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Are China’s regional agricultural productivities converging: How and why?
Soil & Tillage Research 195 (2019) 104433
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Different ammonia oxidizers are responsible for nitrification in two neutral paddy soils Zhihui Wanga, Yanqiang Caoa,b, Alan L. Wrightc, Xiuli Shid, Xianjun Jianga,
T
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a
College of Resources and Environment, Southwest University, 2 Tiansheng Road, Beibei, Chongqing, 400715, China Guangxi Key Laboratory of Plant Conservation and Restoration Ecology in Karst Terrain, Institute of Botany, Guangxi Zhuang Autonomous Region and Chinese Academy of Sciences, Guilin, 541006, China c Indian River Research & Education Center, University of Florida-IFAS, Fort Pierce, FL, 34945, USA d College of Horticulture and Landscape Architecture, Southwest University, 2 Tiansheng Road, Beibei, Chongqing, 400715, China b
A R T I C LE I N FO
A B S T R A C T
Keywords: Ammonia oxidizers Soil nitrification Niche differentiation
Human intervention in agriculture, such as farming and fertilization, could result in major changes of ammonia availability that shapes niche-specific occupied communities of soil ammonia oxidizers. The aim of this study was to investigate nitrification and active ammonia oxidizers in N-rich and N-limited soils, which were induced by different management practices. Here, we used DNA-based stable isotope probing (DNA-SIP) microcosm to decipher active ammonia-oxidizing archaea (AOA) and bacteria (AOB) phylotypes involved in ammonia oxidation in the cultivated and uncultivated soils. The results showed that net nitrification rate in the cultivated soil was significantly higher than in the uncultivated soil (6.19 and 1.40 mg N kg−1 dry soil d-1, respectively). Growth of soil AOB and AOA during incubation occurred in the cultivated soil, while in the uncultivated soil, only AOA significantly increased after 56 days. Combining DNA-SIP and sequencing results, we gathered evidence of ammonia oxidation by Nitrososphaera-like AOA, in addition to active AOB, within the classic Nitrosospira group in the cultivated soil. For the uncultivated soil, ammonia oxidation was driven by Nitrososphaera-like and Nitrosopumilus-like AOA, rather than AOB. Our results demonstrate the potential of ammonium concentration for shaping the communities of active AOA and AOB, which imply that ammonia-dependent “diversification” may be important strategies for niche differentiation of sympatric AOA/AOB under different field management conditions.
1. Introduction
AOB (Prosser and Nicol, 2012; He et al., 2012). Many factors could result in the niche differentiation of AOA and AOB in soils, including substrate concentration (Martens-Habbena et al., 2009), pH (GubryRangin et al., 2011), O2, temperature (Hatzenpichler, 2012) and mixotrophy (Prosser and Nicol, 2012). Among them, soil pH and precipitation were suggested to be principal drivers in shaping niche differentiation of microbes at large-scales (Bahram et al., 2018). However, it is unclear whether microbial niche differentiation can be driven by management of soils. Human intervention in agriculture, such as farming and fertilization, could result in major changes in soil physicochemical and biological properties (e.g. soil prokaryotic communities, pH, N availability) (Babin et al., 2019; Das et al., 2019). Numerous studies have suggested that ammonia concentration is an important factor shaping the niche differentiation between AOA and AOB (Verhamme et al., 2011; Sterngren et al., 2015; Daebeler et al., 2015). However, with few exceptions, most of these observations resulted from AOA and AOB amoA
Nitrification in agricultural soils can lead to nitrogen (N) losses following ammonia-based fertilizer application through conversion of ammonium into nitrate (Gruber and Galloway, 2008). Consequently, leaching of nitrate results in pollution of streams and groundwater, and denitrification losses increase nitrous oxide emissions (Ravishankara et al., 2009). Thus, ammonia oxidizers, crucial mediators of nitrification, are widely recognized as key players for sustainable N management of agricultural ecosystems and environmental stewardship. Ammonia-oxidizing bacteria (AOB), first isolated in the 19th century (Winogradsky, 1890), were thought to govern ammonia oxidation exclusively. In 2005, ammonia-oxidizing archaea (AOA) were proven to be a new group capable of oxidizing ammonia (Könneke et al., 2005; Francis et al., 2005). However, genomic studies of archaeal isolates suggested the unique biochemical and genetic traits from their bacterial counterparts, which supposes niche differentiation between AOA and ⁎
Corresponding author. E-mail address: [email protected] (X. Jiang).
https://doi.org/10.1016/j.still.2019.104433 Received 21 December 2018; Received in revised form 13 August 2019; Accepted 25 September 2019 0167-1987/ © 2019 Elsevier B.V. All rights reserved.
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2.2. Microcosm construction
gene quantification, and their abundances alone can not provide enough information about the relative importance of bacteria and archaea for NH3-oxidation because the genes might not have been expressed or enzymes were inactivated. Since not all microbes possessing amoA genes are active oxidizers for NH3-oxidation (Prosser and Nicol, 2008; Stahl and de la Torre, 2012), and most analyses of NH3-oxidizer communities usually include populations not actively contributing to nitrification, DNA-based stable isotope probing (DNA-SIP) techniques can instead be used to directly decipher active AOA and AOB involved with nitrification for in situ field conditions. This technique is thus a powerful means to link functional processes to taxonomic identities of active microorganisms in complex soil environments (Radajewski et al., 2003). We hypothesized that: (i) at the local scale, ammonia availability induced by management practices is the key driver of niche differentiation between AOA and AOB; (ii) the limited nitrogen caused by long-term non-cultivation will result in the predominance of AOA in nitrification for the uncultivated soil due to their higher affinity for substrates. Therefore, 13CO2-based DNA-SIP, quantitative PCR and nucleotide sequencing approach were used to investigate nitrification and active ammonia oxidizers under different agricultural managements, cultivated and uncultivated soils, which represent N-rich and N-limited environments, respectively.
Soil DNA-SIP microcosms were constructed to investigate the active soil ammonia oxidizers community as previously described (Jia and Conrad, 2009). Briefly, soil microcosms were established in 120 mL serum bottles tightly capped with black butyl stoppers which contained sieved soil (8.0 g dry weight soil) and 5% headspace (vol/vol) 13CO2 (99% atom enriched, Sigma-Aldrich Co., St Louis, MO, USA) or 12CO2 (generated by acidifying sodium carbonate). Besides 12CO2-control microcosms and 13CO2-labeled microcosms, an additional treatment included a 13CO2 + C2H2 (100 Pa) microcosm to assess whether 13CO2 assimilation by ammonia oxidizers was dependent on energy generated from ammonia oxidation that could be completely inhibited by C2H2 (Berg et al., 1982). Each treatment received 50 mg NH4+-N kg−1 dry soil weekly and microcosms were incubated at 60% soil water-holding capacity at 28 °C in the dark throughout the eight-week incubation. Each bottle was opened weekly to allow N addition and air exchange to maintain aerobic conditions. Water lost by evaporation was replenished with sterilized water, and 13CO2, 12CO2, and C2H2 in headspace were renewed immediately after bottle resealing. All treatments were performed in triplicate. Soil (2 g) from each bottle was destructively harvested at day 0 and day 56, and stored immediately in a −80 °C freezer (Meiling Biology & Medical, Anhui, China). Remaining soil was used for determination of ammonium and nitrate using a SKLAR continuous-flow analyzer (SKLAR San++, Netherland, 2003). Soil deoxyribonucleic acids were extracted from 0.5-g soil samples using a FastDNA spin kit for soil (MP Biomedicals, Cleveland, OH, USA) according to the manufacturer’s instructions. Extracted DNA was used for subsequent SIP and quantification of amoA genes.
