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Article

Liming Alters Microbial Communities Affecting Nitrification in the Rhizosphere of Camellia sinensis

by
Chang Zhao
1,
Xiaoxiang He
2,
Han Jiang
1,
Xiaoyan Wang
1,
Jinjuan Gao
1 and
Chanjuan Hu
1,*
1
Institute of Geographical Science, Henan Academy of Science, Zhengzhou 450052, China
2
Institute of Resources, Environment and Soil Fertilizer, Fujian Academy of Agricultural Sciences, Fuzhou 350013, China
*
Author to whom correspondence should be addressed.
Agronomy 2026, 16(1), 77; https://doi.org/10.3390/agronomy16010077 (registering DOI)
Submission received: 25 November 2025 / Revised: 18 December 2025 / Accepted: 24 December 2025 / Published: 26 December 2025
(This article belongs to the Topic Effective Strategies for Rangeland Conservation and Sustainable Management)
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Abstract

After years of cultivation, tea garden soils gradually become acidified and compacted. Liming to ameliorate soil acidification alters the soil microbial activity and community structure, ultimately affecting the soil nitrogen cycling and nitrogen use efficiency of tea plants. In this study, ammonium-preferring tea plants (Camellia sinensis) were cultivated in lime-amended soils across a pH gradient (pH 4.5, pH 5.5, pH 6.5, and pH 7.5) to assess the impacts of soil liming on the structure and composition of the rhizosphere microbial community. The results demonstrated that, as the soil pH increased, the diversity and richness of both bacterial and fungal communities exhibited a gradual decline. The biomasses of AOA and AOB were dominated in acid soils and alkaline soils, respectively. The abundance of Proteobacteria, Actinobacteria, and Bacteroidetes in pH 6.5 and pH 7.5 were 16.56–22.56%, 16.29–18.09%, and 1.65–4.52% times higher than those in pH 4.5 and pH 5.5 soils. Ascomycota and Basidiomycota were accounting for 89.93% of all the species. Across the rising pH gradient, the relative abundance of Ascomycota significantly increased by 9.64–20.49%, whereas Basidiomycota decreased by 1.11–15.01%. The RDA analysis results showed the soil pH was the main effect factor for the differences in the structure and composition of bacterial and fungal communities. This conclusion provides theoretical support for the optimization of acidic soil improvement techniques after long-term tea cultivation.

1. Instruction

Soil microorganisms play a crucial role in nitrogen turnover, maintaining energy flow, and material cycling [1]. In soil–plant ecosystems, the soil pH and tree species alter soil physicochemical parameters, influencing the diversity and composition of bulk soil bacterial and fungal communities [2,3]. Plant roots establish a complex and resource-rich hotspot, selectively recruiting microbial communities in the rhizosphere. The uptake of ions and water by roots, combined with the exudation of carbon-rich compounds, leads to the formation of a rhizosphere where the microbial cycling of nitrogen is rapid, dynamic, and competitive with that of the bulk soil [4]. While the effects of pH and the rhizosphere environment on microbiomes have often been studied independently, comprehending their interactions is crucial for understanding the assembly and ecology of rhizosphere microbial communities.
In the acidic forest soils of subtropical humid regions, competition for inorganic nitrogen occurs between plants and microorganisms [5]. This competition drives plants to allocate more of their photosynthetic products as root exudates, thereby enhancing the rhizosphere microbial activity. Consequently, this process promotes the net nitrogen mineralization rates and satisfies the inorganic nitrogen requirements of both plants and microorganisms. Previous studies have demonstrated that, compared to bulk soils, planted soils exhibit an increase in the activity of the microbial community and the rate of nitrogen cycling [6,7,8]. Currently, the research on the effects of vegetation on the soil microbial community structure primarily focuses on comparisons between different natural ecosystems [9], such as wetlands [10], forests [11], and grasslands [12]. However, studies on the impact of plants under various soil pH conditions on the rhizosphere microbial community structure are still insufficient.
In artificially managed tea gardens, agronomic practices such as fertilization [13,14] and continuous cropping [15] can influence the stability of soil microbial communities. After years of tea plantation, the soil experiences an accumulation of exchangeable Al3+ and phenolic compounds, leading to soil acidification [16]. Short-term liming impacts are detected in soil biological processes, such as increasing the N availability for plant uptake [17]. Therefore, it offers an efficient and economical strategy for improving soil quality and nitrogen cycling [18]. Nitrification is a critical process that regulates the primary forms of inorganic nitrogen in soil, with archaeal ammonia oxidizers (AOA) and bacterial ammonia oxidizers (AOB) being the dominant microorganisms responsible for driving autotrophic nitrification. AOA exhibit a higher acid tolerance, whereas AOB are the primary ammonia-oxidizing microorganisms in nitrogen-rich soils under neutral to alkaline conditions [19,20]. Variations in archaeal and bacterial communities’ composition are considered to be related to many soil biotic and abiotic factors [21]. Liming results in changes to both the nutrient availability and edaphic characteristics potentially having substantial effects on archaeal and bacterial communities. Additionally, fungi exhibit a higher acid tolerance compared to bacteria [22]. Additionally, fungal hyphae play a crucial role in aiding plants to assimilate organic nitrogen from the soil [23]. Among the fungal groups known for their heterotrophic nitrification capabilities are Aspergillus, Penicillium, Streptomyces, Mortierella, and Trichoderma. Arbuscular mycorrhizal fungi play a critical role in plant growth because they can provide necessary nutrients and help plants resist biotic and abiotic stresses [24]. Ref. [25] found that liming could enhance AM fungal production, potentially alleviating the environmental stress to the host plants. However, the results of previous studies on how liming influences bacterial and fungal types remain inconclusive. Therefore, the potential effects of liming on the microbial community composition need further evaluation.
Tea (Camellia sinensis) is among the most significant industrial crops and economic beverage in the globe [26]. In order to increase tea production, people have been applying chemical fertilizers in large quantities for a long time, neglecting the combined application of organic fertilizers and the balance of other elements, which has led to the serious acidification of tea soil and the imbalance of nutrients and other problems. To deeply understand the impact of lime amendment on the structure and diversity of the microbial community in acidic soils, this study focused on the acidic forest soils in the subtropical humid region of China. This research examined tea plants cultivated in lime-amended soils with varying pH levels. Using quantitative real-time polymerase chain reaction (qPCR) and amplicon high-throughput sequencing, we investigated the soil microbial communities. These techniques allowed for a comprehensive analysis of the microbial community structure and composition changes. The primary objective of our study is to try to explain how liming modifies the rhizosphere and affects microbial communities, impacting the focus of nitrification among archaea and bacteria in the tea rhizosphere. This study may provide a theoretical basis for optimizing tea plant cultivation strategies and increasing production.

