Influence of Various Biochars on the Rhizosphere Microenvironment and Allelopathic Effects of Polygonatum cyrtonema Hua: Microbial Community Modulation and Enhancement of Plant Quality

Abstract

Polygonatum cyrtonema Hua (PCH) is traditionally recognized as both an edible and medicinal food source. Its rhizomes contain numerous bioactive compounds, notably polysaccharides and flavonoids, which serve as key constituents in functional food development. However, the cultivation of PCH is often hindered by allelopathic effects, which diminish its quality and restrict its industrial application. To mitigate these allelopathic influences, three types of biochars derived from maize straw (MB), rice husk (RB), and tea stem (TB) were applied at concentrations of 0%, 2%, and 4%. Initially, the physicochemical properties of these biochars were characterized, followed by an evaluation of their impact on (1) the synthesis of quality-related components, secondary metabolites, and allelochemicals within PCH rhizomes and (2) the fundamental physicochemical properties and bacterial community structure of the PCH rhizosphere soil. The findings indicated that the application of 4% RB significantly enhanced the content of total polysaccharides by 48.5%, total flavonoids by 30.2%, total saponins by 28.6%, and total polyphenols by 18.3%, while concurrently reducing protein (PRO) and free amino acid (FAA) concentrations in the rhizomes. Non-targeted metabolomic analyses revealed that biochar amendments (1) upregulated metabolites involved in the citrate cycle and galactose metabolism pathways, thereby facilitating energy supply and precursors for polysaccharide biosynthesis; (2) downregulated metabolites involved in the arginine biosynthesis pathway, which is unfavorable for protein and amino acid synthesis; (3) decreased the abundance of six identified allelochemicals, including 5-hydroxy-L-tryptophan and andrographolide, with the most pronounced effect observed in the 4% TB treatment (T2); (4) improved soil physicochemical parameters such as pH, soil organic matter (SOM), total nitrogen (TN), and available potassium (AK); and (5) altered the rhizosphere bacterial community by enriching beneficial phyla, notably Myxococcota and Gemmatimonadota. These modifications in soil properties and bacterial community composition were closely associated with enhanced rhizome quality and a reduction in allelochemical accumulation. Collectively, the results of this study elucidate the potential mechanisms linking biochar application to allelopathy mitigation, optimization of soil microbial communities, and improvement of PCH rhizome quality. This research provides a theoretical basis for the production of high-quality PCH while concurrently minimizing allelochemical accumulation in its rhizomes.

1. Introduction

Polygonatum cyrtonema Hua (PCH) represents a significant source of traditional Chinese medicinal materials [1]. This medicinal and edible species is rich in bioactive constituents, including polysaccharides and flavonoids, which exhibit immunomodulatory and antioxidant properties [2]. The increasing market demand for health products and functional foods derived from Polygonatum has necessitated the expansion of its cultivation, primarily due to the limited availability of wild germplasm resources. Nevertheless, continuous monoculture practices may detrimentally affect the quality of PCH rhizomes, a phenomenon largely attributed to allelopathic effects arising from the accumulation and subsequent release of metabolites, such as phenolic acids [3], carboxylic acids, saponins, and flavonoids, into the surrounding environment [4,5]. These compounds can exert allelopathic influences on both conspecific and heterospecific organisms [6]. Despite this, investigations into the specific allelochemicals present in PCH rhizomes remain scarce.

Allelochemicals released into the rhizosphere soil can adversely impact soil physicochemical characteristics and modify the composition of soil microbial communities. For example, certain carboxylic acids, including coumaric acid, have been reported to acidify soil, diminish soil organic matter (SOM) content, disrupt soil aggregate stability, and reduce soil aeration [7]. Additionally, these compounds may facilitate nutrient leaching, affecting essential elements such as nitrogen (N), phosphorus (P), and potassium (K) [8]. Polyphenolic compounds detected in the rhizosphere soils of tea plants have been shown to inhibit beneficial plant growth, promoting bacteria, including members of the genera Bacillus and Sphingomonas [9]. In the rhizosphere of Panax notoginseng, saponins have been implicated in promoting the proliferation of pathogenic microorganisms (such as Fusarium) while suppressing beneficial taxa (such as Saitozyma and Streptomyces), thereby increasing vulnerability to root rot disease [10]. Furthermore, flavonoids like quercetin dihydrate, quercitrin, and rutin have been found to enhance the growth of pathogenic fungi in the rhizosphere soil of the potato plant, including Alternaria solani, Botrytis cinerea, Fusarium solani, and Verticillium dahliae [11]. Collectively, these alterations in the physicochemical and biological properties of rhizosphere soil exert feedback effects that ultimately influence plant quality [12].

Biochars, produced through the pyrolysis of organic substrates such as wood, straw, and rice husk under high temperature and oxygen-limited conditions [13], have demonstrated multiple beneficial effects. These include (1) mitigation of allelopathic effects in various medicinal plants, such as Panax ginseng [14], Panax quinquefolius [15], Panax notoginseng [16], and Chrysanthemum [17]; (2) enhancement of soil nutrient status, optimization of soil structure, and modulation of soil pH [12]; (3) promotion of growth in medicinal plants like Sophora tonkinensis Gagnep [18] and Panax ginseng [14]; and (4) augmentation of the maximum sorption capacity of plant roots by increasing the number of functional groups on root cell walls, thereby altering root metabolite profiles [19]. However, biochars derived from different feedstocks often exhibit distinct physicochemical properties. For instance, biochars produced from rice straw, tea waste, and other agricultural residues vary in parameters such as pH, ash content, and nutrient composition [20]. Consequently, the application of different biochars may differentially influence the quality of PCH rhizomes and modulate allelopathic effects. Nonetheless, empirical evidence supporting this hypothesis remains limited.

To address these gaps, the present study employed a pot experiment utilizing the PCH cultivar Minchangjing 1 and three biochars derived from maize straw (MB), rice husk (RB), and tea stems (TB). The investigation focused on assessing (1) the concentrations of key compounds in the PCH rhizomes, including total polysaccharides (TPS), total flavonoids (TF), total saponins (TS), total polyphenols (TPP), total proteins (PRO), and free amino acids (FAAs); (2) the abundance of significant metabolites, encompassing allelochemicals, in PCH rhizomes (including allelochemicals) through non-targeted metabolomic analysis; and (3) the physicochemical properties and bacterial community composition of PCH rhizosphere soil. The overarching objective was to evaluate the potential of different biochars to enhance PCH rhizome quality while concurrently mitigating allelopathic effects, thereby promoting the sustainable cultivation of PCH.

2. Materials and Methods

2.1. Experimental Design

This study utilized the PCH cultivar Minchangjing 1, officially registered in 2024 by our research team. Sandy loam soil (depth < 20 cm) was collected from a cultivation site in Guangze County, Fujian Province, China (117.53° E, 27.18° N) [21]. Three commercial biochars (designated MB, RB, and TB) were purchased from Henan Lize Environmental Protection Technology Co., Ltd. (Zhengzhou, Henan Province, China). These biochars were produced via pyrolysis at temperatures ranging from 500 to 550 °C under oxygen-limited conditions.

