Marine fungi degrade plastic and can be conditioned to do it faster

Abstract

Plastics are a prevalent and persistent pollutant in the environment. As plastic production increases, finding ways to degrade these recalcitrant polymers is paramount. Many terrestrial fungi, across the kingdom, degrade various types of plastic. Plastics are the fastest-growing habitat in the oceans, and we hypothesized that fungi isolated from the ocean would demonstrate high success rates in degrading polyurethane (PU). To test this, visual degradation assays were performed by inoculating 1% PU-medium with 68 different fungal strains cultured from marine habitats. The area of clearance of the fungus was measured periodically, to determine a relative degradation rate. Of the 68 fungal strains, 42 demonstrated the ability to degrade PU. We conditioned the nine fastest PU degraders through serial inoculations into liquid media with increasing concentrations of PU, starting at 1% and going up to 12%. The growth rates of the original and conditioned fungi were then compared in new inoculation trials, and results show that three of the nine conditioned fungi demonstrate higher PU degradation rates than their unconditioned counterparts. Marine fungi, coupled with conditioning, show promise for developing novel mycoremediation technologies.

Introduction

Plastics are among the most prevalent pollutants in the ocean (Gunawan et al. 2022; NOAA 2021; Zeghal et al. 2021). New models suggest that 500 kilotons of plastic enter the ocean annually (Kaandorp et al. 2023) and that number is expected to continue rising exponentially in the foreseeable future (Gunawan et al. 2022). Of all the waste in the oceans, 60–80%, by mass, is plastic (Sangale et al. 2019). Plastics are cheap, strong and useful for consumers, but plastic waste is problematic because rather than decomposing, it is very long-lived, breaking into microplastics (5 mm in size or less) under environmental stressors such as UV radiation, heat, and physical stress (Hammer et al. 2012; NOAA 2021; Urbanek et al. 2018; Yogalakshmi and Singh 2020). For this reason, microplastics resulting from fragmentation of larger plastics exposed to the environment have become the most abundant marine debris (Roman et al. 2021).

Plastics are also detrimental to marine ecosystems (Gunawan et al. 2022; Thompson 2017; Yogalakshmi and Singh 2020). Toxins resulting from harmful additives used during production, can be released into aquatic environments creating detrimental by-products (Yogalakshmi and Singh 2020). Furthermore, plastics can adsorb and concentrate dangerous chemicals in the environment, such as phthalates and bisphenol A, which can disrupt the hormones of organisms and lead to cascading negative ecosystem effects (Hammer et al. 2012; Mohanan et al. 2020; Urbanek et al. 2018; Yogalakshmi and Singh 2020). Plastics can also cause physical harm or be ingested and lead to the starvation of marine animals due to malnutrition (Yogalakshmi and Singh 2020). In fact, entanglement and ingestion are the two most common ways in which plastics harm marine organisms (Hammer et al. 2012; Thompson 2017). Over 1,400 marine species are known to ingest marine debris, and flexible plastics are one of the most lethal types (Roman et al. 2021). Conservationists and managers need a method to effectively dispose of plastics and remediate areas already impacted by plastic pollution.

To this end, various microbes have been tested for their ability to degrade plastics in the hopes that an aspirational biotechnology can, one day, be deployed at ecologically relevant scales (Yogalakshmi and Singh 2020). Results from such studies are promising, and fungi and bacteria, in particular, show great potential as agents of plastic remediation (Brunner et al. 2018; Kavitha and Bhuvaneswari 2021; Matavuly and Molitoris 2009; Mohanan et al. 2020; Russell et al. 2011; Sangale et al. 2019; Urbanek et al. 2018; Yamada-Onodera et al. 2001; Yogalakshmi and Singh 2020).

