Microbial Inoculant‐Driven Degradation of Microplastics Associated With Garbage Environment and Assessment of Degradation Product Toxicity

ABSTRACT

This investigation revealed that the application of specific microbial inoculants could facilitate the effective biodegradation of polyethylene terephthalate (PET) microplastics collected from urban garbage sites, resulting in non‐toxic end products. We employed Azotobacter chroococcum (MTCC 3853), Rhizobium leguminosarum (MTCC 9766), Azospirillum brasilense (MTCC 4036), and Trichoderma viride (MTCC 9681) for PET microplastics degradation and assessed their degradation efficacy through a series of controlled in vitro batch experiments. The study encompassed quantitative analysis of PET weight loss, detailed chemical profiling of degradation intermediates and products, biofilm formation assessment, microbial growth monitoring, and measurement of plastic‐degrading enzyme induction. To comprehensively evaluate environmental safety, phytotoxicity assays were performed on Vigna mungo and Vigna radiata, while zebrafish embryos and adults were subjected to acute and embryonic toxicity tests. A. chroococcum (MTCC 3853) was identified as the most efficient strain, showing the greatest reduction in PET mass, enhanced biofilm formation, sustained microbial growth, and peak enzymatic activity, with no detrimental effects on plant or aquatic models, confirming the safety of the biodegradation process. These results underscore the potential of A. chroococcum (MTCC 3853) as a powerful and environmentally friendly solution for microplastic remediation in urban environments.

Microplastics degradation by microbial inoculants and screening of biocompatibility and eco‐toxicity.

1. Introduction

Within the evolving landscape of anthropogenic environmental threats, microplastics—defined as plastic particles < 5 mm—have emerged as both a scientific and policy conundrum. Their diminutive scale facilitates pervasive infiltration into ecological niches previously considered protected from plastic contamination, and complicates efforts to quantify, monitor, and mitigate their impact. Their environmental discharge releases hazardous chemicals, such as phthalates, polybrominated diphenyl ethers, and tetrabromobisphenol A, thereby increasing risks to the ecosystem and human health (Avinash et al. 2023). The incineration of plastics further intensifies global warming and disseminates organic pollutants, impacting aquatic species and humans alike. Microplastics are produced by the breakdown of larger plastics or through the manufacturing of microbeads, and appear as fibers, domains, or particles (Hale et al. 2020).

Long‐term environmental exposure to microplastics has been linked to toxicological effects in a range of organisms and at multiple ecological scales. According to Campanale et al. (2020), less than 0.3% of particles (< 150 μm) and 0.1% of those particles (> 10 μm) can traverse gastrointestinal, placental, and neural barriers (Barboza et al. Barboza, Vieira, Branco, Carvalho & Guilhermino 2018). These pollutants have been detected in aquatic, terrestrial, and airborne environments (Li et al. 2023). In soil−plant systems, microplastics disrupt seedling emergence, root extension, and resource absorption, and act as carriers for toxicants that induce cellular and molecular harm (Qi et al. 2018; de Souza Machado et al. 2018; Mattsson et al. 2017).

Mitigation of microplastic contamination utilizes physical (adsorption, filtration) (Mohammad R and Qusay 2023), chemical (hydrolysis, alcoholysis, acidolysis, glycolysis, aminolysis) (Bhuvaneswari 2018), electrochemical, and photocatalytic strategies (Namasivayam and Avinash 2024). Microbial biodegradation is increasingly recognized for its efficiency and environmental compatibility. The effectiveness of biological degradation is influenced by substrate characteristics, surface topology, morphology, and molecular mass (Albertsson et al. 1987; Ammala et al. 2011; Zeenat et al. 2021; Dawson et al. 2018). Polyethylene degradation has been attributed to a variety of microbes, including Achromobacter, Acinetobacter, Arthrobacter, Aspergillus, Bacillus, Comamonas, Delftia, Micrococcus, Nesiotobacter, Paenibacillus, Pseudomonas, Rahnella, Staphylococcus, and Stenotrophomonas (Sangeetha Devi et al. 2019; Lu et al. 2022; Zhai et al. 2023).

Indicators of biodegradation include decreased mass, chemical transformation, and CO₂ evolution (Chaudhary and Vijayakumar 2020; Mohan et al. 2016). The engineering of plastic‐degrading microorganisms and enzymes is enabling the conversion of plastic residues into carbon‐rich or alkane products, facilitating recycling and reducing pollution (Liu et al. 2023; Paço et al. 2017; Kunlere et al. 2019; Mohanan et al. 2020; O'Toole et al. 2000; L. Wang et al. 2024). Recent studies demonstrate that Azotobacter vinelandii with biochar improves nitrogen retention on PBAT plastics, while Azospirillum brasilense can reduce indole‐3‐acetic acid by 50%–66% within 10 days, depending on plastic characteristics. Despite the growing body of research, there are limited comprehensive studies on the effects of microplastics within biotic and abiotic systems. Qi et al. (2018) found that LDPE and PET did not impact the growth of Triticum aestivum, while pullulan reduced it. Zhao et al. (2023) observed that cadmium in combination with HDPE and PS impeded maize growth and increased Cd accumulation.

This investigation focuses on the in vitro degradation of microplastics by microbial inoculants (Azotobacter chroococcum [MTCC 3853], Rhizobium leguminosarum [MTCC 9766], Azospirillum brasilense [MTCC 4036], Trichoderma viride [MTCC 9681]). Evaluations included weight reduction, biofilm production, enzyme activities (PEG Dehydrogenase, Esterase, Lipase, Cutinase), and chemical changes (Fourier‐transform infrared [FTIR], GCMS). The toxicity and compatibility of crude metabolites were further tested in Vigna radiata, Vigna mungo, and Danio rerio, with molecular docking of PET metabolites in V. radiata and Homo sapiens.

Rather than focusing on organisms engineered for plastic degradation, this study scrutinizes commonly used agricultural bioinoculants—A. chroococcum (MTCC 3853), R. leguminosarum (MTCC 9766), A. brasilense (MTCC 4036), and T. viride (MTCC 9681)—for their possible role in PET breakdown. These microbes have been selected for their remarkable metabolic flexibility, including the production of extracellular esterases and lipases essential for cleaving PET's ester linkages. The investigation advances a dual‐benefit framework: alongside their established plant growth‐promoting effects, such as nitrogen‐fixation and phytohormone synthesis, their potential in microplastic remediation is systematically evaluated. Ultimately, the research aims to determine if these accessible, single‐strain inoculants can be repositioned as central tools in both environmental and agricultural spheres.

Through a multi‐pronged experimental protocol, the research quantifies not only the rate and extent of microplastic mass loss but also systematically evaluates biofilm formation capacity and the catalytic spectrum of PEG dehydrogenase, esterase, lipase, and cutinase. Molecular transformations are interrogated through advanced FTIR spectroscopic mapping and GC‐MS profiling, revealing both qualitative and quantitative shifts in polymer structure and degradation byproducts. The environmental and biological safety profiles of crude microbial metabolites are stringently assessed through ecotoxicological assays performed on V. radiata, V. mungo, and D. rerio, ensuring the environmental compatibility of bioremediation products. Furthermore, leveraging in silico molecular docking, the research elucidates the mechanistic underpinnings of PET‐degraded metabolite interaction with receptor proteins in both plant (V. radiata) and human (H. sapiens) systems. Collectively, this integrative approach—which synergizes high‐resolution analytical chemistry, robust toxicological screening, and predictive computational modeling, establishes a paradigmatic shift—enabling a comprehensive, multidimensional appraisal of the ecological and biological implications attendant to microplastic degradation.

2. Methodology

2.1. Sample Collection and Processing

The challenge of PET microplastic persistence in the environment drove the meticulous design of this study's materials and methods. Sampling commenced with the targeted collection of PET bottles from waste management zones in Pallikaranai, Chennai, with inclusion criteria mandating a uniform wall thickness of 1 mm. The rationale for PET selection was its chemical recalcitrance and overwhelming presence in polluted environments, which render it a high‐priority target for biodegradation research. At the outset, the PET bottles were cut into five fragments, each measuring precisely 1 cm². These fragments were desiccated in a controlled drying chamber until their mass stabilized, after which their baseline weights were determined using a high‐precision analytical balance with a sensitivity of 0.1 mg. The sterilization sequence entailed immersion in a 0.1% HgCl₂ solution, followed by exhaustive rinsing in sterile distilled water to mitigate any risk of residual sterilant interfering with subsequent microbial assays. Following chemical sterilization, UV irradiation at 254 nm was applied for 30 min while the samples were positioned in a laminar airflow cabinet, in accordance with the standardized protocol of Mohan et al. (2016). To rigorously isolate the specific impact of microbial enzymatic activity from potential abiotic modifications, a parallel control group was established; these PET samples were subjected to identical UV treatment without microbial inoculation. Empirical observations confirmed the validity of the control: weight loss was restricted to less than 0.05%, and FTIR spectra indicated an absence of significant functional group transitions. These findings corroborated that UV exposure primarily conferred sterilization and a modest increment in hydrophilicity, rather than instigating substantive PET degradation. The implementation of this robust, multi‐step sterilization and preparation protocol resulted in PET microplastics that were both microbiologically inert and physically conditioned for enhanced microbial colonization. The hydrophilicity enhancement imparted by UV irradiation is recognized to facilitate initial microbial binding and subsequent biofilm development, prerequisites for effective PET biodegradation. This methodical approach ensured that any observed experimental changes in PET mass or structure could be attributed with confidence to microbial degradation mechanisms, rather than to procedural artifacts or uncontrolled abiotic influences.

