A Foundation for Advancing Studies of the Biodegradation of Polyethylene Surrogates by Environmental and Model Laboratory Microbes

ABSTRACT

Polyethylene represents a particularly recalcitrant class of plastics that persist for decades in the natural environment when released as the result of failed waste management policies. In this report, we present a detailed survey of microbes with varying abilities to degrade either branched or linear waxy hydrocarbons that serve as a surrogate for the study of polyethylene biodegradation. This analysis includes measurement of the degree of branching for the surrogates. We further monitored the growth of individual isolates as an indication of substrate preference. We sequenced the genomes for each of our isolates that showed significant rates of growth to accommodate future biochemical studies, and provide a general characterisation of each strain. The vast majority of microbes that we isolated and identified as part of this study were Actinomycetes. However, a small selection of gram‐negative microbes were identified that resulted in degradation of the surrogates. Importantly, our results further identified the model microbes Acinetobacter baylyi and Rhodococcus jostii as strains that were particularly good at degrading all three of the model polyethylene surrogates employed in this study. The results of this study should serve as a detailed genetic and biochemical resource to the research community investigating polyethylene biodegradation.

A polyethylene surrogate sample containing paraffin, polywax and polyethylene was exposed to selected microbes identified in this study. This image shows the surface of this surrogate following a six‐week incubation with Rhodococcus jostii .

1. Introduction

Plastics are synthetic polymers primarily derived from non‐renewable fossil fuels, and were first mass‐produced in the nineteenth century. Their durability, lightweight nature and high malleability make them an ideal material for various industrial applications, ranging from packaging to agriculture, and they are now a staple of everyday living. The relatively low cost and ease of production have further contributed to their extensive use across diverse applications (Andrady and Neal 2009; Biron 2016; Geyer et al. 2017). However, the same properties that make plastics highly versatile also make them exceptionally recalcitrant to degradation. Nearly 77% of global plastics production consists of polymers with carbon–carbon backbones, which include polyethylene (PE), polypropylene (PP) and polystyrene (PS). These polymers are particularly challenging to degrade due to their chemically inert structures. It is projected that approximately 12 billion metric tons of plastic waste will have accumulated on Earth by 2050 (Geyer et al. 2017; Law and Narayan 2022).

Plastics are significant pollutants in oceans, freshwater systems, and even the air, where they persist due to their very slow degradation. In the environment, larger plastic waste fragments into microplastics under the influence of UV radiation and mechanical forces (Andrady 2011; Thompson et al. 2009). These particles infiltrate ecosystems, disrupting marine life and entering food webs. Microplastics also originate from primary sources like synthetic textiles, cosmetic microbeads and tire wear (Ivar do Sul et al. 2014; John et al. 2023; Möhlenkamp et al. 2018; Musa et al. 2024). Plastic waste is currently managed using various physical and chemical methods. Incineration generates energy but can release harmful pollutants, while landfilling, though widely used, can cause environmental issues like leachate contamination and greenhouse gas emissions (Monni 2012; Siddiqua et al. 2022). Pyrolysis breaks plastics into solid, liquid or gaseous fuels through high‐temperature processing in the absence of oxygen (Jeswani et al. 2021; Kabeyi and Olanrewaju 2023). Mechanical recycling involves shredding and repurposing plastics into new products, and chemical recycling uses chemical processes to break certain plastics into monomers for reuse. Despite these efforts, challenges like inefficiency, high costs, the requirement of harsh chemicals and harmful gas emissions hinder the effective management of plastic waste through abiotic methods (Jarosz et al. 2024; Jiang et al. 2022).

When all other forms of waste management have failed, biodegradation of plastics becomes the last defence against the persistence of these commodity products when introduced into the natural environment. Understanding the true duration that will be required for biological processes to fully degrade these materials back to simple compounds that can be recycled into the global carbon cycle becomes difficult because experiments to test these processes take so long to perform (Albertsson et al. 1987; Alshehrei 2017). Although most plastics are generally regarded as non‐biodegradable, recent studies have demonstrated the potential of specific microbes and their enzymes to degrade plastics (Bahl et al. 2021).

Over the past few years, our laboratory sampled and enriched various communities of microbes using a polyethylene surrogate (PE‐surrogate) cocktail. Through this process and prior work, we isolated a variety of microbes from these communities and tested them for the ability to degrade these pseudo‐polymers as individual isolates. In this current work, we grew individual strains with a selection of PE‐surrogate components with various degrees of chain length and branching. We further sequenced the genomes of each of these strains and provide a table of growth capabilities along with these sequences to serve as a resource for future work to further characterise the aspects of these microbes that enable them to degrade solid‐state hydrocarbons which share features of polyethylene.

2. Methods

2.1. Sample Collection and Enrichment

Bacterial strains were isolated from diverse environmental sources across Minnesota between 2018 and 2023. Freshwater samples were obtained from urban lake systems using a custom‐designed sampling apparatus optimised for littoral zone collection (Camur et al. 2024). To maintain sample viability, specimens were preserved in sterile collection bags with sufficient native water to prevent desiccation. Soil samples were collected from three distinct ecological zones in Minnesota: agricultural soils (0–15 cm depth), residential garden plots and oil‐contaminated sites. Additionally, plastic wastes encountered during sampling were also collected and used for microbial enrichment.

