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Soil Biology and Biochemistry
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Fate of N-labelled ammonium nitrate with or without the new nitrification ■. inhibitor DMPSA in an irrigated maize crop
Guillermo Guardia^*, Antonio Vallejoa, Laura M. Cardenasb, Elizabeth R. Dixonb, Sonia García-Marco
a ETSI Agronómica, Alimentaria y de Biosistemas, Universidad Politécnica de Madrid, Ciudad Universitaria, 28040 Madrid, Spain b Rothamsted Research, North Wyke, Devon, EX20 2SB, UK
ARTICLE INFO
ABSTRACT
Keywords:
Fertilizer use efficiency Nutrient cycling Nitrogen losses N recovery 15N labelling
Nitrification inhibitors, originally proposed for nitrate leaching mitigation, are recommended as effective nitrous oxide (N2O) mitigation strategies. Several compounds have been trialled and used in the past including di-cyandiamide (DCD) or 3,4-dimethylpyrazole phosphate (DMPP). Yet, little is known about the new nitrification inhibitor 2-(3,4-dimethyl-1H-pyrazol-1-yl) succinic acid isomeric mixture (DMPSA). A field experiment using
15N single-labelled ammonium nitrate (15NH4NO3 or NH415NO3) was conducted to understand the effectiveness of DMPSA on a biochemical basis in an irrigated maize (Zea mays L.) crop. Gaseous fluxes, i.e. N2O, 15N2O, 15N2, methane (CH4), and carbon dioxide (CO2) were measured, as well as soil mineral N, yield components and 15N recovery in plant and soil. During the maize cropping period, the use of DMPSA significantly reduced cumulative
analyses revealed that most N2O losses (particularly during the emission peak) came from 15NH4NO3 (i.e. nitrification, nitrifier denitrification and/or coupled nitrification denitrification) rather than NH415NO3 in this calcareous low C-content soil. As expected, DMPSA decreased N2O losses from 15NH4+ oxidation, but an effect on non-target microorganisms was noticed, as shown by the significant reduction of respiration rates and N2O losses coming from 15NO3-. No significant effect of DMPSA on CH4 oxidation or 15N2 fluxes was observed. The DMPSA did not lead to a significant improvement of the dry weights of grain or biomass, although an increment of root biomass by 64% was found. This compound also tended to increase plant N recovery (average 67.8%) and to decrease soil N recovery (average 18.3%) but differences were not statistically significant. Conversely, the nitrification inhibitor significantly reduced the residual fertilizer-N in the 15-30 cm and 30-45 cm soil layers. The use of DMPSA was confirmed as a highly effective tool to reduce N2O emissions from irrigated crops in semiarid areas.
N2O emissions (118 g N ha 1) compared to ammonium nitrate without inhibitor (231 g N ha 1). The N
1. Introduction
Finding strategies to maximize nitrogen (N) use efficiency (NUE) and reduce N losses is a major goal for sustainable crop production (Sutton et al., 2013). These N losses involve water-soluble forms (e.g. nitrate, NO3-) and gaseous emissions (e.g. ammonia, NH3, nitric oxide, NO, nitrous oxide, N2O). Nitrous oxide is a persistent and harmful greenhouse gas (GHG) whose global warming potential is 265 times that of carbon dioxide (CO2) (IPCC, 2014). One strategy is the use of nitrification inhibitors, which delay the oxidation of ammonium (NH4 + ) to NO3-, thus increasing the opportunity for plant N uptake and reducing leaching of NO3- and emissions of N oxides (Quemada et al., 2013; Qiao et al., 2015), which are derived directly or indirectly from the nitrification process. Therefore, the use of nitrification
inhibitors appears to be a good strategy to achieve an optimal balance between farm and environmental costs (Abalos et al., 2014a; Thapa et al., 2016; Yang et al., 2016).
The efficiency of nitrification inhibitors reducing N2O losses has been demonstrated under several pedo-climatic and management conditions (Feng et al., 2016; Gilsanz et al., 2016). Most previous studies have focused on dicyandiamide (DCD) or 3,4-dimethylpyrazole phosphate (DMPP). Conversely, little is known about the new inhibitor 2-(3,4-dimethyl-1H-pyrazol-1-yl) succinic acid isomeric mixture (DMPSA). This novel nitrification inhibitor differs from DMPP in the presence of the succinic group instead of phosphate. The combination with succinic acid results in (i) less volatilization of DMP; (ii) a smoother and prolonged availability of DMP in the soil; (iii) combination with any mineral fertilizer (e.g. urea, calcium ammonium nitrate,
* Corresponding author. E-mail address: guillermo.guardia@upm.es (G. Guardia).
http://dx.doi.org/10.10167j.soilbio.2017.10.013
Received 23 May 2017; Received in revised form 7 September 2017; Accepted 14 October 2017
0038-0717/ © 2017 Published by Elsevier Ltd.
diammonium phosphate) and (iv) the same inhibitory effect as established nitrification inhibitor for a lower concentration (CA 2933591 A1 2015/06/18 Patent; Pacholski et al., 2016).
