[21] Phloem transport

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Transport

B y GABRIELE ORLICH a n d EWALD K O M O R

The term phloem transport refers to the flow of assimilates from their site of synthesis or storage to their site of consumption, a process which, for methodological reasons, may be separated into three phases: phloem loading, long-distance transport, and phloem unloading. To gain a better understanding of phloem transport, a close relationship between structural and functional information is necessary. Therefore, methods for studying phloem transport should include anatomical, histological, and physiological approaches. In this chapter a selection of techniques is presented with special emphasis on their methodological value, and open questions are briefly outlined on occasion. The different techniques are introduced by those who are working in the respective field to provide firsthand information. P h l o e m Loading General Comments The term phloem loading as used in this chapter denotes the transfer of assimilates from the sites of CO: fixation (or the sites of mobilization of storage carbohydrates in stems or roots) to the phloem region via symplastic and/or apoplastic routes as well as the process of sucrose uptake into the sieve tube-companion cell complex as the final, most important step in this sequence, the mechanism of which may be termed either symplastic, if accomplished by plasmodesmatal connections between adjacent cells, or apoplastic, if mediated by a plasma membrane-bound carder system. Symplastic versus apoplastic pathways through the tissue and symplastic versus apoplastic uptake of sucrose into the sieve tube-companion cell complex depend on species-specific properties of tissues, such as the arrangement of different cell types in mature source leaves, their enzyme and carrier composition, including cell wall-bound invertase (p-fruetofuranosidase), and the frequency of plasmodesmata between different cell types. There is a common consensus that sucrose is translocated within the sieve tubes. The mechanism of sucrose uptake into the sieve tubes, however, has been debated since the 1960s 1,2 for reasons which, as we see it, R. T. Giaquinta, Annu. Rev. Plant Physiol. 34, 347 (1983). 2 A. L. Kursanov, "Assimilate Transport in Plants," p. 81. Elsevier, Amsterdam, 1984.

METHODS IN ENZYMOLOGY, VOL. 174

Copyright© 1989by AcademicPress,Inc. All fightsof reproductionin any formreserved.

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must be ascribed to the experimental problem of either quantifying or eliminating the contribution of nonphloem cells to the available data on sucrose movement through leaf tissues. So, quite consistently, more recent conceptual assessments3 (see also earlier discussions4,5) again concentrate on the fate of sucrose along its way to the phloem, in an attempt to elucidate its compartmentation at the cell type level and thereby trying to determine a potential regulatory role of assimilate transfer through the tissue in sucrose uptake at the sieve tube-companion cell complex. The choice of the ideal object with which to investigate phloem loading, as well as the kind of pretreatment of a selected plant material, should depend on the question under study; thus, to trace the pathway(s) of assimilates (sucrose) from their site of synthesis or application to the site of sucrose uptake, strict methodology would dictate the use of intact, nonwounded tissue, whereas isolated, enriched or purified cells, protoplasts or membranes of a distinct cell type should be used to characterize mechanisms of carder-mediated sugar uptake. Isolating viable phloem cells remains a technical problem; on the other hand, and more importantly, many of the efforts to analyze compartmentational and mechanistic aspects of phloem loading suffer from methodological restrictions and therefore often do not allow clear interpretation. In these introductory remarks we have tried to connect the pertinent questions with the available techniques and thus elucidate their methodological value. Uptake of [l~C]sucrose The preferential labeling of veins in autoradiographs after application of [~4C]sucrose to a pretreated mature green leaf has been taken as evidence that phloem loading has occurred; therefore p4C]sucrose uptake experiments have become a widely used technique to study compartmentational, kinetic, and energetic aspects of phloem loading. Interpretation of results from [~4C]sucrose uptake experiments with leaf tissue is often impeded for the following reasons: (1) Sucrose applied via the apoplast may not follow a route in the tissue that is identical to the in vivo pathway after CO2 fixation5 (see also Giaquinta et al. 6 for the pathway of unloading). (2) The leaf tissue has to be made accessible for [~4C]sucrose either by abrading the leaf surface or by cutting small leaf disks. These pretreatments, especially in 3 W. J. Lucas, in "Regulation of Carbon Partitioning in Photosynthetic Tissue" (R. L. Heath and J. Preiss, eds.), p. 254. American Society of Plant Physiologists, Rockville, Maryland, 1985. 4 B. R. Fondy and D. R. Geiger, Plant Physiol. 59, 953 (1977). 5 M. Madore and J. A. Webb, Can. J. Bot. 59, 2550 (1981). 6 R. T. Giaquinta, W. Lin, N. L. Sadler, and V. R. Franceschi, Plant Physiol. 72, 362 (1983).

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combination with prolonged incubation times to reduce internal sugar levels, may cause wound responses 7 such as ethylene-induced inactivation and formation of (new) carder systemss or synthesis of (additional) invertase 9,1° thereby changing the relative contributions of sugar pathways to phloem supply in the tissue.11 For maize leaves, Heyser et aL 12 have developed a technique for supplying sucrose from the apoplast that avoids severe wounding by introducing sucrose into the xylem via the transpiration stream. The cotyledons of castor bean have the advantage of being accessible to externally supplied sucrose without abrasion of the epidermis since they take up sugars from the apoplast (endosperm) during germination. 13 However, because of the different physiological (heterotrophic) status in terms of simultaneous source and sink characteristics, the results obtained with the cotyledon may not adequately reflect compartmentational properties of phloem loading in a green source leaf. (3) Sucrose is not exclusively taken up by phloem cells. Although autoradiographs show that label is preferentially accumulated in the veins of leaf tissue, Fondy and Geiger,4 in an analysis of labeled products following [14C]sucrose uptake by sugar beet leaves and quantitative autoradiography, found that label was also present in the mesophyll cells, part of which had been identified as sucrose. This indicates that sucrose is not exclusively taken up by phloem ceils. Alternatively, either mesophyll cells can take up sucrose as well or part of the sucrose is hydrolyzed by a cell wall-bound invertase and hexoses are taken up from which sucrose is resynthesized: in both cases [14C]sucrose uptake does not exclusively reflect "phloem loading" (in the traditional sense of uptake into the sieve tube-companion cell complex). Therefore, interpretation of kinetic data from sucrose uptake experiments with whole leaf tissue to characterize the presumed sucrose carrier in the sieve tube-companion cell complex is ambiguous. Preferential labeling of veins in autoradiographs after [~4C]sucrose application may indicate preferential sucrose uptake into the sieve tubecompanion cell complex from the apoplast but would also be consistent with the interpretation that sucrose is taken up by mesophyll cells exclusively and subsequently loaded symplastically into the sieve tube-

7 R. F. M. van Steveninck, Annu. Rev. Plant Physiol. 26, 237 (1975). s G. Abraham and L. Reinhold, Planta 150, 380 (1980). 9 M. Turkina and S. Sokolova, Soy. Plant Physiol (Engl. Transl.) 15, 1 (1968). ~oC. J. Pollock and E. J. Lloyd, Z. Pflanzenphysiol. 90, 79 (1978). tl R. Lemoine, S. Delrot, and E. Auger, PhysioL Plant. 61, 571 (1984). ~2 W. Heyser, O. Leonhard, R. Heyser, E. Fritz, and W. Esehrich, Planta 122, 143 (1975). 13 p. Kriedemann and H. Beevers, Plant Physiol. 42, 161 (1967).

