Isolation and Characterization of Ammonia-Oxidizing Bacterium N.eA1: Insights into Nitrogen Conversion and N₂O Emissions in Varied Environmental Conditions

Abstract

While temperature, pH, DO, and ammonia nitrogen concentration are known to affect nitrous oxide (N₂O) emissions from ammonia-oxidizing bacteria (AOB), the specific responses of individual AOB species to these environmental variables have yet to be fully elucidated. The present study reports the isolation and pure culture of a new AOB strain, designated as N.eA1, from a stable CANON bioreactor. The strain’s denitrification and N₂O emission were systematically evaluated through a comprehensive analysis of growth kinetics, morphological characteristics, genetic composition, and nitrogen transformation under various environmental processes. Our results indicated that N.eA1 shares 95.33% sequence homology with Nitrosomonas europaea H1 AOB3, and exhibited higher nitrite (NO₂⁻-N) conversion efficiency. Morphological examination revealed white, semi-transparent spherical colonies. The bacterial growth kinetics included adaptation phase (0–12 h), exponential growth phase (12–36 h), stationary phase (36–72 h) and decline phase (after 72 h). Under optimal cultivation conditions (30 °C, DO concentration of 7.3 mg∙L⁻¹, pH 8.0, and NH₄⁺-N concentration of 260 mg∙L⁻¹), the culture achieved a maximum growth rate of 0.0723 h⁻¹, a maximum ammonia oxidation rate (AOR) of 10.74 mg∙(MLVSS∙h)⁻¹, and a minimum doubling time of 9.59 h. The peak time of nitrogen conversion was earlier than that of N₂O emission, with a maximum N₂O-N conversion from NH₄⁺-N of 1.039%.

1. Introduction

Global warming has garnered extensive attention in recent years [1], with the reduction of greenhouse gas (GHG) emissions emerging as a primary strategy to address this challenge [2]. Among these gases, N₂O, a significant contributor to stratospheric ozone depletion, has attracted increasing scientific scrutiny [3]. Previous studies indicate that N₂O exhibits a global warming potential 298 times greater than that of carbon dioxide (CO₂) over a 100-year period, with its atmospheric concentration continuing to rise at an annual rate of 0.26% [4]. Wastewater treatment processes have been identified as one of the principal sources contributing to this phenomenon. In these processes, the formation of N₂O is driven by multiple microbial processes, including nitrification and denitrification [5,6]. Therefore, understanding the mechanisms of microbial N₂O production in biological treatment systems is crucial for regulating N₂O emissions in wastewater treatment processes.

Biological treatment has gained prominence in nitrogen removal from wastewater due to its cost-effectiveness and minimal secondary pollution [7,8]. Within these systems, ammonia-oxidizing bacteria (AOB) are crucial. AOB are classified as Gram-negative chemolithoautotrophic microorganisms that utilize ammonia oxidation as their primary energy source [9]. These organisms exhibit distinctive physiological characteristics, including stringent growth requirements with a preference for neutral to slightly alkaline conditions, and grow slowly [10,11]. Therefore, AOB affect the nitrogen content of the entire ecosystem [12,13]. Their significance extends beyond wastewater treatment, as they represent crucial mediators in the global nitrogen cycle and constitute essential members of the microbial consortia present in wastewater treatment processes [10]. AOB participate in both nitrification and denitrification. Nitrification is an important N cycling process in ecosystems [14], which oxidizes ammonia (NH₃) to nitrate (NO₃⁻-N) via intermediate products, i.e., hydroxylamine (NH₂OH) and nitrite (NO₂⁻-N) in a two-step process, and can consequently emit N₂O as a by-product under certain conditions [12,15]. Ammonia oxidation, the oxidation of NH₃ to NO₂⁻-N via NH₂OH, is the rate-limiting step in nitrification and can be carried out by both AOB and ammonia-oxidizing archaea (AOA). Nitrification-related processes (the oxidation of NH₂OH and NO₂⁻-N reduction) mediated by AOB are recognized to be a main pathway of N₂O emission [16].