2. Materials and Methods

2.1. Experiment Site and Sampling

The soil samples utilized in this study were collected from the Longhushan Nature Reserve, situated within the subtropical monsoon climate zone of China. This region is characterized by an annual mean temperature of 17.5 °C and an annual precipitation of 1750 mm. The soil type in this region is red soil formed from the Quaternary red clay, and the dominant vegetation types are Chinese fir (Cunninghamia lanceolata) and Masson pine (Pinus massoniana). Three sampling areas were chosen in this design; the areas of each region is about 10 m × 10 m. The surface litter and weeds were removed, and soil samples were collected from the 0 to 20 cm depth, excluding the humus layer. The three areas’ soil samples were thoroughly mixed to form a composite sample and transferred into sterile ziplock bags. For transportation, the bags were placed in insulated sampling boxes containing ice packs to ensure continuous refrigeration. Upon arrival at the laboratory, the soil samples were passed through a 2 mm sieve. A portion of the sieved soil was used for the determination of soil chemical properties, while the remaining soil was stored at 4 °C for subsequent experimental use. The measured soil chemical properties, including soil moisture content, pH, soil organic carbon (SOC), and total nitrogen (TN), were 26.52%, 4.37, 20.07 g kg−1, and 1.09 g kg−1, respectively.
The acidic soil was amended with 100 mesh CaO (Sinopharm Chemical Reagent Co. Ltd., Shanghai, China) powder at four application rates, 0%, 0.2%, 0.4%, and 0.6% (w/w), for pH adjustment. The soil was thoroughly mixed with CaO at varying application rates, then sealed and incubated at room temperature (25 ± 1 °C) for a 14-day stabilization period. Subsequent pH measurements of the amended soils showed progressive neutralization, with recorded values of 4.5, 5.5, 6.5, and 7.5 across the treatment gradients. For this study, the tea plant cultivar “Longjing 43” with uniform growth (approximately 13–15 cm height) was selected as the experimental plant. The seedlings’ roots were meticulously washed with distilled water to eliminate the original growth medium. Subsequently, the tea was planted in four distinct soil pH treatments (dry soil weight: 200 g, and pot dimensions: 5 × 7 × 10 cm3). To ensure uniformity in ammonium and nitrate concentrations across all treatments, we used a Hoagland solution (YaJi biology, Shanghai, China) without nitrogen element containing magnesium sulfate and potassium dihydrogen phosphate, and 5 mL was added to soils every two days. This enabled the tea plants to grow in an environment where all soil factors were consistent, except for pH. Three experimental treatments were established: control (no tea plant, CK), tea cultivation for 30 days (D30), and tea cultivation for 60 days (D60). A total of 36 soil samples were obtained (3 replicates × 3 treatments × 4 pH gradient). All treatments were maintained under identical greenhouse conditions (constant temperature of 25 °C, 12 h of UV light per day, and 60% soil water-holding capacity). Every three days, all the samples were rotated 90° in a clockwise direction, to ensure the samples stayed in a uniform state and avoided growth deviations caused by factors such as light exposure. Based on the changes in soil ammonium and nitrate concentrations, as well as the growth status of the tea plants, soil samples were collected from each treatment at 30 and 60 days of cultivation. The soil samples were divided into two portions and stored at 4 °C and −80 °C, respectively, for subsequent physicochemical analysis.

