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Biodegradation of polyethylene microplastics by the marine fungus Zalerion maritimum
Science of the Total Environment 586 (2017) 10–15
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Short Communication
Biodegradation of polyethylene microplastics by the marine fungus Zalerion maritimum Ana Paço a,⁎, Kátia Duarte a,b, João P. da Costa a,b, Patrícia S.M. Santos a,b, R. Pereira c, M.E. Pereira a,b, Ana C. Freitas d, Armando C. Duarte a,b, Teresa A.P. Rocha-Santos a,b a
Department of Chemistry, University of Aveiro, Campus de Santiago, 3810-193 Aveiro, Portugal CESAM, University of Aveiro, Campus de Santiago, 3810-193 Aveiro, Portugal Department of Biology, Faculty of Sciences of the University of Porto, CIIMAR – Interdisciplinary Centre of Marine and Environmental Research & Green UP/CITAB-UP, Porto, Portugal d Universidade Católica Portuguesa, CBQF - Centro de Biotecnologia e Química Fina – Escola Superior de Biotecnologia, Rua Arquiteto Lobão Vital, Apartado 2511, 4202-401 Porto, Portugal b c
H I G H L I G H T S
G R A P H I C A L
A B S T R A C T
• Plastic pollution is of growing concern as it accumulates in the environment. • Multiple microorganisms have been described as potential bioremediation solutions. • The potential of Zalerion maritimum in the biodegradation of polyethylene was tested. • Z. maritimum used polyethylene as evidenced by FTIR, NMR and SEM results. • The results highlight the prospective use of Z. maritimum as a bioremediation tool.
a r t i c l e
i n f o
Article history: Received 15 December 2016 Received in revised form 2 February 2017 Accepted 3 February 2017 Available online xxxx Editor: D. Barcelo Keywords: Microplastics Polyethylene Biodegradation Fungi Zalerion maritimum FTIR-ATR NMR
⁎ Corresponding author. E-mail address: [email protected] (A. Paço).
http://dx.doi.org/10.1016/j.scitotenv.2017.02.017 0048-9697/© 2017 Elsevier B.V. All rights reserved.
a b s t r a c t Plastic yearly production has surpassed the 300 million tons mark and recycling has all but failed in constituting a viable solution for the disposal of plastic waste. As these materials continue to accumulate in the environment, namely, in rivers and oceans, in the form of macro-, meso-, micro- and nanoplastics, it becomes of the utmost urgency to find new ways to curtail this environmental threat. Multiple efforts have been made to identify and isolate microorganisms capable of utilizing synthetic polymers and recent results point towards the viability of a solution for this problem based on the biodegradation of plastics resorting to selected microbial strains. Herein, the response of the fungus Zalerion maritimum to different times of exposition to polyethylene (PE) pellets, in a minimum growth medium, was evaluated, based on the quantified mass differences in both the fungus and the microplastic pellets used. Additionally, molecular changes were assessed through attenuated total reflectance Fourier transform Infrared Spectroscopy (FTIR-ATR) and Nuclear Magnetic Resonance (NMR). Results showed that, under the tested conditions, Z. maritimum is capable of utilizing PE, resulting in the decrease, in both mass and size, of the pellets. These results indicate that this naturally occurring fungus may actively contribute to the biodegradation of microplastics, requiring minimum nutrients. © 2017 Elsevier B.V. All rights reserved.
A. Paço et al. / Science of the Total Environment 586 (2017) 10–15
1. Introduction Microplastics, according to the National Oceanic and Atmospheric Administration (NOAA), are plastic particles with a diameter b 5 mm, while those b1 μm are defined as nanoplastics (GESAMP, 2015; NOAA, 2015). Microplastics can be classified according to their origin as either primary or secondary microplastics. The former refers to particulates originally produced with sizes b 5 mm, in the form of pellets, plastic-based granulates for the cosmetics industry, or as a vector for drugs in medicine. Secondary microplastics are those generated from the fragmentation of larger plastic particles, including macro- and mesoplastics, such as glass bottles and plastic bags (Cole et al., 2011; Rocha-Santos and Duarte, 2015; Wright et al., 2013). This fragmentation can be caused by chemical and physical aging, as well as through (bio)degradation mechanisms. The latter is the most usual process in the evolution of macroplastics to microplastics in the ocean (Lambert et al., 2014; Sivan, 2011). Recently, there have been numerous reports detailing the prominent increase in the amount of these particles in the oceans, with notorious consequences to the environment and biota (Browne et al., 2008; Cole et al., 2013; Ivar do Sul and Costa, 2014), including commercial species (Avio et al., 2015; Neves et al., 2015). Microplastics can be found throughout the globe, including in remote locations, such as sub-Antarctic islands or deep seas, suggesting a transfer from discharge areas to deposition zones (Caruso, 2015; Desforges et al., 2014; Woodall et al., 2014). Many works have, hence, described the concomitant ingestion of such particulates by multiple organisms, including zooplankton, which stands at the basis of the foodchain (Cole et al., 2013; Desforges et al., 2015; Long et al., 2015; Setälä et al., 2014). This poses a risk not only due to the inherent physical harm caused by microplastics, but also due to the possible presence of contaminants and pollutants that adhere and/or are adsorbed in microplastics, which may lead to bioaccumulation and bioamplification phenomena (da Costa et al., 2016). When in the environment, (micro)plastics are subject to biotic and/ or abiotic degradation mechanisms. Biotic degradation, or biodegradation, is mediated by microorganisms and, hence, is defined as a “process which is capable of decomposition of materials into carbon dioxide, methane, water, inorganic compounds, or biomass in which the predominant mechanism is the enzymatic action of microorganisms, that can be measured by standard tests, in a specified period of time, reflecting available disposal conditions” (ASTM, 2010). Biodegradation is influenced by both the characteristics of the polymer, including chemical and physical properties, and environmental factors, such as light (UV), heat, humidity and the presence of chemicals (Shah et al., 2008). Several fungi have been shown to be able to use plastics as the sole source of nutrients (Russell et al., 2011; Yamada-Onodera et al., 2001), including in solid matrices, such as soil (Bhardwaj et al., 2013) and compost (Zafar et al., 2013), thus highlighting the potential use of these organisms in the bioremediation of plastics, an increasing environmental concern, namely, in aquatic systems (Cole et al., 2011; da Costa et al., 2016) Herein, we demonstrate the capacity of Z. maritimum, a naturally occurring fungus in the marine environment and present in Portuguese coastal waters (Barata, 2006), as a biodegradation agent of polyethylene, one of the most widely used polymers. These results pave the way for developing new, innovative strategies aiming at mitigating the environmental impacts of (micro)plastics with low required investment, namely, the minimal nutrients need for its growth, making this an economically feasible approach. 2. Materials and methods 2.1. Microplastics Polyethylene pellets (PE, linear formula H(CH2CH2)nH), with a melt index of 1.0 g/10 min (190 °C/2.16 kg) were acquired from Sigma-
