Effects of Co-Application of Biochar and Nitrogen Fertilizer on Soil Properties and Microbial Communities in Tea Plantation

Abstract

Soil acidification reduces the abundance and activity of beneficial microorganisms, impairs tea plant growth, and ultimately leads to a decline in tea quality. Maintaining healthy soil is critical for sustainable tea agriculture. However, the interactive effect of biochar and nitrogen fertilizer on the microbial community structure and function in acidic tea plantation soils remains unclear. This study was designed to explore whether the co-application of biochar and fertilizer could enhance soil properties and maintain microbial health in tea plantations. Three treatments were set up through a controlled pot experiment: no fertilizer or biochar application (B₀N₀), fertilizer without biochar (B₀N₁), and biochar with fertilizer (B₁N₁). High-throughput sequencing technology was used to investigate the characteristics of soil microbial communities in tea plantations. Biochar amendment increased soil pH by 0.8 units, organic matter and total nitrogen by 13.5% and 21.4%, and reduced NH₄⁺-N and NO₃⁻-N leaching by 10.8% and 12.9%, respectively. It also modulated microbial community structure, enhanced the abundance of nitrogen-cycling genes (e.g., narB, nirK, nosZ), and influenced nitrogen availability through adsorption. Nitrate was identified as the main factor shaping microbial communities under fertilization. These results highlight the potential of biochar as a sustainable amendment to improve soil health and nitrogen retention in tea cultivation systems. Further field studies are warranted to validate its efficacy in enhancing tea productivity and reducing environmental nitrogen losses under real-world conditions.

1. Introduction

Tea is a traditionally important crop in China, with cultivation area reaching 3.05 million ha in 2017, accounting for 45.9% of the world’s total area [1]. To maintain high yields and improve tea quality, long-term intensive fertilization has been widely adopted in tea plantations. However, excessive use of nitrogen fertilizer has intensified soil acidification, leading to reduced soil quality and fertility, loss of essential nutrients, and posing serious threats to crop growth and potentially exacerbating environmental pollution [2]. In economically important systems such as tea plantations, soil acidification is a major obstacle to sustainable development. Therefore, it is critical to implement soil management practices such as applying organic fertilizers and amendments like biochar to restore soil health and support sustainable tea cultivation [3].

In recent years, biochar has gained significant attention as a soil amendment and in ecological remediation due to its unique physicochemical properties. Characterized by a high carbon content (60–85%), a well-developed porous structure, and abundant surface functional groups (e.g., carboxyl, hydroxyl), biochar exhibits exceptional adsorption capacity and chemical stability, making it highly effective for enhancing soil fertility, promoting carbon sequestration, and mitigating environmental pollution [4]. Its porous structure enables the adsorption of H⁺ and Al³⁺, thereby regulating soil pH, reducing nutrient losses, and increasing cation exchange capacity (CEC) through surface charge interactions [5]. In terms of soil C-N cycling, biochar has been shown to significantly enhance total organic carbon (TOC) and total nitrogen (TN) while reducing the mobility of dissolved organic carbon (DOC) [6]. Moreover, it influences soil enzyme activities (e.g., β-glucosidase, protease) and reshapes microbial community structure, offering significant potential for the restoration of degraded soils and the advancement of sustainable agriculture practices [7]. The porous matrix also provides favorable habitats for microorganisms and selectively enriches specific bacterial groups, with pH adjustment playing a crucial role in shaping bacterial abundance [8]. Therefore, optimizing biochar production parameters and application strategies is key to maximizing soil improvement and ecological benefits.

