Mitigating Nicosulfuron Injury and Remediating Herbicide Residues in Maize Fields: Degradation by the Immobilized Microbial Agent IM-2NN and Soil Metabolomic Analysis

Abstract

Long-term nicosulfuron application can cause herbicide injury in maize and lead to residue accumulation in agricultural soils. To develop an effective strategy for reducing nicosulfuron toxicity and promoting soil recovery, we combined two highly efficient nicosulfuron-degrading bacterial strains, Klebsiella jilinsis 2N3 and Serratia marcescens N80, at a 1:1 ratio. The mixed bacterial culture achieved a nicosulfuron degradation rate of 92.15%. To improve its stability and field applicability, the strains were immobilized on a biochar carrier prepared from discarded fungal substrate, generating the immobilized microbial agent IM-2NN. This formulation significantly enhanced the environmental adaptability and stress tolerance of both strains. Flow cytometry further showed that immobilization prolonged the colonization period and increased the abundance of strains 2N3 and N80 in maize rhizosphere soil. Analysis of nicosulfuron degradation products in maize field soil indicated that IM-2NN may degrade nicosulfuron primarily through sulfonylurea bridge cleavage and hydrolysis of amide bonds. Soil metabolomic analysis showed that IM-2NN application altered several metabolic pathways, including tryptophan and pyruvate metabolism, suggesting that microbial remediation may support the functional recovery of contaminated soils by regulating amino acid metabolism and energy-conversion networks. In maize seedling assays, IM-2NN also alleviated nicosulfuron-induced phytotoxicity by increasing antioxidant enzyme activities and reducing malondialdehyde accumulation, thereby effectively limiting oxidative damage during detoxification. Together, these findings show that IM-2NN can efficiently degrade nicosulfuron residues, improve soil microecological function, and alleviate herbicide-induced stress in maize. This study provides a practical and environmentally friendly microbial strategy for remediating nicosulfuron-contaminated maize fields and supporting sustainable maize production.

Keywords

maize / Klebsiella jilinsis 2N3 / Serratia marcescens N80 / nicosulfuron / immobilized mixed bacterial agent / metabolomics

1 INTRODUCTION

Nicosulfuron is a sulfonylurea herbicide widely used in maize fields to control annual and perennial weeds. Since its introduction in China in 1998, it has become a common post-emergence herbicide because of its high selectivity, broad weed-control spectrum, and low mammalian toxicity (Liu et al., 2022). However, improper or excessive application can cause serious phytotoxicity in maize, including delayed growth, impaired root development, inhibited reproductive development, reduced yield and quality, and, in severe cases, complete crop loss. The severity of nicosulfuron injury depends on multiple agronomic and environmental factors, including application dose, timing, maize genotype, growth stage, and weather conditions (Wu et al., 2022; Silva et al., 2020; Li et al., 2022). Mechanistically, nicosulfuron inhibits acetolactate synthase (ALS), interferes with photosynthesis, and induces oxidative stress, thereby suppressing plant growth and yield formation. Differences in herbicide tolerance among maize varieties are associated mainly with ALS sensitivity, metabolic detoxification capacity, including cytochrome P450 and glutathione S-transferase activities, and the strength of antioxidant responses. Resistant varieties can partially alleviate herbicide injury by regulating physiological processes and stress-related gene expression, whereas sensitive varieties are more prone to irreversible damage (Lu et al., 2024). In addition to its direct effects on maize, overuse of nicosulfuron can lead to residue accumulation in crops, soil, and water, posing risks to crop safety and environmental health (Kniss, 2017; Xiao et al., 2024). Therefore, effective remediation strategies are urgently needed to reduce nicosulfuron residues, protect maize productivity, and support the sustainable management of maize production systems.

Because the photolytic degradation efficiency of sulfonylurea herbicides is generally low (Benzi et al., 2011), nicosulfuron degradation in soil and water occurs mainly through microbial degradation and chemical hydrolysis. Among these processes, microbial degradation is regarded as an environmentally friendly approach for reducing residual sulfonylurea herbicides in agricultural environments (Li et al., 2022; Lei et al., 2023). Microorganisms are particularly suitable for bioremediation because of their small size, wide distribution, rapid reproduction, strong adaptability, and diversity. These characteristics allow microbial degradation to overcome several limitations of physical and chemical remediation methods and to improve the efficiency of nicosulfuron removal (Zhong et al., 2023). In general, microbial degradation depends on enzymes and other metabolites that directly or indirectly modify the nicosulfuron structure, ultimately reducing or eliminating its biological activity (Zhang et al., 2020e). To date, several nicosulfuron-degrading strains have been reported, including Pseudomonas nitroreducens NSA02 (Zhao et al., 2018), Bacillus velezensis CF57 (Zhang et al., 2020c), Pseudomonas aeruginosa B9 (Pourbabaee et al., 2018), Bacillus subtilis YB1 (Zhang et al., 2020d), Klebsiella sp. Y1 (Wang et al., 2016), Alcaligenes faecalis ZWS11 (Zhao et al., 2015), Pseudomonas fluorescens SG-1 (Carles et al., 2017), and Aspergillus niger YF1 (Lu et al., 2012). These findings indicate that microorganisms provide an important biological resource for nicosulfuron remediation, although the performance of individual strains may remain limited under complex field conditions.

To improve degradation efficiency and environmental robustness, increasing attention has been given to mixed microbial communities. Compared with single strains, mixed cultures can combine diverse metabolic capacities, enzyme systems, and ecological functions, enabling cooperative degradation or co-metabolism of organic pollutants (Yu et al., 2020; Yang et al., 2021). Such microbial interactions may accelerate the breakdown of nicosulfuron and increase tolerance to high pollutant concentrations or adverse environmental conditions (Yuan et al., 2016; Li et al., 2021). Previous studies have shown that mixed cultures often outperform individual strains. For example, a mixed culture of Klebsiella variicola FH1 and Arthrobacter sp. NJ-1 degraded 98.23% of 50 mg/L atrazine within 9 days at pH 9 and 30 ℃, with a cell concentration of 1.6 × 10⁸ CFU/mL, whereas strain FH1 alone degraded only 81.5% after 11 days (Zhang et al., 2019b). Similarly, Enterobacter sp. BD19 and E. cloacae complex sp. BD17 individually degraded 71.10% and 76.19% of 200 mg/L pyridine within 72 h, respectively, whereas their mixed culture achieved 91.70% degradation within 36 h (Nie et al., 2021). In nicosulfuron remediation, the mixed cultivation of B. subtilis YB1 and A. niger YF1 also achieved a substantially higher degradation rate than either single culture under the same conditions (Lu et al., 2012). These examples suggest that mixed bacterial systems are a promising strategy for enhancing the bioremediation of herbicide-contaminated soils.

However, the field application of free microbial cells is often constrained by poor survival, unstable colonisation, and reduced activity under environmental stress. Immobilisation provides a practical way to overcome these limitations by protecting microbial cells and improving their persistence in soil. Biochar is a particularly effective immobilisation carrier because of its large specific surface area, strong adsorption capacity, and high porosity, which help maintain high microbial density and permeability within the remediation system (Zhang et al., 2020b). After immobilisation on biochar, microorganisms generally show improved tolerance to toxic pollutants and reduced sensitivity to fluctuations in pH, temperature, and salinity (Chen et al., 2022). In addition, polyvinyl alcohol (PVA) and sodium alginate (SA) have been widely used as immobilisation matrices for the removal of target pollutants (Wen et al., 2021; Chen et al., 2021; Xiang et al., 2022; Landreau et al., 2020). For instance, Bacillus cereus WL08 immobilised in a bamboo charcoal-SA matrix efficiently degraded dimethomorph in both soil and water. Immobilisation also improved the tolerance of WL08 to adverse conditions and increased its stability during reuse and storage (Zhang et al., 2020b). Together, these findings indicate that combining biochar with polymer-based immobilisation materials may improve both microbial performance and practical applicability in contaminated soils.

In our previous studies, two highly efficient nicosulfuron-degrading bacterial strains, Klebsiella jilinsis 2N3 and Serratia marcescens N80, were isolated under controlled laboratory conditions (Zhang et al., 2010; Zhang et al., 2012; Zhai et al., 2023; Zhang et al., 2021). Preliminary experiments showed that co-cultivation of these two strains markedly improved nicosulfuron biodegradation compared with either strain alone. Building on these findings, the present study aimed to develop an immobilised mixed-bacterial agent and evaluate its capacity to enhance nicosulfuron degradation in soil. Biochar was prepared from spent mushroom substrate at different pyrolysis temperatures, and the most suitable biochar was selected according to its adsorption capacity. The selected biochar was then combined with SA and PVA to immobilise the mixed bacterial strains 2N3 and N80, generating the immobilised microbial agent IM-2NN. We further optimised the nicosulfuron biodegradation conditions of this system, assessed the colonisation capacity of IM-2NN in soil, and evaluated its effects on nicosulfuron degradation, soil metabolic recovery, and maize phytotoxicity under contaminated conditions. This study provides a microbial remediation strategy for nicosulfuron-contaminated maize fields and offers a potential route for reusing agricultural solid waste as a carrier for immobilised bacterial consortia.

