Suppressor of cytokine signaling (SOCS)5 ameliorates influenza infection via inhibition of EGFR signaling

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Abstract

Influenza virus infections have a significant impact on global human health. Individuals with suppressed immunity, or suffering from chronic inflammatory conditions such as COPD, are particularly susceptible to influenza. Here we show that suppressor of cytokine signaling (SOCS) five has a pivotal role in restricting influenza A virus in the airway epithelium, through the regulation of epidermal growth factor receptor (EGFR). Socs5-deficient mice exhibit heightened disease severity, with increased viral titres and weight loss. Socs5 levels were differentially regulated in response to distinct influenza viruses (H1N1, H3N2, H5N1 and H11N9) and were reduced in primary epithelial cells from COPD patients, again correlating with increased susceptibility to influenza. Importantly, restoration of SOCS5 levels restricted influenza virus infection, suggesting that manipulating SOCS5 expression and/or SOCS5 targets might be a novel therapeutic approach to influenza.

DOI: http://dx.doi.org/10.7554/eLife.20444.001

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Influenza, commonly referred to as the flu, is a highly contagious disease caused by a virus. When an infected person coughs or sneezes, droplets containing the virus are released into the air. Other individuals nearby may breathe in the virus, which then enters the cells lining the lungs and multiplies.

Some people are more susceptible to the influenza virus than others. In particular, individuals with chronic obstructive pulmonary disease (COPD) often suffer much worse flu symptoms and are more likely to be admitted to hospital. COPD results from smoke exposure, including cigarette smoke and, in developing countries, the smoke from cooking fires, but it is not clear why individuals with COPD are more susceptible to the influenza virus.

The influenza virus gains entry to lung cells by manipulating receptors on the cell surface. A protein called SOCS5 is present inside these cells and has been suggested as a potential regulator of these receptors. Here, Kedzierski et al. reveal that SOCS5 plays a critical role in protecting lung cells in mice and humans from the virus.

The experiments show that mice lacking the gene that encodes SOCS5 were more susceptible to infection by the influenza virus, had more severe symptoms of disease and increased amounts of virus in their lungs. Further experiments in lung cells collected from human volunteers show that SOCS5 levels increased in both healthy smokers and non-smokers in response to influenza infection. Conversely, SOCS5 levels in lung cells of smokers with COPD remained low after infection. This suggests that SOCS5 might be an important factor in the susceptibility of these patients to influenza.

The next is step is to understand exactly how SOCS5 works, which may make it possible to develop new treatments that boost SOCS5 activity in influenza patients.

DOI: http://dx.doi.org/10.7554/eLife.20444.002

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Neutrophil Extracellular Traps Contain Calprotectin, a Cytosolic Protein Complex Involved in Host Defense against Candida albicans

Constantin Urban, David Ermert, Monika S. Schmid … (2009)

