Evaluation of Microbial Degradation of Thermoplastic and Thermosetting Polymers by Environmental Isolates

Abstract

In this study, a microbial–enzymatic strategy was pursued to address the challenge of degrading thermoplastic and thermosetting polymers. Environmental microorganisms were isolated, and their enzymatic activities were assessed using colorimetric assays to evaluate their potential for producing enzymes capable of degrading these polymers. Microorganisms demonstrating higher positivity in the enzymatic assays were selected for a 30-day biodegradation experiment, in which epoxy resins, polyethylene terephthalate, or polystyrene served as the sole carbon source. The effectiveness of biodegradation was assessed through the ATR-FTIR analysis of the chemical composition and the SEM examination of surface characteristics before and after degradation. The results indicated that thermoplastic compounds were more susceptible to microbial degradation, exhibiting greater changes in absorbance. In particular, PET treated with Stenotrophomonas sp. showed the most significant efficacy, achieving a 60.18% reduction in the area under the curve with a standard error of ± 3.42 when analyzed by FTIR spectroscopy. Significant alterations in surface morphology were noticed in thermoplastic compounds. In contrast, thermosetting compounds demonstrated lower reactivity, as evidenced by the absence of band shifts in FTIR spectra and minor changes in bond absorbance and surface morphology.

1. Introduction

Nowadays, the importance of recycling thermoplastic and thermosetting polymers has never been more pressing. With mounting concerns over environmental degradation, resource depletion, and the growing issue of plastic pollution, the need to find innovative and efficient methods for managing these materials has become crucial. Plastics and synthetic polymer materials, ubiquitous in modern society, have permeated every facet of our lives, from packaging and consumer goods to the construction and automotive industries. Similarly, fiber-reinforced polymers, especially carbon-fiber-reinforced polymers (CFRPs), prized for their lightweight and high-strength properties, have found widespread application in the aerospace, automotive, and renewable energy sectors. By 2030, the demand for CFRPs is projected to increase up to 500,000 tons annually [1].

Currently, thermoplastic and thermosetting recycling [2] can be broadly categorized into three methods: mechanical, thermal, and chemical [3,4,5,6]. Mechanical recycling is technologically the most mature polymer recycling method, which involves multiple steps to reduce the size of the waste. Thermal recycling methods utilize high temperature and/or fluids to break down the matrix, categorized as pyrolysis and fluidized bed recycling [4,6]. Chemical and electrochemical recycling methods employ various solvents [5], including supercritical and subcritical ones, often operating at ambient pressures. However, these methods require expensive equipment and may be the cause of adverse effects on the environment and human health due to the waste substances released into the environment [5]. Notably, all these technologies variably diminish the physical and mechanical properties of the recycled product, especially of thermosetting.

Despite their utility, the disposal and inefficient recycling of plastics and CFRPs pose significant environmental challenges. These materials persist in the environment for centuries, polluting oceans, soil, and air, and posing threats to wildlife and human health [7]. Moreover, the traditional recycling methods, especially for CFRPs, often entail power-consuming processes and yield recycled materials of inferior quality, limiting their applicability and exacerbating the demand for virgin resources [8].

Considering these challenges, there is a growing recognition of the potential of microbial degradation as a promising alternative for recycling plastics and CFRPs [9,10]. Microorganisms exhibit an extraordinary capability to enzymatically break down intricate organic materials into simpler compounds. Leveraging the metabolic potential of microorganisms may expedite the degradation of plastics and CFRPs by breaking complex molecules such as polymers into smaller, more readily decomposable oligomers or monomers, thereby contributing to the closure of the material lifecycle loop.

The use of microorganisms might offer several advantages over conventional recycling methods. Microbial degradation is inherently eco-friendly due to its operation under mild conditions, minimal consumption of energetic resources, and negligible production of harmful by-products, unlike the solvents and chemicals utilized in more mature industrial recycling methods. Furthermore, microorganisms are also utilized in the biological treatment process designed to degrade and eliminate pollutants from wastewater, contributing to environmental remediation [11,12]. Regarding CFRPs, conventional methods such as mechanical or chemical recycling pose significant challenges, primarily due to their propensity to exert aggressive actions on the embedded carbon fibers within the matrix. Consequently, the integrity of the fibers is significantly compromised during recycling processes. This deterioration often limits their secondary use potential, as the materials lose their original value and may not meet the required standards for high-performance applications [8].