Previous meta-analysis studies provided an average inhibition efficacy (ranging from 30 to 42%), but it has been demonstrated that the effectiveness of nitrification inhibitors in abating N2O emissions is highly dependent on soil type and weather (Mahmood et al., 2011; Misselbrook et al., 2014; Abalos et al., 2017). Environmental conditions affect the relative influence of the distinct biochemical processes which cause the release of N2O. Therefore, it is important to accurately determine the dominant processes leading to N oxide emissions in each specific cropping area, in order to i) understand the mechanisms which drive nitrification inhibitors efficiency; and ii) find if other alternative strategies (e.g. changing N source) can be implemented with similar mitigation performance and lower monetary costs than nitrification inhibitors (Migliorati et al., 2014; Harty et al., 2016). In this sense, the use of stable isotope 15N techniques emerged as an accurate and useful tool to discriminate between different possible N2O sources (Stevens et al., 1997). When using the two single-labelled ammonium nitrate (NH4NO3) forms (i.e. 15NH4NO3 and NH415NO3) it is possible to quantify the importance of the processes derived from 15NH4 + i.e. nitrification, nitrifier denitrification or coupled nitrification-denitrifica-tion; and 15NO3- i.e. denitrification or dissimilatory NO3- reduction to NH4+ (DNRA, Wrage et al., 2001; Medinets et al., 2015).
Additionally, the use of single 15N labelled fertilizers in nitrification inhibitor experiments provides an opportunity to determine if nitrification inhibitors have a short-term effect on non-target microorganisms (Florio et al., 2016), particularly on the denitrifying community (thus affecting N oxide losses from denitrification). So far, little information has been published with regards to the effect of nitrification inhibitors on denitrifiers, but the recent study of Kou et al. (2015) reported a decline in the copy numbers of the genes encoding the nitrite reductase (which catalyzes the reduction of nitrite to gaseous nitric oxide), i.e. NirS and NirK, in the presence of a pyrazole-based inhibitor (DMPP). The measurement of N2 can also help to elucidate if nitrification inhibitors affect the denitrification process and the release of the end product of this stepwise reduction reaction under propitious conditions, e.g. irrigated soils. So far, the measurement of N2 has been hardly addressed because of its high atmospheric background concentrations (Van Groenigen et al., 2015). The 15N gas flux method appears as a more reliable quantification method than the acetylene (C2H2) inhibition technique (Stevens and Laughlin, 1998), but only a few field studies using 15N labelling have been carried out so far in field conditions, particularly in semi-arid agro-ecosystems (Buchen et al., 2016).
In this context, a field experiment was carried out aiming to measure the effect of the new inhibitor DMPSA on N2O losses in an irrigated crop. Concurrently, methane (CH4) and soil respiration were measured to detect if changes on soil mineral N due to the application of DMPSA could affect CH4 uptake capacity (Aronson and Helliker, 2010) and nontarget microorganisms (not only those involved in the N cycle), respectively. Single-labelled 15NH4NO3 and NH415NO3 were applied in parallel microplots to study the main biochemical processes involved in the emission of N2O, aiming to discover the most effective mitigation strategies. At the end of the experiment, 15N in the plants and the residual 15N in the soil were also measured. We hypothesized that DMPSA would be efficient in the abatement of N2O emissions in a C-poor soil, in which processes derived from NH4+ oxidation would be confirmed as the main N2O contributors, even under irrigated conditions (Aguilera et al., 2013).
2. Materials and methods 2.1. Site description
The study was carried out at "El Encin" field station in Madrid
(latitude 40° 32'N, longitude 3° 17'W). The soil was a Calcic Haploxerept (Soil Survey Staff, 1992) with a sandy clay loam texture (clay, 28%; silt, 17%; sand, 55%) in the upper horizon (0-28 cm) with vermiculite as a dominant clay mineral. Some relevant characteristics of the top 0-28 cm soil layer are as follows: total organic C, 8.1 ± 0.3 g kg-1; pHH2O, 7.6; bulk density, 1.1 ± 0.1 g cm-3; and CaCO3, 13.2 ± 0.4 g kg-1. At the beginning of the experimental period, the NH4+content was 1.0 ± 0.7 mg NH4+-N kg soil-1 and the NO3- content was 9.6 ± 1.1 mg NO3--N kg soil-1. The site has a semiarid Mediterranean climate with a dry and hot summer period, and the mean annual temperature and rainfall (over the last 10 years) in this area is 13.2 °C and 460 mm, respectively. The soil temperature was monitored using a temperature probe (SKTS 200, Skye Instruments Ltd., Llandrindod Wells, UK) inserted 10 cm into the soil.
2.2. Experimental design and management
The experiment was conducted from April to October 2015 in a maize crop. A total of 12 microplots (1 m x 1 m) were selected and arranged in a complete randomized design with three replicates. Each microplot received labelled 15NH4NO3 or NH415NO3 (98 atom % 15N, Campro Scientific GmbH, Berlin, Germany) and unlabelled NH4NO3, with and without the nitrification inhibitor 2-(3,4-dimethyl-1H-pyr-azol-1-yl) succinic acid isomeric mixture (DMPSA), provided by EuroChem Agro. Those treatments were applied as a top-dressing (on 24th June) at a rate of 180 kg total N ha-1 (half as 15NH4NO3 or NH415NO3 and the other half as NH4NO3). A solution of 2 L (24.5 atom % 15N) of each treatment was prepared and homogenously applied over the soil surface of the microplots with a hand sprayer. The proportion of DMPSA in the fertilizer was that recommended by the manufacturer, i.e. 0.8% of the NH4+-N applied. The treatments applied, their designation and application rate are summarized in Table 1. Additionally, a control treatment with no N fertilization (C) was included to compare GHG emissions. Those microplots were installed in the middle of larger plots (covering an area of 45 m2 each) that were fertilized with un-labelled commercial fertilizers (CAN, CAN + DMPSA and no N fertilization), following the same rate as that applied in its appropriate microplot. Each microplot was surrounded by an outside squared boundary (1 m x 1 m) and a row of six maize plants was in the middle of the microplot.
