Uranium exposure induces nitric oxide and hydrogen peroxide generation in Arabidopsis thaliana

Environmental and Experimental Botany 120 (2015) 55–64

Contents lists available at ScienceDirect

Environmental and Experimental Botany journal homepage: www.elsevier.com/locate/envexpbot

Uranium exposure induces nitric oxide and hydrogen peroxide generation in Arabidopsis thaliana Rajesh Tewaria,1, Nele Horemansa,b,1,* , Robin Nautsa , Jean Wannijna , May Van Heesa , Hildegarde Vandenhovea a b

Biosphere Impact Studies, Belgian Nuclear Research Center (SCK
CEN), Boeretang 200, Mol 2400, Belgium Hasselt University, Centre for Environmental Sciences, Agoralaan gebouw D, B-3590 Diepenbeek, Belgium

A R T I C L E I N F O

A B S T R A C T

Article history: Received 4 March 2015 Received in revised form 22 July 2015 Accepted 19 August 2015 Available online 22 August 2015

Uranium (U) is a nonessential and toxic heavy metal and radioactive element. Nitric oxide (NO) and hydrogen peroxide (H2O2) are intimately involved in the growth, development and metabolic function of plant cells. In addition to possible toxic activities, a role for these signalling molecules in the regulation of plant responses to toxic metals has been proposed. Here, we investigated the synthesis of NO and H2O2 in Arabidopsis thaliana seedlings exposed to 25 mM U. We demonstrated that U induces NO and H2O2 production in the roots and leaves. Addition of the NO releasing compound, sodium nitroprusside (SNP) as well as the NO synthase inhibitor (L-NAME) to the plants, resulted in a decrease of the U-induced NO production and both alleviated U toxicity as indicated by plant growth (leaf area and biomass). Treatment with L-NAME further resulted in the disappearance of the U-induced H2O2 in the shoots of the plants. It appears that generation of NO and H2O2 is intimately linked in the toxicity response of U-exposed A. thaliana plants. ã 2015 Elsevier B.V. All rights reserved.

Keywords: Arabidopsis thaliana Hydrogen peroxide Lipid peroxidation Nitric oxide Uranium toxicity

1. Introduction Uranium, is a metal and radioactive element existing as a mixture of three radioisotopes U238, U235, U234 (Sheppard et al., 2005). Among uranium isotopes, U238 is most prevalent in natural occurrence (99.27%) (Sheppard et al., 2005). Uranium concentration in the earth’s crust may range from 1 to 4 mg kg1 in sedimentary rocks to even hundreds of mg kg1 in phosphate rich deposits or in U ore deposits (Srivastava et al., 2010). Due to industrial activities like U-mining and milling, the phosphate industry and the application of this heavy metal in nuclear energy production, there exists a risk of leakage of uranium into the terrestrial and aquatic environment. Uranium exposure adversely affects plant growth, leads to an altered photosynthesis efficiency

Abbreviations: cPTIO, 2-(4-carboxy-2-phenyl)-4,4,5,5-tetramethyl-imidazoline1-oxyl-3-oxide; DAF-2DA, 4,5-diaminofluorescein diacetate; DAF-2T, DAF-2 triazole; H2DCF-DA, 2,7-dichlorodihydrofluorescein diacetate; L-NAME, Nv-nitro-Larginine methyl ester hydrochloride; MB, methylene blue; NO, nitric oxide; SNP, sodium nitroprusside. * Corresponding author at: Biosphere Impact Studies, Belgian Nuclear Research Center (SCK
CEN), Boeretang 200, Mol 2400, Belgium. E-mail addresses: [email protected] (R. Tewari), [email protected] (N. Horemans), [email protected] (R. Nauts), [email protected] (J. Wannijn), [email protected] (M. Van Hees), [email protected] (H. Vandenhove). 1 Both authors contributed equally. http://dx.doi.org/10.1016/j.envexpbot.2015.08.004 0098-8472/ ã 2015 Elsevier B.V. All rights reserved.

and induces oxidative damage as evident by increased lipid peroxidation in Arabidopsis thaliana (Saenen et al., 2013; Vandenhove et al., 2006; Vanhoudt et al., 2011c, 2008). Nitric oxide (NO) is a biologically active, diffusive, water and lipid soluble diatomic gaseous free radical. It has been shown to be a ubiquitous signalling molecule in animal, plant and microbial systems effective even at nanomolar concentration (1.0 nmol L1) but so far its involvement in U-induced stress responses has not been studied. NO is shown to be a key modulator of antioxidant defense system and plays a crucial role in plant survival under toxicities of various metals such as Al (He et al., 2012; Tian et al., 2007), Cd (Singh et al., 2008), Cu (Wang et al., 2010) and Zn (Xu et al., 2010). Nitric oxide is known to form complexes with transition metal ions in aqueous media (Begara-Morales et al., 2013). Metal-nitrosyl complexes may act as links between the different redox states of NO under neutral physiological conditions. Studies with exogenous application of the NO donor sodium nitroprusside (SNP) revealed that NO can alleviate copper toxicity-induced effects by up regulating the antioxidant defense system and keeping accumulation of H2O2 under control (Tewari et al., 2008a). SNP application protected breakdown of chloroplastic pigments and improved biomass of plants under copper toxicity (Wang et al., 2010). An exogenous application of NO releasing compound along with heavy metal also up-regulated transcript levels of ferritin (Murgia et al., 2004) and metallothionein (Wang et al., 2010).

