domain: chim sdu

text:
F Poitrasson
email: franck.poitrasson@get.omp.eu

Araus Jl
email: jaraus@ub.edu

Cristina Caldelas
email: criscaldelas@ub.edu

José Luis

Jérôme Viers
email: jerome.viers@get.omp.eu

Stable Zn isotopes reveal the uptake and toxicity of zinc oxide engineered nanomaterials in Phragmites australis

The Zn stable isotope composition of plants demonstrates that ZnO engineered nanomaterials dissolve before their uptake and accumulation by the roots (brightest inclusions in root cortex above).

Environmental Significance Statement

Our understanding of zinc oxide nanomaterials interaction with wetland plants is hampered by the lack of scientific consensus about their uptake and toxicity mechanisms in these species. This is a serious concern given the alarming global increase in the discharge of these nanomaterials into the environment and the key ecological roles of wetland plants. The Zn isotopic signature of plant tissue integrates all the Zn metabolic pathway throughout the plant's life, giving insight about the form of Zn taken up, even if this later transforms into another Zn species. Thus, our findings clarify the exposure routes and the mechanisms of action of zinc oxide engineered nanomaterials in wetland plants while advancing the toolbox for plant physiology and environmental studies.

Introduction

An estimated 34,000 tonnes year -1 of ZnO-ENMs are emitted into the environment globally, of which 3,000 are directly discharged into water bodies [START_REF] Keller | Global life cycle releases of engineered nanomaterials[END_REF] . Models predict that sediments will receive most of the ZnO-ENMs released into water bodies (up to 1 mg Kg -1 year -1 ) [START_REF] Sun | Comprehensive probabilistic modelling of environmental emissions of engineered nanomaterials[END_REF] , especially near large cities and related industry, in areas that favour deposition like wetlands and marshes. This may cause chronic toxicity in emergent wetland plants (helophytes), which play vital ecological roles [START_REF] Caffrey | Factors influencing the distribution of aquatic plant communities in Irish canals[END_REF] . Exposure to ZnO-ENMs decreases photosynthesis, antioxidant activity, and growth in aquatic plants, and increases Zn bioaccumulation and oxidative stress [START_REF] Song | Phytotoxicity and accumulation of zinc oxide nanoparticles on the aquatic plants Hydrilla verticillata and Phragmites Australis: leaf-type-dependent responses[END_REF][START_REF] Zarate-Cruz | Phytotoxicity of ZnO nanoparticles on the aquatic fern Azolla filiculoides Lam[END_REF][START_REF] Zhang | Phytotoxicity and bioaccumulation of ZnO nanoparticles in Schoenoplectus tabernaemontani[END_REF][START_REF] Hu | Evaluation of growth and biochemical indicators of Salvinia natans exposed to zinc oxide nanoparticles and zinc accumulation in plants[END_REF][START_REF] Hu | Biochemical responses of duckweed (Spirodela polyrhiza) to zinc oxide nanoparticles[END_REF][START_REF] Thwala | The oxidative toxicity of Ag and ZnO nanoparticles towards the aquatic plant Spirodela punctata and the role of testing media parameters[END_REF][START_REF] Chen | Time Matters: the Toxicity of Zinc Oxide Nanoparticles to Lemna minor L. Increases with Exposure Time[END_REF] . However, studies on helophytes are few and the uptake and toxicity of ZnO-ENMs, especially after chronic exposure, are poorly characterised in these plants.

There is an unresolved controversy about the capacity of entire ZnO-ENMs to enter roots. In several crops, ZnO-ENMs have been reported in the root epidermis, cortex, endodermis, stele, lateral roots, and on the root surface [START_REF] Chen | Phytotoxicity and bioaccumulation of zinc oxide nanoparticles in rice (Oryza sativa L.)[END_REF][START_REF] Wang | Fate of ZnO Nanoparticles in Soils and Cowpea (Vigna unguiculata)[END_REF][START_REF] Da Cruz | Shedding light on the mechanisms of absorption and transport of ZnO nanoparticles by plants via in vivo X-ray spectroscopy[END_REF][START_REF] Lv | Accumulation, speciation and uptake pathway of ZnO nanoparticles in maize[END_REF][START_REF] Lin | Root Uptake and Phytotoxicity of ZnO Nanoparticles[END_REF][START_REF] Hernandez-Viezcas | Situ Synchrotron Xray Fluorescence Mapping and Speciation of CeO2 and ZnO Nanoparticles in Soil Cultivated Soybean (Glycine max)[END_REF] . At the cellular level, ZnO-ENMs have accumulated in the intercellular spaces, along the plasma membrane, and in the cytoplasm, vacuoles and nuclei [START_REF] Chen | Phytotoxicity and bioaccumulation of zinc oxide nanoparticles in rice (Oryza sativa L.)[END_REF][START_REF] Lv | Accumulation, speciation and uptake pathway of ZnO nanoparticles in maize[END_REF] . It has been suggested that ZnO-ENMs in the root apoplast can enter the symplast by endocytosis [START_REF] Chen | Phytotoxicity and bioaccumulation of zinc oxide nanoparticles in rice (Oryza sativa L.)[END_REF] and pass to the xylem directly from the apoplast, through lateral roots with immature Casparian bands [START_REF] Zhang | Phytotoxicity and bioaccumulation of ZnO nanoparticles in Schoenoplectus tabernaemontani[END_REF][START_REF] Lv | Accumulation, speciation and uptake pathway of ZnO nanoparticles in maize[END_REF] . However, few studies have confirmed either the elemental composition of the candidate ENMs or the Zn speciation in roots. Wang et al found 65% ZnO and 32% Zn-histidine in the roots of hydroponically grown cowpea exposed to ZnO-ENMs [START_REF] Wang | Fate of ZnO Nanoparticles in Soils and Cowpea (Vigna unguiculata)[END_REF] .

Nonetheless, Da Cruz et al demonstrated that ZnO was only found inside roots when they had been previously damaged [START_REF] Da Cruz | Shedding light on the mechanisms of absorption and transport of ZnO nanoparticles by plants via in vivo X-ray spectroscopy[END_REF] . Accordingly, ZnO-ENMs normally do not reach the shoots unless roots are damaged [START_REF] Wang | Fate of ZnO Nanoparticles in Soils and Cowpea (Vigna unguiculata)[END_REF][START_REF] Da Cruz | Shedding light on the mechanisms of absorption and transport of ZnO nanoparticles by plants via in vivo X-ray spectroscopy[END_REF][START_REF] Lv | Accumulation, speciation and uptake pathway of ZnO nanoparticles in maize[END_REF][START_REF] Lin | Root Uptake and Phytotoxicity of ZnO Nanoparticles[END_REF] . Intact roots generally contain no ZnO but do contain other Zn species like Zn-phosphates, Zn-citrate, Zn-malate, Zn-histidine, or Zn-nitrate [START_REF] Da Cruz | Shedding light on the mechanisms of absorption and transport of ZnO nanoparticles by plants via in vivo X-ray spectroscopy[END_REF][START_REF] Lv | Accumulation, speciation and uptake pathway of ZnO nanoparticles in maize[END_REF][START_REF] Hernandez-Viezcas | Situ Synchrotron Xray Fluorescence Mapping and Speciation of CeO2 and ZnO Nanoparticles in Soil Cultivated Soybean (Glycine max)[END_REF] . In the aquatic plant Schoenoplectus tabernaemontani, ZnO-ENMs were reportedly found around plastids in the roots, arranged like beads on a string [START_REF] Zhang | Phytotoxicity and bioaccumulation of ZnO nanoparticles in Schoenoplectus tabernaemontani[END_REF] . However, this observation could be better explained as endoplasmic reticulum wrapped around plastids [START_REF] English | Interconnections with Other Organelles[END_REF] .

