• E-mail
• Print
• Comment
• Font Size
• Digg
• del.icio.us
• Discuss article

Phloem Loading in Two Scrophulariaceae Species. What Can Drive Symplastic Flow Via Plasmodesmata?1

Posted on: Thursday, 9 February 2006, 06:00 CST

By Voitsekhovskaja, Olga V; Koroleva, Olga A; Batashev, Denis R; Knop, Christian; Et al

To determine the driving forces for symplastic sugar flux between mesophyll and phloem, gradients of sugar concentrations and osmotic pressure were studied in leaf tissues of two Scrophulariaceae species, Alonsoa meridionalis and Asarina barclaiana. A. meridionalis has a typical symplastic configuration of minor-vein phloem, i.e. intermediary companion cells with highly developed plasmodesmal connections to bundle-sheath cells. In A. barclaiana, two types of companion cells, modified intermediary cells and transfer cells, were found in minor-vein phloem, giving this species the potential to have a complex phloem-loading mode. We identified all phloem-transported carbohydrates in both species and analyzed the levels of carbohydrates in chloroplasts, vacuoles, and cytoplasm of mesophyll cells by nonaqueous fractionation. Osmotic pressure was measured in single epidermal and mesophyll cells and in whole leaves and compared with calculated values for phloem sap. In A. meridionalis, a 2-fold concentration gradient for sucrose between mesophyll and phloem was found. In A. barclaiana, the major transported carbohydrates, sucrose and antirrhinoside, were present in the phloem in 22- and 6-fold higher concentrations, respectively, than in the cytoplasm of mesophyll cells. The data show that diffusion of sugars along their concentration gradients is unlikely to be the major mechanism for symplastic phloem loading if this were to occur in these species. We conclude that in both A. meridionalis and A. barclaiana, apoplastic phloem loading is an indispensable mechanism and that symplastic entrance of solutes into the phloem may occur by mass flow. The conditions favoring symplastic mass flow into the phloem are discussed.

Phloem transport and partitioning of assimilates are essential determinants of plant productivity in agriculture. In leaves, the mode of phloem loading depends on organization of the interface between mesophyll and phloem that has the maximal surface in leaf minor veins (van Bel and Gamalei, 1991). Nearly 1,000 dicotyledonous plant species have been studied with respect to their minor-vein anatomy (Pate and Gunning, 1969; Turgeon and Webb, 1975; Gamalei, 1990; Gamalei, 2000). Knowledge of the structure of phloem companion cells in minor-vein phloem is very informative. The potential of the plant to use a symplastic and/or apoplastic route for phloem loading is indicated by plasmodesmal abundance between phloem companion cells and bundle-sheath cells. Traditionally, the division into apoplastic and symplastic loaders was made on the basis of plasmodesmal abundance between phloem companion cells and bundle- sheath cells (a structural definition). However, functional analysis is necessary to make conclusions about the real contribution of each mechanism. We will use the terms putative symplastic loader and putative mixed loader to indicate the potential for symplastic phloem loading as reflected by plasmodesmal abundance and to provide a link between traditional terms in structural studies and functional analysis. Two types of companion cells show the highest degree of specialization of their structural features with regard to symplastic and apoplastic loading, respectively. These are intermediary cells (ICs) with highly developed plasmodesmal connections to bundlesheath cells (40-60 plasmodesmata per m^sup 2^ cell surface; Gamalei, 1991) and transfer cells (TCs) that have very few plasmodesmata (less than 0.01 plasmodesma per m^sup 2^ cell surface; Gamalei, 1991) but possess cell wall protuberances that increase the cell surface contacting the apoplast (Pate and Gunning, 1969). Many plant species are difficult to classify into putative apoplastic or symplastic phloem loaders on the basis of the structure of their phloem companion cells. This fact is illustrated by the diagram in Figure 1 where data on plasmodesmal frequencies between phloem companion cells and bundle sheath are shown for 60 plants from 34 families of dicots (Gamalei, 1991). In some species, companion cells have less-abundant plasmodesmata than do ICs, and no cell wall protuberances. In other plants, both different structural types of companion cells are present within one minor vein, e.g. in Asarina scandens (Turgeon et al., 1993). In such plants, both apoplastic and symplastic routes can be expected to operate in phloem loading to a comparable extent.

For apoplastic phloem loading, a reliable model has been provided that shows that the primary event is the active uptake of Sue from the apoplast into the phloem by proton symporters (Riesmeier et al., 1994). The mechanism of symplastic loading of sugars is not yet fully understood. It might be expected that symplastic transfer of sugars from mesophyll into the phloem is nonselective and does not depend on synthesis of specific forms of transport carbohydrates. Yet a correlation between the abundance of plasmodesmata in phloem companion cells and the amounts of raffinose family oligosaccharides (RFOs) in the phloem sap has been noticed (Gamalei, 1984; Turgeon et al., 1993). This correlation was explained by the polymer-trap model of symplastic phloem loading (Turgeon, 1991, 1996). The model postulates that raffinose and stachyose are synthesized from Sue and galactinol in ICs. According to the polymer-trap model, the size exclusion limit of plasmodesmata connecting ICs to the bundle sheath enables the passage of disaccharides such as Sue from mesophyll into the phloem, whereas tri- and tetrasaccharides raffinose and stachyose remain trapped in the phloem. Thus, the synthesis of raffinose and stachyose in ICs should provide favorable concentration gradients for Suc to move from mesophyll into the phloem. Recently, polymer trapping was also assigned a role in helping to reduce solute leakage from the phloem due to lower membrane permeability of RFOs (Ayre et al., 2003).

Figure 1. Plasmodesmal frequencies between mesophyll bundlesheath cells and companion cells as determined in 60 species selected from 34 families of dicotyledonous plants (data taken from Gamalei, 1991). Black circles, ICs; white circles, OCs and other companion- cell types; white diamonds, TCs. Species are distributed along the horizontal axis in decreasing order of plasmodesmal numbers.

In compliance with the polymer-trap model, Sue is expected to be present in the cytosol of mesophyll cells of symplastic species in higher concentrations than in the phloem. Assuming that the passage via plasmodesmata is not regulated, an indiscriminate movement of other appropriately sized molecules should also take place between the phloem and the cytoplasm of mesophyll cells along their concentration gradients. Comparison of concentrations of sugars in subcellular compartments of leaf cells and in the phloem could give an insight into the mode of symplastic phloem loading. However, little information is available on the compartmentation of soluble carbohydrates in mesophyll cells in putative symplastic phloem loaders as compared to apoplastic ones. Thus far, no phloemloading model for putative mixed loaders has been created but two such species, Clethra barbinervis and Liquidambar styraciflua, were suggested to load completely from the apoplast (Turgeon and Medville, 2004). For this study, two Scrophulariaceae species, Alonsoa meridionalis and Asarina barclaiana, were chosen. A. meridionalis has ICs in minor veins (Knop et al., 2001; Knop et al., 2004) and translocates mainly stachyose in the phloem, which indicates that the phloem loading in this plant could occur symplastically by the polymer-trap mechanism. In A. barclaiana, two types of companion cells in minor veins were observed, implying that this species has a complex phloem-loading mode (Knop et al., 2001).

In this study, we aimed at determining the major force(s) governing the symplastic exchange of sugars between mesophyll and phloem. This knowledge is of primary importance for understanding how symplastic phloem loading can operate. For symplastic movement of solutes, two possibilities have been considered; the diffusion of solutes and the mass flow of solution (Tyree, 1970; Mnch, 2003). Diffusion is the mechanism most often implied when symplastic phloem loading is discussed (Turgeon and Medville, 2004). We addressed the following questions: What are the patterns of subcellular compartmentation and concentration gradients for Sue in leaf tissues of the putative symplastic phloem loader A. meridionalis? What are these gradients for the phloem-transported carbohydrates in putative mixed phloem loader A. barclaiana? What are the osmotic pressure gradients in the leaf tissues of both plants?

