Biodegradation of nonylphenol polyethoxylates by litter-basidiomycetous fungi

Journal of Environmental Chemical Engineering 7 (2019) 103316

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Biodegradation of nonylphenol polyethoxylates by litter-basidiomycetous fungi

T

⁎

Julieta Mallermana, , Raúl Itriaa,b, Paola Babayc, Mario Saparratd,e, Laura Levina a

Laboratorio de Micología Experimental, Departamento de Biodiversidad y Biología Experimental, Facultad de Ciencias Exactas y Naturales, Universidad de Buenos Aires, InMiBo UBA-CONICET, 1428, Buenos, Aires, Argentina b Instituto Nacional de Tecnología Industrial, Avenida General Paz 5445, 1650, San Martín, Buenos Aires, Argentina c Gerencia Química, Centro Atómico Constituyentes, Comisión Nacional de Energía Atómica, Av. General Paz 1499, 1650, San Martín, Buenos Aires, Argentina d Instituto de Fisiología Vegetal (INFIVE), UNLP, CCT-La Plata-CONICET, Diag. 113 y 61, CC 327, 1900, La Plata, Argentina e Instituto de Botánica Spegazzini, Facultad de Ciencias Naturales y Museo, UNLP, 53 # 477, 1900, La Plata, Argentina

A R T I C LE I N FO

A B S T R A C T

Keywords: Biodegradation Surfactant Litter-decomposing fungi Extracellular enzymes Laccase Manganese peroxidase

Nonylphenol polyethoxylates (NPnEOs) are widely used surfactants whose degradation products are a matter of concern due to their greater persistence in the environment, toxicity and endocrine-disrupting effects in wildlife and humans. The saprotrophics Gymnopus luxurians and Hypholoma fasciculare, and the ectomycorrhizal Xerocomellus chrysenteron were selected among nineteen litter-decomposing fungi due to their capacity to tolerate up to 10 g L−1 of NP10EO. While X. chrysenteron was unable to remove NP10EO when amended to agar medium at 1 g L-1, G. luxurians and H. fasciculare were efficient degraders, reaching 37.5 ± 3.1% and 74.4 ± 4.4% of elimination after 15 days, respectively. Under solid-state fermentation using Ligustrum lucidum leaf-litter as substrate, G. luxurians and H. fasciculare removed correspondingly 71.3 ± 3.8% and 96.3 ± 1.4% of the surfactant after 90 days. Fungal degradation ability was related to the secreted titers of the ligninolytic enzymes laccase and manganese peroxidase. Degradation pathway involved the elimination of the shorter homologues (n ≤ 7) while carboxylated products were not detected, consequently potentially toxic metabolites did not accumulate. Therefore, these litter-basidiomycetous fungi showed as promising tools for detoxifying nonylphenol polyethoxylates and other related chemical compounds with endocrine disrupting activity (such as nonylphenol).

1. Introduction Nonylphenol polyethoxylates (NPnEOs) are among the most extensively employed non-ionic surfactants [1]. Their main applications include the manufacture of detergents and cleaners, cosmetics, spermicides, paints and their use as dispersing or emulsifying agents in pesticides and herbicides [2]. Most of them are introduced into the environment as aqueous solutions through industrial and domestic effluents and discharged to surface waters either directly or after waste treatment in sewage treatment plants. Many technologies have been applied in order to degrade or remove these contaminants from wastewater, among them membrane treatment using biological (membrane bioreactors) or physical processes (i.e. nanofiltration), biotechnological-based methods (biofilms, immobilized enzymes, etc.), adsorptionoriented and advanced oxidation processes (i.e. photocatalysis),

combined methods are interesting as well, since they benefit from synergistic effects [3,4]. Biological transformation of NPnEOs has been shown to occur in both, natural ecosystems and wastewater treatment plants, though their elimination is not complete. Degradation products like alkylphenol diethoxylate (NP2EO), alkylphenol monoethoxylate (NP1EO) and nonylphenol (NP) are recalcitrant to further microbial attack, and due to their hydrophobic character they accumulate in sediments, groundwater, and sewage sludge [5,6]. These compounds are receiving increasing attention because of their demonstrated aquatic toxicity and harmful effects on biota, such as endocrine disruption in several species [7,8] even at low concentrations. Because of their estrogenic effects, ubiquitous nature, toxicity and persistence, the use of NPnEOs has been banned in Europe [9] and Canada [10]; however, their use continues worldwide due to their low cost and good performance in a broad number of applications, in

Abbreviations: LDF, litter-decomposing fungi; MnP, manganese peroxidase; NP, nonylphenol; NpnEOs, nonylphenol polyethoxylates; SSF, solid-state fermentation ⁎ Corresponding author. Present address: Instituto de Investigaciones Fisiológicas y Ecológicas Vinculadas a la Agricultura (IFEVA), Facultad de Agronomía, Universidad de Buenos Aires, CONICET, 21 Av. San Martín 4453, Buenos Aires, C1417DSE, Argentina. E-mail address: [email protected] (J. Mallerman). https://doi.org/10.1016/j.jece.2019.103316 Received 3 April 2019; Received in revised form 13 July 2019; Accepted 23 July 2019 Available online 25 July 2019 2213-3437/ © 2019 Elsevier Ltd. All rights reserved.

