Discerning the sources of silver nanoparticle in a terrestrial food chain

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Discerning the sources of silver nanoparticle in a terrestrial food chain by stable isotope tracer technique Fei Dang, Yuanzhen Chen, Yingnan Huang, Holger Hintelmann, Youbin Si, and Dong-Mei Zhou Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/acs.est.8b06135 • Publication Date (Web): 12 Mar 2019 Downloaded from http://pubs.acs.org on March 17, 2019

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Environmental Science & Technology

1

Discerning the sources of silver nanoparticle in a terrestrial food

2

chain by stable isotope tracer technique

3

Fei Dang †*, Yuan-Zhen Chen†, §, Ying-Nan Huang†, Holger Hintelmann‡, You-Bin

4

Si§, Dong-Mei Zhou† *

5

†

6

Science, Chinese Academy of Sciences, Nanjing 210008, P.R. China

7

‡

8

K9J 0G2, Canada

9

§

Key Laboratory of Soil Environment and Pollution Remediation, Institute of Soil

Water Quality Centre, Trent University, 1600 West Bank Drive, Peterborough, ON

School of Resources and Environmental Science, Anhui Agricultural University,

10

Hefei 230036, China

11

*Corresponding

12

Fei Dang; Dong-Mei Zhou, Key Laboratory of Soil Environment and Pollution

13

Remediation, Institute of Soil Science, Chinese Academy of Sciences, Nanjing

14

210008, P.R. China

15

Tel: +86-25-86881179

16

E-mail: [email protected]; [email protected]

authors:

17 18 19 20 21 22 1

ACS Paragon Plus Environment


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Abstract. The increasing use of silver-containing nanoparticles (NPs) in commercial

24

products has led to NP accumulation in the environment and potentially in food webs.

25

Identifying the uptake pathways of different chemical species of NPs, such as

26

Ag2S-NP and metallic AgNPs, into plants is important to understanding their entry

27

into food chains. In this study, soybean Glycine max L. was hydroponically exposed

28

to Ag2S-NPs via their roots (10 − 50 mg L-1) and stable-isotope-enriched

29

via their leaves [7.9 µg (g fresh weight)−1]. Less than 29% of Ag in treated leaves (in

30

direct contact with

31

whereas almost all of the Ag in soybean roots and untreated leaves sourced from

32

Ag2S-NPs. Therefore, Ag2S-NPs are phytoavailable and translocate upwards. During

33

trophic transfer the Ag isotope signature was preserved, indicating that accumulated

34

Ag in snails most likely originated from Ag2S-NPs. On average, 78% of the Ag in the

35

untreated leaves was assimilated by snails, reinforcing the considerable trophic

36

availability of Ag2S-NPs via root uptake. By highlighting the importance of root

37

uptake of Ag2S-NPs in plant uptake and trophic transfer to herbivores, our study

38

advances current understanding of the biogeochemical fate of Ag-containing NPs in

39

the

109AgNP)

109AgNPs

was accumulated from root uptake of Ag2S-NPs,

terrestrial

2


environment.

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INTRODUCTION

41

The rapidly expanding use of engineered silver (Ag)-containing nanoparticles (NPs)

42

as fungicides and bactericides has raised concerns regarding their impact on wildlife

43

and on humans. Sulfidation is a major transformation process of Ag-containing NPs

44

in wastewater treatment and in soils.1-5 As a result, Ag2S builds up in agricultural soils.

45

Silver sulfidation leads to a marked reduction in their bioavailability and toxicity.6-8

46

Plants may be simultaneously exposed to metallic AgNPs via foliar exposure, given

47

that foliar spraying of AgNPs is an attractive approach to fight plant pathogens in

48

agriculture.9,

49

wind, thus reaching unintended targets.11 How and to what extent the various NP

50

sources, defined herein as NP exposure pathways, each associated with a different

51

chemical speciation (i.e., foliar uptake of AgNPs and root uptake of Ag2S-NPs),

52

contribute to NP phytoavailability and trophic transfer remain largely unknown. We

53

recently showed that foliar exposure to metallic AgNPs leads to greater Ag

54

accumulation but to less toxicity to crop plants compared to root exposure at similar

55

exposure doses.12 That study illustrated the pathway-specific effect on plants.

