Anionic Cerium Oxide Nanoparticles Protect Plant Photosynthesis

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Anionic Cerium Oxide Nanoparticles Protect Plant Photosynthesis from Abiotic Stress by Scavenging Reactive Oxygen Species Honghong Wu, Nicholas Tito, and Juan Pablo Giraldo ACS Nano, Just Accepted Manuscript • DOI: 10.1021/acsnano.7b05723 • Publication Date (Web): 03 Nov 2017 Downloaded from http://pubs.acs.org on November 3, 2017

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Anionic Cerium Oxide Nanoparticles Protect Plant Photosynthesis from Abiotic Stress by Scavenging Reactive Oxygen Species Honghong Wu, Nicholas Tito, Juan P. Giraldo*

Department of Botany and Plant Sciences, University of California, Riverside, 92521 * Corresponding author: Dr. Juan P. Giraldo, Department of Botany and Plant Sciences, University of California, Riverside, 92521, telephone: +1 9518273583, email: [email protected]

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Abstract Plant abiotic stress leads to accumulation of reactive oxygen species (ROS) and a consequent decrease in photosynthetic performance. We demonstrate that a plant nanobionics approach of localizing negatively charged, sub-11 nm, spherical cerium oxide nanoparticles (nanoceria) inside chloroplasts in vivo augments ROS scavenging and photosynthesis of Arabidopsis thaliana plants under excess light (2000 µmol m-2 s-1, 1.5 h), heat (35 °C, 2.5 h), and dark chilling (4 °C, 5 days). Poly (acrylic acid) nanoceria (PNC) with a hydrodynamic diameter (10.3 nm) - lower than the maximum plant cell wall porosity - and negative zeta potential (-16.9 mV) exhibit significantly higher colocalization (46 %) with chloroplasts in leaf mesophyll cells than aminated nanoceria (ANC) (27 %) of similar size (12.6 nm) but positive charge (9.7 mV). Nanoceria are transported into chloroplasts via non-endocytic pathways, influenced by the electrochemical gradient of the plasma membrane potential. PNC with a low Ce3+/Ce4+ ratio (35.0 %) reduce leaf ROS levels by 52 %, including hydrogen peroxide, superoxide anion, and hydroxyl radicals. For the latter ROS there is no known plant enzyme scavenger. Plants embedded with these PNC that were exposed to abiotic stress exhibit an increase up to 19 % in quantum yield of photosystem II, 67 % in carbon assimilation rates, and 61 % in Rubisco carboxylation rates relative to plants without nanoparticles. In contrast, PNC with high Ce3+/Ce4+ ratio (60.8 %) increase overall leaf ROS levels and do not protect photosynthesis from oxidative damage during abiotic stress. This study demonstrates that anionic, spherical, sub-11 nm PNC with low Ce3+/Ce4+ ratio can act as a tool to study the impact of oxidative stress on plant photosynthesis and to protect plants from abiotic stress.

Keywords:

nanoceria,

reactive

oxygen

species,

chloroplast,

oxidative

stress,

photosynthesis, Rubisco, quantum yield.


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Plant nanobionics is an approach that seeks to enable plant organelles, tissues, and whole organisms with augmented functions through the use of nanomaterials.1,2 This interdisciplinary field at the interface of nanotechnology and plant biology has the potential to augment tolerance to abiotic stress of wild type plants by embedding nanoparticles within photosynthetic tissues and organelles. Although significant progress has been made towards understanding plant-nanoparticle interactions, numerous challenges and opportunities remain to use nanotechnology as a tool to study and engineer plant function.3,4,5,6 The impact of nanoparticles having distinct size-dependent optical, electronic, and catalytic properties on plant photosynthesis is poorly understood. A plant nanobionic approach demonstrated enhanced photosynthetic electron transport rates in extracted chloroplasts and leaves as a result of the spontaneous penetration of semiconducting single walled carbon nanotubes within the chloroplast thylakoid membranes.1 Nanoceria (cerium oxide nanoparticles) were also shown to act as catalytic scavengers of reactive oxygen species (ROS) in isolated chloroplasts.1 ROS accumulation can lead to oxidation of proteins, lipids, carbohydrates, and DNA.7,8 Thus nanoceria are well positioned to protect plant photosynthesis from the detrimental effects of ROS accumulation during abiotic stress.

Nanoceria are a family of cerium oxide nanoparticles with sizes ranging from a few to hundreds nanometers, diverse shapes (e.g. sphere, rod, and nano-sheets), and zeta potentials (neutral, negative, and positive)9,10,11,12 that has been widely used as antioxidant in biomedical research.13,14,15 Previous studies have reported that nanoceria with different size, charge, and exposure regime (dose, timing, and media) have distinct effects on plant photosynthesis and growth. Conway et al. (2015) found that positively charged uncoated cubic nanoceria (231 ± 16 nm, 32.8 ± 1.0 mV; 100 mg/L, four weeks, soil) decreased plant photosynthetic CO2 assimilation efficiency in herbaceous annual plants (Clarkia unguiculata).16 Du et al. (2015) reported that the final biomass in winter wheat grown under field conditions was not affected by CeO2 nanoparticles (231 ± 16 nm, zeta potential not reported; up to 400 mg/L, seven months, soil).17 In contrast, Rico et al. (2014) demonstrated that negatively charged rods of nanoceria (231 ± 16 nm, -22.8 ± 4.5 mV; 500 mg/L, three months, soil) significantly improve wheat growth and shoot


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biomass in potted plants grown in greenhouse conditions.18 Structural properties of nanoceria such as size and charge may affect their transport through plant cell walls with a porosity less than ~13 nm,19 and cell lipid bilayers with membrane potentials about 140 mV.20,21 Recently, it was reported that negatively or positively charged but not neutrally charged nanoparticles spontaneously penetrate the lipid envelopes of extracted chloroplasts.22 Spielman-Sun et al. (2017) observed higher root to leaf translocation of negatively charged than positively charged nanoceria.23 Furthermore, Pulido-Reyes et al. (2015) determined that the percentage of surface content of Ce3+ sites is the main driver of toxicity or non-toxicity of nanoceria which may also explain the variety of roles that have been attributed to nanoceria in plants.11 Nanoceria with high Ce3+/Ce4+ ratio (e.g. 58%) show superoxide dismutase mimetic activity, produces hydrogen peroxide, and impairs growth of green algae whereas nanoceria at the same concentration (10 mg/L) and exposure regime (72 h) but with low Ce3+/Ce4+ ratio (e.g. 36%) exhibit catalase mimetic activity, scavenges hydrogen peroxide to water and molecular oxygen, and is not toxic to green algae.11 ROS accumulation is a common response of plants to almost all abiotic stresses.7,24 More than 96% of global rural land area is affected by abiotic stresses that strongly inhibit plant growth and lead to about half of crop yield loss worldwide.25,26 Climate change is predicted to exacerbate extreme abiotic stress events in agricultural land.27,28 Reflecting the urgency to understand and mitigate the impact of abiotic stress on agriculture, there is an increasing research effort in this area.25 Abiotic stresses are well known to cause ROS accumulation and oxidative damage to the plant photosynthetic machinery.29,30 Plant abiotic stress limits CO2 fixation and inhibit electron transport in chloroplasts, inducing • –

ROS formation of hydrogen peroxide (H2O2), superoxide anion (O ), singlet oxygen (1O2), and hydroxyl radicals (OH•).31 The latter is the most destructive and toxic ROS and cannot be scavenged by any known enzymes in biological systems.32

Chloroplast photosynthetic performance is highly sensitive to abiotic stresses including excess light, heat, and chilling that lead to accumulation of ROS,33 impaired chlorophyll biosynthesis,34 reduced chloroplast electron transport,35 perturbation of thylakoid


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membrane fluidity,36 and reduced Ribulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco) activity.37 Nanoceria are well positioned to protect plants against oxidative damage caused by abiotic stresses. Unlike other antioxidants, nanoceria can catalytically reduce oxidative stress in plants by regenerating the sites in the cerium oxide lattice that scavenge ROS.1,38 Nanoceria forms oxygen vacancies resulting in dynamic defect sites with dangling Ce3+ bonds which can effectively scavenge oxygen radicals.39,40 Nanoceria can catalyze the quenching of hydroxyl radicals, superoxide anion, and hydrogen peroxide to hydroxyl groups, oxygen, and water.12,41,42,43,44 Nanoceria catalytic scavenging of ROS can protect the light and carbon reactions of photosynthesis in plants under abiotic stress by minimizing oxidative damage to chloroplast photosystems, pigments, lipid membranes, and enzymes involved in carbon fixation. Thus interfacing nanoceria with chloroplasts augments plant ROS scavenging by enabling the quenching of superoxide anion and hydrogen peroxide, and introduces a new function by scavenging the highly damaging hydroxyl radicals. For the latter there is no known scavenging enzymatic pathway.32

We investigated the mechanisms of anionic and cationic nanoceria transport to chloroplasts in vivo to create Arabidopsis thaliana plants with augmented ROS scavenging and higher photosynthetic performance. We studied if nanoceria uptake by leaf mesophyll cells and delivery to chloroplasts are facilitated by coating the nanoparticles with negatively or positively charged polymers. We determined if the transport of nanoceria through leaf mesophyll cell membranes into chloroplasts is endocytosis dependent and affected by the plasma membrane potential. We hypothesized that localizing nanoceria with low Ce3+/Ce4+ ratio within chloroplasts of leaf mesophyll cells significantly reduces ROS accumulation in plants under abiotic stress, enabling higher quantum yield of photosystem II and carbon assimilation rates.

