Introduction

Agricultural soils have become the main source of anthropogenic nitrous oxide (N2O), a powerful greenhouse gas and the single most important substance depleting stratospheric ozone1. Delaying the conversion of ammonium (NH4+) to nitrate (NO3−), nitrification inhibitors (NIs) have been suggested as a means to reduce N2O emissions from agricultural soils. NIs demonstrated their efficacy across different cropping soils2, but results vary widely, and in particular in pasture soils the use of NIs had no or little effect on N2O emissions3,4,5. Despite a growing body of research on NIs, mechanisms and factors determining their efficacy to reduce N2O emission remain poorly understood6. The challenges to understand these mechanisms derive from the fact that N2O is formed via several different pathways in the soil matrix7, tightly coupled to different processes of N supply and consumption8. Critically, N2O can be further reduced to N2 via the microbial-mediated process of denitrification, and the sole quantification of N2O as affected by NIs provides therefore only a limited insight into mechanisms of N2O mitigation using NIs.

Microbial metabolic pathways can contribute via a wealth of different processes to N2O production and consumption, i.e. the reduction to N2 in soils. Apart from abiotic processes, N2O formation can be categorized into nitrification-mediated pathways, denitrification and biotic formation of hybrid N2O9. Denitrification is generally assumed to be the main process contributing to overall N2O production from agricultural soils7,10,11,12 and is also the main process reducing N2O into environmentally benign N2 via the N2O reductase, the enzyme encoded by the functional nosZ gene. The reduction of N2O to N2 does not reduce overall N losses but limits the environmental impact of denitrification losses from agricultural soils. A reduction of N2O emissions by NIs can be attributed to (a) reduced N2O production via nitrification mediated pathways, (b) reduced N2O production via denitrification (c) increased consumption of N2O via denitrification, i.e., a shift in the N2O:N2 ratio towards N2. As these effects may overlap, a mechanistic understanding of the effects of NIs on N2O production and consumption processes needs to be based on N2O source partitioning, and the direct quantification of N2.

Most of the NIs inhibit the first and rate-limiting enzymatic step of nitrification, the conversion of NH4+ to hydroxylamine (NH2OH) via the ammonia monooxygenase13. The inhibition of nitrification means a reduced supply of N into the NO3− pool as the source pool of denitrification, but also an increase in NH4+ availability, leading to an increase of fertilizer N immobilization11 and mineralization/immobilization turnover rates14,15. Availability of N for N2O producing processes determines both production, but also consumption of N2O, as high NO3− availability shifts the N2O:N2 ratio of denitrification towards N2O16. The link between N transformation rates and N2O and N2 emissions is therefore critical to understand the effects of NIs in agricultural soils.

Typically, pulses of N2O are observed after fertilization and irrigation events. These pulses are short-lived and can account for more than 90% of cumulative N2O emissions from agro-ecosystems17, defining the critical time-window which determines the efficacy of NIs to mitigate N2O emission. Building on extensive research at the field scale conducted across different agro-ecosystems5,11,18,19,20, this study investigated the short-term effect of 3,4-dimethylpyrazole phosphate (DMPP) on N-turnover and N2O and N2 emissions from two contrasting agricultural soils in response to N-fertilization. We combined a 15N tracing analysis with the direct quantification of N2 and N2O emissions using the 15N gas flux method, complemented with the quantification of the nosZ gene via quantitative polymerase chain reaction (qPCR) in a soil microcosm study to constrain factors determining the efficacy of the NI DMPP to mitigate N2O emissions from agricultural soils.

Results

Physical and chemical properties for the two soils used in this experiment are shown in Table 1. The contrasting soils, a horticultural and a pasture soil, are henceforth referred to as sandy clay loam (sandy CL) and loam, according to their texture from 0–10 cm.