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The copyright holder for this preprint (which was not peer-reviewed) is the . https://doi.org/10.1101/2020.04.08.031179 doi: bioRxiv preprint

The present research aims to give a comprehensive insight on ZnO-ENM uptake and chronic toxicity in P. australis at the levels estimated by models in current sediments, with an ample margin for a future increase in concentrations. This is crucial because emissions are expected to continue growing. To do so, we determined whether entire ZnO-ENMs enter P. australis roots by combining transmission electron microscopy (TEM), X-ray microanalysis, dark-field mapping, and Zn stable isotope analysis in different plant organs. Zinc stable isotopes are valuable to understand Zn uptake and toxicity in plants [START_REF] Caldelas | Zinc isotopic fractionation in Phragmites australis in response to toxic levels of zinc[END_REF][START_REF] Aucour | Fractionation of stable zinc isotopes in the zinc hyperaccumulator Arabidopsis halleri and nonaccumulator arabidopsis petraea[END_REF] . The Zn isotopic signature of plant tissue integrates all the processes causing Zn fractionation throughout a plant's life and it is a proxy of Zn speciation and the activity of Zn membrane transporters [START_REF] Caldelas | Zinc Homeostasis and isotopic fractionation in plants: a review[END_REF] . Hence, Zn isotopes can be used to determine whether Zn was first taken up as either Zn 2+ or ZnO-ENMs, even after they form other Zn species. Additionally, we unravelled the mechanisms of ZnO-ENM toxicity on the photosynthetic apparatus and fully characterised the physiological response of P. australis to ZnO-ENMs.

Methods

Nanomaterials and non-nano ZnO

Four ZnO materials were purchased from Sigma-Aldrich: non-nano ZnO (96479 Fluka, ACS reagent ≥99.9%); spherical particle nanopowder <100 nm in diameter (544906 Aldrich, ≈80%); <50 nm nanopowder particles of various shapes (677450 Aldrich, >97%); and nanowires 50 nm in diameter x 300 nm long (773980 Aldrich). These materials will be referred to as Bulk, NP100, NP50, and NW, respectively. To determine their chemical composition, 2 ml HNO 3 (69.0-70.0% Instra reagent, 9598-34 Baker) was added to 50 mg aliquots and digested overnight at 90°C. The extracts were brought to 100 ml with 1% HNO 3 and analysed for Al, Cd, Cr, Cu, Fe, Ni, Pb, and Zn content by inductively coupled plasma optical emission spectrometry (ICP-OES) using a Perkin Elmer Optima-8300 (Waltham, MA, USA). Three replicates were analysed for each ZnO material. To study particle size and shape, samples were observed on a field emission scanning electron microscope (FESEM) JEOL JSM 7100F at 20.0 kV, and secondary electron images were taken.

Plant growth conditions

Phragmites australis (Cav.) Trin. Ex Steud plants were purchased from a local nursery (Tres Turons, Sabadell, Spain). Plant cultivation took place in the greenhouse of the Experimental Field Services of the University of Barcelona (UB). Roots were washed to remove the original substrate and shoots cut to induce new growth. Plants were initially grown in trays filled with modified half-strength Hoagland's solution as detailed in [START_REF] Caldelas | Zinc isotopic fractionation in Phragmites australis in response to toxic levels of zinc[END_REF] . The pH was adjusted to 6.5 and solution was replaced every three days. After 7 weeks, plants were rinsed in distilled water, selected within a small range of fresh weight (11.99 ± 0.23 g, mean ± standard error, n = 95) and height (27.54 ± 0.34 cm), and placed in individual 1-gallon (3.895 l) glass pots. These were filled with modified half-strength Hoagland's solution with no added Zn according to [START_REF] Emanuel | Mineral nutrition of plants: principles and perspectives[END_REF] (Supplementary Table 1), at pH 5.9. Each pot was wrapped in aluminium foil to limit algal growth. After 10 days, the ZnO materials described above were added at a concentration of 0, 0.1, 1, 10, 100, or 1000 mg l -1 . Nanowires were only tested up to 10 mg l -1 due to their high cost.

Four pots per treatment were distributed randomly on the greenhouse table, and the nutrient solution was replenished three times a week. Five pots without plants but containing the same solution were included. Plants were grown for 14 weeks, from November 2016 to February 2017. The mean temperature was 17.9±0.22°C, the relative humidity 51.0±1.06%, and the maximum PPFD (photosynthetic photon flux density) ~500 µmol m -2 s -1 . The pH and electrical conductivity (EC) of the solutions were controlled weekly in a sub-sample of 10 pots (results not shown) and recorded for the whole experiment on weeks 8 and 12 (Supplementary Table 2).

Elemental analyses

At the end of the experiment, 10 ml aliquots of the growth solutions were centrifuged for 10 minutes at 9065 relative centrifugal force in a Hettich 32R centrifuge (Tuttlingen, Germany) with a 1620A angle rotor. Supernatants were filtered using Whatman® Grade GF/A glass microfibre filters (WHA1820042) and 100 µl HNO 3 was added (Baker Instra-Analysed Reagent, 69.0-70.0%, 9598-34). The Zn and Al contents of the solutions were measured by ICP-OES.

Plants were thoroughly washed in distilled water, patted dry, and weighed. Roots and shoots were separated using a scalpel. Samples were oven-dried at 60°C for 48h and finely ground for 2 min in a Retzch MM400 mixer mill at a frequency of 20 s -1 . Next, 0.1 g of the ground material was digested overnight at 90°C in a mixture of 2 ml HNO 3 and 0.5 ml H 2 O 2 (30% Suprapur, 1.07298.1000 Merck). Then 50 µl of HF (40% reagent grade, AC10511000 Scharlab) were added to each sample and digested for an additional 2h at 90°C. Digests were diluted in 30 ml MilliQ water (18.2 Ω) before analysis. Per every 30 samples, 3 blanks and 2 aliquots of the BCR-60 reference material (Lagarosiphon major) (Community Bureau of Reference, Brussels, Belgium) were digested using the same protocol. We obtained 303.1±5.3 (n=6) µg g -1 Zn, in line with the certified values (313.0±8.0). The Al, Ca, Cu, Fe, K, Mg, Mn, P, S, and Zn contents of the extracts were then determined by ICP-OES. To study Zn mass balance at the end of the experiment, Zn in mg was calculated for each reservoir in our culture system (Equation 1):

Zn watering + Zn ZnO = Zn root + Zn shoot + Zn solution + Zn solid The Zn input contributed by the nutrient solution (Zn watering ) was calculated by multiplying the [Zn] of the initial nutrient solution by the total volume of solution added and by the sum of all waterings (E). Zinc input from ZnO materials (Zn ZnO ) was obtained from the [ZnO] of each treatment (0.1, 1, 10, 100, and 1000 mg l -1 ) multiplied by the Zn concentration of each material.

Zinc extracted by roots (Zn root ) and shoots (Zn shoot ) was determined by multiplying [Zn] by dry weight (DW). The total Zn in the final solutions (Zn solution ) was calculated by multiplying [Zn] in the final solutions by the volume of the container. Finally, the Zn in the solid phase of the nutrient media (Zn solid ) was inferred from subtracting Zn root , Zn shoot , and Zn solution from the sum of all Zn inputs. The same procedure was followed to calculate the Al mass balance.

Zinc separation and isotopic analyses

Zinc isotope analyses were carried out in the facilities of the Géosciences Environnement, Toulouse, France. In the clean lab (ISO3), 100 mg of sample were weighed in Teflon beakers.

