Marine fungi are generally understudied, which is also reflected by the relatively limited number of studies targeting fungi on marine plastic debris (MPD) (Jacquin et al., 2019). Oberbeckmann et al. (2016) conducted a seasonal comparison of microbial communities including both bacteria and fungi on submerged PET bottles, glass slides, and seawater in the North Sea. Fungal communities were represented by Ascomycota, Basidiomycota and Chytridiomycota. The study showed that the PET-attached eukaryotic communities (fungi among them) varied significantly with season and location. Kettner et al. (2017, 2019) conducted exposure experiments with PE and PS in the Baltic Sea and the river Warnow and investigated colonization of these plastic surfaces by fungi. The fungal genus Chytridium, as well as fungi-like Rhinosporideacae, Rhizidiomyces, and Pythium taxa, had a high read count on both plastic types at both locations. However, a substantial number of sequences was assigned as unclassified fungi. The fungal community composition was significantly influenced by location but not polymer type. Furthermore, the alpha diversity was significantly lower for PE and PS compared to the surrounding water and wood particles. De Tender et al. (2017) conducted a 44-week experiment during which PE plastic sheets and dolly ropes were weighed down close to the sediment at a harbor and an offshore location in the North Sea. Similar to the Baltic Sea studies by Kettner et al. (2017, 2019), they found that the Ascomycota were highly abundant followed by a smaller fraction of Basidiomycota and Mucoromycota. Furthermore, a minor fraction of members of Ascomycota from the Lecanoromycetes class (Physconia, Candelariella, Caloplaca) were identified. Analyses on the beta diversity of the fungal community composition showed statistically significant effects of sample type (natural substrate vs. plastic polymers), environment, and exposure time. Some of the detected species were previously identified as potential PE degraders in terrestrial environments: Cladosporium cladosporioides (Bonhomme et al., 2003) and Fusarium redolens (Albertsson, 1980; Albertsson and Karlsson, 1990).

Recently, Lacerda et al. (2020) studied the fungal diversity associated with plastics in the surface waters of the Western South Atlantic and the Antarctic Peninsula by investigating three different molecular marker genes for fungal identification: ITS2 (Internal transcribed spacer), the variable regions V4 and V9 of the 18S rRNA gene sequence. To date, this is the only study aiming to resolve fungal diversity on plastic using multiple marker genes. Across the tested marker genes, a total of 64 different fungal orders were associated with plastics. The primers targeting the 18S rRNA gene (V4 and V9 regions) were able to detect a higher number of Chytridiomycota. Some taxa that were totally omitted by the ITS2 marker were detected, such as Rozellomycota, Zoopagomycota, Aphelidomycota, and Blastocladiomycota. Across all samples, the genus Aspergillus was the most abundant. Some of its OTUs were assigned at the species level, identifying A. vitricola, A. restricus, and A. wentii. None of the identified strains were previously reported as plastic degraders, although other Aspergillus species have been shown to be able to oxidize different plastic types (Table 2). Lacerda and colleagues also reported on fungal taxa, such as Aphelidomycota, Zoopagomycota, Mucoromycota, and Blastocladiomycota, that had previously not been detected to colonize plastic in the marine environment.

In contrast to environmental studies, Kirstein et al. (2018) studied the microbial composition formed on microplastics under controlled laboratory conditions. Their set-up consisted of a flow-through system with North Sea water in which HDPE (High Density Polyethylene), LDPE (Low Density Polyethylene), PP, PS and PET were incubated in the dark. The 18S rRNA gene sequencing analysis of the eukaryotic community of biofilms revealed that the highest fungal read abundances belonged to Chytridiomycota (up to 3% of sequences on PET). This can be explained by this taxa’s dominance throughout aquatic environments (Comeau et al., 2016) or the compatibility of biofilm presence on PET and chytrid’s parasitic lifestyle.

Biodegradation is the degradation of compounds and substrates mediated by living organisms, most commonly microorganisms. The soluble products of biodegradation (typically low molecular weight compounds) are absorbed or assimilated by the microorganisms. The biodegradation can be partial or complete. Complete biodegradation results in the formation of CO₂ and is also referred to as biomineralization. On the other hand, degradation of organic matter without a terminal electron acceptor, conditions that are countered in some reduced environments, leads to the formation of CH₄ and/or other short-chain hydrocarbons. In natural environments, biodegradation is mediated by enzymes or by other compounds (such as acids and peroxides), secreted by microorganisms.

