E. coli strains (Supplementary Table S1) were grown on liquid or solid Luria−Bertani (LB) medium at 37°C for 16–18 h with shaking at 200 rpm for liquid media. Haemophilus influenzae (Hi) 2019 (Campagnari et al., 1987) (NCBI acc. no.: NZ_CP008740.1), NTHi 86-028NP (Bakaletz et al., 1989) (NCBI acc. no.: NC_007146.2) and derivative strains (Supplementary Table S1) were grown on Brain Heart Infusion broth (BHI, BBL) or agar supplemented with hemin (10 μg/mL) and NAD (10 μg/mL) at 37°C for 16 h and 200 rpm shaking for broth cultures. Chemically Defined Medium (CDM) was also used and contained 10 mM glucose, 1 mM sodium pyruvate, 0.08 mg/mL uracil, 0.17 mg/mL inosine, 10 μg/mL β-NAD, 25 mM HEPES, pH 7.4, 10 μg/mL hemin and 2 mg/mL NaHCO₃ in RPMI1640 (Coleman et al., 2003). The sBHI and CDM media were also used to cultivate H. influenzae clinical isolate strains from our collection. Actinobacillus pleuropneumoniae (strain 4074) (Supplementary Table S1) was cultivated on CDM medium under microaerobic conditions at 37°C and 45°C, respectively. Ampicillin (100 μg/mL E. coli), kanamycin (100 μg/mL E. coli; 10 μg/mL Hi) and tetracycline (10 μg/mL E. coli, 1 μg/mL Hi) were added to culture media when needed.

NTHi growth experiments (Hosmer et al., 2022) were conducted in microtiter plates, each well containing 200 μL of CDM medium. Three biological replicates per strain were incubated at 37°C and 200 rpm using a Clariostar multimode plate reader (BMG LabTech). The atmospheric control unit was set up to allow approximately 20% O₂ for aerobic conditions, 2.8% O₂ for microaerobic conditions, and 5% CO₂ with no oxygen for anaerobic conditions (no shaking). Growth rates were calculated according to the method described in Kurokawa and Ying (2017).

NTHi transformation was performed as in Poje and Redfield (2003). Briefly, H. influenzae cultures (20 mL) were grown to an OD_600nm of 0.25 using sBHI, harvested, and washed twice in freshly prepared MIV solution (Poje and Redfield, 2003) before resuspension in 5 mL MIV and incubation at 37°C with shaking for 100 min to develop competence. One microgram of linearized plasmid was added to 1 mL of competent cells and incubated at 37°C for 30 min before 1 mL of sBHI was added. Samples were incubated at 37°C for 1 h before plating on sBHI plates with 20 μg/mL kanamycin. MIV solution was prepared by combining 25 mL solution 21 (per liter: 4 g L-aspartate, 0.2 g L-glutamate, 1 g fumarate, 4.7 g NaCl, 0.87 g K₂HPO₄, 0.76 g KH₂PO₄, 0.2 mL Tween 80, pH 7.0) with 0.25 mL solution 22 (4 mg L-cytosine, 10 mg L-tyrosine dissolved in 1 mL 1 M HCl at 37°C followed by addition of 9 mL H₂O and 6 mg L-citrulline, 20 mg L-phenylalanine, 30 mg L-serine, 20 mg L-alanine), 0.25 mL 0.1 M CaCl₂, 0.25 mL 0.05 M MgCl₂, and 0.25 mL 5% Difco vitamin-free Casamino acids.

Biofilm formation in 96-well microtiter plates used the protocol described in Schembri and Klemm (2001), and bacterial survival in biofilms was determined as in Dhouib et al. (2016) using proteinase K digestion and serial dilution. Briefly, non-typeable H. influenzae strains were grown to OD_600nm of 0.2–0.3 under microaerobic conditions in sBHI broth at 37°C with shaking. Then, the cultures were diluted to an OD_600nm of 0.05 and dispensed into 96-well microtiter plates (100 μL per well, 96-well plate: Techno Plas, cat no: SMPSL) and incubated at 37°C. Following incubation, planktonic cells were removed by washing the wells twice with sterile water, and the wells were subsequently stained using 0.1% crystal violet for 10 min. After plate drying, 30% acetic acid was applied to dissolve the biofilm, and the optical density at 550 nm (OD_550nm) was measured using a plate reader. To determine viable cell counts, the wells of the biofilm plate were rinsed twice with sterile water, followed by incubation with 100 μL of 0.1 mg/mL protease K for 10 min at room temperature to dissolve the biofilm. Subsequently, samples were collected, serially diluted in 1xPhosphate Buffered Saline (PBS), and plated on sBHI agar to quantify the colony forming units (CFU) per well. Each strain was analyzed with three biological replicates, and several technical replicates were used for each biological replicate.