For enrichment studies, modified Bristol's medium (Bold 1949; Knutson et al. 2018) adjusted to pH 7.6 was supplemented with three distinct hydrocarbon substrates serving as PE‐surrogates and sole carbon sources: low‐melting paraffin (Sigma‐Aldrich P/N: 327204, melting point: 53°C–57°C), high‐melting paraffin (Sigma‐Aldrich P/N: 411663, melting point: ≥ 65°C) and Polywax500 (NuCera, melting point: ≥ 88°C). The medium composition and enrichment protocol were adapted from a previously established methodology (Camur et al. 2024). Cultures were incubated at 30°C with agitation at 180 rpm.

2.2. Isolation and Identification of Microorganisms

Bacterial isolates were obtained through cultivation on selective media, including lysogeny broth (LB), nitrogen‐supplemented and nitrogen‐deficient Burk's medium and Reasoner's 2A medium. All media were solidified with bacteriological agar for plate preparation. Chemical reagents were sourced from Sigma‐Aldrich (St. Louis, MO) and Fisher Scientific (Pittsburgh, PA) unless otherwise stated. Enrichment cultures were serially diluted before plating onto selective media. Colony selection was based on distinct morphological features such as shape, margin definition, elevation, pigmentation and surface texture.

Taxonomic identification was performed through 16S rRNA gene amplification using universal primers: forward (5′‐GAACGCTGGC GGCRKGCYTW AYACATGC‐3′) and reverse (5′‐TACGGYTACC TTGTTACGAC TTM‐3′). Individual colonies were suspended in 100 μL deionised water for use in colony PCR. PCR amplification was conducted in 30 μL reaction volumes containing 15 μL of 2× Buffer G (FailSafe PCR reagent), 10 pmol of each primer, 0.5 μL FailSafe enzyme mix and 1 μL cell suspension, then brought to 30 μL with distilled water. Thermal cycling parameters consisted of initial denaturation at 95°C for 2 min, followed by 35 cycles of denaturation (95°C, 30 s), annealing (60°C, 45 s) and extension (68°C, 3 min), concluding with a final extension at 68°C for 2 min. Amplification products were visualised by electrophoresis using 0.5% (w/v) agarose gels in TBE buffer. PCR products were purified using the DNA Clean and Concentrator kit (Zymo Research, Cat. No. D4033) and subjected to Sanger sequencing (ACGT Inc). Taxonomic assignments were determined through BLAST analysis of the sequences against NCBI reference databases.

2.3. Characterisation of Paraffin and Polywax Surrogates

For gas chromatography (GC) analysis, 50 mg of each PE‐surrogate was individually extracted using a solvent mixture comprising methylene chloride, tetrahydrofuran and hexane in equal parts (1:1:1) (Barney et al. 2012). The extraction process involved sonication with a Misonix XL‐2000 for 60 s, repeated three times with a total of 15 mL of the solvent mixture.

The extracts were treated with N‐Methyl‐N‐(trimethylsilyl) trifluoroacetamide (MSTFA) to prepare them for analysis. Gas chromatography was conducted by injecting 1 μL of the treated extract into the GC system. A flame ionisation detector (FID) was used for GC/FID analysis due to its superior sensitivity compared to mass spectrometry (MS). A high‐temperature RTX‐Biodiesel TG column (15 m, 0.32 mm inner diameter, 0.10 μm film thickness; Restek, Bellefonte, PA) was employed, along with a Shimadzu GC‐2010 gas chromatograph equipped with a programmed temperature vaporisation (PTV) injector. The carrier gas was helium, flowing at a linear velocity of 20 cm/s. The temperature program began at 60°C, held for 1 min, followed by a ramp of 10°C/min to 370°C, which was maintained for 15 min (Barney et al. 2012). This method enabled effective separation and detection of hydrocarbon components in the PE‐surrogates.

Branching in the PE‐surrogates was quantified using nuclear magnetic resonance (NMR) (Jung et al. 2016). The analysis was performed on a Bruker 700‐MHz Avance or Bruker 850‐MHz Avance III spectrometer, both equipped with 5‐mm TXI probes with z‐gradient capability. Samples of wax (10–50 mg) were dissolved in 1 mL of CDCl₃ with tetramethylsilane (TMS) as the internal reference. All experiments were conducted at room temperature, except for Polywax 500, which required a temperature of 40°C due to its partial solubility at room temperature.

Peaks of the branches of 1–6 carbons in the waxes were identified using a proton–carbon heteronuclear multiple quantum coherence (¹H–¹³C HMQC) pulse program. A homodecoupling pulse program at 1.319 MHz was applied to decouple protons in the 0.9 MHz region, isolating branch methyl peaks from the main carbon chain methyl peaks and improving integration accuracy. The degree of branching was calculated by normalising the main chain methyl peak to six protons. Integration of the branch methyl peaks was divided by three, yielding the percentage of branching relative to the main carbon chain (Jung et al. 2016). This approach provided a reliable measure of the branching density within the PE‐surrogates.

