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Neutrophils deposit antimicrobial proteins, such as myeloperoxidase and proteases on chromatin, which they release as neutrophil extracellular traps (NETs). Neutrophils also carry key components of the complement alternative pathway (AP) such as properdin or complement factor P (CFP), complement factor B (CFB), and C3. However, the contribution of these complement components and complement activation during NET formation in the presence and absence of bacteria is poorly understood. We studied complement activation on NETs and a Gram-negative opportunistic bacterial pathogen Pseudomonas aeruginosa (PA01, PAKwt, and PAKgfp). Here, we show that anaphylatoxin C5a, formyl-methionyl-leucyl-phenylalanine (fMLP) and phorbol myristate acetate (PMA), which activates NADPH oxidase, induce the release of CFP, CFB, and C3 from neutrophils. In response to PMA or P. aeruginosa, neutrophils secrete CFP, deposit it on NETs and bacteria, and induce the formation of terminal complement complexes (C5b–9). A blocking anti-CFP antibody inhibited AP-mediated but not non-AP-mediated complement activation on NETs and P. aeruginosa. Therefore, NET-mediated complement activation occurs via both AP- and non AP-based mechanisms, and AP-mediated complement activation during NETosis is dependent on CFP. These findings suggest that neutrophils could use their “AP tool kit” to readily activate complement on NETs and Gram-negative bacteria, such as P. aeruginosa, whereas additional components present in the serum help to fix non-AP-mediated complement both on NETs and bacteria. This unique mechanism may play important roles in host defense and help to explain specific roles of complement activation in NET-related diseases.
Neutrophils play a central role in the innate immune system and function in inflammation and immune surveillance. At sites of inflammation, neutrophils kill pathogens via phagocytosis and release of proteolytic enzymes (1, 2). Recently, the ability of neutrophils to form web-like neutrophil extracellular traps (NETs) has been identified as an additional strategy for antimicrobial defense. The process of NET formation (i.e., NETosis) is a specific form of cell death, in which nuclear DNA undergoes decondensation with subsequent expulsion of chromatin that is coated with cytotoxic granular proteins, such as myeloperoxidase (MPO), elastase, and other proteases (3). NETs are released in response to a variety of stimuli, including NADPH oxidase (Nox) agonist, such as phorbol-12-myristate-13-acetate (PMA), inflammatory stimuli, and bacteria (4, 5). Two major types of NETosis have been reported to date: Nox-dependent NETosis and Nox-independent NETosis, in which reactive oxygen species (ROS) are generated by Nox and mitochondrial complexes, respectively (6–9). In both of these types of NETosis, neutrophil release chromatin coated with granular proteins as NETs. In the presence of C5a, GM-CSF-primed neutrophils undergo a vital NETosis, in which cells do not die, but release mitochondrial DNA. This type of NETosis is regulated by mitochondrial ROS production (10).
Once formed, NETs ensnare pathogens and expose them to high localized concentrations of antimicrobial proteins (11). NETs can also be cytotoxic and have been shown to contribute to thrombosis, sepsis, cystic fibrosis, asthma, systemic lupus erythematosus (SLE), rheumatoid arthritis (RA), and anti-neutrophil cytoplasmic antibody (ANCA)-associated vasculitis (AAV) (12–24). Complement and infections have been implicated in the pathogenesis and exacerbation of many of these diseases. Although it has recently been described that NETs can activate and deposit complement alternative pathway (AP) components (25), the involvement of the different complement pathways and their components in the context of NETosis and bacterial infection has not been fully understood. This fundamental knowledge is essential for understanding molecular mechanisms involved in NET-related pathobiology.