While the role of AOB in nitrogen cycling has been well established, their environmental sensitivity remains a critical factor influencing both their performance and N₂O emissions. Environmental parameters, including temperature, DO, and pH, significantly affect AOB activity and N₂O production [17]. Jiang [18] demonstrated that low-temperature conditions reduce the activity of AOB by impairing functional gene clusters, thereby suppressing N₂O emissions in AOB-dominated communities. Guo [19] utilized online monitoring to link real-time N₂O fluxes with microbial community dynamics in an anoxic–oxic (A/O) wastewater treatment system. Their findings underscore the critical role of pH regulation, recommending an influent pH of 6–8 as optimal for minimizing emissions during anaerobic processes. Rathnayake [20] observed a positive correlation between dissolved oxygen (DO) levels (0.5–7.3 mg∙L⁻¹) and N₂O production rates in autotrophic partial nitrification (PN) granules, attributing this phenomenon to enhanced ammonia oxidation at higher DO concentrations. Sarkar [21] investigated the effects of historical land use and precipitation regimes on the metabolic niches of ammonia-oxidizing archaea (AOA) and AOB in terrestrial ecosystems, demonstrating that drought stress amplifies N₂O fluxes through niche partitioning between these microbial guilds.

These studies provide valuable insights into the environmental adaptability of AOB and their contributions to N₂O emissions. However, technical limitations have confined most prior investigations to mixed microbial consortia, inherently masking strain-specific responses to operational parameters [22]. Furthermore, the slow growth rates, fastidious nutritional requirements, and susceptibility of AOB to environmental perturbations have posed substantial challenges in obtaining stable pure cultures [23]. For instance, in partial nitrification systems, elevated N₂O fluxes are often ambiguously attributed to AOB-dominated processes, without distinguishing contributions from individual species or their interactions with heterotrophs [24]. Systematic comparisons of AOB ammonia oxidation kinetics and N₂O emissions under well-controlled multi-parametric conditions remain scarce. Critical unresolved questions include the following: How do individual AOB strains balance ammonium oxidation efficiency against N₂O emission potential across environmental gradients? Can pure culture-derived thresholds for pH, DO, and temperature be directly translated to complex bioreactor ecosystems?

This study aims to address these persistent challenges by unraveling the strain-specific contributions of AOB to N₂O emissions. The primary objectives are as follows: establishing axenic cultures of an AOB strain isolated from a CANON bioreactor; systematically quantifying its ammonia oxidation kinetics and emission responses across environmental gradients, including temperature, pH, DO, and NH₄⁺-N concentrations, and exploring the temporal dynamics of nitrogen transformation; developing threshold-based operational guidelines to optimize nitrogen removal efficiency while mitigating N₂O emissions. The experimental design progresses sequentially from microbial isolation and genomic characterization to controlled-environment physiological profiling, ultimately exploring the nitrogen transformation pathway. Throughout this workflow, we prioritize decoupling strain-specific behaviors from mixed-culture effects—a critical gap in previous studies on microbial consortia. By isolating AOB strain-specific responses to environmental stressors, this study eliminates confounding interactions with heterotrophic denitrifiers or AOA, thereby resolving conflicting observations in previous mixed-culture research. Furthermore, we offer a predictive framework for optimizing CANON bioreactors. By integrating kinetic modeling with phylogenetic analysis, we demonstrate that phylogenetically similar AOB strains exhibit divergent N₂O emission potentials under identical conditions, highlighting the need for strain-level resolution in greenhouse gas inventories. This study provides critical insights for optimizing AOB performance and mitigating N₂O emissions, thereby contributing to the development of more sustainable nitrogen removal technologies.

2. Materials and Methods

2.1. Separation, Purification and Enrichment Culture

Original sludge was sourced from a stable CANON bioreactor operating in a laboratory (Guizhou, China). The inlet water of the bioreactor is simulated industrial sewage, and its water quality is characterized by high NH₄⁺-N and low C/N ratio. The bacterial strain was introduced into 100 mL of liquid culture medium (

Table 1. Ammonia-oxidizing bacteria purification medium.

Serial Number	Reagent	Addition Amount (μg∙L−1)
1	NaCl	300
2	MgSO4·7H2O	140
3	FeSO4·7H2O	30
4	KH2PO4	20
5	NH4HCO3	282
6	NaHCO3	1600
7	CuSO4·5H2O *	0.075
8	ZnSO47H2O *	0.03
9	CoCl2 6H2O *	0.375
10	MnCl2·2H2O *	0.3
11	H3BO4 *	0.014
12	EDTA *	0.5
13	NaMoO4·2H2O *	0.22

Note: Those marked with * are trace elements; Reagents are all from Sinopharm Chemical Reagent Co., Ltd., Shanghai, China.