Introduction Neutrophils are an essential component of the innate immune response since neutropenia or impairment of neutrophil function results in microbial infections that are often fatal [1]. Microbes engulfed by neutrophils are efficiently killed by reactive oxygen species (ROS) and antimicrobial proteins within vacuoles [2]. Additionally, neutrophils [3] and two other granulocytes, mast cells [4] and eosinophils [5], release web-like extracellular traps that ensnare and kill microbes. Neutrophil Extracellular Traps (NETs) are released during a novel form of cell death that requires ROS produced by the NADPH-oxidase complex [6]. During this process, the nucleus decondenses and intracellular membranes disintegrate allowing the mixing of nuclear and cytoplasmic components. Eventually, the plasma membrane ruptures to release NETs, structures that contain chromatin and granule proteins. The overall composition of NETs has not been explored. Neutrophils of several species make NETs [7],[8],[9] and they might be important in the immune defense against bacteria and fungi [10],[11],[12],[13]. Whereas bacteria [3] and parasites [14] probably are killed by histones in NETs, in a previous study we found that purified histones did affect Candida albicans in vitro only poorly [13]. Thus, it remains to be determined whether histones or other antifungal effectors in NETs kill or inhibit fungi. This seems to be particularly of importance since previous reports have demonstrated that histones and histone peptides kill different fungal species such as Cryptococcus neoformans and Candida tropicalis [15],[16],[17]. Fungal pathogens, in particular C. albicans, cause an increasing number of severe infections with high mortality rates [18]. C. albicans is an opportunistic pathogen that can be part of the normal microbial flora of humans. In immunosuppressed patients the microbe can use a variety of virulence factors that enables it to exploit various host niches and to cause different diseases ranging from cutaneous to systemic infections [19]. A key characteristic of C. albicans is the ability to change growth morphology from budding yeast to filamentous forms: pseudohyphae and true hyphae [20]. A variety of external stimuli have been shown to induce the yeast-to-hyphae transition, such as serum, alkaline pH and temperatures above 37°C [21]. The ability to reversibly switch between different morphologies upon external stimuli appears to be essential for the virulence of C. albicans [22],[23]. Using a proteomic approach, we analyzed the qualitative and quantitative protein composition of NETs. We identified 24 different proteins, including the cytoplasmic calprotectin protein complex (also called Mrp8/14-complex or S100A8/A9) that has been shown previously by several groups to have potent antimicrobial properties [24],[25],[26]. S100A8 and S100A9 belong to the large group of S100 calcium-binding proteins and form a heterodimer, calprotectin, which is abundant in neutrophils, monocytes and early differentiation stages of macrophages [27]. In other cell types, such as keratinocytes and epithelial cells, the expression can be induced under inflammatory conditions [28]. The antibacterial and antifungal activity of the complex is reversible by Zn2+ [29] and does not require direct contact to the microbe [30],[31]. Therefore, it is thought that calprotectin chelates divalent metal ions that are required for microbial growth. This defense mechanism has been termed nutritional immunity [32]. Recently, Sroussi et al. proposed that the antifungal activity of calprotectin may be increased by oxidative stress [33]. Calprotectin is elevated in the extracellular fluids of patients with inflammatory disorders such as rheumatoid arthritis and vasculitis. Indeed, this complex is now used as a marker for inflammation [34]. Recently, calprotectin was described as a potential endogenous Toll-like receptor 4 (TLR-4) activator that promotes lethal endotoxic shock [35]. Despite all these important extracellular functions the complex lacks a secretion signal. A non-classical and tubulin-dependent secretion mechanism was shown in monocytes activated by inflammatory cytokines [36]. The mechanism by which this cytoplasmic dimer derived from neutrophils is able to interact with extracellular microbes in vivo is not completely understood. Here we show that calprotectin is released as the major antifungal protein in NETs. Our data indicate that at infection sites NET formation is a mechanism which ensures the interaction between cytoplasmic calprotectin and extracellular microbes at high local concentrations. Results Qualitative analysis of the protein composition of NETs We isolated neutrophils from healthy donors and induced them to form NETs using phorbol myristate acetate (PMA). After gently washing the NETs twice to remove unbound proteins, we solubilized NET-bound proteins with DNase-1 (Figure S1). NET proteins were digested with trypsin and analyzed by nano-scale liquid chromatography coupled matrix-assisted laser desorption/ionization mass spectrometry (nano LC-MALDI-MS). The identification quality is represented by the MS/MS spectrum of S100A9, a subunit of calprotectin found with this approach (Figure S2 A). A protein was considered to be associated to NETs when the identification criteria (see Methods) were fulfilled in at least two from a total of three independent samples. We identified 24 different proteins (Table 1, NET Database: http://web.mpiib-berlin.mpg.de/cgi-bin/pdbs/lc/index.cgi), nine of which were previously shown to localize in NETs by immunofluorescence microscopy [3],[37], which correlates well with the results of our approach. 10.1371/journal.ppat.1000639.t001 Table 1 Summary of identified NET proteins. Cellular localization Protein name Gene name Swissprot/TREMBL Granules Leukocyte elastase ELA2 P08246 Lactotransferrin LTF P02788 Azurocidin AZU1 P20160 Cathepsin G CTSG P08311 Myeloperoxidase MPO P05164 Leukocyte proteinase 3 PR3 P24158 Lysozyme C LYZ P61626 Neutrophil defensin 1 and 3 DEFA-1 and -3 P59665, P59666 Nucleus Histone H2A H2A Q9NV63+ Histone H2B: a) Histone H2B H2B Q16778+ b) Histone H2B-like H2B Q3KP43, Q6GMR5 Histone H3 H3 Q71DI3+ Histone H4 H4 P62805+ Myeloid cell nuclear differentiation antigen MNDA P41218 Cytoplasm S100 calcium-binding protein A8 S100A8 P05109 S100 calcium binding protein A9 S100A9 P06702 S100 calcium-binding protein A12 S100A12 P80511 Cytoskeleton Actin ( and/or ) ACTB, ACTG1 P60709, P63261 Myosin-9 MYH-9 P35579 Alpha-actinin (1 and/or -4) ACTN1, ACTN4 P12814, O43707 Plastin-2 LCP1 P13796 Cytokeratin-10 KRT-10 P13645 Peroxisomal Catalase CAT P04040 Glycolytic enzymes Alpha-enolase ENO1 P06733+ Transketolase TKT P29401 Proteins that localize to NETs. Proteins are organized by their localization in unstimulated neutrophils. Those identified to be NET-associated for the first time in this report are shown in blue. Swissprot/TREMBL accession numbers marked with a “+” denote possible forms of the protein that cannot be discriminated by this analysis. We identified two distanced groups of histone H2B. The larger group of individual histone H2B types we refer to as H2B and the other type that is less well characterized we refer to as “H2B-like”. All possible accession numbers can be found in the NET Database (http://web.mpiib-berlin.mpg.de/cgi-bin/pdbs/lc/index.cgi). We identified proteins that have a nuclear, granular or cytoplasmic localization in unstimulated neutrophils. Among the nuclear components, we confirmed the presence of all four subtypes of core histones and newly found the myeloid cell differentiation antigen (MNDA). We also identified eight granular proteins, five of which (neutrophil elastase (NE), lactotransferrin (LTF), cathepsin G (CG), myeloperoxidase (MPO) [3] and more recently proteinase 3 (PR3) [37]) were previously found associated with NETs. Azurocidin, lysozyme C (LysC) and α-defensins were not known to be NET components. Notably, we found eleven cytoplasmic proteins; two glycolytic enzymes, catalase, five cytoskeletal proteins and three S100 proteins. We confirmed NET association of twelve proteins found in this study by indirect immunofluorescence. Notably, the linker histone H1, bactericidal/permeability increasing protein (BPI), pentraxin 3 (PTX-3) and cathelicidin (CAP-18), were described as NET-associated [3],[38],[39], but we did not find them with this approach. Immunoblot analysis, however, confirmed the presence of BPI, but not of PTX-3 or CAP-18 (Figure S2 B–D). To evaluate the specificity of our approach, we analyzed all purification steps. As previously described, activated neutrophils release many unbound proteins into the supernatant [40] (Figure 1A and B, lane 2) which were removed by two washes with culture medium (Figure 1A and B, lane 3 and 4). NET-associated proteins were specifically released by Dnase-1 (Figure 1A and B) with or without protease inhibitor cocktail (lane 7 and 8 respectively). As an additional control for the specificity of our approach we demonstrated that two cytoplasmic proteins, glyceraldehyde-3-phosphate dehydrogenase (GAPDH) and lactate dehydrogenase (LDH), were present in the supernatant (Figure 1B, lane 2), but not in the nuclease-digested NETs (Figure 1B, lanes 7 and 8). 10.1371/journal.ppat.1000639.g001 Figure 1 Identification of NET-associated proteins. (A) Silver stained SDS-PAGE and (B) immunoblots with samples from NET protein purification procedure. Human neutrophils were stimulated to form NETs. Supernatants from unstimulated (lane 1) and stimulated (lane 2) neutrophils; first wash (lane 3); second wash (lane 4); medium containing DNase-1 incubated with unstimulated neutrophils (lane 5); DNase-1-free medium incubated with washed NETs (lane 6); medium containing DNase-1 incubated with washed NETs (lane 7); medium containing DNase-1 incubated with washed NETs including protease inhibitor cocktail (lane 8). To control the efficiency of the washing steps we increased the number of washes from two to nine (1 ml/wash, Text S1) in similar experiments as described above and determined the amount of calprotectin, a very abundant protein in neutrophils, in each fraction with ELISA. After NET formation calprotectin was present in similar amounts in culture supernatants and in wash 1 (Figure S2 E). The calprotectin concentration dropped in wash 2. These data agreed well with the immunoblot data presented in Figure 1B. However, we still detected minor amounts of calprotectin in wash 3 (55 ng/ml) and 4 (21 ng/ml). The amount decreased further in washes 5–7 until the calprotectin concentration reached the detection limit (1.6 ng/ml) of the assay in wash 8 and 9. Notably, from digestion of these intensely washed NETs with 1 ml RPMI containing 5 U/ml MNase we obtained approximately 170 ng/ml calprotectin which equals 23% of the total amount (720 ng/ml) according to our determination with the same ELISA under the same conditions. This is 20-fold more calprotectin than in wash 7 suggesting that calprotectin is bound to NETs. Taken together, this indicates that we purified and selectively enriched NET-associated proteins and that the NETs contain at least 24 proteins 15 of which were identified as NETs component for the first time (Table 1, marked in blue, and NET Database). Quantitative analysis of the protein composition of NETs To determine the relative amounts of NET proteins we quantified 15 of the 24 NET-associated proteins by immunoblotting (Table 2, Figure S3, NET Database). We purified NET proteins from neutrophils isolated from ten healthy donors. On average, NETs derived from 1012 neutrophils contained 3.58+/−0.28 g of protein and 2.24+/−0.51 g of DNA. This indicates a ratio of 1.67+/−0.26 g of protein per gram of DNA. 10.1371/journal.ppat.1000639.t002 Table 2 Quantified NET proteins. Protein name Molecular weight (KDa) µmoles (per g NET-DNA) % Molar amount mg protein (per g NET-DNA) Histone H2A 16.1 23.60±1.07 26.29 379.3±17.3 Histone H2B 15.3 21.50±0.77 23.95 298.9±10.7 Histone H3 13.9 13.02±1.39 14.50 199.2±21.3 Neutrophil elastase 25.4 5.24±0.85 5.84 133.0±21.5 Histone H4 11.4 3.96±0.39 4.41 45.2±4.5 S100A8 10.8 3.59±0.48 4.00 38.8±5.2 Lactotransferrin 76.0 2.46±0.28 2.74 186.6±21.6 Azurocidin 23.8 2.35±0.44 2.62 55.9±10.6 Cathepsin G 26.6 2.22±0,11 2.47 59.2±2.7 S100A9 13.2 1.27±0.12 1.41 16.7±1.6 Myeloperoxidase 78.4 0.91±0.07 1.01 71.3±5.3 Proteinase 3 24.0 0.64±0.13 0.71 15.5±3.2 Actin 41.8 0.15±0.02 0.17 6.3±0.9 Lysozyme C 14.5 0.12±0.036 0.13 1.8±0.5 Catalase 59.8 0.02±0.001 0.02 1.25±0.07 Quantified NET proteins are listed according to their µmolar amounts normalized to the amount of NET-DNA. Additionally, the calculated amount as a percentage of total NET protein content and the mass per g NET-DNA are shown. The data are representative mean values±mean deviation (n = 3) from two independent experiments. The core histones H2A, H2B, H3 and H4, were the most abundant proteins and account for 70% of all NET-associated proteins. The molecular mass of these proteins is decreased by approximately 2–5 kDa, when compared to histones present in the nucleus (Figure 2A). This modification is specific, since the masses of non-nuclear proteins do not change upon association with NETs. (Figure 2A, S100A8 and NET Database). Moreover, the stoichiometry of the four core histones is different in NETs as compared to intact nuclei. In unstimulated neutrophils, the core histones are present in similar amounts (Figure 2A, lane 2). In contrast, on NETs, H3 and H4 are found in lower amounts than H2A and H2B (Figure 2A, lane 7) and their molarity per gram NET-derived DNA is different (Table 2, NET Database). These observations correlated to high resolution Field Emission Scanning Electron Microscope (FESEM) analyses (Figure 2B and C). NETs consist of “smooth” stretches, probably only composed of histones and DNA (Figure 2B, white box), interspersed with globular domains that contain granular proteins [3]. The smooth stretches showed periodical signal intensities that are approximately 5 nm thick and 10 nm wide (Figure 2C). These dimensions are similar to those of nucleosomes [41] suggesting that smooth NET stretches are composed of stacked cylindrical nucleosomes. On different areas we found intensities with a similar horizontal periodicity but a lower vertical dimension consistent with the notion that the histone composition was changed during NET formation (Figure 2A). 10.1371/journal.ppat.1000639.g002 Figure 2 Histones are altered during NET formation. NETs from human neutrophils were washed and digested with DNase-1. (A) The NET-fraction (N) and the remaining pellet after DNase-1 digest (P) were analyzed by immunoblotting at the indicated time points. Unstimulated neutrophils served as controls. All core histones have a reduced molecular mass (2–5 kDa less) in NETs compared to the pellet fraction and the unstimulated control. A representative experiment out of three in total is shown. (B) High-resolution SEM analysis of NETs which consist of smooth fibers (white box) and globular domains (diameter 25–50 nm, arrows), scale bar = 100 nm. (C) High-resolution FESEM analysis of smooth stretch of a singular NET-fiber. Signal intensities were profiled vertically and horizontally showing similar diameters to nucleosomes (depicted as cartoon structure models taken from [41], with approximate horizontal and vertical diameters of 5 nm and 10 nm, respectively). One experiment out of two is shown. The most abundant non-histone NET protein was NE and the least abundant protein detected was catalase (5.24 and 0.02% of total protein, respectively). The 15 proteins we quantified represent 90.3% of the total protein in NETs. Thus, the remaining nine proteins and those that were potentially not identified in our approach represent 9.7% of the total NET proteins. NETs contained 39 ng of S100A8 and 17 ng of S100A9 per µg of DNA after 2 washes (Table 2 and NET database). Since each well yielded in average 3.8 µg DNA, we calculated 148 ng S100A8 and 65 ng S100A9 per well. Both proteins add up to a total of 213 ng/well. This correlates well with the quantification of calprotectin by ELISA (Figure S4 E) which yielded 170 ng/well. It is possible that the difference observed is due to the different number of washes in the two experiments. Taken together, the different quantification methods are comparable. Calprotectin localizes in NETs The association of calprotectin to NETs suggests that the complex is released during a specific form of holocrine secretion [42] referred to as NETosis [43]. Using indirect immunofluorescence we verified that calprotectin is released through NET formation in vitro (Figure 3A–F). A calprotectin-heteroduplex-specific antibody [44] confirmed that the dimer was cytoplasmic (red), and it overlapped with granular MPO (green) in the compact cytoplasm of unstimulated cells (Figure 3A). There was also a faint calprotectin signal within the nucleus (DNA stain, blue) consistent with the fact that cytoplasmic proteins with a mass below 30 kDa can diffuse through the nuclear pore complex. Thirty minutes after stimulation the neutrophils flattened as a sign of activation revealing a granular staining for MPO and a more dispersed cytoplasmic staining for calprotectin (Figure 3B). After stimulation for 1 hour, we observed partial colocalization of MPO and calprotectin in the cytoplasm (Figure 3C). This increased two hours after activation, when the chromatin decondensed and the nuclear membrane disassembled. At this time point calprotectin, MPO and DNA colocalized (Figure 3D). A proportion of the MPO signal remained in the area between the plasma membrane and the decondensed nucleus. Three and 4 hours post-activation, when the plasma membrane ruptured, calprotectin, DNA and MPO colocalized on NETs (Figure 3E–F). 