# Nanoparticle surface charge influences translocation and leaf distribution in vascular plants with contrasting anatomy

Eleanor Spielman-Sun, Astrid Avellan, Garret Bland, Ryan Tappero, Alvin

Acerbo, Jason Unrine, Juan Pablo Giraldo, Gregory Lowry

► To cite this version:

Eleanor Spielman-Sun, Astrid Avellan, Garret Bland, Ryan Tappero, Alvin Acerbo, et al.. Nanoparticle surface charge influences translocation and leaf distribution in vascular plants with contrasting anatomy. Environmental science. Nano, 2019, 6 (8), pp.2508-2519. 10.1039/C9EN00626E. hal-03707346

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# Nanoparticle surface charge influences translocation and leaf distribution in vascular plants with contrasting anatomy

E. Spielman-Sun, R. Tappero

To be published in "ENVIRONMENTAL SCIENCE NANO"

July 2020

Photon Sciences

**Brookhaven National Laboratory**

**U.S. Department of Energy**

USDOE Office of Science (SC), Basic Energy Sciences (BES) (SC-22)

Notice: This manuscript has been authored by employees of Brookhaven Science Associates, LLC under Contract No. DE-SC0012704 with the U.S. Department of Energy. The publisher by accepting the manuscript for publication acknowledges that the United States Government retains a non-exclusive, paid-up, irrevocable, world-wide license to publish or reproduce the published form of this manuscript, or allow others to do so, for United States Government purposes.
DISCLAIMER

This report was prepared as an account of work sponsored by an agency of the United States Government. Neither the United States Government nor any agency thereof, nor any of their employees, nor any of their contractors, subcontractors, or their employees, makes any warranty, express or implied, or assumes any legal liability or responsibility for the accuracy, completeness, or any third party's use or the results of such use of any information, apparatus, product, or process disclosed, or represents that its use would not infringe privately owned rights. Reference herein to any specific commercial product, process, or service by trade name, trademark, manufacturer, or otherwise, does not necessarily constitute or imply its endorsement, recommendation, or favoring by the United States Government or any agency thereof or its contractors or subcontractors. The views and opinions of authors expressed herein do not necessarily state or reflect those of the United States Government or any agency thereof.
# Nanoparticle surface charge influences translocation and leaf distribution in vascular plants with contrasting anatomy†

Eleanor Spielman-Sun, IDª Astrid Avellan, IDª Garret D. Bland, IDª

Ryan V. Tappero, IDᵇ Alvin S. Acerbo,ᵇᶜ Jason M. Unrine, IDᵈ

Juan Pablo Giraldo IDᵉ and Gregory V. Lowry IDª*ᵃ

Root uptake and translocation of engineered nanoparticles (NPs) by plants are dependent on both plant species and NP physicochemical properties. To evaluate the influence of NP surface charge and differences in root structure and vasculature on cerium distribution and spatial distribution within plants, two monocotyledons (corn and rice) and two dicotyledons (tomato and lettuce) were exposed hydroponically to positively-charged, negatively-charged, and neutral ~4 nm CeO₂ NPs. Leaves were analyzed using synchrotron-based X-ray fluorescence microscopy to provide lateral Ce spatial distribution. Surface charge mediated CeO₂ NP interactions with roots for all plant species. Positively charged CeO₂ NPs associated to the roots more than the negatively charged NPs due to electrostatic attraction/repulsion to the negatively charged root surfaces, with the highest association for the tomato, likely due to higher root surface area. The positive NPs remained primarily adhered to the roots untransformed, while the neutral and negative NPs were more efficiently translocated from the roots to shoots. This translocation efficiency was highest for the tomato and lettuce compared to corn and rice. Across all plant species, the positive and neutral treatments resulted in the formation of Ce clusters outside of the main vasculature in the mesophyll, while the negative treatment resulted in Ce primarily in the main vasculature of the leaves. Comparing leaf vasculature, Ce was able to move much further outside of the main vasculature in the dicot plants than monocot plants, likely due to the larger airspace volume in dicot leaves compared to monocot leaves. These results provide valuable insight into the influence of plant structure and NP properties on metal transport and distribution of NPs in plants.

## Environmental significance

Plant nanobiotechnology promises transformative solutions to the most vexing problems threatening global food security, e.g. drought, disease, and soil nutrient deficiencies. However, the lack of cost-effective methods to deliver the nanomaterials to precise locations in plants where they are needed to be active, e.g. inside vascular tissues, or into plant organelles, impedes these technological innovations. These findings not only provide insight into how plant structural features influence NP behavior but also how surface charge can be tailored for targeted delivery of nutrients to specific plant organs.

