Denitrification is the major nitrous acid production pathway in boreal agricultural soils

Bhattarai, Hem Raj; Wanek, Wolfgang; Siljanen, Henri M. P.; Ronkainen, Jussi G.; Liimatainen, Maarit; Hu, Yuntao; Nykänen, Hannu; Biasi, Christina; Maljanen, Marja

doi:10.1038/s43247-021-00125-7

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Published: 05 March 2021

Denitrification is the major nitrous acid production pathway in boreal agricultural soils

Communications Earth & Environment volume 2, Article number: 54 (2021) Cite this article

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Abstract

Nitrous acid (HONO) photolysis produces hydroxyl radicals—a key atmospheric oxidant. Soils are strong HONO emitters, yet HONO production pathways in soils and their relative contributions are poorly constrained. Here, we conduct ¹⁵N tracer experiments and isotope pool dilution assays on two types of agricultural soils in Finland to determine HONO emission fluxes and pathways. We show that microbial processes are more important than abiotic processes for HONO emissions. Microbial nitrate reduction (denitrification) considerably exceeded ammonium oxidation as a source of nitrite—a central nitrogen pool connected with HONO emissions. Denitrification contributed 97% and 62% of total HONO fluxes in low and high organic matter soil, respectively. Microbial ammonium oxidation only produced HONO in high organic matter soil (10%). Our findings indicate that microbial nitrate reduction is an important HONO production pathway in aerobic soils, suggesting that terrestrial ecosystems favouring it could be HONO emission hotspots, thereby influencing atmospheric chemistry.

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Introduction

Up to 60% of primary production of atmospheric hydroxyl (OH) radicals can be attributed to photolysis of nitrous acid (HONO)¹. Hydroxyl radicals are vital for the removal of pollutants and reactive gases, such as carbon monoxide and methane^2,3 from the atmosphere, and they are also an essential precursor of cloud formation by transforming volatile organic compounds into secondary organic aerosols^4,5. The current global HONO budget is not balanced when all known sources and sinks are included, which suggests that there is a missing source of HONO^6,7,8. Although soils have only recently been identified as important HONO emitters^{9,10,11,12,13,14,15,16,17}, soils have been proposed as a key component of this missing HONO source. Soil nitrite (NO₂⁻) may be the key HONO precursor⁹ but this has not yet been directly demonstrated using ¹⁵N tracer approaches and accounting for primary microbial pathways producing it. Nitrite is mainly produced during microbial ammonia (NH₃) oxidation (autotrophic and heterotrophic nitrification) and nitrate (NO₃⁻) reduction (denitrification and dissimilatory nitrate reduction to ammonium (DNRA)) pathways^18,19,20. In addition, direct release of HONO by NH₃ oxidizers (autotrophic nitrifiers) has also been demonstrated^10,21. Although the mechanisms that underpin HONO production are not yet fully understood, several studies have demonstrated the importance of microbial pathways in soil HONO production^{9,10,11,12,13,14,15,16,17}. However, abiotic pathways (e.g., hydroxylamine+water = HONO) have been proposed but have not been investigated in soils to date²¹.

The first evidence on a potential link between soil HONO emissions and underlying microbial processes was provided conducting ¹⁵N tracer experiments, showing that nitrifiers outpaced denitrifiers in soil HONO emissions²². Another study²³, proving the concept, also provided evidence on the participation of microbial processes in soil HONO emissions, especially the involvement of NH₃ oxidizers (nitrifiers) by using ¹⁵N-labeled urea. To date, only one systematic study has addressed the pathways of soil HONO formation using a tracer approach (labeled ammonium (¹⁵NH₄⁺)), highlighting the sole importance of NH₃ oxidizers in HONO production and excluding the role of NO₃⁻ reduction in aerobic soils¹¹. However, microbial NO₃⁻ reduction is a widespread phenomenon in soils of many ecosystems²⁴, with NO₂⁻ produced as an intermediate of the process. Recently, the participation of denitrification processes in water-saturated soils (~100% water-holding capacity (WHC)) was demonstrated for soil HONO formation, suggesting that NO₃⁻ reduction to NO₂⁻ under anoxic conditions could be crucial for soil HONO emissions²⁵. Nevertheless, besides water-saturated conditions, soil aggregates can also generate anoxic microsites that can favor microbial NO₃⁻ reduction in water-unsaturated soils, e.g., in agricultural soils^26,27. In addition, the relative contribution of different microbial pathways to HONO production may vary in soils, as—beyond soil moisture and oxygen saturation—microbial activity is also influenced by soil pH, organic matter (OM) content, soil carbon to nitrogen (C:N) ratio, and temperature^19,28,29. Thus, HONO production could occur at the microscale level and by contrasting pathways that need to be further investigated. Moreover, the proposed abiotic pathway²¹ needs to be addressed in soils. Thus, in order to get a better understanding of the processes that contribute to HONO emissions from soils and their impact on the global HONO budget, the expected link between the soil inorganic N cycle and HONO formation requires further investigation^9,30,31. Agricultural soils, which cover 50% of the global habitable land area^32,33, possess a strong potential to release HONO because of enhanced N fertilizer use, which is expected to increase by ~2–4-fold by 2050³⁴. Therefore, it is crucial to understand HONO production pathways in agricultural ecosystems.

