Protoplasma (2007) 230: 203–215
DOI 10.1007/s00709-006-0234-7
PROTOPLASMA
Printed in Austria
Annexins: putative linkers in dynamic membrane–cytoskeleton
interactions in plant cells
D. Konopka-Postupolska
Laboratory of Plant Pathogenesis, Institute of Biochemistry and Biophysics, Polish Academy of Sciences, Warsaw
Received January 10, 2006; accepted March 14, 2006; published online April 24, 2007
© Springer-Verlag 2007
Summary. The plasma membrane, the most external cellular structure,
is at the forefront between the plant cell and its environment. Hence, it is
naturally adapted to function in detection of external signals, their transduction throughout the cell, and finally, in cell reactions. Membrane
lipids and the cytoskeleton, once regarded as simple and static structures,
have recently been recognized as significant players in signal transduction. Proteins involved in signal detection and transduction are organised
in specific domains at the plasma membrane. Their aggregation allows to
bring together and orient the downstream and upstream members of signalling pathways. The cortical cytoskeleton provides a structural framework for rapid signal transduction from the cell periphery into the
nucleus. It leads to intracellular reorganisation and wide-scale modulation of cellular metabolism which results in accumulation of newly synthesised proteins and/or secondary metabolites which, in turn, have to be
distributed to the appropriate cell compartments. And again, in plant
cells, the secretory vesicles that govern polar cellular transport are delivered to their target membranes by interaction with actin microfilaments.
In search for factors that could govern subsequent steps of the cell response delineated above we focused on an evolutionary conserved protein family, the annexins, that bind in a calcium-dependent manner to
membrane phospholipids. Annexins were proposed to regulate dynamic
changes in membrane architecture and to organise the interface between
secretory vesicles and the membrane. Certain proteins from this family
were also identified as actin binding, making them ideal mediators in
cell membrane and cytoskeleton interactions.
Keywords: Plant annexin; Actin microfilament; Stress response; Annexin–actin interaction; Exocytosis.
Annexins in plant cells
Annexins constitute a family of ubiquitous, calcium- and
membrane-binding proteins. They have been intensively
studied since the identification of the first annexin in animal
tissues (Creutz et al. 1978) and subsequent recognition of
* Correspondence and reprints: Laboratory of Plant Pathogenesis, Institute of Biochemistry and Biophysics, Polish Academy of Sciences,
Pawinskiego 5A, 02-106 Warsaw, Poland.
proteins with similar characteristics in plant cells (Boustead
et al. 1989, Blackbourn et al. 1991). In vertebrates the annexins were grouped into 13 families. In contrast, plant annexins
seem to represent a relatively simpler, smaller, and less
diverse family of proteins. Nevertheless, in all analysed
plant species, at least two distinct proteins with molecular
masses between 33 and 36 kDa have been discovered
(Smallwood et al. 1990, Shin et al. 1995, Proust et al. 1996,
Thonat et al. 1997). However, a search through the
Arabidopsis thaliana genome, in which seven annexin
genes were found (Clark et al. 2001), showed that this
number can be even larger. Summarising, it seems to be
true for both vertebrates and plants that more than one annexin is usually expressed at a given moment and in a particular cell type (so-called annexin fingerprint). This
indicates that in spite of significant functional homology,
individual annexins support distinct and divergent functions. Some data suggest that individual annexins are associated with different cellular compartments possibly
conferring specificity of cellular response to the given stimulus. A special family of plant vacuolar annexins, VCaBP,
with a slightly larger molecular mass (ca. 42 kDa) was
discovered in various plant species of the families
Solanaceae and Brassicaceae (Seals et al. 1994, Seals and
Randall 1997). In mustard plants (Sinapis alba), annexin
p28 was shown to be a part of the chloroplast translation
apparatus (Pfannschmidt et al. 2000). Finally, the presence
of nuclear annexins has also been documented (Clark et al.
1998, Kovacs et al. 1998).
Annexins have an evolutionary conserved overall structure, with an about 70-amino-acid motif repeated four
times within the molecule, and contain a discrete (neither
EF-hand nor C2) calcium binding site (Fig. 1). Calcium
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D. Konopka-Postupolska: Annexins in regulation of membrane–cytoskeleton dynamics
D. Konopka-Postupolska: Annexins in regulation of membrane–cytoskeleton dynamics
binding induces structural changes resulting in protein
translocation from the cytoplasm to the cell periphery.