The complement system consists of more than 30 proteins distributed in the circulation and on endothelial cells, and functions primarily in microbial defense and clearance of immune complexes and injured cells (26). Complement can be constantly active (via the complement AP) or become activated by immune complexes and dying cells [via the C1q-mediated classical pathway (CP)] or carbohydrate ligands on microorganisms [via the lectin pathway (LP)] (26). Complement factor P (CFP), the only positive complement regulator, acts as stabilizer of the AP convertase (C3bBbP) and selective pattern recognition molecule of certain microorganisms and host cells (i.e., apoptotic/necrotic cells) by serving as a platform for the assembly of the AP C3 convertase (27). Complement progression includes the activation of complement proteins C3 and C5 (to form the potent anaphylatoxins C3a and C5a and the opsonins C3b and C5b) and the subsequent activation of the terminal pathway with the formation of the potentially lytic membrane attack complex (MAC), C5b–9. AP activation is critically enhanced by the C3 convertase C3bBbP, and requires tight regulation to maintain the balance between necessary activation and harmful over-activation (26). Bacteria are capable of inducing Nox-dependent NETosis (9, 28–30), and we aimed to identify possible links between NETosis, bacteria, and the complement system, in particular, the possibility that neutrophils mount a targeted complement response to infectious agents via the formation of NETs and deposition of complement components on NETs and microbial pathogens.
Informed written consent was obtained from all donors. The study protocol was approved by the Research Ethics Board at The Hospital for Sick Children, Toronto, ON, Canada.
All buffer salts and reagents were obtained from Sigma-Aldrich (St. Louis, MO, USA) unless stated otherwise.
Pseudomonas aeruginosa (mPA01, PAKwt, and PAKgfp) cultures were grown overnight in LB-broth. PAKgfp was maintained in 30μg/ml gentamicin. The concentration of bacteria was calculated using [CFU]×108=(OD600) 30.88−99,607. For NETosis assays, P. aeruginosa sub-cultured for 3h was used at a multiplicity of infection (MOI) of 10 or 100.
Human peripheral neutrophils were purified from whole blood (20ml) collected in BD EDTA-vacutainers from healthy donors using Polymorphprep™ (Axis-shield, Oslo, Norway). After lysing erythrocytes with hypotonic buffer, neutrophils were resuspended in RPMI 1640 (Wisent Bioproducts, Montreal, QC, Canada) (31). To obtain neutrophil lysate, cell pellets were resuspended in a lysis buffer [1% (v/v) Triton X-100, 50mM Tris, pH 7.4, 10mM KCl containing 2× complete, mini protease inhibitor cocktail (Roche Diagnostics, Laval, QC, Canada) supplemented with 0.5mM EDTA, 25μM leupeptin, 25μM pepstatin, 25μM aprotinin, 1mM levamisole, 1mM Na3VO4, 25mM NaF, 1mM PMSF], sonicated (VWR Sonics model 50D), and incubated for 15min at 4°C. Neutrophil lysates were centrifuged at 25,000×g for 30min at 4°C and stored at −80°C for future analysis.
Neutrophils (2×107 cells/ml) were resuspended in RPMI 1640 with 10mM Hepes, pH 7.4, and activated with C5a (CompTech, Tyler, TX, USA) (1μM), formyl-methionyl-leucyl-phenylalanine (fMLP) (1μM) or PMA (20nM), and incubated [37°C, 5% (v/v) CO2] for 30min. Stimulation was terminated by incubating these cells at 4°C for 5min. Neutrophils were pelleted (1000×g for 10min) and the supernatant was collected and further centrifuged at 25,000×g for 10min at 4°C, immediately placed in 2×neutrophil Laemmli sample buffer, heated at 95°C for 5min and stored at −80°C for future analysis. Neutrophil lysates were prepared from the remaining cell pellet as described in the Section “Neutrophil Isolation and Preparation of Neutrophil Lysates.” To determine respiratory burst, neutrophils (1×106 cells/ml) were pre-loaded with dihydrorhodamine (DHR) 123 (10μM), treated with the agonists as above and analyzed by flow cytometry (Gallios, Beckman Coulter, Mississauga, ON, Canada). Cells were first gated with forward and side scatters, and further gated for Hoechst (1μg/ml) using 405/450 BP 40 filter channel. ROS was detected using 488/429 BP 28.25 filter channel.
