Improving elms performance under drought stress: The pretreatment with abscisic acid

Environmental and Experimental Botany 100 (2014) 64–73

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Improving elms performance under drought stress: The pretreatment with abscisic acid Maria Celeste Dias ∗ , Helena Oliveira, Armando Costa, Conceic¸ão Santos Laboratory of Biotechnology & Cytometry, Department of Biology & Centre for Environmental and Marine Studies (CESAM), University of Aveiro, 3810-193, Aveiro, Portugal

a r t i c l e

i n f o

Article history: Received 3 September 2013 Received in revised form 11 December 2013 Accepted 21 December 2013 Keywords: Antioxidant activity Flow cytometry Photosynthesis Transpiration Phytohormones Proline

a b s t r a c t Hormonal conditioning of plants in order to increase photosynthetic performance and reduce oxidative stress may improve plants’ tolerance to stress. This study aims to elucidate the effects of ABA pretreatment on the photosynthetic apparatus and antioxidant battery of Ulmus minor plants under well watered (WW) and drought stress (DS) conditions. Leaves were sprayed with ABA (50 and 100 ␮M). After 25 days of treatment DS was initiated by withholding water for 6 days. Water deficit decreased the RWC, induced stomatal closure and impaired net CO2 assimilation rate (A). However, independently of the water regime, ABA pretreatment increased plant DW accumulation, A, carotenoids and Chl a contents and reduced water loss. DS induced oxidative stress, but ABA application increased DS tolerance by the enhancement of the antioxidant system. Under WW conditions, the benefits of ABA application in reducing the cell membrane damages were noticeable. ABA pretreatment and DS induced changes in U. minor cell cycle of leaf cells, with a delay in S phase and an increase of FPCV coefficient. We propose that ABA pretreatment improves plant performance by increasing plant DW accumulation and augmenting the antioxidant system of U. minor plants, not only under DS conditions, but also under WW conditions. The use of ABA as pretreatment to alleviate the negative effects of DS seems to be a promising strategy to reduce plant’s water loss and improve plant productivity in drought prone habitats. © 2013 Elsevier B.V. All rights reserved.

1. Introduction Water deficit is one of the main environmental stress factors that negatively affect plant growth and final yield performance of a crop. Due to the global climate changes, world water scarcity is increasing. It is estimated that up to 45% of the world agricultural lands are already exposed to continuous or frequent drought (Ashraf and Foolad, 2007; IPCC, 2007). Unfortunately, according to the climate changes predictions this scenario is expected to increase by the end of this century (IPCC, 2007). Hence, the knowledge of

Abbreviations: A, net CO2 assimilation rate; ABA, abscisic acid; APX, ascorbate peroxidase; C, group of plants under WW condition; C50, group of plants treated with 50 ␮M ABA under WW condition; C100, group of plants treated with 100 ␮M ABA under WW condition; CAT, catalase; Ci , intercellular CO2 concentration; Chl, chlorophyll; DS, drought stress; DW, dry weight; E, transpiration rate; F0 , minimum Chl fluorescence; FCM, flow cytometry; Fm , maximum Chl fluorescence; Fv , variable fluorescence; Fv /Fm , maximum quantum yield of PSII; gs , stomatal conductance; JA, jasmonic acid; JA-Ile, jasmonyl-isoleucine; PI, propidium iodide; MDA, malondialdehyde; RWC, relative water content; S, group of plants under DS condition; S50, group of plants treated with 50 ␮M ABA under DS condition; S100, group of plants treated with 100 ␮M ABA under DS condition; SA, salicylic acid; SOD, superoxide dismutase; WW, well watered. ∗ Corresponding author. Tel.: +351 234 370 200; fax: +351 234 370 985. E-mail address: [email protected] (M.C. Dias). 0098-8472/$ – see front matter © 2013 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.envexpbot.2013.12.013

physiological and biochemical mechanisms that are negatively affected by drought stress (DS) as well as the plant’s strategies to mitigate those effects deserve considerable attention (e.g., Slama et al., 2007). In plants, low water availability gives rise to several physiological and biochemical responses. The loss of turgor and osmotic adjustment decrease leaf water potential and induce stomatal closure. Consequently, the limitation of gas exchange reduces transpiration and photosynthesis, ultimately limiting plant growth and development (Shao et al., 2008). As the key process of primary metabolism, photosynthesis plays a key role in plant performance under DS conditions (e.g., Brito et al., 2003; Sperdouli and Moustakas, 2012). When net photosynthesis decreases, the excess of excitation energy in the photosystem II (PSII) leads to an impairment of photosynthetic function and to an accumulation of reactive oxygen species (ROS) causing oxidative damages in plants (Wilhelm and Selmar, 2011). ROS, such as O2 •− , H2 O2 and OH radicals, can directly attack the membrane lipids, inactivate enzymes and damage the nucleic acids leading, in many cases, to cell death (Azevedo et al., 2005; Monteiro et al., 2012). To cope with oxidative stress, plant cells developed a highly efficient defense system, with both antioxidant enzymes (e.g., SOD, CAT) and antioxidant metabolites (e.g., ascorbate) that can neutralize free radicals and reduce the potential damages of ROS. The accumulation of some organic

