# Millimeter-scale niche differentiation of N-cycling microorganisms across the soil-water interface has implications for N2O emissions from wetlands

Yu-Jia Cai, Hong-Yang Zhang, Xiao-Ran Hu, Yu-Chen Yang, Christina

Hazard, Graeme Nicol, Ji-Zheng He, Ju-Pei Shen, Zi-Yang He, Lu Zhang, et al.

► To cite this version:

Yu-Jia Cai, Hong-Yang Zhang, Xiao-Ran Hu, Yu-Chen Yang, Christina Hazard, et al. Millimeter-scale niche differentiation of N-cycling microorganisms across the soil-water interface has implications for N2O emissions from wetlands. The International Society of Microbiological Ecology Journal, 2025, 19 (1), pp.109791. 10.1093/ismejo/wraf062. hal-05409044

HAL Id: hal-05409044

https://hal.science/hal-05409044v1

Submitted on 10 Dec 2025

HAL is a multi-disciplinary open access
archive for the deposit and dissemination of sci-
entific research documents, whether they are pub-
lished or not. The documents may come from
teaching and research institutions in France or
abroad, or from public or private research centers.

L'archive ouverte pluridisciplinaire HAL, est
destinée au dépôt et à la diffusion de documents
scientifiques de niveau recherche, publiés ou non,
émanant des établissements d'enseignement et de
recherche français ou étrangers, des laboratoires
publics ou privés.

<figure><img src="image_1.png" /></figure>
# Millimeter-scale niche differentiation of N-cycling microorganisms across the soil-water interface has implications for N₂O emissions from wetlands

Yu-Jia Cai<fnref n="1" />,<fnref n="2" />, Hong-Yang Zhang<fnref n="1" />, Xiao-Ran Hu<fnref n="1" />, Yu-Chen Yang<fnref n="1" />, Christina Hazard ID<fnref n="3" />, Graeme W. Nicol<fnref n="3" />, Ji-Zheng He<fnref n="4" />,<fnref n="5" />, Ju-Pei Shen<fnref n="4" />, Zi-Yang He<fnref n="5" />, Lu Zhang<fnref n="1" />, Jing-Hui Zhang<fnref n="1" />, Hao Liu<fnref n="1" />, Sha Zhang<fnref n="1" />, Zheng Chen ID <fnref n="1" />,∗

## Abstract

Wetlands can be a significant source of N₂O under current global climate change regime with the soil-water interface representing a biogeochemical hotspot for microbial activity. However, the role of soil-water interface in controlling N₂O emissions remains poorly understood. We hypothesized that the millimeter-scale redox gradient across the soil-water interface generates corresponding distinct niche for N-cycling microorganisms that collectively regulate the production and consumption of N₂O over the same spatial scale. The abundance, transcriptional activity and spatial organization of different N-cycling guilds across the soil-water interface were characterized in mesocosms from three different paddy soils with different N₂O emissions. Results demonstrated millimeter-scale stratification of N-cycling microbial activity across the soil-water interface, and in particular within the first 10 mm of flooded soils. Ammonia-oxidizing microorganisms were only transcriptionally active in the top 4 mm, suggesting a previously underestimated contribution to N₂O emissions from wetlands. Variation in N₂O accumulation was observed across the soil-water interface, with the highest concentrations measured at either the soil-water interface or in the deeper anoxic layer of paddy soils. Despite this difference, N₂O-reducing microorganisms exhibited high transcriptional activity at the soil-water interface in all soils, suggesting that there is a microbial-mediated sink for N₂O across the soil-water interface that can reduce N₂O produced from both oxic and anoxic layers. This work demonstrate an underappreciated and essential role of the microbial hot zones at soil-water interface in regulating N₂O emissions from wetlands.

**Keywords:** soil-water interface; Millimeter-scale; N₂O emissions; wetlands; nitrogen cycling; niche partitioning; ammonia oxidizing microorganisms

## Introduction

Microbial communities contribute to the regulation the greenhouse gas (GHG) emissions from wetland soils [1–3]. Although wetlands cover only 8% of the Earth’s terrestrial surface, they represent one of the largest organic carbon stocks [3, 4]. Warming and hydrological fluctuation associated with a changing climate may enhance microbial degradation of organic carbon, and increase GHG emissions from wetlands [3, 5–8]. Among the GHGs emission from wetlands, N₂O, a potent GHG and a depletor of stratospheric ozone layer, has received less attention compared to methane and CO₂. However, fluxes of N₂O from wetlands, such as paddy fields, riparian and intertidal zones, are highly variable in both space and time due to the complexity of N-cycling processes and N₂O dynamics [3, 9–11]. Production of N₂O occurs through microbial-driven pathways during both nitrification- and denitrification-related processes [12, 13]. The extent that these different

pathways contribute to N₂O production largely depends on the microbial community and environmental conditions. For example, nitrification-related N₂O dominates in drier and aerated upland soils [12, 14] whereas denitrification-derived N₂O production dominates in soils with high or saturated water content [15–18].

Recent studies have suggested that nitrification, particularly the role of Ammonia-oxidizing microorganisms (AOM) in wetlands, have been underappreciated [3, 19, 20]. Aerobic chemolithoautotrophic AOM consist of ammonia-oxidizing archaea (AOA), ammonia-oxidizing bacteria (AOB), and the more recently discovered complete ammonia oxidizers (comammox Nitrospira) [21, 22]. AOM can thrive at the thin oxidized layer of surface wetland soils or in the rhizosphere of vascular plants [23–25]. During ammonia oxidation N₂O is produced via both biotic and abiotic mechanisms [26–28]. AOB can also produce N₂O

<footnote n="1">¹Department of Health and Environmental Sciences, Xi'an Jiaotong-Liverpool University, 111 Ren'ai Road, Suzhou, Jiangsu Province 215123, China</footnote>

<footnote n="2">²Department of Geography and Planning, School of Environmental Sciences, University of Liverpool, Brownlow Hill, Liverpool L697ZX, United Kingdom</footnote>

<footnote n="3">³Université Claude Bernard Lyon 1, CNRS, INRAE, VetAgro Sup, Laboratoire d'Ecologie Microbienne, Villeurbanne 69622, France</footnote>

<footnote n="4">⁴Key Laboratory of Humid Subtropical Eco-geographical Process of Ministry of Education, School of Geographical Sciences/School of Carbon Neutrality Future Technology, Fujian Normal University, Fuzhou, Fujian Province 350007, China</footnote>

<footnote n="5">⁵School of Agriculture, Food and Ecosystem Sciences, Faculty of Science, The University of Melbourne, Parkville, VIC 3010, Australia</footnote>

<footnote>*Corresponding author. Department of Health and Environmental Sciences, School of Science, Xi'an Jiaotong-Liverpool University, 111 Ren'ai Road, Suzhou, Jiangsu Province, 215123, China. E-mail: ebiogeochem@outlook.com, Zheng.Chen@xjtlu.edu.cn</footnote>
through the denitrification pathway [12, 20]. N₂O that is produced
can be directly released to the atmosphere as AOM reside in
surface soils. Other N₂O producing pathways in wetlands include
incomplete denitrification and chemodenitrification [12, 29, 30],
adding to the intricate and diverse means of N₂O production.

In wetlands, water flooding reduces the aerobic layer in soil and generates a soil-water interface (SWI) that bridges the oxic overlaying water with the anoxic soil compartment. Due to the slow oxygen diffusion from the atmosphere to water and high oxygen consumption by microorganisms, a distinct oxygen gradient forms across the SWI [31–34]. When oxygen is depleted in soils microorganisms use alternative electron acceptors such as nitrate (NO₃⁻), iron, manganese, and sulfate during respiration [35]. Such oxygen gradient can lead to the niche differentiation of nitrifiers and denitrifiers. Denitrification therefore occurs in deeper anoxic layer, with aerobic nitrification favored at the SWI where oxygen is relatively abundant [25]. Two zones for N₂O production may exist in flooded soils, consequently creating a niche for N₂O-consuming microorganisms across the SWI.

