Biokinetic Characterization and Activities of N2O-Reducing Bacteria in Response to Various Oxygen Levels

Nitrous oxide (N₂O)-reducing bacteria, which reduce N₂O to nitrogen in the absence of oxygen, are phylogenetically spread throughout various taxa and have a potential role as N₂O sinks in the environment. However, research on their physiological traits has been limited. In particular, their activities under microaerophilic and aerobic conditions, which severely inhibit N₂O reduction, remain poorly understood. We used an O₂ and N₂O micro-respirometric system to compare the N₂O reduction kinetics of four strains, i.e., two strains of an Azospira sp., harboring clade II type nosZ, and Pseudomonas stutzeri and Paracoccus denitrificans, harboring clade I type nosZ, in the presence and absence of oxygen. In the absence of oxygen, the highest N₂O-reducing activity, V_m,N2O, was 5.80 ± 1.78 × 10⁻³ pmol/h/cell of Azospira sp. I13, and the highest and lowest half-saturation constants were 34.8 ± 10.2 μM for Pa. denitirificans and 0.866 ± 0.29 μM for Azospira sp. I09. Only Azospira sp. I09 showed N₂O-reducing activity under microaerophilic conditions at oxygen concentrations below 110 μM, although the activity was low (10% of V_m,N2O). This trait is represented by the higher O₂ inhibition coefficient than those of the other strains. The activation rates of N₂O reductase, which describe the resilience of the N₂O reduction activity after O₂ exposure, differ for the two strains of Azospira sp. (0.319 ± 0.028 h⁻¹ for strain I09 and 0.397 ± 0.064 h⁻¹ for strain I13) and Ps. stutzeri (0.200 ± 0.013 h⁻¹), suggesting that Azospira sp. has a potential for rapid recovery of N₂O reduction and tolerance against O₂ inhibition. These physiological characteristics of Azospira sp. can be of promise for mitigation of N₂O emission in industrial applications.

Introduction

Nitrous oxide (N₂O) is an ozone-depleting and greenhouse gas (Ravishankara et al., 2009), therefore it is important to decrease N₂O emissions from natural ecosystems, agriculture, and industrial systems. A large fraction of N₂O is emitted from agricultural croplands (IPCC, 2013; Harter et al., 2014). Emissions from industrial systems, mainly wastewater treatment plants, have become more significant as a result of upgrading of biological nitrogen removal processes, i.e., nitrification–denitrification or partial nitrification–anammox processes (Law et al., 2012). N₂O is produced via multiple biological and abiotic pathways, e.g., in denitrification as an intermediate (Philippot et al., 2011; Wunderlin et al., 2011; Ishii et al., 2014), in the nitrifier denitrification of ammonia-oxidizing microorganisms (Zhu et al., 2013; Ali et al., 2016), and in chemical oxidation of hydroxylamine (Soler-Jofra et al., 2016; Terada et al., 2017). It is consumed mainly by denitrifying bacteria harboring a N₂O reductase system (Nos) (Henry et al., 2006; Zumft and Kroneck, 2006; Jones et al., 2008; Pauleta et al., 2013). In subsequent denitrification steps, N₂O reduction is severely affected by environmental factors, i.e., pH, availability of electron donors, and dissolved oxygen (DO) (Law et al., 2012; Pan et al., 2013). Although the physiological traits on N₂O reduction by canonical denitrifying species, i.e., the genera Pseudomonas and Paracoccus (Zumft, 1997; Vollack and Zumft, 2001; Philippot, 2002; Read-Daily et al., 2016), have been studied to date, knowledge is still limited. Comprehensive and thorough physiological research on N₂O-reducing bacteria under various environmental conditions is therefore warranted.

Recent metagenomic analyses have shown that N₂O-reducing bacteria that harbor Nos can be classified into two clade types: clade I and clade II (Sanford et al., 2012; Jones et al., 2013). It has been reported that the abundances of the two clades potentially depend on environmental conditions, e.g., pH, concentration of calcium ion, and C/N ratio, and niche differentiation probably occurs because of their physiological characteristics (Jones et al., 2014; Domeignoz-Horta et al., 2015; Juhanson et al., 2017). Some clade II type N₂O-reducing bacteria reportedly have lower half-saturation constants for N₂O than do those affiliated to clade I type bacteria, suggesting that a low N₂O concentration favors growth of clade II type N₂O-reducing bacteria (Yoon et al., 2016). Additionally, most non-denitrifying N₂O-reducing bacteria, which are unable to reduce nitrite and nitrate, are clade II type (Sanford et al., 2012; Hallin et al., 2018). Given these traits of clade II type bacteria, reports suggest that they potentially play an important role as N₂O sinks (Jones et al., 2014; Orellana et al., 2014).

