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Effects of graphene oxides on soil enzyme activity and microbial biomass
Science of the Total Environment 514 (2015) 307–313
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Science of the Total Environment journal homepage: www.elsevier.com/locate/scitotenv
Effects of graphene oxides on soil enzyme activity and microbial biomass Haegeun Chung a,⁎, Min Ji Kim b, Kwanyoung Ko a, Jae Hyeuk Kim a, Hyun-ah Kwon a, Inpyo Hong a, Nari Park a, Seung-Wook Lee b, Woong Kim b a b
Department of Environmental Engineering, Konkuk University, Seoul 143-701, Republic of Korea Department of Materials Science and Engineering, Korea University, Seoul 136-713, Republic of Korea
H I G H L I G H T S • • • • •
The effects of graphene oxide (GO) on soil microbial activity was studied via a 59-day incubation experiment. Up to 1 mg GO g− 1 soil was applied and the soil enzyme activities and microbial biomass were measured. Soil enzyme activity was lowered by 15–50% under 0.5–1 mg GO g− 1 soil, but the effect subsided afterwards. Soil microbial biomass showed little change in response to GO treatment. GO can negatively affect soil enzyme activity in short term upon entrance to soils.
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Article history: Received 22 October 2014 Received in revised form 24 January 2015 Accepted 24 January 2015 Available online xxxx Editor: D. Barcelo Keywords: Nanomaterial Microbial biomass Soil incubation Extracellular enzymes Soil microorganisms
a b s t r a c t Due to recent developments in nanotechnology, nanomaterials (NMs) such as graphene oxide (GO) may enter the soil environment with mostly unknown consequences. We investigated the effects of GO on soil microbial activity in a 59-day soil incubation study. For this, high-purity GO was prepared and characterized. Soils were treated with up to 1 mg GO g−1 soil, and the changes in the activities of 1,4-β-glucosidase, cellobiohydrolase, xylosidase, 1,4-β-N-acetyl glucosaminidase, and phosphatase and microbial biomass were determined. 0.5–1 mg GO g−1 soil lowered the activity of xylosidase, 1,4-β-N-acetyl glucosaminidase, and phosphatase by up to 50% when compared to that in the control soils up to 21 days of incubation. Microbial biomass in soils treated with GO was not significantly different from that in control soils throughout the incubation period, and the soil enzyme activity and microbial biomass were not significantly correlated in this study. Our results indicate that soil enzyme activity can be lowered by the entry of GO into soils in short term but it can be recovered afterwards. © 2015 Elsevier B.V. All rights reserved.
1. Introduction Graphene oxides (GOs) are layered graphene sheets that have oxygen-containing functional groups including epoxide, carboxyl, carbonyl, and hydroxyl group (Lerf et al., 1998). Among a wide range of nanomaterials (NMs) that are being manufactured, GO or chemically modified GO are considered to be highly promising novel material due to their superior electrical characteristics, colloidal property, and large surface area (Geim and Novoselov, 2007). These excellent material properties allow GO to be used in various applications such as energystorage materials, paper-like materials, and bioenvironmental materials (Park and Ruoff, 2009; Wang et al., 2011; Zhao et al., 2012). For example, adsorbents and photocatalysts that are based on GO are employed in removing pollutants from the environment (Zhao et al., 2012). For ⁎ Corresponding author. E-mail address: [email protected] (H. Chung).
http://dx.doi.org/10.1016/j.scitotenv.2015.01.077 0048-9697/© 2015 Elsevier B.V. All rights reserved.
GO to be successfully applied in such diverse fields, it is important to determine its fate, distribution, and potential environmental impacts (Anjum et al., 2013). Previous studies have determined the fate and toxicity of other carbon-based NMs including carbon nanotubes (CNTs) and fullerenes in the soil environment (Avanasi et al., 2014; Li et al., 2013a, 2013b; Navarro et al., 2013), but few studies have investigated the impacts of GO on the soil environment. Compared with other carbon-based NMs, GO showed higher mobility in sand, which suggests that highly mobile GO may increase the environmental risks once they enter the soil environment (Qi et al., 2014). However, how GO may impact soil microbial activity that is important for nutrient cycling in soil ecosystems remains to be investigated. It has been shown via culture studies using model microorganisms such as Escherichia coli (E. coli) that GO has strong antimicrobial effect (Akhavan and Ghaderi, 2010; Hu et al., 2010; Liu et al., 2011). For example, a study using GO nanowalls showed that direct contact of bacteria with GO nanowalls can lead to cell damage; after 1 h of bacteria-GO
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nanowall contact, only 41% survived. This cytotoxicity was attributed to the sharp edges of the nanowalls (Akhavan and Ghaderi, 2010). In another study, it was shown that after 2-h contact of bacteria with GO nanosheets, the cell metabolic activity was decreased to approximately 70% and 13% at 0.02 mg ml− 1 and 0.085 mg ml− 1, respectively. The bacterial cell membranes were shown to have been severely damaged and the cytoplasm was flowing out (Hu et al., 2010). Additionally, GO treated on bacteria at 0.005–0.08 mg ml−1 for up to 4 h showed timeand concentration-dependent antibacterial activity (Liu et al., 2011). Antimicrobial effects of GO on microorganisms other than model microorganisms, such as some plant pathogens, have also been shown (Chen et al., 2013; Wang et al., 2014). GO showed 94% cytotoxicity against phytopathogenic bacteria causing infections in rice even at a low concentration. This high antimicrobial efficiency was ascribed to the extremely sharp edges of GO and generation of reactive oxygen species (ROS) (Chen et al., 2013). The antifungal activity of GO was also shown for two important plant pathogenic fungi (Wang et al., 2014). While the antimicrobial effects of GO have been demonstrated in culture studies, only a few studies determined the effects of GO on microbial communities inhabiting environmental samples (Ahmed and Rodrigues, 2013; Wang et al., 2013). GO exhibited a toxic effect on the wastewater microbial communities at concentrations from 0.05 to 0.3 mg ml− 1, and reduced their metabolic activity by 20–70% in a concentration-dependent manner. The potential mechanism for this toxicity was ascribed to the ROS generated by GO because GO at high concentrations produced higher levels of ROS compared to control samples (Ahmed and Rodrigues, 2013). On the other hand, one study reported enhanced bacterial activity by GO (Wang et al., 2013). The activity of anaerobic ammonium-oxidizing (anammox) bacteria that removes nitrogen from wastewater increased up to 10% and the production of carbohydrate, protein, and total extracellular polymeric substances increased in a dose-dependent manner when GO was treated within the concentration of 0.05–0.1 mg ml− 1. However, at 0.15 mg GO ml−1, anammox bacterial activity and extracellular polymeric substance production decreased (Wang et al., 2013). Because both toxic and nontoxic effects of GO were observed as such, generalized conclusions on safety risks associated with GO are yet to be made (Seabra et al., 2014). In this study, we determined the effects of GO exposure on soil microbial activity. Soil enzyme activity and microbial biomass are sensitive indicators of changes in soil ecosystem function under soil disturbance caused by nanomaterials, heavy metal, and organic pollutants (Chung et al., 2011; Kuperman and Carreiro, 1997; Liu et al., 2009; Shrestha et al., 2013). Therefore, the alterations in extracellular enzyme activity and microbial biomass were determined in soils that were treated with thoroughly characterized GO at 0.1–1 mg GO g−1 soil and incubated for 59 days, a point at which the effects of GO on microbial parameters could no longer be detected. We report for the first time in our knowledge the effects of GO on soil microbial activity. When soils were exposed to 0.5–1 mg GO g−1 soil, significant decrease was observed in the activities of xylosidase, 1,4-β-N-acetyl glucosaminidase, and phosphatase which are soil enzymes that mediate C, N, and P cycling, respectively. These effects subsided afterwards, however. Our results suggest that GO may have negative effects on soil enzyme activity in short term.
