Download Annexins: multifunctional components of growth and adaptation

Journal of Experimental Botany, Vol. 59, No. 3, pp. 533–544, 2008
doi:10.1093/jxb/erm344 Advance Access publication 10 February, 2008
REVIEW ARTICLE
Annexins: multifunctional components of growth and
adaptation
Jennifer C. Mortimer1, Anuphon Laohavisit1, Neil Macpherson1, Alex Webb1, Colin Brownlee2,
Nicholas H. Battey3 and Julia M. Davies1,*
1
Department of Plant Sciences, University of Cambridge, Downing Street, Cambridge CB2 3EA, UK
2
Marine Biological Association, The Laboratory, Citadel Hill, Plymouth Pl1 2PB, UK
School of Biological Sciences, Plant Science Laboratories, University of Reading, Whiteknights, Reading RG6
6AS, UK
3
Received 10 October 2007; Revised 3 December 2007; Accepted 4 December 2007
Abstract
Plant annexins are ubiquitous, soluble proteins capable of Ca2+-dependent and Ca2+-independent binding
to endomembranes and the plasma membrane. Some
members of this multigene family are capable of
binding to F-actin, hydrolysing ATP and GTP, acting
as peroxidases or cation channels. These multifunctional proteins are distributed throughout the plant and
throughout the life cycle. Their expression and intracellular localization are under developmental and
environmental control. The in vitro properties of
annexins and their known, dynamic distribution patterns suggest that they could be central regulators or
effectors of plant growth and stress signalling. Potentially, they could operate in signalling pathways involving cytosolic free calcium and reactive oxygen
species.
Key words: Annexin, calcium, channel, GTP, peroxide, stress.
Introduction
Plants sense and respond to a range of environmental,
metabolic, and developmental signals. Operation and
control of interacting signal transduction pathways will
involve cell and endomembranes, and integral, peripheral,
and soluble proteins. Downstream responses may require
remodelling of the cytoskeleton and changes to exocytotic
machinery and walls. One family of plant proteins appears
to have the capacity to function at all of those levels—the
annexins. What are they? First discovered in plants
(tomato) by Boustead et al. (1989), research interest in
plant annexins has been spasmodic. In contrast, animal
annexins have been studied extensively. Animal annexins
are involved in signal transduction, free cytosolic Ca2+
([Ca2+]cyt) homeostasis, exo- and endocytosis, membrane
organization, cytoskeletal dynamics, cell cycle control,
and water permeability (Gerke and Moss, 2002; Hill et al.,
2003; Gerke et al., 2005). Analogous functions in plants
could place annexins centre stage in signalling and
adaptation. They are already implicated in cold, oxidative,
saline, and abscisic acid (ABA) stress responses. What do
we know about them and could they be important
components of signalling and response? Here, plant
annexin proteins, their localization, and possible roles are
reviewed.
Annexins of the plant kingdom
Annexins are a multigene, multifunctional family of
soluble proteins with a broad taxonomic distribution. Over
200 unique annexin sequences have been described in >65
species covering plants, fungi, protists, higher vertebrates,
and recently a prokaryote (reviewed by Gerke and Moss,
2002; Hofmann, 2004; Moss and Morgan, 2004; Morgan
et al., 2006). Most of what we know comes from studies
on mammalian annexins. Little is known about the
phylogenetically distinct plant annexins (Fig. 1). They
have been found in all monocot and dicot plants tested to
date (reviewed by Delmer and Potikha, 1997; Hofmann,
2004), including the model plant Arabidopsis (Clark
et al., 2001) and the model legume Medicago (Kovács
et al., 1998; de Carvalho-Niebel et al., 1998, 2002).
* To whom correspondence should be addressed. E-mail: [email protected]
ª The Author [2008]. Published by Oxford University Press [on behalf of the Society for Experimental Biology]. All rights reserved.
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534 Mortimer et al.
Genome sequencing has revealed seven annexin genes
in Arabidopsis (with an eighth evident; Cantero et al.,
2006) and nine in rice (Moss and Morgan, 2004).
Annexins are expressed throughout the body and lifespan
of the plant; embryo (Yu et al., 2005), seedlings (Clark
et al., 1992, 2001; Proust et al., 1996; Cantero et al.,
2006), roots and tubers (Gidrol et al., 1996; Carroll et al.,
1998; de Carvalho-Niebel et al., 1998; Kovács et al., 1998;
Lim et al., 1998; Clark et al., 2001, 2005a, b; Bassani
et al., 2004; Hoshino et al., 2004; Bauw et al., 2006;
Cantero et al., 2006), stems, hypocotyls, and coleoptiles
(Blackbourn et al., 1991; Thonat et al., 1997; Kovács
et al., 1998; Hoshino et al., 2004; Cantero et al., 2006),
cotyledons and leaves (Kovács et al., 1998; Santoni et al.,
1998; Hofmann et al., 2000; Seigneurin-Berny et al.,
2000; Hoshino et al., 2004; Cantero et al., 2006),
inflorescence (Blackbourn et al., 1992; Kovács et al.,
1998), and fruit (Wilkinson et al., 1995; Proust et al.,
1996; Hofmann et al., 2000). In addition to expression in
the vasculature (Clark et al., 2001), annexin proteins have
been found in phloem sap (Barnes et al., 2004; Giavalisco
et al., 2006). It is estimated that annexins can comprise
0.1% of plant cell protein (Blackbourn et al., 1991).