Table 1 Selected soil characteristics (0–10 cm) for a horticultural (Sandy clay loam) and a pasture soil (Loam) from subtropical Australia.
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Nitrogen transformations and soil microbial parameters

Gross N transformation rates were quantified with a 15N tracing model (Fig. 1) and differed markedly between soils when N-fertilizer was applied without the NI DMPP, referred to as the fertilizer only treatment (Table 2). Gross mineralization rates (Mtot) in the loam exceeded those in the sandy CL by a factor of 39. In the loam, Mtot was dominated by the mineralization of labile N (MNlab), while the mineralization of recalcitrant organic N (Mrec) dominated in the sandy CL. Gross nitrification (Nittot) was higher in the loam with 18.7 ± 0.03 μg N g−1 soil day−1 compared to 5.8 ± 0.03 μg N g−1 soil day−1 in the sandy CL. Autotrophic nitrification (ONH4) was the main pathway of NO3− production in both soils, as heterotrophic nitrification of organic N (ONrec) accounted for only 7% of Nittot in the sandy CL, and was negligible for Nittot in the loam. Immobilization of NH4+ (INH4tot) and NO3− (INO3) was higher in the sandy CL compared to the loam, and was dominated by INO3. In the sandy CL, only minor amounts of NO3− were recycled in the NH4+ pool via dissimilatory NO3− reduction to NH4+ (DNRA, referred to as DNO3 in the 15N tracing model), while DNO3 contributed with more than 2 μg N g−1 soil day−1 to NH4+ production in the loam. Microbial C (Cmic) and N (Nmic) as indicators for the size of the soil microbial biomass (SMB) were higher in the loam, exceeding Cmic and Nmic in the sandy CL by a factor of 5 and 7, respectively (Table 3).

Figure 1
figure 1

Conceptual 15N tracing model for the analysis of N gross transformations with the respective N transformations.

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Table 2 Gross soil N transformations (average ± standard deviation) in a horticultural (Sandy clay loam) and a pasture soil (Loam) after the application of NH4NO3 with and without the nitrification inhibitor DMPP. Means denoted by a different letter indicate significant differences for a specific N transformation across soils and treatments (i.e. no overlap of 95% confidence intervals).
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Table 3 Soil mineral N concentrations 30 minutes and 48 hours after N fertilizer application with and without the nitrification inhibitor DMPP; and dissolved organic C and soil microbial C and N prior and 48 hours after fertilizer application with and without DMPP in a horticulture and a pasture soil.
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Effect of DMPP on N-transformations and soil parameters

The application of N-fertilizer with DMPP had no significant effect on N transformations in the loam but changed N-turnover dynamics in the sandy CL (Table 2). DMPP reduced ONH4 only by 6% in the loam, but reduced ONH4 by more than 60% in the sandy CL. In the sandy CL, both Mtot and INH4tot increased, as well as the relative contribution of MNrec to Mtot, accounting for 80% of Mtot. INO3 decreased by 31%, while DNO3 increased by a factor of >5. DMPP did not affect the soil microbial biomass (SMB) but increased dissolved organic carbon (DOC) by 50% and 32% in the sandy CL and loam, respectively (Table 3).

Emissions of N2O and N2

The dominant N2O production pathway in both soils was denitrification, accounting for more than 90% of the N2O produced (Fig. 2). Over 48 hours, 0.24 ± 0.03 and 1.46 ± 0.38 μg N2O - N g−1 soil were emitted from the sandy CL and the loam, respectively. Both N2O emissions via denitrification (N2Od) and nitrification (N2On) were higher from the loam, exceeding those from the sandy CL by a factor of >8 (Fig. 2). Over the two day incubation period, 0.47 ± 0.09 μg N2 - N g−1 soil and 0.87 ± 0.11 μg N2 - N g−1 soil were emitted as N2 from the sandy CL and the loam, respectively. The main product of denitrification (N2Od + N2) from the sandy CL was N2, with N2Od accounting for 36% of total denitrification losses. Denitrification losses from the loam however were dominated by N2Od, accounting for 75% of total denitrification. There was no indication for hybrid production of N2O or N2.

Figure 2
figure 2

Cumulative emissions of N2O derived from nitrification (N2On) and denitrification (N2Od), cumulative N2 emissions, the product ratio of denitrification (N2O/(N2Od + N2) and the abundance of the nosZ gene encoding the N2O reductase from a horticultural soil (Sandy clay loam) and a pasture soil (Loam) after the application of NH4NO3 with and without the nitrification inhibitor DMPP.