10.1371/journal.ppat.1000639.g003 Figure 3 Neutrophils release calprotectin by forming NETs. (A–F) Confocal images of human neutrophils without stimulation (A), after 0.5 h (B), 1 h (C), 2 h (D), 3 h (E) and 4 h (F) after activation. Samples were stained with antibodies specific for the calprotectin heteroduplex (red) and for MPO (green). DNA was stained with DRAQ5 (blue). Calprotectin localizes to the cytoplasm and partially to the nucleus (A, arrow). After stimulation for 0.5 h (B) the neutrophils flattened and formed numerous vacuoles. This reveals a granular staining for MPO and a more dispersed cytoplasmic staining for calprotectin. After stimulation for 1 h (C) the neutrophils round up slightly. The MPO and calprotectin stain partially overlap in the cytoplasm. After stimulation for 2 h (D), calprotectin, MPO and nuclear DNA start to colocalize in the decondensed nucleus (purple). After 3 h (E) and more so after 4 h (F) of stimulation, the cell membrane ruptures and calprotectin is released in NETs colocalizing with MPO and DNA. Scale bar = 10 µm; one experiment out of two is shown. (G–I) Subunits of calprotectin S100A8 and S100A9 are released after cell death during NET formation and not by degranulation. NET formation was induced with PMA and degranulation using formyl-met-leu-phe (f-MLP). (G) Neutrophil death was monitored by quantification of LDH activity in supernatants calculated as means±s.d. (n = 3). (H) Release of S100A8, S100A9, lactotransferrin (LTF) and myeloperoxidase (MPO) were analyzed by immunoblotting. one experiment out of two is shown. (I) Quantification of immunoblots using 2D densitometry analyzing S100A9 protein preparations from supernatants (lane 1), MNase-digested NETs (lane 2) and cell remnants indigestible for MNase (lane 3). Values were calculated as means±s.d. (n = 3) from one experiment out of two. To determine whether calprotectin is released during NET cell death or before the plasma membrane ruptures, we compared neutrophils that were treated either with PMA to stimulate NET formation [3] or with the microbial peptide formyl-met-leu-phe (f-MLP) to stimulate degranulation [40]. At the indicated time points we monitored neutrophil cell death by quantifying extracellular LDH (Figure 3G) and solubilized NET-bound proteins by adding Dnase-1 (Figure 3H). We subsequently collected the supernatants containing both unbound and NET-bound proteins to determine the total release of the indicated proteins during degranulation and NET formation. We analyzed the samples by immunoblotting and probed for the calprotectin subunits S100A8 and S100A9 as well as for two granular proteins, MPO and LTF (Figure 3H). Thirty minutes after f-MLP treatment, neutrophils released MPO and LTF, but not S100A8 nor S100A9. Four hours after f-MLP treatment, minor amounts of S100A8 and S100A9 appear in the supernatants and less than 10% of the cells were dead. During NET formation, MPO and LTF were secreted in low amounts before S100A8 and S100A9 were released confirming that neutrophils also degranulate upon PMA-activation [40]. Significant amounts of extracellular calprotectin were only found 3 to 4 hours after stimulation, when 40 and 70% of the neutrophils respectively were dead. Furthermore, we determined how much of the total calprotectin amount present in the neutrophil actually binds to NETs. After stimulation we quantified S100A9 released into the supernatant, bound to NETs and remaining in the cell remnants by quantitative immunoblots. Approximately 30% of the total S100A9 was bound to NETs (Figure 3I). Equivalent amounts of S100A9 were found in the supernatants and in the nuclease-resistant cell remnants (Figure 3I). This agrees with a previous report showing that after nuclease treatment, a proportion of chromatin remains within the debris of the neutrophil [6]. We obtained similar results for S100A8 (data not shown). Moreover, a calprotectin specific ELISA (Figure S2 E) confirmed the finding that similar amounts of calprotectin are in NETs (170 ng/ml) compared to unbound in the culture supernatant (200 ng/ml) and that this corresponds to 23% and 28% of the total amount of calprotectin respectively. Taken together, this demonstrates that approximately one third of calprotectin in vitro is released from neutrophils in NETs. Calprotectin is a major antifungal component of NETs As we have reported previously NETs can control the growth of C. albicans yeast and hyphal forms [13] and more recently this has been demonstrated for Aspergillus nidulans conidia and hyphae as well [45]. Additionally, we have now added Cryptococcus neoformans to this list of NET inhibited fungi (Figure S4 A). However, the mechanism behind the antifungal activity remained unclear. In this study we confirmed the previous results by exposing C. albicans to NETs at different multiplicities of infection (MOI) and at different temperatures to induce yeast-form and hyphal growth (Figure S4 B and C). Under all conditions the antifungal activity of NETs was very similar reducing CFU counts approximately 100-fold, but not when the NETs were degraded with nucleases (Figure S4 D). We confirmed the antifungal activity of NETs against hyphae macro- and microscopically (Figure S4 E and F). As expected, hyphae grew in media but this growth was inhibited when they were incubated with NETs. We corroborated the anti-candidal activity of NETs and showed similar results measuring the viability of C. albicans hyphae with the tetrazolium dye XTT (Figure S4 G–I). This method, unlike CFU enumeration, does not require the dispersion of the hyphae [46]. We confirmed that NETs made by neutrophils stimulated with C. albicans contain calprotectin, lactotransferrin and catalase (Figure S4 J). This suggests that NETs have a qualitatively similar composition regardless of the stimulus used to activate the neutrophil. Calprotectin chelates essential metal ions, such as Zn2+ and Mn2+ resulting in reduced microbial growth [30],[31]. Consistent with this mechanism of action the antifungal activity of NETs was inhibited by increasing concentrations of Zn2+ (Figure 4A) or Mn2+ (Figure 4B). The direct role of calprotectin in NETs was tested with immunodepletion experiments. We purified NET proteins by DNase-1 treatment and concentrated the samples 10-fold with 3.5 kDa cut-off membranes. The samples were incubated with a mixture of immobilized antibodies directed against the individual subunits S100A8 and S100A9 (Figure 4C). Immunodepletion of calprotectin, but not treatment of NET preparations with isotype-matched controls, completely abrogated the growth-inhibitory activity of NET proteins showing that this dimer is a major antifungal component of NETs. It is important to note that immunoblotting of the supernatant showed that S100A9, but not lactotransferrin, was depleted (Figure 4C). A complementary assay where C. albicans viability was measured using XTT confirmed the result of the CFU based assay (Figure S4 H). Furthermore, NETs made by calprotectin-deficient mice inhibited C. albicans growth less efficiently than wild-type mouse NETs (Figure 4D) which was confirmed with an XTT assay as well (Figure S4 I). Notably, neutrophils from both genotypes made NETs with similar efficiency (Figure S5 A–F). 10.1371/journal.ppat.1000639.g004 Figure 4 Calprotectin is a major antifungal component in NETs. Human neutrophils were induced to make NETs, washed and infected with C. albicans (MOI 0.01) and incubated overnight at 30°C (yeast-form growth). Antifungal activity was determined by counting CFU. Increasing concentrations of ZnSO4 (A) and of MnCl2 (B) abolished the antifungal activity of human NETs. Shown are means±s.d. (n = 3) from one representative experiment out of three. (C) Purified human NET proteins were concentrated and depleted of calprotectin using immobilized anti-S100A8 and anti-S100A9 combined. Controls were incubated with mouse IgG1 isotype matched antibodies. C. albicans was incubated with these extracts overnight at 37°C (hyphal growth) and CFU were determined. Shown are means±s.d. (n = 3) from one experiment out of two. The inset shows an immunoblot confirmation of the depletion. The blots were probed for S100A9 and lactotransferrin (LTF). The lanes are arranged in the same order as indicated for the antifungal assay below the graph. (D) Mouse neutrophils isolated either from wild-type or calprotectin-deficient mice were induced to make NETs and then infected with C. albicans (MOI 0.02) incubated overnight at 37°C (hyphal growth) and CFU counts were determined; n. d. = no detectable CFU. Shown are means±s.d. (n = 3) from one experiment out of two. (E) Co-precipitation assays of indicated fungi incubated with NET-bound calprotectin (produced by MNase treatment) or soluble calprotectin. Microbes were pelleted, supernatants removed and washed pellets were analyzed using immunoblots against S100A9. Shown are means±s.d. (n = 3) of one representative experiment out of three. The antimicrobial activity of calprotectin does not require direct contact between microbe and protein, which is consistent with its inability to bind microbial surfaces [30],[31]. Regardless, at high concentration this chelator is more efficient. Therefore, we tested whether NET-bound calprotectin binds to microbes. Seeded neutrophils were induced to release NETs. We collected the supernatant after NET formation that contained unbound and soluble calprotectin. The NETs were subsequently washed twice and digested with MNase, a non-processive nuclease, to generate NET fragments. We exposed the indicated fungi to these NET fragments or supernatants containing soluble calprotectin and pelleted the microbes afterwards. The microbial pellets were washed three times. Immunoblotting in Figure 4E shows that NET-associated, but not soluble S100A9, binds to the fungi. We obtained similar results for S100A8 (data not shown). Taken together, these data indicate that calprotectin is a major antifungal NET component. We conclude that the presentation of this dimer in NETs provides a high local concentration on the surface of microbes. Calprotectin is required for innate antifungal defense We addressed the role of calprotectin in C. albicans infections comparing wild-type to S100A9 knockout mice. These animals transcribe the mRNA for S100A8 but are deficient in S100A8 and S100A9 protein [47]. C. albicans exploits different host niches and we investigated three of them: (i) subcutaneous inoculation, which leads to confined abscesses; (ii) intranasal infection, which causes pulmonary candidiasis and (iii) intravenous challenge, which mimics disseminated systemic candidiasis. The dimensions of the abscess lesions were measured at indicated time points. On average, the area of the lesions in calprotectin –deficient mice were twice as large (200 mm2) as compared to those of wild-type animals (100 mm2) measured at two, four and six days after inoculation (Figure 5A–E). At day four, 60% of the abscesses in calprotectin-deficient mice ulcerated as indicated by extensive necrosis which also included the epidermis (Figure 5A). Consistent with a less severe progression, abscesses of wild-type animals were restricted to subcutaneous areas without involvement of outer skin layers (Figure 5D). Eight days after inoculation, the size of the abscesses in calprotectin-deficient and wild-type mice was similar, indicating that calprotectin is essential for the initial phase of the disease but that eventually other mechanisms clear the infection. Interestingly, in about 30% of the infected knockout mice, but not in wild-type controls, infection spread from the original abscess to surrounding areas of the skin (Figure S6). In our experimental settings neutrophil recruitment (Figure S5 G) and NET formation (Figure S5 A–F) was similar in knockout and wild-type animals. 10.1371/journal.ppat.1000639.g005 Figure 5 Calprotectin is required for antifungal immunity. (A–D) Subcutaneous abscesses induced with C. albicans of representative calprotectin -deficient (a and b) and wild-type (c and d) mice at days 2, 4, 6 and 8 post infection (p.i.). (E) the area of the abscess lesions of calprotectin-deficient animals compared to wild type was monitored over 8 days p.i. (n = 10); abscesses of calprotectin-deficient animals are significantly larger at days 2 (P = 0.0007), 4 (P = 0.0012), 6 (P = 0.0003) but not at day 8. (F) calprotectin-deficient mice are more susceptible to intranasal challenge with C. albicans than wild-type mice (n = 10, P = 0.0001). (G) Fungal load in the lungs after intranasal challenge with C. albicans is significantly higher in calprotectin-deficient compared to wild-type mice (n = 11; P = 0.048). (H) Calprotectin-deficient mice are more susceptible to intravenous challenge with C. albicans than wild-type mice (n = 10, P = 0.0159). In pulmonary candidiasis, wild-type mice show disease symptoms within three days but recover and survive. In contrast, calprotectin-deficient mice carried a significantly higher fungal load than wild-type controls (Figure 5F) and succumbed to C. albicans (Figure 5G). Intravenous challenge is lethal in both calprotectin-deficient and wild-type mice. However, the knockout animals died significantly earlier (Figure 5H), suggesting that also in deep-seeded infection sites, the antimicrobial activity of calprotectin contributes to containment of fungal growth. This is consistent with a previous report that calprotectin reduces bacterial load in systemic staphylococcal infection [31]. We conclude that calprotectin is required for an effective acute antifungal response. NETs released during C. albicans infection contain calprotectin We tested whether NET formation is a route of calprotectin release in vivo during subcutaneous (Figure 6A–F) and pulmonary C. albicans infection (Figure 6G–M). Histological analysis of 6 day old abscesses showed fungal foci surrounded by neutrophils (Figure 6A). Using staining with hematoxylin and eosin (H &E), we observed abundant extracellular DNA in web-like structures in these areas (Figure 6B). To confirm that these structures were NETs, the samples were labeled with antibodies directed against MPO (green) and histones (blue) and analyzed by immunofluorescence microscopy. Labeling with anti-S100A9 antibodies demonstrated that these structures also contain calprotectin (Figure 6, C–F). 10.1371/journal.ppat.1000639.g006 Figure 6 Calprotectin is present in C. albicans induced NETs in vivo. Wild-type mice were challenged subcutaneously (A–F) or intranasally (G–L). (A,B, G and H) Hematoxylin & Eosin (H & E) stainings of sections in areas with strong neutrophil infiltration show extracellular DNA (hematoxylin positive) representing NETs and indicated with arrows, scale bars 50 µm in (A,G) and 20 µm in (B,H). Confocal images of indirect immunofluorescence from sections of abscesses 6 days after subcutaneous challenge (C–F) and lungs 24 h after intranasal challenge (I–L) stained with primary antibodies against S100A9 (red), MPO (green) and histone (blue). NETs are web-like and diffuse areas where all signals superimpose indicated by white arrows. Scale bars = 20 µm. In infected lungs, there was a strong neutrophil infiltration into C. albicans colonized bronchioles one day after challenge. These areas showed web-like structures of extracellular DNA (Figure 6 G and H). MPO and histones (Figure 6M, arrow), as well as calprotectin(Figure 6I–l) colocalized in these structures that unfold into the lumen of the bronchioles. To demonstrate that C. albicans and NETs interact in the lumen of bronchioles we analyzed these sections by Scanning Electron Microscopy (SEM) (Figure 7A). Areas colonized by C. albicans show web-like structures that cover fungal surfaces (Figure 7B and C). These structures have a very similar morphology and dimension to those observed for NETs in vitro. Taken together these observations demonstrate that NETs and C. albicans interact in vivo and that these NETs contain calprotectin. We propose that release and NET-association of calprotectin from neutrophils could contribute to contain fungal infections. 