Proteomic studies now show that plant and oomycete
annexins exist in the cell wall as well as the cytoplasm
(Kwon et al., 2005; Meijer et al., 2006). To date, plant
studies have focused on annexin structure and in vitro
protein function. Such studies have revealed a capacity for
annexins to be multifunctional and point towards possible
in vivo roles.
Plant annexin structure differs from that of
animal annexins
Fig. 1. Phylogenetic tree containing annexins from both plant and
animal species. Plant annexins form a distinct monophyletic group.
Identifiers after species names refer to the common annexin designation.
AnxD nomenclature for plant annexins (where established) is included
in parentheses. Multiple sequence alignment was carried out using
Clustal X (Thompson et al., 1997) and the tree was drawn with
TreeView (Page, 1996). NCBI identifiers are as follows (RefSeq where
available, otherwise GI): A. thaliana AnxAt1, NP_174810.1; A.
thaliana AnxAt2, NP_201307.1; A. thaliana AnxAt3, NP181410.1;
A. thaliana AnxAt4, NP_181409.1; A. thaliana AnxAt5, NP_564920.1;
A. thaliana AnxAt6, NP_196584.1; A. thaliana AnxAt7, NP_196585.1; A.
thaliana AnxAt8, NP_568271.2; Z. mays AnxZm33, 6272285; Z. mays
AnxZm35, 1370603; Fragaria3ananassa AnxFa4, 6010777; O. sativa
AnxOs1, NP_001061839; T. aestivum AnxTa1, 38606205; C. annuum
AnxCa24, 75319682; M. truncatula AnxMt1, 3176098; C. elegans Nex-1,
NP_497903; H. sapiens AnxA1, NP_000691; H. sapiens AnxA5,
NP_001145; H. sapiens AnxA6, NP_001146; M. musculus AnxA1,
NP_034860; M. musculus AnxA2, NP_031611; M. musculus
AnxA3, NP_038498; M. musculus AnxA11, NP_038497; M. musculus
AnxA13, NP_081487; D. rerio AnxA1a, NP_861423; R. norvegicus AnxA1,
NP_037036; R. norvegicus AnxA2, NP_063970; R. norvegicus AnxA3,
NP_036055.
Plant annexins have a molecular weight in the region of
32–42 kDa and, although sharing a common evolutionary
ancestor, differ structurally from their animal counterparts.
Animal annexins consist of a conserved a-helical core and
a variable N-terminal region. Their annexin core is
constructed from annexin domains (usually found repeated four times within the protein) each comprising five
short a-helices. The annexin domain, of ;70 amino acids,
contains the conserved endonexin fold (K-G-X-G-T-{38}D/E) and is able to bind Ca2+ (Fig. 2). Structurally, each
annexin domain forms a slightly curved disc. The convex
side contains the Ca2+-binding sites (described as type II
and type III) and faces the membrane surface when an
annexin is associated with lipid (Gerke and Moss, 2002).
Ca2+ ions form a bridge between the annexin and
negatively charged membrane phospholipids. The concave
side faces towards the cytosol, and is available for
interactions both with other parts of the protein and with
other molecules within the cytosol (Fig. 3A). Mammalian
annexins show great variability in the length and sequence
of the N-terminal region. It is thought that this region
Plant annexins 535
Fig. 2. Multiple sequence alignment of a selection of plant annexins. Sequences in the alignment, and the highlighted features, are discussed in the
text. Sequence shading: red box and white character, strict identity; red box and red character, similarity in a group; blue box, similarity across
groups. Features described in the text are highlighted. Green triangle, His40 key peroxidase residue; blue square, IRI actin-binding motif; yellow star,
GXXXXGKT, DXXG putative GTP-binding motif; purple square, KGXGT-38-D/E Ca2+-binding sites; black triangle, putative S3 cluster; turquoise
circle, conserved tryptophan required for membrane binding. GenBank accession numbers: AnxZm33, CAA66900; AnxZm35, CAA66901;
AnxCa32, CAA10210; AnxGh1, AAC33305; AnxAt1, NP_174810; AnxAt2, NP_201307; AnxLe35, AAC97493; AnxMt1, CAA75308. Sequence
alignment was generated using Clustal X (Thompson et al., 1997; default settings). Features were added using ESPript (Gouet et al., 2003).
regulates the stability of different annexin conformations,
determines interactions with other proteins, and hence is
responsible for the functional diversity of mammalian
annexins (Gerke and Moss, 2002). It is the site of
secondary modification, including phosphorylation, nitrosylation, S-glutathiolation, and N-myristoylation, indicative of regulation by several distinct signalling pathways
(reviewed by Gerke and Moss, 2002; Gerke et al., 2005).
For plant annexins, typically only the first and fourth
repeated domains have the characteristic endonexin
sequence (Fig. 2). Crystal structures have been described
(Hofmann et al., 2000, 2003). Plant annexins have, on
average, a larger surface area than mammalian annexins
(Clark et al., 2001). This is due to extra grooves and
clefts, perhaps suggesting a wide range of interaction
partners and a broad range of roles within the cell.