Samples were digested in several steps: i) 24h at room temperature in 1.5ml HNO 3 and 1 ml H 2 O 2 ; ii) 24h on a hotplate at 80°C with the same mixture, then evaporated; iii) 24h at 80°C in a mixture of 1.2ml HF and 1.2 ml HNO 3 , then evaporated; and finally iv) 24h on a hotplate at 115°C in 20 drops of HCl and 10 drops of HNO 3 , and evaporated. The digests were then refluxed in 15 ml 10% HNO 3 for Zn quantification in an ICP-MS Agilent 7500 (Santa Clara, USA). Three aliquots of BCR-60 were ground and processed in the same manner as the samples to quantify the Zn contribution of the stainless-steel jars and balls. We obtained 298.2±2.1 µg g - 1 Zn for the ground BCR-60 and 301.3±6.9 for the non-ground. These values are slightly lower than the certified 313.0±8.0 µg g -1 Zn, which is likely due to the cleanroom being a much cleaner environment than a regular chemistry laboratory. The sample residual was weighed and refluxed in 1 ml of 7N HCl + 0.001% H 2 O 2 overnight at 40°C. HCl and HNO 3 were double distilled, HF was 41-51% (Fluka A513-P500 suprapur) and H 2 O 2 30% (Merk 1.07298 suprapur). Plastic and Teflon material was acid-washed before use. Zinc separation was performed in Poly-Prep chromatography columns (BIO-RAD, 731-1550) containing 2 ml of anion exchange resin AGMP-1M (100-200 mesh, chloride form) (BIO-RAD, 1411841). The resin was first cleansed three times with 10 ml 18.2Ω water followed by 7 ml 0.5N HNO 3 , then conditioned with 6 ml 7N HCl + 0.001% H 2 O 2 before loading the samples. Matrix elements and Cu were eluted with 30 ml 7N HCl + 0.001% H 2 O 2 . Iron (Fe) was eluted with 10 ml 2N HCl + 0.001% H 2 O 2 and the resin was rinsed in 2 ml of 0.5N HNO 3 . Finally, Zn fractions were eluted on 8 ml of 0.5N HNO 3 and evaporated. Three aliquots of the reference material BCR 281 (ryegrass) were processed in the same manner to check the accuracy of the isotope measurements. Digested ZnO materials were diluted up to 1:3000. Column yield was checked from the Zn intensities during isotope analysis and was 96.6 ± 18.0 % (mean ± 2SD, n=48) for plant samples and 89.7 ± 5.6 (n=12) for ZnO materials. Three samples with bad yields were rejected and were not included in this study. Zinc fractions were concentration-matched to within 10% and measured for Zn isotopic composition in a High-Resolution Multi-Collector ICP Mass Spectrometer (Neptune, Thermofinnigan). Instrument settings are shown in Supplementary Table 3. The Neptune cup configuration was: L4 ( 62 Ni), L2 ( 63 Cu), L1 ( 64 Zn), Central ( 64 Cu), H1 ( 66 Zn), H2 ( 67 Zn), and H3 ( 68 Zn). To correct for mass bias, the samplestandard bracketing procedure as described in [START_REF] Peel | A simple combined sample-standard bracketing and inter-element correction procedure for accurate mass bias correction and precise Zn and Cu isotope ratio measurements[END_REF] was applied, with AA ETH Zn as bracketing standard and copper (Cu) NIST SRM 976 as the external element. Sample and standard measurements were obtained from 40 cycles of 8-second integrations. Samples were measured 2-3 times on different sessions. Repeatability throughout the three sessions was 0.002‰ ± 0.035 (2SD; n = 150), calculated from the bracketing standard. Isotope compositions are expressed in the δ notation (Equation 2):

!" = $% & ' & '( ) -1, × 1000
For a given chemical element (X), R s is the ratio of the heavy isotope to the light isotope of the sample and R st of the standard. All δ 66 Zn in this study were calculated using the 66 and 64 isotopes and expressed relative to JMC Zn. The δ 66 Zn JMC for AA ETH Zn and ryegrass were 0.27 ± 0.04 (2SD; n = 150), and 0.42 ± 0.06 (2SD; n = 6) respectively, which was in agreement with the literature [START_REF] Archer | Inter-calibration of a proposed new primary reference standard AA-ETH Zn for zinc isotopic analysis[END_REF][START_REF] Smolders | Isotopic fractionation of Zn in tomato plants suggests the role of root exudates on Zn uptake[END_REF] .

Root anatomy and root cell ultrastructure

Root tips were cut from controls and plants treated with either 1000 mg l -1 (Bulk, NP100, NP50)

or 10 mg l -1 (NW) for 9 weeks. Root tips were rinsed in distilled water and fixed in an ice-cold mixture of 2.5% glutaraldehyde and 2% paraformaldehyde in 0.1M phosphate buffer (pH 7.4), then stored at 4°C. Samples were exposed to vacuum for 1h to remove air bubbles, washed in phosphate buffer, and postfixed and stained with 1% osmium tetroxide and 0.8% potassium ferrocyanide for 1h. Stained samples were washed in distilled water and dehydrated in an acetone series of increasing concentration to achieve 100%. All the fixation steps were carried out at 4°C. Fixed samples were polymerised in epoxy Spurr resin for 48h at 60°C. For light microscopy, 1 μm semi-thin cross-sections were stained with methylene blue and photographed with a light microscope (Olympus 175 CX41, Tokyo, Japan) coupled with a digital camera (Olympus DP70). For transmission electron microscopy (TEM), 70 nm ultrathin sections were cut with a Reichert-Jung Ultracut E ultramicrotome (C. Reichert AG, Vienna, Austria), stained with uranyl acetate and lead citrate, and observed under a Jeol JEM 1010 (Tokyo, Japan) operated at 100 kV. Images were taken with a Gatan Orius camera (Gatan, Pleasanton, USA).

For microanalysis, unstained cuts were dried and mounted on titanium grids. Cuts were analysed using a JEOL JEM-2100 LaB6 transmission electron microscope equipped with an Energy Dispersed Analysis of X-ray Spectrometer (EDXS), operating at 200 kV in STEM mode using the dark field detector. The beam size used in this mode was around 15 nm. The spectrometer is an INCA x-sight (Oxford Instruments, Abingdon, UK), with a Si (Li) detector.

Micrographs were obtained using a Gatan Orius SC1000 CCD camera with Digital Micrograph Version 1.71.38 software. Map acquisition was accomplished using the INCA Microanalysis Suite version 4.09 software. X-ray maps were obtained by selecting Zn kα1 as the characteristic X-ray peak.

Evapotranspiration and photosynthetic performance

To calculate the water consumption from each pot (E, in g), the nutrient solution added during each watering was weighed. Chlorophyll content on a leaf area basis was measured with a SPAD-502 portable chlorophyll meter (Minolta, Illinois, USA), as reported elsewhere [START_REF] Krugh | The solid-state chlorophyll meter: a novel instrument for rapidly and accurately determining the chlorophyll concentrations in seedling leaves[END_REF] . Five representative mature leaves of each plant were measured at 2 cm from the base. The leaf gas exchange and chlorophyll fluorescence were determined using a Li-COR 6400 portable photosynthesis system (Li-COR Inc., Lincoln, NE, USA) running OPEN version 4.06. The third fully developed leaf of the healthiest shoot of each plant was measured at approximately 2 cm from the base. Leaves were first dark-adapted for 30 min to measure maximum quantum yield (F v /F m ). The same leaves were then re-acclimated to environmental light until stabilised (up to 45 min) to determine relative quantum yield (F v '/F m '), quantum yield of photosystem-II photochemistry (ΦPSII) [START_REF] Genty | The relationship between the quantum yield of photosynthetic electron transport and quenching of chlorophyll fluorescence[END_REF] , quantum yield of CO 2 fixation (ΦCO 2 ), the electron transport rate (ETR, μmol m -2 s -1 ), photochemical (qP) and non-photochemical quenching (qN, NPQ), the light-saturated net CO 2 assimilation rate (A s , μmol CO 2 m -2 s -1 ), stomatal conductance to water (g s , mol H 2 O m -2 s -1 ), intercellular CO 2 concentration (C i , μmol CO 2 mol air -1 ), the transpiration rate (E, mmol H 2 O m -2 s -1 ), and water vapour pressure deficit of the leaf (VPD, kPa).