The microbial degradation of polymeric molecules typically targets chemical bonds that are susceptible to enzymatic hydrolysis or oxidation, such as ester bonds, ether linkages, and amide bonds found in various polymers. In the context of carbon-fiber-reinforced polymers (CFRPs), the epoxy resin matrix commonly used presents a promising solution for microbial degradation, as microorganisms can selectively degrade the resin while preserving the integrity of the carbon fibers. The structural characteristics of carbon fibers, including their highly crystalline and graphitic nature, render them resistant to enzymatic breakdown by microorganisms [13]. Moreover, the inert and tightly packed nature of carbon–carbon bonds in these fibers further impedes microbial colonization and enzymatic activity, contributing to their inherent resistance to biodegradation. Presently, there is a scarcity of research on microorganisms and their interaction mechanisms, especially towards thermosetting polymers. This study’s findings promise to provide valuable insights into how microorganisms engage with and potentially degrade complex materials such as the polymers under investigation. Such understanding could lead to the development of novel strategies for environmental cleanup and sustainable waste handling.

The scope of this paper incorporates investigating the impact of environmental microorganisms on polyethylene terephthalate (PET), polystyrene (PS), and two epoxy resin systems. These materials serve as representative samples for thermoplastic and thermosetting polymers, respectively. The objective is to isolate and identify microorganisms capable of growing in conditions of nutrient deprivation, where the sole carbon source is represented by the compounds selected for this study. The characterization methods for measuring the effectiveness of the degradation process include spectroscopic techniques such as Fourier transform infrared spectroscopy (FTIR), which identifies chemical changes in the materials. Additionally, scanning electron microscopy (SEM) is employed for morphological analysis, observing alterations in the surface structure of the degraded materials. Understanding the degradative capacity of the isolated microorganisms and their impact on the selected compounds will provide a deeper insight into their enzymatic properties. Enhancing understanding will be essential for future research endeavors aimed at investigating how microorganisms able to degrade thermoplastic and thermosetting compounds could improve current technologies and be applied across different fields.

2. Materials and Methods

2.1. Target Materials

Epoxy samples were prepared using two different resin systems (supplied by COMEC Innovative S.r.l., Chieti, Italy). The first system (RS1) was made of Araldite^®. The second system (RS2) was made of Epikote™. The components of each system were mixed according to the specified mixing ratio and cured based on the curing conditions provided in the respective data sheet of the manufacturer. The cured samples were 5 mm × 5 mm × 5 mm in dimensions, achieved by using a silicone mold.

Polyethylene terephthalate (PET; Polymer Institute, SAS, Bratislava, Slovakia) samples were prepared by dissolving granular PET in dimethyl terephthalate (DMT). The solution was cast onto a flat surface using circular molds to ensure uniform thickness. After allowing the solvent to evaporate, thin PET samples resembling paper, 3 cm in diameter, were obtained for various analyses and applications.

Similarly, polystyrene (PS; Polymer Institute, SAS) samples were prepared by dissolving granular PS in tetrahydrofuran (THF), deposited onto flat supports using circular molds, with a diameter of 3 cm, and then subjected to THF evaporation to yield thin, paper-like PS samples suitable for a range of applications.

2.2. Soil and Oil Refinery Activated Sludge (ORAS) Suspensions

The microorganisms were isolated from two samples (soil and ORAS); consequently, two different suspensions were prepared. The first one was prepared by harvesting fresh undergrowth soil, adding it to 360 mL of 0.9% NaCl in a sterile flat-bottomed flask, and placing it in an orbital shaker incubator ES-20 (Biosan, Riga, Latvia) at 100 RPM, room temperature. The suspension was thus filtered and named soil suspension (SS). The second one was prepared using oil refinery activated sludge (ORAS) residue collected from industrial waste oil, adding it to 250 mL of 0.9% NaCl in a sterile flat-bottomed flask, and placing it in an orbital shaker incubator ES-20 (Biosan) at 100 RPM. The supernatant was collected and named ORAS suspension (ORAS-S).

2.3. Agar Media and Isolation of Microorganisms

To isolate the microorganisms contained in the two suspensions (SS and ORAS-S), the following agar plates were prepared: Luria Bertani agar (LBA; Sigma-Aldrich, Burlington, VT, USA), Pseudomonas Isolation agar (PIA; Sigma-Aldrich), Czapek Dox agar (CZDA; Sigma-Aldrich) supplemented with chloramphenicol (25 µg/mL; Sigma-Aldrich); Azotobacter agar (Sigma-Aldrich). All the media were sterilized by autoclaving at 121 °C for 15 min. A series of dilutions of the SS and ORAS-S were prepared starting with a 1:10 dilution, where 1 part of the sample was mixed with 9 parts of 0.9% NaCl, and additional dilutions were prepared up to 1:10,000. One hundred µL of each dilution was plated on the above-mentioned agar plates. The plates were placed at room temperature until the growth of the microorganisms. The individual isolates were selected according to their diverse morphological characteristics.

