Environ. Sci. Technol. 2009, 43, 4530–4536
Impact of Silver Nanoparticle Contamination on the Genetic Diversity of Natural Bacterial Assemblages in Estuarine Sediments ADAM BRADFORD,† RICHARD D. HANDY,‡ JAMES W. READMAN,† ANDREW ATFIELD,‡ AND ¨ H L I N G * ,† MARTIN MU Plymouth Marine Laboratory, Prospect Place, The Hoe, Plymouth PL1 3DH, U.K., and School of Biological Sciences, University of Plymouth, Plymouth, PL4 8AA, U.K.
Received January 20, 2009. Revised manuscript received April 12, 2009. Accepted April 14, 2009.
Recent advancements in the nanotechnology industry have seen a growing interest in integrating silver nanoparticles (AgNPs) into consumer and medical products. To date, there has been little research into the toxicological impact that AgNPs will have when they enter the marine realm. Of particular concern are the possible effects of Ag-NPs on natural bacterial assemblages, given the antimicrobial activity of silver. In this study, estuarine sediment samples were dosed in triplicate for 20 days from a stock solution of Ag-NPs, with a final cumulative treatment of either 0 µg L-1 (control), 25 µg L-1 or 1000 µg L-1. The experimental tanks were left for a further 10 days to allow for any recovery. Inductively coupled plasma-optical emission spectrometery (ICP-OES) of water and sediment samples confirmed that the Ag-NPs concentration in the aqueous phase decreased after each dosing and were transported to accumulate in the surface layer of the sediment (∼top 3 mm). The overall concentration of AgNPs in the water column, however, increased steadily during the 20 days of dosing but decreased rapidly during the following 10 days without dosing. Nevertheless, the AgNPs did not have any impact on the prokaryotic abundance in the water column over the incubation period (ANOVA, P < 0.05). Environmental DNA was extracted from sediment samples and a two-step nested PCR-denaturing gradient gel electrophoresis (DGGE) approach, using PCR primers specific to the phylum Bacteria, was adopted to assess their diversity. Multivariate statistical analyses of presence/ absence matrices produced from DGGE profiles revealed negligible differences in bacterial diversity between treatments, suggesting that, under the selected experimental regime, AgNPs present little or no impact on estuarine sediment bacterial diversity. Possible reasons for this could include environmental factors, in particular the chloride ions in estuary water affecting the chemistry and behavior of Ag-NPs.
* Correspondence author e-mail:
[email protected]. † Plymouth Marine Laboratory. ‡ University of Plymouth. 4530
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Introduction Nanotechnology is a novel field of science and engineering which utilizes engineered nanoparticles (ENPs). ENPs can be broadly defined as having at least one dimension between 1 and 100 nm (1). ENPs often display unique physicochemical properties and reactivities due to their small size and homogeneous composition, structure, or surface composition (2). These unique properties are not often present at the larger scale, and consequently govern the increasing commercial interest in ENPs (2). The global nanotechnology market is currently growing at an exponential rate and is estimated to be worth $1 trillion by 2015, a huge increase from the current market value of $10.5 billion (2, 3). However, while the rise in the nanotechnology industries’ share of the global market is likely to result in an increase of the environmental release of ENPs, little or nothing is known about the environmental impact caused by ENPs released into the environment, and this is of growing concern among the scientific and regulatory community (2, 4-8). One group of ENPs increasingly receiving attention are silver nanoparticles (Ag-NPs). Silver has been widely accepted to have antibacterial properties, which have been demonstrated in studies with different species of bacteria (8, 9). These findings are mainly concluded from laboratory-based studies of cultures treated with Ag-NPs (8, 10-13). For example, Cho et al. (12) observed significant inhibition of the gram-negative and gram-positive species of bacteria, Escherichia coli and Staphylococcus aureus, at concentrations exceeding 5 µg mL-1 and 10 µg mL-1, respectively. Complete bacterial inhibition has also been demonstrated with E. coli, Bacillus subtilis, and S. aureus, with concentrations of 50, 70, and 50 µg mL-1, respectively (8, 11, 12). However, the exact antibacterial mechanism of Ag-NPs is not fully understood (10). Possible antibacterial mechanisms of Ag-NPs have been speculated to be broadly similar to that of silver ions (Ag+) (10), but Ag and AgCl were also shown to have antimicrobial effects (14). Reports on the antibacterial mechanism of Ag+ have shown that upon