For scanning electron microscopy imaging, a PE‐surrogate mixture containing 25 mg of low‐melting paraffin, 25 mg of high‐melting paraffin, 25 mg of Polywax500, 25 mg of Polywax1000 (Nucera) and 50 mg of polyethylene (Average M_W ~4000, P/N 427772, Sigma‐Aldrich) were combined, and autoclaved. The mixture was heated to 130°C and coated on the interior surface of an Erlenmeyer flask and cultured with growth media for a period of 6 weeks. After 6 weeks of incubation, several plastic flakes that had separated from the glass surface were retrieved and mounted onto microscope slides. Samples were fixed with 2.5% glutaraldehyde in 0.05 M cacodylate buffer for 1.5 h at 4°C. Following fixation, samples were washed with the same buffer, post‐fixed with osmium tetroxide and dehydrated through a graded ethanol series (50%, 70%, 95% and 100%). Dehydrated samples were sputter‐coated with a gold layer (Edwards S150B) and imaged using a Hitachi SU3800 scanning electron microscope. Sample preparation and imaging were performed at the UMN Imaging Center.

2.4. Genomic and Phylogenetic Analysis

Genomic DNA was extracted using the Quick‐DNA Fungal/Bacterial Kit (Zymo Research, Cat. No. D6005) following the manufacturer's protocol. Whole‐genome sequencing was performed using various sequencing platforms depending on the sample. Strains PSBB011, PSBB049 and PSBB011 were sequenced using the PacBio RS II system, with de novo assembly conducted using Flye v.2.6 (Kolmogorov et al. 2019). Strains PDNC001, PDNC005, PDNC012 and PDNC014 were sequenced using PacBio Sequel II, with assembly also performed using Flye v.2.6 (Kolmogorov et al. 2019). Strain PSC013 was processed using the PacBio Sequel II platform, employing Hifiasm v.0.16 (Cheng et al. 2021) for assembly to optimise contig generation and genome completeness. The remaining strains PDNC009, PDNC017, PDNC021, PDNC024, PSBB026 and PSBB082 were sequenced using Oxford Nanopore technology and assembled using Flye v.2.6 (Kolmogorov et al. 2019).

Whole‐genome sequences were employed to construct a phylogenetic tree, offering insights into the evolutionary relationships among the bacterial strains. Phylogenetic analysis was conducted using the BV‐BRC Bacterial Genome Tree service (Olson et al. 2023). Protein sequences were aligned using MUSCLE (Edgar 2004), while nucleotide coding genes were aligned using BioPython's Codon_align function (Cock et al. 2009). A concatenated alignment of all proteins and nucleotides was written to a PHYLIP‐formatted file, followed by the generation of a partitions file for RAxML (Stamatakis 2014). This partitions file described the alignment in terms of protein sequences and the first, second and third codon positions. Support values were calculated using 100 iterations of RAxML's ‘Rapid’ bootstrapping option (Stamatakis et al. 2008), ensuring robust statistical confidence in the phylogenetic tree. The resulting phylogenetic tree was visualised using iTOL v6 (Letunic and Bork 2024).

2.5. Growth Studies

For growth measurements, 46 mg of each PE‐surrogate was added to 125 mL Erlenmeyer flasks separately as the sole carbon source. PE‐surrogates included low‐melt paraffin, high‐melt paraffin and Polywax500. The flasks were autoclaved and subsequently heated to 120°C to melt the PE‐surrogates. While still hot, the PE‐surrogates were evenly coated across the liquid‐accessible surface of the glassware up to the 50 mL mark by rotating the flask to ensure uniform distribution. Cultures were inoculated with 50 mL of modified Bristol's medium and incubated at 30°C with agitation at 180 rpm. Growth was monitored for 2 weeks in the cases of low‐ and high‐melt paraffin and for 5 weeks with Polywax500. Optical density (OD) measurements were recorded at 600 nm at ~3‐day intervals using a spectrophotometer to assess bacterial growth. In addition to strains isolated during this study, several common laboratory bacterial strains were sourced from established repositories and collaborators to provide a diverse set of organisms for comparative analysis. Marinobacter aquaeolei VT8, Acinetobacter baylyi ADP1, Azotobacter vinelandii DJ, Bacillus subtilis subsp. subtilis 168 and Gluconacetobacter diazotrophicus PA15 were acquired from the American Type Culture Collection (ATCC). Additionally, Klebsiella sp. M5al was kindly provided by Ruth Schmitz‐Streit and Corynebacterium glutamicum ATCC 13032 was kindly provided by Claudia Schmidt‐Dannert. These strains were selected for their reported or potential hydrocarbon‐degrading capabilities or as controls representing common model organisms for non‐oil‐degrading bacteria.

To investigate the potential to degrade true polyethylene, a selection of strains was selected to test with a low molecular weight polyethylene polymer. Samples were prepared as described above in 125 mL Erlenmeyer flasks containing 100 mg of polyethylene (average M_W ~4000) as the sole carbon source, and were autoclaved and then heated to 135°C to cover the accessible surface of the flask. Cultures were inoculated with 50 mL of modified Bristol's medium and incubated at 30°C with agitation at 180 rpm for 6 weeks. Due to the much slower rate of growth, samples were monitored visually for separation of the polyethylene from the glass walls of the flask, and then tested at the end of the experiment by measuring optical density at 600 nm.

