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There is accumulating evidence that following bacterial infection, the massive recruitment and activation of the phagocytes, neutrophils, is accompanied with the extracellular release of active neutrophil elastase (NE), a potent serine protease. Using NE-deficient mice in a clinically relevant model of Pseudomonas aeruginosa-induced pneumonia, we provide compelling in vivo evidence that the absence of NE was associated with decreased protein and transcript levels of the proinflammatory cytokines TNF-α, MIP-2, and IL-6 in the lungs, coinciding with increased mortality of mutant mice to infection. The implication of NE in the induction of cytokine expression involved at least in part Toll-like receptor 4 (TLR-4). These findings were further confirmed following exposure of cultured macrophages to purified NE. Together, our data suggest strongly for the first time that NE not only plays a direct antibacterial role as it has been previously reported, but released active enzyme can also modulate cytokine expression, which contributes to host protection against P. aeruginosa. In light of our findings, the long held view that considers NE as a prime suspect in P. aeruginosa-associated diseases will need to be carefully reassessed. Also, therapeutic strategies aiming at NE inhibition should take into account the physiologic roles of the enzyme.
Inflammation is a physiological reaction of the host to protect itself from insulting agents such as pathogens. This complex and dynamic process is characterized by an innate immune response, which involves a coordinated expression of inflammatory cytokines and implication of various cell types particularly immune cells aimed at clearing the pathogenic agent.
In the setting of respiratory bacterial infections (e.g. bacterial pneumonia), the host innate immune response is characterized by the initial recognition of invading microbes by host “sentinel” cells via Toll-like receptors (TLRs)3 or other pattern recognition molecules (1). Subsequently, this results in the production of an array of inflammatory mediators including early responsive cytokines. Another hallmark of innate host lung defense, especially when the first lines of defense: epithelial barrier and resident macrophages, are breached is the massive recruitment of polymorphonuclear neutrophils (PMN) to the infected site (2). PMNs are efficient phagocytes whose main function upon activation is thought to be the clearance of infecting bacteria. To do so, these cells are equipped with a myriad of antimicrobial molecules grouped into oxidative and nonoxidative systems (3, 4).
Regarding the nonoxidative system, we and others have identified the PMN-specific serine protease, neutrophil elastase (NE), as a key antimicrobial enzyme (5). NE catalytic activity relies on the His57-Asp102-Ser195 triad (chymotrypsin numbering system), where Ser represents the active residue (6). NE, a cationic glycoprotein, is stored in readily active form in PMN primary granules at concentrations exceeding millimolar range making it a major component of PMN (7). Gene targeting of NE in mice revealed that the enzyme contributes significantly to host protection against microbial infections (8). In determining NE-mediated defense against Gram-negative bacteria, it was shown that active enzyme degrades the major outer membrane protein (Omp), structural proteins localized on the cell wall (9, 10). NE also has the capacity to attenuate the pathogenicity of invading microbes by targeting their virulence factors (11). In recent years, it was shown that NE binds to PMN-derived chromatin structures, termed neutrophil extracellular traps, and exerts its antimicrobial function (12).
Although NE has been always regarded as pathogenic in Pseudomonas aeruginosa-associated tissue inflammatory and destructive diseases, we have recently provided compelling in vivo evidence that the enzyme contributes considerably to PMN-mediated host protection in a mouse model of P. aeruginosa-induced pneumonia (10). There is also accumulating evidence that during P. aeruginosa infections, active NE is released in the extracellular milieu by recruited PMNs (13, 14). A number of in vitro studies have suggested that NE has the potential to change biologic activities of various inflammatory mediators (15). Altogether, these observations prompted us to hypothesize that the NE role in host defense against P. aeruginosa lung infection may not be only limited to just killing bacteria. Here, we report that in vivo, extracellular active NE has the capacity to induce mRNA expression of the early responsive proinflammatory cytokines, tumor necrosis factor-α (TNF-α), macrophage inflammatory protein-2 (MIP-2), and interleukin-6 (IL-6); an induction that is mediated at least in part through TLR-4. These findings were further confirmed following exposure of cultured macrophages to purified NE. Our studies reveal for the first time that extracellularly released NE can have physiologic inflammatory properties that contribute to host defense against P. aeruginosa.
Purified NE and elastin were obtained from Elastin Products Co. (Owensville, MO). NE activity was determined spectrophotometrically using the specific chromogenic substrate N-methoxysuccinyl-Ala-Ala-Pro-Val-pNA (Elastin Products Co.) according to the manufacturer's recommendations. Pefabloc SC®, 4-(2-aminoethyl)-benzenesulfonyl-fluoride (AEBSF) and secretory leukocyte proteinase inhibitor (SLPI) were from Roche Applied Science and R&D Systems, respectively. RPMI 1640 medium, DMEM, fetal bovine serum (FBS), penicillin, streptomycin, and PBS were obtained from Invitrogen. Ketamine hydrochloride and medotomidine hydrochloride were obtained from CEVA Santé Animale (Libourne, France). Primers for semiquantitative and real time RT-PCR were purchased from Operon Biotechnologies (Cologne, Germany). SYBR Green for real time RT-PCR was obtained from Invitrogen. Polyclonal anti-murine NE antibody was produced in rabbit (16). Antibody raised against amino acids 198–395 of mouse TLR-4 (sc-293072) were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). All other chemicals were reagent grade and purchased from Sigma, unless otherwise stated. Of note, glassware used in the study were LPS-free and when appropriate, reagents especially purified for NE were tested for LPS contamination by the Limulus amebocyte-lysate assay (E-toxate, Sigma).