Crystallographic data reveal that membrane binding occurs via formation of a ternary complex between annexin,
calcium, and the membranes (Swairjo et al. 1995). Annexins have been shown to preferentially bind to negatively
charged membrane phospholipids (e.g., phosphatidylserine). Plant annexins are fairly abundant cellular proteins
(Clark and Roux 1995) and should thus be considered as a
very important element of calcium signalling pathways.
On the basis of immunocytochemical experiments, it has
been concluded that in plant cells, annexins are localised
mainly in the cytoplasm. When calcium levels increase,
they are moved towards the cytoplasmic surface of certain
membrane structures, mainly the plasma membrane, but
the presence of an annexin that binds specifically to the
outer membrane of the chloroplast envelope was also reported (Seigneurin-Barny et al. 2000). On the other hand,
experimental data show that particular proteins, although
lacking defined targeting sequences, are present in nonstimulated cells in different cellular compartments. Proteomic analysis revealed that, e.g., AnnAt1 from A. thaliana
was present in the fraction of cell wall proteins (P. Wojtaszek, Adam Mickiewicz University, Poznan, Poland, pers.
commun.), integral membrane proteins (Santoni et al. 1998,
S. Lee et al. 2004), in central, vegetative vacuoles (Carter
et al. 2004), as well as in the nuclear matrix (A. Jerzmanowski, Institute of Biochemistry and Biophysics, Polish
Academy of Sciences, Warsaw, Poland, pers. commun.).
Mimosa annexin exhibits day–night changes in distribution:
during the daytime it is localised on the cell periphery,
while at night it stays in the cytoplasm (D. Hoshino et al.
2004). Additional analyses are necessary to establish
whether this microcompartmentalisation is also true for the
other plant annexins.
Despite several years of investigation, the primary
physiological function for annexins has not yet been elucidated. It is generally assumed that annexins are implicated in several processes related to membranes, including
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regulation of membrane organisation, membrane trafficking, interactions with the cytoskeleton, and secretion. In
time it became clear that certain plant annexins can also
function in plant stress response. Expression of different
annexins was induced by osmotic stress (AnnMs2 from
alfa alfa [Kovacs et al. 1998]; AnnAt1 from A. thaliana
[S. Lee et al. 2004 and our unpubl. data]), which suggests
that they might participate in drought resistance. Some
data indicate also that AnnAt1 can act at a crossroad between auxin and abscisic acid signalling (Bianchi et al.
2002). Enhanced expression of annexin was also reported
after different treatments that led to accumulation of reactive oxygen species during defence response in tomato
plants (Xiao et al. 2001), and after salicylic acid and hydrogen peroxide treatment in Arabidopsis plants (Gidrol
et al. 1996). It is worth mentioning that those two annexins, namely, p34 from tomato cells and AnnAt1 from
A. thaliana, represent close homologs (84% of homology
on protein level). AnnAt1 also has the ability to protect
heterologous cells from the consequences of oxidative
stress. Molecular mechanisms of this protection have not
been elucidated, although different hypotheses are considered, beginning from an intrinsic peroxidase activity of
AnnAt1 (Gidrol et al. 1996). An indirect effect via modulation of calcium signalling that results in the lowering of
superoxide production and reduction of protein kinase C
activity was also proposed (Kush and Sabapathy 2001,
Janicke et al. 1998). It is also possible that it can be a nonspecific consequence of membrane lipid protection
against oxidative stress. Mammalian annexin A5 was
shown to bind, with affinity similar to that of phosphatidylserine, to malondialdehyde adducts, a major product of lipid peroxidation generated by the nonenzymatic
reaction of polyunsaturated fatty acids with molecular
oxygen (Balasubramanian et al. 2001). In contrast to free
radicals, lipid peroxides are long-lived and can thus diffuse from the site of origin and exert deleterious effects
and/or activate stress-related pathways in surrounding tissues. If AnnAt1, and possibly other annexins, shares this
Fig. 1. Alignment of the deduced amino acid sequences of human (A1, A2, A4, A5, A6, A7, A11, A13) and plant (Arabidopsis thaliana AnnAt1 to
-7, Capsicum annuum Ca_p32 and p38, Lycopersicon esculentum Le_p34 and p35, Gossypium hirsutum Gh1, Nicotiana tabacum Nt_VCaBP, Solanum
tuberosum St_p34, and Medicago sativa MsAnn annexin genes obtained using T-COFFEE (Notredame et al. 2000). Potential functional domains are
indicated as follows: Ca2-binding sites of type II G-X-GTD-{ca. 38}-E/D are shown in black, potential actin binding motifs (IRI and/or LLYLCG
GDD) are shown in italics, putative PIP2 binding domain (R/K)LXXX(K)X(K)(R/K) is underlined, generic motif for N-myristoylation site in
AnxA13. GenPept accession numbers of human annexins are as follows: AnxA1, NP_000691; AnxA2, NP_001002858; AnxA4, NP_001144;
AnxA5, NP_001145; AnxA6, NP_004024; AnxA7, NP_004025; AnxA11, NP_665876; and AnxA13, NP_004297. Locus numbers of Arabidopsis
annexins are as follows: AnnAt, At1g357201; AnnAt2, At5g65020; AnnAt3, At2g38760; AnnAt4, At2g38750; AnnAt5, At1g68090; AnnAt6,
At5g10220; and AnnAt7, At1g10230. GenPept accession numbers of plant annexins are as follows: Ca_p32, CAA10210; Ca_p38, CAA10261;
Le_p34, AAC97494; Le_p35, AAC97493; Nt_VCaBP42, AAD24540; St_p34, ABB02651; and MsAnn, CAA72183. Due to lack of space only the
first half of the doubled annexin AnxA6 molecule is depicted in the scheme
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D. Konopka-Postupolska: Annexins in regulation of membrane–cytoskeleton dynamics
malondialdehyde-binding activity, it could protect cells
from oxidative stress by hampering the propagation of
signalling molecules derived from polyunsaturated fatty
acids.
