Overdose fertilization induced ammonia-oxidizing

Science of the Total Environment 650 (2019) 1787–1794

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Science of the Total Environment journal homepage: www.elsevier.com/locate/scitotenv

Overdose fertilization induced ammonia-oxidizing archaea producing nitrous oxide in intensive vegetable fields Pengpeng Duan a, Changhua Fan b, Qianqian Zhang a, Zhengqin Xiong a,⁎ a b

Jiangsu Key Laboratory of Low Carbon Agriculture and GHGs Mitigation, College of Resources and Environmental Sciences, Nanjing Agricultural University, Nanjing 210095, China Environment and Plant Protection Institute, Chinese Academy of Tropical Agricultural Sciences, Hainan 571737, China

H I G H L I G H T S

G R A P H I C A L

A B S T R A C T

• Soil pH induced by N gradients controlled the abundance/contribution of AOA and AOB. • AOA-derived N2O were greater than AOB in the control and overdose fertilized soils. • AOB-derived N2O dominated in conventionally fertilized vegetable soils.

a r t i c l e

i n f o

Article history: Received 13 August 2018 Received in revised form 24 September 2018 Accepted 26 September 2018 Available online 27 September 2018 Editor: Jay Gan Keywords: Soil acidification Ammonia-oxidizing archaea (AOA) Ammonia-oxidizing bacteria (AOB) Nitrous oxide Nitrite

⁎ Corresponding author. E-mail address: [email protected] (Z. Xiong).

https://doi.org/10.1016/j.scitotenv.2018.09.341 0048-9697/© 2018 Elsevier B.V. All rights reserved.

a b s t r a c t Little is known about the effects of nitrogen (N) fertilization rates on ammonia-oxidizing archaea (AOA) and ammonia-oxidizing bacteria (AOB) and their differential contribution to nitrous oxide (N2O) production, particularly in greenhouse based high N input vegetable soils. Six N treatments (N1, N2, N3, N4, N5 and N6 representing 0, 293, 587, 880, 1173 and 1760 kg N ha−1 yr−1, respectively) were continuously managed for three years in a typically intensified vegetable field in China. The aerobic incubation experiment involving these field-treated soils was designed to evaluate the relative contributions of AOA and AOB to N2O production by using acetylene or 1-octyne as inhibitors. The results showed that the soil pH and net nitrification rate gradually declined with increasing the fertilizer N application rates. The AOA were responsible for 44–71% of the N2O production with negligible N2O from AOB in urea unamended control soils. With urea amendment, the AOA were responsible for 48–53% of the N2O production in the excessively fertilized soils, namely the N5–N6 soils, while the AOB were responsible for 42–55% in the conventionally fertilized soils, namely the N1–N4 soils. Results indicated that overdose fertilization induced higher AOA-dependent N2O production than AOB, whereas urea supply led to higher AOB-dependent N2O production than AOA in conventionally fertilized soils. Additionally, a positive relationship existed between N2O production and NO2− accumulation during the incubation. Further mechanisms for NO2−-dependent N2O production in intensive vegetable soils therefore deserve urgent attention. © 2018 Elsevier B.V. All rights reserved.

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P. Duan et al. / Science of the Total Environment 650 (2019) 1787–1794