NE-deficient (NE−/−) mice were generated by targeted mutagenesis as previously described (5). C57BL/6 NE−/− mice and their wild-type (WT) littermates and C3H/HeJ (TLR4−/−) and control C3H/HeN mice (Harlan Laboratories, Indianapolis, IN) were used in this study. C3H/HeJ mice have a naturally nonresponsive TLR4. Mice were sex and age (8–10 weeks) matched and maintained in the animal barrier facility with a 12-h light/dark cycle and provided with water and food ad libitum. Animal handling and procedures were approved by the Animal Studies Committee at our institution (Health and Animal Protection Office, Châlons-en-Champagne, France, authorization number 51-31) in accordance with the guidelines of the Federation of European Laboratory Animal Science Associations (FELASA) and following the European Directive 2010/63/EU on the protection of animals used in scientific procedures.
P. aeruginosa H103 was kindly provided by Dr. Hancock (Vancouver, BC, Canada). This strain is noteworthy in its inability to produce Pseudomonas metalloelastase (17). An overnight bacterial culture (1 ml) was grown in Luria Bertani broth (10 ml) at 37 °C to late exponential phase (3 h). Bacteria were washed twice with PBS (pH 7.4) and the optical density (OD) of the cultures was determined at 600 nm (1 OD600 nm ≈1 × 109 bacteria/ml).
Two types of lung infection experiments were carried out. In the first set of experiments, to monitor the release of free active NE in function of time in the setting of infection, NE−/− and WT mice (n = 24 mice/genotype) were challenged intranasally with bacteria and sacrificed at fixed time points. Briefly, mice were anesthetized by intraperitoneal (intraperitoneal) injection of ketamine hydrochloride (75 mg/kg) and medotomidine hydrochloride (1 mg/kg). Next, mice were challenged intranasally with 50 μl of saline buffer (PBS) containing a predetermined dose of bacteria that killed 50% of WT mice (107 CFUs/per mouse, LD50 ≈ 107 CFUs) (10). One NE−/− mouse died during intranasal instillation. Control mice (n = 8 mice/genotype) were challenged with 50 μl of sterile PBS alone. WT mice and NE−/− mice that survived bacterial challenge were sacrificed at the designated time points of 4, 12, and 24 h post-infection (4 mice per each time point). Mouse lungs were analyzed for histology, inflammatory cell recruitment, and detection of free active NE. Of note, at the onset all NE−/− appeared clinically worse than WT mice. By 12 h, NE−/− mice exhibited signs of distress, which were markedly less severe in WT mice. At 24 h, both types of mice displayed a morbid state that was more marked in NE−/− mice. At this time point, whereas no death was recorded in WT mice 11 NE−/− mice (~45%) died. The survival of the remaining mice was monitored up to 72 h later and in accordance with our previous report (10), ~50% of WT mice survived against ~20% of NE−/− mice.
In the second set of experiments, groups of WT and NE−/− mice were challenged using experimental conditions of lung infection as above. The surviving mice (24 WT mice and 10 NE−/− mice) were then sacrificed at the 24-h time point, a time that corresponded to maximal PMN infiltration and maximal release of free active NE (Fig. 1) (10). Control mice were sacrificed 24 h post-saline instillation. Bronchoalveolar lavage fluids (BAL) of control and surviving mice were pooled as the following: 3 pools of control BALs (2 pools of 3 BALs and 1 of 2 BALs), 3 pools of infected WT BALs (4 BALS per pool), and 3 pools of infected NE−/− BALs (2 pools of 3 BALs and 1 of 4 BALs). In all mouse experiments, lungs were lavaged in situ using 1 ml of PBS (pH 7.4), cycled three times (18). Identical recoveries of BAL (700 μl/mouse) were obtained for each of the experimental groups. Both BALs and recovered lungs were processed for analysis as described below. These infection experiments were carried out twice for data reproducibility.
Mouse lungs corresponding to 24 h post-infection were processed for histology analysis as previously described (18). Total cell counts from BAL fluids were immediately performed by a hemacytometer. For differential counts, cells were cytospun, Wright-stained, and identified using standard morphological criteria. The remaining BAL samples were centrifuged for 5 min at 4 °C and supernatants were snap-frozen and stored at −80 °C until use.
J774 mouse macrophage cell line was seeded on a 24-well plate (ATGC Biotechnologies SA, Marne-la-Vallée, France) at 1 × 105 cells/well and grown for 3 days to confluence at 37 °C in 5% CO2 in DMEM supplemented with 1% FBS and 1% penicillin/streptomycin (19). In separate experiments, primary macrophages were elicited to the peritonea of C57BL/6 WT mice or C3H/HeJ (TLR4−/−) and control C3H/HeN mice following intraperitoneal injection with 3 ml of 3% Brewer thioglycollate broth (Difco Laboratories, Detroit, MI) (20). Four days later, the peritonea were lavaged with 10 ml of RPMI 1640 (Invitrogen). Cells were collected and washed twice in RPMI 1640 and resuspended in the medium supplemented with 1% autologous serum. Macrophages represented >95% of the cell population and >98% were viable as judged by differential counting and trypan blue dye exclusion, respectively. Cells were seeded on a 24-well tissue culture plate at 5 × 105 cells/well. Of relevance, to minimize PMN contamination and the potential confounding effect of endogenous NE, cells were cultured for 6 h at 37 °C in 5% CO2 to allow macrophages to adhere and washed twice with 1 ml of medium to remove nonadherent cells including PMNs. Cells were cultured for an additional 24 h to ensure for maximal purity of adherent macrophages and absence of PMN-derived NE as judged by light microscopy of cultured cell cytospins and elastin zymography on concentrated cell-free culture media (Nanosep filter units, Life Sciences, VWR International, Radnor, PA).