To answer these questions, we performed a comprehensive analysis of minor-vein structure in A. meridionalis and A. barclaiana, identified all carbohydrates present in the phloem of both species, and analyzed their subcellular localization and apoplastic concentrations. To understand water fluxes within the leaves, gradients of osmotic pressure in epidermis, mesophyll, and phloem were studied in both species.

RESULTS

Comprehensive Analysis of Minor-Vein Anatomy of A. barclaiana and A. meridionalis

The minor veins of A. barclaiana consist typically of two companion cell-sieve element (CC-SE) complexes (Fig. 2, A and B). Companion cells of the \two CC-SE complexes show contrasting structural features. The adaxial CC-SE complex contains one sieve element with two large companion cells that are connected to the neighboring bundle-sheath cells by plasmodesmata organized in small plasmodesmal fields (Fig. 2, C [black arrowheads] and E). Also, the cell wall forms protuberances on the side of the bundle sheath (Fig. 2C, white arrowheads). Companion cells of similar structure have been reported for A. scandens by Turgeon et al. (1993) and termed modified intermediary cells (MICs). The abaxial SE-CC complex comprises one sieve element connected to two TCs lacking plasmodesmata at the bundle-sheath side but possessing cell wall ingrowths (Fig. 2D). The plastids of phloem parenchyma cells are chloroplasts (data not shown), but in both types of companion cells they are represented by leucoplasts (Fig. 2F). Thus, the minor veins of A. barclaiana contain two structurally different CC-SE complexes in the phloem. First, they have the adaxial complex, potentially specialized in both symplastic and apoplastic loading as suggested by the occurrence of plasmodesmal fields and of cell wall protuberances. second, they have the abaxial complex potentially specialized only for apoplastic loading, as there are almost no plasmodesmata available for symplastic transfer.

Figure 2. Minor-vein anatomy and companion-cell structure in A. barclaiana (A-F) and A. meridionalis (G and H) as studied by TEM. A, General view of a minor vein of A. barclaiana, 1,700. B, Enlarged view of the minor vein shown in A, 3,500. MIC, Modified intermediary cell; TC, transfer cell; SE, sieve element. C, MIC, 6,500. Black arrowheads point on plasmodesmal fields, white arrowheads point on cell wall protuberances. D, TC, 10,000. E, Plasmodesmal field at the interface of a MIC and a bundle sheath, 25,000. F, Plastid in a companion cell (arrow), 12,000. G, General view of a minor vein of A. meridionalis, 2,000. IC, Intermediary cell; OC, ordinary companion cell; SE, sieve element. H, Plasmodesmal field at the interface of an IC and a bundle-sheath cell, 14,000.

Minor veins of A. meridionalis (Fig. 2G) have a typical symplastic configuration with two laterally positioned ICs with extensively developed plasmodesmata organized in plasmodesmal fields like the one shown in Figure 2H. The associated abaxial companion cell can be classified as an ordinary companion cell (OC; Fig. 2G; Turgeon et al., 1993) as it possesses multiple, single nonbranched plasmodesmata not arranged into plasmodesmal fields (data not shown).

Levels of Soluble Carbohydrates in Whole Leaves of A. meridionalis and A. barclaiana

HPLC analysis of the leaf extracts of A. barclaiana and A. meridionalis has shown that GIc, Fru, Sue, and two of the RFOs, raffinose and stachyose, as well as the precursors of their synthesis, the cyclitol myo-inositol and its galactoside galactinol, were present in both plants (Knop et al., 2001; Table I). In A. barclaiana, a sugar alcohol mannitol and an unknown compound were also found. The latter was isolated and its chemical structure was determined by NMR as described by Tietze et al. (1980). It was identified as the iridoid glucoside antirrhinoside, which has already been reported from another Asarina species, A. scandens (Gowan et al., 1995). Antirrhinoside made up nearly half of the total content of soluble carbohydrates of A. barclaiana leaves (Table I). We determined that the antirrhinoside concentration in the phloem sap of A. barclaiana was about 800 mM (Table II). Calculated from the data in Table II, antirrhinoside was the second major compound in the phloem of A. barclaiana and accounted for 39% of total carbon transported in the phloem, whereas Sue and mannitol accounted for 48% and 3%, respectively.

Distribution of Carbohydrates between Subcellular Compartments of Mesophyll Cells

The distribution of sugars between chloroplastic, cytoplasmic, and vacuolar compartments of mesophyll cells of A. meridionalis and A. barclaiana is shown in Table III. It should be mentioned that the non-aqueous fractionation technique does not resolve between the cytosol and endomembrane compartments. For instance, fractionation of leaves expressing a single-chain antibody anchored to the endoplasmic reticulum by the KDEL amino acid sequence showed that the antibody was entirely confined to the cytosol (Straufi et al., 2001). We designate this compartment cytoplasm, which includes also mitochondria, nuclei, peroxisomes, and endomembrane compartments other than vacuoles. In both plants, GIc and Fru were confined entirely to vacuoles. Myo-inositol was present in all three compartments but the highest amounts were found in chloroplasts. In A. meridionalis, Sue was distributed between all three compartments with the highest portion being in the cytoplasm. Galactinol was mostly concentrated in the vacuoles and its distribution pattern resembled that of hexoses.

Table I. Sugar contents in the leaves of A. barclaiana and A. meridionalis after 2 h of the light period

Mean values from four independent measurements SD are shown. FW, Fresh weight.

In A. barclaiana, antirrhinoside was mostly present in the vacuoles but chloroplasts and cytoplasm also contained significant proportions. Sue was found in all compartments but half was confined to the vacuoles. Mannitol was distributed between all compartments.

It was not possible to assign raffinose and stachyose to certain compartments in A. barclaiana and A. meridionalis because their concentrations in whole leaves were too low (data not shown). Also, the amount of galactinol in A. barclaiana leaves was reduced to an undetectable level after fractionation (data not shown).

Table II. Phloem sap concentrations determined in A. meridionalis and A. barclaiana

Mean values SD; n = 9 (for antirrhinoside n = 3). Asterisk (*) indicates data taken from Knop, 2001.

Estimation of the Volumes of Mesophyll Tissue in A. meridionalis and A. barclaiana

Knowledge of the volumes of whole mesophyll tissues in leaves of A. meridionalis and A. barclaiana is required for the assessment of subcellular concentrations of carbohydrates in mesophyll cells using the approach of Winter et al. (1993). The direct determinations by morphometric analysis in A. meridionalis and A. barclaiana proved to be extremely laborious. However, the completed morphometric analyses of two dicots, spinach (Spinacia oleracea) and potato (Solanum tuberosum), revealed that the volumes of the mesophyll relative to the leaf weight calculated per milligram of chlorophyll (ChI) were strikingly similar in these species (74% and 75% of the leaf water content per milligram of ChI for spinach and potato, respectively; Winter et al, 1994; Leidreiter et al., 1995). Thus, we applied this relationship to the leaves of A. barclaiana and A. meridionalis. We determined the average water content of 394 L (mg ChI)^sup -1^ for leaves of A. meridionalis and of 498 L (mg ChI)^sup -1^ for leaves of A. barclaiana, and estimated the mesophyll volumes as 296 L (mg ChI)^sup -1^ and 374 L (mg ChI)^sup -1^ for leaves of A. meridionalis and A. barclaiana, respectively.

