Effects of high salinity irrigation on growth, gas-exchange, and photoprotection in date palms (Phoenix dactylifera L., cv. Medjool)

Environmental and Experimental Botany 99 (2014) 100–109

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Effects of high salinity irrigation on growth, gas-exchange, and photoprotection in date palms (Phoenix dactylifera L., cv. Medjool) Or Sperling a , Naftali Lazarovitch a , Amnon Schwartz b , Or Shapira b,c,∗ a The Wyler Department of Dryland Agriculture, French Associates Institute for Agriculture and Biotechnology of Drylands, J. Blaustein Institutes for Desert Research, Ben-Gurion University of the Negev, Israel b The Robert H. Smith Institute of Plant Sciences and Genetics in Agriculture, Faculty of Agriculture, Food and Environment, The Hebrew University of Jerusalem, Rehovot, Israel c Northern R&D, Migal, Israel

a r t i c l e

i n f o

Article history: Received 1 August 2013 Received in revised form 15 October 2013 Accepted 18 October 2013 Keywords: Irrigation Palms Photoprotection Photosynthesis Salinity Solar irradiance

a b s t r a c t Date palms are widely cultivated in arid Mediterranean regions and require large quantities of water to produce commercial fruit yields. In these regions the plantations are commonly irrigated with lowquality water, which results in reduced growth and yields. To study the effect of using saline water for irrigation, date palm seedlings (cv. Medjool) were subjected to long-term irrigation treatments with water containing between 2 and 105 mM NaCl. The effect of saline irrigation was determined according to leaf gas exchange, chlorophyll a fluorescence, growth parameters and the distribution of key minerals in different plant organs. High salinity decreased plant growth and increased Na+ accumulation in the roots and lower stem. However, Na+ ions were mostly excluded from the sensitive photosynthetic tissues of the leaf. Thus, the reduction in the CO2 assimilation rate was primarily attributed to a reduced stomatal conductance. Consistent with this finding, the photosynthetic response to variable intercellular CO2 concentrations (A/Ci curves) revealed no permanent damage to the photosynthetic apparatus and implicated developed photoprotective mechanisms. Independent of salinity treatment, 80% of the energy absorbed by the leaf was directed to non-photochemical quenching, as presented in electronequivalent units. Functioning at full capacity, the non-photochemical mechanism could not compensate for all the excess irradiance. Thus, of the remaining absorbed energy, a significant portion was directed to photochemical O2 related processes, rather than CO2 prevented photoinhibition. The exclusion of toxic ions and O2 -dependent energy dissipation maintained photosynthetic efficiency and supported survival under salt stress. © 2013 Elsevier B.V. All rights reserved.

1. Introduction Date palms are more adapted to salinity stress compared to most cultivated trees with a threshold of 4 dS m−1 and a reduction of 3.6% yield per unit of electrical conductivity [EC (dS m−1 )] (Maas and Hoffman, 1990). Notwithstanding, salinity stress is usually associated with growth inhibition and yield reduction (Tester, 2003; Tripler et al., 2011) due to the osmotic effect on water uptake, reduced water conductivity of the roots, disrupted ion homeostasis in cells, inhibited metabolism, damaged membranes, and divergence of energy to salt-protection (Greenway and Munns, 1980; Frans et al., 1996; Tester, 2003; Tripler et al., 2011). Date palm

∗ Corresponding author at: The Robert H. Smith Institute of Plant Sciences and Genetics in Agriculture, Faculty of Agriculture, Food and Environment, The Hebrew University of Jerusalem, Rehovot, Israel. Tel.: +972 505469004. E-mail address: [email protected] (O. Shapira). 0098-8472/$ – see front matter © 2013 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.envexpbot.2013.10.014

growth is inhibited by irrigation salinity (commonly expressed in electric conductivity units – EC) above 9 dS m−1 , the yield is reduced by half (EC50 ) at a salinity of 18 dS m−1 (Alrasbi et al., 2010), and leaf elongation is suppressed at a salinity over 4 dS m−1 (Tripler et al., 2011). However, these findings are in contrast to previous studies performed by Furr and Armstrong (1962), who reported insignificant changes caused by 24 dS m−1 irrigation salinity. Most salt-resistance mechanisms reported previously included the exclusion of toxic ions (e.g., Na+ and Cl− ) from highly sensitive tissues (e.g., photosynthetic mesophyll) (Tester, 2003). Toxic solutes may be retained in the roots, or directed to alternative accumulation sites, such as the parenchymatous tissue or leaf margins (Stelzer, 1981; Shapira et al., 2009). Because Na+ toxicity is primarily caused by competition for K+ -binding sites in the mesophyll, a major protective factor is high K+ concentration in the shoots, which is expressed as a high K+ -to-Na+ ratio (K:N) (Tester, 2003). Na+ can be disposed from the roots back into the soil via plasma membrane Na+ /H+ anti-ports, or compartmentalized in the