2. Materials and methods 2.1. Study site description and soil collection The study site, Purple Soil Ecology Experimental Station of Southwest University, is located in the southwest of China (30° 26′ N, 106° 26′ E). The climate is subtropical with a mean annual temperature of 18.2 °C and a mean annual total rainfall of 1105 mm. Both soils developed from the purple mudstone and are classified as Eutric Regosol (FAO/UNESCO, 1988). The tested neutral paddy soils were collected from two long-term treatments established in 1990: the cultivated soil was in a rice (Oryza sativa L.) and oilseed rape (Brassica napus L.) cropping rotation, with conventional tillage as previously described (Li et al., 2015). The amount of fertilizer applied to the cultivated soil annually is urea 270 kg ha−1, superphosphate 500 kg ha−1, potassium chloride 150 kg ha-1. The fertilization strategy is that 60% of N, 100% of P and 50% of K are applied as base fertilizer before planting rape and rice, and then 40% of N and 50% of K as topdressing at the 5–6 leaf stage for rape or at the tillering stage for rice. The uncultivated field was left uncultivated and unfertilized, and the main vegetation of this fallow field was weeds (Gramineae). Both cultivated and uncultivated fields were under the same water management: fields were irrigated after rape harvest in May and kept flooded with a layer of water until rice maturation, and after rice harvest in the middle of August, fields were drained and remained dry for rape planting until the next May. The properties for the cultivated soil are: pH 7.3 (H2O), 16.5 g organic matter kg−1 dry soil, 1.37 g total N kg−1 dry soil, 10.7 mg NH4+-N kg−1 dry soil, and 13.6 mg NO3–N kg−1 dry soil. Properties for the uncultivated soil are: pH 7.3 (H2O), 20.5 g organic matter kg−1 dry soil, 0.921 g total N kg−1 dry soil, 4.08 mg NH4+-N kg−1 dry soil, and 9.50 mg NO3–N kg−1 dry soil. Each field contains three replicated plots of dimension 4 m × 5 m. Five soil cores were collected at a 0–20 cm depth from each plot using a 13 cm diameter soil auger in November 2017, and then mixed thoroughly to reduce heterogeneity and pooled to form composite samples. Three composite samples were collected from each sampling site generating three replicates for each treatment. Soil samples were transferred to the laboratory on ice and passed through a 2 mm sieve to remove roots and stones. Then each soil sample was divided into two parts: one was stored at 4 °C until SIP incubation, another was air dried to analyze the chemical properties with the same methods as we previously described (Wang et al., 2019).
2.3. SIP gradient fractionation The total DNA extract was subjected to isopycnic centrifugation to separate 13C-DNA from native 12C-DNA as previously described (Jia and Conrad, 2009). About 3.0 μg of total DNA extracts were thoroughly mixed with a CsCl solution to reach an initial CsCl buoyant density of 1.725 g mL−1 and placed in a 5.1 mL Beckman polyallomer tube. Suspensions were centrifuged in a Vti65.2 vertical rotor (Beckman Coulter Inc., Palo Alto, CA, USA) at 177,000 g for 44 h at 20 °C. Fifteen DNA fractions (∼380 μL) were obtained by displacing the gradient medium from the top of the ultracentrifuge tube with sterile water using a syringe pump (New Era Pump Systems Inc., Farmingdale, NY, USA). The buoyant density of each DNA fraction was measured indirectly using an AR200 digital hand-held refractometer (Reichert Inc., Buffalo, NY, USA). The fractionated DNA was precipitated and purified as Jia and Conrad (2009) described and dissolved in 30 μL TE buffer. 2.4. Quantification of amoA genes Real-time quantitative PCR (qPCR) was performed to estimate the abundance of AOA and AOB during the eight-week incubation, and the abundance of 13CO2-labeled amoA-carrying ammonia oxidizers in each CsCl fraction on a QuantStudio 6 Flex Real-Time PCR Systems (Applied Biosystems of Life Technologies, Singapore). Bacterial and archaeal amoA genes were amplified by primer pairs amoA-1 F/amoA-2R with amplicon length of 491 bp (Rotthauwe et al., 1997) and Arch-amoAF/ Arch-amoAR with amplicon length of 635 bp (Francis et al., 2005), respectively. The reaction was performed in a 20 μL mixture containing 10 μL SYBR Premix Ex Taq (TaKaRa, Dalian, China), 0.5 μL of each primer (10 μM), 0.4 μL of ROX Reference Dye II (50×) and 1 μL of DNA template (1–10 ng), and 7.6 μL sterile water. QPCR was performed in biological triplicates with three technical replicates. A plasmid DNA from one representative clone containing AOA or AOB amoA genes was used as the template to generate the standard curve, and the detailed steps and the subsequent amplification condition were the same as 2
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previously described (Jiang et al., 2015). The final amplification efficiencies were 95%–103% with R2 values of 0.993-0.999. Additionally, a melting curve analysis was used to check the specificity of the amplification products at the end of each qPCR run.
bacteria in both soils. 3.3.
13
CO2-labeled DNA in AOA and AOB
CO2-labeled DNA in the ‘heavy’ fractions was separated from native 12CO2-control DNA in the ‘light’ fractions for each SIP microcosm, and qPCR assessed amoA genes of each buoyant density of the SIP gradient fraction. AOA or AOB populations assimilating 13C into their genomes are regarded as being active ammonia oxidizers (Jia and Conrad, 2009; Wang et al., 2015b). Two peaks of archaeal and bacterial amoA gene abundance were observed in the gradient ranges of DNA fractions for 13CO2-labeled microcosms of cultivated soil, and considerable AOA and AOB were detected in the ‘heavy’ fraction (buoyant densities 1.725–1.735 g mL−1), implying archaeal and bacterial populations were labeled by 13CO2. For 12CO2-control microcosms, archaeal and bacterial amoA genes peaked in the native ‘light’ fractions with buoyant densities of 1.70 g mL-1 (Fig. 2a, c). These results demonstrated substantial 13CO2 assimilation by both AOA and AOB in the cultivated soil. However, in the uncultivated soil, archaeal amoA genes peaked in the ‘light’ fractions and the ‘heavy’ fractions for the control and labeled microcosms, respectively (Fig. 2b), while bacterial amoA genes peaked in the native ‘light’ fractions for both the control and labeled microcosms (Fig. 2d), indicating that only AOA was active during nitrification. 13
2.5. Sequencing and phylogenetic analysis MiSeq sequencing of archaeal and bacterial amoA genes in the fractionated DNA from 13CO2 labeled SIP microcosms were carried out on an Illumina MiSeq sequencer instrument (USA) at a Personal Biotechnology Co., Ltd. (Shanghai, China). The amplification of the fractionated DNA was performed using the primers amoA-1 F/amoA-2R and Arch-amoAF/Arch-amoAR with specific barcode sequences for AOA and AOB amoA genes, respectively. The PCR procedures and quality control of the obtained raw sequences were based on methods of Zhao et al. (2015). Reads with quality score lower than 20, ambiguous bases and improper primers were discarded. The resulting high-quality sequences were subgrouped to obtain operational taxonomic units (OTUs) at 97% sequence similarity levels. Phylogenetic trees of AOA and AOB amoA genes were constructed with MEGA 4.0 package through a neighbor-joining algorithm (Kimura 2-parameter distance model) using 1,000 bootstrap replicates (Tamura et al., 2007). 2.6. Statistical analyses One-way analysis of variance was performed to check the variance between different treatments followed by Fisher’s protected least significant difference test. Net nitrification rate (mg N kg−1 dry soil d−1) was calculated as the difference between the final and the initial nitrate concentrations divided by the incubation time (Robertson et al., 1999). All analyses were conducted using SPSS 13.0 package (SPSS, USA), and P < 0.05 was considered to be statistically significant.
