Ability of fungi isolated from plastic debris floating in the shoreline of a lake to degrade plastics

Plastic waste in the environment is a significant threat due to its resistance to biological processes. Here we report the ability of fungal strains found on floating plastic debris to degrade plastics. In particular, we wanted to know which fungi grow on plastic debris floating in the shoreline, whether these fungi have the ability to degrade plastics, whether the plastic-degrading fungi can degrade other complex C-polymers such as lignin, and whether lignin-degraders vice versa can also break down plastics. Overall, more than a hundred fungal strains were isolated from plastic debris of the shoreline of Lake Zurich, Switzerland, and grouped morphologically. Representative strains of these groups were then selected and genetically identified, altogether twelve different fungal species and one species of Oomycota. The list of fungi included commonly occurring saprotrophic fungi but also some plant pathogens. These fungal strains were then used to test the ability to degrade polyethylene and polyurethane. The tests showed that none of the strains were able to degrade polyethylene. However, four strains were able to degrade polyurethane, the three litter-saprotrophic fungi Cladosporium cladosporioides, Xepiculopsis graminea, and Penicillium griseofulvum and the plant pathogen Leptosphaeria sp. A series of additional fungi with an origin other than from plastic debris were tested as well. Here, only the two litter-saprotrophic fungi Agaricus bisporus and Marasmius oreades showed the capability to degrade polyurethane. In contrast, wood-saprotrophic fungi and ectomycorrhizal fungi were unable to degrade polyurethane. Overall, it seems that in majority only a few litter-saprotrophic fungi, which possess a wide variety of enzymes, have the ability to degrade polyurethane. None of the fungi tested was able to degrade polyethylene.


Introduction
Plastic debris in the environment poses a significant threat because of its resistivity to photooxidative, thermal, mechanical and biological processes [1,2]. Although overlooked for many years, the amount of plastic debris accumulating in the environment has been steadily increasing as a result of the material's durability and lightweight nature [3,4]. Once discarded on land, PLOS  plastic debris makes its way to water bodies that act as sinks for low-density litter [5][6][7][8].
Topography, wind and water currents, and proximity to pollution sources control the amount and types of plastics along shorelines, whereas degradation processes determine how long plastic debris remains on beaches [9,10]. An estimated 300 million tons of plastic are produced yearly [11]. Plastics are human-made materials manufactured from polymers or long chains of repeating molecules. They are derived from oil, natural gas, and, increasingly, from plants like corn and sugarcane. About four percent of the world's petroleum is used to make plastic, and another four percent is used to power plastic manufacturing processes [12]. Polyethylene (PE) represent about a third of total plastic production, with PE is largely utilized in packaging [11].
Plastic debris, an inevitable consequence of living the 'Plastic Age', is dominating our lakes and oceans and poses a worldwide threat to aquatic wildlife [3,13]. Floating or drifting plastic creates environmental hazards including the risks of plastic ingestion, starvation, and entanglement of aquatic organisms [5,9]. Plastic debris, as recently published from the Lake Geneva, consisted of various size and colour, including bottles, bottle tops, cotton buds, pens, toys, straws, and pieces or blocks of expanded polystyrene or polyurethane foam [14]. Plastic debris also provides novel aquatic vehicles for a wide range of rafting species, such as bacteria, fungi, algae, or insects, posing a potential threat to introduce invasive species [13]. Once plastics are discharged into aquatic environments, they can persist for up to 50 years, and their complete mineralisation may take hundreds or thousands of years [15].
In 2011 US researchers discovered an endophytic fungal species, which was able to degrade polyurethane (PU), a plastic which is widely used in the manufacture of e.g. high-resilience foam seating, rigid foam insulation panels, or tires such as skateboard wheels [16]. This discovery obtained a high attention in the media (e.g. http://www.dailymail.co.uk/sciencetech/ article-2146224/Could-fungi-break-plastic-stop-modern-scourge.html). The question arises, what about the fungi which can be found on plastic debris? We had the hypothesis that at least some fungi that grow on plastic debris have the potential to degrade plastics, and that the fungi that can degrade plastics are more generalists than specialists. In particular, we wanted to answer the following questions: (1) Which fungi grow on plastic debris floating in the shoreline? (2) Do the fungi isolated from floating plastics have the ability to degrade PE or PU? (3) Do fungi which are able to degrade plastics also have the ability to degrade other complex Cpolymers such as lignin? (4) Do fungal lignin-degraders vice versa have the capability to degrade PE or PU? With this study we also had the intention to clarify the abilities of the various trophic modes of fungi (saprotrophs, pathotrophs, symbiotrophs) with its functional guilds (e.g. plant pathogens, wood saprotrophs, [17]) for the degradation of PE or PU.

