Bundle-sheath cell regulation of xylem-mesophyll water transport via aquaporins under drought stress: a target of xylem-borne ABA?

Authors

  • Arava Shatil-Cohen,

    1. Faculty of Agriculture, Food and Environment, The Robert H. Smith Institute of Plant Sciences and Genetics in Agriculture, The Hebrew University of Jerusalem, Rehovot 76100, Israel
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  • Ziv Attia,

    1. Faculty of Agriculture, Food and Environment, The Robert H. Smith Institute of Plant Sciences and Genetics in Agriculture, The Hebrew University of Jerusalem, Rehovot 76100, Israel
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  • Menachem Moshelion

    Corresponding author
      (fax 972 8 9489899; e-mail moshelio@agri.huji.ac.il).
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(fax 972 8 9489899; e-mail moshelio@agri.huji.ac.il).

Summary

The hydraulic conductivity of the leaf vascular system (Kleaf) is dynamic and decreases rapidly under drought stress, possibly in response to the stress phytohormone ABA, which increases sharply in the xylem sap (ABAxyl) during periods of drought. Vascular bundle-sheath cells (BSCs; a layer of parenchymatous cells tightly enwrapping the entire leaf vasculature) have been hypothesized to control Kleaf via the specific activity of BSC aquaporins (AQPs). We examined this hypothesis and provide evidence for drought-induced ABAxyl diminishing BSC osmotic water permeability (Pf) via downregulated activity of their AQPs. ABA fed to the leaf via the xylem (petiole) both decreased Kleaf and led to stomatal closure, replicating the effect of drought. In contrast, smearing ABA on the leaf blade, while also closing stomata, did not decrease Kleaf within 2–3 h of application, demonstrating that Kleaf does not depend entirely on stomatal closure. GFP-labeled BSCs showed decreased Pf in response to ‘drought’ and ABA treatment, and a reversible decrease with HgCl2 (an AQP blocker). These Pf responses, absent in mesophyll cells, suggest stress-regulated AQP activity specific to BSCs, and imply a role for these cells in decreasing Kleaf via a reduction in Pf. Our results support the above hypothesis and highlight the BSCs as hitherto overlooked vasculature sensor compartments, extending throughout the leaf and functioning as ‘stress-regulated valves’ converting vasculature chemical signals (possibly ABAxyl) into leaf hydraulic signals.

Introduction

Competition for light and the need for effective spore dispersal in terrestrial plants prompted the evolution of increased height and a vascular system – the xylem – to provide water and minerals from the soil to the photosynthetic cells in the leaf: the mesophyll (reviewed by Pittermann, 2010). Thus, water and dissolved minerals flow from roots to shoots within the hollow dead xylem trachea elements, then from the xylem into the mesophyll (radial flow) across a pipe-like structure consisting of dense parenchymatous cells making up the bundle sheath (BS; see also Figure S1). A layer of bundle-sheath cells (BSCs) enwraps the entire vascular tissue of the leaf, except for the leaf edges where the xylem elements end at the hydathodes, allowing for guttation (Aloni et al., 2003; Leegood, 2008; Pilot et al., 2004), or the marginal vessels in some leaf laminas, such as those of banana (Musa sp., Musaceae; Shapira et al., 2009). Based on the anatomy of the leaf, the BSCs are considered to be a distinct cell type that developed in close association with vascular cells (Kinsman and Pyke, 1998).