However, in contrast to their animal counterparts, the
N-terminal region of all known plant annexins is short
(;10 amino acids). The crystal structure of bell pepper
annexin (AnxCa32) revealed that the short N-terminal
536 Mortimer et al.
Fig. 3. Annexin structure and localization. (A) Animal annexin (blue)
membrane association in the presence of Ca2+ ions (green). Each of the
four conserved annexin domains (I–IV) is predicted to bind a single
Ca2+ ion, forming a slightly concave disc. In plants, typically only the
first and fourth repeated domains have the characteristic endonexin
sequence. The N-terminal region (N) is the site of secondary modification and in plants is only ;10 amino acids long. (B) Subcellular
localization of plant annexin proteins (blue circles) in an idealized plant
cell. The localization depends on many factors, including plant species,
tissue type, and [Ca2+]cyt, as described in the text. A, actin;
C, chloroplast; CW, cell wall; G, Golgi; M, mitochondria; N, nucleus;
P, peroxisome; PM, plasma membrane; R-ER, rough endoplasmic
reticulum; S-ER, smooth endoplasmic reticulum; V, vacuole.
region interacts with the annexin core, suggesting that
some regulatory function of this region is conserved in
plant annexins (Hofmann et al., 2000).
Ca2+-dependent and -independent membrane
binding
Characteristically, both animal and plant annexins bind
Ca2+ and, in the presence of (micromolar) Ca2+, will bind
to negatively charged phospholipids including phosphati-
dylserine, phosphatidylinositol, and phosphatidic acid
(Blackbourn et al., 1991; Balasubramanian et al., 2001).
Binding can be reversed by addition of Ca2+ chelators. An
annexin may be membrane associated or even membrane
inserted, depending on the [Ca2+]cyt, pH, lipid composition, and voltage (reviewed by Gerke and Moss, 2002).
Certain annexins such as AnxB12 from Hydra have the
capacity to be soluble, peripheral, and integral proteins
(Ladokhin and Haigler, 2005). Using molecular simulation
and site-directed mutagenesis, Montaville et al. (2002)
identified a consensus phosphatidylserine-binding site ([R/
K]XXXK-BC-helices-[R/K]XXXXDXXS[D/E]+Ca2+) in
mammalian annexins. Found in domain I and sometimes
additionally in domain II (but never in domains III or IV),
the site overlaps the Ca2+-binding domains.
Sequence alignment with mammalian annexins
revealed that the phosphatidylserine-binding site is only
poorly conserved in plant annexins (Hofmann et al.,
2000). Despite divergence in amino acid sequence,
phosphatidylserine-binding activity is conserved in at
least some plant annexins including maize, bell pepper,
and cotton (Blackbourn et al., 1991; Hofmann et al.,
2000; Dabitz et al., 2005). Strict sequence conservation
does not appear to be necessary for membrane-binding
function. This is supported by the observation that both
plant and animal annexins bind to a range of negatively
charged phospholipids in addition to phosphatidylserine,
including phosphatidylinositol, phosphatidic acid, and
malonaldehyde-conjugated lipids (Blackbourn et al.,
1991; Balasubramanian et al., 2001). In plants, hydrophobic interactions are also involved in membrane
binding. AnxCa32 attachment to membranes involves
the hydrogen bonding of several amino acid residues to
the phospholipid headgroup and glycerol backbone
(Dabitz et al., 2005). Site-directed mutagenesis of
recombinant tomato annexin p35 (AnxLe35) revealed
that the fourth repeat of the core domain was critical to
lipid binding (Lim et al., 1998).
Although Ca2+ is required for membrane binding at
neutral pH, at acidic pH (<pH 6.0) some animal annexins
bind to membranes independently of Ca2+ (Köhler et al.,
1997; Langen et al., 1998; Rosengrath et al., 1998;
Golczak et al., 2004). Plant annexins also appear capable
of Ca2+-independent membrane binding. Recently, it has
been reported that ;20% of the annexin protein (AnxGh1
and AnxCa32) remains bound to lipid vesicles in the
absence of Ca2+ at neutral pH, and the proteins can be
released following addition of detergent (Dabitz et al.,
2005). However, rather than Ca2+-independent membrane
binding, as suggested by Dabitz et al. (2005), this may
represent a proportion of the population undergoing
membrane insertion, hence the requirement for detergent
to release the protein. In addition to promoting
Ca2+-independent binding, acidic pH can reduce the Ca2+
requirement for phosphatidylserine binding (Blackbourn
Plant annexins 537
et al., 1991). The mechanism of plant annexin
Ca2+-independent membrane binding is still unclear, although a pair of conserved tryptophans (W35/W107) is
involved (Dabitz et al., 2005). Additionally, it has been
suggested that Ca2+-independent membrane binding
serves as a platform for an annexin population whose
membrane binding is Ca2+ dependent. Given that sequential and co-operative binding of annexins has been shown,
the two modes of membrane binding may be intimately
linked (Dabitz et al., 2005).
Although the majority of plant annexins tend to be
found in the cytosol (Blackbourn et al., 1992; Clark et al.,
1992, 1994; Thonat et al., 1997), they are also found
bound to (or in some cases inserted into) both plasma
membranes and endomembranes (Fig. 3B). Annexins can
localize to the plant vacuolar membrane (Seals et al.,
1994; Seals and Randall, 1997; Carter et al., 2004), the
Golgi, and Golgi-derived vesicles (Clark et al., 1992).