The soil, biochars, and chicken manure were air-dried and sieved through a 2 mm mesh. Subsequently, the soil was thoroughly homogenized with basal fertilizers, biochars, and chicken manure according to the following specifications: (1) chicken manure was applied at 2% (w/w) per pot; (2) basal fertilizers were supplied at rates of 350 mg kg⁻¹ N [(NH₄)₂SO₄], 320 mg kg⁻¹ P₂O₅ [Ca(H₂PO₄)₂·CaSO₄], and 100 mg kg⁻¹ K₂O (KH₂PO₄); and (3) biochars were incorporated at concentrations of 0% (control, CK), 2% MB (M₁ or MB₁), 2% TB (T₁ or TB₁), 2% RB (R₁ or RB₁), 4% MB (M₂ or MB₂), 4% TB (T₂ or TB₂), and 4% RB (R₂ or RB₂). This resulted in seven treatment groups, each with three biological replicates (one replicate per pot), and six plants per pot. Statistical analyses were performed at the pot level. Plant-level measurements were averaged per pot prior to statistical analysis. The prepared mixtures (2 kg per pot) were placed into plastic pots (14 cm length × 10 cm height) and allowed to stabilize at ambient temperature for three weeks.

2.2. Determination of Basic Physicochemical Properties of Biochars

The physicochemical characteristics of the biochars were assessed following the protocols described by Ma et al. [22]. The pH and electrical conductivity (EC) of biochar suspensions (solid-to-liquid ratio of 1:20) were measured using a pH meter (PHSJ-3F, Leici Technology Co., Ltd., Shanghai, China) and an EC meter (DDSJ-307F, Leici Technology Co., Ltd., Shanghai, China), respectively. Ash content was quantified via high-temperature ashing. Cation exchange capacity (CEC) was determined using the barium chloride–sulfuric acid method. Total N, P, and K contents were measured after digestion with H₂SO₄-H₂O₂, employing the Kjeldahl method for N, the molybdenum–antimony anti-ascorbic acid colorimetric method for P, and flame spectrophotometry for K, respectively. Detailed methodologies are provided in Supplementary Materials File S2. The results for pH, EC, ash content, CEC, total N (TN), total P (TP), and total K (TK) are presented in Table S1.

2.3. Structural Characterization of Biochars

The biochars were further characterized for several physicochemical parameters, including Brunauer–Emmett–Teller (BET) surface area, micropore volume and area, average pore size, and N₂ adsorption–desorption isotherms (Figure 1). Surface morphology was examined via scanning electron microscopy (SEM, JSM-7610F Plus, JEOL Ltd., Kyoto, Japan) (Figure 2a), surface elemental composition was analyzed using energy-dispersive X-ray spectroscopy (EDS, X-MaxN 80T, Oxford Instruments plc, Oxford, UK) (Figure S1), and surface functional groups were identified through Fourier transform infrared spectroscopy (FTIR, Nicolet iS 50 ⁺ Continuum, Thermo Fisher Scientific Inc., Waltham, MA, USA) (Figure 2b,c) and X-ray photoelectron spectroscopy (XPS, Kratos Axis Supra⁺ X, Shimadzu Corporation, Kyoto, Japan) (Figure 3). The specific analytical procedures are detailed in Supplementary Materials File S2.

2.4. Cultivation and Maintenance of PCH

Uniform and healthy PCH rhizomes of three-year-old PCH plants were collected and surface-sterilized through immersion in 0.1% potassium permanganate solution for 10 min, followed by thorough rinsing with deionized water. Six rhizomes were transplanted into each plastic pot, with a total of 21 pots arranged randomly within a greenhouse. Plants were cultivated under a red-to-blue LED light ratio of 3:1, with a photosynthetic photon flux density of 170 μmol m⁻² s⁻¹ and a photoperiod of 16 h light/8 h dark. Temperature was controlled at 26 ± 1 °C during the day and 18 ± 1 °C at night, with relative humidity maintained between 60% and 70% via an automated humidification/dehumidification system. Soil moisture was maintained at 70% of field capacity using the gravimetric method.

2.5. Collection and Analysis of PCH Rhizosphere Soil

2.5.1. Rhizosphere Soil Sampling and Physicochemical Property Analysis

After six months of biochar treatment, PCH rhizomes were harvested. Soil particles adhering to the rhizomes were gently brushed off and collected and sieved to remove large debris, defined as the rhizosphere soil. A total of 21 soil samples were collected (7 treatments × 3 replicates), with soil from six rhizomes per pot combined into a single sample. Portions of these samples were air-dried for physicochemical analyses conducted according to Lu [23]. Soil pH and EC were measured at a soil-to-water ratio of 1:2.5 using a digital pH meter (PHS-25, Leici Technology Co., Ltd., Shanghai, China) and a portable conductivity meter (DDSJ-308F, Leici Technology Co., Ltd., Shanghai, China), respectively. CEC was determined by means of ammonium acetate extraction followed by inductively coupled plasma optical emission spectrometry (Thermo iCAP 7000, Thermo Fisher Scientific, Waltham, MA, USA). Soil organic matter (SOM) content was quantified via oxidation with potassium dichromate in concentrated sulfuric acid, followed by titration to determine the oxidant consumption. Total N (TN) was measured after digestion with concentrated sulfuric acid and subsequently using an automatic Kjeldahl nitrogen analyzer (K1100, Hanon Future Technology Group Co., Ltd., Jinan, China). Available N (AN) was determined by means of sodium hydroxide hydrolysis and boric acid absorption. Total P (TP) was quantified following digestion with perchloric acid–sulfuric acid, and available P (AP) was extracted using 0.03 mol/L ammonium fluoride solution (pH 8.5), with both analyzed spectrophotometrically via the molybdenum–antimony anti-ascorbic acid colorimetric method (UV-1601, Beijing Beifen-Ruili Analytical Instrument (Group) Co., Ltd., Beijing, China). Total K (TK) was determined by means of high-temperature fusion with sodium hydroxide and available K (AK) was extracted using ammonium acetate solution (pH 7.0), with concentrations measured by means of flame photometry (AA-3800, Metash Instruments Co., Ltd., Shanghai, China).

2.5.2. Microbial Community Analysis of Rhizosphere Soil

A subset of rhizosphere soil samples was immediately frozen in liquid nitrogen for DNA extraction and subsequent bacterial community profiling at Majorbio Bio-Pharm Technology Co., Ltd. (Shanghai, China). Genomic DNA was extracted from 0.5 g of frozen soil samples using the E.Z.N.A.^® Soil DNA Kit (Omega Bio-tek, Norcross, GA, USA). The V3–V4 hypervariable regions of the 16S rRNA gene were amplified via polymerase chain reaction (PCR) using primers 338F (5′-ACTCCTACGGGAGGCAGCAG-3′) and 806R (5′-GGACTACHVGGGTWTCTAAT-3′) [21]. PCR products were purified, quantified, and used to construct sequencing libraries with the NEXTFLEX Rapid DNA-Seq Kit (Revvity, Inc., Waltham, MA, USA), involving adapter ligation, magnetic bead-based removal of adapter dimers, PCR enrichment, and final purification. Paired-end sequencing was performed on an Illumina NextSeq2000 platform (Illumina, San Diego, CA, USA). Raw sequencing data were deposited in the NCBI Sequence Read Archive (SRA) under accession number PRJNA1394316. Bioinformatic processing included quality filtering, amplicon sequence variant (ASV) inference via DADA2 (v. 1.28.0) denoising, taxonomic annotation against the Silva 16S rRNA database (v. 138), and analyses of microbial alpha and beta diversity.

