Introduction

Ocean plastic pollution is an environmental problem of exponentially increasing magnitude [1,2,3,4,5,6]. However, uncertainty exists on the importance of individual pathways through which plastic is transported from land to the ocean [7,8,9,10,11], how much plastic remains in coastal areas [12], the open ocean surface [13, 14], the mid-water column [15,16,17], or sinks to the ocean floor [18, 19]. Current estimates of the total amount of plastic at the ocean surface only account for less than 1% of the estimated amount of all plastics that have ever been released into the sea [20, 21]. Thus, an unknown sink mechanism apparently removes detectable plastic debris from the ocean surface [2, 21]. This might be an abiotic process, such as fragmentation to micro and nanoscale particles [21] or photodegradation, which destabilizes the polymer structure [22] and causes leaching of dissolved organic carbon [3, 23,24,25]. In addition, the removal mechanism can also be biotic mineralization, mediated by microbes such as bacteria, archaea, or fungi. Plastic degradation by bacteria has been well documented for terrestrial and marine environments. For example, the terrestrial bacterium Ideonella sakaiensis hydrolyzes polyethylene terephthalate [26,27,28], while Rhodococcus ruber strain C208 has been found to degrade polyethylene (PE) and polystyrene (PS) [29,30,31]. In addition, the marine species Bacillus sphericus and Bacillus cereus have been shown to degrade polyethylene [32]. In contrast to bacteria, our knowledge of the potential role of fungi-mediated plastic degradation is in its infancy, especially in the marine environment. Due to their genetic and metabolic capabilities, fungi are best known as the main degraders of natural polymers such as wood, plants, cellulose, and lignin. Furthermore, fungi are efficient degraders of various complex hydrocarbons, including polycyclic aromatic hydrocarbons [33], oil, and alkanes [34, 35]—i.e., compounds that to some extent chemically resemble plastic. Fungi harbor powerful enzymatic machineries comprising, for example, manganese peroxidases, lignin peroxidases, and laccases [36,37,38], which have also been linked to plastic degradation [39, 40]. With respect to plastic degradation, multiple Aspergillus and Penicillium species, isolated from soils and gut microbiomes, were shown to degrade polyethylene [41,42,43,44,45,46]. Though fungi are common plastic colonizers in the ocean [47,48,49,50,51], only two species, Zalerion maritimum [52] and Alternaria alternata [53], have been identified as polyethylene degraders in the marine realm.

A general problem in measuring microbial plastic degradation and comparing results from different studies stems from methodological challenges and limitations. The chemical configuration of the used polymers, for example, the degree of crosslinking, crystallinity, and the addition of additives, as well as the size of the plastic particles and environmental/incubation conditions, likely affect microbial degradation [54]. The most commonly applied methods for investigating microbial plastic degradation include measuring the weight loss of polymers gravimetrically or determining polymer oxidation with Fourier-transform infrared spectroscopy [55]. In addition, scanning electron microscopy or laser scanning confocal microscopy has been applied to visualize ongoing plastic fragmentation and biofilm formation on the plastic surface [56, 57]. These approaches can typically not distinguish between abiotic and biotic degradation pathways, are not sensitive, and/or require time-consuming experimental procedures. Furthermore, none of these methods are suitable to directly trace carbon from the polymer into degradation products or microbial biomass (which would provide unambiguous proof for biodegradation) and they are also not suitable for determining microbial degradation kinetics.

Here, we show that the fungus Rhodotorula mucilaginosa, which we isolated from plastic debris from a North Sea laboratory microcosm, is capable of degrading UV-treated polyethylene. We incubated this fungus with virgin (-UV) and UV-irradiated 13C-labeled polyethylene and traced the polyethylene-derived carbon from the polymer source to the terminal oxidation product CO2 and measured polyethylene mineralization rates with unprecedented sensitivity. Furthermore, we visualized the assimilation of plastic-derived 13C into single cells of R. mucilaginosa by nanometer scale secondary ion mass spectrometry (nanoSIMS). This approach allows following the fate of plastic-derived carbon in marine ecosystems.