In C4 plants (such as Zea mays), BSCs play an important role in CO2 fixation, and have been suggested to conduct aquaporin (AQP)-facilitated transmembrane water flow (Heinen et al., 2009). However, in C3 plants (such as Arabidopsis), BSCs have been considerably less studied. Accumulating evidence suggests a role for BSCs in these plants in controlling the fluxes of solutes and water from the xylem to the mesophyll, and in particular, fluxes of water by regulating the radial hydraulic conductivity of the leaf (Kleaf) (which exists in series with stomatal water vapor conductance). The dynamic regulation of Kleaf has been demonstrated by some researchers (Cochard et al., 2007; Levin et al., 2007), with several studies showing (albeit without explicitly focusing on the BS layer) that Kleaf decreases dramatically in response to many environmental and abiotic stresses (e.g. Martre et al., 2002; reviewed in Sack and Holbrook, 2006). Early studies speculated that the osmotic water permeability coefficient (Pf) of the BSC membrane is too low to support the transpiration stream, and that the apoplastic route therefore serves as the major pathway for xylem-sap transport (Boyer, 1974). This notion gradually changed after the discovery of plant AQPs, suggesting that the plasma membrane might support rapid water transport (Frangne et al., 2001; Kaldenhoff and Eckert, 1999; Lee et al., 2009; Sack and Holbrook, 2006). In some plants, anatomical evidence suggests the involvement of BSCs in the regulation of radial transport: suberin deposits in the apoplast of leaf BSCs appear to render the apoplast impermeable to xylem sap (reviewed in Sack and Holbrook, 2006). There is also physiological evidence: the specific radial transport capability of the BSCs was revealed through the deletion of GDU1, the glutamine transporter in Arabidopsis vascular tissues, which resulted in the enhanced secretion of glutamine via hydathodes, in guttation droplets (Pilot et al., 2004). Thus, the BSCs are highly likely to be the site of the ‘solute-transport-regulating barrier’. The existence of an ion-selective barrier surrounding the xylem was recently demonstrated in the leaves of banana (a C3 plant), where it was shown that K+, but not Na+, ions cross easily from xylem to mesophyll cells (Shapira et al., 2009). In addition, there have been hints of an isolating barrier surrounding the vascular bundle: in Arabidopsis, stress-induced H2O2 was secluded in the vascular parenchyma cells and the surrounding intercellular spaces of BSCs, but remained excluded from the mesophyll (Galvez-Valdivieso et al., 2009). In the boron-tolerant barley (Hordeum vulgare) varietal Sahara, up to 80% of the daily absorption of boron (by the roots) was eliminated via guttation (Oertli, 1962; Sutton et al., 2007).

Logically, the anatomically intermediate location of the BS layer should expose it to long-distance (chemical and/or hydraulic) signals coming from the roots. As soil becomes dry, these signals – important for plant adaptation to water stress – would be transported via the xylem to the leaves, eventually resulting in reduced water loss (stomatal closure) and decreased leaf growth (Schachtman and Goodger, 2008). Indeed, in gradually dehydrated plants, such as tomato (Solanum lycopersicum), sunflower (Helianthus annuus), maize (Zea mays), poplar (Populus sp.) and Arabidopsis, the concentration of the stress phytohormone ABA progressively increases in the xylem sap (Parent et al., 2009; Holbrook et al., 2002; Tardieu and Simonneau, 1998), very likely from its site of production in the vascular parenchyma tissue (Endo et al., 2008; Galvez-Valdivieso et al., 2009). Surprisingly, however, the xylem-sap ABA (ABAxyl) did not have a direct effect on stomatal closure under stress conditions in Arabidopsis or tomato plants (Christmann et al., 2007; Holbrook et al., 2002). Nonetheless, a recent study reported ABA effects on AQP content and changes in hydraulic conductivity in maize plants (Parent et al., 2009). Thus, the role of ABAxyl in the long-distance root-to-shoot signaling pathway, and its involvement in controlling Kleaf, are not fully understood. Finally, a very recent study has suggested that radial flow to the leaf is not only directly dependent on stomatal movement, but may be controlled by the vascular BS (Ache et al., 2010). Using transmission electron microscopy, Ache et al. revealed numerous symplastic connections between Arabidopsis BS and mesophyll cells, which are absent between the BS and vascular cells. They therefore proposed that the BS is the bottleneck in leaf hydraulic conductance.

Motivated by the evidence and suggestions discussed, in this study we tested the hypothesis that the BSCs of Arabidopsis (a model C3 plant) play a key role in controlling Kleaf in response to stress signals arriving through the vascular system. The specific elements of this hypothesis are: (i) under drought stress the increasing concentration of ABAxyl decreases the hydraulic water conductivity of the BS layer by reducing the the BSC membrane water permeability; and (ii) the decrease in membrane water permeability results from the reduced activity of BSC AQPs. Our data support this hypothesis and suggest a key role for the BS as a ‘smart pipe’ that dynamically controls Kleaf and leaf water potential (Ψleaf).

