impact of physico-chemical properties of …

University of Kentucky University of Kentucky

UKnowledge UKnowledge

Theses and Dissertations--Toxicology and Cancer Biology Toxicology and Cancer Biology

2018

IMPACT OF PHYSICO-CHEMICAL PROPERTIES OF IMPACT OF PHYSICO-CHEMICAL PROPERTIES OF

MANUFACTURED NANOMATERIALS ON PLANT UPTAKE AND MANUFACTURED NANOMATERIALS ON PLANT UPTAKE AND

TROPHIC TRANSFER TROPHIC TRANSFER

Jieran Li University of Kentucky, [email protected] Author ORCID Identifier:

https://orcid.org/0000-0002-1785-6652 Digital Object Identifier: https://doi.org/10.13023/etd.2018.474

Right click to open a feedback form in a new tab to let us know how this document benefits you. Right click to open a feedback form in a new tab to let us know how this document benefits you.

Recommended Citation Recommended Citation Li, Jieran, "IMPACT OF PHYSICO-CHEMICAL PROPERTIES OF MANUFACTURED NANOMATERIALS ON PLANT UPTAKE AND TROPHIC TRANSFER" (2018). Theses and Dissertations--Toxicology and Cancer Biology. 24. https://uknowledge.uky.edu/toxicology_etds/24

This Doctoral Dissertation is brought to you for free and open access by the Toxicology and Cancer Biology at UKnowledge. It has been accepted for inclusion in Theses and Dissertations--Toxicology and Cancer Biology by an authorized administrator of UKnowledge. For more information, please contact [email protected].

http://uknowledge.uky.edu/

http://uknowledge.uky.edu/

https://uknowledge.uky.edu/

https://uknowledge.uky.edu/toxicology_etds

https://uknowledge.uky.edu/toxicology_etds

https://uknowledge.uky.edu/toxicology

https://orcid.org/0000-0002-1785-6652

https://uky.az1.qualtrics.com/jfe/form/SV_9mq8fx2GnONRfz7

mailto:[email protected]

STUDENT AGREEMENT: STUDENT AGREEMENT:

I represent that my thesis or dissertation and abstract are my original work. Proper attribution

has been given to all outside sources. I understand that I am solely responsible for obtaining

any needed copyright permissions. I have obtained needed written permission statement(s)

from the owner(s) of each third-party copyrighted matter to be included in my work, allowing

electronic distribution (if such use is not permitted by the fair use doctrine) which will be

submitted to UKnowledge as Additional File.

I hereby grant to The University of Kentucky and its agents the irrevocable, non-exclusive, and

royalty-free license to archive and make accessible my work in whole or in part in all forms of

media, now or hereafter known. I agree that the document mentioned above may be made

available immediately for worldwide access unless an embargo applies.

I retain all other ownership rights to the copyright of my work. I also retain the right to use in

future works (such as articles or books) all or part of my work. I understand that I am free to

register the copyright to my work.

REVIEW, APPROVAL AND ACCEPTANCE REVIEW, APPROVAL AND ACCEPTANCE

The document mentioned above has been reviewed and accepted by the student’s advisor, on

behalf of the advisory committee, and by the Director of Graduate Studies (DGS), on behalf of

the program; we verify that this is the final, approved version of the student’s thesis including all

changes required by the advisory committee. The undersigned agree to abide by the statements

above.

Jieran Li, Student

Dr. Jason M. Unrine, Major Professor

Dr. Isabel Mellon, Director of Graduate Studies

IMPACT OF PHYSICO-CHEMICAL PROPERTIES OF MANUFACTURED

NANOMATERIALS ON PLANT UPTAKE AND TROPHIC TRANSFER

________________________________________

DISSERTATION

________________________________________

A dissertation submitted in partial fulfillment of the

requirements for the degree of Doctor of Philosophy in the

College of Medicine

at the University of Kentucky

By

Jieran Li

Lexington, Kentucky

Director: Dr. Jason M. Unrine, Associate Professor

Lexington, Kentucky

2018

Copyright © Jieran Li 2018

https://orcid.org/0000-0002-1785-6652

https://orcid.org/0000-0002-1785-6652

ABSTRACT



Large quantities of manufactured nanomaterials (MNM) are released into the

environment by human activity each year. The entry of MNM into the terrestrial food webs,

which has the potential for far-reaching impacts, begins with the uptake by plant species

from the soil. These processes can be affected by MNM physico-chemical properties such

as size, chemical composition, surface charge, etc., of which our knowledge is still

incomplete. To bridge some of the gaps in our understanding of these processes and,

specifically, to determine whether the physico-chemical properties of the MNM are

predictive of their behavior in terrestrial food chains, we conducted a series of experiments

using different MNM and model organisms.

First, we synthesized functionalized CeO2 MNM having different charges and

exposed tomato plants (Solanum lycopersicum cv Micro-Tom) to them. We found that

plant growth and the rate of root-to-shoot translocation were functions of surface charge

and exposure concentration. Mechanisms of entry into roots were examined using recent

advances in high-resolution synchrotron X-ray microscopy to show that a combination of

apoplastic and symplastic routes was used by the particles to penetrate to the interior of the

roots. Our results also illustrate that these particles have drastically different tissue

distribution patterns depending on their surface charges.

Second, we exposed tomato plants with these CeO2 MNM and fed the leaves to the

tobacco hornworm (Manduca sexta). Differential trophic transfer was observed as a

function of the surface charge of the particles. An uptake and elimination study was

conducted to obtain a time course of Ce dynamics. Despite no observed overall

biomagnification across trophic levels, these differentially charged CeO2 MNMs had

greater bioaccumulation factors than that of ionic Ce3+. The uptake-elimination dynamics

were influenced by the surface charge of the NPs. Positively charged NPs had higher

bioaccumulation factors and assimilation efficiencies but lower elimination rate than

neutral and negatively charged CeO2 MNMs.

Finally, to determine if studies conducted with highly purified, lab synthesized

materials, were predictive of behavior of commercial nanopesticide formulations, we

studied the dietary uptake of Cu(OH)2 MNMs by hornworms feeding on surface-

contaminated tomato leaves. We compared lab-synthesized copper hydroxide (Cu(OH)2)

nanowire with the widely used fungicide KOCIDE® 3000, whose active ingredient is nano-

needles of copper hydroxide (Cu(OH)2). The difference in their toxicity and

accumulation/elimination dynamics was found to correlate with the solubility of the

materials.

We have shown that the physico-chemical properties of MNM affect the toxicity,

bio-distribution and trophic transfer of MNM in terrestrial ecosystems. With the increase

of MNM release into the environment as a result of the rapid development of

nanotechnology, these results have important implications for the evaluation of

environmental risks associated with these MNMs and may help the application of

nanotechnology to evolve to be more environmentally friendly.

KEYWORDS: Nanotechnology, Plant, Environment, Agriculture, Toxicology, Insect

Jieran Li

(Name of Student)

12/13/2018

Date



By

Jieran Li

Jason M. Unrine

Director of Dissertation

Isabel Mellon

Director of Graduate Studies

12/13/2018

Date

iii

ACKNOWLEDGMENTS

This dissertation was supported by the National Science Foundation under Grants

1530594 and 1266252. This research also used the Hard X-ray Nanoprobe (HXN) Beamline

at 3-ID and the Submicron Resolution X-ray Spectroscopy (SRX) Beamline at 5-ID

National Synchrotron Light Source II, a U.S. Department of Energy (DOE) Office of

Science User Facility operated for the DOE Office of Science by Brookhaven National

Laboratory under Contract No. DE-SC0012704. Portions of this work were performed at

GSECARS (The University of Chicago, Sector 13), Advanced Photon Source (APS),

Argonne National Laboratory. GSECARS is supported by the National Science Foundation

– Earth Sciences (EAR-1128799) and Department of Energy – Geosciences (DE-FG02-

94ER14466). APS facility is supported by DOE under Contract No. DE-AC02-

06CH11357. The author acknowledges Y-C. Chen-Wiegert, L. Li, J. Thieme, W. Rao, J.

Begley, S. Sutton, M. Newville, and A. Lanzirotti. The author also wishes to thank his

dissertation committee (Olga Tsyusko, S. Reddy Palli, David McNear, Isabel Mellon) and

outside examiner (Jason DeRouchey) for their valuable guidance.

iv

TABLE OF CONTENTS

ACKNOWLEDGMENTS .................................................................................................................. iii

TABLE OF CONTENTS .................................................................................................................... iv

LIST OF TABLES ............................................................................................................................ vii

LIST OF FIGURES ......................................................................................................................... viii

CHAPTER 1. Introduction ......................................................................................................... 1

1.1 Nanotechnology and nanotoxicology ................................................................................. 1

1.2 Abiotic transformation of MNMs in the environment ........................................................ 2

1.3 Bioaccumulation of MNMs in plants ................................................................................... 4

1.3.1 Uptake and translocation ........................................................................................... 4

1.3.2 Phytotoxicity of MNMs ............................................................................................. 6

1.4 Dietary uptake and trophic transfer of MNMs ................................................................... 8

1.4.1 Trophic transfer of MNMs in aquatic ecosystem ...................................................... 9

1.4.2 Trophic transfer of MNMs in terrestrial ecosystems ............................................... 10

1.5 Dissertation outline ........................................................................................................... 11

CHAPTER 2. Effect of CeO2 nanomaterial surface functional groups on tissue and subcellular

distribution of Ce in tomato (Solanum lycopersicum) † ............................................................................ 13

2.1 Synopsis ............................................................................................................................ 14

2.2 Introduction ...................................................................................................................... 15

2.3 Methods ............................................................................................................................ 18

v

2.4 Results and discussion ...................................................................................................... 22

2.5 Conclusion ......................................................................................................................... 31

2.6 Tables and figures ............................................................................................................. 33

2.7 Supplemental information ................................................................................................ 42

CHAPTER 3. Trophic transfer of differentially charged CeO2 nanoparticles from tomato

(Solanum lycopersicum) plants to tobacco hornworm (Manduca sexta) larvae ..................................... 52

3.1 Synopsis ............................................................................................................................ 52

3.2 Introduction ...................................................................................................................... 53

3.3 Methods ............................................................................................................................ 55


3.5 Conclusion ......................................................................................................................... 63


CHAPTER 4. Comparing trophic transfer of lab-synthesized nano-Cu(OH)2 to a commercial

nano-Cu(OH)2 fungicide formulation from tomato (Solanum lycopersicum) leaves to tobacco hornworm

(Manduca sexta) larvae .......................................................................................................................... 70

4.1 Synopsis ............................................................................................................................ 70

4.2 Introduction ...................................................................................................................... 71

4.3 Methods ............................................................................................................................ 73


4.5 Conclusion ......................................................................................................................... 81


vi

CHAPTER 5. Conclusions and future directions ...................................................................... 92

REFERENCES ................................................................................................................................ 96

VITA .......................................................................................................................................... 111

vii

LIST OF TABLES

Table 2.1 Characterization of CeO2 MNMs ..................................................................... 33

Table 4.1 Dissolution of the nanomaterials ....................................................................... 82

viii

LIST OF FIGURES

Figure 2.1 Biomass reduction ............................................................................................ 34

Figure 2.2 Damage to the root tips .................................................................................... 35

Figure 2.3 Bulk Ce concentration in tissue ........................................................................ 36

Figure 2.4 Ce distribution at root tips ................................................................................ 37

Figure 2.5 Evidence of cytoplasmic distribution ............................................................... 39

Figure 2.6 Ce speciation at the root tip .............................................................................. 40

Figure 2.7 Ce distribution and speciation in leaf tissue ..................................................... 41

Figure 2.8 Chlorosis .......................................................................................................... 45

Figure 2.9 Ionic control ..................................................................................................... 46

Figure 2.10 Ce concentration in stem vs. leaf tissue ......................................................... 47

Figure 2.11 Additional XRF coarse graphs ....................................................................... 48

Figure 2.12 Example of linear combination fitting of the XANES spectra ....................... 51

Figure 3.1 Growth of insect larvae .................................................................................... 65

Figure 3.2 Accumulation of Ce ......................................................................................... 66

Figure 3.3 Uptake and elimination dynamics .................................................................... 67

Figure 3.4 Ce distribution in the midgut tissue.................................................................. 68

Figure 3.5 Ce speciation in the midgut tissue .................................................................... 69

Figure 4.1 Growth retardation and mortality ..................................................................... 83

Figure 4.2 Bulk Cu in tissue .............................................................................................. 84

Figure 4.3 Trophic transfer and assimilation efficiency .................................................... 85

Figure 4.4 Correlation between trophic transfer factors and free Cu2+ .............................. 86

Figure 4.5 Biodynamic model ........................................................................................... 87

Figure 4.6 TEM graphs of nCu(OH)2 and Kocide ............................................................. 88

Figure 4.7 Ingestion rate .................................................................................................... 89

Figure 4.8 Bulk Cu in the food .......................................................................................... 90

Figure 4.9 Growth curve fitting ......................................................................................... 91

1

CHAPTER 1. INTRODUCTION

1.1 Nanotechnology and nanotoxicology

Manufactured nanomaterials (MNMs) are used to enhance the properties of many

manufactured products. Manufactured nanomaterials are defined as having at least one

dimension under 100 nm. In this size range, the number of atoms exposed on the surface

increases exponentially to give the materials new properties that differentiate them

drastically from the bulk materials 1. Manufactured nanomaterials have diverse chemical

compositions and applications, with a large proportion of them being metal and metal

oxides. The most well studied metal and metal oxides, in terms of their fate and effects in

ecosystems are Au, Ag, TiO2, ZnO, CeO2, and Cu/CuO/Cu(OH)2. Gold MNMs are used

in biomedical and clinical research as drug delivery systems, bio-imaging agents,

chemotherapeutics, biosensors, and as catalysts for fuel cells 2-11. Silver MNMs are widely

used as chemical catalysts, chemotherapeutics, antimicrobial drugs, and additives in

various consumer products 12-20. Zinc oxide MNMs are used as sensors, components of

electronic devices, antimicrobial and as ultraviolet absorber in sunscreens 21-25. Titanium

dioxide MNMs are used as additives in a wide range of consumer products such as

sunscreens, plastics and pains 26-28. Cu-based MNMs such as nano-Cu and CuO are used

as sensors in electronics and biochemical analysis, as chemical catalysts, and as pesticide

in agricultural practices 29-36. CeO2 MNMs are used as pharmaceuticals, polishing agents,

and diesel fuel additives 37-40. Carbon MNMs such as fullerene and carbon nanotubes are

widely used in biomedical research and engineering as mechanical parts, sensors, micro-

scaffolds, energy conversion and storage in fuel cells, supporting scaffolds, and electric

conducting materials 41-45.

2

With rapid increase of MNMs being produced and released into the environment,

their impact on ecosystems is only beginning to be investigated 46, 47. Even for MNMs

sharing the same chemical composition, their behaviors in the environment are difficult to

predict due to the diverse physico-chemical properties such as size, shape and surface

chemistry, which are often susceptible to change under different environmental conditions.

Physical and chemical transformations constantly occur after the MNMs enter the

environment and are influenced by extrinsic factors such as pH, redox potential, ionic

strength, and the presence of ligands. Furthermore, the bioavailability and toxicity of these

MNMs will vary due to the great variety of organisms that have different routes of being

exposed to the MNMs in the environment. Ecotoxicology of MNMs is beginning to attract

enormous attention especially as MNMs often exhibit special properties and bio-reactivity

in contrast to the corresponding bulk materials and ions.

1.2 Abiotic transformation of MNMs in the environment

MNM undergo a variety of transformation processes in the environment 48. As

colloidal suspensions dispersed in aqueous phase of a wide range of environmental media,

MNMs tend to form aggregates 49, 50, which can be either homoaggregates, formed between

particles of the same type, or heteroaggregates, formed between different types of MNMs

or naturally occurring nanoparticles such as soil colloids. Aggregation might also occur as

the particles are adsorbed on the surface of microorganisms 51. Aggregation of MNMs can

dramatically alter their surface properties and reactivity such as crystal phases and the

number of atoms per surface area, which will have great influence on how they further

interact with the environment and the organisms exposed to them 52. Dissolution is another

3

important process that can alter the properties of the MNMs in the environment. Ag, Cu

and ZnO MNMs are soluble under most environmentally relevant conditions 53-56. The ions

released from the particles, Ag+, Zn2+ and Cu2+, can be precipitated by inorganic ligands

such as S2-and PO43-. Sometimes the highly insoluble compounds form as a shell that

shields the surface of the MNMs from the external environment preventing further

reactions 56-58. Other MNMs such as Au, CeO2 and TiO2 tend to resist dissolution in the

environment although some studies suggest dissolution of Ce and Au is possible 59-63.

Chemical and physical transformations of MNMs in the environment are highly

complex processes whose kinetics depend on both the intrinsic properties of the materials

and extrinsic conditions such as pH, ionic strength, redox potential, and natural organic

matter (NOM). Dissolution of coated Ag MNMs was shown to be influenced by the size

of the particles 54. Dissolution of Cu based MNMs is accompanied by redox reactions and

interaction with organic acids and is greatly influenced by pH 58. The rate of the dissolution

and sulfidation of Ag MNMs in biosolids is a function of pH, dissolved oxygen

concentration, and NOM 64. Association of CeO2 MNMs with biomass in wastewater

treatment plants is accompanied by the reduction of Ce(IV) particles to Ce(III) species65.

Often the properties of MNMs are altered by these transformation processes that the ways

they interact with the environment are also changed, causing further transformation to

occur. Silver MNMs that developed a Ag2S shell on the surface became unstable and were

prone to aggregation 57. In the presence of organic acids, Zn2+ released from the ZnO

MNMs were precipitated by the phosphate, causing the MNMs to be gradually transformed

into various mineral phases of zinc phosphate with diverse morphology and crystal

structures 66. Sunlight, the presence of divalent cations such as Ca2+ and Mg2+ and NOM

4

were key parameters for the morphology change and aggregation of AgNPs in surface

water 67. The complexity of the transformation processes means that environmental

conditions and the physico-chemical properties of the MNMs themselves must be

specifically defined in order to understand and predict their behaviors.

1.3 Bioaccumulation of MNMs in plants

1.3.1 Uptake and translocation

There are two primary pathways by which materials are taken up and transported by

the plant tissue: apoplastic transport through cell walls and the intercellular space vs.

symplastic transport or transcellular transport where the materials enter the interior of the

cells. The porous network of the plant cell wall consists of polysaccharide fiber matrices

with various pore sizes ranging from 4 to over 10 nm depending on the species which

allows the movement of water and solutes 68. MNMs of the appropriate sizes can thus move

through this network with minimum resistance 69, 70. In the case of foliar exposure, it was

observed that MNMs entered through the pores of stomata and were translocated to

different parts of the leaf by the phloem 71. Physical barriers, such as the Casparian strip in

the root endodermis, exist in the plant tissue to regulate the movement of water and solutes

through the apoplast 72, 73. Symplastic routes can be taken by the MNMs to bypass these

barriers. A confocal microscopy study by Zhao et al. demonstrated that ZnO MNMs

penetrated the epidermis and cortex of the root of corn plants through the apoplastic

pathway, but aggregates of particles were also found in the xylem, suggesting that these

MNMs or aggregates pass through the endodermis via the symplastic pathway 74. One

mechanism by which symplastic or transcellular movement of MNMs may occur is via

5

endocytosis, which can be either dependent or independent of clathrin and endosome 75.

Internalization of Au MNMs by both clathrin dependent and independent pathways was

shown in tobacco protoplasts 76. Similarly, quantum dots were found to be taken up by

plant cells via receptor-mediated clathrin-dependent as well as non-specific clathrin-

independent endocytosis 77. Furthermore, it has been shown that multi-walled carbon

nanotube are taken up by plant cells through endocytic pathways with or without

endosomal processes 78, 79. Transcellular movement of MNMs can also be achieved via the

plasmodesmata which is likely a size-limit transport pathway. Ag and Au MNMs < 40 nm

were found to accumulate at plasmodesmata and the longitudinal channels in the cell walls

and both symplastic and apopastic pathways were suggested as transit routes 80, 81.

The presence and translocation of MNMs to distant parts of the plants from the site

of uptake via the vascular system was well documented. Transport of MNMs through the

xylem with the water flux is very efficient and is only limited by the pore size of the xylem

structural components, such as the filter plates 81-85. In a study with pumpkin plants, long

range transport of graphite-coated Fe MNMs by xylem was shown to be size-dependent,

where Fe MNMs found at the distant site had similar a size of 40 nm 84. Phloem transport

of MNMs has also been noted, which is allows for the downward transport from the leaf

86-89. Sometimes MNMs rely on both xylem and phloem for multidirectional transport in

the plant. It was demonstrated that in corn plants, the root-to-shoot translocation of CuO

MNMs was mediated by xylem, and the phloem was responsible for transfer in the reverse

direction from shoot to root 85. The transport of MNMs by phloem is essential for the

accumulation of MNMs in fruits and transgenerational effects passed down to the offspring

90, 91.

6

1.3.2 Phytotoxicity of MNMs

Although beneficial effects of MNMs have been reported by some studies in plants,

the majority of the research shows various toxic effects. Macroscopic changes on the

organism level resulting from the exposure to MNMs include reduced biomass

accumulation, lower rate of growth and seed germination, etc., with symptoms varying

from study to study depending on plant species and the types of MNMs 92-96. Exposure to

ZnO MNMs but not Al2O3 and SiO2 MNMs reduced seed germination rate and general

growth of Arabdiopsis thaliana 97. Cu and Ag MNMs had no effects on the seed

germination but significantly reduced the biomass and root elongation in Cucurbita pepo

98. CeO2 MNMs were shown to reduce the growth and biomass accumulation of Glycine

max 99. Other rare earth metal oxide MNMs such as La2O3, Gd2O3, and Yb2O3 were

observed to induce root growth inhibition in Brassica napus, Raphanus sativus, Lactuca

sativa, Lycopersicum esculentum and Brassica oleracea 100. Tomato plants exposed to TiO2

and ZnO MNMs through both the root and foliar routes exhibited reduced root growth and

biomass accumulation, with the root exposure being more efficacious 90. TiO2 MNMs were

shown to reduce root biomass and inhibit the development of lateral roots 101. Nair et al.

showed that exposure to Ag MNMs decreased root and shoot biomass of Oryza sativa 102.

Similarly, AgNPs caused biomass reduction in C. pepo 103. Furthermore, O. sativa exposed

to carbon MNMs had retarded growth and flowering was significantly delayed 91.

These macroscopic changes are usually underscored by pronounced damage at the

cellular level as well as alterations in metabolism, biochemistry and gene expression.

Exposure to Ag and ZnO MNMs was found to decrease cell size and turgidity as a result

of shrinkage of the vacuole in B. oleracea and Zea mays 104. Cucumber plants exposed to

7

ZnO MNMs were also found to have similar shrinkage of the vacuoles and collapsed cortex

and epidermis 105. Likewise, when exposed to ZnO MNMs, collapse of the root tip tissue

was observed in several crop plants due to extensive loss of cells in the epidermis and

cortex 106. This study also demonstrated clogging of the apoplast and plasmodesmata by

the MNMs, resulting in obstructed intercellular trafficking. Excessive accumulation of

MNMs in the apoplast can cause reduction in water flux and nutrient uptake. Exposure to

TiO2 MNMs blocked transpiration and decreased hydraulic conductivities in Z. mays 83.

Similarly, exposure to Ag MNMs reduced transpiration in C. pepo 107. In addition, ZnO

MNMs were found to cause nutrient deficiency and translocation hindrance in cowpea 108.

Nair et al. demonstrated that exposure of O. sativa to Ag MNMs affected leaf area

and reduced contents of chlorophyll and carotenoids which are important for

photosynthesis 102. Similarly, Mukherjee et al. (2014) showed that green peas (Pisum

sativum) exposed to ZnO MNMs had decreased chlorophyll content 109. Also, CeO2 MNMs

caused a decline in chlorophyll production in Arbidopsis 110. Chlorophyll and carotenoids

contents in rice seedlings were found to be significantly decreased as a result of exposure

to Ag MNMs 102. A similar reduction of chlorophyll content was also observed in cucumber

plants and Phaseolus vulgaris treated with TiO2 MNMs and in P. sativum treated with ZnO

MNMs 109, 111, 112.

