Nanoparticle Uptake in Plants: Gold Nanomaterial Localized in Roots

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Nanoparticle Uptake in Plants: Gold Nanomaterial Localized in Roots of Arabidopsis thaliana by X-Ray Computed Nanotomography and Hyperspectral Imaging Astrid Avellan, Fabienne Schwab, Armand Masion, Perrine Chaurand, Daniel Borschneck, Vladimir Vidal, Jérôme Rose, Catherine Santaella, and Clément Levard Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/acs.est.7b01133 • Publication Date (Web): 07 Jul 2017 Downloaded from http://pubs.acs.org on July 8, 2017

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Environmental Science & Technology

Nanoparticle Uptake in Plants: Gold Nanomaterial

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Localized in Roots of Arabidopsis thaliana by X-Ray Computed

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Nanotomography and Hyperspectral Imaging

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Astrid Avellan1,2,3, Fabienne Schwab1, 2, Armand Masion1,2, Perrine Chaurand1,2, Daniel

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Borschneck1,2, Vladimir Vidal1,2, Jérôme Rose1,2, Catherine Santaella2,3 and Clément

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Levard1,2*

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(1) Aix Marseille Univ., CNRS, IRD, Coll. France, CEREGE, Aix en Provence, France.

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(2) iCEINT, International Center for the Environmental Implications of Nanotechologies,

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CNRS - Duke University, Europôle de l’Arbois, 13545 Aix-en-Provence, France.

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(3) Aix-Marseille Univ, CEA, CNRS, Laboratory of Microbial Ecology of the Rhizosphere

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and Extreme Environments (LEMIRE), Biosciences and biotechnology Institute of Aix-

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Marseille (BIAM) ECCOREV FR 3098, CEA/Cadarache, St-Paul-lez-Durance, France.

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Keywords: Nanoparticle detection, Environment, Complex matrices, Uptake, Translocation,

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Roots, Mucilage.

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*Corresponding author: Tel: +33 7 77 97 50 80, [email protected]

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ACS Paragon Plus 1Environment


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Abstract

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Terrestrial plants can internalize and translocate nanoparticles (NPs). However, direct

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evidence for the processes driving the NP uptake and distribution in plants is scarce at the

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cellular level. Here, NP-root interactions were investigated after 10 days of exposure of

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Arabidopsis thaliana to 10 mg.L-1 of negatively or positively charged gold NPs ((-/+)Au-NPs,

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13.4±1.3 nm, and 12.1±0.8 nm, respectively) in gels. Two complementary imaging tools were

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used: X-ray computed nanotomography (nano-CT), and enhanced dark-field microscopy

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combined with hyperspectral imaging (DF-HSI). The coupled use of these emerging

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techniques improved our ability to detect and visualize NP in complex plant tissue: by

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spectral confirmation via DF-HSI, and in three dimensions via nano-CT. The resulting

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imaging provides direct evidence that detaching border-like cells (i.e. sheets of border cells

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detaching from the root) and associated mucilage can accumulate and trap NPs irrespective of

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particle charge. On the contrary, border cells on the root cap behaved in a charge-specific

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fashion: positively charged NPs induced a higher mucilage production and adsorbed to it,

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which prevented translocation into the root tissue. Negatively charged NPs did not adsorb to

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the mucilage and were able to translocate into the apoplast. These observations provide direct

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mechanistic insight into NP-plant interactions, and reveal the important function of border

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cells and mucilage in interactions of plants with charged NPs.


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TOC Art

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1

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Understanding the interactions of nanoparticles (NPs) with terrestrial plants is

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essential to predict their fate in terrestrial environments and their possible accumulation in the

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food chain.1 Plant roots can internalize and accumulate NPs, presumably primarily via the

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apoplastic pathway along the cell walls and intercellular spaces into the vasculature. From the

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vasculature, NPs may further translocate into stems and leaves.2,3 Initial studies on metal-

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based NP interactions with roots suggest a charge-dependent NP uptake and translocation

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rates. Positively charged NPs have been found to associate to a greater extent with roots.

