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Haemophilus influenzae efficiently colonizes and persists at the human nasopharyngeal mucosa, causing disease when it spreads to other sites. Nitric oxide (NO) represents a major antimicrobial defense deployed by host cells in locations colonized by H. influenzae during pathogenesis that are likely to vary in oxygen levels. Formate-dependent nitrite reductase regulator (FNR) is an oxygen-sensitive regulator in several bacterial pathogens. We report that fnr of H. influenzae is required for anaerobic defense against exposure to NO donors and to resist NO-dependent effects of gamma interferon (IFN-γ)-activated murine bone marrow-derived macrophages. To understand the mechanism of resistance, we investigated the role of FNR-regulated genes in defense against NO sources. Expression analysis revealed FNR-dependent activation of nrfA, dmsA, napA, and ytfE. Nonpolar deletion mutants of nrfA and ytfE exhibited sensitivity to NO donors, and the ytfE gene was more critical for survival. Compared to the wild-type strain, the ytfE mutant exhibited decreased survival when exposed to macrophages, a defect that was more pronounced after prior stimulation of macrophages with IFN-γ or lipopolysaccharide. Complementation restored survival of the mutant to the level in the parental strain. Increased sensitivity of the ytfE mutant relative to that of the parent was abrogated by treatment of macrophages with a NO synthase inhibitor, implicating YtfE in resistance to a NO-dependent pathway. These results identify a requirement for FNR in positive control of ytfE and indicate a critical role for ytfE in resistance of H. influenzae to reactive nitrogen species and the antibacterial effects of macrophages.
Haemophilus influenzae is a gram-negative bacterial pathogen of humans that colonizes the nasopharynx of healthy individuals, causing primarily asymptomatic infections. Disease occurs by extension of bacteria to privileged anatomical sites, leading to otitis media, meningitis, septicemia, or respiratory infections (59). Despite the availability in developed countries of an effective vaccine against type b strains, which cause meningitis, H. influenzae type b remains a significant cause of disease worldwide (13). The incidence of infection by nontypeable H. influenzae strains (NTHi) appears to be unaffected by the type b vaccine, and NTHi remain a frequent cause of otitis media (65). NTHi is also a significant medical concern in recurring bacterial exacerbations of chronic obstructive pulmonary disease and cystic fibrosis (33, 43, 58, 60). The mechanisms by which this bacterium survives and causes disease at diverse sites of infection are not well understood.
In different tissues or stages of pathogenesis, bacteria must recognize environmental signals as cues for appropriate regulation of gene expression. The results of recent studies suggest that levels of oxygen availability generate signals leading to modulation of virulence factor expression in H. influenzae. Modification of H. influenzae's lipooligosaccharide with phosphorylcholine, a virulence-associated structure that is also targeted by host immunity (86), is upregulated when oxygen availability is decreased (89). Genes similar to arcA and fnr, global genetic regulatory genes critical to the response to anaerobiosis in other species, are present in H. influenzae. The H. influenzae ArcA/ArcB system is a two-component signal transduction system that is active under low-oxygen conditions and functions via a phosphorelay from a sensor kinase protein, ArcB, to activate the DNA binding response regulator, ArcA, analogous to the well-characterized signal-transduction mechanism of ArcA/ArcB of Escherichia coli (30, 45). Phosphorylated ArcA then controls a set of genes involved in adaptation to respiratory conditions of growth that is similar but not identical to the ArcA/ArcB regulon of E. coli (47, 62, 72, 91). In support of the hypothesis that H. influenzae encounters a low-oxygen environment in vivo in the host, H. influenzae mutants lacking ArcA are attenuated for survival in murine models of pathogenesis (19, 91).
In E. coli and other bacteria, transitions between aerobic and anaerobic environments also involve the global regulator FNR (formate-dependent nitrite reductase regulator), a cyclic AMP receptor protein/catabolite activator protein-like transcriptional regulatory protein that activates and represses the expression of multiple genes under anaerobic conditions (75). FNR's functional role in H. influenzae has not been characterized; however, its predicted amino acid sequence is 79% identical to E. coli FNR, which has been studied intensively. Unlike ArcB, which indirectly senses oxygen via changes in the reduction/oxidation status of the quinone pool (52), FNR of E. coli is considered to be a direct oxygen sensor by virtue of its iron-sulfur center, which is required for maintaining an active conformation that promotes dimerization and DNA binding (46). Upon oxidation, the iron-sulfur center undergoes a transition leading to the conversion of FNR to its inactive monomeric form (16, 20). FNR and FNR-like anaerobic regulators have been implicated in the pathogenesis of Salmonella enterica serovar Typhimurium (28), Neisseria meningitidis (8), Actinobacillus pleuropneumoniae (5), and Bordetella pertussis (92). The mechanisms by which these regulators contribute to virulence are not fully understood; however, FNR-regulated genes that participate in virulence properties have been identified in several organisms. A. pleuropneumoniae HlyX is an FNR-like regulator that controls dmsA and aspA, genes involved in anaerobic respiration that are required for virulence (39). In addition, genes of nitrosative defense constitute an important class of FNR-regulated genes that have been identified in diverse bacterial species, including Campylobacter jejuni (25), S. enterica serovar Typhimurium (28, 32), and E. coli (14, 64, 67, 68).
During immune responses to bacteria, host cells produce the free radical gas nitric oxide (NO) via metabolism of l-arginine by inducible NO synthase (iNOS) (for reviews, see references 10 and 51). NO is highly reactive and unstable in biological systems, especially in the presence of oxygen at neutral pH, which results in its conversion to nitrite (87). Additional reactions with NO generate a collection of other reactive nitrogen species (RNS), such as S-nitrosothiols, which are thought to exceed the cytotoxicity of NO, leading to diverse mechanisms of antimicrobial activity (11, 26). Alcohol dehydrogenase (adhC) has been implicated in defense of H. influenzae against one type of RNS, S-nitrosoglutathione (GSNO), under normoxic conditions (44); however, information regarding how H. influenzae defends itself from host-derived RNS under oxygen-limiting conditions has not been reported. Genome analysis suggests that H. influenzae possesses a single nitrite reductase, encoded by the nrfABCD operon, comprised of nitrite reductase genes similar to those implicated in NO metabolism in other species (66, 76). Also present in the H. influenzae genome is the ytfE gene, predicted to encode a di-iron protein similar to a family of iron-sulfur repair proteins involved in RNS and reactive oxygen species resistance in E. coli and a diverse group of additional bacterial species (63).
