Low Concentrations of Silver Nanoparticles in Biosolids

Cause Adverse Ecosystem Responses under Realistic

Field Scenario

Benjamin P. Colman1,9*, Christina L. Arnaout2,9, Sarah Anciaux3,9, Claudia K. Gunsch2,9,

Michael F. Hochella Jr.4,9, Bojeong Kim4,9, Gregory V. Lowry5,9, Bonnie M. McGill1

, Brian C. Reinsch5,9,

Curtis J. Richardson6,9, Jason M. Unrine7,9, Justin P. Wright1

, Liyan Yin8,9, Emily S. Bernhardt1,9

1 Department of Biology, Duke University, Durham, United States of America,, 2 Civil and Environmental Engineering Department, Duke University, Durham, United States

of America, 3 Department of Chemistry, Coe College, Cedar Rapids, United States of America, 4Department of Geosciences, Virginia Tech, Blacksburg, United States of

America, 5 Department of Civil and Environmental Engineering, Carnegie Mellon University, Pittsburgh, United States of America, 6 Duke Wetland Center, Nicholas School

of the Environment, Duke University, Durham, United States of America, 7 University of Kentucky, Department of Plant and Soil Sciences, Lexington, United States of

America, 8 Key Laboratory of Aquatic Botany and Watershed Ecology, Wuhan Botanical Garden, Chinese Academy of Sciences, Wuhan, People’s Republic of China,

9 Center for the Environmental Implications of Nanotechnology (CEINT), Duke University, Durham, United States of America

Abstract

A large fraction of engineered nanomaterials in consumer and commercial products will reach natural ecosystems. To date,

research on the biological impacts of environmental nanomaterial exposures has largely focused on high-concentration

exposures in mechanistic lab studies with single strains of model organisms. These results are difficult to extrapolate to

ecosystems, where exposures will likely be at low-concentrations and which are inhabited by a diversity of organisms. Here

we show adverse responses of plants and microorganisms in a replicated long-term terrestrial mesocosm field experiment

following a single low dose of silver nanoparticles (0.14 mg Ag kg21 soil) applied via a likely route of exposure, sewage

biosolid application. While total aboveground plant biomass did not differ between treatments receiving biosolids, one

plant species, Microstegium vimeneum, had 32 % less biomass in the Slurry+AgNP treatment relative to the Slurry only

treatment. Microorganisms were also affected by AgNP treatment, which gave a significantly different community

composition of bacteria in the Slurry+AgNPs as opposed to the Slurry treatment one day after addition as analyzed by TRFLP analysis of 16S-rRNA genes. After eight days, N2O flux was 4.5 fold higher in the Slurry+AgNPs treatment than the

Slurry treatment. After fifty days, community composition and N2O flux of the Slurry+AgNPs treatment converged with the

Slurry. However, the soil microbial extracellular enzymes leucine amino peptidase and phosphatase had 52 and 27% lower

activities, respectively, while microbial biomass was 35% lower than the Slurry. We also show that the magnitude of these

responses was in all cases as large as or larger than the positive control, AgNO3, added at 4-fold the Ag concentration of the

silver nanoparticles.

Citation: Colman BP, Arnaout CL, Anciaux S, Gunsch CK, Hochella MF Jr, et al. (2013) Low Concentrations of Silver Nanoparticles in Biosolids Cause Adverse

Ecosystem Responses under Realistic Field Scenario. PLoS ONE 8(2): e57189. doi:10.1371/journal.pone.0057189

Editor: Stephen J. Johnson, University of Kansas, United States of America

Received June 13, 2012; Accepted January 22, 2013; Published February 27, 2013

Copyright: 2013 Colman et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits

unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Funding: This study was supported by the National Science Foundation (NSF) and the Environmental Protection Agency (EPA) under NSF Cooperative

Agreement EF-0830093, and the Center for the Environmental Implications of Nanotechnology (CEINT). Any opinions, findings, conclusions, or recommendations

expressed in this paper are those of the authors 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. Funding was also provided for BCR by the Bertucci Graduate Fellowship at Carnegie Mellon University. The

funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Competing Interests: While one of our coauthors, JW, is a PLOS ONE Editorial Board member, this fact does not alter the authors’ adherence to all the PLOS

ONE policies on sharing data and materials, as detailed online in the PLOS ONE guide for authors.

* E-mail: benjamin.colman@duke.edu

Introduction

Engineered silver nanoparticles (AgNPs) are an emerging

environmental contaminant of concern for regulators and

consumer advocates because of their antimicrobial properties.

AgNP production and incorporation into consumer products is

increasing rapidly[1,2,3], and the majority of AgNPs released from

consumer products are expected to enter terrestrial ecosystems

through land-application of biosolids[4]. Given the critical role of

microbial communities in organic matter and nutrient cycling in

ecosystems, environmental exposures of AgNPs have the potential

to alter ecosystem productivity and biogeochemistry[5].

Our knowledge of AgNP toxicity is largely drawn from

controlled laboratory experiments with single strains of bacteria

or fungi. These studies have shown that AgNP exposure leads to

membrane damage, oxidative stress, and significant mortality

[6,7,8,9,10]. AgNP toxicity is not limited to bacteria and fungi;

recent studies have reported that AgNPs significantly reduced the

growth of the annual grass, Lolium multiflorum [11], and reduced

photosynthesis by the green alga, Chlamydomonas reinhardtii [12].