Studies of the oxygen effects are of fundamental and engineering importance. The DO concentration determines N₂O emissions from soils and wastewater treatment plants via mainly nitrifier–denitrification and heterotrophic denitrification (Tallec et al., 2006; Morley et al., 2008; Riya et al., 2012). Studies on gene transcription and enzyme expression have suggested that Nos expression is regulated by the DO concentration (Körner and Zumft, 1989; Bergaust et al., 2012) and, more importantly, that the Nos enzyme is inactivated by oxygen (Pauleta et al., 2013). In addition, the susceptibility of N₂O-reducing activity to the oxygen concentration is distinct at species or strain levels, as reported for the genera Thauera and Pseudomonas (Miyahara et al., 2010; Liu et al., 2013; Zheng et al., 2014). However, the influence of O₂ on N₂O reduction by clade II type N₂O-reducing bacteria has not been comprehensively studied, except Gemmatimonas aurantiaca strain T-27 (Park et al., 2017). To enable their use in engineering applications as N₂O sinks, the effects of oxygen on the N₂O reduction activities of clade I and II type N₂O-reducing bacteria need to be compared based on biokinetic analysis.

In this study, we, for the first time, compared the N₂O reduction kinetics of clade I and clade II type N₂O-reducing bacteria in the presence and absence of oxygen. Pseudomonas stutzeri and Paracoccus denitrificans, harboring nosZ clade I type, and two strains of Azospira sp., isolated from a N₂O-fed enrichment device (unpublished data) inoculated with municipal wastewater treatment biomass, were subject to biokinetic comparison.

Materials and Methods

Bacterial Strains and Culture Conditions

Two Azospira sp. strains and two strains of canonical denitrifiers, i.e., Ps. stuzeri strain JCM5965 (ATCC17588) and Pa. denitrificans strain NBRC102528 (ATCC17741), were used in this study. Azospira sp. strains I09 and I13, classified as Betaproteobacteria, were isolated from enrichment devices supplying N₂O as an electron acceptor (unpublished data). Ps. stutzeri and Pa. denitrificans were chosen because they are widely used as canonical denitrifiers (Körner et al., 1987; Baumann et al., 1996; Miyahara et al., 2010; Black et al., 2016). For comparison with other biokinetic studies, weight of a bacterial cell was determined as 0.388, 0.604, 0.743, and 0.787 pg-dry weight/cell for Azospira sp. strain I09, I13, Ps. stutzeri and Pa. denitrificans, respectively.

All the strains were aerobically pre-grown in an autoclaved nutrient medium containing (per liter of distilled water) 5.0 g of Bacto Peptone (BD-Difco, NJ), 3.0 g of Oxoid™ Lab-Lemco meat extract (Thermo Scientific, MA), and 5.0 g of NaCl.

Chemical Analyses

Dissolved organic carbon and dissolved total nitrogen were determined with a TOC analyzer, installing a total nitrogen measurement unit (5000A, Shimadzu, Kyoto, Japan). pH was measured using a pH meter (F-52, HORIBA, Kyoto, Japan). The gaseous N₂O concentration was determined using a gas chromatograph with an electron-capture detector (GC-14B, Shimadzu, Kyoto, Japan) instrument. The measurement conditions were described in a previous paper (Terada et al., 2013).

Activity Measurements

The pre-grown bacterial strains were harvested in centrifuge tubes at the early stationary phase. Two strains of Azospira spp. (I09 and I13) were washed twice with 0.05 × phosphate-buffered saline (PBS) by centrifugation at 5,000 rpm for 5 min, and re-suspended in the experimental medium. Our preliminary experiment showed that the N₂O-reducing activities of Ps. stutzeri JCM5965 and Pa. denitrificans NBRC102528 were significantly reduced by washing by centrifugation (data not shown). The pre-incubated cell suspensions of these two strains were therefore diluted with the experimental medium instead of washing. The experimental medium was placed in a conical flask, and the flask was sealed with a silicone cap (Shin-Etsu Polymer Co., Tokyo, Japan). The medium was mixed by shaking at 100 rpm at 30°C. The phosphate buffer medium, devoid of electron donors, contained (per liter of distilled water) 100 mg of KH₂PO₄, 6.6 mg of NaCl, 8.20 mg of MgSO₄·7H₂O, 13.4 mg of KCl, 115 mg of NH₄Cl, 188 mg of NaHCO₃, and 1 mL of a trace element solution consisting of (per liter of distilled water) 10 g of FeSO₄ 7H₂O, 10 g of FeCl₃ 7H₂O, 2 g of ZnSO₄ 7H₂O, 4 g of CuSO₄ 7H₂O, 0.5 g of NaMoO₄ 2H₂O, 0.1 g of MnCl₂ 4H₂O, 0.1 g of H₃BO₄, 0.3 g of Na₂SeO₃, and 10 g of citric acid (Miyahara et al., 2010). After sterilization, pH of the medium was adjusted to 7.5 with 1 M HCl.