2.2. Soil Property Analyses

Soil pH was measured at a 1:2.5 (soil/distilled water) ratio, stirred with a magnetic stirrer for 2 min, and then stilled 10 min. Complete the measurement with a DMP-2 mV/pH detector (Quark Ltd., Nanjing, China) within 1 h and ensure the measurement environment temperature remains at 25 °C. Soil dissolved organic carbon (DOC) content was measured in a 1:4 (soil/deionized water) ratio, and then 100 mL deionized water was added to 25 g fresh soil in 200 mL plastic bottle, and shaken on a reciprocating shaker for 30 min (280 r/min), and then transferred the extract to a centrifuge tube and centrifuge for 20 min. Filter the supernatant through a 0.45 μm filter membrane. Then, transfer the filtrate to an Analyzer Multi N/C (Analytic Jena, Jena, Germany) for determination. The concentrations of ammonium (NH4+) and nitrate (NO3−) were determined using a SAN Plus continuous flow analyzer (Skalar San++, Breda, The Netherlands).

2.3. Soil DNA Extraction and Real-Time PCR Assay

Genomic DNA was extracted from 0.5 g fresh soil according to the manufacturer’s instructions of a PowerSoil DNA Isolation Kit (MoBio Laboratories, Inc., Carlsbad, CA, USA). The DNA quality and quantity were measured by NanoDrop 2000 spectrophotometer (Thermo, Waltham, MA, USA). Quantitative real-time polymerase chain reaction (qPCR) amplifications were performed in biological triplicates on a CFX-96 thermocycler (Bio-Rad Laboratories Inc., Hercules, CA, USA). The abundance of bacteria and fungi were measured using general primer sets (338F/518R for bacteria, and ITS1F/2R for fungi, respectively) and the details of qPCR reaction were listed in Table 1. The reaction mixture for both fungi and bacteria was set according to [27] and the amplification efficiencies were 98% and 115%, respectively. The R2 value of the standard curves was 0.9984 for bacteria and 0.9982 for fungi. All samples were analyzed in triplicates to ensure accuracy.

2.4. Microbial Community Analysis by High-Throughput Sequencing

Soil microorganisms were characterized by high-throughput sequencing. The bacterial 16S rRNA V4–V5 region was amplified using the primer set 515F (5′-GTGCCAGCMGCCGCGGTAA-3′) and 907R (5′-CCGTCAATTCMTTTRAGTTT-3′). Primers ITS1F (5′-CTTGGTCATTTAGAGGAAGTAA-3′) and ITS2R (5′-GCTGCGTTCTTCATCGATGC-3′) were used to amplify the ITS region [28]. The samples were sequenced on the Illumina MiSeq PE 250 platform from Genesky Biotechnologies Inc., Shanghai, China. High-throughput raw sequences have been deposited in DDBJ/EMBL/GenBank as BioProject ID PRJNA673937.
Raw sequence data were processed and analyzed using the QIIME (Version 1.9.0, http://qiime.sourceforge.net/ (accessed on 12 April 2024)). All the sequences with low quality score (<20) or not match the primers and the barcodes were removed before downstream application. The quality-filtered sequences were clustered using 95% sequence similarity for operational taxonomic units (OTUs) based on the Greengenes 13_8 database for bacteria and the UNITE database for fungi, respectively.

2.5. Statistical Analysis

Statistical analysis was performed using IBM SPSS Statistics 22 (SPSS Inc. Chicago, IL, USA) and R × 32 (3.6.0). Differences in soil properties, gene copy numbers, and microbial alpha-diversity among samples were tested using a one-way analysis of variance (ANOVA) in IBM SPSS Statistics 19 for Windows (Armonk, NY, USA).
Principle coordinates analysis (PCoA) was performed to examine beta-diversity (Bray–Curtis distances) between individual samples by ‘vegan’ (Version 2.6-4) and ‘ggplot2’ (Version 3.3.3) packages. Analysis of similarity (ANOSIM) was used to determine the significance of community composition differences among treatments. The core bacteria and fungi were selected using Similarity Percentage (SIMPER) analysis by ‘simper’ (Version 0.8.5) package. The ‘phyloseq’ (Version 3.4.3) package was used to estimate the optimal environmental factors. The “rda” function was used to conduct Redundancy Analysis (RDA, Redundancy Analysis) between the optimal environmental factors and the significantly different microbial communities to explore the environmental factors that contribute the most to the explanatory degree of community differences.

3. Results

3.1. Effects of Liming on Soil Physicochemical Properties

Throughout the experiment of 30-day and 60-day tea cultivation, the soil moisture content was maintained at 24.56–29.78%. The soil pH across all gradients was shown to be relatively stable. However, a progressive acidification trend emerged with a prolonged planting duration, as evidenced by the gradually decreasing pH values. In CK soils, the physicochemical properties varied significantly among pH gradients (Table 2), the NH4+ concentration decreased with increasing pH, the NO3− concentration first decreased then increased with rising pH, and DOC showed the opposite trend (first increase, and then decrease). For D30 and D60 soils, the NH4+ concentration first increased and then decreased, reaching its maximum of 18.53 mg kg−1 in pH5.5. The NO3− concentration showed a significant increase in D30, ranging from 43.53 mg kg−1 to 76.38 mg kg−1, with the increasing pH. But, in D60, there was first a decrease, and then an increase, reaching its lowest concentration of 42.2 mg kg−1 in pH 5.5. The DOC content represented an increasing trend with increasing pH, increasing by approximately 25.32% and 34.65% in D30 and D60.