Plant annexins can also cause aggregation of liposomes
and secretory vesicles, implicating them in membrane
organization (Blackbourn and Battey, 1993). Medicago
truncatula annexin1 (AnxMt1; de Carvalho-Niebel et al.,
1998) localizes to the nuclear membrane, and M. sativa
(the model legume) AnxMs2 has been shown to localize
in the nucleolus under stress conditions even though the
protein shows no typical nuclear localization signal
(Kovács et al., 1998). A nuclear localization has also been
reported for pea annexin (Clark et al., 1998). A putative
spinach annexin has been identified in chloroplast envelope membranes (Seigneurin-Berny et al., 2000). Arabidopsis AnxAt1 has been found in chloroplasts (Peltier
et al., 2002, 2006; Friso et al., 2004; Kleffmann et al.,
2004; Renaut et al., 2006) but also as a tonoplast protein
(Carter et al., 2004) and an integral plasma membrane
protein ostensibly under non-stressed conditions (Santoni
et al., 1998; Alexandersson et al., 2004). Its membrane
association is prompted by salinity stress (Lee et al.,
2004). Arabidopsis AnxAt4 has also been identified as
a plasma membrane protein (Alexandersson et al., 2004).
An annexin from Bryonia diocia relocates from the
cytoplasm to the plasma membrane following mechanostimulation (Thonat et al., 1997). In wheat exposed to low
temperatures, two wheat annexins accumulate in the
plasma membrane (Breton et al., 2000). Moreover, they
are integral membrane proteins, which cannot be released
by addition of Ca2+ chelators (Breton et al., 2000). That
annexin association with or insertion into membranes can
be dynamic and responsive to environmental change is
consistent with their involvement in signalling and
adaptation.
Actin binding
Actin filaments help shape a cell, are essential for the
development of certain plant cell types, and act in
signalling (reviewed by Drøbak et al., 2004). Actin
binding is limited to a small number of animal annexins
and, as a generalization, actin–annexin interaction appears
to be restricted to regions close to animal membranes
(Hayes et al., 2004). The molecular mechanism of actin
binding is poorly understood, but a C-terminal motif
(LLYLCGGDD) has been implicated; this is not present
in plant annexins (reviewed by Hayes et al., 2004;
Konopka-Postupolska, 2007). Evidence for binding of
plant annexins to actin is mixed, and appears to be species
specific. Tomato and mimosa annexins both undergo
Ca2+-dependent F-actin binding in vitro (Calvert et al.,
1996; Hoshino et al., 2004). Two plasma membraneassociated annexins from zucchini bind to zucchiniderived F-actin (Hu et al., 2000). Mimosa annexin
organizes F-actin into thick bundles in the presence of
2 mM Ca2+ in vitro (Hoshino et al., 2004). Cotton, bell
pepper, and maize annexins have all been extensively
tested and show no affinity for actin, in either the presence
or absence of calcium (Blackbourn et al., 1992; Delmer
and Potikha, 1997; Hoshino et al., 2004). However, the
latter all possess the IRI motif (needed in F-actin binding
to mysosin) implicated in actin binding by Lim et al.
(1998) for tomato annexin. This suggests that the
structural requirement for actin binding is more complex.
As yet annexins from two of the most studied species,
Arabidopsis and Medicago, have not been tested for
F-actin binding, although Arabidopsis sequences contain
a full or partially conserved F-actin-binding motif (Clark
et al., 2001). The functional significance of annexin–actin
interaction is unknown, but has been postulated to be involved in exocytosis and signalling (Konopka-Postupolska,
2007).
Peroxidase activity
The crystal structure of cotton annexin AnxGh1 revealed
two adjacent cysteine residues which, in combination with
a nearby methionine residue, form an S3 cluster. Although
no function has been demonstrated for this cluster, it has
been suggested that it may have a role in transfer of
electrons to an oxidizing molecule, potentially a reduced
reactive oxygen species (ROS) (Hofmann et al., 2003).
Although experimental studies have been limited to
a single Arabidopsis annexin, it is possible that several
plant annexins may be able to act as peroxidases.
Heterologous expression of Arabidopsis annexin AnxAt1
in the Escherichia coli DoxyR (peroxide-sensitive) mutant
or overexpression in mammalian cells afforded protection
against peroxide-mediated oxidative stress (Gidrol et al.,
1996; Kush and Sabapathy, 2001). Peroxidase activity
was demonstrated in vitro using both recombinant
AnxAt1 and AnxAt1 purified from Arabidopsis tissue
(Gidrol et al., 1996; Gorecka et al., 2005). Post-translational
modification of AnxAt1 may be required for peroxidase
538 Mortimer et al.
activity as recombinant AnxAt1 purified from E. coli had
lower activity than that purified from Nicotiana benthamiana, the activity of which was decreased by dephosphorylation (Gorecka et al., 2005). Indeed, secondary
modification of plant annexins is suggested by SDS–
PAGE analysis. For example, the theoretical size and
isoelectric point of AnxAt1 are 36 kDa and pI 5.2,
respectively, whereas following two-dimensional electrophoresis Lee et al. (2004) found two microsomal AnxAt1
spots: 40 kDa, pI 5.2; and 40 kDa, pI 5.3. Santoni et al.