Prior to NE treatment, cell line or primary macrophages were washed twice with 1 ml of PBS and cultured for 1 h with corresponding serum-free medium. To determine optimal experimental conditions for maximal expression of the mediators, experiments of enzyme dose-response and time course of cell exposure to NE and culture post-treatment were carried out. J774 cells were treated with various concentrations of NE (100, 300, and 500 nm) for different times (1, 2, and 4 h), washed twice with 1 ml of PBS, and cultured at 37 °C in 5% CO2 in fresh complemented medium for different periods of time (2 and 4 h). The identified optimized conditions were applied also to primary macrophages. To ensure maximal removal of NE, culture media was checked for the absence of active NE against its specific substrate (21). In parallel experiments, NE was preincubated with AEBSF (1 mm) for 5 min at 37 °C prior to addition to cells. Controls included cells cultured in the absence of NE. Under these experimental conditions, over 90% of the cells were alive, as judged by morphologic criteria and trypan blue dye exclusion. Each treatment condition was performed in triplicate and cell culture experiments were repeated at least 3 times.
Cell lysates were prepared using RIPA buffer (50 mm Tris-HCl, 150 mm NaCl, 0.1% SDS, 0.5% sodium deoxycholate, 1% Triton X-100, 2 mm EDTA, 1 mm dithiothreitol (DTT), pH 7.4) (Sigma) and centrifuged at 600 × g for 30 min at 4 °C to remove cell debris. Next, lysates were exposed to a fixed NE concentration (500 nm) for the defined time periods (0.25, 0.5, 1, and 5 min). In parallel experiments, NE was preincubated with SLPI (10 μg) for 5 min at 37 °C or heat inactivated for 10 min at 65 °C prior to addition to cell lysates.
Cell-free WT and NE−/− BAL fluids of the 24-h time point post-infection were processed to assess the levels of various cytokines using the Raybio® Mouse Cytokine Antibody Array III according to the manufacturer's instructions (RayBiotech, Tebu-Bio, Le Perray-en-Yvelines, France) (22). Briefly, equal volumes of cell-free BAL fluids (400 μl) were added to antibody-coated membranes and detection of immunoreactive cytokines was carried out following sequential incubations of the membranes with biotinylated anti-cytokine antibodies and streptavidin-horseradish peroxidase and visualization by enhanced chemiluminescence. Images were obtained with a ChemiDoc XRS imaging system (Bio-Rad). Densitometric analysis was performed on captured images using Quantity One one-dimensional analysis software (version 4.5.2) (Bio-Rad). Spots of interest were normalized to an internal control after subtraction of the representative background sample. Next, to determine the concentration of cytokines of interest (TNF-α, MIP-2, and IL-6), equal volumes of WT and NE−/− cell-free BAL fluids (50 μl) were processed using a multiplex bead-based immunoassay kit (Bio-Rad). Cytokine assays were performed as described by the manufacturer's protocol. Each reaction in the kit was performed in triplicate. Both cytokine antibody array and multiplex assays were performed on all BAL pools.
NE or TLR-4 antigens were detected by Western blotting (18). Briefly, samples (cell-free BAL fluids (20 μl) and lung or cell lysates (35 μg)) were resolved by SDS-PAGE under reducing conditions and transferred to polyvinylidene difluoride (PVDF) membranes (Millipore Corp., Bedford, MA). The membranes were sequentially incubated with primary antibodies to mouse NE (dilution, 1:2,000) or TLR-4 (dilution, 1:500) followed by their respective horseradish peroxidase-conjugated secondary antibodies. When indicated, the membranes were stripped (100 mm β-mercaptoethanol, 2% SDS, and 62.5 mm Tris-HCl, pH 6.7, for 30 min at 50 °C) and immunoblotted using primary rabbit polyclonal anti-mouse albumin antibody (Rockland, Gilbertsville, PA) and its corresponding horseradish peroxidase-conjugated secondary antibody. Immunoreactive bands were visualized by enhanced chemiluminescence (ECL, Amersham Biosciences).
NE activity was determined by α-elastin zymography (23). Briefly, cell-free BAL fluids (20 μl) were electrophoresed under nonreducing conditions at 4 °C on 12% SDS-PAGE gels containing 1 mg/ml of elastin. Following electrophoresis, gels were soaked in 2% Triton X-100 for 30 min (twice), rinsed briefly, and incubated at 37 °C for 72 h in 50 mm Tris-HCl (pH 8.2), containing 5 mm CaCl2. The gels were then stained with Coomassie Blue and destained in 5% acetic acid and 10% methanol. Active NE appears as a transparent lysis band at ~29 kDa. NE activity in cell-free BAL fluids was further confirmed using conventional chromogenic peptide assays as previously described (21).
