Preparation of AS-SAMDC plants (N. tabacum cv Xanthi) was described previously (Moschou et al., 2008b). In order to achieve synchronous germination, seeds of both genotypes, WT and AS-SAMDC were stratified (4°C, 7 days), and immersed in KNO₃ (1%) solution for 12 h. Seeds were surface sterilized with bleach for 10 min, washed four times with sterile water and plated out on a filter paper in plates containing half strength Murashige and Skoog medium [Duchefa Biochemie, Haarlem, Netherlands; supplemented with 0.05% MES/KOH, pH 5.7 (2-(N-morpholino) ethanesulfonic acid; Sigma-Aldrich, St. Louis MO, USA), Gamborg B5 vitamins and micronutrient mixture (Duchefa), 2% (w/v) sucrose]. Seeds were germinated and grown under long day conditions [16/8 h photoperiod; 22/18°C, 110 μEm⁻²s⁻¹ PAR supplied by cool-white fluorescent tungsten tubes (Osram, Berlin, Germany)] in a plant growth chamber for 13 days. Seedlings were transplanted in plates with the same medium containing 0 or 100 mM NaCl for 2 more days. The seedlings were used immediately or weighed, frozen in liquid nitrogen and stored at −80°C for further use.

Responses at the whole plant level were studied using potted plants, which received NaCl treatments for 21 days. Stratified seeds, were sown in individual 10-cm diameter pots filled with peat (Terraplant, Compo) and perlite (2:1) mixture, and placed under plastic cover for the first 3 days to sustain high relative humidity. At the four-true-leaf stage, 120 seedlings per genotype were transplanted in 0.25 L pots, filled with the aforementioned mixture and placed in a completely randomized design (split-split-plot arrangement). At the 4–5 pair true leaf stage (70 days-old plants), five salt treatments (0, 50, 100, 200, 300 mM NaCl) were applied (24 plants per genotype). All plants were subjected to salt treatment through irrigation either with 100 mL aqueous solution of the respective NaCl concentration or with 100 mL water (controls) at 0, 4, 7, 11, 14, and 18 days (days after treatment, DAT). Preliminary experiments showed that this was an efficient method for sustaining the respective salinity levels in the substrate throughout the entire 21 days experimental period. At four sampling times (0, 7, 14, 21 DAT) six plants (each considered as a single replication) per genotype and treatment were harvested. Growth, agronomical, and physiological parameters were evaluated on three plants; phytochemical indices were determined on the other three.

At each sampling time (0, 7, 14, 21 DAT), individual plant height was recorded from the soil surface to the base of the petiole of the youngest fully expanded leaf. Then, the roots were separated from the shoots, and the shoot fresh weight was determined. The leaves were counted and subsequently removed from the stems for further study. The leaf area (LA) was evaluated from digital images that were acquired using a flatbed scanner. The acquired images were analyzed using ImageJ (Schneider et al., 2012). The samples (leaves and stems) were then oven-dried at 70°C till stable weight and their dry weight was determined.

Total RNA isolation was carried out using the Trizol reagent (Chomczynski, 1993). The extracted RNA was subsequently used for RT-PCR using the Takara Prime Script kit. Alternatively, total RNA was extracted from 100 mg of tissue with TRItidy G^TM (AppliChem). cDNA was synthesized in a 20 μl reaction mixture containing ±4 μg denatured RNA, 50 pM oligo (dT₁₇) primer (VBC-Biotech, Vienna, Austria), 1 mM dNTPs mixture (Qiagen, Germany), 4 μl 5X buffer (PrimeScript, Takara, Shiga, Japan), 20 Units of RNase inhibitor (Biolabs), and 100 Units reverse transcriptase (PrimeScript, Takara, Shiga, Japan). cDNA synthesis conditions were as follows: 5 min at 65°C, 60 min at 42°C, 70°C for 15 min. cDNA samples were diluted 2.5x. PCR was performed in a final volume of 25 μl containing 1.25 units Taq DNA polymerase (Life Technologies, Rockville, Md or Invitrogen, Carlsbad, California), 0.5–6 μl cDNA, 0.3 pM gene—specific primer, 0.3 mM dNTPs mixture (Qiagen), 1.5 mM MgCl₂ (Invitrogen), and 1x PCR reaction buffer minus Mg²⁺ (Invitrogen). Conditions of PCR amplification were as follows: 2 min at 94°C, 1 min at 92°C, 1 min at the appropriate annealing temperature, 35 s at 72°C, and a final extension step of 5 min at 72°C. The primers used were AGCCCATCTTCCTCTCTGTTC and CCTCAAAGCCAATAGCCA AG for SPMS, CCAACCACAACAACGACAAC and TCCAAAACAAGCACCTTTCC for SPDS, and GTCTGGTGATGGTGTTAGC and CCTATCAGCAATTCCAGGAAAC for ACTIN (33, 33, and 30 cycles, respectively). All amplicons were separated by 1.4% (w/v) agarose gel electrophoresis, stained with EtBr and visualized under UV light. The total density of individual bands was measured with GelEval software package (v1.37, Frog Dance Software).

