Sorption-Mediated Carbon Stabilization and Bacterial Assembly Regulated by Biochar Derived from Invasive Solanum rostratum in China

Abstract

The surface chemistry of biochar plays a pivotal role in the adsorption and stabilization of soil organic carbon (SOC); however, sorption-mediated mechanisms remain insufficiently understood for biochars derived from invasive plants. In this study, Solanum rostratum biomass, an aggressive invasive weed in northern China, was pyrolyzed at 400–600 °C in 2023 to produce biochars with varying surface functionalities and structural features. FTIR, Raman, XPS, and SEM analyses revealed that increasing pyrolysis temperature led to decreased oxygen-containing functional groups and enhanced aromatic condensation, reflecting a transition from hydrogen bonding to π–π and hydrophobic sorption mechanisms. Soil incubation experiments using sandy loam soil showed that biochar produced at 500 °C significantly increased the stable carbon pool (SCP) to 52.4%, compared to 30.6% in unamended soils. It also reduced cumulative CO₂ release from 1.74 mg g⁻¹ to 1.21 mg g⁻¹ soil, indicating improved carbon retention. Bacterial 16S rRNA gene sequencing revealed that biochar amendments significantly altered community composition and increased deterministic assembly, particularly under 500 °C biochar, suggesting a sorption-driven niche filtering effect. These findings demonstrate that S. rostratum-derived biochar, especially at intermediate pyrolysis temperatures, enhances both carbon sequestration and microbial habitat structure. This has direct implications for improving degraded soils in arid farming regions, offering a dual strategy for invasive biomass management and climate-resilient agriculture.

1. Introduction

Soils represent the largest terrestrial carbon (C) reservoir, storing more C than the atmosphere and terrestrial vegetation combined [1]. Maintaining or enhancing soil organic carbon (SOC) stocks is therefore a critical strategy for supporting soil fertility and mitigating climate change through long-term C sequestration [2,3]. However, SOC is vulnerable to microbial mineralization and decomposition, particularly under increasing anthropogenic pressures and environmental changes [2,3]. Among various strategies, biochar, a carbon-rich byproduct generated by pyrolyzing biomass under limited oxygen conditions, has emerged as a promising amendment for enhancing SOC stability and improving overall soil health [4,5]. Its physicochemical attributes, such as high surface area, porosity, and diverse functional groups, endow biochar with excellent capacity to modify soil structure, nutrient dynamics, and microbial activity [6,7].

A distinctive feature of biochar is its adsorptive capability, which stems from its hierarchical pore network and chemically reactive surface [6,7,8]. These features allow biochar to retain organic molecules, microbial exudates, and root-derived compounds in soil [8,9], thus influencing microbial access to labile substrates and regulating carbon cycling processes [10]. Such sorption-mediated interactions can act as an “adsorptive filter,” altering microbial habitat availability and community structure [10]. Specifically, biochar may promote the dominance of oligotrophic microbial taxa adapted to nutrient-poor microenvironments while reducing the abundance of copiotrophic groups, thereby enhancing microbial C use efficiency and promoting long-term C retention.

Pyrolysis temperature plays a decisive role in shaping the sorption-related characteristics of biochar. Low-temperature biochars (300–400 °C) typically retain more polar groups and labile carbon, enhancing chemical reactivity, whereas high-temperature biochars (≥600 °C) become more aromatic and structurally condensed, improving their longevity and resistance to decomposition [11,12,13,14]. Therefore, determining the optimal pyrolysis condition is essential for balancing immediate bioavailability with long-term carbon sequestration [15,16].

Furthermore, the sorptive properties of biochar link its chemical composition to microbial community assembly patterns. By adsorbing extracellular enzymes, signaling compounds, and potential inhibitors, biochar can shape the biochemical microenvironment, exerting selective pressure on microbial colonization [17,18,19]. This niche filtering effect increases the role of deterministic processes in community succession, which can be detected using quantitative indices such as the β-nearest taxon index (βNTI) and Raup-Crick dissimilarity (RC_bray) [20,21,22].

From a feedstock sustainability perspective, the valorization of invasive or weedy plants into biochar presents a dual-benefit strategy. Invasive species such as S. rostratum (buffalo bur) produce significant biomass while threatening biodiversity and agricultural productivity [23,24]. Notably, prior to this study, S. rostratum has not been widely explored as a biochar feedstock. Transforming its biomass into biochar aligns with circular economy principles and provides an eco-friendly means of environmental remediation [25].

While the potential of biochar to stabilize SOC is well recognized, limited research has focused specifically on the mechanisms by which biochars derived from invasive plant biomass, such as S. rostratum, influence microbial community structure and SOC partitioning [26,27]. In particular, the connections among surface functional evolution, microbial niche filtering, and labile versus SCP remain poorly elucidated.

This study aims to address these gaps by investigating the effects of S. rostratum-derived biochars produced at 400 °C, 500 °C, and 600 °C on SOC dynamics and microbial communities. We hypothesized that (i) pyrolysis temperature-induced changes in biochar sorptive properties influence bacterial community structure and assembly patterns, and (ii) stronger adsorption capacity promotes SOC stabilization by enhancing microbial C use efficiency and necromass retention. To test these hypotheses, we conducted soil incubation experiments in Liaoning Province, China (2023–2024), and applied a two-pool kinetic model to quantify carbon mineralization and partitioning. This work contributes to our understanding of how invasive-plant biochars act as both carbon sinks and microbial habitat filters, with implications for soil carbon management and sustainable agriculture, particularly on marginal lands.

2. Materials and Methods

2.1. Soil and Biomass Sampling and Preparation

Soil and plant biomass were collected in July 2023 from riparian shelterbelts along the Daling River in western Liaoning Province, Northeast China (41°12′–41°18′ N, 120°39′–120°45′ E), a region characterized by semi-arid continental monsoon climate with an average annual temperature of ~8.5 °C and total precipitation between 450 and 550 mm, mostly occurring during July–September. The site is part of the Three-North Shelterbelt Program, dominated by Salix matsudana and Populus spp. in the afforestation zones, with understory frequently invaded by S. rostratum Dunal, a noxious annual weed known for its adaptability to disturbed soils [23,24].

