Introduction

Plastic pollution, a critical environmental issue in the 21st century, has detrimental consequences for ecosystems and public health. The global projection for plastic waste (PW) indicates a surge from 353 million tonnes in 2019 to 1014 million tonnes by 2060, a challenge exacerbated by inadequate global waste management capabilities [1]. Landfilling and incineration collectively account for approximately 70% of plastic waste disposal, with a mere 9% being effectively recycled in 2019, leaving the remainder to infiltrate unregulated dumpsites, burn in open pits, or contaminate soil and aquatic habitats [2]. Although recycling is one of the best ways for PW treatment compared with the disposal method, most synthetic plastics are derived from hydrocarbon raw materials (e.g. natural gas, crude oil, and plants). Some chemicals, plasticisers, and additives are added to some synthetic plastics to help them be used in specific applications, which makes PW one of the most complex material mixtures from a recycling perspective [3].

Currently, there are two main strategies for PW disposal: landfilling and thermal treatment (incineration, pyrolysis, and gasification). Landfilling is the most common PW disposal method as it is costless and convenient. However, the formation of secondary microplastics (MPs) pollution and gaseous products can cause a significant threat to the environment and climate. Even though thermal treatment could reduce waste volume whilst producing energy, financial and environmental issues limited their wide application.

Biodegradation could overcome the disadvantages of those conventional methods, such as cost and pollution. Plastic biodegradation is the process in which living microorganisms utilise plastic as a carbon source and break organic substances into more minor compounds and residual biomass [4,5,6]. The biodegradation of plastic treatment strategy is the most environmentally friendly way due to creation of less secondary pollutants compared to other disposal methods. Moreover, biodegradation can be optimised and be employed in large-scale operations in the form of reactor systems, which are already extensively used to treat organic wastes [7]. Numerous studies have identified microorganisms in various environments with the capacity to degrade plastic [8, 9]. However, the slow degradation of synthetic plastics in nature is influenced by physicochemical properties and environmental factors [10, 11]. The microbial decomposition of plastic involves five key processes: colonisation, biodeterioration, biofragmentation, absorption, and mineralization [12]. Plastics are relatively large molecule's size, hydrophobic and have stable, functional groups, such as phenyl and alkane groups, which resist microbial colonisation and prevent transportation into the cells [13, 14]. Additionally, the absence of favourable functional groups for oxidative reactions further hampers microbial growth on conventional plastics [15, 16].

Efforts to enhance plastic biodegradation have explored various pretreatments [17, 18]. Studies suggest that physicochemical pretreatments, including UV, thermo-oxidative, and chemical treatments, can induce surface oxidation, creating carbonyl, carboxyl, and ester functional groups [19, 20], thereby reducing surface hydrophobicity and promoting microbial biofilm formation, thus improving biodegradation efficiency [21, 22]. Additionally, gamma irradiation and heat treatment have been shown to enhance the degradation of certain plastics. Novotný et al. (2018) show the presence of an aldehyde. Ketone, ether or ester groups formed after gamma-irradiated cum heat-treated PE and degraded better than the untreated sample. Taghavi et al. (2021) found that UV pretreatment of PE and PS results in a higher roughness, lower hydrophobicity, higher biofilm colonisation and a more significant physical and molecular weight reduction compared to untreated samples. Chemical pretreatment of plastic is efficient at hydrolysing the material and reducing its deterioration rate. However, chemical pretreatments raise environmental concerns due to chemical usage, cost, and the recovery of chemicals post-treatment [23]. Present pretreatment techniques were not effective enough in enhancing degrading efficiency. Thus, there is a need for more sustainable and effective technologies as a pretreatment method before biodegradation.

Cold atmospheric plasma (CAP) application is a practical, cheap, and environmentally friendly technology for surface modification of polymeric materials. CAP treatment can alter surface chemical compositions and physical properties without affecting the bulk properties of polymers [24]. Previous research demonstrated that plasma treatment leads to hydrogen abstraction and surface etching and introduces chemically reactive functional groups or free radicals at the polymer surface. The chemical changes, as well as the surface roughening resulting, showed significant improvement in surface hydrophilicity [25, 26], cell adhesion and proliferation [27, 28]. Based on the mechanisms of plasma irradiation, it is believed that CAP can improve microbial colonisation and degradation efficiency by increasing plastic surface hydrophilicity and inducing polymer oxidation [29].