2.6. Collection and Analysis of Pch Rhizomes: Biomass, Metabolomics, and Key Compounds

2.6.1. Collection of PCH Rhizomes and Biomass Determination

All six rhizomes per pot were washed with tap water to remove soil and debris; fibrous roots were excised, followed by rinsing with deionized water and blotting dry. Three rhizomes were randomly selected per pot and pooled as one experimental replicate. Fresh weight was measured using an electronic analytical balance (ME204E, Mettler Toledo, Greifensee, Switzerland). Samples were then deactivated at 105 °C for 30 min and oven-dried at 60 °C to constant weight, after which dry weight was recorded (n = 3 biological replicates). The remaining three rhizomes per pot were pooled, sectioned, rapidly frozen in liquid nitrogen, and stored at −80 °C for metabolomic analysis.

2.6.2. Metabolomic Profiling of PCH Rhizomes

Metabolomic analyses were conducted at Majorbio Bio-Pharm Technology Co., Ltd., following the protocol of Zhu et al. [24]. Approximately 100 mg of frozen rhizome tissue was extracted with 800 μL of methanol–water solution (4:1, v/v) using low-temperature ultrasonic treatment. Chromatographic separation was performed on a ACQUITY UPLC system (Waters Corporation, Milford, MA, USA) equipped with a BEH C18 column (100 mm × 2.1 mm, 1.7 μm particle size). Mobile phases consisted of 0.1% formic acid in water (phase A) and acetonitrile with 0.1% formic acid (phase B). Further methodological details are provided in Supplementary Materials File S2.

2.6.3. Quantification of Key Compounds in PCH Rhizomes

Remaining rhizome samples were oven-dried at 60 °C, ground into powder, and sieved through a 50-mesh screen for the determination of quality indices including TPS, TF, TS, TPP, PRO, FAA, water-soluble extracts (WSEs), and dry matter content (DMC). TPS content was measured according to the Chinese Pharmacopoeia [1]. TF concentrations were quantified using a rutin standard curve [25]. TS content was determined following the method of Peng et al. [26]. TPP concentration was assessed using a gallic acid standard curve [27]. PRO and FAA contents were determined in accordance with the National Food Safety Standard [28] via the Kjeldahl method. WSE content was determined as per the Chinese Pharmacopoeia [1]. Fresh rhizomes were sliced thinly, oven-dried at 60 °C to constant weight, and DMC was calculated as the percentage of dry weight relative to initial fresh weight. Detailed protocols are available in Supplementary Materials File S2.

2.7. Data Analysis

Physicochemical properties of soil and biochar, as well as key compound contents, were expressed as means ± standard error (SE, n = 3). Statistical analyses were performed using SPSS version 22.0. One-way analysis of variance (ANOVA) followed by Tukey’s multiple comparison test was employed to assess significant differences among treatments (p ≤ 0.05). Graphical representations were generated using Origin 2021 and Microsoft Office Visio 2016.

Microbial community and metabolomic data (n = 3) were analyzed and visualized using the cloud platform of Majorbio Bio-Pharm Technology Co., Ltd. The associated raw data were present in https://www.majorbio.com/. Differentially accumulated metabolites (DAMs) were identified based on three integrated criteria: (1) variable importance in projection (VIP) score > 1.0 from orthogonal partial least squares-discriminant analysis (OPLS-DA, SIMCA-P v14.1), validated though seven-fold cross-validation and permutation testing (R²Y = 0.996, Q² = 0.986); (2) statistical significance with a false discovery rate (FDR) < 0.05, where FDR was adjusted from the original p-values of Student’s t-test using the Benjamini–Hochberg method; and (3) an absolute log₂ fold change (|log₂FC|) ≥ 1.0. Additional details on microbiome data processing and metabolomic analyses are provided in Supplementary Materials File S2.

3. Results

3.1. Isothermal Adsorption–Desorption Curve Analysis and Specific Surface Area Analysis

Maize straw biochar (MB) exhibited the highest BET specific surface area of 13.87 m² g⁻¹, along with a micropore volume of 0.0015 cm³ g⁻¹, and a micropore surface area of 3.27 m² g⁻¹ (Table in Figure 1). These characteristics indicate a well-developed porous structure with abundant active sites. In contrast, rice husk biochar (RB) and tea stem biochar (TB) demonstrated comparatively lower micropore volumes and surface areas. RB had the largest average pore diameter of 21.79 nm, suggesting a more open pore architecture that facilitates material transport. TB and MB had average pore sizes of 11.89 nm and 7.40 nm, respectively.

Increased relative pressure (P/P0) enhanced the nitrogen (N₂) adsorption capacity on the pore surfaces of the three biochars (Figure 1a–c). At lower pressures, N₂ adsorption capacity increased gradually, corresponding to monolayer formation (Figure 1a–c). When the pressure exceeded 0.9 P/P0, a rapid increase in N₂ adsorption capacity was observed, indicating multilayer adsorption and subsequent N₂ condensation within the pores (Figure 1a–c). Notably, the adsorption and desorption curves for all three biochars exhibited poor overlap, resulting in a hysteresis loop (Figure 1a–c). This phenomenon suggests the presence of mesoporous structures with capillary condensation effects.

Among the biochars, MB demonstrated the highest N₂ adsorption capacity, while TB exhibited the lowest (Figure 1a–c). At P/P0 values nearing 1.0, MB’s adsorption capacity reached approximately 16 cm³ g⁻¹ (STP) (Figure 1a), compared to 7 cm³ g⁻¹ for RB (Figure 1b) and 2 cm³ g⁻¹ for TB (Figure 1c). The pronounced hysteresis loop observed in MB (Figure 1a) indicates the presence of mesopores (2–50 nm) and a complex pore architecture. In contrast, RB and TB displayed less distinct hysteresis loops (Figure 1b,c), suggesting a lower abundance of mesopores and less structural complexity, particularly in TB. The gradual increase in adsorption capacity at low P/P0 for MB implies a microporous structure with abundant adsorption sites for N₂. Conversely, the lower adsorption capacities and slower rates of increase for RB and TB suggest smaller specific surface areas and fewer adsorption sites.

Pore volume distribution analysis revealed that MB exhibited a rapid decrease in pore volume (dV/dD) within the small pore size range (<50 nm) (Figure 1d). This observation indicates a predominance of small pores such as micropores or small mesopores, with the presence of minimal large pores (>50 nm). In contrast, RB and TB displayed irregular pore volumes for pores smaller than 50 nm and a decrease in pore volume for pores larger than 50 nm (Figure 1e,f). These findings suggest that RB and TB possess a more heterogeneous pore size distribution and a higher proportion of large pores compared to MB.