## Introduction

Nanotechnology has the potential to become a valuable tool for improving agro-ecosystem resiliency against major environmental stressors (e.g. drought, salinity, disease) and efficiency by enhancing crop productivity and reducing nutrient losses (e.g. through controlled release of agrochemicals and target-specific delivery).¹⁻⁵ Firstly, nanoparticles (NPs) are small enough to cross important plant mechanical barriers (e.g. cuticle and cell walls).⁶ NPs can also cross cellular and organelle membranes and move in between cells which enables them to load into the vasculature either *via* apoplastic (extracellular) or symplastic (intracellular) pathways, as both

<footnote>ªCivil and Environmental Engineering, Carnegie Mellon University, Pittsburgh, Pennsylvania 15213, USA. E-mail: glowry@cmu.edu; Tel: (+412) 268 2948</footnote>

<footnote>ᵇNational Synchrotron Light Source II, Brookhaven National Laboratory, Upton, NY 11973, USA</footnote>

<footnote>ᶜCenter for Advanced Radiation Sources University of Chicago, Chicago, IL 60637, USA</footnote>

<footnote>ᵈDepartment of Plant and Soil Sciences, University of Kentucky, Lexington, Kentucky 40546, USA</footnote>

<footnote>ᵉDepartment of Botany and Plant Sciences, University of California, Riverside, CA 92521, USA</footnote>

<footnote>†Electronic supplementary information (ESI) available. See DOI: 10.1039/c9en00626e</footnote>
mechanisms have been proposed for translocation of NPs in plants.<sup>4,7,8</sup> Secondly, NP surfaces are easily modifiable with a variety of coatings and, similar to drug delivery, NP surface properties can theoretically be tuned to deliver them to specific tissues or organelles in plants.<sup>9,10</sup> Combined with the inherent rate limited dissolution of many metal and metal oxide NPs, NPs could provide slow release<sup>11,12</sup> of the constituent metals in the desired locations of plants, which can be less phytotoxic than soluble forms of metals applied at the same dose.<sup>13</sup> Finally, NPs have been shown to increase photosynthesis,<sup>14,15</sup> biomass production rates,<sup>16,17</sup> plant stress tolerance,<sup>18</sup> plant resistance to diseases,<sup>19,20</sup> and agrochemical utilization efficiency.<sup>21</sup> Despite the great potential of NPs, there remains limited understanding of how NP physical and chemical properties (e.g. size, charge, solubility, coating, chemical composition) dictate NP-plant interactions and translocation behavior in plants. A better understanding of these NP-plant interactions is needed to design targeted and controlled delivery, which has the potential to reduce the number of applications of fertilizers and pesticides, decrease nutrient losses from fertilizers, and increase yields through optimized nutrient management.

Surface charge is an important property dictating NP fate in plants. While positively-charged CeO₂ NPs have been shown to more readily attach to roots of wheat (*Triticum aestivum* cv. *Shield*)²² and tomato (*Solanum lycopersicum* cv. *Micro-Tom*),²³ negatively charged CeO₂ NPs more efficiently translocate to the shoots. The same trend has been observed for gold nanoparticles (AuNPs) in radish, ryegrass, rice and pumpkin²⁴ and in tomato and rice.²⁵ Because charge can influence NP interactions with charged biological structures²⁶ and therefore ability to cross biological membranes,²⁷ it is desirable to better understand precisely how charge affects the spatial distribution of NPs that have translocated in plants. Few studies characterize the spatial distribution of the NPs in leaves beyond total metal analysis, particularly at the whole-leaf scale.²²,²³,²⁸

Though many published studies have focused on NP uptake by plant roots, the observations made in one plant species are often difficult to generalize to other plants. Often differences in the plant's structural features, e.g. root or leaf architecture, are hypothesized to explain the observed differences in NP uptake and translocation, but this hypothesis has yet to be systematically evaluated. Flowering plants (angiosperms) can be classified by anatomical differences into two categories: monocotyledon (monocot) and dicotyledon (dicot). In general, monocots are more resistant to heavy metal NP uptake than dicots.<sup>29-32</sup> Differences in NP uptake between monocots and dicots could be due to differences in vasculature and structural features (fibrous vs. taproot system) leading to different surface areas interacting with the environment, greater binding capacity and transpiration<sup>33</sup> in dicots, and differences in root exudation profiles,<sup>34-37</sup> as well as mucilage production at the NPs-root tip interface.<sup>38</sup> However, the relative importance of each of these differences has yet to be thoroughly investigated.

CeO₂ NP interactions with different plant species have been widely studied. However, there are contradictory reports on whether CeO₂ NPs may act as an oxidative stress inducer or antioxidant. CeO₂ NPs have been shown to protect cells *in vitro* against reactive oxygen species (ROS)-induced damage,<sup>39-41</sup> including isolated chloroplasts<sup>14,42</sup> due to oxygen vacancies in the crystalline lattice that readily enable cycling between Ce³⁺ and Ce⁴⁺ oxidation states. However, there are limited examples of this potential *in vivo*. CeO₂ NPs with low Ce(III)/Ce(IV) ratios (50 mg L⁻¹) delivered *via* foliar infiltration have also been shown to improve plant photosynthetic rates under heat<sup>43</sup> and salinity<sup>18,44</sup> by serving as antioxidants. In contrast, decreased photosynthetic rate and CO₂ assimilation efficiency, increased lipid peroxidation, and other stress measures have also been observed in a variety of plants exposed to CeO₂ NPs, particularly at high doses greater than 500 mg kg⁻¹ soil<sup>45-49</sup> and NPs with higher Ce(III)/Ce(IV) ratios.<sup>50</sup> A better understanding of the impacts of NP properties on plant photosynthesis and respiration is needed to fully leverage their benefits.