Total RNA isolation was performed using MasterPureTM RNA Purification kit as described by the manufacturer's protocol (Epicenter, Biotechnologies, Madison, WI) (19). Briefly, mouse lung tissues (half-lobe) or cultured macrophage cell pellets were lysed and RNA was extracted. Purified RNA was resuspended in RNase inhibitor-containing TE Buffer. RNA concentration and purity were determined by spectrophotometry (A260/A280 ratio). Integrity of RNA samples was verified by electrophoresis on 2% agarose gels and visualization under UV light. RNA aliquots of individual mice were pooled in the same manner as for BAL fluids at equal concentrations.
Total RNA samples (1 μg) were reverse-transcribed using the SuperScript First Strand Synthesis System (Invitrogen). Next, cDNAs were amplified by PCR using specific primers (40 cycles starting with DNA denaturation for 2 min at 94 °C; each cycle corresponded to denaturation for 15 s at 94 °C, primer annealing at 60 °C for 30 s and extension at 72 °C for 30 s). RT-PCR products were analyzed by electrophoresis on 1.5% agarose gels and quantified by densitometry using Quantity One Software (Bio-Rad). Levels of cytokine mRNA transcripts were normalized to the internal control 28 S mRNA. RT-PCR was performed in quadruplicate within each pool.
Forward and reverse primers for TNF-α, MIP-2, IL-6, and 28 S were designed as follows: TNF-α forward, 5′-GGG-ACA-GTG-ACC-TGG-ACT-GT-3′; TNF-α reverse, 5′-CTC-CCT-TTG-CAG-AAC-TCA-GC-3′; MIP-2 forward, 5′-CCA-CTC-TCA-AGG-GCG-GTC-AA-3′; MIP-2 reverse, 5′-CCC-CTT-ATC-CCC-AGT-CTC-TTT-CAC-3′; IL-6 forward, 5′-GAT-GCT-ACC-AAA-CTG-GAG-ATA-AAT-C-3′; IL-6 reverse, 5′-GGT-CCT-TAG-CCA-CTC-CTT-CTG-TG-3′; 28 S forward, 5′-CGG-AAT-TCG-CCA-CCA-GCC-GCC-TG-3′; 28 S reverse, 5′-CGT-CTA-GAC-TTT-CTC-CGT-TTA-CTT-GC-3′.
Briefly, total RNA of cultured cells was reverse-transcribed into cDNA using the SuperScript First Strand Synthesis System (Invitrogen). Next, real time-PCR amplification was performed on the ABI PRISMTM 7500 Sequence Detection System (PE Applied Biosystems, Carlsbad, CA) using Platinum SYBR® Green qPCR SuperMix-UDG with ROX (Invitrogen). PCR amplification conditions were as follows: initial DNA denaturation for 5 min at 94 °C, primer annealing at 60 °C for 1 min, and extension at 72 °C for 1 min for a total 40 cycles. Data analysis was performed with the SDS Software (Applied Biosystems). Real time RT-PCR was performed in triplicate and repeated at least 3 times in each experimental condition for data reproducibility.
Forward and reverse primers for TNF-α, MIP-2, IL-6, and GAPDH were designed as follows: TNF-α forward, 5′-GGC-AGG-TTC-TGT-CCC-TTT-CA-3′; TNF-α reverse, 5′-CTG-TGC-TCA-TGG-TGT-CTT-TTC-TG-3′; MIP-2 forward, 5′-GTG-AAC-TGC-GCT-GTC-AAT-GC-3′; MIP-2 reverse, 5′-ACT-CAA-GCT-CTG-GAT-GTT-CTT-GAA-3′; IL-6 forward, 5′-CAA-CCA-CGG-CCT-TCC-CTA-CTA-3′; IL-6 reverse, 5′-GTT-GGG-AGT-GGT-ATC-CTC-TGT-GA-3′; GAPDH forward, 5′-CAG-CCT-CGT-CCC-GTA-GAC-AA-3′; GAPDH reverse, 5′-CCC-AAT-ACG-GCC-AAA-TTC-G-3′.
Unless specified, data are expressed as mean ± S.E. Where appropriate, statistical differences between groups were tested using Student's unpaired t test. Differences were considered significant at p ≤ 0.05.
NE−/− and WT mice that survived infection were sacrificed at the designated time points (4, 12, and 24 h). Aliquots with equal volumes of cell-free NE−/− and WT BAL fluids were subjected to elastin zymography. As reflected by the intensity of band transparency, Fig. 1A (upper panel) shows the presence of active NE in cell-free WT, but not NE−/−, BAL fluids as early as 12 h that were markedly enhanced at the 24-h time point. Detection of active NE was further confirmed by incubation of cell-free BAL fluids with synthetic NE specific substrate (data not shown). Of relevance, BAL pools within each genotype showed similar NE profile. Moreover, the identity of the active band was confirmed by Western blotting using anti-mouse NE antibody (Fig. 1A, lower panel). Of note, there was no evidence of active NE in BAL fluids derived from saline-treated mice (data not shown).
