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Article

Study on Microbial Community Structure and Soil Nitrogen Accumulation in Greenhouse Vegetable Fields with Different Planting Years

by
Luzhen Li
1,
Changsheng Zhao
1,*,
Qingfeng Chen
2,
Ting Liu
1,
Lei Li
1,
Xuzhen Liu
1 and
Xiaokai Wang
1
1
Shandong Analysis and Test Center, Qilu University of Technology (Shandong Academy of Sciences), Jinan 250014, China
2
College of Geography and Environment, Shandong Normal University, Jinan 250300, China
*
Author to whom correspondence should be addressed.
Agronomy 2022, 12(8), 1911; https://doi.org/10.3390/agronomy12081911
Submission received: 1 June 2022 / Revised: 6 August 2022 / Accepted: 11 August 2022 / Published: 14 August 2022
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Abstract

:
Soil microbial communities are an important part of the soil ecosystem in greenhouse vegetable fields, where ammonia-oxidising microorganisms play a key role in nitrogen conversion. The health and stability of the ecological environment of greenhouse vegetable fields are affected by the number of years of continuous cultivation. We used real-time PCR amplification and 16S rRNA gene sequencing to analyse the changes in soil microbial community structure and diversity in different planting years (0, 3, 9, and 13). The content of environmental factors increased with the increase of planting year; the NO3−-N content in the 0–20 cm soil layer showed a cumulative trend, peaking to 1290–1390 mg/kg in year 13. The abundance of operational taxonomic units (OTUs) in the microbial community gradually decreased, and the OTUs of 0–20 cm soil layer in year 13 decreased by 52.2% compared to year 0. The Shannon and Simpson indices indicated a substantial decrease in microbial diversity after year 9. The dominant phyla in the soil microbial community mainly included Firmicutes (23.6%), Actinobacteria (23.2%), Proteobacteria (17.6%), Crenarchaeota (83.4%), and Euryarchaeota (2.7%). Nitrosopumilus and Nitrososphaera in the ammonia-oxidising archaeal (AOA) community and Nitrolancea and Nitrospira in the ammonia-oxidising bacterial (AOB) community dominated the ammonia-oxidising microorganisms. With the increase in planting years in greenhouse vegetable fields, the structure of soil microbial community changed significantly, with soil biomass and diversity significantly decreasing in years 9 and 13. Reasonable fertilization and planting year would improve microbial activity and provide a basis for sustainable utilization and high-quality production in greenhouse vegetable fields.

1. Introduction

The facility vegetable planting method has been developing rapidly in China, providing more than 4 million ha of cultivated area in 2021. Shandong Province is the main vegetable producing area, comprising 10% of the vegetable planting region in China. Furthermore, the region with greenhouse vegetable fields covers about 25% of the nation, and is especially concentrated in Shouguang City, the birthplace of greenhouse vegetable fields and the largest vegetable distribution centre in China [1]. However, 1500–3000 kg N/ha is applied to greenhouse vegetable fields, which is four to six times higher than the amount used in other crops. Nevertheless, the average utilization rate is less than 20% [2]. Owing to the enclosed environment, high fertilizer application, relatively homogeneous planting methods, and frequent irrigation, these systems may cause environmental problems such as NO3−-N leaching, water pollution, and soil quality degradation with prolonged cultivation and lead to changes in the community structure, abundance, and function of soil microorganisms, which seriously hinders the sustainable use of soil [3,4].
The main processes of the nitrogen cycle include ammonification, nitrification, and denitrification, all of which are driven by specific functional microorganisms [5]. Ammonia-oxidising microorganisms regulate the process of ammonia oxidation. These microorganisms are an important part of greenhouse vegetable field systems and are involved in nitrogen conversion [6,7,8,9]. In agricultural production, nitrogen fertilizer plays a critical role in maintaining high-quality and healthy crops. Studies on short-term and long-term vegetable cultivation patterns revealed that the amount of ammonia-oxidising bacteria (AOB) increases significantly with increasing nitrogen application, and AOB dominate over ammonia-oxidising archaea (AOA) in the nitrification process. Furthermore, the denitrification rate is significantly higher in greenhouse vegetable field systems than in other cultivation patterns and results in increased release of N2O [10,11]. Long-term fertilization significantly affects ammonia monooxygenase subunit A (amoA) gene diversity and abundance in bacterial communities; however, small populations of AOB and denitrifying bacteria that play an important role in the nitrogen cycle are rarely reported [12,13]. In addition, domestic and foreign scholars found that factors such as soil water content, temperature, temporal and spatial changes, and nitrogen fertilizer application can lead to changes in microbial community structure and abundance [14,15,16,17,18,19,20]. Soil water content significantly affects the growth of microbial community, such as ammonia-oxidising microorganisms and denitrifying bacteria, and elevated water content leads to the increase in functional genes [5]. With increasing number in planting years in greenhouse vegetable fields, the variability between soil physicochemical properties and other agricultural soils increases, which also leads to significant differences in functional microorganisms, such as AOB and denitrifying bacteria [21]. At present, a large number of studies mainly focus on the analysis of physicochemical properties, microbial quantity, and enzyme activity. The research on the change of soil microbial community diversity is still weak. Moreover, the reported research results show inconsistent conclusions due to the soil conditions in different regions, especially with the extension of planting years of greenhouse vegetable fields, the dynamic change characteristics of soil microbial community structure are not clear.
The present study investigated the impact of the number of planting years on the structure and diversity of soil microbial community in greenhouse vegetable fields using real-time PCR (qPCR) amplification and 16S rRNA gene sequencing technology. The aim of the study was to explore the intrinsic causes of soil quality degradation in order to preserve a healthy and stable soil ecological environment and provide a theoretical foundation for sustainable use of soil, soil microbial community stability, and high-quality production of vegetables in greenhouse vegetable fields.

