Oxidative biodegradation of single-walled carbon nanotubes by partially purified lignin peroxidase from Sparassis latifolia mushroom

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Oxidative biodegradation of single-walled carbon nanotubes by partially purified lignin peroxidase from Sparassis latifolia mushroom Gayathri Chandrasekaran a, Soo-Kyung Choi a, Young-Chul Lee b, Geun-Joong Kim c, Hyun-Jae Shin a,* a b c

Department of Chemical and Biochemical Engineering, Chosun University, Gwangju 501-759, Republic of Korea Department of Biological Engineering, College of Engineering, Inha University, Incheon 402-751, Republic of Korea Department of Biological Science, College of Natural Sciences, Chonnam National University, Gwangju 500-757, Republic of Korea

A R T I C L E I N F O

A B S T R A C T

Article history: Received 10 September 2013 Accepted 8 December 2013 Available online xxx

Two types of carbon nanotubes (usually single-walled carbon nanotubes (SWCNTs) and multi-walled carbon nanotubes (MWCNTs)) have been intensively focused on academic researches and massproduced for wide applications such as composite materials, biosensors, and drug delivery systems. However, due to oxidative stress-dependent and physically-induced cellular toxicity of CNTs, many efforts to render biocompatible and biodegradable properties in CNTs have been highlighted. Thus, taking into the consideration of exposure in human health and the environment, biodegradation of CNTs as a potential disposal is highly addressed. In this study, lignin peroxidase (LiP) was isolated and partially purified from the fruiting bodies of the edible mushroom Sparassis latifolia (S. latifolia). The biodegradation of raw grade and thermally-treated carboxylated SWCNTs (denoted as ASA and AST) with this enzyme was investigated, prior to more biodegradation-resistant MWCNTs. The interactions between the SWCNTs and LiP were investigated using various techniques, and the intermediate byproducts of the LiP degradation were identified. Our findings demonstrated that both ASA and AST were efficiently degraded by LiP where the producing radicals by the LiP played a critical role in the biodegradation of SWCNTs. The final degraded products were confirmed with the generation of CO2 gas. Conclusively, the low extraction cost of partially purified enzyme from mushrooms can make this approach a promising alternative in environmental bioremediation as a practical application. ß 2013 Published by Elsevier B.V. on behalf of The Korean Society of Industrial and Engineering Chemistry.

Keywords: Single-walled carbon nanotubes Lignin peroxidase Sparassis latifolia Biodegradation Bioremediation

1. Introduction Carbon-based nanotechnology has become one of the most important and exciting aspects of research in various fields and especially in engineering and biology [1–6]. Presently, the environmental and human health concerns over engineered nanomaterials (ENMs), especially on single-walled carbon nanotubes (SWCNTs) and multi-walled carbon nanotubes (MWCNTs), are increasing [7–10], an important issue to be taken into consideration. Moreover, it was stated that carbon nanotubes (CNTs) affect the environment as much as they affect humans [11– 13], and the levels of risk are proportional to the amount of ENMs produced globally. Various reports have stated that the CNTs cause skin cancer, oxidative stress, granuloma formation, fibrosis, lung cancer, genotoxicity, and mutagenicity [12]. Moreover, the dermal toxicity

* Corresponding author. Tel.: +82 62 2307518; fax: +82 62 2307226. E-mail address: [email protected] (H.-J. Shin).

of the SWCNTs causes oxidative stress to the skin. Various types of CNT products, including raw grade and thermally-treated grades, are available; they contain iron, nickel, and yttrium, which exhibited a proven pulmonary toxicity [9,12]. Although they contain lower amounts of catalysts, the overall research has revealed that CNTs tend to cause epithelioid granulomas and interstitial inflammation in animals [14]. Recently, it has been demonstrated that CNTs can trigger some biological responses similar to those of asbestos. For example, MWCNTs (less than 20 mm) are short and soft, which could cause serious health problems with the identical mechanism [15]. As a result, such functionalized CNTs to reduce toxicity and grant biocompatible and biodegradable characteristics have been developed [16]. However, it is still stressed that the disposal of CNTs is critical issues. Concerning the environmental hazards of CNTs, those trials are reported that SWCNTs and MWCNTs could be degraded by enzyme-catalyzed oxidations within a few weeks or months [17]. Among several peroxidases which are activated by H2O2 to generate unstable radicals for degradation of carbonaceous nanomaterials, white-rot fungi (WRF) can degrade a wide range

1226-086X/$ – see front matter ß 2013 Published by Elsevier B.V. on behalf of The Korean Society of Industrial and Engineering Chemistry. http://dx.doi.org/10.1016/j.jiec.2013.12.022

