Short communication: The anatomy of the ... - CSIRO Publishing

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Functional Plant Biology, 2005, 32, 367–374

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Short communication: The anatomy of the pathway of sucrose unloading within the sugarcane stalk Kerry B. WalshA,C , Russell C. SkyA and Sharon M. BrownA,B A Plant

Sciences Group, School of Biological and Environmental Sciences, Central Queensland University, Rockhampton, Qld 4702, Australia. B Current address: School of Land and Food, University of Queensland, Brisbane, Qld 4072, Australia. C Corresponding author. Email: [email protected]

Abstract. The physical path of sucrose unloading in the sugarcane stalk is described. About 50% of the vascular bundles in the internodes were located within 3 mm of the outside of the stalk. These bundles were inactive in long distance sucrose transport, as assessed by dye tracers of phloem flow. A sheath of fibres isolates the phloem apoplast from that of the storage parenchyma. In bundles associated with long distance transport (i.e. in the central region), the fibre sheath is narrowest to either side of the phloem fibre cap, and consists of living cells with plasmodesmata within pits in the secondary wall. Plasmodesmata were also arranged into pit fields between cells of the storage parenchyma. Since the vascular apoplast is isolated from the apoplast of the storage parenchyma, sucrose must move through the symplast of the fibre sheath. The calculated flux of sucrose through plasmodesmata of this cell layer was at the low end of reported values in the literature. Sucrose unloading within the storage parenchyma may also follow a symplastic route, with unloading into the apoplast of the storage parenchyma occurring as part of a turgor mechanism to increase sink strength. Keywords: apoplastic movement, plasmodesmata, sucrose unloading, sugarcane, symplastic movement.

Introduction Sugar levels vary within harvested sugarcane, and can be manipulated by choice of genotype or by agronomic practices (e.g. timing of harvest, nitrogen status at harvest, use of inhibitors of vegetative growth such as ethrel and glyphosphate). Understanding the pathway of sucrose movement from phloem to storage tissue in the sugarcane stem is integral to the study of control and subsequent genetic modification of sucrose unloading and storage in the sugarcane stem (e.g. Patrick in Wilson 1992). For example, the work by Ma et al. (2000) on the metabolic engineering of cell wall invertases in sugarcane was based on the premise that in the sugarcane stem, sucrose unloaded from the phloem passes through the apoplast to the cytoplasm of storage cells and is then loaded to vacuoles, with flux limitation either by uptake by storage cells or loading to vacuoles. However, the flux of sucrose may be path limited, for example, if there is a symplastic step in the unloading pathway, thus, a study focused on plasmodesmatal function is warranted. The pathway of sucrose unloading in the sugarcane stem may be through the symplast and / or apoplast. The symplastic

pathway involves movement directly from cell to cell via plasmodesmata. The apoplastic pathway involves movement via the cell wall space, with carriers on the cell membrane responsible for loading and unloading between the cell and the apoplast. Sugarcane stems contain high levels of sugars in the apoplast, leading Hawker and Hatch (1965) to suggest that sucrose was unloaded from the phloem complex directly into the apoplast, with uptake of inverted sugars from the apoplast by the storage parenchyma cells. However, the vascular bundles (VBs) of the cane stalk are surrounded by a sheath of fibre cells, which serve to isolate the xylem water from the apoplast of stalk storage tissues (Jacobsen et al. 1992; Welbaum et al. 1992). Welbaum et al. (1992) observed plasmodesmata between all cell types in the pathway from phloem to storage parenchyma and suggested that sucrose must follow a symplastic path, with subsequent release into the apoplast of storage parenchyma. However, the observation of a plasmodesmatal connection is not proof that the majority of solute flux is symplastic. A quantitative analysis of plasmodesmatal frequency and sucrose fluxes can be used

Abbreviations used: CF, 5,6-carboxy fluorescein; FW, fresh weight; LYCH, Lucifer yellow-CH; pd, plasmodesmata; VB, vascular bundle. © CSIRO 2005