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companion cell complex. If the sugar carrier composition of the different cell types are known, this would give a clear-cut answer in case no sucrose carrier is found in the plasmalemma of the sieve tube-companion cell complex: in this case a completely symplastic loading mechanism must be operative (see discussion in van Bel et al.~4). However, the presence of a sucrose carrier in the plasmalemma of the sieve tube-companion cell complex cannot be taken as proof for an exclusively apoplastic final uptake step since the carrier may function as a retrieval mechanism for sucrose leaked out into the apoplast, t5 while the bulk of assimilates is supplied symplastically. Although there have been some attempts to isolate tissue enriched in phloem either by mechanical separation of veins from leaves ~6 and petioles ~s,~7 or by enzymatic digestion of cell walls of leaves with subsequent separation of parenchyma and vein-type cells, ta'19 all procedures still yield a mixture of different cell types so that the requirements for an unequivocal localization of a sucrose carrier in the plasmalemma of the sieve tube-companion cell complex are not yet met. As long as cell cultures containing pure phloem cells are not available, the only advantage of a differentiated cell culture as compared to leaf tissue may be that phloem cells can be more readily separated and isolated by using a phloem-enriched (maximum 10%) cell clump as the starting material. 2° It must also be taken into consideration that the composition of carriers and enzymes in a hormone-treated cell culture may not be identical to the protein pattern of the same cell type after differentiation in the intact plant tissue. Isolation of mesophyll cells and comparison of their uptake kinetics with those of leaf disks is another approach to reveal the uptake characteristics of phloem cells and the involvement of mesophyll cells during phloem loading (van Bel et al.~4; see also Ref. 21 for comparison of parenchymatous suspension cells and cotyledons of castor bean). If isolated cells or protoplasts of a single cell type are used to study uptake activities, it should be kept in mind that enzymatic digestion of cell walls may damage transport sites and modify membrane energetic parameters, or lead to loss of potential binding proteins. Therefore properties of iso-

~4A. J. E. van Bel, A. Ammedaan, and G. Blaauw-Jansen, J. Exp. Bot. 37, 1899 (1986). ~5j. W. Maynard and W. J. Lucas, Plant PhysioL 70, 1436 (1982). 16 M. J. Brovchenko, Soy. Plant Physiol. (Engl. Transl.) 12, 230 (1965). ~7j. Daie, J. Am. Soc. Hortic. Sci. 111, 216 (1986). ~8C. Wilson, J. W. Oross, and W. J. Lucas, Planta 164, 227 (1985). ~9A. J. E. van Bel and A. J. Koops, Planta 164, 362 (1985). 20 R. D. Sjrlund and C. Y. Shih, J. Ultrastruct. Res. 82, 111 (1983). 2~ B.-H. Cho and E. Komor, J. Plant Physiol. 118, 381 (1985).

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lated cells and protoplasts should be compared with those of intact tissue and isolated cells, respectively. 14,22-24 To characterize the sugar carrier composition of a distinct cell type it has to be established whether the p4C]sucrose applied is taken up unhydrolyzed or whether (part of) it is split to hexoses by a cell wall-bound invertase. As long as there is no specific inhibitor for a cell wall-bound invertase available, the uptake rates of differently labeled sucrose (glucose versus fructose versus uniformly labeled sucrose) can be compared in order to distinguish between sucrose and hexose uptake. If the sucrose is taken up without previous hydrolysis, the uptake rates of the three differently labeled sucrose molecules should be identical, whereas if the sucrose is hydrolyzed prior to uptake the rates will be different. A prerequisite for this type of experiment is, in case the cells can take up hexoses, too, that the uptake kinetics for glucose and fructose be different. Another approach to reveal whether sucrose is hydrolyzed during [14C]sucrose u p t a k e is t o supply asymmetrically labeled sucrose (labeled either in the glucose or fructose moiety) and subsequently analyze the distribution of the label in the sucrose recovered. Sucrose which is not hydrolyzed will remain asymmetrically labeled, whereas sucrose which is resynthesized from the products of [~4C]sucrose hydrolysis will be labeled more or less randomly [in case the activity of the cytoplasmic glucose-6phosphate isomerase (phosphoglucoisomerase) is not rate-limiting]. However, if the asymmetry of the label is retained, it still cannot be concluded that no sucrose hydrolysis has occurred unless the sucrose recovered is approximately 100% of the incorporated label. For instance, if hexoses derived from sucrose hydrolysis are not entirely resynthesized to sucrose, this proportion of the label would be neglected, although it might contribute rather substantially to total sucrose uptake. (Giaquinta 25 showed that for sugar beet leaves the sucrose recovered retained asymmetry of the label to 95% but only amounted to about 60% of the total sucrose taken up.) By comparing uptake characteristics of sucrose and hexoses in isolated mesophyll cells and their protoplasts the involvement of cell wall-bound invertase during sucrose uptake will also show up (see Refs. 23 and 24 for this kind of experiment with heterotrophic tissues). Recently the sucrose analog fluorosucrose has been introduced as a new tool to study sucrose uptake, without interference of invertase activity26; 22 R. J. Henry, A. Schibeci, and B. A. Stone, Aust. J. Plant Physiol. 11, 119 (1984). 23 W. Lin, M. R. Schmitt, W. D. Hitz, and R. T. Giaquinta, Plant Physiol. 75, 936 (1984). 24 M. Stanzel, R. D. Sjolund, and E. Komor, Planta 174, 201 (1988). 25 R. Giaquinta, Plant Physiol. 59, 750 (1977). 26 W. D. Hitz, M. R. Schmitt, P. J. Card, and R. T. Giaquinta, Plant Physiol. 77, 291 (1985).

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fluorosucrose is not hydrolyzed by invertase but is taken up by sucrose carrier systems. This compound will be very useful for further studies investigating the sugar carrier composition of different cell types, also in combination with microautoradiography.

Techniques of Loading Sucrose into Leaves* A method for labeling and autoradiography of leaves is given that can be adapted with variation for different plant species. For sugar beet leaves, 27 several different methods of phloem-loading measurements are possible: reverse flap feeding, abraded epidermis feeding, or uptake by leaf disks. Only the latter two are described here. A sugar beet leaf (usually kept dark overnight) is abraded on the upper surface over an area of 10 cm 2 by rubbing gently with 300-mesh ceroxide paste and rinsed with water, or the lower epidermis is removed by cracking the upper epidermis and leaf by bending the leaf past a 90 ° angle. The leaf piece is used as a tab to peel the lower epidermis from the other piece. With practice, areas several centimeters in length can be peeled in one motion. The leaf portion is then sealed with cord-type caulking compound into a Plexiglas incubation chamber, so that the lower surface of the leaf is aerated by an air stream whereas the upper surface is supplied with a solution of radioactive sucrose in 5 m M potassium phosphate, pH 6.5. The sucrose concentration, the specific radioactivity, and the duration of incubation are varied according to the experimental aims. A time course of approximately 30 min is appropriate for sugar beet. In sugar beet the uptake rates are of the order of 0.5/tmol sucrose min -~ dm -2 at 100 m M sucrose applied. After the radioactive incubation, the surface of the leaf is usually rinsed carefully with water or with the same solution without the radioactive sugar. However, the time needed for wash of free space must be determined by following the time course of exit of labeled material. In sugar beet, 10 min suffices. Another method is incubation of leaf disks, e.g., of I cm 2, cut from the interveinal areas with a razor blade. The disks, after a short rinsing in buffer, are floated in 5 m M potassium-phosphate, pH 6.5, and labeled sucrose (ranging from l0 to 400 mM), e.g., 50 btl of labeled solution, on which one disk is allowed to float for 30 min. The experiment is terminated by removing the disk, rinsing it with water, and digesting it with H202 and perchloric acid or extracting it with boiling ethanol, etc., and the

* Section prepared by B. R. Fondy, D. R. Geiger, and S. Delrot. 27 S. A. Sovonick, D. R. Geiger, and R. J. Fellows, Plant Physiol. 54, 886 (1974).

