Download Exocytosis and cell polarity in plants exocyst and recycling domains

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Exocytosis and cell polarity in plants –
exocyst and recycling domains
Author for correspondence:
Viktor Žárský
Tel: +420 221 951 685
Emails: [email protected],
[email protected]
Viktor Žárský1,2, Fatima Cvrcková1, Martin Potocký2 and Michal Hála2
1
Department of Plant Physiology, Charles University, Vinicná 5, 128 44 Praha 2, Czech Republic;
2
Institute of Experimental Botany, Academy of Sciences of the Czech Republic, Rozvojová 263, 165 02
Praha 6, Czech Republic
Received: 18 February 2009
Accepted: 6 April 2009
Contents
Summary
255
I.
Introduction
256
II.
Donor compartments for plant exocytotic vesicles: where do
they come from?
256
V.
VI.
III.
IV.
Mechanisms of exocytotic vesicle formation:
is the truth naked?
257
Building the tracks for exocytotic vesicles: cytoskeleton in
exocytosis and cell polarization
259
Manysidedness of a plant cell: delimiting functional
plasmalemma domains within a cell
260
Vesicle tethering, docking and fusion
262
VII. Polarity by recycling: exocytosis, endocytosis and recycling
domains in plant cells
264
VIII. Conclusions: constitutive or regulated secretion in
plant cells?
266
Acknowledgements
267
References
267
Summary
New Phytologist (2009) 183: 255–272
doi: 10.1111/j.1469-8137.2009.02880.x
Key words: cell polarity, Exo70, exocyst,
exocytosis, GTPases, membrane recycling,
recycling domain, secretory pathway.
© The Authors (2009)
Journal compilation © New Phytologist (2009)
In plants, exocytosis is a central mechanism of cell morphogenesis. We still know
surprisingly little about some aspects of this process, starting with exocytotic vesicle
formation, which may take place at the trans-Golgi network even without coat
assistance, facilitated by the local regulation of membrane lipid organization. The
RabA4b guanosine triphosphatase (GTPase), recruiting phosphatidylinositol-4-kinase
to the trans-Golgi network, is a candidate vesicle formation organizer. However, in
plant cells, there are obviously additional endosomal source compartments for
secretory vesicles. The Rho/Rop GTPase regulatory module is central for the initiation
of exocytotically active domains in plant cell cortex (activated cortical domains).
Most plant cells exhibit several distinct plasma membrane domains, established and
maintained by endocytosis-driven membrane recycling. We propose the concept of
a ‘recycling domain’, uniting the activated cortical domain and the connected
endosomal compartments, as a dynamic spatiotemporal entity. We have recently
described the exocyst tethering complex in plant cells. As a result of the multiplicity
of its putative Exo70 subunits, this complex may belong to core regulators of recycling
domain organization, including the generation of multiple recycling domains within
a single cell. The conventional textbook concept that the plant secretory pathway
is largely constitutive is misleading.
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Abbreviations: ABA, abscisic acid; ACD, activated cortical domain; AGP, arabinogalactan protein; ATPase, adenosine triphosphatase; BODIPY, 4,4-difluoro-5,7-dimethyl4-bora-3a,4a-diaza-S-indacene; CDPK, calcium-dependent protein kinase; DAG,
diacylglycerol; DRP, dynamin-related protein; EE, early endosome; GAP, GTPaseactivating protein; GDI, GDP dissociation inhibitor; GEF, guanine nucleotide exchange
factor; GPI, glycosylphosphatidylinositol; GTPase, guanosine triphosphatase; LE, late
endosome; MVB, multivesicular body; NOX, NADPH oxidase; PA, phosphatidic acid;
PKD, protein kinase D; PLC/D, phospholipase C/D; PPI, phosphoinositide; PtdIns4P,
phosphatidylinositol-4-phosphate; PtdIns-P, phosphatidylinositol-phosphate; PtdIns4P5K3, PtdIns4P,5-kinase isoform 3; RD, recycling domain; RE, recycling endosome; RLK,
receptor-like serine/threonine kinase; ROS, reactive oxygen species; sec-GFP, secreted
green fluorescent protein; TGA, tip growth apparatus; TGN, trans-Golgi network;
TPC, TGN-to-plasma membrane carrier; XET, xyloglucan endotransglycosylase.
I. Introduction
Exo- and endocytosis in plant cells should be understood as
inseparable phases of the same dynamic secretory process
(Battey et al., 1999). Surprisingly, we still know less about
plant exocytosis than about transport to the vacuole or
endocytosis (Rojo & Denecke, 2008). Foresti & Denecke
(2008) recently noted: ‘Solving the mystery of how secreted
proteins reach the plasma membrane remains a formidable
challenge for the plant field’. We thus may need to speculate
more often than usual for this type of review.
Here, we focus on the last exocytosis step between donor
compartments and the plasmalemma. Details on endomembrane
compartment relationships, cellular and molecular machineries
of the ‘core’ secretory pathway and other relevant phenomena
not covered here can be found in recent reviews, including
whole special issues of journals [Plant Physiology 147(4), 2008;
Current Opinion in Plant Biology 11(6), 2008]. However, we
must include some aspects of endocytosis, important in the
context of cell polarity regulation and in the proposed concept
of recycling domains (RDs). We apologize to the authors of
many relevant reports not covered because of space limitations.
We use the terms ‘exocytotic vesicle’ and ‘secretory vesicle’
as synonyms denoting any membrane container competent to
fuse with the plasmalemma in vivo. Specifically for exocytotic
carriers from the trans-Golgi network (TGN) to the plasmalemma, we adopt the previously proposed term ‘TGN-toplasma membrane carriers’ (TPCs; Bard & Malhotra, 2006).
II. Donor compartments for plant exocytotic
vesicles: where do they come from?
Exocytotic containers vary in size, appearance and contents,
indicating possible diverse compartment origins. The largest
secretory compartments may be multivesicular bodies (MVBs;
Février & Raposo, 2004) or whole trans-cisternae of the Golgi
apparatus in some unicellular algae. A typical exocytotic vesicle
diameter ranges between 60 and 150 nm; in Arabidopsis
pollen tubes, it is c. 180 nm, whereas, in root hairs, it is only
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c. 70 nm (Ketelaar et al., 2008). Chara has two classes of
exocytotic vesicle – 200 nm light and 180 nm dark (Limbach
et al., 2008). Even budding yeast has two types of 100 nm
secretory vesicle – those containing the Bgl2 glucantransferase
and plasma membrane proteins and those carrying periplasmic
invertase. Although Bgl2 vesicles travel directly from TGN to
the plasmalemma, invertase is transported through endosomal
compartments (Harsay & Schekman, 2002).
Which plant endomembrane system compartments produce exocytotic vesicles? The answer depends on the cell type;
however, TGN is clearly not the only source. The structural
complexity of angiosperm tissues, as well as the molecular
complexity of, for example, the plant RabA guanosine
triphosphatase (GTPase) subfamily (Rutherford & Moore,
2002), suggests multiple pathways to the cell surface. The
diversity of the Arabidopsis plasmalemma SNAREs is consistent
with at least three exocytotic pathways (Uemura et al., 2004).
Different pathways are used for secreted proteins versus cell
wall polysaccharides, and two closely related syntaxins, SYP121
and SYP122, participate in two independent exocytotic
pathways in tobacco (Leucci et al., 2007; Rehman et al., 2008).
1. The Golgi and trans-Golgi network (I, II)
Exocytosis from TGN is considered the default: cargos
without specific sorting signals are secreted. In addition, in
plants, soluble cargos travel to the apoplast by default, as
documented, for example, for secreted green fluorescent
protein (sec-GFP) (Batoko et al., 2000) or the Clv3 peptide
(Rojo et al., 2002). However, at least some plasmalemma
proteins are possibly sorted at the TGN (Bard & Malhotra,
2006; Wang et al., 2006; Foresti & Denecke, 2008).
RabE/Rab8 GTPases may characterize a Golgi/TGN compartment with direct exocytotic connection to the plasmalemma (Zheng et al., 2005; pathway II in Fig. 1), although
they may also alternatively function together with RabA in
pathway I (see below; Woollard & Moore, 2008).
