Interactions of Silver Nanoparticles with Pseudomonas putida Biofilms

Oct 30, 2009 - Xu , X.; Brownlow , W.; Kyriacou , S.; Wan , Q.; Viola , J. Real-time probing of membrane transport in living microbial cells using sin...
1 downloads 0 Views 2MB Size
Download PDF
Environ. Sci. Technol. 2009 43, 9004–9009

Interactions of Silver Nanoparticles with Pseudomonas putida Biofilms JULIA FABREGA,† JOANNA C. RENSHAW, AND JAMIE R. LEAD* School of Geography, Earth and Environmental Sciences, University of Birmingham, Edgbaston, Birmingham, B15 2TT, United Kingdom

Received June 10, 2009. Revised manuscript received October 14, 2009. Accepted October 24, 2009.

Silver nanoparticles (Ag NPs) may present a risk to the environment due to their expected toxicity and wide exposure. The interactions between Ag NPs and laboratory-grown Pseudomonas putida biofilms were investigated under a range of environmentally relevant conditions (pH 6 and 7.5; presence and absence of Suwannee River fulvic acid (SRFA)) over 4 days. In the absence of SRFA, there was extensive sloughing of the biofilm bacteria into suspension implying NP-bacterial interactions and potential effects on NP transport in the environment. In the presence of SRFA, sloughing of cells into suspension was reduced under some conditions and Ag NPs and their aggregates were observed and quantified on and in the bacterial cells in the biofilm. Viability of the cells in all cases appear unchanged by the presence of Ag NPs. Cell viability was independent of the concentration of NPs in solution, but sloughing rates varied substantially, sometimes in a dose-dependent manner. The results suggest that biofilms are impacted by Ag NPs when SRFA was not present, and that SRFA increases uptake and bioaccumulation of Ag NPs to biofilms, perhaps resulting in longer term effects, which need further investigation.

Introduction The antimicrobial properties of silver nanoparticles (Ag NPs) and relatively low manufacturing cost have made them the most extensively used NPs in a range of products and processes (1). They are present in industrial applications and consumer goods such as medical devices (2), fabrics (3, 4), drinking water filters (5), food sprays (6), toys, containers, and electrical goods (7). The sharp increase in products containing Ag NPs in the past few years has increased the likelihood of their release into the environment (8). However, few studies have addressed the behavior of NPs in the environment (9, 10) and the effects on aquatic and sedimentary organisms (1, 11-13). The toxicity of NPs to organisms has been attributed to their large specific surface area (14, 15), chemical composition, surface structure, solubility (16), shape (17, 18), and charge (19), as well as aggregation state (20). These same properties will also control their stability and dispersion in aquatic environments (12). Previous research has suggested that the toxicity of Ag NPs to microorganisms is due to the release of Ag ions into the * Corresponding author e-mail: [email protected]. † Current address: School of Biosciences, Geoffrey Pope Building, University of Exeter, Stocker Road, Exeter, EX4 4QD, UK. 9004

9

ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 43, NO. 23, 2009

media (21), although other studies have indicated that particle size and shape may play a role in toxicity (15, 17). Proteomic studies have shown a definite effect of the Ag NP separate from the dissolved ion and this toxicity was observed at concentrations which are orders of magnitude lower than those for the dissolved ion (22). Our recent work (23) has shown a similar Ag NP specific toxicity but shown the Ag NPs to be somewhat less toxic than the dissolved Ag. Such differences may be rationalized by studies using NPs with systematically altered properties along with full characterization and such studies are urgently required. In this work, we studied the interactions of wellcharacterized Ag NPs (23) on single species biofilms of Pseudomonas putida under environmentally relevant conditions, investigating NP toxicity to biofilms and biofilm effects on NP movement.

