# Nanoparticle Size and Coating Chemistry Control Foliar Uptake Pathways, Translocation, and Leaf-to-Rhizosphere Transport in Wheat

Astrid Avellan, Jie Yun, Yilin Zhang, Eleanor Spielman-Sun, Jason Unrine, Juergen Thieme, Jieran Li, Enzo Lombi, Garret Bland, Gregory Lowry

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## ► To cite this version:

Astrid Avellan, Jie Yun, Yilin Zhang, Eleanor Spielman-Sun, Jason Unrine, et al., Nanoparticle Size and Coating Chemistry Control Foliar Uptake Pathways, Translocation, and Leaf-to-Rhizosphere Transport in Wheat. ACS Nano, 2019, 13 (5), pp.5291-5305. 10.1021/acsnano.8b09781. hal-03708335

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# Nanoparticle Size and Coating Chemistry Control Foliar Uptake Pathways, Translocation, and Leaf-to-Rhizosphere Transport in Wheat

A. Avellan, J. Thieme

To be published in "ACS NANO"

May 2019

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 Size and Coating Chemistry Control Foliar Uptake Pathways, Translocation, and Leaf-to-Rhizosphere Transport in Wheat

Astrid Avellan,<sup>*</sup>,†<sup>●</sup> Jie Yun,<sup>†</sup> Yilin Zhang,<sup>†</sup> Eleanor Spielman-Sun,<sup>†</sup>● Jason M. Unrine,<sup>‡</sup>‡<sup>●</sup>
Juergen Thieme,<sup>§</sup> Jieran Li,<sup>‡</sup> Enzo Lombi,<sup>||</sup> Garret Bland,<sup>†</sup>● and Gregory V. Lowry<sup>*,†</sup>

**ABSTRACT:** Nanoenabled foliar-applied agrochemicals can potentially be safer and more efficient than conventional products. However, limited understanding about how nanoparticle properties influence their interactions with plant leaves, uptake, translocation through the mesophyll to the vasculature, and transport to the rest of the plant prevents rational design. This study used a combination of Au quantification and spatial analysis to investigate how size (3, 10, or 50 nm) and coating chemistry (PVP *versus* citrate) of gold nanoparticles (AuNPs) influence these processes. Following wheat foliar exposure to AuNPs suspensions (~280 ng per plant), adhesion on the leaf surface was increased for smaller sizes, and PVP-AuNPs compared to citrate-AuNPs. After 2 weeks, there was incomplete uptake of citrate-AuNPs with some AuNPs remaining on the outside of the cuticle layer. However, the fraction of citrate-AuNPs that had entered the leaf was translocated efficiently to the plant vasculature. In contrast, for similar sizes, virtually all of the PVP-AuNPs crossed the cuticle layer after 2 weeks, but its transport through the mesophyll cells was lower. As a consequence of PVP-AuNP accumulation in the leaf mesophyll, wheat photosynthesis was impaired. Regardless of their coating and sizes, the majority of the transported AuNPs accumulated in younger shoots (10–30%) and in roots (10–25%), and 5–15% of the NPs <50 nm were exuded into the rhizosphere soil. A greater fraction of larger sizes AuNPs (presenting lower ζ potentials) was transported to the roots. The key hypotheses about the NPs physical–chemical and plant physiology parameters that may matter to predict leaf-to-rhizosphere transport are also discussed.

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**KEYWORDS:** foliar application of nanoparticles, uptake pathways, in planta translocation, root exudation, agriculture, enhanced dark-field microscopy coupled with hyperspectral imaging, X-ray fluorescence mapping

The application of agrochemicals to crops is notoriously inefficient. Up to 95% of applied micronutrients¹ and 99.9% of applied pesticides² never reach their targets and are wasted. The active ingredients can be degraded prior to achieving their effects, accumulate in soil or runoff, causing collateral environmental damage, degrading soils, and wasting the water and energy used in their production and application. Foliar applications of agrochemicals with delivery systems based on engineered nanomaterials (ENMs) could lead to greater efficiency,³,⁴ thus reducing these losses and decreasing environmental risks. Due to properties that their sizes confer, ENMs could be promising for designing foliar-applied

products. Foliar delivery could replace less efficient soil applications for micronutrients and pesticides. It could be better to deliver agrochemicals where and when the plants need them, at the right doses, and with greater bioavailability. This would also decrease application rates and lower the resulting detrimental impacts.⁵

Various studies have shown that foliar application of nanoparticles (NP) can increase plant growth,⁶ protect

<footnote><sup>†</sup>Civil and Environmental Engineering, Carnegie Mellon University, Pittsburgh, Pennsylvania 15213, United States</footnote>

<footnote><sup>‡</sup>Department of Plant and Soil Sciences, University of Kentucky, Lexington, Kentucky 40546, United States</footnote>

<footnote><sup>§</sup>NSLS-II, Brookhaven National Laboratory, Brookhaven, New York 11973, United States</footnote>

<footnote><sup>||</sup>Future Industries Institute, University of South Australia, Mawson Lakes, South Australia 5095, Australia</footnote>
photosynthesis from reactive oxygen species⁷ or improve photosynthesis rate,⁸ enhance plant resistance to disease,⁹ or increase plant biomass⁶ and fruit production.¹⁰ The foliar application of pesticide carriers is also a promising tool for managing plant diseases¹¹,¹² or reducing crop losses in infested soils.¹³ To date, the literature demonstrates that metals, as salts or particules, can be taken up and further translocate through the leaves¹⁴,¹⁵ into the vasculature¹⁶ and be transported to other plant compartments.¹¹,¹⁶⁻¹⁸ The majority of these studies on foliar uptake and mobility of NPs are using metal- or metal-oxide-based NPs that can potentially release ions outside or inside the plants. It thus remains unclear what form of the metals was taken up, if they transformed *in planta*, and if they are translocated as NPs, dissolved metal, or other precipitated species. Furthermore, the fraction of applied metals from NPs that are transported from leaves to roots is generally not quantified, which makes assessing the potential for effective delivery of NPs into the plants vasculature difficult.

The pathways of entrance of foliar applied NPs are not yet clearly understood. Stomatal and cuticular uptake pathways are two potential routes of entrance into the leaves (see Figure 1).