3.4. Phylogenetic analysis of active nitrifying communities For cultivated soil, the obtained reads were binned into 10 and 8 OTUs for the archaeal and bacterial amoA genes, respectively. Phylogenetic analysis of amoA showed that 95.8% of active AOA fell within the group I.1b-Nitrososphaera lineage. OTU1 was the most abundant species which was closely related to Nitrososphaera sp. JG1 (JF748723) (with 96.9% identities), accounting for 80.7% of all active AOA reads. The remaining 4.2% was affiliated with group I.1aNitrosopumilus lineage (Fig. 3). As for active AOB in cultivated soil, 78.9% of reads were grouped into Nitrosospira cluster 3a.1, with genus Nitrosospira sp. Nsp2 (AJ298719) as the closest cultured relative. 21.0% of reads were affiliated with Nitrosospira cluster 3a.2, which was related to Nitrosospira multiformis (X90822), and the remaining small proportion was clustered into Nitrosospira cluster 9 (Fig. 4). For uncultivated soil, the high quality reads of archaeal amoA genes were binned into 10 OTUs and equally distributed into two clusters—50% reads were affiliated with the soil group I.1b-Nitrososphaera lineage and the other 50% fell into the marine group I.1a-Nitrosopumilus lineage. The most abundant OTU was closely related to an uncultured archaeon in sediments (KJ005105) in group I.1a-Nitrosopumilus lineage, accounting for 37.7% of the total amoA gene reads (Fig. 5).
3. Results 3.1. Nitrification activity For both soils, significant accumulation of ammonium was observed at day 56 in the presence of C2H2 compared to day 0 (P < 0.05). In the absence of C2H2, ammonium did not differ between Day 56 and Day 0 for cultivated soil (P > 0.05), but ammonium increased significantly at Day 56 for uncultivated soil (P < 0.05) (Fig. 1a). For cultivated soil, NO3−-N concentrations averaged 13.6 mg NO3−-N kg-1 at day 0, and increased significantly to 359 and 361 mg NO3−-N kg-1 at day 56 for no C2H2 treatments (Fig. 1c). For uncultivated soil without C2H2, NO3−-N increased by 78.4 mg kg-1 after 56 days, showing a lower net nitrification rate (1.40 mg N kg-1 dry soil d-1) compared to the same treatment with cultivated soil (6.19 mg N kg-1 dry soil d-1). Soil nitrate production was negated with C2H2 addition for both cultivated and uncultivated soils (Fig. 1c), indicating that microbial NH3-oxidation was inhibited by C2H2 during the incubation. No significant nitrification was detected over the course of the 56-day incubation in the 13CO2+C2H2 soil microcosms.
4. Discussion The DNA-SIP technique provided strong evidence that niche differentiation of sympatric ammonia oxidizers occurred in these two neutral soils, as both archaea and bacteria played roles in autotrophic nitrification in cultivated soil, but only AOA were active in autotrophic nitrification for uncultivated soil. Phylogenetic analysis clearly showed that the dominant active archaea and bacteria in cultivated soils were affiliated with Nitrososphaera subcluster 1.1 within the soil group 1.1b lineage and Nitrosospira cluster 3a.1, respectively, however the dominant ammonia oxidizers in uncultivated soil was only AOA in Nitrososphaera subcluster 1.1 within the soil group 1.1b lineage and Nitrosopumilus cluster within the marine group I.1a lineage. For uncultivated soil, there were concurrent low numbers of AOB amoA genes at the beginning and the end of incubation (Fig.1d), indicating that AOB could not account for the observed nitrification rate. This result was also supported by the SIP data suggesting that no bacterial amoA genes were labeled by 13CO2 (Fig. 2d). However, AOA
3.2. AOA and AOB abundance For the cultivated soil, in the absence of C2H2, amoA gene abundance increased from 3.60 × 107 g−1 dry soil at Day 0 to 8.80 × 107 g−1 dry soil at Day 56 and 2.85 × 106 g−1 dry soil at Day 0 to 1.18 × 107 g−1 dry soil at Day 56, corresponding to 1.45- and 3.11 fold increases for AOA and AOB after incubation (Fig. 1b, d). For the uncultivated soil, AOA increased significantly from 2.85 × 106 g−1 dry soil at Day 0 to 1.17 × 107 g−1 dry soil at Day 56 in the absence of C2H2 (P < 0.05), while AOB showed no significant change from Day 0 (1.50×105 g−1 dry soil) to Day 56 (1.46 × 105 g−1 dry soil) (P > 0.05) (Fig. 1b, d). The addition of C2H2 inhibited growth of archaea and 3
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Fig. 1. The concentrations of soil ammonium (a) and nitrate (c), and the abundances of archaeal (b) and bacterial (d) amoA genes in cultivated and uncultivated soil microcosms over a 56-days incubation at 28 °C. The error bars indicate standard errors of the mean of the triplicate microcosms. The different letters above the columns indicate a significant difference (P < 0.05).
Fig. 2. Quantitative distribution of the relative abundances of archaeal (a, b) and bacterial (c, d) amoA genes across the entire buoyant density gradient of the DNA fractions retrieved from 12CO2 and 13CO2 56-day DNA-SIP microcosms of cultivated and uncultivated soil, respectively. Error bars represent standard errors from three replicates.
was 183 and was significantly higher than cultivated soil (12.6). Similarly, in a survey from 12 pristine and agricultural soils of three climatic zones, the ratio of AOA/AOB amoA gene copies ranged from 1.5 to 232, and the highest ratio occurred for non-fertilized soil (Leininger et al., 2006). This result is not surprising because the uncultivated neutral purple soil has not received cultivation or N fertilization for about 30 years, which created a N-limited environment
abundance was significantly higher than its bacterial counterpart and increased with nitrification (Fig. 1b). We can therefore assume that AOA plays a role in ammonia oxidization in uncultivated soil, which is in line with SIP results (Fig. 2b). These results are consistent with previous studies attributing ammonia oxidization to AOA in nutrientlimited, non-fertilized soils (Daebeler et al., 2014, 2015; Sterngren et al., 2015; Shi et al., 2018). The ratio of AOA/AOB in uncultivated soil 4
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Fig. 3. Phylogenetic analysis of the archaeal amoA genes from the 13CO2-labeled microcosms at the end of incubation in the neutral cultivated soil. The designation AOA-OTU1 (80.7%) indicates that OTU1 account for 80.7% of the AOA amoA reads from the labeled microcosms. One representative sequence from each OTU was selected using the mothur software for the tree construction. Archaeal amoA subclusters were classified according to Pester et al. (2012).