This study provides an innovative eco-friendly way for PW disposal. For the first time, the effect of CAP as a pretreatment on microbial colonisation and biodegradation of thermoplastics was investigated. Two fungal strains, Aspergillus flavus and Penicillium glaucoroseum were selected for this study. The choice was based on a proven ability to degrade PS and PE [30], the ability to survive in harsh environments with low nutrient and moisture levels and a greater source of hydrolytic enzymes compared with bacteria [31]. The strains were isolated from soil, worm excreta, and active sludge, based on their capacity for microbial colonisation and biofilm development on plastic film surfaces [30]. High-density polyethylene (HDPE) and PS, both untreated and CAP-pre-treated, were incubated with these microorganisms for 50 days, with various analytical methods applied to assess the impact of CAP on the physical and chemical properties of the polymers.

Materials and methods

Materials

The chemicals used in this study were KH2PO4, K2HPO4, MgSO4·7H2O, NH4NO3, NaCl, FeSO4·7H2O, MnSO4·H2O, crystal violet (CV), and potato dextrose broth (PDB). These compounds are of analytical quality and were purchased from Sigma-Aldrich (New Zealand). The stock cultures (A. flavus and P. glaucoroseum) were preserved at 4 °C in potato dextrose agar (PDA), and the mix cultures suspension was prepared according to standard ASTM G21-15 [32]. Argon gas (purity ≥ 99.9%) was used for plasma generation.

White HDPE film (the shopping bag) with 0.12 mm thickness and white PS foam with 2 mm thickness (one side glaze and one side mate (drinking cup)) were purchased locally. The proximate and ultimate analyses of PS and HDPE samples are shown in Table 1. The proximate analysis of PS and HDPE was carried out according to ASTM D3173-75 and the ultimate analysis was performed using a Vario EL cube elemental analyser (Elementary Analysensysteme GmbH, Germany).

Table 1 Proximate and ultimate analysis results of PS and HDPE
Full size table

The plastic samples were cut into 15 × 15 mm square films (96 pieces). Before incubation, these films were immersed in 75% (v/v) ethanol solution for 30 min, washed with sterilised Milli-Q water, and oven-dried at 50 °C overnight in clean Petri dishes with lid on to avoid any airborne microbial contamination. The Petri dishes were not wrapped with Parafilm to allow moisture to escape. A control group was carried out to eliminate the possibility of migration of plastic additives into the ethanol solution during the immersing step before and at the end of the incubation period.

Media preparations

Nutrient-salts broth (NSB) was prepared according to ASTM G21-15 (Standard Practice for Determining Resistance of Synthetic Polymeric Materials to Fungi) protocol [33] and the quantities of materials are shown in Table 2. The test medium was sterilised by autoclaving at 121 °C for 20 min, and the media pH after sterilisation was measured. The original pH was measured using leftover media after adding media to the experiment flasks to avoid potential contamination from the pH metre to the media. Other nutrient-rich media potato dextrose agar (PDA) and potato dextrose broth (PDB) were prepared according to the manufacturer's (Sigma-Aldrich) descriptions.

Table 2 Composition of nutrient-salts broth (NSB)
Full size table

Cold plasma-pretreatment

The HDPE and the PS films were exposed to a low-power, atmospheric-pressure plasma by PlasmaTact (PM01-15AR0) using argon gas and microwave energy. Based on the previous work, the test condition was performed under three different exposure times (the 90, 120 and 180 s) with two distances (1 and 2 cm) [26, 34,35,36]. The applied microwave energy and argon gas flow rate were 15 W and 2.5 standard litres per minute (SLM), respectively. After completion, treated samples were analysed for weight reduction, hydrophobicity changes and chemical transformation to determine the preferred conditions for the biodegradation experiment. The goal of pretreatment was to decrease the surface hydrophobicity and induce free radicals. The samples treated with 120 s exposure time and 1 cm distances from the torch showed the most significant reduction of water contact angle and chemical changes. Therefore, PE and PS samples with CAP treated time of 120 s and 1 cm distances from the torch were selected for the biodegradation experiment. The cold atmospheric plasma pretreatment and biological treatment conditions are presented in Table 3.