<figure><img src="image_4.png" /><figcaption>Figure 1. Diagram of a wheat leaf cross-section along with the hypothesized routes for NP uptake following foliar exposure. Cuticular entrance may occur through small hydrophilic pores or *via* cuticle diffusion, disruption (direct), or both. The stomatal pathway likely takes place through a passive stomatal loading.</figcaption></figure>

While the stomatal pathway has been the most-investigated route,¹⁴,²⁰ stomatal loading capacity is highly variable and is often reported as the rate limiting step for NPs entry into plants.¹ Stomatal uptake of NPs will likely be limited by the stomata size (ranging from 10 to 100 µm),²¹,²² by the stomatal density on plant leaves, and by stomatal aperture cycles. The significantly greater amount of cuticle area on the leaves compared to stomata area make cuticular uptake potentially a more efficient pathway for NP uptake. Leaf epidermis is covered by cuticle, a waxy and porous layer designed to prevent water loss and protect plants from pathogen infection.¹⁹ While cuticle adsorption of ionic species through hydrophilic cuticular pores has been demonstrated,¹⁶ these pores are small and have a reported size-exclusion limit in the

nanometer range.²³ One study hypothesized that 4–100 nm NPs could cross the cuticle by disruption of the waxy layer,²⁴ and another described fluorescent accumulation from tagged NP (>50 nm) in the epidermis layer underneath the cuticle in areas devoid of stomata.²⁵ These studies suggest that NPs larger than the cuticular pores can cross the cuticle. Knowledge about the physical–chemical NPs properties that would facilitate NP entrance through the cuticle and how they could further transport to other areas in the plants is lacking. One property that could modify NP foliar uptake pathways is its lipophilicity, as previously observed for molecules.²⁶ Different leaf features (as stomata, trichomes, or cuticle) present a broad range of lipophilicity, with stomatal guard cells being most protein-like and hydrophilic or amphiphilic, and leaf cuticle being the most lipophilic. It is thus possible that the amphiphilicity and lipophilic properties of NPs surfaces will play a role into NP uptake through the cuticle.

After crossing the cuticle, NPs will encounter epidermis and mesophyll tissues (see Figure 1). NPs surfaces will likely interact with these tissues and structures and influence NP translocation to the leaf vasculature. Internal leaf cells will present a broad range of functional groups that could interact to the NPs surfaces.²⁷ The capacity of the NPs surface to hydrogen bond with cell wall features will likely modulate their translocation as well. It is then very likely that NP coating may influence their translocation in leaves. Furthermore, several internal leaves features will present different size exclusion limits, e.g. apoplastic transport in the intracellular space between cells, or endocytosis that is required for symplastic translocation of NPs into phloem. For instance, it has been proposed that the route for cellular translocation (apoplastic versus symplastic; see Figure 1) may be influenced by the size and/or the shape of NPs.²⁸ Thus, the size of NPs should also be a critical property that could affect leaf uptake and translocation in the vasculature bundles.²⁹

While the NP size and surface properties are likely to play important roles in their interaction with plant leaves, their uptake into the leaf, and their transport in *planta*, no study has systematically investigated these processes. Here, gold nano-particle (AuNPs) with different sizes (herein referred to as 3, 10, or 50 nm) and surface coatings (citrate versus polyvinylpyrrolidone, PVP) were foliarly applied to wheat. AuNPs were used as a model NP providing good chemical stability, control over size distribution, and low biological background. Impact of NPs size and coating on their foliar uptake, translocation, leaf-to-rhizosphere transport, and plant health was investigated.

## RESULTS AND DISCUSSION

**AuNP Characterization.** AuNPs of different sizes were coated with either PVP or citrate (later named PVP-AuNPs or citrate-AuNPs). Citrate is a small ionic molecule at neutral pH, while PVP is an amphiphilic polymer soluble in water and alcohols (e.g. methanol and ethanol) as well as relatively hydrophobic solvents (1,2-dichloroethane and trichloromethane).³⁰ The primary particle size distribution (measured by TEM, Figures 2A and S1) of the AuNPs were unaffected by their coatings. The average diameters were 2.6 ± 1.0 and 2.7 ± 0.2 nm (for 3-citrate and 3-PVP); 9.5 ± 0.9 and 9.2 ± 1.5 nm (for 10-citrate and 10-PVP); or 53.4 ± 11.4 and 59.1 ± 14.8 (for 50-citrate and 50-PVP). While each particle has a size distribution, these were statistically significantly distinct (Table S1). The hydrodynamic diameters of the PVP-coated particles
<figure><img src="image_5.png" /><figcaption>Figure 2. Gold nanoparticle (AuNPs) size characterization. (A) Primary particle diameter distributions based on TEM images (at least 100 particles; see Figure S1 for TEM images) and (B) hydrodynamic diameter.</figcaption></figure>

were slightly higher than their citrate-coated counterparts (Figure 2B) but not significantly different (Table S1), as also suggested by the UV-vis absorbance of the suspensions (see Figure S2). This is likely due to the higher molecular weight of PVP (MW = 40 000 g mol⁻¹) than the citrate molecules (MW= 188 g mol⁻¹).

The ζ potentials of the AuNPs were all strongly negative, ranging from -31.1 to -69.2 mV (at 10 mg/L in 1 mM KCl and pH 6.8; see Table S1). The interaction of the PVP molecules with AuNPs occurs through the nonbonding electron transfer from the O atoms of the carbonyl groups to the surface of AuNP, resulting in a negatively charged AuNP surface.³¹ Citrate adsorption on the AuNP surface is made through carboxylate groups of the molecules, where free carboxylate groups confer the negative charge on the citrate-AuNPs.³² This weak bond between citrate and AuNPs may be easily desorbed over a range of ionic strength and pH. Both the size and the type of coating influenced the ζ potentials of the AuNPs. Smaller AuNPs had a more-negative ζ potential. Furthermore, for the same size, PVP-AuNP had lower ζ potential than citrate-AuNPs.

The citrate coating induces more hydrophilic character to the AuNPs³³ than PVP that is amphiphilic.³⁴ PVP is indeed often used for its lipophilic properties in drug delivery systems to improve active ingredient dispersion in non-aqueous phases.³⁵ While there is no standard method to measure the hydrophobicity of a NP, 40% and 85% of the 3 and 10 nm PVP-coated AuNP adsorbed at the dodecane-water interface, while <15% of the citrate-coated AuNPs of similar size did (see Figure S3). This PVP-enhanced affinity for the oil phase was not observed for the largest sizes, possibly because of higher sedimentation properties due to their size. These results illustrate that the PVP- and citrate coatings confer different properties to the AuNPs.

## Interactions and Adhesion with Wheat Leaf Surfaces.

The initial interaction of foliar applied NPs will be with the cuticle. For both coatings, the percentage of Au associated with the wheat leaf after washing was higher for smaller sizes (Figure 3A). For equivalent sizes though, PVP-AuNPs induced higher Au association with the leaf than the citrate-coated counterparts. The percentage of Au associated with leaves represents the adhering AuNP, as well as the fraction that potentially entered the plant. Enhanced dark-field microscopy coupled to hyperspectral imaging (EDF-HSI; see Figure S4 for hyperspectral libraries) of the AuNPs deposited on leaves

(Figure 3B) indicates that, before rinsing, AuNPs were homogeneously distributed on the leaf surface, irrespective of their coating, as were the 3 nm after rinsing. The 10 nm PVP-AuNPs, after rinsing, were mainly associated with the cuticle with less PVP-AuNPs co-localized with the stomata guard cells, while 10 nm citrate-AuNPs were more homogeneously distributed on the leaf surface. Larger sizes were washed off easily, and only a few AuNP were detected on the leaf. These results are in accordance with the measured percentages of Au associated with the leaves after rinsing presented on Figure 3A.