Results

Kleaf response to xylem-fed ABA or HgCl2

To study the effect of [ABA]xyl on Arabidopsis Kleaf, we used a ‘detached-leaf’ approach that allowed us to feed ABA directly to the xylem via the petiole. Leaves excised before dawn were immediately immersed (petiole-deep) in artificial xylem sap (AXS) with or without ABA. The transpiration rate (E) and water potential (ψleaf) of the leaves were measured (see Experimental procedures), yielding a calculated Kleaf (ratio of E to ψleaf) for well-watered plants of roughly 12 mm m−2 sec−1 MPa−1 (Figure 1a). The high similarity to previously reported values (reviewed by Sack and Holbrook, 2006) validates our ‘detached-leaf’ approach. Feeding the leaf via its petiole with 10 μm ABA or 50 μm of the AQP blocker HgCl2 added to the AXS decreased Kleaf by nearly 50% (Figure 1a). These treatments considerably reduced ψleaf, making it 79 and 111% more negative, respectively, than controls (Figure 1c). In contrast, smearing ABA on the leaf surface, while reducing transpiration, had no effect on Kleaf or ψleaf (Figure 1a,c). Both ABA treatments – xylem (petiole) feeding and leaf smearing – reduced the leaf transpiration rate, by 24 and 22%, respectively, of control values (Figure 1b), indicating the partial closure of the stomata. As ψleaf represents the balance between water entering the leaf via the xylem and water leaving the leaf as vapor via the stomata, a decrease in this parameter signifies increased water deficit in the leaf. In the ABA-smeared leaf, the diminished water loss from the leaf resulting from the partial closure of the stomata could not reverse the water deficit, i.e. it did not decrease ψleaf.

Figure 1.

 Leaf hydraulic conductance (Kleaf) decreases in response to xylem-fed ABA or HgCl2. Leaves picked before dawn were either fed through the petiole (‘xylem fed’) with 10 μm ABA or 50 μm HgCl2, or both leaf surfaces were smeared (using a fine paint brush) with 10 μm ABA or artificial xylem sap (as a control). After 2–3 h, Kleaf.
(a) was calculated for each individual leaf by dividing the whole leaf transpiration rate E.
(b) by the absolute value of the leaf water potential Ψleaf.
(c) Different letters above the columns represent significant differences between treatments (Student’s t-test, P < 0.05). Data are means (±SEs) of values from the indicated number of repeats from at least three independent experiments.

In the xylem (petiole)-fed leaf, the reduction of ψleaf (the increase in water deficit) relative to controls must have stemmed from a change in balance: depressed water influx into the leaf relative to water efflux from the leaf. In other words, xylem-fed ABA and HgCl2 reduced Kleaf mainly by reducing water influx into the leaf via the radial ‘trans-BSC (xylem-to-mesophyll) pathway’ (and thus by decreasing ψleaf), and affected Kleaf less by decreasing water vapor efflux via stomatal closure (i.e. by decreasing E).

As the relevant effects of ABA and HgCl2 are on the cell membrane, they can only affect the overall hydraulic conductance of the leaf if the water permeability of the apoplastic element in the radial xylem-to-mesophyll path is negligible compared with the water permeability of the parallel transmembranal element. To test this, we assayed the permeability of BSC apoplast to a water solution of the cell-wall-specific dye Calcofluor White (CW) (see Experimental procedures).

BSC apoplast permeability to radial sap flow

The CW dye spread readily with the transpiration stream through all of the leaf veins, including the densely branching minor veins within the mesophyll, and vascular staining in the whole leaf was complete within less than 1 h (Figure 2a,b). In contrast, the mesophyll cells, invisible on the dark background outside the veins, did not stain, even after 48 h of submerging the leaf petiole in a high concentration of the dye (1 g l−1; Figure 2b,h), suggesting no leakage of the dye outside the veins. Only a few more strongly stained dots appeared on the leaf margins, very likely associated with guttation from hydathodes at the open ends of the xylem veins (compare Figure 2b,c). Notably, a 1-min exposure to the dye (after peeling away the epidermis), followed by intensive washing, sufficed to demonstrate that the mesophyll cell walls are capable of immediate strong staining with CW (Figure 2g). The absence of radial leakage of the dye from the veins thus attests to a tight apoplastic barrier sealing off the vascular bundle. As water does flow through this tight barrier from the xylem into the mesophyll, it must flow exclusively via the BSC membranes. This suggests the BSC membrane as a potential site for Kleaf regulation.

Figure 2.