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The response of the N2O reductase gene nosZ to fertilization and the use of DMPP

The abundance of nosZ prior to fertilization differed markedly between soils (Fig. 2). Copy numbers of nosZ in the loam exceeded those in the sandy CL by a factor of 6. After fertilization and the increase in soil moisture from 50% to 75% water-filled pore space (WFPS), nosZ copy numbers decreased in both soils, with a reduction by 77% and 32% for the sandy CL and the loam, respectively. DMPP did not affect nosZ abundance in the loam. DMPP however increased nosZ copy numbers by 227% compared to the fertilizer only treatment in the sandy CL.

Effect of DMPP on N2O and N2 emissions

DMPP significantly reduced N2O emission from the sandy CL but had no effect on N2O emissions from the loam (Table 4). DMPP reduced N2Od from the sandy CL by 46% (P < 0.05), but did not affect N2On (Fig. 2). There was no effect of DMPP on N2 emissions from the two soils. In the sandy CL, DMPP shifted the product ratio of denitrification (N2Od /(N2Od + N2)) to N2, decreasing the percentage of denitrification emitted as N2Od from 36% to 20%.

Table 4 Cumulative emissions of N2, N2O and CO2 from a horticultural soil (Sandy clay loam) and a pasture soil (Loam) after the application of NH4NO3 with and without the nitrification inhibitor DMPP.
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Discussion

The fertilization and irrigation of agricultural soils triggers a cascade of N transformations associated with pulses of N2O and N2 emissions. These short-term events are critical to understand the effects of NIs on N2O production and consumption in agricultural soils. Linking N turnover to emissions of N2O and N2 and the abundance of the N2O reductase gene nosZ in a short-term incubation demonstrated (a) that increasing NO3− availability after fertilization suppressed nosZ abundance, (b) that nosZ abundance, nitrification and N2 + N2O emissions remained largely unaffected by DMPP in the loam and (c) that DMPP decreased nitrification and increased nosZ abundance in the sandy CL, shifting the N2:N2O ratio towards N2. Our findings highlight the short-term effect of DMPP as highly soil specific, and show that reduced nitrification by DMPP can limit the suppression of the N2O reductase by high NO3− concentrations in the soil, enabling complete denitrification to N2.

Nitrogen transformation rates identified the loam as the more active soil regarding N turnover compared to the sandy CL (Table 2). Gross mineralization rates (Mtot) of more than 8 μg N g−1 soil day−1 together with a low immobilization of mineral N (INH4tot and INO3) denote high mineral N availability due to the rapid mineralization of organic N. This is further supported by the dominant contribution of the labile organic N pool to mineralization (MNlab), representing the microbial biomass and low molecular organic N compounds with a fast turnover. The high nitrification rates in the loam (>18 μg N g−1 soil day−1) denote rapid conversion of mineralized N to NO3− and show the dominant role of NH4+ oxidation for N-turnover in this soil. Gross mineralization was markedly lower in the sandy CL with Mtot at only 0.21 μg N g−1 soil day−1 and dominated by the mineralization of recalcitrant organic N, indicating limited and slower supply of mineral N via mineralization. Mineralization accounted for only 4% of nitrified N in the sandy CL, as compared to 45% in the loam, implying a rapid depletion of the NH4+ pool in the sandy CL. The observed differences between soils are consistent with microbial C and N contents (Table 3), indicating a larger soil microbial biomass in the loam and reflect the impact of perennial versus short term/annual and tilled versus undisturbed plant-systems on soil organic matter and microbial activity: Intensive tillage and irrigation in horticultural systems lead to loss of soil organic C21, while an extensive root system under permanent pasture is likely to promote microbial activity through constant inputs of C and N. These findings establish the differences in magnitude and relative importance of N transformations and microbial activity between the two contrasting soils.