3.2. Effects of Liming on Soil Microbial Biomass

The soil bacterial biomass across treatments ranged between 1.61–2.41 × 1010 copies g−1 dry soil (Figure 1A). At pH 4.5 and 7.5, CK had a higher bacterial biomass than D30 and D60; at pH 5.5 and 6.5, D30 exceeded CK and D60. The fungal biomass (2.09–4.14 × 108 copies g−1 dry soil) was significantly lower than bacterial biomass (Figure 1B). D60 had a higher fungal biomass than CK and D30 across pH gradients, and the bacterial-to-fungal ratio (54.17–99.25, Figure 1C) was higher in D30 than CK and D60 at all pH except 7.5.
The AOA (ammonia-oxidizing archaea) and AOB (ammonia-oxidizing bacteria) biomasses differed significantly across pH gradients (Figure 1D,E). AOA exhibited significantly higher levels than AOB in the pH 4.5 and 5.5 treatment, but, in pH 6.5 and 7.5, the situation was reversed (p < 0.05). The AOA-to-AOB ratio was higher at pH 4.5 and 5.5 than at 6.5 and 7.5, with the tea-planted soils diminishing as the pH increased (Figure 1F).

3.3. Effects of Liming on Soil Microbial Diversity

By clustering at the 97% similarity level, 8153 bacterial and 6325 fungal OTUs were obtained from 3,236,358 16s rRNA gene sequences and 4,135,608 ITS sequences, respectively. Lime application significantly reduced the α-diversity of both bacterial and fungal communities (Figure 2A,C), with the most pronounced reduction observed in the soil without tea plant cultivation. For bacterial diversity, the indices at D30 and D60 were significantly lower in alkaline soil than in acidic soil. In contrast, the fungal diversity results indicated no significant differences at D30 and D60 after lime application.
The principal coordinate analysis (PCoA) based on the Bray–Curtis distance (β-diversity) suggested that both the bacterial and fungal community compositions were first separated by lime treatments and then by the tea planted (ANOSIM, p < 0.05, Figure 2B,D). In addition, for fungi, the community compositions revealed a significant separation between the alkaline treatments (pH 6.5 and pH 7.5) and the other two treatments (pH 4.5 and pH 5.5), while the results for bacteria were the opposite of those for fungi.

3.4. Effects of Liming on Soil Microbial Community Composition

The relative abundance of the soil bacterial and fungal community at the phylum level were showed in Figure 3. The main phyla in the bacterial communities of all soils were largely the same. Proteobacteria, Acidobacteria, Chloroflexi, Actinobacteria, and Bacteroidetes were the dominant phyla, accounting for 18.6–43.9%, 3.89–21.01%, 5.97–15.74%, 6.59–11.12%, and 0.86–19.88% of all the sequences, respectively (Figure 3A). The abundance of Proteobacteria, Actinobacteria, and Bacteroidetes in pH 6.5 and pH 7.5 were increased by 16.56–22.56%, 16.29–18.09%, and 1.65–4.52% compared to those in pH 4.5 and pH 5.5 soils. Furthermore, with the application of liming, the relative abundance of Proteobacteria, Actinobacteria, Bacteroidetes, Gemmatimonadetes, and Armatimonadetes increased, whereas that of Acidobacteria and Chloroflexi decreased.
There was a significant difference in the fungal communities in all treatments at the phylum level (Figure 3B). Ascomycota and Basidiomycota were the dominant phyla in the 12 soils, accounting for 89.93% of all the sequences, respectively. With the application of liming, the relative abundance of Ascomycota increased, whereas that of Basidiomycota, Mortierellomycota, and Mucoromucota decreased. Furthermore, the relative abundance of Ascomycota were significantly decreased with the tea planted compared with CK.
The LEfSe analysis confirmed significant bacterial compositional differences across pH gradients (Figure 4A, phylum-to-family level, p < 0.05, LDA > 4.0). In pH 6.5 and 7.5 soils, dominant families (>1%) like Chitinophagaceae, Hyphomicrobiaceae, Micromonosporaceae, Cytophagaceae, Haliangiaceae, Sphingomonadaceae, Rhodospirillaceae, and Xanthomonadaceae were enriched compared to those in pH 4.5 and 5.5 (Figure 4B). Conversely, Thermogemmatisporaceae, Koribacteraceae, and Burkholderiaceae were depleted by approximately 8.47%, 3.61%, and 2.21%, respectively. Fungal community differences were also significantly influenced by tea cultivation (Figure 4C,D; p < 0.05). With the increase in pH, the relative abundances of the fungal dominant groups Penicillium, Aspergillus, and Uc_Nectriaceae in pH 7.5 soil increased significantly by approximately 3.88, 3.67, and 2.24%, respectively, compared with those in pH 4.5 soil. The relative abundance of Uc_Orbiliaceae shows an increasing trend as the pH increases. The relative abundances of Mortierella, Talaromyces, Trichoderma, and Uc_Helotiales in pH 4.5 soil were significantly reduced by approximately 2.21, 8.34, 1.34, and 0.78%, respectively, compared to those in pH 7.5 soil (p < 0.01).