) at an inoculation volume ratio of 2% (v/v). Cultivation was performed in a shaking incubator (VRERA, Shanghai, China) maintained at 30 °C with an agitation rate of 120 r∙min⁻¹, without light throughout the incubation period. Medium pH was adjusted to 8.0 using 1 mol∙L⁻¹ hydrochloric acid (HCl) (Sinopharm Chemical Reagent Co., Ltd., Shanghai, China) and sodium hydroxide (NaOH) (Sinopharm Chemical Reagent Co., Ltd., Shanghai, China) solutions. NO₂⁻-N accumulation was monitored at 24 h intervals. Positive cultures exhibiting significant NO₂⁻-N accumulation were subcultured into fresh medium at a 1% (v/v) inoculation ratio for five consecutive rounds. Subsequently, positive samples were subjected to three rounds of streak/spread plate isolation on solid medium, followed by selection of single colonies for incubation in a shaking incubator (Figure 1). Throughout the experimental period, pH was regularly monitored and maintained at 8.0. For solid medium preparation, 1.5% (w/v) agar was supplemented. The medium was subsequently autoclaved at 121 °C for 30 min and stored until use.

2.2. Growth Characteristic Experiment

The bacterial enrichment culture obtained from Section 2.1 was inoculated into fresh liquid medium at a 2% (v/v) inoculation ratio. Initial cultivation parameters were established as follows: NH₄⁺-N concentration of 260 mg∙L⁻¹ and pH 8.0. The cultures were maintained in an orbital shaking incubator under light-excluded conditions at 30 °C with continuous agitation at 120 r·min⁻¹. Quantitative measurements of NH₄⁺-N concentration, NO₂⁻-N concentration, and bacterial growth (monitored as optical density at 600 nm) were conducted at 12 h intervals throughout the 96 h cultivation period. All experimental conditions were established with duplicate control groups.

2.3. Environmental Factor Experiment

The bacterial enrichment culture obtained from Section 2.1 was inoculated into fresh liquid medium at a 2% (v/v) inoculation ratio. The experimental design incorporated the investigation of four environmental parameters: temperature, pH, DO concentration and NH₄⁺-N concentration (

Table 2. Single factor experiment.

Serial Number	Factors	Level	Cultivation Conditions	Measuring Time Interval
1	Temperature	25 °C	120 rpm; in dark	12 h
2		30 °C
3		35 °C
4	pH	6.5	120 rpm; in dark	12 h
5		8.0
6		9.5
7	DO concentration	2.5 ± 0.5 mg∙L−1	120 rpm; in dark	6 h
8		4.5 ± 0.2 mg∙L−1
9		5.8 ± 0.2 mg∙L−1
10		6.5 ± 0.2 mg∙L−1
11		7.3 ± 0.2 mg∙L−1
12	NH4+-N concentration	36.5 ± 0.5 mg∙L−1	120 rpm; in dark	12 h
13		71.0 ± 0.5 mg∙L−1
14		133.0 ± 0.5 mg∙L−1
15		281.0 ± 0.5 mg∙L−1

). Throughout the environmental factor studies, all parameters were maintained at constant levels except for the specific variable under investigation. All experimental conditions were established with duplicate control groups. In pH experiments, medium pH was adjusted through titration with 1 mol∙L⁻¹ HCl and NaOH solutions. A phosphate buffer system was incorporated to ensure pH stability throughout the experimental period. pH was monitored at 6 h intervals.

For DO experiments, aeration was facilitated through a glass tube apparatus extending to the vessel bottom, with the external air inlet equipped with two serially-connected 0.22 μm filters to prevent microbial contamination from ambient air. DO concentrations were regulated via a controlled aeration system (Aquasystems International, Halle, Belgium), implementing distinct aeration durations (5, 20, 40, and 60 min) at 2-h intervals. A non-aerated control group was maintained throughout the experiment, with DO levels continuously monitored using a calibrated DO meter (YSI, Yellow Springs, OH, USA). Gas samples were collected over 1 min intervals from each experimental group at 6 h intervals during aeration periods. Prior to subsequent aeration cycles, cultures were subjected to agitation at 200 r∙min⁻¹ for 20 min, followed by collection of headspace gas samples. In the NH₄⁺-N concentration experiment, while the theoretical NH₄⁺-N concentrations were set at 50, 100, 200, and 400 mg∙L⁻¹, the actual measured concentrations were 36.5, 71.0, 133.0, and 281.0 mg∙L⁻¹, respectively.