Because the vast majority of fungal research focuses on terrestrial taxa, these constitute the majority of isolates tested for their ability to degrade polyurethane-type plastics (Zeghal et al. 2021; Russell et al. 2011). Marine fungi promise a novel, and promising target for research into plastic degrading microorganisms (Zenghal et al. 2021). Chemists estimate that around 71% of marine natural products are not expressed by organisms in terrestrial environments (Kamat et al., 2023). Marine fungi, in particular, have proven a rich source of novel chemistry and enzymes (Bonugli-Santos et al. 2015; Carroll et al. 2024). As an example, a pioneering study demonstrated the polyethylene degradation ability of a marine yeast, Rhodotorula mucilaginosa, using  13C labelled PE substrate (Vaksmaa et al. 2023). Further, marine fungi are proximate to continental-scale masses of plastic “garbage patches,” a new and large carbon source which they might exploit. With fewer than 1% of marine fungi currently identified (Gladfelter et al. 2019), marine fungi offer a promising and largely untapped group to study.

Plastics vary in their chemical structures, and in this study, we focused on polyurethane (PU). PU comprises 6–7% of global plastic production and is widely used for various purposes (Biffinger et al. 2015; Schulzke et al. 2018; Simón et al. 2018). PU is produced in the form of a flexible foam, and is also common in medical and industrial products, such as insulation in buildings. It is also found in other products such as adhesives, foams, tires, and sponges (Biffinger et al. 2015; Hammer et al. 2012; Mohanan et al. 2020; Schulzke et al. 2018; Simón et al. 2018; Zafar and Sharmin 2012). Structurally, PU has both carbon and non-carbon atoms including six-membered ring intermediates (Fig. S1; Mohanan et al. 2020; Schulzke et al. 2018; Simón et al. 2018). While some advances have been made in identifying the enzymes involved in degrading various types of plastics, there is still a large gap in our knowledge. For this project, we assayed PU in the form of an aqueous dispersion called Impranil^®.

Plastic degradation by marine fungi could provide insights into their possible future roles within the ecosystem, in addition to offering hope in remediating the ocean of this pervasive type of pollution. We hypothesized that marine fungi might represent novel targets for PU degraders. Further, we set out to test whether fungi can be experimentally conditioned to decompose plastic faster.

Methods

Marine Fungal Isolations

Fungi were isolated from a wide array of marine samples collected from coastal waters around Oʻahu, including sessile invertebrates, sediments, and water (see Table S1 for details). Malt Extract Agar (MEA) was prepared by combining 10g +/− 0.05 g of agar, 2.5 g malt extract, 2.5 g yeast extract, and 500 mL water, autoclaving the mixture, adding 25 mg of kanamycin and 25 mg of ampicillin, and pouring the mixture to cool into 100 mm Petri dishes (VWR, Radnor, Pennsylvania, USA). Colonies were serially transferred onto new plates, and single, clean cultures were isolated on MEA plates. Duplicate plugs of these colonies were stored submerged in ultrapure H₂0 at 4°C and in 10% glycerol solutions at −80°C.

Isolates were identified by sequencing the ITS1, 5.8S, and ITS2 regions of the nuclear ribosomal DNA using the Sanger method with ITS1F and ITS4 primers (White et al. 1990). The resulting sequences were classified using the naïve Bayesian classifier (Wang et al. 2007) with reference to the UNITE database (version 10.5.21, Nilsson et al. 2018). A subset of 68 isolates were chosen for this study to represent a phylogenetic diverse subset within Basidiomycota and Ascomycota, and to include replicates from genera previously noted to have degradative abilities (Zenghal et al. 2021; Table 1). In cases where sequences diverge from species present in the database, the classifier identifies the lowest (most granular) taxonomic rank in which probability of a correct classification is achieved according to default settings. Many, perhaps the majority, of fungi encountered in marine environments are not obligately marine but might also be encountered in terrestrial or freshwater habitats (Amend et al. 2019; El Baidouri et al. 2021).

Table 1.

Degradative abilities of all fungal isolates tested.

