Microbial Bioavailability of Covalently Bound Polymer Coatings on

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Microbial Bioavailability of Covalently Bound Polymer Coatings on Model Engineered Nanomaterials Teresa L. Kirschling,†,‡ Patricia L. Golas,†,§ Jason M. Unrine,†,3 Krzysztof Matyjaszewski,§ Kelvin B. Gregory,†,|| Gregory V. Lowry,*,†,|| and Robert, D. Tilton*,†,‡,^ †

Center for the Environmental Implications of Nanotechnology (CEINT), ‡Department of Chemical Engineering, Department of Chemistry, Department of Civil and Environmental Engineering, and ^ Department of Biomedical Engineering Carnegie Mellon University, Pittsburgh, Pennsylvania 3 Department of Plant and Soil Sciences, University of Kentucky, Lexington, Kentucky )

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bS Supporting Information ABSTRACT: By controlling nanoparticle flocculation and deposition, polymer coatings strongly affect nanoparticle fate, transport, and subsequent biological impact in the environment. Biodegradation is a potential route to coating breakdown, but it is unknown whether surface-bound polymers are bioavailable. Here we demonstrate, for the first time, that polymer coatings covalently bound to nanomaterials are bioavailable. Model poly(ethylene oxide) (PEO) brush-coated nanoparticles (densely cross-linked bottle brush copolymers) with hydrophobic divinyl benzene cross-linked cores and hydrophilic PEO brush shells, having ∼30 nm hydrodynamic radii, were synthesized to obtain a nanomaterial in which biodegradation was the only available coating breakdown mechanism. PEO-degrading enrichment cultures were supplied with either PEO homopolymer or PEO brush nanoparticles as the sole carbon source, and protein and CO2 production were monitored as a measure of biological conversion. Protein production after 90 h corresponded to 14% and 8% of the total carbon available in the PEO homopolymer and PEO brush nanoparticle cultures, respectively, and CO2 production corresponded to 37% and 3.8% of the carbon added to the respective system. These results indicate that the PEO in the brush is bioavailable. Brush biodegradation resulted in particle aggregation, pointing to the need to understand biologically mediated transformations of nanoparticle coatings in order to understand the fate and transport of nanoparticles in the environment.

’ INTRODUCTION The rapidly growing number and variety of nanomaterials with current or envisioned commercial application have motivated research to understand their fate and impact in the environment.1 3 Life cycle analyses indicate various routes for engineered nanoparticles to enter the environment in significant quantities.4,5 Most nanomaterials are manufactured with organic macromolecular coatings to control their colloidal stability, transport, chemical functionality, compatibility with polymer matrices, or biocompatibility.6 12 Even uncoated nanomaterials will acquire macromolecular natural organic matter coatings upon release into the environment.10 Both engineered and natural coatings influence the fate and impact of nanomaterials in environmental systems, especially their aggregation tendencies, cellular uptake, and cytotoxicity.8,13 18 Loss of the macromolecular coating from particle surfaces would likely promote nanoparticle aggregation and deposition, thereby greatly affecting their transport and fate in the environment. Understanding r 2011 American Chemical Society

how engineered nanoparticles are transformed following environmental release is necessary to predict their environmental impacts.19 A wide variety of polymer coatings are used to stabilize engineered nanomaterials. Polymers can be physisorbed, grafted to, or grafted from the particle surface, or they can be covalently incorporated during synthesis (Figure 1). Different coating methods produce different polymer grafting densities and conformations on the particle surface, both of which affect particle particle and particle surface interaction forces. The magnitude of these interaction forces will affect microbe-nanoparticle adhesion and subsequent biological effects.8 Different degrees of exposure of polymer end groups in nanoparticle coatings may also Received: March 7, 2011 Accepted: May 3, 2011 Revised: May 2, 2011 Published: May 24, 2011 5253

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Figure 1. (a) Schematic of a physisorbed polymer on a nanoparticle. The polymer adopts a classic “loop-train-tail” conformation on the particle surface. (b) Engineered nanoparticle modified by a “grafting-to” method. This method yields a higher mass of coating on the nanoparticle compared to physisorption, but a lower polymer grafting density than particles modified by either a one step synthesis method or a “graftingfrom” method pictured in (c).