Materials and methods

Assays with 13C-polyethylene and Rhodotorula mucilaginosa

Details of the experimental setups are provided in the Supplementary appendix. In brief, Rhodotorula mucilaginosa (Supplementary Table S1) was isolated from a 350 L laboratory seawater microcosm containing a variety of plastic items mostly retrieved from the North Sea [58]. Two separate assays were performed to test the ability of the cultured R. mucilaginosa cells to mineralize polyethylene and assimilate polyethylene-derived carbon. The first approach involved incubating R. mucilaginosa cells in sealed bottles containing ~1–2 mg of 13C-labeled polyethylene (≥99% 13C, Sigma-Aldrich), either UV-treated or untreated, and measuring the transfer of 13C-label into the CO2 pool and cellular biomass (<1 week of incubation). The first assay was performed using 13C-labeled polyethylene as the sole energy and organic carbon source. In this experiment, the cultured cells were washed twice with sterile and autoclaved seawater, centrifuged (4000×g) at room temperature for 5 min, and “starved” for a week in sterile seawater at 25 °C to allow consumption of potentially remaining sucrose from medium prior to the incubation with polyethylene. In the second assay, we tested the ability of R. mucilaginosa to utilize polyethylene in the presence of another, more labile carbon source. In this setup, in addition to 13C-labeled polyethylene, the incubations also contained sucrose-based culture medium (10% of total liquid volume, 90% of seawater). Both assays included three treatments, each performed in triplicates: (i) 13C-labeled polyethylene and R. mucilaginosa, (ii) UV-treated 13C-labeled polyethylene and R. mucilaginosa, and (iii) uninoculated control with UV-treated 13C-labeled polyethylene.

Quantification of polyethylene degradation rates

The rate of polyethylene degradation in assays with polyethylene as the sole carbon source was determined from the increase in the total amount of 13C-CO2 in the incubation bottles over time. Firstly, the identity and concentration of the headspace CO2 gas were measured using gas chromatography with quadrupole mass spectrometry and flame ionization detection, respectively. The isotopic composition of headspace CO2 was then determined by isotope ratio mass spectrometry (GC-IRMS; details in the Supplementary Material). The pH of the liquid phase was determined with a pH meter (Mettler-Toledo Seven compact S210) at the end of the incubation. Concentrations of dissolved inorganic carbon (DIC) in the liquid phase were then determined from headspace CO2 and pH measurements [59, 60]. CO2 and DIC concentrations were then converted to the total amount of CO2 per incubation bottle (∑CO2). Excess 13C in the headspace CO2 and DIC (liquid phase) pool was calculated from the change in δ13C-CO2, which is equivalent to a change in the fractional abundance of 13C (13F [61]).

$${\,}^{13}F_{{{{{{\rm{sample}}}}}}} = \frac{{\left[ {\left( {\frac{{{\,}^{13}{{{{{\rm{C}}}}}}}}{{{\,}^{12}{{{{{\rm{C}}}}}}}}} \right)_{{{{{{{{\rm{standard}}}}}}}}} \times \left( {\frac{{\delta {\,}^{13}{{{{{\rm{C}}}}}} - {{{{{\rm{CO}}}}}}_2}}{{1000}} + 1} \right)} \right]}}{{1 + \left\lfloor {\left( {\frac{{{\,}^{13}{{{{{\rm{C}}}}}}}}{{{\,}^{12}{{{{{\rm{C}}}}}}}}} \right)_{{{{{{{{\rm{standard}}}}}}}}} \times \left( {\frac{{\delta {\,}^{13}{{{{{\rm{C}}}}}} - {{{{{\rm{CO}}}}}}_2}}{{1000}} + 1} \right)} \right\rfloor }}$$
(1)

Here, δ13C-CO2 is the measured stable isotope composition of headspace CO2, and 13C/12Cstandard is the stable carbon isotope ratio of the Vienna PeeDee Belemnite standard (VPDB). The only 13C-organic carbon source in our incubations was polyethylene. Hence, an increase in 13F in the ∑CO2 pool is caused by an excess amount of 13C (13Cex) originating from the added substrate:

$${\,}^{13}{{{{C}}}}_{ex }= ({\,}^{13}F_{t_{n}} -{\,}^{13}F_{t_{o}}) \times \sum {{{{{\rm{CO}}}}}}_2$$
(2)

The change over time in 13Cex (Δ13Cex) was calculated from the change in δ13C (thus 13F), i.e., the slope of δ13C (Fig. 1A) and ∑CO2 at the endpoint of the experiment. Δ13Cex is proportional to the mineralization rate of polyethylene-derived carbon. Mineralization rates were expressed as %-degradation of the initially added 13C-polyethylene (% d–1,% yr–1 Supplementary Table 2).

