Living in a bottle: Bacteria from sediment‐associated Mediterranean waste and potential growth on polyethylene terephthalate

Abstract Ocean pollution is a worldwide environmental challenge that could be partially tackled through microbial applications. To shed light on the diversity and applications of the bacterial communities that inhabit the sediments trapped in artificial containers, we analyzed residues (polyethylene terephthalate [PET] bottles and aluminum cans) collected from the Mediterranean Sea by scanning electron microscopy and next generation sequencing. Moreover, we set a collection of culturable bacteria from the plastisphere that were screened for their ability to use PET as a carbon source. Our results reveal that Proteobacteria are the predominant phylum in all the samples and that Rhodobacteraceae, Woeseia, Actinomarinales, or Vibrio are also abundant in these residues. Moreover, we identified marine isolates with enhanced growth in the presence of PET: Aquimarina intermedia, Citricoccus spp., and Micrococcus spp. Our results suggest that the marine environment is a source of biotechnologically promising bacterial isolates that may use PET or PET additives as carbon sources.

Plastic residues in landfills are exposed to wind and water flows, which transport them into rivers and streams and, ultimately, into the oceans (Lebreton et al., 2017). Moreover, other direct sources such as beach littering, aquaculture, or fishing are also responsible for the accumulation of plastic in marine environments (GESAMP, 2016).
Due to the generally low temperature and limited UV exposure in marine conditions, plastic degradation is considered to take longer in the sea (Gewert et al., 2015;Napper & Thompson, 2019). Plastic waste tends to fragment and spread in small particles (<5 mm) commonly known as microplastics (Arthur et al., 2009), which are easily ingested by marine wildlife, entering this way the trophic chain, and finally being ingested by humans (Setälä et al., 2014). Several studies have revealed the presence of plastic particles in fish, crustaceans, and mollusks (Neves et al., 2015;Van Cauwenberghe et al., 2015;Watts et al., 2014), and even in dietary salt (Iñiguez et al., 2017). This may have an impact on human health because of its physical accumulation as well as the toxicity of the additives used in plastic industries and the organic pollutants that plastic can adsorb in the marine environment (Bouwmeester et al., 2015;Rochman et al., 2013;Teuten et al., 2009). Moreover, not only the entrance of these microplastics on the trophic chain but also the enrichment of potentially pathogenic multidrug-resistant bacterial strains in the plastisphere is a major health problem to face (Wang et al., 2021).
However, the amount of plastic estimated to enter into marine ecosystems does not correlate with the accumulation found by sampling techniques (Eriksen et al., 2014;Jambeck et al., 2015). Although there could be biases in sampling specific areas, this fact could also indicate that either physical or chemical plastic degradation is taking place in these ecosystems and/or microbial biodegradation is involved (Auta et al., 2017;Gewert et al., 2015;Sole et al., 2017;Zrimec et al., 2021). In recent years, plastic debris has proved a niche for specific plastic-associated microbial communities to flourish, generally known as the "plastisphere" (Agostini et al., 2021;Zettler et al., 2013). Microbial growth on the plastisphere usually takes place in the shape of a biofilm on the plastic surface (Lobelle & Cunliffe, 2011). Although meta-analyses are suggesting that a significant enrichment of potentially plastic biodegrading microorganisms in the plastisphere is detected (Wright, Langille, et al., 2021), there are still contradictory reports on the specificity of the composition of the microbial plastisphere. Specifically, some studies have shown that nonbiodegradable plastics, such as polyethylene terephthalate (PET), are colonized by a general biofilm rather than plasticspecific species (Oberbeckmann et al., 2016;Pinnell & Turner, 2019). Therefore, microbial biofilms attached to plastic surfaces in the marine environment seem to be composed of complex communities where some microorganisms, although not being the primary producers, may have evolved or adapted to degrade plastic polymers or plasticizers (Pinnell & Turner, 2019).
In the last decades, there has been a rapid rise in the use of PET to produce disposable packaging, such as single-use plastic bottles.
This has led to a dramatic increase in PET waste generation, which is now one of the most common plastics polluting marine environments (PlasticsEurope, 2020;Ritchie & Roser, 2018). PET is a polymer made from raw petroleum-derived monomers, terephthalic acid, and ethylene glycol. Its high content in aromatic compounds makes it chemically inert and subsequently very robust against biodegradation (Sinha et al., 2010).
In this context, bioprospecting microbial species able to in situ biodegrade plastic has arisen as a potentially useful tool for tackling the plastic contamination problem in the oceans (Danso et al., 2018). The first bacterium that demonstrated an effective PET-degrading activity due to the expression of a lipase (PETase) was Ideonella sakaiensis, isolated from the sediments of a plasticrecycling industry, which can hydrolyze this polymeric compound (Yoshida et al., 2016). However, these enzymes capable of PET hydrolysis have also been detected in other bacterial and fungal isolates, such as Thermobifida fusca, Streptomyces spp. or Fusarium solani, among others (Carr et al., 2020), and have been mainly described as cutinases, lipases, and esterases which are carboxylic ester hydrolases (Kawai et al., 2020).
Here, we show a complete characterization of the microbial communities associated with marine residues from the Mediterranean Western coast with a dual culture-dependent and -independent approach. We have studied the biofilm morphology on plastic and aluminum debris through scanning electron microscopy (SEM), characterized the microbial communities of their inner sediments by 16S and 18S ribosomal RNA (rRNA) genes sequencing, and established a microbial collection of mainly culturable bacteria and some yeasts, whose ability to grow on media supplemented with PET as sole carbon source has been characterized.

