Metatranscriptomics of microbial biofilm succession on HDPE foil: uncovering plastic-degrading potential in soil communities

doi:10.21203/rs.3.rs-3014809/v1

Download PDF

Research Article

Metatranscriptomics of microbial biofilm succession on HDPE foil: uncovering plastic-degrading potential in soil communities

https://doi.org/10.21203/rs.3.rs-3014809/v1

This work is licensed under a CC BY 4.0 License

Version 1

posted

You are reading this latest preprint version

Background

Microbial communities in soil are a complex and sensitive system which secures soil health, nutrient cycling and the degradation of natural and xenobiotic substances. Even though plastic pollution is increasing worldwide, very little is known about microbial processes that take place once plastic debris gets incorporated into the soil matrix. In this study, we conducted the first metatranscriptome analysis of polyethylene (PE)-associated biofilm communities in a highly polluted landfill soil and compared their gene expressions to those of a forest soil community within a 53-day period.

Results

Our findings indicate that the microbial population present in soil contaminated with plastic debris carries a predisposition to both inhabit and degrade plastic surfaces. Surprisingly, the microbial community from an undisturbed forest soil contained a diverse array of plastic-associated genes (PETase, alkB etc.), indicating the presence of an enzymatic machinery capable of plastic degradation. Plastic-degrading taxa were upregulated in the early stages of biofilm and the PE-degrading enzymes alkB1/alkM and transporters such as FadL, livG, livF, livH and livM and fatty acid β-oxidation pathway were active during the maturation of the biofilm. We also found an increase in nitrogen fixation genes in the plastic soil community (but not in forest soil), indicating an essential metabolic adaptation of biofilm communities in the plastisphere.

Conclusion

With this study, we address the underlying patterns of gene expression during biofilm development of a PE-associated plastisphere in soil and address the pressing question whether or not natural microbial communities carry the potential to biodegrade petrochemical-based plastic in the (soil) environment.

plastisphere

soil

metatranscriptomic

biofilm

plasticdegradation

microorganisms

Plastic has become a ubiquitous human-made material found in nature. It is known to accumulate as pollutant in terrestrial ecosystems through a variety of sources, such as improper disposal, blow-offs from landfills and large remains of mulching foils and sewage sludge used in agricultural applications [1, 2]. Its presence in soil is currently estimated to be 4–23 times higher than in the ocean. Since microorganisms are generally the first responders to anthropogenic perturbations, their interactions with plastic debris are crucial for the resilience of soil ecosystems [3]. Recent literature has shown that plastic debris can shape the associated soil microbial community similar to microbial communities found on plastic debris in aquatic ecosystems, which characterises the terrestrial plastisphere as a distinct microbial habitat [4–10].

Once littered to the ground and incorporated into the soil matrix, plastic debris can compromise a number of important healthy soil properties such as soil aggregation [11], hydraulic conductivity and water holding capacity [12]. It furthermore shows negative dose-dependent effects on agricultural crop growth [13, 14]. Most plastic waste is generated in the packaging sector in which polyethylene forms the biggest fraction of plastic primary production [15]. In the year 2015, 97 million tons of polyethylene (PE) were disposed either into landfills, incinerators or have eventually entered natural environments [16]. PE is also commonly used as mulching film or foil in agricultural fields, where fragmented PE residues accumulate in soil over the years forming a tremendous threat to global soil health [17, 18].

PE is a petrochemical-based polymer consisting of long-chains (~ 4000–40 000 carbon atoms) of -CH₂-CH₂- functionalities derived from ethylene monomers whose carbon backbone is either branched, as in low density polyethylene (LDPE), or linear as in high-density polyethylene (HDPE). Due to its high molecular weight and hydrophobicity, a successful microbial degradation requires the organisms to possess cellular properties which allow an interaction with the recalcitrant PE surface [19]. Even though a surprisingly large number of bacterial and fungal genera were found associated with PE degradation (Acinetobacter, Arthrobacter, Aspergillus, Bacillus, Brevibacillus, Klebsiella, Nocardia, Pseudomonas, Rhodococcus, Staphylococcus, Streptomyces) [19–25], so far only a few studies proposed biochemical mechanisms or enzymes expressed within PE-associated microorganisms [26–28]. Studies that screened microbial metagenomes for potential plastic-degrading enzymes from uncultivated microorganisms, have demonstrated the great potential of such Meta’omics techniques to unravel underlying mechanisms of PE degradation [29, 30].

Crucial for microbial colonization of plastics and potential degradation, is an initial abiotic deterioration driven by UV-radiation, heat and mechanical forces which lead to macromolecular changes in the polymer structure and an increase of hydrophilic groups on the surface [31]. This creates favourable conditions for plastic-associated biofilm formation. Due to the complexity of the enzymatic processes involved in a complete bio-degradation of long-chained polymers like PE, it must be assumed that different microorganisms are responsible for the various steps of fragmentation and deterioration of the plastic material up to the assimilation and mineralization of plastic-derived carbon [32–34]. The formation of mixed microbial biofilms was noted to play a major role in the decomposition of plastics in soil environments, promoting direct contact and possible degradation processes to occur at the interface of biofilm and the plastic surface [35].

Like formation of any other biofilms, the colonization of plastic occurs through different successional stages: (i) During an initial attachment phase, pioneering microorganisms that possess physiological features to adhere to the plastic surface (e.g. high cell hydrophobicity) will colonize the plastic surface rapidly [36]. At that stage, physico-chemical surface properties of the polymer, such as polymer type, hydrophobicity, roughness and degree of surface weathering, play a major role [37]. Studies suggest that after this initial attachment stage, a selection phase follows in which plastic-degrading taxa (e.g. Rhodococcus, Micrococcus, Pseudomonas and Brevibacillus spec.) will be enriched and compete for the released PE oligomers and/or leaching additives as carbon source from the plastic itself [38–40]. Extracellular enzymes such as hydroxylases, oxygenases, laccases and other candidate enzymes cleave the polymeric backbone into smaller oligomers, like aliphatic carboxylic acids, aldehydes and alkanes which can later be assimilated and metabolized by bacteria or fungi [41, 42]. Once the biofilm matures further, more cells grow as a secondary biofilm layer, forming irreversibly attached cell aggregates with the excretion of extracellular polymeric substances (EPS) which stabilizes and protects the multicellular communities. During this mature succession phase (iii), the plastic-derived carbon sources will be exhausted, and the polymer properties no longer play a major role in shaping the microbial community. Phototrophy, grazing and cross-feeding become the major fuels for microbial growth which out-compete the pioneering specialized plastic-associated taxa [43–46].

Here, we aimed to track microbiome gene expression patterns during biofilm formation on HDPE foil (hereafter PE foil) by conducting the first metatranscriptomic study of the terrestrial plastisphere communities under laboratory conditions. To do so, we performed a controlled soil incubation experiment in which the biofilm formation on PE foil was compared to that on glass slides over a course of 53 days. Total RNA was extracted directly from the colonised surfaces at five time points (T₁: 2 days, T₂: 7 days, T₃: 14 days, T₄: 21 days and T₅: 53 days). Furthermore, by imaging the biofilm succession with scanning electron microscope (SEM), we compared the surface interaction and the biofilm coverage on both glass and PE during the course of time. As early biofilm succession selects microorganisms capable to utilize the polymer as substrate of growth, we expected an enrichment of microbial taxa and genes associated with PE degradation to occur within the early sampling points.

Previous studies showed that once in contact with the soil microbiome, plastic debris enriches a specialized microbial community, distinct in their taxonomic composition and adaptation to plastic as substrate[7, 9, 10, 47]. We therefore hypothesized, that microbial communities with a higher exposure to plastic debris in their soil matrix, would likewise colonize and possibly utilize a newly introduced PE foil more efficiently. In order to test this hypothesis, we compared the potential of two very different soil types and their native microbial communities. Namely a natural forest soil community and a community derived from a highly plastic polluted landfill. We predicted the landfill microbial community to be more effective than the forest soil community in colonizing the PE surface and to express a larger variety of genes and taxa related to PE degradation. With this study, we want to elucidate the underlying patterns of gene expression that take place during biofilm development of a PE-associated plastisphere in soil. Furthermore, in the light of an ever-increasing plastic production worldwide, our research tackles the pressing question whether or not natural microbial communities have the potential to biodegrade petrochemical-based plastics in the (soil) environment.

Plastic Material and UV-Weathering

The HDPE polyethylene foil (hereafter PE) used for the incubation experiments was kindly provided by ‘Bundesanstalt für Materialforschung und -prüfung’ (BAM), Berlin. In order to mimic outdoor weathering conditions and to increase the foil’s susceptibility to microbial attachment, the PE foil was UV-weathered for 100 h at 75° C in a UV weathering chamber (Global-UV Test 200, Weiss Umwelttechnik, Vienna, Austria) with UV-fluorescence lamps according to DIN EN ISO 4892-3, type 1A (UVA-340). HDPE samples received a total surface energy of 14 MJ m^-2 (40W m^-2 x 100 h).