Neutrophils (3×104 cells) were seeded onto 96-well plates in the presence of cell-impermeable Sytox Green DNA-binding dye (5μM) and were activated with agonists or three stains of P. aeruginosa (mPAO1, PAKwt, and PAKgfp) at MOIs of 10 and 100. For inhibition studies, neutrophils were preincubated with 2μM of Nox inhibitor, diphenyleneiodonium (DPI) for 1h before activation. Fluorescence intensity was measured by the POLARstar Omega microplate reader (BMG Labtech, Ortenberg, Germany) with excitation/emission (485/520), every 30min. NET formation was normalized to total neutrophils DNA content determined by permeabilizing the cells with 0.5% (w/v) Triton X-100.
Neutrophil lysates (50μg) were size-fractionated in 10% (w/v) SDS-polyacrylamide gels, transferred to nitrocellulose membranes, blocked with 5% (w/v) skim-milk+0.05% (v/v) Tween-20 (TBST), probed with goat polyclonal antibody to complement proteins (1:1000 dilution; Complement Technology, Tyler, TX, USA) or mouse monoclonal antibody to β-actin (BA3R, 1:10,000 dilution; Thermo Fisher Scientific, Rockford, IL, USA) in 5% (w/v) skim-milk in TBST, washed, and incubated with secondary antibody in 5% (w/v) skim-milk in TBST. Proteins were detected using Western Lighting™ Plus-ECL, Enhanced (PerkinElmer, Waltham, MA, USA) and developed on radiographic film on a Kodak X-Omat 2000a processor.
Phorbol-12-myristate-13-acetate (20nM) activated neutrophils at 240min were fixed with 4% (w/v) paraformaldehyde (Electron Microscopy Sciences, Fort Washington, PA, USA), blocked with 3% (v/v) cold water fish skin gelatin and incubated with anti-complement antibodies: rabbit polyclonal antibody to CFP (SC-68366, 1:50 dilution; Santa Cruz Biotechnology, Dallas, TX, USA), rabbit polyclonal antibody to complement factor B (CFB) (SC-67151, 1:50 dilution; Santa Cruz Biotechnology, Dallas, TX, USA), and rabbit polyclonal antibody to complement C3 (ab97462, 1:100 dilution; Abcam, Cambridge, MA, USA). Neutrophils activated with PMA were co-stained with mouse monoclonal antibody to MPO (ab25989, 1:500 dilution; Abcam, Cambridge, MA, USA). Complexes were detected with donkey anti-primary antibodies conjugated with Alexa Fluor® 555 (Invitrogen, Eugene, OR, USA). Specimens were mounted with Dako Fluorescence Mounting Media (Dako Canada, Burlington, ON, Canada) for analysis with spinning-disk confocal microscopy.
After 240min of neutrophil activation, culture plates were centrifuged at 200×g for 5min at 4°C. The media was replaced with 500μl of 20% (v/v) Refludan® (Bayer Healthcare, Wayne, NJ, USA) fresh frozen PPP prepared in RPMI 1640 media or AP buffer (20mM Hepes, pH 7.4, 144mM NaCl, 7mM MgCl2, and 10mM EGTA). For experiments with AP buffer, a buffer exchange was performed prior to the addition of plasma (Bayer Healthcare, Wayne, NJ, USA). After 15min [37°C, 5% (v/v) CO2], specimens were fixed with 4% (w/v) paraformaldehyde (Electron Microscopy Sciences, Fort Washington, PA, USA). In some experiments, DNase I (50μg/ml) was added to digest DNA. After blocking with 3% (v/v) gelatin, mouse monoclonal antibody to C5b–9 (DIA 011-01, 1:200 dilution; Antibody Shop, Gentofte, Denmark) was incubated, washed, and further incubated with donkey anti-mouse secondary antibody conjugated with Alexa Fluor® 555 (Invitrogen, Eugene, OR, USA). PAKgfp signal was enhanced with Alexa Fluor® 488 conjugated rabbit polyclonal antibody to GFP (A21311, 1:400 dilution; Invitrogen, Eugene, OR, USA). For CFP inhibitor assays, optimal concentration of a mouse monoclonal anti-CFP antibody (Anti Factor P#1, A233; Quidel Corporation, San Diego, CA, USA) was determined by rabbit red blood cell lysis assay (32). An antibody concentration of 4μg/ml was used in the final assays. Presence of C5b–9 was detected as described above. Wheat germ agglutinin was used for labeling neutrophil membrane. Specimens were mounted and analyzed with spinning-disk confocal microscopy.