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compatible compounds, such as l-proline, plays a significant role in plant protection and adaptation to a broad range of stress (Ashraf and Foolad, 2007; Dias et al., 2013a). Besides its important role as an osmolyte for osmotic adjustment, proline also contributes to the detoxification of ROS, protection of membrane integrity and stabilization of enzymes/proteins (Ashraf and Foolad, 2007). Several phytohormones, such as abscisic acid (ABA), salicylic acid (SA), jasmonic acid (JA) and jasmonyl-isoleucine (JA-Ile, the most biologically active form of jasmonates) (Fonseca et al., 2009), are pivotal for plant growth and development, but also play an important role in integrating several stress signals and controlling downstream stress responses (Corcuera et al., 2012). The phytohormone ABA is well known as one of the main internal plant signals that play a critical role in the regulation of plant responses from the whole plant level to the cellular level (Efetova et al., 2007). ABA regulates the plant’s adaptive response to a wide range of environmental stresses, such as DS, via diverse physiological and developmental processes. Under DS conditions, ABA primarily promotes stomatal closure to minimize water loss by transpiration and then mitigates stress damage through the activation of many stress-responsive genes that collectively increase plant stress tolerance (Aroca et al., 2008). In particular, this phytohormone interacts with membrane phospholipids to stabilize the membranes under stress conditions and enhances plant tolerance to oxidative stress by increasing the activity of antioxidant enzymes (Guschina et al., 2002). The stress-related responses of plants by ABA usually occur before any change of plant water status during soil drying and are considered the first line of defense as soil water deficits starts (Liu et al., 2005). The effects of ABA on cell cycle have been studied mostly in roots and in in vitro cells, supporting that ABA inhibits DNA replication and cell division, which results in retarded plant growth. Cell cycle changes were described in e.g., Pisum sativum, Nicotiana tabacum or Zea mays after ABA treatment (Levi et al., 1993; Muller et al., 1994; Swiatek et al., 2002). ABA has been shown to be involved in promoting drought tolerance through the enhancement of the antioxidant system by exogenous application in several species (e.g., Duan et al., 2007; Wang et al., 2011). However, at the photosynthetic level little is known, and the available data demonstrates that A values increased in Phaseolus vulgaris, Beta vulgaris, N. tabacum and Z. mays plants previously treated with ABA (before the onset of DS) (Pospíˇsilová and Batková, 2004). Trouverie et al. (2003), Ma et al. (2008), Li et al. (2004), Yin et al. (2004), Duan et al. (2007) and Wang et al. (2011) also studied the effects of exogenous ABA application but, in their studies, the onset of the ABA treatment coincided with the DS treatment. These authors found that the simultaneous exposure to DS + ABA negatively affected plant height, biomass accumulation and A in Malus domestica, Populus davidiana, Populus kangdingensis, Populus cathayana, Picea asperata and Actinidia deliciosa. This raised a question if, instead of simultaneous treatment, a preconditioning of plants with ABA might improve their tolerance to DS. Elms (Ulmus sp.) are widely used as ornamental and as timber source. Among other ornamental woody species, elms present a moderate tolerance to DS, for example, being considered to be more drought tolerant than willows or cottonwoods. Within this genus, the U. minor species is an elegant model in studying biotic and abiotic stresses (e.g., Oliveira et al., 2009; Conde et al., 2008; Dias et al., 2011, 2013b). Moreover, Solla and Gil (2002) described that DS could influence the development of Dutch Elm Disease symptoms in this species, with consequences on elm resistance and breeding (Desprez-Loustau et al., 2007). Therefore, ABA pretreatment to prevent/reduce plant loss under DS conditions could be a promising strategy that deserves to be further investigated.

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We propose here that ABA must be applied prior the stress imposition (pretreatment), and not during the stress, in order to allow the plant to trigger defense mechanisms necessary to deal with stress. We also hypothesize that some of these key mechanisms involve improved photosynthesis and oxidative protection. We tested this hypothesis of ABA-pretreatment, using elms and we analyzed several parameters: plant growth (DW accumulation), RWC, photosynthesis, pigments, H2 O2 , phytohormones, proline, antioxidant enzymes, lipid peroxidation and cell membrane stability. Considering the contribution of cell division to plant growth we also analyzed the effects of ABA on cell cycle and on ploidy levels. We demonstrate here that ABA pretreatment improved plant’s photosynthesis and antioxidant system, and ultimately improved plant growth. 2. Material and methods 2.1. Plant material and experimental conditions U. minor Mill. seedlings were grown in 500 cm3 pots containing an autoclaved mixture of peat and perlite (3:2, v/v) in a controlled climate chamber (Phytotron, Snijders, Tilburg) with a photoperiod of 16-h, a temperature of 22 ± 2 ◦ C and a photosynthetic photon flux density of 200 ± 10 ␮mol m−2 s−1 . Two month old plants with average values of 48.4 ± 6.5 g DW (corresponding to plants with 10.0 ± 2.1 cm of stem height with 7–9 developed leaves) were randomly separated in three groups: in Group 1 – each plant was sprayed with 50 ml of distillated water; in Group 2 – each plant was sprayed with 50 ml of 50 ␮M ABA; and in Group 3 – each plant was sprayed with 50 ml of 100 ␮M ABA. ABA application was performed twice a week during 4 weeks. After 25 days of ABA treatment, two water regimes were employed in the three groups: well watered (WW) and drought stress (DS). In the WW treatment the pots were watered at 100% field capacity (FC) by replacing the amount of water transpired every second day (water loss was measured by weighing the pots). In the DS treatment, water was withholding for 6 days. For the water regime experiments, each Group (1, 2 and 3) was separated in two sub-groups with 20 plants each. The plants of the Group 1 were randomly separated in two sub-groups, C and S (C: plants previously sprayed with water under WW condition; and S: plants previously sprayed with water under DS condition); the plants of Group 2 were randomly separated in two sub-groups, C50 and S50 (C50: plants previously sprayed with 50 ␮M ABA under WW condition; and S50: plants previously sprayed with 50 ␮M ABA under DS condition); and the plants of Group 3 were randomly separated in two sub-groups, C100 and S100 (C100: plants previously sprayed with 100 ␮M ABA under WW condition; and S100: plants previously sprayed with 100 ␮M ABA under DS condition). After 6 days under WW and DS conditions, the RWC, gas-exchange parameters, chlorophyll a fluorescence, cytometry analysis, Rubisco activity, cell membrane permeability were analyzed. At the same time, leaf samples were collected, frozen immediately in liquid nitrogen and stored at −80 ◦ C for further quantification of pigments, phytohormones, proline, antioxidant enzymes, H2 O2 content and MDA. 2.2. Plant water status and plant growth Plant water status was assessed through the determination of the RWC of leaf discs. RWC was calculated as 100 × (FW − DW)/(TW − DW), where FW is the fresh weight of leaf discs, TW is their turgid weight (determined after floating for 180 min leaf discs on water at 5 ◦ C) and DW is the dry weight (determined after drying the leaf discs at 80 ◦ C for 48 h). Total plant DW