Oxygen and other biogeochemical gradients across the SWI generally manifest at micro- or millimeter scales [31, 32, 34, 36]. Heterogeneity in microbial communities at these scales may also be present but this is often obscured by typical bulk sampling approaches [37–39]. Considering the size of prokaryotes and the scale at which cellular interactions can occur, millimeter-scale is a biologically relevant and feasible scale to study the activities of microbial communities across the SWI [37, 38]. Although microbiome analyses have been successfully performed at this scale in soil crust [40], microbial mats [41, 42], and sediment [43], less has been done across the SWI [38].

This study aimed to examine the potential partitioning of N₂O production pathways across the SWI to gain insight into N cycling and N₂O dynamics in wetlands. It was hypothesized that the redox gradient across the SWI generates different niches for N-cycling microorganisms that influences the production and consumption of N₂O across the SWI. A flooded mesocosm experiment was performed to profile the vertical distribution of N-cycling microorganisms across the SWI at the millimeter-scale. Utilizing our previously developed approach to effectively sample at the millimeter-scale [38], we assessed microbial community structure, abundance of N-cycling microorganisms, and transcriptional activity vertically at 2 mm spatial resolution. With incorporation of in-situ and fine-scale sampling methods for soils and sediments [44–46], spatial and temporal heterogeneity in soil water, gas, and solid phase were determined, which enabled distinguishing N-cycling processes occurring across the SWI.

Materials and methods
Soil collection and characterization

Wetland soils including paddy soils were collected from ten sites
across China in 2017 (Table S1). The upper 20 cm layer of soil was
collected. Soil samples were air-dried, sieved (2 mm) for homog-
enization, and preserved at room temperature. A subsample of
each soil was used for soil physicochemical analyses (Table S1).
Soil pH was measured in a soil suspension at a soil:water ratio
of 1:2.5. Soil NH₄⁺, NO₂⁻, and NO₃⁻ -N were extracted with
0.5 M K₂SO₄ and determined using colorimetric assays with a
microplate reader (Tecan, Austria) [47].

A preliminary soil microcosm was carried out to examine
the N₂O emissions of the ten wetland soils. Three paddy soils,
Kunshan (KS), Shaoguan (SG), and Wenshan (WS) (Table S1), were
selected for the subsequent soil mesocosms experiment. Due

to the quantity of soil required for these experiments, KS and
WS paddy soils were re-collected according to the original GPS
information in February 2023 and May 2023, respectively. Soil
physicochemical analyses were measured as previously described
(Table S2).

Soil microcosm experiment to determine N₂O emissions from wetland soils

Soil microcosms were established in triplicate under flooded conditions to determine N₂O emissions from the ten wetland soils. Briefly, each 40 mL glass vial was filled with 10 g dry soil and ultrapure water to form a 3 cm water layer and an 18 mL headspace air. Microcosms were sealed with breathable sealing film and incubated in the dark at 25°C for 45 days. 1 mL headspace air was sampled for N₂O analysis every week.

Soil mesocosm experiment with selected soils

Of the ten soils tested, two demonstrated high rates of N₂O emissions (KS and WS) and were used in a subsequent mesocosm experiment with a non N₂O-emitting soil (SG) to examine N₂O emissions across the SWI (Table S1). Soil was added into lightproof pot containers (12 cm length × 12 cm width × 15 cm height) to form a 10 cm soil layer (approximately 1300 g). Deionized water was added to the soil until saturation with a 3 cm layer of standing water. Three replicate pots were established for WS and SG soil, and four replicate pots for KS soil. Mesocosms were placed in an automated platform (Fig. S1) and covered with aluminum foil to exclude light during incubation. Room temperature was maintained at 25°C for the duration of the experiment. Water levels were checked twice a week and adjusted if needed to maintain the standing water layer. The redox potential (Eh) gradient across the SWI was determined every week using an Eh microelectrode. For KS soil, when the redox gradient across the SWI reached 400 mV after 38 days of incubation, the four mesocosms were randomly divided into two groups for light–dark cycling to imitate the natural environment. One group, designated as “light–dark”, was subjected to a 12 h light- 12 h dark cycle, whereas the other group “all–dark” was kept in constant darkness. The light–dark cycling was maintained for 6 days to prevent the formation of phototrophic microbial biofilm. At the end of the incubation (44 days for KS, 40 days for SG and WS soils), soil cores within each mesocosm were destructively sampled for analyses of soil chemistry (e.g., NH₄⁺ and NO₃⁻), microbial community structure, and transcriptional activity.

Determination of N concentrations and N₂O
fluxes across SWI

Soil NH₄⁺, NO₂⁻, and NO₃⁻ -N concentrations were determined in either the porewater or soil extracts with soil porewater collected using the integrated porewater injection (IPI) sampler [44, 46] (Detailed in Supplementary text). The IPI sampler, allowing in situ and continuous sampling, was used in the KS soil to monitor the change of dissolved NH₄⁺, NO₂⁻, and NO₃⁻ -N across the SWI during incubation. Whereas for SG and WS soils, soil cores were sampled for N analysis at the end of incubation. Briefly, soil cores were sampled using a 20 mL modified syringe, carefully inserted into the soil to prevent disturbance of the SWI before withdrawing to remove a core [38]. After sampling, the soil core was pushed out at 2 mm intervals and slices removed using a sterile ceramic knife. The 2 mm soil slice were put to inorganic N extraction at a soil: 1 M KCl ratio of 1:5.

Soil gases (e.g., N₂O and CO₂) were analyzed after diffusing
into silicone tubing (5 mm outer diameter × 115 mm length). The
silicone tubing was attached to the inside of the pot using silicone sealant while isolating the water [48–50]. A gas tight syringe was inserted into the silicone tubing and 1 mL gas was collected. The spatial resolution for the soil N₂O profile was 10 mm.

To determine N₂O fluxes in the mesocosms, gas was collected once or twice a week using the static chamber technique with transparent polymethylmetacrylate (PMMA) chambers (10 cm length × 10 cm width × 6 cm height). During sampling, the over-laying water within each mesocosm seals the chamber (Fig. S1C). Preliminary tests showed that a 2 h interval was appropriate for sealing chambers to measure N₂O emissions (data not shown). During the 12 h light-12 h dark cycling, the N₂O flux measurement was performed 6 h into the lighting period of incubation every day.

The N₂O flux was calculated using linear regression analysis [51, 52]:

$$F = \left( \frac{\Delta c}{\Delta t} \right) \times \frac{V}{22.4} \times \left( \frac{273.15}{273.15 + T} \right) \times \frac{M}{A}$$

Where F is the N₂O flux (ng N h⁻¹ dm⁻²), Δc/Δt is the rate of N₂O accumulation in the chamber (nmol mol⁻¹ h⁻¹), V is the effective sampling volume of the chamber (L), 22.4 is the molar gas constant at 0°C (L mol⁻¹), T is the temperature in Celsius (°C), M is the molecular weight of N₂O-N (g mol⁻¹), and A is the surface area of the chamber (dm²).

For N₂O analysis, 1 mL of gas sample was injected into a GC fitted with an ECD (Agilent 7890B, USA). Carboxen 1010 columns were kept at 150°C (oven temperature), and ultrapure N₂ was used as carrier gas. The ECD current was 250 mV, and the ECD cell was kept at 300°C.