Verification by Direct Measurements in Single-Cell Samples

To confirm the accuracy of the calculated volumes of the mesophyll, the concentrations of GIc, Fru, and Sue were measured in mesophyll cells by single-cell sampling and compared to equivalent values estimated on the basis of calculated cell volumes. Both methods produced similar results for both hexoses and Sue (Table IV), which justifies the assumptions made for calculations of cell volumes. This allowed us to proceed with the determination of the sugar levels in mesophyll cell compartments based on the partial volumes of compartments estimated from transmission electron microscopy (TEM) photographs.

Table III. Distribution of carbohydrates between stromal (Ch), cytoplasmic (Cyt), and vacuolar (Va) compartments of leaf cells in species with a different phloem-loading mode as determined by nonaqueous fractionation

Estimation of Subcellular Volumes of Mesophyll Cells in A. meridionalis and A. barclaiana

The partial volumes of the vacuolar, chloroplastic, and cytoplasmic compartments made up 72%, 20%, and 8% on the TEM micrographs of mesophyll cells in A. meridionalis and 70%, 23%, and 7% in A. barclaiana, respectively. From these values, the volumes of the stromal, cytoplasmic, and vacuolar compartment of mesophyll cells were determined as 85,26, and 261 L (mg ChI)^sup -1^ for A. barclaiana and 59,24, and 213 L (mg ChI)^sup -1^ for A. meridionalis, respectively. To calculate subcellular concentrations on the basis of these volumes, a correction has to be made that takes into account the volume of the epidermis. This is because vacuoles occupy up to 99% of the volume of epidermal cells (Winter et al., 1993, 1994) and thus, during fractionation of whole leaves, metabolites from the epidermis contribute only to the vacuolar fraction of the mesophyll cells that would result in overestimated vacuolar concentrations. The levels of carbohydrates measured in the epidermal cells of A. barclaiana and A. meridionalis by single-cell sampling and analysis were comparable with the carbohydrate levels in the mesophyll cells (data not shown). Again, as described above, the morphometric relations in potato leaves were used where the epidermis made up 74 L, (mg ChI)^sup -1^ (Leidreiter et al., 1995), and corresponding epidermal volumes were 70 /u,L (mg ChI)" for A. barclaiana and 55 L (mg ChI)^sup -1^ for A. meridionalis, respectively. The addition of the volume of the epidermis to the volumes calculated for mesophyll vacuoles resulted in 331 L (mg ChI)^sup -1^ in A. barclaiana and 268 L (mg ChI)^sup -1^ in A. meridionalis, respectively. These values were taken for the calculation of vacuolar concentrations in mesophyll cells.

Table IV. Concentrations (mM) of CIc, Fm, and Suc in mesophyll cells \of A. meridionalis and A. barclaiana as estimated on the basis of HPLC and calculated mesophyll volumes or measured by single- cell sampling after 3-h-light period

For single-cell sampling, mean values of four independent measurements SD are shown.

Sugar Contents and Concentrations in Compartments of Mesophyll Cells

Sugar contents in compartments of mesophyll cells in A. meridionalis and A. barclaiana are shown in Table V. Altogether, vacuoles contained the highest amounts of total soluble carbohydrates (82% in A. barclaiana and 69% in A. meridionalis), while cytoplasm and chloroplasts contained much less. Subcellular concentrations were calculated for each substance based on the volumes of chloroplastic, cytoplasmic, and vacuolar compartments estimated as described above. These data are shown in Figure 3.

In the chloroplasts of A. meridionalis leaves, myoinositol dominated, followed by Sue. The highest level of galactinol in A. meridionalis was found in vacuoles. In the vacuoles of A. meridionalis mesophyll cells, hexoses were the carbohydrates present at the highest concentrations. In the cytoplasm, Sue was the dominating carbohydrate, followed by myo-inositol.

In chloroplasts of A. barclaiana leaves, the calculated values for mannitol and antirrhinoside were the highest, followed by myo- inositol (Fig. 3). In vacuoles, antirrhinoside concentration was highest, followed by GIc and Sue in much lower concentrations. The dominating carbohydrate in the cytoplasm was also antirrhinoside, followed by mannitol, Sue, and myoinositol.

Table V. Sugar contents in the chloroplastic, cytoplasmic, and vacuolar compartments of mesophyll cells from A. barclaiana and A. meridionalis

Osmotic Pressure in Leaf Extracts, Mesophyll, Epidermis, and the Phloem

The osmolality of the whole leaf sap and of single cells sampled from mesophyll and epidermis was determined at the beginning of the light period (Fig. 4). The values measured in whole leaf saps were 549 mOsmol kg^sup -1^ for A. bardaiana and 365 mOsmol kg^sup -1^ for A, meridionalis. In A. bardaiana, the single-cell osmolalities were 629 mOsmol kg^sup -1^ for the epidermis and 682 mOsmol kg^sup -1^ for the mesophyll. In A. meridionalis, the osmolalities were 491 mOsmol kg^sup -1^ and 487 mOsmol kg^sup -1^ for the epidermis and the mesophyll, respectively. Thus, in both species, osmotic pressure measured in epidermal cells did not differ significantly from that in mesophyll cells.

For osmotic pressure in the phloem sap, the approximate values were calculated from the known concentrations of sugars and amino acids (summarized in Table II). These values were 950 mOsmol kg^sup - 1^ for A. meridionalis and 2,260 mOsmol kg^sup -1^ for A. bardaiana. We consider these estimations to be similar to the real values because the ion concentrations so far determined in the phloem were much lower than the total concentrations of either amino acids or sugars in maize (Zea mays) and Arabidopsis (Arabidopsis thaliana; Lohaus et al., 2000; Deeken et al., 2002), and the osmotic coefficients for sugars are very close to 1.

Sugar Levels in the Apoplast of Leaves with Normal and Inhibited Phloem Translocation

Antirrhinoside (8 mM) and mannitol (4 mM) dominated the apoplast of A. bardaiana leaves, while Suc, GIc, and Fru were present at concentrations below 2 mM (Fig. 5). The apoplastic levels of all these sugars except mannitol increased several fold after the exposure of detached leaves to continuous light for 24 h (Fig. 5). The concentration of antirrhinoside rose up to 24 rnM and that of Sue to 10 mM. In A. meridionalis, the only sugars found in the apoplast were hexoses and their levels did not exceed 1 mM (Fig. 5). After 24 h of exposure of detached leaves to continuous light, apoplastic hexose levels increased to 4.5 mM and 5 mM for GIc and Fru, respectively, and Sue accumulated in the apoplast to a concentration of only 1 mM (Fig. 5).

DISCUSSION

In this study we determined the concentrations of soluble carbohydrates and the osmotic pressures in mesophyll cells and phloem in plants where these tissues are connected via plasmodesmata. The phloem concentrations for most phloem- translocated sugars were found to be at least twice as high as those in the cytoplasm of mesophyll cells indicating that symplastic phloem loading, were this to occur in these species, is unlikely to do so by diffusion. However, the data are consistent with the solute mass flow from mesophyll into the phloem via pressure-regulated plasmodesmata openings.