exposure DNA loses its ability to replicate and express ribosomal subunit proteins, along with potentially causing the deactivation of other cellular proteins and enzymes essential to the production of ATP (15, 16). It is believed that upon contact with a bacterial cell Ag+ are released from the surface of the NP, which in turn interact with the elements of bacterial membranes and inhibit respiratory enzymes. This, as well as Ag and AgCl (14), in turn facilitates the generation of reactive oxygen species that cause structural changes to the cells, which then leads to the dissipation of the proton motive force and eventually cell death (8, 10, 17). This observation is in agreement with the recent electron microscopic analysis that showed that AgNPs interact with bacterial membranes, causing structural changes that ultimately cause cell death (8). However, while such studies are useful in validating a toxic effect of a chemical, they do not take environmental physiochemical factors into consideration. Consequently, the understanding of the potential impacts of Ag-NPs on the bacteria in the environment is poor. The lack of this scientific knowledge coupled with the increased use of Ag-NPs in manufactured goods, has led to an increasing concern among the scientific community of the potential environmental impact Ag-NPs present. In particular, there is great apprehension about the loss of vital bacteria that are essential in numerous activities that are important in the support and continuation of all forms of life, such as the degradation of organic matter, transformation 10.1021/es9001949 CCC: $40.75
2009 American Chemical Society
Published on Web 04/29/2009
of elements, and recycling of nutrients (18). The most probable environmental exposure route of Ag-NPs is predicted to be through wastewaters and surface waters (6, 19). A recent study by Benn and Westerhoff (20) observed that Ag-NPs can be readily released from commercially available sock fabrics previously impregnated with Ag-NPs, a treatment used to, for example, protect fabrics against odor-producing bacteria. The study demonstrated that up to 1360 µg Ag-NPs could be absorbed in 1 g of sock fabric which then could leach as much as 650 µg of silver in 500 mL of distilled water during a 24-h washing cycle. Other potential sources of AgNPs in the aquatic environment have been identified to come from the use or degradation of the following products: shampoo, soap, toothpaste, paints, clothing, cosmetics, paints, washing powders, drinks, plastic food container, wound dressings, and biocidal coatings (20, 21). Despite this, the utilization of Ag-NPs is fast increasing. Currently there are no legally enforcable standards for the release of NPs in general into the environment, mainly due to the lack of an understanding of their potentially toxic effects. This is particularly of concern in the case of nano-Ag, given its antimicrobial activity. Therefore, we decided to investigate the potential impact of nano-Ag on the genetic diversity of natural bacterial assemblages. In England and Wales, it has been estimated that approximately 37.5% of the total sewage treatment plant effluents are discharged into estuarine and coastal environments (22). Once released into the coastal environment, Ag-NPs are likely to precipitate onto the surface of sediment due to the high density of Ag. Based on this assumption, we designed a 30-day microcosm study in which we added Ag-NPs to estuary water overlying estuarine sediment cores. Changes in bacterial diversity in the sediment were investigated using denaturing gradient gel electrophoresis (DGGE), while overall prokaryotic abundance in the overlaying seawater was determined by flow cytometry.
Material and Methods Aqueous Ag-NPs Preparation and Dosing. The Ag-NP test material (Sigma Aldrich Silver nano; < 100 nm) was selected as the Ag-NPs were representative in size to those used by Yoon et al. (11) and Pal et al. (10). A stock solution of 100 mg L-1 Ag-NPs was prepared in a pyrex bottle using Millipore water (ion free) and sonicated in a Sonicleaner bath (Lucas Dawe Ultrasonics) for 5 h. The Ag-NP stock solution was sonicated for a further 30 min prior to each addition to the experimental tanks. Dispersion in stock solutions was tested by transmission electron microscopy (TEM, JEOL 1200EX II) and spectral scans (Lambda Bio 20 UV/vis Spectrometer, Perkin-Elmer). The spectral scan of the sonicated Ag-NPs showed a well-defined plasmon band at 420 nm (Supporting Information (SI) Figure S1a), which has been shown to be characteristic of Ag-NPs (8, 10). TEM revealed that the sonicated Ag-NPs stock solution was well dispersed (SI Figure S1b) and that the individual particles (if assumed to be spheroidal) had an average diameter of 58.6 nm (standard deviation (SD) ) 