Table 3 Cold atmospheric plasma pretreatment and biological treatment conditions
Full size table

Biodegradation

Three sets of controls (Table 3) were employed in this work to distinguish between the impact of cold plasma and biodegradation. Control-1 contained CAP-pretreated plastic samples incubated in NSB without inoculation. Control-2 contained untreated plastic samples incubated in NSB with strain present and Control-3 contained non-inoculated NSB with untreated plastic samples. The microorganism inoculum was enriched individually in 15 mL sterilised fresh potato dextrose broth (PDB) at 26 °C for 24 h. After 24 h of enrichment, the microbial suspension was removed and centrifugated at 5000 × g (8 min).

For media adaptation, the mixed microbial cultures were suspended in NSB for 2 h. The suspension was centrifuged under identical conditions, and the pellets produced were combined to create a mixed microbial suspension (co-culture). The co-culture was vortexed and added into 250 mL flasks, each flask containing 75 mL NSB with a 5 mL of microbial co-culture. Upon incubation, six pieces of each plastic type were added to each 250 mL flask and incubated at 30 °C at 90 rpm for 50 days. The biodegradation was conducted according to the conditions set out in Table 3. All experiments were duplicated under the same conditions. The results were then contrasted and shown as mean ± standard deviation.

Analyses

Different techniques were used to assess the changes in the treated polymer's physical, chemical, and surface properties. Weight, wettability, chemical structure and media pH were examined before and after the CAP pretreatment and biodegradation. Surface roughness and morphology analysis using atomic force microscopic (AFM) and scanning electron microscope (SEM) were also performed on the samples after biodegradation.

Biofilm formation and media pH variation

The crystal violet (CV) assay was used as an indirect method for assessing the biofilm formation on the plastic's surface, and the media's pH was measured before and at the end of the incubation period.

After 50 days of incubation, the plastic samples were removed from each flask and washed gently with sterilised Milli-Q water to remove unattached cells. Simultaneously, the pH of the media was measured. For biofilm measurement, plastic films were immersed into 3 mL 1% (w/v) CV for 15 min. Upon completion of the immersion, the samples were gently washed with Milli-Q water to remove the excess stain on the surface. After washing, samples were air-dried for 30 min with the plastic films placed into 46 well plates individually. Plastic samples were then air-dried (30 min), and the formed biofilm was extracted by adding 0.5 mL 95% (v/v) ethanol in 46 well-plates containing the dried plastic sample. After that, the absorbance was read by a microplate reader at 595 nm (PerkinElmer, MLD2300, USA).

All readings were repeated twice, and results were expressed as mean ± standard deviation.

Physical weight loss

The physical weight loss of the polymer directly measures the degradation of the polymer. After biofilm measurement, samples were rewashed with distilled water to remove the extract excess stain, followed by immersion in sodium dodecyl sulphate (SDS) solution (2% w/v) for 24 h and further washed with distilled water. The cleaned samples were then oven dried at clean petri dishes with no lid at 55 °C for 24 h, and then a 6-digit balance examined weight loss. All measurements were repeated three times, with the results expressed as mean ± standard deviation. The percentage of the weight loss Eq. (1) was calculated below [37]:

$${\text{WeightLoss}} \left(\%\right)=(\frac{{{W}_{{\text{i}}}-W}_{{\text{e}}}}{{W}_{{\text{i}}}})\times 100$$
(1)

where Wi is the sample's initial dried weight (g), and We is the weight (g) of the sample after being cleaned and dried.

Hydrophobicity

Hydrophobicity was determined using a contact angle analyser (Attension Theta Flex Optical Tensiometer, Biolin Scientific, Sweden). Each plastic sample was tested by dropping a droplet of distilled water (4 µL) on its flat surface. The static angle was measured based on Young's equation which assumes that interfacial forces are thermodynamically stable. Each plastic sample was examined at three different sites, and the results were expressed as mean ± standard deviation.

FT-IR spectroscopy

Structural modifications and chemical transformations of plastic samples were examined by FT-IR (PerkinElmer, USA). The spectra were scanned on both sides at a resolution of 4 cm−1 in the frequency range of 4000–400 cm−1.

Surface morphology and roughness

The surface topography of the polymer was investigated with an atomic force microscope (AFM, Asylum Cypher- ES, USA) and scanning electron microscope (SEM) (FEI Quanta 200f, USA). For AFM analysis, samples were mounted on a 15 mm diameter AFM specimen disc and scanned at a 5 × 5 µm scale. For SEM analysis, the same samples were sputter-coated with platinum (Quorum, Q150RS) and were scanned under low vacuum at 10 kV. The images taken after AFM microscopy were analysed by Gwyddion software (Czech Metrology Institute) for surface roughness analysis.