Sampling of plastic debris
Plastic debris was collected in the shoreline of Lake Zurich close to Wädenswil (UTM coordinates 32T 474250 5231960) at September 2 nd 2015. The plastic pieces either floated on the water or were found in a depth of up to 20 cm in the reed belt. The pieces of plastic were picked up with a pair of tweezers and a 0.7 cm x 0.7 cm piece was cut out with a pair of scissor. That piece was then placed into a sterile 50 ml Falcon tube. Before use, tweezers and scissors were dipped into 70% ethanol and flamed over a lighter for sterilization. The falcon tubes were kept sealed in a refrigerated bag and transported the same day to the lab where they were kept at 4˚C until use. In total, 13 pieces of plastics were sampled out of the water of the lake, and one was found close to the water (No. 16, Table 1). One piece was a hard-plastic chunk and only a 2.5 cm 2 fragment could be turned off with the tweezers.

Isolation of fungi
In the laboratory, 2 ml of sterile water was added to each Falcon tube containing one plastic debris piece. Then the tubes were mixed using a vortex mixer for about 10 s to allow the fungal hyphae and spores to separate from the plastic samples. In the sterile bench, 100 μl of water from each Falcon tube was taken with a sterile pipette, released into a Petri dish containing modified Melin-Norkrans (MMN) nutrient agar, and spread with a flamed glass rod on the surface of the agar (compare also [18]). Per Falcon tube, four Petri dishes were incubated. The plates were then incubated at room temperature in the dark until after a few days the first fungal colonies were visible. Emerging fungal colonies were then punched out with a flamed hook, transferred onto a malt agar in glass tubes ('test-tubes') and incubated at room temperature in the dark.

Identification of fungi
Once the fungal mycelia in the glass tubes covered about half of the agar surface, they were transferred to 4˚C to stop growth. In order to select fungi for DNA identification, the isolated fungal strains were morphologically grouped according to their external appearance in terms of colour and texture. Representative strains of these groups were selected, and a part of the nuclear small subunit rDNA was sequenced. Samples of the fungal mycelia were directly placed into the wells of 96-well PCR-plates containing 100 μl DNAse/RNAse free PCR-grade water per well. Then the fungal hyphae were frozen by submerging the plates into liquid N 2 and thawed at room temperature at least three times in order to break up the cells and to release the DNA. This solution was then 1:10 diluted in PCR-grade water and used as template for the PCR reaction, performed with the G2 Hot Start Polymerase (Promega AG, Dübendorf, Switzerland), MgCl 2 , dNTP, BSA and the primer pair ITS3/ITS4 [19] similar as in [20]. The resulting PCR products were then sequenced by a company (GATC Biotech, Köln, Germany), and the obtained nucleotide sequences blasted using the National Centre of Biotechnology Information (NCBI) database to obtain the closest species match.
In order to obtain longer DNA fragments for a more precise identification, the fungi, which were able to degrade PE or PU, were sequenced again (Macrogen Europe Amsterdam, the Netherlands) with the primer pair ITS1/ITS4 [19]. These nucleotide sequences were deposited at the NCBI GenBank.
The ability of the fungi to degrade a complex C-polymer other than plastic, e.g. lignin, was tested with the 'Bavendamm' assay in Petri dishes on agar medium [23]. This assay uses polyphenols as a lignin substitution. The agar medium contained 20 g L -1 malt extract and 15 g L -1 agar, and as a polyphenol, 0.5 g L -1 tannic acid (TA) was added to the solution before autoclaving [23].
Into each of the Petri dishes, three inoculi per fungal strain were placed on the media (compare also Fig 1). Then, the dishes were sealed with plastic paraffin film and incubated at room temperature in the dark. The Petri dishes were visually inspected every few days.