A cultivator pass was performed before sowing (13th April 2015). Maize (Zea mays L. FAO class 600) was sown on 17th April 2015 with a plant density of 7.50 plants m-2. A basal fertilization was applied on 14th April 2015, by hand at the rate of 50 kg P ha-1 and 150 kg K ha-1 as Ca(H2PO4)2 and K2SO4, respectively, in all plots. Irrigation was carried out using a 12 m x 12 m sprinkler irrigation system at a height of 2.5 m. A total amount of 705 mm of water was applied from late May to early September in 31 irrigation events. The field was kept free of weeds, pests and diseases, following local practices (e.g. herbicides, pesticides, etc.). The maize was harvested on 28th October 2015.
2.3. GHG sampling and analyses
Fluxes of N2O, CH4 and CO2 were measured during the maize cropping cycle using opaque manual circular static chambers as
Table 1
Treatments applied in 15N microplots (N rates correspond to the labelled and unlabelled AN applied).
Treatment applied Designation N rate (kg N ha-1)
15NH4NO3 + h NH4NO3 15AN 90 + 90
nh415no3 + h NH4NO3 A15N 90 + 90
15NH4NO3 + h NH4NO3 + DMPSA 15AN + DMPSA 90 + 90
nh415no3 + h NH4NO3 + DMPSA A15N + DMPSA 90 + 90
described in detail by Sanz-Cobena et al. (2014). The chambers (diameter 35.6 cm, height 11 cm) were hermetically closed for 1 h, by fitting them into stainless steel rings, which were inserted at the beginning of the study into the soil to a depth of 10 cm to minimize the lateral diffusion of gases and avoid the soil disturbance associated with the insertion of the chambers in the soil (Hutchinson and Livingston, 2001). The rings were only removed during management events. One chamber was located in each microplot.
Gas samples were taken 6 times per week during the first week after fertilization. During the second week after fertilization, GHGs were sampled three times. The gas sampling frequency was then gradually decreased until the end of the experiment, but samples were taken after all irrigation or rainfall events and at the same time of day (10-12 a.m.) in order to minimize any effects of diurnal variations in the emissions (Reeves and Wang, 2015). Gas samples were taken in 20 ml vials at 0, 30 and 60 min to test the linearity of gas accumulation in each chamber. Measurements of N2O, CO2 and CH4 concentrations in the vials were made to estimate the emissions. The increases in N2O, CH4 and CO2 concentrations within the chamber headspace were generally linear (> 90% of cases, particularly when the highest fluxes or emission peaks were reported, R2 > 0.90) during the sampling period (1 h). In the case of nonlinear fluxes, linear regressions were performed, since it has been described as the recommended option for three sampling points (Venterea et al., 2012).
The concentrations of N2O, CO2 and CH4 were quantified by gas chromatography, using a HP-6890 gas chromatograph (GC) equipped with a headspace autoanalyzer (HT3), both from Agilent Technologies (Barcelona, Spain). HP Plot-Q capillary columns transported the gas samples to a63Ni electron-capture detector (ECD) to analyze the N2O concentrations and to a flame-ionization detector (FID) fitted with a methanizer to analyze the CH4 and CO2 concentrations. Cumulative gas emissions during the experimental period were calculated as described by Menéndez et al. (2006), by multiplying the average flux of two successive determinations by the length of the period between sampling and adding that amount to the previous cumulative total.
2.4. Soil and plant sampling and analyses
To relate gas emissions to soil properties, soil samples were collected from two depths (0-2 cm and 2-10 cm) during the experimental period on almost all gas-sampling days. One soil core was randomly sampled close to the ring in each microplot and then mixed and homogenized in the laboratory. The soil NH4 + -N and NO3--N concentrations were analyzed using 8 g of soil extracted with 50 mL of KCl (1 M) and measured by automated colorimetric determination using a flow injection analyzer (FIAS 400 Perkin Elmer) with a UV-V spectro-photometer detector. Soil dissolved organic C (DOC) was determined by extracting 8 g of homogeneously mixed soil with 50 mL of deionized water. Afterwards, DOC content was analyzed using a total organic C analyzer (multi N/C 3100 Analytik Jena) with an IR detector. Water-filled pore space (WFPS) was calculated by dividing the volumetric water content by the total soil porosity, assuming a particle density of 2.65 g cm-3. The gravimetric water content was determined by oven-drying soil samples at 105 °C with a MA30 Sartorius® moisture analyzer.
In each microplot there were six maize plants, and only the four middle plants were sampled and used in the analysis to avoid the border effects. Aerial biomass was cut by hand at soil level, and then above-ground biomass (stem + leaf + cob) was separated from the grain. Also, a root situated in the middle of each microplot was sampled and washed with water to separate roots from soil. Dry weights of the different parts of maize plant were determined, those of the roots after drying at 60 °C. After maize harvesting, soil samples at three depths (0-15 cm, 15-30 cm and 30-45 cm) were also collected. Both, soil and maize samples were air-dried and finely ground to determine total N, 15N in plant and residual 15N in soil as described in Section 2.5.2.