O. Sperling et al. / Environmental and Experimental Botany 99 (2014) 100–109

cell vacuoles of insensitive tissues. With regard to solute accumulation in date palm tissues, Furr and Armstrong (1962) found no increase in Na+ or Cl− concentrations in the leaf matter of date palms, whereas Aljuburi and Al-Masry (1996) reported a correlation between excess concentrations and soil water salinity. More recently, Tripler et al. (2007) found elevated Na+ and Cl− concentrations in the roots of date palms irrigated with saline water. However, these differences were not evident at the leaf level, where the solute concentrations were remarkably low. The CO2 assimilation rates and cell growth are among the main processes inhibited by drought or salinity stress (Chaves, 1991). This may be due to direct limitations (e.g., reduced stomatal and mesophyll conductance) or secondary effects, such as oxidative stress and damage of photosynthetic pigments and proteins (Flexas et al., 2004). Stomatal restriction is a major photosynthesis inhibitor under drought conditions (Cornic and Briantais, 1991), yet nonstomatal limitations such as mesophyll conductance (Chaves et al., 2009) or a reduction in photosynthetic capacity (Tezara, 2002) contributes to a reduction in CO2 assimilation in some cases. Long-term water stress combined with high solar irradiance can result in photoinhibition (i.e., rate of photodamage exceeds the repair process capacity) of the photosynthetic system (PS) and severe damage to the plant. However, established plants growing in their natural habitat will, in most cases, recover from photoinhibition overnight. Thus, decreased photosynthesis is not necessarily associated with permanent physiological damage (Anderson et al., 1997; D’Ambrosio et al., 2006). Photoprotection in leaves is attributed to either photochemical processes [e.g., photorespiration, Mehler reaction or cyclic electron transport around PSI (Badger et al., 2000)] or non-photochemical processes. Nevertheless, on a larger scale, a high capacity for electron transport will better protect plants from photodamage compared to inhibition of the electron transport chain (Ort and Baker, 2002). Photorespiratory O2 uptake via Rubisco or the Mehler reaction promotes non-assimilatory electron transport and stimulates photon utilization under CO2 -limited conditions and bright light. Thus, photoprotection mainly affects the alternate electron sinks and thermal dissipation of excess photons (Niyogi, 2000). Furthermore, photoprotection mechanisms are dependent on environmental conditions. Low temperatures can enhance non-photochemical quenching (NPQ), similar to non-saturating light conditions (Ort and Baker, 2002), whereas higher temperatures induce photorespiration (D’Ambrosio et al., 2006). Several studies consider photorespiration a wasteful process (Peterhansel et al., 2010); nevertheless it was found to be an electron sink under drought and stress conditions (Wingler et al., 2000), making it an effective excess energy eliminator. Photoprotection has been reported to minimize the reduction in photosynthesis and enable overnight recovery of chlorophyll fluorescence in the Scheelea palm under high-light conditions (Araus and Hogan, 1994). Date palms are widely cultivated in the arid regions of the Mediterranean and are of essential economic value in these harsh environments. In Israel, date palms are predominantly cultivated in the Arava Valley; a hyper arid region where temperatures reach 42 ◦ C, relative humidity is usually low (15%), summer midday air vapor deficits (VPD) averages at 5 kPa, and the annual precipitation is below 25 mm. Moreover, palm plantations in these regions require over 2000 mm y−1 to produce commercial yields (Tripler et al., 2011) delivered by irrigation. Thus, date palm cultivation in these regions is applicable due to the palm tolerance to salinity (Maas and Hoffman, 1977) and the extensive use of marginal water in irrigation. This study was undertaken with two hypotheses in mind: (1) the salinity-tolerance of date palms is attributed to control of uptake, distribution, and accumulation of solutes in the sensitive photosynthetic tissues of the plant; and (2) under low

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stomatal conductance and high solar-irradiance the photosynthetic system is further protected from photoinhibition by non-photochemical and O2 -dependent photochemical mechanisms. The objectives of this study were to (1) describe the irrigation-water solute distribution in date palms and its effect on plant growth and photosynthesis; (2) differentiate between osmotic and toxic stress; and (3) investigate the protective mechanisms contributing to the date palm’s durability under low water qualities. 2. Materials and methods 2.1. Experimental facility and treatments The experiment was performed between May and October 2011, inside a greenhouse at the Faculty of Agriculture, Food and Environment of the Hebrew University of Jerusalem (Rehovot, 31◦ 54 N, 34◦ 48 E). The greenhouse was completely covered and plants only received water from irrigation throughout the experiment. Measurements were performed during the summer months, when the air temperature inside the greenhouse peaked at approximately 40 ◦ C at midday (occasionally) and the relative humidity (RH) averaged approximately 45%. Sixteen 2-year-old date seedlings (Phoenix dactylifera L., cv. Medjool) were planted in 10-L pots filled with highly porous organic planting soil and placed on 1-m high metal mesh tables, thus allowing maximum drainage. Irrigation was provided via 2 L h−1 compensated drippers (Netafim, Tel Aviv, Israel), with two drippers per pot. The final irrigation solutions were prepared in advance in 1-m3 tanks connected to a 16-mm irrigation pipe by a filter and the pump was controlled by electric timers. The irrigation quantities were set by weighing the drainage water collected in plastic tubs, which was placed beneath the pots. A leaching factor of 0.3 (drainage mass divided by irrigation mass) was maintained throughout the experiment, thereby avoiding water stress and ensuring adequate leaching of the root zone. Date seedlings were subjected to four irrigation salinities, which represented the range of possible water qualities of municipal recycled wastewater used for irrigation in Israel. To distinguish between the osmotic and salt-specific effects, NaCl was used to generate the salinity levels. The treatments contained 2 (control), 30, 75 and 105 mM NaCl. All of the plants received fertilizer at a concentration of 0.5 g L−1 (Multi NPK, Haifa Chemicals Ltd., Israel). During the measurements, the plants were deliberately exposed to two environmental conditions in the laboratory; VPDs of 1.5 and 4 kPa, which represented the winter and summer conditions of the major cultivation regions, respectively. The external conditions were 28 ◦ C and 50% RH for the low VPD, and 38 ◦ C and 20% RH for the high VPD. Acclimation of the sample plants to the environmental conditions was obtained after 24 h of pre-exposure. 2.2. Gas-exchange and fluorescence measurements Hourly measurements of stomatal conductance were performed throughout the entire day (10 measurements a day on average) every 7th day using a null-balance porometer (LI-1600, LiCor Inc., Lincoln, NE, USA). The measurements were performed under greenhouse conditions and on leaves that were exposed to direct sunlight. Gas-exchange measurements were performed using the Li-6400 portable photosynthesis and fluorescence system (6400-40 leafchamber fluorometer; LiCor Inc.). The measuring chamber enclosed a circular 2-cm2 leaf area and evaluated the gas fluxes on both sides of the leaf. The air-flow rate was maintained at a constant rate of 500 ␮mol s−1 and the reference CO2 concentration was

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400 ␮molCO2 mol−1 air (ppm). The light intensity was detected prior to each set of measurements and kept constant (10% blue light). Although the trees were grown in the greenhouse, the measurements were performed under laboratory conditions (plants were moved on a trolley), thus enabling the instrumentation to perform with maximum accuracy and provided control of the environmental conditions. The gas exchange and fluorescence performance were measured at ambient O2 concentration (i.e., 21%) and at nonphotorespiratory conditions of 2% O2 . Low O2 concentrations were obtained by connecting the sensor to a 2% O2 pressure tank. The ingoing air passed through a water-saturated air chamber to restore the ambient humidity conditions. The plant response to varying intercellular CO2 concentrations, commonly referred to as A/Ci curves, was examined for two irrigation treatments; 2 and 105 mM NaCl. The measurements were performed at the saturating light intensity of 1600 ␮molphotons m−2 s−1 , with an air-flow of 500 ␮molair s−1 , and constant leaf temperature (27 ◦ C). The initial CO2 concentration was 400 ppm and was reduced to 300, 200, 100, and 50 ppm; the readings were taken immediately upon stabilization of the concentration to allow rapid measurements. Fast measurements were particularly important at the lower CO2 concentrations to avoid significant changes in Rubisco activity due to low photosynthetic rates (Long and Bernacchi, 2003). The CO2 concentration was then returned to 400 ppm and increased to 600, 800, 1200, and 1600 ppm. Leaf photosynthesis–irradiance response curves were determined for two irrigation treatments: irrigation salinities of 2 and 105 mM. Repeating the measurements under both ambient and non-photorespiratory conditions, i.e., 2% oxygen in total air inflow, produced the quantum yield to electron transport calibration (Gilbert et al., 2011). Response curves were determined under ambient CO2 conditions (400 ppm) and a constant air-flow of 500 ␮molair s−1 . The light intensities were 50, 200, 400, 700, 1000, 1500, and 2000 ␮molphotons m−2 s−1 . Readings were taken after the conditions had stabilized with the limiting factor, i.e., stomatal conductance, reaching a steady state only after 20–30 min. The plants were subjected to long periods of darkness (approximately 1 h) to measure the basic fluorescence (F0 ), followed by a single light-saturating flash for the maximum fluorescence reading used to calculate the ratio of variable to maximum fluorescence (Fv /Fm ) and the non-photochemical quenching (NPQ) according to Stern–Volmer.