Nguyen et al., 2019; Kong et al., 2019). Soil ammonia concentration is regarded as one of the most critical factors shaping the sympatric differentiation of archaea and bacteria communities (Martens-Habbena et al., 2009; Prosser and Nicol, 2012). Our results provide compelling evidence for substrate-induced niche differentiation of sympatric AOA/AOB for these two soils. For the cultivated soil (N-rich condition), the dominant active AOA was affiliated with Nitrososphaera subcluster 1.1 and the major active AOB were affiliated with Nitrosospira cluster 3a.1. For the uncultivated soil (N-limited condition), only AOA was labeled and the dominant active AOA was affiliated with Nitrososphaera subcluster 1.1 and Nitrosopumilus cluster with a closely cultured relative of genus Nitrosopumilus maritimus which possesses an extremely high substrate affinity. These results are consistent with previous findings that AOA is the dominant NH3-oxidizer in oligotrophic environments, and possibly may be the only driver of NH3-oxidation in soils with deficient available nitrogen (Daebeler et al., 2014, 2015; Sterngren et al., 2015). On the contrary, AOB appears to dominate NH3 oxidation after addition of N fertilizer at high concentrations (Jia and Conrad, 2009; Zhong et al., 2016; Xiang et al., 2017). It is worth noting that in the present study, only 4.2% of active AOA belong to the group I.1a-Nitrosopumilus lineage in cultivated soil, but in the uncultivated soil, contributions were up to 50% (Figs. 3 and 5), indicating that substrate can shift the predominance of phylotypes within AOA or AOB, not only between AOA and AOB. Nitrosopumilus-
favoring AOA rather than AOB owing to a much higher NH3 affinity of archaea (Prosser and Nicol, 2012; Shi et al., 2018). The relative importance of archaea and bacteria in ammonia oxidation is influenced by many environmental factors, including pH (Gubry-Rangin et al., 2011), ammonia concentration (Prosser and Nicol, 2012), temperature (Avrahami and Conrad, 2003), water content (Hu et al., 2015), and dissolved oxygen levels (Santoro et al., 2008). Numerous studies have shown that AOA is an active player in acidic soils, while AOB is the driver of nitrification in neutral and alkaline soils (Taylor et al., 2012; Jiang et al., 2015; Shi et al., 2018). However, the diversity and abundance of AOA increases with rising soil pH (Gubry-Rangin et al., 2011), suggesting that some phylotypes of archaea may be active in NH3-oxidation in neutral or alkaline soils. In the cultivated soil, both AOA and AOB amoA gene abundances significantly increased and were paralleled by an increase in nitrate concentrations (347 mg N kg−1 d.w.s) (Fig. 1), indicating that both AOA and AOB contributed to nitrification. This result was subsequently confirmed by 13 CO2-labeled NH3-oxidizers, demonstrating that substantial 13CO2 was assimilated into archaea and bacteria (Fig. 2). The phylogenetic analysis further revealed that the dominant active AOA and AOB fell into Nitrososphaera and Nitrosospira clusters, respectively, which provided strong evidence that both Nitrososphaera-like AOA and Nitrosospira-like AOB are active regulators of nitrification in this neutral soil (Figs. 3 and 4). This result was similar to those found in previous studies showing a cooperation of AOA and AOB in neutral soils (Wang et al., 2015a;
Fig. 4. Phylogenetic analysis of the bacterial amoA genes from the 13CO2-labeled microcosms at the end of incubation in the neutral cultivated soil. The designation AOB-OTU2 (18.2%) indicates that OTU1 account for 18.2% of the AOA amoA reads from the labeled microcosms. One representative sequence from each OTU was selected using the mothur software for the tree construction. Bacterial amoA clusters were classified according to Avrahami and Conrad (2003).
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Fig. 5. Phylogenetic analysis of the archaeal amoA genes from the 13CO2-labeled microcosms at the end of incubation in the neutral uncultivated soil. The designation AOA-OTU1 (13.5%) indicates that OTU1 account for 13.5% of the AOA amoA reads from the labeled microcosms. One representative sequence from each OTU was selected using the mothur software for the tree construction. Archaeal amoA subclusters were classified according to Pester et al. (2012).
respectively). The qPCR of amoA genes showed higher AOA abundance than AOB for both soils, with AOA/AOB for the cultivated and uncultivated soils averaging 12.6 and 183, respectively. Obvious growth of soil AOB and AOA occurred during incubation in the cultivated soil, while in the uncultivated soil, only AOA increased significantly. Our results provide robust evidence for niche differentiation of active AOA and AOB phylotypes in the cultivated and uncultivated soils. The phylogenetic analysis reveals that Nitrososphaera-like AOA and Nitrosospira-like AOB drive ammonia oxidization in concert in the Nrich cultivated soil, but in the N-limited uncultivated soil, the active drivers are soil group I.1b Nitrososphaera-like and marine group I.1a Nitrosopumilus-like AOA lineages. These results suggest that soil ammonia oxidization is carried out by diverse physiologically versatile ammonia-oxidizing phylotypes in soils with different management, indicating that ammonium-dependent “diversification” may be an important strategy for niche differentiation of sympatric AOA/AOB under variable field conditions.
like AOA are often found in aquatic environments, such as the open ocean, but are less common in soils (Offre et al., 2009; Pester et al., 2012). The seasonal flooding of the paddy soil causes oxygen stress and favors Nitrosopumilus lineage archaea which has a higher affinity for O2 compared to Nitrososphaera lineage (Martens-Habbena et al., 2009; Jung et al., 2011). Interestingly, many previous DNA SIP studies of fertilized paddy soils found that active AOA was affiliated with group I.1b-Nitrososphaera lineage, rather than group I.1a-Nitrosopumilus lineage (Wang et al., 2015a; Zhao et al., 2015). However, some studies indeed directly link group I.1a-Nitrosopumilus lineage AOA to active ammonia oxidization in an ammonia-limited volcanic grassland soil and an agricultural soil in Scotland with low ammonia concentration (Daebeler et al., 2014; Zhang et al., 2010). These results indicated that substrate limitation selected active I.1a-Nitrosopumilus lineage. Some phylotypes such as Nitrosopumilus lineage could successfully compete with its sympatric Nitrososphaera lineage when substrates become more deficient, suggesting a higher ammonia affinity of archaea in Nitrosopumilus cluster than in Nitrososphaera cluster just like their isolated cultured strains (Martens-Habbena et al., 2009; Jung et al., 2011). In this study, although 50 mg N/kg d.s.w/week ammonium sulfate was added to soil, no obvious growth of AOB was found during incubation, but AOA increased significantly. Given that AOB has a faster growth rate than AOA (Jiang and Bakken, 1999; Tourna et al., 2011), we can conclude that AOA was already active under the field conditions. The oligotrophic condition led to the inactivation of AOB in ammonia oxidization and selectively prospered the Nitrosopumilus cluster AOA, implying that AOA/AOB may develop their “diversity” to adapt to different environments based upon long-term evolutionary shifts (Lennon and Jones, 2011). The diversity of archaeal or bacterial lineages may be a result of evolution based upon specific adaptations to soil pH, ammonia availability or oxygen content, and thus show niche specialization. The recent discovery of “comammox”, which occurs when one nitrifying microorganism carries out the complete oxidation of ammonia to nitrate (van Kessel et al., 2015v; Daims et al., 2015), contributed to a growing appreciation that nitrifiers are more versatile than formerly believed (Santoro, 2016). Understanding the niche differentiation of AOB and AOA to substrates can thus improve our ability to manage N more efficiently for agriculture.