2.5. 15N isotope analyses and calculations
2.5.1. 15N2O and 15N2
Three 12 ml vials (Exetainers, Labco, Lampeter, UK) containing gas from the headspace at 60 min per microplot and per sampling date were collected during the 2 weeks following N addition. One of these replicated vials was used to measure N2O flux as explained in section 2.3. The other two vials corresponding to 3, 4, 6, 8, 10 and 13 days after fertilization (during the N2O emission peaks) were sent to Rothamsted Research North Wyke to measure the 15N enrichment of N2O and N2, by using a TG2 trace gas analyzer interfaced to a 20-22 isotope ratio mass spectrometer (both from SerCon Ltd., Crewe, UK). For the N2O analysis, solutions of 6 and 30 atom% ammonium sulphate ((NH4)2SO4) were prepared and used to generate 6 and 30 atom% N2O (Laughlin et al., 1997) and used as reference and quality control standards. During the experiment, the mean natural abundance of atmospheric N2O (0.3663 atom% 15N) was subtracted from the 15N enrichment of the samples to calculate the atom percent excess (ape). To obtain the N2O flux that was derived from fertilizer (N2O —Nff), the following equation was used (Senbayram et al., 2009):
>Tn .T .T „ {N2 OapeSample)
N2O - Ndff = N2O - N X I-— I
apefertilizer I
in which 'N2O — N' is the N2O emission from soil expressed as mg m-2 d-1, 'N2O apesample' is the 15N atom% excess of emitted N2O, and 'apefertjljzer' is the 15N atom% excess of the applied fertilizer (Senbayram et al., 2009). To obtain the N2O flux that was derived from soil (N2O —Ndfs), the following equation was used:
N2O — Ndfs = (N2O-N) - (N2O-Ndf15AN) - (N2O-NdfA15N)
In the above equation, 'N2O -Ndf15AN and N2O -NdfA15N' is the N2O flux that was derived from fertilizer 15AN or A15N, respectively.
The amount of N2 evolved was calculated from the ratios of the intensity of ion beams at mass to charge ratios 28, 29 and 30, using the equations of Stevens and Laughlin (1998).
N2 evolved (mg) = d x N2 in headspace / (1-d)
in which:
'N2 in headspace (mg)' was calculated using the ideal gas law (PV=nRT), where P = 1 atm, T = 294.3 K (mean temperature of closed chambers), V = headspace volume x 78.085% (percentage of N2 in the air) and molecular weight of N2 is 28.013 g.
d' is the fraction proportional to the amount of 15N-labelled N2 in the headspace:
d = 5 R / ( Xn)2
15Xn = 2(530R / 529R) / (1+ 2530R / 529R)
' 15Xn' is the mole fraction of NO3 in the soil that is being denitrified; S29R = 29N2/28N2 in the headspace - 29N2/28N2 in atmospheric air; S30R = 30N2/28N2 in headspace - 30N2/28N2 in atmospheric air. The amount of N2 evolved was marked as below the detection limit if S29R < 8.0 x 10-6 or S30R < 3.2 x 10-7 (Stevens and Laughlin, 2001). The approximate detection limit for N2 fluxes was 12.8 mg N-N2 m- 2 d- 1 for this chamber type, enclosed for 1 h and with a total enrichment in the soil of 24.5% atom 15N.
2.5.2. Total N and 15N in plant and soil
Samples of maize plants (separately for roots, grain and above-ground biomass) and soil were air-dried and finely ground (< 100 mm) with a ball mill (Retsch MM 400, Retsch GmbH, Haan, Germany). Total N and 15N content in the soil and plants was determined in at the Interdepartmental Investigation Service (SIdl) at Autónoma University of Madrid by combustion of the samples using an Elemental Analyzer Thermo 1112 Flash HT hyphenated interfaced to an Isotope Ratio Mass
Spectrometer Thermo Delta V Advantage (Bremen, Germany).
The contribution of each source (15AN/A15N) in each treatment (AN or AN + DMPSA) was calculated following Reddy and Reddy (1993). Briefly, the N in each part of maize or in each soil layer that was derived from fertilizer 15AN or A15N was:
%15AN (or NdfA!5N) = TN X-
Atom% 15N excess . Atom% 15N excessso
Where 'TN' is the total N uptake in the corresponding part of the plant or the total N in each soil layer (taking into account the layer thickness and the bulk density). The 'Atom % 15N excess' was calculated by subtracting the atom% 15N in the natural standard (air) to the atom% 15N in the soil, plant tissue or fertilizer (NH4NO3). The total fertilizer recovery in maize plants was calculated as the sum of the total (i.e. roots + green biomass + grain) Ndf15AN and NdfA15N. The residual fertilizer in the soil was calculated by adding N df15AN and NdfA15N, in the three sampled layers (i.e. 0-45 cm depth, see section 2.4).
Fig. 1. Soil water-filled pore space (WFPS) and soil temperature at 10 cm depth during the first 36 days after N fertilization (during the N2O peak, see Fig. 3). Vertical bars indicate standard errors.
2.6. Statistical methods
Data analysis was performed using Statgraphics Plus v. 5.1. Analyses of variance (one-way ANOVA for 15N in plant and soil, two-way ANOVA for 15N2O and 15N2) were performed for all variables throughout the experiment. Data distribution normality and variance uniformity were previously assessed by the Shapiro-Wilk test and Levene's statistic, respectively, and log-transformed before analysis when necessary. The means were separated by the LSD test at P < 0.05. For non-normally distributed data, the Kruskal-Wallis test was used on non-transformed data to evaluate the differences at P < 0.05. Linear correlations were carried out to determine the relationships between the gas fluxes (N2O, N2, CH4, and CO2) and WFPS, soil temperature, DOC, NH4 + -N and NO3--N. These analyses were performed using the mean/cumulative data of the replicates of all treatments (n = 12), and also for all the days when the soil and GHG were simultaneously sampled (n = 22).