1. Introduction China accounts for about 52% of the world's vegetable production. The area devoted to vegetable cultivation increased from 3.3 × 106 ha in 1976 to 24.7 × 106 ha in 2012 (FAO, 2015). It has become a common phenomenon for excessive fertilizer application to produce higher yield in greenhouse vegetable field. According to the review by Hu et al. (2017), fertilization in greenhouse vegetable areas were 2–5 times higher than those in open fields. The nitrogen (N) amount of urea could be up to 510–948 kg N ha−1 two crops yr−1 in the greenhouse vegetable system in Nanjing, far beyond the nutrient demands for cropping (Yang et al., 2016; Zhong et al., 2016), which has greatly contributed to serious environmental consequences such as soil acidification (Guo et al., 2010) and nitrous oxide (N2O) emissions (De Rosa et al., 2016; Li et al., 2016; Fan et al., 2017). The direct N2O emissions from intensive greenhouse vegetable fields are believed to contribute approximately 20% of the total N2O emissions from Chinese cropland (Wang et al., 2011) and the main source of global warming potentials judged by the life cycle assessment (Jia et al., 2012). To predict the consequences of this input, there is a pressing need to understand the basic mechanisms that underlie microbial N transformations (Kuypers et al., 2018). Various processes are associated with N2O production in soils following urea or organic N addition with or without enzyme catalysis, i.e. nitrification (Beeckman et al., 2018), nitrifier denitrification (Stieglmeier et al., 2014), codenitrification (Spott et al., 2011), chemodenitrification (Homyak et al., 2017) and heterotrophic denitrification (Hallin et al., 2018). The ammonia (NH3) oxidation pathway converts NH3 to nitrite (NO2−) as the first and rate-limiting step in nitrification and is therefore the main contributor to the ammonium (NH4+):nitrate (NO3−) balance in soil (Beeckman et al., 2018). There are three distinct groups of aerobic autotrophic microorganisms that oxidize ammonia: NH3 oxidizing bacteria (AOB) (Prosser, 1990), NH3 oxidizing archaea (AOA) (Konneke et al., 2005) and the newly discovered complete ammonia oxidation “comammox” bacteria (Daims et al., 2015; van Kessel et al., 2015; Kits et al., 2017). However, their relative contribution is difficult to estimate, as different environmental factors can affect their abundance and activity (Prosser and Nicol, 2012). For example, although the relative importance of AOA might be low in N-rich soils, their nitrification rate is at maximum and, therefore, should not be ignored (Beeckman et al., 2018; Li et al., 2018), especially in acidic soils, where the NH3 concentration is low, high substrate affinity might give them a competitive advantage (Prosser and Nicol, 2012). Recently, Kits et al. (2017) have indicated that oligotrophic AOB, many AOA and comammox bacteria are adapted to low ammonium concentrations and are inhibited by higher concentrations, but comammox bacteria has a higher ammonia affinity than all cultured terrestrial ammonia-oxidizing archaea. There remains a paucity of evidence regarding how the amendment of soil with different N fertilization rates influences their relative importance in NH3 oxidation. The niche differentiation of AOA, AOB and comammox bacteria associated with NH4+ supply has the potential to influence, significantly, N2O emissions due to their apparently distinct physiological processes (Beeckman et al., 2018; Lehtovirta-Morley, 2018). Hink et al. (2017, 2018) reported that the N2O production was lower following ammonia oxidation by AOA and the N2O production of AOB and the N2O/NO2− product ratio of nitrification increased by increasing ammonium concentration in agricultural soils. Traditionally, the AOB produce N 2O enzymatically by two mechanisms: incomplete oxidation of hydroxylamine (NH 2 OH) to NO2− (Arp and Stein, 2003), and via nitrifier denitrification, the sequential reduction of NO2− to nitric oxide (NO) and N2O (Kozlowski et al., 2014). However, recent studies revealed two other routes for the N2 O production from the AOB N. europaea under anaerobic conditions: the direct oxidation of NH2OH to N2O by the enzyme cytochrome (cyt) P460 and nitrification intermediate NO (Caranto et al., 2016; Caranto and

Lancaster, 2017). Moreover, the mechanisms of N2O production by AOA appear to differ from that of AOB, as AOA lack genes encoding a canonical hydroxylamine dehydrogenase and NO reductase, which are involved in N2O production by AOB (Walker et al., 2010; Tourna et al., 2011). It is thought that AOA produce hybrid N2O (with one N sourced from ammonium and the other from nitrite) during NH3 oxidation through an abiotic reaction between NH2OH and NO, an intermediate of the AOA ammonia oxidation pathway (Stieglmeier et al., 2014; Lehtovirta-Morley, 2018), which has been demonstrated for pure cultures (Kozlowski et al., 2016). Given that nitrite and nitrous acid are in a pH-dependent equilibrium, AOA strains grown in acidic pH media produce proportionally much more N2O than in neutral pH, although it is unclear whether this is due to effects of pH or strain physiology (Jung et al., 2014). This indicates that acidic soil has a higher potential to cause AOAdependent N2O production. However, the mechanisms of N2O production by AOA under oxic conditions remain unclear. Hence, considering the soil pH as an important factor and the different mechanisms of AOA and AOB in N2O production can provide more insights for the associated kinetic properties influencing N2O pathways in soils with different N fertilization rates. As described by Taylor et al. (2013), 1-octyne is a newly developed inhibitor, which specifically inhibits AOB growth and activity while acetylene can effectively inhibit both AOA and AOB at low concentrations. Therefore, 1-octyne and acetylene are adopted to assess the relative contribution of AOA and AOB to N2O production from six levels of N fertilization treated vegetable soils after 3-years. We hypothesize that the AOA would be responsible for the majority of N2O production, given the acidic conditions which induced by intensive and overdose fertilization.