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Aldrich (USA). These exhibited spheroid morphology and were approximately 2–4 mm in size. The polyethylene (PE) pellets were mechanically cut and the resulting fragments sieved to obtain particles (microplastics) with a size range 250 μm b MP b 1000 μm. Microplastics were analysed by optical and electron microscopies (Fig. 1) and FTIRATR spectroscopy. 2.2. Biological material Zalerion maritimum, also commonly referred to as Z. varium, (ATTC 34329) was grown at 25 °C in a growth medium containing 20 g/L of glucose, 20 g/L of malt extract and 1 g/L of peptone (Rao, 2012), as well as 35 g/L of sea salts (Sigma-Aldrich, 2016), under stirring for 5 days prior to the microplastics' removal assays. 2.3. Culture conditions Batch reactors (100 mL Erlenmeyer flasks) with 0.130 g of microplastics and 50 mL of 10 times diluted minimum growth medium (2 g/L of glucose, 2 g/L of malt extract and 0.1 g/L of peptone with 35 g/L sea salts) was inoculated with 0.500 g of filtered Z. maritimum mycelium. This biomass was obtained from Z. maritimum grown in ideal conditions (Section 2.2). Batch reactors were incubated (HWY-200D, Lan Technics, USA) in the dark and stirring was maintained at 120 rpm's for a maximum of 28 days. Temperature was kept at a constant 25 °C. Three batch reactors (replicas) were sampled after incubation periods of 7, 14, 21 and 28 days. Six additional batch reactors were kept throughout the experience time, 28 days, as controls. Three contained only microplastics (control_microplastics) in the diluted minimal growth medium, in order to evaluate any potential effects derived from their presence in the growth medium. Three additional batches contained only fungi (control_fungus) in the diluted minimal growth medium, in order to evaluate the growth of the fungi in the medium without the microplastics. The replicate samples were retrieved from the shaker and the fungus biomass and microplastics were separated from the medium by filtration, using 47 mm diameter glass fibre filters (Whatman plc, UK). After, the biomass was inspected in order to identify and extract any plastic materials attached and, when necessary, examinations were aided by the use of an optical microscope. The biomass (wet weight) was recorded after filtration and was then subjected to an overnight drying process, at 100 °C (Binder, Germany) and the dry weight was recorded. Then, a sample of the initial fungus (0 days, to be used as reference), and those recovered after the tested times (7, 14, 21 and 28 days) were frozen and lyophilized for further analysis by FTIR-ATR spectroscopy, NMR and optical and electron microscopies. 2.4. Analytical techniques FTIR-ATR analyses were carried out using a Perkin Elmer (USA) Spectrum BX FTIR instrument. The pellets were analysed at a 4 cm−1 resolution within the 4000–550 nm range. Air was used for the background spectrum. For 1H NMR analyses, 20 mg of lyophilized mycelium were suspended in 700 μL of D2O and subsequently agitated in a vortex for 10 min. A sample volume of approximately 650 μL was placed in 5 mm NMR tubes (Aldrich528PP, 5 mm). Spectra were acquired using a Bruker Avance-III 400 MHz spectrometer, operating at 9.4 T. The 1H NMR spectra were recorded at room temperature, at a spinning rate of 15 kHz and with a pulse length of 90° and 3 μs. Optical microscopy of was carried out using an Olympus BX51 Polarizing Microscope. The samples were prepared by direct deposition of aliquots on glass slides. Electron microscopy morphological analyses were performed using a Field Emission Gun (FEG) - SEM Hitachi S4100 microscope (Japan), operated at 25 kV. The samples were prepared by
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Fig. 1. Optical (A) and electron microscopy (B) of the surface of a sample of the microplastic particles used. Note that, in B, the identifiable streak may have been caused by the mechanical cutting process.
deposition directly onto the carbon tape and then coated by carbon evaporation (Emitech K950X, France).
3. Results and discussion Fig. 2 shows both the observed biomass variation and the percentage of removed plastics in the tested periods. A positive correlation was noted, as an increase in weight percentage for Z. maritimum corresponded to an increase in the mass loss percentage of the plastic particles. This was particular evident for the samples exposed to the PE microplastics for 14 days, in which a biomass variation of 82.0% ± 2.1% was accompanied by a mass variation of the polymeric materials of 56.7% ± 2.9%, i.e., a removal exceeding 43%. Such parallel variations have been previously noted for both bacterial and fungal strains (Russell et al., 2011; Shah et al., 2013a; Shah et al., 2013b; Zafar et al., 2013), though these pertained to the degradation of polymers in solid matrices (soil). It is therefore possible to determine the kinetic parameters for the degradation of PE as well as the order of reaction. This was done by choosing the best adjustment to the speed of the reaction, as described by Barreto and co-workers (Barreto et al., 2011). Considering the equations describing zero ([A] − [A]0 = − kt), first (ln[A] − ln[A]0 = − kt) and second order (1/[A] − 1/[A]0 = kt) reactions, where [A] is the concentration at time t and [A]0 is the initial
concentration at zero time, the correlations coefficients were determined for each case and are summarized in Table 1. The correlation coefficients appear to indicate that the degradation of PE by Z. maritimum follows a fractional order of reaction, which is not surprising, as this is likely a complex reaction mechanism (Boudart and Djéga-Mariadassou, 2014), involving more than one enzyme, and thus not amenable to be described by simpler enzymatic reaction mechanisms (Barreto et al., 2011). Taken together with the measured growth for the control sample, results suggest that the slightly higher mass increase of Z. maritimum after 7 days, in the presence of PE pellets, may be attributable to the use of this polymeric material by the fungus as a nutrient source. Supporting this rationale is the fact that the majority of fungi biomass was formed in the first 7 days (83.1% ± 4.3% biomass variation) due to the nutrients present in the growth media; in this period the microplastics removal was practically null. After this initial high growth rate, the production of biomass deaccelerated but continued probably due to the degradation of polymeric material of the pellets whose higher removal rate was in fact observed between 7 and 14 days. Although the biomass variation was recorded, it was also necessary to ascertain whether these showed similar chemical profiles. Hence, FTIR-ATR analyses were carried out for the Z. maritimum mycelium following freeze-drying. Fig. 3 shows the FTIR-ATR spectra of Z. maritimum before, during and after exposure to PE microplastics. The spectrum for Z. maritimum was similar to those reported for other fungi (Mularczyk-Oliwa et al., 2013; Salman et al., 2010). Characteristically, the region between 3700 and 3050 cm−1 was noticeable, which is attributed to the bond vibrations of carboxyl, hydroxyl or phenol groups, and to amides' N\\H vibrations (Coates, 2000; Socrates, 2004). Other peaks were also present, namely, the 3050–3000 cm−1 peak, commonly attributed to C\\H bonds from lipids, two peaks in the 2996–2800 cm−1 region, caused by vibrations of CH2 and CH3 functional groups from lipids or proteins. Additionally, peaks within the 1800–1700 cm−1 region, caused by the C _O bonds, typically from lipids, and two other peaks found between 1700 and 1500 cm−1, caused by amides in proteins, were also clearly present. Carbon-Nitrogen bonds (C\\N) found in proteins contributed to the detected peaks 1500–1400 cm−1, with amines contributing to the two peaks noted at the 1450–1250 cm−1
Table 1 Kinetic parameters determined for the degradation of PE by Z. maritimum. The correlation coefficients are shown.