Study design and incubation

In order to compare the native inhabiting microbial community from natural forest soil with that of a highly plastic polluted soil (hereafter plastic soil), both soil types were sampled in the summer months (August 2018, September 2021) and stored in sterile Schott bottles and frozen in -20°C until further handling. Soils were thawed over a week at an increasing temperature from 8°C to 16°C in the dark to acclimate and re-establish microbial growth. From each of the two soil types, 15 g of material was placed in a sterile Schott bottle (100 ml) and wetted with demineralized water at approximately 60% of water holding capacity (WHC).

Bottles were handled horizontally to increase the soil surface area to its maximum. We sterilized both the PE foil and the glass-slides by submerging them into 70% ethanol for 15 min and letting them dry under a Lamina flow chamber. The PE foil was cut into pieces of roughly 1.0 x 0.5 cm. In each bottle, three pieces of PE foil and three glass-slides were carefully placed and buried with soil material. Bottles were incubated at 28°C with an LED light (SanLight model S2W) which mimicked a 6 hours day-night cycle. Bottles were incubated over a period of T₁: 2 days, T₂: 7 days, T₃: 14 days, T₄: 21 days and T₅: 53 days.

SEM sampling and fixation

For SEM analysis of biofilms, both glass and PE foils were gently rinsed with demineralized water before being fixated in Methacarn (60% Methanol, 30% Chloroform and 10% Acetic acid) solution for at least 24 hours. After fixation, samples were dehydrated in a graded series of ethanol 30% (5 min), 50% (10 min), 70% (15 min), and 99% (20 min) according to an adjusted protocol by Dassanayake et al, 2020 [48]. Samples were coated with 12 nm of carbon using LEICA EM ACE600 coater. SEM imaging was performed on a FEI Quanta 3D SEM at accelerating voltage of 10 kV, beam current 23–93 pA, working distance 6–10 mm using secondary electron ET detector. To compare surface coverage 4 images per sample per time point and type of soil, biofilm growth of both PE and glass were obtained and analysed. Surface coverage, cell-morphology and general organic features were documented and compared for the different soils and time points.

RNA and DNA extraction

At each of the five time points, bottles from both soil types were sampled in duplicates and handled under sterile and nuclease-free conditions. Out of each bottle, PE foils (n = 3) and glass slides (n = 3) were carefully recovered with sterilized metal tweezers and rinsed with nuclease-free water (sterile DEPC treated water, Carl Roth GmbH) to remove external soil particles. PE foil and glass slides were weighed and placed directly into ZYMO Research Bead Lysis tubes for immediate RNA extraction using the ZYMO Research Quick-RNA ^TM Soil Microprep Kit. Extraction samples were lysed and homogenized with a FastPrep-24^™ beat beating grinder at two cycles of 25 sec (6 m s^-2) and placed on ice immediately after. Extracted RNA was eluted in 15 µL nuclease-free water and stored at − 80°C prior to final quantification of total RNA yield.

For a metagenome sequencing of the bulk microbial communities present in the two different soils, we took DNA samples from each bulk material (plastic soil and forest soil) before the incubation at timepoint T₀ and after 53 days of incubation at the final timepoint T₅. The DNA extraction was performed using Roboklon EurX Soil DNA Purification Kit with 250–300 mg of bulk soil material per sample and further handled according to the protocol below.

Quality control and metatranscriptome/metagenomic sequencing

To remove any residual DNA from the RNA extracts, eluted RNA samples were treated with DNAse using the TURBO DNA-free kit (Thermo Fisher Scientific, Germany) before further quantification and library preparation. All RNA samples were then analyzed using a Bioanalyzer RNA 6000 Nano Assay to determine yield and integrity. RNA extracts of each triplicate from a single bottle were pooled and concentrated to a final concentration of 150 ng RNA in 15 µL using vacuum centrifugation (Concentrator plus, Eppendorf Germany) for metatranscriptome sequencing. This way, we obtained a biological duplicate per time-point and soil type for further sequencing. Library preparation and RNA/DNA sequencing were performed by Eurofins Genomic Germany GmbH (Anzinger Str. 7a, 85560 Ebersberg, Germany) on an Illumina NovaSeq6000 platform aiming for 30 million paired-end reads of length 150nt. The final sequencing depth differed between samples since the obtained RNA concentrations were variable and generally low due to the limitation of attached biomass especially during early biofilm succession. To compensate for the overall low biomass of the attached biofilms, we used triplicates of samples in each experimental unit and pooled their RNA yield as described above. In the following all raw reads were processed using the ATLAS metagenome pipeline (QC module) to obtain dereplicated, quality controlled, and trimmed reads [49].

De Novo assembly of metagenomics reads and functional annotation

The ATLAS metagenome pipeline (module assembly) was used to generate contigs from the DNA reads. From these contigs, genes were predicted using prodigal v2.6.3 [50] and annotated using the eggNog emapper v2.0.1 [51] with a database from October 2020 [52]This resulted in contigs and annotated genes for each of the 4 DNA samples (forest/soil for 2d and 53d).

KEGG annotation of RNA reads

RNA reads were mapped to the DNA contigs (53d of the corresponding soil) and the reads were counted per gene. The obtained counts were normalized as follows: First, gene counts were corrected for sequencing depth with bowtie2 v2.4.2 [53]by normalizing to the total mapped sequencing reads of each sample (per million mapped reads). Second, to correct for different gene length, counts were normalized by kilobases of gene length (e.g. for a gene with 2000nt length, the count is divided by 2). These normalized gene counts are comparable between different genes within and between samples. The functional annotation with EGGNog (see above) also annotates KEGG IDs. To obtain summarized KEGG counts, the normalized counts were simply summed-up by KEGG ID.

Taxonomic and functional assignment of active plastisphere community

To obtain a detailed taxonomic assignment of the active microbial species, quality-controlled RNA/DNA reads were mapped to the SILVA SSU database v138 [54] to calculate the abundance of microbial ASV’s in the plastisphere community. Mapping was performed using bowtie2 v2.4.2 [53]. Taxon abundance was finally obtained by summing gene counts by taxonomic level of interest. For the functional profiling of active transcripts within the plastisphere community, we used our KEGG annotated metatranscriptomic data and compared it to a set of 35 known enzymes involved in PE- and/or general polymer colonization, five genes related to nitrogen fixation (nif genes) as well as 133 biofilm related enzymes involved in lipopolysaccharide biosynthesis and extracellular matrix synthesis (Supplementary table 2) [20, 27, 55].

Data analysis

Data analysis and plotting was performed using RStudio version 2022.07.0 + 548 "Spotted Wakerobin" Release for Ubuntu Bionic. R packages used for sequence data analysis included vegan, phyloseq and genefilter. We performed SEM image analysis and the calculation of the surface coverage with Fiji ImageJ version 1.53q using a Kuwabara filter (radius 3) and the Triangle algorithm for threshold analysis of the biofilm coverage. Graphics were adjusted using Inkscape version 0.92.5.

RNA sequencing and taxonomic annotation

A total of 263 059 818 raw reads were obtained from 24 RNA samples which eventually passed quality control, with an average of 10 960 826 reads per sample. After merging and the removal of chloroplasts and mitochondria reads, we obtained a total of 15 029 taxonomic units from our RNA-Seq library from which 1 059 of the taxonomic units were filtered out due to low relative read numbers abundance (read number mean abundance < 10^⁻5). The microbial diversity and community structure were thus analyzed based on a total of 9 594 prokaryotic and 4 376 taxonomic units were assigned to the Eukaryota (Supplementary table 1).

With our original experimental design, we aimed for a total of 38 pooled RNA samples obtained as duplicates from five time points in time (T₁ = 2 days, T₂ = 7 days, T₃ = 14 days, T₄ = 21 days and T₅ = 53 days). However, due to low RNA yield from the biofilm specimen, only 24 samples (none of T₄ = 21 days) passed the RNA sequencing quality control with an average amount of 28.32 ng RNA per sample from four sampling points in time.

Soil and time dependent biofilm succession on PE and glass

We tested whether we could observe different stages of biofilm succession on soil-buried PE foil and performed a PERMANOVA analysis (Table 1) in order to identify the different conditions shaping this biofilm community. To do so, we used the taxonomic composition assigned from our RNA transcripts and calculated their relative abundance within the active portion of the sequenced community to calculate a PCoA plot based on Bray-Curtis measures of dissimilarity (Fig. 1).