Distribution Patterns of Carbohydrates and Their Levels in Compartments of Mesophyll Cells Were Similar in Species with Different Minor-Vein Anatomy

The distribution of sugars among compartments of mesophyll cells was studied by nonaqueous fractionation in the putative symplastic phloem loader A. meridionalis and the putative mixed loader A. barclaiana. The data show that the vacuoles are the primary compartments accumulating carbohydrates in the mesophyll cells of these species, as 82% and 69% of the total sugar contents were found in the vacuoles in A. bardaiana and A. meridionalis, respectively (Table V). The individual distributions for each carbohydrate between subcellular compartments of mesophyll cells have been analyzed in A. barclaiana and A. meridionalis and compared with previously obtained figures for apoplastic loaders (Table III). The rationale for this comparison was the assumption that in apoplastic loaders, Sue destined for the phloem transport has to be released from mesophyll cells into the apoplast, whereas in symplastic loaders, sugars synthesized in mesophyll and destined for the phloem transport, are expected to stay in cytoplasm. Thus, it could be expected that the percentage of these sugars in the cytoplasm of mesophyll cells in putative symplastic loaders would be much higher than in the apoplastic loaders, and that this might be compensated by the changes in the subcellular distribution of other metabolites in putative symplastic loaders. However, comparison of the patterns of subcellular distribution of metabolites has shown that they are similar in all plants studied, irrespective of their phloem-loading mode (Table III). This was found for sugars destined for phloem transport, as well as for carbohydrates, which are not translocated. Hexoses, galactinol, and antirrhinoside were mostly located in vacuoles. Suc and mannitol showed more-or-less equal distribution among all three subcellular compartments. Myo-inositol was the only carbohydrate in which the highest proportion is found in chloroplasts.

Figure 3. Sugar concentrations (mM) in subcellular compartments of mesophyll cells of A. barclaiana (black columns, left side) and A. meridionalis (white columns, right side). 1, Myo-inositol; 2, galactinol; 3, mannitol; 4, antirrhinoside; 5, GIc + Fru; and 6, Suc.

Galactinol had been found previously in the mesophyll vacuoles of the apoplastic phloem loader Antirrhinum majus (Moore et al., 1997). Although galactinol-synthesizing enzyme is thought to be cytosolic (Bachmann and Keller, 1995), in A. meridionalis galactinol occurred predominantly in the vacuoles of mesophyll cells, similar to the situation in Antirrhinum. This suggests that the pool of galactinol in the mesophyll is not directly related to RFO synthesis in the phloem. It is possible that in A. meridionalis galactinol is produced not only in the mesophyll but also within ICs where it is used for the synthesis of raffinose and stachyose. This was shown for Ajuga reptans, a plant with two isoforms of galactinol synthase in leaves, one mesophyll specific and one IC specific (Sprenger and Keller, 2000). Also, in Cucurbita pepo, immunolocalization of galactinol synthase protein showed it to locate in ICs (Beebe and Turgeon, 1992). For the synthesis of galactinol in ICs, Sue could be used after its hydrolysis by Sue synthase producing UDP-GIc that can be further converted by UDP-Glc-4 epimerase into UDP-GaI, which is, together with myoinositol, a substrate for galactinol synthase.

The subcellular distribution of the iridoid glucoside antirrhinoside present in A. bardaiana was studied for the first time. Antirrhinoside was mostly located in vacuoles, but, in contrast to hexoses, a significant proportion of it (10%) was distributed between the cytoplasm and chloroplasts. The high proportion of antirrhinoside in chloroplasts also resulted in the calculation of its high concentration for this compartment.

Figure 4. Osmolalities of leaf extracts and of single cells of epidermis and mesophyll in A. barclaiana and A. meridionalis. Samples were taken from plants after a 3-h-light period. Mean values from five independent measurements SD are shown. For comparison, estimations of the osmolalities of the phloem saps of A. barclaiana and A. meridionalis are shown as calculated from the data of Table II.

Figure 5. Apoplastic sugar levels in leaves of A. bardaiana and A. meridionalis at the beginning of the light period (10 AM) and after 24-h exposure of detached leaves to continuous light.

The question arises whether chloroplasts really accumulate antirrhinoside as well as soluble carbohydrates. The chloroplastic pool of myo-inositol probably originates from its synthesis by the stromal isoform of the myo-inositol synthesizing enzyme myo- inositol phosphate synthase (Adhikari et al., 1987). Other carbohydrates located in chloroplasts by the nonaqueous fractionation technique might represent the fraction associated with the chloroplastic outer membrane (Heber, 1957).

In A. meridionalis, Concentration of Sue in the Phloem Was Higher Than in Mesophyll Cell Compartments

Comparison of sugar concentrations in cytoplasm of mesophyll cells and in the phloem sap is shown in Table VI. It should be pointed out that the nonaqueous fractionation technique was developed for determination of subcellular concentrations of metabolites, such as Calvin cycle intermediates, that are \exclusively located in the mesophyll (Gerhardt and Heldt, 1984). For other metabolites such as sugars, which also are present in tissues other than the mesophyll, it tends to somewhat overestimate concentrations in the mesophyll cell compartments, because the amounts of metabolite in other tissues are assumed to be negligible. However, even overestimated subcellular concentrations would not affect our conclusions about the direction of diffusion gradients between mesophyll and phloem as discussed below.

A. meridionalis is the only species with ICs in minorvein phloem that has been studied using the non-aqueous fractionation technique with respect to the ratio of sugar concentrations in the phloem sap and in the cytoplasm of mesophyll cells. Based on its minorvein anatomy and the predictions of the polymer-trap model, Sue is expected to enter the phloem in this species along its concentration gradient via plasmodesmata. However, in A. meridionalis, the Sue concentration in the phloem sap was twice as high as in the cytoplasm of mesophyll cells (Table VI). This difference is statistically significant and indicates that diffusion cannot be a mechanism for symplastic loading of Sue in A. meridionalis. At the same time, this ratio is much lower than those determined for apoplastic phloem loaders (e.g. 16 for spinach; Lohaus et al., 1995) and resembles the values estimated for the putative symplastic loaders peach (Prunus persica; 1.7; Moing et al., 1997) and melon (Cucumis melo; 0.7; Haritatos et al., 1996). The 2-fold concentration gradient is obviously easier to smooth down or even reverse, depending on the metabolic situation, than the much steeper gradients found in apoplastic phloem loaders, so the possibility of symplastic phloem loading of Sue by simple diffusion under some conditions cannot be completely ruled out.

Apoplastic levels of Sue were found to be negligible but showed some increase to about 1 mM after phloem translocation from leaves was blocked for 24 h. An active uptake of Sue from the apoplast into the phloem has been strongly suggested for A. meridionalis by Knop et al. (2004) where the H+/Sue transporter AmSUTl was shown to be present in the plasma membrane of sieve elements and companion cells of A. meridionalis. Together, the data available thus far support apoplastic transfer of Sue into the phloem.

In A. barclaiana, Concentrations of Sue and Antirrhinoside Were Several Times Higher in the Phloem Than in the Cytoplasm of Mesophyll Cells, whereas Mannitol Levels Were Similar

Our study of the minor-vein anatomy of A. barclaiana has shown that there are two companion-cell types in its minor veins, one of which, based on its structural features, is expected to specialize in both symplastic and apoplastic loading (MICs) and another one in apoplastic loading only (TCs). The data on concentration gradients for each translocated carbohydrate are important to reach conclusions about the preferred routes.

In A. barclaiana, the ratio of Sue concentrations between phloem and cytoplasm of mesophyll cells was as high as 22 (Table VI). Similarly high, or somewhat lower ratios have been previously estimated for Suc in the apoplastic phloem loaders spinach (16; Lohaus et al, 1995), barley (Hordeum vulgare; 5; Lohaus et al, 1995), and maize (5; Lohaus et al., 1998). The concentration of antirrhinoside in the phloem sap of A. barclaiana was 6 times higher than in the cytoplasm of mesophyll cells, and almost as high as the phloem concentration of Sue (Table VI). Therefore, the loading of both Sue and antirrhinoside into the phloem in A. barclaiana must be energized. This is consistent with the following observations: First, H+/Sue symporters from A. barclaiana were cloned and their function in Sue uptake confirmed by functional expression and complementation in yeast (Saccharomyces cerevisiae; Knop, 2001); second, transfer of glucosides via the plasma membrane against their concentration gradient has been demonstrated in another Scrophulariaceae species, Digitalis lanata (Christmann et al., 1993), and recently, the Sue transporter AfSUC2 was shown to accept a broad range of glucosides as substrates (Chandran et al., 2003); and third, antirrhinoside and Sue were both present in the apoplast and their apoplastic concentrations in leaves with blocked translocation increased 3-fold and 6-fold, respectively. Thus, in A. barclaiana, Sue and antirrhinoside are likely to be actively loaded into the phloem from the apoplast, perhaps mainly via TCs.

