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CSIRO PUBLISHING Functional Plant Biology, 2007, 34, 314–331 www.publish.csiro.au/journals/fpb Review: Nutrient loading of developing seeds Wen-Hao ZhangA , Yuchan ZhouB,E , Katherine E. DibleyB,E , Stephen D. TyermanC , Robert T. FurbankD and John W. PatrickB,F A Key Laboratory of Photosynthesis and Environmental Molecular Physiology, Institute of Botany, The Chinese Academy of Sciences, Beijing 100093, China. B School of Environmental and Life Sciences, The University of Newcastle, Callaghan, NSW 2238, Australia. C School of Agriculture, Food and Wine, Adelaide University, Waite Campus, PMB #1, Glen Osmond, SA 5064, Australia. D CSIRO Plant Industry, GPO Box 1600, Canberra, ACT 2601, Australia. E These authors contributed equally to this work. F Corresponding author. Email: [email protected] This paper originates from an International Symposium in Memory of Vincent R. Franceschi, Washington State University, Pullman, Washington, USA, June 2006. Abstract. Interest in nutrient loading of seeds is fuelled by its central importance to plant reproductive success and human nutrition. Rates of nutrient loading, imported through the phloem, are regulated by transport and transfer processes located in sources (leaves, stems, reproductive structures), phloem pathway and seed sinks. During the early phases of seed development, most control is likely to be imposed by a low conductive pathway of differentiating phloem cells serving developing seeds. Following the onset of storage product accumulation by seeds, and, depending on nutrient species, dominance of path control gives way to regulation by processes located in sources (nitrogen, sulfur, minor minerals), phloem path (transition elements) or seed sinks (sugars and major mineral elements, such as potassium). Nutrients and accompanying water are imported into maternal seed tissues and unloaded from the conducting sieve elements into an extensive post-phloem symplasmic domain. Nutrients are released from this symplasmic domain into the seed apoplasm by poorly understood membrane transport mechanisms. As seed development progresses, increasing volumes of imported phloem water are recycled back to the parent plant by process(es) yet to be discovered. However, aquaporins concentrated in vascular and surrounding parenchyma cells of legume seed coats could provide a gated pathway of water movement in these tissues. Filial cells, abutting the maternal tissues, take up nutrients from the seed apoplasm by membrane proteins that include sucrose and amino acid/H+ symporters functioning in parallel with non-selective cation channels. Filial demand for nutrients, that comprise the major osmotic species, is integrated with their release and phloem import by a turgorhomeostat mechanism located in maternal seed tissues. It is speculated that turgors of maternal unloading cells are sensed by the cytoskeleton and transduced by calcium signalling cascades. Additional keywords: membrane transport, nutrients, phloem transport, remobilisation, seeds, symplasmic transport. Introduction Seeds are heterotrophic organs, totally dependent on nutrients imported (nutrient loading) from the parent plant for their growth and development. Nutrient loading of seeds influences seed number at seed set and determines their final size, which are properties of key biological and agronomic significance. In terms of reproductive success, an evolutionary trade-off exists between seed number and size (Fenner 2005). Prolific production of small seeds increases the probability of some progeny dispersing into favourable microhabitats. Large seed size, modulated by parental imprinting (for a review, see Gehring et al. 2004), confers stress tolerance and competitive ability of the resulting seedling (Fenner 2005). For agronomic purposes, final crop yield is a function of average seed size summed © CSIRO 2007 over total seed number. In this context, the central importance of seeds for human consumption is illustrated by the 2005 world harvest of seeds. This comprised 2 billion metric tons of cereal grains, 146 million metric tons of oil seeds and nuts and 62 million metric tons of pulse seeds (see http://faostat.fao.org, accessed 14 February 2007). In addition, developing seeds of cereals and pulses offer tractable experimental models to study physiological contexts in which high nutrient fluxes are exchanged between two disjunct but proximal symplasmic compartments (Patrick and Offler 1995). Examples of these situations include apoplasmic phloem loading (Lalonde et al. 2003), phloem unloading in sinks that accumulate soluble sugars to high concentrations and biotrophic relationships (Patrick 1997). Together, these imperatives have driven an 10.1071/FP06271 1445-4408/07/040314 Nutrient loading of developing seeds interest to discover the processes contributing to nutrient loading of developing seeds and, particularly those of agronomic significance such as cereals and pulses. Owing to a number of functional genomic opportunities offered by Arabidopsis, there is an increasing focus on nutrient loading in seeds of this species despite technical challenges posed by their small size. Nutrient loading of seeds is a spatially and temporally dynamic process. The latter is inextricably linked with their development (for a recent review of seed development see Weber et al. 2005). Of relevance to nutrient loading, seeds progress through three main stages of development. Immediately following setting, seed development is dominated by cell division before a storage phase when most nutrients are loaded as their cells expand. For a period, cell division, cell expansion and storage overlap temporally as sequential waves of these phenomena progress through the tissues of developing seeds in distinctive spatio-termporal patterns (Jenner et al. 1991; Weber et al. 2005). Nutrient loading rates per seed reach a plateau coincident with attaining final seed volume. Thereafter, loading rates are constant during the remainder of storage product accumulation before declining precipitously as seed maturity is approached. During seed development, dominant sinks for nutrient loading shift from maternal tissues early in development, to filial tissues during later stages of development. However, exceptions to this generalisation exist, as illustrated by predominant allocation of biomass into maternal fibre cells of cotton seeds (Ruan 2005). During the storage phase, nutrients are partitioned between endosperm and embryo. In non-endospermic seeds of dicots the endosperm functions as a nutrient source for the developing embryo during the prestorage phase of seed development and, by the onset of the storage phase, is completely depleted (Weber et al. 2005). By contrast, endospermic seeds harbour a diminutive embryo and the endosperm is the major sink for nutrient loading throughout seed development. This developmental pattern is characteristic of all cereal grains (e.g. Jenner et al. 1991). Nutrient exchange between maternal, endosperm and embryo compartments of developing seeds has directed attention to membrane transport events and how these interface with nutrient import by, and nutrient metabolism/compartmentation within, developing seeds. These issues have been addressed in past reviews of nutrient loading of developing seeds with a particular focus on sugar transport (see Thorne 1985; Wolswinkel 1992; Weber et al. 1997b; Patrick and Offler 1995, 2001). The present review builds on this information base by drawing on more recent findings that contain discoveries at the molecular level. In addition, there is a growing body of research output contributing to the understanding of mineral element transport to and within developing seeds. Including these aspects, we review nutrient loading of seeds in a developmental context. Informed by the principles of phloem transport, our analysis begins by exploring roles played by the source/path/sink system in regulating nutrient fluxes to developing seeds. Regulation along the source/sink/path sink system – lessons from phloem translocation Most nutrients, except calcium, are imported into seeds through the phloem (Patrick and Offler 2001). The general consensus Functional Plant Biology 315 is that phloem translocation occurs by bulk flow. Hydrostatic pressure differences established along the phloem path drive bulk flow from source leaves to developing seeds. Phloem path pressure differences result from loading of osmotically-active nutrients into sieve tubes within source leaves and unloading of these solutes from sieve tubes located in developing seeds. Thus, nutrient import rates (Rf ) into developing seeds can be predicted from the relationship: Rf = Lp (Psource − Pseed ) × A × C, (1) where Lp is path hydraulic conductivity, P is hydrostatic pressure at source and seed ends of the phloem path, A is path crosssectional area and C is the concentration of transported nutrient. Hydrostatic pressures developed at source and seed ends of phloem paths are a function of phloem sap osmolality. Equation 1 defines how source, phloem path and seed sink properties contribute to phloem import rates of a nutrient species into developing seeds. Hydrostatic pressure differences along the path are regulated by phloem loading of the major osmotic species (sucrose and potassium, Patrick and Offler 2001) in source leaves and their unloading from the phloem in seed sinks. Phloem loading in source leaves and along the axial path (re-mobilisation from storage reserves; xylem–phloem transfer) sets the phloem sap concentration of each transported nutrient. In addition, path properties determine hydraulic conductivity (sieve pore dimensions) and cross-sectional area (number of sieve elements in path cross section). Thus, import rates of major sap osmotic species may be influenced by source, path and seed processes. In contrast, variation in import rates of minor osmotic species relies solely on their phloem sap concentrations. This is determined by rates of phloem loading in source leaves and along the transport path. Unloading of minor osmotic species in seed sinks will have little influence on depressing hydrostatic pressures of importing sieve elements. Key controls of nutrient import by seeds alter during their development Manipulating production of, and inter-sink competition for, photoassimilates (sucrose) has been used to assess whether seed development is carbon supply- (i.e. source, path) or sink-limited (e.g. Borrás et al. 2004). These studies show that the pre-storage phase of seed development is acutely sensitive to alterations in carbon supply. Source photosynthesis is unlikely to impose this limitation as the relative temporal shifts in photosynthetic rates do not quantitatively match the explosive increase in dry matter import by seeds once seed filling commences (Evans and Wardlaw 1996; Pate and Armstrong 1996). This observation points to a contribution of photoassimilate partitioning between developing seeds and their competing sinks. Under conditions of supply-limited growth, partitioning can be under path control (see Eqn 1). At the pre-storage phase of seed development, photoassimilates are delivered symplasmically in provascular strands over distances of up to 1–2 mm (Esau 1965). The hydraulic conductivity and cross-sectional area of these strands are predicted to be considerably less than those of the interconnecting sieve elements and, as such, would dominate path conductance (Eqn 1). Procambial stand conductance will be determined mainly by diameters of their interconnecting 316 W.-H. Zhang et al. Functional Plant Biology plasmodesmal canals. These are dynamic structures that are highly regulated in ways that are just beginning to be understood (Roberts and Oparka 2003). In contrast, during the subsequent storage phase of seed development alterations in leaf photosynthetic rates exert minimal impact on seed biomass gain leading to the conclusion that seed growth is predominantly sink-limited (Borrás et al. 2004). Additionally, conductive capacity of the vascular system (Eqn 1) does not impose any limitation on carbon supply (Evans and Wardlaw 1996). Together these observations point to regulation mediated by phloem turgor in seeds (Pseed , Eqn 1). In this context, turgor pressures of developing seeds are strikingly independent of changes in water relations of the parent plant and are regulated by local factors influencing seed water potentials (Fisher and Cash-Clark 2000b; Shackel and Turner 2000). Water in protoplasmic and apoplasmic compartments in developing seeds appears to be in quasi equilibrium (Jenner and Jones 1990). Hence, water potential (ψ) of sieve element sap approximates that of their surrounding apoplasm (a) such that: PSE − !SE = Pa − !a , (2) PSE = (!SE − !a ) + Pa , (3) and, as a consequence, where ! is sap osmotic pressure and P is hydrostatic pressure. The minimal evaporative loss of water from seeds will cause Pa to be relatively small and negative. The small negative hydrostatic pressures are conjectured to be generated by tensions transmitted in the xylem water columns connecting the transpiring parent plant with their magnitudes attenuated by the low hydraulic conductivities of intervening xylem discontinuities. As a consequence, sap osmotic pressures are likely to be the major factors influencing PSE (Eqn 3). Thus differences between !SE and !a , and hence PSE (Eqn 3), will be governed by phloem unloading rates of the major osmotic nutrients. Sources of phloem imported nutrients Carbon Proximal leaves are the principal sources of current photosynthetically-reduced carbon imported by developing seeds, with lesser contributions from photosynthesis of reproductive structures (Evans and Wardlaw 1996; Pate and Armstrong 1996). As the proximal leaves senesce, carbon is remobilised from storage sites in leaf sheaths, stems and reproductive structures (Evans and Wardlaw 1996; Pate and Armstrong 1996). Further, for some cereal crops, remobilisation of stored carbohydrates may contribute substantially to final grain yield (Schnyder 1993), especially under drought conditions (Yang and Zhang 2006). Transition from polysaccharide synthesis to remobilisation is accompanied by changes in expression and activity profiles of carbohydrate metabolising enzymes (Takahashi et al. 2005). Intercellular transport of sucrose between sieve tubes and storage cells could also change. For instance, in wheat internodes, membrane-impermeant