influence their availability for degradation by microorganisms. For example, higher molecular weight water-soluble polymers are less biodegradable than lower molecular weight polymers presumably because the polymer adopts a random coil conformation in solution shielding the end groups of the larger polymers.20,21 Once released in the environment, coatings may potentially be removed from engineered nanoparticles by several mechanisms. Certain polymers may hydrolyze in water, whereas others may photochemically decompose.22 Physisorbed polymers may desorb,23 albeit slowly. Little is known about biologically mediated coating removal and transformation mechanisms for engineered nanoparticles, and to date no study has demonstrated whether or not covalently bound macromolecules can be removed from nanoparticles by cells. Microorganisms are the most likely candidates to mediate this coating removal. Some macromolecules are broken down extracellularly by secreted enzymes,24 26 but the degradation of water-soluble synthetic polymers by bacteria often requires the molecule to be drawn into the cell membrane.21,24 Since nanoparticles larger than 10 nm may not be internalized by bacteria with intact membranes,27 it is not obvious that biodegradable polymers can be transformed when covalently bound to a nanoparticle. Another complicating factor is that some polymer coatings, especially anionic polyelectrolytes or certain hydrophilic nonionic polymers such as polyethylene oxide (PEO), may decrease cell contact via steric

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and/or electrostatic repulsions between the particle and the bacterial cell.8,28,29 The objective of this work was to determine whether or not macromolecular stabilizers covalently bound to engineered nanoparticles can be bioavailable to microorganisms. The choice of nanoparticle was critical to ensure that polymer was degraded from the particle surface by direct biological action, without abiotic hydrolysis or spontaneous desorption. Model PEO brushcoated nanoparticles with hydrophobic divinyl benzene crosslinked cores and hydrophilic poly(ethylene oxide) brush shells, having ∼30 nm hydrodynamic radius, were custom synthesized to obtain a nanomaterial in which biodegradation was the only available coating breakdown mechanism. These nanoparticles have a hydrophobic, nonhydrolyzable divinylbenzene crosslinked core resembling a polystyrene nanoparticle, surrounded by a brush of end-grafted, 2000 MW PEO chains. Most importantly, the stable C C bond between the core and the brush assures that the only removal mechanism under environmental conditions is biodegradation of nanoparticle-bound polymers. PEO was chosen as the model polymer coating as it is widely used as a nanoparticle stabilizer,28 30 in homopolymer form it is biodegradable by a variety of microorganisms,20,21,24 and it resists hydrolysis in water under environmental conditions.24 Given the broad biocompatibility and protein repellency of PEO,28 30 the PEO brush nanoparticles were expected to exhibit minimal cytotoxicity to bacteria. Here we demonstrate for the first time that poly(ethyleneoxide) (PEO) brushes, covalently bound to an engineered nanoparticle, are utilized as a carbon source for growth of bacteria enriched from urban river water. Biomass and CO2 production by an enrichment culture supplied with PEO brush nanoparticles as the sole added carbon source indicate that the polymer brush is bioavailable. Brush biodegradation resulted in particle aggregation, indicating that understanding biotransformations of nanoparticle coatings will play an important role in understanding nanoparticle fate in the environment.

’ MATERIALS AND METHODS Culture Enrichment and Characterization. A mixed culture of PEO-degrading bacteria was enriched from Monongahela River water in carbon-free minimal media (media composition listed in Supporting Information) augmented with 0.1 wt % 2000 g/mol dimethoxy PEO homopolymer (Sigma Aldrich, St. Louis, MO) as the sole added carbon source. Briefly, 500 mL of river water, collected in Pittsburgh, Pennsylvania, was filtered onto a sterile 0.22 μm filter. Filters were cut in half and added to BOD bottles containing minimal media and 100 mg/L PEO. Bottles were incubated at 30 C. After the dissolved oxygen in the BOD bottles was depleted to below 5% saturation, cultures were transferred to fresh media containing 500 mg/L of PEO. Cultures were incubated at 30 C in 125 mL flasks stirred at 100 rpm with a magnetic stir bar. To determine community composition, DNA was extracted from the mixed culture and the 16S rDNA gene was cloned and sequenced (methods in Supporting Information). The mixed PEO degrading community contained 78% Novosphingobium sp., 13% Pseudomonas sp., and 5% Hydrogenophaga sp. Certain members of the families Pseudomonadaceae (which includes Pseudomonas sp.) and Sphingomonadaceae (which includes Novosphingobium sp.) are known to degrade PEO.21 PEO Brush Nanoparticle Synthesis and Characterization. PEO brush nanoparticles (cross-linked bottle brush copolymers) were synthesized by atom transfer radical polymerization (ATRP). 5254

