donors.

Despite differences in priming capacity between PVL and LukGH, each induces formation of NETs (24). NETs are DNA-based structures containing anti-bacterial proteins such as elastase, cathepsin G, or MPO, which are typically contained within primary/azurophilic granules but are released during cell lysis and associate with NETs (Supplemental Fig. 3) (27, 28, 35). Induction of NETs by LukGH or PVL ((24); and our unpublished data) is time and concentration dependent. NETs were originally reported to have bactericidal activity (27, 35). However, our data (Fig 5A) and others (36, 37) suggest NETs ensnare/trap bacteria rather than contributing directly to PMN bactericidal activity. Although LukGH failed to promote bactericidal activity directly, it is possible that trapping of bacteria by NETs promotes enhanced clearance of the pathogen in vivo. Indeed, it is known that molecules from lysed neutrophils—including during NET formation—attract other professional phagocytes such as macrophages (38, 39), which could expedite clearance of bacteria.

On the other hand, formation of NETs and associated PMN lysis has potential negative outcomes for the host, including non-specific damage to host tissues, and a contribution to autoimmune disorders or vascular dysfunction (40, 41). For example, McDonald et al. (25) demonstrated recently that NETs entrap bacteria during blood stream infections, a process that could be beneficial for the host despite concomitant host organ damage (42). These data are consistent with our previous studies, in which we found decreased survival of mice infected with a USA300 isogenic ΔlukGH/Δpvl deletion mutant strain compared with the wild-type USA300 strain (5). Despite recent progress toward our understanding of the role of PVL and LukGH during S. aureus-host interaction, further studies are needed to elucidate detailed mechanisms underlying the enhanced host response to infection mediated by these interesting molecules.

On a final note, we found that formation of NETs can occur as a consequence of non-specific physical damage to the neutrophil plasma membrane (Supplemental Fig. 2). The implication of these findings is that NETs can form simply from non-specific PMN lysis that is not dependent on specific signal transduction events.

Supplementary Material

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ACKNOWLEDGEMENTS

We thank Rocky Mountain Laboratories National Institute of Allergy and Infectious Diseases (NIAID) colleagues Anita Mora for photography and Kevin Braughton for technical assistance.

This research was supported by the Intramural Research Program of the National Institute of Allergy and Infectious Diseases, National Institutes of Health.

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