Plant TGN is partially a Golgi apparatus-independent
organelle, as TGN and Golgi SNARE markers are often
© The Authors (2009)
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Fig. 1 Sources of plant exocytotic vesicles from indisputable (filled
heavy arrow) to hypothetical (broken light arrow). Marker proteins
for various compartments are shown in bold. The Golgi apparatus
and trans-Golgi network (TGN) can be viewed as at least three
partially overlapping subcompartments – TGN/early endosome (EE)/
recycling endosome (RE) (I), TGN/Golgi (II) and a CC domain (coat
in red). TGN-to-plasma membrane carriers (TPCs) may arise without
specific coat proteins as a result of lipid modification-induced
membrane deformation (blue halo) or by dynamin-induced
membrane scission (purple rings). Other possible sources of secretory
vesicles include GNOM and other putative RE compartments (III), the
prevacuolar compartment (PVC) [multivesicular body (MVB)/late
endosome (LE); IV; IVb shows whole MVB exocytosis with exosome
vesicle release to the apoplast), and ‘kiss-and-run’ vesicles that may
originate from various compartments (V). Relationships with other
endomembrane compartments are not shown.
separated (Uemura et al., 2004). The notion that plant TGN
itself is an early (or recycling) endosome is now generally
accepted (Dettmer et al., 2006; Robinson et al., 2008; Woollard & Moore, 2008). TGN consists of at least three partially
overlapping domains, one of which, characterized by the
presence of the V-adenosine triphosphatase (ATPase) subunit
paralogue VHA-a1, syntaxin SYP41 and SCAMP1 (Dettmer
et al., 2006; Lam et al., 2007), represents a TGN/early endosome (EE) compartment, partially overlapping with the
RabA-2/3 compartment (Chow et al., 2008). TGN may
function also as a recycling endosome (RE); it is unclear
whether plant TGN/EE serves also as a sorting endosome (for
example, Foresti & Denecke, 2008; Robinson et al., 2008;
Woollard & Moore, 2008). Different TGN subdomains may
harbour not only different Arf and Rab GTPases as specific
membrane domain organizers (Zerial & McBride, 2001), but
also specific local modifications of membrane lipid composition. The maturation of TGN-derived compartments may
involve Rab conversion, that is subsequent replacement of
Rab GTPase isoforms (Rink et al., 2005). Within the framework of our RD concept (see below), we may expect diversification of TGN/EE/REs, resulting in the coexistence of
differentially equipped/matured TGNs within the same cell.
Review
GNOM-dependent compartment participating in the
constitutive recycling of PIN auxin transporters (Geldner
et al., 2003), driven by clathrin-dependent PIN endocytosis
(Dhonukshe et al., 2007). Two endocytotic pathways, one
clathrin-dependent and the other clathrin-independent, have
been documented recently in tobacco. Endocytosed nanogold
particles first enter the early endosome compartment with a
tubulo-vesicular structure, and pulse-chase experiments have
proven that most of the internalized marker is recycled back
to the plasma membrane (Onelli et al., 2008). Several different
REs (with different Arfs/Rabs and their regulators) may coexist within the same plant cell.
3. The prevacuolar compartment (multivesicular body,
late endosome; IV)
The auxin influx carrier Aux1 is recycled by a mechanism
distinct from the PIN1 recycling pathway, involving the
SNX1 endosome characterized by the presence of sorting
nexin 1 (AtSNX1; Jaillais et al., 2006), and possibly identical
with the late endosome (LE) or MVB, characterized by the RabF
GTPases, the PEP12 SNARE and phosphatidylinositol-3phosphate-rich membranes (Robinson et al., 2008).
Exocytosis from LE/MVB or even lysosomes (for example,
D. Li et al., 2008) is well documented in metazoans, especially in
the form of whole MVB fusion with the plasmalemma, resulting
in exosome vesicle release into the apoplast (Février & Raposo,
2004; Fig. 1, IVb). In plants, possible direct exocytosis from
MVB to the plasma membrane is accepted as a speculative
possibility (see Foresti & Denecke, 2008; Robinson et al.,
2008). Plant proteins recycle to the plasmalemma mostly
indirectly via TGN, employing the retromer salvage pathway
for vacuolar sorting receptors (see Robinson et al., 2008;
Woollard & Moore, 2008). The operation of the MVB–
exosomal pathway (Fig. 1, IVb) in plants was recently indicated by Meyer et al. (2009) as participating in the formation
of pathogen-induced cell wall compartments.
4. ‘Kiss-and-run’ vesicles (V)
‘Kiss-and-run’ exocytosis, immediately followed by vesicle
retrieval, has been documented in plant protoplasts (Weise
et al., 2000). Kiss-and-run vesicles might also be refilled with
a solute cargo, such as, for example, auxin, in the cytoplasm
(Baluška et al., 2005).
Different cargos might use the same endosome as early,
sorting or recycling, making finite categorization of endosome
compartments impossible.
2. The recycling endosome (III)
III. Mechanisms of exocytotic vesicle formation:
is the truth naked?
The first plant RE candidate characterized at the molecular
level was the Arf guanine nucleotide exchange factor (GEF)
Although most, if not all, vesicles are believed to form with the
aid of coat proteins, coat-like structures are observed rarely on
© The Authors (2009)
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presumed plant secretory vesicles (for example, Staehelin &
Moore, 1995). The mechanisms of TPC formation belong to
major ‘unsolved mysteries in membrane traffic’ (Pfeffer, 2007).
Local modifications of membrane lipid composition, such
as local production of diacylglycerol (DAG), may contribute
to coat-less vesicle formation (Bard & Malhotra, 2006; Helling
et al., 2006). Recent evidence implicates phosphatidylinositol4-phosphate (PtdIns4P), traditionally understood only as a
precursor of PtdIns(4,5)P2, in membrane trafficking control.
In plants, PtdIns4P makes up 80% of the total PtdInsP pool
(see Meijer & Munnik, 2003), and was recently found in the
Golgi and plasma membrane, but not in LEs (Vermeer et al.,
2009).
PtdIns4P is produced from PtdIns by PtdIns4-kinases
(PtdIns4K or PI4K). Two mammalian PtdIns4K isoforms,
PI4KIIα and PI4KIIIβ, the latter interacting with Arf1, localize
to the Golgi apparatus, together with PtdIns4P-binding
proteins (Wang et al., 2003). In metazoans, PtdIns4P and DAG
work in concert in TPC formation (Bard & Malhotra, 2006).
The conical shape of DAG favours membrane bilayer curvature,
contributing to vesicle initiation (Szule et al., 2002; Fig. 2).
The only two known Arabidopsis PtdIns4-kinases, AtPI4Kβ1
and AtPI4Kβ2, belong to class IIIβ. Both interact with
RabA4b, a GTPase controlling post-Golgi to plasmalemma
trafficking in root hair tips. A double mutant lacking both
genes exhibits reduced TPC formation, resulting in enlarged
vacuoles and aberrant root hair growth (Preuss et al., 2006).
Secretory proteins accumulate in the Golgi apparatus in cells
expressing mutant rice RabGAP; this effect is relieved by
RabA but not RabE overproduction. RabGAP thus may facilitate trafficking from the TGN both to the plasmalemma and
the vacuole by increasing RabA recycling (Heo et al., 2005).
Efficient PtdIns4K recruitment to the Golgi may require this
recycling. An Arabidopsis root hair mutant (rhd4) exhibiting
enlarged Golgi cisternae and major polarity defects carries a
mutation in a PtdIns4P phosphatase, again suggesting an
essential role for PtdIns4P turnover in Golgi organization and
TPC formation (Thole et al., 2008). Thus, PtdInsP metabolism at the TGN may have a prominent role in plant TPC
formation, with RabA4b serving as a major organizer of the
vesicle formation domain. Related mechanisms may participate
in the formation of endosome-derived exocytotic vesicles.
Loss of COW1 and SFH1, Arabidopsis homologues of the
yeast phosphatidyl transfer protein Sec14p, disrupts root hair
elongation, with disorganization of tip-directed PtdIns(4,5)P2,
perturbation of the cytoskeleton and dispersal of secretory
vesicles from the tip cytoplasm (Böhme et al., 2004; Vincent
et al., 2005). The mammalian Sec14 homologue Nir2 participates in secretory vesicle formation via the regulation of
DAG metabolism at the Golgi apparatus (Litvak et al., 2005,
Bard & Malhotra, 2006; Fig. 2).