This study focused on the understanding of HONO production and emissions from agricultural soils of the boreal region, which are typically acidic (pH < 7)⁹. To date, only three studies^12,13,15 have investigated HONO emissions from boreal agricultural soils. Yet, the processes and pathways behind those HONO emissions have not been demonstrated. In this study, we performed a series of ¹⁵N tracer experiments to understand HONO production pathways in two distinct boreal agricultural soils in Finland; one with a low organic matter (LOM) content and the other with a high organic matter (HOM) content (Supplementary Table 1). First, we aimed to distinguish between biotic and abiotic HONO production pathways by adding ¹⁵N-labeled NO₂⁻(LNi) to both, live, and sterile soils. Second, to determine the contribution of the two main biotic HONO production pathways (microbial NH₃ oxidation and NO₃⁻ reduction), we performed separate experiments using ¹⁵N-labeled ammonium (NH₄⁺, LA) and NO₃⁻ (LNa) in live (non-sterilized) soils. The HONO flux rates were then compared with data compiled in a global HONO flux rate synthesis, which was generated for the first time here. In addition, gross NO₂⁻ production rates across soil pH levels, and the contribution of nitrification and denitrification to the soil NO₂⁻ pool were also compared with published data.

Results

Time kinetics of added ¹⁵NO₂ ⁻ and HO¹⁵NO production in live and sterile soils

We sterilized the soil samples by autoclaving to ensure that the biotic pathways that produce HONO via NO₂⁻ or NH₃-oxidizing microbes were completely blocked. Autoclavation is the most effective sterilization method in soils, though autoclavation can alter the structure of OM content and cause disaggregation, for example, increasing the availability of dissolved organic matter³⁵. We examined the ¹⁵N atom percent excess (APE) of emitted HONO (HO¹⁵NO APE) and of soil NO₂⁻ (¹⁵NO₂⁻ APE) in live and sterile soil samples from time zero to 240 minutes, after the addition of labeled ¹⁵NO₂⁻ (LNi) and unlabeled NO₂⁻ (control) (Fig. 1).

**Fig. 1: The ¹⁵N atom percent excess (APE) of nitrous acid (HONO) and soil nitrite (NO₂⁻) in labeled ¹⁵NO₂⁻ (LNi) and control treatments.**

In the LNi treatment under live conditions, labeled ¹⁵NO₂⁻ gradually declined (Fig. 1a, b), indicating the presence of active microbial pathways that produce and consume NO₂⁻. Further, ¹⁵NO₂⁻ consumption was not equally accompanied by ¹⁵NO₃⁻production (by nitrifiers) under live conditions (Supplementary Figure 1). Microbial NO₂⁻ production pathways led to a 50% decrease in ¹⁵NO₂⁻ APE by the end of the experiments in both soils (Fig. 1a, b, Supplementary Table 2–3). Both soils also showed a steady decrease in HO¹⁵NO APE under live conditions, with the HOM soils exhibiting a stronger decrease than the LOM soils. Under sterile conditions, ¹⁵NO₂⁻APE remained stable at the approximate initial values during the whole incubation period, confirming the blockage of biotic pathways that produce NO₂⁻ (Fig. 1c, d). Despite stable ¹⁵NO₂⁻ APE, HO¹⁵NO APE decreased towards the end of the incubation period under sterile conditions (Fig. 1c, d), although the ¹⁵N depletion of HO¹⁵NO occurred later in time and was less pronounced (at least for LOM) than under live conditions.

Relationship between biotic pathways and HONO production

To further disentangle the biotic HONO production pathways (in addition to ¹⁵NO₂⁻ (LNi; see experiment above)), we added ¹⁵NH₄⁺ (LA) and ¹⁵NO₃⁻ (LNa) to the live soil samples and measured the subsequent change in ¹⁵NO₂⁻ and emissions of HO¹⁵NO. We also quantified gross production and gross consumption rates of NH₄⁺, NO₂⁻, and NO₃⁻ in the live soil samples, based on ¹⁵N pool dilution approaches, to better explain the relationship between the microbial pathways that use ¹⁵NH₄⁺ and ¹⁵NO₃⁻ to produce HONO. Production (i.e., N mineralization) and consumption rates of NH₄⁺ were relatively low and did not differ between soils. Production (i.e., nitrification) and consumption rates of NO₃⁻ were 6–24 times higher than gross NH₄⁺ turnover rates and were significantly higher in the HOM soil than the LOM soil (Fig. 2a). Gross NO₂⁻ production and consumption rates ranged between the values observed for nitrification and mineralization, and were not significantly different between the two studied soils. Production of ¹⁵NO₂⁻ increased gradually in the LNa treatment in both soils, which exhibited high NO₃⁻ consumption rates (Fig. 2a–c, Supplementary Table 4), indicating that the reduction of NO₃⁻ to NO₂⁻ is a crucial biotic pathway in both water-unsaturated soils. Despite high nitrification rates (22 µg N g_dw ⁻¹ d⁻¹) in the HOM soil, the magnitude of ¹⁵NO₂⁻ and HO¹⁵NO production during nitrification was smaller compared with NO₃⁻ reduction (Fig. 2b, treatments LA vs. LNa), which suggests that the capability to nitrify by the NH₃ oxidizers, which reside in the HOM soil, was lower compared with the NO₃⁻ reducers that shared the same environment.