The O₂ and N₂O consumption behaviors were investigated using an O₂ and N₂O micro-respiration system with amperometric microsensors (Unisense, Aarhus, Denmark). This system consisted of a Clark-type N₂O sensor (N2O-MR), O₂ sensor (OX-MR), double-port chamber (10 mL, MR-Ch double port), and stirrers installed in a sensor stand (Figure 1). The temperature during the experiments was controlled at 30 ± 0.2°C by a water bath. The experimental sequences of O₂ and N₂O consumption are shown in Figure 1B. The chamber was filled with the cell suspension, in which electrodes were inserted. Highly concentrated N₂O water (27.05–24.09 mM at 20–25°C and 0% salinity Weiss and Price, 1980) was prepared by pouring 25 mL of deionized water in a 50 mL vial, supplying pure N₂O gas to the water for 10 min at a room temperature and subsequently sealing the vial with a butyl rubber stopper. The highly concentrated N₂O water with a volume of 15–20 μL was injected into the chamber from the injection port (Figure 1A) using a Hamilton syringe, to replace the bacterial cell suspension and to give an initial dissolved N₂O concentration of approximately 50 μM. Then 250 mM sodium acetate solution (25 μL) was added to adjust the initial concentration to 625 μM. To achieve a homogeneous condition immediately, the glass-coated stir bar, was stirred at 600 rpm for 5 min after N₂O and carbon source injection and then at 300 rpm during the measurements. O₂ and N₂O concentrations were continuously monitored using SensorTrace Suite ver.2.8.0 (Unisense, Aarhus, Denmark). After the N₂O was completely consumed, highly concentrated N₂O water was injected again. The number of N₂O injections was changed depending on the distinct trends in the N₂O reduction rates observed among the tested strains. After the measurements, a sample of the cell suspension was immediately taken from the chamber and fixed with a 2% glutaraldehyde solution to determine the cell number, as previously described (Lunau et al., 2005). Briefly, the cell-counting procedure was as follows. Ethanol was added to the fixed cell suspension to achieve a final concentration of 10%. The mixture was homogenized for 10 s at 10 W (VP-050, Taitec, Tokyo, Japan), followed by dilution with 0.05 × PBS. The cells were trapped on a 0.2 μm membrane filter (Isopore, Merck Millipore, Germany), and washed twice with 1 mL of sterilized Tris-acetate EDTA buffer. The cells on the filter were stained with Moviol-SYBR Green I (Thermo Fisher Scientific, MA) and enumerated under a fluorescence microscope (BZ-8100, Keyence, Osaka, Japan).

FIGURE 1

Figure 1. (A) Diagram of double-port chamber and (B) experimental sequence of N₂O and O₂ respiration measurements.

Kinetic Parameter Estimation

The O₂ and N₂O concentration profiles were smoothed with the function of Sigma Plot 13.0 (Systat software, CA) to remove high-frequency noise and instantaneous reduction rate (Δ = 3–5 s) was calculated and normalized with cell number (Vs [pmol/h/cell]). The maximum O₂ and N₂O uptake rates (V_m,O2 and V_m,N2O [pmol/h/cell]) and the half-saturation constants for O₂ and N₂O (K_m,O2 and K_m,N2O [μM]) were determined by fitting the profiles to the Michaelis–Menten equation (Equation 1) (Martens-Habbena et al., 2009). The V_m,N2O value was calculated from the last measurement of the N₂O profiles.

$\begin{array}{cc}{V}_s=\frac{V_{\text{m},\text{S}}·S}{K_{\text{m},\text{S}}+S}& (1)\end{array}$

where S [μM] is the concentration of either O₂ or N₂O. To compare the N₂O reduction activities among the strains, the specific affinity for N₂O (a_0,N2O [L/cell/h]) was calculated as follows (Equation 2):

$\begin{array}{cc}{a}_{\text{0},\text{N2O}}=\frac{V_{\text{m},\text{N2O}}}{K_{\text{m},\text{N2O}}}\times {10}^{-6}& (2)\end{array}$

Statistical analysis was performed with ANOVA (Tukey HSD) in SPSS Statistics (IBM, NY), and statistical significance was evaluated by p-value below 0.05 as a threshold.