3. Results
3.1. Environmental conditions, soil mineral N and DOC
The mean soil temperature (at 10 cm depth) during the maize cropping period was 20.5 °C, a typical value in the experimental area. During the whole maize cycle (6 months), soil WFPS ranged from 18 to 76% (data not shown), while values from one day before N fertilization to 36 days after N fertilization (during the N2O emission peak, see section 3.2) ranged from 35 to 63% (Fig. 1).
Soil NH4 + contents increased after N addition, decreasing thereafter to values below 10 mg N kg soil-1 from 22 days after fertilization until the end of the experimental period (Fig. 2a). During the first two months after N fertilization (Fig. 3a), the nitrification inhibitor DMPSA significantly increased the average NH4+ content in both 0-2 cm and 2-10 cm layers (by 48% and 60%, respectively). This was also observed in almost all sampling dates (Fig. 2a). The NO3- contents, were significantly reduced in the AN + DMPSA treatment compared to AN. That was observed in almost all sampling dates (except at 1, 8 and 10 days after N fertilization, Fig. 2b) and for average concentrations (Fig. 3b) in both 0-2 cm (43% reduction, P < 0.05) and 2-10 cm soil layers (42% reduction, P < 0.05). The NH4+ and NO3- contents in the upper layer exceeded those of the 2-10 cm layer by an average factor of 3.3 and 2.0, respectively. During the maize cropping period, DOC contents of the topsoil (0.10 cm) ranged from 76 to 123 mg C kg dry soil-1, and no significant differences were found between treatments (data not shown).
3.2. GHG emissions
During maize cropping period, N2O emissions ranged from 0.0 to 4.2 mg N m-2 d-1 (Fig. 2c). Before N fertilization, and from one month after N addition to the end of the experimental period, fluxes were < 0.1 mg N m-2 d-1; therefore, only the days during the peak N2O emission have been represented in Fig. 2c in order to improve its visualization. Fluxes increased 3 days after N addition, and the greatest N2O peak occurred 6 days after fertilization. The nitrification inhibitor DMPSA significantly decreased (by 49%) cumulative N2O fluxes at the end of the experimental period: these were 231 and 118gNha-1 for AN and AN + DMPSA, respectively. Fluxes from the AN + DMPSA treatment did not differ significantly to those from the C treatment. Nitrous oxide emissions were strongly correlated with respiration fluxes (P < 0.001, n = 22, r = 0.90) and WFPS (P < 0.05, n = 22, r = 0.59) throughout the maize cropping period.
Negative CH4 fluxes were generally observed throughout the maize cropping season. A positive correlation between CH4 emissions and NH4+ concentrations was found (P < 0.05, n = 22, r = 0.54). Since no significant differences were observed between treatments for cumulative CH4 oxidation (data not shown), the nitrification inhibitor also resulted in a significant mitigation of CO2 equivalent emissions (52% reduction), taking into account the global warming potential of N2O (265 times that of CO2) and CH4 (28 times that of CO2) over 100 years (IPCC, 2014). Respiration rates ranged from 0.5 to 4.2 g C m-2 d- 1 (data not shown). Maximum fluxes were observed 6 days after N fertilization, as for N2O emissions. Consequently, a significant and high correlation between both fluxes (P < 0.001, n = 20; r = 0.90) was obtained throughout cropping season. As for N2O emissions, cumulative respiration fluxes were significantly lower in AN + DMPSA (1.2 Mg C ha-1) than in AN (1.6 Mg C ha-1).
3.3. Contribution of different sources to N2O emissions
The amount of N2O coming from exogenous NH4 + , exogenous NO3- and endogenous soil N during the week in which the peak occurred is shown in Fig. 4. In the AN treatment, the 15N2O analysis revealed that 6 days after fertilization (when the highest N2O increment was noticed, see Fig. 4a) the percentages of N2O losses coming from 15AN, A15N and soil N were 90, 10 and 0%, respectively. On day 3 after N fertilization (when a smaller peak occurred), those relative contributions were 41, 12 and 46% for 15AN, A15N and soil, respectively. Therefore, the relative contribution of exogenous NH4+ to N2O losses was significantly higher than that of exogenous NO3- , during the days when 15N2O was analyzed.
In the case of the AN + DMPSA, endogenous soil N was the main
Fig. 2. NH4+ (a) and NO3- (b) contents in the 0-10 cm soil layer; and daily N2O fluxes (c) during the first 36 days after N fertilization for the different treatments: ammonium nitrate (AN), AN + DMPSA (AN + DMPSA) and control (C). Vertical bars indicate standard errors.
source of N2O losses even during the days of highest N2O emissions (Fig. 4b), and its relative contribution ranged from 70 to 80% of N2O during the first two weeks after N fertilization. No significant differences between the amount coming from exogenous NH4+ and exogenous NO3- were observed in this treatment. Comparing both fertilized treatments (AN and AN + DMPSA), the use of DMPSA decreased N2O losses from NH4+ (P < 0.05), and emissions from exogenous NO3- were also significantly decreased in the DMPSA-based treatment. Furthermore, the nitrification inhibitor also tended to lower the emissions coming from endogenous soil N, but differences were not statistically significant.