56

R. Tewari et al. / Environmental and Experimental Botany 120 (2015) 55–64

Hydrogen peroxide (H2O2) is the two electron reduction product of molecular oxygen (O2). It is a potentially reactive oxygen species (ROS), but not a free radical. However, in the presence Fe2+, it produces highly reactive OH radical in the Fenton reaction, leading to the peroxidation of membrane lipids (Cheeseman, 2007). On the other hand, the membrane-permeable H2O2 has been shown to play a key role as a signalling molecule, involved in growth, development, and response to environmental stimuli. Also an H2O2-dependent long-distance signal has been shown to exist and to be induced by various abiotic stimuli (Miller et al., 2009). The plant respiratory burst oxidase homologues (RBOHs) are cell membrane-associated proteins involved in the generation of superoxide anion (O2) outside of cell membrane in the apoplast (Baxter et al., 2014; Kobayashi et al., 2007). RBOHs were shown to be required for the initiation and self-propagation of a rapid cell to cell systemic signal that is dependent upon H2O2 accumulation in the extracellular spaces (apoplast) to generate a ‘ROS wave’  (Mittler et al., 2011). The O2  dismutates to H2O2 spontaneously or catalytically by the action of superoxide dismutases (Munoz et al., 2005). Hydrogen peroxide accumulation has often been reported in plants exposed to various stresses such as summer drought (Jubany-Mari et al., 2008), hypoxia (Vergara et al., 2012), mineral nutrient deficiencies (Shin and Schachtman, 2004; Tewari et al., 2013a, 2006, 2004) and heavy metals toxicity caused by Cd (Kumar et al., 2008a), Cu (Kumar et al., 2008b), Fe (Kumar et al., 2008b), Ni (Kumar et al., 2007) and Zn (Tewari et al., 2008b). However, information on U-induced H2O2 generation or their interaction with the induction of toxic responses in U-exposed plants is unavailable in the literature. As the U toxicity originates from its chemotoxicity as a heavy metal rather than its radiotoxicity (Bleise et al., 2003), it was hypothesised that as for other metals NO and H2O2 play a role in signal transduction leading to the plant response to U. This hypothesis was tested by measuring H2O2 and NO generation in A. thaliana plants exposed to 25 mM of uranyl nitrate and the implications of exogenous supply or scavenging of NO on the observed U-induced growth retardation. 2. Materials and methods 2.1. Chemicals All chemicals were purchased from Sigma–Aldrich unless otherwise stated. Depleted uranium salt, Uranyl nitrate [UO2(NO3)2
6H2O] was purchased from SPI Chemicals, USA. 2.2. Plant material and growth conditions A. thaliana Col-0 seeds were stratified and grown in aerated hydroponic culture condition with modified Hoagland medium (final concentration of salts: 1.0 mM KNO3, 0.3 mM Ca(NO3)2, 0.2 mM MgSO4, 0.1 mM NH4H2PO4, 1.62 mM FeSO4, 0.78 mM Na2EDTA.2H2O, 4.6 mM H3BO3, 0.9 mM MnCl2, 32 nM CuSO4, 56.6 nM H2MoO4, 76.5 nM ZnSO4) in a growth chamber held at day/night temperature 22/18  C, light/dark period 14/10 h, humidity 65% and light intensity 150 mmol m2 s1. Plants were exposed to 0 mM or 25 mM U under different NO producing conditions either control Hoagland medium or with 200 mM sodium nitroprusside (SNP, NO producer) or 200 mM methylene blue (MB, NO scavenger), or 200 mM Nv-nitro-L-arginine methyl ester (L-NAME, a NO synthase inhibitor). The U concentration of 25 mM was chosen based on previous experiments in which it was shown to induce a decrease in growth but not plant dead within the exposure time used here (Saenen et al., 2013). As a NO scavenger 24-carboxyphenyl-4,4,5,5-tetramethylimidazoline-1-oxyl-3-oxide (cPTIO) is often used in contrast to MB. However, to expose

hydroponic plant cultures for 3 days to different NO modulating compounds the use of MB was chosen above cPTIO for a number of reasons. First, it has been shown that cPTIO is not very stable and can degrade within minutes in Arabidopsis cell cultures (D’Alessandro et al., 2013). Secondly, cPTIO has been shown to sometimes interfere with NO detection (D’Alessandro et al., 2013; Vitecek et al., 2008) and finally out of economic considerations as cPTIO is a very expensive chemical. At the beginning of the experiment the pH of Hoagland medium was adjusted to 5.5 with NaOH, if required. After 3 days or 5 days of U-exposure plants were analysed for leaf area and fresh weight and snap-frozen in liquid nitrogen in 50–100 mg fractions, except for those stated otherwise. Frozen samples were stored at 80  C until further biochemical analyses. NO and H2O2 was localized in freshly harvested non-frozen roots exposed for 3 or 5 days to U as stated in the figure legends. 2.3. Determination of total leaf area Pictures of A. thaliana rosette were taken after 3 days of U-exposure and images were analyzed for total leaf area using Image J software (http://imagej.nih.gov/ij/) as described previously (Vanhoudt et al., 2014). 2.4. Determination of uranium concentration Uranium concentration in leaf and root was measured exactly as described previously by Vanhoudt et al. (2014) with ICP-MS (PerkinElmer Elan 5000 utilizing a cross flow nebulizer and a ryton spray chamber, Perkin–Elmer, Waltham, MA, USA). 2.5. Hydrogen peroxide H2O2 concentration was determined as H2O2–titanium complex as described previously (Brennan and Frenkel, 1977; Kumar et al., 2010). Briefly, freshly harvested plant tissue (100 mg fresh weight) was snap-frozen in liquid nitrogen and powdered by using tungsten carbide beads and mixer mill for 3.5 min at 30 Hz. Subsequently hydrogen peroxide was extracted by addition of 200 mL of ice cold acetone:water (2:1) mixture and clarified by short centrifugation. To the acetone extract (100 mL) 10 mL of titanium reagent (20 % (v/v) titanic tetrachloride in concentrated HCl) was added followed by addition of 20 mL of concentrated NH4OH to precipitate H2O2–titanium complexes. The H2O2– titanium complexes were subsequently pelleted by centrifugation for 5 min at 10,000  g and were repeatedly washed with ice cold acetone to remove the pigments. Finally the pellet was solubilized in 2 N H2SO4 and absorbance of the solution was read at 415 nm. 2.6. H2O2 localization in roots Plants were placed in loading buffer (5 mM MES-KOH, pH 5.7, 0.25 mM KCl, 1 mM CaCl2 0.5 M sorbitol) containing 10 mM 2,7dichlorodihydrofluorescein diacetate (H2DCF-DA) for 30 min. Samples were washed three times for 15 min in 5 mM MES buffer (pH 5.7) and were mounted on the microscope slides using same buffer. DCF-fluorescence was monitored with a Nikon Eclipse Ti fluorescence microscope using a FITC filter; excitation, 490 nm; emission, 515 nm. Similarly treated unstained roots were used as controls to check autofluorescence, if any, for microscope setting. 2.7. Nitrate and nitrite measurements Liquid N2 frozen leaf and root tissue (100 mg) were powdered using tungsten carbide beads and mixer mill for 3.5 min at 30 Hz. Powdered sampled were mixed in 2 mL of 50 mM acetate buffer pH

