Cadmium sulfide quantum dots impact Arabidopsis thaliana physiology and morphology

Journal Pre-proof Cadmium sulfide quantum dots impact Arabidopsis thaliana physiology and morphology Marta Marmiroli, Francesca Mussi, Luca Pagano, Davide Imperiale, Giacomo Lencioni, Marco Villani, Andrea Zappettini, Jason C. White, Nelson Marmiroli PII:

S0045-6535(19)32095-8

DOI:

https://doi.org/10.1016/j.chemosphere.2019.124856

Reference:

CHEM 124856

To appear in:

ECSN

Received Date: 16 July 2019 Revised Date:

10 September 2019

Accepted Date: 13 September 2019

Please cite this article as: Marmiroli, M., Mussi, F., Pagano, L., Imperiale, D., Lencioni, G., Villani, M., Zappettini, A., White, J.C., Marmiroli, N., Cadmium sulfide quantum dots impact Arabidopsis thaliana physiology and morphology, Chemosphere (2019), doi: https://doi.org/10.1016/ j.chemosphere.2019.124856. This is a PDF file of an article that has undergone enhancements after acceptance, such as the addition of a cover page and metadata, and formatting for readability, but it is not yet the definitive version of record. This version will undergo additional copyediting, typesetting and review before it is published in its final form, but we are providing this version to give early visibility of the article. Please note that, during the production process, errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain. © 2019 Published by Elsevier Ltd.

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Cadmium sulfide quantum dots impact Arabidopsis thaliana physiology and

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morphology

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Marta Marmiroli,1* Francesca Mussi,1 Luca Pagano,1 Davide Imperiale,2 Giacomo Lencioni,1

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Marco Villani,3 Andrea Zappettini,3 Jason C. White,4 Nelson Marmiroli.2

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1 - Department of Chemistry, Life Sciences and Environmental Sustainability, University of Parma,

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Parma, Italy.

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2 - Consorzio Interuniversitario Nazionale per le Scienze Ambientali (CINSA), University of

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Parma, Parma, Italy.

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3 - IMEM-CNR, Parma, Italy.

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4 - The Connecticut Agricultural Experiment Station, New Haven, CT, USA.

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*, corresponding author, Parco Area delle Scienze 33/A, 43123 Parma, Italy, +39 0521905698,

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[email protected]

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Abstract

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The differential mechanisms of CdS QDs (Quantum Dots) and Cd ion toxicity to Arabidopsis

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thaliana (L.) Heynh were investigated. Plants were exposed to 40 and 60 mg L-1 for CdS QDs and

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76.9 and 115.2 mg L-1 CdSO4·7H2O and toxicity was evaluated at 5, 20, 35 (T5, T20, T35) days

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after exposure. Oxidative stress upon exposure was evaluated by biochemical essays targeting non-

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enzymatic oxidative stress physiological parameters, including respiration efficiency, total

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chlorophylls, carotenoids, ABTS and DPPH radicals reduction, total phenolics, GSH redox state,

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lipid peroxidation. Total Cd in plants was measured with AAS. Root and leaf morphology and

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element content were assessed in vivo utilizing low-vacuum Environmental Scanning Electron

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Microscopy (ESEM) with X-ray microanalysis (EDX). This integrated approach allowed

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identification of unique nanoscale CdS QDs toxicity to the plants that was distinct from CdSO4

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exposure. The analyses highlighted that CdS QDs and Cd ions effects are modulated by the

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developmental stage of the plant, starting from T20 till T35 the plant development was modulated

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by the treatments, in particular CdS QDs induced early flowering. Both treatments induced Fe

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accumulation in roots, but at different intensities, while CdS QDs was associated with Mn increase

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into plant leaf. CdSO4 elicited higher levels of oxidative stress compared with QDs, especially the

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former treatment caused more intense respiration damages and reduction in chlorophyll and

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carotenoids than the latter. The two types of treatments impact differently on root and leaf

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morphology.

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Keywords: CdS QDs, Arabidospis thaliana, oxidative stress, morphology, Iron, ESEM/EDX.

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Highlights

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CdS QDs and Cd ion impact differently on A. thaliana morphology and physiology.

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CdS QDs damage mostly roots and induce early flowering.

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CdS QDs and Cd ion modulate Fe concentration in roots.

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1.Introduction

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Engineered nanomaterials (ENMs) are becoming widely diffused in many industrial products of

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everyday use. Nanomaterial interactions with biota continue to be a topic of significant scientific

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interest, with more than 260,000 published papers since 2010 (https://www.scopus.com). One class

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of engineered nanomaterials (ENM) recently gaining increasing attention is Quantum dots (QDs).

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Quantum dots are semiconductor nanocrystals with diameters range between 2 and 10 nm

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(Alivisados et al., 1998), and importantly, their chemical and physical properties are largely size-

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dependent. Given their unique optical properties, QDs have achieved a number of important

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applications (Brus, 1983; Frecker et al., 2016). For example, QDs are key constituents of innovative

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nanotechnology-enabled tools for medical diagnostic and ex vivo imaging (Padmanabhan et al.,

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2016; Wang et al., 2016). Quantum dots have also been used to improve energy efficiency of

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diodes, color gamut and resolution in digital cameras, TVs, computers and smartphones displays;

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and to augment energy conversion efficiency in quantum solar cells (QDSSCs) (Nurmikko, 2015;

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Zhu et al., 2015). Consequently, the increasing uses of QDs-enabled products are expected to result

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in the release of these materials to the environment (Gensch et al., 2016; Vance et al., 2015).

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However, although some QDs effects on single cells have been elucidated, particle impacts on

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whole organisms are still poorly understood (Paesano et al., 2016; Wu et al., 2016, Wang et al.,

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2016; Pagano et al., 2018). 3

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Regulatory guidance within the European Union regarding engineered nanomaterials (ENM) is

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delineated in the REACH legislation (REGULATION (EC) No 1907/2006). QDs utilization is also

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regulated by the RoHS Directive (DIRECTIVE 2002/95/EC) and by further exemption requests

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(Gensch et al., 2016). Pursuant to this legislation, only the amount of Cd within the nanocrystals is

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considered relevant. However, a number of publications have indicated disparate results regarding

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the causes of Cd-based QDs toxicity: several studies have implicated Cd release as the main toxicity

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factor but under other circumstances, particle properties such as shape and size seem to be more

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relevant (Wu et al, 2014; de Carvalho et al., 2016; Marmiroli et al., 2014; Marmiroli et al., 2016).

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Importantly, the influence of exposure duration on Cd-based QDs toxicity is poorly understood h

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(Oh et al., 2016; Yan et al., 2019).

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To date, few studies have investigated and compared the impact of CdS QDs and Cd ions on whole

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organism phenologic development, as well as on organ, tissue and cellular structure (Wang et al.,

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2018; Rocha et al., 2017; Majumdar et al., 2019). The goal of the current work was to understand

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the mechanisms of CdS QDs toxicity relative to that of Cd ions, and to determine how exposure

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impacts overall plant growth at different life stages. In order to localize Cd and other elements, and

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to detect morphological variations in roots and leaves, Environmental Scanning Electron

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Microscopy (ESEM) coupled with X-ray microanalysis (EDX) was used (Donald, 2003). Given that

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induction of cellular ROS and the ensuing oxidative stress are considered major causes of ENM

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toxicity (Yan et al., 2013; Khanna et al., 2015; Oh et al., 2016; Wu et al., 2016), a comprehensive

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picture of oxidative stress caused by Cd in the form of QDs or ions was obtained by means of

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biochemical essays targeting non-enzymatic oxidative stress physiological parameters, including

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respiration efficiency, total chlorophylls, ABTS and DPPH for radicals reduction, total phenolics,

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carotenoids, GSH redox state, lipid peroxidation.