Materials and Methods Preparation of Stock Solutions and Characterization. Silver nanopowder was purchased from Stanford Materials Corp. (Aliso Viejo, CA). The average diameter of Ag NPs was 65 ( 30 nm, the reported purity was 99.9%, and SSA was 2.40 ( 0.18 m2 g-1. A stock solution of Ag NPs in 0.25 mM Na citrate was prepared as previously described (23). Suwannee River Fulvic Acid (SRFA) was purchased from the International Humic Substances Society (IHSS, St. Paul, MN). Full details of characterization of size, morphology, aggregation, surface properties, and dissolution under appropriate conditions have been previously reported (23). SRFA stock solutions were prepared by dissolving 1 g L-1 and shaking at room temperature for 24 h to fully solubilize. Stocks were kept at 4 °C. Bacterial Strain and Media. Pseudomonas putida OUS 82 was obtained from Prof. W.T. Liu from the National University of Singapore. The media used for the Ag NPs exposure assay was minimal Davis medium (MDM, ionic strength of 0.055 M) (23). Previous research (23-26) suggests that this media is adequate for nanoparticle assays due to a relatively low salt concentration, although some aggregation occurs, which is reduced in the presence of SRFA (23). A known volume of Ag NPs stock was dispersed into MDM to reach the appropriate concentrations. SRFA was added (10 mg L-1) if required. Mixtures were shaken for 24 h at 25 °C. Media were prepared at three different concentrations of Ag NPs (20, 200, and 2000 ppb), at two different pH values (pH 6 and 7.5), and with and without 10 mg L-1 SRFA. Biofilm Reactor. All the material was acid washed with 10% HNO3, rinsed with water, and sterilized before use. Biofilms were grown in a Perspex flow cell (37.8 mm long × 19.8 mm wide × 5 mm thick). The complete flow cell reactor consisted of a total of 8 such flow cells independently connected to a flask containing MDM. A peristaltic pump was used to provide a constant flow of media at a rate of 0.1 mL min-1. Biofilms grown in the flow cell were analyzed for Ag uptake and biofilm biomass. Eluent from the biofilm was collected for further analysis of NPs or sloughed biofilm cells. For quantification of Ag uptake, three sterile plastic coverslips (13 mm diam.) were attached to the bottom of each flow cell prior to biofilm growth. Biomass of biofilms was also analyzed directly on these square plastic coverslips of the flow cell (27). Biofilm Growth. All experiments were performed at 25 °C. Cells of P. putida were grown in MDM (pH 6 or 7.5) to exponential phase, collected by centrifugation, washed, and resuspended in MDM at the appropriate pH. The optical density at 595 nm (OD 595; Lightwave WPA Diode array 25 10.1021/es901706j CCC: $40.75

 2009 American Chemical Society

Published on Web 10/30/2009

FIGURE 1. TEM micrographs of 4-d-old P. putida biofilm exposed for 24 h to (A) 0 ppb Ag NPs; (B) 2000 ppb Ag NPs; and (C) 2000 ppb of Ag NPs and 10 mg L-1 SRFA. Dark agglomerates in image C are Ag NPs. (D) Closeup of Ag NP interaction in C. Line represents 0.5 µm (A-C). Spectrophotometer) of the cell suspension was measured and adjusted to 0.1 by the addition of MDM. An aliquot of 3 mL of bacterial solution was injected under sterile conditions in the flow cell and left for 2 h, to allow adhesion of a small number of bacterial cells to the surface (28). Thereafter, MDM was passed through the flow cells at a flow rate of 0.1 mL min-1 for 3 days. Flasks were restocked with fresh and sterile MDM daily. After 3 days, solutions of Ag NPs in MDM with or without SRFA were passed over the 3-d old biofilm using a flow rate of 0.1 mL min-1 for 24 h. Control flow cells continued receiving MDM or MDM with SRFA. Metal Analysis. The Ag content in the media flowing through the reactor was measured by graphite furnace atomic absorption (AAS; Perkin-Elmer AA 600 Analyst graphite furnace spectrometer). Initial samples were checked by analysis by inductively coupled plasma mass spectroscopy (ICP-MS, Agilent 7500ce Octopole Reaction System) with excellent agreement. Samples of media were taken downstream and upstream of the flow cell where the biofilm was grown. The upstream sample verified the initial Ag concentration, while the downstream sample contained the material (bacterial cells and Ag NPs) detached from the biofilm as well as those Ag NPs not taken up by the biofilm. All downstream samples were kept on ice for the duration of the sampling to prevent bacterial growth. The downstream sample was filtered using a 0.22 µm pore size nitrocellulose disk (Millipore, diameter 25 mm) and ultrafiltered (Ultrafiltration system Amicon Stirred Cells, models 8400, Millipore; membranes of regenerated cellulose: 76 mm diameter, NMWL: 1 kDa), and the filtrates were analyzed for silver to determine the dissolved phase Ag (from NP dissolution) as previously reported (23). Solubility was measured on samples containing Ag NPs only, and not with SRFA due to the possibility of membrane-SRFA artifacts and inaccurate