Bactericidal assays were conducted following the protocol described in Nasreen et al. (2020) and used 150 and 175 μM HOCl, 0.5–1.5 mM NCT. In all experiments, HOCl was applied as NaOCl. H. influenzae 2019 strains, obtained from freshly grown sBHI plates, were resuspended to an OD_600nm of 1.0 (∼6 × 10⁸ CFU/mL) in 1xPBS. Subsequently, 900 μL of bacterial culture were combined with 100 μL of a freshly prepared 10x stock of the test compound or sterile water as a control. The reactions were incubated at room temperature for 60 min with orbital shaking, followed by immediate serial dilution and plating on sBHI plates for CFU/mL determination.

For S-oxide reductase activity assays in H. influenzae strains 2019 and 86-028NP, cultures growing microaerobically in CDM were treated with 0.4 mM HOCl or 10 mM paraquat for 60 min at mid-exponential phase before harvesting.

Nitrosative stress susceptibility was assessed following the method of (27). NTHi strains, with three biological replicates grown on sBHI agar plates, were inoculated in 50 mL sBHI broth in 50 mL tubes and incubated at 37°C in a CO₂ incubator for 16–18 h. These cultures were then used to inoculate sBHI broth buffered to pH 5.5 at an initial OD_{600 nm} of 0.07. 150 μL of this bacterial suspension were added to each well of a round-bottom 96-well microtiter plate (Costar 3799), followed by addition of 150 μL of 20 mM or 30 mM NaNO₂ to each reaction to achieve final concentrations of 10 mM and 15 mM. Sterile water was added to control wells. Plates were incubated at 37°C, 5% CO₂, for 16 h in an anaerobic jar using Anaerocult A catalysts (Merck) to remove oxygen. After 2 h of incubation, the cultures were serially diluted and plated on sBHI plates for CFU/mL determination. Growth rate determinations used the program GrowthRates v.6.2.1 (Hall et al., 2013).

Standard methods were used throughout (Ausubel et al., 2005). All chemicals were purchased in analytical grade or equivalent. Genomic DNA was isolated using DNAzol reagent (Thermo Fisher Scientific). GoTaq Green MasterMix (Promega) was used for general PCR (primer sequences, Supplementary Table S2), the GeneJET Plasmid Miniprep Kit and GeneJET PCR Purification kit (both Thermo Fisher Scientific) for plasmid and PCR product purification. Restriction enzymes were from Thermo Fisher Scientific, T4 ligase (NEB) was used for ligations. Cell-free extracts of NTHi were generated using BugBuster Mastermix (Novagen) according to the manufacturer’s instructions (750 μL were used for pellets from 50 mL culture). Protein concentrations were determined using the BCA-1 kit (Sigma Aldrich).

The previously constructed plasmid, pGEM-HidmsA::kan (Dhouib et al., 2021) was linearized using NcoI and transformed into competent NTHi 86-028NP using the method described above.

The tet resistance cassette, amplified from pRK415 using primers from Supplementary Table S2, was ligated into two previously constructed plasmids: pGEM-HimtsZ (Dhouib et al., 2016) and pBlu-HimsrAB (Nasreen et al., 2020). This resulted in the construction of pGEM-HimtsZ::tet and pBlu-HimsrAB::tet plasmids. The linearized pGEM-HimtsZ::tet plasmid was transformed into Hi2019^ΔdmsA and Hi2019^ΔmsrAB to generate Hi2019^ΔdmsAΔmtsZ and Hi2019^{ΔmsrABΔmtsZ}. Additionally, linearized pBlu-HimsrAB::tet was used to create double mutant Hi2019^{ΔdmsAΔmsrAB}.

Samples for RNA isolation from bacteria were obtained during the mid-exponential growth phase (OD_600nm Hi, Ap ∼ 0.45) from cultures growing on CDM under microaerobic conditions. Additionally, RNA samples were collected at 30, 60, and 120 min after treating cultures with 200 μM HOCl, 150 μM H₂O₂, or 5 mM paraquat, following the method outlined in reference (Pauwels et al., 2004).

For each sample, 2 mL of culture was preserved using 2 mL of RNAprotect Bacteria Reagent (Qiagen) before RNA isolation using the Illustra RNAspin Mini RNA isolation kit (Cytiva). RNA from snap-frozen mouse lung tissue was isolated by homogenization in 1 mL Trizol (Thermo Fisher Scientific), followed by centrifugation to remove large cellular debris and RNA isolation according to the manufacturer’s instructions. Genomic DNA was removed using the Turbo DNAfree kit (Thermo Fisher Scientific), and RNA concentrations were determined using the Quant-it RNA HS kit (Thermo Fisher Scientific). The absence of genomic DNA was confirmed by PCR using 16S primers and β-actin (ACTB, mouse) primers (Supplementary Table S2), with purified RNA serving as the template. Subsequently, cDNA was synthesized from 500 ng of genomic DNA-free RNA using random hexamer primers and either the Superscript IV VILO Master Mix (Thermo Fisher Scientific) or the Lunascript RT SuperMix kit (New England Biolabs), following the manufacturer’s instructions.