Acknowledgments We are grateful to the editor and three anonymous referees for their careful reviews and constructive suggestions. This research was financially supported by the National Natural Science Foundation of China (41671232), the National Key Research and Development Program of China (2016YFD0300901), the Graduate Research and Innovation Program of Chongqing (CYB19105). Zhihui Wang thanks the China Scholarship Council (CSC) for her scholarships. References Avrahami, S., Conrad, R., 2003. Patterns of community change among ammonia oxidizers in meadow soils upon long-term incubation at different temperatures. Appl. Environ. Microbiol. 69, 6152–6164. Babin, D., Deubel, A., Jacquiod, S., Sørensen, S.J., Geistlinger, J., Grosch, R., Smalla, K., 2019. Impact of long-term agricultural management practices on soil prokaryotic communities. Soil Biol. Biochem. 129, 17–28. Bahram, M., Hildebrand, F., Forslund, S.K., Anderson, J.L., Soudzilovskaia, N.A., Bodegom, P.M., Bengtsson-Palme, J., Anslan, S., Coelho, L.P., Harend, H., HuertaCepas, J., Medema, M.H., Maltz, M.R., Mundra, S., Olsson, P.A., Pent, M., Põlme, S., Sunagawa, S., Ryberg, M., Tedersoo, L., Bork, P., 2018. Structure and function of the global topsoil microbiome. Nature 560, 233–237. Berg, P., Klemedtsson, L., Rosswall, T., 1982. Inhibitory effect of low partial pressures of acetylene on nitrification. Soil Biol. Biochem. 14, 301–303. Daebeler, A., Bodelier, P.L., Yan, Z., Hefting, M.M., Jia, Z., Laanbroek, H.J., 2014. Interactions between Thaumarchaea, Nitrospira and methanotrophs modulate autotrophic nitrification in volcanic grassland soil. ISME J. 8, 2397–2410. Daebeler, A., Bodelier, P.L.E., Hefting, M.M., Laanbroek, H.J., 2015. Ammonia-limited conditions cause of Thaumarchaeal dominance in volcanic grassland soil. FEMS
5. Conclusions Net nitrification rate of the cultivated soil was significantly higher than the uncultivated soil (6.19 and 1.40 mg N kg−1 dry soil d−1, 6
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Different ammonia oxidizers are responsible for nitrification in two neutral paddy soils Zhihui Wanga, Yanqiang Caoa,b, Alan L. Wrightc, Xiuli Shid, Xianjun Jianga,
T
⁎
a
College of Resources and Environment, Southwest University, 2 Tiansheng Road, Beibei, Chongqing, 400715, China Guangxi Key Laboratory of Plant Conservation and Restoration Ecology in Karst Terrain, Institute of Botany, Guangxi Zhuang Autonomous Region and Chinese Academy of Sciences, Guilin, 541006, China c Indian River Research & Education Center, University of Florida-IFAS, Fort Pierce, FL, 34945, USA d College of Horticulture and Landscape Architecture, Southwest University, 2 Tiansheng Road, Beibei, Chongqing, 400715, China b
A R T I C LE I N FO
A B S T R A C T
Keywords: Ammonia oxidizers Soil nitrification Niche differentiation
Human intervention in agriculture, such as farming and fertilization, could result in major changes of ammonia availability that shapes niche-specific occupied communities of soil ammonia oxidizers. The aim of this study was to investigate nitrification and active ammonia oxidizers in N-rich and N-limited soils, which were induced by different management practices. Here, we used DNA-based stable isotope probing (DNA-SIP) microcosm to decipher active ammonia-oxidizing archaea (AOA) and bacteria (AOB) phylotypes involved in ammonia oxidation in the cultivated and uncultivated soils. The results showed that net nitrification rate in the cultivated soil was significantly higher than in the uncultivated soil (6.19 and 1.40 mg N kg−1 dry soil d-1, respectively). Growth of soil AOB and AOA during incubation occurred in the cultivated soil, while in the uncultivated soil, only AOA significantly increased after 56 days. Combining DNA-SIP and sequencing results, we gathered evidence of ammonia oxidation by Nitrososphaera-like AOA, in addition to active AOB, within the classic Nitrosospira group in the cultivated soil. For the uncultivated soil, ammonia oxidation was driven by Nitrososphaera-like and Nitrosopumilus-like AOA, rather than AOB. Our results demonstrate the potential of ammonium concentration for shaping the communities of active AOA and AOB, which imply that ammonia-dependent “diversification” may be important strategies for niche differentiation of sympatric AOA/AOB under different field management conditions.
1. Introduction
AOB (Prosser and Nicol, 2012; He et al., 2012). Many factors could result in the niche differentiation of AOA and AOB in soils, including substrate concentration (Martens-Habbena et al., 2009), pH (GubryRangin et al., 2011), O2, temperature (Hatzenpichler, 2012) and mixotrophy (Prosser and Nicol, 2012). Among them, soil pH and precipitation were suggested to be principal drivers in shaping niche differentiation of microbes at large-scales (Bahram et al., 2018). However, it is unclear whether microbial niche differentiation can be driven by management of soils. Human intervention in agriculture, such as farming and fertilization, could result in major changes in soil physicochemical and biological properties (e.g. soil prokaryotic communities, pH, N availability) (Babin et al., 2019; Das et al., 2019). Numerous studies have suggested that ammonia concentration is an important factor shaping the niche differentiation between AOA and AOB (Verhamme et al., 2011; Sterngren et al., 2015; Daebeler et al., 2015). However, with few exceptions, most of these observations resulted from AOA and AOB amoA
Nitrification in agricultural soils can lead to nitrogen (N) losses following ammonia-based fertilizer application through conversion of ammonium into nitrate (Gruber and Galloway, 2008). Consequently, leaching of nitrate results in pollution of streams and groundwater, and denitrification losses increase nitrous oxide emissions (Ravishankara et al., 2009). Thus, ammonia oxidizers, crucial mediators of nitrification, are widely recognized as key players for sustainable N management of agricultural ecosystems and environmental stewardship. Ammonia-oxidizing bacteria (AOB), first isolated in the 19th century (Winogradsky, 1890), were thought to govern ammonia oxidation exclusively. In 2005, ammonia-oxidizing archaea (AOA) were proven to be a new group capable of oxidizing ammonia (Könneke et al., 2005; Francis et al., 2005). However, genomic studies of archaeal isolates suggested the unique biochemical and genetic traits from their bacterial counterparts, which supposes niche differentiation between AOA and ⁎
Corresponding author. E-mail address: [email protected] (X. Jiang).
https://doi.org/10.1016/j.still.2019.104433 Received 21 December 2018; Received in revised form 13 August 2019; Accepted 25 September 2019 0167-1987/ © 2019 Elsevier B.V. All rights reserved.
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2.2. Microcosm construction
gene quantification, and their abundances alone can not provide enough information about the relative importance of bacteria and archaea for NH3-oxidation because the genes might not have been expressed or enzymes were inactivated. Since not all microbes possessing amoA genes are active oxidizers for NH3-oxidation (Prosser and Nicol, 2008; Stahl and de la Torre, 2012), and most analyses of NH3-oxidizer communities usually include populations not actively contributing to nitrification, DNA-based stable isotope probing (DNA-SIP) techniques can instead be used to directly decipher active AOA and AOB involved with nitrification for in situ field conditions. This technique is thus a powerful means to link functional processes to taxonomic identities of active microorganisms in complex soil environments (Radajewski et al., 2003). We hypothesized that: (i) at the local scale, ammonia availability induced by management practices is the key driver of niche differentiation between AOA and AOB; (ii) the limited nitrogen caused by long-term non-cultivation will result in the predominance of AOA in nitrification for the uncultivated soil due to their higher affinity for substrates. Therefore, 13CO2-based DNA-SIP, quantitative PCR and nucleotide sequencing approach were used to investigate nitrification and active ammonia oxidizers under different agricultural managements, cultivated and uncultivated soils, which represent N-rich and N-limited environments, respectively.
Soil DNA-SIP microcosms were constructed to investigate the active soil ammonia oxidizers community as previously described (Jia and Conrad, 2009). Briefly, soil microcosms were established in 120 mL serum bottles tightly capped with black butyl stoppers which contained sieved soil (8.0 g dry weight soil) and 5% headspace (vol/vol) 13CO2 (99% atom enriched, Sigma-Aldrich Co., St Louis, MO, USA) or 12CO2 (generated by acidifying sodium carbonate). Besides 12CO2-control microcosms and 13CO2-labeled microcosms, an additional treatment included a 13CO2 + C2H2 (100 Pa) microcosm to assess whether 13CO2 assimilation by ammonia oxidizers was dependent on energy generated from ammonia oxidation that could be completely inhibited by C2H2 (Berg et al., 1982). Each treatment received 50 mg NH4+-N kg−1 dry soil weekly and microcosms were incubated at 60% soil water-holding capacity at 28 °C in the dark throughout the eight-week incubation. Each bottle was opened weekly to allow N addition and air exchange to maintain aerobic conditions. Water lost by evaporation was replenished with sterilized water, and 13CO2, 12CO2, and C2H2 in headspace were renewed immediately after bottle resealing. All treatments were performed in triplicate. Soil (2 g) from each bottle was destructively harvested at day 0 and day 56, and stored immediately in a −80 °C freezer (Meiling Biology & Medical, Anhui, China). Remaining soil was used for determination of ammonium and nitrate using a SKLAR continuous-flow analyzer (SKLAR San++, Netherland, 2003). Soil deoxyribonucleic acids were extracted from 0.5-g soil samples using a FastDNA spin kit for soil (MP Biomedicals, Cleveland, OH, USA) according to the manufacturer’s instructions. Extracted DNA was used for subsequent SIP and quantification of amoA genes.