subsamples to be incubated were placed in glass jars. The soil was a sandy loam, and the weight proportion of sand, silt, and clay was 52.77 (±2.12) (average (± one standard error), n = 3) %, 39.23 (±2.67) %, and 8.00 (±1.00) %, respectively. The pH of the soil was 4.62 (±0.05). The organic C and N concentrations in soil were 12.16 (±0.49) g C kg−1 soil and 0.80 (±0.03) g N kg−1 soil, respectively. The ratio of C:N in soil was 15.22 (±0.14). 2.2. Characteristics of GO GO was prepared by modified Hummers method (Hummers and Offeman, 1958). Graphite flake (1 g, 99.8%, Alfa Aesar) and NaNO3 (1 g, ≥99.0%, Sigma-Aldrich) were dissolved in H2SO4 (50 ml, 95–97%, Sigma-Aldrich), and KMnO4 (6 g, ≥99.0%, Sigma-Aldrich) was slowly added. The solution mixture was magnetically stirred for 1 h at 35 °C on a hot plate. Deionized (DI) water (80 ml) was added to the solution and heated for 30 min at 90 °C. Subsequently, DI water (200 ml) and H2O2 (6 ml, 30 wt.%, Sigma-Aldrich) were successively added to the solution. Brownish GO powder was obtained by filtration with a glass microfiber filter (CHMLAB, GF4), and stored in vacuum at room temperature. The filtered GO cake was dispersed in 5 wt.% HCl aqueous solution with mild stirring for 12 h in order to remove the metal ions. Subsequently, the GO solution was allowed to settle for 1 day, and the supernatant was decanted away. The completely precipitated GO was purified via dialysis for 3 days through which the residual metal ions and acid were removed. After dialysis, the gel-type GO was redispersed in ethanol and then the mixture was centrifuged at 13,000 rpm for 60 min to separate the solid GO. Finally, the solid product obtained was dried in air. The morphology of GO was investigated by scanning electron microscopy (SEM; Hitachi S-4800, HITACHI, Japan) and high-resolution transmission electron microscopy (HRTEM; Tecnai 20, FEI, USA). The HRTEM image of GO was obtained by drying drops of GO solution on holey carbon grid. The structure of GO was characterized by X-ray diffraction (XRD:D/MAX-2500V/PC, Rigaku INC., Japan) with Cu Kα radiation (λ = 1.5406 Å) at the scanning rate of 4° min−1. Fourier transform infrared spectroscopy (FTIR; Varian 640-IR, Agilent Technologies, USA) was performed to identify chemical functional groups over the wave number range of 800–3800 cm−1 using KBr pellets. X-ray photoelectron spectroscopy (XPS) spectra were obtained with an Automated XPS Microprobe PHI X-tool (ULVAC-PHI, Japan). Raman spectroscopy (Labram ARAMIS IR2, HORIBA, USA) was used to analyze the crystallinity (e.g., crystal structure, disorder, and defects) of GO (Kudin et al., 2008). Thermogravimetric analysis (TGA) was performed (DSC2010, TA Instrument, USA) to confirm the content of metal catalysts and purity of GO by heating it up to 1000 °C at the rate of 10 °C min− 1 under air atmosphere. The specific surface area of GOs was determined by the Brunauer, Emmet, and Teller (BET) method (Brunauer et al., 1938) using BEL SORP-mini II (BEL Japan, Japan) at 350 °C. The suspension behavior of GO at 7.14 mg ml− 1, which was the concentration used for 0.5 mg GO g−1 soil treatment, was analyzed by taking images of GO suspension at various time points after it was sonicated for 1.5 h (Fig. SI-1). In addition, to compare the suspension behavior of GO with that of single-walled CNTs (SWCNTs), SWCNT suspension at 7.14 mg ml−1 was prepared and analyzed in the same way as the GO suspension (Fig. SI-2).
2. Materials and methods 2.3. Soil incubation 2.1. Soil sampling Soil samples were collected in October 2013 from top 15 cm of a site dominated by deciduous trees in Konkuk University campus. This site was chosen for our study because it can represent the urban ecosystem and NMs are most likely to enter soils in an urban environment (Chung et al., 2011; Jin et al., 2013). Upon collection, soil samples were sieved with an 8-mm sieve and kept frozen. Subsequently, 60-g soil
The soil incubation experiment was conducted using suspended form of GO which was prepared by bath-sonicating (Branson, USA) the mixture of GO and DI water at room temperature for 1.5 h. GO suspension was added to 60 g of field-moist soil subsamples placed in glass jars and mixed. The concentrations of GO applied to soils were 0 (DI water only), 0.1, 0.5, and 1.0 mg g−1 soil, and there were 4 replicates for each concentration. In preparing the GO suspension of all
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concentrations, equal volume of DI water was used and this volume was sufficient to disperse the GO throughout each 60-g soil subsample. Control soils also received same volume of DI water. After the GO was added to soils, the soils were incubated at 25 °C and the soil moisture content was maintained at the initial content which was 18.4% of the dry soil weight by supplying DI water regularly. 2.4. Extracellular enzyme assays The activity of 5 extracellular enzymes that mediate C, N, and P cycling in soil ecosystems was determined. Cellobiohydrolase and β-1,4-glucosidase degrade cellulose, and xylosidase breaks down hemicellulose. β-1,4-N-acetylglucosaminidase degrades chitin and acid phosphatase cleaves phosphoester bonds. Fluorogenic substrate methods were used in analyzing extracellular enzyme activities (DeForest, 2009; Saiya-Cork et al., 2002). Briefly, 2 g of soil sample from soils within the incubation jar (4 replicates per concentration) and 120 ml sodium acetate buffer were homogenized, and these soil slurries were put into the black 96-well microplate. Substrates and standards were added and enzyme activities were determined using Synergy HT Multi-Mode Microplate Reader (BioTek, USA). The wavelengths were set at 355 nm for excitation and 430 nm for emission. These extracellular enzyme assays were carried out at 0, 7, 13, 21, 31, 45, 59 days after the application of GO to the soils. The enzyme activities are expressed as nmol 4-MUB g−1 h−1. Control experiment was implemented to determine the effect of GO on enzyme assay. For this, GO suspension of different concentrations (0 (buffer only), 0.1, 0.5, and 1.0 mg g−1 soil) were prepared and enzyme assay was performed. The results showed that GO at different concentrations did not influence the enzyme assay. More specifically, the enzyme activity measured with buffer only and that with different concentrations of GO suspension added was not different (data not shown). 2.5. Microbial biomass C analyses Soil microbial biomass was analyzed by chloroform fumigation– extraction method according to Vance et al. (1987) at 0, 7, 13, 21, 31, 45, 59 days since the soils were treated with various concentrations of GO (4 replicates per concentration). 5 g of soils was fumigated with chloroform in a vacuum desiccator for 3 days in dark. The soils were then extracted with 0.5 M K2SO4 by shaking for 1 h, and the extractant was analyzed for its total organic carbon (TOC) concentration using SIEVERS 5310C Laboratory TOC Analyzer (GE Analytical Instruments, USA). Microbial biomass is expressed as μg C g−1 soil. 2.6. Statistical analyses Analyses of variance (ANOVA) were conducted using SAS version 9.3 (SAS Inc., USA). One-way ANOVA were employed to determine the effect of GO treatment at different concentrations (4 replicates per concentration) on soil enzyme activity and microbial biomass measured at 7 different time points. Significant effects of GO treatment were accepted at α = 0.05. When the effect of GO treatment was significant for enzyme activity or microbial biomass analyzed at each time point, Tukey's honestly significant difference test was used to further determine which means differ from other means within a group (P b 0.05). The correlation between soil microbial biomass and enzyme activity was determined via regression analyses using SPSS version 21 (IBM, USA).
