Responses of Ammonia-Oxidizing Bacteria and Archaea in Two Agricultural Soils to Nitrification Inhibitors DCD and DMPP: A Pot Experiment

Pedosphere 23(6): 729–739, 2013 ISSN 1002-0160/CN 32-1315/P c 2013 Soil Science Society of China  Published by Elsevier B.V. and Science Press

Responses of Ammonia-Oxidizing Bacteria and Archaea in Two Agricultural Soils to Nitrification Inhibitors DCD and DMPP: A Pot Experiment∗1 GONG Ping1,2 , ZHANG Li-Li1,∗2 , WU Zhi-Jie1 , CHEN Zhen-Hua1 and CHEN Li-Jun1,3 1 Institute

of Applied Ecology, Chinese Academy of Sciences, Shenyang 110016 (China) of Chinese Academy of Sciences, Beijing 100049 (China) 3 State Key Laboratory of Forest and Soil Ecology, Institute of Applied Ecology, Chinese Academy of Sciences, Shenyang 110164 (China) 2 University

(Received December 27, 2012; revised July 22, 2013)

ABSTRACT Taking two important agricultural soils with different pH, brown soil (Hap-Udic Luvisol) and cinnamon soil (Hap-Ustic Luvisol), from Northeast China, a pot culture experiment with spring maize (Zea mays L.) was conducted to study the dynamic changes in the abundance and diversity of soil ammonia-oxidizing bacteria (AOB) and ammonia-oxidizing archaea (AOA) populations during maize growth period in response to the additions of nitrification inhibitors dicyandiamide (DCD) and 3,4-dimethylpyrazole phosphate (DMPP) by the methods of real-time polymerase chain reaction (PCR) assay, PCR-denaturing gradient gel electrophoresis (DGGE), and construction of clone library targeting the amoA gene. Four treatments were established, i.e., no urea (control), urea, urea plus DCD, and urea plus DMPP. Both DCD and DMPP inhibited growth of AOB significantly, compared to applying urea alone. Soil bacterial amoA gene copies had a significant positive linear correlation with soil nitrate content, but soil archaeal amoA gene copies did not. In both soils, all AOB sequences fell within Nitrosospira or Nitrosospira-like groups, and all AOA sequences belonged to group 1.1b crenarchaea. With the application of DCD or DMPP, community composition of AOB and AOA in the two soils had less change except that the AOB community composition in Hap-Udic Luvisol changed at the last two growth stages of maize under the application of DCD. AOB rather than AOA likely dominated soil ammonia oxidation in these two agricultural soils. Key Words:

ammonium, clone library, denaturing gradient gel electrophoresis, nitrate, real-time polymerase chain reaction

Citation: Gong, P., Zhang, L. L., Wu, Z. J., Chen, Z. H. and Chen, L. J. 2013. Responses of ammonia-oxidizing bacteria and archaea in two agricultural soils to nitrification inhibitors DCD and DMPP: A pot experiment. Pedosphere. 23(6): 729–739.

INTRODUCTION Soil nitrification, one of the key processes of the N cycle, oxidizes ammonia to nitrite and then to nitrate, enhancing losses of ammonium and ammoniumproducing fertilizers applied to soil by leaching and denitrification of nitrate, and being a main source of N2 O and NO to the atmosphere, as shown by the low nitrogen use efficiency in crop production (Ram et al., 1996; Pasda et al., 2001; Zerulla et al., 2001; Macadam et al., 2003; Majumdar, 2005). The suppression of soil nitrification can be an important strategy for minimizing N losses from agricultural systems (Sahrawat and Keeney, 1985, Prasad and Power, 1995). Nitrification inhibitors offer the potential for decreasing soil nitrate losses via retarding the microbial transformation of soil ammonium to soil nitrate (Patra et al., 2006; Zaman ∗1 Supported

et al., 2009). Ammonia oxidation, as the first step in the nitrification process, is often thought to be the rate-limiting step in autotrophic nitrification (Barraclough and Puri, 1995, De Boer and Kowalchuk, 2001). Ammoniaoxidizing bacteria (AOB) are traditionally considered to be mainly responsible for ammonia oxidation, but the discovery of the greater abundance of ammoniaoxidizing archaea (AOA) than AOB in 12 soils (Leininger et al., 2006) brought a new debate on the contribution of AOA to soil nitrification. Tourna et al. (2008) reported that AOA but not AOB played a role in soil nitrification since the community structure of active AOA changed with temperature during nitrification. Other researchers suggested control of nitrification by AOA rather than AOB in two agricultural acidic soils (Gubry-Rangin et al., 2010). However,

by the National Natural Science Foundation of China (No. 41101242), the National Basic Research Program (973 Program) of China (No. 2011CB100504) and the National Key Technology R&D Program of China (Nos. 2011BAD11B04 and 2012BAD14B04). ∗2 Corresponding author. E-mail: [email protected].