results. To estimate the uptake of Ag by biofilms, 2 or 3 13mm coverslips were recovered from each flow cell, rinsed by dipping three times into sterile and fresh MDM, and gently dried to remove loose bacteria and Ag NPs. The biofilm samples were digested in a sealed vial containing 1 mL of concentrated HNO3 in a water bath at 60 °C for a period of 7 h. The samples were then diluted with 0.25 mM citrate solution to prevent aggregation or sorption of residual NPs. The total Ag content of the samples was quantified by AAS. Analysis of Biofilm Biomass and Cellular Viability. A standard LIVE/DEAD BacLight Bacterial Viability Kit (Invitrogen, Molecular probes) was used to monitor the viability of the biofilm population and to estimate total bacterial biomass, and was applied as described by the manufacturer. Biofilms were examined with a Leica DMIRE2 inverted confocal laser scanning microscope coupled with a constant temperature chamber maintained at 25 °C. Images were obtained with a 63×/1.4 N.A. Plan-Apochromat oil immersion DIC lens. A minimum of 30 microscopic images were taken for each treatment (n ) 3) and controls (n ) 3) and were analyzed by the software Image Structure Analyzer-2 (ISA-2) (29). The software was used to calculate biovolume per unit substrate (µm3) (BVSA) of viable cells and dead cells in each treatment. Addition of both viable and dead BVSA gave an estimate of the total biofilm volume per unit substrate surface area. Electron Microscopy. TEM with EDX was used to study the interaction of Ag NPs with P. putida biofilms. Coverslips from each treatment and control were sampled at the end of each assay, washed three times by gently dipping them into sterile and fresh MDM, and fixed with a solution of 2.5% glutaraldehyde in PBS for 3 h, followed by a secondary fixation and staining with OsO4 for 1 h. Samples were dehydrated with a series of ethanol:water mixtures, and finally embedVOL. 43, NO. 23, 2009 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

9

9005

ded into Mollenhauer resin and left to polymerize at 60 °C for 24 h. Polymerized samples were then sectioned in ∼60 nm slices. Samples were visualized using TEM JEOL 1200 EX. For EDX analysis of biofilms and Ag NPs, selected samples were visualized and analyzed using a TEM JEOL 7000 FEGSEM coupled to an Inca EDS at 20 kV laser intensity. Statistical Analysis. Statistical analysis was performed using a one-way analysis of variance followed by a Tukey’s posthoc test (SPSS Inc., Chicago IL). P values less than 0.05 were considered statistically significant.

Results and Discussion Characterization of Ag NPs. The results of Ag NP characterization in MDM are described in detail elsewhere (23). Particle size distribution ranged from 5 to 150 nm with a mean size of 65 ( 30 nm and good agreement among several different techniques. Particles were negatively charged, with electrophoretic mobility measurements of -2 and ca. -3.5 cm2 V-1 s-1 at pH 6 and 7.5, respectively, without natural organic matter (NOM) present, and values of less than -4.0 cm2 V-1 s-1 for both pH values with NOM present. NOM was shown qualitatively to cause disaggregation of aggregates of the NPs at low concentrations and short time periods and to have a protective effect on short-term toxicity of Pseudomonas sp (23). Visualization of Biofilms and Ag NPs. Figure 1 shows TEM images of biofilm cells unexposed to Ag NPs (control, Figure 1A), exposed to 2000 ppb Ag NPs (Figure 1B), and exposed to 2000 ppb Ag NPs with SRFA present (Figure 1C). In the controls (no Ag NPs with and without SRFA), cells were observed, but no Ag NPs, as expected. Ag NPs nor bacterial cells could be readily visualized or detected in the biofilm in the absence of SRFA at either pH and surface associated cells were less obvious. However, in the presence of SRFA, Ag NP aggregates and dispersed NPs were frequently detected in the EPS matrix and attached to the bacterial cells (Figure 1C), at both pH values. The aggregates observed were relatively large (∼ 0.5 µm), and closely associated with the bacterial cell surface (Figure 1C and D, Figure 2), with apparent invagination of the cell wall/membrane where in contact with the Ag NPs. In addition, these cells had a central component which was considerably less electron dense than the controls or after exposure to the Ag NPs alone (without SRFA), indicating some changes in cell structure caused by exposure to Ag NP with a SRFA coating (we have previously shown that HS forms a coherent film on these NPs (23)). This is consistent with, but not identical to, previous research which has, for instance shown membrane pitting in bacteria exposed to Ag NPs (30). The less electron dense cell cytoplasm has previously been described elsewhere (15) when E. coli cells were exposed to Ag nitrate solution, but to our knowledge has not been detected upon exposure to Ag NPs. EDX mapping analysis was used to confirm the presence of Ag (Figure 3), this is taken as indicative of the presence of Ag NPs. In addition to aggregates associated with the cell surface, a large number of particles were also detected within the cell (Figure 2) in biofilms exposed to Ag NPs and SRFA. EDX analysis showed that the majority of these are staining artifacts but a small number were disaggregated Ag NPs, indicating that a small fraction of the Ag NPs was taken up intracellularly, and confirming some of our qualitative TEM observations. The interaction of Ag NPs with planktonic bacteria, internalization, and accumulation has been previously described and related to NP size (15, 17, 30-32). However, similar work on biofilms has been lacking until now. Ag NP Uptake by Biofilms. Table 1 summarizes the uptake of Ag NPs by the biofilm under different concentrations and pH values, with the data given in more detail in Figure 4. 9006