2.3. Leaf water potential The leaf water potential was measured at the final stage of the experiment on September 29, 2011. The pressure was measured in a pressure chamber (Model 600, PMS Inc., Albany, OR, USA) at midday. The leaves were covered 1 h prior to cutting with a bag that was impermeable to water vapor and light, thus allowing for water equilibrium with the whole plant and avoiding water loss at cutting. The pressure chamber’s aperture was specific for wide leaves, such as palm leaves, thus enabling maximum sealing.

2.4. Plant growth and dry mass On October 20, 2011, after all the physiological measurements were performed and the growing season was over, the plants were cut down. Each plant was separated into its different parts – roots, stems, leaf axes, and leaflets. Each of the plant parts was weighed immediately after cutting (fresh mass) and packed into a paper bag. The samples were dried in a 65 ◦ C oven for 3 days prior to reweighing for dry mass.

Table 1 Parameters described in Sharkey et al. (2007) for fitting A versus Ci reaction curves of C3 plants and the partitioning of stomatal and non-stomatal limitations to photosynthesis.

Salinity Vcmax J TPU Ls Lns KC KO *

Units

Control

High salinity

Comments

mM ␮mol m−2 s−1 ␮mol m−2 s−1 ␮mol m−2 s−1 % % Pa Pa Pa

2 145 156 11.77 22

105 127 148 11 29 9 33.86 17.67 4

Treatments P < 0.167 P < 0.255 P < 0.213

33.86 17.67 4

Predetermined variables of the model

2.5. Mineral allocations After the plants were cut and separated into different parts, the samples were obtained for chemical analyses. The samples were rinsed twice in tap water, once in HCl solution (1 mL L−1 ), and finally in double-distilled water. Next, the samples were ovendried, ground in an electric mill, weighed out into 150-mg portions, and digested with 5 mL of reagent-grade nitric acid at 130 ◦ C. The volume of the digest was brought to 50 mL with distilled water and stored in the dark at 4 ◦ C. The Na+ and K+ concentrations were measured using inductively coupled plasma-atomic emission spectroscopy (ICP-AES, Arcos, Spectro, Kleve, Germany). The Cl− ions were extracted independently from the dry material using doubledistilled water and filtering, and measured using a chloridometer (chloride analyzer 926, Corning, Medfield, MA, USA). 2.6. Theory and calculations The maximum carboxylation rate by RuBP (Vcmax ), the rate of photosynthetic electron transport (Jmax ), triose phosphate use (TPU), and day respiration (RD ) were determined using the A/Ci curve-fitting utility (Sharkey et al., 2007). This utility requires 5 pairs of A/Ci values and an educated assessment of the carboxylation-limiting factor for Sharkey’s equation: A = Vcmax





CC −  ∗ − RD CC + KC (1 + (O/KO ))

(1)

where CC is the partial CO2 pressure of Rubisco for carboxylation, KC is the Rubisco Michaelis constant for carbon dioxide, O is the partial pressure of O2 at the Rubisco, KO is the Rubisco O2 inhibition constant, and  * is the photosynthetic compensation point (the constants were fitted using the model, see Table 1). The stomatal limitation to photosynthesis was calculated as previously described by Farquhar and Sharkey (1982): Ls =

ACi =400 − ACa =400

(2)

ACi =400

where ACa =400 and ACi =400 are the CO2 assimilation rates under external and internal CO2 concentrations of 400 ppm, respectively. Non-stomatal limitations to photosynthesis were evaluated according to the following equation: Lns =

A2 mM NaCl, Ci =400 − A105 A2

mM NaCl, Ci =400

(3)

mM NaCl, Ci =400

where A2 mM NaCl, Ci =400 and A105 mM NaCl, Ci =400 are the photosynthetic rates at the internal CO2 concentrations of the control and high salinity treatments, respectively (Matthews and Boyer, 1984). The proportion of non-cyclic electron transport directed to O2 dependent processes is represented by Pdiss (Peterson, 1990): Pdiss =

˚s − ˚s ˚s

(4)

JT = JC + JO .

500 Dry Weight (g)

where ˚s and ˚s are the observed quantum efficiencies of CO2 fixation at O2 concentrations of 2% and 21%, respectively. Partitioning of the total electron flow (JT , ␮molelectrons m−2 s−1 ) to carboxylation and oxygenation cycles (JC and JO , respectively, ␮molelectrons m−2 s−1 ) driven by Rubisco was calculated as previously described by Valentini et al. (1995). Assuming that all other light-consuming processes are either constant or negligible (Cornic and Briantais, 1991), the distribution of electron flow is as follows: (5)

JT = ˚PS2 PARi 0.5˛

(6)

where PARi is the photon flux density (Valentini et al., 1995) and ˛ = 0.84 is the leaf absorbance (D’Ambrosio et al., 2006). The quantum yield of PSII (PS2 ) was calculated according to: ˚PS2 =

 −F Fm t  Fm

1 [J 3 T

+ 8(A + RD )]

(8)

2 [J 3 T

− 4(A + RD )]

(9)

where A is the net CO2 assimilation rate and RD is the mitochondrial day respiration rate (Valentini et al., 1995). JT was measured using the fluorescence detector while A and RD were assessed using the gas-exchange sensor. Valentini’s (Valentini et al., 1995) analysis was adequate for this study as confirmed by the calibration between the photosynthesis rate and electron transport under nonphotorespiratory conditions. The electron equivalent rate of xanthophyll-regulated and pH energy dissipation, JNPQ , was calculated as previously described (Hendrickson et al., 2004): JNPQ =

F

s  Fm

−

Fs Fm



PARi 0.5˛

(10)

 are the dark and light acclimated maximum where Fm and Fm chlorophyll fluorescence, respectively. Fs is the steady-state lightacclimated chlorophyll fluorescence yield, and PARi is the external light intensity. The apparent in vivo specificity factor of Rubisco (S) was derived as previously described (Valentini et al., 1995):

S=

0.8

300

0.6

200

0.4

100

0.2 20

40 60 80 100 Irrigation [NaCl] (mM)

0

Fig. 1. The final plant dry mass (filled circles) and root-to-shoot ratio (empty circles) at the end of the experiment. Each data point represents the average ± SE of 4 plants per treatment. The regression significance is marginal (P < 0.0668).

light curves (e.g., Pn to I) were fitted to the equation using ‘best fit’ algorithms in MATLAB software.