56

However, the chemical speciation of Ag-containing NPs was not considered, which

57

impeded an extension of the data to more environmentally relevant settings.

10

In addition, a fraction of the released AgNPs can be dispersed by

58

Identification of the major sources of Ag-containing NPs in organisms is

59

fundamental to an understanding of the biogeochemical cycling of NPs and to

60

controlling their risk to wildlife. Trophic transfer of metal-based NPs (e.g., CeO2,

61

La2O3, TiO2 and AuNPs) in terrestrial food chains has been reported.13-17 A recent 3



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study showed that dietary ingestion, rather than direct uptake of metallic AgNPs from

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the surrounding environment, was the dominant process of NP accumulation in a

64

terrestrial snail and thus revealed the previously unrecognized role of the trophic

65

transfer of AgNPs.18 However, many questions regarding the factors regulating the

66

trophic transfer of NPs remain unanswered. An especially important issue is the

67

relative importance of the different sources of Ag-containing NPs during trophic

68

transfer.

69

The paucity of information can be attributed to a lack of suitable methods to

70

quantitatively distinguish NP sources. Parallel exposures, as well as the variable

71

background metal contents among different individuals or systems, complicate the

72

detection of small changes in NP uptake based on measurements of metal

73

concentrations alone.19,

74

methodologies. The stable isotope tracer method is a powerful approach that has been

75

successfully manipulated to allow assessments of the fate of metal-based NPs in

76

exposure media21,

77

concentrations.19,

78

concentrations as low as 10 ng L−1 AgNPs, thus demonstrating the advantage of the

79

isotope tracer approach.24 More importantly, this tracer methodology could be used to

80

quantitatively differentiate metal sources in a single system.26,

81

attempt has rarely been made on NPs in literature.28

82 83

22

20

Thus, there is a clear need to explore alternative tracing

as well as monitoring of the biouptake of NPs at realistic

23-25

Quantitative detection of AgNPs has been achieved at

In this study, we used isotopically enriched

109Ag

27

However, such

as a tracer to track specific

AgNP source, thus extending our prior work on single-exposure scenario to include 4


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84

parallel exposures of NPs containing different Ag species and the investigation of a

85

complex environmental system, by discriminating between the different geochemical

86

sources and quantifying their relative importance in plant uptake. More importantly,

87

we were able to further elucidate the effects of the particle sources on the dietary

88

transfer of NPs internalized by plants to the terrestrial snail Achatina fulica. Taken

89

together, the results improve our understanding of the biogeochemical cycle of

90

Ag-containing NPs and are fundamental to the control of their risk to wildlife, by the

91

identification of the major sources of Ag-containing NPs in organisms.

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MATERIALS AND METHODS

93

Nanoparticle synthesis and characterization.

94

isotopically enriched metallic Ag foil (109Ag at 98.56%, Oak Ridge National Lab.

95

USA) using a reported method.21, 29 Ag2S-NPs were synthesized by reacting AgNO3

96

with elemental sulfur.5 Both nanoparticles were purified by 3-kDa ultracentrifugation

97

filter at 4000 × g filtration (maximum pore size ~3 nm, Amicon Ultra-15, Millipore,

98

USA),30 redispersed in Milli-Q water and stored in the dark at 4°C. For details see the

99

Supporting Information (SI). Silver concentrations were determined by HNO3

100

digestion with ICP-MS (iCAP QC, Thermofisher, USA). The morphology, diameter

101

and elemental composition of NPs were determined by transmission electron

102

microscopy coupled with energy dispersive X-ray spectrometry at an accelerating

103

voltage of 200 kV (TEM-EDS, JEOL JEM-2100, Japan), and particle size was

104

calculated based on the results of at least 300 particles using Nano Measure software.