fluorescent dyes move symplasmically from the phloem to storage parenchyma cells during carbohydrate accumulation (Aoki et al. 2004). At the onset of carbohydrate remobilisation in rice leaf-sheaths, OsSUT1 expression increases (Hirose et al. 1999) suggesting that this sucrose transporter may function in apoplasmic phloem loading during the remobilisation phase. Amino nitrogen and sulfur In contrast with carbon, nitrogen import into developing seeds is largely derived from re-mobilisation of amino acids sourced as proteolytic products of proteins synthesised before the onset of reproductive development (Jenner et al. 1991; Staswick 1994; Tilsner et al. 2005). For monocarpic species, current assimilation from the soil solution contributes minimally to seed nitrogen levels as soil pools proximal to the root surfaces are depleted and root growth slows commensurate with seed fill (Jenner et al. 1991; Schiltz et al. 2005). Leaves are the principal source of amino acids destined for seed import, with remobilisation from stems and reproductive structures making a lesser contribution (Good et al. 2004; Schiltz et al. 2005). Rubisco accounts for 50% of the total protein content of leaves, and, together with photosystem proteins, is a major source of nitrogen for remobilisation (Schiltz et al. 2004). Glutamine synthetase may play a key role in rendering nitrogen available for remobilisation from senescing leaves (Good et al. 2004; Tabuchi et al. 2005). In certain species, and in particular grain legumes, an additional source of remobilisable proteins are two glycoproteins that accumulate to high levels in leaves, stems and pod walls; the so-called vegetative storage proteins (VSP) (Staswick 1994). In leaves, VSP primarily accumulates in vacuoles of a specialised layer of mesophyll cells, the paraveinal mesophyll, located between the palisade and spongy mesophyll (Klauer et al. 1996). During seed fill, VSP is hydrolysed and the resulting amino acid products are moved symplasmically to the vascular bundles for phloem loading and export to developing seeds (Franceschi et al. 1983; Lansing and Franceschi 2000). However, VSP appears to play a supplementary, rather than primary role, in nitrogen storage for seed filling (Staswick et al. 2001). In other plant species, excess amino acids are stored in vacuoles of importing leaf cells before remobilisation during seed development (Tilsner et al. 2005). Total amino acid concentrations of phloem saps fall in the range of 50–200 mM for most plant species, except Brassica spp. with phloem saps containing up to 650 mM (Lohaus and Moellers 2000; Tilsner et al. 2005). However, the contribution of amino acids to sap osmolality is surprisingly no more than 20% (Lohaus and Moellers 2000) and, hence, only exerts a modest impact on hydrostatic pressure differences driving phloem translocation (see Eqn 1). This conclusion predicts that nitrogen loading of developing seeds is likely to be regulated by phloem sap concentrations of amino acids; a prediction verified for Brassica species (Lohaus and Moellers 2000; Tilsner et al. 2005). Phloem sap concentrations are a function of phloem loading rates, which, for apoplasmic phloem loaders, depend on activities of amino acid transporters located in sieve element/companion cell complexes (cf. Tilsner et al. 2005). Sulfur is delivered to seeds through the phloem as sulfate (grain legumes) or in reduced forms such as glutathione (rice), S-methylmethionine (wheat, Tabe et al. 2002). Sulfur-reducing enzymes are present in developing seeds and account for all the reduced sulfur accumulated in mature seeds of grain legumes (Sexton and Shibles 1999; Tabe and Droux 2001, 2002) and Nutrient loading of developing seeds wheat (Fitzgerald et al. 2001). In contrast, rice appears to rely on imported amino sulfur (Hagan et al. 2003). Sulfate retrieval by phloem-localised sulfate transporters contributes significantly to seed import (Awazuhara et al. 2005). Major mineral salts and micronutrients Similar to carbon and amino nitrogen, reliance on an external supply to the fruit/inflorescence is the principal source of minerals accumulated by the filial generations of grain legume (Hocking and Pate 1977) and cereal (Pearson et al. 1998) seeds. Import of transition elements, such as iron and zinc, is of particular interest to re-dress their deficiencies in seeds used for human consumption (Marentes and Grusak 1998) and is the focus of the discussion presented below. Relative delivery of minerals to developing seeds, not surprisingly, reflects their phloem mobility (Hocking and Pate 1977; Pearson et al. 1998). Transition metal ions precipitate at alkaline pHs characteristic of phloem saps. This points to a requirement for chelation to ensure their phloem transport. Nicotianamine (NA) is a mineral chelating candidate that is ubiquitous throughout higher plants. This non-protein amino acid is a strong chelator of transition elements and the resulting complexes are very stable at alkaline pH (von Wiren et al. 1999). The role of NA in mobilisation of Fe to seeds has been shown using a NA-deficient tobacco. Here a barley nicotianamine amino transferase (NAAT) enzyme was overexpressed to deplete the NA pool (Takahashi et al. 2003). Seeds of naat transformants exhibited depressed Fe levels and this phenotype could be rescued by exogenous applications of NA to leaves. Significantly, one of three rice NA synthetase genes (OsNAS3) is expressed selectively in phloem and bundle sheath cells of rice (Inoue et al. 2003). This provides a source of NA to chelate transition metals before their entry into the phloem translocation stream. A maize mutant, that exhibits interveinal chlorosis (yellow stripe, ys), led to the discovery of a membrane transporter, ZmYS1. The ZmYS1 transporter, along with an Arabidopsis homologue (AtYSL2), has been shown to be capable of transporting Fe-NA complexes (Curie et al. 2001). Single (ysl1, Le Jean et al. 2005) and double (ysl1ysl3, Waters et al. 2006) knockout mutants of Arabidopsis exhibit reduced import of transition metals by seeds. Moreover, Fe import could be restored by expression of the wild type AtYS1 gene in the ysl1 mutant background (Le Jean et al. 2005). Together, these findings suggest that YSL transporters function in loading of NA-transition metal complexes into leaf phloem for seed import. Pathways of nutrient and water flows in maternal seed structures Phloem entry and post-phloem symplasmic domains Differentiated vascular systems are restricted to maternal seed tissues and are primarily comprised of phloem with no or limited xylem (Patrick and Offler 1995). In addition, these vary from highly branched networks of veins (e.g. Glycine, Phaseolus spp.) to truncated vascular bundles terminating at the pedicel/funicle junction (e.g. Zea mays, Patrick and Offler 1995; Arabidopsis, Stadler et al. 2005). However, seed vascular systems do not appear to constrain rates of nutrient import as their extensiveness are unrelated to final seed size (Patrick and Offler 1995). Functional Plant Biology 317 Circumstantial evidence suggests that phloem-imported nutrients, and phloem water, are unloaded from sieve elements through symplasmic routes (Patrick and Offler 2001; Fig. 1). Plasmodesmata, interconnecting sieve elements with adjacent vascular parenchyma, exhibit unusually large size exclusion limits in wheat grains (Fisher and Cash-Clark 2000a) and Arabidopsis seeds (Stadler et al. 2005). Their diametres, possibly as large as 42 nm (Fisher and Cash-Clark 2000a), may confer hydraulic conductivities of sufficient magnitude to permit sieve element unloading by bulk flow driven down large hydrostatic pressure gradients operating across this interface (∼1 MPa, Fisher and Cash-Clark 2000b). Bulk flow is the simplest mechanism to ensure homeostasis of hydrostatic pressures in the importing sieve tubes by coupling water and nutrient unloading rates (cf. Murphy 1989). Subsequent symplasmic passage through plasmodesmata of adjacent ground tissues, with smaller size exclusion limits (Fisher and CashClark 2000a; Stadler et al. 2005), is likely to be dominated by diffusion deduced from measures of turgor and concentration gradients (Fisher and Cash-Clark 2000a, 2000b). Irrespective of transport mechanism, current evidence suggests that control of the symplasmic nutrient flux is mediated by plasmodesmal conductivities at sieve element boundaries (Fisher and CashClark 2000a, 2000b; Thomas et al. 2000). In addition, the extensiveness of post-phloem symplasmic domains varies between species (Patrick and Offler 2001) and during seed development (Stadler et al. 2005). For large seeds, symplasmic movement must rely on specialised transport mechanisms to sustain the observed high rates of nutrient transport over considerable distances (cm) through the post-phloem pathway (Patrick and Offler 2001). The nature of the proposed transport mechanisms are unknown and would merit investigation. For instance, plasmodesmal gating (Ruan et al. 2001) could contribute to developmental shifts in post-phloem symplasmic domains. In relation to nutrient delivery function, post-phloem domains abut (e.g. cereals, Vicia, Pisum, Fisher and Cash-Clark 2000a; Patrick and Offler 2001; van Dongen et al. 2003), or are separated from (e.g. Arabidopsis, Stadler et al. 2005; cotton, Ruan et al. 2001; Phaseolus, Patrick et al. 1995), the underlying filial tissues (Fig. 1). For the latter cellular organisation, it is unresolved whether further inward radial movement of nutrients to the filial interface follows an apoplasmic (cf. Wang et al. 1995) or additional symplasmic routes arranged in series with apoplasmic steps (Stadler et al. 2005). Cellular site(s) of exchange to the seed apoplasm Identifying cells responsible for nutrient release from maternal tissues of developing seeds has relied largely on indirect evidence. This evidence includes detecting cell morphologies specialised for membrane transport (i.e. transfer cells) and apoplasmic barriers that isolate putative unloading cells (e.g. cuticle layer in barley and wheat grains), estimating cell/tissue type capacities for membrane transport based on their membrane surface areas (Patrick and Offler 1995) and localisation of membrane proteins considered to mediate membrane transport to the seed apoplasm (Patrick and Offler 2001). With reference to the latter approach, most investigations have assumed that a component of sucrose release is secondary active and coupled 318 W.-H. Zhang et al. Functional Plant Biology Filial tissue Cotyledon Seed apoplasm Maternal tissue Seed coat Actin [Ca2+]c ATP V ATP IP3 [Ca2+] H+ AA c H+ Hex Actin PM PM Vascular phloem Efflux cell Apoplasm Influx cell Fig. 1. Schematic diagram showing membrane transporters involved in transferring phloem-imported nutrients from seed coats to cotyledons of developing grain legume seeds. Phloem-imported nutrients are transported through plasmodesmata (PD) to the seed coat efflux cells. Here, membrane transporters in plasma membranes (PM) lining efflux cells release nutrients to the seed apoplasm. Currently known transporters are: (1) non-selective channel; (2) sucrose/H+ antiporters; (3) H+ -ATPase; (4) sucrose facilitators; (5) aquaporins; (6) sucrose/H+ symporters; (7) pulsing Cl− channel. Nutrients are taken up from the seed apoplasm by membrane transporters located in plasma membranes of cotyledon dermal cell-complexes. Currently known transporters are: (8) non-selective cation channel; (9) sucrose/H+ symporters; (10) H+ -ATPase; (11) amino acid/H+ symporters and (12) hexose/H+ symporters. An elevated cell turgor (dart), due to an enhanced uptake of nutrients from the seed apoplasm, activates Cl− and non-selective channels by interacting with the actin cytoskeleton. The elevated turgor signal may also activate IP3 -dependent Ca2+ release, leading to an increase in the cytosolic Ca2+ activity [Ca2+ ]c , which serves as signals to activate sucrose/H+ antiporters and Cl− channels. with H+ -ATPase activities. For example, strong proton pump activities have been detected in plasma membranes of ground parenchyma cells proximal to vascular bundles and thin-wall parenchyma transfer cells of Phaseolus and Vicia seed coats, respectively (Wang et al. 1995). That thin-walled parenchyma transfer cells of Vicia seed coats are enriched in transporter proteins was verified by histochemical detection of high densities of H+ -ATPases, sucrose binding proteins (Harrington et al. 1997a) and an amino acid permease, VfAAP1 (Miranda et al. 2001). Similar studies have established that sucrose transporter proteins and active H+ -ATPases are enriched in nucellar tissues of developing grains of barley (Weschke et al. 2000), wheat (Bagnall et al. 2000) and rice (Furbank et al. 2001). However, these studies cannot exclude the possibility of unloading from all cells located in post-phloem symplasmic domains (cf. van Dongen et al. 2003). In this context, cells located along the postphloem symplasmic pathway may function in retrieval of sucrose (Ritchie et al. 2003; VfSUT1, Weber et al. 1997a; PsSUT1, Tegeder et al. 1999) and amino acids (PsAAP1, Tegeder et al. 2000a; VfAAP3, Miranda et al. 2001) leaked to the seed apoplasm while in transit to the principal site(s) of release. Unequivocal identification of the unloading cells ultimately depends on cloning transporter proteins responsible for effluxing the various nutrient species. These products then can be used in immunolocalisation and protein expression studies to detect these cells. The encoding genes for these membrane proteins are yet to be discovered. Nutrient efflux mechanisms from maternal tissues – pores, channels and carriers Sucrose and amino acids Mechanisms of sucrose release could well alter during seed development. Early in development, imported sucrose released from maternal tissues is cleaved by an extracellular invertase in faba bean (Weber et al. 1997a), rice (Hirose et al. 2002), barley (Weschke et al. 2003) and maize (Chourey et al. 2006). This is conjectured to result in steepened transmembrane gradients of sucrose concentration to drive passive release of sucrose to the seed apoplasm. Storage product accumulation coincides with a marked upward shift in the sucrose/hexose composition of the seed apoplasm (Patrick and Offler 2001; Weschke et al. 2003). Nutrient loading of developing seeds Functional Plant Biology The latter is caused by a sharp decline in extracellular invertase activity (Weber et al. 1997a; Weschke et al. 