In this study, we investigated the role of FNR in regulation of the defense of H. influenzae from RNS generated in vitro and by activated macrophages. A single-copy, chromosomal reporter fusion to nrfA was generated to monitor expression from its endogenous promoter. The fusion strain was used to identify a range of low-oxygen conditions that are required for FNR-mediated transcriptional activation in H. influenzae. Control of candidate FNR-regulated genes, including the nrfA and ytfE genes, which have not been investigated in this bacterium, was assessed by mRNA analysis. Mutants lacking FNR or genes that it regulates were evaluated for resistance to two chemical donors of NO in vitro, implicating each of these genes in RNS defense. To investigate the context in which anaerobic sensitivity to RNS could influence pathogenesis, the effect of macrophage activation with lipopolysaccharide (LPS) or gamma interferon (IFN-γ) on the survival of wild-type, fnr, and ytfE H. influenzae strains was determined, and the role of NO in this process was addressed by using an inhibitor of NOS. We report that FNR is required for anaerobic defense of H. influenzae against exposure to NO donors and IFN-γ-treated macrophages. Furthermore, the results demonstrate positive control by FNR of H. influenzae ytfE and reveal a critical role for ytfE in resistance of this bacterium to RNS and the antibacterial effects of activated macrophages.
(Parts of the manuscript have been included in a Ph.D. dissertation by Jane Harrington.)
H. influenzae Rd BA042, a capsule-deficient derivative obtained from the serotype d source that was used to generate Rd KW20 (89), and NTHi clinical isolate NT127 (provided by Robert N. Husson), were grown in brain heart infusion broth (BHI) supplemented with 10 μg/ml hemin, 10 μg/ml NAD, 2 mM d-xylose (sBHI) or on sBHI agar plates at 35°C. For growth in semidefined medium lacking detectable nitrate or nitrite sources, bacteria were grown in MIc, a low-nutrient medium capable of supporting growth of H. influenzae (7). To generate a condition approximating anaerobiosis, strains were grown in anaerobic chambers with BBL GasPak Plus generators to deplete oxygen (Becton Dickenson and Company, Sparks, MD); for simplicity, these are termed low-oxygen conditions. Development of competence for transformation of H. influenzae was accomplished as previously described (7). For selection of Rd-derived strains, antibiotics at the following concentrations were used: 8 μg/ml tetracycline (Tc), 20 μg/ml kanamycin (Km), and 10 μg/ml gentamicin (Gm). Selection of NT127 strains was conducted at the same antibiotic concentrations, except that Gm was used at 20 μg/ml.
Plasmids and PCR products were constructed using standard molecular biology techniques (3). For complementation of mutants, DNA fragments were amplified by PCR and cloned between adjacent SapI restriction sites of the chromosomal delivery vector pXT10, which does not replicate in H. influenzae (90). The pXT10-based plasmids contain upstream (xylF) and downstream (xylB) homologous regions flanking the SapI cloning sites that allow precise fusion of genes of interest to the xylose-inducible xylA promoter, as previously described (90). Recombination at the xylose catabolic locus replaces the endogenous xylA gene with the cloned fragment and the tetAR Tc resistance (Tcr) cassette. Plasmids were linearized by digestion with ApaLI and introduced into competent H. influenzae, and tetracycline-resistant recombinants were selected on sBHI agar plates. Double-crossover homologous recombination between the H. influenzae chromosome and constructs for each mutation in this study was confirmed by PCR with primers specific for sequences outside the inserted recombinant region. All genes introduced for complementation of mutations were verified by DNA sequence analysis and were identical to corresponding genes in H. influenzae Rd KW20 (GenBank accession number L42023, updated 21 December 2005).
For gene replacement of nrfA with a hemagglutinin (HA)-tagged reporter fusion construct, a 1,360-bp fragment corresponding to the region immediately 5′ of the nrfA translational start site was amplified by PCR using primers rtnrfARI and 5nrfAexp, with Rd genomic DNA as template. Primer 5nrfAexp, which includes a BssHII restriction site at its 5′ end, binds to the nrfA translational start codon and to the sequence directly upstream of the start codon. A second fragment was generated by PCR using primers 3nrfArevexp and lt1068nrfARI, which amplify the 725-bp product of a corresponding region downstream of nrfA starting at the termination codon. Primer 3nrfArevexp carries a StuI restriction site at its 5′ end and binds the nrfA stop codon, the nrfB start codon, and an nrfB sequence immediately downstream of the start codon. Primers 5nrfAexp and 3nrfArevexp include 17 bp of overlapping sequence, allowing them to be joined by overlap extension PCR using primers rtnrfARI and lt1068nrfARI. The resulting product was cloned into the EcoRI site of vector pBR322, generating vector pBNA1. The overlap region added by primers 5nrfAexp and 3nrfArevexp introduced BssHII and StuI cloning sites, allowing the hel gene coding sequence, which was amplified by PCR from Rd genomic DNA with primers 5′helORF and 3′helORF, to generate an 863-bp product incorporating a C-terminal HA epitope tag to be precisely cloned from the translation initiation to the termination codon of nrfA, resulting in vector pBNheltag. Finally, a fragment containing the nrfA promoter upstream of the hel-HA fusion gene flanked by sequences identical to those flanking nrfA was generated by PCR with primers rtnrfARI and lt1068nrfARI, using pBNheltag as a template. The coding sequence of the fusion protein was verified by DNA sequence analysis. The resulting 3-kb product was used to transform Rd, and recombinants were screened nonselectively for loss of the ability to reduce nitrite, generating strain RdHA. This strain contains an HA epitope-tagged reporter controlled by the 5′ nrfA transcriptional and translational signals. In addition, the H. influenzae lipoprotein P4, encoded by the hel gene, is reported to be accessible to antibodies at the cell surface (35), potentially allowing detection applications that do not require cell lysis.
To complement the nrfA mutation in strain RdHA with the wild-type nrfA, introduced at the xyl locus, the nrfA gene coding sequence and upstream promoter elements were amplified from Rd by PCR using primers 5′nrfAsapI and 3′nrfAsapI to generate a 1.8-kb fragment. SapI restriction sites at the 5′ ends of the primers were used to clone the resulting fragments into the adjacent SapI sites of pXT10, resulting in vector pXTnrfAC. To generate strain RnrfAC, pXTnrfAC was digested with ApaLI, gel purified, and transformed into RdHA, followed by selection on sBHI containing Tc.
