The comparative plastisphere microbial community profile at Kung Wiman beach unveils potential plastic-specific degrading microorganisms

Collection of seawater, sand, and plastics samples

Sixteen samples of plastic debris were collected from the beach surface. Togethering with, four sand samples were collected from four different locations on Kung Wiman Beach according to Fig. S1. All were kept in zip lock plastic bag. For microbiome analysis, one-gram samples of plastics and sand were preserved in ampoules filled with DNA/RNA Shield solution (Zymo Research, Tustin, CA, USA) and stored at 4 °C. Seawater was collected from the near-shore area and stored in a gallon bottle at room temperature for media preparation, dilution, and salinity experiment. Then, all samples were transported to the laboratory in an iced container.

Determination of the physical properties of sand and seawater

Four sand samples were incubated at 60 °C for 24 h until completely dry. Next, approximately 500 g of the dried sand was sifted through a sieve shaker with six sieves of different sizes (63, 125, 250, 500 μm, 1, and 2 mm) for 10 min. Then, the sand samples remaining on each sieve were weighed, and the particle size fraction was calculated.

The organic carbon content of the sand particles was determined using the modified Walkley–Black method (Loring & Rantala, 1992). The dried sand samples were sifted through a 2 mm sieve and then finely and homogeneously crushed using an agate mortar. The accuracy of the organic carbon content in the sand samples was evaluated using dextrose (C₆H₁₂O₆) as the standard. It is important to note that 0.01 g of dextrose contains 39.99% carbon.

The pH and salinity measurements were conducted using a pH meter and salinity refractometer, respectively. The specific gravity was also calculated.

Plastics type analysis by Fourier-transform infrared (FT-IR) spectroscopy

Sixteen plastic samples were categorized based on their collection location. The plastic types were examined by attenuated total reflectance FT-IR (ATR-FTIR) using a Nicolet 6700 FT-IR spectrometer (Thermo Fisher Scientific, Waltham, MA, USA). The analysis was conducted within a wave number range of 650–4,000 cm⁻¹, with a resolution of 32 cm⁻¹. Each sample was analyzed 32 times. The obtained spectra were compared with the standard peaks of various plastics (Jung et al., 2018).

Culture-dependent method

Microorganisms were extracted from the samples using the ultrasonic method. Plastic samples were cut into three–four small pieces, placed into 10 mL of sterile seawater, and vortexed for 1 min. The extraction of microorganisms from the plastic sheet was performed in an Ultrasonic Cleaner E100H (Elma, Singen, Germany) for 5 min, followed by vortexing for 1 min. For the sand samples, approximately 1 g or 1 spatula of sand was added to 9 mL of sterile seawater and vortexed for 1 min. Therefore, a microbial solution was obtained from each plastic and sand sample. A total of 20 samples of the extracted microbial solution underwent ten-fold serial dilution and 100 µl of the microbial solution were spread on marine agar (MA) supplemented with cycloheximide to inhibit fungal growth and potato dextrose agar (PDA) supplemented with nalidixic acid to inhibit bacteria growth. The cultured microorganisms were incubated at room temperature for 1 month. Then, the distinct colony characteristics of microorganisms from each sample were collected and counted, and the number of microbial colonies (colony-forming unit (CFU)/mL) was calculated.