Table 1

PERMANOVA testing of factors explaining differences in biofilm community
	df	Sums of square	Pseudo F	R²	P value	Significance
PE vs. glass	1	0.4566	1.2581	0.05	0.266	-
Forest soil vs. Plastic soil	1	1.8062	5.9899	0.21	0.001	***
Days	3	2.0328	2.1151	0.24	0.002	**
Early biofilm (Day 2 vs. rest)	1	0.5118	1.4202	0.06	0.152	-
Early mid biofilm (Day 7 vs. rest)	1	0.7724	2.2162	0.09	0.015	*
Late mid biofilm (Day 14 vs. rest)	1	0.3082	0.8338	0.04	0.584	-
Late biofilm (Day 53 vs. rest)	1	1.1902	3.6116	0.14	0.001	***

According to PERMANOVA analysis, the type of soil explained 21% of the observed differences between the biofilm communities (Table 1), whereas the type of substrate (PE vs. glass) did not contribute to a significant shift of the biofilm community structure (R² = 0.05, p = 0.266).

We observed a clear separation of the sequenced biofilm communities incubated in plastic soil versus forest soil, which deviated along the primary principal axis (Fig. 1). According to PERMANOVA analysis, the type of soil explained 21% of the observed differences between the biofilm communities. Within the two soil types, the most apparent separation of the microbial communities was explained by the different successional stages of the biofilm according to days of incubation (Fig. 1). In both plastic soil and forest soil, the microbial communities changed along the progression of the biofilm from an early biofilm (Day 2) to a mid- (Day 7) and late biofilm succession (Days 14–53). The most advanced point of our biofilm incubation experiment (Day 53) showed also the most distinct microbial beta diversity, which differed significantly from the rest of the sequenced time points in both forest- and plastic soil (PERMANOVA: R² = 0.146, p = 0.0012).

Most striking was the community succession of the incubated biofilm in plastic soil: All four time points formed individual clusters along the two axes of the PCoA (Fig. 1) and a development of the biofilm communities was observed based on their taxonomic shifts from early- to mature colonization. In contrast to that, the community structure of the biofilms in forest soil clustered closely together during the early- and mid-biofilm successional stages and showed a significant shift in community structure only on day 53 (Fig. 1).

To test the activity and function of the biofilm microbial taxa, we compared their expression levels of 16S/18S rRNA gene transcripts for taxonomic assignment (Fig. 1–3) at the given points in time as well as the expression levels of specific genes relevant f or the PE-plastisphere (Supplementary table 2) for functional profiling (Fig. 4–5). By using this metatranscriptomic approach, we were able to follow only the active portion of the PE-associated plastisphere community during their biofilm succession over 53 days in the two different soils.

Plastic soil

The PE-associated biofilm communities in plastic soil developed over the course of time into a less diverse community compared to their initial successional stage (Fig. 2). After an initial increase in diversity at day 7 (Median_Chao1 = 4732.7, Median_H’ = 5.42, Median_D’ = 0.99), the microbial diversity decreased significantly within the biofilm of the late successional stage at day 53 (Median_Chao1 = 3899.8, Median_H’ = 2.85, Median_D’ = 0.78 ). We found no significant differences in alpha diversity between the biofilm community on PE vs. glass at any successional stage. Concerning the taxonomic makeup of the attached microbial communities, however, we did observe slight differences between the microbial community attached to PE when compared to the biofilm community on glass (Fig. 2). A preference of known plastic-associated taxa (e.g Rhodococcus, Nocardioides and Streptomyces) to PE surfaces was observed in the mid-biofilm stages as compared to glass, however, none of them were exclusively active on PE (Fig. 2).

In the early- to mid-biofilm stages (between 2 and 7 days), transcripts of the Actinobacteriota and Firmicutes were the most abundant prokaryotic phyla of the biofilm communities in plastic soil (Fig. 2). Transcripts of the genera Lamia (4.9% of mean relative abundance), Mycobacterium (3.3%), Nocardioides (2.2%), Rhodococcus (2.4%) and Streptomyces (2.0%), were dominant in the early stages of biofilm succession, followed by the members of the Firmicutes phylum Sporosarcina (1.4%) and Bacillus (1.1%). All of them decreased in their SSU rRNA abundance during biofilm succession (Fig. 2) and disappeared in the late biofilm stage (e.g Proteobacteria) or were otherwise present in abundances below 0.1% (Firmicutes). Other present SSU rRNA transcripts in early biofilm succession were Chloroflexia JG30-KF-CM45 (2.0%) of the Chloroflexi and Pedosphaeraceae of the Verrucomicrobiota phylum (1.2%). Proteobacteria were represented by transcripts of the genera Amaricoccus (0.7%), Mesorhizobium (0.8%) and Rhodobacteraceae (0.6%).

The active eukaryotic fraction of the biofilm community consisted of the Ascomycota Aspergillus (0.9%), single-celled protists like Cercozoa (1.0%) and Cercomonas (0.3%) (Supplementary Figure S1 A). Other abundant Eukaryotic members of the community were the saprotrophic fungi Mortierella (1.8%), as well as the protozoa (Ciliophora) and the order of Agaricales (gilled mushroom). Trancriptomic reads originating from Embryophyta (plants) and Acanthamoeba (amoeba) were likewise present across all communities (Supplementary Figure S1 A).

Moving further in time, the most apparent shift in activity of the microbial community was detected between Day 14 and 53 of biofilm succession. The transition from a mid-succession stage to a late biofilm community was accompanied by a strong increase of Cyanobacterial transcripts (up to 49% of all sequenced reads per sample) and a decrease of other early-to mid-biofilm members (Fig. 2). Highly abundant were autotrophic genera of the Nostocaceae, e.g Nostoc_PCC and Tolypthrix. Other eukaryotic late colonizers were the green algae Chlorophycae (7%) and Trebouxiophycae (2.5%) of the Chlorophyta phylum (Supplementary Figure S1). Firmicutes members of the late biofilm succession were Bacillus (1.0%) and Sporosarcina (0.9%), whereas the Chloroflexi phylum was represented by the genus A4b of the Anaerolineae (1.3%), JG30-KF-CM45 (0.6%) and Caldilineaceae (0.5%). Also, phylum members of the Planctomycetota were a dominant part of the late biofilm community with Gemmataceae (1.24%), SM1A02 (1.2%) and Pirellulaceae (0.67%). From the Actinobacteriota only Microbacterium (1%) and Microtrichaceae (1.5%) were present at that late succession stage. Higher abundant eukaryotic transcripts were again derived from Embryophyta (17%) and reads of the grazing protists Acanthamoeba (1%) and Vermamoeba (0.6%).

Forest soil

Similar to the biofilm communities in plastic soil, we saw a first slight increase in diversity peaking at day 7 of incubation (Median_Chao1 = 4279 Median_H’ = 4.14, Median_D’ = 0.94) and a gradual drop in diversity towards the late successional stage after 53 days (Fig. 3). The differences in forest soil where, however, smaller than in plastic soil and none of them statistically significant (Fig. 3). The sequenced biofilm community in forest soil stayed overall more stable during biofilm succession, the taxonomic differences were also less distinct between the different biofilm stages (Fig. 3).

The early biofilm community was again dominated by Actinobacteriota such as Acidothermus (32.9%), Mycobacterium (6.9%), Conexibacter (3.5%) and Acidimicrobiia (3.6%), which where the top initial colonizers in the biofilm community on both substrates (Fig. 3). However, the early biofilms derived from forest soil contained more members of the Proteobacteria, such as Elsterales (5.9%), Acetobacteraceae (1.9%) and Roseiarcus (1.6%), as well as members of the phylum Planctomycetota, e.g the cellulose degrader Gemmataceae (2.6%).

Compared to plastic soil, there were less taxonomic changes during biofilm succession until day 53 of incubation in forest soil. During the late succession stage of the biofilm, there was a tremendous decrease in taxonomic richness on glass compared to that of PE (Fig. 3 and Figure Supplementary Figure S1 B). Both the Bacterial- and Eukaryotic fraction of the biofilm on glass consisted of a few highly abundant taxa: The Actinobacteriota Acidothermus was highly abundant with up to 43.4% of all reads in a given sample. Equally abundant were the genera Gemmataceae and Pirellulaceae of the Planctomycetota phylum. A reduced taxonomic diversity on glass-associated biofilm was mostly visible in the top 50 prokaryotic taxa (Fig. 3), as only seven of the top genera were present in the mature biofilm (day 53) on glass compared to the 31 unique genera of higher abundance present within the PE-associated biofilm of day 53.