The effect of O₂ on the N₂O-reducing activity was investigated by fitting the experimental profiles to a mathematical model. The heterotrophic denitrification model, incorporating the O₂ inhibition (Ni et al., 2011), was used to estimate the degree of O₂ inhibition to N₂O uptake rates. The terms of an electron donor, nitric oxide (NO) and ammonium (${\text{NH}}_4^{+}$) were excluded in the model proposed by Ni et al. (2011) (Equation 3) because theoretically NO is not produced and organic carbon and ${\text{NH}}_4^{+}$ were in excess in the medium (i.e., addition of ${\text{NH}}_4^{+}$, organic carbon, and N₂O under anoxic conditions).

$\begin{array}{cc}\frac{d{S}_{\text{N2O}}}{dt}=V_{\text{m},\text{N2O}}\frac{S_{\text{N2O}}}{K_{\text{m},\text{N2O}}+S_{\text{N2O}}}\frac{K_{\text{I},\text{O2}}}{K_{\text{I},\text{O2}}+S_{\text{O2}}}X\times {10}^{-6}& (3)\end{array}$

where X [cells/L] is the concentration of bacterial cells in the chamber, and K_I,O2 [μM] is the O₂ inhibition coefficient. For Azospira sp. strains I09, I13, and Ps. stutzeri, K_I,O2 values were estimated by fitting the 1st-spiked N₂O profile to the model based on the least-squares method using the solver function of Microsoft Excel ver. 15.26. For Pa. denitrificans, determination of K_I,_O2 was not feasible because the trend of N₂O consumption is not explainable by the model. The lowest K_I,O2 value detected in this study was 0.1 μM due to the detection limit of an O₂ microelectrode.

The relative activity of N₂O reduction rate E [dimensionless] under an anoxic condition was calculated by Equation (4).

$\begin{array}{cc}E=\frac{V_{\text{N2O}}}{V_{\text{m},\text{N2O}}}& (4)\end{array}$

V_N2O was attained from an N₂O profile with the concentration range from 10 to 40 μM under an anoxic condition. The Nos activation rate (V_Nos [h⁻¹]) was defined as a degree of activity recovery of N₂O reduction after changing an aerobic to anoxic condition. V_Nos value was acquired by linear approximation of E as a function of elapsed time after DO concentration becomes zero (t_anoxic). The analysis was performed with Sigma Plot 13.0.

Quantifying Transcripts of nosZ Gene

Dynamics of nosZ gene transcripts of Azospira sp. strain I09 were quantified by reverse-transcription quantitative PCR (RT-qPCR). A medium, adding (per liter of distilled water) 0.20 g of sodium acetate and 0.050 g of NH₄Cl to the synthetic medium for the respirometric test, was autoclaved, followed by pH adjustment at 7.5 with 1 M HCl. Azospira sp. strain I09 was inoculated and aerobically pre-grown at 30°C in a 500 mL bottle until the late-exponential growth phase (OD600 = 0.121 after incubation for 23 h). After the aerobic pre-growth, different gases were supplied via a sterilized filter (HEPA-VENT, GE Healthcare, UK) at three different phases: 1 L/min of air in Phase 1, 0.5 L/min of N₂ in Phase 2, and 0.5 L/min of 100 ppm N₂O/N₂ in Phase 3. The gas flow rate was controlled using a mass flow controller (HORIBA, Kyoto, Japan). DO concentration in the medium was monitored using a DO meter (FDO Multi3410, WTW, Weilheim in Oberbayern, Germany). At each sampling point, cell suspension (10 mL) was transferred to a 15 mL tube and centrifuged at 10,000 rpm for 5 min at 4°C, and the supernatant was subsequently decanted. One milliliter of RNApro solution (FastRNA Pro Blue Kit, MP Biomedicals, CA) as a retardant of RNA degradation was immediately added to completely re-suspend the pellet according to the manufacture's protocol, followed by storage at 4°C until RNA extraction. RNA was extracted with a FastRNA Pro Blue Kit (MP Biomedicals, CA) and quantified by a UV-Vis spectrophotometer (Nanodrop 2000c, Thermo Fisher Scientific, MA). The extracted RNA was reverse-transcribed to complementary DNA with a QuantiTect Reverse Transcription Kit (QIAGEN, Hilden, Germany). The clade II type nosZ and 16S rRNA genes were quantified by real-time qPCR using a CFX96 Real-Time PCR Detection System (BioRad Laboratories, CA). The primer sets for clade II type nosZ and 16S rRNA genes were nosZ-II-F (5′-CTIGGICCIYTKCAYAC-3′)—nosZ-II-R (5′-GCIGARCARAAITCBGTRC-3′) (Jones et al., 2013) and 341f (5′-CCTACGGGAGGCAGCAG-3′)−517r (5′-ATTACCGCGGCTGCTGG-3′) (Muyzer et al., 1993), respectively. PCR for 16S rRNA gene amplification was initiated with initial denaturation at 95°C for 2 min, followed by 40 cycles at 95°C for 30 s, 60°C for 30 s, and 72°C for 30 s. PCR for the clade II type nosZ gene was initiated with denaturation at 95°C for 1 min, followed by 50 cycles at 95°C for 15 s, 54°C for 30 s, 72°C for 30 s, and 80°C for 30 s. The reaction buffer for the 16S rRNA gene consisted of 10 μL of SsoFast™ EvaGreen^® Supermix (BioRad Laboratories, Hercules, CA), 1 μL of 10 mM forward and reverse primers, 5 μL of template, and 3 μL of distilled water. The PCR buffer for the clade II type nosZ gene consisted of 10 μL of SYBR Premix Ex Taq II (Tli RNaseH Plus, Takara Bio, Shiga, Japan), 2 μL of 10 mM forward and reverse primers, 1 μL of distilled water, and 5 μL of template. Plasmid DNA (pGEM-T Easy Vector, Promega, WI) containing each gene was transformed into Escherichia coli competent cells (Competent High DH5α, Toyobo, Osaka, Japan) and the plasmids were isolated for each standard solution with a plasmid extraction kit (MagExtractor -Plasmid-, Toyobo, Osaka, Japan). After confirmation of insertion of each functional gene by Sp6 and T7 primers (Huang et al., 2005), the plasmid DNA as a positive control was diluted to obtain standard solutions containing 1.0 × 10⁸ to 1.0 × 10² copies per 5 μL in series. To increase the PCR efficiency, the plasmid containing the nosZ clade II amplicon was linearized with the restriction enzyme EcoRI (Takara bio, Kyoto, Japan) and used as a standard solution. The gene transcripts of nosZ were normalized with the amount of total RNA and gene transcripts of 16S rRNA to trace dynamics of nosZ gene expression. The transcripts of 16S rRNA gene was in the same order of magnitude during the tested three phases (Data not shown).