Table Vl. Sugar concentrations (Mm) in cytoplasm of mesophyll cells and in the phloem sap in A. meridionalis and A. barclaiana

For cytoplasm, mean values from five independent nonaqueous fractionations SE are shown. For phloem concentrations, mean values from three to nine independent measurements SE are shown.

The measured total concentration of solutes in A. barclaiana phloem was very high, and antirrhinoside made a significant contribution to it. Also, the osmolalities in cells other than phloem in A. barclaiana were higher than in A. meridionalis, which could be due to the high amounts of antirrhinoside found in all tissues in A. barclaiana. The remarkable difference in total phloem solute concentrations between A. barclaiana and other plants studied in this respect might be explained by the fact that other plants do not translocate glycosides in the phloem. Subtracting the antirrhinoside concentration from the phloem sap of A. barclaiana results in a total concentration of 1,480 m.M, which is still higher than the value for A. meridionalis (950 mM).

The concentrations of mannitol in the phloem sap of A. barclaiana and in the cytoplasm of mesophyll cells were not significantly different (Table VI). This implies that there may be no concentration gradient for the diffusion of mannitol from mesophyll cells into the phloem via the symplastic pathway. Mannitol was found in the apoplast of A. barclaiana leaves but its level did not increase in leaves with blocked translocation, which may indicate that most phloem loading of mannitol occurs symplastically.

Symplastic Phloem Loading Is More Likely to Occur by Mass Flow via Plasmodesmata Than by Diffusion

A. meridionalis has a structural potential for symplastic phloem loading of assimilates as indicated by the presence of plasmodesmal fields in its ICs. The apoplastic loading of Sue is likely to occur in this plant mainly via OCs in both minor and larger veins. It is tempting to speculate that different companion-cell types within the same minor vein may use different loading mechanisms. However, in our present work we focused on the question of which forces could drive symplastic loading of Sue, if this were to occur in two species under consideration. From a thermodynamic point of view, the difference between Sue concentrations in mesophyll and phloem determines the possibility of its diffusion in one direction or another. Companion cells and sieve elements, being connected via pore-plasmodesma units, represent one symplastic domain (phloem) with the same concentration of sugars. Our present data show that the phloem in A. meridionalis has at least 2-fold higher concentration of Sue than the cytoplasm of mesophyll cells. Regardless of the way the Sue was loaded into the phloem, this would prevent any further diffusional movement of Sue from mesophyll into the ICs. In A. barclaiana, the similarity of mannitol concentrations in the cytoplasm of mesophyll cells and the phloem suggests that mannitol could enter the phloem by diffusion through the plasmodesmata. In such a case, however, the concentration gradients for Sue and antirrhinoside would simultaneously favor their diffusion from the phloem into the mesophyll until the concentrations on both sides become equal. As the phloem concentrations of Sue and antirrhinoside remained high, despite the fact that the size exclusion limit of plasmodesmata does not allow the effective retention of disaccharides in the phloem, we conclude that in A. barclaiana also no diffusion of any sugar via plasmodesmata typically takes place between the mesophyll and phloem companion cells during the photoperiod. It should be emphasized that these conclusions apply only to the conditions under which the experiments were performed, i.e. to the mature exporting leaves in the middle of the light period. It is reasonable to assume that the situation will be different under conditions that change the carbohydrate and water status of the plant, e.g. in the night or after prolonged shading or drought stress. Further study is necessary to investigate these situations.

Turgeon and Medville (2004) suggested that in putative symplastic phloem loaders lacking ICs, plasmodesmata are held fully or partially closed by the pressure gradient at the companion-cell/ bundlesheath interface, so that phloem loading in such species can occur only from the apoplast. They further suggested that plasmodesmata in ICs remain open under similar pressure (no gradients). Our data show that if the opening of plasmodesmata in ICs were not regulated in A. meridionalis, then Sue would diffuse back into mesophyll rather than enter the phloem. Unfortunately, it is impossible to predict the exclusion properties of plasmodesmata from electron micrographs. Since the plasmodesmata of ICs and MICs are not exactly alike in appearance, it is reasonable to suppose that the loading mechanisms are different and that the plasmodesmata have different properties. However, it is also plausible to assume that, irrespective of the plant species, plasmodesmata are regulated such that no leakage occurs from the phloem. Hence, two questions arise: (1) What else, if not diffusion, may be the driving force for sugar loading into the phloem via plasmodesmata? (2) What is the mechanism that regulates opening/closure of plasmodesmata at the companion-cell/bundle-sheath interface?

Two major mechanisms have been pro\posed to account for the symplastic movement of solutes via plasmodesmata: the mass flow of solution, and the diffusion of solutes (Tyree, 1970; Munch, 1930). Munch provided convincing evidence that in the case of opposite forces mass flow will overcome the diffusion. Overall, our observations can be explained if the symplastic solute flow is considered as mass flow between the mesophyll and the phloem. The assumption that the gradients of the turgor pressure and of the water potential can direct the flux into the phloem can be supported by the following considerations. To maintain a hydrostatic pressure- driven mass flow from source to sink, persisting water influx into the phloem of source leaves is necessary. That means that a steep gradient of water potential between the phloem and surrounding tissues should exist. As the translocation flux runs through the phloem, and therefore the system is not in static equilibrium, it is reasonable to suggest that the osmotic pressure in functional sieve elements is not balanced by the hydrostatic pressure, thus creating a steep water potential gradient between the phloem and other tissues. This model is supported by available data on osmotic pressure and turgor pressure in transport phloem of barley and sow thistle (Sonchus oleraceus). The measured osmotic pressure in sieve elements of sow thistle source-leaf petioles was 2.0 to 3.0 MPa and the hydrostatic pressure was in the range of 1.0 to 1.5 MPa (Gould et al, 2005). In sieve elements of barley roots, osmotic pressure was in the range of 1.9 to 2.6 MPa and hydrostatic pressure in the range of 0.8 to 1.4 MPa during the day (Gould et al., 2005). Hence, water potential for transport phloem in these plants is at the level of -1.0 MPa or lower. A similar value of water potential should be expected for barley collection phloem, and therefore it is most likely that its real turgor pressure values are significantly lower than those ordinarily estimated from the solute concentrations with the assumption of water potential equilibrium. In the same time, the water potential measured in bundle-sheath cells of barley leaves had a value close to -0.1 MPa and the turgor pressure was raising from 1.0 to 1.3 MPa during the photoperiod (Koroleva et al., 2002). Therefore it is very probable that the turgor pressure in the phloem of barley leaves is similar to the turgor pressure in the bundle- sheath cells and sometimes lower. In these circumstances, the water influx into the phloem should be strongly favored due to imbalance between osmotic and hydrostatic pressures in sieve elements.