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and FW were measured at the end of the DS period. DW was determined after drying the samples in an oven at 80 ◦ C till constant weight. 2.3. Phytohormone analyses and proline content determination Hormone extraction and analysis were carried out as described in Durgbanshi et al. (2005). Briefly, leaf tissue (0.4 g) was extracted in ultrapure water using a tissue homogenizer. Before extraction, the samples were spiked with deuterated standards of every compound. After centrifugation, the pH of the supernatant was adjusted to 3.0 with 30% acetic acid and partitioned twice against diethyl ether. The organic layer was recovered and evaporated under vacuum in a centrifuge concentrated. The dry residue was then resuspended in a 10% MeOH solution by gentle sonication. The resulting solution was filtered and injected in an HPLC system. Hormones were separated in a reversed-phase C18 column using a linear gradient of MeOH and water supplemented with 0.01% acetic acid at a flow rate of 300 ␮l min−1 . The mass spectrometer, a Quattro LC triple quadrupole, was operated in a negative ionization electrospray mode, and the hormones were detected according to their specific transitions using a multiresidue mass spectrometric method. Free proline was extracted and determined as described by Hamid et al. (2003). Leaf samples were extracted with 3% sulphosalicylic acid. Extracts were maintained for 1 h at 100 ◦ C with the addition of 2 ml glacial acetic acid and 2 ml acid ninhydrin. Cold toluene was added afterwards and shaken. Absorbance was read at 520 nm. The amount of proline was determined from a standard curve. 2.4. Quantification of chlorophylls and carotenoids Pigments were extracted from the leaf disc in a cold acetone/50 mM Tris buffer (pH 7.8; 80:20, v/v) and centrifuged at 2800 × g for 5 min as described by Sims and Gamon (2002). Absorbance at 470, 537, 647 and 663 nm was determined with a Thermo Fisher Scientific (Waltham, USA) spectrophotometer (Genesys 10-uv S). The contents of chlorophyll (Chl) a, Chl b and carotenoids were calculated according to Lichtenthaler (1987). 2.5. Chlorophyll a fluorescence, gas exchange parameters and Rubisco activity Chl a fluorescence parameters were measured on the adaxial side of the leaf using a pulse amplitude modulation system (MiniPam, Walz). Minimum Chl fluorescence (F0 ) was measured in 30 min dark-adapted leaves by applying a weak modulated light, and maximum Chl fluorescence (Fm ) was measured after applying a 0.7 s saturating pulse of white light (>1500 ␮mol m−2 s−1 ) to the same leaves. The maximum quantum yield of photosystem II (PSII) was calculated as Fv /Fm = (Fm − F0 )/Fm as described by Van Kooten and Snel (1990). In situ leaf gas exchange measurements (net CO2 assimilation rate: A, transpiration rate: E, and the intercellular CO2 concentration: Ci ) were performed using a portable infrared gas analyser (LCpro+, ADC, Hoddesdon, UK), operating in open mode under growth chamber conditions. The stomatal conductance (gs ) was automatically calculated according to Von Caemmerer and Farquhar (1981). Measurements were always performed in the middle of the photoperiod at growth temperature (24 ± 2 ◦ C) and atmospheric CO2 concentration in the youngest fully developed leaf. For Rubisco (EC 4.1.1.39) activity, leaf samples were ground to a powder in a mortar with liquid nitrogen as described by Dias and

Brüggemann (2010). Rubisco maximal activity was measured as described by Lilley and Walker (1974). This assay followed NADPH oxidation measured spectrophotometrically at 340 nm. Total activity was achieved after incubation in 20 mM MgCl2 and 10 mM NaHCO3 for 20 min. 2.6. Flow cytometry analysis U. minor leaves were used as source material for ploidy and cell cycle analysis. Nuclear suspensions were obtained from approximately 30 mg of plant material according to the protocol previously described by Loureiro et al. (2007a). In brief, nuclei were released from cells by chopping with a razor blade in 1 ml of Woody Plant Buffer (WPB) (0.2 M Tris·HCl, 4 mM MgCl2 ·6H2 O, 2 mM EDTA Na2 ·2H2 O, 86 mM NaCl, 10 mM sodium metabisulfite, 1% PVP-10, 1% (v/v) Triton X-100, pH 7.5). For ploidy analysis U. minor leaves were chopped together with the same amount of P. sativum cv. Ctirad (cv = 9.09 pg DNA; kindly provided by J. Dolezel, Laboratory of Molecular Cytogenetics and Cytometry, Institute of Experimental Botany, Olomouc, Czech Republic) leaves used as internal reference standard (Loureiro et al., 2007b). To minimize release of cytosolic compounds, chopping was quick (less than 30 s) and not very intense. Also, in order to reduce suspension viscosity, samples were incubated for 2 min before filtration, as described by Loureiro et al. (2007b). Nuclear suspension was filtered with a 50 ␮m nylon mesh. After that, nuclei were stained with 50 ␮g/ml propidium iodide (PI, Fluka) and to avoid PI staining of RNA, 50 ␮g/ml of RNAse (Sigma, St. Louis, USA) was also added to nuclei suspension. After 5 min of incubation, nuclei were analyzed in a Beckman Coulter EPICS XL (Beckman Coulter, Hialeah, FL, USA) flow cytometer. The instrument was equipped with an air-cooled argon-ion laser tuned at 15 mW and operating at 488 nm. Integral fluorescence together with fluorescence pulse height and width emitted from nuclei was collected through a 645 dichroic long-pass filter and a 620 band-pass filter and converted on 1024 ADC channels. Prior to analysis, the instrument was checked for linearity with fluorescent beads (Coulter Electronics), and the amplification settings were kept constant throughout the experiment. Data was acquired using the SYSTEM II software (version 3.0, Beckman Coulter). The results were obtained in the form of three graphics: linear fluorescence light intensity (FL), forward angle light scatter (FS) versus side angle light scatter (SS) and FL pulse integral versus FL pulse height. The latter was used to eliminate partial nuclei and other debris, nuclei with associated cytoplasm and doublets (these events have a higher pulse area but the same pulse height as single nuclei). In each replicate at least 5000 nuclei were analyzed. The nuclear DNA content was calculated according to the formula: 2C nuclear DNA content of sample (pg) =

