Functional Characterization and Catalytic Mechanism of an Amidase Involved in Acetaminophen Degradation from Acinetobacter sp.

Abstract

The accumulation of acetaminophen (APAP) in agricultural soils and water systems poses risks to ecosystems and public health. This study characterizes Acinetobacter sp. DL27, an APAP-degrading strain with broad temperature and pH adaptability, demonstrating its bioremediation potential in soil and wastewater. Through high-resolution mass spectrometry, we identified three novel metabolic intermediates, thereby refining the bacterial APAP degradation pathway. Multiomics analysis elucidated metabolic coordination and stress-tolerance mechanisms, leading to the identification of a novel amidase (AdA). Recombinant AdA exhibited activity over 10–60 °C and pH 4.0–10.0, with a K _m of 8.96 ± 1.03 μM and a k _cat/K _m of 9.04 μM^–1s^–1, indicating higher catalytic efficiency than previously reported APAP amidases. Molecular dynamics simulations and site-directed mutagenesis confirmed that a Ser161–Ser185–Lys82 triad constitutes the catalytic center driving amide cleavage. These findings provide mechanistic insights into bacterial APAP biodegradation and highlight the practical application potential of strain DL27 and AdA.

1. Introduction

The manufacture of drug substances and the continuous release of thousands of pharmaceutical and personal care products (PPCPs) have posed a significant impact on the environment and human health over the past few decades. Acetaminophen (APAP), a representative pharmaceutical and personal care product, remains one of the most widely prescribed antipyretic and analgesic drugs worldwide. Notably, APAP is listed in the WHO Model List of Essential Medicines. With global population growth, rapid urbanization, and improved healthcare facilities, the production and consumption of APAP have increased substantially, making it one of the ten most widely consumed medicines globally. , In 2023, the global demand for APAP was almost 145,000 tons, which is estimated to reach 235,000 tons by 2034. Furthermore, increased consumption during the COVID-19 pandemic led to the widespread use of APAP for the treatment of fever and myalgia, resulting in a significant surge in its production and use. Although APAP provides significant public health benefits, its potential environmental risks have increasingly attracted attention. Due to its widespread use and incomplete metabolism in the human body, large amounts of APAP are released into the environment through human excretion such as feces and urine, emissions from hospitals and pharmaceutical factories, and improper disposal of excess drugs. APAP exhibits mobility and persistence in aquatic systems due to its high aqueous solubility. Consequently, it has been widely detected in sediments, wastewater, , surface water, groundwater, and drinking water systems across different environments at concentrations ranging from nanograms per liter to milligrams per liter. Strikingly, a study investigated the concentrations of acetaminophen in the influent of wastewater treatment plants across 25 countries and regions, with values ranging from ND to 66.4 mg/L. Furthermore, APAP has been detected in agricultural soils. Correspondingly, various crop species, such as mustard, lettuce, cucumber, carrot, spinach, and cabbage, have been shown to take up and accumulate this compound within their edible tissues. APAP residues have also been identified in fruits. Consequently, these findings raise concerns regarding the potential health risks of APAP exposure via the food chain.

As an active ingredient, APAP poses risks to aquatic organisms and ecosystems, even at low concentrations. For example, APAP induces the generation of reactive oxygen species (ROS) in the fat body of fruit flies, leading to cell death in a dose-dependent manner, and it affects the reproductive and endocrine systems of Daphnia magna. The adverse effects of APAP in fish mainly include reproductive and endocrine impairment, hepatotoxicity, oxidative stress, neurotoxicity, and developmental impairment. − Beyond aquatic systems, the accumulation of APAP in agricultural soils due to wastewater irrigation poses a threat to the agricultural ecosystems. The growth of crops, such as lettuce, maize, and wheat, has been shown to be inhibited. , It also affects the morphology, physiological activity, and nutritional quality of spinach. The APAP exposure can alter the abundance of beneficial soil microorganisms that support crop growth. In addition, APAP may also affect human health through bioaccumulation and transfer within the food chain. Excessive exposure to APAP can lead to serious health effects, including liver damage, fetal developmental defects, abnormalities of the genitourinary system, myocardial infarction, and neurotoxicity. ,,

The ubiquitous occurrence of APAP has led to its recognition as an emerging contaminant of global concern. Considerable efforts have been devoted to developing effective remediation strategies, among which bioremediation has emerged as a promising approach owing to its cost-effectiveness, operational simplicity, and environmental compatibility. , A variety of APAP-degrading bacterial genera have been reported, including Delftia, Enterobacter, Pseudomonas, Bacillus, Klebsiella, Paracoccus, Flavobacterium, Dokdonella, Methylophilus, and Shinella. ,− The degradation pathway generally begins with the amidase-mediated hydrolysis of APAP to produce p-aminophenol (PAP), followed by its conversion into hydroquinone (HQ), 1,2,4-benzenetriol (BT), and catechol, which subsequently enter the TCA cycle for complete mineralization. , In recent years, significant progress has been made in characterizing microbial-derived APAP-degrading amidases. Early studies isolated an arylamidase AAA from soil bacteria, which exhibited optimal activity under alkaline conditions (pH 10.0) and a K _m of 0.32 mM for APAP. Subsequently, a broad-spectrum arylamidase AmpA was cloned from Paracoccus sp. FLN-7 and demonstrated efficient hydrolysis of various amide compounds, including APAP, with a notably higher substrate affinity (K _m = 30.4 μM, k _cat = 43.2 s^–1) than that of AAA. Similarly, the arylamidase Mah from Ochrobactrum sp. PP-2 was found to mediate APAP degradation, exhibiting a K _m of 13.1 μM. More recently, two AS-family amidases, ApaH1 and ApaH2, were identified in Pseudomonas taiwanensis AP-1. Among these, ApaH1 plays a primary role in APAP degradation, characterized by a K _m of 8.38 μM and a k _cat of 64.23 s^–1. The identification of these diverse enzymes and the characterization of their catalytic kinetics provide valuable insights into the microbial metabolic pathways of APAP, establishing a foundation for potential environmental bioremediation applications. Despite these advances, current studies have largely focused on strain isolation, degradation performance, and preliminary metabolite identification. The key functional genes and regulatory networks governing APAP biodegradation remain poorly characterized. Moreover, most investigations have been conducted under controlled laboratory conditions, whereas the performance of APAP-degrading microorganisms in complex environmental matrices remains insufficiently explored.