The goal of the present study was to systematically assess whether plants with different morphologies, two monocots (corn and rice) and two dicots (tomato and lettuce) similarly accumulate CeO₂ NPs and how NP charge influences spatial distribution in leaves with contrasting architectures. We use short-term, well-controlled hydroponic exposure scenarios to determine whether differences in NP charge and plant anatomy and physiology influence the translocation, speciation changes, and spatial distribution of Ce within the plant leaf tissue. A better understanding of the impact of NP charge on translocation routes and distribution in leaves can inform future efforts to design NPs for delivery to specific locations in plant tissues.

# Materials and methods

## Material characterization

Cerium dioxide NPs with three different charges were synthesized as reported previously in Collin *et al.*<sup>51</sup> Briefly, uncharged dextran coated CeO₂ NPs (CeO₂ NP(0)) with a nominal 4 nm primary particle diameter were synthesized then further functionalized with either diethylaminoethyl groups to create a net positive charge (CeO₂(+)) or with carboxymethyl groups to create a net negative charge (CeO₂(-)). The particles were diluted to 50 mg L⁻¹ as Ce in a basal salt solution (1 mM CaCl₂ and 5 µM H₃BO₃, pH = 5.6) and probe sonicated (550 Sonic Dismembrator, Fisher Scientific) for 1 min at 10 s intervals to ensure dispersion. The hydrodynamic diameter and electrophoretic mobility of the NPs in the exposure medium at the exposure concentration (50 mg-Ce per L) were measured using a Nano Zetasizer (Malvern Instruments, Malvern).

## Plant growth and exposure

Crops commonly grown in the United States and easy to cultivate in lab were chosen as model plants. Corn (*Zea mays* cv. *Trinity*) and lettuce (*Lactuca sativa* cv. *Buttercrunch*) seeds
were obtained from Johnny's Selected Seeds (Winslow, ME), and tomato (Solanum lycopersicum cv. Roma) from Burpee Seeds (Warminster, PA). Rice (Oryza sativa cv. Nipponbare) were obtained from the USDA-ARS Dale Bumpers National Rice Research Center (Stuttgart, AR). Seeds were surface sterilized with commercial bleach for 10 min and then thoroughly rinsed with DI water. The sterilized seeds were germinated on deionized water-moistened filter paper in a Petri dish for 4 days for corn, 6 days for tomato and lettuce, and 7 days for rice. Germination was staggered so that all plants were transferred to hydroponic containers on the same day. Each 100 mL container was filled with 80 mL of 1/4 strength Hoagland's medium and covered with a plastic lid with five holes. Five seedlings were transplanted to five of the holes with the roots suspended in a continuously aerated solution. Plants were grown in a controlled environment chamber (Binder™ Model KBWF 729; day/night photoperiod 16 h/8 h, day/night temperature 25 °C/21 °C and 60% humidity). Solution was renewed every 3 days with fresh 1/4 strength Hoagland's medium. After 2 weeks, the plants were hydroponically exposed to 50 mg-Ce per L of CeO₂ NPs as CeO₂(+), CeO₂(0), or CeO₂(−) in a continuously aerated basal salt solution (1 mM CaCl₂ and 5 µM H₃BO₃, pH = 5.6) for 48 h. Exposures were performed in this solution to reduce phosphate interference⁵² and over a short period of time to focus on the plant's initial response to particles with different charges. After exposure, plant roots were rinsed for 30 s in Ce-free medium to remove loosely adhered Ce. This exposure protocol was used for all subsequent measurements.

## Plant health measurements

At the end of the 48 h exposure period, photosynthetic CO₂ quantum yield (ΦCO₂; µmol CO₂ µmol photon⁻¹), photosystem II quantum yield (ΦPSII; mol e⁻ µmol⁻¹), transpiration rates (E; mol H₂O m⁻² s⁻¹), electron transport rates (ETR; µmol photon m⁻² s⁻¹), and stomatal conductance (gsw; H₂O mol m⁻² s⁻¹) were measured in quadruplicate on light-adapted leaves using a LI-6800 portable gas analyzer and fluorometer (Li-COR Bio-sciences, Lincoln, NE). The leaf chamber conditions were: light intensity 600 µmol m⁻² s⁻¹ photosynthetically active radiation (PAR), humidity 60%, leaf temperature 25 °C, flow 500 µmol s⁻¹, and CO₂ concentration 400 µmol mol⁻¹. Measurements were made between 3 to 5 h after sunrise to ensure similar stomatal aperture between samples. Leaves were left to equilibrate for 2 min in the Li-COR chamber before reading. Leaf PSII fluorescence was measured using a fluorometer using a flash of saturated light (1500 µmol m⁻² s⁻¹). For plant root surface area (SA) approximations, four sets of plant roots per species were scanned using an EPSON Perfection V19 scanner. The images were processed using ImageJ software (v 1.52h) to calculate the 2-D surface area.