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other studies showed that AOB but not AOA dominates soil nitrification (Di et al., 2009, Jia and Conrad, 2009; Xia et al., 2011). Nitrification inhibitors, dicyandiamide (DCD) and 3,4-dimethylpyrazole phosphate (DMPP), are widely used in agricultural soils (Xu et al., 2000; Cookson and Cornforth, 2002, Vallejo et al., 2005; Moir et al., 2007). In some previous laboratory experiments and pot trials with ryegrass or cauliflower, AOB was found to be significantly affected by DCD and/or DMPP, with the reduction in AOB population size and activity, while the AOA in the presence of the two nitrification inhibitors remained largely unchanged in their population abundance (Di et al., 2009, 2010; O’Callaghan et al., 2010; Di and Cameron, 2011; Kleineidam et al., 2011). However, limited information is available about the effects of DCD and DMPP on AOA and AOB in agricultural soils with spring maize, which is main crop in Northeast China. Soil pH is a crucial factor affecting the abundance and diversity of ammonia oxidizers (Nicol et al., 2008). With decreasing soil pH, the availability of soil ammonia decreased, and the activity of the isolated AOA (Nitrosopumilus maritimus) increases with the lower half-saturation constant of ammonia than cultivated AOB (Martens-Habbena et al., 2009). In addition, there is a significant correlation between the degradation rate of DCD and soil pH (Rodgers et al., 1985). Brown soil (Hap-Udic Luvisol in the FAO (Food and Agriculture Organization of the United Nations)WRB (World Reference Base) classification system), and cinnamon soil (Hap-Ustic Luvisol in the FAOWRB system) are the main agricultural soils with different pH in Northeast China. In the present study, a pot culture experiment was conducted with the two agricultural soils to investigate the dynamic changes in the abundance and diversity of soil AOB and AOA populations during the growth period of spring maize (Zea mays L.) in response to the additions of nitrification inhibitors DCD and DMPP by the methods of real-time polymerase chain reaction (PCR) assay, PCR-denaturing gradient gel electrophoresis (DGGE), and construction of clone library targeting the amoA gene. MATERIALS AND METHODS Soils and experimental design Surface soil samples (0–20 cm) of a brown soil (Hap-Udic Luvisol) and a cinnamon soil (Hap-Ustic

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Luvisol) were collected from the Experimental Station of Shenyang Agricultural University (41◦ 49 N, 123◦ 33 E) and Chaoyang City (41◦ 49 N, 122◦ 48 E) in Liaoning Province of Northeast China, respectively, with their physical and chemical properties shown in Table I. The mean maximum and minimum temperatures during the growth of maize (May–October) are 18 and 7 ◦ C. TABLE I Selected physical and chemical properties of the test soils Soil property

Hap-Udic Luvisol

Hap-Ustic Luvisol

Sanda) (%) Silta) (%) Claya) (%) pH (soil:H2 O = 1:2.5) Organic Cb) (g kg−1 ) Total Nc) (g kg−1 ) d) (mg kg−1 ) NH+ 4 -N − NO3 -Nd) (mg kg−1 ) Nitrification potentiale) −1 soil h−1 ) (mg NO− 2 -N kg

31 49 20 6.1 9.6 1.0 4.3 13.3 0.19

43 38 19 7.3 11.5 1.1 3.6 15.3 0.40

a) Rapid

sieving method (Kettler et al., 2001). oxidation-reduction method (Mebius, 1960). c) Kjeldahl digestion (Bremner, 1965a). d) 2 mol L−1 KCl extraction (Bremner, 1965b). e) Chlorate inhibition method (Hart et al., 1994). b) K