With respect to immune cell recruitment in response to P. aeruginosa, there was a sharp increase of inflammatory cells in infected WT and NE−/− BAL fluids in function of time. However, total cell counts were comparable in both genotypes at all periods of infections (4, 12, and 24 h) (Fig. 1B). By comparison to time points 4 and 12 h, the influx of recruited cells was considerably high at 24 h. Morphologic analyses and differential counts of BAL cytospins revealed the predominance of PMNs in both types of BAL fluids (Fig. 1C). There was no detectable defect in the recruitment of other immune cells (macrophages and T cells) in NE−/− mice (data not shown). In accordance with our previous work (10, 23), H&E-stained lung sections of both NE−/− and WT mice showed similar patchy neutrophilic infiltrates, typical of pneumonia and there was a decrease in the recruitment of PMN at 48 h (data not shown). Regarding host defense against P. aeruginosa, no death was recorded in both types of mice at the 4-h time point. At 12- and 24-h time points, of the 24 NE−/− mice 3 and 8 mice died, respectively, and no WT mice succumbed to infection. The number of viable bacteria in NE−/− lung tissues was 2-fold greater than that seen in WT tissues at the 24-h time point (data not shown). These findings further reinforce the relative importance of NE for maximal PMN killing of P. aeruginosa.
It must be emphasized that, unlike our previous studies, this is the first study where we focused on the protective role of extracellularly released NE in the setting of lung infection. Curiously, the contribution of free active NE to antibacterial defense of the lungs has not been clarified. Because of the relatively high level of active NE by 24 h post-challenge, all subsequent studies were carried out with mouse BAL fluids and lung tissues corresponding to this time point.
To explore in vivo, the impact of released NE on cytokine production, we assessed the levels of various cytokines in equal aliquots of infected WT and NE−/− cell-free BAL fluids using cytokine antibody array approach. Our data revealed changes in the levels of a number of mediators in the absence of NE (supplemental Fig. S1). In this study, we focused on TNF-α, MIP-2, and IL-6, part of the early responsive cytokine network of infection. Remarkably, whereas their levels increased as expected in response to infection, semiquantitative analysis by densitometry found a marked increase of the proteins in cell-free WT BAL fluids when compared with those of NE−/− BAL fluids (Fig. 2A). Such an increase coincided with enhanced NE activity (Fig. 1A). Of note, cytokine levels in BALs of mice that received PBS were insignificant regardless of mouse genotype and their values were subtracted from those of infected BALs. Next, we employed the multiplex approach to quantitatively compare the levels of TNF-α, MIP-2, and IL-6. As shown in Fig. 2B, protein concentrations of these mediators were significantly low in NE−/− cell-free BALs by comparison to WT cell-free BALs confirming further the cytokine antibody array data. Of importance, these cytokine profiles were similar among BAL pools within each genotype.
The decreased levels of TNF-α, MIP-2, and IL-6 in cell-free NE−/− BALs could be attributed to protein degradation, which is unlikely or changes in transcript expression of these mediators involving NE. To test the latter hypothesis, mRNA expression of TNF-α, MIP-2, and IL-6 was examined by RT-PCR. Gel micrographs and densitometry histogram analyses demonstrate that levels of mRNAs encoding for TNF-α, MIP-2, and IL-6 were lower in infected NE−/− lungs when compared with WT lungs (Fig. 3, A–C). Of note, detected levels of TNF-α, MIP-2, and IL-6 transcripts in saline-instilled WT and NE−/− mice were insignificant (data not shown) and were subtracted from those of infected samples for densitometric analyses. Taken together, these data strongly suggest a role for NE in inducing the expression of TNF-α, MIP-2, and IL-6.
Macrophages are known producers of TNF-α, MIP-2, and IL-6. To begin to understand the role of NE in inducing TNF-α, MIP-2, and IL-6 expression, mouse J774 macrophages were exposed to purified NE and mRNA expression of TNF-α, MIP-2, and IL-6 was assessed by real time RT-PCR as described under “Experimental Procedures.” First, in dose-response and time course experiments, incubation of the cells with NE at 500 nm for 1 h followed by a 4-h incubation after removal of NE led to maximal mRNA expression of TNF-α (Fig. 4A). Using these optimized experimental conditions and by comparison to untreated cells, exposure of J774 cells to NE led to 2.2-, 3-, and 3.1-fold increases of TNF-α, MIP-2, and IL-6 mRNA transcripts, respectively (Fig. 4B). Of importance, NE affects neither the integrity not the viability of cell monolayers (data not show). These results support in vivo findings and indicate that NE can mediate the induction of TNF-α, MIP-2, and IL-6 expression in macrophages.
To further ensure that NE has a similar effect on primary macrophages, we employed thioglycollate-elicited peritoneal macrophages. Using similar experimental conditions as above, incubation of macrophages with purified NE resulted in a significant increase of TNF-α, MIP-2, and IL-6 mRNA expression by comparison to untreated cells (Fig. 5, A and B). Importantly, preincubation of NE with the AEBSF, a relatively stable serine protease inhibitor abrogated the inducing activity of the enzyme (Fig. 5A). Of relevance, cell monolayers were not disrupted in the presence of NE (supplemental Fig. S2). These data clearly indicate that NE utilizes its catalytic activity to induce mRNA expression of TNF-α, MIP-2, and IL-6 in primary macrophages.