The main source of N2O in both soils was denitrification, accounting for more than 90% of N2O produced (Fig. 2), which is in line with previous results from both field11 and laboratory studies10,12. The ability of soils to act as an N2O sink, i.e. the trait to reduce N2O to N2 has been linked to the abundance of nosZ, used as proxy for microorganisms involved in the reduction of N2O. In the study presented here, we compared nosZ abundance with direct measurements of N2 and N2O, evaluating the influence of DMPP on of microorganisms reducing N2O. The abundance of nosZ prior to fertilizer addition was higher in the loam, which is consistent with the reported positive correlation of nosZ copy numbers with soil organic C22. The synthesis of the N2O reductase is promoted by anoxic conditions23, and the increase in soil moisture together with the addition of fertilizer should have increased nosZ abundance. However, nosZ abundance decreased in both soils in the fertilizer only treatment (Fig. 2), indicating that increased NO3− availability due to fertilization and nitrification promoted the reduction of NO3− rather than N2O, shifting the N2Od/(N2Od + N2) ratio towards N2O. The magnitude and N2O:N2 partitioning of denitrification losses is consistent with the nitrification rates in both soils and as such shows the N2Od/(N2Od + N2) ratio as a function of soil intrinsic N – turnover. Cumulative N2Od losses of >2 μg N g−1 soil and 75% of denitrification (N2Od + N2) emitted as N2O from the loam show increased substrate availability for denitrification and simultaneous suppression of nosZ abundance by high NO3− availability (Fig. 2). In turn, lower denitrification losses with only 36% emitted as N2Od reflect slower N turnover in the sandy CL. These findings suggest that the suppression of the N2O reductase and increased N substrate availability are responsible for the large pulses of N2O from agricultural soils observed after fertilization and irrigation. Our results denote an increased risk of N2O loss from highly productive agricultural soils19, where increased mineralization of soil organic N due to fertilization, i.e., priming is likely to amplify the preferential reduction of NO3−, and as such the production of N2O via denitrification.

DMPP reduced N2O emissions from the sandy CL by more than 54% (Table 4). This is reflected in DMPP’s effect on autotrophic nitrification (ONH4) showing a reduction of 63% in the sandy CL (Table 2). The minor reduction of ONH4 by DMPP had however no effect on N2O emissions from the loam. In both soils, N2O derived from nitrification mediated pathways accounted for less than 15% of overall N2O, showing no response to the DMPP treatment. For the sandy CL, this suggests that DMPP primarily affected N2O production pathways indirectly, that is by reducing NO3− availability for denitrification, demonstrated by the reduction of N2O derived from denitrification by 46%. DMPP increased nosZ abundance in the sandy CL by a factor >2 compared to the fertilizer only treatment (Fig. 2). In the absence of direct N2 measurements, this effect has been interpreted as a shift of denitrification losses towards N224. Experimental evidence linking increased nosZ abundance with DMPP to N2 and N2O emissions25 is based on the acetylene inhibition method, which has been shown to lead to an irreproducible underestimation of denitrification rates9. Furthermore, acetylene itself is a potent NI, questioning the use of this method when investigating the effects of NIs on the magnitude and the N2Od/(N2Od + N2) ratio of denitrification. In the study presented here, DMPP reduced the N2Od/(N2Od + N2) ratio by 44% in the sandy CL, demonstrating a significant shift towards N2 (Fig. 2). These results link the increase of nosZ abundance in response to DMPP in the sandy CL to a shift in the N2Od:N2 ratio towards N2, based on direct measurements of N2 and N2Od using the 15N gas flux method. In contrast to previous incubation studies investigating N2O:N2 partitioning in response to DMPP26,27, emissions of N2O and N2 were quantified after incubation under atmospheric O2 conditions and without adding an easily available C source to stimulate denitrification, as these conditions would have altered short-term N dynamics in response to DMPP. Importantly, the shift towards N2 was not observed for the loam, where DMPP had a negligible effect on nitrification. Our findings indicate that the reduction of nitrification by DMPP in the sandy CL reduced the suppression of the N2O reductase after fertilization, enabling complete denitrification to N2. Emissions of N2O produced via nitrification mediated pathways were not affected by DMPP in this soil, showing the reduction of N2O emissions by DMPP as an indirect effect limiting NO3− availability for denitrification.

