Evidence for microbial-induced transformation of acrylonitrile-butadiene-styrene (ABS) and styrene-acrylonitrile (SAN) polymer blends by plastic-degrading bacteria

Abstract

Poly(acrylonitrile-butadiene-styrene) (ABS) and styrene-acrylonitrile (SAN) are extensively used in cosmetic and industrial packaging due to their mechanical strength and durability. However, their intrinsic resistance to microbial and enzymatic attack underlies their long-term environmental persistence. To date, there is limited experimental evidence regarding the microbial transformation of ABS/SAN polymers. Here, we investigated whether four previously reported polyethylene-degrading bacterial strains—Comamonas sp., Delftia sp., Stenotrophomonas sp., and Alcaligenes sp.—can grow on an ABS/SAN blend and induce measurable physicochemical modifications. Each strain was incubated for 90 days in a minimal medium containing ABS/SAN as the sole added carbon and energy source. Bacterial growth, viability, and polymer modification were assessed by fluorescence microscopy, gravimetric analysis, and attenuated total reflectance Fourier-transform infrared spectroscopy (ATR-FTIR). All strains sustained active growth and remained viable throughout incubation. Stenotrophomonas sp. and Delftia sp. exhibited the strongest responses, with approximately 6-fold and 2.5-fold increases in viable cell counts and corresponding mass losses of 2.20% and 1.27%, respectively. ATR-FTIR profiles revealed strain-specific chemical changes on the polymer surface, including reductions in nitrile and carbonyl content consistent with partial oxidation and assimilation of ABS/SAN fragments. Comparative genomic analysis further identified genes encoding putative catabolic functions that may facilitate polymer transformation, such as nitrilases, amidases, phenylacetate and aromatic compound degradation pathways, redox-active enzymes, and efflux transporters. Together, these findings provide a novel experimental evidence that select bacterial strains can interact with and induce measurable modifications in ABS/SAN polymer blend, highlighting new perspectives for the biological degradation of recalcitrant synthetic plastics.

Sustainability Statement.

This study supports global sustainability by providing the first experimental evidence that microorganisms can potentially transform and partially degrade ABS/SAN packaging materials, polymers traditionally regarded as environmentally persistent. By demonstrating biologically driven modification of these recalcitrant plastics, the work directly contributes to Sustainable Development Goal (SDG) 12 (Responsible Consumption and Production) by advancing knowledge that can enable biotechnology-based strategies for reducing plastic waste and improving material circularity. These findings also have broader relevance to environmental protection and innovation, also contributing to SDG 14 and 15, as well as to SDG 9 and 17.

Introduction

The cosmetics industry produces more than 120 billion units of packaging annually, yet only a small fraction enters formal recycling streams. A substantial proportion of these materials ultimately accumulates in landfills or the environment, reflecting a broader global crisis in plastic waste management (British Beauty Council, 2026). Globally, plastic waste more than doubled between 2000 and 2019, rising from 156 to 353 million tons, with only a limited fraction effectively recycled (OECD 2022a). Under a business-as-usual scenario, plastic consumption is projected to reach 1231 million tons by 2060, with waste generation exceeding one billion tons (OECD 2022b). This growing imbalance between plastic production and effective waste management is particularly critical for complex and multi-component materials, which are difficult to recycle and tend to persist in the environment.

Among these materials, styrenic copolymers such as acrylonitrile-butadiene-styrene (ABS) and styrene-acrylonitrile (SAN) play a central role in consumer goods and cosmetic packaging due to their mechanical strength, durability, and surface finish (Gilbert 2016, Jiang et al. 2024). The global market for these polymers is projected to grow from 12.53 million tons in 2025 to 18.88 million tons by 2031, reflecting their increasing industrial relevance. SAN is particularly valued for its optical transparency, whereas ABS combines rigidity with impact resistance, making both materials widely used in high-end packaging applications.

Styrenic polymers originate from the polymerization of styrene (C₈H₈), whose vinyl group enables the formation of copolymers such as SAN and ABS with enhanced mechanical and thermal properties (Canevarolo 2013, Callister and Rethwisch 2022). This structural versatility, while advantageous for material performance, contributes to their chemical complexity and resistance to degradation, further complicating end-of-life management. Recycling rates for engineering plastics such as ABS and SAN remain substantially lower than those of conventional packaging plastics, reaching only 18.8%–20% compared to approximately 37.8% in the packaging sector in Europe (Plastics Europe 2024). In addition, the widespread use of carbon black pigments hinders automated sorting, as these materials are not detected by near-infrared (NIR) systems and are therefore diverted to residual waste streams (WMW 2026). Consequently, ABS and SAN are disproportionately directed to landfills or incineration, reinforcing their environmental persistence and impact.

Mechanical recycling remains the predominant route for ABS and SAN recovery, yet it is frequently constrained by thermo-oxidative degradation, chain scission, and contamination with other polymers or additives, ultimately leading to downcycled products of inferior quality (Jakubowicz and Yarahmadi 2024). Solvent-based approaches have emerged as complementary strategies, enabling polymer separation while preserving molecular integrity, with reported reductions in energy consumption and CO₂ emissions relative to virgin ABS production (Lu and Chen 2023). Thermochemical processes such as pyrolysis also offer potential for valorizing complex blends (Rathsack et al. 2025). However, these approaches are frequently energy-intensive, economically constrained, or limited in scalability, particularly for heterogeneous waste streams.

In parallel, advances in microbial biotechnology have renewed interest in bio-based approaches for polymer valorization. Biodegradation pathways are well established for certain polymers, including polyethylene terephthalate (PET), which is enzymatically depolymerized by PETase and MHETase, and polyurethane (PU), which is naturally susceptible to microbial cleavage of urethane linkages (Mohanan et al. 2020). In contrast, styrenic polymers such as polystyrene (PS), ABS, and SAN remain largely unexplored in the context of biological degradation, despite their environmental prevalence and industrial relevance.

Current studies on PS biodegradation demonstrate that styrene can be utilized as a carbon source for microbial growth, undergoing oxidation to phenylacetate and subsequent assimilation via central metabolic pathways (Ru et al. 2020, Mohanan et al. 2020, Lv et al. 2024). However, these findings primarily involve monomers or simplified systems and are not directly applicable to intact polymer matrices. Recalcitrant non-hydrolyzable polymers such as PS generally require prior abiotic oxidation to enable subsequent microbial attack (Wright et al. 2020, Mohanan et al. 2020). The additional chemical complexity of ABS and SAN, including nitrile groups and polybutadiene domains, further increases resistance to degradation and limits direct microbial accessibility.

As a result, current evidence for the biological transformation of intact ABS/SAN polymer blends remains scarce and largely restricted to surface-level oxidation or physicochemical modifications, rather than clear microbial assimilation or mineralization (Gaytán et al. 2021, Lv et al. 2024). This gap highlights the need for experimental systems capable of probing early-stage interactions between microorganisms and these recalcitrant materials under controlled conditions.

Here, we investigate the potential for oxidative surface modification and partial biodegradation of unpretreated cosmetic packaging composed of ABS, CB, and SAN, using bacterial strains previously implicated in plastic degradation, including Comamonas sp., Delftia sp., Stenotrophomonas sp., and Alcaligenes sp. (Peixoto et al. 2017, Peixoto et al. 2026). Importantly, these strains have been previously characterized under carbon-free conditions, where they exhibit rapid loss of viability in the absence of an external carbon source, typically within 12–20 days of incubation (Peixoto et al. 2017). This behavior provides an important baseline for interpreting their persistence in the presence of polymeric materials. To evaluate whether these microorganisms can induce measurable physicochemical modifications under carbon-limited conditions, bacterial activity was assessed through fluorescence-based viability measurements, while polymer transformation was evaluated using gravimetric analysis and attenuated total reflectance Fourier-transform infrared spectroscopy (ATR-FTIR).

We hypothesize that, although complete biodegradation and mineralization are unlikely under the tested conditions, microbial activity may promote surface oxidation, structural modification, and partial utilization of accessible polymer-associated fractions. By integrating complementary analytical approaches, this study provides a mechanistically grounded assessment of early-stage biodegradation processes in styrenic plastics, contributing to the development of biologically informed strategies for plastic waste management in the cosmetics sector.

Materials and methods

Bacterial strains and culture conditions

Four bacterial strains isolated from the Brazilian savanna-like (Cerrado) soil—Comamonas sp., Delftia sp., Stenotrophomonas sp., and Alcaligenes sp.—were selected for this study, based on previous reports demonstrating their ability to degrade synthetic polymers (Peixoto et al. 2017, Peixoto et al. 2022, Frederico et al. 2025). Strains were maintained at −80°C in Nutrient Broth (NB; Difco, Netherlands) supplemented with 15% (v/v) glycerol. For experimental use, cryopreserved stocks were streaked on NB agar and incubated at 28°C for 48 h to ensure an axenic culture. A single isolated colony from each strain was inoculated into 150 mL of NB and incubated at 28°C and 150 rpm until reaching late exponential phase (OD₆₀₀ ≈ 0.5), generating the standard inoculum for biodegradation assays.