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of organopollutants whose degradable ability is closely related to the lignin-degrading systems and catalyze the oxidation of nonphenolic and various organic compounds. Specifically, this fungus produces mainly two types of extracellular peroxidases, lignin peroxidases (LiP) (EC.1.11.1.14) and manganese dependent peroxidase (MnP) (EC.1.11.1.13), while others secrete laccase and versatile peroxidise [18–20]. LiP consists of a single polypeptide chain, with an iron protoporphyrin prosthetic group. It has a unique ability to degrade lignin polymer through an oxidative electron transfer mechanism. Based on properties of WRF, they were applied in the paper and pulp industries as well as in the textile industry for the decolorization of dyes and transformation of polyaromatic hydrocarbons [21]. Recently, a proteomic analysis of the fruiting bodies of this mushroom and the economic importance of lignin structures was reported [20]. As a result, the microbial degradation by lignolytic fungi has been intensively studied over the past few years due to the irregular structure of lignin; the lignolytic fungi produce extracellular enzymes with low substrate specificity that allows degradation of substrate compounds. Peroxidase enzymes can be an alternative that provides a new strategy to detoxicify organic pollutants from wastewater and soils. However, the bioremediation of carbonaceous materials by LiP extracted from this mushroom is not reported yet despite the low extraction cost of partially purified enzyme. Based on the first report on the enzymatic biodegradation of CNTs by horseradish peroxidase (HRP) [17,22], in this study, the biodegradation of two different types of carboxylated SWCNTs including both a raw grade and thermallytreated using LiP, which is partially purified from the fruiting bodies of the mushroom Sparassis latifolia (formerly S. crispa) was carried out. The findings showed that LiP could play an important role in the biodegradation of carboxylated SWCNTs. We demonstrate feasibility that LiP, partially extracted from mushroom S. latifolia, could be a promising option in large-scale settings as an economic and ecofriendly bioremediation for carbonaceous nanomaterials.

wheat flour + 10% corn syrup solution, (d) Larix leptolepis (80%) + wheat flour (20%) + 10% corn syrup solution, and (e) Larix leptolepis (80%) + wheat flour (10%) + corn pellet (10%) + 10% corn syrup solution. S. latifolia fruiting bodies samples were prepared using these media, and their LiP activities also were tested. 2.3. Enzyme extraction The fresh fruiting body of the mushroom was isolated and freeze-dried. Each frozen fresh fruiting body (50 g) of S. latifolia mushroom was ground to a fine powder in liquid nitrogen using a pre-chilled ceramic mortar pestle. Next, the mushroom powders were extracted in acetate buffer (pH 5.0) separately, phosphate buffer (pH 7.0), and Tris–HCl buffer (pH 9.0) containing 2 mM EDTA, 1 mM MgCl2, and 1 mM phenylmethylsulfonyl fluoride (PMSF) at 4 8C. The concentrations of all the buffers were 10 mM. The mixture was centrifuged at 10,000 rpm for 10 min at 4 8C. The resultant supernatants were used as crude enzymes [23]. 2.4. Protein purification The enzyme was purified from the crude supernatants by the following steps: 1) ammonium sulfate precipitation and 2) DEAE– Sepharose anion exchange chromatography. The crude extracts were precipitated by addition of ammonium sulfate to 65% saturation. The solution was then centrifuged at 8000 rpm for 15 min at 4 8C. The precipitate was dissolved in a Tris–HCl buffer (10 mM, pH 9.0) and desalted over a PD-10 desalting column in the identical buffer. The filtered enzyme solution was applied to a DEAE–anion exchange column (5 ml) that had been equilibrated with a Tris–HCl buffer (100 mM, pH 9.0) containing 0.15 M NaCl and then eluted with a linear salt gradient at a flow rate of 0.5 ml/ min. The fractions with a high LiP specific activity were collected, concentrated, and used with following studies. 2.5. Enzyme assay

2. Experimental 2.1. Materials SWCNTs purchased from Hanwha Nanotech Corporation (Incheon, Korea) were used. The product name of a raw grade of ASA-100 F is manufactured by the arc-discharge process (denoted as ASA) while that of AST-100 F is prepared by thermal treatment (denoted as AST). The S. latifolia mushroom fruiting bodies were collected from Forest Resources Research Institute (Naju, Jeonnam, Korea). Lyophilized HRP type VI (analytical grade) and hydrogen peroxide (30 wt%, analytical grade), and all the substrates including pyrogallol, 2,6-dimethoxy phenol, veratryl alcohol, 2,20 -azino-bis (3-ethylbenzothiazoline-6-sulphonic acid) (ABTS), and guaiacol and methanol, formic acid, and acetonitrile of analytical grade were purchased from Sigma–Aldrich (St. Louis, MO, USA). DEAE–Sepharose Fast Flow and PD-10 desalting columns were acquired from Amersham Biosciences (GE Healthcare, Sweden). Unless otherwise stated, all of the chemicals in this study were used as received from the supplier. Distilled deionized water was utilized through the experiments (resistance > 18 mV, DI water). 2.2. Media compositions of S. latifolia mushroom culture For the media compositions and their effects on enzyme production, five different sawdust media compositions with controlled percentages of media moisture were used such as: (a) Larix leptolepis (100%) + DI water, (b) Larix leptolepis (100%) + 10% corn syrup solution, (c) Larix leptolepis (90%) + 10%

