Three-year old Q. ilex saplings grown under field conditions were potted in 6.5-L pots with growing medium containing a mixture of standard soil Einhetserde Topfsubstrat ED 63 (Sinntal-Altengronau, Germany) and sand (3.5:1, in volume), according to Cotrozzi et al. (2016b). Two weeks before the beginning of the O₃ treatment, 42 plants (WS) received 20% of the effective daily evapotranspiration (calculated by the average 24-h weight loss of five well-watered plants), whereas another 42 plants (WW) were kept at field water capacity. The two groups of plants were then subdivided into four sets (WW-O₃, WS-O₃, WW+O₃, WS+O₃; 21 plants per set) and transferred into four controlled fumigation facilities (temperature 23 ± 1°C, relative humidity 85 ± 5% and photon flux density of 530 μmol photons m^-2 s^-1 at plant height provided by incandescent lamps with L/D 14:10 photoperiod; lights were switched on from 7:00 to 21:00 to simulate environmental light conditions).

WW-O₃ and WS-O₃ plants were randomly distributed into two chambers, whereas WW+O₃ and WS+O₃ plants were randomly distributed in the other two chambers. After one week of acclimation, WW+O₃ and WS+O₃ plants were exposed to an acute O₃ stress (200 nL L^-1, 5 h day^-1, in the form of a square wave between the 2nd and the 7th h of the light period). On the other hand, WW-O₃, WS-O₃ plants were maintained under charcoal-filtered air, in which the O₃ concentration was less than 5 nL L^-1. During the O₃-exposure, environmental factors were maintained as reported above.

The O₃ exposure was performed according to Lorenzini et al. (1994) with minor modifications to avoid pseudo-replications. At the end of the drought exposure, plant water status was evaluated. Photosynthetic parameters were measured at 0, 5, 24 and 48 h from the beginning of the O₃ exposure (FBE, From the Beginning of Exposure). Five fully expanded mature leaves per plant per treatment were taken at 0, 1, 2, 5, 8, and 24 h FBE, stored at –20°C and subsequently used for chemical analyses, with the exception of ET determination, which was performed immediately. At the same measuring times, staining, and microscopic assays were also performed on fresh material.

Pre-dawn leaf water potential (PDΨ_W) was measured on three plants per treatment (one fully expanded mature leaf per plant) with a pressure chamber (PMS model 600, PMS Instrument Company, Albany, OR, USA). On the very same plants, relative water content (RWC) was calculated (one fully expanded mature leaf per plant) as: RWC (%) = (FW-DW)/(TW-DW) × 100, where FW is the fresh weight, TW is the turgid weight after rehydrating samples for 24 h, and DW is the dry weight after oven-drying samples at 85°C for 24 h.

Gas exchange and chlorophyll a fluorescence measurements were determined between 10:00 and 13:00 (solar time) on one fully expanded mature leaf per plant, on three plants per treatment. CO₂ assimilation rate (A), stomatal conductance to water vapor (g_s) and intercellular CO₂ concentration (C_i) in light-saturated conditions and ambient CO₂ concentration were measured using an Infrared Gas Analyzer (LI-COR Inc., Lincoln, NE, United States) as reported by Cotrozzi et al. (2016b). Modulated chlorophyll a fluorescence of photosystem II (PSII) was measured with a PAM-2000 fluorometer (Walz, Effeltrich, Germany) on the same leaves used for the gas exchange after 40 min of dark adaptation using a dark leaf clip provided by the producer. The maximal PSII photochemical efficiency [F_v/F_m = (F_m–F₀)/F_m] and the photochemical efficiency in light conditions [Φ_PSII = (F_m’–F_s)/F_m’)] were calculated (Genty et al., 1989). Maximal fluorescence, F_m, when all PSII reaction centers were closed, was determined by applying a saturating light pulse (0.8 s) at 8,000 μmol m^-2 s^-1 in dark-adapted leaves. Fluorescence was induced with actinic light (about 480 μmol m^-2 s^-1), superimposed with 800 ms saturating pulses (10,000 μmol m^-2 s^-1) at 20 s intervals to determine maximal fluorescence in the light-adapted state (F’_m). Minimal fluorescence in the light-adapted state (F’₀) was determined immediately after turning off the actinic source in the presence of a far-red (>710 nm) background for 10 s to ensure maximal oxidation of PSII electron acceptors. The saturation pulse method was used to analyze the quenching components, as described by Schreiber et al. (1986).

