Role of Nanoparticles in Controlling Arsenic Mobilization from

May 22, 2014 - College of Chemistry and Life Science, Quanzhou Normal University, Quanzhou, P. R. China. §. Environmental Science Research Center, ...
1 downloads 0 Views 6MB Size
Download PDF
Article pubs.acs.org/est

Role of Nanoparticles in Controlling Arsenic Mobilization from Sediments near a Realgar Tailing Guowen Dong,†,‡,§ Yaohua Huang,† Qiangqiang Yu,† Yuanpeng Wang,*,† Haitao Wang,† Ning He,† and Qingbiao Li*,†,∥,§ †

Department of Chemical and Biochemical Engineering, College of Chemistry and Chemical Engineering, and The Key Laboratory for Synthetic Biotechnology of Xiamen City, Xiamen University, Xiamen, P. R. China ‡ College of Resources & Chemical Engineering, Sanming University, Sanming, P. R. China ∥ College of Chemistry and Life Science, Quanzhou Normal University, Quanzhou, P. R. China § Environmental Science Research Center, College of the Environment & Ecology, Xiamen University, Xiamen, P. R. China S Supporting Information *

ABSTRACT: Microcosm experiments were conducted to investigate the mechanism of microbial-mediated As mobilization from high arsenic tailing sediments amended with nanoparticles (NPs). The addition of SiO2 NPs could substantially stimulate arsenic mobilization in the sodium acetate amendment sediments. However, the addition of Fe2O3 and Fe3O4 NPs restrained arsenic release because these NPs resulted in Fe−As coprecipiate. Moreover, NP additions in sediments amended with sodium acetate as the electron donor clearly promoted microbial dissimilatory iron reduction. Nearly 4 times the Fe(II) (11.67−12.87 mg· L−1) from sediments amended with NPs and sodium acetate was released compared to sediments amended with only sodium acetate (3.49 mg·L−1). Based on molecular fingerprinting and sequencing analyses, the NP additions could potentially change the sediment bacterial community composition and increase the abundance of Fe(III) and As(V) reduction bacteria. Several potential NP-stimulated bacteria were related to Geobacter, Anaeromyxobacter, Clostridium, and Alicyclobacillus. The findings offer a relatively comprehensive assessment of NP (e.g., Fe2O3, Fe3O4, and SiO2) effects on sediment bacterial communities and As mobilization.

1. INTRODUCTION Arsenic (As) is widely recognized as one of the most toxic chemical elements and is considered the primary naturally occurring carcinogen in the environment. Due to its serious toxicity, many studies on the fate of As in tailings sediments have been performed, which include the As distribution and geochemistry in tailings.1,2 Nevertheless, the actual As environmental risk is associated with its mobile or bioavailable chemical forms in tailing sediments. Among the As chemical forms or species, the inorganic form is most damaging, especially As(III) in sediments. To date, biogeochemical investigations on the fate of As have revealed that bacteria play an important role in controlling As geochemistry. Numerous studies offer direct evidence that anaerobic metal-reducing bacteria play an important role in the formation of toxic, mobile As(III) in tailing sediments.3−5 Some anaerobic bacteria, especially the family Geobacteraceae,6 are able to gain energy by coupling As(V) reduction to the oxidation of organic carbon, that is, the capability to reduce As(V) to As(III) by means of an As respiration system (arrA).7 Moreover, As bacterial reduction can enhance As mobility in sediment−water systems because As(III) is more mobile than As(V). Many factors affect the As microbial reduction process. © 2014 American Chemical Society

For example, electron acceptors (e.g., insoluble Fe(III) or Mn(IV) oxides), electron-shuttling compounds, organic carbon, and some foreign material, especially nanomaterials, have large effects on microbial reduction. As nanotechnology makes a greater appearance in the field of environmental remediation, widespread nanoparticle (NP) use will result in their release into aquatic and soil systems. NPs are not exclusively anthropogenically synthesized.8 For example, different colloidal NPs are naturally produced from microorganisms and the weathering of metal oxides and silicates. These NPs may play an important role in the fate, transport, transformation, and bioavailability of environmentally relevant substances, which may be taken up by plants and soil organisms. Moreover, NPs may enhance the risks associated with other pollutants by their presence.9,10 For example, the As(V) microbial reduction process may be affected by transferring electrons with NPs acting as electron conduits. Pure-culture studies have shown that dissimilatory metalReceived: Revised: Accepted: Published: 7469

December 11, 2013 April 7, 2014 May 22, 2014 May 22, 2014 dx.doi.org/10.1021/es4055077 | Environ. Sci. Technol. 2014, 48, 7469−7476