Proteins were extracted and treated as described in Papadakis and Roubelakis-Angelakis (2005). For native electrophoresis and in-gel assays, proteins were resolved by native PAGE and then stained according to each enzyme.

NADPH oxidase activity was determined as described by Papadakis and Roubelakis-Angelakis (2005). A spectrophotometric method was used for PAO assays (Federico et al., 1985). CAT activity was determined by measuring the initial rates of H₂O₂ decomposition at 240 nm (Havir and Mchale, 1987). Total SOD activity was determined using the photochemical assay (Misra and Fridovich, 1977).

PA titers were determined as described previously (Kotzabasis et al., 1993) using an HP 1100 high-performance liquid chromatographer (Hewlett-Packard).

In situ accumulation of H₂O₂ was detected using the method of Thordal-Christensen et al. (1997). The endogenous levels of H₂O₂ content of tissues were determined as described by Sahebani and Hadavi (2009). The content of H₂O₂ was estimated by a standard curve.

Quantification of total soluble phenol content and antioxidant capacity was performed over a 21 DAT period in WT and transgenic plants treated with 0, 50, 100, 200, and 300 mM NaCl. Approximately 500 mg of leaf material (fifth leaf from the apex) was homogenized at 4°C with 80% MeOH containing 1% HCl. Total soluble phenolic compounds in MeOH leaf extracts were determined using the Folin-Ciocalteu reagent (Chem-Lab NV, Belgium) in a 7.5% Na₂CO₃ solution. The extraction mixture was placed in a water bath at 50°C for 5 min and the absorbance was then read at 760 nm with a UV-Vis spectrophotometer (Shimadzu UV-1601). Results were expressed as gallic acid equivalents (GAE) per fresh weight. In addition, total phenolic compounds were determined, qualitatively and quantitatively, in leaf tissues from plants treated for 24 h with 100 mM NaCl. Phenolics were extracted according to Tsiri et al. (2009). All extracts were qualitatively analyzed by Thin Layer Chromatography (TLC) using a stable phase 0.25 mm silica gel 60 and two different mobile phases, nButanol:Acetic-acid:water (3:1:1) and Ethyl Acetate:Acetic acid:Formic acid:water (100:11:11:26). Total phenolic compounds were visualized under UV light after staining with Naturstoff (1% methanolic diphenyl boryl oxyethylamine, followed by 5% ethanolic PEG₄₀₀₀). Quercetin dehydrate, rutin and quercetrin were used as standards in all TLC plates. Quantification of total phenolics was performed using the aluminum chloride colorimetric method (Chang et al., 2002). For determining the total antioxidant capacity of leaves, the ferric reducing antioxidant potential (FRAP) assay was used (Benzie et al., 1999) in MeOH leaf extracts prepared similarly to the determination of total soluble phenolic compounds. Absorbance of the mixture was read at 593 nm with a UV-Vis spectrophotometer (Shimadzu UV-1601). Results were expressed as ascorbic acid equivalents (AAE) per fresh weight. All samples were analyzed in triplicates.