2. Materials and methods 2.1. Site description, field treatment and soil collection The field experiment was conducted in a typically managed vegetable agroecosystem in a suburban area of Nanjing, Jiangsu Province, China (32°01 N, 118°52 E) from April 30, 2013 to May 10, 2016 covering eleven crops. Vegetables have been cultivated continuously at this site for more than ten years. The region has a subtropical monsoon climate with an annual mean precipitation of 1107 mm and a mean air temperature of 15.4 °C. The soil is classified as a FimiOrthic Anthrosol (RGCST 2001) consisting of 64.7% silt (0.002–0.05 mm), 30.1% clay (b0.002 mm) and 5.2% sand (N0.05 mm). The soil bulk density is 1.22 g cm−3 with an initial soil pH of 5.1. The soil has a total carbon content of 19.2 g kg−1 soildw, a total N content of 1.4 g kg−1 soildw, and a cation exchange capacity of 31.2 cmol kg−1. The six field fertilizer treatments (each replicated three times) included 0 kg N ha −1 yr−1 (N1), 293 kg N ha −1 yr−1 (N2; 1/3 of the conventional N rate), 587 kg N ha−1 yr−1 (N3; 2/3 of the conventional N rate), 880 kg N ha−1 yr−1 (N4; the conventional N rate), 1173 kg N ha−1 yr−1 (N5; 4/3 of the conventional N rate) and 1760 kg N ha −1 yr−1 (N6; 6/3 of the conventional N rate). These urea rates were selected to cover a range of N fertilizer application practices across different vegetable production patterns. Phosphate (P) and potassium (K) fertilizers as calcium/magnesium phosphate and potassium chloride, respectively, were applied at a rate of 880 kg P2 O 5 ha −1 yr−1 and 880 kg K2 O ha −1 yr−1 . These basal fertilizers were evenly distributed to each of the eleven crops and incorporated into the soil. Soil samples from each replicated plot at a depth of 0–20 cm in November 2016, were divided into three parts: the first part was stored at −80 °C for DNA extraction, the second part was stored at 4 °C prior to determination of exchangeable NH4+, NO3− and NO2− and other chemical properties, the third part was used for the incubation.


2.2. Incubation experiment We assume that the initial carbon source derived from the crop rhizosphere and debris, and the residual fertilizer N from the previous field fertilization application, and the in situ soil temperature would impact the incubation results to a large extent. Thus, a 28-day pre-incubation was established to ensure the uniform condition and true treatment effects following the procedures. The fresh soil samples were passed through a 2 mm screen and visible plant residues were removed by hand. Moisture was adjusted to around 50% of water filled pore space (WFPS) and incubated at 25 °C for 28-day. The WFPS is obtained using the following equation: WFPS ¼

volumetric water content total soil porosity

ð1Þ

Here, total soil porosity is equal to [1-(soil bulk density (g cm−3)/2.65)], with an assumed soil particle density of 2.65 (g cm−3). The total soil bulk density was determined by using the cutting ring method from 0 to 10 cm depth according to Lu (2000). Then the soil properties were measured once again using the abovementioned methods and the aerobic incubation experiment was carried out as follows: In accordance with the procedure described by Taylor et al. (2017), urea was added to the six N-fertilized soils with acetylene (0.01% v/v) or 1-octyne (0.03% v/v) as treatments. 100 mg N-urea kg−1 soildw in incubation is representative of field broadcasting application rate and would also result in low increase of NH4+ concentration in a preliminary soil microcosm experiment over 27 days. Therefore, soil microcosms each containing 15 g of dry weight soil in 120-ml serum bottle, were adjusted to 60% WFPS with urea solution (100 mg N-urea kg−1 soildw) or without (control) in the absence or presence of acetylene or 1-octyne and incubated at 28 °C for 27 days. Three microcosms for each treatment were destructively sampled after incubation at 0, 7, 13, 21 and 27 days and stored at 4 °C and −80 °C for chemical and DNA analyses, respectively. A total of 540 serum bottles (6 field soils × 3 inhibitors × 2 additives × 3 replicates × 5 samplings) were used. The serum bottles were all tightly capped with butyl rubber stoppers and metal crimp tops. Oxic conditions were maintained by opening the bottles for 20 min at intervals of 2–3 days when both water and acetylene or 1octyne were adjusted to maintain the designed conditions. Before each gas sampling, the serum bottles were opened for 20 min to refresh the atmosphere and corresponding inhibitors and then sealed to realize complete inhibition for 8 h. Afterwards, gas samples (10 ml) were collected from the headspace with syringe at 0, 1, 3, 5, 7, 13, 21, and 27 days after the start of incubation. Net N nitrification rate was also calculated as the changes in NO2−-N + NO3−-N concentrations within 7 days of incubation and expressed as mg N kg−1 soil d−1. 2.3. Analyses of soil chemical properties and N2O production The soil pH was measured in deionized water at a soil-to-water volume ratio of 1:5 with a PHS-3C mv/pH detector (Shanghai Kangyi Inc. China). The electric conductivity (EC) was measured at a soil-to-water volume ratio of 1:5 using a Mettler-Toledo instrument (FE30-K, Shanghai, China). Using an ultraviolet spectrophotometer (HITACHI, UV-2900, Tokyo, Japan), soil extracts with 2 M KCl (Maynard et al., 1993) were measured by following two-wavelength ultraviolet spectrometry for NO3−-N concentration, and indophenol blue methods for exchangeable NH4+-N concentration. Nitrite was extracted with deionized water to avoid underestimation by 2 M KCl (Homyak et al., 2015) and the NO2−-N concentration of the soil extract measured by N-(1-naphthyl) ethylenediamine dihydrochloride, using an ultraviolet spectrophotometer (HITACHI, UV-2900, Tokyo, Japan) (Duan et al., 2018). Soil NH2OH concentrations were determined following the procedure described by Liu et al. (2014). The N2O concentration in the headspace was analyzed