2. Materials and methods 2.1. Study site description and soil collection The study site, Purple Soil Ecology Experimental Station of Southwest University, is located in the southwest of China (30° 26′ N, 106° 26′ E). The climate is subtropical with a mean annual temperature of 18.2 °C and a mean annual total rainfall of 1105 mm. Both soils developed from the purple mudstone and are classified as Eutric Regosol (FAO/UNESCO, 1988). The tested neutral paddy soils were collected from two long-term treatments established in 1990: the cultivated soil was in a rice (Oryza sativa L.) and oilseed rape (Brassica napus L.) cropping rotation, with conventional tillage as previously described (Li et al., 2015). The amount of fertilizer applied to the cultivated soil annually is urea 270 kg ha−1, superphosphate 500 kg ha−1, potassium chloride 150 kg ha-1. The fertilization strategy is that 60% of N, 100% of P and 50% of K are applied as base fertilizer before planting rape and rice, and then 40% of N and 50% of K as topdressing at the 5–6 leaf stage for rape or at the tillering stage for rice. The uncultivated field was left uncultivated and unfertilized, and the main vegetation of this fallow field was weeds (Gramineae). Both cultivated and uncultivated fields were under the same water management: fields were irrigated after rape harvest in May and kept flooded with a layer of water until rice maturation, and after rice harvest in the middle of August, fields were drained and remained dry for rape planting until the next May. The properties for the cultivated soil are: pH 7.3 (H2O), 16.5 g organic matter kg−1 dry soil, 1.37 g total N kg−1 dry soil, 10.7 mg NH4+-N kg−1 dry soil, and 13.6 mg NO3–N kg−1 dry soil. Properties for the uncultivated soil are: pH 7.3 (H2O), 20.5 g organic matter kg−1 dry soil, 0.921 g total N kg−1 dry soil, 4.08 mg NH4+-N kg−1 dry soil, and 9.50 mg NO3–N kg−1 dry soil. Each field contains three replicated plots of dimension 4 m × 5 m. Five soil cores were collected at a 0–20 cm depth from each plot using a 13 cm diameter soil auger in November 2017, and then mixed thoroughly to reduce heterogeneity and pooled to form composite samples. Three composite samples were collected from each sampling site generating three replicates for each treatment. Soil samples were transferred to the laboratory on ice and passed through a 2 mm sieve to remove roots and stones. Then each soil sample was divided into two parts: one was stored at 4 °C until SIP incubation, another was air dried to analyze the chemical properties with the same methods as we previously described (Wang et al., 2019).
2.3. SIP gradient fractionation The total DNA extract was subjected to isopycnic centrifugation to separate 13C-DNA from native 12C-DNA as previously described (Jia and Conrad, 2009). About 3.0 μg of total DNA extracts were thoroughly mixed with a CsCl solution to reach an initial CsCl buoyant density of 1.725 g mL−1 and placed in a 5.1 mL Beckman polyallomer tube. Suspensions were centrifuged in a Vti65.2 vertical rotor (Beckman Coulter Inc., Palo Alto, CA, USA) at 177,000 g for 44 h at 20 °C. Fifteen DNA fractions (∼380 μL) were obtained by displacing the gradient medium from the top of the ultracentrifuge tube with sterile water using a syringe pump (New Era Pump Systems Inc., Farmingdale, NY, USA). The buoyant density of each DNA fraction was measured indirectly using an AR200 digital hand-held refractometer (Reichert Inc., Buffalo, NY, USA). The fractionated DNA was precipitated and purified as Jia and Conrad (2009) described and dissolved in 30 μL TE buffer. 2.4. Quantification of amoA genes Real-time quantitative PCR (qPCR) was performed to estimate the abundance of AOA and AOB during the eight-week incubation, and the abundance of 13CO2-labeled amoA-carrying ammonia oxidizers in each CsCl fraction on a QuantStudio 6 Flex Real-Time PCR Systems (Applied Biosystems of Life Technologies, Singapore). Bacterial and archaeal amoA genes were amplified by primer pairs amoA-1 F/amoA-2R with amplicon length of 491 bp (Rotthauwe et al., 1997) and Arch-amoAF/ Arch-amoAR with amplicon length of 635 bp (Francis et al., 2005), respectively. The reaction was performed in a 20 μL mixture containing 10 μL SYBR Premix Ex Taq (TaKaRa, Dalian, China), 0.5 μL of each primer (10 μM), 0.4 μL of ROX Reference Dye II (50×) and 1 μL of DNA template (1–10 ng), and 7.6 μL sterile water. QPCR was performed in biological triplicates with three technical replicates. A plasmid DNA from one representative clone containing AOA or AOB amoA genes was used as the template to generate the standard curve, and the detailed steps and the subsequent amplification condition were the same as 2
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previously described (Jiang et al., 2015). The final amplification efficiencies were 95%–103% with R2 values of 0.993-0.999. Additionally, a melting curve analysis was used to check the specificity of the amplification products at the end of each qPCR run.
bacteria in both soils. 3.3.
13
CO2-labeled DNA in AOA and AOB
CO2-labeled DNA in the ‘heavy’ fractions was separated from native 12CO2-control DNA in the ‘light’ fractions for each SIP microcosm, and qPCR assessed amoA genes of each buoyant density of the SIP gradient fraction. AOA or AOB populations assimilating 13C into their genomes are regarded as being active ammonia oxidizers (Jia and Conrad, 2009; Wang et al., 2015b). Two peaks of archaeal and bacterial amoA gene abundance were observed in the gradient ranges of DNA fractions for 13CO2-labeled microcosms of cultivated soil, and considerable AOA and AOB were detected in the ‘heavy’ fraction (buoyant densities 1.725–1.735 g mL−1), implying archaeal and bacterial populations were labeled by 13CO2. For 12CO2-control microcosms, archaeal and bacterial amoA genes peaked in the native ‘light’ fractions with buoyant densities of 1.70 g mL-1 (Fig. 2a, c). These results demonstrated substantial 13CO2 assimilation by both AOA and AOB in the cultivated soil. However, in the uncultivated soil, archaeal amoA genes peaked in the ‘light’ fractions and the ‘heavy’ fractions for the control and labeled microcosms, respectively (Fig. 2b), while bacterial amoA genes peaked in the native ‘light’ fractions for both the control and labeled microcosms (Fig. 2d), indicating that only AOA was active during nitrification. 13
2.5. Sequencing and phylogenetic analysis MiSeq sequencing of archaeal and bacterial amoA genes in the fractionated DNA from 13CO2 labeled SIP microcosms were carried out on an Illumina MiSeq sequencer instrument (USA) at a Personal Biotechnology Co., Ltd. (Shanghai, China). The amplification of the fractionated DNA was performed using the primers amoA-1 F/amoA-2R and Arch-amoAF/Arch-amoAR with specific barcode sequences for AOA and AOB amoA genes, respectively. The PCR procedures and quality control of the obtained raw sequences were based on methods of Zhao et al. (2015). Reads with quality score lower than 20, ambiguous bases and improper primers were discarded. The resulting high-quality sequences were subgrouped to obtain operational taxonomic units (OTUs) at 97% sequence similarity levels. Phylogenetic trees of AOA and AOB amoA genes were constructed with MEGA 4.0 package through a neighbor-joining algorithm (Kimura 2-parameter distance model) using 1,000 bootstrap replicates (Tamura et al., 2007). 2.6. Statistical analyses One-way analysis of variance was performed to check the variance between different treatments followed by Fisher’s protected least significant difference test. Net nitrification rate (mg N kg−1 dry soil d−1) was calculated as the difference between the final and the initial nitrate concentrations divided by the incubation time (Robertson et al., 1999). All analyses were conducted using SPSS 13.0 package (SPSS, USA), and P < 0.05 was considered to be statistically significant.