When interacting with cells and tissues, some MNMs produce reactive oxygen

species (ROS) and can cause oxidative stress, damage to proteins and DNA, lipid

peroxidation, and electrolyte leakage 113. For example, carbon- and metal-based MNMs

were shown to induce oxidative stress via ROS generations in various plant species,

causing cell cycle arrest, apoptosis and abnormal chromatin structures 114-118. CuO MNMs

8

caused DNA damage in radish and perennial ryegrass 119. Similarly, A. thaliana exposed

to PVP-coated Ag MNMs showed elevated ROS levels across the plant 95. Higher activity

of vital antioxidants such as superoxide dismutase (SOD), catalase and glutathione (GSH)

as well as elevated lipid peroxidation were noted in tomato plants treated with NiO MNMs

120. Similar to this, Jacob et al. (2013) showed that activities of key enzymatic antioxidants

in P. vulgaris were modified by TiO2 MNMs 112. Moreover, A. thaliana exposed to

CdSe/ZnS quantum dots showed reduction in GSH levels 121. MNMs have been shown to

differentially modify the antioxidant system in a context-dependent fashion, where the

types and concentrations of MNMs, exposure media and plant species exposed are the

major parameters in defining the antioxidant response. For example, activities of

glutathione reductase (GR) and ascorbate peroxidase (APX) were found to be reduced in

Vicia faba by TiO2 MNMs 122. Howerver, TiO2 MNMs exposed A. thalia had elevated

activities of SOD, catalase, APX, and guaiacol peroxidase (GPX) 97. Enhanced activities

of SOD, catalase and GPX were also noted in Lemma minor exposed to TiO2 MNMs 123.

Further research is needed to elucidate the foundations of antioxidant response to MNMs

in plants.

1.4 Dietary uptake and trophic transfer of MNMs

A critical part of the ecotoxicity of environmental contaminants is their potential of

transfer between different trophic levels in food webs. This creates possible routes for

species at higher trophic levels to be exposed to these contaminants, sometimes

significantly with the occurrence of biomagnification, i.e., higher concentration of

contaminants in consumer than in the food source.

9

1.4.1 Trophic transfer of MNMs in aquatic ecosystem

With their varied composition and manifold properties, it is not surprising to find

that MNMs exhibit a similar complexity of behavior in terms of their transfer in the food

web comparable to the traditional contaminants discussed above. Only a limited number

of studies have examined trophic transfer of MNMs in the aquatic environment. Holbrook

et al. demonstrated that functionalized quantum dots can be transferred to rotifers through

the dietary uptake of protozoans 140. Bioconcentration of quantum dots by the microbe was

at a low level without any evidence of biomagnification in the upper trophic level.

Similarly, Zhu et al. found that TiO2 MNMs can transfer from daphnids to zebrafish (Danio

rerio) via dietary exposure but with extremely low trophic transfer factor (0.024) 141. In

contrast, bioconcentration by the zebrafish from the aqueous media was very high.

However, the authors showed that dietary exposure resulted in significantly higher body

burden of Ti in the zebrafish than aqueous exposure because the Ti concentration in

Daphnia was much higher than in the media, suggesting that dietary transfer could still be

an important route of exposure for these particles at higher trophic levels. In another study

the transfer of ZnO MNMs from D. magna to D. rerio was much more efficient than from

direct uptake in the aqueous phase and bioaccumulation of ZnO MNMs was significantly

higher than ionic Zn 142. Moreover, CdSe quantum dots were shown to biomagnify from a

simple microbe food chain from the bacteria Pseudomonas aeruginosa to protozoan

Tetrahymena thermophile 143. Intact quantum dots were observed in the vacuole of the

protozoan and could be available for uptake by higher trophic levels. Differences in trophic

transfer with regard to the food source was shown for Au MNMs 144. The authors found

that bioaccumulation of Au in D. magna was higher from dietary uptake of exposed

10

Euglena. gracilis than from Chlamydomonas. reinhardtii. It was argued that this difference

could be attributed to the cell wall of C. reinhardtii, which, as a physical barrier, might

limited the internalization of the particles by the phytoplankton. Ferry et al. investigated

the transfer of Au nanorods from water column to a complex marine food web containing

species at multiple trophic levels 145. In these simulated estuarine mesocosms,

bioaccumulation was found to be greater in clams and biofilms than in sea grass, shrimp,

snails and fish, with the greatest fraction of Au present in the sediment. Similarly, CeO2

MNMs were found to be associated with the sediments in a freshwater ecosystem 146.

However, Ce bioaccumulation declined with increasing trophic level with no observed

biomagnification. Furthermore, Kulacki et al. demonstrated that TiO2 MNMs altered the

composition of the biofilm formed by various types of algae species which in turn

influenced the bioaccumulation of Ti by the snail Physa acuta feeding on the biofilm 147.

1.4.2 Trophic transfer of MNMs in terrestrial ecosystems

As efforts were dedicated to the study of MNMs in the aqueous food webs, trophic

transfer of MNMs in terrestrial ecosystems have also become a focus of research in recent

years, although it is much less understood. The first evidence of trophic transfer of MNMs

in terrestrial ecosystem was provide by Judy et al. where transfer of Au MNMs from the

tomato and tobacco plants to herbivorous tobacco hornworms Manduca sexta was observed

148, 149. The authors demonstrated that dietary uptake of CeO2 MNMs by the hornworms

feeding on surface contaminated tomato leaves was much less efficient than from leaves

of plants pre-exposed to MNMs in hydroponic media. The studies provided the first

evidence of biomagnification of nanomaterials in a terrestrial food chain. The same group

also investigated the transfer of Au MNMs along a food chain from soil and earthworms

11

(Eisenia fetida) to bullfrogs (Rana catesbeina) 150. Although Ce concentrations decreased

by 100 fold for each transfer to a higher trophic level, the assimilation efficiency of Au in

the bullfrogs from ingested E. fetida was still much higher than from oral gavage of Au

MNMs in DI water. Hawthrone et al. studied the transfer of CeO2 MNMs and bulk CeO2

in a terrestrial food chain consisting of zucchini (C. pepo L.) plants, herbivores (crickets,

Acheta domesticus) and carnivores (wolf spiders, Lycosidae) 151. Crickets and wolf spiders

from the MNMs exposure group had significantly higher body concentrations and trophic

transfer factors of Ce than those from the bulk-exposed group. In a more recent study,

transfer of with CeO2 MNMs and bulk CeO2 from kidney bean plant (P. vulgaris) to

Mexican bean beetle (E. varivestis) and subsequently to spined soldier bug (Podisius

maculiventris) was examined 152. Biomagnification of Ce occurred for transfer at each

trophic level but the bioaccumulation factors were not different between the nano and bulk

exposure groups. Similar to this result, De la Torre Roche et al. investigated the trophic

transfer of La2O3 MNMs and bulk La2O3 from soil through a food chain consisting of

producers (lettuce), primary consumers (crickets) and secondary consumers (darkling

beetles, Tenebrionoidea) 153. In this study, no difference in tissue La concentrations was

found between the La2O3 MNMs and bulk La2O3, after 7 d of elimination and there was no

evidence for biomagnification.

1.5 Dissertation outline

In spite of these findings showing potential plant uptake, trophic transfer, and

biomagnification of MNMs in terrestrial ecosystems, our poor understanding of the

principles underlying the varied results warrants further research effort.

12

The objectives of this dissertation research were to investigate the influence of MNM

properties on plant uptake and trophic transfer. Chapters 2 and 3 focus on the role of

surface charge CeO2 nanomaterials in determining plant uptake and biodistribution in the

tomato (Lycopersicum solanum cv Microtom) and subsequent trophic transfer to the

tobacco hornworm (M. sexta). CeO2 nanomaterials are used primarily as fuel catalysts and

in manufacturing of semiconductors and optical components. They provide an interesting

model system because of their limited solubility, low background concentrations (allowing

them to be tracked using imaging techniques), and their redox activity. Chapter 4 focuses

on nanoagrochemicals, using a common fungicide containing Cu(OH)2 nanomaterials

(Kocide 3000). This study compared the commercial Kocide formulation with a similar,

lab synthesized Cu(OH)2 material. The vast majority of studies of the environmental

behavior and effects of MNMs utilized highly-purified, lab-synthesized materials. It is

unclear if studies of these materials would predict the behavior of commercial

formulations. Given the anticipated rapid expansion of the use of nanomaterials in

agriculture 154, this study is particularly timely and critical for developing adequate risk

assessment frameworks for this new class of pesticides.

CHAPTER 2. EFFECT OF CEO2 NANOMATERIAL SURFACE FUNCTIONAL GROUPS ON TISSUE

AND SUBCELLULAR DISTRIBUTION OF CE IN TOMATO (SOLANUM LYCOPERSICUM) †

Jieran Li1, Ryan V. Tappero

2, Alvin S. Acerbo

3, Hanfei Yan

2, Yong Chu

2, Gregory V.

Lowry4 and Jason M. Unrine

1,5, *

1. Department of Toxicology and Cancer Biology, University of Kentucky, Lexington, KY,

40546, USA

2. National Synchrotron Light Source II, Brookhaven National Laboratory, Upton, NY,

11961, USA

3. Consortium for Advanced Radiation Sources, University of Chicago, Chicago, IL, 60637,

USA

4. Department of Civil and Environmental Engineering, Carnegie Mellon University,

Pittsburgh, PA, 15123, USA

5. Department of Plant and Soil Sciences, University of Kentucky, Lexington, KY, 40546,

USA

†This chapter is published online in Environmental Science:Nano

* Corresponding Author: [email protected]

mailto:[email protected]

14

2.1 Synopsis

Using recent advances in X-ray microscopy, this study aimed to elucidate

mechanisms of uptake, subcellular distribution, and translocation of functionalized CeO2

MNM (manufactured nanomaterials), having different charges, by tomato plants (Solanum

lycopersicum cv Micro-Tom). We found that plant growth and Ce concentrations in tissues

were functions of surface charge and exposure concentration. Mechanisms of entry into

roots and translocation within plants were examined using X-ray nano- and microprobes.

There were dramatic differences in the tissue and subcellular distributions of Ce in plant

roots exposed to dextran-coated CeO2 nanoparticles conjugated with positive, neutral and

negative functional groups. The results suggest that particles have drastically different tissue

distribution patterns across the root cross section depending on their surface charges. Ce likely

entered through the gaps between epidermal cells being sloughed off during root growth

and penetrated deeper into the interior of the roots (vasculature) via a combination of

apoplastic and symplastic transport. Evidence of symplastic Ce transport was observed with

the neutrally and negatively charged particles. We observed evidence of endocytosis as the

mechanism for entry into the symplast allowing for entry into the xylem. This study

provides critical information on how particle surface chemistry influences the

biodistribution and cellular localization of nanomaterials in plants and is to date the highest

resolution X-ray imaging of nanomaterials in plant cells.

Key words: CeO2 nanoparticles, surface charge, plant uptake, biodistribution

15

2.2 Introduction

In recent years, there has been intense interest in understanding how manufactured

nanomaterials (MNM) interact with plants84, 115, 117, 155-161. This interest initially developed

over concerns about entry of manufactured nanomaterials into terrestrial food chains148, but

more recently, there has been a surge in interest over the use of nanomaterials as targeted

fertilizers and pesticides117, 154, 162.

Identification of the properties that dictate how MNM interact with plants is crucial

to both assessing ecological risks and developing agricultural applications for nanomaterials

(phytonanotechnology). Evidence of bioaccumulation of metal and metal oxide MNM by

plants is abundant, which usually shows nano-specific effects which differ from treatments

with metal ions or bulk materials that are of the same chemical composition 93, 163.

There is still much to be learned about mechanisms behind uptake of nanomaterials

in plants and how nanomaterial properties influence these mechanisms. Uptake and

translocation of MNM applied to roots requires entry into the xylem. For MNM to move

from soil to the xylem, they must cross multiple biological barriers, such as the mucilage

and the Casparian strip 164. The path from the periphery of the root towards the vascular

tissue in the center, is critical for translocation of MNM into the leaves and other above

ground tissues in a plant. There are three potential routes for movement of water and

associated materials from the soil to the xylem: apoplastic, symplastic and transcellular 165.

Many have suggested that the apoplastic route through the extracellular spaces in roots is

the primary route of transport of MNM to the xylem 166, given that this route is the path of

least resistance without entry into the cells; however, the presence of particles within

intracellular compartments has also been observed 80, 84, 162, 167-169. Further, the Casparian

16

strip, a water impermeable barrier located in the endodermis which blocks apoplastic

transport, makes it likely that MNM must at some point enter the protoplast of cells to gain

entry to the xylem and be transported to aerial portions of the plant. It should be noted,

however, the Casparian strip is poorly developed in growing root tips, so its effectiveness

as a barrier is unclear.

It is widely known that MNM surface chemistry plays a key role in determining the

uptake, toxicity, biodistribution and subcellular localization of MNM in organisms 156, 170-

172. The influence of surface chemistry on the mechanisms of uptake and biodistribution in

plants has not been well explored and deserves systematic investigation 173. Such

investigations have been previously hampered by a lack of analytical techniques that have

the specificity, spatial resolution, and sensitivity required for the task. Fortunately, new

tools which combine these capabilities have recently become available at the newest

generation of synchrotron light sources. We imaged the subcellular distribution of MNM

in plant roots using a novel hard X-ray nanoprobe (HXN) with unprecedented spatial

resolution (< 15 nm) 174, 175. This was complimented by ultra-high sensitivity, spatially

resolved X-ray absorption near edge spectroscopy (XANES) to determine speciation of the

MNM in fixed plant tissues at ng/g dry mass concentrations.

This study utilizes a set of CeO2 MNM with identical cores and surface polymers,

differing only in functional group substitution and resulting impact on zeta potential.

Previous research has demonstrated that these changes in CeO2 MNM surface chemistry

have profound effects on uptake, subcellular localization and toxicity 170, 176. CeO2 serves as

good model material for studying plant-MNM interactions. It is relatively insoluble in

typical plant growth media and can be easily measured and tracked by a number of different

17

imaging and spectroscopic methods. CeO2 MNM are among the most widely used

nanomaterials found in fuel additives, polishing agents, industrial catalysts, and have

recently received interest for use as plant growth promoters in agricultural production 177-

180. In human health research, CeO2 MNM have received attention both as a potential

therapeutic and as a toxicant. The ability of Ce in CeO2 MNM to undergo reversible

transitions between III/IV valence states allows them to behave both as either a pro- or anti-

oxidant depending on context 37. While adverse effects were suggested for CeO2 MNM in

various biological systems 60, 181, 182, the impact of key physico-chemical properties affecting

the environmental fate and ecotoxicity of these materials such as size and surface coatings

have yet to be established 183. A few in vitro studies have investigated the effects of surface

valence state, size, and surface charge on the localization, intracellular trafficking and

cytotoxicity of CeO2 nanoparticles 172, 176, 184. These studies showed that CeO2 MNM with

neutral polymers localized primarily in the cytoplasm and had minimal toxicity, whereas

positively and negatively charged particles were found in lysosomes and caused

cytotoxicity 176. Our previous research demonstrated that positively charged,

diethylaminoethyl dextran (DEAE) coated CeO2 MNM (2-5 nm) were taken up much more

efficiently and elicited far greater toxicity in nematodes (Caenorhabditis elegans) than

neutral dextran coated CeO2 MNM or negatively charged carboxymethyl dextran (CM)

coated CeO2 MNM. We also demonstrated that the macroscopic distribution of neutral and

negatively charged CeO2 NPs in the leaves of wheat plants was different 170.

The objective of this study was to investigate the mechanisms of uptake and

biodistribution of a set of well-defined CeO2 MNM, differing only in surface functional

groups and by extension zeta potential, in the model organism tomato (Lycopersicum

18

solanum CV Micro Tom). This species is a good model vegetable crop which is easily maintained

in the laboratory and has a sequenced genome. This objective required observing the

localization and chemical speciation of the materials across spatial scales ranging from the

nm scale (10-15 nm) to the mm scale at ng/g concentrations. We hypothesized that surface

chemistry would have a profound influence on tissue distribution of CeO2 MNMs, and that

these differences would have a cellular basis. This study has far reaching implications for

a general understanding of how surface chemistry influences the behaviors of nanoscale

objects taken up by plants. It also provides a powerful demonstration of how material

properties can be tailored to control MNM distribution within plants at the tissue and

subcellular level for potential phytonanotechnology applications.

2.3 Methods

Nanoparticle synthesis and characterization

Dextran coated CeO2 MNM with a nominal 2-4 nm primary particle diameter (DEX-

CeO2) were synthesized using a previously described procedure 170. In this study, primary

particle diameter was determined by transmission electron microscopy (TEM; JEOL 2010F,

Tokyo, Japan). This dextran coating was then functionalized with diethylaminoethyl groups

to confer either a net positive charge (diethylaminoethyl dextran; DEAE-CeO2), or

carboxymethyl groups to confer a net negative charge (carboxymethyl dextran; CM-CeO2).

Details of the synthesis and functionalization of the particles can be found in the Supporting

Information (SI). Mean hydrodynamic diameters and electrophoretic mobilities of the MNM

treatment suspensions were measured using a Nano-ZS zetasizer (Malvern Instruments,

Malvern, United Kingdom) using dynamic light scattering (DLS) and phase analysis light

scattering (PALS), respectively. The measurements were made at a suspension

19

concentration of 30 mg Ce/L CeO2 MNMs suspended in 10% Hoagland’s solution. The zeta

potential was calculated using the Hückel approximation using the Malvern software.

Measurement of dissolution from CeO2 in media

The nanoparticle treatments were suspended in 10 mL 10% Hoagland’s media at the

same concentrations used for the experiment (10, 30 and 100 mg/L) and incubated for 24

hours. 5mL of the mixture was filtered through centrifugal devices (Amicon Ultra-4, pore

size 3kD equivalent to 1 nm) and the filtrate was collected. To avoid binding of Ce ions to

the filter media, which could reduce recovery, the filter was pre-conditioned with 1mM

CuSO4 before use. Ultrapure HNO3 was then added to the filtrate to a final concentration of

0.15 M HNO3. The unfiltered aliquot was digested by the microwave-assisted method

described in the exposure section below. Ce content of the filtrate (5mL) as well as the

unfiltered aliquot was analyzed using ICP-MS using previously described methods 170. The

dissolved Ce fraction was calculated as (ng Ce/L in filtrate)/(ng Ce/L in the unfiltered

suspension)*100%.

Exposures

Tomato (Solanum lycopersicum cv Micro-Tom) plants were germinated on a Phytagel

plate (Phytagel, Sigma-Aldritch, St. Louis, MO USA) after which they were transferred and

grown hydroponically in 10% Hoagland’s solution with a photoperiod of 12 h under

fluorescent lights (135 ± 3 μmol photon m−2 s−1 light intensity) at 25°C. Plants were exposed

to particles surface-modified with polymer coatings (DEX-CeO2, DEAE-CeO2, CM-CeO2)

two weeks post germination. Exposure media contained high (100 mg/L) medium (30

mg/L) and low (10 mg/L) concentrations of Ce and the exposure duration was 14 d. Fifteen

20

independent biological replicates per treatment were performed. These exposure

concentrations were chosen based on the principle that a broad range of sub-lethal

concentrations could be tested which caused detectable Ce accumulation in the tissues. The

plants were also exposed under the same conditions to various concentrations of CeCl3 to

determine the uptake of ionic Ce from the media. Plants were then rinsed with DI water and

tissues were collected for determining bulk metal concentration using ICP-MS and for

imaging using synchrotron light sources. Untreated plants were maintained in the same

conditions as negative control.

ICP-MS analyses

Freeze-dried plant tissues (roots and shoots collected separately) were digested with

0.75mL concentrated ultra-pure nitric acid and 0.25mL hydrogen peroxide using a

microwave system. Samples were then diluted with 5% ultra-pure nitric acid for ICP-MS

(Agilent, 7500cx, Santa Clara, CA) analysis. Details of our analytical methods have been

described in detail previously 170.

Synchrotron XRF and XANES

Tissues within 1 mm of the tip of the primary roots were dissected with a razor blade,

fixed in sodium acetate buffer (pH 7.2) containing 4% glutaraldehyde and 1%

formaldehyde and embedded in epoxy resin. Thin sections were cut from the embedded

samples at ~1 µm thickness and mounted on a specially made silicon dioxide slide with a

platinum finding grid for ultra-high-resolution X-ray fluorescence imaging using the Hard

X-ray Nanoprobe (HXN) Beamline 3-ID at NSLS-II, Brookhaven National Laboratory

(BNL). NSLS-II is a medium energy synchrotron operating at 3 GeV. Beamline 3-ID uses

multi-layer Laue lenses (MLLs) for final focusing to provide a minimum focus size smaller

21

than 15 nm. For this work, we used various sampling resolutions of 10~100 nanometers per

pixel, depending on the resolution and throughput needs for the experiment, albeit for a field-

of-view of a few microns. For a complete picture, root tissue cross-sections were imaged at

~1 m resolution using the GSECARS beamline 13-ID-E at (Advanced Photon Source)

APS, Argonne National Laboratory (ANL). The APS is a high energy ring operating at 7

GeV. In addition, spatially-resolved X-ray absorption near edge structure (XANES)

spectroscopy was performed at 13-ID-E to interrogate the Ce speciation in root cells. Lastly,

the second leaf that emerged from plants in three treatments were clipped intact and mounted

with metal-free Kapton tape to aluminum mounts for analysis at Submicron Resolution X-

ray Spectroscopy (SRX) Beamline at beamline 5-ID at NSLS-II, BNL. The SRX had a focus

size of slightly less than 1 m. Details are provided in the supplementary information. All

three beamlines use undulator insertion devices as the X-ray source.

Spectral Fitting and Image Processing HXN and SRX imaging data

Spectral fitting of HXN and SRX data was performed using the PyXRF spectral fitting

program 185. Briefly, a summed spectrum fit by a user-assisted fitting approach determined

global fitting parameters, including the position and peak width of fluorescence peaks,

background, pileup and escape peak and Compton and elastic scatter peaks. Then, elemental

maps were generated based on the integrated fluorescence intensities determined by the

spectral fitting.

Collection of XANES spectra and data reduction

All XANES spectra were plotted and normalized to the region 15-150 eV post edge

(the Ce L3 edge at 5726 eV) using Athena 186. For the root sections, XANES spectra were

collected from regions of interest across the entire root and were presented as merged spectra

22

for each treatment. For leaf samples, three XANES spectra were acquired for multiple

regions of interest. With the DEAE-CeO2 and CM-CeO2 treatments, spectra were merged

for all spots examined. We used bulk CeO2 and CeCl3 as standards for Ce IV and Ce III,

respectively. The standards were finely ground to a particle size which corresponded to less

than one absorption unit thickness (to avoid self-absorption effects) and mounted on

cellulose acetate adhesive tape. Layers of the adhesive tape were stacked until the standards

resulted in an approximately 1 absorption unit edge-step.

Statistical analyses

The SAS software package was used for statistical analysis. The data were tested for

normality using the Shapiro-Wilk test. The distribution if shoot concentrations of Ce was

non-normal hence a non-parametric Kruskal−Wallis one-way ANOVA was used followed

by pairwise comparison using the Mann−Whitney U-test. Data points for biomass and root

Ce concentrations were found to be normally distributed and were analyzed by one-way

ANOVA followed with post-hoc Tukey’s test for pairwise comparisons. Difference with

p<0.05 was considered to be statistically significant. We used 15 replicates per treatment

and all were included in statistical analyses.

2.4 Results and discussion


We utilized a set of CeO2 MNMs that differed only by zeta potential, as conferred by

substituting different functional groups on their surface polymers. The synthesis and

characterization of these MNM has been described previously 170. We synthesized dextran

coated CeO2 MNMs (DEX-CeO2) and then substituted diethylaminoethyl groups (DEAE)

into the dextran to confer positive charge or carboxymethyl groups, (CM) to confer negative

23

charge. The primary crystallite size of the CeO2 cores, as measured by transmission emission

microscope (TEM), ranged from 2-4 nm. Dynamic light scattering (DLS) and phase analysis

light scattering (PALS) showed that the hydrodynamic diameters of the particles were

similar in the exposure media (ranging from 16-22 nm), but the apparent zeta potentials

differed as expected (Table 2.1). The zeta potentials were +13, -3 and -15 for the DEAE-

CeO2, DEX-CeO2, and CM-CeO2 treatments, respectively. Dissolution of the CeO2, as

determined by ultrafiltration (1 nm nominal pore size) and inductively coupled plasma mass

spectrometry (ICP- MS), showed that negligible dissolution (<0.31%) occurred in the

exposure media for all three particle types over the timescales of the experiments (Table 1).

Toxicity

One-week after germination, tomato seedlings were transferred from germination

plates to exposure solutions containing 10, 30, and 100 mg Ce/L in the forms of DEX-CeO2,

DEAE and CM-CeO2 for 14 d. Compared to control, tissue dry mass was significantly

reduced in particle-treated groups for both roots and shoots after two weeks of exposure (Fig

2.1). Positively charged DEAE-CeO2 treatment caused significantly greater growth

inhibition than the other two treatment groups. Light microscopy of the root tips revealed

cell damage (Fig 2.2). Lesions resulting from cell death can be clearly seen in the root tip

regions of plants in the DEAE-CeO2 treatment. Cell shrinkage was observed in roots from

the DEX treatment. Such cellular injury can severely hamper the function of the roots, such

as the ability to absorb and transport minerals, which might explain the chlorotic symptoms

observed in the leaves (Fig. 2.S1).