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(NPnEO with an average number of 10 ethoxy units, Proquimia S.A., Chile) were used as analytic standards. Solvents were HPLC-grade and all other chemicals were reagent-grade. For solid-phase extraction (SPE), glass columns and PTFE frits were from Merck; Octadecylsilica sorbent was CEC-18 (UCT, USA).

addition to the lack of regulations in most countries. Hence, the search for organisms capable of reducing their concentration in the environment is of great relevance [11]. Bioremediation using ligninolytic fungi can be a valuable alternative approach with respect to conventional physical and chemical methods [12] or can be suitable in combination with other wastewater cleaning techniques such as membrane bioreactors [4]. White rot fungi (WRF) secrete extracellular enzymes to decompose plant cell wall polymers as carbon and energy sources. Among them, ligninolytic enzymes with low substrate specificity [e.g. laccase and manganese peroxidase (MnP)] are a distinctive feature of saprotrophic basidiomycetes, that allow them to degrade the more recalcitrant fractions of organic matter, and an ample array of organopollutants with structural resemblance to lignin [13]. Intracellular cytochrome P-450 monooxygenase proved to be involved in fungal degradation of several xenobiotics including NP as well [14]. Removal of NP has been recently documented using the WRF Phanerochaete chrysosporium, Phanerochaete sordida, Pycnoporus sanguineus, Pleurotus ostreatus, Trametes versicolor, Bjerkandera sp., Coriolopsis polyzona and Coriolopsis gallica [15–19]. NP degradation was determined using crude extracts [17] or partly or totally purified laccase [15,20] and MnP [15,21]. Studies using laccase of C. gallica and C. polyzona [17,18] described the formation of dimeric and higher transformation NP products lacking estrogenic-activity. Tsutsumi et al. [15] suggested similar transformation products following oxidation with MnP of P. chrysosporium. Syed et al. [22] proposed another transformation mechanism involving intracellular monooxygenases from P. chrysosporium: the oxidation of alkylphenols including NP at the terminal alkyl chain carbon followed by the removal of the terminal carbons via the β-oxidation pathway. The possible entry into the basal metabolism is also sustained by Dubroca et al. [23], who observed mineralization of NPs by a culture of T. versicolor using 14C-labelled. Though the ability to degrade NP was also demonstrated in some ligninolytic non-wood rotting fungi, particularly in two aquatic ascomycetes [24], and another isolate from soil [20], the exploration of other ecophysiological groups of fungi is limited. In this sense, there is scarce data available about the degradation of NPnEOs or other surfactants by litter-decomposing fungi (LDF). Like WRF, many LDF secrete laccase and MnP, and demonstrated capacity of degrading several xenobiotics such as polycyclic aromatic hydrocarbons (PAHs), polychlorinated biphenyls (PCBs), pesticides and herbicides [25,26]. Recently the capability of the LDF Stropharia rugosoannulata to degrade alkylphenol and oxyethylated alkylphenol representatives was demonstrated. Degradation was accompanied by laccase and MnP production, suggesting their involvement in the process [27]. The ability of LDF to colonize and survive in soil competing with other microorganisms, can make them even more appropriate for bioremediation applications compared with WRF, which have poor capability to grow in soil. Even so, a restricted number of species has been evaluated so far [13]. Therefore, the aim of this work was to evaluate the ability of selected LDF to tolerate and degrade NP10EO, the most broadly used form of NPnEOs. Furthermore, we investigated a possible association between NP10EO degradation and oxidative laccase and MnP enzyme production. The degradative ability of the selected fungi was studied in agar cultures and under solid-state fermentation (SSF) using Ligustrum lucidum leaf-litter as substrate. The present research would contribute to establish new potential strategies for degrading and detoxifying NPnEOs as well as to clarify the mechanisms involved in their degradation by fungi.

2.2. Fungal isolates and identification Fungal strains were collected from litter samples from Delta of the Paraná River and Platense Riverside region (Argentina) during 2013 and 2014. Either mycelium from the inner part of the basidiome, or in the case of small ones a spore printing, were transferred under sterile conditions onto agar plates containing 20 mL of malt extract agar medium (MEA: glucose 10, malt extract 12.7 and agar 20 g L−1), and incubated at 25 °C. Some strains from the same area were provided by BAFCcult (Culture collection of Natural and Exact Sciences Faculty at the University of Buenos Aires). Inoculum consisted of one core of agar (9 mm diameter) obtained from the border of a 7-day old agar slant culture. Stock cultures were preserved on MEA at 4 °C in BAFCcult. Taxonomic identification of the litter-decomposing specimens was performed through the study of the morphological characteristics of the basidiomes and confirmed by sequencing the internal transcribed spacer (ITS) region of the ribosomal DNA. For this, total DNA was extracted from fresh mycelia using the Microbial DNA Kit (MoBio, USA) and then used for PCR amplification (Bio-Rad Thermal Cycler, USA) with the fungal-specific primer pair ITS1/ITS4 [28]. When mycelium could not be isolated from the agarized medium, a fragment of the basidiome was used for amplification. The PCR products were analyzed by electrophoresis in 1.5% agarose gel and sent to the sequencing facility (University of Buenos Aires). Sequences of the isolates were compared using BLAST with homologous sequences in NCBI-GenBank database (www.ncbi.nlm.nih.gov) and deposited under accessions numbers listed in Table 1.

2.3. Culture conditions for the screening of NP10EO tolerant strains and ligninolytic enzyme activities Fungal strains were inoculated in 9-cm Petri dishes containing 20 mL of GA agarized medium (glucose 10 g, asparagine 4 g, agar 20 g, MgSO4·7H2O 0.5 g, KH2PO4 0.5 g, K2HPO4 0.6 g, CuSO4·5H2O 0.4 g, MnCl2·4H2O 0.09 mg, H3BO3 0.07 mg, NaMoO4·2H2O 0.02 mg, FeCl3 1 mg, ZnCl2·H2O 2.5 mg, biotin 0.005 mg, thiamine hydrochloride 0.1 mg, and distilled water up to 1 L) supplemented with 1 g L−1 NP10EO. Every 2 days, the diameter of the fungal colonies was measured and growth rates were calculated from the linear phase range (mm day-1). Control cultures in GA plates were used to evaluate the effect of the surfactant on fungal growth. For estimating laccase and MnP activities, MEA medium was supplemented respectively with 2,6dimethoxyphenol (DMP) [29] or MnCl2.4H2O 1 mM [30], and color changes due to DMP or manganese oxidation were registered. Enzyme efficiency (E) was calculated as the ratio between the activity zone and colony diameter. All experiments were performed in duplicate.