2 Cr2 O7

The soil samples were sieved (5 mm) and put into pots (diameter 28 cm and height 26 cm), with a mass weight of 15 kg dry soil pot−1 . Four treatments was established, i.e., no urea (control), urea (U), urea plus DCD (U+DCD), and urea plus DMPP (U+DMPP). Each treatment had 12 replicates. The crop was spring maize (Zea mays) cv. Fuyou-1. In all treatments, monocalcium phosphate and potassium chloride were applied with an application rate of 0.1 g P2 O5 kg−1 soil and 0.2 g K2 O kg−1 soil, respectively. In treatments U, U+DCD, and U+DMPP, the application rate of urea was 0.15 g N kg−1 soil, and those of DCD and DMPP were 5% and 1% of urea, respectively (Xu et al., 2001; Gioacchini et al., 2002; Yu et al., 2007). The fertilizers and inhibitors were thoroughly mixed with soil samples, which were packed and irrigated to make the soil keep constant moisture content (60% field water capacity). About 5 cm height space was kept from the soil surface to pot top. Watering was carried every day during maize growth period. Three seeds were sown in each pot, and the seedlings were thinned to one per pot after germination and seedling establishment (about 15 days). DCD and DMPP were supplied by Shanghai Chemical Institute, China, with a purity of 99.5% and 97%, respectively.

RESPONSES OF AOB AND AOA TO INHIBITORS

Sampling of soils A destructive collection of the soils in pots was made at maize seedling stage, stem elongation stage, tasselling stage, and maturity stage 21, 49, 77 and 132 days after sowing, respectively, with three replicates for each treatment. For each pot, ten soil cores were collected by using a soil auger (2.5 cm in diameter), and bulked into a composite sample. The samples were packed with ice pack and transported to laboratory, and passed through a 2-mm sieve after removing debris. Part of the samples were stored at 4 ◦ C for chemical analysis, and part of them were stored at −60 ◦ C for DNA extraction. For molecular biological analyses, about 10 g fresh soil was collected from each replicate. Determination methods − Soil NH+ 4 -N and NO3 -N contents were immediately determined by an AutoAnalyzer III continuous flow analyzer (Bran+Luebbe, Norderstedt, Germany) after extraction with 2 mol L−1 KCl solution. Soil potential ammonia oxidation (PAO) was determined by a chlorate inhibition method (Hart et al., 1994). Soil DNA was extracted from 0.5 g soil by using a FastDNA Spin kit for soil (Qbiogene, Inc., Irvine, USA) according to the manufacturer’s instructions. The purity of DNA was checked by using a Nanodrop ND-1000 UV-Vis spectrophotometer. The extracted soil DNA in triplicates was then pooled and stored at −20 ◦ C for subsequent molecular analysis. According to the protocol described in previous studies (Chen et al., 2010), briefly, the copy numbers of archaeal and bacterial amoA genes were determined by real-time PCR, using an iCycler iQ 5 thermocycler (Bio-Rad, USA) with the fluorescent dye SYBRGreen I. The DNA extracts were ten-fold diluted, and used as template with a final content of 1–9 ng in each reaction mixture. The primer pairs (amoA 1F/amoA 2R and Arch-amoA AF/Arch-amoA AR) targeted the amoA gene of the AOB and AOA DNA. Amplifications were carried out in 25 μL reaction mixtures, including 12.5 μL SYBR Premix Ex TaqTM (TaKaRa, Japan), 1 μL bovine serum albumin (25 mg mL−1 ), 0.5 μL each primer (10 μmol L−1 ) and 1 μL DNA template. Cycling conditions were 95 ◦ C for 1 min, followed by 40 cycles of 10 s at 95 ◦ C, 30 s at 55 ◦ C for AOB or 53 ◦ C for AOA, 1 min at 72 ◦ C and plate read at 83 ◦ C. A melting curve analysis was performed to check PCR products specificity after amplification, and PCR products were confirmed by standard 1% agarose gel electrophoresis. PCR products