Phosphatidic acid (PA), produced by phospholipase D (PLD),
may be another regulator of plant TPC formation – directly
or as a precursor for DAG production by PA phosphatase.
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Fig. 2 Role of lipid metabolism in exocytotic vesicle formation.
(a) Hypothetical mechanism of trans-Golgi network (TGN)-to-plasma
membrane carrier (TPC) formation, based on the mammalian model.
Differences in the molecular shapes between phosphatidylinositol-4phosphate (PtdIns4P) (formed by RabA4b-recruited type III
phosphatidylinositol-4-kinases) and diacylglycerol (DAG) [formed
from phosphatidic acid (PA), produced by phospholipase D (PLD)],
and their phase separation within the plane of the membrane, aided
by Sec14-related proteins and flippases, supports curvature
formation and vesicle initiation. Dynamin might perform final
scission. (b) Inhibition of PLD by n-butanol inhibits TPC production in
tobacco pollen tube tips (electron micrograph by Martin Potocký,
Mieke Wolters-Arts and Jan Derksen, Radbout University, Nijmegen,
the Netherlands). Bars, 2 µm.
PA stimulates pollen tube growth, whereas PLD inhibition
by n-butanol inhibits it, causing a loss of vesicles from the clear
zone (Potocký et al., 2003; Fig. 2). Butanol also induces the
release of the GTPase Arf1 from Golgi membranes in tissue
culture cells (Langhans & Robinson, 2007). 4,4-Difluoro-5,7dimethyl-4-bora-3a,4a-diaza-S-indacene (BODIPY® FL)labelled PA localized, on prolonged exposure, into Golgi bodies
in pollen tubes (Potocký et al., 2003). Arabidopsis mammalianlike PLD isoforms participate in the cycling of the PIN2 auxin
transporter (Li & Xue, 2007), vesicle-mediated auxin transport
(Mancuso et al., 2007) and root hair development (Ohashi
et al., 2003).
The translocation of phospholipids between sheets of the
membrane bilayer by lipid flippases may participate in vesicle
formation, as documented for the yeast Drs2p enzyme (D.
Liu et al., 2008). A functionally related Arabidopsis P4-ATPase/
flippase, ALA3, localizes to the Golgi apparatus and participates
in secretory vesicle formation (Poulsen et al., 2008; Fig. 2).
© The Authors (2009)
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Tansley review
In metazoans, vesiculation of the whole Golgi complex
takes place during mitosis; it can also be induced by ilimaquinone (Bard & Malhotra, 2006). This process, as well as TPC
formation, requires trimeric G-proteins and a serine/threonine protein kinase D (PKD) interacting with DAG membrane
domains (Bard & Malhotra, 2006). However, plant cells
retain the Golgi apparatus throughout their cell cycle and
apparently lack this mechanism, as we found no PKD homologues in plant sequence searches. Other lipid-modifying
activities might contribute to plant TPC formation.
Membrane tube pulling by membrane-anchored microtubule motor kinesin results in the tube breaking into vesicles
(Roux et al., 2006; ‘pull and cut’ model of Bard & Malhotra,
2006). The presence of dynamin and GTP was sufficient to
achieve fragmentation of artificial tubular membrane protrusions into 60–80 nm vesicles (Bashkirov et al., 2008). Plant
dynamin-related proteins (DRPs), known to participate in
cytokinesis and endocytosis, might assist exocytotic vesicle
formation (Kang et al., 2003).
It is possible that plant TPCs might also form without the
assistance of coat proteins, consistent with the default plasmalemma destination of the bulk of exocytotic cargos. Sorting,
however, appears to act on some plasmalemma proteins at the
TGN (Bard & Malhotra, 2006). Exocytosis of the yeast chitin
synthase Chs3p involves a nonconventional coat (exomer), an
activated Arf1 GTPase and acidic phospholipids (Wang et al.,
2006). The discovery of nonconventional coats thus cannot
be excluded also for some plant TPCs.
If exocytotic vesicle-generating compartments are modelled
by DRPs and actin, consortia of nonseparated vesicles connected by short tubular necks might participate in exocytosis.
In vivo observations of larger exocytotic entities at growing
pollen tube tips using video-enhanced microscopy indeed
suggest that plant TPCs may include not only isolated vesicles
but also more complex structures (Bard & Malhotra, 2006;
Ovecka et al. 2008; M. Potocký et al., unpublished).
IV. Building the tracks for exocytotic vesicles:
cytoskeleton in exocytosis and cell polarization
Much of our understanding of cytoskeleton involvement in
exocytosis comes from studies on tip-growing cells – root hairs
or pollen tubes. In these cells, actin filaments form a fine
dynamic network in the extending tip, merging into thicker
cables along the shank of the hair or tube. As the cell ceases to
grow, cables replace the fine meshwork, suggesting a causal
role of fine actin arrays in tip growth (Baluška et al., 2000;
Ketelaar et al., 2003; Lovy-Wheeler et al., 2005), and the
perturbation of actin dynamics often results in characteristic
phenotypes (see Ren & Xiang, 2007; Cheung & Wu, 2008).
The cytoskeleton participates in long-distance organelle
and vesicle movement. In opisthokonts, motor proteins aid
Rab-assisted loading of vesicles onto microtubule or actin
tracks (Deneka et al., 2003). Similar mechanisms probably
© The Authors (2009)
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Review
operate also in plants. Arabidopsis class XI myosins participate
in organelle trafficking, with a specific myosin function
related to vesicle transport in root hair elongation (Ojangu
et al., 2007; Prokhnevsky et al., 2008). A myosin-independent
‘comet’ mechanism based on F-actin polymerization might
also participate in secretory vesicle transport; in plants, there
is circumstantial evidence for this mechanism in endosome
movement in growing root hair tips (Voigt et al., 2005).
Dynamin-mediated scission of mammalian endocytotic
vesicles requires actin polymerization and fine actin filaments
may also coat exo- and endocytotic vesicles (Tsujita et al.,
2006). Myosin-dependent transport of membrane organelles
in vivo requires continuous dynamic turnover of F-actin
(Semenova et al., 2008).
Microtubules are not essential for tip growth in budding
yeast or filamentous fungi; however, in fission yeast, they
determine the direction of tip growth via the delivery of
‘landmark’ complexes. Some of the responsible proteins have
homologues in plants (Sieberer et al., 2005). In addition, in
plant tip-growing cells, microtubules control growth direction
rather than growth itself (Bibikova et al., 1999); microtubule
disruption by oryzalin induces wavy root hair or pollen tube
growth (see Sieberer et al., 2005; Gossot & Geitmann, 2007).
Tubulin depletion causes ectopic root hair formation and
branching (Bao et al., 2001).
A reciprocal relationship between mechanical stress and
microtubule organization, long known in plants (for example,
Lintilhac, 1984), has been documented recently in fission
yeast, where microtubules induce actin reorganization on
mechanical bending (Minc et al., 2009). Mechanical induction of new secretory domains and RDs (see below) may be a
first step towards organ initiation, as demonstrated by new
auxin maxima and lateral root initiation in roots subjected to
bending (Ditengou et al., 2008).
In diffusely growing plant cells, cortical microtubules
delimit expanding areas (see Wasteneys, 2004). Microtubules
also determine the position of secretory domains in nonexpanding cells, such as Arabidopsis pectin-depositing seed coat
cells, where a dense microtubule meshwork marks domains
for pectin insertion (McFarlane et al., 2008). In addition,
de novo localization of PIN1 at the cross-walls of root cortex
cells after cytokinesis requires microtubules, whereas actin
participates in the recycling of PIN1 and in the localization of
AUX1 (Geldner et al., 2003; Boutté et al., 2006; Kleine-Vehn
et al., 2006). Actin bundling elicited by a mouse talin-derived
fusion protein in cultured tobacco cells produced a phenotype
suggesting the perturbation of auxin transport (Maisch &
Nick, 2007), and some auxin transport inhibitors perturb
actin (Dhonukshe et al., 2008a). Interestingly, at least two
Arabidopsis actin organizers from the formin family localize to
cross-walls of the root cortex (Deeks et al., 2005).