The mixture consists of 50 mM of pyrogallol (2.5 ml), 100 mM of sodium acetate buffer pH 5.4 (16 ml), and 50 mM of H2O2 (1.5 ml). Approximately 250 ml of the mixture was aliquotted to each well in the 96-well microtiter plate; 10 ml of the enzyme sample (0.05 U/ml) was added and incubated at 37 8C for a few minutes. The appearance of a dark-color indicated the presence of peroxidase activity [24,25]. The activity of LiP was quantitatively assayed by the reported method [26] using veratryl alcohol as a substrate and monitoring the formation of veratraldehyde at 310 nm spectrophotometrically. The reaction solution (1 ml) consisted of 2 mM of veratryl alcohol and 0.4 mM of H2O2 in 50 nM of sodium tartarate buffer, pH 3, at 25 8C. The reaction was begun by adding 50 ml of the enzyme solution. The MnP and laccase were then assayed by the reported methods [27,28]. 2.6. Electrophoretic and zymographic analysis Sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS–PAGE) was performed using a 10% polyacrylamide gel [29]. A high-range molecular weight marker (EBM-1031, Elpis Biotech Inc.) was used as a standard. The proteins were quantified by the bicinchoninic acid (BCA) assay using bovine serum albumin (BSA) as a standard [30]. For the assay of peroxidase activity, a zymogram analysis was performed in the polyacrylamide gels; the protein samples were dissolved in loading buffer without SDS or thiolreducing agents and separated on a 7.5% Native-PAGE gel. After the protein separation, the gels were equilibrated for 30 min in 50 mM of sodium acetate buffer (pH 5.4) prior to their incubation with

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80 mM of pyrogallol and 80 mM of H2O2 in fresh sodium acetate buffer. 2.7. Carboxylation and its examination of SWCNTs by scanning electron microscopy (SEM) observation SWCNTs were carboxylated as described previously [31]. Briefly, approximately 25 mg of each ASA and AST was sonicated in H2SO4/H2O2 (30%) at a ratio of 3:1 for 24 h at 0 8C. After 10-15 h, 2.0 ml H2O2 was added to the reaction to replace the spent H2O2. The final dispersion was then heated at 70 8C for 10 min, and subsequently diluted 10-fold and filtered through a 0.22 mm Teflon membrane filter; the sample was washed with copious amounts of water to become a neutral pH. The carboxylated ASA and AST were subjected to an SEM (Hitachi S-2400N, Japan) analysis. This reaction is known as carboxylation, and it was confirmed through the SEM studies. 2.8. Transmission electron microscopy (TEM) measurement The samples in PBS solution were centrifuged at 3400 rpm for 3 h to remove salts from the buffer [17,22]. The supernatant was removed and the pellet was re-suspended in DMF and sonicated for 1 min. The sample was dropped on a lacey carbon grid (pacific-Grid Tech) and allowed to dry for 1 hr and subsequently take TEM images (FEI Morgagni, 80 keV or JEOL 2100F, 200 keV). 2.9. Treatment of SWCNTs with LiP and H2O2 Approximately 1 mg of each carboxylated ASA and AST was added to 4.0 ml of phosphate buffered saline (PBS) and sonicated for 1 min. As a positive control, 0.385 mg/ml of lyophilized HRP type VI was solubilized in PBS. The negative control was performed with buffer alone. The experimental samples consisted of the LiP purified from S. latifolia and solubilized in PBS at the identical concentration. Four ml of each enzyme solution was added to the vials containing the carboxylated SWCNTs and brought to a total volume of 8 ml. All the vials were then sealed with a septum and wrapped with parafilms. From the first day of incubation, 8 ml of 40 mM H2O2 was added through 20 daily additions of 250 ml by syringes to the all vials; during the 20 days, the vials were kept at 25 8C [17,22]. Also crude LiP was incubated under the identical condition except that the incubation temperature was lowered to 4 8C.