2.2. Selection of Microbial Bioinoculants

Microbial strains used in this study, namely A. chroococcum (MTCC 3853), R. leguminosarum (MTCC 9766), A. brasilense (MTCC 4036), and T. viride (MTCC 9681), were all acquired from the Microbial Type Culture Collection (MTCC) Gene Bank, Chandigarh, India, reflecting a commitment to scientific rigor and reproducibility. The preservation of each strain was achieved through regular sub‐culturing on the specified media, with all relevant compositions and handling details provided in Table 1, eliminating a significant source of experimental variability.

Table 1.

Description of selected bioinoculants.

S. no.	Microbial strain	Growth medium	Preservation suspending medium	Optimum condition
1	Rhizobium leguminosarum (MTCC 9766)	Yeast Mannitol Agar (Mannitol—10 g, KH2PO4—0.5 g, MgSO4.7H2O—0.2 g, NaCl—0.1 g, Yeast extract—0.4 g, CaCO3—4 g, Agar—15 g, Distilled water—1 L)	10% Glycerol	Aerobic (Incubation: 48 h, Temperature: 30°C)
2	Azotobacter chroococcum (MTCC 3853)	Azotobacter Medium (K2HPO4—1 g, MgSO4.7H2O—0.2 g, NaCl—0.2 g, FeSO4—5 mg, Soil extract—100 mL, Agar—15 g, Distilled water—900 mL, Mannitol—2%, Soil—77 g, Na2CO3—0.2 g, Distilled water—200 mL)	10% Glycerol	Aerobic (Incubation: 24 h, Temperature: 30°C)
3	Azospirillum brasilense (MTCC 4036)	LB Medium (Tryptone—10 g, Yeast extract—5 g, NaCl—10 g, Distilled water—1 L)	10% Glycerol	Aerobic (Incubation: 48 h, Temperature: 30°C)
4	Trichoderma viride (MTCC 9681)	PDA Agar (Potato—200 g, Dextrose—20 g, Agar—15 g, Distilled water—1 L)	10% Glycerol	Aerobic (Incubation: 1 w, Temperature: 25°C)

2.3. Microplastics Degradation Studies

2.3.1. Shake Flask Assay

Sterilized polyethylene terephthalate microplastics (PET‐MPs) were combined with 250 mL MSM broth (K₂HPO₄—1.73 g/L, KH₂PO₄—0.68 g/L, MgSO₄·7H₂O—0.1 g/L, FeSO₄·7H₂O—0.03 g/L, NH₄NO₃—1 g/L, CaCl₂·2H₂O—0.02 g/L) in 500 mL Erlenmeyer flasks and inoculated with 1 mL of activated microbial suspension. Both controls (without PET‐MPs) and experimental groups (with PET‐MPs) were established in triplicate and incubated at 37°C under continuous shaking for 28 days. At selected intervals up to 240 h, samples were taken to monitor microbial growth curves at 600 nm. Following incubation, the cultures were centrifuged at 8000 rpm for 10 min, and the supernatants were reserved for downstream analyses, including gravimetric loss, biofilm assessment, identification of degradation products, enzyme activity measurement, biocompatibility studies, in silico docking procedures, and two‐way ANOVA for statistical evaluation.

2.3.2. Percentage of Weight Loss Calculation

Quantification of microplastic degradation was operationalized using a sequential gravimetric protocol, initiated by retrieving PET‐MP samples after a predetermined 28‐day incubation under controlled conditions. The subsequent removal of adherent microbial matrices was achieved by immersing in a 2% sodium dodecyl sulfate solution to maximize biofilm disruption without compromising polymer integrity. Post‐cleansing, samples were exposed to a desiccator to facilitate moisture equilibration, with mass readings acquired at designated intervals until stability was documented. Weight loss fractions were then extracted by applying the analytical frameworks delineated by Mohan et al. (2016) and Liu et al. (2023):

$$\%\text{weight loss}=\frac{\text{Initial Weight – Final Weight}}{\text{Initial Weight}}\times 100$$

A precision balance with 0.1 mg sensitivity underpinned all gravimetric assessments, thereby safeguarding the reliability of degradation measurements.

2.3.3. Determination of Biofilm Colonization

To quantify biofilm development, a 96‐well plate assay with crystal violet and spectrophotometric analysis, adapted from O'Toole et al. (2000), was employed. A 100 µL portion of inoculant was added to each well, followed by 72 h incubation at 37°C. After standard processing, absorbance at 520 nm was recorded. Biofilm quantification on microplastic surfaces involved placement in petri dishes, inoculation with 100 µL of bioinoculant, and execution of all subsequent detection and measurement steps as established in the microtiter protocol, with the absorbance measured at 570 nm. Biofilm production was compared based on the difference in OD between microtiter plate (A₁) and petri dish (A₂) methods.

$$\text{Biofilm Formation Efficacy}(\%)=\frac{{\text{OD}}_{A_1}\text{–}{\text{OD}}_{A_2}}{{\text{OD}}_{A_1}}\times 100$$

${\text{OD}}_{A_1}$ is the optical density of ethanol‐solubilized biofilm of PET‐MP coated with bioinoculants derived from a microtiter plate. ${\text{OD}}_{A_2}$ is the Optical Density of ethanol‐solubilized biofilm of PET‐MP coated with bioinoculants derived from a petri plate

The examination of biofilm architecture on PET‐MP surfaces was structured around a multi‐stage analytical regimen. Initial detection and semi‐quantitative evaluation of biofilm biomass were accomplished via the crystal violet assay. Following this, microplastic samples were anchored onto brass stubs through the application of carbon adhesive tape to prevent movement during further processing. A uniform, 60‐s platinum sputter coating was then administered, optimizing the specimens for subsequent high‐fidelity scanning electron microscopy (SEM). Surface morphological features were investigated using the JEOL JSM‐IT800 Nano SEM, with fields of view deliberately chosen to capture intra‐sample heterogeneity. Analysis was repeated across 3 independent biological replicates to afford robust comparative insights. Each electron micrograph was annotated using calibrated scale bars, thereby facilitating rigorous quantification of biofilm spatial organization and complexity.

2.3.4. Screening of Biodegraded Products

2.3.4.1. FTIR Analysis

After experimental degradation, the structural and compositional evolution of PET‐MP was rigorously evaluated through FTIR spectroscopy, utilizing KBr as the pelletizing medium to ensure optimal spectral clarity. Data collection spanned the 400–4000 cm⁻¹ region, with each spectrum derived from the averaged response of 16 scans at an 8 cm⁻¹ resolution. Advanced computational analysis, executed via Thermo Scientific OMNIC Paradigm Desktop Software, enabled the resolution of overlapping vibrational modes and the systematic identification of emergent degradation signatures. For quantitative assessment of oxidative transformation, the Carbonyl Index was computed as the ratio of the absorbance at the carbonyl stretching frequency (approximately 1715 cm⁻¹) to the absorbance at the C─H stretching frequency (approximately 2960 cm⁻¹). This index provides the foundation for rigorous statistical interrogation of polymer oxidation, facilitating direct comparison across heterogeneous microbial consortia and enabling mechanistic insights into degradation pathways.

2.3.4.2. GCMS Analysis

A comprehensive profiling of degradation end products in the supernatant was undertaken using Gas Chromatography–Mass Spectrometry (GCMS). Experiments were performed on a SHIMADZU GC‐MS QP 2010 plus instrument, featuring a DB‐5MS capillary column of 30 m length, 0.25 mm diameter, and 0.25 μm film thickness. The separation protocol relied on a temperature program increasing from 45°C to 300°C. The mobile phase consisted of helium gas with a purity of 99.9999%. System stability was maintained by keeping the injection port and mass analyzer at 280°C throughout the analysis.

2.4. Evaluation of PET‐MPs Degrading Enzymes

2.4.1. Qualitative Determination

PET microplastic degradation by enzymatic means was assessed qualitatively through the use of specifically prepared agar substrates: Polyethylene Glycol Dehydrogenase was assayed with nutrient agar supplemented by 2% polyethylene glycol and Coomassie Brilliant Blue, Esterase with 1% nutrient agar and 1% Tween‐80 plus Phenol Red, Lipase with tributyrin agar made from 5 g/L peptone, 3 g/L yeast extract, 12 g/L agar, and 10 g neutral tributylene, and Cutinase with 0.4% tomato cutin in 100 mL sterile nutrient agar. With solidification complete, wells were aseptically introduced and inoculated with 100 μL of enzyme solution. The plates were then incubated at 37°C for 48–96 h, and hydrolysis was quantified by measuring the diameter of the clear zones surrounding each well.