Results and discussion Mechanisms of nanoceria transport to chloroplasts in vivo Transmission electron microscopy images indicated that the cerium oxide core of nanoceria is spherical with an average diameter of 4.6 ± 1.6 nm (Figure 1a). Dynamic


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light scattering measurements (Nano S, Malvern) showed negatively charged PNC1, PNC2, and positively charged aminated poly (acrylic acid) nanoceria (ANC) monodisperse solutions of similar hydrodynamic diameter, 10.3 ± 1.3, 9.1 ± 1.1 and 12.6 ± 2.6 nm, respectively (P > 0.05) (Figure 1b). Zeta potential characterization (Nano ZS 90, Malvern) confirmed the presence of negative charge for PNC1, -16.9 ± 6.1 mV, PNC2, -14.6 ± 7.1 mV, and positive charge for ANC, 9.7 ± 1.2 mV (Figure 1c). PNC1, PNC2, and ANC have peaks of absorbance corresponding to nanoceria at 271, 265, and 260 nm, respectively (Figure S1a). The polymer surface coating of PNC1, PNC2, and ANC was characterized by Fourier transform infrared spectroscopy (FTIR) analysis indicating the presence of C=O-OH bonds in PNC and NH-C=O bonds in ANC (Figure S1b).

The delivery of nanoceria particles to leaf mesophyll chloroplasts was performed by a simple method of infiltration through the stomata pores into the leaf lamina (Figure 2a). A nanoceria concentration of 450 µM (~50 mg/L) was chosen for leaf infiltration. This is an optimal concentration that did not have a significant impact on leaf chlorophyll content and lifespan under normal growth conditions (Figure S2). For confocal imaging of nanoparticles in leaf mesophyll cells, nanoceria was labeled with the fluorescent dye 1,1'-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine

perchlorate

(DiI)

and

the

autofluorescence of chloroplast photosynthetic pigments was detected. We performed confocal z-stacks in leaf mesophyll cells with a 2 µm optical resolution (Video S1), smaller than the diameter of Arabidopsis chloroplasts (~5-10 µm).45 After 3 hours of leaf infiltration with nanoceria, the PNC1 and PNC2 showed a mean colocalization rate with chloroplasts of 46.0 ± 5.3 % and 41.2 ± 0.9 %, respectively, whereas ANC colocalization with these organelles was significantly lower (27.3 ± 2.9 %) (Figure 1d-f). No fluorescence signal was detected in leaves infiltrated with TES buffer solution (10 mM TES, 10 mM MgCl2, pH 7.5) and non-labeled nanoceria (Figure S3a). Nanoceria appear to move rapidly from leaf cell extracellular spaces, through mesophyll cell walls and plasma membranes into chloroplasts in vivo (Figure 2a). Nanoceria were found in chloroplasts only two minutes after leaf infiltration with the nanoparticles. The percentage of PNC1, PNC2, and ANC colocalization with chloroplasts in leaf mesophyll


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cells increased linearly from 5 min to 60 min (Figure 1f, Figure S3b). Nanoceria uptake into chloroplasts reaches a plateau about 180 min after leaf infiltration with the nanoparticles (Figure 1g, Figure S3b).

Before reaching the chloroplasts in vivo, nanoceria necessarily have to cross a leaf mesophyll cell membrane. We observed almost two times higher colocalization of negatively charged PNC than positively charged ANC within chloroplasts in leaf mesophyll cells. We propose that these differences in colocalization can be in part explained by the interaction between the nanoparticle’s zeta potential and the plasma membrane potential. The outside of the plasma membrane has a net positive charge that preferentially binds to anionic nanoparticles. We performed a depolarization of the plasma membrane potential to investigate its role in the transport of nanoceria across the leaf mesophyll cell membrane. The plant plasma membrane potential is primarily built by H+ electrochemical gradient20,46 and can be depolarized by applying NaCl.47,48 Plasma membrane potential is approximately -140 mV in plants at non-stress conditions.20,21 After applying 50-100 mM NaCl the recovered steady state plasma membrane potential is two to three folds lower than the non-stressed one.47,48 In this study, we observed a significant increase of 27 % (P < 0.05) in colocalization of chloroplasts with PNC when the leaf mesophyll cell membranes were depolarized with 100 mM NaCl (Figure 2b,c). In contrast, ANC colocalization with chloroplasts significantly decreased (P < 0.05) from 27.3 % ± 2.9 % (ANC) to 18.0 % ± 2.9 % (ANC + NaCl) (Figure 2b,c). To eliminate possible confounding effects of osmotic change by applying 100 mM NaCl, a non-ionic isotonic solution (170 mM mannitol) was used for plant infiltration together with PNC and ANC. Similar chloroplast colocalization levels (P > 0.05) were found between plants infiltrated with PNC and PNC + mannitol (Figure 2b,c; Figure S4) or ANC and ANC + mannitol (Figure 2b,c, Figure S4).

Nanoceria transport through leaf mesophyll cell membranes appears to be endocytosis independent. Nanoceria colocalization with chloroplasts was not significantly different in plants infiltrated with PNC or ANC at temperatures ranging from 24 ºC down to 14 ºC and 4 ºC (P > 0.05) (Figure 2d, Figure S5a). The lack of temperature effect on nanoceria


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colocalization with chloroplasts indicates that the transport of these nanoparticles occurs via non-endocytic pathways. We also suppressed endocytosis in leaf mesophyll cells with a well-known inhibitor auxin.49,50 No significant changes in nanoceria chloroplast uptake were found between plants infiltrated with nanoceria and nanoceria plus auxin (Figure 2e, Figure S5b). Together, these results demonstrate that the transport of nanoceria into chloroplasts is apparently independent of endocytosis in leaf mesophyll cells.

Our results indicate that Arabidopsis leaf mesophyll cells, like mammalian cells, preferentially uptake anionic nanoceria51 with a higher affinity to cell membranes52 than cationic nanoceria. NaCl induced depolarization of the plasma membrane potential reduces the electrical gradient opposing the transport of negatively charged nanoparticles into the leaf mesophyll cells thus favoring the cellular uptake of anionic PNC. Transport of nanoceria occurs through non-endocytic pathways in vivo whereas uptake of nanoceria by mammalian cells is governed by energy dependent endocytosis.53 Other nanomaterials such as single walled carbon nanotubes and nanosheets are transported into cultured plant cells by endocytosis or internalized in plant root cells via non-endocytic pathways.54,55 Channels or transporters in the plasma membrane could act as alternative pathways for entry of nanoparticles into plant cells. Silver nanoparticles can activate mechanosensitive channels in Arabidopsis protoplasts.56 The size of opened mechanosensitive channel pores ranges between 3-4 nm after undergoing one of the largest conformational changes known in membrane proteins.57,58 However, these channels are not large enough to transport nanoceria having average hydrodynamic diameter of 10.3 ± 1.3 nm. Likewise, chloroplast porins of 2.5-3 nm are unlikely to enable nanoceria entry into these organelles.59 Temporary pores can be formed in the plasma membrane after the application of nanoparticles including quantum dots60 and silica nanospheres.61 Single walled carbon nanotubes have being shown to passively penetrate chloroplast membranes by disrupting lipid bilayers.1,22 Like carbon nanotubes, nanoceria also localize inside the envelopes of chloroplasts for which no endocytosis mechanisms have been reported. The formation of temporary pores by nanoparticle disruption of lipid membranes may provide a passive transport mechanism of cell and organelle nanoceria uptake that is independent of plant metabolic activity.