16S rRNA/ITS gene amplification and identification

The genomic DNA from the microorganism was extracted by the microwave method with adapted condition from the previous study (Dashti et al., 2009). Bacterial colonies were dissolved in 500 µl of 1X TE buffer and were heated for 10 s by microwave oven. After centrifugation for 5 min at 13,000 rpm, the supernatant was used for the template of PCR. Amplification reactions were prepared to final volume of 20 µl containing 2X PCR Master mix solutions (i-Taq™; iNtRON Biotechnology, Seongnam, Korea), 10 pmols of forward and reverse primer, and template DNA. For bacteria, the 16S rRNA region was polymerase chain reaction (PCR)-amplified using the primers 27F (5′-AGAGTTTGATCMTGGCTCAG-3′) and 1492R (5′-TACGGYTACCTTGTTACGACTT-3′). For fungi, the ITS region was PCR-amplified using the primers ITS4 (5′-TCCTCCGCTTATTGATATGC-3′) and ITS5 (5′-GGAAGTAAAAGTCGTAACAAGG-3′). The PCR conditions consisted of an initial denaturation at 95 °C for 5 min, followed by 30 cycles involving denaturation at 95 °C for 30 s, annealing at 52 °C for 30 s, and extension at 72 °C for 30 s. The process concluded with a final extension step at 72 °C for 5 min. The PCR products were verified through agarose gel electrophoresis and subsequently purified using the AccuPrep PCR Purification Kit (Bioneer, Daejeon, Korea). The purified PCR products were subjected to Sanger sequencing, performed by U2Bio Company (Bangkok, Thailand). The resulting partial 16S rRNA and ITS sequences had approximate lengths of 1,500 and 700 bp, respectively. The bacterial and fungal sequences were identified using EzBiocloud (Yoon et al., 2017) and UNITE (Kõljalg et al., 2013), respectively. The nucleotide sequences have been deposited in the GenBank database under accession numbers OR185572–OR185595 (bacteria) and OR205218–OR205249 (fungi).

A phylogenetic tree was constructed using MEGA11 software (Tamura, Stecher & Kumar, 2021). The evolutionary history was inferred using the neighbor-joining method (Saitou & Nei, 1987). The bootstrap consensus tree inferred from 1,000 replicates was taken to represent the evolutionary history of the analyzed taxa. Branches corresponding to partitions reproduced in <50% of bootstrap replicates are collapsed. The evolutionary distances were computed from 58 nucleotide sequences using the maximum composite likelihood method and were in the units of the number of base substitutions per site. The analysis included all codon positions (1^st, 2^nd, 3^rd, and noncoding). All ambiguous positions were removed for each sequence pair (pairwise deletion option). The final dataset comprised 852 positions.

Preparation of MSM agar with emulsified plastic

Plastic degradation properties were tested on mineral salts medium (MSM) agar supplemented with 0.5% emulsified plastic: PS (Sigma-Aldrich, St. Louis, MO, USA), PVC (Sigma-Aldrich, St. Louis, MO, USA), PLA (Goodfellow, England), or bis (2-hydroxyethyl) terephthalate (BHET; Sigma-Aldrich, St. Louis, MO, USA). Two grams of each plastic was put into a 250 mL flask and dissolved with 60 mL of dichloromethane (Honeywell, Charlotte, NC, USA) and heated at 55 °C and agitated with a magnetic stirrer before 15 mL of 2% sarkosyl (Wako Pure Chemical, Osaka, Japan) was added. For BHET was dissolved with dimethyl sulfoxide (Thermo Fisher Scientific, Waltham, MA, USA) and heated only. The plastic mixtures were processed with a VC/VCX 750 High-Intensity Ultrasonic Processor (Sonics, Oklahoma City, OK, USA) at 60% amplitude, 20 s run time, and 10 s rest time for a total time of 10 min. Next, 100 mL of distilled water was added to each plastic solution and mixed for 10 min until it turned milky white. Then, the plastic solution was adjusted in volume to 400 mL with distilled water to obtain a 0.5% plastic emulsion for all four plastic types.

The MSM agar was prepared by adding 0.5 g of yeast extract, 2 g of (NH₄)₂SO₄, 1 g of K₂HPO₄, 0.5 g of KH₂PO₄, 0.25 g of MgSO₄.7H₂O, and 7.5 g of agar in 400 mL of distilled water. MSM agar supplemented with PS or PVC was prepared by adding 100 mL of emulsified PS or PVC to the 400 mL of MSM before sterilization in an autoclave. MSM agar supplemented with PLA or BHET was prepared by adding 100 mL of emulsified PLA or BHET plastic to the 400 mL of MSM after sterilization in an autoclave.