2.4.2. Quantitative Determination

For the quantitative analysis of PET‐MPs degrading enzymes, 100 mL of substrate‐supplemented nutrient broth was inoculated with 1 mL microbial culture, incubated under agitation for 48–96 h, centrifuged at 8000 rpm for 15 min to obtain supernatants for use in the enzyme assay, and subsequently analyzed by UV‐Vis spectrophotometry across 400–899 nm, assessing PEG Dehydrogenase at 580 nm and Esterase, Lipase, and Cutinase at 610 nm.

2.5. Non‐Target Effects of PET‐MP Degraded Metabolites

2.5.1. Phytotoxicity Assessment

Following the acquisition of healthy seeds of black gram (V. radiata) and green gram (V. mungo) from an organic supplier in Neelangarai, ECR, Chennai, these materials were brought to the laboratory to enable the commencement of subsequent experimental procedures.

2.5.2. Treatment

According to protocols modified from Sahasa et al. (2023) and Avinash et al. (2024), Petri plates autoclaved at 121°C for 15 min were placed in a Laminar Air Flow Chamber, with healthy seeds being exposed to a 10‐minute immersion in either microbial supernatant or distilled water, transferred to the plates by forceps, and incubated at ambient temperature for 7 days after application of distilled water; post‐incubation, both seedling length and seed morphology were measured for further analysis.

2.5.3. Germination Index

This investigation systematically explored germinative responses across a spectrum of PET‐MP‐derived metabolite concentrations, deploying degraded supernatant at 25%, 50%, 75%, and 100% (v/v) alongside a distilled water control in an effort to construct a comprehensive dose‐response profile. Seeds were subjected to continuous germination monitoring, generating a temporal dataset from which the germination index (%) was meticulously derived for each treatment arm. This quantitative endpoint served as a robust indicator for comparing the biological impact of each graded exposure, allowing the research to delineate finely resolved, concentration‐dependent effects.

$$\text{Germination Index}(\%)=\frac{\text{No. of seeds in control}}{\text{No. of seeds in treatment}}\times 100$$

The careful calibration of this experimental array ensured that all conclusions regarding the safety or potential adverse outcomes of PET‐MP‐derived metabolites were rigorously circumscribed within the tested interval of 25%–100% (v/v), thus upholding methodological integrity and preventing unsupported extrapolation beyond the empirical evidence established by the experimental design.

2.5.4. Proximate Composition

Utilizing the procedural benchmarks set forth by AOAC Kavanagh (1981), both control and PET‐MP metabolite‐exposed seedlings were analyzed for proximate composition, in which key constituents such as moisture, ash, protein, carbohydrate, fiber, and fat were quantitatively analyzed.

2.6. Evaluation of Embryonic and Acute Toxicity in Zebrafish

2.6.1. Collection and Maintenance

To study the biological impacts of PET microplastic degradation products, adult zebrafish were obtained from a regional aquarium outlet and introduced into 35 L glass aquaria containing dechlorinated, aerated water, with aeration provided by stone diffusers and a mechanical compressor. The environmental temperature was precisely maintained at 26°C ± 1°C. Commercial feed was administered twice to the fish during the acclimatization phase. All procedures involving animal care and maintenance were conducted in strict accordance with laboratory animal welfare guidelines.

2.6.2. Zebrafish Embryonic Toxicity

2.6.2.1. Breeding Setup

To systematically study the embryonic toxicity of PET‐MP metabolites, D. rerio adults (3.05 ± 0.04 cm, 1.00 ± 0.5 g) were kept in gender‐specific aquaria for 24 h, then combined at a 2:1 male‐to‐female proportion in breeding tanks lined with nylon mesh to intercept eggs, and after a 16 h dark period, eggs were harvested and immersed in E3 medium (5 mM sodium chloride, 0.17 mM potassium chloride, 0.33 mM calcium chloride, 0.33 mM magnesium chloride, pH set to 7.2–7.3) in Petri plates, with unfertilized eggs subsequently removed via microscopic screening.

2.6.2.2. Developmental Stages

To systematically investigate non‐targeted toxicological consequences associated with PET‐MP breakdown, zebrafish embryos were exposed to a graded series of supernatant dilutions—specifically 25%, 50%, 75%, and 100% v/v. The experimental methodology prioritized simultaneous monitoring of multiple developmental benchmarks: survival and mortality rates were recorded at regular intervals, enabling dynamic assessment of embryonic viability; the onset of cardiac activity was pinpointed through precise temporal tracking; intra‐chorionic movements were observed in real time to identify early neuromuscular anomalies; and blood circulation patterns were evaluated during pivotal stages of embryogenesis.

2.6.2.2.1. Survival Rate

The equation below served as the basis for calculating the percentage of surviving zebrafish embryos after PET‐MP degraded metabolite exposure:

$$\text{Survival Rate}(\%)=\frac{\text{No. of live embryos}}{\text{Total no. of embryos}}\times 100$$

2.6.2.2.2. Heartbeat Count

For both PET‐MP metabolite‐exposed and control zebrafish fry groups, heart rate data were manually collected by observing each fry under a phase contrast microscope at 48–96 h after hatching, ensuring precise interval‐based measurement.

2.6.2.3.

2.6.2.3.1. Hatching Rate

For each temporal evaluation, the hatching rate of zebrafish embryos exposed to PET‐MP degraded metabolites was derived using the mathematical approach indicated below:

$$\text{Hatching Rate}(\%)=\frac{\text{No. of hatched embryos}}{\text{Total no. of embryos}}\times 100$$

2.6.3. Zebrafish Acute Toxicity

2.6.3.1. Experimental Model

A cohort of 12 adult zebrafish of both sexes and comparable weight was assigned to four separate groups to study the acute toxicity of PET microplastic degradation products. For the acute toxicity investigation of PET microplastic degradation products in adult zebrafish, 12 fish of both sexes and similar weight were distributed into four groups per experimental run. The first group served as an untreated reference, while the experimental groups were exposed to specific concentrations (0.25%, 0.50%, and 0.75% v/v) of the degraded product to establish a dose‐response profile. On the 8th day, euthanasia was performed for biochemical and histopathological evaluation. The subsequent safety narrative is bounded strictly by these tested concentrations. Upon completion of the 8‐day treatment, euthanasia was performed to facilitate blood collection for biochemical markers—AST, ALT, ALP, glucose, total cholesterol, triglycerides, urea, and creatinine—and to allow for the removal and formalin fixation of liver and kidney tissues for histopathological analysis. The safety analysis is restricted to the concentrations used during the study.

2.6.3.2. Estimation of Biochemical Blood Parameters

Assessment of blood biochemical indices in zebrafish under acute toxicity conditions relied on the use of Biosystems Diagnostic biochemistry kits, specifically catalog numbers 11830 (Aspartate Aminotransferase), 11832 (Alanine Aminotransferase), 11592 (Alkaline Phosphatase), 11803 (glucose), 11805 (total cholesterol), 11828 (triglycerides), 11536 (blood urea), and 11802 (creatinine). The analytical methodology for each marker was executed exactly in line with the manufacturer's prescribed protocols.

2.7. Docking Studies

Addressing the intricate issue of non‐target biological interactions posed by metabolites originating from polyethylene terephthalate‐microplastic (PET‐MP) degradation, this study constructed a rigorously detailed molecular docking framework. The investigation commenced with the systematic extraction of atomic‐resolution structures for two key proteins: the membrane aquaporin from V. radiata (PDB: 6AFS) and the cytosolic regulatory protein from H. sapiens (PDB: 6JFV), each sourced from the Protein Data Bank. Employing only models resolved at sub‐2.0 Å, the protocol was designed to maximize structural fidelity and provide a robust basis for subsequent computational docking and interaction profiling. A strategic, principle‐driven process guided the selection of protein receptors for docking. Central to this approach was the prioritization of proteins with established significance in cellular homeostasis and documented vulnerability to xenobiotic interference. In the botanical context, the V. radiata aquaporin (PDB: 6AFS) was chosen due to its essential role in sustaining water equilibrium and facilitating nutrient transport across the plasma membrane. Its identity as PIP2;1 underscores its criticality for plant osmoregulation, and the prospect of its disruption by PET‐MP‐derived metabolites portends adverse consequences such as reduced seedling vigor and physiological stress. Complementarily, the human cytosolic protein (PDB: 6JFV) was incorporated to serve as an exemplar of regulatory signaling, providing a well‐resolved structural template to interrogate the binding affinity and inhibition potential of PET‐associated metabolites on key metabolic enzymes. The deliberate use of sub‐2.0 Å resolution for both protein models was instrumental in ensuring that docking simulations would illuminate the molecular underpinnings of potential interference within both plant and human systems.