10.1371/journal.ppat.1000639.g007 Figure 7 Fine structure of C. albicans induced NETs in pulmonary infection. (A–C) SEM analysis of sections from C. albicans infected mouse lungs 24 h after intranasal challenge. (A) Image shows a bronchiole colonized with C. albicans and infiltrated by host immune cells, b = bronchiole. (B) High resolution image of boxed area in (n) shows respiratory epithelium of the bronchiole colonized with C. albicans yeast-form (arrow) and hyphae (arrowhead). (C) High resolution image of boxed area in (O) showing NETs covering fungal surfaces (arrow). Scale bars in (N) = 100 µm, in (O) = 10 µm and in (P) = 2 µm. Discussion The analysis presented here identified 24 NET-associated proteins. Nine of the 24 proteins were described previously as NET-associated which correlates well to our approach. Fifteen proteins were hitherto unknown to be NET-bound and their association to NETs was confirmed by immunoblotting and indirect immunofluorescence (NET Database). The data underscore the specificity and reproducibility of the analysis. Moreover, the composition of the identified NET proteins was very similar among different neutrophil donors (NET Database). The small number of identified NET proteins is surprising since the neutrophil cell membrane ruptures during the process of NET release. Additionally, we used mild washing conditions for the isolation. Therefore, the protein incorporation into NETs appears to be selective. We found that, in addition to five nuclear proteins, the NETs contain eight cationic granule proteins. Furthermore, we identified eleven cytoplasmic proteins which have neutral to acidic isoelectric points. This suggests that charge is not an exclusive requirement for binding to NETs. In previous reports histone H1 [3], bactericidal/permeability increasing protein (BPI) [3], cathelicidin (CAP-18) [37],[39] and pentraxin 3 (PTX-3) [38] have been described as NET-associated proteins. In contrast, we did not find these proteins in our MS approach. We further investigated these proteins by immunoblot (Figure S2B–C). We did not obtain clear results for histone H1 because commercially available anti H1-antibodies are cross-reactive with other histones (data not shown). This remains to be clarified. We could, however, detect the 25 kDa cleavage product of BPI in both neutrophil lysates and NET extracts. The failure of MS analysis to detect BPI remains unclear (Figure S1B). Notably, we detected CAP-18 and PTX-3 in neutrophil lysates but not in NET extracts. Therefore the presence of these two proteins in NETs should be further investigated. It is possible that these proteins are very loosely attached to NETs, that they are present in very low amounts, that they were lost during the isolation procedure or that the MS analysis failed to detect them. Notably, neither by immunoblotting nor by MS did we find GAPDH, a very abundant cytoplasmic and highly cationic protein. This supports the assumption that NET binding is not exclusively mediated by charge. Quantification by immunoblots showed that 15 NET proteins comprised 90% of the total protein amount in NETs, indicating good coverage. Detection of 1.25 ng of catalase per µg NET-DNA underscored the sensitivity of the approach. As the amounts of NET proteins were similar in different donors the approach was also reproducible (NET Database). More importantly this suggests that NETs do not assemble randomly and are similar in many individuals. The core histones are the major protein components in NETs as shown by the quantitative analysis. However, we found a reduction of the molecular mass of NET-associated compared to chromatin histones, possibly caused by post-translational modifications. Indeed, histone H3 is deiminated during NET formation in HL60 cells but this remains to be confirmed in primary neutrophils [48]. Additionally, the stoichiometry of the four histones is different in NETs when compared to chromatin. There is less H3 and less H4 in NETs than in chromatin, but the significance of these differences remain unclear. High resolution FESEM correlated with this finding. We determined a transverse periodicity in individual NET-fibers that was similar to the actual dimensions of intact and partially degraded nucleosomes. The role of some of the identified cytoplasmic proteins in NETs is unknown but there are indications for potential functions. Enolase, for example, has been found to be a plasminogen activator on leukocyte surfaces, although the secretion mechanism is unknown [49]. NET-association could explain how enolase is released allowing it to participate in tissue remodeling at inflammatory sites. We identified S100A8 and S100A9 that form the heterodimer calprotectin as NET-associated proteins by MS analysis and by indirect immunofluorescence. The dimer lacks a secretion signal and localizes to the cytoplasm as well as partially to the nucleus of unstimulated neutrophils. Therefore, the proteins must be released in order to function as an antimicrobial protein against extracellular pathogens. The release of calprotectin by neutrophils [50] and other cells [36] in response to specific stimuli was previously reported. Our data, however, establish NET formation as a hitherto unrecognized mechanism of calprotectin release in neutrophils. Interestingly, incubation of C. albicans with neutrophils was reported to increase extracellular calprotectin and, simultaneously decrease neutrophil viability [51]. The release mechanism, however, remained unknown. Consistent with these data we reported that C. albicans induces neutrophils to form NETs and die in the process [13],[52]. Here, we demonstrate that NET formation induced by C. albicans during infection is a novel route for release and presentation of calprotectin in vivo. We additionally showed that calprotectin tightly binds to NETs as several intense washes cannot remove calprotectin from NETs (Figure S2 E) or NET-bound calprotectin from fungal surfaces (Figure 4E). In contrast the unbound proportion of calprotectin released during NET formation does not adhere to fungal surfaces. These findings suggest that calprotectin, although it is abundant in unstimulated neutrophils, specifically interacts with NETs. We determined that 30 and 23% of the total calprotectin was NET-associated by immunoblot densitometry (Figure 3I) and ELISA (Figure S2E respectively). These values indicate that a significant proportion of the protein complex associates with NETs. Additionally, approximately the same amount of calprotectin is released to the supernatant. Thus, probably 50–60% of total cellular calprotectin might become extracellular during NET formation. It is important to note that with other methods previous publications determined the total amount of calprotectin to be approximately 5 µg/106 human neutrophils [53],[54]. Here we report approximately a ten-fold lower amount of calprotectin (0.43 µg/106). This difference could be due to the methodology, donor variation and the technique to count cells. Our functional assays on the inhibition of C. albicans growth by NETs suggest that calprotectin in NETs might have a biological implication due to the high local concentration and close contact to the microbe. In agreement with the findings discussed above we showed that calprotectin is a major antifungal protein in NETs affecting C. albicans and C. neoformans. Addition of Zn2+ or Mn2+ and immunodepletion of calprotectin abrogated the antifungal activity of NETs. This is consistent with previous reports showing that calprotectin is microbiostatic in vitro because it chelates ions [24],[25],[26],[30]. Notably, an enhancement of calprotectin's antifungal activity by oxidative stress has been reported recently [33]. This could as well be the case in NETs, since the release requires a robust oxidative burst to occur before [6]. Other NET proteins were not affected by calprotectin depletion (Figure 4C), but nonetheless NETs contain more antifungal proteins, such as azurocidin and LTF. Interestingly, addition of Fe2+ did not have an impact on C. albicans in our experimental settings (data not shown), suggesting that calprotectin, rather than LTF, controls C. albicans under these conditions. This is probably also true for C. neoformans. Fractionations of neutrophil compartments revealed previously that azurophilic granule extract inhibited cryptococcal growth to 80% and cytoplasmic extract to 70% [55]. The group identified calprotectin as the one single effector in the cytoplasm responsible for the strong inhibition correlating well with our findings. Whether other proteins contribute to the antifungal activity of NETs still remains to be determined (Figure 4D). Moreover, our data demonstrate for the first time that calprotectin is required for the innate immune defense to C. albicans infections. We detected abundant NETs in subcutaneous abscesses (Figure 6A–F), lungs after pulmonary challenge (Figure 6 G–M) and kidneys from systemic candidiasis (data not shown), and we propose that they prevent the spread of C. albicans. This might be in particular important for C. albicans hyphae that are too large to be engulfed and is consistent with our previous finding that hyphae are more potent NET inducers as yeasts [52]. Thus, we conclude that NET formation could serve as an additional neutrophil-mediated anti-hyphal mechanism complementing the previously described damage of hyphae by oxidative products and the myeloperoxidase system [56],[57]. The NET-released calprotectin reduces C. albicans growth correlating well with the more severe disease symptoms we observed in calprotectin-deficient mice as compared to wild type: increased abscess sizes upon subcutaneous challenge and higher fungal loads in lungs upon intranasal challenge. Indeed, the importance of NETs in reducing dissemination seems likely to be similar to that observed in streptococcal pneumonia [10]. This is consistent with a stronger survival phenotype of calprotectin in pulmonary compared to systemic infection, where C. albicans was disseminated from the outset. We and others determined neutrophils as a major source of calprotectin in infected tissue under the given conditions (data not shown, [31]). However, calprotectin has been reported to be expressed by other myeloid and epithelial cells and to be involved in numerous other immune-related functions, such as chemotaxis, regulation of the NADPH oxidase complex, activation of Toll-like receptor 4 and induction of apoptosis [35],[58],[59],[60]. Any of these functions may contribute to the phenotypes in C. albicans infection we report here. So far we have not identified a component that is exclusively required for NET formation but not other neutrophil functions. Therefore, we do not have a tool to determine the actual contribution of NET-associated calprotectin in the described infection phenotypes. We propose that differential localization of calprotectin, e. g. unbound and NET-bound, may enable the protein to be multi-functional. Our analysis revealed a simple, stable and reproducible composition of NETs. These data are likely to help investigate the functions of NETs in infections and in other disorders such as autoimmune diseases. Materials and Methods Mice, cells and strains S100A9 −/− mice [47] that are deficient in both calprotectin subunits S100A8 and S100A9 protein were backcrossed 7 times into C57 BL/6. These mice and C57 BL/6 controls were bred in our animal facility. All animal experiments were in compliance with the German animal protection law in a protocol approved by the Landesamt fur Gesundheit und Soziales, Berlin. Human peripheral blood neutrophils and mouse bone marrow derived neutrophils were isolated as described [6],[61]. We used C. albicans clinical isolate SC 5314 [62]. C. neoformans strain DSM 11959 was obtained from the German Collection of Microorganisms and Cell Cultures (DSMZ). C. albicans was cultured overnight in YPD (1% yeast extract, 2% bacto peptone and 2% glucose) at 30°C and C. neoformans in the same medium at 37°C. Cells were washed 3 times in PBS prior to the experiments. Cell numbers were calculated by OD600 correlation (C. albicans: 1 OD600 = 3×107 cells/ml; C. neoformans: 1 OD600 = 6×107 cells/ml). For experiments with neutrophils and NETs C. albicans was either incubated at 37°C to induce hyphae or at 30°C to preserve yeast-form growth. C. neoformans was always incubated at 37°C. All infection assays were performed in RPMI medium. Purification of NET proteins Human neutrophils were seeded in 12-well tissue culture plates to a density of 1.7×106 ml−1 (RPMI 1640 without phenol red). 1.7×106–3.4×106 (according to 1–2 wells) and 1.7×107 (according to 10 wells) neutrophils were used for analytical and MS identification respectively. Neutrophils were activated with 20 nM PMA for 4 h at 37°C in a 5% CO2 atmosphere. Each well was carefully washed twice after removing the supernatant by pipetting 1 ml of fresh and pre-warmed RPMI into the well along the wall of the well. Each wash was incubated for 10 min at 37°C. Subsequently the NETs were digested for 20 min in 1 ml RPMI with 10 U/ml DNase-1 (Worthington). DNase-1 was stopped with 5 mM EDTA (final concentration). The samples were centrifuged at 300×g to remove whole cells and then at 16,000×g to remove debris. Proteins were acetone precipitated. Four matched samples from different wells were pooled together and transferred to a 30 ml glass corex tube and 16 ml of ice-cold acetone (−20°C) was added. For precipitation samples were incubated overnight at −20°C and then centrifuged at 10 000×g for 30 min at 4°C. The protein pellet was washed with 1 ml 80% acetone buffered in 20 mM Tris-HCl pH 8.0 and solubilized in 120 µl SDS loading buffer or prepared for MS identification as described below. For verification of our purification procedure we followed each step of the isolation procedure by analysis of the respective sample using silver stained SDS electrophoresis and immunoblotting. We harvested all steps including the supernatant after 4 h incubation, the washes 1 and 2, and the NET digests. As controls (i) NETs were mock-digested with nuclease-free RPMI and (ii) unstimulated neutrophils that did not release NETs were washed twice and incubated with RPMI containing 10 U/ml Dnase-1 for 20 min at 37°C. Each sample was derived from one well containing 1.7×106 neutrophils in a volume of 1 ml. Four samples out of 4 wells were pooled, acetone precipitated, solubilized in 120 µl SDS loading buffer and boiled for 3 min. To account for potential protein loss due to proteolytic activity in the samples a complete purification procedure was performed in the presence of protease inhibitor cocktail (Sigma P1860; 1∶200) added to the wells 2 h after stimulation start as described above. Protease inhibitor cocktail was additionally present in all media used for washing and digestion. For analysis of the purification steps we loaded 30 µl of each pooled sample (equaling 1 well and 1.7×106 neutrophils) on SDS protein gels (Tris-HCl 10–20%). The gels were either silver stained as described elsewhere [63] or immunoblotted by transferring to a PVDF membrane (Immobilon 40; Millipore). The immunoblots were performed as described in detail for the quantitative blots below. The following primary antibodies were used: α-elastase (Calbiochem 481001, 2 µg/ml), α-azurocidin (Sigma N5662, 0.5 µg/ml), α-histone H2B (Upstate 07-371, 1 µg/ml), α-S100A8 combined with α-S100A9 (Acris BM4029 and BM4027, both at 2.5 µg/ml), α-catalase (Sigma C0979, 1 µg/ml), α-GAPDH (Labfrontier LF-PA0018, 0.5 µg/ml), α-LDH (Abcam ab7639, 1 µg/ml). As secondary antibodies horseradish-peroxidase-conjugated F(ab')2 fragments (Jackson ImmunoResearch) were used in 1∶20 000 dilutions. For MS analysis NET proteins from 3 different donors were purified independently in the absence (donor 1–3, named sample 1–3) or presence of the protease inhibitor cocktail indicated above (donor 1–2, named sample 4 and 5). DNA-concentration, neutrophil elastase activity and reactive oxygen species (ROS) were measured as described previously [6]. Sample preparation and digestion for MS Samples 1–5 (acetone precipitates from NET-purifications) were solubilized in 40 µl of 500 mM triethylammonium bicarbonate buffer pH 8.5 (TEAB) and reduced with 2 µl of 50 mM tris-(2-carboxyethyl)phosphine (TCEP) for 60 min at 60°C. After alkylation with 1 µl of 200 mM methyl methanethiosulfonate (MMTS) at RT for 10 min each sample was incubated overnight at 37°C with 10 µl of a 200 µg/ml trypsin solution, solubilized in 500 mM TEAB. The reaction was stopped with 1 µl of a 10% TFA solution to obtain a final concentration of TFA of approximately 0.2%. The sample was centrifuged for 10 min at 13800×g and the supernatant used for LC/MS analysis. Nano-LC/MALDI-MS The samples were analyzed by bottom-up nano-LC/MALDI-MS as described in detail in the NET Database. Proteins were digested with trypsin and the resulting peptides separated by nano-LC (Dionex). Peptides were fractionated (Probot microfraction collector, Dionex) and analyzed with a 4700 Proteomics Analyzer (Applied Biosystems) MALDI-TOF/TOF instrument. The criterion for the identification of a protein was a minimum number of 3 peptides fulfilling the Mascot homology criteria. Candidates with two peptides fulfilling these criteria were verified by checking the fragmentation rules, such as hypercleavage sites (Asp, Glu, Pro), the appearance of common immonium masses and mass losses [64]. A protein was considered as localizing to NETs only when found in at least 2 independent samples from different donors. Exceptions are MNDA, actinin and lysozyme C. MNDA and actinin were identified with one peptide in independent samples only, however the peptide is unique to both proteins within the IPI-database. Presence of lysozyme C in NETs was verified by immunoblotting. The MS analysis is described in more detail on the NET database and in the supporting materials. Quantification of NET proteins NET-associated proteins were quantified by immunoblot as described [65]. Neutrophils were purified from 10 different healthy donors, proteins were quantified using the DC assay (Biorad) and DNA was quantified with Pico Green™ (Invitrogen) [6]. We isolated NET proteins as described in ‘Purification of NET-proteins’. From each donor we prepared 12 times 1.7×106 neutrophils each seeded in 1 ml RPMI per well in a 12 well tissue culture plate. NETs from 10 wells were digested with 5 U/ml MNase (Fermentas), a non-processive nuclease that cuts DNA at linker sites. 