Pseudomonas aeruginosa SG01: A Novel Polyethylene-Degrading Bacterium in Petrochemical Wastewater

TL;DR

This study identifies a new bacterium, Pseudomonas aeruginosa SG01, that can break down polyethylene, a common plastic pollutant, and change its surface properties.

Contribution

The discovery of Pseudomonas aeruginosa SG01 as a novel polyethylene-degrading bacterium in petrochemical wastewater.

Findings

SG01 showed the highest growth rate when using polyethylene as its sole carbon source.

Treated PE films had reduced mass and increased hydrophilicity, as shown by contact angle and FTIR analysis.

SEM images revealed surface erosion and cracks on PE films after bacterial treatment.

Abstract

Microbial degradation technology presents a sustainable approach to address the environmental persistence of polyethylene (PE). In this study, a consortium of PE-degrading strains was isolated from sludge in the production wastewater of a PE-manufacturing plant. Among these strains, Pseudomonas aeruginosa SG01 demonstrated the highest cellular growth rate in culture medium, indicating its capacity to efficiently degrade PE and utilize it as the sole carbon source. Following treatment with SG01, the PE films exhibited a significant reduction in mass along with a clear decrease in surface contact angle, suggesting an improvement in hydrophilicity. Fourier transform infrared spectroscopy (FTIR) analysis detected the formation of new absorption bands on the treated PE films, corresponding to hydroxyl, carboxyl, and amide functional groups. Scanning electron microscopy (SEM) observations further revealed the presence of erosion pits and network-like cracks on the film surface. This study confirms that Pseudomonas aeruginosa SG01 can effectively degrade PE and modify its surface properties, offering a novel microbial resource for the bioremediation of PE contamination.

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Full text

1. Introduction

Polyethylene (PE), a representative thermoplastic polyolefin, features a molecular structure composed of regular [–CH_2_–CH_2_–]n repeating units [1]. It ranks as the most extensively produced synthetic polymer worldwide, with broad applications in packaging, agriculture, and construction [2]. Recent studies demonstrate that environmental PE waste undergoes progressive physicochemical degradation through photo-oxidation, mechanical abrasion, and hydraulic forces, ultimately fragmenting into microplastics (MPs; 1–5000 μm) [3,4]. These secondary MPs exhibit remarkable environmental persistence, with estimated half-lives ranging from 50 to 100 years. Critically, their hydrophobic surfaces and high specific surface area enable selective adsorption of persistent organic pollutants (POPs) and heavy metals, forming composite contaminants [5,6]. Upon biological uptake, these contaminants undergo trophic transfer with potential for biomagnification, inducing oxidative stress, inflammatory responses, and metabolic dysregulation [7,8]. Epidemiological evidence further indicates a significant positive association (p < 0.05) between exposure to PE-derived MPs and cardiovascular disease incidence. Notably, petrochemical wastewater treatment systems have been identified as major accumulation hubs for such MPs [9,10,11], providing a mechanistic basis for bioaugmentation-mediated remediation strategies.

Conventional physicochemical treatment methods, including landfill, incineration, and mechanical recycling, are not only inefficient in decomposition and pose risks of soil and groundwater contamination but also consume substantial land resources [11,12]. For example, the incineration of PE plastics generates persistent organic pollutants such as polycyclic aromatic hydrocarbons, while incompletely degraded agricultural mulch films can impair soil fertility. In recent years, enzymatic and microbial degradation of plastics has attracted growing attention due to its environmental compatibility and cost-effectiveness. This approach produces neither harmful gases nor leachate, and the resulting degradation products can be repurposed as fertilizers or other reusable resources [13,14]. Using metagenomic sequencing, researchers have identified several microbial taxa with PE-degrading potential in natural water bodies, sediments, and landfills, including genera such as Rhodococcus and Pseudomonas [15,16]. These findings provide a crucial foundation for developing environmentally friendly biodegradation technologies targeting PE microplastics (PE-MPs), with important implications for advancing lifecycle-based management of plastic pollution. To date, numerous bacteria, fungi, and actinomycetes—such as waxworm gut bacteria, Pseudomonas, filamentous fungi, and Bacillus—have been reported to participate in PE degradation [17,18,19,20]. For instance, PARK et al. [21] isolated a thermophilic bacterial consortium from landfills capable of degrading PE. A number of studies have also demonstrated that bacteria derived from the guts of insects like whiteflies, superworms, mealworms, yellow mealworms, and wax moth larvae exhibit PE-degrading activity [22,23,24].