The spatial coverage of nitrifying microsites by the inhibitor is critical for efficient inhibition of nitrification. Limited diffusion of DMPP may explain the he observed inefficacy of DMPP to reduce autotrophic nitrification in the loam, which is consistent with reports from other pasture soils15. The amount of DMPP applied with N fertilizer is small, and the initial sorption to organic matter and uneven distribution of DMPP may hinder its short-term effectiveness to reduce nitrification in specific micro sites. Sorption of DMPP is likely to be more pronounced in the loam as a pasture soil with higher organic matter content as compared to the sandy CL owing to the positive correlation of DMPP sorption with organic C28,29. The high microbial activity in the loam also infers a larger number of microsites with nitrifying activity compared to the sandy CL, suggesting the spatial separation of DMPP from nitrifiers may be responsible for the short-term inefficacy of DMPP to reduce autotrophic nitrification in the loam. This theory is further supported by a study where DMPP did not affect the initial pulse of N2O after fertilization and irrigation from the loam, but reduced denitrification losses after that initial period11. This shows a delayed effect of DMPP in this soil, demanding further research on how diffusion in the soil matrix, sorption and distribution affects DMPPs efficacy to reduce autotrophic nitrification.

DMPP also affected non-targeted N transformation in the sandy CL: Mineralization and immobilization turnover was stimulated by DMPP, demonstrated by the five-fold increase of total mineralization (Mnrec + Mnlab) and the simultaneous increase of NH4+ immobilization (INH4rec + INH4lab) by a factor > 2 (Table 2). Increased mineralization/immobilization turnover has been reported after the application of DMPP15 and dicyandiamide (DCD)14 and can be attributed to higher NH4+ availability, stimulating microbial immobilization of NH4 (INH4lab) and mineralization of labile Norg (MNlab) to NH4+. This effect may further prime the mineralization of recalcitrant N (MNrec) in response to DMPP30. Interestingly, DMPP increased DOC availability in both soils, confirming previous results from a wheat-maize cropping system31 (Table 3). Increased MNrec in the sandy CL indicates mineralization of organic matter induced by DMPP contributed to higher DOC availability, but no such effect was observed for MNrec in the loam. Based on the data available, it remains unclear what caused the increase in DOC in response to DMPP. This increase has however important implications for N-turnover, in particular for the sandy CL as soil with limited labile C availability. DMPP increased DNRA by a factor >5 in the sandy CL, suggesting labile C promoted NO3− consumption via DNRA10,23. DNRA competes with denitrification for available NO3−, but the magnitude of DNRA in the sandy CL was insignificant regarding NO3− availability for denitrification. More importantly, labile C affects denitrification32, by supplying a reductant for denitrifiers, or through the stimulation of heterotrophic soil respiration, decreasing soil O2 levels and thus promoting denitrification. Furthermore, readily decomposable C can decrease the N2Od/(N2Od + N2) ratio of denitrification23. The increase in DOC observed in this study demonstrates an important non-targeted effect of DMPP, which can alter both rate and N2O:N2 partitioning of denitrification losses and therefore warrants further research.

Nitrification activity during pre-incubation increased NO3− levels in both soils. In the loam, NO3− levels were above those measured at the respective field site, which is also reflected in higher N2Od/(N2Od + N2) ratios11. This phenomenon often occurs in incubation studies, where the absence of plant uptake, pre-incubation33,34, and the addition of glucose26 increases NO3− levels in the soil. It is therefore important to consider N substrate availability when interpreting the effects of NIs on rate and N2O:N2 partitioning of denitrification losses. The mineral N levels in both soils indicate no N substrate limitation for denitrification regardless of the treatment. Under these conditions, DMPP had no effect on overall denitrification losses in both soils. The minor reduction of nitrification by DMPP in the loam did not reduce NO3− availability to a degree that limited preferential reduction of NO3−. The high initial NO3− values in the loam are also likely to have overwritten a significant reduction of nitrification. The reduction of N2O emissions, together with the increase of nosZ abundance in the sandy CL suggests however that DMPP lowered NO3− availability below a soil specific treshold35, limiting the preferential reduction of NO3− over N2O. The results from the sandy CL confirm the proposed mechanism of N2O reduction via a shift in the N2:N2O ratio26, and show that DMPPs inhibitory effect on nitrification can limit the suppression of the N2O reductase, promoting complete denitrification to N2.