2. Materials and Methods

2.1. Sample Collection and Determination of Physicochemical Properties

The soil used in the study was collected from the greenhouse vegetable field subjected to high-intensity fertilization and located in Shouguang City, Shandong Province, China (34°7′24′′ N, 113°7′24′′ E). The soil type is moist soil, and cucumber and balsam pear are planted for a long time. The fertilizer applied in the soil of greenhouse vegetable fields was compound fertilizer and organic fertilizer mainly made of chicken manure and rice husk. Soil samples were collected on 20 August 2021. Mixed samples of different soil layers were collected using the plum blossom distribution method from the surface layer (0–20 cm, GSS) and deep layer (20–40 cm, GDS) of greenhouse vegetable fields that were cultivated for years 0, 3, 9, and 13. Each sample was repeated three times.
Soil pH was determined by an acidometer, and electrical conductivity (EC) was determined by a DDBJ-350 conductivity meter. NH4+-N and NO2−-N concentrations were determined using the KCl solution extraction-UV spectrophotometer method. The concentration of NO3−-N was determined using ion chromatography. The soil samples were extracted and filtered with 0.5 M NaHCO3 solution, and the available phosphorus (Avail P) was determined using the Mo-Sb anti-spectrophotometric method. Available potassium (Avail K) was determined using ICP-AES. To determine soil organic matter (SOM) content, a K2Cr2O7-H2SO4 solution was added to the air-dried soil sample for heating and digestion and then titrated using 0.1 M (NH4)2Fe(SO4)2 solution after digestion. SOM content was calculated from the volume of (NH4)2Fe(SO4)2 [22,23].

2.2. DNA Extraction and qPCR Amplification

Soil DNA was extracted using a DNA kit (Omega Bio-Tek, Norcross, GA, USA) according to the manufacturer's instructions. DNA concentration and purity were detected using a micro UV-VIS spectrophotometer (NanoDrop 2000; Thermo Fisher Scientific, Waltham, MA, USA). PCR amplification was conducted using the PCR amplification primers and circulation conditions of genes (Table 1) in an ABI GeneAmp® 9700 thermal cycler (Applied Biosystems, Foster City, CA, USA).

2.3. Analysis of 16S rRNA High-Throughput Sequencing

The V3–V4 region of the bacterial and V4–V5 region of the archaeal 16S rRNA gene were amplified by paired-end sequencing on the Illumina MiSeq platform to establish MiSeq libraries. The obtained sequences were subjected to operational taxonomic unit (OTU) clustering, denoising, and de-chimerisation to form a complete OTU species abundance profile.