Annexins and membrane dynamics
Plant cellular membranes contain striking amounts of
both structurally and functionally varied lipids that are
Fig. 2 A–C. Schematic representation of raft
structures and mechanism of their interactions with annexins. Rafts represent laterally
organised lipid microdomains enriched in
sphingolipids and sterols (liquid order phase).
A specific set of proteins interact with rafts
through transmembrane domains or lipid anchor (GDI-tail for extracellular and double
acyl for intracellular proteins). Due to their
ability to concentrate signaling molecules
and to interact with the actin cytoskeleton,
rafts were proposed to function as signaling
centers. In animal cells, annexins were proposed to be important for raft function, governing their aggregation and influencing cytoskeleton interactions. There are some data
suggesting that in plant cells, annexins may
support similar functions. A Upon initial calcium elevation, different calcium-binding
proteins, including annexins, are activated.
This can result in triggering of specific intracellular transduction pathways, e.g., phosphoinositide signaling. At the same time, a
particular annexin (A1) with the highest calcium affinity is translocated to the plasma
membrane and binds to negatively charged
phospholipids in the inner leaflet. B Once
bound, annexin A1 can oligomerise on membrane surfaces, thus initiating raft clustering
and stabilisation. In turn, further calcium elevation can lead to the activation of another
protein from this family, with slightly different characteristics and binding preferences.
After translocation, A2 can bind to another
type of negatively charged phospholipid,
PIP2, that is enriched within sterol-rich domains. C In this localisation, annexin A2 can
serve as a platform for actin assembly, promote cytoskeleton rearrangement, and regulate its binding with membranes. Thus, new
signaling pathways can be constituted. Moreover, raft clusters (macrorafts) are further stabilised by hindering their lateral mobility by
association with the cytoskeleton. Additionally, transporting vesicles that moved along
microfilaments can be directed to the center
of exocytosis
D. Konopka-Postupolska: Annexins in regulation of membrane–cytoskeleton dynamics
distributed asymmetrically among different membranes.
For example, cholesterol and phytosterols are localised
mainly in the plasma membrane, within the outer leaflet
of the bilayer, while phospholipids are more or less uniformly distributed in the inner leaflet. Moreover, even
within a single membrane, lipids can be organised laterally to form discrete domains that can support particular
cellular functions. One of the most widely studied types
of such lateral organisation results from the association of
sphingolipids parallel to one another, probably through
weak interactions between their carbohydrate heads.
Empty spaces between associated sphingolipids are filled
with stiff sterol molecules arranged perpendicularly to the
bilayer forming a so-called liquid ordered structure, called
rafts (Simons and Ikonen 1997, Harder and Simons 1997,
Munro 2003, Edidin 2003) (Fig. 2). In such an ordered
arrangement, the lateral movement of lipids is not impaired so that rafts can easily move in the plane of the surrounding, more fluid bilayer and aggregate into larger
domains. Rafts are especially abundant in the plasma
membrane but can also be found in secretory and endocytic
pathways (Simons and Toomre 2000). Although rafts develop exclusively in the outer leaflet of a bilayer, they are
able to organise, in a still unknown way, the lipids of the inner leaflet. Recently, Gri et al. (2004) found that in Jurkat
T cells the domains at the outer and inner leaflets are physically coupled and this coupling requires cholesterol.