Fungi are able to degrade synthetic compounds; e.g., persistent organic pollutants (POPs) (Singleton, 2001), polycyclic aromatic hydrocarbons (PAHs) (Cerniglia and Sutherland, 2001), benzene, toluene, ethylbenzene and xylenes (BTEX compounds) (Buswell, 2001) and pesticides (Pinto et al., 2012). The metabolic versatility of fungi and ability to degrade complex compounds indicates that biodegradation of plastics in the environment could be a potential metabolic trait of some fungi (Vaksmaa et al., 2021a). To date, some plastic degrading fungi have indeed been identified mostly from the Ascomycete phylum to which also Aspergillus, Fusarium, and Penicillium belong (Table 2 Fusarium sp. and F. oxysporum isolated from soil provoked weight loss of Nylon, PE, and PU) (Tachibana et al., 2010; Raghavendra et al., 2016). F. solani from a collection hydrolyzed PET to terephthalic acid (TPA) via a cutinase (Ronkvist et al., 2009). Furthermore, several strains belonging to Penicillium are considered as potential plastic degraders. P. chrysogenum, P. oxalicum, P. simplicissimum isolated from soil (Yamada-Onodera et al., 2001; Sowmya et al., 2012; Ojha et al., 2017) and Penicillium sp. isolated from seawater (Alshehrei, 2017) showed potential to degrade PE. Similar to Penicillium, different species belonging to the genus Aspergillus were found to be potential plastic degraders as well (Table 2). A. flavus isolated from soil (Deepika and Madhuri, 2015), wax moth gut (Zhang et al., 2020) and a marine environment (Sangeetha Devi et al., 2015; Alshehrei, 2017) exhibited the potential to degrade PE and A. niger isolated from soil (Deepika and Madhuri, 2015; Raghavendra et al., 2016) and seawater (Alshehrei, 2017) showed potential to degrade PE and PU. Also, A. terreus isolated from soil (Zahra et al., 2010), mangrove sediments (Ameen et al., 2015; Sangale et al., 2019) and seawater (Alshehrei, 2017) were potentially able to degrade PE. In addition to these species, several other types of Aspergillus strains were considered as potential PE and PU degraders. Many of the plastic degrading species that were isolated from terrestrial environments (Table 2) are also found in marine habitats. However, it has not been confirmed if all of the plastic degrading strains found in terrestrial environments perform equally well in the marine realm.

The enzymes utilized by plastic degrading fungi in the environment are typically not constrained. However, fungi produce a wide range of enzymes that have the potential to break down the chemical bonds of the plastic polymers (Figure 1). Amongst these are manganese peroxidase (MnP) and lignin peroxidase (LiP), which are commonly associated with lignin degradation (Xu et al., 2013). These enzymes catalyze oxidation-reduction reactions, involving free radicals, transforming several compounds into oxidized or polymerized products (Wei and Zimmermann, 2017). Peroxidase are also used in industrial applications for degrading recalcitrant organic pollutants, PAHs, industrial dyes and chlorophenols (Qin et al., 2014). Lignin peroxidase is characterized by a high redox potential, and enables oxidation of non-phenolic aromatic compounds. Another enzyme that might be involved in plastic degradation is laccase, a multicopper oxidase, which is a well classified lignin-modifying enzyme (Mehra et al., 2018) and mediates the oxidation of the polymers’ carbon backbone (Amobonye et al., 2021).

Elevated laccase, manganese peroxidase and lignin peroxidase activities were observed during PE degradation by a fungal consortium in a mangrove (Ameen et al., 2015). It appears likely that these enzymes also play a role in potential plastic degradation in the ocean. Marine adapted fungi have the ability to regulate the expression of their enzymes according to salinity. Cultivated in marine conditions, Peniophora sp., for example, showed multigene transcription of ligninolytic laccase enzymes (Otero et al., 2017). Fungi isolated from marine environments can also produce enzymes that allow them to grow in liquid media with sole carbon sources such as agar, alginate, carrageenans, laminarians, and ulvans, i.e., polymers which are common in the marine realm (Wang et al., 2016). However, whether these compounds are also utilized by marine fungi in situ, i.e., in the ocean, still needs to be shown. Similarly to several biopolymers in the terrestrial realm, these marine polymers also feature chemical similarities with some plastic types (presence of aromatic compounds, presence of carbonyl groups). The ability of marine fungi to regulate their metabolism and to adapt to different environments and substrates indicates that the number of marine fungi with the potential to degrade plastics could be underestimated.