2003). Extracellular invertase activity is likely to be regulated by a transcription factor (AP2) that causes changes in the hexose/sucrose ratio and seed size in Arabidopsis (Ohto et al. 2005). Elements of the phenomenon have been detected in non-endospermic dicot seeds such as common plantain (Gahrtz et al. 1996), canola (King et al. 1997), cotton (Ruan et al. 2001) and Arabidopsis (Ruuska et al. 2002). However, for all tropical grasses (Patrick and Offler 1995; Chourey et al. 2006) and endospermic dicot seeds that store oil (e.g. tobacco, Tomlinson et al. 2004), extracellular invertase activity is retained throughout seed filling. Most studies of nutrient release have focused on the storage phase of grain legume and wheat seed development. Early models for sucrose unloading envisaged a pump leak system whereby passively released sucrose is taken up by sucrose symport that functions to regulate net rates of sucrose release. Such a model is consistent with observed expression of sucrose symporter genes in maternal tissues of developing seeds of both monocots (Bagnall et al. 2000; Weschke et al. 2000; Furbank et al. 2001) and dicots (Weber et al. 1997a; Tegeder et al. 1999; Meyer et al. 2004; Zhou et al. 2007). However, there are several observations that do not support a pump/leak model of sucrose release. First, estimates of sucrose release by simple diffusion at best only account for 25% of the observed sucrose flux (Table 1). Second, sucrose symport activity is not detectable in maternal tissues of developing wheat grains (Wang and Fisher 1995; Bagnall et al. 2000) and only marginally so in seed coats of pea (de Jong et al. 1996; de Jong and Borstlap 2000) and of two bean species at low (<10 mM) external sucrose concentrations (Ritchie et al. 2003). Depending on the proton motive force (PMF) and the opposing transmembrane sucrose concentration difference, it has been shown that sucrose/H+ symporters can function in an efflux mode (Carpaneto et al. 2005). Assuming a sucrose/H+ stoichiometry of one, the following derivative of the Nernst equation predicts the intracellular (Ci ) and external (Co ) sucrose concentration differences at which sucrose/H+ symport will Table 1. Comparison of predicted fluxes of sucrose release by simple diffusion with those observed to occur across plasma membranes of cells considered to release nutrients from maternal tissues of developing seeds of wheat, French and broad bean Predicted flux by simple diffusion was computed as the product of membrane permeability (10−10 m s−1 , Cram 1984) and observed transmembrane differences in sucrose concentration Plant species Transmembrane concentration difference (mM) Predicted flux (10−8 mol m−2 s−1 ) Observed flux (10−8 mol m−2 s−1 ) Wheat French bean Broad bean 50A 10C 40C 0.5 0.1 0.4 2.2B 8.8D 7.0E A Fisher and Wang (1995). et al. (1995). C Patrick (1994). D Offler and Patrick (1984). E Offler and Patrick (1993). B Wang 319 reverse for a given proton motive force comprised of membrane potential ("ψ) and proton ("pH) differences: Log10 Ci = Log10 Co − ["ψ +59 mV"pH]/59 mV. (4) Membrane potentials of seed coats range from −40 to −50 mV (Walker et al. 1995; van Dongen et al. 2001), with proton differences of one pH unit (Walker et al. 1995). These generate an inward directed PMF of −100 mV for the lesser membrane potential. Estimates of sucrose concentrations in seed apoplasmic spaces of grain legumes range from 5 to 200 mM (Patrick and Offler 2001), with bean seed coats at 80 mM sucrose. For a PMF of −100 mV, Eqn 4 predicts sucrose/H+ symport will reverse to an efflux mode at intracellular sucrose concentrations of 251 or 3981 mM when apoplasmic sucrose concentrations are 5 or 80 mM, respectively. These predicted intracellular sucrose concentrations exceed estimates of 100–120 mM for seed coat tissues (Patrick and Offler 2001). Therefore, under these conditions, sucrose symporters are likely to function in sucrose retrieval modes in non-vascular cells of seed coats (cf. Ritchie et al. 2003). However, assuming a phloem sucrose concentration of 500 mM, sucrose symporters could function as effluxers from sieve elements/vascular parenchyma for apoplasmic concentrations up to 10 mM (predicted from Eqn 4). Owing to the relatively small membrane surface areas of these cells, symporters transporting at maximal rates might account for 10% of the released sucrose (Patrick and Offler 2001). Thus, sucrose/H+ symporters in seed coats may play a minor role in modulating rates of net sucrose release from seed coats and, under depleted apoplasmic sucrose concentrations, could contribute to efflux from sieve elements/vascular parenchyma cells (Fig. 1). In developing pea seeds, release of sucrose (de Jong et al. 1996) and amino acids (de Jong et al. 1997) from their coats has been suggested to occur through non-selective pores (also see van Dongen et al. 2001). A carrier-mediated facilitated diffusion may account for sucrose release from maternal nucellar cells of developing wheat grains (Wang and Fisher 1995). In contrast, studies with whole seed coats of two bean species (Fieuw and Patrick 1993; Walker et al. 1995, 2000) and membrane vesicles derived from coat tissues (C. Niemietz, J. W. Patrick and S. D. Tyerman, unpublished data), have yielded evidence indicating that sucrose efflux has both passive and energised components. Energy coupling is likely to be secondary active through a sucrose/H+ antiport mechanism (Fieuw and Patrick 1993; Walker et al. 1995; C. Niemietz, unpublished data; Fig. 1). High densities of H+ -ATPases, localised to plasma membranes of seed coat cells putatively responsible for nutrient release (Wang et al. 1995; Harrington et al. 1997a), could function to generate the PMF driving sucrose/H+ antiport (Fig. 1). The molecular identities of non-selective pores (cf. Schuurmans et al. 2003) and sucrose/H+ antiporters remain to be identified. Predicted losses of sucrose by simple diffusion (Table 1) only account for 2–10% of the observed energy-independent and sulfhydryl reagent insensitive sucrose release from bean seed coats (Fieuw and Patrick 1993; Walker et al. 1995). Interestingly a low affinity and sulfhydryl modifier-independent facilitated transport of sucrose has been shown to function at high external concentrations (Ritchie et al. 2003). This transport behaviour is consistent with transport properties of three sucrose facilitators 320 W.-H. Zhang et al. Functional Plant Biology (SUFs) recently cloned from coats of developing pea and French bean seeds (Zhou et al. 2007; Fig. 1). The SUFs exhibited high apparent Km values (30–100 mM sucrose) and transport activities that were independent of sulfhydryl reagents when expressed in yeast cells. Modest outward-directed gradients of sucrose concentrations across the plasma membranes of seedcoat unloading cells (Table 1) could drive energy-independent sucrose efflux (Fieuw and Patrick 1993; Walker et al. 1995) through these SUFs. Evidence for facilitated diffusion, with comparable high apparent Km values (∼228 mM derived from fig. 5 of Wang and Fisher 1995) to those found for bean seed coats (∼400–500 mM sucrose , Ritchie et al. 2003) has been reported as a suggested mechanism for sucrose release from the nucellus projection of developing wheat grains (Wang and Fisher 1995). Minerals In addition to sucrose, large amounts of mineral nutrients, notably K+ and Cl− , are released from legumes seed coats to the seed apoplasm (e.g. Walker et al. 1995). To elucidate mechanisms of mineral nutrient release, protoplasts of seed coat cells of Phaseolus vulgaris L. (Zhang et al. 2002) and Vicia faba L. (Zhang et al. 1997), putatively responsible for nutrient release, were isolated and identified. Ionic channels in plasma membranes of these protoplasts were characterised using patchclamp techniques. Two types of non-selective channels and one pulsing anion channel have been identified in plasma membranes of ground parenchyma cells of P. vulgaris seed coats (Zhang et al. 2000, 2002, 2004a; Fig. 1). The non-selective channels were permeable to univalent cations including inorganic cations such + as K+ , Na+ , NH+ and large organic cations, TEA+ 4 , Cs + and choline (Zhang et al. 2000, 2002). The non-selective characteristics of the channels are unique among ionic channels characterised so far in plant cells (cf. Maathuis et al. 1997). The channels are distinguished by their activation kinetics with one activating ∼50 times faster than the other (Zhang et al. 2002). The fast-activating channel displays weak voltage-dependence and activates over a wide range of voltages (Zhang et al. 2002). By comparing the pharmacological profiles of the two types of channels with those of K+ efflux from intact seed coats, it was concluded that the fast-activating channel is likely to play a role in mediating K+ efflux from intact seed coats. This function is analogous to that of K+ -selective channels of xylem parenchyma cells that load xylem elements (Wegner and Raschke 1994; Roberts and Tester 1995; Wegner and de Boer 1997; Gaymard et al. 1998). The slowly-activating, nonselective channel exhibits a higher permeability to Ca2+ than the fast-activating channel (Zhang et al. 2002), implying that this channel may play a role in transduction of Ca2+ -dependent signal (and see further on). In addition, the permeability of the slowlyactivating, non-selective channel to Ca2+ may also provide a route for efflux of other divalent cations such as Mg2+ (e.g. Wolswinkel et al. 1992), Zn2+ and Mn2+ . Release of organic cations (choline+ , histidine+ ) from pea seed coats has been suggested to occur through non-selective membrane pores (van Dongen et al. 2001). The same nonselective pores may also account for efflux of neutral solutes such as sucrose, mannitol, glucose, valine and histidine (de Jong et al. 1996, 1997; van Dongen et al. 2001). The putative non-selective membrane pores in pea seed coats (van Dongen et al. 2001) and non-selective channels in P. vulgaris seed coats (Zhang et al. 2002) may share the same molecular identity. Furthermore, a component of the passive and energy-independent sucrose efflux from coats of P. vulgaris seeds may occur through the nonselective channels. This assertion is based on the observation that Gd3+ and La3+ block current flow through non-selective channels (Zhang et al. 2002) also inhibit K+ and sucrose efflux from P. vulgaris seeds (Walker et al. 2000). In addition to non-selective channels, a pulsing Cl− -permeable anion efflux channel recently has been characterised in developing P. vulgaris seed coat cells (Zhang et al. 2004a). The channel displayed a spontaneous activation and time-dependent inactivation, and was sensitive to La3+ and neomycin, but not Gd3+ (Zhang et al. 2004a). Similar pharmacological profiles were observed for Cl− efflux from excised seed coats (Zhang et al. 2004a), suggesting that the pulsing Cl− channel is likely to act as a route for Cl− release. Neomycin inhibition points to involvement of IP3 in channel activation, possibly through an IP3 -dependent intracellular Ca2+ release. In this context, a similar pulsing Cl− efflux channel, sensitive to neomycin and La3+ , has been characterised in Chara, in which the IP3 -dependent liberation of intracellular Ca2+ occurs as an all-or-none event (Wacke and Thiel 2001). The mechanism underlying the IP3 -dependent activation of Cl− channel in seed coat cells is unknown. Like the non-selective channels, the gene(s) encoding the Cl− channel remains to be cloned. Developing cotyledons of grain legumes are capable of assimilating sulfate into sulfur amino acids (Tabe and Droux 2001) presumably acquired from the seed apoplasm following release from seed coats. In this context, a voltage-dependent SO4 2− -permeable channel was identified in plasma membranes of protoplasts derived from coats of developing chickpea (Cicer arietinum L.) seeds. The channel was inhibited by ∼60% with anion channel blocker, niflumate. The same channel blocker reduced SO4 2− efflux from chickpeas seed coats from 4.8 ± 2.4 to 1.9 ± 2.4 µmol g−1 FW h−1 (W.-H. Zhang, unpublished data). Together, these observations suggest that this channel is likely to provide a low resistance route for efflux of the phloemimported SO4 2− from seed coats to the seed apoplasm. The SO4 2− -permeable channel in the chickpeas seed coats displayed comparable activation and deactivation kinetics to those characterised in Arabidopsis hypocotyl (Frachisse et al. 1999) and Arabidopsis root cells (Kiegle et al. 2000). A putative sulfate transporter gene (CaSultr3-1) is expressed in seed coats but not in developing embryos of chickpea (Tabe and Droux 2003). Whether CaSultr3-1 encodes a low affinity sulfate transporter underlying the observed sulfate-permeable channel remains to be verified. Water At later stages of seed development most of the water flow into the seed apoplasm is recirculated to the parental plant via the xylem (Pate et al. 1985). This occurs despite an apparent independence of seed water potential from that of the parent plant (Wang and Fisher 1994; Shackel and Turner 2000). Furthermore Nutrient loading of developing seeds the high apoplasmic solute concentration (300–400 mOsmol) (Patrick and Offler 2001) needs to be confined to the unloading site and not be advected to the xylem in the return pathway of water flow. There may be a high hydraulic resistance in the xylem (Jenner 1985) and retention of apoplasmic solutes by a semipermeable apoplasmic barrier (Bradford 1994). The putative barrier may allow some water movement and the hydraulic conductivity of the barrier may be variable, for example it may decrease under drought. In summary, in accounting for water flows: (1) water must be able to exit the symplasm at high rates to account for the mass flow over the available membrane surface area (Murphy 1989); (2) at the same time, water must flow back to the xylem which in some systems (e.g. bean) are relatively close to the phloem sieve elements; (3) high solute concentrations are retained in the apoplasm; and (4) the seed appears to be somewhat hydraulically isolated from the parent plant. A mechanism that can account for all these observations has yet to be proposed and experimentally tested. To develop a hypothetical mechanism one must understand the possibilities and the constraints. In this respect recent information about the regulation of aquaporins (water channels) provides some alternatives that have not been previously considered. Aquaporins are major