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2.Materials and methods

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2.1. CdS QDs synthesis and characterization

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Uncoated Cadmium Sulfide Quantum Dots (CdS QDs) were synthesized by IMEM-CNR (Parma,

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Italy), following the method of Villani et al. (2012). The CdS QDs were characterized in deionized

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water by transmission electron microscopy (TEM) (Hitachi HT7700, Hitachi High Technologies

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America, Pleasanton, CA) (Pasquali et al., 2017). Average static diameter was 5 nm, and the crystal

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structure was that of wurtzite (ZnS) with approximately 78% Cd. Average particle size (dh) of the

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aggregates and zeta potential (ζ) in ddH2O were estimated in deionized water at 196.0 nm and +15.2

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mV, respectively. (Pagano et al., 2017) (Zetasizer Nano Series ZS90, Malvern Instruments,

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Malvern, UK). Additional particle characterization data is provided in the Supplementary Materials,

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and Figures S1-S2. We also performed a 5-day experiment to asses Cd release from CdS QDs in

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Murashige and Skoog (MS) medium with and without the presence of plant roots. Details are in the

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Supplementary Materials.

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2.2. Experimental set-up

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Wild type seedlings of Arabidopsis thaliana (L.) Heynh ecotype Landsberg erecta (Ler-0) were

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grown on a Murashige and Skoog (MS) nutrient medium (Duchefa Biochemie, Haarlem, NED)

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containing 1% w/v sucrose and solidified with 0.8% w/v agar at 24 °C, 30% relative humidity, and

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16-h photoperiod (light intensity 120 µM m−2 s−1 photosynthetic photon flux). After 10 days of

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growth on non-treated MS medium, the seedlings were transferred to media treated with a range of

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concentrations of CdS QDs or CdSO4 (Sigma-Aldrich, St. Louis, MO); untreated controls were

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maintained unamended media. The treatments (Table S1) were established as follows, with the

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minimum growth inhibiting concentration (MIC) established by Marmiroli et al. (2014): CdS QDs 5

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½ MIC = 40 mg L-1, CdS QDs ¾ MIC = 60 mg L-1, CdSO4 ·7H2O ½ MIC = 100µM or 76.9 mg L-1

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CdSO4, CdSO4 ·7H2O ¾ MIC = 150µM or 115.35 mg L-1 CdSO4 where reported in table S1. Plants

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were sampled for the different endpoints as described below beginning on the day of transfer to

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treated media (T0), after 5, 20, 35 days (T5, T20, T35), as reported in Table S2. All reagents were

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purchased from Sigma-Aldrich (St. Louis, MO, USA) unless stated otherwise.

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2.3. Cadmium (Cd) content determination

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The plant tissue Cd content was determined by FA-AAS (Flame-Atomic Absorption Spectrometry)

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(AA240FS, Agilent Technologies, Santa Clara, CA, USA) at 228.8 nm. Plants harvested after

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exposure to either CdS QDs or CdSO4 were thoroughly washed to remove residual particles/ions

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and were then dried at 50°C for 24 h. A 300 mg (dry weight) aliquot of ground plant material was

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digested in 10 mL 14.6 M HNO3 for 20 min at 165°C followed by 30 minutes at 230°C. The

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resulting solution was subsequently diluted to 6.7 M HNO3 using distilled water. The recording

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absorbance for each sample was converted to Cd concentrations via a standard curve based on a

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standard solution of high purity (>99%) Cd (Agilent Technologies, TO, Italy). All analyses were

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performed in triplicate.

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2.4. Leaf pigment content

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Chlorophyll and carotenoids content in the leaves of treated plants were evaluated according to Ni

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et al. (2009) with minor modification. Briefly, an a 300 mg aliquot of flash-frozen in liquid

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nitrogen and ground leaves was suspended in 800 µL 95% acetone (Ni et al. 2009). After

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incubation for 10 min on ice, the samples were centrifuged at 1000 g and 4°C for 10 min. The

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following parameters were determined in the supernatant by spectrophotometric analysis (Varian

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Cary 50, Agilent Technologies, TO, Italy): chlorophyll a: 662 nm, chlorophyll b: 647 nm, total

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carotenoids: 480 nm (Porra et al. 1989; Wellburn et al.. 1994).

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2.5. TTC assay

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The TTC (2,3,5-triphenyltetrazolium chloride) reduction assay was used as a quantitative method to

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evaluate tissue viability through respiration activity (Porter et al., 1994). A 200 mg aliquot of the

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aerial tissue was immersed in 3 ml of TTC buffer (TTC 0.18 M, 78% Na2HPO4·H2O 0.05 M, 22%

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KH2PO4 0.05 M). The samples were incubated at 30°C for 15 hours and the supernatant was then

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drained off, following by two additional washes with deionized water. The resulting formazan

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formed by the reaction was extracted by adding 10 ml of 95% ethanol for 10 minutes at 80°C.

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Formazan was then quantified spectrophotometrically (Varian Cary 50, Agilent Technologies, TO,

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Italy) at 530 nm.

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2.6. Plant extract preparation

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Leaf methanol extracts were used to estimate the total phenolic content and antioxidant activity

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(Capanoglu et al., 2008). Leaves were harvested, snap frozen in liquid nitrogen, ground and 100

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mg aliquots of the powder were stored at −80°C. Each sample was sonicated (Transsonic T460,

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Elma Schmidbauer GmbH, Singen, Germany) for 15 minutes at 35 kHz in 1 ml of 75% methanol

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and then centrifuged (Microfuge 22R, Beckman-Coulter, CA, USA) at 15000 g for 10 min at 4°C.

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The supernatant was collected and the pellet was subjected to a second round of the extraction

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procedure. The two supernatants were then combined for analysis as described below.

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The aqueous leaf extracts were used to estimate the glutathione (GSH) content (Gondim et al.,

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2013). Samples were harvested, snap frozen in liquid nitrogen, ground and 500 mg aliquots were

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homogenized in 1 ml of sterile distilled water and were centrifuged at 15000 g for 15 min at 4°C.

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The supernatant was collected and stored at -20°C until use as described below.

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2.7. Total phenolic content

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The total phenolic content of plant methanol extracts was determined by the Folin-Ciocalteu

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spectrophotometric method (Singleton and Rossi, 1965). Briefly, 100 µl of sample was mixed with 7

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0.75 ml of Folin-Ciocalteu reagent and allowed to stand at 22 °C for 5 min; 0.75 ml of sodium

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bicarbonate (60 g L-1) solution was then added to the mixture and after 90 min at 22°C, absorbance

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was measured (Varian Cary 50 spectrophotometer, Agilent Technologies) at 725 nm. The total

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phenolic content was calculated from a calibration curve using a gallic acid (GA) standard (1:100

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µg ml-1).

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2.8. ABTS assay

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The 2,2’-azinobis-(3-ethylbenzothiazoline-6-sulfonic acid) assay (ABTS assay) is a colorimetric

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method to assess the antioxidant activity of lipophilic and hydrophilic antioxidants. Following the

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method of Re et al. (1999), a 7 mM ABTS aqueous solution was oxidized by adding 2.45 mM

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potassium persulfate at 4°C for 16 h in the dark before use. The radical solution was heated to 30°C

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in a water bath and diluted in methanol to reach an absorbance between 0.65 and 0.75 at 734 nm.

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After the addition of 1 ml of diluted ABTS•+ solution to 10 µl of the methanol plant extracts, the

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samples were heated to 30°C for 5 min and the absorbance was recorded (Varian Cary 50

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spectrophotometer, Agilent Technologies) after 5, 10, 15 and 20 min. Appropriate solvent blanks

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were included in each assay. Trolox (6-hydroxy-2,5,7,8-tetramethylchroman-2-carboxylic acid) was

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used as reference standard and was added to the radical solution at 0, 1, 5, 10, 20, 50 µM to obtain

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the concentration/response curve. The radical inhibition percentage (I %) was calculated as ((A

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ABTS

•+

- A sample)/ A ABTS•+) x 100.

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2.9. DPPH assay

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The 2,2-Diphenyl-1-picrylhydrazyl assay (DPPH assay) is a colorimetric method to measure the

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free radical scavenging activity of antioxidant compounds. According to the method of Brand-

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Williams et al. (1995), a solution of 0.06 mM DPPH in methanol was prepared daily. An aliquot of

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50 µl of sample of methanol-plant extract was added to 1.95 ml of DPPH solution and after 30 8

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minutes at ambient temperature, the absorbance at 520 nm was measured (Varian Cary 50

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spectrophotometer, Agilent Technologies). The absorbance was also read after 40 and 50 minutes of

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incubation to verify that a steady state was achieved (plateau in the curve). Appropriate solvent

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blanks were run in each assay. Trolox was used as reference standard and was added to the radical

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solution at 0, 1, 5, 10, 20, 50 µM to obtain the concentration/response curve. The radical inhibition

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percentage was calculated as: (ADPPH - A sample)/ADPPH) x 100.