Statistical analyses

To ascertain the statistical significance of the effect of pretreatments on the observed differences in the results. Two-way ANOVA was conducted using SPSS software on all the data.

Results and discussion

Biofilm formation and media pH variation

Biofilms are a primary form of microbial growth. Cells that are affiliated with biofilms are better able to modify plastic surfaces and use them as a food source [27, 38]. A biofilm is a collection of microbial cells encased in a polysaccharide-based matrix and permanently attached to a surface, which cannot be removed through mild washing. The solid plastic surface and the aqueous medium (NSB) interface provide an ideal environment for the attachment and growth of microorganisms [39]. The properties of the media, including pH, nutrient levels, and temperature, as well as the substrate's roughness and wettability, influence the intricate attachment process [40]. Two isolated fungus species, A. flavus and P. glaucoroseum were incubated in NSB media with PS and PE as sole carbon substrates. Noticeably, visible colonisation started to appear on PS and PE surfaces after 5 days, as the mycelial growth enabled rapid substrate colonisation when the fungus was isolated [20].

Moreover, PS had higher optical density reading (OD 595 nm) than PE regardless of treatment type, which agrees with Taghavi et al. [41, 42]. The higher OD reading may be due to the polystyrene foam structure, as PS comprises numerous similar-shaped beads, each compactly positioned adjacent to the previous one. Some CV dye may sink inside the gaps between beads. The CV dye on the PE and PS surfaces washed away quickly. Nevertheless, the results (Fig. 1). provided strong evidence that CAP-pre-treated plastic samples (T1) exhibited more significant microbial colonisation than untreated plastics for both PS (p < 0.05) and PE (p < 0.005). Therefore, the higher biofilm reading was not due to the more trapped stain amongst PS structure. Indeed, it was for higher microbial colonisation there. The formed biofilm at T1, C1, C2, and C3 for PS foam were 0.56, 0.29, 0.44, 0.19 at OD 595 nm, and for PE film, were 0.19, 0.11, 0.15, 0.07 at OD 595 nm, respectively.

Fig. 1
figure 1

Quantification of biofilm formation on PS and PE within 50 days. The asterisk brackets indicate significant differences between samples

Full size image

Moreover, compared to the pH of the control set, a reduction in solution pH was observed for T1 samples (from pH 6.8 to pH 6.63). No pH change was observed in control samples (C1, C2, and C3). The degradation of plastics by fungus involves the action of microbial enzymes on the surface of the plastics. A lowering in the pH value results from the secretion of enzymes and acids from the microorganisms [39]. The reduction in pH suggested that the microbial trains incubated with CAP-treated plastics performed higher enzyme secretion and were metabolically active.

Weight loss

Analysis of PE samples T1 (5.1 ± 0.6%) and C2 (4.85 ± 0.25%) showed statistically significant higher gravimetric weight loss compared with strains-free samples C1 (3.2 ± 0.3%) and C3 (2.75 ± 0.35%). The results indicated that A. flavus and P. glaucoroseum have the ability to degrade PE. In contrast, the PS sample's weight reduction was insignificant (Fig. 2). The fragmentation of plastic films during biological treatment or residues of microbial biofilm that were firmly adhered to the plastic substrate material occasionally reduced the precision of the gravimetric approach. The microbial debris and colonisation inside the PS bead cavities would have also contributed to weight gain instead of weight loss, similar behaviour also was seen in another study by Taghavi et al. 2021. Additionally, as PS has an aromatic ring in the chemical structure, it takes more enzyme-mediated oxidation steps to open the aromatic ring and create acetyl-Co A or succinate for the TCA cycle and PHA for energy generation [40, 43]. Therefore, compared to PE, their biodegradation rates may be slower due to their more complex enzyme systems [44].