| Sampling
Plastic residues and cans were collected from the Malva-rosa beach (València, Spain; 39°27′48.3″N 0°19′07.6″W) in September 2017 ( Figure 1). The sampling was carried out at 20 m from the coastline and 3 m in depth. Four PET plastic bottles (labeled as P1-4) and four metallic beverage cans (labeled as M10-13) were collected and transported to the laboratory into sterile plastic bags. All the residues were originally submerged or half-buried in the marine sediments and they were thus partially filled with sand, mollusk shells, and marine plants (Posidonia oceanica) debris. Three samples of control seabed sediments (CS4-6) from the same area where plastic and aluminum residues were collected, which consisted of similar materials like sand, little stones, and shells, were also collected. Furthermore, some of the marine residues collected were still labeled with the expiration date of the product; therefore, an approximate age for these bottles or cans can be deduced: aluminum can M10 (expiration date 2003), aluminum can M12 and M13 (expiration date 2018), plastic bottle P1 (expiration date 2010).
Samples from the insides of each recipient (sediments) were collected under sterile conditions in the laboratory and stored at −20°C until required. To obtain samples from the plastic surface biofilms, recipients P1-4 were shortly rinsed with sterile water and then cut into small pieces which were shaken together with glass beads in phosphate-buffered saline (PBS; pH 7.4; in g/L: 8.0 NaCl, 0.2 KCl, 1.42 Na 2 HPO 4 , 1.80 KH 2 PO 4 ), at 500 rpm, for an hour. A total of 150 ml of the resulting suspension were collected and centrifuged at 4500 rpm for 15 min (sample P12) and stored at −20°C until required. Sample 12 was only analyzed in terms of culturable bacteria and it was not included in the high-throughput 16S rRNA gene sequencing.
Two replicates were incubated under aerobic conditions and the other two replicates in anaerobic conditions by placing the dishes inside a hermetic container without oxygen (N 2 atmosphere).
Individual colonies were picked according to morphological traits (color, shape, and size) and restreaked on fresh media until a pure culture was obtained. The strains were named after a code composed of a letter and a number associated with its origin (P1-4 and P12: plastic bottles; M10-13: aluminum cans; CS4-6: external sediments), followed by a unique number for each strain and a letter referring to the incubation conditions (X: aerobic conditions; A: anaerobic conditions). For example, P1.1X means the first colony isolated from bottle P1 that grew under aerobic conditions. The strains were stored in cryotubes with 20% glycerol at −80°C until used.
Amplicons were precipitated overnight in isopropanol 1:1 (v:v) and potassium acetate 3 M, pH 5, 1:10 (v:v) at −20°C. After centrifuging at 12,000 rpm for 10 min, DNA pellets were washed in 70% ethanol and resuspended in the required amount of sterile Milli-Q water.  (Karnovsky, 1965). The fixation solution was changed after five hours and samples were stored in this solution at 4°C until required. For SEM, the pieces were washed in phosphate buffer 0.1 M, pH 7.4 (PB, in g/L: 3.1 NaH 2 PO 4 ·H 2 O, 10.9 Na 2 HPO 4 ) to remove the fixative and progressively dehydrated in increasing ethanol concentrations. Samples were placed inside microporous specimen capsules (30 μm pore size) immersed in absolute ethanol, followed by critical point drying in an Autosamdri 814. The fragments were then arranged on SEM aluminum stubs using carbon tape and coated with Au/Pd sputtered in argon gas. The observation was carried out in a Scanning Electron Microscope Hitachi S-4800 at the electron microscopy service of the University of València (SCSIE).