## Total Ce association and translocation

After exposure, plants were harvested, and roots and shoots separated and lyophilized. Dried plant tissue samples were

digested overnight at room temperature in concentrated HNO₃ and 30% H₂O₂, then heated to 95 °C for 30 minutes, then allowed to cool down and 30% H₂O₂ was added to obtain a 2:1 HNO₃:H₂O₂ (v/v) ratio and heated again at 95 °C for 2 h (protocol adapted from EPA Method 3050b⁵³). Following digestion, the samples were diluted to 5% (v/v) HNO₃ using deionized water and filtered through a 0.45 µm filter before analysis using inductively coupled plasma mass spectrometry (ICP-MS) (Agilent 7700x, Santa Clara, CA). Blanks and standard reference material (Environmental Express, Charleston, SC) were used to validate the digestion and analytical method. The calibration curve consisted of the following concentrations: 0, 1, 10, 50, 100, 500, 1000 µg kg⁻¹. All samples either fell within the range of the calibration curve or were diluted to be within the range. Blanks were run every 10 samples. The detection limit was 0.5 µg kg⁻¹. Samples were measured five times and averaged to give an output concentration with an RSD.

## X-ray absorption spectroscopy

After exposure, rinsed roots from two plants were lyophilized, combined, ground and homogenized, pressed into a pellet, and sealed in Kapton® tape. Cerium LIII X-ray absorption near edge structure (XANES) spectroscopy data were collected at the Stanford Synchrotron Radiation Lightsource (SSRL) on Beamline 11-2. Beam energy was calibrated using a Cr foil (5.989 keV). A double crystal monochromator (Si [220], crystal φ = 90) equipped with a harmonic rejector was used in conjunction with a 100-element solid-state Ge detector. Measurements were collected at 77 K using a liquid N₂ cryostat. All scans were energy calibrated, deadtime corrected, and averaged using the SIXPACK software package (v1.2.10).⁵⁴ Scans were then background subtracted, normalized, and fit using linear combination fitting (LCF) using ATHENA (Demeter v0.9.24).⁵⁵ For the purposes of LCF, we assume that the starting materials are all Ce(IV) oxidation state.⁵⁶ Ce(III) acetate (Sigma-Aldrich, St. Louis, MO) was used as a model compound for Ce(III).

## X-ray fluorescence (XRF) imaging and µ-XANES collection and analysis

After exposure, fresh plant leaves were placed between two pieces of 4 µm-thick Ultralene®, which formed a seal around the plant tissue to minimize dehydration. µ-XRF maps and µ-XANES were acquired at National Synchrotron Light Source (NSLS-II) at Brookhaven National Laboratory on SRX (5-ID) for the CeO₂(0) and CeO₂(−) NP exposures and XFM (4-BM) for the CeO₂(+) NP exposures *via* fly-scanning. On SRX, samples were oriented 45° to incoming beam and to a three-element Vortex-ME3 silicon-drift detector. Elemental maps with an incident energy of 14 keV were collected *via* fly-scanning using a step size of 4 µm and a dwell time of 0.1 s, and spectral fitting was performed using the PyXRF spectral fitting program.⁵⁷ On XFM, samples were oriented 45° to incoming beam and to a four-element Vortex-ME4 silicon-drift detector. Large area (>1 mm)
elemental maps with an incident energy of 11 keV were cre-
ated using a step size of 20 µm and a dwell time of 0.2 s, and
spectral fitting was performed using GSE MapViewer in Larch
(v 0.9.40).⁵⁸ µ-XANES were then collected at locations of inter-
est across the Ce L<sub>III</sub>-edge (5.623–5.823 keV) and data analysis
was performed using ATHENA as detailed above.

**Results and discussion**

NP characterization

NPs have previously been characterized by transmission
electron microscopy (TEM), Fourier-transform infrared spectro-
scopy (FTIR), and X-ray diffraction (XRD) by Collin et al.⁵¹ The
primary crystallite diameters, as measured by TEM, are be-
tween 2 and 4 nm. Here, the number-weighted average hydro-
dynamic diameters of the particles in the exposure medium
were 30.3 ± 2.8, 22.9 ± 2.2, and 27.9 ± 2.2 nm for the CeO₂(+),
CeO₂(0), and CeO₂(−) particles, respectively. Volume- and
intensity-weighted distributions and averages are presented in
Fig. S1.† The electrophoretic mobility of the particles in the nu-
trient solution were +1.69 ± 0.50, -0.14 ± 0.50, and -2.48 ± 0.60
µm cm V⁻¹ s⁻¹ for the CeO₂(+), CeO₂(0), and CeO₂(−) particles,
respectively. This corresponds to apparent zeta potentials using
the Hückel approximation of +32.2 ± 9.6 mV, -2.6 ± 8.6 mV,
and -52.3 ± 12.7 mV, for the CeO₂(+), CeO₂(0), and CeO₂(−) par-
ticles, respectively. Generally, a ζ-potential between -10 mV and
+10 mV is considered to be relatively neutral, while values
greater than ±10 mV to be cationic or anionic, respectively.⁵⁹ At
the end of the exposure, <0.1% of the Ce remaining in the ex-
posure solution was dissolved (Table S1†).

Total Ce uptake

The Ce concentrations associated with plant roots and shoots
from the three different treatments are shown in Fig. 1A.