routes for water flux across cell membranes (Tyerman et al. 2002). Plant aquaporins are divided into four subfamilies: plasma membrane intrinsic proteins (PIPs), tonoplast intrinsic proteins (TIPs), NOD26like membrane intrinsic proteins (MIPs) (NIP) and small basic intrinsic proteins (SIPs). The PIP subfamily is further separated into two groups PIP1 and PIP2 (Johanson et al. 2001). Functionally, MIPs falls into two groups: aquaporins and glycerol facilitators (Park and Saier 1996). Certain plant aquaporins could be permeable to ammonia (Niemietz and Tyerman 2000; Jahn et al. 2004), urea (Gerbeau et al. 1999), boron (Dordas et al. 2000; Ruiz, 2001), hydrogen peroxide (Henzler and Steudle 2000), small molecular weight alcohols (Henzler and Steudle 2000), carbon dioxide (Uehlein et al. 2003) and silicon (Ma et al. 2006). Aquaporins of the PIP family are gated (opened/closed, respectively) by phosphorylation/dephosphorylation (Johansson et al. 1998), alkaline/acid cytoplasmic pH (Tournaire-Roux et al. 2003) and raised/lowered cytoplasmic calcium (Alleva et al. 2006; Törnroth-Horsfield et al. 2006). The functional activity of some aquaporins may also be controlled by the formation of heterotetramers (Fetter et al. 2004). The density in the membrane is controlled by transcriptional regulation and targeting which has been shown to be diurnally regulated (Lopez et al. 2003). These general characteristics allow for various possibilities in control of water and solute flows in the seed coat. In previous discussions of water flow in seed coats it has generally been assumed that the hydraulic conductivity is invariant in time. However, aquaporin activity can change in response to a variety of signals and via several mechanisms summarised above. For instance, it is possible that water transfer to the xylem may not happen at exactly the same time that water unloading from the symplasm occurs. This could involve alternative opening and closing of aquaporins corresponding to phloem unloading and xylem recycling. Pate et al. (1985) showed diurnal patterns of xylem recycling, that they attributed to variations in transpirational demand and changes in phloem Functional Plant Biology 321 transport. Such time dependent regulation of aquaporins may also result in apparent disequilibrium of water potential between the seed and parent plant. Another possibility is that the aquaporins are sensitive to the osmotic potential in the apoplasm. It has been recently been shown that a membrane rich in a NIP aquaporin is exquisitely sensitive to osmotic potential (Vandeleur et al. 2005) and other membranes rich in aquaporins have shown water permeability that is sensitive to osmotic potential (Niemietz and Tyerman 1997; Ye et al. 2005). In the case of the NIP aquaporin, the water permeability is reduced with decreased osmotic potential from a very high level to almost zero over a range of 2.5 MPa (Vandeleur et al. 2005). A NIP aquaporin has been identified in pea seed coats (Schuurmans et al. 2003), but we have not been able to detect a NIP orthologue in bean seed coats (Y. Zhou, N. Setz, C. Niemietz, S. D. Tyerman and J. W. Patrick, unpublished data; Fig. 1). It is possible that PIP aquaporins, which have been located in seed coats of both pea (Schuurmans et al. 2003) and bean (Y. Zhou, N. Setz, C. Niemietz, S. D. Tyerman and J. W. Patrick, unpublished data), may also be sensitive to apoplasmic osmotic potential (Ye et al. 2005). If osmotic sensitivity were to occur with seed coat aquaporins, it could allow autoregulation of unloading and water recycling to the xylem as follows: (1) at high apoplasmic solute concentration there would be a closure of the aquaporins. This would reduce the rate of unloading from the coat symplasm. Water flow would continue to return to the xylem via the coat apoplasm with ongoing reabsorption of solutes into the cotyledon symplasm. (2) Once the apoplasmic build-up of solutes has been relieved, the aquaporins open, allowing resumption of mass flow from the phloem via the symplasmic connections. It is also intriguing that PIP aquaporins are gated by variation in cytoplasmic calcium, which has been implicated in turgor homeostasis of seed coat unloading cells (see below). The turgor homeostasis mechanism may involve the PIP aquaporins such that for a high efflux from the unloading cells, corresponding to downregulation of turgor, there is a concomitantly high water permeability (i.e. the same end result as the osmotic regulation mechanism described above). Thus it is temping to suggest that some of the aquaporins, with highly specific water permeability, may mediate both unloading and water recycling from the seed apoplasm by providing hydraulic coupling between xylem and neighbouring cells in seed coats (Maurel et al. 1997). Further investigation is required to resolve the pathway and kinetics by which the apoplasmic water is recycled and the likely involvement of aquaporins. Nutrient uptake by filial tissues is mediated by an array of membrane transporters Sugars High affinity hexose/H+ symporters have been cloned from faba bean cotyledons (VfSTP1, Weber et al. 1997a) and developing barley grains (HvSTP1 and HvSTP2, Weschke et al. 2003) at the pre-storage phase of development (Fig. 1). Here, hexose transporter activity accounts for broad bean cotyledon biomass gains but only a minor component once cotyledons enter the storage phase (Harrington et al. 2005). During the storage phase of seed fill, sucrose influx from the seed apoplasm exhibits a saturable component at low 322 W.-H. Zhang et al. Functional Plant Biology concentrations while a linear component dominates at higher concentrations (Patrick and Offler 1995). The saturable component has been characterised as a sucrose/H+ symport mechanism (Patrick and Offler 1995). Therefore, sucrose transporter genes expressed in filial tissue of dicots and monocots have been shown to function as sucrose/H+ symporters (Table 2; Fig. 1). A hint that these cloned sucrose transporters account for the saturable component of sucrose uptake by seeds is that their apparent Km values of 10 mM or less (see papers cited in Table 2) correspond with those for sucrose/H+ symport by developing grain legume seeds (Patrick and Offler 2001). Of the cloned sucrose/H+ symporters, only AtSUC5 exhibits a seed specific expression pattern (Baud et al. 2005). The remaining transporters are expressed, to varying degrees, throughout the parent plant body (see papers cited in Table 2). Applying the phylogenetic classification by Lalonde et al. (2004), the dicot sucrose transporters expressed in developing seeds cluster in all three sucrose transporter Clades with Clade I transporters being the most strongly represented. Clade I principally contains high affinity sucrose/H+ symporters (Lalonde et al. 2004) but not exclusively so (e.g. PvSUF1, PsSUF1, Zhou et al. 2007). Furthermore, seeds of some species have been found to express low affinity sucrose symporters (Barth et al. 2003) and facilitators (Zhou et al. 2007) that cluster in Clades II and III. In contrast, most sucrose transporters expressed in monocot seeds cluster in Clade III and all of these characterised thus far function as high affinity sucrose/H+ symporters (Table 2; Lalonde et al. 2004). The functional role of sucrose/H+ symporters in biomass gain by seeds only has been shown for rice. Here, antisense suppression of OsSUT1, that encodes a high affinity sucrose symporter, resulted in a shrivelled grain phenotype and, without phloem loading in leaves being compromised, points to a seed-specific role of the transporter (Scofield et al. 2002). Current evidence for dicot seeds is less convincing. Knocking out seed-specific AtSUC5 resulted in a biochemical seed phenotype containing less oil (Baud et al. 2005). However, seed size was unaffected suggesting that other transporters also contributed to sucrose import by Arabidopsis seeds. Consistent with a regulatory role of sucrose transporters in nutrient loading, selective overexpression of a high affinity potato sucrose/H+ symporter (StSUT1) in storage parenchyma cells of pea cotyledons enhanced their rates of sucrose influx and biomass gain (Rosche et al. 2002). Together, these observations are consistent with high affinity sucrose/H+ symporters, contributing to sink limitation observed during the storage phase of seed development (Borrás et al. 2004). Transporters contributing to the non-saturable component of sucrose transport have yet to be identified with certainty. In this context, a 62-kD soybean sucrose binding protein (SBP), first isolated by photoaffinity labelling (Ripp et al. 1988) and localised to tissues actively engaged in sucrose transport (Grimes et al. 1992), was shown to mediate a non-saturable component of sucrose uptake (Overvoorde et al. 1996; Delú-Filho et al. 2000). However, the latter findings could not be repeated with a faba bean SBP that was shown to be a storage protein (Heim et al. 2001). These disparities may be reconciled by the fact that the various isoforms of SBP could perform different functions (Hajduch et al. 2005). Another set of candidates that may contribute to the non-saturable component of sucrose uptake is a suite of sucrose transporter genes that function as low affinity SUFs. These are expressed in developing cotyledons of pea (PsSUF1, PsSUF4) and French bean (PvSUF1, Zhou et al. 2007; Table 2). Further investigations are required to provide more insight into the physiological roles, if any, of SBPs and SUFs may play in sucrose uptake by filial tissues of developing seeds. Table 2. Sucrose transporters expressed in developing seeds Clades are as per Lalonde et al. (2004). SUT, sucrose/H+ symporter; SUF, sucrose facilitator Class Clade Dicot I Monocot II III II III Plant species Arabidopsis thaliana Arabidopsis thaliana Glycine max Phaseolus vulgaris Phaseolus vulgaris Pisum sativum Pisum sativum Vicia faba Pisum sativum Plantago major Hordeum vulgaris Oryza sativa Hordeum vulgaris Oryza sativa Triticum aestivum Sucrose transporters AtSUC5 AtSUC3 GmSUT1 PvSUT1 PvSUF1 PsSUT1 PsSUF1 VtSUT1 PsSUF4 PmSUC3 HvSUT2 OsSUT2 HvSUT1 OsSUT1 OsSUT3 OsSUT4 OsSUT5 TaSUT1a TaSUT1b TaSUT1d References Baud et al. (2005) Meyer et al. (2000) Aldape et al. (2003) Zhou et al. (2007) Zhou et al. (2007) Tegeder et al. (1999) Zhou et al. (2007) Weber et al. (1997a) Zhou et al. (2007) Barth et al. (2003) Weschke et al. (2000) Aoki et al. (2002) Weschke et al. (2000) Hirose et al. (1997), Aoki et al. (2002) Aoki et al. (2002) Nutrient loading of developing seeds Amino acids Storage protein is a major reserve in legume seeds and its accumulation depends on amino nitrogen availability and uptake (Weber et al. 2005). At early stages of seed development, glutamine, alanine, threonine and valine are the main forms of amino N released from seed coats (Lanfermeijer et al. 1992), with increasing amounts of asparagine unloaded in later stages of development (Rochat and Boutin 1991). Amino N uptake into young cotyledons is mediated by a facilitated diffusion (non-saturable) mechanism. As storage product accumulation progresses, a saturable low affinity (Km = 5 mM) and high capacity uptake system predominates (Patrick and Offler 2001). Genes coding for broad-spectrum amino acid permeases (AAPs and cf. Fischer et al. 1998) have been cloned from pea and broad bean cotyledons (Tegeder et al. 2000a; Miranda et al. 2001; Fig. 1). Expression of VfAAP1 reaches a maximum before the beginning of storage protein accumulation, suggesting a role in providing cotyledons with amino acids used for the synthesis of storage proteins (Miranda et al. 2001). Consistent with this conclusion, overexpression of VfAAP1 in pea seeds resulted in higher seed protein levels (Rolletschek et al. 2005). In addition to amino acid transport, peptide transporters (PTR) may play a role in providing peptides for protein deposition during seed development (Miranda et al. 2003). Mineral ions Unlike sucrose and amino acids, few studies have been conducted to elucidate mechanisms of uptake of mineral nutrients in general, and K+ uptake in particular. These studies mainly have focused on legume cotyledons, which accumulate large amounts of K+ (Laszlo 1994) from an enriched seed apoplasm containing K+ concentrations of up to 100 mM (Patrick 1994). Thus, a low affinity K+ channel would be expected to mediate K+ influx across plasma membranes of cotyledon dermal cells. In this context, voltage- and time-dependent K+ -selective inward rectifying channels belonging to Shaker-like K+ channels have been characterised in many types of plant cells (Hedrich and Dietrich 1996). This type of K+ channel includes Arabidopsis KAT1 (Nakamura et al. 1995), AKT1 (Hirsch et al. 1998), potato KST1 (Müller-Röber et al. 1995) and maize ZMK1 channel (Philippar et al. 1999). A cationselective channel, highly selective for K+ over Cl− and Ca2+ , has been characterised in plasma membranes of protoplasts derived from dermal cell complexes in developing P. vulgaris cotyledons (Zhang et al. 2004b; Fig. 1). Unlike the KAT1 channels, this cation channel exhibits weak rectification and a strong dependence on the external Ca2+ and pH (Zhang et al. 2004b). The inhibitory effect of external Ca2+ on K+ current was dependent on voltage, with a greater inhibition observed at more negative membrane potentials. This causes a sigmoid shaped I-V curve with a negative conductance at membrane potentials between –220 and −140 mV (Zhang et al. 2004b). These characteristics are in contrast to the KAT1 channels, but they do resemble those of the phloem-related K+ inward rectifying channels such as AKT2/3 (Marten et al. 1999), ZMK2 (Bauer et al. 2000) and VFK1 (Ache et al. 2001). Moreover, the cotyledon cation channel differs from the characterised plant KAT1 and ATK2/3 channels in terms of selectivity for Functional Plant Biology 323 univalent cations. For example, the AKT2/3 channels display high selectivity for K+ over other univalent cations such as Na+ (Marten et al. 1999). By contrast, the cotyledon cation channel was relatively non-selective among univalent cations + (i.e. NH+ and Rb+ ) (Zhang et al. 2004b). Thus this 4 , Na cation channel can be classified into the category of plant nonselective cation channels (Demidchik et al. 2002). However, the conductance of the channel was markedly reduced when the external K+ was substituted by other univalent cations (Zhang et al. 2004b). These findings suggest that the cation channel allows both K+ and general