9

ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 43, NO. 23, 2009

FIGURE 2. TEM micrographs of (A) Ag NPs dispersed in fresh MDM; (B) MDM containing Ag NPs coated with extrapolymeric substances collected after exposure to P. putida biofilm; (C) closeup of interaction of Ag NPs with P. putida cell membrane; and (D) (inset) interaction of Ag NPs with P. putida biofilm. Line represents 0.2 µm Uptake of Ag NP by biofilms includes material within the cell, attached to the cell, and in the biofilm EPS. The total mass of Ag NPs to which the biofilms were exposed to over

FIGURE 3. (A) EDX mapping and spectrum of Ag in the P. putida biofilm (pH 6 with SRFA); (B) TEM of P. putida biofilm mapped with EDX.

TABLE 1. Uptake of Ag (µg, Mean ± Standard Deviation) by the Biofilm Grown in the Flow Cell Reactor (Number in Parentheses Represents the Mass (µg) to Which Biofilms Were Exposed over 24 h; AAS Data Were Normalized to µg Per Surface Area of Biofilm and the Total Ag NPs Uptake by the Area Covered by Biofilm Was Estimated from This Value) Ag NPs (ppb) and total (µg) Ag NPs exposed over 24 h treatment pH pH pH pH

6 6 SRFA 7.5 7.5 SRFA

20 (2.28 µg)

200 (22.8 µg)

2000 (228 µg)

2.12 ( 0.2 1.1 ( 0.2 0.05 ( 0.01 0.12 ( 0.05

1.2 ( 0.3 1.2 ( 0.6 0.1 ( 0.02 1.8 ( 0.3

9.9 ( 1.3 23.2 ( 2.6 7.1 ( 0.7 15.9 ( 1.1

24 h was 2.28, 22.8, and 228 µg for the 20, 200, and 2000 ppb solutions and clearly only a small fraction (ca. 10% maximum) of this was retained by the biofilms under all conditions studied. We have previously shown that only 1% of the total Ag is dissolved from the NPs (23), so essentially all measured Ag is expected to be in the nanoparticulate form, at least before any potential biotransformations. There are indications that SRFA causes greater bioaccumulation of the Ag NPs; this trend was most obvious at the highest concentration

(2000 ppb), with the presence of SRFA more than doubling Ag uptake. Uptake was also higher at pH 6 with and without SRFA, compared to pH 7.5; the higher pH is closer to optimal growth conditions and perhaps the biofilms were better able to remove uptaken NPs or prevent uptake. Effect of Ag NPs on Biofilms. Interactions between the Ag NPs and cells were observed by TEM in the presence of SRFA only and quantification of silver uptake indicated a significantly higher uptake with SRFA than without. Nevertheless in the presence of SRFA, no decrease in biomass was observed compared to control biofilms, while in the absence of SRFA small but significant decreases in biomass were observed (Figure 4). A decrease of BVSA by ca. 8% (compared to control biofilms) at pH 6 for concentrations from 20 to 2000 ppb and at pH 7.5 at 200 and 2000 ppb (Figure 4, ANOVA p < 0.05) occurred. Regardless of the Ag NP concentration, when SRFA was present, there was no decrease in biomass at either pH value (Figure 4; ANOVA, p < 0.05). As previously shown (23), SRFA coats the Ag NPs with a coherent nanoscale film and this appears to “passivate” the NPs in terms of their biological effect. We have speculated (23) that this may be due to an antioxidant effect of the SRFA or protection of the cell wall and membrane from physical damage by the NP. However, until the exact mechanisms of toxicity are understood, this remains speculation. Despite