temperature in gas permeable Teflon tubing. The tubing was filled with 10 ml of sample, folded over, and placed in the ESR quartz tube with an open 3.0 mm internal diameter. The ESR solutions were prepared by incubating LiP with 0.02 mg/ml SWCNTs in PBS for 1 min at room temperature; pyrogallol was then added and the peroxidase reaction was initiated by adding H2O2 (80 mM). As a control, HRP (0.35 mM) was incubated with SWCNTs (0.02 mg/ml) in PBS for 1 min at room temperature, and ascorbate (100 mM) was then added. The ESR spectra of the radicals were recorded 1 min after addition of H2O2. The spectra of these radicals were recorded using the following conditions: 3270 G, center field; 10 G, sweep width; 10 mW, microwave power; 0.4, field modulation; 103, receiver gain; 0.1 s, time constant; 1 min, scan time [22]. 2.12. Liquid chromatography–mass spectrometry (LC–MS) Approximately 3 ml of the aqueous samples of both ASA and AST were acidified by the addition of 500 ml of 0.1 M of HCl and extracted with dichloromethane (3 ml). The dichloromethane was then removed and the products were then re-dispersed in pure MeOH (500 ml). Approximately 5 ml of the concentrated sample was injected onto a C18 column (100  2.1 mm, 1.7 mm) at 40 8C in a mobile phase of 20:80 (v/v, formic acid and water:formic acid and acetonitrile). The samples were then analyzed for positive ions using electrospray mass spectrometry. An accurate mass measurement was performed with a Synapt high-definition mass spectrometry system (HDMS; Waters Co.). Leucine enkephalin was used as an independent reference lock-mass via the LockSpray to ensure mass accuracy and reproducibility. The LC–MS profiling was performed on an Acquity UPLC system (Waters Co., Milford, MA) equipped with a binary solvent delivery system and an autosampler. The chromatographic separation was performed on an Acquity UPLC BEH [17,22]. 2.13. Gas chromatography (GC) Approximately 2 ml headspace of each biodegradation for ASA and AST sample (total headspace volume: 5 ml) was taken through the septum of the vials and injected into a Shimadzu QP5050A GCMS unit with an XFI-F capillary column. The temperature program was set to hold at 100 8C for 1 min, followed by temperature ramping at a rate of 10 8C/min until a maximum temperature of 325 8C was achieved and held for an additional 10 min.

2.10. Visible near infrared (vis-NIR) and Raman spectroscopy

3. Results and discussion

150 ml of the aqueous samples of both carboxylated SWCNTs were analyzed in 2 ml-glass cuvettes using a Lambda 900 spectrophotometer (Perkin-Elmer, Norwalk, CT). The SWCNTs were scanned from 600 to 1300 nm. Six samples as mentioned above were subjected to the vis-NIR spectroscopy. All the samples were centrifuged at 3400 rpm for 3 h to remove salts from the buffer. The precipitated samples were then treated with MeOH following a 2 min sonication. Approximately 20 ml of the samples were placed on the microscope slide and dried. All the spectra were collected on a Renishaw inVia Raman microscope using an excitation wavelength of 633 nm. The samples were scanned from 1000 to 1800 cm1 to visualize the changes in the D and G band intensities resulting from the degradation process. All the spectra were collected using a 15 s exposure period and averaged 5 scans per sample [17,22].

3.1. Identification and partial purification of LiP

2.11. Electron spin resonance (ESR) spectroscopy ESR spectra were recorded on a JES-FA ESR spectrometer (JEOL, Tokyo, Japan). The measurements were performed at room

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The crude S. latifolia mushroom fruiting bodies exhibited a peroxidase activity toward the substrate pyrogallol in the 96-wellplate. The appearance of a dark reddish-brown color in the 96well-plate indicated the presence of the peroxidase activity. Generally, there are three types of oxidoreductase enzymes in the fungal sources; therefore, we sought to screen for the enzymes types including LiP, MnP, and laccase using the spectrophotometric method. The screening results indicated that veratrylaldehyde was formed by LiP activity. Other enzymes such as MnP and laccase were not found. The relative specificity of LiP toward the substrates was ranked in the order of pyrogallol > veratryl alcohol > ABTS > guaiacol (Supplementary data, Table S1). S. latifolia LiP was partially purified by ammonium sulfate precipitation followed by DEAE column chromatography (Fig. 1a). Furthermore, the peroxidase protein of the 65% fraction was purified by anion exchange chromatography from the other proteins present. The unbound fraction (Fig. 1b lane 1) demonstrates the absence of the LiP band, but two major peaks were

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Fig. 1. SDS-PAGE (a) M, standard protein marker; lane 1, 40% ammonium sulfate precipitate; lane 2, 65% ammonium sulfate precipitate. SDS-PAGE (b) M, standard protein marker; lane 1, DEAE unbound fraction; lane 2, DEAE 1st bound fraction; lane 3, DEAE 2nd bound fraction. Native PAGE (7.5%) zymography (c) of peroxidase from S. latifolia.

observed within the bound fractions. The first peak did not exhibit LiP activity (Fig. 1b lane 2), while the second peak showed the presence of LiP that was confirmed by the enzyme assay (Fig. 1b lane 3). Therefore, the DEAE fractions were used in the characterization and application studies without further purification. The zymographic analysis strongly suggested that the protein retained its peroxidase activity (Fig. 1c). Furthermore, the lower molecular weight protein at 26 kDa was purified and subjected it to the enzyme activity assay; there was no LiP enzyme activity present (see Supplementary data, Fig. S1). The LiP of S. latifolia was purified from its fruiting bodies. SDS– PAGE revealed that the molecular mass of the LiP was approximately 45 kDa (Fig. 1b). Some fungal LiP, including LiP isoenzyme 41 with a 44 kDa enzyme from Irpex lacteus and a 45 kDa from Phanerochaete sordida YK-624, have been reported with a molecular mass similar to that of the S. latifolia LiP [32,33].