Sample G0/G1 mean FL Reference standard G0/G1 mean FL

× 2C nuclear DNA content of reference standard

Conversion of mass values into base-pair numbers was done according to the factor 1 pg = 978 Mbp (Doleˇzel et al., 2003). For cell cycle analysis, the proportion of cells in each phase, the full peak coefficient of variation (FCPV) and the half- peak coefficient of variation (HPCV) of the G0/G1 peak were collected from the fluorescence intensity (FL) histograms (5 plants per condition). Raw data were exported as Static Dimension data files LMD files (Listmode File Format) and analyzed using FlowJo (Tree Star Inc., Ashland, OR, USA).

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2.7. Cell membrane permeability and lipid peroxidation

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contained 0.1 M potassium phosphate buffer (pH 7.0) and enzyme extract. To start the reaction, 20 mM H2 O2 (ε = 39.4 mM−1 cm−1 ) was added and the decrease of absorbance at 240 nm was recorded. APX activity was determined at 25 ◦ C by recording the decrease in absorbance at 290 nm due to ascorbic acid (ε = 2.8 mM−1 cm−1 ) oxidation to dehydroascorbate by H2 O2 , according to the method of Nakano and Asada (1981). The reaction mixture contained 100 mM potassium phosphate buffer (pH 7.0), 0.5 mM ascorbic acid, and an aliquot of enzyme extract, and 0.5 mM H2 O2 was added to start the reaction. One unit of APX activity was expressed as mmol of ascorbic acid oxidized per minute, using its extinction coefficient (ε) 2.8 mM−1 cm−1 . H2 O2 concentration was measured according to Zhou et al. (2006). Briefly, leaves (0.5 g) were ground in 5 ml precooled 5% TCA (w/v) and activated charcoal. After centrifugation, extracts were adjusted to pH 8.4 with 17 M ammonia, and H2 O2 was spectrophotometrically quantified following its reaction with 4-aminoantipyrine and phenol to form a stable red product in the presence of 150 U mg−1 peroxidase. Blanks containing 8 ␮g CAT were run for each sample as well as for the calibration with H2 O2 standards, which were added to the extraction medium in parallel to the samples.

Electrolyte leakage was used to assess cell membrane permeability as described by Silva et al. (2013). Leaf segments were detached, washed with deionized water, placed in closed vials containing 20 ml of de-ionized water and incubated over night at 25 ◦ C, on a rotary shaker. Electrical conductivity of the bathing solution (Lt ) was determined after 24 h. Samples were then autoclaved at 120 ◦ C for 20 min and a last conductivity reading (L0 ) was obtained upon equilibration at 25 ◦ C. The electrolyte leakage was defined as Lt /L0 and expressed as percentage. Lipid peroxidation on leaves was obtained by measuring malondialdehyde (MDA) production. Approximately 0.5 g of leaves were homogenized with 5 ml of 0.1% TCA (w/v) and centrifuged at 1000 × g for 10 min at 4 ◦ C (Santos et al., 2004). After centrifugation, 1 ml of supernatant was mixed with 4 ml 20% TCA (w/v) in 0.5% TBA (w/v) and incubated for 30 min at 95 ◦ C. The extract was then cooled immediately on ice to stop the reaction and centrifuged at 1000 × g for 10 min at 4 ◦ C. MDA concentration was estimated by subtracting the nonspecific absorption at 600 nm from the absorption at 532 nm using an absorbance coefficient of extinction (ε), 155 mM−1 cm−1 .

2.9. Statistical analysis 2.8. Antioxidant enzyme activity and H2 O2 content All the determinations were obtained with randomly chosen plants and the experiments were conducted with 20 plants per treatment. Data were analyzed by a one-way analysis of variance (ANOVA) using the Sigma Stat program (Windows, version 3.1). Comparisons between means were evaluated by Post Hoc Test (Tukey’s multiple comparison test) at a significant level set to 0.05.