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2.10. Glutathione redox state

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Reduced and total glutathione content in the leaf aqueous extracts was determined according to

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Griffith (1980). The reduced glutathione (GSH) content was measuerded by adding to 20 µl of

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sample to 180 µl of reaction mixture containing 0.5 N potassium phosphate buffer (pH 7.5), 0.1

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mM ethylenediamine tetra acetic acid (EDTA), and 6 mM 5,5-dithiobis-(2-nitrobenzoic acid)

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(DTNB). After 10 min of incubation at 30°C, the absorbance was read with a microplate reader at

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412 nm (iMarkTM Microplate Absorbance Reader, Bio-Rad; MI, Italy). Total glutathione

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[GSH+oxidized glutathione disulphide (GSSG)] was measured after reduction of GSSG to GSH by

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adding 2 mM nicotinamide adenine dinucleotide phosphate (NADPH) and 1U glutathione reductase

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to the reaction mixture. The content of GSH was estimated using GSH as a standard and the

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glutathione redox state was calculated as [(GSH)/(GSH+GSSG) ×100].

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2.11. Lipid peroxidation

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The thiobarbituric acid (TBA) test was performed to evaluate lipid peroxidation. According to the

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method of Murshed et al. (2008), 200 mg of sample was ground, homogenized in 1 ml of 0.1%

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(w/v) TCA and centrifuged for 15 min at 12000g. A 500 µl aliquot of the supernatant was added to

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1 ml of 0.5% (w/v) TBA in 20% (w/v) TCA. The mixture was incubated at 95°C on a thermoblock

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(Termo Block 780, Asal, Firenze, Italy) for 30 min and the reaction was halted by placing the 9

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sample tubes in an ice bath. The samples were then briefly vortexed and the absorbance was read at

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532 nm (Varian Cary 50 spectrophotometer, Agilent Technologies). The value for non-specific

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absorbance at 600 nm was subtracted and a standard curve using commercial MDA was used to

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determine sample MDA content.

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2.12. ESEM /EDX

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To overcome the drawbacks of high vacuum, Environmental-Low vacuum (Lo-vac, 60Pa)

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ESEM/EDX was utilized. Whole plants were analyzed fresh with no fixation or staining; the plants

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were carefully washed, gently blotted, and were positioned on 2 cm diameter stainless-steel sample

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holder (stub) covered with adhesive carbon tape. The scanning microscope ESEM FEG2500 FEI

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(FEI Europe, Eindhoven, The Netherlands), operating in low-vacuum (60 Pa) with LFD (Large

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Field Detector) allowed optimal Secondary Electron (SE) imaging. The cone PLA (Pressure

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Limiting Aperture) 500µm improved the signal available to the Bruker X-ray detector, QUANTAX

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XFlash® 6 | 30 Detector with energy resolution ≤126 eV FWHM at Mn Kα., and a highly efficient,

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versatile mid-size 30 mm² SDD (Silicon Drift Detector) for nano-analysis and high count rate

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spectral imaging (Bruker Nano GmbH, Berlin, Germany). SE imaging was performed at 10 KeV

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with a beam size of 2.5 µm, EDX analysis at 20 KeV acceleration voltage, final lens aperture of 40

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µm, and beam size of 4 µm. The working distance was about 10 mm, and the scanning time was 60

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s. The software xT microscope Control, xT microscope Server and FEI User Management software

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were used for imaging, Esprit 1.9 package was used for X-ray spectra acquisition and analysis was

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conducted during two acquisition modes: Point/Area analysis and Linescan. X-ray spectra

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deconvolution

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(Peak/Background evaluation matrix with atomic number (Z), absorption (A), and secondary

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fluorescence (F) correction) interactive method supported by Esprit 1.9 “Quantify Method Editor”

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option (Goldstein, et al., 2003). SE images and EDX spectra were collected for samples treated with

and

standard-less

quantification

was

performed

using

the

P/B-ZAF

10

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the highest concentrations of CdS QDs and CdSO4, as well as for the untreated controls, at all three

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time points (T5, T20, T35). For roots and leaves, each of the three biological replicates were

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analyzed where at least 6 EDX point spectra were collected for the standard-less analysis. The

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detection limit in our working conditions were of 0.01% for elements with N ≥ 21 and 0.005% for

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N < 21.

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2.13. Statistical analysis

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Statistical calculations were based on routines implemented either in IBM SPSS v. 24.0 (Chicago,

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Illinois, USA; http://ibm.com/analytics/us/en/technology/spss/) or in R v3.3.1 (www.r-project.org).

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Two-way and three-way ANOVAs were performed after establishing non-significance of Levene’s

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test on the individual variables. The threshold for the multivariate and univariate p was set at 0.05.

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Post hoc tests were performed for each dependent variable (D.V.) within each independent variable

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(I.V.) and their combinations; the post-hoc Tukey’s HSD test was performed as suggested by

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literature (Field 2013). For the dimension reduction analysis, a principal component analysis (PCA)

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was performed. To ascertain the suitability of the datasets, the KMO (Kaiser-Mayer-Olkin) index

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was calculated; since this yielded 0.775, dimension reduction was considered to be justifiable. The

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number of components extracted was set by applying an eigenvalue threshold of λ = 1, and this was

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verified by use of rawpar.sps program (O'Connor, 2000).

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3.Results and Discussion

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3.1. Total Cd concentration in plants

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The total Cd content of A. thaliana roots and aerial tissues was measured by flame atomic

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absorption spectrometry (FA-AAS). The amount of Cd depended both on the concentration and

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duration of exposure. Significant increases in Cd concentration were detected after the exposure to 11

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both CdSO4 doses. For CdS QDs treatment the range of Cd concentrations at T20 was between 1.0

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and 1.1 mg g-1 Cd on dw; for the CdSO4 at the same time the range was between 2.5 and 3.8 mg g-1

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Cd on dw. The highest Cd concentrations were measured after the treatment with CdSO4,

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specifically with the 115.35 mg L-1 salt concentration and in the long-term treatment T35, when the

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concentration was above 7 mg g-1 Cd on dw. At T35 for the higher treatment with CdS QDs (60 mg

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L-1), the Cd concentration in plants was 1.7 mg g-1 Cd on dw. For the lower CdS QDs (40 mg L-1)

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treatment the concentration of Cd at T35 was 1.1 mg g-1 Cd on dw (Figure 1A). These results were

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in line with our earlier work on A. thaliana and S. cerevisiae indicating that CdS QDs did not

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release Cd ions (Marmiroli et al., 2014; Pasquali et al., 2017, Ruotolo et al. 2018) and enter into the

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plant cells as whole nanoparticles. Once inside the plant cell the CdS QDs cause toxicity primarily

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with the generation of ROS (reactive oxygen species) (Pagano et al., 2018).

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3.2. Physiological and biochemical assays

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3.2.1 Leaf chlorophyll and carotenoids concentrations

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Leaf chlorophyll and carotenoid content showed a significant time-dependent decrease with both

280

CdS QDs and CdSO4 treatments. In particular, under the higher treatments of both types (60 mg L-1

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CdS QDs and 115.35 mg L-1 CdSO4) chlorophyll decreased significatively from T20 to T35;

282

starting at T20 from 0.5 and 0.56 mg g-1 fw for CdS QDs and CdSO4 respectively reaching at T35

283

below 0.48 and 0.25 mg g-1 fw respectively. Interesting, the negative effect on the plant pigments at

284

the two concentrations of CdS QDs treatments followed the same trend as the Cd salt treatments:

285

constantly decreasing from T5 till T35 (Figures 1B-C). These findings agreed with the literature,

286

where a number of studies have shown that ENMs exposure can damage the photosynthetic

287

apparatus and deplete chlorophyll content (Zsiros et al., 2019, Hatami et al., 2016). Furthermore,

288

from past work, we know that the CdS QDs target the chloroplast structure and functionality 12

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causing an overall decrease in the chlorophylls pool (Pagano et al., 2018). On the other hand, Cd

290

affects photosynthesis by inhibition of different reaction steps of the Calvin cycle and not by

291

interaction with photochemical reactions located on the thylakoid membranes (Dijebali et al.,

292

2005).