2 MATERIALS AND METHODS

2.1 Chemicals and media

Nicosulfuron was purchased from Shanghai Aladdin Biochemical Technology Co., Ltd. (Shanghai, China). All other chemicals were of analytical grade or higher. Mineral salts medium (MSM) contained the following components (g/L): 0.20 g MgSO₄·H₂O, 1.0 g KH₂PO₄, 1.0 g K₂HPO₄, 0.2 g NaCl, 5.0 g C₆H₁₂O₆, and 1.6 g NH₄NO₃. The pH was adjusted to 7.0. Lysogeny broth (LB) medium contained the following components (g/L): 10.00 g NaCl, 10.0 g tryptone, and 5.00 g yeast extract. The pH was adjusted to 7.0−7.5.

S. marcescens N80 (GenBank accession number GQ351502) and K. jilinsis 2N3 (GenBank accession number FJ422735) were provided by the Pesticide Department of Jilin Agricultural University, China. The discarded fungal substrate was supplied by the Engineering Research Center of the Chinese Ministry of Education for Edible and Medicinal Fungi, Jilin Agricultural University, China.

The test soil, classified as meadow black soil, was collected from an experimental maize field at Jilin Agricultural University. Fresh soil samples were collected from the plough layer at a depth of 0−20 cm. The samples were naturally air-dried and passed through a 40-mesh sieve before experimental analysis. The soil had a pH of 6.4 and an organic matter content of 2.65%, and had no previous history of nicosulfuron application. The maize variety used in this study was JS416.

2.2 Screening for the optimal bacterial mixing ratio and microbial culture conditions

Single colonies of strains 2N3 and N80 were selected and transferred to liquid LB medium. The cultures were incubated at 30 ℃ and 150 rpm for 12 h and then centrifuged at 7,000 rpm for 5 min. The bacterial suspensions of 2N3 and N80 were mixed at different volume ratios, namely 1:1, 2:1, 1:2, 3:2, and 2:3 (Gao et al., 2020). To evaluate degradation capacity, the mixed bacterial cultures were inoculated at 5% (v/v) into MSM containing 50 mg/L nicosulfuron. The nicosulfuron concentration in the medium was measured by high-performance liquid chromatography (HPLC) every 12 h, and the degradation rate was calculated. The optimal mixing ratio of the two bacterial strains was then selected by comparing the nicosulfuron degradation efficiencies of the different mixed bacterial cultures.

2.3 Preparation and characterization of biochar

Biochar was prepared from discarded edible mushroom culture substrate. The substrate was rinsed with distilled water to remove dust, stones, and other impurities and then air-dried. The dried material was cut into small pieces, placed in a crucible, and pyrolyzed in a box-type muffle furnace (SX2-4-10). Pyrolysis was performed at a heating rate of 10 ℃/min and maintained for 3 h at 300 ℃, 450 ℃, or 600 ℃. The resulting biochar was cooled to room temperature, rinsed with sterile water, and adjusted to neutral pH. The samples were then dried at 70 ℃, ground, and passed through a 2-mm sieve. The biochars prepared at 300 ℃, 450 ℃, and 600 ℃ were designated BC300, BC450, and BC600, respectively.

The morphological characteristics of the biochar samples were examined by scanning electron microscopy (SEM; Zeiss EVO18) (Li et al., 2022).

2.4 Adsorption of nicosulfuron by carrier materials

2.4.1 Adsorption kinetics

A sterile background solution containing 0.01 M CaCl₂ and 200 mg/L NaN₃ was prepared for the adsorption experiments. For each treatment, 0.2 g of biochar was added to 100 mL of 50 mg/L nicosulfuron solution in a 250-mL glass vial. Four treatments were established: BC300, BC450, BC600, and a blank control without biochar. The vials were incubated in a shaker at 30 ℃ and 150 rpm. Samples were collected after 1, 2, 4, 6, 8, 10, 12, 24, and 48 h, filtered through a 0.22-µm membrane filter, and analyzed by HPLC. All experiments were performed in triplicate, and the results are presented as the mean ± standard deviation (SD) (Tao et al., 2020; Wang et al., 2020).

2.4.2 Adsorption isotherms

Based on the adsorption kinetics results, adsorption isotherm experiments were conducted using initial nicosulfuron concentrations of 0.5, 5, 10, 20, and 50 mg/L. The samples were incubated in a shaker at 150 rpm under three temperature conditions: 20 ℃, 25 ℃, and 30 ℃. Adsorption was allowed to proceed for 48 h to reach equilibrium, and samples were collected at 0, 24, and 48 h. All experiments were performed in triplicate (Lonappan et al., 2018). Detailed calculation methods and equations are provided in the Supplementary Materials.

2.5 Preparation of the immobilized bacterial mixture

Based on the adsorption kinetics and isotherm results, BC600 was selected for subsequent experiments. Sterile BC600 was added to the 2NN bacterial suspension and incubated at 150 rpm for 4 h to allow bacterial adsorption onto the biochar. The mixture was then centrifuged at 10,000 rpm for 10 min to obtain bacteria-loaded biochar (BC-2NN), which was mixed with 3% SA and 6% PVA (Jiang et al., 2022). The resulting mixture was extruded dropwise into sterile 3% calcium chloride solution using a syringe and crosslinked at 4 ℃ for 24 h. Immobilized carriers without bacteria (IM) were prepared using the same procedure and stored at 4 ℃ until further use (Sun et al., 2020).

2.6 Degradation characteristics of nicosulfuron

A series of experiments was conducted to determine the optimal conditions for nicosulfuron degradation. Each assay was performed in 100 mL of MSM, with inoculum concentration, nicosulfuron concentration, pH, and incubation temperature varied as experimental variables. The tested bacterial concentrations were 4.0 × 10⁷, 8.0 × 10⁷, 1.2 × 10⁸, 1.6 × 10⁸, and 2 × 10⁸ CFU/mL. The tested nicosulfuron concentrations were 10, 20, 50, 80, and 100 mg/L. The pH values were adjusted to 5, 6, 7, 8, and 9, and the incubation temperatures were set to 20 ℃, 25 ℃, 30 ℃, 35 ℃, and 40 ℃. After incubation for 3 d at 150 rpm, the residual nicosulfuron concentration and bacterial cell density (OD₆₀₀) were measured to identify the optimal biodegradation conditions (Bhatt et al., 2022). Bacterial cell numbers in the suspensions were determined using the dilution plate method.

2.7 Cell viability assay of bacterial strains

The immobilized bacterial carrier was placed in 10 mL of phosphate buffer (pH 7.6) and incubated for 15 min with shaking to release the bacterial cells. The cells were then washed three times with precooled phosphate-buffered saline (PBS), collected, and stained using an Annexin V-PE/7-AAD apoptosis detection kit according to the manufacturer’s instructions (Beijing Solarbio Biotechnology Co., Ltd., Beijing, China) (Alimohammadi et al., 2021).

2.8 Reusability and storage stability of IM-2NN

The reusability of IM-2NN for nicosulfuron degradation was evaluated in repeated-batch degradation assays. In the first cycle, 10 g of IM-2NN was added to 100 mL of MSM containing 50 mg/L nicosulfuron and incubated for 72 h in the dark at 35 ℃ with shaking at 150 rpm. After each cycle, the IM-2NN beads were collected, washed repeatedly with sterile distilled water, and transferred to fresh MSM containing 50 mg/L nicosulfuron for the next degradation cycle. To evaluate storage stability, IM-2NN was stored at 25 ℃ for 120 d. At 20-d intervals, samples of stored IM-2NN were collected and incubated with MSM containing 50 mg/L nicosulfuron to determine the remaining degradation efficiency (Zhu et al., 2021).