It should be noted that while Au measurements on the leaves exposed to 10 nm citrate-AuNPs (Figure 3A) indicated that ~20% of Au mass remained associated with the leaves, only a few AuNPs were detected by EDF-HSI, indicating that (i) the microscope imaging and hyperspectral data treatment did not allow for the detection of some particles (either too small or with a changing hyperspectral signal), (ii) a portion of citrate-AuNPs were relatively rapidly (<2 h) taken up into the leaf and thus was not detected at the leaf surface, or both.

Large NPs are then washed off more easily, as previously demonstrated with different Cu formulations.³⁶ Additionally, these results demonstrate that coating properties affect NP adhesion strength and location on the leaf surface. A higher percentage of PVP-AuNPs remained associated with the leaf after rinsing, and 10 and 50 nm PVP-AuNP seemed to preferentially interact with the cuticle. It is noteworthy that this adhesion does not correlate with the measured ζ potentials, indicating that it is likely not driven by electrostatic interaction. This enhanced adhesion for PVP-AuNPs could be due to enhanced hydrophobic interactions between the amphiphilic PVP-AuNPs surfaces and the lipophilic cuticle. Further investigation is needed to elucidate the impact of hydrophilic-hydrophobic interactions on NPs adhesion to cuticle.

## Uptake Pathways and Translocation to the Leaf Vasculature.
The uptake of 10 nm citrate- and PVP-AuNPs into wheat was assessed at the leaf surface with hyperspectral microscopy and in the leaves by synchrotron micro X-ray fluorescence (μ-XRF) mapping on leaf cross-sections underneath the deposited drop. After 2 h of exposure, all treatments (the deposition of 4 drops of 7 μL was allowed to completely dry) induced a similar homogeneous coverage of the leaf surface (see Figures 3B and 4). After 7 days of exposure to the citrate-AuNPs, a significant amount of AuNPs remained at the surface of the (unrinsed) leaves (Figure 4A,B), suggesting that hydrophilic citrate-AuNPs are not efficiently taken up into the
<figure><img src="image_6.png" /><figcaption>Figure 3. Adhesion of gold nanoparticles (AuNPs) on wheat leaves. (A) Percentage of Au associated with the leaves 2 h after drop deposition and 1 min of rinsing in 10⁻³ M KCl measured by leaf digestion and ICP-MS measurements. Different lower-case letters indicate significant differences between the mean of the groups (ANOVA test followed by Fisher's LSD test for multiple comparisons, *p* ≤ 0.05). (B) Hyperspectral imaging of the leaves before or after 1 min of rinsing. Red pixels are AuNPs detected using their hyperspectral signature (see Figures S4 and S5 for more details; SAM, 0.085 rad).</figcaption></figure>

leaves. The dark-field microscope images and the hyperspectral analysis also indicate the absence of citrate-AuNPs where the stomata are located (Figures 4A and S6), especially for the 3 nm citrate-AuNPs, which may indicate a preferential stomatal uptake. The μ-XRF on cross-sections of dried tissue (Figure 4B) confirmed that citrate-AuNPs accumulated at the surface of the leaf where the AuNP suspension was deposited. Furthermore, no Au was detected inside the leaf underneath the drop, suggesting that any AuNPs that had entered the leaves had already translocated to the plant vasculature. Evidence of the uptake of AuNPs into other compartments of the plant are presented below.

Observations are very different regarding the PVP-AuNPs, for which no NP were detected at the leaf surface after 7 days (Figure 4A,B) or very few in the case of the largest particles, while AuNPs aggregated when drying (Figure S1), and citrate-

and PVP-coated AuNP covered the leaf surface at day 0 (Figure 3). The PVP-coated AuNPs were not detectable after 7 days on the leaf surface (without rinsing) (Figure 4), indicating cuticular uptake of PVP-coated AuNPs for all sizes. It is noteworthy that the size exclusion limit of plant cuticle for polar compounds through hydrophilic cuticular pores is below the nanometer range²³ (Figure 1). Our results indicate that uptake of PVP-coated AuNPs occurs at least partly via disruption and/or diffusion through the cuticle layer, and possibly through stomata as well (Figure 1). Furthermore, the μ-XRF imaging of a leaf cross-section where the drop was deposited (Figure 4B) indicated that while no PVP-AuNPs remained at the surface of the leaf, PVP-AuNPs that entered the mesophyll remained trapped, likely during their apoplastic and/or symplastic translocation to the plant vasculature. In
<figure><img src="image_7.png" /><figcaption>Figure 4. Gold nanoparticle (Au NPs) interaction with leaves after drop deposition. (A) Enhanced dark-field microscopy images of the drop-deposition area on wheat leaves after 7 days of exposure (no rinsing). Pixels in red are the results of hyperspectral analysis showing locations of AuNPs. (B) Sub-micro X-ray fluorescence mapping (incident beam: 14 keV) collected on 5-ID at NSLSII on dried leaf cross-sections underneath the drop deposition (flash-frozen, microtomed, and freeze-dried). Colors represent the fluorescence emission lines on fitted spectra of (i) in red Au-Lα and Au-Lβ, respectively, at 9.7 and 11.5 keV (ii) in green K-Kα at 3.3 keV and (iii) in blue Mn-Kα at 5.9 keV. Mn and K fluorescence were displayed with Au fluorescence to highlight the different leaf structures.<sup>44,45</sup> Ep.: epidermis cells; p., parenchyma cells; v., vasculature bundle. Optical images of the cross-section can be found in Figure S7.</figcaption></figure>

contrast, 10 nm citrate-AuNPs accumulated at the leaf surface but were not present in the mesophyll.

There are several possible explanations for the less-efficient translocation of AuNPs coated with PVP in comparison to the citrate-coated AuNPs. First, both the surface amphiphilicity³⁷ and charge density³⁸ have been shown to influence NP capacity to cross lipid membranes, which will likely influence the potential for citrate- *versus* PVP-coating to interact with cell membranes and thus modify NP mobility. Second, it is likely that when crossing plant biological barriers and membranes, AuNP surfaces will acquire additional organic coatings,³⁸,³⁹ including proteins, lipids, and carbohydrates present in phloem. This corona formation may change the AuNP's potential for aggregation as well as cellular uptake, trans-