Optical evaluation of the degradation
The media in Petri dishes containing PE or PU were both milky and not transparent. The PE polymers, however, floated on the top of the medium during agar solidification, whereas PU polymers remained homogeneously distributed within the medium after agar solidification. According to Russell et al. [16] it was expected that fungi capable of degrading the plastic polymers would display a zone of clearance ('halo') around the growing cultures as a result of enzymatic plastic degradation by diffusing enzymes excreted by the fungal hyphae, or in the case of PE, grow on the plastic granules [21]. The media with the TA, however, was expected to change the colour from light brown to dark brown as a result of an enzymatic oxidative reaction of the TA by diffusing enzymes excreted by the fungal hyphae [23].
In order to have a control strain for the degradation of PU, Pestalotiopsis microspora was purchased from the Westerdijk Fungal Biodiversity Institute (CBS No. 364.54; CBS-KNAW, Utrecht, The Netherlands). P. microspora is able to degrade PU [16].

Fungal strains isolated from plastic debris
Fungal strains usually grew within a few days after dispersing the water from the Falcon tubes on the Petri dishes. In total, more than one hundred fungal strains were isolated. According to their external appearance, the fungal strains were grouped into morphological groups. From these groups, one or two fungal strains per group were selected, in total 24 fungal strains, and a part of the nuclear small subunit rDNA was sequenced. After blasting the sequences with the NCBI database, the names of the closest species match were listed. In several cases, identical names appeared. The final list of organisms isolated and sequenced from plastic debris contained twelve different fungal species belonging to the Ascomycota and one species to the Oomycota (Pythium) ( Table 2). The fungal names were checked and approved using the 'Index Fungorum' (http://www.indexfungorum.org). A good identification of fungal names is given when the nucleotide identity was equal or above 97% [25]. If the identity was below 97%, then the names have to be taken with caution, and they might not be correct ( Table 2).

Ability of the fungal strains from plastic debris to degrade PE and PU
In total, twelve fungus species and one species of Oomycota were tested to degrade PE or PU ( Table 2). After at least three weeks of growth, neither signs of 'halos' were visible around the inoculi nor growth of the inoculi was recorded in the PE degradation assay. I contrast, 'halos' were visible in the PU degradation assay after at least three weeks of growth around the inoculi of four fungal species Cladosporium cladosporioides, Xepiculopsis graminea, Penicillium griseofulvum, and Leptosphaeria sp. (Table 2, Fig 1). The most efficient fungi for PU degradation was C. cladosporioides with an approximate growth of the halo of 4 mm/d (Fig 1). To ensure the species names, some of these taxa were sequenced again with the primer pairs ITS1 and ITS4 to obtain longer sequences, which then were deposited at the NCBI database under the accession numbers MF327241-MF327243 (see also Table 2).

Ability of fungal strains from various fungal guilds to degrade PU and TA
Overall, none of the tested fungi was able to degrade PE (data not shown). However, three fungal species were able to degrade PU: Agaricus bisporus, Marasmius oreades, and Pestalotiopsis microspora (Table 3). Surprisingly, none of the highly specialised lignin-decomposing fungi such as the saprotrophic white-rot fungi or the plant pathogens were able to degrade PU. Similarly, the ectomycorrhizal fungi as well as the saprotrophic brown-rot fungi were not able to degrade PU. From the common saprotrophs, who all were able to degrade TA, only the two species A. bisporus and M. oreades were able to degrade additionally PU (Table 3).
Of the four fungal species isolated from the plastic debris and able to degrade PU, Xepiculopsis graminea was the only species that was able to degrade TA. The endophytic P. microspora, our PU-degradation reference strain [16], was not able to degrade TA (Table 3).
Besides the PU-degrading fungi reported in the present study, 15 ascomycete fungi are reported to potentially degrade PU ( Table 4). The best-known fungi are members of the genera Aspergillus, Penicillium, and Trichoderma. Two ascomycete and two basidiomycete fungi from this study are newly reported to be able to degrade PU.