2.9 Degradation of nicosulfuron in soil by 2NN and IM-2NN

The test soil was collected from the experimental field of Jilin Agricultural University. The soil had an organic matter content of 2.71% and a pH of 6.41, and had not been treated with nicosulfuron for at least two years. The ability of the immobilized bacterial agent to degrade nicosulfuron in soil was then evaluated. Nicosulfuron was added to 500 g of soil and mixed thoroughly to obtain a final concentration of 5 mg/kg. The soil water content was adjusted to 20% using sterile water. Six treatments were established: sterilized soil containing nicosulfuron (A), sterilized soil containing nicosulfuron and free 2NN bacteria (B), sterilized soil containing nicosulfuron and IM-2NN (C), unsterilized soil containing nicosulfuron (D), unsterilized soil containing nicosulfuron and free 2NN bacteria (E), and unsterilized soil containing nicosulfuron and IM-2NN (F). Sterilized soil was autoclaved at 121 ℃ for 1 h. Each treatment was incubated at 25 ℃. Soil samples (20 g) were collected after 0, 1, 3, 7, 14, 21, and 28 d to determine the residual nicosulfuron content.

To quantify nicosulfuron residues in soil, 20 g of soil was mixed with 50 mL of acetonitrile and 10 mL of water. The mixture was transferred to a 250-mL Erlenmeyer flask and shaken at 30 ℃ for 40 min. The extract was then collected by vacuum filtration and transferred to a 100-mL stoppered bottle containing approximately 8 g of NaCl. The bottle was shaken vigorously 100 times and left to stand for 30 min, then shaken again 150 times and left to stand for 1 h. The upper acetonitrile phase (25 mL) was collected and evaporated under reduced pressure at 40 ℃ using a rotary evaporator until nearly dry. The residue was dissolved in 1 mL of acetonitrile, vortexed for 1 min, filtered through a 0.22-µm PTFE filter, and analyzed by HPLC (Cueff et al., 2021).

2.10 Degradation pathway of nicosulfuron in soil

To investigate the degradation mechanism of nicosulfuron in soil, degradation products generated after IM-2NN treatment were characterized. Briefly, 20 g of nicosulfuron-contaminated soil treated with IM-2NN was mixed with 20 mL of dichloromethane in a separatory funnel and shaken vigorously for 10 min. The organic phase was collected, dehydrated with anhydrous sodium sulfate, and transferred to a round-bottom flask. The extraction was repeated three times. The dichloromethane extracts were combined and evaporated to dryness, and the residue was redissolved in 1 mL of acetonitrile. The solution was then filtered through a 0.22-μm organic-phase filter membrane before analysis of degradation products.

Degradation products were analyzed using an Agilent 6545 LC/Q-TOF system. The mass spectrometry conditions were as follows: the electrospray ionization interface was operated in both positive and negative ion modes; the voltage in positive-ion mode (ESI+) was set to 2 kV and the voltage in negative-ion mode (ESI−) was set to 1.5 kV; the capillary temperature was set to 325 ℃; the heater temperature was set to 350 ℃; the drying gas flow rate was 10 L/min; the nebulizer pressure was 35.0 psi; the drying gas temperature was 300 ℃; and the scan range was m/z 50 to 700. The compounds were identified by comparison with the LC-MS library.

2.11 Extraction and analysis of soil metabolites

Soil samples were freeze-dried and ground into a fine powder in liquid nitrogen. Ground soil samples (1.0 g) were extracted by sonication in 1 mL of methanol/water (1:1, v/v) at 4 ℃ for 10 min. The extraction was repeated three times, and the extracts were centrifuged at 10,000× g for 10 min at 4 ℃. The supernatant was transferred to a 2-mL centrifuge tube and freeze-dried. The dried sample was resuspended in 300 μL of methanol/water (1:1, v/v), vortexed for 30 s, and filtered through a 0.22-μm PTFE syringe filter. Metabolites were detected using a Q Exactive Orbitrap Plus mass spectrometer (Thermo Fisher Scientific, USA), and the data were processed and statistically analyzed using Metabolomics data were processed and statistically analyzed in R, with multivariate analysis, differential metabolite screening, heatmap visualization, pathway enrichment analysis, and metabolite−metabolite correlation network construction performed using the corresponding R packages.

2.12 Colonization of immobilized mixed bacteria in soil

The collected soil was air-dried and sterilized at high temperature to obtain sterile soil for the colonization experiment. For each treatment, 500 g of sterile soil was placed in a disposable seedling bag measuring 16 cm in diameter and 16 cm in height. Deionized water was added to maintain the soil at 60% water-holding capacity. Two treatments were established: T1, soil inoculated with free 2NN at 10⁶ CFU/g; and T2, soil inoculated with IM-2NN. Each treatment included three replicates. Soil samples were collected at 0, 1, 3, 5, 7, 14, 21, and 28 d after inoculation and stored at −80 ℃ until further analysis. To assess the colonization of 2NN and IM-2NN in soil, total DNA was extracted from the collected soil samples.

The soil samples were centrifuged at 12 000 rpm for 20 min, and the supernatant was discarded. The remaining soil material was freeze-dried to remove water. Subsequently, 0.5 g of dried soil was weighed accurately, and total DNA was extracted using the FastDNA SPIN Kit for Soil (MP Biomedicals, CA, USA) according to the manufacturer’s instructions. The extracted soil DNA was stored at −20 ℃. Specific primers were used to amplify marker genes from the genomic DNA of the tested strains and from total soil DNA. The ssuD gene of strain 2N3 was amplified using the primers ssuD-F (forward: 5'-GGCTGAGTTCCTTGTCTCTCTC-3'; reverse: 5'-CGCTGCCCGGGTGACCAGATTG-3') (Zhang et al., 2019a). The tenA gene of strain N80 was amplified using the primers tenA (forward: 5'-TCCATCACCACCCCTGTGTGCTTC-3'; reverse: 5'-AAATGCGCGACTCCAGATTGTG-3') (Hou et al., 2022). The PCR program consisted of initial denaturation at 95 ℃ for 3 min, followed by 30 cycles of denaturation at 95 ℃ for 30 s, annealing at 56 ℃ for 30 s, and extension at 72 ℃ for 90 s, with a final extension at 72 ℃ for 5 min and a hold at 4 ℃. A total of 10 μL of each PCR product was separated by electrophoresis on a 2% agarose gel. The presence of target bands was used to indirectly assess the colonization of free 2NN and immobilized IM-2NN in soil.

2.13 Remediation of nicosulfuron injury in maize

The experiment evaluating the remediation of nicosulfuron-induced phytotoxicity in maize included six treatments: S1, untreated control without nicosulfuron (CK); S2, nicosulfuron only; S3, nicosulfuron plus free mixed bacterial culture 2NN; S4, nicosulfuron plus the immobilized bacterial agent IM-2NN; S5, nicosulfuron plus a 2,500-fold dilution of brassinolide (Ma et al., 2022); and S6, nicosulfuron plus cyprosulfamide (Giannakopoulos et al., 2020). The nicosulfuron concentration in the soil of treatments S2−S6 was 0.1 mg/kg. Uniformly germinated maize seeds were selected and sown in the corresponding treatment soils. Each pot contained 20 seeds, and the plants were cultivated under controlled conditions at 25 ℃ with adequate light. On day 30, root growth was evaluated. The activities of peroxidase (POD), superoxide dismutase (SOD), and catalase (CAT), as well as the malondialdehyde (MDA) content, were measured by spectrophotometry (Singh et al., 2015; Silva et al., 2004; Generozova et al., 2022).

2.14 Chemical analysis

The nicosulfuron concentration was determined by HPLC using an Agilent 1260 system. Detection was performed at 240 nm using a UV detector and a reversed-phase C18 column (5 μm, 250 mm × 4.6 mm; Agilent Zorbax RRHD Eclipse Plus). The mobile phase consisted of acetonitrile and 0.1% acetic acid (70:30, v/v), with a flow rate of 1.0 mL/min. The column temperature was maintained at 30 ℃, and the injection volume was 10 μL.

2.15 Data analysis

The results are presented as the mean ± standard deviation (SD) for each treatment. One-way analysis of variance (ANOVA) was conducted using DPS software version 7.05 to assess significant differences among treatments. Statistical significance was set at P < 0.05. All experiments were conducted with three biological replicates.

3 RESULTS

3.1 Selection of the optimal mixing ratio of bacterial strains

The total number of introduced bacterial cells was kept constant across all treatments throughout the experiment (Fig. S1). The nicosulfuron-degrading strains 2N3 and N80 were then mixed at different volume ratios using bacterial suspensions adjusted to OD₆₀₀ = 1.0, and their degradation performance was assessed after 72 h of incubation. Among all tested combinations, the 1:1 mixture showed the strongest degradation capacity, reaching a nicosulfuron degradation rate of 92.15%. This value was higher than those observed for the other mixed-ratio treatments and clearly exceeded the degradation efficiencies of the individual 2N3 and N80 suspensions, which reached 74.22% and 75.88%, respectively. These results indicate that combining 2N3 and N80 improved nicosulfuron biodegradation compared with either strain alone, with the 1:1 ratio providing the most effective balance between the two strains. Therefore, this ratio was selected for subsequent experiments.