location, and effects.³⁹ Such effects have been, to date, largely unexplored. One has found the presence of an organic coating around TiO₂ NPs in plant mesophyll after leaf exposure.²⁴ The ability of organics to bind to AuNPs surface will not be similar for citrate versus PVP-coated surfaces.⁴⁰ Citrate weakly associates with the AuNP through carboxyl bonding that may be readily displaced by other molecules including thiols, amines, larger-molecular-weight polymers, antibodies, and proteins.⁴¹,⁴² Citrate also contains free carboxyl groups that can serves as a bridge and bind to proteins. In contrast, PVP will associate more strongly with the AuNP surface because of its carbonyl bonding to Au surface and its high molecular weight.⁴³ All of these hypothesis could explain the difference of fate of PVP- *versus* citrate-coated AuNP in the mesophyll.
<figure><img src="image_8.png" /><figcaption>Figure 5. Au translocation on wheat leaves grown hydroponically and foliarly exposed to 10 nm citrate-AuNPs (A, top) or 10 nm PVP-AuNP (B, bottom) for 12 or 36 h. The different panels present a. X-ray fluorescence mapping (incident beam 15 keV) (μ-XRF) of the Au-Lα fluorescence emission lines (centered in 9.7 keV) of fitted spectra. The leaves were fresh (no rinsing) and mounted between ultralene layers, still attached to the plant during and in between the scans (see Figure S9 for a picture of the setup). Scans were made at the exact same locations and μ-XRF were thresholded the same way. b. Average Au-Lα fluorescence counts every 20 μm over the transversal box presented in panels a after 12 h (red) or 36 h (black) of exposure. c. Correlation plots between the Au fluorescence surfaces presented in panel b and their Zn fluorescence counts (Zn Kα centered at 8.7 keV of fitted spectra). All of the correlation plots are scaled equally. Blue dotted line represents the threshold above which Zn is mainly present in the leaf veins (see Figure S8). The pixels due to background contribution were identified from panel b and removed from the correlation plots. Fluorescence counts were normalized by 10 prior to plotting. μ-XRF acquisition were done with similar slits aperture and distance of the detector.</figcaption></figure>

Altogether, these results indicate that citrate-AuNPs remained largely on the cuticle outside of the leaf, with some

uptake, potentially preferentially occurring through stomata. In contrast, the PVP-AuNPs were able to cross the cuticle more
<figure><img src="image_9.png" /><figcaption>Figure 6. Au translocation in wheat leaves after 48 h of foliar exposure to 10 nm citrate and 10 nm PVP-AuNPs. The leaves and roots were harvested (no rinsing) and mounted fresh between Kapton and Ultralene with no further treatments. X-ray fluorescence maps (incident beam: 14 keV) collected on 5-ID at NSLS-II are presented, along with high-resolution maps (panels a–e). Colors represent the fluorescence emission lines of (i) in red Au-Lα and Au-Lβ, respectively, at 9.7 and 11.5 keV (ii) in green Ca-Kα at 3.7 keV and (iii) in blue Zn-Kα at 8.7 keV. Ca and Zn fluorescence were displayed with Au fluorescence to highlight the different leaf structures. All of the color thresholds between images are identical.</figcaption></figure>

efficiently. After entering the plant leaf, PVP-AuNPs remained trapped in the leaf mesophyll, but the citrate-AuNPs were not, likely due to differences in coating affinity with cell structures and macromolecules that will modulate the AuNPs reactivity at the bionano interface. Based on these uptake and translocation results, different transport behavior in other plant compartments were expected and are presented below.

**Leaf Mesophyll Translocation vs Accumulation of AuNPs.** The time scale for the *in vivo* translocation of foliar applied 10 nm AuNPs was studied with μ-XRF on fresh tissues (Figures 5). Plotting Au fluorescence counts over a transverse section of an exposed leaf area (Figure 5b) indicates that while the Au content in leaves exposed to 10 nm citrate-AuNPs decreases uniformly after 12 and 36 h of exposure, this trend is not as clear for 10 nm PVP-AuNPs. The average Au

fluorescence counts over the transverse section identified in Figure 5b were plotted against the Zn fluorescence (Figure 5c). Because Zn concentrations are higher in the leaf vasculature,⁴⁶ Au co-localized with high Zn counts can be assumed to be primarily located in the plant vasculature (above a given count threshold, Zn is only found in the vasculature; see the blue dotted lines in Figure 5). These Au-Zn correlation plots indicate that 94% of the 10 nm citrate-AuNPs were present in vein tissue, while this was the case for 64% of the 10 nm PVP-AuNPs (see Table S2 for calculations).

These results also indicate that 10 nm citrate-AuNPs could be moving at a higher rate at the early time of exposure. Indeed, between 12 and 36 h of exposure, the Au signal from the selected region of interest decreased by 31% versus 18% for 10 nm citrate-AuNPs and 10 nm PVP-AuNPs, respectively.
<figure><img src="image_10.png" /><figcaption>Figure 7. Gold nanoparticle (AuNPs) transport in wheat. A. Au mass balance in wheat 7 days after foliar exposure to 10 µg/l Au suspensions (4 × 7 µL drop deposition). Au percentage is calculated based on the mass of Au added in the drop. B. Differential accumulation of gold in roots (root tissues + adhering soil) relative to the Au mass in shoots. Different small-case letters indicate significant differences between the mean of the groups (ANOVA test followed by Fisher's LSD test for multiple comparisons, *p* ≤ 0.05).</figcaption></figure>

Furthermore, the Au from 10 nm citrate-AuNPs sprayed on leaves are mainly (~95%) moving through the leaf vasculature when 30% of the 10 nm PVP-AuNPs moves through other compartment of the leaves with low Zn concentration, likely between the cuticle and the epidermis, through the mesophyll, or both, as highlighted in Figure 4. It is noteworthy that these AuNP movements were investigated for short time points after the exposure, the time at which the translocation is likely still occurring. While similar time frames were reported for the transport of ionic Zn species,⁴⁷ the maximum rate of translocation of Pb NPs in leaves was reached after several days.⁴⁸

The AuNPs were translocated both upstream and downstream of the drop deposition as well as to the roots (Figure 6). Consistent with leaf cross-sections, Figures 4B and 6d suggest that 10 nm citrate-AuNPs did not accumulate close to the drop deposition, indicating an efficient translocation of the fraction of the citrate-AuNP taken up. In contrast, PVP-AuNPs accumulated in and near the vasculature bundle (Figure 6e). AuNPs then further translocated in the leaves, and, for both coatings, AuNPs were detected in the tips of the exposed leaves (Figure 6) and at the base of some trichomes (Figure 6a-c).

Recent studies have shown the importance of trichomes for the adsorption of ionic species on leaf surfaces,¹⁶ and metal accumulation has been observed at the base of trichomes after root exposure to nanoparticulate⁴⁹ and ionic metal exposure.⁵⁰,⁵¹ The present study indicates that similar accumulation of NPs on the bases of trichomes can be observed after a foliar exposure. Furthermore, μ-XRF indicates that the PVP coating induces Au accumulation co-localized with the leaf vasculature bundle, while citrate coating did not (Figure 6d,e). However, either coating enabled AuNPs to translocate to plant compartments away from the drop-deposition zone over a short time scales. The amounts and locations of transported AuNPs are discussed later.