Soil microorganisms are core participants in ecosystems, playing central roles in carbon and nitrogen cycling and in maintaining overall soil health [9]. The porous structure of biochar provides habitats for microorganisms, while its rich organic carbon and mineral nutrients (e.g., P, K, Ca) promote the enrichment of bacterial taxa (e.g., Actinobacteria and Acidobacteria) and functional genes (e.g., the nifH gene (involved in nitrogen fixation) and the nosZ gene (denitrification gene)) [10]. These microbial shifts can enhance nitrogen cycling efficiency. However, some studies have reported contrasting effects on soil fungi. For example, Wu et al. [11] observed that biochar suppresses fungal proliferation, whereas Peng et al. [12] found that it promoted the growth of mycorrhizal fungi (e.g., Rhizophagus). Such discrepancies may arise from differences in biochar properties and soil environmental conditions. In summary, biochar can improve soil fertility and crop stress resistance through microbial-mediated regulation, though its effects must be tailored to soil types and fertilization strategies to support sustainable agricultural development.

Although biochar is known to ameliorate soil acidity and influence microbial activity, the interactive effects of biochar and nitrogen fertilizer on the structure and function of microbial communities in acidic tea plantation soils remain poorly understood. To address this knowledge gap, this study investigated the effects of the co-application of biochar and fertilizer on the soil physicochemical properties, enzyme activities, and microbial community structure of acidic tea plantation soil. By exploring the interactions among biochar, soil microbiota, and microbial biomass, the research aims to provide a theoretical foundation for developing scientifically grounded and engineering-oriented solutions to address challenges in tea plantation ecosystems. We hypothesized that the co-application of biochar and fertilizer would improve soil physicochemical properties, which in turn would alter microbial community structure and diversity and maintain the activity of relevant nitrogen-converting enzymes. The findings are expected to contribute to strategies for mitigating agricultural non-point source pollution, improving nitrogen use efficiency (NUE), enhancing soil quality, and ensuring both yield and quality of tea. This research holds substantial practical value for advancing the sustainable development of tea plantation ecosystems in southern China.

2. Materials and Methods

2.1. Study Area

The experimental site of this study is located in Banmian Town, Youxi County, Sanming City, Fujian Province (118°14′ E, 26°10′ N). It is located in the transitional zone between the Wuyi and Daiyun Mountain ranges, an area characterized by a typical mid-subtropical monsoon climate influenced by both continental and maritime air masses. The region has a mean annual temperature of 18.9 °C (January average: 9.3 °C; July average: 27.9 °C) and receives a mean precipitation of 1600 mm (range: 1400–1800 mm). The soil is an acidic red Ultisol with a sandy loam texture (62% sand, 29% silt, 9% clay) and a deep, loose profile, with the following basic properties: pH 4.1, organic carbon 17.2 g kg⁻¹, total N 1.8 g kg⁻¹, total P 0.96 g kg^−1, available P 173.2 mg kg⁻¹, and available K 322.8 mg kg⁻¹. The biochar used in this study was purchased from Sanli New Energy Co., Ltd., Shangqiu, China, and produced using a specialized biochar carbonization furnace. It was derived from corn straw and was pyrolyzed at 500 °C. The resulting biochar exhibited the following characteristics: pH 9.6, organic carbon 430.5 g kg⁻¹, total N 84.3 g kg⁻¹, total P 21.3 g kg⁻¹ and total K 162.1 g kg⁻¹.

2.2. Experimental Design

Soil was collected from a local tea plantation, with visible impurities such as leaves and stones removed. The air-dried soil was sieved through a 2 mm mesh and then placed into plastic pots (30 cm in inner diameter × 30 cm in height). Three treatments were set up: a blank control without biochar or fertilizer (B₀N₀), 3 g of nitrogen fertilizer without biochar (B₀N₁), and 3 g of nitrogen fertilizer with biochar (B₁N₁). Each treatment was replicated three times in a completely randomized design in a plastic greenhouse. The experimental design was based on a conversion from conventional tea plantation fertilization rates [13]. For the biochar treatment (B₁N₁), 15 kg soil was thoroughly mixed with 1% (w/w) biochar. The experiment utilized ‘Fuyun No. 6’ tea cultivar (aged <1 year), with three similarly sized seedlings transplanted into each pot in June 2024 under conventional cultivation practices. Urea, potassium sulfate, and calcium superphosphate were applied as nitrogen (N), phosphorus (P), and potassium (K) sources, respectively. A single application of N:P:K fertilizer at a mass ratio of 2.5:1:1 was added to each pot.