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Nanoceria augmentation of plant ROS scavenging ROS are mainly produced by plant cells in chloroplasts, mitochondria, peroxisomes, and the apoplast,33 with illuminated chloroplasts being a major source of ROS production.62,63 Superoxide anion and hydroxyl radicals have short lifetimes64 constraining their diffusion within chloroplasts. Thus, delivering nanoceria inside chloroplasts is a promising way to provide augmented ROS scavenging and enhanced photosynthesis to plants (Figure 3a). The PNC1 and PNC2 show almost two times higher colocalization levels with chloroplasts than ANC (P < 0.001, Figure 1f). The PNC1 Ce3+/Ce4+ ratio of 35.0 % ± 4.4 % (Figure 3b, Figure S6) is within the range of nanoceria scavenging capacity for both superoxide anion and hydrogen peroxide.11 In contrast, the high Ce3+/Ce4+ ratio (60.8 ± 7.6 %) of PNC2 indicates ability for superoxide anion scavenging with hydrogen peroxide generation (Figure 3c, Figure S6).11 ANC have a Ce3+/Ce4+ ratio of 41.3 % ± 3.2 %, slightly higher than that of PNC1 (Figure 3d). These nanoceria can catalytically • –

scavenge chloroplast generated hydroxyl radicals (OH•), superoxide anion (O ), and hydrogen peroxide (H2O2) via the following reactions:1,42,43,65,66 Ce3+ ↔ Ce4+ + e– Ce3+ + OH• → Ce4+ + OH‒ • –

Ce4+ + O → Ce3+ + O2 H2O2 + 2Ce4+ + 2OH‒ → 2H2O + O2 + 2Ce3+ We monitored ROS generation in leaf mesophyll cells infiltrated with PNC1, PNC2 and ANC. We used DHE (dihydroethidium) dye for confocal imaging of superoxide anion and H2-DCFDA (2',7'-dichlorodihydrofluorescein diacetate) for hydrogen peroxide (H2O2). The DHE can freely permeate cell membranes and form a red fluorescent product 2-hydroxyethidium upon reaction with superoxide anions.67 The H2-DCFDA dye is converted to its fluorescent form DCF (2',7'-dichlorofluorescein) after interaction with ROS such as H2O2. We induced ROS generation via exposing leaf discs to UV-A light (405 nm) during confocal experiments. In vivo monitoring of DHE fluorescence showed that both PNC1 and PNC2 infiltrated plants have significantly less superoxide anion in leaf mesophyll cells than buffer infiltrated leaves (controls) after 1 min of UV-A stress (P


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< 0.05) (Figure 3e,f). However, changes in DCF fluorescence intensity in leaf mesophyll cells indicated that PNC1 have a strong scavenging effect of ROS such as H2O2 whereas PNC2 lead to a significant higher accumulation of H2O2 in infiltrated leaves under UV exposure (Figure 3e,f). In agreement with previous reports,11,41,68 superoxide dismutase (SOD) and catalase (CAT) mimetic activity assays in vitro indicated that PNC1 (low Ce3+/Ce4+) scavenge both superoxide anion and H2O2 whereas PNC2 (high Ce3+/Ce4+) scavenge mainly superoxide anion but not H2O2 (Figure S6). Both DCF and DHE fluorescence intensity in ANC infiltrated plants were lower but not significantly different than controls (P > 0.05) (Figure 3e,f). The low ANC ROS scavenging efficiency in leaf mesophyll cells can be attributed to the significantly lower colocalization rate of ANC (27.3 ± 2.9 %) with the sites of ROS generation inside chloroplasts relative to that of PNC1 (46.0 ± 5.3 %) (Figure 1f), and lower catalase (CAT) mimetic activity of ANC compared with PNC1 (P < 0.05) (Figure S6b). Overall, our results demonstrate that anionic PNC1 with low Ce3+/Ce4+ are potent in vivo non-enzymatic ROS scavengers in plants.

Nanoceria enhancement of photosynthesis under excess light, heat, and dark chilling Accumulation of ROS under excess light leads to damage of susceptible components in the chloroplast photosynthetic machinery such as the D1 protein, oxygen evolving complex in PSII, thylakoid membrane lipids, and chloroplast DNA.8,32,69 Herein, we assessed the impact of excess light on photosynthetic parameters in leaves infiltrated with PNC1 (PNC1-Leaves), PNC2 (PNC2-Leaves) and ANC (ANC-Leaves) using a GFS3000 gas exchange analyzer and fluorometer (Walz). The PNC1-Leaves exposed to excess light for 1.5 h (2000 µmol m-2 s-1 of photosynthetic active radiation (PAR), similar to full sunlight levels70,71) showed up to 19 % higher (P < 0.05) quantum yield of PSII (QY) than controls without nanoparticles (NNP-Leaves). In contrast, the QY of PNC2Leaves is not significantly different than NNP-Leaves except at the maximum light intensity of 2000 µmol m-2 s-1 (P > 0.05) (Figure 4a). Thus, PNC1 with low Ce3+/Ce4+ ratio enhance the proportion of photons used in photochemistry under excess light whereas PNC2 with high Ce3+/Ce4+ ratio had no significant impact at most light levels (P > 0.05). Similarly, PNC1-Leaves exhibited a 10 % higher maximum yield of PSII


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(Fv/Fm) than controls infiltrated with buffer (P < 0.05) compared to no significant change in PNC2-Leaves (P > 0.05) (Figure 4b). The higher QY and Fv/Fm values in leaves with PNC1 relative to controls reflect improved quantum efficiency of PSII, an indicator of plant photosynthetic performance. However, the Fv/Fm values for PNC1Leaves (0.65 ± 0.03), PNC2-Leaves (0.60 ± 0.05) and NNP-Leaves (0.59 ± 0.04) were lower than optimal values around 0.83 for most plant species72 indicating a degree of light induced stress across all treatments (Figure 4b). ANC-Leaves do not have significantly different QY and Fv/Fm than NNP-Leaves (P > 0.05) (Figure 4a,b).

Photosynthetic CO2 assimilation (A) light curves measured after leaves were exposed to excess light for 1.5 h showed A values up to 40 % higher in PNC1-Leaves (at 50 µmol m2

s-1 PAR, P < 0.05) and up to 38 % lower in PNC2-Leaves relative to controls (Figure

4c). In PNC1-Leaves, increased A was observed at a broad range of PAR both within the photosynthesis light limited region and beyond the light saturation point of 1200 µmol m2

s-1 PAR (Figure 4c), whereas A in PNC2-Leaves decreased. Differences in A across

treatments between PNC1-Leaves and NNP-Leaves were not associated to changes in stomatal conductance (Gs, Figure S7a). In contrast, a significant reduction in Gs was found in PNC2-Leaves whereas ANC-Leaves have higher Gs compared with NNPLeaves (P < 0.05) (Figure S7a). PNC1-Leaves exhibited significantly higher quantum efficiency of CO2 uptake (φCO2) (0.0247 ± 0.0029) relative to NNP-Leaves (0.0215 ± 0.0032) (P < 0.05), whereas PNC2-Leaves showed lower φCO2 than NNP-Leaves (Figure 4d). Under excess light, the response of A to internal intercellular mole fraction of CO2 (Ci) (A-Ci curve) showed a significantly higher A (up to 19% increase) in PNC1-Leaves than NNP-Leaves from 160 to 745 ppm ci. In comparison, A in PNC2-Leaves decreased (Figure 4e), indicating that PNC1 but not PNC2 enhance in vivo Rubisco carboxylation activity (Vcmax). PNC1-Leaves exhibited significantly higher Vcmax (137.7 ± 28.0 µmol CO2 m-2 s-1) relative to NNP-Leaves (111.2 ± 28.1 µmol CO2 m-2 s-1). In contrast, the Vcmax (69.7 ± 10.1 µmol CO2 m-2 s-1) in PNC2-Leaves is significantly lower than the NNP-Leaves (P < 0.05). Maximum carbon assimilation rates for PNC1-Leaves were also higher in the region limited by Ribulose-1,5-bisphosphate (RuBP) regeneration, above ~300 ppm ci. No significant differences in these photosynthetic parameters were found


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between ANC-Leaves and NNP-Leaves (Figure. 4c-f). These results indicate that PNC1 scavenging of both superoxide anion and hydrogen peroxide at the sites of ROS generation in the chloroplasts protects key enzymes and intermediates of the carbon reactions of photosynthesis from ROS damage such as Rubisco enabling more efficient carboxylation under excess light. Despite superoxide anion scavenging by PNC2, the associated increase in hydrogen peroxide levels has a detrimental effect on both carbon assimilation rates and gas exchange. The significantly lower chloroplast uptake of ANC compared to PNC1 (Figure 1f) limits ANC ability to protect most chloroplasts in leaf mesophyll cells from the degradation of the photosynthetic machinery caused by accumulation of ROS under excess light. Plants often encounter light intensities that exceed their photosynthetic capacity73 and induce damaging ROS accumulation.74 In Arabidopsis, diurnal excess light above 1300 µmol m-2 s-1 leads to a significant drop of Fv/Fm.75 Continuous light has also a severe negative impact on plant health that is associated with ROS generation.76 The negative effects of continuous light include plant chlorosis, necrosis, and reductions in photosynthetic capacity, Rubisco carboxylation, and quantum yield of PSII.76,77 We report that PNC1 infiltrated Arabidopsis plants were better at tolerating continuous excess light (1300 µmol m-2 s-1 PAR) than controls without nanoparticles (Figure 4g,h). After only 1 day of exposure to continuous excess light, plants infiltrated with PNC1 exhibited higher chlorophyll content index (CCI) than controls (P < 0.05) (Figure 4g,h). PNC1 scavenging of hydrogen peroxide and superoxide anion inside chloroplasts reduces oxidative damage to chlorophyll under excess continuous light. In contrast, plants infiltrated with PNC2 had lower CCI than controls after 3 days of continuous excess light (P < 0.05). The accumulation of hydrogen peroxide generated by PNC2 superoxide anion scavenging activity (Figure 3, Figure S6A) leads to a reduction in chlorophyll content. No significant differences in CCI were found between plants infiltrated with ANC and controls without nanoparticles as expected from the low levels of colocalization of ANC with chloroplast pigments in leaf mesophyll cells.