Furthermore, TEM analyses were performed to compare the degradation of the sample during a 20 day incubation with LiP and H2O2 (Fig. 2b–d): the non-degraded AST on the initial day, after 10 days, the degradation of the carboxylated AST by LiP with H2O2; this degradation is indicated by the reduced lengths of the SWCNTs. When compared with the initial stage, the 10 day and 20 day samples showed the presence of non-tubular structures, indicating the degradation of the SWCNTs. However, it was unable to discern complete oxidation at day 20, which would be indicated by the presence of the intermediates of carbonaceous products. To observe the longevity of the enzyme, a microtiter plate assay was performed using pyrogallol as a substrate for the LiP. This assay was performed for each sample after the 20 day incubation, and the results showed that enzyme activity was nearly retained (Supplementary data, Table S1). 3.3. Visible near infrared and Raman spectra

3.2. Enzymatic degradation of SWCNTs SWCNTs treatment imparts carboxylic acid groups and improves the CNTs dispersion in aqueous solution [31] and removes the residual metal catalyst. The SEM analysis (Supplementary data, Fig. S2) clearly shows both ASA and AST were carboxylated, and observed that the metal impurities are significantly reduced after the acid treatment, compared to the pristine SWCNTs. After the acid treatment, the SWCNTs are not only cut into short pipes but also purified because the acid mixture is known to intercalate and exfoliate graphite. As-prepared shorten and carboxylated SWCNTs were used throughout the study. The photograph demonstrating the enzyme degradation is shown (Fig. 2a). The vial (1) contained the carboxylated AST without degradation at day 1, while vial (2) is a positive control containing HRP, vial (3) contains the AST with S. latifolia LiP. The vials (2) and (3) exhibited the noticeable color change that indicates degradation after 20 days incubation. This result confirmed that both enzymes degrade the AST. In regard to the degradation of the pristine AST, the visual observation indicated that complete oxidation did not occur, corresponding to TEM image (Supplementary data, Fig. S3). Because the degradation rate was considerably lower than that of the carboxylated SWCNTs. Thus the degradation studies of that material have not further pursued. In addition, pristine and carboxylated ASA showed a similar trend, compared to AST cases (data not shown).

The visible near infrared (vis-NIR) and Raman spectroscopic methods can be used to monitor the degradation of CNTs [17]. The degradation of ASA and AST was monitored using a vis-NIR spectroscopy after a 5 days incubation with the partially purified LiP and 40 ml H2O2 (aq). Non-degraded carboxylated SWCNTs are mixtures of various diameters and helicities, exhibiting metallic and semiconducting electronic properties (Fig. 3). It shows the spectral range of the M1 metallic band between 650 and 750 nm, as well as the broad S2 semiconducting band of the CNTs absorbing between 1000 and 1100 nm [34]. By monitoring these bands over the 5 days incubation period with partially purified LiP and H2O2 at 25 8C, the M1 and S2 band of the CNTs decreased. The graphitic structure of the CNTs diminished and completely disappeared as a result of the enzymatic degradation during the 10 days incubation period (Fig. 3a and b). Also, observed results on the 20th day confirmed this phenomenon, indicating similar results for both SWCNTs. Furthermore, when the incubation of the SWCNTs with crude LiP and H2O2 took place at 4 8C, the degradation process slowed and required 60 days (Supplementary data, Fig. S4). This result indicated that at 25 8C, the degradation was faster than at 4 8C. The results are similar to those of the HRP reaction that serve as a control. Temperature has a considerable effect on the ability of the HRP to degrade CNTs [17]. Because approximately 60 days at a lower temperature are required to achieve the same level of degradation, this demonstrates that CNTs degradation was slower compared with the degradation at 25 8C.

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Fig. 2. Photograph (a) of the enzymatic degradation of the carboxylated AST: vial 1, carboxylated AST; vial 2, after 10 days of incubation of carboxylated AST with HRP; vial 3, after 10 days of incubation of carboxylated AST with S. latifolia LiP. TEM micrographs (b–d) confirming the degradation of the carboxylated AST according to incubation time (1, 10, and 20 days), scale bars = 200 nm.

The degradation of SWCNTs was also confirmed by Raman spectroscopy, and the results showed that at 4 8C, the carboxylated AST display D and G bands. After 60 days incubation with crude LiP, these bands were decreased, indicating the degradation of the

SWCNTs (Fig. 4). The D and G bands disappeared after the incubation with HRP. When the samples were incubated with LiP, however, the bands did not completely disappear; due to variations in the samples, some fluctuations in the D:G ratios

Fig. 3. Vis-NIR spectra of carboxylated ASA (a) degraded by HRP and S. latifolia LiP after 5, 10, and 20 days at 25 8C and carboxylated AST (b) degraded by HRP and S. latifolia LiP after 5, 10, and 20 days at 25 8C.