The mechanistic insights from these single-species experiments are

vital to understanding AgNP impacts on organisms, but extrapolating from them to multi-species communities in complex

environments is not practical. For example, the small number of

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laboratory AgNP exposure experiments conducted in environmental media (i.e., soil, streamwater, sediment) with diverse native

microbial communities have shown that AgNPs had no

effects[13,14], limited sublethal effects[15], or reduced effects in

comparison to dissolved Ag[14,16].

Additionally, the dominant route by which AgNPs will enter

natural ecosystems is as aged particles through the landapplication of wastewater treatment biosolids[4], not as Ag0 in

freshly synthesized well dispersed nanoparticles. Biosolids are the

processed or refined sewage sludge from wastewater treatment

plants which are used in agricultural lands and rangelands as a

nutritional soil additive. Recent work has shown that in biosolids,

Ag is predominantly present as Ag2S, [17,18] and sulfidation

dramatically alters the properties of AgNPs including their surface

charge, the ability to release Ag+

[19], and toxicity[20].

In this paper, we present a field experiment examining the

ecosystem level impacts of AgNPs by using AgNP-dosed biosolids

in outdoor mesocosms with diverse plant and microbial communities. We sought to answer three questions: 1) What is the

environmental fate of Ag under this exposure scenario? 2) How do realistic

additions of AgNPs affect plant productivity, microbial community composition,

and microbially-mediated biogeochemical cycling? 3) To what extent are the

fate and biological impacts of AgNPs distinct in magnitude or direction from

those of Ag+? To address these questions we added biosolids alone,

or biosolids amended with either polyvinylpyrrolidone-coated

AgNPs or AgNO3 to a total of 24 replicate grassland mesocosms

representative of abandoned agricultural fields found throughout

the piedmont of the southern United States (Figure 1A, B).

Setup and AgNP Addition

All mesocosms were filled with ,81 kg of local floodplain soils

that had been sieved (10 mm) to homogenize them and remove

large rocks. Each mesocosm was planted with live plugs of three

individuals of five commonly occurring plant species (Carex lurida,

Juncus effusus, Lobelia cardinalis, Microstegium vimineum, and Panicum

virgatum) on June 23, 2009 (Figure 1A). Plants were allowed to

establish for 2 months prior to initiation of treatments. On August

25th, 2009 (Day 0), four experimental treatments were applied

(Figure 1B): ‘‘Controls’’ received 1.5 L of deionized water;

‘‘Slurry’’ treatments received 1.5 L of biosolid slurry (200 g Class

A biosolids mixed in deionized water, with a background of

1.5 mg Ag); ‘‘Slurry+AgNPs’’ received 1.5 L of biosolids containing 9.9 mg of Ag as AgNPs (total 11.4 mg Ag); and ‘‘Slurry+AgNO3’’ received 1.5 L of biosolid slurry containing 44 mg of

Ag as AgNO3 to serve as a positive control (total 45.5 mg Ag).

Assuming homogeneous mixing with the soil, these loading rates

would give concentrations of 0.02, 0.14, and 0.56 mg Ag kg21 soil

in Slurry, Slurry+AgNPs, and Slurry+AgNO3 treatments, respectively. Each treatment was applied to six randomly-selected

replicate mesocosms. This biosolid application rate of 870 g m22

was based on US EPA guidelines [21], and Ag additions were

based on a recent national analysis of biosolids contaminants[22].

Specifically, our AgNP treatment was at the 95% confidence

interval of total Ag in biosolids, and our AgNO3 treatment was 4-

fold higher to provide a positive control, while still remaining

within the range of measured biosolid silver values.

For our AgNP treatment, we used a polyvinylpyrrolodinecoated AgNP powder that is representative of AgNPs that are

commercially available for incorporation into consumer products

(Nanoamorphous Materials, Los Alamos, USA). TEM analyses

revealed particle diameters to be 21617 nm (Figure 2A, B).

Detailed characterization of these particles has been previously

reported[23]. Briefly, particles in the stock suspension (see methods

for stock suspension preparation) had a left skewed particle size

distribution, were polydisperse, had a pH of 4.5, a zeta potential of

222.5 mV, and were about 10% Ag2O, and 90% Ag0 as

measured by extended x-ray absorption fine structure [24].

To examine ecosystem level effects, we determined treatment

differences in soil microbial community biomass, composition, and

activity; plant biomass and photosynthesis; and trace gas fluxes

from soils. Photosynthesis was measured on Days 8 and 30, while

soil gas flux was measured on Days 8 and 50. Three replicate 1 cm

diameter soil cores were collected for microbial community

composition and microbial biomass on days 0 (before applying

treatments), 1, and 50 of the experiment. On day 50 following gas

flux measurements, all aboveground plant biomass was clipped at

the soil surface and sorted by species. Following plant harvest, we

collected three replicate 5 cm diameter soil cores, which were

immediately separated in the field into depth classes (0–1, 1–5, 5–

10 cm) and were subsequently used to determine root biomass, soil

Ag content, microbial biomass, and extracellular enzyme activity.