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Environmental Science & Technology Additional synthesis information is included in Supporting Information. The reagents were used as received from Sigma Aldrich (St. Louis, MO) unless otherwise indicated. PEO methacrylate (PEOMA2000) (subscript refers to approximate polymer molecular weight) (5.11 g, 2.5 mmol), CuBr (0.0705 g, 0.49 mmol), and tris[(2-pyridyl)methyl]amine (TPMA) (0.1427 g, 0.49 mmol) were added to a Schlenk flask, which was deoxygenated by six cycles of vacuum and backfill with N2. Anisole (15 mL) was injected, and the reaction components were allowed to dissolve. Divinyl benzene (1.3 mL of 80% solution) 7.4 mmol) was then injected, and the polymerization was initiated by injection of ethyl 2-bromoisobutyrate (EBiB) (72 μL, 0.49 mmol). The flask was placed into an oil bath thermostatted at 80 C. Samples were periodically withdrawn via nitrogen-purged syringe and analyzed by size-exclusion chromatography (SEC). The reaction was quenched after 110 h by dilution with ∼10 mL THF. GPC traces indicated a 75% conversion of the PEOMA. The PEO brush nanoparticles exhibited apparent Mn = 26 600 g/mol and Mw/Mn = 1.11, as determined by conventional SEC using a series of Styragel columns (Polymer Standards Services, 105 Å, 103 Å, 100 Å). The SEC eluent was THF (35 C) at a flow rate of 1.0 mL/min. Linear polystyrene standards were used as calibration for the particles. The significant discrepancy between the SEC determined molecular weight and that determined by light scattering (below) indicates a very compact structure, as expected. The copper catalyst was removed by passing the solution through a column filled with neutral alumina. PEO brush nanoparticles were purified by dialysis against methanol for one day, followed by dialysis against deionized water for 14 days (membrane molecular weight cut off = 25 000) with once daily water changes. The solution was filtered through a 0.2 μm sterile Nylon syringe filter, and the final concentration of the polymer was determined gravimetrically. The hydrodynamic radius of PEO brush nanoparticles was determined in batch mode by dynamic light scattering using a Malvern Instruments Zetasizer Nano ZS. Analysis of intensity autocorrelation functions by the CONTIN algorithm provided an intensity weighted distribution of diffusion coefficients, which was converted to a hydrodynamic size distribution using the Stokes Einstein equation for spheres. The radius of gyration, molar mass and hydrodynamic radius of the particles were also determined in separations mode using asymmetrical flow field flow fractionation coupled to refractive index detection and multiangle laser light scattering (AF4-RIMALLS). The particles were separated using a Wyatt Technologies Eclipse 3 AF4 system. Light scattering intensity was measured at 14 angles and integrated every second using a Wyatt DAWN HELEOS II MALLS. Additional information on the channel and the analysis is available in Supporting Information. Transmission electron microscopy (TEM) imaging of PEO brush nanoparticles was performed by misting droplets of a 20 g/L nanoparticle suspension onto a carbon coated grid. Particles were imaged using a Hitachi H-7100 TEM. Degradation Experiments. Polymer degradation experiments were conducted in 15 mL cultures in 125 mL Erlenmeyer flasks. All glassware was baked at 500 C overnight to remove any residual organic carbon and autoclaved prior to use. Duplicate cultures from the seventh transfer of the enrichment culture were amended either with 500 mg/L PEO homopolymer or PEO brush nanoparticles. Carbon-free control cultures received no carbon source amendment. Cultures were incubated at 30 C with constant stirring at 100 rpm. Cultures were seeded to an initial cell density of 2  106 cells per mL (determined by direct