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by chromatography (Agilent 7890A, Agilent Ltd., Shanghai, China) equipped with an electron capture detector (ECD) and quantified by comparing the peak areas with the reference gases (Nanjing Special Gas Factory, Nanjing, China). The carrier gas was argon-methane (5%) at a flow rate of 40 ml min−1. The N2O production rate was calculated by the linear increase of headspace N2O concentration and cumulative N2O production was calculated from the individual rate and the weighted incubation time. 2.4. DNA extraction and quantitative PCR (qPCR) analyses The subsamples collected at 0, 7, 13, 21 and 27 days were used to determine the amoA abundance of AOA and AOB populations (Table S1). Details about the soil DNA extraction and qPCR analyses are provided in the Supplementary section. Since N2O emissions at 13-d were higher, the results of qPCR at 13-d were selected for analyses. 2.5. Calculations and statistical analyses Since acetylene can completely inhibit ammonia oxidation by both AOA and AOB while 1-octyne can specifically inhibit AOB, the total N2O production from AOA + AOB is calculated by subtracting the N2O production from the urea + acetylene treatment from that in the urea treatment. The N2O production from AOA is determined by subtracting the N2O production in the urea + acetylene treatment from that in the urea + 1-octyne treatment. The N2O production from AOB is then calculated as the difference between the N2O production from AOA + AOB and from AOA. The relative contributions of N2O production by AOA and AOB are deduced from their corresponding production and the total production. NH3 rather than NH4+ is thought to be the substrate oxidized by AOA and AOB. NH3 concentrations depend not only on the amount of NH4+ in a given environment but also on the pH of that environment (Norman and Barrett, 2014). Assuming the pKa for NH3/NH4+ of 9.25 based on the Henderson-Hasselbalch equation, the amount of NH3 available is estimated as follows: ðpH−9:25Þ NH 3 ¼ NH þ 4 10

ð2Þ

The N2O production (‰) is expressed as the N2O-N per NO2−-N + NO3−-N produced by treatments where ammonia oxidation was not inhibited by acetylene as follows: N2 Oproduction ¼

N2 O−N 1000 − NO− 2 −N þ NO3 −N

ð3Þ

The statistical analyses are performed using SPSS 20.0 software (IBM Co., Armonk, NY, USA). An ANOVA is performed to analyze the differences in N2O production, amoA gene abundance and exchangeable NH4+, NO2− and NO3− concentrations. Pearson's correlation procedure is used to evaluate the relationship among the edaphic factors and the abundances of AOB and AOA. The effects of substrate and treatment on the amoA genes, mean N2O production, and net nitrification rate are analyzed by one-way ANOVA and least significant difference test at a 0.95 confidence level. The linear fit for N2O production rate and the NO2− concentration is analyzed using OriginPro 8.1 (OriginLab., USA). 3. Results 3.1. Soil properties and correlation analyses with measured variables N fertilization rates showed significant effects on soil EC, pH and AOA-, AOB-amoA abundances (p b 0.05) after 3-year application of urea (Table S2). Soil pH was significantly higher under N1 and N2 than under N3, N4, N5 and N6 soils before and after the pre-incubation (p b

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increased gradually (Fig. 2), indicating that mineralization of native organic N from vegetable soils was lower than nitrification. The NO3− did not accumulate in +acetylene treatments, indicating that ammonia oxidation was completely inhibited (Fig. 2). When 1-octyne was present, both exchangeable NH4+ and NO3− concentrations remained similar in the absence of 1-octyne, indicating that the ammonia oxidation in the control was not influenced by the 1-octyne. In the urea treatment without inhibitors, the exchangeable NH4+ concentration decreased and the NO3− concentration increased rapidly whereas NO2− accumulated at the early incubation period (1–7 d), indicating a rapid nitrification process occurred. When 1-octyne was present, the rates of exchangeable NH4+ decrease and NO3− concentration increase were between the urea and the urea + acetylene treatment, indicating a partial inhibition by 1-octyne on the NH3 oxidation, due to the inhibition of AOB (Fig. 2). There was clear difference in N2O emission in the presence or absence of 1-octyne and acetylene after 27 days of incubation (Fig. 2). N2O emission significantly increased with urea addition, and when urea was applied with 1-octyne or acetylene, cumulative N2O emission was dramatically reduced, especially in N1–N4 soils (Table S3). However, significant difference in N2O production in acetylene-treated microcosms with or without added urea in N5–N6 soils (Table S3). 3.3. The relative contribution of AOA and AOB to total N2O production