To address the need for PE pollution remediation, this study isolated and screened the strain Pseudomonas aeruginosa SG01 from PE production wastewater within a petrochemical facility, demonstrating significant PE-degrading potential. The degradation mechanisms of this strain on PE substrates were systematically investigated using comprehensive characterization techniques, including scanning electron microscopy (SEM), Fourier-transform infrared spectroscopy (FTIR), and contact angle analysis. Results indicate that SG01 effectively alters the surface morphology and chemical properties of PE through bio-oxidation processes. This work not only enriches the microbial germplasm resources available for PE degradation in petrochemical environments but also provides theoretical and technical support for in situ bioremediation of plastic waste in the petrochemical industry.

2. Materials and Methods

2.1. Soil, Sewage, Sludge, and PE Plastic Products

Soil, wastewater, and sludge samples were collected from exposed surfaces and wastewater treatment tanks at the Qilu Petrochemical Plastics Plant (Zibo, Shandong Province, China). Sampling was performed at four random sites across workshops producing high-density polyethylene (HDPE), low-density polyethylene (LDPE), polypropylene (PP), and general polyethylene (PE), resulting in four soil samples. Subsurface soil (10–15 cm depth) was collected using a five-point sampling method and stored in sterile glass containers under temperature-controlled transport (4 °C) to preserve sample integrity. Sludge and wastewater were collected from the central zone of treatment tanks, allowed to settle, and then filtered to separate settled sludge from supernatant. This process yielded one sludge sample and one wastewater sample. The collected sludge was stored in clean glass containers under controlled temperature and humidity. PE plastic products used in the study were supplied by Huayuan Polymer Co., Ltd. (Qingdao, Shandong Province, China).

2.2. Culture Media and Main Chemical Reagents

The mineral salt base medium was formulated with 3000 mg/L Na_2_HPO_4_, 1200 mg/L KH_2_PO_4_, 800 mg/L MgSO_4_·7H_2_O, and 500 mg/L (NH_4_)2_SO_4. Trace elements were supplemented as follows: 0.001 mg/L CuCl_2_·2H_2_O, 0.5 mg/L Na_2_EDTA, 0.03 mg/L H_3_BO_3_, 0.02 mg/L CoCl_2_·6H_2_O, 0.2 mg/L FeSO_4_·7H_2_O, 0.01 mg/L ZnSO_4_·7H_2_O, 0.003 mg/L MnCl_2_·4H_2_O, 0.003 mg/L Na_2_MoO_4_·2H_2_O, and 0.002 mg/L NiCl_2_·6H_2_O. For bacterial cultivation, Lysogeny Broth (LB) medium containing 11.5 g/L NaCl, 11.5 g/L tryptone, and 5 g/L yeast extract was used. The selective screening medium was prepared by adding 15 g/L agar and 3 g/L of sterilized PE particles (<100 μm) to the base medium. The mixture was autoclaved at 120 °C (100.0 kPa) for 110 min, after which 10% (v/v) sterile LB medium was aseptically incorporated.

2.3. PE Pretreatment

PE microplastic particles (100 μm in diameter) were sterilized by ultraviolet (UV) irradiation. The procedure involved evenly dispersing the particles in sterile glass Petri dishes and exposing them to UV light (254 nm, 30 W) for a total of 6 h. During sterilization, the particles were stirred hourly with a sterile glass rod to ensure uniform exposure. After treatment, the particles were spread onto LB agar plates and incubated at 30 ± 0.5 °C for 72 h. The absence of microbial growth (CFU = 0) confirmed complete sterilization, and the particles were thus approved for use in subsequent degradation experiments.

A multi-step sterilization protocol was implemented. Original PE films were precisely cut into 3.0 cm × 3.0 cm specimens. Surface cleaning was performed by immersion in anhydrous ethanol (≥99.7% purity) and ultrasonication in ultrapure water (resistivity ≥18.2 MΩ·cm), followed by UV irradiation (254 nm, 30 W, 6 h). Sterilization effectiveness was verified by incubating the treated films on nutrient agar at 30.0 ± 0.1 °C for 72 h, with sterility confirmed based on colony counting (detection limit: <1 CFU/cm^2^).