R. Tewari et al. / Environmental and Experimental Botany 120 (2015) 55–64

57

3.6, containing 4% (w/v) zinc diacetate. Pigments were removed by mixing leaf tissue extract (2 mL) with activated charcoal (25 mg) using vortex mixer and subsequently centrifuged twice at 10,000  g for 10 min each (Zhou et al., 2005). Nitrite content was determined by incubating equal amounts of tissue extract and Greiss reagent for 90 min at 37  C as described previously (Tewari et al., 2013b). Absorbance was determined at 540 nm. Similarly, nitrate plus nitrite was measured in the same extract by including VCl3 for reduction of nitrate to nitrite. Nitrite concentration was calculated by comparison to a standard curve of NaNO2.

diaminofluorescein-2 diacetate (DAF-2DA) for 1 h in the dark, and subsequently washed with loading buffer 3 times for 5 min each. The fluorescence of specific regions of the roots was monitored using a fluorescein isothiocyanate (FITC) filter; excitation, 490 nm; emission, 515 nm with a Nikon Eclipse Ti fluorescence microscope (Nikon, Japan). Similarly treated unstained roots were used to control autofluorescence of samples, if any. At least five different roots were scanned for each treatment. Images were analyzed by selecting region of interest using ImageJ software (http://imagej. nih.gov/ij/) and data were normalized against background.

2.8. Nitric oxide localization in roots

2.9. Lipid peroxidation

At time of harvest plants grown in hydroponic culture were placed in a loading buffer [5 mM 4-morpholineethanesulfonic acid (MES)-KOH, pH 5.7, 0.25 mM KCl, 1 mM CaCl2] containing 5 mM

Plant tissues (100 mg fresh weight) were homogenized in 1 mL of 0.1% TCA and centrifuged at 10,000  g for 10 min. Lipid peroxidation was determined in terms of thiobarbituric acid

Fig. 1. Uranium concentration (A), dry weight (B) and lipid peroxidation (C) in the shoots and roots of mock-treated (control) or uranyl nitrate (25 mM)-treated Arabidopsis seedling for 3 day, H2O2 localization by H2DCF-DA in the roots (D) and H2O2 concentration by Ti-reagent in the shoots (E) and roots (F) of mock-treated (control) or uranyl nitrate (25 mM)-treated Arabidopsis seedling for 5 days. Data are mean of six replicates (SE). Bars carrying different letters in an individual organ are significantly different (p  0.05) by the Fisher-LSD method.

58

R. Tewari et al. / Environmental and Experimental Botany 120 (2015) 55–64

reactive substances (TBARS) (Kumar et al., 2010). The amount of TBARS was calculated from the difference in absorbance at 532 nm and 600 nm using extinction coefficient of 155 mM 1 cm 1. 2.10. Statistical analysis Significant differences between means were determined by ANOVA and subsequent multiple pairwise comparison by 95% Fisher LSD method using Sigma Stat (Systat Software Inc., San Jose, CA). 3. Results and discussion 3.1. Uranium exposure adversely affects shoot growth Treating A. thaliana plants with 25 mM U resulted in an U accumulation in the shoots and roots (Fig. 1A), and inhibited plant growth as indicated by decreased shoot dry weight (Fig. 1B). Most of the U was retained by the roots and less than 0.04 % of the total U was transferred to the shoots which caused a significant inhibition in shoot growth. A small root-to-shoot transport of U is in consonance with previous studies conducted on A. thaliana exposed to 25 mM U at various pH regimes (Saenen et al., 2013; Vanhoudt et al., 2014, 2008). A similar inhibition in the shoot growth of U-exposed A. thaliana plants has been reported

previously (Saenen et al., 2013; Vanhoudt et al., 2011a, 2014, 2008). In the roots, U exposure, surprisingly, did not cause any decrease in dry weight (Fig. 1B), despite of several-thousands (2786)-fold higher U accumulation in the roots. The weight of the roots of U-treated plants was even slightly higher than untreated control plants (Fig. 1A). Increased root weight of U-treated plants has been observed previously in studies with A. thaliana exposed to 25 mM U (Saenen et al., 2013). However, higher concentrations of U have been shown to inhibit root growth in A. thaliana plants (Vanhoudt et al., 2014, 2011c). This differential effect of U to the roots is probably related to differences in the applied U concentrations. Together, present observations and previous studies with A. thaliana suggest that U-induced effects are more prevalent in the shoots compared to the roots at least during the first 3-days of U exposure. The U toxicity of the shoots can possibly be linked to its influence on the uptake and distribution of several nutrients that will lead to hampered nutrient profiles in the plants as was suggested by Vanhoudt et al. (2011b). Uranium exposure for 3-days did not affect lipid peroxidation either in the shoots or in the roots significantly (Fig. 1C) indicating limited or no oxidative damage induced by 25 mM U. An increased lipid peroxidation in shoots at pH 4.5 for 25 mM (Saenen et al., 2013) and at pH 5.5 for 50 mM (Vanhoudt et al., 2014) has already been reported. However, as shown in Saenen et al. (2013) plants exposed to similar conditions as used here, namely pH 5.5 and 25 mM of U, no

Fig. 2. NO localization by DAF2-DA in the roots (A), and nitrite concentration in the shoots (B) and roots (C) of mock-treated (control) or uranyl nitrate (25 mM)-treated Arabidopsis seedling for 3 days. Data are mean of six replicates (SE). Bars carrying different letters in an individual organ are significantly different (p  0.05) by the FisherLSD method.

R. Tewari et al. / Environmental and Experimental Botany 120 (2015) 55–64

significant difference in lipid peroxidation between control and treated plants was observed. 3.2. Uranium exposure induces H2O2 generation Hydrogen peroxide concentration estimated as H2O2–Ti complexes, was enhanced in the shoots but not in roots (Fig. 6D) of three day U-exposed plants compared to controls. An extension of the U-exposure to five days, however, significantly enhanced H2O2 levels both in shoot and roots (Fig. 1E and F). Additionally, in the roots DCF-fluorescence was increased in 5-days U-exposed plants compared to control plants (Fig. 1D). These observations suggest that U exposure induced H2O2 generation in A. thaliana. An accumulation of H2O2 in the leaves of maize plants exposed to different heavy metals such as Cd (Kumar et al., 2008a), Cu (Kumar et al., 2008b; Tewari et al., 2008a), Fe (Kumar et al., 2008b) and Ni (Kumar et al., 2007) has already been reported. Higher H2O2 accumulation in U-exposed roots and leaves may be associated with the increases in expression of RBOH-C,RBOH-D, and FSD1 (Vanhoudt et al., 2011c).