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(Fig. 1(b)). Fig. 2 presents physicochemical characterization of GO. The XRD result is represented in Fig. 2(a). GO displays a strong sharp peak at around 2θ = 9.84°, and this corresponds to d-spacing of 0.898 nm. On the other hand, graphite flake exhibits a prominent peak at around 2θ = 27.64°, and this corresponds to d(002) = 0.322 nm. The FTIR spectra of GO (Fig. 2(b)) shows water peak stretching at 3415 cm−1, carbonyl group stretching at 1730 cm− 1, C\O epoxide group stretching at 1224 cm− 1 and 1053 cm− 1, and C_C aromatic ring stretching at 1621 cm− 1. C1s XPS spectrum of GO is shown in Fig. 2(c). Binding energy estimated through curve fitting showed four peaks that correspond to C\C, C\O, C_O, and O_C\O (Fig. 2(d)). Data shown in Fig. 2(a–d) suggest that there are carbons bonded with oxygen on GO. Raman spectrum displays two well-known peaks, D and G bands at 1339 cm − 1 and 1582 cm− 1, respectively, and the I D /IG ratio is calculated to be 1.15 (Fig. 2(e)). TGA plot consisted of three weight losses (Fig. 2(f)). First of all, there is a mass loss (~ 15.08%) upon heating below 131 °C, corresponding to the elimination of loosely bound or absorbed water and gas molecules. Second, the largest weight loss (~78.69%) is observed around 200 °C, yielding CO, CO2, and various gases that have labile oxygen-carbon bonding. The last weight loss is due to pyrolysis of carbon skeleton (Fig. 2(f)). The specific surface area of GO powder determined through BET measurement was 120.78 (±1.15) m2 g−1 (n = 3). The suspension of GO was evenly dispersed for 24 h, which was the last time point at which the image of GO suspension was taken (Fig. SI-1). On the other hand, SWCNTs suspended in DI water precipitated 1 h after the sonication process (Fig. SI-2). 3.2. Extracellular enzyme activity in soils treated with GO Xylosidase activity was significantly lowered by 0.5 and 1 mg GO g−1 soil on day 21 (P = 0.014). More specifically, GO treatment at 0.5– 1 mg GO g− 1 soil decreased the activity of xylosidase by 34–37% (Fig. 3(a)). On the other hand, on day 31, xylosidase activity was significantly higher under 0.1 mg GO g− 1 soil when compared to that under 1 mg GO g−1 soil (P = 0.047). Xylosidase activity was not affected significantly by GO treatment on days other than 21 and 31 (Fig. 3(a)). Cellobiohydrolase or β-1,4-glucosidase activity was not affected significantly by GO treatment within the 59-day incubation period (Fig. 3(b) and (c)). High concentrations of GO significantly reduced β-1,4-Nacetylglucosaminidase activity on days 0 (P = 0.001), 7 (P = 0.044), and 21 (P = 0.002). More specifically, β-1,4-N-acetylglucosaminidase activity was lowered by 0.5–1 mg GO g−1 soil by 30–40% 2 h after the addition of GOs (day 0). β-1,4-N-acetylglucosaminidase was also reduced under 0.5–1 mg GO g−1 soil by up to 15% at day 7, and by up to 50% at day 21. On the other hand, on day 31, GO treatment at 0.1 mg GO g−1 soil significantly increased the β-1,4-N-acetylglucosaminidase activity when compared to that in control soils (P = 0.018) (Fig. 4(a)). GO treatment significantly lowered the activity of acid phosphatase on days 0 (P = 0.003) and 21 (P = 0.002). At day 0, when acid phosphatase activity was measured 2 h after the GOs were applied, 1 mg GO g− 1 soil decreased enzyme activity by 19% compared to control soils. At day 21, acid phosphatase activity decreased up to 22% under 0.5–1 mg GO g− 1 soil when compared to control soils. On the other hand, phosphatase activity under 0.1 mg GO g−1 soil was higher than that in control soils at day 45 (P = 0.030) (Fig. 4(b)). 3.3. Microbial biomass in soils treated with GO
3. Results 3.1. Characterization of GO The HRTEM images (Figs. 1(a) and SI-3) show a wrinkled transparent sheet and folding structure of the prepared GO. GO has a layered structure of stacked sheets as shown in the cross-section SEM image
GO had no significant effect on soil microbial biomass except for that determined at day 7. More specifically, the microbial biomass was larger under 0.1 mg GO g−1 soil than that under 0.5 and 1 mg GO g−1 soil at day 7 (P = 0.017). On the other hand, the microbial biomass showed a tendency to increase under GO treatments on days 21, 31, and 45, but these results were not statistically significant (P N 0.05) (Fig. 5).
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Fig. 1. The morphology of graphene oxide (GO). (a) Transmission electron microscope (TEM) image and (b) scanning electron microscope (SEM) image of GO.
The microbial biomass was not significantly correlated to any of the soil enzyme activity determined in this study (P N 0.05).
Reduced soil enzyme activity under high GO concentrations suggests that the antimicrobial activity of GO determined in culture studies may apply to soils as well. The antimicrobial properties of GO have been investigated via culture studies using microorganisms that are relevant to human and environmental health including coliform bacteria, plant pathogens, and marine microorganisms (Akhavan and Ghaderi, 2010; Chen et al., 2013; Pretti et al., 2014; Wang et al., 2014). Our findings demonstrate that in addition to these microorganisms in culture, GO may have negative impacts on the functions of environmental microorganisms inhabiting complex matrix such as soils in short term. Our results are also in agreement with a recent report on the negative effects of GOs on the microbial communities within wastewater (Ahmed and Rodrigues, 2013).
4. Discussion The changes in enzyme activity and microbial biomass of soils exposed to GO were determined in this study. High concentrations of GO lowered soil enzyme activity up to 21 days of incubation, but these effects were transient. The microbial biomass in soils treated with GO was not different from that in the control soils. These results suggest that the effect of GO on soil microbial activity may be dependent on time, and underscore the importance of determining the long-term response of soil microorganisms to GO.
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Fig. 2. Characterization of physicochemical properties of graphene oxide (GO). (a) X-ray diffraction (XRD) pattern of GO. (b) Fourier transform infrared spectroscopy (FTIR) spectrum of GO. (c) X-ray photoelectron spectroscopy (XPS) spectrum of GO. (d) C1s spectrum and binding energy of GO estimated via curve fitting. (e) Raman spectrum. (f) Thermal gravimetric analysis (TGA) curve obtained in air atmosphere at the heating rate of 10 °C min−1 of GO.
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Fig. 3. Activity of extracellular enzymes that mediate C cycling in soils treated with up to 1 mg GO g−1 soil during a 59-day incubation period. Activity of (a) xylosidase, (b) cellobiohydrolase, and (c) β-glucosidase. Means followed by the same letters within a day are not significantly different at α = 0.05. Error bars indicate one standard error of the mean (n = 4).