2.4. Enrichment and Screening of PE-Degrading Bacteria

A comprehensive screening system for PE-degrading bacteria was established as follows (Figure 1A). First, 4.5 g of soil sample was precisely weighed and thoroughly mixed with 0.5 g each of wastewater and sludge samples. This mixture was then introduced into 50 mL of sterile mineral salt medium (composition: Na_2_HPO_4_·12H_2_O 1.5 g/L, KH_2_PO_4_ 0.5 g/L, NH_4_Cl 1.0 g/L, MgSO_4_·7H_2_O 0.2 g/L, CaCl_2_ 0.01 g/L; pH 7.0 ± 0.2). Incubation was carried out in a constant-temperature shaker at 30 ± 0.5 °C with shaking at 180 ± 5 rpm for 72 h. After allowing 24 h for sedimentation, the supernatant was passed through a 0.22 μm membrane filter to obtain the primary bacterial suspension.

The experiment was performed in 250 mL Erlenmeyer flasks. For the experimental group, each flask contained 2.00 ± 0.05 g of PE plastic film (thickness: 50 ± 5 μm), 45 mL of the mineral salt medium described previously, and 5 mL of the primary bacterial suspension. The control group contained 2.00 ± 0.05 g of PE film and 50 mL of mineral salt medium only. Each group was set up with four sampling time points (15, 30, 45, and 60 days) and three replicates per time point. All flasks were incubated under constant shaking (180 ± 5 rpm) at 30 ± 0.5 °C.

During sampling, 1 mL of culture was accurately collected and serially diluted ten-fold (10^−1^ to 10^−6^). Preliminary tests indicated that a 10^−4^ dilution using sterile mineral salt medium produced an optimal colony density of 30–300 CFU/mL. A 100 μL aliquot from the 10^−4^ dilution was spread onto screening solid medium (1.5% agar) supplemented with 1% (w/v) PE powder (100 ± 10 μm). The plates were inverted and incubated under precisely controlled conditions at 30 ± 0.5 °C for 72 h (±2 h). Single colonies were subsequently purified through streak plating for at least three generations to obtain axenic cultures.

2.5. Identification of PE-Degrading Bacteria

The PE-degrading capability of isolated strains was validated through a selective culture approach [25]. Purified colonies were inoculated onto sterile mineral salt medium plates containing PE microplastic particles (100 ± 5 μm in diameter) as the sole carbon source and incubated at 30 ± 0.5 °C for 72 ± 2 h. Candidate strains showing visible growth (colony diameter ≥ 1 mm, regular morphology) underwent secondary purification via streak plating. Positive strains were preserved in 20% (v/v) glycerol at −80 °C for subsequent 16S rRNA gene sequencing and assessment of degradation performance.

Target strains were identified by 16S rRNA gene sequencing. Genomic DNA was extracted using a bacterial DNA isolation kit and amplified by PCR with the universal primers 27F (5′-AGAGTTTGATCCTGGCTCAG-3′) and 1492R (5′-TACGGCTACCTTGTTACGACTT-3′). The 50 μL PCR reaction mixture contained: 25 μL of Taq DNA polymerase premix, 2 μL each of forward and reverse primers (final concentration 0.4 μmol·L^−1^), 3 μL of genomic DNA (50 ng·μL^−1^), and 18 μL of nuclease-free water. PCR cycling conditions were as follows: initial denaturation at 95 °C for 90 s; 35 cycles of denaturation at 95 °C for 10 s, annealing at 58 °C for 10 s, and extension at 72 °C for 70 s; followed by a final hold at 4 °C. Amplified products were verified by 1% agarose gel electrophoresis and sequenced by Sangon Biotech (Shanghai, China). Sequences were submitted to NCBI GenBank and analyzed using BLAST (2.16.0) for phylogenetic identification.

