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Shifts in Nitrobacter- and Nitrospira-like nitrite-oxidizing bacterial communities under long-term fertilization practices
Soil Biology and Biochemistry 124 (2018) 118–125
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Shifts in Nitrobacter- and Nitrospira-like nitrite-oxidizing bacterial communities under long-term fertilization practices
T
Shun Hana, Luyang Zenga,b, Xuesong Luoa,b, Xiang Xionga,b, Shilin Wenc, Boren Wangc, Wenli Chena,∗, Qiaoyun Huanga,b,∗∗ a
State Key Laboratory of Agricultural Microbiology, Huazhong Agricultural University, Wuhan, 430070, China Key Laboratory of Arable Land Conservation (Middle and Lower Reaches of Yangtze River), Ministry of Agriculture, College of Resources and Environment, Huazhong Agricultural University, Wuhan, 430070, China c Hengyang Red Soil Experimental Station, Chinese Academy of Agricultural Sciences, Hengyang, 421001, China b
A R T I C LE I N FO
A B S T R A C T
Keywords: Fertilization treatment Nitrite-oxidizing bacteria (NOB) Nitrospira-like NOB Nitrobacter-like NOB Potential nitrite oxidation activity (PNO)
Nitrite-oxidizing bacteria (NOB) are key players in the second step of nitrification, which is an important process in the soil nitrogen (N) cycle. However, the ecology of nitrite oxidizers and their response to disturbances such as long-term fertilization practices are scarcely known in agricultural ecosystems. We used samples from a Red soil subject to a long-term chemical and organic fertilization experiment, including control without fertilizer (CK), swine manure (M), chemical fertilization (NPK), and chemical/manure combined fertilization (MNPK) treatment, to explore how agricultural practices impact the community structure, abundance, and potential activity of nitrite oxidizers (PNO). The abundance of Nitrobacter was significantly increased in the M and MNPK plots, whereas the abundance of Nitrospira was significantly reduced in the M and NPK treatment plots and less inhibited in the MNPK treatment. The PNO showed a similar trend to that for Nitrobacter abundance. The diversity of Nitrobacter increased in the M-treated plots, while that of Nitrospira increased in the M and MNPK plots and decreased in the NPK plots. Non-metric multidimensional scaling (NMDS) revealed that the Nitrobacter- and Nitrospira-like NOB community was shift in these four fertilization treatments. Redundancy analysis showed that pH+SOC (soil organic carbon) and pH+TN (total nitrogen) significantly explained the variation in the composition of Nitrobacter and Nitrospira, respectively. In addition, the Nitrospira/Nitrobacter abundance ratio and community structure of Nitrobacter- and Nitrospira-like NOB are responsible for the changes of soil PNO. Collectively, these data suggest that the nitrite-oxidation process in the red soil is possibly controlled by both Nitrospira and Nitrobacter-like NOB, which were shaped by pH+TN and pH+SOC, respectively.
1. Introduction Nitrification, the microbiological oxidation of ammonia (NH4+) to nitrite (NO2−) and subsequently to nitrate (NO3−), influences the fate of nitrogen (N) in terrestrial systems. The first limiting step of nitrification, NH4+ oxidation to NO2−, is mediated by ammonia-oxidizing bacteria (AOB) of the β- and γ-Proteobacteria, ammonia-oxidizing archaea (AOA) of the Thaumarchaeota (Kowalchuk and Stephen, 2001; Leininger et al., 2006; Schleper and Nicol, 2010; Norton and Stark, 2011). The second step of nitrification, NO2− oxidation to NO3−, is catalyzed by nitrite-oxidizing bacteria (NOB), which play an important role in the biogeochemical N cycle in many terrestrial ecosystems such as soils (Prosser, 1989; Daims et al., 2015). NOB are broadly distributed among the α-, β-, γ-, and δ-Proteobacteria as well as the Nitrospira classs
∗
(Gould and Lees, 1960; Teske et al., 1994; Alawi et al., 2007; Sorokin et al., 2012). Comammox Nitrospira complete oxidation of ammonium to nitrate for the nitrification (Daims et al., 2015; van Kessel et al., 2015). In recent years, many studies have focused on the ecology of AOA and AOB in a broad range of soil environments (Kowalchuk and Stephen, 2001; Webster et al., 2005; Chu et al., 2007; Chen et al., 2008; Shen et al., 2012; Bertagnolli et al., 2016; Zhang et al., 2017). Ammonia oxidation is often assumed to be the rate-limiting step of nitrification as nitrite generally does not accumulate in the environment. In fact, nitrite oxidation can also become the limiting step for nitrification in disturbed soil systems (Gelfand and Yakir, 2008; Roux-Michollet et al., 2008). In view of the importance of nitrite oxidizers, it is urgently needed to decipher the ecological response of nitrite oxidizer community to any
Corresponding author. State Key Laboratory of Agricultural Microbiology, Huazhong Agricultural University, Wuhan 430070, China. Corresponding author. State Key Laboratory of Agricultural Microbiology, Huazhong Agricultural University, Wuhan 430070, China. E-mail addresses: [email protected] (W. Chen), [email protected] (Q. Huang).
∗∗
https://doi.org/10.1016/j.soilbio.2018.05.033 Received 22 November 2017; Received in revised form 30 May 2018; Accepted 31 May 2018 0038-0717/ © 2018 Elsevier Ltd. All rights reserved.
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2016. Six soil cores (approximately 5 cm in diameter) were taken from each plot and mixed to form one composite sample. Samples were placed in a sterile plastic bag for transport to the laboratory within 24 h after collection. Each soil sample was divided into three portions. One portion was used for DNA extraction and stored at −80 °C, another was stored at 4 °C for measuring PNO, and the third was air-dried at room temperature for analysis of soil chemical properties.
type of environmental disturbances, such as different fertilization regimes in agricultural soils. Nitrobacter-like NOB are considered to be the key functional players within the NOB community, exhibiting high activity levels in tillage/ no-tillage agricultural systems with high N availability (Attard et al., 2010). In addition, soil potential nitrite oxidation activity (PNO) is strongly positively correlated with the abundance of Nitrobacter-like NOB as well as weakly negatively correlated with the abundance of Nitrospira-like NOB. Possible changes were also observed for the abundance and community of Nitrobacter and Nitrospira in agricultural soils in response to tillage practices. In contrast, fertilization causes rapid shifts in the structure of the Nitrobacter-like NOB community, which dominates nitrification in forest soils (Wertz et al., 2011). The application of manure in a Merzenhausen agricultural soil was found to have decreased the diversity of Nitrobacter community in the rhizosphere (Ollivier et al., 2013). Nitrite oxidation in surface agricultural soils may be predominantly driven by Nitrospira spp. (Ke et al., 2013). Han et al. (2017) showed that Nitrospira might be more responsive than Nitrobacter in soils under a rapeseed-rice rotation. In addition, the Nitrospira-like NOB community was found to be significantly shaped by soil pH, moisture, and NH4+ content, whereas the Nitrobacter-like NOB community was not. In acidic forest soils, long-term fertilization increased AOB and Nitrobacter-like NOB abundances but did not influence AOA and Nitrospira-like NOB abundances (Wertz et al., 2011). However, the responses of the community structure and diversity of NOB to long-term fertilization regimes and their links with nitrite oxidation activities in agricultural soils are still not well understood. The ways in which NOB populations respond to changes in soil pH, specifically the acidification of terrestrial environments, have rarely been investigated. In this study, samples from a Red soil in the Qiyang Long Term Soil Experimental Station, Hunan Province, China was used to investigate the ecological effects of long-term fertilization practices on the Nitrobacter- and Nitrospira-like bacterial community and their potential activity. Quantitative PCR (qPCR) and high-throughput sequencing of marker genes were performed to investigate the abundance, community diversity, and population composition of Nitrobacter- and Nitrospira-like NOB in the soil. We hypothesized that (1) chemical fertilization may have a negative impact on the activity and diversity of NOB; (2) manure would have a positive effect on NOB community structure, as it provides both inorganic and organic nutrients for NOB and does not lead to acidification; (3) the combined use of manure and chemical fertilizers would represent a balanced fertilization treatment, inhibiting soil acidification and balancing the negative effect of chemical fertilizers on NOB activity and diversity.