The effects of other carbon-based NMs including CNTs and fullerenes on soil microbial activity have been studied to some extent (Chung et al., 2011; Jin et al., 2013; Shrestha et al., 2013; Tong et al., 2007) but no such studies have been conducted for GOs. In previous studies regarding the effects of CNTs on soil microbial activity conducted within approximately 3 weeks, CNT treatments at 1 mg SWCNTs g−1 soil and 5 mg multiwalled CNTs g−1 soil repressed soil enzyme activity (Chung et al., 2011; Jin et al., 2013); on the other hand, CNT treatment had little effect on soil enzyme activity determined at 3 months (Shrestha et al., 2013). In this study, decrease in soil enzyme activity was observed under 0.5 mg GO g−1 soil at day 21, and this is at a lower concentration than that reported for CNTs. GO has many hydrophilic functional groups (Fig. 2), and this allows GO to be better dispersed in water than CNTs that have significantly less functional groups (Fig. SI-1, 2). Therefore,
the contact of GO, which was dispersed in water and applied to soils, with soil microorganisms is likely to be larger than that of CNTs which were also dispersed in water and added to soils (Ahmed and Rodrigues, 2013; Liu et al., 2011). This may explain the decrease in soil enzyme activity observed under GO treatment at a lower concentration than CNTs. Lowered activities of soil enzymes that decompose hemicellulose, chitin, and organic phosphate produced by soil bacteria and fungi under GO treatment show that in short term, GO may inhibit microbial activity related to C, N, and P cycling in soil ecosystems. However, these effects were transient, and this suggests that soil microbial communities may have recovered from stress exerted by GO, as has been reported for a soil incubation study where the effects of CNTs on soil microorganisms were analyzed (Rodrigues et al., 2013). Soil microorganisms may have
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Fig. 4. Activity of extracellular enzymes that mediate N and P cycling in soils treated with up to 1 mg GO g−1 soil during a 59-day incubation period. Activity of (a) N-acetylglucosaminidase and (b) acid phosphatase. Means followed by the same letters within a day are not significantly different at α = 0.05. Error bars indicate one standard error of the mean (n = 4).
mechanisms to mitigate the toxicity of carbon-based NMs, for instance, via alterations in microbial community composition that includes microorganisms that are more resistant to stress exerted by GO (Rodrigues et al., 2013; Shrestha et al., 2013). The recovery of soil microbial communities and possible changes in their composition, however, needs to be confirmed via further study. In studies that determined the antimicrobial effects of GO, either in culture or in environmental samples, only short-term response was
studied; most of the measurements were made within 48 h (Ahmed and Rodrigues, 2013; Akhavan and Ghaderi, 2010; Carpio et al., 2012). Results from our study that was conducted for 59 days indicate that GO toxicity may be transient. This suggests that acute effects of GO may be different from long-term effects, and thus long-term incubation may be required for determining the effects of GO on the activity of microorganisms inhabiting complex environmental matrix such as soils. On the other hand, the activity of enzymes in control soils increased at
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day 7 and decreased afterwards in this study, and similar trends have been reported in studies where soils were incubated for a designated period of time and soil enzyme activity was measured at different time points (Allison and Vitousek, 2005; Perucci, 1990). 5. Conclusion We demonstrate that high concentrations of GOs, when treated to soils, can decrease soil enzyme activity in short term. On the other hand, soil microbial biomass changed little in response to GO treatment. Altogether, our results suggest that soil enzyme activity is relatively a sensitive indicator of soil disturbance caused by the entry of GOs, and that high concentrations of GO may have adverse effects on soil microbial activity in short term. Acknowledgments This work was supported by the faculty research fund of Konkuk University in 2011. Appendix A. Supplementary data Supplementary data to this article can be found online at http://dx. doi.org/10.1016/j.scitotenv.2015.01.077. References Ahmed, F., Rodrigues, D.F., 2013. Investigation of acute effects of graphene oxide on wastewater microbial community: a case study. J. Hazard. Mater. 256–257, 33–39. Akhavan, O., Ghaderi, E., 2010. Toxicity of graphene and graphene oxide nanowalls against bacteria. ACS Nano 4, 5731–5736. Allison, S.D., Vitousek, P.M., 2005. Responses of extracellular enzymes to simple and complex nutrient inputs. Soil Biol. Biochem. 37, 937–944. Anjum, N.A., Singh, N., Singh, M.K., Shah, Z.A., Duarte, A.C., Pereira, E., Ahmad, I., 2013. Single-bilayer graphene oxide sheet tolerance and glutathione redox system significance assessment in faba bean (Vicia faba L.). J. Nanopart. Res. 15, 1770. Avanasi, R., Jackson, W.A., Sherwin, B., Mudge, J.F., Anderson, T.A., 2014. C60 fullerene soil sorption, biodegradation, and plant uptake. Environ. Sci. Technol. 48, 2792–2797. Brunauer, S., Emmett, P.H., Teller, E., 1938. Adsorption of gases in multimolecular layers. J. Am. Chem. Soc. 60, 309–319. Carpio, I.E.M., Santos, C.M., Wei, X., Rodrigues, D.F., 2012. Toxicity of a polymer-graphene oxide composite against bacterial planktonic cells, biofilms, and mammalian cells. Nanoscale 4, 4746. Chen, J., Wang, X., Han, H., 2013. A new function of graphene oxide emerges: inactivating phytopathogenic bacterium Xanthomonas oryzae pv. Oryzae. J. Nanopart. Res. 15, 1658. Chung, H., Son, Y., Yoon, T.K., Kim, S., Kim, W., 2011. The effect of multi-walled carbon nanotubes on soil microbial activity. Ecotoxicol. Environ. Saf. 74, 569–575. DeForest, J.L., 2009. The influence of time, storage temperature, and substrate age on potential soil enzyme activity in acidic forest soils using MUB-linked substrates and L-DOPA. Soil Biol. Biochem. 41, 1180–1186. Geim, A.K., Novoselov, K.S., 2007. The rise of graphene. Nat. Mater. 6, 183–191. Hu, W., Peng, C., Luo, W., Lv, M., Li, X., Li, D., Huang, Q., Fan, C., 2010. Graphene-based antibacterial paper. ACS Nano 7, 4317–4323.