Results

Activity Measurements and Biokinetic Comparison

Activity measurement of each strain was conducted in triplicate. The representative O₂ and N₂O concentrations profiles are shown in Figure 2 and the two other replicates in SI (Figures S1–S4). All the bacterial strains showed facultative N₂O-reducing activities, consuming O₂ prior to N₂O. However, the O₂ concentration at which N₂O-reducing activity was initiated was not consistent for each strain. Except in the case of Pa. denitrificans, the first-spiked N₂O was completely consumed, followed by injection of N₂O-concentrated liquid. For Azospira sp. strains I09 and I13, the maximum N₂O reduction rate reached a plateau at the second or third additional N₂O injection. The maximum N₂O reduction rate of Azospira sp. I09 after the second injection was normalized to 114 ± 8% (n = 3) higher than that after the first injection. The analogous trend was attained for Azospira sp. I13, displaying that the maximum N₂O reduction rates after the second and third injections were 132 ± 14% (n = 3) and 132 ± 8% (n = 2) of the rate after the first one, respectively. For Ps. stutzeri, the maximum N₂O reduction rates after the additional injections further increased: 224 ± 69% (n = 3) for the second, 267 ± 62% (n = 3) for the third, 280 ± 40% (n = 2) for the fourth, and 294 ± 29% (n = 2) for the fifth, higher than the rate after the first injection. Pa. denitrificans was spiked with N₂O once because the reduction rate slowed down below 50 μM N₂O, entailing 10 h to consume the first-spiked N₂O (Figure S4). The initial N₂O concentration hampered accurate measurement for the N₂O reduction activity of Pa. denitrificans; therefore, the volume of injected N₂O solution was adjusted to ensure a higher N₂O concentration (100–150 μM) (Figure 2D and Figure S4). The maximum N₂O uptake rate (V_m,N2O) and half saturation constant for N₂O (K_m,N2O) were estimated by fitting the Michaelis–Menten equation to the N₂O profile at the final injection in each run (Table 1). Azospira sp. I13 showed the highest V_m,N2O, i.e., 5.80 ± 1.78 × 10⁻³ pmol/h/cell, among all the strains, followed by Ps. stutzeri (1.64 ± 0.34 × 10⁻³ pmol/h/cell), Azospira sp. I09 (6.34 ± 0.8 × 10⁻⁴ pmol/h/cell), and Pa. denitrificans (5.01 ± 1.0 × 10⁻⁴ pmol/h/cell). The highest and lowest K_m,N2O values were 34.8 ± 10.2 μM for Pa. denitirificans and 0.866 ± 0.29 μM for Azospira sp. I09, respectively. The other two strains displayed comparable K_m,N2O values of about 4 μM. The N₂O affinities (a_0,N2O) of Azospira sp. strains I13 and I09 were higher than those of Ps. stutzeri and Pa. denitirificans because of the inherently higher V_m,N2O (I13) or lower K_m,N2O (I09).