Genus	Species	ID	Degradation Visible by Day 96	Days to Full Clearancea
Acremonium	brachypenium	635	N
Acremonium	citrinum	702	Y	96
Acremonium	vitellinum	364	N
Allophoma	tropica	867	Y	96
Angustimassarina	sp.	609	N
Aspergillus	caespitosus	749	Y
Aspergillus	ficuum	354	N
Aspergillus	hongonkensis	372	Y	62
Aspergillus	sclerotioniger	209	N
Aspergillus	sp.	349	N
Aspergillus	sp.	378	Y
Aspergillus	sp.	396	Y	96
Aspergillus	sp.	501	N
Aspergillus	sp.	503	Y	96
Aspergillus	sp.	512	Y
Aspergillus	sp.	530	Y	96
Aspergillus	sp.	614	Y	96
Aspergillus	terreus	456	Y
Aspergillus	varians	179	Y	62
Aspergillus	varians	214	N
Aspergillus	westerdijkiae	321	Y
Aspergillus	westerdijkiae	887	Y
Aureobasidium	sp.	51	Y	96
Candida	orthopsilosis	605	N
Chaetomium	sp.	408	Y
Cladosporium	dominicanum	195	Y
Cladosporium	halotolerans	268	N
Cladosporium	sp.	460	Y
Fusarium	sp.	135	Y	62
Fusarium	sp.	256	N
Fusarium	sp.	328	Y	62
Fusarium	sp.	834	Y	62
Fusicolla	sp.	384	Y
Gibberella	intricans	330	Y	19
Graphium	jumulu	201	N
Hortaea	werneckii	490	Y
Musicillium	theobromae	737	Y
Neocosmospora	falciformis	241	N
Neocosmospora	falciformis	361	N
Novel (Family Nectriaceae)	sp.	496b	Y	62
Penicillium	brocae	118	N
Penicillium	coccotrypicola	59	Y
Penicillium	commune	46	N
Penicillium	hetheringtonii	244	N
Penicillium	sp.	84	N
Penicillium	sp.	223	Y
Penicillium	sp.	276	N
Penicillium	sp.	424	Y
Penicillium	sp.	483	Y
Penicillium	sp.	528	Y
Penicillium	sp.	674	Y
Penicillium	sp.	707	Y
Penicillium	steckii	204	Y
Penicillium	steckii	381	Y
Penicillium	sumatraense	740	N
Purpureocillium	lilacinum	100	Y	62
Rhodosporidiobolus	sp.	274	N
Rhodotorula	dairenensis	448	Y
Rhodotorula	mucilaginosa	106	N
Rousoella	sp.	590	N
Roussoella	sp.	21	N
Sarocladium	bifurcatum	334	Y
Sarocladium	kiliense	67	Y
Selenophoma	linicola	318	Y
Sporothrix	ramosissima	127	N
Stilbocrea	walteri	229	N
Trichoderma	lixii	986	N
Volutella	sp.	336	N

^aFor the fungi that were able to clear the entire plate, the upper number of days of the time range within which the full plate was cleared (0–19 days; 20–62 days; 63–96 days).

^bNovel species 496 has been identified down to the family Nectriaceae.

PU Degradation Assays

To prepare the media for the PU degradation assays, 3 g/L of NH₄ NO₃, 5 g/L of K₂ HPO₄, 1 g/L of NaCl, 0.2 g/L of MgSO₄ 7H₂O, and 15 g/L of agar were combined with sterile water. The mixture was autoclaved, allowed to cool to 55°C, and then 10 mL/L of Impranil^®DLN (Covestro), 60 mg/L of streptomycin, 60 mg/L of kanamycin, and 100 mg/L of chloramphenicol were added. The medium was then poured into 100 mm Petri dishes (VWR).

Once solidified, each plate was inoculated with two plugs of each of the 68 isolates. Each plug was considered an observational pseudoreplicate (so that there were duplicates of each isolate but grown on a single plate), and incubated in the dark at room temperature (20–23°C) for 96 days. Clearance was measured every 2–3 days for the first month, and then at longer intervals. Initially, percent clearance was estimated by measuring the average distance between the two furthest points in the zone of clearance. However, as the fungi grew, their growth became less uniform, making the measurements less reliable, so percent clearance was visually estimated instead. Because PU refracts light, absence of PU can be measured as the area where agar is no longer opaque in the medium (Fig. 1). Diameter of clearance is a standard method to determine breakdown of PU and other hydrocarbons (Russell et al. 2011; Biffinger et al. 2015; Bollinger et al. 2020; Molitor et al. 2020; Kim et al. 2022; Khruengsai et al. 2022; Park et al. 2023).