Quantitative reverse transcription-PCR (qRT-PCR) was conducted following the procedures outlined in reference (Othman et al., 2014). Each qRT-PCR reaction (10 μL) comprised diluted cDNA (1:100) as the template, QuantiNova SYBR Green qPCR Master Mix from Qiagen, and specific oligonucleotide primers (Conc 2 μM) listed in Supplementary Table S2. Reference genes used included gyrase (gyrA) genes for H. influenzae, Actinobacillus pleuropneumoniae, and β-actin (ACTB) for mouse cDNA. Data analysis and normalization were performed as in Kappler et al. (2005). The cycle threshold (CT) value for all samples was determined using QuantstudioTM Real-time PCR software version 1.3 (Thermo Fisher Scientific). The PCR efficiencies were determined using LinRegPCR software version 2016.0 (Ruijter et al., 2009).

S- and N-oxide reductase activity assays were performed anaerobically in 20 mM sodium phosphate buffer (pH 6.8) containing 0.2 mM Methyl viologen, 0.3 mM sodium dithionite and one of the following terminal electron acceptors: 10 mM Dimethyl Sulfoxide (DMSO), 10 mM L-Methionine Sulfoxide (L-MetSO), 10 mM S-Biotin Sulfoxide (S-BSO), 10 mM racemic methyl p-tolyl sulfoxide (MPTS), 10 mM R-MPTS & S-MPTS, 2 mM Pyrimidine N-Oxide (Pyr N-Ox), and 5 mM Nicotinamide N-Oxide (Nic N-Ox). The re-oxidation of reduced methyl viologen [ɛ₆₀₀ = 13.7 mM⁻¹ cm⁻¹ (Watanabe and Honda, 1982)] was monitored at 37°C in a Cary60 Spectrophotometer (Agilent Technologies) as described in Dickie and Weiner (1979). For determination of kinetic parameters, the concentrations of the substrates were varied. Specific enzyme activities are given as μmoles of substrate reduced per min (U) and mg of protein present. Kinetic data were fitted using Prism 9 (GraphPad). For enzyme assays using purified H. influenzae MtsZ, the protein was prepared as described in Dhouib et al. (2016) and Struwe et al. (2021).

The Human Bronchial Epithelial 16HBE14 cell line was cultured using Minimal Essential Medium (MEM) supplemented with 10% heat-inactivated Fetal Bovine Serum (FBS). Adherence and invasion assays for NTHi were conducted following the procedures outlined in Dhouib et al. (2015). Normal Human Nasal Epithelia (NHNE) were prepared following the protocol outlined in Yeo et al. (2019) using primary nasal cells obtained from healthy donors at the Child Health Research Centre, UQ, Brisbane. Seven days before infection, NHNE were transferred to steroid- and antibiotic-free PneumaCult-ALI medium (Stemcell). NTHi for infection were resuspended in 1xPBS and applied at a Multiplicity of Infection (MOI) of 10:1. Uninfected NHNE were exposed to an equal amount of sterile 1xPBS. Following a 24-h infection period at 37°C with 5% CO₂, the inoculum was removed by washing with 1xPBS, and the basal medium was replaced.

Post-infection, NHNE apical surfaces were washed every second day using 100 μL 1xPBS, and the basal medium was replaced. Lysates were generated by a 10 min incubation of NHNE with 1% saponin at 37°C. Intracellular bacterial Colony Forming Units (CFU) were determined after treating the NHNE surface with gentamycin (100 μg/mL) for 1 h at 37°C. Gentamycin was then aspirated, and the apical compartments were washed five times with 200 μL of 1xPBS before lysing the NHNE with a 10 min incubation at 37°C with 1% saponin. Bacterial loads in lysates were determined by plating serial dilutions on sBHI agar.

NHNE transepithelial electrical resistance (TEER) was measured every 2 days using a Millicell ERS-2 Volt-ohm meter. For competition assays, NHNE were infected with an equal mixture of Hi2019^WT and one mutant strain (total MOI 10:1), and bacterial loads were analyzed as described above. However, serial dilutions were plated on sBHI to determine total bacterial numbers and on sBHI with 20 μg/mL kanamycin to assess the load of NTHi carrying a mutation. Hi2019^WT cell numbers were determined by subtracting CFU/mL of kanamycin-resistant (mutant) NTHi from total NTHi CFU/mL for the same time point. Each infection and co-infection involved two biological replicates for each strain or strain combination, with three technical replicates used for CFU/mL determination in each biological replicate.