## N-cycling microbial community structure and transcriptional activity across the SWI

Approximately 1.0 g per soil slice (2 mm) was used for immediate RNA extraction and another 0.4 g was stored –20°C for subsequent DNA extraction. Total RNA was extracted using the PowerSoil Total RNA Isolation Kit (Qiagen, USA) according to the manufacturer's instructions. Removal of genomic DNA and reverse transcription were performed with the PrimeScript RT reagent kit with gDNA Eraser (Takara, Japan). DNA extraction was performed using the DNeasy PowerSoil Kit (Qiagen, USA), following the manufacturer's instructions. Nucleic acid concentration and purity were determined using a NanoDrop One spectrophotometer (Thermo Scientific, USA).

Quantitative PCR (qPCR) was conducted in a Light Cycler 480 II (Roche, Switzerland) with three technical replicates per sample. Primer sets used for qPCR of N-cycling and 16S rRNA genes are given in Table S3. For the nitrite-oxidizing bacteria (NOB), the primer set nxrB169f/nxrB638r was designed based on Nitrospira nxrB gene and unlikely detect other NOB here due to the dominance of Nitrospira NOB in low-NO₂⁻ and low-oxygen environments (e.g., wetland) [53, 54]. The reaction mixture (10 µl per sample) consisted of 5 µl TB Green premix (Takara, Japan), 0.5 µl for forward and reverse primer (final concentration 0.5 µM), 1.5 µl total cDNA/DNA, and 2.5 µl DNase-free water. Standard curves for quantification were generated using 10-fold serial dilutions of plasmid DNA containing a fragment of each target gene. Amplification efficiency of standards ranged from 95%–110% with an R² > 0.99. Melt curve analysis and gel electrophoresis were conducted to assess the specificity of qPCR products.

## Statistical analysis

Pearson correlation analysis was performed for the abundance of N cycling microorganisms using the R packages "Hmisc" [55] and

"corplot" [56]. ANOVA and t-test were performed using IBM SPSS Statistics (Version 29.0.0.0).

## Results

### N₂O emission and depth profiles vary in wetlands

The emissions of N₂O from the wetland soil samples, collected from rice paddies, coastal intertidal wetland, and other wetlands across China, were measured after 45 days of incubation under water-flooding conditions (Table S1). Two soils that produced detectable N₂O emissions (KS and WS) and one soil (SG) that did not were selected for the subsequent mesocosm experiments.

Various parameters (Eh, N₂O, NH₄⁺, NO₂⁻, and NO₃⁻) were measured across the SWI (from the SWI to the 50 mm soil depth) at the end of the incubation experiment (Fig. 1, S2, and S3). The Eh dropped at ~300 mV from the water layer to the ~10 mm soil layer in all three soils (Fig. 1A), representing a steep redox gradient across the SWI at the millimeter scale. Based on this redox gradient, the soil was divided into 3 layers: an oxic layer (0–2 mm), a transition layer (2–10 mm), and an anoxic layer (below 10 mm).

During the incubation, only the KS and WS soils produced N₂O with emission rates peaking with 7.00 ± 2.10 and 1.32 ± 0.70 µmol mol⁻¹ h⁻¹, which then decreased to 1.11 ± 0.50 and 0.13 ± 0.08 µmol mol⁻¹ h⁻¹ after 40 days, respectively (Fig. 2). The depth profiles of N₂O concentrations varied with different N₂O accumulation zone across the SWI. N₂O was more accumulated in the 20–40 mm soil layer of KS soil, whereas in WS soil the accumulation occurred at the SWI, and no accumulation was found in SG soil (Fig. 1B).

In the flooded soils, the concentration of NH₄⁺ was greater than that of NO₂⁻ and NO₃⁻ across the depth profile (Fig. S2). Concentrations of NH₄⁺ increased with depth in KS and WS soils (Fig. S2A). Temporal change of dissolved NH₄⁺ profile was tracked in KS soil using the IPI sampler (detailed in Supplementary Material). The gradient of NH₄⁺ concentrations across the SWI developed and reached its steepest slope at day 25, then decreased to 0.5–3.5 mg L⁻¹ (Fig. S2A and S3), which was higher than the original NH₄⁺ content in bulk soil.

## Abundance of nitrifier functional genes and transcripts across the SWI

The quantification of nitrifier abundance and transcriptional activity indicated that nitrification primarily occurred within the top 4 mm (Fig. 3A-C and 4A-C). The AOA ammonia monooxygenase sub-unit A (amoA), AOB amoA, and Nitrospira nitrite oxidoreductase beta sub-unit (nxrB) genes and transcript abundances were highest in the top 0–2 mm surface soil (P < 0.001), with mRNA transcripts only detected in 0–4 mm layers in KS and WS soils, and no nitrification transcriptional activity detected in SG soil profile. No depth-related trends were observed for total prokaryotic 16S rRNA gene and transcript abundances (Fig. S4). Among the three soils, nitrifiers were most abundant in WS soils, followed by KS and SG soils (F = 59.2, P < 0.001).

## Abundance of NO₃⁻ and NO₂⁻ reductase genes and transcripts across the SWI

We quantified marker genes for denitrifiers, including those encoding the membrane-bound and periplasmic NO₃⁻ reductase sub-units (narG and napA), as well as the cytochrome cd1-containing and copper-dependent NO₂⁻ reductase sub-units (nirS and nirK). Unlike the nitrifiers, KS soil harbored more NO₃⁻ and NO₂⁻ reducing microorganisms followed by WS and
<figure><img src="image_2.png" /><figcaption>**Figure 1.** Depth profiles of eh (A) and N₂O (B) across the soil-water interface at the end of incubation experiment. Additional graphs showing the eh gradient change during incubation of KS soil can be found in Supplementary Fig. S4A. Different symbols in (A) and shades of the color in (B) represent independent replicates.</figcaption></figure>

SG soil (F = 190.3, P < 0.001). These denitrifier associated genes demonstrated different distribution patterns compared to those of nitrifiers, as well as differed within their N-cycling functional equivalent and distinct groups (Fig. 3, 4, and S5). Changes in transcript abundance of *narG* and *napA* genes demonstrated similar depth-dependent patterns in KS and WS soil profiles, with *napA* gene more transcriptionally active in the transition zone (Fig. 3D and E). No spatial changes in gene abundance were observed (Fig. 4D and E).

For NO₂⁻ reducers, *nirK* gene and transcripts were generally lower (or absent) throughout the profile compared to *nirS* (Fig. 3F,G and 4F,G). The transcriptional activity of *nirS* and *nirK* genes varied in different soil profiles (Fig. 3F,G and S5G,H). Transcripts of *nirS* were more abundant at 2–6 mm in KS soil but decreased with increasing soil depth in WS soil (Fig. 3F and S5G). The transcripts of *nirK* guild were only detected at the SWI of KS soil, and were below detection in SG and WS soil profiles.

## N₂O-consuming microorganisms across the SWI

The N₂O reductase (NosZ) genes are separated into two distinct clades and are associated with different functional roles that

clade I microorganisms are primarily denitrifiers, whereas clade II are non-denitrifying pure N₂O reducers [57]. The abundance of N₂O-reducing microorganisms varied significantly (F = 711.7, P < 0.001) in the order WS > SG > KS soil, but corresponding order was not found in their transcriptional activity with few active N₂O reducers found in SG soil. Both nosZ clades were transcriptionally active throughout the SWI in KS and WS soils, and were more active closer to the SWI (Fig. 3H,I and S5I,J). No obvious depth-related changes in gene abundance were observed. The two clades exhibited divergent transcriptional patterns across the SWI. In KS soil, clade I nosZ transcripts were detected throughout the profile, whereas the activity of clade II was restricted to the first 6 mm. In contrast, the transcriptional activity of both clades decreased with increasing depth in WS soil, with the highest activity observed at the SWI.