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radioactivity in the samples is determined. Analysis for either method can be by autoradiography or by chemical means. Chemical Analysis of Metabolic Products of Sucrose Feeding. The tissue is quickly immersed in a small test tube with 20 ml of chloroformmethanol (1:4, v/v) at 65 ° for 10 min (refluxed in the tube with a stainless steel bolt to condense the solvent). The extract is collected, and the extraction is repeated with 80% methanol. The extracts are combined (rinse the tubes with chloroform) and may be used for thin-layer chromatography (TLC), enzymatic assay, or ion-exchange separation. For ion-exchange separation, chloroform is added to the extract to give two phases (after overnight in the refrigerator and centrifugation at 1000 g for 10 min). The chloroform layer is discarded, and the extract is dried on an evaporating block at 60 ° with a stream of dry nitrogen. The sample is then dissolved in 5 ml of 500/0 ethanol in water and subjected to ion-exchange Sepharose chromatography. The extract can be chromatographed on cellulose TLC plates (prepare 2 cm wide lanes with a razor blade and spot the extract, maximally 0.5 cm in diameter). Up to 80/zg of total sugar can be applied to 500-/zm plates. Lipids in the sample are moved to the solvent front by placing the plates in a tray of chloroform and allowing the front to advance 2 - 3 cm. This procedure is repeated twice, with evaporation of the solvent between runs. The plates are finally developed in ethyl acetate-acetic acid-formic acid-water (18:3:1:4) (for separation of mannitol, sucrose, glucose, and fructose) or in butanol-acetic acid-water (3:3: 1) (for separation of stachyose, raffinose, sucrose, glucose, and fructose). The separation is improved by repeating the development 2 to 3 times in these solvent systems. Labeled sugars are visualized with a fast-reacting X-ray film (e.g., Kodak Type AA) with a cumulative radiation of 4 × 10 6 counts/cm 2. The labeled spots are quantified by stripping off the labeled thin layer [apply a stripping mixture (7 g cellulose acetate, 3 g diethylene glycol, 2 g camphor, 25 ml n-propanol, 75 ml acetone) and the spots will curl after the mixture dries]. The bands can be mixed with the scintillation fluid and the radioactivity determined. Enzymatic analysis of the sugar in the extract can be performed by common methods, starch analysis by the method of Outlaw and Manchester. 2s

Microautoradiography A technique for tracing the route of assimilates from mesophyll to phloem tissue at the cellular level is microautoradiography after feeding a 2s

W. H. Outlaw and J. Manchester, Plant Physiol. 64, 79 (1979).

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leaf with 14CO2. In a pulse-chase experiment the cells involved in the pathway will show up by successive appearance of silver grains with increasing duration of the chase (see also Outlaw and Fisher~9 and Outlaw et al. 3° for details of this approach). A cell type actively accumulating assimilates may be identified by the level of grain density, if certain considerations are noted. (l) A transient increase in grain density is expected in each cell type involved in the pathway during a pulse-chase experiment; therefore, the relative proportions of grain densities in the different cell types have to be analyzed over a time-course experiment. (2) The highest relative grain density always means that active assimilate uptake has occurred, but maximal relative grain density cannot be expected in each cell type actively accumulating labeled assimilates, because an active uptake site is masked if the rate-limiting step in the pathway is localized before an active uptake site, thus always generating a lower level of grain density in the following steps and cells, respectively. In this respect, experimental conditions to provide slow fluxes of label (extensive predarkening 3~ or low-light intensity during 14CO2uptake 32) would probably prevent rate-limiting steps from being detected (or rather avoid masking of actively accumulating cell types) from the outset, because fluxes higher than those determined by a rate-limiting step might thus be suppressed. (3) The rate-limiting step in a sequence usually is a target for regulation; therefore, varying the photosynthetic rate or translocation rate may change the pattern of grain density distribution in a way that could help elucidate this interesting aspect of phloem loading. Although the cell types involved in the pathway of assimilates to the phloem can be identified by microautoradiography, the method does not reveal whether the assimilates are transported via the symplast or the apoplast. Geiger et al. a3 have tried to discriminate these routes by inhibition of fluxes of label through plasmodesmata using plasmolyzed leaf cells, but since disruption of cytoplasmic connections was not complete and photosynthetic activity was also impaired, this approach did not yield clear-cut results. If application of [~4C]sucrose is used in microautoradiographic studies, the restrictions of interpretation as outlined in the section o n [14C]sucrose uptake should be noted. Comparing the labeling pattern after t4CO2 feeding and [14C]sucrose uptake, Fritz et al. 3t have produced results which 29 W. H. Outlaw, Jr., and D. B. Fisher, Plant Physiol. 55, 699 (1975). 30 W. H. Outlaw, Jr., D. B. Fisher, and A. L. Christy, Plant Physiol. 55, 704 (1975). 31 E. Fritz, R. F. Evert, and W. Heyser, Planta 159, 193 (1983). 32 X.-D. Wang and M. J. Canny, Plant, CellEnviron. 8, 669 (1985). 33 D. R. Geiger, S. A. Sovonick, T. L. Shock, and R. J. Fellows, Plant Physiol. 54, 892 (1974).

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indicate that the pathway of [14C]sucrose applied to a green leaf from the apoplast may be different from the pathway of assimilates after 14CO2 fixation. Technique ofMicroautoradiography. Microautoradiography of watersoluble compounds poses a serious technical problem since precise localization of the label on a cellular basis requires that neither loss nor dislocation of the water-soluble label must occur; that is, once the water is removed from the tissue either by freeze-drying or by freeze substitution, contact with water has to be avoided during the further processing of the tissue. A procedure for microautoradiography which uses freeze-drying has been described in detail by Fritz. a4 A detailed procedure which uses freeze substitution with special emphasis on the description of "dry" processing is given by Altus and Canny 35 and Wang and Canny. 32 The reader is also referred to the articles of Fisher~6 and Fisher and Housley37 in which the advantages and drawbacks of the two procedures are evaluated. In brief, the procedure comprises the following steps: 1. Application of label: either ~4CO2 fixation or uptake of [~4C]sucrose may be used (see previous and following sections). 2. Freezing: quick freezing in isopentane, liquid propane, or Freon 22 cooled by liquid nitrogen without cryoprotectant is usually sufficient for preserving the tissue structure at the light microscope level. 3. Removal of water and infiltration of organic solvent and plastic: two techniques can be used, either freeze-drying or freeze substitution. FREEZE-DRYING. The frozen tissue is freeze-dried at low temperature and high vacuum. The dry tissue can be stored for several days without dislocation of the label, provided air humidity is kept low (~<25%). The problems of infiltrating freeze-dried tissue are overcome by infiltrating the tissue with diethyl ether and plastic under high pressure. 34 FREEZE SUBSTITUTION. The frozen tissue is transferred to a plastic miscible solvent (acetone, propylene oxide, or diethyl ether with 10% acrolein) containing a molecular sieve, cooled at - 70 °, and maintained at that temperature for several days. The volume ratio of solvent to tissue has to be high. 36 After slowly warming to room temperature, the tissue is (immediately) processed for embedding in plastic by conventional procedures with the added precaution that all solutions be dried over molecular sieves. 4. Cutting: sections of 1-2/zm are cut dry with a glass knife and transferred to microscopic slides coated with gelatin. 34 E. Fritz, Ber. Dtsch. Bot. Ges. 93, 109 (1980). 35 D. P. Altus and M. J. Canny, Plant, CellEnviron. 8, 275 (1985). 36 D. B. Fisher, Plant Physiol. 49, 161 (1972). 3s D. B. Fisher and T. L. Housley, Plant Physiol. 49, 166 (1972).