3.5. The Correlation Between Dominant Microbial Groups and Environmental Factors

The RDA analysis of the bacterial families and environmental factors (Figure 5A) revealed positive correlations of Cytophagaceae, Chitinophagaceae, Hyphomicrobiaceae, Micromonosporaceae, Haliangiaceae, and Sphingomonadaceae with pH, DOC, and NO3−, and negative correlations with NH4+. Opposite trends were observed for Rhodospirillaceae, Thermogemmatisporaceae, Koribacteraceae, and Burkholderiaceae. The soil variables explained 77.6% of the community variance (Figure 5B), with pH, NH4+, and NO3− as the primary drivers (95.34%, 56.99%, 54.54%, respectively).
The RDA analysis of the dominant fungal genera and soil environmental factors revealed two diametrically opposed ecological adaptation strategies (Figure 5C): genera such as Uc_Nectriaceae and Aspergillus gained competitive advantages by preferring environments with a high pH, high DOC, and high NO3−, while genera including Mortierella and Trichoderma were more likely to colonize under acidic conditions with high NH4+. The soil properties collectively explained 48.8% of the variation in the fungal genus composition (Figure 5D). Among these, pH emerged as the primary driver (80.46% explanatory power), followed by NO3− (68.36%) and NH4+ (28.81%). This hierarchical influence highlights how edaphic factors partition fungal niches through both direct (pH) and indirect (nitrogen speciation) mechanisms (Figure 5D).

4. Discussion

4.1. Liming Significantly Influenced the Activity of Soil Microorganisms

In this study, the bacterial biomass in CK soils decreased, and then stabilized, while the fungal biomass increased progressively across pH 4.5 to pH 7.5. This trend may indicate a higher pH sensitivity in bacteria than fungi, with fungi showing a broader pH tolerance [29]. In tea-planted soils, the bacterial biomass reached the peak in pH 6.5, and then declined in D30 and D60 treatments, differing significantly from CK. However, fungal abundance increased significantly with extended planting duration, suggesting that rhizosphere effects play a critical role in regulating the soil microbial activity [30]. Many plant roots release labile carbon, stimulating the R-selected bacteria that use exudates and available nitrogen for rapid growth. As resources deplete, the K-selected fungi enhance the biomass by decomposing the organic matter and extending the hyphae to access low-availability nutrients [31]. In our study, the microbial biomass may reflect from the side the adaptive strategies of soil microorganisms to resource availability. This shift not only influences the nutrient availability for tea plants but also contributes to the overall stability of the soil ecosystem under long-term tea cultivation.
AOA and AOB are the dominant microbial groups mediating soil ammonia oxidation. As the rate-limiting step in autotrophic nitrification, they act as critical bioindicators for NH4+ and NO3− concentrations. In acidic soils (pH 4.5 and pH 5.5), the abundance of AOA was significantly greater than that of AOB. As the soil pH increased to pH 6.5 and pH 7.5, the AOB abundance surpassed that of AOA. This observation is consistent with the majority of findings from studies on ammonia-oxidizing microorganisms in subtropical acidic forest soils [32,33]. AOA possess a high substrate affinity for NH4+. Under acid pH conditions, the bioavailability of NH4+ is reduced [34], a scenario that favors AOA proliferation. The activity of AOB is typically positively correlated with increasing soil pH and elevated available nitrogen derived from nitrogen inputs, particularly in neutral to slightly alkaline (pH 6.5 and pH 7.5) agricultural ecosystems. In acidic conditions, the high NH4+ concentrations may exert a suppressive effect on AOB.

4.2. Liming Significantly Influenced the Dominant Taxa of Bacteria and Fungi

In the present study, the bacterial community displayed a higher sensitivity to pH variations compared to tea cultivation. With increasing pH, both bacterial diversity and richness exhibited a decreasing trend in CK treatments, suggesting that the pH imposes a selective pressure on the bacterial community, whereby taxa incapable of adapting to pH fluctuations are promptly replaced by those adapted to the altered conditions. Across the four soil pH gradients, the dominant bacterial phyla at the phylum level primarily included Proteobacteria, Actinobacteria, Acidobacteria, Chloroflexi, and Bacteroidetes, which is consistent with the previous research findings on subtropical acidic forest soils [35]. Specifically, the bacterial community compositions at pH 4.5 and 5.5 were comparable and significantly distinct from those at pH 6.5 and 7.5.
The relative abundances of Proteobacteria, Actinobacteria, and Bacteroidetes increased significantly with rising pH, whereas the relative abundances of Acidobacteria and Chloroflexi showed an opposite trend. Previous studies have identified Proteobacteria, Actinobacteria, and Bacteroidetes as copiotrophic groups, playing pivotal roles in decomposing and transforming rhizodeposits [36]. In contrast, Acidobacteria and Chloroflexi are oligotrophic taxa, dominating in oligotrophic environments with a low nitrogen use efficiency and primarily involved in organic matter decomposition [35]. In neutral and slightly alkaline soils (pH 6.5 and 7.5), it is mainly due to the fact that Nitrospirae and Gemmatimonadetes enhance organic matter degradation and nitrification processes, thereby providing increased carbon, nitrogen, and other nutrients to support tea plant growth.
Across the rising pH gradient, the relative abundance of Ascomycota increased significantly, whereas Basidiomycota decreased. This phylum level shift reflected their divergent niches and adaptations: Ascomycota, with a broad pH tolerance and efficient carbon utilization, gained an advantage in higher-pH soils. In contrast, Basidiomycota that specialized in decomposing recalcitrant organic matter (e.g., lignin) and are prevalent in acidic environments showed suppressed growth under elevated pH. Notably, this pattern aligns with Weber et al. [37], who reported similar pH-driven shifts in temperate forest soil fungal communities. The consistency across acidic soil ecosystems (temperate forests and tea plantations) underscores the pH as a general driver of fungal dynamics. Our observations are congruence-validated by this study, with the pH identified as a key driver of the fungal assemblage composition. Furthermore, the phylum-specific pH responses imply conserved physiological and ecological mechanisms, supporting the established theories on edaphic-factor-mediated microbial community assembly.