Based on the results of previous laboratory experiments in

environmental media [13,14] we expected to see minimal effects

from AgNPs in our mesocosm experiment compared to our

AgNO3 positive control. Contrary to our expectations, microorganisms and plants showed predominantly adverse effects from the

AgNP treatment, which in almost all cases were as large as or

larger than the response to treatment with AgNO3 added at a fourfold higher concentration. Figure 1. Terrestrial mesocosms in the Duke Forest, Durham,

NC, USA. Mesocosms A on the day of planting, and B 63 days later

(Day 0 of the experiment) mesocosms being amended with biosolid

slurry

doi:10.1371/journal.pone.0057189.g001

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Results and Discussion

Silver in plants and soil

Across all three biosolid treatments, Ag recovery averaged

58625% (mean 6 SD). This incomplete recovery is possibly due

to a combination of factors including loss due to raindrop splash of

biosolids, wind erosion of dessicated biosolid crust, movement of

silver below 10 cm, and insect removal activity. Soils were the

major sink for Ag in all cases (Figure 3A), which contained nearly

99% of the recovered Ag in all treatments. Aboveground and

belowground plant tissues accumulated high concentrations of Ag

in the Slurry+AgNPs and Slurry+AgNO3 treatments relative to

Control (Table S1), but contained less than 1% of the recovered

Ag (Figure 3A). Regardless of the initial form in which Ag was

added (native Ag in Slurry, Slurry+AgNPs, or Slurry+AgNO3),

concentrations and recoveries (Figure 3A) were highest in surface

Figure 2. TEM characterization of particles and aggregates. TEM

image A of a mixture of primary particles and particle aggregates B

particle size distribution, and C detailed view of AgNP aggregate. (A

and B are adapted from reference 23, and reproduced by permission of

the Copyright Clearance Center on behalf of Elsevier)

doi:10.1371/journal.pone.0057189.g002

Figure 3. Silver fate in terrestrial mesocosms. A Recovery of silver

by ecosystem compartment after 50 days exposure to biosolid Slurry

(white bars), Slurry+AgNPs (gray bars), or Slurry + AgNO3 (black bars),

and B EXAFS linear combination fit (k-space) of AgNPs after 15 minute

exposure to biosolid slurry. In B, Lines indicate the data (black line), the

linear combination fit (light gray dashed line), and the individual fit

components Ag0 (gray line) and Ag2S (dark gray line) are shown, and

represent 7562% and 2566% percent of the silver, respectively. The

model R-factor = 0.0672, chi2 = 86.64, and the reduced chi2 = 0.4867

(parameters describing goodness of fit of the model to the data). Error

bars in panel A are standard errors of the mean (n = 6). Since all

treatments showed the same pattern in ANOVA post-hoc testing,

differences for each treatment within each ecosystem compartment

were denoted with brackets with letters, where shared letters denote

no significant difference at p,0.05 between ecosystem compartments

within a treatment

doi:10.1371/journal.pone.0057189.g003

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(0–1 cm) soils and decreased with depth to 10 cm where they were

equivalent to concentrations measured in control treatment soils.

Given the low concentrations used in our experiment, direct

observations of the physical and chemical properties of Ag

recovered in our mesocosm soils and plants (e.g., particle size,

aggregation state, speciation) using techniques including transmission electron microscopy (TEM) and extended x-ray absorption

fine structure (EXAFS) proved to be elusive. However, two lines of

evidence suggest that AgNPs added to biosolids were rapidly

transformed. Our first line of evidence comes from EXAFS data

from a separate batch study of these same AgNPs and biosolids at

a 10-fold higher concentration. The EXAFS data showed that

after just fifteen minutes of exposure at 22 uC, 25% of the Ag was

transformed into Ag2S (Figure 3B).The second line of evidence

comes from the detection of Ag intimately associated with TiO2-

NP aggregates[25] in our surface soils in the Slurry+AgNPs

treatment (Figure S1), suggesting the potential for partial oxidation

of Ag2S or AgNPs over the course of the experimental exposure,

and that TiO2-NPs and other metal-oxide nanoparticles found in

soils and biosolids may serve as a sink and/or carrier for Ag in the

environment. Both lines of evidence are consistent with literature

demonstrating oxidation and sulfidation of Ag in soils [20,24] and

biosolids [17,18].

All five plant species accumulated significantly more Ag in

aboveground tissues in the Slurry+AgNO3 treatment than in the

Controls or Slurry (Table S1). Ag bioaccumulation tended to be

lower for the AgNP treatment, yet all species except for M.

vimineum and L. cardinalis had significantly higher tissue Ag

concentration in the AgNP treatments than in Control or Slurry

treatments. Interestingly, M. vimineum and L. cardinalis had the

highest tissue Ag concentrations of any plants in both Ag

treatments, but the high variability obscured statistical differences

in the AgNP treatments. Some fraction of the silver associated with

plants could have been due to rain associated spray, though the

similar concentrations in the low creeping M. vimineum and the

erect and tall L. cardinalis may suggest that it was not simply due to

raindrop spray.

AgNPs impacted above and belowground plant biomass

Out of the 5 species we planted in the mesocosms, only

Microstegium vimineum showed sensitivity to the AgNP treatment,

growing 32% less aboveground biomass in the Slurry+AgNP

treatment as opposed to the Slurry treatment (Figure 4A). While

Ag toxicity may be tied to uptake in M. vimineum, Lobelia cardinalis

had similar Ag concentrations in aboveground biomass, yet

showed no evidence of toxicity. Thus uptake does not necessarily

lead to growth inhibition, and it may be that different species of

plants have different susceptibilities to Ag induced toxicity [26].

The Slurry+AgNP treatment also affected root biomass, in that

there were significantly more roots in the shallowest (0–1 cm) soils

of the Slurry+AgNP mesocosms than in the Slurry alone

mesocosms (Figure 4B). However, total root biomass did not

differ significantly between slurry treatments (Figure S2). Given

the difficulty in separating roots out by species, it is not known

whether these effects were due to a change in rooting depth and

allocation of one, many, or all species.