2.4. Bioinfomatics Analysis

The enriched flocculent biomass was placed in 1.5 mL centrifuge tubes and centrifuged at 4500 rpm for 5 min. After discarding the supernatant, sterile water was added to restore the original volume. This centrifugation process was repeated three times. The resulting biomass was stored at −20 °C and sent to Guiyang Weilai Testing Co., Ltd. (Guiyang, China) for bio-electron microscopy analysis.

The enrichment culture from a single colony was centrifuged, and genomic DNA was extracted using the Ezup Column Bacterial Genomic DNA Extraction Kit (Shanghai Sangon Biotech, Shanghai, China). PCR amplification was done using the TC1000-G PCR machine (SCILOGEX, Rocky Hill, CT, USA). PCR amplification was performed using primers 27F (5′-AGAGTTTGATCMTGGCTCAG-3′) and 1541R (5′-AAGGAGGTGATCCA GCC-3′) as forward and reverse primers, respectively. The PCR products were verified by 1% (w/v) agarose gel electrophoresis and then sent to Sangon Biotech (Shanghai, China) for sequencing. The sequencing results were analyzed using BLAST (https://blast.ncbi.nlm.nih.gov/Blast.cgi, accessed on 14 November 2023) against the NCBI database, and a phylogenetic tree was constructed using the Neighbor-Joining method implemented in MEGA11.

2.5. Analytical Methods

NH₄⁺-N, NO₂⁻-N concentrations and OD600 were measured using a UV-visible spectrophotometer (Shimadzu, Kyoto, Japan). NH₄⁺-N was measured using the Nessler’s reagent spectrophotometric method [25], with a wavelength of 420 nm. NO₂⁻-N was measured using the N-(1-naphthyl)-ethylenediamine spectrophotometric method [26], with a wavelength of 520 nm. N₂O was collected at each phase using a gas sampler and analyzed using a GC9790⁺ gas chromatograph (Foley Instrument Co., LTD, Zhejiang, China). High-purity N₂ (99.999%) was used as the carrier gas at a flow rate of 21 mL·min⁻¹. The temperatures of the ECD detector and column oven were set at 250 °C and 70 °C, respectively. Data analyses and visualizations were performed using Python (Version 3.8.5, Python Software Foundation). The maximum growth rate (μ_max) was determined through linear regression analysis of the natural logarithm of OD600 values during the exponential growth phase (12–36 h) using scipy.stats module.

The maximum specific growth rate (μ_max) (h⁻¹) and minimum doubling time (t_d) (h) were calculated as follows:

t_d = ln (2)/μ_max

(1)

The maximum AOR [mg∙(L∙h)⁻¹] was calculated by linear regression of NH₄⁺-N consumption data during the period of highest activity; the unit conversion was performed using the following formula (the specific formula is in Appendix A):

V_max = v/C_mlvss

(2)

where:

V_max: Maximum AOR [mg∙(MLVSS∙h)⁻¹]

v: Maximum AOR [mg∙(L∙h)⁻¹]

C_mlvss: 0.38—biomass concentration (g∙L⁻¹)

The N₂O conversion percentage was determined by the ratio of N₂O-N (sum of N₂O in liquid and gas phases) to the total nitrogen change (ΔN). The conversion percentage of N₂O was calculated as follows (the specific formula is in Appendix A):

P = (N₂O_liquid + N₂O_air)/ΔN × 100%

(3)

where:

N₂O_liquid: N₂O in liquid (mg∙L⁻¹)

N₂O_air: N₂O in air (mg∙L⁻¹)

ΔN: Change in total nitrogen (mg∙L⁻¹)

In the measurement of NH₄⁺-N, nitrogen exists in both ammonium ion (NH₄⁺) and free ammonia (FA) forms, which undergo dynamic interconversion. An equilibrium relationship exists between NH₄⁺ and FA in wastewater. The calculation formula for FA concentration was derived by incorporating the ionization constants of ammonia and water into the equilibrium equation [27,28]:

FA = 17 (NH₄⁺-N) × 10^pH/14e^6334/(273+t) + 10^pH

(4)

Results are presented as mean values of triplicate experiments with standard deviation (mean ± SD, n = 3). Time series analyses were conducted to evaluate the temporal dynamics of NH₄⁺-N degradation, NO₂⁻-N accumulation, and N₂O emissions in different environmental conditions.