Local balance between G-actin, fine F-actin arrays and Factin cables may control exocytosis, with cortical actin acting
as a barrier preventing secretory vesicles from reaching the
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Fig. 3 The cytoskeleton participates in the delimitation of
exocytotically (and endocytotically) active plasmalemma domains
(light blue). While the cortical microtubular network (blue) is
permeable for secretory vesicles and may even direct them to the sites
of exocytosis, the fine actin mesh (red) acts as a barrier to exocytosis.
plasmalemma and thus delimiting the secretory domain
(as proposed for metazoan cells by Valentijn et al., 1999),
whereas microtubules may provide a vesicle-permeable framework at the sites of exocytosis or endocytosis in plant cells
(Karahara et al., 2009; McFarlane et al., 2008; Fig. 3). The
concerted action of Rop GTPases (and effectors), microfilaments and microtubules defining plant epidermal cell lobes
(Fu et al., 2005) is consistent with this model, which might
represent an evolutionarily conserved polarity regulation
module.
V. Manysidedness of a plant cell: delimiting
functional plasmalemma domains within a cell
A mere look into a microscopic section of plant tissue suggests
the presence of multiple exocytotic domains in a single cell.
For example, a leaf epidermal cell has a ‘distal’ plasmalemma
domain oriented towards the environment and a ‘proximal’
domain facing palisade parenchyma cells. Tip-growing cells
exhibit dramatic surface diversity, with a pectinaceous wall
at the growing tip and a callose-rich subapical nongrowing
domain. A cell attacked by a fungal pathogen re-polarizes to
form a callose, lignin and suberin-rich cell wall papilla
adjacent to the invading appressorium (Schmelzer, 2002).
Local differences in cell wall and plasmalemma protein
composition have been well documented by immunodetection
techniques (see, for example, Knox, 2008). Examples include
the polarized localization of PIN and AUX1 carriers, glycosylphosphatidylinositol (GPI)-anchored protein COBRA and
cuticle protein BODYGUARD (see, for example, Schindelman et al., 2001; Kurdyukov et al., 2006; Wisniewska et al.,
2006). We use the term ‘activated cortical domain’ (ACD) to
denote plasmalemma domains poised to or actually performing exocytosis (and endocytosis), not necessarily related to cell
growth (Fig. 4).
1. Rop guanosine triphosphatases and related
signalling pathways in the definition of the activated
cortical domain
Local activation of Rho family GTPases (Rac, Rho or Cdc42
in opisthokonts, Rop in plants), organizing both the
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Fig. 4 Schematic view of activated cortical domain (ACD)
establishment. The activation of Rop guanosine triphosphatase
(GTPases), for example, by receptor-like serine/threonine kinases
(RLKs) via a plant-specific Rop nucleotide exchanger protein
(PRONE-GEF) is central to the assembly of an ACD. Rop controls
the local activity of lipid-modifying enzymes [which generate a
phosphatidylinositol-4,5-bisphosphate (PtdIns(4,5)P2)-enriched
membrane domain that can be recognized by the Exo70 exocyst
subunit], mediates the assembly of the exocyst via the effector Icr1
interacting with Sec3, and controls cytoskeletal organization.
Inhibition of Rop, for example, by GTPase-activating proteins (GAPs)
or GDP dissociation inhibitors (GDIs, GEFs), and lipid modification by
phospholipase C (PLC), restricts lateral spreading of the secretory
domain. Activated Rops and RLKs are sequestered into lipid rafts,
thus promoting signalling interactions. Expansin- and xyloglucan
endotransglycosylase (XET)-mediated cell wall loosening takes place
adjacent to the ACD. PM, plasma membrane.
cytoskeleton and the secretory pathway, is central to
eukaryotic cell polarization; unfortunately, we know little
about plant Rop effectors and regulators (see Ridley, 2006;
Kost, 2008; Yalovsky et al., 2008). Receptor-like serine/
threonine kinases (RLKs), a large family of versatile signalling
proteins whose localization, at least in some cases, requires
Rop-regulated exocytosis (Lee et al., 2008), and a novel class
of Rop GEFs (PRONE-GEF; Berken et al., 2005), which are
activated by RLKs (Zhang & McCormick, 2007), are prime
candidates for local regulators of Rop activity and ACD
formation. The multiplicity of angiosperm RLKs may
provide a means for the formation of diverse ACDs during
development or in response to external signals, including
interactions with pathogens. The recently discovered adaptor
protein ICR1, which interacts with GTP-charged Rop and
is able to recruit the exocyst subunit Sec3 (see below), is
important for ACD initiation (Lavy et al., 2007); the same
protein is recruited into pollen tube tips by a Rop-dependent
mechanism (S. Li et al., 2008; Fig. 4).
The overexpression of Rops in tip-growing cells often
causes tip depolarization (for example, Fu et al., 2001;
Cheung et al., 2003). Wild-type, constitutively active and
dominant-negative Rop mutants all elicit tip swelling to a
varying extent, and some Rops inhibit tip growth when
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overexpressed (Kost, 2008; Yalovsky et al., 2008). Microinjected nonhydrolysable analogues of GTP or GDP cause
swelling in pollen tubes (Eliáš et al., 2001). This suggests
either a requirement for Rop cycling between the GTP and
GDP-bound states, or involvement of multiple pathways
(perhaps employing different Rops or Rop effectors) in the
maintenance of tip-focused growth.
Negative regulators of Rop may control the spread of
ACDs. Mutation of the Arabidopsis Rop-associated GDP dissociation inhibitor (GDI) SCN1 results in multiple, short,
swollen root hairs per trichoblast (Carol et al., 2005), and the
loss of either GDI or a GTPase-activating protein (GAP)
causes pollen tube swelling (Klahre & Kost, 2006; Klahre
et al., 2006).
Metazoan Rhos induce actin cables in stress fibres, filopodia
and lamellipodia, partly by direct binding to formins (see
Ridley, 2006). However, angiosperm formins contain no
known GTPase-interacting motifs, although moss and
lycophyte formins possess a candidate Rho-binding domain
(Grunt et al., 2008). Thus, plant Rops may control actin indirectly or via other effectors, such as SCAR/WAVE proteins
regulating Arp2/3-mediated actin nucleation (see Deeks &
Hussey, 2005; Mathur, 2005) or a PtdIns-P kinase (Kost et al.,
1999, see below). The first known Rop effectors were the
small CRIB domain-containing RIC proteins (Wu et al., 2001).
Arabidopsis RIC3 and RIC4 are responsible for ‘translating’
oscillations in apical [Ca2+] into periodical growth rate
changes, with the accumulation of secretory vesicles on RIC4driven F-actin assembly and their simultaneous discharge on
RIC3-induced calcium-regulated actin disassembly (Gu et al.,
2005; Lee et al., 2008).
The Rop/RIC module is also involved in diffuse cell expansion (Panteris & Galatis, 2005). AtROP2 controls the development of lobed epidermal pavement cells through two
complementary pathways, one stimulating actin assembly via
RIC4 in outgrowing lobes and the other inducing RIC1dependent microtubule reorganization in indentations of the
adjacent cell, whilst locally inhibiting ROP2 (Fu et al., 2005).
Antagonistically acting RICs may partly explain the diversity
of Rop-associated phenotypes.
Local cell wall relaxation [involving expansins or xyloglucan
endotransglycosylases (XETs)] might serve as an example of
mechanical stress inducing new ACDs, resulting in the initiation of root hair bulges on rhizodermal trichoblasts or ectopic
leaf initiation at the apical meristem (Reinhardt et al., 1998;
Baluška et al., 2000; Vissenberg et al., 2001).
2. The role of membrane lipid composition and
modifications
Eukaryotic plasma membranes exhibit lateral heterogeneity
based on cholesterol-enriched lipid microdomains (detergentresistant membranes or rafts) organized by membrane
proteins (Salaun et al., 2004; Grossmann et al., 2007).
© The Authors (2009)
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Review
Arabidopsis sterol mutants indicate the involvement of
membrane sterols in endocytosis, exocytosis and cell polarization (Souter et al., 2002; Willemsen et al., 2003; Men et al.,
2008). GTP-bound type I Rops are sequestered into
detergent-resistant membranes as a result of hydrophobic
modification by reversible covalent binding of palmitic or
stearic acids in addition to standard C-terminal geranylation
(Sorek et al., 2007; Yalovsky et al., 2008).