In this study, a highly efficient APAP-degrading strain, Acinetobacter sp. DL27, was successfully isolated from the activated sludge. Its degradation characteristics and bioremediation potential in agricultural soil and wastewater were evaluated. The APAP degradation pathway by strain DL27 was identified using UHPLC-Q-Orbitrap HRMS. In addition, multiomics analyses elucidated the molecular mechanisms of APAP degradation, and the key enzyme amidase (AdA) was verified through heterologous expression. Collectively, these findings offer valuable bioresources for mitigating APAP pollution and establish a theoretical foundation for green bioremediation.

2. Materials and Methods

2.1. Chemicals and Media

APAP (purity ≥ 98%) and other analytical reagents were obtained from Aladdin Biochemical Technology Co., Ltd. (Shanghai, China). HPLC-grade methanol was purchased from Kermel Chemical Reagent Co., Ltd. (Tianjin, China). The components of the Luria–Bertani (LB) medium used for bacterial growth and the mineral salt medium (MSM) used for screening APAP-degrading bacteria were provided in Text S1.

2.2. Isolation and Identification of APAP-Degrading Bacteria

The activated sludge obtained from a municipal wastewater treatment plant was enriched and acclimated for 120 d in an MSM with APAP. The microbial consortium obtained in this study, which could effectively degrade APAP, was subsequently subjected to metagenomic sequencing by GENEWIZ Biotechnology (Suzhou, China). The metagenomic sequencing data have been submitted to the NCBI database under accession number PRJNA1434051. Pure bacterial strains exhibiting robust APAP degradation capacities were subsequently isolated. The morphological analysis and 16S rRNA gene sequencing were used for the characterization of these isolates, with detailed procedures provided in Text S2.

2.3. Biodegradation Characteristics of APAP

Single-factor experiments were conducted to assess the effects of environmental variables (temperature, pH, inoculum size, and agitation speed) on the APAP degradation. The degradation assays were performed in MSM with 150 mg/L of APAP and incubated for 30 min. APAP concentrations were determined by HPLC. The experimental design and HPLC conditions are provided in Texts S3 and S4. The effect of the initial substrate concentration on APAP degradation was evaluated by cultivating strain DL27 in MSM containing 15, 45, 75, 150, 225, 300, and 450 mg/L APAP. Cultures were sampled every 10 min for APAP quantification. Experiments were conducted in triplicate with uninoculated control.

2.4. Bioremediation of APAP by Strain DL27 in Agricultural Soil and Wastewater

To assess the bioremediation potential of strain DL27, we tested its ability to degrade APAP in agricultural soil and wastewater. Experiments were conducted using sterilized and nonsterilized groups. The strain was inoculated at a 10% (v/v) ratio into soil and wastewater containing APAP at initial concentrations of 15, 45, and 75 mg/kg and 15, 45, and 75 mg/L, respectively. Uninoculated controls were maintained in parallel with all treatments. APAP concentrations were quantified by HPLC. The experimental procedures are provided in Text S5.

2.5. Localization of Degradation Enzymes

To determine whether the degrading enzymes are exclusively expressed under APAP-induced conditions, the DL27 strain was cultured to the logarithmic phase with and without APAP, maintaining an identical OD₆₀₀ value across treatments. To isolate extracellular enzymes, the cultures were centrifuged, and the resulting supernatants were filter-sterilized through a 0.22 μm membrane. To extract intracellular enzymes, the harvested cell pellets were lysed using a Scientz-IID ultrasonic cell disruptor (Ningbo, China) according to previously established parameters, followed by centrifugation to collect the clarified intracellular lysate. Subsequently, these four crude enzyme extracts were individually added to PBS supplemented with APAP. Samples were collected at 30 min intervals to quantify residual APAP concentrations and assess their respective degradation activities. Heat-inactivated enzyme controls were included across all assays to account for potential abiotic degradation.

2.6. Identification of Products from APAP Biodegradation

To determine the intermediate products of APAP degradation by strain DL27, three experimental conditions were established: (i) strain DL27 in MSM supplemented with 150 mg/L APAP; (ii) MSM containing 150 mg/L APAP without strain DL27; and (iii) strain DL27 in MSM without APAP. Samples were collected after 20 min and 2 h of incubation. The degradation products were examined by UHPLC-Q-Orbitrap HRMS. The instrumental parameters and the elution gradient program are shown in Text S6 and Table S1, respectively.

2.7. Multiomics Analysis

2.7.1. Whole Genome Analysis

Whole genome sequencing was conducted on both the Illumina and PacBio platforms at GENEWIZ Biotechnology Co., Ltd. (Suzhou, China). The predicted genes were functionally annotated using several databases including COG, GO, KEGG, and CARD. The genomic sequencing data have been submitted to the NCBI database with an accession number of PRJNA1432674.

2.7.2. Transcriptomic Analysis and qRT–PCR

Strain DL27 was cultured in MSM containing 150 mg/L of APAP for 20 min, while an APAP-free culture was used as the control. RNA samples were sent to GENEWIZ Biotechnology Co., Ltd. (Suzhou, China) for cDNA synthesis and library construction. The libraries underwent stringent quality control prior to sequencing on an Illumina NovaSeq 6000 platform. The clean reads were aligned to the genome by using Bowtie2 (v2.2.6). Differentially expressed genes (DEGs) were then identified by using the DESeq2 package. The experimental procedures for qRT-PCR and the primer sequences are provided in Text S7 and Table S2, respectively. The transcriptomic sequencing data have been deposited in the NCBI database under accession number PRJNA1433608.