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were gel-purified and ligated into the pGEM-T Easy vector (Promega, Madison), and the resulting ligation products were transformed into Escherichia coli JM109 competent cells. The positive clones were selected to extract plasmid DNA after re-amplification with the vector-specific primers T7 and SP6. The concentration of plasmid was determined on a Nanodrops ND-1000 UV-Vis spectrophotometer and the copy numbers of amoA genes were calculated directly from the contribution of the extracted plasmid DNA. A ten-fold dilution series of a known copy number of the plasmid DNA in triplicate was used as a standard. PCR efficiency and correlation coefficients for standard curves were 98.2% and R2 = 0.992 for AOB and 93.8% and R2 = 0.991 for AOA, respectively. All results are expressed on the basis of oven-dried soil weight (105 ◦ C, 24 h). The PCR for DGGE analysis was performed with the primer pairs containing a 40 bp GC-clamp in one primer. The components of 50 μL reaction mixtures, PCR strategy, and DGGE procedure were same to previous researchers (Chen et al., 2010). The results were analyzed by using the Quantity One software (Bio-Rad Laboratories Inc., Hercules, California, USA). Construction of amoA gene fragment libraries The construction of amoA gene fragment libraries was performed as described by He et al. (2007). According to the analysis of DGGE, ten clone libraries of the soil samples collected at maturity stage were constructed in all, due to limited resource. The ten libraries contained four treatments (control, U, U+DCD, and U+DMPP) amoA gene fragment libraries for AOB in Hap-Udic Luvisol, two (control and U+DMPP) for AOB in Hap-Ustic Luvisol, and two (control and U+DMPP) for AOA in Hap-Udic Luvisol and in Hap-Ustic Luvisol. Positive clones (about 50) were randomly selected for PCR re-amplification. The amplicons were analyzed with restriction endonuclease BstHHI and RsaI (Bio Basci, Canada). Phylogenetic analysis was performed by using MEGA version 4.0 (Tamura et al., 2007). The neighbor-jointing tree was constructed, with 1 000 replicates to produce bootstrap values. All amoA gene sequences determined in this study were deposited in GenBank nucleotide sequence database under accession numbers JF500113 to JF500142 for AOB and JF500143 to JF500166 for AOA. Statistical analysis All statistical analyses were performed by using

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SPSS version 16.0. Significant differences (P < 0.05) in obtained data between treatments were determined using analysis of variance (ANOVA) followed by Duncan’s test. Regressions were performed to understand relationships between soil nitrate contents and AOB and AOA populations. RESULTS Soil mineral nitrogen contents and PAO Fig. 1 showed that in the two test soils, NH+ 4 -N content was significantly higher in treatments U+DCD and U+DMPP than in treatments control and U at elongation stages, but the differences between the four treatments was less at tasselling and maturity stages. In Hap-Udic Luvisol, the NH+ 4 -N content in treatments U+DCD and U+DMPP had less difference at seedling and elongation stages, but in Hap-Ustic Luvisol, the NH+ 4 -N content in treatment U+DMPP was significantly higher at seedling stage but significantly lower at elongation stage, as compared with that in treatment U+DCD. The NO− 3 -N content in the two soils in all treatments had the same change trend with the growth of maize (Fig. 1), i.e., decreasing after an initial increase, with the peak at elongation stage. The soil NO− 3 -N content at seedling and elongation stages was signifi-

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cantly lower in treatments U+DCD and U+DMPP than in treatment U. The PAO in the two test soils in whole maize growth period was stimulated significantly by urea addition, as compared with that in the control (Fig. 2). DCD and DMPP suppressed the PAO significantly, comparing with applying urea alone. In Hap-Udic Luvisol, DMPP showed significant inhibition of the PAO than DCD at seedling stage, but it was adverse at elongation and tasselling stages. The PAO was higher in Hap-Ustic Luvisol than in Hap-Udic Luvisol (Fig. 2). Abundance of soil AOB and AOA The total bacterial amoA gene copy number in the two soils ranged from 2.9 (± 0.19) × 106 to 9.5 (± 1.4) × 107 copies g−1 dry soil, and that in the control changed little with time (Fig. 3). Treatment U had the largest AOB population size. Applying urea plus DCD or DMPP inhibited AOB growth significantly, as compared with applying urea alone. The AOB population density in the two soils had no significant difference among the treatments control, U+DCD, and U+DMPP during the four sampling stages, except that in Hap-Udic Luvisol at maturity stage. The AOB population density in all treatments was significantly higher in Hap-Ustic Luvisol than in Hap-Udic Luvisol.