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to calculate the flux rate per plasmodesmata. Comparison of such a value relative to that of other published studies can be used as an index to the capacity of plasmodesmata to handle the sucrose flux in the system under consideration. For example, fluxes through plasmodesmata (pd) are reported at 1.5 × 104 for wheat leaves (Kuo et al. 1974), 8.4 × 105 in Abutilon nectaries (Gunning and Hughes 1976), 2.26 × 104 − 1.2 × 106 in Phaseolus stems (Hayes et al. 1985) and 2.7 × 107 pmol sucrose cm−2 pd s−1 in the endodermis of soybean nodules (Brown et al. 1995). Values are often reported on a per unit cross-sectional area to accommodate differences in plasmodesmatal radius (with the assumption that flow through tubes of such small radii will not obey the Hagen–Poiselle law, that flux is proportional to the fourth power of the radius). In this report, we quantify the flux rate of sucrose through the plasmodesmata of the fibre sheath of the VBs, and compare this rate to other published reports of flux through plasmodesmata to assess the potential of the symplastic pathway to support observed rates of sugar accumulation in the cane stalk.

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Apoplastic discontinuity Apoplastic discontinuity was assessed in stem tissue by monitoring movement of the membrane-impermeant fluorescent stain Lucifer yellow-CH (LYCH) (Sigma, St Louis, MO) along stem vascular traces. Lengths (200 mm) of stem were cut at the internodes and placed into a beaker containing 1% LYCH (aqueous). A vacuum (85 kPa) was imposed for 1 min at room temperature (25◦ C). Cubes (5 mm per side) of the tissue were fixed overnight in 5% glutaraldehyde in 0.05 M sodium phosphate buffer (pH 6.9). Hand sections were cut for observation by epifluorescence microscopy, using blue excitation wavelengths. The movement of LYCH applied to storage parenchyma of hand-cut (fresh) sections of the fourth stalk internode was also monitored. Phloem transport Phloem transport was assessed by monitoring movement of the phloemmobile, membrane-impermeant dye, 5,6-carboxy-fluorescein (CF) (Sigma), applied in the membrane-permeant diacetate form. Dye was applied to an abraded area (15 cm2 ) of the leaf, and covered with parafilm to limit evaporation. Feeds (n = 4) were undertaken to the fourth leaf (counting from the top visible dewlap) using separate stalks. After 2 h, each stalk was harvested and transverse hand sections of the leaf lamina, sheathing base and internodes were prepared for observation by epifluorescence microscopy, using blue excitation wavelengths.

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Saccharum officinarum L. (var. Q124) plants were maintained on the Central Queensland University, Rockhampton campus, fertilised monthly with Greenland 44 (equivalent of 300 kg N ha−1 year−1 ). Stalk material (three stems) was harvested at monthly intervals from March to August 1994. At each harvest, internodes were juiced, and soluble sugar content estimated in a 1 : 5000 dilution by the anthrone method (Yem and Willis 1954), using sucrose as a standard. For a given cane, leaves and internodes were numbered with reference to the first visible dewlap leaf. Vascular bundle counts To allow counts of vascular bundles within the stalk, transverse hand sections of all internodes were stained with 1% (w / v) phloroglucinol according to the methodology of Jacobsen et al. (1992). Tissue was observed by epifluorescence microscopy (Eclipse E600, Nikon, Tokyo), using blue excitation wavelengths. Suberisation and lignification Suberisation and lignification of cells in stem tissue was examined using the fluorescent stains Auramine O and berberine sulphate (Ajax Chemicals, Crown Scientific, Brisbane, Qld). Fresh sections were cut by hand and stained for 30 s on a glass slide with 1% Auramine O. Tissue was observed by epifluorescence microscopy, using blue excitation wavelengths. Tissue (∼1 mm3 ) containing VBs was also dissected from internode 4 (during June), from a depth of at least 5 mm from the epidermis and fixed for 2 h at 4◦ C in phosphate-buffered 2.5% glutaraldehyde containing 0.6 M sucrose (pH 7.2), post-fixed in 2% osmium tetroxide for 2 h, then dehydrated in an acetone series, embedded in LR White resin (ProSciTech, Qld), and sectioned. Serial sections (2 µm thick) were stained for suberin and lignin using the berberine-sulphate staining protocol (Brundrett et al. 1988). All fluorescent microscopy was photographed with 100ASA Kodak TMAX film (Kodak, Rochester, NY) exposed at 800ASA.