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cell counting with acridine orange according to the methods of Kepner and Pratt31). Flasks were incubated aerobically, but sealed, with a sodium hydroxide trap consisting of a test tube filled with 1 mL of 50% NaOH, to absorb CO2 and prevent autotrophic growth. For protein analysis, samples were centrifuged, the supernatant removed, and the cell pellet disrupted by three freeze thaw cycles ( 78 and 100 C) in 50 mM NaOH. Protein content in this solution was determined by the microBCA assay (Thermo Fisher, Pittsburgh, PA), with additional details provided in Supporting Information. PEO homopolymer concentration was determined by the phosphomolybdic acid (PMA) assay according to the Stevenson method.32 Cells were removed by centrifugation and the supernatant was filtered through a 0.2 μm filter, as bacterial cells interfere with the PMA assay. A 250 μL aliquot of this solution was added to 750 μL deionized water in a 1.5 mL centrifuge tube. PEO was precipitated by the addition of 30 μL 2.4 M HCl (Fisher Scientific, Pittsburgh, PA), 20 μL of 10% barium chloride solution (Sigma Aldrich, St. Louis, MO), and 20 μL 10% phosphomolybdic acid (Sigma Aldrich, St. Louis, MO). The supernatant was removed. The precipitate was washed twice with deionized water and dissolved in 200 μL sulfuric acid. Deionized water was added to bring the volume to 1 mL, and 167 μL of 35% stannous chloride and 83 μL 5% ammonium thiocyanate (Sigma Aldrich, St. Louis, MO) was added. The absorbance was measured at 470 nm after 40 min of incubation at room temperature. Results were compared to a standard curve prepared with 2000 g/mol dimethoxy PEO in minimal media. A separate set of experiments was performed simultaneously to measure CO2 production in 160 mL serum bottles fitted with a CO2 trap filled with 1 mL 50% NaOH. Triplicate cultures were sacrificed at two time points, immediately after assembly (to determine initial CO2 concentrations) and after 7 days of incubation, by injecting 0.5 mL of 12 M HCl into the culture media. Acidified cultures were incubated overnight to allow liberated CO2 to be absorbed in the trap. CO2 from the sodium hydroxide trap was measured on an OI Analytical 1010 liquid TOC analyzer. Particle size distributions were monitored over time throughout the duration of the degradation experiments. One mL samples from PEO brush nanoparticle cultures were centrifuged for 5 min at 16 000 g to remove bacterial cells. The average hydrodynamic diameter of PEO brush nanoparticles in the supernatant was measured by dynamic light scattering using a Malvern Instruments Zetasizer Nano ZS as described above. As a control experiment, PEO brush nanoparticles were incubated in sterile media and DLS measurements were taken at the same set of time points. PEO brush nanoparticles were also incubated in the filter-sterilized supernatant from a 7-day culture of PEO-degrading bacteria that had been growing on 2000 MW PEO homopolymer to verify that changes in the nanoparticle size distributions were not a result of extracellular products in the culture. Supernatants from PEO homopolymer degrading cultures and no-carbon controls were also measured to verify that there were no entities in suspension similar in size to the PEO brush nanoparticles or aggregates. TEM was also used to independently verify the presence of aggregates in PEO brush nanoparticle cultures. PEO brush nanoparticle cultures were centrifuged for 5 min at 16 000  g. Pellets were fixed in 2% glutaraldehyde, followed by 1% osmium tetroxide buffered with PBS. After washing the bacteria 5255

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Figure 3. (a,b) TEM images of PEO brush nanoparticles prior to incubation with bacterial cultures. (c) TEM images showing PEO brush nanoparticle aggregates after 30 day incubation with bacteria. Most aggregated particles do not remain in direct contact with bacteria. No similar objects were observed in TEM images from cultures grown on PEO homopolymer after 30 days. (d) Close-up image of an aggregate comprising numerous PEO brush nanoparticles.

Figure 2. (a) Protein production by the mixed culture supplied with 500 mg/L of either PEO brush nanoparticles (filled diamond, filled circle) or 2000 MW dimethoxy PEO homopolymers (open triangle, open right-facing triangle) in solution. The no added carbon control is represented by the asterisk. (b) Corresponding degradation of the PEO homopolymer quantified by the phosphomolybdic acid assay. PEO brush nanoparticles could not be quantified by the same assay because they could not be separated adequately from cells and cell debris. Error bars are the standard error based on triplicate protein or PEO assays.

with distilled water and ethanol solutions of increasing strength (50%, 70%, 95%, and 100%), they were placed overnight in a 1:1 mixture of LR White resin and propylene oxide. The mixture was then replaced with 100% LR White resin and the samples were sliced into sections (100 nm thick). Thin sections were stained with 1% uranyl acetate and Reynold’s lead citrate, and viewed on a Hitachi 7100 transmission electron microscope. Digital images were obtained using AMT Advantage 10 CCD Camera System and NIH Image software. Unaggregated PEO brush nanoparticles incubated in sterile media did not centrifuge out of solution, and therefore could not be analyzed by this particular method. PEO homopolymer cultures were also observed by TEM following the same procedure.