## Effect of light exposure on N-cycling microorganisms at the SWI

To investigate the response of N-cycling microorganisms at the SWI to light exposure, KS soil mesocosms were incubated under a diurnal cycle for 6 days at day 38. Relatively lower N₂O emission
<figure><img src="image_3.png" /><figcaption>**Figure 2.** N₂O flux of the (A) Kunshan (KS), (B) Shaoguan (SG) and (C) Wenshan (WS) soil during the soil incubation experiment. In (A) KS soil, the four mesocosms were incubated in the dark and can be considered as replicates until day 38 before separated into the all-dark (blue) and light–dark (red) groups. The N₂O flux during light exposure was zoomed in on the top of (A).</figcaption></figure>

were observed during light exposure (Fig. 2A), concurrent with increased growth and activity of N-cycling microorganisms at the SWI (Fig. S5). Microbial responses to light exposure varied in terms of gene abundances and transcriptional levels. For example, after 6-day light–dark cycling, a congruent stimulation of both gene

abundances and transcriptional levels in AOA, AOB, *napA* NO₃⁻-reducers, NO₂⁻-reducers, and clade I *nosZ* N₂O reducers were observed (Fig. S5). Clade II *nosZ* N₂O reducers were only stimulated at the transcriptional level. Light exposure had no effect on *Nitrospira* *nxrB* and *narG* NO₃⁻ cohorts. A greater number of significant positive correlations were found between the abundances of N-cycling microorganisms in the light–dark group compared to the all-dark group (Fig. S6).

## Discussion

The millimeter-scale variation of N-cycling microbial community confirmed our hypothesis that distinct spatial distributions of active N-cycling microorganisms exist across the narrow SWI, suggesting their collective impact in regulating the N₂O emission from wetland soils (Fig. 5). Alike, the vertical stratification of microbial communities at millimeter scale is observed in diverse ecosystems, as evidenced by studies of paddy soil [33], soil crust [40], microbial mat [41, 42], and sediment [43]. Whereas these works focused on carbon, sulfur cycling and total microbial community shifts, we emphasized on the novel insights into functional redundancy and N₂O emission through these transcriptional stratification of N-cycling genes.

## Role of SWI in N₂O emission from wetland

According to the currently known N₂O producing pathways, we postulated that N₂O would be produced in deeper anoxic zones where it is suitable for denitrification, or at the SWI where nitrifiers were predicted to thrive. In the N₂O producing soils, N₂O accumulated in deeper anoxic zone of KS soil and at the SWI of WS soil. Such accumulation of N₂O was also found in biofilm [58–60] and a Danish wetland [61], where the oxygen gradient manifests at micrometers and millimeters. Correspondingly, the activity of AOM were restricted at the SWI together with low ammonium concentrations implying that there was nitrifier-related N₂O production in this zone. Nitrifiers produce N₂O via two pathways: through ammonia oxidation and nitrifier denitrification by AOB and NOB [12, 14, 27, 62]. Intriguingly, *nirK* NO₂⁻-reducers were more active at the SWI, relative to the AOM. The similar spatial distribution between *nirK* transcript and the activity of nitrifying microorganisms may be attributed to the close evolutionary history of *nirK* orthologue gene among nitrifiers [63]. In metagenomics studies, the *nirK* gene has been found in AOA and NOB from marine systems [26, 64, 65]. The catalytic product of NO₂⁻-reductase—NO is not only an essential intermediate during ammonia oxidation [66], but also a signaling molecule that enhances the mutualistic interaction between AOA and *Nitrospira* [54].

The N₂O accumulation at the SWI of WS soil was related to active N cycling microorganisms. However, the activity of denitrifying populations at the end of the incubation was not within the N₂O accumulation zone in KS soil. We speculate that this might be attributed to the 1) distinct spatial distribution of NO reducers compared to NO₂⁻- reducers [62]; 2) contribution of abiotic N₂O production such as chemodenitrification [30]; 3) incomplete denitrification during the formation of the redox gradient and peak N₂O emissions, and 4) low activity of N₂O-consuming microorganisms. Moreover, *nir* genes may not reflect activity of upstream or downstream processes of denitrification and therefore cannot be used as a proxy for all N oxide reductive processes [67]. During N₂O formation, NO production is usually concomitant with N₂O [12, 29, 58]. However, current primers cannot be used to detect and quantify *norB* genes of NO reducers from a diverse range
<figure><img src="image_4.png" /><figcaption>**Figure 3.** Abundance of transcripts across the soil-water interface of genes encoding for N-cycling processes. Error bar represents the standard error of the mean of technical replicates of KS soil microcosm and biological replicates (n = 3) of SG and WS mesocosms. No biological replicates are available for KS soil. Additional graphs showing the transcript abundance of N-cycling genes in the KS 0–50 mm soil profile are presented in *Supplementary Fig. S6*.</figcaption></figure>

of taxonomic groups [68]. Current findings have limitation in distinguishing the source of N₂O from the wetlands and the contribution of different N-cycling guilds, which may be addressed in future research with the application of stable isotope techniques and omics-based approach.

Combining the observed N₂O accumulation across the SWI, SWI showed potential in reducing N₂O emission through the highly active N₂O-reducing microorganisms in the top 10 mm soil layer or at the SWI. This spatial separation suggests that these microorganisms act as a microbial N₂O sink, consuming N₂O either diffused from deeper soil or produced by nitrifier metabolism. A similar stratification between N₂O and N₂O-consuming microorganisms has been found in oxic marine water, where N₂O consumption rates are higher compared to the anoxic zone [65, 67]. In addition, these aerobic N₂O consumers can consume more N₂O once shifted to anoxic conditions [65, 69–71]. This rapid N₂O consumption under oxic condition is surprising, as it is typically favored in anoxic environments, suggesting that oxygen plays a key regulatory role. Studies have demonstrated that *nosZ* gene expression is regulated by oxygen level, and N₂O reduction characterizes a life strategy for aerobic microorganisms to survive temporary anoxia [69, 71, 72]. Oxygen tolerance in N₂O

consumers is not related to the phylogeny or the structure of nitrous oxide reductase (NOS), but depends on their ability to quickly scavenge oxygen and/or the activity of accessory proteins encoded by the *nos* cluster, which maintain low oxygen levels to prevent NOS inactivation [73]. Though aerobic N₂O reduction is promising, it remains challenging in practical applications [74]. Further research is needed to investigate the oxidative damage to denitrification reductases using detailed proteomic analyses [71]. Overall, the balance between nitrifiers and N₂O-consuming microorganisms at the SWI plays a critical role in controlling N₂O emission from flooded wetland soils, highlighting the underappreciated importance of the SWI.

## Niche differentiation of the N-cycling microorganisms across the SWI

N-cycling microbial activity is stratified at the millimeter-scale across the SWI, with most activity restricted within the top 10 mm of the flooded soils, and particularly at the SWI (Fig. 4). These findings are observed in biofilm [60] and microbial mat [75], yet rarely found in analogous environments due to the trade-off between fine-scale soil/sediment sampling and active microbiome analysis. The spatial variations reflect the physiological
<figure><img src="image_5.png" /><figcaption>**Figure 4.** Abundance of genes across the soil-water interface encoding for N-cycling processes. Error bar represents the standard error of the mean of technical replicates of KS soil mesoocosm and biological replicates (n = 3) of SG and WS mesocosms. No biological replicates are available for KS soil. Additional graphs showing the gene abundance of N-cycling genes in the KS 0–50 mm soil profile are presented in Supplementary Fig. S6.</figcaption></figure>

features of different N-cycling guilds. For example, the ammonia oxidizers, which require oxygen for energy conservation, thrived at the oxygen-rich SWI, whereas denitrifiers preferred the transition layer in the flooded soils.