 The apoplast of the bundle-sheath cell (BSC) layer has limited permeability to radial sap. Arabidopsis leaves picked just before dawn were immersed upon excision (petiole deep) in concentrated (1 g l−1) Calcofluor White (CW), a fluorescent dye that stains cellulose in the cell walls. The leaf CW fluorescence (380-nm excitation; 475-nm emission) was imaged:
(a) 1 h after lights were turned on; (b) after 48 h in the dye. Note the lack of CW staining outside of the veins, except at a few points on the leaf margins (arrows), probably associated with hydathodes.
(c) Guttation through the hydathodes in an intact leaf (arrows) appearing just before dawn. The fact that the pressurized xylem sap (leading to guttation) did not flood the air spaces within the mesophyll (as judged by the uniformly light color of the leaf) serves as additional evidence that the BSC layer is hydraulically isolated, and that the apoplastic water route across this layer is negligible.
(d) A fluorescent image (488-nm excitation; 520-nm emission) of a leaf expressing GFP under the Scarecrow promoter. Boxed areas were enlarged to show: (e) BSCs of secondary vessels and (f) primary BSCs of the midrib.
(g) Fluorescent image (380/475 nm, as above) of mesophyll tissue from a leaf with peeled off epidermis stained directly by CW for 1 min, and then washed intensively. Note the well-stained cell walls of the mesophyll cells.
(h) Larger magnification of a leaf area (of the size of the top square in d), demonstrating the absence of radial CW leakage (from veins to mesophyll) after 48 h. Note that CW stains only the vein-bordering inner cell walls of the ‘pipe’ formed by the BSCs, invisible here and seen in (i).
(i) Fluorescent image of the GFP-labeled BSCs superimposed on a bright-field image of the mesophyll. Note the confinement of GFP to the tight layer of BSCs around the secondary veins, and its absence in the adjacent mesophyll cells.
(j) BS protoplast in transmitted light, and (k) showing green (GFP) fluorescence surrounding the autofluorescence of the chloroplasts (488 nm excitation and two emission wavelengths: green, 520 nm; red, 650 nm). Scale bars: a–d, 1 mm; e–f, 200 μm; g–i, 100 μm; j–k, 10 μm.

To examine the link between BSCs and the regulation of Kleaf, we measured the osmotic water permeability (Pf) of the BSC membrane directly. The BS protoplasts could be distinguished from mesophyll protoplasts within the isolated protoplast mixture (for convenience, we will henceforth use ‘cells’ and ‘protoplasts’ interchangeably), by their GFP label (Figure 2d–f,i–k; see Experimental procedures) (Kupper et al., 2007; Wysocka-Diller et al., 2000).

Physiological and morphological comparison of BS and mesophyll cells

Within the BSCs themselves, those surrounding the leaf midrib veins (‘main BSCs’) and those surrounding the secondary, tertiary and quaternary veins (‘secondary BSCs’; obtained separately as described in Experimental procedures) did not differ in size, number of chloroplasts or membrane Pf (Figure S2). Because they were more numerous, we mainly used protoplasts from secondary BSCs.

Bundle-sheath protoplasts were about 50% smaller than mesophyll protoplasts [the mean diameters of those used in the assays were, respectively, 23.6 ± 1.2 (SE) μm, n = 38 and 48.0 ± 1.0 μm, n = 46], and contained less chloroplasts than the mesophyll cells (on average, respectively, 20.5 ± 0.9, n = 33, and 48.5 ± 1.7, n = 33).

The mean Pf of the BSCs was significantly lower than that of the mesophyll cells (5.5 ± 0.4 μm sec−1, n = 126 and 9.3 ± 1.8 μm sec−1, n = 80, respectively; Figure 3a). The frequency distribution of the Pf values (which we examined because of the much smaller standard deviation in BSCs than in mesophyll cells, 4.4 and 15.9 μm sec−1, respectively; Figure 3c,d) was much more symmetrical in the BSCs than in the mesophyll cells. We therefore sorted the mesophyll cells (crudely) into two groups: low Pf (<25 μm sec−1), comprising ∼90% of the population; and high Pf (≥25 μm sec−1), comprising ∼10% of the population. Whereas the mean Pf of the high-Pf group of mesophyll cells was 58 μm sec−1, that of the low-Pf group was 5.4 ± 0.6 μm sec−1 (n = 74), equal to the Pf of the BSCs (Figure 3b).

Figure 3.

 The osmotic water permeability, Pf, in bundle-sheath cell (BSC) and mesophyll cell protoplasts.
(a) Mean Pf (±SE) of BSC protoplasts (n = 126) compared with the mean Pf of the whole mesophyll protoplast population (n = 80). *Significant difference (Student’s t-test, P < 0.05).
(b) Mean Pf of BSC protoplasts compared with the mean Pf of the ‘low-Pf group’ of mesophyll protoplasts (with Pf ≤ 25 μm sec−1, n = 74; i.e. comprising over 90% of the population).
(c, d) Frequency distributions of the Pf data of the cells in (a).
Note the ‘tail’ of the outstanding Pf values (>25 μm sec−1) of the ‘high-Pf’ group of mesophyll protoplasts. Data are from at least five independent experiments.