Fig. 2. Z. maritimum biomass variation (dry weight, %) and microplastic removal (%) during the experiment's interval. Note that the latter corresponds to a loss in mass. For visualization purposes, the absolute values were plotted. Data were acquired in weekly intervals. Error bars are shown.
Order of reaction
Equation
Correlation coefficient (R2)
Zero First Second
y = −2.909× + 100.39 y = −0.0506× + 4.6591 y = 0.001× + 0.008
0.870 0.908 0.937
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microplastics, it is possible to assume that the fungi used the microplastics as source of nutrients, in particular after 7 days. Based on the aforementioned observations, the biomass of Z. maritimum at the beginning and following a 28 day exposure to microplastics and grown only in minimum medium (control) were further analysed through 1H NMR spectroscopy. Results are shown in Fig. 4A. Based on the chemical shift assignments present in the organic compounds, it is possible to consider four regions: δH = 0.6–1.8 ppm from aliphatic protons, H\\C, \\CH N \\CH2 N \\CH3; δH = 1.8– 3.2 ppm from aliphatic protons bound to carbon atoms in α position to unsaturated groups in allylic groups (H\\Cα\\H_), carbonyl (H\\Cα\\C_ O) or imino groups (H\\Cα\\C_ N); δH =3.2–4.1 ppm from aliphatic protons on carbon atoms singly bound to oxygen atoms (H\\C\\O: H\\C\\O\\CO\\R N H\\C\\OH or H\\C\\O\\C) in alcohols, polyols, ethers and esters; δH = 4.1–6.0 ppm from anomeric protons of glycosidic structures (H\\C(\\O)\\O) (Lopes et al., 2015; Santos et al., 2012). Peaks at 5.25 ppm and 3.25 were observed, which have been associated to α- and β-glucose, respectively, as well as a peak at approximately 4 ppm, attributable to fructose (Caligiani et al., 2007). In fact, this is the reason why some authors refer to this region as the sugar or carbohydrate region (4.1–3.3 ppm) (Kutyshenko et al., 2015). The observed peaks between 3.3 ppm and 1.7 ppm correspond to different amino acids, and the peak at 1.28 ppm has been assigned to lipids. Lastly, the peak at 0.9 ppm corresponds to the methyl groups of lipids and proteins (Kutyshenko et al., 2015). When compared, the intensities of these peaks in the different spectra in the region 6 to 3.3 ppm appear to increase with exposure. The peaks' intensities are also higher for the control sample, as evidenced in Fig. 4B, showing the determined relative areas for each identifiable region. These observations are consistent with the FTIR results, as they support the hypothesis of an increase in the content of carbohydrates with exposure time. Moreover, this is accompanied by a decrease in the 3.3 to 0.9 ppm region, which is also consistent with the previous findings, suggesting a decreasing concentration of lipids and proteins along the exposure to microplastics. In order to assess the potential assimilation of microplastics by Z. maritimum, a sub-sample of the fungus was subjected to lyophilisation and to subsequent visual analysis in order to identify potentially assimilated microplastics in the mycelium. As depicted in Fig. 5, the PE plastic particles were recovered from replicates at every sampling time (7, 14, 21 and 28 days) and, as observable, some of these were reduced to threads. Optical and electron microscopy results also revealed morphological changes to the PE particles, when compared to the results shown in Fig. 1. As evidenced in Fig. 5A, the polymeric particles seem to have been shredded, and the surface smoothness and regularity exhibited in Fig. 1B appears to have given place to a pronounced irregularity at the surface of the pellets. Noteworthy, electron microscopy results (Fig. 5B) also clearly indicate the presence of biological material at the pellets' surface, thus corroborating previous reports describing the growth of fungi at the surface of polyethylene (Esmaeili et al., 2013; Pramila and Ramesh, 2011). These noted differences may stem from the exposure of these particles to the fungal biomass. Identical findings have been reported by other authors, when assessing the biodegradation of polymeric materials (Jecu et al., 2012), thus highlighting the
Fig. 3. FTIR-ATR spectra for the different samples of Z. maritimum, including at the time of inoculation, grown in the presence of microplastics for 7 (F7), 14 (F14), 21 (F21) and 28 days (F28). The control sample corresponds to Z. maritimum grown for 28 days with no contact with the microplastic particles.
