Subscriber access provided by UNIVERSITY OF CONNECTICUT
Article
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
Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a free service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are accessible to all readers and citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.
Environmental Science & Technology is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.
Page 1 of 28
Environmental Science & Technology
Nanoparticle Uptake in Plants: Gold Nanomaterial
1 2
Localized in Roots of Arabidopsis thaliana by X-Ray Computed
3
Nanotomography and Hyperspectral Imaging
4
Astrid Avellan1,2,3, Fabienne Schwab1, 2, Armand Masion1,2, Perrine Chaurand1,2, Daniel
5
Borschneck1,2, Vladimir Vidal1,2, Jérôme Rose1,2, Catherine Santaella2,3 and Clément
6
Levard1,2*
7
(1) Aix Marseille Univ., CNRS, IRD, Coll. France, CEREGE, Aix en Provence, France.
8
(2) iCEINT, International Center for the Environmental Implications of Nanotechologies,
9
CNRS - Duke University, Europôle de l’Arbois, 13545 Aix-en-Provence, France.
10
(3) Aix-Marseille Univ, CEA, CNRS, Laboratory of Microbial Ecology of the Rhizosphere
11
and Extreme Environments (LEMIRE), Biosciences and biotechnology Institute of Aix-
12
Marseille (BIAM) ECCOREV FR 3098, CEA/Cadarache, St-Paul-lez-Durance, France.
13 14
Keywords: Nanoparticle detection, Environment, Complex matrices, Uptake, Translocation,
15
Roots, Mucilage.
16
*Corresponding author: Tel: +33 7 77 97 50 80,
[email protected] 17
ACS Paragon Plus 1Environment
Environmental Science & Technology
18
Abstract
19
Terrestrial plants can internalize and translocate nanoparticles (NPs). However, direct
20
evidence for the processes driving the NP uptake and distribution in plants is scarce at the
21
cellular level. Here, NP-root interactions were investigated after 10 days of exposure of
22
Arabidopsis thaliana to 10 mg.L-1 of negatively or positively charged gold NPs ((-/+)Au-NPs,
23
13.4±1.3 nm, and 12.1±0.8 nm, respectively) in gels. Two complementary imaging tools were
24
used: X-ray computed nanotomography (nano-CT), and enhanced dark-field microscopy
25
combined with hyperspectral imaging (DF-HSI). The coupled use of these emerging
26
techniques improved our ability to detect and visualize NP in complex plant tissue: by
27
spectral confirmation via DF-HSI, and in three dimensions via nano-CT. The resulting
28
imaging provides direct evidence that detaching border-like cells (i.e. sheets of border cells
29
detaching from the root) and associated mucilage can accumulate and trap NPs irrespective of
30
particle charge. On the contrary, border cells on the root cap behaved in a charge-specific
31
fashion: positively charged NPs induced a higher mucilage production and adsorbed to it,
32
which prevented translocation into the root tissue. Negatively charged NPs did not adsorb to
33
the mucilage and were able to translocate into the apoplast. These observations provide direct
34
mechanistic insight into NP-plant interactions, and reveal the important function of border
35
cells and mucilage in interactions of plants with charged NPs.
ACS Paragon Plus 2Environment
Page 2 of 28
Page 3 of 28
36
Environmental Science & Technology
TOC Art
37 38
ACS Paragon Plus 3Environment
Environmental Science & Technology
39
1
40
Understanding the interactions of nanoparticles (NPs) with terrestrial plants is
41
essential to predict their fate in terrestrial environments and their possible accumulation in the
42
food chain.1 Plant roots can internalize and accumulate NPs, presumably primarily via the
43
apoplastic pathway along the cell walls and intercellular spaces into the vasculature. From the
44
vasculature, NPs may further translocate into stems and leaves.2,3 Initial studies on metal-
45
based NP interactions with roots suggest a charge-dependent NP uptake and translocation
46
rates. Positively charged NPs have been found to associate to a greater extent with roots.