univalent cation permeation. However, in the case of cations other than K+ , there is a stronger binding in the channel that limits permeation. Another feature of the cotyledon non-selective cation channel is that the Ca2+ -dependence of the current disappeared when K+ in the external solution was substituted with other univalent cations. For example, the conductance was greater in the presence of + + NH+ 4 and Na than in the presence of K at more hyperpolarised membrane potentials and at high external Ca2+ (Zhang et al. 2004b). These characteristics indicate that both external Ca2+ and membrane potentials play a role in modulating influx of K+ and other univalent cations. In addition, since cotyledon dermal cells are highly hyperpolarised under in vivo conditions (Zhang et al. 2004b), the Ca2+ - and pH-dependent cation channels would be fully activated, and thus support influx of phloem-imported K+ as well as other univalent cations into developing cotyledon. In developing P. vulgaris seeds, the molar ratio of sucrose and K+ released from their coats approximates to one (Walker et al. 2000). It is expected that influx of the two solutes is coordinated to maintain a constant concentration of sucrose and K+ in the seed apoplasm. In this context, both K+ influx through the nonselective cation channel (Zhang et al. 2004b) and sucrose/H+ symporter activity (Boorer et al. 1996; Zhou and Miller 2000) are dependent on membrane potential. Thus, it is likely that membrane potential is an important parameter that coordinates K+ and sucrose fluxes across plasma membranes of cotyledon dermal cells. In addition to membrane potential, pH-dependence of the cation channel may also play a role in regulating the fluxes of sucrose and K+ into the cotyledon cells. In this case, assuming that the rate of proton pumping continues unchanged, a decrease in the apoplasmic pH resulting from a reduction of the activity of the sucrose/H+ symporter would inhibit the channel-mediated K+ influx. A similar interaction between ATK2/3 K+ channel and sucrose/H+ symporter, via the phloem membrane potential, has been reported to regulate sucrose and K+ loading into the phloem (Deeken et al. 2002). Pathways of nutrient transport in filial tissues Cellular sites of nutrient uptake from the seed apoplasm have been deduced from a range of indirect observations similar to those used to discover cells responsible for nutrient release from maternal tissues of developing seeds (Patrick and Offler 1995, 2001). However, in marked contrast to nutrient release, transporters for sugar (hexose and sucrose) and amino acid uptake have been cloned from developing monocot and dicot seeds (for more details see previous section). This has allowed precise identification of cells expressing these transporters and, 324 Functional Plant Biology hence, a clearer understanding of nutrient transport pathways followed in filial tissues of developing seeds. During the pre-storage phase of seed development in faba bean (VfSTP1, Weber et al. 1997a) and barley (HvSTP1, HvSTP2, Weschke et al. 2003), hexose transporters are coordinately expressed with extracellular invertases to recover hexoses from the seed apoplasm (Harrington et al. 2005). Hexose transporters are expressed most highly in filial cells bordering the seed apoplasmic space (Weber et al. 1997a; Weschke et al. 2003; cf. Harrington et al. 2005). Presumably hexose transporter expression and activity persists in those seeds in which extracellular invertase activity is retained during the storage phase (Tomlinson et al. 2004; Chourey et al. 2006). Further, hexose transporters may not be expressed in developing Arabidopsis seeds. Here sucrose transporters are expressed in the cellularised endosperm (AtSUC5, Baud et al. 2005), suspensor and embryonic root tips (AtSUC3, Meyer et al. 2004; Stadler et al. 2005). The suspensor is considered to be a primary symplasmic route for delivering endosperm-derived nutrients to the developing embryo (Yeung and Meinke 1993; Stadler et al. 2005). In this context, an amino acid transporter is coexpressed in suspensor and endosperm cells (AtAAP1, Hirner et al. 1998) with AtSUC5 (Baud et al. 2005) and AtSUC3 (Meyer et al. 2004), respectively. Expression of sucrose transporters at these early stages of seed development (PsSUT1, Rosche et al. 2002) suggests that some released sucrose escapes extracellular hydrolysis and is loaded intact into endosperm and suspensor as well as embryo cells (Rosche et al. 2002; Stadler et al. 2005). As the embryo develops, symplasmic movement becomes more restricted as plasmodesmal size exclusion limits are downregulated (Kim et al. 2005; Stadler et al. 2005). However, these alterations in plasmodesmal conductivity may be more to direct movements of macromolecular signals than nutrient flows (Kim et al. 2005). In monocot and non-endospermic dicot seeds, declining extracellular invertase activity at the onset of the storage phase results in marked increases in apoplasmic sucrose concentrations (Patrick and Offler 2001). These events are accompanied by elevated expression levels of sucrose transporter genes in developing seeds of dicots (Weber et al. 1997a; Harrington et al. 1997b; Rosche et al. 2002; Aldape et al. 2003; Baud et al. 2005) and monocots (Hirose et al. 1997; Weschke et al. 2000; Aoki et al. 2002; Table 2). Strong expression, and high densities, of sucrose transporters have been detected throughout all dermal cell complexes of French [sucrose/H+ symporter (SUT), SBP] (Tegeder et al. 2000b) and broad (SBP) (Harrington et al. 1997a, 1997b) bean cotyledons. A broad specificity amino acid transporter (PsAAP1) co-expresses with the sucrose transporter (PsSUT1) in pea cotyledons (Tegeder et al. 1999, 2000a). This contrasts with faba bean cotyledons, where expression of an amino acid transporter is confined to the storage parenchyma cells (VfAAP1, Miranda et al. 2001) that could function in retrieval of amino acids leaked to the seed apoplasm. Moreover, a peptide transporter is localised to the adaxial epidermal cells (VfPTR1, Miranda et al. 2003) where it may play a primary role in peptide acquisition by the embryo (e.g. Stacey et al. 2002). In cereals, sucrose transporters are expressed in the aleurone/subaleurone endospermal transfer cells of wheat (SPB and SUT) (Bagnall et al. 2000), barley (HvSUT1; HvSUT2 – W.-H. Zhang et al. Weschke et al. 2000) and rice (OsSUT1, Furbank et al. 2001) grains. A more recent study shows that HvSUT2 is localised to tonoplast rather than plasma membranes (Endler et al. 2006). Transporter expression in wheat (Bagnall et al. 2000) and barley (Weschke et al. 2000) seeds is restricted to cells abutting their endosperm cavities. For rice grains, that lack an endosperm cavity, transporter expression spreads down two-thirds of the aleurone following the opposing nucellar tissues (Furbank et al. 2001). However, except for sucrose transporters in pea (PsSUT1, Tegeder et al. 1999) and amino acid transporters in faba bean (VfAAP1, Miranda et al. 2001) cotyledons, sucrose and amino acid transporters are localised to the outermost cell layers of the filial tissues that abut maternal sites for nutrient release (see previous section; Fig. 1). These uptake cells exhibit ultra-structural characteristics of cells committed to a transport function. These features include a dense cytoplasm, extensive networks of endoplasmic reticulum, mitochondria aligned to plasma membranes and, in some cases, invaginated wall ingrowths typical of a transfer cell morphology (Patrick and Offler 2001). This cellular localisation of transporters is accompanied by notably high resistances for solute diffusion through sub-surface layers of the filial apoplasm of both monocot and dicot seeds (Patrick and Offler 2001). The apoplasmic resistance is arranged in parallel with a symplasmic pathway, containing high densities of interconnecting plasmodesmata capable of supporting observed fluxes of sucrose to the storage cells (Patrick and Offler 2001). Integration of filial nutrient demand with membrane transport and phloem import Feed forward control of nutrient loading under supply limitation is regulated by nutrient levels reaching the developing seeds. Falling into this category are minor elements and amino acids (see earlier discussion). Feedback control, generated by filial demand, is less well understood. This involves transmission of regulatory signal(s) from sites of metabolism/compartmentation within developing filial tissues to control nutrient transport and transfer processes located along the source/path/sink system. In broad terms this control could be open (mediated by signals outside nutrient flows such as hormones) or closed (mediated by nutrient levels) loop. The former is illustrated by altering filial demand for Fe by overexpressing ferritin biosynthesis in filial seed tissues. In some instances this has caused an enhanced accumulation of Fe in seeds (e.g. rice grains, Qu et al. 2005). In this case, depletion in Fe levels in the filial cytoplasm must be transmitted back to and activate source loading mechanisms (see earlier discussion) to supply the additional Fe. Since Fe is a minor osmotic specie, signalling cannot involve a hydrostatic pressure change transmitted along the phloem path and must rely on the transmission of a putative hormonal signal from seed sink to source (Grusak 1995). This form of mechanism must apply to all nutrient species that make a minor contribution to osmolality of the phloem sap (see earlier discussion). In the case of major osmotic species the following closed-loop control may apply in developing legume seeds. Depletion of sucrose pools by increased rates of polymer formation in pea cotyledons appears to ease a substrate-regulated repression of PsSUT1 expression and, as a result, increase Nutrient loading of developing seeds sucrose uptake from the seed apoplasm (K. Chan, C. E. Offler and J. W. Patrick, unpublished data); a phenomenon demonstrated in vitro by exposing cotyledons to an excess supply of sucrose (Weber et al. 1997a). Resulting decreases in apoplasmic sap osmolality has been shown to stimulate efflux of nutrients from seed coats regulated by a turgor-homeostat mechanism (Patrick and Offler 1995; Walker et al. 2000; Fig. 1). Here, an increased turgor, resulting from an enhanced uptake of nutrients from the small apoplasmic pool by the cotyledons acts, as an error signal to elicit an immediate and compensatory increase in solute efflux to maintain a homeostatic turgor of the seed coat unloading cells (Patrick 1994; Zhang et al. 1996). The turgor-dependent increase in nutrient efflux results in a rapid downward turgor regulation in the unloading cells (Zhang et al. 1996). This downwardly regulated turgor maintains a constant hydrostatic pressure difference between source and sink, thus, allowing a sustained phloem import into seed coats to match rates of nutrient uptake by cotyledons. As discussed above it is possible that PIP aquaporins are also involved in this regulation. Therefore, turgor of unloading cells acts as a key modulator to link demand for major osmotic nutrients by filial cotyledons to nutrient supply from maternal seed coats. However, little is known about the mechanism(s) of turgor-induced efflux of nutrients from coats of developing grain legume seeds. Based on observations in other systems, a hypothetical scenario is outlined below. Turgor-dependent nutrient release from seed coats is likely to involve sensing of turgor by a receptor, transducing the turgor signal, which ultimately activates target membrane transporters. Cytoplasmic free Ca2+ activity ([Ca2+ ]c ) is a candidate signal to activate turgor-elicited efflux of nutrients as deduced from their responses to affectors of Ca2+ signalling (Walker et al. 2000; Fig. 1). A transient, biphasic increase in [Ca2+ ]c in response to a hypo-osmotic treatment has widely been observed in plant cells and is suggested to be a prerequisite for hypo-osmotically induced turgor regulation (Takahashi et al. 1997). The first phase increase in [Ca2+ ]c is likely to result from Ca2+ influx through mechano-sensitive Ca2+ channels (Cessna and Low 2001) and the second phase in [Ca2+ ]c elevation is probably mediated by intracellular Ca2+ release (Cessna et al. 1998; Cessna and Low 2001). In mammalian cells, the cytoskeleton plays an important role in sensing, transduction and regulation of osmotically-induced volume changes (Pedersen et al. 2001) through cytoskeletal re-organisation affecting [Ca2+ ]c (Lange and Brandt 1996). In contrast with animal cells, little is known about the role of cytoskeleton in regulating [Ca2+ ]c in plant cells (e.g. Liu and Luan 1998; Thion et al. 1998; Wang et al. 2004). In this context, we found that there was a marked increase in outward cation efflux when protoplasts derived from bean seed coat unloading cells were exposed to hypo-osmotic solutions (W.-H. Zhang, J. W. Patrick and S. D. Tyerman, unpublished data). The pulsing Cl− efflux also occurred more frequently under hypo-osmotic conditions (Zhang et al. 2004a). These osmotic responses were restricted to seed coat cells consistent with coat ‘efflux cells’, but not cotyledon ‘influx cells’ of developing P. vulgaris seeds, being capable of turgor regulation (Zhang et al. 1996). Treatment of protoplasts with cytochalasin D, under isoosmotic conditions, led to similar Functional Plant Biology 325 effects on outward and inward currents to that induced by hypo-osmotic treatments (W.