2 wells were mock-digested with nuclease-free medium as controls. We measured 3 independent pools made from 3 different donors each to average samples. For each quantification, the 3 averaged samples and 6 different concentrations of the respective purified protein, used as a standard, were loaded on the same SDS-protein gel (Tris-HCl 10–20%). The gel was transferred to a PVDF-membrane (Immobilon 40; Millipore) in a liquid transfer system for 2 h at 80 V. The membrane was blocked for 1 h in PBS with 5% skim milk powder, washed 3 times in PBS with 0.05% Tween-20 (PBST). Primary antibodies (listed in the NET Database) were diluted in PBST with 1% BSA (PBST-BSA) and incubated with the membrane for 1h at room temperature or at 4°C over night. After 3 washes in PBST horseradish-peroxidase-conjugated F(ab')2 fragments (Jackson) were used as secondary antibodies and incubated for 30 min. The blots were developed with chemiluminescence and detected in a LAS 3000 camera (Fujifilm Europe). The signal intensities of the bands were analyzed by 2D densitometry (array imager software 4.15, Raytest) and the concentration calculated based on the purified protein standards in a linear range of analysis. The amount of sample was adjusted to be within this range. To confirm the reproducibility of our quantification approach, we prepared a separate batch of pooled NET proteins from additional 5 donors and quantified again 7 randomly chosen NET proteins producing comparable results (NET Database). Purified proteins as standards for quantitative immunoblots Recombinant human S100A8 and S100A9 for quantification were purified as previously described [66]. cDNA of human S100A8 and S100A9 in the plasmid pQE32 (C. Kerkhoff) were amplified by standard PCR using primers with HindIII and NdeI restriction sites at either end (S100A8: AGTCCTAAGCTTCTACTCTTTGTGGCTTTCTT, ATTACACATATGATGTTGACCGAGCTGGA; S100A9: ATCTAACATATGATGACTTGCAAAATGTCGCAGC, ATCTTCAAGCTTTTAGGGGGTGCCCTCCC). The PCR products were cloned into the expression vector pET28a+ (Invitrogen) and confirmed by sequencing. We purchased recombinant core histones from Upstate, catalase purified from human erythrocytes (Sigma-Aldrich) and all other proteins purified from human neutrophils (Athens Research and Technology). High resolution FESEM analysis of NET fine structure Human neutrophils were seeded on 12-mm cover slips and stimulated with PMA for 4 hours. After fixation (2.5% glutardialdehyde), specimens were contrasted using repeated changes of 0.5% OsO4 and 0.05% tannic acid. Specimens were then dehydrated in a graded ethanol series and subjected to critical point drying. After coating with platinum/carbon, specimens were analyzed in a Leo 1550 field emission scanning electron microscope (FESEM, Zeiss SMT). Peripheral NET areas with individual NET fibers were recorded at high magnification and the obtained images were analyzed using the SmartSEM software (Zeiss SMT). Release of calprotectin 1.7×106 ml−1 human neutrophils were seeded in 1 ml RPMI medium containing 10 U/ml DNase-1 in 12 well tissue culture plates. For each condition to be tested three wells were seeded with neutrophils (n = 3). They were stimulated to form NETs using 20 nM PMA, or to degranulate using 5 µM f-MLP at 37°C in a 5% CO2 atmosphere similar to [40]. Dnase-1 was present to collect all released proteins, also the ones that are bound to NETs. At the indicated time points the supernatants were collected. Hundred µl aliquots were used to quantify LDH activity in the samples using the Cytotoxicity Assay™ (Promega) as a marker for cell death [67],[68]. As 100% control for LDH we lysed the same amount of neutrophils in 1 ml RPMI with 0.1% Triton X-100 and measured 100 µl. The rest of the samples were acetone precipitated. Precipitates were boiled in 30 µl SDS sample buffer and analyzed by immunoblotting using anti-S100A8 (Acris BM4029, 2.5 µg/ml), anti-S100A9 (Acris BM4027, 2.5 µg/ml), anti-LTF (Sigma-Aldrich L3262, 2 µg/ml) and anti-MPO (DAKO A0398, 2 µg/ml) antibodies. To determine the relative amounts of calprotectin in the supernatant after NET formation and in NETs we seeded 1.7×106 neutrophils per well and induced them to make NETs for 4 hours. We collected the supernatant and washed the cells twice. The washes were discarded. Then we digested the NETs with nuclease in the same volume and removed the supernatant again after 20 min. Subsequently, we added 200 µl reducing SDS protein sample buffer to the remaining debris in the wells and scratched them thoroughly and boiled for 3 minutes. As 100% control we lysed the same amount of neutrophils in 400 µl protein sample buffer and boiled as well. The supernatant and the NET digest were lyophilized overnight and also resuspended in 200 µl sample buffer. 40 µl of each fraction was loaded and subjected to SDS PAGE and immunoblotting with an anti-S100A9 antibody. The signal intensities of the bands were analyzed by 2D densitometry as described under “Quantification of NET proteins”. The relative amounts were calculated. Antimicrobial activity of NETs 5×105 ml−1 human or murine neutrophils were stimulated with 20 nM PMA for 4 h at 37°C in a 5% CO2 atmosphere to form NETs in 24 well tissue culture plates. Supernatants were then removed, NETs were washed twice with 1 ml RPMI and microbes were added at a multiplicity of infection (MOI) of 0.01 if not stated differently in 500 µl of RPMI per well. Samples C. albicans were incubated overnight at 30°C to induce yeast-form growth or at 37°C to induce hyphal growth. NETs showed to be similarly antifungal at both temperatures (Figure S4) and purified calprotectin as well (data not shown). Assays with C. neoformans were incubated only at 37°C. After incubation overnight the microbes were plated on YPD agar plates to determine colony forming units (CFU) or an XTT assay was performed as shown in Figure S4. For assays in Figure 4 A & B C. albicans and NETs were incubated at 30°C. Immunodepletion of calprotectin from NETs Twenty times 5×105 human neutrophils were seeded into 24 well tissue culture plates each in 500 µl RPMI and stimulated with 20 nM PMA for 4h at 37°C in a 5% CO2 atmosphere to form NETs. Supernatants were removed, NETs were washed twice with 1 ml RPMI and digested with 500 µl 10 U/ml DNase-1 each. The digested NET proteins were pooled into two 5 ml fractions and concentrated 10-fold on filter columns with a 3.5 kDa cut-off to a volume of 500 µl. The concentrated samples were incubated for 2 h either with a combination of anti-S100A8 and anti-S100A9 (Acris BM4029 and BM4027, 10 µg each), or isotype mouse IgG1 as a control (Sigma M5284), immobilized on 100 µl magnetic sepharose beads (Pierce). After incubation, the samples were diluted back to the original volume using fresh medium to complement metal ions. CFU were determined after incubation of these samples overnight with C. albicans. Viability of C. albicans was also monitored using the XTT assay as stated in supporting material Figure S4. The immunodepletion was evaluated by immunoblotting. Samples were tested for the presence of calprotectin (Acris BM4027, 2.5 µg/ml) and lactotransferrin (Sigma-Aldrich L3262, 2 µg/ml) as a control. Co-precipitation assay of free and NET-bound calprotectin 1.7×106 ml−1 human neutrophils were stimulated with 20 nM PMA for 4h at 37°C in a 5% CO2 atmosphere to form NETs in 1 ml RPMI. NET-free supernatants after incubation were removed and kept. NETs were washed twice with 1 ml RPMI and digested in 1 ml RPMI with 5 U/ml MNase for 20 min to obtain fragments of NETs. NET-free supernatants and digested NET fragments were incubated on a rolling wheel with 3×107 C. albicans yeasts and C. neoformans from YPD overnight cultures previously washed 3 times in PBS. After 30 min the microbes were pelleted, the supernatants removed and washed three times in fresh medium. These new supernatants were acetone-precipitated. Precipitates from supernatants and microbial pellets were boiled in SDS sample buffer and analyzed by immunoblotting using anti-S100A8 and anti-S100A9 antibodies (Acris BM4029 and BM4027, both at 2.5 µg/ml) as described above. Infections and immunohistochemistry For analysis of abscess formation 10 anesthetized animals per group were shaved on the back and subcutaneously infected with 5×107 C. albicans. Abscesses were measured with a caliper at the indicated time points. Pulmonary candidiasis was induced by intranasal challenge of anesthetized animals with 5×107 C. albicans and systemic candidiasis by intravenous injection to the tail vein with 5×105 C. albicans in 10 animals per group. Inoculation doses were determined by CFU counts after plating. For intranasal and intravenous challenge, survival was monitored daily. For fungal load, lungs from 11 mice per group were macerated and CFU determined 3 days after intranasal challenge. For histology of pulmonary infections lungs were removed 24 h after intranasal challenge and 6 days after subcutaneous challenge abscesses were removed. Tissue samples were fixed in 2% formalin, dehydrated, embedded in paraffin, sliced to 5 µm, rehydrated and stained with hematoxylin and eosin (H & E). For immunostainings, samples were rehydrated, subjected to antigen retrieval and incubated with primary antibodies directed against calprotectin subunits S100A8 and S100A9 (produced in house), MPO (DAKO A0398) and histone (Santa Cruz 8030). These were detected with secondary antibodies coupled to Cy2, Cy3 or Cy5. DNA was detected with DRAQ5™ (Biostatus). For fine structural analysis of paraffin-embedded tissue samples, 5 µm sections were rehydrated, postfixed with glutaraldehyde, contrasted using repeated changes of 0.5% OsO4 and 0.05% tannic acid, dehydrated in a graded ethanol series, critical-point dried and coated with 5 nm platinum/carbon. For immunostainings with human neutrophils, 1×105 cells were seeded on 13 mm glass cover slips, stimulated with 20 nM PMA and fixed in 2% formalin at the indicated time points. Specimens were blocked with 3% cold water fish gelatin, 5% donkey serum, 1% BSA, 0.25% TWEEN 20 in PBS, incubated with primary antibodies directed against S100A8/A9 complex (Acris BM4025) and MPO (DAKO A0398) and then washed. Primary antibodies were detected with species-specific secondary antibodies and DNA with DRAQ5™. Specimens were analyzed using a SP5 confocal microscope (Leica). Statistical analysis One-way analysis of variance (ANOVA) with Bonferroni post-tests was applied when multiple groups were compared and two-tailed Student's t-test was used for analysis of two groups. For non-parametrically distributed data, the two-tailed Mann-Whitney test was used. Survivals of infected mice were determined by the log-rank test. Differences were considered statistically significant at P<0.05. All statistical tests were performed using GraphPad Prism version 4.02. Supporting Information Figure S1 Optimization of NET protein purification. Human neutrophils were induced to make NETs. (A) The optimal time point for isolation of NET proteins was determined by monitoring DNA amount and neutrophil elastase (NE) activity at the indicated time points. Four hours after stimulation, both DNA amount and NE activity reached a maximum. This time point was chosen for purification and identification of NET proteins. (B–C) Different nucleases were compared. DNase-1 (B) and MNase (C), a non-processive nuclease, digest NETs to release a maximum of 2.5 µg/ml DNA. Thus we used 10 U/ml Dnase-1 for protein identifications. For a stable DNA concentration in all quantitative analyses we used 5 U/ml MNase and normalized protein concentration to DNA concentration. NETs were digested for different amounts of time. Both Dnase-1 (D) and MNase (E) released a maximum of DNA before 10 min of digest. To ensure complete degradation of NETs 20 min were used for all experiments. The concentration (F) of Dnase-1, as well as the time of digest (G), was confirmed in a silver-stained SDS-PAGE analysis to be optimal at 10 U/ml and 20 min for a maximal protein yield. Shown are means±s.d. (n = 3) of representative experiments from two. (4.40 MB TIF) Click here for additional data file. Figure S2 Mass spectrometry (MS) identification quality and immunoassays to verify the absence or presence of proteins in NETs. (A) Representative identification of NET protein S100A9 by LC-MS/MS. We obtained 51% sequence coverage from 8 MS/MS spectra and a Mascot ion score of 102. We show a representative MS/MS spectrum of one peptide mass (1806.9532). In this case, identification was confirmed by 9 y-ions in series and 11 b-ions, and the immonium ion of His (H). We evaluated the NET association of (B) bactericidal/permeability increasing protein (BPI), (C) pentraxin 3 (PTX-3) and (D) cathelicidin CAP-18 that have been described as NET-associated [3],[4],[37],[38], but were not found in our MS approach. BPI and CAP-18 are endogenously cleaved by neutrophil proteinases into a 25 kDa and a 5 kDa (LL-37) cleavage product [2]. We immunoblotted neutrophil (PMN) lysates, NET extracts and purified proteins as positive controls. Human neutrophil granular extract (hNGE) was used as positive control for the presence of BPI, as well as recombinant human PTX-3 and purified human LL-37 peptide. We confirmed NET association of BPI, but not of PTX-3 and CAP-18. (E) Quantification of calprotectin. We confirmed that calprotectin was a bona fide NET protein by washing the NETs until unbound calprotectin was not detectable using an enzyme-linked immunosorbent assay (ELISA) from Hycult. The data are the average of two independent experiments (means±s.d., n = 4), bdl = below detection limit (1.6 ng/ml). (0.42 MB TIF) Click here for additional data file. Figure S3 NET Protein quantification. Representative quantitative immunoblots for S100A8 (A) and MPO (E) showing six different protein amounts of the standard protein as well as a NET sample in triplicate. The signal intensities (B and F) were plotted against the amount of standard protein within a linear range (C for S100A8 and G for MPO). The resulting equation was used to calculate the respective amounts of proteins as means from three different samples (D and H). The protein amounts (D and H) are specified as mg or µmoles referred to the amount of DNA within the sample. The DNA amount is the mean of all samples used for this quantification. One sample equals the amount of NETs isolated from 1.7×106 human neutrophils. Similar analyses of all the quantified proteins are available on the NET database. (0.98 MB TIF) Click here for additional data file. Figure S4 Antifungal activity and composition of NETs is similar under different conditions (MOI or temperature) and stimuli (PMA or C. albicans). Human neutrophils were induced to make NETs and washed twice. C. albicans and NETs were incubated overnight in all assays either at 30°C to preserve yeast-form growth or at 37°C to induce hyphal growth. (A) Cryptococcus neoformans or C. albicans were added to NETs with a MOI of 0.04, incubated overnight at 37°C and CFU determined, for C.n.+/−NETs P<0.01, for C.a.+/−NETs P<0.05. (B) C. albicans was added to NETs and incubated to induce hyphal growth. NETs inhibit C. albicans hyphae at a MOI of 0.04 and 0.01 similarly. (C) C. albicans was added to NETs and incubated to preserve yeast-form growth. NETs inhibit C. albicans yeast at a MOI of 0.2, 0.04 and 0.01 similarly. (D) Addition of MNase to NETs significantly reduced inhibition of C. albicans yeast, P<0.01. (A–D) Shown are means±s.d. (n = 3) from one representative experiment out of three. (E) Representative tissue culture plate and (F) light microscopic image with growing C. albicans hyphae in the absence (top) or presence of NETs (bottom). (G–I) Antifungal NET assays using XTT [46] we confirmed that NETs also reduce C. albicans growth under hyphae inducing conditions at 37°C only in the absence of 0.5 µM Zn2+ for +/−NETs at 0 µM Zn2+ P<0.001 (G). NET protein extracts inhibited hyphal growth significantly stronger than calprotectin-depleted extracts, for +/− depletion P<0.01 (H). NETs from wild-type mouse neutrophils reduced hyphal growth 20-fold whereas S100A9-knockout neutrophils only reduced growth 3-fold, for WT vs. KO P<0.001 (I). Shown are means±s.d. (n = 3) from one experiment out of two. (J) Human neutrophils were induced to make NETs overnight using either 20 nM PMA or C. albicans (MOI 2) in the presence of protease inhibitors. NET proteins were purified and protein concentrations of samples were determined and equal amounts loaded. Immunoblots probed for lactotransferrin (LTF), cathepsin G (CG), calprotectin and catalase indicated a similar protein composition of NETs independent of the stimulus. Differences in amounts of individual proteins might be caused by an overall higher protein concentration in samples from C. albicans induced NETs due to secreted C. albicans proteins. (4.12 MB TIF) Click here for additional data file. Figure S5 Calprotectin-deficient mice release similar amounts of NETs as wild-type mice in vitro and in vivo. (A) NET-DNA release was induced with PMA and measured by Sytox Green™. There were no significant differences between calprotectin-deficient and wild type mouse neutrophils. (B) H & E stain of an abscess section from calprotectin-deficient mice, 6 days after infection. NETs are indicated by arrows, scale bar = 50µm. (C–F) Confocal images of indirect immunofluorescence with antibodies against (C) S100A9 (red), (D) MPO (green), (E) histone (blue) and (F) superimposition of all signals. Arrows indicate NETs. No stain can be detected for the anti-S100A9 antibody verifying its specificity. Scale bar = 50 µm. (G) Neutrophil recruitment was counted from images of representative abscess sections for calprotectin-deficient and wild type mice and calculated as neutrophils/area (mm2). Shown are means μ s.d. of triplicates from three independent abscesses per group. (4.10 MB TIF) Click here for additional data file. Figure S6 C. albicans infection spread after subcutaneous challenge in calprotectin-deficient but not in wild type mice. In calprotectin-deficient mice, abscess lesions spread to other locations in approximately 30% of the infected animals (n = 10). Spreading did not occur in wild type animals. (A) Shown are representative abscesses from a calprotectin-deficient and (B) from a wild type mouse at day 21 p.i. (5.00 MB TIF) Click here for additional data file. Text S1 Supporting Methods (0.05 MB DOC) Click here for additional data file.