To date, the most commonly applied technique to assess plastic biodegradation is based on gravimetric measurement of weight/mass loss of the polymer over a specific time period during which the plastic is exposed to an environmental matrix or to cultured microbes. Weight loss of plastic has been demonstrated both in studies where plastics were exposed to soils, the marine water column and sediments (Syranidou et al., 2017; Welden and Cowie, 2017) as well as in studies focusing on the capability of a single strain to degrade specific plastic polymers (Table 2). However, in environmental studies, it remains unclear to which extent the observed plastic breakdown was due to solely fungal activity. In the environment, the observed breakdown may be facilitated by microorganisms other than fungi and/or by microbial consortia, and/or by abiotic factors, such as mechanical degradation. These factors remain inseparable from potential fungal biodegradation. In contrast, laboratory studies with single strains decrease the number of biotic and abiotic factors, that may contribute to the plastic degradation. Fungal isolates recovered from municipal solid waste revealed that Fusarium oxysporum, Aspergillus fumigatus, Lasiodiplodia crassispora, Aspergillus niger, Penicillium sp., and Trichoderma harzianum were all able to cause a weight loss of PE as well as PU (Raghavendra et al., 2016). A. niger showed the highest biodegradation efficiency leading to a weight loss of 2.9, 4.3, and 5.1% of LDPE sheets and 0.8, 1.5, and 2.2% of PU sheets exposed for 30, 60 and 90 days, respectively. Fusarium oxysporum as well as Aspergillus sp. have previously been shown to cause weight loss of LDPE (Das and Kumar, 2014). Trichoderma viride and Aspergillus nomius, isolated from landfill soil, caused weight loss of LDPE films of 5.1 and 6.6%, respectively, after 45 days (Munir et al., 2018). Cladosporium tenuissimum caused 25.9 and 65.3% of weight loss of PE-PU foams with and without flame retardants, respectively (Álvarez-Barragán et al., 2016). The marine fungus Zalerion maritimum exposed to PE microplastics caused mass loss of 56.7 ± 2.9% of the plastic, corresponding to 43% of removal after 2 weeks of exposure (Paço et al., 2017).

Although determining the weight loss of plastic is a straightforward method, it requires a long monitoring time, ranging from months to years to yield measurable gravimetric changes, while short term studies often yield inconclusive results (Lee et al., 1991). Gravimetric measurements require post incubation treatments, i.e., the removal of the biofilm from the incubated plastic. This can easily cause measurement artifacts; e.g., the incomplete removal of biofilm or accidental removal of a polymer respectively leads to an under or overestimation of weight loss. Weight loss measurements indicate that plastic polymers indeed disintegrate over time, however, this type of measurement does not reveal whether the polymer is broken down physiochemically (e.g., plastics release lower molecular weight compounds as a result of photooxidation; Wayman and Niemann, 2021) or if the plastic polymer was hydrolyzed and metabolized by microbes. Some plastics (e.g., polyvinyl chloride) contain high quantities of additives which, if soluble, can be released, and thus bias weight loss measurements. Also, biomass figures (i.e., cell numbers, culture dry weight) are typically not reported for culture-based studies, making these investigations quantitatively non-repeatable, thus, severely limiting comparability. Hence, gravimetric measurements are insufficient to identify plastic degrading organisms and microbial kinetics, but should be accompanied by other methods.

Determining microbial biomass growth on plastic as the sole carbon source (e.g., monitoring cell numbers) has been used to infer the biodegradability of a specific polymer. Together with monitoring the weight loss of the polymer, some studies have measured an increase in fungal biomass as an indicator for the activity of fungi (Paço et al., 2017), and interpreted that the increase in fungal biomass occurred at the expense of carbon originating from the plastics.