Preparation of each receptor followed a stringent protocol: all crystallographic water molecules were excised, polar hydrogen atoms were appended, and Kollman partial charges were introduced to prime the proteins for optimal ligand accommodation. Ligands, emulating the spectrum of PET‐MP degradation products identified via GCMS, were systematically retrieved from PubChem. Their chemical structures underwent conversion from SDF to PDB format using PyMOL version 1.5.7, followed by transformation to PDBQT format through AutoDock Vina version 1.5.7. This preparatory sequence ensured compatibility with the docking software and preserved structural integrity throughout the analysis. Molecular docking simulations were performed using AutoDock Vina, which enabled the exhaustive assessment of binding energies and the precise spatial mapping of ligand–protein interactions. The evaluation leveraged a sophisticated grid‐based scoring system, capable of capturing atomic‐scale determinants of binding specificity. Post‐docking, detailed visualization and interpretative analysis of the resulting complexes were accomplished using PyMOL, affording high‐resolution insight into the structural and energetic landscape of ligand–receptor engagement. The research protocol integrated the resources of NCBI, PubChem, and PDB, and was executed in strict accordance with methodologies articulated in the latest scholarly literature (Xue et al. 2022). Throughout, the investigation was governed by a commitment to transparency, reproducibility, and the highest standards of computational molecular science.

2.8. Statistical Analysis

To achieve a robust quantitative assessment, all gravimetric and other characterization endpoints were measured three times independently, and the values were expressed as mean ± SD. Statistical processing was performed in OriginPro software (version 8.5), employing a two‐way ANOVA that allowed for the evaluation of both principal effects and their interactions. The identification of significant differences among microbial treatments as well as control groups was accomplished using both Tukey's and Dunnett's post hoc tests. At every stage, the criterion for significance was set at p < 0.01, reflecting a conservative threshold to ensure scientific rigor and reliability in the interpretation of results.

3. Results and Discussions

3.1. In Vitro Batch Degradation Studies

To rigorously determine the specific biodegradation capabilities of A. chroococcum (MTCC 3853), R. leguminosarum (MTCC 9766), A. brasilense (MTCC 4036), and T. viride (MTCC 9681) against PET‐MPs under standardized batch culture conditions with mineral salt medium, the study implemented a multifaceted analytical approach encompassing quantitative measurements of microbial growth kinetics, precise determination of PET‐MP mass reduction, comprehensive evaluation of biofilm formation on microplastic surfaces, and sensitive detection of metabolic byproducts attributable to PET‐MP catabolism.

3.1.1. Growth Curve

Growth curves for each microbial species were constructed by recording optical density at standardized intervals post‐PET‐MP inoculation, affording a granular view of their respective proliferation patterns (Figure 1). A. chroococcum (MTCC 3853) rapidly exited the lag phase and maintained a prolonged logarithmic phase (p < 0.01). This phenotype likely arises from a combination of constitutive and inducible expression of a broad spectrum of PET‐hydrolyzing enzymes, enabling efficient initial depolymerization of PET‐MP into intermediates that can be further catabolized. Equally important is the apparent proficiency of A. chroococcum (MTCC 3853) in integrating PET‐MP‐derived monomers into its central metabolic architecture, such as the assimilation of ethylene glycol and terephthalic acid into glycolytic or tricarboxylic acid cycle intermediates, which sustains cell division and biomass accumulation over prolonged periods. The organism's resilience against growth inhibition suggests effective detoxification strategies, including the action of efflux pumps, enzymatic neutralization of reactive intermediates, and the deployment of molecular chaperones or antioxidant systems to counteract oxidative or chemical stress induced by microplastic degradation. Whereas the profiles for R. leguminosarum (MTCC 9766), A. brasilense (MTCC 4036), and T. viride (MTCC 9681) were characterized by extended lag times and a short‐lived log phase, culminating in an early stationary phase. These strains may be hampered by narrow substrate specificity, weak affinity for PET‐MP, or the presence of regulatory mechanisms that suppress catabolic gene expression in the face of unfamiliar substrates. Once the lag phase crosses, the brevity of their log phase underscores a further limitation: the rapid accumulation of degradation byproducts—such as oligomers or aromatic acids—that the cells are unable to effectively process or neutralize. These toxic intermediates can interfere with membrane integrity, disrupt cellular signaling, and inhibit metabolic enzymes, ultimately triggering stress responses that curtail further growth and precipitate entry into the stationary phase. In addition, the metabolic inflexibility of these species may prevent them from rerouting metabolic fluxes or compensating for the toxic effects of accumulated intermediates, further exacerbating their inability to achieve sustained microplastic degradation. Taken together, these findings underscore the importance of a holistic physiological strategy—encompassing rapid enzyme induction, efficient substrate assimilation, and robust stress management—for successful PET‐MP bioconversion, a strategy that A. chroococcum (MTCC 3853) appears to possess to a far greater extent than its counterparts in this experimental paradigm.

Figure 1.

Growth curve of PET‐MPs treated with microbial bioinoculants (statistically significant [*p < 0.01]).

3.1.2. Weight Reduction

Assessing the efficiency of in vitro microplastic degradation predominantly relies on systematic measurement of sample weight, as microbial populations colonizing PET‐MPs initiate enzymatic fragmentation, culminating in a gradual yet measurable decline in mass throughout the experimental timeline (Figure 2). A highly significant reduction in PET weight was achieved by day 25 (p < 0.01), confirming active microbial metabolism of the plastic. Notably, A. chroococcum (MTCC 3853) was responsible for the greatest reduction, achieving 89.1% mass loss, with shorter incubation periods—20, 15, 10, and 5 days—yielding sequentially lower reductions. The impacts of R. leguminosarum (MTCC 9766) and A. brasilense (MTCC 4036) were marginal (8.1% at day 10% and 7.0% at day 15), and no significant differences were observed at other intervals (p < 0.01). T. viride (MTCC 9681) was the least efficient, reaching a maximum of 6.2% at day 15, distinctly lower than all other treatments (p < 0.01). The superior capacity of A. chroococcum (MTCC 3853) for PET degradation likely reflects its evolved enzymatic systems tailored for synthetic polymer catabolism, while the limited reductions seen with other strains point to the necessity of optimizing microbial consortia for targeted bioremediation. The consistently poor performance of T. viride (MTCC 9681) further emphasizes how microbial specificity and enzyme complement determine the overall success of plastic breakdown under experimental conditions. This is consistent with the findings of Chaudhary and Vijayakumar (2020), who observed limited weight loss in polystyrene samples following fungal incubation.

Figure 2.

Weight reduction of PET‐MPs treated with microbial bioinoculants (statistically significant [*p < 0.01]).

3.1.3. Biofilm Formation Efficacy

Biofilms constitute structured microbial ecosystems capable of colonizing both organic and inorganic surfaces, as noted by Hamdy (2024). When formed on microplastics, these communities induce marked shifts in the surface's physical and chemical landscape, thereby enabling extensive microbial colonization (Hall‐Stoodley et al. 2004). Quorum sensing (QS) mediates these interactions, utilizing species‐specific signal molecules to synchronize population activities in response to community composition. In this study, A. chroococcum (MTCC 3853) emerged as the most effective biofilm‐forming agent, achieving 76.4% efficacy at 25 days. A. brasilense (MTCC 4036) and R. leguminosarum (MTCC 9766) produced biofilms at much lower rates (7.2% and 6.4%), and T. viride (MTCC 9681) was completely ineffective in biofilm production (p < 0.01) (Figure 3). To visualize the biofilm structures, SEM was employed. SEM investigation revealed that A. chroococcum (MTCC 3853) generated substantial, matrix‐embedded cell clusters, whereas the other bacterial species produced only weak, thin biofilm layers populated by scattered, unattached cells. The absence of biofilm structures in T. viride (MTCC 9681) was evident, with microscopy showing merely a few isolated spores (Figure S1a–e).

Figure 3.

Biofilm formation efficacy of PET‐MPs treated with microbial bioinoculants (statistically significant [*p < 0.01]).

The preeminent biofilm‐forming ability of A. chroococcum (MTCC 3853) on microplastics can be attributed to a multifactorial synergy of cellular adaptations, regulatory networks, and ecological strategies. This organism is adept at sensing and rapidly responding to abiotic surfaces via the induction of adhesion molecules and EPS‐producing genes, which are integral not only to initial attachment but to the subsequent formation of a densely packed, protective biofilm matrix. The biochemical complexity of the EPS produced—encompassing polysaccharides, proteins, lipids, and extracellular DNA—confers both mechanical stability and functional versatility, enabling the biofilm to sequester nutrients, neutralize toxins, and maintain hydration in a physically challenging environment. At the ecological level, competitive dominance of A. chroococcum (MTCC 3853) is further enhanced by its ability to outcompete other microbes for space and resources on the microplastic surface, in part due to its rapid growth kinetics and efficient resource utilization. Its QS circuitry ensures coordinated expression of biofilm‐promoting genes, facilitating the rapid transition from initial adhesion to mature, structured community. In stark contrast, A. brasilense (MTCC 4036) and R. leguminosarum (MTCC 9766) are likely restricted by less effective adhesion mechanisms, lower EPS production capacity, and potential vulnerability to environmental stresses, all of which limit their capacity to establish persistent biofilms. The inability of T. viride (MTCC 9681) to even initiate biofilm formation underscores fundamental deficits in both surface recognition and stress adaptation systems, likely stemming from evolutionary divergence in cell wall composition, lack of suitable enzymes, or incompatibility of signaling cascades with synthetic polymer environments. Thus, the pronounced difference in biofilm formation efficacy among these strains highlights the critical interplay between molecular, physiological, and ecological attributes in determining successful microplastic colonization and potential biodegradation.