2.4. Tolerance towards increasing concentrations of NP10EO Fungi that showed the utmost radial growth rate in GA plates amended with 1 g L−1 NP10EO were inoculated in GA plates containing different concentrations of the surfactant (0.01, 0.1, 0.5, 1, 5 and 10 g L−1), and incubated at 25 °C for 15 days. Growth rates were calculated as described above. Non-inoculated plates served to evaluate abiotic removal. The effective concentration 50 (EC50): the concentration that caused nearly 50% reduction in growth, was determined.

2. Materials and methods 2.1. Chemicals Technical grade NP (85% 4-NP, Fluka, Switzerland), a technical mixture of 4-NP1EO and 4-NP2EO (Igepal CO-210, 99.5% min. active, with an average number of 1.5 ethoxy units, Aldrich, USA) and NP10EO 2

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Table 1 Growth rates and in-vitro production of ligninolytic enzymes on MEA medium supplemented with specific substrates: 2,6-dimethoxyphenol to detect laccase activity, MnCl2.4H2O to detect MnP; enzyme efficiencies (E = radial growth /enzyme halo) of LDF strains in the different media assayed. Fungal taxa selected from this screening are highlighted in grey.

The values are the mean of two replications ± standard deviation. Different letters in superscript indicate significant differences among fungal taxa (within columns) from DGC post hoc comparisons (P < 0.001). * Strains requested to the BAFCc. ** [61].

volumes were of 50 μL. A binary gradient elution was performed for the simultaneous measurement of both short and longer oligomers. Mobile phases were solvent A: hexanes (95% n-hexane - 5% branched hexanes) and solvent B: 2-propanol. The program started isocratically with 96% A - 4% B for 15 min, with a linear gradient through 50% A–50% B in 27 min; there was finally 6 min of isocratic hold with 100% B. Flow rate was 1.0 mL min-1. Compounds with less than 3 ethylene oxide units were identified with standard spikes of t-NP, 4-NP1EO and 4-NP2EO, while the higher ethoxymers were assigned from their elution sequence in NP10EO. Complementarily, NP10EO conversion by cultures of Hypholoma fasciculare (the strain that achieved the higher degradation percentages) was evaluated by Liquid chromatography - Mass spectrometry (LC–MS). LC–MS system consisted of a Shimadzu LCMS 2020 instrument, equipped with an ESI source and a single quadrupole mass spectrometer, using a C18 reverse phase column (Gemini NX3 μm 100 x 4 mm) at 50 °C and isocratic conditions with water: acetonitrile 99: 1, as mobile phase, at a flow rate of 1.2 mL min-1 with flow derivation to allow an entrance of 0.4 mL min-1 to the ESI. Oxidation products were identified by comparing the retention times, mass spectra and UV-spectra with standards and with literature data.

2.5. Biodegradation of NP10EO by selected fungal strains To estimate NP10EO degradation, 9-mm agar plugs from GA plates amended with different NP10EO concentrations were cut out from the mycelial growing zone at day 15 of incubation and placed in 2 mL glass test tubes. Residual NP10EO was extracted by adding 0.5 mL of ethyl acetate (EtAc) and sonicating for 30 min at room temperature. Two aliquots of 0.5 mL of EtAc were used to wash the agar plug, and were collected in the same tube. The extracts were evaporated to dryness under N2 and re-dissolved in the High performance liquid chromatography (HPLC) elution mixture for analysis. Sterile controls were redissolved with different elution volumes to achieve a unique final concentration of 30 mg L−1. Performance of the extraction procedures with non-inoculated control samples was evaluated through recovery assays. Composition of oligomers was confirmed to be uniformly extracted for every NP10EO concentration. Residual NP10EO was calculated as the percentage of the chromatographic peak area resulting from fungal inoculated samples in comparison with the non-inoculated control. 2.6. Chemical analysis of degradation products

2.7. Enzyme assays The HPLC system used consisted of a Spectra SERIES P200 binary pump (Thermo Separation Products, CA, USA). Detection was performed with a Linear LC-305 fluorescence detector (Linear Instruments, NE, USA) at 230 nm excitation wavelength and 300 nm emission wavelength. Data were acquired and analyzed with the Konikrom 5.2 software (Konik Instruments, Barcelona, Spain). Column heater Eppendorf CH-30 and controller Eppendorf TC-50 (Alltech, IL, USA) were used to control column temperature within 0.1 °C. Standard solutions of t-NP, Igepal and NP10EO for HPLC were prepared by dilution of 1 g L−1 stock solutions of the individual compounds with hexane: 2PrOH (96:4) for t-NP, 2 g L-1 for Igepal and 10 g L−1 for NP10EO surfactant. All solutions were stored at 4 °C. Separations were performed on a 5 μm particle aminopropyl silica column (250 x 4.6 mm) with 10 x 4.0 mm guard-column (Pinnacle II, Restek, USA) at 35 °C. Injecting