3.2 Characterization of biochar and IM-2NN

The characteristics of BC300, BC450, BC600, BC-2NN, and IM-2NN were examined by scanning electron microscopy (SEM; Fig. 1). Biochars produced at different pyrolysis temperatures showed clear differences in pore structure and surface morphology. BC300 had a relatively compact surface, with few visible pores or ridges (Fig. 1−A). By contrast, BC450 exhibited a rougher and more porous tubular structure (Fig. 1−B), whereas BC600 showed the most developed pore network among the three biochars (Fig. 1−C). These structural differences suggest that increasing the pyrolysis temperature promoted pore formation and improved the potential of the biochar to act as a microbial carrier. After loading with the mixed bacterial culture, bacterial cells were observed on the surface and within the pores of BC-2NN (Fig. 1−E), indicating that the biochar provided suitable attachment sites for strain retention. In IM-2NN, the bacterial cells were further embedded within immobilized beads (Fig. 1−F), which may protect the strains from environmental stress while maintaining contact with the surrounding soil environment. IM-2NN also displayed abundant wrinkles and pores, features that are favourable for pollutant adsorption, microbial colonisation, and substrate exchange. Together, these observations indicate that IM-2NN provides a suitable immobilization structure for the mixed bacterial strains and may facilitate the transport of oxygen, nutrients, and nicosulfuron during biodegradation.

3.3 Adsorption kinetics and isotherms of biochar

The adsorption behaviour of nicosulfuron differed among the three biochars and was strongly associated with pyrolysis temperature. As shown in Fig. S2A, BC300, BC450, and BC600 all adsorbed nicosulfuron rapidly during the first 1−6 h, after which the adsorption rate gradually slowed and reached equilibrium within 48 h. The adsorption kinetics fitted the pseudo-second-order model better than the pseudo-first-order model, with coefficients of determination (R²) above 0.99 (Fig. S2−B; Table S1). The equilibrium adsorption capacities of BC300, BC450, and BC600 were 2607.7, 3234.6, and 4982.7 mg/kg, respectively, indicating that a higher pyrolysis temperature markedly improved the adsorption capacity of biochar for nicosulfuron. The adsorption isotherms further confirmed this trend (Fig. S2−C). As the initial nicosulfuron concentration increased, adsorption gradually approached saturation. The experimental data fitted the Langmuir model well, suggesting that nicosulfuron adsorption on biochar mainly occurred as monolayer adsorption on relatively homogeneous adsorption sites. According to the Langmuir equation, the maximum adsorption capacities of BC300, BC450, and BC600 were 2638.98, 3305.09, and 5013.56 mg/kg, respectively, following the order BC600 > BC450 > BC300. In addition, the Freundlich nonlinearity coefficient (n) decreased with increasing pyrolysis temperature (Table S2), which was consistent with the enhanced adsorption capacity observed for biochars produced at higher temperatures. Together, these results indicate that BC600 had the strongest nicosulfuron adsorption capacity and was therefore the most suitable biochar carrier for subsequent immobilization experiments.

3.4 Degradation of nicosulfuron by free 2NN and IM-2NN

The degradation efficiencies of both free 2NN and IM-2NN were affected by bacterial concentration, initial nicosulfuron concentration, temperature, and pH (Fig. 2). Under the tested conditions, IM-2NN generally showed higher degradation activity than free 2NN, indicating that immobilization improved the stability and performance of the mixed bacterial strains. The effect of bacterial concentration was first evaluated. When the cell concentration reached 9 × 10⁷ CFU/mL, the degradation rates of nicosulfuron by free 2NN and IM-2NN reached their highest values after 72 h, at 85.73% and 90.81%, respectively (Fig. 2−A). Lower bacterial concentrations resulted in weaker degradation, likely because the number of active cells was insufficient for efficient nicosulfuron degradation. However, further increases in inoculum concentration did not improve degradation and may have limited bacterial activity because of competition for nutrients, oxygen, or available space. These results suggest that 9 × 10⁷ CFU/mL was the optimal cell concentration for nicosulfuron degradation under the tested conditions.

The initial nicosulfuron concentration also influenced degradation performance (Fig. 2−B). After 72 h, free 2NN and IM-2NN showed their highest degradation rates, at 88.39% and 91.03%, respectively, at 50 mg/L nicosulfuron. At the lower concentration of 10 mg/L, degradation efficiency was reduced, possibly because the available substrate was insufficient to maintain high degradation activity. By contrast, exposure to 100 mg/L nicosulfuron decreased degradation efficiency, suggesting that excessive herbicide concentration inhibited bacterial growth or metabolic activity. Notably, IM-2NN maintained higher degradation activity than free 2NN under high nicosulfuron concentrations, indicating that the immobilization matrix protected the strains and helped preserve their degradation capacity under herbicide stress.

Temperature had a clear effect on nicosulfuron degradation by the mixed bacterial strains (Fig. 2−C). With a 5% inoculum and an initial nicosulfuron concentration of 50 mg/L, degradation efficiency first increased and then decreased as temperature rose. The highest degradation rates were observed at 35 ℃, reaching 86.24% for free 2NN and 90.06% for IM-2NN after 72 h. This pattern indicates that moderate warming enhanced bacterial activity and promoted nicosulfuron degradation. However, temperatures above 35 ℃ substantially reduced degradation efficiency, likely because excessive heat impaired bacterial growth or enzyme activity. Therefore, 35 ℃ was identified as the optimal temperature for nicosulfuron degradation by the mixed bacterial system.

The degradation efficiency of free 2NN and IM-2NN also varied with pH (Fig. 2−D). As pH increased, both bacterial growth and nicosulfuron degradation initially improved and then declined. The highest degradation rates were observed at pH 7.0, reaching 85.31% for free 2NN and 88.61% for IM-2NN within 72 h. Acidic and alkaline conditions reduced the growth and degradation capacity of the mixed bacterial strains, indicating that neutral pH was most favourable for nicosulfuron biodegradation. Under the same pH conditions, IM-2NN consistently showed stronger degradation activity than free 2NN. This advantage was likely due to the protective effects of the biochar-SA-PVA immobilization matrix, which helped the mixed bacterial strains tolerate unfavourable environmental conditions.

3.5 Cell viability assay of bacterial strains

The viability of the bacterial strains under nicosulfuron stress was evaluated by flow cytometry using an Annexin V-PE/7-AAD dual-staining kit (Fig. 3). In the scatterplots, Q1 represented mechanically damaged cells, Q2 represented late apoptotic cells, Q3 represented early apoptotic cells, and Q4 represented live cells. Flow cytometry showed that the proportion of apoptotic cells increased as the nicosulfuron concentration increased, indicating that high herbicide concentrations imposed clear stress on the bacterial cells. However, at 100 mg/L and 200 mg/L nicosulfuron, IM-2NN retained a higher proportion of live cells than free 2NN. These results indicate that immobilization improved the tolerance of the mixed bacterial strains to nicosulfuron stress and helped maintain cell viability under adverse conditions.

3.6 Reusability and storage stability of IM-2NN

The reusability of IM-2NN was evaluated over ten consecutive degradation cycles (Fig. S3−A). During the first six cycles, IM-2NN maintained high nicosulfuron degradation activity, with degradation rates of 89.21%, 88.03%, 85.32%, 82.76%, 79.82%, and 74.54%, respectively. The immobilized beads ruptured after the sixth cycle, after which degradation efficiency declined more rapidly. During cycles 7−10, the degradation rates decreased to 62.34%, 53.26%, 45.61%, and 42.22%, respectively. These results indicate that IM-2NN retained strong reusability for up to six cycles, but that structural damage to the immobilized beads limited its long-term repeated use.

The storage stability of free 2NN and IM-2NN was then compared (Fig. S3−B). After 90 d of storage, IM-2NN retained a nicosulfuron removal rate of 76.54%, whereas free 2NN showed only 8.11% removal activity. No detectable nicosulfuron degradation by free 2NN was observed after 90 d, indicating that the activity of the non-immobilized bacterial culture was largely lost during storage. By contrast, IM-2NN maintained degradation rates of 72.60% and 64.49% after 120 and 150 d of storage, respectively. Thus, immobilization greatly improved the storage stability of the mixed bacterial strains, likely because the biochar-SA-PVA carrier protected the cells from environmental stress and helped preserve their degradation capacity over time.