For the detection of dead cells, Evan’s blue staining was used according to Tonelli et al. (2015). Leaf material was boiled for 1 min in a mixture of phenol, lactic acid, glycerol and distilled water (1:1:1:1, in vol.) containing 20 mg mL^-1 Evan’s blue and, after clarification with an aqueous chloral hydrate solution, examined under a light microscope (DM 4000 B, Leica, Wetzlar, Germany). To detect H₂O₂ accumulation, fresh leaf samples were stained with 3,30-diaminobenzidine (DAB) following Tonelli et al. (2015). Leaf parts were incubated for at least 8 h in a DAB solution (1 mg mL^-1) in HCl adjusted to pH 5.6. The samples were then soaked in 70% (v/v) boiling ethanol and clarified overnight in a solution of 2.5 g L^-1 aqueous chloral hydrate solution. The cellular H₂O₂ accumulation was visualized under a light microscope as a reddish-brown precipitation.

H₂O₂ production was measured using the Amplex Red Hydrogen Peroxide/Peroxidase Assay Kit (Molecular Probes, Invitrogen, Carlsbad, CA, United States), according to Pellegrini et al. (2013). Spectrofluorimetric determinations were performed with a fluorescence/absorbance microplate reader (Victor3 1420 Multilabel Counter Perkin Elmer, Waltham, MA, United States) at 530 and 590 nm (excitation and emission resorufin fluorescence, respectively). O2– concentration was measured according to Tonelli et al. (2015), after extraction with a Tris-HCl buffer (50 mM, pH 7.5), with a spectrophotometer (6505 UV-Vis, Jenway, United Kingdom) at 470 nm, and using a buffer solution as a blank.

Two minutes after excision, ET production was measured by enclosing six intact leaves (cut a few millimeters below the petiole by a scalpel) in air-tight glass containers (80 mL). Gas samples (2 mL) were taken from the headspace of containers after 1 h incubation at room temperature. Separations were performed with a gas chromatograph (HP5890, Hewlett-Packard, Ramsey, MN, United States) equipped with a stainless steel column (150 × 0.4 cm i.d. packed with Hysep T) and a flame ionization detector. Analytical conditions were as follows: injector and transfer line temperature at 70 and 350°C, respectively, and carrier gas nitrogen at 30 mL min^-1 (Pellegrini et al., 2013). SA was determined according to Vitti et al. (2016) with some minor modifications. High performance liquid chromatography (HPLC) separations were performed with a liquid chromatograph (Dionex, Sunnyvale, CA, United States) equipped with a reverse-phase Dionex column (Acclaim 120, C18 5 μm particle size, 4.6 mm i.d. × 150 mm length) and RF 2000 Fluorescence Detector. Analytical conditions were as follows: excitation and emission at 305 and 407 nm, respectively, mobile phase containing 0.2 M sodium acetate buffer, pH 5.5 (90%) and methanol (10%), and the flow-rate at 0.8 mL min^-1. JA was determined according to Pellegrini et al. (2013). HPLC separations were performed with the Dionex column described above and a UVD 170U UV/VIS detector. Analytical conditions were as follows: absorbance at 210 nm, mobile phase containing 0.2% (v/v) acidified water, and the flow-rate at 1 mL min^-1. ABA was measured after extraction in distilled water (water:tissue ratio, 10:1) overnight at 4°C. The indirect ELISA determinations, based on the use of DBPA1 monoclonal antibody, raised against S(+)-ABA, as described by Trivellini et al. (2011), were performed at 415 nm with a microplate reader (MDL 680, Perkin-Elmer, Waltham, MA, United States).

Proline content was measured as reported in Cotrozzi et al. (2016b), after extraction with sulfosalicylic acid (3%, v/v). Spectrophotometric determinations were performed at 520 nm, using toluene as a blank.