Biomass changes in the PE-degrading bacteria were monitored in an inorganic salt liquid medium with PE as the sole carbon source. The isolated degrading strains were first activated in LB medium and shaken at 30 °C and 180 r/min. The logarithmic growth phase was determined based on the OD_600_ growth curve [26]. After reaching this phase, cells were harvested by centrifugation at 8000 r/min for 10 min. The pellet was resuspended in inorganic salt medium and centrifuged again for 10 min; this washing step was repeated three times. Finally, the bacterial suspension was adjusted with inorganic salt medium to an OD_600_ of 1.0. This inoculum was then added at 20% (v/v) to a screening medium containing 100 μm PE powder. A control group consisting of sterile screening medium with the same mass of 100 μm PE powder was prepared in parallel. Three replicates were set up for each group. All cultures were incubated in a shaker at 30 °C and 180 r/min, and OD_600_ was measured every 48 h.

The activated degrading strain (OD_600_ = 1.0 ± 0.05) was inoculated at 10% (v/v) into a 250 mL Erlenmeyer flask (pre-autoclaved at 121 °C for 20 min), consisting of 5 mL of bacterial suspension and 45 mL of inorganic salt medium. Each experimental group included four time points (0, 15, 30, and 45 d) with three biological replicates. A precisely weighed PE film (2.000 ± 0.005 g, thickness 50 ± 5 μm) was added to each flask. After sealing, the flasks were incubated in a constant-temperature shaker at 30 ± 0.5 °C and 180 ± 5 rpm. To accurately determine mass changes in the PE film, the following standardized post-treatment procedure was established: samples were soaked in 75% (v/v) ethanol for 36 h, ultrasonically cleaned three times with ultrapure water (resistivity ≥ 18.2 MΩ·cm) for 10 min each, dried in the dark at room temperature for 24 h, and finally weighed using an analytical balance with an accuracy of 0.1 mg. Preliminary experiments confirmed that the weighing error of this method was within ±0.5%.

The surface morphology of PE films after microbial degradation was characterized by SEM. After drying at 60 °C, samples were mounted on a specimen holder with conductive double-sided tape and observed using a Hitachi S3400 microscope (Hitachi High-Technologies Corporation, Tokyo, Japan). An acceleration voltage of 15 kV was applied to systematically examine surface changes at different magnifications. Representative areas were selected for high-resolution imaging. All steps were performed under standard laboratory conditions to ensure result reliability.

FTIR analysis was performed to characterize changes in the chemical structure of PE films following microbial degradation. The procedure was as follows: after drying at a constant temperature of 60 °C, the treated commercial PE film samples were placed directly into the FTIR sample chamber for measurement. Prior to analysis, the system was purged with high-purity nitrogen to minimize environmental interference, and baseline correction was conducted using an air background. During measurement, the infrared beam was accurately focused on the sample. Spectra were collected over a range of 4000–400 cm^−1^ with a resolution of 4 cm^−1^, and 32 cumulative scans were averaged to obtain high-quality infrared spectra. Alterations in functional groups on the PE molecular chains were identified by analyzing shifts in characteristic absorption peaks.

The surface contact angle of PE samples was measured following a standardized protocol. The procedure was carried out as follows: each PE sample was securely mounted on the stage of a contact angle analyzer. By adjusting the stage height and optical focus, a clear image of the sample surface was obtained on the monitor. A micro-dispenser (accuracy ± 0.1 μL) was used to deposit a droplet of deionized water (resistivity ≥ 18.2 MΩ·cm) with a volume of 3.0 ± 0.5 μL onto the sample surface. Droplet shape was recorded in real time using a high-speed camera system (frame rate 1000 fps, resolution 1920 × 1080). To ensure reliability, three random locations on each sample were selected, and each location was measured three times independently. The average of all measurements was reported as the final contact angle value.

2.6. Statistical Analysis

All experiments were performed in triplicate, and the results are presented as the mean ± standard deviation (SD). Statistical analysis was performed using SPSS 26.0. To compare the differences between the experimental and control groups, specifically the weight loss rate of the PE film and the contact angle measurements, an unpaired two-tailed Student’s t-test was employed.