Strain Rfnr containing a transposon insertion mutation in fnr was generated by targeted in vitro transposon mutagenesis of the fnr locus with the Himar1 derivative magellan1, followed by identification of the desired mutant by genetic footprinting as previously described (1). Rfnr contains a magellan1 insertion ~170 bp from the 5′ end of the 770-bp fnr protein-coding region. To generate strain RHAfnr, a fragment consisting of 1.4 kb upstream of the transposon insertion in fnr to 1.3 kb downstream of the insertion was amplified by colony PCR using Rfnr as a template with primers 5′FNRupstreamCK and 3′FNRdownstreamCK. The resulting product was used to transform strain RdHA, resulting in strain RHAfnr. Kanamycin-resistant recombinants were selected on sBHI agar, and the resulting mutation was verified by PCR. To complement fnr mutants, a 1,286-bp PCR product containing the HI1425 gene and its putative promoter region upstream (468 bp 5′ of initiation codon ATG of fnr coding region) was amplified from Rd with primers FNRMlu1FORWARD and FNRMlu1REV. This PCR product was digested with MluI and cloned into the AscI site of pXT10. This plasmid, pXTfnrC, was introduced into the xyl locus of strain Rfnr or RHAfnr by double-crossover homologous recombination, and transformants were selected on sBHI agar containing Tc to create strains RfnrC and RHAfnrC, respectively.
A nonpolar, in-frame deletion of HI677 (ytfE) was created by replacing the protein-coding sequences with the aacC1 gentamicin resistance cassette to create strain Rytfe by overlap extension PCR as follows. A 1,042-bp PCR product containing the 5′ flanking region of HI1677 was amplified from Rd with primers 1kb5′1677 and HI1677-5′out. A 1,044-bp PCR product containing the 3′ flanking region of HI1677 was amplified from Rd with primers HI1677-3′out and 1kb3′1677. A 536-bp fragment containing the aacC1 gentamicin resistance cassette was amplified with primers aacC1-5ORF and aacC1-3ORF from pBSL182 (2). The 1,042-bp, 1,044-bp, and 536-bp products were combined in a PCR with primers 1kb5′1677 and 1kb3′1677. The resultant 2.6-kb product was introduced into Rd, and Gmr transformants were selected on sBHI agar containing Gm to create strain Rytfe.
To generate RytfeC, the ytfE gene, including its native promoter, was amplified by PCR using Rd as template with primers 1677upcomp and 1677-3′sap, which introduce SapI restriction sites flanking ytfE and its predicted promoter region. The resulting 1.4-kb product was digested with SapI and cloned into SapI-digested pXT10. The resulting plasmid, pXTPytfeC, was linearized with ApaLI and transformed into Rytfe with selection for Tcr.
The NT127 xyl locus differs from that of Rd in that it contains xylAB but lacks the xylHGF genes. To facilitate complementation of mutants by introduction of genes at the NT127 xyl locus with our exchange vector, pXT10, the xylHGF genes were cloned into a Kmr-marked derivative of pXT10 and introduced into NT127 at the xyl locus as they are arranged in Rd, providing sequences for homologous recombination with pXT10 and its derivatives. The resulting strain was transformed with pXT10 or the ytfE mutation to generate strains NTV (“vector only” strain) and NytfE (ytfE mutant), respectively. To generate strain NytfeV containing a ytfE mutation and “empty vector” sequences, the ytfE mutation was amplified with primers 3′HI1676 and 3′ytfe.down using RytfE as a template. The resulting 3-kb PCR product was transformed into the parental strain NTV, and transformants selected for Gmr under anaerobic conditions. Sequence analysis verified that the ytfE-flanking regions of Rd and NT127 strains are identical. For complementation of the ytfE mutation in the NT127 background, primers 5′ytfEc and 1677-3′sap were used to amplify the ytfE protein-coding sequence, followed by digestion with SapI and cloning into the SapI sites of pXT10, placing ytfE under the control of the xylA promoter. The resulting plasmid was transformed into Nytfe with selection for Tcr. RfnrV, RHAfnrV, and RytfeV were generated by transforming an ApaLI-digested linear pXT10 fragment into Rfnr, RHAfnr, and RytfE, respectively, with selection for Tcr.
To measure reduction of nitrite by H. influenzae, triplicate cultures of MIc in 25-ml DeLong flasks (Bellco) were inoculated from standing overnight cultures to a starting optical density at 600 nm (OD600) of 0.02. To minimize aeration, flasks were filled to the necks to a 30-ml total volume (25-ml markings denote the typical maximum culture volume, but flasks hold 30 ml when filled to the neck). The resulting cultures were incubated at 35°C with shaking at 250 rpm. When cultures reached mid-log phase (OD600 of 0.4 to 0.5), 7.5 OD600 equivalents of cells were harvested by centrifugation at 5,000 × g for 5 min and resuspended in 25 ml MIc supplemented with NaNO2 to a starting concentration of 0.25 mM in 25-ml Erlenmeyer flasks. These cultures were then incubated at 35°C with shaking at 200 rpm, and 200-μl aliquots were removed at selected time points and stored on ice in a 96-well flat-bottom dish until completion of the assay. The 96-well dish was centrifuged at 5,000 × g at 4°C for 5 min to pellet cells. The presence of nitrite in cell-free-supernatant medium was quantified by using the Griess reaction. Supernatants were incubated with 50 μl of a solution of 5% phosphoric acid, 1% sulfanilamide for 5 min, followed by the addition of 50 μl of aqueous 0.1% N-N-1-napthylethylenediamine dihydrochloride for 5 min. Subsequently, the OD540 was recorded with a VersaMax microplate reader (Molecular Devices, Sunnyvale, CA) and nitrite concentrations were calculated using a standard curve. For assays of nitrite reduction after a 24-hour incubation, cultures were inoculated in 200 μl sBHI supplemented with NaNO2 at a concentration of 1.0 mM to a starting OD600 of 0.01 in a 96-well dish. Cultures were incubated for 24 h at 35°C, and remaining nitrite concentrations in the supernatants determined as described above.
To determine the role of FNR in reporter expression, strains were grown to mid-log phase in 30 ml of sBHI in 25-ml DeLong flasks (filled to the neck) and were lysed in sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) sample buffer, boiled for 5 min, centrifuged for 1 min at 16,000 × g, and then resolved on 12% SDS-PAGE gels (0.4 OD600 equivalents per lane). Gels were electrotransferred to Immobilon-P membranes (Millipore Corporation, Billerica, MA). Aliquots of each sample were analyzed in parallel on a gel stained with Coomassie blue (3) to verify equal sample concentrations. Membranes were blocked with 1% Tris-buffered saline-0.1% Tween (TBS-T) supplemented with 1% instant milk, probed with a 1:1,000 dilution of anti-HA.11 (Covance, Berkeley, CA) for 60 min, washed with TBS-T, probed with a 1:5,000 dilution of secondary antibody goat anti-mouse immunoglobulin G-horseradish peroxidase conjugate (Upstate, Lake Placid, NY) for 30 min in TBS-T, and washed with TBS-T. The Pnrf-HA-encoded reporter protein was then detected with a West-One Western blot detection system (iNtRON Biotechnology).