GPI-anchored proteins, including plant cell wall arabinogalactan proteins (AGPs, Morel et al., 2006), are characteristic inhabitants of lipid microdomains. AGPs might be
responsible for the lateral differentiation of plant plasmalemma,
partly via the anchoring of detergent-resistant microdomains
to the cell wall. Indeed, some plant membrane microdomains
are nonmotile, as shown for the KAT1/SYP121 markers (Sutter
et al., 2007; Grefen & Blatt, 2008). This may contribute to
plasmalemma domain specification, as illustrated by the
distinct localization of the COBRA GPI protein to the lateral
cell wall domain, required for correct cell expansion (Schindelman et al., 2001). However, not all plant GPIs are polarized;
the Arabidopsis SKU5 protein covers more or less evenly root
cell surfaces, yet mutants suffer from root polarity defects
(Sedbrook et al., 2002). Thus, even the plasmalemma of a single
cell may harbour diverse membrane microdomains which
may combine to form distinct, but possibly overlapping, plasmalemma sectors or ACDs with different recycling kinetics
(documented for yeast by Grossmann et al., 2007). As Rops,
RLKs, SNAREs, ion transport channels, GPIs and other
proteins involved in exocytosis and polarity signalling are all
enriched in plant detergent-resistant membranes (Morel
et al., 2006), these membrane microdomains may facilitate
crowding of cell polarity regulators and enhance their mutual
interactions (Yalovsky et al., 2008; Fig. 4).
Phosphoinositides (PPIs) also act as local membrane
domain organizers and regulators. PPI metabolism is localized,
with various kinases and phosphatases active at distinct compartments (see Thole & Nielsen, 2008). PPIs bind to specific
protein motifs (for example, PH, PX, FYVE, etc.) and additional binding sites are often required to engage membraneresident proteins (Lemmon, 2008). In tip-growing plant cells,
PtdIns(4,5)P2 is localized to the apex and spreads throughout
the plasma membrane on growth cessation (for example, Kost
et al., 1999; Vincent et al., 2005; Ischebeck et al., 2008; Kusano
et al., 2008; Stenzel et al., 2008). PtdIns4,5-kinase isoform 3
(PtdIns4P-5K3), a key enzyme producing PtdIns(4,5)P2, is
preferentially expressed in root hairs, and localized to the
periphery of their apex, possibly associated with the plasmalemma or exocytotic vesicles. Mutants exhibit reduced root
hair growth, whereas overexpression leads to multiple hairs
per trichoblast and the loss of cell polarity, i.e. disturbances of
ACD regulation (Kusano et al., 2008; Stenzel et al., 2008).
Analogous observations were made for PtdIns4P-5K4/5 in
pollen, whose loss inhibits pollen germination and pollen
tube growth (Ischebeck et al., 2008; Sousa et al., 2008). In
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growing pollen tubes, both proteins localize to a distinct tip
plasmalemma domain, overlapping with the distribution of
PtdIns(4,5)P2; overexpression induces tip branching and
large plasma membrane invaginations, again suggesting a
direct role of PtdIns4P-5Ks in ACD regulation. Negative
regulators are important to restrict PtdIns(4,5)P2 to a distinct
plasmalemma domain. In pollen tubes, this is achieved by the
localization of PtdIns(4,5)P2-consuming PLC to the border
of the ACD just behind the growing tip (Dowd et al., 2006;
Helling et al., 2006). Plants have multiple PtdIns-P kinases
producing PtdIns(4,5)P2 (five isoforms are co-expressed in
Arabidopsis pollen; Žárský et al., 2006; Ischebeck et al., 2008).
Mutant phenotypes point to a differential effect, implying the
existence of different PtdIns(4,5)P2 pools (Ischebeck et al.,
2008; Sousa et al., 2008). Multiple PtdIns(4,5)P2 pools with
distinctive fatty acids in the same cell were observed on stress
treatment (König et al., 2008), implying their possible participation in defining distinct ACDs (see Section VII).
So far, we have treated ACD almost as an isolated plasmalemma domain. However, the establishment and function of
an exocytotically active ACD involves dynamic membrane
recycling. In Section VII, we propose a general concept of a
‘recycling domain’ (RD) as a dynamic entity connecting
the ACD with a specific subset of recycling compartments/
endosomes.
VI. Vesicle tethering, docking and fusion
The ACD may be viewed as a docking platform for exocytotic
vesicles. In opisthokonts, distinct phases of vesicle tethering
and docking at the target membrane, preceding SNARE
complex formation and vesicle fusion, have been described.
Tethering links the vesicle at a distance of more than one-half
of its diameter from the target membrane, whereas docking
sensu stricto holds the two membranes within a bilayer’s
distance (< 5–10 nm). In plants, such resolution has not yet
been achieved; therefore we shall refer to both tethering and
docking as tethering sensu lato. The last phase of vesicle fusion
has been exhaustively reviewed recently, focusing especially on
SNARE function (for example, Lipka et al., 2007).
Vesicle tethering is an important targeting step, mediated
either by long tethering proteins or by multisubunit tethering
complexes (Cai et al., 2007). In opisthokonts, exocytotic
vesicle tethering involves the octameric exocyst complex, originally discovered in yeast as effector of the Rab GTPase Sec4
(TerBush et al., 1996), and later found also in animals (for a
review, see Hsu et al., 2004; Cai et al., 2007; Wu et al., 2008).
The exocyst localizes exocytotic vesicles to specific plasmalemma domains (ACDs), facilitating early events of polarized
secretion. We have recently described the angiosperm exocyst
complex using genetic, biochemical and cytological methods
(Hála et al., 2008), confirming the presence of homologues of
all eight subunits (Sec3, Sec5, Sec6, Sec8, Sec10, Sec15,
Exo70, Exo84). Surprisingly, Exo70 forms an extremely large
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family of paralogues in land plants (Eliáš et al., 2003; Synek
et al., 2006). T-DNA insertions into the Exo70A1 gene,
encoding the most abundant Exo70 isoform among the 23
Arabidopsis paralogues, resulted in a discernible phenotype
with a semi-dwarf stature, reduced apical dominance, failure
of polar growth of root hairs and stigmatic papillae, and
delayed senescence and lateral root initiation (Synek et al.,
2006). Mutants in SEC5, SEC6, SEC8 and SEC15a show
defective pollen germination and pollen tube growth, and
Sec6, Sec8 and Exo70A1 colocalize at growing tobacco pollen
tube tips (Cole et al., 2005; Hála et al., 2008). A maize
mutant (rth1) in the Sec3 subunit was discovered independently in a forward screen for root hair defects (Wen et al.,
2005). All of these observations strongly implicate the plant
exocyst in cell polarity and morphogenesis regulation.
Although the mechanism of exocyst action remains somewhat unclear (for example, Wu et al., 2008; Songer & Munson,
2009), the Exo70 and Sec3 subunits obviously provide a
crucial activated Rho-dependent landmark for assembling the
rest of the complex on the plasmalemma (Boyd et al., 2004).
The complete exocyst probably assembles on arrival of the
remaining subunits carried by vesicles to the target membrane
marked by the Exo70/Sec3 landmark, which is delivered to
the ACD independently from other exocyst subunits and
possibly also from actin (Novick et al., 2006). However, an
alternative model has been proposed, involving the local activation of preassembled exocyst by the Rho GTPase (Wu et al.,
2008). In mammals, insulin-induced delivery of the GLUT4
transporter to the plasmalemma depends on binding between
Exo70 and Rho GTPase TC10 (Inoue et al., 2003).
PtdIns(4,5)P2 also participates in vesicle tethering, in
particular in attaching Exo70 and Sec3 to the plasmalemma.