The results of several studies suggest that this lateral
assembly of sphingolipids and sterols may be involved in
transport of newly synthesised material to the cell surface
in polarised and nonpolarised vertebrate cells (Musch
et al. 1996, Simons and van Meer 1988, Yoshimori et al.
1996, Zegers and Hoekstra 1998) and in organising the
cellular machinery for exo- and endocytosis (Schnitzer
et al. 1996). Due to this ability to selectively aggregate
special proteins, rafts were also proposed to function in
cell signaling (Parton and Simons 1995, Simons and
Toomre 2000). Protein association with lipid rafts is mediated by glycosylphosphatidylinositol (GPI) anchoring
(Mayor et al. 1994), which results in coclustering of double acylated proteins on the cytosolic surface (Resh 1999).
Association with the inner leaflet could also be governed
by direct protein interaction with cholesterol (Parton et al.
1994). The newest measurements, using single-particle
tracking, showed that at steady state, rafts are generally
small, with a diameter of about 50 nm (which corresponds
to about 3500 lipids [Pralle et al. 2000]), exist for less
than 1 min, and contain no more than 10–30 protein molecules (Varma and Mayor 1998). Thus, single rafts are too
small and have to coalesce into bigger domains (macro-
207
rafts) to fulfill the signalling function. Clustering can be
accomplished from both sides of the plasma membrane,
and some experimental evidence indicates that annexins,
acting from the inner leaflet side, promote lipid raft formation and influence their dynamics. Indeed, certain vertebrate annexins (A2, A6, and A13b) were shown to be
present in a Triton X-100-insoluble membrane fraction
that contains raft domains (Harder and Gerke 1994, Lafon
et al. 1998, Draeger et al. 2005). It was proposed that annexins oligomerise on the membrane surface resulting in
the formation of a two-dimensional lattice (Lambert et al.
1997) that can serve as a platform for raft aggregation into
larger domains (Oliferenko et al. 1999). It is worth stressing that several plant annexins were shown to form
homodimers and oligomers in solutions, which is a prerequisite for raft aggregation (Hoshino et al. 1995, Hofmann
et al. 2002). Moreover, the presence of a protein sheet on
membrane surface can induce segregation of certain proteins and modulate enzyme functions (Andree et al. 1992,
Dubois et al. 1998). In turn, raft binding can also modify
the properties of annexins. For example, annexin A2 was
identified as a one of the main cellular substrates for
pp60c-Src kinases (Hubaishy et al. 1995), which is concentrated in cholesterol-rich domains.
One has to keep in mind that a single cell expresses
several annexins at the same time and they may act in synergy to support cellular processes, although their mode of
action could be slightly different. The most detailed and
compelling picture depicting the function of annexins in
membrane dynamics emerged from smooth muscle cells
(Babiychuk and Draeger 2000, Draeger et al. 2005). The
initial calcium signal (up to 300 nM) induces annexin A2
translocation to the rafts, which promotes their aggregation and formation of macrorafts. As the calcium concentration increases further (up to 600 nM), the second annexin,
A6, translocates to the plasma membrane, where it binds
in the vicinity of the macrorafts and induces the formation
of a mechanical link coupling the actin-based cytoskeleton and the membrane (Babiychuk et al. 1999), thus stabilising the macrorafts. Further elevation of the calcium
signal above 1000 nM results in annexin A5 binding to
glycerophospholipids, which may additionally stabilise
the macrorafts. It is not clear if a similar step-by-step mechanism also operates in other less specialised cell types, but
the fact that single cells usually express several annexins differing in calcium affinity and other properties (e.g., the ability to bind to the cytoskeleton) supports this notion (Fig. 2).
The presence of lipid microdomains in plant cell membranes has been recognised only recently. With respect to
lipid and protein contents, they seem to have characteris-
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D. Konopka-Postupolska: Annexins in regulation of membrane–cytoskeleton dynamics
tics similar to those of animal and yeast rafts, although
discrete differences have also been reported (Peskan et al.