region. Lastly, the peak between 1200 and 1100 cm−1 is frequently attributed to the vibrations of C\\O bonds, found in carbohydrates, as well as the peak at 1080–1000 cm−1. Also noteworthy was the peak at 1100–1000 cm−1, assigned to the PO2 group present in nucleic acids (Sivan et al., 2006; Yamada-Onodera et al., 2001). As noted in Fig. 3, there was an overall decrease in the intensities of theses peaks throughout the duration of the experiment, except for the peak associated to the C\\O vibrations of carbohydrates (1200–1100 cm−1). These results suggest that there is a reduction in the lipidic and proteic content of Z. maritimum when exposed to microplastics. Moreover, it is possible to conjecture that this reduction was likely due to the reduced nutrient medium in which the fungus was grown (Weete, 2012), leading to the use of lipids as an energy source (Prasad and Ghannoum, 1996). The reduction of proteins and increase of carbohydrates can be explained by the changes caused in the fungal metabolism when exposed to a medium with a reduced carbon or nitrogen source. These induce fungi to produce proteolytic enzymes that degrade the intracellular proteins, as the deamination of amino acids provides the C skeletons required for energy purposes (McIntyre et al., 2000; Nitsche et al., 2012). These also induce nutrient recycling, mediated by glycosyl hydrolases and other enzymes that are synthesized aiming at the release of carbohydrates present in the cell wall by remodelling it, leading to an increase in the accumulation of carbohydrates (McIntyre et al., 2000; Nitsche et al., 2012; Pollack et al., 2008). Hence, for the control sample (Z. maritimum grown in minimum medium without microplastics), the lack of an alternative carbon source resulted in an endogenous search for energy. This ultimately led to the substantial differences observed in the FTIR spectra, which are made evident by the calculated corresponding areas, shown in Table 2. The overall behaviour seen in the spectra can therefore be attributed to the lack of nutrients in the medium, which in the case of the control fungus, Z. maritimum not exposed to microplastics, lead to a search for endogenous sources of carbon, and to a variation in the spectrum more significant. In the case of the Z. maritimum exposed to
Table 2 Relative areas for the different regions of the FTIR-ATR spectra (in triplicate) of Z. maritimum, grown in the presence of microplastics for 7 (F7), 14 (F14), 21 (F21) and 28 days (F28). The control sample corresponds to Z. maritimum grown for 28 days with no contact with the microplastic particles. Region (cm−1)
Inoculum
F7
F14
F21
F28
Control
3700–3000 3000–2800 1800–1700 1700–1500 1500–1250 1200–1100 1100–1000
64 ± 2 16.5 ± 0.8 1.2 ± 0.5 20.79 ± 0.04 17 ± 1 5±1 23 ± 2
21 ± 2 4.1 ± 0.7 0.9 ± 0.8 10.3 ± 0.4 6.3 ± 0.2 9.0 ± 0.4 9.6 ± 0.2
23.3 ± 0.3 2.4 ± 0.2 0.2 ± 0.2 9.1 ± 0.1 3.6 ± 0.6 7.5 ± 0.6 13.4 ± 0.8
34 ± 3 7±1 0.2 ± 0.1 12.9 ± 0.3 7.7 ± 0.6 8.5 ± 0.1 13 ± 1
24 ± 2 2.74 ± 0.09 0.3 ± 0.4 10 ± 1 5.0 ± 0.7 7.8 ± 0.5 10.4 ± 0.6
28.6 ± 0.9 4.8 ± 0.5 0.6 ± 0.4 10.7 ± 0.9 7.5 ± 0.3 7±1 15 ± 2
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A. Paço et al. / Science of the Total Environment 586 (2017) 10–15
Fig. 6. Infrared spectra in the region 600–4000 cm−1 of microplastics exposed to Z. maritimum for different periods of time. The observed differences in the spectra highlight the different degrees of degradation.
increasing vibrations. Such observations are further corroborated by the calculated areas for the aforementioned regions, shown in Table 3, clearly highlighting the chemical modifications detected through FTIRATR spectroscopy. 4. Conclusions
Fig. 4. In A), 1H NMR spectra of the lyophilized samples of Z. maritimum. In B), the corresponding relative areas from specific regions of the 1H NMR spectra obtained, corresponding to different classes of biomolecules (see text), are shown.
biological action of such organisms over these materials and their resulting biodegradation potential. The particles recovered from the samples grown for 14, 21 and 28 days were subsequently analysed by FTIR-ATR spectroscopy and the results are shown in Fig. 6. As evidenced in Fig. 6, there was a gradual increase in band intensities for peaks at 3700–3000 cm−1, caused by the hydroperoxide and hydroxyl groups. Similarly, areas in the region of 1700–1500 cm−1 and 1200–950 cm−1, caused by carbonyl groups and by double bonds, respectively, also gradually increased. These may result from different oxidation reactions with, for example, the functional groups present in the polymer (Barbeş et al., 2014; Ołdak et al., 2005), thus yielding the noted
This study underlines the high potential of Z. maritimum for the biodegradation of (micro)plastics, thus responding to the current and urgent need of alternative routes to minimize and abate the presence of these materials in aquatic systems and, more specifically, in the oceans. This organism was evidenced to grow in a minimum growth medium in the presence of microplastic particles, which, based on the observed modifications, point to their use as substrate by Z. maritimum. This assertion is based on the mass variations, as well as on the FTIR and NMR results. These support such conclusion, as the noted reduction in lipid and protein content, accompanied by an increase in carbohydrate concentration occurred to a larger extent in the fungus not exposed to microplastics. Due to the natural occurrence of this organism in maritime coastal waters, this study underlines the high potential of this fungus in future bioremediation strategies aiming at curtailing the increasing presence of (micro)plastics in the environment. Further studies are required in order to evaluate the likely mechanisms at the core of this demonstrated capacity of Z. maritimum to use polyethylene as a substrate and these are already underway. Acknowledgements This work was supported by national funds through FCT/MEC (PIDDAC) under project IF/00407/2013/CP1162/CT0023. Thanks are
Fig. 5. Optical (A) and electron (B) microscopy images of particles exposed to Z. maritimum for 28 days. Despite less evident in A, electron microscopy results (B) highlight the marked differences between the microplastics before and after exposure to Z. maritimum, which show pronounced surface irregularities and also clearly indicate the presence of biological material at the pellets' surface.
