Download Review: Nutrient loading of developing seeds

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Functional Plant Biology, 2007, 34, 314–331
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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
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Manuscript received 26 October 2006, accepted 30 January 2007
http://www.publish.csiro.au/journals/fpb