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5. Processing for autoradiography: the slides are coated with nuclear emulsion (Ilford K5 or L4, or Kodak NTB2) by gently blowing a thin film of emulsion dried in a metal loop. 35 The slides are then kept in a dry box for several days or weeks, depending on the amount of radioactivity incorporated, and developed according to conventional procedures. Electron Microscopy

As a structural prerequisite for symplastic movements of assimilates, the plasmodesmatal frequency between different cell types has to be sufficiently high to account for the observed translocation rates. Only a few quantitative electron microscopic studies are available to show that the number of plasmodesmata between mesophyll cells, bundle sheath cells, and phloem parenchyma allow for symplastic movement of assimilates along this route. The frequency of plasmodesmata between phloem (vascular) parenchyma and the sieve tube-companion cell complex is different in different plant species. 3s-42 For instance, in maize as and A m a r a n t h u s 39 leaves the sieve tube-companion cell complex is virtually isolated symplastically from the surrounding parenchyma, a fact that can hardly be reconciled with symplastic loading, whereas in sugar beet4° and Cucurbita 4' leaves plasmodesmata between phloem parenchyma and companion cells are abundant, i.e., an entirely symplastic pathway of assimilates could be assumed. On principle, it is premature to take the existence of cytoplasmic connections as proof for symplastic transport because our knowledge of the function and the regulation of plasmodesmata is still too scant. The technique of injecting a membrane-impermeable fluorescent dye into the cytoplasm of a cell and following the spreading of the dye is a promising approach to study functional symplastic continuity. The dye may be injected either d i r e c t l y , 43 with the risk of vacuolar injection, or encapsulated into liposomes~ which after injection into the vacuole can fuse with the tonoplast membrane and release the dye into the cytoplasm. By coinjection of compounds like c a l c i u m 43 and variation of environmental conditions more information on the regulation of the functional status of the plasmodesmata may be gained. 3s R. F. Evert, W. Eschrich, and W. Heyser, Planta 138, 279 (1978). 39 D. G. Fisher and R. F. Evert, Planta 155, 337 (1982). 40 D. R. Geiger, R. T. Giaquinta, S. A. Sovonick, and R. J. Fellows, Plant Physiol. 52, 585 (1973). 4, R. Turgeon, J. A. Webb, and R. F. Evert, Protoplasma 83, 217 (1975). 42 D. G. Fisher, Planta 169, 141 (1986). 43 M. G. Erwee and P. B. Goodwin, Planta 158, 320 (1983). 44 M. A. Madore, J. W. Oross, and W. J. Lucas, Plant Physiol. 82, 432 (1986).

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pH Electrode and Membrane Potential Measurements The mechanism of active sucrose uptake during phloem loading is postulated to be a sucrose-H + symport energized by the electrochemical gradient of protons across the plasma membrane of the sieve t u b e companion cell complex. Therefore, a transient alkalinization of the medium is expected during sucrose uptake, and this has been shown in several cases with leaf tissue. 45-47 Again, this technique does not allow one to identify the cell type responsible for the pH change. This can be accomplished, however, by measuring membrane potential changes with a microelectrode. By putting the tip of a microelectrode into the sieve tube sap exuded from a cut aphid stylet, Wright and Fisher measured a decrease of the membrane potential after supplying sucrose to bark strips of willow, a finding consistent with a sucrose-H + symport mechanism for sucrose uptake into the sieve tube. 48 An advantage of this technique is the precise knowledge of the site of the electrode impalement; on the other hand no cell type but the sieve tube is accessible for such a measurement. Without the use of aphids, the cell type can be identified by labeling the cell with a (charged) fluorescent dye (e.g., Lucifer yellow) injected into the cell by a current pulse via the microelectrode after recording the membrane potential change. Long-Distance T r a n s p o r t

General Comments The translocation of substances within the sieve tubes from the phloem loading site to the sink region is covered under the term long-distance transport. Several features of long-distance transport can be determined, for example, the speed and mass transfer rate of transport; the direction of the flow of substances from a particular leaf, leaf part, or storage organ; the nature of substances which are transported and their concentrations; the factors (e.g., hormones, sink strength, temperature, water availability, and transpiration) which control or modulate transport speed and direction; and the specialization and diversity of sieve tubes within a bundle, or of bundles within an organ, with respect to transport speed, transport direction, and composition of the transported compounds.

45 E. Komor, M. Rotter, and W. Tanner, Plant Sci. Lett. 9, 153 (1977). 4~ W. Heyser, Bet. Dtsch. Bot. Ges. 93, 221 (1980). 47 S. Delrot, Plant Physiol. 68, 706 (1981). J. P. Wright and D. B. Fisher, Plant Physiol. 67, 845 (1981).

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Two types of methods are described in detail: measurement of the translocation profile and sieve tube sap collection. The translocation profile of a particular transported compound can give information about the speed of transport and about its direction in case several plant parts or organs are monitored simultaneously. Measuring translocation profiles in plant parts either by external monitors or by extracting plant tissue will yield the translocation averaged over all bundles and sieve tubes. The collection of phloem sap by incision of only a small bark area or by an aphid which is feeding on one sieve tube, yields translocation profiles of particular bundles or sieve tubes. Collection of phloem sap also allows analysis of the composition of phloem sap and the concentration of the transported compounds. The disadvantage of phloem sap collection is that it is an invasive technique involving opening of the sieve tube and allowing maximal outflow of sap, which probably changes several parameters of transport such as flow speed, sap concentration, and, perhaps, even shifting of the direction of phloem transport. The direction of the flow of substances is also dependent on the vasculature of the plant and its organs and the place and frequency of connections between sieve tubes. Some morphological methods and results are described in Refs. 4 9 - 51.

Translocation Profile of Long-Lived Isotopes The flow of assimilates can be followed in the conventional way by feeding ~4COz to a leaf, cutting the petiole and the stem in portions of equal length a certain time after feeding, and determining the radioactivity in the cut pieces. This method, used in steady-state labeling or pulse-labeling experiments, requires, however, an enormous amount of plant material in parallel sets to follow a detailed time course. The method of feeding the leaf is described by Geiger. 52 The extraction of tissue is principally the same as described before (chemical analysis). When high amounts of ~4C label are used, the translocation can also be followed with externally positioned detectors on the leaf blade. 53 Quantitative analysis of the radioactivity profile data can be complicated and is described elsewhere2 4'55 49 j. Stieber and H. Beringer, Bot. Gaz. (Chicago) 145, 465 (1984). 5o T. C. Vogelmann, P. R. Larson, and R. E. Dickson, Planta 156, 345 (1982). 5t j. T. Colbert and R. F. Evert, Planta 156, 136 (1982). 52 D. R. Geiger, this series, Vol. 69, p. 561. 53 D. R. Geiger and B. R. Fondy, Plant Physiol. 64, 361 (1979). 54 p. E. H. Minchin and J. H. Troughton, Annu. Rev. Plant Physiol. 31, 191 (1980). 5s p. Young, "Recursive Estimation and Time-Series Analysis: An Introduction." SpringerVerlag, Berlin and New York, 1984.