2.4. Data Processing and Statistical Analysis

Data were processed and plotted using SPSS 25.0 (IBM Corp., Armonk, NY, USA), Excel 2010, and Origin 2018 (OriginLab, Northampton, MA, USA) software for statistical analysis. QIIME2 core-diversity plug-in was used for colony diversity analysis. The R package “vegan” and redundancy analysis method were used to describe the association between microbial communities and relevant environmental factors.

3. Results

3.1. Changes in Soil Physicochemical Properties at Different Planting Years

The results showed that pH, EC, NH4+-N, NO3−-N, NO2−-N, Avail P, Avail K, and SOM in the soil increased with increasing planting years, reaching the highest values in years 9 or 13 (Table 2). The values of all the physicochemical parameters, except pH, were higher in GSS than in GDS in years 3, 9, and 13. Avail P content increased approximately six-fold in GSS. The NO3−-N content was higher than that of NO2−-N, and the NO3−-N content of GSS13 reached 1290–1390 mg/kg.

3.2. Soil microbial Community at Different Planting Years

3.2.1. Changes in the Abundance and Diversity of Soil Bacterial and Archaeal Communities

The data in Table 3 showed that the abundance of bacteria-associated OTUs significantly decreased with increasing planting year; the OTUs of GSS0 was 1430. The abundance of archaea in the soil was significantly lower than that of the bacteria. The OTUs of GSS13 and GDS13 decreased by 52.2% and 60.3%, respectively, when compared to their values at year 0.
The Shannon and Simpson indices showed that soil microbial community diversity was the highest in year 0 and the lowest in years 9 or 13; the Shannon indices of bacteria and archaea decreased by 21.1% and 67.4%, respectively, compared to year 0. This confirmed that continuous cultivation decreases the richness and diversity of soil microbial community, reaching their lowest values in year 9 for bacterial and year 13 for archaeal communities. In conclusion, microbial richness and diversity in greenhouse vegetable fields significantly varied with planting years, which has indirectly affected the nitrogen cycle.

3.2.2. Bacterial Community Structure

The sequencing analysis results of dominant bacteria in the soil microbial community of the greenhouse vegetable field at the phylum, class, and genus levels in different planting years are shown in Figure 1. The dominant phyla comprised Firmicutes, Actinobacteria, Proteobacteria, Chloroflexi, Gemmatimonadetes, and Acidobacteria (Figure 1a). Firmicutes in GSS9 and GDS9 accounted for 28.7% and 46.4% of the phyla, respectively, which values were higher than those in other planting years. The percentage of Actinobacteria in GSS9 (50.6%) was significantly higher than that in GDS9 (19.1%). The percentage of Chloroflexi in GDS was significantly higher than that in GSS, except in year 0.
The dominant bacterial classes included Bacilli, Actinobacteria, Alphaproteobacteria, Gammaproteobacteria, Chloroflexia, and Gemmatimonadetes (Figure 1b). The total number of the six dominant classes in GDS decreased as the increase of planting year. The number of the six dominant classes in GDS significantly decreased compared with that in GSS, except for year 0.
The dominant bacterial genera consisted of Bacillus, Streptomyces, RB41, Truepera, Paenibacillus, Thermomonospora, and Nitrolancea (Figure 1c). The number of dominant genera in GSS first increased and then decreased with an increase in planting year, accounting for about 61% of the genera in GSS9. However, the dominant genera in GDS decreased and then increased in the same period. The genus Bacillus was the most dominant, accounting for 5.2–19.3% of the genera.

3.2.3. Archaeal Community Structure

The sequencing analysis results of dominant archaea in the soil microbial community of the greenhouse vegetable field at the phylum, class, and genus levels in different planting years are shown in Figure 2. Crenarchaeota and Euryarchaeota were the most dominant phyla (Figure 2a), with Crenarchaeota accounting for 52.3–96.9% and Euryarchaeota for 1–8% of the archaea in the greenhouse vegetable field in different planting years. The largest number of Crenarchaeota was recorded in GSS0, while the lowest number was in GDS9; the largest number of Euryarchaeota was detected in GDS0.
The dominant classes of archaea included Thaumarchaeota in Crenarchaeota and Thermoplasmata in Euryarchaeota (Figure 2b). Thaumarchaeota accounted for 51.3% to 96.8% of the archaea in different planting years, whereas Thermoplasmata accounted for relatively small and was almost absent at year 13.
The dominant genus was Nitrososphaera (Figure 2c), belonging to Crenarchaeota, accounting for 61.5–93.7% of the total archaea in the plot. The abundance of other genera showed a large variation in different years and soil layers.