The procedure for isolating Bone Marrow Macrophages (BMM) from C57 Black mice was adapted from Tushinski et al. (1982). In summary, mice were sacrificed, and femurs and tibias were collected, with muscles and surrounding tissues removed. The tips of the femurs and tibias were excised using surgical scissors, and the bone cavity was flushed with complete macrophage medium [RPMI1640 containing 10% heat-inactivated FBS 2 mM L-glutamine (GlutaMAX, Thermo Fisher Scientific) and 150 ng/mL recombinant colony-stimulating factor-1 (rCSF-1, produced at Protein Expression Facility, The University of Queensland)].

Bone marrow cells were washed twice with complete medium (centrifuged at 300 × g for 5 min each time) before being evenly distributed into eight sterile, 100 mm square petri dishes, each containing 15 mL of complete medium. Penicillin/streptomycin (Stock: Penicillin: 10,000 Units/mL, Streptomycin: 10,000 mg/mL) was added to the media to a final concentration of 1%, and the plates were incubated for 6 days at 37°C with 5% CO₂. On day 3, fresh rCSF-1 (final conc.: 150 ng/mL) was added. On day 6, the growth medium was discarded, and BMMs were washed off the plate using 10 mL of 1xPBS. BMMs were then centrifuged at 300 × g for 5 min and resuspended in antibiotic-free BMM medium. Subsequently, 1 × 10⁵ cells/well were seeded into 24-well tissue culture plates for the infection assay.

The infection assays used a MOI of 1,000:1. After 1 h, the inoculum was removed, and complete macrophage medium with 7 μg/mL polymyxin B was added to the wells for 60 min to eliminate extracellular bacteria. For 2 h infections, cell layers were washed twice with RPMI before 100 μL of 0.1% saponin was used to lyse the macrophages. Intracellular bacteria were quantified through dilution plating. For 4 and 6 h infections, the medium containing 7 μg/mL polymyxin B was replaced with medium containing 0.5 μg/mL polymyxin B for 2 h and 4 h before processing the cells to determine intracellular bacteria.

For NTHi pulmonary infection, the mouse model of lung infection we described previously was used (Morey et al., 2013; Dhouib et al., 2016). Hi2019 and Hi2019 ^ΔdmsA were cultured on sBHI plates for 16 h at 37°C with 5% CO₂. A bacterial suspension in 1xPBS, containing 10⁷ CFUs in 30 μL, was intranasally inoculated into female BALB/c mice aged 5 to 6 weeks. Groups of 5 mice were euthanized and necropsied at 0 h, 6 h, 24 h, 48 h, and 72 h, as described in Hosmer et al. (2022). Lungs were aseptically removed from sacrificed mice, and lung tissue was either snap-frozen in liquid nitrogen for RNA isolation or homogenized in 1 mL 1xPBS. The homogenized samples were serially diluted and plated on sBHI agar to determine CFU/mL. Additionally, bronchoalveolar lavage fluid (BALF) was collected, and CFU/mL were determined. CFUs per lung were calculated as described in Essilfie et al. (2012). Giemsa staining of BALF was performed according to the manufacturer’s instructions (Sigma-Aldrich). Statistical comparisons of mean CFU/lung and BALF were carried out using two-tailed t-tests integrated into the Prism 9 software package.

Mouse BALF was used for ELISA assays (Thermo Fisher Scientific) according to the manufacturer’s instructions to detect the presence of mouse TNFα and IL-6. Data were collected for all mice in each treatment group. The statistical significance of the detected differences among the groups was assessed using Prism 9 (GraphPad).

Statistical analyses were carried out using Prism versions 9 and 10 (GraphPad). Statistical comparisons of mean CFU/lung and BALF following mouse lung infection experiments were carried out using two-tailed t-tests. All other analyses used either 1-Way of 2-Way ANOVA with an appropriate multi-comparison correction. All analyses used a p < 0.05 to determine statistical significance. Details of the analyses are provided in the Figure legends.

Experimental animal procedures were conducted in strict accordance with the recommendations outlined in the QLD Animal Care and Protection Act (2001) and the Australian Code of Practice for the Care and Use of Animals for Scientific Purposes, 8th edition. The protocols were approved by the Animal Care and Ethics Committees of QIMR Berghofer and the University of Queensland (QIMR/050/19).

Human nasal cell samples were obtained from healthy donors at the Child Health Research Centre, UQ, Brisbane, under the approval of the University of Queensland Ethics Committee (Approval #2017000520). Clinical isolates of Haemophilus influenzae were collected from routine pathology samples according to approvals 2021/HE001644, 2014/HE000067.