18.6 nm; n ) 64). Environmental Sample Collection. Estuarine sediment samples were collected from an intertidal area at the mouth of the Tamar Estuary in Plymouth Sound (“St John’s Lake” mud flats, OSGB grid ref SX412539). Sediment samples were collected by placing a standard core (L 28 cm × W 17 cm × D 7 cm) into the sediment, then carefully cutting the sediment slab out, and sliding a Perspex sheet underneath the core. Each core (with the Perspex sheet underneath) was lowered slowly onto the bottom of an experimental tank (L 39 cm × W 27 cm × D 32 cm) which contained 20 L of estuary water taken from the sampling site from the previous high tide to sediment sampling. Care was taken to ensure that there was minimal disturbance of the surface layer and stratification
of the sediment, and that any large invertebrates were avoided during collection. Sediment samples were approximately 3.8 ( 0.2 kg (mean ( S.E.M, n ) 9 sediment cores) in weight. Samples were allocated randomly to treatments to minimize any potential sampling bias. Experimental Design. Estuarine sediment samples were exposed in triplicate batch tanks to one of the following three treatments: 0 µg L-1 of Ag-NPs in overlying estuary water (control; tanks T01, T02, T03), 25 µg L-1 (T04, T05, T06), and 1000 µg L-1 (T07, T08, T09) Ag-NPs. The 25 µg L-1 concentration was chosen as it represents the lower end of the detection limits while the 1000 µg L-1 concentration was selected as it is much higher than predicted future environmental concentrations (19). Therefore, if no effect on the bacterial diversity was to be seen at this concentration then it can be assumed that Ag-NPs do not affect bacterial diversity in the natural estuarine sediment under the experimental conditions. To achieve a final concentration of 0 µg L-1 (Control), 25 µg L-1, and 1000 µg L-1, the experimental tanks were given a cumulative dose of Ag-NPs over 20 days, with daily doses of 1/20th of the final concentration. After addition of each daily dose of Ag-NPs, the experimental tanks were stirred slowly for 5 s to aid the aeration system in dispersion of the NPs. The experimental tanks were left for a further 10 days (i.e., 30 days in total) without dosing with Ag-NPs. All glass and plastic wear, including the equipment used for the tank set-ups and for sampling, were thoroughly acid washed (0.5% nitric acid, overnight) to avoid any contamination with trace metals. Glass tanks were always kept covered, aerated, and in a constant-temperature culture room at 15 °C, operated on a 12 h light:12 h dark cycle. Light was provided by basic white fluorescent tubes (Osram 36W) irradiating the tanks with an average of 2.2 ( 0.8 (n ) 9) µmol photons m-2 s-1 photosynthetic active radiation (PAR). Each tank consisted of one sediment sample and 20 L of overlying water. Sediment subsamples were taken randomly from each of the sediment cores prior to the addition of the daily doses on days 0, 1, 4, 10, 15, 20, 25, 30 and stored at -80 °C until further analysis. This was done by inserting a 5 mL syringe casing (without the plunger) into the sediment, to remove a representative core sample of sediment. A plastic bung was then placed into the resulting hole in the sediment in order to identify where previous samples were taken, and to prevent subsidence. Water samples (10 mL) were also collected at each sampling point for metal analysis before and immediately after the dosing with Ag-NPs. Further 25 mL water samples were taken for nutrient analysis. Nutrient concentrations (phosphate, nitrite, nitrate, silicate, ammonium) were determined with a Technicon segmented flow colorimetric autoanalyser using the method described by Woodward (23). Additionally, water quality parameters were measured on the eight sampling days: salinity (using a WTW LF197 probe WTW GmbH, Weilheim, Germany), dissolved oxygen (Oxygen meter, Model 781, Strathkelvin Instruments, Glasgow, UK), pH (pH meter AR15, Fisher Scientific, Loughborough, UK) and temperature. Metal Analysis. Trace metal concentrations in sediment samples in aqua regia digests and acidified water samples were analyzed by inductively coupled plasma-optical mass spectrometery (ICP-OES) using a Varian 725-ES spectrometer (Varian, Inc., Palo Alto, CA). The instrument settings were calibrated using mixed standards prepared in Milli-Q water with a known concentration of yttrium (Y (24)) to determine instrumental drift and variations in plasma conditions, thus allowing to correct subsequent measurements. Samples were analyzed for the following elements: Ag, Ca, Cu, Fe, K, Mg, Na, Zn. The 10-mL water samples taken before and after Ag-NP dosing were acidified with 50 µL of trace analysis grade nitric acid (trace analysis grade, Fisher). Sediment samples from VOL. 43, NO. 12, 2009 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
