TY - JOUR
AU - Zwieniecki, Maciej, A
AB - Abstract Working in tandem with root exclusion, stems may provide salt-tolerant woody perennials with some additional capacity to restrict sodium (Na) and chloride (Cl) accumulation in leaves. The Pistacia genus, falling at the nexus of salt tolerance and human intervention, provided an ideal set of organisms for studying the influences of both variable root exclusion and potentially variable discontinuities at the bud union on stem processes. In three experiments covering a wide range of salt concentrations (0 to 150 mM NaCl) and tree ages (1, 2 and 10 years) as well as nine rootstock-scion combinations we show that proportional exclusion of both Na and Cl reached up to ~85% efficacy, but efficacy varied by both rootstock and budding treatment. Effective Na exclusion was augmented by significant retrieval of Na from the xylem sap, as evidenced by declines in the Na concentrations of both sap and wood tissue along the transpiration stream. However, while we observed little to no differences between the concentrations of the two ions in leaves, analogous declines in sap concentrations of Cl were not observed. We conclude that some parallel but separate mechanism must be acting on Cl to provide leaf protection from toxicity specific to this ion and suggest that this mechanism is recirculation of Cl in the phloem. The presented findings underline the importance of holistic assessments of salt tolerance in woody perennials. In particular, greater emphasis might be placed on the dynamics of salt sequestration in the significant storage volumes offered by the stems of woody perennials and on the potential for phloem discontinuity introduced with a bud/graft union. Introduction Climate change is further stressing already taxed freshwater distribution systems (IPCC 2008). A scarcity of freshwater both exacerbates reliance on field-level salinity management practices (e.g., by increasing leaching requirements) and limits the potential of these practices to sustain viable farming conditions for at least three reasons. (i) Alternatives to fresh surface water are often more saline (groundwater) or more energy intensive (desalinated water). (ii) Soil quality inevitably declines with exposure to any irrigation (Pitman and Läuchli 2002), but much more quickly when irrigation is highly saline (Rengasamy 2006), a likely scenario when it is exhumed from great depths (Schoups et al. 2005, Kang and Jackson 2016). (iii) Desalinated water is not yet available, economical or environmentally sound on a commercial scale in most parts of the world (Ziolkowska 2015). Given projected fresh water scarcity and the drawbacks or limitations of alternatives, understanding and improving the salinity tolerance of crop plants may provide tools critical to extending the productivity of irrigation-dependent agricultural landscapes. However, while there has been significant progress towards understanding mechanisms and improving the salinity tolerance of short-lived annual crops, particularly cereals (Munns and Tester 2008), the parameters of tolerance outlined for annuals may not adequately capture the context of long-lived woody perennials (e.g., tree and vine crops). Distinguished from short-lived herbaceous species by the perenniality and significant volumes of their stems, salt-tolerant trees may partially rely on the same strategies, that annuals employ to tolerate or combat osmotic stress compartmentalize harmful ions or restrict the accumulation of specific ions in leaf blades, but also include supplementary or exaggerated stem-specific strategies, which enhance salinity tolerance over longer timescales. Some salt inclusion—uptake from soil solution and xylem loading of inorganic osmolytes—is not only biochemically inevitable (Amtmann and Beilby 2010, Teakle and Tyerman 2010), but also biochemically essential. Potential damage to leaves must be weighed against the maintenance of osmotic gradients that favor water movement from root to transpiring tissues despite low soil pore water potentials and charge balance between apoplast and symplast along the way (Bolaños and Longstreth 1984). Although rarely quantified (Raven 1985), the potential energetic costs of exclusive reliance on compatible carbon solutes to achieve the same osmotic adjustment effects, as well as the costs of wasteful cycling of sodium (Na) in and out of root cortical cells, are at the forefront of salinity tolerance discussions (Tyerman et al. 2019). Both of these costs may be mitigated by permitting some managed entry and distribution throughout the plant of ions highly active in solution. While it is accurate to classify the most salt tolerant of the world’s agricultural species like barley (Shabala et al. 2010), and species of both socioeconomic and ecological importance like mangroves (Krishnamurthy et al. 2014, Reef and Lovelock 2015), as excluders, it is more accurate to classify them as highly regulated Includers. Salt is not absent in their tissues when exposed. Indeed, mangroves require some salts for the maintenance of growth (Ball 1988, Cram et al. 2002). Still, salt tolerance does depend, at least in part, on protecting a plant’s photosynthetic capacity by restricting the inflow of environmentally pervasive yet minimally essential ions like Na or chloride (Cl) to the leaf mesophyll (Schactman and Munns 1992) and not all plants are as adept as mangroves in sequestering or excreting excess. Between roots and leaves, in trees, is a stem of enormous volume, contributing 72–75% of the total biomass (Reich et al. 2014). Much of this volume is composed of living cells (xylem parenchyma) associated with storage of non-structural carbohydrates (Bazot et al. 2013, Plavcová et al. 2016), among other functions (Holbrook and Sinclair 1992, Secchi and Zwieniecki 2011, Broderson and McElrone 2013, Morris et al. 2016b, von Arx et al. 2017). Could these cells also be used to store salts? A recent meta-analysis found that an average of 10% (conifers) to 30% (angiosperm trees) of wood is occupied by parenchyma cells, with many species falling out well above these means (Morris et al. 2016a). Given the volumes they occupy in trees and their location along vessels, we hypothesized that the stem’s xylem parenchyma may also act as an ion exchange column essential to our understanding of tree salt tolerance, providing significant desalinization of the xylem sap and salt storage that extends the protective capacity of roots. There is a paucity of investigations linking storage in the xylem parenchyma with sequestration capacity. If they are linked, in a cropping system that relies on budding or grafting (as most tree cropping systems do), a rootstock that contributes some tolerance by excluding high percentages of the salts in soil solution may not in and of itself protect scions from toxic ion accumulation, raising interesting questions about the cumulative contributions to salt tolerance of both rootstock and scion as well as the impacts of their interaction. The Pistacia genus is native to the arid Mediterranean climates of Eurasia and salt tolerant relative to other pomological crops (Ferguson et al. 2005). Declines in production for all other commercially successful tree crops, with the exception of dates (a monocot), occur at salinities 5–10 times lower (Grieve et al. 2012). However, degrees of attributed tolerance may vary by species or cultivar (Picchioni et al. 1990, Ferguson et al. 2002, Chelli-Chaabouni et al. 2010). We capitalized on pistachio’s general but variable tolerance to test root and stem contributions to salt tolerance, following patterns of Na and Cl distribution and accumulation in the xylem sap, roots, wood, bark and leaves of several cultivars (nine rootstock–scion combinations) exposed to concentrations of NaCl between 0 and 150 mM NaCl. Our highest salinity treatment approached the margins of acceptability in agricultural systems (Pitman and Läuchli 2002). Methods We first examined the limits of Na tolerance in one pistachio rootstock (Experiment 1) and later used several rootstocks and budding treatments to test the efficacy of exclusion from the stem and filtration by the stem for both Na and Cl (Experiment 2). We also tested long-term salt storage in the stems of pistachio established in and regularly irrigated with saline ground water over 10 years. Experiment 1 Plant material and growing conditions In late spring (May 15), 1-year-old clonal pistachio rootstocks (Pistacia atlantica Desf. x Pistacia integerrima J. Stewart ‘UCB1’ (UCB1) were transplanted into 11.5 l tree