We have previously reported that H. influenzae DmsABC is required for virulence in a mouse model of infection (Dhouib et al., 2021), and to better understand this process, we investigated whether the presence or absence of DmsABC affected the host response to H. influenzae infection. Using a mouse model of lung infection (Dhouib et al., 2021), the Hi2019^ΔdmsA strain was cleared significantly faster than the wildtype, with first differences becoming apparent at 24 h post-infection (p.i.) (Figure 1A). Compared to the wildtype, the total lung-associated populations of Hi2019^ΔdmsA were reduced 10x, 28x, and 18x times at 24 h, 48 h, and 72 h post-infection (p.i.) (total CFU/lung: p < 0.0001). Attenuation was similar for bacteria detected in bronchoalveolar lavage fluid (BALF) (Supplementary Figure S1A). We then tested the host immune response to infection using qPCR for RNA isolated from lung tissue and ELISA for BALF samples. The faster clearance of Hi2019^ΔdmsA was associated with reduced expression of the pro-inflammatory markers IL6, TNFα and IL1β in mouse lung tissue, with expression levels returning to near baseline by 48 h p.i. In contrast, for infections with the wildtype strain, expression of these genes was still elevated at 72 h p.i. (Figures 1B–D). Similar expression patterns were also observed for Hif1α, TGFβ and the BIRC3 gene that encodes an anti-apoptotic effector and has shown increased expression in human tissue cells infected with wildtype H. influenzae (Nasreen et al., 2020) (Supplementary Figures S1B–D). In mouse BALF, a reduced immune response was also apparent for TNFα, but not IL6 levels (Figure 1F; Supplementary Figure S1E). These data suggested a reduced production of chemo-attractants during infection with Hi2019^ΔdmsA that was accompanied by a reduced influx of immune cells into the lungs of infected mice (Figure 1E; Supplementary Figures S1F–H). The difference was most prominent at 6 h p.i., but persisted up to 72 h p.i. and was primarily due to lower neutrophil numbers. Together with the reduced recovery of Hi2019^ΔdmsA from mouse lungs, this could indicate reduced survival of the Hi2019^ΔdmsA strain in contact with resident lung immune cells.

To investigate this possibility, we tested whether Hi2019^ΔdmsA showed increased susceptibility to killing by bone marrow-derived mouse macrophages. While we had previously demonstrated slightly increased susceptibility of Hi2019^ΔdmsA to neutrophil-mediated killing (Dhouib et al., 2021), in macrophage killing assays, this strain showed a statistically significant increase in intracellular survival compared to the wild type at 6 h p.i. (Supplementary Figure S2). As neutrophil numbers were low in Hi2019^ΔdmsA infection BALF samples, we conclude that the increased clearance of Hi2019^ΔdmsA in the mouse lung was not due to a greater sensitivity to killing by phagocytic cells such as tissue-resident macrophages.

The human respiratory tract is the only known natural niche of H. influenzae (Van Eldere et al., 2014), and as a result, H. influenzae infections in healthy mice are not stable and of short duration (Figure 1A). To determine whether the dmsA mutation also reduces H. influenzae survival in contact with human respiratory epithelia, we infected normal human nasal epithelia (NHNE) with either the Hi2019^WT or the Hi2019^ΔdmsA strain. NHNE are pseudostratified epithelia that are derived from primary human nasal cells (Pezzulo et al., 2011; Hosmer et al., 2022). NHNE are differentiated at the air-liquid interface and closely resemble primary human respiratory epithelia (Pezzulo et al., 2011; Hosmer et al., 2022).

In NHNE single-strain infections, compared to the wildtype, attenuation of Hi2019^ΔdmsA increased from 3.6- and 2.2 -fold for total and intracellular bacteria on day 3 p.i. to 3 × 10³- and 1 × 10⁴- fold, respectively, by day 7 p.i. (Figures 2A,B). Transepithelial migration of Hi2019^ΔdmsA was also reduced, with only ~0.3% of total bacteria recovered for the mutant strain on day 7 p.i., compared to ~1.1% for the wildtype strain (Supplementary Figure S2C).

We then tested the survival of Hi2019^ΔdmsA in competitive infections of NHNE. Using an equal mixture of Hi2019^WT and Hi2019^ΔdmsA, attenuation of the mutant strain was ~100 fold by day 3 p.i., and this increased to ~1,000 fold by day 7 p.i. for both total and intracellular bacterial populations (Figures 2E–G). Neither infection type appeared to cause significant damage to NHNE, as indicated by the consistency of TEER values between infected and uninfected control NHNE (Figure 2D; Supplementary Figure S3). These data confirmed that DmsABC is also required for the survival and virulence of H. influenzae in human nasal epithelia, its primary niche.

Together with our previous investigations that documented the absence of an in vitro phenotype of Hi2019^ΔdmsA (Dhouib et al., 2021), the data above confirm that DmsABC is required under conditions that only exist when H. influenzae is in contact with host cells. This then suggested that host-produced stressors might play a role in controlling expression of dmsABC. To investigate this, we monitored expression of dmsA following exposure of actively growing, microaerobic cultures of Hi2019 to hypochlorite, hydrogen peroxide and or paraquat, a superoxide-producing agent. Hypochlorite caused a 3.3-fold increase in dmsA expression within 30 min of exposure, followed by a slow decrease in gene expression, while hydrogen peroxide had no or very little effect on dmsA expression (Figure 3A). Interestingly, the highest levels of dmsA expression were observed following paraquat exposure; however, induction of gene expression only occurred 60 min post-exposure (Figure 3A).