-H. Zhang, J. W. Patrick and S. D. Tyerman, unpublished data). These findings suggest that the actin cytoskeleton acts as an osmosenser to modulate turgor-dependent solute efflux from seed coats (Fig. 1). In addition, the pulsing Cl− inward current was sensitive to neomycin, an antagonist of phospholipase C, suggesting this channel is modulated by IP3 -dependent intracellular Ca2+ release (cf. Wacke and Thiel 2001). We propose the following model (Fig. 1) based on these findings and the close link of actin filaments to phospholipase C activity in mammalian cells (van Haelst and Rothstein 1988). Disruption of actin filaments by hypo-osmotic treatment stimulates IP3 production through inducing phospholipase C activity. This in turn causes a transient increase in cytoplasmic Ca2+ activity through IP3 -mediated intracellular Ca2+ release which leads to activating a Ca2+ -dependent transient Cl− channel (Zhang et al. 2004a). Future prospects Progress is being made towards discovering the underlying molecular mechanisms regulating nutrient loading of seeds during their storage phase of development. Among the remaining key questions is to identify the membrane proteins mediating nutrient release from maternal seed tissues and their cellular location(s). Biochemical and biophysical approaches to study transport mechanisms in native membranes suffers from uncertainties introduced by working with membrane fractions sourced from a range of cell types that may perform differing transport functions. The least equivocal experimental approach will be to clone genes encoding membrane proteins responsible for nutrient efflux. Opportunities exist for cloning by homology for ion channels or using innovative functional expression systems for carriers. These proteins function in series with plasmodesmata to regulate nutrient flows in to and out of the maternal seed tissues. Mechanisms by which these maternal transport functions are coupled with filial demand for nutrients are only beginning to emerge, and present a rich environment for exciting future discoveries. Together these discoveries will contribute to understanding how the potential for nutrient accumulation by the filial tissues is realised. In the long term, focus needs to be brought to addressing the essentially unexplored area of nutrient loading during the pre-storage phase of seed development. Here the important steps of determining final seed number and potential size are clearly influenced by unknown nutrient transport and transfer processes. Acknowledgements This review is dedicated to the memory of Vincent Franceschi, a great friend and colleague who contributed significantly to conceptual advances in understanding nutrient loading of seeds. In particular, he made seminal findings as to the role paraveinal mesophyll play in assimilate partitioning and compartmentation (1983–2000). By a generous sharing of his immense intellect and innovative use of imaging and microtechniques, Vince has provided a continuing legacy that inspires and underpins efforts to discover mechanisms regulating nutrient transport to and within developing seeds. Studies reported from the authors’ laboratories were supported by grants from the Australian Research Council, Natural Science Foundation of China (30570136) and Grain Research and Development Corporation. 326 Functional Plant Biology References Ache P, Becker D, Deeken R, Dreyer I, Weber H, Fromm J, Hedrich R (2001) VFK1, a Vicia faba K+ channel involved in phloem unloading. The Plant Journal 27, 571–580. Aldape MJ, Elmer AM, Chao WS, Grimes HD (2003) Identification and characterization of a sucrose transporter isolated from the developing cotyledons of soybean. Archives of Biochemistry and Biophysics 409, 243–250. doi: 10.1016/S0003-9861(02)00631-8 Alleva K, Niemietz CM, Sutka M, Maurel C, Parisi M, Tyerman SD, Amodeo G (2006) Plasma membrane of Beta vulgaris storage root shows high water channel activity regulated by cytoplasmic pH and a dual range of calcium concentrations. Journal of Experimental Botany 57, 609–621. doi: 10.1093/jxb/erj046 Aoki N, Whitfield P, Hoeren F, Scofield G, Newell K, Patrick J, Offler C, Clarke B, Rahman S, Furbank RT (2002) Three sucrose transporter genes are expressed in the developing grain of hexaploid wheat. Plant Molecular Biology 50, 453–462. doi: 10.1023/A:1019846832163 Aoki N, Scofield GN, Wang X-D, Patrick JW, Offler CE, Furbank RT (2004) Expression and localisation analysis of the wheat sucrose transporter TaSUT1 in vegetative tissues. Planta 219, 176–184. doi: 10.1007/s00425-004-1232-7 Awazuhara M, Fujiwaa T, Hayashi H, Watanabe-Takahashi A, Takahashi H, Saito K (2005) The function of SULTR2;1 sulfate transporter during seed development in Arabidopsis thaliana. Physiologia Plantarum 125, 95–105. doi: 10.1111/j.1399-3054.2005.00543.x Bagnall N, Wang X-D, Scofield GN, Furbank RT, Offler CE, Patrick JW (2000) Sucrose transport-related genes are expressed in both maternal and filial tissues of developing wheat grains. Australian Journal of Plant Physiology 27, 1009–1020. Barth I, Meyer S, Sauer N (2003) PmSUC3: characterization of a SUT2/SUC3-type sucrose transporter from Plantago major. The Plant Cell 15, 1375–1385. doi: 10.1105/tpc.010967 Baud S, Wuilleme S, Lemoine R, Kronenberger J, Caboche M, Lepiniec L, Rochat C (2005) The AtSUC5 sucrose transporter specifically expressed in the endosperm is involved in early seed development in Arabidopsis. The Plant Journal 43, 824–836. doi: 10.1111/j.1365313X.2005.02496.x Bauer CS, Hoth S, Haga K, Philippar K, Aoki N, Hedrich R (2000) Differential expression and regulation of K+ channels in the maize coleoptile: molecular and biophysical analysis of cells isolated from cortex and vasculature. The Plant Journal 24, 139–145. doi: 10.1046/ j.1365-313X.2000.00844.x Boorer KJ, Loo DDF, Frommer W, Wright EM (1996) Transport mechanism of the cloned potato H+ /sucrose cotransporter StSUT1. Journal of Biological Chemistry 271, 25139–25144. doi: 10.1074/ jbc.271.41.25139 Borrás L, Slafer GA, Otegui ME (2004) Seed dry weight response to source–sink manipulation in wheat, maize and soybean: a quantitative re-appraisal. Field Crops Research 86, 131–146. doi: 10.1016/ j.fcr.2003.08.002 Bradford KJ (1994) Water stress and the water relations of seed development: a critical review. Crop Science 34, 1–11. Carpaneto A, Geiger D, Bamberg E, Sauer N, Fromm J, Hedrich R (2005) Phloem-localized, proton-coupled sucrose carrier ZmSUT1 mediates sucrose efflux under control of the sucrose gradient and the proton motive force. Journal of Biological Chemistry 280, 21437–21443. doi: 10.1074/jbc.M501785200 Cessna SG, Low PS (2001) Activation of the oxidative burst in aequorintransformed Nicotiana tabacum cells is mediated by protein kinase- and anion channel-dependent release of Ca2+ from internal stores. Planta V214, 126–134. Cessna SG, Chandra S, Low PS (1998) Hypo-osmotic shock of tobacco cells stimulates Ca2+ fluxes deriving first from external and then internal Ca2+ stores. Journal of Biological Chemistry 273, 27286–27291. doi: 10.1074/jbc.273.42.27286 W.-H. Zhang et al. Chourey PS, Jain M, Li Q-B, Carlson SJ (2006) Genetic control of cell wall invertases in developing endosperm of maize. Planta 223, 159–167. doi: 10.1007/s00425-005-0039-5 Cram WJ (1984) Manitol transport and suitability as an osmoticum in root cells. Physiologia Plantarum 61, 396–404. doi: 10.1111/j.13993054.1984.tb06346.x Curie C, Panaviene Z, Loulergue C, Dellaporta SL, Briat JF, Walker EL (2001) Maize yellow stripe1 encodes a membrane protein directly involved in Fe(III) uptake. Nature 409, 346–349. doi: 10.1038/ 35053080 Deeken R, Geiger D, Fromm J, Koroleva O, Ache P, Langenfeld R, Sauer N, May ST, Hedrich R (2002) Loss of the AKT2/3 potassium channel affects sugar loading into the phloem of Arabidopsis. Planta 216, 334–344. doi: 10.1007/s00425-002-0895-1 Delú-Filho N, Pirovani CP, Pedra JHF, Matrangolo FSV, Macedo JNA, Fontes EPB (2000) A sucrose binding protein homologue from soybean affects sucrose uptake in suspension-cultured transgenic tobacco cells. Plant Physiology and Biochemistry 38, 353–361. doi: 10.1016/S09819428(00)00752-X Demidchik V, Davenport RJ, Tester M (2002) Non-selective cation channels in plants. Annual Review of Plant Biology 53, 67–107. doi: 10.1146/ annurev.arplant.53.091901.161540 van Dongen JT, Laan RGW, Wouterlood M, Borstlap AC (2001) Electrodiffusional uptake of organic cations by pea seed coats. Further evidence for poorly selective pores in the plasma membrane of seed coat parenchyma cells. Plant Physiology 126, 1688–1697. doi: 10.1104/pp.126.4.1688 van Dongen JT, Ammerlaan AMH, Wouterlood M, Van Aelst AC, Borstlap AC (2003) Structure of the developing pea seed coat and the post-phloem transport pathway of nutrients. Annals of Botany 91, 729–737. doi: 10.1093/aob/mcg066 Dordas C, Shrispeels MJ, Brown PH (2000) Permeability and channelmediated transport of boric acid across membrane vesicles isolated from squash roots. Plant Physiology 124, 1349–1361. doi: 10.1104/ pp.124.3.1349 Endler A, Meyer S, Schelbert S, Schneider T, Weschke W, Peters SW, Keller F, Baginsky S, Martinoia E, Schmidt UG (2006) Identification of a vacuolar sucrose transporter in barley and Arabidopsis mesophyll cells by a tonoplast proteomic approach. Plant Physiology 141, 196–207. doi: 10.1104/pp.106.079533 Esau K (1965) ‘Vascular differentiation in plants.’ (Holt, Rinehart and Winston: New York) Evans LT, Wardlaw IF (1996) Wheat. In ‘In photoassimilate distribution in plants and crops. Source–sink relationships’. (Eds E Zamski, AE Schaffer) pp. 501–518. (Marcel Dekker Inc.: New York) Fenner M (2005) Seed size and chemical composition: the allocation of minerals to seeds and their use in early seedling growth. Botanical Journal of Scotland 56, 163–173. Fetter K, van Wilder V, Moshelion M, Chaumont F (2004) Interactions between plasma membrane aquaporins modulate their water channel activity. The Plant Cell 16, 215–228. doi: 10.1105/tpc.017194 Fieuw S, Patrick JW (1993) Mechanism of photosynthate efflux from Vicia faba L. seed coats. Journal of Experimental Botany 44, 65–74. doi: 10.1093/jxb/44.1.65 Fisher DB, Cash-Clark CE (2000a) Gradients in water potential and turgor pressure along the translocation pathway during grain filling in normally watered and water-stressed wheat plants. Plant Physiology 123, 139–147. doi: 10.1104/pp.123.1.139 Fisher DB, Cash-Clark CE (2000b) Sieve tube unloading and postphloem transport of fluorescent tracers and proteins injected into sieve tubes via severed aphid stylets. Plant Physiology 123, 125–138. doi: 10.1104/pp.123.1.125 Fisher DB, Wang N (1995) Sucrose concentration gradients along the postphloem transport pathway in the maternal tissues of developing wheat grains. Plant Physiology 109, 587–592. Nutrient loading of developing seeds Fischer WN, Andre B, Rentsch D, Krolkiewicz S, Tegeder M, Brenner ML, Frommer WB (1998) Amino acid transport in plants. Trends in Plant Science 3, 188–195. doi: 10.1016/S1360-1385 (98)01231-X Fitzgerald MA, Ugalde TD, Anderson JW (2001) Sulfur nutrition affects delivery and metabolism of S in developing endosperms of wheat. Journal of Experimental Botany 52, 1519–1526. doi: 10.1093/ jexbot/52.360.1519 Franceschi VR, Wittenbach VA, Giaquinta RT (1983) Paraveinal mesophyll of soybean leaves in relation to assimilate transfer and compartmentation. III. Immunohistochemical localization of specific glycopeptides in the vacuole after depodding. Plant Physiology 72, 586–589. Frachisse JM, Thomine S, Colcombet J, Guern J, Barbier-Brygoo H (1999) Sulfate is both a substrate and an activator of the voltage-dependent anion channel of Arabidopsis hypocotyl cells. Plant Physiology 121, 253–262. doi: 10.1104/pp.121.1.253 Furbank RT, Scofield GN, Hirose T, Wang XD, Patrick JW, Offler CE (2001) Cellular localisation and function of a sucrose transporter OsSUT1 in developing rice grains. Australian Journal of Plant Physiology 28, 1187–1196. Gahrtz M, Schmelzer E, Stolz J, Sauer N (1996) Expression of the PmSUC1 sucrose carrier gene from Plantago major L. is induced during seed development. The Plant Journal 9, 93–100. doi: 10.1046/j.1365313X.1996.09010093.x Gaymard F, Pilot G, Lacombe B, Bouchez D, Bruneau D, Boucherez J, Michaux-Ferrière N, Thibaud J-B, Sentenac H (1998) Identification and disruption of a plant shaker-like outward channel involved in K+ release into the xylem sap. Cell 94, 647–655. doi: 10.1016/S00928674(00)81606-2 Gehring M, Choi Y, Fischer RL (2004) Imprinting and seed development. The Plant Cell 16, S203–S213. doi: 10.1105/tpc.017988 Gerbeau P, Guclu J, Ripoche P, Maurel C (1999) Aquaporin Nt-TIPa can account for the high permeability of tobacco cell vacuolar membrane to small neutral solutes. The Plant Journal 18, 577–587. doi: 10.1046/j.1365-313x.1999.00481.x Good AG, Shrawat AK, Muench DG (2004) Can less yield more? Is reducing nutrient input into the environment compatible with maintaining crop production? Trends in Plant Science 9, 597–603. doi: 10.1016/j.tplants.2004.10.008 Grimes HD, Overvoorde PJ, Ripp K, Franceschi VR, Hitz WD (1992) A 62-kD sucrose binding protein is expressed and localized in tissues actively engaged in sucrose transport. The Plant Cell 4, 1561–1574. doi: 10.1105/tpc.4.12.1561 Grusak MA (1995) Whole-root iron (III)-reductase activity throughout the life cycle iron-grown Pisum sativum L. (Fabaceae): relevance to the iron nutrition of developing seeds. Planta 197, 111–117. doi: 10.1007/BF00239946 van Haelst C, Rothstein TL (1988) Cytochalasin stimulates phosphoinositide metabolism in murine B lymphocytes. Journal of Immunology 140, 1256–1258. Hagan ND, Upadhyaya LM, Tabe LM, Higgins TJV (2003) The redistribution of protein sulphur in transgenic rice expressing a gene for a foreign, sulfur-rich protein. The Plant Journal 34, 1–11. doi: 10.1046/j.1365-313X.2003.01699.x Hajduch M, Ganapathy A, Stein JW, Thelen JJ (2005) A systematic proteomic study of seed filling in soybean. Establishment of highresolution two-dimensional reference maps, expression profiles, and an interactive proteome database. Plant Physiology 137, 1397–1419. doi: 10.1104/pp.104.056614 Harrington GN, Franceschi VR, Offler CE, Patrick JW, Tegeder M, Frommer WB, Harper JF, Hitz WD (1997a) Cell specific expression of three genes involved in plasma membrane sucrose transport in developing Vicia faba seed. Protoplasma 197, 160–173. doi: 10.1007/BF01288025 Functional Plant Biology 327 Harrington GN, Nussbaumer Y, Wang X-D, Tegeder M, Franceschi VR, Frommer WB, Patrick JW, Offler CE (1997b) Spatial and temporal expression of sucrose transport-related genes in developing cotyledons of Vicia faba L. Protoplasma 200, 35–50. doi: 10.1007/BF01280733 Harrington GN, Dibley KE, Ritchie RJ, Offler CE, Patrick JW (2005) Hexose uptake by developing cotyledons of Vicia faba: physiological evidence for transporters of differing affinities and specificities. Functional Plant Biology 32, 987–995. doi: 10.1071/FP05081 Hedrich R, Dietrich P (1996) Plant K+ channels: similarity and diversity. Botanica Acta 109, 94–101. Heim U, Wang Q, Kurz T, Borisjuk L, Golombek S et al. (2001) Expression patterns and subcellular localization of a 52-kDa sucrosebinding protein homologue of Vicia faba (VfSBPL) suggest different functions during development. Plant Molecular Biology 47, 461–474. doi: 10.1023/A:1011886908619 Henzler T, Steudle E (2000) Transport and metabolic degradation of hydrogen peroxide in Chara corallina: model calculations and measurements with