4.3. Potential Links Between Microbes and Different Edaphic Properties

The soil pH and tea cultivation significantly affect the fungal community structure, highlighting the interactions between the edaphic properties, plants, and microbial assemblages. This indicates that the plant-derived carbon allocation, including root exudates, leaf litter, and mucilage from tea plants, may regulate the fungal taxa activity and distribution. These carbon sources, varying in quantity, quality, and spatial distribution, selectively favor specific fungal groups to shape the community composition. For instance, tea root exudates (phenolics, carbohydrates, and amino acids) created a microhabitat that promote some fungal species while inhibiting others. In our study, liming alters the soil pH: it changes the availability and solubility of nutrients, and further alters the microbial community. The influence of tea cultivation on the fungal community is achieved by altering the soil structure to create more suitable ecological niches for the colonization and growth of fungi.
Moreover, the alteration in soil pH due to tea cultivation practices such as fertilization and liming can further modify the fungal community by changing the availability of nutrients and the solubility of toxic elements. This dynamic interplay between the soil chemistry and biological processes underscores the complexity of ecosystem responses to agricultural management. Additionally, the influence of tea plants on fungal communities extends beyond direct carbon inputs, as they can also alter the soil structure and moisture regimes, creating unique niches for fungal colonization and growth. Understanding these multifaceted interactions is crucial for optimizing tea production systems that are both productive and ecologically sustainable.
The redundancy analysis (RDA) showed that environmental variables explained 77.6% of the variance in the bacterial community structure, with soil pH, NH4+, and NO3− as significant drivers (p < 0.05). This shows that both the direct pH effects on the bacterial taxa and the indirect impacts from the nitrogen availability shifts due to tea plants’ preferential NH4+ uptake. Most bacterial families are positively correlated with pH and NO3−, but negatively with NH4+. Notably, Thermogemmatisporaceae, Koribacteraceae, and Burkholderiaceae, known as beneficial taxa that metabolize rhizodeposited carbon and mediate nitrogen cycling, are positively correlated with NH4+. It indicates that a rhizospheric environment promotes the growth of beneficial microorganisms, mitigating the NH4+ competition between microbes and plants and aiding in the production of compounds suppressing soil-borne pathogens [38].
Considering the substantial disparities in the soil developmental geography, climatic conditions, and nutrient status between the study site and temperate forests, an RDA analysis was performed to identify the primary factors driving the variations in the fungal community composition. Soil pH, NH4+, and NO3− concentrations were identified as significant determinants, with the relative abundances of Russula and Talaromyces showing significant negative correlations with soil pH and positive correlations with NH4+. These results were consistent with those reported by Zhang et al. [39], suggesting that the low available nitrogen content in acidic soils stimulates fungal activity and hyphal elongation to meet plant nitrogen requirements. Collectively, the soil environmental factors explained 48.75% of the variance in the fungal community structure at the genus level. The observed moderate explanatory power likely stems from plant–fungal symbioses, with ectomycorrhizal associations being a key driver. Such symbioses enable bidirectional nutrient transfer, exchanging soil nitrogen for plant-derived carbon, which is essential for plant nitrogen acquisition in nitrogen-depleted environments [40]. This study revealed that both the soil pH and tea planting were identified as key factors shaping the structure and activity of bacterial and fungal communities. These findings elucidate the dominant drivers of microbial changes, providing a theoretical basis for acidic soil remediation following long-term tea cultivation.