Plastic degrading properties test

Each isolated bacterium/fungus colony was tested on MSM agar supplemented with each emulsified plastic. Bacterial colonies were streaked on the agar, while fungal colonies were cut off and placed on the agar. The plate was further incubated at room temperature for 1 month, then the hydrolysis capacity (HC) was calculated as follows:

$$\mathrm{H}\mathrm{C}\mathrm{v}\mathrm{a}\mathrm{l}\mathrm{u}\mathrm{e}={\displaystyle \frac{\mathrm{d}\mathrm{i}\mathrm{a}\mathrm{m}\mathrm{e}\mathrm{t}\mathrm{e}\mathrm{r}\mathrm{o}\mathrm{f}\mathrm{t}\mathrm{h}\mathrm{e}\mathrm{c}\mathrm{l}\mathrm{e}\mathrm{a}\mathrm{r}\mathrm{z}\mathrm{o}\mathrm{n}\mathrm{e}\mathrm{a}\mathrm{r}\mathrm{o}\mathrm{u}\mathrm{n}\mathrm{d}\mathrm{t}\mathrm{h}\mathrm{e}\mathrm{b}\mathrm{a}\mathrm{c}\mathrm{t}\mathrm{e}\mathrm{r}\mathrm{i}\mathrm{a}\mathrm{l}\mathrm{c}\mathrm{o}\mathrm{l}\mathrm{o}\mathrm{n}\mathrm{y}}{\mathrm{d}\mathrm{i}\mathrm{a}\mathrm{m}\mathrm{e}\mathrm{t}\mathrm{e}\mathrm{r}\mathrm{o}\mathrm{f}\mathrm{t}\mathrm{h}\mathrm{e}\mathrm{b}\mathrm{a}\mathrm{c}\mathrm{t}\mathrm{e}\mathrm{r}\mathrm{i}\mathrm{a}\mathrm{l}\mathrm{c}\mathrm{o}\mathrm{l}\mathrm{o}\mathrm{n}\mathrm{y}}}$$

16S and ITS amplicon sequencing

Four plastic debris from each plastic type (PS, PP, PET) and four sand samples from Kung Wiman Beach were performed in 16S and ITS amplicon sequencing analysis. These samples were processed and analyzed by Zymo Research Corporation using the ZymoBIOMICS Targeted Sequencing Service (Zymo Research, Tustin, CA, USA). The ZymoBIOMICS-96 MagBead DNA Kit (Zymo Research, Tustin, CA, USA) was used to extract DNA using an automated platform. The targeted bacterial 16S ribosomal RNA sequencing library was prepared using the Quick-16S NGS Library Prep Kit (Zymo Research, Tustin, CA, USA); the 16S primers (341F; 5′-CCTACGGGDGGCWGCAG-3′ and 806R; 5′-GACTACNVGGGTMTCTAATCC-3′) amplified the V3–V4 region of the 16S rRNA gene. The targeted fungal ITS sequencing library was also prepared using the Quick-16S NGS Library Prep Kit but with the 16S primers replaced by custom ITS2 primers (ITS3-F; 5′-GCATCGATGAAGAACGCAGC-3′ and ITS4-R; 5′-TCCTCCGCTTATTGATATGC-3′). The sequencing library was generated through a novel library preparation process, wherein PCR reactions were performed in real-time PCR machines to control cycles and, consequently, limit the formation of PCR chimeras. Quantification of the final PCR products was performed using quantitative PCR fluorescence readings, and the products were combined based on equal molarity. The resulting pooled library underwent purification using the Select-a-Size DNA Clean & Concentrator Kit (Zymo Research, Tustin, CA, USA) and was subsequently quantified using TapeStation (Agilent Technologies, Santa Clara, CA, USA) and Qubit (Thermo Fisher Scientific, Waltham, MA, USA). The ZymoBIOMICS Microbial Community Standard (Zymo Research, Tustin, CA, USA) served as a positive control for each DNA extraction. Additionally, the ZymoBIOMICS Microbial Community DNA Standard (Zymo Research, Tustin, CA, USA) was employed as a positive control for each specific library preparation. Negative controls were included to evaluate the bioburden level throughout the wet-lab process. The final library was subjected to sequencing on an Illumina MiSeq system, using the v3 reagent kit (600 cycles) along with a 10% PhiX spike-in. The 16S and ITS amplicon sequencing raw sequence reads have been deposited in the GenBank database under accession numbers PRJNA997562.