A. Paço et al. / Science of the Total Environment 586 (2017) 10–15 Table 3 Relative area from the identified regions from the FTIR-ATR spectra for the recovered microplastic particles (in triplicate). Region (cm−1)
PE
PE_14Days
PE_21Days
PE_28Days
3700–3000 3000–2800 1750–1500 1500–1400 1400–1300 1200–1000 750–650
1±1 24.1 ± 0.2 0.2 ± 0.3 3.8 ± 0.5 0.3 ± 0.2 0.3 ± 0.2 2.5 ± 0.3
15 ± 2 10.1 ± 0.2 5.5 ± 0.2 3.07 ± 0.08 1.7 ± 0.5 5.0 ± 0.3 2.5 ± 0.4
20 ± 2 22.3 ± 0.2 5.5 ± 0.5 4.71 ± 0.07 1.6 ± 0.5 7.8 ± 0.1 3.8 ± 0.4
57 ± 2 15 ± 1 1±1 4.6 ± 0.1 2.5 ± 0.8 6±2 3.4 ± 0.5
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Mularczyk-Oliwa, M., Bombalska, A., Kaliszewski, M., Włodarski, M., Kwaśny, M., Kopczyński, K., et al., 2013. Rapid discrimination of several fungus species with FTIR spectroscopy and statistical analysis. Biuletyn Wojskowej Akademii Technicznej 62, 71–80. Neves, D., Sobral, P., Ferreira, J.L., Pereira, T., 2015. Ingestion of microplastics by commercial fish off the Portuguese coast. Mar. Pollut. Bull. 101, 119–126. Nitsche, B.M., Jørgensen, T.R., Akeroyd, M., Meyer, V., Ram, A.F., 2012. The Carbon Starvation Response of Aspergillus niger during Submerged Cultivation. NOAA, Administration NOAA, 2015. Microplastics One Pager. Department of Commerce, MD, USA. Ołdak, D., Kaczmarek, H., Buffeteau, T., Sourisseau, C., 2005. Photo-and bio-degradation processes in polyethylene, cellulose and their blends studied by ATR-FTIR and Raman spectroscopies. J. Mater. Sci. 40, 4189–4198. Pollack, J.K., Li, Z.J., Marten, M.R., 2008. Fungal mycelia show lag time before re-growth on endogenous carbon. 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Science of the Total Environment journal homepage: www.elsevier.com/locate/scitotenv
Short Communication
Biodegradation of polyethylene microplastics by the marine fungus Zalerion maritimum Ana Paço a,⁎, Kátia Duarte a,b, João P. da Costa a,b, Patrícia S.M. Santos a,b, R. Pereira c, M.E. Pereira a,b, Ana C. Freitas d, Armando C. Duarte a,b, Teresa A.P. Rocha-Santos a,b a
Department of Chemistry, University of Aveiro, Campus de Santiago, 3810-193 Aveiro, Portugal CESAM, University of Aveiro, Campus de Santiago, 3810-193 Aveiro, Portugal Department of Biology, Faculty of Sciences of the University of Porto, CIIMAR – Interdisciplinary Centre of Marine and Environmental Research & Green UP/CITAB-UP, Porto, Portugal d Universidade Católica Portuguesa, CBQF - Centro de Biotecnologia e Química Fina – Escola Superior de Biotecnologia, Rua Arquiteto Lobão Vital, Apartado 2511, 4202-401 Porto, Portugal b c
H I G H L I G H T S
G R A P H I C A L
A B S T R A C T
• Plastic pollution is of growing concern as it accumulates in the environment. • Multiple microorganisms have been described as potential bioremediation solutions. • The potential of Zalerion maritimum in the biodegradation of polyethylene was tested. • Z. maritimum used polyethylene as evidenced by FTIR, NMR and SEM results. • The results highlight the prospective use of Z. maritimum as a bioremediation tool.
a r t i c l e
i n f o
Article history: Received 15 December 2016 Received in revised form 2 February 2017 Accepted 3 February 2017 Available online xxxx Editor: D. Barcelo Keywords: Microplastics Polyethylene Biodegradation Fungi Zalerion maritimum FTIR-ATR NMR
⁎ Corresponding author. E-mail address: [email protected] (A. Paço).
http://dx.doi.org/10.1016/j.scitotenv.2017.02.017 0048-9697/© 2017 Elsevier B.V. All rights reserved.
a b s t r a c t Plastic yearly production has surpassed the 300 million tons mark and recycling has all but failed in constituting a viable solution for the disposal of plastic waste. As these materials continue to accumulate in the environment, namely, in rivers and oceans, in the form of macro-, meso-, micro- and nanoplastics, it becomes of the utmost urgency to find new ways to curtail this environmental threat. Multiple efforts have been made to identify and isolate microorganisms capable of utilizing synthetic polymers and recent results point towards the viability of a solution for this problem based on the biodegradation of plastics resorting to selected microbial strains. Herein, the response of the fungus Zalerion maritimum to different times of exposition to polyethylene (PE) pellets, in a minimum growth medium, was evaluated, based on the quantified mass differences in both the fungus and the microplastic pellets used. Additionally, molecular changes were assessed through attenuated total reflectance Fourier transform Infrared Spectroscopy (FTIR-ATR) and Nuclear Magnetic Resonance (NMR). Results showed that, under the tested conditions, Z. maritimum is capable of utilizing PE, resulting in the decrease, in both mass and size, of the pellets. These results indicate that this naturally occurring fungus may actively contribute to the biodegradation of microplastics, requiring minimum nutrients. © 2017 Elsevier B.V. All rights reserved.
A. Paço et al. / Science of the Total Environment 586 (2017) 10–15
1. Introduction Microplastics, according to the National Oceanic and Atmospheric Administration (NOAA), are plastic particles with a diameter b 5 mm, while those b1 μm are defined as nanoplastics (GESAMP, 2015; NOAA, 2015). Microplastics can be classified according to their origin as either primary or secondary microplastics. The former refers to particulates originally produced with sizes b 5 mm, in the form of pellets, plastic-based granulates for the cosmetics industry, or as a vector for drugs in medicine. Secondary microplastics are those generated from the fragmentation of larger plastic particles, including macro- and mesoplastics, such as glass bottles and plastic bags (Cole et al., 2011; Rocha-Santos and Duarte, 2015; Wright et al., 2013). This fragmentation can be caused by chemical and physical aging, as well as through (bio)degradation mechanisms. The latter is the most usual process in the evolution of macroplastics to microplastics in the ocean (Lambert et al., 2014; Sivan, 2011). Recently, there have been numerous reports detailing the prominent increase in the amount of these particles in the oceans, with notorious consequences to the environment and biota (Browne et al., 2008; Cole et al., 2013; Ivar do Sul and Costa, 2014), including commercial species (Avio et al., 2015; Neves et al., 2015). Microplastics can be found throughout the globe, including in remote locations, such as sub-Antarctic islands or deep seas, suggesting a transfer from discharge areas to deposition zones (Caruso, 2015; Desforges et al., 2014; Woodall et al., 2014). Many works have, hence, described the concomitant ingestion of such particulates by multiple organisms, including zooplankton, which stands at the basis of the foodchain (Cole et al., 2013; Desforges et al., 2015; Long et al., 2015; Setälä et al., 2014). This poses a risk not only due to the inherent physical harm caused by microplastics, but also due to the possible presence of contaminants and pollutants that adhere and/or are adsorbed in microplastics, which may lead to bioaccumulation and bioamplification phenomena (da Costa et al., 2016). When in the environment, (micro)plastics are subject to biotic and/ or abiotic degradation mechanisms. Biotic degradation, or biodegradation, is mediated by microorganisms and, hence, is defined as a “process which is capable of decomposition of materials into carbon dioxide, methane, water, inorganic compounds, or biomass in which the predominant mechanism is the enzymatic action of microorganisms, that can be measured by standard tests, in a specified period of time, reflecting available disposal conditions” (ASTM, 2010). Biodegradation is influenced by both the characteristics of the polymer, including chemical and physical properties, and environmental factors, such as light (UV), heat, humidity and the presence of chemicals (Shah et al., 2008). Several fungi have been shown to be able to use plastics as the sole source of nutrients (Russell et al., 2011; Yamada-Onodera et al., 2001), including in solid matrices, such as soil (Bhardwaj et al., 2013) and compost (Zafar et al., 2013), thus highlighting the potential use of these organisms in the bioremediation of plastics, an increasing environmental concern, namely, in aquatic systems (Cole et al., 2011; da Costa et al., 2016) Herein, we demonstrate the capacity of Z. maritimum, a naturally occurring fungus in the marine environment and present in Portuguese coastal waters (Barata, 2006), as a biodegradation agent of polyethylene, one of the most widely used polymers. These results pave the way for developing new, innovative strategies aiming at mitigating the environmental impacts of (micro)plastics with low required investment, namely, the minimal nutrients need for its growth, making this an economically feasible approach. 2. Materials and methods 2.1. Microplastics Polyethylene pellets (PE, linear formula H(CH2CH2)nH), with a melt index of 1.0 g/10 min (190 °C/2.16 kg) were acquired from Sigma-