59

2011). Despite these drawbacks, DAFs dyes are still in frequent use for visualizing NO production in biological systems (Corpas and Barroso, 2014; Hu et al., 2014; Li et al., 2013; Wei et al., 2014) due to ease of its application and detection with generally available instrumentation. Moreover, an increase of DAF-2T fluorescence of the roots treated with NO releasing compound, SNP (Fig. 4A and B), compared to untreated control roots suggest that DAF-2DA indeed detected endogenous levels of NO in our experimental system. Together with the elevation in nitrite levels these data were taken as the first indication that NO is being induced in A. thaliana plants exposed to U for three days. Prevention of DAF-2T fluorescence of the roots pre-incubated with a NO scavenger, methylene blue (MB) (Fig. 2A) prior to staining with DAF-2DA, further suggests NO generation in the roots of U-treated plants. In this study, MB was chosen as a NO scavenger as it has been shown to be a more effective NO scavenger compared to well-known NO scavenger, c-PTIO (Vandana et al., 2012). Methylene blue inhibits soluble guanylate cyclase and thereby the action of NO and cGMP (Juffermans et al., 2010). Nonetheless, MB also inhibits NO production by inhibiting inducible-NO synthase (Juffermans et al., 2010).

3.3. Uranium exposure induces NO generation NO generation was measured in DAF-2T stained roots or as concentration of nitrite, an oxidation product of NO (Tewari et al., 2013b) in root and shoot tissue of both control and three day Utreated plants. Uranium exposure induced NO generation as indicated by increases in DAF-2T fluorescence (Figs. 2 A and 4 A,B) in the roots and concentration of nitrite, an oxidation product of NO (Tewari et al., 2013b), in the shoots (Figs. 2 B and 5 B) and roots (Figs. 2 C and 5 B). These results are comparable to the response of Panax ginseng exposed to 50 mM Cu where a similar increase in DAF-2T fluorescence and in nitrite concentration was observed in the adventitious roots (Tewari et al., 2008a). However, the specificity of DAFs reaction with NO has recently been questioned as DAF-reactive compounds were prevalently identified as reaction products of H2O2 plus apoplastic peroxidase and only minor peak was detected for DAF-2T, a reaction product of DAFs with NO (Rumer et al., 2012). On the other hand, O-diamine moiety of DAF2DA has been shown to be more selective for NO over NO3, ONOO, H2O2, and O2 under aerobic conditions and maintains a 5 nM NO detection limit (Pluth et al., 2011). Measurement of NO by DAF dye may also be perturbed by the presence of dehydroascorbic acid or ascorbic acid, glutathione and homocysteine (Pluth et al.,

3.4. Effects of exogenous NO modulators on U uptake in roots and shoots To further study the possible NO production and its role in the response of A. thaliana plants to U, NO-modulators were added to the growth medium during the whole U-exposure period in an attempt to artificially modulate NO levels. As a NO-producer 200 mM SNP was used whereas MB (200 mM) and L-NAME (200 mM) were added to the growth medium as NO-scavenger and NO-synthase inhibitor respectively. At the end of the three day exposure plants were scored for NO and H2O2 production, U-uptake and growth responses. The NO producer, SNP, did not affect the accumulation of U in the shoots (Fig. 3A), but it reduced U uptake with 67% U compared to controls roots (100 %) (Fig. 3B). As such, a higher root-to shoot transfer is observed. SNP has been shown to inhibit Al-uptake in wheat and rye roots exposed to 50 mM Al (He et al., 2012) and Cduptake in Medicago truncatula seedling exposed to 50 mM Cd (Xu et al., 2009). MB did not show any significant effect either on uptake or transport of U (Fig. 3A and B). However, most of the studies related with exogenous application NO modulating compounds on metal toxicity as cited above did not provide

Fig. 3. Uranium concentration in the shoot (A) and roots (B) of Arabidopsis plants either mock-treated (control) or treated with 200 mM SNP and 200 mM MB with or without 25 mM uranyl nitrate. Data are mean of six replicates (SE). Bars carrying different letters are significantly different (p  0.05) by Fisher-LSD method for an individual organ (shoot or root).

60

R. Tewari et al. / Environmental and Experimental Botany 120 (2015) 55–64

information about the effects of these compounds on the concentration of metal under investigation in the shoots. These studies are basically focussed on metal concentration in the roots only. 3.4.1. Uranium induced nitric oxide production in plants grown in the presence of nitric oxide modulators The application of 200 mM MB to the A. thaliana plants during three days turned out to induce severe toxicity and oxidative stress as evidenced by stunted growth of the plants (Fig. 6A and B), a high lipid peroxidation in the shoots (Fig. 6C) and an over accumulation of H2O2 in the shoots as well in roots (Fig. 6D). A. thaliana plants treated with MB plus U experienced even more pronounced oxidative stress, which was also reflected by a further decrease in plant growth (Fig. 6A and B) and further increase in lipid peroxidation (Fig. 6C) in the shoots. MB is a known photosensitizer and therefore, it produces singlet oxygen in the presence of light and oxygen. Singlet oxygen may subsequently react with alkenes to form organic peroxides via Diels–Alder reaction. Therefore, an enhanced level of lipid peroxide and H2O2 are expected for plants co-treated with MB and U. A similar inhibition of plant growth along with increase in ROS levels and loss of membrane integrity have been reported in M. truncatula seedlings co-treated with Cd and NO scavenger, c-PTIO (Xu et al., 2009). On the other hand, the toxicity induced by MB may also be attributable to a complete abolishment of NO production measured as DAF-2T fluorescence (Fig. 4A and B) or to a disturbed nitrate/nitrite balance. As such, MB-treated shoots irrespective of U-treatment showed decreased nitrate and increased nitrite concentrations (Fig. 5A and B). MBtreated roots exhibited a decrease in nitrite concentration. The difference in root-shoot nitrite production upon MB treatment is possibly explained by MB induced ROS production interfering with