For ligninolytic enzyme determination a 9-mm agar plug with mycelium or an appropriately diluted enzyme solution was added to the reaction buffer containing the enzyme substrate. Laccase (EC 1.10.3.2) activity was determined spectrophotometrically measuring the increase at 469 nm due to the oxidation of 5 Mm DMP in 0.1 M Na-acetate buffer (pH 3.6) at 30 °C (ε469 = 27.5 mM−1 cm−1) [31]. Absorbance was monitored for 2 min under continuous vortex agitation. MnP activity (E.C.1.11.1.13) was measured by the reaction product at 610 nm (ε610 = 22 mM−1 cm-1) obtained from phenol red oxidation. The reaction mixture contained 0.05 M succinate buffer pH 4.5, 0.1 mM MnSO4, 0.1 mM H2O2 and 0.01% phenol red [32]. Reactions were halted by adding NaOH 5 N after 10 min of continuous stirring at 30 °C, centrifuged for 10 min and increase in A610 was measured. One unit of 3

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enzyme activity (U) was defined as the quantity of enzyme required to oxidize 1 μmol of substrate per min in assay conditions.

level. Total NP10EO degradation was analyzed with a two-way ANOVA with fungal strain and NP10EO concentration as main factors.

2.8. Fungal cultures on litter under SSF conditions

3. Results and discussion

Ligustrum lucidum senescent leaves obtained in the studied area, were air-dried at room temperature and stored at 4 °C until utilize. 2.5 g of leaves cut into pieces (size > 20 mm) were placed in 125 mL Erlenmeyer flasks, and moistened either with 20 mL of distilled water, or water with 1 g L−1 NP10EO. The flasks were autoclaved for 20 min at 120 °C, cooled and inoculated with 2 agar mycelial plugs of 9-mm diameter, obtained from the border of a vigorously growing colony. Cultures were incubated at 25 °C in darkness for 3 months and samples were taken every 30 days. Non-inoculated flasks functioned as controls and to estimate water loss during incubation. Sterile water was replenished when sampling. Per treatment, triplicates were conducted. A water soluble fraction (WSF) was collected at each sampling time, from both non-treated and fungal transformed litter, according to Saparrat et al. [33]. For this, 20 mL of distilled water was added to each flask, mixing it afterwards on a rotary shaker at 150 rpm for 1 h. The content was subsequently filtered through gauze and centrifuged at 5000 × g per 10 min. Supernatants were used as enzyme sources and for pH measurements. Litter solid fraction (SF) was dried at 80 °C and stored at 4 °C before use. Finally, surfactant extraction was carried out in each of the fractions and the amount of residual NP10EO was assessed by thin layer chromatography.

3.1. Screening of tolerant LDF strains Nineteen basidiomycetous fungi were isolated and identified from Delta of Paraná River and Platense Riverside region, in order to evaluate their potential for NP10EO degradation and their ligninolytic ability (Table 1). Among them, 18 were saprotrophic basidiomycetes belonging to the order Agaricales, members of 6 different families, while one strain namely Xerocomellus chrysenteron, was an ectomycorrhizal fungus from the order Boletales. This last basidiomycete was incorporated because it is related to the litter environment and although it is not considered an efficient lignin degrader, proved its ability to mineralize DDT [35]. All 19 litter fungal strains tested in the present study were significantly affected by NP10EO 1 g L−1 addition to the culture medium. Half of the strains were incapable of growing or developed very slowly in GA supplemented plates. X. chrysenteron showed the highest radial growth rate (2.99 mm day−1), followed by H. fasciculare, Coprinus comatus, Gymnopus luxurians and Marasmiellus candidus. Fungal growth in GA-NP10EO significantly correlated with fungal growth in a nutritive common medium such as MEA (data not shown; r = 0.54; P = 0.0003), and therefore the 5 strains selected for further studies, were those with a general vigorous growth. While almost all isolates (except for X. chrysenteron) gave positive for laccase activity, only 60% of them revealed MnP activity. Laccase production was inversely associated with fungal growth in the same medium (r = -0.46; P = 0.0029), thus strains with higher growth rates were less efficient in enzyme production per mycelium unit. On the contrary, MnP production was independent of the fungal growth in the same medium (P = 0.73). Inhibition in growth by the addition of the alkylphenol polyethoxylate was accompanied by macroscopic changes in the morphology of the colonies (data not shown), probably in response to this stressful environment. Mycelial cords formation (an aggregation of hyphae parallel orientated), observed in H. fasciculare and G. luxurians, was reported to allow better interconnection of the mycelium providing alternative transport routes with a consequent increased resilience to damage [36]. X. chrysenteron manifested changes in colony pigmentation, from intense violet to cream with violet tints. Some fungal species have been reported to produce pigmented metabolites associated with a greater tolerance for the survival under chemical stress [37,38].

2.9. Thin layer chromatography (TLC) Ten μL of sample solutions were spotted onto pre-activated normalphase plates coated with silica gel 60 F254 on aluminum sheets (Merck, Germany). The mobile phase solvent system used for the detection of surfactant homologs was a mixture of EtAc, glacial acetic acid and water (70:16:15 v/v). The sample migration distance was 10 cm. After development, the plate was completely dried using an air drier and spots were revealed by spraying it with modified Burger reagent [34]. Images were captured electronically by a desktop scanner and processed with the ImageJ 1.45 s analysis program (NHI, National Institutes of Health, USA). Densitometric scanning was performed and the area of resolved peaks was recorded. In addition, retention factor (Rf) values were calculated for each spot. Degradation of NP10EO was calculated by integrating and adding the areas corresponding to the different ethoxymers in one sample and comparing them with the noninoculated control (100% NP10EO). Serial dilutions were made from stock solution of standard NP10EO in order to calculate the mass limit of detection on TLC plates that is, the minimum mass of a substance that can be detected in a sample, which was found at 2 μg. A calibration curve was performed with the standard NP10EO (0–30 μg), and the total area of the different ethoxymers was plotted as a function of the mass of NP10EO. The curve was adjusted to a linear regression, showing a determination coefficient (R2) of 0.9999.