2.2. Soil chemical analytical procedures Soil total carbon (TC) and total nitrogen (TN) contents were analyzed using a Vario Max element analyzer (Elementar Vario PYRO cube and Isoprime100, Germany). Soil exchangeable ammonium (NH4+-N) and nitrate (NO3−-N) contents were determined on a FIAstar 5000 Analyzer (Foss Tecator, Denmark) after extraction from fresh soil with 2 M KCl (w/v, 1:5). Soil pH was determined at a soil/water ratio of 1:2.5, and soil organic content (SOC) was determined by the K2Cr2O7 oxidation method. 2.3. Assays for determination of PNO PNO was determined using the method described by Wertz et al. (2007), modified from Smorczewski and Schmidt (1991). Briefly, samples of fresh soil (5 g equivalent dry mass) were incubated with 50 ml of a solution of NaNO2 (5 μg of N-NO2 g−1 dry soil) dissolved in autoclaved deionized water for 30 h with gentle shaking (180 rpm) at 28 °C. During incubation, 2.0-ml aliquots of the suspensions were sampled at 0, 4, 8, 12, 24, and 30 h and centrifuged (5000 rpm for 5 min). The supernatants were filtered (0.2-μm pore size) and analyzed for nitrite (N-NO2-) concentration on a spectrophotometer (L6, Shanghai, China) at 520 nm using Griess reagent at room temperature (Smorczewski and Schmidt, 1991). Levels of nitrite generally decreased linearly with time throughout the first 12 h until nitrite was depleted. Rates were calculated from the linear decrease and taken as the PNO. In this study, according to the literature, we used nitrite solution to measure the potential value directly without considering the ammonia oxidation process (Smorczewski and Schmidt, 1991; Wertz et al., 2007; Attard et al., 2010). The nitrite produced from ammonia oxidation process during incubation was not taken into account. 2.4. Soil DNA extraction DNA was extracted from 0.5 g soil using Lysing MatrixB tubes (Bio101) as described previously by Griffiths et al. (2000). Humic acids, a PCR inhibitor, were removed from the soil DNA using DNA-EZ Reagents M Humic Acid-Be-Gone B (Sangon Biotech, Shanghai, China). Purification was performed according to the manufacturer's instructions. The quality and concentration of DNA were determined using a NanoDrop spectrophotometer (NanoDrop Technologies, Wilmington, DE, USA).
2. Material and methods 2.1. Experimental site and sampling The experimental site was located at the Qiyang Red Soil Experimental Station (26°45′N, 111°52′E), Hunan Province, China. This site represents a typical agricultural region of subtropical China. It has a subtropical monsoon climate with an annual rainfall of 1300 mm and annual average temperature of 18 °C. A long-term fertilizer experiment was established in 1990 with a winter wheat (Triticum aestivum L.) and summer maize (Zea mays L.) rotation system, including three replicates of four treatments in a randomized plot design: control without fertilizers (CK); swine manure (M); chemical fertilization (nitrogen, phosphate, and potassium fertilizers, NPK); and chemical/organic combined fertilization (nitrogen, phosphate, potassium, and swine manure fertilizers, MNPK). The NPK-treated soil was severely acidified. The nitrogen fertilizer was applied as urea or swine manure at 300 kg N ha−1, phosphate (P) as a single application of superphosphate [Ca(H2PO4)2] at 53 kg P ha−1, and potassium (K) as potassium chloride (KCl) at 100 kg K ha−1. Soil samples were collected at a depth of 0–20 cm in November
2.5. Measurement of Nitrobacter- and Nitrospira-like NOB abundance by qPCR QPCR assays were conducted using an ABI7500 FAST Real-time PCR system with the nxrA primers F1norA and R2norA (Attard et al., 2010) for Nitrobacter-like NOB and the nxrB gene primers nxrB169f and nxrB638r (Pester et al., 2014) for Nitrospira-like NOB. The 20-μl PCR reaction mixtures contained 10 μl SYBR Premix Ex Taq II (2×) (Takara, Bio Inc., Shiga, Japan), 1.0 μl of a 10 mM solution of each primer, 6.0 μl DEPC-treated water, and 2.0 μl soil extract (diluted to 5 ng/μl) or 2.0 μl standard plasmid. A standard curve was generated using ten-fold serial dilutions of a plasmid containing a copy of the target gene. The following program was used for the nxrA gene: 95 °C for 10 min followed by 40 cycles of denaturation at 95 °C for 30 s, annealing at 55 °C for 30 s, and elongation at 72 °C for 30 s. For the nxrB gene, the program 119
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Table 1 Soil physicochemical properties of the studied soil samples. Treatments
pH (1:2.5)
CK M NPK MNPK
6.87 6.52 4.83 5.90
± ± ± ±
TC (%) 0.02a 0.11a 0.05c 0.29b
0.64 1.56 1.02 1.36
± ± ± ±
0.05d 0.05a 0.06c 0.05b
0.11 0.21 0.15 0.19
NO3− (mg/kg)
NH4+ (mg/kg)
TN (%) ± ± ± ±
0.004d 0.003a 0.008c 0.006b
9.03 7.62 9.95 8.06
± ± ± ±
2.03a 0.14a 0.33a 0.24a
3.43 19.8 15.9 17.4
± ± ± ±
SOC (g/kg) 0.51b 2.18a 4.07a 5.12a
17.3 33.3 23.1 28.2
± ± ± ±
0.43d 1.36a 0.81c 1.41b
Numbers in the same column followed by different letters are significantly different (Tukey's test, P < 0.05, n = 3).
the correlation between the rank similarity matrices for PNO and NOB community structure (Attard et al., 2010). Multivariate regression tree (MRT) analysis (De'ath, 2002) was performed using the R package mvpart (with default parameters) to relate the relative abundances of lineages to the site characteristics. A general linear model in combination with quasi-likelihood shrunken dispersion estimates was implemented in the R package edgeR 3.6.8 (Robinson et al., 2010; Lund et al., 2012) to identify which OTUs were stimulated or repressed by soil pH. Multiple sequence alignments of genes were performed using Muscle based alignments of the translated amino acid sequences. Phylogenetic trees were constructed by neighbor-joining using the selected sequences (relative abundance > 1%) with 1000 bootstrap replicates of MEGA 6. The reliability of the tree topology was evaluated by 1000 replicates of bootstrap resampling.
consisted of: 95 °C for 10 min followed by 40 cycles of denaturation at 95 °C for 30 s, annealing at 56.2 °C for 45 s, and elongation at 72 °C for 45 s. A negative control was included in each run using water instead of soil DNA extract. Melting curve and gel electrophoresis analyses were carried out to confirm amplification specificity. The dilution of DNA extracts is a commonly used method to reduce levels of inhibition (Wang et al., 2017), and several DNA samples were diluted to 5 ng/μl based on a preliminary experiment to determine the optimal dilution concentration. 2.6. High-throughput sequencing and bioinformatics analysis The sequences of the PCR products of the nxrA and nxrB genes were obtained using the Illumina MiSeq PE300 platform and the primers described above. To distinguish amplicons originating from different soil samples, barcode oligonucleotides 7 bp in length were ligated to the ends of the purified PCR products. The PCR products were checked by agarose gel electrophoresis and cleaned by Agencourt AMPure XP (Beckman Coulter, Inc. S. Kraemer Boulevard Brea, CA, USA). The DNA concentration of the purified PCR product was measured using the Quant-iT™ PicoGreen dsDNA BR Assay Kit (Invitrogen) and a microplate reader (BioTek FLx800) according to the manufacturer's protocol. Next, an equal amount of PCR product for each sample was combined in a single tube to be run on the sequencing platform. Raw reads were filtered based on the barcodes using QIIME (Caporaso et al., 2010), and low-quality sequences (quality score < 20) and ambiguous bases were removed using USEARCH (http://www.drive5.com/usearch/). The remaining sequences were further screened for frame shifts using FrameBot from the RDP FunGene Pipeline (http://fungene.cme.msu.edu/ FunGenePipeline). The remaining quality-screened sequences were clustered into operational taxonomic units (OTUs) using UCLUST (Edgar, 2010) based on a 95% nucleic acid sequence identity cutoff. In addition, OTUs with abundances less than 0.001% of the total sample sequence were removed (Bokulich et al., 2013), and then the OTU table was used for subsequent analysis.
3. Results 3.1. Soil physicochemical properties The physicochemical properties of the collected soil samples are shown in Table 1. Significant differences (P < 0.05) in soil pH were observed among all treatments. The CK and M plots had higher pH values (Table 1), while the lowest pH values (4.83) were recorded from the NPK samples. There were also significant differences (P < 0.05) in soil TC, TN, and SOC contents among all treatments, with the highest levels in the M treatment, followed by those in the MNPK, NPK, and CK treatments. The CK soil NO3− content was the lowest among all treatments. Ammonia content was relatively stable among the different fertilization regimes. 3.2. Soil PNO Soil PNO values ranged from 0.07 to 0.37 μg NO2−-N kg−1 dry soil h (Fig. 1). PNO was significantly (P < 0.01) elevated by about 3.9-, 3.7-, and 4.6-fold under the M, NPK, and MNPK treatments, −1
2.7. Statistical analysis One-way analysis of variance (ANOVA) was used to analyze the effects of fertilization treatments on the abundances of Nitrobacter- and Nitrospira-like NOB, PNO, and soil variables and automatic liner modeling was performed at the confidence level of 95% by using SPSS 19.0 statistical software (IBM Co., Armonk, NY USA). Spearman's correlation was used to determine whether the correlation between the abundances of Nitrobacter- and Nitrospira-like NOB, PNO, and each soil variable was significant. The program Mothur (http://www.mothur.org/wiki/ MainPage) was used to determine α-diversity indices. Redundancy analysis (RDA) was carried out using the Canoco 4.5. Monte Carlo permutation test with 999 unrestricted permutation to determine the extent of the environmental parameter(s) that were able to explain the variation in the nitrifier community. Non-metric multidimensional scaling (NMDS) ordination plots were used to identify the differences in NOB community composition using the R package vegan. Spearman correlation coefficient and associated p significance level (obtained by a permutation test using 999 permutations) were computed to quantify
Fig. 1. Soil potential nitrite oxidation activity (PNO) under four fertilization treatments. Error bars represent standard error and are accompanied by lowercase letters indicating significant differences according to Tukey's test (P < 0.05). 120
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Nitrobacter population size was detected in the MNPK treatment, followed by the M, CK, and NPK treatments in a significantly (P < 0.05) decreasing order (Fig. 2A). In contrast, the Nitrospira population sizes were significantly (P < 0.05) higher in the CK treatment than in the other fertilization treatments (Fig. 2B). Fertilization exerted considerable (P < 0.01) effects on the gene copy numbers of both Nitrobacter and Nitrospira. The ratio of nxrB/nxrA gene copy numbers ranged from 7.6 to 168.2, with the highest ratio observed for CK treatment, followed by NPK, MNPK, and M treatments, indicating a significant (P < 0.05) negative impact of fertilization on the nxrB/nxrA ratio. Spearman's rank correlation showed that soil TC and TN had significant (P < 0.05) positive and negative impacts on the abundances of nxrA and nxrB, respectively (Table S1). In addition, PNO was found to be highly positively correlated with Nitrobacter-like nxrA copy number (ρ = 0.664, P < 0.05) except for NPK treatment, but was not correlated with Nitrospira-like nxrB copy number (ρ = −0.559, P = 0.059). However, PNO were strongly and negatively (ρ = −0.923, P < 0.01) correlated with the nxrB/nxrA abundance ratio.