Microbiome analyses

Unique amplicon sequence variants were identified from raw reads through the DADA2 pipeline (Callahan et al., 2016) by Zymo Research Corporation. The DADA2 pipeline was also employed to eliminate potential sequencing errors and chimeric sequences. Taxonomy assignment was carried out using Uclust from QIIME (v.1.9.1) with the Zymo Research Database, a 16S database, or ITS internally designed and curated as a reference. Composition visualization, alpha-diversity, and beta-diversity analyses were conducted with QIIME (v.1.9.1) (Caporaso et al., 2010). In cases where applicable, taxonomies exhibiting significant abundances in different groups were identified through linear discriminant analysis effect size (LEfSe) (Segata et al., 2011) using default settings. Other analyses were performed with internal scripts, including heatmaps, Taxa2ASV Decomposer, and principal coordinates analysis (PCoA) plots. To re-analyze non-classified fungal sequences, the raw ITS sequence reads in FASTQ format (provided by Zymo Research Corporation, Tustin, CA, USA) were further processed using the DADA2 pipeline to generate amplicon sequence variant (ASV). The total ITS amplicon sequences in each sample were determined using QIIME 2 (v.2022.11). Subsequently, the consensus sequences from each ASV were analyzed using the BLAST tool against the GenBank database (Altschul et al., 1990).

The statistical analysis was conducted by our researcher group using QIIME 2 (v.2022.11). Alpha diversity, including Shannon indices, was assessed using Kruskal-Wallis rank-sum tests to evaluate differences in microbial diversity among each sample. Principal Coordinates Analysis (PCoA) based on Bray-Curtis dissimilarities was employed to visualize beta diversity, assessing variations in microbial diversity between samples, and permutational multivariate analysis of variance (PERMANOVA) was employed to assess statistical significance. A p-value of <0.05 was considered statistically significant in all statistical experiments.

Sand, seawater, and plastics type analysis

The illustration showing the experimental design was shown in Fig. 1. Twenty samples of different plastic types and sand were collected at four different locations around Kung Wiman Beach in Chanthaburi Province, Thailand (Fig. S1): GPS point 1 = 12°60′29.50′N 101°87′71.24′E; GPS point 2 = 12°60′33.29″N 101°87′68.61″E; GPS point 3 = 12°60′45.31″N 101°87′51.91″; and GPS point 4 = 12°60′33.43″N 101°87′68.62″E.

The particle size of the four sand samples was mostly fine sand (125–250 µm) and medium sand (250–500 µm), with the organic carbon content between 0.04% and 0.09% (Fig. S2 and Table S1). The seawater’s pH was 7.63 and its salinity was 29 psu.

Sixteen plastic samples (Table S2) were identified by ATR-FTIR. By comparing the plastic samples’ ATR-FTIR peaks with the standard peaks of various plastics as follows: Polypropylene (2,950, 2,915, 2,838, 1,455, 1,377, 1,166, 972, 840, and 808 cm⁻¹), Polystyrene (3,024, 2,847, 1,601, 1,492, 1,451, 1,027, and 694 cm⁻¹), Polyethylene terephthalate (1,713, 1,241, 1,094, and 720 cm⁻¹), and high-density PE (2,915, 2,845, 1,462, 730, and 717 cm⁻¹), revealed that five samples were PP (sample codes: 1.3, 1.4, 4.1, 4.2, and 4.4), four were PS (sample codes: 2.1, 2.2, 2.3, and 2.4), four were PET (sample codes: 3.1, 3.2, 3.4, and 4.3), two were HDPE (sample codes: 1.2, and 3.3), and one was an unidentified plastic (sample code: 1.1) (Fig. S3).