In contrast to our observations, previous studies have shown moderate toxicity for

CeO2 NPs in various plant species 100, 118, 187. Sometimes, even beneficial effects were noted

24

for these particles on seed germination and seedling growth 187. However, particles used in

these studies were without surface coatings and considerably larger in size (>20 nm) as

compared to particles used in this study (2-4 nm) 100, 118, 187. The increased toxicity of our

materials relative to those used in previous studies may relate to increased reactivity of CeO2

particles of <10 nm in diameter, enhanced colloidal stability of polymer coated CeO2 and/or

the charged functional groups, particularly the positively charged DEAE groups.

Tissue distribution

Root Ce concentrations were drastically different among the treatments (Fig 2.3), with

the DEAE-CeO2 (positive) and DEX-CeO2 (neutral) treatments having much higher Ce

concentrations than CM-CeO2 (negative). The outermost surface of the root tip is covered

with a mucilage layer which consists of polysaccharides and proteinaceous substances.

Under typical soil pH values, this layer gives the root a negatively charged surface, which

would therefore tend to have a greater affinity for neutral and positively charged MNM than

negatively charged MNM. This result is consistent with a similar trend of root association

as influenced by surface coating and charge noticed in previous studies 188-190, suggesting a

consistent mechanism governing the interaction. Our previous study with the same particles

used in this study revealed a similar trend of tissue distribution and similar root masses of

Ce within wheat plants (Triticum aestivum) exposed for up to 34h 191.

Analysis of the shoot tissues revealed that accumulation in the roots did not predict

the extent to which Ce was translocated to the aerial part of the plants. Despite having lower

root concentration, a greater quantity of the CM-CeO2 (negatively charged) coated MNM were

translocated to the shoots than the other two treatments (Fig. 2.3C). This, together with the

fact that only a small fraction of Ce was translocated from the roots to the shoots, indicates

25

that the majority of Ce associated with the root tissue did not enter the xylem, which is the

only route by which materials are carried from root to shoot.

We also examined the consequences of exposure to ionic Ce using dissolved CeCl3

(Fig 2.S2). Growth retardation from the ionic Ce(III) treatment was insignificant when

exposure concentrations were below 5 mg Ce/L. Given the low dissolution rate of CeO2

shown in Table 1, the toxicity and Ce bioaccumulation are very likely the result of particle

uptake. Furthermore, under the highest CeCl3 exposure concentration (15 mg Ce/L), which

was toxic to the plants, less Ce was translocated to the shoots, whereas for the particle

treatments, translocation rates remained greater even with plants suffering from the severest

toxicity (DEAE-CeO2-treated). Given that the Hoagland’s medium contains phosphate, it is

possible that CePO4 precipitate formed in the CeCl3 treatment, limiting uptake.

Subcellular distribution and speciation of Ce in root tips

Numerous previous studies have demonstrated that a variety of plant species take up

various types of MNM. However, the mechanism of uptake has remained enigmatic. A

recent study investigating exposure of alfalfa (Medicago sativa) to Ag and Ag2S

nanoparticles indicated that the root tip is the site where nanoparticles likely enter the plant

166. Data for the overall distribution of Ce in cross sections of the root tip was obtained at

~1 µm resolution showed that distributions of Ce in the tissue are different for the three

treatments. Ce appears to be co-localized with the cell wall/cell membrane boundaries in

DEX-CeO2-treated root at this resolution (Fig. 2.4 A, B). In the CM-CeO2-treated root, Ce

was found almost across the entire root cross section. Scattered hotspots were visible as

well as regions with a more diffuse distribution (Fig. 2.4 F, G). For the DEAE-CeO2-treated

26

root (positive charge), Ce was found primarily on the epidermis, with little in the interior of

the root (Fig. 2.4 K, L).

Images showing the cellular and subcellular distribution of Ce within the root tips

using the HXN are shown in

Fig 2.4 C, D, E, H, I, J, M and Fig. 2.S4. These images show

entry of Ce into plant roots at the cellular level. Clearly observed in the DEX-CeO2 (neutral)

treatment, is Ce present within spaces between the epidermal cells that are partially separated

from the epidermis, presumably as a result as cells are being shed from the epidermis during

root tip growth (Fig. 2.4C, 2.S4). The presence of Ce in the apoplastic spaces between cells

(Fig 2.4D, E and Fig. 2.S4), which are approximate 100-200 nm across, suggests that

movement through these spaces is the route of passage from the epidermis to the vascular

tissue. The presence of intracellular Ce hotspots in the cytoplasm of cells near the vascular

bundle in the center of the root suggests that Ce was actively internalized by the cells within

this region (Fig 2.4E). Within the stele itself, cells appear to show Ce both within the

apoplast and throughout the cytoplasm within intracellular vesicles which are likely

endosomes (Fig 2.4E and Fig 2.5). The Casparian strip is a barrier in the endodermis that

prevents mass transfer from the cortex into the xylem. It has been an open question as to

how particles traverse this barrier. One hypothesis was that they move through poorly

developed regions of the Casparian strip in the root tip. Without completely ruling out that

hypothesis, we conclude based on the evidence presented here that the particles are also

endocytosed within the inner regions of the cortex and transferred across the endodermis

symplastically.

XANES spectroscopy is very useful in distinguishing between the two primary

oxidation states of Ce (Ce(III) and Ce(IV)), with Ce(III) producing a solitary absorption

27

peak or “white line” at 5722 keV, and Ce(IV) producing two peaks at 5725 and 5732 keV.

The particles mostly consist of Ce (IV) as-synthesized, although a small percentage of the

atoms are in the Ce (III) state due to oxygen vacancies as shown in our previous work 192.

An important question is whether this material is intact DEX-CeO2 or transformation

products. We conclude that at least a portion of this material is intact CeO2. The proportion

of Ce(IV) in the epidermis, cortex, and vasculature is 47, 32 and 36 percent, respectively

(Fig 2.6). Given that the percentage of Ce(IV) is similar in the cortex, which is where Ce

was primarily located in the apoplastic space, was similar to the vasculature, where Ce was

primarily found in the cytoplasm, it is likely that both intact CeO2 and transformation

products were internalized into cells. However, Fig 2.5 clearly shows, that if present, these

transformation products must have been nanoscale or smaller, given that the size of the Ce

hotspots in the cytoplasm was less than 20 nm. CePO4 particles seen in previous studies

as transformation products are typically larger than this 126, 193. Dextran-coated CeO2

particles within this size range (<5 nm) can accommodate Ce (III) atoms within their crystal

structure and these atoms can reversibly transition between the +III and +IV oxidation

states 194, 195, so it is possible that these <20 nm hotspots contain CeO2 particles with a large

percentage of oxygen vacancies on the surface giving them a greater percentage of Ce (III)

194. The presence of Ce ions bound to macromolecules is unlikely as they would likely

have had a much more diffuse distribution as has been seen with Cu ions in tissues versus

Cu oxides 53.

A previous study suggested that the primary site of biotransformation is on the

surface of the epidermis 193. Given that the percentage of Ce (IV) was similar in epidermis

and the vasculature, we cannot rule this out. It is important to note that the particles used

28

in our study are far smaller at 2-4 nm, than particles used in this previous study 193 which

used 25 nm particles, and are likely more readily reduced 194, 195, so they may behave

differently intracellularly. In our previous study, differences in Ce(IV)/Ce(III) ratios were

observed for hydroponic exposures to wheat plants, with a higher Ce(IV)/Ce(III) ratio being

observed in areas with greater total Ce concentration, and a lower Ce(IV)/Ce(III) ratio

observed in areas with less total Ce 188. This suggests differential transport of transformation

products within the tissues, relative to intact materials.

A very different distribution pattern emerged in the CM-CeO2 (negative) treatment

(Fig 2.4G). In addition to the apoplast, Ce was observed in the cytoplasm throughout the

cells in the cortex (Fig 2.4 H&I). High resolution HXN imaging clearly identified multiple

circular/spherical structures of around 100 nm in diameter within the cells containing Ce

(Fig. 2.4 J), again suggestive of the presence of particles within endosomes. Thus, it appears

that the negatively charged particles are more actively endocytosed within all regions of the

cortex and exist in both the symplastic and apoplastic compartments of the root tissue. This

suggests to us that the cortical cells preferentially endocytose negatively charged particles.

It is possible that the DEX-CeO2 particles began to be endocytosed within the inner regions

of the cortex as they gained a negative charge due to the formation of a protein corona. It

has been shown that acquisition of a protein corona, or surface coating of proteins, confers

a net negative charge to nanoparticles with a wide variety of initial surface chemistries 196,

197. Alternatively, endocytosis is less charge-specific within the inner cortex.

The percentage of Ce (III) in CM-CeO2 treated root tissue decreased from the

epidermis (54%) to the cortex (49 %) and vasculature (24%) (Fig 2.6). This suggests that

untransformed particles were preferentially transported to the cortex from the epidermis,

29

and it does tend to support the hypothesis that the majority of transformation of the CM-

CeO2 particles occurred in the epidermis.

For the DEAE-CeO2 (positive) treatment, the root was completely coated with a thick

(>1µm) layer containing high concentrations of Ce (Fig 2.4 K, L, and M), which correlates

with the high bulk Ce concentration detected in roots by ICP-MS relative to the other two

treatments (Fig 2.3). Low concentrations of Ce within the cortex and vascular tissues made

it impossible to discern the tissue and subcellular level distribution of the DEAE-CeO2

particles. This explains the relatively poor translocation efficiency observed for the DEAE-

CeO2 treatment. It remains unclear whether the necrosis in root tissue observed in this

treatment resulted from impairment of water or nutrient uptake through the root hairs and

epidermis or was primarily caused by internalized DEAE-CeO2 particles, which only

account for a small fraction of the observed Ce. Necrosis in the root tissue is likely a

significant cause of observed decreased growth and foliar symptoms. Our previous studies

of the same materials with the nematode C. elegans found that the DEAE-CeO2 particles

were several orders of magnitude more toxic than the CM-CeO2 or DEX-CeO2 particles,

even after factoring in differences in bioavailability among the particle types 170, 173.

The percentage of Ce (III) was slightly higher in the cortex (34%) than the epidermis

(22%) for DEAE-CeO2 treated plants (Fig 2.6). This suggests two possibilities, either CeO2

continued to be transformed within the cortex, or transformed particles were preferentially

transported to the cortex from the epidermis. Although preferential transport of

transformation products seems unlikely given their larger size, we cannot rule out this

hypothesis.

Cerium distribution and speciation within leaf tissue

30

Because of the low concentrations of Ce found within the shoots, and even lower

concentrations in the leaf tissue itself (Fig 2.S3), we required lower detection limits for

imaging than were possible at HXN. We utilized the Submicron Resolution X-ray

Spectroscopy (SRX) Beamline at NSLS-II which enabled micron-scale imaging and

XANES spectroscopy at low to sub- mg/kg Ce concentrations. Again, surface charge greatly

affected the distribution of Ce within leaves (Fig 2.7). In the CM-CeO2 treatment, the

majority of the Ce was found within the vascular tissue in relatively large foci with few

smaller foci within the mesophyll. The Ce in the DEX-CeO2 and DEAE-CeO2 treatment was

distributed primarily as small foci. XANES spectra collected at the foci from the DEAE-

CeO2 leaf sample, which located primarily in the mesophyll, indicated the predominance of

Ce (IV) (Fig 2.7D). In contrast, almost all the Ce regions in the CM-CeO2 leaf were shown

to contain the reduced form, Ce(III). For the DEX-CeO2 treatment, Ce(III) was still the

major form in the veins while small spots in the mesophyll produced characteristic Ce(IV)

peaks.

Dissolved Ce3+

is likely released through reductive dissolution of CeO2 they

precipitate as relatively large Ce(III) phosphate crystallites. These large elongated

hexagonal Ce (III) phosphate crystals have been previously observed in the apoplast of

cucumber plants exposed to CeO2 MNM 126. While Ce L-edge XANES is not capable of

discriminating between different Ce (III) compounds, Zhang et al. 126 used FTIR

spectroscopy to confirm that these crystals, which they observed using TEM, were CePO4.

The spatially resolved XANES spectroscopy presented here also show that the

untransformed CeO2 is accumulated primarily within the mesophyll, while Ce (III)

accumulated within the vasculature (Fig 2.7). This is most clearly shown for the DEX-CeO2

31

treatment where merged spectra from the mesophyll are 73% Ce (IV), while the veins

contain only 46% Ce (IV). These very small Ce (IV) hotspots may be intact particles while

the larger elongated structures observed in the veins may be the hexagonal CePO4 structures

observed by Zhang et al. 126.

2.5 Conclusion

Surface functionalization had a profound influence on the uptake, tissue and

subcellular distribution CeO2 MNM in the plants. Endocytosis also appears to be more

efficient for negatively charged CeO2 than positively charged CeO2 or neutral CeO2 in the

plant root tip cells. This likely explains the dramatic difference in translocation of these

differentially charged CeO2 MNM. When MNM age in natural soil, they are likely to

ultimately gain a coating of natural organic matter, conferring a net negative charge. This

may ultimately enhance their translocation to leaves regardless of initial surface chemistry.

Finally, the differential tissue and cellular distribution as a function of particle surface

chemistry suggests that targeted, MNM-mediated delivery of agrochemicals to different

biological compartments within the plant is possible at both the tissue and subcellular levels.

Acknowledgements

This material is based upon work supported by the National Science Foundation under

Grants 1530594 and 1266252. This research also used the Hard X-ray Nanoprobe (HXN)

Beamline at 3-ID and the Submicron Resolution X-ray Spectroscopy (SRX) Beamline at 5-

ID National Synchrotron Light Source II, a U.S. Department of Energy (DOE) Office of


32






06CH11357. The authors acknowledge Y-C. Chen-Wiegert, L. Li, J. Thieme, W. Rao, J.

Begley, S. Sutton, M. Newville, and A. Lanzirotti.

33

2.6 Tables and figures

Table 2.1 Characterization of CeO2 MNMs

Size distribution, zeta potential, and dissolution of polymer coated CeO2 nanoparticles in

10% Hoagland’s medium (DEAE = diethylaminoethyl dextran coated CeO2; DEX =

dextran-coated CeO2; CM= carboxymethyl dextran-coated CeO2). Dissolved Ce measured

after 24 hours. Data are means with standard deviations in parenthesis.

Nanoparticle

treatments

Volume weighted

hydrodynamic

diameter

Apparent zeta

potential

Dissolved Ce

in media (%)

DEAE-CeO2 16 nm (2.3 nm) +13 mV (0.67 mV) 0.22 (0.07)

DEX-CeO2 22 nm (3.5 nm) -3 mV (0.36 mV) 0.31 (0.06)

CM-CeO2 20 nm (3.8 nm) -15 mV (1.23 mV) 0.26 (0.04)

34

Figure 2.1 Biomass reduction

Reduction in biomass production for shoots (A) and roots (B) upon treatment with CeO2

nanoparticles coated with diethylaminoethyl dextran (DEAE, +), dextran (DEX, N) or

carboxymethyl dextran (CM, -). Statistical significance detected by post-hoc Tuckey’s

tests on the coating factor is denoted by * (between treatment groups) and # (treatment vs.

control), p<0.05 and p<0.01 levels are indicated by single and double symbols.

35

Figure 2.2 Damage to the root tips

Micrographs showing histological lesions at the root tips. A, control, B, DEAE, C, DEX.

Damage to the meristem is pointed out by arrows in B&C. There are extensive cellular

abnormalities and void necrotic regions around the meristem. Scale bar = 50 µm.

36

Figure 2.3 Bulk Ce concentration in tissue

Bulk tissue concentration of Ce in root and shoot was determined for treatment groups

DEAE, DEX, and CM after two weeks of treatment. See text for details. The same

statistical procedures were used to analyze the data set as in Fig 2.1. Significant differences

detected by post-hoc Tuckey’s tests on the coating factor are denoted by * (p<0.05) and **

(p<0.01).

37

Figure 2.4 Ce distribution at root tips

X-ray fluorescence micrographs of root tip cross sections for DEX (A-E), CM (F-J), and

DEAE (K-M). B, G, and L are 1 µm resolution coarse maps of Ce distribution throughout

the root collected at Advanced Photon Source sector 13. Images C, D, and E represent

magnified view of the rectangular regions in B, and were collected at the Hard X-ray

nanoprobe (HXN) beamline of the National Synchrotron Light Source II at a 60 nm

sampling resolution. Image E is further described in Figure 2.5. These images show

subcellular features near the epidermis, cortex and stele, respectively. Images H, I are 60

nm sampling resolution images of the rectangular regions in G, showing distribution in the

cortex and at the endodermis, respectively. The squared region in J was mapped with ultra-

high 15 nm resolution (H), showing signs of endocytic transport of Ce (putative vesicular

structures indicated by arrows). Image M is a 60 nm resolution image of the rectangular

region in L, at the epidermis. A, F and K are light micrographs of the root sections. Scale

bars: B,G, and L-50 m, C,D,G,H and L- 5 m, E- 2 m, I- 200 nm. Cy = cytoplasm; Ap

= apoplast; Va = vacuole; En = endosome. (Figure on next page)

39

Figure 2.5 Evidence of cytoplasmic distribution

A 60 nm sampling resolution image collected at NSLS-II from near the stele of the root

from a DEX treated plant. Brighter areas indicate areas of increased Ce La fluorescence

intensity. Clearly visible in this image is Ce within the apoplast and within putative

endosomes located in the cytoplasm surrounding the vacuole (annotated at the right).

40

Figure 2.6 Ce speciation at the root tip

In situ Ce L3-edge XANES spectra of the root cross sections. A-C) Merged spectra from

hot spots in different regions of the section. DEAE: #1-12, epithelium; #13, cortex. DEX:

#7-8, epithelium; #5,6,9, cortex; #1-4,10, vasculature. CM: #1,2,7-11, epithelium; #3-4,

cortex; #5-6, vasculature. D) Linear combination was used to fit the spectra with the two

standards. The respective contributions of Ce (III) and Ce (IV) are not forced to sum to

100%. Quality of the fit is denoted by reduced chi-squared. Epi, epithelium; Cor, cortex;

Vas, vasculature. Scale bar in the XRF graph = 50 µm. Examples of individual fits are

shown in the Fig S5. The horizontal lines in figure C are from scattered X-rays from the

platinum grid on the sample holder, not Ce.

41

Figure 2.7 Ce distribution and speciation in leaf tissue

XRF scans and in situ Ce L3-edge XANES spectra of the leaf samples from three

treatments. A) CM, B) DEX, C) DEAE. Signals of Ce and K are represented in the red and

green channel of the images, respectively. Regions from which XANES spectra shown in

Fig 2.6 were collected are indicated by numbered squares. Scale bar = 20 µm. D) Merged

XANES spectra from leaf areas denoted by squares. DEX1 are collected from hotspots

within veins (#2,3&6 in Fig.2.7B), DEX2 from hotspots in the mesophyll (#1,4,5&7 in Fig

2.7B). E) Linear combination was used to fit the spectra with the two standards. The

respective contributions of Ce (III) and Ce (IV) are not forced to sum to 100%. Quality of

the fit is denoted by reduced Chi-squared.

42

2.7 Supplemental information

Additional details for synchrotron X-ray methods

The goal of the synchrotron-based micro X-ray Fluorescence experiments (XRF) was

to evaluate the effect of surface charge/coating on localization of nano-Ce in roots and

leaves of tomato plants. Roots of tomato plants exposed to nano-Ce having neutral,

negative or positive surface charge were harvested and prepared as thin sections by routine

electron microscopy (EM) processing methods: fixation in 4% paraformaldehyde + 1%

glutaraldehyde mixture followed by dehydration series in methanol and then resin

infiltration series in LR white resin. Root cross-sections approximately 1 mm from the

apex were cut to 1 micron thickness using an ultramicrotome and mounted onto silicon

dioxide backing material for -XRF; serial sections were prepared on glass microscope

slides for light microscopy and stained with toluidine blue dye to define cellular

boundaries. Routine EM processing is assumed to cause redistribution of mobile elements

(e.g., aqueous K+ ion) in cells and tissues; however, it is unlikely to alter the localization

of discrete or agglomerated particles such as CeO2.

Ultra-high spatial resolution (circa 15 nm) -XRF images of root cells were acquired

at the Hard X-ray Nanoprobe (HXN) beamline at the National Synchrotron Light Source

II (NSLS-II), and -XRF images of entire root sections were acquired at GSECARS

Beamline 13-ID-E (The University of Chicago) at the Advanced Photon Source (APS).

Additionally, fresh tomato leaves from the third position on the apical stem were harvested

and mounted onto Kapton tape and select regions were imaged at the Sub-micron

Resolution X-ray (SRX) beamline at the National Synchrotron Light Source II (NSLS-II).

43

For -XRF experiments, the incident beam energy was fixed at 12 keV. At the HXN

beamline, samples were oriented 15° to the incident beam and scanned on-the-fly in the

transverse direction using nano-positioning stages with laser interferometer feedbacks.

HXN is an undulator-based, scanning X-ray microscope using a new class of X-ray

nanofocusing optics -known as multilayer Laue lenses (MLLs) -to enable imaging

experiments at a resolution of 10 nm, with an ultimate goal of about 1 nm1. Images were

typically collected for a 1 to 10 m2 area using a pixel size of 10 nm (fine resolution) to

100 nm (course) and a transit time of 50 to 100 ms. At the GSECARS 13-ID-E beamline,

samples were oriented 45° to the incident beam and scanned on-the-fly in the transverse

direction. Beamline 13-ID-E is an undulator-based hard X-ray microprobe instrument

which utilizes dynamically-figured, grazing incidence silicon mirrors placed in Kirkpatrick

Baez geometry for X-ray focusing.2 Images were collected for areas up to 350 m2 using

a pixel size of 1 m and a transit time of 100 ms. At the SRX beamline, samples were

oriented 45° to the incident beam and sub-millimeter regions of tomato leaf were scanned

with a step size of 5 to 20 microns using a dwell time of 1 second. Full X-ray fluorescence

spectra for each pixel were acquired using Quantum Detectors Xspress3 Electronics with

a Hitachi Vortex ME3 silicon drift diode (SDD) detector (HXN and SRX) or a Hitachi

Vortex ME4 SDD detector (GSECARS) sitting at 90° to the incident beam and in the plane

of the storage ring. The fluorescence signal was normalized to the changes in intensity of

the X-ray beam (I0). Data processing for HXN and SRX was performed with NSLS-

II/PyXRF software (https://github.com/NSLS-II/PyXRF) and with U. Chicago

CARS/LARCH software (https://github.com/xraypy/xraylarch).

44

1. Chu, Y.S., H. Yan, E. Nazaretski, S. Kalbfleisch, X. Huang, K. Lauer and N.

Bouet. 2015. Hard X-ray nanoprobe facility at the National Synchrotron Light Source II.

SPIE.

2. Lanzirotti, A., M. Newville, L. Manoukian, and K. Lange. 2016. High-speed,

coupled microbeam XRD/XRF/XAFS mapping at GSECARS: APS Beamline 13-ID-E.

The Clay Minerals Society Workshop Lectures Series, 21, 53-64.

45

Figure 2.8 Chlorosis

Fig 2.S1. Signs of chlorosis in leaves (treatment level mg/L). A. DEX and CM, B. DEAE,

C. Control.

46

Figure 2.9 Ionic control

Fig 2.S2. Uptake of ionic cerium from CeCl3 (0.5, 1.5, 5, 15 mg/L Ce). A, dry masses of

shoots and roots. B, C, tissue concentration of Ce.

47

Figure 2.10 Ce concentration in stem vs. leaf tissue

Fig 2.S3. Cerium concentration in the leaves is much lower than in the stem. DEAE(+),

DEX(N), CM(-).

48

Figure 2.11 Additional XRF coarse graphs

Fig 2.S4. Additional XRF coarse tiling over a large area were used to locate specific

regions of interest for high-resolution scans. A, DEAE, B, DEX (aside is a corresponding

light microscope image of the part of the section scanned by XRF), C, CM. Scale bars are

10 µm in A and C and 5 µm in B.

49

Figure 2.S4. (continued)

50

Figure 2.S4. (continued)

51

Figure 2.12 Example of linear combination fitting of the XANES spectra

Fig 2.S5. Fit of XANES spectra from the epithelial regions of the cross sections A)

DEAE, B) DEX and C) CM.