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To investigate if protection from oxidative damage by nanoceria extends to different types of abiotic stresses besides excess light, we conducted experiments on temperature stresses such as heat and dark chilling. Heat and dark chilling have been demonstrated to cause a decrease of PSII abundance, chlorophyll biosynthesis, electron flow, and Rubisco activity.35,78 Our results indicate that heat and dark chilling exposed PNC1-Leaves exhibit higher carbon assimilation rates and quantum yield of CO2 uptake than NNP-Leaves but not PNC2-Leaves or ANC-Leaves (Figure 5a-b,e-f). Maximum enhancement of carbon assimilation rates in A-light curves is 67% and 46% in PNC1-Leaves under heat and dark chilling stress, respectively. Similarly, PNC1-Leaves show significantly higher carbon assimilation rates per given internal carbon dioxide concentration and maximum Rubisco carboxylation rates than NNP-leaves under heat (Figure 5c,d) and dark chilling stress (Figure 5g,h). PNC1-leaves have 61% and 49% higher A in heat and dark chilling, respectively, than NNP-Leaves. However, heat and dark chilling exposed PNC1-Leaves, PNC2-Leaves, or ANC-Leaves have an either marginal or non-significant improvement in Fv/Fm and quantum yield of PSII (Figure S8). Similar to leaves exposed to excess light, PNC2 generation of hydrogen peroxide reduces photosynthetic performance under temperature stress. ANC exhibit low colocalization with chloroplasts and as a result do not have a significant impact on the light and carbon reactions of photosynthesis of leaves exposed to heat or dark chilling stress. Overall, our results indicate that PNC1 with low Ce3+/Ce4+ ratio protect the enzymatic carbon reactions of photosynthesis in plants under temperature stress but not the photon absorption efficiency of PSII (Figure 5, Figure S8).

Plant ROS are mainly produced by chloroplasts, mitochondria, peroxisomes, NADPH oxidases, and class III peroxidases.7,33 Among these, chloroplasts are a source of hydroxyl radical production in leaves under stress conditions.7,33,79 The cerium oxide lattice in PNC with large surface to volume ratios catalytically scavenges ROS produced by the chloroplasts such as superoxide anion, hydrogen peroxide, and hydroxyl radicals, the most destructive ROS in plant cells.12,39,41,79 Unlike superoxide anion and hydrogen peroxide, no specific scavenging enzyme for hydroxyl radicals has been found in plants.32 Light exceeding the chloroplast’s photosynthetic capacity results in levels of ROS that cannot be controlled by the natural scavenging mechanisms of plants.74,76 Heat and


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chilling cause oxidative damage to chloroplast components inhibiting the repair of PSII, the most temperature-sensitive component of the photosynthetic apparatus.29,80–82 PNC1 protect both the light and carbon reactions of photosynthesis from ROS damage in plants under excess light while only improving Rubisco carboxylation rates and quantum efficiency of CO2 uptake in plants exposed to heat and dark chilling. Temperature stress leads to both enhanced chloroplast ROS generation31 and membrane destabilization.26,83 Although, PNC1 protect chloroplasts from oxidative damage by scavenging ROS, these nanoparticles are not able to prevent the dysfunction of light energy absorption and conversion to electron flow that arises from chloroplast thylakoid membrane destabilization.

Engineering the plant antioxidant defense system may be an effective mean to improve tolerance to excess light, heat and dark chilling stresses.31,33,84,85 Arabidopsis as a plant model system provides molecular tools that complemented with low Ce3+/Ce4+ ratio PNC1 will increase our understanding of the role of intracellular ROS communication in regulating fine-tuned abiotic stress responses.33,86 Although genetic engineering of Arabidopsis and tomato plants has improved tolerance to abiotic stress,75,87 plant nanobionics offers an alternative approach to enhance abiotic stress tolerance in nonmodel systems and wild-type plants. The method of plant ROS manipulation via nanoceria infiltration through the stomata pores in the leaf lamina is well suited for research on diverse broad-leaf plant species both in the laboratory and the field. Developing foliar sprayed formulations for large-scale PNC1 delivery could increase crop photosynthetic performance under a variety of abiotic stresses responsible for worldwide decline in crop yields.

Conclusions Utilizing a plant nanobionic approach, we demonstrate that anionic spherical sub-11 nm nanoceria with low Ce3+/Ce4+ ratio (PNC1) are potent ROS scavengers in leaf mesophyll cells protecting the chloroplast photosynthetic machinery from abiotic stresses. Nanoceria transport from leaf extracellular spaces to chloroplasts occurs via non-endocytic pathways influenced by the leaf mesophyll plasma membrane potential. Negatively


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charged PNC1 exhibit almost two-fold higher colocalization rate with chloroplasts than positively charged ANC. The PNC1 with low Ce3+/Ce4+ ratio localized in chloroplasts boost plant light energy absorption efficiency under excess light by shielding vulnerable chloroplast photosystems and chlorophyll pigments from oxidative damage. The PNC1 also enhance the carbon reactions of photosynthesis by enabling higher Rubisco carboxylation rates and photosynthetic capacity under both light and temperature stresses. As a result, Arabidopsis plants augmented with low Ce3+/Ce4+ ratio PNC1 have improved photosynthetic performance under excess light, continuous excess light, heat and dark chilling.

Experimental methods Plant material Four weeks old Arabidopsis thaliana (Columbia 0) plants were used in this study. Seeds were sown in 2 x 2 inch disposable pots filled with standard soil mix (Sunshine, LC1 mix). About 32 of these disposable pots were placed in a plastic tray. Only one individual was kept in each pot after one week of germination. Plants were grown in Adaptis 1000 growth chambers (Conviron) at 200 µmol m-2 s-1 photosynthetic active radiation (PAR), 24 ± 1 ºC, 60% humidity, and 14/10 h day/night regime. Plants were hand-watered by pouring tap water directly on the plastic tray containing the disposable pots once every two days.

Synthesis and characterization of PNC and ANC The poly (acrylic acid) nanoceria (PNC1) were synthesized using the methodology described previously with some modifications.9 Briefly, 1.0 M cerium (III) nitrate (2.17 g, Sigma Aldrich, 99%) in molecular biology grade water (5.0 mL, Corning, Mediatech, Inc.) was mixed with an aqueous solution (10 mL) of 0.5 M poly (acrylic acid) (1,800 MW, 9 g, Sigma Aldrich). Then the solution was mixed thoroughly at 2000 rpm for 15 min using a vortex mixer (model no 945415, Fisher). The resulting mixture was then added dropwise to an ammonium hydroxide solution (30.0 mL, 30%, Sigma Aldrich) under continuous stirring at 500 rpm (RCT basic, IKA) at ambient temperature. After 24 hr, the solution was transferred to a 50 mL falcon tube and centrifuged at 4,000 rpm for 1