5. Conclusions

This study evaluated how liming and the pH gradient structure bacterial and fungal/archaeal communities in the rhizosphere of Camellia sinensis. Key findings revealed that soil pH, NH4+, and NO3− content were the primary drivers of bacterial and fungal community variations. Both the microbial diversity and richness declined progressively as the soil pH increased. As the tea plant cultivation time increases, the relationship between pH and diversity still needs further study. Notably, bacterial communities showed a greater sensitivity to pH changes than fungal communities. The tea rhizosphere exhibited the selective enrichment of bacterial families, including Rhodospirillaceae, Koribacteraceae, and Burkholderiaceae, usually as plant-growth-promoting bacteria. Concurrently, we observed a significant increase in the relative abundance of fungal genera such as Mortierella, Penicillium, and Aspergillus. There still are many main limitations in this study, such as the absence of functional measurements (enzymatic activity, nitrification rates, and exudation), the lack of evaluation of plant growth, and the short experimental period, which restrict a more in-depth interpretation of the results. In practical applications, the results can be helpful in better adjusting the liming doses in acidic soils, and also guiding fertilization management, and pH-sensitive microbial groups have potential as indicators with which to monitor soil health.

Author Contributions

Conceptualization, writing—original draft, writing—review, editing, study design, formal analysis, funding acquisition, methodology, investigation, validation, and visualization, C.Z.; study design, methodology, visualization, and data curation, X.H.; writing—review, editing, methodology, formal analysis, and validation, H.J.; resources, methodology, investigation, visualization, and data curation, X.W.; writing—review, editing, investigation, resources, and visualization, J.G.; and writing—review, editing, methodology, supervision, and funding acquisition, C.H. All authors have read and agreed to the published version of the manuscript.

Funding

This study was financially supported by the Joint Fund of Henan Province Science and Technology R&D Program (225200810047, 225200810119), and the Basic Research Projects of Henan Academy of Sciences (20250601009, 20250601003).

Data Availability Statement

The original contributions presented in this study are included in the article. Further inquiries can be directed to the corresponding author.

Acknowledgments

During the preparation of this work, the authors used DeepSeek (Version 3.2) tools only for language refinement, specifically for grammar checking, spelling correction, and improving overall fluency. This tool was not involved in generating scientific content, interpreting data, or formulating conclusions. All aspects of this research, include study design, data collection, analysis, interpretation of results, and core intellectual arguments, were conducted solely by the authors.

Conflicts of Interest

The authors declare no conflicts of interest.