For the determination of antioxidant enzyme activities, leaves (0.5 g) were homogenized in 5 ml of extraction buffer in a prechilled mortar and pestle by liquid nitrogen as described by Silva et al. (2013). The extraction buffer contained 0.1 M potassium phosphate buffer (pH 7.5), 0.5 mM Na2 EDTA, 1% PVP (w/v), PMSF 1 mM, 0.2% Triton X-100 (v/v) and 2 mM DTT. The homogenate was filtered through four layers of cheesecloth and centrifuged at 8 000 × g for 20 min at 4 ◦ C. The supernatant was used for enzyme assays: ascorbate peroxidase (APX, EC 1.11.1.11), catalase (CAT, EC 1.11.1.6) and superoxide dismutase (SOD, EC 1.15.1.1) and total soluble protein quantification. Protein concentration was determined according to the method of Bradford (1976) using the Total Protein Kit, Micro (Sigma, Germany). SOD activity was assayed at 25 ◦ C by monitoring the decrease of absorbance at 560 nm generated by the inhibition of the reduction of NBT (nitroblue tetrazolium chloride) following the method of Agarwal et al. (2005). The reaction mixture contained 13.3 mM methionine, 63 ␮M NBT, 0.1 mM EDTA, 50 mM potassium phosphate buffer (pH 7.8), 50 mM Na2 CO3 , and an aliquot of extract. Reaction was started by adding 2 ␮M riboflavin and placing the tubes under a 15 W fluorescent lamp for 15 min. A reaction mixture without the extract was used for control. To stop the reaction, the light was switched off and the tubes were maintained in the dark. For enzymatic and control assays a non-irradiated correspondent reaction mixture, with or without extract respectively, was used as blank. One unit of SOD activity was defined as the amount of enzyme required to cause 50% inhibition of the reduction of NBT in comparison with control. CAT activity was assayed at 25 ◦ C as described by Beers and Sizer (1952). Assay mixture

3. Results 3.1. RWC and total plant DW The maximum decrease of RWC (P < 0.05) was found in plants not treated with ABA under DS conditions (18% decrease in plants of S treatment compared with C treatment). The decrease of RWC in DS plants of treatments S50 and S100 was only 10%, as compared to the respective WW control plants (C50 and C100) (Table 1). Independently of the water status condition (WW and DS), plants not treated with ABA showed a lower total DW (P < 0.05) compared to those treated with ABA (Table 1). No significant differences were observed in the total plant DW of ABA treated plants under WW and DS conditions. 3.2. Plant phytohormone content and proline content The level of ABA in plants previously treated with ABA was significantly higher than that of plants not treated with ABA (Fig. 1A). Moreover, the application of the highest ABA concentration (100 ␮M) resulted in a higher accumulation of this hormone in leaves (P < 0.05). In plants not treated with ABA, plants under DS (S

Table 1 Relative water content (RWC), total plant DW, pigment content, Fv /Fm , Rubisco activity, and proline content in WW (C, C50 and C100) and DS (S, S50 and S100) U. minor plants. Values are means ± SD (n = 10). Different letters indicate significant differences between treatments at a significant level equal to 0.05. Parameters RWC (%) Plant DW (g DW) Chl a (mg g−1 FW) Chl b (mg g−1 FW) Carotenoids (mg g−1 FW) Fv /Fm Rubisco (␮mol m−2 s−1 ) Proline (nmol g−1 FW)

C 92.1 60.0 4.1 2.1 1.0 0.84 11.9 32.0

S ± ± ± ± ± ± ± ±

a

1.1 8.0a 0.14a 0.18a 0.09a 0.04a 2.4a 6.0a

75.1 53.0 3.6 2.1 1.2 0.84 5.5 66.0

C50 ± ± ± ± ± ± ± ±

b

2.9 10.0a 0.23b 0.29a 0.09b 0.02a 1.3b 6.0b

91.8 86.0 5.2 2.1 2.4 0.84 16.8 34.0

S50 ± ± ± ± ± ± ± ±

a

1.2 3.0b 0.77c 0.18a 0.14c 0.03a 2.5c 4.0a

82.3 83.0 4.5 2.0 2.5 0.87 10.3 96.0

C100 ± ± ± ± ± ± ± ±

c

1.1 9.0b 0.27c 0.11a 0.41c 0.04a 2.8a 10.0c

91.3 85.0 5.2 2.4 2.2 0.85 15.3 40.0

S100 ± ± ± ± ± ± ± ±

a

0.7 5.0b 0.40c 0.24a 0.54c 0.2a 1.9c 6.1a

81.9 81.0 4.9 2.4 2.6 0.84 8.4 130.0

± ± ± ± ± ± ± ±

1.0c 8.0b 0.43c 0.41a 0.25c 0.05a 1.5a 16.4d

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Fig. 1. Contents of ABA (A); SA (B); JA (C) and JA-Ile (D) in WW (C, C50 and C100) and DS (S, S50 and S100) U. minor plants. Values are means ± SD (n = 6). Different letters indicate significant differences between treatments at a significant level equal to 0.05.

treatment) showed a significantly higher ABA content than plants under WW conditions (C treatment), contrarily to other conditions where the DS did not affect the level of ABA with respect to the corresponding controls. No significant differences were observed in the level of SA (Fig. 1B). DS plants showed the highest levels of JA (P < 0.05), which was not influenced by the pretreatment with ABA (Fig. 1C). Plants under DS conditions of treatments S and S50 showed a significant higher level of JA-Ile than plants under WW conditions from treatments C, C50 and C100 (Fig. 1D). No significant differences were observed in proline content in plants under WW condition (C, C50 and C100) (Table 1). Under DS conditions, plants previously treated with 50 ␮M ABA (S50) showed the highest proline content while those not treated with ABA (S) showed the lowest content of this amino acid (P < 0.05).

significantly in DS plants of treatments S, S50 and S100 respectively to WW plants of treatments C, C50 and C100 (Fig. 2B and C). Plants treated with ABA (C50, C100, S50 and S100) showed a significantly lower E and gs compared to the respective controls without ABA treatment (C and S). Plants from the C treatment (WW conditions) showed the highest Ci , while the lowest Ci was observed in S50 and S100 treatments (DS conditions) (P < 0.05) (Fig. 2D). No significant differences in the Ci were observed between the treatments S, C50 and C100. Plants treated with ABA under WW conditions (C50 and C100) showed the highest Rubisco activities (P < 0.05) (Table 1). The activity of Rubisco was significantly lower in plants under DS conditions (S, S50 and S100) when compared to the respective control under WW conditions (C, C50 and C100). The activities of Rubisco in the DS plants of treatments S50 and S100 were similar to C treatment of the WW conditions (P > 0.05).