Concerning the eukaryotic fraction of the biofilm community, fungi were more abundant in forest soil than in plastic soil. The genera Aspergillus (3.4%) and Penicillium (5.3%) of the phylum Ascomycota were abundant in the early- to mid-successional stages of the biofilm community, yet decreased in their abundance within the mature biofilm community (Supplementary Figure S1 B). Contrasting to the eukaryotic community in plastic soil, we detected no members of the Chlorophyta phylum (green algae) in the early- to mid-biofilm stage but a strong domination of the late biofilm stages by these genera (Supplementary Figure S1 B). The changes in bacterial composition and activity were less pronounced compared to plastic soil. In addition, no Cyanobacteria were observed in the forest soil biofilm community.

PE-specific gene expression marks successional biofilm stages on PE and glass

In search of specific pathways during the succession of PE-associated biofilms, we used our KEGG annotated metatranscriptomic data and compared it to a set of 35 known enzymes involved in PE- and/or general polymer colonization (Figs. 4 and 5), five nif genes related to nitrogen fixation (Fig. 6) as well as 133 biofilm related enzymes involved in lipopolysaccharide biosynthesis and extracellular matrix synthesis, respectively (Supplementary Figure S2). Gene expression patterns of the analysed PE-specific genes varied over the course of biofilm succession. Some of the involved pathways showed an overexpression only at specific successional stages, which we verified in detail by DESEQ2 analysis of differential abundance (Supplementary table 3). In general, we detected a later onset of PE-specific pathways in forest soil when compared to the same genes in plastic soil biofilm communities (Fig. 4A and 4B).

During the early colonization in plastic soil (day 2–7) of PE foils, we found a high activity of genes involved in fatty acid degradation while in forest soil this occurred later during the mid-successional stage (day 7–14). For example, transcripts of the acyl-CoA dehydrogenase (ACADM), a putative acyltransferase (atoB), 3-ketoacyl-CoA thiolase (FadA) and a fatty acid oxidation complex subunit alpha (FadJ) were expressed and overly abundant in this initial biofilm stage on PE foil in plastic soil (Fig. 4A). All of these genes are involved in either aerobic long-chain fatty acid degradation (fadB and fadA), or the anaerobic equivalent of the same pathway (fadJ). Even though transcripts of atoB and FadA were detected in one glass sample at day 2, fatty acid degradation was less prevalent on glass than on PE foil in plastic soil for the rest of the biofilm stages (Fig. 4A).

The initial biofilm stage on day 2 was also shaped by genes encoding membrane transporters such as the ATP-binding proteins (ABC transporters) LivF, LivH, LivM and LivG which were upregulated in both glass- and PE biofilm communities in forest- and plastic soil (Fig. 4A and 4B). The long-chain fatty acid transporter FadL and the TRAP-like transport protein yiaO, both involved in the transport of aliphatic hydrocarbons, were active in two initial biofilm samples on PE foil in plastic soil during day 2 and day 7, as well as in a mature PE-associated biofilm sample of day 53 (Fig. 4A). In forest soil, these two transporters were active only in the biofilm samples of day 7–14 (Fig. 4B).

Genes involved in the degradation of n-alkanes were the alkane-monooxygenase (alkM), the S-(hydroxymethyl)glutathione dehydrogenase (frmA), an aldehyde dehydrogenase (ALDH) and the long-chain alkane monooxygenase (ladA), all of which were also active during the early successional stages (day 2–7) of PE-associated biofilm from plastic soil (Fig. 4A) and a bit later (day 7–14) in biofilm communities from forest soil (Fig. 4B). The expression of genes related to n-alkane degradation were therefore highest in early- to mid-successional stages and dropped in activity towards the late biofilm communities. Other enriched pathways were genes involved in the breakdown of secondary plant metabolites and natural terpenes, such as geraniol and citronellol (atuC,atuD and atuF), all of which were active during day 7–14 in forest soil and during the initial successional stage (day 2) in plastic soil (Fig. 4). The only exception was the related isohexenylglutaconyl-CoA hydratase (atuE), which was active exclusively during the late successional stage of day 53 in both soils (Fig. 4).

The mid successional stage in forest soil was characterized by the expression of enzymes involved in fatty acid beta-degradation, such as fadJ (3-hydroxyacyl-CoA dehydrogenase), fadB (Fatty acid oxidation complex subunit alpha) and ACADM (acyl-CoA dehydrogenase), which had their peak of expression on day 14 (Fig. 4). Also, genes from the pathways of geraniol degradation were overexpressed during this phase of biofilm development. Noteworthy in the context of possible PE-degrading pathways, was the overexpression of the extracellular lipase lip, as well as the Acyl-CoA dehydrogenase DCAA, capable of degrading caprolactam – the monomer of Nylon 6. Moreover, we found almost all genes involved in the N-alkane degradation pathway to be overly active at certain biofilm stages (Fig. 4). Identical to genes expressed in plastic soil, the Alkane 1 monooxygenase alkM (candidate for PE-degradation) was active during day 7 and day 14 of biofilm formation and remained downregulated during both the initial biofilm phase (day 2) and the late biofilm succession (day 53). To our surprise, we found the polyethylene terephthalate degrading enzyme PETase to be present and expressed by the biofilm community in their mid-successional stage in forest soil, yet entirely absent in plastic soil.

Overall, the late biofilm samples (day 53) showed only a few genes to be expressed above the average (Fig. 4). With the exception of a single PE sample in plastic soil (Fig. 4A, PE1 day 53) we observed an overall decline in the expression of genes relevant for PE-degradation towards the late successional phases of biofilm growth (Fig. 5). In fact, most genes that were highly active in the preceding biofilm stages were underexpressed in the mature biofilm communities. Out of genes involved in fatty acid beta-oxidation, only fadB and fadN (both Fatty acid oxidation complex subunit alpha) were slightly above the average in the late biofilm stage of plastic soil and forest soil, respectively (Figs. 4 and 5). Other earlier abundant enzymes, such as ABC transporter genes or the PE-specific candidate genes Alkane 1 monooxygenase alkM and the extracellular lipase lip were remarkably underrepresented in the late biofilm stage. Moreover, nonspecific genes involved in the biosynthesis of secondary metabolites (3-oxoacyl-[acyl-carrier protein] reductase FabG and a putative enoyl-CoA hydratase atuE) were overrepresented on day 53 of incubation in both soils. One sample of the biofilm community on PE foil (PE1day53), showed a remarkably high expression of PE-specific genes compared to other samples from the same sampling day (Fig. 4A). These highly expressed genes included ABC transporters (e.g LivH, LivF and LivG/CysA), as well as genes involved in fatty acid metabolism including the extracellular lipase lip (Lactonizing lipase precursor) and fadA (3-ketoacyl-CoA thiolase). In the forest soil, we observed a clear functional pattern that distinguished the different successional biofilm stages from each other in regard to PE-specific genes. (Fig. 4B). Especially genes encoding dehydrogenases like fadE (acyl-CoA dehydrogenase) and frmA (S-(hydroxymethyl)glutathione dehydrogenase) were considerably active compared to later biofilm stages. Unique for the sequenced PE biofilm communities in this early stage, was an overexpression of the extracellularly secreted carboxylic-ester hydrolase phaZ (Esterase PHB depolymerase) in PE communities compared to the biofilm on glass (Fig. 4).

Plastic soil microbial community showed high expression of nif genes to overcome nitrogen limitation on PE in the mature biofilm

Apart from genes related to PE degradation, we detected the expression of genes responsible for microbial nitrogen fixation (Fig. 6) in the biofilm community. Most striking was the increase in expression of a number of nif genes over the course of biofilm succession in plastic soil, which encode a variety of nitrogenases and are an important part in the microbial nitrogen fixation pathway (Fig. 6A).

In plastic soil, the expression of nifD, nifH, nifK, nifN and nifT was the highest in the mature biofilm stage, with nifH reaching the highest gene expression on PE foil after 53 days of incubation (Fig. 5A1). Transcripts of nifD, nifH and nifK were, however, present during the mid-successional biofilm stage on two PE samples (Fig. 6) and on a single glass sample on day 2 of incubation. The highest expression levels were reached on the PE samples at this stage of biofilm succession, even though the biofilm of a single glass sample also expressed four of the nif genes in low abundances. To our surprise, we could not detect a single transcript of nif genes related to nitrogen fixation in our forest soil biofilm community.

Biofilm succession and surface coverage using SEM image analysis

In addition to the transcriptomic data of the microbial biofilm communities on soil-buried PE foil, we used SEM image analysis to visually follow growth and successional development of the biofilm stages in both soils (Fig. 7 and Fig. 8). In order to quantify the surface coverage, we used a threshold-based image analysis which depicted biofilm structures from the background surface and calculated the extent of microbial coverage (Supplementary Figure S3).

An initial biofilm growth was visible on day 2 of incubation in both soil types. Over the course of time, we observed a higher surface coverage of biofilms in plastic soil (Fig. 7) compared to the biofilm growth in forest soil (Fig. 8). Biofilm complexity on the PE surface increased with time of incubation in both soils with a variety of morphological biofilm features. Structures of extracellular polymeric substances (EPS) were detected within the biofilm, where strings connected the thick microbial mats with the surface (Fig. 7: day 53).