293

A greater decrease in terpenic pigment concentration was observed under the CdSO4 high dose

294

treatment than under the higher dose of CdS QDs treatments. At T20, for both treatments the

295

concentrations of carotenoids were 0.8 and 1.1 mg g-1 fw respectively, whilst at T35 the

296

concentrations dropped to 0.05 and 0.06 mg g-1 fw respectively. Carotenoids were surely utilized to

297

quench free radicals caused by the treatments by both Cd and CdS QDs (Gallego et al., 2012), but

298

the capacity of the plant to replace them was scarce. Probably because many terpenes are

299

synthesized in the plastoglobuli contained in the chloroplasts (van Wijk and Kessler, 2017) which in

300

turn are under the negative influence of Cd and CdS QDs. Not surprisingly, a significant decrease in

301

terpenenes was observed in the untreated sample at T35, indicative of the natural senescence

302

process (Zimmermann et al., 2006). As evident in Table 1, there is significant evidence for a strong

303

interaction between Cd type and time of treatment (Table 1). Yan et al., 2019 have found that in

304

algae, the interaction of time and treatment was of a critical factor to the extent of damage caused

305

by CdSe quantum dots.

306

3.2.2 Leaf respiration (TTC assay)

307

The respiratory efficiency of plants was evaluated by measuring the reduction of 2,3,5-

308

triphenyltetrazolium chloride (TTC) in the aerial tissues. TTC reduction occurs at the end of the

309

mitochondrial respiratory chain in complex IV and, therefore, reflects the total electron flow,

310

including the alternative oxidative respiratory pathway (Rich et al.,2001). CdS QDs exposure had a

311

significantly different effect on respiration efficiency than did Cd salt treatments at all exposure

312

times (T5, T20, T35) (Figure 1D). A marked decrease in respiration was observed for both types of

313

treatments, but the pattern of that decrease was markedly different. The highest decrease occurred 13

314

for both CdS QDs (40 and 60 mg L-1) treatments at T0 and T5 where the respiration index went

315

from 0.6 OD to 0.2 OD. During the same time span for the two concentrations of CdSO4 (115.35

316

and 76.9 mg L-1) the respiration index went from 0.6 to 0.41 OD. Considering the time span from

317

T5 to T20 the respiration index increased for the CdS QDs treatments, conversely for the CdSO4

318

treatments the respiration index further decreased. During the last time span from T20 to T35 for

319

the CdS QDs treatments the respiration index decreased, whilst for the CdSO4 treatments the

320

respiration index was stable (Figure 1D).

321

The respiration rate in the control ultimately decreased as much as the treatments, although this was

322

a function of the natural senescence process (Buchanan-Wollaston et al., 2003). CdS QDs have

323

been shown to impair respiration in both plant species and yeast (Marmiroli et al., 2014; Pagano et

324

al., 2018; Pasquali et al 2018), but in the aforementionned published studies, the time-dependent

325

nature of the response was unknown. It is likely that the opposite effect observed for the two

326

treatments, detrimental for CdSO4, slightly positive for CdSQDs, T20 was less influenced by

327

inherent senescence at T35, when for both treatments respiration decreased, highlighting the

328

importance of differential response based on substrate type within the first 5 days of exposure.

329

330

3.2.3 Total phenolics content

331

Phenolic compounds are secondary plant metabolites of considerable physiological and

332

morphological importance and are known to be important to plant growth, reproduction and defense

333

against biotic and abiotic stress (Ignat et al., 2011). These biomolecules are characterized by

334

significant antioxidant activity, mainly due to their redox properties, which can play an important

335

role in binding and neutralizing free radicals, quenching singlet and triplet oxygen, or decomposing

336

peroxides (Tsuda et al., 2000).

14

337

We observed a significant time- and dose-dependent increase of the total phenolic (TP) content with

338

both treatment types. Phenolics in the control plants were consistently lower than in the treated

339

samples, with the exception being T5 for CdS QDs treatments. At any given treatment time, total

340

phenolics were consistently higher under CdSO4 treatment (both levels) than with CdS QDs (Figure

341

1E). From T0 to T35 TP, for the higher CdSO4 (115.35 mg L-1) treatment, increased steadily.

342

During the same time span, TP for the lower CdSO4 treatment (76.9 mg L-1) increased till T20, but

343

then decreased to 39.5 mg GA eq g-1 fw at T35. On the contrary, for both CdS QDs treatments the

344

greatest increase in TP was from T0 to T5. However, from T5 to T35 there was little or no increase

345

in TP under these CdS QDs treatments, whilst for Cd there was a substantial increase in these

346

molecules reflecting a higher amount of ROS produced by the Cd treatment, to be quenched within

347

the plant cells to protect photosynthesis and respiration. (Hatami et al., 2016).

348

349

3.2.4 ABTS and DPPH assays

350

The leaf antioxidant activity was assessed using two methods: the ABTS and DPPH assays, which

351

indicate the total capacity of cells to reduce two synthetic radicals (ABTS•+ radical cation and

352

DPPH• neutral radical). These two parameters account for all the ROS scavenging molecules, not

353

only phenols and carotenoids, but also ascorbic acid, small organic acids, sugars and amino acids

354

(Mittler, 2002). However, it is possible that the metabolite extraction method for the tissues may

355

cause partial loss of photosensitive antioxidative molecules, such as tocopherols, and this must be

356

taken into consideration.

357

The ABTS and DPPH radical inhibition activity increased over time, following similar trends

358

(Figures 1F-G). In both cases, the highest scavenging activity occurred for the 115.2 mg L-1 CdSO4

359

treatment and increased over the time. The lowest antioxidant activities for ABTS and DPPH were

360

observed for the control and for the CdS QDs treatment at 40 mg L-1 at all treatment times. During 15

361

the treatment, the plant did not increase anti-ROS activity as a function of CdS QDs exposure. In

362

addition, at T35 antioxidant activity in all treatments was increased significatively in comparison

363

with T20; this is consistent with what was observed for the phenolic compounds that was associated

364

with excessive ROS production in the plant cells caused mostly by Cd ions and in a lower quantity

365

by CdS QDs (Pagano et al., 2018), confirming that Cd ions produce more ROS than QDs.

366

367

3.2.5 Glutathione redox state (GSH/GSSG)

368

Glutathione is a key molecule in defense pathways against stressors such as heavy metals and is a

369

precursor in the synthesis of phytochelatins (PCs) (Cobbett and Goldsbrough, 2002). GSH also

370

scavenges heavy metal-induced ROS by the ascorbate–glutathione cycle and is involved in the

371

regulation of intracellular redox homeostasis and signaling (Noctor et al., 2012).

372

We observed a significant increase in the GSH redox state, particularly evident at T20, for both

373

CdSO4 concentrations, but only for the higher dose of CdS QDs (60 mg L-1) (Figure 1H).

374

Importantly, the 40 mg L-1 CdS QDs treatment did not significantly change the GSH redox state

375

with respect to control, suggesting that this dose is has minimal toxicity. When plants were

376

challenged with the higher nanoparticle concentrations (60 mg L-1), GSH/GSSG reached the control

377

level at T35 (22%). The higher percentage of the GSH redox state was reached at T20 for all

378

treatments, although with significant differences between CdSO4 (almost 100%) and CdS QDs

379

(60% for the treatment with 60 mgL-1). Therefore, for GSH there was a strong interaction between

380

both treatment and time, which suggests the need for cells to modulate their redox state not only as

381

consequence of exposure, but also as a function of upcoming senescence. It has to be noted that

382

treatment with Cd ions consistently induces the synthesis of higher concentrations of GSH due to

383

the high induction of ROS and direct chelation of Cd ions in comparison to CdS QDs. In addition,

384

we note that an increase in thiols is a typical response of plant exposure to ENMs (Du et al., 2017). 16

385

However, during the first 5 days of treatment (T0-T5), when senescence had not begun yet, the two

386

types of treatment elicited an opposite behavior of the GSH redox state: in the case of CdSO4 the

387

value increased, but for CdS QDs, the level decreased (40% and 19% respectively) (Figure 1H).