Cells were harvested by centrifugation at 5500 × g for 20 min at 10°C, washed three times with sterile saline solution (0.9% NaCl) to remove residual organic carbon, and resuspended in 150 mL of carbon-free minimal mineral medium (MMM) (Peixoto et al. 2017). The MMM was supplemented with 1% (w/v) of disinfected ABS/SAN polymer blend as the sole carbon and energy source. Cultures were incubated at 28°C under orbital agitation of 150 rpm for up to 90 days. Abiotic controls containing sterile MMM and polymeric fragments, but lacking bacterial inoculation, were prepared under identical conditions. All experimental treatments were conducted in biological triplicates.

ABS/SAN characterization and preparation

The polymeric material used in this study consisted of fragments of commercial cosmetic packaging composed of an ABS/SAN blend compounded with approximately 1% CB. The material was supplied by Grupo Boticário (Curitiba, Brazil) and reflects a typical formulation used in rigid cosmetic containers. The characterization of both SAN and ABS/CB polymers—which compose the blend—was performed using a Fourier-transform infrared (FTIR) spectrometer (PerkinElmer, USA) equipped with an Attenuated Total Reflectance (ATR) accessory. The analysis aimed to identify the characteristic absorption bands of the ABS/SAN blend. Spectra were collected over a range of 400–4000 cm⁻¹, with a spectral resolution of 4 cm⁻¹ and 32 accumulated scans per sample to enhance the signal-to-noise ratio (Supplementary Fig. S1; Supplementary Table S1).

Prior to the biodegradation assays, the polymer was mechanically ground into irregular fragments using a high-speed rotary grinder equipped with a stainless-steel chamber to increase surface area and improve microbial accessibility. The resulting material was subjected to a standardized cleaning and disinfection procedure to eliminate surface contaminants. Fragments were first washed with 2% (w/v) sodium dodecyl sulfate (SDS) for 4 h under gentle agitation, followed by a 4 h treatment in 70% (v/v) ethanol. Samples were then thoroughly rinsed with sterile deionized water and oven-dried at 60°C to constant weight (Hadar and Sivan 2004, Peixoto et al. 2017).

Dried fragments were individually weighed on an analytical balance to determine their initial dry weight and stored in sterile glass containers under laminar airflow until use. For biodegradation assays, 1.5 g of disinfected polymer was added to 150 mL of culture medium, corresponding to a final concentration of 1% (w/v). This preparation procedure ensures sterility and reproducibility across replicates while preserving the polymeric integrity and physicochemical characteristics (Hadar and Sivan 2004, Peixoto et al. 2017).

Biodegradation assays

Biodegradation experiments were performed in carbon-free minimal mineral medium (MMM) formulated to support bacterial growth using the polymer blend as the sole carbon and energy source (Pridham and Gottlieb 1948, Peixoto et al. 2017). The medium was autoclaved at 121°C for 15 min and cooled to room temperature prior to the addition of the polymer. Cultures were inoculated with washed bacterial suspensions prepared as described in section “Bacterial strains and culture conditions.” Flasks were incubated at 28°C with constant orbital shaking (150 rpm) for 90 days in the dark to prevent photochemical effects on the polymer surface.

Abiotic controls containing sterile MMM and ABS/SAN fragments but no bacterial inoculum were prepared and incubated under identical conditions to account for potential physical or chemical alterations unrelated to microbial activity. Cultures were sampled at 30 and 90 days of incubation for the assessment of cell viability and biofilm formation on the polymer surface. After 90 days, plastic fragments were recovered for (i) gravimetric analysis to determine total dry weight loss, and (ii) ATR-FTIR analysis to evaluate potential chemical modifications induced by microbial activity. All assays were performed in biological triplicates.

Assessment of bacterial growth and viability

Bacterial adhesion to the polymer surface and cell viability throughout the biodegradation assay were evaluated using fluorescence microscopy after staining with LIVE/DEAD™ BacLight™ Bacterial Viability Kit (Thermo Fisher Scientific, USA). This dual-staining system enables the simultaneous detection of viable and non-viable cells based on membrane integrity, using SYTO9 (green-fluorescent nucleic acid stain) and propidium iodide (red-fluorescent nucleic acid stain).

At 30 and 90 days of incubation, 1 mL aliquots were collected from each culture flask and gently vortexed to disperse both suspended and surface-associated cells. Samples were washed in sterile phosphate-buffered saline (PBS; 10 mM, pH 7.4) and stained according to the manufacturer’s instructions using final dye concentrations of 5 µM SYTO9 and 30 µM propidium iodide (Life Technologies, USA). After 15 min of incubation in the dark at room temperature, stained suspensions were mounted on glass slides and visualized under a Zeiss Axio Fluorescence Microscope equipped with FITC and Texas Red filter sets (Carl Zeiss, Germany).

Fluorescence micrographs were acquired using the ZEN Blue software (Carl Zeiss, Germany) with identical exposure parameters across all treatments to ensure signal comparability. Images were analyzed to qualitatively evaluate bacterial viability and to quantify the proportion of viable (green) and non-viable (red) cells. Quantification was based on at least ten randomly selected fields per sample and processed in ImageJ (Schneider et al. 2012) using pixel-based segmentation and live/dead ratio analysis. This analytical approach allowed monitoring of bacterial persistence, adhesion dynamics, and viability over time, providing insights into the ability of each strain to colonize and potentially metabolize the ABS/SAN surface during long-term incubation.

Dry weight loss

After 90 days of incubation, the entire culture volume was processed to recover the polymeric material for gravimetric analysis. The contents of each flask were filtered through sterile qualitative cellulose fiber filters to separate polymer fragments from the biomass and culture medium. Retained fragments were rinsed with sterile dH₂O to remove loosely attached cells and medium components. Subsequently, recovered fragments were disinfected by immersion in 2% (w/v) sodium dodecyl sulfate (SDS) for 4 h, followed by treatment in 70% (v/v) ethanol for an additional 4 h under mild agitation to ensure the removal of adsorbed organic matter and residual biomass. The fragments were then rinsed with sterile deionized water and oven-dried at 60°C to constant weight. Final dry mass was measured using an analytical balance with a precision of 0.0001 g (Shimadzu AUW220D, Japan). Percentage mass loss was calculated using the initial dry weight (W_i​) and the final dry weight (W_f) according to the following equation:

All measurements were performed in biological triplicate. Abiotic controls incubated under identical conditions were used to correct for non-biological changes in mass (e.g. leaching, swelling, or fragmentation). This gravimetric analysis provided a quantitative endpoint assessment of polymer degradation and complemented the spectroscopic and microscopic evaluations described below.

ATR-FTIR analysis

Chemical modifications of the dried disinfected polymer fragments recovered after 90 days of incubation were analyzed by attenuated total reflectance Fourier-transform infrared spectroscopy (ATR-FTIR). Spectra were acquired using a Bruker Alpha II spectrometer (Bruker, Germany) equipped with a ZnSe ATR crystal and operated through OPUS 7.5.18 software (Bruker, Germany). Measurements were performed at room temperature with a spectral range of 4000–650 cm⁻¹, a resolution of 4 cm⁻¹, and an average of 32 scans per sample.

Raw spectra were exported as ASCII files and numerically processed. Diagnostic absorption bands characteristic of ABS and SAN were monitored, including the nitrile stretching vibration (C≡N) near 2237 cm⁻¹, aromatic C=C stretching around 1600 cm⁻¹, carbonyl stretching (C=O) near 1715 cm⁻¹, and the broad O–H/N–H stretching region around 3400 cm⁻¹. For semi-quantitative analysis, absorbance values were averaged within predefined wavenumber windows corresponding to each functional group (Table 1).

Table 1.

FTIR band assignments and wavenumber windows used for the analysis of ABS/SAN samples.