2.4. DNA Extraction, PCR, and Sequencing

The DNA from selected bacteria was extracted using the DNeasy UltraClean kit (Qiagen, Hilden, Germany) according to the manufacturer’s instructions. For fungal DNA extraction, the DNeasy Plant Mini kit (Qiagen) was employed. Bacterial identification was performed by amplifying the 16S rRNA gene with primers 27f (5′-AGA GTT TGA TCC TGG CTC AG-3′) and 685r (5′-TCT ACG CAT TTC ACC GCT AC-3′ [14]). Fungal identification involved amplifying and sequencing the internal transcribed spacer (ITS) region using primers ITS1 (5′-TCC GTA GGT GAA CCT GCG G-3′) and ITS4 (5′-TCC TCC GCT TAT TGA TAT GC-3′ [15]).

The PCR mixture (25 μL) contained 50 pmol of each primer, 200 μmol L⁻¹ of dNTP (Life Technologies, Gaithersburg, MD, USA), 1.5 U HotStar Taq plus DNA polymerase (Qiagen), 1× PCR buffer, and 3 μL of the extracted DNA. The PCR protocol included an initial denaturation at 94 °C for 5 min, followed by 30 cycles of denaturation at 94 °C for 30 s, annealing at 54 °C for 45 s, extension at 72 °C for 1 min, and a final extension at 72 °C for 10 min.

PCR products were purified using ExoSAP-IT (Affymetrix, Cleveland, OH, USA) and sequenced at Eurofins Genomics (Ebersberg, Germany). The sequences were compared with those in GenBank using the BLAST program (http://blast.ncbi.nlm.nih.gov/Blast.cgi (accessed on 4 July 2022)) and were deposited under accession numbers PP486466-PP486480 (bacterial isolates) and PP488307-PP488309 (fungal isolates).

2.5. Colorimetric Agar Assays

The lipolytic activity was assessed on commercial Spirit Blue agar (SBA; HiMedia Laboratories, Mumbai, India). Spirit Blue agar was prepared by suspending 32.15 g in 1000 mL distilled water, autoclaved, cooled down to 50 °C and supplemented with 30 mL of lipase substrate (1 mL of Tween 80, 400 mL of warm distilled water, and 100 mL of linseed oil; sterilized by autoclaving), slowly mixed, and poured onto Petri dishes [16]. One hundred µL of broth medium (Luria Bertani broth, Sigma-Aldrich) containing a specific isolate was plated onto the SBA and left at room temperature over a week and visually examined for the formation of a bleaching ring around the colonies.

The haloalkane dehalogenase activity of isolates was also evaluated. The specific medium (dehalogenase pH indicator agar) was prepared by suspending 5.00 g of LB broth and 15 g of Bacteriological agar (HiMedia, Mumbai, India) in 1000 mL distilled water. The pH was adjusted to 8.2 with 1 M NaOH prior to autoclaving. To indicate pH changes, a mixture of phenol red (PhR) and bromothymol blue (BT) (each at a final concentration of 13 μg/mL) was added to the medium. The medium was sterilized by autoclaving, cooled down to 50 °C, and poured onto the Petri dishes. Each freshly grown isolate was streaked onto a sterile 0.2 μm-pore-size nitrocellulose filter and grown overnight on an LB agar plate. Then, the filters were transferred to the dehalogenase pH indicator plates. Pure 1,2-dichloromethane was applied to the lids of the plates, which were then sealed shut with parafilm. The plates were incubated at room temperature over a week and visually examined for color change [17].

The cutinase activity was detected by using polycaprolactone (PCL; Polymer Institute, SAS), which was priorly dissolved in acetone to favor the dissolution of the compound and therefore added to Minimal Medium (MM; KNO₃ 0.5 g/L, KH₂PO₄ 0.1 g/L, KH₂PO₄ 1 g/L, MgSO₄ 0.5 g/L, KCl 0.5 g/L, Bacteriological agar 15 g/L) previously autoclaved before plating. One hundred µL of broth medium (Luria Bertani broth, Sigma-Aldrich) containing a specific isolate was inserted into the middle of a PCL plate and left at room temperature for two/three days. An aqueous solution of safranin was prepared by dissolving 2.5 g of safranin in 10 mL of 95% ethanol and then adding it to 100 mL of distilled water. After incubation, the microorganisms on PCL agar plates were flooded with the safranin stain solution and allowed to stand for 20–30 min with gentle shaking. Stained plates were washed off thrice with 1.0 M NaCl solution for 15–20 min. Then, all the plates were observed for clear zone formation [18].