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Negatively charged NPs can be taken up more easily and translocate more efficiently into

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shoots.4–9 Until now, phytotoxicity of NPs has gained more interest than the underlying

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mechanisms of NP uptake and the associated mobility in plants.10

Introduction

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A major challenge in studying NP-plant interactions is to obtain direct and

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unambiguous evidence of NP uptake.11 Initial studies assessed NP-root association and

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translocation via metal analysis in roots and shoots, and visualization tools such as electron

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microscopy to confirm the NP uptake. Using a single visualization technique to localize NPs

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in plant tissue often results in conflicting data, especially at low, environmentally relevant

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concentrations. The few observable NPs can be almost indistinguishable from naturally

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occurring NPs or other background signals.12,13 Moreover, complex and destructive sample

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preparation protocols as labeling, staining, sputter-coating, and ultrathin cutting, can all

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introduce artifacts.14 Less invasive sample preparation and the coupling of complementary

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techniques, e.g. elemental analysis coupled with NP identification and mapping techniques,

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can help to reduce such artifacts.15

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One emerging technique with great potential to be included in such an

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interdisciplinary approach is the enhanced resolution dark–field microscopy16 and

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hyperspectral imaging (DF-HSI). This 2D visualization tool requires minimal sample ACS Paragon Plus 4Environment

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preparation, and can detect and map the NP-specific reflectance spectra of a material in a

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complex environmental matrices17 at the nano-scale, in relatively short time (minutes-hours),

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and in a narrow focus plane. The current spectral resolution of DF-HSI is 1.5 nm, and the

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spatial resolution is about 90 nm.16,17 Importantly, although 3D imaging could be done with

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the most recent setups, DF-HSI only enabled 2D visualization and could fail in identifying

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NPs biological barrier crossing. Objects smaller than the spatial resolution (≈10 nm) can also

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be detected if they possess a strong light scattering signal. Dark–field hyperspectral imaging

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has already been used to study NP-organism interactions, e.g. in vitro interactions of NPs with

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cell;18,19 or in vivo interactions of NPs with unicellular organisms such as protozoa,20

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bacteria,21–23, and green algae,24,25 or in entire organisms such as fishes26 or worms.27,28

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Despite its demonstrated usefulness to provide information of NP location in cells and small

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organisms, DF-HSI has not yet been used to detect NPs in terrestrial plants.

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An other promising technique for NP imaging in plants providing valuable

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complementary information besides DF-HSI is X-ray computed tomography (CT). This

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mature 3-dimensional (3D) imaging technique is based on the X-ray attenuation by a sample.

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CT requires neither cutting nor labeling or staining of the samples, which greatly reduces the

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risk of artifacts from sample preparation. CT imaging techniques have resolutions of ~1 µm

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and ~50nm for micro- and nano-CT, respectively. Micro-CT has been successfully used to

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visualize microscopic plant features in 3D with a resolution of few µm29. Nano-CT was first

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developed on synchrotron beamlines,30 and has recently been adapted to benchtop

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systems.31,32 Nano-CT could provide valuable 3D information with a good resolution on the

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roots-NP interaction (adsorption vs. internalization), and the scanning of large volumes can

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simplify the detection of low NP concentrations. However, unambiguous identification of

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NPs can become challenging in natural heterogeneous sample potentially composed of



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impurities with X-ray attenuation similar to that of metal-based NPs. A cross-validation

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identification with a NP-specific technique is then required.

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In the present study, a novel methodology based on these two complementary 2D and

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3D imaging techniques (DF-HSI and nano-CT) was proposed to perform a systematic

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characterization of NP-plant interactions at the cellular level. We aimed to explore the

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capabilities and limitations of the combination of DF-HSI and nano-CT to obtain two and

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three-dimensional and cross-validated information on distribution of small (~12 nm in

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diameter) Au-NPs on and in plant roots. Arabidopsis thaliana served as a model plant and

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was exposed to negatively and positively charged gold NPs (-/+ Au-NPs). This approach

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allowed an evaluation of hypotheses on NP uptake mechanisms at the cellular level.2,4,6

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2.1

Experimental Section

Au-NPs

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Negatively ((-)Au-NPs) or positively ((+)Au-NPs) charged Au-NPs stabilized by a

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coating of citrate, or branched polyethyleneimine, respectively, were purchased from

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nanoComposix Inc., Czech Republic. Diameters of (-/+)Au-NPs provided by the

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manufacturer were 13.4±1.3 nm, and 12.1±0.8 nm, respectively. TEM images and distribution

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histograms are shown in Figure S1 of the supplemental information (SI). Zeta potentials and

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hydrodynamic diameter of the NPs were measured in ultrapure water by electrophoretic- and

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dynamic light scattering (Zetasizer nanoZS, Malvern Inc., UK). The resulting hydrodynamic

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diameters and zeta potential for (-)Au-NPs and (+)Au-NPs at pH 7±0.2 in stock suspensions

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diluted in ultrapure water, were 18.6±7.1nm and -32.1mV; and 47.6±11.3nm and +46.3mV,

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respectively. The latter results confirmed those provided by the manufacturer.