The demonstrated link between nosZ and directly measured N2O and N2 emissions suggests that DMPP promotes the abundance of nosZ carrying denitrifiers. Including a comprehensive assessment of abundance and activity of nitrifying and denitrifying microbial communities in future research could further help to understand mechanisms of N2O mitigation by DMPP. Our study shows N dynamics in response to DMPP on a soil microcosm scale. This approach does not account for plant-microbe interactions and plant N uptake under field conditions but enables to isolate effects of NIs on key N transformations, with practical implications for the use of NIs in different agricultural soils. The relative magnitude of N2O emissions reflects cumulative losses observed from the same soils in the field, demonstrating a larger N2O mitigation potential for the pasture soil as compared to the horticultural soil. The short term inefficacy of DMPP to reduce nitrification in the pasture soils demands therefore improved strategies regarding rate and application of NIs. In soils with high organic matter content, and high soil intrinsic N turnover, repeated applications of DMPP, increasing the rate of DMPP, and/or the application of DMMP prior to fertilization may increase DMPPs efficacy, limiting the effect of N fertilizer priming on N2O emissions. Decreased nosZ abundance after fertilization and irrigation indicates suppression of the N2O reductase by increased NO3− availability, identifying NO3− availability as the control for the reduction of NO3− vs. N2O, which determines the magnitude of N2O losses. These findings apply to conditions of non-limiting NO3− availability for overall denitrification, which can be found in agricultural soils after N fertilization and irrigation when plant N uptake is limited. Under these conditions, the efficacy of NIs to mitigate N2O emissions depends on their ability to limit the suppression of the N2O reductase by high NO3− concentrations in the soil, enabling complete denitrification to N2.

Material and Methods

Soils and site

Soil samples (0–10 cm) were collected randomly (n = 4) from a vegetable cropping site (Gatton, Qld)20 and an intensively managed dairy pasture (Gympie, Qld)11 in subtropical Australia, referred to according to their texture in the first 10 cm as sandy clay loam (sandy CL) and loam, respectively. Site characteristics including physical and chemical soil properties are shown in Table 1. Soil samples were bulked, air dried and sieved to <4 mm and stored in a cold room at 4 °C.

Soil microcosms

Before treatment application, the soils were incubated in bulk for 4 days at a gravimetric water content of 30%. The experimental design consisted of two soils and two treatments: ammonium nitrate (NH4NO3) and NH4NO3 with DMPP (DMPP), each with four different 15N label combinations and four replicates. The NH4NO3 was applied with either (a) the NH4+ (15NH4NO3−) or (b) the NO3− (NH415NO3−) labeled at 10 atom %. NH415NO3− at 60 atom % (c) was used to quantify N2 emissions36, while non-labeled NH4NO3 (d) was used for the quantification of the SMB, DOC, and nosZ abundance. For the incubation, soil microcosms were established in centrifuge tubes (50 ml) using the equivalent of 8 g oven dry soil at a soil bulk density of 1 g cm−3. NH4NO3 equivalent to 35 µg N g−1 soil was applied in solution (1 ml) with 0.6% DMPP (w/w) added for the DMPP treatment. Additional water was applied to achieve the water-filled pore space (WFPS) of 75%. Water and fertilizer solutions were applied dropwise on two layers of 4 g of soil to ensure homogenous 15N labeling. After fertilization, centrifuge tubes were closed with Suba-seals (Sigma Aldrich) and were kept closed in an incubator at a constant temperature of 25 °C between gas sampling events. Additional soil microcosms (a and b, n = 4) were established for destructive sampling 30 minutes after fertilizer application.