We found various shapes of bacterial cells (e.g. singular cocci and rod-shaped bacteria as well as streptococci and tetrads) in close arrangements on the surface and embedded within surface grooves of the PE foil (Figs. 7 and 8). Most apparent were the large fungal hyphae structures from day 14 on and a number of fungal spores on the PE foil. In the late biofilm stage within the plastic soil, we found an increased number of filamentous bacteria as well as streptococci and chains of bacilli (Fig. 7: day 53). In general, the surface coverage of PE samples with biofilm in plastic soil was higher than those incubated in forest soil. The PE surface in plastic soil was covered in thick biofilm after 53 days of incubation, where mean coverage reached 39% of the PE surface (Fig. 7). Especially in the middle stages of biofilm growth (14 days), there were numerous bacterial cell aggregates visible on PE, as well as fungal structures like hyphae and spores which formed a substantial microbial layer on top of the surface. In forest soil, mean surface coverage was highest at day 14 with 2.3% (Fig. 8: day 14) and did not reach higher coverages throughout the incubation. Considering these low coverage numbers, we detected an earlier onset of biofilm development in plastic soil compared to the succession in forest soil similar to the patterns visible in the metatranscriptome data.

Maturation of biofilm outcompetes microbial PE degradation in soil

In both soils, we found a time-dependent change in the taxonomic composition of the microbial community and their PE-specific gene expression pattern. PERMANOVA analysis showed that the period of incubation had a significant effect on the community structure of the active biofilm community, despite the origin of the colonizing microbial community (plastic soil vs. forest soil). The sequenced PE-associated biofilm communities developed over the course of time into a less diverse community compared to their initial successional stage. The successional age of our biofilm had therefore a strong effect on our microbial community composition, independent of the source community (plastic soil vs. forest soil). These findings are consistent with other studies, that showed that bacterial communities on PE surfaces become significantly less diverse over the course of succession due to plastic-specific dynamics within the attached plastic-associated biofilm community [40, 46, 56, 57].

Biofilm succession refers to the gradual change in the composition and structure of a biofilm community over time. In our study, we observed the most significant changes in community structure within the biofilm between day 14 and the last day of incubation (day 53). The microbial community in both soil types became less diverse over time and clustered apart from all other successional stages. Similarly, Harrison and colleagues (2014) demonstrated in their study that in soil bacterial biofilms on LDPE, successional shifts occurred only after a longer time period [56]. In their study from 2021, Wright and colleagues reviewed a similar time-dependent dynamic in microbial biofilms on plastic debris in the ocean [58]. According to their observation, a microbial biofilm forming on plastic debris in aquatic medium can be considered matured and independent from the material’s surface characteristics after two weeks of incubation. Our data showed that indeed most transcripts of plastic-associated taxa were present within the first two sampling points in time (Day 2 and Day 7). In plastic soil, we observed the plastic-degrading taxa Aspergillus, Rhodococcus, Bacillus, Nocardioides, Streptomyces and Nocardia to be present in the early successional stage of the PE biofilm yet in decreasing abundance as the biofilm matured over the course of time. We assume that even though the initial colonization of plastic debris happens more rapidly in aquatic ecosystems, the time required for plastic degradation and adaptations to physico-chemical properties of the polymer are the major ecological drivers of the community composition is similar in the terrestrial plastisphere.

During the process of biofilm succession, there are various selective pressures that can result in such a shift in community composition: It is assumed that the initial phase is dominated by bottom-up selection, where the availability of heterotrophic resources (e.g plastic derived carbon) plays a key role in regulating the community. However, as the biofilm matures, it was shown that both bottom-up and top-down (e.g competition by higher trophic levels) mechanisms play an equal role in regulating the biofilm community [59]. After a first enrichment of plastic-associated taxa during the early successional stages, our biofilm community shifted towards a dominance of phototrophic Cyanobacteria and green algae (Chlorophycae and Trebouxiophyceae), as well as biofilm grazers like Acanthamoeba and Vermamoeba. At this stage of biofilm development, the system was most probably already exhausted of easily available plastic-derived organic matter which led to autotrophic taxa and predators taking over as an example of top-down selection. Although plastisphere research has traditionally focused on heterotrophic bacteria, a few other aquatic studies observed high abundances of a few opportunistic cross-feeders, grazers and phototrophs during biofilm maturation [43, 60, 61]. From our and other study observations, heterotrophic plastic-degrading microorganisms are often contributing to the microbial community in rather low abundances, which makes it hard for them to compete against this top-down pressure by the dominant autotrophs [40, 62]. Furthermore, as biofilm growth becomes thicker, the contact to the plastic surface decreases, which reduces the possibility for direct microbial degradation of the polymer. Therefore, we conclude that once biofilm succession on plastic surfaces shift into a mature stage (day 14–53), dominance of (photo)autotrophs will eventually prevent an efficient plastic-degradation due to top-down selection pressure. Our research shows that this applies not only to marine plastic debris [36] but also for the terrestrial plastisphere.

However, unlike in aquatic systems, our data suggests the development of a soil crust-like community during biofilm maturation. Natural soil crusts are well-described model ecosystems, which are composed of microbial filaments, aggregates, cyanobacteria and algae, embedded in an organic matrix [63, 64]. Due to their extreme tolerance to desiccation and nutrient deficiencies and their high photosynthetic activity, they are able to accumulate organic matter and built a micro habitat independent from the original surface properties [65, 66]. Nevertheless, biological soil crusts need the exposure to sunlight in order to photosynthesise. The PE samples used in our soil incubations were submerged in a shallow soil layer and UV-weathered prior to incubations. The availability of light is therefore a factor that needs to be considered when microbial strategies to the presence of plastic debris in soil are being investigated. This certainly applies to communities attached to plastic debris deposited in the upper soil layers, where photosynthesis is an available autotrophic microbial energy source. We, however, speculate that plastic-associated biofilms in deeper soil layers will underly other selection mechanisms in the microbial community when no photosynthesis is possible. Plastic-associated biofilm communities in deeper soil layers might therefore take a longer time for this metabolic shift, which could give plastic degrading taxa a prolonged time to act on the plastic surface. A detailed analysis of plastic-attached microbial communities in soil considering also the depth of the soil layer might help to further understand the different strategies in the terrestrial plastisphere.

Microbial community from plastic soil is more efficient in colonizing PE surface than forest soil

We predicted the two soil types and their native microbial community to differ in both their colonizing potential and their PE-specificity which was indeed consistent with our observations. Based on our SEM image analysis, a more rapid colonization of the PE foil by microorganisms from the plastic soil community was observed. This trend was reflected by the overall higher biofilm coverage of the biofilm samples from plastic soil and less variation in biofilm associated genes. Recent mining of environmental metagenomes has revealed a correlation between the levels of pollution in a given ecosystem and the abundance of plastic-degrading enzymes, suggesting that areas of high pollution are already driving microbiomes to evolve a potential for plastic degradation [25, 67]. In accordance with these findings, we assume that the exposure to plastic debris had shaped our studied microbial community in plastic soil towards a community of highly efficient colonizers, able to attach to the PE surface quite rapidly. As soil matrices consist of numerous available surfaces on which biofilms can be established (e.g. soil particles and biotic structures), the competition for “attachment” might be less severe, yet biofilm associated bacteria in soil were shown to have fitness advantages when compared to planktonic organisms [68]. In highly plastic polluted soil matrices, however, the competition for a rapid biofilm succession might be higher than in natural soils due to the scarcity of available substrates and the selective pressure to develop a functioning biofilm community including photoautotrophs which fixate carbon and nitrogen into the system. Earlier research has shown that nitrogen fixation is the rate-limiting step to microbial polymer degradation in soil which can be overcome by the establishment of a functioning N-uptake system provided by phototrophs and other N-fixing microorganisms [69, 70].

In our case, we saw an increase of N-fixing phototrophs in the late stages of biofilm succession in plastic soil but not in the forest soil. This increase of the cyanobacterial genera Nostoc, Desmonostoc and Tolypothrix was accompanied by an increase in genes responsible for N fixation like nifD, nifH, nifK, nifN and nifT, all of which were absent in the natural forest soil community. This is also supported by the observation that the N-fixing bacterial taxa Gemmataceae and Gemmata followed the same trend and increased in abundance during the late biofilm stage (day 14–53). This is an interesting finding, as it coincides with earlier research that found cyanobacteria and other N-fixing microorganisms to actually facilitate polymer degradation by providing the missing nitrogen for other hydrocarbon-degrading species [71]. So, in addition to selecting for plastic-degrading capabilities, plastic debris also imposes selective pressure on microbial communities to adapt and overcome the scarcity of heterotrophic energy sources within the plastisphere and in particular nitrogen limitation. Therefore, the establishment of a functional biofilm community on plastic debris is not only dependent on plastic degradation but also on the ability to overcome nutrient limitations, further highlighting the important role of (photo)autotrophs within a plastic-associated biofilm community.