3.2. Tolerance to increasing concentrations of the surfactant Strains selected from the screening were cultivated in GA media supplemented with different NP10EO concentrations. EC50 values are depicted in Table 2. H. fasciculare, G. luxurians and X. chrysenteron were able to grow throughout the whole concentration range. X. chrysenteron exhibited an abrupt reduction in mycelial growth up to concentration of 0.1 g L−1 NP10EO, and no major changes were detected from this point (P < 0.05).

2.10. Statistical analysis One-way analysis of variance (ANOVA) was used to evaluate differences in fungal growth rates in GA-NP10EO 1 g L−1 among LDF. Mean values were compared with a DGC post-hoc test with 5% of significant level. Pearson’s correlation coefficient (r) was calculated to test linear relationships between growth in GA-NP10EO 1 g L−1 medium and growth in MEA. In tolerance tests, dose-response curves were constructed by plotting the growth rates against the logarithmic concentration of NP10EO. The EC50 values were calculated through a nonlineal regression using the “log (inhibitor) vs. normalized response – variable slope” model included in GraphPad Prism version 5.01 (GraphPad Software, La Jolla, CA, USA). EC50 among strains were compared with a one-way ANOVA and a Tukey’s test at 5% significant

Table 2 Tolerance levels of the selected fungal strains to the toxicant (EC50). Fungal strain

EC50

R2

IC 95%

C. comatus G. luxurians H. fasciculare M. candidus X. chrysenteron

0.024cd 0.046cd 0.086bc 0.016d 0.583a

0.991 0.934 0.964 0.986 0.658

0.021 0.026 0.058 0.014 0.144

to to to to to

0.028 0.079 0.125 0.019 2.365

Different superscript letters indicate significant differences from Tukey´s Test comparisons (P < 0.0001). The values are the mean of two replications. 4

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ethoxylated counterparts [44]. The endpoint of the degradation is nonylphenol. In effluent treatment plants, 50–99% removal efficiency has been reported for NPnEOs [5]. Strains tested in this work showed substantial removal of NP10EO, even at high concentrations such as 10 g L−1, proving the degradative potential of these fungi. The capacity of NP degradation by WRF such as T. versicolor and Bjerkandera sp., has been already demonstrated, removing most of NP 0.1 g L−1 (97%) after 25 days [16]. Cabana et al. [17] reported more than 95% of NP 0.005 g L−1 removal after 4 hs of incubation with an enzyme preparation from C. polyzona. As an example, chromatograms obtained under the tested experimental conditions with H. fasciculare are depicted in Fig. 2. The profile of the NP10EO standard solution showed a Poisson distribution with maximum intensity for the homologues NP8EO and NP9EO and a bilateral decrease in the intensity for the rest of the homologues. Only the areas of the oligomers with n ≤ 13 were integrated because at higher retention times the resolution became diffuse. The degradation results showed that all NP ethoxylates were biologically degradable. Furthermore, a differential removal of oligomers was observed as a function of NP10EO concentration and fungal strains employed (Fig. 3). At lower NP10EO concentrations, especially with H. fasciculare and G. luxurians that exhibited high percentages of degradation, ethoxymers with n ≤ 7 were completely degraded, ethoxymers with an intermediate lengh 8 < n < 10 were depleted with respect to standard NP10EO, and the proportion of longer ethoxymers with n ≥ 10 increased. At higher NP10EO concentrations, relative ethoxymer composition was similar to that of standard NP10EO. Therefore, in the present study as a result from NP10EO fungal transformation, at any of the concentrations and surfactant exposure times tested, alkylphenol metabolites (NP and NP1EO-NP3EO) accumulation was not detected. Conversely, a trend towards the elimination of short homologs was observed (Fig. 3). Additionally, while analyzing H. fasciculare NP10EO degradation products by LC–MS, toxic nonylphenol ethoxycarboxylates (NP1EC and NP2EC) as a possible alternative pathway of surfactant conversion, were not detect (data not shown). Thus, NP10EO degradation by LDF involves a mechanism that differs from the one usually proposed during aerobic and anaerobic wastewater treatment of alkylphenol ethoxylates [45]. This is an important issue as alkylphenols intermediates are more toxic and persistent than parent compounds and usually accumulate not only in the sedimentable fractions of treatment plants but also in treated effluents [5,43]. NP10EO 60 mg L−1 biodegradation in aerobic systems of activated sludge resulted in an accumulation of the metabolites NP1EO, NP2EO and NP3EO [46]. Hayashi et al. [47] reported that while NP10EO was the only oligomer adhered initially to the system, 48% NP2EO and 18% of NP3EO were detected, after 25 days of culture with an inoculum coming from an effluent. Ethoxymers composition resulting from fungal degradation varied as a function of the NP10EO concentration employed (Fig. 3). At lower concentrations, short ethoxylates (n ≤ 6) were more easily removed even though they were not the major compounds in the initial NP10EO mixture, whereas there was an enrichment in larger homologues (n > 9). This selectivity in degradation diminished as surfactant concentration increased, showing the initial distributions profiles of the different homologues and those resulting from fungal degradation, scarce differences. This behavior may be due to the differential hydrophobicity of the homologues, with lower lipophilicity as the number of ethoxy groups increases [48]. Fungal cell wall surface has a high number of potential binding sites exhibited by free carboxyl, amino, hydroxyl, phosphate and mercapto groups [49], and thus provides a matrix for the interaction, mainly with the shorter ethoxymers, through the formation of hydrophobic complexes, in a mechanism called biosorption. A schematic representation of the possible mechanisms involved in nonylphenol polyethoxylates removal by litter-basidiomycetous is depicted in Fig. 4. The same effect was reported previously when testing the bioaccumulation and biodegradation of a mixture of NPnEOs (n = 1–12) by the microalga Chlorella vulgaris [50]. The higher biosorption of short homologues may have