(1998) reported two plasma membrane forms of AnxAt1
with the following migration characteristics: 39 kDa, pI
5.0; and 34 kDa and pI 5.1. Additionally, annexin nitrosylation has been observed in Arabidopsis (Lindemeyr
et al., 2000).
The peroxidase activity of AnxAt1 is suggested to be
due to a region of the first annexin domain in the
N-terminal region (centring on a conserved histidine residue;
His40) which has a strong sequence similarity to the ;30
amino acid haem-binding motif of plant peroxidases,
typified by horseradish peroxidase (Clark et al., 2001;
Gorecka et al., 2005). Consistent with this idea, mutation
of the conserved histidine to alanine substantially reduced
in vitro peroxidase activity (Gorecka et al., 2005). The
putative haem-binding motif is conserved in several plant
annexins including those of maize, cotton, and bell pepper
(Fig. 2). Haem-containing peroxidases catalyse the oxidation of a substrate by the removal of a single electron and
reduce hydrogen peroxide to water. However, to date
there have been no reports of haem binding to plant
annexins to confer peroxidase function. Neither are the
potential targets of annexin peroxidase activity identified
as yet. It remains feasible that in vitro peroxidase activity
is a consequence of metal-binding ability generating
artificial catalytic sites. It is feasible that in common with
other types of peroxidases, annexins are capable of
generating ROS. However, the sheer numbers of peroxidases in plants and potential for redundancy may make
this aspect of annexin physiology particularly difficult to
elucidate.
ATPase and GTPase activity
Binding to ATP and GTP is a common feature of animal
annexins even though they do not contain the Walker
A and B motifs. Rather, the nucleotide-binding motif is
thought to be FXXKYD/EKSL (Bandorowicz-Pikula
et al., 2003). Plant annexins not only bind purine
nucleotides but also hydrolyse them (maize, McClung
et al., 1994; tomato, Calvert et al., 1996; Lim et al., 1998;
cotton, Shin and Brown, 1999). In contrast to animal
annexins, nucleotide binding and hydrolysis may depend
on a Walker A motif (GXXXXGKT/S) and a GTPbinding motif typical of the GTPase superfamily (DXXG).
These sequences have been found in the fourth repeat of
AnxGh1; C-terminal deletion and loss of the fourth repeat
abolished its GTPase activity (Shin and Brown, 1999). In
Arabidopsis, the greatest similarity is in the fourth repeat
of AnxAt2 and AnxAt7 (Clark et al., 2001).
Both maize and tomato annexins are able to hydrolyse
ATP and GTP at a similar rate, but GTP is the preferred
substrate for cotton annexin AnxGh1. Tomato annexin
GTPase activity still proceeds when the protein is bound
to actin (Calvert et al., 1996), suggesting that cytoskeletal
association may specifically locate the annexin’s GTPase
function in the cell. Ca2+ has an inhibitory effect on cotton
annexin GTPase activity but not the ATPase/GTPase
activity of maize annexin AnxZm33/35 (McClung et al.,
1994; Shin and Brown, 1999). Alignments of the primary
sequence of cotton annexin, AnxAt2, and AnxZm33/35
showed that the GTP-binding motifs overlap the
Ca2+-binding motif of the fourth endonexin domain
(McClung et al., 1994; Shin and Brown, 1999). Ca2+ and
GTP may therefore compete for binding. Ca2+-mediated
phospholipid binding has been shown to inhibit hydrolytic
activity of tomato annexin (Calvert et al., 1996). Mutagenesis of the Ca2+-binding sites does not impair GTPase
activity of the soluble form (Lim et al., 1998), demonstrating that membrane binding prevents GTP from reaching its catalytic site. Overall, modulation of enzyme
activity by Ca2+ and membrane binding may afford spatiotemporal definition of annexin function. That tomato,
maize, and cotton annexins have different requirements,
despite catalysing the same reaction, may reflect the
diverse roles that plant annexins fulfil.
Sensor and channel activity
Ca2+ binding is a defining annexin characteristic, but
studies on animal annexins show them to be capable of
sensing and regulating free cytosolic calcium (Hawkins
et al., 2000; Watson et al., 2004). Mammalian AnxA6 has
been shown to act as a Ca2+ sensor, mediating membrane
association of a GTPase-activating protein (GAP) that
then regulates a monomeric GTPase (Grewal et al., 2005).
In addition to controlling trafficking of ion transporters to
their target membranes and regulating their activity
(reviewed by Gerke et al., 2005), animal annexins can
also form Ca2+-permeable ion channels themselves. This
ability was first demonstrated with bovine AnxA7 which
forms a highly selective, voltage-gated Ca2+ channel in
phosphatidylserine bilayers (Pollard and Rojas, 1988).
Since then, many other animal annexins have been shown
to form ion channels, although their properties (particularly selectivity) differ (reviewed by Hawkins et al., 2000;
Kourie and Wood, 2000). ATP, GTP (Kirilenko et al.,
2006), cAMP, hydrogen peroxide (Kubista et al., 1999),
and low pH (Köhler et al., 1997; Langen et al., 1998;
Rosengarth et al., 1998) have all been shown to regulate
annexin channel activity. Animal annexins can cause Ca2+
Plant annexins 539
2+
influx across animal plasma membrane and mobilize Ca
from internal stores (Kubista et al., 1999; Watson et al.,
2004) placing them at the core of Ca2+-mediated signal
transduction and Ca2+ homeostasis.
