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First published online December 23, 2005; 10.1104/pp.105.068312 Plant Physiology 140:383-395 (2006) © 2006 American Society of Plant Biologists Phloem Loading in Two Scrophulariaceae Species. What Can Drive Symplastic Flow via Plasmodesmata?1Albrecht-von-Haller-Institute for Plant Sciences, Plant Biochemistry, 37077 Goettingen, Germany (O.V.V., C.K., H.-W.H., G.L.); Komarov Botanical Institute, Russian Academy of Sciences, Plant Ecological Physiology, 197376 St. Petersburg, Russia (O.V.V., D.R.B., Y.V.G.); and School of Biological Sciences, University of Wales, Bangor, Gwynedd, Wales LL57 2UW, United Kingdom (O.A.K., A.D.T.)
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 bundle-sheath cells (40–60 plasmodesmata per µm2 cell surface; Gamalei, 1991) and transfer cells (TCs) that have very few plasmodesmata (less than 0.01 plasmodesma per µm2 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 Suc 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 Suc 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 Suc 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).
In compliance with the polymer-trap model, Suc 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 phloem-loading 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
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 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.
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)
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).
HPLC analysis of the leaf extracts of A. barclaiana and A. meridionalis has shown that Glc, Fru, Suc, 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
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 nonaqueous 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 (Strauß et al., 2001
In A. barclaiana, antirrhinoside was mostly present in the vacuoles but chloroplasts and cytoplasm also contained significant proportions. Suc 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).
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)
To confirm the accuracy of the calculated volumes of the mesophyll, the concentrations of Glc, Fru, and Suc 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 Suc (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.
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 µL (mg Chl)–1 for A. barclaiana and 59, 24, and 213 µL (mg Chl)–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
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, myo-inositol dominated, followed by Suc. 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, Suc 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 Glc and Suc in much lower concentrations. The dominating carbohydrate in the cytoplasm was also antirrhinoside, followed by mannitol, Suc, and myo-inositol.
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–1 for A. barclaiana and 365 mOsmol kg–1 for A. meridionalis. In A. barclaiana, the single-cell osmolalities were 629 mOsmol kg–1 for the epidermis and 682 mOsmol kg–1 for the mesophyll. In A. meridionalis, the osmolalities were 491 mOsmol kg–1 and 487 mOsmol kg–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–1 for A. meridionalis and 2,260 mOsmol kg–1 for A. barclaiana. 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.
Antirrhinoside (8 mM) and mannitol (4 mM) dominated the apoplast of A. barclaiana leaves, while Suc, Glc, 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 mM and that of Suc 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 Glc and Fru, respectively, and Suc accumulated in the apoplast to a concentration of only 1 mM (Fig. 5).
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.
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. barclaiana 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, Suc 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.
Galactinol had been found previously in the mesophyll vacuoles of the apoplastic phloem loader Antirrhinum majus (Moore et al., 1997 The subcellular distribution of the iridoid glucoside antirrhinoside present in A. barclaiana 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.
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
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
A. meridionalis is the only species with ICs in minor-vein phloem that has been studied using the nonaqueous fractionation technique with respect to the ratio of sugar concentrations in the phloem sap and in the cytoplasm of mesophyll cells. Based on its minor-vein anatomy and the predictions of the polymer-trap model, Suc is expected to enter the phloem in this species along its concentration gradient via plasmodesmata. However, in A. meridionalis, the Suc 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 Suc 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 Suc by simple diffusion under some conditions cannot be completely ruled out.
Apoplastic levels of Suc 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 Suc from the apoplast into the phloem has been strongly suggested for A. meridionalis by Knop et al. (2004)
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 Suc 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 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 mM, 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.
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 Suc 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 Suc, if this were to occur in two species under consideration. From a thermodynamic point of view, the difference between Suc 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 Suc than the cytoplasm of mesophyll cells. Regardless of the way the Suc was loaded into the phloem, this would prevent any further diffusional movement of Suc 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 Suc 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 Suc 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)
Two major mechanisms have been proposed to account for the symplastic movement of solutes via plasmodesmata: the mass flow of solution, and the diffusion of solutes (Tyree, 1970
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 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 Suc in exporting leaves of A. meridionalis and of Suc and antirrhinoside in A. barclaiana 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 Suc, and by synthesis of raffinose and stachyose in the ICs. In A. barclaiana, we propose that apoplastic loading of Suc 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. barclaiana, 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
Plants Alonsoa meridionalis O. Kuntze and Asarina barclaiana Pennell (Scrophulariaceae) were grown in a greenhouse on pot soil at 600 to 700 µmol m–2 s–1 photon fluence rate, 14-h-light/10-h-dark period, and 22°C/14°C 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 20°C in a 16-h-light/8-h-dark cycle and a photon flux of 500 µmol m–2 s–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.
After shock freezing in liquid nitrogen, plant tissue was ground in a mortar and extracted on ice in a chloroform:methanol 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 µm cellulose-acetate; Schleicher and Schuell), and stored at –80°C.
100 g A. barclaiana 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 34°C. Cations 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 x 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 34°C. 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 Göttingen, as described by Tietze et al. (1980)
Apoplastic fluid was obtained from leaves according to the method of Speer and Kaiser (1991)
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 –25°C for 5 d. The nonaqueous fractionation was performed as described in Knop et al. (2001)
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)
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.
For carbohydrate analysis by HPLC, an anion-exchange column (CarboPAC10; Dionex) was used for the determination of mono-, di-, and oligosaccharides and an MA1 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 CarboPAC10 column was eluted isocratically with 80 mM NaOH with a flow rate of 1 mL min–1 and the MA1 column with 600 mM NaOH with a flow rate of 0.4 mL min–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).
Discs were cut from leaves, placed in 1.5 mL Eppendorf tubes, and frozen at –20°C, 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 5100B.
Single-cell sap was extracted from individual epidermal and mesophyll cells by the glass microcapillary technique (Tomos et al., 1994
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
Sieve tube sap was obtained from severed stylets of the green-peach aphid, Myzus persicae, as described in Knop et al. (2001)
The significance of difference between mesophyll and phloem concentrations of each sugar was analyzed using Student's t test (Zar, 1996
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; returned for revision October 31, 2005; accepted November 3, 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.). 
2 Present address: John Innes Centre, Colney Lane, Norwich NR4 7UH, UK. The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Olga V. Voitsekhovskaja (ovoitse{at}yandex.ru). Article, publication date, and citation information can be found at www.plantphysiol.org/cgi/doi/10.1104/pp.105.068312. * Corresponding author; e-mail ovoitse{at}yandex.ru; fax 7–812–2344512.
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