AgNPs impacted microbes

Microorganisms had broad sensitivity to the AgNP treatment,

and we observed significant changes in their abundance, function,

and community composition across our experiment. At the end of

the experiment (Day 50), microbial biomass in 0–1 cm soils in the

Slurry+AgNP treatment was significantly lower than the Slurryonly treatment (Figure 5A). Given that microbial biomass typically

increases with increasing root biomass[27], and given the increase

in root biomass in the 0–1 cm soils in the Slurry+AgNPs, this

decrease in microbial biomass is opposite of our expectations. We

hypothesize that the significant reductions in microbial biomass

and changes in microbial activity were due to Ag+ released from

partially sulfidized AgNPs[19,20].

The most notable changes in microbial activity in the

Slurry+AgNPs treatment were an increase in the flux of nitrous

oxide (N2O) as measured on Day 8 (Figure 5B), and lower

extracellular enzyme activity as compared to the Slurry treatment

at Day 50 (Figure 5C, D). The N2O flux was 350% higher in the

Slurry+AgNPs treatment than in the Slurry only treatment on Day

8, a dramatic increase given that N2O is both an important

greenhouse gas with 296 times the global warming potential of

CO2[28], and N2O is also the dominant stratospheric ozone

depleting substance[29]. We did not observe this difference in

N2O flux on Day 50 (data not shown), however the activity of

extracellular enzymes on Day 50 showed that differences persisted

in microbial activity. These enzymes are often used as indicators of

the microbial potential to decompose organic matter [30]. Both

leucine aminopeptidase (degrades amines; Figure 5C) and

phosphatase (degrades phospho-ester bonds; Figure 5D) decreased

in concert with microbial biomass in the Slurry+AgNPs treatment

compared to Slurry-only. Though neither difference was significant as measured by ANOVA post-hoc tests, both phosphatase

activity and leucine aminopeptidase activity were tightly correlated

to microbial biomass (r2= 0.61, p,0.0001 and 0.56, p,0.0001

respectively). This suggests that these changes in activity may have

been driven by changes in microbial abundance.

Changes in microbial biomass and activity were accompanied

by changes in microbial community composition. We examined

bacterial community composition using a genetic fingerprinting

technique (terminal restriction fragment length polymorphism; TRFLP) to assess the similarity in the bacterial 16S-rRNA genes

from 0–1 cm soils sampled on Days 0, 1, and 50. Overall, after

silver addition treatments differed in operational taxonomic unit

richness (OTU; each fragment corresponded to an OTU,

analogous to the species concept in other organisms) and bacterial

community composition (Figure 5E). An examination of OTU

Figure 4. Mesocosm plant aboveground and belowground

biomass affected by Ag. A Aboveground plant biomass of

Microstegium vimineum, B root biomass in 0–1 cm soils. Error bars are

standard error of the mean, and shared letters denote no significant

difference at p,0.05 between treatments differences (n = 6)

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richness showed that, prior to biosolid treatment, all treatments

were statistically equivalent with an average of 150 6 9 OTUs.

On Day 1, the Slurry+AgNPs richness declined to 101 6

22 OTUs, significantly lower than the control richness (142 6

23 OTUs) or Slurry only treatment (136 6 25 OTUs). By Day 50,

the richness for all treatments was statistically similar again, with

an average of 112.9 6 17.1 OTUs. Our NMS ordination

complemented our OTU richness trends, and showed that just

one day post-dosing, the Slurry+AgNPs treatment community was

significantly different from Slurry as seen in the increased distance

between treatment averages (Fig 5E; Bray-Curtis ANOSIM value

R = 0.3444, p,0.0117). By Day 50, the treatments were still

divergent from their respective Day 0 and Day 1 values, but were

no longer significantly different from each other.

Effects of AgNO3 treatment # AgNP treatment

While AgNO3 was added at a four-fold higher concentration to

act as a positive control, its impact on plants and microbes was

never greater than that of the AgNP treatment. In terms of

observed plant effects, both the decrease in M. vimineum biomass

and the increase in 0–1 cm root biomass relative to the Slurry

treatment were of similar magnitude for both Slurry+AgNPs and

Slurry+AgNO3 (Figure 4A, B). The magnitude of the AgNO3

treatment effect on microbial abundance, community composition, and function was also consistently equal to or less than the

effects of AgNP treatment. Microbial biomass and enzyme activity

on Day 50 were not significantly different between Slurry+AgNO3

and Slurry only (Figure 5A, C, D). Nitrous oxide fluxes on Day 8

were not significantly different between the Slurry and Slurry+AgNO3 treatments. Similar to Slurry+AgNPs, bacterial OTU

richness declined (from 150 6 9 down to 109 6 11 OTUs), and

bacterial community composition as indicated by T-RFLP was

also different between Slurry and Slurry+AgNO3 on Day 1 (BrayCurtis ANOSIM value R = 0.31, p,0.005).

The similar or stronger effects of AgNP treatment on activity,

abundance, and composition of plants and microbes compared to

AgNO3 treatment—even though AgNO3 was applied at a 4-fold

higher total Ag concentration—was unexpected given earlier

comparative studies[14]. We hypothesize that the AgNP treatment

Figure 5. Microbial abundance, activity, and composition affected by Ag. A Microbial biomass in 0–1 cm soils on Day 50 of the experiment;

B N2O flux from soil on day 8; C activity of the proteolytic extracellular enzyme leucine aminopeptidase (LAP), on day 50; D activity of the

organophosphorous degrading enzyme phosphatase on day 50; E NMS ordination of bacterial community composition with day of experiment

designated by shapes: Day 0 (triangles), Day 1 (squares), and 50 (circles); and treatment designated by colors: Control (white), Slurry (black),

Slurry+AgNPs (gray), and Slurry+AgNO3 (red). All error bars are standard error of the mean, and shared letters denote no significant difference at

p,0.05 between treatments in panels A–D (n = 6)

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provided a slow release of Ag+ to the ecosystem, whereas Ag added

as AgNO3 may have been immediately sequestered, perhaps as

Ag2S or Ag-sulhydryl compounds [31]. This is consistent with the

findings that partially sulfidized Ag NPs remained toxic to

Escherichia coli in pure culture[20] and thus may remain

bioavailable to plants and microbes [31].