3.7 Degradation of nicosulfuron in soil by IM-2NN

The degradation of nicosulfuron in soil was promoted by both free 2NN and the immobilized microbial agent IM-2NN, but the effect was stronger in unsterilized soil and after immobilization (Fig. S4). In soil contaminated with 5 mg/kg nicosulfuron, free 2NN degraded 60.56% of nicosulfuron in sterilized soil and 78.37% in unsterilized soil after 28 d. Under the same conditions, IM-2NN further increased the degradation efficiency to 68.32% in sterilized soil and 89.68% in unsterilized soil. These results indicate that immobilization improved the soil degradation performance of the mixed bacterial strains. The higher degradation efficiency observed in unsterilized soil also suggests that native soil microorganisms may interact with the introduced 2NN strains and contribute to nicosulfuron removal. Thus, IM-2NN appears to enhance nicosulfuron degradation not only by maintaining the activity of the inoculated strains, but also by supporting cooperative interactions within the soil microbial environment.

3.8 Degradation pathways of nicosulfuron in soil

To clarify the possible degradation mechanism of nicosulfuron in soil, degradation products were analyzed by HPLC-MS/MS. Eleven compounds were detected in the samples (Fig. 4). Compound A showed a molecular ion peak at m/z 411, consistent with the parent compound nicosulfuron, and was therefore identified as nicosulfuron. On the basis of the detected intermediates, several possible transformation routes were proposed. Cleavage of the C−C bond in nicosulfuron may generate M1, identified as N-((4,6-dimethoxypyrimidin-2-yl)carbamoyl)pyridine-2-sulfonamide (m/z 338). At the same time, cleavage of the sulfonylurea bridge may produce M2, identified as 2-((4,6-dimethoxypyrimidin-2-yl)amino)nicotinic acid (m/z 274), and M5, identified as 2-isocyanato-4,6-dimethoxypyrimidine (m/z 182). Further transformation of M2 through C−N bond cleavage may generate M3, identified as 2-aminosulfonyl-N,N-dimethylnicotinamide (ASDM, m/z 230). M3 may then undergo S−N bond cleavage to form M4, identified as 3-(dimethylcarbamoyl)-pyridine-2-sulfinate (m/z 213). In another branch of the pathway, M5 may be converted to M6, identified as 2-amino-4,6-dimethoxypyrimidine (ADMP, m/z 156), followed by demethylation and deamidation to produce M7, identified as 4,6-dihydroxypyrimidine (m/z 112), and M8, identified as 4,6-dimethoxypyrimidine (m/z 141). In addition, C−S bond cleavage and removal of the sulfur dioxide group from nicosulfuron may generate M9, identified as 2-(1-(4,6-dimethoxypyrimidin-2-yl)ureido)-N,N-dimethylnicotinamide (m/z 347), which may subsequently undergo C−N bond cleavage to form M10, identified as 2-((4,6-dimethoxypyrimidin-2-yl)amino)-N,N-dimethylnicotinamide (m/z 304). Overall, the putative degradation pathway of nicosulfuron in soil involves sulfonylurea bridge cleavage and the cleavage of S−N, C−N, C−C, and C−S bonds (Fig. 4). These transformations suggest that IM-2NN-mediated degradation reduces nicosulfuron through sequential bond cleavage and conversion into smaller heterocyclic and sulfonamide-containing intermediates.

The effects of nicosulfuron and IM-2NN on maize rhizosphere soil metabolites were evaluated using principal component analysis (PCA), partial least-squares discriminant analysis (PLS−DA), and orthogonal partial least-squares discriminant analysis (OPLS−DA). The CK, YM, and IM groups were clearly separated, with no overlap among treatments, whereas replicates from the same treatment clustered closely together (Fig. S6). These results indicate good reproducibility within each group and clear metabolic differences among treatments. Differential metabolite analysis further showed that nicosulfuron and IM-2NN caused distinct changes in the maize rhizosphere soil metabolome. In the YM vs. CK comparison, 1,200 metabolites were screened, of which 165 were up-regulated and 157 were down-regulated in the YM group relative to the CK group (Fig. 5A−1). In the IM vs. CK comparison, 533 metabolites were up-regulated and 172 metabolites were down-regulated in the IM group relative to the CK group (Fig. 5A−2). In the YM vs. IM comparison, 103 metabolites were up-regulated and 513 metabolites were down-regulated in the YM group relative to the IM group (Fig. 5A−3). Overall, IM-2NN treatment caused broader metabolic reprogramming than nicosulfuron alone, and the large number of metabolites differing between YM and IM suggests that microbial remediation substantially reshaped the rhizosphere metabolic environment under nicosulfuron stress. To explore potential functional relationships among these metabolites, a metabolite-metabolite correlation network was constructed (Fig. 5−D). The network was mainly composed of organic acids and lipid compounds, indicating that carbon metabolism and lipid-related pathways may be important components of the soil metabolic response to IM-2NN treatment.

3.9 Colonization of bacterial strains in soil

The colonization capacity of the introduced bacterial strains was evaluated by conventional PCR using total DNA extracted from soil samples as the template (Fig. S6−A and S6−B). Specific DNA bands corresponding to the ssuD gene of strain 2N3 and the tenA gene of strain N80 were still detectable in soil samples after 28 d. These results indicate that the introduced degrading strains were able to persist in soil for at least 28 d. The detection of both marker genes also suggests that immobilization with the biochar-based carrier supported the survival of the functional strains in the soil environment. Sustained colonization by these degrading bacteria may contribute to continued nicosulfuron removal, thereby improving remediation efficiency and reducing the potential risk of herbicide residues in maize production systems.

3.10 Alleviation of nicosulfuron phytotoxicity in maize

The protective effect of the inoculated bacterial strains against nicosulfuron-induced oxidative stress was evaluated in maize seedlings. Without bacterial inoculation, seedlings exposed to nicosulfuron (S2) showed a significantly higher MDA content than untreated seedlings (S1), indicating that nicosulfuron induced lipid peroxidation in maize roots (Fig. S7−A). Bacterial inoculation markedly reduced MDA accumulation. The MDA levels in the S3 and S4 treatments decreased to 6.57 ± 0.53 mmol/g and 6.32 ± 0.45 mmol/g, respectively, compared with 12.8 ± 0.71 mmol/g in the S2 treatment. Brassinolide (S5) and the nicosulfuron safener cyprosulfamide (S6) also reduced MDA content, indicating that both chemical protection and microbial inoculation can alleviate nicosulfuron-induced membrane damage.

To determine whether the inoculated strains also affected antioxidant defense responses, the activities of POD, CAT, and SOD were measured in maize roots. POD and CAT showed similar response patterns (Fig. S7−B and −C). Their activities were significantly higher in the S2 treatment than in the S1 treatment, consistent with the activation of antioxidant responses under nicosulfuron stress. After bacterial inoculation, POD and CAT activities decreased in the S3 and S4 treatments, suggesting that the oxidative burden was reduced. By contrast, SOD activity was significantly inhibited by nicosulfuron in the S2 treatment compared with that in S1, whereas bacterial inoculation markedly restored SOD activity in maize roots (Fig. S7−D). Brassinolide and cyprosulfamide also alleviated nicosulfuron injury, with cyprosulfamide showing the strongest protective effect among the tested treatments. Together, these results indicate that IM-2NN mitigated nicosulfuron-induced phytotoxicity in maize by reducing lipid peroxidation and helping rebalance antioxidant enzyme activities.

4 DISCUSSION

In this study, both the free mixed bacterial culture 2NN and the immobilized microbial agent IM-2NN significantly enhanced nicosulfuron degradation in maize field soil. However, IM-2NN consistently showed stronger degradation performance than free 2NN, indicating that immobilization improved the effectiveness and stability of the degrading strains under soil conditions (Fig. S4). This effect is likely associated with the slow and continuous release of bacteria from the immobilized matrix, which can help maintain microbial activity and prolong colonization in soil (Zhang et al., 2020a; Qi et al., 2020). Consistent with this explanation, the marker genes of both introduced strains remained detectable after 28 d, suggesting that IM-2NN supported the persistence of the degrading bacteria in the soil environment (Fig. S6). Biochar may also contribute to this effect by providing a porous habitat for bacterial attachment, improving nutrient and oxygen exchange, and buffering microorganisms against environmental stress. Previous studies have shown that biochar application can activate indigenous microorganisms, enhance microbial community diversity, improve soil fertility, and accelerate the recovery of soil physicochemical properties (Rafiq et al., 2019). In the present study, nicosulfuron degradation was higher in unsterilized soil than in sterilized soil, suggesting that native soil microorganisms may cooperate with the introduced 2NN strains to promote herbicide removal. Therefore, the remediation effect of IM-2NN likely reflects both direct nicosulfuron degradation by the introduced strains and indirect improvement of the soil microecosystem.