photochemical processes and hence also with ferrodoxin production. The latter is an essential electron donor of nitrate reductase. Thus, higher nitrite levels in shoots could hypothetically be explained by a changed nitrite reductase activity. The accumulation of nitrite in the shoots of MB and MB plus U-treated plants (Fig. 5B) suggests that MB probably either inhibited the pathway for further reduction of nitrite into ammonium and its assimilation into amino acids or available nitrate in shoots was reduced to nitrite by reduced-MB (NO3 + MBH2 ! NO2 + MB+ + H2O) (Egami and Taniguchi, 1965). The concentration of nitrite in plants is regulated by rate of reduction of both nitrate and nitrite apart from oxidation of NO to nitrite. The MB, like PTIO, a well-known NO scavenger, may oxidize NO to nitrite (NO + MB+ + H2O ! NO2 + MBH2) (Arita et al., 2006) and therefore, shoots co-treated with MB and U over-accumulated nitrite as P. ginseng roots co-treated with salicylate and PTIO accumulated large amount of nitrite (Tewari and Paek, 2011). Nitrite is well-known for its toxicity in plants (Oke, 1966). It is therefore conceivable that excessive accumulation of nitrite in shoot can cause nitrite toxicity and result in a seriously hampered plant growth. As the MBtreatment applied here resulted in high plants toxicity it is difficult to discriminate between MB- or U- induced effects or to study the involvement of NO in these responses and therefore MB-treated plants are no longer considered in the following paragraphs. Treating A. thaliana plants with the exogenous NO-producer SNP resulted in an increased DAF-2T fluorescence indicating increased NO production. However, contradictory to the expectations in plants exposed to a combination of a SNP and U, a suppressed DAF-2T fluorescence was observed compared to the individual treatment of SNP or to the U-treatment without SNP (Fig. 4A and B). Adventitious roots of Panax ginseng also behaved similarly with co-application of Cu and SNP (Tewari et al., 2008a).

Fig. 4. Effect of U on NO production as measured by DAF-2T fluorescence in roots of Arabidopsis. (A) Arabidopsis seedling were either mock-treated (control) or treated with 200 mM SNP, 200 mM MB, and 200 mM L-NAME with or without 25 mM uranyl nitrate and stained with the NO-sensitive fluorescent dye, DAF-2DA for 1 h. Scale bars: 100 mm. (B) Relative DAF-2T fluorescence in the roots as described above. Data are mean of five replicates (SE). Bars carrying different letters are significantly different (p  0.05) by the Fisher-LSD method.

R. Tewari et al. / Environmental and Experimental Botany 120 (2015) 55–64

61

Fig. 5. Concentration of nitrate (A) and nitrite (B) in the shoot and roots of Arabidopsis plants either mock-treated (control) or treated with 200 mM SNP, 200 mM MB, and 200 mM L-NAME with or without 25 mM uranyl nitrate. Data are mean of six replicates (SE). Bars carrying different letters for an individual organ (shoot or root) are significantly different (p  0.05) by the Fisher-LSD method.

Down-regulated NO levels in the roots co-treated with SNP and U, might be associated with feedback down-regulation due to the presence of an external NO producer, SNP. In Cd-exposed A. thaliana roots, it was reported previously that treatment with 5 mM of the NO synthase inhibitor L-NAME abolished DAF-2T fluorescence (Besson-Bard et al., 2009). In contrast to our expectations, L-NAME (200 mM) treatment of roots did not result in a significant change in NO-production measured as DAF-2T fluorescence compared to non-treated control plants (Fig. 4A and B). The lack of effect of L-NAME on DAF-2T fluorescence may be attributed to 25-times-lower concentration of L-NAME (200 mM) applied in our experiment [compared to 5 mM L-NAME used by Besson-Bard et al. (2009)]. However, the increased NO-production observed in U-treated plants as compared to control plants was no longer present in plants treated with L-NAME and U simultaneously when compared to those treated only with L-NAME. This indicates that U-induced NO-production was no longer measurable in L-NAME treated plants and further supports the idea that NO production is induced by U. Nitrite concentrations were measured in both shoots and roots of the plants treated with different NO-modulators with or without U (Fig. 5B). Nitrite, as an oxidation product NO in the plants (Pluth et al., 2011) is taken here as a measure for NO production. However, as indicated above for MB, care must be taken as modulators of NO might also modulate N-metabolism in different ways independent of NO production. In the roots differences measured in nitrite concentrations between the different treatments were small but a significant increase in nitrite production compared to the control

plants is present in U and SNP+U treated roots (Fig. 5B). It must be taken into account that the applied nitrite measurements do not detect degradation products of nitrite like ammonia that might be formed in the chloroplast. Therefore, nitrite production, and hence NO production, as measured here might be underestimated. No significant difference in nitrite production between SNP and SNP +U could be observed (Fig. 5B). For the shoots the picture is identical to the results obtained in the DAF-2T stained roots. Namely, compared to non-treated controls a marked increase in nitrite concentration was found in plants treated with either SNP or U (Fig. 5B). However, treatment with both SNP and U simultaneously resulted in a decrease in production when compared to SNP or U alone. Also for L-NAME the results for nitrite concentration in the shoots and roots were comparable to the picture found with DAF-2T in the roots namely that no U-inducible increase in nitrite could be observed. However, in contradiction to the expectations nitrite levels were higher in L-NAME treated plants compared to non-treated controls (Fig. 5B). Taken together, these results further support an U-inducible NO production. 3.4.2. Effects of exogenous NO modulators on uranium toxicity and hydrogen peroxide production To test the toxicity of U in plants treated simultaneously with different NO-modulators fresh weight of root and shoot (Fig. 6A), the leaf surface area (Fig. 6B) and lipid peroxidation was measured (Fig. 6C). Whilst an increasing trend in the amount of MDA formed in the shoots was shown, no significant differences between U

62

R. Tewari et al. / Environmental and Experimental Botany 120 (2015) 55–64

Fig. 6. Fresh weight of shoot and roots (A), leaf area (B), lipid peroxidation (C) and H2O2 (D) in the shoot and roots of Arabidopsis plants either mock-treated (control) or treated with 200 mM SNP, 200 mM MB, and 200 mM L-NAME with or without 25 mM uranyl nitrate. Data are mean of six replicates (SE). Bars carrying different letters are significantly different (p  0.05) by the Fisher-LSD method for an individual organ (shoot or root).

treated or plants treated with different NO-modulators could be observed. Small differences were also found in root fresh weight (Fig. 6A). As was reported before by Saenen et al. (2013), root fresh

weight of A. thaliana plants treated with 25 mM U was higher than that of controls after 3 days. This increase in fresh weight was less prevalent in L-NAME+U treated plants and absent in SNP+U treated