3.3. Soil AOA and AOB Communities

The analysis of soil ammonia-oxidising microbial community composition in the greenhouse vegetable field (Figure 3) showed that the predominant genera of the AOA community were Nitrosopumilus of Thaumarchaeota and Nitrososphaera of Crenarchaeota, whereas the predominant genera of the AOB community were Nitrolancea, Nitrospira, Azospirillum, Anammox, Nitrosospira, and Nitrosomonas; the number of these genera varied greatly among years and soil layers.
Nitrososphaera accounted for more than 95% of the microbial communities in different planting years and soil layers and played an important role in the soil nitrogen transformation process. The proportion of Nitrolancea in the AOB community increased and that of Nitrospira decreased with the increase in planting year; both genera were dominant in the soil samples.

3.4. Clustering Analysis of Species Abundance in Soil Microbial Community

The cluster heat map analysis of the top 20 genera by relative abundance in the bacterial community resolved GSS3, GSS9, GSS13, GDS3, and GDS13 into one group and GSS0, GDS0, and GDS9 into another group (Figure 4a). Except for Microbacterium, Brevibacterium, Symbiobacterium, and Nocardioides, which were relatively abundant in GDS9, the relative abundance of bacteria was higher in GSS than in GDS. Pseudomonas was relatively abundant in both GSS0 and GDS0, whereas Steroidobacter was more abundant in GSS3 and GSS13.
In the cluster heat map of the top 20 genera based on their relative abundance in the soil archaeal community, each genus was clustered into individual categories, except for GDS3 and GDS13, which formed a single clade (Figure 4b). In GDS, the most abundant genera of GDS0 and GDS9 were Halococcus and Pilimelia, respectively. There were almost no abundant archaeal genera among different years, indicating a significant difference in soil microbial community between different years. These results suggested that the increase in planting year substantially changed the structure of soil microbial community in greenhouse vegetable fields.

3.5. Correlation Analysis of Soil Environmental Factors and Planting Years

In the RDA, the correlation between soil environmental factors and microbial community structure explained 46.0% of the total variation in microbial community structure (Figure 5a). RDA axis 1 clearly differentiated GSS0 and GDS0 from other years, and axis 2 separated GDS9 from other years. The soil environmental factors affecting the structure of bacterial microbial community were ranked as follows: pH > NO2−-N > NH4+-N > Avail P > SOM > EC > NO3−-N > Avail K. Thus, pH, NO2−-N, NH4+-N, and Avail P were the main environmental factors influencing the changes in bacterial structure.
High-throughput sequencing and RDA analysis of archaeal communities showed that environmental factors could explain 47.3% of the variation in microbial community structure (Figure 5b). GSS13 was clearly differentiated from other years along RDA axis 1 or axis 2, but the separation or aggregation effect was not obvious in other years. The results indicated a positive correlation among soil environmental factors, with EC, NO3−-N, NH4+-N, and Avail P being the key factors influencing the changes in archaeal community structure.