Images were taken on an Olympus IX81 inverted fluorescence microscope using a 60×/1.35 oil immersion objective equipped with a Hamamatsu C9100-13 back-thinned EM-CCD camera and Yokogawa CSU X1 spinning-disk confocal scan head (with Spectral Aurora Borealis upgrade). The unit is equipped with four separate diode-pumped solid state laser lines (Spectral Applied Research, 405, 491, 561, and 642nm) with emission filters: 447±60, 525±50, 593±40, 620±60, 676±29, and 700nm±75, and 1.5× magnification lens (Spectral Applied Research). Confocal images were taken with an Improvision Piezo Focus Drive. Z-stacks were taken at 0.25μm. Images taken using the spinning-disk confocal microscope were deconvolved by iterative restoration using Volocity Software (PerkinElmer, Waltham, MA, USA) with confidence limit set to 95% and iteration limit set to 20.
Student’s t-test, or one-way or two-way ANOVA with Tukey’s multiple comparison test was used for statistical comparison as needed. A p-value was set at 0.05, 0.01, or 0.001 for statistical significance. All statistical analyses were performed using GraphPad Prism (GraphPad Software, La Jolla, CA, USA) statistical analysis software (Version 6.0).
Reactive oxygen species is considered to be important for NETosis. However, different agonists induce ROS to different degrees. Therefore, to determine whether ROS was sufficient to induce NETosis, ROS production was measured 30min post neutrophil stimulation using DHR 123 and flow cytometry. C5a (1–2μM) did not generate ROS above baseline levels (Figure S1A in Supplementary Material); however, similar concentrations of fMLP led to a 1.5-fold increase in ROS compared to baseline values (p<0.05; Figure S1B in Supplementary Material). The use of 20nM PMA produced a ninefold increase of ROS in comparison to non-treated control neutrophils (p<0.05; Figure S1C in Supplementary Material). The overall ability of these agonists to induce ROS production was PMA>>fMLP>C5a.
Although both PMA and fMLP induced ROS production, whether fMLP can induce NETosis is uncertain. Therefore, to identify the ability of PMA, fMLP, and C5a to independently elicit NETosis, we treated neutrophils with varying concentrations of these reagents. NETosis was monitored using a plate reader assay by measuring the production of extracellular DNA. This assay monitors the fluorescence generated by the binding of cell-impermeable DNA-binding Sytox Green fluorescent dye to NET DNA. Neither C5a nor fMLP (up to concentration of 2μM) induced NET formation within the observed 300-min time period (Figure (Figure1).1). However, stimulation of neutrophils with PMA resulted in NET generation after approximately 120min, as determined by Sytox Green plate reader assay (Figure (Figure1)1) and nuclear morphology changes (Figure S2 in Supplementary Material). As expected, the use of the NADPH inhibitor DPI abrogated PMA-induced NET formation with levels remaining near baseline. This confirms that PMA induces NETosis via the Nox-dependent pathway, and that C5a and fMLP on their own do not induce NETosis.