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Excessive Neutrophils and Neutrophil Extracellular Traps Contribute to Acute Lung Injury of Influenza Pneumonitis

Teluguakula Narasaraju, Edwin Yang, Ramar Samy … (2011)

Complications of acute respiratory distress syndrome (ARDS) are common among critically ill patients infected with highly pathogenic influenza viruses. Macrophages and neutrophils constitute the majority of cells recruited into infected lungs, and are associated with immunopathology in influenza pneumonia. We examined pathological manifestations in models of macrophage- or neutrophil-depleted mice challenged with sublethal doses of influenza A virus H1N1 strain PR8. Infected mice depleted of macrophages displayed excessive neutrophilic infiltration, alveolar damage, and increased viral load, later progressing into ARDS-like pathological signs with diffuse alveolar damage, pulmonary edema, hemorrhage, and hypoxemia. In contrast, neutrophil-depleted animals showed mild pathology in lungs. The brochoalveolar lavage fluid of infected macrophage-depleted mice exhibited elevated protein content, T1-α, thrombomodulin, matrix metalloproteinase-9, and myeloperoxidase activities indicating augmented alveolar-capillary damage, compared to neutrophil-depleted animals. We provide evidence for the formation of neutrophil extracellular traps (NETs), entangled with alveoli in areas of tissue injury, suggesting their potential link with lung damage. When co-incubated with infected alveolar epithelial cells in vitro, neutrophils from infected lungs strongly induced NETs generation, and augmented endothelial damage. NETs induction was abrogated by anti-myeloperoxidase antibody and an inhibitor of superoxide dismutase, thus implying that NETs generation is induced by redox enzymes in influenza pneumonia. These findings support the pathogenic effects of excessive neutrophils in acute lung injury of influenza pneumonia by instigating alveolar-capillary damage.