PE-degrading enzymes detected

An earlier metagenomic study of plastic debris in water, concluded that the surface of plastic debris enriches taxa which possess discrete sets of genes compared to those in the surrounding bulk waters [21]. Applying this notion to the terrestrial realm, we expected an enrichment of PE-specific genes on the buried PE foil when compared to the tested glass slides or genes active in the surrounding bulk soils. Therefore, we predicted the plastic soil community to express a larger variety of genes related to PE degradation. In their study from 2021, Zrimec and colleagues found that a high abundance of plastic-degrading microbial enzymes was correlated with a higher degree of plastic pollution in a system [67].

In terms of potentially PE-degrading enzymes, we found the alkane monooxygenase alkB1_2/alkM to be upregulated during the early stages of biofilm succession in plastic soil, which suggests an active hydroxylation of polymer derived compounds during this early phase of the biofilm succession. This coincides with our observed presence of plastic-degrading taxa mostly in the early successional stages, even though we did not identify the taxonomical origin of AlkB and its related genes. Other PE-degrading enzymes worth testing in our metatranscriptome were the laccase derived from Rhodococcus ruber [72] and the manganese peroxidase (mpn) expressed by two lignin-degrading fungi, Phanerochaete chrysosporium and TrarnetesTrametes versicolor [73]. Even though Rhodococus was abundant in this early successional stage, we could not detect transcripts of the previously described laccase in our metatranscriptome.

As a second step in PE degradation and assimilation, Priya and colleagues suggested the existence of dedicated transport systems for short length-PE fragments after external hydroxylation [74]. In this context, different active membrane transporters were found to facilitate the uptake of hydroxylated hydrocarbon compounds [75]. The long-chain fatty acid transporter (FadL) is known to play a major role in alkane transport and was found present in bacteria involved in the biodegradation of xenobiotics [76]. Next to this, Zadjelovic and colleagues showed in their extensive metaproteome study, that both TRAP-like membrane transporters and ATP-binding proteins (ABC transporters) were upregulated in the presence of weathered PE [27]. Moreover, the actinomycete Rhodococcus rhodochrous was reported to assimilate PE oligomers the size of 600 Da through cassettes of ATP-binding proteins [77]. Our detection of high expression of transporter genes (ABC transporters) in our PE-associated microbial biofilm is consistent with these findings, which hints towards an active uptake of plastic-derived oligomers from the PE surface.

Our metatranscriptomic data shows that during the early successional stages, plastic-degrading taxa were present along with the upregulation of plastic-degrading enzymes. We observed the PE-degrading enzymes alkB1/alkM and transporters such as FadL, livG, livF, livH and livM and fatty acid β-oxidation pathway to be active during the maturation of the biofilm. To our surprise, the number of PE-related genes present and expressed in the natural forest soil biofilm community were very similar to the genes expressed in the highly polluted plastic soil community, in contrast to earlier prediction. Moreover, the transcriptional response of the PE-attached biofilm community followed a clear successional pattern over the course of time with distinct phases in which only certain PE-specific pathways were active. Overall, the presence and expression of these enzymes suggest that the microbial community in the plastic soil has the potential to degrade plastic compounds, although their expression patterns over time also suggest that other metabolic dynamics may be involved in shaping the microbial community in the biofilm. We can further conclude from our findings that even pristine forest soils which lack a major exposure to plastic debris, still harbour a variety of plastic-associated genes (PETase, alkB etc..) and therefore carry the enzymatic potential for microbial plastic degradation. In the quest for plastic-degrading enzymes, these findings might shift the attention from highly disturbed ecosystems (e.g landfills) towards natural and diverse soil ecosystems where PE-specific enzymes seem to be likewise present.

In conclusion, our metatranscriptome study provides new insights into the taxonomic and functional dynamics of microbial biofilm succession in the terrestrial plastisphere. We observed a time-dependent shift in the composition of the biofilm community, with plastic-degrading taxa being present in the early successional stage and phototrophic organisms and grazers dominating the later stages. Here we observed a soil crust-like biofilm community as the most matured successional stage on PE in soil, most likely resulting from top-down selection pressure rather than actual plastic-degradation once nitrogen fixation by (photo)autotrophs became the limiting step. These observations also highlight the importance of considering the availability of light as a factor that influences the microbial strategies to plastic pollution in soil. Further research on plastic-attached microbial communities in soil, considering the depth of the polluted soil layer, will be needed to better understand the different physiological strategies of such systems. Our results also imply that even pristine forest soils without prior exposure to plastic debris harbour and express a variety of plastic-associated genes (PETase, alkB etc..). However, despite the apparent metabolic potential of such natural soil communities, microbiomes which were previously exposed to plastic debris seem to be primed towards the colonization of PE and most likely also PE degradation. Furthermore, we observed an increase in genes related to nitrogen fixation in the plastic soil community but not in the forest soil community. With regard to this observation, we demonstrated that the high abundance of nitrogen fixation genes is another essential metabolic adaptation of biofilm communities in highly plastic polluted soil environments. Overall, these findings highlight the complex and dynamic nature of plastic-associated biofilms and suggest that different stages of biofilm development may be associated with different microbial functions and potential for plastic degradation. This understanding is crucial in the ongoing quest for the discovery of plastic-degrading microorganisms and shows that the whole community dynamics should be considered for mitigating plastic pollution in soil ecosystems.

ASV

Amplicon sequence variant

Dalton (unit)

HDPE

High-density polyethylene

Polyethylene

PET

Polyethylene terephthalate

SEM

Scanning electron microscopy

Acknowledgements

We would like to thank the interdisciplinary consortium of the BMBF research focus “Plastics in the environment” and the BMBF for their generous support. Special thanks go to the BAM Division 6.6 who provided us with the HDPE material and to Volker Wachtendorf from BAM Division 7.5 for the excellent UV-weathering of the material. We are also very grateful for the contribution of Liane G. Benning, who has helped us with her SEM expertise and to the PISA facility and Helmholtz Recruiting Initiative (Award No. I-044-16-0).

Authors’ contributions

JM designed the study, collected samples, analysed the data and wrote the manuscript. AB analysed the data and wrote the manuscript. RB performed SEM analysis and contributed to the SEM manuscript section. SL contributed resources and expertise. DW contributed resources and expertise. All authors edited the manuscript and approved the final draft.

Ethics approval

Not applicable

Availability of data and material

All data generated or analysed during this study are included in this published article [and its supplementary information files]. Metagenomic and metatranscriptome raw data was uploaded to the ENA under project accession number PRJEB57718.

Funding

This research was generously funded by Bundesministerium für Bildung und Forschung (ENSURE, grant number 02WPL1449D). SEM analysis was provided and facilitated by PISA facility and Helmholtz Recruiting Initiative (Award No. I-044-16-0)

Competing Interests statement

The authors declare no conflict of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript, or in the decision to publish the results.