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Internode Fig. 1. Sugar and dry matter content, and extent of storage cell wall lignification / suberisation of internodes within canes. (A) Sugar distribution within canes of sugarcane, during maturation of the crop. Data is the mean of three canes (typical coefficient of variation 8%; , March; , May; , June; , July; 䊏, August harvests). (B) Dry matter content ( ) and extent of parenchyma cell lignification / suberisation ( , assessed on a cell wall length basis within a field of view using microscopic examination of berberine sulfate stained sections) within a cane harvested during July.

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Sucrose unloading within the sugarcane stalk

Symplastic continuity was also assessed by microinjection of CF into cells within the vascular bundle (hand-cut section, under a layer of silicon oil). Electron microscopy For electron microscopy (EM), gold coloured sections were cut using glass knives, collected on Formvar-coated copper grids, and stained with 2% uranyl acetate for 30 min, then 0.2% lead citrate for 5 min. Electron micrographs were taken with a JEOL 100C transmission electron microscope (TEM; JEOL 100C, JEOL Australia, Sydney, NSW). A crude form of freeze–fracture was used for scanning electron microscopic (JEOL JSM-5300 LV, JEOL Australia) examination, with tissue pieces plunged into liquid nitrogen, then split with a sharp razor blade before viewing in the scanning electron microscope. A freeze– fracture replica method was used for TEM observations, following the method by Shaw and Stowe (1982). Calculations and estimation of associated error Standard errors of means were compounded in calculations involving the means by the general formulae: A / B[(SE2A / A)2 + (SEB / B)2 ]0.5 , for the division of A and B, and: A × B[(SE2A / A)2 + (SEB / B)2 ]0.5 , for the multiplication of A and B.

Results Stalk sugar content increased as winter progressed, with first the lower internodes, and then upper internodes, accumulating a maximum of 14% soluble sugar (sucrose equivalents, FW basis). The maximum rate of sugar


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accumulation occurred in the fourth internode below the node of the first visible dewlap leaf during the month of June (Fig. 1; internode 4 became internode 7 during this month, accumulating 76 mg g−1 FW) (see also Table 1, items 1–4). The number of vascular bundles in a cross section of stalk increased from ∼850 in internode 1 to ∼1100 in internode 10 (data not shown). Vascular bundle sheaths were lignified / suberised in all internode age classes examined, but the extent of lignification / suberisation of the storage parenchyma increased from 0% in internode 1 to >60% in internode 10 (Fig. 1). The dry matter content of tissue also increased with internode age, reaching a plateau after internode 10 (Fig. 1). About half of VBs (i.e. ∼400) noted in internode cross section were tightly packed around the periphery (outer 3 mm) of the stem, with the remainder relatively widely scattered in the remaining ‘central’ region (Table 1, item 5; Jacobsen et al. 1992). In the central region where VB density was lower, ∼10 storage parenchyma cells separated the bundles. The number of VBs per internode did not vary greatly with internode number (data not shown). All VBs possessed a complete sheath of thick-walled, living (as judged from the presence of cytoplasmic contents) fibre cells (Fig. 2A), although in VBs of the peripheral region this sheath was much more extensive. Following feeding of CF, the symplastic marker, to leaf 4, tracer was noted in leaf VBs (data not shown), and in central VB of internodes subtending the leaf (Fig. 2C). Following placement of a cut stalk into a solution of