2.7.3. Proteomic Analysis

Following the sampling protocols for RNA sequencing, bacterial cells were collected for protein extraction. A 100 μL sample was mixed with 200 μL of RIPA lysis buffer (Beyotime, China) and steel beads. Mechanical disruption was then performed for 2 min at 35 Hz, followed by ultrasonication using a Scientz-IID ultrasonic cell disruptor (Ningbo, China) for 20 min. Protein concentration was determined by BCA Protein Assay Kit (Beyotime, China). Subsequently, proteins were reduced, alkylated, and digested into peptides. Samples were analyzed by LC–MS/MS in positive ion mode using data-independent acquisition. The parameters are provided in Text S8.

2.8. Expression and Purification of Recombinant Protein

The ada gene was amplified from strain DL27 and cloned into the pET-28a vector to generate a recombinant plasmid. The plasmid was transformed into Escherichia coli BL21­(DE3) after sequence verification. The transformed cells were cultured in LB medium containing 100 mg/L kanamycin at 37 °C until the OD₆₀₀ reached approximately 0.6. Protein expression was induced with 0.2 mM IPTG at 16 °C for 16 h. The recombinant protein was purified using Ni-NTA resin (GenScript, USA) and analyzed by SDS-PAGE.

2.9. APAP Degradation by the Recombinant Protein

The reaction system for the enzyme activity assay consisted of a total volume of 1 mL, containing 10 mM PBS (pH 7.0), 300 μM APAP, and 0.5 μg of recombinant protein. Enzyme activity was measured over a temperature range of 10–60 °C and a pH range of 4.0–10.0 to determine the optimal conditions for biochemical characterization. Buffers at a concentration of 20 mM were used, including citric acid–sodium citrate (pH 4.0–6.0), NaH₂PO₄–Na₂HPO₄ (pH 7.0–8.0), and NaHCO₃–Na₂CO₃ (pH 9.0–10.0). To evaluate the effects of various metal ions (K⁺, Na⁺, Mg²⁺, Fe³⁺, Ca²⁺, Cu²⁺, Ni²⁺, and Zn²⁺) on enzymatic activity, the enzyme was incubated with each ion at a final concentration of 1 mM at 35 °C and pH 8.0 for 5 min. A reaction mixture without any added metal ions served as the control. After incubation at 35 °C for 5 min, the reaction was terminated by adding 1 mL of acetonitrile, and the products were identified using UHPLC-Q-Orbitrap HRMS. , The enzymatic kinetics of the AdA were determined as a function of APAP concentration ranging from 0 to 100 μM.

2.10. Molecular Docking and Dynamics Simulation

According to AlphaFold3, the three-dimensional structure of the amidase AdA was predicted, while the ligand APAP was retrieved from the PubChem database. Subsequently, the molecular recognition between APAP and AdA was analyzed through molecular docking simulations by AutoDock Vina. The resulting binding modes, along with the key interacting residues, were visualized by PyMOL. The docked APAP–AdA complex was subjected to molecular dynamics simulations using GROMACS 2023.2. Detailed operational steps are provided in Text S9.

2.11. Site-Directed Mutagenesis

The site-directed mutagenesis using PCR was performed to elucidate the functions of putative key amino acid residues involved in the interaction between APAP and AdA. In brief, the recombinant plasmid pET-28a-ada was used as a template, and specific residues were substituted with alanine using a site-directed mutagenesis kit (TianGen, China) along with sequence specific primers (Table S3). The PCR products were subsequently treated with DpnI to digest the parental template DNA. The resulting mutated plasmids were then transformed into E. coli DH5α and verified by DNA sequencing. The correctly mutated plasmids were further introduced into E. coli BL21­(DE3) for heterologous expression, enabling comparative analysis of the enzymatic activities between the wild-type and mutant proteins.

2.12. Statistical Analysis

All experiments were performed in triplicate, and data were presented as mean ± standard deviation. Statistical analyses were performed using one-way analysis of variance (ANOVA) and t-tests (SPSS 22.0), and a p-value <0.05 was considered statistically significant.

3. Results and Discussion

3.1. Isolation and Characterization of APAP-Degrading Bacterium

Of the 12 APAP-degrading strains isolated from activated sludge, all degraded over 50% of 150 mg/L APAP within 30 min at 30 °C and pH 7.0 (Figure A). Strain DL27 was selected for further study due to its highest degradation efficiency of 93.04%. TEM and SEM revealed that DL27 cells are coccobacillary and nonflagellated (Figure B,C). BLAST analysis of the 16S rRNA gene sequence against the EzBioCloud database showed the highest similarity to members of the genus Acinetobacter (Figure D). Furthermore, phylogenetic analysis showed that strain DL27 is presently closely related to Acinetobacter johnsonii CIP 64.6^T and Acinetobacter oryzae B23^T. Therefore, strain DL27 was identified as Acinetobacter sp. based on combined morphological and molecular evidence. While Acinetobacter species have been reported to participate in the remediation of several pollutants, including benzopyrene, phthalic acid esters, ochratoxin A, polystyrene, and phenol. This study markedly extends their metabolic flexibility to the effective degradation of APAP.

1.

Identification of highly efficient degrading strains. (A) Comparison of APAP degradation rates of different isolated strains. (B) Morphology of strain DL27 under SEM. (C) Morphology of strain DL27 under TEM. (D) Phylogenetic tree based on 16S rRNA sequence of strain DL27. Strain DL27 is indicated by a red asterisk. A phylogenetic tree was constructed using the neighbor-joining method, and bootstrap values were estimated from 1000 replicate analyses.