−1 Fig. 1. The net nitrification rates (mg NO− soildw) (A) in field soils after urea 3 -N kg addition and AOA-, AOB-amoA gene abundances (B) in field soils with or without urea addition at 0-d and 13-d incubation. N1, N2, N3, N4, N5 and N6 are soils treated with 0, 1/3, 2/3, 3/3, 4/3 and 6/3, respectively of the conventional N rate of 880 kg N ha−1 yr−1. Error bars represent the SE (n = 3). Asterisks (B) represent that significant higher values of amoA abundance between treatments within each soil, ⁎p b 0.05, ⁎⁎p b 0.01. Different letters above the bars denote significant difference and the same letters denote no significant difference.

0.05). In contrast, the AOA- and AOB-amoA abundances of all the fertilization treatments were not strongly affected by N application rates after pre-incubation, while overdose fertilization increased the AOA-amoA abundances (p N 0.05). Net N nitrification rate differed significantly in the following patterns: N2 = N3 N N1 = N4 N N5 N N6 (p b 0.05) (Fig. 1A), and being significantly positively correlated with the soil pH (p b 0.01) (Table 1). Following application of urea, both AOA and AOB amoA gene abundances significantly increased in N1 (p b 0.05), with the greatest increase observed for AOB (Fig. 1B). The AOA amoA abundance showed a significant negative correlation with the soil pH and NH3 concentration (p b 0.05), whereas AOB amoA abundance showed a significant positive correlation with the pH (p b 0.05) (Table 1). 3.2. Kinetics of soil exchangeable NH4+, NO3−, and NO2− and N2O fluxes as affected by inhibitors In the control treatment without urea or inhibitor addition, the exchangeable NH4+ concentration remained consistently low while NO3− Table 1 Pearson correlation analyses between net nitrification rate, amoA gene abundance from incubation and soil properties after pre-incubation.

pH EC a NH+ 4 -N NO− 3 -N NO− 2 -N NH3

Net nitrification rate

AOA-amoA

AOB-amoA

0.824⁎⁎ 0.254 −0.282 0.867⁎⁎ −0.009 0.807⁎⁎

−0.578⁎ −0.485 0.446 −0.639⁎ 0.015 −0.627⁎

0.513⁎ 0.372 −0.390 0.810⁎⁎ 0.090 0.765⁎

⁎ and ⁎⁎ indicate significant correlation at p b 0.05 and 0.01, respectively. a

Exchangeable.

The N2O produced ranged from 23.42 ± 7.84 to 78.76 ± 6.21 ng N g−1 soildw in the unamended control soil (Table S3). When 1-octyne was present for inhibiting AOB, no difference in N2O production was found between the absence and presence of the 1-octyne (p N 0.05). When acetylene was present for inhibiting both AOB and AOA, N2O production decreased from 23.42–78.76 to 12.58–27.29 ng N2O-N g−1 soildw. The AOA were responsible for ~44–71% of N2O production whereas other processes being responsible for ~28–55% of the N2O production in the unamended control soils (Fig. 3A). When urea was added to the field soils, N2O production was stimulated by 2–5 folds, ranging from 127.41 ± 25.3 to 264.00 ± 30.81 ng N g−1 soildw (Table S3). When 1-octyne was present, significant lower NO3− concentration and N2O production than without the inhibitor was found, and the N2O production was significantly reduced when acetylene was present. The AOB were responsible for ~42–56% while AOA were responsible for ~32–39% of the N2O production in the N1–N4 soils. On the contrary, the AOA were responsible for ~50–53% while AOB were responsible for ~17–24% of the N2O production in the N5–N6 soils (Fig. 3B). The remaining ~11–30% came from other processes in the N1–N6 soils. 3.4. AOA- and AOB-dependent N2O production and relationship between N2O production rate and NO2− accumulation The AOA- and AOB-dependent N2O production were significantly higher in the urea treatment than in the control treatment in all fertilized field soils (p b 0.05), except N1–N3 soils, where no significant difference in AOA-dependent N2O production between the control and urea treatment (p N 0.05). The AOA-dependent N2O production were significantly higher than AOB under urea addition in the N5 and N6 soils than other soils (p b 0.05) (Fig. 4A). In non-amended microcosms, where ammonia oxidation and N2O production resulted mainly from activity of AOA, the N2O production was ~0.06–0.28‰. In ureaamended, non-inhibited microcosms, where both AOA and AOB were active, the AOA-dependent N2O production was ~0.06–0.55‰, and the AOB-dependent N2O production was ~0.18–0.37‰ (Fig. 4A). NO2− was accumulated during the whole incubation period in the N1–N6 soils (Fig. 2). The N2O production rate was significantly positively correlated with NO2− accumulation across N1–N6 soils (R2 = 0.69, p b 0.01) (Fig. 4B).