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Translocation Profile of Short-Lived Isotopes* The application of short-lived isotopes with long-ranging radiation has the advantage that the tracer flow can be followed with monitors located along the outside of the plant shoot and can be repeated under slightly varied experimental conditions on the same plant, thus saving plant material and allowing excellent comparison of data. Carbon-11 has a half-life of 20.4 min and decays emitting a fl+ (positron) which has a maximum path length in water of about 4 m m (mean of 1 mm) and is annihilated with a fl- (electron), producing a pair of antiparallel 7 rays of 511 keV. Carbon-11, in the chemical form of HCO2, can readily be produced in quantities adequate for phloem research using a low-energy particle accelerator such as a 3-MeV Van de Graaff accelerator. 56 Carbon-1 1 can be observed either by detecting the charged fl+ particle or by detecting the annihilation radiation. Geiger-Mi~ller (GM) tubes are charged particle detectors and have been used with carbon-l 1.57 The 7 rays can also be detected with scintillation detectors, followed by pulse-height analyzers. This method gives a very large dynamic range that is extended by the use of attenuators which are sequentially removed as the tracer level decays. The main disadvantage of scintillation detection is that large quantities of lead shielding are needed to collimate the 51 l-keV 7 rays. Coincidence counting, in which both of the annihilation 7 rays are detected, 58 partially overcomes the difficulties of shielding but requires duplication of expensive detectors and their associated electronics. The 51 1-keV 7 radiation is not appreciably absorbed by plant material, soil, or air, so in vivo measurements are readily carried out. When following the tracer levels within phloem source or sink regions, it is possible to follow the carbon-I 1 for up to about 12 half-lives (i.e., about 250 min). When following the movement of ~C-labeled photosynthate pulses along the phloem pathway, only 1 h or less of useful data can be obtained. The major disadvantage of ~C is that it must be produced near the site of usage. Access to a particle accelerator is essential, but often small accelerators are

* Section prepared by P. E. H. Minchin and M. R. Thorpe. s6 G. J. McCaUum, G. S. McNaughton, P. E. H. Minchin, R. D. More, M. R. Presland, and J. D. Stout, Nucl. Sci. Appl. 1, 163 (1981). 57 D. S. Fensom, E. J. Williams, D. Aikman, J. E. Dale, J. Scobie, K. W. D. Ledingham, A. Drinkwater, and J. Moorby, Can. J. Bot. 55, 1787 (1977). 5s C. E. Magnuson, T. Fares, J. D. Goeschl, C. E. Nelson, B. R. Strain, C. H. Jaeger, and E. J. Bilpuch, Radiat. Environ. Biophys. 21, 51 (1982).

[21]

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301

available that are not being fully used as they have been superseded by larger machines. The experimental method of observing the movement patterns of l~Clabeled photosynthate is as follows. A clear Perspex chamber is sealed over part or all of the leaf, using a lanolin-calcium carbonate paste. Air is continuously drawn through the chamber using a suction pump vented well away from the experimental area. To load the leaf with l~CO2, the pump is turned off, and l~CO2 is injected into the chamber. After 110 min normal air flow is resumed, thus flushing unfixed ~CO2 from the chamber. The loading time is varied so as to obtain reasonable t~CO2 uptake. With maize 100% uptake is usual, while with wheat as little as 10% may be absorbed. Uptake depends on the area of leaf exposed to the HCO2 . Radiation detectors are placed at various areas around the plant to monitor ~lCO2 levels. A detector monitoring the loaded region will observe ~lC-labeled photosynthate prior to phloem loading as well as that just loaded into the sieve tubes. Since phloem transport occurs at a speed of about 1 cm min-~ in most species and the sieve tubes are very small, little of the observed tracer in the load zone will be within the phloem pathway. The shape of the tracer profiles seen downstream of the loaded zone is determined by the loading processing, while changes in profile shape seen between two pathway detectors depends on the phloem transport process. 59 A detector monitoring the entire plant downstream of any point will register tracer accumulation into this downstream sink. Many variations of this simple system are possible: for example, removal of the epidermis from a bean stem allows [t~C]sucrose to be exuded from the stem apoplast. Using surgically modified pea seeds still attached to the pod, phloem unloading into the seed coat can be followed. Direct quantitative comparison between consecutive experiments is not simple. Since ~C-labeling studies are necessarily short-term, and when pulse-labeling is used, t tC levels and accumulation rates rarely have time to become constant. More complex, and more clear-cut, methods of analysis are needed. Immediate changes in tracer profiles resulting from a treatment can be seen using a qualitative approach. Quantitative analysis based on simple heuristic measurements of tracer profile movement have been used and criticized. 59,6°Analysis based on the assumptions of compartmental analysis have been used and more recently model-independent methods have begun to be employed. 54,55

59 p. E. H. Minchin and J. H. Troughton, Annu. Rev. Plant Physiol. 31, 191 (1980). 60 M. J. Canny, Biol. Rev. Cambridge Philos. Soc. 35, 507 (1960).

302

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[21]

Collection of Sieve Tube Sap by Cutting* The collection of phloem exudate (the mobile fluid of sieve tubes) is a process that is still not fully understood and must, for this reason, be regarded as something of an art. The following review of techniques gives guidance on the following components: (1) plant material, (2) wounding techniques, (3) sap collection methods, and (4) prolongation and control of exudate collection. Not all plants exude phloem sap on wounding; indeed, most species generally exude only insignificant amounts of sap, regardless of how skillfully they are incised. The capacity to exude varies among species and even varieties. Nevertheless, a great many plants have yielded analyzable amounts of exudate. It is advisable to select species and varieties which have proven to yield consistent and reliable amount of phloem sap. A short list is given in Table I. Material selection is important, and it is essential that it be in good physiological state. Generally, plants should be capable of vigorous sieve tube transport. Hence, all conditions which promote rapid growth are beneficial--good supplies of water, nutrients, light, and an appropriate temperature regime are all important factors. The rate of growth does not in itself appear to be the predominant factor, however, since many plants exude extremely well even after rapid growth has been suspended through the imposition of water stress. Depending on the toughness and magnitude of the tissue involved, several methods are used to incise sieve tubes. Large trees, e.g., Manna ash and palms (e.g., coconut and Arenga), are cut with machetes, sickles, or strong knives. 61 Small plants, e.g., Yucca, or woody herbs, e.g., Ricinus, can be incised conveniently using razor blades. Monocotyledonous tissues are cut transversely in toto, but dicotyledonous species are normally cut to the depth of the cambium so that the secondary phloem, but not the xylem, is severed. In all techniques a diagonal cut is made at 30 ° to 45 ° to the horizontal so that exudate will drain laterally to a collection point. It is essential that the blade severing the phloem be very sharp: experience indicates that slicing is more effective than a crushing action. Many tissues contain mineral crystals, e.g., oxalates and silica, which can blunt blades, and therefore their renewal or resharpening is very important. The slicing action should "skate" over the cambium and xylem quite delicately. Recently, solid needles or syringe needles have been used successfully * Section prepared by J. A. Milburn. 61 j. Kallarackal, Sci. Rep. 12, 172 (1975).