The experiment was randomly arranged. Linear and non-linear regression analyses were performed using MATLAB. The analysis of variance (one-way ANOVA) for significant variations in response to irrigation salinity was assessed and means were separated according to the Tukey–Kramer HSD (P < 0.05) test using JMP 5 (SAS Institute Inc.). Separations of the means for two salinity treatments were analyzed using Student’s t-test (P < 0.01) in the A/Ci curves and one-way ANOVA (P < 0.05) in the light response curves, with the introduction of the VPD factor. 3. Results 3.1. Vegetative growth High salinity irrigation dramatically reduced the growth of the date palm saplings. The total dry mass of the salinity-induced plants decreased by ca. 40%, and averaged 500 g for control plants versus 300 g for 105 mM NaCl treated palms (Fig. 1). The dry mass reduction is described by the linear regression: DW = −1.7 IS + 472 (r2 = 0.87, P = 0.00668); where DW represented the total dry mass (g) and IS represented the irrigation salinity [NaCl] (mM). In contrast, the root-to-shoot ratio (i.e., root mass divided by the aboveground plant mass) exhibited no significant differences between treatments; approximately 33% of the total dry mass was allocated to the root zone.

(11)

where c (CO2 mole fraction at the carboxylation site) and o (O2 mole fraction at the oxygenation site) are assumed to equal Ci and O2 mole fraction in air, respectively, multiplied by the water solubility ratio. The light response curves were analyzed using a nonrectangular hyperbolic model (Marshall and Biscoe, 1980; Lieth, 1990): Pn =

Root : Shoot

3.2. Stomatal conductance

JO /JT c/o

˛I + Pm −

1

2.7. Statistical analyses

and JO =

Dry Weight

(7)

 is the maximal where Ft is the steady-state fluorescence and Fm fluorescence in the light-adapted state. The two flows can then be derived as:

JC =

400

0 0

The electron transport rate was derived from the fluorescence measurements (␮molelectrons m−2 s−1 ):

r2=0.87

103

Root : Shoot (-)

O. Sperling et al. / Environmental and Experimental Botany 99 (2014) 100–109



(˛I + Pm )2 − 4eIPm  − RD 2

(12)

where Pn is the net photosynthesis rate (␮molCO2 m−2 s−1 ), I is the light intensity (␮molelectrons m−2 s−1 ), Pm is the photosynthetic rate under saturated light conditions (␮molphotons m−2 s−1 ), e is the photosynthetic efficiency (␮molCO2 m−2 s−1 ),  is the convexity (–), and RD is the leaf’s day respiration (␮molCO2 m−2 s−1 ). The

The stomatal conductance in date palms varied throughout the day; it increased during the morning hours, peaked at approximately 13:00 h, and gradually decreased throughout the afternoon (Fig. 2). At 14:00 h, there was a temporal reduction in stomatal conductance, which is commonly referred to as midday depression. However, the stomatal conductance varied significantly between salinity treatments at all times except at 18:00 h. In the morning, i.e., from 08:00 to 10:00 h, the stomatal conductance was inversely related to irrigation salinity; i.e., 0.4(A), 0.35(AB), 0.3(BC), and 0.2(C) molH2 O m−2 s−1 , for 2, 30, 75, and 105 mM, respectively. The uppercase letters indicated significant differences between treatments at P < 0.05 (Tukey HSD). Throughout the rest of the day, the plants subjected to irrigation salinity of 30 mM exhibited a higher stomatal conductance compared to the control plants. However, as the salinity increased over 30 mM, the stomatal conductance decreased. Thus, at midday, the stomatal conductance was 0.5(AB), 0.6(A),

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O. Sperling et al. / Environmental and Experimental Botany 99 (2014) 100–109

(A)

1.5 B A A A

Na+ (mol kg-1)

gs (molH2O m-2 s-1)

0.6

0.4

0.2

0

2 30 75 105 (mM) NaCl

8

10

12 14 16 Time of Day (h)

1

0.5 B AB AB A

C B B A

18 B AB A A

0

3.3. Mineral analysis The Na+ and Cl− concentrations differed between the plant compartments despite their equal concentrations in the irrigation solution (Fig. 3). Na+ ions were found at very high concentrations in the roots, reaching 1 mol kg−1 in the 75 mM- and 105 mM-treated plants (Fig. 3A). However, in the stem and leaf axis, the Na+ concentration decreased by 75% with a clear correlation to water salinity. Finally, in the leaflet, the Na+ concentration decreased to negligible values, averaging at 0.05 mol kg−1 . The Cl− concentration was significantly elevated (P < 0.05) due to irrigation salinity in the root zone alone, and averaged at 0.6 mol kg−1 and 1.4 mol kg−1 for the 2 mM- and 75 mM-treated plants, respectively (Fig. 3B). In the canopy, the Cl− concentrations were high and did not differ between treatments; 0.4 mol kg−1 in the stem, 0.7 mol kg−1 in the leaf axis, and 0.3 mol kg−1 in the leaflet. The ratio between the K+ and Na+ concentrations (K:Na) varied inside the plant (Fig. 3C); it was as low as 0.5 in the roots due to high Na+ concentrations, except in control plants, where Na+ ions were scarce (K:Na = 5). Because Na+ ions were excluded from the plant canopy, the K:Na ratio increased in the stem and leaf axis to 10. However, the major increase in the K:Na ratio was at the leaflet where, due to nearnegligible Na+ concentrations, it averaged at 80. The K:Na ratio was inversely correlated to irrigation water salinity (P < 0.05) and was particularly high in the control plants (due to low Na+ levels), which reached 160 in the leaflets. 3.4. Leaf water potential The midday leaf water potential was correlated to stomatal conductance (Fig. 4) (r2 = 0.99, P < 0.001). The stomatal conductance exhibited a sensitivity to midday leaf water potential, decreasing from 0.6 to 0.2 mol m−2 s−1 with a minor loss in water potential (0.3 MPa). 3.5. A/Ci curves The CO2 assimilation rates varied with changes in the intercellular CO2 (Ci ) concentration, irrigation salinity, and external O2 concentration (Fig. 5). CO2 assimilation in high-salinity-treated plants (Fig. 5B) and under photorespiratory conditions (21% O2 ) was limited (P < 0.01) compared to the control plants (Fig. 5A).