A number of models have been proposed to explain the
mechanism whereby animal annexin proteins (which are
too small to span a membrane) form ion channels. At
acidic pH, the short helices of the monomer may join to
form longer helices to allow spanning of the membrane.
In invertebrates this has been observed for AnxB12,
which undergoes large-scale conformational changes under
conditions that result in membrane insertion (Langen
et al., 1998; Isas et al., 2000). Recently, freeze-fracture
electron microscopy revealed images of AnxB12 as an
integral membrane protein at pH 4 but not at higher pH
(Hegde et al., 2006). This membrane-inserted form of
AnxB12 appears to be monomeric, unlike the trimeric
membrane-attached form (Ladokhin and Haigler, 2005).
Another model suggests that an annexin monomer
associates with the membrane and causes an electrostatic
disruption which allows passage of ions through the
central pore of the protein, although it is not clear how
this pore would be selective for Ca2+ (reviewed by Kourie
and Wood, 2000). Regardless of the mechanism, annexin
ion channels challenge the view of ion channels as solely
integral membrane proteins. As unusual as this is,
membrane insertion of ion channels from the aqueous
phase is not unprecedented. In addition to annexin
channels, an animal chloride channel (CLIC-1; Tulk
et al., 2002) also moves directly from the cytosol into
a membrane.
An ability of plant annexins to form or regulate Ca2+
channels in plasma and endomembranes would enable
signal transduction and amplification (Kovács et al., 1998;
White et al., 2002). The putative pore region of the human
AnxA5 channel contains two salt bridges (Asp92–Arg117
and Glu112–Arg271) that regulate selectivity for calcium
and channel opening in response to voltage (Liemann
et al., 1996). Both salt bridges are quite well conserved in
plant annexins. It has been proposed that plant annexins
could act as the plasma membrane Ca2+-permeable
channels that mediate Ca2+ entry into the cell at very
negative (hyperpolarized) membrane voltage (White et al.,
2002). Such channels are strongly implicated in growth
and signalling. To date, AnxCa32 has been shown to
mediate passive Ca2+ flux in fura-2-loaded vesicles
(Hofmann et al., 2000), supporting the general concept of
channel function. Maize annexins AnxZm33/35 contain
the putative salt bridges, and a partially purified preparation formed hyperpolarization-activated Ca2+-permeable
channels in planar lipid bilayers (Nichols, 2005). As only
one gene has been verified as encoding a plant Ca2+
channel (TPC1 encoding a vacuolar channel; Peiter et al.,
2005), it will be of great interest to see whether plant
annexins purified to homogeneity support Ca2+ channel
activity. Of the eight Arabidopsis annexins, AnxAt1 has
been found to form K+-permeable channels in bilayers,
with channel formation favoured at low pH (Gorecka
et al., 2007). In common with AnxA5 and AnxB12, acidic
pH increases the hydrophobicity of AnxAt1 and promotes
the oligomerization thought necessary for transport activity.
The Ca2+ permeability of the AnxAt1 channel remains to
be determined, as does its in vivo function, but AnxAt1
illustrates clearly the multifunctional nature of annexins—
potentially a peroxidase and a channel in one.
Function in exocytosis, growth, and development
Annexin expression is dynamic even under normal growth
conditions. Transcript levels of annexin genes in Arabidopsis vary depending on tissue type and age, suggesting
specific purposes at different developmental stages in
different tissue (Clark et al., 2001, 2005a; Hoshino et al.,
2004; Cantero et al., 2006). Annexin expression increases
during fruit ripening and gall ontogeny, implying hormonal control (Wilkinson et al., 1995; Proust et al., 1996;
Vandeputte et al., 2007). Nod factors induce M. truncatula annexin1 (AnxMt1) expression, and co-localization
studies using AnxMt1–GFP (green fluorescent protein)
have suggested that it may be involved in the early stages
of cell division required for nodule formation (de
Carvalho-Niebel et al., 2002).
The in vitro ability to bind membranes, Ca2+, purine
nucleotides, and actin predicts critical roles for both
animal and plant annexins in membrane trafficking and
signal transduction. Animal annexins are clearly involved
in exo- and endocytosis, and the targeting of proteins to
specific membrane sites (reviewed by Gerke and Moss,
2002; Gerke et al., 2005). The clearest demonstration to
date of annexin function in planta has been the stimulation of Ca2+-dependent vesicle fusion to the plasma
membrane of maize root cap protoplasts by AnxZm33
and 35 (Carroll et al., 1998). Plant annexins are abundant
and underlie the plasma membrane in cells associated with
high secretion rates. Annexins are prominent at the apex
of cells undergoing polar elongation, such as root hairs,
pollen tubes, and fern rhizoids (Blackbourn et al., 1992;
Blackbourn and Battey, 1993; Clark et al., 1995, 2001,
2005a). Annexin expression has been detected in the root
elongation zone of maize (Carroll et al., 1998; Bassani
et al., 2004) and Arabidopsis (Clark et al., 2005a, b). As
well as possibly being involved in primary and root hair
elongation growth, a specific annexin in Arabidopsis
(AnxAt2) is implicated in lateral root development (Clark
et al., 2005a), implying upstream regulation by growth
regulators. Cotton annexin (AnxGh1) mRNA is upregulated during cotton fibre elongation, and the protein
may be associated with Golgi-derived coated vesicles
mediating fibre elongation (Shin and Brown, 1999; Bulak
Arpat et al., 2004).