Irrespective of plant species, CeO₂(+) NPs adhered more read-
ily to the plant roots than CeO₂(−) NPs due to electrostatic at-
traction to the negatively charged root surface or repulsion
for the negatively charged particle, which is consistent with
numerous other studies comparing the impact of surface
charge in plants.²²⁻²⁵,³⁸,⁶⁰ Across the plant species, the to-
mato accumulated the most Ce in/on the roots for all NP
treatments, with the highest being from the CeO₂(+) NP treat-
ment (47 300 ± 3100 mg kg⁻¹). Neutral particles had an inter-
mediate degree of interaction.

The dicots generally show more Ce in the shoots than the
monocots (Fig. 1). This trend is consistent with trends ob-
served by Lopez-Moreno, et al.⁶¹ between dicots (alfalfa, to-
mato, cucumber) and a monocot (corn) exposed hydroponi-
cally to 7 nm CeO₂ NP and by Schwabe et al.³³ between a
dicot (pumpkin) and a monocot (wheat) to 9 nm CeO₂ NPs.
With regards to surface charge, corn, rice, and lettuce
followed previously observed statistically significant
trends,²²⁻²⁴ in which plants accumulated higher amounts of
metal in the shoots from the negatively charged NP exposure
compared to the positively charged NP exposure. The tomato
plant, however, followed the opposite trend, with the highest
Ce accumulation from the CeO₂(+) NP treatment and the low-
est from the CeO₂(−) NP treatment. This is likely due to high
accumulation of Ce in/on the roots from the CeO₂(+) NP ex-
posure compared to the CeO₂(−) NP which enabled more Ce
to translocate, albeit less efficiently. The speciation of Ce that
is translocating is discussed later in the paper.

Translocation efficiency was also calculated as a ratio of
total Ce in shoots to total Ce in/on roots to better compare
the capability of different particles to move from the roots to
shoots (Fig. 1B). All plants had the highest translocation effi-
ciencies for the CeO₂(−) NP treatment and the lowest for the
CeO₂(+) NP treatment, further suggesting that the positively

<figure><img src="image_2.png" /><figcaption>**Fig. 1** (A) Ce concentration (mg-Ce per kg of dried plant tissue) on/in dried roots (bottom) and shoots (top) and (B) translocation efficiency (%, Tot Ce<sub>shoots</sub>/Tot Ce<sub>roots</sub>) of corn (yellow), rice (orange), tomato (light blue), and lettuce (dark blue) after 48 h of hydroponic exposure to 50 mg-Ce per L as CeO<sub>2</sub>(+), CeO<sub>2</sub>(0), or CeO<sub>2</sub>(−) NPs. Roots were rinsed for 30 s in Ce-free medium prior to lyophilization and analysis. The means are averaged from four replicates. Error bars correspond to standard deviation. Significant differences [based on ANOVA and Tukey HSD *post hoc* tests (p < 0.05)] between plant species for the same NP treatment for either the roots or shoots are indicated by capital letters.</figcaption></figure>
charged particles adhere too strongly to the root surface to translocate. The CeO₂(−) NP treatment for lettuce had the largest value (24 ± 4%). Regarding the high Ce leaf concentration in the CeO₂(+) NP exposure in tomato, the lower translocation efficiency for the positive treatment than the negative treatment further corroborates the hypothesis that negatively charged particles are able to more efficiently translocate than positively charged particles. The two dicots translocated Ce more efficiently than the monocots for all particle types (though the tomato neutral treatment was not statistically significantly higher). This is likely due to the high transpiration rate in the dicots compared to monocots (see later discussion). The trends in uptake observed here for the NPs follow the trends observed in a field study using soil contaminated with Cd, Pb, Cu, and Zn with ten different plants.⁶² Lettuce and other leaf vegetables were shown to have higher translocation factors than tomato and other fruit vegetables, which were higher than corn and other grains.

Calculated root SA for the corn, rice, tomato, and lettuce were 6.6 ± 1.2 cm², 0.6 ± 0.2 cm², 11.3 ± 3.2 cm², and 1.3 ± 1.0 cm², respectively (Table S2†). There was no correlation between root surface area and Ce root uptake/attachment for the CeO₂(+) NP or CeO₂(−) NP exposure (Fig. 2), emphasizing the importance of this electrostatic attraction/repulsion between the charged NPs and the charged root surface. The roots of dicots generally have greater cation exchange capacities than monocots,⁶³ which likely explains the higher Ce association for the tomato and lettuce compared to the corn and rice for the CeO₂(+) NP treatment. In contrast, higher root surface area correlated with higher Ce root attachment/uptake for the CeO₂(0) NP exposure (Fig. 2), suggesting primarily a sorption interaction when NPs are relatively uncharged.

## Plant response

Physiological measurements of plant health are presented in (Fig. 3). No statistically significant changes in dry biomass were observed for exposed vs. control plants, likely due to the short-term exposure and low Ce dose. No differences between the exposed and control plants were observed for any of the photosynthesis parameters measured for rice, again likely due to the low Ce transport into its leaves compared to the other plants. In contrast, the most significant changes to plant photosynthesis (ΦCO₂, ΦPSII, ETR) and gas exchange (E, gsw) were observed for corn with all NP treatments. Interestingly, similar changes were also observed in the positive NP treatment despite accumulating ~10 times less Ce in the shoots than the neutral and negative NP treatments. We hypothesize that the NPs induce changes to the root water potential, permeability, or conductivity to water. This in turn resulted in a higher stomatal conductance and therefore increased CO₂ uptake and subsequently ΦPSII.