Sensitivity Analysis Using Standardized Regression Coefficients (SRC) [29] quantifies the effect of each independent variable on the dependent variable (N₂O emissions) in a regression model. This method is widely used to understand how variations in input variables impact model outputs [30]. The regression model can be expressed as follows:

$$y=b_0+{\sum }_{k=1}^n{b}_k{X}_k$$

(5)

where:

$y$: N₂O emissions (dependent variable)

$X_{1,...,k}$: The environmental factors (temperature, pH, DO, and NH₄⁺-N)

$b_{1,...,k}$: The regression coefficients

The data for each independent variable (X) were standardized by subtracting the mean and dividing by the standard deviation. This ensures that each variable contributes equally to the analysis, allowing for comparison of their relative effects on N₂O emissions.

The Standardized Regression Coefficients (SRC) are given by the following equation:

$${SRC}_i={\displaystyle \frac{{\beta }_i·SD\left(X_i\right)}{SD\left(Y\right)}}$$

(6)

where:

${\beta }_i$: The regression coefficient for each independent variable

$SD\left(X_i\right)$: The standard deviation of the independent variable

$X_i$$SD\left(Y\right)$: The standard deviation of the dependent variable $Y$

The sensitivity analysis was carried out using Python 3.8 and the statsmodels package for regression analysis. Data were first standardized, and then an Ordinary Least Squares (OLS) regression was performed. The SRC values were calculated to quantify the influence of each environmental factor on N₂O emissions.

The R² value of the regression model was also calculated to ensure the validity of the results. The R² value indicates the proportion of variance in N₂O emissions explained by the model. A value above 0.7 was considered acceptable for determining the effectiveness of the SRC coefficients.

3. Results and Discussion

3.1. Growth Characteristics, Genetic Identification and Morphological Analysis

A comparative analysis on the NCBI website revealed a 95.33% homology with Nitrosomonas europaea H1 AOB3 (Figure 2), and the strain was designated as N.eA1. Colonies appeared as white, semi-translucent, spherical structures, morphologically similar to the Nitrosomonas colonies isolated from activated sludge [7]. SEM images revealed rod-shaped bacterial cells, measuring 8–10 µm × 5–6 µm, enveloped in extracellular polymeric substances (EPS) (Figure 3b,d). Specifically, EPS serves to protect AOB against environmental stressors [31] and provides nutrient reserves during starvation periods, thereby enhancing microbial stability and function [32,33]. However, excessively thick EPS layers can impede oxygen and ammonia diffusion, potentially reducing nitrogen removal efficiency [34]. Therefore, maintaining optimal EPS thickness is crucial for maximizing CANON system performance.

The adaptation, exponential growth, stationary, and decline phases occurred at 12 h, 12–36 h, 36–72 h, and above 72 h post-inoculation, respectively. During the logarithmic phase, the strain exhibited enhanced growth rates (Figure 3a), with a maximum growth rate of 0.0723 h⁻¹ and a minimum doubling time of 9.59 h, similar to Nitrosomonas europaea [35,36]. Furthermore, under similar NO₂⁻-N concentrations, the N₂O emission rate of the N.eA1 strain (3.494 × 10⁻⁵ mg N₂O/mg NO₂⁻-N) was higher than that of Nitrosomonas europaea (1.048 × 10⁻⁵ mg N₂O/mg NO₂⁻-N) [37], suggesting that the N.eA1 strain may maintain a higher metabolic activity state under similar NO₂⁻-N concentrations.

Throughout the growth process, the maximum AOR was 10.74 mg∙(MLVSS∙h)⁻¹, with an average AOR of 5.08 mg∙(MLVSS∙h)⁻¹. When the initial NH₄⁺-N concentration was 260 mg·L⁻¹, more than 50% of NH₄⁺-N degradation occurred within 72 h (Figure 3c). This rapid degradation phase was likely driven by the high metabolic activity of the strain during the initial stages of cultivation, where NH₄⁺-N served as the primary substrate for energy production and growth. This finding is consistent with previous studies indicating that AOB can efficiently oxidize ammonia under optimal conditions [38,39]. Correspondingly, NO₂⁻-N accumulation increased over time (Figure 3e). In the later stages of cultivation, the NH₄⁺-N concentration gradually leveled off until it remained constant, potentially due to NO₂⁻-N accumulation leading to reduced bacterial activity and inhibition of the ammonia oxidation reaction. It was observed that when NO₂⁻-N exceeded 90 mg·L⁻¹, it significantly inhibited AOB activity, as high nitrite levels can lead to toxic conditions for AOB, interfering with their metabolic processes [40]. Interestingly, the decline in NO₂⁻-N during the later stages suggested a possible shift in the metabolic activity of the strain towards denitrification. Increased nitrite reductase (NIR) enzyme expression in response to high NO₂⁻-N concentrations can enhance the conversion of nitrite to gaseous products such as N₂O and N₂ [41].