3.4. 15N2 fluxes
Daily 15N2 fluxes ranged from 50 to 150 mg N m-2 d-1 (Fig. 5), and were highest after the N2O emission peaks (6 days after fertilization, Fig. 2c). Cumulative fluxes in the first two weeks after N addition
ranged from 9.6 to 11.7 kg N ha-1, and no significant differences were observed between treatments. The same accounted for daily N2 flux in most of the sampling dates. Only at 3 and 4 days after N fertilization, significantly higher fluxes of 15N2 came from AN than from AN + DMPSA (P < 0.05). Daily 15N2 and N2O fluxes were strongly negatively correlated (P < 0.05, n = 6, r = -0.80).
3.5. Dry weight and 15N in plant and soil
The new inhibitor DMPSA tended to increase the dry weigh of grain, green biomass and root biomass, but it was only significant in the latter (Fig. 6). On average, 67.8% and 18.3% of the N applied through the fertilizer was recovered in maize (roots + aboveground biomass) and soil, respectively (Table 2). The DMPSA tended to increase the plant recovery and to decrease soil recovery, but it was not statistically significant. The tendency of soil recovery to reduce in the AN + DMPSA treatment was caused by the significant reduction of residual N from
Fig. 3. Average NH4 + -N (a) and NO3- -N (b) concentrations in the 0-2 cm and 2-10 cm soil layers during the first two months after application of ammonium nitrate (AN) with and without the inhibitor DMPSA. Different letters within columns indicate significant differences within each period of the year, by applying the LSD test at P < 0.05. Vertical bars indicate standard errors.
fertilizer in the 15-30 and 30-45 cm layers (Fig. 7).
4. Discussion
4.1. Effect of DMPSA and N substrate on gaseous N fluxes
The nitrification inhibitor DMPSA showed high effectiveness in reducing N2O losses (49% abatement). Even though other nitrification inhibitors such as DCD, DMPP or nitrapyrin have been broadly evaluated and showed positive effect (Akiyama et al., 2010), information of the performance of this new inhibitor is scarce. Our results revealed even higher mitigation efficiency of DMPSA than that reported by Gilsanz et al. (2016) for DCD (average 42%) and DMPP (average 40%) in a meta-analysis from 39 recent studies. This result could be due to i) higher efficacy of inhibitors under management conditions which lead to high N2O losses (e.g. irrigated crops in semi-arid areas, with continuous drying-rewetting cycles) (Abalos et al., 2014a; Guardia et al., 2017a); ii) notable effect of nitrification inhibitors on sandy-clay loam soils, with an average 50.0% ± 12.4 mitigation (Gilsanz et al., 2016); and iii) the chemical properties of DMPSA (succinic group), as explained in Section 1 (Introduction). The study of Huérfano et al. (2016) evaluated the use of DMPSA with ammonium sulphate in a rainfed wheat crop under Mediterranean humid conditions, finding that this inhibitor reduced N2O losses (by 11-56%). Our results suggest that the use of DMPSA could be a promising mitigation option under humid or irrigated conditions, which normally lead to higher N2O emission factors in semi-arid areas (Cayuela et al., 2017).
As expected, the 15N2O analysis revealed that the application of
Fig. 4. Amount of N2O coming from 15AN, A15N (see Table 1) and soil N in (a) ammonium nitrate (AN) and (b) AN + DMPSA. Vertical bars indicate standard errors.
Fig. 5. Daily 15N2 fluxes for the different treatments (see Table 1). Vertical lines indicate standard errors. "NS" and * mean no significant and significant at P < 0.05, respectively.
Fig. 6. Dry weight of grain, green biomass (stem + leave + cob) and root biomass in ammonium nitrate (AN) and AN + DMPSA.
DMPSA significantly decreased the amount of N2O emissions coming from 15NH4NO3 (Fig. 4). Therefore, the inhibitor was an effective technique to reduce N2O emissions arising from direct NH4 + oxidation.
Table 2
Recovery of N applied through ammonium nitrate (AN) or AN with the nitrification inhibitor DMPSA (AN + DMPSA) in maize (roots and aboveground biomass) and soil.
Recovery from fertilizer (%) N unaccounted for
Plant Soil Total
Treatment NS NS NS NS
AN 66.2 23.2 89.4 10.6
AN + DMPSA 69.4 13.4 82.9 17.1
S.E. 2.5 2.2 3.2 3.2
The Standard Error of the mean (S.E.) is given for each effect. "NS" means no significant at P < 0.05.
N from fertilizer (kg N ha1)
0 10 20 30 40 50
■ AN OAN+DMPSA
Fig. 7. Amount of residual N in soil coming from ammonium nitrate (AN) or AN with the nitrification inhibitor DMPSA (AN + DMPSA) in the different soil layers. The horizontal lines indicate standard errors and * mean significant at P < 0.05.
These processes could have been: i) nitrification (predominant under aerobic conditions and low C-content soils) (Firestone and Davidson, 1989); ii) coupled nitrification-denitrification, which can occur if favorable conditions for both processes occur in soil neighboring microhabitats, e.g. varying soil moisture conditions (irrigated areas with drying-rewetting cycles) (Guardia et al., 2017b); or iii) nitrifier deni-trification, which could be relevant under soil organic C concentrations and moisture conditions which are sub-optimal for denitrification (Kool et al., 2011). Even though we could not discriminate between all these potential processes, all of them involve the oxidation of NH4 + to hy-droxylamine (first step of nitrification), so the inhibition of the ammonium monooxygenase is expected to mitigate the emissions coming from all of these biochemical processes (i.e. from exogenous 15NH4 + ), as we actually observed (Fig. 4). We speculate that these results could be a consequence of the inhibition of growth of NH3-oxidizing bacteria and archaea, as observed with other pyrazole-based inhibitors (e.g. DMPP) by other authors (Di and Cameron, 2011; Shi et al., 2016; Duncan et al., 2017; Barrena et al., 2017). The effectiveness on the inhibition of NH4+ oxidation was also supported by average soil mineral contents, since the DMPSA increased NH4 + and consequently decreased NO3- concentrations (Figs. 2a,b and 3).