Binding of Exo70 to PtdIns(4,5)P2 is confined to a motif at
the C-terminus of the rod-like molecule (He et al., 2007b),
conserved in many, but not all, Arabidopsis Exo70 paralogues
(Fig. 5). In addition, yeast Sec3 binds to PtdIns(4,5)P2 in the
membrane (Zhang et al., 2008), but its PPI-binding site is
located in the nonconserved N-terminal part of the protein
and may thus represent a lineage-specific feature. In yeast,
genetic evidence suggests that Sec8p, Sec10p and Sec15p
exocyst subunits and the Sec9p plasmalemma t-SNARE are
regulated by PtdIns(4,5)P2 (Routt et al., 2005). Although, in
opisthokonts, activated Rho interacts directly with exocyst
subunits (Guo et al., 2001; Wu et al., 2008), in Arabidopsis,
the Rop–Sec3 interaction appears to be mediated by the adaptor
protein ICR1 (see above, Lavy et al., 2007). We have observed
interaction between Arabidopsis Sec3 and Exo70A1 subunits
(Hála et al., 2008), which, together with the predicted ability
of Exo70A1 (and some other Exo70 paralogues) to bind
membrane phospholipids, suggests that plant Exo70s might
serve as landmarks for exocyst vesicle targeting, coordinated
with local Rop activity via Sec3–ICR1 interactions (Fig. 4).
In multicellular Characean algae, or in the first land plants,
genes encoding Exo70 underwent a series of duplications,
© The Authors (2009)
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Review
Fig. 6 Hypotheses on possible primary functional diversification of
land plant Exo70 paralogues. The three assumed Exo70 paralogues in
ancestral plants may have divided their function either among three
spatial directions (or planes of division) within the multicellular body
– basal–apical, radial and tangential (a), or between three kinds of cell
polarity phenomena – tip growth, nonisodiametric cell expansion and
protein deposition, and cytokinesis (b). Distinct Exo70 paralogues
may be responsible for maintaining multiple recycling domains per
cell (c).
Fig. 5 Plant Exo70 isoforms harbour C-terminal conserved
PtdIns4,5P2 binding (red) and Arp2/3 complex binding (blue) sites.
(a) Structural model produced by threading of the major
housekeeping Arabidopsis isoform, AtExo70A1, on the empirically
determined structure of mouse Exo70 (PDB:2pft) shows that the
binding sites nearly overlap. (b) PROSITE pattern representation of
the same binding motifs in selected yeast and metazoan Exo70
proteins and all Arabidopsis Exo70 isoforms reveals a diversity of
presumed binding abilities among the plant paralogues (GenBank
accessions: yeast, ScExo70, NP_012450.1; rat, RnExo70,
NP_073182.1; Drosophila, DmExo70, NP_648222.3; Arabidopsis –
AtExo70A1, NP_001119162.1; AtExo70A2, NP_200047.3;
AtExo70A3, NP_200048.2; AtExo70B1, NP_200651.1; AtExo70B2,
NP_172181.1; AtExo70C1, NP_196819.1; AtExo70C2,
NP_196903.1; AtExo70D1, NP_177391.1; AtExo70D2,
NP_175811.1; AtExo70D3, NP_566477.2; AtExo70E1,
NP_189586.1; AtExo70E, NP_200909.1; AtExo70F1, NP_199849.2;
AtExo70G1, NP_194882.2; AtExo70G2, NP_175575.1; AtExo70H1,
NP_191075.2; AtExo70H2, NP_181470.1; AtExo70H3,
NP_187564.1; AtExo70H4, NP_187563.1; AtExo70H5,
NP_180432.2; AtExo70H6, NP_683286.2; AtExo70H7,
NP_200781.1; AtExo70H8, NP_180433.1).
© The Authors (2009)
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generating three major Exo70 families found in all land plants
beginning with mosses (Eliáš et al., 2003, Synek et al., 2006).
Multiple Exo70 isoforms within the same cell may contribute
to specific landmarks, and thus to the initiation and maintenance of specific ACDs and RDs (Fig. 6).
Yeast Exo70 participates in the regulation of actin dynamics
by interaction with the Arp2/3 nucleation complex via a motif
almost overlapping with the PtdIns4,5P2-binding site (Zuo
et al., 2006). Among Arabidopsis Exo70 paralogues, only
Exo70A1 exhibits a perfect match to the mammalian Arp2/3binding motif (Fig. 5). The diversity of C-terminal-binding
motifs in plant Exo70 proteins suggests diverse protein or
lipid interactions (Fig. 5).
As the exocyst is currently the only plant candidate for
vesicle tethering at the plasmalemma, we suggest that Exo70
paralogues may, apart from cell type-specific functions, create
a variety of exocyst complexes within the same cell, poised to
specific ACDs or endomembrane destinations (Fig. 6).
Together with other factors, such as a variety of GTPases,
lipids, formins or SNAREs, this may be central for the establishment of the dynamic manysidedness of the plant cell.
In opisthokonts, vesicle tethering, followed by activation
and formation of the SNARE complex, involves a Rab
GTPase cascade. The yeast exocytotic Rab GTPase Sec4 is
activated by GDP/GTP exchange, aided by the Sec2 GEF
that is recruited by the upstream Golgi-based Rab GTPase
Ypt31/32 (Novick et al., 2006). Active Sec4 recruits the
exocyst by interacting with its Sec15 subunit, which also
transiently binds Sec2 (Novick et al., 2006). Sec4 and Exo84
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then attract Sro7/77, a WD40 family protein implicated in
maintaining cell polarity, which, in turn, binds and regulates
the t-SNARE Sec9p (Lehman et al., 1999). Arabidopsis has
two putative Sro7 homologues, At5g07860 and At3g35560,
both carrying N-terminal syntaxin-binding WD40 repeats
and C-terminal R-SNARE domains, suggesting that they
bind SNARE proteins. It is not known yet whether they also
interact with Rab GTPases. The yeast Exo84 exocyst subunit
binds to Sec1, whose interaction with the SNARE complex
initiates membrane fusion (Novick et al., 2006). The Arabidopsis Sec1 homologue KEULE is involved in cytokinesis
(Assaad et al., 2001). Exocyst-like structures in plant cytokinesis have been observed (Otegui & Staehelin, 2004;
Seguí-Simmaro et al., 2004), and phenotypes of Arabidopsis
exo70a1 mutants also implicate the exocyst in meristem cell
division regulation (Synek et al., 2006).
Lipids participate in both vesicle fission (see above) and
fusion. PA may promote membrane fusion by influencing
membrane topology. Yeast PLD, encoded by Spo14p, is
required for prospore membrane formation, and PA controls
the localization of the SNAP25-like t-SNARE Spo20p during
sporulation (Liu et al., 2007). PA production by Spo14p is
also activated by the nonclassical PtdIns transfer protein
Sfh5p, which, in turn, promotes PtdIns(4,5)P2 production at
the plasmalemma (Routt et al., 2005). Moreover, in animal cells,
PA stimulates membrane fusion governed by the t-SNARE
complexes syntaxin 1A/SNAP25 and syntaxin 4/SNAP23
(Vicogne et al., 2006; Lam et al., 2008). PLD activation by
the small GTPase ARF6 stimulates PtdIns(4,5)P2 production
and exocytosis (Béglé et al., 2009).
In opisthokonts, PtdIns(4,5)P2 is crucial for both ATPdependent vesicle priming and Ca2+-dependent fusion during
regulated exocytosis. This involves the binding of PtdIns(4,5)P2
to several proteins, including SCAMP2, SNAP25/syntaxin
1A, exophilin 4 and synaptotagmins, which all have homologues in plants. Arabidopsis AtSYT1, a plasmalemma-localized
synaptotagmin homologue, is indispensable for maintaining
plasmalemma integrity under salt or freezing stress, presumably
through exocytotic membrane resealing (Schapire et al., 2008).
The role of [Ca2+]i gradients in plant exocytosis, and especially
in tip growth, has been the subject of many reviews (for example,
Hepler, 2005). As in metazoan neurones, in plant cells, calcium
has been suggested to regulate the switch from constitutive to
regulated exocytosis (see Homann, 2006). Calcium-dependent
candidates for plant exocytosis regulators include calmodulin,
calcium-dependent protein kinases (CDPKs), NADPH
oxidases (NOX), actin-binding proteins (see Cole & Fowler,
2006), and annexins and synaptotagmins (Mortimer et al.,
2008, Schapire et al., 2008). Dynamic proton transport
loops at the tip of pollen tubes may play a prominent regulatory role upstream of calcium (Certal et al., 2008).
In tip-growing cells, a tip-focused calcium gradient may be
maintained by a positive feedback loop involving calcium
operating at the activated exocytotic domain: the Rop-activated
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NOX catalyses reactive oxygen species (ROS) production,
which then stimulates Ca2+ influx carriers, bringing about
calcium-induced NOX activity (Carol et al., 2005; Žárský
et al., 2006; Potocký et al., 2007; Takeda et al., 2008).