FIGURE 2

Figure 2. N₂O and O₂ respiration profiles of isolates: (A) Azospira sp. strain I09, (B) Azospira sp. strain I13, (C) Pseudomonas stutzeri JCM5965, and (D) Paracoccus denitrificans strain NBRC102528. Black line: O₂, gray line: N₂O. Vertical arrows indicate N₂O spikes.

TABLE 1

Table 1. Biokinetic parameters for the tested strains.

Effects of O₂ on N₂O Reduction

The effects of O₂ on N₂O reduction were compared on the basis of the relative activities of N₂O reduction, E (Equation 4). E as a function of O₂ concentration is shown in Figure 3. The effect of the DO concentration on E differed among the tested strains. Although the activity was lower (10% of V_m,N2O) than that in the absence of oxygen, Azospira sp. I09 showed N₂O consumption activity under microaerophilic conditions at DO concentrations of 100–110 μM. In contrast, the DO concentration needed to initiate N₂O consumption by Azospira sp. I13 and Pa. denitrificans was much lower (25 μM O₂). The N₂O consumptions of these three strains recovered significantly after complete consumption of O₂. Ps. stutzeri did not consume N₂O until the O₂ was completely depleted.

FIGURE 3

Figure 3. Relative N₂O consumption rates as function of O₂ concentration.

The time series for the N₂O relative reduction rates under anoxic conditions are shown in Figure 4. Pa. denitrificans was excluded from the analysis because the inherently low N₂O affinity of Pa. denitrificans hampered accurate measurement of the N₂O reduction rate (Table 1). Relative activity of N₂O reduction rate increased linearly in the three strains. The two Azospira sp. strains showed the same trend in E-values, and regained 0.8 of the initial value in 1.93 h (I09) and 1.38 h (I13), respectively. In contrast, Ps. stutzeri required 4.32 h for 80% recovery of E (Figure 4). The Nos activation rates (V_Nos), i.e., the slopes in Figure 4, were estimated to be 0.319 ± 0.028, 0.397 ± 0.064, and 0.200 ± 0.013 h⁻¹ for Azospira sp. strain I09, I13, and Ps. stutzeri, respectively, indicating the highest V_Nos value for Azospira sp. strain I13.

FIGURE 4

Figure 4. Recovery of N₂O reduction activity after O₂ exposure. Plots are derived from N₂O consumption tests, performed in duplicate or triplicate: t_anoxic = 0 is elapsed time after DO is completely depleted. Solid and broken lines represent approximated activity recovery for N₂O reduction in each strain and the 95% confidence interval of each approximated line, respectively. For the approximation, triplicate and duplicate data were used for the two Azospira sp. strains and Ps. stutzeri, respectively.

The effect of O₂ on the N₂O reduction activity was quantitatively compared by determination of biokinetic parameters (Figure S5). K_I,O2 was estimated by the model fitting for N₂O profiles (Figure S5). Azospira sp. strain I09 showed the highest K_I,O2 value of 2.33 ± 1.7 μM while those of Azospira sp. strain I13 and Ps. stutzeri were below 0.5 μM as summarized in Table 1.

nosZ Gene Transcription of Azospira sp. Strain I09

Dynamics of nosZ transcription of Azospira sp. strain I09, displaying N₂O uptake even at above 100 μM O₂, was monitored under different redox conditions (Figure 5). During Phase 1 for aerobic pre-incubation by air bubbling, nosZ transcription level slightly increased. Switching to the anoxic condition by pure N₂ bubbling decreased the nosZ transcription level after 40 min by 74%. Subsequently, mixing N₂O with N₂ gas to keep N₂O concentration of 100 ppmv (equivalent dissolved N₂O of 2.16 μM at 30°C) in Phase 3 stimulated nosZ transcription, reaching the comparable level in Phase 1.

FIGURE 5

Figure 5. Dynamics of nosZ gene expression in Azospira sp. I09 under three different redox conditions. The gaseous compositions were ambient air (N₂O: 0.34 ppm) in Phase 1, N₂ (N₂O: <0.005 ppm) in Phase 2, and N₂ + N₂O (N₂O: 100 ppm) in Phase 3, respectively. The gas flow rate was consistently 0.5 L/min.

Discussion

This study, for the first time, showed the effects of oxygen on the N₂O reduction activities of nosZ clade I and II type N₂O-reducing bacteria in a comparative manner. Monitoring N₂O consumption dynamics has always been challenging because of the difficulty in setting up conditions under which the N₂O and O₂ concentrations are always transitioning. A common batch experiment, measuring the headspace gas in a vial (Bueno et al., 2015), tackled the difficulty of monitoring N₂O and O₂ concentrations within short intervals. The micro-respirometric system using DO and N₂O microsensors used in this study enabled real-time monitoring of these dissolved gases at a much higher resolution (second level) than in conventional methods. The highly resolved profiles detect N₂O-reducing activity even under aerobic conditions. We found that physiological characteristics do not clearly distinguish clade I and II type N₂O-reducing bacteria.