52

CHAPTER 3. TROPHIC TRANSFER OF DIFFERENTIALLY CHARGED CEO2 NANOPARTICLES

FROM TOMATO (SOLANUM LYCOPERSICUM) PLANTS TO TOBACCO HORNWORM

(MANDUCA SEXTA) LARVAE

Jieran Li1 and Jason Unrine1,2

Departments of 1Toxicology and Cancer Biology, and 2Plant and Soil Sciences, University

of Kentucky, Lexington, KY USA

3.1 Synopsis

We exposed tomato plants with differentially charged CeO2 MNM and fed the leaves

to the tobacco hornworm (Manduca sexta). Differential trophic transfer was observed as a

function of the surface charge of the particles. An uptake and elimination study was

conducted to obtain a time course of Ce dynamics. Despite no observed overall


higher bioaccumulation factors than that of ionic Ce3+. The uptake-elimination dynamics

were influenced by the surface charge of the NPs. Positively charged NPs had higher

bioaccumulation factors and assimilation efficiencies but lower elimination rate than

neutral and negatively charged CeO2 MNMs. These results highlight the importance of

MNM surface chemistry in determining their trophic transfer. This is the first study to show

that the effects of MNM surface chemistry on bioavailability persist through trophic

transfer.

53

3.2 Introduction

Previous studies have shown that various types of manufactured nanomaterials can

be taken up by the plants and transported to different tissues and organs 127, 158, 160, 198, with

adverse effects on many crop plants including morphological changes, altered growth,

biochemical and physiological changes, and genetic modifications 83, 95, 160, 199-203. As plants

are primary producers in terrestrial ecosystems, there is a concern about the trophic transfer

of MNMs from plants to consumers. While many studies have examined trophic transfer

of MNMs in the aquatic environment 142, 144, 147, 204-206, only a handful of studies have

explored trophic transfer of MNMs in terrestrial systems 149-153, 207. Although they used a

wide range of metal-based nanomaterials, these studies have shown a common trend that

exposure to nanomaterials results in higher body burden and trophic transfer rate than

exposure to the corresponding metal ions or bulk metal compounds. These studies have

also not systematically investigated how surface chemical properties, such as surface

charge affect trophic transfer, despite the profound effects that surface charge has on plant

uptake and translocation of MNMs from roots to shoots 124, 208.

CeO2 MNMs are a group of materials widely used as diesel fuel additives, polishing

agents, chemical-mechanical planarization agents, and redox catalysts. While sharing a

core chemical composition, these materials also possess unique properties such as size,

crystalline phase and surface functionalization 178, 180, 209, 210. CeO2 MNMs released into the

environment have been shown to have potential impacts on ecosystems, to various degrees

182, 211, 212. A number of studies provided evidence of how CeO2 MNMs, having various

physico-chemical properties, could interact with biological systems. In vitro cultured

mammalian cells were found to endocytose CeO2 MNMs and transfer them to various

54

intracellular compartments via different mechanisms based on their size and surface charge

176, 184. Polymer-coated particles with positive or negative charges on the surface co-

localized with lysosomes and had greater cytotoxicity than particles coated with neutrally

charged polymer, which were primarily found in the cytoplasm 176. Studies using whole

organisms found that the charge of the polymer coating had a great impact on the uptake

efficiency and toxicity of CeO2 MNMs, whereas the toxicity of uncoated CeO2 MNMs was

primarily dictated by the valence state of Ce on the surface 170, 213. Furthermore, our

previous research on tomato plants demonstrated the uptake, translocation, sub-cellular

distribution and biotransformation of CeO2 MNMs were functions of surface charge

(Chapter 2). We made a key observation that different spatial distribution patterns of

oxidized vs. reduced forms of Ce in the leaves of the plants might be indicative of the

biotransformation processes that occurred in the roots after uptake. Scattered small foci

containing Ce in the mesophyl tissue and larger foci of Ce in the vasculature (leaf vein)

were found to contain primarily Ce(IV) and Ce(III) species, respectively. Given that insect

larvae, such as hornworms, primarily assimilate the mesophyll tissue, they may

preferentially take up the untransformed CeO2 materials, which contain primarily Ce (IV)

versus the Ce (III) transformation products.

Trophic transfer of CeO2 MNMs from the plants to herbivores such as insects has

not been well documented. Upon entry into the digestive system of insects, the uptake or

retention of particles occurs under the influence of complex biochemical and physiological

factors of the alimentary canal such as pH, redox potential, and enzyme activity 214-216. Our

previous research found evidence of uptake of Au nanoparticles (5, 10, and 15 nm) in the

midgut of tobacco hornworm 149, 207. The alkaline environment of the midgut (pH > 8.0)

55

can hypothetically help stabilize the negative surface charges on the particles and reducing

aggregation. In two other studies, it was shown that nano-sized CeO2 MNMs had greater

potential for bioaccumulation and trophic transfer than bulk CeO2 151, 152. These results

illustrate the importance of particle size in the context of trophic transfer of MNMs, as

smaller particles may have a greater chance of crossing the major physical barrier in the

alimentary canal known as the peritrophic matrix 217.

In this study we examined the trophic transfer of a set of well-defined polymer-coated

CeO2 MNMs, differing only in the functional groups and therefore charges carried by the

polymer coatings, in a simulated food chain. Herbivorous tobacco hornworm larvae

(Manduca sexta) were fed with leaves of tomato plants (Lycopersicum solanum CV Micro

Tom) that were pre-exposed to these CeO2 MNMs. We show that the rate of trophic transfer

and bioaccumulation is influenced by surface charges. By tracking the body concentrations

of Ce in tobacco hornworms, we were able to study the uptake and elimination dynamics

of Ce, which in combination determined the overall trophic transfer efficiency. This is the

first investigation into the effect of surface charge on the trophic transfer of Ce from CeO2

nanoparticles in a terrestrial food chain.

3.3 Methods


Nanoparticles were synthesized and characterized according to the procedure we

described previously 170. Briefly, CeO2 MNMs with a dextran coating were synthesized

using a modified alkaline precipitation method. Dextran coated CeO2 (DEX) was then

functionalized with either diethylaminoethyl groups (DEAE) or carboxymethyl groups

56

(CM) to confer net positive and net negative charges, respectively, over a wide pH range.

Monodispersed particles in deionized water at 30 mg Ce/L were deposited on copper grids

and dried for Transmission Electron Microscopy (TEM). Hydrodynamic diameters and

electrophoretic mobilities of the MNMs were measured by Dynamic Light Scattering

(DLS) and Phase Analysis Light Scattering (PALS), respectively, on a Nano-ZS zetasizer

(Malvern Instruments, Malvern, United Kingdom) at 30 mg Ce/L MNMs suspended in

deionized water. The zeta potential was estimated from electrophoretic mobility using the

Huckel approximation.

Exposure of tomato plants

Tomato (Solanum lycopersicum cv Micro-Tom) plants were germinated on a

Phytagel plate (Phytagel, Sigma-Aldritch, St. Louis, MO USA). Seedlings were transferred

and grown hydroponically in 10% Hoagland’s solution with a photoperiod of 12 h under

broad-spectrum fluorescent lights at 25°C. Treatments started two weeks post germination

with particles surface-modified with polymer coatings (DEX-CeO2(0), DEAE-CeO2(+),

CM-CeO2(-)) (30mg Ce/L) and CeCl3 (10 mg/L). Exposure lasted for two weeks. 10%

Hoagland’s media was used to grow control plants.

Exposure of the primary consumer

Tobacco hornworm (Manduca sexta) eggs were purchased from Carolina Biological

Supply (Burlington, NC, USA) and hatched on tomato leaves from control plants. Once

the larvae reached the second instar, they were randomly divided into groups which were

fed on fresh leaves from plants exposed to DEAE, DEX, CM, and CeCl3 as described

above. The feeding lasted 7 days followed by a 7-day period of elimination when

hornworms were fed with clean leaves from control plants. Larvae and feces were collected

57

for all treatments at 1, 3, 4, 5, 7, 8, 9, 10, 11 and 14 days. For larvae collected from the

exposure phase (days 1-7), clean food was provided for a 12h period to allow clearing of

Ce in undigested plant tissues before the larvae were prepared for analysis.

ICP-MS analysis

Freeze-dried leaves and insect tissues were digested with 0.75mL concentrated nitric

acid (OmniTrace Ultra®) and 0.25mL hydrogen peroxide using a microwave digestion

system in metal free polypropylene centrifuge tubes at 100 C for 10 minutes. Samples

were then diluted to 0.75 M HNO2 with 18 M deionized water (DI) prior to analysis using

inductively coupled plasma mass spectrometry (ICP-MS). Details of this method have

been described previously 170.

Synchrotron XRF mapping of the midgut tissue and XANES analysis

At the end of the exposure (day 7), larvae were dissected and the midgut was

removed, fixed with neutral buffered formalin, and embedded in paraffin wax. The tissues

were sectioned to 10 m using a microtome equipped with a surgical steel knife and

mounted on quartz glass slides. These slides were mounted for analysis at beamline 5-ID

at the National Synchrotron Light Source -II, Brookhaven National Laboratory, Upton,

NY, USA. X-Ray fluorescence (XRF) microscopy experiment was conducted with an

incident X-ray at 10 keV. Micrographs were obtained for DEAE, DEX and CM treatments

using the PyXRF software. The Lα1 and Lβ1emission lines were used to fit the Ce in the

spectra. For three selected areas of elevated pixel intensity, we collected micro-focused X-

ray absorption near edge structure (XANES) spectra. The XANES spectra in each sample

and merged for all spots examined. We used bulk CeO2 and CeCl3 as standards for Ce IV

and Ce III, respectively. The standards were finely ground to a particle size which

58

corresponded to less than one absorption unit and mounted on cellulose acetate adhesive

tape. Layers of the adhesive tape were stacked until the standards resulted in an

approximately one absorption unit edge-step. XANES spectra were plotted and normalized

to the region 15-150 eV post edge (the Ce L3 edge at 5726 eV) using Athena 186.

Data analysis

Assimilation efficiency (AE) was calculated as the ratio of the tissue Ce (ng) to the

amount of Ce (ng) in the consumed food by day 1. Trophic transfer factor (TTF) was

calculated as the Ce concentration in larvae divided by the Ce concentration in leaves. The

SAS software package was used for statistical analysis. The data were tested for normality

using the Shapiro-Wilk test (SAS procedure MULTIVARIATE-Shapiro-Wilk). All data

points for Ce concentrations were found to be normally distributed and were analyzed by

one-way ANOVA with repeated measures followed by post-hoc Tuckey’s test for pairwise

comparisons (SAS procedures ANOVA and GLM). Linear regression analysis was

performed on natural log-transformed data points to estimate the elimination rate constants

(SAS procedure GLM-Parameter Estimate). Fifteen replicates per treatment were used for

the experiment and difference with p<0.05 was considered to be statistically significant.


Characterization of the MNMs

The DEAE, DEX and CM-CeO2 MNMs synthesized for this study share the same

polymer-coated CeO2 core differing only in the surface functionalization. The primary

particle size as measured by TEM was 3.28 ± 0.33 nm (n = 50). The mean volume weighted

59

hydrodynamic diameters in 10% Hoagland’s solution were measured by DLS for the

DEAE, DEX and CM-CeO2 MNMs were 20.7 ± 3.03, 18.5 ± 3.56, and 23.6 ± 2.27 nm

respectively. The apparent zeta potentials as measured by PALS for the DEAE, DEX and

CM-CeO2 MNMs are +16.2 ± 0.74, -2.45 ± 0.21, -18.26 ± 0.69 mV (n = 5). These MNMs

were very similar from those used in our previous study which were synthesized by the

same method (Li et al., Chapter 2).

Surface functionalization profoundly affects trophic transfer of CeO2 MNMs

Second-instar Manduca larvae were randomly divided into four groups and fed

respectively with leaves from plants grown in control media and media containing DEAE,

DEX, CM-coated CeO2 MNMs, and CeCl3. The exposure lasted for 7 days followed by 7

days of elimination where leaves from unexposed plants were provided as food. The

growth of hornworm larvae was not affected by any of the treatments (Fig 3.1). Bulk Ce

concentrations in hornworms at the end of the exposure phase (7 days) were 4~14 fold

lower than in the food (Fig 3.2A). They were orders of magnitudes lower at the end of the

experiment (14 days). The trophic transfer factors (TTF) for the treatments at 7 days were

0.25, 0.13, 0.11, and 0.07 for DEAE, DEX, CM CeO2 MNMs and CeCl3, respectively (Fig

3.2B). TTF values for DEAE, DEX, and CM-CeO2 MNMs decreased dramatically to

0.035, 0.005, and 0.001 at 14 days, with no detectable Ce levels in animals exposed to

CeCl3. The bio-dilution (TTF < 1) of CeO2 nanoparticles is in line with previous studies

showing low trophic transfer of a variety of metal compounds in aquatic as well as

terrestrial environment 140, 141, 146, 151. In contrast, some studies have demonstrated

biomagnification for Au nanomaterials 145, 148, 149. Differences in the trophic transfer

potential of nanomaterials are likely dependent on environmental conditions and material

60

properties, but the physiology and life history traits of the organisms involved also play a

key role 150. For example identical tannate coated Au MNMs biomagnify from tobacco

plants to tobacco hornworm larvae 207 but biodilute from earthworms to bullfrogs 150.

Furthermore, in our study, the particles having different surface chemistries were subject

to differential biotransformation processes in the plants prior to consumption by the insect

larvae, which might dictate the uptake and elimination rates. Thus it is very likely that the

surface chemistry or chemical composition of the particles is altered during the trophic

transfer process. We previously showed that trophic transfer tends to increase the uptake

of Au MNMs, relative to direct exposure of as-synthesized particles administered via oral

gavage 150. In that study, it is unlikely that the Au cores themselves were biotransformed,

but rather their surface chemistry may have been altered such as the acquisition of a protein

corona. For CeO2, however, low pH and the ubiquitous presence of ligands with high

affinity for Ce3+, such as phosphate, shifts the equilibrium of dissolution and re-

precipitation to the formation of insoluble compounds, particularly CePO4 218, potentially

serving as a detoxification mechanism 219. Our previous work (Li et al., Chapter 2) showed

that the surface chemistry of these materials had a profound influence on the extent of

biotransformation of these materials in the tomato plant. We found that CM-CeO2 and DEX

CeO2 were more extensively converted to Ce(III) species than the DEAE particles. We

attributed this to their increased tendency to be endocytosed by cells at the endodermis of

the roots. In the same study, foci consisting primarily of Ce(IV) were found in the

mesophyll tissues of the leaves in DEAE and DEX treated tomato plants, whereas in the

CM treatment Ce(III) was distributed ubiquitously in the vascular system of the leaves.

The vascular tissue is more fibrous and likely more difficult for the hornworms to digest,

61

hence the lower bioavailability of Ce associated with it. Although there are many studies

that presented varied results on the extent of biotransformation of metal-containing

nanoparticles within plant tissues 81, 127, 218, 220, results comparing trophic transfer potentials

under the same conditions were lacking. Our study is the first evidence of the impact of

surface charges on the trophic transfer dynamics of nanoparticles in a terrestrial food chain.

Assimilation efficiencies and elimination rates

The assimilation efficiency (AE) values calculated for the treatments were also found

to follow the same trend as the TTF, in the order of DEAE (4.3%), DEX (2.8%), CM

(2.1%), and CeCl3 (0.2%) (Fig 3.2C). As discussed in the previous section, distinct

distribution patterns and biotransformation of Ce for the DEAE, DEX and CM-coated

CeO2 MNMs in the leaves of exposed plants likely indicates Ce pools of different

bioavailability to the insect larvae. The difference in AEs observed here further suggests

the biodistribution and biotransformation processes occurring in the plant tissue are to

some extent predictive of dietary uptake rate.

We observed a universal increase of body Ce concentration in the exposure phase

followed by a rapid decrease after day 7, when larvae were switched to clean leaves for

elimination (Fig 3.3A). The first-order elimination rate constants were calculated using

four data points in the initial period of elimination (Fig 3.3B). Elimination rates during the

initial elimination phase were also correlated with the trophic transfer factors. Elimination

rate constants ranged from -0.62 to -2.40 in the order of CeCl3 > CM > DEX > DEAE (Fig

3.2). Body Ce content dropped rapidly in the first 72 h and then became asymptotic towards

the end of the experiment. Similar to this result, previous studies on Au and CeO2

nanoparticles showed a lack of elimination for these insoluble materials 150, 221. However,

62

Ce was detected from the feces collected at these time points, suggesting trace amount of

Ce was still being eliminated (Fig 3.3C), possibly from residual food particles. A previous

study observed a similar bi-phasic elimination curve for quantum dots in Daphnia 222,

where the rapid clearing of the GI tract coupled with slow elimination of internalized

quantum dots. Furthermore, it was shown that in caterpillars the peritrophic matrix, a

proteinaceous meshwork synthesized by the insect gut epithelia to enclose the food bolus,

has a relatively long gut transit time whereas undigested food is usually eliminated as feces

within hours 217, 223. The fast elimination of Ce in the initial 72 h after the cessation of

dosing could therefore be a combination of voiding of undigested food and the peritrophic

matrix turnover.

Distribution and speciation of Ce in the midgut

To investigate the anatomical basis of the elimination process, we imaged the

distribution of Ce using a synchrotron-based X-ray microprobe. For all treatments, Ce was

found in scattered regions on the luminal side of the epithelia and within the epithelial cells

(Fig 3.4). The luminal Ce containing particles are likely partially digested plant tissue. Ce

was not detectable on the basal side of the epithelial cells; however, bulk concentrations of

Ce in the larvae after cessation of dosing were below 1 µg Ce/g, which is the detection

limit of the microprobe. It is likely that the initial voidance of the gut contents was followed

by elimination of Ce as the gut epithelial cells, which contained the highest Ce

concentrations, were shed. The remaining portion of Ce that is not eliminated may have

been transported away from the gut epithelium and internalized. In the CM treatment, there

appeared to be either a membrane layers enriched in Ce, possibly the peritrophic matrix

63

lining the luminal face of the epithelia, or fibers in the undigested vascular tissue, which

was not observed in the other treatments.

We obtained XANES spectra at these hotspots to further understand Ce speciation.

The Ce spectra for all three treatments showed that they had the similar oxidation states in

lumen and epithelia for DEX and CM treatments and Ce was present as a combination of

Ce (III) and Ce(IV) with approximately 34 and 63% in the Ce (IV) oxidation state in the

epithelial cells for DEX and CM, respectively (Fig 3.5). The DEAE treatment had different

percentages of Ce (IV) in the epithelial cells and lumen with approximately 51 and 60%,

respectively. Ce (IV) species are very likely intact CeO2, given that Ce is reduced to the

Ce (III) oxidation state during dissolution and may be precipitated as secondary Ce (III)

minerals, such as CePO4. This suggests that Ce is being taken up by the hornworms both

as Ce (III) species and intact CeO2 species. The fact that uptake and elimination rates are

dependent upon initial surface chemistry, also reinforces the conclusion that intact CeO2

was taken up, as the influence of surface chemistry would have been removed once the

CeO2 MNMs have dissolved.

3.5 Conclusion

This study investigated the trophic transfer of CeO2 nanoparticles from tomato leaves

to tobacco hornworms and is the first to reveal the influence of surface charges on the

uptake and elimination dynamics. While several studies have investigated the effect of

MNM surface chemistry on uptake in plants and other organisms, this is the first study

showing that surface chemistry affects uptake along a simulated food chain. The fact that

the influence of surface chemistry persists even after uptake and translocation to plant

64

leaves, is a novel finding. Overall, negatively charged CeO2 MNMs had lower

bioaccumulation and assimilation efficiency than the positively charged and neutral

particles as a result of higher elimination rate. This effect ran counter the opposite trend in

translocation efficiency in plants observed in Li et al., (Chapter 2), where negatively

charged CeO2 was translocated more efficiently than positively charged CeO2. As a result,

the net accumulated body concentrations were similar among treatments, despite lower

concentrations of Ce in the leaves of the DEAE and DEX treated plants, relative to CM

treated plants. These results highlight the importance of MNM surface chemistry in

determining their trophic transfer. This is the first study to show that the effects of MNM

surface chemistry on bioavailability persist through trophic transfer.

Acknowledgements

This material is based upon work supported by the National Science Foundation under

Grants 1530594 and 1266252. This research also used the Hard X-ray Nanoprobe (HXN)

Beamline at 3-ID and the Submicron Resolution X-ray Spectroscopy (SRX) Beamline at 5-

ID National Synchrotron Light Source II, a U.S. Department of Energy (DOE) Office of







06CH11357. The authors acknowledge Y-C. Chen-Wiegert, L. Li, and J. Thieme.

65


Figure 3.1 Growth of insect larvae

Growth of M. sexta larvae throughout the experiment. No significant difference was found

between the treatments and with control (ANOVA with post hoc Tukey’s test, n = 15).

66

Figure 3.2 Accumulation of Ce

A, trophic dilution of CeO2 MNMs from S. lycopersicum cv Micro-Tom to M. sexta

larvae. The concentrations of Ce in the larvae at 7 and 14 days are much lower than in the

leaves of the respectively treated plants. Comparison of the TTF (trophic transfer factor)

(B) and the percent of Ce assimilated from the food (C) for the treatments. Means with the

same letter are not significantly different at Tukey’s test (p < 0.01, n = 15).

67

Figure 3.3 Uptake and elimination dynamics

A, Ce concentrations in M. sexta from the DEAE, DEX, CM and CeCl3 treatments. B, Ce

concentration was adjusted for growth dilution and natural Log-transformed to determine

the elimination constants, assuming first order elimination. The first four points of the

elimination phase were selected as the linear range (dashed rectangle), beyond which no

further reduction of Ce content was observed. Calculated elimination constants (K1-K4)

are shown on the graph. C, Fecal Ce concentrations plotted on a Log scale.

68

Figure 3.4 Ce distribution in the midgut tissue

XRF scans shows the distribution of Ce in the midgut of the larvae from the three

treatments (A-B, DEAE; C-D, CM; E-G, DEX). Signals of Ce and Zn are represented in

the red and blue channel of the images, respectively. Regions from which XANES spectra

shown in Fig 3.4 were collected are indicated by numbered squares. Different anatomic

compartments are labeled: epi = gut epithelia and lum = lumen of the gut.

A lum

epi

epi

F

epi

epi

lum

B C

D

A

G

lum

lum

epi

epi

epi

epi

epi

lum

lum

E

lum

epi

69

Figure 3.5 Ce speciation in the midgut tissue

In situ Ce L3-edge XANES spectra collected from hotspots denoted by numbered squares

in Fig 3.3: A, DEAE, B, DEX and C, CM. Spectra from the same anatomic compartment

are merged and plotted with two standards (Ce(III) nitrate and DEX-CeO2 MNM). D,

Linear combination fitting of the spectra with the two standards shows the ratio of Ce (III)

and Ce (IV) components in the mixture. The respective contributions of Ce (III) and Ce

(IV) to the spectra are not forced to sum to 100%. Quality of the fit is denoted by reduced

chi-squared.

D

A B

C

70

CHAPTER 4. COMPARING TROPHIC TRANSFER OF LAB-SYNTHESIZED NANO-CU(OH)2 TO A

COMMERCIAL NANO-CU(OH)2 FUNGICIDE FORMULATION FROM TOMATO (SOLANUM

LYCOPERSICUM) LEAVES TO TOBACCO HORNWORM (MANDUCA SEXTA) LARVAE

Jieran Li1, Sonia Rodrigues2, and Jason M. Unrine1,3

1. Department of Toxicology and Cancer Biology, University of Kentucky,

Lexington, KY USA

2. Department of Chemistry, University of Aveiro, Aveiro, Portugal

3. Department of Plant and Soil Sciences, University of Kentucky, Lexington, KY,

USA

4.1 Synopsis

To determine if studies conducted with highly purified, lab synthesized materials,

were predictive of behavior of commercial nanopesticide formulations, we studied the

dietary uptake of Cu(OH)2 MNMs by hornworms feeding on surface-contaminated tomato

leaves. We compared lab-synthesized copper hydroxide (Cu(OH)2) nanowire with the

widely used fungicide KOCIDE® 3000, whose active ingredient is nano-needles of copper

hydroxide (Cu(OH)2). The difference in their toxicity and accumulation/elimination

dynamics was found to correlate with the solubility of the materials. Our result provides

valuable knowledge on whether the ecotoxicity of commercial MNM products such as

Kocide can be compared to lab-synthesized counterparts. Nevertheless, future research is

needed to determine to what extent environmental factors such as pH, ionic strength, and

the presence of inorganic ligands and natural organic matter play a role.

71

4.2 Introduction

Modern agriculture has relied heavily on copper-based products for control of fungal

and microbial infections in crop plants since the introduction of Bordeaux mixture (mixture

of CuSO4 and Ca(OH)2) . Recently, materials like cupric hydroxide (Cu(OH)2), copper

oxychloride (Cu(OH)2∙CuCl2), cuprous oxide (Cu2O), and especially engineered

nanomaterials (MNMs) containing copper, have been developed as replacement of

Bordeaux mixture for their higher efficacy, making the accumulation of copper in

ecosystem a serious problem 34-36. Many of these materials are developed and applied as a

mixture of complex ingredients whose specific formulations are a matter of proprietary

interest. This leads to a poor understanding of how they interact with ecological receptors,

making the associated environmental risks difficult to assess.