Figure 1.

Example of the zone of clearance in a PU assay. Area of clearance can be seen around the fungal plug and is indicated by the medium turning clear and/or filled with hyphal growth. Pictured is the growth of an unidentified Volutella species isolated from a sponge in the genus Spheciospongia.

Experimental Conditioning of PU Degraders

To experimentally condition the fungal isolates, liquid medium was prepared as indicated for the PU plates, excluding the addition of agar, and adding 10 g/L of NaCl to simulate seawater conditions more closely. We note that growth of fungi isolated from marine habitats are generally unaffected by salt concentrations in lab conditions (Jones & Jennings, 1964), but suspected that addition of NaCl might help survivorship in increasingly extreme conditions. From the nine isolates with the fastest growth rate on solid PU medium, a circular plug was removed from an area of active growth on solid media and transferred to 10 mL of 1% PU liquid medium in 15mL Falcon tubes. Tubes were inspected for clearance of visible PU particles. Once the tubes appeared completely cleared, serial transfers were made to 1.5 mL tubes with PU concentrations increasing to 5%, 10% and finally 12% PU liquid medium. Fungi were eliminated from the experiment if no signs of clearance were observed at the higher PU concentrations.

To determine whether conditioning impacted fungal clearance rates on PU media, we grew hyphae retained from the 1% concentration medium alongside the mycelium conditioned on 12% concentration medium and compared rates of clearance. We compared the growth in three separate formats: 1) in 1 mL of liquid medium, 2) on 60 mm Petri dishes (each isolate on its own plate), and 3) with both the original and conditioned strain growing together on one large 100 mm Petri dish. Inoculants were observed for 12 days.

Results

PU Degradation Assays

In the PU degradation assays, 42 of the 68 fungal isolates (62%) produced detectable levels of PU degradation within 96 days (Table 1; Table S2). Of 26 genera examined, 17 contained isolates with demonstrated plastic degradation ability (Fig. S2). Colony morphologies varied markedly among the isolates, and some prolific sporulators produced plates that were dotted with small areas of growth/degradation (Fig. S3). The fastest degrader was isolate 330, Gibberella intricans; the two replicates/plugs cleared the entire plate in 19 days (Table 1). Five other fungi cleared the entire plate in 42–62 days, and another six fungi were able to clear the entire plate between 62–96 days (Table 1). The other thirty fungal isolates demonstrated degradative abilities but were considerably slower and unable to clear the entire plate within 96 days. Overall, the rate at which the plates were cleared varied widely across the isolates (Fig. 2).

Figure 2.

The percentage of clearance on a 100 mm Petri dish containing 1% PU medium by two pseudoreplicates (fungal plugs) per plate of selected fungi. The isolate numbers for each fungus are indicated in parentheses in the legend.

There was some (putatively fungal) contamination that was observed on eight of the plates. The earliest sign of contamination was at day 35 on the plate of isolate 51 (Aureobasidium sp.) plate. Three other plates showed signs of contamination at day 62, and the other four plates showed signs of contamination between days 63 and 96 (Supplements Table S2).

Experimental Conditioning in Liquid PU Medium

Results of the experimental conditioning were mixed. Two of the species, Sarocladium kiliense (isolate 67) and Gibberella intricans (isolate 330), stopped growing at the highest concentration (12%) of PU. Three of the conditioned fungi showed faster rates of degradation than their unconditioned counterparts during the 12-day long growth comparisons; Aureobasidium sp. (isolate 51) cleared 19% more of the plate, Penicillium coccotrypicola cleared 20% more, and Penicillium steckii cleared 11% more of the plate. Of those, the conditioned Aureobasidium sp. (isolate 51) had fastest conditioned degradation rate, clearing 95% of a 60 mm Petri dish in 12 days, compared to the 80% degraded by the unconditioned isolate (Table 2). The other four conditioned fungi did not demonstrate faster growth (based on percent clearance of the plate) compared to their unconditioned counterparts (Table 2; Supplements Fig. S4).