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− − Fig. 2. The kinetics of the exchangeable NH+ 4 -N, NO3 -N, NO2 -N and N2O from field soils with or without urea addition and incubated for 27 days in combination with or without inhibitors (acetylene or 1-octyne). N1, N2, N3, N4, N5 and N6 are soils treated with 0, 1/3, 2/3, 3/3, 4/3 and 6/3, respectively of the conventional N rate of 880 kg N ha−1 yr−1. The data are the means with the SE (n = 3).

4. Discussion 4.1. Relative contributions of AOA and AOB to the total N2O production Although we primarily focus on acetylene-sensitive AOA and AOB activities, the non-NH3 oxidizers N2O production occurred from control (~25–53% of the N2O production, Fig. 3A) and amended N1–N6 soils (~11–30%), through other processes probably denitrification and abiotic processes. N2O accumulated slowly in acetylene-treated microcosms,

Fig. 3. The relative contributions of AOA and AOB to the total N2O production from field soils with or without urea addition and incubated for 27 days in combination with or without inhibitor acetylene or 1-octyne. N1, N2, N3, N4, N5 and N6 are soils treated with 0, 1/3, 2/3, 3/3, 4/3 and 6/3, respectively of the conventional N rate of 880 kg N ha−1 yr−1.

where NH3 oxidizers growth and activity were inhibited (Fig. 2, Table S3). A decrease in NO3− concentration, and accompanied by increasing exchangeable NH4+ concentration after 21 days of incubation was observed in acetylene treatment (Fig. 2), indicating a possible increase in denitrification activity and/or dissimilatory nitrate reduction to ammonium (DNRA) and/or immobilization. High rates of nitrification might cause oxygen-limiting micro-hot-spots and the accumulated NO2−/NO3− may induce the occurrence of heterotrophic denitrification in acidic soils although the current incubation was set to be oxic at WFPS of 60%. Qin et al. (2017) reported that excessive N fertilization potentially increased N2O emissions by reducing N2O reductase activity in soils. Qu et al. (2014) observed that soil pH was the master variable controlling the percentage of denitrified N emitted as N2O under high fertilizer N levels conditions. The process of DNRA might influence N2O flux, as nitrate-ammonifying and nitrate-denitrifying microorganisms compete for NO3− in soils (Minick et al., 2016). Abiotic N2O production served as another possible process particularly in N5 and N6 soils, where the intermediate NO2− was significantly accumulated (Fig. 2). Homyak et al. (2017) reported that acidic conditions favored chemodenitrification, when biological processes become stressed. In line with previous studies (Wang et al., 2016; Hink et al., 2017; Hink et al., 2018), NH4+ strongly stimulated AOB-driven N2O production rather than AOA in N1–N6 soils (Fig. 3B). The AOB growth are favored by high-NH4+-N concentration in soil (Jia and Conrad, 2009; Giguere et al., 2015), whereas the AOA growth rates are not affected (Di et al., 2009) or suppressed (Norman and Barrett, 2014; Wang et al., 2017). Here, in the conventionally N fertilized field soils (N1–N4), the stimulation of AOB-amoA by NH4+ occurred rather than AOA-amoA (p b 0.05,

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Fig. 4. The AOA, AOB-dependent N2O production (A) and correlation between N2O and NO− 2 -N accumulation (B) from field soils treated with or without urea. The N2O production is − averaged and expressed as the N2O-N per (NO− 3 -N + NO2 -N) produced for treatments without acetylene. N1, N2, N3, N4, N5 and N6 are soils treated with 0, 1/3, 2/3, 3/3, 4/3 and 6/3, respectively of the conventional N rate of 880 kg N ha−1 yr−1. Error bars represent the SE (n = 3). Asterisks (A) represent that significant higher values of N2O production between treatments within each soil, ⁎p b 0.05, ⁎⁎p b 0.01. Asterisks (B) represent regression significance ⁎⁎p b 0.01. Confidence and prediction intervals at 95% confidence level are indicated by shaded bands and dotted lines, respectively.