The methods used for measuring extents of energy quenching (qE) component of non-photochemical quenching (NPQ), rates of LEF, and the relative extents of proton motive force (pmf) components were as described in Kanazawa and Kramer (2002) except that a newly developed instrument was used. This instrument was based on the non-focusing optics spectrophotometer (NoFOSpec; Kramer and Sacksteder, 1998) but has been modified to allow near-simultaneous measurements of absorbance changes at four different wavelengths, by aiming four separate banks of light-emitting diodes (HLMP-CM15, Agilent Technologies, Santa Clara, CA), each filtered through a separate 5-nm band-pass interference filter (Omega Optical, Brattleboro, VT), into the entrance of a compound parabolic concentrator (Hall et al., 2013). Each complete set of three pulses was deconvoluted by using the procedure described previously to obtain estimates of Electrochromic Shift (ECS; Avenson et al., 2004; Ioannidis et al., 2012a). The instrument was also used to measure changes in chlorophyll a fluorescence yield by using the 520-nm light-emitting diode bank as a probe beam. Saturation pulses (>7000 μmol of photons m⁻²s⁻¹ PAR) were imposed by using light from the two red actinic LEDs, filtered through heat-absorbing glass. Actinic light was filtered out by using an RG-695 Schott glass filter. Saturation pulse-induced fluorescence yield changes were interpreted as described in Genty et al. (1989). The qE component of NPQ was calculated from the saturation pulse-induced maximum fluorescence yields during steady-state illumination (Fm′) and 10 min (Fm″) after switching off the actinic light (Navakoudis et al., 2007).

Measurements of net photosynthesis (A_net, μmol m⁻²s⁻¹), stomatal conductance (Gs, mol m⁻²s⁻¹), photosynthetic yield (QY), and chlorophyll content index (CCI) were performed on tobacco plants treated with 0, 50, 100, 200, and 300 mM NaCl over a 21 DAT period. Measurements were performed on the fifth fully expanded leaf (counting from the apex) of 70 days-old plants (time 0), 77, 84, and 91 days-old plants (7, 14, 21 DAT, respectively). Three plants (measurements) were used per genotype, parameter and treatment at each sampling time. Both parameters were monitored using a portable photosynthesis system (LI-6200, LI-COR Inc. Lincon NE, USA). Chlorophyll fluorescence parameters were recorded on the fifth leaf (chosen as above) using a direct portable fluorometer (Photosynthesis yield analyzer MINI–PAM, Walz, Effeltrich, Germany). Photosynthetic yield [effective quantum yield of photochemical energy conversion in PSII or simply quantum yield (QY) was computed in terms of the efficiency of the energy harvesting by open PSII reaction centers in the light as previously described (Kadoglidou et al., 2014). CCI was measured using an Opti-Sciences CCM-200 chlorophyll content meter (OptiSciences Inc., Tyngsboro MA). For both QY and CCI, 10 measurements were taken per genotype and treatment at each sampling day. To avoid photoinhibition, all physiological measurements were performed in a 2 h time span, approximately 3 h after sunrise.

Proton Flux and pmf can provide useful information on the photosynthetic performance. These analyses were performed by newly introduced techniques that allow us to non-invasively probe the “proton circuit” of photosynthesis. For the theoretical framework for these methods see (Avenson et al., 2004; Ioannidis et al., 2012a and refs therein). We probed the ECS, an index of pmf by using a previously described technique called dark-interval relaxation kinetic analysis (Sacksteder et al., 2000), in which steady-state photosynthesis is perturbed by short (up to 0.5 s) dark intervals, allowing the photosynthetic apparatus to relax in ways that reveal information about the system in the steady state. The amplitude of the light–dark ECS signal (ECSt) parameter was obtained by taking the total amplitude of the rapid phase of ECS decay from steady state to its stable level after ~300 ms of darkness (Kanazawa and Kramer, 2002). For the deconvolution, all traces were normalized to the initial dark value (i.e., before actinic) and then the following equation was used (Ioannidis et al., 2012a): ECS₅₂₀ = A₅₂₀−0.5 × A₅₃₅−0.5 × A₅₀₅.

NPQ was determined in samples exposed to actinic light of 110 μmol photons m⁻²s⁻¹ using PAM-210 fluorometer (Heinz Walz, Germany). Samples were incubated in the dark for at least 10 min prior to measurement. The NPQ-parameter was calculated according to the equation: NPQ = (FM−FM′)/FM′ (Bilger and Björkman, 1990). For the fluorescence induction measurements, the portable Plant Efficiency Analyzer, PEA (Hansatech Instruments) was used as previously described (Kotakis et al., 2014). The method is based on the measurement of a fast fluorescent transient with a 10 μs resolution in a time span of 40 μs–1 s. Fluorescence was measured at a 12 bit resolution and excited by three light-emitting diodes providing an intensity of 3000 μmol photons m⁻²s⁻¹ of red light (650 nm). For the estimation of Ka, DCMU inhibited electron transport after QA and analysis of the area closure was performed according to the method of Melis as modified (Ioannidis et al., 2009).