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each of the nine experimental tanks were dissected and analyzed by depth. A sediment slice of ∼3 mm in depth was removed at the surface, 10 and 20 mm down the core. The aliquots were air-dried (60 °C for 12 h) and weighed. Samples were resuspended in 4 mL of aqua regia (three parts HCl to one part HNO3) and incubated at 70 °C for 2 h (24), followed by dilution with Milli-Q water to 18 mL. A certified estuarine reference material (LGC 6137, Laboratory of the Government Chemist) was treated likewise in order to evaluate the efficacy of this extraction. Results for the estuarine sediment reference material (LGC 6137) were within 10% of the expected range of certified values, thus validating the measurements taken in this study. Flow Cytometric Anaylsis. The prokaryotic abundance in the water column of each of the experimental tanks was determined using a Becton Dickenson flow cytometer (FACSort). Bacterial cells in 10-mL subsamples of the experimental tanks were fixed with 1% glutaraldehyde (final concentration), and stored at -20 °C until analysis. An aliquot of each sample was stained with the nucleic acid stain SYBR Green-I. Fluorescent beads (3.6 µm diameter) were added to each sample to normalize cell fluorescence and light scatter values. Clades were defined on cytogram plots of side scatter versus green fluorescence to define high nucleic acid (HNA) and low nucleic acid (LNA) cells. DNA Extraction. Genomic DNA was isolated from sediment samples based on the method described by Smith et al. (25). In brief, this involved the surface layer being removed from the frozen sample and aliquoted into approximately 300-mg quantities. The samples were resuspended in a mix of 0.5 mL of 0.1 M Na2HPO4 (pH 8), 0.5 mL of phenol: chloroform:isoamyl alcohol (25:24:1) and 0.5 g of 106 µm and 0.1 g of 212-300 µm acid washed glass beads (Sigma, UK) using a vortex mixer (MO BIO laboratories, Vortex-Genie 2, 30 s pulse). After centrifugation for 5 min at 16.1g at 4 °C genomic DNA in the supernatant was purified by subsequent treatments with equal volumes of phenol:chloroform:isoamyl alcohol (25:24:1) and chloroform:isoamyl alcohol (25:24) followed by precipitation with 2 volumes ethanol and 1 volume 10 M sodium acetate overnight. The DNA was pelleted by centrifugation at 16.1g for 20 min at 4 °C, and washed twice with 200 µL of ice-cold 70% (v/v) ethanol. Further DNA purification was achieved using a modified method described by Murray and Thompson (26), which is described in detail in SI. PCR-DGGE. Bacterial diversity was assessed using a nested PCR-DGGE approach as described by Dar et al. (27) with modifications introduced by Mu ¨ hling et al. (28). In essence, 1500-bp fragments of the 16S rRNA gene were amplified from the environmental DNA using the Bacteria-specific primer 9bfm and the universal primer 1512uR (28). Aliquots of these PCR products were used as templates for reamplification with the nested Bacteria-specific primer pair 341f-GC/518r (27-29). This approach resulted in the explicit amplification of 16S rRNA gene fragments of members of the phylum Bacteria. DGGE analysis of the 16S rRNA gene fragments was based on that of Muyzer et al. (29) and Mu ¨ hling et al. (28). Statistical Analysis. Results obtained from ICP-OES and flow cytometery were analyzed using Mini-tab 15. As no tank effects were observed within treatments, data were pooled by treatment for statistical analysis. Data were checked for kurtosis, skewedness, and unequal variance (Bartlett’s test), subsequent to data being analyzed by one-way analysis of variance (ANOVA), with treatment as a factor. Differences at the 99% of silver has been reported from seawater onto aggregated particles (33). Additionally, organic, humic acid, carboxyl, phenolic, amino, and sulfidic ligand interactions can occur (33, 34). It has been shown that salinity can also lead to an increase in aggregation of Ag-NPs with a concurrent loss in antibacterial activity (35). Also, it is generally accepted that silver in seawater results in the formation of free ionic silver and chloro-complexes (Ag+, AgCl°, AgCl2-, and AgCl32-) which, compositionally, are regulated by salinity. Of these species, the free Ag+ cation is generally responsible for most of the antimicrobial properties of Ag (36, 37). At a salinity of 25‰, Ag+, AgCl°, AgCl2-, and AgCl32- constitute 0.004, 1.4, 49, and 49%, respectively (38), though this was determined using AgNO3 rather than Ag-NPs. Thus, at this salinity, the highly toxic free ion contributes less than one thousandth of the dosed concentration. Although we did not determine the chemical transformation of the Ag-NPs in the experimental tanks we believe that Ag cation formation at the surface of the