Extrapolating this situation to putative symplastic loaders, we propose that the water potential gradient between bundle-sheath cells and phloem cells could well be the driving force for symplastic water flow from bundle-sheath cells into the phloem in putative symplastic loaders. We further propose that the hydrostatic pressure gradient between the bundle-sheath cells and the phloem can drive symplastic mass flow of solutes via plasmodesmata connecting bundle-sheath cells and the phloem companion cells in putative symplastic loaders. This flux can be directed into and out of the phloem depending on the turgor pressure gradient. The degree of the turgor pressure difference might be the mechanism regulating opening/ closure of the plasmodesmata. Oparka and Prior (1992) have demonstrated that the low hydrostatic pressure difference between the cells of tobacco (Nicotiana tabacum) trichomes (less than 0.2 MPa) kept plasmodesmata between these cells open, whereas the rise of the hydrostatic pressure gradient at this interface above 0.2 MPa closed the plasmodesmata (Oparka and Prior, 1992). We suggest that the low hydrostatic pressure difference between the phloem and the bundle-sheath cells could keep plasmodesmata at the interface of these tissues open and allow symplastic mass flow into the phloem, whereas the rise of the hydrostatic pressure gradient at this interface could close the plasmodesmata, preventing symplastic outflow of phloem solutes. The pressure-dependent regulation of plasmodesmata openings might be achieved by means of the dilatation and contraction of desmotubules (Gamalei et al., 1994). This mechanism could efficiently prevent the equilibration of sugar concentrations in phloem and mesophyll by diffusion.

The importance of apoplastic loading as the mechanism that creates high sugar concentrations in the phloem should be emphasized. Our experimental data have shown that in the light period, phloem concentrations of Sue in exporting leaves of A. meridionalis and of Sue and antirrhinoside in A. bardaiana are higher than those in mesophyll cytoplasm. Thus, the loading of these sugars occurs against the concentration gradient and ought to be energized. We propose that this is achieved by using energy from proton motive force. From the data available so far, the osmotic pressure in the phloem of A. meridionalis is likely to be built up by apoplastic phloem loading of Sue, and by synthesis of raffinose and stachyose in the ICs. In A. bardaiana, we propose that apoplastic loading of Sue and antirrhinoside occurs mostly via transfer companion cells, and it is mainly responsible for increasing the osmotic pressure in the phloem. The small amounts of raffinose and stachyose found in the phloem sap of this species might be synthesized in MICs. In both species, symplastic inflow of solutes via plasmodesmata probably dilutes only slightly the phloem sap, whereas apoplastic loading continues to operate. Symplastic entry of solutes into the phloem is thus likely to represent an additional route for supplying sugar and water to the phloem.

In conclusion, our data for two Scrophulariaceae species, A. meridionalis and A. bardaiana, present evidence that diffusion of sugars along their concentration gradient cannot be the main mechanism that determines symplastic exchange of carbohydrates between mesophyll and phloem during daily export of photoassimilates from the leaves. Questions remain as to (1) Why do some plants need plasmodesmal connections at the mesophyll/phloem interface and potentially symplastic movement of solutes into the phloem while others do not? (2) What mechanism underlies the positive correlation between the number of plasmodesmata at the mesophyll/phloem interface and the amount of RFOs in the phloem sap (Gamalei, 1984; Turgeon et al., 1993)? (3) What is the role of RFO synthesis in the phloem? Further studies should shed light on these issues.

MATERIALS AND METHODS

Plants

Alonsoa meridionalis O. Kuntze and Asarina bardaiana Pennell (Scrophulariaceae) were grown in a greenhouse on pot soil at 600 to 700 mol m^sup -2^ s^sup -1^ photon fluence rate, 14-h-light/10-h- dark period, and 22C/14C temperature period. We used plants at the same stage of development (grown for 4 weeks and used before the transition to flowering). Phloem sap was collected over several hours during the light period, and the other samples were taken within this period. For nonaqueous fractionation, plants were sampled at different times within the first 5 h of the light period, and the samples produced similar results. For the experiment shown in Figure 4, plants were grown on pot soil in a controlled- environment chamber (Sanyo Gallenkamp) at 20C in a 16-h-light/8-h- dark cycle and a photon flux of 500 mol m^sup -2^ s^sup -1^ and 0.035% CO2. All physiological studies, extractions of apoplastic sap, and sugar analyses in whole leaves and in single-cell samples were performed using mature, fully expanded leaves, usually the third leaf from the top of the branch.

Chloroform:Methanol Extraction

After shock freezing in liquid nitrogen, plant tissue was ground in a mortar and extracted on ice in a chloroforrrumethanol mixture (3:7, v/v). The homogenate was then extracted twice with water. The aqueous phases were combined and evaporated in a rotatory evaporator. The dried residue was dissolved in ultra-pure water (Millipore), syringe filtrated (0.45 m celluloseacetate; Schleicher and Schuell), and stored at -80C.

Preparative Isolation and Confirmation of the Chemical Structure of Antirrhinoside

100 g A. bardaiana leaves were ground to a powder in liquid nitrogen and extracted in 1.5 L of a chloroform:methanol mixture (3:7, v/v) on ice for 30 min. The homogenate was extracted twice with 500 mL distilled, deionized water. The volume of aqueous phase was reduced to 100 mL in a rotary evaporator at 34C. Canons were removed from the extract by adding a cation exchanger Dowex AG 50W- X8 (in H+ form) and Polyclar AT (Sigma) was used to remove polyphenols and polysaccharides. The volume of the extract was reduced to 10 mL in the rotary evaporator and the extract was applied to an anion-exchange column of 50-cm length and 2.8-cm diameter filled with Dowex 1 8 (OH- form). Passing the extract through the column allowed the removal of anions and at the same time led to a separation of the extract into fractions with different sugar composition. The elution was performed with 0.2 M NaOH at a rate of 2.5 mL/min. The sugar composition of the fractions was determined by HPLC. The antirrhinoside-containing fractions were combined and neutralized using 5 M HCl and the volume was reduced to 10 mL in the rotary evaporator at 34C. Antirrhinoside was purified by descendent paper chromatography. After elution from paper chromatograms with distilled, deionized water in an ultrasonication bath for 20 min and concentration in a rotary evaporator, 35 mL of approximately 30 mM antirrhinoside solution were obtained (i.e. approximately 350 mg antirrhinoside). From this solution, 5 mL (approximately 50 mg antirrhinoside dissolved in water) were used for the analysis of the chemical structure of antirrhinoside by NMR at the Institute for Inorganic Chemistry, the University of Gottingen, as described by Tietze et al. (1980).

Extraction of Apoplastic Sap

Apop\lastic fluid was obtained from leaves according to the method of Speer and Kaiser (1991) and Lohaus et al. (2001). Leaves were detached from the plants and infiltrated with ice-cold 50 mM CaCl^sub 2^ solution using a 60-mL syringe. The leaves were then carefully blotted dry, positioned into a 10-mL vessel located on top of a centrifuge tube and centrifuged for 5 min at 80g to 160g and 4C. Apoplastic sugar concentrations in the leaves were determined on the basis of the dilution factor F = (V^sub apoplast^ + V^sub gas Space^)/V^sub apoplast^. The volumes of the liquid apoplast ( V^sub apoplast^) have been determined as described in Knop et al. (2001). The absence of cellular contamination of the apoplastic extracts was proven by measurements of the activity of mala te dehydrogenase in the samples.

Nonaqueous Fractionation of Leaves

Leaves were cut from the plants after 5 h of the light period. The middle rib was removed, and the samples were ground to a fine powder in liquid nitrogen in a precooled mortar. The leaf tissue powder was lyophilized at -25C for 5 d. The nonaqueous fractionation was performed as described in Knop et al. (2001). For determination of metabolite concentrations in the gradient fractions, the dried sediments were extracted in chloroformmethanol mixture as described above and used for HPLC analyses.