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Hummers Jr., W.S., Offeman, R.E., 1958. Preparation of graphitic oxide. J. Am. Chem. Soc. 80, 1339. Jin, L., Son, Y., Yoon, T.K., Kang, Y.J., Kim, W., Chung, H., 2013. High concentrations of single-walled carbon nanotubes lower soil enzyme activity and microbial biomass. Ecotoxicol. Environ. Saf. 88, 9–15. Kudin, K.N., Ozbas, B., Schniepp, H.C., Prud'homme, R.K., Aksay, I.A., Car, R., 2008. Raman spectra of graphite oxide and functionalized graphene sheets. Nano Lett. 8, 36–41. Kuperman, R.G., Carreiro, M.M., 1997. Soil heavy metal concentrations, microbial biomass and enzyme activities in a contaminated grassland ecosystem. Soil Biol. Biochem. 29, 179–190. Lerf, A., He, H., Forster, M., Klinowski, J., 1998. Structure of graphite oxide revisited. J. Phys. Chem. B 102, 4477–4482. Li, S., Irin, F., Atore, F.O., Green, M.J., Cañas-Carrell, J.E., 2013a. Determination of multi-walled carbon nanotube bioaccumulation in earthworms measured by a microwave-based detection technique. Sci. Total Environ. 9–13, 445–446. Li, S., Turaga, U., Shrestha, B., Anderson, T.A., Ramkumar, S.S., Green, M.J., Das, S., CañasCarrell, J.E., 2013b. Mobility of polyaromatic hydrocarbons (PAHs) in soil in the presence of carbon nanotubes. Ecotoxicol. Environ. Saf. 96, 168–174. Liu, F., Ying, G.G., Yang, L.H., Zhou, Q.X., 2009. Terrestrial ecotoxicological effects of the antimicrobial agent triclosan. Ecotoxicol. Environ. Saf. 72, 86–92. Liu, S., Zeng, T., Hofmann, M., Burcombe, E., Wei, J., Jiang, R., Kong, J., Chen, Y., 2011. Antibacterial activity of graphite, graphite oxide, graphene oxide, and reduced graphene oxide: membrane and oxidative stress. ACS Nano 9, 6971–6980. Navarro, D.A., Kookana, R.S., Kirby, J.K., Martin, S.M., Shareef, A., Du, J., McLaughlin, M.J., 2013. Behaviour of fullerene (C60) in the terrestrial environment: potential release from biosolids-amended soils. J. Hazard. Mater. 262, 496–503. Park, S., Ruoff, R.S., 2009. Chemical methods for the production of graphenes. Nat. Nanotechnol. 4, 217–224. Perucci, P., 1990. Effect of the addition of municipal solid-waste compost on microbial biomass and enzyme activities in soil. Biol. Fertil. Soils 10, 221–226. Pretti, C., Oliva, M., Pietro, B.D., Monni, G., Cevasco, G., Chiellini, F., Pomelli, C., Chiappe, C., 2014. Ecotoxicity of pristine graphene to marine organisms. Ecotoxicol. Environ. Saf. 101, 138–145. Qi, Z., Zhang, L., Wang, F., Hou, L., Chen, W., 2014. Factors controlling transport of graphene oxide nanoparticles in saturated sand columns. Environ. Toxicol. Chem. 33, 998–1004. Rodrigues, D.F., Jaisi, D.P., Elimelech, M., 2013. Toxicity of functionalized single-walled carbon nanotubes on soil microbial communities: implications for nutrient cycling in soil. Environ. Sci. Technol. 47, 625–633. Saiya-Cork, K.R., Sinsabaugh, R.L., Zak, D.R., 2002. The effects of long term nitrogen deposition on extracellular enzyme activity in an Acer saccharum forest soil. Soil Biol. Biochem. 34, 1309–1315. Seabra, A.B., Paula, A.J., de Lima, R., Alves, O.L., Durán, N., 2014. Nanotoxicity of graphene and graphene oxide. Chem. Res. Toxicol. 27, 159–168. Shrestha, B., Cox, S.B., Das, S., Green, M.J., Li, S., Cañas-Carrell, J.E., 2013. An evaluation of the impact of multiwalled carbon nanotubes on soil microbial community structure and functioning. J. Hazard. Mater. 261, 188–197. Tong, Z., Bischoff, M., Nies, L., Applegate, B., Turco, R.F., 2007. Impact of fullerene (C60) on soil microbial community. Environ. Sci. Technol. 41, 2985–2991. Vance, E.D., Brookes, P.C., Jenkinson, D.S., 1987. An extraction method for measuring soil microbial biomass C. Soil Biol. Biochem. 19, 703–707. Wang, Y., Li, Z., Wang, J., Li, J., Lin, Y., 2011. Graphene and graphene oxide: biofunctionalization and applications in biotechnology. Trends Biotechnol. 29, 205–212. Wang, D., Wang, G., Zhang, G., Xu, X., Yang, F., 2013. Using graphene oxide to enhance the activity of anammox bacteria for nitrogen removal. Bioresour. Technol. 131, 527–530. Wang, X., Liu, X., Chen, J., Han, H., Yuan, Z., 2014. Evaluation and mechanism of antifungal effects of carbon nanomaterials in controlling plant fungal pathogen. Carbon 68, 798–806. Zhao, G., Wen, T., Chen, C., Wang, X., 2012. Synthesis of graphene-based nanomaterials and their application in energy related and environmental-related areas. RCS Adv. 2, 9286–9303.
Contents lists available at ScienceDirect
Science of the Total Environment journal homepage: www.elsevier.com/locate/scitotenv
Effects of graphene oxides on soil enzyme activity and microbial biomass Haegeun Chung a,⁎, Min Ji Kim b, Kwanyoung Ko a, Jae Hyeuk Kim a, Hyun-ah Kwon a, Inpyo Hong a, Nari Park a, Seung-Wook Lee b, Woong Kim b a b
Department of Environmental Engineering, Konkuk University, Seoul 143-701, Republic of Korea Department of Materials Science and Engineering, Korea University, Seoul 136-713, Republic of Korea
H I G H L I G H T S • • • • •
The effects of graphene oxide (GO) on soil microbial activity was studied via a 59-day incubation experiment. Up to 1 mg GO g− 1 soil was applied and the soil enzyme activities and microbial biomass were measured. Soil enzyme activity was lowered by 15–50% under 0.5–1 mg GO g− 1 soil, but the effect subsided afterwards. Soil microbial biomass showed little change in response to GO treatment. GO can negatively affect soil enzyme activity in short term upon entrance to soils.
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Article history: Received 22 October 2014 Received in revised form 24 January 2015 Accepted 24 January 2015 Available online xxxx Editor: D. Barcelo Keywords: Nanomaterial Microbial biomass Soil incubation Extracellular enzymes Soil microorganisms
a b s t r a c t Due to recent developments in nanotechnology, nanomaterials (NMs) such as graphene oxide (GO) may enter the soil environment with mostly unknown consequences. We investigated the effects of GO on soil microbial activity in a 59-day soil incubation study. For this, high-purity GO was prepared and characterized. Soils were treated with up to 1 mg GO g−1 soil, and the changes in the activities of 1,4-β-glucosidase, cellobiohydrolase, xylosidase, 1,4-β-N-acetyl glucosaminidase, and phosphatase and microbial biomass were determined. 0.5–1 mg GO g−1 soil lowered the activity of xylosidase, 1,4-β-N-acetyl glucosaminidase, and phosphatase by up to 50% when compared to that in the control soils up to 21 days of incubation. Microbial biomass in soils treated with GO was not significantly different from that in control soils throughout the incubation period, and the soil enzyme activity and microbial biomass were not significantly correlated in this study. Our results indicate that soil enzyme activity can be lowered by the entry of GO into soils in short term but it can be recovered afterwards. © 2015 Elsevier B.V. All rights reserved.
1. Introduction Graphene oxides (GOs) are layered graphene sheets that have oxygen-containing functional groups including epoxide, carboxyl, carbonyl, and hydroxyl group (Lerf et al., 1998). Among a wide range of nanomaterials (NMs) that are being manufactured, GO or chemically modified GO are considered to be highly promising novel material due to their superior electrical characteristics, colloidal property, and large surface area (Geim and Novoselov, 2007). These excellent material properties allow GO to be used in various applications such as energystorage materials, paper-like materials, and bioenvironmental materials (Park and Ruoff, 2009; Wang et al., 2011; Zhao et al., 2012). For example, adsorbents and photocatalysts that are based on GO are employed in removing pollutants from the environment (Zhao et al., 2012). For ⁎ Corresponding author. E-mail address: [email protected] (H. Chung).
http://dx.doi.org/10.1016/j.scitotenv.2015.01.077 0048-9697/© 2015 Elsevier B.V. All rights reserved.