3.3. Pigment contents 3.5. Flow cytometric analyses Plants treated with ABA showed the highest levels of Chl a, while the lowest levels were observed in plants not treated with ABA under DS conditions (P < 0.05) (Table 1). No significant differences were observed in Chl b content (Table 1). Independently of the water regime, plants treated with ABA showed the highest concentration of carotenoids (P < 0.05) (Table 1). Concerning the plants not treated with ABA, the S treatment induced an increase of carotenoids content compared to C treatment. 3.4. Photosynthetic parameters and Rubisco activity Independently of the ABA pretreatment, no significant differences were observed in the Fv /Fm between DS and WW (Table 1). The A values decreased (P < 0.05) in plants of treatments S, S50 and S100 respectively to their controls (C, C50 and C100; Fig. 2A). Plants under WW conditions treated with ABA, C50 and C100, showed the highest and significant A mean, while the lowest A mean was observed in plants under DS conditions without ABA (S treatment) (P < 0.05). Similar to the observed for A, also the E and gs decreased

Leaf flow cytometry (FCM) histograms showed a main peak corresponding to G0/G1, a secondary peak corresponding to G2 and between those peaks a region corresponding to cells in S phase (in control represented 84%, 3.6 and 12.6%, respectively) (Fig. 4). Plants treated with ABA and under DS (S50 and S100 treatments) showed an increase in the percentage of cells in S phase comparatively to the plants under WW conditions of treatment C. These FCM fluorescence histograms displayed an excellent value of HPCV (ranging from 2.5% to 3.2%) much lower than the maximum HPCV value for this type of analyses (HPCV < 5%) and supporting the reliability of the protocol. For the DNA content value, Table 2 shows the 2C nuclear DNA content of U. minor plants. Plants of treatment C (WW conditions) showed a genome size of 4.06 ± 0.01 pg/2C, while for the DS conditions and/or ABA treatments these values ranged from 4.04 to 4.07 pg/2C. However none of these affected the ploidy level in U. minor leaves, which was confirmed by the DNA fluorescence Index (DI = 2C U. minor/2C P. sativum) that did not change independently of the treatment (Table 2).

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Fig. 2. In situ net photosynthetic rate (A) (A), transpiration rate (E) (B), stomatal conductance (gs ) (C) and intercellular CO2 concentration (Ci ) (D) in WW (C, C50 and C100) and DS (S, S50 and S100) U. minor plants. Values are means ± SD (n = 10). Different letters indicate significant differences between treatments at a significant level equal to 0.05.

Treatment with 50 ␮M of ABA (C50 − WW conditions) induced a significant increase of FPCV comparatively to the treatment S (DS conditions) (Table 2). 3.6. Lipid peroxidation, cell membrane permeability, antioxidant enzyme activities and H2 O2 content Plants under DS conditions showed a higher H2 O2 content than those under WW conditions (P < 0.05) (Fig. 3A). However, for the plants of DS conditions the H2 O2 content was significantly higher in the treatment S than in the S50 and S100 treatments. No significant differences were observed in the content H2 O2 in plants under WW conditions. The activity of SOD was significantly higher in the DS conditions than in the WW conditions (Fig. 3B). DS plants of the treatment S50 and S100 showed the highest APX activity, while the plants under WW conditions exposed to treatment C showed the lowest APX activities (P < 0.05) (Fig. 3C). For CAT, the highest activity was observed in S50 treatment, while the treatments C and C100 displayed the lowest activities (P < 0.05) (Fig. 3D). Lipid peroxidation was measured in terms of MDA content and cell membrane permeability was determined by measuring cell Table 2 Nuclear DNA content, and full peak coefficient of variation (FPCV) of WW (C, C50 and C100) and DS (S, S50 and S100) U. minor plants. Values of nuclear DNA content are given as a mean and standard deviation of the mean of the DNA index relative to the internal standard P. sativum cv. Ctirad, as a mean and standard deviation of the mean of the nuclear DNA content (pg/2C) and as 1C genome size of U. minor. Values are means ± SD (n = 6). Different letters indicate significant differences between treatments at a significant level equal to 0.05. Treatment