An interesting result was that in spite of the fact that N2O losses from A15N were low, these were also significantly reduced in the AN + DMPSA treatment, compared to AN. Our results suggest that in addition to the indirect effect of nitrification inhibitors on denitrifica-tion (due to the delay in the nitrification and the reduced availability of the substrate for denitrification) (Ruser and Schulz, 2015), these compounds could also have a direct effect on denitrifiers. Previous incubation studies based on the acetylene (C2H2) blockage method (Vallejo et al., 2001) or gas chromatography (using a He ionization detector to analyze N2, Hatch et al., 2005) did not find any significant effects of nitrification inhibitors (e.g. DCD or DMPP) on direct losses from denitrification. In contrast, our approach involves the use of stable isotopes (15N) at field conditions, representing a more accurate and
representative way to identify the effect of nitrification inhibitors on the N2O release from the heterotrophic reduction of NO3 -. Dong et al. (2013) or Kou et al. (2015) found that DMPP reduced the copy numbers of genes encoding the enzymes that catalyse the reduction of soluble nitrite (NO2-) to gaseous nitric oxide (NO): nirK and/or nirS. Moreover, Barrena et al. (2017) also found that DMPP decreased the NO3- reductase (narG) and N2O reductase (nosZ) gene abundance. However, a NH4 + -N based fertilizer (urea) was employed in these experiments, so these authors attributed their results to a restricted denitrification due to the lower NO3- availability.
Previous findings could help to explain this effect of DMPSA on denitrifying microorganisms. For instance, Maienza et al. (2014) found that DMPP caused a depletion of fungal growth. Since fungal denitrifier communities generally lack the gene encoding N2O reductase, they are associated with an increment of the N2O/N2 ratio (Mothapo et al., 2015). Consequently, a negative effect of pyrazole-based compounds on fungal community could be associated to a reduction of N2O losses coming from denitrification. In addition, if pyrazole compounds act as metal chelators (Ruser and Schulz, 2015), we hypothesized that the nitrification inhibitors could have affected the reduction of NO2- to NO, since this step can be catalyzed by two different nitrite reductases: NirS (which contains a cytochrome cd1 active site) and NirK (a multi-Cu oxidase metalloprotein family) (Jones et al., 2008; Glass and Orphan, 2012). Moreover, Garbeva et al. (2007) reported that ammonium oxidizing bacteria contain the nirK gene, so the effect of DMPP on this community of nitrifiers (Ruser and Schulz, 2015) could decrease the abundance of nirK copy numbers. A possible effect of the inhibitor in the stepwise reduction of NO3- to N2 could be noticed through the measurement of N2 fluxes. In our case, a reduction of N2 fluxes was detected, but only at 3 and 4 days after fertilization (Fig. 5), coinciding with the N2O peak but with lower N2 fluxes. Further research involving changes in microbial function is required to confirm our findings and understand the possible influence of pyrazole-based nitrification inhibitors on denitrifying communities and the final products of deni-trification. With regards to endogenous soil N, a tendency (non-significant) to reduce the amount of N2O losses coming from this source was found. The N2O losses from endogenous soil N may have come mostly from the reduction of residual NO3-, since the NH4 + concentration before N addition was low.
In addition to the effectiveness of DMPSA, our results also showed that the N source must also be considered as a mitigation strategy. In the AN treatment, the amount of N2O losses coming from NO3- was much lower than that from NH4 + , particularly at the emission peak (Fig. 4). This suggests that the choice of N fertilizers containing NO3--N can reduce N2O emissions in irrigated crops (Abalos et al., 2014b; Zhang et al., 2016; Guardia et al., 2017b). Even though the water availability is not a limiting factor for denitrification in irrigated semiarid areas (Aguilera et al., 2013), the high soil temperatures during summer lead to continuous drying-rewetting cycles, favoring coupled nitrification-denitrification (Guardia et al., 2017b). The positive correlation of N2O fluxes with WFPS (see section 3.2) supports this hypothesis. The importance of this biochemical process is supported by the lack of significant differences between the amount of 15N2 coming from A15N or AN, suggesting an intense nitrifying activity followed by the reduction of NO3 -. The quantitatively higher N2 fluxes (compared to that of N2O, see section 3.4), could also suggest that denitrification rates may not be marginal. The WFPS conditions in the micropores after irrigation events may have promoted the reduction of N2O to N2 (Friedl et al., 2016), thus decreasing N2O losses from A15N. When compared with other climatic areas, daily N2 fluxes in this experiment (50-150 mg m-2 d-1) were, as expected, lower than those in the experiment of Buchen et al. (2016) (reaching 911.5 mg m-2 d-1) in a grassland site with favorable conditions for denitrification. Therefore, the recommended use of NO3- -N synthetic sources under conditions similar to those of our experimental area should not be extrapolated to all type of soils and climates. In cropping areas with a predominance of
denitrification (e.g. temperate humid grasslands), urea (an NH4 + -N based fertilizer) resulted in lower N2O losses than AN (Roche et al., 2016). In addition, some side effects (e.g. reduced N immobilization in soil, increased N leaching potential in well-drained soils) should also be considered when choosing N fertilizers containing NO3- -N, instead of NH4+ -N based ones, to optimize crop N use efficiency (Fageria and Baligar, 2005; Quemada et al., 2013). Best mitigation practices should be adopted taking into account the ecological and management conditions, as well as side effects on crop yields and farm costs (which are influenced by the change of N source, including the use of nitrification inhibitors), in order to find sustainable strategies which are potentially suitable for farmers.