FIGURE 4. Effects of a 24 h exposure of Ag NPs on biovolume per surface area (BVSA, solid bars) of a P. putida biofilm at pH 6 (A1); pH 6 with 10 mg L-1 SRFA (A2); pH 7.5 (B1) and pH 7.5 with 10 mg L-1 SRFA (B2). The total Ag NPs uptake (µg) by the biofilm is represented by the solid line. Bars represent 1 standard deviation. Asterisks (*) represent treatments in which Ag NPs had a significant effect on BVSA compared to control biofilms (ANOVA, p < 0.05). VOL. 43, NO. 23, 2009 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

9

9007

FIGURE 5. Biomass of biofilm (OD595) sloughed off due to Ag NPs after initial exposure (0-4 h) (A) and from 8 to 24 h (B). Asterisks (*) represent treatments in which the biomass sloughed off was significantly different from control biofilms (ANOVA, p < 0.05). the evident thinning of the biofilm after exposure to Ag NPs only (no SRFA), cellular viability of the biofilm cells remaining after exposure was not significantly different among pH values, concentrations of exposure, and presence or absence of SRFA (data not shown). The reduced biofilm volume, no change in viability and “bioaccumulation” of Ag NPs suggest the biofilms may be acting in such a way as to remove the toxicant (Ag NPs) from the biofilm but that the presence of SRFA serves to mask the NPs and decrease the biofilm response. Further work is required to elucidate the longer term effects of NPs on biofilms. In particular, does this bioaccumulation within the biofilm pose long-term hazards for biofilms and their environmental functions? The thinning of the biofilms was observed semiquantitatively (TEM) and quantitatively (confocal microscopy). The mechanism for this thinning appears to be at least in part due to sloughing of biofilm material into suspension as shown in Figure 5. Initially (0-4 h), a dose-dependent increase in biomass detaching from the biofilms was observed with exposure to Ag NPs in the absence of SRFA at pH 7.5 (Figure 5A). At pH 7.5 this sloughing was suppressed with the addition of SRFA, consistent with the picture developed. However, negligible sloughing occurs at pH 6 without SRFA and greater sloughing occurs with SRFA present, but in a non-dose-dependent manner, and the presence of SRFA caused a significant increase in the amount of biomass detaching, both in the controls and the biofilms exposed to Ag NPs. The removal of bacterial cells at pH 6 appears to be related to the SRFA and not the AgNPs, indicating the complexity of the system studied and the need for more detailed analysis to investigate the effects of natural nanoparticles in addition to the manufactured nanoparticles. The effect of the Ag NPs on the biofilm appeared to vary over time of exposure, with significantly less biomass detaching under all conditions during later exposure to Ag NPs (8-24 h; Figure 5B), compared to the initial exposure. It is important to note that the minimal number of points in a time series (0-4 h and 8-24 h, forced on us by feasibility considerations) may mask trends in data. Overall, although no changes in cell viability were detected, biofilm behavior was substantially affected by the addition of a typical natural organic matter, with changes in surface chemistry and dispersion (23) substantially altering NP-cell interactions and sloughing. This study is, to our knowledge, the first to quantitatively discuss the interactions between well quantified and characterized bacterial biofilms and silver nanoparticles. Bacteria and biofilms are fundamental in providing many environmental services and nanosilver is an emerging pollutant of 9008

9

ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 43, NO. 23, 2009

enormous potential. The impacts of nanosilver on biofilm detachment and of SRFA on nanosilver bioaccumulation have many implications for environmental behavior, although much further work is required.

Acknowledgments This work was funded by EU Marie Curie Early Stage Training awarded to J.R.L. (MEST-CT-2004-504356). We thank the Centre for Electron Microscopy from the University of Birmingham for their help on TEM sample preparation.