Fig. 2
figure 2

Physical weight loss after biodegradation of PE and PS. The asterisk brackets indicate significant differences between samples

Full size image

The synergistic effects of CAP pretreatment and biodegradation of plastic samples (condition C3 and T1) showed a significant difference (p < 0.005) in weight reduction for PE. Also, the trend of biofilm formation and weight reduction are proportional. Increased biofilm formation correlates to increased weight loss. These relationships may be explained by the biofilm-associated cells colonising the plastic surface by generating an extracellular polymeric substance (EPS) matrix. The EPS matrix mainly consists of polysaccharides forming a sheath bonded to the polymer. This helps transport the depolymerising enzymes to its surface. Therefore, more biofilm formation on the surface could enhance the weight reduction rate.

Wettability change

The surface hydrophobicity can be determined by its contact angle, θ (theta). A contact angle of ≥ 90° is considered hydrophobic, whilst 0° ≤ θ ≤ 90° is said to be hydrophilic, and θ ≥ 150° becomes superhydrophobic [45]. The hydrophobic polymer surface vigorously protects the plastic from the effect of enzymes. Therefore, one of the primary purposes of using cold plasma as a pretreatment method is to increase the plastic surface hydrophilicity. The pristine PE and PS films (Fig. 1c) showed hydrophobic properties with a WCA of 95° (± 4° SD) and 104° (± 1° SD), whereas the CAP pretreated PE and PE films had a more hydrophilic surface with a mean CA of 78 (± 2° SD), and 91 (± 2° SD). An Oxygen containing group (C–O) was incorporated after CAP treatment onto the polystyrene surface (Fig. 3). The water contact angle is reduced as oxygen content rises with additional polar groups, leading to increased polarity, surface tension, and surface free energy, resulting in increased hydrophilicity and wettability of the film surface. This is consistent with previous studies [35, 46, 47]. Another likely cause of increasing hydrophilicity for the PE sample is the surface roughness brought on by the etching process, which helps the liquid spread across the surface rather than adding free radicals to the structure [41].

Fig. 3
figure 3

Water contact angle variation of PE and PS before and after biodegradation. The asterisk brackets indicate significant differences between samples

Full size image

Furthermore, after biodegradation, both plastic types were more hydrophilic at the end of the experiment than the pristine plastic samples. In every instance, the surfaces had become more hydrophilic due to exposure and the attachment of microorganisms to the polymer surface, followed by surface degradation.

FT-IR analysis

The FT-IR analysis showed that regardless of the pretreatment condition, the spectra pattern of the PE sample before biodegradation and after biodegradation was almost the same (Fig. 4). However, C2 observed a new peak attributed to alkoxy groups (C–O) at 1000–1050 cm−1 after biodegradation, which could suggest the biodegradation of HDPE samples [18, 44]. The results are in agreement with other study where C–O chemical bond formed after the biodegradation [30]. In PS spectra, whilst there is a lack of alkoxy group (C–O) at 1200–1275 cm−1 in C2 and C3 conditions, this functional group detected in T1 and C1 revealed the interaction between oxygen-containing species in the plasma and the substrate. During the CAP treatment, plasma caused the scission of R–H and C–C bonds [42]. At or near the surface, hydrogen atoms are removed by inert gas ions and ground-state atoms, leaving behind metastable polymer radicals that form the active sites on the polystyrene surface [48]. PS readily incorporated oxygen due to the aromatic ring structures in the polymer chains [49]. The results from this study agree with the previous reports [50,51,52]. When the treated polymer exposes to the laboratory atmosphere, ambient oxygen can interact with the active sites and bond to them to form the hydrophilic functional groups, like ether or alcohol C–O and carbonyl C = O.

Fig. 4
figure 4

FT-IR spectra of pretreated sample and control set, PS (a), PE (b) after 50 days of biodegradation

Full size image

Surface morphology and roughness analysis

Atomic force microscope (AFM) and Scanning electron microscope (SEM) analysis revealed the possible surface degradation of plastic samples after CAP pretreatment and biodegradation. The surface roughness of each condition is compared to C2 to assess the effectiveness of both pretreatment and biodegradation. The intention was to highlight the changes in surface roughness as a measure of the combined effects of cold plasma pretreatment and biodegradation. It was found that the surface roughness was changed after CAP and biological treatment for both PE and PS samples (Fig. 5). Comparing C1 to C3, the average surface roughness (Ra) profile indicated an increased surface roughness for both PE and PS after CAP treatment. After plasma treatment, the topology and roughness of the virgin PE films' smooth surfaces were altered. This observation confirmed that CAP pretreatment increases PE surface roughness by generating pits and pores. After 50 days of biodegradation with microbes, biological treatment C2 samples showed considerable increases in the roughness of more than 32 and 36% for PE and PS compared to C3, respectively. Whilst the addition of CAP (over and above biological treatment) further increased the roughness compared to biological treatment alone (C2). The increased surface roughness was calculated to be 10 and 35% for PE and PS, respectively.