Fig. 1: Results of polyethylene degradation assays with R. mucilaginosa.
figure 1

A Development of δ13C-CO2 values in incubations with with 13C-polyethylene (PE) as the sole carbon source with prior UV-treatment (+UV) and without (-UV) with R. mucilaginosa (RM) and with 13C-polyethylene (PE) with prior UV-treatment without R. mucilaginosa. The strong increase in δ13C-CO2 in incubations with R. mucilaginosa provides evidence for the mineralization of polyethylene-derived 13C. B CO2 in the headspace (15 mL) of the incubations. All data are shown as averages ± standard deviation (n = 3). C, D Microscopic images of R. mucilaginosa stained with acridine orange and visualized with ×100 magnification, C R. mucilaginosa cells grown in sucrose-based medium (control), D R. mucilaginosa cells of incubation assay with untreated polyethylene as sole carbon source. Cells are attached to the polyethylene particles and form densely packed aggregates with polyethylene particles. The scale bar is equal to 5 µm.

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Quantification of polyethylene-derived carbon assimilation

Small aliquots (150 µL) of the liquid with the R. mucilaginosa biomass collected at the end of incubation experiments were filtered onto polycarbonate filters (0.2 µm pore size, Millipore), washed three times with 1×PBS, and placed in a desiccator at room temperature to dry. Chemical fixation was not performed, thus avoiding dilution of the isotope label. These samples were subsequently measured with nanoSIMS to quantify the 13C labeling of individual R. mucilaginosa cells (details in the Supplementary Material). In this analysis, R. mucilaginosa cells from the inoculum were used as controls. NanoSIMS data were processed and analyzed using Look@NanoSIMS [62].

Results

Polyethylene degradation rates

We conducted activity assays with 13C-labeled polyethylene as the sole organic carbon and energy source to investigate the potential of R. mucilaginosa to degrade and mineralize polyethylene. We used either untreated 13C-polyethylene or 13C-polyethylene that was irradiated prior to incubation with a UV A/B dose corresponding to ~50 and ~125 days of UV irradiation at the sea surface in the subtropical and temperate regions, respectively [63].

We found that the δ13C-values in the headspace CO2 pool increased linearly by 117‰ over 5 days of incubation when the experiment was conducted with R. mucilaginosa and UV-treated 13C-polyethylene (Fig. 1A). In contrast, during the same time interval, the δ13C-CO2 values increased by 16‰ in the experiments where the UV-treated 13C-polyethylene was incubated without R. mucilaginosa. The δ13C-CO2 values did not increase in incubations with untreated 13C-polyethylene and R. mucilaginosa. The CO2 concentrations in the headspace remained relatively constant, ~1900 ppm in incubations with fungal inoculum and ~2400 ppm in incubations without fungi (Fig. 1B). With respect to the headspace volumes, this amounts to a total of ~1.2 and 1.5 µmol CO2, respectively. Plastic mineralization was determined from the total amount of CO2 per incubation bottle (∑CO2), comprising CO2 in the headspace and DIC in the liquid phase. This, combined with the change in δ 13C-CO2, translates to an excess production of 13C in the carbonate system of the incubations (13F, see Materials and methods). The 13C excess production is equivalent to the mineralization rate of polyethylene-derived carbon. Based on linear regression analysis, the increase in δ13C in incubations with UV-treated 13C-polyethylene and with R. mucilaginosa was 31.5‰ d–1 (Fig. 1A). This is equivalent to an absolute change in 13F (Δ13F) of 0.000345 d–1 (Supplementary Table S2). Together with an average ∑CO2 of 57.7 µmol, this translates to a Δ13Cex of 0.0199 µmol d–1. In control incubations with UV-treated 13C-polyethylene but without R. mucilaginosa the increase in δ13C of 3.9‰ d–1 translates to a Δ13F of 0.000043 d–1 and together with a ∑CO2 value of 151.7 µmol to a Δ13Cex of 0.0064 µmol d–1. We attribute this excess production to ongoing radical chain reactions leading to polymer oxidation and CO2 production even after UV exposure has stopped [3, 24]. In contrast, δ13C increased insubstantially by only 0.01‰ d–1 in incubations with R. mucilaginosa but where the polyethylene was not irradiated with UV light prior to incubation. In these incubations, the Δ13F was 0.0000001 d–1, ∑CO2 was 48.2 µmol and Δ13Cex was hence 0.000006 µmol d–1. For the incubations with UV-treated polyethylene, we further calculated net polyethylene mineralization rates. Considering the ~1.8 and ~1.7 mg 13C-polyethylene added to incubations with and without fungi, the Δ13Cex in these incubations translates to polyethylene mineralization rates of 0.016% d–1 and 0.006% d–1, respectively. Hence, the net R. mucilaginosa mediated polyethylene degradation amounts to 0.01% d–1, or 3.8% yr–1. The R. mucilaginosa cell counts in this incubation were 3.32 × 105 ± 1.27 × 104 ml–1 resulting in mineralization rates of 6.05 × 10–5 nmol polyethylene cell–1 d–1 or 0.022 nmol polyethylene cell–1 yr–1.