H. fasciculare and G. luxurians showed a staggered inhibition (Fig. S1). Maximal EC50 values were obtained with X. chrysenteron, although the R2 was very low. In order to reduce the number of strains selected for degradation studies, both C. comatus and M. candidus were discarded due to their greater sensitivity to the surfactant, which was revealed by the lowest EC50 registered and complete growth inhibition above 5 g L−1 (Fig. S1). There is no data available for NP10EO EC50 in fungi or other organisms. Existing records report EC50 for nonylphenol, the most toxic metabolite, resulting from natural biological degradation of NP10EO. To compare our toxicity results with those of other studies, we used the toxic equivalence factor (TEF) between the NP and NPnEOS, assuming in 0.005 the factor for polyethoxylates with n ≥ 9, meaning that the latter would be about 200 times less toxic than NP [39]. Karley et al. [40] informed that NP had negative effects on filamentous fungi at 1.3–6.2 mg L-1 concentrations, via multiple physiological effects as respiration dissociation, whereas Kollmann et al. [41] stated that NP had no negative effects on fungi at environmental concentrations (0.08–1.45 mg L-1). In this work, EC50 for NP10EO were respectively 583, 86 and 46 mg L-1 for X. chrysenteron, H. fasciculare and G. luxurians, and therefore negative effects might be expected, at least in some specimens of this functional group of fungi growing next to urban sites. Toxicity data in which nonylphenol polyethoxylates were used, were reported in Killi fish with an EC50 value of 12 mg L-1 for NP9EO, and an LC50 of 6.6 mg L-1 in the fathead minnow Pimephales promelas after 96 h [39]. 3.3. Biodegradation of NP10EO in agarized medium Both G. luxurians and H. fasciculare were efficient in NP10EO degradation after 15 days of culture in Petri dishes (Fig. 1). In contrast, X. chrysenteron was the most tolerant strain but was not able to remove it (Figs. S1 and 1). The highest degradation was obtained with H. fasciculare, respectively 88.8 ± 1.76%, 77.7 ± 1.67% and 74.4 ± 4.43% removal at 0.1, 0.5 g L−1 and 1 g L−1 NP10EO. Although degradation efficiency decreased at increasing surfactant concentrations, a considerable removal was attained even at 10 g L−1 (38.3 ± 7.9%). G. luxurians exhibited a mean contaminant removal of 31.8% with no significant differences in degradation ability at the whole concentration range evaluated. It is well established that alkylphenol ethoxylates parent compounds present a rapid biodegradation however, NPnEOs are more slowly degraded in sediments under anaerobic conditions than in the presence of oxygen [42]. NPnEOs degradation carried out by different phylogenetic groups proceeds via separate pathways in oxic and anoxic conditions, as demonstrated in different tests using water from rivers, sediments, effluents and soils [42,43]. Aerobic sequences generate carboxylated NPs while anaerobic ones produce the

Fig. 1. Biodegradation of NP10EO by selected LDF: G. luxurians ( ), H. fas) and X. chrysenteron ( ). Data are the mean of two replicates, ciculare ( one standard deviation of the mean is represented. 5

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Fig. 2. Chromatograms of standard NP10EO (black line), and those obtained after 15-day H. fasciculare cultivation in GA-NP10EO at the indicated concentrations. The labels above the peaks indicate the number (n) of ethoxy groups in the corresponding NPnEO oligomer.

Fig. 3. Relative ethoxymer composition (%) after 15 day cultivation in GA-NP10EO at concentrations: 0.5 g L−1 (a), 1 g L−1 (b), 5 g L−1 (c) and 10 g L−1 (d) with ). Data are the mean of two selected fungal strains G. luxurians ( ), H. fasciculare ( ) and X chrysenteron ( ). Non-inoculated control composition is shown in ( replicates, one standard deviation of the mean is represented.

3.4. Enzyme activities

favored their interaction with ligninolytic enzymes attached to the cell wall or secreted extracellularly, but with limited diffusibility in agar medium [51], which might be involved in surfactant degradation. On the other hand, especially for non-ionic detergents, water solubility significantly influences the adsorption degree [52], thus the more polar longer homologs could have had a greater tendency to remain dissolved in the aqueous agar medium. However, in the presence of high surfactant concentrations, a larger amount of long polyethoxylates spatially close to fungal hyphae, could sterically forbid the degradative action of ligninolytic enzymes on shorter ethoxymers.

Extracellular enzyme activities laccase and MnP secreted by the selected fungi were analyzed at day 15 of the cultivation period (Fig. 5). Both H. fasciculare and G. luxurians were able to produce laccase, and enzyme titers depend on NP10EO concentrations employed. For both strains, from 0.5 g L−1 of NP10EO (P < 0.05) laccase production was significantly higher than the obtained in the control without surfactant. Activity was maximal with 1 g L−1 of NP10EO (1.1 U mL-1 in H. fasciculare, 1.2 U mL-1 in G. luxurians) corresponding respectively to 3- and 25-fold increase in the activity recorded in the controls. Higher 6

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Fig. 4. Possible mechanisms involved in nonylphenol polyethoxylates removal by litter-basidiomycetous fungi.