3.2. Degradation Characteristics of APAP by Strain DL27

Most degrading strains exhibit a specific optimal temperature range. A temperature range of 30–40 °C is suitable for APAP degradation by strain DL27, with the highest degradation rate of 97.90% observed at 35 °C (Figure A). Notably, strain DL27 also exhibited considerable activity at 15 and 50 °C. Such efficient degradation across a wide temperature range is uncommon among previously reported APAP-degrading bacteria. The biodegradation of pollutants is influenced by pH, as it affects both cell membrane permeability and enzyme activity. The optimal pH range was determined to be 7.0–9.0, with the highest degradation rate of 97.51% at pH 8.0 (Figure B). This finding is consistent with previous studies, in which Acinetobacter sp. XS-4 and Glutamicibacter sp. S2 demonstrated significant degradation of benzopyrene and cephalexin at pH 8.0. , Inoculum size is a critical parameter for the practical application of bioremediation. , Under inoculum sizes ranging from 5% to 25%, the degradation rate of APAP exceeded 80% after 30 min of cultivation (Figure C). The APAP degradation rate peaked at an optimal inoculation level of 15%, driven by enhanced biomass and enzymatic activity. Agitation speed is a key parameter in the bacterial degradation process, mainly affecting the dissolved oxygen level. Strain DL27 achieved an APAP degradation efficiency above 94% across a range of 50–250 rpm, with little variation (Figure D). The consistent APAP degradation by strain DL27 across a wide range of agitation speeds underscores its practical utility.

2.

Optimization of conditions for APAP degradation by strain DL27, kinetic analysis, and its application in agricultural soil. (A–D) Optimization of single-factor degradation conditions. Effect of temperature (A), pH (B), inoculum size (C) and agitation speed (D) on the APAP degradation rate, respectively. Bars represent the means of triplicate replicates (n = 3). Error bars depict the standard deviation of the means. (E, F) Degradation kinetics analysis. (E) First-order kinetic models of APAP degradation at various initial concentrations. (F) Relationship between initial APAP concentration and specific degradation rate. (G–I) Degradation in agricultural soil at initial concentrations of 15 mg/kg (G), 45 mg/kg (H), and 75 mg/kg (I). SS: sterile soil control; SS+DL27: sterile soil inoculated with strain DL27; NS: natural soil control; NS+DL27: natural soil inoculated with strain DL27.

Strain DL27 efficiently degraded APAP at concentrations of 15–450 mg/L (Figure S1). Complete degradation occurred within 30 min at 15–45 mg/L, and over 90% efficiency was maintained up to 150 mg/L. Beyond 225 mg/L, the degradation rate decreased markedly, which can be attributed to substrate-induced metabolic inhibition. ,, The degradation process at all concentrations fitted well to the pseudo-first-order kinetic model (R ² > 0.90, Figure E), showing the shortest half-life (t _1/2 ) at 45 mg/L (Table S4). Furthermore, the relationship between the specific degradation rate (q) and initial APAP concentration was well described by a substrate inhibition kinetic model (R ² = 0.9837, Figure F). The maximum specific degradation rate (qmax) was determined to be 0.1169 min^–1, half-saturation constant (K_s ) was 22.3812 mg/L, and substrate inhibition constant (K _i ) was 36.0682 mg/L. Similar substrate inhibition kinetics have been documented in other bacterial systems.

3.3. Bioremediation of APAP by Strain DL27 in Agricultural Soil and Municipal Wastewater

To evaluate the application potential of strain DL27 in complex, real-world environments, this study investigated its degradation kinetics of APAP at various initial concentrations in agricultural soil and municipal wastewater. In the uninoculated control groups, negligible APAP degradation occurred over the experimental period, indicating that abiotic degradation or indigenous microbiota-mediated degradation was highly limited (Figures G–I and S2). In contrast, upon inoculation with strain DL27, APAP in both soil and wastewater was rapidly degraded, with the degradation processes fitting the first-order kinetic model (Tables S5 and S6). In both soil and wastewater systems, the degradation rates in the nonsterile inoculated groups were significantly higher than those in the sterile inoculated groups. For instance, in the 45 mg/L wastewater system, the half-life (t _1/2 ) of APAP in the nonsterile group (8.98 min) was shortened by nearly half compared to that in the sterile group (17.86 min), suggesting a strong synergistic degradation effect between strain DL27 and the indigenous microorganisms. Compared to the soil system, APAP degradation in the wastewater system was more rapid. In the nonsterile wastewater inoculated groups at 15 mg/L and 45 mg/L, the half-lives of APAP were only 9.86 and 8.98 min, respectively. This could be attributed to the more sufficient contact between bacterial cells and pollutant molecules in the liquid matrix as well as the higher bioavailability of the pollutant in the aqueous phase. As the initial pollutant concentration increased, the degradation rate exhibited an initial increase, followed by a subsequent decrease. When the substrate concentration reached 75 mg/kg (or 75 mg/L), the degradation rate slowed due to substrate inhibition or toxic effects; nevertheless, DL27 maintained a robust tolerance and eliminated the majority of the pollutant within a short period. Several studies have evaluated the potential of microbial strains for pollutant remediation in actual environments. ,,

In summary, strain DL27 exhibited excellent biodegradation activity in both agricultural soil and municipal wastewater, showing a distinct synergistic degradation effect with indigenous microorganisms in nonsterile natural environments. These findings demonstrate that strain DL27 possesses promising application potential and practical value for the bioremediation of agricultural soils and municipal wastewater in complex real-world environments.

3.4. Degradation Enzyme Localization of Strain DL27

The enzymes produced by microorganisms for pollutant degradation can be either intracellular or extracellular. Extracellular enzymes act directly on pollutants, whereas intracellular enzymes require pollutants to be adsorbed and transported across the cell membrane before degradation can occur. , To determine the subcellular localization and expression pattern of the APAP-degrading enzymes in strain DL27, the degradation capacities of intracellular and extracellular crude enzyme extracts were evaluated under both induced and noninduced conditions. Results revealed that regardless of APAP preinduction, APAP concentrations in the extracellular extract group remained largely unchanged within 90 min, indicating negligible extracellular degradation activity (Figures S3A and S4A). In contrast, the intracellular degradation system proved to be strongly induced by APAP. Under noninduced conditions, intracellular extracts exhibited minimal activity, yielding a degradation rate of only 10.94% at 90 min (Figure S3B). Upon APAP preinduction, however, intracellular enzymatic activity was markedly enhanced in a time-dependent manner, achieving a degradation rate of 17.00% at 30 min and reaching 81.26% by 90 min (Figure S4B). Overall, APAP degradation in strain DL27 is predominantly mediated by intracellular inducible enzymes, whose expression and catalytic activity are significantly upregulated in the presence of APAP. These findings align with those of the previous studies. For example, during the biodegradation of BDE-47 by Stenotrophomonas sp. WZN-1, intracellular enzymes were identified as the primary drivers, whereas extracellular enzymes exhibited negligible activity.