Discussion
The list of organisms, which have been isolated from plastic debris, included commonly occurring saprotrophic fungi but also some plant pathogens. Commonly occurring saprotrophic fungi were Penicillium griseofulvum and Cladosporium cladosporioides [26,27]. Some fungi are known to live as saprotrophs in soils and sediments such as Xepiculopsis graminea and Phialemoniopsis curvata [28,29]. Some fungi are known to live in association with grasses or with plants growing in the littoral zones of lakes, e.g. Arthrinium arundinis, Leptosphaeria sp. and Phoma sp. [30,31]. Some fungal species are highly specialised to the common reed (Phragmites australis) such as the endophytic fungus Stagonospora neglecta [32]. Botryotinia fuckelinana is known as a necrotrophic fungus that affects many plant species [33]. The fungal species, which only had a low identity (Exophiala bonariae, Pseudorobillarda texana, Setophoma vernoniae), were isolated by others either from rocks or from leaves of exotic plants [28,34]. The only organism not belonging to the fungi was the oomyceteous Pythium phragmitis which is a pathogen for the common reed (Phragmites australis) [35].
The four fungal species isolated from plastic debris showed a 'halo' in the PU assay: C. cladosporioides, P. griseofulvum, X. graminea, and Leptosphaeria sp. Cladosporium cladosporioides had been observed already by others to be able to degrade PU. Á lvarez-Barragán et al. [36]  found that the six best PU-degrading strains using an Impranil assay belonged to the C. cladosporioides complex, with identities between 99% and 100%. Further BLAST analysis of the actin and translation elongation factor from these six strains showed the highest matches with the C. pseudocladosporioides, C. tenuissimum, C. asperulatum, and C. montecillanum [36]. Some reports, in contrast to our study, stated that C. cladosporioides is able to degrade PE as well (e.g. [37,38]). However, their results based not on the formation of a 'halo' in a Petri dish after PE degradation, but on observing erosion of the PE film surface in the vicinity of the fungal hyphae as well as formation of oxidation products in the surface of the polymer film measured by FTIR (Fourier-transform infrared spectroscopy). Penicillium ochrochloron, a different species than our isolated P. griseofulvum, had been observed already by other authors to have the capability to degrade PU [39]. These authors applied similarly as described above the PU assay using Impranil for soil fungi which they isolated from soil-buried PU pieces. For Xepiculopsis graminea and Leptosphaeria sp., in contrast, no references were found in the literature. Thus, this is the first report on the ability of these two fungi to degrade PU. There is evidence from the literature that microorganisms capable of degrading complex C polymers such as lignin can also degrade plastics [40]. Such degradation potential is based on lignin-degrading enzymes, e.g. oxidases, laccases and peroxidases, which are used in various industries and which are also reported to be involved in the degradation of xenobiotic compounds and dyes [41]. Overall, the three saprotrophic fungi Agaricus bisporus, Marasmius oreades and Xepiculopsis graminea remain the only fungi in our study which were able to degrade PU as well as TA. At least A. bisporus is known to possess a wide variety of enzymes including enzymes involved in xylan, cellulose, pectin, and protein degradation, as well as heme-thiolate peroxidases and β-etherases, which are distinctive from other wood-decayers and suggest a broad attack on decaying lignin and related metabolites found in humic acidrich environment [42]. The catabolic ability of A. bisporus agrees with the presence of a large set of genes encoding CAZymes [43] acting on cell wall polysaccharides including glycoside hydrolases, polysaccharide lyases, and carbohydrate esterases [42]. Carbohydrate esterases are suited in A. bisporus to break down the cell wall polysaccharides xylan, chitin, and pectin [44].
Marasmius oreades is known to produce fairy rings in grasslands. Fairy rings are characterised by two or three adjacent concentric zones of abnormal turf. Within the zone of most intense fungal growth, the grass is often killed, and this effect has been attributed to a lack of moisture and to hydrocyanic acid produced by the fungus [45,46]. The occurrence of fairy rings in natural vegetation has simultaneous contrasting effects of both stimulation and a parasitisation of plant species in adjacent zones, producing concentric regular bands of lush and scorched vegetation [47]. In soils colonized by M. oreades, degradation of plant roots in the presence of fungal cell-wall degrading enzymes increased the content of dissolved organic carbon [48]. Interestingly, similar as M. oreades, members of the Agaricus genus form fairy ring as well, e.g. A. arvensis [49]. Thus, it can be assumed, that members of both, Agaricus and Marasmius, possess similar enzymatic capabilities to break down complex carbohydrate polymers.
Besides the PU-degrading fungi reported in the present study, a series of other ascomycete fungi are reported to degrade PU ( Table 4). The best-known fungi are members of the genera Aspergillus and Trichoderma, all of which are known to be used in biotechnological processes [54]. Members of Aspergillus are used to produce the enzymes amylases, glucoamylases, glucose oxidase, invertase, pectinase, and proteinases, whereas members of Trichoderma are used to produce cellulase [55].
Although we have in the present study not investigated ourselves the enzymes produced by the PU-degrading fungi, there are several studies which report that enzymes involved in PU degradation are most likely esterases and hydrolases. Alvarez-Barragan et al. [36] postulated that Cladosporium cladosporioides complex were the best PU degraders among the fungi tested, whereas Aspergillus fumigatus and Penicillium chrysogenum were the least degrading strains. Besides Impranil, the fungal isolates of Cladosporium spp. degraded PU foam as well. FTIR spectroscopy and GC-MS analysis showed that ester and urethane groups were attacked through the activity of fungal enzymes. During PU degradation, considerable activities of esterases were detected, but only low urease and no protease activities [36]. Loredo-Treviño et al. [56], isolating 32 fungal strains from sand samples contaminated with PU, reported 22 strains being able to grow using PU as nutrient source. Among the genera found were Aspergillus, Trichoderma, Penicillium, and Fusarium. Almost all of the PU-degrading fungi showed urease activity, whereas esterase, protease, and laccase activities were present only in a lower amount of the fungi. For the PU-degrader species Pestalotiopsis microspora, Russell et al. [16] suggested a serine hydrolase-like enzyme being responsible for PU degradation.