3.2. X-Ray Photoelectron Spectroscopy (XPS) Analysis

Full-spectrum XPS analysis (Figure 3) revealed distinct elemental compositions among the biochars: TB contained carbon (C), oxygen (O), nitrogen (N), fluorine (F), and potassium (K); RB comprised C, O, N, sulfur (S), and silicon (Si); and MB included C, O, N, S, and iron (Fe) (Figure 3a–c).

High-resolution C 1s XPS spectra revealed that TB contained the following chemical bonds: C=C (284.2 eV, 21.7%), C=C (283.68 eV, 29.14%), C–C/C–H (285.31 eV, 15.57%), C–F (292.31 eV, 3.67%), and C=O (287.82 eV, 7.21%) (Figure 3d). In contrast, RB was characterized predominantly by C=C (283.76 eV, 66.77%), C–C/C–H (284.78 eV, 9.35%), and C–H/C–O (285.16 eV, 18.79%) bonds (Figure 3e). MB exhibited C=C (283.87 eV, 59.22%), C–C (284.59 eV, 10.85%), C–H (284.99 eV, 21.00%), C=O (288.28 eV, 1.78%), and C–F (292.58 eV, 7.15%) bonds (Figure 3f).

Analysis of the high-resolution N 1s spectra revealed that TB contained multiple nitrogen-containing bonds. These included C–N–H (399.48 eV, 21.03%), C–N–H (400.3 eV, 11.95%), C–N=O (401.05 eV, 2.75%), and various C=N–C configurations (398.76 eV, 9.03%; 397.92 eV, 9.62%; 396.34 eV, 0.92%) (Figure 3g). In contrast, RB exhibited bonds such as –C–NH–C– (399.28 eV, 7.18%), C–N–H (397.67 eV, 43.83%), C=N–C (398.31 eV, 21.52%), –NH₂ (400.12 eV, 6.44%), –NH₃⁺ (401.22 eV, 3.68%), and –N–OH (402.55 eV, 17.36%) (Figure 3h). MB presented bonds including –NO₂ (402.1 eV, 0.76%), C–N–H (399.8 eV, 43.65%), C=N–C (398.5 eV, 13.66%), C=N–C (397.54 eV, 6.79%), and Fe–N (394.42 eV, 1.38%) (Figure 3i).

The high-resolution S 2p spectra revealed that TB exhibited bonds corresponding to FeS (157.76 eV, 15.37%), S²⁻ (162.25 eV, 7.65%), –SH (163.54 eV, 4.65%), C–S–C (164.46 eV, 5.74%), SO₃²⁻ (165.57 eV, 8.75%; 167.41 eV, 21.98%; 167.69 eV, 4.51%), SO₄²⁻ (168.29 eV, 11.85%; 170.11 eV, 6.49%), and –SO₃H (169.11 eV, 13.01%) (Figure 3j). In contrast, RB contained FeS (161.56 eV, 6.33%), S⁰ (162.86 eV, 2.69%), C–S–C (163.78 eV, 6.79%), S–S (164.72 eV, 5.19%), –SO– (165.9 eV, 9.95%), SO₃²⁻ (167.54 eV, 21.06%), SO₄²⁻ (168.3 eV, 10.05%), and S₂O₈²⁻ (169.35 eV, 16.88%) bonds (Figure 3k). Additionally, MB displayed FeS₂ (160.96 eV, 3.10%), C–S–C (163.95 eV, 30.52%), –SO– (167.52 eV, 16.11%), SO₄²⁻ (168.28 eV, 8.32%; 168.99 eV, 6.29%), –SO₃H (170.26 eV, 8.49%), and S₂O₈²⁻ (171.05 eV, 8.49%) bonds (Figure 3l).

3.3. Scanning Electron Microscopy (SEM) and Fourier Transform Infrared (FTIR) Analysis

At a magnification of 100 μm (Figure 2a), MB exhibited a loose and aggregated structure characterized by numerous strips and debris. RB displayed blocky and strip-shaped aggregates, whereas TB demonstrated a continuous layered or honeycomb-like dense structure. At 10 μm magnification (Figure 2a), MB revealed irregular pores and an overall loose architecture; RB showed a uniform honeycomb-like porous structure with well-connected pores; and TB exhibited a uniform and compact pore arrangement. At the highest magnification of 1 μm (Figure 2a), MB presented rough, fragmented fibers and flaky features; RB had smooth pore walls with a regular pore structure; and TB showed a higher density with relatively fine texture.

FTIR analysis (Figure 2b) indicated that all three biochars contained C–O bonds (associated with alcohols and ethers), C=C bonds (aromatic ring structures), and O–H bonds (hydroxyl groups), confirming the presence of oxygen-containing functional groups and/or aromatic rings. Specifically, MB exhibited Si–O–Si bonds (465/800 cm⁻¹), CO₃²⁻ functional groups (870 cm⁻¹), and O=C=O bonds (2357 cm⁻¹) (Figure 2b). RB showed Si–O–Si bonds (460 cm⁻¹), C–H bonds (786 cm⁻¹), and C≡N bonds (2197 cm⁻¹). TB lacked Si–O–Si bonds but contained C–H bonds (656 cm⁻¹), O=C=O bonds (2357 cm⁻¹), and hydroxyl groups (3450 and 3762 cm⁻¹), which were absent in MB and RB. These variations in functional groups likely influence the surface chemical properties of the biochars and, consequently, their adsorption capacities.

3.4. Quality Assessment of PCH Rhizomes

All biochar treatments significantly enhanced both fresh and dry weights of PCH rhizomes, with M_1/2 and T₁ treatments demonstrating the most pronounced yield improvements (Figure 4a,b). Biochar application generally increased TPS content, with the R₂ treatment eliciting a significant 48.5% increase relative to the control (Figure 4c). TF content was significantly elevated under T_1/2, R_1/2, and M₂ treatments, particularly in the M₂, R₂, and T₁ groups (Figure 4d). TS content was significantly augmented by M_1/2 and R₂ treatments compared to the control (Figure 4e). TPP content increased significantly following M₁, R_1/2, and T₁ treatments, with the M₁ treatment showing the greatest effect (Figure 4f).

Conversely, PRO content was significantly reduced by all treatments except R₁, especially at higher application rates of MB, RB, and TB (Figure 4g). FAA content decreased significantly under R₂ and T_1/2 treatments, notably in R₂ (Figure 4h). DMC was significantly increased by R₂ and T_1/2 treatments (Figure 4j). Collectively, these findings suggest that the R₂ treatment notably enhanced fresh and dry weights, TPS, TF, TS, TPP, and DMC, while reducing FAA and PRO levels (Figure 4).

3.5. Identification of Key Metabolites in PCH Rhizomes

Non-targeted metabolomic profiling identified a total of 959 DAMs across all biochar treatments (Figures S2 and S3), with carboxylic acids and their derivatives being the most abundant metabolite class according to HMDB classification (Figure 5a). KEGG pathway enrichment analysis revealed that the citrate cycle, arginine biosynthesis, and galactose metabolism pathways were the top 3 significantly enriched pathways (p ≤ 0.05) (Figure 5b).