GO to be successfully applied in such diverse fields, it is important to determine its fate, distribution, and potential environmental impacts (Anjum et al., 2013). Previous studies have determined the fate and toxicity of other carbon-based NMs including carbon nanotubes (CNTs) and fullerenes in the soil environment (Avanasi et al., 2014; Li et al., 2013a, 2013b; Navarro et al., 2013), but few studies have investigated the impacts of GO on the soil environment. Compared with other carbon-based NMs, GO showed higher mobility in sand, which suggests that highly mobile GO may increase the environmental risks once they enter the soil environment (Qi et al., 2014). However, how GO may impact soil microbial activity that is important for nutrient cycling in soil ecosystems remains to be investigated. It has been shown via culture studies using model microorganisms such as Escherichia coli (E. coli) that GO has strong antimicrobial effect (Akhavan and Ghaderi, 2010; Hu et al., 2010; Liu et al., 2011). For example, a study using GO nanowalls showed that direct contact of bacteria with GO nanowalls can lead to cell damage; after 1 h of bacteria-GO
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nanowall contact, only 41% survived. This cytotoxicity was attributed to the sharp edges of the nanowalls (Akhavan and Ghaderi, 2010). In another study, it was shown that after 2-h contact of bacteria with GO nanosheets, the cell metabolic activity was decreased to approximately 70% and 13% at 0.02 mg ml− 1 and 0.085 mg ml− 1, respectively. The bacterial cell membranes were shown to have been severely damaged and the cytoplasm was flowing out (Hu et al., 2010). Additionally, GO treated on bacteria at 0.005–0.08 mg ml−1 for up to 4 h showed timeand concentration-dependent antibacterial activity (Liu et al., 2011). Antimicrobial effects of GO on microorganisms other than model microorganisms, such as some plant pathogens, have also been shown (Chen et al., 2013; Wang et al., 2014). GO showed 94% cytotoxicity against phytopathogenic bacteria causing infections in rice even at a low concentration. This high antimicrobial efficiency was ascribed to the extremely sharp edges of GO and generation of reactive oxygen species (ROS) (Chen et al., 2013). The antifungal activity of GO was also shown for two important plant pathogenic fungi (Wang et al., 2014). While the antimicrobial effects of GO have been demonstrated in culture studies, only a few studies determined the effects of GO on microbial communities inhabiting environmental samples (Ahmed and Rodrigues, 2013; Wang et al., 2013). GO exhibited a toxic effect on the wastewater microbial communities at concentrations from 0.05 to 0.3 mg ml− 1, and reduced their metabolic activity by 20–70% in a concentration-dependent manner. The potential mechanism for this toxicity was ascribed to the ROS generated by GO because GO at high concentrations produced higher levels of ROS compared to control samples (Ahmed and Rodrigues, 2013). On the other hand, one study reported enhanced bacterial activity by GO (Wang et al., 2013). The activity of anaerobic ammonium-oxidizing (anammox) bacteria that removes nitrogen from wastewater increased up to 10% and the production of carbohydrate, protein, and total extracellular polymeric substances increased in a dose-dependent manner when GO was treated within the concentration of 0.05–0.1 mg ml− 1. However, at 0.15 mg GO ml−1, anammox bacterial activity and extracellular polymeric substance production decreased (Wang et al., 2013). Because both toxic and nontoxic effects of GO were observed as such, generalized conclusions on safety risks associated with GO are yet to be made (Seabra et al., 2014). In this study, we determined the effects of GO exposure on soil microbial activity. Soil enzyme activity and microbial biomass are sensitive indicators of changes in soil ecosystem function under soil disturbance caused by nanomaterials, heavy metal, and organic pollutants (Chung et al., 2011; Kuperman and Carreiro, 1997; Liu et al., 2009; Shrestha et al., 2013). Therefore, the alterations in extracellular enzyme activity and microbial biomass were determined in soils that were treated with thoroughly characterized GO at 0.1–1 mg GO g−1 soil and incubated for 59 days, a point at which the effects of GO on microbial parameters could no longer be detected. We report for the first time in our knowledge the effects of GO on soil microbial activity. When soils were exposed to 0.5–1 mg GO g−1 soil, significant decrease was observed in the activities of xylosidase, 1,4-β-N-acetyl glucosaminidase, and phosphatase which are soil enzymes that mediate C, N, and P cycling, respectively. These effects subsided afterwards, however. Our results suggest that GO may have negative effects on soil enzyme activity in short term.
subsamples to be incubated were placed in glass jars. The soil was a sandy loam, and the weight proportion of sand, silt, and clay was 52.77 (±2.12) (average (± one standard error), n = 3) %, 39.23 (±2.67) %, and 8.00 (±1.00) %, respectively. The pH of the soil was 4.62 (±0.05). The organic C and N concentrations in soil were 12.16 (±0.49) g C kg−1 soil and 0.80 (±0.03) g N kg−1 soil, respectively. The ratio of C:N in soil was 15.22 (±0.14). 2.2. Characteristics of GO GO was prepared by modified Hummers method (Hummers and Offeman, 1958). Graphite flake (1 g, 99.8%, Alfa Aesar) and NaNO3 (1 g, ≥99.0%, Sigma-Aldrich) were dissolved in H2SO4 (50 ml, 95–97%, Sigma-Aldrich), and KMnO4 (6 g, ≥99.0%, Sigma-Aldrich) was slowly added. The solution mixture was magnetically stirred for 1 h at 35 °C on a hot plate. Deionized (DI) water (80 ml) was added to the solution and heated for 30 min at 90 °C. Subsequently, DI water (200 ml) and H2O2 (6 ml, 30 wt.%, Sigma-Aldrich) were successively added to the solution. Brownish GO powder was obtained by filtration with a glass microfiber filter (CHMLAB, GF4), and stored in vacuum at room temperature. The filtered GO cake was dispersed in 5 wt.% HCl aqueous solution with mild stirring for 12 h in order to remove the metal ions. Subsequently, the GO solution was allowed to settle for 1 day, and the supernatant was decanted away. The completely precipitated GO was purified via dialysis for 3 days through which the residual metal ions and acid were removed. After dialysis, the gel-type GO was redispersed in ethanol and then the mixture was centrifuged at 13,000 rpm for 60 min to separate the solid GO. Finally, the solid product obtained was dried in air. The morphology of GO was investigated by scanning electron microscopy (SEM; Hitachi S-4800, HITACHI, Japan) and high-resolution transmission electron microscopy (HRTEM; Tecnai 20, FEI, USA). The HRTEM image of GO was obtained by drying drops of GO solution on holey carbon grid. The structure of GO was characterized by X-ray diffraction (XRD:D/MAX-2500V/PC, Rigaku INC., Japan) with Cu Kα radiation (λ = 1.5406 Å) at the scanning rate of 4° min−1. Fourier transform infrared spectroscopy (FTIR; Varian 640-IR, Agilent Technologies, USA) was performed to identify chemical functional groups over the wave number range of 800–3800 cm−1 using KBr pellets. X-ray photoelectron spectroscopy (XPS) spectra were obtained with an Automated XPS Microprobe PHI X-tool (ULVAC-PHI, Japan). Raman spectroscopy (Labram ARAMIS IR2, HORIBA, USA) was used to analyze the crystallinity (e.g., crystal structure, disorder, and defects) of GO (Kudin et al., 2008). Thermogravimetric analysis (TGA) was performed (DSC2010, TA Instrument, USA) to confirm the content of metal catalysts and purity of GO by heating it up to 1000 °C at the rate of 10 °C min− 1 under air atmosphere. The specific surface area of GOs was determined by the Brunauer, Emmet, and Teller (BET) method (Brunauer et al., 1938) using BEL SORP-mini II (BEL Japan, Japan) at 350 °C. The suspension behavior of GO at 7.14 mg ml− 1, which was the concentration used for 0.5 mg GO g−1 soil treatment, was analyzed by taking images of GO suspension at various time points after it was sonicated for 1.5 h (Fig. SI-1). In addition, to compare the suspension behavior of GO with that of single-walled CNTs (SWCNTs), SWCNT suspension at 7.14 mg ml−1 was prepared and analyzed in the same way as the GO suspension (Fig. SI-2).