For tomato, increases to ΦCO₂, ΦPSII, E, ETR, and gsw were observed for the CeO₂(0) NP treatment. Negatively charged CeO₂ NPs have been previously reported to boost photosynthesis rates in soil-grown soybean under non-

stressed conditions (soil exposure; ζ-potential = -51.57 mV)⁶⁴ and salt-stressed canola (hydroponic; ζ-potential = -51.8 mV),¹⁸ and boost carbon assimilation rates and ΦPSII *Arabidopsis* plants exposed to salt-stress, heat, and high light (foliar infiltration; ζ-potential = -17 ± 2.7 mV).⁴⁴ Wu *et al*.⁴³ observed almost two times higher colocalization of negatively charged (ζ-potential = -16.9 ± 6.1 mV) than neutral/moderately-positive (ζ-potential = +9.7 ± 1.2 mV) CeO₂ NPs within chloroplasts in *Arabidopsis* leaf mesophyll cells exposed via foliar infiltration. Though the observed increase in plant health was not statistically significant for the CeO₂(−) NP treatment, Ce accumulation from this exposure was almost three times lower than from the CeO₂(0) NP exposure. The reported increases in plant photosynthesis by CeO₂(−) NP were observed in stressed *Arabidopsis* plants experiencing

<figure><img src="image_3.png" /><figcaption>Fig. 2 Correlation between root surface area (SA) and Ce associated with roots of corn (yellow), rice (orange), tomato (light blue), and lettuce (dark blue) after 48 h of hydroponic exposure to 50 mg-Ce per L as CeO₂(+), CeO₂(0), or CeO₂(−) NPs. Roots were rinsed for 30 s in Ce-free medium prior to lyophilization and analysis. The means are averaged from four replicates. Error bars correspond to standard deviation. Raw values are reported in Table S2.†</figcaption></figure>
<figure><img src="image_4.png" /><figcaption>Fig. 3 Measurements of quantum yield of CO₂ uptake (ΦCO₂) and photosystem II quantum yield (ΦPSII), transpiration rates (E), dry biomass, electron transport rate (ETR), and stomatal conductance (gsw) after 48 h of hydroponic exposure to 50 mg-Ce per L as CeO₂(+), CeO₂(0), or CeO₂(−) NPs. The means are averaged from four replicates. Error bars correspond to standard deviation. Asterisks indicate statistically significant differences relative to the control (2-sample t-test; *p* ≤ 0.05, **p** ≤ 0.01).</figcaption></figure>

ROS accumulation whereas in this study, plants were exposed to CeO₂ NPs under normal growing conditions. It is likely that oxidative stress levels in tomato were not high enough for CeO₂ NPs to provide a beneficial impact on plant health through ROS scavenging. In general, the dicots have higher transpiration rates, indicating higher water uptake, which could contribute to their higher Ce uptake.

Negative impacts on ΦCO₂, E, and gsw were observed for the lettuce CeO₂(+) NP exposure. Positively charged CeO₂ NPs (ζ-potential = +32.8 ± 1.0 mV) have been shown to decrease ΦCO₂ and intercellular CO₂ concentration in *Clarkia unguiculata*.⁴⁵ Considering ΦCO₂ and not ΦPSII is impacted, the NPs are likely causing the plant to divert energy for stress response mechanisms rather than the typical plant processes.⁶⁵ Cationic NPs in general have been shown to be more toxic in a variety of cells compared to their neutral or anionic counterparts.⁵¹,⁵⁶,⁶⁶,⁶⁷ Asati et al.⁶⁷ observed that CeO₂ surface charge influenced toxicity in normal and cancer mammalian cell lines: positively charged nanoceria would generally localize in lysosomes and release ROS-generating Ce³⁺ due to an acidic microenvironment, while neutral particles localized in the cytoplasm and remained untransformed and displayed no toxicity. In this study, the highest Ce accumulation in leaves from the CeO₂(+) NP treatment was observed in tomato and lettuce, which were the only treatments that observed decreases to plant health, though this decrease was only statistically significant for the lettuce exposure. Lettuce has also been shown to be more sensitive to CeO₂ NPs compared to cabbage, wheat, cucumber, radish, tomato, and rape.³⁰,⁶⁸

## Ce reduction in roots

No evidence for Ce reduction in/on roots was observed in any plant for the CeO₂(+) NP treatment, while the CeO₂(0) and CeO₂(−) NP treatments show up to ~30% reduction to Ce(III), with the most reduction observed for lettuce. In agreement with these results, previous XANES maps on wheat roots exposed to these same particles show no reduction from the CeO₂(+) NP treatment and ~15% from the CeO₂(0) and CeO₂(−) NP treatments.²² Furthermore, bulk XANES on hydroponically exposed cucumber (ζ-potential = −10 mV) roots have been shown to undergo some reduction to Ce(III) (<20%) (Fig. 4).⁶⁹⁻⁷¹