Several factors were unresponsive to silver treatments. Total

aboveground and belowground plant biomass did not differ

significantly between the three slurry treatments (Figure S2),

despite the significant decrease in M. vimineum biomass in both Ag

treatments. Similarly, for the four plant species for which we

measured photosynthesis (Day 8 and Day 30), we saw no evidence

of a significant difference in photosynthesis in Slurry+AgNP or

Slurry+AgNO3 as compared to Slurry (Table S2). The impacts of

Ag exposure on microbes and plant roots were confined to the

shallowest soil layers, with microbial and root biomass in soils

below 1 cm showing no Ag effects (Figures S2 and S3). Although

slurry additions led to increased soil nutrient concentrations

(nitrate and phosphate; Figure S4), there were no differences

between the three treatments that received biosolid slurry, despite

changes in microbial abundance, community composition, and

activity. There were also no significant differences in CO2 or CH4

flux from soils receiving Slurry, Slurry+AgNO3, or Slurry+AgNPs

on either Day 8 or Day 50 (data not shown).

Conclusions

An estimated 60% of the average 5.6 million tons of biosolids

produced each year in the United States is land applied [32], and

represents an important and understudied route of exposure of

natural ecosystems to engineered nanoparticles. Our results show

that biosolids amended with AgNPs at environmentally relevant

concentrations and added to a diverse terrestrial ecosystem caused

ecosystem-level impacts. Specifically, the AgNP treatment led to

an increase in N2O fluxes, changes in microbial community

composition, biomass, and extracellular enzyme activity, as well as

species specific effects on aboveground plant biomass (i.e., on M.

vimineum). Moreover, for all of these parameters, the effect of the

AgNP treatment was as large as or larger than that of the

treatment with AgNO3 added at a four-fold higher concentration,

though whether this is due to differences in bioavailability of

AgNPs and AgNO3 when added to biosolids, saturated bioavailability of the AgNO3, or some other factor is beyond the scope of

this experiment. Importantly, given that this experiment used a

one-time application of biosolids, these impacts could be enhanced

by repeated applications, with further changes expected in

microbial and plant community composition and functioning in

the ecosystem.

Our results also suggest that while AgNPs may be transformed

in biosolids through oxidation and sulfidation, they still had an

impact on plants and microbes. Whether these impacts were

through direct or indirect plant or microbially mediated mechanisms remains unexplored. We have also demonstrated the

potential for several species of plants to take up Ag from AgNPs

in soils, though the extent to which different plant species

accumulated Ag varied greatly. Uptake and incorporation of Ag

into plant biomass suggests the potential for trophic transfer of Ag.

Future studies are aimed at identifying the bioavailable forms of

Ag and on the speciation of Ag taken up into plants.

Materials and Methods

Soils

The soils used in this experiment were surface mineral soils from

the floodplain of the Sandy Creek Restoration in Durham,

NC[33], which is located in the Duke Forest Teaching and

Research laboratory. Soils are of the Cartecay series (Coarseloamy, mixed, semiactive, nonacid, thermic Aquic Udifluvents)

and Chewacala series (Fine-loamy, mixed, active, thermic

Fluvaquentic Dystrudepts) and had 63.5% sand, 10.5% silt, 26

% clay, and 1.8% loss on ignition. Mesocosms were established in

the Duke Forest by first screening soils for materials larger than

10 mm, and then adding ,81 kg of soil to 21.5 gallon

polyethylene tubs (Rubbermaid, Wooster, USA) equipped with

drains at 10 cm and at the bottom.

Plants

Mesocosms were planted on June 23, 2009 with fifteen plants

(three replicates from each of five species) placed in random order

in fifteen of the sixteen slots in a 4 64 grid. Plants included four

species of plants native to NC meadows (Carex lurida, sedge; Juncus

effusus, rush; Lobelia cardinalis, forb; and Panicum virgatum, grass;

purchased from Mellow Marsh Farm, Silk Hope, USA), as well as

the non-native invasive C4 grass, Microstegium vimineum, collected

from the Duke Forest on June 23, 2009. For collection of M.

vimineum, no specific permits were required, and permission for

collection was granted by Judson Edeburn, Duke Forest Resource

Manager. Plants were watered daily with groundwater until they

were well established. The mesocosms were weeded twice before

the experimental treatments were applied to keep non-target

plants to a minimum. Non-target plants were collected when

mesocosms were sampled and were grouped together as ‘‘Others’’.

Biosolids

The biosolids used in these experiments were rated Class A EQ,

and were obtained from the South Cary Water Reclamation

Facility (Apex, USA) on July 15th, 2009 as dried pellets for use as

fertilizer (7.47 % moisture) with total N reported as 7.5%. The

addition rate was calculated to meet plant N demands during the 8

weeks of the experiment using established guidelines[21] adapted

for our 0.23 m2 mesocosms.

These biosolid pellets were used for their ease of transport,

storage, handling, and homogeneity, but they are not the typical

form of biosolids that are land-applied in many agroecosystems

and rangelands. They are dried, which decreases their volume.