Table 2.

Comparison of the growth of conditioned and unconditioned fungi on 1% PU plates.

Fungus	Isolate Number	T1a Percent Plate Clearance at Day 12	T4b Percent Plate Clearance at Day 12	T4 Conditioned Speed Compared to T1
Aureobasidium sp.	51	80	95	Faster
Fusarium sp.	135	100	100	Same
Fusarium sp.	328	100	100	Same
Gibberella intricans	330	100	0	Died
Novel sp.; Nectriaceae	496	100	100	Same
Penicillium coccotrypicola	59	50	60	Faster
Penicillium steckii	204	90	100	Faster
Sarocladium kiliense	67	15	0	Died
Selenophoma linicola	318	95	95	Same

^aTransfer 1 (T1) refers to the original fungal isolate grown in 1% liquid PU medium.

^bTranfser 4 (T4) refers to the conditioned fungi that were transferred from 1%, to 5%, to 10%, and finally to 12% liquid PU medium.

Discussion

Our results support those of other plastic degradation studies and offer new insights into the potential utility of marine fungi for plastic degradation. We note that a remarkably high proportion of our isolates (62%, 42 of 68 marine fungal isolates) exhibited some ability to degrade PU over 96 days. Previous research, which focused on terrestrial fungi exclusively, showed significantly lower proportions of strains with degradation ability. For example, in one study of fungi isolated off of PU biofilms found in the soil, a lower proportion of isolates (10–30%) were able to degrade PU (Barratt et al. 2003). We note that our selection of isolates was not random, and we purposely included a greater number of isolates from genera that are known to degrade plastics and other polymers. These selections support previous results, showing that terrestrial isolates of Aspergillus spp., Fusarium spp., and Penicillium spp. (Magnin et al. 2018; Srikanth et al. 2022) can degrade PU.

Results of the experimental conditioning demonstrated that in some, though not all cases, conditioning via serial transfer resulted in faster degradation rates. The conditioned Aureobasidium sp. (isolate 51) degraded 15% more of a plate in 12 days compared to the unconditioned counterpart (Table 2). Metabolic adaptation is the most parsimonious explanation for the change in degradation rate resulting from the conditioning experiment. In future studies, we might compare the genomes of conditioned and original isolates to look for selective mutations in gene involved in PU degradation.

In the PU degradation assays, there was some contamination observed on eight of the plates (Supplements Table S2). However, in all cases, the contamination was only observed on day 62 or later. This delayed appearance might be an example of the priming effect, which is generally associated with an increase in organic matter microbial mineralization rates (Bengtsson et al. 2018). Here, the occurrence of contamination at such a late stage may suggest that the target inoculant was necessary in the initial metabolism of the available substrate, after which bi-products of the degradation were used as energy sources by the contaminant. Experiments focusing on coculture and degradation bi-products may help answer some of the existing questions about the dynamics observed here.

There is a high burden of proof to demonstrate that fungi directly degrade or metabolize plastic, and additional lines of evidence might lend mechanistic insights into specific enzymes and pathways that marine fungi are using to this end. Having identified candidate strains in this study, we hope that future experiments might identify genes that are upregulated in the presence of plastic substrates, compared with other carbon sources, in order to investigate and confirm key degradation genes, and their products. Another line of research might be to quantify production of known plastic degrading enzymes. Laccases and peroxidases, among others, are known plastic degraders (Zenghal et al. 2021), and measuring their activity in degradation assays might further refine our insights into known and novel fungal degradation mechanisms.

Extracellular enzymatic activity of fungi offer promise for bio-engineering solutions to emergent pollution problems. Marine fungi are a largely untapped source of novel chemistry and metabolic potential. Enhancing this innate capacity within the framework of metabolic adaptation framework shows promise to an emerging and difficult problem of high priority. Fungi may be the key to solving the problem of plastic pollution in oceans and waterways.