The interplay between microplastic type and microbial biofilm architecture, as explored by Ayush et al. (2022) and Yuan et al. (2020), reinforces the findings of substrate‐specific biofilm formation and microbial adaptation observed in this research.

3.2. Screening of Biodegraded Products

3.2.1. FTIR Analysis

In the present study, FTIR spectroscopy, performed in the range of 4000–400 cm⁻¹ using Thermo Scientific OMNIC Paradigm Desktop Software, was orchestrated to dissect the molecular transformations imparted to PET‐MPs following exposure to selected bacterial and fungal strains. The experimental paradigm was grounded in the hypothesis that the secretion of extracellular enzymes by these microbes would catalyze the cleavage of specific polymer bonds, thereby yielding new or modified functional groups, a process which manifests as discrete shifts or masking effects in the resulting FTIR spectra. The analytical approach centered on identifying the vibrational frequency changes that correspond to such biochemical transformations, with a particular emphasis on the spectral consequences of treatments involving A. chroococcum (MTCC 3853), R. leguminosarum (MTCC 9766), A. brasilense (MTCC 4036), and T. viride (MTCC 9681), each of which was anticipated to leave a unique molecular fingerprint upon the PET matrix.

The FTIR spectrum of untreated PET microplastics, serving as a control, was distinguished by prominent absorption features at 3331 cm⁻¹, attributable to secondary amine and alkyne functionalities, a pronounced band at 1631 cm⁻¹ corresponding to cyclic alkene and amine moieties, and a signature within the 422–417 cm⁻¹ region indicative of metal–oxygen interactions, collectively delineating the baseline chemical architecture of the polymer. Upon challenge with R. leguminosarum (MTCC 9766), the PET‐MPs displayed a modest yet discernible shift in the principal 3331 cm⁻¹ peak to 3320 cm⁻¹, reflecting the transformation of secondary amine groups and the emergence of alcohol functionalities; the cyclic alkene/amine signal observed at 1631 cm⁻¹ shifted to 1633 cm⁻¹, while the metal–oxygen signature broadened from 422–417 to 425–409 cm⁻¹, collectively evidencing the biochemical reactivity of this bacterial strain. A. chroococcum (MTCC 3853) treatment precipitated a dramatic migration of the 3331 cm⁻¹ peak to 3412 cm⁻¹, an alteration interpreted as the formation of aliphatic primary amines and alcohol groups, accompanied by the appearance of numerous new absorption bands between 2209 and 405 cm⁻¹, which are assignable to isothiocyanate, carbodiimide, nitrile, azide, allene, aromatic, halo, and substituted benzene functional groups, signaling a profound restructuring of the polymer matrix. In the case of A. brasilense (MTCC 4036), the FTIR analysis registered a slight elevation of the 3331 cm⁻¹ peak to 3333 cm⁻¹, alongside the manifestation of alcohol signatures and fresh absorption features within the 1712–1239 and 1093–721 cm⁻¹ intervals, corresponding to carboxylic acid, aromatic, nitro, alkane, sulfate, secondary alcohol, aliphatic ether, substituted benzene, fluoro, and alkyl halide groups, thus underscoring the diversity of enzymatic transformation pathways accessible to this bacterium. The application of T. viride (MTCC 9681) gave rise to both pronounced shifts and an array of novel absorptions from 3431 to 2968 cm⁻¹ (alkane, amine salt, alcohol) and within the 1713–1240 and 1094–500 cm⁻¹ intervals, introducing carboxylic acid, aromatic, cyclic alkene, nitro, sulfonate, alkyl aryl ether, fluoro, amine, alkene, halo, substituted benzene, alkyl halide, and metal–oxygen groups, reflecting the complex oxidative and hydrolytic alterations orchestrated by fungal metabolism (Figure 4a–e).

Figure 4.

FTIR spectra of (a) control PET‐MP and PET‐MP degraded with (b) Rhizobium leguminosarum (MTCC 9766), (c) Azotobacter chroococcum (MTCC 3853), (d) Azospirillum brasilense (MTCC 4036), and (e) Trichoderma viride (MTCC 9681).

Through comparative FTIR assessment, A. chroococcum (MTCC 3853) emerged as the most potent agent of PET‐MPs degradation, its treatment culminating in the predominance of alcohol, primary amine, alkyne, isothiocyanate, aromatic, halogen, and alkene functionalities. The observed pattern of peak shifts, emergent and masked bands, and broader spectral modifications directly attests to the efficiency and mechanism of microbially mediated bond scission and functional group evolution, in harmony with the findings reported by Torena et al. (2021), Ayush et al. (2022), and Tegariyanto et al. (2024), thereby reinforcing FTIR as an indispensable modality for biodegradation studies.

3.2.2. GCMS Analysis

The employment of GCMS enabled a comprehensive analysis of the spectrum of degradation products uniquely associated with A. chroococcum (MTCC 3853)‐mediated conversion of PET‐MPs. The resultant chromatograms, as outlined in Figure 5, confirmed the dominance of Monomethyl phthalate, Ethyl‐4‐hydroxyphenylacetate, Isophthalic acid, Methyl‐2,4‐di‐O‐methyl·β·l‐arabinopyranoside, 13‐Docosenamide, Crotonoyl bromide, and Heptasiloxane, while comparative analysis of the control group revealed a different profile, featuring 2‐Ethoxyethyl acetate, 1,4‐Benzenedicarboxylic acid, Heptasiloxane, Phenol, 13‐Docosenamide, and Supraene.

Figure 5.

GCMS spectra of (a) control, (b) Azotobacter chroococcum (MTCC 3853) degraded PET‐MPs.

The present research expands the scientific understanding of microplastic biodegradation by focusing on the metabolic fate of PET‐MPs subjected to enzymatic and non‐enzymatic transformations by A. chroococcum (MTCC 3853). Employing advanced GCMS methodologies, the study meticulously mapped the array of metabolites produced, revealing not only the expected fragmentation products but also the emergence of monomethyl phthalate and isophthalic acid—two entities known for their chemical persistence and capacity to modulate endocrine functions in aquatic and terrestrial organisms. These findings acquire heightened significance when juxtaposed with the results of acute toxicity assessments conducted on V. radiata seeds and D. rerio embryos, in which no overt signs of toxicity, lethality, or developmental arrest were observed at the tested metabolite concentrations, as measured across 96‐hour exposure intervals. Despite these ostensibly reassuring outcomes, the ecological narrative is rendered more complex by the recognition that phthalate‐derived compounds and aromatic acids have a well‐established proclivity for inducing adverse effects through mechanisms that do not necessarily manifest as acute toxicity. Chronic exposure scenarios, characterized by low‐level, sustained contact with these metabolites, have been implicated in the disruption of endocrine signaling pathways, leading to reproductive dysfunction, altered growth dynamics, and compromised population viability over time. The literature further underscores the potential for such compounds to undergo biomagnification within food webs, thereby amplifying their ecological impact beyond what is captured by short‐term, organism‐level bioassays. To address these multifaceted risks, the study underscores the necessity for a paradigm shift in microplastic risk assessment and management. This includes the implementation of secondary and tertiary treatment interventions—such as advanced oxidation processes, bioreactor enhancements, or phytoremediation modules—capable of achieving near‐complete mineralization of recalcitrant metabolites. Furthermore, the adoption of comprehensive environmental monitoring frameworks, integrating both chemical analytics and biological effect markers, is recommended to facilitate the early detection of sublethal and chronic ecological effects. Collectively, these strategies are essential to mitigate the latent, long‐duration hazards associated with the environmental dissemination of PET‐MP biotransformation products, thus safeguarding ecosystem health and resilience.

3.3. Microplastics Degrading Enzymes Production Potential

The intricate network of biological processes responsible for the degradation and cycling of organic matter in natural systems relies heavily on the action of specialized enzymes. Recent concerns regarding the ubiquitous dispersal of microplastics have intensified the scientific pursuit of enzymatic solutions for plastic depolymerization (Tegariyanto et al. 2024; Othman et al. 2021). Enzymes expedite the conversion of robust synthetic polymers such as PET‐MPs by the microbial release of enzymes, including PEG Dehydrogenase, Esterase, Lipase, and Cutinase (Ramamurthy et al. 2024).

By juxtaposing the enzymatic potential of various microbial inoculants, the study identified A. chroococcum (MTCC 3853) as the premier producer, with significantly larger hydrolysis zones in qualitative assessments and the highest U/mL values in quantitative enzyme assays (p < 0.01).