Impact of abiotic stress signals on BS and mesophyll cell Pf

To study the effect of abiotic stress signals on the Pf of BS and mesophyll cells, we exposed the plants to ‘drought’ for 8–10 days (Experimental procedures). This treatment resulted in a 28% reduction in BSC Pf (Figure 4a), but had no effect on mesophyll cell Pf (Figure S3; Table 1). An even stronger response (42% reduction in Pf) was seen in BSC protoplasts isolated from untreated control plants and incubated for 1–4 h with 1 μm ABA (Figure 4b). As before, mesophyll cells remained unaffected by this treatment (Figure S3; Table 1). A few minutes of treatment with 1 μm ABA did not affect Pf in either cell type (Table 1).

Figure 4.

 Drought, ABA and HgCl2 treatments decrease osmotic water permeability (Pf) of bundle-sheath cell (BSC) protoplasts.
(a) Mean Pf (±SE) of BSC protoplasts isolated from well-irrigated Arabidopsis plants (control), and from plants subjected to 8–10 days of ‘drought’ (averaged over the indicated numbers of repeats).
(b) Mean Pf (±SE) of control BSC protoplasts and protoplasts pre-treated (for 1–4 h) with 1 μm ABA.
(c) Effects of HgCl2 on Pf (mean Pf ± SE). The first assay was performed in a control solution after a wash with control solution or after pre-treatment (for up to 10 min) with 50 μm HgCl2. The second assay of the same cells was performed after a 10-min flush with isotonic solution containing 2 mm DTT.
*Significant difference from control (Student’s t-test, P < 0.05). Data from at least three independent experiments for each treatment.

Table 1.   Effects of drought and ABA on the osmotic water permeability (Pf) in protoplasts isolated from bundle sheath cells (BSCs) and mesophyll cells (MCs)
TreatmentBSCsMCs
Low Pf populationWhole population
ControlTreatedControlTreatedControlTreated
  1. Plants were subjected to 8–10 days of drought, or protoplasts were subjected directly to 1 μm ABA; n is the number of protoplasts assayed.

  2. *Significant difference between treated protoplasts and the corresponding control group (Student’s t-test, P < 0.05). Data are from at least three independent experiments.

Drought4.4 ± 0.6 (n = 20)2.9 ± 0.4* (n = 15)5.29 ± 1.0 (n = 30)4.7 ± 0.7 (n = 33)25.1 ± 4.5 (n = 46)21.2 ± 4.4 (n = 46)
ABA (1–4 h)5.3 ± 0.70 (n = 42)3.1 ± 0.5* (n = 28)6.6 ± 1.1 (n = 30)6.1 ± 1.4 (n = 23)14.8 ± 3.8 (n = 35)10.3 ± 3.2 (n = 25)
ABA (≤10 min)4.5 ± 0.8 (n = 14)4.5 ± 1.0 (n = 14)6.8 ± 1.6 (n = 12)5.5 ± 1.8 (n = 12)10.4 ± 3.2 (n = 14)15.4 ± 6.9 (n = 14)

Relevance of Pf to transpiration and involvement of AQPs in BSC Pf changes

Can a BSC Pf of ∼5 μm sec−1 support the transport of water though the whole plant? According to our estimate (Appendix S1), BSCs with this Pf can support the transpiration of 4.6 ml g−1 leaf (fresh weight) per day. The measured transpiration of the whole Arabidopsis plant was ∼2 g water g−1 (fresh weight) per day (see Experimental procedures), hence the BSCs – in their conducting mode – would not constitute a limiting factor to water flow through the leaf.

The distinct Pf changes in the BSCs in response to stress signals led us to address the involvement of their AQPs in these responses. Treatment with the AQP inhibitor HgCl2 (50 μm) nearly halved the rate of BSC swelling (Figure S4), and reduced Pf by threefold (Figure 4c). The reduction in Pf was specific to AQPs, rather than a general effect of HgCl2 toxicity, as we used only a short, up to 10-min treatment, and the effect was reversed by a 10-min wash with 2 mm DTT (Figures 4c and S4). The leftward shift of the Pf distribution histograms of the HgCl2-treated BS protoplasts (Figure S5) indicates a homogeneous effect on the whole BSC population. Blockage by HgCl2, and even more so its reversal, strongly indicated that it is the regulation of AQPs in BSCs that underlies the changes in their Pf values. Unfortunately, mesophyll cells were overly sensitive to the same HgCl2 treatment, precluding a determination of their Pf values.