Literature Cited (1) Klaine, S. J.; Alvarez, P. J. J.; Batley, G. E.; Fernandes, T. F.; Handy, R. D.; Lyon, D. Y.; Mahendra, S.; McLaughlin, M. J.; Lead, J. R. Nanomaterials in the environment: Behavior, fate, bioavailability, and effects. Environ. Toxicol. Chem. 2008, 27 (9), 1825– 1851. (2) Roe, D.; Karandikar, B.; Bonn-Savage, N.; Gibbins, B.; Roullet, J.-B. Antimicrobial surface functionalization of plastic catheters by silver nanoparticles. J. Antimicrob. Chemother. 2008, 61 (4), 869–876. (3) Kulpinski, P. Bioactive cellulose fibres with silver nanoparticles. e-Polym. 2007; Art. 068. (4) Lee, H. J.; Yeo, S. Y.; Jeong, S. H. Antibacterial effect of nanosized silver colloidal solution on textile fabrics. J. Mater. Sci. 2003, V38 (10), 2199. (5) Jain, P.; Pradeep, T. Potential of silver nanoparticle-coated polyurethane foam as an antibacterial water filter. Biotechnol. Bioeng. 2005, 90 (1), 59–63. (6) An, J.; Zhang, M.; Wang, S.; Tang, J. Physical, chemical and microbiological changes in stored green asparagus spears as affected by coating of silver nanoparticles-PVP. Food Sci. Technol. 2008, 41 (6), 1100–1107. (7) www.nanotechproject.org; Project on emerging nanotechnologies; 2008. (8) Luoma, S. N. Silver Nanotechnologies and the Environment: Old Problems and New Challenges?; Woodrow Wilson International Center for Scholars or The PEW Charitable Trusts: Washington DC, September, 2008; p 72. (9) Baalousha, M.; Manciulea, A.; Cumberland, S.; Kendall, K.; Lead, J. R. Aggregation and surface properties of iron oxide nanoparticles: Influence of pH and natural organic matter. Environ. Toxicol. Chem. 2008, 27 (9), 1875–1882. (10) Diegoli, S.; Manciulea, A. L.; Begum, S.; Jones, I. P.; Lead, J. R.; Preece, J. A. Interaction between manufactured gold nanoparticles and naturally occurring organic macromolecules. Sci. Total Environ. 2008, 402 (1), 51–61. (11) Blaise, C.; Gagne´, F.; Fe´rard, J. F.; Eullaffroy, P. Ecotoxicity of selected nano-materials to aquatic organisms. Environ. Toxicol. 2008, 23 (5), 591–598. (12) Navarro, E.; Baun, A.; Behra, R.; Hartmann, N. B.; Filser, J.; Miao, A. J.; Quigg, A.; Santschi, P. H.; Sigg, L. Environmental behavior

(13) (14) (15) (16)

(17)

(18)

(19)

(20) (21)

(22)