The experimental soils were derived from sandy loam, classified as Typic Haplustalf under the USDA Soil Taxonomy and WRB (World Reference Base) system. These soils are particularly vulnerable to aeolian erosion, and thus representative of marginal lands targeted for ecological restoration. The choice of sandy loam reflects a practical scenario where biochar amendments could be most impactful. A map showing the study region and sampling locations is provided (Figure 1) to enhance spatial clarity and reproducibility.

Surface soil (0–20 cm) was collected from multiple plots (~20 m²) using a stainless-steel auger and homogenized into a composite sample. Visible debris, stones, and roots were removed manually. Soil was air-dried, sieved to <2 mm, and stored at room temperature prior to incubation.

The aboveground biomass of S. rostratum, which densely colonized degraded riverbanks and abandoned fields in the same zone was harvested at full flowering stage. Plants were cut at the base, transported in sealed polyethylene bags, and washed with tap water followed by distilled water to eliminate soil and contaminants. Cleaned biomass was sun-dried for 7 days until constant weight, ground using a high-speed mill, and passed through a 0.25 mm mesh for uniform particle size. The processed feedstock was stored in airtight containers for subsequent pyrolysis. Prior to this study, S. rostratum had not been widely investigated as a biochar precursor, despite its abundance and lignocellulosic characteristics, which justify its potential as a carbon-rich biomass source [23].

2.2. Biochar Preparation

Processed biomass of S. rostratum was subjected to slow pyrolysis at three target temperatures, 400 °C, 500 °C, and 600 °C, to generate biochars with distinct physicochemical and surface characteristics. For each pyrolysis batch, approximately 35 g of oven-dried biomass was placed in lidded ceramic crucibles and heated in a muffle furnace (SX2-4-10A, Shanghai Yiheng Instruments, Shanghai, China) under oxygen-limited conditions. The temperature was increased at a constant heating rate of 10 °C min⁻¹, with a residence time of 4 h at the designated final temperature. This heating rate was chosen based on established pyrolysis protocols for lignocellulosic biomass, as it provides a compromise between maximizing carbonization efficiency and preserving the porous structure and functional surface groups of the resulting biochar [11,14].

Importantly, the biochars were not washed after pyrolysis to retain water-soluble salts and ash components, which are known to influence sorption characteristics and microbial interactions in soil [6,7]. This decision reflects practices in similar studies focused on field-relevant biochar behavior and is especially relevant in the context of marginal land restoration.

The final Solanum rostratum-derived (SRD) biochar samples were labeled according to pyrolysis temperature: SRD400, SRD500, and SRD600. Uncharred biomass was referred to as SR. All materials were stored in airtight containers at room temperature prior to characterization and soil incubation experiments.

2.3. Soil Incubation Experiment

A controlled soil incubation experiment was conducted to evaluate the effects of S. rostratum-derived biochars on soil carbon mineralization and the potential for sorption-mediated carbon stabilization. Air-dried and sieved soil (<2 mm) collected from the Daling River riparian shelterbelts was homogenized and allocated into 250 mL airtight glass incubation jars. Four treatments were established:

• SS: unamended soil (control);
• BC400: soil amended with SRD400 biochar at 10% (w/w);
• BC500: soil amended with SRD500 biochar at 10% (w/w);
• BC600: soil amended with SRD600 biochar at 10% (w/w).

Each treatment was replicated four times (n = 4), in line with standard replication levels in similar soil carbon incubation studies to ensure statistical robustness while remaining logistically feasible [11,28].

The biochar application rate of 10% (w/w) was intentionally selected to simulate a high-input scenario tailored for marginal or degraded lands where agricultural productivity is not a concern. In such contexts, high-dose amendments may be environmentally safe and practically desirable to accelerate soil restoration and maximize in situ biochar sequestration [6,15]. We acknowledge that this rate exceeds typical agronomic field application rates, and thus, interpretive caution is warranted regarding broader field-scale generalization.

All jars were adjusted to 60% of the soil’s water-holding capacity (WHC) and sealed with gas-permeable parafilm to maintain aerobic conditions. Incubation was conducted in the dark at 25 ± 1 °C for 60 days. Carbon mineralization was tracked via CO₂ emission measurements on days 1, 3, 5, 7, 10, 15, 20, 30, 45, and 60 using the alkali absorption-acid titration method. Specifically, 20 mL of 0.1 mol L⁻¹ NaOH was placed in each jar to trap evolved CO₂. At each time point, NaOH traps were replaced and titrated with standardized HCl after precipitating carbonates using BaCl₂. Blank jars (NaOH without soil) were included to correct for background CO₂ absorption, and alkali trapping efficiency was periodically verified using known CO₂ standards [29].

To differentiate labile versus SCPs, cumulative CO₂ release data were fitted to a two-pool first-order kinetic model:

$$M_t=C_l(1-e^{-k_{l}t})+C_s(1-e^{-k_{s}t})$$

(1)

where $M_t$ is the cumulative mineralized carbon at time $t$, $C_l$ and $C_s$ are the sizes of labile and SCPs, and $k_l$, $k_s$ are their decomposition rate constants, respectively. The percentage of stable carbon was then calculated using:

$$\mathrm{StableC} (\%)={\displaystyle \frac{C_s}{C_l+C_s}}\times 100$$

(2)

Nonlinear regression analysis was performed using Python (v3.10), with SciPy (v1.8.1) and Matplotlib (v3.5.2) libraries, enabling extraction of kinetic parameters and comparison of carbon stabilization potential across biochar treatments [30].