3. Results

3.1. Characterisation of PE‐Surrogates for Biodegradation Studies

Polyethylene (PE) is a polymer with remarkable resistance to biodegradation, which presents a significant challenge for studying its biodegradation within practical time frames (Mohanan et al. 2020). Paraffins and polywaxes serve as effective surrogates for studying polyethylene biodegradation over a reasonable period of time due to their chemical similarity to PE (Camur et al. 2024). The chemical backbones of paraffin, polywax and PE are structurally analogous, consisting of long hydrocarbon chains (Albertsson et al. 1987), and all remain solids under typical physiological conditions. However, these substrates differ from PE and one another in terms of hydrocarbon chain lengths, degree of branching and crystallinity (Jung et al. 2016; Liu et al. 1999; Ronca 2017). Low‐melting paraffin has a maximum chain length of 32 carbons and a melting point of 55°C, whereas high‐melting paraffin has a maximum chain length of 40 carbons and a melting point of 65°C. Polywax500, on the other hand, features a maximum chain length of 72 carbons and a melting point of 88°C. Each of these heterogeneous mixtures of hydrocarbons can be tracked using high‐temperature gas chromatography techniques, as illustrated in Figure 1.

FIGURE 1.

Hydrocarbon surrogates for polyethylene (PE‐surrogates) with varying chain lengths. This figure provided representative gas chromatograms of long‐chain hydrocarbon polymers used in this study, illustrating differences in percent branching (PB), melting points (MP) and carbon chain lengths (CL).

The crystallinity of paraffin, polywax and polyethylene, which plays a crucial role in determining their physical properties and biodegradation potential, is strongly influenced by the density of branching in their molecular structures (Liu et al. 1999). The branching density of the PE‐surrogates used in this study was determined using high‐magnetic‐field ¹H‐NMR spectroscopy experiments (Jung et al. 2016). The methods employed effectively differentiated short‐chain branches (1–6 carbons) from the main carbon chain of the waxes but could not distinguish long‐chain branches (> 6 carbons) from the main chain. In the homodecoupled ¹H NMR spectrum (Figure 2, left panel), the main chain methyl groups were identified by a prominent peak at approximately 0.90 ppm, while the small‐chain branch methyl groups appeared upfield between 0.83 and 0.89 ppm. To better resolve these differences, we conducted a 2D NMR experiment using proton–carbon heteronuclear multiple quantum coherence (¹H–¹³C HMQC) (Figure 2, right panel). Low‐melting paraffin exhibited a branching density of 38.7%, while high‐melting paraffin showed a higher branching density of 53.0%. Polywax500, in contrast, is a purely linear hydrocarbon with no branching. The HMQC spectra of these PE‐surrogates, shown in Figure 2, illustrate these differences and provide a simple method to compare and contrast the complexity of these polymer chains. This gradation in chain length, melting point and branching density offers a versatile range of substrates for evaluating microbial degradation capabilities under controlled conditions.

FIGURE 2.

Homodecoupled ¹H NMR (left panel) and ¹H–¹³C HMQC (right panel) spectra of PE‐surrogates. Left panel shows the 1D NMR spectra and representative chemical structures while the right panel displays the 2D NMR spectra of Polywax500, high‐melt and low‐melt paraffin (top to bottom). Branching is evident in the spectra of low‐melting (bottom) and high‐melting (middle) paraffin wax samples, while no branching was observed in the Polywax500 sample (top).

3.2. Diversity and Phylogeny of Bacterial Strains for Biodegradation Studies

Over the past 5 years, we undertook various efforts to isolate bacteria from multiple environments as potential candidates for plastic biodegradation. From our collection of isolates, those belonging to the Actinomycetes and Pseudomonads were selected for growth studies using different PE‐surrogates. Additionally, three well‐characterised oil degraders, Marinobacter aquaeolei VT8 (Huu et al. 1999), Rhodococcus jostii RHA1 (Gibu et al. 2019; Patrauchan et al. 2005; Patrauchan et al. 2008) and A. baylyi (Ratajczak et al. 1998; Wentzel et al. 2007), were included to evaluate their potential for solid‐state hydrocarbon degradation. To provide a broader comparative perspective and validate the methodology, several non‐oil‐degrading or common laboratory strains of bacteria were also incorporated into the study, including Klebsiella sp. M5al, Azotobacter vinelandii , Corynebacterium glutamicum , Gluconacetobacter diazotrophicus , Escherichia coli K‐12 and Bacillus subtilis . The phylogenetic tree in Figure 3 illustrates the evolutionary relationships among all the organisms included in this study.

FIGURE 3.

Phylogenetic tree of all bacterial strains used in this study. The tree was constructed using whole‐genome sequences with 100 iterations of the “Rapid” bootstrapping method in RAxML to ensure robust statistical support. Branch lengths represent the expected number of substitutions per site, with the scale bar corresponding to one substitution per site (branch length = 1). The tree illustrates the evolutionary relationships among the strains included in the analysis.

3.3. Growth Profile and Biodegradation Potential of Bacterial Isolates on Different PE‐Surrogates

Table 1 summarises the growth profiles of all of the bacterial strains we tested on the three PE‐surrogates used in this study. Growth was categorised based on OD₆₀₀ measurements as follows: no growth (−) [OD₆₀₀: 0.0–0.1], low growth (+) [OD₆₀₀: 0.11–0.3], moderate growth (++) [OD₆₀₀: 0.31–1.00] and high growth (+++) [OD₆₀₀: > 1.0]. Growth was monitored for 2 weeks for low‐ and high‐melt paraffin and for 5 weeks for Polywax500. The table categorises each strain based on the highest recorded OD₆₀₀ value during the observation period. In addition to growth data, the table provides the NCBI accession numbers of whole‐genome sequences for all isolates, as well as their gram‐staining phenotypes.