In recent years, in vitro studies have reported the implication of TLR4 in NE-induced expression of IL-8 in human bronchial epithelial cells (24). Given that macrophages also express TLR4, we assessed the role of this receptor in NE-mediated induction of TNF-α, MIP-2, and IL-6 expression. Equivalent numbers of peritoneal WT macrophages and macrophages with nonfunctional TLR4 (TLR4−/−) were incubated with NE as described above. Analyses of real time RT-PCR data found that mRNA expression of TNF-α, MIP-2, and IL-6 increased by 2.2-, 2.6-, and 1.6-fold, respectively, in NE-treated WT cells when compared with untreated cells supporting further data of Fig. 4B. Of note, IL-6 expression was less impressive in primary WT C3H/HeN macrophages than cell line-derived macrophages, a finding that was reproducible with WT C57BL/6 cells. Although we are unable to explain such an observation, one plausible explanation could be related to differences in intracellular signaling events of IL-6 between the J774 cell line and primary macrophages. More importantly, there were no significant changes in mRNA expression of TNF-α, MIP-2, and IL-6 by TLR4−/− macrophages when compared with untreated WT or TLR4−/− cells (Fig. 5B). These findings indicate that NE-mediated induction of TNF-α, MIP-2, and IL-6 expression by macrophages involves, at least in part, TLR4.
Next, we sought to determine the faith of TLR-4 following exposure of macrophages to NE. Using an antibody against the N-terminal part of TLR-4, Western blot analysis revealed the presence of intact TLR-4 at the expected size in control cells, which disappeared in NE-treated cells (Fig. 6A). Next, immunoblotting analysis following incubation of WT macrophage lysates with NE found progressive loss of TLR-4 antigen in function of exposure time to NE (Fig. 6B). Interestingly, pretreatment of NE with its physiologic inhibitor, SLPI, or its denaturation by heat prevented degradation of TLR-4 (Fig. 6B and data not shown). Noteworthy, discrete TLR-4 cleavage products were detected by immunoblotting following migration of ×10 concentrated cell-free culture media on 16% SDS-PAGE gel (supplemental Fig. S3).
To determine the in vivo significance of our findings, we looked for evidence of TLR-4 degradation in cell-free BAL fluids and lung lysates of infected WT mice by immunoblotting. Fig. 7A shows the presence of distinct degradation fragments in cell-free WT BALs and only intact TLR-4 in lung lysates. To assess the contribution of NE to TLR-4 cleavage, we first observed the absence of immunoreactive TLR-4-derived fragments in cell-free BALs of both NE−/− and WT control mice (Fig. 7C, left panel, and data not shown). Next, Western blotting analysis of infected cell-free NE−/− BALs revealed a TLR-4 cleavage pattern that appears different from that seen in infected WT BALs. Also, protein levels of intact TLR-4 in infected lavaged NE−/− lungs was higher than that detected in infected lavaged WT lungs (supplemental Fig. S4), a finding that was corroborated by cell culture data in this work and previously reported studies (Fig. 6 and Ref. 24). Altogether, these data suggest the contribution of NE and potentially other enzymes to TLR-4 proteolysis. Of note, all immunoblotting micrographs of in vivo studies showed two unspecific bands that were prominent in infectious conditions despite the stringent membrane washing conditions. Also, high molecular bands above the intact TLR-4 band could be seen in some instances. These could be either unspecific immunoreactive fragments or high molecular bands resulting from binding of TLR-4 or its degraded fragments with BAL proteins. At any rate, detection of cleaved TLR-4 and active NE (Fig. 1A) suggest that both proteins co-localize in infected lungs and support the hypothesis that NE could target TLR-4 in vivo.
A number of in vitro studies have suggested that proteolytic modifications of cytokines by neutrophil serine proteases including NE change their biologic activities (15, 25). In the present work, we show that in the setting of P. aeruginosa lung infection, NE deficiency resulted in changes of protein levels of a variety of cytokines coinciding with increased mortality of mutant mice to infection. The increased or decreased levels of cytokine levels (supplemental Fig. S1) suggest the NE pleiotropic effect in infected situations. Probably, the purpose of these NE-mediated effects is to bring into harmony the levels of cytokines for host lung protection against P. aeruginosa. Focusing on TNF-α, MIP-2, and IL-6, we went on to provide compelling in vivo evidence that released active NE has the capacity to induce mRNA expression of these inflammatory mediators. Cell culture experiments were crucial in that not only did they allow us to circumvent the confounding stimulatory effects of bacteria, but clearly support the role of NE in inducing cytokine expression. It must be emphasized that this is the first report that shows both in in vivo and cell culture studies that active NE induces mRNA expression of TNF-α, MIP-2, and IL-6 resulting in the increase of their protein levels.