Table 1. Calculation of rate of sucrose flux through plasmodesmata (pd) of the fibre layer of the bundle sheath of vascular bundles of Saccharum officinarum L. var. Q124 Data is presented of the internode and period estimated to have the highest rate of sucrose accumulation (internode 4 became internode 7 during the month of June). Data presented as mean ± SE. Sucrose accumulation, internode fresh weight and internode length were estimated from hand cut transverse sections, n = 3 (separate canes). Sucrose accumulation per internode calculated by: (item 1) × (item 2). Number of VB in the stalk central region was estimated from hand-cut transverse sections, n = 3 (separate canes). Fibre sheath ‘phloem shoulder’ perimeter was estimated from resin embedded transverse sections (e.g. Fig. 2), n = 10 (vascular bundles). Surface area of fibre sheath was calculated by: (item 6) × (item 5) × (item 4) / 1000. Pit frequency data from outer tangential walls of fibre sheath in resin embedded transverse sections of 2 µm thickness, n = 10 (vascular bundles). Plasmodesmata per pit field data from freeze–fracture electron micrographs, n = 8 (pit fields). Bundle sheath pd per internode calculated by: (item 9) × (item 8) × (item 7) × 106 µm3 mm−3 . Sucrose flux through bundle sheath pd calculated by: (item 3) / 342 mg mmol−1 × 109 mmol pmol−1 × 24 h d−1 / (item 10). Diameter of plasmodesmatal neck data of TEM sections, n = 10 (plasmodesmata). Sucrose flux through bundle sheath calculated by: (item 11) × 3600 s h−1 / [3.14 × (item 12 × 10−7 )2 ] Item 1 Sucrose accumulation 2 Internode fresh weight 3 Sucrose accumulation per internode 4 Internode length 5 Number of VB in the stalk central region 6 Fibre sheath ‘phloem shoulder’ perimeter 7 Surface area of fibre sheath 8 Pit frequency 9 Plasmodesmata (pd) per pit field 10 Bundle sheath pd per internode 11 Sucrose flux through bundle sheath pd 12 Diameter of plasmodesmatal neck 13 Sucrose flux through bundle sheath (area basis)

Value 2.71 ± 0.18 29.1 ± 3.2 79 ± 10 50.5 ± 4.3 420 ± 80 120 ± 35 2,550 ± 910 0.008 ± 0.003 42 ± 13 8.6 × 108 ± 5.2 × 108 0.011 ± 0.0069 16 ± 3 3.9 × 105 ± 2.5 × 105

Unit FW d−1 g mg internode−1 d−1 mm internode−1 µm vascular bundle−1 mm2 internode−1 pits µm−2 pd pit−1 pd internode−1 pmol pd−1 h−1 nm pmol sucrose cm−2 pd s−1 mg g

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the membrane-impermeant dye, lucifer yellow, was dye was observed in upper internodes within VBs, but not in the storage parenchyma (Fig. 2B). Similarly, lucifer yellow introduced to the apoplast of storage parenchyma did not

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pass the fibre sheath into vascular bundles (Fig. 3B). When microinjected into cells within the vascular bundle, lucifer yellow dye was noted to rapidly move between cells in the vascular bundle, but the fibre sheath retarded movement (Fig. 3A). The fibre cell walls were ∼2 µm thick and were lignified or suberised, as indicated by berberine-sulphate staining (Fig. 2A). In the VBs of the central region, the sheath was characterised by a fibre cap directly ‘above’ the phloem (i.e. towards the epidermis), but was reduced to a layer only about one or two cells deep on the two flanks of the phloem fibre cap (Fig. 2A). The perimeter of VB fibre sheath in the phloem flank position was measured for SEM-visualised transverse sections and multiplied by internode length to give area (Table 1, items 6, 7). Plasmodesmata were observed at all cell interfaces in the phloem to storage parenchyma pathway (e.g. Fig. 5A). The plasmodesmata linking VB fibre cells to adjacent storage parenchyma cells were located within large pits (up to 1 µm diameter) (Fig. 4A, B). Pit frequency in the fibre cells of the phloem shoulder–storage parenchyma interface was measured of resin sections using light microscopy (Table 1, item 8). Plasmodesmata were also clustered into pit fields between storage parenchyma cells (with ∼40 per pit, Fig. 5; Table 1, item 9). As it was not possible to reliably section through the pit fields of the VB fibres, it was assumed that the plasmodesmatal frequency in these pits was as measured of the storage parenchyma pit fields. The number of plasmodesmata between the phloem flank fibres and the storage parenchyma for internode 4 (Table 1, item 10) was calculated by multiplying surface area of the phloem flank fibres, pit frequency in these fibres, and the estimate of plasmodesmata per pit (Table 1,

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Fig. 2. Transverse sections of sugarcane stalk (internode 4). (A) Resin sections stained with berberine sulphate for localisation of lignin and suberin and viewed with blue excitation under epifluorescent optics. Vascular bundles in the inner zones of the stalk have a single layer of fibre cells in the phloem ‘shoulder’ region (between arrows). (B) Section of fixed sugarcane stalk (internode 4) of a stalk placed in a solution of lucifer yellow and allowed to transpire. Section viewed with blue excitation under epifluorescent optics. Lucifer yellow movement was contained within the vascular bundles, constrained by the fibre layer. (C) Hand-cut fresh section of sugarcane stalk from a plant on which a leaf was fed carboxy fluorescein. Transport was restricted to the phloem tissue of scattered vascular bundles in the central region of the stalk (arrow). Scale bar represents 50 µm for panel A and 270 µm for panels B and C.