Measurements of ion content in soil and in plants treated with 0, 50, 100, 200, and 300 mM NaCl were performed over a 21 days period. In addition, electrical conductivity (EC_se) and sodium adsorption rate (SAR) in substrates were determined (Supplemental Table 1). On each sampling day, soil substrates (i.e., the peat: perlite mixture) from three pots per genotype and treatment were dried at 70°C until constant weight. The dried substrates were then water saturated and the pH of the created pastes measured. Thereafter, aqueous saturation extracts were obtained through vacuum filtration and EC_se, water soluble K⁺, Na⁺, Ca²⁺, Mg²⁺, and SAR were determined (Rhoades, 1996). Ion content (K⁺, Na⁺, Ca²⁺, and Mg²⁺) was determined following NaCl treatment. Shoot samples (0.5–1 g) were burnt in a muffle furnace at 500°C for at least 4 h and the residue was dissolved in a 3.6 M HCl, 1.4 M HNO₃ aqueous solution. Monovalent ions were determined using a flame photometer (Jenway PFP 7, Gransmore Green, Felsted England), and the divalent using an atomic absorption spectrophotometer (ShimadzuAA 6300, Japan).

Wherever applicable, the experiments were designed at a completely randomized pattern in a split-split-plot arrangement, with the genotype, the NaCl dosage and the sampling time as independent factors. Each combined factor had three replicates in all parameters except for the CCI and QY that had 10 replicates. The analysis of variance (ANOVA) and the Tukey's test on the results obtained over the entire experimental period (up to 21 DAT) were performed by the statistical package SPSS (version 22). Duncan's test was done using SigmaPlot 12.0. Student's test was done using JMP (SAS, version 11 pro). Multiple average comparisons, graphs and data processing were performed in EXCEL. Fisher's Protected LSD procedures were used to determine statistical differences between genotypes and treatments at a 5% significance level.

Previously, we used young transgenic seedlings of AS-SAMDC and showed that NaCl stress can be used to monitor the effects of altered ratio of biosynthesis/catabolism of PAs and also the contribution of PAO-generated H₂O₂ in the adaptation of plants to stress and in developmental processes (Moschou and Roubelakis-Angelakis, 2014). We estimated the mRNA abundance of the PA biosynthetic enzymes Spd synthase (SPDS) and Spm synthase (SPMS) in control and NaCl treated WT and AS-SAMDC plants. Under control conditions (0 mM NaCl), mRNA levels of SPDS and SPMS were significantly higher in AS-SAMDC plants by almost two-fold compared to WT plants. During NaCl stress (100 mM), both SPDS and SPMS mRNA levels increased in WT (ca two-fold), while they significantly decreased in AS-SAMDC, when compared to their corresponding controls (Figure 2A). Under control conditions, WT and AS-SAMDC exhibited similar PAO activity. Twenty four hours post-treatment with 200 mM NaCl, PAO activity decreased by 60% in WT whereas increased by 80% in the AS-SAMDC plants. In the WT it increased by 20% and in AS-SAMDC by two-fold 72 h post-treatment (Figure 2B). Following a decrease at 24 h (50%), PAO activity was transiently increased (two-fold) in WT 48 h post-treatment with 300 mM NaCl, while at 72 h it was similar to the corresponding control levels. In AS-SAMDC at 300 mM NaCl PAO showed a constant increase.

Next, we determined the titers of endogenous PAs in WT and AS-SAMDC grown in perlite. PAs were significantly lower in AS-SAMDC, in agreement with previous results (Figure 2C; Moschou et al., 2008b). WT contained higher titers of Put (four-fold), Spm (two-fold), and Spd (20%) compared to the AS-SAMDC. NaCl resulted to a significant increase in PAs in WT (107, 20, and 46% increase in Put, Spd and Spm, respectively, whereas in AS-SAMDC only Put increased (29%); Spd and Spm titers did not change (Figure 2C).