The growth of fungi at the expense of plastic can be monitored visually with degradation assays based on plates coated with agar and solubilized plastic. Inoculation at the surface and utilization of the plastic leads to the formation of clearance zones, referred to as ‘halos.’ Appearance of ‘halos’ on plates with plastics as the sole carbon source is used as an indicator for plastic degradation. The fungal species Cladosporium cladosporioides, Xepiculopsis graminea, and Penicillium griseofulvum and Leptosphaeria sp., isolated from plastic debris from the lake Zurich, formed clearance zones on PU. However, none of these strains were able to degrade polyethylene (Brunner et al., 2018). Attempts to use polyethylene and clearance zone formation have been successful for Aspergillus niger and Aspergillus flavus (Deepika and Madhuri, 2015).

Clearance zone tests allow for rapid visual observations. However, the technique has the drawback that it requires dissolving the polymer in an organic solvent that can be applied to the petri dish at relatively cold temperatures to avoid agar melting. Furthermore, remnants of the solvents (rather than the plastic) may act as a carbon source for the fungi, leading to false-positive results. In addition, bias associated with clearance zone formation is relatively large, as clearance zones are not uniform in size or shape. Finally, fungi with hyphae may grow through the plastic, thus gain access to the agar below, which can also lead to false-positive results. As agar also represents a carbon source, this test alone does not prove microbial plastic degradation.

Scanning electron microscopy (SEM) is used to create a surface image by directing a high-intensity electron beam at the surface and scanning over this surface. SEM allows high magnification, thus offers high resolution at the nanometer range. SEM-based observations are used to examine and evaluate the colonization of plastic films or particles by microorganisms and to simultaneously visualize cracks, pits and deformations on the plastic surface (Zettler et al., 2013; Vaksmaa et al., 2021a), which in return can indicate if the polymer is degraded. SEM has been applied in several studies to investigate fungi on plastics, for example, to visualize the growth of C. tenuissimum and C. pseudocladosporioides hyphae within PE-PU foams (Álvarez-Barragán et al., 2016). Furthermore, Paço et al. (2017) visualized the attachment of Z. maritimum on PE. With SEM, also surface roughness of single plastic fragments can be visualized. Floating marine plastics often feature signs of abrasion, cracking and ongoing fragmentation (Zettler et al., 2013; Vaksmaa et al., 2021b). Also, plastics exposed to the marine environment developed such signs during the incubation. For example, Welden and Cowie (2017) exposed PE, PP and Nylon to marine sediment for 12 months and found breaks and increased fraying on nylon ropes, surface scratching and roughening on PE filament rope, cracks, fissures and finer surface fibers scaling off from PP.

Scanning electron microscopy is a rapid technique, and allows to visualize surface attachment and morphological microstructures. Observations by SEM without chemical fixation can be achieved by applying FIB-SEM (Focused Ion Beam milling combined with Scanning Electron Microscopy). However, SEM does not allow phylogenetic identification of the microbes (unless the strain-specific morphological characteristics allow this) and the formation and attachment of biofilms is not necessarily an indication for biodegradation. Furthermore, though SEM is a valuable tool to visualize surface defects (e.g., cracks) of the plastic, it does not allow to scale in Z-direction (i.e., to measure the depth of cracks). Alternatively, this may be achieved by atomic force microscopy (AFM). For example, an increase in surface roughness and formation of cracks and grooves on PE films were investigated with AFM after exposure to two different Penicillium strains (Ojha et al., 2017).

Fourier-transform infrared spectroscopy (FTIR) is a technique used to obtain an infrared spectrum of absorption, emission, and photoconductivity of a material allowing to determine the chemical identity of most polymers. Furthermore, the FTIR spectrum enables detection and semi-quantification of changes of the original polymer configuration, for example, the introduction of carbonyl groups during polymer oxidation (Xu et al., 2019; Almond et al., 2020). The degree of carbonylation can be enumerated by determining the carbonyl index (calculated from the ratio between the integrated band absorbance of the carbonyl and that of the methylene peaks). Carbonyl indexes, as a measure of degradation, has been applied for a variety of polymers, such as PU (Filip, 1979; Álvarez-Barragán et al., 2016), PE (Paço et al., 2017), PS (Tian et al., 2017), and PP (Sheik et al., 2015). PE degradation has been evaluated by FTIR in co-cultures of bacteria and fungi, Lysinibacillus xylanilyticus and A. niger, isolated from soil (Esmaeili et al., 2013). For example, Penicillium variabile CCF3219 strain decreased the carbonyl peaks of pre-oxidized 14C-βPS after 16 weeks of incubation and ozonation pre-treatment enhanced subsequent biodegradation (Tian et al., 2017). An additional advantage of FTIR is that it can be used for small plastic particles (the diffraction limit in IR spectroscopy is ∼10–20 μm).