− Fig. 1 Soil NH+ 4 -N and NO3 -N contents in Hap-Udic Luvisol (a and c) and Hap-Ustic Luvisol (b and d) during maize growth period. Error bars represent standard deviations of the means (n = 3). U = urea; DCD = dicyandiamide; DMPP = 3,4-dimethylpyrazole phosphate.

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The archaeal amoA gene copy number was significantly higher in Hap-Udic Luvisol than in Hap-Ustic Luvisol, and had no significant difference among the four treatments except the treatment control at the four sampling stages (Fig. 3). Relationships between soil nitrate content and soil AOB and AOA abundance Soil nitrate concentration had a significant positive linear correlation with soil bacterial amoA gene copy number (Fig. 4), but no significant correlation with soil archaeal amoA gene copy number (data not shown). Community composition of soil AOB and AOA

Fig. 2 Potential ammonia oxidation (PAO) in Hap-Udic Luvisol (a) and Hap-Ustic Luvisol (b) during maize growth period. Error bars represent standard deviations of the means (n = 3). U = urea; DCD = dicyandiamide; DMPP = 3,4dimethylpyrazole phosphate.

The DGGE analysis showed that the AOA community composition in the two soils at the four sampling stages was less affected by the application of urea and its combination with DCD or DMPP than the treatment control. Similarly, no significant change was observed in the AOB community composition among the treatments in Hap-Ustic Luvisol (Fig. 5). However, the AOB community composition in Hap-Udic Luvisol showed some changes, especially at the last two sampling stages (Fig. 5).

Fig. 3 Abundance of ammonia-oxidizing bacteria (AOB) and ammonia-oxidizing archaea (AOA) in Hap-Udic Luvisol (a and c) and Hap-Ustic Luvisol (b and d) at different stages of maize growth. Error bars represent standard deviations of the means (n = 3). U = urea; DCD = dicyandiamide; DMPP = 3,4-dimethylpyrazole phosphate.

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Although there were some variations in the DGGE profile among the bacterial amoA gene fragments retrieved from Hap-Udic Luvisol, most of the bacterial amoA gene sequences were affiliated with the fragment patterns belonging to clusters 3a1 and 3a2 (Table II). TABLE II Fragment patterns of bacterial amoA genes in the Hap-Udic Luvisol and the proportion of sequence types affiliated with amoA cluster in the clone libraries Treatmenta)

AOB pattern numberb)

Cluster 3a1

3a2

3b

9

1

4

0 0 0 6.0

0 4.0 0 0

%c) Control U U+DCD U+DMPP

10 8 6 7

36.2 8.0 26.0 34.0

48.9 74.0 66.0 36.0

0 0 8.0 0

14.9 14.0 0 24.0

a) U = urea; DCD = dicyandiamide; DMPP = 3,4-dimethylpyrazole phosphate. b) Number of molecular fragments specific for ammonia-oxidizing bacteria (AOB) following digestion with BstHHI. c) Percentage of analyzed clones affiliated with the fragment patterns belonging to the relative cluster among the total clone numbers analyzed in the clone library.

Fig. 4 Relationships between soil NO− 3 -N contents and ammonia-oxidizing bacteria (AOB) abundance in Hap-Udic Luvisol (a) and Hap-Ustic Luvisol (b) during maize growth period. Each point represents the mean of 3 replicates.

Most of the archaeal amoA gene sequences in treatment control could be found in treatment U+DMPP, indicating no significant community shift under DMPP and urea amendments.

Phylogenetic analysis showed that all the bacterial amoA gene sequences in the two soils fell within Nitrosospira or Nitrosospira-like species (Fig. 6), and all the archaeal amoA gene sequences belonged to group 1.1b crenarchaea.

DISCUSSION Soil mineral nitrogen Applying urea plus DCD or DMPP resulted in higher NH+ 4 concentrations in the two soils at stem elo-

Fig. 5 Denaturing gradient gel electrophoresis (DGGE) analysis of bacterial (ammonia-oxidizing bacteria) amoA fragments retrieved from Hap-Udic Luvisol (a) and Hap-Ustic Luvisol (b) under four treatments during maize growth period. U = urea; DCD = dicyandiamide; DMPP = 3,4-dimethylpyrazole phosphate.