Niche differentiation exists in functional equivalent denitrifiers. For example, the *narG* guild predominated in NO₃⁻ reducers, which is also observed in estuarine sediment or flooded paddy soils [76, 77]. The prevailing transcriptional activity through the soil profile confirms NO₃⁻ respiration one of the ancient energy-conserving pathway in microorganisms, and the NO₃⁻ reductase involved are biochemically distinct [78, 79]. The membrane-bound NO₃⁻ reductase can provide energy through the proton motive force during NO₃⁻ reduction, whereas periplasmic NO₃⁻ reductase does not [80]. High NO₃⁻ concentrations seem favorable for the *narG* microbial cohorts, whereas populations possessing *napA* prefer low NO₃⁻ concentrations and oxic conditions [79, 81, 82].

The *nirS* guild was observed as the dominant NO₂⁻ reducing microorganisms in the flooded soils. Genomic analysis and experimental results suggest that the *nirS* guild dominates over the *nirK* guild under conditions that are favorable for denitrification [83–85]. Similarly, their transcriptional activity was dominant across the SWI whereas *nirK* transcripts were only detected at

the SWI. Niche partitioning between *nirS* and *nirK* NO₂⁻ reducers may be influenced by their tolerance to oxygen with *nirS* more sensitive to oxygen than the *nirK* carriers [83, 86–89].

## Influence of day-time light on activity of N-cycling microorganisms at the SWI

Light exposure affected the abundance and transcriptional activity of N-cycling microorganisms and slightly decreased N₂O emissions, but did not influence N₂O accumulation in the flooded soils. More significant correlations were found in the light–dark group, suggesting enhanced microbial interactions within the N-cycling microbial community under light. Studies have revealed the difference in N₂O emissions under light exposure, which may be related to the diurnal change in microbial activity, and confirmed it a prevalent phenomenon across terrestrial ecosystems, including agricultural, forest, and wetland soils [90–92]. The underlain mechanisms remain less studied, but recent research from riverine water columns reported strong inhibition of N₂O emissions with increasing light irradiation, potentially attributed to the change in N-cycling microbial activity [93]. However, the robustness of the stimulation effect on N-cycling microorganisms
<figure><img src="image_6.png" /><figcaption>**Figure 5.** Schematic description of active N transformation processes across the soil-water interface. The size of the N species label indicates relative differences in concentration. Solid arrows indicate microbial transformations and dotted arrows indicate N diffusion or exchange. Microbial guilds possessing the same gene type are described with the size of the label indicating the relative difference of guild activity. *nxrB* represents Nitrospira microorganisms which thrive in low-NO₂⁻ and low-oxygen environments [54]. Lightning symbol next to the label indicates that the microbial N cohort is sensitive to light exposure.</figcaption></figure>

here requires further experimental confirmation with more repli-
cates. The biological replicates are limited in KS soil as there was
a trade-off between requirements of RNA extraction and massive
sample number when investigating the SWI at fine-scale.

These light-driven changes in microbial community structure
and activity could be associated with the formation of photo-
chemically produced reactive intermediates (e.g., singlet oxygen
(¹O₂), peroxide hydrogen (H₂O₂), and hydroxyl radicals (·OH)) [93].
The oxygen gradient and redox oscillation makes the SWI a
previously unappreciated hotspot for production of reactive oxy-
gen species, influencing microbial activity and N-cycling [94, 95].
Though reactive oxygen species can inhibit or kill cells, light
exposure in this study stimulated the N-cycling microbial activity
and interaction, indicating microbial response to oxygen stress
is dose- and exposure-time dependent [96]. Metatranscriptomics
may help to examine whether microorganisms at the SWI are
coping with oxidative stress in the daytime.

Conclusion

This study provides a detailed perspective on the dynamic pro-
cesses of nitrification and denitrification across the SWI at the
millimeter scale. Sampling at an appropriate spatial scale may
help clarify the ecological processes driven by microbial activities
and lead to more reliable inferences of the functional capability of
microbiomes [37, 97, 98]. Findings demonstrate that the SWI is an
ideal environment to disentangle the complexities of N₂O emis-
sions from wetlands. The SWI may harbor an underappreciated

"microbial N₂O barrier", where the balance between nitrification-
related N₂O production and N₂O consumption collectively con-
trols the release of N₂O to the atmosphere. The SWI in wetland
systems characterizes as a fine-scale, sensitive, and represen-
tative oxic-anoxic environment in where soils are subjected to
intermittent or permanent water flooding. Drainage, warming,
and land use change of wetland soils disturb the SWI, which may
cause negative effects on wetland ecosystem services through
accelerating microbial activity and consequently promoting GHG
production or release from deeper preserved soils.

Acknowledgements

We thank our lab technicians Ms. Xiao-Ping Xie, Ms. Wen Jiang, and Ms. Xiao Zhou for their assistance in the lab. We thank Dr. Jianhong Liang from Institute of Karst Geology, China for the sampling and collection of the WS soil.

Author contributions

Cai YJ and Chen Z conceptualized this study. Cai YJ, Zhang HY, Hu
XR, Yang YC, and Zhang JH performed the experiment and data
analyses. Cai YJ drafted the manuscript with the methodological
advice and guidance from Chen Z, Hazard C, Nicol GW, He JZ, Shen
JP, He ZY, and Zhang L. Liu H provided guidance for the perfor-
mance of the automated platform. Zhang S provided guidance for
the performance of IPI samplers. All authors provided feedback
and proofread the manuscript.
## Supplementary material

Supplementary material is available at *The ISME Journal* online.

## Conflicts of interest

The authors declare no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

## Funding

This work was supported by the National Natural Science Foundation of China (No. 42477116, 41977320). Cai YJ acknowledges Xi'an Jiaotong-Liverpool University for the PhD studentship that supported her PhD studies.

## Data availability

All data needed to evaluate the conclusions in the paper are presented in the paper and/or the Supplementary Information.

## References

1. Tian HQ, Xu RT, Canadell JG. et al. A comprehensive quantification of global nitrous oxide sources and sinks. *Nature* 2020;586:248-56. https://doi.org/10.1038/s41586-020-2780-0

2. Zhang WS, Li HP, Pueppke SG. et al. Restored riverine wetlands in a headwater stream can simultaneously behave as sinks of N₂O and hotspots of CH₄ production. *Environ Pollut* 2021;284:117114. https://doi.org/10.1016/j.envpol.2021.117114

3. Bahram M, Espenberg M, Pärn J. et al. Structure and function of the soil microbiome underlying N₂O emissions from global wetlands. *Nat Commun* 2022;13:1430. https://doi.org/10.1038/s41467-022-29161-3

4. Mitsch WJ, Gosselink JG. *Wetlands*. New York: John Wiley & Sons, Inc., 2015.

5. Pärn J, Verhoeven J, Butterbach-Bahl K. et al. Nitrogen-rich organic soils under warm well-drained conditions are global nitrous oxide emission hotspots. *Nat Commun* 2018;9:1135. https://doi.org/10.1038/s41467-018-03540-1

6. Hergoualc'h K, Dezzeo N, Verchot LV. et al. Spatial and temporal variability of soil N₂O and CH₄ fluxes along a degradation gradient in a palm swamp peat forest in the Peruvian Amazon. *Glob Chang Biol* 2020;26:7198-216. https://doi.org/10.1111/gcb.15354

7. Zhang TB, Liu X, An Y. Fluctuating water level effects on soil greenhouse gas emissions of returning farmland to wetland. *J Soils Sediments* 2020;20:3857-66. https://doi.org/10.1007/s11368-020-02730-9

8. Tangen B, Bansal S. Prairie wetlands as sources or sinks of nitrous oxide: effects of land use and hydrology. *Agric For Meteorol* 2022;320:108968. https://doi.org/10.1016/j.agrformet.2022.108968