47
Negatively charged NPs can be taken up more easily and translocate more efficiently into
48
shoots.4–9 Until now, phytotoxicity of NPs has gained more interest than the underlying
49
mechanisms of NP uptake and the associated mobility in plants.10
Introduction
50
A major challenge in studying NP-plant interactions is to obtain direct and
51
unambiguous evidence of NP uptake.11 Initial studies assessed NP-root association and
52
translocation via metal analysis in roots and shoots, and visualization tools such as electron
53
microscopy to confirm the NP uptake. Using a single visualization technique to localize NPs
54
in plant tissue often results in conflicting data, especially at low, environmentally relevant
55
concentrations. The few observable NPs can be almost indistinguishable from naturally
56
occurring NPs or other background signals.12,13 Moreover, complex and destructive sample
57
preparation protocols as labeling, staining, sputter-coating, and ultrathin cutting, can all
58
introduce artifacts.14 Less invasive sample preparation and the coupling of complementary
59
techniques, e.g. elemental analysis coupled with NP identification and mapping techniques,
60
can help to reduce such artifacts.15
61
One emerging technique with great potential to be included in such an
62
interdisciplinary approach is the enhanced resolution dark–field microscopy16 and
63
hyperspectral imaging (DF-HSI). This 2D visualization tool requires minimal sample ACS Paragon Plus 4Environment
Page 4 of 28
Page 5 of 28
Environmental Science & Technology
64
preparation, and can detect and map the NP-specific reflectance spectra of a material in a
65
complex environmental matrices17 at the nano-scale, in relatively short time (minutes-hours),
66
and in a narrow focus plane. The current spectral resolution of DF-HSI is 1.5 nm, and the
67
spatial resolution is about 90 nm.16,17 Importantly, although 3D imaging could be done with
68
the most recent setups, DF-HSI only enabled 2D visualization and could fail in identifying
69
NPs biological barrier crossing. Objects smaller than the spatial resolution (≈10 nm) can also
70
be detected if they possess a strong light scattering signal. Dark–field hyperspectral imaging
71
has already been used to study NP-organism interactions, e.g. in vitro interactions of NPs with
72
cell;18,19 or in vivo interactions of NPs with unicellular organisms such as protozoa,20
73
bacteria,21–23, and green algae,24,25 or in entire organisms such as fishes26 or worms.27,28
74
Despite its demonstrated usefulness to provide information of NP location in cells and small
75
organisms, DF-HSI has not yet been used to detect NPs in terrestrial plants.
76
An other promising technique for NP imaging in plants providing valuable
77
complementary information besides DF-HSI is X-ray computed tomography (CT). This
78
mature 3-dimensional (3D) imaging technique is based on the X-ray attenuation by a sample.
79
CT requires neither cutting nor labeling or staining of the samples, which greatly reduces the
80
risk of artifacts from sample preparation. CT imaging techniques have resolutions of ~1 µm
81
and ~50nm for micro- and nano-CT, respectively. Micro-CT has been successfully used to
82
visualize microscopic plant features in 3D with a resolution of few µm29. Nano-CT was first
83
developed on synchrotron beamlines,30 and has recently been adapted to benchtop
84
systems.31,32 Nano-CT could provide valuable 3D information with a good resolution on the
85
roots-NP interaction (adsorption vs. internalization), and the scanning of large volumes can
86
simplify the detection of low NP concentrations. However, unambiguous identification of
87
NPs can become challenging in natural heterogeneous sample potentially composed of
ACS Paragon Plus 5Environment
Environmental Science & Technology
88
impurities with X-ray attenuation similar to that of metal-based NPs. A cross-validation
89
identification with a NP-specific technique is then required.
90
In the present study, a novel methodology based on these two complementary 2D and
91
3D imaging techniques (DF-HSI and nano-CT) was proposed to perform a systematic
92
characterization of NP-plant interactions at the cellular level. We aimed to explore the
93
capabilities and limitations of the combination of DF-HSI and nano-CT to obtain two and
94
three-dimensional and cross-validated information on distribution of small (~12 nm in
95
diameter) Au-NPs on and in plant roots. Arabidopsis thaliana served as a model plant and
96
was exposed to negatively and positively charged gold NPs (-/+ Au-NPs). This approach
97
allowed an evaluation of hypotheses on NP uptake mechanisms at the cellular level.2,4,6
98
2
99
2.1
Experimental Section
Au-NPs
100
Negatively ((-)Au-NPs) or positively ((+)Au-NPs) charged Au-NPs stabilized by a
101
coating of citrate, or branched polyethyleneimine, respectively, were purchased from
102
nanoComposix Inc., Czech Republic. Diameters of (-/+)Au-NPs provided by the
103
manufacturer were 13.4±1.3 nm, and 12.1±0.8 nm, respectively. TEM images and distribution
104
histograms are shown in Figure S1 of the supplemental information (SI). Zeta potentials and
105
hydrodynamic diameter of the NPs were measured in ultrapure water by electrophoretic- and
106
dynamic light scattering (Zetasizer nanoZS, Malvern Inc., UK). The resulting hydrodynamic
107
diameters and zeta potential for (-)Au-NPs and (+)Au-NPs at pH 7±0.2 in stock suspensions
108
diluted in ultrapure water, were 18.6±7.1nm and -32.1mV; and 47.6±11.3nm and +46.3mV,
109
respectively. The latter results confirmed those provided by the manufacturer.