R. Tewari et al. / Environmental and Experimental Botany 120 (2015) 55–64

plants. Shoot fresh weight on the other hand is adversely affected in both U treated plants as in SNP+U or L-NAME+U plants. However the relative percentage reduction in shoot fresh weight, calculated to the respective controls without U, is less for U+SNP (16.72 %) and U+L-NAME (13.18 %) compared to U-treatment alone (23.26 %). These results concord with the hypothesis for the functioning of NO in metal stress suggested by He et al. (2014) namely, that exogenous NO production leads to an alleviation of the metal toxicity whereas endogenous NO production actually leads to higher toxicity. As second growth parameter leaf surface area was measured on pictures taken at day three after onset of U exposure. U-treated plants were clearly smaller (20.27%) compared to control plants without U. For SNP+U and L-NAME+U plants however leaf surface area was not significantly different from the SNP or L-NAME treated controls, respectively. It is conceivable that alterations induced by NO-modulators lead to changes in transpiration rates. Stomatal closure is a highly regulated process in plants and can be altered by different (a) biotic stressors. A rise in NO-levels in guard cells was indicated as a signalling response downstream of ROS production to induce an upregulation of Ca-fluxes that lead to stomatal closure (for review, Gayatri et al., 2013). However, if the absence of a U-induced NO production in the presence of SNP or L-NAME leads to altered stomatal closure and hence to a different leaf surface/fresh weight ratio, still needs to be further investigated. Taken together based on the results of both fresh weight and leaf surface measurements it can be suggested that lower U-induced NO production in the presence of either SNP or L-NAME results in a lower U toxicity measured as shoot fresh weight as well as leaf surface area and further indicates a role of NO in the U-induced toxic response. The effect of the different NO-modulators on the production of H2O2 in roots and shoots measured as H2O2–Titanium-complexes, was also analysed (Fig. 6D). For roots no significant H2O2 production was induced by U, L-NAME+U or SNP+U treatment. Also in the shoots without U there was no significant induction of H2O2 by either L-NAME or SNP although for both treatments an increasing trend was present. Uranium induced a significant increase in H2O2 levels in the shoots both in the presence and absence of SNP. However, in contrast no additional production of H2O2 was found when plants were simultaneously treated with U and L-NAME. Cross talk between signalling of H2O2 and NO has been studied extensively (for review, Qiao et al., 2014). Hydrogen peroxide production has in a number of cases been shown to be upstream of NO production for example in guard cells leading to stomatal closure (Bright et al., 2006; Gayatri et al., 2013; and for review, Qiao et al., 2014). In contrast in A. thaliana cell suspension cultures exposed to Cd (100–150 mM) it was shown that inhibiting NO production with the NO-synthase inhibitor NG-mono-methylArg monoacetate (L-NMMA) completely prevented the Cd-induced H2O2 production possibly by an upregulation of catalase and ascorbate peroxidase activities (De Michele et al., 2009). The results presented in this study namely the prevention of U-induced H2O2 production after treatment with L-NAME seem to indicate a similar role for NO in U as in Cd toxicity; however, further experimental evidence like testing of antioxidative enzyme activities is needed to corroborate this hypothesis. 4. Conclusion The possible role of NO and H2O2 in U toxicity was studied in 18 day-old A. thaliana seedlings exposed for three days to 25 mM U. It is shown, for the first time, that U induces the production of both NO and H2O2 in roots and in shoots. Effects in shoots were more pronounced despite the fact that most U is retained in the roots. Using NO modulators that induce NO or inhibit the activity of NO

63

synthase it was shown that the endogenous NO produced under U stress seems to enforce U toxicity whereas exogenous supply of NO alleviates the toxicity. These results are similar to observations made for other metal stresses (He et al., 2014). It was further shown that inhibiting NO synthase activity results in a decrease of the U-induced H2O2 production indicating an interplay between NO and H2O2 in the U toxicity response. Taken together, the data support the idea that NO and H2O2 generation is a primary plant response to U stress. As discussed, both H2O2 and NO are unstable molecules and therefore their detection is not always evident. Future experiments using mutants in NO or H2O2 production might further unravel the role of these signalling molecules and their cross talk in the U toxicity in plants. Acknowledgements The authors wish to thank the Belgian Science Policy (BELSPO) for financially supporting the post-doctoral fellowship of R. Tewari. Contributions of co-authors: Conception and design: Rajesh Tewari, Nele Horemans. Analysis and interpretation of the data: Rajesh Tewari, Nele Horemans, May Van Hees. Drafting of the article: Rajesh Tewari, Nele Horemans. Critical revision of the article for important intellectual content: Rajesh Tewari, Nele Horemans, Hildegarde Vandenhove. Final approval of the article: Rajesh Tewari, Nele Horemans, Robin Nauts, Jean Wannijn, May Van Hees. Statistical expertise: Nele Horemans. Obtaining of funding: Nele Horemans, Hildegarde Vandenhove. Technical support: Robin Nauts, Jean Wannijn. Collection and assembly of data: Rajesh Tewari, Robin Nauts, Jean Wannijn. References Arita, N.O., Cohen, M.F., Tokuda, G., Yamasaki, H., 2006. Fluorometric detection of nitric oxide with diaminofluoresceins (DAFs): applications and limitations of plant NO research. In: Lamattina, L., Polacco, J.C. (Eds.), Nitric Oxide in Plant growth, Development and Stress Physiology, Plant Cell Monographs, 6. Springer-Verlag Berlin, Heidelberg, pp. 269–280. Baxter, A., Mittler, R., Suzuki, N., 2014. ROS as key players in plant stress signalling. J. Exp. Bot. 65, 1229–1240. Begara-Morales, J.C., Chaki, M., Sánchez-Calvo, B., Mata-Pérez, C., Leterrier, M., Palma, J.M., Barroso, J.B., Corpas, F.J., 2013. Protein tyrosine nitration in pea roots during development and senescence. J. Exp. Bot. 64, 1121–1134. Besson-Bard, A., Gravot, A., Richaud, P., Auroy, P., Duc, C., Gaymard, F., Taconnat, L., Renou, J.P., Pugin, A., Wendehenne, D., 2009. Nitric oxide contributes to cadmium toxicity in Arabidopsis by promoting cadmium accumulation in roots and by upregulating genes related to iron uptake. Plant Physiol. 149, 1302–1315. Bleise, A., Danesi, P.R., Burkart, W., 2003. Properties, use and health effects of depleted uranium (DU): a general overview. J. Environ. Radioact. 64, 93–112. Brennan, T., Frenkel, C., 1977. Involvement of hydrogen-peroxide in regulation of senescence in pear. Plant Physiol. 59, 411–416. Bright, J., Desikan, R., Hancock, J.T., Weir, I.S., Neill, S.J., 2006. ABA-induced NO generation and stomatal closure in Arabidopsis are dependent on H2O2 synthesis. Plant J. 45, 113–122. Cheeseman, J.M., 2007. Hydrogen peroxide and plant stress: a challenging relationship. Plant Stress 1, 4–15. Corpas, F.J., Barroso, J.B., 2014. Peroxynitrite (ONOO) is endogenously produced in arabidopsis peroxisomes and is overproduced under cadmium stress. Ann. Bot. 113, 87–96. D’Alessandro, S., Posocco, B., Costa, A., Zahariou, G., Lo Schiavo, F., Carbonera, D., Zottini, M., 2013. Limits in the use of cPTIO as nitric oxide scavenger and ERR probe in plant cells and seedlings. Front. Plant Sci. 4, 7. De Michele, R., Vurro, E., Rigo, C., Costa, A., Elviri, L., Di Valentin, M., Careri, M., Zottini, M., di Toppi, L.S., Lo Schiavo, F., 2009. Nitric oxide is involved in cadmium-induced programmed cell death in Arabidopsis suspension cultures. Plant Physiol. 150, 217–228. Egami, F., Taniguchi, S., 1965. Nitrate. In: Bergmeyer, H.U. (Ed.), Methods of Enzymatic Analysis. Elsevier Science, 636. Gayatri, G., Agurla, S., Raghavendra, A.S., 2013. Nitric oxide in guard cells as an important secondary messenger during stomatal closure. Front. Plant Sci. 4, 11. He, H.Y., He, L.F., Gu, M.H., Li, X.F., 2012. Nitric oxide improves aluminum tolerance by regulating hormonal equilibrium in the root apices of rye and wheat. Plant Sci. 183, 123–130.