pots filled with a 50:50 mixture of fine sand and peat moss. Trees were then kept on raised beds outdoors at a UC Davis facility (38.542379, −121.762495). Starting on August 20 (treatment day 0), eight replicate treatments of 0, 10, 50 and 150 mM NaCl were applied daily at 16:00 h. Pots were flushed with solutions beyond soil water holding capacity such that soil solution was replaced daily, at least one-third of the applied irrigation leaking from perforations at the bottom each pot. We increased the NaCl concentration of our highest salinity treatment in three daily steps of 50 mM (starting on August 18) to avoid osmotic shock. In vivo measurement Weekly measurements included predawn stem water potential, leaf Na concentration, stomatal conductance and photographs to quantify canopy development. For determination of predawn stem water potential, mature leaves were sealed into small plastic bags in the field 30 min before dawn and allowed to equilibrate for ~10 min prior to excision with a razor. Additional equilibration time was allowed in transport to the lab where pressure bomb measurements were completed. Leaves were then rinsed in a series of three baths of ultrapure water (<0.05 dS m−1), dried at approximately 80° C for 48 h, ground and analyzed for Na (see below). Stomatal conductance (Porometer, Decagon Devices, Inc., Pullman, Washington) measurements were taken from 08:00 until 16:00 h and averaged over the course of the day (four to six measurements per individual per day, once every 1.5–2 h). Weekly photographs of each plant were used as a proxy for leaf area analysis. Post-harvest analysis At final harvest (September 23, treatment day 34), seven trees from each treatment were destructively sampled and sorted into roots, stems and leaves then weighed fresh. Three leaf samples from each tree were then selected for Na analysis from the healthiest available leaves matured prior to the start of treatment, matured during treatment or that were immature. Coarse (>0.5 mm) and fine (<0.5 mm) root samples were collected as were stem samples 2 cm in length taken at the root crown, the stem midpoint (mature) and 5 cm below the apical meristem (immature). Xylem sap was extracted from the remaining stem segments between these samples (proximal and distal) for five to six individuals from each treatment using a vacuum method (Secchi and Zwieniecki 2012). All tissues were then dried at ~80° C for 48 h and reweighed. Stem samples used for analysis were stripped of bark before drying; root samples were washed thoroughly with deionized and then ultrapure water before drying. After drying, leaf tissues were ground with a mortar and pestle then sifted through a 0.5 mm sieve. Wood and root samples were ground and sifted (0.5 mm) with a Wiley Mill (Thomas Scientific). Homogenized tissue (15.0 ± 1.0 mg) was then placed in a 2 ml microtube with 1.5 ml of ultrapure water (milliQ), agitated and centrifuged. The supernatant was then further diluted in ultrapure water (1:10 v/v) to accommodate the diameter of our Na specific electrode (Thermo Scientific). Measured mV values were converted to mmol g−1 dry weight (DW) using a standard calibration curve. Sap samples were centrifuged and diluted in ultrapure water (1:10 v/v) for measurement with the same Na specific electrode. Statistical analysis Linear mixed effects models with tree ID as a random effect were applied in conjunction with an ANOVA to analyze the effect of time on repeated measurements of physiology, growth and leaf Na accumulation. Measurements of physiology, leaf area and leaf Na concentrations at a particular date, as well as endpoint basal sap concentrations (exclusion) and endpoint tissue concentrations were separated by Tukey’s HSD (P < 0.05). Significance values for the difference between treatment and proximal sap concentrations (exclusion) as well as proximal and distal sap concentrations for each treatment (retrieval) were determined using the Wilcoxon rank-sum test. Experiment 2 Plant material and growing conditions One-year-old seedling rootstocks (P. integerrima x P. atlantica ‘PGII’ (PGII), P. integerrima ‘PGI’ (PGI), and P. atlantica x P. integerrima ‘UCB1’ (UCB1)) were removed from their nursery pots in early summer, washed of any medium and transplanted into 38 l plastic bins (BRUTE), which had each been perforated with six drainage holes 2 cm in diameter, lined with a doubled nylon mesh and filled with coarse sand. Forty-five individuals of each rootstock were distributed among 15 irrigation blocks—five blocks for each of three future salinity treatments, three individuals of each rootstock to each block. After budding treatments (described below), the experiment was arranged as a split plot, with one of each rootstock–budding combination (nine total) randomized within each irrigation block and the arrangement of irrigation/salinity blocks also randomized. Pots were placed above trays ~10 cm deep and 45 cm across filled with washed pea gravel and connected into a gravity-driven effluent recapture and redistribution system. Buds were collected from a single mature Pistacia vera L. (Kerman) individual in the fall and T-budded at ~50 cm height to a third of the individuals of each rootstock so that one rootstock-Kerman combination was present within each of the 15 blocks. The same was done for a second third of each rootstock, but with buds taken from higher up on the rootstock itself. The last third of each rootstock type was left unbudded. Shortly after budding, the apical meristems of each seedling were removed to disrupt apical dominance. This included the unbudded treatment for consistence. Any buds that were not scion, or a selected single scaffold that approximated the scion in the unbudded treatment (all budding treatments collectively referred to in the text as `scion’), were pinched as soon as they were observed and the rootstock was cut just above the bud as soon as leaves of the new shoot began to expand in the spring. In the budding process, we lost one Kerman-budded PGI rootstock, four self-budded PGI rootstocks, three Kerman-budded PGI rootstocks and three Kerman-budded UCB1 rootstocks. We distributed the remaining individuals among salinity treatments such that our replications were as high as possible for each treatment group (Table S1 available as Supplementary Data at Tree Physiology Online). Trees were irrigated daily throughout the first summer (preceding budding) and fall with a modified half Hoagland’s solution (low salinity, Table S2 available as Supplementary Data at Tree Physiology Online) from six 2 l per hour inline drip emitters starting at 16:00 h in six 15-min bursts separated by 15 min of infiltration, achieving saturation and drainage. After senescence, irrigation was reduced to weekly in the absence of rainfall. Daily irrigation began again as soon as leaf buds began to swell the following spring using the same modified Hoagland’s solution. Salinity treatments (see below) began in early July after at least one fully expanded leaf was present on each individual and continued until mid-September. We added calcium chloride (CaCl2) to NaCl treatments in an effort to avoid confounding results with symptoms of Na-induced calcium deficiency (for CaCl2 concentrations see below). Weekly additions of reverse osmosis water to replace evapotranspiration and electroconductivity (EC) measurements of drainage ensured that our low salinity treatment (modified Hoagland’s solution +0 mM NaCl) was maintained between 1 and 2 dS m−1, the moderate salinity treatment (modified Hoagland’s solution + 50 mM NaCl + 10 mM CaCl2) was maintained between 6 and 7 dS m−1 and the high salinity treatment (modified Hoagland’s solution + 100 mM NaCl +20 mM CaCl2) was maintained between 11.5 and 12.5 dS m−1. When beginning the treatments we added CaCl2 with nutrients on day 1 and then 50 mM NaCl 2 days in a row to reduce the likelihood of osmotic shock. In vivo measurements Weekly or biweekly measurements included predawn and midday water potential, stomatal conductance, leaf chlorophyll content and three separate growth measurements. Leaves for predawn water potential were collected weekly in the same manner described for predawn collections in Experiment 1. Leaves for midday stem water potential were collected in the same way, but bagging began 6 h after sunrise. Stomatal conductance was also measured weekly from 08:00 to 16:00 h, so that we completed one morning, one midday and one afternoon measurement for each seedling, each measurement separated by 2–3 h. Rootstock and scion diameter were measured 