2.3. Sample Collection and Analysis

After one year of cultivation (June 2025), rhizosphere soil samples were collected as follows: tea seedlings were carefully uprooted, and loosely attached soil was gently removed from the roots. Roots with tightly adhered soil were placed into a 250 mL sterile flask containing 100 mL of sterile water and shaken vigorously to separate roots from the soil. The resulting soil-water suspension was transferred to 50 mL centrifuge tubes and centrifuged at 8000 rpm for 10 min at 4 °C. The pelleted material was collected and designated as rhizosphere soil. Soil pH was measured using a digital pH meter (PH220, Horiba, Ltd., Kyoto, Japan) at a soil-to-water ratio of 1:2.5 (w/v). Soil organic matter was measured by the external heating potassium dichromate method [14]. The soil total nitrogen (TN), ammonium nitrogen (NH₄⁺-N), and nitrate nitrogen (NO₃⁻-N) concentrations were quantified using a continuous flow analyzer (Skalar San++, Skalar Analytical B.V., Breda, The Netherlands).

2.4. High-Throughput Sequencing Analysis

2.4.1. DNA Extraction, Library Construction, and Metagenomic Sequencing

Genomic DNA was isolated from 0.5 g fresh soil samples employing the MoBio PowerSoil™ DNA Isolation Kit (MoBio Laboratories, Carlsbad, CA, USA) per the manufacturer’s protocol. The purified DNA was quantified for concentration and purity using a TBS-380 system (TurnerBioSystems, The Bay Area, CA, USA) and NanoDrop 2000 spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA), respectively. DNA integrity was assessed via electrophoresis on 1% agarose gels. For library preparation, the DNA was enzymatically sheared to ~350 bp fragments using a Covaris M220 ultrasonicator (Gene Company Limited, Beijing, China) to enable paired-end sequencing. The fragmented DNA was subsequently processed into paired-end libraries using the NEXTFLEX Rapid DNA-Seq Kit (Bioo Scientific, Austin, TX, USA). During this process, adapters with complete sequencing primer hybridization sequences were ligated to the DNA fragment ends. Final paired-end sequencing was conducted on an Illumina NovaSeq™ X Plus platform (Illumina Inc., San Diego, CA, USA) at Majorbio Bio-Pharm Technology Co., Ltd. (Shanghai, China), utilizing the NovaSeq X Series 25B Reagent Kit strictly following the manufacturer’s guidelines (http://www.illumina.com (accessed on 20 June 2025)).

2.4.2. Sequence Quality Control and Genome Assembly

Raw sequencing reads were processed using the Majorbio Cloud Platform. Quality control, assembly, gene prediction, taxonomic assignment, and functional annotation were conducted with standard pipelines. Detailed procedures, including software, parameters, and reference databases, are described in the Supplementary Information (Sections S1 and S2).

2.5. Statistical Analysis

Statistical analyses were performed using SPSS 27.0 for Windows (SPSS Inc., Chicago, IL, USA). One-way analysis of variance (ANOVA) was conducted followed by post hoc comparisons using Tukey’s test at p < 0.05. All data are presented as mean ± standard error (SE). Differences between treatment means were considered statistically significant at p < 0.05.

3. Results

3.1. Soil Physicochemical Characteristics

Table 1. Effect of different treatments on soil physicochemical properties.