**Leaf-to-Rhizosphere Transport.** The overall translocation of AuNPs was assessed by measuring Au concentrations in

different parts of the plants, e.g., in the exposed leaf tips and in younger or more mature leaves than the exposed one, as well as in plant roots, and rhizosphere soil adhered to the roots (Figure 7A,B). Although differences in transport through the mesophyll and vasculature were observed between the different NPs coatings, coatings did not strongly affect AuNP translocation to other parts of the plant (see Figure S10 for statistical analysis). A similar accumulation of Au (~10–25%) was found in plants roots, regardless of the AuNP coating, for similar sizes. This indicates that despite the impact of the coating for the first steps of entry into the plants (adhesion, uptake, and transport to the vasculature), AuNPs that reach the vasculature were similarly translocated to other parts of the plant, possibly because this mobile fraction has been coated by an organic corona, as hypothesized earlier. One observed difference induced by the different coatings was that PVP-AuNPs of <50 nm accumulated in mature leaves, i.e., older than the exposed leaves, while citrate-AuNPs did not, possibly because of the above observed higher interaction of PVP-AuNPs with the plant leaf structures.

A significant fraction of the dosed Au (between 10 and 20% depending on the treatments) was found in plant roots. The speciation of Au in the roots of the treatment leading to the highest Au accumulation (50 nm citrate and 50 nm PVP) indicates that the Au in roots was 100% metallic Au (see Figure S11), suggesting the transport of Au as intact AuNPs to the roots. The partial dissolution of AuNPs on the leaves, followed by ion transport to the roots and subsequent reduction of ionic Au back to metallic Au, cannot be strictly ruled out. However, this mode of transport is unlikely given the relative stability of AuNPs in oxic environments⁵² as well as the absence (in our controlled system) of a source of complexing agents⁵²,⁵³ that could induce AuNP oxidative dissolution in wheat (e.g., cyanide).⁵⁴ Au was also detected in the rhizosphere soil adhering to the roots for AuNPs of a nominal size smaller than 50 nm, regardless of their coating, indicating that AuNPs can not only reach plant roots but also be exuded into the surrounding soil. It is noteworthy that, in wheat, 10–15% of
<figure><img src="image_11.png" /><figcaption>Figure 8. Impact of foliar applied gold nanoparticle (AuNP) suspensions and Au ions on plant health (applied Au was 280 ng per exposed leaf). The proxies used were the plant dry biomass as well as respiration and photosynthesis traits. Stars indicate significant differences relative to the deionized water (DIW) control (ANOVA test followed by a Dunnett post-test for two-sided comparisons). Single asterisks indicate *p* ≤ 0.05, and double asterisks indicate **p** ≤ 0.01 (relative to the control treatment).</figcaption></figure>

the photosynthetically fixed carbon is transferred into the rhizosphere through root exudates.⁵⁵ Coincidentally, about 15% of the applied Au NPs were also exuded into the rhizosphere soil, suggesting that NP transport and transport of fixed carbon may be related.

The largest AuNPs were not exuded into the soil, even though the concentration of Au associated with the roots was similar, possibly due to sieving by the root cells preventing exudation. The overall ratio of Au accumulation in roots and adhering soil relative to the shoots (Figure 7B) confirmed that the coating did not play a significant role in the AuNPs translocation from wheat leaves to roots. A greater fraction of the larger sizes (10 and 50 nm) were transported to the roots compare with the smallest sizes and the ionic Au. This transport correlated to the number of NPs applied (see Figure S12A), as previously observed for root uptake by tobacco of AuNPs.⁵⁶ However, while a greater number of the smallest AuNPs were transported, a higher mass fraction of the applied AuNPs reached the root system for larger sizes. Size may not be the only parameter explaining for this increased translocation. Plotting the rate of AuNPs reaching the root system against the ζ potential of each AuNP (Figure S12B) indicates that AuNP transport may be facilitated by lower surface charges. One way this differential transport could happen is through differential cell uptake for NPs of different sizes and charges.⁵⁷ Previous research on the influence of NPs diameter

on their endocytosis in mammalian cells indicated that larger sizes (approximately 50–60 nm in diameter) induced higher cellular uptake through endocytosis.⁵⁸ Although endocytosis in plants serves different purposes than in mammalian cells, it is possible that a similar phenomenon is observed here. Furthermore, it has been demonstrated that passive diffusion through lipid layers (cell membranes) was enhanced for NPs presenting higher charge densities.³⁸

While previous studies have shown translocation of metal from poorly soluble foliar applied NPs to roots,¹⁷,⁵⁹,⁶⁰ or the exudation of Cu into hydroponic systems,⁶¹ none, to the best of our knowledge, reported the exudation of foliar applied NPs into rhizosphere soils. It is noteworthy that phloem loading and transport can be metal- and species-dependent,⁶² and thus, these transport potentials could be different across different metal-core NPs and plant species. Finally, root exudates have been shown to shape the rhizosphere soil bacterial community structure⁶³ and plant–microorganism interaction.⁶⁴ It is thus likely that root exudation of foliar-applied compounds could impact the root–rhizosphere microbiome, especially for NPs that are antimicrobial such as Cu or Ag.

**Treatment Effects on Plant Photosynthesis.** The impact of these AuNP treatments on plant health were assessed by measuring the dry mass of the plant as well as different respiration and photosynthesis traits of the leaves exposed to 280 ng of the different Au treatments for 3 and 7
days (Figure 8). The photosynthesis rate represents the quantity of assimilated CO₂ used for the photosynthesis. The stomatal conductance is the rate of water vapor exiting the stomata of the leaf. Photochemical versus nonphotochemical quenching ($q_p$-to-$q_N$ ratio) represent the fraction of photons used for photosynthesis or as a photoprotective mechanism preventing formation of highly reactive forms of oxygen. While the overall decrease of all traits in all plants (including the control plant) between 3 and 7 days is due to natural leaf aging, treatments impaired plant photosynthesis differently.

Overall, the ionic Au treatment impaired plant net photosynthesis the most, which correlated with the appearance of cell necrosis at the drop-deposition area. The citrate-AuNPs increased the stomatal conductance and the photosynthesis rate of the exposed leaves, correlating with a statistically significant gain of dry biomass for shoots (3 and 10 nm) and roots (10 and 50 nm) of the exposed plants. PVP-AuNPs (especially the larger sizes) had an opposite effect, showing a tendency to lower the net photosynthesis rate, the stomatal conductance, and the $q_p$-to-$q_N$ ratio after 7 days but did not modify the respiration traits much at that time point. This suggests that PVP-AuNPs induced a stress that disturbed the photosynthesis system of the exposed leaves, which is likely to be the consequences of PVP-AuNP accumulation in the leaf mesophyll (Figure 4B). This impairment of the photosynthesis system for the PVP-coated AuNPs could be due to their amphiphilic properties. Amphiphilic coatings are known to increase the potential for NPs to cross the phospholipid membranes of protoplasts and chloroplasts,⁶⁵ potentially interacting more directly with the thylakoids of the chloroplasts, where light-dependent photosynthesis reactions take place. Indeed, AuNPs has been shown to be able to quench chlorophyll fluorescence.⁶⁶ The citrate (more-hydrophilic)-AuNPs may not reach the chloroplasts as much. Furthermore, a previous study reported that foliar application of AuNPs can increase the leaf chlorophyll content; accelerating photochemical reactions and improving CO₂ fixation.⁶⁷ A similar mechanism explaining the beneficial impacts of the citrate-coated AuNPs may be observed here. These tendencies are statistically significantly different from the control for the 50 nm PVP-AuNPs treatment, even 7 days after exposure. However, these impairments of the photosynthesis systems had no significant impact on the dry biomass of the exposed plants.