3.2. Effect of Temperature on Strain N.eA1

At 30 °C, the AOR and the NO₂⁻-N accumulation rate were significantly higher compared to the other two temperatures. The average AORs at the three temperatures were 3.13 mg∙(MLVSS∙h)⁻¹, 5.00 mg∙(MLVSS∙h)⁻¹ and 4.92 mg∙(MLVSS∙h)⁻¹, respectively. The maximum N₂O emission occurred between 24 and 36 h at 35 °C, reaching 1030.21 μg∙L⁻¹. The highest N₂O conversion percentage occurred at 24 h, after which the conversion percentage decreased. The total N₂O release concentrations after 48 h reached 1223.07 μg∙L⁻¹, 2406.48 μg∙L⁻¹ and 2699.36 μg∙L⁻¹, respectively (Figure 4).

Temperature could be one of the key factors contributing to N₂O emissions during nitrification and denitrification processes [42]. The maximum AOR occurs at 30 °C, indicating that increased temperature promoted NH₄⁺-N degradation by the AOB strain. However, the excessively high temperatures inhibited bacterial activity [43]. During the logarithmic growth phase (12–36 h), the bacteria exhibited a high demand for oxygen, which may have resulted in the incomplete oxidation of NH₂OH and the release of N₂O. Additionally, the high bacterial density during this phase likely contributed to a significant portion of N₂O production through the denitrification process. Furthermore, the apparent relationship between N₂O emissions and temperature has been investigated in several studies. The N₂O solubility decreases as temperature increases (being about two times lower at 25 °C than at 5 °C), therefore the liquid phase N₂O is more easily stripped into the gas phase, leading to the enhancement of N₂O emissions [44]. The peak N₂O conversion percentage was recorded at 24 h. The N₂O emission concentration peaked at 36 h, indicating that during this period (24–36 h), although N₂O emissions continued to rise, more NH₄⁺-N was being converted into NO, NO₂⁻-N, and NO₃⁻-N. It can also be seen from the change of nitrite concentration that the accumulation of nitrite increased significantly during this period. The maximum AOR also appeared in this period.

3.3. Effect of DO Concentration on Strain N.eA1

As shown in Figure 5, the consumption of NH₄⁺-N and the accumulation of NO₂⁻-N both increased with the rise in DO concentration. In the non-aerated control group, the NH₄⁺-N conversion percentage at 30 h was 9.8%, with a conversion amount of 14.8 mg·L⁻¹, and the accumulated NO₂⁻-N was 8.0 mg·L⁻¹. At DO concentration of 7.3 mg·L⁻¹, the 30 h conversion rate of NH₄⁺-N reached 45.7%, with a conversion amount of 68.8 mg·L⁻¹, and the accumulation of NO₂⁻-N was 35.7 mg·L⁻¹. The N₂O emissions during the aeration stage were significantly lower than those during the unaerated stage. During the unaerated stage, N₂O emissions increased over time. However, during the aeration stage, when the DO concentration exceeded 4.5 mg·L⁻¹, N₂O emissions peaked between 12 and 18 h and subsequently declined, suggesting that high DO concentrations (>4.5 mg·L⁻¹) could significantly reduce N₂O emissions while maintaining high ammonia oxidation efficiency. In the non-aerated control group, the N₂O release was relatively lower than in other experimental groups.

DO concentration simultaneously affects the rates of both nitrification and denitrification [45], and DO is considered an important parameter affecting N₂O emissions [46]. According to Park [47], the higher the DO concentration, the faster the accumulation rate of NO₂⁻-N and the lower the amount of N₂O released. In another study, Law [48] investigated how varying DO concentrations affected AOR and N₂O production in an enriched AOB culture. They observed minimal N₂O production at low DO levels (0.05–0.2 mg·L⁻¹), which increased as DO levels rose (2.4 mg·L⁻¹). This finding differs from our results, possibly due to different DO concentration settings. In low DO conditions (<2.4 mg·L⁻¹), increased DO promoted AOR, leading to more rapid production of intermediates (NH₂OH, NO). The accumulation of these intermediates resulted in increased N₂O production. In high DO conditions, sufficient oxygen favored complete oxidation, reducing the risk of intermediate accumulation and thereby suppressing the expression of denitrification-related enzymes, ultimately leading to decreased N₂O emissions. The DO control group exhibited higher N₂O accumulation and conversion percentage, as the low-oxygen environment induced denitrification by AOB, and the absence of the NoS encoding gene led to the accumulation of N₂O [49]. In the aeration stage, when DO concentration exceeded 4.5 mg·L⁻¹, the N₂O emissions decreased in the later stages. This may have been due, in part, to the more complete oxidation of N₂O at high DO concentrations, reducing the emission of some N₂O. Additionally, high DO concentrations limited the denitrification of AOB [50]. This further demonstrated that high DO concentrations were beneficial for reducing N₂O emissions in ammonia oxidation processes in AOB pure culture systems [51].