110
2.2
Plant culture, exposure to Au-NPs and growth
111
Arabidopsis thaliana (Columbia ecotype) seeds were grown in gel as described
112
elsewhere.33 Seeds were surface-sterilized for 5 min in Tween® 20, rinsed twice with ethanol, ACS Paragon Plus 6Environment
Page 6 of 28
Page 7 of 28
Environmental Science & Technology
113
and air-dried sterilely. Ten seeds were sown in square plates (12x12 cm2) containing sterile
114
one half-strength Hoagland's solution solidified with Phytagel (Sigma Aldrich, United States).
115
Recipes of these solutions are shown in Table S1 in the SI).
116
Au-NPs were mixed with the nutrient/Phytagel solution prior to solidification to obtain
117
a concentration of 10.0 mg Au-NPs.L-1. Control plants without exposure to Au-NPs were also
118
prepared. The plates were sealed with Micropore® tape (3M, USA) to limit water evaporation
119
and allow gas exchange, and incubated vertically for 10 days at a photon flux from the top of
120
150μmol.m-2 s-1 and under a light:dark cycle of 16:8 h and 21:19°C. The germination rate
121
was determined on ten seeds 10 days post-sowing, and the length of the roots measured using
122
digital images and ImageJ software34.
123
2.3
Preparation of roots for analysis
124
Per treatment, five plants were analyzed in total: two plants by DF-HSI, one by nano-
125
CT, and two by µ-XRF. For DF-HSI, the roots were separated from the shoots using a sterile
126
razor blade immediately prior to analysis, rinsed 3 times with a sterilized solution containing
127
10-3 mol.L-1 KCl, and were directly mounted between glass slide and coverslip with a 200 µL
128
drop of the KCl solution.
129
For µ-XRF and nano-CT root analysis, harvested roots were washed three times for
130
5 min with pure sodium phosphate buffer (PPB, 0.10M, pH 7.2) and fixed using 2.5% (v/v)
131
glutaraldehyde in PPB at room temperature for 12 h. Fixed roots were then dehydrated by
132
immersion in ethanol series of 25, 50, 70, 90, 90, 100, 100 (% v/v) for 20 min each. The
133
ethanol-saturated roots were dried using a CO2 supercritical point dryer (EM CPD 3000, Leica
134
Microsystems Inc., USA) to preserve the cellular structure of the root tissue, and to limit
135
drying artifacts.35 Dried roots were then slipped into a polyimide tube (Kapton®, Cole-
136
Parmer, USA) mounted on a pin-type sample holder. For µ-XRF measurements, the
137
supercritical point dried roots were fixed with polyimide tape on a clean silicon wafer.
ACS Paragon Plus 7Environment
Environmental Science & Technology
Elemental micro-analysis of roots by µ-XRF
138
2.4
139
We performed micro-X-ray fluorescence (µ-XRF) on two different roots to identify
140
the root regions containing the highest Au content. Semi-quantitative analysis of Ca and Au in
141
roots was performed using a custom-built laboratory-scale µ-XRF microscope named
142
HERMES (High X-ray Energy Resolution Microscope for Environmental Sciences). High
143
sensitivity was achieved through a high flux incident X-ray beam (Mo rotating anode, 50 kV,
144
24 mA, 1-2×1011 photons mm-2 s-1, spot diameter of 400 µm, high-flux optics from XENOCS,
145
Chantilly, USA), and a 4 elements X-ray detector (Vortex®-ME4, Hitachi, Japan), allowing
146
for the detection of relatively low local element concentrations (0.05). After 10 days, the average root length and their
233
standard deviation were 2.47±0.47cm for the control root, 2.60±0.57cm for the (-)Au-NP
234
exposed root and 4.25±2.37cm for the (+)Au-NP exposed root.
Results Growth and elemental micro-analysis of roots
235
µ-X-ray fluorescence (spot analysis of 400 µm) was primarily used to detect the
236
presence of Au associated with the roots. Two zones were analyzed: the apical zone including
237
the root cap, and the root tip ~1 cm above the elongation zone. No Au was detected in the
238
control roots. In both (-/+)Au-NP exposed roots, Au was detected (Table 1, Figure S2). When
239
normalizing the Au XRF signal by the Ca signal, the Au distribution within the root was
240
different for the two Au-NPs. While the Au signal of the (-)Au-NP exposed root was equally
241
distributed between the two analyzed regions, Au in the (+)Au-NPs exposed root was mostly
242
detected in the apical zone, while in the elongation zone, the Au signal was 5 times lower.
243
Table 1: Normalized µ-XRF intensities (XRF, a.u.) in roots of Arabidopsis thaliana,
244
means and standard deviations. The roots were analyzed in (i) the apical zone including the root
245
cap; and (ii) ~1 cm above the root tip in the elongation zone. n.d.: not detected. The limit of
246
detection was 0.2 a.u. Groups presenting different letters (a, b, c) are significantly different
247
(ANOVA, Turkey HSD test, p