Though similar amounts of Au were transported in plant compartments regardless of coating, different plant health impacts were observed. This suggests that plant health impacts depend on where, and at what rates, the AuNPs were accumulating in and on the plant leaves. citrate-AuNP, mainly accumulating on the outside of the plant leaves had overall neutral or beneficial impacts, primarily on respiration traits, while PVP-AuNPs, crossing the cuticle layer and interacting with (in or on) the mesophyll cells were detrimental for plant photosynthesis.

## CONCLUSIONS

This study demonstrates that the chemical properties of the NP's coating plays a major role on the interaction of AuNPs with wheat leaves, their path of entry, and the subsequent health impacts. The size of the AuNPs affected their adhesion on the leaves. The relatively more hydrophobic PVP coating increased the interaction of AuNPs with the leaf surface. It also influences the ability of AuNP to cross the cuticle, enter the

mesophyll, and move through the plant vasculature. These differences in AuNPs accumulation in leaf tissues underneath the drop deposition correlated with different plant health responses, with PVP-coated AuNPs impairing leaf photosynthesis.

While the coating chemistry plays a role for the adhesion, uptake, and translocation of AuNPs, their transport to different plant compartments was similar, regardless of their coatings. The partitioning of AuNPs in wheat roots relative to the Au present in the shoots was impacted by AuNPs primary size, with a lower fraction of small NPs (ionic or 3 nm AuNPs) reaching the wheat root system. It was not clear here if this behavior should be attributed to AuNPs size, their surface charge, or both. The fact that similar transport was observed for a given size (for the range of sizes studied) suggests that AuNP transport may be regulated by a force that limits the vasculature loading and transport. These limiting steps could be (i) the rate of generation of products from the photosynthesis entering the phloem and their flux, (ii) an optimum size (at the larger end)⁵⁸ or charge density³⁸ for objects' endocytosis, (iii) the formation of a corona around the NP (made of lipids, proteins, or carbohydrates) while crossing biological barriers and compartments that may facilitate their transport, or a combination of these. Finally, while all AuNPs reached the root systems, the biggest (>50 nm) AuNPs were not detectable in soil, suggesting a root-sieving step that is preventing the exudation of larger objects.

This study improves our understanding of NP-leaf interactions and of the potential impacts of foliar applied NPs. Altogether, these results indicate that larger objects than previously thought (up to 50 nm) can enter plant leaves by cuticular uptake pathways. The foliar application of NPs with appropriate sizes and coating properties can enable targeting to specific plant compartments (e.g., adhering to the leaf surface, accumulating in the mesophyll, and delivery to the roots or the rhizosphere soil) in more-efficient ways than applications of ionic or chelated metal species. These could lead to alternative designs and uses of agrochemicals, potentially (i) reducing the amount of chemical needed, and thus environmental pollution and side effects by agrochemicals better reaching their target, or (ii) increasing the bioavailability of poorly soil-bioavailable nutrients.

These results also raise questions about unintended side-effects. It is likely that corona formation around the NPs happens in planta, e.g., with proteins or lipids when crossing cell membranes or with polysaccharides when entering the phloem. It remains unclear what coronas may form and how it will be influenced by different coatings. The influence of the corona on NP fate in plants, e.g., aggregation, biotransformation, and impacts, is also unknown. Furthermore, giving the importance of symbiotic relationships of plants with microorganisms, it remains unknown how foliar applied NPs, especially those known to be antimicrobial, could impact plant microbiome (phyllosphere, endophytic, and rhizosphere) and, thus, plant functions. Investigations are also needed to understand the mechanisms of cuticular uptake. It is unclear what coating properties enable cuticular uptake, or if disruption of the waxy cuticle layer is needed for efficient uptake. Better understanding the interaction between NPs of different coating hydrophobicities and cuticles of different composition is needed to better predict NP foliar uptake. Finally, the location of the NP suspension drop deposition was fixed, but exposing different parts of leaves at different
maturities may have an impact on NP adhesion, uptake and mobility, as differences in leaf surface properties, such as stomatal density or cuticle thickness, can change with age. These could also be influenced by the plant species and the conditions in which they were grown. Finally, because different metals will present different solubility, phloem loading, or plant homeostasis and regulation, there is a need to understand if similar coatings on different metal-core NPs will induce similar translocation in plants with different metal-uptake and transport strategies.

## MATERIALS AND METHODS

**AuNPs Synthesis and Characterization.** *Synthesis of 10 nm Citrate-Coated AuNPs.* The 10 nm Au-NPs were synthesized following the Turkevich method.⁶⁸ A pair of solutions of 100 mL of 1 mM HAuCl₄ and 0.2 M sodium citrate solution were prepared. HAuCl₄ was heated and boiled with reflux and magnetic stirring. After reflux started, 2 mL of citrate solution was added all at once. The reaction was continued under reflux and vigorous stirring for 20 min. The resulting gold colloid solution was removed from heat and stirred continuously until cooled to room temperature.

*Synthesis of 50 nm Citrate-Coated AuNPs by the Seeding Approach.* The procedure for the synthesis of 50 nm AuNPs was modified from previous 40 nm AuNP synthesis method.⁶⁹ A total of 1 mL of 200 mg/L 10 nm AuNP was mixed into 94.5 mL of DI water to prepare a AuNPs seed solution. A total of 1 mL of 40 mM sodium citrate solution was added to the AuNP seed solution under stirring and heating. As the reflux started, 0.5 mL aliquots of 0.01 M HAuCl₄ solution were added every 15 min until a total volume of added HAuCl₄ reached 7.5 mL (15 aliquots). The reaction was continued with reflux and stirring for 15 min under heat and removed from heat and continued stirring until cooled.

*Surface Modification of 10 and 50 nm Citrate-Stabilized Au-NPs.* The citrate coating on surface of AuNP was replaced by the strong stabilizer polyvinylpyrrolidone.⁷⁰ The surface modification was realized by adding 0.05 g of PVP into 50 mL of citrate-stabilized Au-NPs and stirred for 4 days. After that, the 10 nm nanoparticles were centrifuged at 10500g for 90 min and the 50 nm nanoparticles at 6000g for 15 min and resuspended in DI water to remove the free citrate in solution.