3. Results and Discussion

3.1. Isolation and Purification of Degrading Bacteria

By screening for PE-degrading bacteria in soils, sewage, and sludge from plastic manufacturing facilities using PE as the sole carbon source, 49 single colonies capable of growing on MSM agar plates supplemented with PE were obtained. Analysis of 16S rRNA gene sequences from the isolated strains (32 species in total) and comparison with closely related organisms revealed that six strains—SG01, SG15, SG19, SG25, SG27, and SG29—showed a 1.8–2.5-fold increase in OD_600_ compared to the initial values after 50 days of culture in liquid MSM containing PE. Phylogenetic analysis demonstrated that SG01, SG15, SG19, SG25, SG27, and SG29 formed separate branches with Pseudomonas aeruginosa, Serratia marcescens, Pseudomonas monteilii, Enterobacter kobei, Stenotrophomonas maltophilia, and Micrococcus luteus, respectively.

3.2. Growth Curves of Degrading Bacteria

The growth kinetic profiles of microorganisms in inorganic salt medium (MSM) with PE film as the sole carbon source exhibited distinct differences. P. aeruginosa displayed a typical sigmoidal growth curve, characterized by a lag phase of about 3 days, a rapid logarithmic growth phase (3–15 days), and entry into stationary phase at 27 days (OD_600_ = 1.15 ± 0.05), as shown in Figure 1B. In contrast, S. maltophilia showed a biphasic growth pattern, with peaks at 12 and 24 days (OD_600_ = 0.60 and 0.72), suggesting activation of secondary metabolic pathways. While both S. marcescens and P. monteilii exhibited single-peak growth curves, S. marcescens displayed clear growth inhibition after 18 days (OD_600_ decreased by 12.5%), possibly due to accumulation of metabolic by-products. Notably, M. luteus and E. kobei sustained only low-level growth throughout the incubation (OD_600_ < 0.30), indicating limited metabolic capacity for PE. P. aeruginosa demonstrated a notable growth advantage, with a maximum specific growth rate 2–3 times higher than other strains and the highest biomass accumulation (OD_600_ = 1.15 ± 0.05) during the culture period, likely attributable to its unique rhamnolipid secretion system and P450 oxygenase activity. The growth curves of all experimental groups were significantly higher than those of the control group, confirming the PE-degrading potential of the tested strains.

To systematically evaluate the biodegradation efficiency of P. aeruginosa SG01 on PE, three PE-dose experimental groups (0.5, 1.0, and 2.0 g) were established, along with two control groups: PE-positive/strain-negative (Control 1) and PE-negative/strain-positive (Control 2). Results showed that the 1 g PE group displayed optimal growth kinetics, reaching peak biomass (OD_600_ = 0.800 ± 0.05) at 27 days of cultivation (Figure 1C). This value was significantly higher than those of the 0.5 g group (0.550 ± 0.04) and the 2 g group (0.600 ± 0.05) (p < 0.05), confirming that the most favorable metabolic balance between PE degradation and bacterial growth occurred at this concentration. Notably, the 2 g PE group exhibited marked growth retardation during the early cultivation period (0–9 days), with biomass 46.7% lower than that of the 1 g group, likely due to substrate mass-transfer limitations and hydrophobic surface effects caused by excessive PE loading.

3.3. Degradation Efficiency of Degrading Bacteria

3.3.1. Effect of Degrading Bacteria on Weight Loss Rate of PE Film

Strain SG01 obtained through screening exhibited significant degradation capacity for PE. Experimental data showed that under standard culture conditions, the weight loss rate of PE film (1 g) was the highest and continuously increased with prolonged culture time: 15 d (5.62 ± 0.23%), 30 d (9.33 ± 0.31%), 45 d (12.45 ± 0.28%), and 60 d (14.25 ± 0.35%), all of which were significantly higher than the control group (0.30 ± 0.05%) (p < 0.001). The influence of different PE concentrations on degradation efficiency is presented in Figure 2A. These results confirm that strain SG01 can effectively degrade PE materials, and the degradation efficiency is positively correlated with time (r = 0.982, p < 0.001).

3.3.2. Analysis of Surface Morphology Characteristics

Ultra-structural observation and analysis of the enriched PE plastic film were conducted using a SEM. The results showed that the control group’s PE film, which was not inoculated with degrading bacteria and statically cultured for 60 d, exhibited a smooth and dense physical morphology, retaining the regular texture structure formed by the production process, but no obvious cracks or perforations were observed.