TABLE 1.

Growth of different strains on various polyethylene surrogates.

Acc. number	Strain ID	Species	Gram stain	Waxes			LMWPE
				Low‐melt paraffin	High‐melt paraffin	Polywax500
GCF 016839185.1	PSBB011	Rhodococcus aetherivorans	+	+++	++	++	++
GCF 017068295.1	PSBB049	Rhodococcus sp.	+	+++	++	++
GCF_017068255.1	PSBB066	Rhodococcus sp.	+	++	++	++
GCF 017068255.1		Rhodococcus jostii RHA1	+	+++	+++	++	++
GCF_048282645.1	PDNC024	Pimelobacter simplex	+	+++	++	+	++
GCF_048282625.1	PDNC017	Pimelobacter sp.	+	++	++	+
GCF 016919385.1	PDNC005	Gordonia sp.	+	+++	+++	++	+
GCF 016919465.1	PDNC001	Tsukamurella tyrosinisolvens	+	++	++	++	+
GCF_049532645.1	PCS013	Mycolicibacterium sp.	+	+++	+++	+	+++
GCF 000015005.1		Mycolicibacterium smegmatis	+	++	++	+
GCF_048282635.1	PDNC021	Mycolicibacterium sp.	+	+	+	−
GCF 016919325.1	PDNC014	Mycolicibacterium nivoides	+	+++	+	+
GCF 016919345.1	PDNC012	Mycolicibacterium septicum	+	++	−	+
GCF 000011325.1		Corynebacterium glutamicum	+	−	−	−
GCF 000009045.1		Bacillus subtilis 168	+	−	−	−
GCF 000015365.1		Marinobacter aquaeolei VT8	−	−	++	−
GCF 000046845.1		Acinetobacter baylyi ADP1	−	+++	+++	++	+
GCF_051244225.1	PSBB082	Pseudomonas alcaligenes	−	++	++	+	+
GCF_051246325.1	PSBB026	Pseudomonas putida	−	−	−	−
GCF 002090195.1		Klebsiella sp. M5al	−	−	−	−
GCF 000021045.1		Azotobacter vinelandii DJ	−	−	−	−
GCF 000021325.1		Gluconacetobacter diazotrophicus	−	−	−	−
GCF 904425475.1		Escherichia coli K‐12	−	−	−	−

Note: Growth levels for waxes were categorised based on OD₆₀₀ measurements as follows: No growth (−) [OD₆₀₀ : 0.0–0.1], low growth (+) [OD₆₀₀ : 0.11–0.3], moderate growth (++) [OD₆₀₀ : 0.31–1.00] and high growth (+++) [OD₆₀₀ : > 1.0]. Growth studies on low‐melt and high‐melt paraffin were conducted for 2 weeks, while Polywax500 studies were extended to 5 weeks. For low‐molecular‐weight polyethylene (LMWPE), growth levels were categorised based on OD₆₀₀ measurements as follows: low growth (+) [OD₆₀₀ : 0.05–0.15], moderate growth (++) [OD₆₀₀ : 0.16–0.29] and high growth (+++) [OD₆₀₀ : > 0.3]. LMWPE biodegradation was only measured for a small selection of strains (8 total) that were cultured for 6 weeks.

All Rhodococcus species exhibited strong growth on all three PE‐surrogates, with R. jostii RHA1 emerging as the most effective degrader among the Rhodococcus strains tested. The highest growth was observed on low‐melting paraffin, which has the shortest chain length but less branching compared to high‐melting paraffin. These findings suggest that chain length plays a more significant role than branching in their degradation mechanism. To confirm that the growth rate correlated with a consumption of the specific substrate utilised, we also tested specific strains with each individual PE‐surrogate to confirm biodegradation of the provided carbon source (Figure 4). These results confirmed significant decreases in the presence of remaining substrates following growth with R. jostii RHA1.

FIGURE 4.

Polyethylene surrogate decomposition by selected microbial strains. Shown above are gas chromatograms of initial (red) and final (blue) samples following growth with Acinetobacter baylyi (left) and Rhodoccous jostii (right), with low‐melt paraffin (top), high‐melt paraffin (middle) and Polywax500 (bottom). Strains were cultured under the same conditions as used for the assessment in Table 1. Values listed on each graph are the percentage (total peak area) of the specific surrogate found upon extraction of the entire sample at the end of the specified growth period (2 weeks for low‐melt and high‐melt paraffin, and 5 weeks for Polywax500).

For Pimelobacter species and Gordonia sp., a similar trend was observed, with stronger growth on substrates with shorter chain lengths. However, the overall degradation mechanism of Pimelobacter species for long‐chain hydrocarbons appears less robust when compared to Rhodococcus and Gordonia. Interestingly, Tsukamurella tyrosinisolvens displayed moderate growth across all substrates, showing no distinct preference for chain length or branching density, indicating a more generalised degradation capability.

Among the Mycolicibacterium species, PSC013 demonstrated the best overall degradation capabilities. Most Mycolicibacterium species, except for M. septicum , tended to degrade shorter chains more effectively than longer ones. However, M. septicum failed to grow on high‐melting paraffin, highlighting a drastic phenotypic difference within the genus. This observation is worthwhile for further investigation to understand the underlying biodegradation mechanisms.