2.4. Biochar Characterization and Physicochemical Properties

To elucidate the sorption-relevant physicochemical properties of S. rostratum-derived biochars and their potential contributions to soil carbon stabilization, a comprehensive suite of analytical techniques was employed.

pH and Water-Holding Capacity (WHC): Biochar pH was measured in a 1:20 (w/v) suspension using deionized water. The mixture was shaken for 1 h, allowed to settle, and the supernatant pH was recorded with a calibrated digital pH meter (Mettler Toledo, Zurich, Switzerland), following standard protocols [1]. WHC was determined by saturating 1 g of biochar with distilled water for 24 h, draining it on pre-weighed filter paper for 20 h, and calculating the difference between the wet and oven-dried (105 °C, 12 h) masses [2,31].

Elemental Composition and Surface Morphology: Surface morphology and microstructure were examined using a field-emission scanning electron microscope (FESEM; GeminiSEM 500, ZEISS, Oberkochen, Germany). Elemental composition was determined via energy-dispersive X-ray spectroscopy (EDX) attached to the SEM and reported as both atomic and weight percentages, enabling insight into the distribution of major elements such as C, O, N, and minerals [3].

Spectroscopic Analysis: Fourier-transform infrared spectroscopy (FTIR; Nicolet iS50, Thermo Fisher, Waltham, MA, USA) was used to identify functional groups across the 4000–400 cm⁻¹ range, providing qualitative insights into oxygen-containing groups and aromatic functionalities [5]. Raman spectroscopy (inVia, Renishaw, Wotton-under-Edge, Gloucestershire, UK) with a 532 nm laser was applied to assess graphitization and structural disorder (D and G bands) [6]. X-ray diffraction (XRD; SmartLab, Rigaku, Tokyo, Japan) with Cu Kα radiation (λ = 1.5418 Å) was used to detect crystalline versus amorphous carbon phases. X-ray photoelectron spectroscopy (XPS; ESCALAB 250Xi, Thermo Fisher, Waltham, MA, USA) further characterized surface elemental composition and bonding environments, with high-resolution spectra deconvoluted using XPSPEAK 4.1 software [7].

Total Element Content and Ash: Total carbon (TC), total organic carbon (TOC), and total nitrogen (TN) contents were determined using a Vario EL III elemental analyzer (Elementar, Langenselbold, Germany). TOC was measured after removing inorganic carbonates with acid treatment. Ash content was quantified by combusting 1 g of biochar at 750 °C for 6 h in a muffle furnace, which also provided indirect information on mineral residue and its potential influence on nutrient dynamics and sorption processes [8].

These integrated analyses enabled a comparative assessment of aromaticity, functional group diversity, surface area, and porosity across SRD400, SRD500, and SRD600 biochars, thereby offering mechanistic understanding of their potential to retain and stabilize organic carbon within soil matrices.

2.5. Soil Physicochemical Analyses

At the conclusion of the 60-day incubation, composite soil subsamples from each treatment (SS, BC400, BC500, and BC600) were collected to evaluate the impacts of biochar amendments on key soil physicochemical properties and carbon stabilization dynamics.

Elemental Composition and pH: Total carbon (TC) and total nitrogen (TN) contents were determined using an elemental analyzer (Vario EL III, Elementar, Langenselbold, Germany) following standard dry combustion procedures [29]. Total organic carbon (TOC) was measured via the potassium dichromate oxidation method with external heating. Soil pH was assessed in a 1:2.5 (w/v) soil-to-deionized water suspension using a calibrated digital pH meter (Mettler Toledo, Zurich, Switzerland) after equilibration for 30 min.

Moisture Content and Stable Carbon Estimation: Gravimetric soil moisture was determined by oven-drying approximately 5 g of fresh soil at 105 °C for 24 h. The proportion of the SCP (StableC, %) was derived from the kinetic parameters $C_s$ obtained through fitting the cumulative CO₂ mineralization data to the two-pool first-order model (Equation (1) in Section 2.3). To ensure accuracy, blank jars (containing only NaOH traps without soil or biochar) were included for each batch to correct for ambient CO₂ interference. The efficiency of alkali absorption was verified periodically using known CO₂ gas standards, and corrections were applied [29].

These measurements enabled a comparative assessment of how different biochar treatments influenced total nutrient status, acid-base properties, and the partitioning of labile versus stable carbon fractions in the sandy loam soil.

Soil Baseline Description: The baseline properties of the unamended soil (SS) included a pH of 6.47, TOC of 6.52%, and TN of 0.64%. These parameters reflect the relatively low fertility and weak buffering capacity of the sandy loam soils typical of riparian shelterbelts in this region, reinforcing their suitability for evaluating biochar-based soil enhancement strategies.

2.6. Microbial Community Analysis

To elucidate microbial community responses and their potential roles in carbon stabilization following biochar amendment, soil samples from each treatment group (SS, BC400, BC500, and BC600) were collected after 60 days of incubation for microbial analysis.

DNA Extraction and 16S rRNA Sequencing: Total genomic DNA was extracted from 0.5 g of fresh soil using the FastDNA™ Spin Kit for Soil (MP Biomedicals, Irvine, CA, USA), following the manufacturer’s protocol. DNA integrity was verified on 1% agarose gel, and concentration was quantified using a NanoDrop 2000 spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA). The bacterial 16S rRNA gene (V3–V4 region) was amplified using universal primers 338F (5′-ACTCCTACGGGAGGCAGCAG-3′) and 806R (5′-GGACTACHVGGGTWTCTAAT-3′). Amplicons were purified with AMPure XP beads (Beckman Coulter, Brea, CA, USA), quantified using Qubit dsDNA HS Assay Kit (Thermo Fisher, Waltham, MA, USA), and pooled in equimolar concentrations. Paired-end sequencing (2 × 300 bp) was performed on the Illumina MiSeq PE300 platform (Illumina, San Diego, CA, USA) at Majorbio Bio-Pharm Technology Co., Ltd (Shanghai, China).