Jambeck JR, Geyer R, Wilcox C, Siegler TR, Perryman M, Andrady A, et al. Plastic waste inputs from land into the ocean. Science (1979) 2015; 347: 768–771.
Nizzetto L, Futter M, Langaas S. Are Agricultural Soils Dumps for Microplastics of Urban Origin? Environ Sci Technol . 2016.
Jacoby R, Peukert M, Succurro A, Koprivova A, Kopriva S. The role of soil microorganisms in plant mineral nutrition—current knowledge and future directions. Front Plant Sci 2017; 8.
Thoha TB, Thomas BT, Olanrewaju-Kehinde DSK, Popoola OD, James ES. Degradation of Plastic and Polythene Materials by Some Selected Microorganisms Isolated from Soil. World Appl Sci J 2015; 33: 1888–1891.
Huang D, Xu Y, Lei F, Yu X, Ouyang Z, Chen Y, et al. Degradation of polyethylene plastic in soil and effects on microbial community composition. J Hazard Mater 2021; 416.
Sajjad M, Huang Q, Khan S, Khan MA, Liu Y, Wang J, et al. Microplastics in the soil environment: A critical review. Environ Technol Innov . 2022. Elsevier B.V. , 27: 102408
Kublik S, Gschwendtner S, Magritsch T, Radl V, Rillig MC, Schloter M. Microplastics in soil induce a new microbial habitat, with consequences for bulk soil microbiomes. Front Environ Sci 2022; 10: 1478.
Ju Z, Du X, Feng K, Li S, Gu S, Jin D, et al. The Succession of Bacterial Community Attached on Biodegradable Plastic Mulches During the Degradation in Soil. Front Microbiol 2021; 12: 3949.
Sun Y, Shi J, Wang X, Ding C, Wang J. Deciphering the Mechanisms Shaping the Plastisphere Microbiota in Soil. mSystems 2022; 7.
Maclean J, Mayanna S, Benning LG, Horn F, Bartholomäus A, Wiesner Y, et al. The terrestrial plastisphere: Diversity and polymer-colonizing potential of plastic-associated microbial communities in soil. Microorganisms 2021; 9.
Lehmann A, Leifheit EF, Gerdawischke M, Rillig MC. Microplastics have shape-and polymer-dependent effects on soil aggregation and organic matter loss-an experimental and meta-analytical approach.
Jazaei F, Chy TJ, Salehi M. Can Microplastic Pollution Change Soil-Water Dynamics? Results from Controlled Laboratory Experiments. 2022.
Moreno‐Jiménez E, Leifheit EF, Plaza C, Feng L, Bergmann J, Wulf A, et al. Effects of microplastics on crop nutrition in fertile soils and interaction with arbuscular mycorrhizal fungi. Journal of Sustainable Agriculture and Environment 2022; 1: 66–72.
Abel De Souza Machado A, Lau CW, Kloas W, Bergmann J, Bachelier JB, Faltin E, et al. Microplastics Can Change Soil Properties and Affect Plant Performance. 2019.
Brussels. COMMUNICATION FROM THE COMMISSION TO THE EUROPEAN PARLIAMENT, THE COUNCIL, THE EUROPEAN ECONOMIC AND SOCIAL COMMITTEE AND THE COMMITTEE OF THE REGIONS A European Strategy for Plastics in a Circular Economy. COM 2018; 16.
Geyer R, Jambeck JR, Law KL. Production, use, and fate of all plastics ever made. 2017.
Steinmetz Z, Wollmann C, Schaefer M, Buchmann C, David J, Tröger J, et al. Plastic mulching in agriculture. Trading short-term agronomic benefits for long-term soil degradation? Science of the Total Environment 2016; 550: 690–705.
Hou L, Xi J, Liu J, Wang P, Xu T, Liu T, et al. Biodegradability of polyethylene mulching film by two Pseudomonas bacteria and their potential degradation mechanism. Chemosphere 2022; 286: 131758.
Restrepo-Flórez JM, Bassi A, Thompson MR. Microbial degradation and deterioration of polyethylene - A review. Int Biodeterior Biodegradation . 2014.
Amobonye A, Bhagwat P, Singh S, Pillai S. Plastic biodegradation: Frontline microbes and their enzymes. Science of the Total Environment 2021; 759.
Bryant JA, Clemente TM, Viviani DA, Fong AA, Thomas KA, Kemp P, et al. Diversity and Activity of Communities Inhabiting Plastic Debris in the North Pacific Gyre. Ecological and Evolutionarz Science 2016; 1: e00024-16.
Danso D, Schmeisser C, Chow J, Zimmermann W, Wei R, Leggewie C, et al. New insights into the function and global distribution of polyethylene terephthalate (PET)-degrading bacteria and enzymes in marine and terrestrial metagenomes. Appl Environ Microbiol 2018; 84: AEM.02773-17.
Pinto M, Zhao Z, Klun K, Libowitzky E, Herndl GJ. Microbial Consortiums of Putative Degraders of Low-Density Polyethylene-Associated Compounds in the Ocean. mSystems 2022; 7.
Mitzscherling J, MacLean J, Lipus D, Bartholomäus A, Mangelsdorf K, Lipski A, et al. Paenalcaligenes niemegkensis sp. nov., a novel species of the family Alcaligenaceae isolated from plastic waste. Int J Syst Evol Microbiol 2022; 72.
Mitzscherling J, MacLean J, Lipus D, Bartholomäus A, Mangelsdorf K, Lipski A, et al. Nocardioides alcanivorans sp. nov., a novel hexadecane-degrading species isolated from plastic waste. Int J Syst Evol Microbiol 2022; 72.
Sowmya H V., Ramalingappa, Krishnappa M, Thippeswamy B. Degradation of polyethylene by Penicillium simplicissimum isolated from local dumpsite of Shivamogga district. Environ Dev Sustain 2015; 17: 731–745.
Zadjelovic V, Erni-Cassola G, Obrador-Viel T, Lester D, Eley Y, Gibson MI, et al. A mechanistic understanding of polyethylene biodegradation by the marine bacterium Alcanivorax. J Hazard Mater 2022; 436: 129278.
Zampolli J, Zeaiter Z, Di Canito A, Di Gennaro P. Genome analysis and -omics approaches provide new insights into the biodegradation potential of Rhodococcus. Appl Microbiol Biotechnol 2019; 103: 1069–1080.
Gravouil K, Ferru-Clément R, Colas S, Helye R, Kadri L, Bourdeau L, et al. Transcriptomics and Lipidomics of the Environmental Strain Rhodococcus ruber Point out Consumption Pathways and Potential Metabolic Bottlenecks for Polyethylene Degradation. Environ Sci Technol 2017; 51: 5172–5181.
Pérez-García P, Danso D, Zhang H, Chow J, Streit WR. Exploring the global metagenome for plastic-degrading enzymes. Methods in Enzymology. 2021. Academic Press Inc., pp 137–157.
Dussud C, Ghiglione J-F. La dégradation des plastiques en mer - sfecologie.org. 2014.
Lucas N, Bienaime C, Belloy C, Queneudec M, Silvestre F, Nava-Saucedo JE. Polymer biodegradation: Mechanisms and estimation techniques - A review. Chemosphere 2008; 73: 429–442.
Syranidou E, Karkanorachaki K, Amorotti F, Franchini M, Repouskou E, Kaliva M, et al. Biodegradation of weathered polystyrene films in seawater microcosms. Sci Rep 2017; 7: 1–12.
Pathak VM, Navneet VM. Review on the current status of polymer degradation: a microbial approach. Bioresour Bioprocess 2017; 4: 15.
Puglisi E, Romaniello F, Galletti S, Boccaleri E, Frache A, Cocconcelli PS. Selective bacterial colonization processes on polyethylene waste samples in an abandoned landfill site. Sci Rep 2019; 9: 14138.
Rummel CD, Lechtenfeld OJ, Kallies R, Benke A, Herzsprung P, Rynek R, et al. Conditioning Film and Early Biofilm Succession on Plastic Surfaces. Cite This: Environ Sci Technol 2021; 55: 11006–11018.
Renner LD, Weibel DB. Physicochemical regulation of biofilm formation. Materials Research Society 2011.
Datta MS, Sliwerska E, Gore J, Polz MF, Cordero OX. Microbial interactions lead to rapid micro-scale successions on model marine particles. Nat Commun 2016; 7: 11965.
Montazer Z, Najafi MBH, Levin DB. Challenges with verifying microbial degradation of polyethylene. Polymers (Basel) 2020; 12.
Pinto M, Langer TM, Hüffer T, Hofmann T, Herndl GJ. The composition of bacterial communities associated with plastic biofilms differs between different polymers and stages of biofilm succession. PLoS One 2019; 14: e0217165.
Gewert B, Plassmann MM, Macleod M. Pathways for degradation of plastic polymers floating in the marine environment. Environmental Science 2015; 17: 1513.
Joshi SJ, Kant Bhatia S, University K, Korea S, Wu W-M, Levin DB, et al. Microbial and Enzymatic Degradation of Synthetic Plastics. 2020.
Dey S, Rout AK, Behera BK, Ghosh K. Plastisphere community assemblage of aquatic environment: plastic-microbe interaction, role in degradation and characterization technologies. Environ Microbiome 2022; 17: 1–21.
Harcombe W, Mitri S, Mccully AL, Shorten PR, Smith NW, Altermann E, et al. The Classification and Evolution of Bacterial Cross-Feeding. Frontiers in Ecology and Evolution | www.frontiersin.org 2019; 1: 153.
Wright RJ, Bosch R, Langille MG, Christie-Oleza JA. A Multi-OMIC Characterisation of Biodegradation and Microbial Community Succession Within the PET Plastisphere. 2020.
Oberbeckmann S, Labrenz M. Marine Microbial Assemblages on Microplastics: Diversity, Adaptation, and Role in Degradation. 2019.
Huang Y, Zhao Y, Wang J, Zhang M, Jia W, Qin X. LDPE microplastic films alter microbial community composition and enzymatic activities in soil. Environmental Pollution 2019; 254: 112983.
Dassanayake Id RP, Falkenberg SM, Stasko JA, Shircliff AL, Lippolis JD, Briggs RE. Identification of a reliable fixative solution to preserve the complex architecture of bacterial biofilms for scanning electron microscopy evaluation. 2020.
Kieser S, Brown J, Zdobnov EM, Trajkovski M, McCue LA. ATLAS: A Snakemake workflow for assembly, annotation, and genomic binning of metagenome sequence data. BMC Bioinformatics 2020; 21.
Hyatt D, Chen G-L, Locascio PF, Land ML, Larimer FW, Hauser LJ. Prodigal: prokaryotic gene recognition and translation initiation site identification. 2010.
Huerta-Cepas J, Szklarczyk D, Heller D, Hernández-Plaza A, Forslund SK, Cook H, et al. EggNOG 5.0: A hierarchical, functionally and phylogenetically annotated orthology resource based on 5090 organisms and 2502 viruses. Nucleic Acids Res 2019; 47: D309–D314.
Cantalapiedra CP, Hern̗andez-Plaza A, Letunic I, Bork P, Huerta-Cepas J. eggNOG-mapper v2: Functional Annotation, Orthology Assignments, and Domain Prediction at the Metagenomic Scale. Mol Biol Evol 2021; 38: 5825–5829.
Langmead B, Salzberg SL. Fast gapped-read alignment with Bowtie 2. Nat Methods 2012; 9: 357–359.
Yilmaz P, Parfrey LW, Yarza P, Gerken J, Pruesse E, Quast C, et al. The SILVA and ‘all-species Living Tree Project (LTP)’ taxonomic frameworks. Nucleic Acids Res 2014; 42.
Yoshida S, Hiraga K, Takehana T, Taniguchi I, Yamaji H, Maeda Y, et al. A bacterium that degrades and assimilates poly(ethylene terephthalate). Science (1979) 2016; 351.
Harrison JP, Schratzberger M, Sapp M, Osborn AM. Rapid bacterial colonization of low-density polyethylene microplastics in coastal sediment microcosms. BMC Microbiol 2014; 14: 1–15.
Wallbank JA, Lear G, Kingsbury JM, Weaver L, Doake F, Smith DA, et al. Into the Plastisphere, Where Only the Generalists Thrive: Early Insights in Plastisphere Microbial Community Succession. Front Mar Sci 2022; 9: 626.
Wright RJ, Langille MGI, Walker TR. Food or just a free ride? A meta-analysis reveals the global diversity of the Plastisphere. ISME J 2021; 15: 789–806.
Tobias-Hünefeldt SP, Wenley J, Baltar F, Morales SE. Ecological drivers switch from bottom–up to top–down during model microbial community successions. ISME J 2021; 15: 1085–1097.
Wright RJ, Langille MGI, Walker TR. Food or just a free ride? A meta-analysis reveals the global diversity of the Plastisphere. ISME J 2021; 15: 789–806.
Kesy K, Oberbeckmann S, Kreikemeyer B, Labrenz M. Spatial Environmental Heterogeneity Determines Young Biofilm Assemblages on Microplastics in Baltic Sea Mesocosms. Spatial Environmental Heterogeneity Determines Young Biofilm Assemblages on Microplastics in Baltic Sea Mesocosms Front Microbiol 2019; 10: 1665.
Kirstein IV, Wichels A, Gullans E, Krohne G, Gerdts G. The plastisphere – Uncovering tightly attached plastic “specific” microorganisms. PLoS One 2019; 14: 1–17.
Belnap J, Büdel B, Lange OL. Biological Soil Crusts: Characteristics and Distribution. Ecological Studies .
Eldridge D. Ecology and Management of Biological Soil Crusts: Recent Developments and Future Challenges. Source: The Bryologist . 2000. Winter.
Kuang X, Peng L, Chen S, Peng C, Song H. Immobilization of metal(loid)s from acid mine drainage by biological soil crusts through biomineralization. J Hazard Mater 2023; 443: 130314.
Abed RMM, Al-Kindi S. Effect of disturbance by oil pollution on the diversity and activity of bacterial communities in biological soil crusts from the Sultanate of Oman. Applied Soil Ecology 2017; 110: 88–96.
Zrimec J, Kokina M, Jonasson S, Zorrilla F, Zelezniak A. Plastic-Degrading Potential across the Global Microbiome Correlates with Recent Pollution Trends. mBio 2021; 12.
Kragh KN, Hutchison JB, Melaugh G, Rodesney C, Roberts AEL, Irie Y, et al. Role of multicellular aggregates in biofilm formation. mBio 2016; 7.
Osono T, Takeda H. Decomposition of organic chemical components in relation to nitrogen dynamics in leaf litter of 14 tree species in a cool temperate forest. Ecol Res 2005; 20: 41–49.
Tanunchai B, Kalkhof S, Guliyev V, Wahdan SFM, Krstic D, Schädler M, et al. Nitrogen fixing bacteria facilitate microbial biodegradation of a bio-based and biodegradable plastic in soils under ambient and future climatic conditions. Environ Sci Process Impacts 2022; 24: 233–241.
Abed RMM, Köster J. The direct role of aerobic heterotrophic bacteria associated with cyanobacteria in the degradation of oil compounds. Int Biodeterior Biodegradation 2005; 55: 29–37.
Santo M, Weitsman R, Sivan A. The role of the copper-binding enzyme - laccase - in the biodegradation of polyethylene by the actinomycete Rhodococcus ruber. Int Biodeterior Biodegradation 2013; 84: 204–210.
Johansson T, Nyman PO. A cluster of genes encoding major isozymes of lignin peroxidase and manganese peroxidase from the white-rot fungus Trametes versicolor. Gene 1996; 170: 31–38.
Priya A, Dutta K, Daverey A. A comprehensive biotechnological and molecular insight into plastic degradation by microbial community. Journal of Chemical Technology and Biotechnology 2022; 97: 381–390.
Rosa LT, Bianconi ME, Thomas GH, Kelly DJ. Tripartite ATP-Independent Periplasmic (TRAP) Transporters and Tripartite Tricarboxylate Transporters (TTT): From Uptake to Pathogenicity. Front Cell Infect Microbiol 2018; 8.
Van Den Berg B. The FadL family: unusual transporters for unusual substrates. Curr Opin Struct Biol 2005; 15: 401–407.
Mercier A, Gravouil K, Aucher W, Brosset-Vincent S, Kadri L, Colas J, et al. Fate of eight different polymers under uncontrolled composting conditions: relationships between deterioration, biofilm formation and the material surface properties. 2017.