PEG Dehydrogenase initiates the oxidative breakdown of PEG chains, serving as a gateway enzyme in PET‐MP catabolism (Sahu et al. 2023). PEG Dehydrogenase activity profiles revealed that A. chroococcum (MTCC 3853) exhibited a steady, extended rise, peaking at 80 U/mL at 168 h. R. leguminosarum (MTCC 9766) showed a moderate, early peak of 31.2 U/mL at 96 h before plateauing, while T. viride (MTCC 9681) displayed a rapid, low‐intensity maximum of 10% at 48 h with a swift subsequent decline (Figure 6a). The comparative persistence and magnitude of PEG Dehydrogenase activity in A. chroococcum (MTCC 3853) likely result from a confluence of molecular, regulatory, and ecological factors. Genetically, A. chroococcum (MTCC 3853) may possess multiple, highly inducible copies of the PEG Dehydrogenase, facilitated by strong promoters and efficient mRNA processing. Physiologically, the organism's secretion systems and chaperone networks might be especially adapted to ensure enzyme stability and export under xenobiotic stress. In contrast, moderate output shown by R. leguminosarum (MTCC 9766) could be limited by tighter regulatory controls that shut down enzyme production upon reaching threshold substrate or product concentrations, while the weak and short‐lived response by T. viride (MTCC 9681) may be reflective of insufficient induction signals, rapid enzyme degradation, or metabolic competition with other cellular processes. Ecological adaptation is also paramount: The superior performance by A. chroococcum (MTCC 3853) may reflect historical adaptation to environments with high synthetic polymer load, conferring a selective advantage to strains with persistent depolymerizing capabilities. From a biotechnological viewpoint, these insights direct attention towards both natural screening for high‐performing isolates and the rational engineering of less efficient strains to express robust enzyme systems. In addition, deploying consortia that integrate strains with complementary kinetics could help maintain steady‐state degradation rates in complex habitats, maximizing the environmental impact of microplastic remediation efforts. Asees and Divya (2024) and Rosario and Baburaj (2017) found comparable inter‐strain differences among mangrove isolates degrading PVC microplastics, with only certain taxa consistently achieving high PEG Dehydrogenase production.

Figure 6.

Enzyme activity of PET‐MP degraded metabolites (a) PEG dehydrogenase (b) esterase (statistically significant [*p < 0.01]).

Esterase is central to the enzymatic fragmentation of ester‐rich microplastics, producing intermediates for further metabolism (Asees and Divya 2024). A clear performance gradient was observed: A. chroococcum (MTCC 3853) achieved the highest esterase activity, rising to 81.2 U/mL at 168 h; R. leguminosarum (MTCC 9766) peaked at 22.2 U/mL at 96 h; T. viride (MTCC 9681) reached only 8 U/mL at 48 h [Figure 6b]. The sustained esterase output by A. chroococcum (MTCC 3853) is emblematic of deep evolutionary and mechanistic optimization for plastic degradation. At the molecular level, the organism's genome may be enriched with esterase isozymes, each fine‐tuned for slightly different substrate configurations, collectively providing a broad and adaptable toolkit for attacking diverse ester bonds found in PET‐MPs. These isozymes may be regulated by a suite of environmental sensors and signaling pathways, allowing dynamic modulation of enzyme synthesis in response to polymeric stress or the appearance of hydrolysis intermediates. Transcriptionally, esterase produced by A. chroococcum (MTCC 3853) is likely under the control of strong, inducible promoters, possibly responsive to specific small‐molecule effectors derived from plastic breakdown. This ensures both rapid upregulation at the onset of substrate exposure and sustained expression as long as the degradation process continues. Efficient mRNA processing and stability could further amplify the rate of protein synthesis, while translational controls may prioritize esterase production under conditions of environmental challenge. At the metabolic level, A. chroococcum (MTCC 3853) may coordinate esterase synthesis with central metabolic fluxes, ensuring that the energy and precursors needed for robust enzyme production are available even during extended periods of plastic degradation. The organism's secretion pathways are presumably adapted for high‐volume export, and its extracellular environment may be actively managed—via pH regulation or the production of stabilizing agents—to preserve enzyme functionality over time. Ecologically, long‐term selective pressure in polluted or plant‐rich environments would favor strains capable of maintaining high esterase activity for extended durations, outcompeting less resilient organisms and dominating ecological niches where persistent polymers are a primary carbon source. Methodologically, these insights highlight the importance of integrating systems biology approaches—combining genomics, transcriptomics, proteomics, and metabolomics—to uncover the regulatory and metabolic architecture enabling such high performance. This information can be harnessed to engineer new strains or consortia with tailored esterase output, designed to match the demands of specific environmental or industrial plastic degradation scenarios. Looking ahead, the lessons learned from the esterase system generated by A. chroococcum (MTCC 3853) can inform the design of synthetic operons, the selection of microbial consortia for bioaugmentation, and the optimization of process parameters in bioreactors or field‐scale interventions. By emulating or enhancing these natural strategies, the field is poised to develop robust, efficient solutions for the ever‐growing challenge of microplastic pollution. El‐Morsy (2017) and Plou et al. (1998) similarly demonstrated that only select fungal isolates achieve high esterase activity for PU degradation, with most others exhibiting much lower responses.

Lipases, with their ability to hydrolyze diverse ester bonds, are key biocatalysts in both natural and synthetic polymer degradation (El‐Morsy 2017). Among the isolates, A. chroococcum (MTCC 3853) produced a consistent, prolonged increase in lipase activity, reaching 81 U/mL at 168 h. R. leguminosarum (MTCC 9766) achieved a delayed maximum (27.3 U/mL at 144 h), while T. viride (MTCC 9681) showed a rapid but brief spike at 14 U/mL at 48 h (Figure 7a). The pronounced superiority of A. chroococcum (MTCC 3853) in lipase activity can be attributed to a sophisticated interplay of genetic, metabolic, and evolutionary adaptations. At the genomic level, this strain may possess an expanded repertoire of lipase genes, potentially including inducible and constitutive variants, thereby enhancing rapid and sustained responses to changing substrate availability. The regulatory architecture controlling these genes likely features responsive promoter regions and transcription factors attuned to the presence of PET‐MPs and their breakdown products, resulting in a finely tuned, environmentally responsive expression profile. Metabolically, A. chroococcum (MTCC 3853) may allocate a significant fraction of its cellular resources to the biosynthesis, folding, and secretion of lipase enzymes when in the presence of plastic‐derived esters. This prioritization could involve the upregulation of protein chaperones and secretion pathway components, ensuring that newly synthesized lipases are efficiently exported and maintained in an active conformation. In addition, the strain's ability to metabolize the hydrolysis products of PET‐MPs may serve as a positive feedback mechanism, further enhancing lipase production and substrate turnover. On an ecological timescale, persistent exposure to lipid‐rich or plastic‐contaminated environments could drive the evolution of strains with high basal and inducible lipase activity. Such environments would select for organisms capable of exploiting diverse carbon sources, including complex, recalcitrant polymers. In contrast, R. leguminosarum (MTCC 9766) and T. viride (MTCC 9681), with their more modest and transient lipase responses, may be less well‐adapted to such niches, either due to limited genetic capacity, less responsive regulation, or prioritization of primary metabolism over secondary catabolic functions. From a methodological standpoint, the kinetic differentiation among strains underscores the importance of deploying microbial consortia with complementary activity profiles. For example, a combination of high‐output, sustained performers like A. chroococcum (MTCC 3853) and rapid, early responders could maximize the efficiency and continuity of plastic degradation. Furthermore, the use of omics technologies to map the regulatory and metabolic networks underlying lipase production will enable targeted genetic enhancements, either through traditional selection or advanced genome editing. Strategically, future efforts could focus on engineering regulatory modules from A. chroococcum (MTCC 3853) into other industrially relevant hosts, optimizing fermentation conditions for maximal lipase yield, and tailoring enzyme cocktails to match the composition and temporal dynamics of environmental microplastic pollution. Similar findings by Mohanan et al. (2020) reported that LIP1 from Pseudomonas chlororaphis PA23 surpassed LIP2 in PHA hydrolytic efficiency.

Figure 7.

Enzyme activity of PET‐MP degraded metabolites (a) lipase (b) cutinase (statistically significant [*p < 0.01]).