9. Bruland G, Richardson C, Whalen S. Spatial variability of denitrification potential and related soil properties in created, restored, and paired natural wetlands. *Wetlands* 2006;26:1042-56. https://doi.org/doi:10.1672/0277-5212(2006)26[1042:SVODPA]2.0.CO;2

10. Groffman P, Butterbach-Bahl K, Fulweiler R. et al. Challenges to incorporating spatially and temporally explicit phenomena (hotspots and hot moments) in denitrification

models. *Biogeochemistry* 2009;93:49-77. https://doi.org/10.1007/s10533-008-9277-5

11. Rosentreter J, Al-Haj A, Fulweiler R. et al. Methane and nitrous oxide emissions complicate coastal blue carbon assessments. *Glob Biogeochem Cycles* 2021;35:e2020GB006858. https://doi.org/10.1029/2020GB006858

12. Hu H, Chen D, He J. Microbial regulation of terrestrial nitrous oxide formation: understanding the biological pathways for prediction of emission rates. *FEMS Microbiol Rev* 2015;39:729-49. https://doi.org/10.1093/femsre/fuv021

13. Kuypers MMM, Marchant HK, Kartal B. The microbial nitrogen-cycling network. *Nat Rev Microbiol* 2018;16:263-76. https://doi.org/10.1038/nrmicro.2018.9

14. Hink L, Gubry-Rangin C, Nicol G. et al. The consequences of niche and physiological differentiation of archaeal and bacterial ammonia oxidisers for nitrous oxide emissions. *ISME J* 2018;12:1084-93. https://doi.org/10.1038/s41396-017-0025-5

15. Rudaz A, Davidson E, Firestone M. Sources of nitrous-oxide production following wetting of dry soil. *FEMS Microbiol Ecol* 1991;85:117-24. https://doi.org/10.1111/j.1574-6968.1991.tb04703.x-11

16. Holtan-Hartwig L, Dörsch P, Bakken L. Low temperature control of soil denitrifying communities: kinetics of N₂O production and reduction. *Soil Biol Biochem* 2002;34:1797-806. https://doi.org/10.1016/S0038-0717(02)00169-4

17. Xing GX, Cao YC, Shi SL. et al. Denitrification in underground saturated soil in a rice paddy region. *Soil Biol Biochem* 2002;34:1593-8. https://doi.org/10.1016/S0038-0717(02)00143-8

18. Palmer K, Biasi C, Horn M. Contrasting denitrifier communities relate to contrasting N₂O emission patterns from acidic peat soils in arctic tundra. *ISME J* 2012;6:1058-77. https://doi.org/10.1038/ismej.2011.172

19. Morse JL, Bernhardt ES. Using ¹⁵N tracers to estimate N₂O and N₂ emissions from nitrification and denitrification in coastal plain wetlands under contrasting land-uses. *Soil Biol Biochem* 2013;57:635-43. https://doi.org/10.1016/j.soilbio.2012.07.025

20. Yuan D, Zheng L, Liu Y. et al. Nitrifiers cooperate to produce nitrous oxide in plateau wetland sediments. *Environ Sci Technol* 2023;57:810-21. https://doi.org/10.1021/acs.est.2c06234

21. Prosser JI, Nicol GW. Archaeal and bacterial ammonia-oxidisers in soil: the quest for niche specialisation and differentiation. *Trends Microbiol* 2012;20:523-31. https://doi.org/10.1016/j.tim.2012.08.001

22. Daims H, Lebedeva EV, Pjevac P. et al. Complete nitrification by Nitrospira bacteria. *Nature* 2015;528:504-9. https://doi.org/10.1038/nature16461

23. Chen XP, Zhu YG, Xia Y. et al. Ammonia-oxidizing archaea: important players in paddy rhizosphere soil? *Environ Microbiol* 2008;10:1978-87. https://doi.org/10.1111/j.1462-2920.2008.01613.x

24. Li YL, Wang XX. Root-induced changes in radial oxygen loss, rhizosphere oxygen profile, and nitrification of two rice cultivars in Chinese red soil regions. *Plant Soil* 2013;365:115-26. https://doi.org/10.1007/s11104-012-1378-1

25. Small GE, Cotner JB, Finlay JC. et al. Nitrogen transformations at the sediment-water interface across redox gradients in the Laurentian Great Lakes. *Hydrobiologia* 2014;731:95-108. https://doi.org/10.1007/s10750-013-1569-7

26. Stein L. Insights into the physiology of ammonia-oxidizing microorganisms. *Curr Opin Chem Biol* 2019;49:9-15. https://doi.org/10.1016/j.cbpa.2018.09.003

27. Kits K, Jung M, Vierheilig J. et al. Low yield and abiotic origin of N₂O formed by the complete nitrifier Nitrospira inopinata.
Nat Commun 2019;10:1836. https://doi.org/10.1038/s41467-019-09790-x

28. Sun D, Tang X, Li J. et al. Chlorate as a comammox Nitrospira specific inhibitor reveals nitrification and N₂O production activity in coastal wetland. Soil Biol Biochem 2022;173:108782. https://doi.org/10.1016/j.soilbio.2022.108782

29. Zhu-Barker X, Cavazos A, Ostrom N. et al. The importance of abiotic reactions for nitrous oxide production. Biogeochemistry 2015;126:251-67. https://doi.org/10.1007/s10533-015-0166-4

30. Buessecker S, Sarno A, Reynolds M. et al. Coupled abiotic-biotic cycling of nitrous oxide in tropical peatlands. Nat Ecol Evol 2022;6:1881-90. https://doi.org/10.1038/s41559-022-01892-y

31. Reimers CE, Fischer KM, Merewether R. et al. Oxygen micropro- files measured in situ in deep ocean sediments. Nature 1986;320: 741-4. https://doi.org/10.1038/320741a0

32. Sweerts JPRA, Bargilissen MJ, Cornelese AA. et al. Oxygen- consuming processes at the profundal and littoral sediment water interface of a small meso-eutrophic Lake (lake Vechten, the Netherlands). Limnol Oceanogr 1991;36:1124-33. https://doi.org/10.4319/lo.1991.36.6.1124

33. Lüdemann H, Arth I, Liesack W. Spatial changes in the bacte- rial community structure along a vertical oxygen gradient in flooded paddy soil cores. Appl Environ Microbiol 2000;66:754-62. https://doi.org/10.1128/AEM.66.2.754-762.2000

34. Corzo A, Jiménez-Arias JL, Torres E. et al. Biogeochemical changes at the sediment-water interface during redox transitions in an acidic reservoir: exchange of protons, acidity and electron donors and acceptors. Biogeochemistry 2018;139:241-60. https:// doi.org/10.1007/s10533-018-0465-7

35. Borch T, Kretzschmar R, Kappler A. et al. Biogeochemical redox processes and their impact on contaminant dynamics. Environ Sci Technol 2010;44:15-23. https://doi.org/10.1021/es9026248

36. Mitsunobu S, Toda M, Hamamura N. et al. Millimeter-scale topsoil layer blocks arsenic migration in flooded paddy soil. Geochim Cosmochim Acta 2020;274:211-27. https://doi.org/10.1016/j.gca.2020.01.038

37. Armitage DW, Jones SE. How sample heterogeneity can obscure the signal of microbial interactions. ISME J 2019;13:2639-46. https://doi.org/10.1038/s41396-019-0463-3

38. Cai YJ, Liu ZA, Zhang S. et al. Microbial community structure is stratified at the millimeter-scale across the soil-water interface. ISME Commun 2022;2:53. https://doi.org/10.1038/s43705-022-00138-z