3.5. Degradation Pathway of APAP in Strain DL27

High-resolution mass spectrometry plays an important role in the study of pollutant biodegradation. , The intermediates of the APAP biodegradation process by strain DL27 in the MSM were characterized using UHPLC-Q-Orbitrap HRMS. The parent compound APAP (m/z 152.07) was found in the control samples at an RT of 4.68 min. A total of seven major degradation products were identified, with their mass spectra shown in Figure S5, based on which a comprehensive biodegradation pathway was proposed (Figure ). The cleavage of the amide bond is catalyzed by amidase, initiating the degradation process. In this step, deacetylation of APAP produces DP1 (m/z 110.06), which is identified as PAP. This initial step is consistent with previous observations of APAP degradation by Pseudomonas taiwanensis AP-1. Subsequently, DP1 undergoes degradation via three pathways. First, deamination of DP1 produces DP2 (HQ, m/z 111.04), which is further hydroxylated to produce DP3 (BT, m/z 127.03). Second, intermolecular coupling of DP1 leads to the formation of a dimer, DP4 (m/z 202.09), which is subsequently oxidized to a benzoquinone-like compound, DP5 (m/z 200.07). Third, polycondensation of multiple DP1 molecules generates a nitrogen-containing polycyclic compound, DP6 (m/z 322.12), which is further oxidized at the phenolic hydroxyl group to produce DP7 (m/z 320.10). We noticed that the key products produced by different strains during the APAP degradation process reveal differences in their metabolic pathways. ,, Taken together, these findings demonstrate that APAP biodegradation by strain DL27 proceeds through a series of reactions, including amide hydrolysis, deamination, hydroxylation, redox reactions, and condensation. The cleavage of the amide bond is the initiating step. Notably, DP4, DP5, and DP7 were identified in this study for the first time, providing novel insights into the microbial metabolic pathways of APAP.

3.

Proposed pathways for APAP biodegradation by strain DL27. The chemical structures highlighted in red represent the novel APAP degradation products proposed in this study.

3.6. APAP Degradation Mechanisms Based on Multiomics Analyses

3.6.1. Genomic Analysis of Strain DL27

Genomic analysis provides valuable insights into the key microbial enzymes involved in pollutant degradation and facilitates the identification of the degradation pathways. , The assembled genome consists of one chromosome and two plasmids, with a total size of 3,544,417 bp and a GC content of 41.53% (Figure S6 and Table S7). The gene prediction analysis indicated the presence of a total of 3, 336 genes that account for 85.97% of the entire genome. In this study, the proposed APAP biodegradation pathway is initiated by amidase that catalyzes the hydrolysis of APAP to PAP. Using previously characterized APAP amidases as query sequences, three amidase homologous sequences, gene1845, gene2495, and pa16, were identified. In addition, several other enzymes potentially involved in downstream APAP degradation were annotated, including putative deaminase, phenol 2-monooxygenase, NAD­(P)­H-dependent oxidoreductase, and catechol 1,2-dioxygenase.

3.6.2. Transcriptomic Analysis of APAP Degradation by Strain DL27

The overall distribution of gene expression across all samples is shown in Figure A. Principal component analysis and intersample correlation analysis showed significant transcriptional changes between the APAP-treated group and control group, along with high consistency among biological replicates (Figure B,C). The analysis of differential expression identified a total of 1,032 DEGs in strain DL27 after APAP treatment, including 501 upregulated and 531 downregulated genes (Figure D,E). The upregulation of specific genes appears to enhance the metabolism of APAP.

4.

Transcriptomic analysis of APAP degradation by strain DL27. (A) Gene expression distribution across all samples. The vertical axis represents gene expression levels. The horizontal axis represents sample names. (B) Principal component analysis of the transcriptome data. Red circles and blue triangles represent the control and treatment groups, respectively. (C) Analysis of correlation between samples. The color bar on the right indicates the correlation coefficients (0.4 to 1.0), with red and blue representing high and low correlations, respectively. (D) Volcanic plot of the distribution of DEGs between the APAP treatment group and the control group. Red and blue dots represent significantly upregulated and downregulated genes, respectively, while gray dots indicate genes without significant differences. The thresholds for DEGs were set as |log₂FoldChange| ≥ 1 and p < 0.05. (E) Number of DEGs between control group and APAP treatment group; (F) KEGG and (G) GO enrichment of the DEGs between control group and APAP treatment group. The horizontal and vertical axes represent the Rich factor and the enriched pathways, respectively. Bubble size and color correspond to the number of DEGs and the statistical significance, respectively.

KEGG enrichment analysis revealed that most of the upregulated genes were associated with metabolism and the environmental response (Figure F). Specifically, enrichment in aromatic compound degradation pathways suggests that DL27 possesses specialized enzymes for APAP catabolism. Additionally, the activation of glutathione metabolism points to an active response to oxidative stress to maintain cellular homeostasis. Regarding environmental adaptation, the enrichment of bacterial secretion systems and protein export pathways suggests enhanced transmembrane transport for enzyme secretion, while the enriched quorum-sensing pathway likely coordinates nutrient bioavailability, transport, and transformation processes. GO enrichment analysis further characterized the molecular response of strain DL27 during APAP degradation (Figure G). The highest enrichment of GO terms indicates that the catalytic activity may involve a wide range of enzymes in APAP degradation. The significant enrichment of phenol 2-monooxygenase activity indicates the involvement of monooxygenase-mediated hydroxylation reactions in APAP degradation. Bacterial oxidoreductases are well-known for their ability to degrade aromatic pollutants, and this study further supports their indispensable role in APAP degradation, as evidenced by the significant enrichment of oxidoreductase-related functions. Additionally, transmembrane transport and energy-coupling processes, such as ATPase activity, the proton-transporting ATP synthase complex, and various ion channel activities, were significantly enriched.