Isolation of microorganisms using culture-dependent methods

Fifty-four and 62 isolates of bacteria and fungi were successfully isolated from plastic and sand samples on MA supplemented with cycloheximide and PDA supplemented with nalidixic acid, respectively. The bacterial colonies ranged from 2.5 × 10³–5.10 × 10⁷ CFU/g, while fungal colonies were <2.5 × 10² CFU/g.

After checking viability during cultivation, this study selected only 27 bacterial and 38 fungal isolates, based on distinct colony characteristic and the source of isolation (PP, PS, PET, and sand samples). These bacterial isolates were categorized into four groups as follows: PP plastic (eight isolates), PS plastic (six isolates), PET plastic (eight isolates), and sand (five isolates). In case of fungal isolates, they were categorized as follows: PP plastic (ten isolates), PS plastic (nine isolates), PET plastic (eleven isolates), and sand (eight isolates). The strain codes assigned to the bacteria and fungi in each isolate are shown in Table 1.

The 16S and ITS regions were successfully molecularly classified. Thirteen bacterial genera were identified through 16S rRNA analysis with EzBiocloud: Marinobacter, Pseudoalteromonas, Salipiger, Planococcus, Psychrobacter, Shewanella, Exiguobacterium, Marinomonas, Cobetia, Jonesia, Bacillus, Kocuria, and Halomonas. The isolated fungi were classified into nine genera based on their ITS region: Aspergillus, Penicillium, Pestalotiopsis, Peacilomyces, Cochliobolus, Absidia, Fusarium, Diplodia, and Petromyces (Tables S3 and S4). The phylogenetic trees of the isolated bacteria and fungi are presented in Figs. 2A and 2B.

Microorganism degradation on plastic-containing agar plates

One bacterium and 15 fungi exhibited clear zones on MSM agar containing emulsified PLA, indicating their ability to degrade this plastic. In addition, 14 bacteria and 19 fungi demonstrated the ability to degrade BHET on MSM-emulsified plastic agar. However, none of the microorganisms demonstrated degradation of PS or PVC on MSM-emulsified plastic agar. The identification of bacteria and fungi through 16S rRNA and ITS region revealed the capability of seven bacterial genera (Salipiger, Planococcus, Psychrobacter, Shewanella, Jonesia, Bacillus, and Kocuria) and five fungal genera (Aspergillus, Penicillium, Peacilomyces, Absidia, and Cochliobolus) to degrade plastics. The results of clear zone are shown in Figs. 3A, 3B and 4. Additionally, the HC values of bacteria range from 1 to 1.64. Notably, Planococcus sp. PP5 obtained from a PP sample had the highest HC value of 1.64 ± 0.12, followed by Salipiger and Kocuria with HC values of 1.57 ± 0.18 and 1.55 ± 0.15, respectively (Table S3).

Microbiome analysis

The 16S and ITS amplicon sequencing analysis in plastic and sand samples revealed the Shannon indices in Fig. 5, assessing the alpha diversity of the bacterial (Fig. 5A) and fungal (Fig. 5B) communities. Alpha diversity serves as a measure of the microbial diversity of each sample. The results indicated that the sand samples exhibited the highest alpha diversity for both bacteria and fungi, followed by PP, PS, and PET samples for bacteria and PET, PP, and PS samples for fungi. Additionally, the statistical analysis of Shannon indices for different groups was evaluated using the Kruskal-Wallis test, revealing significant differences in microbial communities among sample groups (p value < 0.05).