Cutinase bridges the gap between esterase and lipase activity, facilitating the complete degradation of diverse polyester substrates (Bharathi et al. 2019). A. chroococcum (MTCC 3853) achieved the most robust cutinase activity, ascending to 79.2 U/mL at 168 h, with R. leguminosarum (MTCC 9766) and T. viride (MTCC 9681) trailing at 34 U/mL and 17 U/mL at earlier respective time points (Figure 7b). Cutinase activity exhibited by A. chroococcum (MTCC 3853) exemplifies the integration of genetic breadth, regulatory finesse, and ecological specialization. At the genetic level, the strain likely possesses multiple cutinase genes, each equipped for rapid induction and sustained expression. These may be complemented by neighboring genes encoding transporters, accessory enzymes, or regulatory factors that collectively enhance the organism's capacity to sense, access, and degrade polyester substrates. Regulatory complexity is a hallmark of the cutinase system by A. chroococcum (MTCC 3853), with multiple signal transduction pathways converging on cutinase gene promoters. These pathways may respond to a variety of environmental and metabolic signals, including the presence of microplastic surface features, degradation intermediates, or stress‐related cues, ensuring that enzyme production is finely calibrated to both immediate and long‐term needs. The metabolic architecture of A. chroococcum (MTCC 3853) supports the persistent allocation of resources toward cutinase synthesis, export, and function. High‐efficiency secretion systems, possibly coupled with specific chaperones or stabilizing factors, allow for the accumulation of active enzyme in the extracellular matrix, where it can act on recalcitrant polymers over extended periods. This stability is essential for maintaining high levels of enzymatic activity in the face of environmental variability, potential inhibitors, and substrate heterogeneity. Ecologically, the organism's historical exposure to environments saturated with natural or synthetic polyesters would exert selective pressure for the evolution of both the genetic and regulatory machinery necessary for persistent, high‐output cutinase production. This trait confers a decisive advantage in microplastic‐rich habitats, allowing A.chroococcum (MTCC 3853) to outcompete other taxa and drive long‐term degradation processes. Methodologically, an integrated research strategy‐combining temporal enzyme profiling, regulatory network mapping, and synthetic biology—will be key to unlocking the full potential of the cutinase system. Such insights can be translated into engineered strains or microbial consortia optimized for in situ or industrial‐scale plastic remediation, with the flexibility to adapt to the evolving landscape of plastic pollution. Ultimately, the study of cutinase output displayed by A. chroococcum (MTCC 3853) provides a template for the development of next‐generation biocatalysts and microbial platforms, capable of tackling the challenges of persistent microplastic contamination through sustained, high‐efficiency enzymatic action. Gomes et al. (2013) and Gururaj et al. (2021) demonstrated that recombinant expression in E. coli can yield high cutinase activity, but only when host regulatory and metabolic backgrounds are optimized.

3.4. Toxicity Assessment of Biodegraded Products

Despite extensive research underscoring the disruptive potential of PET‐MPs to soil ecosystems, including their capacity to alter key soil functions, hinder resource uptake, and negatively influence plant growth trajectories—effects with substantial consequences for both environmental stability and agricultural output, as delineated by Rillig et al. (2019) and Gomes et al. (2013)—there is a conspicuous lack of studies evaluating the risks associated with the degradation products generated through microbial action on these microplastics; accordingly, this study was comprehensively designed to interrogate the effects of PET‐MPs breakdown products on the seedling physiology of V. mungo and V. radiata by employing robust phytotoxicity methodologies, and to systematically examine the embryonic and acute toxicological responses in zebrafish, thereby offering new perspectives on the broader implications of microplastic pollution.

3.4.1. Phytotoxicity Studies

The germinative phase of crop development is critically dependent on the seed coat's ability to shield the embryo from harmful agents, as highlighted by Souza and Marcos‐Filho (2001) and Rillig et al. (2019). The widespread distribution of microplastics in the environment, notably as outlined by Sahasa et al. (2023), has necessitated research into their effects on plant growth. Accordingly, this study scrutinized the effects of PET‐MPs degradation metabolites on both the germination and early growth of V. mungo and V. radiata.

In the context of PET‐MPs stress, the capacity of different microbial supernatants to mediate seed germination was found to differ substantially. Notably, only the 100% v/v supernatant from A. chroococcum (MTCC 3853) sustained germination rates of 99% (p < 0.01), fully restoring viability to levels indistinguishable from the control. This outcome contrasts starkly with the 82% germination observed under A. brasilense (MTCC 4036) treatment and the 71% rate recorded for T. viride (MTCC 9681) (p < 0.01), as depicted in Figure 8. Such variation underscores the uneven distribution of PET‐MPs remediation capacities among the microbes tested. The superior performance of A. chroococcum (MTCC 3853) likely reflects its ability to both degrade PET‐MPs and suppress the accumulation of phytotoxic intermediates, thus maintaining a non‐inhibitory environment for germination. Conversely, the inferior efficacy of T. viride (MTCC 9681) suggests incomplete detoxification, leading to the retention of toxic substances that inhibit seed emergence.

Figure 8.

Germination index of PET‐MP degraded metabolites (a) Vigna mungo (b) Vigna radiata (statistically significant [*p < 0.01]).

Seedling growth data mirrored germination trends, with A. chroococcum (MTCC 3853) fostering seedling lengths of 9.5 cm (p < 0.01). Intermediate results were seen with R. leguminosarum (MTCC 9766) at 7.5 cm, while T. viride (MTCC 9681) was linked to stunted seedlings of 2.5 cm (p < 0.01) (Figure 9 and Figure S2). This pattern reinforces the significance of thorough PET‐MPs biodegradation for promoting plant vigor. Ineffective degradation, as with T. viride (MTCC 9681), perpetuates the presence of inhibitory byproducts, thus curtailing seedling elongation and potential biomass accumulation.

Figure 9.

Seedling length of degraded PET‐MP degraded microbial metabolites (a) Vigna mungo (b) Vigna radiata (statistically significant [*p < 0.01]).

No significant differences were detected in the proximate analysis—moisture, ash, carbohydrate, protein, fat, and fiber content—between treated and control seeds (Figure S3a–f). The lack of measurable changes in seed composition supports the conclusion that PET‐MP‐degraded metabolites act primarily on developmental processes, rather than altering the intrinsic macronutrient composition of the seeds.

Sahasa et al. (2023) documented that V. mungo seeds exposed to polyethylene microplastics at concentrations of 0.25%, 0.5%, 0.75%, and 1% exhibited a decrease in germination of 12%, 16%, and 20% at the respective concentrations after 24 h. After 48 h, germination rates rose to 90% at all concentrations. In Solanum lycopersicum, polyethylene microplastics resulted in marked reductions in germination at multiple concentrations, with variable decline observed at 4 and 6 days. However, germination rates converged and stabilized by day 7.

3.4.2. Zebrafish Toxicity Studies

In this investigation, the inherently advantageous characteristics of zebrafish—most notably, their transparent embryonic development and swift life cycle progression—were strategically utilized to facilitate a rigorous, real‐time toxicological evaluation of PET microplastic degradation products, with the A. chroococcum (MTCC 3853)‐mediated metabolites serving as the focal toxicants; through the systematic analysis of both embryonic and acute toxicity responses, this study not only elucidated the pathways by which such microplastics are assimilated, distributed, and metabolized, but also provided critical insights into their broader ecological implications for aquatic organisms and interconnected trophic systems, building upon the investigative framework established by Pitt et al. (2018) and Souza and Marcos‐Filho (2001).

3.4.2.1. Embryonic Toxicity

The reliability of the acute toxicity paradigm was unmistakably confirmed by the precipitous and total mortality of the positive control within 6 h, as established in the 0–96 h experimental timeline. Exposure to the degraded sample at 1.0% and 1.5% concentrations led to an abrupt inflection in survival metrics, with significant and enduring declines apparent from 6 h onward, extending throughout the observation period (p < 0.01) (Figure 10). This pronounced, threshold‐dependent deviation from baseline is emblematic of a complex interplay of pathogenic mechanisms, as the data suggest that the biodegraded agents do not act through singular pathways but rather through overlapping cellular insults—including oxidative perturbation, mitochondrial destabilization, and failure of ionic regulation. The ensuing inability of embryos to re‐establish homeostasis under continuous insult drives a progressive unraveling of organismal viability, underscoring the necessity to integrate both acute and cumulative toxicity into robust environmental risk models.

Figure 10.

Survival rate of zebrafish embryo exposed to PET‐MP degraded metabolites at different time intervals (statistically significant [*p < 0.01]).

The hatching process, occurring predominantly between 48 and 66 h, was significantly impaired in both 1.0% and 1.5% treatment groups compared to controls, achieving statistical significance (p < 0.01) (Figure S4). Successful hatching in zebrafish relies on a complex interplay between gene‐regulated enzyme production, proper osmotic regulation, and physical integrity of the chorion. The observed reduction in hatching rate likely arises from disruption at one or several of these levels: the test compounds may inhibit choriolytic enzyme activity, interfere with osmoregulatory pathways, or create a hostile chemical environment via the accumulation of toxic metabolites, thereby directly impeding the mechanical or biochemical steps required for eclosion. Additionally, persistent exposure may induce developmental delays, further reducing the proportion of embryos able to transition from the protected embryo stage to free‐swimming larva.

Heartbeat analysis at 48, 72, and 96 h revealed that only the highest concentrations (1.0% and 1.5%) induced a statistically significant increase in heart rate over controls (p < 0.01) (Figure S5), highlighting the sensitivity of the embryonic cardiovascular system to chemical disturbance. This tachycardia may reflect direct toxicity to cardiac pacemaker cells, upregulation of adrenergic signaling in response to metabolic stress, or a generalized physiological adaptation to maintain tissue perfusion under toxic conditions. Such early cardiac alterations are particularly concerning, as they may compromise oxygen and nutrient delivery to essential organs at critical stages of development, increasing the risk for subsequent morphological and functional defects. The persistent nature of the observed heartbeat elevation supports its utility as a biomarker for sublethal stress and developmental toxicity in aquatic models.