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2.2

Plant culture, exposure to Au-NPs and growth

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Arabidopsis thaliana (Columbia ecotype) seeds were grown in gel as described

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elsewhere.33 Seeds were surface-sterilized for 5 min in Tween® 20, rinsed twice with ethanol, ACS Paragon Plus 6Environment

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and air-dried sterilely. Ten seeds were sown in square plates (12x12 cm2) containing sterile

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one half-strength Hoagland's solution solidified with Phytagel (Sigma Aldrich, United States).

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Recipes of these solutions are shown in Table S1 in the SI).

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Au-NPs were mixed with the nutrient/Phytagel solution prior to solidification to obtain

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a concentration of 10.0 mg Au-NPs.L-1. Control plants without exposure to Au-NPs were also

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prepared. The plates were sealed with Micropore® tape (3M, USA) to limit water evaporation

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and allow gas exchange, and incubated vertically for 10 days at a photon flux from the top of

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150μmol.m-2 s-1 and under a light:dark cycle of 16:8 h and 21:19°C. The germination rate

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was determined on ten seeds 10 days post-sowing, and the length of the roots measured using

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digital images and ImageJ software34.

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2.3

Preparation of roots for analysis

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Per treatment, five plants were analyzed in total: two plants by DF-HSI, one by nano-

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CT, and two by µ-XRF. For DF-HSI, the roots were separated from the shoots using a sterile

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razor blade immediately prior to analysis, rinsed 3 times with a sterilized solution containing

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10-3 mol.L-1 KCl, and were directly mounted between glass slide and coverslip with a 200 µL

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drop of the KCl solution.

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For µ-XRF and nano-CT root analysis, harvested roots were washed three times for

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5 min with pure sodium phosphate buffer (PPB, 0.10M, pH 7.2) and fixed using 2.5% (v/v)

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glutaraldehyde in PPB at room temperature for 12 h. Fixed roots were then dehydrated by

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immersion in ethanol series of 25, 50, 70, 90, 90, 100, 100 (% v/v) for 20 min each. The

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ethanol-saturated roots were dried using a CO2 supercritical point dryer (EM CPD 3000, Leica

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Microsystems Inc., USA) to preserve the cellular structure of the root tissue, and to limit

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drying artifacts.35 Dried roots were then slipped into a polyimide tube (Kapton®, Cole-

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Parmer, USA) mounted on a pin-type sample holder. For µ-XRF measurements, the

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supercritical point dried roots were fixed with polyimide tape on a clean silicon wafer.



Elemental micro-analysis of roots by µ-XRF

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2.4

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We performed micro-X-ray fluorescence (µ-XRF) on two different roots to identify

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the root regions containing the highest Au content. Semi-quantitative analysis of Ca and Au in

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roots was performed using a custom-built laboratory-scale µ-XRF microscope named

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HERMES (High X-ray Energy Resolution Microscope for Environmental Sciences). High

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sensitivity was achieved through a high flux incident X-ray beam (Mo rotating anode, 50 kV,

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24 mA, 1-2×1011 photons mm-2 s-1, spot diameter of 400 µm, high-flux optics from XENOCS,

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Chantilly, USA), and a 4 elements X-ray detector (Vortex®-ME4, Hitachi, Japan), allowing

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for the detection of relatively low local element concentrations (0.05). After 10 days, the average root length and their

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standard deviation were 2.47±0.47cm for the control root, 2.60±0.57cm for the (-)Au-NP

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exposed root and 4.25±2.37cm for the (+)Au-NP exposed root.

Results Growth and elemental micro-analysis of roots

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µ-X-ray fluorescence (spot analysis of 400 µm) was primarily used to detect the

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presence of Au associated with the roots. Two zones were analyzed: the apical zone including

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the root cap, and the root tip ~1 cm above the elongation zone. No Au was detected in the

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control roots. In both (-/+)Au-NP exposed roots, Au was detected (Table 1, Figure S2). When

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normalizing the Au XRF signal by the Ca signal, the Au distribution within the root was

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different for the two Au-NPs. While the Au signal of the (-)Au-NP exposed root was equally

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distributed between the two analyzed regions, Au in the (+)Au-NPs exposed root was mostly

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detected in the apical zone, while in the elongation zone, the Au signal was 5 times lower.

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Table 1: Normalized µ-XRF intensities (XRF, a.u.) in roots of Arabidopsis thaliana,

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means and standard deviations. The roots were analyzed in (i) the apical zone including the root

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cap; and (ii) ~1 cm above the root tip in the elongation zone. n.d.: not detected. The limit of

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detection was 0.2 a.u. Groups presenting different letters (a, b, c) are significantly different

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(ANOVA, Turkey HSD test, p

Nanoparticle Uptake in Plants: Gold Nanomaterial Localized in Roots

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