There has been open discussion on whether clade II type N₂O-reducing bacteria have inherently high N₂O affinities compared with those of their clade I type counterparts (Yoon et al., 2016; Hallin et al., 2018). Current research suggested the existence of unique electron transport module in clade II type microorganisms, i.e., Wolinella succinogenes (Hein et al., 2017). The feature is advantageous for low-energy translocation pathways and effective energy conversion within the N₂O respiration compared to nosZ clade I type bacteria. Our biokinetic analyses of the clade II type N₂O-reducing bacterium, i.e., two Azospira sp. strains, show that their half-saturation constants (K_m,N2O) are lower than those of the two clade I type denitrifying bacteria (Table 1). This is in agreement with previous studies, in which clade II type strains, namely Dechloromonas aromatica strain RCB and Anaeromyxobacter dehalogenans strain 2CP-C, gave lower K_m,N2O and higher biomass yields than did clade I type strains (Yoon et al., 2016). Further comparison, with the substrate affinity a_0,N2O as an indicator of N₂O-reducing activity, shows that clade II type Azospira sp. strain I13 is a promising N₂O sink (Table 1). However, even for the same Azospira sp., the two isolates had distinct a_0,N2O values. Furthermore, the recent report demonstrated that a chemostat system, supplying N₂O as a sole electron acceptor, enriched clade I type N₂O reducing bacteria with high a_0,N2O (Conthe et al., 2018). Given that taxonomic proximity is not necessarily linked to physiology, N₂O-reducing bacteria having very high affinity to N₂O could be broadly distributed irrespective nosZ clade type. Comprehensive biokinetic analysis focusing on not only clade II type but also clade I type is needed.

The N₂O reduction activities in the presence of O₂ differed among the four bacterial strains, irrespective of their nosZ clade types. Azospira sp. strain I09 exhibited N₂O-reducing activity at a DO concentration of 110 μM (3.52 mg/L) and higher O₂ inhibition coefficient (2.33 μM), confirming the participation of aerobic N₂O reduction. Previous physiological studies using isolates or mixed cultures implied that some bacteria are capable of denitrifying and respiring N₂O under aerobic or microaerobic conditions, but the level of DO concentration needed for detectable N₂O-reducing activity depends on the bacterial strain (Liu et al., 2013). Thauera sp. strain 63 expresses nosZ mRNA under microaerobic conditions (<10 μM), although T. terpenica strain DSM12139 does so under obligate anaerobic conditions (Liu et al., 2013). The Nos of Ps. stutzeri strain ATCC14405 is expressed at a DO concentration of 119 μM (Körner and Zumft, 1989) and Ps. stutzeri strain TR2 showed N₂O-reducing activity at a DO concentration of 35 μM (Miyahara et al., 2010). In a mixed culture, ${\text{NO}}_2^{-}$ reduction activity by microorganisms in soils appears at a DO concentration of 80 μM (Morley et al., 2008), and denitrification gene transcription is continuously detected under O₂-remaining conditions in coastal sediments (Marchant et al., 2017). On the basis of previous reports, the DO level for the emergence of N₂O consumption by Azospira sp. strain I09 (110 μM) is higher than those of Thauera sp. strain 63 and Ps. stutzeri strain TR2, and comparable to that of Ps. stutzeri strain ATCC14405. Thorough, holistic investigations are needed, but our results suggest that Azospira sp. strain I09 has promise as a N₂O consumer under microaerophilic conditions.

The activity tolerance of O₂ inhibition was reasonably described applying the Nos activation rate, V_Nos (Figure 4). The higher values of V_Nos for the Azospira sp. strains (0.319 ± 0.028h⁻¹ for I09 and 0.397 ± 0.064 h⁻¹ for I13) than that for Ps. stutzeri (0.200 ± 0.013 h⁻¹) indicate that the recovery of N₂O reduction activity by the Azospira sp. strains is faster than that by Ps. stutzeri. In addition, E-values for the Azospira sp. strains higher than 0 at t_anoxic = 0 suggests their superior N₂O uptake resilience to O₂ exposure, which was not observed for Ps. stutzeri (Figure 4). In previous reports, two mechanisms of O₂ inhibition to N₂O reduction activity have been hypothesized, namely regulation of the RNA transcription level (Arai, 2011; Bergaust et al., 2012) and Nos inactivation (Otte et al., 1996; Miyahara et al., 2010). The quantification of nosZ clade II type mRNA transcripts for Azospira sp. strain I09 under different redox conditions likely provided Nos reactivation as a limiting factor to determine N₂O uptake rate. The application of RT-qPCR demonstrated that expression of the nosZ mRNA by Azospira sp. strain I09 was suppressed in the absence of O₂ and N₂O but not in the presence of N₂O and O₂ (Phase 1 and Phase 3 in Figure 5). This agrees with the trend that nosZ gene is expressed even in the presence of O₂ in coastal sediments (Marchant et al., 2017). The consistent trend corroborates that Nos is probably synthesized even under aerobic or microaerobic conditions. Studies on extracted and purified Nos support Nos recovery after switching from aerobic to anoxic conditions (Ghosh et al., 2003; Chan et al., 2004). On the basis of these observations, it is likely that recovery of N₂O consumption depends on the rate of Nos reactivation.