**Fig. 2: Gross transformation rates of mineral nitrogen and production of ¹⁵NO₂⁻ and HO¹⁵NO in live soils.**

The contribution of the microbial pathways that use NH₄⁺ and NO₃⁻ to total production and emissions of HONO was evident. In general, reduction of ¹⁵NO₃⁻ affected the production of HO¹⁵NO via ¹⁵NO₂⁻ formation, as ¹⁵N APE values of HONO (on average ~1–18% in the LOM soil and 1.2–18% in the HOM soil) and of NO₂⁻ increased gradually with time only in the LNa treatment in both soils (Fig. 2b–c). The strong relationship between produced HO¹⁵NO and added ¹⁵NO₂⁻ (LNi treatment), or with ¹⁵NO₂⁻produced from ¹⁵NO₃⁻ (LNa treatment) (Supplementary Figure 2), further confirmed that HONO was generated mostly from NO₂⁻ produced via microbial NO₃⁻ reduction in both soils. In addition, the observation that HONO emission rates decreased with decreasing NO₂⁻ concentrations (Supplementary Figures 3–6 and Supplementary Table 4) would also suggest that HONO production and emissions are directly linked to soil NO₂⁻, especially in LOM soil. The HOM soil showed a curvilinear relationship between HONO emissions and soil NO₂⁻ concentrations (Supplementary Figure 6a), indicating rapid NO₂⁻ consumption processes that competed tightly with HONO production via the soil NO₂⁻ pool. In contrast, microbial NH₄⁺ oxidation occurred only in the HOM soil but not in the LOM soil. Production of HO¹⁵NO (from 0.1 ± 0.1 at 30 min to 0.4 ± 0.1% ¹⁵N APE at 240 min) and ¹⁵NO₂⁻ (from 0.2 ± 0.3 at 90 min to 1.6 ± 1.3% ¹⁵N APE at 240 min) from ¹⁵NH₄⁺ (LA treatment) was less than NO₃⁻ reduction (LNa treatment), and showed a weak, insignificant association (Supplementary Figure 2).

Quantification of HONO emissions by biotic and abiotic processes

In the presence of microbial activity in the live soil samples, a two-source isotope mixing model³⁶ showed that NO₂⁻ contributed 100% to HONO production in the LOM soil and 81% in the HOM soil. In the latter, (in addition to NO₂⁻) an additional biotic HONO source contributed 19% to HONO production towards the end of the incubation period (Fig. 3a). Despite the presence of a consistent HONO precursor (i.e., NO₂⁻), the dynamics of biotic HONO emissions in the LOM soil differed from the HOM soil (Fig. 3b). Biotic HONO emissions in the LOM soil peaked at 180 min and then decreased slightly until the end of the incubation, whereas biotic HONO emissions increased gradually in the HOM soil, and saturated after 180 min (Fig. 1, Fig. 3b). Microbial processes produced 10.5 ng N m⁻² s⁻¹ and 24.3 ng N m⁻² s⁻¹ of HONO in the HOM and LOM soils, respectively (Fig. 4).

**Fig. 3: Source contribution and nitrous acid (HONO) emission rates from biotic and abiotic sources.**

**Fig. 4: Total soil nitrous acid (HONO) emission rates via abiotic and biotic processes, with the relative contribution of biotic pathways to HONO production as inset.**

Although ¹⁵N enrichment of NO₂⁻ remained constant in the sterile soil samples (as expected if only biotic processes form NO₂⁻), HO¹⁵NO declined, which indicates the presence of an unknown abiotic HONO source (Fig. 3c, d). In the absence of microbial activities in the sterile soils, the unknown HONO production source increased with time (Fig. 3c). Furthermore, the unknown HONO source was more pronounced in the HOM soil than in the LOM soil (Fig. 3c). In the latter, the contribution of the unknown source to total HONO flux peaked at the end of the incubation, reaching a maximum of 22%. In the HOM soil, the contribution from the unknown source increased gradually, exceeding HONO production from soil NO₂⁻ at the end of the incubation, and contributed up to 52%. In both soils, the fractional contributions of the unknown source and abiotic HONO emissions showed similar increasing trends (Fig. 3c, d). In comparison with the LOM soil, the HOM soil had a greater contribution of an unknown source and also greater abiotic HONO emission rates (Fig. 3d).

Discussion

Soil HONO emissions in our boreal agricultural soils (10.5–24.3 ng N m⁻² s⁻¹, Fig. 4) were relatively high compared to a global synthesis of soil HONO emission rates (median 5.0 ng N m⁻² s⁻¹, n = 199; Fig. 5, Supplementary Data 1), but were reflective of managed soils, where fertilization or grazing promote N cycling, which increases soil HONO emissions (cropland ≥ grassland ≥ forest, and peatland, Fig. 5). Only deserts showed greater HONO emissions, indicating arid biological soil crusts being hotspots of HONO emissions globally³⁷.

**Fig. 5: Global synthesis of soil nitrous acid (HONO) emissions rates, gross nitrite (NO₂⁻) production, and the contribution of biotic pathways to the soil NO₂⁻ pool.**

