Nanoparticle Uptake in Plants: Gold Nanomaterial Localized in Roots

Jul 7, 2017 - Terrestrial plants can internalize and translocate nanoparticles (NPs). However, direct evidence for the processes driving the NP uptake...
0 downloads 0 Views 1MB Size
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