We examined the effect of SAMDC depletion on the development and NaCl stress tolerance. The ANOVA test indicated that all developmental parameters examined were significantly (P ≤ 0.001) affected by sampling time and by the five different NaCl concentrations. In most cases, the effect of the genotype, and the effect of two-way interactions on the various parameters were also significant (Supplemental Table 2). In order to facilitate comparisons, we examined either the effects of each NaCl treatment over a 21 DAT, or the means across the four sampling time points were averaged per NaCl concentration over the entire experimental period (up to 21 DAT). The latter is exemplified throughout Supplemental Figures. Control AS-SAMDC were on average 1.2-fold taller than the respective WT over the 21 days experimental period (Supplemental Figure 1A). Under NaCl stress, AS-SAMDC plants were less resilient and an abrupt decrease of their height was observed (Figure 3; Supplemental Table 2; genotype × NaCl interaction, P < 0.05). At the highest NaCl concentration (300 mM), plant height was reduced by 52% in AS-SAMDC plants, and only 29.5% in WT plants, compared to the respective control plants (0 mM) at the same time point (Figure 3; 21 DAT). Similar trend was observed at lower NaCl concentrations (100 and 200 mM). The dose-response curves over the whole experimental period (Supplemental Figure 1A) show that only the highest concentration of NaCl (300 mM) affected the height of WT while all NaCl concentrations negatively affected the height of AS-SAMDC. NaCl stress caused a reduction in leaf area per plant (LA; cm²) in both genotypes (Supplemental Table 2; Supplemental Figure 2), which was greater in AS-SAMDC (Supplemental Figures 1B, 2). This reduction in AS-SAMDC (reaching a 42% decrease) was evident at all NaCl concentrations, whilst in WT only at the highest ones (Supplemental Figure 1B). Control AS-SAMDC had significantly higher number of leaves throughout the experimental period (Supplemental Figure 1C). NaCl stress led to a significant decrease in leaf number regardless genotype (Supplemental Table 2). Although AS-SAMDC had more leaves at all NaCl concentrations, their leaf number abruptly declined upon NaCl treatment (Supplemental Figure 1C).

Generally, the above ground fresh weight (FW) of controls was not significantly different between the two genotypes neither at 0 DAT nor during the experimental period (Supplemental Figure 3). NaCl resulted to a reduction of FW, yet this effect was similar in both genotypes (Supplemental Table 2; genotype × NaCl interaction). Notably, the highest NaCl concentrations affected more the FW of AS-SAMDC (65 vs. 57% in WT compared to controls) at 21 DAT (Supplemental Figure 3). Interestingly, when the above ground biomass was expressed as dry weight (DW), the control AS-SAMDC plants showed increased biomass compared to the WT from 14 DAT and onwards (Figure 4; 25%). NaCl significantly impaired DW in both genotypes but to a different extent (Figure 4; Supplemental Table 2; genotype × NaCl interaction, P < 0.001; Supplemental Figure 4). The same trend was observed at 50 mM NaCl even though the differences were not statistically significant. At high NaCl concentrations however, a trade-off between AS-SAMDC DW and NaCl tolerance was evident (a 64% decrease in AS-SAMDC at 300 mM, and a 34% decrease in WT at 300 mM Figure 4).

Considering the increased DW of AS-SAMDC plants under normal growth conditions, we examined whether this correlated to increased photosynthetic potential. We assessed maximal photosynthetic rates in terms of CO₂ fixation for the 21 DAT period. Net photosynthetic rate, A_net, expressed as the rate of net CO₂ assimilated per leaf area was severely impaired by NaCl in both genotypes regardless of the concentration used (Figure 5; Supplemental Table 2). Interestingly though, at 21 DAT AS-SAMDC showed ca. 50% lower A_net at 300 mM NaCl compared to WT (AS-SAMDC 1.6 vs. 3.2 mol m⁻²m⁻¹ for WT). At the same time point, the highest NaCl concentration affected more A_net of AS-SAMDC (up to 72%) than of WT (up to 50%) compared to corresponding controls. On the other hand, at lower NaCl concentrations (e.g., 50 mM) or earlier time points (e.g., 200 mM, 14 DAT), an inverse trend could be observed. Stomatal conductance (mol m⁻²s⁻¹) was not significantly affected by NaCl, genotype or the interaction between these two factors (Supplemental Table 2), although the AS-SAMDC showed in some cases a slightly reduced response under NaCl (Supplemental Figure 5). Generally, WT and AS-SAMDC controls showed comparable values of QY (Supplemental Figure 6). Yet, NaCl slightly reduced QY especially at concentrations over 200 mM, regardless of the genotype, but to a different extent. In particular, at 21 DAT the reduction in QY compared to control plants was greater in the NaCl-treated AS-SAMDC compared to WT plants at the highest NaCl concentration (a maximal 4.4% decrease vs. 1.3%, respectively), suggesting that SAMDC has a role in sustaining the efficiency of light use in Photosystem II (PSII) during photosynthesis.