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Fig. 3. Transverse hand sections of sugarcane stalk (internode 4), viewed with blue excitation under epifluorescent optics. Scale bar represents 150 µm. (A) The membrane-impermeant dye lucifer yellow was microinjected into the cells within the phloem tissue. Over a period of minutes dye moved between cells within the phloem, but did not move beyond the fibre layer. (B) Lucifer yellow introduced into the apoplast of the storage parenchyma spread rapidly within the storage parenchyma tissue, but did not pass the vascular bundle fibre layer.


items 7, 8, 9). Plasmodesmata were typically constricted in the region of the connection to the cell membrane (Fig. 4B, diameter 32 ± 2.7 nm, n = 10), and dilated within the cell wall region (diameter 64 ± 1.4 nm, n = 12) (cf. 37 nm diameter of straight-channelled plasmodesmata measured in soybean nodule vascular endodermis by Brown et al. 1995).


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Fig. 5. Plasmodesmata in pit fields of storage parenchyma cell interfaces, viewed by transmission electron microscope. Plasmodesmatal fields are indicated by arrows. Scale bar represents 500 nm. (A) Transverse section of plasmodesmatal field (arrow). (B) Freeze–fractured replica of cell wall surface between two cells.

Fig. 4. Transverse section of resin embedded sugarcane stalk (internode 4) examined using transmission electron microscopy. (A) Sclereid cell in the phloem ‘shoulder’ region of a vascular bundle, showing a pit containing plasmodesmata. Scale bar represents 2 µm. (B) Detail of a pit shown in (A), illustrating plasmodesmata through the primary cell wall (small arrowheads). The section is oblique to the direction of the pit pair, such that the connecting pit through the secondary wall of the lower cell (large arrowhead) is not completely demonstrated. Scale bar represents 50 µm.

Finally, the rate of sucrose flux through the plasmodesmata of the fibre sheath of a vascular bundle was estimated by dividing the assessed maximum rate of sucrose accumulation per internode by the estimated number of plasmodesmata per internode (Table 1, item 11) and the cross-sectioned area of the plasmodesmata (Table 1, item 12). Storage parenchyma cell walls lignified during internode maturation (Fig. 1B). Lignification began in cells adjacent to the vascular bundle (with less than 5% of storage cells lignified in internode 4), and extended outwards with time (with over 60% of cells lignified

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in internode 10). Certain cells never lignified (Fig. 6). Fluid was present in the apoplast of storage parenchyma tissue (Fig. 7). Discussion We conclude that the vascular bundles in the peripheral region of the stem serve to strengthen the stalk (Wilson 1990) but were not active in long distance phloem transport. This transport role was served by VBs of the central region. Vascular bundles were surrounded by a fibre sheath with lignified and / or suberised cell walls, which isolated the vascular apoplast from that of the storage cell parenchyma. This observation is consistent with the reports by Jacobsen et al. (1992) and Welbaum et al. (1992), where xylem water was isolated from the apoplast of storage tissue. Thus, sucrose cannot travel via a strictly apoplastic path between the phloem complex and storage parenchyma.

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B Fig. 6. Fresh hand section of internode storage parenchyma, stained with auramine O and viewed with epifluorescent optics. Transverse (A) and longitudinal (B) section demonstrating cells with reduced cell wall thickening and lignification. Scale bar represents 100 µm.

Fig. 7. Scanning electron micrograph of frozen, fractured internode storage parenchyma. Intercellular spaces contain ice crystals. Scale bar represents 10 µm.