Bioinformatics and Taxonomic Annotation: Raw sequences were processed using QIIME2 (v2022.8) with DADA2 denoising to identify high-resolution amplicon sequence variants (ASVs). Sequences were clustered with 100% similarity, and chimeras were removed. Taxonomic annotation of ASVs was conducted against the SILVA 138.1 database using a naive Bayes classifier. Sequence data quality control and processing followed the QIIME2 official pipeline guidelines.

Diversity Indices and Ecological Analysis: Alpha diversity indices (Chao1 richness, Shannon index, and Pielou’s evenness) were calculated in QIIME2. Beta diversity was assessed using Bray–Curtis dissimilarity and visualized via non-metric multidimensional scaling (NMDS) and principal coordinate analysis (PCoA), conducted in R (v4.2.1) with the vegan and ggplot2 packages. Functional annotation of prokaryotic taxa was inferred using the Functional Annotation of Prokaryotic Taxa (FAPROTAX) database.

Community Assembly and Environmental Correlations: To explore community assembly mechanisms, β-nearest taxon index (βNTI) and Raup-Crick dissimilarity (RC_bray) were calculated as described by Stegen et al. (2013) using the picante and iCAMP packages [21]. Redundancy analysis (RDA) and variance inflation factor (VIF) diagnostics were performed to identify key environmental drivers of microbial composition. These multivariate analyses clarified the links between biochar-modulated soil conditions and bacterial assembly patterns.

2.7. Statistical Analysis

All quantitative data were expressed as mean ± standard deviation (SD). Prior to statistical testing, assumptions of normality and homogeneity of variance were verified using the Shapiro–Wilk and Levene’s tests, respectively, in R (v4.2.1). One-way analysis of variance (ANOVA) was performed to evaluate the effects of biochar treatments on soil carbon fractions, physicochemical properties, and microbial diversity indices.

Where ANOVA indicated significant effects (p < 0.05), Tukey’s honestly significant difference (HSD) test was used for post hoc comparisons if homogeneity of variances was met. Otherwise, the Games-Howell test was applied. Pearson correlation analysis was conducted to explore associations among biochar physicochemical traits, soil carbon pools, and microbial indices.

To identify key environmental drivers of microbial community structure and carbon stabilization, redundancy analysis (RDA) and variance inflation factor (VIF) diagnostics were conducted using the vegan package in R. All graphical outputs, including boxplots, cumulative CO₂ mineralization curves, heatmaps, and ordination plots (e.g., PCoA, RDA), were generated using ggplot2 and associated visualization libraries in R.

A replication size of n = 4 per treatment group was selected based on previous peer-reviewed studies in soil incubation and microbial ecology. This replication strategy has been shown to provide sufficient statistical power to detect treatment-level differences while remaining practical for resource-intensive biochar studies.

3. Results

3.1. Morphological and Structural Properties of Biochars

The morphological and structural characteristics of biochars play a pivotal role in their adsorption behavior and carbon stabilization potential. Biochars prepared at 400 °C, 500 °C, and 600 °C exhibited distinct physical and chemical transformations that reflect pyrolysis-driven evolution of pore structures, surface chemistry, and aromatic ordering.

3.1.1. Surface Morphology and Pore Development

SEM images (Figure 2) revealed pronounced changes in the surface structure of S. rostratum biomass after pyrolysis. The raw straw (SR) displayed compact, fibrous tissue with no observable porosity. In contrast, the SRD400 biochar (400 °C) showed initial pore formation and surface fragmentation. The SRD500 biochar (500 °C) exhibited the most developed mesopores, with widespread distribution of spherical microvoids and fragmented walls. At 600 °C (SRD600), some surface sintering and pore collapse were observed, suggesting overheating may compromise pore integrity.

Notably, arrows in Figure 2 indicate characteristic structures such as fragmented walls, developed mesopores, and sintered surfaces. These morphological transitions suggest that SRD500 provides the most favorable surface for sorption due to its rich mesopore structure and roughened surface topography, which may enhance both physical adsorption and microbial colonization.

Previous studies on herbaceous feedstock-derived biochars report BET areas ranging from 20 to 60 m²/g at 400 °C, and up to 150–200 m²/g at 500–600 °C. Based on morphological trends, SRD500 likely corresponds to a peak in mesoporosity and surface area.

3.1.2. Surface Functional Groups

FTIR spectra (Figure 3a) showed a progressive loss of oxygen-containing functional groups with increasing pyrolysis temperature. In SRD400, peaks were observed at 3326 cm⁻¹ (–OH), 2928 cm⁻¹ (C–H), 1771 cm⁻¹ (C=O), 1580 cm⁻¹ (aromatic C=C), 1388 cm⁻¹ (C–OH), and 1008 cm⁻¹ (C–O), indicating the presence of polar groups relevant for chemical sorption. These signals diminished notably in SRD500 and were nearly absent in SRD600, confirming progressive deoxygenation.

The reduction in functional group abundance reflects a trade-off between chemical and physical sorption: SRD400 is rich in polar moieties for chemical adsorption, while SRD500 and SRD600 show increased structural stability and hydrophobicity, favoring physical adsorption.

3.1.3. Aromaticity and Carbon Ordering

Raman spectroscopy (Figure 3b) revealed the evolution of carbon structural order. The intensity ratio of the D-band (disordered carbon) to G-band (graphitic carbon), ID/IG, was 1.47 for SRD400, 0.89 for SRD500, and 1.08 for SRD600. The sharp decrease in ID/IG from 400 to 500 °C reflects enhanced graphitization and ordering, while the slight rebound at 600 °C may indicate localized structural collapse.

These observations suggest that SRD500 exhibits the highest degree of structural order, which may promote π–π interactions and van der Waals adsorption of organic molecules.

3.1.4. Crystallinity and Aromatic Condensation

XRD patterns (Figure 3c) displayed broad humps at ~22–24° (2θ), characteristic of amorphous carbon. Sharp peaks corresponding to SiO₂, graphite-like structures, and minor CaCO₃ phases became more defined at higher pyrolysis temperatures. Aromatic condensation is inferred from the narrowing and intensification of these peaks, particularly in SRD500 and SRD600, which is favorable for enhancing the carbon sequestration potential of biochars through increased chemical stability.