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Fig. 4. Raman spectra of carboxylated AST after 60 days incubation at 25 8C with S. latifolia crude LiP (black), HRP (red), and buffer alone (green). (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

were observed that were consistent with another study [35]. However, this result indicated the LiP reduced the D and G bands. It is indicated that the degradation does occur but rate is low, compared to the partially purified samples (data not shown). Clearly, the crude LiP was sufficient to initiate and promote the degradation process. The results are consistent with other reported studies for the enzymatic degradation by crude MnP of polyaromatic hydrocarbons [36]. 3.4. ESR spectra ESR measurement of the LiP and HRP activity in the presence of the SWCNTs was used to detect the radicals formed during the oneelectron oxidation of pyrogallol and ascorbate compounds by LiP and HRP, respectively. The addition of H2O2 to LiP in the presence of pyrogallol produced the characteristic ESR signals of a pyrogallol oxygen radical (pyrogallol-O*). The free radical spectrum is characterized by a signal at g = 2.00 (Fig. 5a, 1 and 2). The signal of the ascorbate radical was detected upon the incubation of HRP with ascorbate in the presence of H2O2 (Fig. 5b, 1 and 2). The addition of the SWCNTs to the incubation mixture did not change the ESR signals of the ascorbate and pyrogallol radical (#2 in Fig. 5a and b). In the absence of LiP and H2O2 or HRP and H2O2, the free radical signal was several-fold lower, thus confirming that the oxidation of pyrogallol and ascorbate occurred mainly via the peroxidase reaction (Fig. 5a and b, 3 and 4). The spectra were identical in the presence and in the absence of SWCNTs. LiP produced free radical signals from the substrate in the presence of H2O2 similarly to that produced by HRP. Therefore, both enzymes produced strong signals from the respective peroxidase substrates, thus demonstrating that CNTs did not inactivate the enzymes. Other studies reported that ESR signals showed that cation radicals were produced during the degradation of organopollutants, which proves the cation radical oxidizes these compounds [37]. The catalytic cycle of LiP is similar to that of other peroxidase wherein ferric enzyme is first oxidized by H2O2 to generate the twoelectron oxidized intermediate, Compound I [38]. The degradation mechanism occurs through a classical compound I spectrum, similar to that described for other peroxidases and indicative of the formation of a porphyrin p cation radical [39]. Generally, Compound I contains two oxidizing equivalents; one is stored in the enzyme as an oxoferryl moiety [Fe (IV) = 0]2+ and the other is OH radicals. It is interesting that in all the peroxidases and other heme-containing proteins that react with H2O2, the radical is a porbital delocalized porphyrin radical. Moreover, there is a report

Fig. 5. The ESR spectra to characterize the peroxidase activity of HRP (a) and S. latifolia LiP (b) in the presence and absence of SWCNTs. In (a), the radicals produced by the HRP with ascorbate as a substrate. (1) HRP and H2O2; (2) SWCNTs, HRP, and H2O2; (3) SWCNTs and H2O2; (4) SWCNTs and HRP. In (b), the radicals produced by the S. latifolia LiP with pyrogallol as a substrate. (1) LiP and H2O2; (2) SWCNTs, LiP, and H2O2; (3) SWCNTs and H2O2; (4) SWCNTs and LiP.

that porphyrins physisorb onto SWCNTs providing close proximal contact with the iron site, which further promotes degradation [17]. The enzymatic breakdown of the raw grade and thermally treated SWCNTs is proposed to occur through a mechanism of action that includes the generation of an aryl cation radical by reaction with H2O2. It has been proven that the biodegradation of SWCNTs is induced by free radicals which aid in the oxidation of SWCNTs [40]. Conclusively, the interaction of LiP with H2O2 molecules produces free radicals. Those unstable radicals could degrade SWCNTs non-selectively. Alternatively, there is a report on the induction of extracellular OH radicals production in WRF through quinone redox cycling. This mechanism can be explained by the production of OH radicals from the generation of Fe2+ and H2O2 (i.e., Fenton reaction). Like the WRF, S. latifolia as a brown rot fungi (BRF) [41] can also able to produce OH radicals during the depolymerization of cellulose. However, the production of reactive oxygen species (ROS) in the extracellular environment in both WRF and BRF is evidence of quinine redox cycling. This was explained as a simple strategy for the WRF to induce extracellular OH radicals production [42,43]. 3.5. LS–MS analysis To identify the final products of the degradation of the ASA and AST by LiP and HRP, LC–MS was performed, with monitoring for negative ions with a positive detector (Supplementary data, Fig. S5). The results revealed that the major degraded products of both ASA and AST are acidic by-products and aldehydes. For example, in this study we found salicylic acid as the degradation product; this identification was also confirmed later with an authentic standard. Thus, these compounds are within a class of compounds similar to