Previous studies have demonstrated copper-based MNMs are taken up by plants and

elicit a variety of phytotoxic effects, including suppression of growth and seed germination,

changes in physiology and metabolism, and altered gene expression 36, 159, 160, 203, 224-230.

While these studies provide abundant evidence on different routes by which copper is

bioaccumulated by plants, whether higher trophic levels will be impacted by these

materials remains an open question. Although phytotoxicity is carefully avoided when

these copper-based pesticides are intentionally deposited on the aerial parts of crop plants,

a risk that the entire food web lying above these plants might be exposed to copper

contamination exists. Trophic transfer has been shown to occur for MNMs in aquatic and

terrestrial food chains 142, 145, 146, 149, 150, 152, 205, 207, 219. While these studies suggest that the

efficiency of transfer can be influenced by the environmental conditions as well as the

physico-chemical properties of the MNMs, there is a question whether we can make

72

reliable predictions about how commercial pesticide formulations containing MNMs will

interact with the ecosystem components based on studies of the properties of highly

purified, lab-synthesized nanomaterials.

In this study we investigated the trophic transfer of Cu from tomato plants to tobacco

hornworms following the application of the commercially available pesticide, Kocide

3000®, which contains Cu(OH)2 nano-needles as its active ingredient (Kocide) to lab-

synthesized copper hydroxide nano-needles (nCu(OH)2). CuSO4 was used as an ionic

control. Kocide is one of the most widely used commercial pesticides designed to protect

crop plants from fungal and bacterial infection. As the product is prepared in suspension

and sprayed in the field, the nano-sized copper hydroxide provides sustained protection

against fungal infection by a continual release of ionic copper from the slowly dissolving

copper hydroxide particulates on the foliage, while keeping phytotoxicity at a low level 231,

232.

We designed exposures to mimic the realistic scenario in the field, where treatments

were sprayed on the leaves of the plants which then were fed to the insect larvae. By

tracking the amount of copper that remained in the body of the larvae along multiple time

points, the uptake and elimination dynamics of copper from exposure to different materials

was established. From parameters extracted from the simple setting, we built a biodynamic

model that describes potential trophic transfer as a result of long-term exposure. The

toxicity and bioaccumulation of copper sulfate and two forms of nano-copper hydroxide

was also compared using this model, to determine if we can predict the behavior of

composite nano-products with unknown formulation in the environment by using their

active ingredients as surrogates.

73

4.3 Methods

Nanoparticle characterization

Nano copper fungicide Kocide 3000 (hereafter as Kocide) was purchased from

DuPont Inc. The active ingredients were shown to be Cu(OH)2 nano-sheets comprised of

layers of Cu(OH)2 nano-needles, which dissolve to produce sustained release of Cu2+ to

kill microbial pathogens 231, 232. Copper hydroxide nano-needles (nCu(OH)2) was

synthesized using alkaline precipitation methods adapted from a protocol described

previously 233. Briefly, 1.6 mL of 1 M KOH solution was dropped into 4 mL of 0.1 M

CuSO4 solution at room temperature (∼21 °C) with constant stirring, followed by dropping

of 0.6 mL of 12.5 wt % ammonia solution. Then the mixture was stirred for 3 min and

incubated for 12 h (at room temperature). The resulting precipitate was filtered, washed

with ultrapure water, and dried in vacuum for 12 h. The material was characterized by

TEM, SEM and XRD. Materials were dispersed in deionized water at 50 mg Cu per L and

analyzed for hydrodynamic diameters and electrophoretic mobilities using Dynamic Light

Scattering (DLS) and Phase Analysis Light Scattering (PALS), respectively, on a Nano-

ZS Zetasizer (Malvern Instruments, Malvern, United Kingdom).

Measuring the dissolution of MNMs

nCu(OH)2 and Kocide were dispersed in 10mL deionized water at 50 mg Cu/L. The

amount of dissolved Cu2+ in a 5mL aliquot was quantified at 0h, 24h, and 48h by

centrifuging the suspension at maximum speed (3220 × g) for 4 hours on an Eppendorff

5804R centrifuge and collecting the supernatant. Ultrapure HNO3 was then added to the

supernatant to a final concentration of 0.15 M. The other 5mL aliquot of the whole

suspension was digested and analyzed using ICP-MS as described in section 2.3. The

74

fraction of dissolution was calculated as (ng Cu in the 5mL supernatant) / (ng Cu in the

5mL whole suspension aliquot) × 100%.

Surface contamination of tomato leaves

Tomato (Solanum lycopersicum cv Micro-Tom) plants were germinated on a

Phytagel plate (Phytagel, Sigma-Aldritch, St. Louis, MO USA). Seedlings were transferred

and grown hydroponically in 10% Hoagland’s solution with a photoperiod of 12 h under

broad-spectrum fluorescent lights at 25°C. True leaves emerging above the dicot leaves in

the third week post germination were collected from the seedlings. A spray chamber was

constructed using an concentric nebulizer to deliver micro-droplets of nanomaterial

suspensions to the leaves. Test sprays were conducted to manipulate the amount and spatial

distribution of Cu deposited on the leaves. The amount Cu per unit area was adjusted to

mimic the actual application rate of fungicide in the field as specified on the Kocide 3000

label (1.5 kg/ha or 1.34 lb/acre). Leaves were treated with Kocide 3000, lab-synthesized

Cu(OH)2 nano-needles. The droplets were air-dried on to the surface of the leaves.

Exposure of the primary consumer

Tobacco hornworm (Manduca sexta) eggs were purchased from Carolina Biological

Supply (Burlington, NC, USA) and hatched on untreated leaves. Treated leaves were then

fed to M. sexta larvae, starting from the first (5 replicates for each data point) and second

instar (12 replicates for each data point) stages. For the first instar larvae, the exposure

lasted 6 days followed by an 8-day n period in which clean tomato leaves were provided

as food to determine elimination rate. For the second instar larvae, the time spans of the

uptake and elimination phases were 7 days and 10 days, respectively. Larvae and feces

were collected starting from the first day of treatment for all the second instar groups at 2-

75

day intervals. For the first instar groups, caterpillars were sacrificed and analyzed for bulk

copper only at day 12 and day 20, the beginning and end point of elimination. For all larvae

collected at or before the start point of elimination, clean food was provided for a 12h

period to allow clearing of Cu in undigested plant tissues before the larvae were sacrificed.

ICP-MS analysis

Oven-dried leaves and insect tissues were digested in 0.75mL concentrated nitric

acid (OmniTrace Ultra®) using a microwave digestion system in metal free polypropylene

centrifuge tubes at 100 C for 10 minutes. Samples were then diluted to 0.75 M HNO3 with

18 M deionized water (DI) prior to analysis using inductively coupled plasma mass

spectrometry (ICP-MS).

Data analysis

The SAS software package was used for statistical analysis. The data were tested for

normality using the Shapiro-Wilk test and for homoscedascity using Barlett’s test. All data

points for Cu concentrations were found to be normally distributed and homoscedastic

(SAS procedures MULTIVARIATE and GLM). Data were analyzed by one-way ANOVA

followed by Tuckey’s post-hoc pairwise comparisons (SAS procedure ANOVA). Survival

time analysis was used to compare the mortality rates of the treatments and control group

(SAS procedure LIFETEST). Difference with p<0.05 was considered to be statistically

significant.

Trophic transfer factor (TTF) was calculated as the Cu concentration (µg/g dry

tissue) in the hornworm divided by the Cu concentration (µg/g dry tissue) in the food at

the end of the experiment. AE was calculated as the ratio of the tissue Cu (ng) to the amount

of Cu (ng) in the consumed food at the end of the first day of the dosing. We extrapolated

76

the short exposures of the first and second instar larvae (6 and 7 days, respectively) to the

entire span of the larvae stage to estimate bioaccumulation of the contaminants used a

modified biodynamic model 150. Trophic transfer factor at time t, TTF(t), was expressed as

a function of time according to the following equations:

M(t) = Mmax × e –k, (1)

k = b × e -ct, (2)

A(t) = A(t-Δt) + AE × CF × IR × M(t) × Δt, (3)

TTF(t) = A(t)/M(t)/CF, (4)

Where A(t) is the amount of Cu in the larvae at time t, M(t) is the mass of the larvae

at time t, and CF is the Cu concentration in the food which is treated as a constant. M(t)

was described by a truncated Gompertz function (Equation 1), where Mmax is the

asymptotic maximum body mass of M. sexta larvae and the exponential growth rate

gradually declines (Equation 2) ) 234, 235. The last instar larva will stop feeding and enter

the wandering stage prior to pupation, hence Mmax is never reached in the real scenario 235.

A growth curve was fitted to the data points using this equation and nonlinear regression

with a least square estimator was employed to determine the constants b and c, which

correlate to the inflection point and steepness of the curve, respectively. A(t) was calculated

using the modified biodynamical model (Equation 3), where AE is the assimilation

efficiency and IR is the ingestion rate as percentage of the body weight per day averaged

throughout the experiment. The rate of change in A(t) and TTF(t) was calculated

recursively by taking infinitesimal time increments (Δt = 0.2 d).

77


Characterization of the particles

The nCu(OH)2 exhibited a particle size of a mean length of 1442 nm (n=335) and 16

nm diameter (n=32) (Fig 4.S1). Due to interference from the product formulation of the

Kocide, we could not quantitatively assess the particle size distribution; however,

qualitatively, the nano-needles are 10-15 nm in diameter and 300-600 nm in length (Fig

4.S1). The nano-needles in both materials tend to stack to form small nano-sheets (Fig

4.S1). The mean volume-weighted hydrodynamic diameters of the Kocide and nCu(OH)2

were measured at 50mg Cu/L in deionized water to be 232 ± 28 nm (mean ± standard

deviation, n = 3) and >2000 nm, respectively, with substantial aggregation of the lab-

synthesized Cu(OH)2 nano-needles. The apparent zeta potentials were -38 ± 3.5 and +13 ±

4.7 mV (mean ± standard deviation, n = 3), respectively, for the Kocide and nCu(OH)2 (pH

= 6.5). It has been shown that Kocide contains considerable amount of non-copper

ingredients 236, which might modify the surface of nano Cu/Cu(OH)2 to give the Kocide a

net negative charge. Dissolved Cu was also measured at 4, 24 and 48hours after the

suspension was prepared (Table 4.1). Within 4 hours after the particles were dispersed in

suspension, 26.33% and 5.61% of the total Cu was detected, respectively, as Cu ions in the

Kocide and nCu(OH)2 samples. The dissolved Cu stayed unchanged from till 48 hours.

Aggregation of nCu(OH)2 may have contributed to its lower dissolution rate as compared

to the Kocide particles.

Toxicity of CuSO4, Kocide and nCu(OH)2

First and second-instar Manduca larvae were randomly divided into treatment groups

and fed with tomato leaves contaminated with CuSO4, Kocide and nCu(OH)2 as described

78

in the Methods section. Retardation of growth appeared immediately with the start of

feeding on the contaminated leaves for both the first and second instar larvae (Fig 4.1A).

Ingestion rate was not affected by any of the treatments (Fig 4.S2), suggesting that either

energy assimilation was decreased or the basal metabolic rate was increased 237. Compared

to Kocide and nCu(OH)2, a more significant reduction in biomass was caused by the CuSO4

treatment. And the differential effects progressed steadily with time. Previous studies have

shown that for soluble metal oxide MNMs such as Ag, CuO and ZnO particles, the

corresponding ionic forms of the metal are more toxic and bioavailable to various

organisms and the toxic effects of the MNMs can be largely attributed to ions released by

dissolution 36, 224, 225, 238-243. Given the differences in dissolution of the materials, dissolved

ions, or ions released in vivo, might be the major source of toxicity for these MNMs.

Growth suppression increased with dissolved Cu2+ concentrations in the treatments.

Excessive exposure to Cu2+ have been shown to elicit various detrimental effects in many

species. Cu2+ is highly potent in changing the redox environment of biological systems,

causing damage to macromolecules and disrupting cellular function and metabolism 119, 244,

245.

For all of the treatments, exposure of the first instar larvae resulted in an increase in

mortality compared to control, which was not observed on the second instar larvae (Fig

4.1B). The drastic difference in toxicity on the first and second instar larvae agrees with

previous studies which revealed that the toxicity of MNMs depend on the life stage of the

organism at the time of exposure 80. In addition to being more physiologically developed,

the second instar larvae used for the experiment were healthy individuals surviving the

initial phase of rapidly rising mortality (16.7 % in the untreated caterpillar population, Fig

79

4.1B). In our previous study with M. sexta exposed to Au nanoparticles, the mortality rate

in the control population was even higher and is likely party attributable to stress from

handling the small, fragile hatchlings 149.

Uptake and elimination of CuSO4, Kocide and nCu(OH)2

Analysis of bulk Cu content in the second instar larvae shows the gradual

accumulation of Cu in the dosing phase followed by a rapid decrease at the beginning of

the elimination phase, after which the body burden became asymptotic for an extended

period (Fig 4.2A). This indicates that after the voiding of the gut of residual Cu-containing

food and feces which dominated the initial phase of the elimination period, followed by a

period of Cu homeostasis. This result is consistent with our previous studies on Au and

CeO2 MNMs 149, 150, 221. For both the first and the second instar larvae exposed to CuSO4,

Kocide and nCu(OH)2, the body Cu concentrations at the end of the elimination period

were much higher than the background Cu in the insect despite growth dilution ( Fig 4.2B).

CuSO4 had the highest trophic transfer factors (TTF) followed in order by Kocide

and nCu(OH)2 (Fig 4.3), suggesting that differential accumulation of Cu as a result of these

treatments could be explained by their respective solubilities. Indeed, TTF at the end of the

experiment were correlated with concentrations of soluble Cu2+ in the food (Fig 4.4). It has

been shown that the most bioavailable fraction of soluble metal oxide MNMs is the

dissolved ions 95, 220, 225, 240. TTF for the more vulnerable first instar larvae were

significantly higher than the second instar larvae. All the treatments had AE significantly

higher than control (background Cu in the leaf tissue), in the descending order of CuSO4,

Kocide and nCu(OH)2, and the first instar larvae had higher AE than the second instar

larvae but the difference was not statistically significant (Fig 4.3B). Given that bulk Cu

80

concentrations in the contaminated leaves were ~200 times higher than in the control (Fig

4.S3), the slight difference between the AE of treatment versus control indicates the dietary

uptake of Cu was closely regulated in the larvae. Despite having lower TTF than the second

instar larvae which might be a result of rapid growth at early life stage, the first instar larvae

had drastic reduction of growth and high mortality rate when the dosing started. Insects

employ various strategies to elimination and detoxify heavy metals, such as

compartmentalizing the contaminant in granules thus reducing their reactivity, or

minimizing their bio-reactivity by synthesizing proteins that can bind and sequester metal

ions 246, 247. The sensitivity of the first instar larvae may be due to a lack of effective

detoxification mechanisms for heavy metal ions and/or the energetic demands of early

growth and development.

Biodynamic model of the bioaccumulation of Cu

Although TTF was far below 1 after the initial voiding of the gut content, the lack of

elimination and high AE suggests that significant bioaccumulation could still occur if the

animal were to be continually exposed to the contaminants throughout the entire span of

their larval stage where ingestion rate is high. We used a modified biodynamic model to

simulate the outcome under this scenario 150, 248. We assumed AE for each treatment to be

a constant through time based on the results above showing no significant difference

between AE of the first and second instar larvae. Ingestion rate was also fixed as a constant

based on observation. The fitting on the sigmoid growth curve for the control using a

Gompertz function are shown in Fig 4.5A (see Method section for details). Using the

respective growth model generated for each treatment (Fig 4.S4), TTF was plotted as a

function of time. The body mass of M. sexta larvae rarely reach the asymptotic maximum

81

but molting will continue if the critical weight for pupation is not reached 235, 249. This gives

the larvae with slower growth a longer time of exposure and hence higher maximum TTF

(Fig 4.5B). For simplicity, we truncated the TTF function for the treatments at the different

time when they each reach the critical body mass of 11.3g to show the actual maximum

TTF for each treatment that can be reached 234. TTF for all treatments become significant

under this model, with CuSO4 crossing the line of TTF = 1, suggesting biomagnification

could be possible for caterpillars continually feeding on contaminated leaves. CuSO4 still

has the highest TTF followed by Kocide and nCu(OH)2, but difference between the three

expands with time, with the TTF of CuSO4 being twice as high as that of nCu(OH)2 at their

maxima.

4.5 Conclusion

The trend of bioavailability and toxicity observed for CuSO4, Kocide and nCu(OH)2

correlates with the amount of free Cu2+. It seems that although the complex formula of

Kocide makes its primary properties such as hydrodynamic diameter size and zeta potential

different from those of lab-synthesized nCu(OH)2, the overall impact of these materials on

the insects dietarily exposed to them is still comparable and can probably be predicted

based on solubility. Previous studies showed the correlation between dissolution and the

toxicity of Cu based MNMs 159, 231, 232. However, comparing the ecotoxicity of commercial

MNM products such as Kocide and lab-synthesized counterparts has not been well studied.

Our result provides valuable insights into this area. Nevertheless, future research is needed

to determine to what extent environmental factors such as pH, ionic strength, and the

presence of inorganic ligands and natural organic matter play a role.

82


Table 4.1 Dissolution of the nanomaterials

Dissolution of Kocide and nCu(OH)2 after 4, 24, and 48h as measured in deionized water

at 50mg Cu/L. Values are % dissolved Cu with standard deviations in the parenthesis.

4h 24h 48h

Kocide 26.33 (2.74) 24.72 (3.48) 25.53 (2.91)

nCu(OH)2 5.61 (0.24) 6.23 (0.35) 5.88 (0.22)

83

Figure 4.1 Growth retardation and mortality

Growth retardation A) and % mortality B) in 1st and 2nd instar M. sexta larvae exposed to

CuSO4, Kocide and nCu(OH)2.The 1st instar larvae were more vulnerable to the treatments

than the 2nd instar larvae. Background mortality rate in the control reached 16% at the end

of the experiment, which was not significantly different from the 2nd instar curves.

84

Figure 4.2 Bulk Cu in tissue

A Change of Cu content in the 2nd instar larvae through time. Caterpillars were switched to

clean diet for depuration after day 20, causing a rapid decrease in Cu. Elimination was

negligible after 4 days of feeding on clean food. B Cu concentrations in the body for all

the treatments at the end of depuration period (day 20 for the 1st instar and 30 for the 2nd

instar) were significantly higher than control. Means with the same letter are not

significantly different at Tukey’s test (p < 0.05, n = 12).

85

Figure 4.3 Trophic transfer and assimilation efficiency

A Trophic transfer factors (TTF) at the end of the depuration period (day 20 for the 1st

instar and day 30 for the 2nd instar). No biomagnification was observed. B Assimilation

efficiency (AE) calculated for all the treatment groups. Means with the same letter are not

significantly different at Tukey’s test (p < 0.05, n = 12).

86

Figure 4.4 Correlation between trophic transfer factors and free Cu2+

Trophic transfer factors (TTF) plotted as a function of dissolved Cu2+ in the food for

CuSO4, Kocide and nCu(OH)2.

87

Figure 4.5 Biodynamic model

Biodynamic model for M. sexta larvae exposed CuSO4, Kocide and nCu(OH)2. A) The

sigmoid growth curve from the control population was fitted with a Gompertz function,

with the 95% confidence interval (CI) shown on the graph. B) Trophic transfer factors

(TTF) were modeled for the entire span of the larval stage, which was different for each

treatment and was determined by the growth rate, assuming pupation occurs at the

maximum body mass. Biomagnification is possible under this model. See text for details.

88

Figure 4.6 TEM graphs of nCu(OH)2 and Kocide

Fig 4.S1. Transmission electron micrograph of synthesized nCu(OH)2 (A) and Kocide

3000 (B).

89

Figure 4.7 Ingestion rate

Fig 4.S2. Ingestion rate as a function of day.

90

Figure 4.8 Bulk Cu in the food

Fig 4.S3. Cu concentration in the leaves of the tomato plant (Solanum lycopersicum cv

Micro-Tom) surface-contaminated with CuSO4, Kocide and nCu(OH)2.

91

Figure 4.9 Growth curve fitting

Fig 4.S4. Individual fittings of the growth curves for the treatments. Statistics of the fit as

output from SAS are listed in the table below, b and c are parameters of the Gompertz

function. See Method for details.

Para-

meters

Estimates Standard

Error

95% Confidence

Limits

Skewness

Control b 8.824 0.00325 8.063 10.34 0.0531

c 0.0782 0.00462 0.0732 0.0855 0.0163

1st instar

CuSO4

b 9.461 0.00415 9.043 10.513 0.0434

c 0.0629 0.00326 0.0536 0.0709 0.0363

1st instar

Kocide

b 9.446 0.00313 8.744 10.257 0.0279

c 0.0635 0.00279 0.0564 0.0722 0.0321

1st instar

nCu(OH)2

b 9.508 0.00386 8.576 9.945 0.0365

c 0.0676 0.00203 0.0584 0.0758 0.0263

2nd instar

CuSO4

b 6.773 0.00323 6.239 7.364 0.0204

c 0.0638 0.00442 0.0527 0.0693 0.0217

2nd instar

Kocide

b 6.725 0.00276 6.348 7.228 0.0142

c 0.0626 0.00213 0.0558 0.0746 0.0165

2nd instar

nCu(OH)2

b 7.385 0.00281 6.832 7.929 0.0139

c 0.0694 0.00309 0.0601 0.0796 0.0177

92

CHAPTER 5. CONCLUSIONS AND FUTURE DIRECTIONS

We have conducted three groups of experiments to investigate the impact of physico-

chemical properties of the MNMs on plant uptake and trophic transfer. Firstly, plant uptake

of positively, negatively and neutrally charged CeO2 MNMs was investigated using the

model organism Solanum lycopersicum cv Micro-Tom. Plant growth, Ce concentrations in

tissues and root-shoot translocation rates were functions of surface charge and exposure

concentrations of the MNMs. Results from high-resolution X-ray microscopy suggest that

particles have drastically different tissue distribution patterns across the root and leaf

tissues accompanied by various degrees of biotransformation depending on their surface

charges. They entered through the gaps between epidermal cells being sloughed off during

root growth and penetrated deeper into the interior of the roots (vasculature) via a

combination of apoplastic and symplastic transport. Evidence of endocytosis was found at

the endodermis of the root tip, which appears to lead to dissolution of the CeO2 MNMs,

with subsequent re-precipitation as Ce (III) containing minerals found in root and leaf

tissues.

The impact of surface charge on the uptake and biodistribution of MNMs by the plant

has several important implications. MNMs in the soil become coated with natural organic

matter, a net negative charge is acquired, which might enhance their translocation to the

aerial part of the plant. Moreover, this net negative charge also makes dissolution and

biotransformation during symplastic transport more like for the MNMs. Furthermore, from

the perspective of nanotechnology, the differential tissue and cellular distribution as a

function of particle surface chemistry suggests that targeted, MNM-mediated delivery of

93

agrochemicals to different biological compartments within the plant is possible at both the

tissue and subcellular levels.

Uptake of MNMs by the primary producer means that the upper trophic layers could

also be exposed to these contaminants. In the second experiment, we investigated the

dietary uptake and elimination of the same CeO2 MNMs carrying different surface charges

in a simulated food chain consisting of the tomato plant (Solanum lycopersicum cv Micro-

Tom) and the tobacco hornworm (Manduca sexta). Despite no observed overall


higher trophic transfer factors than those of ionic Ce3+. The uptake-elimination dynamics

were influenced by the surface charge of the MNMs. Positively charged MNMs had higher

trophic transfer factors and assimilation efficiencies but lower elimination rate than neutral

and negatively charged CeO2 MNMs. X-ray microscopy also revealed the impact of

extensive biotransformation on the uptake of these materials by the insect digestive system.

These results provide important evidence that surface properties of MNMs such as

surface charge not only influence plant uptake but also the transfer of MNMs in the

terrestrial food web. The more extensive biotransformation processes experience by the

negatively charged particles, as compared to positively charged ones, likely contribute to

this difference. The net negative charge that is conferred by the coating of natural organic

matter as the MNMs age in the soil will diminish their impact on the upper trophic levels.

In the third experiment, we investigated the trophic transfer of copper from Cu(OH)2

MNMs in the same model food chain. Leaves of tomato plants were surface-contaminated

by copper sulfate (CuSO4), copper hydroxide nanoneedle (nCu(OH)2), and a commercial

pesticide Kocide® 3000, whose active ingredient is Cu(OH)2 nanosheets. These

94

contaminated leaves were provided to M. sexta larvae as food. Differential toxicity on the

caterpillar and trophic transfer of Cu occurred in the order of CuSO4 > Kocide > nCu(OH)2.