388

This trend reflected the different actions of the two types of treatment on the early developmental

389

stages of the plants, with the Cd ions generating a greater response than the QDs. It is also likely

390

that CdS QDs cannot be chelated as effectively by GSH as can Cd

391

suggest that CdS QDs detoxification rests less on GSH than does ionic Cd2+, and that additional S-

392

bearing molecules may contribute to this phase of QDs detoxification. In recent studies on ENMs-

393

effects on plants, the role of GSH has been clearly been linked to the fact that ENMs produce ROS,

394

which GSH then reduces through oxidation-reduction cycles with reduced and oxidized ascorbate

395

(ASA and DHA) (Soares et al. 2018). Here we confirm the important role of GSH in exposure, but

396

highlight that the changes in oxidation state are modulated according to the type of treatment, with

397

the ion having a greater effect on the cell oxidative state than the QDs (Figure 1H).

2+

. Taken together these results

398

399

3.2.6 Lipid peroxidation

400

As noted above, both metals and ENM induce oxidative stress in plants through the overproduction

401

of ROS, subsequently leading to DNA, protein and lipid damage (Sharma et al, 2012; Marslin et al.,

402

2017). According to this evidence, a time- and dose-dependent increase in lipid peroxidation was

403

induced by both treatments. The effect of CdS QDs treatments on lipid peroxidation was lower than

404

that of CdSO4 at any time point. It should be noted that, lipid peroxidation followed a similar trend

405

in response to that of total phenolics, GSH redox state, DPPH and ABTS assays (Figure 1I). For

406

example, there was a minimal increase from T0 to T35 of the levels of lipid peroxidation index for

407

the two treatments with CdS QDs (40 and 60 mgL-1). Conversely, for CdSO4 (115.35 and 76.9 mg

408

L-1) treatments the increase in lipid peroxidation index was high especially from T5 to T35. It

409

should be of notice that from T0 to T5 the behavior of lipid peroxidation followed that of GSH 17

410

redox state for the two types of treatments, i.e. their trends were opposite: increase for the ion

411

treatment, decrease for the QDs treatment (Figures 1H-1I ). Soares and colleagues (2018), in a

412

recent review on the effects of ENM on plants oxidative stress, found that lipid peroxidation is

413

constantly increasing in many plant species treated with different types of nanomaterials, especially

414

those based on metals. However here the toxicity of CdSO4 treatments is overwhelming in

415

comparison with that of CdS QDs at any concentration and time point.

416

417

3.3. Multivariate statistics

418

3.3.1 Two-way ANOVA

419

The two-way ANOVA for the independent variable (I.V.) “Treatment time” was highly significant

420

for all dependent variables (D.V.) of the physiological assays, with p ≤ 0.05 and ɳ2 >70%. The I.V.

421

“treatment type”, discriminating between CdS QDs or CdSO4 without considering concentration,

422

was highly significant for all the D.V.s with p≤0.05 and ɳ2 >70% for all variables, except for

423

carotenoids which had p≤0.07 and ɳ2 >35% and chlorophyll which had p≤0.08 and ɳ2 =30%. The

424

I.V.s interaction “treatment time x treatment type” was highly significant for the variance of all the

425

D.V.s (with the same parameters p and ɳ2 values as for the “treatment type”, i.e. p≤0.05 and ɳ2

426

>70%) (Table 1). From these results, the importance of plant developmental stage on the treatment

427

effects as measured by all the physiological parameters considered and on the amount of

428

bioaccumulated Cd was clearly stated (Figure 1). Similarly, Yan et al (2019) reported in algae that

429

the interaction of time and treatment was highly influential on CdSe quantum dot toxicity (Yan et

430

al., 2019). In our case from Figure 1 it appeared that the intensity of the action of the treatment with

431

CdSO4 (both concentrations) on all the parameters was greater than that of the CdS QDs (both

432

concentrations). In particular, during the first 5 days of experiment (T0-T5) the action of the two

433

types of treatment elicited opposite response in GSH redox state and in lipid peroxidation: increase 18

434

for the ion, decrease for the QDs. On the other hand, both respiration activity and chlorophyll

435

concentration decreased under both treatments starting from T0, even though with a significant

436

greater extent for the ion treatments in respect to the QDs treatments. It is known from previous

437

work that CdS QDs and Cd ions affect negatively both mitochondria and chloroplasts causing

438

diminished performances of these vital organelles (Pasquali et al. 2017; Pagano et al., 2018).

439

Indeed, also in this case we observed physiological and morphological decline of the plants caused

440

both by time and type of treatment (Tab1, Fig1 and Fig 2).

441

442

3.3.2 Dimensions reduction: Principal Components Analysis (PCA)

443

To ascertain the suitability of the dataset for dimension reduction, the KMO (Kaiser-Mayer-Olkin)

444

index was calculated; since this yielded 0.775, factor reduction was considered justifiable. Two

445

components with eigenvalues λ >1 were extracted for PCA (Table S3); PCA analysis explained

446

68% of total variance (Table S3). Vectors for dependent variables “cadmium”, “GAE”, “MDA”,

447

“GSH”, “ABTS” were present primarily as the first component; vectors for “Chlorophyll”,

448

“Carotenoids”, “Respiration” were represented as the second component (Table S4). The vector

449

corresponding to the dependent variable DPPH was almost equally present on both components

450

(Figure S3, Table S4). These results suggest that the treatments exerted different mechanisms of

451

toxicity on the most important cellular reactions linked to respiration and photosynthesis as

452

compared to ROS detoxification activities such as the synthesis of phenolics, carotenoids and GSH .

453

In our case, it was possible to partially pool together experimental data points according to type of

454

treatment (Non treated, CdS QDs, CdSO4) in separate groups (Figure S4A).

455

significant interaction between time and type of treatment was notable, and after 35 days, the “time

456

of treatment” showed a stronger effect than the “treatment type” (Figure S4A). In Figure S4B, data

457

points of the “treatment time” T35 were removed, which allowed to compact clusters of points

However, the

19

458

according to the variable “treatment type”. Taken together these data mean that, even though each

459

single parameter showed a specific trend during time, the interaction between “time” and “type” of

460

treatment became stronger as the plant progressed toward the later stages of its development (T35).

461

462

3.4. Low-vacuum ESEM coupled with microanalysis

463

Low vacuum scanning microscopy allowed analysis of plant tissues in vivo with minimal sample

464

preparation. Organ morphology and development of treated plants was compared to controls at the

465

three time-points; T5, T20, and T35. The analyses were performed only for the highest treatment

466

concentrations for both CdSO4 and CdS QDs (115.35 mg L-1 and 60 mg L-1).

467

3.4.1 Modifications in plant organs structure

468

To measure alterations in plant structure and development, several parameters were considered: i)

469

main and lateral roots size and shape; ii) upper leaf features such as trichomes, wax deposition, and

470

stomatal number; iii) flowering time and development. As compared with controls, CdSO4

471

treatment caused faster and stronger changes in organ development and morphology than CdS QDs;

472

however, these changes differed in type and features.

473

CdS QDs treatment (60 mg L-1) caused deformities in lateral roots that worsened exposure. After 5

474

days of treatment, the lateral roots started to swell, becoming more enlarged at T20. At T35, the

475

lateral roots seemed to burst as the tissues split apart (Figure 2, Figures S5A-S5C). This

476

phenomenon could likely be ascribed to the partial clogging of the roots cause by the CdS QDs

477

nanostructures when they enter the root epidermis as has been reported for other metal-based ENMs

478

(Zuverza-Mena et al., 2017). Moreover, root splitting was consistent with the high degree of lipid

479

peroxidation associated with cell membrane disruption and breakdown. (Figure 1). Conversely,

480

CdSO4 treatment (115.35 mg L-1) negatively affected the main and lateral root development,

481

causing general thinning, after 35 days, lateral roots ceased to develop and root tips became necrotic 20

482

(Figures 2, Figures S5A-S5C) (Gallego et al., 2012). There are thus different ways in which CdS

483

QDs and Cd ions damage the root development in accordance with the fact that the source of

484

toxicity for nanoparticles is not the release of Cd ions, but their structure and reactivity.