2.6. Biodegradation Experiment

Samples of thermosetting and thermoplastic materials, including RS1, RS2, PET, and PS, were prepared as the object of study. Minimal Medium (MM) was used as the culture medium for separately inoculating the materials with five different microorganisms. Each compound underwent individual exposure to all five selected microorganisms, with each inoculation conducted in separate Erlenmeyer flasks. The five most promising microorganisms were selected from the isolated strains, based on positive outcomes observed in colorimetric agar tests. The flasks containing MM, plastic compounds and microorganisms were then left in a 100 rpm orbital shaker at room temperature for 30 days. On the thirtieth day, all the samples were taken out of the flasks and placed in a small Eppendorf tube to be sterilized by heat prior to further analysis. The experiment was deliberately designed to last only 30 days, as the primary goal was to evaluate the feasibility of implementing a biodegradation process on an industrial scale.

2.7. Fourier Transform Infrared Spectrometry Analysis

All spectra were measured within the mid-IR range (4000–400 cm⁻¹), using the NICOLET 8700™ spectrophotometer (purchased from ThermoFisher Scientific, Madison, WI, USA). The spectrum of each sample was acquired from three different points on the surface of the material after 30 days of incubation with a selected microorganism. Following the acquisition of three spectra for each sample, an arithmetic mean of the obtained values was calculated to generate a composite spectrum as the result for each sample. This was carried out to mitigate any potential differences in degradation that occurred on the surface. The resolution was set to 4 cm⁻¹ and the average number of scans was between 100–200. Spectra were baseline-corrected [19] and automatically smoothed using Omnic software Ver. 8.1.0.10. (ThermoFisher Scientific, Madison, WI, USA). Peak integration of the FTIR spectra was carried out for PET samples using OriginPro 2023 software Ver. 10.0.0.154. (OriginLab Corporation, Northampton, MA, USA). Integration of the peaks was performed using various methods available in OriginPro, including simple summation, trapezoidal integration, or other appropriate algorithms depending on the shape of the peaks and the nature of the data. This integration process allowed for the calculation of the area under each peak, providing a quantitative measure of the intensity or concentration represented by the peak. The peaks associated with particular and distinctive vibrational modes of chemical bonds in the compounds under investigation were pinpointed and individually adjusted to extract their respective areas. The integrated area under each peak was then calculated and compared to that of the pristine sample, and then used for quantitative analysis of functional groups present in the samples subjected to biodegradation.

2.8. Scanning Electron Microscopy (SEM)

The surface of samples subjected to microbial attack was examined using a JSM Jeol 6610 Scanning Electron Microscope (SEM) (JEOL, Tokyo, Japan) with an accelerated voltage of 15 kV. To prepare the non-conductive materials for observation, a thin gold layer was applied using a Balzers SCD 040 sputter coater (Balzers Union Limited, Balzers, Liechtenstein). The gold layer helps dissipate charged electrons from the material. SEM images were captured using AzTec software Ver. 2.1. (Springfield, NJ, USA), and later processed with ImageJ software Ver. 1.8.0., which is freely available. (LOCI, University of Wisconsin, Madison, WI, USA).

3. Results and Discussion

3.1. Target Materials

The two three-component epoxy resin systems utilized in this study, RS1 and RS2, are both covered by industrial patents, making it challenging to precisely identify their chemical structures. Very limited detailed information regarding the composition and formulation of the resins is publicly available. The RS1 consists of Araldite LY 3508, a bisphenol-A-based epoxy resin known for its high molecular weight and superior thermal stability; Aradur 1571, a curing agent that contains dicyandiamide (DICY), which acts as a latent hardener providing extended workability at room temperature and reacts upon heating to form a highly cross-linked network; and Accelerator 1573, which is also based on DICY, serving to expedite the curing process and ensure uniform and complete polymerization. The RS2 employs Epikote 05910, another bisphenol-A-based epoxy resin characterized by its excellent chemical resistance and durability; Epikure Curing Agent 05900, a DICY-based latent hardener similar to Aradur 1571; and Epikure Catalyst 05900, a homogeneous, agglomerate-free dispersion of DICY in liquid epoxy resin that functions as an accelerator to ensure consistent and efficient curing. These insights are derived from publicly available materials and laboratory analyses. Additionally, the chemical structures of polyethylene terephthalate and polystyrene are known, as they are not protected by industrial patents.

3.2. Isolate Identification and Their Enzymatic Characterization

Isolated microorganisms (18 microorganisms) from soil and ORAS were identified by DNA sequencing (

Table 1. DNA identification of isolates and their enzymatic properties. In bold, the isolates utilized in the biodegradation experiment.