Biotic pathways of soil HONO formation and significance

A consistent decrease in the ¹⁵N enrichment of HONO, in parallel with that in NO₂⁻ in the live soil samples, and much higher rates of HONO production in the live soil samples compared with the sterile soil samples confirmed that microbial processes are the most important HONO sources in soils. Although soil HONO emissions have been previously linked to soil NO₂⁻ based on physicochemical reasoning (acid-base equilibria between H⁺ and NO₂⁻-producing HONO)⁹, here for the first time we unequivocally and causally link the soil NO₂⁻ pool with HONO emissions, based on ¹⁵N tracing and liquid chromatography-high resolution mass spectrometry. The findings further demonstrate that microbial NO₃⁻ reduction dominated HONO production and by far exceeded the concurrently accepted major HONO production pathway, i.e., NH₃ oxidation by autotrophic nitrifiers, in aerobic, water-unsaturated soils, here boreal agricultural soils (Figs. 2a–c, 3b). Among the three soil NO₂⁻-producing microbial pathways, only NO₃⁻ reduction triggered significant ¹⁵NO₂⁻ and HO¹⁵NO production in both soils. Soil NO₃⁻ reduction was solely linked to the presence of the periplasmic NO₃⁻ reductase (napA) gene, as the cytoplasmic NO₃⁻ reductase (narG) gene³⁸ was absent in both soils (Supplementary Figure 7). The lack of ¹⁵NH₄⁺ production from ¹⁵NO₃⁻ in the LNa treatment (Supplementary Figure 1) suggests that DNRA (dissimilatory nitrate reduction to ammonium) was absent and that denitrification dominated microbial NO₃⁻ reduction and soil HONO emissions. This is further confirmed by the absence of the key DNRA gene, i.e., nitrite reductase (nrfA)^18,39, whereas the genes of key denitrifying genes were abundant, e.g. napA, nirK, nirS, and nosZ (Supplementary Figure 7). Nitrification only contributed to a small extent to soil HONO production (Fig. 4), and was only found in the HOM soil (Fig. 2b). Nitrification could be attributed to the presence of bacterial and archaeal NH₃ oxidizers^10,21, as suggested by higher gene copy numbers of bacterial (and archaeal) ammonia monooxygenase (amoA) in the HOM soil (Supplementary Figure 7), and these are capable of directly producing HONO¹⁰ before accumulating NO₂⁻ (Supplementary Figure 2). Decoupling of microbial NO₃⁻ reduction (high ¹⁵NO₂⁻ output, Fig. 2a) and biotic HONO production (Fig. 4) in the HOM soil could be associated with the combined effect of microbial NO₂⁻ consumption and abiotic NO₂⁻ reactions with soil OM via nitration and nitrosation⁴⁰. In these soils, we observed a curvilinear relationship between soil NO₂⁻ concentrations and HONO emission rates (Supplementary Figure 6a), which indicates a gradual conversion of NO₂⁻ to NO₃⁻ (Supplementary Figure 1) and concurrently occurring chemical reactions between NO₂⁻ and soil OM⁴⁰, the latter of which surpass HONO formation via NO₂⁻ after reaching an optimum.

Soil moisture is an important environmental driver of soil HONO emissions. Maximum soil HONO emissions were identified in a wide range of ecosystems under low soil moisture contents (“dry” peak at 20–30% WHC)^9,10,25. Such HONO emission peaks occurred during progressive reduction in soil moisture in the experiments and were linked to nitrifiers as NH₃ oxidation is an aerobic pathway and requires oxygen. Therefore, it was assumed that with soil moisture reduction and progressively developing the aerobic condition, NH₃ oxidation^10,21 dominates soil HONO production and emissions. However, the ¹⁵NH₄⁺ supplemented LOM soil did not emit HO¹⁵NO, even until the end of the experimental period, despite a slight reduction in soil moisture (from 60%_start to 51%_end WHC; Supplementary Figure 8). In contrast, ¹⁵NH₄⁺ supplemented HOM started to emit HO¹⁵NO, presumably by NH₃ oxidizers^10,21, from the middle of the experiment (Fig. 2b, Supplementary Table 2–3), despite a smaller reduction in soil moisture (from 60%_start to 54%_end WHC) compared with the LOM soil (Supplementary Figure 8). In our study, we did not measure HONO emissions for longer periods and therefore did not reach lower soil moisture regimes, for example, 20–30% WHC, at which the contribution of nitrifier pathways could increase^10,21. For example, Wu et al.²³ showed that soil HONO emissions (at 20–30% WHC) were driven by NH₃ oxidation in a pH neutral soil from the temperate region. On the other hand, Kubota and Asami²² estimated that ~69% of the soil emitted HONO originated from NH₃ oxidation at 60% WHC from sub-tropical acidic soils (pH = 4.8–5.8) incubated at 30°C for 30 days. Our estimation shows that NH₃ oxidation contributed only 0–10% to the total biotic HONO emissions (at 60–54% WHC range) from slightly acidic (pH = 6.1) boreal agricultural soils incubated at 21 °C for 48 hours. The likely reason for higher contributions of nitrifiers to soil HONO emissions in the two earlier studies^22,23 could be differences in soil moisture and temperature between studies, which are known to control the activities of NH₃ oxidizers²⁹ and thereby of soil HONO emissions^10,23,25. Nevertheless, our results obtained from the LA treatments in two boreal agricultural soils suggest that the capabilities of NH₃ oxidizers to produce HONO^10,21 vary according to soil type and/or climatic zone, which requires further studies.