Environmental Science & Technology

Article

2.3. Batch Incubation Assays Preparation. Approximately 20 g of sediment was mixed with 24 mL deionized water. A sediment/water slurry was placed in 105 mL serum bottles, N2-bubbled for 30 min and fitted with a butyl rubber stopper, sealed with an aluminum clamp under an N 2 atmosphere and incubated at 30 °C in the dark. The experimental conditions were as follows: (i) anaerobic, amended with 0.88 mM sodium acetate; (ii) anaerobic, amended with 0.88 mM sodium acetate and 0.2 g of Fe2O3, Fe3O4, and SiO2 microparticles (a few micrometers); (iii) anaerobic, amended with 0.88 mM sodium acetate and 0.2 g of Fe2O3, Fe3O4, and SiO2 NPs (25−40 nm); (iv) anaerobic, amended with deionized water; and (v) the aforementioned anaerobic sterile samples were autoclaved, respectively. Microcosms were subsampled at discrete time points, maintaining the same sediment/water ratio throughout the experiment. Each microcosm was prepared in triplicate. Analytical-grade or better chemicals were used in this study. 2.4. Chemical Analyses. The sample As (total), As(III) and Fe(II) concentrations were analyzed. For microcosm sediment cultures, approximately 1.0 mL of sample supernatant was withdrawn from the microcosm culture at each time in an anaerobic chamber. The supernatant was then passed through a 0.45-μm filter. Furthermore, 100 μL of filtered aqueous sample was immediately anaerobically mixed with 4.9 mL ferrozine (1 g·L−1) solution at room temperature for 10 min and analyzed for Fe(II) using a modified ferrozine-based method at 562 nm on a UNICAM UV-300 spectrophotometer (Thermo Spectronic).16 The remaining filtered aqueous sample was analyzed for total As using ICP-MS and for As(III) using HG-AFS (AF610B).17 2.4. PCR-DGGE. After 7 weeks of anaerobic incubation, the microbial community phylogenetic diversity in different treatment sediments was evaluated with PCR amplification of 16S rDNA genes and denaturing gradient gel electrophoresis (DGGE). Sediment DNA was extracted from approximately 0.5 g of sediment using the Fast DNA Spin Kit for soil (MP Biomedical) according to the manufacturer’s instructions. The 16S rRNA fragment (200 bp) was amplified by PCR from extracted DNA samples using universal bacterial primers as described by Liu.18 Briefly, the forward primer was F338-GC, and the reverse primer was R518. The presence of PCR products was confirmed by analyzing 5 μL of the product on a 1.5% agarose gel that was stained with SYBR Green I. A DCode Universal Mutation Detection System (Bio-Rad, Hercules, CA) was used for DGGE analysis. Here, 20-μL PCR samples were electrophoresed on an 8% polyacrylamide gel (37.5:1 acrylamide: bis(acrylamide)) with a denaturing gradient that ranged from 30% to 70%(where the 100% denaturation corresponds to 7 M urea and 40% formamide). All gels were run in 1 × TAE buffer (40 mM Tris base, 20 mM sodium acetate, and 1 mM EDTA) at a constant 150 V and 60 °C. After 5 h of electrophoresis, the gel was stained with SYBR Green I for 30 min and then visualized using a Gel imager with a CCD camera (Gel Logic 200 Kodak). Dominant DGGE bands were excised and eluted overnight in sterilized Milli-Q water at 4 °C and reamplified using F338 and R518 primers without GC clamps. PCR products were purified using Qiaquick PCR cleanup columns (Qiagen, Valencia, CA) before cloning. To profile Geobacteraceae communities in the DGGE, a nested PCR approach was applied as described by Lin.19 First, a Geobacteraceae-specific PCR was performed to amplify a 0.8 kb 16S rRNA gene fragment using bacterial forward primer 8F (5′-

reducing bacteria can utilize iron oxide NPs as electron ́ conduits to reduce distant terminal acceptors.11 DominguezGaray et al. reported that silica colloid formation enhanced the performance of sediment microbial fuel cells in a low conductivity soil.12 An electrically conductive network between microbes and minerals in sediments using NPs as a longdistance electron transfer conduit may exist. He et al. reported that the addition of iron oxide magnetic NPs could potentially stimulate bacterial growth and change the soil bacterial community structure while the bacterial abundance remains the same.13 However, there are limited and inconsistent data regarding the effect of NPs on sediment microbial communities. Furthermore, relatively few investigations on the effect of foreign NPs on microbial activities and arsenic release in sediments exist. Therefore, a better understanding of how microorganisms respond to NPs can help address environmental and health concerns that arise from nanomaterial use. Therefore, the purpose of this work is to evaluate the potential of an anaerobic microbial consortium to biologically mobilize As from sediments amended with NPs (i.e., Fe3O4, Fe2O3, and SiO2) at an abandoned sulfur mine in Shimen County, Hunan Province, China.