Calculation of the Subcellular Distribution of Metabolites

For the evaluation of the subcellular distribution of sugars and amino acids between the stromal, cytoplasmic, and vacuolar compartments, a calculation procedure according to Riens et al. (1991) was used. The calculations were based on mean values obtained from measurements from five independent, density-gradient fractionations for each species. The protein concentrations were measured in gradient fractions from A. meridionalis and A. bardaiana leaves according to Lowry et al. (1951).The Chl:protein ratio was determined for each preparation of leaf tissue powder. Based on the known Chl:protein ratio, the amounts of metabolites were recalculated as micromoles per milligram of ChI, which allowed the comparison between several gradients from independent preparations of leaf tissue powder.

Estimation of the Partial Volumes of Compartments of Mesophyll Cells

The micrographs of mesophyll cells were obtained by conventional electron microscopy and the partial volumes of the chloroplast, vacuoles, and cytoplasm were determined by image-analysis technique (Bioscan Optimas). The measurements were carried out using 15 to 20 sections of the mesophyll tissue for each species.

Carbohydrate Analysis

For carbohydrate analysis by HPLC, an anion-exchange column (CarboPAClO; Dionex) was used for the determination of mono-, di-, and oligosaccharides and an MAl column from the same firm for the determination of polyols and cyclitols. Both columns were eluted with NaOH (Baker) using the LC-9A pump from Shimatzu. The CarboPACIO column was eluted isocratically with 80 mM NaOH with a flow rate of 1 mL min^sup -2^ and the MAl column with 600 mM NaOH with a flow rate of 0.4 mL min^sup -1^. Sugars were detected by a thin layer amperometric cell (ESA, model 5200) with a gold electrode and a pulse amperometric detector (Coulochem II). The evaluation of chromatograms was performed with the integration program Peaknet 5.1 (Dionex).

Determination of the Osmolality of the Leaf Sap

Discs were cut from leaves, placed in 1.5 mL Eppendorf tubes, and frozen at -20C, then thawed on ice and centrifuged to extract cell sap from the tissue. This sap was used for the determination of the osmolality using the osmometer Wescor 510OB.

Sampling of Single Cells

Single-cell sap was extracted from individual epidermal and mesophyll cells by the glass microcapillary technique (Tomos et al., 1994). Prior to use, a microcapillary was back filled with low- viscosity water-saturated paraffin oil (Sigma). Ejection of the single-cell sample under the oil allowed the determination of osmotic pressure by picolitre osmometer (Tomos et al., 1994) and sugars (GIc, Fru, and Sue) by an enzymatic assay (Koroleva et al., 1998). The measurements were highly reliable in the used concentration range between 2 and 200 mM.

Inhibition of Phloem Translocation

To interrupt phloem translocation, leaves were detached and the leaf petioles were kept in 2 mM CaCl2 solution, leading to the formation of callose and sealing of the phloem, thus preventing the exudation of sugars from leaves (King and Zeevaart, 1974). During this time, the leaves were kept under continuous light of 500 mol photons m^sup -2^ s^sup -1^.

Collection of Sieve Tube Sap

Sieve tube sap was obtained from severed stylets of the green- peach aphid, Myzus persicae, as described in Knop et al. (2001). About 10 aphids were caged for about 3 h on the mid portion of the leaf. Their stylets were cut using a laser beam. The exuding phloem sap was collected in sterile microcapillaries (total volume 0.5 L) and the volume of the exudates was determined by measuring the length occupied by the solution. Evaporation of the phloem sap was prevented by ensuring that the front edge of the capillary was in close contact with the leaf surface and the back end was surrounded by a plastic cap to minimize air circulation. The humidity around the capillary was about 80%. In this case evaporation from reference capillaries was not detectable. The samples were injected into 100 L of distilled sterile water and stored at -80C.

Statistical Treatment of the Data

The significance of difference between mesophyll and phloem concentrations of each sugar was analyzed using Student's t test (Zar, 1996).

ACKNOWLEDGMENTS

We are grateful to Lutz F. Tietze for the determination of the chemical structure of antirrhinoside, and to Ulrich Heber and Katharina Pawlowski for helpful discussions. We wish to thank the anonymous reviewer for the helpful comments on the manuscript.

Received July 15, 2005; revised October 31, 2005; accepted November 3, 2005; published December 23, 2005.

1 This work was supported by a grant of the Deutscher akademischer Austauschdienst (to O.V.V.), by the Russian Foundation for Basic Research (grant no. 04-04-48388 to D.R.B. and O.V.V.), and by the Deutsche Forschungsgemeinschaft (to G.L.).

LITERATURE CITED

Adhikari J, Bhaduri TJ, DasGupta S, Majumder AL (1987) Chloroplast as a locale of L-myo-inositol-1-phosphate synthase. Plant Physiol 85: 611-614

Ayre BG, Keller F, Turgeon R (2003) Symplastic continuity between companion cells and the translocation stream: Long-distance transport is controlled by retention and retrieval mechanisms in the phloem. Plant Physiol 131: 1518-1528

Bachmann M, Keller F (1995) Metabolism of the raffinose family oligosaccharides in leaves of Ajuga replans L.: intercellular and intracellular compartmentation. Plant Physiol 109: 991-998

Beebe DU, Turgeon R (1992) Localization of galactinol, raffinose, and stachyose synthesis in Ciicurbita pepo leaves. Planta 188: 354- 361

Chandran D, Reinders A, Ward JM (2003) Substrate specificity of the Arabidopsis thaliana sucrose transporter AtSUC2. ] Biol Chem 278: 4432044325

Christmann J, Kreis W, Reinhard F, (1993) Uptake, transport and storage of cardenolides in foxglove: Cardenolide sinks and occurrence of cardenolides in the sieve tubes of Digitalis lanata. Bot Acta 106: 419-427

Deeken R, Geiger D, Fromm J, Koroleva O, Ache P, Langenfeld- Heyser R, Sauer N, May ST, Hedrich R (2002) Loss of the AKT2/3 potassium channel affects sugar loading into the phloem of Arabidopsis. Planta 216: 334-344

Flora LL, Madore MA (1996) Significance of minor-vein anatomy to carbohydrate transport. Planta 198: 171-178

Gamalei YuV (1984) The structure of leaf minor veins and the types of translocated carbohydrates. Dokl Akad Nauk 277: 1513-1516

Gamalei YuV (1990) Leaf Phloem. Nauka, Leningrad

Gamalei YuV (1991) Phloem loading and its development related to plant evolution from trees to herbs. Trees (Berl) 5: 50-64

Gamalei YuV (2000) Comparative anatomy and physiology of leaf minor veins and bundle sheath parenchyma in leaves of dicotyledonous plants. Botan Zhurnal 85: 34-49

Gamalei YuV, van Bel AJE, Pakhomova MV, Sjutkina AV (1994) Effects of temperature on the conformation of the endoplasmic reticulum and on starch accumulation in leaves with the symplastic minor vein configuration. Planta 494: 443-453

Gerhardt R, Heldt HW (1984) Measurement of subcellular metabolite levels by fractionation of freeze-stopped material in nonaqueous media. Plant Physiol 75: 542-547

Gould N, Thorpe MR, Koroleva O, Minchin PEH (2005) Phloem hydrostatic pressure relates to solute loading rate: a direct test of the Munch hypothesis. Functional Plant Biology 32: 1019-1026

Gowan E, Lewis BA, Turgeon R (1995) Phloem transport of antirrhinoside, an iridoid glycoside, in Asarina scandens. J Chem Ecol 21: 1781-1788

Haritatos E, Keller F, Turgeon R (1996) Raffinose oligosaccharide concentrations measured in individual cell and tissue types in Cucumis melo L. leaves: implications for phloem loading. Planta 198: 614-622