Sample	Isolate	DNA Identification, Sequence Similarity	Lipases Assay	H. D. Assay	Cutinases Assay
Soil	S1	HQ634928 Acinetobacter sp. 99.49%	+	−	−
	S2	LT547828 Flavobacterium oncorhynchi 100%	+	−	−
	S3	MH512740 Flavobacterium succinicans 100%	+	+	−
	S4	MK965149 Flavobacterium sp. 100%	+	−	++
	S5	MK368666 Rahnella sp. 99.84%	+	++	++
	S6	KY117494 Stenotrophomonas sp. 100%	+	++	++
	S7	MN193369 Luteibacter sp. 99.69%	+	−	−
	S8	OP811704 Luteibacter pinisoli 99.84%	+	−	−
	S9	MT502791 Sporobolomyces roseus 99.62%	+	++	−
	S10	MH595326 Bullera alba 100%	+	++	+
Oil Refinery Activated Sludge	ORAS1	JN378750 Pseudomonas stutzeri 100%	+	−	−
	ORAS2	ON688707 Pseudomonas stutzeri 100%	+	−	−
	ORAS3	ON706951 Pseudomonas linyingensis 99.37%	+	−	−
	ORAS4	OL348500 Scedosporium boydii 100%	+	−	+
	ORAS5	LT629780 Pseudomonas guangdongensis 99.84%	+	−	−
	ORAS6	OL445008 Bacillus tropicus 99.69%	+	++	++
	ORAS7	AF094741 Pseudomonas putida 99.83%	+	−	+
	ORAS8	MZ519906 Acinetobacter sp. 100%	+	−	−

H. D.: haloalkane dehalogenase; (−): negative; (+): positive; (++): strongly positive compared to others.

). The soil was characterized by the presence of members of the genus Flavobacterium, while in the ORAS sample members of the genus Pseudomonas were predominant. Flavobacterium and Pseudomonas are two bacterial genera that have been associated with the degradation of plastics and polymers. Studies have indicated that Pseudomonas possess the capabilities to degrade and metabolize synthetic plastics such as polystyrenes and polyethylenes [20]. Furthermore, Pseudomonas, along with Flavobacterium, have been identified as dominant in the degradation of polypropylene carbonate (PPC) plastic films, further emphasizing their role in plastic degradation processes [21]. Three fungi were also isolated: Sporobolomyces roseus and Bullera alba from soil samples, and Scedosporium boydii from ORAS.

All microorganisms subjected to the Spirit Blue agar (SBA) assay demonstrated positive results (Table 1). Lipases and esterases are the key enzymes assessed in this test, and the positive outcomes across the board suggest a widespread presence of these enzymes or their related activity among the tested microorganisms. Lipases are known for their ability to hydrolyze ester bonds, which are commonly found in thermosetting and thermoplastic structures, and they have been shown to effectively hydrolyze a variety of substrates containing ester linkages. For instance, some research conducted by Jaeger [22] and Satti [23] highlighted that bacterial lipases exhibit a preference for hydrolyzing primary ester bonds in triacylglycerol substrates. Esterases have been observed to play a role in breaking down cellulose, indicating that nature may be developing analogous decomposition strategies for synthetic plastics with similar structures, such as PET [24]. Moreover, investigations have demonstrated the effectiveness of enzymes such as cutinases and lipases, belonging to the subclass of esterases, in the degradation of PET, as reported by Ahmaditabatabaei [25]. The enzyme facilitates the cleavage of ester bonds, leading to the depolymerization of the molecules.

Initially, SBA plates typically exhibit a dark greenish-blue hue; however, as the incubation progresses, microorganisms capable of producing lipase enzymatically break down the tributyrin present in the agar. The transformation is observable as a shift from the initial greenish-blue color towards a clear halo surrounding the bacterial colonies (Figure S1), indicating the hydrolysis of tributyrin and the subsequent acidification of the surrounding medium.

Haloalkane dehalogenases (HLDs) constitute a class of enzymes that play a pivotal role in biodegradation processes by facilitating the cleavage of carbon–halogen bonds in various halogenated organic compounds [26]. This could potentially contribute to the breakdown or modification of thermosetting and thermoplastic polymers, but the specific mechanisms and efficiency would depend on the enzyme’s substrate specificity and the chemical structure of the polymer [26]. The indicator plates used in the experiment had a violet color (Figure S2). After the application of the nitrocellulose filter on which the microorganisms had been grown overnight and following further incubation, some plates changed color. The toning, by some microorganisms, was more accentuated, bringing the coloring of the plates to a pink/orange, while others did not cause any toning of the indicator plates, indicating a negativity of the expression of HLDs (Figure S2). The test results, indicating whether positivity was observed or not, are presented in Table 1.