To monitor the expression of the Pnrf-hel-HA fusion under a range of aeration conditions, volumes of 7.5, 10, 30, 45, or 60 ml (filled to the neck for 60 ml) of sBHI in 50-ml DeLong flasks (nominal volume) were inoculated in triplicate from overnight cultures at a starting OD600 of 0.005 and incubated at 35°C with shaking at 250 rpm. When cultures reached mid-log phase, 0.25 OD600 equivalents of each sample were centrifuged for 5 min at 16,000 × g, the supernatant was removed, and the resulting pellets were resuspended in Hank's balanced salt solution. Subsequently, samples were lysed in SDS-PAGE sample buffer and proteins were separated by 12% SDS-PAGE. The upper portion of the gel (~98 to 64 kDa) was removed and stained with Coomassie blue to verify equal loading. The lower portion of the gel was electrotransferred to Immobilon-P and analyzed via Western blotting as described above, except that blocking was conducted with TBS-T containing 2% instant milk and secondary probing with goat anti-mouse immunoglobulin G ReserveAP phosphatase-labeled antibody (KPL, Gaithersburg, MD) at a 1:30,000 dilution. The Pnrf-HA-encoded reporter protein was then detected with CDP-Star detection reagent (GE Healthcare). Densitometry was performed on the original scanned image using ImageJ (National Institutes of Health, Bethesda, MD), and local background was measured and subtracted for each individual lane. The contrast of the displayed blot was adjusted (applied to entire image) using Adobe Photoshop 10.0.1 (Adobe Systems, Incorporated). In parallel with sample collection, for each of the three cultures per condition, percent oxygen saturation levels were measured with a Clark-type oxygen probe (model DO-166; Lazar Research labs) calibrated according to the manufacturer's instructions.
To determine the sensitivity of fnr, nrfA, and ytfE mutants to RNS, strains were cultured overnight in an anaerobic chamber with BBL GasPak Plus generators and the cultures were used to inoculate 5 ml of sBHI broth in 18- by 150-mm culture tubes to an initial OD600 of 0.02 and then incubated in an anaerobic chamber at 35°C with shaking at 120 rpm until they reached mid-log phase. Triplicate 200-μl aliquots of each culture were incubated in sBHI in 96-well flat-bottom microtiter plates overnight in an anaerobic chamber in the presence or absence of RNS donors and then diluted and plated on sBHI agar for enumeration of CFU. For acidified sodium nitrite (ASN) exposure, 5 × 105 cells were incubated in sBHI adjusted to pH 6.5 in the presence or absence of a final concentration of 10 mM NaNO2. GSNO (Sigma-Aldrich St. Louis, MO) was used at a final concentration of 5 mM with 5 × 104 cells in sBHI at pH 7.5.
To evaluate the dynamics of killing by ASN, time-kill curves were generated. Anaerobic overnight cultures of strains were used to inoculate sBHI adjusted to pH 5.5 for an initial OD600 of 0.01. Cells were then inoculated in triplicate 200-μl cultures into 96-well flat-bottom microtiter plates and treated with the addition of NaNO2 to a final concentration of 15 mM or not treated. For low-oxygen conditions, dishes were incubated at 35°C in individual BD GasPak EZ anaerobe gas-generating pouches. For higher-oxygen conditions, microtiter plates were incubated in ambient air (35°C). Bacteria were diluted and plated for enumeration of CFU at various time points throughout the assay.
Putative FNR binding sites were identified upstream of nrfA, napA, dmsA, and ytfE based on previously reported candidate FNR binding sequences for E. coli and H. influenzae (31, 69, 83). DNA sequencing verified that promoter regions in Rd BA03 were identical to those of Rd KW20 (GenBank accession number L42023, updated 21 December 2005). Putative binding sequences were used to generate a sequence logo diagram (http://weblogo.berkeley.edu/logo.cgi/).
To quantify mRNA of FNR-controlled genes, total RNA was isolated from 40-ml cultures of Rd, RfnrV, and RfnrC grown to mid-log phase in anaerobic chambers by using TRIzol reagent (Invitrogen). Isolation of intact RNA for each sample was verified by agarose gel electrophoresis with ethidium bromide staining. RNA was quantified by measuring UV absorption at 260 and 280 nm with a SmartSpec plus spectrophotometer (Bio-Rad, Hercules, CA). Samples were then treated with DNase I (Ambion, Austin, TX) and phenol extracted. To verify the absence of DNA contamination, 1 μg of the RNA sample was used as template for real-time PCR containing primers HI1069-5′ and HI1069-3′ (89) and iQ SYBR green supermix (Bio-Rad Laboratories, Hercules, CA) with fluorescence in real-time PCR measured with a DNA Engine Opticon II system (MJ Research, Waltham, MA). No DNA contamination was detected with a limit of detection equal to 0.001 ng, as determined via a genomic DNA standard curve. The RNA (5 μg total) samples served as a template for cDNA synthesis using random primers (New England Biolabs, Beverley, MA) and SuperScript II reverse transcriptase (Invitrogen, Carlsbad, CA). PCR was performed using iQ SYBR green Supermix, with fluorescence measured in real-time PCR by using the DNA Engine Opticon II system. Amounts of 1/10 of the cDNA reactions were used as a template for PCR with primers specific for nrfA (HI1069-5′ and HI1069-3′; 89), dmsA (HI1047-5′ and HI1047-3′; 89), napA (5′napA-RT and 3′napA-RT), ytfE (5′ytfe-RT and 3′ytfe-RT), and rpoA (HI0802-5′ and HI0802-3′; 91). Approximate transcript values were calculated from a standard curve generated with each primer set. Relative expression was standardized by determining the ratio of the transcript value of each gene to the rpoA transcript value. Control reactions were performed in parallel with mock cDNA reactions generated without reverse transcriptase to verify specific amplification, and the presence of single products of correct approximate sizes was verified by agarose gel electrophoresis and ethidium bromide staining. Real-time cycler conditions were as described previously (91).
Bone marrow-derived macrophages (BMM) were generated by differentiating C57BL/6J bone marrow cells in a complete bone marrow medium (Dulbecco's modified Eagle's medium, 10% heat-inactivated fetal calf serum [Invitrogen], 100 U/ml penicillin [Invitrogen], 100 μg/ml streptomycin [Invitrogen], and 10% L929 fibroblast-conditioned medium as a source of macrophage colony-stimulating factor) for 6 days in 10-cm petri dishes (VWR). Prior to exposure to bacteria, BMM were washed in antibiotic-free bone marrow medium and incubated with or without 100 U/ml IFN-γ (Peprotech) for 24 h, 100 μM Nω-nitro-l-arginine methyl ester HCl (L-NAME) for 24 h, or 100 ng/ml LPS for 2 h.