Conclusions
The majority of fungi isolated from plastic debris in the shoreline of a lake in Switzerland do not seem to be able to degrade the plastic they grew on. None of the fungi was able to degrade PE, whereas at least a few fungi isolated had the ability to degrade PU. Three of these fungi were saprotrophs, and one was a plant pathogen. Thus, we could only partially confirm the previously formulated hypothesis that at least some of the fungus can degrade plastic, but only PU and not especially PE. The search for additional fungal species isolated from other substrates than from plastics or from fruiting bodies revealed that they were in general as well unable to degrade PE. Only two saprotrophic fungi, Agaricus bisporus and Marasmius oreades, were able to degrade PU.
It seems that the biological degradation of PE still remains a challenge. Although there have recently been several review articles that fungi can degrade PE (e.g. [1,40,[57][58][59][60][61][62]), these reports are no more than vague hints. Otherwise, the plastic waste would not be transported vertically across oceans and landscapes, mechanically fragmented, and eventually accumulated as micro-or nanoplastic in the sediments and environment if fungi and other microorganisms were efficient in degradation. Although the majority of plastic debris that has entered the ocean since 1950 has settled to depths below the ocean surface layer, it is estimated that 0.3 million tons of plastic are floating on the ocean surface, of which an estimated 14% is microplastic (0.335-5 mm) and 2.5% is nanoplastic (<0.335 mm) [63]. These small plastic fragments in particular are problematic, because they enter into the food webs and accumulate potentially in animals [64][65][66][67][68].
Knowing the ecological guild would facilitate the search for potential fungi which are able to degrade plastics. Not long ago a Japanese group found in a similar way a bacterium that degrades poly(ethylene terephthalate) PET [69]. If such microorganisms could be found, their spores or their plastic degrading enzymes could be incorporated into the plastic material during manufacturing and, when the plastic waste would come into contact with lake-or seawater, the fungi would start to grow and to degrade the plastic.