References

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Figure 1. Characteristics of soil microbial biomass in different pH soils after 30 days and 60 days of cultivation. CK, the control soil without tea planted; D30 and D60, the soils after tea was planted for 30 and 60 days, respectively. Different lowercase letters indicate significant differences (p < 0.05, LSD test). (A) Gene copies number of bacterial biomass. (B) Gene copies number of fungal biomass. (C) Ratio of bacteria to fungi. (D) Gene copies number of AOA biomass. (E) Gene copies number of AOB biomass. (F) Ratio of AOA to AOB.
Figure 1. Characteristics of soil microbial biomass in different pH soils after 30 days and 60 days of cultivation. CK, the control soil without tea planted; D30 and D60, the soils after tea was planted for 30 and 60 days, respectively. Different lowercase letters indicate significant differences (p < 0.05, LSD test). (A) Gene copies number of bacterial biomass. (B) Gene copies number of fungal biomass. (C) Ratio of bacteria to fungi. (D) Gene copies number of AOA biomass. (E) Gene copies number of AOB biomass. (F) Ratio of AOA to AOB.
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Figure 2. The difference in microbial alpha diversity ((A) bacterial community and (C) fungal community) and beta diversity ((B) bacterial community and (D) fungal community) in different pH soils. CK, the control soil without tea planted; D30 and D60, the soils after tea was planted for 30 and 60 days, respectively.
Figure 2. The difference in microbial alpha diversity ((A) bacterial community and (C) fungal community) and beta diversity ((B) bacterial community and (D) fungal community) in different pH soils. CK, the control soil without tea planted; D30 and D60, the soils after tea was planted for 30 and 60 days, respectively.
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Figure 3. Relative abundance (%) of soil microbial community composition at phylum level in all treatments: (A) bacterial community, and (B) fungal community. CK, the control soil without tea planted; D30 and D60, the soils after tea was planted for 30 and 60 days, respectively.
Figure 3. Relative abundance (%) of soil microbial community composition at phylum level in all treatments: (A) bacterial community, and (B) fungal community. CK, the control soil without tea planted; D30 and D60, the soils after tea was planted for 30 and 60 days, respectively.
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Figure 4. Differences in the composition of bacterial (A,B) and fungal (C,D) taxa in different pH soils by LDA Effect Size analysis. LDA score threshold of bacteria was 4.0. (A,C) represent the significantly different taxa of bacterial and fungal communities in different pH soils. (B,D) shows that the relative abundance of dominant bacterial family community and fungal genera community significantly varied in different planted treatments of pH regulation. CK, the control soil without tea planted; D30 and D60, the soils after tea was planted for 30 and 60 days, respectively.
Figure 4. Differences in the composition of bacterial (A,B) and fungal (C,D) taxa in different pH soils by LDA Effect Size analysis. LDA score threshold of bacteria was 4.0. (A,C) represent the significantly different taxa of bacterial and fungal communities in different pH soils. (B,D) shows that the relative abundance of dominant bacterial family community and fungal genera community significantly varied in different planted treatments of pH regulation. CK, the control soil without tea planted; D30 and D60, the soils after tea was planted for 30 and 60 days, respectively.
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Figure 5. Redundancy analysis (RDA) between soil bacterial (A) and fungal (C) dominant taxa and environmental factors, and the explanatory rate of environmental variables for soil bacterial (B) and fungal (D) community structure variations. * means the significant important (p < 0.05). The black solid arrows represent the microbial community. The red hollow arrows represent environmental factors.
Figure 5. Redundancy analysis (RDA) between soil bacterial (A) and fungal (C) dominant taxa and environmental factors, and the explanatory rate of environmental variables for soil bacterial (B) and fungal (D) community structure variations. * means the significant important (p < 0.05). The black solid arrows represent the microbial community. The red hollow arrows represent environmental factors.
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Table 1. Primers and PCR conditions used for real-time PCR.
Table 1. Primers and PCR conditions used for real-time PCR.
Target GenePrimer SetSequence (5′-3′)Thermal Profile
Bacterial 16SEub338 (F)ACTCCTACGGGAGGCAGCAG2 min at 95 °C, followed by 40 cycles of 10 s at 95 °C, 20 s at 53 °C, and 30 s at 72 °C
Eub518 (R)ATTACCGCGGCTGCTGG
Fungal ITSITS1 (F)TCCGTAGGTGAACCTGCGG2 min at 95 °C, followed by 40 cycles of 10 s at 95 °C, 20 s at 53 °C, and 30 s at 72 °C
ITS2 (R)CGCTGCGTTCTTCATCG
Table 2. Soil physicochemical properties in different pH soils.
Table 2. Soil physicochemical properties in different pH soils.
Soil TypepHNH4+ (mg kg−1)NO3− (mg kg−1)DOC (mg kg−1)SWC
CK4.1531.01 ± 0.66 cC48.47 ± 1.14 bA66.86 ± 1.01 cC25.83 ± 0.19 aA
pH 4.5D304.1816.66 ± 0.64 bC43.53 ± 2.48 aA37.42 ± 3.89 bA25.06 ± 0.47 aA
D604.0910.38 ± 1.31 aA50.05 ± 5.31 bB17.98 ± 3.38 aA28.7 ± 1.15 bA
CK5.3630.02 ± 0.71 cC44.25 ± 4.32 aA73.94 ± 2.96 cD26.38 ± 0.25 bA
pH 5.5D305.3518.53 ± 1.63 bC52.37 ± 4.77 bB40.87 ± 11.56 bA24.56 ± 0.12 aA
D605.1912.35 ± 0.66 aB42.2 ± 1.14 aA21.21 ± 1.01 aA28.32 ± 0.19 cA
CK6.8314.92 ± 0.12 cB106.04 ± 4.8 bB52.33 ± 2.06 bA26.01 ± 0.26 bA
pH 6.5D306.8912.14 ± 0.17 bB71.24 ± 2.79 aB49.57 ± 1.86 bA24.79 ± 0.34 aA
D606.7311.27 ± 0.72 aAB65.93 ± 6.45 aC39.87 ± 7.01 aB29.69 ± 1.26 cA
CK7.7413.44 ± 0.83 bA109.88 ± 2.09 bB59.15 ± 2.49 aB25.25 ± 0.17 aA
pH 7.5D307.799.79 ± 0.45 aA76.38 ± 13.11 aB62.74 ± 5.03 aB25.02 ± 0.05 aA
D607.749.57 ± 0.57 aA72.49 ± 11.98 aC52.63 ± 5.83 aC29.78 ± 0.33 bA
CK, the control soil without tea planted; D30 and D60, the soils in which tea was planted for 30 and 60 days, respectively. NH4+, the content of soil ammonia; NO3−, the content of soil nitrate; DOC, the dissolved carbon; SWC, the content of soil water. Values were the mean ± standard deviations, n = 3. Different lowercase letters indicate significant differences in various planted treatments of the same pH soils, and uppercase letters indicate significant differences in various pH soils of the same planted treatments (p < 0.05, LSD test).
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Zhao, C.; He, X.; Jiang, H.; Wang, X.; Gao, J.; Hu, C. Liming Alters Microbial Communities Affecting Nitrification in the Rhizosphere of Camellia sinensis. Agronomy 2026, 16, 77. https://doi.org/10.3390/agronomy16010077

AMA Style

Zhao C, He X, Jiang H, Wang X, Gao J, Hu C. Liming Alters Microbial Communities Affecting Nitrification in the Rhizosphere of Camellia sinensis. Agronomy. 2026; 16(1):77. https://doi.org/10.3390/agronomy16010077

Chicago/Turabian Style

Zhao, Chang, Xiaoxiang He, Han Jiang, Xiaoyan Wang, Jinjuan Gao, and Chanjuan Hu. 2026. "Liming Alters Microbial Communities Affecting Nitrification in the Rhizosphere of Camellia sinensis" Agronomy 16, no. 1: 77. https://doi.org/10.3390/agronomy16010077

APA Style

Zhao, C., He, X., Jiang, H., Wang, X., Gao, J., & Hu, C. (2026). Liming Alters Microbial Communities Affecting Nitrification in the Rhizosphere of Camellia sinensis. Agronomy, 16(1), 77. https://doi.org/10.3390/agronomy16010077

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