Region (cm⁻¹)	Assignment	Symbol
3600–3200	O–H/N–H stretching;	AOH
1760–1680	C=O stretching (carbonyl/oxidized groups)	AC=O
1620–1580	Aromatic C=C stretching (styrene ring, internal reference)	AAr
2260–2215	C≡N stretching (acrylonitrile unit)	ACN
1659–1540	Amide I and II (protein/EPS residues)	AAmI, A(AmII

To correct for variations in ATR crystal contact and sample heterogeneity, absorbance values were normalized using two internal reference bands: (i) the aromatic C = C band (A₁₆₀₀) and (ii) the nitrile band (A₂₂₃₇), both reported to be chemically stable (Reggio et al. 2020, Teixeira et al. 2023). From these normalized absorbances, the following diagnostic spectral indices were calculated: I_CN/Ar = A₂₂₃₇/A₁₆₀₀; I_C=O/Ar = A₁₇₀₂/A₁₆₀₀; I_OH/Ar = A₃₄₀₇/A₁₆₀₀; I_C=O/CN = A₁₇₀₂/A₂₂₃₇; I_OH/CN = A₃₄₀₇/A₂₂₃₇; I_AmI/CN = A_AmI/A₂₂₃₇; I_AmII/CN = A_AmII/A₂₂₃₇; and I_OH/C=O = A₃₄₀₇/A₁₇₀₂. Normalization to the aromatic ring band minimized variability in crystal-sample contact, whereas normalization to the nitrile band enabled tracking of changes relative to the acrylonitrile content of the polymer. All spectra were acquired under identical contact pressure and baseline conditions to ensure comparability among treatments. This approach provides a semi-quantitative framework for comparing relative changes in functional groups across treatments, rather than absolute quantification of chemical composition.

Relative variations in the intensities of the C≡N, C=O and aromatic C=C bands were used to assess potential oxidation, nitrile depletion, and structural rearrangements of the ABS/SAN backbone. This spectroscopic approach provided molecular-level evidence of chemical transformations on the polymer surface and complemented the gravimetric and microscopic measurements of biodegradation.

Genomic analysis and functional gene targets

Draft genomes of Comamonas sp., Delftia sp., Stenotrophomonas sp., and Alcaligenes sp. were analyzed to identify genes potentially involved in the biodegradation of ABS/SAN polymer blend. Genome sequencing and assembly were previously described in Peixoto et al. (2022). Functional annotation was performed using Prokka v.1.14.6, followed by refinement through mapping against the eggNOG 6.0, KEGG, and UniProt databases.

Targeted searches were focused on genes associated with (i) nitrile metabolism, including nitrile hydratase (nhhA, nhhB), amidase (amiE), and nitrilase (nit); (ii) styrene degradation, involving styrene monooxygenase (styA, styB), styrene oxide isomerase (styC), phenylacetaldehyde dehydrogenase (padA), and the phenylacetate catabolon (paaABCDEFGHIJK); (iii) polybutadiene and alkene oxidation, including alkane monooxygenases (alkB, ladA, almA), FMN-dependent monooxygenases, and epoxide hydrolases (epx, eh); (iv) oxidative and peroxidative enzymes such as peroxidases (dyp), catalase-peroxidases (katG), cytochrome P450 (cyp), and multicopper oxidases (mco); and (v) aromatic ring-cleavage pathways, including catABC, xylE, mhpB, pcaGH, pcaB, pcaC, and pcaD. Additional searches targeted other oxidoreductases, as well as efflux and transport systems implicated in hydrocarbon tolerance, including RND, ABC, and major facilitator superfamily (MFS) transporters.

Protein sequences were queried using HMMER 3.3.2 (Eddy 2011) against curated PFAM and TIGRFAM hidden Markov model (HMM) profiles. Significant matches were defined by thresholds of E ≤ 1e⁻²⁰, sequence coverage ≥ 70%, and identity ≥ 35%. Orthology was validated using reciprocal best BLAST hits and contextualized using KEGG ortholog assignments. Presence-absence matrices were generated to compare the distribution of degradation-relevant pathways across the four genomes.

Statistical analysis

All experiments were performed in biological triplicates, and results are presented as mean values ± standard deviation (SD). Gravimetric data were obtained by direct measurement of polymer dry mass before and after incubation, and percentage mass loss was calculated individually for each replicate. Abiotic controls were included and processed under identical conditions to account for non-biological variations in mass, such as handling effects, washing procedures, or potential leaching. These control measurements were used as a baseline to estimate and interpret biologically induced mass changes.

For microscopy-based analyses, cell counts and viability estimates were obtained from at least ten independent fields per sample, ensuring robust representation of each replicate. Similarly, ATR-FTIR spectra were collected from multiple random regions of each sample (n = 10 spectra per replicate), reducing variability associated with sample heterogeneity and ATR contact. Spectral processing and visualization were performed in R using the packages hyperSpec, dplyr, tidyr, and ggplot2. Relative absorbance values of replicates were used to compute diagnostic spectral indices and to promote a quantitative comparison of chemical modifications across treatments. Given the exploratory nature of this study, descriptive statistics (mean ± SD) were used to characterize trends across treatments. Inferential statistical testing was not applied to avoid overinterpretation of limited datasets. Instead, consistency across independent analytical approaches (i.e. gravimetry, spectroscopy, and microscopy) was used to support the robustness of the observed patterns.

Results

Bacterial growth and viability in the presence of ABS/SAN polymers

All four bacterial strains were able to grow in MMM supplemented with 1% (w/v) ABS/SAN as the sole carbon and energy source, demonstrating their capacity to persist under stringent nutrient limitation (Table 2). After 90 days of incubation, all cultures showed a measurable increase in total cell density, indicating sustained metabolic activity in the presence of the polymeric matrix. Based on quantitative cell counts, the relative increase in viable cells reached approximately 157% for Alcaligenes sp., 69% for Comamonas sp., 258% for Delftia sp., and 594% for Stenotrophomonas sp., the latter exhibiting the most pronounced population increase.

Table 2.

Percentage of viable cells after 30 and 90 days of cultivation in minimal mineral medium supplemented with 1% (w/v) ABS/SAN (mean ± SD, n = 3).

Bacterial strain	30 days (%)	90 days (%)
Alcaligenes sp.	85.7 ± 1.5	64.8 ± 11.3
Comamonas sp.	75.3 ± 2.6	73.2 ± 3.7
Delftia sp.	73.5 ± 18.4	82.5 ± 5.1
Stenotrophomonas sp.	96.9 ± 2.1	87.5 ± 3.2

Viability assessments performed at 30 and 90 days further confirmed the long-term physiological maintenance of the cultures (Table 2; Figs. 1 and 2). After 30 days, viability remained high across all strains, ranging from 73.5% ± 18.4% (Delftia sp.) to 96.9% ± 2.1% (Stenotrophomonas sp.). By 90 days, Delftia sp. (82.5% ± 5.1%) and Stenotrophomonas sp. (87.5% ± 3.2%) continued to display the highest proportions of viable cells, while Comamonas sp. maintained stable viability (73.2% ± 3.7%). Alcaligenes sp. showed a moderate decline in viability (64.8% ± 11.3%), yet still retained a predominantly viable population capable of sustaining metabolic activity throughout the incubation period.

Figure 1.

Fluorescence microscopy-based viability analysis of Alcaligenes sp., Comamonas sp., Delftia sp., and Stenotrophomonas sp. after 30 days of cultivation in minimal medium supplemented with 1% ABS/SAN. Live cells (+) are visualized in green, dead cells (–) in red, and merged images (+/–) depict the spatial distribution of viable and non-viable cells. All strains maintained predominantly viable populations after long-term incubation with the polymer blend.

Figure 2.

Fluorescence microscopy-based viability analysis of Alcaligenes sp., Comamonas sp., Delftia sp., and Stenotrophomonas sp. after 90 days of cultivation in minimal medium supplemented with 1% ABS/SAN. Live cells (+) are visualized in green, dead cells (–) in red, and merged images (+/–) depict the spatial distribution of viable and non-viable cells. All strains maintained predominantly viable populations after long-term incubation with the polymer blend.

Indeed, fluorescence microscopy corroborated these results (Figs. 1 and 2; Supplementary Figs. S2–S5). Live/dead staining consistently revealed dense populations dominated by live (i.e. green-fluorescent) cells after 90 days, with non-viable (i.e. red-fluorescent) cells representing only a minor fraction. Merged images showed spatially structured microcolonies and biofilm-like assemblies, indicative of surface-associated growth.

Collectively, these findings demonstrate that all four isolates remain physiologically active in the presence of ABS/SAN and can sustain growth for at least three months when the polymer blend is the sole available carbon source. Notably, previous studies have demonstrated that these strains undergo rapid loss of viability when incubated in carbon-free MMM, typically within 12–20 days, due to the absence of an external carbon and energy source (Peixoto et al. 2017). Therefore, the sustained viability and significant increases in cell density observed over 90 days in the present study cannot be explained by endogenous reserves or transient recycling of lysed cells alone. These observations provide qualitative insight into the temporal persistence of bacterial activity, although they do not allow a quantitative assessment of degradation rates over time.