11
Aldrich (USA). These exhibited spheroid morphology and were approximately 2–4 mm in size. The polyethylene (PE) pellets were mechanically cut and the resulting fragments sieved to obtain particles (microplastics) with a size range 250 μm b MP b 1000 μm. Microplastics were analysed by optical and electron microscopies (Fig. 1) and FTIRATR spectroscopy. 2.2. Biological material Zalerion maritimum, also commonly referred to as Z. varium, (ATTC 34329) was grown at 25 °C in a growth medium containing 20 g/L of glucose, 20 g/L of malt extract and 1 g/L of peptone (Rao, 2012), as well as 35 g/L of sea salts (Sigma-Aldrich, 2016), under stirring for 5 days prior to the microplastics' removal assays. 2.3. Culture conditions Batch reactors (100 mL Erlenmeyer flasks) with 0.130 g of microplastics and 50 mL of 10 times diluted minimum growth medium (2 g/L of glucose, 2 g/L of malt extract and 0.1 g/L of peptone with 35 g/L sea salts) was inoculated with 0.500 g of filtered Z. maritimum mycelium. This biomass was obtained from Z. maritimum grown in ideal conditions (Section 2.2). Batch reactors were incubated (HWY-200D, Lan Technics, USA) in the dark and stirring was maintained at 120 rpm's for a maximum of 28 days. Temperature was kept at a constant 25 °C. Three batch reactors (replicas) were sampled after incubation periods of 7, 14, 21 and 28 days. Six additional batch reactors were kept throughout the experience time, 28 days, as controls. Three contained only microplastics (control_microplastics) in the diluted minimal growth medium, in order to evaluate any potential effects derived from their presence in the growth medium. Three additional batches contained only fungi (control_fungus) in the diluted minimal growth medium, in order to evaluate the growth of the fungi in the medium without the microplastics. The replicate samples were retrieved from the shaker and the fungus biomass and microplastics were separated from the medium by filtration, using 47 mm diameter glass fibre filters (Whatman plc, UK). After, the biomass was inspected in order to identify and extract any plastic materials attached and, when necessary, examinations were aided by the use of an optical microscope. The biomass (wet weight) was recorded after filtration and was then subjected to an overnight drying process, at 100 °C (Binder, Germany) and the dry weight was recorded. Then, a sample of the initial fungus (0 days, to be used as reference), and those recovered after the tested times (7, 14, 21 and 28 days) were frozen and lyophilized for further analysis by FTIR-ATR spectroscopy, NMR and optical and electron microscopies. 2.4. Analytical techniques FTIR-ATR analyses were carried out using a Perkin Elmer (USA) Spectrum BX FTIR instrument. The pellets were analysed at a 4 cm−1 resolution within the 4000–550 nm range. Air was used for the background spectrum. For 1H NMR analyses, 20 mg of lyophilized mycelium were suspended in 700 μL of D2O and subsequently agitated in a vortex for 10 min. A sample volume of approximately 650 μL was placed in 5 mm NMR tubes (Aldrich528PP, 5 mm). Spectra were acquired using a Bruker Avance-III 400 MHz spectrometer, operating at 9.4 T. The 1H NMR spectra were recorded at room temperature, at a spinning rate of 15 kHz and with a pulse length of 90° and 3 μs. Optical microscopy of was carried out using an Olympus BX51 Polarizing Microscope. The samples were prepared by direct deposition of aliquots on glass slides. Electron microscopy morphological analyses were performed using a Field Emission Gun (FEG) - SEM Hitachi S4100 microscope (Japan), operated at 25 kV. The samples were prepared by
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A. Paço et al. / Science of the Total Environment 586 (2017) 10–15
Fig. 1. Optical (A) and electron microscopy (B) of the surface of a sample of the microplastic particles used. Note that, in B, the identifiable streak may have been caused by the mechanical cutting process.
deposition directly onto the carbon tape and then coated by carbon evaporation (Emitech K950X, France).
3. Results and discussion Fig. 2 shows both the observed biomass variation and the percentage of removed plastics in the tested periods. A positive correlation was noted, as an increase in weight percentage for Z. maritimum corresponded to an increase in the mass loss percentage of the plastic particles. This was particular evident for the samples exposed to the PE microplastics for 14 days, in which a biomass variation of 82.0% ± 2.1% was accompanied by a mass variation of the polymeric materials of 56.7% ± 2.9%, i.e., a removal exceeding 43%. Such parallel variations have been previously noted for both bacterial and fungal strains (Russell et al., 2011; Shah et al., 2013a; Shah et al., 2013b; Zafar et al., 2013), though these pertained to the degradation of polymers in solid matrices (soil). It is therefore possible to determine the kinetic parameters for the degradation of PE as well as the order of reaction. This was done by choosing the best adjustment to the speed of the reaction, as described by Barreto and co-workers (Barreto et al., 2011). Considering the equations describing zero ([A] − [A]0 = − kt), first (ln[A] − ln[A]0 = − kt) and second order (1/[A] − 1/[A]0 = kt) reactions, where [A] is the concentration at time t and [A]0 is the initial
concentration at zero time, the correlations coefficients were determined for each case and are summarized in Table 1. The correlation coefficients appear to indicate that the degradation of PE by Z. maritimum follows a fractional order of reaction, which is not surprising, as this is likely a complex reaction mechanism (Boudart and Djéga-Mariadassou, 2014), involving more than one enzyme, and thus not amenable to be described by simpler enzymatic reaction mechanisms (Barreto et al., 2011). Taken together with the measured growth for the control sample, results suggest that the slightly higher mass increase of Z. maritimum after 7 days, in the presence of PE pellets, may be attributable to the use of this polymeric material by the fungus as a nutrient source. Supporting this rationale is the fact that the majority of fungi biomass was formed in the first 7 days (83.1% ± 4.3% biomass variation) due to the nutrients present in the growth media; in this period the microplastics removal was practically null. After this initial high growth rate, the production of biomass deaccelerated but continued probably due to the degradation of polymeric material of the pellets whose higher removal rate was in fact observed between 7 and 14 days. Although the biomass variation was recorded, it was also necessary to ascertain whether these showed similar chemical profiles. Hence, FTIR-ATR analyses were carried out for the Z. maritimum mycelium following freeze-drying. Fig. 3 shows the FTIR-ATR spectra of Z. maritimum before, during and after exposure to PE microplastics. The spectrum for Z. maritimum was similar to those reported for other fungi (Mularczyk-Oliwa et al., 2013; Salman et al., 2010). Characteristically, the region between 3700 and 3050 cm−1 was noticeable, which is attributed to the bond vibrations of carboxyl, hydroxyl or phenol groups, and to amides' N\\H vibrations (Coates, 2000; Socrates, 2004). Other peaks were also present, namely, the 3050–3000 cm−1 peak, commonly attributed to C\\H bonds from lipids, two peaks in the 2996–2800 cm−1 region, caused by vibrations of CH2 and CH3 functional groups from lipids or proteins. Additionally, peaks within the 1800–1700 cm−1 region, caused by the C _O bonds, typically from lipids, and two other peaks found between 1700 and 1500 cm−1, caused by amides in proteins, were also clearly present. Carbon-Nitrogen bonds (C\\N) found in proteins contributed to the detected peaks 1500–1400 cm−1, with amines contributing to the two peaks noted at the 1450–1250 cm−1
Table 1 Kinetic parameters determined for the degradation of PE by Z. maritimum. The correlation coefficients are shown.