39. Jin C, Sengupta A. Microbes in porous environments: from active interactions to emergent feedback. Biophys Rev 2024;16:173-88. https://doi.org/10.1007/s12551-024-01185-7

40. Garcia-Pichel F, Johnson SL, Youngkin D. et al. Small-scale verti- cal distribution of bacterial biomass and diversity in biological soil crusts from arid lands in the Colorado plateau. Microb Ecol 2003;46:312-21. https://doi.org/10.1007/s00248-003-1004-0

41. Harris JK, Caporaso JG, Walker JJ. et al. Phylogenetic stratigraphy in the Guerrero negro hypersaline microbial mat. ISME J 2013;7: 50-60. https://doi.org/10.1038/ismej.2012.79

42. Ohkubo S, Miyashita H. A niche for cyanobacteria producing chlorophyll f within a microbial mat. ISME J 2017;11:2368-78. https://doi.org/10.1038/ismej.2017.98

43. Melton ED, Stief P, Behrens S. et al. High spatial resolution of distribution and interconnections between Fe- and N-redox processes in profundal lake sediments. Environ Microbiol 2014;16: 3287-303. https://doi.org/10.1111/1462-2920.12566

44. Yuan ZF, Gustave W, Bridge J. et al. Tracing the dynamic changes of element profiles by novel soil porewater samplers with

ultralow disturbance to soil-water interface. Environ Sci Technol 2019;53:5124-32. https://doi.org/10.1021/acs.est.8b05390

45. Li C, Ding SM, Chen MS. et al. Visualizing biogeochemical heterogeneity in soils and sediments: a review of advanced micro-scale sampling and imaging methods. Crit Rev Environ Sci Technol 2023;53:1229-53. https://doi.org/10.1080/10643389. 2022.2128239

46. Zhang S, Yuan Z, Cai Y. et al. Dissolved solute sampling across an oxic-anoxic soil-water interface using microdialysis profilers. J Vis Exp 2023;193:e64358. https://doi.org/10.3791/64358-v

47. Sims GK, Ellsworth TR, Mulvaney RL. Microscale determination of inorganic nitrogen in water and soil extracts. Commun Soil Sci Plant Anal 1995;26:303-16. https://doi.org/10.1080/00103629 509369298

48. Jacinthe PA, Dick WA. Use of silicone tubing to sample nitrous oxide in the soil atmosphere. Soil Biol Biochem 1996;28:721-6. https://doi.org/10.1016/0038-0717(95)00176-X

49. Clough TJ, Kelliher FM, Wang YP. et al. Diffusion of ¹⁵N-labelled N₂O into soil columns: a promising method to examine the fate of N₂O in subsoils. Soil Biol Biochem 2006;38:1462-8. https://doi.org/10.1016/j.soilbio.2005.11.002

50. Jorgensen CJ, Struwe S, Elberling B. Temporal trends in N₂O flux dynamics in a Danish wetland- effects of plant-mediated gas transport of N₂O and O₂ following changes in water level and soil mineral-N availability. Glob Chang Biol 2012;18:210-22. https:// doi.org/10.1111/j.1365-2486.2011.02485.x

51. Collier SM, Ruark MD, Oates LG. et al. Measurement of green- house gas flux from agricultural soils using static chambers. J Vis Exp 2014;90:e52110. https://doi.org/10.3791/52110

52. Wang YQ, Bai R, Di HJ. et al. Differentiated mechanisms of biochar mitigating straw-induced greenhouse gas emissions in two contrasting paddy soils. Front Microbiol 2018;9:02566. https:// doi.org/10.3389/fmicb.2018.02566

53. Pester M, Maixner F, Berry D. et al. NxrB encoding the beta subunit of nitrite oxidoreductase as functional and phylogenetic marker for nitrite-oxidizing Nitrospira. Environ Microbiol 2013;16: 3055-71. https://doi.org/10.1111/1462-2920.12300

54. Daimis H, Lücker S, Wagner M. A new perspective on microbes formerly known as nitrite-oxidizing bacteria. Trends Microbiol 2016;24:699-712. https://doi.org/10.1016/j.tim.2016.05.004

55. Harrell Jr F. Hmisc: Harrell Miscellaneous. R package version 5.1-1. 2023. http://CRAN.R-project.org/package=Hmisc

56. Wei T, Simko V. Corrplot: Visualization of a Correlation Matrix (Version 0.92). 2021. http://github.com/taiyun/corrplot

57. Hallin S, Philippot L, Löffler F. et al. Genomics and ecology of novel N₂O-reducing microorganisms. Trends Microbiol 2018;26: 43-55. https://doi.org/10.1016/j.tim.2017.07.003

58. Schreiber F, Loeffler B, Polerecky L. et al. Mechanisms of transient nitric oxide and nitrous oxide production in a complex biofilm. ISME J 2009;3:1301-13. https://doi.org/10.1038/ismej.2009.55

59. Kinh CT, Riya S, Hosomi M. et al. Identification of hotspots for NO and N₂O production and consumption in counter- and co- diffusion biofilms for simultaneous nitrification and denitrifica- tion. Bioresour Technol 2017;245:318-24. https://doi.org/10.1016/j. biortech.2017.08.051

60. Kinh CT, Suenaga T, Hori T. et al. Counter-diffusion biofilms have lower N₂O emissions than co-diffusion biofilms during simultaneous nitrification and denitrification: insights from depth-profile analysis. Water Res 2017;124:363-71. https://doi. org/10.1016/j.watres.2017.07.058

61. Jorgensen CJ, Elberling B. Effecsts of flooding-induced N₂O pro- duction, consumption and emission dynamics on the annual
N₂O emission budget in wetland soil. Soil Biol Biochem 2012;53: 9–17. https://doi.org/10.1016/j.soilbio.2012.05.005

62. Schmidt I, van Spanning RJM, Jetten MSM. Denitrification and ammonia oxidation by Nitrosomonas europaea wild-type, and NirK- and NorB-deficient mutants. Microbiology-Sgm 2004;150: 4107–14. https://doi.org/10.1099/mic.0.27382-0

63. Cantera JJL, Stein LY. Molecular diversity of nitrite reductase genes (nirK) in nitrifying bacteria. Environ Microbiol 2007;9:765–76. https://doi.org/10.1111/j.1462-2920.2006.01198.x

64. Lücker S, Nowka B, Rattei T. et al. The genome of Nitrospina gracilis illuminates the metabolism and evolution of the major marine nitrite oxidizer. Front Microbiol 2013;4:00027. https://doi.org/10.3389/fmicb.2013.00027

65. Sun X, Jayakumar A, Tracey J. et al. Microbial N₂O consumption in and above marine N₂O production hotspots. ISME J 2021;15: 1434–44. https://doi.org/10.1038/s41396-020-00861-2

66. Kozlowski JA, Steidlmeier M, Schleper C. et al. Pathways and key intermediates required for obligate aerobic ammonia-dependent chemolithotrophy in bacteria and Thaumarchaeota. ISME J 2016;10:1836–45. https://doi.org/10.1038/ismej.2016.2

67. Zhang IH, Sun X, Jayakumar A. et al. Partitioning of the denitrification pathway and other nitrite metabolisms within global oxygen deficient zones. ISME Commun 2023;3:76. https://doi.org/10.1038/s43705-023-00284-y

68. Ma Y, Zilles JL, Kent AD. An evaluation of primers for detecting denitrifiers via their functional genes. Environ Microbiol 2019;21: 1196–210. https://doi.org/10.1111/1462-2920.14555

69. Qu Z, Bakken LR, Molstad L. et al. Transcriptional and metabolic regulation of denitrification in Paracoccus denitrificans allows low but significant activity of nitrous oxide reductase under oxic conditions. Environ Microbiol 2015;18:2951–63. https://doi.org/10.1111/1462-2920.13128