Cl- (mol kg-1)

0.4(B), and 0.3(B) mol m−2 s−1 for salinities of 2, 30, 75, and 105 mM, respectively (P < 0.05).

(B)

1.5

A A A A

1 B A A A A A A A

.5

A A A A

0 2 (mM) NaCl 30 75 105

150

K+: Na+ (-)

Fig. 2. Hourly stomatal conductance (gs ) for the four irrigation salinity treatments: 2 mM (filled circles), 30 mM (empty circles), 75 mM (filled triangles) and 105 mM (empty triangles). Each data point represents the average ± SE of 4 plants per treatment (2 leaflets per plant). The statistical analysis revealed significant differences between treatments for all hours except 18:00 h at P < 0.05 (Tukey HSD; the significance marks are not shown for clarity).

(C)

100 A B B B

50 A B B B A B B B A B B B

0

Roots

Stem Leaf axis Leaflet

Fig. 3. Accumulation of Na+ (A), Cl− (B), and the K+ :Na+ ratio (C) in the different plant compartments. The uppercase letters indicate significant differences between treatments at P < 0.05 (Tukey HSD). Data presented for the four salinity treatments: 2 mM (black columns), 30 mM (dark gray), 75 mM (light gray), and 105 mM (white). Each column represents the average ± SE of 4 plants per treatment.

Nevertheless, as Ci approached 1000 ppm, both treatments averaged at 29 ␮molCO2 m−2 s−1 . However, under non-photorespiratory conditions (i.e., 2% O2 ), as Ci surpassed 600 ppm, the CO2 assimilation rates in the control plants were equal under ambient and non-photorespiratory conditions, while the saline-treated plants maintained a near constant differential in CO2 assimilation (4 ␮molCO2 m−2 s−1 ). The proportion of non-cyclic electron transport directed to O2 -dependent processes, Pdiss was significantly enhanced (P < 0.01) due to high salinity irrigation; a 0.18 increase between the control and 105 mM NaCl-treated plants for Ci concentration above 300 ppm (Fig. 6). Moreover, Pdiss of the saline-treated plants was not eliminated despite saturated CO2 concentrations. The A/Ci curve-fitting utility (Sharkey et al., 2007) only exhibited nearly significant differences in the maximum rate of carboxylation, and no significant differences in the electron transfer rate, or triose phosphate use due to irrigation salinity. Nevertheless, the

O. Sperling et al. / Environmental and Experimental Botany 99 (2014) 100–109

0.7

1

* * *

Pdiss (-)

2 (mM) NaCl 105 (mM) NaCl

0.6 0.4 0.2

2

gs (molH O m-2 s-1)

0.8 0.5

105

0.3

0 0

r2 = 0.99

500 1000 Ci (µmol mol-1)

1500

0.1 -1.4

-1.6

-1.8

Fig. 4. Midday (13:00) stomatal conductance and leaf water potential with significant linear regression (P < 0.001).

A/Ci curves showed that the stomatal limitation to photosynthesis increased from 22% in the control treatment to 29% in the salinetreated plants (Table 1). Finally, an additional 10% reduction in photosynthesis was attributed to non-stomatal limitations in the saline-treated plants (Table 1). 3.6. Light curves The stomatal conductance and CO2 assimilation rate response to light intensities (Fig. 7) showed a significant interaction with the external VPD and irrigation salinity (P < 0.05). At a VPD of 1.5 kPa, the stomatal conductance increased with light intensity equally in both treatments (Fig. 7B), reaching maximum rates of 0.25 molH2 O m−2 s−1 . However, at a VPD of 4 kPa, the salinity treatment had a significant effect (P < 0.05); the stomatal conductance of saline-treated plants ceased to increase for light intensities over 400 ␮molphotons m−2 s−1 , and maintained a mean value of 0.12 molH2 O m−2 s−1 (Fig. 7B). The induction of a single stress did not affect the stomatal conductance rates because the control plants maintained regular stomatal performance, despite elevated VPD (Fig. 7B). The CO2 assimilation rates in the plants exposed to

(A)

(B)

30

both salinity and VPD stress were significantly reduced (P < 0.05) (Fig. 7A). However, each of these stresses, when imposed alone, showed only a moderate and non-significant (P > 0.05) effect on CO2 . Fitted parameters of the non-rectangular hyperbolic model (Marshall and Biscoe, 1980; Lieth, 1990) for the light–response curves are presented in Table 2. According to this model, the photochemical efficiency (e) was not affected by either the VPD levels or the salinity treatments. However, the photosynthetic rate under saturated light conditions (Pm ) was reduced by the increases in VPD and irrigation salinity (P < 0.05). NPQ increased with light intensity, mainly due to excess irradiance (Fig. 8). For moderate light intensities, NPQ did not vary (P > 0.05) with irrigation salinities or VPD. However, irrigation salinity did have an effect above 1000 ␮molphotons m−2 s−1 ; NPQ of the 105 mM-treated plants increased at VPD of 1.5 kPa (P < 0.05). Nevertheless, at a VPD of 4 kPa, NPQ was repressed and did not differ between treatments (P > 0.05). The calculated non-photochemical electron-equivalent energy (JNPQ ) was over three times greater

gs (mol H O m-2 s-1)

2% O 2

105 (mM) NaCl

2 (mM) NaCl

5 0 0

15 10 5

* *

1.5 kPa, 2 mM 1.5, 105 4, 2 4, 105

*

*

0.2

2

A (µmol m-2 s-1)

21% O 2

15 10

(A)

0 0.3 (B)

25 20

20

2

-1.2

Midday Leaf Water Potential (MPa)

A (µmol CO m-2 s-1)

-1

Fig. 6. The proportion of non-cyclic electron transport directed to O2 -dependent, Pdiss , presented for two salinity treatments; 2 mM (filled symbols) and 105 mM (empty symbols) in response to increasing intercellular CO2 concentrations. Each data point represents the average ± SE of 4 plants per treatment (1 leaflet per plant). Significant differences are marked by asterisks (Student’s t-test; P < 0.01).