Key metabolites involved in the citrate cycle (including pyruvic acid, isocitric acid, L-malic acid, and citric acid) were significantly upregulated by M₂, R_1/2, and T_1/2 treatments (Figure 5c and Figure S4a–d). Within the galactose metabolism pathway, UDP-D-galactose abundance was significantly increased by all biochar treatments (Figure 5c and Figure S4e). Conversely, levels of galactan, fructose-6-phosphate, and galactonic acid were significantly downregulated by M₂, R_1/2, and T_1/2 treatments (Figure 5c and Figure S4f–h). Regarding arginine biosynthesis, L-(+)-arginine abundance was significantly reduced by all biochar treatments (Figure 5c and Figure S4i). Additionally, argininosuccinic acid levels were significantly decreased by M_1/2 treatments, and N-acetyl-L-glutamyl 5-phosphate abundance was significantly lowered by M₂ and R₂ treatments (Figure 5c and Figure S4j,k).

These results collectively suggest that the citrate cycle may be enhanced while arginine biosynthesis may be suppressed under biochar amendments (MB, RB, TB). Furthermore, in galactose metabolism, the accumulation of UDP-D-galactose was promoted under the condition of biochar application. Among the treatments, R₂ induced the most pronounced upregulation of UDP-D-galactose.

3.6. Abundance of Allelochemicals in PCH Rhizomes

Among the DAMs previously identified, eight well-characterized, allelopathic compounds were detected, including 5-hydroxy-L-tryptophan [29], andrographolide [30], pinocembrin [31], 3-coumaric acid [32], cellobiose [33], 3-feruloylquinic acid [34], sclareol [35], and salicylic acid [36].

Relative to the CK, the abundance of 5-hydroxy-L-tryptophan was significantly decreased by the T_1/2, M_1/2, and R₂ treatments, whereas the R₁ treatment did not produce a statistically significant effect on its abundance (Figure 6a). Similarly, the abundance of andrographolide was markedly decreased by the T_1/2, R₁, and M₂ treatments (Figure 6b).

The T₂ treatment significantly elicited reductions in the abundances of pinocembrin, 3-coumaric acid, and cellobiose by 82.34%, 79.23%, and 28.14%, respectively, compared to the CK (Figure 6c–e). Additionally, the M_1/2 treatments significantly diminished 3-coumaric acid levels, while cellobiose abundance was notably lowered by the R₂ treatment. The relative abundance of 3-feruloylquinic acid was significantly reduced under the R₁ and T₂ treatments (Figure 6f), and the abundance of sclareol was significantly decreased by the M₁, R_1/2, and T_1/2 treatments (Figure 6g). In contrast, salicylic acid abundance was significantly elevated following the M₁ treatment (Figure 6h).

Collectively, these findings indicate that biochar application substantially suppressed the abundance of the majority of identified allelochemicals, with the T2 treatment demonstrating the most pronounced inhibitory effect.

3.7. Effects of Biochars on the Physicochemical Properties of Rhizosphere Soil

Application of biochar significantly enhanced soil pH, SOM, TN, and TP relative to the CK, and the values of pH, SOM, TN, and TP increased with these parameters increasing concomitantly with higher biochar application rates (Figure 7a–d). Notably, soil pH values in the R₂ and T₂ treatments were significantly elevated compared to other treatments (Figure 7a). The highest concentrations of SOM, TN, and TP were observed under the T₂ treatment (Figure 7b–d). The TK content was significantly reduced only by the M₂ treatment (Figure 7e). The CEC was significantly increased by all biochar treatments relative to the CK, particularly under the R₁ and T₂ treatments (Figure 7f). The EC was elevated by the M_1/2, R₁, and T_1/2 treatments but decreased under the R₂ treatment (Figure 7g). The AN content was significantly diminished by all biochar treatments, with the most pronounced reduction observed in the M₁ treatment (Figure 7h). The AP was significantly increased by the M_1/2 and T₂ treatments (Figure 7i). The AK content was significantly enhanced by all biochar applications, especially under the T₂ and M₂ treatments (Figure 7j).

These results collectively suggest that biochar amendments markedly improved most physicochemical properties of the rhizosphere soil, with the exception of AN and TK. Among the treatments evaluated, T₂ exhibited the greatest effectiveness in augmenting SOM, TN, TP, CEC, and pH.

3.8. Bacterial Community Structure in the Rhizosphere Soil of PCH

3.8.1. Alpha and Beta Diversity

High-throughput sequencing of 21 soil samples yielded a total of 22,041 ASVs (Table S2). Analysis of alpha diversity revealed that biochar amendments significantly increased the Shannon diversity index (Figure S5a) while concurrently decreasing the Simpson index relative to the CK (p ≤ 0.05) (Figure S5b), indicating an enhancement in both bacterial richness and community evenness. Principal coordinates analysis (PCoA) based on Bray–Curtis dissimilarities demonstrated a distinct separation of bacterial communities at the genus level across different treatments (ANOSIM, R = 0.7659, p ≤ 0.05) (Figure S5c,d), suggesting that biochar application substantially altered the structure of the rhizosphere bacterial community.

3.8.2. Bacterial Community Composition

Across all treatments, 252 microbial taxa were shared, with the T₂ treatment harboring the highest number of unique taxa (1182), whereas the CK treatment contained 576 unique taxa (Figure S6a). Linear discriminant analysis effect size analysis (LEfSe) was employed to identify biomarker taxa distinguishing the treatments (LDA score > 3, p ≤ 0.05, Figure S6b,c). Notably, the phylum Actinobacteriota was enriched in the CK treatment; Firmicutes in R₁; the family Rhizobiaceae in R₂, the genus Chujaibacter in M₁, the order Sphingomonadales in M₂, the order Streptomycetales in T₁, and the phylum Proteobacteria in T₂, each representing key biomarkers differentiating the respective treatment groups.

The predominant bacterial phyla identified were Actinobacteriota (22.55–69.81%), Proteobacteria (11.97–36.34%), and Chloroflexi (4.04–27.16%) (Figure S7a). At the genus level, dominant taxa included Streptomyces (0.84–8.75%), norank_o_Gaiellales (0.28–8.50%), and Sphingomonas (1.81–9.89%) (Figure S7b).

3.8.3. Impact of Biochar on Bacterial Abundance at the Phylum and Genus Levels

To elucidate the principal shifts in soil bacterial community structure induced by biochar application, the top 10 taxa at both phylum and genus levels exhibiting significant differences in relative abundance compared to the CK (p ≤ 0.05) were selected for further analysis (Figure 8). At the phylum level, the relative abundance of Actinobacteriota (Figure 8a) was significantly reduced by all biochar treatments. Conversely, the relative abundances of several phyla, including Firmicutes (Figure 8d), Bacteroidota (Figure 8f), Gemmatimonadota (Figure 8g), Myxococcota (Figure 8h), and Patescibacteria (Figure 8i), were significantly elevated in most biochar-amended soils.