Transcriptomic analysis revealed that among the three APAP-degrading amidases identified in the genome of strain DL27, the gene pa16 exhibited a 167.26-fold upregulation. In addition, genes encoding putative deaminase, phenol 2-monooxygenase, and catechol 1,2-dioxygenase were upregulated by 2.88-, 2.63-, and 9.00-fold, respectively. An NAD­(P)­H-dependent oxidoreductase exhibited the highest increase, with a 383.66-fold upregulation, while several other oxidoreductases also showed elevated expression levels. These results indicate that amidase catalyzes the initial hydrolytic cleavage of APAP, while the other enzymes participate in the downstream steps of the degradation pathway. The expression patterns of key DEGs identified by qRT-PCR were consistent with the RNA-seq data (Figure S7), validating the transcriptomic analysis and further supporting the important roles of these genes in APAP degradation.

3.6.3. Proteomic Analysis of APAP Degradation by Strain DL27

Proteomic analysis can reveal the mechanistic roles of key enzymes involved in complex biodegradation pathways. ,, Principal component analysis revealed that APAP-treated and control groups exhibited distinct global protein expression patterns (Figure S8A). Exposure of strain DL27 to APAP resulted in 608 differentially expressed proteins (DEPs), including 195 upregulated and 413 downregulated proteins (Fold change ≥2.0 or ≤ 0.5) (Figure S8B). Subcellular localization analysis showed that nearly all upregulated proteins (191 proteins) were localized in the cytoplasm, with very few detected in the periplasm or extracellular space (Figure S8C). This finding indicates that intracellular enzymatic activity is primarily responsible for APAP degradation in strain DL27. KEGG enrichment analysis of the proteomic data revealed that nitrogen metabolism was the most significantly enriched pathway, suggesting its potential involvement in APAP degradation (Figure S8D). Additionally, enhanced biosynthesis of branched-chain amino acids (valine, leucine, and isoleucine) and sulfur-containing amino acids (cysteine and methionine) not only supports energy production but may also improve bacterial tolerance to oxidative stress by promoting reduced glutathione synthesis. The enrichment of degradation pathways for toluene, chlorocyclohexane, and chlorobenzene suggests that strain DL27 may produce enzymes capable of degrading APAP and its aromatic intermediates.

GO enrichment analysis of the DEPs further elucidated the functional responses of strain DL27 during APAP degradation (Figure S8E). Increased activities of phenol 2-monooxygenase and oxidoreductases were consistent with transcriptomic results, highlighting their important roles in APAP degradation. , Importantly, proteins involved in amino acid biosynthesis, particularly branched-chain and pyruvate-family amino acids, were significantly upregulated, suggesting that DL27 enhances amino acid production to meet the metabolic energy demands. Enhanced expression of proteins involved in small-molecule and ion binding may contribute to maintaining ion homeostasis and facilitating metabolite transport, thereby improving APAP degradation efficiency.

Notably, pa16 was significantly upregulated at the transcript level, while its encoded protein showed an 18.60-fold increase in abundance in the proteomic analysis, further supporting its key role in the initial step of APAP degradation. This gene was designated as ada for further studies. Furthermore, putative deaminases, phenol 2-monooxygenase, and catechol 1,2-dioxygenase were upregulated 4.60-, 2.24-, and 2.60-fold, respectively, at the protein level. An NAD­(P)­H-dependent oxidoreductase showed a 3.66-fold increase along with several other oxidoreductases. These results collectively highlight the crucial roles of these enzymes in the APAP degradation.

Multiomics approaches offer key insights into the biodegradation of environmental contaminants. ,, This study integrated transcriptomic and proteomic analyses to elucidate the mechanisms of APAP degradation in the strain DL27. The results indicate that APAP is initially imported into the cell via an upregulated, ATP-dependent transmembrane transport system. Once intracellular, APAP is degraded through pathways involving key enzymes, including amidase, putative deaminase, phenol 2-monooxygenase, catechol 1,2-dioxygenase, and NAD­(P)­H-dependent oxidoreductases, supported by upregulated cofactor biosynthesis (riboflavin, porphyrin, and biotin metabolism). To mitigate oxidative stress induced by APAP and its intermediates, strain DL27 enhances its antioxidant capacity by upregulating the glutathione metabolism, amino acid biosynthesis, and antioxidant proteins. Additionally, enhanced energy metabolism and ion transport maintain the cellular homeostasis. These findings clarify the stress-responsive biodegradation mechanisms of strain DL27, offering a theoretical basis for its potential applications in bioremediation.

3.7. Phylogenetic and Sequence Analyses Reveal Mechanistic Insights into the Conserved Catalytic Domains of AdA

To investigate the evolutionary relationship and phylogenetic novelty of AdA from Acinetobacter sp., we constructed a phylogenetic tree using homologous sequences retrieved from the UniProtKB/Swiss-Prot database (Figure A). The retrieved homologues originated from taxonomically diverse bacterial taxa, predominantly spanning the Pseudomonadota, Actinomycetota, and Bacillota. Notably, phylogenetic analysis revealed that AdA from Acinetobacter sp. belongs to the upper major clade. Although it shares an evolutionary ancestry with the cluster containing Corynebacterium ammoniagenes (A0ABN0ABT5) and Pseudochrobactrum asaccharolyticum (A0A366E0P5), AdA was resolved as a highly distinct, deeply branching lineage. Taken together, these phylogenetic features suggest that AdA from Acinetobacter sp. represents a novel, functionally uncharacterized member of this protein family.

5.

Phylogenetic analysis and multiple sequence alignment of AdA. (A) Phylogenetic tree of AdA from Acinetobacter sp. DL27 and its homologous proteins from various bacterial species. (B) Multiple sequence alignment.