The exact location of this biotransformation and the mechanisms occurring are still under debate. One hypothesis is that the plant roots are taking up the CeO₂ NPs, after which the NPs undergo reductive dissolution intracellularly to Ce(III), as discussed below. The Ce(III)/Ce(IV) equilibrium mostly involves the atoms on the surface of CeO₂ NPs,⁷²,⁷³ thus we posit that the Ce(III) is not truly dissolved, but rather that Ce(IV) reduction happens at the NP surface, likely as CePO₄. HR-TEM images by Singh et al.⁷⁴ showed no significant changes to average crystal size of CeO₂ NPs incubated in PBS buffer for 72 h, but the XPS and UV-vis spectra suggest the formation of amorphous Ce(III) phosphate at the particle surface. This was further corroborated by Schwabe et al.⁷⁵ who found less released Ce when phosphate was present in the media, indicating that the Ce(III) is not released from the surface but most likely trapped by the formation of CePO₄ on the NP surface. Where majority of the particles remained
<figure><img src="image_5.png" /><figcaption>Fig. 4 Change in Ce oxidation state, presented as pie charts, in root tissue of corn (yellow), rice (orange), tomato (light blue), and lettuce (dark blue) after 48 h of hydroponic exposure to 50 mg-Ce per L as CeO₂(+), CeO₂(0), or CeO₂(−) NPs. Roots were rinsed for 30 s in Ce-free medium prior to lyophilization and analysis. Normalized Ce L III XANES experimental spectra (solid) are presented with LCF fits (dotted). Fitting statistics are provided in Table S3.†</figcaption></figure>

adhered to the root outer surface from the CeO₂(+) NP exposure, no reduction was observed, while Ce reduction was observed in the CeO₂(0) and CeO₂(−) NP exposed roots where more particles were likely internalized to a greater degree (as suggested by translocation efficiencies in Fig. 1B).

However, reductive dissolution at the root surface is also an important mechanism. Plant roots exude a variety of bio-molecules (e.g. organic acids, amino acids) that can promote the dissolution of metal oxide NPs and/or the precipitation of metals.<sup>76,77</sup> It has been proposed that CeO<sub>2</sub> NPs are first reduced then released as Ce(III) with the assistance of reducing substances,<sup>70,78</sup> and then is often precipitated with phosphate.<sup>52,79</sup> Though the particles are stable in solution and do not significantly dissolve (see Table S2†), CeO<sub>2</sub> NP dissolution has been observed in the presence of low molecular weight organic acids,<sup>37,75</sup> and studies have confirmed both CeO<sub>2</sub> NP and Ce(III) ion uptake in hydroponic exposures by radish,<sup>37</sup> and sunflower, wheat, and pumpkin.<sup>34</sup> Additionally, Schwabe *et al.*<sup>75</sup> observed greater solution acidification for the dicot (pumpkin) compared to the monocot (wheat) exposed to CeO<sub>2</sub> NPs.

## Ce distribution in leaves

Both NP charge and plant vasculature affected the distribution of CeO₂ NPs in plant leaves. XFM maps of exposed monocots (corn and rice) are shown in Fig. 5. For corn, the Ce from the CeO₂(−) NP exposure accumulated in parallel lines with Zn, suggesting the Ce is primarily located in the leaf veins.<sup>80</sup> No Ce fluorescence signal was detected in the leaves of plants for the CeO₂(0) NP treatment. For rice, Ce fluorescence signal from the CeO₂(−) NP treatment formed clusters within the leaf veins as ~30% Ce(III). The CeO₂(0) NP

treatment in rice induced a non-uniform distribution of Ce; a large aggregate (100 µm × 30 µm) of Ce was detected outside the vasculature. These results are consistent with our previous results in wheat using the same particles where the neutral treatment resulted in clusters outside of the main vasculature and the negative resulted in Ce accumulation in the

<figure><img src="image_6.png" /><figcaption>Fig. 5 Tri-colored XRF maps of monocot leaves showing Ce (red), Zn (blue), and Cl (green) distribution in corn and rice after 48 h of hydroponic exposure to 50 mg-Ce per L as CeO₂(0) or CeO₂(−) NPs. Ce signal in the leaves exposed to CeO₂(+) NPs was too low for imaging. White boxes indicate where μ-XANES were acquired, with the LCF results presented as a pie chart. Ce signal was too low to acquire μ-XANES for either corn exposure. XRF maps of individual elements and XANES spectra and fitting statistics are provided in Fig. S2–S5.† Scale bar = 200 µm.</figcaption></figure>
veins primarily as Ce(iv) with some reduction (~20%) outside of the vasculature.²²