2. MATERIALS AND METHODS 2.1. Site Characteristics and Sampling. Arsenic-contaminated tailing sediment samples were collected from an abandoned mine area in Shimen, a former realgar mine site in Hunan province, China. This mine contained the largest source of realgar (As4S4) ore resources in Asia and was exploited for approximately 1500 years. However, after the closure, the mine was abandoned without receiving any treatment to minimize environmental impacts. In three villages within the polluted area, soil and river water arsenic levels were 84.17−296.19 mg· kg−1 and 0.5−14.5 mg·L−1, respectively. Moreover, 167 of the 648 residents in the three villages have chronic As poisoning symptoms.14 Tailing sediment samples were carefully collected and stored in sterile polyethylene bags and transported to the laboratory. Some of the samples were air-dried and sieved with a 2 mm mesh for physicochemical characterization; other samples were not dried for microbiological analysis and microcosm set up. To maintain the original environment for the survival of indigenous bacteria, moist tailing sediment samples were stored in polyethylene vinyl containers at 4 °C. 2.2. Elemental Characterization of Tailing Sediment Samples. The main elemental characteristics of the sieved tailing sediments were measured. The carbon, nitrogen, and sulfur contents of the samples were determined using VarioELIII (Elementar, Germany). The total elemental composition of the studied samples was determined using acid digestion in a microwave oven. Each sample (0.25 g) was weighed in Teflon vessels; a mixture of 7.5 mL HNO3, 2.5 mL HCl, and 2 mL HF was added to the vessels. The samples were digested in an MARS Xpress microwave system (CEM) at 190 °C for 15 min.15 Appropriate dilutions were made when required. The final samples were filtered through 0.45-μm membrane filters. Sequential extraction techniques were used to examine the chemical speciation of As fractions in the tailing sediments.15 The sediment As, aluminum (Al), iron (Fe), and manganese (Mn) contents resulting from the total acid digestions and As in the solutions resulting from every extraction step were analyzed using ICP-MS (Agilent 7700x). 7470

dx.doi.org/10.1021/es4055077 | Environ. Sci. Technol. 2014, 48, 7469−7476

Environmental Science & Technology

Article

Table 1. Selected Physical and Chemical Properties of the Sediments Used in the Experiments sample

C (%)

tailing sediment

0.79 exchangeable As(mg·kg1−) 6.41 ± 0.04

N (%)

S (%)

0.10 1.16 reducible As(mg·kg 1− ) 5.87 ± 0.09

T-As (mg·kg1−)

T-Al (g·kg1−)

T-Mn (g·kg1−)

T-Fe (g·kg1−)

425.82 52.86 oxidizable As(mg·kg1−)

1.35 17.52 residual fraction (mg·kg1−)

324.16 ± 11.77

89.39 ± 17.48

T: total.

Figure 1. As and Fe(II) release from sediments amended with different NPs. (a) Fe2O3 NPs; (b) Fe3O4 NPs; (c) SiO2 NPs; (d) Fe(II) release from sediments amended with different NPs. No detectable concentrations of soluble Fe(II) were found in the sediments amended with Fe2O3 and all autoclave controls sediments (X: abiotic controls; NaAc: sodium acetate).

AGAGTTTGATCCTGGCTCAG-3′) and Geobacteraceae specific reverse primer 825R (5′-TACCCGCRACACCTAGT-3′) in the first round of a seminested PCR protocol; F338-GC and R518 were used in the second round. The PCR conditions of the first-round were as previously described.19 2.5. Cloning, Sequence and Phylogenetic Analysis. The purified products were cloned into a pMD18-T vector (TaKaRa, Japan) and transformed into Escherichia coli JM109 competent cells. Subsequently, the cloned 16S rDNA gene fragments were sequenced using Sangon (Shanghai, China). Sequences were analyzed against the NCBI database (http:// www.ncbi.nlm.nih.gov) using the BLAST program and matched to known 16S rRNA gene sequences. In addition to the BLAST searches, the bacterial 16S rRNA gene sequences were analyzed

using the RDP (Ribosomal Database Project) classifier (http:// rdp.cme.msu.edu/). The sequence alignment and phylogenetic analysis were performed using Mega 4.1 software. The phylogenetic tree was constructed in Mega using the neighbor-joining method.20 2.6. Nucleotide Sequence Accession Numbers. Sequences obtained in this study were deposited in the GenBank database under accession numbers JX090253−JX090269 and KC713806−KC713819. 2.7. Quantitative Real-Time PCR. To confirm whether arsenate-respiring bacteria or Fe(III)-reducing bacteria could play a role in arsenic release from the sediments, primers targeting the functional genes involved in the bacterial As(V) respiration (arrA)21,22 and specific to Geobacteraceae species 7471

dx.doi.org/10.1021/es4055077 | Environ. Sci. Technol. 2014, 48, 7469−7476

Environmental Science & Technology

Article

Figure 2. SEM-EDX image of Fe2O3 and Fe3O4 NP precipitation on the sediment surfaces. (a) Fe2O3 NP SEM image; (b) Fe3O4 NP SEM image; (c) Fe2O3 NP EDX spectrum; and (d) Fe3O4 NP EDX spectrum.