Discussion

The BSCs: chemical–hydraulic transduction

We demonstrated that the leaf BSCs enclose and isolate the dead xylem conduits from the major tissue of the leaf: the mesophyll. In our experiments, the BSCs acted as a barrier not only to a small solute, CW, but also, notably, to water; we were able to manipulate the permeability to water experimentally. The fact that the pressurized xylem sap at the end of the night did not flood the air spaces within the mesophyll, but released water as guttation via hydathodes, serves as additional evidence that the BS layer is hydraulically isolated, and that the apoplastic route across this layer is negligible, even for water. Interestingly, experiments with a mutant Arabidopsis line (ost1-2), defective in the ABA-signaling pathway that closes stomata, and therefore with constitutively wide-open stomata, also indicated the existence of a hydraulic barrier within the leaf (Ache et al., 2010). Nevertheless, the series of events observed with ABA feeding does not necessarily reflect the series of events initiated by drought stress. The changes in Kleaf caused by the direct application of ABA or HgCl2 to the leaf xylem attest to live cells, rather than apoplast, comprising the hydraulic barrier, which is the target of these effectors. That this barrier consists of the BSC layer finds support in the same effects produced in protoplasts isolated from the BSCs: decreased permeability to water by drought and by direct application of ABA or HgCl2.

Notably, we calculated the Pf equivalent of the Kleaf of well-watered plants, assuming that the barrier characterized by Kleaf is a semipermeable membrane (Appendix S2), to be about 30 μm sec−1. This is in the same ballpark as the Pf determined experimentally in BSC protoplasts (about 5 μm sec−1)!

In addition to identifying the BSCs as the underlying elements of the intraleaf barrier, with modifiable hydraulic water permeability, the sensitivity of this barrier to blockage by HgCl2, paralleling the similar – and reversible – HgCl2 sensitivity of the isolated BSC protoplasts, indicates the participation of AQPs as the molecular water conduits in the BSC membranes, and the targets of modification of the hydraulic barrier by physiological signals.

Taken together, these results suggest a fresh view of BSCs: these cells, tightly wrapped around the entire vascular system, act as the live walls of ‘smart pipes’ arranged in a network: they sense the stress signals within the xylem sap and respond by changing their ‘across-the-pipe wall’ (radial) hydraulic conductivity. This is highly likely to occur via the downregulation of their AQP activity. This, in turn, results in reduced water flow into the leaf, and, consequently, to the decline in ψleaf. Whereas a higher abundance of AQPs in the BSCs compared with their adjacent mesophyll and parenchyma cells has been reported in a relative of Arabidopsis –Brassica napus (Frangne et al., 2001) – to the best of our knowledge, there is no experimental evidence in the literature linking ABA simultaneously to the macroscopic regulation of Kleaf and the cellular-level regulation of Pf and AQPs in BSCs.

Conclusions

The flow of water in the soil–plant–atmosphere continuum is driven by a potential gradient, and is also governed by the hydraulic conductivity of the pathway. In this study, we establish a role for BSCs as a stress signal-sensing ‘control center’ for this water flow (confirming the hypothesis formulated by Sack and Holbrook, 2006). This control center converts the xylem chemical signal – which we assume is ABAxyl– to a leaf water potential (ψleaf) signal. We posit that it is this lower ψleaf that later leads to stomatal closure. In this new role as a vascular control center, the BSCs serve as a ‘first line of defense’ that, in response to drought-stress signals from the root, might temporarily block the vasculature–mesophyll water pathway. According to this concept, the vascular BSCs act in series with the stomata. If the ‘BSC block’ precedes stomatal closure, the water-use efficiency of the whole plant may be increased (Figure 5).

Figure 5.

 The bundle sheath (BS) as a valve controlling the passage of water through a transpiring leaf: the ‘smart pipe’ model. Artist’s rendering of a leaf section (towards the leaf tip), with the cuticle-covered epidermis (Ed) partially removed, featuring the BS ‘smart pipe’ supported by the scaffold of dead xylem trachea (Xt) elements and illustrating regulation by the BS cells (BSCs) of the radial transport of xylem sap (Xs).
(a) At night, stomata and their guard cells (S/GC) are closed and high pressure builds up in the pipe, pushing the xylem sap (Xs) towards the hydathodes to be exuded as guttation drops (GD; (Aloni et al., 2003; Pilot et al., 2004). The leaf water potential (Ψleaf) is high and the leaf maintains full turgor.
(b) During the day, stomata are open and evaporation (E) of water out of the stomata is faster than radial water influx via the BSCs (RI). Note that in our model most of the radial flow (RI) is transcellular, via the BSCs, and not extracellular between them (tiny horizontal blue arrows). Leaf water potential (Ψleaf) and turgor are lower than in (a). Increasing the levels of ABA in the xylem sap (e.g. under drought) reduces the radial symplastic influx to the leaf via the BS and decreases leaf hydraulic conductance (Kleaf). The stomata are still relatively open at this early stage of drought, and although transpiration continues, Ψleaf decreases.
(c) Recapitulation of the water path, from xylem to atmosphere. The flow of water is determined by the water-potential differences along the path. The flow rate is limited by Kleaf. The BSCs tightly envelop the vascular bundle containing the leaky water tube (xylem; the BS ‘smart pipe’ is shown here with the front part removed). Water channels (aquaporins, AQPs) of the BSCs control their membrane osmotic water permeability (Pf) and determine the flow of water across the BS. Thus, the BS acts like a dynamic ‘valve’ or ‘control center’ regulating Kleaf in response to signals from the xylem. As the concentration of the plant stress hormone, ABA, increases in the xylem, AQP activity in the BSCs is downregulated, reducing water flow into the leaf.