Treatment	pH	Organic Matter (g kg−1)	Total Nitrogen (g kg−1)	NH4+-N (mg kg−1)	NO3−-N (mg kg−1)
B0N0	4.5 ± 0.03 b	32.8 ± 1.4 b	1.1 ± 0.02 b	24.7 ± 3.2 c	11.8 ± 2.1 b
B0N1	4.8 ± 0.01 b	35.6 ± 1.5 b	1.4 ± 0.04 b	53.9 ± 7.8 b	36.2 ± 2.8 a
B1N1	5.3 ± 0.03 a	40.4 ± 1.7 a	1.7 ± 0.06 a	48.1 ± 18.6 a	31.5 ± 2.4 a

B₀N₀, no biochar or fertilizer; B₀N₁, fertilizer without biochar; B₁N₁, biochar with fertilizer. Different lowercase letters in a column indicate significant differences in different treatments at p < 0.05.

shows that the B₁N₁ significantly increased soil pH by 0.5–0.8 units compared to both B₀N₁ and B₀N₀. Relative to B₀N₀, adding N fertilizer (B₀N₁) increased the concentrations of NH₄⁺-N and NO₃⁻-N, whereas the co-application of biochar and N fertilizer (B₁N₁) decreased the concentrations of NH₄⁺-N and NO₃⁻-N by 10.8% and 12.9%, respectively.

3.2. Soil Microbial Community Diversity and Structure

3.2.1. Soil Bacterial Community Diversity

The Shannon index showed significant differences between the B₀N₁ and B₀N₀ treatments (p < 0.05), while no significant differences were observed among biochar-amended treatments (Figure 1A). Venn diagrams were used to count the number of operational taxonomic units (OTUs) that are shared among or unique to different experimental treatments. Figure 1B showed that 5305 bacterial species shared across all three treatments. Non-biochar treatments (B₀N₀ and B₀N₁) exhibited significantly higher numbers of unique bacterial species compared to the biochar-amended treatment (B₁N₁) (p < 0.05). Furthermore, the N-fertilized treatment (B₀N₁) contained significantly more unique bacterial species than the non-N control (B₀N₀) (p < 0.05). The PCoA analysis of the fixed microbial communities under three different treatments showed that PC1 and PC2 explained 87.2% and 5.6% of the variation in bacterial community composition, respectively, accounting for 92.8% of the total variation (Figure 1C). Both biochar and N fertilizer addition altered the bacterial community structure in tea plantation soils. Hierarchical clustering analysis based on genus-level composition further confirmed significant differences among the three treatments (Figure 1D). Specifically, the bacterial community in N fertilizer-treated soil (B₀N₁) was more distinct from that of the untreated control (B₀N₀) but more closely resembled that of the biochar plus N fertilizer treatment (B₁N₁).

3.2.2. Soil Fungal Community Diversity

The Shannon index showed no significant differences among treatments (p > 0.05) (Figure 2A). Venn diagram analysis revealed 730 fungal species shared across all three treatments (Figure 2B). Non-biochar treatments (B₀N₀ and B₀N₁) exhibited significantly more unique fungal species compared to the biochar-amended treatment (B₁N₁) (p < 0.05). The N-fertilized treatment (B₀N₁) contained the same unique fungal species as the non-N control (B₀N₀). The PCoA analysis demonstrated that PC1 explained 88.08% of the variation in fungal community composition among treatments, while PC2 explained 5.32%, accounting for a cumulative 93.4% of the total variation (Figure 2C). Both biochar and N fertilizer significantly restructured the soil fungal community in the tea plantation ecosystem. Hierarchical clustering analysis revealed treatment-specific assemblages, with three distinct clusters corresponding to the different fertilization regimes (Figure 2D). Notably, N-amended soils (B₀N₁) formed a separate cluster from the unfertilized treatment (B₀N₀), while the combined biochar-N treatment (B₁N₁) showed intermediate compositional features. At the genus level, Rhizophagus exhibited a higher relative abundance in B₁N₁, Penicillium was more abundant in B₀N₁, and Paraglomus demonstrated higher abundance in B₁N₁.