494F (5′-AGGAAGCACCGGCTAACTCC−3′) and 825R23,24 were used. The copy numbers of the As(V) respiratory reductase genes (arrA) from the arsenate-respiring bacteria and the16S rRNA gene from the Fe(III)-reducing family Geobacteraceae were determined using real-time PCR (ABI Step One Plus). Assays were set up using the SYBR Premix Ex-Taq Kit (TaKaRa) according to the manufacturer’s instructions. For arrA quantification, the following real-time PCR protocol was applied: 95 °C for 30 s and 40 cycles of 95 °C for 5 s and 61 °C for 30 s. Moreover, for Geobacteraceae quantification, the following procedure was followed: 95 °C for 30 s and 40 cycles of 95 °C for 5 s and 62 °C for 30 s. To generate a standard curve, a single clone containing the correct bacterial 16S rRNA gene was grown in LB medium; thereafter, plasmid DNA was extracted, purified and quantified. A 10-fold dilution series of the plasmid DNA was made to generate a standard curve of bacterial 16S rRNA gene covering 5 orders of magnitude from 1.0 × 104 to 1.0 × 108 copies of the template per assay. Realtime PCR was performed in triplicate; amplification efficiencies of 91.3−103.8% were obtained with R2 values of 0.971−0.996. SYBR Green I assays were always performed together with a melting curve analysis to verify the PCR specificity. 2.8. Characterization of Nanoparticles. For the scanning electron microscope (SEM) characterization, NPs suspended in supernatant and precipitated on the sediments were collected.

Furthermore, SEM observations were performed using an LEO-1530 electron microscope (LEO, Germany). 2.9. Statistical Analysis. Data were analyzed using Excel 2003 and SPSS. Each data point shown in the figures represents an average value. The standard deviations (SD) for the triplicate tests are shown in the figures as error bars. Data were analyzed by analysis of variance (ANOVA) using the SPSS statistical package version 13.0 with a 95% confidence limit.

3. RESULTS AND DISCUSSION 3.1. Chemical Characteristics of Tailing Sediments. The main elemental properties of the tailing sediments sample are summarized in Table 1. High As contents (425.82 mg·kg−1) in the Shimen sediments were much higher than natural background values in this region. Sequential extraction results indicated that As in the sediments occurred primarily in oxidizable species (75%), that is, bound to organic matter or sulfides. Although 1.4% of total As was associated with the Fe fraction, this portion of As mobility is significantly affected by the processes mediating Fe behavior, such as microbially catalyzed Fe reduction. The total organic carbon content in the sediment was 0.79%. 3.2. Roles of NPs in Microbially Mediated As(III) Mobilization. All microcosms (including autoclave controls) had similar arsenic concentrations (approximately 10.0 μg·L−1) 7472

dx.doi.org/10.1021/es4055077 | Environ. Sci. Technol. 2014, 48, 7469−7476

Environmental Science & Technology

Article

NPs. Therefore, the presence of iron oxide NPs during the microbial reduction of As(V) was critical in determining whether reduced As(III) were sorbed and retained in solution. When SiO2 NPs were used in the sodium acetate amendment microcosm experiments, the indigenous bacteria substantially increased the rate of As release into solution under anaerobic conditions. Meanwhile, silicate was able to competitively displace As(III) sorbed on iron oxides. The As bioavailability and mobility were governed by biological and physicochemical factors in sediments amended with SiO2 NPs. Compared with iron oxide NPs, silicate is a potentially effective ligand for inhibiting adsorption or promoting As(III) desorption. We provided the evidence that solid phase SiO2 NPs can generate a few dissolved silicate (SiO44−, 26.2 mg/L, about 0.3% of the total) under acidic conditions (pH = 6.7), but SiO2 Non-NPs cannot (Figure S8). The results indicated that dissolved silicate were able to displace the reduced As(III) through competitive adsorption, which suggested that indigenous bacteria in As-contaminated sediments could increase As mobility from the solid media when organic substrates were supplied, particularly with the addition of SiO2 NPs. The SiO2 NPs present during microbial reduction of As(V) was found to be critical in controlling arsenic mobilization, whether it was reductive dissolution or ligand-promoted mobilization, might be dominantly responsible for the As release. Microbial arsenic reduction plays a very important role in biogeochemical arsenic cycling. It was reported that some metal-reducing bacteria have been isolated from different environments, including sediments, alkaline and saline lakes, and hot springs.27 These anaerobic bacteria, including As(V)respiring bacteria and Fe(III)-reducing bacteria, were thought to be responsible for arsenic release to the environment and increased arsenic mobility. In the present work, the bacteria were enriched from arsenic-rich sediments and can reduce As(V) to As(III) in anaerobic conditions. Additional factors that may add further complications to anion displacement mechanisms from sediments include the predicted sorbed arsenic mobilization by phosphate and silicate. Silicate is commonly present in the environment. Therefore, silicate was able to competitively displace As(III) sorbed on iron oxides, affecting As(III) mobility and potential bioavailability in natural environments, which was a significant mechanism for contaminant sequestration. In this study, the microcosm experiment results suggest that microbial Fe(III) reduction was further enhanced by adding NPs, which demonstrates that nanosized iron oxides strongly enhance respiratory iron reduction by indigenous bacteria. The surface area and mineral structure are recognized as crucial parameters for microbial reduction rates of bulk, macroaggregate iron minerals. However, a large fraction of iron oxide minerals in the subsurface should be present as nanosized colloids.28 The increased reactivity was not only due to the large colloidal aggregate surface areas but also the higher reactivity per unit area. Different particle aggregate sizes might affect iron oxide bioavailability in microbial reduction. Nanosized aggregates appearing in colloidal suspensions might be spatially more accessible for microorganisms than large aggregates flocculating in a bulk phase. In addition, Bosch and colleagues have demonstrated that nanosized iron oxides strongly enhance respiratory iron reduction by G. sulf urreducens.29 Nakamura et al. reported that Shewanellaloihica PV-4 has the ability to self-organize an electrically conductive network using semiconductive minerals as a long-distance electron