In our study, we focused on the 24-h time point post-infection because it corresponds to a sharp increase of neutrophil numbers and enhanced NE activity by comparison to other time points (23). But, the associated changes in cytokine expression could be seen at any time as long as significant amounts of free active NE are available and the microenvironment allows interaction of the enzyme and its target cells. Of interest, activated PMNs also release two other members of the neutrophil serine protease family, cathepsin G (CG) and proteinase 3 (PR3). As for NE, the relative importance of CG and PR3 in modulating cytokine expression (e.g. induction of TNF-α, MIP-2, and IL-6 transcript expression) will be best defined using mice that are deficient in all these proteases. Regarding cell culture studies, the concentration of NE used in this study is relevant because it can be reached or even exceeded in the lungs in diseased conditions (26). In fact, these suggested estimates do not even take into account possible evanescent quantum bursts of pericellular NE activity, local microenvironment, or the half-life of NE in tissues, which might increase the effective concentration of the enzyme (7). Importantly, enzyme inhibition experiments revealed that NE must be catalytically active to induce mRNA expression of TNF-α, MIP-2, and IL-6. In support of this observation, sivelestat, a selective synthetic neutrophil elastase inhibitor, has been reported to suppress the production of TNF-α and MIP-2 by LPS and/or NE-stimulated leukocytes in whole blood culture (27, 28). We have purposely used the synthetic inhibitor AEBSF instead of physiologic inhibitors such as SLPI because these latter have been reported to down-regulate the expression of inflammatory mediators (29, 30). Recently, NE has been shown to cleave SLPI (31).
TNF-α, MIP-2, and IL-6 are expressed following host recognition of pathogens and triggering of NF-κB signaling pathways. They have been reported to participate in the recruitment and/or activation of immune cells, particularly neutrophils in the setting of bacterial infections (32, 33). Surprisingly, in vivo changes in the levels of these mediators in the absence of NE has no statistically significant bearing on the recruitment of immune cells in response to P. aeruginosa suggesting the presence of a multitude of host and pathogen-derived factors that call in inflammatory cells.
The role of TNF-α in mediating host defense against bacterial infection has been reported in different studies using gene targeting and ligand or receptor blocking antibody approaches (34, 35). Also, this cytokine stimulates inflammatory mediator production such as MIP-2 and IL-6 that contribute to the recruitment and activation of inflammatory cells (36–38), strengthens the bactericidal capacity of phagocytic cells by an as yet undefined mechanism (39), and behaves as a secretagogue for neutrophils to secrete their granule content (32). Although MIP-2 is known to selectively attract PMNs, it is considered as a potent inducer of degranulation and enhancer of bacterial killing by phagocytic cells including PMNs themselves (40–42). In vivo studies using anti-MIP-2 serum in an infection model with Klebsiella pneumoniae revealed that this chemokine is an important mediator for effective bacterial clearance. Like TNF-α, IL-6 can be detected readily in stressed conditions as in infection. Regarding host defense against invading pathogens, genetically engineered mice deficient in IL-6 displayed impaired antibacterial protection (43). Collectively, these reported studies show the relative importance of TNF-α, MIP-2, and IL-6 in host protection against bacterial infections inferring that the observed decrease in their levels in our study should contribute to the increased susceptibility of NE-deficient mice to P. aeruginosa infection. It would be of interest to determine the role of NE in expression of the remaining cytokines and how this might relate to host susceptibility to infection.
The capacity of NE to induce gene expression of TNF-α, MIP-2, and IL-6 suggests that the enzyme interacts with a membrane-associated recognition receptor that signals cytokine expression. Although cytokine expression is mediated by TLRs among others pattern recognition receptors, we narrowed our focus in this study to TLR-4 whose implication in TNF-α, MIP-2, and IL-6 gene expression is largely documented (44–46). Moreover, it was reported that active NE induces IL-8 expression in vitro via TLR4 (24). To determine the implication of this latter in NE-induced cytokine expression, we sought to employ WT macrophages with intact TLR-4 and C3H/HeJ macrophages bearing nonfunctional TLR4 (47). There was a significant expression increase of TNF-α, MIP-2, and IL-6 mRNA expression in WT cells by comparison to mutant cells following cell exposure to NE. These data suggest that TLR-4 mediates NE-induced expression of these mediators, at least in mice. In line with our findings, a study by Tsujimoto and colleagues (28) showed that pretreatment of murine macrophage RAW 264.7 cells with an anti-TLR-4 antibody blocked NE-induced MIP-2 production in a dose-dependent manner. Indirectly, Calkins et al. (48) reported that TLR-4 was required for the production of MIP-2 because C3H/HeJ mice were unresponsive to LPS, the major ligand of the receptor. Also, TLR-4 deficiency in mice was accompanied by impaired production of TNF-α in response to lung challenge with P. aeruginosa (49).
To begin to elucidate the underlying basis for the NE-TLR-4 interaction, we demonstrate that the protease cleaves the receptor either in intact cells or present in cell protein extracts resulting in the generation of distinct fragments. More importantly, our in vivo findings using the murine P. aeruginosa pneumonia model indicate that NE could encounter TLR-4 in the lungs and contributes to its cleavage. Of relevance, TLR-4 was also cleaved in the absence of NE implying that other proteases might cleave the receptor as well. In this regard, whether CG and PR3 are as potent as NE to degrade TLR-4 needs to be assessed. This could be of important biological relevance given that these proteases are all released from neutrophil granules, but show different susceptibilities to inactivation by physiological protease inhibitors such as SLPI (50). Interestingly, inspection of the primary structure of TLR-4 especially within the putative extracellular domain (amino acid sequence from 26 to 638, NCBI accession number Q9QUK6) reveals the presence of peptide bonds that are preferred by the enzymes (51). Based on the estimated sizes of TLR-4 fragments detected by Western blotting and the epitope (amino acid sequence 198–395) that was used to raise anti-TLR-4 antibody, NE might likely cleave the receptor within its extracellular region. Of relevance, there exists a soluble small form of TLR-4 (sTLR-4) that has been reported to attenuate TLR-4 receptor functions (52). As such, the possibility that NE targets sTLR-4 preventing its interference with LPS-induced NF-κB activation and mediator production may not be ruled out.