After 15 days of enriched culture of the degrading bacterial community in a selective medium with PE film as the sole carbon source, the original production textural features were significantly weakened, and characteristic elliptical void structures began to appear on the surface, with irregular erosion traces at the edges. At 30 days of culture, the void structures on the PE film surface underwent obvious morphological evolution: the major axis diameter of the elliptical voids expanded significantly, the boundary contours became clear and sharp, and the color depth was enhanced due to increased electron density in the eroded areas. After 45 days of enriched culture, the void structures exhibited hierarchical degradation characteristics: the area of main voids further expanded, secondary erosion pits were visible at the edges, and partially unpenetrated thin film structures formed in some regions. Quantitative analysis of the temporal sequence changes in void area confirmed that the erosion process of PE film by degrading bacteria followed a bimodal mechanism, achieving material degradation through the synergistic radial expansion of voids and progressive thinning in the film thickness direction. At 60 days of culture, the PE film surface presented a porous network structure, with the void density significantly increasing compared to the 45-day group, and adjacent voids gradually merging to form composite erosion regions (Figure 2C). Comprehensive analysis of the spatiotemporal dynamic evolution characteristics observed by SEM showed that with the extension of enriched culture time, the biological erosion of PE film by the degrading bacterial community was characterized by an exponential increase in the number of voids, revealing the temporal correlation between microbial metabolic activities and the microstructural damage of plastic materials.

3.3.3. Evolution of Functional Groups on PE Film Surface During Microbial Degradation

FTIR analysis revealed significant changes in functional group characteristic peaks of PE films during degradation (0, 15, 30, 45, and 60 d), as presented in Figure 2B. Major absorption peaks were observed at 2914.03 cm^−1^ (asymmetric C–H stretching), 2846.53 cm^−1^ (symmetric C–H stretching), 2362.46 cm^−1^, 1641.83 cm^−1^ (C=O stretching), and 719.34 cm^−1^ (long-chain alkane –CH_2_ bending). Notably, the enhancement of the C=O peak around 1641.83 cm^−1^ indicated oxidative modification of PE during degradation, likely linked to oxygen incorporation through microbial metabolism. Furthermore, new absorption bands emerged in the 1650–2000 cm^−1^ range, attributable to the formation of amino (–NH_2_) or hydroxyl (–OH) moieties, suggesting potential substitution of carbonyl groups (C=O) in PE chains by amino groups. The appearance of amino groups suggests that the nitrogen source was likely derived from the culture medium, which contained ammonium sulfate ((NH_4_)2_SO_4, 500 mg/L) as a nitrogen source (see Section 2.2). We propose two possible mechanisms for the incorporation of these nitrogen-containing groups onto the PE film surface: first, residual biofilm components or microbial secretions containing nitrogen may remain adhered to the PE surface despite the washing procedure; and second, reactive intermediates generated during PE oxidation may react with ammonium ions (NH_4_^+^) in the medium, leading to the formation of amide or amine groups at the chain ends. Therefore, the emergence of these nitrogenous groups suggests a complex interplay between microbial metabolism and the polymer surface. These findings not only confirm the metabolic capacity of the degrading bacteria but also demonstrate that PE undergoes substantial chemical restructuring under microbial activity. Given that pure PE contains only carbon and hydrogen, the emergence of these oxidized products provides further evidence that the strain facilitates PE degradation via an oxidative pathway.

Multiple related studies have similarly demonstrated the formation of oxidative functional groups on the surface of polyolefin films following bacterial degradation. Yang et al. [27] used FTIR analysis to show that the carbonyl index (C=O) of PE films degraded by intestinal bacteria of Galleria mellonella increased significantly by 2.8-fold. Similarly, Bombelli et al. [21] employed gravimetric measurements to determine that the larval gut microbiota of G. mellonella degraded 13 ± 2% of PE films within 12 h (p < 0.05); FTIR spectroscopy further revealed a 3-fold increase in the intensity of the carbonyl absorption peak at 1712 cm^−1^, confirming the microbially mediated oxidative degradation mechanism.