These observations indicate that all strains remained metabolically active in the presence of ABS/SAN and were able to persist under carbon-limited conditions. However, while sustained growth suggests interaction with the polymer system, it does not by itself confirm direct utilization of polymer-derived carbon, as alternative sources such as leachates or trace organic compounds cannot be fully excluded. The persistently high viability levels, coupled with significant increases in cell density, support the interpretation that these strains interact metabolically with the ABS/SAN matrix, enabling prolonged colonization and potentially contributing to the physicochemical modifications described in the following sections.

Quantitative assessment of polymer dry weight loss

Gravimetric analysis demonstrated cumulative measurable reductions in ABS/SAN dry weight after 90 days of incubation for all bacterial strains, although the extent of mass loss varied among isolates (Table 3). Importantly, abiotic controls processed under identical conditions exhibited minimal mass variation (≤ ~2 mg), confirming that the observed changes in experimental treatments exceed baseline variations associated with handling, washing, or experimental procedures. It is important to note, however, that gravimetric changes alone cannot distinguish between biodegradation, physical fragmentation, additive leaching, or the release of low-molecular-weight compounds from the polymer matrix. Thus, these results should be interpreted together with the other analyses performed.

Table 3.

Dry weight variation of ABS/SAN fragments after 90 days of cultivation with each bacterial strain (mean ± SD; n = 3).

Bacterial strain	Final dry weight (g)	Mean reduction (mg)	Total mass reduction (%)
Alcaligenes sp.	1.496 ± 0.004	4 ± 4	0.27 ± 0.27
Comamonas sp.	1.485 ± 0.008	15 ± 8	1.00 ± 0.53
Delftia sp.	1.481 ± 0.018	19 ± 18	1.27 ± 1.20
Stenotrophomonas sp.	1.467 ± 0.029	33 ± 30	2.20 ± 2.00

Initial dry weight = 1.5 g.

Mass loss values were calculated relative to these controls, providing a corrected estimate of biologically associated mass reduction. Stenotrophomonas sp. and Delftia sp. showed the highest reductions (approximately 2.20% and 1.27%, respectively), while Comamonas sp. and Alcaligenes sp. exhibited more modest effects. Although these reductions are quantitatively measurable, they remain relatively low, as expected for highly recalcitrant polymers such as ABS and SAN. Given the relatively small magnitude of mass loss observed, these values should be interpreted with caution, as they may partially overlap with experimental variability associated with long-term incubation and sample handling. Therefore, gravimetric data were interpreted conservatively and not as definitive evidence of complete polymer biodegradation. Instead, mass loss is considered a supporting quantitative indicator of polymer-associated changes, which must be considered in association with spectroscopic and microbiological evidence. In general, these results suggest that bacterial activity contributes to measurable modifications of the material under nutrient-limiting conditions, although the precise contribution of specific mechanisms such as surface erosion, additive leaching, or partial polymer breakdown cannot be fully resolved within the scope of this study.

FTIR analysis reveals chemical modifications of the ABS/SAN matrix

Surface-level chemical changes in ABS/SAN polymer fragments after 90 days of incubation with the four bacterial strains were evaluated using ATR-FTIR spectroscopy (Fig. 3). Diagnostic absorption bands were monitored at 2237 cm⁻¹ (C≡N stretching of acrylonitrile), 1715 cm⁻¹ (C=O stretching), 1600 cm⁻¹ (aromatic C=C), 3400 cm⁻¹ (O–H/N–H stretching), and 1540–1650 cm⁻¹ (amide I and II). To minimize variability resulting from differences in ATR crystal contact, all band intensities were normalized using two internal reference bands: (i) the aromatic C=C ring vibration (A₁₆₀₀) and (ii) the nitrile band (A₂₂₃₇).

Figure 3.

ATR-FTIR spectra of ABS/SAN after 90 days of incubation with Alcaligenes sp. (blue), Comamonas sp. (purple), Delftia sp. (burgundy), and Stenotrophomonas sp. (pink), compared with the sterile abiotic control (red). All bacterial treatments showed detectable alterations in relative band intensities and spectral profiles.

Quantitative spectral indices revealed clear strain-specific differences relative to the abiotic control (Table 4). These normalized indices allow a semi-quantitative comparison of functional group variations across treatments, supporting the interpretation of relative chemical changes in the polymer matrix. The nitrile-to-aromatic ratio (I_CN/Ar = A₂₂₃₇/A₁₆₀₀) decreased in all biotic samples (≈0.83–0.90) compared with the control (≈0.96), indicating a consistent reduction in the relative contribution of SAN-associated nitrile groups on the polymer surface. The most pronounced changes were observed in Comamonas sp. (0.8339 ± 0.24) and Stenotrophomonas sp. (0.8291 ± 0.0071), while Delftia sp. showed similarly reduced but highly reproducible values (0.846 ± 0.0016). The derived nitrile retention index (R_N-Ar; I_CN/Ar, sample/I_CN/Ar, control) ranged from 0.864 to 0.938, reinforcing widespread attenuation of nitrile signatures following microbial exposure.

Table 4.

Diagnostic ATR-FTIR spectral indices of ABS/SAN fragments after 90 days of incubation (mean ± SD, n = 3).

Sample	ICN/Ar	IC=O/Ar	IOH/Ar	IC=O/CN	IOH/CN
Abiotic control	0.9597 ± 0.031	2.071 ± 0.77	0.8948 ± 0.22	2.146 ± 0.73	0.9368 ± 0.25
Alcaligenes sp.	0.8997 ± 0.049	1.834 ± 2.2	1.145 ± 0.59	2.13 ± 2.7	1.298 ± 0.74
Comamonas sp.	0.8339 ± 0.24	0.5194 ± 0.054	0.7896 ± 0.066	0.6525 ± 0.15	0.9968 ± 0.26
Delftia sp.	0.8460 ± 0.0016	0.5231 ± 0.0012	0.7754 ± 0.0066	0.6184 ± 0.0026	0.9165 ± 0.0061
Stenotrophomonas sp.	0.8291 ± 0.0071	0.5214 ± 0.00 016	0.7586 ± 0.0014	0.6288 ± 0.0056	0.9150 ± 0.0062

Indices normalized to the aromatic band include ICN/Ar = A2237/A₁₆₀₀, IC = O/Ar = A1702/A₁₆₀₀, and IOH/Ar = A3407/A₁₆₀₀. Indices normalized to the nitrile band include IC = O/CN = A1702/A2237, IOH/CN = A3407/A2237, IAmI/CN, and IAmII/CN. RN-Ar compares nitrile retention across treatments relative to the abiotic control (RN-Ar = ICN/Ar, sample ÷ ICN/Ar, control).

Carbonyl-based indices exhibited the clearest and most consistent differences among the strains. The carbonyl-to-aromatic ratio (I_C=O/Ar = A₁₇₀₂/A₁₆₀₀) decreased sharply in Comamonas sp., Delftia sp. and Stenotrophomonas sp. treatments (≈0.519–0.523), compared with the abiotic control (2.071 ± 0.77). A similar pattern occurred for the carbonyl-to-nitrile ratio (I_C=O/CN = A₁₇₀₂/A₂₂₃₇), which dropped from 2.146 ± 0.73 in the control to ≈ 0.618–0.652 after the microbial treatments. These reductions suggest biological consumption of carbonyl-containing oxidized moieties of the polymer, which are consistent with the modest but measurable mass loss observed. In contrast, Alcaligenes sp. retained higher carbonyl indices (I_C=O/Ar = 1.834 ± 2.2; I_C=O/CN = 2.13 ± 2.7), suggesting milder oxidation without substantial assimilation of oxidized polymer components.

Hydroxyl-related indices provided additional insight. The hydroxyl-to-nitrile ratio (I_OH/CN = A₃₄₀₇/A₂₂₃₇) increased most strongly in Alcaligenes sp. (1.298 ± 0.74) and moderately in Comamonas sp. (0.9968 ± 0.26), while Delftia sp. and Stenotrophomonas sp. treated samples showed more modest increases. These differences might relate to the efficiency in the metabolization of oxidized fragments to support microbial growth. In addition, the hydroxyl-to-carbonyl ratio (I_OH/C=O = A₃₄₀₇/A₁₇₀₂) increased in all biotic treatments (≈1.455–1.523) compared to the control (0.4904 ± 0.24), reflecting a general shift toward higher surface polarity and evidence of biologically mediated oxidation.

The spectral patterns demonstrate that all four strains induced measurable and strain-specific chemical modifications on the ABS/SAN surface. Alcaligenes sp. produced the clearest accumulation of oxygen-containing groups (C = O and O–H), consistent with mild oxidation followed by limited assimilation of the resulting oxidized fragments. In contrast, Comamonas sp., Delftia sp., and Stenotrophomonas sp. displayed marked decreases in both aromatic- and nitrile-normalized indices, together with reductions in I_CN/Ar. This combination supports a model of preferential utilization of low-molecular-weight oxidized fragments alongside de novo oxidation.