3.4. Effect of pH on Strain N.eA1

At pH 8.0, the degradation rate of NH₄⁺-N and accumulation rate of NO₂⁻-N reached their maximum values. Within 84 h, 98% of NH₄⁺-N was degraded, while NO₂⁻-N accumulated to 158.1 mg∙L⁻¹. N₂O emission peaked between 24 and 36 h, reaching 228.86 μg∙L⁻¹, with a conversion rate of 0.021%. At pH 9.5, the AOR decreased significantly. After 84 h, only 42.1% was degraded, with a conversion amount of 82 mg∙L⁻¹ and NO₂⁻-N accumulation of 39.9 mg∙L⁻¹. This indicates that high pH inhibits the strain’s ammonia oxidation capacity, resulting in significantly reduced N₂O emissions. The cumulative N₂O release was only 103.8 μg∙L⁻¹, with a maximum conversion rate of 0.017% (Figure 6).

The influence of pH on N₂O production during denitrification can be mechanistically explained through the structure of enzymes [52]. The NoS enzyme contains two copper centers (CuA and CuZ), where the electron transfer between these sites significantly impacts N₂O reduction efficiency. Within the pH range of 4–8, this electron transfer serves as the rate-limiting step in N₂O reduction [53]. Previous studies have established that maintaining pH between 7 and 8 during denitrification is optimal for minimizing N₂O emissions [52]. Consistent with previous research, we observed that NO₂⁻-N accumulation inhibited subsequent ammonia oxidation, likely due to its toxic effects on AOB and the consequent reduction in available substrate [54]. Notably, at pH 9.5, the N₂O emissions were significantly reduced and the conversion rate of N₂O was increased, exhibiting significantly reduced ammonia oxidation capacity. FA formation is favored at higher pH levels, which has been demonstrated to substantially inhibit bacterial activity and consequently reduce ammonia oxidation rates [55]. On the other hand, given that nitrification and denitrification pathways constitute the primary routes for N₂O production [56], elevated pH levels can compromise these processes through NOs enzyme inactivation and suppression of microbial activity [57].

3.5. Effect of NH₄⁺-N Concentration on Strain N.eA1

At initial NH₄⁺-N concentrations of 36, 71, 133 and 281 mg∙L⁻¹, the degradation amounts after 48 h were 34.8, 55.9, 81 and 73 mg∙L⁻¹, respectively, while NO₂⁻-N accumulated to 42.6, 50.6, 111.3, and 89.9 mg∙L⁻¹, respectively. Respective N₂O emissions at 36 h reached 111.91, 195.11, 142.08 and 249.02 μg∙L⁻¹, with emission peaks observed at 36 h across all concentrations (Figure 7). The experimental group with an initial NH₄⁺-N concentration of 281 mg∙L⁻¹ exhibited the highest ammonia oxidation rate of 5.07 mg∙(MLVSS∙h)⁻¹, corresponding to the maximum NO₂⁻-N accumulation. Calculations using Formulas (2)–(4) yielded theoretical FA values of 3.08, 6.24, 11.55, and 24.48 mg∙L⁻¹ for the four respective NH₄⁺-N concentrations.