105

AgNP suspensions were characterized by UV-vis spectrophotometry at 300−600 nm

109AgNPs

5


were prepared with an


106

(UV-2700, Shimadzu, Japan). Ag2S-NP suspensions were freeze-dried (Alpha 1-2

107

LDplus, Germany) and verified using X-ray diffraction analysis (XRD, Rigaku

108

Ultima IV diffractometer with a Cu Kα source). Hydrodynamic diameters

109

(intensity-weighted and number-weighted) and zeta potential of Ag2S-NPs were

110

measured in the plant growth medium using dynamic light scattering (DLS,

111

NanoBrook 90 Plus PALS, Brookhaven, US) and a Zetasizer Nanos ZS (Marvern,

112

UK). The dissolution of Ag2S-NPs was determined in the growth medium after 3 days

113

using a 3-kDa centrifuge filter to separate the particles from the dissolved species.

114

Plant exposure. Soybean seedlings (Glycine max L.) were hydroponically grown for

115

21 days, as described in the Supporting Information. They were concurrently exposed

116

to Ag2S-NPs via their roots and to

117

Information (SI) Figure S1). Thus, the experiment consisted of four treatments: i) a

118

control without Ag2S-NPs or

119

7.9 µg

120

7.9 µg 109AgNP g−1), and iv) a High group (50 mg Ag2S-NPs L−1 and 7.9 µg 109AgNP

121

g−1). The control was used to account for the interference of background Ag levels.

122

Each treatment was replicated four times, with two seedlings per polypropylene pot.

109AgNP

109AgNPs

109AgNPs,

via their leaves for 7 days (Supporting

ii) a Low group (10 mg Ag2S-NPs L−1 and

(g fresh weight)−1]), iii) a Medium group (20 mg Ag2S-NPs L−1 and

123

The use of Ag2S-NPs in the growth medium was to represent sources for in situ

124

formation and/or biosolid application. The Ag2S-NP concentrations (10, 20 and 50 mg

125

Ag L−1) were comparable to those used in previous hydroponic experiments,31, 32 but

126

likely much higher than those typically expected in pore water of soils amended with

127

Ag2S-NPs or Ag-containing sludge,2,

5, 33

as well as the likely environmental

6


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concentrations of nano-Ag in sludge treated soil in the EU (in the range of µg kg-1).34

129

The Ag2S-NP suspensions were renewed every 3 days.

130

Simultaneously, the 13th to 15th leaves (referred as treated leaves hereafter) from

131

the apex of V6 stage soybeans35 were carefully sprayed with 109AgNPs three times per

132

day (total volume of 60 mL per day) using acid-washed polypropylene aerosol

133

sprayers.12 The high spraying frequency minimized Ag loss resulting from spraying

134

suspension dripping off the leaves.12 An isotope-enriched

135

freshly prepared each day to minimize potential particle aggregation and dissolution.

136

The foliar dose of

137

used during treatments for plant pathogens.10 Untreated leaves were covered by black

138

plastic only during spraying, to avoid unintentional exposure to

139

were covered with foil to avoid potential

140

and separated from each other to avoid cross-contamination. The natural abundance of

141

Ag isotopes in growth medium spiked with Ag2S-NPs indicated negligible

142

contamination.

109AgNPs

109AgNP

suspension was

was 7.9 µg Ag (g fresh weight)−1, comparable to that

109Ag

109AgNP.

The pots

contamination of the growth medium

109AgNP

143

After a 7-day exposure, the maximum photochemical activity of PSII (Fv/Fm), the

144

performance index based on the absorption of light energy (PIABS), and the absorption

145

flux per reaction center (ABS/RC) were measured in the dark-adapted 13th leaves

146

using PAR-FluorPen FP 100-MAX-LM-D (Photon Systems Instruments, Brno, Czech

147

Republic) (n = 4). No attempt was made to compare these chlorophyll fluorescence

148

parameters between treated and untreated leaves within one plant. Treated leaves (13th

149

to 15th leaves in direct contact with 109AgNPs), untreated leaves (leaves with no direct 7



150

contact to 109AgNPs) and roots were harvested and weighed. Samples serving as snail

151

food were rinsed thoroughly with Milli-Q water and kept fresh at 4 oC. Samples for

152

TEM-EDS and single-particle ICP-MS (spICP-MS) analysis were immediately

153

prepared. Samples for Ag stable isotope analysis were washed as described below,

154

frozen in liquid nitrogen and stored at -80 oC.