Next, we examined in more detail the photosynthetic apparatus in seedlings grown under more defined conditions (i.e., growth in perlite with MS). We determined the photo-protection capacity via NPQ, and the qE at 200 mM NaCl at four different light intensities. These mechanisms are employed by cells to dissipate the adverse effects imposed by high light intensity (Figures 6A,B; and Supplemental Table 4). NPQ is the sum of the total effect of high qE, state transitions, and photoinhibition (pI). At low light intensities NaCl affected slightly the NPQ and qE (about 27% in WT and 33% in AS-SAMDC compared to controls). However, with increasing light intensity the differences between WT and AS-SAMDC increased (NPQ by 40 and 91%, respectively). By estimating qE values under these conditions we confirmed that the effect in NPQ is largely due to qE and not to qI or state transitions. NaCl led to a profound increase of qE only in the AS-SAMDC (207 vs. 27% in WT; Figure 6B). In addition, Electrochromic Shift (ECS), an index of pmf was determined. The techniques used take advantage of the ECS (known also as Δ⁵²⁰ or ΔA⁵¹⁸) of certain carotenoid species that naturally occur in thylakoidal membranes. The ECS is a linear indicator of changes in trans thylakoid ΔΨ and is particularly useful since it responds to the trans thylakoid movement of protons, as well as other charged species. ECS was significantly increased in AS-SAMDC (Figure 6C). In particular, NaCl exerted a mild effect, if any, on WT whereas the same NaCl concentration increased thylakoidal pmf of AS-SAMDC (by 32%). On the other hand, NaCl hindered linear electron flow (LEF) at higher light almost at the same level for both genotypes (21% in WT and 25% AS-SAMDC, Figure 6D). The apparent conductivity of the thylakoidal ATPase was much more sensitive in AS-SAMDC (about 19% at low light) than in WT (practically similar values of gH⁺ between 0 and 200 mM NaCl; Figure 6E). Only at higher light intensities, there was a clear decrease in gH⁺ values in WT. NaCl decreased the total proton flux (UH+) from lumen to stroma (19% in WT and up to 23% in AS-SAMDC, Figure 6F). Furthermore, these changes were associated with significant reductions of Put and Spd in AS-SAMDC (Figure 6G).

The leaves of AS-SAMDC were significantly greener at the start of the experiment (0 DAT, 0 mM NaCl), as indicated by the significantly higher CCI at pre-treatment (Supplemental Figure 7). Yet, this difference between controls of the two genotypes became non-significant when examined throughout the experimental period. Interestingly, at 21 DAT, NaCl treatment significantly increased CCI in both genotypes, compared to controls, an increase becoming evident from 100 mM NaCl onwards in WT, but only at the highest NaCl concentration in the AS-SAMDC plants (Supplemental Figure 7).

Considering the paramount role of PAs in the regulation of H₂O₂ homeostasis (Andronis et al., 2014), and the effect of ROS in plant development and stress defense, we examined main oxidant/antioxidant machineries and H₂O₂ titers in WT and AS-SAMDC control or stressed plants. Hydrogen peroxide and superoxides (O2.-) are tightly inter-dependent in cells and elaborate machinery regulates the balance between these two ROS. One of the major oxidative machineries (i.e., ROS producers) is that of the plasma membrane localized nicotinamide adenine dinucleotide phosphate-oxidase (NADPH-oxidase) that produces O2.-. The mRNA abundance of the NADPH-oxidase subunits, the respiration burst homolog rbohD increased in the AS-SAMDC control plants, while that of rbohF was reduced under control conditions (Supplemental Figure 8; P < 0.05). NaCl induced the accumulation of rbohD and rbohF mRNA in both genotypes in a dose-dependent manner, with WT plants showing higher induction. On the other hand, NADPH-oxidase activity in AS-SAMDC was significantly higher under both control and NaCl conditions (Figure 7A).