3.1.5. Biochar-Derived Carbon Pool Distribution

The distribution of carbon pools in S. rostratum-derived biochars was strongly influenced by pyrolysis temperature (Figure 4). At 400 °C (SRD400), approximately 32.1% of the initial biomass carbon was retained as stable carbon, which increased slightly to 34.6% and 34.2% at 500 °C (SRD500) and 600 °C (SRD600), respectively. The oxidized carbon fraction (representing chemically reactive carbon species prone to microbial transformation) remained relatively low across treatments, ranging from 4.6% in SRD400 to 2.7% in SRD600. In contrast, thermal decomposition accounted for the majority of carbon loss, with volatilization-related losses increasing slightly at higher temperatures.

These trends align with the increasing aromatic condensation and graphitization observed in spectroscopic analyses (Section 3.1.2 and Section 3.1.3), supporting the conclusion that high-temperature biochars enhance carbon stability through structural recalcitrance. However, the modest increase in stable carbon fraction between 500 °C and 600 °C suggests diminishing returns in carbon retention at very high temperatures, possibly due to pore collapse and loss of surface area.

This compositional shift in carbon pools confirms that pyrolysis at 500 °C achieves a favorable balance between maximizing stable carbon content while preserving porous structure and adsorption capacity-properties essential for long-term carbon sequestration in soil.

3.2. Effects of Biochar on Soil Carbon Dynamics and Physicochemical Properties

3.2.1. Soil TOC, pH, Moisture, and TC

The application of S. rostratum-derived biochars significantly modified key soil properties (Figure 5a). Total carbon (TC) increased from 5.9% in the control soil (SS) to 8.1%, 8.3%, and 7.1% under BC400, BC500, and BC600 treatments, respectively. Similarly, total organic carbon (TOC) content rose from 1.08% in SS to 5.88%, 6.83%, and 3.91% in the respective biochar treatments. The highest TOC and TC values were observed in the BC500 treatment, suggesting improved carbon retention driven by sorptive interactions between biochar surfaces and labile organic matter.

Soil gravimetric moisture content also increased markedly from 10.4% (SS) to 14.2% and 14.3% in BC400 and BC500 treatments, respectively, likely due to the porous structure and hydrophilic functional groups of the biochar facilitating water retention. pH remained within a slightly alkaline range across treatments, increasing slightly from 7.8 (SS) to 8.1–8.3 in biochar-amended soils.

These results demonstrate that BC500 in particular achieved a favorable combination of surface activity, moisture retention, and pH buffering, supporting enhanced carbon stabilization potential in the soil matrix.

3.2.2. Stability of Soil Carbon Pools

The SCP, modeled via a two-pool kinetic approach (Figure 5b), increased from 31.3% in SS to 50.2%, 52.4%, and 44.3% under BC400, BC500, and BC600 treatments, respectively. Among these, BC500 showed the highest SCP proportion, indicating enhanced protection of carbon via sorptive and structural stabilization mechanisms. The improved SCP in BC500 likely results from a synergistic balance between its condensed aromatic framework and residual functional groups, which both support carbon retention.

3.2.3. SOC Mineralization Dynamics

Temporal measurements of SOC mineralization rate (R_SOC) revealed peak values around days 10–20 across all treatments (Figure 5c). The highest rates were recorded under BC400, reaching up to 0.18 mg CO₂ g⁻¹ soil d⁻¹, followed by BC500 and BC600. In contrast, the control soil (SS) exhibited lower mineralization rates throughout, indicating reduced microbial stimulation in the absence of biochar.

Cumulative carbon mineralization (M_SOC) over 60 days followed a similar trend (Figure 5d). BC400 released the highest amount of CO₂ at 1.37 mg CO₂ g⁻¹ soil, while BC500 and BC600 released 1.28 and 1.13 mg CO₂ g⁻¹ soil, respectively. SS remained lowest at 0.74 mg CO₂ g⁻¹ soil. These results suggest that biochars, particularly BC400, stimulated microbial activity and co-metabolism of labile carbon, while BC500 offered a moderate balance between mineralization and carbon stabilization.

3.3. Soil Physicochemical Properties and Their Linkages to Carbon Stabilization

Pearson correlation analysis (Figure 6) revealed strong positive associations between the SCP and other soil parameters. SCP showed nearly perfect correlations with TOC (r = 0.99), moisture content (r = 0.98), and SOC mineralization rate R_SOC (r = 0.99), suggesting that higher TOC and water availability promote the adsorption and retention of labile organic matter on biochar surfaces.

In contrast, cumulative mineralization (M_SOC) was more strongly correlated with pH (r = 0.97) and TC (r = 0.93), implying that bulk carbon content and alkalinity may facilitate overall microbial activity and decomposition, but are less directly tied to short-term stabilization via adsorption.

These findings support the hypothesis that biochar facilitates carbon sequestration not only through direct carbon input but also via the physical and water-mediated protection of labile organic substrates, reducing their accessibility to microbial degradation and promoting longer-term stability in soil systems.

3.4. Biochar-Mediated Adsorptive Filtering of Microbial Communities

Application of S. rostratum-derived biochars significantly influenced soil microbial community diversity and composition, consistent with an adsorption-mediated filtering effect (Figure 7). Among alpha diversity indices, Pielou evenness decreased significantly in the BC400 treatment (p = 0.049) compared to the control (SS), suggesting a decline in community uniformity possibly driven by selective colonization under lower-temperature biochars (Figure 7c). In contrast, Chao1 richness (Figure 7a) and Shannon diversity (Figure 7b) did not show significant differences across treatments, indicating that species richness remained largely unchanged despite shifts in distribution.

Multivariate ordinations provided further evidence of biochar-induced restructuring of microbial assemblages. Principal Coordinates Analysis (PCoA; Figure 7d) and Non-Metric Multidimensional Scaling (NMDS; Figure 7e) both revealed distinct treatment-specific clustering. Notably, microbial communities in BC500 and BC600 formed separate clusters from those in SS and BC400, highlighting the influence of pyrolysis temperature on microbial community assembly.