Besides soil moisture, soil pH also has a significant role in HONO emissions. Although nitrification (especially by bacterial ammonia oxidizers) decreases towards low soil pH levels⁴¹, at the same time, regardless of its source, rapid protonation of NO₂⁻ in acidic soils⁹ can promote soil HONO formation and release. The reported HONO release from soils by acidophilic and high NH₃ affinity archaea^11,41,42 was lower than HONO emissions via microbial NO₃⁻ reduction (Supplementary Figure 9), further supporting our finding that soil NO₂⁻ via microbial NO₃⁻ reduction is crucial for HONO production, especially in soils with pH < 7. In addition, denitrification demands the presence of protons (2NO₃⁻ + 10e⁻ + 12H⁺→N₂ + 6H₂O) and therefore shows a pH optimum under acidic conditions unlike nitrification, which does the opposite, i.e., it releases H⁺ (NH₃ + 2O₂ → NO₃⁻ + H⁺ + H₂O) into the soil and therefore shows a pH optimum at neutral to alkaline conditions. Therefore, acidic soils are likely to favor NO₃⁻ reduction via denitrification under suitable conditions, such as ample NO₃⁻ availability and anoxic microsites. Moreover, our data synthesis clearly shows that gross NO₂⁻ production rates increase in acidic soils (Fig. 5b, Supplementary Data 2). In published studies that have examined the contribution of nitrifiers and denitrifiers to soil NO₂⁻ production, an average 48% and 57% of the NO₂⁻ source was contributed by denitrifiers (n = 82, Fig. 5c, Supplementary Data 2), when organic N oxidation to NO₂⁻ by heterotrophic nitrifiers was included or excluded. The relationship between soil NO₃⁻ reduction by denitrifiers and soil HONO emissions as found in our study is in agreement with Wu et al.²⁵ who recently demonstrated high HONO emissions from soils under water-saturated, anaerobic conditions (“wet” peak caused by microbial NO₃⁻ reduction at ~100% WHC). However, here we show the importance of microbial NO₃⁻ reduction for HONO production under water-unsaturated, aerobic soil conditions (between 51% and 60% WHC). This suggests that not only water-saturated, anoxic soils, but also anoxic microsites in aerobic soils that are commonly generated in the interior of soil aggregates²⁶ are key to the promotion of microbial NO₃⁻ reduction and soil HONO formation. Therefore, soil HONO production via microbial NO₃⁻ reduction in aggregates is likely a more widespread phenomenon because of the anoxic soil microsites and the ubiquity and diversity of NO₃⁻ reducers (archaea, fungi, bacteria) in a wide range of ecosystems^24,43, and across a wide range of soil pH and moisture contents.

Abiotic pathway of soil HONO formation

Decreasing ¹⁵N enrichment in HONO in sterile soils, despite the constant ¹⁵N enrichment in NO₂⁻, suggests the presence of an abiotic HONO production pathway. With consistent ¹⁵N enrichment in NO₂⁻ and zero microbial production and consumption of NO₂⁻ (Fig. 1), we can only assign the observed HONO flux to an unknown abiotic source, not related to the soil NO₂⁻ pool. As our HONO measurements were conducted in Teflon coated (inner wall) chambers in the dark, heterogeneous reactions that require nitrogen dioxide (NO₂) and light conditions to form HONO^44,45 are expected to be insignificant. Also, the absence of microbial NO₂⁻ production in sterile conditions precludes the possibility of HONO production via self-decomposing HNO₂, as its products (NO₂ and nitric oxide (NO) gases) must pass through the soil NO₂⁻ pool⁴⁶. Recently, a reaction between soil hydroxylamine and H₂O has been suggested to produce HONO abiotically on glass beads²¹, although there is little evidence for this phenomenon in soils. Nevertheless, abiotic production of HONO was twofold higher (Fig. 4) in the HOM soil (1.3 ng N m⁻² s⁻¹) compared with the LOM soil (0.6 ng N m⁻² s⁻¹), which indicates that soil OM may play a role in the abiotic formation of HONO. This unidentified abiotic HONO pathway, which does not require passage through the soil NO₂⁻/HNO₂ pool, clearly needs further investigation.

Conclusion

We confirmed that microbial processes are essential for HONO emissions from agricultural soils, as they contributed 8–42 times more to soil HONO emissions than abiotic processes (Fig. 4). Using ¹⁵NO₂⁻, ¹⁵NO₃⁻, and ¹⁵NH₄⁺ tracers, we showed, for the first time, that microbial NO₃⁻ reduction (denitrifiers) is driving soil HONO emissions in aerobic soils, by fueling the soil NO₂⁻. Other microbial pathways contributed little or not to HONO production, i.e., nitrification (NH₃ and organic N oxidation) and DNRA (Figs. 4, 6). We conclude that microbial processes are essential for soil HONO emissions, and that the microbial NO₃⁻ reduction pathway could be a significant contributor in aerobic soils of many ecosystems. Moreover, abiotic HONO production pathways that have remained elusive could exist in soils, thereby contributing to notable emissions of HONO to the atmosphere, although microbial pathways are dominant. Therefore, in order to better understand the impact of soil emitted HONO on the seasonally changing atmospheric HONO budget, atmospheric OH radical production and associated atmospheric chemistry^30,31,50 (at present and in the future under a changing climate), we suggest that future studies should assess the relative contribution of microbial pathways, in tandem with concurrent abiotic processes (Fig. 6), to soil HONO production, by applying appropriate ¹⁵N tracer approaches.

**Fig. 6: Conceptual model showing nitrous acid (HONO) production pathways associated with the nitrogen cycle in agricultural soils.**

Methods

Soil sampling and sample preparation

Two soils (HOM and LOM) with distinct edaphic properties (Supplementary Table 1) were sampled from two separate agricultural fields maintained by the Natural Resources Institute Finland (LUKE) in eastern Finland, Maaninka (63°09ʹ N, 27°20ʹ E). According to the World Reference Base of soils⁵¹, the LOM soil is classified as a Dystric Regosol, which covers 2% of the global land surface and the HOM soil is classified as a Histosol, typical of the northern latitudes, and covers 2.5% of the global land area, though 30% in Finland. The sampled bulk soils (0–20 cm soil depth) were immediately transported to the laboratory where the roots and remaining plant parts were removed manually. Then the soils were homogenized, sieved (4 mm), and stored at 4 °C until the start of the experiments. ¹⁵N tracer experiments were conducted with live (non-sterile) and sterile (autoclaved) soil conditions. More details are described in the Supplementary Methods.