nanoSIMS

In all sample sets, R. mucilaginosa cells featured similar cellular properties with two distinct subpopulations: enlarged cells of ~4 µm (likely growing/dividing cells or polyploids), and smaller cells with a diameter of ~2 µm (Fig. 2). Fluorescence microscopy showed a homogenous cell suspension (without formation of cell aggregates) of R. mucilaginosa cells in the original inoculum (Fig. 1C). In contrast, R. mucilaginosa cells adhered to plastic particles and clumped together in the incubations with added polyethylene (Fig. 1D). For nanoSIMS measurements of R. mucilaginosa cells, we analyzed three sample sets: (i) the original inoculum, (ii) after incubation with 13C-labeled polyethylene without and (iii) with UV-treatment. We measured a total of 1144 regions of interest (ROIs, i.e., corresponding to 1144 individual cells; Table 1). In control incubations with no added polyethylene, both cell types had mean 13F-values of 0.0105 and 0.0106, respectively (Figs. 2, 3 and Table 1). Similarly, small cells in incubations with untreated 13C-polyethylene showed 13F-values of 0.0105, but larger cells were slightly, yet significantly 13C enriched with 13F-values of 0.0107 (p < 0.001, Dunn’s Kruskal–Wallis test with Bonferroni correction). The small cells in incubations with UV-treated 13C-polyethylene had similar 13C-enrichment with 13F-values of 0.0107. However, the most substantial 13C-enrichment with 13F-values of 0.024 was found in large R. mucilaginosa cells with UV-treated 13C-polyethylene. This was significantly higher than any other cell group, irrespective of the added substrate (p < 0.001, Dunn’s Kruskal–Wallis test with Bonferroni correction). Results of the statistical analysis are presented in Supplementary Table S3.

Fig. 2: NanoSIMS images of R. mucilaginosa cells.
figure 2

A Secondary electron image of R. mucilaginosa cells in the medium (Control), B R. mucilaginosa cells with untreated 13C-polyethylene (-UV), C R. mucilaginosa cells with UV-treated 13C-polyethylene (+UV). The respective 13F-values of R. mucilaginosa cells are presented on panels D–F. Highest incorporation of 13C label was found in R. mucilaginosa cells with UV-treated 13C-polyethylene as depicted by warm colors in panel F.

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Table 1 13F-values of cells in the medium (Control), R. mucilaginosa cells with untreated 13C-polyethylene (13C-PE) and UV-treated 13C-polyethylene (UV 13C-PE).
Full size table
Fig. 3: Box and whiskers plot of 13F-values of small (SC) and larger (LC) R. mucilaginosa cells.
figure 3

A Cells in medium (Control), with untreated 13C-polyethylene (13C-PE), UV-treated polyethylene (UV 13C-PE). B A zoom in of Control and 13C-polyethylene treatments. Differences in 13F-values were statistically significant for 13C-polyethylene and UV 13C-polyethylene for small and enlarged cells. p values (t-test) are indicated in the plot. Boxplots depict the median, the first and third quartiles, the upper/lower whiskers that extend from the hinge to the largest/smallest value no further than 1.5× of the interquartile range from the hinge. Outliers are represented as big dots.

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To investigate if R. mucilaginosa also metabolizes polyethylene-derived carbon in the presence of other, potentially more accessible carbon sources, we additionally incubated R. mucilaginosa in seawater containing 10% of MS medium, with and without UV-treated 13C-polyethylene. NanoSIMS analysis revealed slightly 13C-enriched cells (13F-values of ~0.012) in incubations where UV-treated 13C-labeled polyethylene was added (Supplementary Fig. S1). In these incubations, the 13C-enrichment was more homogenous over all cell types. However, the 13C-enrichment was much lower compared to the incubations where R. mucilaginosa cells where exposed to UV-treated polyethylene as the sole carbon source and where large cells showed 13F-values of 0.024 (see above). No incorporation of the 13C to R. mucilaginosa cells was recorded in incubations where the 13C-polyethylene was not UV-treated.