540 Mortimer et al.
Recent work on a Saprolegnia annexin has revealed an
ability to stimulate (1–3)-b-D-glucan synthase activity,
implying a role in the regulation of wall synthesis
(Bouzenenza et al., 2006). This is in contrast to the
inhibitory effect of cotton annexin AnxGh1 (Andrawis
et al., 1993) and suggests that different annexins play
distinct regulatory roles. Arabidopsis AnxAt7 expression is
up-regulated by oxylipin treatment that induces callose
formation and causes wavy roots and lateral root inhibition (Vellosillo et al., 2007). It will be interesting to
see whether this annexin regulates glucan synthase. In
addition to roles in exocytosis and wall synthesis, individual annexins could be acting as Ca2+ or GTP sensors
to co-ordinate growth. A role as a Ca2+ sensor has been
proposed for vacuole-associated annexins (Seals et al.,
1994). Vacuolar biogenesis is a key component of cell
expansion, and expression of the vacuole-associated
tobacco annexin VCaB42 correlates with vacuolar biogenesis in expanding cells (Seals and Randall, 1997). The
annexin is associated with a ROP GTPase (Lin et al.,
2005), a type of protein viewed as master regulators of
growth. Addition of GTP inhibited annexin-mediated
exocytosis in root cap protoplast (Carroll et al., 1998),
suggesting that annexin function is co-ordinated by local
GTP and Ca2+ levels.
Light response, nyctinastic movement, and
gravitropism
As well as being linked to growth and development,
annexin expression and distribution can also change in
response to environmental stimuli. Light affects the
expression of certain annexins in Arabidopsis (Cantero
et al., 2006). In hypocotyls, AnxAt5 expression is increased by red light and this is reversible by application of
far red light; in cotyledons, AnxAt6 has a similar red/far
red response (Cantero et al., 2006). These results point to
annexin expression being downstream of phytochrome A,
and further dissection of this relationship is awaited.
Phytochrome action has previously been implicated in the
regulation of polarized annexin distribution in fern
rhizoids (Clark et al., 1995). Nyctinastic (night time)
movement of the Mimosa pulvinus provides a beautiful
example of temporal regulation of annexin abundance and
positioning. The amount of annexin is more significant at
night (when the leaf droops) and the protein is largely
cytosolic, whilst in the morning (when the leaf is held up)
it has redistributed to the outermost periphery of the motor
cells in the pulvinus (Hoshino et al., 2004). Quite what
the annexin does in either position remains unknown but
its abundance is positively regulated by ABA (Hoshino
et al., 2004), which suggests that annexins link stress
responses with nyctinastic and possibly seismonastic
(touch-induced) movement. Both types of movement
involve Ca2+ influx from the apoplast (Campbell and
Thomson, 1977), and perhaps this is involved in annexin
relocation. Mimosa annexin binds F-actin in vitro (Hoshino
et al., 2004). Actin filaments in Mimosa are thought to
control pulvinus movement, in conjunction with altered
osmotic pressures, through decreased tyrosine phosphorylation of the actin, leading to tissue bending. However,
since Mimosa annexin distribution does not precisely
follow actin distribution in vivo, the relationship between
the two remains unclear (Hoshino et al., 2004; Kanzawa
et al., 2006).
Although there are no reports as yet for involvement of
annexins in gravisensing, they are implicated in mediating
the resultant differential growth response. Prior to a gravitational stimulus, annexins are asymmetrically distributed
in the direction of gravity at the periphery of cells just
below the apical meristem of etiolated pea plumules
(Clark et al., 2000). Using immunofluorescence, annexins
were found to redistribute within 15 min of a gravitational
stimulus (and prior to the onset of plumule curvature),
occupying a more evenly distributed peripheral position
(Clark et al., 2000). This has been interpreted in terms of
an annexin involvement in redirecting materials for
growth (Clark et al., 2000). In gravistimulated Arabidopsis roots, the abundance of AnxAt1 increases in roots
(Kamada et al., 2005) and predominates in epidermal cells
that would undergo the greatest growth rate to bend the
root (Clark et al., 2005b). This distribution largely
matched that of polysaccharide secretion, supporting a role
for annexins in the gravistimulated growth response
(Clark et al., 2005b). In gravistimulated hypocotyls,
AnxAt2 was detected preferentially in the epidermis that
would grow the fastest (Clark et al., 2005b). Determination of how these distributions are caused and link to
gravistimulated changes in [Ca2+]cyt, actin, and ROS is
increasingly within the technical range of plant biologists.
Overall, the ability to bind membrane, GTP, and actin
suggests the involvement of annexins in (differential)
growth and places them downstream of a wide range of
signal transduction pathways.