Measurements were taken under a saturating light (photosynthetic photon flux density of 1200 µmol photons m -2 s -1 ), 400 µmol mol -1 of CO 2 , and an air temperature of 25.7±0.1°C.

Carbon and nitrogen isotopic relation in plants

For each plant sample, 0.8-0.9 mg of finely ground dry matter were weighed in tin capsules (Lüdiswiss, Flawil, Switzerland). The total C and N contents were analysed using an Elemental Analyser (EA, Carlo Erba 2100, Milan, Italy), which was interfaced with an Isotope Ratio Mass Spectrometer (IRMS, Thermo-Finnigan Deltaplus Advantage, Bremen, Germany) to analyse the

Results

Characterisation of ZnO sources

Zinc purity was very similar in all four ZnO sources: 94.2% for Bulk, 96.3% for NP100, 90.2%

for NP50, and 94.5% for NW (Supplementary Table 4). According to the manufacturer, Zn purity was 99.9%, ~80%, and 91% for Bulk, NP100, and NP50 respectively (no information was given for NW). Besides, 6% Al was reportedly added to NP50 as a dopant but only 2.2%

was measured. Traces of other unreported elements were also found: up to 27.2 µg g -1 Pb (NP50), 8.5 µg g -1 Fe, 4.5 µg g -1 Cu, 3.4 µg g -1 Ni, 1.2 µg g -1 Cd, and 0.7 µg g -1 Cr. Average particle length was 215±4 nm (mean±SE, n = 946) for bulk ZnO, with 10% of particles in the nano range. For NP100, the mean particle length was 99±2 nm (n = 956), but 35% of the particles were >100 nm. Similarly, NP50 diameter was 48.7±0.57 nm (n = 467), with 39% of the particles in the 50-100 nm range. Finally, the mean diameter of NW was 59±0.5 nm (n = 223).

Zinc and Al dissolution and uptake

Zinc concentrations in the growth solution ([Zn] sol ), roots ([Zn] root ), and shoots ([Zn] shoot ) increased with increasing ZnO supply (Fig. 1, Supplementary Table 5), reaching 20x, 231x, and 60x those of the controls at 1000 mg l -1 , respectively (P<0.001). Controls had similar [Zn] root and [Zn] shoot (~30 µg g -1 ). By contrast, plants treated with ZnO accumulated Zn preferentially in the roots (7.6 mg g -1 in roots vs. 1.8 mg g -1 in shoots at the highest ZnO supply) (Fig. 1B,C). At 100 and 1000 mg l -1 , NP50 released up to twice as much Zn into solution as bulk and NP100 (P =0.019 and 0.011, respectively) (Fig. 1A).

[Zn] shoot in plants treated with 1000 mg l -1 NP50 reached 3.1 mg g -1 , twice as much as bulk and NP100 plants (P =0.034) (Fig. 1C), and [Zn] root reached 10.4 mg g -1 Zn, 60% higher than bulk and 80% higher than NP100 plants (Fig. 1B). The mass balance was calculated to assess the fate of Zn in our system. The proportion of Zn in the solid fraction at the end of the experiment (Zn solid ) increased with increasing [ZnO] (P<0.001) (Fig. 2). At 100 and 1000 mg l -1 , the majority of Zn (85 -99 %) remained in the solid fraction.

By contrast, the dissolved Zn fraction (Zn sol ) decreased with increasing ZnO (P<0.001). The fraction of Zn stored in roots (Zn root ) and shoots (Zn shoot ) gradually increased up to 10 mg l -1 and decreased at the two highest concentrations (P<0.001). At 100 and 1000 mg l -1 , Zn sol was the highest in NP50 treatments (9.8 and 2.9%, respectively). The maximum Zn root was 16% and Zn shoot 8%, both at 10 mg l -1 NP50.

To study the uptake of doping elements by plants and the capacity of ENMs to retain metals from solution, Al content and mass balance were investigated. At 1000 mg l -1 ZnO (P<0.001) (Supplementary Table 5) the [Al] roots was 3.5x higher than the control. The only Al-doped material, NP50, resulted in the highest [Al] root (up to 195 mg g -1 at 1000 mg l -1 , P=0.022). The

[Al] shoots increased from the 10 mg l -1 ZnO treatments and beyond, with a maximum increase of 30% at 1000 mg l -1 ZnO (all P<0.001). However, the mass balance calculations indicated that the Al missing from solution mostly ended up in the solid fraction (Al solid ), which increased progressively with [ZnO] (P<0.001) (Supplementary Fig. 1). NP50 showed the highest Al solid (P<0.001, Supplementary Fig. 1C), starting with 98.9% at 0.1 mg l -1 . For the rest of the ZnO materials, at 10 mg l -1 the Al solid was 42% in Bulk (Supplementary Fig. 1A), 59% in NP100 (Supplementary Fig. 1B), and 78% in NW (Supplementary Fig. 1D). The solution pH progressively increased in pots containing plants, reaching 8.2 by the end of the experiment, while pots without plants had a pH value of 5.5. At week 8, the pH of the 1000 mg l -1 solutions was 1 pH unit higher than controls and close to neutrality (P<0.001) (Supplementary Table 2).

The pH was highest in NP100 treatments and lowest with NW (P=0.011). At the end of the experiment, the solution pH was higher for plants under 1000 mg l -1 ZnO and controls than for the rest of concentrations, with no difference between sources.

Zn isotopic fractionation

The δ 66 Zn of all ZnO materials was very similar, on average 0.33 ± 0.04 ‰ (n = 10, Fig. 3, Supplementary Table 6). By contrast, the δ 66 Zn root showed clear differences between treatments (P=0.003). Bulk and NP100 plants had the lightest δ 66 Zn root (0.02 and 0.04‰ a , respectively), followed by NP50 (0.13‰ ab ), and NW and controls (0.35 and 0.38‰ b , respectively). The δ 66 Zn shoot also differed between ZnO treatments (P=0.027, Fig. 3). The shoots of plants treated with bulk ZnO had the lightest δ 66 Zn shoot (-0.61 ‰ a ), while the rest of the ZnO treatments ranged -0.5 to -0.32‰ ab , and controls had the heaviest δ 66 Zn shoot (0.27‰ b ). The root-to-shoot fractionation (ΔZn shoot-root ) was very small in controls (-0.08‰), which had no Zn added to the nutrient solution. For the rest of the treatments, ΔZn shoot-root ranged from -0.71 to -0.52‰, with no statistically significant difference among them.

Plant growth and evapotranspiration

ZnO caused severe, dose-dependent effects on plant growth and evapotranspiration (ET). Fresh weight (FW) was reduced by up to 74% in whole plants, 66% in roots, and 88% in shoots, whereas FW root / FW shoot was up to 2.6x higher (all P<0.001) (Supplementary Table 7). In agreement, dry weight (DW) greatly decreased in response to ZnO, up to 61% in roots and 83% in shoots (both P<0.001). The DW root / DW shoot ratio increased up to 2.1x (P<0.001). Plant height decreased up to 49% in response to ZnO (P<0.001). Root length also decreased with increasing ZnO, with a 61% reduction at 1000 mg l -1 (P<0.001) (Supplementary Table 7). Mean root length was lowest in NP50 plants (P =0.005). The rest of the growth parameters measured did not show any significant differences between sources. The ET decreased up to 59% with ZnO (P<0.001). ET from plants at 1000 mg l -1 was 1078±41 g of solution (n = 14), which was very similar to pots without plants (992±24, n = 5), evidencing a strong inhibition of plant transpiration. Significant effects on growth and ET started from 1 mg l -1 (DW shoot , DW root / DW shoot , ET), 10 mg l -1 (all FW, DW root , height), or 100 mg l -1 (root length), but the trend was often present at 0.1 mg l -1 . The δ 13 C increased in shoots at [ZnO] ≥100 mg l -1 (P<0.001) (Supplementary Table 8). A similar trend was observed in roots but it did not attain significance.