Experimental procedures

Plant materials and growth conditions

Arabidopsis thaliana plants (ecotype Landsberg) comprise the LHG4-10OP two-component system in which the synthetic transcription factor LhG4 is expressed under the promoter of interest. TRANS-activation lines were generated by transcriptional fusion of the promoter Scarecrow (SCR, BS-specific promoter) in front of the chimeric LhG4 and ER-GFP subcloned behind an operator array in the BJ36 vector (Moore et al., 1998). The SCR:LhG4 line and the 10OP:ER-GFP line were a generous gift from Prof. Yuval Eshed, Weizmann Institute of Science, Israel. Those lines were crossed to generate T1 plants in which GFP is expressed specifically in the endodermis and BSCs.

All plants were grown in soil containing (w/w) 30% vermiculite, 30% peat, 20% tuff and 20% perlite (Shacham, Israel). The plants were kept in a growth chamber under short-day conditions (10-h light) and a controlled temperature of 20–22°C, with 70% humidity and light intensity of 75–105 μm m−2 sec−1.

Under well-watered conditions the plants were irrigated at least three times a week, maintaining a high soil water content of ∼60 ± 0.3% (w/w), whole-plant transpiration rate of 77 ± 6.2 ml m−2 h−1 [0.20 ml g−1 (fresh weight) h−1] and leaf water potential of 0.044 ± 0.005 MPa (mean ± SE, n = 23).

On days 8–10 of drought treatment, the non-irrigated plants reached a soil water content of 39.6 ± 1.2% (w/w), and the whole-plant transpiration was reduced approximately by half, i.e. down to 40 ± 1.4 ml m−1 h−1. As expected, the leaf water potential did not change compared with the control irrigated plants: 0.047 ± 0.005 MPa. Because of the isohydric water balance regulation of the Arabidopsis plant (Levin et al., 2007; Martre et al., 2002), values are means ± SEs, n = 21.

GFP localization in the T1 leaf

The specific expression of GFP in the BSCs was verified by epifluorescence using an inverted microscope (Olympus-IX8 Cell-R; Olympus, http://www.olympus-global.com), with the following set of objective lenses: plan apochromat, 60× water immersion, n.a. = 1.2; 20×, n.a. = 0.75; 4×, n.a. = 0.16; and 1.25×, n.a. = 0.04. The CCD cameras were a 12-bit Orca-AG (Hamamatsu, http://www.hamamatsu.com) and DP71 (Olympus). The filter sets were GFP-3035B-000 and TXRED-4040B, with zero pixel shift (Semrock, http://www.semrock.com). All images were processed using Olympus imaging software cell-r for windows.

Protoplast isolation

The lower leaf epidermis was peeled off at the leaf center (when extracting the midrib BSCs, the epidermis right above it was peeled off), and the peeled leaves were cut into small squares and incubated in enzyme solution [∼3.3% w/w of an enzyme mix containing the following enzymes in the given proportions: 0.55 cellulase (Worthington, http://www.worthington-biochem.com), 0.1 pectolyase (Karlan, http://www.karlan.com), 0.33 polyvinylpyrrolidone K 30 (Sigma-Aldrich, http://www.sigmaaldrich.com), 0.33 BSA (Sigma-Aldrich)] in solution containing (in milli molar) 10 KCl, 1 CaCl2, 540 d-sorbitol and 8 2-(N-morpholine)-ethanesulphonic acid (MES), pH 5.7. After 20 min of incubation at 28°C, the leaf tissue was transferred to the same solution, but without the enzymes, agitated at 100 rpm for 5 min or until all protoplasts were released into the solution. The remaining tissue pieces (such as of bottom epidermis) were removed and the remaining solution containing the protoplasts was collected with a cut tip into a 1.5-ml tube. This protoplast isolation procedure resulted in a very high yield of protoplasts (∼20 million protoplasts per g leaf).