In this study, multiple sequence alignment of AdA was performed against previously reported APAP-degrading enzymes (Figure B). Secondary structure prediction indicates that AdA adopts a complex, mixed α/β fold. The full-length protein is predicted to contain 15 α-helices, 15 β-strands, and five 310-helices. This alternating α/β topology is a characteristic feature shared by various hydrolase and amidase families. Multiple sequence alignment of the AdA protein sequence with identified APAP-degrading amidases revealed that the Gly/Ser-rich motif (GGSS­[GS]­G) and the Ser-Ser-Lys catalytic triad are highly conserved, indicating that AdA is an amidase belonging to the amidase signature family. ,

3.8. Heterologous Expression, Purification, and Functional Characteristics of AdA

Following the expression of the recombinant protein in E. coli BL21 (DE3), the target protein was purified via Ni-NTA affinity chromatography and eluted with PBS buffer containing 300 mM imidazole. The target protein, AdA, comprises 511 amino acids, with a theoretical molecular weight of 55.2 kDa and a predicted isoelectric point of 5.71. SDS-PAGE analysis indicated that purified AdA was resolved as a single band with a molecular weight of approximately 55.0 kDa, which is close to its predicted value of 55.2 kDa (Figure A).

6.

Functional identification and enzymatic characterization of AdA. (A) SDS-PAGE analysis of AdA. Lane M: Protein marker; Lane 1: the purified protein AdA. Effect of temperature (B), pH (C) and metal ions (D) on the enzymatic activity of the AdA. (E) Michaelis–Menten kinetics of AdA on APAP hydrolyzation.

Characterizing the biochemical properties of proteins provides key insights for their practical application. To evaluate the industrial and practical potential of the purified AdA, its biochemical properties were systematically characterized. The enzyme exhibited catalytic activity across a wide temperature range of 10–60 °C (Figure B). The maximum relative activity was achieved at 35 °C. Notably, AdA retained over 50% of its peak activity even at the temperature extremes of 10 and 60 °C, suggesting a relatively broad temperature tolerance that could be advantageous in fluctuating environmental conditions.

The effect of pH on the catalytic performance of AdA is illustrated in Figure C. The enzyme showed a clear preference for neutral to weakly alkaline environments with the optimum activity observed at pH 8.0. High relative efficiency (>90%) was maintained at pH 9.0. Conversely, the enzymatic activity declined progressively under acidic conditions, dropping to approximately 28.80% of its maximum value at pH 4.0, indicating that the active site or conformation of the enzyme may be sensitive to protonation at lower pH values.

Metal ions are critical factors that often modulate enzymatic performance. Therefore, the effects of various metal ions on the activity of AdA were assessed (Figure D). Compared to the control group (without metal ions), the addition of K⁺ slightly stimulated the enzymatic reaction, increasing the relative activity to approximately 107.76%. In contrast, the presence of Mg²⁺ exerted no statistically significant effect on the enzyme’s catalytic capability. Other tested divalent and trivalent cations exhibited varying degrees of inhibitory effects; Fe³⁺, Ca²⁺, and Ni²⁺ moderately suppressed the activity (retaining 72.98%–84.58% relative activity), while Zn²⁺ reduced the activity to around 56.72%. Most notably, Cu²⁺ acted as a potent inhibitor, severely diminishing the relative activity to approximately 32.47%. The pronounced inhibitory effect of Cu²⁺ on AdA may result from metal-induced perturbation of the enzyme structure or catalytic environment, although the precise molecular mechanism requires further investigation.

Kinetic analysis determined the K _m , V _max, and k _cat of AdA to be 8.96 ± 1.03 μM, 1.62 ± 0.03 μM/s, and 80.99 s^–1, respectively (Figure E). According to Table S8, the substrate affinity of previously reported hydrolases for APAP followed the order of ApaH1 > AdA > Mah > ApaH2 > AmpA > AAA. However, their catalytic efficiencies (k _cat/K _m ) ranked as AdA > ApaH1 > Mah > ApaH2 > AmpA > AAA. In particular, the catalytic efficiency of AdA was 1.18, 5.87, 6.19, 6.37, and 1.51 × 10⁶-fold higher than those of ApaH1, Mah, ApaH2, AmpA, and AAA, respectively. − Consequently, although ApaH1 exhibited a slightly higher substrate affinity, AdA possessed higher catalytic efficiency. In conclusion, we purified the amidase AdA from strain DL27 and demonstrated its exceptional ability to hydrolyze the amide bond of APAP. Given its higher catalytic efficiency compared to known APAP amidases, AdA represents a highly promising candidate for the bioremediation of APAP contamination.

3.9. Amidase Hydrolysis of APAP by AdA to Produce PAP

As shown in the HPLC chromatograms (Figure A), the APAP control group exhibited a prominent peak at a retention time of approximately 5.11 min. Following incubation with the enzyme AdA, the peak corresponding to APAP was substantially reduced, while a new product peak emerged at a shorter retention time of ∼1.03 min. The mass spectrum of the substrate peak displayed a dominant signal at m/z 152.0704, which is consistent with the protonated molecular ion of APAP (Figure B). For the newly formed product peak, a major ion signal was observed at m/z 110.0600, corresponding to the protonated form of PAP (Figure C).

7.

Amidase hydrolysis of APAP by AdA. (A) HPLC analysis of the transformation of APAP by purified AdA. Mass spectrometric analysis of APAP (B) and the catalytic product (C) in the reaction system.

3.10. Structural Prediction and Molecular Docking Analysis of AdA

The three-dimensional structure of the recombinant protein AdA was predicted using AlphaFold3. As illustrated in Figure A, the protein displays a well-defined and regular secondary structure, characterized by cyan α-helices, purple β-sheets, and pink loops. The core region is composed of multiple parallel or antiparallel β-sheets, flanked by peripheral α-helices, which adopts the typical α/β-hydrolase fold topology. This compact 3D conformation not only provides a structural basis for the protein’s stability in aqueous environments but also establishes the necessary spatial framework for the formation of its catalytic pocket.

8.