Three repeated experiments were set up following a randomized design and the experimental plot consisted of one plant per container. Ecophysiological and biochemical measurements were carried out on three replicates for each treatment. The normality of data was preliminary tested by the Shapiro–Wilk W test. The effects of drought exposure vs. well-watering were analyzed by the Student’s t-test. The effects of O₃ on ecophysiological parameters were tested using one-way repeated measures ANOVA with treatment (WW+O₃, WS+O₃) as the variability factor. The effects of O₃ on biochemical parameters were evaluated by two-way ANOVA with treatment and time as variability factors. For both ecophysiological and biochemical analyses, Fisher’s LSD was used as the post-test, with a significance level of P ≤ 0.05. Since data obtained by control plants maintained in filtered air (WW-O₃ and WS-O₃) did not show significant differences during the time course (data not shown), a comparison among means was carried out using as WW+O₃ and WS+O₃ plants controls before beginning the fumigation. Analyses were performed by NCSS 2000 Statistical Analysis System Software (Kaysville, UT, United States).

After 15 days of drought, plants did not show visible foliar injury. Physiological responses are reported in Table 1. In WS plants, PDΨ_W decreased significantly to -4.0 MPa at the end of water deprivation compared to WW controls (–0.5 MPa). However, no changes in RWC were recorded in WS leaves. The net carbon gain was reduced by about 73% in response to drought, which was a larger effect compared with the reduction of g_s (–50%). Values of C_i increased in WS leaves (+7%). Chlorophyll fluorescence measurements revealed a reduction in Φ_PSII (–39%) and qP (–18%) in WS compared to WW leaves, but no changes in the F_v/F_m ratio. An increase of qNP (+30%) was found after drought stress.

The biochemical responses at the end of drought exposure are summarized in Table 2. In comparison to the controls, H₂O₂ levels did not change in WS leaves, while accumulation of O2– was 1.6-fold higher under drought. A strong increase in Pro content (+39%) was observed in WS leaves. The endogenous concentration of ABA and SA measured in WS leaves decreased significantly at the end of the experimental period (–33% and –39%, respectively). However, the JA and ET amounts accumulated by WS leaves increased significantly (about 7-fold and +66%, respectively).

Although drought induced a strong decrease in PDΨ_W (reaching lower values than those reported in a previous study; Cotrozzi et al. (2016b), attributable to different growing seasons between experiments), the RWC of WS leaves did not significantly change in comparison to the WW leaves. This indicates that a good level of leaf hydration was also maintained under drought conditions, RWC being a reliable indicator of leaf water content (Rosales-Serna et al., 2004). This result is in accordance with the accumulation of Pro, a metabolite that is considered an important compatible solute which (i) facilitates water absorption by increasing the cell osmotic potential (Ashraf and Foolad, 2007), and (ii) reduces cell damage (Filippou et al., 2014). The important role of Pro in response to water stress has already been reported in this species where Pro played a key role in the high plasticity of Q. ilex under a long period of moderate water stress (Cotrozzi et al., 2016b).

In WS plants, the strong decline in CO₂ photo-assimilation was attributable to coordinated and concomitant stomatal and mesophyll limitations, which is in line the results obtained by several authors (e.g., Centritto et al., 2009; Flexas et al., 2013). The drop in A levels was higher than that in g_s, thus leading to lower values of intrinsic water use efficiency (data not shown), as already reported in this species when subjected to water stress (Cotrozzi et al., 2016b).

These outcomes indicate that CO₂ assimilation was strongly influenced not only by stomatal behavior but also by mesophyll limitations, as demonstrated by the increase in C_i. Drought compromised the PSII photochemical efficiency in light adapted leaves with decreases in Φ_PSII and qP levels, although this inhibition did not determine PSII photoinhibition, as confirmed by unchanged values in the F_v/F_m ratio. In WS conditions, the lower CO₂ assimilation rate induced, in turn, a reduced consumption of ATP and NADPH synthesized into the chloroplasts and, consequently, led to an excess of excitation energy in the thylakoid membrane, which was only partially dissipated, via non-photochemical mechanisms (increase in qNP). The remaining excess of reducing power in WS plants altered the ROS levels (although H₂O₂ did not change, a strong increase of O2– was observed). This led to a modification in phytohormones and other signaling molecule cross-talk in terms of (i) promoting oxidative stress and (ii) modulating leaf senescence (Munné-Bosch and Alegre, 2004; Miller et al., 2010; Baxter et al., 2014), which is a defense commonly activated in response to both a plethora of abiotic and biotic stresses (Wingler and Roitsch, 2008).