This is the first time that a specific, fast-acting inducer for dmsA expression other than general anaerobiosis has been identified in any bacterium, and we therefore tested whether hypochlorite induction of dmsA expression also occurs in other H. influenzae strains and other bacteria in which dmsABC genes are present such as Actinobacillus pleuropneumoniae. A. pleuropneumoniae is another bacterial pathogen of the Pasteurellaceae family for which a link between virulence and the presence of DmsABC has been previously reported (Baltes et al., 2003) (Figures 3B,C) Hypochlorite induced dmsA expression in all bacteria tested, but expression patterns and maximal induction levels varied, especially for the three H. influenzae strains. In most cases, increases in dmsA expression were measurable after 30 min, and peak expression occurred between 60 and 90 min post HOCl exposure. The smallest relative change, ~2-fold induction at 60 min, was observed for A. pleuropneumoniae (Figure 3C). The data clearly establish hypochlorite-induction as a feature of dmsABC regulation, suggesting a general role of DmsABC in bacterial stress responses during host-bacteria interactions in H. influenzae and other Pasteurellaceae (Baltes et al., 2003; Dhouib et al., 2021).

As H. influenzae dmsA gene expression levels were very strikingly different between the three strains tested above, we hypothesized that this might correlate with the presence or absence of genes encoding the MtsZ MetSO reductase that is another S-oxide reductase in H. influenzae but is not strictly conserved. Accordingly, we compared S-oxide reductase activity levels in five genotypically dmsABC⁺/ mtsYZ⁺ strains and two dmsABC⁺/ mtsZ⁻ strains from our H. influenzae isolate strain collection. Hi2019 and Hi 86-028NP were used as reference strains for the two groups, respectively, and had 0.11 and 1.8 U/mg S-oxide reductase activity, respectively. Following anaerobic growth, total S-oxide reductase activities between 0.09 and 1.8 U/mg for the DmsABC-only group, and 0.11–3.44 U/mg for the DmsABC/MtsZ positive group of strains were determined (Figure 3D). Together, this indicates that under standard anaerobic conditions, S-oxide reductase activity varied significantly in a strain- but not genotype-dependent manner. It also shows that high S-oxide reductase activity is not a specific trait of recent clinical isolate strains.

To confirm that the observed induction of gene expression in the presence of HOCl and paraquat leads to an increase in S-oxide reductase activity, we then exposed the two reference strains to HOCl or paraquat prior to S-oxide reductase activity determination. This experiment confirmed induction of activity by 1.5- and 3.2-fold for Hi2019 and 86-028NP following HOCl exposure and 3.2- and 14.4-fold, respectively, for paraquat exposure (Figure 3E).

While the dmsABC⁺/ mtsZ⁻-genotype is only present in about 20% of H. influenzae isolate strains, these strains offer a unique opportunity to better understand the physiological role of DmsABC in H. influenzae pathogenesis and we therefore created and characterized a Hi86-028NP ΔdmsA strain.

The mutation was confirmed using PCR, and enzyme assays showed that in crude extracts from the mutant strain, S-oxide reductase activity was reduced by ~80% (Figure 4A). As the Hi2019^ΔdmsA strain lacked a distinct in vitro growth phenotype but showed increased biofilm formation, we then tested biofilm formation in Hi86028NP^ΔdmsA to confirm a similarity of phenotypes. Under microaerobic conditions, the Hi86028NP^ΔdmsA strains showed a 5-fold increase in biofilm formation (1-Way ANOVA, Dunnett, p < 0.0001) and similar viable cell counts to the wildtype (Figures 4B,C). In addition to oxidizing compounds, reactive nitrogen species are also commonly found at sites of infection (Ulfig and Leichert, 2021), and we therefore tested the susceptibility of the Hi2019 and 86-028NP dmsA mutant strains to NO stress, however, both strains behaved the same as the respective wildtype strains (Supplementary Figure S4).

In infection assays using 16HBE14 bronchial epithelial cells, the 86-028NP dmsA mutant strain showed a reduction in total cell-associated bacteria of 1.6- and 7.7-fold relative to the wildtype strain at 4 h and 24 h p.i. (2-Way ANOVA, Sidak, p = 0.0255 & 0.023), while intracellular bacterial loads were reduced 8-fold at 4 h and 100-fold at 24 h p.i. (2-Way ANOVA, Sidak, p = 0.0001 & 0.001) (Figures 4D,E). Planktonic cell numbers showed no statistically significant differences (Figure 4F). These results resemble the phenotype we reported for Hi2019^ΔdmsA (Dhouib et al., 2021), except that for the 86-028NP ΔdmsA strain, the infection phenotype was somewhat more pronounced. In infections of mouse primary macrophages, the 86-028NP ΔdmsA strain showed no statistically significant difference in survival compared to the WT strain, which differs from the behavior of the corresponding Hi2019 derivative strain that showed increased survival at 6 h post-infection (Supplementary Figure S2). The slight differences in infection phenotypes may be due to the absence of the MtsZ methionine sulfoxide reductase that provides some functional redundancy in H. influenzae 2019.