64

R. Tewari et al. / Environmental and Experimental Botany 120 (2015) 55–64

He, H.Y., He, L.F., Gu, M.H., 2014. The diversity of nitric oxide function in plant responses to metal stress. Biometals 27, 219–228. Hu, W.J., Chen, J., Liu, T.W., Liu, X., Wu, F.H., Wang, W.H., He, J.X., Xiao, Q., Zheng, H.L., 2014. Comparative proteomic analysis on wild type and nitric oxideoverproducing mutant (nox1) of Arabidopsis thaliana. Nitric Oxide 36C, 19–30. Jubany-Mari, T., Munne-Bosch, S., Lopez-Carbonell, M., Alegre, L., 2008. Hydrogen peroxide is involved in the acclimation of the Mediterranean shrub, Cistus albidus L., to summer drought. J. Exp. Bot. 60, 107–120. Juffermans, N.P., Vervloet, M.G., Daemen-Gubbels, C.R., Binnekade, J.M., de Jong, M., Groeneveld, A.B., 2010. A dose-finding study of methylene blue to inhibit nitric oxide actions in the hemodynamics of human septic shock. Nitric Oxide 22, 275–280. Kobayashi, M., Ohura, I., Kawakita, K., Yokota, N., Fujiwara, M., Shimamoto, K., Doke, N., Yoshioka, H., 2007. Calcium-dependent protein kinases regulate the production of reactive oxygen species by potato NADPH oxidase. Plant Cell Online 19, 1065–1080. Kumar, P., Tewari, R. Kumar, Nand Sharma, P., 2007. Excess nickel-induced changes in antioxidative processes in maize leaves. J. Plant Nutrit. Soil Sci. 170, 796–802. Kumar, P., Tewari, R.K., Sharma, P.N., 2008a. Cadmium Enhances generation of hydrogen peroxide and amplifies activities of catalase, peroxidases and superoxide dismutase in maize. J. Agron. Crop Sci. 194, 72–80. Kumar, P., Tewari, R.K., Sharma, P.N., 2008b. Modulation of copper toxicity-induced oxidative damage by excess supply of iron in maize plants. Plant Cell Rep. 27, 399–409. Kumar, P., Tewari, R.K., Sharma, P.N., 2010. Sodium nitroprusside-mediated alleviation of iron deficiency and modulation of antioxidant responses in maize plants. AoB Plants 2010, plq002. Li, H., Song, J.B., Zhao, W.T., Yang, Z.M., 2013. AtHO1 is involved in iron homeostasis in an NO-dependent manner. Plant Cell Physiol. 54, 1105–1117. Miller, G., Schlauch, K., Tam, R., Cortes, D., Torres, M.A., Shulaev, V., Dangl, J.L., Mittler, R., 2009. The plant NADPH oxidase RBOHD mediates rapid systemic signaling in response to diverse stimuli. Sci. Signal. 2, ra45. Mittler, R., Vanderauwera, S., Suzuki, N., Miller, G., Tognetti, V.B., Vandepoele, K., Gollery, M., Shulaev, V., Van Breusegem, F., 2011. ROS signaling: the new wave? Trends Plant Sci. 16, 300–309. Munoz, I.G., Moran, J.F., Becana, M., Montoya, G., 2005. The crystal structure of an eukaryotic iron superoxide dismutase suggests intersubunit cooperation during catalysis. Protein Sci. 14, 387–394. Murgia, I., Concetta de Pinto, M., Delledonne, M., Soave, C., De Gara, L., 2004. Comparative effects of various nitric oxide donors on ferritin regulation, programmed cell death, and cell redox state in plant cells. J. Plant Physiol. 161, 777–783. Oke, O.L., 1966. Nitrite toxicity to plants. Nature 212, 528. Pluth, M.D., Tomat, E., Lippard, S.J., 2011. Biochemistry of mobile zinc and nitric oxide revealed by fluorescent sensors. Annu. Rev. Biochem. 80, 333–355. Qiao, W.H., Li, C.N., Fan, L.M., 2014. Cross-talk between nitric oxide and hydrogen peroxide in plant responses to abiotic stresses. Environ. Exp. Bot. 100, 84–93. Rumer, S., Krischke, M., Fekete, A., Mueller, M.J., Kaiser, W.M., 2012. DAFfluorescence without NO: elicitor treated tobacco cells produce fluorescing DAF-derivatives not related to DAF-2 triazol. Nitric Oxide 27, 123–135. Saenen, E., Horemans, N., Vanhoudt, N., Vandenhove, H., Biermans, G., Van Hees, M., Wannijn, J., Vangronsveld, J., Cuypers, A., 2013. Effects of pH on uranium uptake and oxidative stress responses induced in Arabidopsis thaliana. Environ. Toxicol. Chem. 32, 2125–2133. Sheppard, S.C., Sheppard, M.I., Gallerand, M.O., Sanipelli, B., 2005. Derivation of ecotoxicity thresholds for uranium. J. Environ. Radioact. 79, 55–83. Shin, R., Schachtman, D.P., 2004. Hydrogen peroxide mediates plant root cell response to nutrient deprivation. Proc. Natl. Acad. Sci. 101, 8827–8832. Singh, H.P., Batish, D.R., Kaur, G., Arora, K., Kohli, R.K., 2008. Nitric oxide (as sodium nitroprusside) supplementation ameliorates Cd toxicity in hydroponically grown wheat roots. Environ. Exp. Bot. 63, 158–167. Srivastava, S., Bhainsa, K.C., D’Souza, S.F., 2010. Investigation of uranium accumulation potential and biochemical responses of an aquatic weed Hydrilla verticillata (L. f.) Royle. Bioresour. Technol. 101, 2573–2579. Tewari, R.K., Paek, K.-Y., 2011. Salicylic acid-induced nitric oxide and ROS generation stimulate ginsenoside accumulation in Panax ginseng roots. J. Plant Growth Regul. 30, 396–404.