Photosynthetic performance

The chlorophyll content of mature leaves decreased in response to all ZnO materials and was 25% lower than controls at 1000 mg l -1 (P<0.001, Supplementary Table 9). Accordingly, photosynthetic efficiency was severely affected. A s and ΦCO 2 decreased from 10 mg l -1 and dropped by 79% and 75% respectively at 1000 mg l -1 (P<0.001) (Fig. 4, Supplementary Table 9). The ΦPSII, qP, and ETR also decreased dramatically, up to 80% at 1000 mg l -1 (all P<0.001). By contrast, F v /F m had only a minor reduction (6%) (P=0.002), whereas NPQ increased by 4.4x at 1000 mg l -1 (P=0.005). The ΦCO 2 , F v /F m , ΦPSII, qP, and ETR were lower in plants under the Bulk treatment, followed by the NP50 treatment (P=0.046, 0.038, 0.002, 0.014, and 0.002, respectively, Fig. 4, Supplementary Table 9). Gas exchange was also affected by ZnO: g s and E decreased up to 78% and 72%, respectively (both P<0.001), while VPD was 20% higher than controls at 1000 mg l -1 (P=0.002). Finally, F v '/F m ' was not affected by ZnO.

Root anatomy and ultrastructure

Initial light microscope exploration revealed extensive damage caused by ZnO to the roots: detachment of the epidermis, disorganisation and thickening of cell walls, and vacuolisation and death of root cells, which is indicative of increased aerenchyma formation (Supplementary Fig. 2). Plants treated with NW showed the most severe effects, with generalised apoptosis of the cortex cells that made further characterisation difficult. On the 100Kev TEM, the root epidermis showed significant loss of cell wall material in the Bulk, NP100, and NP50 treatments (Supplementary Fig. 3). The epidermal cells in the Bulk and NP50 treatments exhibited loss of turgor, protoplasm shrinkage, and accumulation of electron-dense granules in the vacuoles. In the cortex, cell-wall thickening, protoplasm shrinkage, and a high number of starch granules or amyloplasts were observed in all three treatments (Supplementary Fig. 4). Additionally, electron-dense precipitates accumulated in the intercellular spaces (Supplementary Fig. 5).

Microanalysis confirmed the presence of Zn in various locations: i) a few granules on the root surface in NP50 and NP100 that are morphologically consistent with NPs (Fig. 5); ii) large amorphous precipitates between cell walls and plasma membrane in the cortex of NP50, NP100, and Bulk plants (in order of abundance, Fig. 67), where most of Zn accumulated; and iii) small vacuoles containing Zn, P, and sometimes Ca in the rhizodermis and cortex of NP50 plants (Fig. 6).

Nutrient content and distribution

Nitrogen content changed in response to ZnO, with a N shoot increase of up to 24% (P<0.001, Supplementary Table 8). The C shoot /N shoot ratio decreased progressively in response to increasing ZnO, reaching a 17% reduction at 1000 mg l -1 (P<0.001). ZnO slightly increased C root (~5% at 1000 mg l -1 , P =0.023) and decreased C shoot (~3% at 100 mg l -1 , P =0.013). The C root /C shoot ratio increased progressively with increasing [ZnO], up to 7% (P =0.023). In the roots, [Mn] and [Fe] greatly increased in response to ZnO, up to 79% and 99%, respectively, at 1000 mg l -1 (P=0.012 and 0.004, Supplementary Table 9). Conversely, [K] showed a progressive decrease that reached 41% at 1000 mg l -1 ZnO (P<0.001). Phosphorus in the roots increased in response to low ZnO, then decreased up to 25% at 1000 mg l -1 (P<0.001). In the shoots, [P] was higher than controls at 10 mg l -1 ZnO and above, with a peak 34% increase at 100 mg l -1 (P<0.001).

Besides, both [S] and [K] increased in response to 10-100 mg l -1 ZnO, attaining levels 32% and 14% higher, respectively, than controls at 100 mg l -1 (P<0.001 and P=0.020). Copper in shoots increased at 0.1 and 1 mg l -1 ZnO, then gradually decreased across treatments, reaching a 30% reduction at 1000 mg l -1 (P<0.001). Manganese in shoots progressively decreased in response to ZnO and was 66% lower than controls at the highest ZnO level (P<0.001). However, [Mn] shoot was lower in the NP50 and NP100 treatments compared to the other ZnO sources (P<0.001).

Discussion

Route of plant exposure to ZnO-ENMs

Our results show that ZnO-ENMs dissolve slowly in the chosen experimental conditions and that the majority was still solid at the end of the experiment (Fig. 2). Dissolved Zn reached only 4.3-7.2 mg l -1 in 100 mg l -1 ZnO-ENM treatments (Fig. 1). This range closely agrees with dissolution experiments by Reed and co-workers, where 100 mg l -1 ZnO-NP suspended in water for 60 days released 2.2-7.4 mg l -1 into solution [START_REF] Reed | Solubility of nano-zinc oxide in environmentally and biologically important matrices[END_REF] . Smaller NPs are known to dissolve more easily due to their larger surface area [START_REF] Mudunkotuwa | Dissolution of ZnO Nanoparticles at Circumneutral pH: A Study of Size Effects in the Presence and Absence of Citric Acid[END_REF] , which explains why [Zn] sol was higher in the NP50 treatment than in the NP100 and bulk treatments (Fig. 1). Solution pH was ~6 initially but gradually increased up to 7.5-8.2 in the presence of plants. This pH range is typical of wetland waters [START_REF] Gerla | Can pH and electrical conductivity monitoring reveal spatial and temporal patterns in wetland geochemical processes?[END_REF] . In our study, the proportion of undissolved Zn and Al increased with increasing ZnO concentrations and solution pH. Accordingly, removal of Zn, Cd, Cu, Hg, Ni, and Pb from solution by ZnO-ENMs has been reported to increase with pH and increasing sorbent mass [START_REF] Mahdavi | Removal of heavy metals from aqueous solutions using Fe3O4, ZnO, and CuO nanoparticles[END_REF][START_REF] Sheela | Kinetics and thermodynamics studies on the adsorption of Zn(II), Cd(II) and Hg(II) from aqueous solution using zinc oxide nanoparticles[END_REF] .

The effect of pH on metal removal is explained by the increased deprotonation of the surface and attraction between negative hydroxyl sites and cations, while the effect of the sorbent mass is due to the increased number of binding sites. Aluminium is frequently added as a dopant to ZnO-NPs to enhance their electrical and optical properties [START_REF] Zhang | Aluminum-doped zinc oxide powders: synthesis, properties and application[END_REF] , so the Al mass balance in our system can serve as a proxy for the fate of doping elements. In our study, most of the Al ended up in the solid fraction, which increased with increasing ZnO, while the dissolved Al fraction decreased in the presence of ZnO (Supplementary Fig. 1). However, [Al] was higher in plants grown at high [ZnO], especially for Al-doped NP50. This is consistent with the "Trojan horse effect" hypothesis, which states that ENMs might bind to other chemicals in solution and enhance their uptake by plants, leading to increased phytotoxicity. Aluminium is toxic to plants, with a reduction in root elongation as the main symptom [START_REF] Imadi | Plant Metal Interaction[END_REF] . The increased uptake of Zn and Al could explain the stronger reduction in root length in NP50 plants as compared with other treatments (Supplementary Table 6). Additionally, the capacity of ZnO-ENMs for metal removal from solution might have affected the bioavailability of essential nutrients, as seen from the changes in nutrient content.