To evaluate the survival of fnr and ytfE mutants in the presence of BMM, strains were cultured anaerobically as described above for the RNS sensitivity assay. Cultures were then diluted in Dulbecco's modified Eagle's medium, and 2.5 × 105 bacterial cells were aliquoted into each well of a 96-well dish containing 5 × 104 adherent BMM per well. The 96-well dish was centrifuged for 5 min at 200 × g, sealed in a BD GasPak EZ anaerobe gas-generating pouch, and incubated for 0 or 30 min at 36°C, 5% CO2. Following incubation, wells were treated with saponin to a final concentration of 0.1%, a concentration that does not influence bacterial viability, and samples were triturated vigorously by pipetting to lyse macrophages and release bacteria. Samples were diluted and plated for CFU enumeration. This assay measures the ability of BMM to limit survival of H. influenzae by killing or growth inhibition during a transient incubation. Relative survival is calculated as the ratio of the number of bacteria recovered after treatment with macrophages compared to the number of bacteria recovered from parallel cultures incubated in medium alone.
As a starting point for investigating whether FNR of H. influenzae participates in the regulation of defense against RNS, including NO, we evaluated the role of FNR as an anaerobically active regulator of nitrite reductase in H. influenzae. The H. influenzae genome contains a single putative nitrite reductase locus that is comprised of the nrfABCD genes (57%, 50%, 64%, and 48% amino acid identity, respectively, to their putative homologs in E. coli) (18, 29, 37). To verify the predicted role of nrfA in reduction of nitrite by H. influenzae, we compared the parental strain Rd, a mutant containing a nonpolar deletion of nrfA (strain RdHA), and a complemented strain in which nrfA was provided in trans to the deletion (RnrfAC) for their ability to reduce nitrite within 24 h in the presence of 1 mM nitrite under microaerobic conditions (Fig. (Fig.1A).1A). At 24 h, nitrite levels from cultures of Rd and the nrfA-complemented strains were below the limit of detection. In contrast, the nrfA mutant was incapable of reducing nitrite and the concentrations at the end of the assay remained equivalent to the initial values. These results indicate that nrfA is required for nitrite reduction by H. influenzae in this assay.
To determine whether FNR is required for nitrite reduction, we compared microaerobically grown cultures of the parental strain (Rd); an fnr disruption mutant (Rfnr); an fnr mutant carrying vector sequences in the xyl locus (RfnrV); and an fnr mutant complemented with a wild-type copy of the fnr gene, provided at the xyl locus (RfnrC), for the ability to deplete nitrite from culture supernatants during a 1-h assay (Fig. (Fig.1B).1B). Nitrite was steadily depleted from supernatants of Rd and RfnrC cultures during the first 45 min, at which time residual nitrite became undetectable. In contrast, the concentrations of nitrite did not detectably decrease in culture supernatants of the fnr mutants (Rfnr and RfnrV), providing evidence that FNR is required for appreciable activity or expression of nitrite reductase under these conditions.
To facilitate studies of nitrite reductase gene expression, we utilized strain RdHA, a nrfA deletion mutant which also serves as a reporter strain, as it contains the nrfA promoter precisely fused to the hel gene encoding H. influenzae protein P4. A fusion of an influenza virus HA epitope to the C terminus of P4 allows immunological detection of the reporter fusion with monoclonal antibody HA.11. To preserve the native expression elements controlling nrfA, the reporter was designed to replace the native nrfA by homologous recombination at its endogenous locus. A markerless allelic exchange procedure (see Materials and Methods) was used to introduce this construct into H. influenzae Rd to create strain RdHA.
This reporter strain was used to assess whether FNR is required for expression from the nrfA promoter. RdHA derivatives were generated (Table (Table1)1) that contained the fnr mutation (RHAfnr); the fnr mutation and “empty vector” sequences at the xyl locus (RHAfnrV); or the fnr mutation and a wild-type copy of fnr, provided at the xyl locus (RHAfnrC). These strains were assayed by Western blotting for levels of the fusion protein after growth under low-oxygen conditions predicted to activate FNR (Fig. (Fig.2A).2A). The fusion protein was readily detectable as a band migrating at ~29 kDa in lysates from strains RdHA and RHAfnrC that contain fnr, consistent with the predicted 29.5-kDa molecular mass of the mature fusion protein. As expected, no reactivity with monoclonal antibody HA.11 was detected in the parental strain Rd, which lacks the fusion construct. Lysates from fnr mutant strains RHAfnr and RHAfnrV contained no detectable fusion protein, indicating that FNR is required for expression of the fusion from the nrfA promoter.
The dependence of the Pnrf-hel-HA reporter fusion on FNR for its expression suggested that this system could be used to investigate the aeration conditions influencing FNR activity in H. influenzae. A range of oxygen conditions was generated by varying the volume of culture medium in a series of flasks of the same size. As the culture volume increases, the liquid/air interface decreases, thus yielding lower rates of oxygen supply (21). Genomic transcription-profiling experiments with H. influenzae have shown that this range of conditions modulates the expression of nrfA and other genes predicted to be controlled by FNR (89). The effect of these conditions on oxygen levels was verified by measuring dissolved oxygen levels in cultures at the time of sample harvesting (see Materials and Methods). In parallel, fusion protein levels in cells from the same cultures were assessed by Western blot analysis. Increased expression of Pnrf-hel-HA was observed in samples with decreased oxygen levels (Fig. 2B and C) Although the intracellular oxygen concentration sensed by FNR is related to a complex function of the oxygen consumption rate and other factors not measured here, the results demonstrate that activation of the nrfA promoter occurs under low-oxygen conditions but does not require complete anaerobiosis. Together, these results indicate that FNR of H. influenzae increases nrfA expression in response to conditions of decreased culture aeration.
In other bacteria, FNR has been implicated in both negative and positive control of genes for nitrosative stress defense. To investigate the potential role of FNR in NO resistance by H. influenzae, we evaluated the sensitivity of the fnr mutant to two NO donors, GSNO and nitrite. GSNO acts by slow release of NO, causing bacterial damage through protein transnitrosation, a covalent transfer of a NO group to free thiol groups (77). Under acidic conditions, nitrite is a NO donor, commonly called ASN (73). Strains Rd, RfnrV, and RfnrC were compared for survival after challenge with GSNO under anaerobic conditions. Recovery of CFU of the fnr mutant strain RfnrV was decreased by 600-fold relative to the rate of recovery of parental strain Rd, and complementation with fnr in strain RfnrC restored resistance to wild-type levels (Fig. (Fig.3A).3A). Treatment of cultures with GSNO in the presence of oxygen resulted in no measurable growth inhibition for all strains (data not shown), consistent with the instability of NO in the presence of oxygen, particularly in biological systems (34). Exposure to ASN yielded results with the fnr mutant RfnrV that were comparable to those obtained with GSNO, exhibiting a 40-fold decrease in recoverable CFU compared to the number of CFU recovered for the parental strain, Rd; complementation with the wild-type fnr in strain RfnrC restored resistance to parental levels (Fig. (Fig.3B3B).