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the identified biodegradation products for the SWCNTs by HRP. However, the entire spectra of the products might be likely impossible to identify or interpret. As shown in Supplementary data, Fig. S5a, the mass to charge (m/z) value, 136.93, was observed for the LiP-degraded SWCNTs, indicative of salicylic acid. The product identified was similar to those observed in the studies of the bioremediation of polyaromatic hydrocarbons (PAHs) [21]. The molecular masses of the degraded products were very similar in the studies of the HRP- and LiP-treated CNTs. This indicated that the enzymes perform a similar cleavage of the CNTs. However, it is important to note that mass spectrometry cannot distinguish between molecular ions and fragments, and the preparative scale experiments involving product separation still must be performed. Furthermore, the LiP purified from Gloeophyllum sepiarium MTCC-1170, which degrades coal humic acid, and the mechanism involves the generation of an aryl cation radical during the reaction with H2O2 [44]. It has been proven that P. chrysosporium oxidizes PAH compounds through a radical mechanism. HRP also degrades the PAHs through cation radical intermediates. The first enzyme to attack lignin-type compounds was the LiP isolated from P. chrysosporium. This lignolytic capacity makes most taxa of fungi for use in bioremediation. Pollutants such as chlorophenols, nitrophenols, and polyaromatic hydrocarbons can be transformed by the lignolytic enzymes due to the free radical reactions [8]. 3.6. GC analysis GC was used to analyze the final degradation products of the SWCNTs. Furthermore, the evolution of CO2 gas in the sample headspace on the10th day of incubation was monitored and also compared with the degradation process of the carboxylated ASA and AST incubated with LiP and H2O2 or with HRP and H2O2 (data not shown). Both ASA and AST in the presence of H2O2 did not produce any significant concentration of CO2 in the headspace over the course of 10 days. In contrast, when carboxylated ASA and AST were incubated with LiP and H2O2 and HRP and H2O2, CO2 was measured in the headspace, which was evidence of the degradation of the SWCNTs. In particular, it was observed that the AST produced more CO2 gas than the ASA. The ASA incubated with HRP and H2O2 exhibited a higher CO2 gas concentration, compared to those incubated with the LiP and H2O2. The analysis of products indicates that complete degradation produces CO2 gas. A previous study proposed that the lignolytic fungus Pleurotus ostreatus degrades phenanthrene, producing intermediate products such as 9,10-phenanthrenequinone and 2,2-diphenic acid before forming CO2 gas [39]. Therefore, the results were agreed with the other findings and proposed that most of the fungal degradation products for the organopollutants are oxidized to carboxylic acids and finally to form CO2 gas [45]. 4. Conclusions The crude fruiting bodies of S. latifolia can potentially be used in bioremediations for carbon-based nanomaterials. S. latifolia crude extracts could be a highly efficient and low cost source of the materials for environmental remediation. Furthermore, the results are consistent with other studies that show crude purification of the enzyme is sufficient to degrade organic pollutants. For example, crude MnP was sufficient to initiate and promote the degradation of anthracene and pyrene [46]. Similarly, another study reports that crude LiP from WRF P. chrysosporium can degrade pharmaceutically active compounds such as carbamazepine and diclofenac [47]. In summary, LiP isolated from the S. latifolia mushroom catalyzed the oxidation of a raw grade and thermally-treated