Discussion

Microbial degradation, in conjunction with physicochemical processes, may be an important pathway in breaking down plastic litter in the marine environment. Yet, little is known about potential microorganisms degrading marine plastic litter because most quantitative techniques do not allow resolving degradation in the sub-percent range, nor do the experimental setups provide unambiguous proof of microbial utilization of plastics. In this work, we present data from short (<1 week) experimental assays that allowed us to quantitatively measure plastic mineralization rates. In addition, we traced and visualized the assimilation of plastic-derived carbon into individual cells of R. mucilaginosa. Central to this approach is the utilization of isotopically labeled polyethylene, containing a high degree of 13C instead of 12C, and tracing the polyethylene-derived 13C in the degradation product CO2 and biomass.

Degradation of 13C-polyethylene

Fungal mineralization of polyethylene-derived carbon occurred in incubations with UV-treated polyethylene, indicating that initial photooxidation enhances fungal plastic degradation in the marine environment. UV-induced photooxidation leads to the formation of carbonyl and hydroxyl moieties in the polymer [24, 64]. This facilitates access for microbial enzymes and thus further plastic degradation [65]. Moreover, UV-induced photodegradation results in the leaching of a plethora of lower molecular weight degradation products [3, 10, 23, 25, 66], which, at least to some degree, stimulate microbial activity [23, 25]. In our experiments, we did not measure photooxidation products other than CO2, and we can thus not determine which compounds were utilized by R. mucilaginosa. Nevertheless, the UV-treated 13C-polyethylene was washed and dried before incubation, which likely removed volatile compounds that were generated during the UV treatment. Longer chain degradation compounds and polymers with carbonyl and hydroxyl moieties [24], however, will likely have remained accessible for the fungus. For measuring degradation kinetics, we quantitatively traced 13C from isotopically labeled 13C-polyethylene into the terminal oxidation product CO2. Mineralization rates of UV-treated polyethylene with R. mucilaginosa amounted to 0.01% d–1; extrapolated, this translates to degradation rates of 3.8% yr–1 of the initially added polyethylene. These microbially mediated degradation rates are difficult to compare with previously reported rates from literature. Polymer weight loss over time is a frequently used parameter for measuring plastic degradation rates [67,68,69]. Polymer mass may, however, be lost as a result of fragmentation, and during clean-up procedures, so gravimetric methods do not allow a clear-cut distinction between biotic and abiotic degradation. Furthermore, resolving environmental plastic degradation rates of a few percent per year necessitates long incubation times of months to years in order to detect gravimetric changes. Also, Fourier-transform infrared spectroscopy allowing detection of carbonyl and hydroxyl groups in the polymer backbone [68, 70] does not allow a distinction between biotic and abiotic degradation. Chemical alteration of the initial polymer matrix can be the result of microbial degradation, but also caused by physical and chemical processes such as photooxidation. In contrast, using isotopically labeled polymers offers the advantage of directly and quantitatively tracing plastic-derived carbon into the terminal degradation product CO2. Our methodological approach allows resolving δ13C-CO2 values of ≳1‰ with confidence, which is equivalent to a change in the 13C content in the ∑CO2 pool of ~0.001%. This detection limit is determined by the background CO2 level as well as the added 13C-substrate. For a setup resembling ours (0.1 mmol ∑CO2, 1 mg of 99 atom % 13C-labeled polyethylene), degradation of ~0.002% of the added polyethylene is detectable. This method is consequently orders of magnitude more sensitive than more commonly used methods and also allows measuring plastic degradation rates in much shorter time periods of days to weeks as opposed to months to years.