Together, these experiments demonstrate that the phenotype of the H. influenzae 86-028NP ΔdmsA strain is similar to what we previously described for the equivalent mutation in Hi2019 (Dhouib et al., 2021), but in the tissue cell infection model, the H. influenzae 86-028NP ΔdmsA strain showed an increased colonization defect for tissue cell adherent bacteria.

Our data so far indicate that DmsABC is important for full virulence of H. influenzae, with strong phenotypes observed only during infection. This strongly suggests that the substrate that DmsABC converts is only available during contact with host cells. As the enzyme is produced in the presence of hypochlorite and paraquat-mediated stress, this suggests that the natural substrate could be an oxidatively damaged biomolecule. To better understand the catalytic properties of HiDmsABC, we used cell extracts of the MtsZ negative Hi86-028NP wildtype strain that has high S-oxide reductase activity (Figures 3D,E). Initial experiments used the artificial substrate Methyl-p-tolyl-sulfoxide (MPTS) to determine the HiDmsABC stereospecificity. Activity with racemic RS-MPTS was 2.23 U/mg, while R- and S-MPTS resulted in 1.86 U/mg and 0.54 U/mg of activity (Figure 5A). This confirmed that, like its E. coli counterpart (SimalaGrant and Weiner, 1996), HiDmsABC is specific for R-sulfoxides and thus has the opposite stereospecificity to HiMtsZ, which is an S-sulfoxide-specific enzyme (Dhouib et al., 2016). We then tested HiDmsABC activity with different S- and N-oxides that could occur in the human respiratory tract and DMSO as a control substrate. HiDmsABC easily converted L-methionine S/R sulfoxide, nicotinamide-N-oxide, pyrimidine-N-oxide and can also catalyze reduction of adenine-N-1-oxide, while S-Biotin Sulfoxide was not converted and was not investigated further (Figure 5B). As expected, due to the higher reactivity of N-oxides compared to S-oxides, DmsABC activities were highest with nicotinamide-N-oxide and pyrimidine-N-oxide (~4 U/mg), but interestingly the K_M-values for both substrates were in the low micromolar ranges with 32 ± 13 μM, and 76 ± 17 μM, and were, in fact, in a similar range as the K_M-value for L-methionine S/R sulfoxide (35 ± 9 μM) (Figures 5C–E). These catalytic parameters make all three compounds excellent candidates as natural DmsABC substrates. In contrast, adenine-N-1 oxide had a K_M-value of 793 ± 138 μM, making it an unlikely natural substrate for DmsABC. Effective conversion of nicotinamide- and pyrimidine N-oxide by DmsABC has not previously been described but is an important discovery as H. influenzae is an auxotroph for both pyrimidine and NAD, for which nicotinamide is a precursor. With the exception of adenine-N-1-oxide, all K_M-values are not only within a concentration range that is highly likely to be physiologically relevant, but the K_M-value for L-methionine sulfoxide is also comparable to the value of 90 μM that has been previously reported for DmsABC from E. coli (SimalaGrant and Weiner, 1996).

In addition to DmsABC, H. influenzae strains contain two other periplasmic sulfoxide reductases, the MtsZ methionine sulfoxide reductase and the MsrAB peptide methionine sulfoxide reductase (Dhouib et al., 2016; Nasreen et al., 2020, 2022). The three enzymes reduce either protein-bound methionine sulfoxide (MsrAB) or free S-/N-oxides (MtsZ, DmsABC), and their expression is upregulated when the bacteria are exposed to hypochlorite (Dhouib et al., 2016; Nasreen et al., 2020). We hypothesized that together, these three enzymes form a new system for protection of H. influenzae from oxidative stress, and to fully define the roles of these three methionine sulfoxide reductases in this process, we constructed H. influenzae 2019 double mutant strains that each lack two of the three S-oxide reductases.

S-oxide reductase activity assays were used to confirm the contribution of DmsA and MtsZ to the reduction of small-molecule S-oxides in the double mutant strains. As expected, double mutants that lacked MtsZ showed only very low levels of S-oxide reductase activity with both L-methionine sulfoxide and DMSO (Supplementary Figures S5A,B), with lowest activities observed for the ΔdmsAΔmtsZ strain. Unexpectedly, the ΔdmsAΔmsrAB strain in which MtsZ is the only sulfoxide reductase showed about 35% reduced activity, which is about 4x more than expected based on the activity levels (7% of WT) detected in the ΔmsrABΔmtsZ strain and could indicate some synergistic effects.