Fig. 1B), supporting that AOB play a critical role in N2O production in the conventionally N fertilized field soils (Fig. 3B). However, the N2O production by AOA was higher than that from AOB (Fig. 3B) when supplemented with excessively urea to the N fertilized N5 and N6 soils. N in the form of ammonium or urea inputs into soils are usually accompanied by an increase in the N availability and a decrease in the soil pH, which cause feedbacks on the abundance and activities of AOB and AOA as already mentioned (Ying et al., 2017). A significantly positive correlation was observed for net nitrification rate with soil pH (p b 0.01, Table 1), which is consistent with previous findings that potential ammonia oxidation decreased significantly with decreasing soil pH in the greenhouse vegetable land (Zhong et al., 2016). Higher application rates of N fertilization stressed environmental conditions, such as soil secondary salinization and acidification (Guo et al., 2010). Previous results showed that the pH was the predominant factor affecting nitrification (Sahrawat, 2008; Yao et al., 2013), since the pH influences substrate availability, niche differentiation and the abundances of AOA and AOB (Hallin et al., 2009; Liu et al., 2017a; Liu et al., 2018). However, the low soil pH caused by N application rates was more beneficial to the AOA than to the AOB (p b 0.01, Table 1), which is supported by reports of Lehtovirta-Morley (2018) that AOA would still grow when the amount of available NH3 is extremely low in acidic conditions. These results suggest that AOA might play a more important role than AOB in nitrification and N2O production under acidic condition which induced by overdose N fertilization. Thus, the dominant contribution of AOA to N2O production in N5–N6 soils was probably due to low pH and accompanying low soil solution NH3 concentration. Lu et al. (2012) also reported that AOA dominated nitrification in acidic soils. The isolation of ‘Candidatus Nitrosocosmicus franklandus’, a novel ureolytic soil archaeal ammonia oxidizer tolerating high ammonia concentrations by Lehtovirta-Morley et al. (2016), suggested the major contribution to nitrification in fertilized soils. In line with previous studies, the N2O production by AOA was dominant while negligible by AOB without exogenous N amendment (Fig. 3A). In such soils, AOA activity may be tightly linked to organic N mineralization (Strauss et al., 2014; Carey et al., 2016; Hink et al., 2017). It is worth noting that the AOA and AOB contribution to N2O production may have been overestimated when the incubations become long (e.g., 27 days during incubation) though 1-octyne was effective in distinguishing N2O production from AOA and AOB. Changes in the activity and competitive ability of AOA may have been resulted from changes in community composition, potentially selecting, for example, for NH3tolerant phylotypes such as N. franklandus (Lehtovirta-Morley et al., 2016). Hink et al. (2017) reported that the AOA amoA transcript abundances increased in the NH4+ + 1-octyne treatment at 20 day, which

is likely due to a selection for AOA population that could only grow in the absence of competition with AOB. We also observed that AOA grew better by 1-octyne inhibition when competition with AOB was removed. As such, the AOA contribution to N2O production may have been overestimated in the absence of any inhibitor. For example, compared to soil AOA a higher substrate affinity of the comammox bacterium N. inopinata was reported at low ammonia concentrations (Kits et al., 2017) and comammox sequences even represented about 25% of the ammonia oxidizers in rice paddy soils (Pjevac et al., 2017). 4.2. Potential N2O production processes involved in ammonia oxidation In the conventionally fertilized N1–N4 soils, N2O production was dominated by AOB while AOA-dependent N2O production dominated in the control and excessively fertilized N5–N6 soils (Fig. 4A). Typically, AOB-driven N2O production involved enzymatic mechanisms of incomplete NH2OH oxidation, nitrifier denitrification (Kozlowski et al., 2014), and direct oxidation of NH2OH to N2O by the enzyme cytochrome (cyt) P460 and nitrification intermediate NO (Caranto et al., 2016; Caranto and Lancaster, 2017). The AOA N. viennensis and N. maritimus are reported to be incapable of canonical nitrifier denitrification, but produce N2O via hybrid formation (with one N sourced from ammonium and the other from nitrite) (Stieglmeier et al., 2014; Kozlowski et al., 2016), which relies more on abiotic hybrid via nucleophilic nitrosation of NH2OH (Liu et al., 2017b). Jung et al. (2014) also reported that archaea were capable of producing similar, even higher N2O production as the bacterial nitrifiers using five AOA strains from soil enrichment culture. Given the lower pH of the N5 and N6 soils in this study, the high production of acidophile AOA strains is possibly due to their high sensitivity to NO2−, which can inhibit the growth of Nitrosotalea devanaterra and strain CS at 40 μM and 100 μM, respectively (Lehtovirta-Morley et al., 2011; Jung et al., 2014). However, it still remains unclear what mechanism is used by acidophilic AOA in ammonia oxidation pathway (Beeckman et al., 2018; Lehtovirta-Morley, 2018). Moreover, we found a positive correlation between N2O production rate and NO2− accumulation (Fig. 4B). Since N2O, unlike NO2−, is not the major nitrogenous product during ammonia oxidation, the impact of the pH on the N2O production may be decoupled from its impact on the ammonia oxidation, which has been prevailing in agricultural soils (Ma et al., 2015; Venterea et al., 2015; Giguere et al., 2017; Duan et al., 2018; Giguere et al., 2018; Lu et al., 2018; Tierling and Kuhlmann, 2018;). However, AOA or AOB-dependent N2O production was also observed in soils where NO2− was not detected (Wang et al., 2016; Hink et al., 2017; Hink et al., 2018). The rate of NH3 oxidation is generally thought to limit the rate of NO2− oxidation, however, the