********* 500 1000

1500 500 Ci (µmol mol-1)

0.1 0 0

1000

1500

Fig. 5. The CO2 assimilation rates (A) of control plants (left panel) and 105 mM NaCl-treated plants (right panel) versus intercellular CO2 concentrations (Ci ) in photorespiratory conditions (21% O2 ; filled circles) and non-photorespiratory conditions (2% O2 ; empty circles). The photosynthetic data were fitted using Sharkey’s utility (Sharkey et al., 2007). Measurements were performed at an air-flow of 500 ␮mol s−1 , constant leaf temperature (27 ◦ C), and air relative humidity of ca. 50%. Each data point represents the average ± SE of 4 plants per treatment (1 leaflet per plant). Significant differences are marked by asterisks (Student’s t-test; P < 0.01).

*500 * 10*00 PPFD (µmol

photons

*

1500

*

2000

m-2 s-1)

Fig. 7. (A) Response of CO2 assimilation rates (A) and (B) stomatal conductance (gs ) to increasing light intensities. The control plants (filled symbols) and 105 mMtreated plants (empty symbols) under two atmospheric conditions are presented: VPD of 1.5 kPa (circles) and 4 kPa (triangles). Measurements were performed at CO2 concentration of 400 ppm and a constant air-flow of 500 ␮molair s−1 . The air temperatures were 28 ◦ C and more than 50% relative humidity (RH) for the low VPD, and 38 ◦ C with 20% RH for the high VPD conditions. Each data point represents the average ± SE of 7 measured leaflets. Measurements are accompanied by hyperbolic curves and significant differences are marked by asterisks (one-way ANOVA; Tukey HSD, P < 0.05).

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Table 2 Analysis of the light–response curve using a non-rectangular hyperbolic model (Marshall and Biscoe, 1980; Lieth, 1990) and the rate of xanthophyll-regulated and pH energy dissipation, JNPQ.

VPD IS e Pm  RD JNPQ PPFD=1500

2.5

Control, low VPD

High salinity, low VPD

Control, high VPD

High salinity, high VPD

kPa mM ␮mol m−2 s−1 ␮mol m−2 s−1 (–) ␮mol m−2 s−1 ␮mol m−2 s−1

1.5 2 0.242 28.2 −0.0338 1.2 326.5

1.5 105 0.299 24.6 −0.3244 1.3 349.3

4 2 0.323 24.8 −1.098 1.0 324.3

4 105 0.256 17.1 −0.0671 0.8 329.4

1.5 kPa, 2 mM 1.5, 105 4, 2 4, 105

2

NPQ (-)

Units

1.5 1 0.5

*

0 0

500 1000 1500 PPFD (µmolphotons m-2 s-1)

* 2000

Fig. 8. Response of NPQ to increased light intensities for the control plants (filled symbols) and 105 mM-treated plants (empty symbols) under two atmospheric conditions: VPD of 1.5 kPa (circles) and 4 kPa (triangles). Each data point represents the average ± SE of 4 plants per treatment (1 leaflet per plant). Significant differences are marked by asterisks (one-way ANOVA; Tukey HSD, P < 0.05).

than the total electron transport under saturated light intensities (Tables 1 and 2). According to Valentini’s (Valentini et al., 1995) analysis, the oxygenation portion of electron flow (JO /JT ) increased with light intensity (Fig. 9B), from 0.2 at low light intensity to 0.3 under

S (-)

120

1.5 kPa, 2 mM 1.5, 105 4, 2 4, 105

(A)

80 40

JO / JT (-)

0 0.4 (B)

*

*

*

*

*

*

0.3 0.2 0.1 0 0

250 500 750 1000 PPFD (µmolphotons m-2 s-1)

Fig. 9. Apparent in vivo specificity factor of Rubisco (A) and the oxygenation share of total electron flux (B) response to increasing light intensities. Measurements were performed on the control plants (filled symbols) and 105 mM-treated plants (empty symbols) under two atmospheric conditions: VPD of 1.5 kPa (circles) and 4 kPa (triangles). Each data point represents the average ± SE of 4 plants per treatment (1 leaflet per plant). Significant differences are marked by asterisks (one way ANOVA; Tukey HSD, P < 0.05).

saturated light conditions. While irrigation salinity had no significant effect (P > 0.05) on the JO /JT at a VPD of 1.5 kPa, it increased the oxygenation portion of electron flux by ca. 20% at VPD of 4 kPa and saturating light intensities (P < 0.05). The Rubisco specificity for CO2 assimilation (S) was reduced under high light intensity, decreasing from an average value of 100 (no units) at low light conditions to 70 at saturated light intensity (Fig. 9A). Up to an irradiance of 600 ␮molphotons m−2 s−1 , Rubisco specificity (S) was only VPD reliant; decreasing by 20% at a VPD of 4 kPa. However, at saturated light intensities, S exhibited an additional 10% reduction due to irrigation salinity at both VPD conditions. 4. Discussion Salt tolerance is a complex trait involving the regulation of numerous physiological and biochemical mechanisms (Munns, 2002; Chaves et al., 2009; Shabala et al., 2010). Despite being a nonhalophytic species, date palm varieties are considered salt-tolerant. Previous studies have even suggested that date palms can withstand seawater irrigation (Alhammadi and Kurup, 2012). Thus, the present study examined the effect of saline irrigation on 2-year-old, greenhouse-grown, potted date palm (cv. Medjool) seedlings. The irrigation salinity levels represented the range of water qualities used for irrigation in Israel. High salinity irrigation affects 2-year-old date palm saplings; it restricts plant growth and results in a major loss in the dry mass of trees irrigated with 105 mM (NaCl) solution (Fig. 1). This is consistent with findings obtained from previous studies in dates (Aljuburi and Al-Masry, 1996; Tripler et al., 2011), tomatoes (Karlberg et al., 2006), olives (Chartzoulakis, 2005), citrus (Garcia-Legaz et al., 1993) and avocado (Bar et al., 1997). However, salinity, which is commonly associated with toxic effects and damage to the photosystem (PS), does not cause permanent damage or efficiency loss in date palms. Typical symptoms of damage by salinity, e.g., chlorosis of the leaf margins or tip burns (Kozlowski, 1997; Shapira et al., 2009), were not evident in any of the salinity treated plants. In fact, a large proportion of the solutes did not penetrate the plants (Fig. 3). Solute accumulation in the leaves is largely dependent on the ability of the roots to exclude ions and in the compartmentalization of salts in the root and stem tissues. Na+ , which is more toxic of all the components (Tester, 2003), is accumulated mainly in the root system and excluded from the sensitive leaflet tissues of the date palms (i.e., 0.05 mol kg−1 for the highest salinity treatment). In contrast, Cl− is allocated to all plant parts, independent of salinity treatment; thus indicative of an elevated efficiency for Na+ exclusion, and indicating that Cl− was below hazardous concentrations to date palms in this research. However, previous field studies performed by Tripler et al. (2007) or Alrasbi et al. (2010) found higher leaf Na+ concentrations (0.23 and 0.11 mol kg−1 , respectively). These differences were most likely due to the shorter period of examination in the present research [9 months compared to 7 years (Tripler et al., 2007) and 2 years (Alrasbi et al., 2010)], and the reduced water use imposed by the internal greenhouse micro-climate. However, elevated Cl− concentrations in the leaf, which was previously thought