At the genus level (Figure 8k–t), biochar amendments significantly increased the relative abundance of several dominant genera. For instance, Sphingomonas (Figure 8m) and Gemmatimonas (Figure 8q) were notably enriched by the M₂ and R₂ treatments, whereas Streptomyces abundance was significantly enhanced by the R₂ and T₁ treatments (Figure 8k). In contrast, the relative abundances of Humibacter (Figure 8p) and norank_o_Gaiellales (Figure 8l) were generally diminished following biochar application.

Redundancy analysis (RDA) was conducted to explore the relationships between bacterial community composition and soil physicochemical properties. At both phylum and genus levels, the first two RDA axes explained substantial cumulative variances of 83.99% and 72.23%, respectively. Among soil parameters, pH and AN emerged as the most influential factors associated with biochar-driven modifications in bacterial community structure (Figure 8u,v).

3.9. Functional Prediction Analysis

Functional predictions conducted using BugBase indicated that the application of biochar, particularly under the T₂ treatment, may lead to an increased relative abundance of Gram_negative bacteria, as well as potential for mobile genetic elements, biofilm formation, and stress tolerance. Conversely, biochar application appeared to reduce the relative abundance of the facultative anaerobes and Gram_positive bacteria (Figure 9a).

Similarly, functional annotation via FAPROTAX suggested that biochar treatment may promote the relative abundance of chemoheterotrophic bacteria, while potentially diminishing the functions associated with nitrate_reducing bacteria (Figure 9b).

3.10. Correlation Analysis Among Key Compounds and Allelochemicals in PCH Rhizomes and Soil Parameters

3.10.1. Relationships Between Allelochemicals/Key Compounds in PCH Rhizomes and Soil Physicochemical Properties

The abundance of allelochemicals exhibited significant negative correlations with most soil physicochemical properties (Figure 9c). Specifically, Cellobiose was significantly negatively correlated with TK and AN. Andrographolide showed significant negative correlations with SOM, TN, AK, and CEC. Additionally, pinocembrin was significantly negatively correlated with EC, TP, and CEC; 3-coumaric acid with AK; 3-feruloylquinic acid with SOM and TN; and 5-hydroxy-L-tryptophan was significantly negatively correlated with pH, EC, TP, and AP. Notably, cellobiose was positively correlated with AP. Furthermore, 3-coumaric acid was positively correlated with TK (Figure 9c).

The TPS content in PCH rhizomes was significantly positively correlated with soil SOM, TN, AK, and CEC (Figure 9c). TF, TS, and TPP were significantly positively correlated with soil pH, whereas FAAs and PRO were negatively correlated. TPS showed positive correlations with SOM and TN, but negative correlations with TF and PRO in relation to SOM and with FAAs and PRO in relation to TN. TS and TPP were positively correlated with TP, negatively correlated with AN, and, along with WSEs, positively correlated with AP. PRO exhibited inverse trends, showing negative correlations with TP and AP and a positive correlation with AN. TPS was positively correlated with AK, while TF and PRO were negatively correlated with this parameter.

3.10.2. Associations Between Allelochemicals/Key Compounds in PCH Rhizomes and Soil Microbial Phyla

As illustrated in Figure 9d, the relationships between soil microbial phyla and the synthesis of allelochemicals and key quality compounds in PCH rhizomes are as follows: (1) The phylum Actinobacteriota appears to negatively influence the biosynthesis of allelochemicals such as pipecolic acid, pinocembrin, and 5-hydroxy-L-tryptophan, as well as the quality compound PRO, while promoting the formation of quality compounds such as TS and TPP. (2) Proteobacteria may enhance the synthesis of allelochemicals including andrographolide, pinocembrin, 3-coumaric acid, and 3-feruloylquinic acid, alongside quality compounds such as TF and PRO. (3) The phylum Chloroflexi may facilitate the production of allelochemicals like pipecolic acid and 5-hydroxy-L-tryptophan and may suppress the accumulation of quality compounds TS and TPP. (4) Firmicutes may contribute to increased synthesis of 3-feruloylquinic acid. (5) Acidobacteriota may negatively affect the formation of the quality compound TPP. (6) Bacteroidota may promote the biosynthesis of allelochemicals such as andrographolide, pipecolic acid, pinocembrin, and 3-coumaric acid, as well as quality compound TF. (7) Gemmatimonadota may stimulate the production of certain allelochemicals, including pipecolic acid and 5-hydroxy-L-tryptophan, and enhance the synthesis of FAAs and PRO, yet may inhibit the formation of TS and TPP. (8) Myxococcota appears to suppress the synthesis of allelochemicals such as cellobiose, as well as quality compounds TF, TS, and TPP, but may promote the formation of pipecolic acid, 5-hydroxy-L-tryptophan, FAA, and PRO. (9) Patescibacteria may encourage the biosynthesis of allelochemicals, including pipecolic acid, pinocembrin, and 5-hydroxy-L-tryptophan. (10) The phylum WPS-2 seems to be unfavorable for the formation of allelochemicals such as pipecolic acid and 5-hydroxy-L-tryptophan, as well as for FAA (Figure 9d) production.

3.10.3. Associations Between Allelochemicals/Key Compounds in PCH Rhizomes and Soil Microbial Genera

According to Figure 9e, the relationships between specific soil microbial genera and the synthesis of allelochemicals and key quality compounds in PCH rhizomes are detailed as follows: (1) The genus Streptomyces may not support the biosynthesis of allelochemicals such as pipecolic acid and pinocembrin. (2) The genus norank_o_Gaiellales may not be conducive to the synthesis of allelochemicals, including pipecolic acid, pinocembrin, 3-feruloylquinic acid, and 5-hydroxy-L-tryptophan, yet may facilitate the synthesis of TPP. (3) The Sphingomonas may negatively affect the production of allelochemicals such as cellobiose, as well as quality compound TS, but may enhance the synthesis of 5-hydroxy-L-tryptophan and PRO. (4) The genera unclassified_c_Actinobacteria, Humibacter, and unclassified_o_Frankiales generally do not facilitate the synthesis of pipecolic acid (specifically for Humibacter), pinocembrin (excluding Humibacter), 3-feruloylquinic acid (specifically for unclassified_o_Frankiales), and 5-hydroxy-L-tryptophan, nor the formation of FAAs (excluding unclassified_c_Actinobacteria) and PRO. However, these genera may stimulate the synthesis of quality compounds of TS and TPP. (5) The genus Nitrolancea may promote the formation of cellobiose and TF. (6) The genera Gemmatimonas and norank_c_Acidimicrobiia may not support the formation of cellobiose (only Gemmatimonas) or the synthesis of quality compounds TS and TPP (only Gemmatimonas), but they may facilitate the biosynthesis of allelochemicals such as pipecolic acid, pinocembrin, and 5-hydroxy-L-tryptophan, as well as the synthesis of FAAs (only Gemmatimonas) and PRO. (7) Ochrobactrum may enhance the synthesis of andrographolide, 3-feruloylquinic acid, and WSEs, while potentially inhibiting the synthesis of TPS in PCH rhizomes (Figure 9e).