| DNA purification and high-throughput 16S rRNA gene sequencing
Internal sediments from the marine residues collected were subjected to DNA extraction. In particular, 1 g of sediments of each sample (2 × 250 bp) was employed for sequencing the samples. All the library preparation and sequencing steps were carried out by Novogene.

| Bioinformatic analysis
Raw Illumina sequences were analyzed using Qiime2 (v. 2020.8) (Bolyen et al., 2019). Briefly, the quality of the reads was assessed with the Demux plugin, and the sequences were subsequently corrected, trimmed, and clustered into amplicon sequence variants (ASVs) via Dada2 (Callahan et al., 2016). The taxonomy of each sequence variant was assigned employing the classify-Sklearn module from the feature-classifier plugin (Bokulich et al., 2018). SILVA (v. 138) was used as a reference for the 16S rRNA gene assignment (Quast et al., 2013). The phyloseq R package (McMurdie & Holmes, 2013) was used for analyzing and visualizing the data. All the α-diversity tests were carried out using ASVs and rarefying to the lowest library size (128,327 sequences).

| Plastic degradation assay in solid medium
Plastic degradation was assessed through qualitative assays by comparing the growth of the bacterial strains on minimal marine medium (MMA), enriched marine medium (MME), and marine medium supplemented with plastic (MMP). MMA consisted of water from the Mediterranean Sea and 15 g/L agar, whereas MME consisted of seawater and, in g/L, 1.0 yeast extract, 5.0 bacteriological peptone, and 15 agar. MMP was prepared by using seawater, supplemented with 9.3 g/L of ground PET of approximately 0.5 mm in size, from a commercial PET water bottle (brand Cortes) and 15 g/L of agar, which was then sterilized at 121°C for 30 min. The PET bottle was ground in a coffee grinder for 5 min at maximum speed. As plastic particles tended to sediment on the bottom of the dishes, the media was stirred by using sterile spatulas before solidification.
Before the incubation with PET, bacterial isolates were grown on solid MMA for 4 days at room temperature. Cell suspensions with an Optical Density at 600 nm (OD 600 ) of 1 were prepared in PBS and 4 µl of the suspensions were placed on Petri dishes containing MMA, MME, and MMP (in duplicate). The dishes were incubated for 16 days at 18°C.
Isolates with a more vigorous growth (as determined by colony diameter and cell density) in MMP than in MMA were selected as potential plastic degrading bacteria and tested again in the same media conditions but using a 10-fold dilution of the bacterial suspensions (OD 600 of 0.1).

| Plastic degradation assay in liquid medium
Assay tubes were prepared with 3 ml of seawater and 0.400 ± 0.001 g of particles of PET from a new water bottle (brand Cortes), of 3 mm in size (cut by hand to obtain homogeneous size), and sterilized by autoclaving at 121°C for 30 min. Bacterial strains were grown on solid MA for 4 days at room temperature. Cell suspensions were prepared in PBS and adjusted to a final OD 600 of 0.05. The assay was carried out in duplicate by incubating the tubes at 18°C under shaking (200 rpm) for 3 months. Control tubes consisted of sterile seawater inoculated with the microbial cultures, as well as seawater and plastic particles but without inoculated bacteria.
At the end of the incubation period, PET fragments were rinsed with sterile water and vortexed for 2 min in distilled water. The process was repeated three times and the washed plastic particles were dried at 65°C for 48 h. Finally, the remaining plastic particles were weighted in a precision balance. To finally compare the colony-forming units (CFU) in each condition, the recovered supernatants of each tube were diluted in serial dilutions and 50 µl of each dilution was inoculated in duplicate into MA plates.