Fig. 2. Z. maritimum biomass variation (dry weight, %) and microplastic removal (%) during the experiment's interval. Note that the latter corresponds to a loss in mass. For visualization purposes, the absolute values were plotted. Data were acquired in weekly intervals. Error bars are shown.
Order of reaction
Equation
Correlation coefficient (R2)
Zero First Second
y = −2.909× + 100.39 y = −0.0506× + 4.6591 y = 0.001× + 0.008
0.870 0.908 0.937
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microplastics, it is possible to assume that the fungi used the microplastics as source of nutrients, in particular after 7 days. Based on the aforementioned observations, the biomass of Z. maritimum at the beginning and following a 28 day exposure to microplastics and grown only in minimum medium (control) were further analysed through 1H NMR spectroscopy. Results are shown in Fig. 4A. Based on the chemical shift assignments present in the organic compounds, it is possible to consider four regions: δH = 0.6–1.8 ppm from aliphatic protons, H\\C, \\CH N \\CH2 N \\CH3; δH = 1.8– 3.2 ppm from aliphatic protons bound to carbon atoms in α position to unsaturated groups in allylic groups (H\\Cα\\H_), carbonyl (H\\Cα\\C_ O) or imino groups (H\\Cα\\C_ N); δH =3.2–4.1 ppm from aliphatic protons on carbon atoms singly bound to oxygen atoms (H\\C\\O: H\\C\\O\\CO\\R N H\\C\\OH or H\\C\\O\\C) in alcohols, polyols, ethers and esters; δH = 4.1–6.0 ppm from anomeric protons of glycosidic structures (H\\C(\\O)\\O) (Lopes et al., 2015; Santos et al., 2012). Peaks at 5.25 ppm and 3.25 were observed, which have been associated to α- and β-glucose, respectively, as well as a peak at approximately 4 ppm, attributable to fructose (Caligiani et al., 2007). In fact, this is the reason why some authors refer to this region as the sugar or carbohydrate region (4.1–3.3 ppm) (Kutyshenko et al., 2015). The observed peaks between 3.3 ppm and 1.7 ppm correspond to different amino acids, and the peak at 1.28 ppm has been assigned to lipids. Lastly, the peak at 0.9 ppm corresponds to the methyl groups of lipids and proteins (Kutyshenko et al., 2015). When compared, the intensities of these peaks in the different spectra in the region 6 to 3.3 ppm appear to increase with exposure. The peaks' intensities are also higher for the control sample, as evidenced in Fig. 4B, showing the determined relative areas for each identifiable region. These observations are consistent with the FTIR results, as they support the hypothesis of an increase in the content of carbohydrates with exposure time. Moreover, this is accompanied by a decrease in the 3.3 to 0.9 ppm region, which is also consistent with the previous findings, suggesting a decreasing concentration of lipids and proteins along the exposure to microplastics. In order to assess the potential assimilation of microplastics by Z. maritimum, a sub-sample of the fungus was subjected to lyophilisation and to subsequent visual analysis in order to identify potentially assimilated microplastics in the mycelium. As depicted in Fig. 5, the PE plastic particles were recovered from replicates at every sampling time (7, 14, 21 and 28 days) and, as observable, some of these were reduced to threads. Optical and electron microscopy results also revealed morphological changes to the PE particles, when compared to the results shown in Fig. 1. As evidenced in Fig. 5A, the polymeric particles seem to have been shredded, and the surface smoothness and regularity exhibited in Fig. 1B appears to have given place to a pronounced irregularity at the surface of the pellets. Noteworthy, electron microscopy results (Fig. 5B) also clearly indicate the presence of biological material at the pellets' surface, thus corroborating previous reports describing the growth of fungi at the surface of polyethylene (Esmaeili et al., 2013; Pramila and Ramesh, 2011). These noted differences may stem from the exposure of these particles to the fungal biomass. Identical findings have been reported by other authors, when assessing the biodegradation of polymeric materials (Jecu et al., 2012), thus highlighting the
Fig. 3. FTIR-ATR spectra for the different samples of Z. maritimum, including at the time of inoculation, grown in the presence of microplastics for 7 (F7), 14 (F14), 21 (F21) and 28 days (F28). The control sample corresponds to Z. maritimum grown for 28 days with no contact with the microplastic particles.