Metabolomic analysis further showed that nicosulfuron contamination caused broad changes in the metabolic profile of maize rhizosphere soil. Compared with the untreated control (CK), nicosulfuron-treated soil (YM) showed clear changes in metabolites associated with microbial nutrient acquisition, energy metabolism, membrane structure, oxidative stress, secondary metabolism, and cell-to-cell communication (Fig. 5). The decrease in the iron carrier nigrifactin may indicate disruption of microbial iron uptake, whereas changes in orthophosphate suggest disruption of energy metabolism (Awodi et al., 2017). In addition, reductions in choline phosphate, a membrane phospholipid precursor, and uric acid, a purine metabolite, may reflect impaired membrane-related metabolism and cellular stress. By contrast, increases in biotin and choline may represent adaptive metabolic responses that support coenzyme metabolism and methyl-group transfer under nicosulfuron pressure. These results suggest that nicosulfuron residues not only persist as chemical pollutants but also reshape the functional metabolic state of the rhizosphere microbial community. Stress-related secondary metabolism was also strongly affected by nicosulfuron. The accumulation of antioxidant metabolites, including trans-resveratrol and dihydropinosylvin, suggests activation of oxidative stress responses in contaminated soil, whereas increases in antimicrobial alkaloids, such as isoquinoline and elymoclavine, may reflect intensified microbial competition or defence-related metabolism under pollutant stress (Jiang et al., 2025; Zhao et al., 2025). Lipid-related changes further support this interpretation. The increase in linoleic acid may be associated with lipid remodelling or membrane repair, whereas the decrease in arachidic acid suggests that saturated fatty acid metabolism was impaired. At the same time, changes in allantoate, phenylglyoxal, 2-keto valeric acid, and azelaic acid indicate that nitrogen turnover and carbon metabolism were also affected (Zhao et al., 2025). The opposite trends observed for N-(3-oxododecanoyl)-homoserine lactone and N-(7Z-tetradecenoyl)-homoserine lactone further suggest that nicosulfuron may disturb quorum-sensing-related microbial communication. Together, these metabolic features indicate that nicosulfuron contamination imposed a combined burden on microbial metabolism, membrane integrity, redox balance, and community signalling.

Following IM-2NN application, the soil metabolome showed a distinct pattern that was consistent with microbial activation and partial metabolic recovery. Compared with CK, the IM group exhibited substantial changes in metabolites related to coenzyme metabolism, energy conversion, antioxidant defence, lipid remodelling, and carbon and nitrogen metabolism. The marked increase in biotin suggests that the microbial agent may have promoted coenzyme-associated metabolism, whereas changes in orthophosphate and choline phosphate indicate modulation of energy metabolism and membrane-related processes (Pan et al., 2025). In the IM group, the accumulation of antioxidant compounds, including trans-resveratrol and dihydropinosylvin, may reflect enhanced redox regulation mediated by the microbial agent. Similarly, the increase in isoquinoline suggests that IM-2NN may have influenced antimicrobial defence or competitive interactions within the soil microbial community (Li et al., 2025). Changes in linoleic acid and palmitoleic acid further suggest that membrane lipid remodelling contributed to microbial adaptation after remediation. In addition, altered levels of allantoate, D-asparagine, 2-keto valeric acid, and azelaic acid indicate that the microbial agent affected nitrogen metabolism and carbon allocation. These results suggest that IM-2NN did not simply return the soil metabolome to the untreated state; rather, it actively reshaped rhizosphere metabolism under nicosulfuron stress.

A direct comparison between contaminated soil (YM) and microbially remediated soil (IM) provided further insight into the biochemical basis of remediation. Several putative xenobiotic-like and stress-associated metabolites, including anthracene, 2-naphthaldehyde, S-nicotine, cotinine, N-(4-chlorophenethyl)-1-adamantanecarboxamide, xanthone, and purpurin, were lower in the IM group than in the YM group. These reductions are consistent with enhanced degradation or reduced accumulation of toxic compounds in the rhizosphere, although the identities and origins of these metabolites should be further verified. The decrease in N-(4-chlorophenethyl)-1-adamantanecarboxamide may indicate decomposition of sulfonamide-related compounds or other pesticide-derived metabolites (Hu et al., 2022). Meanwhile, the increase in R-lactate together with the decrease in pyruvic acid suggests that IM-2NN altered pyruvate metabolism and may have increased glycolytic flux, thereby improving energy supply for microbial growth and pollutant transformation (Dong et al., 2024). The accumulation of D-proline may enhance microbial tolerance to environmental stress, while changes in taurine and cystathionine ketimine suggest remodelling of sulfur-related metabolism. These responses indicate that microbial remediation involves both pollutant transformation and broader reorganization of microbial metabolic activity. The metabolite changes also provide a possible explanation for the reduced maize phytotoxicity observed after IM-2NN treatment. The decrease in S-abscisic acid may indicate reduced stress signalling in the rhizosphere, whereas changes in 4-vinylguaiacol sulfate, E-p-coumaric acid, and biotin suggest altered defence-related metabolism and plant−microbe interactions (Hua et al., 2023). In addition, the increase in allantoate may contribute to nitrogen recycling or reactive oxygen species scavenging, whereas changes in hexanal are consistent with altered lipid peroxidation. Microbial functional markers also changed after remediation. For example, the down-regulation of N-(3-hydroxy-pentanoyl)-homoserine lactone suggests altered quorum sensing (Chen et al., 2024), while changes in 2,3-di-O-acyl trehalose may be related to biofilm formation and bacterial colonization. These metabolic shifts are consistent with the physiological results showing that IM-2NN reduced malondialdehyde accumulation and helped rebalance antioxidant enzyme activities in maize roots. Thus, IM-2NN may alleviate nicosulfuron injury through two linked mechanisms: reducing herbicide pressure in the rhizosphere and improving the metabolic environment that supports root stress resistance. This interpretation is also consistent with previous evidence that plant-associated microorganisms can promote stress adaptation, nutrient acquisition, and rhizosphere recovery through coordinated changes in microbial metabolism and plant−microbe signalling (Compant et al., 2025).

Pathway enrichment analysis highlighted amino acid metabolism and energy conversion as important components of IM-2NN-mediated remediation. In the IM group, tryptophan metabolism was activated relative to CK, with changes in metabolites associated with indole-3-acetic acid biosynthesis (Fig. 5C−2; Agus et al., 2018). Because tryptophan-derived metabolites are closely linked to microbial auxin production and root development, activation of this pathway may contribute to maize root recovery after herbicide stress. Changes in D-amino acid metabolism, including D-proline accumulation, may further improve microbial tolerance to adverse conditions (Wang et al., 2025). Histidine metabolism may also participate in detoxification because histidine and related metabolites can contribute to metal binding and stress adaptation. In addition, enhanced pyruvate metabolism, reflected by increased lactate accumulation, may provide energy and reducing power for microbial growth, biofilm formation, and pollutant transformation. These pathway changes suggest that IM-2NN supports remediation by reinforcing microbial stress tolerance, energy supply, and plant-growth-promoting metabolic activity. By contrast, the YM group showed pathway-level features consistent with membrane injury and energy imbalance. Nicosulfuron contamination inhibited unsaturated fatty acid biosynthesis, as reflected by reduced arachidic acid, while increased linoleic acid metabolism and hexanal accumulation suggested lipid peroxidation and oxidative damage to membrane lipids (Du et al., 2025). The decrease in choline phosphate further supports disruption of membrane-associated metabolism. These changes indicate that nicosulfuron may impair microbial cell structure by interfering with lipid metabolism and increasing oxidative stress. The YM vs IM comparison showed that several pathways shifted in opposite directions after microbial remediation. In particular, IM-2NN appeared to reactivate tryptophan metabolism, restore osmoprotective D-amino acid metabolism, enhance histidine-related detoxification, and redirect pyruvate metabolism towards lactate production. These pathway inversions suggest that remediation involves targeted regulation of amino acid metabolism and energy-conversion networks, rather than only passive pollutant removal (Cluntun et al., 2021).

Overall, this study shows that IM-2NN improves nicosulfuron degradation, supports the persistence of functional degrading bacteria in soil, reshapes the rhizosphere metabolome, and alleviates herbicide-induced oxidative damage in maize. Compared with free bacterial cells, immobilization on a biochar-SA-PVA carrier appears to improve bacterial survival, environmental tolerance, degradation stability, and storage performance. The metabolomic results further suggest that IM-2NN-mediated remediation involves coordinated reconstruction of microbial metabolism, redox balance, membrane homeostasis, amino acid metabolism, energy conversion, and plant−microbe interactions. These findings support the potential use of IM-2NN as a practical microbial agent for remediating nicosulfuron-contaminated maize fields. Future studies should verify key metabolite identities using authentic standards, examine changes in microbial community composition and functional genes, and evaluate the long-term performance of IM-2NN under different field conditions.