4. Discussion

4.1. The Three Biochars Demonstrate Superior Physicochemical Properties, Enhancing Soil Physicochemical Properties and Promoting PCH Rhizome Development

Biochar, a commonly employed soil amendment, modulates soil physicochemical attributes, thereby fostering a conducive rhizosphere microenvironment that supports plant growth [37]. Ng et al. [38] observed that the incorporation of 3% phosphorus-modified peanut shell biochar elevated soil AP levels and significantly increased TPS content in Pseudostellaria heterophylla tubers by 44%. This enhancement was attributed to improved phosphorus bioavailability in soil, which augmented photosynthetic efficiency and consequently supplied greater carbon substrates necessary for polysaccharide biosynthesis. In this study, three distinct biochars, namely, MB, RB, and TB, were applied to assess their efficacy in improving the quality of PCH rhizomes and mitigating allelochemical accumulation within these rhizomes. Comparable stimulatory effects on PCH growth (Figure 4a–f) and soil physicochemical properties (Figure 7a–j) were observed, potentially attributable to the following factors.

Firstly, all three biochars exhibited alkaline characteristics, which can effectively alleviate soil acidification [39]. Specifically, MB, RB, and TB displayed elevated pH values of 9.72, 9.85, and 9.76, respectively (Table S1). These properties likely contributed to an increase in soil pH by 0.27 to 0.88 units relative to the CK treatment (initial soil pH: 4.18), with the R₂ treatment notably raising soil pH to approximately 5.0 (Figure 7a). Previous studies have identified the optimal soil pH range for PCH growth as 4.52 to 6.81 [40]. Therefore, the observed elevation in soil pH values may represent a critical factor stimulating PCH growth (Figure 4a,b) and thriving of more microbial communities (enhanced Shannon index in Figure S5a).

Secondly, the biochars provided abundant nutrients essential for PCH development. These biochars are characterized by high contents of ash, TN, TP and TK (Table S1), as well as key elements including C, Si, O, K, Ca, and S (Figure S1). Such nutrient richness likely supports both plant growth and microbial activity. For instance, biochar application significantly increased soil SOM (Figure 7b), TN (Figure 7c), TP (Figure 7d), CEC (Figure 7f), EC (Figure 7g), and AK (Figure 7j) in the rhizosphere soil of PCH, particularly under the T₂ treatment. Notably, TB is recognized for its superior adsorption capacity [41], enabling effective nutrient retention and reducing nutrient leaching. Given that N, P, and K are vital for plant growth, with K playing a pivotal role in photosynthesis [42], the combined effects of optimized soil pH and enhanced nitrogen bioavailability can improve photosynthetic carbon fixation [43,44]. This, in turn, supplies adequate carbon skeletons and metabolic energy necessary for the biosynthesis of secondary metabolites, especially TPS, within PCH rhizomes.

Thirdly, the three biochars possess a porous architecture (Figure 3a), abundant micropores with high specific surface areas (Figure 1), and numerous functional groups (Figure 3b and Figure S1). These features likely enhance water retention and nutrient adsorption capacities, as supported by N₂ adsorption analysis (Figure 1a–f). For example, RB and MB uniquely contain C≡N and CO₃⁻ functional groups, respectively, whereas TB lacks the Si–O–Si functional group (Figure 3b). The C≡N bond is a strongly polar covalent triple bond, exhibiting partial negative charge at the N-terminus and partial positive charge at the C-terminus [45]. The CO₃²⁻ group can complex with cations such as Ca²⁺, Fe³⁺, Zn²⁺, and Mn²⁺, thereby potentially reducing the bioavailability of these elements [46,47]. Additionally, the surface of silicon oxides (Si–O–Si) is highly polar and can bind metal cations, including Ca²⁺, Fe³⁺, and Al³⁺, through hydrogen bonding, electrostatic adsorption, and coordination bonds [48,49].

4.2. Biochar Amendments Enhance Soil Physicochemical Properties, Facilitating the Synthesis of Quality Components in PCH Rhizomes

The present study demonstrated that biochar application markedly increased the concentrations of key quality compounds in PCH rhizomes, such as TPS; notably under the R₂ treatment (Figure 4c), TF (under M₂, R_1/2, and T_1/2 treatments, Figure 4d), TS (M_1/2, R₂, and T₂ treatments, Figure 4e), and TPP (under M₁ and R_1/2 treatments, Figure 4f). These enhancements were closely linked to biochar-induced improvements in the physicochemical properties of the rhizosphere soil. Specifically, (1) soil SOM, TN, AK, and CEC exhibited significant positive correlations with TPS content in PCH rhizomes (Figure 9c), with TPS content increasing by 48.5% under the R₂ treatment (Figure 4a); (2) TF content was positively correlated with soil pH, but negatively correlated with SOM, AK, and CEC; and (3) TS and TPP showed significant positive correlations with soil pH, TP, and AP, while negatively correlating with AN. These findings align with prior studies, such as Yang et al. [50], who reported that RB enhanced saponin accumulation in Panax quinquefolius roots, and Zulfiqar et al. [51], who observed that wheat straw biochar increased total phenol and TF content in Alpinia zerumbet. Collectively, these results suggest that the application of various biochars optimizes soil physicochemical characteristics, particularly for soil pH, thereby promoting the accumulation of TPS, TF, TS, and TPP in PCH rhizomes.

4.3. Biochar-Induced Activation of the Citrate Cycle and Provision of Galactose Donors May Support Polysaccharide Biosynthesis in PCH Rhizomes

The citrate cycle constitutes a fundamental metabolic pathway for energy generation via aerobic respiration [52]. In this investigation, all three biochar treatments significantly elevated the abundance of key citrate cycle metabolites (Figure 5c), including pyruvic acid (Figure S4a), isocitric acid (Figure S4b), L-malic acid (Figure S4c), and citric acid (Figure S4d). These results indicate that biochar amendments were associated with the stimulation of energy metabolism in PCH rhizomes, thereby enhancing ATP production. Consistent with these findings, previous research has demonstrated that peanut shell biochar promotes plant growth by activating galactose metabolism pathways in tomatoes [53]. UDP-D-galactose, a critical glycosyl donor derived from galactose metabolism, plays a pivotal role in plant polysaccharide biosynthesis [54]. The abundance of UDP-D-galactose was significantly increased by all biochar treatments, particularly under the R₂ treatment (Figure S4e). Therefore, biochar application, especially the R₂ treatment, may augment energy supply by upregulating citrate cycle metabolites and facilitate polysaccharide synthesis through increased UDP-D-galactose availability in PCH rhizomes.

4.4. Biochar Application Reduces FAA and PRO Concentrations in PCH Rhizomes

Biochar amendments have been reported to influence amino acid and carbohydrate metabolic pathways [55,56]. For example, RB modulated the synthesis of carboxylic acids involved in the citrate cycle (e.g., citrate, malate, acetate, and oxalate) in rice roots [57], while 0.25% wheat straw biochar inhibited amino acid metabolism in wheat roots [58]. In the current study, several metabolites associated with the arginine biosynthesis pathway were significantly downregulated across all biochar treatments (Figure 5c and Figure S4i–k). These observations corresponded with reductions in FAAs (under R₂ and T_1/2 treatments, Figure 4h) and PRO (under M_1/2, R₂, and T_1/2 treatments, Figure 4g) in PCH rhizomes. Similar decreases in FAA content have been documented in spinach [59] and protein content in soybean [60]. It is plausible that biochar amendments enhance soil nitrogen uptake and reduce substrate availability for amino acid biosynthesis [61,62].