the pressure probe suggest transport of H2 O2 across water channels. Journal of Experimental Botany 51, 2053–2066. doi: 10.1093/jexbot/51.353.2053 Hirner B, Fischer WN, Rentsch D, Kwart M, Frommer WB (1998) Developmental control of H+ /amino acid permease gene expression during seed development of Arabidopsis. The Plant Journal 14, 535–544. doi: 10.1046/j.1365-313X.1998.00151.x Hirose T, Imaizumi N, Scofield GN, Furbank RT, Ohsugi R (1997) cDNA cloning and tissue specific expression of a gene for sucrose transporter from rice (Oryza sativa L.). Plant & Cell Physiology 38, 1389–1396. Hirose T, Endler A, Ohsugi R (1999) Gene expression of enzymes for starch and sucrose metabolism and transport in leaf sheaths of rice (Oryza sativa L.) during the heading period in relation to the sink to source transition. Plant Production Science 2, 178–183. Hirose T, Takano M, Terao T (2002) Cell wall invertase in developing rice caryopsis: molecular cloning of OsCIN1 and analysis of its expression in relation to its role in grain filling. Plant & Cell Physiology 43, 452–459. doi: 10.1093/pcp/pcf055 Hirsch RE, Lewis BD, Spalding EP, Sussman MR (1998) A role for the AKT1 potassium channel in plant nutrition. Science 280, 918–921. doi: 10.1126/science.280.5365.918 Hocking PJ, Pate JS (1977) Mobilization of minerals to developing seeds of legumes. Annals of Botany 41, 1259–1278. Inoue H, Higuchi K, Takahashi M, Nakanishi H, Mori H, Nishizawa NK (2003) Three rice nicotianamine synthase genes, OsNAS1, OsNAS2, and OsNAS3 are expressed in cells involved in long-distance transport of iron and differentially regulated by iron. The Plant Journal 36, 366–381. doi: 10.1046/j.1365-313X.2003.01878.x Jahn TP, Moller ALB, Zeuthen T, Holm LM, Klaerke D, Mohsin B, Kuhlbrandt W, Schjoerring JK (2004) Aquaporin homologues in plants and mammals transport ammonia. FEBS Letters 574, 31–36. doi: 10.1016/j.febslet.2004.08.004 Jenner CF (1985) Transport of tritiated water and 14 C-labeled assimilate into grains of wheat: III. Diffusion of THO through the stalk. Australian Journal of Plant Physiology 12, 595–608. Jenner CF, Jones GP (1990) Diffusion of water in the wheat grain: nuclear magnetic resonance and radioisotopic methods provide complementary information. Australian Journal of Plant Physiology 17, 107–118. Jenner CF, Ugalde TD, Aspinall D (1991) The physiology of starch and protein deposition in the endosperm of wheat. Australian Journal of Plant Physiology 18, 211–226. Johansson I, Karlsson M, Shukla VK, Chrispeels MJ, Larsson C, Kjellbom P (1998) Water transport activity of the plasma membrane aquaporin PM28A is regulated by phosphorylation. The Plant Cell 10, 451–459. doi: 10.1105/tpc.10.3.451 328 Functional Plant Biology de Jong A, Borstlap AC (2000) A plasma membrane-enriched fraction isolated from the coats of developing pea seeds contains H+ -symporters for amino acids and sucrose. Journal of Experimental Botany 51, 1671–1677. doi: 10.1093/jexbot/51.351.1671 Johanson U, Karlsson M, Johansson I, Gustavsson S, Sjövall S, Fraysse L, Weig AR, Kjellbom P (2001) The complete set of genes encoding major intrinsic proteins in Arabidopsis provides a framework for a new nomenclature for major intrinsic proteins in plants. Plant Physiology 126, 1358–1369. doi: 10.1104/pp.126.4.1358 de Jong AJ, Koerselmann-Kooij JW, Schuurmans JAMJ, Borstlap AC (1996) Characterisation of the uptake of sucrose and glucose by isolated seed coat halves of developing pea seeds. Evidence that a sugar facilitator with diffusional kinetics is involved in seed coat unloading. Planta 199, 486–492. doi: 10.1007/BF00195177 de Jong A, Koerselman-Kooij JW, Schuurmans JAMJ, Borstlap AC (1997) The mechanism of amino acid efflux from seed coats of developing pea seeds as revealed by uptake experiments. Plant Physiology 114, 731–736. Kiegle E, Moore CA, Haseloff J, Tester MA, Knight MR (2000) Celltype-specific calcium responses to drought, salt and cold in the Arabidopsis root. The Plant Journal 23, 267–278. doi: 10.1046/j.1365313x.2000.00786.x Kim I, Cho E, Crawfors K, Hempel FD, Zambryski PC (2005) Cell-to-cell movement of GFP during embryogenesis and early seedling development in Arabidopsis. Proceedings of the National Academy of Sciences USA 102, 2227–2231. doi: 10.1073/pnas.0409193102 King SP, Lunn JE, Furbank RT (1997) Carbohydrate content and enzyme metabolism in eveloping canola siliques. Plant Physiology 114, 153–160. Klauer SF, Franceschi VR, Ku MSB, Zhang D (1996) Identification and localization of vegetative storage proteins in legume leaves. American Journal of Botany 83, 1–10. doi: 10.2307/2445947 Lalonde S, Tegeder M, Throne-Holst M, Frommer WB, Patrick JW (2003) Phloem loading and unloading of sugars and amino acids. Plant, Cell & Environment 26, 37–56. doi: 10.1046/j.1365-3040. 2003.00847.x Lalonde S, Wipf D, Frommer WB (2004) Transport mechanisms for organic forms of carbon and nitrogen between source and sink. Annual Review of Plant Biology 55, 341–372. doi: 10.1146/ annurev.arplant.55.031903.141758 Lanfermeijer FC, Oene MA, Borstlap AC (1992) Compartmental analysis of amino-acid release from attached and detached pea seed coats. Planta 187, 75–82. doi: 10.1007/BF00201626 Lansing AJ, Franceschi VR (2000) The paraveinal mesophyll: a specialised path for intermediary transfer of assimilates in legume leaves. Australian Journal of Plant Physiology 27, 757–767. Lange K, Brandt U (1996) Calcium storage and release properties of F-actin: evidence for the involvement of F-actin in cellular calcium signalling. FEBS Letters 395, 137–142. doi: 10.1016/00145793(96)01025-3 Laszlo JA (1994) Changes in soybean fruit Ca2+ (Sr2+ ) and K (Rb+ ) transport ability during seed development. Plant Physiology 104, 937–944. Le Jean M, Schikora A, Mari S, Briat J-F, Curie C (2005) A loss-offunction mutation in AtYSL1 reveals its role in iron and nocotianamine seed loading. The Plant Journal 44, 769–782. doi: 10.1111/j.1365313X.2005.02569.x Liu K, Luan S (1998) Voltage-dependent K+ channels as targets of osmosensing in guard cells. The Plant Cell 10, 1957–1970. doi: 10.1105/tpc.10.11.1957 Lohaus G, Moellers C (2000) Phloem transport of amino acids in two Brassica napus L. genotypes and one B. carinata genotype in relation to their seed protein content. Planta 211, 833–840. doi: 10.1007/s004250000349 W.-H. Zhang et al. Lopez M, Bousser AS, Sissoeff I, Gaspar M, Lachaise B, Hoarau J, Mahe A (2003) Diurnal regulation of water transport and aquaporin gene expression in maize roots: contribution of PIP2 proteins. Plant & Cell Physiology 44, 1384–1395. doi: 10.1093/pcp/pcg168 Ma JF, Tamai K, Yamaji N, Mitani N, Konishi S, Katsuhara M, Ishiguro M, Murata Y, Yano M (2006) A silicon transporter in rice. Nature 440, 688–691. doi: 10.1038/nature04590 Maathuis FJM, Ichida AW, Sanders D, Schroeder JI (1997) Roles of higher plant K+ channels. Plant Physiology 114, 1141–1149. doi: 10.1104/pp.114.4.1141 Marentes E, Grusak MA (1998) Iron transport and storage within the seed coat and embryo of developing seeds of pea (Pisum sativum L.). Seed Science Research 8, 367–375. Marten I, Hoth S, Deeken P, Ache P, Ketchum KA, Hedrich R (1999) ATK3, a phloem-localized K+ channel, is blocked by protons. Proceedings of the National Academy of USA 96, 7581–7586. doi: 10.1073/pnas.96.13.7581 Maurel C, Chrispeels M, Lurin C, Tacnet F, Geelen D, Ripoche P, Guern J (1997) Function and regulation of seed aquaporins. Journal of Experimental Botany 48, 421–430. Meyer S, Melzer M, Truernit E, Hummer C, Besenbeck R, Stadler R, Sauer N (2000) AtSUC3, a gene encoding a new Arabidopsis sucrose transporter, is expressed in cells adjacent to the vascular tissue and in a carpel cell layer. The Plant Journal 24, 869–882. doi: 10.1046/j.1365313x.2000.00934.x Meyer S, Lauterbach C, Niedermeier M, Barth I, Sjolund RD, Sauer N (2004) Wounding enhances expression of AtSUC3, a sucrose transporter from Arabidopsis sieve elements and sink tissues. Plant Physiology 134, 684–693. doi: 10.1104/pp.103.033399 Miranda M, Borisjuk L, Tewes A, Heim U, Sauer N, Wobus U, Weber H (2001) Amino acid permeases in developing seeds of Vicia faba L.: expression precedes storage protein synthesis and is regulated by amino acid supply. The Plant Journal 28, 61–71. doi: 10.1046/j.1365313X.2001.01129.x Miranda M, Borisjuk L, Tewes A, Dietrich D, Rentsch D, Weber H, Wobus U (2003) Peptide and amino acid transporters are differentially regulated during seed development and germination in faba bean. Plant Physiology 132, 1950–1960. doi: 10.1104/pp.103.024422 Müller-Röber B, Ellenberg J, Porvart N, Willmitzer L, Busch H, Becker D, Dieterich P, Hoth S, Hedrich R (1995) Cloning and electrophysiological analysis of KST1, an inward rectifying K+ channel expressed in potato guard cells. EMBO Journal 14, 2409–2416. Murphy R (1989) Water flow across the sieve tube boundary: estimating turgor and some implications for phloem loading and unloading. IV. Root tips and seed coats. Annals of Botany 63, 571–579. Nakamura RL, Mckendree WL, Hirsch RE, Sedbrook JC, Gaber F, Sussman MR (1995) Expression of an Arabidopsis potassium channel gene in guard cells. Plant Physiology 109, 371–374. doi: 10.1104/pp.109.2.371 Niemietz CN, Tyerman SD (1997) Characterization of water channels in wheat root membrane vesicles. Plant Physiology 115, 561–567. Niemietz CM, Tyerman SD (2000) Channel-mediated permeation of ammonia gas through the peribacteroid membrane of soybean nodules. FEBS Letters 465, 110–114. doi: 10.1016/S0014-5793(99)01729-9 Offler CE, Patrick JW (1984) Cellular structures, plasma membrane surface areas and plasmodesmatal frequencies of seed coats of Phaseolus vulgaris L. in relation to photosynthate transfer. Australian Journal of Plant Physiology 11, 79–99. Offler CE, Patrick JW (1993) Pathway of photosynthate transfer in the developing seed of Vicia faba L.: a structural assessment of the role of transfer cells in unloading from the seed coat. Journal of Experimental Botany 44, 711–724. doi: 10.1093/jxb/44.4.711 Nutrient loading of developing seeds Ohto M, Fischer RL, Goldberg RB, Nakamura K, Harada JJ (2005) Control of seed mass by APETALA2. Proceedings of the National Academy of Sciences USA 102, 3123–3128. doi: 10.1073/pnas.0409858102 Overvoorde PJ, Frommer WB, Grimes HD (1996) A soybean sucrose binding protein independently mediates nonsaturable sucrose uptake in yeast. The Plant Cell 8, 271–280. doi: 10.1105/tpc.8.2.271 Park JH, Saier MH (1996) Phylogenetic characterization of the MIP family of transmembrane channel proteins. Journal of Membrane Biology 153, 171–180. doi: 10.1007/s002329900120 Pate JS, Armstrong EL (1996) Pea. In ‘Photoassimilate distribution in plants and crops. Source–sink relationships’. (Eds E Zamski, AE Schaffer) pp. 625–642. (Marcel Dekker Inc.: New York) Pate JS, Peoples MB, van Bel AJE, Kuo J, Atkins CA (1985) Diurnal water balance of the cowpea fruit. Plant Physiology 77, 148–156. Patrick JW (1994) Turgor-dependent unloading of assimilates from coats of developing legume seed. Assessment of the significance of the phenomenon in the whole plant. Physiologia Plantarum 90, 645–654. doi: 10.1111/j.1399-3054.1994.tb02519.x Patrick JW (1997) Phloem unloading: sieve element unloading and postsieve element transport. Annual Review of Plant Physiology and Plant Molecular Biology 48, 191–222. doi: 10.1146/annurev.arplant.48.1.191 Patrick JW, Offler CE (1995) Post-sieve element transport of sucrose in developing seeds. Australian Journal of Plant Physiology 22, 681–702. Patrick JW, Offler CE (2001) Compartmentation of transport and transfer events in developing seeds. Journal of Experimental Botany 52, 551–564. doi: 10.1093/jexbot/52.356.551 Patrick JW, Offler CE, Wang XD (1995) Cellular pathway of photosynthate transport in coats of developing seed of Vicia faba L. and Phaseolus vulgaris L. I. Extent of transport through the coat symplast. Journal of Experimental Botany 46, 35–47. doi: 10.1093/jxb/46.1.35 Pearson JN, Rengel Z, Jenner CF, Graham RD (1998) Dynamics of zinc and manganese movement into developing wheat grains. Australian Journal of Plant Physiology 25, 139–144. Pedersen SF, Hoffmann EK, Mills JW (2001) The cytoskeleton and cell volume regulation. Comparative Biochemistry and Physiology 130, 385–399. Philippar K, Fuchs I, Luthen H, Hoth S, Bauer CS et al. (1999) Auxininduced K+ channel expression represents an essential step in coleoptile growth and gravitropism. Proceedings of the National Academy of Sciences USA 96, 12186–12191. doi: 10.1073/pnas.96.21.12186 Qu LQ, Yoshihara T, Ooyama A, Goto F, Takaiwa F (2005) Iron accumulation does not parallel the high expression of ferratin in transgenic rice seeds. Planta 222, 225–233. doi: 10.1007/s00425-005-1530-8 Ripp KG, Viitanen PV, Hitz WD, Franceschi VR (1988) Identification of membrane protein associated with sucrose transport into cells of developing soybean cotyledons. Plant Physiology 88, 1435–1445. Ritchie RJ, Fieuw-Makaroff S, Patrick JW (2003) Sugar retrieval by coats of developing seeds of Phaseolus vulgaris L. and Vicia faba L. Plant & Cell Physiology 44, 163–172. doi: 10.1093/pcp/pcg022 Roberts AG, Oparka KJ (2003) Plasmodesmata and the control of symplastic transport. Plant, Cell & Environment 26, 103–124. doi: 10.1046/j.13653040.2003.00950.x Roberts SK, Tester M (1995) Inward and outward K+ selective currents in the plasma membrane of protoplasts from maize root cortex and stele. The Plant Journal 8, 811–825. doi: 10.1046/j.1365-313X.1995.8060811.x Rochat C, Boutin J-P (1991) Metabolism of phloem-borne amino acids in maternal tissues of fruit of nodulated or nitrate-fed pea plants (Pisum sativum L.). Journal of Experimental Botany 42, 207–214. doi: 10.1093/jxb/42.2.207 Rolletschek H, Hosein F, Miranda M, Heim U, Gotz K-P, Schlereth A, Borisjuk L, Saalbach I, Wobus U, Weber H (2005) Ectopic expression of an amino acid transporter (VfAAP1) in seeds of Vicia narbonensis and pea increases storage protein. Plant Physiology 137, 1236–1249. doi: 10.1104/pp.104.056523 Functional Plant Biology 329 Rosche E, Blackmore D, Tegeder M, Richardson T, Schroeder H, Higgins TJV, Frommer WB, Offler CE, Patrick JW (2002) Seed-specific overexpression of a potato sucrose transporter increases sucrose uptake and growth rates of developing pea cotyledons. The Plant Journal 30, 165–175. doi: 10.1046/j.1365-313X.2002.01282.x Ruan Y-L (2005) Recent advances in understanding cotton fibre and seed development. Seed Science Research 15, 269–280. doi: 10.1079/SSR2005217 Ruan YL, Llewellyn DJ, Furbank RT (2001) The control of singlecelled cotton fiber elongation by developmentally reversible gating of plasmodesmata and coordinated expression of sucrose and K+ transporters and expansin. The Plant Cell 13, 47–60. doi: 10.1105/tpc.13.1.47 Ruiz JM (2001) Aquaporin and its function in boron uptake. Trends in Plant Science 6, 95. doi: 10.1016/S1360-1385(00)01837-9 Ruuska SA, Girke T, Benning C, Ohlrooge JB (2002) Contrapuntal networks of gene expression during Arabidopsis seed filling. The Plant Cell 14, 1191–1206. doi: 10.1105/tpc.000877 Schiltz S, Gallardo K, Huart M, Negroni L, Sommerer N, Burstin J (2004) Proteome reference maps of vegetative tissues in pea. An investigation of nitrogen mobilization from leaves during seed filling. Plant Physiology 135, 2241–2260. doi: 10.1104/pp.104.041947 Schiltz S, Munier-Lolain N, Jeudy C, Burstin J, Salon C (2005) Dynamics of exogenous nitrogen partitioning and nitrogen remobilization from vegetative organs in pea revealed by 15 N in vivo labelling throughout seed filling. Plant Physiology 137, 1463–1473. doi: 10.1104/pp.104.056713 Schnyder H (1993) The role of carbohydrate storage and redistribution in the source–sink relations of wheat and barley during grain filling – a review. New Phytologist 123, 233–245. doi: 10.1111/j.14698137.1993.tb03731.x Schuurmans JAMJ, Van Dongen JT, Rutjens BPW, Boonman A, Pieterse CMJ, Borstlap AC (2003) Members of the aquaporin family in the developing pea seed coat include representatives of the PIP, TIP, and NIP subfamilies. Plant Molecular Biology 53, 655–667. doi: 10.1023/B:PLAN.0000019070.60954.77 Scofield GN, Hirose T, Gaudron JA, Upadhyaya NM, Ohsugi R, Furbank RT (2002) Antisense suppression of the rice sucrose transporter gene, OsSUT1, leads to impaired grain filling and germination but does not affect photosynthesis. Functional Plant Biology 29, 815–826. doi: 10.1071/PP01204 Sexton PJ, Shibles RM (1999) Activity of ATP sulfurylase in reproductive soybean. Crop Science 39, 131–135. Shackel KE, Turner NC (2000) Seed coat cell turgor in chickpea is independent of changes in plant and pod wall water potential. Journal of Experimental Botany 51, 895–900. doi: 10.1093/jexbot/51.346.895 Stacey G, Koh S, Granger C, Becker JM (2002) Peptide transport in plants. Trends in Plant Science 7, 257–263. doi: 10.1016/S13601385(02)02249-5 Stadler R, Wright KM, Lauterbach C, Amon G, Gahtz M, Feuerstein A, Oparka KJ, Sauer N (2005) Expression of GFP-fusions in Arabidopsis companion cells reveals non-specific protein trafficking into sieve elements and identifies a novel post-phloem domain in roots. The Plant Journal 41, 319–331. doi: 10.1111/j.1365-313X.2004.02298.x Staswick PE (1994) Storage proteins of vegetative plant tissues. Annual Review of Plant Physiology and Plant Molecular Biology 45, 303–322. doi: 10.1146/annurev.pp.45.060194.001511 Staswick PE, Zhang Z, Clements TE, Specht JE (2001) Efficient down regulation of the major vegetative storage protein genes in transgenic soybean does not compromise plant productivity. Plant Physiology 127, 1819–1826. doi: 10.1104/pp.127.4.1819 Tabe LM, Droux M (2001) Sulfur assimilation in developing lupin cotyledons could contribute significantly to the accumualtion of organic sulfur reserves in seed. Plant Physiology 126, 176–187. doi: 10.1104/pp.126.1.176 330 Functional Plant Biology Tabe LM, Droux M (2002) Limits to sulfur accumulation in transgenic lupin seeds expressing a foreign sulfur-rich protein. Plant Physiology 128, 1137–1148. doi: 10.1104/pp.010935 Tabe LM, Droux M (2003) Sulphate transporters in developing seeds. ComBiol. Conference 143. (Australian Society for Biochemistry and Molecular Biology Inc.: Kent Town, South Australia) Tabe LM, Hagan N, Higgins TJV (2002) Plasticity of seed protein composition in response to nitrogen and sulfur availability. Current Opinion in Plant Biology 5, 212–217. doi: 10.1016/S13695266(02)00252-2 Tabuchi M, Suglvama K, Inoue E, Sato T, Takahashi H, Yamaya T (2005) Severe reduction in growth rate and grain filling of rice mutants lacking OsGS1;1, a cytosolic glutamine synthetase1;1. The Plant Journal 42, 641–651. doi: 10.1111/j.1365-313X.2005.02406.x Takahashi K, Isobe M, Muto S (1997) An increase in cytosolic calcium ion concentration precedes hypoosmotic shock-induced activation of protein kinases in tobacco suspension culture cells. FEBS Letters 401, 202–206. doi: 10.1016/S0014-5793(96)01472-X Takahashi M, Terada Y, Nakai I, Nakanishi H, Yoshimura E, Mori S, Nishzawa NK (2003) Role of nicotianamine in the intracellular delivery of metals and plant reproductive development. The Plant Cell 15, 1263–1280. doi: 10.1105/tpc.010256 Takahashi S, Ishimaru K, Yazaki J, Fujii F, Shimbo K, Yamamoto K, Sakata K, Sasaki T, Kishimoto N, Kikuchi S (2005) Microarray analysis of sink–source transition in rice leaf sheaths. Breeding Science 55, 153–162. doi: 10.1270/jsbbs.55.153 Tegeder M, Wang X-D, Frommer WB, Offler CE, Patrick JW (1999) Sucrose transport into developing seeds of Pisum sativum L. The Plant Journal 18, 151–161. doi: 10.1046/j.1365-313X.1999.00439.x Tegeder M, Offler CE, Frommer WB, Patrick JW (2000a) Amino acid transporters are localized to transfer cells of developing pea seeds. Plant Physiology 122, 319–325. doi: 10.1104/pp.122.2.319 Tegeder M, Thomas M, Hetherington L, Wang X-D, Offler CE, Patrick JW (2000b) Genotypic differences in seed growth rates of Phaseolus vulgaris L. – II. Factors contributing to cotyledon sink activity and sink size. Australian Journal of Plant Physiology 27, 119–128. Thion L, Mazars C, Nacry P, Bouchez D, Moreau M, Ranjeva R, Thuleau P (1998) Plasma membrane depolarizationactivated calcium channels, stimulated by microtubuledepolymerizing drugs in wildtype Arabidopsis thaliana protoplasts, display constitutively large activities and a longer half life in ton 2 mutant cells affected in the organization of cortical microtubules. The Plant Journal 13, 603–610. doi: 10.1046/j.1365313X.1998.00062.x Thomas M, Hetherington L, Patrick JW (2000) Genotypic differences in seed growth rates of Phaseolus vulgaris L. I. General characteristics, seed coat growth factors and comparative roles of seed coats and cotyledons. Australian Journal of Plant Physiology 27, 109–118. Thorne JH (1985) Phloem unloading of C and N assimilates in developing seeds. Annual Review of Plant Physiology 36, 317–343. doi: 10.1146/annurev.pp.36.060185.001533 Tilsner J, Kassner N, Struck C, Lohaus G (2005) Amino acid contents and transport in oilseed rape (Brassica napus L.) under different nitrogen conditions. Planta 221, 328–338. doi: 10.1007/s00425004-1446-8 Tomlinson KL, McHugh S, Labbe H, Grainger JL, James LE, Pomcroy KM, Mullin JW, Miller SS, Dennis DT, Miki BLA (2004) Evidence that the hexose-to-sucrose ratio does not control the switch to storage product accumulation in oilseeds: analysis of tobacco seed development and effects of overexpressing apoplastic invertase. Journal of Experimental Botany 55, 2291–2303. doi: 10.1093/jxb/erh251 Törnroth-Horsefield S, Wang Y, Hedfalk K, Johanson U, Karlsson M, Tajkhorshid E, Neutze R, Kjellbom P (2006) Structural mechanism of plant aquaporin gating. Nature 439, 688–694. doi: 10.1038/nature04316 W.-H. Zhang et al. Tournaire-Roux C, Sutka M, Javot H, Gout E, Gerbeau P, Luu DT, Bligny R, Maurel C (2003) Cytosolic pH regulates root water transport during anoxia stress through gating of aquaporins. Nature 425, 393–397. doi: 10.1038/nature01853 Tyerman SD, Niemietz CM, Bramley H (2002) Plant aquaporins: multifunctional water and solute channels with expanding roles. Plant, Cell & Environment 25, 173–194. doi: 10.1046/j.00168025.2001.00791.x Uehlein N, Lovisolo C, Siefritz F, Kaldenhoff R (2003) The tobacco aquaporin NtAQP1 is a membrane CO2 pore with physiological functions. Nature 425, 734–737. doi: 10.1038/nature02027 Vandeleur R, Niemietz CM, Tilbrook J, Tyerman SD (2005) Roles of aquaporins in root responses to irrigation. Plant and Soil 274, 141–161. doi: 10.1007/s11104-004-8070-z Wacke M, Thiel G (2001) Electrically triggered all-or-none Ca 2+ liberation during action potential in the giant alga Chara. Journal of General Physiology 118, 11–21. doi: 10.1085/jgp.118.1.11 Walker NA, Patrick JW, Zhang W-H, Fieuw S (1995) Efflux of photosynthate and acid from developing seed coats of Phaseolus vulgaris L.: a chemiosmotic analysis of pump driven efflux. Journal of Experimental Botany 46, 539–549. doi: 10.1093/jxb/46.5.539 Walker NA, Zhang W-H, Harrington G, Holdaway N, Patrick JW (2000) Effluxes of solutes from developing seed coats of Phaseolus vulgaris L. and Vicia faba L.: locating the effects of turgor in a coupled chemisosmotic system. Journal of Experimental Botany 51, 1047–1055. doi: 10.1093/jexbot/51.347. 1047 Wang N, Fisher D (1994) The use of fluorescent tracers to characterize the post-phloem transport pathway in maternal tissues of developing wheat grains. Plant Physiology 104, 17–27. Wang N, Fisher DB (1995) Sucrose release into the endosperm cavity of wheat grains apparently occurs by facilitated diffusion across the nucellar cell membranes. Plant Physiology 109, 579–585. Wang X-D, Harrington G, Patrick JW, Offler CE, Fieuw S (1995) Cellular pathway of photosynthate transport in coats of developing seed of Vicia faba L. and Phaseolus vulgaris L. II. Principal cellular site(s) of efflux. Journal of Experimental Botany 46, 49–63. doi: 10.1093/jxb/46.1.49 Wang Y-F, Fan L-M, Zhang W-Z, Zhang W, Wu W-H (2004) Ca2+ permeable channels in the plasma membrane of Arabidopsis pollen are regulated by actin microfilaments. Plant Physiology 136, 3892–3904. doi: 10.1104/pp.104.042754 Waters BM, Chu H-H, DiDonato RJ, Roberts LA, Eisley RB, Lahner B, Salt DE, Walker EL (2006) Mutations in Arabidopsis yellow stripe-like1 and yellow stripe-like3 reveal their roles in metal homeostasis and loading of metal ions in seeds. Plant Physiology 141, 1446–1458. doi: 10.1104/pp.106.082586 Weber H, Borisjuk L, Heim U, Sauer N, Wobus U (1997a) A role for sugar transporters during seed development: molecular characterization of a hexose and sucrose carrier in faba bean seeds. The Plant Cell 9, 895–908. doi: 10.1105/tpc.9.6.895 Weber H, Borisjuk L, Wobus U (1997b) Sugar import and metabolism during seed development. Trends in Plant Science 2, 169–174. doi: 10.1016/S1360-1385(97)85222-3 Weber H, Borisjuk L, Wobus U (2005) Molecular physiology of legume seed development. Annual Review of Plant Biology 56, 253–279. doi: 10.1146/annurev.arplant.56.032604.144201 Wegner LH, de Boer AH (1997) Properties of two outward-rectifying channels in root xylem parenchyma cells suggest a role of in K+ homeostasis and long distance signalling. Plant Physiology 115, 1707–1719. Wegner LH, Raschke K (1994) Ion channels in the xylem parenchyma of barley roots. Plant Physiology 105, 799–813. Nutrient loading of developing seeds Functional Plant Biology Weschke W, Panitz R, Sauer N, Wang Q, Neubohn B, Wobus U (2000) Sucrose transport into barley seeds: molecular characterization of two transporters and implications for seed development and starch accumulation. The Plant Journal 21, 455–467. doi: 10.1046/j.1365313x.2000.00695.x Weschke W, Panitz R, Gubatz S, Wang Q, Radchuk R, Weber H, Wobus U (2003) The role of invertases and hexose transporters in controlling sugar ratios in maternal and filial tissues of barley caryopses during early development. The Plant Journal 33, 395–411. doi: 10.1046/j.1365313X.2003.01633.x von Wiren N, Klair S, Bansal S, Briat J-F, Khodr H, Shiori T, Leigh RA, Hider RC (1999) Nicotianamine chelates both FeIII and FeII. Implications for metal transport in plants. Plant Physiology 119, 1107–1114. doi: 10.1104/pp.119.3.1107 Wolswinkel P (1992) Transport of nutrients into developing seeds: a review of physiological mechanisms. Seed Science Research 2, 59–73. Wolswinkel P, Ammerlaan A, Koerselman-Kooij J (1992) Effect of the osmotic environment on K+ and Mg2+ release from the seed coat and cotyledons of developing seeds of Vicia faba and Pisum sativum. Evidence for a stimulation of efflux from the vacuole at high cell turgor. Journal of Experimental Botany 43, 681–693. doi: 10.1093/jxb/43.5.681 Yang J, Zhang J (2006) Grain filling of cereals under soil drying. New Phytologist 169, 223–236. doi: 10.1111/j.1469-8137.2005.01597.x Ye Q, Muhr J, Steudle E (2005) A cohesion/tension model for the gating of aquaporins allows estimation of water channel pore volumes in Chara. Plant Cell & Environment 28, 525–535. doi: 10.1111/j.13653040.2004.01298.x Yeung EC, Meinke DW (1993) Embryogenesis in angiosperms: development of the suspensor. The Plant Cell 5, 1371–1381. doi: 10.1105/tpc.5.10.1371 Zhang W-H, Atwell BJ, Patrick JW, Walker NA (1996) Turgor-dependent efflux from of assimilate from coats of developing seeds of Phaseolus vulgaris L.: water relations of the cells involved in efflux. Planta 199, 25–33. 331 Zhang WH, Walker N, Tyerman S, Patrick J (1997) Mechanisms of solute efflux from seed coats: whole-cell K+ currents in transfer cell protoplasts derived from coats of developing seeds of Vicia faba L. Journal of Experimental Botany 48, 1565–1572. Zhang W-H, Walker NA, Tyerman SD, Patrick JW (2000) Fast activation of a time-dependent outward current in protoplasts derived from coats of developing Phaseolus vulgaris seeds. Planta 211, 894–898. doi: 10.1007/s004250000391 Zhang W-H, Skerrett M, Walker NA, Patrick JW, Tyerman SD (2002) Non-selective currents and channels in plasma membranes of protoplasts from coats of developing seeds of bean seeds. Plant Physiology 128, 388–399. doi: 10.1104/pp.128.2.388 Zhang W-H, Walker NA, Tyerman SD, Patrick JW (2004a) Pulsing Cl− channels linked to hypoosmotically-induced turgor regulation in coat cells of developing bean seeds. Journal of Experimental Botany 55, 993–1001. doi: 10.1093/jxb/erh120 Zhang W-H, Walker NA, Tyerman SD, Patrick JW (2004b) Ca2+ -dependent K current in dermal cells of developing bean cotyledons. Plant, Cell & Environment 27, 251–262. doi: 10.1046/j.1365-3040.2004. 01152.x Zhou JJ, Miller AJ (2000) Comparison of the transport properties of three plant sucrose carriers expressed in Xenopus oocytes. Australian Journal of Plant Physiology 27, 725–732. Zhou Y, Qu H, Dibley KE, Offler CE, Patrick JW (2007) A suite of sucrose transporters expressed in coats of developing legume seeds includes novel pH-independent facilitators. The Plant Journal 49, 750–764. doi: 10.1111/j.1365-313X.2006.03000.x Manuscript received 26 October 2006, accepted 30 January 2007 http://www.publish.csiro.au/journals/fpb