Heber U (1957) Zur frage der lokalisation von loslichen zuckern in der pflanzenzelle. Ber Dtsch Bot Ces 70: 371-382

Heineke D, Wildenberger K, Sonnewald U, Willmitzer L, Heldt HW (1994) Accumulation of hexoses in leaf vacuoles: studies with transgenic tobacco plants expressing yeast-derived invertase in the cytosol, vacuole or apoplasm. Planta 194: 29-33

King RW, Zeevaart JAD (1974) Enhancement of phloem exudation from cut petioles by chelating agents. Plant Physiol 53: 96-103

Knop C (2001) Zur bedeutung von saccharose-transportern in pflanzen mit offener phloemanatomie. PhD thesis. Universitt Gottingen, Gottingen, Germany

Knop C, Stadler R, Sauer N, Lohaus G (2004) AmSVTI, a sucrose transporter in collection and transport phloem of the putative symplastic phloem loader Alonsoa meridionalis. Plant Physiol 134: 204-214

Knop C, Voitsekhovskaja O, Lohaus G (2001) Sucrose transporters in \two members of the Scrophulariaceae with different types of transport sugar. Planta 213: 80-91

Koroleva OA, Farrar JF, Tomos AD, Pollock CJ (1998) Carbohydrates in individual cells of epidermis, mesophyll, and bundle sheath in barley leaves with changed export or photosynthetic rate. Plant Physiol 118: 1525-1532

Koroleva OA, Tomos AD, Farrar JF, Pollock CJ (2002) Changes in osmotic and turgor pressure in response to sugar accumulation in barley source leaves. Planta 215: 210-219

Leidreiter K, Kruse A, Heineke D, Robinson DG, Heldt HW (1995) Subcellular volumes and metabolite concentrations in potato (Solatium tuberosum cv. Dsire) leaves. Bot Acta 108: 439-444

Lohaus G, Buker M, Humann M, Soave C, Heldt HW (1998) Transport of amino acids with special emphasis on the synthesis and transport of asparagine in the Illinois Low Protein and Illinois High Protein strains of maize. Planta 205: 181-188

Lohaus G, Hussmann M, Pennewiss K, Schneider H, Zhu JJ, Sattelmacher B (2000) Solute balance of a maize (Zea mays L.) source leaf as affected by salt treatment with special emphasis on phloem re-translocation and ion leaching. J Exp Bot 51: 1-12

Lohaus G, Pennewiss K, Sattelmacher B, Hussmann M, Muhling KH (2001) Is the infiltration-centrifugation technique appropriate for the isolation of apoplastic fluid? A critical evaluation with different plant species. Physiol Plant 111: 457-465

Lohaus G, Winter H, Riens B, Heldt HW (1995) Further studies of the phloem loading process in leaves of barley and spinach: the comparison of metabolite concentrations in the apoplastic compartment with those in the cytosolic compartment and in the sieve tubes. Bot Acta 108:270-275

Lowry OH, Rosebrough NJ, Farr AL, Randall RJ (1951) Protein measurement with the Folin phenol reagent. J Biol Chem 193: 265-275

Moing A, Carbonne F, Zipperlin B, Svanella L, Gaudillere J-P (1997) Phloem loading in peach: symplastic or apoplastic? Physiol Plant 101: 489-496

Moore BD, Palmquist DE, seemann JR (1997) Influence of plant growth at high CO2 concentrations on leaf content of ribulose-l,5- biphosphate carboxylase/oxygenase and intracellular distribution of soluble carbohydrates in tobacco, snapdragon, and parsley. Plant Physiol 115:241-248

Munch E (1930) Material Flow in Plants (translated 2003; JA Milburn, KH Kreeb, University of Bremen, Bremen, Germany). Gustav Fischer Verlag, Jena, Germany

Oparka KJ, Prior DAM (1992) Direct evidence for pressure- generated closure of plasmodesmata. Plant J 2: 741-750

Pate JS, Gunning BES (1969) Vascular transfer cells in angiosperm leaves. A taxonomic and morphological survey. Protoplasma 68: 135- 156

Riens B, Lohaus G, Heineke D, Heldt HW (1991) Amino acid and sucrose content determined in the cytosolic, chloroplastic, and vacuolar compartments and in the phloem sap of spinach leaves. Plant Physiol 97: 227-233

Riesmeier JW, Willmitzer L, Frommer WB (1994) Evidence for an essential roie of the sucrose transporter in phloem loading and assimilate partitioning. EMBO J 13: 1-7

Speer M, Kaiser WM (1991) Ion relations of symplastic and apoplastic space in leaves from Spinacea oleracea L. and Pisum sativum L. under salinity. Plant Physiol 97: 990-997

Sprertger N, Keller F (2000) Allocation of raffinose family oligosaccharides to transport and storage pools in Ajuga reptans: the roles of two distinct galactinol syntheses. Plant J 21: 249-258

Strau M, Kauder F, Peisker M, Sonnewald U, Conrad U, Heineke D (2001) Expression of an abscisic acid-binding single chain antibody influences the subcellular distribution of abscisic acid and leads to developmental changes in transgenic potato plants. Planta 213: 361-369

Tietze LF, Niemeyer U, Marx P, Glusenkamp KH (1980) Determination of the relative configuration and conformation of isomeric iridoid glycosides by proton and carbon-13 NMR and mass spectroscopy. Tetrahedron 36: 1231-1236

Tomos AD, Hinde P, Richardson P, Pritchard J, Fricke W (1994) Microsampling and measurement of solutes in single cells. In N Harris, KJ Oparka, eds, Plant Cell Biology-A Practical Approach. IRL Press, Oxford, pp 297-314

Turgeon R (1991) Symplastic phloem loading and the sink-source transition in leaves: a model. In Bonnemain JL, Delrot S, Lucas W, Dainty J, eds, Recent Advances in Phloem Transport and Assimilate Compartmentation. Ouest Editions, Nantes, France, pp 18-22

Tuigeon R (1996) Phloem loading and plasmodesmata. Trends Plant Sci 1: 418-423

Turgeon R, Beebe DU, Gowan E (1993) The intermediary cell: minor- vein anatomy and raffinose oligosaccharide synthesis in the Scrophulariaceae. Planta 191: 446-456

Turgeon R, Medville R (2004) Phloem loading: a revaluation of the relationship between plasmodesmatal frequencies and loading strategies. Plant Physiol 136: 3795-3803

Turgeon R, Webb JA (1975) Leaf development and phloem transport in Cucurbita pepo: carbon economy. Planta 123: 53-62

Tyree MT (1970) The symplast concept: a general theory of symplastic transport according to the thermodynamics of irreversible process. J Theor Biol 26: 181-189

van Bel AJE, Gamalei YuV (1991) Multiprogrammed phloem loading. In Bonnemain JL, Delrot S1 Lucas W, Dainty J, eds, Recent Advances in Phloem Transport and Assimilate Compartmentation. Ouest Editions, Nantes, France, pp 128-139

Winter H, Robinson DG, Heldt HW (1993) Subcellular volumes and metabolite concentrations in barley leaves. Planta 191: 180-190

Winter H, Robinson DG, Heldt HW (1994) Subcellular volumes and metabolite concentrations in spinach leaves. Planta 193: 530-535

Zar JH (1996) Biostatistical Analysis. Prentice-Hall, Upper Saddle River, NJ, pp 353-356

Olga V. Voitsekhovskaja*, Olga A. Koroleva2, Denis R. Batashev, Christian Knop, A. Deri Tomos, Yuri V. Gamalei, Hans-Walter Heldt, and Gertrud Lohaus

Albrecht-von-Haller-Institute for Plant

Source: Plant Physiology

More News in this Category