485

At T5, leaves did not show any adverse effects as a consequence to treatments. The leaves of plants

486

treated with QDs presented normal growth until T20, comparable to non-treated samples; the only

487

exception being the presence of some leaves with high number of trichomes even if not statistically

488

significant (Figure 2, Figures S5A-S5C). Increased wax deposition on upper leaf surface was

489

observed after 35 days under CdS QDs treatment, accompanied by reduced stomatal and trichome

490

density as compared to controls (Figure 2, Figures S5A-S5C). At T20 under CdSO4 treatment, the

491

leaves of treated plants also showed a decrease in stomatal and trichome density, with appreciable

492

adaxial wax deposition compared to leaves of non-treated plants (Figure 2, Figures S5A-S5C). At

493

T35, under CdSO4 treatment, the upper leaf surface was covered by a thick wax and displayed

494

necrotic areas, similar to Cd toxicity symptoms already reported (Figure S5B) (Shanying et al.,

495

2017). The difference in the toxicity symptoms in leaves between the two types of treatment might

496

be due to a faster translocation of Cd ions to the above ground parts in comparison to the movement

497

of CdS QDs (Pagano et al., 2018).

498

Control plants showed inflorescences after 20 days, which blossomed after 35 days. At T20 in most

499

of the plants treated with CdS QDs, mature flowers were observed bearing fully developed

500

reproductive organs. Importantly, with CdSO4 significantly fewer plants exhibited fully formed

501

flowers at that same time point (Figure S5).

502

Plants can be induced to flower when under water or low nutrient stress (Wada and Takeno, 2010).

503

It is possible that the presence of CdS QDs, which negatively impacted the root structure, impaired

504

nutrient and water uptake capacity of the plant, which consequently increased the stress degree and

505

anticipated flowering. It is consistent with its higher toxicity that the treatment with CdSO4, which

506

had more detrimental effects on all the physiological and morphological parameters in respect to 21

507

CdS QDs treatment, evoked less early flowering. Furthermore, flower development and plant

508

fertility are dependent on iron homeostasis (Takahashi et al., 2003). Small organic molecules such

509

as nicotianamine (NA) and citrate participate in this process by chelating Fe in order to shuttle it

510

throughout the plant organelles especially to chloroplasts (Rellán-Alvarez et al., 2011)

511

512

3.4.2 Detection of Elements in plant organs

513

The elements detected and quantified through EXD include: Ca, Mg K, P, S, Cd, Cl, Cu, Fe, Mn. A

514

three-way ANOVA was performed considering all elements as Dependent Variables (DVs) and the

515

Independent Variables (IVs) included: Organ (roots, leaves), Time (T5 = 5 days, T20 = 20 days,

516

T35 = 35 days), and Treatment (NT= non treated, Cd ions II = high CdS QDs concentration of 60

517

mg L-1, QDs II = high CdSO4 concentration of 115.2 mg L-1) (Table 2).

518

From the three-way ANOVA, it appeared that the elements with the highest variance in all possible

519

conditions and combinations were Cd, S, and Fe; those with the lowest variance were Ca and Mg

520

(Table 2). This means that the elemental content, evaluated according to the semi-quantitative EDX

521

detection, was highly dependent on the type of organ and on treatment time. It is possible to

522

consider this finding as a confirmation of the physiological assay data, where variance was linked

523

more to the combinations of independent variables than to each single independent variable alone.

524

This in vivo description of elemental distribution provides a unique perspective as compared to

525

more traditional isolation of tissues or organs followed by desiccation, acid digestion and ICP-MS

526

analysis.

527

Cadmium was detected in the roots and leaves of treated plants after 5 days of treatment and the

528

amount increased with time; in general, the levels were consistently higher under CdSO4 treatment

529

in comparison to CdS QDs, even though the actual dose of elemental Cd was higher with the CdS

530

QDs treatment than with the CdSO4 (Table S1, Table 3A, B). At T20 the Cd content in the roots 22

531

and leaves of plants treated with CdSO4 was almost 10 times higher than in plants treated with QDs,

532

but at T35, the difference had decreased to two-fold (Table 3A, B). These data obtained with the X-

533

ray emission microanalysis are consistent with the FA-AAS quantification of Cd concentrations in

534

the whole plant (Figure 1A.) At T35 in CdSO4 and CdS QDs treated plants, Cd was the most

535

abundant element in roots and shoots, at times exceeding the main macronutrients (Ca, K, S, P)

536

(Figure 3, Figures S6A-S6E, Table S3A, B). In both roots and leaves, Ca and S increased over

537

time, while Cl, P, and K decreased. It has been reported that TiO2 and ZnO ENM altered the

538

internal ionome of kidney bean (Phaseolus vulgaris) and basil (Ocimum basilicum), respectively. In

539

particular, both types of ENMs were able to change Ca, S, and P internal concentrations roots and

540

shoots (Tan et al., 2017; Medina-Velo et al., 2017). Therefore, it is possible that the very nature of

541

the structure of the CdS QDs caused similar effects. Furthermore, S was significantly influenced by

542

the type of treatment, being consistently higher in treated plants in respect to control (Figure 3,

543

Figures S6A-S6E, Table 3A, B). This finding was likely to reflect that S was exploited in thiol-

544

bearing molecules to detoxify ROS generated by the treatments, as noted with GSH levels in Figure

545

1H (Du et al., 2017).

546

As expected, Mg was higher in leaves than in roots, where levels varied according to time, and the

547

amounts were consistently lower in treated plants than in controls (Figure 3, Figures S6A-S6E,

548

Table S5). This finding is in keeping with the decrease in chlorophyll observed from the

549

physiological analyses and reported in literature regarding ENMs toxicity (Hitami et al., 2016;

550

Zuverza-Mena et al., 2017).

551

Micronutrients such as Cu, Fe, and Mn varied between organs and followed different trends.

552

Modulation of these three micronutrients was also found by Majumdar and colleagues (2019) in

553

roots of soybean treated with coated and pristine CdS QDs. Leaf Cu was significantly lower in the

554

control plants; the levels of Cu were highest in CdSO4 treated plants (at all time points), but in

555

roots, Cu amounts decreased from T5 to T35 under all types of treatment (Figure with TiO2 and 23

556

ZnO ENM (Tan et al., 2017; Medina-Velo et al., 2017). Increased Cu can have a negative impact

557

on chlorophyll and photosynthesis, as we have observed in paragraph 2.1 (Figure 1B) (Apodaca et

558

al., 2017).

559

Fe content was higher in roots than in leaves, Mn followed the opposite trend; however, both

560

increased with time (Figure 3, Figures S6A-S6E, Table 3A, B). Fe increased in the roots whilst Mn

561

increased in the leaves under CdS QDs treatments, especially at T20 and T35 (Figure 3, Figures

562

S6A-S6E, Table 3A, B). Increased Fe in roots after ENMs treatment is consistent with literature

563

reports that utilized different plants (such as Ocimum basilicum L., Triticum aestivum L., Zea mays

564

L., Glycine max L., Lactuca sativa L., Cucumis sativus L., Phaseolus vulgaris L.) and different

565

types of ENM (nCeO2, nCuO, nTiO2, nZnO coated and uncoated) (Tan et al., 2017; Medina-Velo et

566

al., 2017, Du et al., 2017). Although Fe-SOD (Superoxide Dismutase) enzymes are found primarily

567

in chloroplasts, it was possible that the increase of Fe in treated roots reflected the increase in ROS

568

detoxifying enzymes. A similar mechanism could occur in leaves with Mn-SOD (Alscher et al.,

569

2002). On the other hand, the accumulation of Fe in the roots, starting at T20 and increasing at T35,

570

could limit leaf photosynthetic and respiratory activity, if in this way Fe flow to the shoot tissues is

571

obstructed (Briat et al., 2015). The increase in root Fe may also reflect that this metal could directly

572

impact the uptake of either CdSO4 or CdS QDs. In a recent paper by Majumdar et al (2019), it was

573

found that CdS QDs, bare or functionalize, modulate the concentrations of Fe in roots and shoots of

574

soybean. The Authors argue that since Cd2+ translocation in plants is driven by cell membrane metal

575

transporters (NRAMP) and zinc-regulated transporter/iron-regulated transporter related protein

576

(ZIP) families, these transporters can control the movement of metal ions with the same valence as

577

Fe2+, Zn2+, Cu2+. Competitive binding to the same class of transporter can arise in case of Fe2+ and

578

Cd2+ that could impair the uptake of Fe2+ in the plants. In our case the opposite seemed to occur

579

because we observed that the cotransporters of Fe2+ and Cd2+ increase the uptake of both metals

580

within the roots, especially in the case of CdSO4 treatment. Moreover, disturbing Fe homeostasis by 24

581

Fe excess due to Cd treatment leads to alteration of flower biology (Sudre et al., 2013), consistently

582

we observed an anticipation in flowering time in the plants challenged with CdS QDs (Figure S6).