They are sold to agricultural users, landscapers, and consumers for

use in dried and pelletized form [34]. To make a slurry from these

biosolids, 200 g of dried pellets were first rehydrated with 750 ml

of water and homogenized using an immersion blender (KitchenAid, St. Joseph, USA) on high for three minutes. The slurry then

sat for 2 minutes to allow the immersion blender to cool and the

pellets to soften. Pellets were then homogenized for another

3 minutes and the volume was brought to 1.5L with deionized

water.

Silver nanoparticles and dosing concentrations

Instead of freshly synthesized particles, we chose to use a

commercially available AgNP powder representative of what is

available for incorporation into consumer products (Nanoamorphous Materials, Los Alamos, USA). Particles were purchased as a

dry powder, and suspended to make a 250 mg Ag L21 suspension

in deionized water by sonicating them for 10 minutes at 100W

with a Sonicator 4000 equipped with a K inch diameter flat

titanium tip (Misonix, QSonica LLC, Newton, USA). TemperaAgNPs Impact Organisms and Ecosystem

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ture was controlled by placing the beaker used to make the

suspension in an ice-water bath. Particles in suspension had

diameters of 21617 nm as measured by TEM. Detailed

information on particle characterization has been previously

published [23]. For the positive control, we used AgNO3

(SigmaAldrich, St. Louis, USA).

Additions of Ag were chosen to fall within the range of

concentrations of Ag measured in the United States Environmental Protection Agency’s Targeted National Sewage Sludge Survey

(TNSSS)[22], which reported concentrations between 2 and

195 mg Ag kg21

, and a mean 6 SD of 20622 mg Ag kg21,

excluding one outlier at 856 mg Ag kg21

. For AgNPs, we added

9.9 mg Ag per mesocosm of AgNPs in addition to the 1.5 mg Ag

in the 200 g of biosolids (57 mg Ag kg21 biosolids). This rate of

addition represents the top end of the 95% confidence interval of

the TNSSS. With the AgNO3 treatment, we wanted to have a

positive control with a better documented toxin, and so we added

44 mg Ag as AgNO3 per mesocosm in addition to the 1.5 mg Ag

in the 200 g of biosolids (228 mg Ag kg21 biosolid), which is twice

the 99% confidence interval of the TNSSS [22], but lower than

the highest value detected in the TNSSS.

Treatments

For the mesocosm experiment, manipulations were done on

August 25, 2009. There were six replicates of each of four

treatments, which were randomly assigned to each of the 24

mesocosms: ‘‘Controls’’ which received only DI water; ‘‘Slurry’’

which received 200 g biosolid slurry only; ‘‘Slurry+AgNPs’’, which

received 200 g biosolids and 9.9 mg AgNPs; and ‘‘Slurry+AgNO3’’, which received 200 g biosolids and 44 mg Ag as

AgNO3. The biosolids in the Slurry treatment samples were

homogenized on the day of treatment with a milkshake maker

(DrinkMaster Drink Mixer, Hamilton Beach, Southern Pines,

USA) for 1 min just prior to slurry application, then promptly

added to the soil surface of the mesocosms with care taken not to

apply slurry to the foliage. The only difference for the

Slurry+AgNPs and Slurry+AgNO3 treatments from the Slurry

treatments was that the Ag was added during homogenization with

the milkshake maker. Containers used for holding water or slurries

were rinsed with 500 ml well water, which then was added to the

soil surface.

For the separate batch assay for generation of samples that had

a high Ag concentration yielding a strong enough signal to be

analyzed by EXAFS (Figure 3B), a slurry was prepared as for the

mesocosm experiment. Then, to a subsample of biosolids (5 g dry

weight equivalent) 10.8 ml of 250 mg Ag L21 stock of AgNPs was

added, shaken at 120 rpm for 15 minutes, frozen in liquid

nitrogen, and freeze dried.

Sampling

We sampled the top 1 cm of surface soil using three replicate

1 cm diameter cores immediately before treatment (Day 0), and 1

and 50 days after treatment application. These samples were

preserved for molecular and enzymatic analyses by freezing at

280 uC. We destructively harvested all aboveground biomass on

Day 50, and subsequently removed five 5 cm diameter x 15 cm

deep soil cores from each mesocosm to estimate root and

microbial biomass, as well as Ag content for the soil depth

increments 0–1, 1–5 and 5–10 cm.

To sample CO2, CH4, and N2O emissions from soil, we used

10 cm gas collars installed to 10 cm depth placed in the one space

in the 4 64 planting grid which was left empty at planting. Caps

were installed on the gas collars immediately prior to gas

collection, and trace gases were allowed to accumulate. Gases

were sampled every 20 minutes for one hour after capping the

collars by withdrawing 10 ml of gas from the headspace, and

transferring that to 9 ml evacuated serum vials.

In order to obtain an index of available NH4

+

, NO3

2, and

ortho-phosphate throughout the course of this experiment, Plant

Root Simulator (PRSTM) probes were installed (17.5 cm2 area per

probe; Western Ag Innovations, Saskatoon, Canada), with two

anion and two cation probes installed per mesocosm between the

1st and 2nd, or 3rd and 4th rows of plants. PRSTM-probes were

installed for four time periods: one pre-experiment period (8/5/

2011 to 8/26/2009), and three post-treatment time intervals (8/

26/2011 to 9/16/2011, 9/16/2009 to 10/7/2009, and 10/07/

2009 to 10/15/2009). Probes were rinsed in the field when

removed, and fresh probes were installed in their place. Sampled

probes were brought back to the lab in polyethylene bags, and

were then extracted with 0.5M HCl for later analysis

Sample processing and analysis

Soil core increments were sieved (2 mm opening) to remove

rocks and roots, and homogenize samples. Subsamples were also

used for microbial biomass. A separate subsample of soil from each

was dried and ground for Ag analysis. Soils were microwavedigested using EPA method 3052 [35]. All silver concentrations

were measured using ICP-MS (Agilent 7500CX, Santa Clara, CA,

USA) as previously described[24]. The method detection limit

varied by sample due to differences in sample mass. The average

method detection limits were 0.066, 0.037, 0.030, 0.014, and

0.023 mg Ag/kg for 0–1 cm roots, 1–5 cm roots, 5–10 cm roots,

shoots, and soils, respectively.