and ecotoxicity of engineered nanoparticles to algae, plants, and fungi. Ecotoxicology 2008, 17 (5), 372–386. Handy, R. D.; von der Kammer, F.; Lead, J. R.; Hassellov, M.; Owen, R.; Crane, M. The ecotoxicology and chemistry of manufactured nanoparticles. Ecotoxicology 2008, 17 (4), 287–314. Choi, O.; Hu, Z. Size Dependent and Reactive Oxygen Species Related Nanosilver Toxicity to Nitrifying Bacteria. Environ. Sci. Technol. 2008, 42 (12), 4583–4588. Morones, J. R.; Elechiguerra, J. L.; Camacho, A.; Holt, K.; Kouri, J. B.; Ramı´rez, J. T.; Yacaman, M. J. The bactericidal effect of silver nanoparticles. Nanotechnology 2005, 16, 2346–2353. Franklin, N. M.; Rogers, N. J.; Apte, S. C.; Batley, G. E.; Gadd, G. E.; Casey, P. S. Comparative toxicity of nanoparticulate ZnO, bulk ZnO, and ZnCl2 to a freshwater microalga (Pseudokirchneriella subcapitata): The importance of particle solubility. Environ. Sci. Technol. 2007, 41 (24), 8484–8490. Pal, S.; Tak, Y. K.; Song, J. M. Does the antibacterial activity of silver nanoparticles depend on the shape of the nanoparticle? A study of the gram-negative bacterium Escherichia coli. Appl. Environ. Microbiol. 2007, 73 (6), 1712–1720. Poland, C. A.; Duffin, R.; Kinloch, I.; Maynard, A.; Wallace, W. A. H.; Seaton, A.; Stone, V.; Brown, S.; Macnee, W.; Donaldson, K. Carbon nanotubes introduced into the abdominal cavity of mice show asbestos-like pathogenicity in a pilot study. Nat. Nanotechnol. 2008, 3 (7), 423–8. Gratton, S. E. A.; Ropp, P. A.; Pohlhaus, P. D.; Luft, J. C.; Madden, V. J.; Napier, M. E.; DeSimone, J. M. The effect of particle design on cellular internalization pathways. PNAS 2008, 105 (33), 11613– 11618. Brant, J.; Lecoanet, H.; Wiesner, M. R. Aggregation and deposition characteristics of fullerene nanoparticles in aqueous systems. J. Nanopart. Res. 2007, 7, 545–553. Lok, C. N.; Ho, C. M.; Chen, R.; He, Q. Y.; Yu, W. Y.; Sun, H.; Tam, P. K. H.; Chiu, J. F.; Che, C. M. Silver nanoparticles: partial oxidation and antibacterial activities. J. Biol. Inorg. Chem. 2007, 12 (4), 527–534. Lok, C.; Ho, C.; Chen, R.; He, Q.; Yu, W.; Sun, H.; Tam, P.; Chiu, J.; Che, C. Proteomic analysis of the mode of antibacterial action of silver nanoparticles. J. Proteome Res. 2006, 5 (4), 916–624.

(23) Fabrega, J.; Fawcett, S. R.; Renshaw, J. C.; Lead, J. R. Silver nanoparticle impact on bacterial growth: effect of pH, concentration and organic matter. Environ. Sci. Technol. 2009, 43 (19), 7285–7290. (24) Lyon, D. Y.; Adams, L. K.; Falkner, J. C.; Alvarez, P. J. J. Antibacterial Activity of Fullerene Water Suspensions: Effects of Preparation Method and Particle Size. Environ. Sci. Technol. 2006, 40, 4360–4366. (25) Lyon, D. Y.; Fortner, J. D.; Sayes, C. M.; Colvin, V. L.; Hughes, J. B. Bacterial cells association and antimicrobial activity of a C60 water suspension. Environ. Toxicol. Chem. 2005, 24 (11), 2757–2762. (26) Li, D.; Lyon, D. Y.; Li, Q.; Alvarez, P. J. J. Effect of soil sorption and aquatic natural organic matter on the antibacterial activity of a fullerene water suspension. Environ. Toxicol. Chem. 2008, 27 (9), 1888–1894. (27) Bressel, A.; Schultze, J. W.; Khan, W.; Wolfaardt, G. M.; Rohns, H.-P.; Irmscher, R.; Schoning, M. J. High resolution gravimetric, optical and electrochemical investigations of microbial biofilm formation in aqueous systems. Electrochim. Acta 2003, 48 (2022), 3363–3372. (28) Dunne, W. M., Jr. Bacterial Adhesion: Seen Any Good Biofilms Lately? Clin. Microbiol. Rev. 2002, 15 (2), 155–166. (29) Beyenal, H.; Donovan, C.; Lewandowski, Z.; Harkin, G. Threedimensional biofilm structure quantification. J. Microbiol. Methods 2004, 59 (3), 395–413. (30) Sondi, I.; Salopek-Sondi, B. Silver nanoparticles as antimicrobial agent: a case study on E-coli as a model for Gram-negative bacteria. J. Colloid Interface Sci. 2004, 275 (1), 177–182. (31) Xu, X.; Brownlow, W.; Kyriacou, S.; Wan, Q.; Viola, J. Real-time probing of membrane transport in living microbial cells using single nanoparticle optics and living cell imaging. Biochemistry 2004, 43 (32), 10400–10413. (32) Shrivastava, S.; Bera, T.; Roy, A.; Singh, G.; Ramachandrarao, P.; Dash, D. Characterization of enhanced antibacterial effects of novelsilvernanoparticles.J.Nanotechnol.2007,18,225103–225112.

ES901706J

VOL. 43, NO. 23, 2009 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

9

9009