Toward N₂O mitigation in engineered systems, the degree of Nos reactivation after O₂ exposure should be quantitatively estimated. Combination of a respirometric analysis with a mechanistic modeling potentially accelerates selection of highly efficient N₂O-reducing bacteria able to respire N₂O under microaerophilic conditions. The heterotrophic denitrification model, incorporating the O₂ inhibition (Equation 3) (Ni et al., 2011), likely necessitates the extension due to some discrepancies with the experimental results (Figure S5). Validation and verification of the extended models, e.g., an enzyme-explicit denitrification model, where a ratio of inactive Nos to total Nos and its recovery rate determine N₂O uptake rate (Zheng and Doskey, 2015) and an integrated model incorporating the O₂ inhibition and Nos recovery, warrant future intensive studies.

This study shows that strains of the genus Azospira are promising N₂O reducers because of their O₂ inhibition tolerance and high affinity for N₂O. Use of the proposed kinetic parameters, V_Nos and K_I,O2, provides, for the first time, quantitative information on Nos recovery from O₂ inhibition. Application of respirometric biokinetic analysis will be useful in developing N₂O mitigation strategies using N₂O reducers in engineered systems. Comprehensive screening of N₂O-reducing bacteria displaying high N₂O affinities (a_0,N2O) and high resilience against O₂ exposure (V_Nos, K_I,O2) is required.

Conclusion

This study investigated the effects of O₂ on the N₂O consumption biokinetics of bacterial isolates affiliated to nosZ clade I and II types. Respirometric assays showed that the N₂O affinities of two Azospira sp. strains of nosZ clade II type were higher than those of Pa. denitrificans and Ps. stutzeri of nosZ clade I type. However, the clade type does not completely explain the different N₂O affinities, and this suggests that the physiological traits of N₂O-reducing bacteria differ at the species and strain levels. The N₂O consumption activities are also significantly affected by the O₂ levels but the degree of O₂ inhibition differs depending on the species or strain rather than the nosZ type. Azospira sp. strain I09 showed aerobic N₂O-reducing activity and nosZ gene transcription at a DO concentration of 110 μM. Recoveries of the N₂O-reducing activities of Azospira sp. strains I09 and I13 after O₂ exposure are faster than that of Ps. stutzeri of nosZ clade I type. This underscores that the Nos of Azospira spp. is reactivated after switching from aerobic to anoxic conditions. Our results provide new information on the physiological characteristics of N₂O-reducing bacteria, and also a valuable investigative parameter involving a respirometric analysis. This comprehensive and thorough analyses based on a respirometric approach will accelerate the discovery of novel N₂O reducers and contribute to N₂O mitigation in engineering applications.

Author Contributions

AT: designed and led the study. TS: performed the biokinetic estimation and oxygen effect on nitrous oxide reduction of the isolates. TS: wrote the paper with major edits and inputs from SR, MH, and AT.

Funding

We would like to thank the Japan Society for the Promotion of Science (JSPS) for supporting this research with Grants-in-Aid for Challenging Exploratory Research (26630420, 16K12616) and JSPS fellows (16J08601).

Conflict of Interest Statement

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Acknowledgments

We thank Dr. Barth F. Smets (Technical University of Denmark, Kongens Lyngby, Denmark) and Dr. Tomoyuki Hori (National Institute of Advanced Industrial Science and Technology, Ibaraki, Japan) for discussing this research and Ms. Kanako Mori for supporting molecular biology experiments. The microorganism cultures were provided by the RIKEN Biological Resource Center (JCM, Ibaraki, Japan) and NITE Biological Resource Center (NBRC, Chiba, Japan). We thank Helen McPherson, Ph.D., from Edanz Group (www.edanzediting.com/ac) for editing a draft of this manuscript.

Supplementary Material

The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fmicb.2018.00697/full#supplementary-material

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