3.2.3. Soil Bacterial Community Composition

At the phylum level, 20 bacterial phyla were identified, accounting for 97.3–97.7% of the total bacterial community, with unclassified bacteria representing 2.3–2.7% (Figure 3A). Neither biochar nor N fertilizer treatments significantly affected the proportion of unclassified bacteria. The dominant bacterial phyla across all treatments were Actinobacteria, Proteobacteria, Acidobacteria, Chloroflexi, and Planctomycetes. Compared with the control (B₀N₀), both N fertilizer (B₀N₁) and the combined biochar plus N fertilizer treatment (B₁N₁) significantly increased the relative abundance of Actinobacteria and Chloroflexi while decreasing the relative abundance of Proteobacteria and Acidobacteria. Notably, compared with N fertilizer treatment alone (B₀N₁), the combined biochar plus N fertilizer treatment (B₁N₁) reduced the abundance of Actinobacteria and Chloroflexi, while the abundance of Proteobacteria and Acidobacteria increase again. At the genus level (Figure 3B), 20 bacterial genera were identified, representing 25.2–42.8% of the total bacterial community, with unclassified bacteria accounting for 54.8–57.2%. The five most abundant bacterial genera in tea rhizosphere soils were Trebonia (4.7–10.7%), Ktedonobacter (4.9–6.3%), Bradyrhizobium (3.0–5.9%), Candidatus_Sulfotelmatobacter (0.9–4.7%), and unclassified_p__Candidatus_Dormibacteraeota (2.2–2.8%). N fertilizer alone (B₀N₁) significantly increased the relative abundance of Trebonia and Ktedonobacter, but these decreased by 4.1% and 1.1%, respectively, under B₁N₁. Conversely, compared with the control, the N fertilizer treatment alone (B₀N₁) reduced the abundance of Bradyrhizobium and Candidatus_Sulfotelmatobacter, while the addition of biochar with N fertilizer increased their abundances. The abundance of N cycle-related genes was relatively low under untreated soils (B₀N₀), indicating weak microbial N metabolism under natural conditions. Following N fertilizer application (B₀N₁), the relative abundance of nitrate assimilation genes (nasC and nasA) increased markedly (Figure 3C). Notably, the combined application of biochar and N fertilizer (B₁N₁) significantly enhanced the abundance of multiple key functional genes involved in the N cycle, including nitrate reductase (narB), nitrite reductase (nirK), and nitrous oxide reductase (nosZ).

3.2.4. Soil Fungal Community Composition

At the phylum level, a total of 11 fungal taxa were identified across the three treatment groups (Figure 4A). Among these, Ascomycota (33.3–51.3%), Mucoromycota (25.7–51.3%), Basidiomycota (10.4–16.0%), and Chytridiomycota (3.2–4.4%) each exhibited relative abundances exceeding 1% and collectively accounted for over 98% of the total fungal community, thereby representing the dominant fungal phyla. In both B₀N₀ and B₁N₁, Ascomycota demonstrated relative abundances surpassing 50%, establishing it as the most dominant phylum. Under N fertilization (B₀N₁), Ascomycota remained predominant with a relative abundance of 47.6%. Compared with the control (B₀N₀), biochar application (B₁N₁) significantly altered the fungal community structure. The relative abundance of Ascomycota decreased by 35.3% (p < 0.05), while Mucoromycota showed a significant increase in relative abundance exceeding 50%. At the genus level (Figure 4B), dominant genera with relative abundances exceeding 1% included Rhizophagus (8.3–16.7%), Penicillium (3.7–15.1%), Aspergillus (4.8–7.0%), Paraglomus (1.3–10.1%), Rhizopus (2.0–3.4%), Entrophospora (1.6–3.6%). Biochar application significantly (p < 0.05) increased the relative abundance of Rhizophagus and Paraglomus but decreased Penicillium and Aspergillus. Compared with the non-application treatment (B₀N₀), N fertilizer application increased the relative abundance of dominant fungal genera.