2000, Mongrand et al. 2004, Borner et al. 2005). They contain significant amounts of GPI-anchored proteins (Borner
et al. 2005), a large number of signalling molecules (receptor kinases with leucine-rich repeats), several components of downstream signalling pathways (small and
heterotrimeric G proteins), and finally proteins involved in
stress response (similar to hypersensitive-stress-responserelated proteins from Zea mays) (Shahollari et al. 2004,
Borner et al. 2005). Elicitation of tobacco cells with cryptogein results in recruitment of NADPH oxidase to the
phytosterol-rich domains (Mongrand et al. 2004), indicating that upon stimulation, plant rafts are able to mobilise a
specific set of proteins. Recently, Bhat et al. (2005)
showed that infection with a fungal pathogen leads to redistribution and polarisation of sterols in the plasma membrane of epidermal cells. This is accompanied by redistribution of plasma membrane proteins and followed
by focal accumulation of callose deposition at the site of
pathogen entry. In line with the observation that elicitors
induce NADPH recruitment to membrane microdomains
(Mongrand et al. 2004), these data indicate that generation
of phytosterol-rich macrodomains may participate in the
plant defence response.
Far-reaching similarities between plant and animal rafts
raise the possibility that regulatory processes that govern
raft dynamics may also be similar. Thus, plant annexins
could also be involved in the regulation of raft dynamics
in plant cells. So far only a very limited amount of colocalisation data supports this idea. In plant cells, annexins
are concentrated in the region of polarised growth, and in
vertebrate cells, raft domains are known to be especially
abundant in such region. Recently, Clark et al. (2005)
showed colocalisation of AnnAt1 and AnnAt2 with arabinogalactan proteins in Arabidopsis cells. Since classical
arabinogalactan proteins are predicted to have a GPI anchor, they may be a constituent of rafts. This may suggest
that plant annexins could also be connected with specialised raft domains and are therefore involved in their
functions.
However, the very basic question concerning the mechanism of annexin interaction with lipid rafts still remains
open. Only in the case of annexin A13b, this interaction
can be, at least partially, attributed to single-protein
myristoylation (Fig. 1) (Wice and Gordon 1992) (typically
double acylation is necessary to confer raft association),
whereas other proteins from this family do not undergo
even such posttranslational modifications. In principle, annexins bind to negatively charged phospholipids that are
distributed more or less uniformly within the inner leaflet,
while rafts, enriched in sphingolipids and sterols, develop
in the outer leaflet. However, it was shown that under certain conditions, individual annexins could display wider
binding preferences than it had been concluded from
in vitro experiments with artificial membranes. Annexins
A2 and A6 were shown to interact directly with cholesterol in native membranes isolated from chromaffin granules and CHO endosomes, respectively (Ayala-Sanmartin
et al. 2000, Diego et al. 2002). In turn, annexin A2 protects membrane cholesterol from extraction with cyclodextrin (Ayala-Sanmartin et al. 2001, Mayran et al.
2003). Furthermore, even low concentrations of cholesterol-sequestering agents like filipin or digitonin quantitatively released annexin 2 from the membranes (Jost et al.
1996, Harder et al. 1997). These data suggest that at least
certain annexins can bind to rafts through direct interaction with cholesterol. On the basis of recent experiments,
another possibility was also indicated. Annexin A2 was
shown to directly interact with another membrane component known to be located in rafts, phosphatidylinositol
4,5-bisphosphate (PIP2), despite the fact that the protein
lacks a typical PIP2-interacting domain (Hayes et al.
2004, Rescher et al. 2004). Recently, Gokhale et al.
(2005) proposed that PIP2 binding could be governed by a
cluster of cationic residues localised on the convex surface of the AnxA2 molecule (Fig. 1). In addition to the
well established role of PIP2 as a precursor of lipid-derived signalling molecules, it is known to act as a localised regulator of membrane events (vesicle fusion,
fission, and endocytosis) (Cullen et al. 2001, Martin
2001). Hence, Rescher and Gerke (2004) proposed that
recruitment of annexin A2 into rafts could be a two-step
process. After docking to the membrane through interactions with negatively charged phospholipids, the annexin
could be subsequently recruited into rafts via direct binding to PIP2. It remains to be established whether PIP2
binding is a generic feature of the whole family or just the
exclusive property of AnxA2. Since PIP2 is known to be
an important regulator of cytoskeleton function, the ability
of annexins to interact with PIP2 suggests that annexins
can connect rafts with the cortical cytoskeleton and organise the membrane–cytoskeleton interface. In line with this
reasoning, certain members of the vertebrate annexin family were shown to copurify with cytoskeleton proteins
(Gerke and Weber 1984, Shadle et al. 1985, Mangeat
1988), and perturbation of membrane cholesterol resulted
in the release of annexin A2 together with the actin cytoskeleton (Harder et al. 1997, Zeuschner et al. 2001). In
summary, it seems that in vertebrate cells, annexins facili-
D. Konopka-Postupolska: Annexins in regulation of membrane–cytoskeleton dynamics
tate rapid clustering of rafts in response to calcium elevation and stabilise such structures by hindering lateral
movements of lipids within the bilayer by attaching them
to the cytoskeleton. Although the significance of annexin
interaction with the actin cytoskeleton is not completely
clear, the linkage of annexins both to membranes and to
the actin cytoskeleton could be important for cortical cytoskeleton rearrangement in response to different stimuli
as well as for cytoskeleton interaction (see below).