Moreover, alterations in soil physicochemical properties induced by biochar treatments may influence FAA and PRO contents (Figure 8c). Specifically, (1) soil pH and TN exhibited negative correlations with FAA content; (2) soil pH, SOM, TN, TP, AP, and AK were negatively correlated with PRO content; and (3) the AN was positively correlated with both FAA and PRO contents (Figure 9c). Consequently, the generally increases in soil pH (Figure 7a), SOM (Figure 7b), TN (Figure 7c), TP (Figure 7d), AP (Figure 7i), and AK (Figure 7j), coupled with decreases in AN (Figure 7h), appear to be unfavorable for the accumulation of FAAs and PRO in PCH rhizomes.

4.5. Biochar Application Mitigates the Production of Established Allelochemicals in PCH Rhizomes

Medicinal plants frequently contain allelochemicals that are easily exuded, potentially leading to autotoxicity and challenges associated with continuous cropping [61]. The monoculture of Polygonatum species results in the exudation of root-derived allelochemicals, which substantially diminish crop yield and quality [3]. Specific allelochemicals including 5-hydroxy-L-tryptophan [29], andrographolide [30], pinocembrin [31], and 3-feruloylquinic acid [32], have been demonstrated to directly inhibit plant growth. Furthermore, cellobiose fosters the proliferation of pathogenic fungi such as Fusarium [33], while 3-coumaric acid adversely influences soil pH, SOM content, and rhizobacterial diversity [7], thereby degrading the rhizosphere environment. Saha et al. [62] reported that biochar derived from lemongrass distillation residues reduced andrographolide concentrations in Andrographis paniculata leaves. Consistently, the relative abundance of the allelochemicals depicted in Figure 6a–h was generally and significantly diminished by most biochar treatments compared to the CK. These observations may be attributed to several factors:

(1) Inhibition of Key Metabolic Pathways: Biochar treatments generally elevated salicylic acid levels in PCH rhizomes (Figure 6h), which may suppress the jasmonic acid signaling pathway, thereby modulating the biosynthesis of terpenoid allelochemicals (e.g., andrographolide) [63,64]. Furthermore, salicylic acid can interfere with the synthesis and metabolism of 5-hydroxy-L-tryptophan, a metabolite involved in the auxin biosynthesis pathway [65]. This partially accounted for the reduced abundance of this allelochemical.

(2) Optimization of Soil Physicochemical Properties: The abundance of these allelochemicals exhibited significant negative correlations with various physicochemical parameters of PCH rhizosphere soil (Figure 9c), including pH (pipecolic acid and 5-hydroxy-L-tryptophan), EC/TP (pipecolic acid, pinocembrin and 5-hydroxy-L-tryptophan), and AK (andrographolide and 3-coumaric acid). These findings suggest that biochar induces modifications of soil characteristics and contributes to the reduction in the accumulation of allelochemicals in PCH rhizomes.

(3) Modulation of Functional Microorganisms: Wang et al. [66] observed that biochar significantly inhibited the microbial metabolism of cellobiose, thereby reducing plant allelopathy. Shi et al. [12] reported that biochar derived from silkworm excrement enhanced soil bacterial abundance, promoted enrichment of stress-resistant bacterial genera, and decreased the prevalence of pathogenic genera. In the present study, biochar treatments (i) provided a stable habitat and carbon sources for microbes, significantly increasing microbial diversity (Figure S5a); (ii) altered the abundance of Gemmatimonadota (Figure 8g) and Myxococcota (Figure 8h). Li et al. [14] demonstrated that MB increased Gemmatimonas abundance in corn rhizosphere soil. Myxobacteria function as micro-predators preying on diverse bacteria, archaea, and fungi [67], secreting antibacterial compounds such as myxocins that inhibit soil-borne pathogens like Ralstonia solanacearum [68], thereby regulating microbial community assembly. Both Myxococcota and Gemmatimonadota produce extracellular enzymes (e.g., cellulase, ligninase, and chitinase) that degrade complex organic substrates such as chitin and cellulose [69]; (iii) reduced the relative abundance of the phylum Actinobacteriota (Figure 8a) and genus Humibacter (Figure 8p). As heterotrophic ammonifying bacteria, members of Actinobacteriota and Humibacter degrade nitrogenous SOM (e.g., proteins and peptides), releasing ammonium N (NH₄⁺-N) [70]. The decreased abundance of these taxa partially accounts for the significant reduction in AN concentration observed in PCH rhizosphere soil (Figure 7h); and (iv) notably increased the relative abundance of the genus Ochrobactrum under R_1/2 treatments (Figure 8t). Ochrobactrum species synthesize and secrete siderophores that chelate iron, forming soluble iron-siderophore complexes [71], which facilitate photosynthetic product synthesis at appropriate iron levels [72,73].

(4) Restructuring of Soil Microbial Community Composition: Correlation analyses revealed that phyla Myxococcota and Gemmatimonadota were significantly positively correlated with allelochemicals of pipecolic acid and 5-hydroxy-L-tryptophan, as well as contents of FAAs and PRO; conversely, phylum Actinobacteriota exhibited strong negative correlations with pipecolic acid, pinocembrin, and 5-hydroxy-L-tryptophan (Figure 9d). These results suggest a potential regulatory relationship between allelochemical accumulation in PCH rhizomes and the structure of the rhizosphere microbial community under biochar application.

5. Conclusions

The application of biochars significantly enhanced the quality of PCH rhizomes by stimulating the synthesis of TPS, TF, TS, and TPP, although they did not promote the synthesis of PRO and FAAs in PCH rhizomes. Non-targeted metabolomic analyses indicated that biochar amendments increased the abundance of various metabolites involved in the citrate cycle and elevated the abundance of UDP-D-galactose, thereby facilitating polysaccharide biosynthesis. Additionally, biochars suppressed various metabolites involved in the arginine biosynthesis pathway, partially accounting for the decreased PRO and FAA contents in PCH rhizomes. All biochar treatments significantly reduced the relative abundance of multiple allelochemicals, with the T₂ treatment exhibiting the most pronounced effect. Moreover, biochars markedly improved soil physicochemical properties (e.g., pH, SOM, TN, and AK) and restructured the rhizosphere bacterial community by increasing the abundance of beneficial phyla such as Myxococcota and Gemmatimonadota. In summary, the three biochars likely enhance the content of quality components and reduce allelochemical accumulation in PCH rhizomes through the following mechanisms: (1) optimization of soil physicochemical properties; (2) alteration of the abundance of key metabolites such as salicylic acid involved in allelochemical synthesis within PCH rhizomes; (3) modulation of functional microbial abundance in the PCH rhizosphere; and (4) reconstruction of the soil microbial community structure in the PCH rhizosphere.