region. Lastly, the peak between 1200 and 1100 cm−1 is frequently attributed to the vibrations of C\\O bonds, found in carbohydrates, as well as the peak at 1080–1000 cm−1. Also noteworthy was the peak at 1100–1000 cm−1, assigned to the PO2 group present in nucleic acids (Sivan et al., 2006; Yamada-Onodera et al., 2001). As noted in Fig. 3, there was an overall decrease in the intensities of theses peaks throughout the duration of the experiment, except for the peak associated to the C\\O vibrations of carbohydrates (1200–1100 cm−1). These results suggest that there is a reduction in the lipidic and proteic content of Z. maritimum when exposed to microplastics. Moreover, it is possible to conjecture that this reduction was likely due to the reduced nutrient medium in which the fungus was grown (Weete, 2012), leading to the use of lipids as an energy source (Prasad and Ghannoum, 1996). The reduction of proteins and increase of carbohydrates can be explained by the changes caused in the fungal metabolism when exposed to a medium with a reduced carbon or nitrogen source. These induce fungi to produce proteolytic enzymes that degrade the intracellular proteins, as the deamination of amino acids provides the C skeletons required for energy purposes (McIntyre et al., 2000; Nitsche et al., 2012). These also induce nutrient recycling, mediated by glycosyl hydrolases and other enzymes that are synthesized aiming at the release of carbohydrates present in the cell wall by remodelling it, leading to an increase in the accumulation of carbohydrates (McIntyre et al., 2000; Nitsche et al., 2012; Pollack et al., 2008). Hence, for the control sample (Z. maritimum grown in minimum medium without microplastics), the lack of an alternative carbon source resulted in an endogenous search for energy. This ultimately led to the substantial differences observed in the FTIR spectra, which are made evident by the calculated corresponding areas, shown in Table 2. The overall behaviour seen in the spectra can therefore be attributed to the lack of nutrients in the medium, which in the case of the control fungus, Z. maritimum not exposed to microplastics, lead to a search for endogenous sources of carbon, and to a variation in the spectrum more significant. In the case of the Z. maritimum exposed to
Table 2 Relative areas for the different regions of the FTIR-ATR spectra (in triplicate) of Z. maritimum, grown in the presence of microplastics for 7 (F7), 14 (F14), 21 (F21) and 28 days (F28). The control sample corresponds to Z. maritimum grown for 28 days with no contact with the microplastic particles. Region (cm−1)
Inoculum
F7
F14
F21
F28
Control
3700–3000 3000–2800 1800–1700 1700–1500 1500–1250 1200–1100 1100–1000
64 ± 2 16.5 ± 0.8 1.2 ± 0.5 20.79 ± 0.04 17 ± 1 5±1 23 ± 2
21 ± 2 4.1 ± 0.7 0.9 ± 0.8 10.3 ± 0.4 6.3 ± 0.2 9.0 ± 0.4 9.6 ± 0.2
23.3 ± 0.3 2.4 ± 0.2 0.2 ± 0.2 9.1 ± 0.1 3.6 ± 0.6 7.5 ± 0.6 13.4 ± 0.8
34 ± 3 7±1 0.2 ± 0.1 12.9 ± 0.3 7.7 ± 0.6 8.5 ± 0.1 13 ± 1
24 ± 2 2.74 ± 0.09 0.3 ± 0.4 10 ± 1 5.0 ± 0.7 7.8 ± 0.5 10.4 ± 0.6
28.6 ± 0.9 4.8 ± 0.5 0.6 ± 0.4 10.7 ± 0.9 7.5 ± 0.3 7±1 15 ± 2
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Fig. 6. Infrared spectra in the region 600–4000 cm−1 of microplastics exposed to Z. maritimum for different periods of time. The observed differences in the spectra highlight the different degrees of degradation.
increasing vibrations. Such observations are further corroborated by the calculated areas for the aforementioned regions, shown in Table 3, clearly highlighting the chemical modifications detected through FTIRATR spectroscopy. 4. Conclusions
Fig. 4. In A), 1H NMR spectra of the lyophilized samples of Z. maritimum. In B), the corresponding relative areas from specific regions of the 1H NMR spectra obtained, corresponding to different classes of biomolecules (see text), are shown.
biological action of such organisms over these materials and their resulting biodegradation potential. The particles recovered from the samples grown for 14, 21 and 28 days were subsequently analysed by FTIR-ATR spectroscopy and the results are shown in Fig. 6. As evidenced in Fig. 6, there was a gradual increase in band intensities for peaks at 3700–3000 cm−1, caused by the hydroperoxide and hydroxyl groups. Similarly, areas in the region of 1700–1500 cm−1 and 1200–950 cm−1, caused by carbonyl groups and by double bonds, respectively, also gradually increased. These may result from different oxidation reactions with, for example, the functional groups present in the polymer (Barbeş et al., 2014; Ołdak et al., 2005), thus yielding the noted
This study underlines the high potential of Z. maritimum for the biodegradation of (micro)plastics, thus responding to the current and urgent need of alternative routes to minimize and abate the presence of these materials in aquatic systems and, more specifically, in the oceans. This organism was evidenced to grow in a minimum growth medium in the presence of microplastic particles, which, based on the observed modifications, point to their use as substrate by Z. maritimum. This assertion is based on the mass variations, as well as on the FTIR and NMR results. These support such conclusion, as the noted reduction in lipid and protein content, accompanied by an increase in carbohydrate concentration occurred to a larger extent in the fungus not exposed to microplastics. Due to the natural occurrence of this organism in maritime coastal waters, this study underlines the high potential of this fungus in future bioremediation strategies aiming at curtailing the increasing presence of (micro)plastics in the environment. Further studies are required in order to evaluate the likely mechanisms at the core of this demonstrated capacity of Z. maritimum to use polyethylene as a substrate and these are already underway. Acknowledgements This work was supported by national funds through FCT/MEC (PIDDAC) under project IF/00407/2013/CP1162/CT0023. Thanks are
Fig. 5. Optical (A) and electron (B) microscopy images of particles exposed to Z. maritimum for 28 days. Despite less evident in A, electron microscopy results (B) highlight the marked differences between the microplastics before and after exposure to Z. maritimum, which show pronounced surface irregularities and also clearly indicate the presence of biological material at the pellets' surface.
A. Paço et al. / Science of the Total Environment 586 (2017) 10–15 Table 3 Relative area from the identified regions from the FTIR-ATR spectra for the recovered microplastic particles (in triplicate). Region (cm−1)
PE
PE_14Days
PE_21Days
PE_28Days
3700–3000 3000–2800 1750–1500 1500–1400 1400–1300 1200–1000 750–650
1±1 24.1 ± 0.2 0.2 ± 0.3 3.8 ± 0.5 0.3 ± 0.2 0.3 ± 0.2 2.5 ± 0.3
15 ± 2 10.1 ± 0.2 5.5 ± 0.2 3.07 ± 0.08 1.7 ± 0.5 5.0 ± 0.3 2.5 ± 0.4
20 ± 2 22.3 ± 0.2 5.5 ± 0.5 4.71 ± 0.07 1.6 ± 0.5 7.8 ± 0.1 3.8 ± 0.4
57 ± 2 15 ± 1 1±1 4.6 ± 0.1 2.5 ± 0.8 6±2 3.4 ± 0.5
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