155

Snail exposure. Terrestrial snails of the species Achatina fulica (soft tissues of 2.7 ±

156

0.4 g, dry weight) were acclimated under a 14-10 h light/dark cycle at a relative

157

humidity of 80% for one week, and fed Ag-free lettuce (< 4.2 ng g-1) at 4% of the

158

snails’ body weight per day. The snails were not fed 48 h before the experiment to

159

assure their maximum ingestion of food.

160

They were divided randomly into two groups. One group (n = 5) was fed

161

untreated leaves collected from the High group (the mixture of untreated leaves from

162

all plants), while the other group (n = 5) received leaves from the control.

163

Immediately before feeding, leaves were cut into small sections, mixed thoroughly

164

and supplied ad libitum for 4 h per day. Over a period of 5 days, feces were collected

165

cumulatively, and unconsumed leaves were removed and weighed every day. During

166

feeding experiments no snail died. The snails were then removed, rinsed with 10 mM

167

L-cysteine and Milli-Q water,18 and fed Ag-free lettuce for 48 h to allow clearance of

168

the Ag-containing material from the gut before harvest.

169

Sample preparation and Ag determination. Plant tissues for stable isotope analysis

170

were thoroughly rinsed with Milli-Q water, 10 mM HNO3,36 10 mM EDTA37 and

171

finally again with Milli-Q water to remove loosely adsorbed Ag. Plant tissues (n = 4, 8


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both plants per pot), snail soft tissues (n = 5) and feces (n = 5) were freeze-dried

173

(ALPHA 1-2 LD plus, Christ, Germany), digested with 65% HNO3 in a microwave

174

oven (Ethos one, Milestone, Italy) according to EPA2001b, and their Ag stable

175

isotope concentrations determined by ICP-MS. Digestion blanks, matrix spikes and

176

certified reference materials (GBW 10020, citrus leaf, Chinese Academy of

177

Geophysical Sciences, China; LUTS-1, lobster hepatopancreas, National Research

178

Council Canada) were included for QA/QC. Recovery rates were 100.0 ± 11.7% for

179

plant tissues and 94.0 ± 3.2% for snail tissues. Instrument biases were calibrated by

180

comparing the measured isotope ratio with the natural isotope ratio in an Ag ion

181

standard solution.

182

Roots, treated and untreated leaves from the High group were analyzed for

183

Ag-containing NPs by TEM-EDS and spICP-MS (7900, Agilent, USA), as described

184

in our recent study.12 Details can be found in the Supplementary Information.

185

Data analysis. One-way analysis of variance (ANOVA) followed by Turkey’s

186

post-hoc analysis was performed to identify statistical differences among treatments

187

(p < 0.05).

188

An isotope tracing technique was used in this study to source the foliar uptake of

189

109AgNP.

190

abundances of 0.518 and 0.482, respectively.29 A significant difference between the

191

Ag isotope ratio in the sample and the natural value therefore indicates the presence of

192

the isotopic tracer

193

that sample can also be accurately quantified.

Silver has two natural stable isotopes,

109AgNPs

or

109Ag

107Ag

and

109Ag,

derived therefrom. The uptake of

9


with natural

109AgNPs

in


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The total Ag concentration (CAg) in a sample, defined as the sum of the measured

194

107Ag

and

109Ag

isotopes by ICP-MS (referred as C107Ag and

195

concentrations of

196

C109Ag, respectively), consisted of pre-existing Ag (Ccontrol), Ag from the foliar uptake

197

of 109AgNPs (CAgNPs) and Ag from the root uptake of Ag2S-NPs (CAg2S-NPs) [Eq. (1)].

198

C Ag  C107 Ag  C109 Ag  Ccontrol  C AgNPs  C Ag2 S  NPs

(1)

199

For treated leaves in direct contact with the 109AgNP tracer, CAgNPs was calculated

200

according to Eq. (2), following Hintelmann et al.,38 where Rsp and Rnatural are the

201

isotope ratios of

202

manufacturer's reported value) and in nature (indistinguishable from those in roots,

203

see the Results section), and A109Agsp represents the abundance of 109Ag in the tracer

204

solution [i.e., 0.9856, see Eq. (3)]. Once CAgNPs has been determined, CAg2S-NPs in

205

treated leaves can be readily quantified using Eq. (1).