Cutinases represent a diverse group of enzymes that play a crucial role in the breakdown of cutin, a complex polymer that forms the waxy protective layer on the surface of plant leaves and fruits [27]. Polycaprolactone (PCL), a synthetic polyester mostly used in the production of polyurethanes, has a structure analogous to that of cutin, which provides the opportunity of different microorganisms to degrade the polymer [28]. The indicator plates used in the experiment had a milky color (Figure S3a). To better observe the clear zone formation, the plates were stained with an aqueous solution of safranin (Figure S3b) and the results are shown in Table 1. Few microorganisms showed capacity to grow and create a clear area on the plates. Among them, the most promising were Flavobacterium sp. (S4), Rahnella sp. (S5), Stenotrophomonas sp. (S6), and Bacillus tropicus (ORAS6). Some cutinases have shown potential for the breakdown of various classes of polyesters and polyamides, as investigated by Ferrario [27]. The mechanism involves the enzyme attacking ester bonds in the polymer chains. Specifically, in the case of PET, cutinase has shown capacity to cleave ester linkages, breaking it down into its constituent monomers [29].

Five microorganisms [the bacteria: Rahnella sp. (S5), Stenotrophomonas sp. (S6) and Bacillus tropicus (ORAS6); the fungi: Sporobolomyces roseus (S9) and Bullera alba (S10)] were chosen, according to the results of agar colorimetric assays, in order to perform the biodegradative tests on PET, PS, and the two epoxy systems (RS1 and RS2).

3.3. Characterization of Degradation of Plastic Materials by Selected Microorganisms

3.3.1. Polyethylene Terephthalate (PET)

In Figure 1 are reported the FTIR spectra in the “fingerprint” region (below 2000 cm⁻¹), where most of the significant PET bands are [30]. The black line represents the pristine PET sample. Peaks at 727 cm⁻¹ are associated with the interaction of polar ester groups and benzene rings; 1018 cm⁻¹, gauche bending oxy-methylene group; 1099 cm⁻¹, stretching C-O-C group; 1261 cm⁻¹, asymmetric stretch C-C-O group bonded to aromatic ring; 1340 cm⁻¹, trans wagging band -CH2; 1370 cm⁻¹, gauche wagging band -CH2; 1410 cm⁻¹, bending -CH2; 1710–1740 cm⁻¹, stretching -C=O. Comparing the FTIR spectra of the PET samples subjected to degradation activity with the PET control, the most significant bands did not change the wavelength at which they were expressed. However, the absorbance of the bands in the degraded samples was lower compared to the control sample. This may suggest that the degraded samples retained their original chemical structure, losing some of their initial mass. To better compare the data, FTIR spectra were normalized to the stretching of the carbonyl bond present at 1720 cm⁻¹.

SEM images of the PET control surface, captured at magnifications of 500× and 1400×, reveal a generally smooth surface with minor irregularities resulting from the sample preparation process. Among the five microorganisms tested for their degrading activity on PET samples, a particular activity by Rahnella sp. and Stenotrophomonas sp. was noted. Rahnella sp. seems to have caused the formation of crystals, visible at the magnification of 1400×. In relation to the behavior of Stenotrophomonas sp., it appears to have induced the formation of folds on the surface of the material (Figure 2), a phenomenon previously documented during the investigation of low-density polyethylene biodegradation [31]. This indicates that Rahnella sp. and Stenotrophomonas sp. likely exhibit a strong affinity for PET surfaces by expressing specialized esterases, such as cutinases, which facilitate binding to the surface and cleave ester bonds. In the literature, there is limited direct evidence regarding the capability of Rahnella sp. and Stenotrophomonas sp. in degrading PET. Some studies have demonstrated the enzymatic degradation of PET by different bacterial species. For example, Müller [32] discussed the rapid hydrolysis of PET facilitated by a hydrolase from Thermobifida fusca, highlighting the impact of factors such as crystallinity, melting point, and glass transition temperature on the enzymatic breakdown of PET, shedding light on the enzymatic mechanisms involved in PET degradation and the catalytic potential of specific enzymes. Furthermore, Stenotrophomonas sp. has been linked to the biodegradation of diverse compounds, as evidenced by Santos [33], who identified efficient carbendazim-degrading bacteria, including Stenotrophomonas sp. Although carbendazim is a fungicide and not a plastic material itself, the structure of carbendazim contains a benzene ring and an imidazole ring, which are aromatic in nature. This aromaticity and the presence of nitrogen and oxygen atoms in its structure give carbendazim some similarities to certain types of plastics, particularly aromatic polyesters, and polyimides. These findings suggest the metabolic adaptability of Stenotrophomonas sp. in breaking down complex organic compounds, indicating its promising use for thermoplastic and thermosetting compound degradation.