By tracking the uptake and elimination dynamics of Cu in the hornworms, we built a

modified biodynamic model to compare the rates of trophic transfer for these treatments

assuming continued exposure during the entire span of the larval stage. Our results

demonstrated that lab-synthesized Cu(OH)2 MNMs and Kocide have comparable

behaviors. Toxicity and accumulation/elimination dynamics was found to be correlated

with the dissolution rate of the materials. Prediction of the biodynamic model implied

possible biomagnification from continuous exposure throughout the life span of the larvae.

This result provides an example as to how well the ecotoxicity of commercial nano-

products can be differ from lab-synthesized MNMs that mimic their active ingredients.

These nanoproducts usually have a complex formula to aid in specific applications.

Although these specifically engineered properties are diverse and not easily replicated by

lab synthesis of the active ingredients, these lab-synthesized surrogates can be nevertheless

used as a simple model to interrogate their behavior in the ecosystem, provided that the

role of key properties, such as solubility, are understood.

Given the simplicity of our experimental models, future efforts are required to

determine whether these observations can be translated to more realistic scenario such as

exposure from contaminated soil. Key environmental players such as pH and or natural

organic matter need to be investigated to provide a more complete view. Overall, this work

provides important insights into the impact of MNMs on the terrestrial ecosystem which is

still poorly understood. It demonstrates properties such as surface charge and chemical

composition as important parameters governing the ecotoxicology of MNMs. Finally, this

95

knowledge will benefit the design MNMs for targeted delivery of chemicals to plants and

insects such as development of nano-agrochemicals for agricultural practice.

96

REFERENCES

1. Auffan, M.; Rose, J.; Bottero, J. Y.; Lowry, G. V.; Jolivet, J. P.; Wiesner, M. R., Towards

a definition of inorganic nanoparticles from an environmental, health and safety perspective. Nat

Nanotechnol 2009, 4, (10), 634-641.

2. Han, G.; Ghosh, P.; Rotello, V. M., Functionalized gold nanoparticles for drug delivery.

Nanomedicine 2007, 2, (1), 113-123.

3. Qian, X.; Peng, X. H.; Ansari, D. O.; Yin-Goen, Q.; Chen, G. Z., In vivo tumor targeting

and spectroscopic detection with surface-enhanced Raman nanoparticle tags. Nature

Biotechnology 2008, 26, (1), 83-90.

4. Mackey, M. A.; Ali, M. R.; Austin, L. A.; Near, R. D.; El-Sayed, M. A., The most

effective gold nanorod size for plasmonic photothermal therapy: theory and in vitro experiments.

The Journal of Physical Chemistry B 2014, 118, (5), 1319–1326.

5. Sajjadi, A. Y.; Suratkar, A. A.; Mitra, K. K.; Grace, M. S., Short-pulse laser-based

system for detection of tumors: Administration of gold nanoparticles enhances contrast. J

Nanotechnol Eng Med 2012, 3, (2), 235-240.

6. Niidome, T.; Yamagata, M.; Okamoto, Y.; Akiyama, Y.; Takahashi, H.; Kawano, T.;

Katayama, Y.; Niidome, Y., PEG-modified gold nanorods with a stealth character for in vivo

applications. Journal of Controlled Release 2006, 114, (3), 343–347.

7. Hainfeld, J. F.; Slatkin, D. N.; Smilowitz, H. M., The use of gold nanoparticles to

enhance radiotherapy in mice. Physics in Medicine and Biology 2004, 49, (18), 309-315.

8. Zhang, Z.; Chen, Z.; Wang, S.; Qu, C.; Chen, L., On-site visual detection of hydrogen

sulfide in air based on enhancing the stability of gold nanoparticles. ACS Applied Materials &

Interfaces 2014, 6, (9), 6300–6307.

9. Okamoto, T.; Yamaguchi, I.; Kobayashi, T., Local plasmon sensor with gold colloid

monolayers deposited upon glass substrates. Opt Lett 2000, 25, (6), 372-376.

10. Xiao, Y.; Patolsky, F.; Katz, E.; Hainfeld, J. F.; Willner, I., Plugging into Enzymes:

nanowiring of redox enzymes by a gold nanoparticle. 299 2003, 5614, (1877–1881).

11. Kim, B. W.; Voitl, T.; Rodriguez-Rivera, J. G.; Dumesic, J. A., Powering fuel cells with

CO via aqueous polyoxometalates and gold catalyst. Sience 2004, 305, (5688), 1280-1283.

12. Jiang, Z.-J.; Liu, C.-Y.; Sun, L.-W., Catalytic properties of silver nanoparticles supported

on silica spheres. The Journal of Physical Chemistry B 2005, 109, (5), 1730–1735.

13. Ameen, K. B.; Rajasekar, K.; Rajasekharan, T., Silver nanoparticles in mesoporous

aerogel exhibiting selective catalytic oxidation of benzene in CO2 free air. Catalysis Letters

2007, 119, (3-4), 289–295.

14. Liu, J.-H.; Wang, A.-Q.; Chi, Y.-S.; Lin, H.-P.; Mou, C.-Y., Synergistic effect in an

Au−Ag alloy nanocatalyst: CO oxidation. The Journal of Physical Chemistry B 2005, 109, (1),

40-43.

15. Mukherjee, S.; Chowdhury, D.; Kotcherlakota, R.; Parta, S.; Bhadra, M. P.; Sreedhar, B.;

Patra, C. R., Potential theranostic application of biosynthesized silver nanoparticles. Theranostics

2014, 4, (3), 316–335.

16. Hong, R.; Han, G.; Fernández, J. M.; Kim, B.-j.; Forbes, N. S.; Rotello, V. M.,

Glutathione mediated delivery and release using monolayer protected nanoparticle carriers. J Am

Chem Soc 2006, 128, (4), 1078–1079.

17. Shahverdi, A. R.; Fakhimi, A.; Shahverdi, H. Q.; Minaian, S., Synthesis and effect of

silver nanoparticles on the antibacterial activity of different antibiotics against Staphylococcus

aureus and Escherichia coli. Nanomedicine 2007, 3, (2), 168-171.

97

18. Rigo, C.; Ferroni, L.; Tocco, I.; Roman, M.; Munivrana, I., Active silver nanoparticles for

wound healing. Int J Mol Sci 2013, 14, (3), 4817–4840.

19. Jain, P.; Pradeep, T., Potential of silver nanoparticle-coated polyurethane foam as an

antibacterial water filter. Biotechnol Bioeng 2005, 90, (1), 59–63.

20. Cheon, J. M.; Lee, J. H.; Song, Y.; Kim, J., Synthesis of Ag nanoparticles using an

electrolysis method and application to inkjet printing. Colloids and Surfaces A: Physicochemical

and Engineering Aspects 2011, 389, (1-3), 175-179.

21. Gulson, B.; McCall, M.; Korsch, M.; Gomez, L.; Casey, P.; Oytam, Y.; Taylor, A.;

McCulloch, M.; Trotter, J.; Kinsley, L.; Greenoak, G., Small amounts of zinc from zinc oxide

particles in sunscreens applied outdoors are absorbed through human skin. Toxicological Sciences

2010, 118, (1), 140–149.

22. Burnett, M. E.; Wang, S. Q., Current sunscreen controversies: A critical review.

Photodermatology, Photoimmunology & Photomedicine 2011, 27, (2), 58-67.

23. Banoee, M.; Seif, S.; Nazari, Z. E.; Jarafi-Fesharaki, P.; Shahverdi, H. R.; Moballegh, A.;

Moghaddam, K. M.; Shahverdi, A. R., ZnO nanoparticles enhanced antibacterial activity of

ciprofloxacin against Staphylococcus aureus and Escherichia coli. J Biomed Mater Res B 2010,

93, (2), 557-561.

24. Wang, H. T.; Kang, B. S.; Ren, F.; Tien, L. C.; Sadik, P. W.; Norton, D. P.; Pearton, S. J.;

Lin, J., Hydrogen-selective sensing at room temperature with ZnO nanorods. Applied Physics

Letters 2005, 86, (24), 24350.

25. Look, D., Recent advances in ZnO materials and devices. Materials Science and

Engineering B 2001, 80, 383–387.

26. Kessler, R., Engineered nanoparticles in consumer products: Understanding a new

ingredient. Environmental Health Perspectives 2011, 119, (3), 120-125.

27. Cross, S. E.; Innes, B.; Roberts, M. S.; Tzuzuki, T.; Robertson, T. A.; McCormick, P.,

Human skin penetration of sunscreen nanoparticles: In-vitro assessment of a novel micronized

zinc oxide formulation. Skin Pharmacol Physiol 2007, 20, (148-154).

28. Kaegi, R.; Ulrich, A.; Sinnet, B.; Vonbank, R.; Wichser, A.; Zuleeg, S.; Simmler, H.;

Brunner, S.; Vonmont, H.; Burkhardt, M.; Boller, M., Synthetic TiO2 nanoparticle emission from

exterior facades into the aquatic environment. Environ Pollut 2008, 156, (2), 233-239.

29. Luo, X.; Morrin, A.; Killard, A. J.; Smyth, M. R., Application of nanoparticles

inelectrochemical sensors and biosensors. Electroanalysis 2006, 18, 319-326.

30. Yetisen, A. K.; Montelongo, Y.; Vasconcellos, F. D. C.; Martinez-Hurtado, J.; Neupane,

S.; Butt, H.; Qasim, M. M.; Blyth, J.; Burling, K.; Carmody, J. B., Reusable, robust, and accurate

laser-generated photonic nanosensor. Nano Letters 2014, 14, 3587-3593.

31. Ibupoto, Z.; Khun, K.; Beni, V.; Liu, X.; Willander, M., Synthesis of novel CuO

nanosheets and their non-enzymatic glucose sensing applications. Sensors 2013, 13, 7926–7938.

32. Zen, J.; Hsu, C.; Kumar, A. S.; Lyuu, H.; Lin, K., Amino acid analysis using disposable

copper nanoparticle plated electrodes. Analyst 2004, 129, 841.

33. Gawande, M. B.; Goswami, A.; Felpin, F.-X.; Asefa, T.; Huang, X.; Silva, R.; Zou, X.;

Zboril, R.; Varma, R. S., Cu and Cu-based nanoparticles: Synthesis and applications in catalysis.

Chem Rev 2016, 116, (6), 3722-3811.

34. Giannousi, K.; Avramidis, I.; Dendrinou-Samara, C., Synthesis, characterization and

evaluation of copper based nanoparticles as agrochemicals against Phytophthora infestans. RSC

Adv 2013, 3, 21743-21751.

35. Oustriere, N.; Marchand, L.; Roulet, E.; Mench, M., Rhizofiltration of a Bordeaux

mixture effluent in pilot-scale constructed wetland using Arundo donax L. coupled with potential

Cu-ecocatalyst production. Ecol Eng 2017, 105, 296-305.

36. Zuverza-Mena, N.; Medina-Velo, I. A.; Barrios, A. C.; Tan, W.; Peralta-Videa, J. R.;

Gardea-Torresdey, J. L., Copper nanoparticles/compounds impact agronomic and physiological

98

parameters in cilantro (Coriandrum sativum). Environmental science. Processes & impacts 2015,

17, (10), 1783-1793.

37. Celardo, I.; Pedersen, J. Z.; Traversa, E.; Ghibelli, L., Pharmacological potential of

cerium oxide nanoparticles. Nanoscale 2011, 3, (4), 1411-1420.

38. Trovarelli, A.; de Leitenburg, C.; Boaro, M.; Dolcetti, G., The utilization of ceria in

industrial catalysis. Catal Today 1999, 50, (2), 353−367.

39. Yu, T. P., Y. I.; Kang, M. C.; Joo, J.; Park, J. K.; Won, H. Y.; Kim, J. J.; Hyeon, T. ,

Large-scale synthesis of water dispersible ceria nanocrystals by a simple sol-gel process and their

use as a chemical mechanical planarization slurry. Eur J Inorg Chem 2008, 6, 855−858.

40. Milt, V. G.; Querini, C. A.; Miro, E. E.; Ulla, M. A., Abatement of diesel exhaust

pollutants: NOx adsorption on Co, Ba, K/CeO2 catalysts. Journal of Catalysis 2003, 220, 424-

432.

41. Zanello, L. P.; Zhao, B.; Hu, H.; Haddon, R. C., Bone cell proliferation on carbon

nanotubes. Nano Lett 2006, 6, (3), 562-567.

42. Tan, C.; Tan, K.; Ong, Y.; Mohamed, A. R.; Zein, S. H. F.; S., T., Energy and

environmental applications of carbon nanotubes. Environ Chem Lett 2012, 10, (3), 265–273.

43. Belkin, A.; Hubler, A.; Bezryadin, A., Self-assembled wiggling nano-structures and the

principle of maximum entropy production. Sci Rep 2015, 5, 8323.

44. Dillon, A. C.; Jones, K. M.; Bekkedahl, T. A.; Kiang, C. H.; Bethune, D. S.; Heben, M.

J., Storage of hydrogen in single-walled carbon nanotubes. Nature Biotechnology 1997, 386,

(6623), 377-379.

45. Snow, E. S.; Perkins, F. K.; Houser, E. J.; Badescu, S. C.; Reinecke, T. L., Chemical

detection with a single-walled carbon nanotube capacitor. Science 2005, 307, (5717), 1942-1945.

46. Handy, R. D.; Owen, R.; Valsami-Jones, E., The ecotoxicology of nanoparticles and

nanomaterials: current status, knowledge gaps, challenges, and future needs. Ecotoxicology 2008,

17, (5), 315-325.

47. Batley, G. E.; Kirby, J. K.; McLaughlin, M. J., Fate and risks of nanomaterials in aquatic

and terrestrial environments. Acc Chem Res 2013, 46, (3), 854-862.

48. Lowry, G. V.; Gregory, K. B.; Apte, S. C.; Lead, J. R., Transformations of nanomaterials

in the environment. Environ Sci Technol 2012, 46, (13), 6893-6899.

49. Gao, J.; Youn, S.; Hovsepyan, A.; Llaneza, V. L.; Wang, Y.; Bitton, G.; Bonzongo, J. J.,

Dispersion and toxicity of selected manufactured nanomaterials in natural river water samples:

effects of water chemical composition. Environ Sci Technol 2009, 43, (1), 3322-3328.

50. Espinasse, B.; Hotze, E. M.; Wiesner, M. R., Transport and retention of colloidal

aggregates of C60 in porous media: effects of organic macromolecules, ionic composition,and

preparation method. Environ Sci Technol 2007, 41, (2), 7396-7402.

51. Smith, C. J.; Shaw, B. J.; Handy, R. D., Toxicity of single walled carbon nanotubes on

rainbow trout, (Oncorhynchus mykiss): respiratory toxicity, organ pathologies, and other

physiological effects. Aquat Toxicol 2007, 82, (3), 94-109.

52. Hotze, E. M.; Phenrat, T.; Lowry, G. V., Nanoparticle aggregation: challenges to

understanding transport and reactivity in the environment. Journal of Environmental Quality

2010, 39, (6), 1909-1924.

53. Unrine, J. M.; Tsyusko, O. V.; Hunyadi, S. E.; Judy, J. D.; Bertsch, P. M., Effects of

particle size on chemical speciation and bioavailability of copper to earthworms (Eisenia fetida)

exposed to copper nanoparticles. J Environ Qual 2010, 39, (6), 1942-1953.

54. Ma, R.; Levard, C.; Marinakos, S.; Chen, Y.; Liu, J.; Brown, G. E., Jr.; Lowry, G. V.,

Size controlled dissolution of silver nanoparticles. Environ Sci Technol 2012, 46, (2), 752– 759.

55. Liu, J.; Hurt, R., Ion release kinetics and particle persistence in aqueous nano-silver

colloids. Environ Sci Technol 2010, 44, (2), 2169-2175.

99

56. Jitao Lv; Shuzhen Zhang; Lei Luo; Wei Han; Jing Zhang; Ke Yang; Christie∥, P.,

Dissolution and microstructural transformation of ZnO nanoparticles under the influence of

phosphate. Environ Sci Technol 2012, 46, (13), 7215-7221.

57. Levard, C.; Reinsch, B. C.; Michel, F. M.; Oumahi, C.; Lowry, G. V.; Brown, G. E.,

Sulfidation processes of PVP-coated silver nanoparticles in aqueous solution: Impact on

dissolution rate. Environ Sci Technol 2011, 45, (12), 5260-5266.

58. Mudunkotuwa, I. A.; Pettibone, J. M.; Grassian, V. H., Environmental implications of

nanoparticle aging in the processing and fate of copper-based nanomaterials. Environ Sci Technol

2012, 46, (13), 7001-7010.

59. Lopez-Moreno, M. L.; de la Rosa, G.; Hernandez-Viezcas, J. A.; Castillo-Michel, H.;

Botez, C. E.; Peralta-Videa, J. R.; Gardea-Torresdey, J. L., Evidence of the differential

biotransformation and genotoxicity of ZnO and CeO2 nanoparticles on soybean (Glycine max)

plants. Environ Sci Technol 2010, 44, (19), 7315-20.

60. Zhang, H.; He, X.; Zhang, Z.; Zhang, P.; Li, Y.; Ma, Y.; Kuang, Y.; Zhao, Y.; Chai, Z.,

Nano-CeO2 exhibits adverse effects at environmental relevant concentrations. Environ Sci

Technol 2011, 45, 3725-3730.

61. Krauskopf, K., The solubility of gold. Econ Geol 1951, 46, (8), 858-870.

62. Schmidt, J.; Vogelsberger, W., Aqueous long-term solubility of titania nanoparticles and

titanium (IV) hydrolysis in a sodium chloride system studied by adsorptive stripping

voltammetry. Journal of Solution Chemistry 2009, 38, (10), 1267-1282.

63. Cornelis, G.; Ryan, B.; McLaughlin, M. J.; Kirby, J. K.; Beak, D.; Chittleborough, D.,

Solubility and batch retention of CeO2 nanoparticles in soils. Environ Sci Technol 2011, 45, (7),

2777-2782.

64. Liu, J.; Pennell, K. G.; Hurt, R. H., Kinetics and mechanisms of nanosilver

oxysulfidation. Environ Sci Technol 2011, 45, (2), 7345-7353.

65. Barton, L. E.; Auffan, M.; Bertrand, M.; Barakat, M., Transformation of pristine and

citrate-functionalized CeO2 nanoparticles in a laboratory-scale activated sludge reactor. Environ

Sci Technol 2014, 48, (13), 7289-7296.

66. Lv, J.; Zhang, S.; Luo, L.; Han, W.; Zhang, J.; Yang, K.; Christie, P., Dissolution and

microstructural transformation of ZnO nanoparticles under the influence of phosphate. Environ

Sci Technol 2012, 46, (13), 7215-7221.

67. Yin, Y.; Yang, X.; Zhou, X.; Wang, W.; Yu, S.; Liu, J.; Jiang, G., Water chemistry

controlled aggregation and photo-transformation of silver nanoparticles in environmental waters.

J Environ Sci 2015, 34, 116-125.

68. Carpita, N. C.; Gibeaut, D. M., Structural models of primary cell walls in flowering

plants: consistency of molecular-structure with the physical properties of the walls during growth.

Plant J 1993, 3, 1-30.

69. Chen, R.; Ratnikova, T. A.; Stone, M. B.; Lin, S.; Lard, M.; Huang, G.; Hudson, J. S.;

Ke, P. C., Differential uptake of carbon nanoparticles by plant and mammalian cells. Small 2010,

6, (5), 612-617.

70. Lin, C.; Fugetsu, B.; Su, Y.; Watari, F., Studies on toxicity of multi-walled carbon

nanotubes on Arabidopsis T87 suspension cells. J Hazard Mater 2009, 170, 578-583.

71. Eichert, T.; Goldbach, H. E., Equivalent pore radii of hydrophilic foliar uptake routes in

stomatous and astomatous leaf surfacesefurther evidence for a stomatal pathway. Physiol Plant

2008, 132, (4), 491-502.

72. Peterson, C.; Enstone, D., A survey of angiosperm species to detect hypodermal

Casparian Bands. I. Roots with a uniseriate hypodermis and epidermis. Botannical Journal of the

Linnean Society 1990, 103, 93-112.

73. Peterson, C.; Enstone, D., Functions of passage cells in the endodermis and exodermis of

roots. Physiologia Plantarum 1996, 97, 592-598.

100

74. Zhao, L.; Peralta-Videa, J. R.; Ren, M.; Varela-Ramirez, A.; Li, C.; Hernandez-Viezcas,

J. A., Transport of Zn in a sandy loam soil treated with ZnO NPs and uptake by corn plants:

Electron microprobe and confocal microscopy studies. Chem Eng J 2012, 184, 1-8.

75. Etxeberria, E.; Gonzalez, P.; Pozueta, J., Evidence for two endocytic transport pathways

in plant cells. Plant Sci 2009, 177, (4), 341-348.

76. Onelli, E.; Prescianotto-Baschong, C.; Caccianiga, M.; Moscatelli, A., Clathrin-

dependent and independent endocytic pathways in tobacco protoplasts revealed by labelling with

charged nanogold. J Exp Bot 2008, 59, (11), 3051-3068.

77. Etxeberria, E.; Gonzalez, P.; Baroja-Fernandez, E.; Romero, J. P., Fluid phase endocytic

uptake of artificial nano–spheres and fluorescent quantum dots by sycamore cultured cells:

Evidence for the distribution of solutes to different intracellular compartments. Plant Signaling

Behav 2006, 1, 196-200.

78. Serag, M. F.; Kaji, N.; Gaillard, C.; Okamoto, Y.; Terasaka, K.; Jabasini, M.; Tokeshi,

M.; Mizukami, H.; Bianco, A.; Baba, Y., Trafficking and subcellular localization of multiwalled

carbon nanotubes in plant cells. ACS Nano 2010, 5, (1), 493-499.

79. Santos, A. R.; Miguel, A. S.; Tomaz, L.; Malh o, R.;

Maycock, C.; Patto, M. C. V.; Fevereiro, P.; Oliva, A., The impact of CdSe/ZnS quantum dots in

cells of Medicago sativa in suspension culture. J Nanobiotec 2010, 8, (1), 1-11.

80. Geisler-Lee, J.; Wang, Q.; Yao, Y.; Zhang, W.; Geisler, M.; Li, K.; Huang, Y.; Chen, Y.;

Kolmakov, A.; Ma, X., Phytotoxicity, accumulation and transport of silver nanoparticles by

Arabidopsis thaliana. Nanotoxicology 2013, 7, (3), 323-337.

81. Zhai, G.; Walters, K. S.; Peate, D. W.; Alvarez, P. J. J.; Schnoor, J. L., Transport of gold

nanoparticles through plasmodesmata and precipitation of gold ions in woody poplar. Environ Sci

Technol Lett 2014, 1, (2), 146-151.

82. Cifuentes, Z.; Custardoy, L.; de la Fuente, J. M.; Marquina, C.; Ricardo Ibarra, M.;

Rubiales, D., Absorption and translocation to the aerial part of magnetic carbon–coated

nanoparticles through the root of different crop plants. J Nanobiotechnol 2010, 8, (26), 1-8.

83. Asli, S.; Neumann, P. M., Colloidal suspensions of clay or titanium dioxide nanoparticles

can inhibit leaf growth and transpiration via physical effects on root water transport. Plant Cell

Environ 2009, 32, (5), 577-584.

84. Corredor, E.; Testillano, P. S.; Coronado, M. J.; Gonzalez-Melendi, P.; Fernandez-

Pacheco, R.; Marquina, C.; Ibarra, M. R.; de la Fuente, J. M.; Rubiales, D.; Perez-De-Luque, A.;

Risueno, M. C., Nanoparticle penetration and transport in living pumpkin plants: in situ

subcellular identification. Bmc Plant Biology 2009, 9, (45), 1-11.

85. Wang, Z.; Xie, X.; Zhao, J.; Liu, X.; Feng, W.; White, J. C.; Xing, B., Xylem-and

phloem-based transport of CuO nanoparticles in maize (Zea mays L.). Environ Sci Technol 2012,

46, (8), 4434-4441.

86. Zhang, Z. Y.; He, X.; Zhang, H. F.; Ma, Y. H.; Zhang, P.; Ding, Y. Y., Uptake and

distribution of ceria nanoparticles in cucumber plants. Metallomics 2011, 3, (8), 816-822.

87. Larue, C.; Castillo-Michel, H.; Sobanska, S.; Cecillon, L.; Bureau, S.; Barthes, V., Foliar

exposure of the crop Lactuca sativa to silver nanoparticles: Evidence for internalization and

changes in Ag speciation. J Hazard Mater 2014, 264, 98-106.

88. Wang, Q.; Ma, X.; Zhang, W.; Pei, H.; Chen, Y., The impact of cerium oxide

nanoparticles on tomato (Solanum lycopersicum L.) and its implications for food safety.