Osmotic water permeability coefficient (Pf) measurements

To identify the BS protoplasts labeled with GFP, or the mesophyll cells that were not labeled, we screened the protoplast population using the above filter sets.

The Pf was measured in single protoplasts from the initial (recorded) rate of their volume increase in response to hypo-osmotic challenge (a change from 600 mOsm isotonic bath solution to 500 mOsm hypotonic solution). Pf was determined using a numerical approach, an offline curve-fitting procedure using the pffit program, as detailed previously (Moshelion et al., 2002, 2004; Volkov et al., 2007).

Stress and HgCl2 treatments

Protoplasts were incubated in 1 μm ABA (Sigma-Aldrich) for 1–4 h (long treatment) or for 3 min (short treatment). To determine the effect of the drought treatment, the Pf was determined in protoplasts isolated on the first day as a control, and then in protoplasts from plants not irrigated for 10 days. To determine whether AQPs are involved in the water permeability of the BS protoplasts, the protoplasts were incubated for 10 min with 50 μm of the AQP inhibitor, HgCl2 (Sigma-Aldrich), then Pf was determined and the same protoplasts were incubated with 2 mm DTT (Sigma-Aldrich) for 10 min, and the Pf was determined again.

Kleaf measurements

Arabidopsis leaves (6–9 weeks old, ecotype Colombia*) were excised just before dawn and immediately immersed (petiole-deep) in artificial xylem sap (AXS; containing 1 mm KH2PO4, 1 mm K2HPO4, 1 mm CaCl2, 0.1 mm MgSO4, 3 mm KNO3 and 0.1 mm MnSO4 buffered to pH 5.8 with 1 m HCl or KOH; Wilkinson et al., 1998) in control experiments, or AXS supplemented with either 10 μm ABA or 50 μm HgCl2. For the smearing assay, the leaves immersed in AXS were smeared, using a fine paint brush, either with 10 μm ABA or with AXS (as a control). Smearing was repeated at three intervals during the experiment. The immersed leaves were exposed to a light intensity of 52 μm m−2 sec−1, and the experiment was performed 1–5 h after leaves were excised. The transpiration rate (E) was calculated from measurements of the rate of weight (i.e. water) loss by placing the immersed petiole leaf on a sensitive balance (AB135-S, Classic; Mettler-Toledo, http://us.mt.com), and from measurements of the leaf area (at the end of the experiment). The measurement of weight loss was followed immediately by the determination of leaf water potential (ψleaf) using a pressure chamber (ARIMAD-3000; MRC, http://www.mrclab.com/htmls/home.aspx) and home-made silicon adaptor especially designed for Arabidopsis petioles in order to fit the O-ring of the pressure chamber. After placing the leaf in the chamber and tightly closing it, pressure (N2) was gradually applied until the appearance of water at the cut was observed with the help of a binocular microscope (SZ; Olympus) under the illumination of a cool light (Intralux 5000; Volpi AG, http://www.volpi.ch) directed towards the petiole. Kleaf, Eψleaf (Martre et al., 2002; Sack and Holbrook, 2006), was calculated for each individual leaf by dividing the whole leaf transpiration rate, E, by the leaf water potential, ψleaf. (In our calculation ψleaf = Δψleaf, as the leaf petiole was dipped in AXS at a water potential of 0).

The *Kleaf of the Landsberg ecotype, xylem-fed with 10 μm ABA, was reduced by about 50% relative to control, similar to Kleaf of ecotype Colombia. In our experiments we used the Colombia ecotype in order to enable direct comparison with previous reports on Arabidopsis Kleaf values (Levin et al., 2007; Martre et al., 2002).

Calcofluor white staining

Leaves of 5–6-week-old Arabidopsis (ecotype Colombia) were excised a few minutes before the lights were turned on (‘pre-dawn’) and immediately immersed (petiole-deep) in 1 g l−1 Calcofluor White stain in 0.1 phosphate buffer, pH 7 (18909; Fluka, now Sigma-Aldrich) and returned to the Arabidopsis growth chamber for 1–48 h. After the incubations, leaf fluorescence was imaged by epifluorescence inverted microscopy (Olympus-IX8 Cell-R; 380-nm excitation; 475-nm emission) to see the distribution of the dye in the leaf.

Statistical analysis

The Student’s t-test was used for comparison of means, which were considered to be significantly different at P < 0.05.

Acknowledgements

This work was supported by the Israel Science Foundation (ISF), grant no. 953/07. We would like to thank Professors Arie and Nava Moran for their useful comments.

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