3.3.2. Comparison of the Area under the Curve in PET FTIR Spectra Following Biodegradation

The absorption bands of both pristine PET and PET subjected to microbial degradation exhibit overlapping bands without any observable shifts. Consequently, the area under the curve of the six principal absorption bands was computed and found to be comparable. Specifically, the absorption bands corresponding to the interaction of polar ester groups and benzene rings (P1), gauche bending oxy-methylene group (P2), stretching C-O-C group (P3), asymmetric stretch C-C-O group bonded to aromatic ring (P4), bending -CH2 (P5), and stretching -C=O (P6), as reported in Figure 3. The area under the curve for each peak was determined using OriginPro software (OriginLab Corporation) and subsequently compared between the control and degraded samples of PET. This was repeated for each microorganism employed in the study (Figure 3). A tabulated dataset was compiled to illustrate the percentage reduction in area under the curve relative to the PET control after a degradation period of 30 days (

Table 2. Comparison of inoculating PET samples by a single microorganism [Rahnella sp. (PET_S5), Stenotrophomonas sp. (PET_S6), Sporobolomyces roseus (PET_S9), Bullera alba (PET_S10), and Bacillus tropicus (PET_ORAS6)] with PET_C representing the pristine sample. The delta area percentage (ΔA%) is calculated against the control after 30 days.

N° Peak	PET_S5 ΔA%	PET_S6 ΔA%	PET_S9 ΔA%	PET_S10 ΔA%	PET_ORAS6 ΔA%
1	50.66	57.24	31.58	18.42	35.53
2	44.00	44.80	42.40	7.20	32.80
3	57.15	61.16	42.22	24.24	45.91
4	60.57	66.52	44.10	34.22	53.34
5	64.05	64.05	52.81	28.09	47.20
6	64.03	67.31	49.12	32.97	54.72
mean ± standard error	56.74 ± 3.26	60.18 ± 3.42	43.70 ± 2.96	24.19 ± 4.14	44.91 ± 3.68

). Subsequently, the mean percentage reduction in area under the curve for each peak was calculated.

Notably, our findings reveal a consistent reduction in the absorption peaks of all bands following microbial degradation. Among the microorganisms studied, Stenotrophomonas sp. exhibited the most significant efficacy, achieving a 60.18% reduction in the area under the curve, with a standard error of ±3.42 (Table 2). This significant finding indicates the possibility of altering the mass quantity of our samples undergoing degradation, prompting further investigation into potential optimization strategies.

3.3.3. Polystyrene (PS)

The PS spectrum (Figure 1) has dominant peaks at 2926 and 2851 cm⁻¹ from the methylene stretches; out-of-plane C-H bends of the aromatic ring are intense at 697 and 755 cm⁻¹ (mono-substituted); aromatic ring breathing modes appear at 1601, 1493, and 1452 cm⁻¹; C-H wag is present at 1028 and 1068 cm⁻¹; peaks at 3082, 3061, and 3027 cm⁻¹ are absorptions from the aromatic C-H stretches [34]; characteristic bands of C=C stretching of the vinyl group signal are found at 1630–1660 cm⁻¹ [35], particularly in PET samples degraded by Rahnella sp. and Bacillus tropicus. This would indicate a possible change in the polymer chains and structure, as shown in the SEM images acquired of PS, respectively (Figure 4).

The acquired PS samples’ surface images show a seemingly smooth surface at a magnification of 500×, but is instead rough at a magnification of 1400×. Of the five microorganisms with which the sample was subjected to degradation activity, particular degradation activity by Rahnella sp. and Bacillus tropicus was noted. Both of them caused a sort of remodeling of the material with the opening of fenestrations of different sizes, as shown in Figure 4. Rahnella sp. and Bacillus tropicus may exhibit enhanced degradation activity on PS due to their potential to express specific enzymes like esterases or cutinases. These enzymes likely interact with and cleave ester bonds present in PS, facilitating polymer breakdown and structural modification observed in the SEM images. At the moment, the evidence regarding the PS-degrading capabilities of Rahnella sp. and Bacillus tropicus is limited. An interesting study has investigated the biodegradation efficacy of Bacillus tropicus on a low-density polyethylene (LDPE) matrix. The research included experimental methods to assess the biodegradation process, such as FTIR and SEM, also used in this paper. A loss of mechanical properties and weight reduction were among the results observed [36].