583

Mn is essential for photosynthesis because it affects the water-splitting system of photosystem II

584

(PSII), which provides the necessary electron for the transport chain (Broadley et al., 2002). Mn

585

may have been recruited to the leaves to counteract Fe deficiency experienced in the chloroplasts.

586

From the line-scan analysis (Figure 4, Figure S7), it appears that trichomes acted as storage cells for

587

Cd under both treatments; this is consistent with what was already observed for both Cd and other

588

metals in different hyperaccumulator (Alyssum sp.) and non-hyperaccumulator (Nicotiana tabacum)

589

plant species (Broadhurst et al., 2004; Choi et al., 2001). It has been reported that in Cucumis

590

sativus that TiO2 ENM can translocate into leaf trichomes which act as sink or as possible secretory

591

cells for the nanomaterials (Servin et al., 2012). Therefore, it seems likely that CdS QDs could be

592

translocated into the trichomes as a detoxification strategy for both treatment types.

593

594

595

4. Conclusions

596

In the recent literature, there are only few examples of studies that investigate the different effects

597

of CdS QDs Cd ions on whole organisms of (Marmiroli et al., 2016; Wang et al., 2016). There are

598

some published studies on whole plants with pristine or coated QDs over short exposure times that

599

do not encompass all phenological stages (Marmiroli et al., 2014, Wang et al., 2016; Majumdar et

600

al 2019). This work set out with two goals; the first was to clarify the causes, and possibly the

601

mechanisms, of CdS QDs toxicity, with the direct intent to compare this to that of Cd ions. The

602

second goal was to investigate how the plant progress through the life stages was interconnected

603

with the toxicity exerted by CdS QDs treatments. The findings indicate that the detrimental action

604

of Cd as CdS QDs was different from that of Cd ions in a number of important ways. Although in 25

605

both cases elemental Cd was translocated from roots to shoots and produced oxidative stress, the

606

intensity and the physiological manifestations of the stress were quite different. Specifically, plants

607

treated with CdS QDs displayed less oxidative stress than those treated with CdSO4. This result was

608

more likely a function of the limited release of Cd ions from the QDs than of their size and

609

reactivity. The damage observed in vivo to the roots in the case of Cd ions treatment was consistent

610

with what found in literature for coated CdTe QDs and CdSe QDs (Wang et al., 2014; Modlibova et

611

al., 2018), and importantly, this displayed different features from CdS QDs-induced root damage. In

612

the case of CdS QDs, the roots appeared to be structurally damaged and deformed, but did retain

613

some degree of functionality given that the leaf appearance was healthier than that of the CdSO4

614

treatment. There was a clear indication of the importance of the interaction of time and type of

615

treatment in the toxicity assessment of CdS QDs and also Cd ions. Specifically, CdS QDs

616

treatments stimulated early flowering through the modulation of Fe homeostasis (Sudre et al., 2013)

617

and interestingly, senescence interacted with the treatment toxicity not only at the level of root and

618

leaf morphology, but also through the stimulation of changes in the physiological parameters that

619

we have measured. From

620

chlorophyll because of increased ROS-induced lipid peroxidation of the cell membranes. The level

621

of respiration was decreased starting from T5 till T35, likely due to Fe accumulation in the roots.

622

Furthermore, Fe modulation is linked with Cd uptake from the media under both Cd ions and CdS

623

QDs (Majumdar et al 2019) The model plant A. thaliana allowed a detailed investigation of the

624

mechanisms of CdS QDs toxicity in comparison to Cd ions under a full life cycle condition and

625

showed that senescence played a key role in the variation of the parameters chosen to measure this

626

toxicity on the plant organs.

T20, leaf cells started to loose macronutrients, micronutrients, and

627

628

Acknowledgments

26

629

Authors acknowledge the support of the project INTENSE, grant no. 652515. JCW acknowledges

630

USDA NIFA Hatch CONH00147.

631

632

Supplementary Materials

633 634

Methods section related to CdS QDs synthesis and characterization.

635

Figure S1. HRTEM image of ligand-free QDs assembly and XRD spectra

636

Figure S2. ESEM image and EDX spectra of CdS QDs

637

Figure S3. PCA vectors for the physiological analysis.

638

Figure S4. PCA points for the treatment time and types.

639

Figure S5. Low vacuum ESEM SE images of roots, leaves and flowers.

640

Figure S6. EDX spectra for roots and leaves, at all treatment times.

641

Figure S7. Trichome linescan from the CdSO4 treatment.

642

Table S1. Treatment conditions and their effective Cd content.

643

Table S2. Treatment and sampling times.

644

Table S3. PCA vectors’ eingenvalues and explained variance.

645

Table S4. PCA vectors loading coefficients.

646

647

648

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878 879 880

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37

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895

Figure Captions and Tables

896

897

Figure 1. Cd concentrations in the whole plant measured with FA-AAS and physiological assay

898

performed for all times and types of treatments. Following a one-way ANOVA the Tukey’s HSD

899

test was performed. Different lower-case letters correspond to significant differences with p≤0.05.

900

Each assay has been given a different upper-case letter as identified in the text.

901 38

902

Figure 2. Low vacuum ESEM SE images of roots, leaves and flowers acquired at 10 or 5 kV of

903

beam energy, beam size of 2.5 µm, pressure 60 Pa, and a working distance about 10 mm. The white

904

bar at the left of each image indicates the reference measure. Images were acquired only for the

905

highest treatments (60 mg L-1 CdS QDs and 115.2 mg L-1 CdSO4·7H2O) and for the control (NT).

906

Images were acquired at T20.

907

908

Figure 3. EDX spectra collected for the highest treatments (60 mg L-1 CdS QDs and 115.2 mg L-1

909

CdSO4·7H2O) at all treatment times. Spectra were acquired at 20 kV, beam size of 4 µm, working

910

distance was about 10 mm, and scanning time 1-3 µs. The yellow rectangle on the SE images

911

corresponds to the EXD deconvoluted spectra below for T20 root.

912 913

Figure 4. Trichome linescan from the base to the first branching for the 60 mg L-1 CdS QDs

914

treatment. (A) Relative elemental content along the scanned section (green arrow) (B). The Cd

915

linear distribustion (in blue) resulted highly represented throughout the scanned section.

916 917

Table 1: Two-way ANOVAs for physiological parameters. I.V. independent variables; D.V. dependent

918

variables; H.df. Hypothesis degrees of freedom = (I.V. subgroups number)-1; (a) Significance: ***p≤0.001,

919

** p≤0.01, *p≤0.05, ns p>0.05; (b) Univariate Effect Size % partial eta square η2. I.V. main effects and interaction

Treatment Time

D.V.