Two established model oil‐degrading bacteria exhibited distinct growth behaviours on the PE‐surrogates. M. aquaeolei demonstrated moderate growth exclusively on high‐melting paraffin, characterised by a medium chain length and the highest degree of branching among the substrates. This observation suggests that its degradation mechanism is more effective on branched chains compared to non‐branched ones. In contrast, A. baylyi was capable of growing on all three PE‐surrogates, showing high growth on both low‐ and high‐melting paraffins and moderate growth on Polywax500. A. baylyi was also tested for PE‐surrogate biodegradation using GC analysis (Figure 4) with the resulting loss of each individual surrogate correlated with the OD measurements obtained in Table 1. In addition to the model strains, Pseudomonas alcaligenes was the only other gram‐negative bacterium we isolated that successfully grew on the PE‐surrogates, exhibiting moderate growth on both low and high‐melting paraffins and low growth on Polywax500. Among the Gram‐positive strains, Bacillus subtilis and Corynebacterium glutamicum showed no growth on any of the substrates, and were indistinguishable from an abiotic control. Notably, C. glutamicum is the only Actinomycete in our sample group that did not display any PE‐surrogate degradation capability.

These findings reveal species‐specific differences in solid‐state hydrocarbon biodegradation capabilities and provide potential insights into the mechanisms underlying substrate utilisation. This work underscores the importance of substrate characteristics such as chain length and branching density in influencing microbial degradation and highlights promising candidates for further studies on PE‐surrogate biodegradation.

3.4. Growth of Specific Grades of Polyethylene in Contrast to PE‐Surrogates

In order to determine how studies with PE‐surrogates might translate to long‐term studies of polyethylene biodegradation in the natural environment, we also tested a small selection of strains for the ability to grow in the presence of polyethylene alone, using a low‐molecular‐weight polyethylene (LMWPE). For these studies, we used only the 4000 average M_W LMWPE as a carbon source. Eight strains exhibiting good growth with PE‐surrogates were selected from those described in Table 1 and grown for 6 weeks on LMWPE as the sole carbon source. Clear differences were found for the eight strains, with Mycolicibacterium sp. PSC013 demonstrating the highest degree of growth, followed by two species of Rhodococcus and the Pimelobacter strain, while the other four strains showed a lower degree of biodegradation. These assessments of biodegradation were based on OD₆₀₀ measurements, which were a good indication of biodegradation with the PE surrogates and correlated well with loss of substrate (Figure 4). LMWPE is not amenable to analysis using the high‐temperature GC method we employed for PE surrogates.

3.5. Morphological Changes in PE‐Surrogate Surfaces as a Result of Culturing With Specific Strains

As a further confirmation of the ability of specific strains to grow with progressively more complex PE‐surrogates, we constructed an additional PE‐surrogate containing LMWPE (33%) and Polywax1000 (17%) along with equivalent (17%) amounts of low‐melting and high‐melting paraffin and Polywax500. Several strains with a strong potential to degrade PE‐surrogates identified from the studies in Table 1 were selected and cultured for 6 weeks; then flakes of the remaining PE‐surrogate were examined using scanning electron microscopy (Figure 5). Each strain revealed distinct morphological changes to the PE‐surrogate surfaces that had detached from the glassware during the course of the growth, indicating the potential for unique mechanisms in each of these strains for accessing and degrading the PE‐surrogates.

FIGURE 5.

Scanning electron micrographs of PE‐surrogates treated with selected microbial strains. Shown are images of the surface of the polyethylene surrogates cultured with Acinetobacter baylyi (A), Mycolicibacterium sp. PCS013 (B), Pseudomonas alcaligenes PSBB082 (C) and Rhodoccous jostii (D), indicating various degrees of pitting, decomposition and deposition.

4. Discussion

Plastics like polyethylene share certain properties and chemical characteristics with many petroleum hydrocarbons. While generally viewed as recalcitrant molecules due to the time that is required for significant levels of biodegradation to occur in the natural environment, polyethylene does slowly degrade due to both abiotic and biotic processes (Albertsson and Karlsson 1990). A primary obstacle to characterising the process of biological degradation of polyethylene is the duration of time required to run experiments to quantify rates of decay, which can take years, decades or centuries (Ronca 2017). Bridging the interface between polyethylene and simple hydrocarbons such as motor oil are the classes of compounds known as paraffins and polywaxes. These compounds share many of the same characteristics as polyethylene, including higher melting points and longer polymer chain lengths, while the carbon chain lengths are not so long that they take years to fully degrade through biological processes. We previously applied a methodology that utilises combinations of paraffins and polywaxes to enrich communities of microbes from various environments in Minnesota for the ability to fully degrade these PE‐surrogates in a matter of weeks (Camur et al. 2024). The selection of these materials is based on the rationale that any microbe that is capable of degrading the longer polymers found in commercial grades of polyethylene should also degrade these shorter polywaxes and paraffins (Albertsson and Karlsson 1990; Ammala et al. 2011). Other aspects of this method include heating the waxes and then coating the culture vessel surface with the PE‐surrogate materials to maximise surface area available to the microbes while also assuring the substrates and media have been sterilised prior to introduction of single microbial strains.