Among phytohormones, the important roles of ABA and ET in plant responses to drought is well known (Munné-Bosch and Alegre, 2004) as ABA represents the most important regulator of stomata functioning (Wilkinson and Davies, 2002), whereas ET is a shoot growth inhibitor and a promoter of ripening, senescence and abscission (Abeles et al., 1992). However, ET can inhibit ABA-induced stomatal closure (Tanaka et al., 2005). Wilkinson and Davies (2009, 2010) reported that under stress-oxidative conditions, an ET-antagonism of the stomatal response to ABA occurs. This interaction was confirmed by our data (ABA decreased, while ET increased), suggesting that drought-induced ET biosynthesis could be considered a marker of leaf senescence (Dangl et al., 2000; Dolferus, 2014).

In addition, SA and JA have been shown to play a central role in leaf senescence (Abreu and Munné-Bosch, 2008), although they are well known for triggering defense reactions against biotrophic and necrotrophic pathogens such as induced resistance (Barna et al., 2012; Shigenaga and Argueso, 2016). In particular, SA and JA interact at physiological levels in many growth and developmental processes, and they play a role in controlling gene expression during leaf senescence (Abreu and Munné-Bosch, 2008). However, as only the JA levels increased in WS leaves, it is reasonable that only JA participated in senescence-associated signaling and degradative processes of membranes. The significantly higher levels of JA shown by WS compared to WW plants indicate that lipid peroxidation producing substrates for octadecanoid pathways was exacerbated in limited water conditions. In particular, JA could be a promoter of leaf senescence in response to drought, thus leading to stomatal closure and accumulation of osmo-compatible solutes (in our case, only Pro), in line with Dar et al. (2015).

The physiological responses observed in O₃-stressed plants were similar to those shown at the end of water deprivation, and in accordance with a previous study by our research group on Q. ilex exposed to O₃ (Cotrozzi et al., 2016b). At the end of the fumigation, the O₃-induced stomatal closure found in both WW+O₃ and WS+O₃ leaves led to significant reductions in CO₂ assimilation. The more pronounced decrease in A in WW+O₃ compared to WS+O₃ leaves, as well as the lack of a further decrease in A observed in WS+O₃ plants throughout the recovery phase, was probably attributable to the very low CO₂ assimilation rate shown by water stressed plants before the beginning of the fumigation. The increase in C_i level in plants exposed to O₃ indicates that the pollutant gas influenced not only stomatal conductance but, as with after water stress, also the mesophyll activity. In fact, an impairment of PSII activity was recorded. Although F_v/F_m and Φ_PSII values decreased significantly after O₃ exposure in both WW+O₃ and WS+O₃ plants, the reduction was more pronounced in WS+O₃ plants. This behavior was linked to different quenching responses to the leaf-water status of plants. In WW+O₃ plants, where qP did not decrease, a mechanism aimed at dissipating the excess excitation energy was activated (qNP increased). By contrast, in WS+O₃ leaves, the O₃-induced decrease in qP was ascribable to the fact that qNP values were already high (probably at their maximum in relation to the capability of plants to activate this mechanism) after drought, and the leaves were not able to enhance this type of dissipation mechanism. The complete recovery of PSII photochemical activity during the recovery time after drought and O₃ exposure indicates that the decrease in PSII activity was sufficient to prevent the photosynthetic apparatus from undergoing irreversible damage.

Unlike the ecophysiological measurements, microscopic analyses highlighted significant differences between plants exposed (WW+O₃, WS+O₃) or not (WW and WS) to the gaseous pollutant. Although visible symptoms were not shown by any of the plants irrespectively of the applied treatment, DAB staining and Evan’s blue incorporation observed in WW+O₃ and WS+O₃ leaves 5 h FBE indicated that H₂O₂ deposition and cell death had already occurred at the end of exposure. This confirms that O₃ resembles the HR occurring in incompatible plant-pathogen interactions (Iriti and Faoro, 2008; Vainonen and Kangasjärvi, 2015). An integrated perspective has been proposed to explain how phytohormones and signaling molecules might be involved in molecular events (namely lesion initiation, propagation, and containment) leading to O₃-induced HR-mimicking foliar symptoms (Overmeyer et al., 2003, 2005; Kangasjärvi et al., 2005). ROS, phytohormones and other signaling molecules have a pivotal role in both HR-mimicking responses induced by acute O₃ and in promoting leaf senescence under drought (as shown by WS plants). The trends of these molecules were monitored in both well-watered and drought-stressed plants during and after O₃ exposure, in order to test the hypotheses of this work.