| Residue types and samples
Plastic PET bottles and aluminum cans were collected to study their associated microbiota as described in Section 2. The bacterial communities present in the inside-sediments, coming from PET bottles and aluminum cans, were compared with control, non-artificial residuesassociated sediments from the same area. Interestingly, some of the marine residues collected were still labeled with the expiration date of the product; therefore, an approximate age for these bottles or cans can be

| Scanning electron microscopy
The SEM images of the surface of plastic and aluminum marine waste suggest a diverse microbial community attached to these surfaces ( Figure 2). Different microbial morphologies could be differentiated in both cases, including rod-and coccus-shaped cells as well as diatoms and filamentous microorganisms. In particular, spermatozoid-shaped bacteria stood out in Figure 2c,e which may belong to prosthecate bacteria such as Hyphomonadaceae. Interestingly, several samples showed 2 µm fusiform bacilli firmly attached to the plastic surface, to which they were linked through polar fimbriae-like structures

| Taxonomy of the waste-associated bacterial communities
The bacterial community of marine waste was studied by highthroughput 16S rRNA gene sequencing yielding the composition of the taxa in the inside sediments of four PET bottles, inner sediments of four aluminum cans, as well as three samples of control marine sediments. The shape of rarefaction curves revealed that sequencing was deep enough to cover all the microbial diversity for all samples (1.5%), and uncultured Syntrophobacterales (1.4%). Samples CS4 and P2 showed a similar taxa composition to the other samples, but clear F I G U R E 3 Representations of the values of alpha diversity indices in the (a) observed richness at the amplicon sequent variant (ASV) level (number of ASVs), (b) Shannon index of diversity, and (c) Simpson index of diversity. The 11 analyzed samples are represented: insidesediments of cans (green); polyethylene terephthalate inside-sediments (purple); controlsediments of the sea-bed (blue) F I G U R E 4 Principal coordinates analysis (PCoA) based on Bray-Curtis dissimilarities at the genus level in bacterial populations of both inside-sediments of marine residues, plastic (blue), and aluminum cans (red). Sample P2 not included differences in abundance, where Vibrio and Sulfurovum were the dominant genera in each sample, respectively. A test for differential abundance (Table A1) revealed that the phylum Caldatribacteriota was significantly more abundant in plastic sediments than in aluminum sediments. At the same time, it showed that when comparing debris sediments to control sediments, Cyanobacteria and Marinimicrobia were more abundant in can sediments as well as Campilobacteria, Cloacimonadota, and Acetothermia were significantly more abundant in inner plastic sediments.

| Strain collection and identification
Culturing the marine sediments associated with artificial residues yielded a large number of highly diverse microbial colonies, in terms of color and morphology. A total of 170 bacterial strains and one yeast were isolated. All the strains that grew at first under anaerobic conditions showed later the ability to grow in the presence of oxygen. In total, 142 out of 171 strains were identified through colony PCR and 16S and 18S rRNA gene sequencing (Table A2), whereas 29 remained unidentified due to the impossibility to carry out the amplification of these fragments through PCR. The identified bacteria were distributed into four phyla: Firmicutes, Proteobacteria, Bacteroidota, and Actinobacteriota ( Figure 6). Bacillus spp. was by far the most abundant genus (33 species identified), followed by Vibrio spp.
(9), Erythrobacter spp. (8), Planomicrobium spp (7), Sulfitobacter spp. (6) and Sphingorhabdus spp. (5) among other genera. Interestingly, the identification of a large fraction of the microorganisms in the collection revealed that some isolates could represent new species, as they held a percentage of identity with the closest type strain below F I G U R E 5 Barplots showing the taxonomic profiles at the phylum (a), class (b), and genus (c) level of the top 20 most abundant groups in terms of relative abundance of inside-sediments from marine residues (plastic and aluminum cans) and control sediments by high-throughput 16S ribosomal RNA gene sequencing VIDAL-VERDÚ ET AL. | 7 of 23 the 98.7% threshold established to circumscribe a new bacterial species (Chun et al., 2018). In particular, isolates M10.2A, M10.9X, and P4.10X with the closest type strains belonging to the genera Gillisia, Sagittula, and Maritalea, respectively, are potentially new species. Further characterization is needed to determine it.