and no detectable concentrations of soluble Fe(II) in the aqueous phases during the first incubation day. However, the indigenous bacteria substantially increased the rate of As release into solution under anaerobic conditions compared with abiotic controls after sodium acetate was used as a carbon source in the microcosm experiments. Aqueous Fe(II) and As(III) reached concentrations of 3487.6 μg·L−1 (SI Figure S1) and 2338.0 μg· L−1, respectively, after 7 weeks of incubation. Meanwhile, As(III) and Fe(II) did not substantially increase in the sterilized samples (Figure 1), demonstrating that this process was biologically mediated. Potential effects of the NPs on the enhancement of As released from the sediments with additional organic substrates were investigated in batch experiments. After adding sodium acetate as a carbon source, sediments amended with Fe2O3, Fe3O4 and SiO2 NPs incubated under anaerobic sterile controls exhibited a negligible reduction in Fe(III) and As(V) from the sediments with time; the maximum concentrations of As(V) were 20.9 μg·L−1, 20.3 μg·L−1 and 42.1 μg·L−1, respectively (SI Figure S2). The results indicated that As release through desorption from the sediment surface was not an important factor. In contrast, the maximum As contents in the anaerobic samples (nonsterilized) amended with Fe2O3, Fe3O4 and SiO2 NPs were much higher, up to 523.3 μg·L−1, 859.1 μg·L−1 and 6266.0 μg·L−1, respectively, existing predominantly as As(III) (SI Figures S3 and S4). However, the maximum As(III) concentrations in the sediments amended with Fe2O3, Fe3O4, and SiO2 Non-NPs were 23.3 μg·L−1, 763.6 μg·L−1, and 1430.0 μg·L−1, respectively, suggesting that the NP effects on As release from sediments was more important than Non-NP effects. More importantly, SiO2 NP additions to the sediments resulted in a significant increase in As(III) mobilization in the presence of sodium acetate (Figure 1c). Meanwhile, during anaerobic incubation, nearly 4 times the Fe(II) (11.67−12.87 mg/L) (Figure 1d) from sediments amended with NPs and sodium acetate was released compared to sediments amended with only sodium acetate (3.49 mg/L) (SI Figure S1), indicating that Fe(III) reduction was further enhanced by adding NPs in the presence of sodium acetate. Specialist dissimilatory metal-reducing bacteria (e.g., Geobacter, Anaeromyxobacter, Clostridium) were detected by genetic profiling, which have been shown to reduce a wide range of high valence metals with the notable exception of As(V).6,25 These results implied that addition of NPs could potentially stimulate bacterial growth and resulted in a marked stimulation in the rate of Fe(III) and As(V) reduction. Fe2O3 and Fe3O4 NP additions decreased As(III) mobilization compared with no iron additions (Figure 1a and b). The results suggest that the iron oxide NP additions, which are potential sorbents for As(III), caused a decrease in arsenic solution concentrations. Numerous studies have demonstrated that iron oxide NPs have a high affinity for the adsorption of As(III) and As(V).25,26 The undisturbed samples were placed a month after sampling, there was a layer of black precipitate between the solid/solution interfaces amended with Fe2O3 and Fe3O4 NPs, but no precipitate on the sediments amended with SiO2 NPs (SI Figures S5−S7). The valuable information could explain that Fe2O3 and Fe3O4 NP additions decreased As(III) concentration in solution. The SEM-EDX was used to investigate black precipitate. As shown in Figure 2. There were lots of NPs in the black precipitate. The Fe, O, and As signals were included in the EDX spectrum (Figure 2b and d). The results could confirm that As was absorbed by iron oxide 7473