[21 ]

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TABLE I PLANT MATERIALS, REGIONS TAPPED, LOCATION OF INCISION, AND APPROXIMATE MAXIMAL RATES OF FLOW

Category

Genus, species, and variety

Vol. hr -I, Tissue tapped

Incisions

max.

Ref.

Vlonocots Palms

~)icots Manna and American

ashes Castor bean

Yucca, Agave Arenga, Cocos, Phoenix

Inflorescence Inflorescence

Whole Whole and side

Several cm a Many cm 3

a, b

Fraxinus ornus, F.

Main trunk (bark)

5-cm cut

U p to 1 cm 3

c

Stem, bark,

l-em cut/

U p to 1 em 3

d

Up to l cm 3 U p to l0 m m 3

e f

americana Ricinus communis ev. gibsonsii

Squashes

Cucurbita slap.

petioles, peduncles Stems, petioles

Legumes

Lupinus alba

Stems,

whole Whole ½-cm wound

petioles,

pods a j. Kallarackal, Sci. Rep. 12, 172 (1975). b j. Van Die and P. M. L. Tammes, Encycl. Plant Physiol., New Ser. 1, 196 (1975). c M. H. Zimmerman and H. Ziegler, Encycl. Plant Physiol., New Set. 1,480 (1975). a j. A. Milburn, Planta 117, 303 (1974). e A. S. Crafts, "Translocation in Plants." Holt, Rinehart & Winston, New York, 1961. f J. A. Smith and J. A. Milburn, Planta 148, 35 (1980).

to promote exudation. In the cryopuncture technique 62 a needle with a finely pointed tip and previously cooled in liquid nitrogen is used to puncture the tissue, incising the vascular trace, and is held in position for 5 sec or longer. Unfortunately, the cryopuncture method does not work with all species but is effective on cowpea (Vigna unguiculataL.). Palms have been tapped since antiquity by placing the severed organ, often trained to incline downward over a period, into a hollow gourd or large pot. Manna ash is tapped by collecting the dried sap from either a collection bowl or the bark itself. Purer sap can be collected in volumetric glassware or plastic bags if the sap can be led from the collection point via some form of tubing. In the laboratory, phloem sap is most conveniently and accurately collected into precision-fabricated microcapillaries. Surface tension draws the liquid horizontally from the collection point. If a scale is fitted along62 j. S. Pate, M. B. Peoples, and C. A. Atkins, Plant Physiol. 74, 499 (1984).

304

TRANSPORT IN PLANTS

[21 ]

FIG. 1. Collection of phloem sap from bark incisions in a Ricinus interuode. Sap is being collected from third incision in series. The stem has been partially ringed to isolate a longitudinal strip of bark. Fine divisions of the upper scale on the ruler are millimeters. (From Smith and Milburu. 63)

side the tube, the volume of exudate can be measured per unit time, hence exudation rates can be calculated (Fig. 1).63 It may be necessary to rely on the waxy surface on the cuticle or bark to prevent sap from escaping down the stem. Such losses will occur if exudation continues after the horizontal microcapillary becomes overfiUed. One can tilt the capillary slightly and fit a collection bottle to the distal end for prolonged collections. However, the handling of microcapillaries is a delicate matter. Fine tubes hold sap quite well, whereas air bubbles rapidly displace the sap in wider tubes unless the tube is slightly inclined to the horizontal or one end is physically blocked. Various methods can be used to enhance the duration of exudation. Plants cannot exude if the loading capacity for solutes is exhausted, and the composition of the sap may be altered. The availability of materials is 63 j. A. C. Smith and J. A. Milburn, Planta 148, 35 (1980).

[21]

PHLOEM TRANSPORT

305

dependent on stored materials from previous growth and also good growth conditions during the exudation process itself. Cooling the severed flower stalk in an ice-filled Dewar flask has been found to prolong exudation in Yucca, presumably because the sealing processes are slowed at low temperature. The cryopuncture method presumably also stops immediate sealing responses because subsequent exudation can be prolonged for many hours or days, though the quantity of exudate is small. It has been reported that chelating agents can prolong exudation, but more recent evidence indicates that such "exudation" is seepage from phloem parenchyma rather than from sieve tubes themselves. Several methods have been used to ensure that the cross section of sampled sieve tubes remains constant. One is a (previously performed) surgical treatment whereby strips of intervening bark are removed. This ensures that a blade always severs the same remaining cross section of bark through which longitudinal transport takes place. 63 Another technique which may be useful is that of bark compression applied some distance from the bleeding incision which can stop exudation instantly. The advantage of this technique is that, at least in Ricinus and Cocos, the compressed tissue regains its capacity to conduct longitudinally after a recovery period of I h or more. A potential problem of phloem sap collection by exudation from incision is contamination with xylem sap. In case the plant is transpiring the negative pressure in the xylem vessels will prevent xylem sap from exuding (but will lead to some loss of phloem sap), but when a petiole or stem is cut or transpiration is low, root pressure will cause exudation of xylem sap at the wound. Since no unambiguous xylem sap marker can be recommended, the only way to estimate contamination by xylem sap is to stop phloem transport by compression of the bark or by treatment of the wound with steam.

Collection of Sieve Tube Sap by Aphid Stylets* General Techniques with Aphids. The interest of plant physiologists in aphids is based on the fact that these insects feed on sieve tubes of angiosperms or on sieve cells of gymnosperms, pteridophytes, and even mosses. Most aphids prefer the vascular bundles of leaves and young shoots, but some species, the lachnids, feed on secondary phloem of twigs and branches. Most aphids can be maintained year-round on fresh host plants, which are more or less specific for each aphid species.64 Some aphid species * Section prepared by W. Eschrich. 64 D. B. Fisher and J. M. Frame, Planta 161, 385 (1984).

306

TRANSPORT IN PLANTS

[21 ]