107Ag

to

109Ag

in the tracer solution (0.015 based on the

C107 Ag  Rnatural  C109 Ag  ( Rsp  Rnatural )  A109 Ag sp

206

C AgNPs

207

C 109 Ag C 107 Ag 107 A Ag sp  107 , A Ag  107 C Ag  C 109 Ag C Ag  C109 Ag

208

(2)

109

(3)

For untreated leaves and the roots, CAg2S-NPs was determined using Eq. (4), where

209

Rnatural and Rexposed represent the isotope ratios of

210

leaves, and A107Ag represents the abundance of

211

was then determined using Eq. (1).

212

C Ag2 S  NPs 

C 109 Ag  Rexposed  C107 Ag ( Rroot  Rexposed )  A107 Ag

109Ag

107Ag

to

107Ag

in roots and treated

in the samples (Eq. 3). CAgNPs

 Ccontrol

(4)

213

For all tissues, the relative importance of the foliar uptake of AgNPs (ffoliar) could

214

be established based on the ratio of foliar uptake to the overall Ag concentration, as 10


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215 216


described by Eq. (5).

f foliar 

C AgNP s C Ag

100%

(5)

217

RESULTS

218

Nanoparticle characterization.

219

average TEM diameter of 36.8 ± 4.1 nm (Figure 1a). Both EDS profile and surface

220

plasmon resonance absorbance at ~410 nm indicated the presence of metallic AgNPs

221

(Figure 1b, c). Ag2S-NPs in Milli-Q water were elliptical in shape, with width

222

averaging 49.9 ± 12.3 nm (Figure 1d). EDS and XRD analysis confirmed the

223

formation of Ag2S-NPs (Figure 1e).5 The intensity-weighted hydrodynamic diameters

224

of

225

number-weighted and intensity-weighted hydrodynamic diameters of Ag2S-NPs were

226

relatively constant after 2 days in the growth medium (Figure S2). Their zeta

227

potentials were −34.2 and −38.6 mV for 109AgNPs and Ag2S-NPs. Dissolved Ag ions

228

released from Ag2S-NPs were undetectable for 3 days (< 5 ng L-1), after which the

229

medium was renewed.

230

Sub-lethal toxicity to plants. Exposure to Ag-containing NPs tended to decrease

231

both the fresh biomass of soybeans and the fluorescence parameters of the leaves

232

relative to the control groups (Figure 2). Notably, plants in the High group suffered

233

greater toxicity than their counterparts in the other groups, as the fresh biomass of

234

roots decreased by as much as 42% and that of shoots by as much as 43% compared

235

to the control (Figure 2a). In addition, as in our earlier investigation,12 brownish

236

necrosis was observed in the treated leaves (Figure S3). The maximal photochemical

109AgNPs

109AgNPs

in Milli-Q water were spherical with an

and Ag2S-NPs were 56.1 ± 3.7 and 190.5 ± 2.8 nm, respectively. Both

11



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237

activity of PSII (Fv/Fm), the performance index based on the absorption of light

238

energy (PIABS) and the absorption flux per reaction center (ABS/RC) decreased by

239

38.8%, 96.4% and 59.6%, respectively, in treated leaves relative to those in control.

240

These results suggested that soybean leaves are susceptible to AgNPs via foliar

241

exposure.

242

Uptake and translocation of Ag-containing NPs in Plants. Average total Ag

243

concentrations in the control were 0.25 ± 0.071 and 0.67 ± 0.26 µg g−1 in the leaves

244

and roots, respectively. Upon exposure to Ag-containing NPs, total Ag concentrations

245

increased to 7.92−22.84, 0.38−1.45, and 1005−5623 µg g−1 in treated leaves,

246

untreated leaves, and roots, respectively, for the exposed groups. In this study, silver

247

accumulation in plant tissues may have resulted from three geochemical sources:

248

background, root uptake of Ag2S-NPs, and foliar uptake of

249

accumulation in the exposed groups from each geochemical source, determined after

250

subtracting the background Ag level derived from the control (no Ag-containing NPs

251

were added), is presented in Figure 3.