4. Discussion

Microorganisms play a vital role in the soil environment, especially in terms of nutrient and organic matter cycling and soil fertility [26]. Ammonia-oxidising microorganisms in soil drive nitrogen cycling and nitrification [27,28]. Research on the correlation of AOA and AOB with soil potential nitrification potential in acidic soil revealed that AOA had a dominant role in ammonia oxidation and AOB are important in the process of nitrification [29,30,31,32].
The present study showed that the environmental parameters other than pH were greater in GSS than in GDS, except for NH4+-N, Avail K, and SOM in year 0, and most of the indicators increased with the increase in planting year. The content of NH4+-N, NO3−-N, and NO2−-N decreased in the order of NO3−-N > NO2−-N > NH4+-N, which corresponds to the nitrogen conversion process in which nitrogen is transformed into NO3−-N available for crop absorption or converted into N2O gas. However, the high NO3−-N content indicates that NO3−-N was substantially accumulated during the long-term cultivation of vegetables and could easily leach out and pollute water. The physicochemical properties in year 13 differed significantly from those in other years, which may be caused by altered plot conditions and long-term human activities. In addition, inorganic and organic fertilizers in the soil were hydrolysed by a variety of enzymes into mineral nitrogen that could be absorbed and utilized by crops. However, a large amount of nutrients remained in the soil during the long-term planting process. The planting years were negatively correlated with the richness of bacterial OTUs, and the bacterial richness was significantly higher than that of archaea. The soil bacterial richness and diversity were significantly reduced after year 9, as indicated by the Shannon and Simpson indices. A similar trend in microbial community monoculture was reported by Meng et al. [33] and Wang et al. [34] after a long-term vegetable crop succession. It might be due to continuous crop barriers that limit the sustainable and healthy development of the soil microbial community in greenhouse vegetable fields. Liu et al. [35] determined that crop disorders can cause microbial mortality in greenhouse vegetable fields mainly due to the increase in pathogenic bacteria and the self-toxicity of crops.
The patterns of change in community structure and abundance of bacteria, AOB, archaea, and AOA were not consistent in this study. At the bacterial community phylum level, Firmicutes, Actinobacteria, and Proteobacteria were dominant, which was consistent with the results reported by Li et al. [36]. The number of bacteria in GSS first increased then decreased with the increase in planting year, peaking at year 9. The results were consistent with the trend of bacterial changes in greenhouse vegetable fields observed by Wang et al. [21]. At the bacterial community class level, Bacilli and Actinobacteria, which accounted for a larger proportion of bacteria, did not exhibit an obvious pattern of change; however, the proportion of dominant bacteria in GSS was larger than that in GDS. At the genus level, Bacillus was the dominant genus (19.3%) in GDS0, and its number in GSS continuously increased with the increase in planting year. In addition, nitrolancea is a rod-shaped nitrifying bacterium that affect the process of nitrification-denitrification.
In the archaeal communities of soils, Crenarchaeota and Euryarchaeota were two main phyla initially discovered by scientists studying archaeal rRNA gene sequences. The Crenarchaeota have a facilitative role in nitrification and increase the rate of soil nitrification [37,38]. In addition to participating in the carbon cycle (anaerobic methane oxidation), Hadesarchaea reduce nitrite in the nitrogen cycle [39]. At the class level, the percentage of Thaumarchaeota in the soil’s archaeal community in different years and soil layers was above 50%, and present in the highest proportion in GSS0, which may be related to human activities. The high fertilizer input in greenhouse vegetable fields increases the nutrient content of the soil, providing a suitable environment for the survival and reproduction of microorganisms under high temperature and high humidity conditions. At the genus level, Nitrososphaera accounted for more than 60% of the archaeal community, and AOA were the main contributor to nitrification, mainly through increased abundance of individual OTUs of Nitrososphaera. Furthermore, the correlation between AOA and nitrification suggests that Nitrososphaera is inextricably linked to the nitrogen cycle [21,40]. In addition, Li et al. [41] found that Nitrososphaera is the main genus promoting nitrification in acidic soils, confirming the importance of Nitrososphaera for nitrogen transformation.
The cluster analysis of the soil microbial community revealed that more genera were present in GSS than in GDS, and the microbial community varied among different years and soil layers. This suggests that most microorganisms can survive in surface soil, and cultivation over many years will change the structure of the microbial community in greenhouse vegetable fields. According to the RDA of environmental elements, pH was the most important element altering the structure of soil bacterial community, which is consistent with the findings by Chang et al. [42] who determined that environmental elements such as pH and temperature seriously influence soil microbial activity.