Nuclear DNA content pg/2C

C S C50 S50 C100 S100

4.06 4.04 4.06 4.05 4.07 4.05

± ± ± ± ± ±

0.01a 0.02a 0.02a 0.03a 0.01a 0.01a

DI 0.45 0.44 0.45 0.45 0.45 0.45

FPCV ± ± ± ± ± ±

0.001a 0.002a 0.002a 0.001a 0.001a 0.001a

3.87 4.05 5.07 4.98 4.65 4.57

± ± ± ± ± ±

0.68a 0.04a 0.27b 0.99b 0.25ab 0.34ab

electrolyte leakage. Plants under DS conditions (S, S50 and S100) showed higher MDA contents and cell membrane permeability (P < 0.05) than their respective controls (C, C50 and C100) (Fig. 3E and F). Moreover, ABA pretreatment led to a general trend of MDA and cell membrane permeability decreases, compared with plants not treated with ABA. Under DS conditions, 50 ␮M ABA was the most effective treatment in decreasing cell membrane permeability. 4. Discussion In this study we applied ABA prior to the DS, contrarily to other approaches that only applied ABA during the DS treatment (e.g., Ma et al., 2008; Li et al., 2004; Yin et al., 2004; Duan et al., 2007; Wang et al., 2011). The period of ABA pretreatment represents therefore not only the innovative approach of this work, but also revealed to be crucial to the results obtained. As expected, ABA-treated plants showed higher levels of this phytohormone when compared with those not treated with ABA. This finding proved that applied ABA was absorbed by leaves and also demonstrated that the pretreatment resulted in an internal accumulation of ABA in leaves that depends on the concentration applied. It is well known that the levels of ABA increase in plants in response to DS (Corcuera et al., 2012). Similar findings were observed in U. minor plants not treated with ABA under DS conditions. Contrasting, in U. minor plants pretreated with ABA, DS did not induce an increase of the ABA levels. Similar results were reported by Duan et al. (2007) with P. asperata, in which ABA was applied together with DS. However, Yin et al. (2004) described that even when treated with 50 ␮M of ABA, poplar plants under DS showed increased levels of endogenous ABA. It was evident that water withheld induced DS in all the treatments, but ABA pretreatment reduced the levels of DS: U. minor plants not treated with ABA showed a RWC below 80%, indicating that the level of DS to which the plants were subjected was severe, while ABA treated plants maintained RWC values above this value indicating moderate stress. Stomatal closure is among the earliest responses to DS, protecting the plants from extreme water loss, which can result in cell

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Fig. 3. Changes in the H2 O2 content (A), in the activities of SOD (B), APX (C) and CAT (D), in cell membrane permeability (E) and in the content of MDA (F) in WW (C, C50 and C100) and DS (S, S50 and S100) U. minor plants. Values are means ± SD (n = 8). Different letters indicate significant differences between treatments at a significant level equal to 0.05.

dehydration and eventually in plant death (Chaves et al., 2003). DS induced stomatal closure in U. minor to prevent water loss, but also limited the CO2 availability in the intercellular spaces of the mesophyll cells, as evidenced by the low Ci . Although DS affected negatively the A, these effects did not influence total DW. When comparing U. minor plants treated vs not treated with ABA it is evident that ABA acts at the stomatal level reducing stomatal closure, decreasing water loss, increasing A and plant DW. Even, in some

100 90

a

a

a

a

a

80

a

% of cells

70 60

%G0/G1

50

%S

40

%G2

30 20 10 0

a

a

C

S

a

a

C 50

S 50

b

ab

b

ab

ab

a

a

a

C 100

S 100

Fig. 4. Cell cycle dynamics of U. minor leaves in WW (C, C50 and C100) and DS (S, S50 and S100) plants. Values are means ± SD (n = 6). Different letters indicate significant differences between treatments at a significant level equal to 0.05.

cases, plants treated with ABA achieved higher A under reduced E, gs and lower CO2 availability. This finding could be related to a putative higher in vivo Rubisco activity in ABA pretreated plants since this key-enzyme is reported to be very sensitive to oxidative stress (Wang et al., 2011) and ABA application enhances the activity of antioxidant enzymes, reducing oxidative stress (Yoshiba et al., 1995), as also observed in this work. Moreover, ABA seems to affect the expression of Rubisco genes (Bray, 2002), but the effects of ABA on the activity and/or regeneration of Rubisco are not yet confirmed. The results obtained in the present work corroborate those reported by other authors that ABA, in general, improve the stability of the photosynthetic apparatus (Pospíˇsilová and Batková, 2004; Wang et al., 2011). As reported in our study, the pretreatment of P. vulgaris, B. vulgaris and Z. mays with 100 ␮M of ABA, before the onset of DS, improved the A (Pospíˇsilová and Batková, 2004). Curiously, in those species the E and gs were, in general, higher in plants treated with ABA than in plants not treated with ABA under DS conditions. Several other reports demonstrated that when ABA (50 ␮M) was applied simultaneously with the beginning of the DS, plant growth and A were negatively affected (Li et al., 2004; Yin et al., 2004; Duan et al., 2007; Ma et al., 2008). Taking all together it seems that the application of ABA before the onset of DS is more beneficial since it improves A and plant growth. DS and ABA pretreatment did not affect the photochemical efficiency of PSII. ABA pretreatment was effective in increasing the levels of carotenoids and Chl a. Similarly, results were reported by Agarwal et al. (2005) in wheat plants. Carotenoids act as