3.3. Environmental Factors Influencing Microbial Community Structure

Redundancy analysis (RDA) revealed significant correlations between soil physicochemical properties and microbial community structure at the phylum level. The correlation between soil bacteria and environmental factors (Figure 5A) showed that the RDA1 and RDA2 axes explained 89.31% and 6.26% of the community variation, respectively, accounting for a cumulative 95.57% of the total variation. This indicates that the selected environmental factors significantly drove the differentiation of soil microbial community structure. Specifically, TN was identified as the primary environmental factor influencing the microbial community in the B₁N₁ treatment (biochar plus N fertilizer). In contrast, NO₃⁻ showed a strong association with bacterial community distribution in the B₀N₁ treatment (N fertilizer alone), as evidenced by its vector position being closely aligned with the B₀N₁ treatment point while distinctly separated from both the control (B₀N₀) and B₁N₁ treatment points. This demonstrates that NO₃⁻ serves as a key environmental factor regulating bacterial community structure in the B₀N₁ treatment.

The correlation between soil fungal community and environmental factors (Figure 5B) showed that the RDA1 and RDA2 axes explained 50.07% and 27.83% of the community variation, respectively, with a cumulative proportion of 77.9%. The soil fungal community of the N fertilizer treatment (B₀N₁) showed a positive correlation with NO₃⁻. The soil fungal community of the biochar treatment showed a positive correlation with NH₄⁺, TN and TOC.

4. Discussion

4.1. The Effect of Biochar Combined with Fertilizer on Soil Characteristics

In the current study, we found that the co-application of biochar and nitrogen fertilizer significantly influenced soil microbial properties in tea plantations, with broader implications for nutrient cycling and soil health. The observed increase in soil pH is consistent with recent studies [15,16], indicating that the biochar is an effective amendment for improving acidic soil. This effect can be attributed to its inherent alkalinity, surface functional groups that neutralize H⁺ and bind exchangeable Al³⁺ [17,18], and enhanced retention of base cations [19]. Notably, the rise in soil organic matter following biochar application likely stems from both its direct carbon input and its ability to stabilize native organic compounds through adsorption and catalytic polymerization [20]. Moreover, the highly aromatic structure of biochar contributes to long-term carbon sequestration, as documented by Ayito et al. [21]. A key finding was the reduction in NH₄⁺-N and NO₃⁻-N leaching under biochar amendment, consistent with previous studies [22]. A possible explanation for this might be that biochar has a pronounced capacity for physical adsorption within its porous matrix [23], as well as chemical interactions through oxygen- and nitrogen-containing functional groups that enhance ammonium and nitrate retention [24].

4.2. The Effect of Biochar Combined with Fertilizer on Soil Microbial Community Structure and Diversity

Our results demonstrate that both N fertilization alone and biochar addition significantly alter soil microbial composition and function, with important implications for N cycling efficiency and ecosystem sustainability. The increased abundance of Actinobacteria and Chloroflexi under N amendment aligns with previous studies, which suggest their adaptation to high-nutrient conditions, likely due to their copiotrophic taxa and versatile metabolic capacities for complex organic matter decomposition [25]. The rebound of Acidobacteria and Proteobacteria in biochar-amended soil (B₁N₁) may reflect improved habitat heterogeneity and moderated N availability, highlighting biochar’s role in restoring microbial community balance under high-input fertilization [26].

The contrast between N fertilization (enhancing nitrate assimilation genes nasC and nasA) and biochar addition (upregulating nitrification and denitrification genes such as narB, nirK, and nosZ) suggests a fundamental shift in microbial N processing strategies. This implies that biochar facilitates a more modular and complete nitrogen cycle, potentially reducing N losses and enhancing ecosystem retention. The increase in bacterial Shannon diversity following biochar amendment is consistent with its documented role in enhancing microhabitat diversity and nutrient availability [27,28], supporting the hypothesis that biochar serves as both a refuge and a resource for microbial communities.