¹⁵N tracer experiments

In all, 100 g of soil (live or sterilized) were transferred to a sterile petri dish (Ø 0.137 m, h = 0.017 m) with a sterilized spatula, and ¹⁵N tracer solution was added evenly to the soil surface with a pipette, immediately prior to the HONO measurements. We used three different tracers, ¹⁵NO₂⁻, ¹⁵NH₄⁺, and ¹⁵NO₃⁻, thereby establishing three different treatments: labeled nitrite (LNi), labeled ammonium (LA), and labeled nitrate (LNa), respectively. We also established a control treatment, where live and sterile soil samples received all three N forms at natural ¹⁵N abundance. More details are described in the Supplementary Methods.

HONO flux measurements and flux calculation

The ¹⁵N-amended samples were immediately transferred inside an opaque dynamic chamber, and HONO concentrations were measured with a Long Path Absorption Photometer instrument (LOPAP)⁵². The HONO flux was measured as described in Bhattarai et al.¹⁵. More details are provided in the Supplementary Methods.

Isotope pool dilution assays and determination of isotope ratios

The Isotope pool dilution (IPD) assays⁵³ were performed for all three N pools in live and sterile soil samples. For IPD assays, 4 g fresh soil (live or sterile) were weighed into 50 ml sterile polypropylene tubes (CELLSTAR®, Greiner BIO-ONE) and supplemented with 300 µl ¹⁵N tracer mix. Here, concentrations and atom% of tracer solutions and the soil moisture content were the same as used in the HONO flux experiments. After tracer addition, the tubes were vortexed for 2 minutes to allow the tracers to mix sufficiently in the soil matrix. The incubations were stopped by extraction with 30 ml cold (4 °C) 1 M KCl at zero, 30, 60, 90, 120, 180, and 240 minutes. The isotope ratios (¹⁵N/¹⁴N) in NH₄⁺ and NO₃⁻ were determined using the micro-diffusion method⁵³ in the soil extracts collected at zero, 90, 180, and 240 during the IPD experiment. More details are provided in the Supplementary Methods.

Collection and purification of HONO and NO₂ ⁻ azo dyes

The collection and purification of HONO and NO₂⁻ azo dyes were performed following Wu et al.²³, who established a method to analyze ¹⁵N APE in gaseous HONO. In the LOPAP instrument, HONO is reacted to an azo dye (C₁₈H₁₉O₂N₅S). Gaseous HONO is scrubbed in two sequential reactions, first with acidic sulfanilamide solution (R₁) in a two-channel striping coil, and second with N-(1-Naphthyl)ethylenediamine solution (R₂), which was collected separately from channel 1 and channel 2 after photometric detection in the LOPAP⁵². Each 5 ml of HONO azo dye solution was collected in 15 ml sterile polypropylene tubes (CELLSTAR®, Greiner BIO-ONE) at six sampling points that ended at 30, 60, 90, 120, 180, and 240 minutes after connecting the soil samples to the LOPAP. In parallel, NO₂⁻ azo dyes were produced from the soil extracts collected during the IPD experiment and standards using an identical approach for preparing azo dye²³. Immediately upon extract collection at each time point, 5 ml azo dye was generated by reacting 2.5 ml R₁ + R₂ mix (mixed at 1:1 (v:v) ratio) with 2.5 ml soil extract or standards. The R₁ and R₂ solutions used in the LOPAP instrument and for generating NO₂⁻ azo dyes in the soil extracts were of the same concentrations and the same chemical brand. The generation of NO₂⁻ azo dye from the soil extracts in the IPD assays was done for the first time in this study to understand the dynamics in the isotope ratios (¹⁵N/¹⁴N) of NO₂⁻ produced by different microbial N cycling pathways and to relate them to the isotope dynamics of HONO emitted from soils. HONO and NO₂⁻ azo dyes were purified by a reversed-phase extraction, according to Wu et al.²³, except for the volume of acetonitrile and milli-Q H₂O used for the preconditioning and final washing step of SPE (Solid Phase Extraction) columns. We used 1 ml acetonitrile and 2 ml milli-Q H₂O more than used by Wu et al.²³, to ensure the best cleaning of SPE columns (during preconditioning) and to maximize the removal of inorganic ions (in the final washing step). In brief, the pH of the azo dye solutions was adjusted to ~5 with 2 M sodium hydroxide and loaded onto 6 ml pre-conditioned SPE columns (Discovery® DSC-18 6 ML/500MG SPE, Sigma-Aldrich). The SPE columns were pre-conditioned by washing with 3 ml acetonitrile (HPLC grade, ≥99.8%, Thermo fisher Scientific) followed by 4 ml milli-Q H₂O (18.2 MΩ). The SPE columns loaded with azo dye were then washed with 4 ml milli-Q H₂O and stored at −20 °C for further analyses.