Given that O₃ induces an endogenous, active and self-propagating ROS generation in the apoplast and a subsequent cellular oxidative burst, some authors have proposed that short-term O₃ exposure mimics pathogen infection (Rao et al., 2000; Kangasjärvi et al., 2005; Carmody et al., 2016). The two O₃-induced H₂O₂ peaks observed in our saplings, irrespectively of the water conditions, are analogous to the biphasic response usually observed during the establishment of the HR of plants against pathogens. The first H₂O₂ peak usually reflects elicitation by pathogen-associated molecular patterns, and the second reflects the interaction between a pathogen-encoded virulence gene product with a plant resistance gene (Mur et al., 2009). In our study, the first peak observed during the fumigation was attributable to O₃-decomposition, whereas the second peak, in the recovery period, could be entirely ascribable to the plant metabolism, in line with Mahalingam et al. (2006), Di Baccio et al. (2012), and Pellegrini et al. (2013) in herbaceous species.

Although the similarity in H₂O₂ profiles over time between WW+O₃ and WS+O₃ conditions, the divergence in the magnitude of their relative peaks (the second peak of the WW+O₃ plants was much greater than their first peak and greater than the second peak of the WS+O₃ plants, where the two peaks were not significantly different from each other) suggests that drought partially inhibited the response to O₃-stress. As H₂O₂ is one of the most important products of oxidative stress (Gill and Tuteja, 2010), it is reasonable to speculate that the biphasic trend of H₂O₂ observed in Q. ilex, irrespectively of drought stress, might reflect the biphasic oxidative burst in response to O₃, in line with several authors (e.g., Wohlgemuth et al., 2002; Di Baccio et al., 2012).

However, this hypothesis is strengthened by O2– only in WW+O₃ plants, where a biphasic time course of O2– levels was shown concurrently with H₂O₂. Kangasjärvi et al. (2005) reported similar temporal changes in ROS for O₃-sensitive genotypes of several species (e.g., tobacco, tomato, birch), whereas only a modest increase in the first hours of exposure was observed in O₃-tolerant genotypes. Conversely, the different O2– patterns of the WS+O₃ plants suggest a possible dual function of this radical depending on water stress. In fact, the significant decrease in O2– observed in WS+O₃ plants during the first hours of exposure suggests that under drought+O₃ superoxide anion may act as a precursor of H₂O₂ and even more toxic radical derivatives. However, the accumulation of O2– had already been induced by drought before the beginning of O₃-exposure. On the other hand, the marked increase in O2– content at the end of the exposure demonstrates that this radical may also be directly involved in the O₃-oxidative burst.

Reactive oxygen species should not be considered as exclusively deleterious and harmful. They can (i) play a key role in intracellular communication which triggers the acclimation ability, and (ii) indirectly orchestrate PCD (Mittler et al., 2011; Xia et al., 2015; Carmody et al., 2016). The amplification of ROS signals and the complete induction of defense genes seem to require signal molecules (Overmeyer et al., 2003). The differences observed in the present study in O₃-induced ROS extent dynamics in relation to water stress suggest a rather complex network of events in signal transduction, involving other molecules (e.g., phytohormones) and processes. Metabolites such as ET, ABA, SA and JA may interact at the physiological level in many growth and developmental processes, with a key role in controlling gene expression during leaf senescence. Most of the genes regulated by these metabolites are defense-related (Fossdal et al., 2007), participating therefore in responses to O₃ (Xu et al., 2015).

Under both biotic and abiotic stresses, SA is required for the induction of PCD, controlling and potentiating the oxidative burst together with ET, whereas JA is involved in limiting the spread of lesions (van Loon et al., 2006; Shigenaga and Argueso, 2016). There are three phases that highlight the influence of ABA on stress conditions (Rejeb I.B. et al., 2014). First, ABA induces stomatal closure, which leads to a reduction in water loss (in this phase, SA, JA and ET may not yet be activated and ABA can antagonize their induction). In the second step, there is a post-infection reaction- an intact ABA signaling pathway is required to increase callose accumulation in affected plants, and the presence of ABA can induce or repress additional callose accumulation depending on the environmental conditions. The third phase begins when pathogen-associated molecular patterns stimulate the accumulation of specific SA, JA, and ET hormones in order to regulate the defense reaction.