’ RESULTS AND DISCUSSION The mixed culture was capable of growth on PEO homopolymers and PEO brush nanoparticles, using protein as a measure of

biomass production (Figure 2). Protein production after ∼90 h corresponds to approximately 14% and 8% of the total carbon available in the PEO and PEO brush cultures respectively. Neither PEO homopolymers nor PEO brush nanoparticles interfered with the MicroBCA protein assay (Supporting Information Figure 3). Protein production, and therefore microbial growth, was not observed in the no added carbon control. Protein production was also not observed when cultures were incubated in media prepared from water obtained from the final dialysis exchange (25 000 MWCO) of the PEO brush nanoparticles, indicating that any unreacted macromonomer or other carbon residues from nanoparticle synthesis (if present), did not support growth. Figure 2 also shows that protein production correlated with PEO degradation by cultures that were supplied with PEO homopolymer as the carbon source. No such information is available for the PEO brush nanoparticles. This is because the filtration step required to analyze PEO in cell-containing samples by the phosphomolybdic acid assay removed the PEO brush nanoparticles from solution. Physical characterization of the nanoparticles before and after exposure to bacterial cultures, as well as CO2 production, confirms that PEO brushes are degraded. PEO homopolymer cultures produced 8.37 ( 0.09 mM CO2 while PEO brush nanoparticle cultures produced 0.853 ( 0.008 mM CO2 over the course of 7 days. These quantities correspond to 37% and 3.8% of the carbon added to the system, respectively. While a small fraction of the added carbon is converted to CO2 in the PEO brush nanoparticle culture, this amount is quantifiable and indicates that the PEO brush is metabolically available. The protein and CO2 production data suggest that PEO brush nanoparticles are less available than PEO homopolymers for the 5256

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Environmental Science & Technology bacteria to use as a carbon source. The total amount of carbon added in these two systems is nearly identical. The nanoparticles are approximately 80% PEO by weight, so the fact that some of the carbon is bound in a less degradable polystyrene-like core cannot be the sole explanation. The key is likely that the PEO in the brush nanoparticles is attached by one end to the core, while PEO homopolymers each present two free ends to solution. Thus, the brush nanoparticles provide less than half of the total available chain end groups compared to PEO homopolymer. While different bacterial strains have different PEO degradation mechanisms, one work indicates that sphingomonads degrade PEO via successive oxidation of terminal groups. In the case of hydroxy-terminated PEO, the hydroxyl end group is oxidized to an aldehyde followed by a carboxylic acid, and the terminal ethylene glycol unit is cleaved by PEG dehydrogenase.21,33 This process requires that one end of the molecule must be drawn into the cell membrane21,33 so having fewer available end groups for PEO brush nanoparticles may account, in part, for the lower biomass and CO2 production compared to PEO homopolymer. Steric repulsions between densely packed chains in the brush may also hinder end group accessibility. Although polystyrenelike compounds have demonstrated biodegradability by certain mixed cultures,34 the time scales are particularly slow (on the order of 1% degradation over 80 days; compared to 100% PEO homopolymer degradation in a few days by the current enrichment culture). It is unlikely that the particle core is bioavailable in these short experiments. AF4-RI-MALLS indicates PEO brush nanoparticles are polydisperse (polydispersity index = 3.04), have a mass weighted average molar mass (Mw) of 7.35  106 g/mol, a mass weighted root-mean-square radius (Rw) of 23 nm, and a mass weighted hydrodynamic radius (Rh(w)) of 27 nm. Light scattering was complimented by transmission electron microscopy (TEM). Prior to incubation nanoparticles have an oblong shape after deposition on a carbon coated grid (Figure 3), with average long and short axes of ∼90 and 40 nm (estimated from TEM images, n = 25), respectively, in the dry state. Because of the small sample size and probable drying artifacts in TEM, the size determined from bulk scattering measurements is the reliable measure of particle size in aqueous suspension. Incubation of PEO brush nanoparticles with the PEO-degrading culture consistently resulted in aggregation of the particles. Dynamic light scattering analysis of supernatant samples collected from PEO brush nanoparticle-augmented cultures revealed an increase in hydrodynamic diameter over time (Figure 4a). This is consistent with PEO brush nanoparticle aggregation. In longterm incubations (30 days) bimodal size distributions were often observed (Figure 4b), however, conservative measures of aggregated particle sizes reported in Figure 4a are taken from monomodal distributions of particle size (Supporting Information), because a single size average is not a sufficient representation of aggregate characteristics in bimodal samples, where the degree of aggregation is even more extensive. Dynamic light scattering on samples collected from PEO homopolymer-augmented cultures or the no added carbon control revealed no entities in suspension with sizes comparable to the PEO brush nanoparticles or aggregates. Thus, the observed shift in nanoparticle size can be directly attributed to particle aggregation and not to any secreted extracellular products. Aggregation is likely caused by the degradation of the PEO brush which exposes the hydrophobic nanoparticle core and eliminates the steric stabilization provided by the brush. PEO