ASN releases NO rapidly, providing the opportunity to monitor decreases in CFU over a relatively short period of time (73). Cultures of strains Rd, RfnrV, and RfnrC were incubated in acidified medium under oxygen-depleted conditions in the presence or absence of ASN and monitored for recoverable CFU for 120 min. The cultures incubated in this acidified medium in the absence of ASN exhibited negligible growth or loss of viability over the course of the assay (data not shown), indicating that decreases in recoverable CFU when ASN is present are not a direct result of pH sensitivity. In contrast, in the presence of ASN, the fnr mutant (RfnrV) exhibited a dramatic decrease in CFU by 60 min (Fig. (Fig.3C),3C), with CFU dropping by 161- and 81-fold relative to CFU of the parental and complemented strains, respectively, which exhibited only 1.4- and 2.5-fold decreases in survival at this time point relative to their starting CFU. Rd and RfnrC were affected by ASN at later times, exhibiting ~100-fold reductions relative to their starting CFU at 120 min, yet the fnr mutant remained significantly more sensitive than parental and complemented strains, by 69- and 28-fold, respectively (Fig. (Fig.3C).3C). As was observed with GSNO, cultures exposed to ASN under oxygen-rich conditions exhibited no appreciable decreases in CFU (data not shown), consistent with the rapid oxidation of NO to nitrite expected to occur under these conditions (87). Together, these results indicate that FNR is required to promote resistance of H. influenzae to killing by NO donors under low-oxygen conditions.
To identify mechanisms by which FNR could participate in defense against RNS, we examined FNR-mediated transcriptional control of a selected set of genes with potential FNR binding sites in their putative promoters (Fig. (Fig.4A).4A). Included in the analysis were genes with potential roles in NO resistance and, for comparison, several genes predicted to be members of the FNR regulon based on their regulation in other bacteria. Based on results obtained with the Pnrf-hel-HA reporter fusion in nitrite reductase assays and Western blotting (Fig. (Fig.11 and and2),2), FNR was expected to positively control transcription of the nrfA gene. The dmsA operon encodes a putative homolog of dimethyl sulfoxide reductase, which is also positively regulated by FNR in E. coli (15) and was included to further evaluate similarities between the FNR regulon of H. influenzae and those of other bacteria. The predicted H. influenzae napA gene product has 74% amino acid identity to the product of the FNR-regulated napA that is required for nitrate reductase activity in E. coli (80, 81), and a transposon insertion in H. influenzae napA abrogates nitrate reductase activity (J. C. Harrington, unpublished results). The ytfE of E. coli encodes a di-iron protein that repairs nitrosative damage (42). HI1677, the putative homolog of ytfE in H. influenzae, encodes a predicted protein with 57% amino acid identity to E. coli YtfE. The presence of a potential FNR binding site in the promoter region of ytfE in H. influenzae suggested that it could play a role in FNR-mediated RNS resistance.
To evaluate the potential regulation of these genes by FNR, total RNA was obtained from cultures of Rd, RfnrV, and RfnrC grown under low-oxygen conditions, and RT-qPCR was used to measure the transcript levels of each gene, normalized to the transcript level of the housekeeping gene rpoA. The transcript abundance of each of the candidate FNR-regulated genes was decreased in the fnr mutant and restored to wild-type levels in the complemented strain (Fig. (Fig.4B).4B). Specifically, a 70-fold decrease in nrfA mRNA levels was observed in the fnr mutant relative to the level in the parent strain, consistent with the pronounced effect of the fnr mutation on the expression of the Pnrf-hel-HA reporter fusion (Fig. (Fig.2B).2B). The dmsA transcript levels were similarly affected, exhibiting a decrease of 23-fold in the fnr mutant. The napA gene was moderately influenced, and its expression was decreased by threefold in the fnr mutant. This result with napA is consistent with a partial decrease in nitrate reductase activity in the fnr mutant (data not shown). The mRNA levels of ytfE decreased by 8.8-fold in the fnr mutant relative to the level in the wild type. Therefore, unlike E. coli ytfE, which is negatively regulated by FNR (41), ytfE is positively controlled by FNR in H. influenzae.
FNR-dependent transcription of nrfA and ytfE indicated a potential role for these genes in FNR-mediated resistance to RNS in H. influenzae. The nrfA mutant strain, RdHA, and strain RytfeV containing an in-frame, nonpolar deletion of ytfE, were compared to NrfA+ YtfE+ parental strains for resistance to RNS generated by ASN or GSNO. Recovery of CFU of the nrfA mutant strain relative to that of wild-type Rd was decreased by 5.8-fold after exposure to GSNO (Fig. (Fig.5A)5A) and 4.2-fold after 90 min of exposure to ASN (Fig. (Fig.5B).5B). The ytfE deletion mutant, RytfeV, exhibited a 25-fold decrease in CFU relative to its parental strain, RV, after exposure to GSNO, and complementation restored its resistance to parental levels (Fig. (Fig.5A).5A). Similarly, after 60 min of exposure to ASN, 25-fold fewer CFU were recovered from cultures of RytfeV relative to the amount recovered for the wild-type strain, RV, and complementation restored resistance (Fig. (Fig.5B).5B). These results indicate that the nrfA and ytfE deletion mutants exhibit hypersensitivity to RNS exposure. The sensitivity of the nrfA mutant, RdHA, to NO donors was less striking than that of RytfeV, indicating that although nrfA appears to contribute to RNS resistance, the requirement for ytfE is greater under these conditions.
H. influenzae is likely to encounter low-oxygen conditions at sites of infection in which macrophages and other cells are sources of RNS. We investigated the potential role of FNR in defense against macrophages by comparing the rates of recovery of bacteria incubated under low-oxygen conditions in the presence and absence of BMM from C57BL/6 mice. The parent strain, RV, and the fnr mutant, RfnrV, both exhibited ~70% survival when incubated with BMM for 30 min, and no significant difference in survival was observed between these two strains under these conditions (Fig. (Fig.6A).6A). The expression of iNOS and production of NO has previously been shown to increase in IFN-γ-stimulated murine macrophages exposed to bacterial LPS (48), an effect that is enhanced under low-oxygen conditions (53, 54). Therefore, the effect on the survival of H. influenzae of prestimulating BMM with IFN-γ or LPS was evaluated. Prestimulation resulted in decreased survival of both strains; however, the fnr mutant, RfnrV, exhibited a greater decrease than the parent, RV, after exposure to either type of stimulated macrophages, and the attenuation of the mutant was statistically significant in both cases (Fig. (Fig.6A).6A). Based on these data, we concluded that fnr is required for optimal survival of H. influenzae after exposure to activated macrophages.