Elution and analyses of HONO and NO₂ ⁻ azo dyes

The frozen SPE columns were transported to the Terrestrial Ecosystem Research laboratory, University of Vienna, Austria, within one month after collection, where elution and analyses were performed. To avoid temperature effects (if any) during transportation, the frozen samples were transported in a sealed cooling box that contained ice bags. Immediately upon arrival, they were transferred to a −20 °C freezer until elution. The elution and analyses of HONO and NO₂⁻ azo dyes were different from those in Wu et al.²³. The fundamental difference was the instrument used to analyze the isotope ratios (¹⁵N/¹⁴N) in HONO and NO₂⁻ azo dyes. In our study, we analyzed HONO and NO₂⁻ azo dyes with an UPLC system (Ultimate 3000, Thermo Fisher Scientific, Bremen, Germany) coupled to an Orbitrap Exactive HCD MS (Thermo Fisher Scientific) with a mass resolution of 50,000. The choice of the device was made to optimize the detection of the ¹⁵N signal in azo dyes, as suggested by Wu et al.²³. The high resolution, high mass accuracy, and sensitivity of the Orbitrap-MS allowed us to separate the ¹⁵N and ¹³C isotopologue peaks that have a very small mass difference (∆_mass of 0.00632 Da)^54,55. This and similar instruments are widely used to study metabolites, including those in soil extracts⁵⁶. The choice of instrument (with electrospray ionization) eventually also led us to use eluents and reagents different from Wu et al.²³ for elution and chromatographic separation. Prior to elution, the SPE columns were transferred to room temperature for 20 minutes. Azo dyes were eluted with 5 ml eluent [80% methanol (HPLC grade, Sigma-Aldrich, ≥99.9%)+1% formic acid (98–100%, Merck Pro analysis)] using SPE vacuum manifolds. The eluates were diluted at 1:1 (v:v) ratio with milli-Q H₂O and analyzed on the same day of elution. More details are provided in the Supplementary Methods.

Calculation of ¹⁵N enrichment in HONO and NO₂ ⁻ azo dyes

The ¹⁴N and ¹⁵N peak areas in the samples and calibration standards were integrated manually with Xcalibur software and exported into Microsoft Excel sheets for further calculations. After blank correction of the ¹⁴N and ¹⁵N peaks of the samples and standards, we then calculated the concentrations (µM) of the samples using the peak area sum (area of ¹⁴N plus area of ¹⁵N peaks), based on the slope and intercept build from the unlabeled concentration calibration standards using equation 1. Next, we calculated the percentage (%) of isotopically heavy (i.e., ¹⁵N) molecules of total molecules by dividing the peak area of the heavy isotopologues by the peak area sum of heavy and light isotopologues, as shown in Eq. 2. The atom% ¹⁵N of the azo dyes of HONO and NO₂⁻ were quantified from a series of polynomial functions generated from % of the heavy atoms (theoretical ¹⁵N in NO₂⁻ azo dye of isotope calibration standards) and atom% ¹⁵N measured in NO₂⁻ azo dyes by LCMS at two concentration levels of nitrite, 1.56 µM and 25 µM, and their average. These polynomial functions were applied depending upon the concentration and % of heavy isotopologues in the azo dye samples, which adapted within and between samples over time (e.g., LA vs. LNa vs. LNi treatments). The calibration curves for concentration from natural abundance calibration standards and theoretical ¹⁵N atom% ¹⁵N in NO₂⁻ azo dyes and atom% ¹⁵N as measured by LCMS at 1.56 and 25 µM labeled calibration standards are shown in the Supplementary Methods and in Supplementary Figure 10.

$${\mathrm{Azo}}\,{\mathrm{dye}}\,{\mathrm{concentration}}\,(\mu {\mathrm{M}}) = \frac{{{\mathrm{peak}}\,{\mathrm{area}}\,{\mathrm{sum}} - {\mathrm{intercept}}}}{{{\mathrm{slope}}}}$$

(1)

$$\% {\mathrm{of}}\,15{\mathrm{N}}\,{\mathrm{atom}} = \frac{{{\mathrm{peak}}\,{\mathrm{area}}\,{\mathrm{of}}\,15{\mathrm{N}}\,{\mathrm{atom}}}}{{\left( {14{\mathrm{N}} + 15{\mathrm{N}}} \right){\mathrm{peak}}\,{\mathrm{area}}}} \times 100$$

(2)

Gross rates

Gross production and consumption rates of NO₂⁻, NH₄⁺, and NO₃⁻ were calculated according to Kirkham and Batholomew⁵³. More details are provided in the Supplementary Methods.

Source contribution and determination of abiotic and biotic HONO fluxes

The contributions (%) of NH₄⁺ oxidation, NO₃⁻ reduction, and organic nitrogen oxidation (the inset in Fig. 4) were calculated using the slopes (Fig. 2b–c) generated between ¹⁵NO₂⁻ and HO¹⁵NO from the live soil samples of the LNi, LNa, and LA (only in the HOM soil) treatments. The contribution of NH₄⁺ oxidation and NO₃⁻ reduction was obtained by dividing the respective slopes by the slope between ¹⁵NO₂⁻ and HO¹⁵NO in the LNi treatment. The contribution of organic N oxidation was obtained by subtracting the sum of NH₄⁺ oxidation and NO₃⁻ reduction from 100%. The contribution of a potentially unknown abiotic HONO source was quantified from the LNi treatment using Eqs. 3 and 4.

$${\mathrm{atom\% }}_{{\mathrm{mixture}}} = \left( {{\mathrm{atom\% }}_{{\mathrm{source}}1}} \right) \times f_1 + \left( {{\mathrm{atom}}\% _{{\mathrm{source}}2}} \right) \times f_2$$

(3)

$$f_1 = \frac{{({\mathrm{atom}}\% _{{\mathrm{mixture}}} - {\mathrm{atom}}\% _{{\mathrm{source}}2})}}{{({\mathrm{atom}}\% _{{\mathrm{source}}1} - {\mathrm{atom}}\% _{{\mathrm{source}}2})}}$$

(4)