Soil analysis

Soil mineral N

All soil mineral N extractions were conducted in the centrifuge tubes to avoid subsampling errors using 40 ml 2 M KCl (1:5 w/v ratio). Four soil microcosms per soil were extracted before fertilizer application to determine initial conditions. Soil microcosms a and b were extracted with 40 ml 2 M KCl, 30 minutes (t = 0) and 48 h (t = 2 days) after N fertilizer application. The centrifuge tubes were shaken with a horizontal shaker (150 rpm) for one hour, and extracts were filtered through Whatman no. 42 filter paper. After sample dilution, concentrations of NH4+ and NO3− were determined using colorimetric methods, NH4+ with a modified indophenol reaction37 and NO3− with the VCL3/Griess assay38. The 15N enrichments of the NH4+ and NO3− pool were determined for soil microcosms a and b by the diffusion method39.

Quantitative PCR analyses

For qPCR analysis, subsamples of 0.25 g of soil were taken prior to fertilizer application, and 48 h after (t = 2 days) from soil microcosms d and extracted immediately for total DNA using the PowerLyzer® PowerSoil® DNA Isolation Kit from MoBio (Mobio Laboratories, Inc., Carlsbad, CA, USA) according to the manufacturer’s instructions, with some minor modifications. Briefly, the soil was extracted twice by using the same soil and PowerBead Tubes to increase recovery of DNA. DNA concentration and quality were determined spectrophotometrically (NanoDrop 2000, Thermofisher, MA, USA). The two DNA aliquots from each sample were pooled before qPCR. The real-time PCR assay was carried out in a volume of 10 µl, and the assay mixture contained GoTaq® qPCR Master Mix (Promega, USA), 10 µM of each nosZ primer40 and 1 µl of pooled template DNA. Thermal cycling conditions for the nosZ2F (CGCRACGGCAASAAGGTSMSSGT) and nosZ2R (CAKRTGCAKSGCRTGGCAGAA) were as follows: an initial cycle of 95 °C for 3 min, 39 cycles of 95 °C for 15 s, 39 cycles of 60 °C for 45 s, 39 cycles of 72 °C for 45 s and 65 °C and 95 °C for 5 s. Each sample was quantified in triplicates using the iCycler iQ Real-Time PCR Detection System and the iQ 5 Optical System software (Bio-Rad Laboratories, Hercules, CA, USA).

Soil microbial biomass

Microbial C (Cmic) and N (Nmic) were quantified before and two days after fertilizer application using the chloroform fumigation-extraction41. Two aliquots of 3.5 g soil were sampled from each soil microcosm (d) with one aliquot subsequently fumigated with chloroform for 24 h. Fumigated and non-fumigated samples were extracted with 2 M KCl (1:10 w/v) and stored frozen until further analyses. Samples were acidified to remove inorganic C and analyzed for total N and organic C with an automated TOC/TN analyzer (TOC-V CPHE200V) linked with a TN-unit (TNM-1 220 V, Shimadzu Corporation, Kyoto, Japan). Cmic and Nmic were calculated as the difference in N and C between fumigated and non-fumigated samples without using a correction factor42. Dissolved organic C (DOC) was quantified as the amount of total C in the extracts of the non-fumigated samples.

Gas sampling and analysis

Air samples (n = 4) were taken daily before closing the centrifuge tubes to quantify ambient N2O concentrations. Specific background samples were taken above the respective soil microcosms treated with NH415NO3 at 60 atom % (c) for 15N2 analysis before closing the tubes. The entire headspace atmosphere was sampled 24 and 48 h after closure using a gas-tight syringe from soil microcosms a, b and c. After the 24 h gas sampling, the Suba-seals were removed for 10 minutes, allowing the headspace atmosphere to equilibrate10. Gas samples were transferred into pre-evacuated 12 ml exetainer tubes with a double wadded Teflon/silicon septa cap (Labco Ltd, Buckinghamshire, UK) and stored until N2O and CO2 analysis by gas chromatography (Shimadzu GC-2014). Gas samples from soil microcosms c were also analyzed for the isotopologues of N2 (15N14N, 15N15N) and N2O ([14N15N16O + 15N14N16O] and 15N15N16O) using an automated isotope ratio mass spectrometer (IRMS) coupled to a trace gas preparation unit (Sercon Limited, 20–20, UK).