These FTIR signatures substantiate active microbial interaction with the ABS/SAN surface and align with the mass loss trends observed for each strain. Although ATR-FTIR analysis does not directly demonstrate polymer chain scission or mineralization, the use of normalized spectral indices provides a semi-quantitative framework to assess chemical modifications in the polymer matrix, strengthening the interpretation of surface-level transformations. It is important to note that, even though the spectral changes are consistent with chemical modification of the polymer surface, these results are interpreted as evidence of surface-level transformation rather than definitive biodegradation.

Comparative genomic analysis identifies catabolic potential for ABS/SAN biodegradation

Comparative genomic analysis revealed that all isolates possess genes associated with oxidative and hydrolytic processes that could support the transformation of ABS and SAN polymers (Fig. 4; Supplementary Tables S2 and S3). Although the genomes vary in size and gene repertoire, each genome encodes a unique combination of catabolic and redox-related enzymes indicative of potential polymer activation, oxidation, and detoxification. All strains harbored amidases, oxidases, peroxidases, and MFS transporters, which are broadly linked to amide bond hydrolysis, oxidative fragmentation of synthetic polymers, and transport of hydrophobic metabolites. These core features across Stenotrophomonas sp., Delftia sp., Comamonas sp., and Alcaligenes sp. suggest a metabolic basis for plastic surface oxidation and stress mitigation during long-term incubation.

Figure 4.

Comparative heatmap of genes potentially involved in acrylonitrile butadiene-styrene (ABS) and styrene-acrylonitrile (SAN) transformation across Alcaligenes sp., Comamonas sp., Delftia sp., and Stenotrophomonas sp. The heatmap displays the copy number of genes associated with nitrile metabolism, styrene oxidation, aromatic ring cleavage, oxidative and peroxidative stress responses, acrylate detoxification pathways, and transport systems. Color intensity represents log10 (1 + gene copy number), applied to normalize the wide dynamic range of gene abundances across genomes. Gene copy numbers are based on genome annotations and sequence similarity.

The genomes of Delftia sp. and Comamonas sp. encoded multiple enzymes involved in the phenylacetate degradation pathway, including phenylacetate-CoA ligase, phenylacetate oxygenase, and 4-hydroxyphenylacetate isomerase. This pathway enables the conversion of styrene-derived aromatic intermediates into central metabolic substrates, supporting a potential route for assimilation of oxidized styrenic fragments. In Stenotrophomonas sp., genes encoding aminoacrylate peracid reductase, peroxiredoxins, and several oxidative enzyme families were detected, suggesting an enhanced capacity for redox regulation and oxidative attack. Although this genome contained fewer aromatic catabolic enzyme-encoding genes than Delftia or Comamonas, it retained genes for multiple oxidative enzymes belonging to the monooxygenase, peroxidase, cytochrome, and multicopper oxidase families, consistent with a strong potential for polymer surface oxidation.

Additionally, the genome of Alcaligenes sp. combined genes associated with nitrile, acrylate, and aromatic compound metabolism. Detected enzyme-coding genes included nitrilases (aliphatic and arylacetone-specific), isonitrile hydratase, and amidase, together with a complete set of acrylate detoxification enzymes such as methylthioacryloyl-CoA hydratase, acrylyl-CoA reductase, 2-haloacrylate reductase, and the Rut pathway enzymes (RutB, RutC, RutD). In addition, Alcaligenes sp. carried genes encoding phenylacetaldehyde dehydrogenase, phenylacetate-CoA ligase, and p-hydroxyphenylacetate 3-hydroxylase reductase, suggesting metabolic potential for processing both nitrile-derived and styrene-derived intermediates.

Collectively, these genomic features suggest that the studied strains possess metabolic potential to interact with complex hydrocarbon-like substrates, including aromatic and nitrile-containing compounds present in ABS/SAN polymers. In particular, the presence of genes related to oxidoreductases, monooxygenases, and cytochrome P450 enzymes supports the possibility of oxidative transformations, which are consistent with the chemical modifications observed in ATR-FTIR analyses. Additionally, the identification of genes encoding transport systems such as MFS, ABC, and RND families suggests potential mechanisms for substrate uptake and tolerance to hydrophobic or toxic intermediates generated during polymer-associated transformations (Peixoto et al. 2026). Although these findings do not represent a direct evidence of enzymatic activity, they provide a mechanistic framework that supports the observed physicochemical and microbiological results.

Integrated analysis of microbial transformation of ABS/SAN

The integrated evaluation of microbial growth, viability, polymer mass loss, and ATR-FTIR spectroscopy reveals a coherent pattern of strain-dependent interactions with the ABS/SAN polymer blend. Although all four isolates were capable of sustained growth and long-term viability with ABS/SAN as the sole carbon source, the magnitude and nature of polymer modification varied markedly among them. Because only four bacterial strains were evaluated, formal correlation analysis among microbial response, polymer-associated mass loss, and ATR-FTIR indices would have limited statistical power and unstable estimates. Therefore, the datasets were integrated descriptively rather than through inferential correlation testing. This synthesis revealed convergent strain-level patterns: Stenotrophomonas sp. and Delftia sp. showed the strongest biological responses, the highest polymer-associated mass losses, and pronounced alterations in nitrile- and carbonyl-related FTIR indices. In contrast, Alcaligenes sp. displayed the lowest mass loss and a distinct FTIR profile, consistent with a weaker transformation phenotype. These patterns support a qualitative association between sustained microbial activity and ABS/SAN surface transformation, while not demonstrating causality, polymer-derived carbon assimilation, or mineralization.

Stenotrophomonas sp. and Delftia sp. consistently emerged as the most active strains across the experimental endpoints. Both exhibited the largest increases in viable cell numbers over the 90-day incubation and generated the highest polymeric dry-weight reductions (2.20% and 1.27%, respectively). Their ATR-FTIR spectra showed pronounced decreases in both nitrile-normalized and aromatic-normalized carbonyl indices (I_C=O/CN and I_C=O/Ar) together with reductions in I_CN/Ar, indicating a selective depletion of carbonyl- and nitrile-containing functionalities from the polymer surface. These convergent observations suggest a process involving the consumption of oxidized and/or low-molecular-weight fractions originally present on the ABS/SAN matrix.

Comamonas sp. induced an intermediate degree of polymer mass loss (1.0%) and maintained high cell density and viability throughout the incubation. FTIR indices of Comamonas-treated ABS/SAN followed the pattern observed for Delftia sp. and Stenotrophomonas sp., particularly the decreases in I_C=O/CN and I_CN/Ar, though with greater variability across replicates. These observations indicate that Comamonas sp. interacts metabolically with the polymeric surface in a manner consistent with the selective utilization of oxidized fragments, but with lower overall efficiency.

Alcaligenes sp., on the other hand, produced minimal mass loss (0.27%) and showed a moderate decline in viability after 90 days, which likely correlates to insufficient carbon assimilation from the polymer. Indeed, its resulting FTIR profile diverged substantially from those of the other strains, displaying relatively elevated carbonyl-based indices (I_C=O/Ar and I_C=O/CN) and increased hydroxyl-associated absorbance. These evidences suggest that Alcaligenes sp. primarily induces oxidative modifications on the ABS/SAN surface, with limited capacity to assimilate the resulting oxidized fragments.

Integrating these results reveals that the four bacterial isolates interact with ABS/SAN through two main transformation mechanisms. Stenotrophomonas sp., Delftia sp., and Comamonas sp. primarily modify the polymer by (i) forming and then (ii) reducing the abundance of surface carbonyl and nitrile functionalities by assimilation, resulting in measurable mass loss and sustained bacterial growth. Alcaligenes sp., although capable of efficiently colonizing the polymer, induces only minor mass changes and displays a unique IR signature suggestive of superficial oxidative modification rather than substantial chemical depletion. Together, these findings highlight a spectrum of microbial strategies acting on the ABS/SAN matrix, with Stenotrophomonas sp. and Delftia sp. emerging as the strongest candidates for polymer biodegradation under the tested conditions.

Discussion

Bacterial adaptation and growth in the presence of ABS/SAN polymers

The biodegradation of synthetic polymers such as ABS and SAN remains largely unexplored, despite their extensive use in packaging and consumer products and their environmental persistence (Chakraborty et al. 2023, Debbarma et al. 2024). These materials combine aromatic, aliphatic, and nitrile components that confer exceptional mechanical resistance but also render them highly recalcitrant to microbial attack (Wyatt and Knowles 1995, Gewert et al. 2015, Sunarko and Sulistinah 2019, Ellis et al. 2021). To date, no direct evidence has demonstrated bacterial growth or biochemical transformation of ABS/SAN, and proposed mechanisms underlying their biodegradation have been primarily inferred from studies on related materials such as e-waste, polystyrene, and polyacrylonitrile (Mohanan et al. 2020, Gaytán et al. 2021, Zhang et al. 2022, Chakraborty et al. 2023, Debbarma et al. 2024, Lv et al. 2024). In this context, the present study provides experimental evidence that oxidative heterotrophic bacteria are capable of colonizing and modifying an ABS/SAN polymeric surface, thereby expanding the known metabolic repertoire of plastic-degrading microorganisms (Peixoto et al. 2017, Peixoto et al. 2022, Frederico et al. 2025).