All cells in the pathway from phloem to storage parenchyma in VB of the central region were alive and connected by plasmodesmata, as observed by Welbaum et al. (1992). Thus, sucrose may move symplastically from the phloem, through the fibre sheath, and throughout the storage parenchyma. However, CF movement from a fed leaf was restricted to within the stalk VB. Also, movement of Lucifer yellow microinjected into VB cells was restricted to within the bundle sheath. These observations are consistent with a ‘flux restriction’ in the path from phloem to storage parenchyma at the fibre sheath. The fibre sheath around each VB of the central region was reduced to only 1–2 cells adjacent to the phloem fibre cap. We surmise that the majority of sucrose flux from the phloem to the storage parenchyma will pass through this section of the fibre sheath, as a path of least resistance. Sieve element–companion cells–vascular parenchyma interfaces were well endowed with plasmodesmata (data not shown), as were storage parenchyma cells, such that the rate of sucrose flux per plasmodesmata in this tissue would be much lower than that in the fibre sheath. Further, given the apoplastic discontinuity in the bundle sheath, all sucrose movement must be symplastic through these cells layers, while movement within the vascular bundle and storage parenchyma may have an apoplastic component. We therefore conclude that the passage of sucrose through the plasmodesmata of the fibre sheath is a likely point of rate limitation in sucrose transport from phloem to storage cell. This conclusion is similar to the situation in the legume root nodule, where vascular bundle sheaths represent a point of flow restriction between the phloem and infected cells (Brown et al. 1995). The maximum flux of sucrose through the plasmodesmata of fibre cells was estimated at 3.9 × 105 pmol cm−2 pd s−1 (Table 1). This rate is low (e.g. Brown et al. 1995 report


a flux of 2.7 × 107 pmol sucrose cm−2 pd h−1 through the endodermis of soybean nodule vascular bundle), although within the range of rates reported in the literature for movement through pd. However, several of the assumptions made in this calculation deserve attention. If sucrose flux occurred through all of the fibre sheath, rather than just the phloem shoulder area, this estimate would be approximately an order of magnitude high. Plasmodesmatal radius was assessed of the narrow neck present at either end (adjoining cell membrane), rather than the dilated internal region, which may also serve to contribute to an overestimation. Conversely, the calculated rate is based on sucrose accumulation over a 1-month period. This crude measure ignores sucrose metabolised within the stem (through respiration, biosynthesis of cell wall components and other processes), and as such, may be an underestimate. In contrast, the measure of Brown et al. (1995) was based on instantaneous measures of respiration and fixation rates (within the highly metabolic legume nodule). The plasmodesmata observed in cane material were characterised by a narrow neck at each end, which may represent a control point for flux. Viral movement proteins act to dilate plasmodesmata and are suggested to be homologues of a class of plant proteins that could regulate flux (Lucas et al. 1993). However, the flux calculation suggests that the ‘plumbing connection’ as represented by the symplastic pathway is not rate limiting to the storage process. Essentially storage sinks have a relatively low rate of flux, relative to metabolic sinks (e.g. C flux into soybean nodule estimated to be 1167 µmol g−1 dry weight h−1 , cf. 3.2 here, Table 1, item 1). The observation of lignification of storage parenchyma cell walls with internode maturation, with the exception of certain cells, and the presence of fluid in the apoplast of this tissue is in agreement with Welbaum et al. (1992), who suggested that the non-lignified cells might serve in the transport of sucrose between the symplast and apoplast. The presence of fluid in the apoplast is consistent with the unloading of sugar to the apoplast. The control of the storage process is ill defined, but given a symplastic path of unloading, two approaches to increased sucrose storage have merit. The incorporation of a viral movement protein into tobacco altered carbohydrate partitioning (Lucas et al. 1993), and this strategy could be applied to sugarcane. Alternatively, Patrick (1990) has proposed that the sink cell regulates its turgor, and thus the pressure gradient driving phloem import, by controlling unloading to the apoplast. Indeed, Moore and Cosgrove (1991) documented constant turgor in cane storage cells during sugar accumulation. Partitioning to storage may thus be improved by increasing sucrose unloading to the parenchyma apoplast by elevating the levels of transport proteins on the ‘transport’ parenchyma cells in the