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Figure 4. (a) Increase in hydrodynamic diameter of PEO copolymer nanomaterials incubated with PEO degrading bacteria ((, 2). The sterile control is represented by an asterisk. Error bars are the standard error based on triplicate measurements. DLS determined size distributions for each sample and time point are available in Supporting Information. (b) Intensity weighted size distribution by dynamic light scattering of PEO brush nanomaterials in sterile minimal media or incubated with the PEO degrading mixed culture after 30 days of incubation.

brush nanoparticles dispersed in sterile media or in supernatant from a seven-day old PEO homopolymer-augmented culture showed no increase in size by dynamic light scattering (Supporting Information). Thus, the PEO brush nanoparticles are stably suspended without aggregating, even in media containing byproducts of bacterial metabolism, lending further support to the conclusion that nanoparticle aggregation was initiated by biodegradation of the PEO brush. PEO brush nanoparticle aggregates are readily observed in TEM images (Figure 3) of samples obtained by centrifugation of PEO brush nanoparticle-augmented cultures. No such aggregates were observed in cultures grown on PEO homopolymer. Most of the aggregated particles are not found in direct contact with cells, as might be expected for nonspecific steric repulsion by PEO.28 30 Specific interaction between PEO homopolymer and bacteria or PEO brush nanoparticles and bacteria likely occurs given that both are degraded, however, this was not observable by TEM. The TEM preparation involves multiple centrifugation steps. It is postulated that high shear forces during centrifugation may separate cells from weakly adherent PEO brush nanoparticles, particularly if the particles are attached to the bacteria by a 5257

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Environmental Science & Technology single end group (i.e., attached to an enzyme) as would be the case if the culture is degrading PEO chains by the same mechanism postulated for sphingomonads.21 In conclusion, by eliminating other coating degradation mechanisms, it has been demonstrated that PEO brushes covalently bound to engineered nanoparticles are bioavailable and can support the growth of microorganisms. Furthermore, bacterial degradation of the PEO brush layers that stabilize those nanoparticles leads directly to nanoparticle aggregation. These observations will likely extend to other types of macromolecular coatings and core nanoparticles, but are likely to be dependent on the biodegradability of the macromolecule, the underlying particle type, and concentration of particles and organisms present. How the surface modifier is attached to the underlying particle may have a strong influence on biodegradability. For example, if the polymer coating is highly cross-linked it may be less available to organisms which degrade polymer sequentially from a free end group. In the environment, the presence of natural macromolecules (natural organic matter) adsorbing on or within the polymer coating on the particle surface may also alter bioavailability. This work is an initial demonstration that covalently attached polymer nanoparticle coatings can indeed be biodegraded. More work is required to understand how polymer type, particle type, and chain conformation affect degradability. The findings also suggest that microbial modification of engineered nanoparticle coatings, when it occurs, will affect nanoparticle stability against aggregation upon release to the environment. Therefore, biotransformation of nanoparticle coatings should be considered when modeling the fate, transport, and potential ecological or human health risks associated with nanoparticle release into the environment.

’ ASSOCIATED CONTENT

bS

Supporting Information. Additional details on the particle synthesis, minimal media, protein assay, static light scattering, and the AF4-RI-MALLS fractograms. This material is available free of charge via the Internet at http://pubs.acs.org.

’ AUTHOR INFORMATION Corresponding Author

*E-mail: [email protected] (G.V.L.); [email protected] (R.D.T).

’ ACKNOWLEDGMENT The authors thank the PPG Industries Colloids, Polymers, and Surfaces Laboratory at Carnegie Mellon University for use of the Malvern Nanosizer and Brookhaven static light scattering apparatus. Joseph Suhan is thanked for performing the TEM. This material is based upon work supported by the National Science Foundation (NSF) and the Environmental Protection Agency (EPA) under NSF Cooperative Agreement EF-0830093, Center for the Environmental Implications of NanoTechnology (CEINT), EPA Grant R833326 and NSF Grant BES-068646. Any opinions, findings, conclusions or recommendations expressed in this material are those of the author(s) and do not necessarily reflect the views of the NSF or the EPA. This work has not been subjected to EPA review and no official endorsement should be inferred. T.L.K. is partially funded by the Bertucci Graduate Fellowship.

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dx.doi.org/10.1021/es200770z |Environ. Sci. Technol. 2011, 45, 5253–5259