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hr to remove any debris and large agglomerates. Then, 45 mL of supernatant solution was diluted in a total 90 mL with molecular biology grade water and purified from free polymers and other reagents by centrifugation at 3,500 rpm (Allegra X30, Beckman) in at least five cycles (10 min each cycle) using a 30K Amicon cell (MWCO 30K, Millipore Inc.). The suspension was reduced in each cycle to about 10% of the initial volume. The absorbance of eluent in each cycle was measured with an UV-VIS spectrophotometer (UV-2600, Shimadzu) to ensure no free polymers and other reagents in the final PNC1 solution. Pulido-Reyes et al. (2015) showed that negligible amounts of dissolved Ce (ranging from 0.00001 to 0.0008 mg/L) in different types of nanoceria (10 mg/L).11 No dissolved Ce(NO3)3 peaks were found in absorption spectra for purified PNC1 solution (Figure S1c). After purification, the nanoparticle suspension was filtered with a 20 nm pore size syringe filter (Whatman, AnotopTM 25). The absorbance of final PNC1 solution was then measured with the UV-VIS spectrophotometer (UV-2600, Shimadzu) and its concentration was calculated by using Beer-Lambert’s law with absorbance at 271 nm, absorption molar coefficient of 3 cm–1 mM–1,88 and pathway length of 1 cm. The final PNC1 solution was stored in a refrigerator (4 ºC) until further use. The nanoceria size and zeta potential is maintained unchanged for 2 weeks (Figure S1d,e). Another type of nanoceria PNC2 was synthesized following previous methods89 with some modifications. Briefly, poly (acrylic acid) (1,800 MW, Sigma Aldrich) were dissolved in 2.5 mL molecular biology grade water (Corning, Mediatech, Inc.) under stirring at 500 rpm (RCT basic, IKA) for 10 min at ambient temperature. A solution of 0.54 g cerium (III) nitrate (Sigma Aldrich, 99%) in 1.25 mL molecular biology grade water was then added dropwise into the poly (acrylic acid) solution followed by 1000 rpm stirring for 2 hours (RCT basic, IKA). The resulting mixture was added dropwise to a hydrogen peroxide solution (7.5 mL, 30% non-stabilized, Arcos) under continuous stirring at 500 rpm (RCT basic, IKA) at ambient temperature. After 24 hr, the solution was transferred to a 15 mL falcon tube and centrifuged at 4000 rpm for 1 hr to remove any debris and large agglomerates. Approximately 11 mL of supernatant solution was diluted in a total of 30 mL using molecular biology grade water. The solution was then purified from free polymers and other reagents by centrifugation at 4000 rpm (Allegra X30, Beckman) in at least seven cycles (10 min each cycle) using a 10K Amicon cell


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(MWCO 10K, Millipore Inc.). The suspension was reduced in each cycle to about 10% of the initial volume. As described above, the eluents and final PNC2 solution were characterized with UV-VIS spectrophotometer (UV-2600, Shimadzu) to ensure no free polymers and other reagents remained in the PNC2 solution. The nanoparticle suspension was filtered with a 20 nm pore size syringe filter (Whatman, AnotopTM 25) after purification. The absorbance at 265 nm of final PNC2 solution was used to calculate its concentration following Beer-Lambert’s law with an absorption molar coefficient of 3 cm–1 mM–1,88 and pathway length of 1 cm. Synthesis of amino nanoceria (ANC) was also based on the methods by Asati et al. (2010) with modifications.9 Briefly, 3.5 mL of 5 mM (58 mg/L) PNC1 was mixed with 1.5 mL molecular biology grade water at 500 rpm for 2 min at ambient temperature. Then, 80 mM 1-ethyl-3-(3- dimethylaminopropyl) carbodiimide (EDC, Sigma Aldrich) solution (76.7 mg) in 0.5 mL MES buffer (100 mM, pH 6.0) was added dropwise into the mixture during continuous stirring at 500 rpm for 4 min. Then 80 mM Nhydroxysuccinamide (NHS, Sigma Aldrich) solution (46.0 mg) in 0.5 mL MES buffer (100 mM, pH 6.0) was added dropwise into the mixture under continuous stirring at 500 rpm. After 5 min incubation, 400 mM (0.14 mL) Ethylenediamine (EDA, 99%, Sigma Aldrich) in 0.5 mL molecular biology grade water (pH 6.8 adjusted with HCl) was added dropwise to the final reaction mixture under continuous stirring at 500 rpm for an additional 3 hr at ambient temperature. The resulting solution was transferred to a 15 mL falcon tube and centrifuged at 4,500 rpm for 15 min to remove any debris and large agglomerates. The supernatant solution was purified from excess EDA and other reagents by centrifugation at 4,500 rpm (Allegra X30, Beckman) in five cycles (15 min each cycle) using a 10K Amicon cell (MWCO 10K, Millipore Inc.). The resulting ANC solution was filtered by first passing it through a 100 nm pore size filter (Whatman, AnotopTM 25). Then the collected solution was filtered with a 20 nm pore size filter (Whatman, AnotopTM 25). The absorbance at 260 nm of final ANC solution was measured by spectrophotometry (UV-2600, Shimadzu) and its concentration was calculated following Beer-Lambert’s law with an absorption molar coefficient of 3 cm–1 mM–1,88 and pathway length of 1 cm. The final ANC solution was stored in refrigerator (4 ºC) until further use.


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Transmission electron microscope images of nanoceria were collected using a Tecnai12 TEM. TEM samples were mounted on pure C grids, 200 mesh Cu (01840, Ted Pella Inc). The PNC and ANC zeta potential and size were measured by a Malvern Zetasizer (Nano ZS) and Sizer (Nano S), respectively. Chemical characterization by Fourier transformed infrared spectroscopy (FTIR) was performed with Nicolet 6700 FTIR (Thermo Electron Corp.). X-ray photoelectron spectroscopy (XPS) characterization was carried out using a Kratos AXIS ULTRADLD XPS system equipped with an Al Kα monochromated X-ray source and a 165-mm mean radius electron energy hemispherical analyzer. Vacuum pressure was kept below 3 × 10–9 torr during the acquisition. Dry samples of PNC were mounted on a carbon tape for XPS analysis. XPS spectra were deconvoluted and analyzed with CasaXPS software (CasaXPS version 2.3.18, Casa Software Ltd). The peaks in XPS spectra were identified as +3 or +4 states of cerium according to the NIST XPS database and previous reports by e.g. Korsvik et al. (2007).90

The concentration of nanoceria is calculated using Beer-Lambert’s law. A = ɛCL Where A is the absorbance of peak value for a given sample. ɛ is the molar absorption coefficient of nanoceria (cm-1 M-1), which is not modified by polymer coating.91 L is the optical path length, which is the length of the cuvette of 1 cm in this study. C is the molar concentration of measured nanoparticles. Molar concentration of nanoceria was converted from mM to mg/L for comparison with previous studies using the molecular weight of cerium(III) oxide and cerium(IV) oxide weighted by the Ce3+/Ce4+ ratios for PNC1 (35.0 %) and PNC2 (60.8 %).

SOD and CAT mimetic assays The SOD mimetic assay of nanoceria was performed as reported previously.90,92 Briefly, ferricytochrome C (350 µM) in reaction buffer (10 mM TES, pH 7.5) was added into each well. Hypoxanthine (25 µM), CAT (2000 U), nanoceria (60 nM) and xanthine oxidase were added into each well of a total reaction volume of 100 µL. In the presence of superoxide anion, ferricytochrome C is reduced to ferrocytochrome C. Absorbance of ferrocytochrome C at 550 nm was recorded using a plate reader (Victor 2, Perkin Elmer


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Wallac).

The CAT mimetic assay of nanoceria was determined by previously reported methods.41,68 Briefly, 60 nM nanoceria followed by 2 µM H2O2 was added to reaction buffer (10 mM TES, pH 7.5) in each well (Costar white 96 well microplate with flat bottom, Corning). Amplex Red (10-acetyl-3,7-dihydroxyphenoxazine, 100 µM) and HRP (horseradish peroxidase, 0.2 U/ml) were then added into the mixture in each well. In the presence of HRP, Amplex Red reacts with hydrogen peroxide and is converted into resorufin. The absorbance of resorufin, which is indicative of hydrogen peroxide levels, was monitored at 560 nm in a plate reader (Victor 2, Perkin Elmer Wallac). Measurements were conducted at 25 °C in a final reaction volume of 50 µL.

PNC and ANC labeling with DiI fluorescent dye The

PNC

and

ANC

were

labelled

with

1,1'-dioctadecyl-3,3,3',3'-

tetramethylindocarbocyanine perchlorate (DiI) fluorescent dye following methods previously published.9 Briefly, 0.4 mL of 5 mM (58 mg/L) PNC1 and PNC2 (62 mg/L) or equal molarity of ANC solution was mixed with molecular biology grade water into a final 4 mL volume. Then, 200 µL DiI dye solution (24 µL of DiI, 2.5 mg/mL, in 176 µL of DMSO, dimethyl sulfoxide) was added dropwise under continuous stirring (1, 000 rpm) at ambient temperature. The incubation time for PNC1 and PNC2 with DiI was 1 min and for ANC 60 min. The resulting mixture was purified from DMSO and any free DiI by centrifugation at 3,000 rpm (Allegra X30, Beckman) in five cycles (5 min each) using a 30K Amicon cell (MWCO 30K, Millipore Inc.). The absorbance of final DiIPNC1 and DiI-PNC2 and DiI-ANC solution was measured by spectrophotometry (UV2600, Shimadzu) and its concentration was calculated as explained above. The final DiIPNC1, DiI-PNC2 and DiI-ANC solutions were filtered with 20 nm pore size filter (Whatman, AnotopTM 25) and stored in a refrigerator at 4 ºC. The DiI dye does not coat the nanoparticle surface but instead is encapsulated inside the hydrophobic polymer shell of nanoceria.9 No significant changes in hydrodynamic diameter and zeta potential were found between DiI labeled nanoceria and non-labeled nanoceria (Figure S1f,g).