Tewari, R.K., Kumar, P., Tewari, N., Srivastava, S., Sharma, P.N., 2004. Macronutrient deficiencies and differential antioxidant responses—influence on the activity and expression of superoxide dismutase in maize. Plant Sci. 166, 687–694. Tewari, R.K., Kumar, P., Sharma, P.N., 2006. Antioxidant responses to enhanced generation of superoxide anion radical and hydrogen peroxide in the copperstressed mulberry plants. Planta 223, 1145–1153. Tewari, R.K., Hahn, E.-J., Paek, K.-Y., 2008a. Modulation of copper toxicity-induced oxidative damage by nitric oxide supply in the adventitious roots of Panax ginseng. Plant Cell Rep. 27, 171–181. Tewari, R.K., Kumar, P., Sharma, P.N., 2008b. Morphology and physiology of zincstressed mulberry plants. J. Plant Nutrit. Soil Sci. 171, 286–294. Tewari, R.K., Hadacek, F., Sassmann, S., Lang, I., 2013a. Iron deprivation-induced reactive oxygen species generation leads to non-autolytic PCD in Brassica napus leaves. Environ. Exp. Bot. 91, 74–83. Tewari, R.K., Prommer, J., Watanabe, M., 2013b. Endogenous nitric oxide generation in protoplast chloroplasts. Plant Cell Rep. 32, 31–44. Tian, Q.-Y., Sun, D.-H., Zhao, M.-G., Zhang, W.-H., 2007. Inhibition of nitric oxide synthase (NOS) underlies aluminum-induced inhibition of root elongation in Hibiscus moscheutos. New Phytol. 174, 322–331. Vandana, S., Sustmann, R., Rauen, U., Stohr, C., 2012. FNOCT as a fluorescent probe for in vivo localization of nitric oxide distribution in tobacco roots. Plant Physiol. Biochem. 59, 80–89. Vandenhove, H., Cuypers, A., Van Hees, M., Koppen, G., Wannijn, J., 2006. Oxidative stress reactions induced in beans (Phaseolus vulgaris) following exposure to uranium. Plant Physiol. Biochem. 44, 795–805. Vanhoudt, N., Vandenhove, H., Smeets, K., Remans, T., Van Hees, M., Wannijn, J., Vangronsveld, J., Cuypers, A., 2008. Effects of uranium and phosphate concentrations on oxidative stress related responses induced in Arabidopsis thaliana. Plant Physiol. Biochem. 46, 987–996. Vanhoudt, N., Cuypers, A., Horemans, N., Remans, T., Opdenakker, K., Smeets, K., Martinez Bello, D., Havaux, M., Wannijn, J., Van Hees, M., Vangronsveld, J., Vandenhove, H., 2011a. Unraveling uranium-induced oxidative stress related responses in Arabidopsis thaliana seedlings. Part II: responses in the leaves and general conclusions. J. Environ. Radioact. 102, 638–645. Vanhoudt, N., Vandenhove, H., Horemans, N., Bello, D.M., Van Hees, M., Wannijn, J., Carleer, R., Vangronsveld, J., Cuypers, A., 2011b. Uranium induced effects on development and mineral nutrition of Arabidopsis thaliana. J. Plant Nutr. 34, 1940–1956. Vanhoudt, N., Vandenhove, H., Horemans, N., Remans, T., Opdenakker, K., Smeets, K., Bello, D.M., Wannijn, J., Van Hees, M., Vangronsveld, J., Cuypers, A., 2011c. Unraveling uranium induced oxidative stress related responses in Arabidopsis thaliana seedlings. Part I: responses in the roots. J. Environ. Radioact. 102, 630–637. Vanhoudt, N., Horemans, N., Biermans, G., Saenen, E., Wannijn, J., Nauts, R., Van Hees, M., Vandenhove, H., 2014. Uranium affects photosynthetic parameters in Arabidopsis thaliana. Environ. Exp. Bot. 97, 22–29. Vergara, R., Parada, F., Rubio, S., Perez, F.J., 2012. Hypoxia induces H2O2 production and activates antioxidant defence system in grapevine buds through mediation of H2O2 and ethylene. J. Exp. Bot. 63, 4123–4131. Vitecek, J., Reinohl, V., Jones, R.L., 2008. Measuring NO production by plant tissues and suspension cultured cells. Mol. Plant 1, 270–284. Wang, L., Yang, L., Yang, F., Li, X., Song, Y., Wang, X., Hu, X., 2010. Involvements of H2O2 and metallothionein in NO-mediated tomato tolerance to copper toxicity. J. Plant Physiol. 167, 1298–1306. Wei, L., Derrien, B., Gautier, A., Houille-Vernes, L., Boulouis, A., Saint-Marcoux, D., Malnoe, A., Rappaport, F., de Vitry, C., Vallon, O., Choquet, Y., Wollman, F.A., 2014. Nitric oxide-triggered remodeling of chloroplast bioenergetics and thylakoid proteins upon nitrogen starvation in Chlamydomonas reinhardtii. Plant Cell 26, 353–372. Xu, J., Wang, W., Yin, H., Liu, X., Sun, H., Mi, Q., 2009. Exogenous nitric oxide improves antioxidative capacity and reduces auxin degradation in roots of Medicago truncatula seedlings under cadmium stress. Plant Soil 326, 321–330. Xu, J., Yin, H., Li, Y., Liu, X., 2010. Nitric oxide is associated with long-term zinc tolerance in Solanum nigrum. Plant Physiol. 154, 1319–1334. Zhou, B., Guo, Z., Xing, J., Huang, B., 2005. Nitric oxide is involved in abscisic acidinduced antioxidant activities in Stylosanthes guianensis. J. Exp. Bot. 56, 3223–3228.