109AgNPs.

The net Ag

252

Nearly all of the Ag in the roots originated from Ag2S-NP adsorption and

253

internalization; the translocation and accumulation of 109AgNPs via foliar uptake were

254

negligible in the exposure scenario of this study (Figure 3a). This observation was

255

further supported by the isotope ratios of

256

exposed groups, which were indistinguishable from the ratio in the control (0.999 ±

257

0.0008 vs. 0.997 ± 0.0007). An increase in the exposure dose of Ag2S-NPs led to

258

greater Ag accumulation in the roots, with 2- to 3-fold greater Ag concentrations in

107Ag/109Ag

of total Ag in the roots of the

12


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259 260


the High group than in the Low and Medium groups. In treated leaves,

109AgNP

amendment greatly changed the isotopic composition

107Ag/109Ag

261

of total Ag, decreasing the

262

0.0821 ± 0.0207, 0.121 ± 0.0363 and 0.108 ± 0.0140 in the Low, Medium and High

263

groups, respectively, indicating the presence of

264

foliar uptake. Net Ag accumulation via foliar uptake of 109AgNPs increased from 7.51

265

± 0.72 to 12.75 ± 5.44 µg g−1, but the increase was not statistically significant (Figure

266

3b). Root uptake of Ag2S-NPs and the upwards translocation in treated leaves resulted

267

in a dose-dependent net accumulation of 0.54 ± 0.24 to 1.59 ± 0.16 µg g−1 (Figure 3b).

268

The contribution of Ag2S-NPs via root uptake to the accumulated Ag was limited,

269

with this pathway accounting for at most 29% of the Ag in the treated leaves. This

270

result pointed to a dominant role of the foliar uptake of AgNPs in treated leaves.

271

ratio from 0.998 ± 0.012 in the control to

109Ag

derived from

109AgNPs

via

Interestingly, in untreated leaves, which did not come into direct contact with

272

109AgNPs

273

isotopic ratio of total Ag (0.944 ± 0.0244) was between that obtained for treated

274

leaves (averagely 0.0821 to 0.121) and that of the control (0.998 ± 0.012). While this

275

result can be explained by a significant effect (p < 0.01) of foliar uptake and the

276

subsequent mobility of

277

in the untreated leaves originated from root uptake of Ag2S-NPs, which suggested a

278

high mobility of these NPs for translocation. Despite the significant mobility of

279

109AgNPs,

280

foliar exposure levels.

or Ag2S-NPs, the uptake pattern differed from that of treated leaves. The

109AgNPs,

further calculation showed that the majority of Ag

their net accumulation was lower compared to Ag2S-NPs, due to the lower

13



281

Characterization of Ag-containing NPs in plants. To detect Ag-containing NPs,

282

plant tissues of the High group were examined by TEM. NPs were detected in the

283

roots at the intracellular level (Figure 4 a, b); unfortunately, the Ag signal was close to

284

the detection limit. It was not possible to conclusively identify Ag-containing NPs in

285

leaves due to the low sensitivity of TEM-EDS. Instead, plant tissues were digested

286

with macerozyme R-10 and subjected to spICP-MS analysis.12 Ag-containing NPs

287

averaging 43.8 ± 2.0, 29.6 ± 1.0 and 33.9 ± 0.1 nm in size were detected in roots,

288

treated leaves and untreated leaves, respectively (Figure 4).

289

Trophic transfer to snails. To further investigate the effects of geochemical sources

290

of Ag-containing NPs on Ag trophic transfer, snails were fed control and Ag-loaded

291

leaves for 5 days. Untreated leaves of the High group were chosen as the Ag-loaded

292

diet because Ag was internalized rather than adsorbed on the leaf surface. There was

293

no mortality of snails over the exposure period; the weight of the snails exposed to

294

untreated leaves did not differ from that of control snails (p > 0.05). Exposed snails

295

contained 0.93 ± 0.19 µg Ag g−1, compared to 0.11 ± 0.03 µg g−1 in control snails.