Like most polymers, the designation of polyethylene does not represent a single homogeneous chemical compound. There are many different classes and grades of polyethylene that include polymers of different chain lengths and various degrees of branching, where a section of polyethylene branches off from the main backbone of the molecule (Ronca 2017). This degree of branching can dramatically alter the ability of these polymers to pack together, affecting the density of the material, and is one of the primary differences between high‐density and low‐density polyethylenes (HDPE and LDPE). The addition of polymer branching means that the chemistry and the biological mechanisms that would be required to fragment and catabolise these polymers into common metabolites that can feed into general metabolism would need to adapt to accommodate the different carbon backbones. Investigating whether certain microbes prefer branched versus linear polymers was a specific aspect of this report that we wanted to explore. To characterise these features, we first applied advanced NMR techniques to characterise the various PE‐surrogates that were selected for these studies (Jung et al. 2016; Liu et al. 1999). The manufacturer of the Polywax500 (NuCera Solutions) claims that their polywaxes are straight‐chained polymers without any branching. Our results confirmed this to be the case and served as a standard for comparison with the low‐melt and high‐melt paraffins that we obtained from Sigma‐Aldrich. Based on this analysis, it was possible to characterise individual strains for their preferential degradation of branched versus linear polymers.

Our prior studies identified one specific class of microbes that seemed to be particularly good at degrading a mixture of PE‐surrogates. Those microbes were all members of the Mycolicibacterium genus, and were found in multiple communities that were isolated from geographically distinct locations and during different time frames (Camur et al. 2024). Mycolicibacterium species are faster growing than many of their pathogenic relatives, and are often characterised as saprophytes that decay organic matter and potentially hazardous chemicals in the environment (Golubev et al. 2021; Ji et al. 2020; Ren et al. 2021; Sun et al. 2022; Zeng et al. 2010; Zhang et al. 2013), and have also been reported to biodegrade polyethylene (Sun et al. 2022). In addition to these individual isolates, other microbes that are also Actinomycetes were common in our communities, including several members of the Rhodococcus genus, which is another class of microbes often involved in the biodegradation of various complex organic compounds (Patrauchan et al. 2005; Patrauchan et al. 2008; Gilan et al. 2004). Various Rhodococcus strains have been identified by other laboratories to play an important role in the biodegradation of plastics (Gilan et al. 2004; Putcha and Kitagawa 2024; Rong et al. 2024; Tao et al. 2023).

In an effort to expand on our prior methods and isolation of various microbes following several years of work in this area of study, we screened a larger strain library of more than 50 isolates for the ability to degrade high‐melt paraffin in a suitable segment of time (2 weeks) to accommodate further scientific investigations. Strains that showed evidence of biodegradation of high‐melt paraffin were further screened for growth on low‐melt paraffin and Polywax500 to determine if these strains shared the same general pathways to degrade these various substrates, or if individual strains may have a selective preference for specific substrates with varying lengths and degrees of chain branching. In addition to screening strains we had isolated in the past, we also selected several model microbes recognised for their ability to degrade petroleum hydrocarbons in the environment (Huu et al. 1999; Gibu et al. 2019; Ratajczak et al. 1998).

During the course of this study, we isolated several members of the Rhodococcus genus that were found to consume the PE‐surrogates in a reasonable timeframe (Figure 4). Our laboratory has studied the strain R. jostii RHA1 in the past in relation to the production of wax esters (Barney et al. 2012). To compare our isolates to R. jostii RHA1, we included this model strain in our analysis and confirmed that it was able to achieve a similar degree of decomposition and growth using the PE‐surrogates, supporting the notion that this may be a general feature of the Rhodococcus genus (Putcha and Kitagawa 2024).

R. jostii RHA1 is among a small selection of bacteria that also accumulate either triacylglycerides or wax esters (Barney et al. 2012). Many of these microbes are recognised as hydrocarbon‐degrading microbes (Huu et al. 1999; Gibu et al. 2019; Ratajczak et al. 1998), and they span a broader diversity of microbes that extends outside of the Actinomycetes. Our laboratory has also studied Marinobacter aquaeolei VT8 (recently renamed M. nauticus (Tindall 2020)) and A. baylyi ADP1 (Barney et al. 2012; Knutson et al. 2017; Singer et al. 2011; Wahlen et al. 2009; Willis et al. 2011). Both of these microbes accumulate wax esters and/or triacylglycerides, and both of these microbes are associated with biodegradation of petroleum‐contaminated environments (Huu et al. 1999; Fox et al. 1992; Gauthier et al. 1992; Ishige et al. 2000). Our experiments to test the abilities of these two microbes to utilise the PE‐surrogates resulted in vastly different results. While M. aquaeolei was only able to grow moderately well using high‐melting paraffin as a substrate, A. baylyi did remarkably well on all three classes of polyethylene PE‐surrogates. Similar to other strains identified in this study, other laboratories have also identified specific Acinetobacter strains as polyethylene degraders (Kim et al. 2024; Montazer et al. 2019; Sandholm et al. 2025; Zhang et al. 2023). The identification of R. jostii RHA1 and A. baylyi ADP1, two microbes that have been broadly studied in laboratories across the world (Barbe et al. 2004; McLeod et al. 2006), as potential model strains for further biochemical study, should accelerate the potential for further biochemical studies with microbes that have highly‐developed genetic systems. Our prior study identified Mycolicibacterium smegmatis, another highly studied model microbe, as a potential PE‐surrogate degrading strain. M. smegmatis was inferior at degrading the PE‐surrogates in comparison to our natural isolates (Camur et al. 2024), an aspect we further confirmed here (Table 1).