dx.doi.org/10.1021/es4055077 | Environ. Sci. Technol. 2014, 48, 7469−7476

Environmental Science & Technology

Article

transfer conduit.30 These pure-culture studies have shown that dissimilatory metal-reducing bacteria are able to utilize ironoxide NPs as electron conduits for reducing distant terminal acceptors. However, the ecological relevance of this energy metabolism has been poorly understood. Kato et al. reported that iron-oxide NPs could be used as electron conduits for facilitating respiration in soil microbial populations, particularly ́ Geobacter species.11 Furthermore, Dominguez-Garay et al. reported that silica-supplemented soil better utilized the electron donor, either acetate or natural rice root exudates, in electrogenic microbial populations.12 Therefore, adding NPs strongly enhance respiratory As(V) and Fe(III) reduction through better utilized the electron donor such as acetate and natural organic matter by indigenous bacteria in sediments. 3.3. Microbial Community Composition and As Release. The PCR-DGGE procedure is sensitive when analyzing microbial community compositions in soils amended with NPs.31 The bacterial community compositions for sediments amended with NPs, Non-NPs, and NPs with sodium acetate were determined and are shown in SI Figure S9a, In contrast to the control sample, the DGGE results indicate that the number of bands from the amended samples increased substantially. A visual inspection demonstrates that the sediment bacterial community structure changed for NP additions, suggesting that the shifts in bacterial community structure were induced by NPs treatment. In addition, the phylogenetic tree (SI Figure S9b) illustrates that these representative bands were affiliated with known members of Firmicutes and α- and γ-proteobacteria (SI Table S1). To obtain a more specific picture of Geobacteraceae diversity, DGGE and phylogenetic analysis of cloned Geobacteraceae 16S rRNA genes were performed (SI Figure S10a, b and Table S2). Specialist dissimilatory metal-reducing bacteria (e.g., Geobacter, Anaeromyxobacter, Clostridium) were detected, which have been shown to reduce a wide range of high valence metals with the notable exception of As(V).6,27 Geobacteraceae of the δproteobacteria were widely distributed in many metal-reducing environments.32 Fe(III)-reducing microorganisms, most notably those in the Geobacteraceae family, could play an important role in the bioremediation of subsurface environments containing organic or metal contaminants. However, detailed information regarding the relationship between the Geobacter community composition and environmental condition changes, for example, NP additions, is scarce. The results show that the Geobacteraceae community composition was affected by introduced chemicals in As-polluted sediments. Differences in the Fe(III) reduction and As(III) release profiles could be explained in terms of the contrasting microbial communities. 3.4. Quantitative PCR Analysis. To further investigate the effects of Fe2O3, Fe3O4, and SiO2 NPs on the abundance of important sediment bacteria, real-time PCR assays were used to determine the abundance of Fe(III)- and As(V)-reducing bacteria after NP exposure. Typically, arrA genes are widely used as functional markers for As(V)-respiring bacteria;7 the quantitative changes in As(V)-reducing bacteria can be evaluated by measuring the arrA gene copy numbers. The 16S rRNA gene of the Fe(III)-reducing bacterial family Geobacteraceae and the dissimilatory As(V)-reducing gene arrA of As(V)-respiring bacteria were determined (Figure 3a and b). The Geobacteraceae and arrA gene copy numbers increased from 2.6 × 103 and 3.5 × 105 copies·g−1 sediment to 3.3 × 104 and 7.1 × 105 copies·g−1 sediment, respectively, after NPs amendments. These genetic analyses provide molecular

Figure 3. (a) Geobacteraceae and (b) arrA gene copy numbers in tailing sediments amended with NPs.

evidence that not only there exist anaerobic microbial consortium including As(V)-respiring bacteria and Fe(III)reducing bacteria, but the SiO2, Fe2O3, and Fe3O4 NPs used favor the growth of metal-reducing sediment bacteria. It was shown that the main bacterial mechanism was via the simultaneously dissimilatory reduction of Fe(III) and As(V) by anaerobic microbial consortium. Although there was no direct evidence of a decoupling of aqueous Fe(II) and As(III) release. 3.5. The As Release Mechanisms. From the above discussion, a presumed mechanism for As release from sediments by biotic and abiotic interactions is shown in Figure 4. Both As redox states strongly sorb to mineral surfaces in