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H5N1 and 1918 Pandemic Influenza Virus Infection Results in Early and Excessive Infiltration of Macrophages and Neutrophils in the Lungs of Mice

Lucy A. Perrone, Julie K. Plowden, Adolfo García-Sastre … (2008)

Introduction The influenza pandemic from 1918 to 1919 was the most devastating infectious disease pandemic ever documented in such a short period of time, killing nearly 50 million people worldwide [1]. Unlike the epidemiological profiles of most influenza infections, young adults aged 18–35 yrs old had the highest mortality rate, so much so that the average life expectancy during those years was lowered by 10 years [2]. In 1918, severe destruction of lung tissue observed by pathologists at autopsy was unlike that typically seen in cases of pneumonia [3] and histopathological analysis of lung tissue showed severe tissue consolidation with unique destruction of the lung architecture [3],[4]. Human infections with highly pathogenic avian influenza (HPAI) strains of subtype H5N1 since the first outbreak in 1997 have also been particularly severe for children and young adults [5]–[7]. Assessing pulmonary infiltrates in response to influenza H5N1 virus infection has been difficult due to the lack of autopsy material. The basis for the high morbidity and mortality associated with the 1918 virus and recent H5N1 viruses remains inconclusive based on viral genetic analysis alone and accounts of patient lung pathology provide only qualitative information about the host factors contributing to disease [4],[8],[9]. Great concern about a pandemic caused by a novel avian H5 subtype virus warrants comparative studies to better understand the cellular pathology caused by a pandemic virus and potentially pandemic viruses. Identification and quantification of the inflammatory cell types associated with highly pathogenic respiratory infections represent prospective targets for modulation of host innate immune responses. Recent studies using animal models to investigate the mechanism(s) of severe influenza virulence have implicated the innate immune system in complicating lung tissue recovery [10]–[13]. Mouse models of highly pathogenic (HP) H5N1 [14]–[18] and 1918 [19],[20] influenza virus infection confirm histological observations of severe lung pathology in human patients, however, the types of immune cells present during the peak of lung pathology have not been fully elucidated. Excessive immune cell infiltration during an acute lung injury may impair tissue restoration directly by interfering with gas exchange, or indirectly through the release of soluble immune mediators. In the present study, we determined key immune cellular components in the murine lung following infection with matched H5N1 and H1N1 virus pairs that represent high and low virulence infections of each influenza subtype as previously determined in the mouse model [18],[21]. The two H5N1 viruses used in this study (A/Thailand/16/2004 and A/Thailand/SP/83/2004) were isolated in 2004 from fatal human cases in Thailand but have a differential pathogenic outcome in mice, specifically a low and high mouse lethal does 50 (LD50 = 1.7 and 5.6 log10 PFU respectively) [18]. For relevant comparison, we also used a contemporary (non lethal) seasonal H1N1 human isolate from 1991 (A/TX/36/91) and the reconstructed 1918 pandemic virus [21]. A detailed flow cytometry evaluation of lung cells demonstrated that macrophages and neutrophils are the prominent cell types associated with and potentially mediating the severe lung pathology following infection with the highly virulent H5N1 and 1918 viruses. Moreover, inoculation of macrophages and dendritic cells with the HP viruses in vitro or ex vivo reveals that some innate immune cells can themselves serve as targets of viral infection. Results Highly pathogenic H1N1 and H5N1 viruses exhibit early and sustained replication in murine lung tissue following intranasal infection Female BALB/c mice were infected intranasally with either highly pathogenic (HP) or low-pathogenic (LP) influenza viruses (Table 1, Methods) based on known LD50's and phenotypes of disease in mouse [21] and ferret [18],[22] models. As shown in Figure 1, the 1918 (H1N1) and A/Thailand/16/2004 (Thai/16, H5N1) viruses replicated to high titers that were at least 5-fold higher than the respective LP viruses, A/Texas/36/91 (TX/91, H1N1) and A/Thailand/SP/83/2004 (SP/83, H5N1) as early as 1 day post-infection (p.i.) and sustained higher levels of replication than the LP viruses throughout the course of infection. 1918 and TX/91 virus infected lungs reached peak titers on day 3 p.i. and remained elevated for the duration of the study. The HP Thai/16 virus reached peak titers later than the 1918 virus at day 5 pi. while the less mouse-virulent SP/83 virus reached it's highest titers on day 7 p.i.. Lung virus titers were significantly higher (*p 10.1 (3.9)* ˆ ± TX/91 6.9 (0.4) 7.8 (0.3) 6.1 (0.6) 5.5 (0.7) 5.7 (0.5) 5.6 (0.6) NA Neutrophils (CD11b+, CD11c−, Ly6G/C+, CD4−, CD8−) SP/83 7.7 (0.5) 7.5 (2.5) 6.3 (1.5) 7.8 (4.5) 8.4 (5.9) 8.7 (0.8) Thai/16 9.7 (1.1) 8.9 (0.8) 9.3 (3.0) 9.1 (1.3) 15.2 (1.6) 18.8 (0.0) PBS 5.3 (0.8) 1918 3.0 (0.3) 2.5 (0.3) 3.0 (0.4)* 2.3 (0.3)* 2.2 (0.1)* 1.9 (0.2)* TX/91 3.5 (1.2) 3.4 (0.5) 4.7 (0.4) 5.7 (1.0) 3.4 (0.6) 3.7 (0.5) NA Dendritic cells (CD11b−, CD11c+, Ly6G/C−, CD4−, CD8−) SP/83 3.7 (0.8) 4.1 (1.0) 6.2 (1.7) 4.4 (0.4) 3.7 (0.6) 3.6 (0.6) Thai/16 3.2 (0.1) 4.0 (0.4) 4.6 (1.3) 3.4 (0.6) 2.5 (0.4) 2.0 (0.0) PBS 3.8 ( 0.2) 1918 13.4 12.1 5.7 (2.5) 8.3 (0.8) 4.9 (2.7) 10.6 (2.7) 7.2 TX/91 11.4 9.2 4.0 6.7 (0.1) 2.5 (0.1) 8.6 (0.6) 2.2 CD4+ T cells (CD4+, CD8−, CD11b−, CD11c−, Ly6G/C−) SP/83 10.0 14.6 4.9 (3.2) 8.0 (1.6) 3.0 (0.7) 9.6 (1.2) 3.1 Thai/16 10.2 14.1 3.7 (1.0) 5.2 (0.8) 0.7 (0.3) 4.8 3.9 PBS 7.2 (2.2) 1918 3.9 5.4 1.8 4.5 2.6 6.8 4.8 TX/91 2.1 1.1 2.9 4.3 2.5 1.0 0.8 CD8+ T cells (CD4−, CD8+, CD11b−, CD11c−, Ly6G/C−) SP/83 2.3 1.4 2.3 2.2 1.8 5.9 1.5 Thai/16 2.2 2.8 2.7 3.0 1.7 1.3 1.6 PBS 2.0 BALB/c mice were infected intra-nasally with 102 PFU influenza viruses (Table 1). Lung immune cell populations were analyzed by flow cytometry from single cell suspensions from three mice per time point, per virus group. ‡ Numbers of immune cell sub-populations are represented here as percent of the total leukocytes measured in these assays. Standard deviations are shown in parenthesis where available. PBS group % populations represented are daily averages. * p<0.05 between HP (1918, Thai/16) and LP (TX/91,SP/83) infection groups. ˆ p<0.05 between 1918 and TX/91 infection groups. ± p<0.05 between Thai/16 and SP/83 infection groups. † Not analyzed. In contrast, dendritic cell populations as percent of the total leukocyte population declined over the course of infection with HP viruses while percentages increased in LP infections, peaking at days 3 and 5 p.i. (Table 2). Significant differences in percent dendritic cells between HP and LP infection groups were observed day 3 p.i. and at all other subsequent time points (* p<0.05). Percentages of CD4+ T cells in the lungs decreased in all infection groups but no significant differences were observed in CD4+ or CD8+ T cell population dynamics between HP and LP infection groups. Together, these results indicate that macrophages and neutrophils are responsible for the majority increase in total lung cell numbers following infection with the 1918 and HP H5N1 influenza viruses. The 1918 pandemic virus and a highly pathogenic H5N1 virus elicit significantly high levels of pro-inflammatory cytokines in the lung To better understand potential influences on immune cell population dynamics in the lung following HP influenza virus infection as demonstrated in Figure 2B and Table 2, we analyzed 17 chemokines and cytokines in the lungs of mice on days 1 and 4 p.i and report data here on 6 of the analytes that revealed significant differences among the viruses tested (Figure 3). These earlier time points were chosen in an effort to understand the temporal relationship with lung immune cell infiltration in 1918 and Thai/16 infections (Figure 2) and day 4 p.i. is a time when significant differences were observed in lung virus titers between HP and LP infection groups (Figure 1). As shown in Figure 3, on day 1 p.i. TX/91 infected lungs exhibited higher titers of IL-1α, IFN-γ, and KC compared to 1918 virus-infected lungs though levels of MIP-1α, MCP-1, and IL-6 were higher in 1918 virus infected lungs. Cytokine levels were similar between the H5N1 viruses at day 1 p.i.. In contrast, lung tissue cytokine and chemokine levels at day 4 p.i. were higher among 1918 and Thai/16 virus-infected mice compared to those infected with the subtype-matched LP (TX/91 and SP/83) counterpart viruses for all cytokines and chemokines measured. Protein levels of the potent monocyte chemoattractant MCP-1 were 10-fold higher in 1918 and H5N1 virus-infected lungs than TX/91 virus-infected lungs, whereas levels of the MIP-1α chemokine were notably elevated in Thai/16 virus-infected lungs. KC (the mouse equivalent of human IL-8) [20],[23] levels were 10-fold higher in Thai/16 and 3-fold higher in 1918 virus infected lungs compared to TX/91 virus-infected lungs with SP/83 levels similar to 1918 infected lungs. The HP viruses were also potent inducers of IL-1α and IFNγ. IL-6, (a generally pro-inflammatory cytokine) was also elevated in 1918 and H5N1 virus-infected lungs compared to TX/91 levels. As shown in Figure 3, significant differences (ˆ p<0.05) were measured between 1918 and TX/91 virus infected lungs on day 4 p.i. for each cytokine and chemokine, accentuating important immunological differences in the immune response to the H1N1 pandemic virus versus a contemporary H1N1 virus. 10.1371/journal.ppat.1000115.g003 Figure 3 Lung cytokine response. Cytokine levels from infected lungs (n = 3 mice per virus group, days 1 and 4 post-inoculation (p.i.)) were measured individually and in duplicate by the Bioplex Protein Array system lungs. Baseline cytokine levels from PBS inoculated mice (mock) are shown as a dashed line in each cytokine graph. Bars represent means of 3 mice from each infection group±standard deviation (SD). Protein levels of IL-6 in Thai/16 infected animals exceeded the y scale shown (indicated by a dash//, the concentration in Thai/16 infected mice was 11.7 ng/ml). ˆ p<0.05 between 1918 and TX/91 viruses, * p<0.05 between 1918 and Thai/16 (HP) virus groups and SP/83 and TX/91 (LP). Innate immune cells are targets of highly pathogenic influenza virus infection Due to the increased presence of macrophages in the lungs following HP influenza virus infection, replication of paired H1N1 and H5N1 viruses (Table 1) was assayed over time in primary human PBMC-derived macrophages and mouse lung macrophages to address whether these cells are specific targets of viral infection and can productively replicate HP viruses. Mouse lung macrophages were harvested from naive BALB/c mice and infected in vitro (MOI = 0.1, Figure 4A) as described. While the HP 1918 and Thai/16 viruses exhibited a slight increase in log titer very early after inoculation (13–24 hrs p.i.), overall these viruses did not replicate efficiently compared to the growth kinetics of these viruses observed in lung epithelial cells [22],[24]. At 13 hrs p.i. there was 18 fold higher 1918 and Thai/16 virus released from infected cells than TX/91 and SP/83 virus-inoculated cultures and nearly 54 times more 1918 and Thai/16 virus than TX/91 or SP/83 virus at 24 hrs p.i. While virus titers in the 1918, TX/91 and SP/83 infection groups declined after the 48 hr time point, the Thai/16 infected macrophages maintained virus titers and these differences were statistically significant (ˆ p<0.05) at 72 hrs p.i.. The lack of prolific replication of these four viruses in primary mouse lung macrophages was confirmed further when a higher MOI (1.0) was utilized (data not shown). Human macrophages also supported replication of all four viruses (Figure 4B, MOI = 0.1). At 48 hrs p.i. 1918 and Thai/16 infected macrophages exhibited 180 times higher virus titers than TX/91 and SP/83 infected cultures. Interestingly, 1918 virus-infected macrophages exhibited a higher baseline titer soon following infection that was found to be statistically significant when compared to the other infection groups († p<0.05). In summary, while human macrophages are a target of viral replication and support replication well, mouse lung macrophages support low levels of 1918 and Thai/16 virus production early following infection. 10.1371/journal.ppat.1000115.g004 Figure 4 Growth of viruses in primary macrophages. Primary macrophages were harvested from the lungs of healthy BALB/c mice through tissue digestion (A) and developed from human peripheral blood monocytes (B) and infected in vitro (MOI = 0.1) with influenza viruses as described (Table 1, Methods). Mouse lung macrophages were grown in 12 well plates and infected with viruses in duplicate and the supernatants sampled for virus growth. Human macrophages were grown in 6 well plates and also infected in duplicate. Virus titers were determined from supernatants in duplicate by plaque assay on MDCK cells. Graphs are representative of results obtained from three independent infection experiments. ˆ p<0.05 between Thai/16 and 1918, SP/83 and TX/91 viruses 72 hrs p.i (A). * p<0.05 between HP Thai/16 and 1918, and LP SP/83 and TX/91 viruses 48 hrs p.i. (B). † p<0.05 between the 1918 virus and other viruses 2 hours post- infection (B). Additional experimentation with primary human macrophages revealed that levels of pro-inflammatory cytokines were higher for H5N1-infected cells than either 1918 or TX/91 virus infected cells (48 hrs p.i., MOI = 0.1, Figure 5). Significant differences in cytokine levels were observed between H5N1 and H1N1 virus infected macrophages in all cytokines measured (*p<0.05) except IL-8 where all four viruses elicited similar levels of this chemokine. Interestingly, the 1918 virus elicited similar cytokine responses as to TX/91 inoculated cells in every cytokine measured even though 1918 virus titers at this time point post-infection were 2.5 logs greater than TX/91 infected macrophages (Figure 4B). Thai/16 infected cultures elicited at least a 2 fold greater cytokine response than the SP/83 virus infected cultures in every cytokine measured except IL-8 and MCP-1 chemokines where Thai/16 levels were only slightly higher than SP/83 levels. 10.1371/journal.ppat.1000115.g005 Figure 5 Cytokine response from infected human macrophages. Primary macrophages were developed from human peripheral blood monocytes in 6 well plates and infected in vitro (MOI = 0.1) with influenza viruses in duplicate as described (Methods, Table 1). Supernatants were sampled 48 hours post-infection and cytokines measured by Bioplex Protein Array assay (Methods). Macrophages were also treated with bacterial lipopolysaccharide (LPS, 100 ng) and Poly I/C (100 ng) to serve as positive inducers. * p<0.05 between H5N1 and H1N1 viruses. Because of their important role in antigen presentation to T cells, we also assessed the ability of primary dendritic cells to productively replicate pandemic H1N1 and human H5N1 viruses (Figure 6A and B). While the HP Thai/16 H5N1 virus replicated slowly over time in cultured mouse lung dendritic cells (Figure 6A), these cells failed to productively replicate 1918, TX/91 and SP/83 viruses. Significant differences in virus titers were measured between HP (1918 and Tha/16) infected cells and TX/91 and SP/83 infected cells at 24 hrs p.i. (Figure 6A, ˆ p<0.05). Significant differences in virus titers was also observed between Thai/16 inoculated cultures and the other infection groups at 48 and 72 hrs p.i. (* p<0.05). Similar to infected human macrophages, these trends were conserved when primary human dendritic cells were the target of infection with these viruses (Figure 6B). Significant differences in virus titers was observed between Thai/16 inoculated cultures and the 1918, TX/91, and SP/83 infection groups at 24, 48 and 72 hrs p.i. (* p<0.05). 10.1371/journal.ppat.1000115.g006 Figure 6 Growth of viruses in primary dendritic cells. Primary dendritic cells were isolated from the lungs of healthy BALB/c mice and developed from human monocytes as described in Methods and infected in vitro with LP and HP influenza viruses (Table 1). Growth of H5N1 and H1N1 viruses in mouse lung macrophages (A, MOI = 0.1) and in human monocyte-derived dendritic cells (B, MOI = 1.0). (ˆ p<0.05 between HP Thai/16 and 1918 viruses and LP SP/83 and TX/91 viruses. * p<0.05 between Thai/16 and 1918, SP/83 and TX/91 viruses.) Graphs are representative of results obtained from three independent experiments. To determine if innate immune cells are being productively infected in vivo, macrophages and dendritic cells were purified from lungs of infected mice and cultured for infectious virus. Lungs from infected (3 days p.i.) mice were harvested and ex vivo cultures containing either lung macrophages or dendritic cells were sampled for infectious virus over a 65 hour time period. While the seasonal influenza isolate, TX/91 virus was not produced from either macrophages (CD11b+) or dendritic (CD11c+) cells, the 1918 pandemic virus as well as the two human H5N1 isolates were released into the culture supernatant (Figure 7), indicating that these cells are being productively infected in the mouse lung. In CD11b+ (macrophages) cell cultures (Figure 7A), the 1918 and Thai/16 virus infected cells released more infectious virus over time than the LP SP/83 virus infected culture. In CD11c+ (dendritic) cell cultures, the Thai/16 virus infected cultures released more infectious virus than the 1918 and SP/83 virus infected cultures (Figure 7B). These data further demonstrate that mouse lung macrophages and dendritic cells are susceptible to highly pathogenic influenza virus infection in the lung tissue. 10.1371/journal.ppat.1000115.g007 Figure 7 Virus replication from primary lung macrophages and dendritic cells cultured ex vivo. BALB/c mice were infected intranasally with 102 PFU of the indicated viruses. Three days post-inoculation, lungs were removed from 2–3 mice per virus group. Cell suspensions were prepared from pooled samples and macrophages (A) and dendritic cells (B) were isolated by CD11b+ ad CD11c+ MACS column purification, respectively. Supernatants from cultures were sampled over time to measure virus production. Virus was titered in duplicate by plaque assay. Discussion Lung consolidation has been described as a pathological feature of severe influenza virus infection caused by the 1918 pandemic virus and H5N1 viruses in humans [9],[25],[26] as well as in animal models [11],[12],[18]. Using a detailed flow cytometry evaluation, the current study set out to characterize differences in the cellular innate immune response in the mouse lung following highly pathogenic (HP) or low pathogenicity (LP) influenza virus infections. Lungs from mice infected with the HP 1918 H1N1 virus and a recent H5N1 human isolate (A/Thailand/16/04 (Thai/16)) exhibited a significant increase in cellularity in comparison to the LP seasonal H1N1 isolate, A/Texas/36/91 (TX/91). Significant differences in titer between HP and LP virus-infected mice were observed as early as day 1 post-inoculation (p.i.), likely forecasting the dramatic increase in immune cell infiltration in the lungs of these virus-infected mice. Interestingly at day 7 p.i. when peak lung cellularity was observed in HP virus infection groups, differences in virus titers between paired subtype viruses (1918 compared to TX/91 and Thai/16 compared to SP/83) were minimal and were limited to a maximum difference of 1 log (Figure 1), indicating a failure by the immune system to clear the viral infection. Lung cellularity was further investigated by characterizing the comprising immune cell sub-populations. We observed a significant increase in macrophages and neutrophils early following infection with the 1918 and Thai/16 viruses, and their sustained presence in the lung tissue mark a distinction between HP and LP influenza virus infection. These data show that virus replication in the lungs of HP influenza infections are sustained at high levels the first week of infection regardless of the high numbers of immune cells present in the tissue. Although the current study did not determine whether these cells are playing an antiviral role against the HP viruses, it has been previously shown that neutrophils and macrophages assist in the clearance of influenza virus early during infection; these cells appear to be capable of only partially reducing the virus load in the lung despite their presence at high numbers [20]. We also observed a decrease in the percentage of lung–associated dendritic cells and T cell lymphocytes during HP influenza virus infection. A decrease in the number of circulating lymphocyte populations has also been previously observed in the peripheral blood of humans and mice infected with H5N1 viruses [5],[9],[10],[15],[20],[27]. The precise mechanism of leukocyte depletion during H5N1 infections is not well understood, but evidence of apoptosis in the spleen and lungs of HP H5N1-infected mice detected in situ suggests a mechanism for cell loss [16]. Influenza virus growth in the respiratory epithelium and the subsequent release of chemotactic proteins from those cells may encourage the increased presence of macrophages and neutrophils [28],[29]. Macrophages and neutrophils can secrete chemokines and cytokines that can act in an autocrine fashion which in turn can promote the increased migration of those cells and other leukocytes into the lung tissue [30]. Elevated levels of certain chemokines and cytokines have been associated with high viral load and severe disease in H5N1 virus infected patients [26],[31]. These studies show that infection with HP 1918 and Thai/16 H5N1 viruses result in elevated amounts of pro-inflammatory chemokines and cytokines in the lungs of mice day 4 post-infection compared with TX/91 and SP/83 infected mice, a time point that correlates with rising but significantly different lung virus titers between infection groups. Elevated levels of the chemokines MCP-1 [32] and MIP1-α were observed among H5N1 and 1918 virus infected mouse lungs. Although, MIP-1α does not appear to be critical for virus replication and spread in the mouse model [10], this chemokine exhibits a variety of pro-inflammatory activities including macrophage and neutrophil recruitment and has been associated with fatal outcomes in human H5N1 virus infections [31]. Lungs infected with the 1918 pandemic and Thai/16 H5N1 viruses also exhibited significantly higher levels of IFN-γ on day 4 p.i compared to their subtype-paired LP virus counterparts. IFN-γ is known to mediate the increased production of nitric oxide [33] which can subsequently result in the recruitment of more neutrophils and macrophages. Higher levels of IL-6 were measured in 1918 and Thai/16 virus infected lungs, supporting observations obtained with the 1997 H5N1 viruses [10] and has been correlated with systemic illness symptoms and fever in experimental human TX/91 infections [34]. By directly measuring cytokine protein levels, these data provide confirming evidence of a heightened lung cytokine response to 1918 and H5N1 infection in mice [13]. Interestingly, while we reveal marked similarities between the HP 1918 and Thai/16 viruses in overall lung cellularity, virus growth and patterns of immune cell sub-population dynamics over time, Thai/16 virus infection consistently resulted in higher levels of chemokines and cytokines both in mouse lungs and human macrophages. Although it has been shown recently by two independent research groups that the lack of key cytokines (through the use of single cytokine gene knockout mice) had no effect on the overall disease outcome or virus replication among H5N1 virus-inoculated mice [10],[35], the present results along with data from others continue to indicate that pro-inflammatory cytokines correlate with disease outcome [36]. It is thought that these immune mediators do not act singularly in vivo and it will be critical to reveal these concerted interactions (both locally in the lung and systemically) to further our understanding of the pathogenesis of H5N1 infection in animal models and in human patients. Macrophages and dendritic cells play a fundamental role in the lung at all stages of influenza virus infection [20],[37],[38]. We have provided evidence regarding the higher replication efficiency of HP influenza viruses in primary human macrophages and dendritic cells, a property that has also been demonstrated previously in other primary cells [24],[39]. Mouse macrophages were also susceptible to virus infection in vitro, however they did not support productive replication to the level observed in primary human monocyte-derived macrophages (Figure 4B) or lung epithelial cells [21],[24]. However, the cytopathic effects (estimated by visual examination of monolayers) among HP virus-infected mouse and human macrophages was observed in a shorter period of time compared to LP virus infected cells. Interestingly, the 1918 virus exhibited higher baseline titers in human and mouse macrophages as early as 2 hrs p.i. compared to the TX/91 H1N1 virus or the H5N1 viruses, indicating a curious property of this pandemic virus. While the importance of the binding properties of the HA molecule has been demonstrated elsewhere extensively [22],[40],[41], further resolution of this interesting finding and its immunological importance deserves further experimentation. The role of other cell surface molecules on innate immune cells such as the mannose receptor in HP influenza infection should be investigated [42],[43]. The higher viral replication of HP viruses in dendritic cells also correlated with the severe pulmonary disease observed in mice. We also demonstrated definitively that these cells are targets of infection ex vivo by highly pathogenic influenza viruses like the 1918 pandemic virus and recent H5N1 isolates (Figure 7). Thus, it appears that macrophages and dendritic cells may contribute to the pathogenesis of HP virus infection due to their susceptibility to influenza virus infection. An inability to mount an adaptive immune response due to direct infection of important innate immune cells such as macrophages and dendritic cells may be a critical difference in host outcome during influenza virus infection [44]. This coupled with the phenomenon of T cell depletion in infected mice [17] and human patients [9],[31] may allow for uncontrolled viral replication. While reducing viral load through anti-viral intervention remains the best treatment option for H5N1 patients, therapies that moderate immunopathology may help to reduce the high case fatality rate currently associated with virus infection [45]. Materials and Methods Viruses and cells All in vitro and in vivo experiments were performed under the guidance of the U.S. National Select Agent Program in negative pressure HEPA-filtered biosafety level laboratory (BSL-3+) enhanced laboratories and with the use of a battery powered Racal HEPA-filter respirator and according to Biomedical Microbiological and Biomedical Laboratory procedures. Influenza viruses used in these experiments included the reconstructed A/South Carolina/1/18 virus (H1N1) [21], A/Texas/36/91 (H1N1), A/Thailand/16/2004 (H5N1), and A/Thailand/SP/83/2004 (H5N1). These viruses were chosen as pairs within subtypes due to their respective fifty percent lethal dose (LD50) titers in mice and ferrets (Table 1). Low pathogenicity in this manuscript refers to the non-lethal phenotype of the seasonal H1N1 TX/91 virus and the low virulence SP/83 H5N1 isolate [18],[21]. The Thai/16 and SP/83 viruses differ from each other in 13 amino acids in 7 proteins and this sequence comparison has been published previously [18]. The A/Texas/36/91 and H5N1 viruses were grown in 10 day old embryonated hen's eggs and the 1918 viruses grown in MDCK cells. All virus stocks were titered by plaque assay on MDCK cells prior to mouse infections. Human peripheral blood monocytes (PBMC's) were obtained by Histopaque (Sigma-Aldrich, St. Louis, MO) density gradient centrifugation of whole blood donated by healthy donors aged 20–40 yrs old without history of influenza vaccination in the past year. Whole blood was obtained through an approved protocol by both the Emory University Institutional Review Board (IRB) and CDC IRB (Emory University Hospital Blood Bank is an FDA-accredited Blood Bank). Human monocytes were obtained by negative selection column enrichment (Miltenyi Biotech, Auburn, CA) yielding approximately 90% CD14+ purity as determined by FACS analysis. For development of macrophages, monocytes were cultured at 37°C in 6 well plates in Macrophage SFM media (Gibco, Grand Island, NY) with 20% heat inactivated autologous serum for 7 days in the presence of GM-CSF (1000 U) before infection [46]. Human macrophages cultured in this manner typically displayed classical morphology with the phenotype: CD11blow, CD11clow, HLA-DRlow, CD14low, CD40low, CD80low, CD83low, CD86- (Figure 4C and D). For the development of dendritic cells (DC's), monocytes were grown in RPMI (Gibco) with 20% heat inactivated autologous serum for 10 days in the presence of IL-4 (1000U) as previously described [46]. Dendritic cells developed in this manner typically displayed a classical morphology with the presence of dendritic processes with the phenotype: CD11blow, CD11chigh, HLA-DRhigh, CD14low, CD40high, CD80high, CD83high, CD86high (Figure 6B). To obtain primary mouse lung macrophages and dendritic cells, lungs from naïve mice were removed and tissue disrupted as described above through the use of collagenase digestion and cell suspensions prepared. Macrophages (CD11b+) and dendritic cells (CD11c+) were extracted from contaminating cells by selection on magnetic columns (Miltenyi Biotech, Auburn, CA). Cells were washed twice with media containing 20% FCS (Macrophage SFM for macrophages (Gibco) or RPMI (Gibco) for dendritic cells) and cultured for 24 hours before in vitro infection. Primary mouse lung macrophages typically displayed the phenotype: CD11bhigh, CD11c-, MHCIIhigh, CD40high, CD80high, CD83high (Figure 4A and B). Primary mouse lung dendritic cells typically displayed typical morphology with dendritic extensions and the phenotype: CD11b-, CD11chigh, MHCIIhigh, CD40high, CD80high, CD83high (Figure 6A). In vitro infections Primary human and mouse cells were washed 3× with serum free growth media and infected for 1 hour with viruses (Table 1) at a multiplicity of infection (MOI) of 0.1. Following infection, cells were washed 3× with serum free growth media and 1 ml SFM media, containing 1 µg/ml of TPCK-treated trypsin (Sigma-Aldrich), was placed into the wells. Virus growth was measured over time in triplicate wells for each experiment and titered in duplicate by standard plaque assay on MDCK cells. All macrophage and dendritic cell data reflects at least three independent experiments (Figures 4 and 6). Cytokine levels produced from infected human macrophages (MOI = 0.1) were quantitated 48 hrs p.i. by BioPlex assay (Figure 5). Escherichia coli lipopolysaccharide (LPS, 100 ng, Sigma-Aldrich) and Poly I/C (100 ng, Sigma-Aldrich) were used as positive control stimulants. Mice infections All animal research was conducted under the guidance of CDC's Institutional Animal Care and Use Committee and in an Association for Assessment and Accreditation of Laboratory Animal Care International- accredited facility. 8–10 week old female BALB/c mice (Harlan, Indianapolis, IN) were anesthetized with Avertin [20] (Sigma-Aldrich) and infected intranasally (i.n.) with 50 µl of 102 PFU of influenza viruses prepared in phosphate buffered saline (PBS). Avertin was chosen as the anesthetic because it provides consistent mouse infections with the viruses used in these studies. Using the sublethal (102 PFU) inoculum, 1918 and Thai/16 virus infected mice survive a prolonged disease course allowing for the measurement of the influx of inflammatory cells into the lung tissue during a full course (∼7–9 days) of influenza virus infection. At indicated times post-infection (n = 3 mice per virus group) mice were euthanatized and exsanguinated. Lungs were removed from individual mice without PBS perfusion and included total lung cell counts included cells located in the bronchoalveolar airways. Perfusion was not possible in many cases of HP influenza virus infection due to the presence of microvascular hemorrhage. We obtained similar results when performing these lung cell quantitation assays with or without lung perfusion in LP virus infected mice and therefore did not introduce this variable in our high containment laboratory. Whole lung cell suspensions were prepared in Dulbecco's minimal essential media (DMEM) with 20% fetal calf serum following collagenase-DNase treatment and manual disruption [47]. Red blood cells were removed by lysis buffer treatment (Sigma-Aldrich). Total viable lung cell number was determined for each mouse by trypan blue exclusion on a hemocytometer. For the ex vivo experiment, macrophages and dendritic cells were isolated from the lungs of infected BALB/c mice. Two or three mice were infected i.n. with 102 PFU of each of the four viruses described in this study (Table 1). Three days post-inoculation, lungs were removed without perfusion and cell suspensions were prepared as described above. Lungs were pooled from mice in each virus infection group. Macrophages and dendritic cells were isolated by positive selection on CD11b+ or CD11c+ MACS columns. Columns containing bound immune cells were washed extensively (5x) and CD11b+ or CD11c+ cells were eluted off the magnetic columns and cultured in 6-well plates in 5 ml of RPMI containing 5% BSA. Supernatants were collected at the indicated times and virus content was determined in a standard plaque assay on MDCK cells. Flow cytometry and lung immune cell quantitation Lung cell suspensions were incubated with anti-Fc block (anti-mouse CD16/CD32) to reduce non-specific antibody binding for 10 min. prior to staining for 1 hr with fluorophore-conjugated antibodies (BD Biosciences, San Diego, CA) specific for immune cell populations according to standard protocols [48] and included: CD11b-PE (pan-macrophage), CD11c-APC (pan-dendritic cell), Ly6G/C-FITC (neutrophil), CD4-PE and CD8-APC T cell markers (Table 2, Figure 2). Cells were washed twice with PBS and fixed overnight at 4°C with 2% paraformaldehyde. Samples were safety tested for infectious virus and removed from the BSL3+ laboratory. Flow cytometry was performed on a FACSAria flow cytometer (BD Biosciences). To further characterize primary mouse and human macrophage and dendritic cells we utilized the following fluorescently conjugated antibodies for flow cytometric analysis: MHC II (I-A/I-E)-PE, HLA-DR-PE, CD14-FITC, CD40-FITC, CD80-FITC, CD83-APC, and CD86-APC (BD Biosciences) (Figures 4 and 6). Virus titrations and cytokine analysis At various times post-infection (n = 3 mice per virus group) lungs were removed and stored at −70°C until virus and cytokine levels could be quantified. Lungs were homogenized individually in 1 ml PBS. Virus was titered from clarified lung homogenates by standard plaque assay on MDCK cells in duplicate and titers are reported as plaque forming units per ml PBS (PFU/ml, Figure 1). Cytokine protein levels were measured (day 4 post-infection ( p.i.)) by the Bioplex Protein Array system [49] (Bio-Rad, Hercules, CA) using beads specific for mouse G-CSF, IL-1α , IL-1β, IL-3, IL-6, IL-9, IL-12 (p40), IL-12 (p70), IL-13, Eotaxin, TNFα, RANTES, KC, MIP1-α, MIP-1β, MCP-1, and IFN-γ. Cytokine protein levels were measured according to the manufacturers instructions by fluorescently conjugated monoclonal antibodies in duplicate against a standard curve (Figures 3 and 5). Statistical analysis Statistical significance of differences between experimental groups was determined through the use of the unpaired, non-parametric Student's t test. Values of p<0.05 were considered significant.