2.5–3.5 h after sunrise at marked locations on the trunk. For rootstocks, the marked location was ~20 cm above the root crown and for `scions’ the marked location was ~5 cm above the bud union. To observe extension growth, we tagged all branches on the most apical node present at the start of treatment then measured cumulative branch length beyond each tag every 2 weeks. Post-harvest analysis At the conclusion of the experiment, 9 to 15 individuals were destructively sampled each day for nine consecutive days (total of 124 trees). Xylem sap was extracted from 2–4 cm stem segments (increasing acropetally to accommodate lower volumes) starting just above the root crown (position 1, 1 cm long), 4 cm below the transition to scion (position 2, 2 cm long), 1 cm above the transition to scion (position 3, 3 cm long) and 10 cm below the apical meristem (position 4, 4 cm long). All bark was removed from these segments before 1 cm segments were bisected along and across their axes to break xylem vessels and placed in 1.5 ml microtubes then centrifuged at 21,000g for 10 min. We used tubes with tapered bottoms to avoid reabsorption. Sap was then diluted in 3% nitric acid and analyzed for Na and Cl using inductively coupled mass spectrometry (ICPMS). We also sampled wood, bark, roots and leaves. Wood and bark samples were taken from analogous segments just above (positions 1 and 3) or below (positions 2 and 4) sap segments of the same position. All tissues were dried as described in Experiment 1, finely chopped using hand pruners, powdered using a mini-beadbeater (Cole-Palmer) and sieved (0.5 mm). Organic material in all tissue samples was then digested to clarify samples for ICPMS analysis of Na and Cl using a combination of concentrated nitric acid (70%) and hydrogen peroxide (30%) in flat-bottomed 2 ml microtubes, a method modified from one previously described (Hansen et al. 2013) to allow us to work with small tissue quantities. While the cost-effective method used for tissue digestion in preparation for ICPMS is likely to volatilize some Cl, and we did observe Cl values on average 38% lower than those reported by the National Institute of Standards and Technology (NIST) for standard reference material (SRM) 1547, deviation around the mean across digestions for Cl (13%) was only slightly higher than that of Na (10%) for SRM 1573a. Statistical analysis Linear mixed effects models were applied to analyze repeated measures of physiology and growth over time and sap and tissue concentrations along each stem (retrieval) using the LM package in R with individual tree as a random effect. To assess the significance of salinity’s effects on physiology for each treatment combination, least squares means were separated by Tukey's honestly significant difference (HSD, P < 0.05). To assess the significance of salinity’s effects on growth for each treatment combination, slopes (coefficients) of moderate and high salinity treatments were compared with those of the low salinity treatment (control) using the glht function (general linear model hypothesis testing) of R’s multcomp package. To assess the significance of retrieval, slopes (coefficients) of each treatment combination’s model were compared with zero also using the glht function of R’s multcomp package. Exclusion was compared within each budding treatment using an ANOVA; groupings for each salinity and rootstock combination were separated by Tukey’s HSD (P < 0.05). ANOVAs were performed for all treatment combinations with root crown sap concentration (Na and Cl) as the measured variable (exclusion from stem) and for all treatment combinations plus position along the stem with both sap and wood concentrations (Na and Cl) as measured variables (retrieval by stem). Bark concentrations of Na and Cl at the end of the experiment were compared with adjacent wood concentrations of Na and Cl using linear models in conjunction with Wilcoxon rank sum tests. Leaf concentrations of Na and Cl at the end of the experiment compared using a linear model. Experiment 3 Plant material and growing conditions A large-scale long-term salinity field trial established in the San Joaquin Valley in 2004 (described in Sanden et al. 2014) provided oven-dried cross sections across the graft unions of six 10-year-old pistachios (P. vera (Kerman) budded to either P. atlantica x P. integerrima ‘UCB1’ rootstock or P. integerrima ‘PGI’ rootstock). Irrigation with saline well water approximated moderate (10–50 mM) NaCl seedling treatments—averages of 22.8 mM Na and 34.1 mM Cl. Post-harvest analysis At the field site, a section of stem containing the graft union (~40 cm thick) was separated from the trunk using a chainsaw. After smoothing to reveal annual growth rings and using drill bits that matched the ring width of each year, we sampled the most recent 3 years of growth on both sides of the graft union at three random locations within each ring. We also chiseled bark from three locations off of both rootstock and scion and finely chopped using hand pruners. We then combined analogous samples and ground them to a powder, sifting, digesting and analyzing for Na and Cl using ICPMS as described in Experiment 2. Figure 1. Open in new tabDownload slide Salinity responses of water status (a), gas exchange (b), canopy development (c) and leaf Na concentration (d) in unbudded pistachio rootstocks (P. atlantica x integerrima ‘UCB1’). Irrigation treatments of 0, 10, 50 or 150 mM NaCl were applied over 35 days. Data points represent means and SE for eight replicates. Significant effects of treatment and time were observed for each analysis (mixed effects model, P < 0.01). Letters represent significant treatment groups at each time point (Tukey’s HSD, P < 0.05). Figure 1. Open in new tabDownload slide Salinity responses of water status (a), gas exchange (b), canopy development (c) and leaf Na concentration (d) in unbudded pistachio rootstocks (P. atlantica x integerrima ‘UCB1’). Irrigation treatments of 0, 10, 50 or 150 mM NaCl were applied over 35 days. Data points represent means and SE for eight replicates. Significant effects of treatment and time were observed for each analysis (mixed effects model, P < 0.01). Letters represent significant treatment groups at each time point (Tukey’s HSD, P < 0.05). Statistical analysis Linear mixed effects models in conjunction with analysis of variance (ANOVA) were applied to compare tissue concentrations (Na and Cl) in rootstocks and scions as well as across bark and annual growth rings using the LM package in R with individual tree as a random effect. Any groupings were separated by Tukey’s HSD (P < 0.05). Results Physiology and growth The highest applied salt concentration (150 mM NaCl) quickly overwhelmed the capacity of young unbudded pistachios to maintain the same water status, gas exchange, canopy development and leaf Na concentrations observed in the control treatment (0 mM NaCl) of Experiment 1 (Figure 1). Significant physiological differences were observed between 150 mM NaCl and control treatments by approximately treatment day 10 for both predawn stem water potential (Figure 1a) and average daily stomatal conductance (Figure 1b); significant differences in leaf area (Figure 1c) were observed by approximately treatment day 20. In contrast, 50 mM NaCl resulted in predawn stem water potentials and leaf area significantly different from the control along the same timeline as the 150 mM treatment, but no significant differences from the control at any time point in average daily stomatal conductance. No significant differences were observed at any time point for any measurement between the control and 10 mM treatments. When time points were assessed together as repeated measures for a given salinity treatment, the effect of time on predawn stem water potential was significant for all salinity treatments (P < 0.01), including the control. The effect of time on average daily stomatal conductance was only significant for the 150 mM NaCl treatment (P < 0.01). Leaf area increased significantly over time for the control, 10 and 50 mM NaCl treatments (P < 0.01), but not for the 150 mM treatment, indicating no significant canopy development. Measurements of water status and gas exchange in Experiment 2 demonstrated similar trends—with a significant effect of the interaction between salinity and time for predawn water potential (P < 0.01), midday water potential (P = 0.02) and stomatal conductance (P < 0.01). While the effect of rootstock was significant for both water potential measurements and stomatal conductance, its interactions with