Annexins and actin cytoskeleton dynamics
Several vertebrate annexins, namely, A1, A2, A5, and
A6, were shown to bind in a Ca2-dependent manner to
filamentous actin both in vitro and in vivo (Schlaepfer
et al. 1987, Khanna et al. 1990, Traverso et al. 1998,
Babiychuk et al. 1999, Tzima et al. 2000). Actin binding
is regulated also by posttranslational modification of annexin proteins, e.g., phosphorylation (Glenney and Tack
1985, Jost and Gerke 1996), S-glutathionylation (Caplan
et al. 2004) and poly-/multiubiquitination (Lauvrak et al.
2005). Annexins A1 and A2 not only bind but also induce
bundling of actin microfilaments in vitro (Schlaepfer et al.
1987, Glenney et al. 1987, Filipenko and Waisman 2001).
Besides actin, particular annexins can also bind accessory
proteins and hence influence the cytoskeleton dynamics
indirectly. Annexin A1 binds profilin, modulating its inhibitory effect on actin polymerisation. In turn, profilin
strongly inhibits annexin self-association and impairs its
ability to aggregate liposomes (Alvarez-Martinez et al.
1996, 1997). Furthermore, annexins A2 and A6 have been
shown to interact with spectrin, which is a member of the
two-dimensional network that lines the inner surface of
most metazoan cells (Bennet 1990). This raises the possibility that annexins could be engaged in remodelling of
this network, which is a prerequisite for the occurrence of
exocytosis and endocytosis (Kamal et al. 1998).
In contrast, only a few plant annexins, namely, p34 and
p35 from tomato (Calvert et al. 1996), zucchini (Hu et al.
2000), and mimosa plants (D. Hoshino et al. 2004), were
shown to bind filamentous actin (F-actin) but not globular
actin in a Ca2-dependent manner. Attachment to the microfilaments did not impair enzymatic activity of p34 and
p35 (ATPase/GTPase) from tomato in contrast to membrane binding that abolished it completely. Mimosa annexin induced bundling of actin in vitro and was shown to
be involved in pulvinar nyctinastic movements (D. Hoshino
et al. 2004).
It is not clear which protein domain is responsible for
annexin interaction with actin. There is no single proto-
209
typic domain of this type and quite divergent peptides support such function in different proteins. In the case of vertebrate annexin A2, the actin binding domain was mapped
experimentally to the very last C-terminal-sequence nonapeptide LLYLCGGDD (Fig. 1). A second peptide, VLIRIMVSR, localised a little towards the centre of the protein
molecule, was shown to be involved in actin bundling
(Jones et al. 1992, Filipenko and Waisman 2001). The former domain, only slightly modified, is also present in animal annexins A5 and A6 that are known to interact with
microfilaments, but no similar sequence was detected in
plant annexins. Instead, a conserved amino acid motif IRI,
previously identified as the smallest region of the putative
F-actin binding site of myosin (Suzuki et al. 1987), was
proposed to support this function. It was found in the 3rd
endonexin repeat in almost all plant annexins identified up
to now and in a subset of vertebrate annexins (A1, A2, A6,
A10, and A13) (Fig. 1). Direct experimental evidence supporting IRI-mediated attachment is lacking. Interestingly,
maize annexins p33 and p35, which share the IRI domain,
failed to interact with F-actin (Blackbourn et al. 1991).
However, under the experimental conditions of that study,
substantial amounts of actin remained in the supernatant
and could retain annexins and prevent coprecipitation. Besides, in those experiments heterologous actin from rabbit
muscle cells was used, potentially impairing the interaction with plant annexin. Summarising, further experiments
are necessary to precisely characterise the mechanism of
plant annexin interactions with actin. Moreover, it is necessary to undertake colocalisation studies of the annexins
and actin filament distribution to assess the significance of
in vitro binding data for cellular events.
For several in vivo experimental systems employing vertebrate cells, the interactions of annexins with the cytoskeleton were shown to take place exclusively on the
membrane surface. Annexins are specifically recruited to
those areas of cellular membranes that are associated with
the actin cytoskeleton, like nascent cell-to-cell contacts
(Hansen et al. 2002, Burkart et al. 2003, D. Lee et al. 2004).