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SWCNTs. This study proposes that LiP employs a mechanism based on the formation of cation radicals for its enzymatic action. Thus, the partially purified enzyme from the S. latifolia mushroom can economically perform the bioremediation and especially on biodegradation of CNTs [48] in which the partially purified enzymes with a low concentration of H2O2 facilitate the oxidation. Acknowledgements This study was carried out with the support of ‘Forest Science & Technology Projects (Project No. 2009-project-02)’ provided by Korea Forest Service. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at doi:10.1016/j.jiec.2013.12.022. References [1] L. Yan, F. Zhao, S. Li, Z. Hu, Y. Zhao, Nanoscale 3 (2011) 362. [2] Y. Lin, S. Taylor, H. Li, K.A.S. Fernando, L. Qu, W. Wang, L. Gu, B. Zhou, Y.-P. Sun, Journal of Materials Chemistry 14 (2004) 527. [3] N. Ajami, N.B. Panah, Journal of Nanostructure in Chemistry 3 (2013) 76. [4] H. Khani, O. Moradi, Journal of Nanostructure in Chemistry 3 (2013) 73. [5] F. Najafi, M. Norouzi, K. Zare, A. Fakhri, Journal of Nanostructure in Chemistry 3 (2013) 60. [6] F. Taleshi, A.A. Hosseini, Journal of Nanostructure in Chemistry 3 (2012) 4. [7] A.A. Shvedova, V.E. Kagan, Journal of Internal Medicine 267 (2009) 106. [8] I.I. Vlasova, T.V. Vakhrusheva, A.V. Sokolov, V.A. Kostevich, A.A. Gusev, S.A. Gusev, V.I. Melnikova, A.S. Lovach, Toxicology and Applied Pharmacology 264 (2012) 131. [9] A. Kunzmann, B. Andersson, T. Thurnherr, H. Krug, A. Scheynius, B. Fadeel, Biochimica et Biophysica Acta-General Subjects 1810 (2011) 361. [10] J. Russier, C. Me´nard-Moyon, E. Venturelli, E. Gravel, G. Marcolongo, M. Meneghetti, E. Doris, A. Bianco, Nanoscale 3 (2011) 893. [11] V.E. Kagan, N.V. Konduru, W. Feng, B.L. Allen, J. Conroy, Y. Volkov, I.I. Vlasova, N.A. Belikova, N. Yanamala, A. Kapralov, Y.Y. Tyurina, J. Shi, E.R. Kisin, A.R. Murray, J. Franks, D. Stolz, P. Gou, J. Klein-Seetharaman, B. Fadeel, A. Star, A.A. Shvedova, Nature Nanotechnology 5 (2010) 354. [12] A.A. Shvedova, A. Pietroiusti, B. Fadeel, V.E. Kagan, Toxicology and Applied Pharmacology 261 (2012) 121. [13] G.P. Kotchey, J.A. Gaugler, A.A. Kapralov, V.E. Kagan, A. Star, Journal of Materials Chemistry B 1 (2013) 302. [14] D.B. Warheit, Carbon 44 (2006) 1064. [15] V.C. Sanchez, J.R. Pietruska, N.R. Miselis, R.H. Hurt, A.B. Kane, WIREs Nanomedicine and Nanobiotechnology 1 (2009) 511. [16] A. Bianco, K. Kostarelos, M. Prato, Chemical Communications 47 (2011) 10182. [17] B.L. Allen, G.P. Kotchey, Y. Chen, N.V.K. Yanamala, J. Klein-Seetharaman, V.E. Kagan, A. Star, Journal of the American Chemical Society 131 (2009) 17194. [18] P. Baldrian, C. Wiesche, J. Gabriel, F. Nerud, F. Zadrazil, Applied and Environmental Microbiology 10 (2000) 2471. [19] N. Mehboob, M.J. Asad, M. Imran, M. Gulfraz, F.H. Wattoo, S.H. Hadri, M. Asghar, African Journal of Biotechnology 10 (2011) 9880. [20] K. Horie, R. Rakwal, M. Hirano, J. Shibato, H.W. Nam, Y.S. Kim, Y. Kouzuma, G.K. Agrawal, Y. Masuo, M. Yonekura, Journal of Proteome Research 7 (2008) 1819. [21] S.M. Bamforth, I. Singleton, Journal of Chemical Technology and Biotechnology 80 (2005) 723. [22] B.L. Allen, P.D. Kichambare, P. Gou, I.I. Vlasova, A.A. Kapralov, N. Konduru, V.E. Kagan, A. Star, Nano Letters 8 (2008) 3899. [23] G. Chandrasekaran, G.-J. Kim, H-J. Shin, Food Chemistry 124 (2011) 1376. [24] S. Johri, U. Jamwal, S. Rasool, A. Kumar, V. Verma, G.N. Qazi, Plant Science 169 (2005) 1014. [25] K. Periasamy, K. Natarajan, Indian Journal of Biotechnology 3 (2004) 577. [26] M. Tien, T.K. Kirk, Proceedings of the National Academy of Sciences of the United States of America 81 (1984) 2280. [27] A. Pasczyn´ski, V.-B. Huynh, R. Crawford, FEMS Microbiology Letters 29 (1985) 37. [28] M. Mansur, M.E. Arias, J.L. Copa-Patin˜o, M. Fla¨rdh, A.E. Gonza´lez, Mycologia 95 (2003) 1013. [29] U.K. Laemmli, Nature 227 (1970) 680. [30] P.K. Smith, R.I. Krohn, G.T. Hermanson, A.K. Mallia, F.H. Gartner, M.D. Provenzano, E.K. Fujimoto, N.M. Goeke, B.J. Olson, D.C. Klenk, Analytical Biochemistry 150 (1985) 76. [31] Z. Wei, M. Kondratenko, L.H. Dao, D.F. Perepichka, Journal of the American Chemical Society 128 (2006) 3134. [32] N. Rothschild, C. Novotny, V. Sasek, C.G. Dosoretz, Enzyme and Microbial Technology 31 (2002) 627. [33] H. Hirofumi, M. Sugiura, S. Kawai, T. Nishida, FEMS Microbiology Letters 246 (2005) 19.

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