No competing interests reported.

SupplementaryFigureS1.png
Supplementary Figure S1: Taxonomy of the eukaryotic fraction of the biofilm community in plastic soil (A) and forest soil (B). The relative abundance of the top 50 most abundant bacterial ASV’s are shown at the genus level The rows show their corresponding genera, colours represent their phyla. Size of the bubbles depicts the relative SSU rRNA abundance in the metatranscriptome. X-axis labels show PE for polyethylene, GL for glass and number 1 and 2 correspond to the biological replicates.
SupplementaryFigureS2.png
Supplementary Figure S2: Gene expression levels of biofilm-specific genes present in the biofilm communities of forest soil A) and plastic soil B) over the course of time. Gene expression counts of biofilm-specific genes are normalisedby Z score transformation and grouped by metabolic function. Sample replicates per day are depicted as GL (glass) and PE (PE foil).
FigureS3SEM.pdf
Supplementary Figure S3: SEM image analysis of biofilm coverage on PE vs. glass on day 2 and day 53 of incubation in forest soil and plastic soil. Surface coverage of all samples was calculated at the same magnification with Fiji ImageJ version 1.53q using a Kuwabara filter (radius 3) and the Triangle algorithm for threshold analysis of the biofilm coverage. The left column shows the original SEM image and the left column shows the processed image for analysis.
SupplementaryTable1.xlsx
Supplementary Table S1. Metatranscriptome of HDPE-attached vs. glass-attached biofilm in two soil types during 53 days. Unfiltered and filtered number of reads per sample. Filter excluded chloroplasts and mitochondrial RNA reads
SupplementaryTable2.xlsx
Supplementary Table S2: Metatranscriptome of HDPE-attached biofilm. Gene expression analysis of genes relevant for PE-degradation (Table S2a) biofilm formation (Table S2b) and nitrogen fixation (Table S2c) over a course of 53 days in two soil types (forest vs. plastic)
SupplementaryTable3.xlsx
Supplementary Table S3: Differentially expressed gene analysis (DESEQ) of genes expressed in three different biofilm stages (early, middle, mature) in plastic soil and forest soil.

Download PDF

Version 1

posted

You are reading this latest preprint version

Metatranscriptomics of microbial biofilm succession on HDPE foil: uncovering plastic-degrading potential in soil communities

Status:

Version 1

Abstract

Figures

Introduction

Material & Methods

Data analysis

Results

Discussion

Conclusion

Abbreviations

Declarations

References

Additional Declarations

Supplementary Files

Status:

Version 1