At high NH₄⁺-N concentration (281 mg∙L⁻¹), the strain exhibited slightly higher AOR compared to those at 133 mg∙L⁻¹. Nitrosomonas europea can tolerate elevated NH₄⁺-N concentrations [58], where higher NH₄⁺-N levels promote increased ammonia oxidation rates, leading to enhanced AMO gene expression and NH₂OH accumulation, subsequently resulting in N₂O formation through oxidative pathways [59]. Peak N₂O emissions were consistently observed at 36–48 h across all experimental groups. This phenomenon can be attributed to both the logarithmic growth phase of the strain and substrate concentration dynamics [60]. N₂O production was relatively high during the initial reaction phase but declined significantly following substantial NH₄⁺-N consumption. AOB demonstrated higher activity during the early stages of the reaction; however, this activity gradually decreased due to limitations in nutrient availability and DO, consequently affecting N₂O production. Maximum N₂O conversion occurs at 36 h. Maximum AOR also occurs during this period (36–48 h). Notably, at an initial NH₄⁺-N concentration of 133 mg∙L⁻¹, the maximum AOR reached 63.12% of that observed at 281 mg∙L⁻¹. This finding provides valuable economic implications for N₂O emission reduction strategies.

3.6. Sensitivity Analysis of Environmental Factors to N₂O Emission

The Standardized Regression Coefficients (SRC) for each environmental factor were calculated to evaluate their relative impacts on N₂O emissions. The results are summarized in

Table 3. SRC for environmental factors affecting N₂O emission.

Serial Number	Environmental Factor	Standardized Regression Coefficient (SRC)
1	Temperature	0.62
2	pH	0.30
3	DO	0.85
4	NH4+-N	0.45

Note: The R² value for the regression model was found to be 0.946, indicating a strong fit of the model to the data and confirming the validity of the SRC values.

, showing the SRC values for temperature, pH, DO, and NH₄⁺-N concentration.

The sensitivity analysis reveals a distinct hierarchical control of environmental parameters over N₂O emissions in the isolated AOB strain, with DO (SRC = 0.85) emerging as the predominant regulatory factor. This pronounced dependence on DO aligns with enzymatically constrained ammonia oxidation pathways, where elevated O₂ levels accelerate hydroxylamine accumulation through AMO activity, thereby promoting N₂O production via nitrifier denitrification [52]. Temperature exhibits secondary yet substantial influence (SRC = 0.62), displaying Arrhenius-type kinetic behavior between 25 and 35 °C, reflecting the thermal acclimation of ammonia mono-oxygenase [61,62]. The high model determination coefficient (R² = 0.946) underscores the robustness of these univariate relationships, providing a solid theoretical foundation for practical operations.

Thus, maintaining DO between 4.5 and 7.3 mg∙L⁻¹ is beneficial to reduce N₂O emissions without AOR reduction. This could be achieved via high-precision DO sensors and adaptive aeration. Thermal management should consider the non-linear temperature effects on N₂O emissions. Although increasing the temperature from 25 °C to 30 °C can enhance nitrogen removal efficiency (ΔAOR = +38% in this study), continuous operation above 35 °C may destabilize emission pathways.

4. Conclusions

In this study, we found that the N.eA1 strain demonstrated high homology with Nitrosomonas europaea H1 AOB3, and exhibited comparable growth kinetics patterns. The N.eA1 strain showed a higher risk of N₂O emission than that of Nitrosomonas europaea at similar NO₂⁻-N concentration. Environmental factors significantly affected the N₂O emission and ammonia oxidation rate of N.eA1; the conditions helpful to the AOR were pH maintained at 7–8, temperature kept at 25–30 °C, and influent NH₄⁺-N concentration maintained above 133 mg∙L⁻¹.

Additionally, the control of DO concentration is crucial. DO concentrations (4.5–7.3 mg∙L⁻¹) promoted enhanced ammonia oxidation while simultaneously reducing N₂O emissions. In the process of nitrogen conversion under different environmental conditions, peak N₂O emissions consistently occurred during the logarithmic growth phase (12–36 h), but the peak of maximum conversion of N₂O was earlier than this time. The maximum AOR time always occurs in this phase (between the timing of maximum N₂O conversion and maximum N₂O emissions). This finding helps us understand the timing of N₂O emissions and nitrogen conversion. However, several limitations remain. This study was conducted under controlled laboratory conditions, and the effects of field-scale variations in environmental parameters (e.g., fluctuations in DO concentrations over time) were not assessed. Additionally, the specific metabolic pathways contributing to elevated N₂O emissions in the N.eA1 strain, particularly in comparison to Nitrosomonas europaea, require further investigation. Future research should focus on the interactions between multiple environmental factors and explore the role of microbial community dynamics in shaping N₂O emission patterns and AOR. Given that microbial consortia may exhibit distinct behaviors compared to isolated strains, we aim to extend these findings to mixed AOB communities, ultimately contributing to the development of more efficient and environmentally sustainable nitrogen removal technologies.