salinity were not. The three-way interaction between salinity, time and budding treatment was significant, but only for stomatal conductance (P < 0.01). For details on all growth and physiology measurements of each of Experiment 2’s rootstock–scion combinations please see Figures S1–S6 available as Supplementary Data at Tree Physiology Online. Figure 2. Open in new tabDownload slide Sodium concentrations at the final harvest of all tissues of unbudded pistachio rootstocks (P. atlantica x integerrima ‘UCB1’) irrigated for 35 days with 0, 10, 50 or 150 mM NaCl. Data points represent means and SE for seven replicates. Letters indicate significance groups (Tukey’s HSD, P < 0.05) and compare averages within each tissue type (roots, stems, leaves) for a given salinity treatment. Figure 2. Open in new tabDownload slide Sodium concentrations at the final harvest of all tissues of unbudded pistachio rootstocks (P. atlantica x integerrima ‘UCB1’) irrigated for 35 days with 0, 10, 50 or 150 mM NaCl. Data points represent means and SE for seven replicates. Letters indicate significance groups (Tukey’s HSD, P < 0.05) and compare averages within each tissue type (roots, stems, leaves) for a given salinity treatment. Vertical compartmentalization of sodium Root, stem and leaf tissue samples harvested at the end of Experiment 1 demonstrated vertical compartmentalization of Na in all tissue types (Figure 2). Significant declines in tissue concentration of Na along the transpiration stream from fine roots to immature leaves were observed for all but the 150 mM NaCl treatment. Coarse roots demonstrated lower concentrations relative to fine roots for the 0 and 10 mM NaCl treatments (Tukey’s HSD, P < 0.05). Wood samples collected from both the mid and distal stem demonstrated lower concentrations relative to those collected from the root crown for the 0, 10 and 50 mM NaCl treatments. Samples of both mature and immature leaves demonstrated lower concentrations than samples of old (lowest available) leaves also for the 0, 10 and 50 mM treatments (Tukey’s HSD, P < 0.05). However, for the 150 mM treatment we observed no significant differences in Na concentration between fine and coarse roots (also for the 50 mM treatment), from the most proximal (root crown) to the most distal stem samples collected, or from the oldest to the youngest leaves. Maximum Na tissue concentrations appear to be between 0.4 and 0.6 mmol g−1 DW. We observed similar concentrations across rootstocks in the fine roots of Experiment 2's highest salinity treatment. With standard errors (SE) in parentheses, averages in the low significance grouping (P < 0.05) were 0.35 (SE = 0.04), 0.36 (SE = 0.05), and 0.4 (SE = 0.08) mmol g−1 DW for Kerman-budded UCB1, Kerman-budded PGI, and self-budded PGI rootstocks, respectively. Averages in the high significance grouping were 0.63 mmol g−1 DW for self- and unbudded PGII rootstocks (SE = 0.08 and 0.13, respectively). Chloride concentrations in fine roots of Experiment 2’s high salinity treatment differed significantly by rootstock (PGI demonstrating lower concentrations than PGII, but not UCB1), but not by budding treatment. In coarse roots, Cl concentrations also differed significantly by rootstock, but not by budding treatment, UCB1 demonstrating less than the other two rootstocks. Maximum concentrations of Cl averaged ~0.10 mmol g−1 DW in both fine and coarse roots. Figure 3. Open in new tabDownload slide Xylem sap Na concentrations in unbudded pistachio rootstocks (P. atlantica x integerrima ‘UCB1’) extracted after 35 days of irrigation with 0, 10, 50 or 150 mM NaCl. The bold dashed line is the 1:1 line. Exclusion of Na (a) is represented in gray as the difference between the Na concentrations applied (treatment solution) and the measured Na concentrations of sap extracted between the stem’s root crown and midpoint (proximal sap). Data represent the means and SE of proximal sap concentrations for five to six replicates. Letters represent significant differences across treatment groups for proximal sap concentrations (Tukey’s HSD, P < 0.05). Significant differences between treatment and proximal sap Na concentrations were observed within the 50 and 150 mM NaCl treatments (Wilcoxon rank-sum test, P < 0.05). Retrieval of Na in (b) is represented in gray by the difference between the Na concentrations of sap extracted between the stem’s root crown and midpoint (proximal sap) and the concentrations of sap extracted between the stem’s midpoint and apical meristem (distal sap). Data represent the means and SE of proximal and basal sap concentrations for five to six replicates. Asterisks indicate that distal sap concentrations were significantly lower than proximal sap concentrations for both 50 and 150 mM treatments (Wilcoxon rank-sum test, P < 0.05). Figure 3. Open in new tabDownload slide Xylem sap Na concentrations in unbudded pistachio rootstocks (P. atlantica x integerrima ‘UCB1’) extracted after 35 days of irrigation with 0, 10, 50 or 150 mM NaCl. The bold dashed line is the 1:1 line. Exclusion of Na (a) is represented in gray as the difference between the Na concentrations applied (treatment solution) and the measured Na concentrations of sap extracted between the stem’s root crown and midpoint (proximal sap). Data represent the means and SE of proximal sap concentrations for five to six replicates. Letters represent significant differences across treatment groups for proximal sap concentrations (Tukey’s HSD, P < 0.05). Significant differences between treatment and proximal sap Na concentrations were observed within the 50 and 150 mM NaCl treatments (Wilcoxon rank-sum test, P < 0.05). Retrieval of Na in (b) is represented in gray by the difference between the Na concentrations of sap extracted between the stem’s root crown and midpoint (proximal sap) and the concentrations of sap extracted between the stem’s midpoint and apical meristem (distal sap). Data represent the means and SE of proximal and basal sap concentrations for five to six replicates. Asterisks indicate that distal sap concentrations were significantly lower than proximal sap concentrations for both 50 and 150 mM treatments (Wilcoxon rank-sum test, P < 0.05). Figure 4. Open in new tabDownload slide Xylem sap Na (a) and Cl (b) concentrations in Kerman-budded, self-budded and unbudded PGII, PGI and UCB1 rootstocks treated with 0 mM NaCl, 50 mM NaCl + 10 mM CaCl2 or 100 mM NaCl + 20 mM CaCl2. The bold dashed line in each panel is the 1:1 line. Data represent the means and SE of proximal sap concentrations for three to five replicates of each of 27 combinations of rootstock, budding treatment and salinity treatment. Exclusion is represented as the difference between the concentration applied (treatment solution) and the measured concentration of sap extracted between the stem’s root crown and midpoint (proximal sap) for a given rootstock in layers of gray. Figure 4. Open in new tabDownload slide Xylem sap Na (a) and Cl (b) concentrations in Kerman-budded, self-budded and unbudded PGII, PGI and UCB1 rootstocks treated with 0 mM NaCl, 50 mM NaCl + 10 mM CaCl2 or 100 mM NaCl + 20 mM CaCl2. The bold dashed line in each panel is the 1:1 line. Data represent the means and SE of proximal sap concentrations for three to five replicates of each of 27 combinations of rootstock, budding treatment and salinity treatment. Exclusion is represented as the difference between the concentration applied (treatment solution) and the measured concentration of sap extracted between the stem’s root crown and midpoint (proximal sap) for a given rootstock in layers of gray. Sodium and chloride exclusion from the shoot Significant exclusion of Na from shoot sap was observed in both experiments. In Experiment 1, which only analyzed for Na, sap collected from stem segments extending between root crown and stem midpoint (proximal sap) exhibited concentrations significantly lower than those of the treatment solution when both 50 and 150 mM NaCl were applied (P < 0.01, Wilcoxon rank-sum test), with means falling well below the 1:1 line (Figure 3a). Concentrations were as low as ~15% of the applied treatment solution for the 150 mM treatment, indicating that 85% of the Na in treatment solution was either excluded from the plant completely or stored by roots before reaching the shoot in sap. In Experiment 2, which analyzed for both Na and Cl, rootstock (stock), budding treatment (bud) and salinity all exhibited highly significant effects on the Na (Figure 4a) and Cl (Figure 4b) concentrations of proximal sap. All two-way interactions were also