4.2. Effect of DMPSA on CH4 and CO2 fluxes
DMPSA decreased daily respiration rates at all sampling dates (data not shown) and cumulative fluxes at the end of the maize cropping cycle (P < 0.05). A depletion of respiration fluxes could be a result of lower respiration of soil microorganisms or lower root respiration (which is related to lower root biomass). The latest hypothesis seems less plausible, since an increment of root biomass was observed in the plots which received DMPSA. Some previous studies (Weiske et al., 2001; Pfab et al., 2012; Maienza et al., 2014; Florio et al., 2016) have found a reduction of soil respiration after DMPP addition. Since nitrifying and denitrifying communities are only a small percentage of soil microbiota, this result could suggest that pyrazole compounds as DMPSA could have affected the activity of non-targeted microorganisms in the short-term, and this effect should be investigated in further experiments.
An effect of mineral N on methanotrophic activity due to the competitive inhibition of CH4 monooxygenase with the AMO mono-oxygenase has been previously suggested (Dunfield and Knowles, 1995). Therefore, changes in topsoil NH4 + and NO3- availability (Figs. 2a,b and 3) due to the presence of nitrification inhibitors (e.g. DMPSA) could affect methanotrophy rates. The study of Aronson and Helliker (2010) indicated that small N additions tend to stimulate CH4 oxidation, whereas large additions (> 100 kg N ha-1 yr-1) are inhibitory. In agreement, we observed a negative correlation between CH4 oxidation and NH4 + concentrations in our experiment, where 180 kg N ha-1 was added. The same amount of N was added to both AN and AN + DMPSA microplots, and the temporary differences in mineral N contents (Figs. 2a,b and 3) between these treatments did not cause a significant change in CH4 uptake. In agreement with our results, no significant effect of DMPSA on CH4 emissions was found by Huérfano et al. (2016), while Weiske et al. (2001), Pereira et al. (2010), Menéndez et al. (2012) or Rime and Niklaus (2017) did not find any effect of DMPP on cumulative CH4 oxidation.
4.3. 15N recovery in plant and soil
The average crop recovery of the synthetic fertilizer (67.8%) was higher than that reported by Rimski-Korsakov et al. (2012) (56%) or Migliorati et al. (2014) (51%). From an agronomic point of view, the N recovery of fertilizers is an important issue affecting NUE, so further research is recommended under these climatic conditions using different N sources and/or N rates under different cropping systems. With regards to the recovery of fertilizer in the residual soil N, the values obtained in our study (13-23%) were in the same range as those obtained by Rimski-Korsakov et al. (2012) (6-11% in residual NO3-, 8-21% in organic N). The proportion of N fertilizer which remained non-accounted for in our study ranged from 10 to 17%. This non-accounted for N could have been lost through NH3 volatilization, NO3-leaching and complete denitrification to N2 (Sanz-Cobena et al., 2012; Pan et al., 2016).
5. Conclusions
Our results confirmed that the new inhibitor DMPSA was an effective strategy to mitigate total N2O emissions, particularly those coming from NH4+ oxidation, but also those coming from NO3— reduction. When the fertilizer was applied without DMPSA, the majority of N2O fluxes in the emission peaks were generated from 15NH4+ (nitrification and/or denitrification and/or coupled nitrification denitrification). The latest process may have been an important contributor to N2O emissions, as shown by the similar 15N2 fluxes coming from 15NH4+ and 15NO3 —. The predominance of these biochemical processes suggests that the change of NH4+ -N based forms to NO3— -N ones can also be considered as a promising N2O mitigation practice in Mediterranean areas. When the inhibitor was added, the endogenous soil N was the main source of N2O losses, and those also tended to decrease due to DMPSA addition. The effect of DMPSA on non-target microorganisms (as suggested by the effect of the inhibitor on respiration fluxes and N2O losses from denitrification) should be investigated in further experiments. The nitrification inhibitor did not cause any improvement of either dry weight of maize grain, aboveground biomass or of plant N recovery. Conversely, it was confirmed as a highly effective tool to reduce N2O emissions in the days following N application in irrigated crops of semi-arid areas.
Acknowledgements
The authors are grateful to the Spanish Ministry of Economy and Innovation (AGL2015-64582-C3-3-R), the Autonomous Community of Madrid (P2013/ABI-2717), and the FPI grant (Ref. BES-2013-063749, AGL2012-37815-C05-01) for their economic support. Funding for this research was also provided by EuroChem Agro GmbH. We offer a special thanks to the field assistants working with us at the "El Encin" field station (IMIDRA), the technicians (Gemma Andreu, Ana Ros, Paloma Martín, Estrella Revenga, Laura Rubio and Alexandra Muro) at the Department of Chemistry and Food Technology of the ETSIAAB, as well as to Ramón Redondo, Vanessa Peiro (Sidi UAM) and Miguel Repullo (Rothamsted) for the measurements of 15N. This work was done in the frame of the Moncloa Campus of International Excellence (UCM-UPM).
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