H. df.

F exact value

Sign.(a)

η2 %(b)

Chlorophyll (Chl a+b)

3

22.033

***

70

Carotenoids

3

11.222

***

55

Respiration (TTC)

3

142.524

***

94

Cadmium

3

22.182

***

71

ABTS

3

14.268

***

60

DPPH

3

35.059

***

79

Total phenolic

3

52.558

***

85

39

Lipid peroxidation (MDA)

3

32.485

***

78

GSH

3

22.650

***

71

Chlorophyll (Chl a+b)

2

0.506

*

30

Carotenoids

2

7.545

**

36

Respiration (TTC)

2

68.096

***

83

Cadmium

2

35.760

***

72

ABTS

2

17.815

***

56

DPPH

2

60.855

***

81

Total phenolic

2

65.827

***

82

Lipid peroxidation (MDA)

2

81.145

***

85

GSH

2

48.465

***

78

Chlorophyll (Chl a+b)

6

0.210

*

30

Carotenoids

6

2.911

**

38

Respiration (TTC)

6

36.231

***

89

Cadmium

6

6.702

***

59

ABTS

6

7.822

***

63

DPPH

6

9.040

***

66

Total phenolic

6

14.620

***

76

Lipid peroxidation (MDA)

6

18.237

***

80

GSH

6

8.524

***

65

Treatment Type

Time * Type

920

Table 2: Three-way ANOVA on the EDX elemental calculation within plant organs (roots and leaves) for

921

three treatment conditions (NT, Cd ions, QDs) and three treatment times (T5, T20, T35). I.V. Independent

922

Variables; D.V. Dependent Variables; Time: T5= 5 days, T20= 20 days, T35= 35 days; Organ: roots, leaves;

923

Treatment: NT= non treated, Cd ions II= high Cd ions concentration, QDs II= high CdS QDs concentration.

924

ns= not significative, *= p < 0.05, ** = p ≤ 0.01, ***= p ≤ 0.005.

I.V.

D.V.

Cd

Ca

Cl

Cu

Fe

Mg

Mn

P

K

S

Treatment Time

***

ns

***

***

***

ns

*

*

***

***

Organ

***

***

ns

***

***

***

***

ns

***

I.V. Main effect

**

40

Treatment Type

***

ns

***

Time*Organ

***

*

ns

Time*Treatment

***

ns

ns

Organ*Treatment

***

ns

Time*Organ*Treatment

***

ns

ns

ns

ns

ns

ns

***

ns

***

ns

**

*

**

**

*

***

*

ns

ns

***

**

*

ns

ns

ns

ns

ns

ns

***

ns

ns

ns

ns

ns

ns

***

***

I.V. Interactions

***

41

925

TableS3A. Tukey’s HSD post-hoc tests for elements in roots. Element time

Cd type

Mea

Ca s.e.

mean

Cl s.e.

mean

Cu s.e.

mean

Fe s.e.

mean

Mg s.e.

mean

Mn s.e.

mean

P

s.e.

mean

K s.e.

mean

S s.e.

mean

s.e.

11.89

1.79

n NT

0.00

0.00

e Cd ion II

15.88

3.97

2.98

d 3.98

7.43

7.88

1.18

a 2.25

3.21

1.11

0.33

a 0.89

1.19

4.40

2.70

d 0.25

17.88

1.05

0.48

cb 3.55

1.52

0.00

0.00

c 0.36

0.14

11.43

2.62

cb 0.05

10.16

55.24

5.63

a 1.98

29.23

b 4.26

9.65

1.35

T5 c QDs II

1.67

c 4.71

d NT

0.00

33.36

2.66

cd 0.00

e Cd ion II

4.56

b

4.34

9.49

1.06

ab 2.98

cd 5.27

6.26

a

8.08

3.12

0.29

a 1.18

a 2.98

1.16

b

0.62

1.10

2.20

d 0.33

b 1.18

4.92

b

7.82

16.53

0.43

b 3.70

cd 0.33

1.79

a

0.81

1.85

0.00

c 0.48

c 4.70

0.00

c

0.00

0.00

2.35

b 0.00

c 0.48

12.72

d

6.02

8.36

5.04

b 2.62

d 0.00

49.92

cb

61.76

12.89

1.60

a 5.63

a 2.62

14.59

7.85

1.79

c 5.63

9.43

1.79

T20 b QDs II

4.61

c 3.98

d NT

0.00

39.66

2.25

c 0.00

e Cd ion II

7.10

b

16.30

10.27

0.89

b 3.44

a 3.18

4.33

a

3.79

0.72

0.25

b 1.37

b 1.80

0.63

b

0.00

0.26

3.55

b 0.38

c 0.71

19.42

b

3.26

13.40

0.36

b 5.42

d 0.20

1.70

c

3.18

1.68

0.01

b 0.55

a 2.83

0.02

d

0.00

0.00

1.98

c 0.00

c 0.29

10.85

e

18.47

9.31

4.26

c 3.03

a 0.00

36.91

cb

38.16

10.16

1.35

b 6.51

c 1.58

11.71

5.82

2.06

d 3.40

11.47

1.08

T35 a QDs II

19.24 c

bc 3.72

11.56 b

c 2.11

1.32 c

c 0.84

0.78 b

c 0.23

26.45 a

b 3.32

1.70 b

c 0.34

0.00 c

c 0.00

12.21 b

e 1.86

11.73 e

b 3.98

10.89

1.26

b

926 927

Different letters indicate significant differences according to ANOVA followed by Tukey’s HSD post-hoc test with p ≤ 0.05. 42

928

Table S3B. Tukey’s HSD post-hoc tests for elements in leaves. Element time

Cd type NT

mean 0.00

Ca s.e.

mean

s.e.

0.00

18.07

2.91

f Cd ion II

1.95

Cl

a 0.58

18.15

mean 6.26

Cu s.e. 1.28

c 1.99

5.30

mean 0.68

Fe s.e. 0.35

d 0.87

2.00

mean 0.00

Mg s.e. 0.00

b 0.24

0.00

mean 2.52

Mn s.e. 0.65

b 0.00

3.35

mean 0.26

P

s.e. 0.15

cd 0.44

1.07

mean 15.89

K s.e. 1.64

ab 0.31

14.71

mean 42.25

S s.e. 3.76

a 1.12

37.19

mean 8.11

s.e. 1.68

d 2.57

13.43

1.15

T5 e QDs II

1.63

a 0.26

e NT

0.00

14.05

cd 2.32

b 0.00

13.40 b

34.91

1.02

c 2.57

f Cd ion II

6.07

b

8.11

18.77

3.14

4.78

0.28

c 1.13

b 1.71

1.21

b

0.61

3.06

0.00

b 0.31

d 1.38

0.00

ab

0.00

0.00

0.52

ab 0.00

b 0.38

3.35

b

2.05

1.99

0.36

a 0.57

c 0.00

2.16

b

0.44

0.04

1.31

ab 0.40

c 0.70

16.24

b

15.70

2.84

3.00

ab 1.44

ab 0.01

40.21

bc

42.30

19.23

1.34

bc 3.32

a 1.77

13.90

11.57

1.48

c 4.06

11.22

1.82

T20 c QDs II

3.19

a 0.94

d NT

0.00

62.99

1.72

b 0.00

f Cd ion II

13.79

d

5.79

17.89

0.76

a 4.44

c 2.42

10.18

a

0.04

0.78

0.21

c 0.01

f 4.44

1.02

b

0.00

2.17

0.04

a 0.00

e 1.95

0.15

c

0.00

0.00

0.38

bc 0.00

b 0.53

2.36

d

4.63

1.52

0.27

b 0.79

a 0.00

1.04

d

0.00

0.00

0.97

c 0.00

d 0.69

8.61

d

22.02

0.59

2.23

a 2.50

a 0.00

43.45

c

29.59

3.90

0.99

b 5.75

c 1.50

15.99

27.85

2.57

a 1.75

6.79

2.57

T35 a QDs II

39.93 b

ab 1.58

16.48 ab

ef 2.91

4.00 d

b 1.28

1.70 c

b 0.35

0.00 b

c 0.00

3.30 ab

d 0.65

0.51 c

d 0.45

2.86 d

e 1.64

16.96 d

d 3.76

16.57

1.68

b

929 930

Different letters indicate significant differences according to ANOVA followed by Tukey’s HSD post-hoc test with p ≤ 0.05. 43

Highlights •

CdS QDs and Cd ion impact differently on A. thaliana morphology and physiology.

•

CdS QDs damage mostly roots and induce early flowering.

•

CdS QDs and Cd ion modulate Fe concentration in roots.