Fig. 5
figure 5

Roughness analysis of PE and PS after 50 days of incubation compares the control with CAP pretreated samples

Full size image

The highest surface degradation in the "CAP pretreated & biotreated (T1)" samples (Figs. 6g, 7g) was observed, compared with the other three control sets (Figs. 6a, c, e, and 7a, c, e). The maximum distance measured from the crest to the trough was 0.22 and 0.15 μm for PE and PS samples, respectively, in the "CAP-pretreated & bio-treated" samples (Figs. 6g, 7g). Consequently, it was concluded that although CAP treatment might alter surface morphology and promote surface deterioration, microbes were necessary for further surface breakdown and degradation.

Fig. 6
figure 6

AFM images and roughness profile of PE after 50 days of incubation under various conditions. a, b C1, CAP pretreated PE without strains present c, d C2, Unpretreated sample with strains present e, f C3, blank g, h T1, CAP treated PE with strains present

Full size image
Fig. 7
figure 7

AFM images and roughness profile of PS after 50 days of incubation under various conditions. a, b C1, CAP pretreated PS without strains present c, d C2, Unpretreated sample with strains present e, f C3, blank g, h T1, CAP treated PS with strains present

Full size image

The changes in surface morphology of the HDPE and PS films were examined by SEM analysis (Fig. 8). Examining untreated PE foam (Fig. 8e, f) revealed that it was formed of numerous beads with smooth surfaces. Even though no apparent surface alteration of PS was observed under SEM, argon has reportedly been shown to be a potent gas for physically etching the surface through intense ion bombardment [51]. The active species produced from argon plasma cause etching, which also removes low molecular weight impurities from the surface and causes enhanced plastic roughness [52]. Microbial adherence to the PE surface during fungal growth deteriorated the sample surface (Fig. 8g, h). This result is in agreement with Bonhomme et al. 2003 [53], where SEM imaging captured evidence of the surface being physically cracked and degraded, and several significant cavities were formed on the incubated PE films. The SEM results found a significant surface degradation and loss of boundaries between PS beads. The biotreated films became degraded, whereas the untreated film retained a smooth surface. According to Characklis et al. [54], the amount of microbial colonisation tends to increase with surface roughness rises as there is an associated reduction in shear forces present, as well as an increase in surface area for rougher surfaces. Based on the AFM and SEM results (Figs. 5, 6), cold plasma can alter surface morphology and help surface deterioration and microbes' colonisation.

Fig. 8
figure 8

SEM analysis of PE (left) under 50 µm scale and 10 µm scale and PS (right) under 200µm scale and 50µm scale after 50 days of incubation under various conditions. a, b, i, j C1,CAP pretreated sample without strains present c, d, k, l C2, unpretreated sample with strains present e, f, m, n C3, blank g, h, o, p T1, CAP treated sample with strains present

Full size image

Conclusion

The current study demonstrates that cold atmospheric plasma pretreated on HDPE and PS increases surface roughness and hydrophilicity by generating active species and physically etching the sample surfaces, increasing surface roughness. As evidenced by contact angle measurements, improved roughness increases surface area, leading to increased surface energy. Furthermore, FT-IR analysis indicated the formation of additional alkoxy functional groups on the PS surface, suggesting cold plasma may break the polymer chains, creating free radicals, which combine with oxygen in the air and facilitates ongoing biodegradation of the polymer.

Consequently, after being incubated with microorganisms, cold plasma pretreated samples had a higher weight loss, surface degradation, biofilm formation and hydrophilicity. Based on surface deterioration (microscopic) in this study, a maximum of 40% surface degradation was observed in the CAP + biotreated HDPE compared with virgin samples, with an increased biodegradation efficiency of 10% to biotreated HDPE. The CAP + biotreated PS condition results in a maximum surface degradation of 58% compared with virgin samples and an increased biodegradation efficiency of 35% compared to biotreated PS.

Overall, the findings of this study support the hypothesis that cold plasma is a suitable technique for improving biodegradation efficiency by changing the surface characteristics of polymeric HDPE and PS.