2. Materials and methods 2.3. Soil incubation 2.1. Soil sampling Soil samples were collected in October 2013 from top 15 cm of a site dominated by deciduous trees in Konkuk University campus. This site was chosen for our study because it can represent the urban ecosystem and NMs are most likely to enter soils in an urban environment (Chung et al., 2011; Jin et al., 2013). Upon collection, soil samples were sieved with an 8-mm sieve and kept frozen. Subsequently, 60-g soil
The soil incubation experiment was conducted using suspended form of GO which was prepared by bath-sonicating (Branson, USA) the mixture of GO and DI water at room temperature for 1.5 h. GO suspension was added to 60 g of field-moist soil subsamples placed in glass jars and mixed. The concentrations of GO applied to soils were 0 (DI water only), 0.1, 0.5, and 1.0 mg g−1 soil, and there were 4 replicates for each concentration. In preparing the GO suspension of all
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concentrations, equal volume of DI water was used and this volume was sufficient to disperse the GO throughout each 60-g soil subsample. Control soils also received same volume of DI water. After the GO was added to soils, the soils were incubated at 25 °C and the soil moisture content was maintained at the initial content which was 18.4% of the dry soil weight by supplying DI water regularly. 2.4. Extracellular enzyme assays The activity of 5 extracellular enzymes that mediate C, N, and P cycling in soil ecosystems was determined. Cellobiohydrolase and β-1,4-glucosidase degrade cellulose, and xylosidase breaks down hemicellulose. β-1,4-N-acetylglucosaminidase degrades chitin and acid phosphatase cleaves phosphoester bonds. Fluorogenic substrate methods were used in analyzing extracellular enzyme activities (DeForest, 2009; Saiya-Cork et al., 2002). Briefly, 2 g of soil sample from soils within the incubation jar (4 replicates per concentration) and 120 ml sodium acetate buffer were homogenized, and these soil slurries were put into the black 96-well microplate. Substrates and standards were added and enzyme activities were determined using Synergy HT Multi-Mode Microplate Reader (BioTek, USA). The wavelengths were set at 355 nm for excitation and 430 nm for emission. These extracellular enzyme assays were carried out at 0, 7, 13, 21, 31, 45, 59 days after the application of GO to the soils. The enzyme activities are expressed as nmol 4-MUB g−1 h−1. Control experiment was implemented to determine the effect of GO on enzyme assay. For this, GO suspension of different concentrations (0 (buffer only), 0.1, 0.5, and 1.0 mg g−1 soil) were prepared and enzyme assay was performed. The results showed that GO at different concentrations did not influence the enzyme assay. More specifically, the enzyme activity measured with buffer only and that with different concentrations of GO suspension added was not different (data not shown). 2.5. Microbial biomass C analyses Soil microbial biomass was analyzed by chloroform fumigation– extraction method according to Vance et al. (1987) at 0, 7, 13, 21, 31, 45, 59 days since the soils were treated with various concentrations of GO (4 replicates per concentration). 5 g of soils was fumigated with chloroform in a vacuum desiccator for 3 days in dark. The soils were then extracted with 0.5 M K2SO4 by shaking for 1 h, and the extractant was analyzed for its total organic carbon (TOC) concentration using SIEVERS 5310C Laboratory TOC Analyzer (GE Analytical Instruments, USA). Microbial biomass is expressed as μg C g−1 soil. 2.6. Statistical analyses Analyses of variance (ANOVA) were conducted using SAS version 9.3 (SAS Inc., USA). One-way ANOVA were employed to determine the effect of GO treatment at different concentrations (4 replicates per concentration) on soil enzyme activity and microbial biomass measured at 7 different time points. Significant effects of GO treatment were accepted at α = 0.05. When the effect of GO treatment was significant for enzyme activity or microbial biomass analyzed at each time point, Tukey's honestly significant difference test was used to further determine which means differ from other means within a group (P b 0.05). The correlation between soil microbial biomass and enzyme activity was determined via regression analyses using SPSS version 21 (IBM, USA).
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(Fig. 1(b)). Fig. 2 presents physicochemical characterization of GO. The XRD result is represented in Fig. 2(a). GO displays a strong sharp peak at around 2θ = 9.84°, and this corresponds to d-spacing of 0.898 nm. On the other hand, graphite flake exhibits a prominent peak at around 2θ = 27.64°, and this corresponds to d(002) = 0.322 nm. The FTIR spectra of GO (Fig. 2(b)) shows water peak stretching at 3415 cm−1, carbonyl group stretching at 1730 cm− 1, C\O epoxide group stretching at 1224 cm− 1 and 1053 cm− 1, and C_C aromatic ring stretching at 1621 cm− 1. C1s XPS spectrum of GO is shown in Fig. 2(c). Binding energy estimated through curve fitting showed four peaks that correspond to C\C, C\O, C_O, and O_C\O (Fig. 2(d)). Data shown in Fig. 2(a–d) suggest that there are carbons bonded with oxygen on GO. Raman spectrum displays two well-known peaks, D and G bands at 1339 cm − 1 and 1582 cm− 1, respectively, and the I D /IG ratio is calculated to be 1.15 (Fig. 2(e)). TGA plot consisted of three weight losses (Fig. 2(f)). First of all, there is a mass loss (~ 15.08%) upon heating below 131 °C, corresponding to the elimination of loosely bound or absorbed water and gas molecules. Second, the largest weight loss (~78.69%) is observed around 200 °C, yielding CO, CO2, and various gases that have labile oxygen-carbon bonding. The last weight loss is due to pyrolysis of carbon skeleton (Fig. 2(f)). The specific surface area of GO powder determined through BET measurement was 120.78 (±1.15) m2 g−1 (n = 3). The suspension of GO was evenly dispersed for 24 h, which was the last time point at which the image of GO suspension was taken (Fig. SI-1). On the other hand, SWCNTs suspended in DI water precipitated 1 h after the sonication process (Fig. SI-2). 3.2. Extracellular enzyme activity in soils treated with GO Xylosidase activity was significantly lowered by 0.5 and 1 mg GO g−1 soil on day 21 (P = 0.014). More specifically, GO treatment at 0.5– 1 mg GO g− 1 soil decreased the activity of xylosidase by 34–37% (Fig. 3(a)). On the other hand, on day 31, xylosidase activity was significantly higher under 0.1 mg GO g− 1 soil when compared to that under 1 mg GO g−1 soil (P = 0.047). Xylosidase activity was not affected significantly by GO treatment on days other than 21 and 31 (Fig. 3(a)). Cellobiohydrolase or β-1,4-glucosidase activity was not affected significantly by GO treatment within the 59-day incubation period (Fig. 3(b) and (c)). High concentrations of GO significantly reduced β-1,4-Nacetylglucosaminidase activity on days 0 (P = 0.001), 7 (P = 0.044), and 21 (P = 0.002). More specifically, β-1,4-N-acetylglucosaminidase activity was lowered by 0.5–1 mg GO g−1 soil by 30–40% 2 h after the addition of GOs (day 0). β-1,4-N-acetylglucosaminidase was also reduced under 0.5–1 mg GO g−1 soil by up to 15% at day 7, and by up to 50% at day 21. On the other hand, on day 31, GO treatment at 0.1 mg GO g−1 soil significantly increased the β-1,4-N-acetylglucosaminidase activity when compared to that in control soils (P = 0.018) (Fig. 4(a)). GO treatment significantly lowered the activity of acid phosphatase on days 0 (P = 0.003) and 21 (P = 0.002). At day 0, when acid phosphatase activity was measured 2 h after the GOs were applied, 1 mg GO g− 1 soil decreased enzyme activity by 19% compared to control soils. At day 21, acid phosphatase activity decreased up to 22% under 0.5–1 mg GO g− 1 soil when compared to control soils. On the other hand, phosphatase activity under 0.1 mg GO g−1 soil was higher than that in control soils at day 45 (P = 0.030) (Fig. 4(b)). 3.3. Microbial biomass in soils treated with GO
3. Results 3.1. Characterization of GO The HRTEM images (Figs. 1(a) and SI-3) show a wrinkled transparent sheet and folding structure of the prepared GO. GO has a layered structure of stacked sheets as shown in the cross-section SEM image
GO had no significant effect on soil microbial biomass except for that determined at day 7. More specifically, the microbial biomass was larger under 0.1 mg GO g−1 soil than that under 0.5 and 1 mg GO g−1 soil at day 7 (P = 0.017). On the other hand, the microbial biomass showed a tendency to increase under GO treatments on days 21, 31, and 45, but these results were not statistically significant (P N 0.05) (Fig. 5).
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Fig. 1. The morphology of graphene oxide (GO). (a) Transmission electron microscope (TEM) image and (b) scanning electron microscope (SEM) image of GO.
The microbial biomass was not significantly correlated to any of the soil enzyme activity determined in this study (P N 0.05).
Reduced soil enzyme activity under high GO concentrations suggests that the antimicrobial activity of GO determined in culture studies may apply to soils as well. The antimicrobial properties of GO have been investigated via culture studies using microorganisms that are relevant to human and environmental health including coliform bacteria, plant pathogens, and marine microorganisms (Akhavan and Ghaderi, 2010; Chen et al., 2013; Pretti et al., 2014; Wang et al., 2014). Our findings demonstrate that in addition to these microorganisms in culture, GO may have negative impacts on the functions of environmental microorganisms inhabiting complex matrix such as soils in short term. Our results are also in agreement with a recent report on the negative effects of GOs on the microbial communities within wastewater (Ahmed and Rodrigues, 2013).