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to be destructive to trees (Bar et al., 1997), was reported to cause no damage to bananas (Shapira et al., 2009), salt-tolerant olive cultivar (Kalamanta) (Chartzoulakis, 2000), and date palms (Tripler et al., 2007; Alrasbi et al., 2010). Nevertheless, a 0.7 mol kg−1 increase in root Na+ concentration, due to high salinity treatments, indicated that Na+ is compartmentalized in the roots. K+ is a major cationic inorganic nutrient and is essential to all plants. Due to the physicochemical similarities between Na+ and K+ , under conditions of high salinity, Na+ ions tend to substitute for K+ ions on its binding sites. Thus, in the cytoplasm, a K:Na ratio of 1 provides a minimum value for enzymatic activity (Frans et al., 1996; Maathuis, 2006). In the current study, the leaflets’ higher K:Na ratio (28) meets the requirement for high K+ concentrations around the mesophyll tissues, enabling optimal PS functionality (Tester, 2003). Such high K:Na ratios have been previously reported in saline-irrigated wheat (Genc et al., 2007), cotton (Brugnoli and Björkman, 1992), and grapevines (Downton et al., 1990). Taken together, the major losses in date palm growth under high salinity irrigation are attributed to osmotic stress rather than to solute toxicity. The reduced growth of date palms under saline irrigation is partially attributed to repressed stomatal conductance (Fig. 2). Stomatal limitation to photosynthesis increased from 22% to 29% for the 2 and 105 mM treatments, respectively; thus, corresponding to previous studies on date palms (Youssef and Awad, 2007). However, this 7% shift in stomatal regulation caused by salinity irrigation could not account for all the co-regulations of stomata and PS (Escalona et al., 2000); the stomatal limitations should be higher (Chaves, 1991). Osmotic adjustments for maintaining leaf water potential and turgid pressure under minor solute stress could potentially explain the elevated stomatal conductance demonstrated with the 30 mM-NaCl-treatment (Morgan, 1984; Katerji et al., 1997). Nevertheless, a major reduction in stomatal conductance at high salinities (75 and 105 mM) reflects the high osmotic stress experienced by these plants. The difference in the midday leaflet water potential between the control and high salinity treatment was only 0.3 MPa while the range in midday stomatal conductance between these two treatments was considerably broader (0.43 mol m−2 s−1 ) (Fig. 4). Thus, osmotic stress is controlled by the high sensitivity of stomatal conductance to minor changes in the water potential, which is commonly referred to as isohydric behavior (Flexas, 2002). An in situ example is the deviation from the common daily pattern of the stomatal apparatus (where conductance increases throughout the morning and gradually decreases throughout the afternoon), as conductance temporally decreases at approximately 14:00 h. This stomatal closure is caused by the high leaf-to-air water vapor difference and elevated temperature present at this time of the day (Roessler and Monson, 1985). This physiological regulation of the stomata should be chemical and hormonal (e.g., root-to-shoot signaling) as well as hydraulic (Schachtman and Goodger, 2008). Stomatal sensitivity also indicates minimal water reservoirs in the plant due to its premature stage, i.e., nearly no developed stem tissues (Holbrook and Sinclair, 1992a,b). Increased air vapor deficits and solar radiation, which is commonly associated with the harsh environmental conditions of the date’s cultivation regions (VPD exceeds 5 kPa on summer middays), enhances the plants’ physiological response to high salinity irrigation (Fig. 6). As the stomatal conductance and CO2 assimilation rates reach a threshold value for saturating light intensities, most of the absorbed irradiance is directed to non-photochemical processes (Tables 1 and 2). Thus, the role of non-stomatal limitations to photosynthesis is substantial under high salinity irrigation. However, the efficiency of energy removal by the non-photochemical pathway (NPQ) decreased under summer day desert conditions (i.e., high temperatures and light intensities) (Ort and Baker, 2002; D’Ambrosio et al., 2006; Eppel et al., 2013).

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Thus, this harmless thermal dissipation is at peak performance in the Arava region date palms, independent of the salinity treatments (Fig. 8), and thus, the excess energy is dissipated in an alternative photochemical process, as previously proposed by Brugnoli and Björkman (1992) in their similar findings on saline irrigated cotton. Drought-adapted cultivars are commonly characterized by major stomatal limitations to photosynthesis (Chaves, 1991); however, non-stomatal limitations are substantial in date palms. These limitations do not result in permanent damage of the PS in date palms due to potential carboxylation (Table 1), and the electron transfer rates (Fig. 5) are independent of irrigation water salinity. However, the CO2 assimilation rates under non-photorespiratory conditions indicated that the proportion of non-cyclic electron transport directed to O2 -dependent processes increased from 20% to 40% between the control and 105 mM treatments (Fig. 6). These processes, which were previously attributed to photorespiration (Sharkey et al., 1986), have been reported to directly consume 30% of the total electrons (Lovelock and Winter, 1996; Badger et al., 2000). Moreover, palms acclimated to saline irrigation demonstrated a significantly enhanced oxygenation portion of total electron flow under high VPD (Fig. 9B) and reduced Rubisco specificity for CO2 assimilation (Fig. 9A). The shift in CO2 to O2 assimilation ratio should not be solely attributed to changes in solubility with temperature, but to enhanced photorespiration as well (Brooks and Farquhar, 1985). Such a Rubisco-supported alternative photochemical acceptor is efficient only in C3 higher plants (Badger et al., 2000), similar to the date palm. Thus, non-stomatal limitations to photosynthesis should not be related only to photodamage, but may rather enhance the alternative photochemical apparatus competing with photosynthesis. This indicates that the total electron flow should not be reduced by stressful conditions, as demonstrated in grapevines (Flexas et al., 1998; Hochberg et al., 2012) and cotton (Brugnoli and Björkman, 1992). Because O2 dependent systems use excess amounts of electrons (compared to CO2 -based systems), they function as a photoprotective mechanism under extreme light conditions and repressed photosynthetic performance (Badger et al., 2000; D’Ambrosio et al., 2006). Thus, O2 -dependent photochemical activity and the accumulation of anthocyanin differs between desert-adapted and Mediterraneanclimate ecotypes of Hordeum spontaneum, the latter demonstrating elevated NPQ (Eppel et al., 2013). Moreover, photorespiratory enzymes (e.g., glycolate oxidase) are stimulated in the presence of NaCl while CO2 assimilation is suppressed in salt-tolerant plants (Garratt and Janagoudar, 2002). Finally, the linear relationship between the total electron flow and CO2 fixation, with a slope of 0.25 (r2 = 0.94, data not shown), indicates that no alternative electron sinks other than those related to O2 (Krall and Edwards, 1992) contribute substantially to date palm tolerance to salinity.