Membrane capacitance measurements have shown that
calcium-independent vesicle fusion also takes place at the
plant plasmalemma (Homann, 2006). Different dependences
on calcium for vesicle fusion may thus contribute to the diversity of secretory pathways within a single plant cell.
VII. Polarity by recycling: exocytosis,
endocytosis and recycling domains in plant cells
In opisthokonts, secretion restricted to specific plasmalemma
domains is characteristic, for example, of yeast buds or the
apical and basolateral membranes in animal epithelia (Mostov
et al., 2003). Such polarized exocytosis has also been well
documented in plants. New cell wall components are inserted
into specific cortical domains during cell expansion and
secondary cell wall deposition and modification, as
exemplified by differential cell wall thickening in colenchyma,
xylem maturation, Caspari band formation or pectin
deposition in seed coat volcano cells. Cytokinesis and tip
growth can be viewed as extreme examples of direct polarized
cargo delivery as well, although they obviously also involve
membrane recycling.
Most (but not all) plasmalemma constituents are continuously recycled even in fully differentiated plant cells. Fast and
slow recycling takes place in plant cells, for example, via
‘kiss-and-run’ vesicles with a half-time of approximately 1 s
and via clathrin-coated vesicles lasting c. 50 s (Homann, 2006).
In metazoan synapses, an increase in [Ca2+]i correlates with
the switch between fast and slow recycling; this might be
similar in plant cells.
Dynamic recycling of plant membrane proteins has
received attention after the discovery of the constitutive recycling of PIN auxin efflux carriers, which is directly regulated by
auxin itself (Geldner et al., 2003; Paciorek et al., 2005).
Meckel et al. (2004) observed constitutive internalization and
recycling of vesicles carrying the KAT1 potassium channel,
with kinetics quite different from PIN recycling, whereas
a heterologously expressed GFP-tagged human membrane
protein was not recycled, pointing to lateral differentiation of
the plasmalemma with respect to selective endocytotic retrieval
of specific membrane proteins (Homann et al., 2007). Abscisic
acid (ABA) triggers specific KAT1 internalization with slow
recycling back to the plasma membrane (Sutter et al., 2007).
The effects of auxin and ABA on the kinetics of endocytosis
suggest that endocytosis is a regulated step in plant RD
dynamics; it is expected that recycling will also be regulated at
the level of exocytosis (see below).
The polarization of plasmalemma proteins is intimately
linked to endocytotic membrane recycling, as shown by
Valdez-Taubas & Pelham (2003), who recognized two basic
© The Authors (2009)
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Tansley review
Fig. 7 Cell polarization by recycling. (a) A membrane protein
deposited in a polar manner stays polar if it is either prevented from
lateral diffusion by a barrier (possibly anchored to a stable structure,
such as the cell wall) or removed by endocytosis [and possibly
recycled back to the activated cortical domain (ACD)]. (b) A plant cell
may comprise multiple recycling domains (RDs), whose topology is
also influenced by cytoplasmic streaming, the position of the nucleus
and the vacuole. (c) In late interphase, the preprophase band of
microtubules (blue) initiates an ACD of local maximum of endocytosis
at the plasmalemma. This dominant endocytosis belt might establish
two new ‘global’ RDs comprising future daughter cells.
mechanisms of cell polarity. Different plasmalemma domains
may be laterally separated by diffusion barriers, such as, for
example, the septin ring at the neck between bud and mother
cell in yeast. Alternatively, dynamic polarization may be based
on slow lateral membrane protein diffusion (in cells with a cell
wall) and local membrane recycling via an endocytosis/exocytosis cycle (Fig. 7). Even in yeast, the second mechanism is
crucial. Yeast SNARE Snc1 is kinetically polarized by endocytosis, as the mutation of an endocytotic signal results in its
even distribution on the cell surface, whereas the mere addition of an endocytosis sorting motif to the Sso1 SNARE
results in its polarization (Valdez-Taubas & Pelham, 2003).
Studies on pollen tubes, root hairs, the KAT1 channel and
auxin carrier recycling show that this type of plasmalemma
polarization is also dominant in plant cells (Paciorek et al.,
2005; Cole & Fowler, 2006; Sutter et al., 2007; Kost, 2008;
Yalovsky et al., 2008).
The establishment and maintenance of different plasmalemma domains of the same plant cell are largely dependent on
kinetic polarization via recycling. This also implies the multiplicity of RE compartments – a feature already observed for
some membrane proteins related to auxin transport (see
Section II). ACDs at the plasmalemma and related recycling
compartment are inseparably dependent on each others’
dynamics, thus establishing a dynamic superstructure which
we propose to call the ‘recycling domain’ (RD).
The RD is a dynamic spatiotemporal entity based on the
continuous reciprocal exchange of specific membrane constituents between distinct ACDs of the plasmalemma and specific
compartments of the endomembrane system. A typical
manysided plant cell embedded in a multicellular tissue has
multiple RDs.
© The Authors (2009)
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Review
The best studied examples of plant RDs are those involved
in the dynamic polarization of auxin carriers. Tip-growing
cells provide an extreme example of a highly polarized RD
(including cytoskeletal structures), in fungi described as the
tip growth apparatus (TGA; Heath & Geitmann, 2000). We
prefer to use this term also for the polarity machinery recently
described as LENS (Cole & Fowler, 2006) in pollen tubes or
root hairs to acknowledge convergent similarities between
fungal and plant tip growth.
RD equilibrium is sensitive to subtle changes in the exocytosis/endocytosis ratio, affected by diverse signalling pathways.
The inhibition of endocytosis by activated Rop (Bloch et al.,
2005) and auxin (Paciorek et al., 2005) has been described. In
addition, in metazoans, activated Rho GTPases inhibit endocytosis (see Ridley, 2006). The effects of auxin and Rop on
endocytosis might be linked through the activation of Rop by
auxin via the RLKs-PRONE GEF pathway (Žárský & Fowler,
2009). Opposite to auxin, ABA stimulates endocytosis of the
KAT1 channel quite selectively (Sutter et al., 2007); membrane
protein fluxes within RDs thus might be regulated mainly at
the level of endocytosis.
There may be major differences between regulatory
networks governing the dynamics of RDs in growing vs
nongrowing cells. As a result of the lateral differentiation of
the plasmalemma, linked to lipid microdomain formation,
RDs with very different kinetics might partly topologically
overlap, using alternative endocytotic and recycling routes.
A good example of endocytosis-dependent plant plasmalemma protein polarization has been published recently by
Dhonukshe et al. (2008b), who demonstrated that newly
synthesized PIN1 is first localized all over the plasmalemma in
Arabidopsis root stele cells, and only subsequently polarizes via
recycling through a late endosomal compartment. This observation pinpoints the important distinction between RD
establishment and RD maintenance; in this case, the former
requires endocytosis and MVB/LE-dependent transcytosis,
but as soon as polar localization is established, it is maintained
via GNOM RE recycling (Geldner et al., 2003; Dhonukshe
et al., 2008b; Kleine-Vehn et al., 2008).
Continuous endosome (and actin)-dependent recycling,
maintaining the polar localization of PIN (and partly also
AUX1) auxin transporters, endows the system with sensitivity
and adaptivity to external cues, explaining the almost immediate relocalization of PIN transporters on gravistimulation of
roots and a concomitant shift in the ACD position (Ottenschläger et al., 2003). Polarization of the PIN2 auxin carrier is
regulated by sterol-dependent endocytosis (Men et al., 2008),
underpinning again RD-based recycling as the background of
many polarization phenomena in plant cells, as well as the role
of lipid rafts in RD establishment. The relocalization of PIN2
protein to the opposite cell domain on prolonged brefeldin
A inhibition of RE function shows that the disruption of
membrane recycling may result in reversible artificial RD
formation (Kleine-Vehn et al., 2008).
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New secretory domain and RD initiation in response to
pathogen attack provides insights into plant ACD and RD
establishment (Schmelzer, 2002). This process is possibly
initiated by Rop GTPase activation at the attack site, with a
subsequent increase in actin dynamics (Schütz et al., 2006).