These divergences are attributed to the changing physicochemical properties of biochars with increasing pyrolysis temperature. As biochar surfaces become more aromatic, hydrophobic, and porous, they may selectively retain or exclude microbial taxa through physical adsorption and substrate availability. This “adsorptive filtering” likely alters microbial habitats directly while concurrently modifying soil conditions such as TOC and pH, reinforcing niche-based community structuring.

3.5. Microbial Taxonomic Shifts and Functional Responses to Biochar Amendments

Biochar treatments caused marked changes in both the taxonomic composition and functional potential of soil bacterial communities (Figure 8a,b). At the phylum level (Figure 8a, left), Actinobacteriota, Pseudomonadota (formerly Proteobacteria) [32], and Acidobacteriota dominated across all treatments. Biochar amendments, especially at 500 °C and 600 °C, increased the relative abundance of Actinobacteriota (from 26.3% in SS to 34.8% in BC500) and Chloroflexi (from 3.9% to 6.7%), both associated with aromatic compound degradation and environmental stress resilience. In contrast, Pseudomonadota declined with rising pyrolysis temperature (from 28.1% in SS to 18.7% in BC600), indicating a potential reduction in copiotrophic taxa favored by labile carbon.

At the genus level (Figure 8a, right), Solirubrobacter, Amycolatopsis, and Nocardioides_A were notably enriched under BC500 and BC600, taxa known for their capacity to degrade complex organic matter and tolerate adsorptive matrices. Genera such as Sphingomonas and Blastococcus, linked to pollutant degradation and biofilm formation, were also favored under high-temperature biochars, suggesting enhanced recruitment of specialized functional groups adapted to modified microhabitats.

FAPROTAX-based functional predictions (Figure 8b) revealed enriched pathways related to carbon cycling, including methanotrophy, methylotrophy, and aerobic degradation of aromatic compounds, particularly in BC500 and BC600. Simultaneously, functions associated with nitrification and human pathogens were suppressed. Additional enrichment was observed in xylanolysis, cellulolysis, and aerobic chemoheterotrophy, reinforcing the idea that biochar not only reshapes microbial community identity but also modulates ecological functionality. Despite the limitations of predictive tools like FAPROTAX, these trends strongly suggest that biochar amendments induce compositional and functional filtering through combined effects of surface chemistry and substrate adsorption.

3.6. Environmental Determinants and Assembly Processes of Soil Microbial Communities

To further dissect the drivers of microbial community shifts, we assessed deterministic and stochastic assembly processes using β-nearest taxon index (βNTI), Raup-Crick Bray–Curtis dissimilarity (RC_bray), and redundancy analysis (RDA) (Figure 9a–c).

Across all treatments, βNTI values were consistently below −2 (Figure 9a), indicating that deterministic processes (i.e., homogeneous selection) predominantly governed microbial community assembly. Notably, the most negative values occurred under BC600 (−3.1 ± 0.4), suggesting intensified environmental filtering due to the altered physicochemical properties of high-temperature biochars.

RC_bray values centered around or slightly above zero (Figure 9b), further supporting a model of niche-based assembly. Greater variance in RC_bray under BC500 and BC600 implies enhanced niche differentiation and adsorption-driven selection.

RDA (Figure 9c) revealed that SCP, soil moisture, and pH were the most influential environmental variables shaping microbial communities. SCP and moisture were positively associated with BC400 and BC500, suggesting that these treatments offered more favorable conditions for microbial colonization and activity. In contrast, BC600 formed a distinct cluster, likely reflecting lower availability of labile carbon and altered adsorption dynamics.

Collectively, these results confirm that biochar-induced changes in soil conditions, especially via adsorption mechanisms, enhance deterministic microbial assembly and reduce stochastic variability, reinforcing the role of biochar as both habitat modifier and selective microbial filter.

4. Discussion

4.1. Pyrolysis Temperature and Biochar Properties

The physicochemical characteristics of biochar, such as surface functionality, porosity, and aromaticity, are primarily governed by pyrolysis temperature. As pyrolysis temperature increases, there is a marked loss of oxygen-containing surface groups and volatile compounds, which results in the progressive aromatization of the carbon matrix and development of micropores. Numerous studies have documented that high-temperature biochars (≥500 °C) typically exhibit increased BET surface area and greater pore volume, while low-temperature biochars (≤400 °C) retain more labile compounds and polar surface functionalities such as –OH and –COOH groups [33,34].

Our findings are consistent with this trend: biochar produced at 500 °C (SRD500) displayed the most favorable balance between pore development and surface chemistry, thereby enhancing both physical adsorption and environmental persistence. SEM imaging and spectroscopic data revealed substantial pore widening and fragmentation at 500 °C, while FTIR and Raman spectra confirmed the evolution of both aromatic structures and partial retention of functional groups. These structural features are considered critical for supporting carbon stabilization and microbial colonization.

Moreover, the results align with Barszcz et al. (2024), who reported that steam-activated biochars produced at 700–800 °C exhibited higher surface areas and graphitic carbon domains, whereas biochars at 300 °C showed more oxygenated functionalities and less developed porosity [35]. Thus, pyrolysis at intermediate temperatures may provide a structurally and chemically balanced biochar, capable of supporting both adsorption-driven carbon retention and biologically favorable interfaces.

4.2. Adsorption by Biochar: Surface Chemistry and Porosity

Beyond structural porosity discussed previously, biochar’s adsorption capacity is strongly governed by its surface chemical functionalities. While micropores and pore hierarchy facilitate physical entrapment of solutes, chemical groups such as hydroxyl (–OH), carboxyl (–COOH), and carbonyl (C=O) mediate key interactions, hydrogen bonding, electrostatic attraction, and ligand exchange, with a broad spectrum of polar compounds and nutrients [35,36]. Notably, high-temperature pyrolysis promotes the development of aromatic domains, enhancing hydrophobic interactions and contributing to overall sorptive performance.