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Fig. 6 Phylogenetic analysis of bacterial amoA gene sequences (up triangle) retrieved from Hap-Udic Luvisol. Designation of the sequences includes the following information: accession number in GenBank with treatments and number of clones in the parentheses. Bootstrap values of > 50% are displayed. Mean branch lengths are characterized by a scale bar representing the evolutionary distance. U = urea; DCD = dicyandiamide; DMPP = 3,4-dimethylpyrazole phosphate.

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ngation stage. It was substantiated by many researchers, who found DCD and DMPP could suppress soil ammonium oxidation (Xu et al., 2001; Cookson and Cornforth, 2002). DMPP or DCD could maintain more NH+ 4 during vegetative period, when maize needs to adsorb large amount of nitrogen from soil. From elongation to tasselling stage, increased ammonium concentrations in treatments control and U mainly were from mineralization of microorganism nitrogen, which was immobilized after urea addition. In the last two stages, there were no significant differences of ammonium concentrations among four treatments in two soils. This phenomenon is mainly caused by decomposition and/or leaching of DCD (Weiske et al., 2001; Zerulla et al., 2001) and DMPP (Barth et al., 2001; Weiske et al., 2001). Mineral N uptake by maize crop resulted in a decrease in total mineral N contents under pot culture conditions. Soil PAO in Hap-Ustic Luvisol was higher than that in Hap-Udic Luvisol. This may be caused by the higher pH and organic carbon content of the Hap-Ustic Luvisol, which may result in more favorable growth conditions for AOB (Nicol et al., 2008; Schauss et al., 2009). In the two soils, the different effects of DCD and DMPP on soil mineral nitrogen content and PAO may be caused by different sand and silt contents, which are key factors to strongly affect adsorption of DMPP (Barth et al., 2001). Decomposition and/or leaching of DCD and DMPP (Weiske et al., 2001) are another reason. Dynamic populations of bacterial and archaeal ammonia oxidizers The response of AOB to application of urea alone is consistent with the result by Okano et al., who found that AOB growth began immediately after ammonium addition in microcosm and field experiments (Okano et al., 2004). Similar effects of urine on AOB populations also were obtained under incubation conditions in nitrogen-rich grassland soils (Di et al., 2009) and pot culture conditions in a sandy loam soil (O’Callaghan et al., 2010). Rapid growth of AOB in response to addition of inorganic nitrogen fertilizers was also observed in an agricultural soil (Jia and Conrad, 2009). Hence, hydrolysis of urea supplies substrate for AOB to stimulate its growth. The AOB populations decreased significantly after addition of DCD or DMPP in this study. As reported in another study (O’Callaghan et al., 2010), AOB was significantly affected by DCD with reductions in its population size and activity. Some researchers also

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found that bacterial amoA gene copies decreased in the presence of DCD with urine compared with in the application of urine alone, and they noted that AOB activity was inhibited when DCD was applied by quantifying RNA copies of AOB (Di et al., 2009, 2010). No significant effects of DCD and DMPP on abundance of AOB was observed in most periods in the two soils, which might be mainly due to that AOB population was reduced in treatments U+DCD and U+DMPP compared with in control treatment. This is in accord with previous research which reported equally effective inhibition of AOB population and decrease in the ammonium oxidising rates between DCD and DMPP across six grassland sites (Di and Cameron, 2011). In contrast, other researchers have found that DMPP is more effective in reducing ammonium oxidation (Chaves et al., 2006; Pereira et al., 2010), due to differences in temperature, soil properties (Di and Cameron, 2011) and application methods of DMPP in Hap-Udic Luvisol, and bacterial amoA gene copies increase at maturity stage in the presence of DCD. The phenomenon may be caused by rapid mineralization of DCD (Weiske et al., 2001). Growth of AOA seemed to be inhibited by the high dose of urea in urea treatment at seedling stage in both soils. Previous study (Di et al., 2010) provided evidence that AOA community was inhibited by the high dose of urine till 96 days in subsoils. In the present study, however, we found that archaeal amoA copies in urea treatment increased after the seedling stage and then no significant differences were obtained among four treatments in both soils. The difference in findings might be caused by the different dose rates of urea added and the presence of a plant. In this experiment, the amount of urea-N was 0.15 g kg−1 dry soil (approximately 337.5 kg ha−1 ) with the maize growing, while the dose of urine-N in the experiment of Di et al. (2010) was 1 000 kg ha−1 without any plant. So, the higher N input and no uptake by plants may result in inhibition of AOA for a longer time in their experiment. Nitrification inhibitors DCD and DMPP had little impact on the growth of AOA in our study. Similar results were obtained by several researchers elsewhere, who reported no change in the abundance of AOA populations in the presence of DCD (Di et al., 2009, 2010; O’Callaghan et al., 2010) and DMPP (Di and Cameron, 2011; Kleineidam et al., 2011). The different responses of AOB and AOA to nitrification inhibitors are caused mainly by different susceptibilities to inhibitory compounds of bacteria and archaea (Kleineidam et al., 2011).