3.4. Nitrobacter- and Nitrospira-like NOB community structures The community structures of Nitrobacter- and Nitrospira-like NOB were characterized by investigating the nxrA and nxrB genes using high-throughput amplicon sequencing. Sequencing yielded a total of 678,665 high-quality nxrA and 370,887 nxrB gene sequences, corresponding to 2878 and 3243 unique OTUs classified as Nitrobacter- and Nitrospira-like NOB, respectively. A resampling procedure was employed at a depth of 28193 and 16802 sequences per sample to calculate the alpha diversity index for Nitrobacter and Nitrospira, respectively. Table 2 shows the diversity properties of Nitrobacter- and Nitrospiralike NOB. For Nitrobacter-like NOB, there were no significant differences in the ACE or Chao1 indices among the treatments. The highest Shannon and Simpson values were detected in the M plots, while those for the other treatments were similar. For Nitrospira-like NOB, the Shannon and Simpson values were highest in the MNPK and M treatments, followed by the CK and then NPK treatments (Table 2). Analysis of similarities (ANOSIM) of the Nitrobacter- (R = 0.58, P = 0.001) and Nitrospira-like NOB (R = 0.72, P = 0.001) community data indicated that fertilization regime was a significant driver of the NOB community. NMDS analysis reveals that the Nitrobacter- and Nitrospira-like NOB communities clustered separately along NMDS1 and NMDS2 according to fertilization treatments (Fig. 3A and B). A significant correlation was observed between PNO levels and the Nitrobacter- (ρ = 0.47, P = 0.005) and Nitrospira-like (ρ = 0.41, P = 0.015) NOB community structure. The hierarchical clustering of microbial communities also demonstrated that the Nitrobacter- and Nitrospira-like NOB were well grouped according to fertilization treatment (Fig. S1A and S1B). The NPK samples of both Nitrobacter- and Nitrospiralike NOB were well separated from those of the other fertilization treatments.
Fig. 2. Abundances of nitrite oxidizers under four fertilization treatments. (A) Nitrobacter-like NOB abundance (nxrA); (B) Nitrospira-like NOB abundance (nxrB). Ratios of nxrB/nxrA copy numbers are shown in boxes above the chart, and different lowercase letters above the nxrB/nxrA ratio indicate significant differences (P < 0.05). Error bars represent standard error and are accompanied by lowercase letters indicating significant differences according to Tukey's test (P < 0.05).
respectively, as compared with that under CK treatment. In addition, ANOVA indicated that PNO values were significantly (P < 0.01) influenced by fertilization treatment. Positive correlations were found between PNO and soil NO3− (ρ = 0.662, P < 0.01), TC (ρ = 0.825, P < 0.01), TN (ρ = 0.895, P < 0.01), and SOC (ρ = 0.835, P < 0.01) (Table S1). 3.3. Nitrobacter- and Nitrospira-like NOB abundances The abundances of Nitrobacter and Nitrospira were estimated by quantifying their respective nxrA/nxrB gene copy numbers. The highest
Table 2 Diversity indices of Nitrobacter- and Nitrospira-like NOB under different fertilization treatments calculated from high-throughput sequencing data.
Nitrobacter-like NOB
Nitrospira-like NOB
Treatment
ACE
Chao1
CK M NPK MNPK CK M NPK MNPK
494.9 ± 139.8a 728.3 ± 188.3a 745.4 ± 15.2a 704.2 ± 90.0a 875.5 ± 179.6b 1298.9 ± 105.7a 755.9 ± 68.4b 1010.8 ± 205.9ab
431.1 680.0 589.1 612.2 573.0 726.1 423.9 640.3
± ± ± ± ± ± ± ±
136.9a 181.1a 29.1a 83.6a 70.9b 27.6a 32.4c 77.8ab
Numbers in the same column followed by different letters are significantly different (Tukey's test, P < 0.05, n = 3). 121
Shannon
Simpson
4.60 6.36 4.37 4.54 4.03 5.01 2.26 5.31
0.85 0.94 0.78 0.84 0.83 0.92 0.44 0.93
± ± ± ± ± ± ± ±
0.19b 0.51a 0.37b 0.23b 0.23b 0.23a 0.43c 0.06a
± ± ± ± ± ± ± ±
0.03ab 0.01a 0.07b 0.01b 0.01a 0.01a 0.11b 0.00a
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Fig. 3. Nonmetric multidimensional scaling (NMDS) ordinations based on the Bray-Curtis dissimilarity matrices showing the changes in Nitrobacter (A) and Nitrospira (B) community composition under four fertilization treatments. Circle size is proportional to PNO level on the NMDS of the soil samples according to community structure.
Fig. 4. Redundancy analyses of nitrite oxidizer community structures under four fertilization treatments. (A) Nitrobacter-like NOB community structures; (B) Nitrospira-like NOB community structures. Monte Carlo permutation test: **P < 0.01.
were stimulated or repressed by soil pH, we identified OTUs with differential abundances in treatments with high pH (pH > 5.225) and low pH (pH < 5.225) according to the results of MRT analysis using a general linear model (Fig. 5C and D). At low pH, four and one OTUs were significantly (P < 0.05) elevated in the Nitrobacter- and Nitrospira-like NOB communities, respectively, all exhibiting a log2-fold change in relative abundance > 6.3. In contrast, seven and ten OTUs decreased in abundance in the Nitrobacter- and Nitrospira-like NOB communities, respectively, at low pH, all exhibiting a log2-fold change in relative abundance < −4.8. One Nitrospira-like OTU (Botu7) belonging to Nitrospira lineage II was not significantly affected by pH (Fig. S2). The ten pH-inhibited Nitrospira-like OTUs were affiliated with the Nitrospira Namibia soil cluster (Fig. 5D), while the seven pH-inhibited Nitrobacter-like OTUs were dominated by uncultured Nitrobacter-like bacteria (Fig. 5C and Fig. S2).
3.5. Relationship between NOB community composition and environmental variables RDA was used to identify the potential effect of environmental factors on the Nitrobacter- and Nitrospira-like NOB community composition across different fertilization treatments (Fig. 4). Overall, the first two axes explained 34.4% and 41.4% of the total variability in the Nitrobacter- and Nitrospira-like NOB communities (Fig. 4A and B), respectively. Among the environmental factors measured, soil pH was the major factor influencing the community composition of Nitrobacter- and Nitrospira-like NOB (F = 2.23 and 3.52, respectively, P < 0.01). In addition, soil SOC and TN contents significantly explained variations in the Nitrobacter- and Nitrospira-like NOB communities, respectively (F = 2.04 and 2.98, respectively, P < 0.01). The MRT analysis was employed to interpret the relationship between the relative abundances of OTUs and soil environmental conditions by providing a tree with four terminal nodes (Fig. 5). Soil pH appeared to be a strong predictor of relative OTU abundance, which was separated with different fertilization treatment, with samples with low pH levels (< 5.225, NPK treatment) clustering separately from those with higher pH values (> 5.225, CK, M, and MNPK treatment) based on Nitrobacter- and Nitrospira-like NOB communities (Fig. 5A and B). In addition, to identify which OTUs (relative abundance > 1%)
3.6. Predictive importance of abundance, diversity and composition of NOB on PNO An automatic liner modeling analysis was employed to evaluate the predictive importance of the abundance, diversity and composition of NOB (Fig. 6). The results reveal that the nxrB/nxrA abundance ratio (69%) is the best predictor for PNO (Fig. 6A). The abundances of nxrB 122
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Fig. 5. Multivariate regression tree analysis of the relationships between relative abundances of dominant OTUs (relative abundance > 1%) and soil properties in nitrite oxidizer community structures under four fertilization treatments. (A) Nitrobacter-like NOB community structures; (B) Nitrospira-like NOB community structures. Soil properties included pH, TC, TN, NH4+, NO3−, and SOC. Differential abundance analysis of the dominant OTUs that significantly responded to soil pH (FDR-corrected P < 0.05). (C) Nitrobacter-like dominant OTUs; (D) Nitrospira-like dominant OTUs.
and nxrA subsequently explained 23% and 8% of the residual PNO, respectively. Thus, the data strongly implicated that the changes of soil PNO was mediated through the shift of the Nitrospira/Nitrobacter abundance ratio. For alpha-diversity, the Chao1 and ACE index of Nitrobacter exhibit strong effects on PNO (Fig. 6B), followed by ACE index of Nitrospira (14%) and Shannon indices of Nitrobacter (12%). The contribution of the other diversity indexes is limited. For the nitrite oxidizers community structure, the Nitrobacter is the most important predictor for PNO, accounting for approximately 72% (NMDS1 = 69% and NMDS2 = 3%) of the relative influence (Fig. 6C), and Nitrospira shows weaker effects on PNO (28%). 4. Discussions 4.1. Environmental determinants of PNO and nitrite oxidizer abundances Our results show that soil PNO was significantly elevated following fertilization compared with levels in the CK treatment. We also found that NPK treatment resulted in lower PNO values than M and MNPK treatments. This is likely linked to the reduced abundances of Nitrobacter, as nxrA was generally positively correlated with PNO (Attard et al., 2010). It has been shown that a lower soil pH can influence ammonia oxidizer activity by decreasing the availability of ammonia due to an increase in ionization to ammonium as pH decreases (De Boer and Kowalchuk, 2001); however, there was no strong negative correlation between PNO and soil pH (Table S1). Concurrently, PNO was significantly and strongly correlated with the soil NO3−, TC, TN, and SOC contents. These data support our hypothesis (1) that chemical fertilizers have a negative impact on the activity of NOB as they lead to soil acidification. A previous study showed that the abundances of Nitrobacter-like NOB increased linearly with increasing N levels along an N fertilization gradient while that of Nitrospira-like NOB did not change significantly in Tibetan alpine meadows (Ma et al., 2016). Wertz et al. (2011) also addressed that fertilization increased Nitrobacter abundances but not Nitrospira in forest soils. In our study, N fertilizers significantly increased the Nitrobacter abundance, except for NPK treatment. We also
Fig. 6. Predictive importance of abundances (A), alpha-diversity indexes (B) and composition of NOB (C) on soil PNO determined by automatic linear modeling.
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2001). This supports that lower pH values suppress the growth of Nitrobacter- and Nitrospira-like OTUs. Strikingly, in our study, there were still a small number of OTUs at high relative abundance in these low-pH soils. Kuang et al. (2013) found that some Nitrospira (Leptospirillum spp.), identified based on 16S rRNA gene sequences, are adapted to more acidic (pH = 2.6 ± 0.45, mean ± SD) environments. It is probable that the Nitrospira-like NOB taxa in the Red soil exhibit a wider range of pH adaptability (ranging from 4.83 to 6.87). This implies that NOB can adapt to and tolerate acidic soil environments, increasing our understanding of NOB ecology and evolution.
found that fertilization led to a decrease in Nitrospira abundance (Fig. 2). This suggests that N inputs may be a key factor for both Nitrobacter and Nitrospira abundances in this Red soil. Furthermore, changes in the abundances of Nitrobacter- and Nitrospira-like NOB are significantly (P < 0.05) positively and negatively correlated with soil TC and TN content, respectively. It is likely that Nitrobacter have a lower affinity than Nitrospira for N substrates and are thus generally favored by higher N levels (Schramm et al., 1999; Wagner et al., 2002). Nitrobacter is common in nutrient-rich environments like wastewater (Wagner et al., 2002). It has been speculated that Nitrobacter can outcompete Nitrospira at high substrate concentrations (Schramm et al., 1999; Le Roux et al., 2016). This may partially explain the above results and the decreases in the nxrB/nxrA ratio in the fertilized plots, as higher TC and TN contents and fertilizer application generally suggest higher nutrient or N levels. However, the fact that the nxrB/nxrA ratio in the NPK plots was higher than those with manure is interesting. This likely stems from the fact that Nitrospira in NPK plots became more competitive compared with Nitrobacter in the manure treatments. However, whether this phenomenon was caused by low pH is unclear.
4.3. Abundance, diversity and composition of NOB to drive soil PNO Automatic liner modeling analysis deciphers that the nxrB/nxrA ratio is the most important variable that influences PNO. It seems that the interaction between Nitrospira- and Nitrobacter-like NOB community is a driving factor for regulating nitrite-oxidation process in Red soil. In fact, there is compelling evidence that both of Nitrospira- and Nitrobacter-like NOB community structures are significantly correlated to soil PNO (Fig. 3). This is different from what was reported by Han et al. (2017) who showed that PNO is positively correlated only with the abundance of Nitrospira in the soil with a rapeseed-rice rotation system. In an earlier study, PNO was found to be positively and negatively correlated with Nitrobacter and Nitrospira abundance, respectively, in agricultural soils with varying tillage practices (Attard et al., 2010). Indeed, both studies suggested that Nitrospira-like rather than Nitrobacter-like nitrite oxidizers play a major functional role in low activity soils (around 0.5 mg N-NO2- g−1 h−1). Ma et al. (2016) indicated that Nitrobacter play a vital role in nitrification along nitrogen gradients, while Nitrospira also contribute to the whole activity along phosphorus gradients. In the present study, the community structure of Nitrobacter and Nitrospira exert a great effect on soil PNO indicated by the ρ and P values (Fig. 3). In short, our results demonstrate that both Nitrobacter- and Nitrospira-like NOB are the key functional players within the NOB community in Red soils, and the changes in PNO are assigned to the shifts between the two communities.
4.2. Relationship between environmental factors and nitrite oxidizer community structures A previous study reported that the Nitrospira community is more diverse under fertilization compared to that under unfertilized management (Freitag et al., 2005). However, Wertz et al. (2011) revealed no differences in the diversity of the Nitrobacter community and only rapid shifts in the structure of the Nitrobacter-like NOB community between long-term fertilized and unfertilized forest soils. We found significant changes in the α-diversity of the Nitrospira community among different fertilizer treatments, but for Nitrobacter, the α-diversity in CK, NPK, and MNPK soils was similar (Table 2). For Nitrospira, αdiversity was highest in the MNPK and M treatments, followed by CK and NPK treatments, while for Nitrobacter, α-diversity was highest in the M treatment, with similar values in the CK, MNPK, and NPK treatments. This supports our hypotheses that (1) chemical fertilizers have a negative impact on the diversity of NOB as they lead to soil acidification; (2) manure has a positive impact on maintaining diversity by providing both inorganic and organic nutrients for NOB and not leading to soil acidification; and (3) the combined use of M and NPK provides more types of inorganic and organic nutrients and balances the negative effect of chemical fertilizers on NOB diversity. It is not surprising that the composition of NOB was altered by the introduction of different fertilization regimes. Therefore, it was necessary to determine how soil properties affected the composition of these NOBs under different fertilization regimes. RDA indicated that the variability in Nitrobacter-like NOB communities was significantly explained by soil pH and SOC content (Fig. 4A). In addition, Nitrospiralike NOB communities were significantly explained by soil pH and TN content (Fig. 4B). These three variables are probably key factors that shape the NOB community structure in this red soil. This differs from what was observed by Han et al. (2017), who found that the Nitrobacterlike NOB community was not shaped by soil pH in a rotation trial. In contrast, Hu et al. (2014) showed that pH is the most important factor affecting the diversity and community structure of AOA and AOB in Chinese agricultural soils. Considering that the function of ammonia and nitrite oxidation is spatially dependent (Grundmann and Debouzie, 2000; Ke et al., 2013; Stempfhuber et al., 2016), it is probable that the Nitrobacter- and Nitrospira-like NOB communities were also affected by soil pH. Differential abundance analysis indicated that most OTUs (relative abundance > 1%) classified as Nitrobacter- and Nitrospira-like NOB were inhibited by low soil pH (< 5.225). Previous studies have suggested that Nitrospira grows optimally at pH values between 7.6 and 8.3 (Ehrich et al., 1995; Blackburne et al., 2007) and that Nitrobacter grows optimally at pH values between 7.6 and 8.2 (Grunditz and Dalhammar,
Compliance with ethical standards This article does not contain any studies with animals performed by any of the authors. All authors have read and approved the final manuscript. Conflicts of interest The authors declare that they have no competing interests. Acknowledgments This work was supported by the National Basic Research Program of China (973, grant No. 2015CB150504), and the Fundamental Research Funds for the Central Universities (project No. 2662015PY016, 2662015PY116). Sequencing service was provided by Personal Biotechnology Co., Ltd. Shanghai, China. Appendix A. Supplementary data Supplementary data related to this article can be found at http://dx. doi.org/10.1016/j.soilbio.2018.05.033. References Alawi, M., Lipski, A., Sanders, T., Eva Maria, P., Spieck, E., 2007. Cultivation of a novel cold-adapted nitrite oxidizing betaproteobacterium from the Siberian Arctic. The ISME Journal 1, 256–264. Attard, E., Poly, F., Commeaux, C., Laurent, F., Terada, A., Smets, B.F., et al., 2010. Shifts
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Shifts in Nitrobacter- and Nitrospira-like nitrite-oxidizing bacterial communities under long-term fertilization practices
T
Shun Hana, Luyang Zenga,b, Xuesong Luoa,b, Xiang Xionga,b, Shilin Wenc, Boren Wangc, Wenli Chena,∗, Qiaoyun Huanga,b,∗∗ a
State Key Laboratory of Agricultural Microbiology, Huazhong Agricultural University, Wuhan, 430070, China Key Laboratory of Arable Land Conservation (Middle and Lower Reaches of Yangtze River), Ministry of Agriculture, College of Resources and Environment, Huazhong Agricultural University, Wuhan, 430070, China c Hengyang Red Soil Experimental Station, Chinese Academy of Agricultural Sciences, Hengyang, 421001, China b
A R T I C LE I N FO
A B S T R A C T
Keywords: Fertilization treatment Nitrite-oxidizing bacteria (NOB) Nitrospira-like NOB Nitrobacter-like NOB Potential nitrite oxidation activity (PNO)
Nitrite-oxidizing bacteria (NOB) are key players in the second step of nitrification, which is an important process in the soil nitrogen (N) cycle. However, the ecology of nitrite oxidizers and their response to disturbances such as long-term fertilization practices are scarcely known in agricultural ecosystems. We used samples from a Red soil subject to a long-term chemical and organic fertilization experiment, including control without fertilizer (CK), swine manure (M), chemical fertilization (NPK), and chemical/manure combined fertilization (MNPK) treatment, to explore how agricultural practices impact the community structure, abundance, and potential activity of nitrite oxidizers (PNO). The abundance of Nitrobacter was significantly increased in the M and MNPK plots, whereas the abundance of Nitrospira was significantly reduced in the M and NPK treatment plots and less inhibited in the MNPK treatment. The PNO showed a similar trend to that for Nitrobacter abundance. The diversity of Nitrobacter increased in the M-treated plots, while that of Nitrospira increased in the M and MNPK plots and decreased in the NPK plots. Non-metric multidimensional scaling (NMDS) revealed that the Nitrobacter- and Nitrospira-like NOB community was shift in these four fertilization treatments. Redundancy analysis showed that pH+SOC (soil organic carbon) and pH+TN (total nitrogen) significantly explained the variation in the composition of Nitrobacter and Nitrospira, respectively. In addition, the Nitrospira/Nitrobacter abundance ratio and community structure of Nitrobacter- and Nitrospira-like NOB are responsible for the changes of soil PNO. Collectively, these data suggest that the nitrite-oxidation process in the red soil is possibly controlled by both Nitrospira and Nitrobacter-like NOB, which were shaped by pH+TN and pH+SOC, respectively.
1. Introduction Nitrification, the microbiological oxidation of ammonia (NH4+) to nitrite (NO2−) and subsequently to nitrate (NO3−), influences the fate of nitrogen (N) in terrestrial systems. The first limiting step of nitrification, NH4+ oxidation to NO2−, is mediated by ammonia-oxidizing bacteria (AOB) of the β- and γ-Proteobacteria, ammonia-oxidizing archaea (AOA) of the Thaumarchaeota (Kowalchuk and Stephen, 2001; Leininger et al., 2006; Schleper and Nicol, 2010; Norton and Stark, 2011). The second step of nitrification, NO2− oxidation to NO3−, is catalyzed by nitrite-oxidizing bacteria (NOB), which play an important role in the biogeochemical N cycle in many terrestrial ecosystems such as soils (Prosser, 1989; Daims et al., 2015). NOB are broadly distributed among the α-, β-, γ-, and δ-Proteobacteria as well as the Nitrospira classs
∗
(Gould and Lees, 1960; Teske et al., 1994; Alawi et al., 2007; Sorokin et al., 2012). Comammox Nitrospira complete oxidation of ammonium to nitrate for the nitrification (Daims et al., 2015; van Kessel et al., 2015). In recent years, many studies have focused on the ecology of AOA and AOB in a broad range of soil environments (Kowalchuk and Stephen, 2001; Webster et al., 2005; Chu et al., 2007; Chen et al., 2008; Shen et al., 2012; Bertagnolli et al., 2016; Zhang et al., 2017). Ammonia oxidation is often assumed to be the rate-limiting step of nitrification as nitrite generally does not accumulate in the environment. In fact, nitrite oxidation can also become the limiting step for nitrification in disturbed soil systems (Gelfand and Yakir, 2008; Roux-Michollet et al., 2008). In view of the importance of nitrite oxidizers, it is urgently needed to decipher the ecological response of nitrite oxidizer community to any
Corresponding author. State Key Laboratory of Agricultural Microbiology, Huazhong Agricultural University, Wuhan 430070, China. Corresponding author. State Key Laboratory of Agricultural Microbiology, Huazhong Agricultural University, Wuhan 430070, China. E-mail addresses: [email protected] (W. Chen), [email protected] (Q. Huang).
∗∗
https://doi.org/10.1016/j.soilbio.2018.05.033 Received 22 November 2017; Received in revised form 30 May 2018; Accepted 31 May 2018 0038-0717/ © 2018 Elsevier Ltd. All rights reserved.
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2016. Six soil cores (approximately 5 cm in diameter) were taken from each plot and mixed to form one composite sample. Samples were placed in a sterile plastic bag for transport to the laboratory within 24 h after collection. Each soil sample was divided into three portions. One portion was used for DNA extraction and stored at −80 °C, another was stored at 4 °C for measuring PNO, and the third was air-dried at room temperature for analysis of soil chemical properties.
type of environmental disturbances, such as different fertilization regimes in agricultural soils. Nitrobacter-like NOB are considered to be the key functional players within the NOB community, exhibiting high activity levels in tillage/ no-tillage agricultural systems with high N availability (Attard et al., 2010). In addition, soil potential nitrite oxidation activity (PNO) is strongly positively correlated with the abundance of Nitrobacter-like NOB as well as weakly negatively correlated with the abundance of Nitrospira-like NOB. Possible changes were also observed for the abundance and community of Nitrobacter and Nitrospira in agricultural soils in response to tillage practices. In contrast, fertilization causes rapid shifts in the structure of the Nitrobacter-like NOB community, which dominates nitrification in forest soils (Wertz et al., 2011). The application of manure in a Merzenhausen agricultural soil was found to have decreased the diversity of Nitrobacter community in the rhizosphere (Ollivier et al., 2013). Nitrite oxidation in surface agricultural soils may be predominantly driven by Nitrospira spp. (Ke et al., 2013). Han et al. (2017) showed that Nitrospira might be more responsive than Nitrobacter in soils under a rapeseed-rice rotation. In addition, the Nitrospira-like NOB community was found to be significantly shaped by soil pH, moisture, and NH4+ content, whereas the Nitrobacter-like NOB community was not. In acidic forest soils, long-term fertilization increased AOB and Nitrobacter-like NOB abundances but did not influence AOA and Nitrospira-like NOB abundances (Wertz et al., 2011). However, the responses of the community structure and diversity of NOB to long-term fertilization regimes and their links with nitrite oxidation activities in agricultural soils are still not well understood. The ways in which NOB populations respond to changes in soil pH, specifically the acidification of terrestrial environments, have rarely been investigated. In this study, samples from a Red soil in the Qiyang Long Term Soil Experimental Station, Hunan Province, China was used to investigate the ecological effects of long-term fertilization practices on the Nitrobacter- and Nitrospira-like bacterial community and their potential activity. Quantitative PCR (qPCR) and high-throughput sequencing of marker genes were performed to investigate the abundance, community diversity, and population composition of Nitrobacter- and Nitrospira-like NOB in the soil. We hypothesized that (1) chemical fertilization may have a negative impact on the activity and diversity of NOB; (2) manure would have a positive effect on NOB community structure, as it provides both inorganic and organic nutrients for NOB and does not lead to acidification; (3) the combined use of manure and chemical fertilizers would represent a balanced fertilization treatment, inhibiting soil acidification and balancing the negative effect of chemical fertilizers on NOB activity and diversity.
2.2. Soil chemical analytical procedures Soil total carbon (TC) and total nitrogen (TN) contents were analyzed using a Vario Max element analyzer (Elementar Vario PYRO cube and Isoprime100, Germany). Soil exchangeable ammonium (NH4+-N) and nitrate (NO3−-N) contents were determined on a FIAstar 5000 Analyzer (Foss Tecator, Denmark) after extraction from fresh soil with 2 M KCl (w/v, 1:5). Soil pH was determined at a soil/water ratio of 1:2.5, and soil organic content (SOC) was determined by the K2Cr2O7 oxidation method. 2.3. Assays for determination of PNO PNO was determined using the method described by Wertz et al. (2007), modified from Smorczewski and Schmidt (1991). Briefly, samples of fresh soil (5 g equivalent dry mass) were incubated with 50 ml of a solution of NaNO2 (5 μg of N-NO2 g−1 dry soil) dissolved in autoclaved deionized water for 30 h with gentle shaking (180 rpm) at 28 °C. During incubation, 2.0-ml aliquots of the suspensions were sampled at 0, 4, 8, 12, 24, and 30 h and centrifuged (5000 rpm for 5 min). The supernatants were filtered (0.2-μm pore size) and analyzed for nitrite (N-NO2-) concentration on a spectrophotometer (L6, Shanghai, China) at 520 nm using Griess reagent at room temperature (Smorczewski and Schmidt, 1991). Levels of nitrite generally decreased linearly with time throughout the first 12 h until nitrite was depleted. Rates were calculated from the linear decrease and taken as the PNO. In this study, according to the literature, we used nitrite solution to measure the potential value directly without considering the ammonia oxidation process (Smorczewski and Schmidt, 1991; Wertz et al., 2007; Attard et al., 2010). The nitrite produced from ammonia oxidation process during incubation was not taken into account. 2.4. Soil DNA extraction DNA was extracted from 0.5 g soil using Lysing MatrixB tubes (Bio101) as described previously by Griffiths et al. (2000). Humic acids, a PCR inhibitor, were removed from the soil DNA using DNA-EZ Reagents M Humic Acid-Be-Gone B (Sangon Biotech, Shanghai, China). Purification was performed according to the manufacturer's instructions. The quality and concentration of DNA were determined using a NanoDrop spectrophotometer (NanoDrop Technologies, Wilmington, DE, USA).
2. Material and methods 2.1. Experimental site and sampling The experimental site was located at the Qiyang Red Soil Experimental Station (26°45′N, 111°52′E), Hunan Province, China. This site represents a typical agricultural region of subtropical China. It has a subtropical monsoon climate with an annual rainfall of 1300 mm and annual average temperature of 18 °C. A long-term fertilizer experiment was established in 1990 with a winter wheat (Triticum aestivum L.) and summer maize (Zea mays L.) rotation system, including three replicates of four treatments in a randomized plot design: control without fertilizers (CK); swine manure (M); chemical fertilization (nitrogen, phosphate, and potassium fertilizers, NPK); and chemical/organic combined fertilization (nitrogen, phosphate, potassium, and swine manure fertilizers, MNPK). The NPK-treated soil was severely acidified. The nitrogen fertilizer was applied as urea or swine manure at 300 kg N ha−1, phosphate (P) as a single application of superphosphate [Ca(H2PO4)2] at 53 kg P ha−1, and potassium (K) as potassium chloride (KCl) at 100 kg K ha−1. Soil samples were collected at a depth of 0–20 cm in November
2.5. Measurement of Nitrobacter- and Nitrospira-like NOB abundance by qPCR QPCR assays were conducted using an ABI7500 FAST Real-time PCR system with the nxrA primers F1norA and R2norA (Attard et al., 2010) for Nitrobacter-like NOB and the nxrB gene primers nxrB169f and nxrB638r (Pester et al., 2014) for Nitrospira-like NOB. The 20-μl PCR reaction mixtures contained 10 μl SYBR Premix Ex Taq II (2×) (Takara, Bio Inc., Shiga, Japan), 1.0 μl of a 10 mM solution of each primer, 6.0 μl DEPC-treated water, and 2.0 μl soil extract (diluted to 5 ng/μl) or 2.0 μl standard plasmid. A standard curve was generated using ten-fold serial dilutions of a plasmid containing a copy of the target gene. The following program was used for the nxrA gene: 95 °C for 10 min followed by 40 cycles of denaturation at 95 °C for 30 s, annealing at 55 °C for 30 s, and elongation at 72 °C for 30 s. For the nxrB gene, the program 119
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Table 1 Soil physicochemical properties of the studied soil samples. Treatments
pH (1:2.5)
CK M NPK MNPK
6.87 6.52 4.83 5.90
± ± ± ±
TC (%) 0.02a 0.11a 0.05c 0.29b
0.64 1.56 1.02 1.36
± ± ± ±
0.05d 0.05a 0.06c 0.05b
0.11 0.21 0.15 0.19
NO3− (mg/kg)
NH4+ (mg/kg)
TN (%) ± ± ± ±
0.004d 0.003a 0.008c 0.006b
9.03 7.62 9.95 8.06
± ± ± ±
2.03a 0.14a 0.33a 0.24a
3.43 19.8 15.9 17.4
± ± ± ±
SOC (g/kg) 0.51b 2.18a 4.07a 5.12a
17.3 33.3 23.1 28.2
± ± ± ±
0.43d 1.36a 0.81c 1.41b
Numbers in the same column followed by different letters are significantly different (Tukey's test, P < 0.05, n = 3).
the correlation between the rank similarity matrices for PNO and NOB community structure (Attard et al., 2010). Multivariate regression tree (MRT) analysis (De'ath, 2002) was performed using the R package mvpart (with default parameters) to relate the relative abundances of lineages to the site characteristics. A general linear model in combination with quasi-likelihood shrunken dispersion estimates was implemented in the R package edgeR 3.6.8 (Robinson et al., 2010; Lund et al., 2012) to identify which OTUs were stimulated or repressed by soil pH. Multiple sequence alignments of genes were performed using Muscle based alignments of the translated amino acid sequences. Phylogenetic trees were constructed by neighbor-joining using the selected sequences (relative abundance > 1%) with 1000 bootstrap replicates of MEGA 6. The reliability of the tree topology was evaluated by 1000 replicates of bootstrap resampling.
consisted of: 95 °C for 10 min followed by 40 cycles of denaturation at 95 °C for 30 s, annealing at 56.2 °C for 45 s, and elongation at 72 °C for 45 s. A negative control was included in each run using water instead of soil DNA extract. Melting curve and gel electrophoresis analyses were carried out to confirm amplification specificity. The dilution of DNA extracts is a commonly used method to reduce levels of inhibition (Wang et al., 2017), and several DNA samples were diluted to 5 ng/μl based on a preliminary experiment to determine the optimal dilution concentration. 2.6. High-throughput sequencing and bioinformatics analysis The sequences of the PCR products of the nxrA and nxrB genes were obtained using the Illumina MiSeq PE300 platform and the primers described above. To distinguish amplicons originating from different soil samples, barcode oligonucleotides 7 bp in length were ligated to the ends of the purified PCR products. The PCR products were checked by agarose gel electrophoresis and cleaned by Agencourt AMPure XP (Beckman Coulter, Inc. S. Kraemer Boulevard Brea, CA, USA). The DNA concentration of the purified PCR product was measured using the Quant-iT™ PicoGreen dsDNA BR Assay Kit (Invitrogen) and a microplate reader (BioTek FLx800) according to the manufacturer's protocol. Next, an equal amount of PCR product for each sample was combined in a single tube to be run on the sequencing platform. Raw reads were filtered based on the barcodes using QIIME (Caporaso et al., 2010), and low-quality sequences (quality score < 20) and ambiguous bases were removed using USEARCH (http://www.drive5.com/usearch/). The remaining sequences were further screened for frame shifts using FrameBot from the RDP FunGene Pipeline (http://fungene.cme.msu.edu/ FunGenePipeline). The remaining quality-screened sequences were clustered into operational taxonomic units (OTUs) using UCLUST (Edgar, 2010) based on a 95% nucleic acid sequence identity cutoff. In addition, OTUs with abundances less than 0.001% of the total sample sequence were removed (Bokulich et al., 2013), and then the OTU table was used for subsequent analysis.
3. Results 3.1. Soil physicochemical properties The physicochemical properties of the collected soil samples are shown in Table 1. Significant differences (P < 0.05) in soil pH were observed among all treatments. The CK and M plots had higher pH values (Table 1), while the lowest pH values (4.83) were recorded from the NPK samples. There were also significant differences (P < 0.05) in soil TC, TN, and SOC contents among all treatments, with the highest levels in the M treatment, followed by those in the MNPK, NPK, and CK treatments. The CK soil NO3− content was the lowest among all treatments. Ammonia content was relatively stable among the different fertilization regimes. 3.2. Soil PNO Soil PNO values ranged from 0.07 to 0.37 μg NO2−-N kg−1 dry soil h (Fig. 1). PNO was significantly (P < 0.01) elevated by about 3.9-, 3.7-, and 4.6-fold under the M, NPK, and MNPK treatments, −1
2.7. Statistical analysis One-way analysis of variance (ANOVA) was used to analyze the effects of fertilization treatments on the abundances of Nitrobacter- and Nitrospira-like NOB, PNO, and soil variables and automatic liner modeling was performed at the confidence level of 95% by using SPSS 19.0 statistical software (IBM Co., Armonk, NY USA). Spearman's correlation was used to determine whether the correlation between the abundances of Nitrobacter- and Nitrospira-like NOB, PNO, and each soil variable was significant. The program Mothur (http://www.mothur.org/wiki/ MainPage) was used to determine α-diversity indices. Redundancy analysis (RDA) was carried out using the Canoco 4.5. Monte Carlo permutation test with 999 unrestricted permutation to determine the extent of the environmental parameter(s) that were able to explain the variation in the nitrifier community. Non-metric multidimensional scaling (NMDS) ordination plots were used to identify the differences in NOB community composition using the R package vegan. Spearman correlation coefficient and associated p significance level (obtained by a permutation test using 999 permutations) were computed to quantify
Fig. 1. Soil potential nitrite oxidation activity (PNO) under four fertilization treatments. Error bars represent standard error and are accompanied by lowercase letters indicating significant differences according to Tukey's test (P < 0.05). 120
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Nitrobacter population size was detected in the MNPK treatment, followed by the M, CK, and NPK treatments in a significantly (P < 0.05) decreasing order (Fig. 2A). In contrast, the Nitrospira population sizes were significantly (P < 0.05) higher in the CK treatment than in the other fertilization treatments (Fig. 2B). Fertilization exerted considerable (P < 0.01) effects on the gene copy numbers of both Nitrobacter and Nitrospira. The ratio of nxrB/nxrA gene copy numbers ranged from 7.6 to 168.2, with the highest ratio observed for CK treatment, followed by NPK, MNPK, and M treatments, indicating a significant (P < 0.05) negative impact of fertilization on the nxrB/nxrA ratio. Spearman's rank correlation showed that soil TC and TN had significant (P < 0.05) positive and negative impacts on the abundances of nxrA and nxrB, respectively (Table S1). In addition, PNO was found to be highly positively correlated with Nitrobacter-like nxrA copy number (ρ = 0.664, P < 0.05) except for NPK treatment, but was not correlated with Nitrospira-like nxrB copy number (ρ = −0.559, P = 0.059). However, PNO were strongly and negatively (ρ = −0.923, P < 0.01) correlated with the nxrB/nxrA abundance ratio.
3.4. Nitrobacter- and Nitrospira-like NOB community structures The community structures of Nitrobacter- and Nitrospira-like NOB were characterized by investigating the nxrA and nxrB genes using high-throughput amplicon sequencing. Sequencing yielded a total of 678,665 high-quality nxrA and 370,887 nxrB gene sequences, corresponding to 2878 and 3243 unique OTUs classified as Nitrobacter- and Nitrospira-like NOB, respectively. A resampling procedure was employed at a depth of 28193 and 16802 sequences per sample to calculate the alpha diversity index for Nitrobacter and Nitrospira, respectively. Table 2 shows the diversity properties of Nitrobacter- and Nitrospiralike NOB. For Nitrobacter-like NOB, there were no significant differences in the ACE or Chao1 indices among the treatments. The highest Shannon and Simpson values were detected in the M plots, while those for the other treatments were similar. For Nitrospira-like NOB, the Shannon and Simpson values were highest in the MNPK and M treatments, followed by the CK and then NPK treatments (Table 2). Analysis of similarities (ANOSIM) of the Nitrobacter- (R = 0.58, P = 0.001) and Nitrospira-like NOB (R = 0.72, P = 0.001) community data indicated that fertilization regime was a significant driver of the NOB community. NMDS analysis reveals that the Nitrobacter- and Nitrospira-like NOB communities clustered separately along NMDS1 and NMDS2 according to fertilization treatments (Fig. 3A and B). A significant correlation was observed between PNO levels and the Nitrobacter- (ρ = 0.47, P = 0.005) and Nitrospira-like (ρ = 0.41, P = 0.015) NOB community structure. The hierarchical clustering of microbial communities also demonstrated that the Nitrobacter- and Nitrospira-like NOB were well grouped according to fertilization treatment (Fig. S1A and S1B). The NPK samples of both Nitrobacter- and Nitrospiralike NOB were well separated from those of the other fertilization treatments.
Fig. 2. Abundances of nitrite oxidizers under four fertilization treatments. (A) Nitrobacter-like NOB abundance (nxrA); (B) Nitrospira-like NOB abundance (nxrB). Ratios of nxrB/nxrA copy numbers are shown in boxes above the chart, and different lowercase letters above the nxrB/nxrA ratio indicate significant differences (P < 0.05). Error bars represent standard error and are accompanied by lowercase letters indicating significant differences according to Tukey's test (P < 0.05).
respectively, as compared with that under CK treatment. In addition, ANOVA indicated that PNO values were significantly (P < 0.01) influenced by fertilization treatment. Positive correlations were found between PNO and soil NO3− (ρ = 0.662, P < 0.01), TC (ρ = 0.825, P < 0.01), TN (ρ = 0.895, P < 0.01), and SOC (ρ = 0.835, P < 0.01) (Table S1). 3.3. Nitrobacter- and Nitrospira-like NOB abundances The abundances of Nitrobacter and Nitrospira were estimated by quantifying their respective nxrA/nxrB gene copy numbers. The highest
Table 2 Diversity indices of Nitrobacter- and Nitrospira-like NOB under different fertilization treatments calculated from high-throughput sequencing data.
Nitrobacter-like NOB
Nitrospira-like NOB
Treatment
ACE
Chao1
CK M NPK MNPK CK M NPK MNPK
494.9 ± 139.8a 728.3 ± 188.3a 745.4 ± 15.2a 704.2 ± 90.0a 875.5 ± 179.6b 1298.9 ± 105.7a 755.9 ± 68.4b 1010.8 ± 205.9ab
431.1 680.0 589.1 612.2 573.0 726.1 423.9 640.3
± ± ± ± ± ± ± ±
136.9a 181.1a 29.1a 83.6a 70.9b 27.6a 32.4c 77.8ab
Numbers in the same column followed by different letters are significantly different (Tukey's test, P < 0.05, n = 3). 121
Shannon
Simpson
4.60 6.36 4.37 4.54 4.03 5.01 2.26 5.31
0.85 0.94 0.78 0.84 0.83 0.92 0.44 0.93
± ± ± ± ± ± ± ±
0.19b 0.51a 0.37b 0.23b 0.23b 0.23a 0.43c 0.06a
± ± ± ± ± ± ± ±
0.03ab 0.01a 0.07b 0.01b 0.01a 0.01a 0.11b 0.00a
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Fig. 3. Nonmetric multidimensional scaling (NMDS) ordinations based on the Bray-Curtis dissimilarity matrices showing the changes in Nitrobacter (A) and Nitrospira (B) community composition under four fertilization treatments. Circle size is proportional to PNO level on the NMDS of the soil samples according to community structure.
Fig. 4. Redundancy analyses of nitrite oxidizer community structures under four fertilization treatments. (A) Nitrobacter-like NOB community structures; (B) Nitrospira-like NOB community structures. Monte Carlo permutation test: **P < 0.01.
were stimulated or repressed by soil pH, we identified OTUs with differential abundances in treatments with high pH (pH > 5.225) and low pH (pH < 5.225) according to the results of MRT analysis using a general linear model (Fig. 5C and D). At low pH, four and one OTUs were significantly (P < 0.05) elevated in the Nitrobacter- and Nitrospira-like NOB communities, respectively, all exhibiting a log2-fold change in relative abundance > 6.3. In contrast, seven and ten OTUs decreased in abundance in the Nitrobacter- and Nitrospira-like NOB communities, respectively, at low pH, all exhibiting a log2-fold change in relative abundance < −4.8. One Nitrospira-like OTU (Botu7) belonging to Nitrospira lineage II was not significantly affected by pH (Fig. S2). The ten pH-inhibited Nitrospira-like OTUs were affiliated with the Nitrospira Namibia soil cluster (Fig. 5D), while the seven pH-inhibited Nitrobacter-like OTUs were dominated by uncultured Nitrobacter-like bacteria (Fig. 5C and Fig. S2).
3.5. Relationship between NOB community composition and environmental variables RDA was used to identify the potential effect of environmental factors on the Nitrobacter- and Nitrospira-like NOB community composition across different fertilization treatments (Fig. 4). Overall, the first two axes explained 34.4% and 41.4% of the total variability in the Nitrobacter- and Nitrospira-like NOB communities (Fig. 4A and B), respectively. Among the environmental factors measured, soil pH was the major factor influencing the community composition of Nitrobacter- and Nitrospira-like NOB (F = 2.23 and 3.52, respectively, P < 0.01). In addition, soil SOC and TN contents significantly explained variations in the Nitrobacter- and Nitrospira-like NOB communities, respectively (F = 2.04 and 2.98, respectively, P < 0.01). The MRT analysis was employed to interpret the relationship between the relative abundances of OTUs and soil environmental conditions by providing a tree with four terminal nodes (Fig. 5). Soil pH appeared to be a strong predictor of relative OTU abundance, which was separated with different fertilization treatment, with samples with low pH levels (< 5.225, NPK treatment) clustering separately from those with higher pH values (> 5.225, CK, M, and MNPK treatment) based on Nitrobacter- and Nitrospira-like NOB communities (Fig. 5A and B). In addition, to identify which OTUs (relative abundance > 1%)
3.6. Predictive importance of abundance, diversity and composition of NOB on PNO An automatic liner modeling analysis was employed to evaluate the predictive importance of the abundance, diversity and composition of NOB (Fig. 6). The results reveal that the nxrB/nxrA abundance ratio (69%) is the best predictor for PNO (Fig. 6A). The abundances of nxrB 122
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Fig. 5. Multivariate regression tree analysis of the relationships between relative abundances of dominant OTUs (relative abundance > 1%) and soil properties in nitrite oxidizer community structures under four fertilization treatments. (A) Nitrobacter-like NOB community structures; (B) Nitrospira-like NOB community structures. Soil properties included pH, TC, TN, NH4+, NO3−, and SOC. Differential abundance analysis of the dominant OTUs that significantly responded to soil pH (FDR-corrected P < 0.05). (C) Nitrobacter-like dominant OTUs; (D) Nitrospira-like dominant OTUs.
and nxrA subsequently explained 23% and 8% of the residual PNO, respectively. Thus, the data strongly implicated that the changes of soil PNO was mediated through the shift of the Nitrospira/Nitrobacter abundance ratio. For alpha-diversity, the Chao1 and ACE index of Nitrobacter exhibit strong effects on PNO (Fig. 6B), followed by ACE index of Nitrospira (14%) and Shannon indices of Nitrobacter (12%). The contribution of the other diversity indexes is limited. For the nitrite oxidizers community structure, the Nitrobacter is the most important predictor for PNO, accounting for approximately 72% (NMDS1 = 69% and NMDS2 = 3%) of the relative influence (Fig. 6C), and Nitrospira shows weaker effects on PNO (28%). 4. Discussions 4.1. Environmental determinants of PNO and nitrite oxidizer abundances Our results show that soil PNO was significantly elevated following fertilization compared with levels in the CK treatment. We also found that NPK treatment resulted in lower PNO values than M and MNPK treatments. This is likely linked to the reduced abundances of Nitrobacter, as nxrA was generally positively correlated with PNO (Attard et al., 2010). It has been shown that a lower soil pH can influence ammonia oxidizer activity by decreasing the availability of ammonia due to an increase in ionization to ammonium as pH decreases (De Boer and Kowalchuk, 2001); however, there was no strong negative correlation between PNO and soil pH (Table S1). Concurrently, PNO was significantly and strongly correlated with the soil NO3−, TC, TN, and SOC contents. These data support our hypothesis (1) that chemical fertilizers have a negative impact on the activity of NOB as they lead to soil acidification. A previous study showed that the abundances of Nitrobacter-like NOB increased linearly with increasing N levels along an N fertilization gradient while that of Nitrospira-like NOB did not change significantly in Tibetan alpine meadows (Ma et al., 2016). Wertz et al. (2011) also addressed that fertilization increased Nitrobacter abundances but not Nitrospira in forest soils. In our study, N fertilizers significantly increased the Nitrobacter abundance, except for NPK treatment. We also
Fig. 6. Predictive importance of abundances (A), alpha-diversity indexes (B) and composition of NOB (C) on soil PNO determined by automatic linear modeling.
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2001). This supports that lower pH values suppress the growth of Nitrobacter- and Nitrospira-like OTUs. Strikingly, in our study, there were still a small number of OTUs at high relative abundance in these low-pH soils. Kuang et al. (2013) found that some Nitrospira (Leptospirillum spp.), identified based on 16S rRNA gene sequences, are adapted to more acidic (pH = 2.6 ± 0.45, mean ± SD) environments. It is probable that the Nitrospira-like NOB taxa in the Red soil exhibit a wider range of pH adaptability (ranging from 4.83 to 6.87). This implies that NOB can adapt to and tolerate acidic soil environments, increasing our understanding of NOB ecology and evolution.
found that fertilization led to a decrease in Nitrospira abundance (Fig. 2). This suggests that N inputs may be a key factor for both Nitrobacter and Nitrospira abundances in this Red soil. Furthermore, changes in the abundances of Nitrobacter- and Nitrospira-like NOB are significantly (P < 0.05) positively and negatively correlated with soil TC and TN content, respectively. It is likely that Nitrobacter have a lower affinity than Nitrospira for N substrates and are thus generally favored by higher N levels (Schramm et al., 1999; Wagner et al., 2002). Nitrobacter is common in nutrient-rich environments like wastewater (Wagner et al., 2002). It has been speculated that Nitrobacter can outcompete Nitrospira at high substrate concentrations (Schramm et al., 1999; Le Roux et al., 2016). This may partially explain the above results and the decreases in the nxrB/nxrA ratio in the fertilized plots, as higher TC and TN contents and fertilizer application generally suggest higher nutrient or N levels. However, the fact that the nxrB/nxrA ratio in the NPK plots was higher than those with manure is interesting. This likely stems from the fact that Nitrospira in NPK plots became more competitive compared with Nitrobacter in the manure treatments. However, whether this phenomenon was caused by low pH is unclear.
4.3. Abundance, diversity and composition of NOB to drive soil PNO Automatic liner modeling analysis deciphers that the nxrB/nxrA ratio is the most important variable that influences PNO. It seems that the interaction between Nitrospira- and Nitrobacter-like NOB community is a driving factor for regulating nitrite-oxidation process in Red soil. In fact, there is compelling evidence that both of Nitrospira- and Nitrobacter-like NOB community structures are significantly correlated to soil PNO (Fig. 3). This is different from what was reported by Han et al. (2017) who showed that PNO is positively correlated only with the abundance of Nitrospira in the soil with a rapeseed-rice rotation system. In an earlier study, PNO was found to be positively and negatively correlated with Nitrobacter and Nitrospira abundance, respectively, in agricultural soils with varying tillage practices (Attard et al., 2010). Indeed, both studies suggested that Nitrospira-like rather than Nitrobacter-like nitrite oxidizers play a major functional role in low activity soils (around 0.5 mg N-NO2- g−1 h−1). Ma et al. (2016) indicated that Nitrobacter play a vital role in nitrification along nitrogen gradients, while Nitrospira also contribute to the whole activity along phosphorus gradients. In the present study, the community structure of Nitrobacter and Nitrospira exert a great effect on soil PNO indicated by the ρ and P values (Fig. 3). In short, our results demonstrate that both Nitrobacter- and Nitrospira-like NOB are the key functional players within the NOB community in Red soils, and the changes in PNO are assigned to the shifts between the two communities.
4.2. Relationship between environmental factors and nitrite oxidizer community structures A previous study reported that the Nitrospira community is more diverse under fertilization compared to that under unfertilized management (Freitag et al., 2005). However, Wertz et al. (2011) revealed no differences in the diversity of the Nitrobacter community and only rapid shifts in the structure of the Nitrobacter-like NOB community between long-term fertilized and unfertilized forest soils. We found significant changes in the α-diversity of the Nitrospira community among different fertilizer treatments, but for Nitrobacter, the α-diversity in CK, NPK, and MNPK soils was similar (Table 2). For Nitrospira, αdiversity was highest in the MNPK and M treatments, followed by CK and NPK treatments, while for Nitrobacter, α-diversity was highest in the M treatment, with similar values in the CK, MNPK, and NPK treatments. This supports our hypotheses that (1) chemical fertilizers have a negative impact on the diversity of NOB as they lead to soil acidification; (2) manure has a positive impact on maintaining diversity by providing both inorganic and organic nutrients for NOB and not leading to soil acidification; and (3) the combined use of M and NPK provides more types of inorganic and organic nutrients and balances the negative effect of chemical fertilizers on NOB diversity. It is not surprising that the composition of NOB was altered by the introduction of different fertilization regimes. Therefore, it was necessary to determine how soil properties affected the composition of these NOBs under different fertilization regimes. RDA indicated that the variability in Nitrobacter-like NOB communities was significantly explained by soil pH and SOC content (Fig. 4A). In addition, Nitrospiralike NOB communities were significantly explained by soil pH and TN content (Fig. 4B). These three variables are probably key factors that shape the NOB community structure in this red soil. This differs from what was observed by Han et al. (2017), who found that the Nitrobacterlike NOB community was not shaped by soil pH in a rotation trial. In contrast, Hu et al. (2014) showed that pH is the most important factor affecting the diversity and community structure of AOA and AOB in Chinese agricultural soils. Considering that the function of ammonia and nitrite oxidation is spatially dependent (Grundmann and Debouzie, 2000; Ke et al., 2013; Stempfhuber et al., 2016), it is probable that the Nitrobacter- and Nitrospira-like NOB communities were also affected by soil pH. Differential abundance analysis indicated that most OTUs (relative abundance > 1%) classified as Nitrobacter- and Nitrospira-like NOB were inhibited by low soil pH (< 5.225). Previous studies have suggested that Nitrospira grows optimally at pH values between 7.6 and 8.3 (Ehrich et al., 1995; Blackburne et al., 2007) and that Nitrobacter grows optimally at pH values between 7.6 and 8.2 (Grunditz and Dalhammar,
Compliance with ethical standards This article does not contain any studies with animals performed by any of the authors. All authors have read and approved the final manuscript. Conflicts of interest The authors declare that they have no competing interests. Acknowledgments This work was supported by the National Basic Research Program of China (973, grant No. 2015CB150504), and the Fundamental Research Funds for the Central Universities (project No. 2662015PY016, 2662015PY116). Sequencing service was provided by Personal Biotechnology Co., Ltd. Shanghai, China. Appendix A. Supplementary data Supplementary data related to this article can be found at http://dx. doi.org/10.1016/j.soilbio.2018.05.033. References Alawi, M., Lipski, A., Sanders, T., Eva Maria, P., Spieck, E., 2007. Cultivation of a novel cold-adapted nitrite oxidizing betaproteobacterium from the Siberian Arctic. The ISME Journal 1, 256–264. Attard, E., Poly, F., Commeaux, C., Laurent, F., Terada, A., Smets, B.F., et al., 2010. Shifts
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