Metallomics 2012, 4, (10), 1105-1112.

89. Hong, J.; Peralta-Videa, J. R.; Rico, C.; Sahi, S.; Viveros, M. N.; Bartonjo, J., Evidence

of translocation and physiological impacts of foliar applied CeO2 nanoparticles on cucumber

(Cucumis sativus) plants. Environ Sci Technol 2014, 48, (8), 4376-4385.

101

90. Raliya, R.; Nair, R.; Chavalmane, S.; Wang, W. N.; Biswas, P., Mechanistic evaluation

of translocation and physiological impact of titanium dioxide and zinc oxide nanoparticles on the

tomato (Solanum lycopersicum L.) plant. Metallomics 2015, 7, (12), 1584-1594.

91. Lin, S.; Reppert, J.; Hu, Q.; Hudson, J. S.; Reid, M. L.; Ratnikova, T. A.; Rao, A. M.;

Luo, H.; Ke, P. C., Uptake, translocation, and transmission of carbon nanomaterials in rice plants.

Small 2009, 5, 1128-1132.

92. Rico, C. M.; Majumdar, S.; Duarte-Gardea, M.; Peralta-Videa, J. R.; Gardea-Torresdey,

J. L., Interaction of nanoparticles with edible plants and their possible implications in the food

chain. J Agric Food Chem 2011, 59, (8), 3485-3498.

93. Miralles, P.; Church, T. L.; Harris, A. T., Toxicity, uptake, and translocation of

engineered nanomaterials in vascular plants. Environ Sci Technol 2012, 46, (17), 9224-9239.

94. El-Temsah, Y. S.; Joner, E. J., Impact of Fe and Ag nanoparticles on seed germination

and differences in bioavailability during exposure in aqueous suspension and soil. Environ

Toxicol 2012, 27, (1), 42-49.

95. Kaveh, R.; Li, Y. S.; Ranjbar, S.; Tehrani, R.; Brueck, C. L.; Van Aken, B., Changes in

Arabidopsis thaliana gene expression in response to silver nanoparticles and silver ions. Environ

Sci Technol 2013, 47, (18), 10637-10644.

96. Aken Van, B., Gene expression changes in plants and microorganisms exposed to

nanoparticles. Curr Opin Biotech 2015, 33, 206-219.

97. Lee, C. W.; Mahendra, S.; Zodrow, K.; Li, D.; Tsai, Y. C.; Braam, J.; Alvarez, P. J.,

Developmental phytotoxicity of metal oxide nanoparticles to Arabidopsis thaliana. Environ

Toxicol Chem 2010, 29, (2), 669-675.

98. Stampoulis, D.; Sinha, S. K.; White, J. C., Assay-dependent phytotoxicity of

nanoparticles to plants. Environ Sci Technol 2009, 43, (24), 9473-9479.

99. Priester, J. H.; Ge, Y.; Mielke, R. E.; Horst, A. M.; Moritz, S. C.; Espinosa, K.; Gelb, J.;

Walker, S. L.; Nisbet, R. M.; An, Y. J.; Schimel, J. P., Soybean susceptibility to manufactured

nanoparticles with evidence for food quality and soil fertility interruption. Proc Natl Acad Sci

Lett 2012, 109, (37), 2451-2456.

100. Ma, Y.; Kuang, L.; He, X.; Bai, W.; Ding, Y.; Zhang, Z.; Zhao, Y.; Chai, Z., Effects of

rare earth oxide nanoparticles on root elongation of plants. Chemosphere 2010, 78, 273-279.

101. Fan, R.; Huang, Y. C.; Grusak, M. A.; Huang, C. P.; Sherrier, D. J., Effects of nano-TiO2

on the agronomically-relevant Rhizobiumelegume symbiosis. Sci Total Environ 2014, 466, 503-

512.

102. Nair, P. M. G.; Chung, I. M., Physiological and molecular level effects of silver

nanoparticles exposure in rice (Oryza sativa L.) seedlings. Chemosphere 2014, 112, 105-113.

103. Hawthorne, J.; Musante, C.; Sinha, S. K.; White, J. C., Accumulation and phytotoxicity

of engineered nanoparticles to Cucurbita pepo. Int J Phytoremediat 2012, 14, (4), 429-442.

104. Pokhrel, L. R.; Dubey, B., Evaluation of developmental responses of two crop plants

exposed to silver and zinc oxide nanoparticles. Sci Total Environ 2013, 452, 321-332.

105. Zhao, L.; Sun, Y.; Hernandez-Viezcas, J. A.; Servin, A. D.; Hong, J.; Niu, G., Influence

of CeO2 and ZnO nanoparticles on cucumber physiological markers and bioaccumulation of Ce

and Zn: a life cycle study. J Agric Food Chem 2013, 61, 11945-11951.

106. Ma, X.; Geiser-Lee, J.; Deng, Y.; Kolmakov, A., Interactions between engineered

nanoparticles (ENPs) and plants: phytotoxicity, uptake and accumulation. Sci Total Environ 2010,

408, (16), 3053-3061.

107. Musante, C.; White, J. C., 2012. . Environ. Toxicol. 27 (9),; 510e517., Toxicity of silver

and copper to Cucurbita pepo: differential effects of nano and bulk-size particles. Environ Toxicol

2012, 27, (9), 510-517.

102

108. Wang, P.; Menzies, N. W.; Lombi, E.; McKenna, B. A.; Johannessen, B.; Glover, C. J.;

Kappen, P.; Kopittke, P. M., Fate of ZnO nanoparticles in soils and cowpea (Vignaunguiculata).

Environ Sci Technol 2013, 47, (23), 13822-13830.

109. Mukherjee, A.; Peralta-Videa, J. R.; Bandyopadhyay, S.; Rico, C. M.; Zhao, L.; Gardea-

Torresdey, J. L., Physiological effects of nanoparticulate ZnO in green peas (Pisumsativum L.)

cultivated in soil. Metallomics 2014, 6, (1), 132-138.

110. Ma, C.; Chhikara, S.; Xing, B.; Musante, C.; White, J. C.; Dhankher, O. P., Physiological

and molecular response of Arabidopsis thaliana (L.) to nanoparticle cerium and indium oxide

exposure. ACS Sus Chem Engi 2013, 1, (7), 768-778.

111. Servin, A. D.; Morales, M. I.; Castillo-Michel, H.; Hernandez-Viezcas, J. A.; Munoz, B.;

Zhao, L.; Nunez, J. E.; Peralta-Videa, J. R.; Gardea-Torresdey, J. L., Synchrotron verification of

TiO2 accumulation in cucumber fruit: a possible pathway of TiO2 nanoparticle transfer from soil

into the food chain. Environ Sci Technol 2013, 47, (20), 11592-11598.

112. Jacob, D. L.; Borchardt, J. D.; Navaratnam, L.; Otte, M. L.; Bezbaruah, A. N., Uptake

and translocation of Ti from nanoparticles in crops and wetland plants. Int J Phytoremediat 2013,

15, (2), 142-153.

113. Rico, C. M.; Peralta-Videa, J. R.; Gardea-Torresdey, J. L., Chemistry, biochemistry of

nanoparticles, and their role in antioxidant defense system in plants. In Nanotechnology and Plant

Sciences, Springer International Publishing: 2015; pp 1-17.

114. Tan, X.; Lin, C.; Fugetsu, B., Studies on toxicity of multi-walled carbon nanotubes on

suspension rice cells. Carbon 2009, 47, 3479-3487.

115. Lin, D.; Xing, B., Phytotoxicity of nanoparticles: inhibition of seed germination and root

growth. Environ Pollut 2007, 150, (2), 243-250.

116. Shen, C. X.; Zhang, Q. F.; Li, J.; Bi, F. C.; Yao, N., Induction of programmed cell death

in Arabidopsis and rice by single-wall carbon nanotubes. Am J Bot 2010, 97, (10), 1602-1609.

117. Nair, R.; Varghese, S. H.; Nair, B. G.; Maekawa, T.; Yoshida, Y.; Kumar, D. S.,

Nanoparticulate material delivery to plants. Plant Science 2010, 179, (3), 154-163.

118. Rico, C. M.; Hong, J.; Morales, M. I.; Zho, L.; Barrios, A. C.; Zhang, J. Y.; Peralta-

Videa, J. R.; Gardea-Torresdey, J. L., Effect of cerium oxide nanoparticles on rice: A study

involving the antioxidant defense system and in vivo fluorescence imaging. Environ Sci Technol

2013, 47, (11), 5635-5642.

119. Atha, D. H.; Wang, H.; Petersen, E. J.; Cleveland, D.; Holbrook, R. D.; Jaruga, P.;

Dizdaroglu, M.; Xing, B.; Nelson, B. C., Copper oxide nanoparticle mediated DNA damage in

terrestrial plant models. Environ Sci Technol 2012, 46, (3), 1819-1827.

120. Faisal, M.; Saquib, Q.; Alatar, A. A.; Al-Khedhairy, A. A.; Hegazy, A. K.; Musarrat, J.,

Phytotoxic hazards of NiO-nanoparticles in tomato: a study on mechanism of cell death. J Hazard

Mater 2013, 250, 318-332.

121. Navarro, D. A.; Bisson, M. A.; Aga, D. S., Investigating uptake of water-dispersible

CdSe/ZnS quantum dot nanoparticles by Arabidopsis thaliana plants. J Hazard Materials 2012,

211, 427-435.

122. Foltete, A. S.; Masfaraud, J. F.; Bigorgne, E.; Nahmani, J.; Chaurand, P.; Botta, C.;

Labille, J.; Rose, J.; Ferard, J. F.; Cotelle, S., Environmental impact of sunscreen nanomaterials:

ecotoxicity and genotoxicity of altered TiO2 nanocomposites on Vicia faba. Environ Pollut 2011,

159, (10), 2515-2522.

123. Song, G. L.; Gao, Y.; Wu, H.; Hou, W.; Zhang, C.; Ma, H., Physiological effect of

anatase TiO2 nanoparticles on Lemna minor. Environ Toxicol Chem 2012, 31, (9), 2147-2152.

124. Zhao, L. J.; Peralta-Videa, J. R.; Varela-Ramirez, A.; Castillo-Michel, H.; Li, C. Q.;

Zhang, J. Y.; Aguilera, R. J.; Keller, A. A.; Gardea-Torresdey, J. L., Effect of surface coating and

organic matter on the uptake of CeO2 NPs by corn plants grown in soil: Insight into the uptake

mechanism. J Hazard Mater 2012, 225, 131−138.

103

125. Schwabe, F.; Schulin, R.; Limbach, L. K.; Stark, W.; Burge, D.; Nowack, B., Influence of

two types of organic matter on interaction of CeO2 nanoparticles with plants in hydroponic

culture. Chemosphere 2013, 91, 512−520.

126. Zhang, P.; Ma, Y.; Zhang, Z.; He, X.; Zhang, J. Y.; Guo, Z.; Tai, R.; Zhao, Y.; Chai, Z.,

Biotransformation of ceria nanoparticles in cucumber plants. ACS Nano 2012, 6, (11), 9943-9950.

127. Hernandez-Viezcas, J. A.; Castillo-Michel, H.; Andrews, J. C.; Cotte, M.; Rico, C.;

Peralta-Videa, J. R.; Ge, Y.; Priester, J. H.; Holden, P. A.; Gardea-Torresdey, J. L., In situ

synchrotron X-ray fluorescence mapping and speciation of CeO2 and ZnO nanoparticles in soil

cultivated soybean (glycine max). ACS Nano 2013, 7, (2), 1415−1423.

128. Dimpka, C. O.; Latta, D. E.; McLean, J. E.; Britt, D. W.; Boyanov, M. I.; Anderson, A.

J., Fate of CuO and ZnO nano- and microparticles in the plant environment. Environ Sci Technol

2013, 47, 4734−4742.

129. Du, W. C.; Sun, Y. Y.; Ji, R.; Zhu, J. G.; Wu, J. C.; Guo, H. Y., TiO2 and ZnO

nanoparticles negatively affect wheat growth and soil enzyme activities in agricultural soil. J

Environ Monit 2011, 13, (4), 822−828.

130. Hernandez-Viezcas, J. A.; Castillo-Michel, H.; Servin, A. D.; Peralta-Videa, J. R.;

Gardea-Torresdey, J. L., Spectroscopic verification of zinc absorption and distribution in the

desert plant Prosopis julif loravelutina (velvet mesquite) treated with ZnO nanoparticles. Chem

Eng J 2011, 170, 346−352.

131. Larue, C.; Laurette, J.; Herlin-Boime, N.; Khodja, H.; Fayard, B.; Flank, A. M.; Brisset,

F.; Carriere, M., Accumulation, translocation and impact of TiO2 nanoparticles in wheat

(Triticum aestivum spp.): Influence of diameter and crystal phase. Sci Total Environ 2012, 431,

197−208.

132. Larue, C.; Veronesi, G.; Flank, A. M.; Surble, S.; Herlin-Boime, N.; Carriere, M.,

Comparative uptake and impact of TiO2 nanoparticles in wheat and rapeseed. J Toxicol Environ

Health Part A 2012, 75, 722−734.

133. Servin, A. D.; Castillo-Michel, H.; Hernandez-Viezcas, J. A.; Diaz, B. C.; Peralta-Videa,

J. R.; Gardea-Torresdey, J. L., Synchrotron micro-XRF and micro-XANES confirmation of the

uptake and translocation of TiO2 nanoparticles in cucumber (Cucumis sativus) plants. Environ

Sci Technol 2012, 46, 7637−7643.

134. Yin, L.; Cheng, Y.; Espinasse, B.; Colman, B. P.; Auffan, M.; Wiesner, M.; Rose, J.; Liu,

J.; Bernhardt, E. S., More than ions: The effects of silver nanoparticles on Lolium multiflorum.

Environ Sci Technol 2011, 45, 2360−2367.

135. Dimpka, C. O.; McLean, J. E.; Martineau, N.; Britt, D. W.; Haverkamp, R.; Anderson, A.

J., Silver nanoparticles disrupt wheat (Triticum aestivum L.) growth in a sand matrix. Environ Sci

Technol 2013, 47, 1082−1090.

136. Colman, B. P.; Arnaout, C. L.; Anciaux, S.; Gunsch, C. K.; Hochella, M. F.; Kim, B.;

Lowry, G. V.; McGill, B. M.; Reinsch, B. C.; Richardson, C. J.; Unrine, J. M.; Wright, J. P.; Yin,

L.; Bernhardt, E. S., Low concentrations of silver nanoparticles in biosolids cause adverse

ecosystem responses under realistic field scenario. PLoS One 2013, 8, (2), e57189.

137. Kole, C.; Kole, P.; Randunu, K. M.; Choudhary, P.; Podila, F.; Ke, P. C.; Rao, A. M.;

Marcus, R. K., 13, 37., Nanobiotechnology can boost crop production and quality: First evidence

from increased plant biomass, fruit yield and phytomedicine content in bitter melon (Momordica

charantia). BMC Biotechnol 2013, 13, (37).

138. Khodakovskaya, M. V.; Kim, B. S.; Kim, J. N.; Alimohammadi, M.; Dervishi, E.;

Mustafa, T.; Cernigla, C. E., Carbon nanotubes as plant growth regulators: Effects on tomato

growth, reproductive system, and soil microbial community. Small 2013, 9, 115−123.

139. De La Torre-Roche, R.; Hawthorne, J.; Deng, Y.; Xing, B.; Cai, W.; Newman, L. A.;

Wang, C.; Ma, X.; White, J. C., Fullereneenhanced accumulation of p,p′-DDE in agricultural crop

species. Environ Sci Technol 2012, 46, 9315−9323.

104

140. Holbrook, R. D.; Murphy, K. E.; Morrow, J. B.; Cole, K. D., Trophic transfer of

nanoparticles in a simplified invertebrate food web. Nat Nanotechnol 2008, 3, (6), 352 - 355.

141. Zhu, X.; Wang, J.; Zhang, X.; Chang, Y.; Chen, Y., Trophic transfer of TiO2

nanoparticles from daphnia to zebrafish in a simplified freshwater food chain. Chemosphere

2010, 79, (9), 928–933.

142. Skjolding, L. M.; Winther-Nielsen, M.; Baun, A., Trophic transfer of differently

functionalized zinc oxide nanoparticles from crustaceans (Daphnia magna) to zebrafish (Danio

rerio). Aquat Toxicol 2014, 157, 101-108.

143. Werlin, R.; Priester, J. H.; Mielke, R. E.; Krämer, S.; Jackson, S.; Stoimenov, P. K.;

Stucky, G. D.; Cherr, G. N.; Orias, E.; Holden, P. A., Biomagnification of cadmium selenide

quantum dots in a simple experimental microbial food chain. Nature Nanotechnology 2011, 6,

65–71.

144. Lee, W. M.; Yoon, S. J.; Shin, Y. J.; An, Y. J., Trophic transfer of gold nanoparticles

from Euglena gracilis or Chlamydomonas reinhardtii to Daphnia magna. Environ Pollut 2015,

201, 10-16.

145. Ferry, J. L.; Craig, P.; Hexel, C.; Sisco, P.; Frey, R.; Pennington, P. L.; Fulton, M. H.;

Decho, G. S. A. W.; Kashiwada, S.; Murphy, C. J. M.; Shaw, T. J., Transfer of gold nanoparticles

from the water column to the estuarine food web. Nat Nanotechnol 2009, 4, (7), 441-444.

146. Zhao, X.; M., Y.; Xu, D.; Liu, A.; Hou, X.; Hao, F.; Long, X.; Zhou, Q.; G., J.,

Distribution, bioaccumulation, trophic transfer, and influences of CeO2 nanoparticles in a

constructed aquatic food web. Environ Sci Technol 2017, 51, (9), 5205–5214.

147. Kulacki, K. J.; Cardinale, B. J.; Keller, A. A.; Bier, R.; Dickson, H., How do stream

organisms respond to, and influence, the concentration of titanium dioxide nanoparticles? A

mesocosm study with algae and herbivores. Environ Toxicol Chem 2012, 31, (10), 2414−2422.

148. Judy, J. D.; Unrine, J. M.; Bertsch, P. M., Evidence for biomagnification of gold

nanoparticles within a terrestrial food chain. Environ Sci Technol 2011, 45, (2), 776-781.

149. Judy, J. D.; Unrine, J. M.; Rao, W.; Bertsch, P. M., Bioaccumulation of gold

nanomaterials by Manduca sexta through dietary uptake of surface contaminated plant tissue.


150. Unrine, J. M.; Shoults-Wilson, W. A.; Zhurbich, O.; Bertsch, P. M.; Tsyusko, O. V.,

Trophic transfer of Au nanoparticles from soil along a simulated terrestrial food chain. Environ

Sci Technol 2012, 46, (17), 9753–9760.

151. Hawthorne, J.; de la Torre Roche, R.; Xing, B.; Newman, L. A.; Ma, X.; Majumdar, S.;

Gardea-Torresdey, J.; White, J. C., Particle-size dependent accumulation and trophic transfer of

cerium oxide through a terrestrial food chain. Environ Sci Technol 2014, 48, (22), 13102−13109.

152. Majumdar, S.; Trujillo-Reyes, J.; Hernandez-Viezcas, J. A.; White, J. C.; Peralta-Videa,

J. R.; Gardea-Torresdey, J. L., Cerium biomagnification in a terrestrial food chain: Influence of

particle size and growth stage. Environ Sci Technol 2016, 50, (13), 6782-6792.

153. De la Torre Roche, R.; Servin, A.; Hawthorne, J.; Xing, B.; Newman, L. A.; Ma, X.;

Chen, G.; White, J. C., Terrestrial trophic transfer of bulk and nanoparticle La2O3 does not

depend on particle size. Environ Sci Technol 2015, 49, (19), 11866−11874.

154. Rodrigues, S. M.; Demokritou, P.; Dokoozlian, N.; Hendren, C. O.; Karn, B.; Mauter, M.

S.; Sadik, O. A.; Safarpour, M.; Unrine, J. M.; Viers, J.; Welle, P.; White, J. C.; Wiesner, M. R.;

Lowry, G. V., Nanotechnology for sustainable food production: promising opportunities and

scientific challenges. Environ Sci Nano 2017, 4, (4), 767-781.

155. Torney, F.; Trewyn, B. G.; Lin, V. S.; Wang, K., Mesoporous silica nanoparticles deliver

DNA and chemicals into plants. Nat Nanotechnol 2007, 2, 295-300.

156. Birbaum, K.; Brogioli, R.; Schellenberg, M.; Martinoia, E.; Stark, J. D.; Gunther, D.;

Limbach, L. K., No evidence for cerium dioxide nanoparticle translocation in maize plants.

Environ Sci Technol 2010, 44, 8718-8723.

105

157. Wild, E.; Jones, K. C., Novel method for the direct visualization of in vivo nanomaterials

and chemical interactions in plants. Environ Sci Technol 2009, 43, 5290-5294.

158. Lin, D.; Xing, B., Root uptake and phytotoxicity of ZnO nanoparticles. Environ Sci

Technol 2008, 42, (15), 5590-5595.

159. Hong, J. R., C.M.; Zhao, L.; Adeleye, A.S.; Keller, A.A.; Peralta-Videa, J.R.; Gardea-

Torresdey, J.L., Toxic effects of copper-based nanoparticles or compounds to lettuce (Lactuca

sativa) and alfalfa (Medicago sativa). Environmental science. Processes & impacts 2015, 17, (1),

177-185.

160. Lee, W. M.; An, Y. J.; Yoon, H.; Kweon, H. S., Toxicity and bioavailability of copper

nanoparticles to the terrestrial plants mung bean (Phaseolus radiatus) and wheat (Triticum

aestivum): plant agar test for water-insoluble nanoparticles. Environ Toxicol Chem 2008, 27, (9),

1915-1921.

161. Mukherjee, A.; Bandyopadhyay, S.; Rico, C.; Zhao, L. J.; Peralta-Videa, J. R.; Gardea-

Torresdey, J. L., ZnO nanoparticle induced phytotoxicity and differential anti-oxidative stress in

green peas (Pisum sativum L.) cultivated in soil. Abstr Pap Am Chem S 2013, 246.

162. Wang, P.; Zhao, F. J.; Kopittke, P. M., Nanotechnology: A new opportunity in plant

sciences. Trends Plant Sci 2016, 21, (8), 699-712.

163. Klaine, S. J.; Alvarez, P. J.; Batley, G. E.; Fernandes, T. F.; Handy, R. D.; Lyon, D. Y.;

Mahendra, S.; McLaughlin, M. J.; Lead, J. R., Nanomaterials in the environment: behavior, fate,

bioavailability, and effects. Environ Toxicol Chem 2008, 27, 1825-1851.

164. Dietz, K. J.; Herth, S., Plant nanotoxicology. Trends Plant Sci 2011, 16, 582-589.

165. Steudle, E., How does water get through roots. J Exp Bot 1998, 49, 775-788.

166. Stegemeier, J.; Schwab, F.; Colman, B.; Webb, S.; Newville, M.; Lanzirotti, A.; WInkler,

C.; Wiesner, M.; Lowry, G. V., Speciation matters: Bioavailability of silver and silver sulfide

nanoparticles to alfalfa (Medicago sativa). Environ Sci Technol 2015, 49, 8451-8460.

167. Ma, Y.; He, X.; Zhang, P.; Zhang, Z., Phytotoxicity and biotransformation of La2O3

nanoparticles in a terrestrial plant cucumber (Cucumis sativus). Nanotoxicology 2011, 5, 743-753.

168. Aubert, T.; Burel, A.; Esnault, M. A.; Cordier, S.; Grasset, F.; Cabello-Hurtado, F., Root

uptake and phytotoxicity of nanosized molybdenum octahedral clusters. J Hazard Mater 2012,

219-220, 111-118.

169. Sun, D.; Hussain, H. I.; Yi, Z.; Siegele, R.; Cresswell, T.; Kong, L.; Cahill, D. M.,

Uptake and cellular distribution, in four plant species, of fluorescently labeled mesoporous silica

nanoparticles. Plant Cell Rep 2014, 33, 1389-1402.

170. Collin, B.; Oostveen, E.; Tsyusko, O. V.; Unrine, J. M., Influence of natural organic

matter and surface charge on the toxicity and bioaccumulation of functionalized ceria

nanoparticles in Caenorhabditis elegans. Environ Sci Technol 2014, 48, (2), 1280–1289.

171. Goodman, C. M.; McCusker, C. D.; Yilmaz, T.; Rotello, V. M., Toxicity of gold

nanoparticles functionalized with cationic and anionic side chains. Bioconjugate Chem 2004, 15,

(4), 897-900.