Fungal community composition was dominated by saprotrophic phyla including Ascomycota, Mucoromycota, Basidiomycota, and Chytridiomycota. The significant response of these taxa to biochar-induced changes in pH, organic carbon, and N-availability underscores the tight coupling between abiotic soil properties and fungal functional composition [29,30]. Furthermore, the modulation of bacterial communities by biochar—particularly groups promoting mycorrhizal symbiosis and nutrient solubilization—may indirectly shape fungal activity and community assembly [31], suggesting that cross-kingdom interactions play a key role in mediating treatment effects.

The synergistic effect of biochar and N fertilizer on microbial carbon utilization and functional diversity aligns with existing concepts of biochar’s “microbial hotspot” function [32]. We propose that biochar regulates nutrient turnover by adsorbing mineral N and mitigating acidification, thereby supporting a more diverse and active microbiome. The enhanced microbial metabolic activity, particularly for carbohydrates, further supports the idea that combined amendment promotes stable carbon and nitrogen co-cycling [33]. Future studies should focus on explicit linkages between microbial functional shifts and ecosystem-scale processes such as N₂O emissions, organic matter stabilization, and long-term soil health.

4.3. The Effects of Co-Application of Nitrogen Fertilizer and Biochar on the Relationship Between Soil Microorganisms and Environmental Factors

The RDA results confirmed that soil N dynamics (TN, NH₄⁺-N, and NO₃⁻-N) were key drivers of microbial community composition, with biochar amendment playing a moderating role in regulating N availability. Biochar influences microbial community structure through multifaceted mechanisms, including altering soil physicochemical properties and stimulating biological processes. PCoA (Figure 1C and Figure 2C) confirmed that there were significant structural shifts in microbial communities under biochar and N fertilization treatments. RDA further highlighted the importance of nitrogen forms in driving these changes (Figure 5). The strong correlation between NO₃⁻ and the microbial profile in the B₀N₁ treatment suggests that nitrate availability is a major determinant of community assembly in fertilized soils. In contrast, the association between TN and the B₁N₁ community indicates that biochar alters the patterns of nitrogen availability, likely through its adsorption properties that slow N release and reduce leaching, thereby promoting a more balanced nutrient supply [34].

These results offer valuable insights for tea plantation management. The capacity of biochar to modulate microbial community structure and enhance the expression of key N-cycling functional genes underscores its potential as an amendment for improving nitrogen use efficiency and reducing environmental N losses. However, the treatment-specific responses observed emphasize the importance of developing tailored biochar application strategies that account for local soil conditions, biochar characteristics, and management practices to maximize beneficial outcomes. In addition, several limitations should be considered: (i) the controlled environment may overestimate field-scale effects; (ii) the economic feasibility of large-scale application requires evaluation against alternative N-loss mitigation strategies. Future research should prioritize field trials that couple microbial monitoring with direct N-flux measurements to bridge the gap between mechanistic understanding and practical optimization.

5. Conclusions

This study investigated the effects of the co-application of biochar and N fertilizer on soil physicochemical properties and microbial community structure in a tea plantation ecosystem. The results demonstrated that biochar amendment significantly increased soil pH, organic matter, and total N while reducing NH₄⁺-N and NO₃⁻-N leaching, indicating enhanced nitrogen retention and improved soil fertility. Microbial analysis further revealed that biochar application increased bacterial diversity and stimulated the abundance of key N-cycling functional genes (e.g., narB, nirK, nosZ), indicating promoted nitrogen transformation efficiency. Additionally, biochar altered fungal community composition toward beneficial taxa that may facilitate symbiotic nutrient acquisition.

While these results highlight the potential of biochar as a sustainable soil amendment for enhancing microbial ecosystem functions, direct agronomic benefits—such as tea yield and quality parameters—remain to be empirically validated. Therefore, we recommend further long-term field studies to confirm these mechanistic insights under practical agronomic conditions, explore interactions with different fertilization regimes, and evaluate functional outcomes through multi-omics approaches. Such efforts are essential to translate these promising findings into actionable strategies for sustainable tea production.