Fig. 5. Laccase (a) and MnP (b) activities attained when culturing G. luxurians ( ), H. fasciculare ( ) and X. chrysenteron ( NP10EO concentrations. Data are the mean of three replicates, one standard deviation of the mean is represented.

) in GA plates with different

these fungi or their enzymatic systems for bioremediation purposes [53]. Some studies showed enhanced ligninolytic production with nonylphenol as well. Mougin et al. [54] detected a 14-fold rise in T. versicolor laccase activity in a medium containing 0.5 mM NP (≡ 0.11 g L-1). Soares et al. [16] found higher laccase production with NP 0.1 g L-1 (3.6-fold increase); maximal laccase and MnP activities were detected in coincidence with the peak of alkylphenol removal. Total removal of NP was achieved with a purified laccase from a soil-isolated fungus [20], but NP removal was also promoted by ABTS [2,2′-azino-bis (3-ethylbenzothiazoline-6-sulphonic acid)] [17] or 1-hydroxybenzotriazole (HBT) [15] addition to the enzyme preparation, suggesting the involvement of a laccase/mediator system in the process. On the other hand, laccase or MnP activities were not detected in X. chrysenteron cultures; probably mechanisms not involving the

concentrations of the surfactant did not exert greater titers. MnP activity was very low in G. luxurians with an average value of 0.013 ± 0.001 U mL-1, without significant differences between cultures with and without surfactant addition (P < 0.05). In contrast, MnP production by H. fasciculare was highest with 0.5–1 g L-1 of NP10EO (P < 0.05) (0.157 ± 0.019 U mL-1), equivalent to a 20-fold rise in activity in comparison with the medium without surfactant. In concordance, highest removal percentages were attained by H. fasciculare within this range of surfactant concentrations, proving the cooperative action of MnP and laccase in NP10EO removal. At higher NP10EO concentrations, MnP activity was not appreciable, yielding both G. luxurians and H. fasciculare comparable removal. Increased ligninolytic production by WRF and LDF in the presence of xenobiotics has been widely reported and is an attribute that supports the use of 7

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alkylphenol polyethoxylate contaminated water and soils.

surfactant transformation such as an increased antioxidant defense [55,56] allow it to tolerate NP10EO toxicity.

Declaration of Competing Interest 3.5. Fungal cultures on litter under SSF conditions None. To have a more realistic scenario, the ability of G. luxurians and H. fasciculare to degrade NP10EO, was measured in SSF culture conditions using L. lucidum litter. For this, NP10EO content was evaluated by TLC in water (WSF) and solid (SF) fractions of the fungal cultures under SSF. TLC proved to be a useful analytical technique for rapidly and semiquantitatively assessing the ethoxylates pattern composing a commercial mixture of NPnEOs as well as for monitoring its primary degradation [57]. Only 52.1 ± 8.2% of the initially added mass of NP10EO was detected in the SF of the control treatments. Taking into account that the extraction efficiency of nonylphenol polyethoxylates in the SSF system was 50.3 ± 2.2% and that traces of surfactant ethoxymers could not be detected in any of the WSF assayed, probably the detergent adsorbed almost completely to the litter used. When comparing the remaining NP10EO in the solid fraction of SSF cultures inoculated with H. fasciculare with respect to non-inoculated cultures, this isolate was able to remove 51.7 ± 22.1% and 96.3 ± 1.4%, respectively after 30 and 90 days of incubation. G. luxurians degraded 60.0 ± 0.1% and 71.3 ± 3.8% of the surfactant, respectively after 30 and 90 days (Fig. S2). Despite the considerable degradation obtained in SSF cultures, laccase and MnP titers detected in the WSF were very low in both fungi, with no significant differences due to surfactant addition. Maximal activities were detected at the first month of incubation, respectively 0.49 ± 0.29 U g−1 of laccase and 0.04 ± 0.02 U g-1 of MnP in G. luxurians and 0.13 ± 0.21 U g-1 of laccase and 0.06 ± 0.01 U g-1 of MnP in H. fasciculare. pH affects microbial growth and enzyme activity, nevertheless both fungal strains only slightly acidified L. lucidum litter throughout the whole incubation period. Ending pH of the non-inoculated control was 4.93 ± 0.11, while G. luxurians and H. fasciculare cultures displayed a final pH of respectively 4.69 ± 0.26 and 4.38 ± 0.23. It has been reported that the natural adsorption of NP to sediments is the main mechanism of removal of this type of xenobiotics [58]. The use of a complex matrix such as L. ligustrum senescent leaves revealed that NP10EO was recovered mainly from this inert substrate (SF), not being available in the aqueous associated soluble fraction (WSF). This is consistent with the high octanol/water partition coefficient (log Kow) calculated for NP (4.48) and, to a lesser extent, for the NPnEOs (1 ≤ n ≤ 9), with values between 4.3 and 3.59 [59,60] expressing the affinity of the mentioned compounds for the organic matter present in soils or sediments. Therefore, the litter system alone could be useful for NP10EO surfactant immobilization, but ligninolytic fungi naturally growing on these readily available materials, such as H. fasciculare and G. luxurians, increased its degradation. NP10EO 1 g L−1 (≡ 3.8 g Kg−1 of dry leaf litter) removal by these fungi growing in SSF with L. lucidum leaf litter as substrate, was greater than 50% after 30 days and around 70 and 95% with G. luxurians and H. fasciculare respectively after 90 days of incubation (Fig. S2). T. versicolor growing on various substrates removed almost 100% of 4-n-nonylphenol after 20 days through a combination of adsorption and biodegradation; and even in the absence of fungus, the methodology adopted achieved a noteworthy contaminant removal [12].