In all ZnO treatments, roots showed intense vesicularisation at the plasma membrane, but the vesicles did not contain either ZnO-ENMs or Zn. Instead, Zn was found mostly in amorphous precipitates outside the plasma membrane, and in NP50 plants alone, in small vacuoles containing Zn, Ca, and P (Fig. 6). Very few ZnO-ENMs were found attached to the root surface and none were detected in the cytoplasm (Fig. 5). These observations indicate that ZnO-ENMs dissolve and the roots take up Zn 2+ , which they mostly immobilise as Zn precipitates in the apoplast and Zn phosphates in small vacuoles. There is abundant evidence in the literature that plant roots sequester excess Zn as precipitates in the intercellular spaces or store Zn in vacuoles bound to phosphates, organic acids or phytochelatins [START_REF] Straczek | Zinc distribution and speciation in roots of various genotypes of tobacco exposed to Zn[END_REF][START_REF] Song | Phytochelatin-metal(loid) transport into vacuoles shows different substrate preferences in barley and Arabidopsis[END_REF][START_REF] Monsant | In vivo speciation of zinc in Noccaea caerulescens in response to nitrogen form and zinc exposure[END_REF][START_REF] Sarret | Forms of zinc accumulated in the hyperaccumulator Arabidopsis halleri[END_REF][START_REF] Van Steveninck | Deposition of Zinc Phytate in Globular Bodies in Roots of Deschampsia caespitosa Ecotypes; a Detoxification Mechanism?[END_REF][START_REF] Neumann | Silicon and heavy metal tolerance of higher plants[END_REF] . Plants exposed to ZnO-ENMs also accumulate Zn in roots in the form of amorphous phosphates or phytates [START_REF] Wang | Fate of ZnO Nanoparticles in Soils and Cowpea (Vigna unguiculata)[END_REF][START_REF] Lv | Accumulation, speciation and uptake pathway of ZnO nanoparticles in maize[END_REF][START_REF] Moghaddasi | Effects of coated and non-coated ZnO nanoparticles on cucumber seedlings grown in gel chamber[END_REF] . In our study, Zn and P only co-occurred in the vacuoles of NP50 plants (Fig. 6). The higher [Zn] achieved in the roots of NP50 plants might have activated vacuolar sequestration. Alternatively, NPs of this size might have been captured by endocytosis and then dissolved inside the vacuole and sequestered as phosphates. The endocytosis of entire 50 nm ZnO-NPs has been reported in hydroponically grown rice [START_REF] Chen | Time Matters: the Toxicity of Zinc Oxide Nanoparticles to Lemna minor L. Increases with Exposure Time[END_REF] . However, our results do not support this explanation because no NPs were detected inside the cells in our study.

The Zn isotope data support the predominant uptake of Zn 2+ , followed by sequestration of excess Zn as precipitates and complexes in root cells. The main causes of Zn isotopic fractionation in plants are Zn speciation, compartmentalisation, and the activity of membrane transporters [START_REF] Caldelas | Zinc Homeostasis and isotopic fractionation in plants: a review[END_REF] . Divalent Zn uptake by low-affinity transporters at the plasma membrane favours the light isotopes, which diffuse faster across membranes thanks to their smaller size [START_REF] Caldelas | Zinc Homeostasis and isotopic fractionation in plants: a review[END_REF][START_REF] John | Zinc isotope fractionation during highaffinity and low-affinity zinc transport by the marine diatom Thalassiosira oceanica[END_REF] . Hence, the relative accumulation of light Zn isotopes in the Bulk, NP100, and NP50 roots compared to ZnO (Fig. 3) can be explained by ZnO dissolution followed by Zn 2+ uptake mediated by membrane transporters. However, NP50 roots were to some extent enriched in heavy isotopes compared to NP100 and bulk roots, although this tendency was not significant. This can be attributed to a greater number and size of Zn precipitates in the intercellular spaces of NP50 roots, which also contained more Zn relative to other treatments, and to the presence of Zn-rich vacuoles in the cortex (Fig. 67). The formation of Zn precipitates and complexes in roots during the plant response to excess Zn 2+ left an isotopically lighter pool of Zn 2+ for transportation to the shoot and favoured the accumulation of heavy isotopes in the root [START_REF] Fujii | Ab initio calculation of the Zn isotope effect in phosphates, citrates, and malates and applications to plants and soil[END_REF][START_REF] Markovic | Experimental determination of zinc isotope fractionation in complexes with the phytosiderophore 2'-deoxymugeneic acid (DMA) and its structural analogues, and implications for plant uptake mechanisms[END_REF][START_REF] Caldelas | Zinc isotopic fractionation in Phragmites australis in response to toxic levels of zinc[END_REF] . Nevertheless, the δ 66 Zn of NP50 roots remained lighter than the starting ZnO because this isotopic composition was still dominated by the initial transfer through the membrane, which favours light isotopes. NW roots were very damaged and had lost their structural coherence, which explains why these roots had an isotopic composition that equalled the ZnO of the NW. Further, the shoots became equally depleted in heavy isotopes in all treatments relative to the roots: ΔZn shoot-root was -0.64±0.22, -0.52±0.25, -0.57±0.12, and -0.71±0.01 for Bulk, NP100, NP50, and NW50, respectively. The magnitude and direction of ΔZn shoot-root are in the same range as previously observed in P.

australis in response to ZnCl 2 toxicity [START_REF] Caldelas | Zinc isotopic fractionation in Phragmites australis in response to toxic levels of zinc[END_REF] . Depletion of heavy Zn isotopes during Zn transport to the shoot is attributed to preferential sequestration of heavy isotopes in the root, the activity the membrane transporters during Zn loading into the xylem, and bulk flow [START_REF] Caldelas | Zinc isotopic fractionation in Phragmites australis in response to toxic levels of zinc[END_REF][START_REF] Viers | Evidence of Zn isotopic fractionation in a soil-plant system of a pristine tropical watershed (Nsimi, Cameroon)[END_REF][START_REF] Couder | Transpiration flow controls Zn transport in Brassica napus and Lolium multiflorum under toxic levels as evidenced from isotopic fractionation[END_REF] . Conversely, there is no isotopic fractionation when Zn is transported in complexes up the shoot due to the bigger mass of the complexes [START_REF] Weiss | Isotopic discrimination of zinc in higher plants[END_REF] . The same likely applies to ZnO-ENMs. Hence, our isotope data indicate that Zn was predominantly transported up the shoot as Zn 2+ , regardless of the ZnO source, and that plants roots are an effective barrier against ZnO-ENM transport up the shoot, which is in agreement with our TEM observations. If ZnO-ENM had been transported through the plant without dissolution, then no isotopic fractionation would have been detected during Zn uptake and transport in the plants. ZnO-NPs in the apoplast reportedly can reach the symplast by endocytosis [START_REF] Chen | Phytotoxicity and bioaccumulation of zinc oxide nanoparticles in rice (Oryza sativa L.)[END_REF] or through the discontinuity of Casparian bands [START_REF] Zhang | Phytotoxicity and bioaccumulation of ZnO nanoparticles in Schoenoplectus tabernaemontani[END_REF][START_REF] Lv | Accumulation, speciation and uptake pathway of ZnO nanoparticles in maize[END_REF] . However, ZnO-NPs in our study were found on the surface of the root but not in the apoplast, indicating that the rhizodermis was an effective barrier. Similarly, recent research shows that ZnO-NPs are unlikely to reach the shoots unless roots are damaged [START_REF] Da Cruz | Shedding light on the mechanisms of absorption and transport of ZnO nanoparticles by plants via in vivo X-ray spectroscopy[END_REF] . Finally, the fractionation of Zn isotopes in control plants is consistent with the plants' response to Zn-deficiency, which results in Zn uptake and transport to the shoots in the form of Zn complexes, with root exudates enriched in heavy isotopes [START_REF] Arnold | Evidence for the mechanisms of zinc uptake by rice using isotope fractionation[END_REF] . The δ 66 Zn of all the ZnO materials was remarkably similar (0.33‰) and the value was consistent with reported ranges for both ZnO-ENMs (-0.31 to 0.28‰) [START_REF] Larner | Evaluation of stable isotope tracing for ZnO nanomaterialsnew constraints from high precision isotope analyses and modelling[END_REF] and natural Zn minerals like hydrozincite (0-0.30‰) [START_REF] Mondillo | A global assessment of Zn isotope fractionation in secondary Zn minerals from sulfide and non-sulfide ore deposits and model for fractionation control[END_REF] .