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Journal ID (nlm-ta): eLife

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Journal ID (hwp): eLife

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Title: eLife

Publisher: eLife Sciences Publications, Ltd

ISSN (Electronic): 2050-084X

Publication date (Electronic, pub): 14 February 2017

Publication date Collection: 2017

Volume: 6

Electronic Location Identifier: e20444

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[1 ]The Walter and Eliza Hall Institute of Medical Research , Parkville, Australia

[2 ]deptDepartment of Medical Biology , University of Melbourne , Parkville, Australia

[3 ]deptCentre for Innate Immunity and Infectious Diseases , Hudson Institute of Medical Research , Clayton, Australia

[4 ]Monash University , Clayton, Australia

[5 ]deptPriority Research Centre for Asthma and Respiratory Diseases , Hunter Medical Research Institute and The University of Newcastle , Newcastle, Australia

[6 ]deptShanghai Public Health Clinical Center and Institutes of Biomedical Sciences , Fudan University , Shanghai, China

[7 ]deptDepartment of Microbiology and Immunology , University of Melbourne, at the Peter Doherty Institute for Infection and Immunity , Parkville, Australia

[8]RIKEN , Japan

[9]RIKEN , Japan

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philip.hansbro@ 123456newcastle.edu.au (PMH);

snicholson@ 123456wehi.edu.au (SEN)

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These authors contributed equally to this work.

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These authors also contributed equally to this work.

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Michelle D Tate http://orcid.org/0000-0002-0587-5514

Alan C Hsu http://orcid.org/0000-0002-6640-0846

Sandra E Nicholson http://orcid.org/0000-0002-1314-2134

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Publisher ID: 20444

DOI: 10.7554/eLife.20444

PMC ID: 5354519

PubMed ID: 28195529

SO-VID: 598ae9f8-8fe7-44de-99dc-0be29e4e417a

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Date received : 08 August 2016

Date accepted : 06 February 2017

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Funded by: Victorian State Government, Australia;

Award ID: Operational Infrastructure Scheme grant

Award Recipient : Lukasz Kedzierski Tatiana B Kolesnik Edmond M Linossi Laura Dagley Sarah Freeman Simon M Chatfield Jeffrey J Babon Nicholas Huntington Gabrielle Belz Andrew Webb Nicos A Nicola Sandra E Nicholson Giuseppe Infusini

Funded by: FundRef http://dx.doi.org/10.13039/501100000925, National Health and Medical Research Council;

Award ID: IRIISS grant 361646

Award Recipient : Lukasz Kedzierski Tatiana B Kolesnik Edmond M Linossi Laura Dagley Sarah Freeman Giuseppe Infusini Simon M Chatfield Nicholas Huntington Gabrielle Belz Andrew Webb Nicos A Nicola Sandra E Nicholson Jeffrey J Babon

Funded by: FundRef http://dx.doi.org/10.13039/501100000925, National Health and Medical Research Council;

Award ID: Fellowship

Award Recipient : Michelle D Tate Malcolm R Starkey Nicos A Nicola Katherine Kedzierska Nicholas Huntington Philip M Hansbro

Funded by: Australian Federal Government;

Award ID: Australian Postgraduate Award

Award Recipient : Edmond M Linossi

Funded by: FundRef http://dx.doi.org/10.13039/501100000925, National Health and Medical Research Council;

Award ID: Project grant #1045762

Award Recipient : Peter AB Wark Philip M Hansbro

Funded by: FundRef http://dx.doi.org/10.13039/501100000925, National Health and Medical Research Council;

Award ID: Program grant #1016647

Award Recipient : Nicos A Nicola Sandra E Nicholson

Funded by: FundRef http://dx.doi.org/10.13039/501100000923, Australian Research Council;

Award ID: Future Fellowship

Award Recipient : Gabrielle Belz

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

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Author impact statement The intracellular SOCS5 protein has a unique and key role in restraining influenza A infection by regulating epidermal growth factor receptor signaling in airway epithelial cells.

ScienceOpen disciplines: Life sciences

Keywords: socs5,influenza,egfr,copd,pi3k,innate immunity,human,mouse,virus

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ScienceOpen disciplines: Life sciences

Keywords: socs5, influenza, egfr, copd, pi3k, innate immunity, human, mouse, virus

Suppressor of cytokine signaling (SOCS)5 ameliorates influenza infection via inhibition of EGFR signaling

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H5N1 and 1918 Pandemic Influenza Virus Infection Results in Early and Excessive Infiltration of Macrophages and Neutrophils in the Lungs of Mice

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