To identify whether stimulation of neutrophils results in the release of complement factors, Western blot analysis was performed on the cell pellets and cell-free supernatant (Figure (Figure2).2). Protein levels of the complement proteins CFP, C3, and CFB were compared for neutrophils induced for 30min either with C5a, fMLP, and PMA or with buffer. Complement proteins were not detected in the supernatant of non-activated neutrophils, with proteins being identified exclusively in cell pellet samples. The release of all three complement proteins was observed for all induction methods; however, the use of PMA elicited the greatest release of both CFP and CFB within the supernatant (Figures (Figures2A–C).2A–C). Pelleted samples for activated cells also contained a large amount of complement proteins: primarily CFP and CFB (and its activation product Bb). Furthermore, CFB-Bb was only identified in PMA-induced neutrophils (Figure (Figure2C).2C). These results indicate the capability of PMA to not only induce NETosis but also cause the greatest release of complement proteins in comparison to C5a and fMLP.
As the next step, we sought to identify whether CFP was capable of adhering to NETosing neutrophils and NETs. Immunofluorescence analysis performed on PMA-induced NETs showed the deposition of both CFP and MPO (another known NET-associated protein) on the surface of extracellular NETs (Figure (Figure3).3). CFP could also be detected on the neutrophil membranes after NET formation. Deposition pattern of both MPO and CFP shows substantial overlap throughout the extracellular DNA lattice structure. Therefore, neutrophils deposit CFP on NETs.
In order to analyze whether complement activation occurs on NETs, neutrophils were stimulated with PMA, washed and incubated with 20% complement active plasma for 15min in complement competent buffers. Under these conditions, both AP- and non-AP-mediated complement fixation can occur. Immunofluorescence microscopy was performed on these NETs to identify the deposition of terminal complement complex C5b–9. Images show that C5b–9 deposits on NETs (Figures (Figures44 and and5–middle5–middle column; Figures Figures4H4H and and5H5H with 2× magnified insets of representative areas). The use of DNAse I, which removes pre-formed NETs, causes a large decrease in the detection of C5b–9 deposition. These data suggest that the presence of NETs is necessary to activate complement cascade and deposit terminal complement complex.
To further identify the contribution of AP to NETosis, this experiment was repeated except that the plasma incubation step was performed in the presence of AP buffer allowing for AP activation only. Immunofluorescence microscopy images show that C5b–9 could be deposited via AP (Figure (Figure6;6; Figures Figures6I–K6I–K with 2× magnified insets of representative areas). To determine whether blocking CFP is sufficient to prevent complement activation, 4μg/ml anti-CFP antibody was added during complement activation. This antibody concentration was chosen because AP-dependent rabbit erythrocyte hemolysis was inhibited in the presence of >4μg/ml anti-CFP antibody (Figure S3 in Supplementary Material). Immunofluorescence microscopy reveals that blocking CFP fully prevents C5b–9 deposition in AP buffer conditions, but reduced C5b–9 deposition only slightly in complete buffer conditions (Figure (Figure6).6). Therefore, both AP- and non-AP-mediated complement depositions occur on NETs induced by PMA.
To test the ability of complement fixation on pathogens during NETosis, neutrophils were exposed to various strains of P. aeruginosa (mPA01, PAKwt, and PAKgfp). Similar to the PMA experiments described above, NETosis was monitored using Sytox Green plate reader assays. All three strains of these bacteria induced NETosis, and followed similar kinetics in terms of post-infection time response (Figure (Figure7).7). NETosis began at approximately 120min and continued to increase throughout the 300-min experimental time period. Furthermore, this induction is bacterial load dependent with both a faster response and a larger response observed when the MOI was increased from 10 to 100. To determine whether this NETosis induction was dependent on Nox, DPI was included in the media. Nox inhibitor DPI fully abrogated P. aeruginosa-induced NETosis for all three stains at both MOIs (p<0.05). This finding indicates that similar to PMA, bacteria-induced NETosis occurs in a Nox-dependent manner.