*Citrate-Stabilized 3 nm Au-NPs.* The 3 nm Au-NPs with citrate coating were synthesized according to the procedure reported by Jana et al.⁷⁰ A total of 20 mL of aqueous solution containing 0.25 mM HAuCl₄ and 0.25 mM sodium citrate was prepared. The citrate only serves as stabilizer for this colloid solution. Concurrently, an ice-cold 0.1 M NaBH₄ solution was prepared, and 0.6 mL of NaBH₄ solution was injected with vigorous stirring in the growth solution. The solution turned orange immediately, and stirring was continued for 15 min for the reaction to be complete.

*PVP-Stabilized 3 nm Au-NPs.* The PVP-stabilized AuNPs were also synthesized by the NaBH₄ reducing method but employed PVP as stabilizer. First, a 0.4 g/L PVP (MW = 40 000, Aldrich) solution was prepared. Then, a water solution with 19 mL of DI water, 0.5 mL of 0.01 M HAuCl₄, and 0.5 mL of PVP solution was prepared. An ice-cold 0.1 M NaBH₄ solution was prepared simultaneously. A volume of 0.6 mL of ice-cold NaBH₄ solution was added into the growth solution under fast stirring. The solution turned orange immediately, indicating the formation of small size Au-NP, and stirring was continued for another 15 min for the reaction to be complete.

*Nanoparticle Characterization.* Hydrodynamic sizes and zeta potential were measured on AuNPs suspensions of 10 mg/L in 10⁻³ M KCl (pH6.8), using a Zetasizer Nano ZS (Mavern Panalytical). A 7 µL drop of a 10 mg/L AuNP suspension was drop-deposited onto a Cu grid with a carbon film and allowed to dry. TEM measurements were done using a JEOL JEM-2000EX operating at 200 keV. The absence of solubilization of the AuNPs was checked based on the measurement of ionic Au by inductively coupled plasma mass spectrometry (ICP-MS) following a 3 kDa filtration. Relative hydrophobic properties of the AuNPs was characterized by AuNP

dodecane-water partitioning. A total of 4 mL of AuNPs suspension in water with different coatings and sizes were probe-sonicated for 30 s at high energy with 4 mL of deodecane and allowed to settle for 24 h. Au concentrations in the water phase were measured by ICP-MS before sonication and 24 h after sonication.

**Seed Germination, Plant Growth, and Nanoparticle Exposure.** Wheat (*Triticum aestivum* cv. cumberland) seeds were obtained from David Van Sanford at University of Kentucky. Seeds were surface-sterilized using 10% w/v bleach (VWR Analytical), gently shaken for 10 min, rinsed three times with DI water, and allowed to soak in DIW for 30 min. Seeds were then placed on DI water-moistened towel paper in a Petri dish and kept in the dark at 25 °C and humidity of 60% for 5 days of germination.

Seedlings, for total metal measurements, plant health assessment, as well as hyperspectral microscope analysis, were transferred to ~15 cm diameters pots filled with the standardized soil Lufa 2.2 and grown in a growth chamber with a light/dark cycle of 16 h:8 h 25 °C: 21 °C and 60% humidity. Pots were randomized, and positions were changed every other day. The pots were watered as needed with DI water. Seedlings for µ-XRF mapping were grown hydroponically in one-fourth strength Hoagland's medium.⁷¹

DIW or 10 mg-Au/L suspensions (HAuCl₄, 3 nm citrate, 3-PVP, 10 nm citrate, 10 nm PVP, 50 nm citrate, and 50 nm PVP) were applied on 14-day-old plants (3-leaf stage). Au-NP solutions were centrifuged and resuspended in DIW prior to their use to ensure for the absence of remaining free coating. Application were done on the second leaf, with the suspensions being either deposited on four drops of 7 µL (estimated dose: 280 ng) along the central vein except for one measurement in which 30 µL of solution were sprayed on the tip of the leaf (see Table S3) using 2 mL glass-containers sprays. Exposed leaves were allowed to dry before the plants were moved back in the growth chamber.

For total metal measurement, plants were allowed to grow 7 days post-exposure. To measure for Au distribution in tissues using µ-XRF, plants were allowed to grow for 12, 36, or 48 h post-exposure (see Table S3).

**AuNPs Adhesion, Uptake, and Translocation Measurements.** *Sampling and Plant Growth Measurement.* The whole shoots were separated from the root 1 cm above the ground. Leaves were thus separated into the following categories: tip, exposed, young shoots (the shoot above the exposed leaf), mature shoots (the shoot part below the exposed leaf), and the rhizosphere soil strongly adhering to the roots. In addition, the roots, quickly rinsed in 10⁻³ M KCl, were also collected. All of the plant tissues were dried at 105 °C for 72 h before dry weight measurement and acid digestion.

*Adhesion Measurements.* To measure for the adhesion of the NP to the plants, the exposed leaf was cut off, rinsed in a 15 mL tube filed with 10⁻³ M KCl in a vial under agitation for 1 min, dried at 105 °C, digested, and analyzed with ICP-MS for Au masses, as described later.

*Elemental Analysis.* Dried plant tissue samples were digested overnight at room temperature in a 2:1 ratio of concentrated HNO₃ and 30% H₂O₂ and then heated at 95 °C for 30 min. After cooling, HCl was added to match a 2:1 HNO₃/HCl ratio and heated again at 95 °C for 30 min. Following digestion, the samples were diluted to reach 5% HNO₃/2.5% HCl using DI water and filtered using 0.45 PTFE filters. NP suspensions and their dissolved fraction (filtrate of a 3 kDa filter, Amicon) were digested using HNO₃/HCl 3:1 and diluted to 2% HCl and 6% HNO₃.⁷² Au analysis was made using ICP-MS (Agilent 7700x, Santa Clara, CA). Calibration was performed across a range of 2–300 µg/L. Calibration curves presented an R value of >0.999 and all relative standard deviations of measurements were below 5%. The method limits of detection were calculated as the mean of 3 blank samples plus 3 standard deviations of 10 replicate analyses of blank sample and was 1.1 µg/L for Au. Measurement drift was checked by analyzing 2 calibration points and a blank every 12 samples.

**Plant Health Measurement.** Gas-exchange parameters and photosynthesis efficiency were determined using a Li-6800 portable photosynthesis system (LI-COR Biosciences, Lincoln, NE), 3 and 7 days after exposure. The measurements were done on the zone the
AuNPs were deposited. The parameters measured during the gas exchange measurements included net CO₂ assimilation rate (A), intercellular carbon dioxide concentration (Ci), transpiration rate (E), and stomatal conductance (Gs). During the measurements, the chamber was set to a flow speed of 500 µmol/s, humidity at 60%, CO₂ concentration at 400 µmol/mol, fan speed of 10 000 rpm, and photosynthetic photon flux density (PPFD) at 600 µmol/(m²·s). Measurements were performed between 10 a.m. and 2 p.m. to ensure for a homogeneous plant response between treatments. Leaves were left for 3 min to stabilize in the LI-COR chamber before each reading. For the photosynthesis traits, maximum quantum efficiency (Fv/Fm) was measured on dark-adapted leaves (kept 30 min in the dark before measurement), and photochemical quenching (qp) and non-photochemical quenching (qN) were measured on light-adapted leaves. For the calculation of all these traits, fluorescence of the leaf was measured before, during, and after a rectangular flash of saturated light (red target 8000 µmol/(m²·s)) during 1000 ms). For dark-adapted measurement, the dark modulation rate was set at 50 Hz. For light-adapted leaves, PPFD was set to be 600 µmol/(m²·s), and the light and flash modulation rates were set up at 50 and 250 kHz, respectively. Leaves were allowed to stabilize for 3 min in the LI-COR chamber before each fluorescence measurement.