Structure, sequence analysis, and molecular docking of AdA. (A) 3D structural model of AdA. (B) Sequence conservation analysis. (C) Conserved motif analysis. (D) Molecular docking simulation of APAP and AdA.

The sequence conservation of AdA was analyzed and mapped onto its 3D surface model (Figure B). The results indicated that the conservation scores of the surface residues ranged from 1 (highly variable, dark cyan) to 9 (highly conserved, dark magenta). The analysis revealed that highly conserved residues (such as K82, S161, V302, and T431) are predominantly clustered within the clefts or grooves of the protein. These highly conserved patches typically correspond to the catalytically active centers or ligand-binding sites of the enzyme. In contrast, highly variable regions are primarily distributed on the solvent-exposed surfaces. Specifically, the spatial positioning of conserved residues, including K82 and S161, suggests that they may play key roles in substrate recognition, binding, or catalytic reactions.

Conserved motif analysis of this protein identified nine representative motifs (Figure C). The sequence logos of each motif illustrate the frequency of the amino acid conservation at specific positions. Their spatial distribution along the polypeptide chain reveals a well-ordered arrangement and a high level of sequence conservation, representing the signature sequence features of this protein family.

Molecular docking analysis was performed to demonstrate the interaction of AdA with the APAP. The active site of AdA comprises residues Ser185, Ser161, Lys82, Phe85, Ile220, Ala182, and Thr180 (Figure D). Specifically, Phe85 anchors the substrate into the pocket by forming a hydrogen bond with the phenolic hydroxyl group of APAP. The amide bond of APAP forms hydrogen bonds with Ser185, Ser161, and Lys82, stabilizing the structure of the complex. Moreover, the phenyl ring and amide moiety of APAP are stabilized by Ile220, Ala182, and Thr180, which enhances substrate stabilization. This binding arrangement allows APAP to be properly positioned for hydrolysis mediated by the catalytic triad (Ser185, Ser161, and Lys82). The essential amino acid residues in AdA involved in APAP hydrolysis were identified on the basis of these findings.

3.11. Mechanism Underlying the Catalytic Action of AdA on APAP

Root-mean-square deviation (RMSD) is commonly used to quantify conformational deviations in molecular systems during molecular dynamics simulations. The RMSD curve of the APAP–AdA complex reached equilibrium during the 45–75 ns interval, with fluctuations ranging from 0.55 to 0.60 nm (Figure A). A trough appeared at approximately 78 ns, where the RMSD decreased by about 0.35 nm. Subsequently, from 80 to 100 ns, the RMSD increased again and fluctuated between 0.50 and 0.55 nm, indicating that the binding gradually stabilized. Additionally, root-mean-square fluctuation (RMSF) is another key parameter, characterizing the degree of fluctuation for individual amino acid residues. Most amino acid residues exhibit low RMSF values (<0.25 nm), indicating that the overall structure of AdA remains relatively stable after binding with APAP (Figure B). The solvent-accessible surface area of the complex fluctuated between 200 and 220 nm², with a marked reduction in fluctuation amplitude after 40 ns (Figure C). This result indicates that the APAP–AdA complex has reached a dynamic equilibrium state, and its overall structure has not undergone significant changes. The radius of gyration (Rg) is an important parameter for assessing the overall structural compactness of protein–small molecule complexes. The Rg value gradually stabilizes throughout the simulation (Figure D), indicating that the structure of the APAP–AdA complex becomes progressively more stable.

9.

Molecular dynamics simulations and site-directed mutagenesis analysis of AdA in complex with APAP. (A) RMSD curves of APAP-AdA complex. (B) RMSF curves of APAP-AdA complex. (C) SASA curve of the APAP-AdA complex. (D) Radius of gyration curve of the APAP-AdA complex. (E) SDS-PAGE analysis of purified mutants. Lane M: Protein marker; Lane 1–3: Purified S161A, S185A, and K82A mutants, respectively. (F) Relative enzymatic activity analysis of AdA and its mutants.

According to sequence alignment, molecular docking, and molecular dynamics simulations, Ser161, Ser185, and Lys82 were predicted to form a conserved catalytic triad in the active site of AdA. To test this hypothesis experimentally, site-directed mutagenesis was performed to substitute each of these residues with alanine, generating three mutants: S161A, S185A, and K82A. Successful expression and purification of the mutant proteins were confirmed by SDS–PAGE analysis (Figure E). All three mutants exhibited an almost complete loss of catalytic activity toward APAP compared with the wild-type AdA, as determined by enzymatic activity assays (Figure F). The substantial decrease in activity highlights the critical importance of these residues, likely due to disruption of the catalytic pocket, which prevents proper APAP binding. Taken together, these findings provide strong evidence that the Ser161–Ser185–Lys82 catalytic triad is essential for the enzyme’s catalytic activity toward APAP.

Glossary

Abbreviations

PPCPs

pharmaceutical and personal care products

APAP

acetaminophen

PAP

p-aminophenol

HQ

hydroquinone

BT

1,2,4-benzenetriol

DEGs

differentially expressed genes

DEPs

differentially expressed proteins

RMSD

root-mean-square deviation

RMSF

root-mean-square fluctuation

Rg

radius of gyration

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acs.jafc.6c09241.

• Details of medium formulation, isolation and identification of APAP-degrading bacteria, determination of APAP concentration, biodegradation characteristics and soil bioremediation kinetics of APAP, activities of intracellular and extracellular enzymes, identification of biodegradation products, genomic features and chromosome circle map of strain DL27, LC-MS/MS proteomic analysis, primer sequences and qRT-PCR analysis of key genes, and molecular dynamics simulations (PDF)

B.K.: Methodology, investigation, writingoriginal draft, data curation. Y.Y.: Formal analysis, software, writingreview and editing. L.W.: Investigation, writingreview and editing. D.M.: Writingreview and editing. Z.S.: Writingreview and editing. Y.L.: Writingreview and editing. Z.Z.: Writingreview and editing. S.D.: Conceptualization, funding acquisition, supervision, writingreview and editing.

The authors declare no competing financial interest.