To determine CFP deposition during NETosis, specimens were immunostained. In the absence of serum, CFP was detected on both bacteria and NETs, and DNAse I treatment abolished CFP deposition (Figure (Figure8;8; Figure Figure8K8K with 2× magnified inset of a representative area). These results indicate that during P. aeruginosa-induced NETosis CFP is released from the neutrophils and deposits on NETs and bacteria.
To determine whether NET induction mediated by P. aeruginosa results in complement activation, C5b–9 deposition was determined by immunofluorescence microscopy. Images show that C5b–9 was deposited on NETs (Figure (Figure9;9; Figure Figure9K9K with a 2× magnified inset of a representative area). This effect was abolished when the NET DNA lattice was removed by DNAse treatment. Thus, complement deposits on NETs during NETosis.
To determine whether the AP is involved in P. aeruginosa-induced NETosis and C5b–9 formation, we first examined C5b–9 deposition under AP activation conditions. Immunofluorescence microscopy analysis shows that C5b–9 is deposited on NETs (Figure (Figure10;10; Figure Figure10K10K with a 2× magnified inset of a representative area).
Next, we used anti-CFP antibody to test the importance of CFP under AP activation conditions (Figure S3 in Supplementary Material). Induction of NETosis was performed using PAKgfp at an MOI of 100. After NET induction with PAKgfp, samples were maintained in complement competent RPMI media with the addition of 20% (v/v) autologous plasma (Figure (Figure11–first11–first column; Figure Figure11M11M with a 2× magnified inset of a representative area). The addition of anti-CFP antibody to these samples did not change the formation and deposition of C5b–9 (Figure (Figure11–second11–second column; Figure Figure11N11N with a 2× magnified inset of a representative area). Next, we incubated neutrophils with 20% (v/v) plasma in AP buffer. C5b–9 deposition was detected on NETs (Figure (Figure1111 – third column; Figure Figure11O11O with a 2× magnified inset of a representative area). Adding 4μg/ml anti-CFP antibody before incubating bacteria-induced NETs with plasma inhibited C5b–9 deposition on NETs (Figure (Figure11–fourth11–fourth column). Taken together, these data show that complement activation and progression to C5b–9 formation on NETs occurs via AP and non-AP pathways, and that AP activation depends on CFP.
Over the past decade, the ability of neutrophils to generate NETs has led to studies attempting to determine their function and involvement in disease. Although extensive studies have been performed, the exact functions of NETs, and their mechanism of action, remain to be completely elicited. NETs have been identified in several diseases that are associated with complement activation (1, 13, 14, 17, 33). In this study, we sought to understand the involvement of the complement system in the context of bacteria and NETs.
The use of varying reagents to induce NETosis has been established in many studies (4, 5). Applying these conditions, we found that PMA, but not C5a and fMLP, induces NETosis via the production of ROS. In order to obtain a more physiological impression of the mechanism of NETosis and the interplay with components of the complement system, we incorporated the use of P. aeruginosa in further studies. As previously seen (28–30), bacteria are capable of inducing Nox-dependent NETosis in a load-dependent manner.
Neutrophils recruited to sites of inflammation are a major determinant of AP activation (1). Neutrophil stimulation with PMA, C5a, and fMLP resulted in a quick release of the complement proteins CFP, C3, and CFB. These proteins are critical for the assembly of the AP convertase C3bBb, where CFP functions as a stabilizer (26). After demonstrating neutrophil release of complement proteins during NETosis, we confirmed that complement proteins became entangled within the NET structures. C5b–9 was deposited on the NETs, independent of the stimulus (PMA and PAKgfp). Deposition was abrogated by the use of DNAse, signifying the requirement of the NET lattices for the generation of C5b–9. This finding supports a role for NETs in inducing and/or enhancing complement activation, which is in keeping with the recently published observation that NETs can activate and deposit complement AP components (25). CFP also binds to DNA exposed by necrotic or apoptotic cells, and neutrophil-secreted CFP has been linked to a positive feedback loop between neutrophil and complement activation (1, 34–37). We also found CFP deposition on NETs, and the use of an anti-CFP antibody blocking the AP allowed us to identify the complement pathways involved in NET-mediated C5b–9 activation (38).