Phytotoxicity

Chronic exposure to high levels of ZnO-ENMs decreased growth, chlorophyll content, and photosynthetic assimilation in P. australis (Fig. 4, Supplementary Tables 67). Biomass allocation, nutrient uptake and distribution, and fractionation of C isotopes were also altered by ZnO-ENMs (Supplementary Tables 89). Similar effects have been previously described in aquatic macrophytes treated with toxic levels of ZnO-ENMs [START_REF] Song | Phytotoxicity and accumulation of zinc oxide nanoparticles on the aquatic plants Hydrilla verticillata and Phragmites Australis: leaf-type-dependent responses[END_REF][START_REF] Zarate-Cruz | Phytotoxicity of ZnO nanoparticles on the aquatic fern Azolla filiculoides Lam[END_REF][START_REF] Zhang | Phytotoxicity and bioaccumulation of ZnO nanoparticles in Schoenoplectus tabernaemontani[END_REF][START_REF] Hu | Biochemical responses of duckweed (Spirodela polyrhiza) to zinc oxide nanoparticles[END_REF][START_REF] Chen | Time Matters: the Toxicity of Zinc Oxide Nanoparticles to Lemna minor L. Increases with Exposure Time[END_REF][START_REF] Haisel | The effect of nanoparticles on the photosynthetic pigments in cadmium-zinc interactions[END_REF][START_REF] Asztemborska | The Effect of Nanoparticles with Sorption Capacity on the Bioaccumulation of Divalent Ions by Aquatic Plants[END_REF] , Zn 2+ (reviewed by [START_REF] Caldelas | Zinc Homeostasis and isotopic fractionation in plants: a review[END_REF] ), and other metals [START_REF] Caldelas | Accumulation and toxic effects of chromium and zinc in Iris pseudacorus L[END_REF][START_REF] Caldelas | Physiological responses of Eichhornia crassipes [ Mart .] Solms to the combined exposure to excess nutrients and Hg[END_REF][START_REF] Basile | Toxicity, Accumulation, and Removal of Heavy Metals by Three Aquatic Macrophytes[END_REF][START_REF] Costa | Accumulation and effects of copper on aquatic macrophytes Potamogeton pectinatus L.: Potential application to environmental monitoring and phytoremediation[END_REF] . In our previous research on P. australis, exposure to 2 mM ZnCl 2 (131 mg l -1 Zn in solution) for 40 days caused a height reduction of 25% [START_REF] Caldelas | Zinc isotopic fractionation in Phragmites australis in response to toxic levels of zinc[END_REF] . Song and coworkers [START_REF] Song | Phytotoxicity and accumulation of zinc oxide nanoparticles on the aquatic plants Hydrilla verticillata and Phragmites Australis: leaf-type-dependent responses[END_REF] reported a ~50% reduction in height after exposure to 100 mg l -1 ZnO NP50 (only 4.5 mg l -1 Zn in solution) for 35 days 4 . Even though [Zn 2+ ] was lower in the latter study, the effects were more severe and attributed to direct root contact with ZnO-NPs. Accordingly, we report a height decrease of 43% after three months at 100 mg l -1 ZnO (3.5-7.2 mg l -1 in solution), irrespective of particle size. The magnitude of the effect paralleled the Song et al. study despite the longer exposure time and larger particle sizes of the Bulk, NP100, and NW treatments. To sum up, ZnO is more toxic to P. australis than ZnCl 2 . The total ZnO added to the system is the most important factor to explain the impact of ZnO-ENMs on plant growth, rather than [Zn 2+ ] in solution and particle size. The toxicity cannot be attributed to the osmotic effect of the increased [Zn 2+ ]. The conductance (EC) did not increase with increasing ZnO, which remained mostly undissolved. This clearly demonstrates that undissolved ZnO plays a key role in ENM phytotoxicity. To correctly assess the environmental risks of ZnO-ENMs, field studies should include the solid ZnO fraction in sediments, rather than just focus on ZnO-ENM levels in the water. However, particle size was relevant for ZnO-ENM impacts on root growth and [Mn] shoot due to the greater dissolution of smaller NPs.

The lower photosynthetic rates can be explained by stomatal closure and root malfunction leading to water deficit, higher δ 13 C, lower chlorophyll content, and nutritional imbalance. Of particular interest were the changes in Mn content, which greatly decreased in shoots in a dosedependent response to [ZnO] and particle size (Supplementary Table 9). It is unlikely that this decrease in [Mn] shoot is caused by Mn removal from solution by ENMs. The removal efficiency is low for Mn 57 and the [Mn] root was high, proving that Mn uptake was not impaired. We propose that excess Zn 2+ might inhibit or compete for Mn 2+ transporters involved in Mn root-toshoot transport, like AtZIP1 and ATZIP2. These carriers transport Zn and Mn, are expressed in the root vasculature, and mediate Mn radial movement towards the xylem and xylem loading [START_REF] Milner | Transport properties of members of the ZIP family in plants and their role in Zn and Mn homeostasis[END_REF] .

Manganese has a key role as a catalyst of the oxygen-evolving complex of PSII and its deficiency greatly reduces photosynthesis [START_REF] Schmidt | Manganese Deficiency in Plants: The Impact on Photosystem II[END_REF] . Gas exchange and chlorophyll fluorescence data taken at the same time confirm lower electron transport efficiency. E and g s decreased,