5. Conclusions

As the planting year in greenhouse vegetable fields increased, soil microbial community structure and diversity changed, the abundance and diversity of bacterial OTUs significantly decreased, and the community structure varied and became unstable. The soil environmental factors were positively correlated with planting year and showed a cumulative trend. NH4+-N and Avail P were the common key factors affecting community structure and abundance of bacteria and archaea, whereas pH was the most important factor responsible for the change in bacterial structure. The dominant genera in the soil microbial community, Bacillus and Nitrososphaera, were particularly prominent. Among ammonia-oxidising microbial communities, the abundance of Nitrosopumilus and Nitrososphaera in the AOA community differed among planting years and soil layers, and the number of Nitrolancea in the AOB community was proportional to the planting year. In conclusion, for greenhouse vegetable fields continuously cultivated for more than 9 years, reasonable fertilization is recommended to improve soil quality, microbial activity, and diversity for sustainable utilization and high-quality production.

Author Contributions

Conceptualization, L.L. (Luzhen Li) and C.Z.; methodology, L.L. (Luzhen Li) and C.Z.; software, L.L. (Lei Li) and X.W.; validation, Q.C., T.L., and L.L. (Lei Li); formal analysis, L.L. (Luzhen Li) and T.L.; investigation, L.L. (Luzhen Li), C.Z. and Q.C.; resources, Q.C.; data curation, L.L. (Lei Li) and X.L.; writing—original draft preparation, L.L. (Luzhen Li); writing—review and editing, C.Z. and Q.C.; visualization, L.L. (Lei Li); supervision, X.L. and X.W.; funding acquisition, C.Z. and Q.C. All authors have read and agreed to the published version of the manuscript.

Funding

This work was funded by the National Natural Science Foundation of China [Qilu University of technology (Shandong Academy of Sciences), grant number 41877041], the National Natural Science Foundation of China [Shandong Normal University, grant number 42077051] and Qilu University of Technology (Shandong Academy of Sciences) science, education and industry integration innovation pilot project [Qilu University of technology (Shandong Academy of Sciences), grant number 2020KJC-ZD13].

Data Availability Statement

Not applicable.

Acknowledgments

We are grateful to the Shandong Provincial Analysis and Test Center for the experimental platform and the subject staff for their valuable comments on this study.

Conflicts of Interest

The authors declare no conflict of interest.