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storage parenchyma tissue (e.g. Riesmeier et al. 1994, reported antisense expression of a sucrose carrier gene in potato leaves). Acknowledgments Funding support from SRDC (grant UCQ 1S) and CQU, and the input of Dr V Shepherd with light microscopy, and Dr S Stowe, RSBS EM Unit, ANU with freeze–fracture work is gratefully acknowledged. We thank J Wilson, now retired, from CSIRO, for editorial input. An abridged version of this study was presented at the SUGAR2000 Symposium (Brisbane, 1996), and published within the symposium proceedings, ‘Sugarcane: research towards efficient and sustainable production’ (Eds JR Wilson, DM Hogarth, JA Campbell, A Garside). pp. 105–107. (CSIRO Division of Tropical Crops and Pastures). References Brundrett MC, Enstone DE, Petersen CA (1988) A berberine–aniline blue fluorescent staining procedure for suberin, lignin and callose in plant tissue. Protoplasma 146, 133–142. Brown SM, Oparka KJ, Sprent JI, Walsh KB (1995) Symplastic transport in soybean root nodules. Soil Biology and Biochemistry 27, 387–399. doi: 10.1016/0038-0717(95)98609-R Gunning BES, Hughes JE (1976) Quantitative assessment of symplastic transport of prenectar in the trichomes of Abutilon nectaries. Australian Journal of Plant Physiology 3, 619–637. Hawker JS, Hatch MD (1965) Mechanism of sugar storage by mature stem tissue of sugarcane. Physiologia Plantarum 18, 444–453. Hayes PM, Offler CE, Patrick JW (1985) Cellular structures, plasma membrane surface areas and plasmodesmatal frequencies of the stems of Phaseolus vulgaris L. in relation to radial photosynthate transfer. Annals of Botany 56, 125–138. Jacobsen KR, Fisher DG, Maretzi A, Moore PH (1992) Developmental changes in the anatomy of the sugarcane stem in relation to phloem unloading and sucrose storage. Botanica Acta 105, 70–80. Kuo J, O’Brien TP, Canny MJ (1974) Pit-field distribution, plasmodesmatal frequency, and assimilate flux in the mestome sheath cells of wheat leaves. Planta 121, 97–118. doi: 10.1007/BF00388750 Lucas W, Olesinski A, Hull RJ, Haudenshield JS, Deom CM, Beachy RN, Wolf S (1993) Influence of the tobacco mosaic virus 30 kDa movement protein on carbon metabolism and photosynthate partitioning in transgenic tobacco plants. Planta 190, 88–96. doi: 10.1007/BF00195679 Ma H, Albert H, Paull R, Moore P (2000) Metabolic engineering of invertase activities in different subcellular compartments affects sucrose accumulation in sugarcane cells. Australian Journal of Plant Physiology 21, 1021–1030. Moore PH, Cosgrove DJ (1991) Developmental changes in cell and tissue water relations parameters in storage parenchyma of sugarcane. Plant Physiology 96, 794–801. Patrick JW (1990) Sieve element unloading: cellular pathway, mechanism and control. Physiologia Plantarum 78, 298–308. doi: 10.1034/j.1399-3054.1990.780220.x

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Riesmeier JW, Willmitzer L, Frommer WB (1994) Evidence for an essential role of the sucrose transporter in phloem loading and assimilate partitioning. EMBO Journal 13, 1–7. Shaw SR, Stowe S (1982) Freeze–fracture evidence for gap junctions connecting the axon terminals of Dipteran photoreceptors. Journal of Cell Science 53, 115–141. Welbaum GE, Meinzer FC, Grayson RL, Thornham KT (1992) Evidence for and consequences of a barrier to solute diffusion between the apoplast and vascular bundles in sugarcane stalk tissue. Australian Journal of Plant Physiology 19, 611–623.

Wilson JR (1990) Influence of plant anatomy on digestion and fibre breakdown. In ‘Microbial and plant opportunities to improve the utilization of lignocellulose by ruminants’. (Eds DE Akin, LG Ljungdahl, JR Wilson, PJ Harris). pp. 99–117. (Elsevier: New York) Wilson JR (1992) Improvement of yield in sugarcane through increased sucrose accumulation. Workshop Report, CSIRO, DTCP, Brisbane. Yem EW, Willis AJ (1954) The estimation of carbohydrates in plant extracts by anthrone. The Biochemical Journal 57, 508–514.

Manuscript received 2 June 2004, received in revised form and accepted 9 March 2005

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