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Nanoceria leaf infiltration protocol A nanoparticle solution of nanoceria (90 µL of 5 mM PNC1 (58 mg/L), PNC2 (62 mg/L), ANC (58 mg/L)) or DiI labeled nanoceria (900µL of 0.45 mM DiI-PNC1 (51 mg/L), DiI-PNC2 (56 mg/L), DiI-ANC (51 mg/L)) was added to TES (Sigma-Aldrich) infiltration buffer (10 mM TES, 10 mM MgCl2, pH 7.5), and vortexed to make a final 1 mL solution. A solution of 10 mM TES infiltration buffer was used as control. The infiltration solution was then transferred to a 1 mL sterile needleless syringe (NORMJECT®) (tapped to remove air bubbles). Leaves were slowly infiltrated with approximately 200 µL of solution by gently pressing the tip of the syringe against the bottom of the leaf lamina. The excess solution that remained on the surface of leaf lamina was gently wiped out using Kimwipes (Kimtech Science®). The infiltrated leaves were then labeled by wrapping a small thread around the petiole. The infiltrated plants were kept on the bench or refrigerator (4 ºC and 14 ºC treatments) for leaf adaptation and incubation with nanoceria for 3 hr.

Nanoceria imaging in leaf tissues by confocal microscopy Arabidopsis leaves of similar size and chlorophyll content (CCI meter, Apogee) were chosen for infiltration with either TES infiltration buffer, DiI-PNC1, DiI-PNC2, or DiIANC. After 3 hr, leaf discs were taken with a cork borer and mounted on microscopy slides as follows. A well for mounting the leaf discs on the slide (Corning 2948-75X25) was made by rolling a pea-size amount of observation gel (Carolina) to about 1 mm thin. A circular section of gel roughly twice the size of the leaf discs was cut in the center of the observation gel. The well was filled completely with perfluorodecalin (PFD, Sigma) using a Pasteur pipet. The leaf disc was placed in the PFD filled well with the infiltrated side facing up. A coverslip (VWR, cat. no.: 48366 045) was placed on top of the leaf disc to seal it into the well ensuring that no air bubbles remain trapped under. The prepared sample slide was placed on the microscopy and imaged by a Leica Laser Scanning Confocal Microscope TCS SP5 (Leica Microsystems, Germany). The imaging settings were: 40x wet objective (Leica Microsystems, Germany); 514nm laser excitation; ZStack section thickness: 2 nm; Line average: 4; PMT1: 550-615 nm; PMT2: 700-800 nm. Three to eight individuals (4 leaf discs for each plant) in total were used. Z-stacks (“xyz”)


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of two different regions were taken per leaf disc. Colocalization analysis was performed in LAS (Leica Application Suit) AF Lite software. Six line sections were drawn across the so-called “region of interest” (ROI) with the 30 µm interval on the DiI dye images. The corresponded distribution profiles of fluorescence intensity of DiI dye and chlorophyll for each ROI lines were plotted in Excel. The colocalization percentage of nanoceria in chloroplast was counted as the overlapped peaks of fluorescence emission of chloroplasts pigments and DiI labeled nanoceria.

Effect of plasma membrane potential depolarization on nanoceria transport in leaf mesophyll cells A solution of NaCl (100 mM, Fisher Chemical) was used to depolarize the plasma membrane potential.93,94,95,96 Leaves from four weeks old Arabidopsis plants were infiltrated with either NaCl + DiI-PNC1 or NaCl + DiI-ANC. An isotonic non-ionic solution mannitol (170 mM, Sigma) was used as a control for possible osmotic effects. The experiments were conducted at ambient temperature. Confocal imaging and colocalization analysis were performed as explained above.

Impact of temperature and auxin on nanoceria uptake Arabidopsis plants were exposed to 24 ± 1 ºC, 14 ± 1 ºC, and 4 ± 1 ºC before and after infiltration with nanoceria. In the 24 ºC treatment, plants were infiltrated separately with TES infiltration buffer (10 mM TES, 10 mM MgCl2, pH 7.5), DiI-PNC1 (0.4 mM, 45 mg/L, in TES infiltration buffer, pH 7.5), and DiI-ANC (0.4 mM, 45 mg/L, in TES infiltration buffer, pH 7.5) at ambient temperature. Nanoceria was allowed to incubate for 3hr. In the 4 ºC and 14 ºC treatments, plants were pre-adapted in the refrigerator (set to 4 ºC and 14 ºC respectively) for 2 hr under darkness. Then, plants were infiltrated separately with TES infiltration buffer, DiI-PNC1 and DiI-ANC, and kept in the refrigerator for another 3 h. Samples from plants kept at 4 ºC and 14 ºC, were kept in a cooler until confocal imaging. To investigate the role of endocytosis on nanoceria uptake, we used auxin as an endocytosis inhibitor.49,50 Plants were infiltrated with 0.4 mM (45 mg/L) DiI-PNC1 or DiI-ANC and 10 µM auxin (NAA, 1-naphthaleneacetic acid) under


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ambient temperature. Confocal imaging and colocalization analysis were performed as explained above.

In vivo monitoring of ROS scavenging by nanoceria For in vivo ROS detection, leaf discs from the infiltrated plants were incubated separately with 25 µM 2',7'-dichlorodihydrofluorescein diacetate (H2DCFDA, Thermo Fisher Scientific) (in TES infiltration buffer, pH 7.5) and 10 µM dihydroethidium (DHE, Thermo Fisher Scientific) (in TES infiltration buffer, pH 7.5) dyes in 1.5 mL Eppendorf tubes for 30 min under darkness. Both of the dyes are dissolved in DMSO. H2DCFDA is converted to its fluorescence form DCF (2',7'-dichlorofluorescein) upon the cleavage of the acetate groups by ROS. DCF is regarded as an indicator of the degree of general oxidative stress. Likewise, DHE fluorescence (fluorescent product 2-hydroxyethidium) increases upon reaction with superoxide anion. Confocal imaging was performed as explained above with modifications. The confocal microscope was manually focused on a region of leaf mesophylls cells. The leaf discs were exposed to 405 nm UV laser for 3 min. The fluorescence signal from the ROS dyes was collected and recorded. Three to eight individuals (4 leaf discs for each plant) in total were used. Time-series (“xyt”) measurement were taken per leaf disc. The imaging settings were: 496 nm laser excitation; PMT1: 500-600 nm; PMT2: 700-800 nm. ROS imaging with DHE and DCF was analyzed with ImageJ software (NIH). DHE and DCF fluorescence intensity was measured in Image J within the imaged region of spongy mesophyll cells. Relative increase of ROS signal intensity (ΔI) was calculated by the following equation: ΔI = (If-Io)/Io Where Io is the initial ROS signal intensity, and If is the final ROS signal intensity at each time point.

Leaf gas exchange and chlorophyll fluorescence Arabidopsis leaves that filled the entire gas analyzer chamber (2.5 x 1 cm2) were chosen for gas exchange and chlorophyll fluorescence measurements with a GFS-3000 (Walz). Leaves were infiltrated with TES buffer (control), PNC1, PNC2 and ANC followed by a 3 hr incubation period. Infiltrated leaves had similar chlorophyll content


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index (CCI, Apogee). To conduct excess light experiments, leaves were exposed to 2000 µmol m-2 s-1 PAR,97,98 a light intensity that plants experience under full sunlight under natural conditions.70,71 A-Ci curve measurements were performed at 800, 600, 500, 400, 300, 200, 100, 50 ppm Ci under excess light, followed by an A-light curve at decreasing light levels from 2000, 1600, 1200, 900, 600, 400, 300, 200, 100, 50 to 0 µmol m-2 s-1 PAR. In dark chilling stress treatment, plants were kept in 4 °C refrigerator under darkness for 5 days. In heat stress experiments, plants were measured at a cuvette temperature of 35 °C and 40% relative humidity. A-Ci curves were measured at a saturating light of 1200 µmol m-2 s-1 PAR, followed by A-light curve measurements at decreasing light levels from 1200, 900, 600, 400, 300, 200, 100, 50 to 0 µmol m-2 s-1 PAR. The cuvette temperature was set at 23 °C and 50% relative humidity for all treatments except heat stress. Note that cerium nitrate does not affect electron transport rates in chloroplasts.39 A-Ci curves were analyzed using the equation developed by Sharkey et al. (2007):99

A=Vcmax[(Cc ‒ Г*)/(Cc + KC(1 + O/KO))] ‒ Rd where Vcmax is maximum rate of carboxylation, Cc is the CO2 partial pressure at Rubisco, Γ* is photorespiratory compensation point, O is partial pressure of oxygen, Rd is mitochondrial respiration, and KC and KO are Michaelis constant of Rubisco for carbon dioxide and oxygen respectively. Vcmax was calculated by fitting our data to the model built by Sharkey et al. (2007)99 with Ci values below 250 ppm. The φCO2 (quantum yield of CO2 assimilation) was gained by calculating the slope of A response to the light level of 0, 50, 100, and 200 µmol m-2 s-1 PAR.