have been successfully kept on artificial diets for about 40 generations. 65 Their enemies are ladybugs, red spiders, and lacewings (aphid lions). Cages should be lined with nylon fabric of about 300 g m mesh width and irradiated continuously, but not exceeding 60 #Eq m -2 sec-~. Phloem-feeding insects other than aphids seldom are used. ~ Honeydew. The excrement of aphids, honeydew, is secreted in fairly constant intervals. Honeydew is secreted only when the aphid has pierced a sieve element, thus, honeydew consists mainly of sieve tube sap. One droplet of honeydew contains the content of about 5000 sieve elements. Honeydew can be collected on a turntable attached to a 24-hr clock, and when the sieve tubes carry 14C-labeled compounds, the resulting honeydew chronogram can be autoradiographed and their mobility, in principle, can be tested according to the appearance of labeled honeydew. However, phloem mobility can be altered, because the sucking of the aphid can attract the labeled compound not only from far away in a sieve tube, but also from the apoplast and adjacent parenchyma. Aphid Stylets. When an aphid is severed from its mouth parts while sucking sieve tube sap, the high turgor of the sieve tube can cause the stylet stump to continue to produce sieve tube exudate. The exudate can be collected with a glass capillary, fastened to a micromanipulator, under the microscope. The advantages of obtaining stylet exudate are that (1) it is true sieve tube content, not contaminated with juices of the aphids intestines and not altered by the action of the aphids' enzymes, and (2) the styler is buried in a canal of solidified saliva, thus preventing any contamination or dilution with apoplastic solutes of water. The stylet exudate is the purest sieve tube sap which can be obtained. However, it must be considered that any turgor release in a sieve tube causes surging of its contents, and callose deposition. Surging can be accompanied with secretion of material from companion cells, especially enzymes and oligonucleotides. Cutting Stylets. Aphid stylets are cut with (1) the microscope laser64,~s,67 or (2) the radio-frequency microcauter (RFM). 64,6s,69 The aphids usually are directed to certain areas of the experimental plant in clip cages, made from a hair clip to which a suitable aerated cage is attached (Fig. 2). After a few hours or overnight, most aphids have settled and started to produce honeydew. Aphids may reproduce in such cages. They usually feed from 65 p. Eberhardt, Z. Vergl. Physiol. 46, 169 (1962). 66 C. A. Barlow and M. E. MeCully, Can. J. Zool. 50, 1497 (1972). 67 C. A. Barlow and P. A. Randolph, Ann. Entomol. Soc. Am. 71, 46 (1978). 6s N. Downing and D. M. Unwin, Phys. Entomol. 2, 275 (1977). 69 D. M. Unwin, Phys. Entomol. 3, 71 (1978).

[21 ]

PHLOEM TRANSPORT

,

~

~

~

"

~ ' ~ 2 ~

f

-,

307

--grassleaf

felt rim ~/ /(~,~glass slide V ~'~ -aphid ~ / tubing L~'~ nylon fabric

FIG. 2. Cage for directing aphids on a grass leaf. (Drawn by W. Eschrich.)

the lower side of a leaf, the phloem side. On branches, aphids may be encaged simply between two girdles of stiff grease. When using the RFM, the microantenna may be touched to the aphid mouth part in any position. Some aphid species are very nervous, and they will retract the stylets if they are not narcotized before with a gentle stream of CO2. [In some countries, RFM, which operates at citizen band (CB) frequencies, must have a permit from the post office.] The microscope laser is preferentially mounted on a vibration-reduced table, on which also the plant with the aphids are placed. Either the microscope with the attached laser or the aphid must be movable in three dimensions. The microscope is usually mounted in a horizontal position and must have objectives with long operating distances (at least 6 mm). The outlet of the laser beam is attached to the tubes of a microscope camera. Convenient are lasers like neodymium-glass lasers which operate in the near-infrared (> 1000 nm) because this light can be absorbed by thick glass which protects the operator's eyes. The laser beam has to be centered on black target paper with a hairline across inserted in the eye piece. Microscope laser combinations are not yet available on the market. (Zeiss, Oberkochen, FRG, and Lasertechnik GmbH, 6056 Heusenstamm, FRG, produce such combinations on request.) Styler Exudate. Stylet exudate contains sucrose in fairly high concentrations (40% or more), but since collection in capillaries can take hours or even days, considerable evaporation of water occurs. For exact quantitative

308

TRANSPORT IN PLANTS

[9.1 ]

determinations, the exuding stylet stump can be wrapped in a drop o f oil. 7° For determination of amino acids, hormones, and inorganic components in the sieve tube exudate, techniques of gas chromatography (GC), highperformance liquid chromatography (HPLC), and atomic absorption spectroscopy are indispensible. Stylet stumps differ greatly in their exuding capacities, and exudation can vary during the period of collection. Sucrose concentration of two stylet stumps positioned at the same leaf may differ considerably. Phloem Unloading General Comments

The term phloem unloading denotes the pathway of assimilates out of the sieve tube into heterotrophic, growing, or storage tissue like young (sink) leaves, stems, fruits, and roots. The same questions outlined for phloem loading concerning symplastic versus apoplastic pathways, localization of active transport sites, and involvement of cell wall-bound invertase also have to be solved at this stage of phloem transport. Different mechanisms of unloading appear to be operative within different sink tissues. Data from sink leaves (sugar beet7~) and roots (pea72 and corn 6,73) have been interpreted to support the view of an entirely symplastic pathway of assimilates from the sieve tube to the final destination cells, whereas an apoplastic step seems to be involved during unloading in the stem (sugarcaneTM and bean 75) and has to occur in fruits (maize, 76 soybean, 77 and beans 7a,79)because of the lack of symplastic connections between maternal and embryonic tissues. According to the frequency of plasmodesmata between sieve tube-companion cell complexes and parenchyma cells, the apoplastic step has been proposed to be localized in the parenchyma cell region, not at the sieve tube-companion cell complex itself. 76'77'79 For symplastic unloading a sucrose gradient has to be maintained either by metabolic conversion of sucrose in the cytoplasm of the sink cells, 70 S. Kawabe, T. Fukumorita, and M. Chino, Plant Cell Physiol. 21, 1319 (1980). 7t j. Gougler Schmalstig and D. R. Gciger, Plant Physiol. 79, 237 (1985). 72 p. S. Dick and T. Ap Recs, J. Exp. Bot. 26, 305 (1975). 73 R. D. Warmbrodt, Bot. Gaz. (Chicago) 146, 169 (1985). 74 K. T. Glasziou and K. Gaylcr, Bot. Rev. 38, 471 (1972). 75 p. E. H. Minchin and M. R. Thorpe, J. Exp. Bot. 35, 538 (1984). 76 F. C. Felker and J. C. Shannon, Plant Physiol. 65, 864 (1980). 77 j. H. Thornc, Plant Physiol. 67, 1016 (1981). 7s p. Wolswinkel and A. Ammerlaan, Planta 158, 305 (1983). 79 C. E. Offler and J. W. Patrick, Aust. J. Plant Physiol. 11, 79 (1984).

[2 1]

PHLOEM TRANSPORT

309

promoted by neutral invertase or sucrose synthase, or by hydrolysis of sucrose in the vacuole by an acid invertase. The activity of at least one of these enzymes has to be high in tissues where symplastic unloading occurs. Although reasonably postulated, the involvement of carder proteins in apoplastic unloading has not yet been proved, nor has the requirement for metabolic energy been established as a general feature of assimilate release into the apoplast. Recent investigations using protein [p-chloromercuribenzenesulfonic acid (PCMBS)] and metabolic [KCN, carbonyl cyanide m-chlorophenylhydrazone (CCCP), azide] inhibitors have produced differing results. 7a'8°'81 Since it could be deduced from the direction of the concentration gradient at the apoplastic step whether an active transport mechanism has to be postulated or whether a facilitated diffusion mechanism would be sufficient, the sucrose concentrations in the apoplast and the adjacent parenchyma have to be determined. This has been done for the soybean seed coat, s2 and results demonstrate that sucrose release into the apoplast is an active transport process. During apoplastic unloading a cell wall-bound invertase may 74,76 or may not 83 be involved, depending on the species. Therefore, its role as a "reflux valve," as suggested by Eschrich, s4 cannot be generalized. The presence of a cell wall-bound invertase, together with a very active hexose uptake system in tissues where symplastic unloading has been shown to be operative, 6 points to a possible additional pathway of sucrose unloading into the apoplast and retrieval as hexoses, which may undergo a different metabolic fate than incoming sucrose, thereby increasing sink strength and regulating partitioning of incoming assimilates into different metabolic pools in the absence of neutral invertase. In sink tissues as well as in source tissue the cell wall-bound invertase may be associated with a special cell type; this could be identified by immunohistochemical techniques, s5 In principle, the same approaches as described for phloem loading can be followed for phloem unloading. These include radioactive tracer studies, microautoradiography, and electron microscopy. A technique for Studying apoplastic unloading within legume seed coats is presented here. The method should also be applicable to other plant species with relatively large seeds.