The aboveground plant biomass was split by species in the field,

while roots were split by depth increment, not species. All biomass

was then oven dried and weighed, and subsamples were ground

for biomass Ag analysis. To prepare plant samples for Ag analyses,

samples were digested using a modification of published methods

[36] using sequential digestion of ,0.1 g of plant material in 2 ml

of concentrated HNO3 and 0.5 ml of 30% H2O2 heated to 70 uC

for 1 hour, and 6 ml concentrated HCl at 70 uC for 1 hour

[36,37].

The fate of Ag in the soil was investigated beyond just

concentration by using X-ray absorption spectroscopy (XAS)

and transmission electron microscopy (TEM). The extended x-ray

absorption fine structure (EXAFS) region was analyzed with linear

combination fitting (LCF), with EXAFS spectra collected at the

Stanford Synchrotron Radiation Laboratory (SSRL) BL4-1. Both

the EXAFS data collection and LCF analysis have been described

in detail in previously published studies.

For detailed TEM analysis, we used an FEI Titan 80–300 field

emission TEM operating at 200 kV. The microscope is equipped

with an energy dispersive X-ray spectrometer (EDAX r-TEM) for

chemical analysis. The chemistry of individual particles of interest

was examined with a lateral spatial resolution of approximately

1 nm using the Scanning TEM mode. High-resolution TEM

analyses were used to examine the crystal structure of the particles

of interest.

To measure microbial biomass, we used a standard method,

substrate induced respiration (SIR) biomass [38]. Briefly, 4 g of

soil was weighed into 40 ml I-chem vials into which was added

10 ml of yeast extract (12 mg mL21

; BD Difco, Sparks, USA).

Following yeast addition, samples were shaken on an orbital

shaker at 120 rpm and CO2 concentration was measured after

30 min, 2, and 4 hours using an infrared gas analyzer (Li-Cor

6400, Lincoln, USA).

Trace gas fluxes from field soils were determined by analyzing

gas samples from the mesocosm gas collars on a Shimadzu GC

AgNPs Impact Organisms and Ecosystem

PLOS ONE | www.plosone.org 7 February 2013 | Volume 8 | Issue 2 | e57189
with FID and ECD detectors (Shimadzu, Columbia, USA). The

rate of accumulation over time of CO2, CH4, and N2O were first

examined for linearity, with data with an r2

,0.7 being set to zero

if the nonlinearity was due to no slope. Slopes were then converted

into mg of C or N m22 hour21

.

Extracellular enzyme activity was determined for samples taken

in surface 0–1 cm soils sampled at the end of the experiment

(October 14, 2009). Activity of the following three enzymes was

measured using published methods[39]: aryl-sulfatase, leucine

aminopeptidase, and alkaline phosphatase. Samples were frozen

prior to analysis, which may influence the absolute rates of this

potential assay, but should have affected all soils in the same way

leaving comparisons between treatments valid[40].

For each soil sampling time-point and treatment, total DNA was

extracted from four individual 0.25 g replicates using the PowerSoilH DNA Isolation Kit (MoBio Laboratories, Solana Beach,

USA). All DNA extractions were performed following the

manufacturer’s protocol with slight modification. The manufacturer’s bead beating step was extended to 15 min. Following

extraction, DNA was stored in elution buffer at 220 uC until

further use. DNA quality and concentration was verified using an

ND-1000 NanoDrop spectrophotometer (NanoDrop Technologies, Wilmington, USA). DNA concentrations ranged from 5 to

20 ng/mL and quality was verified to ensure minimal phenolic or

protein contamination.

PCR amplification of bacterial 16S SSU rDNA was performed

following the protocol described in Lukow et al. [41] with slight

modification. The forward primer (27F) was fluorescently labeled

with 6-carboxyfluorescein. One uL of template DNA was used in a

100 mL reaction. Bovine serum albumin (10 mg) was added to limit

primer dimer formation and humic acid interference. The number

of cycles was extended to 40 to increase amplification. The

presence of PCR amplicons of the correct length was verified by

visualization on a 1% agarose gel containing 0.1% ethidium

bromide. PCR amplicons were purified using Qiagen PCR

Purification Kits (Qiagen, Hilden, Germany) following the

manufacturer’s protocol. Samples were eluted to a final volume

of 50 mL in elution buffer. Final PCR product concentrations and

purity were verified as described above.

Restriction enzyme digests were performed using previously

described methods [41]. Purified PCR product was digested with

HaeII and MspI (New England Bio-labs Inc., Beverly, USA). The

mixture was incubated at 37 uC for 2 hours, followed by 15 min

heating at 65 uC for enzymatic inactivation. All samples were

desalted by spin column filtration, and fragment analysis was

carried out using an Applied Biosystems 3100 capillary sequencer

(Foster City, USA) with POP6 polymer and ROX-labeled

MapMarker 1000 size standards (BioVentures, Inc., Murfreesboro, USA). All analyses were carried out at the Duke University

DNA Analysis Facility (Durham, USA) following standard

procedures.