significant for Na, while only the two-way interaction between budding treatment and salinity treatment was significant for Cl. The maximum exclusion of Na from the shoot observed was 85–90% (self-budded and unbudded UCB1 rootstocks treated with moderate salinity and Kerman-budded UCB1 rootstocks treated with high salinity). The maximum exclusion of Cl observed was also ~85% (Kerman-budded PGII and UCB1 rootstocks treated with high salinity). Probability values generated by the ANOVA are displayed in Table 1. Interestingly, sap concentrations at position 1 (collected just above the root crown) were not correlated with either Na concentrations in fine or coarse roots (P = 0.22 and 0.27, respectively; corresponding r2 = 0.015 and 0.007). The same was true for Cl concentrations in fine or coarse roots (P = 0.25 and 0.17, respectively; corresponding r2 = 0.010 and 0.027, data not shown). Table 1 Probability values and significance codes from an ANOVA performed on sap concentrations of Na and Cl extracted from 1–2 cm xylem segments collected just above root crowns (position 1) of PGII, PGI and UCB1 rootstocks (rootstock) budded to Kerman, self-budded or left unbudded (bud) and treated with 0 mM NaCl, 50 mM NaCl + 10 mM CaCl2 or 100 mM NaCl +20 mM CaCl2 (salinity). Symbol `:’ represents the interaction between factors. Symbols in the Sig column indicate P <0.001 (***), P < 0.01 (**), P < 0.05 (*), P < 0.1 (.), and not significant (NS). Factor Sap Na Sap Cl P Sig P Sig Rootstock <0.01 *** <0.01 *** Bud <0.01 *** <0.01 *** Salinity <0.01 *** <0.01 *** Rootstock:bud 0.01 * 0.23 NS Rootstock:salinity <0.01 *** 0.30 NS Bud:salinity <0.01 *** <0.01 ** Rootstock:bud:salinity 0.17 NS 0.07 . Factor Sap Na Sap Cl P Sig P Sig Rootstock <0.01 *** <0.01 *** Bud <0.01 *** <0.01 *** Salinity <0.01 *** <0.01 *** Rootstock:bud 0.01 * 0.23 NS Rootstock:salinity <0.01 *** 0.30 NS Bud:salinity <0.01 *** <0.01 ** Rootstock:bud:salinity 0.17 NS 0.07 . Open in new tab Table 1 Probability values and significance codes from an ANOVA performed on sap concentrations of Na and Cl extracted from 1–2 cm xylem segments collected just above root crowns (position 1) of PGII, PGI and UCB1 rootstocks (rootstock) budded to Kerman, self-budded or left unbudded (bud) and treated with 0 mM NaCl, 50 mM NaCl + 10 mM CaCl2 or 100 mM NaCl +20 mM CaCl2 (salinity). Symbol `:’ represents the interaction between factors. Symbols in the Sig column indicate P <0.001 (***), P < 0.01 (**), P < 0.05 (*), P < 0.1 (.), and not significant (NS). Factor Sap Na Sap Cl P Sig P Sig Rootstock <0.01 *** <0.01 *** Bud <0.01 *** <0.01 *** Salinity <0.01 *** <0.01 *** Rootstock:bud 0.01 * 0.23 NS Rootstock:salinity <0.01 *** 0.30 NS Bud:salinity <0.01 *** <0.01 ** Rootstock:bud:salinity 0.17 NS 0.07 . Factor Sap Na Sap Cl P Sig P Sig Rootstock <0.01 *** <0.01 *** Bud <0.01 *** <0.01 *** Salinity <0.01 *** <0.01 *** Rootstock:bud 0.01 * 0.23 NS Rootstock:salinity <0.01 *** 0.30 NS Bud:salinity <0.01 *** <0.01 ** Rootstock:bud:salinity 0.17 NS 0.07 . Open in new tab Sodium, but not chloride retrieval Xylem retrieval of Na further lowered concentrations of sap beyond what was observed in the root crown in both experiments. Consistent with bookending wood measurements (Figure 2), sap extracted between mid-stem and apex (distal sap) also demonstrated significantly lower Na concentrations than sap extracted from stem segments between root crown and mid-stem (proximal sap) in Experiment 1 (Figure 3b). For both the 50 mM and 150 mM NaCl treatments, sap samples collected further along the transpiration stream were significantly lower (Wilcoxon rank-sum test, P = 0.03 and 0.02, respectively), a trend that continued regardless of rootstock or budding treatment in Experiment 2 though with some variation (Figure 5). Sodium concentrations in the sap of seedlings declined significantly up the stem from position 1 at the root crown to position 4 below the apical meristem in the high salinity treatments of all rootstock–budding treatment combinations and the moderate salinity treatments of all PGII and PGI combinations (Figure 5a), demonstrating a highly significant effect of position alone as well as the interaction between position and salinity on sap Na concentrations (Table 2). Sodium concentrations in adjacent wood (Figure 5b) also declined significantly up the stem in almost all rootstock–scion combinations, again demonstrating highly significant effects of both position alone and the interaction between salinity and position. Figure 5. Open in new tabDownload slide Sap (a, c) and wood (b, d) concentrations of Na (a, b) and Cl (c, d) along the stem axes of 2-year-old PGII, PGI and UCB1 rootstocks budded to Kerman, self-budded or left unbudded. Low (L), moderate (M) and high (H) salinity treatments of 0 mM NaCl, 50 mM NaCl + 10 mM CaCl2 or 100 mM NaCl + 20 mM CaCl2, respectively, were added to irrigation and applied for ~10 weeks before sap was extracted from a segment just above the root crown (position 1), just below the transition to scion (position 2), just above the transition to scion (position 3) and just below the apical meristem (position 4). Positions of wood sample indicate a segment taken directly above (position 1, position 3) or below (position 2, position 4) sap samples of the same position. Asterisks in a rootstock’s treatment color indicate that slopes of these models were significantly different from zero (glht, P < 0.05). Figure 5. Open in new tabDownload slide Sap (a, c) and wood (b, d) concentrations of Na (a, b) and Cl (c, d) along the stem axes of 2-year-old PGII, PGI and UCB1 rootstocks budded to Kerman, self-budded or left unbudded. Low (L), moderate (M) and high (H) salinity treatments of 0 mM NaCl, 50 mM NaCl + 10 mM CaCl2 or 100 mM NaCl + 20 mM CaCl2, respectively, were added to irrigation and applied for ~10 weeks before sap was extracted from a segment just above the root crown (position 1), just below the transition to scion (position 2), just above the transition to scion (position 3) and just below the apical meristem (position 4). Positions of wood sample indicate a segment taken directly above (position 1, position 3) or below (position 2, position 4) sap samples of the same position. Asterisks in a rootstock’s treatment color indicate that slopes of these models were significantly different from zero (glht, P < 0.05). Table 2 Probability values and significance codes from an ANOVA comparing sap and wood concentrations of Na and Cl at four positions along the stems of PGII, PGI and UCB1 rootstocks (stock) budded to Kerman, self-budded or left unbudded (bud) and treated with 0 mM NaCl, 50 mM NaCl + 10 mM CaCl2 or 100 mM NaCl + 20 mM CaCl2 (salinity) over 10 weeks. Symbol `:’ represents the interaction between factors. Symbols in the Sig column indicate P <0.001 (***), P < 0.01 (**), P < 0.05 (*), P < 0.1 (.), and not significant (NS). Factor Sap Na Wood Na Sap Cl Wood Cl P Significance P Significance P Significance P Significance Stock <0.01 *** <0.01 *** <0.01 *** <0.01 *** Bud <0.01 *** <0.01 ** <0.01 *** <0.01 ** Salinity <0.01 *** <0.01 *** <0.01 *** <0.01 *** Position <0.01 *** <0.01 *** 0.10 NS 0.42 NS Stock:bud 0.01 * 0.10 . 0.02 * 0.42 NS Stock:salinity <0.01 *** <0.01 *** <0.01 ** <0.01 *** Bud:salinity <0.01 *** 0.02 * <0.01 *** 0.22 NS Stock:position <0.01 *** 0.02 * 0.21 NS 0.14 NS Bud:position 0.02 * <0.01 *** 0.49 NS <0.01 *** Salinity:position <0.01 *** <0.01 *** 0.05 . <0.01 *** Stock:bud:salinity 0.09 . 0.45 NS <0.01 *** <0.01 ** Stock:bud:position 0.52 NS 0.02 * 0.83 NS 0.05 . Stock:salinity:position <0.01 ** 0.05 . 0.33 NS 0.23 NS Bud:salinity:position 0.18 NS 0.02 * 0.59 NS <0.01 *** Stock:bud:salinity:position 0.03 * <0.01 *** 0.58 NS <0.01 ** Factor Sap Na Wood Na Sap Cl Wood Cl P Significance P Significance P Significance P Significance Stock <0.01 *** <0.01 *** <0.01 *** <0.01 *** Bud <0.01 *** <0.01 ** <0.01 *** <0.01 ** Salinity <0.01 *** <0.01 *** <0.01 *** <0.01 *** Position <0.01 *** <0.01 *** 0.10 NS 0.42 NS Stock:bud 0.01 * 0.10 . 0.02 * 0.42 NS Stock:salinity <0.01 *** <0.01 *** <0.01 ** <0.01 *** Bud:salinity <0.01 *** 0.02 * <0.01 *** 0.22 NS Stock:position <0.01 *** 0.02 * 0.21 NS 0.14 NS Bud:position 0.02 * <0.01 *** 0.49 NS <0.01 *** Salinity:position <0.01 *** <0.01 *** 0.05 . <0.01 *** Stock:bud:salinity 0.09 . 0.45 NS <0.01 *** <0.01 ** Stock:bud:position 0.52 NS 0.02 * 0.83 NS 0.05 . Stock:salinity:position <0.01 ** 0.05 . 