**Fresh Leaf Imaging and Hyperspectral Mapping.** Fresh leaves were cut and directly mounted with DIW between glass-slide and a coverslip. The imaging was done using an enhanced dark-field microscope (BX51, Olympus) coupled to a hyperspectral imaging system (Cytoviva Inc.). This microscope was equipped with a 150 W halogen light source (Fiber-Lite, Dolan-Jenner), and a hyperspectral camera (CytoViva Hyperspectral Imaging System 1.4). The leaves were observed at a 10× magnification or in oil immersion at a 60× magnification. Hyperspectral images were acquired using 75% light source intensity and 0.1 to 0.25 s acquisition per line. The hyperspectral libraries were built using images of AuNPs in suspension following steps of spectral data reduction and end-member identification as described earlier$^{72,73}$ as well as in Figure S4. These hyperspectral libraries were used to map the dosed leaves. A spectral angular mapping algorithm (SAM, ENVI 5.2) was used to identify the pixels matching the AuNPs hyperspectral library on the first 185 bands of the images (angles of ≤0.085 rad were considered similar). Each pixel identified that way was highlighted in red. All the hyperspectral images were acquired at cross-section focus. Because of the narrow depth of field (less than a micrometer), signals of Au-NPs were only mapped by SAM in the focus plane shown on the pictures, and out-of-focus Au-NPs adsorbed on top or under the focus plane were not mapped.

**XRF for Element Mapping on Fresh and Dried Tissues.** Following leaf exposure (see Table S3), fresh wheat leaves were sealed between two pieces of 4 µm thick Ultralene to prevent dehydration, while the plant was still growing (see Figure S10 for a picture of the setup). Leaves were scanned at the XRF beamline at the Australian Synchrotron, where an in-vacuum undulator is used to produce an X-ray beam. A Si(111) monochromator and Kirkpatrick-Baez (KB) mirror microprobe are used to obtain a monochromatic beam focused onto the specimen (approximately 2 µm × 2 µm).$^{74}$ The X-ray fluorescence emitted by the specimen was collected using the 384-element Maia detector (Rev C) placed in a back-scatter geometry.$^{75}$ Scans were completed at 15.8 keV. Spectral fitting and data treatment was performed using Geopixie software.

X-ray fluorescence mapping (XRF) images at higher resolution were acquired on fresh leaves (mounted on Ultralene) or dried cross-section. For dried cross-sections, the exposed leaves were harvested, cut, and immediately flash-frozen in cold 2-methylbutan. After being embedded into OCT, the tissues were cut using a cryostat in 30 µm thick slices and mounted in Kapton tape. Slices were then freeze-dried at -53 °C and 0.024 mbars for 48 h. XRF were acquired at Beamline S-ID of the National Synchrotron Light Source II (NSLS-II) at Brookhaven National Laboratory. This beamline uses Kirkpatrick-Baez mirrors to produce a focused spot size of 0.5 µm × 0.5 µm of hard X-rays with tunable energy selecting a 14 keV energy. Samples were oriented at a 45° to the incoming beam and to a three-element

Vortex ME3 silicon drift detector. Coarse elemental maps were collected via fly scanning using a step size of 2.5 µm for coarse maps or 1 µm for fine maps on fresh tissue or 0.1 µm on fixed tissue with dwell times of 0.2 s for the fresh tissue or 0.5 s for the dried fixed tissue. Spectral fitting was performed using the PyXRF spectral fitting program.$^{76}$

The tissues presenting both enough mass of tissue and sufficiently high gold concentration (~10 mg/kg), allowing for bulk X-ray absorption near-edge structure (XANES) measurements were the plant roots exposed to 50 nm AuNPs. Frozen roots or 10 nm citrate-AuNPs suspensions (at -80 °C) were freeze-dried at -53 °C under a vacuum of 0.01 mbars, homogenized, pressed into 7 mm pellets, sealed into Kapton tape, and further stored under an anoxic atmosphere before analysis. Gold nanoparticles (Au⁰) were diluted in sugar prior to homogenization and pelleting. XANES spectra were collected using a liquid nitrogen cryostat at the Au L₃₄-edge (11918 eV) in fluorescence mode using a 100 element Ge detector at Stanford Synchrotron Radiation Light Source on Beamline 11-2. Spectra were energy calibrated using an Au metal foil collected simultaneously with the samples spectra. A total of 3 to 10 individual spectra were acquired per pellet to get representative spectra. To minimize the risk of beam damage, each spectrum was collected at a different pellet location. The spectra were then averaged and normalized using SIXpack data processing software. Linear combination fitting was done using the Demeter processing software.

# AUTHOR INFORMATION

## Corresponding Authors

*E-mail: aavellan@andrew.cmu.edu.

*E-mail: glowry@andrew.cmu.edu.

## ORCID

Astrid Avellan: 0000-0001-6081-4389

Eleanor Spielman-Sun: 0000-0001-9626-2969

Jason M. Unrine: 0000-0003-3012-5261

Garret Bland: 0000-0001-9879-7879

Gregory V. Lowry: 0000-0001-8599-008X

# ACKNOWLEDGMENTS

The synchrotron images were collected on Beamline S-ID at the National Synchrotron Light Source II, a U.S. Department of Energy (DOE) Science User facility operated by the DOE Office of Science by Brookhaven National Laboratory (proposal no. 302315) and at the XFM beamline (AS173/XFM/12373) at the Australian Synchrotron, part of ANSTO. Portions of this work were performed at the Stanford Synchrotron Radiation Lightsource (SSRL) on Beamline 11-2, a Department of Energy supported facility. The TEM images
were completed at the Materials Characterization Facility at Carnegie Mellon University supported by grant no. MCF-677785. We thank M. De Jonge and R. Tappero for their help running the XFM beamline at the Australian Synchrotron and 5-ID at NSLS-II, respectively, as well as D. Van Sanford for providing the wheat seeds. This work was supported by the National Science Foundation (NSF) and the Environmental Protection Agency (EPA) under NSF Cooperative Agreement EF-0830093 and DBI-1266252, Center for the Environmental Implications of NanoTechnology (CEINT), and CBET-1530563 (NanoFARM).

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