interrelationship between NH3 and NO2− oxidizers is more complex and can involve reciprocal feeding and mixotrophy (Daims et al., 2016). The six intensive vegetable soils used in our study displayed a range of NH3oxidizing capacities and NO2− accumulation (Fig. 2), further emphasizing the need for a better understanding why NO2− oxidizing activity is limited and the physiological ecology of soil-borne NO2− oxidizing bacteria in overdose fertilization soils. AOA and AOB have also been shown to produce N2O from NO2− (Jung et al., 2014; Stieglmeier et al., 2014; Giguere et al., 2017; Liu et al., 2017b; Duan et al., 2018). Although AOA and AOB are sensitive to NO2−, which can inhibit their growth, the NO2−-dependent N2O production might be a detoxifying mechanism to protect nitrifiers from NO2− toxicity (Jung et al., 2014). These results raise the question to what extent the N2O production reported in previous studies might have been influenced by NO2− accumulation. Furthermore, the recently discovered comammox bacterium N. inopinata can lead to NO2− accumulation during NH3 oxidation (Kits et al., 2017). Given the possibility of an inhibition effect of 1-octyne on comammox bacteria (Taylor et al., 2017), even higher comammox bacteriadependent N2O production may come from NO2− (0.6–1.4‰) (Liu et al., 2017b). The possibility cannot be excluded that NO2− accumulation was produced by comammox bacteria in addition to AOB in soil samples without 1-octyne addition. Further studies are needed to determine if and when the accumulation of NO2− is a requirement for aerobic N2O production. 5. Conclusion We present preliminary evidence that explains how soil pH induced by an N fertilizer gradient might affect AOA- or AOB-dependent N2O production differently, with AOA dominating N2O production in the control soils and the excessively fertilized soils with exogenous N amendment while AOB still dominates N2O production in the urea amended conventionally fertilized vegetable soils. N2O production rate is significantly positively correlated with NO2− accumulation. Further studies are needed to determine if and when the accumulation of NO2− is a requirement for aerobic N2O production. These findings are significant for understanding and regulating N2O production under soil acidification induced by long term intensive fertilization. Acknowledgements We greatly thank three anonymous reviewers for their valuable comments and critical evaluation on this manuscript. This work was jointly supported by the National Natural Science Foundation of China (41471192), the Special Fund for Agro-Scientific Research in the Public Interest (201503106) and the Postgraduate Research & Practice Innovation Program of Jiangsu Province, China (KYCX18_0678). Appendix A. Supplementary data Supplementary data to this article can be found online at https://doi. org/10.1016/j.scitotenv.2018.09.341. References Arp, D.J., Stein, L.Y., 2003. Metabolism of inorganic N compounds by ammonia-oxidizing bacteria. Crit. Rev. Biochem. Mol. Biol. 38, 471–495. Beeckman, F., Motte, H., Beeckman, T., 2018. Nitrification in agricultural soils: impact, actors and mitigation. Curr. Opin. Biotechnol. 50, 166–173. Caranto, J.D., Lancaster, K.M., 2017. Nitric oxide is an obligate bacterial nitrification intermediate produced by hydroxylamine oxidoreductase. Proc. Natl. Acad. Sci. U. S. A. 114, 8217–8222. Caranto, J.D., Vilbert, A.C., Lancaster, K.M., 2016. Nitrosomonas europaea cytochrome P460 is a direct link between nitrification and nitrous oxide emission. Proc. Natl. Acad. Sci. U. S. A. 113, 14704–14709. Carey, C.J., Dove, N.C., Beman, J.M., Hart, S.C., Aronson, E.L., 2016. Meta–analysis reveals ammonia-oxidizing bacteria respond more strongly to nitrogen addition than ammonia-oxidizing archaea. Soil Biol. Biochem. 99, 158–166. Daims, H., et al., 2015. Complete nitrification by Nitrospira bacteria. Nature 528, 504–509.

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