Fluxes of N2, N2O and CO2

The triple labelling approach generates gas samples from three 15N fertilizer treatments with four replicates: a,b and c. Cumulative N2O and CO2 fluxes given in Table 4, were calculated based on gas samples from 15N fertilizer treatments a,b and c. Fluxes of N2 and N2Od, as well as denitrification losses (N2 + N2Od), were calculated based on the gas samples from treatment c. Calculating cumulative N2O fluxes based on 15N fertilizer treatments a, b and c or c alone did not result in significant differences. The reduction of N2O by DMPP in the sandy CL was significant regardless of the calculation chosen.

The flux rates of N2O and CO2 were calculated from the slope of the linear increase in gas concentration during the closure period and were corrected for temperature and air pressure20. The 15N enrichment of the NO3− pool undergoing denitrification (ap) and the fraction of N2 and N2O emitted from this pool (fp) were calculated following the equations given by Spott, et al.43 detailed in the supplementary material. The headspace concentrations of N2O and N2 were multiplied by the respective fp values giving N2 and N2O produced via denitrification (referred to as N2 and N2Od), with their respective fluxes expressed in g N2 or N2Od –N emitted g−1 soil day−1. Potential hybrid formation of N2 and N2O was found to be irrelevant30. The precision of the IRMS for N2 based on the standard deviation of atmospheric air samples (n = 18) at 95% confidence interval was 4.4 × 10−7 and 6.0 × 10−7 for 29R and 30R, respectively. The corresponding method detection limit ranged from 0.005 µg N2-N g−1 soil with ap assumed at 50 atom % to 0.014 µg N2-N g−1 soil with ap assumed at 20 atom %.

Gross N transformations

Gross N transformations were quantified using a 15N tracing model44 (Fig. 1), which uses a Markov Chain Monte Carlo method optimizing the kinetic parameters for the various N transformations by minimizing the misfit between modeled and observed NH4+ and NO3− concentrations and their respective 15N enrichments (soil microcosms a and b). The model considers five N pools including the NH4+ and NO3− pool, a labile (Nlab) and a recalcitrant (Nrec) organic N pool, and a pool for NH4+ adsorbed to cation exchange sites (NH4+ads). These pools are defined by 10 simultaneous occurring gross N transformations calculated by zero-, first-order or Michaelis-Menten kinetics (Table 2): The mineralization of Nlab and Nrec to NH4+ (Mnlab, MNrec), the immobilization of NH4+ to Nlab and Nrec (INH4-Nrec, INH4-Nlab), the adsorption (ANH4) and release (RNH4a) of NH4+ from NH4+ads, the oxidation of NH4+ to NO3− (ONH4), referred to as autotrophic nitrification; the oxidation of Nrec to NO3− (ONrec), referred to as heterotrophic nitrification; dissimilatory NO3− reduction to NH4+ (DNO3) and INO3, the immobilisation of NO3− to Nrec. Total mineralization was calculated as the sum of Mnlab and MNrec, total nitrification as the sum of ONrec and ONH4 and total immobilization of NH4+ as the sum of INH4-Nrec and INH4-Nlab.

Calculations and statistical analysis

The optimization routine used for the 15N tracing model gives a probability density function for each model parameter, which is used to calculated average values and standard errors of the mean. Average gross N transformation rates are obtained by integrating these values over the incubation period. Differences between N-transformations were assessed testing whether the 95% confidence intervals overlap45. The Benjamini Horchberg (BH) adjustment46 was performed to assess the effect of the different fertilization strategies on microbial C and N, DOC and nosZ gene abundance for each soil type. Analysis of variance was performed to assess differences in cumulative emissions of N2, N2O, total denitrification (N2 + N2O) and CO2 between soils within treatments and within soils between fertilization strategies. All values unless otherwise stated are given as mean ± standard error of the mean.