Ag-NPs, aggregation reactions, complexation, speciation and partitioning afford the potential to regulate the toxicological impact of AgNPs in estuary water. Potential Impact of Environmental Release of Ag-NPs. The antibacterial properties of Ag-NPs are well demonstrated in the literature (8, 10-13). However, the test concentrations that have been used in previous studies were much higher (in the region of 5-100 mg L-1, e.g., refs 8, 10-13) than those used in this study (0.025 mg L-1 and 1 mg L-1) and were added to the bacterial culture in one single dose. Furthermore, a 100% inhibition of growth was noted at concentrations exceeding 50 mg L-1 (8, 10). Although this represents an unrealistic scenario in terms of environmentally relevant exposures, it is routinely used to demonstrate the antibacterial potential of Ag-NPs (8, 10-13). A report by Boxall et al. (19) estimated market penetration data of products incorporating ENPs with the aim to predict ENP exposure routes and to quantify realistic environmental concentrations. With regard to Ag-NP products (biocidal coatings, shampoo, soap and toothpaste) estimates for relevant environmental water concentrations for 10, 50, and 100% market penetration were 0.010, 0.051, and 0.10 µg L-1, respectively. Present market penetration is considered to be less than 10%, though this is likely to increase due to the ever more widespread use and development of Ag-NP based products. Relevant soil concentrations were estimated to be
an order of magnitude higher, with 100% market penetration predicted to be 4.26 µg kg-1 soil. Both of these estimates for 100% market penetration are 5 orders of magnitude less than the lowest concentrations noted to have an inhibitory effect (8, 10-13), and are an order of magnitude less than the lower (i.e., 25 µg L-1) concentration used in our exposure experiment. Based on the results of the present study, the current and future predicted environmental concentrations of AgNPs appear to be well below any impact threshold to the microbial health of the environment. In conclusion, this study provides one of the first insights into how Ag-NPs might affect natural bacterial assemblages. Ag-NPs clearly have antibacterial properties which have been well documented for cultures (8, 10-13). However, the conclusion from our study is that Ag-NPs have a negligible impact on bacterial diversity in estuarine sediments. Notably, the data from this study indicate that Ag-NPs will be transported out of the aqueous phase, possibly through formation of AgCl complexes and aggregation. Although we did not elucidate the precise nature of this pathway or whether the size of the Ag-NPs is altered by the exposure to saline estuary water, we were able to demonstrate that Ag accumulated upon the sediment surfaces. However, these results indicate that physicochemical parameters in the brackish environment (such as salinity and organic matter content) might render Ag-NPs less toxic and unreactive within a short time frame of their release. Overall, the findings question the general concept that Ag-NPs pose a threat to bacteria in the natural environment, though this is based on a 30-day exposure period with samples from one estuary and a longer exposure study or application to different estuarine environments might lead to different results. Therefore, further research using longer exposure times as well as investigating the toxicological effects of Ag-NPs in freshwater systems is needed, as the lack of salinity affects the chemistry and behavior of Ag-NPs differently to that observed in a marine environment.
Acknowledgments This research was supported through a grant (NE/F01192X/ 1) from the Natural Environment Research Council’s (NERC) Environmental Nanoscience Initiative (ENI). We are grateful to Peter Bond and to Dr. Andrew Fisher (University of Plymouth) for their help with TEM and ICP-OES, respectively. Dr. Rob Griffiths (Centre for Ecology and Hydrology, Oxford) is thanked for help using the 2D phoretix image analysis software. We are also indebted to Dr. Paul Somerfield (Plymouth Marine Laboratory) for his help with the multivariate statistical analyses.
Abbreviations 16S rRNA bp DGGE nano-Ag NP PCR ICP-OES
16S rRNA base pair(s) denaturing gradient gel electrophoresis silver nanoparticle(s) nanoparticle polymerase chain reaction inductively coupled plasma-optical emission spectrometery
Note Added after ASAP Publication Reference 28 was modified in the version of this paper published ASAP April 29, 2009; the corrected version published ASAP May 7, 2009.
Supporting Information Available Figure S1 shows absorption spectra of Ag-NPs and TEM micrographs of Ag-NPs. This material is available free of charge via the Internet at http://pubs.acs.org.
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