172. Pulido-Reyes, G.; Rodea-Palomares, I.; Das, S.; Sakthivel, T. S.; Leganes, F.; Rosal, R.;

Seal, S.; Fernandez-Pinas, F., Untangling the biological effects of cerium oxide nanoparticles: the

role of surface valence states. Sci Rep 2015, 5, 15613.

173. Judy, J. D.; Bertsch, P. M., Bioavailability, toxicity, and fate of manufactured

nanomaterials in terrestrial ecosystems. Advances in agronomy 2014, 123, 1-64.

174. Huang, X.; Yan, H.; Nazaretski, E.; Conley, R.; Bouet, N.; Zhou, J.; Lauer, K.; Li, L.;

Eom, D.; Legnini, D.; Harder, R.; Robinson, I. K.; Chu, Y. S., 11 nm hard X-ray focus from a

large-aperture multilayer Laue lens. Scientific Reports 2013, 3, 3562.

175. Nazaretski, E.; Yan, H.; Lauer, K.; Bouet, N.; Huang, X.; Xu, W.; Zhou, J.; Shu, D.;

Hwu, Y.; Chu, Y. S., Design and performance of an X-ray scanning microscope at the Hard X-ray

Nanoprobe beamline of NSLS-II. Journal of Synchrotron Radiation 2017, 24, 1113-1119.

106

176. Asati, A.; Santra, S.; Kaittanis, C.; Perez, J. M., Surface-charge-dependent cell

localization and cytotoxicity of cerium oxide nanoparticles. ACS Nano 2010, 4, (9), 5321-5331.

177. Pang, X.; Li, D.; Peng, A., Application of rare-earth elements in the agriculture of China

and its environmental behavior in soil. Environ Sci Pollut Res Int 2002, 9, (2), 143-148.

178. Reed, K., Exploring the properties and applications of nanoceria: is there still plenty of

room at the bottom. Environ Sci Nano 2014, (1), 390-405.

179. Yu, T.; Park, Y. I.; Kang, M. C.; Joo, J.; Park, J. K.; Won, H. Y.; Kim, J. J.; Hyeon, T.,

Large-scale synthesis of water dispersible ceria nanocrystals by a simple sol-gel process and their

use as a chemical mechanical planarization slurry. Eur J Inorg Chem 2008, 6, 855-858.

180. Trovarelli, A.; de Leitenburg, C.; Boaro, M.; Dolcetti, G., The utilization of ceria in

industrial catalysis. Catal Today 1990, 50, (2), 353-367.

181. Thill, A.; Zeyons, O.; Spalla, O.; Chauvat, F.; Rose, J.; Auffan, M.; Flank, A. M.,

Cytotoxicity of CeO2 nanoparticles for Escherichia coli. Physico-chemical insight of the

cytotoxicity mechanism. Environ Sci Technol 2006, 40, (19), 6151-6.

182. Van Hoecke, K.; Quik, J. T.; Mankiewicz-Boczek, J.; de Schamphelaere, K. A.;

Elsaesser, A.; Van der Meeren, P.; Barnes, C.; McKerr, G.; Howard, C. V.; Van de Meent, D.;

Rydzynski, K.; Dawson, K. A.; Salvati, A.; Lesniak, A.; Lynch, I.; Silversmit, G.; de Samber, B.;

Vincze, L.; Janssen, C. R., Fate and effects of CeO2 nanoparticles in aquatic ecotoxicity tests.


183. Karakoti, A. S.; Munusamy, P.; Hostetler, K.; Kodali, V.; Kuchibhatla, S.; Orr, G.;

Pounds, J. G.; Teeguarden, J. G.; Thrall, B. D.; Baer, D. R., Preparation and characterization

challenges to understanding environmental and biological impacts of ceria nanoparticles. Surf

Interface Anal 2012, 44, (8), 882-889.

184. Rejman, J.; Oberle, V.; Zuhorn, I. S.; Hoekstra, D., Size-dependent internalization of

particles via the pathways of clathrin- and caveolae-mediated endocytosis. Biochem J 2004, 377,

(1), 159-169.

185. Li, L.; Yan, H.; Xu, W.; Yu, D.; Heroux, A.; Lee, W. K.; Campbell, S. I.; Chu, Y. S.,

Python-based X-ray fluorescence analysis package. Proc SPIE 10389, X-ray Nanoimaging:

Instruments and Methods III 2017.

186. Ravel, B.; Newville, M., Athena, Artemis, Hephaestus: data analysis for X-ray absorption

spectroscopy using IFEFFIT. J Synchrotron Rad 2005, 12, (4), 537-541.

187. Thomas, P.; Carpenter, D.; Boutin, C.; Allison, J., Rare earth elements (REEs): Effects on

germination and growth of selected crop and native plant species. Chemosphere 2014, 78, 273-

279.

188. Li, H.; Ye, X.; Guo, X.; Geng, Z.; Wang, G., Effects of surface ligands on the uptake and

transport of gold nanoparticles in rice and tomato. J Hazard Mater 2016, 314, 188-196.

189. Koelmel, J.; Leland, T.; Wang, H.; Amarasiriwardena, D.; Xing, B., Investigation of gold

nanoparticles uptake and their tissue level distribution in rice plants by laser ablation-inductively

coupled-mass spectrometry. Environ Pollut 2013, 174, 222-228.

190. Wang, J.; Yang, Y.; Zhu, H.; Braam, J.; Schnoor, J. L.; Alvarez, P. J., Uptake,

translocation, and transformation of quantum dots with cationic versus anionic coatings by

Populus deltoides x nigra cuttings. Environ Sci Technol 2014, 48, (12), 6754-6762.

191. Spielman-Sun, E.; Lombi, E.; Donner, E.; Howard, D.; Unrine, J. M.; Lowry, G. V.,

Impact of surface charge on cerium oxide nanoparticle uptake and translocation by wheat

(Triticum aestivum). Environ Sci Technol 2017, 51, (13), 7361-7368.

192. Graham Uschi, M.; Tseng Michael, T.; Jasinski Jacek, B.; Yokel Robert, A.; Unrine

Jason, M.; Davis Burtron, H.; Dozier Alan, K.; Hardas Sarita, S.; Sultana, R.; Grulke Eric, A.;

Butterfield, D. A., In vivo processing of ceria nanoparticles inside liver: Impact on free‐radical

scavenging activity and oxidative stress. Chem Plus Chem 2014, 79, (8), 1083-1088.

107

193. Rui, Y.; Zhang, P.; Zhang, Y.; Ma, Y.; He, X.; Gui, X.; Li, Y.; Zhang, J.; Zheng, L.; Chu,

S.; Guo, Z.; Chai, Z.; Zhao, Y.; Zhang, Z., Transformation of ceria nanoparticles in cucumber

plants is influenced by phosphate. Environ Pollut 2015, 198, 8-14.

194. Graham, U. M.; Tseng, M. T.; Jasinski, J. B.; Yokel, R. A.; Unrine, J. M.; Davis, B. H.;

Dozier, A. K.; Hardas, S. S.; Sultana, R.; Grulke, E. A.; Butterfield, D. A., In vivo processing of

ceria nanoparticles inside liver: Impact on free-radical scavenging activity and oxidative stress.

Chempluschem 2014, 79, (8), 1083-1088.

195. Perez, J. M.; Asati, A.; Nath, S.; Kaittanis, C., Synthesis of biocompatible dextran-coated

nanoceria with pH-dependent antioxidant properties. Small 2008, 4, 552-556.

196. Lundqvist, M.; Stigler, J.; Cedervall, T.; Berggard, T.; Flanagan, M. B.; Lynch, I.; Eliaz,

G.; Dawson, K. A., The evolution of the protein corona around nanoparticles: A test study. ACS

Nano 2011, 5, (9), 7503-7509.

197. Walkey, C. D.; Olsen, J. B.; Song, F.; Liu, R.; Guo, H., Protein corona fingerprinting

predicts the cellular interaction of gold and silver nanoparticles. ACS Nano 2014, 8, (3), 2439-

2455.

198. López-Moreno, M. L.; de la Rosa, G.; Hernández-Viezcas, J. A.; Peralta-Videa, J. R.;

Gardea-Torresdey, J. L., X-ray absorption spectroscopy (XAS) corroboration of the uptake and

storage of CeO2 nanoparticles and sssessment of their differential toxicity in four edible plant

species. J Agric Food Chem 2010, 58, (6), 3689–3693.

199. Castiglione, M. R.; Giorgetti, L.; Geri, C.; Cremonini, R., The effects of nano-TiO2 on

seed germination, development and mitosis of root tip cells of Vicianar bonensis L. and Zea mays

L. J Nanopart Res 2011, 13, (6), 2443-2449.

200. Feng, Y.; Cui, X.; He, S.; Dong, G.; Chen, M.; Wang, J.; Lin, X., The role of metal

nanoparticles in influencing arbuscular mycorrhizal fungi effects on plant growth. Environ Sci

Technol 2013, 47, (16), 9496-9504.

201. Ghosh, M.; Bandyopadhyay, M.; Mukherjee, A., Genotoxicity of titanium dioxide (TiO2)

nanoparticles at two trophic levels: plant and human lymphocytes. Chemosphere 2010, 81, (10),

1253-1262.

202. Poborilova, Z.; Opatrilova, R.; Babula, P., Toxicity of aluminium oxide nanoparticles

demonstrated using a BY-2 plant cell suspension culture model. Environ Exp Bot 2013, 91, 1-11.

203. Wang, Z.; Xu, L.; Zhao, J.; Wang, X.; White, J. C.; Xing, B., CuO nanoparticle

interaction with Arabidopsis thaliana: Toxicity, parent-progeny transfer, and gene expression.

Environ Sci Technol 2016, 50, (11), 6008–6016.

204. Conway, J. R.; Hanna, S. K.; Lenihan, H. S.; Keller, A. A., Effects and implications of

trophic transfer and accumulation of CeO2 nanoparticles in a marine mussel. Environ Sci Technol

2014, 48, (3), 1517–1524.

205. Lewinski, N. A.; Zhu, H.; Ouyang, C. R.; Conner, G. P.; Wagner, D. S.; Colvin, V. L.;

Drezek, R. A., Trophic transfer of amphiphilic polymer coated CdSe/ZnS quantum dots to Danio

rerio. Nanoscale 2011, 3, (8), 3080-3083.

206. Pakrashi, S.; Dalai, S.; Chandrasekaran, N.; Mukherjee, A., Trophic transfer potential of

aluminium oxide nanoparticles using representative primary producer (Chlorella ellipsoides) and

a primary consumer (Ceriodaphnia dubia). Aquat Toxicol 2014, 152, 74-81.

207. Judy, J. D. U., J.M.; Bertsch, P.M., Evidence for Biomagnification of Gold Nanoparticles

within a Terrestrial Food Chain. Environ Sci Technol 2011, 45, (2), 776-781.

208. Zhu, Z.; Wang, H.; Yan, B.; Zheng, H.; Jiang, Y.; Miranda, O. R.; Rotello, V. M.; Xing,

B.; Vachet, R. W., Effect of surface charge on the uptake and distribution of gold nanoparticles in

four plant species. Environ Sci Technol 2012, 46, 12391−12398.

209. Pang, X. L., D.; Peng, A., Application of rare-earth elements in the agriculture of China

and its environmental behavior in soil. Environ Sci Pollut Res Int 2002, 9, (2), 143-148.

108

210. Yu, T. P., Y.I.; Kang, M.C.; Joo, J.; Park, J.K.; Won, H.Y.; Kim, J.J.; Hyeon, T., Large-

scale synthesis of water dispersible ceria nanocrystals by a simple sol-gel process and their use as

a chemical mechanical planarization slurry. Eur J Inorg Chem 2008, (6), 855-858.

211. Leung, Y. H.; Yung, M. M.; Ng, A. M.; P., M. A.; Wong, S. W.; Chan, C. M.; Ng, Y. H.;

Djurišić, A. B.; Guo, M.; Wong, M. T.; Leung, F. C.; Chan, W. K.; Leung, K. M.; Lee, H. K.,

Toxicity of CeO2 nanoparticles - the effect of nanoparticle properties. J Photochem Photobiol B

2015, 145, 48-59.

212. Zhang, H. H., X.; Zhang, Z.; Zhang, P.; Li, Y.; Ma, Y.; Kuang, Y.; Zhao, Y.; Chai, Z.,

Nano-CeO2 exhibits adverse effects at environmental relevant concentrations. Environ Sci

Technol 2011, 45, (8), 3725-3730.

213. Pulido-Reyes, G. R.-P., I.; Das, S.; Sakthivel, T. S.; Leganes, F.; Rosal, R.; Seal, S.;

Fernandez-Pinas, F., Untangling the biological effects of cerium oxide nanoparticles: the role of

surface valence states. Sci Rep 2015, 5, 15613.

214. Appel, H. M.; Martin, M. M., Gut redox conditions in herbivorous lepidopteran larvae. J

Chem Ecol 1990, 16, (12), 3277-3290.

215. Dow, J. A., Extremely high pH in biological systems: a model for carbonate transport.

Am J Physiol 1984, 246, (4), 633-636.

216. Dow, J. A., pH Gradients in lepidopteran midgut. J Exp Biol 1992, 172, (1), 355-375.

217. Lehane, M. J., Peritrophic matrix structure and function. Annu Rev Entomol 1997, 42,

525-550.

218. Zhang, P. M., Y., Zhang Z.; He, X.; Zhang, J.; Guo, Z.; Tai, Z.; Zhao, Y.; Chai, Z.,

Biotransformation of Ceria Nanoparticles in Cucumber Plants. ACS Nano 2012, 6, (11), 9943–

9950.

219. Gardea-Torresdey, J. L.; Rico, C. M.; White, J. C., Trophic transfer, transformation, and

impact of engineered nanomaterials in terrestrial environments. Environ Sci Technol 2014, 48,

(5), 2526-2540.

220. Schwabe, F.; Tanner, S.; Schulin, R.; Rotzetter, A.; Stark, W.; von Quadt, A.; Nowack,

B., Dissolved cerium contributes to uptake of Ce in the presence of differently sized CeO2-

nanoparticles by three crop plants. Metallomics 2015, 7, (3), 466-477.

221. Yokel, R.; Au, T.; MacPHail, R.; Hardas, S. S.; Butterfield, D. A.; Sultana, R.; Goodman,

M.; Tseng, M.; Dan, M.; Haghnazar, H.; Unrine, J.; Graham, U.; Wu, P.; Grulke, E., Distribution,

elimination and biopersistence to 90 days of a systemically-introduced 30 m ceria engineered

nanomaterial in rats. Toxicol Sci 2012, 127, (1), 256-268.

222. Feswick, A.; Griffitt, R. J.; Siebein, K.; Barber, B. S., Uptake, retention and

internalization of quantum dots in Daphnia is influenced by particle surface functionalization.

Aquat Toxicol 2013, 130–131, 210-218.

223. Hegedus, D.; Erlandson, M.; Gillott, C.; Toprak, U., New insights into peritrophic matrix

synthesis, architecture, and function. Annu Rev Entomol 2009, 54, 285-302.

224. Cota-Ruiz, K.; Hernández-Viezcas, J. A.; Varela-Ramírez, A.; Valdés, C.; Núñez-

Gastélum, J. A.; Martínez-Martínez, A.; Delgado-Rios, M.; Peralta-Videa, J. R.; Gardea-

Torresdey, J. L., Toxicity of copper hydroxide nanoparticles, bulk copper hydroxide, and ionic

copper to alfalfa plants: A spectroscopic and gene expression study. Environ Pollut 2018, 243(Pt

A), 703-712.

225. Tan, W.; Gao, Q.; Deng, C.; Wang, Y.; Lee, W. Y.; Hernandez-Viezcas, J. A.; Peralta-

Videa, J. R.; Gardea-Torresdey, J. L., Foliar Exposure of Cu(OH)2 Nanopesticide to Basil

( Ocimum basilicum): Variety-Dependent Copper Translocation and Biochemical Responses. J

Agric Food Chem 2018, 66, (13), 3358-3366.

226. Zhao, L.; Huang, Y.; Keller, A. A., Comparative metabolic response between cucumber

( Cucumis sativus) and corn ( Zea mays) to a Cu(OH)2 nanopesticide. J Agric Food Chem 2018,

66, (26), 6628-6636.

109

227. Zhao, L.; Huang, Y.; Hannah-Bick, C.; Fulton, A. N.; Keller, A. A., Application of

metabolomics to assess the impact of Cu(OH)2 nanopesticide on the nutritional value of lettuce

(Lactuca sativa): Enhanced Cu intake and reduced antioxidants. NanoImpact 2016, 3-4, 58-66.

228. Zhao, L.; Hu, Q.; Huang, Y.; Fulton, A. N.; Hannah-Bick, C.; Adeleye, A. S.; Keller, A.

A., Activation of antioxidant and detoxification gene expression in cucumber plants exposed to a

Cu(OH)2 nanopesticide. Environ Sci Nano 2017, 4, (8), 1750-1760.

229. Nair, P. M. G.; Chung, I. M., Study on the correlation between copper oxide

nanoparticles induced growth suppression and enhanced lignification in Indian mustard (Brassica

juncea L.). Ecotoxicol Environ Saf 2015, 113, 302-313.

230. Apodaca, S. A.; Tan, W.; Dominguez, O. E.; Hernandez-Viezcas, J. A.; Peralta-Videa, J.

R.; Gardea-Torresdey, J. L., Physiological and biochemical effects of nanoparticulate copper,

bulk copper, copper chloride, and kinetin in kidney bean (Phaseolus vulgaris) plants. Sci Total

Environ 2017, 599-600, 2085−2094.

231. Keller, A. A.; Adeleye, A. S.; Conway, J. R.; Garner, K. L.; Zhao, L.; Cherr, G. N.,

Comparative environmental fate and toxicity of copper nanomaterials. Nanoimpact 2017, 7, 28-

40.

232. Adeleye, A. S.; Conway, J. R.; Perez, T.; Rutten, P.; Keller, A. A., Influence of

extracellular polymeric substances on the long-term fate, dissolution, and speciation of copper-

based nanoparticles. Environ Sci Technol 2014, 48, (21), 12561-12568.

233. Lu, C.; Qi, L.; Yang, J.; D., Z., Simple template-free solution route for the controlled

synthesis of Cu(OH)2 and CuO nanostructures. J Phys Chem B 2004, 108, (46), 17825-17831.

234. Grunert, L. W.; Clarke, J. W.; Ahuja, C.; Eswaran, H.; Nijhout, H. F., A quantitative

analysis of growth and size regulation in Manduca sexta: The physiological basis of variation in

size and age at metamorphosis. PLoS One 2015, 10, (5), 1-23.

235. Nijhout, H. F.; Davidowitz, G.; Roff, D. A., A quantitative analysis of the mechanism

that controls body size in Manduca sexta. J Biol 2006, 5, (5), 16.

236. Simonin, M.; Colman, B. P.; Tang, W.; Judy, J. D.; Anderson, S. M.; Bergemann, C. M.;

Rocca, J. D.; Unrine, J. M.; Cassar, N.; Bernhardt, E. S., Plant and microbial responses to

repeated Cu(OH)2 nanopesticide exposures under different fertilization levels in an agro-

ecosystem. Front Microbiol 2018, 9, 1769.

237. Congdon, J. D.; Dunham, A. E.; Hopkins, W. A.; Rowe, C. L.; Hinton, T. G., Resource

allocation‐based life histories: A conceptual basis for studies of ecological toxicology. Environ

Toxicol Chem 2001, 20, 1698-1703.

238. Griffitt, R. J.; Luo, J.; Gao, J.; Bonzongo, J. C.; Barber, D. S., Effects of particle

composition and species on toxicity of metallic nanomaterials in aquatic organisms. Environ

Toxicol Chem 2008, 27, (9), 1972-1978.

239. Tavares, K. P.; Oliveira, Á. C. d.; Vicentini, D. S.; Melegari, S. P.; Matias, W. G.;

Barbosa, S.; Kummrow, F., Acute toxicity of copper and chromium oxide nanoparticles to

Daphnia similis. Ecotoxicol Environ Contam 2014, 9, (1), 43-50.

240. Franklin, N. M.; Rogers, N. J.; Apte, S. C.; Batley, G. E.; Gadd, G. E.; Casey, P. S.,

Comparative toxicity of nanoparticulate ZnO, bulk ZnO, and ZnCl2 to a freshwater microalga

(Pseudokirchneriella subcapitata): The importance of particle solubility. Environ Sci Technol

2007, 41, (24), 8484-8490.

241. Aruoja, V.; Dubourguier, H.-C.; Kasemets, K.; AnneKahru, Toxicity of nanoparticles of

CuO, ZnO and TiO2 to microalgae Pseudokirchneriella subcapitata. Sci Total Environ 2009, 407,

(4), 1461-1468.

242. Navarro, E.; Piccapietra, F.; Wagner, B.; Marconi, F.; Kaegi, R.; Odzak, N.; Sigg, L.;

Behra, R., Toxicity of silver nanoparticles to Chlamydomonas reinhardtii. Environ Sci Technol

2008, 42, (23), 8959–8964.

110

243. Beer, C.; Foldbjerg, R.; Hayashi, Y.; Sutherland, D. S.; Autrup, H., Toxicity of silver

nanoparticles—Nanoparticle or silver ion. Toxicol Lett 2012, 208, (3), 286-292.

244. Song, Y. F.; Huang, C.; Shi, X.; Pan, Y. X.; Liu, X.; Luo, Z., Endoplasmic reticulum

stress and dysregulation of calcium homeostasis mediate Cu-induced alteration in hepatic lipid

metabolism of javelin goby Synechogobius hasta. Aquat Toxicol 2016, 175, 20-29.

245. Chenae, Z.; Mengae, H.; Xing, G.; Chen, C.; Zhao, Y.; Jia, G.; Wang, T.; Yuan, H.; Ye,

C.; Zhao, F.; Chai, Z.; Zhu, C.; Fang, X.; Ma, B.; Wan, L., Acute toxicological effects of copper

nanoparticles in vivo. Toxicol Lett 2006, 163, (2), 109-120.

246. Vijver, M. G.; Gestel, C. A. M. v.; Lanno, R. P.; Straalen, N. M. v.; Peijnenburg, W. J. G.

M., Internal metal sequestration and its ecotoxicological relevance: A review. Environ Sci

Technol 2004, 38, (18), 4705–4712.

247. Gomes, F. M.; Carvalho, D. B.; Peron, A. C.; Saito, K.; Miranda, K.; Machado, E. A.,

Inorganic polyphosphates are stored in spherites within the midgut of Anticarsia gemmatalis and

play a role in copper detoxification. J Insect Physiol 2012, 58, (2), 211-219.

248. Luoma, S. N.; Rainbow, P. S., Why is metal bioaccumulation so variable? Biodynamics

as a unifying concept. Environ Sci Technol 2005, 39, (7), 1921−1931.

249. Davidowitz, G.; D'Amico, L. J.; Nijhout, H. F., The effects of environmental variation on

a mechanism that controls insect body size. Evol Ecol Res 2004, 6, (1), 49-62.

111

VITA

JIERAN LI

EDUCATION

Wuhan University: Sep 2009 – Aug 2013, B.Sc. in Biology

PROFESSIONAL POSITIONS

Research Assistant, Department of Toxicology and Cancer Biology, University of Kentucky

Research Assistant, Center of Metabolomics and System Biology, University of Kentucky

Student Intern, Center for Disease Control and Environment Protection, Wuhan, Hubei,

China

SCIENTIFIC PUBLICATIONS

1. Zhao, J., Li, J., Fan, T. W. M. and Hou, S. X., Glycolytic reprogramming through

PCK2 regulates tumor initiation of prostate cancer cells, Oncotarget, 2017 Oct 13;

8(48): 83602–83618.

2. Li, J., Tappero, R. V., Acerbo, A. S., Yan, H. Y., Chu, Y. C., Lowry, G. V. and

Unrine, J. M., Effect of CeO2 nanomaterial surface functional groups on tissue and

subcellular distribution of Ce in tomato (Solanum lycopersicum), Environ Sci

Nano, published online, 2018 Dec, DOI: 10.1039/c8en01287c.

3. Li, J., Unrine, J. M., Tappero, R. V., Acerbo, A. S., Uptake, subcellular distribution,

and translocation of differentially charged CeO2 nanomaterials in Solanum

lycopersicum cv Micro-Tom (tomato), 2016, Proceedings of the 11th International

Conference on Environmental Effects of Nanotechnology and Nanomateirals,

Golden, CO.

MISCELLANEOUS

Scientific Outreach Program. National Synchrotron Light Source II, Brookhaven National

Laboratory

X-ray fluorescence microscopy and environmental toxicology (2017)

Manager of teaching affairs. ErHao Education Group, Chongqing, China

Personalized course design and coordination of teaching affairs (2012 – 2013)

Coordinator for teaching affairs. Educational Department, Wuhan University

Education Communication Exchange program between Wuhan University and Ohio

State University (2013)

impact of physico-chemical properties of …

Documents