Figure 4. Schematic illustration of As mobilization mechanisms. 7474

dx.doi.org/10.1021/es4055077 | Environ. Sci. Technol. 2014, 48, 7469−7476

Environmental Science & Technology

Article

(7) Malasarn, D.; Saltikov, C.; Campbell, K.; Santini, J.; Hering, J.; Newman, D. arrA is a reliable marker for As (V) respiration. Science 2004, 306 (5695), 455−455. (8) Gao, Y.; Luo, Z.; He, N.; Wang, M. K. Metallic nanoparticle production and consumption in China between 2000 and 2010 and associative aquatic environmental risk assessment. J. Nanopart. Res. 2013, 15 (6), 1−9. (9) Christian, P.; Von der Kammer, F.; Baalousha, M.; Hofmann, T. Nanoparticles: Structure, properties, preparation and behaviour in environmental media. Ecotoxicology 2008, 17 (5), 326−343. (10) Li, R.; Stroud, J.; Ma, J.; McGrath, S.; Zhao, F. Mitigation of arsenic accumulation in rice with water management and silicon fertilization. Environ. Sci. Technol. 2009, 43 (10), 3778−3783. (11) Kato, S.; Nakamura, R.; Kai, F.; Watanabe, K.; Hashimoto, K. Respiratory interactions of soil bacteria with (semi)conductive ironoxide minerals. Environ. Microbiol. 2010, 12 (12), 3114−3123. (12) Domiguez-Garay, A.; Berná, A.; Ortiz-Bernad, I.; Esteve-Núñez, A. Silica colloid formation enhances performance of sediment microbial fuel cells in a low conductivity soil. Environ. Sci. Technol. 2013, 47 (4), 2117−2122. (13) He, S.; Feng, Y.; Ren, H.; Zhang, Y.; Gu, N.; Lin, X. The impact of iron oxide magnetic nanoparticles on the soil bacterial community. J. Soil Sed. 2011, 11 (8), 1408−1417. (14) Lei, M.; Zeng, M.; Zheng, Y.; Liao, B.; Zhu, Y. Heavy metals pollution and potential ecological risk in paddy soils around mine areas and smelting areas in Hunan Province. Environ. Sci. China 2008, 28 (6), 1212−1220. (15) Larios, R.; Fernandez-Martinez, R.; Rucandio, I. Comparison of three sequential extraction procedures for fractionation of arsenic from highly polluted mining sediments. Anal. Bioanal. Chem. 2012, 402 (9), 2909−2921. (16) Viollier, E.; Inglett, P.; Hunter, K.; Roychoudhury, A.; Van Cappellen, P. The ferrozine method revisited: Fe (II)/Fe (III) determination in natural waters. Appl. Geochem. 2000, 15 (6), 785− 790. (17) Hou, J.; Gong, Z.; Guo, X.; Huang, B. Direct determination of As(III) and As(V) in seawater with flow injection hydride generation non-dispersion atomic fluorescence spectrometry. Chinese Journal of Analysis Laboratory 2000, 19 (6), 13−16. (18) Liu, Z.; Wang, Y.; He, N.; Huang, J.; Zhu, K.; Shao, W.; Wang, H.; Yuan, W.; Li, Q. Optimization of polyhydroxybutyrate (PHB) production by excess activated sludge and microbial community analysis. J. hazard. mater. 2011, 185 (1), 8−16. (19) Lin, B.; Westerhoff, H. V.; Roling, W. F. M. How Geobacteraceae may dominate subsurface biodegradation: Physiology of Geobacter metallireducens in slow-growth habitat-simulating retentostats. Environ. Microbiol. 2009, 11 (9), 2425−2433. (20) Dong, G.; Wang, Y.; Gong, L.; Wang, M.; Wang, H.; He, N.; Zheng, Y.; Li, Q. Formation of soluble Cr(III) end-products and nanoparticles during Cr(VI) reduction by Bacillus cereus strain XMCr6. Biochem. Eng. J. 2012, 70, 166−172. (21) Saltikov, C. W.; Wildman, R. A.; Newman, D. K. Expression dynamics of arsenic respiration and detoxification in Shewanella sp. strain ANA-3. J. Bacteriol. 2005, 187 (21), 7390−7396. (22) Lear, G.; Song, B.; Gault, A.; Polya, D.; Lloyd, J. Molecular analysis of arsenate-reducing bacteria within Cambodian sediments following amendment with acetate. Appl. Environ. Microbiol. 2007, 73 (4), 1041−1048. (23) Luna, G. M.; Dell’Anno, A.; Corinaldesi, C.; Armeni, M.; Danovaro, R. Diversity and spatial distribution of metal reducing bacterial assemblages in groundwaters of different redox conditions. Int. Microbiol. 2010, 12 (3), 153−159. (24) Schippers, A.; Neretin, L. N. Quantification of microbial communities in near-surface and deeply buried marine sediments on the Peru continental margin using real-time PCR. Environ. Microbiol. 2006, 8 (7), 1251−1260. (25) Tuutijärvi, T.; Lu, J.; Sillanpäa,̈ M.; Chen, G. As (V) adsorption on maghemite nanoparticles. J. Hazard. Mater. 2009, 166 (2), 1415− 1420.

sediment. Adding sodium acetate as the electron donor clearly promoted anaerobic bacterial growth and resulted in the release of Fe(II) and As(III). In the presence of dissolved organic matter (DOM), NPs-supplemented sediments better utilized the electron donor, either acetate or natural organic carbon, in anaerobic microbial populations and markedly stimulated the rate of Fe(III) and As(V) reduction. In addition, in the presence of Fe(III) oxide NPs, microbial reduced As(III) is adsorbed on Fe(III) oxide NPs and then precipitated on the solid−liquid interface. However, in the presence of SiO2 NPs, the As(III) solution concentrations increase because of producing silicate and competitive adsorption. Silicate competes with As for sorption sites at mineral surfaces, potentially increasing As mobility but also favoring As desorption from Fe(III) minerals.