The adsorption potential of biochar spans multiple spatial scales. Biochar derived from vascular plant tissues exhibits a combination of micropores (<2 nm), mesopores (2–50 nm), and macropores (>50 nm), forming a heterogeneous internal surface area that can exceed hundreds of m²/g in high-temperature materials [36]. This diversity enables the physical entrapment of small molecules in micropores, while meso- and macropores can host larger substrates such as enzymes, microbial exo-metabolites, or even small microbial cells. These microsites act as adsorptive niches where resource availability can be elevated or restricted [36].

Recent findings show that biochar’s sorptive surfaces also mediate nutrient and carbon dynamics by creating localized hotspots of dissolved organic carbon (DOC) and root-derived exudates [37]. For instance, added glucose has been shown to preferentially adsorb to biochar surfaces, boosting microbial assimilation but simultaneously reducing the availability of labile carbon in bulk soil, a phenomenon termed negative priming [28]. Furthermore, the adsorption and entombment of microbial residues (necromass) in biochar micropores can significantly delay decomposition and promote long-term stabilization of microbial-derived organic matter.

Therefore, biochar functions as a sorbent matrix that mimics organo-mineral associations commonly seen in natural soils. These properties are particularly relevant in degraded or nutrient-poor soils, where the preservation of labile carbon and enhancement of microbial microhabitats are critical for rebuilding soil health and carbon stocks.

4.3. Adsorptive Filtering of Microbial Communities

The adsorption properties of biochar extend beyond physicochemical processes to influence the composition and assembly of soil microbial communities. By sequestering soluble organic compounds and nutrients onto its surface, biochar generates microsites of differential resource availability, creating spatial heterogeneity in soil that shapes microbial habitat quality and competition [37,38]. This localized enrichment at the biochar-soil interface can favor particle-associated microbes that are adapted to low-nutrient or adsorptive environments, often promoting the development of structured biofilms within pore spaces or surface microzones [39].

Such environments select taxa with specialized traits, such as high carbon use efficiency, the capacity to exploit adsorbed carbon via exoenzymatic activity, or tolerance to osmotic and oxidative stresses. These may include oligotrophic bacteria and fungi capable of utilizing adsorbed low-molecular-weight compounds [37]. For example, Solirubrobacter and Amycolatopsis, enriched in our BC500 and BC600 treatments, have been reported in other studies to be dominant under carbon- or nutrient-constrained conditions and associated with aromatic compound degradation [39,40].

From a community ecology perspective, this selection process corresponds to a shift from stochastic to deterministic assembly. Increasing biochar dosage or surface reactivity tends to suppress random colonization and enhance environmental filtering. Meta-analyses and field experiments consistently report reduced β-nearest taxon index (βNTI) variance and values <−2 following biochar application, indicating stronger homogeneous selection across microbial replicates [39,41]. Lei et al. (2023) demonstrated that increasing biochar content decreased the relative contribution of stochastic processes in microbial community assembly, favoring convergence on stress-adapted and adsorption-tolerant microbial strategies [42]. These observations align with the concept of deterministic microbial assembly, wherein environmental conditions such as nutrient heterogeneity, redox status, or sorptive microhabitats exert selective pressures on microbial colonization. In contrast to stochastic (random) community dynamics, deterministic processes are often inferred from significantly negative β-nearest taxon index (βNTI) values (e.g., βNTI < −2), which indicate phylogenetically similar communities arising from similar environmental filters. In the context of biochar-amended soils, such filters include sorptive surfaces, resource hotspots, and abiotic stress gradients.

In this way, biochar functions as an ecological filter: by altering soil pH, moisture, redox conditions, and adsorptive microhabitats, it constrains microbial colonization to taxa that can tolerate or exploit its surface properties. This niche-based filtering is fundamental to biochar’s role in modifying soil microbiomes and has cascading implications for ecosystem processes such as carbon cycling and nutrient retention.

4.4. Carbon Stabilization Mechanisms

Biochar stabilizes soil organic carbon (SOC) through a dual mechanism: physical protection and microbial mediation. Physically, biochar’s porous structure and surface chemistry adsorb dissolved organic matter (DOM), root exudates, and microbial metabolites, reducing the accessibility of labile carbon to decomposers and promoting a “negative priming effect” in which native SOC decomposition is suppressed [39]. This phenomenon results in the sequestration of carbon in biochar-associated pools, which are less prone to microbial mineralization [28,43].

In addition, biochar forms organo-mineral complexes within soil aggregates, providing a stable environment for microbial colonization. These complexes, particularly biochar-associated SOC, are chemically recalcitrant and spatially protected from decomposition by microorganisms [39]. This protection is further enhanced by biochar’s ability to promote microbial carbon use efficiency (CUE), wherein sorbed low-concentration substrates support efficient microbial metabolism, reducing carbon respiratory losses and fostering biomass production.

The eventual death of these microorganisms contributes to microbial necromass, primarily consisting of amino sugars and cell wall components, which becomes physically trapped within biochar’s pores. Studies by Kalu et al. (2024) and Wang et al. (2025) have shown that high-temperature biochars sequester microbial necromass over time, contributing to a stable carbon pool that enhances long-term SOC retention in the soil [44,45].

Overall, these processes underline biochar’s unique ability to stabilize carbon through both physical adsorption and biological conversion, reinforcing its potential for enhancing soil carbon persistence and its application in sustainable carbon management strategies.

4.5. Biochar-Modified Soil Environment and Community Outcomes

In addition to direct adsorption effects, biochar significantly modifies the soil’s abiotic environment, indirectly influencing microbial persistence, diversity, and ecological functioning. One of the most prominent changes is soil pH elevation due to biochar’s inherent alkalinity and ash content, a phenomenon widely recognized as the liming effect. This pH shift can suppress acidophilic taxa and favor neutrophilic or alkaliphilic bacteria, including many nitrifiers and ammonia oxidizers, thereby reshaping community structure [32]. However, as highlighted by the reviewer, pH effects may confound microbial responses attributed to sorption. Future studies should consider pH-adjusted controls or partitioning techniques (e.g., partial RDA) to disentangle the influence of pH from adsorption-driven changes.