| PET degradation assays
To test the PET degrading activity of the microbial isolates obtained from marine waste, a preliminary qualitative screening was carried out consisting of a drop assay of bacterial culture in MMA and MMP to check differential growth when PET plastic was present (see Section 2.7). From this preliminary screening, differences in terms of growth after the drop assay performed as described in Section 2 are shown in Figure 7. In the first round of selection, 27 out of the 171 strains tested were selected as they showed increased growth in minimal medium supplemented with PET particles compared to the control medium without PET, after 28 days at 18°C. A second assay with the 27 selected strains was then carried out and led to the further selection of 16 strains with the more obvious differential growth on PET-containing media. 16S rRNA complete gene sequences were obtained and compared using EzBioCloud thus allowing the identification at the species level (Table A3) The group of 16 strains selected in the previous assay was incubated for 3 months at 18°C in liquid MMP containing PET particles precisely weighted. The following controls were included in the assay: PET without inoculated bacteria; the medium without neither bacteria nor PET; and each bacterium incubated without plastic. The test resulted in no detectable weight loss of the plastic particles in any sample inoculated with any of the 16 strains. Surprisingly, a small weight loss was detected in the noninoculated controls, in which the liquid became cloudy, appearing a white precipitate ( Figure A3). To discard microbial contamination of the controls, the commercial MA medium was inoculated with the cloudy supernatant, which was also observed under the microscope. Both experiments yielded negative results and contamination of the controls was thus discarded. There was one unit decrease in the pH of these control tubes (7.5 ± 0.1) compared with the tubes inoculated with a microorganism, all of which remained at a pH of 8.5 ± 0.3 and exhibited no turbidity in any inoculated tube.
F I G U R E 6 Bar plots showing the distribution into four phyla of the isolated species within the collection. The different colors in each phylum represent one different genus and the numbers indicate the number of isolates identified, which are only written when the number of isolates per genus is greater than two (see Table A2 for detailed information about each strain identified) Substantial differences in bacterial growth were found in the containing-PET and non-containing-PET medium in four of the strains, by comparing cell number (CFU) of the supernatants in-  (Table A3).