Statistics

To examine data for treatment-level effects, data were

compared using analysis of variance (ANOVA). Data were first

tested for normality using the Shapiro-Wilk test, and then for

equal variance. If data were found to be normal and have equal

variance, data were analyzed using ANOVA with Holm-Sidak

post-hoc comparisons. For data that were either non-normal or

had unequal variance, Kruskal-Wallis ANOVA by ranks was used,

with Student-Newman-Keuls post-hoc tests. These analyses were

all conducted using SigmaStat 4 (Systat Software Inc., San Jose,

USA)[42]. For all ANOVA tests, we used a group-wise a = 0.05.

Restriction fragment profiles (from T-RFLP digests) were

visualized using GeneScan v3.7.1 software (Applied Biosystems,

Foster City, USA). Raw fragment data were imported into T-REX

[43] which was then used to select true peaks in all profiles, align

restriction fragments, and then create presence/absence matrices

for all restriction fragments. All restriction fragments smaller than

50 base pairs were excluded to ensure primer dimer fragments

were excluded, and remaining peaks were also excluded if they did

not meet a peak height threshold of 50 relative fluorescent units.

Operational taxonomic unit (OTU) richness (total number of taxa)

of T-RFLP profiles was determined for all profiles and compared

to control mesocosms using a two tailed student’s t-test (p,0.05).

PC-ORD[44] was used to generate ordinations from presence/

absence data by nonmetric multidimensional scaling (NMS) with

Sørenson distance using the medium auto-pilot function [45].

Each point on the ordination represents an average of the six

experimental replicates for a given day/treatment combination,

with points closer together being more similar than those that are

farther apart. Ordination plots were analyzed with analysis of

similarity (ANOSIM) performed using the PAST statistical

software to determine if ordination clusters by treatment were

statistically similar [46]. The null hypothesis assumes that there are

no differences between community composition at given sampling

dates [47], and is assessed by examining the R value and p-value.

The R-value indicates how closely related each replicate within a

treatment is to other replicates within that treatment, as opposed

to replicates in other treatments, while p-values suggest the

probability that the R-value is due to random chance as opposed

to a treatment effect. R-values range from 21 to 1, and an R-value

between 0 and 1 indicates that the treatments are more similar to

their own replicates than those from other treatments, while an Rvalue of 21 to 0 indicates the replicates in a given treatment are

more similar to replicates from other treatments than replicates

from their own treatment.

Supporting Information

Figure S1 TEM image of TiO2 nanoparticle aggregate

with associated Ag. A TEM image of TiO2 nanoparticle

aggregate from surface soils in Slurry+AgNPs treatment, and B

EDX spectra of area on aggregate highlighted with a white circle

in A showing Ag associated with TiO2-nanoparticle aggregate

(adapted from Figure 5 in reference 25, http://dx.doi.org/10.

1039/C2EM10809G, and reproduced by permission of The

Royal Society of Chemistry)

(TIF)

Figure S2 Total aboveground plant biomass by species,

and belowground root biomass by depth. Shared letters

denote no significant difference at p,0.05 between treatments for

either total aboveground or belowground biomass

(TIF)

Figure S3 Microbial biomass in soils. A 0–1 cm soils, B 1–

5 cm soils, and C 5–10 cm soils. Shared letters denote no

significant difference at p,0.05 between treatments, and error

bars are standard error of the mean (n = 6)

(TIF)

Figure S4 Plant Root SimulatorTM Resin available NO3

2

and phosphate. Shared letters denote no significant difference

at p,0.05 between treatments, and error bars are standard error

of the mean (n = 6)

(TIF)

Table S1 Plant biomass, Ag concentration, and Ag

content by mesocosm. Shared letters denote no significant

AgNPs Impact Organisms and Ecosystem

PLOS ONE | www.plosone.org 8 February 2013 | Volume 8 | Issue 2 | e57189
difference at p,0.05 between treatments within a plant species

and sampling date, and error terms are one standard deviation

(DOCX)

Table S2 Photosynthetic rates for the four plant species

measured. Shared letters denote no significant difference at

p,0.05 between treatments within a plant species and sampling

date, and error terms are standard error of the mean (n = 8

measurements from plants from the six replicate mesocosms per

treatment)

(DOCX)

Acknowledgments

The authors would like to thank Medora Burke-Scoll, Brooke Hassett, Wes

Willis, Jonathan Bills, Sarah Diehl, Kevin He, Eddie Liu, Benjamin

Espinasse, and Judson Edeburn & the Duke Forest staff for their help in the

field and the lab, and Melanie Auffan for preparing our nanoparticle

suspension. Portions of this research were carried out at the Stanford

Synchrotron Radiation Lightsource (SSRL) a national user facility at the

Stanford linear accelerator center (SLAC National Accelerator Laboratory)

operated by Stanford University on behalf of the U.S. Department of

Energy, Office of Basic Energy Sciences. We thank SSRL beamline

scientists John Bargar and Joe Rogers for technical support during our

experiments.

Author Contributions

Editing and revising the manuscript: BPC CLA SA CKG MFH BK GVL

BMM BCR CJR JMU JPW LY ESB. Conceived and designed the

experiments: ESB JPW BPC CKG CLA CJR. Performed the experiments:

ESB BMM JPW BK BPC. Analyzed the data: BMM BK BPC SA JMU

BCR GVL. Contributed reagents/materials/analysis tools: ESB JPW

MFH CJR. Wrote the paper: BPC ESB.

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