Biomass incorporation of polyethylene-derived carbon

We measured stable carbon isotope ratios of single fungal cells using nanoSIMS, a technique that, in scanning mode, allows collecting images of isotopic composition with a spatial resolution in the nanometer scale [71]. Previously conducted incubations with the 13C-labeled biodegradable/compostable plastic poly (butylene adipate-co-terephthalate) (PBAT) and agricultural soil demonstrated incorporation of 13C from the PBAT into fungal hyphae and other unicellular organisms [72]. Also, labeled 13C-polyethylene has been used to trace plastic-derived carbon in food webs of a boreal-lake and artificial humic waters [73]. Our findings of highly 13C enriched cells provide, for the first time, unambiguous proof of the assimilation of polyethylene-derived carbon by a marine-derived fungus. The nanoSIMS measurements show that the incorporation of polyethylene-derived 13C was not homogenous into all fungal cells. We detected two subpopulations of cells, one with significant but low 13C-enrichment and a second population with an extraordinary high enrichment. The cells with higher enrichment were larger in size, thus probably growing or polyploids. Potentially, these cells were in closer proximity to the plastic substrate and incorporated more of the 13C label. The vicinity of the fungal cells to the plastic particles is likely crucial in the degradation process as the plastic substrate is not a soluble compound and is thus not homogenously distributed on a µm scale. Similar to kinetic measurements, we found higher assimilation of polyethylene-derived carbon in R. mucilaginosa cells in incubations with UV-treated polyethylene. Moreover, using nanoSIMS, we could also find a 13C-enrichment in R. mucilaginosa cells in incubation with 13C-polyethylene that was not UV irradiated prior to the incubation experiments. This indicates that the fungus can also degrade the virgin (-UV) polymer, though to a much lesser degree, as shown by the lower biomass 13C-enrichment when compared with our measurements with UV-treated polyethylene. Nevertheless, this will need further confirmation in future studies addressing molecular weight distribution of the polymer and the degree to which it contains short-chain impurities.

Finally, in incubations containing a sucrose-based medium and UV-treated 13C-polyethylene as potential carbon substrates, we observed label incorporation into R. mucilaginosa cells, although to a much lesser degree. However, this shows that R. mucilaginosa utilizes polyethylene-derived carbon also in the presence of more bioavailable carbon substrates that are easier to metabolize. This consequently implies that R. mucilaginosa could also metabolize polyethylene-derived carbon in the marine environment, where plastic only makes up a fraction of the plethora of available organic matter compounds.

Environmental implications

The role and identity of plastic-degrading microbes in the marine environment, specifically fungi is unconstrained. Nevertheless, many different fungi colonize plastic marine debris [49, 51, 74, 75] and thus potentially have access to plastic as a carbon source. Though several plastic-degrading fungi have been identified and isolated from terrestrial and freshwater environments [76,77,78], only two marine fungi belonging to the Ascomycota: Zalerion maritimum and Alternaria alternata are known to degrade plastic [52, 53]. Our isolated fungus, R. mucilaginosa, is a marine yeast belonging to the Basidiomycota. Yeasts have generally not been described as plastic degraders; however, yeasts are generally ubiquitous throughout fresh, marine, and deep sea environments [79]. Reported cell counts range from 10 to 50 cells L–1 in seawater compared to up to 500 cells L–1 in rivers [80]. The genus Rhodotorula is widespread throughout all ecosystems [81], including aquatic environments. R. mucilaginosa has been detected in lakes [82], hypersaline inland seas [83], arctic glaciers [84], and the deep sea [85] and was found as a dominant fungus on marine plastics [51]. R. mucilaginosa is common in bioremediation practices, where R. mucilaginosa strains have shown potential as nitrobenzene [86] and acrylamide degraders [87]. Similarly, R. mucilaginosa has been tested for its ability to remove phenolic compounds from olive mill wastewater [88, 89]. R. mucilaginosa thus seems to be able to break down a variety of hydrocarbon/hydrocarbon-like compounds. Together with our findings, this consequently suggests that Rhodotorula mucilaginosa is a potentially important plastic degrader in a wide range of marine environments. Furthermore, as this yeast has been found in a diversity of aquatic systems globally, it may hence degrade plastics there too.

Conclusion

Fungi in the marine environment are highly understudied despite their prevalence in the ocean. With the aid of stable isotope assays, we provide unambiguous proof that the fungus Rhodotorula mucilaginosa uses polyethylene-derived carbon for cellular incorporation and energy gain. The ability of R. mucilaginosa to utilize plastic-derived carbon in the presence of other, high-energy-yielding carbon substrates also indicates that fungal plastic degradation can indeed proceed in the natural environment. Our results confirm that initial plastic photooxidation is a key process in making plastic available for subsequent microbial degradation. Most produced and discarded plastic types such as polyethylene and polypropylene float at the ocean surface and will consequently be subjected to photooxidation so that fungal degradation can commence there. At least parts of the vast amounts of plastic litter in the ocean may thus serve as a carbon source for fungi and possibly other microbes, too.