8oj. W. Patrick, Z. Pflanzenphysiol. 111, 9 (1983). 81G. A. Porter, D. P. Knievel,and J. C. Shannon,Plant Physiol. 77, 524 (1985). 82R. M. Giffordand J. H. Thorne,Plant Physiol. 77, 863 (1985). s3j. H. Thorne,Plant Physiol. 65, 975 (1980). W. Eschrich,Ber. Dtsch. Bot. Ges. 93, 363 (1980). 8sL. Fayeand A. Ghorbel,Plant Sci. Lett. 29, 33 (1983).

310

TRANSPORT IN PLANTS

m

[21]

i

3cm Fno. 3. Diagrammatic representation of the procedure for obtaining "empty" seed coats of Vicia and measuring the release of sugar and amino acids. (a) The pod is placed in a trough of metal in such a way that both halves of the pod are in a more or less horizontal plane. Subsequently, a window is made in the upper half of the pod wall and the four ovules are surgically treated. (b) As shown in cross section, an "empty" ovule is filled with a buffered solution. During the experiment the bottom of the trough is covered with a layer of tissue paper moistened with water and the trough is covered with aluminum foil to maintain a high humidity. (From Wolswinkel and Ammerlaan. 7s)

Techniques of Measuring Phloem Unloading in Legume Seed Coats* The development of legume seeds is characterized by a continuous transfer of assimilates from tissues of maternal origin (seed coat) to the embryonic tissues. Photosynthate imported from the pod wall must pass the seed coat before it is available for uptake by the embryo. For soybean 77 and Phaseolus vulgaris 79 data on phloem in the seed coat have been published. Recently a technique has been developed to measure unloading of assimilates from the seed coat of developing legume seeds. After removal of the embryo from a developing ovule, the "empty" ovule can be filled with a solution to measure assimilate release from the seed coat (Fig. 3). The sites of sucrose and amino acid unloading in the seed coat are accessible via the seed coat apoplast and can be challenged with inhibitors or other solutes to characterize the unloading process. Seed Coat Preparation. The pod is placed in a small chamber (e.g., a trough) in such a position that both halves of the pod wall are in a more or less horizontal plane. First, an approximately rectangular window is made in the upper half of the pod wall. Care is taken to prevent damage to the * Section prepared by P. Wolswinkel.

[21]

PHLOEM TRANSPORT

31 1

ventral or dorsal vascular bundles of the pod. Subsequently, in experiments with pea or broadbean, a more or less rectangular incision is made in the seed coat of several ovules. The part of the seed coat which is excised from each ovule during this procedure represents about 10% of the seed coat. After removal of the embryo from a number of developing seeds (during this procedure each embryo is cut into pieces), the pod is placed in a horizontal position in the "operating room," and the empty ovules are filled with a buffered solution [e.g., 2 or 10 m M 2-N-morpholinoethanesulfonic acid (MES) buffer, pH 5.5]. During the experiment, the bottom of the chamber is covered with a layer of tissue paper moistened with water, and the chamber is covered with aluminum foil to maintain a high humidity. The empty ovules (without embryo but filled with solution) remain attached to the pod vascular bundles via the funiculus. The formation of a funicular abscission layer in nearly mature seeds can add considerable difficulty to the surgical treatment, but until the stage of development in which the embryo has attained about one-half the final dry weight, this is not a serious problem in Pisum sativum and Viciafaba. Pulse-Labeling Procedures. Several procedures have been described in r e c e n t repol"tS. 78'80'86'87 In most cases, labeled compounds are administered to the plant after the surgical treatment. The exposure of a leaf or a leaf part to 14CO2 in an assimilation chamber will produce a ~4C-labeled photosynthate pulse. Labeled compounds can also be applied to an abraded leaf. In double-label experiments with a mixture of 3H and ~4C isotopes of sucrose and/or amino acids, label can also enter the plant via the petiole subtending the fruit used for the experiment. First, the leaflets of a compound leaf are removed and the rachis is severed about l cm below the site of attachment of the uppermost leaflets. Immediately after severing the midvein, it is placed in a narrow glass tube containing labeled solutes in water (e.g., 0.1 ml in the case of pea). This solution is rapidly taken up into the plant (in most cases within 15 - 3 0 min). The distribution pattern of 3H and 14C introduced in this way indicates a rapid entry from the xylem into phloem cells. Exudate Collection. During the course of an experiment ( 10- 12 hr), the solution filling an empty ovule is regularly collected for analysis and a fresh solution transferred into the seed coat cavity. Standard methods of radioactivity measurements or chemical analysis can be used for measurements of solutes present in the seed coat exudate or solutes present in seed coat extracts. Comments. The rate of phloem transport of sucrose and amino acids 86 j. H. Thorne and R. M. Rainbird, Plant Physiol. 72, 268 (1983). 87 p. Wolswinkel and A. Ammerlaan, Physiol. Plant 61, 172 (1984).

312

TRANSPORT IN PLANTS

[22]

into "empty" ovules is dependent on the solute concentration in the solution filling empty ovules. In experiments with pea, a solute (e.g., sucrose or mannitol) concentration of about 400 m M is optimal for sucrose and amino acid transport into the cavity within the empty ovules. The results were comparable to transport into intact ovules. When empty ovules are excised from the maternal plant by cutting the funiculus, the release of solutes from the seed coat is completely dependent on material present in the seed coat at the moment of cutting. By comparing solute release from excised seed coats with solute release from attached seed coats, it is possible to discriminate between effects on unloading from the seed coat, and effects on assimilate transport into the seed coat. Acknowledgments The authors are extremely grateful to Drs. W. Eschrich (G6ttingen), P. E. H. Minchin and M. R. Thorpe (Lower Hut), B. R. Fondy, D. R. Geiger, and S. Delrot (Dayton), J. A. Milburn (Armidale), P. Wolswinkel (Utrecht), and D. B. Fisher (Pullman) for giving detailed descriptions of methods and comments on them.

[22] Patch Clamp Measurements on Isolated Guard Protoplasts and Vacuoles

Cell

By KLAUS RASCHKE and RAINER HEDRICH

Guard cell protoplasts provide a suitable material for ion transport studies for three reasons. (l) The osmotic motor of the guard cells regulates gas exchange of leaves relatively rapidly. Large ion fluxes per unit area of plasma membrane are necessary to produce the volume changes in the guard cells that are required to open and close the stomatal pores. Channels and pumps in the plasmalemma and tonoplast of guard cells are expected to be numerous or of larger capacity than in other plant cells.~ (2) Because, in the whole leaf, gas exchange has to be regulated in response to changes in external and internal factors, 2 mechanisms that control ion fluxes are presumably highly developed in guard cells. (3) Because ion

' K. Raschke, R. Hedrich, U. Reckmann, and J. I. Schroeder, Bot. Acta 1 0 1 , 2 8 3 (1988). 2 K. Raschke, Encycl. Plant PhysioL, New Ser. 7, 382 (1979).

METHODS IN ENZYMOLOGY, VOL. 174

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