Acknowledgements JM was awarded with a scholarship from CONICET. MCNS and LNL are members of the Research Career CONICET (National Research Council, Argentina). This study was supported by CONICET (PIP11220120100408), Agencia Nacional de Promoción Científica y Tecnológica (PICT2015-1620) and University of Buenos Aires, Argentina (UBACyT20020150200018). Appendix A. Supplementary data Supplementary material related to this article can be found, in the online version, at doi:https://doi.org/10.1016/j.jece.2019.103316. References [1] United States Environmental Protection Agency, Nonylphenol (NP) and Nonylphenol Ethoxylates (NPEs) [RIN 2070-ZA09], (2010). [2] I.-H. Acir, K. Guenther, Endocrine-disrupting metabolites of alkylphenol ethoxylates – a critical review of analytical methods, environmental occurrences, toxicity, and regulation, Sci. Total Environ. 635 (2018) 1530–1546. [3] A. Priac, N. Morin-Crini, C. Druart, S. Gavoille, C. Bradu, C. Lagarrigue, G. Torri, P. Winterton, G. Crini, Alkylphenol and alkylphenol polyethoxylates in water and wastewater: a review of options for their elimination, Arab. J. Chem. 10 (2017) S3749–S3773. [4] A. Sepehri, M.H. Sarrafzadeh, Effect of nitrifiers community on fouling mitigation and nitrification efficiency in a membrane bioreactor, Chem. Eng. Process.-Process Intensif. 128 (2018) 10–18. [5] R.J. Maguire, Review of the persistence of nonylphenol and nonylphenol ethoxylates in aquatic environments, Water Qual. Res. J. Canada 34 (1999) 37–78. [6] T.Y. Chiu, N. Paterakis, E. Cartmell, M.D. Scrimshaw, J.N. Lester, A critical review of the formation of mono- and dicarboxylated metabolic intermediates of alkylphenol polyethoxylates during wastewater treatment and their environmental significance, Crit. Rev. Environ. Sci. Technol. 40 (2010) 199–238. [7] G.G. Ying, B. Williams, R. Kookana, Environmental fate of alkylphenols and alkylphenol ethoxylates- a review, Environ. Int. 28 (2002) 215–226. [8] A. Soares, B. Guieysse, B. Jefferson, E. Cartmell, J.N. Lester, Nonylphenol in the environment: a critical review on occurrence, fate, toxicity and treatment in wastewaters, Environ. Int. 34 (2008) 1033–1049. [9] European Parliament and the Council, Directive 2003/53/EC of the European Parliament and of the Council of 18 June 2003 amending for the 26th time Council Directive 76/769/EEC relating to restrictions on the marketing and use of certain dangerous substances and preparations (nonylphenol, nonylphenol ethoxylate and cement), Off. J. Eur. Union (2003) 24–26. [10] Gazette, Canadian Environmental Protection Act, Proposed Notice Requiring the Preparation and Implementation of Pollution Prevention Plans in Respect of Nonylphenol and Its Ethoxylates Contained in Products, Minister of public works and government services, 2003. [11] P.A. Babay, R.F. Itria, E.E. Romero Ale, E.T. Becquart, E.A. Gautier, Ubiquity of endocrine disruptors nonylphenol and its mono- and di-ethoxylates in freshwater, sediments, and biosolids associated with high and low density populations of Buenos Aires, Argentina, Clean - Soil, Air, Water 42 (2014) 731–737. [12] G. Castellana, E. Loffredo, Simultaneous removal of endocrine disruptors from a wastewater using white rot fungi and various adsorbents, Water Air Soil Pollut. 225 (2014) 1872–1885. [13] A. Anastasi, V. Tigini, G.C. Varese, The bioremediation potential of different ecophysiological groups of fungi, in: E.M. Goltapeh, Y.R. Danesh, A. Varma (Eds.), Fungi as Bioremediators. Soil Biology, 32, Springer, New York, 2013, pp. 29–49. [14] V. Subramanian, J.S. Yadav, Role of P450 monooxygenases in the degradation of the endocrine disrupting chemical nonylphenol by the white rot fungus Phanerochaete chrysosporium, Appl. Environ. Microbiol. 75 (2009) 5570–5580. [15] Y. Tsutsumi, T. Haneda, T. Nishida, Removal of estrogenic activities of bisphenol A and nonylphenol by oxidative enzymes from lignin-degrading basidiomycetes, Chemosphere 42 (2001) 271–276. [16] A. Soares, K. Jonasson, E. Terrazas, B. Guieysse, B. Mattiasson, The ability of whiterot fungi to degrade the endocrine-disrupting compound nonylphenol, Appl. Microbiol. Biotechnol. 66 (2005) 719–725. [17] H. Cabana, J.L.H. Jiwan, R. Rozenberg, V. Elisashvili, M. Penninckx, S.N. Agathos, J.P. Jones, Elimination of endocrine disrupting chemicals nonylphenol and bisphenol A and personal care product ingredient triclosan using enzyme preparation from the white rot fungus Coriolopsis polyzona, Chemosphere 67 (2007) 770–778. [18] C. Torres-Duarte, V.M. Teresa, R. Vazquez-Duhalt, Laccase-mediated transformations of endocrine disrupting chemicals abolish binding affinities to estrogen

4. Conclusions In this work two strains belonging to the litter-decomposing basidiomycetes functional group (G. luxurians and H. fasciculare), demonstrated their capacity to remove NP10EO surfactant without the formation of shorter toxic metabolites (neither nonylphenol, nor the corresponding short chain ethoxylates or ethoxycarboxylates). Further research is needed in order to understand the mechanisms underlying the process and verify if fungal strains inoculated with leaf litter or their culture supernatants can be applied in the bioremediation of 8

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