| DISCUSSION
Artificial residues hold great promise as a source of a huge variety of microorganisms for the bioremediation of plastic waste (Delacuvellerie et al., 2019;Yoshida et al., 2016). Regarding the bacterial communities inhabiting the marine sediments studied in this study, at the β-diversity level, the samples analyzed did not cluster together depending on the type of sediment (cans-inner sediments, plastic-inner sediments, and control-external sediments) (Figures 4 and A2). This suggests that the bacterial profile F I G U R E 7 Differential growth of eight selected strains on minimal marine medium (MMA), minimal marine medium supplemented with polyethylene terephthalate (PET) (MMP), and enriched marine medium (MME). MMA was used as a control for the basal growth of the strain without any supplemented carbon source. MMP was used to compare the growth of the isolates in the presence of PET plastic. MME allowed the normal growth of the strain in a rich nutrient marine medium VIDAL-VERDÚ ET AL. | 9 of 23 Interestingly, we found numerous fusiform bacteria attached to the plastic surface through fimbriae-like structures (Figure 2b). Similar shapes have previously been described to inhabit plastic surfaces in marine environments. For example, Bryant et al. (2016), showed a similar microbial community and also reported a bacillary shape that is attached from one pole to the plastic surface. In another study on the plastisphere of microplastics from the Australian shores, the same bacillary shapes with fimbriae-like structures adhering to the plastic surface were described . Furthermore, the wellknown PET degrading bacteria Ideonella sakaiensis exhibits attaching appendages when growing on plastic (Figure 2f Table A3 Desulfobulbaceae, all of which were found in the sediments analyzed in this study. Interestingly, the abundance of the genus Vibrio is remarkable in all the samples. Pathogenic bacterial species belonging to Vibrio have been widely described in marine environments usually in low abundance and they have also been found in plastic debris (Delacuvellerie et al., 2019;Jacquin et al., 2019;Zettler et al., 2013). Vibrio is very resistant to hard conditions and can perform a rapid growth in marine environments in response to an increase of nutrients (Westrich et al., 2018). Another interesting fact is that PET bottle P2 was dominated by Sulfurovum while this genus remained in low abundance in the other samples. Species from the genus Sulfurovum are chemolithoautotrophic sulfur-oxidizing bacteria that are primary producers in marine sediments communities (Mori et al., 2018) and even have been described to be the dominant taxon in seafloor sediments in some localizations (Sun et al., 2020).
The microbial composition we have found is similar to that reported in a variety of studies carried out on the biofilm that directly Erythrobacteraceae and Rhodobacteraceae, which in our collection are represented by the eight genera: two belonging to Erythrobacteraceae (Altererythrobacter, Erythrobacter) and six belonging to Rhodobacteraceae (Epibacterium, Maliponia, Ruegeria, Sagittula, Sulfitobacter, and Yoonia). Moreover, the eight representative genera of the phylum Bacteroidota belonged to the Flavobacteriaceae family, which is, again, a common plastic debris-associated taxa (Amaral-Zettler et al., 2020;Jacquin et al., 2019). The abundance of Firmicutes is linked to the high number of Bacillus spp, (33 species isolated in total) we found. This genus has been reported as a marine plastic colonizer and degrader (Delacuvellerie et al., 2019;Oberbeckmann et al., 2015;Ribitsch et al., 2011).
The diversity of microorganisms found on artificial debris, the presence of biofilms and plastic adhesion fimbriae-like structures, and the taxonomic identity of some of the taxa suggest a possible role in plastic biodegradation of some of the bacteria of the collection we set and characterized. The quantitative PET degradation assay with the selected strains yielded no significant loss of non-pretreated PET particles weight. However, this is not particularly surprising givien the fact that PET is very resistant to biodegradation due to its compact structure, hence heat or oxidative pretreatments are usually needed to enhance biodegradation (Gewert et al., 2015). Nevertheless, we observed an increased growth (measured as CFU count variation), of seven of the isolates when PET was present as the sole carbon source in the medium, suggesting the capability of some strains to degrade plastic or plastic additives, such as plasticizers, antioxidants, light and heat stabilizers, pigments or slip reagents that are usually added to plastics to enhance their structural properties. These compounds are commonly not covalently bonded to the plastic polymer; therefore, they can more easily leak out from the plastic structure to the liquid phase (Hahladakis et al., 2018). Remarkably, the strain of Micrococcus luteus we tested, showed a 20-fold increase in CFUs when the minimal medium was supplemented with PET particles compared to a non-supplemented-PET medium. This is not the first time that Micrococcus luteus has been described to potentially degrade plastic (Montazer et al., 2018;Sivasankari & Vinotha, 2014), and its degrading ability seems to be associated with its ability to form biofilm in plastic surfaces (Blakeman et al., 2019;Feng et al., 2011). The isolates identified as Idiomarina piscisalsi, Citricoccus alkalitolerans, Aquimarina intermedia, and Microbacterium aerolatum which showed roughly a two-to four-fold increase in growth in PET, have been sparsely studied in previous works regarding plastic-degrading activity. Specifically, Idiomarina has been recently reported to possibly assist in the formation of biofilms on the surface of PET particles, although it showed no significant PET degradation (Gao & Sun, 2021). On the contrary, although there is no previous report on the ability of Bacillus algicola (which showed double CFU count when incubated with PET) to degrade plastic polymers, other species and strains within the genus have been described as degraders of polystyrene, polypropylene, polyethylene, and PET microplastic particles (Auta et al., 2017;Wright, Bosch, et al., 2021) as well as polyvinyl chloride (Giacomucci et al., 2019).
Finally, the yeast Rhodotorula evergladensis, which showed a tiny increase in growth on PET in our study, has been previously reported to degrade plasticizers (Gartshore et al., 2003).
Taken together, our results suggest that the marine wasteassociated microbiota hold potential as a source of biotechnological interesting strains for plastic or plastic-related compounds. On the left, the two replicates of the negative control consisting of marine water with PET fragments. A white precipitate of mineral nature appeared after the incubation time, probably due to the change in pH. The two tubes on the right contained only marine water. (b) A representative example of the assay with the isolate M11.3X. All the tubes were inoculated with the bacterium at the beginning of the assay in duplicate, on marine water supplemented with PET particles (left) and marine water without plastic as control (right) T A B L E A2 List of the strains identified in the collection, with the closest type strain, accession number, ID percentage, and the GenBank accession number for the 16S or 18S rRNA gene sequences obtained in this study Note: The identification code of the strains corresponds to the sediments from which it was isolated (CS, control sediments; M, can inside-sediments; P, plastic inside-sediments; and a number).
T A B L E A3 List of selected isolates that showed enhanced growth in PET-containing medium, with the closest type strain, GenBank accession number for the 16S and 18 rRNA gene sequences, and results obtained in the quantitative assay *Tessaracoccus rhinocerotis yielded an uncountable number of colonies; therefore, its differential growth was not measured.