3.3.4. Changes in Surface Hydrophilicity of PE Films

Water contact angle measurements were performed to evaluate changes in the surface hydrophilicity of PE films after 0, 15, 30, 45, and 60 days of incubation with P. aeruginosa SG01. As shown in Figure 2D, the contact angle decreased progressively with increasing incubation time. The initial contact angle of the untreated PE film was approximately 97.8° ± 0.85°, indicating a highly hydrophobic surface. After 15 days of incubation, the contact angle decreased to 90.6° ± 0.57°, and further declined to 85.1° ± 0.71° at 30 days. Following 45 and 60 days of degradation, the contact angles reached 81.65° ± 0.49° and 76.95° ± 0.21°, respectively. This significant reduction (p < 0.01) demonstrates that P. aeruginosa SG01 effectively enhances the surface wettability of PE films.

The increased hydrophilicity is consistent with the introduction of oxygen-containing functional groups detected by FTIR analysis. The improved surface wettability facilitates further microbial attachment and colonization, creating a positive feedback loop that enhances subsequent degradation. These results are in agreement with previous studies showing that microbial degradation of polyolefins is accompanied by increased surface hydrophilicity due to the formation of polar groups [28,29,30].

Pseudomonas aeruginosa’s ability to degrade PE likely stems from its distinctive metabolic features and enzymatic systems. Research indicates that this bacterium secretes extracellular enzymes, including alkane hydroxylase (AlkB) and P450 monooxygenase, which catalyze the oxidative cleavage of polyolefins such as PE, yielding intermediates with carbonyl (C=O) and hydroxyl (−OH) functional groups [28]. Moreover, biofilms formed by P. aeruginosa enhance bacterial adhesion to plastic surfaces, while its biosurfactants (e.g., rhamnolipids) improve the hydrophilicity of hydrophobic plastics [29,30], thereby jointly facilitating PE breakdown. These synergistic mechanisms establish P. aeruginosa as a promising candidate for microplastic bioremediation.

4. Conclusions

This study successfully isolated and identified Pseudomonas aeruginosa SG01, which exhibited significant PE degradation potential. After 60 days of incubation, strain SG01 achieved a weight loss rate of 14.25 ± 0.35% for PE films. Its degradation mechanism involves both physical erosion and chemical oxidation. Multi-scale characterization techniques confirmed that strain SG01 effectively alters the surface properties and chemical structure of PE materials: scanning electron microscopy observed obvious surface erosion morphology, contact angle measurements showed a significant decrease in hydrophobicity (p < 0.01), and FTIR detected enhanced characteristic absorption peaks of oxygen-containing functional groups such as carbonyl (C=O) and hydroxyl (-OH). These findings are consistent with recently reported microbial-mediated polymer degradation mechanisms, indicating that SG01 may achieve the cleavage of PE long-chains through similar oxidative metabolic pathways.

The degradation efficiency of SG01 is comparable to that of previously reported PE-degrading microorganisms. Bombelli et al. [21] reported that the larval gut microbiota of Galleria mellonella degraded 13 ± 2% of PE films, while Yang et al. [27] demonstrated a 2.8-fold increase in carbonyl index after bacterial degradation. The performance of SG01 is noteworthy, given that degradation was achieved using PE as the sole carbon source without additional surfactants or physical pretreatments.

The industrial environmental origin of SG01 provides inherent adaptive advantages for its application in wastewater treatment systems. Future research should prioritize several key directions: optimizing culture parameters (e.g., C/N ratio, temperature, and pH) using statistical approaches such as response surface methodology; establishing degradation kinetic models; elucidating the key enzyme systems and metabolic pathways involved through metagenomic and metabolomic analyses; and evaluating degradation performance in real wastewater matrices, with particular attention to the influence of co-existing pollutants. Furthermore, detailed characterization and resource-oriented utilization of the degradation products will be crucial for enhancing the economic feasibility of this technology.

This study expands the microbial resource library for PE degradation and provides new experimental evidence for understanding the interaction mechanisms at the bacteria–plastic interface. The ability of the strain to significantly alter PE surface properties in the absence of added surfactants suggests the potential production of specific biosurfactants. These findings establish a theoretical foundation for developing plastic pollution control strategies based on synthetic microbial consortia. As research advances, this approach is expected to serve as an important complement to conventional physicochemical treatment methods, offering novel biological pathways toward achieving a circular economy for plastics.