Moreover, they localise to sites of dynamic actin assembly
platforms at cellular membranes, i.e., motile pinosomes
(Merrifield et al. 2001) and pedestals induced by enteropathogenic Escherichia coli (Zobiack et al. 2002). At
the same time, they are not found in cytosolic, actin-containing structures, like stress fibres (Gerke and Moss 2002)
or actin tails that propel intracellular movement of Listeria
species (Merrifield et al. 2001). In summary, it seems that
in vivo membranes are necessary to provide a platform for
annexin-induced actin assembly. The recent discovery that
annexins (or at least annexin A2) can directly bind PIP2,
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D. Konopka-Postupolska: Annexins in regulation of membrane–cytoskeleton dynamics
which is a well-known regulator of cytoskeleton–membrane adhesion, shed some light on this requirement. It is
widely accepted that activity of several cytoskeleton modulating and anchoring proteins can be regulated by PIP2
(Janmey and Stossel 1987, Yonezawa et al. 1990). Hence,
rapid and localised changes in PIP2 concentration resulting
from Ca2-induced activation of protein kinase C, PIP kinases, and phosphatases can regulate the membrane attachment of the cortical cytoskeleton and facilitate changes in
membrane curvatures. Annexins can mediate actin binding
with membranes via interaction with PIP2, thus anchoring
the actin cytoskeleton in dynamic regions engaged in exoand endocytosis and coupling together membrane and
actin-connected signalling cascades.
Another mechanism ensuring the specificity in actin–annexin interactions is also worth mentioning. It was shown
that in vertebrate cells, annexins can, in certain cases, display selectivity toward particular actin isoforms and thus
bind exclusively to a specific pool of cellular actin. Relocation of annexin A5 to the cytoskeleton in human platelets is
mediated by the specific interaction with actin isoform (Tzima et al. 1995). In smooth muscle cells, annexin A2
was shown to interact in a different way with a spatially
segregated “contractile” and “cytoskeletal” actin pool
(Babiychuk et al. 2002). Preferential association of annexins with a particular type of filaments would result in specificity of cell response to a given stimulus. Although there is
no direct experimental evidence testifying to the presence
of different types of actin microfilaments in plant cells,
there are some data supporting such reasoning. Plants express multiple actin isoforms encoded by an ancient gene
family (Meagher and McLean 1990). In the model organism A. thaliana, there are 8 genes (Meagher and Fechheimer
2003) expressed in a tissue-specific manner which thus can
be grouped in two general classes: generative and vegetative (ACT2, -7, and -8) (An et al. 1996). Of the vegetative
isoforms, ACT2 and ACT8 are the most closely related,
with only a single amino acid substitution (McDowell et al.
1996), but the expression of ACT8 is more restricted and
takes place only in a subset of ACT2-expressing tissues (An
et al. 1996). In contrast, ACT7 is expressed predominantly
in developing tissues, in dividing and expanding cells and
in response to most phytohormones (McDowell et al. 1996,
Kandasamy et al. 2001). This unequal expression of actin
isoforms in individual Arabidopsis leaf cells was confirmed
with single-cell RT-PCR (Laval et al. 2002). There is an ongoing debate concerning the purpose of expressing several
almost identical isoforms in a single cell at the same time.
One of the possibilities is that despite significant similarities, particular isoforms can differ in at least one activity. In
other words, the question is whether the isoforms are equivalent to each other and, if so, to what extent. A number of
observations from yeast and animal cells strongly suggest
that actin isoforms, although displaying about 87% homology, could have unique properties. For example, wild-type
yeast actin polymerised in vitro at a faster rate than muscle
actin because of differences in the nucleation step (Buzan
and Frieden 1996, Kim et al. 1996). For plant actin, the
question regarding the functional equivalency of different
isoforms remains open. The available data come exclusively from in vivo experiments with loss-of-function mutants and are contradictory. Disruption of most of the actin
genes (ACT1, -2, and -4) had only a mild effect on plant
morphology (McKinney et al. 1995, Gilliland et al. 2002),
which suggests that certain isoforms can easily substitute
for each other. Moreover, ectopic expression of both vegetative and generative isoforms complemented phenotype alteration in the act2-1 mutant and restored normal root hair
elongation impaired after mutation (Gilliland et al. 2002).
On the other hand, ectopic expression of generative actin
ACT1 had a great impact on plant morphology, caused
dwarfing and the alteration of some organs (Kandasamy
et al. 2002), hence testifying in favour of functional nonequivalency.