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The population sizes of AOB in Hap-Udic Luvisol were smaller than those in Hap-Ustic Luvisol. However, the population sizes of AOA in Hap-Udic Luvisol were greater than those in Hap-Ustic Luvisol. Soil pH is a key factor affecting abundance of ammonia oxidizers (Nicol et al., 2008). In a microcosm across a pH gradient from 4.9 to 7.5, archaeal amoA gene copy number decreased with increasing pH, while bacterial amoA gene abundance increased with increasing pH (Nicol et al., 2008). In addition, AOB growth is usually thought to be favored by fertile soil conditions, whereas AOA may be particularly adapted to low fertility environments (Martens-Habbena et al., 2009; Schauss et al., 2009; Di et al., 2010). These are in accord with our findings, which indicated that organic carbon content and pH in Hap-Ustic Luvisol were higher than those in Hap-Udic Luvisol.

monia oxidizing communities are more insensitive than bacterial ammonia oxidizing communities in paddy soil (Wang et al., 2009). Likewise, O’Callaghan et al. (2010) showed minimal metabolic response of AOA to DCD. These findings indicate that archaeal ammonia oxidizer community is less sensitive to urea and nitrification inhibitors (DCD or DMPP) than bacterial ammonia oxidizer community in the current study. Phylogenetic analysis of AOA identified that group 1.1b dominated in the two soils, which was coincident with the studies of Jia and Conrad (2009) and Xia et al. (2011), but in contrast to findings of the other researchers who found group 1.1a dominated in their experimental soils (Offre et al., 2009; Zhang et al., 2010; Verhamme et al., 2011). As suggested in a previous study, AOA community compositions are mainly determined by soil types (Chen et al., 2010).

Community analysis of bacterial and archaeal ammonia oxidizers

Relative contribution of AOA and AOB to ammonia oxidation

Phylogenetic trees of bacterial ammonia oxidizers showed that Nitrosospira or Nitrosospira-like species dominated in the two soils, which was in agreement with previous findings (Mendum et al., 1999; Phillips et al., 2000; He et al., 2007). The proportion of each cluster showed that Nitrosospira cluster 3 was dominant in Hap-Udic Luvisol, which is consistent with other studies that Nitrosospira cluster 3 was concluded to be most common in agricultural fields (He et al., 2007; Junier et al., 2009). Visible changes of AOB community composition were not found in U+DCD treatment of Hap-Udic Luvisol until day 77. This was also observed by Avrahami et al. (2002, 2003), who reported no differences in the AOB community composition detected within short periods due to the slow growth of populations. However, no shifts of the DGGE patterns based on AOB community were observed in Hap-Ustic Luvisol. The different responses of AOB community composition to nitrification inhibitors in the two soils may be due to different soil types determining AOB community compositions (Chen et al., 2010). DGGE and phylogenetic analysis showed the addition of urea and nitrification inhibitors (DCD or DMPP) had few effects on the diversity of archaeal ammonia oxidizer community in the short term (until 132 days after application). Similar results was observed by Jia and Conrad (2009), who reported no discernable changes in the AOA community found between samples with and without the application of (NH4 )2 SO4 /NH4 HCO3 and/or nitrification inhibitor C2 H2 . In response to urea application, archaeal am-