That an active protease is involved in TLR-4 function is intriguing. Studies using different cell lines support, however, this finding. Pancreatic elastase exhibits a proinflammatory effect via activation of TLR-4 and NF-κB (55). Mansell and colleagues (56) reported that an active serine protease is required for TLR-4 function and NF-κB activation. McElvaney and colleagues (57) showed that NE-mediated IL-8 induction involves the signal transducers MyD88, IRAK, and TRAF6 leading to NF-κB activation and cytokine induction. Subsequently, the same group published that NE up-regulated IL-8 via TLR-4 and observed by Western blotting that protein levels of the receptor decreased considerably (24). Given that gene expression of TNF-α, MIP-2, and IL-6 is NF-κB-dependent as well, we inferred that these cytokines share at least the same signal transduction as that identified for IL-8. These observations along with our cell culture and in vivo data reinforce the hypothesis that NE may well target TLR-4 triggering by a signaling pathway that leads to NF-κB activation and induction of cytokine expression.
Although we have no clear evidence about the underlying mechanism of the NE-triggered function of TLR-4, among the possibilities by which the enzyme could act are: it could engage TLR-4 directly, especially since the protease has been reported to trigger TLR-4 signal transduction in the absence of the accessory protein MD2 (24). A simple cleavage in the extracellular domain might change the conformation of and activate the receptor. TLR-4 cleavage products could be biologically active and serve as a ligand for other receptor(s) to signal cytokine expression. NE might be involved in the generation of a ligand that signals through TLR-4 (56). The possibility that cleavage of TLR-4 corresponds to an epiphenomenon is plausible as well in that NE may interact with the receptor “rapidly” to generate an intracellular signaling event prior to its proteolysis. In this regard, NE has been shown to bind to the membrane of monocytes/macrophages (58–60). Alternatively, NE could act indirectly on TLR-4 function. In recent years, it was reported that NE activates the metalloprotease, meprin-α, which in turn releases TGF-α, a ligand for the epidermal growth factor receptor. EGF receptor co-localizes with TLR4 initiating a signal transduction cascade triggering activation of NF-κB and increased cytokine expression (61). Studies investigating the interaction mechanism(s) of NE and TLR4 are needed to better our understanding of the ensuing signaling cascade(s) that lead to cytokine expression.
The ability of NE to target TLR-4 and affect cytokine expression represents an important step in our understanding of the host strategies to combat insulting agents. It would be of relevance to determine whether neutrophil serine proteases including NE could target other TLRs such as TLR1, TLR2, TLR5, and TLR6, which are also localized on the cell surface and recognize microbial membrane products (62). Consistent with our previous findings, NE can accumulate in close spatial proximity to local cells at sites of active inflammation (18). This suggests that unchecked NE could interact and activate host cells. In support of this hypothesis, our culture study (this work) in addition to a series of published reports indicates that NE is capable of activating various types of host immune (e.g. dendritic cells) and nonimmune (e.g. epithelial cells) cells in vivo (54, 63). The rationale of investigating macrophages is that this cell population plays a prominent role in lung immunity by orchestrating inflammatory and immune responses (53, 64, 65). In fact, these cells represent the most potent sources for TNF-α, MIP-2, and IL-6 in the setting of bacterial infection. Also, TLR-4 is not presented at the apical surface of lung epithelial cells and does not function in signaling responses to P. aeruginosa (54).
Previously, we have reported that NE is required for maximal PMN intracellular killing of P. aeruginosa. Collectively, the present findings reveal a protective role for extracellular NE against the pathogen. They clearly demonstrate that NE, an endogenous effector, could also participate in the orchestration of lung inflammatory response against P. aeruginosa infection by modulating the expression of cytokines (e.g. induction of the expression of the proinflammatory TNF-α, MIP-2, and IL-6). The enzyme might be required to drive a robust NF-κB pathway activation and subsequent cytokine expression. Thus, the net effect of NE (intra- and extracellularly) on P. aeruginosa and perhaps other Gram-negative bacteria is an “astute” response for better lung protection. In conclusion, unopposed active NE is anticipated in any PMN-rich inflammatory milieu. In light of our findings, the long held view that considers NE as “a prime suspect” in inflammatory and tissue-destructive diseases as in acute or chronic pulmonary diseases will need to be carefully reassessed. More importantly, therapeutic strategies aiming at NE inhibition should take into account the physiologic contribution of the enzyme to host inflammatory response against pathogenic agents.
We thank Véronique Laplace and Christine Terryn for excellent technical assistance. We also thank Prof. Soman Abraham (Duke Bacteriology Research Unit, Department of Pathology, Duke University Medical Center, Durham, NC) for critical reading of the manuscript.
*This work was supported by the Inserm Avenir Program, Fondation pour la Recherche Médicale, Agence Nationale Recherche, Association Régionale Pour I'Aide aux Insuffisants Respiratoires de Champagne Ardenne, and Contrat de Projets Etat-Région.
This article contains supplemental Figs. S1–S4.
3The abbreviations used are: