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Article

Impacts of Biochar and Gypsum on Ammonia-Oxidizing Microorganisms in Coastal Saline Soil

by
Hai Zhu
1,*,
Yuxing Liu
1,2 and
Rongjiang Yao
2,*
1
Engineering Research Center of Ecology and Agricultural Use of Wetland, Ministry of Education, College of Agriculture, Yangtze University, Jingzhou 434025, China
2
State Key Laboratory of Soil and Sustainable Agriculture, Institute of Soil Science, Chinese Academy of Sciences, Nanjing 210008, China
*
Authors to whom correspondence should be addressed.
Agronomy 2024, 14(8), 1756; https://doi.org/10.3390/agronomy14081756
Submission received: 24 June 2024 / Revised: 27 July 2024 / Accepted: 8 August 2024 / Published: 11 August 2024
(This article belongs to the Section Agroecology Innovation: Achieving System Resilience)
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Abstract

:
Nitrification is the core step of the soil nitrogen cycle and directly affects the nitrogen use efficiency in agricultural systems. Biochar and gypsum are two important soil amendments widely used in coastal saline farmland. However, little is known about their effects on nitrification and ammonia-oxidizing microorganisms. A one-year pot experiment with three treatments including biochar application (BC), gypsum application (SG), and no amendment (CK) was conducted, and the responses of the nitrification rate, amoA gene copies, and the diversity and community structure of ammonia-oxidizing archaea (AOA) and bacteria (AOB) to biochar and gypsum were evaluated. The results indicated that biochar and gypsum application both resulted in alterations to the soil properties. They both had inhibiting effects on nitrification and AOB amoA gene copies, whereas they had no significant effect on AOA amoA gene copies. Biochar had no significant effect on the diversity indexes of AOA, but it significantly reduced the Shannon index of AOB. Meanwhile, gypsum had no significant influence on the diversity indexes of both AOA and AOB. Biochar and gypsum did not significantly affect the community structure of AOA but did induce changes in that of AOB. In detail, biochar significantly enhanced the relative abundance of the dominant cluster Nitrosospira, whereas gypsum led to a notable increase in the relative abundance of unclassified_o_Nitrosomonadales. The Shannon index of AOB had a significant negative correlation with soil TOC, TN, and NH4+ content, and soil pH was the first primary environmental factor that affected the AOB community structure. In conclusion, biochar and gypsum inhibited nitrification by suppressing the activities of AOB and changed the diversities and community structure of AOB by altering related soil properties.

1. Introduction

Nitrification, which connects mineralization and denitrification, is the central link of the soil nitrogen cycle and directly or indirectly affects soil nitrogen availability, groundwater contamination, and greenhouse gas emissions [1]. Meanwhile, nitrification is a microbial aerobic process that oxidizes ammonia to nitrate and consists of two continuous processes including ammonia oxidation and nitrite oxidation [2]. In this two-step process, ammonia oxidation is catalyzed by ammonia-oxidizing archaea (AOA) and bacteria (AOB) [3], and nitrite oxidation is facilitated by nitrite-oxidizing bacteria [4]. Specifically, the ammonia oxidation process has been proven as the rate-limiting step [5]. Therefore, the ammonia-oxidizing microorganisms have attracted more attention among the nitrifying microorganisms. In recent years, AOA and AOB have been widely studied in various soil and water environments [6]. The majority of studies conducted in soil environments have focused on arable soil without obstacle issues. However, in some arable soil with degeneration issues, it has a lower nitrogen utilization efficiency and suffers from a greater risk of nitrogen loss [7]. The ammonia-oxidizing microorganisms in such soil environments need more attention.
Soil salinization is a typical type of soil degradation and more than 1 billion hectares of soil resources are salt-affected in the world [8]; furthermore, Hsieh et al. [9] reported that approximately 50% of cultivated land would be affected by salinization in 2050. Among all the saline soils, coastal saline soil is a prototypical example and has attracted more attention from pedologists and ecologists. This is because coastal mudflat is a widely spread land type in most maritime countries and is considered an important reserved land resource [10]. In fact, more than one million hectares of mudflats have been reclaimed as farmland in recent years in eastern China [11]. In this environment, soil salinity tends to be the dominant restrictive factor. Previous studies have proved that soil salinity interferes with crop nitrogen uptake by two approaches; on one hand, it alters the migration and transformation processes of nitrogen and restrains microbial activities in soils [12,13]. On the other hand, it changes the morphology, anatomy, and even the gene expression of crops [14,15]. Therefore, the amendment of salinized soil is vital for the improvement in nitrogen use efficiency and the reduction in nitrogen loss.
Numerous studies have been conducted to alleviate soil salinization and improve soil properties. Specifically, the application of soil amendments is a crucial and effective approach [16]. Among the soil amendments, biochar is a popular material for its high adsorption, porosity, and carbon sequestration and has been widely employed in the improvement of soils [17,18]. Previous studies have proved that biochar contributes to the enhancement of soil physicochemical and biological properties [19,20]. In recent years, it has also been used in the amelioration of saline soil. However, most research mainly focused on its effect on the amelioration of soil salinization and the promotion of crop nutrient uptake, and fewer studies paid attention to its effect on soil microorganisms, especially for ammonia-oxidizing microorganisms in saline soil. Furthermore, the effects of biochar on ammonia-oxidizing microorganisms varied in non-saline soils. For example, He et al. [21] reported that biochar addition promoted nitrification by improving the amoA gene copies for both AOA and AOB and changed the community structure of AOA and AOB in oxisols. However, its effects on AOA and AOB were not significant in cambisols. In addition, Bi et al. [22] found that biochar addition stimulated soil nitrification by improving the abundance and changing the community structure of AOB. However, its effect on AOA was not significant in intensive vegetable soil. On the contrary, Li et al. [23] reported that nitrification was inhibited under biochar application, which is in agreement with the findings of Wang et al. [24], while in saline soil, the soil properties, especially the salt content and ionic compositions, may differ from those of non-saline soil. The effect of biochar on nitrification and ammonia-oxidizing microorganisms may vary, necessitating further exploration.
Gypsum, a by-product of coal-fired power plants, is produced in a large amount every year in China. It is a traditional amelioration material of saline soil and has been widely used for a long time in the restoration of soil salinization [25,26]. Li et al. [27] found that gypsum resulted in a delay in the occurrence of the highest amoA gene copies in a composting experiment. Furthermore, Bossolani et al. [28] reported that amoA gene copies of AOA and AOB were both decreased after the application of gypsum in non-saline soil. While in saline soil, most studies primarily focused on the effect of gypsum on soil properties and crop growth [29], fewer studies attempted to investigate its effect on microorganisms, especially for ammonia-oxidizing microorganisms. AOA has been proven to be more adaptable to acidic environments with low-level nitrogen, whereas AOB was found to be predominant in neutral and alkaline environments with high nitrogen [30,31]. Gypsum, as an acidic soil amendment, may affect ammonia-oxidizing microorganisms by influencing soil pH, which is a key factor that can determine the predominant position of AOA and AOB. However, its effect on nitrification and ammonia-oxidizing microorganisms in coastal saline environments is still unclear.
In summary, despite the existence of some studies investigating the impacts of biochar and gypsum on nitrification and ammonia-oxidizing microorganisms, the conclusions were inconsistent. In the meantime, few studies were conducted in saline soil or saline environments. Against the backdrop of salinization, the impacts of biochar and gypsum on nitrification and ammonia-oxidizing microorganisms may differ from those in non-saline environments. It is still unclear whether the soil amendments promote or inhibit ammonia-oxidizing microorganisms, and whether ammonia-oxidizing bacteria and archaea have the same response to the soil amendments in saline soil. In our study, we conducted a one-year pot experiment with three treatments, including biochar application (BC), gypsum application (SG), and no amendment (CK), to investigate their effects on the nitrification rate, amoA gene copies, and the diversity and community structure of AOA and AOB.

2. Materials and Methods

2.1. Pot Experimental Design and Soil Collection

The soil samples used in the pot experiment were collected from the 0–20 cm soil layer in a coastal saline farmland in eastern China (32°38′42.01″ N, 120°54′8.04″ E). The soil was classified as saline alluvial soil (Fluvisols, FAO). The pot experiment included three treatments, including biochar application (BC), gypsum application (SG), and no soil amendment application (CK). Each treatment contained three duplicates. Biochar was added at rates of 10 g/kg (equivalent to 15 t ha−1), and gypsum was added at rates of 3.5 g/kg (equivalent to 5 t h−1). These two amendments were thoroughly incorporated into the soil before seeding. The cropping pattern was a barley–maize rotation. The fertilizers were monoammonium phosphate and urea. After one year of cultivation, soils were collected from the 0~20 cm soil layer. In each pot, soils were collected from multiple points and thoroughly blended. One part of the soils was frozen for DNA extraction and the other part was used for the measurement of soil properties.

2.2. Determination of Potential Nitrification Rate and Analysis of Soil Properties

The potential nitrification rate (PNR) was measured using the method reported by He et al. [1]. Soil electrical conductivity (EC) and pH (1:5 w/v) were measured using pH combined and conductivity electrodes, respectively (Mettler-Toledo Ltd., Shanghai, China). Soil organic carbon (TOC) was measured using the potassium dichromate external heating method. Soil total nitrogen (TN) was measured using the Kjeldahl method (Lu 2000). Soil NH4+-N and NO3−-N were extracted using 1 M potassium chloride solution (1:10 w/v), and then they were measured using the indigo blue colorimetry and ultraviolet dual wavelength methods, respectively. The soil water content (SWC) was measured using the oven-drying method. The soil bulk density (BD) was measured using the cutting-ring method [32].

2.3. DNA Extraction and Quantitative Real-Time PCR of amoA Genes

Soil DNA was extracted using the E.Z.N.A.® Soil DNA Kit (Omega Bio-tek, Norcross, GA, USA), the DNA content and purity were checked using a NanoDrop 2000 spectrophotometer (Thermo Scientific, Wilmington, NC, USA), and then the DNA quality was checked using 1% agarose gel electrophoresis (AGE).
Primers Arch-amoAF and Arch-amoAR were selected for amplification of the AOA amoA gene, and primers amoA-1F and amoA-2R were selected for amplification of the AOB amoA gene [6]. All the quantitative samples were run on an ABI7500 Fast Real-Time PCR System (Applied Biosystems, Foster, CA, USA). The 20 μL reaction mixtures, both for AOA and AOB, contained 16.5 μL of the ChamQ SYBR Color qPCR Master Mix (2X), 0.8 μL of each primer (5 μM), and 2 μL of the template (DNA). The following thermocycling procedures were conducted: an initial denaturation at 95 °C for 5 min, and then 40 cycles of 5 s at 95 °C, 30 s at 60 °C and 58 °C for AOA and AOB, and 40 s at 72 °C. The completion of each cycle was followed by a detection step. Standard curves were generated by performing the serial tenfold dilution gradient of plasmids which contained cloned amoA genes, and the PCR amplification efficiencies were 97.81 and 98.08% for AOA and AOB, respectively.

2.4. Illumina MiSeq Sequencing and Phylogenetic Analysis

The primers used in qPCR were also used for sequencing. Then, the DNA was treated following amplification and the agarose gel electrophoresis check, before being purified using the AxyPrep DNA gel extraction kit (Axygen, Union City, CA, USA). Next, the PCR products were eluted with Tris–HCl buffer and checked again. Finally, the purified amplicons were mixed in equimolar proportions for sequencing which was conducted on an Illumina MiSeq platform (Illumina, San Diego, CA, USA).
After sequencing, the obtained sequences were subjected to quality control and optimization using Trimmomatic software v. 0.39. Operational taxonomic units (OTUs) were determined based on a 97% similarity threshold using Usearch software. The representative sequences of the main OTUs were selected for alignment in the NCBI database to identify the homologs and the closest sequences.

2.5. Statistical Analysis

The means and standard deviations were calculated using Excel 2016. The differences in each soil property between treatments and the differences in AOA (or AOB) amoA gene copies between treatments were determined using SPSS 22.0 (SPSS Inc., Armonk, NY, USA). Graphs were drawn using Origin 2018 (OriginLab Corporation, USA) and R version 4.3.1.

3. Results

3.1. Soil Physicochemical Properties

The soil physicochemical properties are shown in Table 1. The EC and BC in SG were significantly higher and significantly lower than that in CK. The pH in SG was significantly lower than that in CK, while there was no significant difference in pH between BC and CK. The TOC and TN in BC were significantly higher than those in CK, while those in SG had no significant difference with CK. The NH4+ content in BC and SG was significantly higher than that in CK. The BD in BC was significantly lower than that in CK, while that in SG had no significant difference with CK. In terms of NO3− and SWC, there were no significant differences among each treatment.

3.2. Potential Nitrification Rate and amoA Gene Copies of AOA and AOB

The PNRs in BC and SG were significantly lower than those in CK (Figure 1). For the amoA gene copies of AOA, there were no significant differences among each treatment. At the same time, there were significantly fewer AOB amoA gene copies in BC compared to CK. Similarly, SG also had fewer AOB amoA gene copies compared to CK, even if the difference was not significant. Furthermore, the correlation analysis results showed that AOB amoA gene copies had a significant and positive correlation with nitrification rate; however, there was no significant correlation between AOA amoA gene copies and nitrification rate (Table 2).

3.3. AOA and AOB Diversity

OTUs and the Shannon and Chao indexes were selected to evaluate the diversity of AOA and AOB (Figure 2). For AOA, no significant difference was found among different treatments for all diversity indexes. For AOB, there were no significant differences among different treatments for OTUs and the Chao index. Whereas the Shannon index of BC was significantly lower than that in CK, that of SG was also lower than that in CK but not significant.
Principal coordinate analysis (Pcoa) was employed to evaluate the variations in AOA and AOB communities across different treatments (Figure 3). For AOA, there was no significant separation observed among the communities in BC, SG, and CK (R = −0.045, P = 0.607). However, for AOB, a clear separation was evident between the communities in BC, SG, and CK (R = 0.44, p = 0.039). To be specific, the first two axes could explain 77.19% of the variability; the AOB communities in BC and CK were well separated from each other along the first axis, while those in SG were well separated from those in BC and CK along the second axis.

3.4. AOA and AOB Community Structure

The community compositions of AOA and AOB on the genus level are shown in Figure 4. For AOA, Nitrososphaera was the dominant cluster, and the difference in its relative abundance was not significant among different treatments. In addition, similar results were found for other clusters. For AOB, Nitrosospira was the dominant cluster, and its relative abundance in treatment BC was significantly higher than that of other treatments. Simultaneously, the relative abundance of cluster unclassified_o_Nitrosomonadales in treatment SG was significantly higher than that of other treatments. For other clusters, their relative abundance showed a slight difference among different treatments but was not significant.

3.5. Responses of Soil Microbial Community to Soil Properties

Table 3 shows the correlation coefficient between the soil properties and diversity index of AOA and AOB. For AOA, no significant correlation was found between the soil properties and OTUs and the Shannon and Chao indexes. For AOB, the OTUs and Chao index showed no significant correlation with the soil properties, while the Shannon index exhibited a significant correlation with some soil properties, such as TOC, TN, and NH4+. The three soil properties were both negatively correlated with the Shannon index.
Redundancy analysis (RDA) was employed to assess the relationship between the environmental factors and community compositions of AOA and AOB (on the genus level) (Figure 5). For AOA, the effect of soil properties on its community composition was not significant, because the difference in community compositions was not significant among different treatments. In terms of AOB, the first two axes accounted for 88.70% of the variability. The soil pH, SWC, and TOC were the primary environmental factors that affected the community structure of AOB.

4. Discussion

4.1. Effects of Biochar Addition on Soil Properties, Nitrification, and Ammonia-Oxidizing Microorganisms

Biochar, a popular material in many fields, including soil, environmental, and ecological science, has also been widely used in the field of saline soil amelioration in recent years. In this study, biochar significantly decreased the soil EC (Table 1). This result is consistent with the report of Wang et al. [33], who found through a meta-analysis that biochar addition reduces salinity in salt-affected soils, because biochar possesses a porous structure and large surface area. On one hand, it can adsorb and fix salt ions; on the other hand, it will change the soil structure by decreasing the soil bulk density and facilitating soil aggregation formation. Consequently, biochar application accelerates salt leaching and reduces soil salinity. In addition, biochar application decreased soil BD in our study, which agrees with the result reported by Blanco-Canqui et al. [34], who believed that the reduction in bulk density could be attributed to the low density of biochar. Furthermore, biochar could improve the soil aggregate structure and promote the root growth of plants, thus decreasing the soil bulk density to some extent. Meanwhile, some soil properties were significantly improved after biochar application (Table 1); for example, the TOC, TN, and NH4+ contents increased significantly. Generally, soil organic carbon and total nitrogen are defined as the vital soil factors as they influence other soil physicochemical and biological properties including soil aggregate structure, enzymatic and microbial activities, and so on [35]. The promotion effect could be attributed to the following: (1) Biochar brings in some organic carbon and nitrogen because it contains organic carbon and nitrogen. (2) Biochar facilitates crop growth, consequently increasing the residues of roots and leaves.
The potential nitrification rate in BC was significantly lower than that in CK (Figure 1), which indicated that nitrification was inhibited by biochar application. The result is in agreement with Yao et al. [36], who reported that the nitrification rate and nitrification kinetic parameters were negatively correlated with biochar application. Similarly, Zhu et al. [37] reported that nitrification was inhibited in biochar addition treatment, and the inhibitory effect of biochar on nitrification was aggravated in saline soil as large amounts of salt ions interfered with the adsorption of biochar on NH3 and NH4+. However, there are also some conclusions that differ from our findings. Bi et al. [22] found that biochar plus nitrogen treatments significantly improved the net nitrification rate relative to only nitrogen treatments on the third day of the incubation experiment. He et al. [21] discovered that biochar facilitated nitrification and even changed the nitrification patterns from the zero- to the first-order model when the biochar application rate was 22.5 Mg/ha. These inconsistent conclusions could be ascribed to variations in soil conditions, biochar properties, and addition rates [38]. For the inhibitory effect in our study, some studies believed that biochar increased soil pH due to its alkalinity, thus aggravating NH3 volatilization and decreasing the substrate of nitrification (NH4+). In addition, Yao et al. [36] reported that the salts brought into the soil by biochar should not be ignored. However, biochar did not improved soil EC in our study. Even if biochar brought some salts into the soil, it also improved the soil structure and facilitated salt elimination in the soil.
We further measured the amoA gene copies of AOA and AOB. The latter were always higher than the former (Figure 1), which is in agreement with our previous study [6]. Even if the soil sampling site was located in the coastal mudflat zone, it was usually reported that AOA dominated the nitrification because of the poor nitrogen environment. However, more farmland soil properties were obtained after reclamation and cultivation. For example, a good deal of nitrogen fertilizers were applied to the soil. Much research has proved that AOA prevailed in acidic or low-nitrogen environments, while AOB adapted to high-nitrogen or alkaline environments [39]. Therefore, there were more AOB than AOA amoA gene copies in our study. Furthermore, no significant difference in AOA amoA gene copies was found between BC and CK (Figure 1), which is consistent with the result of Shi et al. [40] who found that biochar application had no significant influence on the abundance of AOA. Zou et al. [41] reported AOA was more unresponsive to nitrogen input than AOB. The changes in nitrogen content induced by biochar application might have no significant influence on AOA. For AOB, we found that there were significantly fewer AOB amoA gene copies in BC compared to those in CK, which indicated that biochar inhibited the growth of AOB. Interestingly, the results of previous studies are inconsistent. Lin et al. [42] indicated that biochar application improved the abundance of AOB significantly. In addition, Chen et al. [43] found that biochar had a significant auxo-action on the development of AOB. They believed that this facilitating effect could be attributed to two reasons. First, biochar application increased soil pH, thus facilitating the growth of AOB. Second, biochar application improved the soil porosity and oxygen availability, thus increasing the abundance of AOB. On the contrary, Yao et al. [38] reported that the numbers of AOB amoA gene copies were negatively correlated with biochar addition. This is in line with our study. Possible explanations could be that (1) biochar decreased the utility of NH4+ because of its very high specific surface area and high adsorption capacity, or that (2) biochar contains a mass of phenols and terpene, which altered the community structure of ammonia-oxidizing microorganisms.
In addition, we found that biochar application reduced the Shannon index of AOB significantly (Figure 2). Previous studies have proved that the richness and diversity of ammonia-oxidizing microorganisms could be affected by biochar. However, the results are inconsistent. For example, Yao et al. [38] found that biochar increased the Shannon index of AOB in salt-affected soil. On the contrary, Bi et al. [22] reported that the Shannon index of AOB decreased significantly after biochar application in vegetable soil. These inconsistent results could be attributed to the difference in soil type, soil salinity and sodicity, biochar source materials, and so on. Meanwhile, biochar changed the community composition of AOB rather than AOA, and the AOB communities in BC were well separated from CK along the first axis (Figure 3). In detail, Nitrosospira was the predominant genus, which is in line with a previous discovery that Nitrosospira is frequently observed in saline environments [44]. Meanwhile, its relative abundance in BC was significantly higher than that in CK (Figure 4). Yao et al. [38] reported that biochar application reduced the relative abundance of Nitrosospira. Conversely, Shi et al. [40] found that biochar treatment had no influence on Nitrosospira in the maize season and slightly increased the relative abundance of Nitrosospira in the wheat season, which is consistent with our study. This can be attributed to the higher responsiveness of Nitrosospira to soil salinity [45], and biochar reduced soil salinity and thus promoted the development of Nitrosospira. In addition, Li et al. [46] and Zhang et al. [47] reported that Nitrosospira was positively related with soil pH. In our study, biochar slightly increased the soil pH even if the difference was not significant, as well as probably facilitating the growth of Nitrosospira. The RDA further proved that soil pH was positively correlated with AOB community (Figure 5).

4.2. Effects of Gypsum Addition on Soil Properties, Nitrification, and Ammonia-Oxidizing Microorganisms

Gypsum is a traditional ameliorant for saline soil due to its capacity to alleviate soil salinization by replacing Na+ with Ca2+ and decreasing soil pH. Bossolani et al. [28] indicated that gypsum led to a small increase in soil properties, and only several soil indexes increased. A similar result was found in our study, where only soil EC and pH in gypsum-treated soil differed significantly to those in control soil. In detail, the soil pH decreased significantly under gypsum application, which is consistent with that reported by Zhao et al. [29], who found that gypsum decreased soil pH significantly across the 0–50 cm soil layers at harvest. This is because gypsum application reduced the ionic concentration of Na+, CO32−, and HCO3− by forming CaCO3, thus alleviating the soil alkalinity [48]. At the same time, the soil EC increased significantly after gypsum application, which is in agreement with the result of Amezketa et al. [49], who attributed this result to the dissolution of gypsum.
Nitrification was also inhibited under gypsum application, which was demonstrated by the result that the potential nitrification rate in SG was significantly lower than that in CK (Figure 1). In addition, Zhu et al. [37] found that gypsum-treated soil had higher NH4+ and lower NO3− contents, which means that gypsum application inhibited nitrification and the inhibitory effect was aggravated with the increase in the rate of gypsum use. In our study, we further analyzed the amoA gene copies of AOA and AOB. The difference in AOA amoA gene copies was not significant between SG and CK, which means that gypsum had no significant influence on AOA. Zhu et al. [6] indicated that soil salinity had a significant inhibitory effect on ammonia-oxidizing microorganisms. In our study, gypsum application significantly increased the soil EC that might inhibit the activity of AOA. Meanwhile, gypsum significantly decreased the soil pH that might promote the activity of AOA due to their preference for acidic or low-nitrogen environments [40]. The positive and negative effects offset each other and thus neutralized the effect of gypsum on AOA. For AOB, there were significantly fewer amoA gene copies in SG compared to those in CK (Figure 1), which indicated that gypsum had an inhibitory effect on the activity of AOB. Bossolani et al. [28] also reported that gypsum application reduced the amoA gene copies of AOB. This could be attributed to the changes in soil chemical properties induced by gypsum application. First, gypsum enhanced soil EC and thus inhibited the activity of AOB. Second, AOB prefers high-nitrogen or alkaline environments [40]. However, soil pH decreased significantly after gypsum application. In other words, gypsum suppresses the activity of AOB by affecting soil properties, thus inhibiting the nitrification rate.
Furthermore, we found that gypsum reduced the Shannon index of AOB to some extent, rather than AOA, even if the difference was not significant (Figure 2). In our study, the NH4+ content in SG was significantly higher than that in CK. In addition, the TOC and TN contents in SG were slightly higher than those in CK. The correlation analysis showed that the Shannon index of AOB exhibited a remarkable negative correlation with TOC, TN, and NH4+ (Table 3), which indicated that gypsum reduced the Shannon index of AOB through its influence on soil properties. Meanwhile, gypsum changed the community composition of AOB rather than AOA, and the AOB communities in SG were well separated from CK along the second axis (Figure 3). In detail, the relative abundance of unclassified_o_Nitrosomonadales in SG was significantly higher than that in CK (Figure 4). Yao et al. [31] and Egbeagu et al. [50] reported that the relative abundance of unclassified_o_Nitrosomonadales was significantly negatively correlated with soil pH. The RDA result indicated that soil pH is the most important factor affecting the AOB community structure (Figure 5). In other words, gypsum significantly decreased soil pH and thus promoted the growth of unclassified_o_Nitrosomonadales.

5. Conclusions

Biochar and gypsum application both resulted in alterations to the soil properties; in detail, biochar decreased soil EC and increased soil TOC, TN, and NH4+ content, and gypsum decreased soil pH and increased soil EC and NH4+ content. Biochar and gypsum both had an inhibiting effect on nitrification and AOB amoA gene copies, whereas they had no significant effect on AOA amoA gene copies. Biochar had no significant effect on the diversity indexes of AOA, but it significantly reduced the Shannon index of AOB. Meanwhile, gypsum had no significant influence on the diversity indexes for both AOA and AOB. Biochar and gypsum did not significantly affect the community structure of AOA, while both amendments induced changes in the community structure of AOB. In detail, biochar significantly enhanced the relative abundance of the dominant cluster Nitrosospira, whereas gypsum led to a notable increase in the relative abundance of unclassified_o_Nitrosomonadales. The Shannon index of AOB had a significant negative correlation with soil TOC, TN, and NH4+ content; soil pH was the first primary environmental factor that affected the AOB community structure. In conclusion, biochar and gypsum inhibited nitrification by suppressing the activities of AOB. In addition, its diversities and community structure were changed, which were induced by the alteration of soil properties resulting from the application of biochar and gypsum.

Author Contributions

Conceptualization, H.Z. and R.Y.; methodology, H.Z.; formal analysis, H.Z. and Y.L.; investigation, R.Y.; writing—original draft preparation, H.Z.; writing—review and editing, H.Z. and Y.L.; supervision, H.Z. and R.Y. All authors have read and agreed to the published version of the manuscript.

Funding

The research is supported by the National Natural Science Foundation of China (No.42207414; 32271720), and the Engineering Research Center of Ecology and Agricultural Use of Wetland, Ministry of Education (KF202115).

Data Availability Statement

The data are available from the corresponding author on reasonable request.