Growth of AOB was stimulated by urea addition and significantly suppressed by nitrification inhibitors, and a significant linear correlation was found between soil nitrate contents in the two soils and AOB rather than AOA populations. Additionally, AOA abundance was less variable between treatments when PAO was suppressed by nitrification inhibitors. These results suggested that AOA had little contribution to ammonia oxidation under the conditions of the present study. These results contrast with those of Offre et al. (2009), who reported that determined ammonia oxidation is mostly due to AOA in a soil with a pH of 7.5 because growth of only archaeal rather than bacterial ammonia oxidizers occurs during active nitrification. At the same site with different soil pH (4.5 and 6), similar results were obtained by other researchers (GubryRangin et al., 2010), who found that nitrate production is strongly correlated with abundance of archaeal but not bacterial ammonia oxidizers, and suggested that archaea rather than bacteria control nitrification under the conditions of their study. Although soil pH varied from 6.1 to 7.3 in our study, ammonia oxidation seemed to be dominated by bacterial ammonia oxidizers. This could be confirmed by the significant positive relationship between soil nitrate contents and AOB but not AOA gene copies. Additionally, some or all group 1.1b (in our study) and group 1.1a archaeal ammonia oxidizers differ fundamentally in their physiological and metabolic characteristics (Zhang et al., 2010). The difference in the groups of AOA present may be an important factor. In addition, further study needs to be undertaken to investigate the dynamic populations of

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active ammonia oxidizers by extracting RNA. ACKNOWLEDGEMENTS We are grateful to the Research Centre for Ecoenvironmental Sciences, Chinese Academy of Sciences for allowing us to use their molecular labs. The authors wish to thank Prof. Zhou Li-Kai and Dr. Zhang ChengGang in our institute for their valuable suggestions on the work and Dr. Shen Ju-Pei and Dr. Zhang Xiao-Li in our institute for the technical assistance. Thanks also to the editors and anonymous reviewers for their insightful and helpful comments on the manuscript. REFERENCES Avrahami, S., Conrad, R. and Braker, G. 2002. Effect of soil ammonium concentration on N2 O release and on the community structure of ammonia oxidizers and denitrifiers. Appl. Environ. Microb. 68: 5685–5692. Avrahami, S., Liesack, W. and Conrad, R. 2003. Effects of temperature and fertilizer on activity and community structure of soil ammonia oxidizers. Environ. Microbiol. 5: 691–705. Barraclough, D. and Puri, G. 1995. The use of 15 N pool dilution and enrichment to separate the heterotrophic and autotrophic pathways of nitrification. Soil Biol. Biochem. 27: 17–22. Barth, G., von Tucher, S. and Schmidhalter, U. 2001. Influence of soil parameters on the effect of 3,4-dimethylpyrazolephosphate as a nitrification inhibitor. Biol. Fert. Soils. 34: 98–102. Bremner, J. M. 1965a. Total nitrogen. In Black, C. A. (ed.) Methods of Soil Analysis. Part 2. Chemical and Microbiological Properties. American Society of Agronomy, Madison, Wisconsin. pp. 1149–1178. Bremner, J. M. 1965b. Inorganic forms of nitrogen. In Black, C. A. (ed.) Methods of Soil Analysis. Part 2. Chemical and Microbiological Properties. American Society of Agronomy, Madison, Wisconsin. pp. 1179–1237. Chaves, B., Opoku, A., De Neve, S., Boeckx, P., Van Cleemput, O. and Hofman, G. 2006. Influence of DCD and DMPP on soil N dynamics after incorporation of vegetable crop residues. Biol. Fert. Soils. 43: 62–68. Chen, X., Zhang, L. M., Shen, J. P., Xu, Z. H. and He, J. Z. 2010. Soil type determines the abundance and community structure of ammonia-oxidizing bacteria and archaea in flooded paddy soils. J. Soil. Sediment. 10: 1510–1516. Cookson, W. R. and Cornforth, I. S. 2002. Dicyandiamide slows nitrification in dairy cattle urine patches: effects on soil solution composition, soil pH and pasture yield. Soil Biol. Biochem. 34: 1461–1465. De Boer, W. and Kowalchuk, G. A. 2001. Nitrification in acid soils: micro-organisms and mechanisms. Soil Biol. Biochem. 33: 853–866. Di, H. J. and Cameron, K. C. 2011. Inhibition of ammonium oxidation by a liquid formulation of 3,4-Dimethylpyrazole phosphate (DMPP) compared with a dicyandiamide (DCD) solution in six New Zealand grazed grassland soils. J. Soil. Sediment. 11: 1032–1039. Di, H. J., Cameron, K. C., Shen, J. P., Winefield, C. S., O’Callaghan, M., Bowatte, S. and He, J. Z. 2009. Nitrification driven

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