The intention of this study was to investigate the ability of a broad selection of isolates to degrade compounds that share characteristics with different classes of polyethylenes. Indeed, certain strains seemed to degrade branched polymers much better than straight‐chain polymers. Additionally, our results should inform future efforts to expand this list of microbes to enable both biochemical and bioinformatics approaches to better characterise the repertoire of enzymes and mechanisms that could improve or increase the biodegradation of polyethylene‐related compounds. By sequencing our natural isolates and comparing these with strains that are commonly studied across the globe, we provide a resource that is accessible to the broader research community.

As a further negative control, we selected a number of common laboratory strains that were not anticipated to degrade complex hydrocarbon substrates in a significant manner. As expected, strains such as Escherichia coli K‐12, Bacillus subtilis 168, Klebsiella sp. M5al, Azotobacter vinelandii and Corynebacterium glutamicum showed no visible signs of growth on these substrates and could potentially serve as negative controls in bioinformatics studies. Since many of the microbes isolated in this study are members of the Actinomycetes, we were particularly interested in whether PE‐surrogate degradation was a general feature of this class of microbes. The poor growth of C. glutamicum seems to indicate that this is not just a general feature of the Actinomycetes, but it would be of interest to test additional Actinomycetes in the future to get a more general picture of the prevalence of this phenotype.

Members of the genera Corynebacterium, Rhodococcus, Gordonia, Tsukamurella and Mycolicibacterium are all known to produce mycolic acids of varying chain lengths, from 22 to 90 total carbons, though this chain length varies among different species (Marrakchi et al. 2014). It was postulated that the ability to synthesise these longer hydrocarbon chains might be linked to the ability to degrade longer hydrocarbons such as those employed in this study, and that strains with longer mycolic acids might be better at degrading the longer polymers used here than those with shorter chain lengths. Tsukamurella and Mycolicibacterium generally have longer mycolic acids than Rhodococcus, which was among the best PE‐surrogate degraders identified in our study, so the ability to degrade these PE‐surrogates does not seem to correlate with the size of the mycolic acids that are synthesised internally by the microbe. Additionally, the inability of C. glutamicum to degrade any of the substrates at appreciable rates indicates that this is not just a general feature of strains that produce mycolic acids. The identity of strains such as A. baylyi and P. alcaligenes as strong degraders of the PE‐surrogates indicates that this is not the only factor involved in determining which strains are the best degraders of these materials.

There have been many reports of bacteria with the potential to degrade polyethylene or similar polymers. Representative strains from the genera Rhodococcus, Acinetobacter, Pseudomonas, Gordonia and Mycolicibacterium are commonly recognised in the literature in relation to reported biodegradation of polyethylene or are found associated with these plastics in the environment (Mohanan et al. 2020; Sun et al. 2022; Tao et al. 2023; Montazer et al. 2019; Ghatge et al. 2020; Jachimowicz et al. 2022; Laha et al. 2018). While genomic data are available for some of these reported strains, this is not always the case, and a direct comparison of strains with one another is often difficult as the methods used to characterise the degree and nature of the biodegradation vary from one report to the next. While our report is not meant to supersede the contributions of these other studies, we do believe that the characterisation of a large number of strains in the same manner and with the provision of the sequenced genomes for these strains represents a valuable resource for further studies and in efforts to establish a benchmark for biodegradation potential.

5. Conclusions

The primary goal of this study was to create a resource of sequenced microbial strains with the potential to degrade PE‐surrogates, and also to characterise these strains in terms of the classes of polymers (branched versus straight‐chain) that they are able to degrade. This broad list of strains, and comparisons to other strains commonly associated with general hydrocarbon degradation, should be a valuable resource and tool in efforts to identify the enzymes and general biological mechanisms that enable certain microbes to degrade hydrocarbons from the environment that are solids at physiological temperatures. Future studies could apply RNA‐Seq, Tn‐Seq and other omics approaches coupled to biochemical studies to identify the pathways and supporting metabolites or surfactants that might be essential to these processes.

Author Contributions

Bilge Bahar Camur: investigation; methodology; formal analysis; writing – original draft; writing – review and editing. Lucas Stolp: investigation; methodology; writing – review and editing. Lydia Mindermann: investigation; writing – review and editing. Natalia Calixto Mancipe: investigation; writing – review and editing. Joseph B. Moidl: investigation; writing – review and editing. Brett M. Barney: conceptualization; funding acquisition; investigation; methodology; supervision; writing – original draft (supporting); writing – review and editing (lead).

Funding

This work was supported by the National Institute of Food and Agriculture, MIN‐12‐070, MIN‐12‐081; Legislative‐Citizen Commission on Minnesota Resources, M.L. 2024, Chp. 83, Sec. 2, Subd. 08g; and Fulbright Association.

Conflicts of Interest

The authors declare no conflicts of interest.

Data Availability Statement

Genome sequences assembled as part of this work are available through the provided accession numbers in Table 1 and are available through the National Center for Biotechnology Information (NCBI).