0.33 NS 0.23 NS Bud:salinity:position 0.18 NS 0.02 * 0.59 NS <0.01 *** Stock:bud:salinity:position 0.03 * <0.01 *** 0.58 NS <0.01 ** Open in new tab Table 2 Probability values and significance codes from an ANOVA comparing sap and wood concentrations of Na and Cl at four positions along the stems of PGII, PGI and UCB1 rootstocks (stock) budded to Kerman, self-budded or left unbudded (bud) and treated with 0 mM NaCl, 50 mM NaCl + 10 mM CaCl2 or 100 mM NaCl + 20 mM CaCl2 (salinity) over 10 weeks. Symbol `:’ represents the interaction between factors. Symbols in the Sig column indicate P <0.001 (***), P < 0.01 (**), P < 0.05 (*), P < 0.1 (.), and not significant (NS). Factor Sap Na Wood Na Sap Cl Wood Cl P Significance P Significance P Significance P Significance Stock <0.01 *** <0.01 *** <0.01 *** <0.01 *** Bud <0.01 *** <0.01 ** <0.01 *** <0.01 ** Salinity <0.01 *** <0.01 *** <0.01 *** <0.01 *** Position <0.01 *** <0.01 *** 0.10 NS 0.42 NS Stock:bud 0.01 * 0.10 . 0.02 * 0.42 NS Stock:salinity <0.01 *** <0.01 *** <0.01 ** <0.01 *** Bud:salinity <0.01 *** 0.02 * <0.01 *** 0.22 NS Stock:position <0.01 *** 0.02 * 0.21 NS 0.14 NS Bud:position 0.02 * <0.01 *** 0.49 NS <0.01 *** Salinity:position <0.01 *** <0.01 *** 0.05 . <0.01 *** Stock:bud:salinity 0.09 . 0.45 NS <0.01 *** <0.01 ** Stock:bud:position 0.52 NS 0.02 * 0.83 NS 0.05 . Stock:salinity:position <0.01 ** 0.05 . 0.33 NS 0.23 NS Bud:salinity:position 0.18 NS 0.02 * 0.59 NS <0.01 *** Stock:bud:salinity:position 0.03 * <0.01 *** 0.58 NS <0.01 ** Factor Sap Na Wood Na Sap Cl Wood Cl P Significance P Significance P Significance P Significance Stock <0.01 *** <0.01 *** <0.01 *** <0.01 *** Bud <0.01 *** <0.01 ** <0.01 *** <0.01 ** Salinity <0.01 *** <0.01 *** <0.01 *** <0.01 *** Position <0.01 *** <0.01 *** 0.10 NS 0.42 NS Stock:bud 0.01 * 0.10 . 0.02 * 0.42 NS Stock:salinity <0.01 *** <0.01 *** <0.01 ** <0.01 *** Bud:salinity <0.01 *** 0.02 * <0.01 *** 0.22 NS Stock:position <0.01 *** 0.02 * 0.21 NS 0.14 NS Bud:position 0.02 * <0.01 *** 0.49 NS <0.01 *** Salinity:position <0.01 *** <0.01 *** 0.05 . <0.01 *** Stock:bud:salinity 0.09 . 0.45 NS <0.01 *** <0.01 ** Stock:bud:position 0.52 NS 0.02 * 0.83 NS 0.05 . Stock:salinity:position <0.01 ** 0.05 . 0.33 NS 0.23 NS Bud:salinity:position 0.18 NS 0.02 * 0.59 NS <0.01 *** Stock:bud:salinity:position 0.03 * <0.01 *** 0.58 NS <0.01 ** Open in new tab We did not observe analogous declines in sap (Figure 5c) or wood (Figure 5d) Cl concentrations. Note that the relationship between the scales of sap and wood axes for Na are reversed for Cl—higher sap values for Cl than for Na, higher wood values for Na than for Cl. Position alone was not a significant predictor of concentration (P = 0.10 and 0.42 for sap and wood, respectively) and although the interaction between position and salinity treatment was significant (P = 0.05 and <0.01 for sap and wood, respectively), modeled slopes of Cl concentration in sap up the stem are not significantly different from 0 for any treatment combination. Concentrations in wood demonstrate declines for some combinations of rootstock, budding treatment and salinity treatment, but generally either increase or show no significant difference. Interestingly, while no Kerman-budded rootstocks show significant declines in sap Cl concentrations, they do show significant declines in wood Cl concentrations for moderate (PGII), high (UCB1) and both (PGI) salinity treatments. Comparing bark sodium and chloride to wood sodium and chloride Comparing bark samples with wood samples representing the three salinity treatments, nine rootstock-scion combinations and four positions, significant positive linear correlations were observed for both Na (Figure 6a, slope = 1.14, SE = 0.06, P < 0.01, r2 = 0.768) and Cl (Figure 6b, slope = 0.95, SE = 0.07, P < 0.01, r2 = 0.598). Bark Na concentration was very slightly, but significantly lower than wood Na concentration (P < 0.01, Wilcoxon rank-sum test), while bark Cl concentration was not significantly different from wood Cl concentration (P = 0.1384, Wilcoxon rank sum test). Figure 6. Open in new tabDownload slide Relationships between bark and wood concentrations of Na and Cl (n = 123). Low (L), moderate (M) and high (H) salinity treatments of 0 mM NaCl, 50 mM NaCl + 10 mM CaCl2 or 100 mM NaCl + 20 mM CaCl2, respectively, were added to irrigation and applied for ∼10 weeks before wood and bark samples were collected at several positions along the along the stem axes of 2-year-old PGII, PGI and UCB1 rootstocks budded to Kerman, self-budded or left unbudded. The data for each tissue type are fit by linear regression (solid lines, r2 = 0.768 and 0.598, slope = 1.14 and 0.95, SE = 0.06 and 0.07 for Na and Cl, respectively, and P < 0.05). Confidence intervals (shaded areas, 95%) and a 1:1 line (bold dashed line) are also provided. Figure 6. Open in new tabDownload slide Relationships between bark and wood concentrations of Na and Cl (n = 123). Low (L), moderate (M) and high (H) salinity treatments of 0 mM NaCl, 50 mM NaCl + 10 mM CaCl2 or 100 mM NaCl + 20 mM CaCl2, respectively, were added to irrigation and applied for ∼10 weeks before wood and bark samples were collected at several positions along the along the stem axes of 2-year-old PGII, PGI and UCB1 rootstocks budded to Kerman, self-budded or left unbudded. The data for each tissue type are fit by linear regression (solid lines, r2 = 0.768 and 0.598, slope = 1.14 and 0.95, SE = 0.06 and 0.07 for Na and Cl, respectively, and P < 0.05). Confidence intervals (shaded areas, 95%) and a 1:1 line (bold dashed line) are also provided. Sodium and chloride in leaves The predictable consequence of ineffective Na or Cl exclusion from and retrieval by stems is the arrival of these salts in leaves. The effect of time on leaf Na concentration was significant for the control in Experiment 1 (P = 0.01), but the trend was negative, i.e., the concentration of Na decreased over time; the effect of time on leaf Na concentration was also significant for 50 and 150 mM NaCl treatments (P < 0.01), differences observed by approximately treatment day 20 (Figure 1d). In Experiment 2, significant positive linear correlations were observed between the concentrations of Na and Cl in both mature (P < 0.01, r2 = 0.568, slope = 0.44, SE = 0.07) and immature (P < 0.01, r2 = 0.780, slope = 0.97, SE = 0.1) leaf blades across rootstock, budding and salinity treatments (Figure 7). Figure 7. Open in new tabDownload slide Relationships between Na and Cl concentrations in the mature and immature leaf blades of all 27 treatment combinations. The data for each leaf type are fit by linear regression (solid lines, r2 = 0.568 and 0.780, slope = 0.44 and 0.97, SE = 0.07 and 0.10 for mature and immature leaf blades, respectively, and P < 0.05). Confidence intervals (shaded areas, 95%) and a 1:1 line (bold dashed line) are also provided. Figure 7. Open in new tabDownload slide Relationships between Na and Cl concentrations in the mature and immature leaf blades of all 27 treatment combinations. The data for each leaf type are fit by linear regression (solid lines, r2 = 0.568 and 0.780, slope = 0.44 and 0.97, SE = 0.07 and 0.10 for mature and immature leaf blades, respectively, and P < 0.05). Confidence intervals (shaded areas, 95%) and a 1:1 line (bold dashed line) are also provided. Sodium and chloride in mature trees Tissue concentrations in 10-year-old ‘Kerman’ grafted trees treated with moderate levels of NaCl in Experiment 3 demonstrated significant declines in Na and Cl across the graft union for both PGI and UCB1 rootstocks (Figure 8, P < 0.01). Significant differences across bark and annual growth rings were observed only for Na concentrations in the tissues of ‘Kerman’ scions budded on PGI rootstocks (P < 0.01), in which bark, ring 10 and ring 9 years were not significantly different, but ring 8 was significantly higher than ring 10 (Tukey’s HSD, P < 0.01). Discussion High exclusion rates, particularly when external concentrations are also high, may be partially responsible for pistachio’s relatively high salinity tolerance. Root exclusion is a well-established first line of defense from salt toxicity and utilized by many of the world’s most famously salt tolerant trees—mangroves exclude between 79.5 and 99.6% of external salts (Scholander et al. 1962, Reef and Lovelock 2015). Here, we have shown that exclusion can account for an up to ~85% reduction in the concentrations of both Na and Cl between soil solution and the root crown xylem sap of young pistachios (Figures 3 and 4). However, how pistachio accomplishes its exclusion clearly varies by ion as might be expected—the two ions vary in charge and should thus, given equal concentrations, be actively or passively