3.4. Phylogenetic analysis of active nitrifying communities For cultivated soil, the obtained reads were binned into 10 and 8 OTUs for the archaeal and bacterial amoA genes, respectively. Phylogenetic analysis of amoA showed that 95.8% of active AOA fell within the group I.1b-Nitrososphaera lineage. OTU1 was the most abundant species which was closely related to Nitrososphaera sp. JG1 (JF748723) (with 96.9% identities), accounting for 80.7% of all active AOA reads. The remaining 4.2% was affiliated with group I.1aNitrosopumilus lineage (Fig. 3). As for active AOB in cultivated soil, 78.9% of reads were grouped into Nitrosospira cluster 3a.1, with genus Nitrosospira sp. Nsp2 (AJ298719) as the closest cultured relative. 21.0% of reads were affiliated with Nitrosospira cluster 3a.2, which was related to Nitrosospira multiformis (X90822), and the remaining small proportion was clustered into Nitrosospira cluster 9 (Fig. 4). For uncultivated soil, the high quality reads of archaeal amoA genes were binned into 10 OTUs and equally distributed into two clusters—50% reads were affiliated with the soil group I.1b-Nitrososphaera lineage and the other 50% fell into the marine group I.1a-Nitrosopumilus lineage. The most abundant OTU was closely related to an uncultured archaeon in sediments (KJ005105) in group I.1a-Nitrosopumilus lineage, accounting for 37.7% of the total amoA gene reads (Fig. 5).
3. Results 3.1. Nitrification activity For both soils, significant accumulation of ammonium was observed at day 56 in the presence of C2H2 compared to day 0 (P < 0.05). In the absence of C2H2, ammonium did not differ between Day 56 and Day 0 for cultivated soil (P > 0.05), but ammonium increased significantly at Day 56 for uncultivated soil (P < 0.05) (Fig. 1a). For cultivated soil, NO3−-N concentrations averaged 13.6 mg NO3−-N kg-1 at day 0, and increased significantly to 359 and 361 mg NO3−-N kg-1 at day 56 for no C2H2 treatments (Fig. 1c). For uncultivated soil without C2H2, NO3−-N increased by 78.4 mg kg-1 after 56 days, showing a lower net nitrification rate (1.40 mg N kg-1 dry soil d-1) compared to the same treatment with cultivated soil (6.19 mg N kg-1 dry soil d-1). Soil nitrate production was negated with C2H2 addition for both cultivated and uncultivated soils (Fig. 1c), indicating that microbial NH3-oxidation was inhibited by C2H2 during the incubation. No significant nitrification was detected over the course of the 56-day incubation in the 13CO2+C2H2 soil microcosms.
4. Discussion The DNA-SIP technique provided strong evidence that niche differentiation of sympatric ammonia oxidizers occurred in these two neutral soils, as both archaea and bacteria played roles in autotrophic nitrification in cultivated soil, but only AOA were active in autotrophic nitrification for uncultivated soil. Phylogenetic analysis clearly showed that the dominant active archaea and bacteria in cultivated soils were affiliated with Nitrososphaera subcluster 1.1 within the soil group 1.1b lineage and Nitrosospira cluster 3a.1, respectively, however the dominant ammonia oxidizers in uncultivated soil was only AOA in Nitrososphaera subcluster 1.1 within the soil group 1.1b lineage and Nitrosopumilus cluster within the marine group I.1a lineage. For uncultivated soil, there were concurrent low numbers of AOB amoA genes at the beginning and the end of incubation (Fig.1d), indicating that AOB could not account for the observed nitrification rate. This result was also supported by the SIP data suggesting that no bacterial amoA genes were labeled by 13CO2 (Fig. 2d). However, AOA
3.2. AOA and AOB abundance For the cultivated soil, in the absence of C2H2, amoA gene abundance increased from 3.60 × 107 g−1 dry soil at Day 0 to 8.80 × 107 g−1 dry soil at Day 56 and 2.85 × 106 g−1 dry soil at Day 0 to 1.18 × 107 g−1 dry soil at Day 56, corresponding to 1.45- and 3.11 fold increases for AOA and AOB after incubation (Fig. 1b, d). For the uncultivated soil, AOA increased significantly from 2.85 × 106 g−1 dry soil at Day 0 to 1.17 × 107 g−1 dry soil at Day 56 in the absence of C2H2 (P < 0.05), while AOB showed no significant change from Day 0 (1.50×105 g−1 dry soil) to Day 56 (1.46 × 105 g−1 dry soil) (P > 0.05) (Fig. 1b, d). The addition of C2H2 inhibited growth of archaea and 3
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Fig. 1. The concentrations of soil ammonium (a) and nitrate (c), and the abundances of archaeal (b) and bacterial (d) amoA genes in cultivated and uncultivated soil microcosms over a 56-days incubation at 28 °C. The error bars indicate standard errors of the mean of the triplicate microcosms. The different letters above the columns indicate a significant difference (P < 0.05).
Fig. 2. Quantitative distribution of the relative abundances of archaeal (a, b) and bacterial (c, d) amoA genes across the entire buoyant density gradient of the DNA fractions retrieved from 12CO2 and 13CO2 56-day DNA-SIP microcosms of cultivated and uncultivated soil, respectively. Error bars represent standard errors from three replicates.
was 183 and was significantly higher than cultivated soil (12.6). Similarly, in a survey from 12 pristine and agricultural soils of three climatic zones, the ratio of AOA/AOB amoA gene copies ranged from 1.5 to 232, and the highest ratio occurred for non-fertilized soil (Leininger et al., 2006). This result is not surprising because the uncultivated neutral purple soil has not received cultivation or N fertilization for about 30 years, which created a N-limited environment
abundance was significantly higher than its bacterial counterpart and increased with nitrification (Fig. 1b). We can therefore assume that AOA plays a role in ammonia oxidization in uncultivated soil, which is in line with SIP results (Fig. 2b). These results are consistent with previous studies attributing ammonia oxidization to AOA in nutrientlimited, non-fertilized soils (Daebeler et al., 2014, 2015; Sterngren et al., 2015; Shi et al., 2018). The ratio of AOA/AOB in uncultivated soil 4
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Fig. 3. Phylogenetic analysis of the archaeal amoA genes from the 13CO2-labeled microcosms at the end of incubation in the neutral cultivated soil. The designation AOA-OTU1 (80.7%) indicates that OTU1 account for 80.7% of the AOA amoA reads from the labeled microcosms. One representative sequence from each OTU was selected using the mothur software for the tree construction. Archaeal amoA subclusters were classified according to Pester et al. (2012).
Nguyen et al., 2019; Kong et al., 2019). Soil ammonia concentration is regarded as one of the most critical factors shaping the sympatric differentiation of archaea and bacteria communities (Martens-Habbena et al., 2009; Prosser and Nicol, 2012). Our results provide compelling evidence for substrate-induced niche differentiation of sympatric AOA/AOB for these two soils. For the cultivated soil (N-rich condition), the dominant active AOA was affiliated with Nitrososphaera subcluster 1.1 and the major active AOB were affiliated with Nitrosospira cluster 3a.1. For the uncultivated soil (N-limited condition), only AOA was labeled and the dominant active AOA was affiliated with Nitrososphaera subcluster 1.1 and Nitrosopumilus cluster with a closely cultured relative of genus Nitrosopumilus maritimus which possesses an extremely high substrate affinity. These results are consistent with previous findings that AOA is the dominant NH3-oxidizer in oligotrophic environments, and possibly may be the only driver of NH3-oxidation in soils with deficient available nitrogen (Daebeler et al., 2014, 2015; Sterngren et al., 2015). On the contrary, AOB appears to dominate NH3 oxidation after addition of N fertilizer at high concentrations (Jia and Conrad, 2009; Zhong et al., 2016; Xiang et al., 2017). It is worth noting that in the present study, only 4.2% of active AOA belong to the group I.1a-Nitrosopumilus lineage in cultivated soil, but in the uncultivated soil, contributions were up to 50% (Figs. 3 and 5), indicating that substrate can shift the predominance of phylotypes within AOA or AOB, not only between AOA and AOB. Nitrosopumilus-
favoring AOA rather than AOB owing to a much higher NH3 affinity of archaea (Prosser and Nicol, 2012; Shi et al., 2018). The relative importance of archaea and bacteria in ammonia oxidation is influenced by many environmental factors, including pH (Gubry-Rangin et al., 2011), ammonia concentration (Prosser and Nicol, 2012), temperature (Avrahami and Conrad, 2003), water content (Hu et al., 2015), and dissolved oxygen levels (Santoro et al., 2008). Numerous studies have shown that AOA is an active player in acidic soils, while AOB is the driver of nitrification in neutral and alkaline soils (Taylor et al., 2012; Jiang et al., 2015; Shi et al., 2018). However, the diversity and abundance of AOA increases with rising soil pH (Gubry-Rangin et al., 2011), suggesting that some phylotypes of archaea may be active in NH3-oxidation in neutral or alkaline soils. In the cultivated soil, both AOA and AOB amoA gene abundances significantly increased and were paralleled by an increase in nitrate concentrations (347 mg N kg−1 d.w.s) (Fig. 1), indicating that both AOA and AOB contributed to nitrification. This result was subsequently confirmed by 13 CO2-labeled NH3-oxidizers, demonstrating that substantial 13CO2 was assimilated into archaea and bacteria (Fig. 2). The phylogenetic analysis further revealed that the dominant active AOA and AOB fell into Nitrososphaera and Nitrosospira clusters, respectively, which provided strong evidence that both Nitrososphaera-like AOA and Nitrosospira-like AOB are active regulators of nitrification in this neutral soil (Figs. 3 and 4). This result was similar to those found in previous studies showing a cooperation of AOA and AOB in neutral soils (Wang et al., 2015a;
Fig. 4. Phylogenetic analysis of the bacterial amoA genes from the 13CO2-labeled microcosms at the end of incubation in the neutral cultivated soil. The designation AOB-OTU2 (18.2%) indicates that OTU1 account for 18.2% of the AOA amoA reads from the labeled microcosms. One representative sequence from each OTU was selected using the mothur software for the tree construction. Bacterial amoA clusters were classified according to Avrahami and Conrad (2003).