296

Total Ag in feces was 2.0 ± 0.46 and 0.38 ± 0.11 µg g−1 for exposed and control

297

snails, respectively. Mass balance showed that 91–133% of the Ag was recovered. On

298

average, 78 ± 17 % of the dietary Ag was assimilated by snails during trophic transfer

299

(calculated as the ratio of the amount of Ag in snail soft tissues to that in ingested

300

leaves).

301

The Ag detected in the snails originated to a large extent from Ag2S-NPs, given

302

that Ag in untreated leaves was almost exclusively derived from the root uptake of 14


Page 14 of 39

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303

Ag2S-NPs (Figure 3), whereas the foliar uptake of 109AgNPs had a minor effect. This

304

result was further supported by the indistinguishable isotopic ratios of total Ag during

305

trophic transfer, i.e., 0.95 ± 0.026 and 0.95 ± 0.011 in untreated leaves and snail

306

tissues, respectively (Figure 5). We thus concluded that the Ag that accumulated in

307

untreated leaves via root uptake of Ag2S-NPs is able to enter the food chain.

308

DISCUSSION

309

Our experiments using an enriched stable isotope tracer addressed the questions

310

whether and how different sources of Ag-containing NPs (foliar uptake of AgNPs vs.

311

root uptake of Ag2S-NPs) affect the phytoavailability and trophic transfer of the

312

particles. Concurrent exposure stimulates the worse-case environmental scenario in

313

the field. We and others have demonstrated that Ag, whether in the form of Ag ions or

314

engineered AgNPs, is transformed into Ag2S-NPs in soils and biosolids.1, 3-5, 39 Sludge

315

containing Ag2S is added as fertilizer to agricultural soil in conjunction with AgNPs

316

as fungicides and bactericides for plant pathogen protection.7 Alternatively, AgNPs

317

via foliar application could arrive at the soil and transform to Ag2S. In any case,

318

concurrent exposure provides more realism than single exposure pathway. Our data

319

indicated that root uptake of Ag2S-NPs is the more important source of NPs to plants

320

and thus to a terrestrial snail.

321

Effects on phytoavailability. Our data highlighted the importance of nanoscale Ag2S

322

as a potential source of the metal to terrestrial organisms. In leaf samples, the mobility

323

of Ag derived from Ag2S-NPs and 109AgNPs was demonstrated (Figure 3). However,

324

given the high exposure doses, Ag2S-NPs were the dominant species that accumulated 15



325

in plants. These particles remained suspended over the entire exposure, with

326

non-observable dissolution (Ksp = 5.92×10-51)39 or aggregation in the bulk suspension

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(Figure S2). However, partial dissolution of Ag2S-NPs in the rhizosphere followed by

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Ag-ion uptake has been proposed in plants.38, 40 spICP-MS results demonstrated the

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presence of Ag-containing NPs averaging 29.6–43.8 nm in size in planta (Figure

330

4c-e), which were similar to the originally dosed AgNPs and Ag2S-NPs (34.9 ± 2.5 vs.

331

44.9 ± 1.2 nm, as measured by spICP-MS). However, the chemical speciation of NPs

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in planta was not analyzed, although Ag2S, metallic Ag and Ag-thiols have been

333

identified on the surface of plant roots treated with Ag2S-NPs using X-ray based

334

techniques.31,

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21%) of Ag2S-NPs was sorbed to the plant tissue in our system, but it was much

336

higher than the < 0.21% reported for natural soils amended with Ag-containing

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biosolids.2,

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Ag2S-NPs herein. The aqueous measurements are different from those in planted

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soil,43, 44 due to the presence of soil solid and the associated ligands as well as the

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aging of Ag2S in natural soil reducing the availability of Ag2S to plants.45 The high

341

doses of Ag2S-NPs in this study may be also responsible.34, 40, 42 Likewise, nanoscale

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Ag2S derived from wastewater treatment processes can still be bioavailable to soil

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microflora and isopods, albeit at lower levels than pristine forms.46-48

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42

32, 41

Mass-balance calculations demonstrated that a small fraction (

Discerning the sources of silver nanoparticle in a terrestrial food chain

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