passed across membranes in opposing directions in pursuit of Nernst equilibrium. In this first step in the path from soil solution to leaves, we observed dramatic differences between the retention of Na (>0.35 mmol g−1 DW) and the retention of Cl (~0.10 mmol g−1 DW) in root tissue. Sodium concentrations in both fine and coarse roots were three to six times higher than those of Cl despite the lower concentration of Na in soil solution (100 mM Na, 140 mM Cl). Unless differences in root retention are entirely related to differences in uptake, an unlikely possibility given the tendency of Cl to leak from soil solution into root symplast at high external concentrations (Teakle and Tyerman 2010), and in the absence of any other mechanism, we would expect differences in the accumulation of both ions in root crown sap and subsequently in leaves; yet, we observe comparable levels of the two ions at both locations (Figures 3 and 7, respectively). Our observation of 30–60 mM Na and Cl concentrations in xylem sap extracted at the root crown, combined with an average transpiration rate of ~2 mmol H2O m−2 s−1 should result in the leaf deposition of 5–10 mmol g−1 DW of each ion over 30 days (assuming 10 h of effective transpiration per day). This is a conservative estimate given the reported transpiration rates of healthy pistachio leaves between 3 and 9 mmol H2O m−2 s−1 (Ghazvini et al. 2007, Banakar and Ranjbar 2010) and no observation of declines in transpiration imposed by 100 mM NaCl treatments (Walker et al. 1988). However, our results show a maximum cumulative leaf deposition of only ~0.6 mmol Na g−1 DW over the same amount of time in Experiment 1’s highest salinity treatment (150 mM NaCl, Figure 2), and much lower levels of both Na and Cl, ~0.2 mmol g−1 DW, in Experiment 2’s highest salinity treatment (100 mM Na, 140 mM Cl) over twice that time (Figure 7). Several lines of evidence suggest that limits to leaf Na accumulation are the result of xylem retrieval and sequestration made possible by equilibrium potentials, which favor flux of the positively charged Na into but not out of the cytoplasm of xylem parenchyma cells, as well as into but not out of the vacuoles of these same cells, while limits to leaf Cl accumulation are the result of phloem recirculation. A diagram illustrating some of the changes in membrane potentials between xylem sap and the cytoplasm of parenchyma cells that we may be observing appears in Figure 9. Figure 8. Open in new tabDownload slide Bark and annual growth ring (years 10, 9 and 8) wood concentrations of Na and Cl above and below the graft unions of mature trees established in and irrigated in the field for 10 years with 20–30 mM NaCl. Data points represent means and SE for three replicates each of ‘Kerman’-budded PGI and UCB1 rootstocks. Figure 8. Open in new tabDownload slide Bark and annual growth ring (years 10, 9 and 8) wood concentrations of Na and Cl above and below the graft unions of mature trees established in and irrigated in the field for 10 years with 20–30 mM NaCl. Data points represent means and SE for three replicates each of ‘Kerman’-budded PGI and UCB1 rootstocks. Figure 9. Open in new tabDownload slide A diagram of possible transient membrane potentials over the course of exposure to high salinity at a single point in a root or stem that is consistent with results. Negatively charged spheres in living cells indicate the proteins, which generally contribute to a negative charge in the symplast. The positivity or negativity of voltages (V) across membranes in the direction of arrows for each ion (Na+ or Cl-) are described by the Nernst equation. White arrows indicate passive transport generated by negative membrane potentials; black arrows indicate active transport generated by positive membrane potentials. The first `+’ or `-’ in an arrow reflects electrical potential (the charge of the ion under consideration). The second `+ or `−’ in an arrow reflects chemical potential (positive when symplastic concentrations are lower than adjacent apoplastic concentrations). Because we are considering flow out of a cell with movement from root to xylem, an additional `−’ factor is included in those calculations outside of the parentheses to reverse the charge of the overall electrochemical potential. If vacuolar compartmentalization of Na and passive efflux of Cl over the course of exposure (from `initial’ to `over’) maintains homeostasis in all cytoplasm after initial exposure, the only tissue in which concentrations change is xylem vessels. As sap concentrations increase, this leads to the passive efflux of Na from the xylem parenchyma only at some point beyond `over exposure’, when vacuolar sequestration capacity is exceeded and cytoplasm concentrations in the xylem parenchyma increase beyond vessel concentrations (perhaps what we observe in the 150 mM treatment of Experiment 1). The reverse is true for Cl, which is likely to move past xylem parenchyma unless treatments are extremely high, leading to transient moments when high sap concentrations trigger Cl’s passive flow into parenchyma cells, but also the halt of Cl’s passive efflux out of root cells into the sap, limiting further parenchyma accumulation. Represented concentrations of Na+ are relative to concentrations of Na+ and concentrations of Cl− are relative to concentrations of Cl− across compartments and exposure levels. Figure 9. Open in new tabDownload slide A diagram of possible transient membrane potentials over the course of exposure to high salinity at a single point in a root or stem that is consistent with results. Negatively charged spheres in living cells indicate the proteins, which generally contribute to a negative charge in the symplast. The positivity or negativity of voltages (V) across membranes in the direction of arrows for each ion (Na+ or Cl-) are described by the Nernst equation. White arrows indicate passive transport generated by negative membrane potentials; black arrows indicate active transport generated by positive membrane potentials. The first `+’ or `-’ in an arrow reflects electrical potential (the charge of the ion under consideration). The second `+ or `−’ in an arrow reflects chemical potential (positive when symplastic concentrations are lower than adjacent apoplastic concentrations). Because we are considering flow out of a cell with movement from root to xylem, an additional `−’ factor is included in those calculations outside of the parentheses to reverse the charge of the overall electrochemical potential. If vacuolar compartmentalization of Na and passive efflux of Cl over the course of exposure (from `initial’ to `over’) maintains homeostasis in all cytoplasm after initial exposure, the only tissue in which concentrations change is xylem vessels. As sap concentrations increase, this leads to the passive efflux of Na from the xylem parenchyma only at some point beyond `over exposure’, when vacuolar sequestration capacity is exceeded and cytoplasm concentrations in the xylem parenchyma increase beyond vessel concentrations (perhaps what we observe in the 150 mM treatment of Experiment 1). The reverse is true for Cl, which is likely to move past xylem parenchyma unless treatments are extremely high, leading to transient moments when high sap concentrations trigger Cl’s passive flow into parenchyma cells, but also the halt of Cl’s passive efflux out of root cells into the sap, limiting further parenchyma accumulation. Represented concentrations of Na+ are relative to concentrations of Na+ and concentrations of Cl− are relative to concentrations of Cl− across compartments and exposure levels. In both sap (Figure 5a) and wood (Figure 5b), Na concentrations drop significantly along the transpiration stream. These parallel declines suggest retrieval and subsequent storage in the stem’s xylem parenchyma, stem filtration working in tandem with root exclusion. The specifics of retrieval or the fate of sequestered Na is not well understood for trees but has been studied extensively in annual crop plants, where tolerance is often improved by the introgression or upregulation of genes that increase a plant’s capacity for retrieving Na with ion transport across plasma membranes or the capacity of adjacent tissues (e.g., roots, stems, petioles, midribs) to compartmentalize these salts into vacuoles with ion transport across tonoplasts. The relevance to salt tolerance of genes associated with xylem retrieval and sequestration of Na has been studied in alfalfa (Sandhu et al. 2017), Arabidopsis (Sunarpi et al. 2005), mustard greens (Rajagopal et