Fourier-transform infrared spectroscopy is a straightforward and reliable technique, however, it often yields non-quantitative results that are difficult to compare between studies. Also, the attachment of biofilm on the plastic surfaces affects the plastics optical properties in the IR range because proteinic and polysaccharide contents of the biomass will change the polymer’s IR spectrum (Bonhomme et al., 2003). Samples must hence be pretreated to remove biofilms, e.g., using hydrogen peroxide (Löder and Gerdts, 2015) or sodium dodecyl sulfate (Zhang et al., 2020). These chemicals have been described as the least aggressive. Nevertheless, chemical treatments can modify the molecular structure of plastic surfaces, which can introduce biases.

Isotopically labeled plastics can be traced into biodegradation products sensitively and quantitatively (Lanctôt et al., 2018; Taipale et al., 2019). The first studies involving isotopically labeled plastics were conducted with the radio isotope ¹⁴C. The fungus Fusarium redolens liberated ¹⁴C-CO₂, originating from pulverized ¹⁴C-labeled HDPE (Albertsson, 1978, 1980; Albertsson et al., 1978). ¹⁴C labeled ¹⁴C-CO₂ formation was also evaluated after photoirradiation of ¹⁴C-αPS and ¹⁴C-βPS, exposed to garden soil and activated sludge, showing higher ¹⁴C-CO₂ formation in treatments with photo oxidized polymer. The authors quantified degradation rates and showed that complete degradation of the ¹⁴C-αPS polymer in garden soil would require 20 to 80 years and in activated sludge from 11 to 24 years (Guillet et al., 1974). ¹⁴C labeled polystyrene was also used to test PS degradation capabilities of 17 different fungal species in a 14 days incubation experiment during which 0 to 0.24% of the ¹⁴C-PS was degraded (Kaplan et al., 1979). Because radioactivity can be measured and quantified extremely sensitively, radio isotope probing allows determining extremely low plastic degradation rates. Nevertheless, the use of radio-chemicals entails the necessity for specialized laboratory facilities and trained personnel, and the radio-labeled base materials for synthesizing polymers are extremely expensive or not available at all. This makes the use of radio isotopes mostly impractical. However, using plastics labeled with stable isotopes (e.g., containing a high degree of ¹³C or ²H) provides a good alternative. Similar to radio isotope probing, stable isotope probing (SIP) offers the advantages to allow tracing plastic derived matter into degradation products (Sander et al., 2019; Taipale et al., 2019), though the ubiquitous presence of ¹³C and ²H in most types of matter makes SIP assays less sensitive compared to approaches with radio isotopes. Nevertheless, no study using ¹³C or ²H labeled conventional plastics (PE, PP, PET, PS, PU, and nylon) for investigating interactions of marine fungi and plastics has been published yet. On the other hand, Zumstein et al. (2018) used ¹³C-PBAT (Polybutylene adipate terephthalate) to study its microbial degradation in soil.

Additional methods to evaluate plastic degradation include the following techniques: (i) Respiratory measurements of O₂ consumption or CO₂ production, which can provide quantitative information of degradation, but cannot discriminate between different respiration pathways. (ii) Tensile resistance alterations of plastic can be used as a measure of the strength/integrity of the plastics, which will decrease as a function of degradation (yet, also physiochemically induced degradation reduces tensile strength). (iii) Thermo Gravimetrical Analysis (TGA) characterizes the thermal stability of a polymer, which can potentially indicate its degradation (similar to tensile strength). (iv) Differential scanning calorimetric (DSC) analysis assesses the thermal properties of synthetic polymers, such as glass transition Temperature (T_g).

Lower T_g temperatures are often related to a decrease in the stability, indicating degradation (Lucas et al., 2008). (v) High-Temperature Gel Permeation Chromatography (HT-GPC) provides information on the molecular weight (Mn%) and molecular weight distribution of the polymer. A decrease in Mn% is evidence of chain cleavage that can be related to microbial degradation. Nevertheless, just as alterations of tensile resistance, glass transition Temperature, thermal stability and the molecular weight will also change in response to physicochemical processes Finally, none of the other methods described in this section is able to unambiguously prove the occurrence of complete microbial biodegradation, from initial depolymerization to mineralization and biomass assimilation.