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Figure 1. Dominant bacteria in soil microbial communities at the phylum (a); class (b); genus (c) levels at different years of cultivation.
Figure 1. Dominant bacteria in soil microbial communities at the phylum (a); class (b); genus (c) levels at different years of cultivation.
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Figure 2. Dominant archaea in soil microbial communities at the phylum (a); class (b); genus (c) levels at different years of cultivation.
Figure 2. Dominant archaea in soil microbial communities at the phylum (a); class (b); genus (c) levels at different years of cultivation.
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Figure 3. Composition of soil ammonia-oxidising archaea (a) and ammonia-oxidising bacteria (b) in microbial communities at different facility planting years.
Figure 3. Composition of soil ammonia-oxidising archaea (a) and ammonia-oxidising bacteria (b) in microbial communities at different facility planting years.
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Figure 4. Cluster heat map analysis of species abundance of soil microbial community at different planting years. (a) the cluster heat map analysis of the top 20 genera by relative abundance in the bacterial community; (b) the cluster heat map analysis of the top 20 genera by relative abundance in the archaeal community.
Figure 4. Cluster heat map analysis of species abundance of soil microbial community at different planting years. (a) the cluster heat map analysis of the top 20 genera by relative abundance in the bacterial community; (b) the cluster heat map analysis of the top 20 genera by relative abundance in the archaeal community.
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Figure 5. Correlation analysis between soil environmental factors and the structure of bacterial microbial community. (a) RDA analysis of bacterial communities; (b) RDA analysis of archaeal communities.
Figure 5. Correlation analysis between soil environmental factors and the structure of bacterial microbial community. (a) RDA analysis of bacterial communities; (b) RDA analysis of archaeal communities.
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Table
Table 1. Amplification primers and reaction conditions of real-time fluorescence-based quantitative PCR.
Table 1. Amplification primers and reaction conditions of real-time fluorescence-based quantitative PCR.
Target Group Primer NameSequence (5′–3′)Length of Amplicon (bp)References
Archaeal amoA 1524F10extF
Arch958RmodR		TGYCAGCCGCCGCGGTAA
YCCGGCGTTGAVTCCAATT		434[24]
Bacterial amoA 2338F
806R		ACTCCTACGGGAGGCAGCAG
GGACTACHVGGGTWTCTAAT		468[25]
1 3 min at 95 °C × 1 cycle; 30 s at 95 °C, 30 s at 60 °C, 45 s at 72 °C × 35 cycles; 10 min at 72 °C, 10 °C until halted by the user; 2 3 min at 95 °C × 1 cycle; 30 s at 95 °C, 30 s at 55 °C, 45 s at 72 °C × 27 cycles; 10 min at 72 °C, 10 °C until halted by the user.
Table
Table 2. Physicochemical properties of the soil in the greenhouse vegetable fields at different planting years.
Table 2. Physicochemical properties of the soil in the greenhouse vegetable fields at different planting years.
Soil Layer (cm)TreatmentpHEC (mS/m)NH4+-N (mg/kg)NO3−-N (mg/kg)NO2−-N (mg/kg)Avail P (mg/kg)Avail K (mg/kg)SOM (g/kg)
0–20GSS05.9 c24.2 d2.1 c104.0 c20.4 c77.5 d282.0 c10.5 c
GSS36.2 b37.3 c28.1 b207.0 b56.1 b255.0 c309.0 bc46.3 b
GSS97.1 a70.1 b53.3 a281.0 b92.3 a446.0 b342.0 b58.6 a
GSS137.2 a99.9 a53.5 a1340.0 a91.9 a468.0 a972.0 a64.4 a
20–40GDS06.1 b20.1 c4.7 d73.8 b10.2 d76.5 d392.0 d14.4 c
GDS36.4 b20.7 c16.8 c83.7 b25.2 c123.0 c309.0 c18.8 b
GDS97.1 a23.3 b11.7 b88.3 b40.7 b167.0 b352.0 b24.3 b
GDS137.3 a35.2 a29.1 a366.0 a50.9 a238.0 a604.0 a20.7 a
Different letters in the same column for the same soil layer meant significant difference (p < 0.05).
Table
Table 3. Statistics of different planting years on the abundance and diversity of soil bacteria and archaea.
Table 3. Statistics of different planting years on the abundance and diversity of soil bacteria and archaea.
TreatmentOTUsChao1 IndexShannon IndexSimpson Index
BacteriaGSS011291606.4310.001.00
GSS311821461.169.231.00
GSS9681847.087.890.98
GSS13790841.008.150.99
GDS014301824.089.911.00
GDS314251694.239.761.00
GDS9786896.008.220.99
GDS13930932.278.300.99
ArchaeaGSS0134169.005.610.96
GSS389123.003.830.76
GSS986147.004.170.87
GSS136493.001.830.48
GDS0131149.005.740.96
GDS3113163.004.530.89
GDS9226257.005.900.97
GDS135268.003.610.86
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Li, L.; Zhao, C.; Chen, Q.; Liu, T.; Li, L.; Liu, X.; Wang, X. Study on Microbial Community Structure and Soil Nitrogen Accumulation in Greenhouse Vegetable Fields with Different Planting Years. Agronomy 2022, 12, 1911. https://doi.org/10.3390/agronomy12081911

AMA Style

Li L, Zhao C, Chen Q, Liu T, Li L, Liu X, Wang X. Study on Microbial Community Structure and Soil Nitrogen Accumulation in Greenhouse Vegetable Fields with Different Planting Years. Agronomy. 2022; 12(8):1911. https://doi.org/10.3390/agronomy12081911

Chicago/Turabian Style

Li, Luzhen, Changsheng Zhao, Qingfeng Chen, Ting Liu, Lei Li, Xuzhen Liu, and Xiaokai Wang. 2022. "Study on Microbial Community Structure and Soil Nitrogen Accumulation in Greenhouse Vegetable Fields with Different Planting Years" Agronomy 12, no. 8: 1911. https://doi.org/10.3390/agronomy12081911

APA Style

Li, L., Zhao, C., Chen, Q., Liu, T., Li, L., Liu, X., & Wang, X. (2022). Study on Microbial Community Structure and Soil Nitrogen Accumulation in Greenhouse Vegetable Fields with Different Planting Years. Agronomy, 12(8), 1911. https://doi.org/10.3390/agronomy12081911

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