In our study, the patterns of phytohormones during and after O₃ treatment were completely different in WW+O₃ and WS+O₃ plants, showing how drought stress has a pivotal role in O₃ responses, and how these signal molecules may be altered in relation to water stress. The ET and ABA accumulations observed throughout the entire period of O₃ exposure occurred only in well-watered conditions. On the other hand, when plants had been previously subjected to water stress, their unchanged values suggest that ET and ABA were not involved in either signaling-responses to O₃, or senescence strategies (as shown for WS plants).

It is worth noting that (i) the maximal ET emission in WW+O₃ plants coincided with the second peak of H₂O₂ and O2–; (ii) the first peak of ABA (during O₃ treatment) preceded that of H₂O₂, suggesting that ABA could act as a stress messenger by inducing H₂O₂ (Jiang and Zhang, 2002) and consequently stomatal closure (as confirmed by the decrease in g_s values observed at the end of exposure), and (iii) the weaker second ABA peak (in the recovery phase) was concomitant with the maximum H₂O₂ and O2– levels and the maximal ET emission. These outcomes confirm a spatial and functional correlation between ROS and the accumulation of these phytohormones.

The SA induction observed, irrespectively of whether the plants had been subjected to drought or not, suggests that this metabolite is also an important modulator of O₃-induced responses (Pasqualini et al., 2002; Horváth et al., 2007). However, the differences between WW+O₃ and WS+O₃ plants show that the functioning of SA is dependent on water stress. In WW+O₃ conditions, the strong increase in SA during the first hours of the treatment and at the end of the O₃ fumigation, confirms the central role of this metabolite in lesion initiation and progression in response to O₃ (Tamaoki, 2008). In addition, the greater SA concentration corresponded with the maximal ABA stimulation and the first increase in ET, thus demonstrating the synergistic action of these hormones in the regulation of defense reactions (Roychoudhury et al., 2013; Wang et al., 2013). By contrast, the biphasic time course of SA (similar to that of H₂O₂) shown by WS+O₃ plants (although slight) recalls the biphasic induction that develops during biotrophic pathogen infection (Mur et al., 2009). Here, the similarity in magnitude of the two SA peaks suggests that the first accumulation of this metabolite (concomitant with the first peak of H₂O₂) did not actuate the second increase in H₂O₂ and hence did not affect the level of plant defense.

The highest concentration of JA observed in WW+O₃ plants during the first hour of fumigation coincided with the initial increase in ET and the maximum accumulation of SA and ABA, thus also demonstrating a spatial and functional correlation between these compounds (Thaler et al., 2012). The significant O₃-induced decrease in JA levels observed in WS+O₃ plants during and after the exposure suggests that JA did not promote leaf senescence in O₃-treated leaves in spite of the high concentrations of this metabolite observed in WS+O₃ plants, not excluding the involvement of JA in senescence-associated signaling (Abreu and Munné-Bosch, 2008). In fact, the JA level in WS+O₃ plants increased again after the end of fumigation, reaching higher values than those found in the WW+O₃ counterpart during the recovery. JA is known to rapidly inhibit the expression of genes involved in photosynthesis by inducing chlorophyll loss and cellular changes that cause less photochemical damage (Santino et al., 2013).

Proline plays several roles in plant responses to abiotic and biotic stresses, and under stress its metabolism is affected by multiple and complex regulatory pathways which can profoundly influence cell death and survival in plants (Zhang and Becker, 2015). The slight increase in Pro observed in WS plants compared to WW likely indicates its role as an osmoprotectant. By the same token the O₃-induced increase in Pro observed in WW+O₃ plants only during the first hours of treatment and coinciding with the maximal ABA, SA and JA stimulation and the first increase in ET, H₂O₂ and O2–, suggests a potential cross-talk among signaling molecules in regulating Pro metabolism, as previously reported by Rejeb K.B et al. (2014). The role of Pro as an ROS scavenger has also been reported (Szabados and Savouré, 2009; Banu et al., 2010; Zhang and Becker, 2015). In WW+O₃ plants the lack of accumulation in Pro at 8 h FBE was concurrent with a strong increase in H₂O₂, whereas in WS+O₃ the huge increase in Pro at 8 h FBE suppressed the increase in H₂O₂ (which remained at the same levels shown at 1 h FBE), thus suggesting an H₂O₂-scavenging role in these water conditions. This mechanism was also confirmed at 24 h FBE. Several studies have attributed an antioxidant feature to Pro, suggesting the capability of Pro in O2– and H₂O₂ quenching (e.g., Szabados and Savouré, 2009; Wang et al., 2009).