5 CONCLUSIONS

In this study, K. jilinsis 2N3 and S. marcescens N80 were combined at a 1:1 ratio and immobilized using a composite carrier prepared from biochar, PVA, and SA. The biochar was derived from discarded fungal culture medium, providing a potential route for the reuse of agricultural waste. Compared with the free mixed bacterial culture 2NN, the immobilized microbial agent IM-2NN showed higher nicosulfuron degradation efficiency, stronger storage stability and reusability, and better adaptability to environmental stress. IM-2NN also exhibited stable colonization in soil and promoted the functional recovery of contaminated rhizosphere soil by regulating soil metabolic networks, thereby creating a more favourable microenvironment for maize roots. These results show that IM-2NN can accelerate nicosulfuron degradation in maize field soil and significantly alleviate herbicide-induced phytotoxicity in maize. By improving herbicide residue removal, supporting soil metabolic recovery, and reducing oxidative damage in maize plants, IM-2NN represents a sustainable and efficient strategy for managing nicosulfuron-contaminated maize fields. Future studies should validate the long-term field performance of IM-2NN and evaluate its integration with maize variety selection and agronomic management practices to optimize herbicide safety and crop productivity.

References

[1] Agus A , Planchais J , Sokol H . Gut microbiota regulation of tryptophan metabolism in health and disease. Cell Host & Microbe, 2018, 23(6): 716–724

[2] Alimohammadi A , Moosavy M H , Amin Doustvandi M . et al. Sodium metabisulfite as a cytotoxic food additive induces apoptosis in HFFF2 cells. Food Chem, 2021, 358: 129910

[3] Awodi U R , Ronan J L , Masschelein J . et al. Thioester reduction and aldehyde transamination are universal steps in actinobacterial polyketide alkaloid biosynthesis. Chemical Science, 2017, 8(1): 411–415

[4] Benzi M, Robotti E, Gianotti V. 2011. HPLC-DAD-MSn to investigate the photodegradation pathway of nicosulfuron in aqueous solution. Anal Bioanal Chem, 399(4): 1705−1714.

[5] Bhatt P, Rene E R, Huang Y, et al. 2022. Indigenous bacterial consortium-mediated cypermethrin degradation in the presence of organic amendments and Zea mays plants. Environ Res, 212(Pt A): 113137.

[6] Carles L, Joly M, Bonnemoy F, et al. 2017. Identification of sulfonylurea biodegradation pathways enabled by a novel nicosulfuron-transforming strain Pseudomonas fluorescens SG-1: Toxicity assessment and effect of formulation. J Hazard Mater, 324(Pt B): 184−193.

[7] Chen L , Li Y , Zhang P . et al. Quorum sensing in anaerobic wastewater treatment: Potential microbiological mechanisms and enhancement strategies. Chemical Engineering Journal, 2024, 499: 156150

[8] Chen W , Zhang H , Zhang M . et al. Removal of PAHs at high concentrations in a soil washing solution containing TX-100 via simultaneous sorption and biodegradation processes by immobilized degrading bacteria in PVA-SA hydrogel beads. J Hazard Mater, 2021, 410: 124533

[9] Chen X , Lin H , Dong Y . et al. Mechanisms underlying enhanced bioremediation of sulfamethoxazole and zinc(II) by Bacillus sp. SDB4 immobilized on biochar. J. Clean. Prod, 2022, 370: 133483

[10] Cluntun A A, Badolia R, Lettlova S, et al. 2021. The pyruvate-lactate axis modulates cardiac hypertrophy and heart failure. Cell Metabolism, 33(3): 629−648. e10.

[11] Compant S, Cassan F, Kostić T, et al. 2025. Harnessing the plant microbiome for sustainable crop production. Nature Reviews. Microbiology, 23(1): 9−23.

[12] Cueff S , Alletto L , Dumény V . et al. Adsorption and degradation of the herbicide nicosulfuron in a stagnic Luvisol and Vermic Umbrisol cultivated under conventional or conservation agriculture. Environ Sci Pollut Res Int, 2021, 28(13): 15934–15946

[13] Dong M, Zhou H, Wang J, et al. 2024. Responses of soil microbial metabolism, function and soil quality to long-term addition of organic materials with different carbon sources. Biochar, 6(1): 80.

[14] Du S , Li R , Liu Y . et al. High ω6/ω3 polyunsaturated fatty acid (PUFA) ratio impairs intestinal mucosal barrier function via ROS/TLR4/NF-κB-mediated aberrant expression of tight junction proteins. Food Bioscience, 2025, 66: 106225

[15] Gao N , Zhang J P , Pan Z Q . et al. Biodegradation of atrazine by mixed bacteria of Klebsiella variicola strain FH-1 and Arthrobacter sp. NJ-1. Bull Environ Contam Toxicol, 2020, 105: 481–489

[16] Generozova I P , Vasilyev S V , Butsanets P A . et al. Combined action of melatonin and water deficiency on growth and MDA content of hypocotyls and roots of lupine and respiration of mitochondria isolated from these organs. Russ. J. Plant Physiol, 2022, 69: 93

[17] Giannakopoulos G , Dittgen J , Schulte W . et al. Safening activity and metabolism of the safener cyprosulfamide in maize and wheat. Pest Manag Sci, 2020, 76: 3413–3422

[18] Hou J , Mao D , Zhang Y . et al. Long-term spatiotemporal variation of antimicrobial resistance genes within the Serratia marcescens population and transmission of S. marcescens revealed by public whole-genome datasets. J Hazard Mater, 2022, 423: 127220

[19] Hu J, Li X, Liu F, et al. 2022. Comparison of chemical and biological degradation of sulfonamides: Solving the mystery of sulfonamide transformation. J Hazard Mater, 424(Pt D): 127661.

[20] Hua C, Zhang Z, Guo Z, et al. 2023. 4-Vinyl Guaiacol exists widely in rhizosphere soil of Lanzhou lily (Lilium davidii var. unicolor) and has a strong negative effect on lily planting. Scientia Horticulturae, 315: 111959.

[21] Jiang Y, Li Z, Li S. 2025. Superoxide-mediated O2 activation drives radical cyclization in ergot alkaloid biosynthesis. Engineering Microbiology, 5(2): 100207.

[22] Jiang Z W , Zheng Z J , Wu J N . et al. Synthesis, characterization and performance of microorganism-embedded biocomposites of LDH-modified PVA/SA hydrogel beads for enhanced biological nitrogen removal process. Process Biochem, 2022, 121: 542–552

[23] Kniss A R . Long-term trends in the intensity and relative toxicity of herbicide use. Nature Communications, 2017, 8: 14865

[24] Landreau M , Byson S J , You H . et al. Effective nitrogen removal from ammonium-depleted wastewater by partial nitritation and anammox immobilized in granular and thin layer gel carriers. Water Res, 2020, 183: 116078

[25] Lei Q , Zhong J , Chen S F . et al. Microbial degradation as a powerful weapon in the removal of sulfonylurea herbicides. Environ Res, 2023, 235: 116570

[26] Li H , Jiang Q , Li R Z . et al. Passivation of lead and cerium in soil facilitated by biochar-supported phosphate-doped ferrihydrite: Mechanisms and microbial community evolution. J. Hazard. Mater, 2022, 436: 129090

[27] Li M , Yin H , Zhu M . et al. Co-metabolic and biochar-promoted biodegradation of mixed PAHs by highly efficient microbial consortium QY1. J Environ Sci, 2021, 107: 65–76

[28] Li Z , Wang C , Shah F . et al. Improvement of oil quality with enhanced fatty acid profiling in rapeseed (Brassica napus L. ) seed via varietal screening and lipidomics (UPLC-MS) analysis. LWT, 2025, 216: 117288

[29] Liu X M , Zhao D , Xu X . et al. Role of ZmGST gene family involved in nicosulfuron stress tolerance revealed by genomic and transcriptomic analyses. Agronomy, 2022, 12: 2598

[30] Lonappan L , Rouissi T , Kaur Brar S . et al. An insight into the adsorption of diclofenac on different biochars: Mechanisms, surface chemistry, and thermodynamics. Bioresour Technol, 2018, 249: 386–394

[31] Lu B , Meng R , Wang Y . et al. Distinctive physiological and molecular responses of foxtail millet and maize to nicosulfuron. Frontiers in Plant Science, 2024, 14: 1308584

[32] Lu X H , Kang Z H , Tao B . et al. Degradation of nicosulfuron by Bacillus subtilis YB1 and Aspergillus niger YF1. Prikl Biokhim Mikrobiol, 2012, 48: 510–515