Monitoring plant chlorophyll content index under continuous excess light Arabidopsis plants with large, broad, and flat leaves were chosen for measuring chlorophyll content index (CCI) with an Apogee chlorophyll content meter. Among these individuals, plants were randomly chosen to be infiltrated with 450 µM (51 mg/L) PNC1, PNC2 (56 mg/L), ANC (51 mg/L) and TES infiltration buffer (10 mM TES, 10 mM MgCl2, pH 7.5) (control). CCI was monitored in three leaves from the rosette of each


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individual. The plants were infiltrated with PNC1, PNC2, and ANC solution by placing the leaf between a sterile needleless syringe (NORM-JECT®) and thumb, adjusting the pressure so as to minimize all damage to the leaf. These steps were repeated with the control group, infiltrating with TES buffer only. Plants were placed in a laboratory made growth chamber with a LED light source (CLG-150-36A, MeanWell) that provided an average 1300 µmol m-2 s-1 of continuous PAR. Three fans (Multifan S3 120mm, AC Infinity) cooled down the chamber to 25 ± 1°C. The individuals were placed in a square area inside the chamber right under the LED light source and the position of each individual randomized each day after measuring. The CCI of each individual was measured by taking 4 measurements per leaf, beginning at the basal end of the leaf and moving each successive measurement towards the apical end of the leaf, as close to the tip of the leaf as possible while still covering the entire measuring area of the Apogee meter with the leaf sample. The plants were under continuous light (no dark) for the duration of the experiment, and only removed once each day to measure CCI. Measuring ceased after seven days.

Statistical analysis All data (mean ± SD, n = biological replicates) were analyzed using SPSS 23.0 (SPSS Inc., Chicago, IL, USA). Comparison between treatments was performed by independent samples t-test (two tailed) or one-way ANOVA based on Duncan’s multiple range test (two tailed). The significance levels were *P < 0.05, **P < 0.01 and ***P < 0.001. Different lower case letters mean the significance at P < 0.05.

Author contributions J.P.G. and H.W. conceived experiments and wrote the paper. H.W. and N.T. conducted experiments. H.W., and J.P.G. performed data analysis. N.T. assisted in data analysis.

Conflict of interest The authors declare no competing financial interest.


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Acknowledgements This work was supported by the University of California, Riverside and USDA NIFA Hatch project No. 1009710. We thank Dr. Jinming Li, Ph.D. student Alex Rajewski, and undergraduate Cristina Moreno-Borja from University of California, Riverside for their technical help. XPS measurements were performed using the Kratos XPS Analytical Facility which is supported by NSF grant DMR-0958796.

Supporting information Supporting information available: . This material is available free of charge via the Internet at http://pubs.acs.org.

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Figure Legends Figure 1. Nanoceria colocalization with chloroplasts in leaf mesophyll cells. (a) Transmission electron microscope (TEM) images of the cerium oxide core of nanoceria. (b, c) Comparison of size and zeta potential between negatively charged poly (acrylic acid) nanoceria (PNC1, PNC2) and positively charged aminated poly (acrylic acid) nanoceria (ANC). (d) Representative confocal images showing colocalization of chloroplast autofluorescence with PNC1, PNC2 and ANC. Nanoceria were labeled with DiI fluorescent dye. (e) Chloroplast and nanoceria fluorescence intensity across a region of interest (ROI) in confocal image overlay. (f) Comparison between percentage colocalization of chloroplasts with PNC1, PNC2 and ANC. (g) Temporal patterns of PNC1, PNC2 and ANC colocalization with leaf mesophyll chloroplasts after leaf infiltration. Mean ± SD (n = 3-5). Statistical comparisons were performed using a oneway ANOVA based on Duncan’s multiple range test. Lower case letters represent significance differences at 0.05 level. Scale bar 50 µm.

Figure 2. Mechanisms of nanoceria transport into chloroplasts in vivo. (a) Schematic showing steps of nanoceria transport from leaf extracellular air spaces to mesophyll chloroplasts. Nanoceria are delivered into the leaf by infiltration through the stomata pores (Step 1). The nanoparticles are transported through leaf mesophyll cell walls (Step 2). Nanoceria binds to the outer side of the leaf mesophyll cell membrane where electrostatic interactions with the positively charged side of the membrane favor the binding of negatively charged PNC1 (Step 3). Nanoceria transport into cell cytosol and


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chloroplasts is not endocytosis dependent, but it is affected by the plasma membrane potential (MP) (Step 4). (b, c) Cell membrane depolarization with 100 mM NaCl influences the colocalization of both PNC1 and ANC with leaf mesophyll chloroplasts. Chloroplast colocalization with PNC1 increases whereas colocalization with ANC decreases. An osmotic control with mannitol (170 mM) does not have a significant impact on the colocalization of chloroplasts with PNC1 and ANC (P > 0.05). (d) Similar colocalization percentages of chloroplasts with PNC1 and ANC were observed in leaves infiltrated at 24 ºC, 14 ºC, and 4 ºC indicating that nanoceria move through a nonendocytic transport pathway. (e) No significant change in chloroplast colocalization with PNC1 and ANC in the presence of auxin, an endocytosis inhibitor. Different lower case letters represent a significant difference at P < 0.05 using a one-way ANOVA based on Duncan’s multiple range test. NS represents no significant difference. Mean ± SD (n = 38). Scale bar 50 µm.

Figure 3. Plant augmentation of ROS scavenging by nanoceria. (a) Schematic showing the mechanisms of nanoceria scavenging of ROS in chloroplasts. Briefly, excess light leads to electron transfer from PSI via ferrodoxin (Fd) to oxygen forming • –

superoxide anions (O ). Superoxide anion is catalyzed to hydrogen peroxide (H2O2) via superoxide dismutase (SOD). Hydrogen peroxide is either transformed to H2O and O2 through the reaction with ascorbate (AsA) and ascorbate peroxidase (APX) forming MDA (malondialdehyde) and H2O or to hydroxyl radical (OH•) via Fenton reaction. Hydroxyl radical is the most destructive ROS in plants and there is no known enzyme able to scavenge it. In the presence of nanoceria, superoxide anions, hydrogen peroxide, and hydroxyl radicals are catalyzed to oxygen, water, and hydroxyl ions, respectively. (b, c, d) Deconvoluted XPS spectra showing the surface valence states (Ce3+, Ce4+) of PNC1, PNC2 and ANC. The peaks at 879.4, 879.7, 879.9, 885.2, 899.2, 903.1 and 904.1 correspond to Ce3+, whereas the peaks at 883.5, 883.6, 898.0, 898.1, 901.2 and 901.6 indicate Ce4+. (e) ROS and superoxide generation were monitored by confocal imaging of DCF and DHE fluorescence, respectively, in leaf mesophyll cells exposed to 3 min of UV-A light (405 nm). Leaves were infiltrated with PNC1, PNC2, ANC and TES buffer as control (no nanoparticles, NNP). (f) Time series of DHE and DCF fluorescence


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ACS Nano

intensity calculated as the change between final (If) and initial (Io) fluorescence intensity normalized by Io. Statistics were performed by independent-samples t-test (SPSS 23, * P < 0.05, ** P < 0.01). Asterisks represent significant differences between leaves with nanoceria and buffer controls (NNP-Leaves). Mean ± SD (n = 4-5). Scale bar 50 µm.

Figure 4. Plant nanobionic enhancement of photosynthesis under excess photosynthetic active radiation (PAR). Response of photosynthetic parameters to excess PAR in leaves infiltrated with nanoceria and TES buffer as control (no nanoparticles, NNP). (a) PNC1 enhances quantum yields of PSII below 900 µmol m-2 s-1 PAR. (b) PNC1 enables higher maximum yields of PSII (Fv/Fm) after exposure to excess PAR. (c, d) CO2 assimilation (A) light curves show that PNC1-Leaves have significantly higher maximum CO2 assimilation rates and quantum efficiency of CO2 uptake (φCO2) (15 %). (e) A versus internal CO2 concentration (ci) curve indicates protection of the carbon reactions of photosynthesis by PNC1. (f) PNC1 promotes higher maximum carboxylation rates (Vcmax). Mean ± SD (n = 8-15). (g) Exposure of Arabidopsis plants to 1300 µmol m-2 s-1 of continuous light led to a decline in leaf chlorophyll content index (CCI). However, plants infiltrated with PNC1 but not PNC2 or ANC maintained a significantly higher CCI than those treated with TES buffer as control (NNP). PNC2Leaves showed significantly less CCI than NNP-Leaves after 3 days of continuous excess light. No significant differences in CCI were found between ANC-Leaves and NNPLeaves (P > 0.05). (h) PNC1 mitigated the damage to the leaf lamina of Arabidopsis plants exposed to continuous excess light. Mean ± SD (n = 15). A one-way ANOVA based on Duncan’s multiple range test was used for statistical analysis (b, d, f). Lower case letters represent significance at 0.05 level. Statistical comparisons in (a, c, e, g) were performed by Independent-Samples t-test by SPSS 23 (* P < 0.05, ** P < 0.01, *** P < 0.001). Asterisks represent significant differences between leaves with nanoceria and buffer controls (NNP-Leaves).