Biochar also improves soil cation exchange capacity (CEC) and nutrient retention, enhancing the availability of essential ions such as NH₄⁺, K⁺, Ca²⁺, and PO₄³⁻ in localized microsites [43]. These nutrient-rich patches can favor copiotrophic taxa (e.g., Proteobacteria/Pseudomonadota), which thrive in high-nutrient environments. However, this effect is bidirectional: the strong sorption capacity of biochar may also immobilize nutrients and render them less available to certain microbes, creating resource bottlenecks that select for specialized taxa.

Moreover, biochar’s porous matrix contributes to improved soil moisture retention, particularly in sandy or coarse-textured soils. This water-holding capacity creates buffered microenvironments that protect microbial cells from desiccation during drought or drying-rewetting cycles. As shown in our study, moisture content was positively correlated with microbial evenness and SCP size, reinforcing the role of water availability in shaping microbial resilience. This finding aligns with reports that biochar enhances hydrological function by improving pore structure and water retention [46].

Taken together, biochar modifies multiple abiotic factors, pH, nutrient availability, and water retention, simultaneously. These abiotic shifts serve as additional environmental filters (see Section 4.3), reinforcing deterministic assembly trajectories observed in microbial communities.

4.6. Implications and Future Directions

This study demonstrates that the capacity of biochar to stabilize soil organic carbon (SOC) and shape microbial community composition is closely tied to its adsorption-mediated properties. Biochars, particularly those produced at intermediate pyrolysis temperatures, combine high surface area, hierarchical porosity, and oxygen-containing functional groups, which enable the sequestration of labile organic matter, dissolved nutrients, and water within protected microhabitats. These microsites promote deterministic bacterial assembly, favoring carbon-efficient and stress-resilient taxa while enhancing microbial carbon use efficiency (CUE) and necromass production [47]. Collectively, these mechanisms contribute to the persistence of SOC and the modulation of biogeochemical cycling.

Importantly, the results of this study highlight that adsorption-mediated processes such as sorptive retention, niche filtering, and microhabitat formation underlie key ecological functions attributed to biochar, namely carbon stabilization, pollutant immobilization, and microbial reprogramming. These findings align with recent empirical evidence that points to sorption as a central mechanism in biochar-soil-microbe interactions.

However, several limitations must be acknowledged. First, microbial functional profiles were predicted using FAPROTAX, which infers potential functions based on taxonomic identity and lacks the resolution of direct metagenomic or metatranscriptomic measurements. This limits confidence in pathway-level interpretations. Future studies should validate these predictions using shotgun metagenomics, stable isotope probing, or enzyme activity assays to more directly link microbial identity with function.

Second, the 60-day incubation period may not fully capture long-term stabilization processes or successional dynamics of microbial communities. While our results offer insights into short- to mid-term mechanisms, longer incubation or field trials are essential to verify the durability of carbon retention and the persistence of deterministic community assembly.

Third, although this study focuses on biochar derived from the invasive weed S. rostratum, comparisons with other biochar feedstocks were not systematically addressed. It remains unclear whether the observed adsorption and microbial filtering effects are unique to this biomass type or shared across biochars from other invasive species. Comparative studies with different lignocellulosic or weedy biomasses (e.g., Eichhornia crassipes, Imperata cylindrica) would help contextualize the generality of our findings.

Lastly, while adsorption is a key mechanism, its interaction with other abiotic changes such as pH modification may also shape microbial responses. Future work should consider using pH-buffered controls or partial redundancy analysis (RDA) to partition the relative contributions of sorption and pH-driven effects.

These considerations suggest that future research should prioritize (i) mechanistic validation of microbial functional predictions, (ii) multi-year field studies to assess persistence, and (iii) comparative evaluations across diverse biochar feedstocks. Integrating physicochemical, microbial, and ecological assessments will better inform the design of biochars optimized for carbon sequestration and soil rehabilitation in degraded or marginal ecosystems.

5. Conclusions

This study reveals that biochars derived from the invasive plant S. rostratum, particularly those produced at intermediate pyrolysis temperatures (notably 500 °C), can significantly enhance soil carbon stability and restructure microbial communities via adsorption-mediated mechanisms. Biochars at 500 °C exhibited an optimal combination of aromaticity, porosity, and residual surface functional groups, which promoted the formation of SCPs and deterministic microbial assembly. Our results emphasize the central role of adsorption in biochar function. Specifically, the immobilization of labile organic compounds on porous biochar surfaces reduced their mineralization, while selective microbial filtering enhanced the abundance of carbon-cycling taxa and related functional pathways. These dual effects resulted in lower SOC mineralization rates, higher stable carbon proportions, and environmentally driven microbial community shifts, demonstrated by βNTI-based deterministic assembly. Importantly, this study highlights that biochar’s carbon stabilization potential is not solely a function of its chemical composition, but also of its capacity to act as an ecological filter. These insights offer a valuable design principle: tailoring biochar porosity and surface chemistry to enhance both abiotic sorption and biotic filtering can maximize carbon sequestration outcomes. We recommend the targeted application of 500 °C S. rostratum-derived biochars in degraded or carbon-depleted soils as a sustainable strategy to enhance long-term carbon retention and support microbial ecosystem function. Such biochars offer an eco-friendly solution for managing invasive biomass while contributing to soil restoration and climate mitigation.

Future studies should investigate long-term field-scale dynamics of sorption and microbial turnover, integrate metagenomic validation of microbial functions, and explore interactions with plant-microbe-biochar systems across diverse soil types. Advancing adsorption-oriented biochar design will be critical for optimizing soil health and carbon management under climate change scenarios.