In Eqs. 3 and 4, atom% represents the values of the mixture and the sources (i.e., source 1 and source 2), and the fractional contributions of source 1 and source 2 are represented by f₁ and f₂, respectively. Here, the mixture was HONO, source 2 was NO₂⁻ and source 1 assumed to be at natural ¹⁵N abundance, i.e., 0.3663 atom% ¹⁵N. The fractional contribution of an unknown source (i.e., f₁) was determined using Eq. 4, with the assumption that the sum of fractional source contributions is 1, and that the contribution of NO₂⁻ (f₂) is determined by 1−f₁. The abiotic and biotic HONO emissions were quantified using the HONO emissions from live and sterile LNi treatment. To obtain an estimate of the abiotic HONO flux, we quantified the HONO flux, which solely originated from NO₂⁻ and was subtracted from the total HONO emitted there, in the LNi treatment that used fractional contributions of NO₂⁻ to HONO emissions (ng N m⁻² s⁻¹) in the sterile soil samples. For the biotic HONO flux, HONO that originated from the NO₂⁻ added to the sterile soils was subtracted from the HONO measured from the live soil samples.

Total HONO

Total abiotic and biotic HONO represents the emission rates of soil HONO production averaged over 240 minutes of HONO flux measurements. To do that, first, total HONO in ng N m⁻² was obtained for all replicates separately. Here, the HONO emission rates (in µg N m⁻² s⁻¹) at each sampling point were multiplied by the time in seconds between the two consecutive sampling points, and their sum was calculated to obtain the total HONO. The total HONO (ng N m⁻²) thus obtained was divided by 14,400 (240 × 60) for all replicates separately to get the HONO emissions rate averaged over 240 minutes in ng N m⁻² s⁻¹. Finally, mean values and standard deviations were calculated for the abiotic (sterile soil samples) and biotic (live soil samples) sources.

Soil analyses

A detailed description of basic soil properties (Supplementary Table 1) and mineral N concentrations (Supplementary Figures 4–5) are provided in the Supplementary Methods.

Nucleic acid extractions, purification, and gene quantification

Nucleic acids in both soils were extracted from the control samples, which were used for HONO flux measurements in accordance with a protocol based on Yeates and Gillings⁵⁷ and Griffiths et al.⁵⁸. Real time quantitative polymerase chain reactions were used for gene quantification. Details on nucleic acid extractions and gene quantification are explained in the Supplementary Methods and in Supplementary Tables 5–6.

Statistics

Statistical analysis and graphical presentations were performed in R statistical software (R version 3.6.1), unless otherwise specified. Before performing a statistical test, the distribution of the data was assessed using the Shapiro–Wilk test, Q–Q plots, and histograms, and were transformed (log or square root) when required. More details are provided in the Supplementary Methods.

Data availability

All data produced in this study are available at https://doi.org/10.5281/zenodo.4503384.

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Acknowledgements

This study was funded by the Academy of Finland under the project “The origin of nitrous acid (HONO) emissions from northern soils and linkages to nitrogen cycle processes” (grant no. 297735) and the doctoral program of the University of Eastern Finland (grant to H. R Bhattarai). We also thank the Academy of Finland Center of Excellence program (project no. 307331) for supporting this study. We thank Jaana Rissanen for laboratory assistance and Pasi Yli-Pirilä for technical assistance. We highly appreciate all the laboratory and technical personnel for their help during our research visit in the Terrestrial Ecosystem Research laboratory at the University of Vienna. We also thank Natural Resources Institute Finland (LUKE) for providing the study sites and for providing data on soil texture.

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These authors contributed equally: Hem Raj Bhattarai, Wolfgang Wanek.

Authors and Affiliations

Department of Environmental and Biological Sciences, University of Eastern Finland, Kuopio, Finland
Hem Raj Bhattarai, Henri M. P. Siljanen, Jussi G. Ronkainen, Maarit Liimatainen, Hannu Nykänen, Christina Biasi & Marja Maljanen
Natural Resources Institute Finland, Production systems, Milk Production Unit, Maaninka, Finland
Hem Raj Bhattarai
Department of Microbiology and Ecosystem Science, Center of Microbiology and Environmental Systems Science, University of Vienna, Vienna, Austria
Wolfgang Wanek & Yuntao Hu
Water, Energy and Environmental Engineering Research Unit, University of Oulu, Oulu, Finland
Maarit Liimatainen
Natural Resources Institute Finland, Production systems, Oulu, Finland
Maarit Liimatainen
Lawrence Berkeley National Laboratory (LBNL), Berkeley, CA, USA
Yuntao Hu

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Hem Raj Bhattarai
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Contributions

H.R.B., W.W., C.B., M.L., H.N. and M.M. designed the experiments. ¹⁵N analyses of HONO and NO₂⁻ were performed by H.R.B, M.L, Y.H. and W.W., H.R.B. and C.B. analyzed the acid traps for ¹⁵NO₃⁻ and ¹⁵NH₄⁺_. H.R.B. measured the HONO flux. H.R.B., M.L. and M.M. analyzed the soil physicochemical properties. J.R. and H.S. did the quantification of functional genes. H.R.B. processed all the data and did the statistical tests and wrote the first draft of the manuscript. W.W. produced the synthesis data on global HONO flux rates and gross nitrite production rates from soils and contributed equally to the writing of the manuscript. All authors contributed, read, and approved the manuscript.

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Bhattarai, H.R., Wanek, W., Siljanen, H.M.P. et al. Denitrification is the major nitrous acid production pathway in boreal agricultural soils. Commun Earth Environ 2, 54 (2021). https://doi.org/10.1038/s43247-021-00125-7

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Received: 03 September 2020
Accepted: 09 February 2021
Published: 05 March 2021
DOI: https://doi.org/10.1038/s43247-021-00125-7

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