[33] Ma Q , Wu E , Wang H . et al. Exogenous 24-epibrassinolide boosts plant growth under alkaline stress from physiological and transcriptomic perspectives: The case of broommaize millet (Panicum miliaceum L. ). Ecotoxicol Environ Saf, 2022, 248: 114298

[34] Nie Z , Yan B , Xu Y . et al. Characterization of pyridine biodegradation by two Enterobacter sp. strains immobilized on Solidago canadensis L. stem derived biochar. J Hazard Mater, 2021, 414: 125577

[35] Pan X , Jian Y , An N . et al. Innovations in phosphate phosphors: Bridging structure-property gaps for enhanced luminescence. Coordination Chemistry Reviews, 2025, 528: 216407

[36] Pourbabaee A , Soleymani A , Torabi E . et al. Degradation and detoxification of nicosulfuron by a Pseudomonas strain isolated from a contaminated cornfield soil. Soil and Sediment Contamination, 2018, 27(8): 756–772

[37] Qi X , Gou J L , Chen X M . et al. Application of mixed bacteria-loaded biochar to enhance uranium and cadmium immobilization in a co-contaminated soil. J Hazard Mater, 2020, 401: 123823

[38] Rafiq M K , Bai Y , Aziz R . et al. Biochar amendment improves alpine meadows growth and soil health in Tibetan plateau over a three year period. Sci Total Environ, 2019, 717: 135296

[39] Silva F B , Costa A C , Müller C . et al. Dipteryx alata, a tree native to the Brazilian Cerrado, is sensitive to the herbicide nicosulfuron. Ecotoxicology, 2020, 29: 217–225

[40] Silva H S A , Romeiro S D R , Macagnan D . et al. Rhizobacterial induction of systemic resistance in tomato plants: non-specific protection and increase in enzyme activities. Biol. Control, 2004, 29: 288–295

[41] Singh R P, Jha P, Jha P N. 2015. The plant-growth-promoting bacterium Klebsiella sp. SBP-8 confers induced systemic tolerance in wheat (Triticum aestivum) under salt stress. J Plant Physiol, 184: 57−67.

[42] Sun T , Miao J B , Saleem M . et al. Bacterial compatibility and immobilization with biochar improved tebuconazole degradation, soil microbiome composition and functioning. J. Hazard. Mater, 2020, 398: 122941

[43] Tao Y , Han S Y , Zhang Q . et al. Application of biochar with functional microorganisms for enhanced atrazine removal and phosphorus utilization. J. Clean. Prod, 2020, 257: 120535

[44] Wang H Q , Wu X J , Xu J . et al. Proline mitigates antibiotic resistance evolution induced by ciprofloxacin at environmental concentrations. Journal of Hazardous Materials, 2025, 489: 137561

[45] Wang L, Zhang X, Li Y. 2016. Degradation of nicosulfuron by a novel isolated bacterial strain Klebsiella sp. Y1: condition optimization, kinetics and degradation pathway. Water Sci Technol, 73: 2896−2903.

[46] Wang Y , Du L , Liu H . et al. Halosulfuron methyl did not have a significant effect on diversity and community of sugarcane rhizosphere microflora. J Hazard Mater, 2020, 399: 123040

[47] Wen Y , Xu X , Wang B . et al. PAHs biodegradation in soil washing effluent by native mixed bacteria embedded in polyvinyl alcohol-sodium alginate-nano alumina gel beads. J Environ Manage, 2021, 297: 113415

[48] Wu Z X , Xu N W , Yang M . et al. Responses of photosynthesis, antioxidant enzymes, and related gene expression to nicosulfuron stress in sweet maize (Zea mays L. ). Environ Sci Pollut Res Int, 2022, 29: 37248–37265

[49] Xiang X , Yi X , Zheng W . et al. Enhanced biodegradation of thiamethoxam with a novel polyvinyl alcohol (PVA)/sodium alginate (SA)/biochar immobilized Chryseobacterium sp H5. J Hazard Mater, 2022, 443: 130247

[50] Xiao T , Yang L , Zhang D . et al. Early detection of nicosulfuron toxicity and physiological prediction in maize using multi-branch deep learning models and hyperspectral imaging. Journal of Hazardous Materials, 2024, 474: 134723

[51] Yang F , Jian H , Wang C . et al. Effects of biochar on biodegradation of sulfamethoxazole and chloramphenicol by Pseudomonas stutzeri and Shewanella putrefaciens: Microbial growth, fatty acids, and the expression quantity of genes. J Hazard Mater, 2021, 406: 124311

[52] Yu Y Y , Yin H , Huang W T . et al. Cellular changes of microbial consortium GY1 during decabromodiphenyl ether (BDE-209) biodegradation and identification of strains responsible for BDE-209 degradation in GY1. Chemosphere, 2020, 249: 126205

[53] Yuan X F , Ma L , Wen B . et al. Enhancing anaerobic digestion of cotton stalk by pretreatment with a microbial consortium (MC1). Bioresource Technology, 2016, 207: 293–301

[54] Zhai Q H , Chen X T , Zhang M . et al. Immobilization of Klebsiella jilinsis strain 2N3 by corn straw biochar enhanced the degradation of nicosulfuron and restores the soil microbiome function and composition. Appl. Soil Ecol, 2023, 189: 104917

[55] Zhang B Y , Ni Y X , Liu J W . et al. Bead-immobilized Pseudomonas stutzeri Y2 prolongs functions to degrade s-triazine herbicides in industrial wastewater and maize fields. Sci. Total Environ, 2020a, 731: 139183

[56] Zhang C , Hao Q , Zhang Z . et al. Whole genome sequencing and analysis of chlorimuron-ethyl degrading bacteria Klebsiella pneumoniae 2N3. Int J Mol Sci, 2019a, 20: 3053

[57] Zhang C , Li J , Wu X . et al. Rapid degradation of dimethomorph in polluted water and soil by Bacillus cereus WL08 immobilized on bamboo charcoal−sodium alginate. J Hazard Mater, 2020b, 398: 122806

[58] Zhang C F , Chen L , Si H L . et al. Study on the characteristics and mechanisms of nicosulfuron biodegradation by Bacillus velezensis CF57. J Basic Microbiol, 2020c, 60: 649–658

[59] Zhang H , Mu W , Hou Z . et al. Biodegradation of nicosulfuron by the bacterium Serratia marcescens N80. J Environ Sci Health B, 2012, 47: 153–160

[60] Zhang H , Zhang X , Mu W . et al. Biodegradation of chlorimuron-ethyl by the bacterium Klebsiella jilinsis 2N3. J Environ Sci Health B, 2010, 45: 501–507

[61] Zhang J P, Liang S, Wang X H, et al. 2019b. Biodegradation of atrazine by the novel Klebsiella variicola strain FH-1. Biomed Res Int, 4756579.

[62] Zhang M , Zhang C , Zhang S . et al. Klebsiella jilinsis 2N3 promotes maize growth and induces resistance to northern maize leaf blight. Biol. Control, 2021, 156: 104554

[63] Zhang Z , Yang D , Si H . et al. Biotransformation of the herbicide nicosulfuron residues in soil and seven sulfonylurea herbicides by Bacillus subtilis YB1: A climate chamber study. Environ Pollut, 2020d, 263: 114492

[64] Zhang Z , Yang D C , Wang J Y . et al. Studies on the interactions between nicosulfuron and degradation enzymes. Process Biochem, 2020e, 91: 99–103

[65] Zhao H, Zhu J, Liu S, et al. 2018 Kinetics study of nicosulfuron degradation by a Pseudomonas nitroreducens strain NSA02. Biodegradation, 29(3): 271−283.

[66] Zhao W , Wang C , Xu L . et al. Biodegradation of nicosulfuron by a novel Alcaligenes faecalis strain ZWS11. J Environ Sci, 2015, 35: 151–162

[67] Zhao W B , An J X , Jin Y R . et al. Indoloquinoline alkaloid neocryptolepine derivative inhibits Botrytis cinerea by targeting thiamine thiazole synthase. Science Advances, 2025, 11(11): eadq5329

[68] Zhong J F , Wu S Y , Chen W J . et al. Current insights into the microbial degradation of nicosulfuron: Strains, metabolic pathways, and molecular mechanisms. Chemosphere, 2023, 326: 138390

[69] Zhu C Y , Zhang C , Zhang M . et al. Degradation characteristics and soil remediation of thifensulfuron-methyl by immobilized Serratia marcecens N80 beads. Environ. Technol. Innov, 2021, 24: 102059

RIGHTS & PERMISSIONS

Mitigating nicosulfuron injury and remediating herbicide residues in maize fields: degradation by the immobilized microbial agent IM-2NN and soil metabolomic analysis