All four tested bacterial strains—Alcaligenes sp., Comamonas sp., Delftia sp., and Stenotrophomonas sp.—were able to grow with ABS/SAN as the sole added carbon and energy source, demonstrating a surprising capacity to adapt to the physicochemical challenges imposed by these highly recalcitrant polymers. Viability assays conducted after 30 and 90 days of incubation revealed that all strains remained predominantly active throughout incubation, as confirmed by fluorescence microscopy. Among the isolates, Delftia sp. and Stenotrophomonas sp. exhibited the highest increase in viable cell density, consistent with their previously reported ability to thrive under long-term nutrient-limited conditions. These microbes consist of oxidative heterotrophs with confirmed strong potential to adhere to and degrade hydrophobic and nitrogenated substrates (Peixoto et al. 2017, Peixoto et al. 2022).

A parsimonious explanation for the sustained microbial activity observed in this study is the utilization of dissolved organic carbon (DOC) or low-molecular-weight compounds leached from the ABS/SAN matrix, including potential additives. These soluble fractions may provide readily accessible substrates that support bacterial growth under nutrient-limited conditions. As no DOC quantification or chemical characterization of leachates was performed, their role cannot be excluded and represents an important direction for future investigation. These bacteria may also have relied on the polymer surface as a nutrient interface—likely through slow oxidative modification of the ABS/SAN matrix and partial utilization of low-molecular-weight degradation products. Yet, the observed physicochemical modifications detected by ATR-FTIR, together with genomic evidence of oxidative and aromatic transformation pathways, suggest that polymer-associated transformation processes may occur, albeit as a secondary or later mechanism.

Importantly, the sustained viability observed in this study contrasts with previous findings showing rapid cell death under carbon-free conditions (Peixoto et al. 2017), supporting the interpretation that the polymer blend contributes, directly or indirectly, to maintaining bacterial metabolic activity. This distinction is critical, as it suggests that the observed biological responses are not solely driven by endogenous reserves or starvation survival strategies. The maintenance of metabolic activity over 90 days also suggests efficient energy management under low-carbon conditions, a trait reported for heterotrophic nitrifiers and denitrifiers (Peixoto et al. 2022). In these organisms, it was shown that redox reactions associated with nitrogen metabolism can be coupled to the oxidation of xenobiotic substrates (Hollenberg 1992, Rana et al. 2019, Peixoto et al. 2022).

Fluorescence microscopy further revealed distinct colonization patterns across strains. Delftia sp., Stenotrophomonas sp., and Comamonas sp. formed compact microcolonies and films attached to the polymer surface, whereas Alcaligenes sp. were mostly dispersed (Fig. 2). This morphological differentiation suggests that biofilm formation plays a crucial role in the biodegradation process by promoting localized redox activity and facilitating the concentrating extracellular enzymes at the polymer-cell interface (Peixoto et al. 2017). Biofilm-mediated degradation is widely recognized as a prerequisite for microbial attack of solid hydrophobic polymers because it enables persistent adhesion and creates microenvironments that promote oxidative depolymerization (Wilkes and Aristilde 2017, Tarafdar et al. 2021).

The growth and viability of Stenotrophomonas sp. and Delftia sp., combined with their metabolic versatility and oxidative stress management genes, likely contributed to their superior performance in promoting mass reduction of ABS/SAN when compared to Alcaligenes sp. In general, all isolates were capable of colonizing and persisting on ABS/SAN, despite its complex polymeric nature and the absence of easily assimilable carbon sources. The ability of Delftia sp. and Stenotrophomonas sp. to maintain high viability, form biofilms, and grow in a carbon-limited environment supports the hypothesis that oxidative heterotrophs equipped with redox-balanced and nitrogen metabolism can initiate and sustain the biological transformation of otherwise non-biodegradable plastics (Peixoto et al. 2022).

Evidence of polymer oxidation and chemical modification

The FTIR spectra of the ABS/SAN fragments recovered after 90 days of incubation revealed clear chemical modifications consistent with the oxidative transformation of the polymer surface. Relative to the abiotic control, all biotic treatments showed a marked reduction in the intensity of the nitrile (C≡N) stretching band at approximately 2237 cm⁻¹, a concomitant increase in the carbonyl (C=O) band near 1715 cm⁻¹ and the appearance or amplification of broad O–H/N–H stretching signals around 3300 cm⁻¹. These spectral changes are consistent with oxidative and hydrolytic processes involving partial cleavage or transformation of nitrile, aromatic, and butadiene-derived functional groups.

Similar FTIR shifts, particularly loss of nitrile signatures and the emergence of oxygenated functional groups, have been reported as diagnostic markers of polymer oxidation during bacterial degradation of polyolefins and nitrile-containing polymers (Wilkes and Aristilde 2017, Peixoto et al. 2017, Urbanek et al. 2018, Peixoto et al. 2022). Although ATR-FTIR provides valuable insight into chemical modifications of the polymer surface, it does not directly demonstrate polymer chain scission, formation of degradation products, or mineralization. The spectral indices used in this study enable semi-quantitative comparison of functional group changes, but these should be interpreted as evidence of surface-level transformation rather than definitive proof of polymer degradation.

The reduction in nitrile (C≡N) band intensity is particularly relevant given that acrylonitrile moieties confer chemical and thermal resistance to SAN copolymers. The concurrent enrichment of carbonyl (C = O) and hydroxyl (O–H) stretching bands is consistent with oxidative transformation pathways, potentially involving hydration of nitrile groups followed by the formation of amides and carboxylates or hydroxylated intermediates, processes that may be mediated by amidases nitrile-hydrating enzymes, or broad-spectrum oxidases acting in the presence of reactive oxygen species (Fischer-Colbrie et al. 2006, Fischer-Colbrie et al. 2007, Matamá et al. 2007, Kim et al. 2024, Menon et al. 2025). This interpretation aligns with the genomic profiles of the isolates, which encode amidases, peroxiredoxins, monooxygenases, peroxidases, cytochromes, dioxygenases, and multicopper oxidases, all of which can contribute to nitrile oxidation, hydroxylation, or redox-mediated bond cleavage.

Alterations in the aromatic regions of the spectra (1500–1600 cm⁻¹ and 700–750 cm⁻¹) further suggest partial disruption of the styrene-derived aromatic rings. Decreases in intensity and minor peak shifts in these regions are consistent with biological oxidation of polystyrene and styrene oligomers, where monooxygenases and dioxygenases introduce hydroxyl groups onto the aromatic ring, destabilizing its conjugated structure and promoting subsequent cleavage (Mooney et al. 2006, Mor and Sivan 2008). The presence of these spectral signatures thus reinforce that both SAN and styrene domains of the polymer are susceptible to oxidative modification by the tested strains.

Ultimately, FTIR evidence indicates that the ABS/SAN surface underwent biologically mediated oxidative modification, producing new polar functional groups that likely enhanced the hydrophilicity of the material and facilitated microbial colonization (Orr et al. 2004, Sivan et al. 2006). The formation of carbonyl, hydroxyl, and amide-type functionalities is recognized as a critical early step in the biological degradation of synthetic polymers, as it enhances surface reactivity, promotes fragmentation, and increases bioavailability of oligomeric intermediates. The congruence between these spectral variations and the oxidative/hydrolytic enzymes encoded in the genomes provides a molecular basis for the hypothesis that these strains initiate ABS/SAN biodegradation through oxygen-dependent surface oxidation and partial hydrolysis of nitrile-containing structures.

It is important to emphasize that these findings are based on indirect but complementary lines of evidence. FTIR-based chemical modifications, gravimetric changes, and sustained microbial viability collectively support the occurrence of polymer-associated transformation processes. However, these approaches do not directly demonstrate polymer chain scission, formation of degradation products, or mineralization. In addition, viability was assessed at both 30 and 90 days, while polymer mass loss and ATR-FTIR analyses were performed only after 90 days of incubation. Therefore, the gravimetric and spectroscopic results presented should be interpreted as endpoint measurements reflecting cumulative polymer-associated changes, rather than as kinetic evidence of transformation rates, temporal dynamics, or distinct degradation phases. Although abiotic controls were used to account for non-biological variations, mass reduction alone cannot discriminate between different mechanisms, including biodegradation, physical fragmentation, additive leaching, or the release of low-molecular-weight compounds.