■

ASSOCIATED CONTENT

S Supporting Information *

Tables S1−S2 and Figures S1−S10. This material is available free of charge via the Internet at http://pubs.acs.org.

■

AUTHOR INFORMATION

Corresponding Authors

*(Y.W.) Phone: +86 592 2185495; fax: +86 592 2184822; email: [email protected] *(Q.L.) Phone: +86 592 2189595; fax: +86 592 2184822; email: [email protected]. Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS This work was supported by the National Basic Research Program of China (2013CB733505), the National Natural Science Foundation of China (41271260, 41276101, 41071302, and 41301346), the Program for New Century Excellent Talents in University (NCET-12-0326), the Development and Reform Commission of Fujian Province, China (2011-1598) and the Science and Technology Program of Xiamen of Fujian Province, China (3502Z20126005).

■

REFERENCES

(1) Koo, N.; Lee, S. H.; Kim, J. G. Arsenic mobility in the amended mine tailings and its impact on soil enzyme activity. Environ. Geochem. Health 2012, 34 (3), 337−348. (2) Garcia-Sanchez, A.; Alonso-Rojo, P.; Santos-Frances, F. Distribution and mobility of arsenic in soils of a mining area (Western Spain). Sci. Total Environ. 2010, 408 (19), 4194−4201. (3) Macur, R. E.; Wheeler, J. T.; McDermott, T. R.; Inskeep, W. P. Microbial populations associated with the reduction and enhanced mobilization of arsenic in mine tailings. Environ. Sci. Technol. 2001, 35 (18), 3676−3682. (4) Dhar, R. K.; Zheng, Y.; Saltikov, C. W.; Radloff, K. A.; Mailloux, B. J.; Ahmed, K. M.; van Geen, A. Microbes enhance mobility of arsenic in Pleistocene aquifer sand from Bangladesh. Environ. Sci. Technol. 2011, 45 (7), 2648−2654. (5) Lee, K. Y.; Kim, K. W.; Kim, S. O. Geochemical and microbial effects on the mobilization of arsenic in mine tailing soils. Environ. Geochem. Health 2010, 32 (1), 31−44. (6) Islam, F. S.; Gault, A. G.; Boothman, C.; Polya, D. A.; Charnock, J. M.; Chatterjee, D.; Lloyd, J. R. Role of metal-reducing bacteria in arsenic release from Bengal delta sediments. Nature 2004, 430 (6995), 68−71. 7475

dx.doi.org/10.1021/es4055077 | Environ. Sci. Technol. 2014, 48, 7469−7476

Environmental Science & Technology

Article

(26) Mayo, J.; Yavuz, C.; Yean, S.; Cong, L.; Shipley, H.; Yu, W.; Falkner, J.; Kan, A.; Tomson, M.; Colvin, V. The effect of nanocrystalline magnetite size on arsenic removal. Sci. Technol. Adv. Mater. 2007, 8 (1), 71−75. (27) Oremland, R. S.; Stolz, J. F. The ecology of arsenic. Science 2003, 300 (5621), 939−944. (28) Waychunas, G. A.; Kim, C. S.; Banfield, J. F. Nanoparticulate iron oxide minerals in soils and sediments: Unique properties and contaminant scavenging mechanisms. J. Nanopart. Res. 2005, 7 (4−5), 409−433. (29) Bosch, J.; Heister, K.; Hofmann, T.; Meckenstock, R. U. Nanosized iron oxide colloids strongly enhance microbial iron reduction. Appl. Environ. Microbiol. 2010, 76 (1), 184−189. (30) Nakamura, R.; Kai, F.; Okamoto, A.; Newton, G. J.; Hashimoto, K. Self-constructed electrically conductive bacterial networks. Angew. Chem., Int. Ed. 2009, 48 (3), 508−511. (31) Nogueira, V.; Lopes, I.; Rocha-Santos, T.; Santos, A. L.; Rasteiro, G. M.; Antunes, F.; Gonçalves, F.; Soares, A. M.; Cunha, A.; Almeida, A. Impact of organic and inorganic nanomaterials in the soil microbial community structure. Sci. Total Environ. 2012, 424, 344− 350. (32) Lovley, D. R.; Holmes, D. E.; Nevin, K. P. Dissimilatory Fe(III) and Mn(IV) reduction. Adv. Microb. Physiol. 2004, 49, 219−286.

7476

dx.doi.org/10.1021/es4055077 | Environ. Sci. Technol. 2014, 48, 7469−7476