We then determined growth of the double mutant strains in the presence and absence of oxygen. Growth of all strains was very similar under microaerobic conditions. However, compared to the WT strain, strains carrying a msrAB gene knockout (ΔdmsAΔmsrAB; ΔmsrABΔmtsZ) showed reduced growth under aerobic conditions, and interestingly, under anaerobic conditions, strains carrying a dmsA gene knockout (ΔdmsAΔmsrAB; ΔdmsAΔmtsZ) showed reduced growth (Figures 6A–C; Supplementary Table S3). For the MsrAB sulfoxide reductase, a similar phenotype was already apparent in the single mutant strain. However, the dmsA single mutant strain did not show a reduced growth rate compared to the WT strain (Nasreen et al., 2020; Dhouib et al., 2021).

As all three enzymes have a role in oxidative stress responses, we also determined the sensitivity of the double mutant strains to oxidative stress caused by hydrogen peroxide or the two reactive chlorine species, HOCl or N-Chlorotaurine (NCT). Compared to the WT strain, the double mutants showed a small but statistically significant increase in H₂O₂ sensitivity at high concentrations that are unlikely to be physiologically relevant. However, when the double mutant strains were exposed to HOCl, strains that lacked the msrAB gene were undetectable following exposure to both 150 and 175 μM HOCl, clearly linking the presence of msrAB to H. influenzae HOCl resistance (Figure 6D). We then tested resistance of the strains to another reactive chlorine species, N-Chlorotaurine (NCT), expecting to see a similar, but weaker phenotype as NCT is less reactive than HOCl. Following exposure to NCT, strains lacking the msrAB gene were again most strongly affected, however, reductions in CFU were more pronounced for the ΔmsrABΔmtsZ strain, where a 10⁶-fold decrease in CFU/ml relative to the wildtype strains was observed at concentrations of both 1 mM and 1.5 mM NCT, compared to 10⁴- and 10⁵-fold reductions for ΔdmsAΔmsrAB. Unexpectedly, attenuation was also observed for the ΔdmsAΔmtsZ strain that was unaffected by HOCl exposure but showed a ~ 600- and 10⁴-fold reduction in CFU/ml at 1 mM and 1.5 mM NCT (Figure 6E). These changes in attenuation demonstrate that while MsrAB is the most important enzyme for defense against RCS-induced stress, the two Mo-containing S-oxide reductases, and particularly MtsZ, also contribute to NTHi survival during NCT stress.

Biofilm formation is a typical bacterial response to cellular stress, and in H. influenzae, biofilm formation has been linked to colonization of host cells and tissues. Under microaerobic conditions, two of the three mutant strains showed altered biofilm formation. The ΔdmsAΔmsrAB mutant strain, in which MtsZ is the only sulfoxide reductase, showed a ~ 50% reduction in biofilm formation relative to the wild type. In contrast, in the ΔmsrABΔmtsZ strain, where DmsABC is the only sulfoxide reductase, biofilm formation was increased ~2-fold (Figure 6F). The third mutant strain, ΔdmsAΔmtsZ, showed wildtype levels of biofilm formation.

The link between the presence and absence of these Mo-containing S-oxide reductases and biofilm formation is currently unknown but warrants further investigation. It is intriguing that when present as the sole S-/N-oxide reductases, MtsZ and DmsABC appear to have opposing effects on biofilm formation.

To assess the impact of the double mutants on H. influenzae virulence, we infected 16HBE14 cultured bronchial epithelial cells with either the WT or one of the mutant strains. At 4 h post-infection, the mutant strains showed modest reductions in total cell numbers of 12% (ΔdmsAΔmtsZ), 21% (ΔdmsAΔmsrAB) and 24% (ΔmsrABΔmtsZ) (Figures 6G,H; Supplementary Figure S6). However, clear differences existed for intracellular populations, which were reduced by 60 and 70% for ΔdmsAΔmsrAB and ΔdmsAΔmtsZ, while for the ΔmsrABΔmtsZ mutant, only a modest reduction of intracellular bacteria by 20% compared to the WT was observed. By 24 h, CFUs for all strains had increased, and all mutant strains showed ~40% reduction in total cell numbers compared to the WT. Intracellular cell numbers, however, showed the same trends as at 4 h, with the greatest reductions seen for the dmsAΔmtsZ mutant (30% reduction), while the ΔmsrABΔmtsZ strain was, again, the least affected (20% reduction) (Figures 6G,H; Supplementary Figure S6). Together, the data indicate that MsrAB (ΔdmsAΔmtsZ strain) is not required for invasion of tissue cells but may affect extracellular survival of H. influenzae. In contrast, the two Mo-containing S-/N-oxide reductases are required for intracellular survival. Of the two enzymes, DmsABC had the greatest impact on tissue cell invasion, while MtsZ contributed to invasion but was less critical than DmsABC, as documented by the reduction of bacterial populations in the ΔdmsAΔmsrAB strain.