As a result, the data reflect cumulative effects rather than degradation rates or dynamic trends over time. Thus, the magnitude of mass loss and the absence of time-resolved chemical analyses limit the ability to infer degradation rates or distinguish between potential mechanisms such as polymer transformation, additive leaching, or abiotic effects. Likewise, no direct identification of degradation products or carbon balance measurements was performed, preventing confirmation of polymer-derived carbon assimilation. Future studies integrating analytical approaches such as GC-MS, HPLC, isotope tracing, or CO₂ evolution assays will be essential to determine the extent of depolymerization and carbon assimilation related to this process.

All experiments were conducted under controlled laboratory conditions using defined media and isolated strains. While this approach is necessary to disentangle microbial contributions from abiotic factors, it does not fully capture the complexity of natural environments, where variables such as microbial community interactions, environmental heterogeneity, and abiotic weathering processes play a critical role. Therefore, the results presented here should not be directly extrapolated to environmental degradation rates or efficiencies, but rather interpreted as an evidence of the intrinsic capacity of selected bacterial strains to interact with and/or modify ABS/SAN polymers under controlled conditions.

Genome-informed mechanistic insights into ABS/SAN transformation

While this study does not experimentally validate specific enzymatic activities or metabolic fluxes, the genomic analysis provides a robust framework to infer potential mechanisms underlying the observed polymer-associated transformations. By integrating genome annotation with physicochemical and microbiological data, it is possible to propose mechanistic hypotheses linking enzymatic potential to the structural modifications detected in the ABS/SAN matrix.

Genomic analyses of Comamonas sp., Delftia sp., Stenotrophomonas sp., and Alcaligenes sp. revealed distinct yet complementary enzymatic repertoires capable of acting on multiple structural domains of ABS and SAN. Although each strain was evaluated individually, the diversity of enzymatic systems encoded across their genomes outlines multiple non-exclusive pathways for oxidative, hydrolytic, and detoxification reactions that could support the biological transformation of these complex materials. Even though microbial degradation of ABS and SAN is limited to e-waste (i.e. aged mixed materials), the enzymatic machinery identified here provides plausible molecular routes for activating and partially modifying their monomeric constituents (Chakraborty et al. 2023, Debbarma et al. 2024).

Across all four genomes, genes encoding amidases, peroxiredoxins, and MFS transporters were consistently detected. These functional groups are broadly associated with xenobiotic tolerance, oxidative surface oxidation of recalcitrant polymers, and transport of hydrophobic or toxic intermediates (Dubbs and Mongkolsuk 2007, de Carvalho et al. 2014, Muntanda et al. 2022, Singh et al. 2024, Rotilio et al. 2025). Amidases catalyze the hydrolysis of amide bonds, and may participate in the conversion of nitrile-derived intermediates from the acrylonitrile units of SAN into carboxylic acids, consistent with the attenuation of the C≡N band observed in the FTIR spectra (Fischer-Colbrie et al. 2006, Matamá et al. 2007). Peroxiredoxins maintain redox homeostasis under oxidative stress and have also been reported to act on several pollutants and complex polymers (Dubbs and Mongkolsuk 2007, Danso et al. 2019, Shen and Wang 2024). MFS transporters may contribute to the transport of oxidized oligomers and reactive intermediates, supporting long-term viability on otherwise recalcitrant substrates (de Carvalho et al. 2014, Muntanda et al. 2022). These genomic features relate to the ability of these strains to remain viable during prolonged incubation under nutrient-limiting conditions, as previously reported for oxidative heterotrophs that degrade polyolefins and polystyrene (Peixoto et al. 2017, Peixoto et al. 2022, Zhang et al. 2022, Frederico et al. 2025).

More specific metabolic distinctions emerged among the strains. Delftia sp. and Comamonas sp. genomes encoded a well-developed phenylacetate degradation pathway, including genes such as phenylacetate-CoA ligase, phenylacetate oxygenase, and 4-hydroxyphenylacetate isomerase. Together, these gene products funnel styrene-derived aromatic intermediates into central carbon metabolism. This pathway corresponds to the downstream segment of the canonical aerobic styrene degradation pathway, wherein phenylacetate is converted to acetyl-CoA through β-oxidation-like steps (Mooney et al. 2006). The presence of this metabolic module provides a mechanistic explanation for the attenuation of aromatic C = C bands in the FTIR spectra, linking the conversion of oxidized styrenic fragments into assimilable intermediates to the microbial central metabolism.

In contrast, the genome of Stenotrophomonas sp., which exhibited the highest average polymer mass reduction (2.20%), lacked a complete phenylacetate pathway but encoded a suite of oxidative and stress-response enzyme-coding genes, including peroxiredoxins, aminoacrylate peracid reductases, monooxygenases, dioxygenases, peroxidases, multicopper oxidases, and cytochrome P450s. This profile suggests a strong capacity for initiating oxidative attack on the polymer surface, fragmenting the polymeric matrix and generating oxidized or low-molecular-weight intermediates that may then be further metabolized. This phenotype is consistent with reports describing the role of Stenotrophomonas sp. in oxidative fragmentation of polyethylene and other hydrophobic molecules (Peixoto et al. 2017, Frederico et al. 2021).

Additionally, the detection of aminoacrylate peracid reductases in Comamonas sp. and Stenotrophomonas sp. suggests the ability to detoxify reactive acrylate species that may arise from the oxidation of acrylonitrile residues (Kim et al. 2010, Asao and Alber 2013). Such reductases participate in the reduction of toxic aminoacrylates and peracrylates, preventing their accumulation and linking oxidative polymer attack with nitrogen metabolism. These findings are consistent with the ability of heterotrophic nitrifiers and denitrifiers to couple redox reactions of xenobiotic compounds with nitrogen turnover, as previously observed in polyethylene-degrading systems (Guo et al. 2018, Peixoto et al. 2022).

Alcaligenes sp. exhibited a combination of both nitrile and aromatic catabolism. Several genes associated with the acrylonitrile-acrylate catabolic route were identified, including methylthioacryloyl-CoA hydratase, acrylyl-CoA reductase (AcuI), 2-haloacrylate reductase, and aminoacrylate pathway enzymes (RutB, RutC, RutD). These enzymes participate in the detoxification and reduction of reactive acrylate intermediates, facilitating their conversion to propionyl-CoA or other central metabolites (Kim et al. 2010). The concurrent presence of multiple nitrilases (aliphatic and arylacetone-specific), isonitrile hydratase, and amidase further supports the ability of Alcaligenes to hydrolyze nitrile groups from the SAN and ABS fractions, yielding amides and carboxylate products. This genetic combination provides a coherent biochemical rationale for the reductions in nitrile bands observed in the FTIR spectra and for the ability of Alcaligenes sp. to maintain viability during extended incubation, as acrylonitrile detoxification is essential for survival in nitrile-rich environments (Guo et al. 2018).

In line with the fluorescence viability assays and FTIR-based spectral indices, Stenotrophomonas sp. promoted the highest apparent mass loss of ABS/SAN after 90 days (2.20%), followed by Delftia sp. (1.27%) and Comamonas sp. (1.0%). Interestingly, this pattern does not strictly mirror the completeness of aromatic catabolic pathways. Delftia sp. encodes a broader phenylacetic acid degradation module than Stenotrophomonas sp., yet the latter drove the most pronounced mass reduction. Together with the broad array of oxidative enzymes detected across Pfam survey—including flavin-binding monooxygenases, peroxidases, cytochrome P450, and laccases—these genes indicate that Stenotrophomonas sp. is particularly efficient in the initial oxidative attack of the polymer surface, catalyzing early stages of oxidative depolymerization and generating soluble intermediates that may then be assimilated rather than the complete mineralization of styrene-derived products.

In this context, there is a promising potential for a cooperative consortia-model, in which Stenotrophomonas sp. and Alcaligenes sp. act as primary oxidizers of the polymer matrix, generating oxidized fragments and amide intermediates, whereas Delftia sp. and Comamonas sp. could contribute to the further transformation and assimilation of these soluble compounds through phenylacetate and acrylate metabolic pathways. This division of potential metabolic roles may explain the overall degradation observed, highlighting that ABS/SAN transformation by plastic-degrading bacteria proceeds as a multi-step process integrating oxidative fragmentation, nitrile hydrolysis, detoxification of reactive intermediates, and downstream aromatic metabolism. Taken together, these findings support a genome-informed mechanistic model in which oxidative activation, nitrile transformation, and aromatic metabolism act in a coordinated manner to enable partial transformation of the ABS/SAN system. However, these mechanisms should be interpreted as biologically plausible hypotheses supported by convergent evidence, rather than experimentally validated pathways.