XFM maps of dicots (lettuce and tomato) are shown in Fig. 6 and 7. Unlike the monocots, where the Ce was located in small clusters or in the vasculature, Ce is found throughout the leaf. Dicots generally have larger airspace volume than monocots,⁸¹ which may have allowed the Ce to spread further out of the vasculature through the leaf. Previous XRF images of tomato exposed to the same CeO₂(−) NP particles for 14 days showed Ce accumulation within the vascular tissue in relatively large foci as ~40% Ce(III) from the CeO₂(−) NP treatment.²³ Similar accumulation in the primary and secondary veins was observed for tomato CeO₂(−) NP treatment (Fig. S9†), though less reduction was observed here (13%), possibly due to the shorter exposure period. Interestingly, similar distinct spots were observed in *Arabidopsis* exposed hydroponically to cationic quantum dots.⁸² In contrast, both the lettuce and tomato CeO₂(+) NP exposures showed minimal Ce in the primary vasculature, instead they have Ce clusters around minor veins, suggesting the Ce migrates out of the vasculature at the end of minor veins and accumulates in the cells at this point of exit. Similar Ce accumulation at the leaf tips and at the ends of vascular bundles was observed in cucumber leaves exposed hydroponically to relatively neutral CeO₂ NPs (ζ-potential = +8.8 mV).⁸³

In the tomato CeO₂(+) NP exposure in Fig. 7, there is evidence of Ce-trichome colocalization (see Fig. S14† for larger images of these regions). Trichomes are involved in various secretory and uptake functions, and it has been proposed

<figure><img src="image_7.png" /><figcaption>**Fig. 6** Tri-colored XRF maps of dicot leaves showing Ce (red), Zn (blue), and Ca (green) distribution in lettuce and tomato after 48 h of hydroponic exposure to 50 mg-Ce per L as CeO₂(0), or CeO₂(−) NPs. White boxes indicate where µ-XANES were acquired, with the LCF results presented as a pie chart. XRF maps of individual elements and XANES spectra and fitting statistics are provided in Fig. S6-S10.† Scale bar = 200 µm.</figcaption></figure>

<figure><img src="image_8.png" /><figcaption>**Fig. 7** Tri-colored XRF maps of dicot leaves showing Ce (red), Zn (blue), and either Ca (green) for lettuce or K (green) for tomato distribution in lettuce and tomato after 48 h of hydroponic exposure to 50 mg-Ce per L as CeO₂(+). White boxes indicate where µ-XANES were acquired, with the LCF results presented as a pie chart. XRF maps of individual elements and XANES spectra and fitting statistics are provided in Fig. S11-S13.† The lettuce map was completed as two scans. Scale bar = 1 mm.</figcaption></figure>

that metal NPs can be excreted through trichomes.⁷ Many types of trichomes have been shown to accumulate internalized or airborne metals as a detoxification mechanism.⁸⁴,⁸⁵ With specific regards to NPs, trichomes have been shown to accumulate various types of NPs, including C-coated nano-FeₓOᵧ,⁸⁶ nano-TiO₂,⁸⁷ and nano-gold.⁶ Thus, the CeO₂ NPs could have been translocated from the roots to the shoots through the vascular tissue before being sequestered in the trichomes of the leaves to be further exuded from the plant.

## Conclusions

Both surface chemistry and plant species have a significant impact on the uptake and distribution of CeO₂ NPs. Positively charged CeO₂ NPs remained primarily adhered to the negatively charged roots *via* electrostatics as Ce(iv), with poor Ce translocation efficiency to the shoots. In contrast, negatively charged CeO₂ NPs accumulated significantly less on the roots but had the highest translocation efficiency. Overall, tomato and lettuce (dicots) were able to translocate Ce more efficiently to the shoots than rice and corn. This correlates with higher transpiration rates, and thus water uptake. Increases in plant photosynthesis were observed in corn plants exposed to CeO₂ NPs of all charges that were accompanied by enhanced stomatal aperture and therefore CO₂ uptake. In contrast some reduction to plant photosynthesis was observed in
plants under CeO₂(+) NP exposure, potentially a result of the different spatial distribution of the CeO₂(+) NPs in the leaves. Once in the leaves, CeO₂(−) remained primarily in the veins, while (0) and (+) formed clusters outside of the vasculature, possibly because of different surface biotransformation in planta (e.g. corona formation, heteroaggregation), and/or differential potential for membrane crossing conferred by charge type and/or density. Further research is needed to understand why NPs with surfaces of different charges showed such a different leaf distribution. All in all these results indicates that, even if influenced by plant morphologies, tuning NP surface charge can allow NPs targeting to different plant compartment following root uptake. The different Ce distribution as a function of particle surface chemistry suggests that NPs may potentially be engineered for targeted, NP mediated delivery of agrochemicals to different plant organs.

## Acknowledgements

This material is based upon work supported by the U.S. National Science Foundation (NSF) and the Environmental Protection Agency (EPA) under NSF Cooperative Agreement EF-1266252, Center for the Environmental Implications of Nano-Technology (CEINT), Nano for Agriculturally Relevant Materials (NanoFARM) (CBET-1530563), and from the NSF Integrated Graduate Education and Research Traineeship Nanotechnology Environmental Effects and Policy (IGERT-NEEP) (DGE-0966227). Parts of this research used the XFM and SRX Beamlines of the 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. A. S. A was supported by a DOE-Geosciences Grant (DE-FG02-92ER14244). We thank J. Thieme for his help running the SRX beamline at NSLS-II. Bulk XAS on root tissue was performed on Beamline 11-2 at the Stanford Synchrotron Radiation Lightsource (SSRL). We thank Jieran Li for synthesis of the NPs.

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