Because our data suggested that the fnr-regulated gene, ytfE, is essential for NO resistance under low-oxygen conditions, we investigated the potential role of ytfE in resistance to killing by macrophages under these conditions. To determine whether increased sensitivity of the ytfE mutant to activated macrophages could involve a NO-dependent mechanism, we pretreated IFN-γ-stimulated and unstimulated BMM with L-NAME, a competitive inhibitor of NOS. The parental strain (RV), the ytfE deletion strain (RytfeV), and the ytfE deletion strain complemented with ytfE at the xyl locus (RytfeC) were evaluated in this assay (Fig. (Fig.6B).6B). Unstimulated BMM reduced the survival of each strain slightly relative to the survival of untreated controls. In contrast, after exposure to IFN-γ-stimulated macrophages, the percent survival of the ytfE mutant (RytfeV) was decreased by a greater extent than that of the parent (RV) or the complemented strain (RytfeC) (Fig. (Fig.6B).6B). Unstimulated BMM treated with or without L-NAME reduced the recovery of all strains slightly and to an approximately equal extent. Treatment of BMM with L-NAME during IFN-γ stimulation eliminated the survival defect of the ytfE mutant relative to the survival of the parent and complemented strains in this assay. Together, these results indicate that the susceptibility of H. influenzae to IFN-γ-stimulated macrophages involves a mechanism dependent upon the ability of macrophages to produce NO.
Results with H. influenzae Rd-derived strains implicated ytfE in the defense of H. influenzae from activated macrophages. To investigate whether this mechanism is conserved in a pathogenic NTHi strain, the ytfE mutation was moved into a clinical NTHi strain, NT127. The resulting mutant, NytfeV, was tested for susceptibility to macrophages that had been preincubated with or without IFN-γ (Fig. (Fig.7).7). Unlike that of Rd-derived strains, the survival of the NTHi parental (NTV) and complemented (NytfeC) strains was unaffected by exposure to macrophages with or without prior stimulation, suggesting that this clinical isolate may evade killing by means of additional mechanisms that are absent in Rd. Nevertheless, the ytfE mutant, NytfeV, exhibited a 27% decrease in survival compared to that of the parent and a 20% decrease relative to that of the complemented strains when exposed to unstimulated BMM. Moreover, in the presence of BMM that had been stimulated with IFN-γ, the survival of the ytfE mutant was decreased by 64% relative to that of the parent and 57% compared to that of the complemented strain. Treatment of BMM with L-NAME concurrently with IFN-γ stimulation completely eliminated the survival defect of the ytfE mutant that was seen with BMM in the absence of inhibitor, implicating ytfE in defense against BMM-derived NO. The marked survival defect of the ytfE mutant relative to the survival of the parental and complemented strains after exposure to macrophages highlights the importance of this gene in the ability of NTHi to evade a major antimicrobial defense mechanism of innate immunity.
The concentration of oxygen present within the microenvironment dramatically affects the reactivity of NO by influencing its stability and the range of additional RNS generated. NO is oxidized to nitrite more rapidly as oxygen levels increase, and therefore, oxygen is considered to be one significant factor in limiting the half-life of NO in biological systems (12, 87). Bacteria growing in high- versus low-oxygen conditions exhibit different expression profiles of RNS-detoxifying enzymes, likely as an adaptation to coordinate the production of appropriate RNS detoxification mechanisms with other metabolic pathways active under these environmental conditions (14, 67, 68). Furthermore, inappropriate expression of enzymes involved in RNS defense can be detrimental to bacterial survival. For example, under aerobic conditions, E. coli and S. enterica serovar Typhimurium (56, 79) detoxify RNS via hmp, which catalyzes oxygen-dependent conversion of NO to nitrate and is repressed by FNR under low-oxygen conditions (17, 28, 56, 57, 79, 94). Deletion of hmp reduces the virulence of S. enterica serovar Typhimurium in mice; however, constitutive expression of hmp can lead to microbial-derived oxidative stress (6), implying that tight regulation of genes involved in nitrosative stress defense is essential for bacteria to withstand challenge from host immunity. Therefore, bacteria must tailor their defense strategies against these reactive species according to the conditions in which they are encountered.
In this study, we investigated the role of H. influenzae FNR in the regulation of resistance to RNS. FNR in other bacterial species plays complex roles in RNS defense. In addition to its well-characterized function in oxygen sensing, FNR of E. coli was also shown to respond directly to NO (68). In Neisseria meningitidis, the FNR response to NO mediates part of a homeostatic regulatory circuit by decreasing the expression of the FNR-activated nitrite reductase AniA (36), which is capable of generating a high level of endogenous NO (71). A denitrification pathway resembling that of N. meningitidis is not present in H. influenzae; however, the nrfA gene in H. influenzae is similar at the predicted amino acid level to the cytochrome c nitrite reductase nrfA of E. coli. Unlike AniA, nrfA has been implicated in NO detoxification (66, 85). We found that the H. influenzae fnr gene is required for appreciable levels of nitrite reductase activity (Fig. (Fig.1B),1B), suggesting positive control by FNR of the nrfABCD operon, predicted to encode the sole nitrite reductase in this species. A nonpolar in-frame deletion of the nrfA gene abrogated the FNR-dependent dissimilatory nitrite reduction by H. influenzae, and complementation restored activity (Fig. (Fig.1A).1A). Studies with a reporter fused to the nrfA gene confirmed FNR-dependent activation of nrfA expression and demonstrated nrfA expression under a range of low-oxygen conditions, consistent with the predicted role of FNR in anaerobic transcriptional regulation (Fig. (Fig.2).2). Under low-oxygen conditions that promote fnr activity, as inferred by its ability to positively control nrfA expression, the fnr mutant was hypersusceptible to NO donors GSNO and ASN, implicating genes of the FNR regulon in RNS defense (Fig. (Fig.3).3). Consistent with this hypothesis, nrfA was also required for resistance to NO donors (Fig. (Fig.5).5). The sensitivity of the nrfA mutant was not as pronounced as that of the fnr mutant, suggesting that additional fnr-regulated genes play a role in RNS defense.