Complement factor P blockade was efficient in preventing terminal pathway activation when experimental conditions limited complement activation (i.e., C5b–9) on NETs to the AP. When the classical and lectin pathways were also allowed to be activated, this AP-specific blocking effect was lost. Fluorescence microscopy data suggest the possibility of CFP-mediated C5b–9 formation on NETs–data consistent with recent studies, indicating that both AP- and non-AP-mediated complement activation can occur on NETs (1, 5, 25, 37, 39, 40). CFP binding to targets via C3 fragments (alone or in context of the C3-/C5-convertases) is also possible.
Taken together, our results demonstrate that the “AP tool kit” present in the neutrophils are released upon stimulus, and deposits on targets, such as NETs. In the presence of plasma, NET formation results in terminal pathway activation via both CFP-dependent and -independent mechanisms. In NET-mediated diseases, the formation of NETs might trigger complement activation and exert secondary effects, such as cell injury and death. This is of great importance, as therapeutic complement inhibitors (e.g., eculizumab) are now available for clinical use (41, 42).
JY designed and carried out the experiments, interpreted data, and wrote the first draft of the manuscript; AC, MR, and FP contributed to designing and carrying out experiments, interpreting data, and writing the manuscript; DD provided technical assistance in designing and carrying out some of the experiments; MU provided the PAKgfp strain and contributed interpreting data; WK contributed designing experiments and interpreting data; NP and CL designed experiments, supervised the study, interpreted data, and wrote the final manuscript.
The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.
We thank the following individuals for technical assistance: Damien Noone, Hailu Huang, Hong Wang, and Jalil Nasiri from The Hospital for Sick Children for their help with phlebotomy. This study was funded by Canadian Institutes of Health Research (MOP-111012 to NP and MOP-259952 to WHAK), Natural Sciences and Engineering Research Council of Canada (436250-2013 to NP), and Cystic Fibrosis Canada (2619 to NP). JY is a recipient of Ontario Graduate Scholarship. This work is part of the MSc thesis of JY (University of Toronto, 2013).
The Supplementary Material for this article can be found online at http://journal.frontiersin.org/article/10.3389/fimmu.2016.00137
PMA and fMLP, but not C5a, induce ROS production in neutrophils. Neutrophils were activated with (A) C5a (1 or 2μM), (B) fMLP (1 or 2μM), and (C) PMA (20nM) and analyzed by flow cytometry for oxidative burst using dihydrorhodamine (DHR) 123. A significant ROS production was only observed for fMLP (1μM) and PMA (20nM). Results are given as median fluorescence intensity (MFI) from three independent experiments. Student’s t-test, *p<0.01.
Confocal images showing PMA-mediated kinetics of NETosis. Neutrophils were activated with PMA (20nM) to induce NET formation. Samples were fixed with 4% (w/v) paraformaldehyde and stained with DAPI for microscopy. (A) Four distinct nuclear morphologies (lobulated, delobulated, decondensed nuclei; NETs) can be identified during NETosis. (B) Percentage difference for nuclear morphologies was identified through manual counting of at least 118–220 cells from five different focal planes at 40× magnification. Data are presented as mean±SEM from three independent experiments. Statistical significance is shown only if percentage of nuclear morphology is significantly different compared to all other morphologies at the same time point. Two-way ANOVA with Tukey’s multiple comparison test, *p<0.05, #p<0.001, $p<0.0001.
Anti-CFP antibody concentrations of >4μg/ml block AP-mediated complement activation. Serial dilutions of a mouse monoclonal anti-properdin antibody were performed to determine the antibody concentration required to completely inhibit complement AP as determined by rabbit erythrocyte lysis. This was achieved using concentrations >4μg/ml. Data are presented as mean±SEM from three independent experiments. Student’s t-test, *p<0.05, **p<0.01.