5. Conclusions High salinity irrigation water reduces the growth of date palms. However, toxic Na+ ions are excluded from the sensitive photosynthetic tissues, and K+ ions remain the abundant ion in the vicinity of chlorophyll. Thus, permanent toxic damage to the PS is avoided and optimal PS performance is attainable. Because osmotic stress affects the plant, the photosynthetic productivity is determined by the water potential imbalance. The date palm encounters water stress, forcing stomatal limitations on photosynthesis, particularly under high atmospheric water demands. The reduced photosynthetic performance further exposes the date palm to excess solar irradiance and necessitates efficient photoprotection mechanisms. However, non-photochemical energy dissipation is inefficient in the extreme habitat of date cultivation, and alternative photochemical pathways are necessary. Thus, oxygen contributes to sustaining the

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required electron flow for carbon metabolism, acting as an alternative electron sink under stressful conditions, and is highly efficient in salt-tolerant species. Oxygen-dependent electron flow is elevated in date palms exposed to solute stress and high atmospheric water demands. These enhanced photoprotective mechanisms are crucial for the durability of date palms to avoid solute toxicity and photoinhibition caused by photosynthetic restrictions (stomatal and non-stomatal), extreme atmospheric water demands, and high solar irradiance. This novel finding requires further research investigating the recovery of date palms from salinity stresses to account for the intact photosynthetic apparatus and solute deployment among palm tissues. Acknowledgments This project was partially supported by a grant (No. 704-000209) obtained from the Chief Scientist of the Israeli Ministry of Agriculture, by the I-CORE Program of the Planning and Budgeting Committee and the Israel Science Foundation (Grant No. 152/11). Additional support was also provided by the RosenzweigCoopersmith Foundation and by ICA in Israel. References Alhammadi, M., Kurup, S., 2012. Impact of salinity stress on date palm (Phoenix dactylifera L.) – a review. In: Sharma, P., Arbol, V. (Eds.), Crop Production Technologies. InTech, Rijeka, pp. 169–178. Aljuburi, H.J., Al-Masry, H.H., 1996. Effects of salinity and indole acetic acid on growth and mineral content of date palm seedlings. Fruits 55, 315–323. Alrasbi, S.A.R., Hussain, N., Schmeisky, H., 2010. Evaluation of the growth of date palm seedlings irrigated with saline water in the sultanate of Oman. In: IV International Date Palm Conference, vol. 882, pp. 233–246. D’Ambrosio, N., Arena, C., Desanto, A., 2006. Temperature response of photosynthesis, excitation energy dissipation and alternative electron sinks to carbon assimilation in Beta vulgaris L. Environmental and Experimental Botany 55, 248–257. Anderson, J.M., Park, Y.-I., Chow, W.S., 1997. Photoinactivation and photoprotection of photosystem II in nature. Physiologia Plantarum 100, 214–223. Araus, J., Hogan, K., 1994. Leaf structure and patterns of photoinhibition in two neotropical palms in clearings and forest understory during the dry season. American Journal of Botany 81, 726–738. Badger, M.R., von Caemmerer, S., Ruuska, S., Nakano, H., 2000. Electron flow to oxygen in higher plants and algae: rates and control of direct photoreduction (Mehler reaction) and rubisco oxygenase. Philosophical Transactions of the Royal Society of London Series B, Biological Sciences 355, 1433–1446. Bar, Y., Apelbaum, A., Kafkafi, U., Goren, R., 1997. Relationship between chloride and nitrate and its effect on growth and mineral composition of avocado and citrus plants. Journal of Plant Nutrition 20, 715–731. Brooks, A., Farquhar, G., 1985. Effect of temperature on the CO2 /O2 specificity of ribulose-1,5-bisphosphate carboxylase/oxygenase and the rate of respiration in the light. Planta 165, 397–406. Brugnoli, E., Björkman, O., 1992. Growth of cotton under continuous salinity stress: influence on allocation pattern, stomatal and non-stomatal components of photosynthesis and dissipation of excess. Planta, 335–347. Chartzoulakis, K., 2000. Comparative study on NACL salinity tolerance of six olive cultivars. In: IV International Symposium on Olive Growing, vol. 586, pp. 497–501. Chartzoulakis, K.S., 2005. Salinity and olive: growth, salt tolerance, photosynthesis and yield. Agricultural Water Management 78, 108–121. Chaves, M.M., 1991. Effects of water deficits on carbon assimilation. Journal of Experimental Botany 42, 1–16. Chaves, M.M., Flexas, J., Pinheiro, C., 2009. Photosynthesis under drought and salt stress: regulation mechanisms from whole plant to cell. Annals of Botany 103, 551–560. Cornic, G., Briantais, J., 1991. Partitioning of photosynthetic electron flow between CO2 and O2 reduction in a C3 leaf (Phaseolus vulgaris L.) at different CO2 concentrations and during drought. Planta 183, 178–184. Downton, W.J.S., Loveys, B.R., Grant, W.J.R., 1990. Salinity effects on the stomatal behaviour of grapevine. New Phytologist 116, 499–503. Eppel, A., Keren, N., Salomon, E., Volis, S., Rachmilevitch, S., 2013. The response of Hordeum spontaneum desert ecotype to drought and excessive light intensity is characterized by induction of O2 dependent photochemical activity and anthocyanin accumulation. Plant Science 201–202, 74–80. Escalona, J., Flexas, J., Medrano, H., 2000. Stomatal and non-stomatal limitations of photosynthesis under water stress in field-grown grapevines. Functional Plant Biology 26, 421–433. Farquhar, G.D., Sharkey, T.D., 1982. Stomatal conductance and photosynthesis. Annual Review of Plant Physiology 33, 317–345.

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