al. 2007), rice (Ren et al. 2005), tomato (Zhang and Blumwald 2001) and wheat (Byrt et al. 2007), among others. Direct observations of declining Na sap concentrations have been made in barley (Wolf et al. 1991) and sweet pepper (Blom-Zandstra et al. 1998). However, studies emphasizing the importance of xylem retrieval as a salt tolerance mechanism in annual crops may not adequately elucidate the strategies of woody perennials, as annual plants differ markedly in both the relative volumes they offer for storage and death at season’s end. In the case of perennial plants, parenchyma cells may represent greater volumes relative to root and leaf tissues, but they may also be required to survive multiple seasons, potentially limiting their Na storage capacity over time. We provide some evidence that both wood and bark in mature trees grafted to the ‘Kerman’ scion provides long-term storage, with concentrations of both Na and Cl either constant or increasing with movement away from the cambium (Figure 8). However, concentrations of these ions in irrigation solution did not approach the maximums we tested in seedling experiments—while Cl levels in the wood, for example, were about half what we observed in analogous seedlings, Na levels were as much as 10-fold lower. We are thus left with several lingering questions. In a higher concentration scenario might short-term (several years) tolerance be improved by increasing the volume of living cells in the secondary xylem or transporters across their membranes, only to backfire with the transition from sapwood to heartwood, when these cells die? Are there differences in storage cycles between deciduous and evergreen species—are some salts released from these cells during senescence and so purged in dropped leaves? Concentrations of Na and Cl in wood were comparable to concentrations in bark regardless of tree age (Figures 6 and 8). Are there variable degrees of connection between xylem and phloem depending on species or cultivar through the ray parenchyma and how does this influence ion storage and transport between wood and bark? Does greater connection bolster or hinder salt tolerance? Chloride was treated differently in Pistacia. As already noted, Na and Cl exclusion between soil solution and stems was comparable, the sap extracted at root crowns having roughly the same concentrations as a percentage of what was applied (Figure 4). Also already noted, the concentrations of Na and Cl that accumulated in leaves over 2 months were comparable, particularly for immature leaves, which demonstrated a clear 1:1 ratio (Figure 7). However, the apparent free passage of Cl through both roots and wood beyond some initial epi- or endodermal gatekeeper coupled with relatively constant xylem sap concentrations throughout the stem, indicates that Cl is not immured in cells like Na. How, then, is its leaf accumulation kept to levels comparable to those of Na? We reason that Cl does not accumulate in leaves due its export from leaves via the phloem. Phloem recirculation of both cations, like Na, and anions, including Cl, has been elegantly reported in castor bean (Jeschke and Pate 1991, Zhong et al. 1998). High concentrations of large negatively charged proteins in the cytoplasm of most cells yields negative Nernst equilibrium potentials so that passive inward anion transport across the plasma membrane is only favored if external concentrations are equal to or higher than internal concentrations. If transporters for active uptake of Cl are minimal, this anion might then passively cycle through a plant continuously, only very slowly accumulating beyond some equilibrium concentration balanced among soil solution, apoplast and symplast. Furthermore, if this reasoning is correct, and phloem recirculation contributes to protecting Pistacia against Cl toxicity (and to a lesser extent Na toxicity), this system may be greatly influenced by budding. The formation of sieve plates from independent mother cells at the rootstock–scion junction, and so the lack of shared middle lamellae, can lead to discontinuities in the phloem (Jeffree and Yeoman 1982). Any such discontinuity would demand the unloading and reloading of phloem contents. Indeed, this is consistent with the observed effects of rootstock–scion combination on both retrieval (Table 2) and exclusion (Table 1) from the xylem sap. The impact of budding is complicated if not confusing, but some general points can be made. Sodium retrieval is maintained along the stem for both 50 and 100 mM Na without any noticeable effect of rootstock–scion combination (Figures 5a and b). However, comparing ungrafted rootstocks with self-grafted rootstocks we observe a jump in Cl sap concentration across the bud union (from position 2 to position 3) in cases where Cl concentrations in sap at the root crown are high. This jump makes sense from the perspective of Cl recirculation in the phloem, which would require active reloading at the union between scion and rootstock and perhaps some greater interaction with the apoplast. We do not observe a visible jump in the Cl concentrations in unbudded trees at any point, supporting the idea that phloem discontinuity may be introduced by budding and influence Cl transport. However, ‘Kerman’-budded trees did not show any dramatic change in Cl concentration at the junction, possibly due to increased exclusion levels, making the overall Cl transport process more difficult to interpret. It may also be that the ‘Kerman’ scion is better equipped to retrieve and sequester Cl with active transporters in its plasma membranes or tonoplasts than the tested self-grafted rootstock species, reducing the Cl recirculated and so the Cl that builds up at the union (Figure 5d). The influence of variable scions on rootstock exclusion or retrieval requires further study, but is part of the reason that we undertook budding treatments and feel that it is important to address that there may be variability in the degree of discontinuity between the sieve elements of one cultivated variety/species and another, complicating ion recirculation even when compatibility is otherwise satisfactory. More decisively, we return to retrieval and sequestration, a system combining the capacities of roots and stems to effectively filter Na from xylem sap, but within limits of external concentration that vary by cultivar or species. After just 1 month of treatment, a constant maximum Na concentration was reached in all tissues (0.5–0.6 mmol Na g−1 DW, Figure 1) in seedling UCB1s, the rootstock with the best-performing dual root and stem filtration system we observed (Figure 5). Furthermore, well before this point (in time and applied salinity) the development or growth of new shoots was recessed (Figure 1c). Experiments comparing woody crop species or cultivar salinity tolerance must clearly focus primarily on the resilience of growth and physiology to low stem water potentials if we want crops that sustain high yields under saline conditions. There may, however, be interactions between the breadth of mechanisms that woody perennials use to minimize specific ion accumulation and the essential maintenance of turgor. Greater consideration of phloem recirculation, for anions especially, may prove fruitful in that circular flow challenges presumptions that what gets into the shoot, and there provides some osmotic benefits, must end up in the leaves, doing harm. Acknowledgments The authors would like to acknowledge Austin Cole of the UC Davis ICPMS laboratory for processing our samples and Emilio Laca of UC Davis for his statistical advice. Undergraduate assistance from Mateusz Zwieniecki, Benjamin Taylor, Eduardo Mazariegos, Jessica Orozco and Henry Calhoun was also essential. Conflict of interest None declared. Funding Funding for this work was provided by the California Pistachio Research Board, the Henry Jastro Graduate Research Award and the UC Davis Horticulture and Agronomy Graduate Group. The authors would also like to thank Duarte, S & J and Pioneer Nurseries for their rootstock donations and David Peters for his assistance with budding. 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TI - Sodium interception by xylem parenchyma and chloride recirculation in phloem may augment exclusion in the salt tolerant Pistacia genus: context for salinity studies on tree crops
JF - Tree Physiology
DO - 10.1093/treephys/tpz054
DA - 2019-08-01
UR - https://www.deepdyve.com/lp/oxford-university-press/sodium-interception-by-xylem-parenchyma-and-chloride-recirculation-in-hS3C6cME0m
SP - 1484
VL - 39
IS - 8
DP - DeepDyve
ER -