Responding to stress stimuli and pathogens
The mechanical stress caused by wind results in short,
bushy plants (thigmomorphogenesis). Annexins respond
to mechanical stress in B. dioica internodes by redistributing from the cytosol to the plasma membrane in
parenchyma cells (sampled 30 min after the stimulus;
Thonat et al., 1997). The chain of events leading to this
remains to be determined. However, mechanical stress is
known to elevate [Ca2+]cyt and this could stimulate
annexin–plasma membrane association. The significance
of annexin relocation is not understood, but as regulators
of growth they may govern the radial expansion that
results from mechanical stress or could be ‘conditioning’
the plasma membrane for further stress responses.
Plant annexins 541
Cold causes increased annexin expression in poplar
leaves (Renault et al., 2006). In wheat, the cold-induced
accumulation of annexins p39 and p22.5 and their
insertion into the plasma membrane could be involved in
sensing or transducing Ca2+ signals or in regulating
[Ca2+]cyt during signalling or acclimation (Breton et al.,
2000). Annexin expression, abundance, and cellular
position can respond to osmotic stress, salinity, drought,
and ABA (Watkinson et al., 2003; Lee et al., 2004;
Buitnik et al., 2006; Vandeputte et al., 2007). Alfalfa
annexin AnxMs2 mRNA increased during tissue development, but the amount of transcripts was elevated significantly upon NaCl treatment or exogenous ABA
application (Kovács et al., 1998). The level of Arabidopsis
AnxAt1 protein changes as the plant is subjected to
osmotic stress, even though its transcript is not affected
(Lee et al., 2004). However, in contrast, Cantero et al.
(2006) found that AnxAt1 mRNA abundance is increased
by salinity stress. The temporary annexin relocation to
membranes in response to ABA and saline stress (Lee
et al., 2004) could reflect a role in signalling, or represent
their regulation of proteins already in the membrane or
annexin-mediated insertion of new proteins to cope with
adverse conditions. AnxAt1 transcript accumulates in
response to phosphate deprivation (Müller et al., 2007)
and in the presence of H2O2 (Gidrol et al., 1996). As
H2O2 accumulates in response to phosphate deprivation
(Shin et al., 2005), this result suggests that AnxAt1
operates downstream in this stress response. AnxAt1
expression is also up-regulated by salicylic acid, which
implicates this annexin in pathogen defence responses
(Gidrol et al., 1996). Expression of Arabidopsis AnxAt4,
tomato AnxLe34, and tobacco AnxNt12 also increases
during pathogen attack but the functional significance
remains unknown (Xiao et al., 2001; Truman et al., 2007;
Vandeputte et al., 2007). Salinity and ABA also induce
AnxNt12 expression, but application of H2O2 does not,
suggesting control by distinct signalling pathways and
specific annexin function (Vandeputte et al., 2007).
requires AnxA5 (Kubista et al., 1999). From this it
follows that channel-forming plant annexins (such as
AnxAt1) are candidates for the ROS-activated channels
identified in several plant cells (Foreman et al., 2003).
Recently, annexins have been identified as protein
components of an M. truncatula plasma membrane lipid
raft alongside signalling and redox proteins (Lefebvre
et al., 2007). They could, in common with raft-associated
animal annexins (Babiychuk and Draeger, 2000), anchor
rafts to the actin cytoskeleton (Konopka-Postupolska,
2007). Alternatively, conceivably, raft-associated annexins
could function as channels or peroxidases operating in
a localized ROS signalling ‘hub’. AnxAt1 is present at the
root hair apex (which is thought to harbour lipid rafts;
Jones et al., 2006) and as a peroxidase could help regulate
the intracellular peroxide generated during polar growth
(Foreman et al., 2003). As peroxidases, annexins could
regulate peroxide generated as an inter- or intracellular
messenger or relay/terminate a signal through peroxidedependent oxidation. A protective role can also be
envisaged. Peroxidase activity of annexins associated with
chloroplast RNA polymerase could protect newly synthesized transcripts from oxidation (Pfannschmidt et al.,
2000).
Future perspectives
Sensors? Skeletal regulators? Channels? Peroxidases?
Plant annexins are potentially multifunctional proteins
in vivo involved in membrane dynamics, actin modelling,
and [Ca2+]cyt and ROS regulation. Swifter resolution of
stimulus-induced annexin relocation to membranes (particularly in single cell paradigms such as the root hair or
guard cell) will help secure definitions of function, as will
the increasing availability of mutants.
Acknowledgements
This work was supported by the BBSRC and the Cambridge
Overseas Trust.
Annexins and reactive oxygen species
Production of ROS is implicated in control of plant
(tropic) growth and development, in some cases controlling [Ca2+]cyt (reviewed by Gapper and Dolan, 2006).
Stress conditions that cause ROS generation (such as
pathogen attack, drought, salinity, cold, hypoxia, and
nutritional restriction) also prompt annexin accumulation
or relocation to membranes. However, it is as yet unclear
whether ROS or elevated [Ca2+]cyt trigger annexin
responses. Membrane oxidation increases membrane binding of animal annexins (Balasubramanian et al., 2001).
Peroxide can induce the channel-forming vertebrate
AnxA5 to be inserted into membranes in vitro, and
peroxide-induced Ca2+ influx in vivo in DT40 pre-B cells
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