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light-harvesting pigments, protect chlorophylls from degradation (Rodriguez et al., 2012) and can also be the precursor to the hormone ABA (Zeevaarl and Creelman, 1988). Thus, in plants not treated with ABA the lower levels of carotenoids under DS conditions may be related to the ABA production as observed by the endogenous ABA levels measured, while in ABA treated plants, which presented higher levels of endogenous ABA, carotenoids content were higher and did not varied. According to our results, ABA treatment only induced an accumulation of proline under DS conditions. Moreover, this accumulation was more significant in ABA treated plants where the lower oxidative damages and the higher RWC were observed. These results suggest that proline may have and important role under DS conditions as an osmoprotectant, in membrane structures protection and as a ROS scavenger (Ashraf and Foolad, 2007). The increase of proline content under stress conditions was reported for several species including U. minor (Corcuera et al., 2012; Dias et al., 2013a). In addition to proline, several phytohormones play an important role in integrating several stress signals and in controlling downstream stress responses (Corcuera et al., 2012). ABA pretreatment did not affect the level of SA, JA and JA-Ile. However, DS induced an increase in the levels of both JA and JA-Ile. This result indicates that these compounds play an important role in the response of U. minor to DS and corroborate with the findings of Corcuera et al. (2012) in Pinus pinaster and Dias et al. (2013a) in U. minor. Contrarily, SA was not affected by DS, which agrees with the reported by Dias et al. (2013a) in the same species and also in other species (Munné-Bosch et al., 2008; Corcuera et al., 2012). SA seems to induce protective reactions to DS by increasing proline (Umebese et al., 2009) and ABA (Yoshiba et al., 1995). DS induced oxidative stress in U. minor plants. However, ABA pretreatment increased DS tolerance by the enhancement of the antioxidant system and oxidative membrane damages prevention. It is possible that the prevention of cell membrane damages (including also those of organelles as plastids and mitochondria) in ABA pretreated plants is due to a reduction in ROS production, as demonstrated by the lower levels of H2 O2 (Alscher et al., 2002). Despite the similar SOD activities observed in all the treatments under DS condition, ABA treated plants showed reduced levels of H2 O2 which could be attributed to higher activity of APX and CAT. Duan et al. (2007) obtained similar results in P. asperata plants treated with ABA under DS conditions. However, these authors described a predominance of APX over CAT in H2 O2 elimination. In general, our results are in accordance with the findings that ABA plays an important role in the enhancement of tolerance to oxidative stress by increasing the activity of antioxidant enzymes (Yoshiba et al., 1995; Agarwal et al., 2005). Also Jiang and Zhang (2001) demonstrated in maize that the exposure to low levels of ABA induced the activity of antioxidant enzymes including CAT and APX, while higher levels could induce oxidative stress. In addition to the positive effects of ABA in reducing oxidative stress under DS condition, this phytohormone also reduce permeability of cell membranes even under WW conditions. Regarding both ABA concentrations tested, despite the similar photosynthetic responses, under DS the use of 100 ␮M seems to be less efficient in oxidative membrane damages prevention. This could be related to the reduced CAT activity and also to the lower levels of proline in these plants compared to those treated with 50 ␮M ABA under DS conditions. So, we demonstrate here the beneficial effects of a pretreatment with ABA (<100 ␮M) that, by early promoting the activity of antioxidant enzymes, leads to a “pre-acclimation process” allowing the plants to handle stress. Genome size of U. minor was estimated as 4.06 ± 0.01 pg/2C this result corroborates previous determinations and (4.25 ± 0.16 pg/2C) (Loureiro et al., 2007b). Pretreatment with ABA and DS conditions did not induce any significant change in

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DNA content of U. minor. However, plants pretreated with ABA under DS conditions showed a delay in S-phase. To our knowledge this is the first study on the effects of ABA in the cell cycle of leaves under stress. Previous studies in other species on other plant organs indicate that ABA affects cell cycle and DNA synthesis. For instance, in tobacco BY-2 cultured cells, Swiatek et al. (2002) found that JA and ABA inhibited DNA replication keeping cells in G1 stage. Muller et al. (1994) described that ABA decreased the percentage of cells in S-phase in maize roots. Also, in cultured pea embryo axes, ABA inhibited the transition from G1 to S phase, by inhibiting DNA synthesis (Levi et al., 1993). Similarly, ABA blocked embryonic cell division at G1 /S phase in Arabidopsis (Finkelstein and Rock, 2002). ABA abolished S-phase-specific activation of telomerase in a concentration- and time-dependent fashion in synchronized tobacco BY-2 cells (Yang et al., 2002). More recently, it was demonstrated that ABA regulates cell cycle by controlling the expression of the cyclin-dependent kinase inhibitor KRP1/ICK1 (Yin et al., 2009). Interestingly, in elm leaves, only the combination of ABA and DS increased cells in S-phase, suggesting that a delay is taking place to repair stress-induced processes. Cell cycle delays have been described for other stresses and are crucial for the cell repairing processes, including the damaged DNA, which may arise from the oxidative stress. Simultaneously, ABA pretreatment induced an increase in FPCV of the G0/G1 peak in ABA treated plants, mostly for the concentration of 50 ␮M ABA. FPCV reflects the whole dispersion of nDNA content and its increase has been correlated with the increase of nDNA breaks (Rayburn et al., 2002; Monteiro et al., 2010; Rodriguez et al., 2011; Silva et al., 2013). Taken together, both evidences of ABA-induced a cell cycle delay in S-phase and increase of DNA breaks may support putative protective effects of ABA in the cell cycle, which may putatively involve an increase of transient DNA breaks. Recent data suggest that ABA increases homologue recombination and activates the expression of retrotransposons which may be important mechanisms for restructuring the genome as an adaptive response to environmental challenge (Yin et al., 2009; Donà et al., 2013). These authors suggested that ABA might affect genome stability through the DNA replisome, therefore, allowing plants to adapt to environmental stresses over the course of evolution. Another hypothesis relies on the putative effect of ABA on the repairing process, which remains an unknown topic. DNA-specific repairing enzymes cut DNA transiently increasing the number of DNA breaks, which also can be seen in an increase of the FPCV value by flow cytometry. The observed increase of FPCV in U. minor plants treated with ABA, together with the delay in S-phase, may support a role of ABA in promoting, in plants under DS, a delay in S-phase allowing the repairing system to take place. However, further studies are needed to dissect the functional and molecular mechanisms of ABA-induced increase of DNA breaks, and delay of S-phase.

5. Conclusions This study demonstrates clearly that, independently of the water status conditions, ABA pretreatment improves the antioxidant system, A and growth (DW accumulation) of U. minor plants. Furthermore, ABA delays S-phase and increases FPCV. The mechanism involved in this process remains unknown but may include transient breaks of DNA. Comparing our results with the available reports, we conclude that the application of ABA before the onset of DS could be more advantageous since it improves photosynthesis and plant growth. The use of ABA as a pretreatment to alleviate the negative effects of DS seems to be a promising strategy to reduce crop loss, increase plant productivity and could also be applied in reforestation and afforestation programs in drought prone areas.

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