4. Discussion The changes in enzyme activity and microbial biomass of soils exposed to GO were determined in this study. High concentrations of GO lowered soil enzyme activity up to 21 days of incubation, but these effects were transient. The microbial biomass in soils treated with GO was not different from that in the control soils. These results suggest that the effect of GO on soil microbial activity may be dependent on time, and underscore the importance of determining the long-term response of soil microorganisms to GO.
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Fig. 2. Characterization of physicochemical properties of graphene oxide (GO). (a) X-ray diffraction (XRD) pattern of GO. (b) Fourier transform infrared spectroscopy (FTIR) spectrum of GO. (c) X-ray photoelectron spectroscopy (XPS) spectrum of GO. (d) C1s spectrum and binding energy of GO estimated via curve fitting. (e) Raman spectrum. (f) Thermal gravimetric analysis (TGA) curve obtained in air atmosphere at the heating rate of 10 °C min−1 of GO.
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Fig. 3. Activity of extracellular enzymes that mediate C cycling in soils treated with up to 1 mg GO g−1 soil during a 59-day incubation period. Activity of (a) xylosidase, (b) cellobiohydrolase, and (c) β-glucosidase. Means followed by the same letters within a day are not significantly different at α = 0.05. Error bars indicate one standard error of the mean (n = 4).
The effects of other carbon-based NMs including CNTs and fullerenes on soil microbial activity have been studied to some extent (Chung et al., 2011; Jin et al., 2013; Shrestha et al., 2013; Tong et al., 2007) but no such studies have been conducted for GOs. In previous studies regarding the effects of CNTs on soil microbial activity conducted within approximately 3 weeks, CNT treatments at 1 mg SWCNTs g−1 soil and 5 mg multiwalled CNTs g−1 soil repressed soil enzyme activity (Chung et al., 2011; Jin et al., 2013); on the other hand, CNT treatment had little effect on soil enzyme activity determined at 3 months (Shrestha et al., 2013). In this study, decrease in soil enzyme activity was observed under 0.5 mg GO g−1 soil at day 21, and this is at a lower concentration than that reported for CNTs. GO has many hydrophilic functional groups (Fig. 2), and this allows GO to be better dispersed in water than CNTs that have significantly less functional groups (Fig. SI-1, 2). Therefore,
the contact of GO, which was dispersed in water and applied to soils, with soil microorganisms is likely to be larger than that of CNTs which were also dispersed in water and added to soils (Ahmed and Rodrigues, 2013; Liu et al., 2011). This may explain the decrease in soil enzyme activity observed under GO treatment at a lower concentration than CNTs. Lowered activities of soil enzymes that decompose hemicellulose, chitin, and organic phosphate produced by soil bacteria and fungi under GO treatment show that in short term, GO may inhibit microbial activity related to C, N, and P cycling in soil ecosystems. However, these effects were transient, and this suggests that soil microbial communities may have recovered from stress exerted by GO, as has been reported for a soil incubation study where the effects of CNTs on soil microorganisms were analyzed (Rodrigues et al., 2013). Soil microorganisms may have
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Fig. 4. Activity of extracellular enzymes that mediate N and P cycling in soils treated with up to 1 mg GO g−1 soil during a 59-day incubation period. Activity of (a) N-acetylglucosaminidase and (b) acid phosphatase. Means followed by the same letters within a day are not significantly different at α = 0.05. Error bars indicate one standard error of the mean (n = 4).
mechanisms to mitigate the toxicity of carbon-based NMs, for instance, via alterations in microbial community composition that includes microorganisms that are more resistant to stress exerted by GO (Rodrigues et al., 2013; Shrestha et al., 2013). The recovery of soil microbial communities and possible changes in their composition, however, needs to be confirmed via further study. In studies that determined the antimicrobial effects of GO, either in culture or in environmental samples, only short-term response was
studied; most of the measurements were made within 48 h (Ahmed and Rodrigues, 2013; Akhavan and Ghaderi, 2010; Carpio et al., 2012). Results from our study that was conducted for 59 days indicate that GO toxicity may be transient. This suggests that acute effects of GO may be different from long-term effects, and thus long-term incubation may be required for determining the effects of GO on the activity of microorganisms inhabiting complex environmental matrix such as soils. On the other hand, the activity of enzymes in control soils increased at
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Fig. 5. Microbial biomass of soils treated with up to 1 mg GO g−1 soil during a 59-day incubation period. Means followed by the same letters within a day are not significantly different at α = 0.05. Error bars indicate one standard error of the mean (n = 4).
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day 7 and decreased afterwards in this study, and similar trends have been reported in studies where soils were incubated for a designated period of time and soil enzyme activity was measured at different time points (Allison and Vitousek, 2005; Perucci, 1990). 5. Conclusion We demonstrate that high concentrations of GOs, when treated to soils, can decrease soil enzyme activity in short term. On the other hand, soil microbial biomass changed little in response to GO treatment. Altogether, our results suggest that soil enzyme activity is relatively a sensitive indicator of soil disturbance caused by the entry of GOs, and that high concentrations of GO may have adverse effects on soil microbial activity in short term. Acknowledgments This work was supported by the faculty research fund of Konkuk University in 2011. Appendix A. Supplementary data Supplementary data to this article can be found online at http://dx. doi.org/10.1016/j.scitotenv.2015.01.077. References Ahmed, F., Rodrigues, D.F., 2013. Investigation of acute effects of graphene oxide on wastewater microbial community: a case study. J. Hazard. Mater. 256–257, 33–39. Akhavan, O., Ghaderi, E., 2010. Toxicity of graphene and graphene oxide nanowalls against bacteria. ACS Nano 4, 5731–5736. Allison, S.D., Vitousek, P.M., 2005. Responses of extracellular enzymes to simple and complex nutrient inputs. Soil Biol. Biochem. 37, 937–944. Anjum, N.A., Singh, N., Singh, M.K., Shah, Z.A., Duarte, A.C., Pereira, E., Ahmad, I., 2013. Single-bilayer graphene oxide sheet tolerance and glutathione redox system significance assessment in faba bean (Vicia faba L.). J. Nanopart. Res. 15, 1770. Avanasi, R., Jackson, W.A., Sherwin, B., Mudge, J.F., Anderson, T.A., 2014. C60 fullerene soil sorption, biodegradation, and plant uptake. Environ. Sci. Technol. 48, 2792–2797. Brunauer, S., Emmett, P.H., Teller, E., 1938. Adsorption of gases in multimolecular layers. J. Am. Chem. Soc. 60, 309–319. Carpio, I.E.M., Santos, C.M., Wei, X., Rodrigues, D.F., 2012. Toxicity of a polymer-graphene oxide composite against bacterial planktonic cells, biofilms, and mammalian cells. Nanoscale 4, 4746. Chen, J., Wang, X., Han, H., 2013. A new function of graphene oxide emerges: inactivating phytopathogenic bacterium Xanthomonas oryzae pv. Oryzae. J. Nanopart. Res. 15, 1658. Chung, H., Son, Y., Yoon, T.K., Kim, S., Kim, W., 2011. The effect of multi-walled carbon nanotubes on soil microbial activity. Ecotoxicol. Environ. Saf. 74, 569–575. DeForest, J.L., 2009. The influence of time, storage temperature, and substrate age on potential soil enzyme activity in acidic forest soils using MUB-linked substrates and L-DOPA. Soil Biol. Biochem. 41, 1180–1186. Geim, A.K., Novoselov, K.S., 2007. The rise of graphene. Nat. Mater. 6, 183–191. Hu, W., Peng, C., Luo, W., Lv, M., Li, X., Li, D., Huang, Q., Fan, C., 2010. Graphene-based antibacterial paper. ACS Nano 7, 4317–4323.
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