Conflicts of Interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

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Agronomy 14 01756 g001
Figure 1. Potential nitrification rate and amoA gene copies of AOA and AOB. (a) Different letters denote significant differences (p < 0.05) between treatments. (b) The capital letters denote a difference in AOA abundance between treatments (p < 0.05); the lowercase letters denote a difference in AOB abundance between treatments (p < 0.05).
Figure 1. Potential nitrification rate and amoA gene copies of AOA and AOB. (a) Different letters denote significant differences (p < 0.05) between treatments. (b) The capital letters denote a difference in AOA abundance between treatments (p < 0.05); the lowercase letters denote a difference in AOB abundance between treatments (p < 0.05).
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Figure 2. The diversity indexes of AOA (a) and AOB (b) among different treatments. Different letters denote significant differences (p < 0.05) between treatments.
Figure 2. The diversity indexes of AOA (a) and AOB (b) among different treatments. Different letters denote significant differences (p < 0.05) between treatments.
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Figure 3. Principal coordinate analysis of ammonia-oxidizing archaea (AOA) (a) and bacteria (AOB) (b) communities.
Figure 3. Principal coordinate analysis of ammonia-oxidizing archaea (AOA) (a) and bacteria (AOB) (b) communities.
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Figure 4. Relative abundance of major taxonomic groups of AOA (a) and AOB (b) at the genus level.
Figure 4. Relative abundance of major taxonomic groups of AOA (a) and AOB (b) at the genus level.
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Figure 5. Redundancy analysis (RDA) of the relationship between environmental factors and community compositions of AOA (a) and AOB (b).
Figure 5. Redundancy analysis (RDA) of the relationship between environmental factors and community compositions of AOA (a) and AOB (b).
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Table 1. Soil physicochemical properties for all the treatments.
Table 1. Soil physicochemical properties for all the treatments.
EC
(μs cm−1)		pHTOC
(g kg−1)		TN
(g kg−1)		NH4+
(mg kg−1)		NO3−
(mg kg−1)		SWC
(g g−1)		BD
(g cm−3)
BC459.43 ± 27.54 c9.10 ± 0.04 a11.25 ± 0.97 a0.61 ± 0.03 a3.79 ± 0.36 a6.55 ± 1.14 a0.27 ± 0.02 a1.21 ± 0.02 b
SG646.33 ± 14.94 a8.71 ± 0.06 b8.82 ± 0.28 b0.58 ± 0.01 ab3.55 ± 0.27 a5.63 ± 0.48 a0.24 ± 0.01 a1.25 ± 0.02 a
CK515.67 ± 20.79 b8.99 ± 0.06 a8.47 ± 0.40 b0.56 ± 0.01 b2.67 ± 0.20 b6.27 ± 0.93 a0.25 ± 0.00 a1.28 ± 0.01 a
Different letters denote significant differences (p < 0.05) between treatments.
Table 2. Pearson correlation between potential nitrification rate and amoA gene copies.
Table 2. Pearson correlation between potential nitrification rate and amoA gene copies.
PNRAOAAOB
PNR1
AOA0.5831
AOB0.707 *0.5971
* denotes that the Pearson correlation is significant (p < 0.05).
Table 3. Pearson correlations among soil properties and diversity indexes of AOA and AOB.
Table 3. Pearson correlations among soil properties and diversity indexes of AOA and AOB.
Soil PropertiesAOAAOB
OTUsShannonChaoOTUsShannonChao
EC−0.348−0.142−0.0670.0820.0110.104
pH0.368−0.0950.0670.094−0.0340.075
TOC0.025−0.2030.127−0.127−0.726 *−0.165
TN0.101−0.4770.2350.036−0.859 *0.005
NH4+−0.276−0.241−0.074−0.094−0.690 *−0.15
NO3−0.435−0.2960.08−0.019−0.4360.022
BD0.322−0.3970.1440.261−0.4720.231
SWC0.1720.1780.1080.2760.6650.333
* denotes that the Pearson correlation is significant (p < 0.05).
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Zhu, H.; Liu, Y.; Yao, R. Impacts of Biochar and Gypsum on Ammonia-Oxidizing Microorganisms in Coastal Saline Soil. Agronomy 2024, 14, 1756. https://doi.org/10.3390/agronomy14081756

AMA Style

Zhu H, Liu Y, Yao R. Impacts of Biochar and Gypsum on Ammonia-Oxidizing Microorganisms in Coastal Saline Soil. Agronomy. 2024; 14(8):1756. https://doi.org/10.3390/agronomy14081756

Chicago/Turabian Style

Zhu, Hai, Yuxing Liu, and Rongjiang Yao. 2024. "Impacts of Biochar and Gypsum on Ammonia-Oxidizing Microorganisms in Coastal Saline Soil" Agronomy 14, no. 8: 1756. https://doi.org/10.3390/agronomy14081756

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