Beta diversity assesses the variations in microbial diversity differences between samples. The principal coordinate analysis (PCoA) plot, as shown in Fig. 6, is generated using the matrix of pairwise distances between samples calculated by Bray-Curtis dissimilarity. It reveals distinctions in the communities of both bacteria and fungi between sand and plastic samples. Meanwhile, the microbial communities within plastic samples (PP, PS, and PET) exhibit similarities. The differences in beta-diversity were statistically assessed through permutational multivariate analysis of variance (PERMANOVA), as shown significant differences (p-value < 0.05) among sample groups.

The microbial community composition in Fig. 7 reveals that Proteobacteria were most abundant in PET samples (59.9%) and more commonly found in plastic samples than in sand samples, similar to Bacteroidetes. Cyanobacteria (36.3%) were more abundant in PS samples than in other samples. On the other hand, Actinobacteria and Acidobacteria exhibited higher abundance in sand samples (15.9% and 16.9%, respectively) than in plastic samples. Regarding fungal communities, Ascomycota (11.6%) and Basidiomycota (2.8%) were the most prevalent and abundant groups in PET samples. However, a substantial portion (95.9%) of all samples could not be classified into specific fungal groups.

The fungal microbiome analysis conducted using the Zymo Research database revealed that 95.9% of the isolates could not be identified. Therefore, further analysis was performed using BLAST with the GenBank database. Blast results were obtained from nine samples (sample names: PP1, PP3, PP4, PS1, PS2, PS3, PS4, PET3, and PET4), while the other seven samples (sample names: Sand1, Sand2, Sand3, Sand4, PP2, PET1, and PET2) did not match any sequences in the database with the current search parameters. From the BLAST results shown in Fig. S4, the dominant group was identified as other eukaryotic groups (61.5%), including ciliates, diatoms, hydrozoans, crustaceans, cercozoans, anemones, tunicates, gastropods, bivalves, chytrids, sponges, green algae, and uncultured eukaryotes, while classified fungi were present in a smaller proportion (18.4%). The fungal class Dothideomycetes (9.23%) was the most abundant, followed by Sordariomycetes (3.08%) and Agaricomycetes (2.46%).

Linear discriminant analysis effect size (LeFSe) analysis helps to identify taxa whose distributions among different sample groups are statistically with p-value < 0.05 and the effect size (LDA score) higher than 2. The bacterial at genus level identified through LDA score, represent the most dominant microbial groups in each sample, revealing 15, 21, 15, and 22 bacterial genera in the PP, PS, PET, and sand samples, respectively (Fig. 8A). Similarly, the fungal at genus level identified through microbial composition bar plot, with results showing 7, 3, 26, and 19 fungal genera in the PP, PS, PET, and sand samples, respectively (Fig. 8B). On the other hand, bacterial groups isolated under culture-dependent conditions represented 5, 4, 6, and 5 genera in the PP, PS, PET, and sand samples, respectively (Fig. 9A), while isolated fungal groups represented 5, 6, 4, and 3 genera in the PP, PS, PET, and sand samples, respectively (Fig. 9B). Moreover, the 16S microbiome analysis of plastic and sand samples revealed the dominant bacterial genera that were absent in the culture-dependent conditions, such as Prochlorococcus in the PP samples, Ruegeria in the PS samples, Erythrobacter in the PET samples, and Rhodopirellula in the sand samples. The ITS microbiome indicated that Cladosporium and Malassezia were the most common fungal genera, while the fungi groups under culture-dependent conditions were mostly Aspergillus and Penicillium. Moreover, six bacterial genera (Marinobacter, Pseudoalteromonas, Marinomonas, Jonesia, Bacillus, and Halomonas) and two fungal genera (Aspergillus and Penicillium) were present in both isolation and microbiome analyses. Furthermore, comparing the microbial communities between plastic samples and sand samples, differences were observed in the bacterial communities of each sample. However, in the fungal group, some samples exhibited fungi from the same group, specifically Cladosporium, found in every sample. Aspergillus was identified in PP, PET, and sand samples; Malassezia was found in PS, PET, and sand samples; Irpex was present in PP and PET samples, and Penicillium, Aureobasidium, Hortaea, Kalmusia, and Curvularia were identified in PET and sand samples.