NADPH-oxidase feeds superoxide dismutase (SOD) in the cytoplasm, mitochondria and chloroplasts with O2.-, to dismutate it to H₂O₂. We observed a slight decrease of MnSOD mRNA levels in AS-SAMDC controls, while in both WT and AS-SAMDC mRNA levels of MnSOD were further decreased (Supplemental Figure 8). CuZnSOD mRNA levels were significantly higher in AS-SAMDC compared to WT, during control and NaCl conditions (Supplemental Figure 8). On the contrary, we observed increase of SOD isoforms and activity in both WT and AS-SAMDC (Figure 7B; Supplemental Figure 8). These results suggest that increased O2.- in AS-SAMDC is compensated by SOD controlled at the post-transcriptional level.

Catalase (CAT) is the main enzyme that scavenges bulk H₂O₂. We observed a small non-significant increase of CAT activity in control AS-SAMDC, and a decrease of the corresponding levels during NaCl stress when compared to the WT (Figure 7C) and marginally increased H₂O₂ titers in leaves (Figures 8A,B). On the contrary, under NaCl AS-SAMDC accumulated significantly more H₂O₂ than WT.

Since antioxidant defense of plants is also complemented by non-enzymatic antioxidants, such as phenolics, we examined their content of 24 h post-treatment with 100 mM NaCl (phenolics; Figure 9A), or over a period of 21 DAT (phenolics and antioxidant capacity; Supplemental Figure 9). Total phenolics were significantly higher in the AS-SAMDC under control and NaCl. No significant alterations in the zonation pattern between WT and AS-SAMDC plants were found (Figure 9A). Upon NaCl treatment, both the WT and AS-SAMDC showed significant increase in total phenolics (two- to three-fold; Figure 9B). These results were further confirmed in NaCl-treated plants over a period of 21 DAT, when significantly higher content of total soluble phenolic compounds of AS-SAMDC plants versus WT at 100 mM NaCl was reconfirmed (20% higher), and further recorded at 300 mM NaCl (15% higher; Supplemental Figure 9A). On the contrary, no particular difference in antioxidant capacity was observed between genotypes at control conditions nor upon NaCl treatment (Supplemental Figure 9B).

PAs play a major role in controlling ion homeostasis (Shabala and Pottosin, 2014), and thus we examined whether AS-SAMDC showed imbalances in ion accumulation that could drive NaCl sensitivity. Under control conditions, at the start of the experiment, WT and AS-SAMDC showed similar ion content in leaves (Supplemental Table 3). Thereafter, leaves of WT contained higher amount of K⁺, Ca²⁺, and Mg²⁺ than those of AS-SAMDC (Figure 10). During NaCl treatment, increasing NaCl concentrations caused in some cases statistically significant increase in ion accumulation. In particular, Na⁺ content in leaves increased in both genotypes in a dose-dependent manner over the experimental period (Figure 10A). Statistically significant differences between the two genotypes were observed only at 300 mM NaCl (AS-SAMDC leaves accumulated 16% more Na⁺ than WT ones). Na⁺ content in stressed WT and AS-SAMDC plants at 300 mM NaCl was ca. 5- and 12-fold higher, respectively compared to the corresponding controls. Leaves of WT contained higher K⁺ content throughout the experimental period compared to AS-SAMDC (Figure 10B). Increasing NaCl concentrations led to increased leaf K⁺ content, regardless of the genotype (at 200 and 300 mM). The pattern of Ca²⁺ accumulation was different between the genotypes (Figure 10C). Control WT plants contained 1.3-fold more Ca²⁺ in leaves than AS-SAMDC. Interestingly, NaCl did not affect leaf Ca²⁺ content in WT, where a rather constant content of Ca²⁺ was maintained, whereas NaCl significantly increased Ca²⁺content in AS-SAMDC (over 35% at >200 mM NaCl). Similarly to Ca²⁺, Mg²⁺ content was 1.3-fold higher in the leaves of WT compared to the AS-SAMDC control plants (Figure 10D). The response of Mg²⁺ accumulation under NaCl conditions followed a similar pattern to Ca²⁺, although the increase of Ca²⁺ content in the latter case was more prominent in the AS-SAMDC plants.

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.