70. Kim DD, Han H, Yun T. et al. Identification of nosZ-expressing microorganisms consuming trace N₂O in microaerobic chemostat consortia dominated by an uncultured Burkholderiales. ISME J 2022;16:2087–98. https://doi.org/10.1038/s41396-022-01260-5

71. Sennett LB, Roco CA, Lim NYN. et al. Determining how oxygen legacy affects trajectories of soil denitrifier community dynamics and N₂O emissions. Nat Commun 2024;15:7298. https://doi.org/10.1038/s41467-024-51688-w

72. Park D, Kim H, Yoon S. Nitrous oxide reduction by an obligate aerobic bacterium, Gemmatimonas aurantiaca strain T-27. Appl Environ Microbiol 2017;83:e00502–17. https://doi.org/10.1128/ AEM.00502-17

73. Wang Z, Vishwanathan N, Kowaliczko S. et al. Clarifying microbial nitrous oxide reduction under aerobic conditions: tolerant, intolerant, and sensitive. Microbiol Spectr 2023;11:e04709–22. https://doi.org/10.1128/spectrum.04709-22

74. Hiis EG, Vick SHW, Molstad L. et al. Unlocking bacterial potential to reduce farmland N₂O emissions. Nature 2024;630:421–8. https://doi.org/10.1038/s41586-024-07464-3

75. Maza-Marquez P, Lee MD, Detweiler AM. et al. Millimeter-scale vertical partitioning of nitrogen cycling in hypersaline mats reveals prominence of genes encoding multi-heme and priscmane proteins. ISME J 2022;16:1119–29. https://doi.org/10.1038/ s41396-021-01161-z

76. Bru D, Sarr A, Philippot L. Relative abundances of proteobacterial membrane-bound and periplasmic nitrate reductases in selected environments. Appl Environ Microbiol 2007;73:5971–4. https://doi.org/10.1128/AEM.00643-07

77. Wu Q, Ma Y, Wang X. et al. Functional players involved in the distinct nitrogen metabolism in two geographically

different paddy soils. Biol Fertil Soils 2023. https://doi.org/10.1007/ s00374-023-01776-8

78. Kraft B, Strous M, Tegetmeyer HE. Microbial nitrate respiration - genes, enzymes and environmental distribution. J Biotechnol 2011;155:104–17. https://doi.org/10.1016/j.jbiotec.2010.12.025

79. Richardson DJ. Bacterial respiration: a flexible process for a changing environment. Microbiology-Sgm 2000;146:551–71. https://doi.org/10.1099/00221287-146-3-551

80. Richardson D, Berks B, Russell D. et al. Functional, biochemical and genetic diversity of prokaryotic nitrate reductases. Cell Mol Life Sci 2001;58:165–78. https://doi.org/10.1007/PL00000845

81. Dong LF, Smith CJ, Papaspyrou S. et al. Changes in benthic denitrification, nitrate ammonification, and anammox process rates and nitrate and nitrite reductase gene abundances along an estuarine nutrient gradient (the Colne estuary, United Kingdom). Appl Environ Microbiol 2009;75:3171–9. https://doi.org/10.1128/ AEM.02511-08

82. Sawers R, Fischer M, Falke D. Anaerobic nitrate respiration in the aerobe Streptomyces coelicolor A3(2): helping maintain a proton gradient during dormancy. Environ Microbiol Rep 2019;11:645–50. https://doi.org/10.1111/1758-2229.12781

83. Nadeau SA, Roco CA, Debenport SJ. et al. Metagenomic analysis reveals distinct patterns of denitrification gene abundance across soil moisture, nitrate gradients. Environ Microbiol 2019;21: 1255–66. https://doi.org/10.1111/1462-2920.14587

84. Yuan Q, Liu P, Lu YH. Differential responses of nirK- and nirS-carrying bacteria to denitrifying conditions in the anoxic rice field soil. Environ Microbiol Rep 2012;4:113–22. https://doi. org/10.1111/j.1758-2229.2011.00311.x

85. Pold G, Bonilla-Rosso G, Saghaï A. et al. Phylogenetics and environmental distribution of nitric oxide-forming nitrite reductases reveal their distinct functional and ecological roles. ISME Commun 2024;4:ycae020. https://doi.org/10.1093/ismeco/ycae020

86. Herold M, Giles M, Alexander C. et al. Variable response of nirK and nirS containing denitrifier communities to long-term pH manipulation and cultivation. FEMS Microbiol Lett 2018;365:fny035. https://doi.org/10.1093/femsle/fny035

87. Sun HS, Jiang SX. A review on nirS-type and nirK-type denitrifiers via a scientometric approach coupled with case studies. Environ Sci Process Impacts 2022;24:221–32. https://doi.org/10.1039/D1 EM00518A

88. Qin HL, Xing XY, Tang YF. et al. Soil moisture and activity of nitrite- and nitrous oxide-reducing microbes enhanced nitrous oxide emissions in fallow paddy soils. Biol Fertil Soils 2020;56: 53–67. https://doi.org/10.1007/s00374-019-01403-5

89. Desnues C, Michotey VD, Wieland A. et al. Seasonal and diel distributions of denitrifying and bacterial communities in a hypersaline microbial mat (Camargue, France). Water Res 2007;41: 3407–19. https://doi.org/10.1016/j.watres.2007.04.018

90. Wu Y, Whitaker J, Toet S. et al. Diurnal variability in soil nitrous oxide emissions is a widespread phenomenon. Glob Chang Biol 2021;27:4950–66. https://doi.org/10.1111/gcb.15791

91. Windham-Myers L, Bergamaschi B, Anderson F. et al. Potential for negative emissions of greenhouse gases (CO₂, CH₄ and N₂O) through coastal peatland re-establishment: novel insights from high frequency flux data at meter and kilometer scales. Environ Res Lett 2018;13:045005. https://doi.org/10.1088/1748-9326/ aaae74

92. Yu Z, Li Y, Deng H. et al. Effect of *Scirpus mariqueter* on nitrous oxide emissions from a subtropical monsoon estuarine wetland. J Geophys Res 2012;117:G02017. https://doi.org/10.1029/2011JG001850
93. Jiang CR, Zhang SB, Wang JF, et al. Nitrous oxide (N₂O) emissions decrease significantly under stronger light irradiance in riverine water columns with suspended particles. *Environ Sci Technol* 2023;**57**:19749–59. https://doi.org/10.1021/acs.est.3c05526

94. Yao J, Fu Y, Yu Z. et al. Interaction of rice root Fe plaque with radial oxygen loss enhances paddy-soil N₂O emission by increasing ●OH production and subsequently inhibiting N₂O reduction. *Soil Biol Biochem* 2024;**195**:109475. https://doi.org/10.1016/j.soilbio.2024.109475

95. Wu BB, Zhou C, Zhao GQ. et al. Enhanced photochemical pro-duction of reactive intermediates at the wetland soil-water

interface. *Water Res* 2022;**223**:118971. https://doi.org/10.1016/j.watres.2022.118971

96. Morris JJ, Rose AL, Lu Z. Reactive oxygen species in the world ocean and their impacts on marine ecosystems. *Redox Biol* 2022;**52**:102285. https://doi.org/10.1016/j.redox.2022.102285

97. Dini-Andreote F, Kowalchuk GA, Prosser JI. et al. Towards meaningful scales in ecosystem microbiome research. *Environ Microbiol* 2021;**23**:1–4. https://doi.org/10.1111/1462-2920.15276

98. Ladau J, Eloe-Fadrosh EA. Spatial, temporal, and phyloge-netic scales of microbial ecology. *Trends Microbiol* 2019;**27**:662–9. https://doi.org/10.1016/j.tim.2019.03.003