indicating that the stomatal opening was strongly inhibited. The reduced gas exchange could lower A s by restricting CO 2 availability. The parallel increase in δ 13 C of shoot and root dry matter also supports this conclusion. However, Ci was not affected in our study. Helophytes take up CO 2 dissolved in water by the roots, and it diffuses towards the photosynthetic tissues via the aerenchyma [START_REF] Brix | Uptake and photosynthetic utilization of sediment-derived carbon by Phragmites australis (Cav.) Trin. ex Steudel[END_REF] . This alternative source of CO 2 can contribute significantly to Ci [START_REF] Constable | High Carbon Dioxide Concentrations in Aerenchyma of Typha latifolia[END_REF][START_REF] Wium-Andersen | Photosynthetic Uptake of Free CO2, by the Roots of Lobelia dortmanna[END_REF] . The small 5% reduction in F v /F m in dark-adapted leaves and the unaltered F v '/F m ' showed that PSII was mostly functional. By contrast, the ΦPSII, qP, ΦCO 2 , and ETR were severely reduced in light-adapted leaves, while qN and NPQ increased. Hence, we attribute the reduced ΦCO 2 and A s to decreased stomatal conductance and lower efficiency of electron transport downstream due to an Mn deficiency that caused PSII to become easily saturated by light. Reduced chlorophyll content, A s , g s , E, C i , F v /F m , qP, and ETR coupled with increased NPQ have been reported in terrestrial plants exposed to toxic levels of ZnO-ENMs [START_REF] Wang | Effects of zinc oxide nanoparticles on the growth, photosynthetic traits, and antioxidative enzymes in tomato plants[END_REF][START_REF] Tripathi | Nitric Oxide Ameliorates Zinc Oxide Nanoparticles Phytotoxicity in Wheat Seedlings: Implication of the Ascorbate-Glutathione Cycle[END_REF] . In agreement, the aquatic plants Azolla filiculoides and Lemna minor have shown severely reduced growth, chlorophyll content and F v /F m after exposure to ZnO-ENMs at pH 4.5-5.5 [START_REF] Zarate-Cruz | Phytotoxicity of ZnO nanoparticles on the aquatic fern Azolla filiculoides Lam[END_REF][START_REF] Chen | Time Matters: the Toxicity of Zinc Oxide Nanoparticles to Lemna minor L. Increases with Exposure Time[END_REF][START_REF] Chen | The toxicity of zinc oxide nanoparticles to Lemna minor (L.) is predominantly caused by dissolved Zn[END_REF] . Remarkably, growth and F v /F m in L. minor were not equally reduced at pH 8, due to the lower NP dissolution [START_REF] Chen | The toxicity of zinc oxide nanoparticles to Lemna minor (L.) is predominantly caused by dissolved Zn[END_REF] . In our study, P. australis progressively basified the growth media, which reached pH~8 at the end of the experiment. This high final pH could explain the limited effect of ZnO on F v /F m . The δ 13 C shoot was within -26.7 to -30.7 ‰, which is in agreement with values previously reported for C3 helophytes [START_REF] Caldelas | Accumulation and toxic effects of chromium and zinc in Iris pseudacorus L[END_REF][START_REF] Chappuis | Decrypting stable-isotope (δ 13 C and δ 15 N) variability in aquatic plants[END_REF] . In C3 plants, diffusion of atmospheric CO 2 through the stomatal pore and C fixation by RuBisCO favour 12 C 67 . During stomatal closure, RuBisCO continues consuming intercellular CO 2 and the δ 13 C shoot increases as [START_REF] Wang | Fate of ZnO Nanoparticles in Soils and Cowpea (Vigna unguiculata)[END_REF] C is depleted [START_REF] Farquhar | On the Relationship Between Carbon Isotope Discrimination and the Intercellular Carbon Dioxide Concentration in Leaves[END_REF] . In our study, δ 13 C shoot and δ 13 C root increased (~1‰) in response to high ZnO. This, in the context of reduced transpiration, is generally caused by Ci exhaustion [START_REF] Farquhar | Carbon Isotope Discrimination and Photosynthesis[END_REF] . However, our gas exchange data do not indicate Ci depletion. While P. australis mainly assimilates atmospheric C, it can also take up CO 2 dissolved in water by the roots [START_REF] Brix | Uptake and photosynthetic utilization of sediment-derived carbon by Phragmites australis (Cav.) Trin. ex Steudel[END_REF] . Plants that assimilate more CO 2 by this route show higher δ 13 C, as seen when comparing submerged aquatic plants relative to helophytes [START_REF] Chappuis | Decrypting stable-isotope (δ 13 C and δ 15 N) variability in aquatic plants[END_REF] . The assimilation of a larger proportion of CO 2 from the roots during stomatal closure can explain a higher δ 13 C while maintaining Ci levels. In agreement, we observed increased root aerenchyma development in response to excess Zn. We found no previous record of δ [START_REF] Da Cruz | Shedding light on the mechanisms of absorption and transport of ZnO nanoparticles by plants via in vivo X-ray spectroscopy[END_REF] C in plants exposed to toxic levels of ZnO-ENMs, although Iris pseudacorus grown in 200 mg l -1 ZnCl 2 showed a similar increase of δ 13 C shoot [START_REF] Caldelas | Accumulation and toxic effects of chromium and zinc in Iris pseudacorus L[END_REF] .

Our experiment shows that ZnO-ENMs are toxic from 1 mg l -1 . Effects at this concentration include reduced growth, allocation of new biomass to the roots and rhizomes instead of the photosynthetic tissues, and restriction of transpiration. These levels are high compared to the few existing records of ZnO-ENMs in surface waters: there are up to 1.84 µg l -1 ZnO NMs in the surface waters of Singapore [START_REF] Majedi | Chemometric Analytical Approach for the Cloud Point Extraction and Inductively Coupled Plasma Mass Spectrometric Determination of Zinc Oxide Nanoparticles in Water Samples[END_REF] and just 5 ng l -1 in Canada 70 . However, recent modelling studies estimate ZnO-ENM concentrations of 10 -6 to 1 µg l -1 in surface waters and 11-32 mg kg - 1 in sediments [START_REF] Sun | Dynamic Probabilistic Modeling of Environmental Emissions of Engineered Nanomaterials[END_REF][START_REF] Garner | Assessing the Risk of Engineered Nanomaterials in the Environment: Development and Application of the nanoFate Model[END_REF][START_REF] Dumont | Nano silver and nano zinc-oxide in surface waters -exposure estimation for Europe at high spatial and temporal resolution[END_REF] . Current levels predicted for the sediments would cause substantial toxic effects on P. australis. Besides, the said models have important limitations, and the real concentrations are likely to be locally higher and to increase in coming years [START_REF] Lead | Nanomaterials in the environment: Behavior, fate, bioavailability, and effects-An updated review[END_REF] .

Conclusion

Emerging nano-pollutants like ZnO-ENM are increasingly discharged into surface waters and they accumulate in sediments. This is a cause of major concern because of our limited understanding of the mechanisms of ZnO-ENM uptake and toxicity in wetland plants, which play key ecological roles in aquatic ecosystems. Besides, there is an ongoing controversy about the capacity of ENMs to enter plant roots and be transported to the shoots. Our research concludes that ZnO-ENM dissolve before uptake by plant roots and that they are not transported to the shoots. Nanoparticles smaller than 50 nm dissolve faster than the larger particles and are more toxic to plants. Exposure to ZnO-ENM takes place mostly through the uptake of dissolved

  Zn2+ , but also through direct contact with the root surface. The mechanisms of action of ZnO-ENM adsorbed onto the root surface require further investigation. In Phragmites australis, ZnO-ENM reduce shoot and root growth, transpiration, and photosynthetic rate, while inducing changes in nutrient uptake and distribution. This could be detrimental to the environmental health of aquatic ecosystems and their capacity to sequester carbon. Wetlands are one of the major carbon sinks globally. Pollutants that compromise the biomass production of wetland plants can increase global carbon emissions, of vital importance in the current context of climate change. In conclusion, current and future ZnO-ENM levels in sediments could pose a significant risk for aquatic plants and ecosystems. Our research also evidences the importance of considering the undissolved fraction, exposure time, and NP size to correctly evaluate the environmental risk of nanomaterials.

Fig. 1 .

 1 Figures

Fig. 2 .

 2 Fig. 2. Distribution of Al across the different pools. Plants were treated with four different ZnO

Fig. 3 .

 3 Fig. 3. Zinc isotopic composition of ZnO materials, plants, and the BCR-281 (RYE) reference material. Plants were treated with four different ZnO sources at 100 mg l -1 : micron-size (Bulk),

Fig. 4 .

 4 Fig. 4. Photosynthetic performance under ZnO stress. Plants were treated with four different

Fig. 5 .

 5 Fig. 5. ZnO-NPs on the root surface. Roots in (a) were treated with NP < 50 nm (NP50), while

Fig. 7 .

 7 Fig. 7. Dark-field micrograph and elemental map of the same region as Fig.6a. The Zn

Acknowledgements

This project has received funding from the European Union's Horizon 2020 research and innovation programme under the Marie Skłodowska-Curie grant agreement NANOREM No 704957. Franck Poitrasson is funded by the French Centre National de la Recherche Scientifique. We are grateful for the technical assistance of Joan Martorell and Luís López with TEM microanalysis; Manuel Henry and Jérôme Chmeleff with Zn isotope analysis; and Josep Mata, Marta Pintó, and Xavier García with greenhouse tasks and photosynthesis measurements.

We thank Sheela Paramjothy for her help with maintaining the experiment.