The nucleus moves towards the nascent RD, i.e. to a new pole
of cell polarity (Schmelzer, 2002). Similarly, the nucleus also
acts as an organizer in the regulation of polar cell expansion
in root hairs, where optical trapping of the nucleus inhibits
tip growth. In the root hair branching mutant cow1-2, the
nucleus periodically migrates between the tips of the branched
hair. Tip growth activity alternates between these two tips,
dependent on the proximity of the nucleus; nuclear dynamics
in this cell type are exclusively dependent on the actin
cytoskeleton (Ketelaar et al., 2002). Dynamic nuclear positioning within the cell thus participates in RD establishment
– as is most apparent in preprophase band and fragmoplastrelated RDs in cytokinesis (Van Damme et al., 2007). The
preprophase band of microtubules directed by nuclear position has been shown to initiate new ACD in late interphase
with a distinct maximum of endocytosis (Dhonukshe et al.,
2006; Karahara et al., 2009). We propose that this local cytoplasmic membrane belt of endocytosis maximum might be
dynamically regulated by membrane recycling via exocytosis
in two halves of the cell delineated by the nuclear position,
resulting in the formation of two global RDs encompassing
future daughter cells (Fig. 7c). Gene expression regulation is
a major source of ‘nuclear control’ above cell polarity, reaching,
however, beyond the scope of our review. In any case, nuclear
migration should result in partial polarization of de novosynthesized mRNAs.
The concept of multiple RDs in a single plant cell not only
implies the presence of several subtypes of TGN/EEs and
other endosomes (RE, LE), but, despite the known mobility of
endosomes, also demands spatiotemporal ‘subcompartmentalization’ of endosomes within the cell. This could be at least
partly organized by the position of the nucleus, the development
of the vacuole and also by domains of cytoplasmic streaming.
An active role of Golgi/TGN location in directed secretion
and regulation of cell polarity has been documented recently
in human HeLa cells under conditions specifically disrupting
Golgi positioning without causing microtubule disassembly
(Yadav et al., 2009). It should be expected that, in plant cells,
the pace and direction of cytoplasmic streaming, carrying multiple Golgi bodies and especially ‘semi-autonomous’ TGNs/
EEs/REs, will be equally important. Actomyosin-driven cytoplasmic streaming induces the hydrodynamic flow of small
molecules; more elongated/differentiated cells exhibit significantly more rapid cytoplasmic streaming (Esseling-Ozdoba
et al., 2008). New RDs are established in cells exiting meristem – during the initiation of elongation and differentiation
(for example, in the root tip transition zone where apical and
basal RDs are formed). During the cell growth phase, the
organization and dynamics of cytoplasmic streaming may
New Phytologist (2009) 183: 255–272
www.newphytologist.org
Fig. 8 Schematic representation of the recycling domain
(RD) concept. RDs are only partly synchronous, as they are
consecutively established (or switched off) during the
differentiation and elongation phases of plant cell ontogenesis.
Endosomal recycling often relies on the cooperation of several
endosomal compartments (for simplification, only one is shown).
The scheme of RD I includes reference boxes relating to the
detailed figures above.
participate in the formation of new RDs or the turning down
of existing RDs. On exit from the elongation zone and terminal
cell differentiation, RDs may become more stable.
The exocyst participates in plasmalemma recycling, as it
also localizes to RE via the GTPases Rab11/RabA, and
regulates animal cell polarity-related recycling (for example,
Langevin et al., 2005). Our current data also suggest similar
relationships for the plant exocyst (H. Toupalová et al.,
unpublished). The multiplicity of plant RabA GTPases
(Rutherford & Moore, 2002) and Exo70 subunits may
substantially contribute to the establishment and maintenance
of multiple RDs in a manysided plant cell, through both recycling
and the recognition of diverse targeting platforms (ACDs).
The single yeast Exo70 controls the secretion of specific
exocytotic vesicles (Bgl2p, see above) at the early stages of bud
growth, suggesting an important general principle that
‘different exocytotic pathways can be specifically regulated at
the target membrane as well as at the TGN’ (He et al., 2007a). The
regulation of diverse plant ACDs and exocytotic pathways may
be generally based on reciprocity between specific plasmalemma
ACDs and recycling endosomal compartments, as components
of dynamic RDs (Fig. 8).
VIII. Conclusions: constitutive or regulated
secretion in plant cells?
The eukaryotic secretory pathway is generally understood as
two different processes – constitutive secretion and regulated
secretion. Constitutive secretion is associated with ‘housekeeping’
cellular functions (for example, metazoan collagen secretion),
whereas regulated secretion features in a situation in which the
© The Authors (2009)
Journal compilation © New Phytologist (2009)
Tansley review
exocytosis of pre-existing stored vesicles is triggered by specific
signal inputs. Surprisingly, only vesicles produced in this
second pathway are commonly called ‘secretory vesicles’ (see
Lodish et al., 2004, p. 703). In this context, the secretory
pathways of yeast and plant cells are considered to be
constitutive (see, for example, Fukuda, 2008). This zoocentric
concept might be wrong, as it implies that, in yeast and plants,
secretion is just a ‘boring’ default. This, obviously, is not the
case – even in the secondarily simplified yeast cell, secretion is
highly regulated. The distinction between ‘constitutive’ and
‘regulated’ secretion is, apart from a different molecular
mechanism, based on different time scales of operation –
although the latter step of regulated secretion is triggered by
an abrupt signal on the scale of milliseconds, ‘constitutive’
secretion is regulated on a scale of seconds and minutes.
Many recently discovered examples of plant secretory pathway
regulation suggest that the concept of constitutive vs regulated
secretion does not fit the distinctive features of the plant
secretory pathway. Thoroughly studied examples of tip growth,
cytokinesis, auxin transport (including tropic responses) or
stomata regulation show that plant secretion is always regulated
– on different time scales, covering both slow and abrupt
signal regulation of vesicle exocytosis/endocytosis, often also
co-regulated by calcium, which is considered to be the major
trigger for regulated exocytosis in animals. Moreover, the
concept of constitutive vs regulated secretion is an oversimplification, as it considers fusion with the plasmalemma to be
the most important controlling step. Thus, despite the fact
that plants lack ‘secretory Rabs’ sensu Fukuda (2008), we
propose a perspective acknowledging that plant exocytosis is
regulated, with essential control units being RDs (see above).
In most plant cell types, several RDs co-exist depending on
the stage of differentiation.
In addition to testing the hypothesis that diverse Exo70
paralogues might localize to different ACDs within the same
cell, future research should focus on the mapping of connections
between distinct ACDs and specific endosomal recycling
pathways, as well as the analysis of RD differentiation in cells
leaving the meristematic stage at the onset and during elongation and differentiation. This should also include a consideration of the role of endomembrane compartment motility
and the organization of cortical microtubules and cytoplasmic
streams during different stages of cell elongation on cell polarity.
Detailed mapping of RD compartments, and the identification of endosomal pathways linked to distinct plasmalemma
proteins (both for RD establishment and maintenance), has
already begun, as exemplified by work on auxin transporters.
To solve the mystery of TPC formation, it may be necessary
to develop plant in vitro systems for vesicle formation and
fusion, analogous to those which, in combination with genetics,
were crucial in unravelling the deep secrets of the secretory
pathway in opisthokonts. Although genetic analysis of plant
secretory pathways is flourishing, we have no such in vitro
systems in plants.
© The Authors (2009)
Journal compilation © New Phytologist (2009)
Review
A systems biology approach to the study of cell polarity is
becoming more than just fashionable also in plants. Models of
auxin polar transport at the cellular level, together with mathematical models of tip-growing cells (for example, de Keijzer
et al., 2009), will certainly lead the field. Online resources for
signalling network model development are becoming available
(see Grierson & Hetherington, 2008). This approach will
facilitate both the integration of the fragmented knowledge
on cell polarity regulation, as well as the identification of
unrecognized parameters and components.
Acknowledgements
We thank Bruno Goud (Institut Curie, Paris, France) and
Marek Eliáš (Charles University, Prague, Czech Republic) for
valuable comments and suggestions. Our work was supported
by MSMT LC06034, MSM 0021620858 and GAAVIAA601110916 projects and the MSMT-KONTAKT-ME841
grant (collaboration with the group of John Fowler, Oregon
State University, Corvallis, OR, USA).
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