A previous study using a bioinformatic approach to identify potential FNR-regulated genes in H. influenzae identified potential FNR binding sites in the promoter for ytfE (83). YtfE is a member of a broadly conserved family of di-iron proteins that participate in the biogenesis and repair of iron-sulfur centers (41, 42, 63). Our transcriptional analysis revealed FNR-dependent positive control over transcript levels of ytfE in H. influenzae (Fig. (Fig.4B).4B). Because ytfE participates in RNS resistance mechanisms in other bacteria, we examined its role in the resistance of H. influenzae to chemical NO donors GSNO and ASN. Deletion of ytfE led to marked attenuation for survival after exposure to these NO donors, and complementation with ytfE, provided in single copy at an ectopic chromosomal site, restored resistance (Fig. (Fig.5).5). Therefore, ytfE of H. influenzae is positively controlled by FNR and mediates resistance to donors of RNS, implicating the activation of ytfE expression as a primary mechanism of the observed fnr-dependent RNS resistance in H. influenzae.
These results indicate a different mode of control in H. influenzae than in a diverse set of pathogenic bacteria in which ytfE regulation has been examined. In E. coli, ytfE is negatively controlled by FNR and by an additional NO-sensitive DNA binding protein, NsrR, which is absent in H. influenzae, based on genome sequence analysis (9, 27, 41). The NsrR protein of E. coli has been implicated as a direct repressor (9, 22), whereas the absence of a recognizable FNR binding site in the E. coli ytfE promoter has suggested that the effect of FNR may be indirect (41). In the presence of NO, both negative regulators become inactive, leading to ytfE expression. Much of this regulatory circuitry and expression pattern appears to be conserved in bacterial species as diverse as Staphylococcus aureus, S. enterica serovar Typhimurium, and N. gonorrhoeae, each of which contains genes with functional and sequence similarity to nsrR, fnr, and ytfE (63). Moreover, the N. gonorrhoeae NsrR has been demonstrated to directly repress target promoters in a NO-sensitive manner (38). In contrast, H. influenzae lacks a gene with any significant similarity to nsrR, and FNR positively controls ytfE, potentially via interaction with the possible FNR binding site located in its promoter. The apparent conservation of the regulatory pathway controlling ytfE in phylogenetically diverse bacteria but not in H. influenzae may reflect unique aspects of the regulation of RNS defense pathways in H. influenzae pathogenesis. One possibility is that H. influenzae primarily encounters RNS in a relatively low-oxygen environment that it likely experiences during invasion of the epithelium or in biofilms that appear to form at the mucosal surface during infection (23). Entry into such stages of colonization could signal the requirement for induction of preemptive defenses against RNS generated when host cells respond to increased concentrations of bacterial products. In this model, an nsrR-mediated response to NO would not be required. Alternatively, the configuration of multiple systems to metabolize toxic reactive oxygen species and RNS in different bacteria may constrain the allowable expression patterns for certain enzymes, as exemplified by the toxic effect of ectopic expression of hmp in Salmonella enterica serovar Typhimurium (6). H. influenzae lacks many enzymes of RNS defense that are present in other bacteria and, therefore, may not require the same degree of complex control to prevent such aberrant interactions.
Control of H. influenzae by phagocytic cells has been shown to be critical during early phases of infection and during adaptive immunity (4, 61, 82, 84, 88). To investigate the potential role of fnr regulation in defense against host cells capable of RNS generation during the immune response, we evaluated the ability of H. influenzae to resist the antibacterial effects of BMM with or without prior stimulation with IFN-γ or LPS, which trigger iNOS production (49). The fnr mutant was attenuated for survival in the presence of activated macrophages (Fig. (Fig.6A).6A). Resistance also required ytfE, both in the laboratory strain Rd and in a clinical NTHi background. Complementation of the ytfE mutants or inhibition of macrophages with L-NAME restored the mutants to wild-type survival levels (Fig. (Fig.6B6B and and7).7). It has been shown that NO can inhibit the growth of E. coli under fully anaerobic conditions by damaging the iron-sulfur centers of branched-chain amino acid synthesis enzymes (70). Such bacteriostatic effects could contribute to the decreased net survival of H. influenzae mutants that we observed in our assays. Nevertheless, it appears likely that RNS also kills H. influenzae under the conditions we used. Experiments measuring sensitivity to ASN during a short time course of exposure revealed a decrease in CFU (Fig. (Fig.3C),3C), whereas control cultures of H. influenzae did not multiply in the same acidified medium in the absence of nitrite during this period, indicating that ASN exerted a bactericidal effect. In either case, a destructive effect of RNS on iron-sulfur centers in H. influenzae would be consistent with the role in RNS resistance that we have detected with mutants deficient in ytfE, whose product is a protein involved in repairing iron-sulfur centers. Therefore, the data indicate that FNR and the ytfE gene are probably required to resist inhibitory or antimicrobial effects of RNS produced by the immune response during infection.
The role of RNS resistance in H. influenzae pathogenesis is at the early stages of investigation. Because NO-producing cells, including macrophages and epithelial cells, are abundant in the human nasopharynx and high levels of NO are present in this primary site of asymptomatic colonization (50), it is likely that resistance to RNS is an important adaptation for this organism. Neisseria meningitidis, another nasopharyngeal colonizer, has been shown to require RNS defenses for viability during coincubation of bacteria with nasopharyngeal explants (78), implicating defense against RNS as an important factor in colonization of human respiratory mucosa. Disease states also expose H. influenzae to RNS derived from phagocytes and other cell types. Macrophages are abundant in the lung, and bacterial infection, implicated in exacerbations of chronic obstructive pulmonary disease, is associated with elevated numbers of neutrophils (74). Otitis media pathology involves an influx of polymorphonuclear leukocytes during acute infection and elevated numbers of macrophages during chronic disease (93), and human middle ear effusions contain NO metabolites, including nitrate and nitrite (40). Therefore, it will be informative to exploit knowledge of RNS resistance pathways in H. influenzae with models such as mucosal tissue explant systems, infections of mice deficient in RNS production, or otitis media models of middle ear infection. Our investigation of FNR-mediated environmental control of defense against RNS and phagocytic killing of H. influenzae suggests that this regulation is likely required for the survival of H. influenzae in an in vivo environment in which oxygen levels are low, conditions that may occur when bacterial density increases on mucosal surfaces or during invasion into submucosal sites. Identifying the specific stages of colonization or disease that involve these pathways will be important for understanding how this pathogen evades host immunity.
This work was supported by a grant from the American Heart Association and by NIH NIAID grant number 1RO1-AI49437 to B.J.A.
We thank Jeffrey Gawronski for helpful comments.
Editor: J. B. Bliska
Published ahead of print on 16 March 2009.