Furthermore, the different analytical approaches used in this study provide complementary insights into the interaction between bacterial strains and the ABS/SAN polymer system. While each dataset alone offers limited interpretative power, their integration enables a more robust understanding of the observed phenomena. Sustained bacterial viability and growth under carbon-limited conditions indicate that the presence of the polymer system supports long-term metabolic activity. Gravimetric analysis reveals cumulative changes in polymer-associated mass, whereas ATR-FTIR spectral indices provide semi-quantitative evidence of chemical modifications at the molecular level.

Importantly, these datasets should not be interpreted in isolation or as direct cause–effect relationships. Instead, they collectively support a model in which bacterial activity is associated with physicochemical transformations of the polymer system, even though direct polymer assimilation or mineralization cannot be conclusively demonstrated within the scope of this study. Building upon these mechanistic findings, it is important to consider the ecological context in which such microbial activities may occur and the biotechnological potential of leveraging these metabolic traits for sustainable plastic management.

Ecological and biotechnological implications

This study provides the first experimental evidence of bacterial transformation of the recalcitrant ABS/SAN polymer blend, demonstrating that selected oxidative heterotrophs have potential to colonize, oxidize, and partially depolymerize these complex polymers under laboratory conditions. These findings broaden the known ecological scope of bacterial plastic degraders, previously focused primarily on polyolefins, polyesters, and polyurethanes (Wilkes and Aristilde 2017, Danso et al. 2019). The ability of Stenotrophomonas sp., Delftia sp., and Comamonas sp. to grow and remain viable in the presence of ABS/SAN, while inducing measurable mass loss and chemical modification of the polymers suggests that these taxa may potentially contribute to the environmental fate of aromatic and nitrile-based plastics.

Compared to polymers such as polyethylene terephthalate (PET) and polyurethane (PU), for which enzymatic degradation pathways have been well characterized, the biodegradation of styrenic polymers remains poorly understood (O’Leary et al. 2002, Mooney et al. 2006, Palm et al. 2019). Most available studies report oxidative surface modification, partial fragmentation or degradation of pre-weathered materials, rather than direct microbial interaction with intact polymers (Gaytán et al. 2021, Rybak et al. 2023, Lv et al. 2024). In particular, ABS and SAN are considered highly recalcitrant due to their complex structure, which combines aromatic, nitrile, and aliphatic domains, limiting enzymatic accessibility and microbial attack (O’Leary et al. 2002). As a result, experimental evidence of their biological transformation remains scarce and often restricted to indirect observations (Rybak et al. 2023).

Within this context, the present study contributes to the field by providing convergent evidence that selected bacterial strains can interact with and induce measurable physicochemical modifications in ABS/SAN under controlled nutrient-limiting conditions. These findings should be interpreted as evidence of early-stage transformation processes, contributing to a growing but still limited body of knowledge on the biodegradation potential of styrenic polymers. This is further supported by genomic evidence revealing metabolic pathways associated with aromatic and nitrile compound transformation, which are consistent with mechanisms described for related xenobiotic substrates (Hollenberg 1992, Dubbs and Mongkolsuk 2007, de Carvalho et al. 2014, Rana et al. 2019, Muntanda et al. 2022, Singh et al. 2024, Rotilio et al. 2025).

From an ecological standpoint, the complementary oxidative and hydrolytic pathways among these bacteria reflects a potential division of metabolic labor that could facilitate the transformation of synthetic polymers in natural communities or engineered microbial consortia. The strong oxidative activity of Stenotrophomonas coupled with the catabolic versatility of Delftia and Comamonas points toward a cooperative degradation mechanism. Such synergy resembles naturally occurring microbial consortia that mediate the stepwise breakdown of recalcitrant organic matter in soil and aquatic environments (Urbanek et al. 2018). These observations suggest that, under specific conditions, environmental bacterial communities could potentially contribute to the slow but measurable transformation of ABS/SAN residues, particularly in nutrient-limited ecosystems where oxidative stress responses are coupled to nitrogen cycling (Peixoto et al. 2022, Frederico et al. 2025).

Beyond their ecological significance, the results carry important biotechnological implications. Demonstration that naturally occurring bacteria can oxidize and chemically modify ABS/SAN polymers under mild conditions provides a scientific foundation for developing microbial or enzymatic strategies for the bioremediation, biotransformation, and bio-upcycling of complex styrenic polymers. The enzymatic systems described in this study—including amidases, oxidoreductases, and transporters—represent promising targets for metabolic engineering, directed evolution, or the design of synthetic consortia optimized for aromatic polymer degradation. These mechanisms can be harnessed to design scalable platforms for aromatic polymers, complementing current mechanical and solvent-based recycling technologies that are energy-intensive and yield low-value products (Haider et al. 2019). Harnessing oxidative and nitrile-transforming microbial metabolisms may therefore aid the development of sustainable low-energy biodegradation platforms for styrenic waste disposal and valorization.

It is important to emphasize that all observations in this study were obtained under controlled laboratory conditions using isolated strains and defined media. Such conditions do not capture the complexity of natural environments, where factors such as microbial community interactions, substrate heterogeneity, abiotic weathering, and fluctuating physicochemical parameters strongly influence polymer transformation. Therefore, the ecological implications proposed here should be interpreted conservatively and not directly extrapolated to environmental degradation rates or efficiencies. Instead, the findings reflect the intrinsic metabolic potential of these strains under simplified conditions.

Overall, this work suggests a potential ecological relevance and biotechnological potential of naturally occurring bacteria in transforming polymers previously regarded as non-biodegradable. Future research should focus on several key aspects to advance the understanding of ABS/SAN transformation. First, the identification and quantification of degradation products using analytical techniques such as GC-MS, HPLC, or isotope tracing will be essential to confirm polymer breakdown and carbon assimilation. Second, time-resolved studies with multiple sampling points are required to establish transformation kinetics and dynamics. Third, functional validation of the genomic pathways identified in this study, through transcriptomic, proteomic, or enzymatic assays, will be necessary to confirm the proposed mechanisms. Finally, evaluating these processes under environmentally relevant conditions, including soil and aquatic systems, as well as in microbial consortia and using pre-weathered polymers, will be critical to assess their ecological significance and practical applicability. Ultimately, the coupling of redox-active nitrogen metabolism with oxidative polymer attack represents a unique metabolic strategy (Peixoto et al. 2022, Peixoto et al. 2026) that broadens our understanding of microbial adaptation to synthetic polymers and provides new perspectives for sustainable plastic waste management.

Author contributions

Franciele Saorin (Conceptualization [equal], Data curation [equal], Funding acquisition [lead], Methodology [equal], Writing – original draft [equal], Writing – review & editing [equal]), Tarcisio Camlofski (Conceptualization [equal], Data curation [supporting], Formal analysis [supporting], Funding acquisition [supporting], Investigation [supporting], Writing – review & editing [equal]), Jade Oliveira (Conceptualization [equal], Data curation [supporting], Investigation [supporting], Methodology [supporting], Writing – review & editing [equal]), Anderson Oliveira da Silva (Data curation [equal], Formal Analysis [equal], Writing – review & editing [equal]), Marcia Porto (Conceptualization [equal], Funding acquisition [equal], Project administration [equal], Resources [equal], Writing – review & editing [equal]), Mariana Mattioli (Data curation [equal], Formal Analysis [equal], Investigation [equal], Methodology [equal], Validation [equal], Writing – review & editing [equal]), Ricardo Krüger (Conceptualization [lead], Data curation [equal], Funding acquisition [lead], Methodology [equal], Resources [lead], Supervision [lead], Writing – review & editing [equal]), Julianna Peixoto (Conceptualization [lead], Data curation [equal], Formal Analysis [equal], Funding acquisition [lead], Investigation [lead], Methodology [lead], Project administration [lead], Resources [equal], Supervision [equal], Validation [lead], Visualization [equal], Writing – original draft [lead], Writing – review & editing [equal])

Conflicts of interest

The authors declare no conflicts of interest. Grupo Boticário participated in the study as scientific collaborators and coauthors and did not influence data interpretation or manuscript outcomes beyond their technical and scientific contributions. R.K. is on the editorial board of Sustainable Microbiology. He was not involved in the review or editorial process for this manuscript, on which he is one of the authors.

Funding

This research was supported by Grupo Boticário, which provided financial support for the experimental work and supplied the ABS/SAN polymeric material used in this study. J.P., R.K., and M.M. also acknowledge the National Council for Scientific and Technological Development (CNPq, Brazil), Coordination for the Improvement of Higher Education Personnel (CAPES, Brazil), and the Federal District Research Foundation (FAPDF) for research support.

Data availability

All data supporting the findings of this study are included within the article and its Supplementary material. Additional information is available from the corresponding authors upon reasonable request.