Figure 5. Nanoceria protection of photosynthesis carboxylation reactions from heat and dark chilling. (a, b) PNC1 infiltrated leaves exposed to heat (35 °C) have significantly higher carbon assimilation rates (A) (67% P < 0.05) and quantum yield of


37

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CO2 uptake (φCO2) (27%, P < 0.05) relative to controls without nanoparticles (NNP). (c, d) PNC1 also enables increased A per internal CO2 concentration (Ci) (61%, P < 0.01) and higher maximum carboxylation rates (Vcmax) (51%, P < 0.05) under heat stress. (e, f) Similarly, dark chilling stressed leaves infiltrated with PNC1 have enhanced A at a broad range of PAR levels (46%, P < 0.05) and higher φCO2 (24%, P < 0.05) than controls. (g, h) The A-Ci curves of dark chilled PNC1 plants show enhanced A per given Ci (49%, P < 0.05) and an increase of Vcmax up to 30% (P < 0.05) relative to leaves without nanoparticles. In contrast, PNC2-Leaves have lower levels in all the photosynthetic parameters described above relative to NNP-Leaves (P < 0.05). ANC-Leaves and NNPLeaves have similar photosynthetic performance under heat and dark chilling. Statistical comparisons in (a, c, e, g) were performed by independent-samples t-test between leaves with nanoceria and buffer controls (NNP-Leaves) (SPSS 23, * P < 0.05, ** P < 0.01, *** P < 0.001). One-way ANOVA based on Duncan’s multiple range test was used in (b, d, f, h). Different lower case letters represent significance at 0.05 level. Mean ± SD (n = 1012).


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PNC1 a

Size, nm

15 a

a

10 5 0

b

0 -10 -20

a

-30 PNC1

Chloroplasts

ANC

10

a

10 nm

(d)

PNC2

(c) 20

(b)

Zeta Potential, mV

20

(a)

Nanoceria

PNC2

ANC (e) 300

Overlay

PNC1

Chloroplasts

PNC1

200 100

ROI

0 0

Intensity, a.u.

300

PNC2

ROI

50

100

150

PNC2

200

Chloroplasts

200

100 0 0

50 100 ANC

150 200 Chloroplasts

0

50

150

300

ANC

200 ROI

100 0

(g) 60

(f) 60

a

b

30 20

Colocalization, %

a 40

200

40

30 20

***

***

***

180

240

***

** ***

10

10

100 µm

PNC1 PNC2 ANC

50

50

Colocalization, %

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58

Figure 1

ACS Nano

0

0 PNC1

PNC2

ANC

0

60

120

Time, min


300

360

Figure 2

ACS Nano

(a)

Chloroplast

Leaf mesophyll cell

Vacuole

Chloroplasts

Nanoceria

Overlay

a

PNC1 ANC

PNC1 + NaCl

(b) Cytosol

- - - MP 3

ANC + NaCl

4

++++

Apoplast (c) 70 2

60 Extracellular air space

1

Abaxial epidermis

Stomata guard cells

Colocalization, %

Cell wall

Nanoceria

50

b b

40 c

c

30

d

20 10 0 Nanoceria

(d) 60

PNC1 ANC

a

50

a

a

40 b

30

b

b

20 10 j

Nanoceria + NaCl

Nanoceria + Mannitol

(e) 60

Colocalization, %

50

Colocalization, %

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58

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a a

40

PNC1 ANC

b b

30 20 10

0

0 24 °C

14 °C

4 °C

Nanoceria


Nanoceria + Auxin

Page 41 of 44

(a)

‒

2O2

2H2O + O2

2OH Nanoceria

Nanoceria •

2OH 2 O•2 – 2O2

O2

Fenton reaction

2OH¯

H2O2

Fd

2MDA 2H+

2AsA

SOD

Stroma

H2O

APX

2e–

Thylakoid membrane

PSI

Lumen

Light

XPS intensity (a.u.)


(c)

883.6

901.6 879.7

899.2

PNC2 Experimental data Background Fitted data Ce4+ Ce3+

ANC Experimental data Background Fitted data Ce4+ Ce3+ 901.2

(d)


PNC1 Experimental data Background Fitted data Ce4+ Ce3+

(b)

883.5 901.6

879.9

898.0

883.5

898.1

879.4

903.1

904.1 885.2 915

905

895

885

875

915

905

NNP

895

885

875

915

905

Binding energy, eV

Binding energy, eV

PNC1

PNC2

895

885

875

Binding energy, eV

ANC (f)

(If - Io) / Io

DHE

DHE

(e)

DCF

1.8 1.5 1.2 0.9 0.6 0.3 0

NNP PNC1 PNC2 ANC *

0

DCF

Fluorescence

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58

Figure 3

ACS Nano

1.8 1.5 1.2 0.9 0.6 0.3 0

**

*

** ****** **** **** * * * * * ** 100 200 NNP PNC1 PNC2 ANC **

0


****

** ** *

**

**

*

** ** ** ** **

** ******* *** 100 200 Time, s

Figure 4

ACS Nano

(a)

0.6 *****

NNP PNC1 PNC2 ANC

* * 0.4

(b)

0.2

1000 1500 PAR, µmol m-2 s-1

12

φCO2

** ** ***

*** NNP PNC1 PNC2 ANC

1000 1500 PAR, µmol m-2 s-1

2000

** ** * ** * *

0

***

**

0

A, µmol CO2 m-2 s-1

25

500

NNP PNC1 PNC2 ANC *

20 15

* *

10 * 5 0

*** ***

-5 0

a

*** ***

***

***

***

ab c

0.02

0.01

0 NNP

PNC1

PNC2

ANC

(f) 200

*

*

a 150

b b

100

c

50

0 300

600

900

NNP

PNC1

PNC2

ANC

ci, ppm

(g) 12

Day 0

Day 3

*

*

*

*

*

*

*

*

4

5

6

6 4

NNP PNC1 PNC2 ANC

2 0 0

1

*

2

3

*

**

NNP

*

PNC1

Chlorophyll content index

(h) 10 8

a

Final

b

-2

(e)

(d) 0.03

**

2

a

0.4

**

4

b

PNC1 ANC

0.6

Initial

*

*

*

*

6

NNP PNC2

0

2000

Vcmax, µmol CO2 m-2 s-1


Excess light

500

8

a

0.2

* 0

10

a

a *

(c)

a

0.8

*

0

1 a

Fv/Fm

PSII quantum yield

*

Continuous excess light

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58

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7

Day


Day 6

Page 43 of 44

8

NNP PNC1 PNC2 ANC

6

*

**

*

600

800 1000 1200

NNP

14

* **

11 8 5 2

PNC2

ANC

a

140 120 100

b b

80 60

c

40 20 0

0

500 ci, ppm

1000

NNP

PNC1

PNC2

ANC

(f) 0.03 NNP PNC1 PNC2 ANC *

8 6 4

*

b

* *

*

2

a

*

φCO2

(e) 10


*

*

*

**

PNC1

(d) 160

*

*

c

0


NNP PNC1 PNC2 ANC

17

-1

b 0.02

PAR, µmol m-2 s-1

(c) 20

b

0.01

*

*

b

0.02 c 0.01

*

*

*

0

0 0

200

-2

400

600

800 1000 1200

NNP

PNC1

PNC2

ANC

PAR, µmol m-2 s-1

(g) 15

**

NNP PNC1 PNC2 ANC

12 9

*

(h) 120

** **

**

*

6 3

*

0 0

* **

**

300

**

600

-3

***

900



0.03

*

* * 200 400

-4


a

*

2 0

(b) 0.04

*

*

4

-2 0


Heat (35°C)

10

*

*

φCO2

(a) 14 12

Dark chilling (4°C)

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58

Figure 5

ACS Nano

100

a b

b

80 60

c

40 20 0 NNP

PNC1

ci, ppm


PNC2

ANC

ACS Nano

Light

Chloroplast

2O2

Light reactions

2e– PSI

Fd 2H+

2O•2 –

(CH2O)n

2O2

Thylakoids

SOD

Nanoceria O2

Fenton reaction H2O2 • 2OH 2AsA 2OH

APX

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42

Page 44 of 44

H2O

Thylakoid membrane

2OH

‒

‒

2H2O + O2

2MDA


CalvinBenson cycle Carbon reactions (stroma)

CO2 + H2O

Anionic Cerium Oxide Nanoparticles Protect Plant Photosynthesis

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