3.3.4. Epoxy System 1 (RS1)

RS1 shows peaks at 829 cm⁻¹ associated with the stretching of C-O-C of the oxirane group; 1037 cm⁻¹, stretching of C-O-C of ethers; 1182 cm⁻¹, asymmetrical C-O stretching band of the ester (missing in RS2); 1246 cm⁻¹, asymmetrical aliphatic C-O stretching; 1296 cm⁻¹, asymmetrical CH₂ deformation; 1458, 1508, and 1607 cm⁻¹, C-C stretching vibration in aromatic and N-H bending of primary amine; 1734 cm⁻¹, bending of primary amine; 2871 cm⁻¹, symmetrical C-H stretching of -CH₃ group; 2931 cm⁻¹, asymmetrical C-H stretching of -CH₂ group; 2961 cm⁻¹, asymmetrical C-H stretching of -CH₃ group [34] (Figure 1).

Upon comparing the SEM images of degraded samples with the one of control sample, it is evident that there is minimal degradation activity. The majority of the samples’ surfaces subjected to degradation did not show any consistent changes and appears as smooth as the control sample. On the sample surface subjected to degradative action by Bullera alba, the presence of a concave area, inside which there were fragments and crystals, was noticed. The presence of a concave region, absent in the control sample, and containing crystals within it, marks an important advancement in proving confirmation, although minimal, of the degradation activity performed by Bullera alba (Figure 5).

In epoxy resins, microorganisms could target specific chemical bonds during degradation. These include the oxirane (epoxide) groups (-C-O-C-), ester bonds, aromatic C-C bonds in aromatic rings, and aliphatic C-O bonds. Enzymatic actions on these bonds, indicated by characteristic FTIR peaks, suggest their potential for biodegradation, impacting material properties and recycling strategies.

In the literature, no evidence that directly associates the capability of Bullera alba and Bacillus tropicus to degrade epoxy resin was found. The complexity of identifying microorganisms capable of attacking and degrading epoxy resins is implied, highlighting the challenges in this area. Nevertheless, the findings of this study potentially serve as an initial stage for future research endeavors.

3.3.5. Epoxy System 2 (RS2)

Given that both systems are epoxy-based, RS2 exhibits peaks resembling those of RS1. However, minor differences can be found at 1734 cm⁻¹, representing the bending of the primary amine. As shown on the plot, this absorption band is almost irrelevant in RS2 (Figure 1).

The SEM images of the degraded samples, when compared to the control sample, indicate minimal degradation activity. Observations revealed a subtle alteration on the surface of the sample exposed to degradation by Rahnella sp. This alteration manifested as variations in color intensity, presenting as both lighter and dark patches. Upon examination, surface alterations were observed on the material subjected to degradation by Bacillus tropicus. Notably, at a magnification of 1400×, surface irregularities were evident, resembling clusters of microorganisms. These clusters seemed to elevate the surface, leading to the detachment of material flakes (Figure 5).

In the literature, no evidence that directly associates the capability of Rahnella sp. and Bacillus tropicus to degrade epoxy resin was found. However, the study conducted by Eliaz [37] elucidates the ability of specific microorganisms to utilize epoxy resin as their primary carbon source. By isolating bacteria proficient in metabolizing epoxy resin, the study sheds light on potential avenues for sustainable waste management and recycling strategies. Notably, the utilization of an epoxy resin from the Araldite^® family, similar to the one used in this study, underscores the relevance of this research to industrial applications and environmental concerns surrounding epoxy-based materials.

4. Conclusions

The comprehensive analysis conducted, including DNA sequencing, enzyme assays, and material degradation studies, offers insights into the adaptive capabilities of microorganisms sourced from diverse environments, such us fresh soil and oil refinery activated sludge. The isolated microorganisms demonstrate a remarkable ability to express enzymes that enable them to thrive in environments where thermoplastic and thermosetting polymers serve as the sole carbon sources. By producing specific enzymes such as lipases, esterases, cutinases, and haloalkane dehalogenases, these microorganisms can effectively utilize the polymers as their primary nutrient source. The ATR-FTIR analysis indicates changes in biodegraded compounds, particularly noticeable in PET, where there is a reduction in infrared light absorbance without significant structural shifts, as confirmed by peak positions. This reduction ranges from a percentage of 24.19 ± 4.14 to 60.18 ± 3.42 across various peaks. SEM images reveal fenestrations in both PET and PS, supporting microbial degradation efficacy and potential mass loss. Less pronounced outcomes were observed for epoxy resins, despite microorganisms demonstrating their ability to thrive using them as their sole carbon source. ATR-FTIR analyses revealed a decrease in infrared light absorption in both RS1 and RS2, while SEM images showed little to no alteration in surface morphology. Even modest results will be valuable for advancing future research. Future studies on this topic will incorporate additional parameters such as sample weight to further explore the biodegradation processes. Future research could explore chemical pre-treatments to enhance microbial action on epoxy resins, advancing applications in environmental remediation and sustainable waste management.