All statistical comparisons were performed using two‐way ANOVA (OriginPro 8.5), confirming strong significance for all endpoints at p < 0.01.

A study conducted by La Pietra et al. (2024) and Pitt et al. (2018) found no survival or hatching alterations at PS‐MPs ≤ 10 mg/L, but reported higher morphological alteration rates and increased heartbeat at 10 mg/L, with 1 µm and 3 µm particles showing distinct alteration profiles.

3.4.2.1.1. Microscopic Examination

Within the framework of a high‐resolution toxicological assessment, the present study deployed a meticulously structured protocol in which zebrafish eggs, harvested at the onset of embryogenesis, were exposed to biodegraded product samples, thereafter undergoing a series of comprehensive microscopic examinations at 2, 4, 6, 24, 48, 72, and 96 h, a temporal arrangement designed to capture the intricate progression of developmental events and to detect any deviation attributable to the tested materials (Figures S6–S12). Initial observations at 2 h post‐exposure revealed that most embryos remained in the oblong stage of cellular division—a period marked by intense proliferative activity—while a subset of the population had advanced to epiboly, thus providing an early indication of the dynamic variation in developmental trajectories among the exposed cohort. By the 4 h interval, a substantial proportion of embryos had progressed to the somite stage, a process sharply contrasted by the complete (100%) mortality observed in the positive control group, a finding that underscored the acute susceptibility of this stage to toxic insult. The 6 h assessment further revealed the thickening of the spinal cord and the simultaneous formation of primordial earbuds and emerging eyelid structures, both of which are indicative of the early phases of organogenesis and neural tissue differentiation. Progressing to the 24 h evaluation, the development of elongated tail structures and the emergence of pigment within the cardiac region were documented, highlighting the advancement of axial patterning and hematopoietic differentiation. Observations conducted at 48 h post‐exposure recorded the presence of blood flow and heartbeat, even in the total absence of a yolk sac, with the tail, yolk sac, and ocular regions exhibiting unremarkable morphology and physiological activity, thus suggesting the preservation of fundamental developmental processes. At the 72 h mark, hatching was observed, accompanied by the initiation of active swimming, signifying both the culmination of embryonic development and the onset of larval locomotion. By 96 h, all examined specimens had undergone complete metamorphosis into larvae, displaying swimming patterns that were indistinguishable from those of unexposed controls, which, in aggregate, provided an exhaustive, time‐resolved account of the effects of biodegraded product exposure on zebrafish embryonic and larval ontogeny, with every critical developmental milestone duly recorded and interpreted within the context of environmental toxicology.

3.4.2.2. Acute Toxicity

A comprehensive assessment of the acute effects of A. chroococcum (MTCC 3853)‐degraded PET‐MP metabolites on zebrafish was conducted using a 7‐day exposure protocol with test concentrations at 0.25%, 0.50%, and 0.75%. The biochemical interrogation included hepatic enzymes (ALT, AST, ALP), metabolic endpoints (blood glucose, total cholesterol, triglycerides), and renal indices (blood urea, creatinine). Consistently across all experimental groups, significant reductions in these parameters were observed when compared with non‐treated controls (p < 0.01), indicating a conspicuous lack of acute systemic toxicity within the studied concentration spectrum (Figures S13–S16). The observed biochemical suppression is hypothesized to result from the generation of smaller, more polar molecules during PET‐MP depolymerization, which are less likely to disrupt cellular homeostasis or elicit stress responses. This is further supported by the absence of compensatory elevation in any measured parameter, suggesting that neither metabolic nor excretory systems were taxed by the metabolite load. These data collectively support the environmental acceptability of deploying A. chroococcum (MTCC 3853)‐mediated bioremediation strategies for PET‐MP contamination, as the resultant metabolites do not impart significant acute toxicity at the tested concentrations. This study also reinforces the imperative for nuanced, multi‐axis toxicity and safety determination processes, particularly as novel biotechnological solutions for microplastic pollution move toward wider implementation.

Results were summarized as mean ± SD and analyzed via two‐way ANOVA with Tukey's multiple comparison test in OriginPro 8.5, which indicated statistically significant differences between groups.

Umamaheswari et al. (2021) and La Pietra et al. (2024) reported persistent increases in LDH and AST, transient decreases in AST (day 14) and LDH (day 21) in the liver, the highest ALT in all groups, and reduced ALP at day 35 after exposure to 10 and 100 μg/L polystyrene microplastics in zebrafish.

3.5. In Silico Docking Studies

The toxic effects of PET microplastic degradation metabolites produced by A. chroococcum (MTCC 3853) were interrogated through an exhaustive in silico docking protocol. Structural data for the V. radiata membrane protein (PDB: 6AFS) and the H. sapiens cytosolic protein (PDB: 6JFV) were extracted from the Protein Data Bank and meticulously processed to remove water molecules and incorporate both Kollman charges and polar hydrogens. Metabolite ligands, identified via GC‐MS and accessed from PubChem, were converted from SDF to PDB through PyMOL and then to PDBQT using AutoDock Vina. The finalized receptor and ligand files were subjected to comprehensive docking simulations, aiming to quantify binding energies and delineate interaction profiles at the proteins’ active sites (Table 2, Figures 11 and 12). The rationale for this advanced computational methodology is rooted in the imperative for methodological rigor and mechanistic clarity. AutoDock Vina's scoring algorithm, with its integration of RMSD‐based pose assessment and grid‐based atomic affinity calculations, provides a robust framework for the accurate prediction of protein–ligand interactions. The preparative steps—dehydration and charge/hydrogen optimization—are indispensable for simulating physiologically relevant conditions, thereby enhancing the translational relevance of the results. PyMOL's visualization capabilities are integral, enabling the detailed analysis and effective communication of binding interactions, which is essential for both internal validation and external dissemination of findings. This comprehensive strategy not only advances our understanding of the interaction dynamics between microplastic degradation products and biological macromolecules but also establishes a methodological benchmark for future studies in computational toxicology and environmental health sciences (Xue et al. 2022).

Table 2.

In silico docking binding energy for Vigna radiata membrane protein (PDB ID: 6AFS) and Homo sapiens cytosolic protein (PDB ID: 6JFV).

S. no	Ligands	Binding energy (kcal/mol) with proteins
		6AFS	6JFV
1	Polyethylene terephthalate (PET)	−7.4	−5.3
2	But‐3‐ynoic acid	−3.4	−2.1
3	Butanoic acid	−3.5	−2.2
4	Cyclopentene, 3‐ethyl	−4.3	−2.9
5	Ethanedioic acid	−3.0	−1.8
6	Formic acid	−2.0	−0.9

Figure 11.

Docked image of Vigna radiata membrane protein (PDB ID: 6AFS) with (a) polyethylene terephthalate (PET), (b) but‐3‐ynoic acid, (c) butanoic acid, (d) cyclopentene, 3‐ethyl, (e) ethanedioic acid, (f) formic acid.

Figure 12.

Docked image of Homo sapiens cytosolic protein (PDB ID: 6JFV) with (a) polyethylene terephthalate (PET), (b) but‐3‐ynoic acid, (c) butanoic acid, (d) cyclopentene, 3‐ethyl, (e) ethanedioic acid, (f) Formic acid.

4. Conclusion

In pursuit of sustainable solutions to microplastic pollution, this study systematically examined the capacity of several microorganisms—specifically A. chroococcum (MTCC 3853), R. leguminosarum (MTCC 9766), A. brasilense (MTCC 4036), and T. viride (MTCC 9681)—to degrade polyethylene terephthalate (PET) microplastics sourced from Pallikaranai, Chennai, India. These microplastics were introduced into a series of batch culture experiments, each designed to facilitate and monitor microbial degradation under controlled laboratory conditions. The experimental design incorporated a multifaceted set of metrics: the percentage reduction in microplastic weight, qualitative and quantitative measures of biofilm development, the characterization of microbial growth patterns, and the analysis of enzymatic activities linked to PET degradation. To provide a holistic assessment of the ecological implications, the study extended its evaluation to include phytotoxicity testing on plant seedlings and acute toxicity assays using zebrafish embryos, ensuring that the safety of the degradation products was thoroughly vetted. Among the candidate species, A. chroococcum (MTCC 3853) consistently demonstrated superior biodegradative performance, achieving the greatest microplastic mass loss, displaying vigorous biofilm formation, supporting prolonged growth, and facilitating high levels of enzyme secretion. The absence of significant toxic effects from its metabolic products in both plant and aquatic systems further reinforces its status as a promising, eco‐compatible agent for PET microplastic degradation.

Funding

The authors have nothing to report.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Supporting File

Data Availability Statement

Data are available on request from the authors.