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Fig. 5. Phylogenetic analysis of the archaeal amoA genes from the 13CO2-labeled microcosms at the end of incubation in the neutral uncultivated soil. The designation AOA-OTU1 (13.5%) indicates that OTU1 account for 13.5% of the AOA amoA reads from the labeled microcosms. One representative sequence from each OTU was selected using the mothur software for the tree construction. Archaeal amoA subclusters were classified according to Pester et al. (2012).
respectively). The qPCR of amoA genes showed higher AOA abundance than AOB for both soils, with AOA/AOB for the cultivated and uncultivated soils averaging 12.6 and 183, respectively. Obvious growth of soil AOB and AOA occurred during incubation in the cultivated soil, while in the uncultivated soil, only AOA increased significantly. Our results provide robust evidence for niche differentiation of active AOA and AOB phylotypes in the cultivated and uncultivated soils. The phylogenetic analysis reveals that Nitrososphaera-like AOA and Nitrosospira-like AOB drive ammonia oxidization in concert in the Nrich cultivated soil, but in the N-limited uncultivated soil, the active drivers are soil group I.1b Nitrososphaera-like and marine group I.1a Nitrosopumilus-like AOA lineages. These results suggest that soil ammonia oxidization is carried out by diverse physiologically versatile ammonia-oxidizing phylotypes in soils with different management, indicating that ammonium-dependent “diversification” may be an important strategy for niche differentiation of sympatric AOA/AOB under variable field conditions.
like AOA are often found in aquatic environments, such as the open ocean, but are less common in soils (Offre et al., 2009; Pester et al., 2012). The seasonal flooding of the paddy soil causes oxygen stress and favors Nitrosopumilus lineage archaea which has a higher affinity for O2 compared to Nitrososphaera lineage (Martens-Habbena et al., 2009; Jung et al., 2011). Interestingly, many previous DNA SIP studies of fertilized paddy soils found that active AOA was affiliated with group I.1b-Nitrososphaera lineage, rather than group I.1a-Nitrosopumilus lineage (Wang et al., 2015a; Zhao et al., 2015). However, some studies indeed directly link group I.1a-Nitrosopumilus lineage AOA to active ammonia oxidization in an ammonia-limited volcanic grassland soil and an agricultural soil in Scotland with low ammonia concentration (Daebeler et al., 2014; Zhang et al., 2010). These results indicated that substrate limitation selected active I.1a-Nitrosopumilus lineage. Some phylotypes such as Nitrosopumilus lineage could successfully compete with its sympatric Nitrososphaera lineage when substrates become more deficient, suggesting a higher ammonia affinity of archaea in Nitrosopumilus cluster than in Nitrososphaera cluster just like their isolated cultured strains (Martens-Habbena et al., 2009; Jung et al., 2011). In this study, although 50 mg N/kg d.s.w/week ammonium sulfate was added to soil, no obvious growth of AOB was found during incubation, but AOA increased significantly. Given that AOB has a faster growth rate than AOA (Jiang and Bakken, 1999; Tourna et al., 2011), we can conclude that AOA was already active under the field conditions. The oligotrophic condition led to the inactivation of AOB in ammonia oxidization and selectively prospered the Nitrosopumilus cluster AOA, implying that AOA/AOB may develop their “diversity” to adapt to different environments based upon long-term evolutionary shifts (Lennon and Jones, 2011). The diversity of archaeal or bacterial lineages may be a result of evolution based upon specific adaptations to soil pH, ammonia availability or oxygen content, and thus show niche specialization. The recent discovery of “comammox”, which occurs when one nitrifying microorganism carries out the complete oxidation of ammonia to nitrate (van Kessel et al., 2015v; Daims et al., 2015), contributed to a growing appreciation that nitrifiers are more versatile than formerly believed (Santoro, 2016). Understanding the niche differentiation of AOB and AOA to substrates can thus improve our ability to manage N more efficiently for agriculture.
Acknowledgments We are grateful to the editor and three anonymous referees for their careful reviews and constructive suggestions. This research was financially supported by the National Natural Science Foundation of China (41671232), the National Key Research and Development Program of China (2016YFD0300901), the Graduate Research and Innovation Program of Chongqing (CYB19105). Zhihui Wang thanks the China Scholarship Council (CSC) for her scholarships. References Avrahami, S., Conrad, R., 2003. Patterns of community change among ammonia oxidizers in meadow soils upon long-term incubation at different temperatures. Appl. Environ. Microbiol. 69, 6152–6164. Babin, D., Deubel, A., Jacquiod, S., Sørensen, S.J., Geistlinger, J., Grosch, R., Smalla, K., 2019. Impact of long-term agricultural management practices on soil prokaryotic communities. Soil Biol. Biochem. 129, 17–28. Bahram, M., Hildebrand, F., Forslund, S.K., Anderson, J.L., Soudzilovskaia, N.A., Bodegom, P.M., Bengtsson-Palme, J., Anslan, S., Coelho, L.P., Harend, H., HuertaCepas, J., Medema, M.H., Maltz, M.R., Mundra, S., Olsson, P.A., Pent, M., Põlme, S., Sunagawa, S., Ryberg, M., Tedersoo, L., Bork, P., 2018. Structure and function of the global topsoil microbiome. Nature 560, 233–237. Berg, P., Klemedtsson, L., Rosswall, T., 1982. Inhibitory effect of low partial pressures of acetylene on nitrification. Soil Biol. Biochem. 14, 301–303. Daebeler, A., Bodelier, P.L., Yan, Z., Hefting, M.M., Jia, Z., Laanbroek, H.J., 2014. Interactions between Thaumarchaea, Nitrospira and methanotrophs modulate autotrophic nitrification in volcanic grassland soil. ISME J. 8, 2397–2410. Daebeler, A., Bodelier, P.L.E., Hefting, M.M., Laanbroek, H.J., 2015. Ammonia-limited conditions cause of Thaumarchaeal dominance in volcanic grassland soil. FEMS
5. Conclusions Net nitrification rate of the cultivated soil was significantly higher than the uncultivated soil (6.19 and 1.40 mg N kg−1 dry soil d−1, 6
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