Another question is whether there are cellular mechanisms that allow sorting of different isoforms to separate
microfilaments. Double mutants act2-1 act7-1 developed a more severe phenotype than single mutants, which
the authors interpreted as evidence of synergism in the action of those two actins and argued against sorting of isovariants into specific filaments (Gilliland et al. 2002). Even
if so, induction of ACT7 gene expression in response to
hormonal treatment may result in formation of different filaments as a simple consequence of its over-representation
of one monomer in the total pool. Hence, it is tempting to
speculate that isoforms that are produced or modified in response to external stimulation (Dantan-Gonzalez et al.
2001, Wan et al. 2005) or hormonal induction (McDowell
et al. 1996, Kandasamy et al. 2001) can give rise to new
types of actin microfilament that are functionally different
or may interact with separate sets of actin-binding proteins. Alternatively, even a small contribution of a certain
isoform to the general pool of monomers can modify filament properties due to its selective recruitment into a given
region of microfilament. It was shown that there are some
mechanisms that govern the formation of microfilaments
with distinct compositions despite the availability of different monomers. Actins from rabbit muscle cytoskeleton and
plant pollen cells do not form heteropolymers in vitro, and
the introduction of rabbit actin into plant cells had a detri-
D. Konopka-Postupolska: Annexins in regulation of membrane–cytoskeleton dynamics
mental effect on the cells’ cytoarchitecture (Jing et al.
2003). Thus, if plant annexins share the ability to preferentially bind with different actin isoforms with animal counterparts, it is possible that they may also differentially
regulate actin interaction with membranes in stress conditions. Further analysis of plant annexin preferences toward
actin isovariants are necessary to elucidate if such a mechanism could really be valid.
Annexins in membrane trafficking
Annexins can also regulate membrane dynamics by influencing membrane turnover during exo- and endocytosis.
The story began with the isolation of the first annexin,
synexin (annexin A7), as a protein participating in Ca2regulated exocytosis in chromaffin cells (Creutz et al. 1992).
Since then new data has demonstrated that this is actually
the case and that annexins function in the process of
vesicular transport in all eukaryotic cells. For plants, immunolocalisation studies showed that annexins are highly
concentrated at the extreme tip of polarised growing cells
like pollen tubes of Lilium longiflorum (Blackbourn et al.
1992), maize roots, and fern rhizoids (Clark et al. 1994,
1995). Their localisation can be changed upon external
stimulation, e.g., in touch response of Bryonia dioica
(Thonat et al. 1993, 1997) and during gravity sensing in
pea plumules (Clark et al. 2000). Annexins were found to
be abundant in secretory cells of different types (outer
cells of root caps in maize and pea plants, developing vascular tissues in Arabidopis seedlings, and maize egg cells)
(Clark et al. 1992, 2000, 2001, 2005; Okamoto et al. 2004).
And finally, annexin expression was elevated during fruit
ripening when massive structural remodelling of the cell
wall took place (Wilkinson et al. 1995, Proust et al. 1996).
Considering all those data, it has been proposed that annexins may play a key role in cell expansion and organ
growth due to their function in Golgi-mediated secretion
of polysaccharide precursors for cell wall synthesis. Such
activity has been confirmed both in vitro and in vivo.
Maize and green pepper annexins were shown to induce
aggregation of artificial liposomes or plant secretory vesicles (Blackbourn and Battey 1993, T. Hoshino et al. 1995).
Additionally, maize annexin p35 potentiated the Ca2-driven exocytosis in maize root cap protoplasts (Carroll et al.
1998). More functional analysis performed recently by
Clark et al. (2005) revealed that such a scenario seemed to
be really operating. Localisation of particular annexins
(AnnAt1 and AnnAt2) in young Arabidopsis seedlings
was monitored by immunocytochemistry with specific antibodies and sites of active secretion of cell wall material
211
were detected by autoradiography of de novo synthesised
cell wall polysaccharides. Comparison of those two patterns revealed significant overlapping, thus indicating that
the localisation of annexins is temporally and spatially
coupled with secretion of cell wall material. Discrete differences that were observed in localisation of particular
annexins, e.g., only AnnAt1 was expressed in developing
sieve elements, additionally confirmed that despite functional equivalency, some annexins also fulfil specific functions in different cell types during plant development.
All those data combined testify that annexins can participate in regulated secretion in plant cells. The question
regarding the exact role of annexins in vesicular trafficking still remains open. To secrete their cargo, transporting
vesicles have to arrive at the target membrane first. In
plant cells this transport is supported by microfilaments
(Taylor and Hepler 1997, Battey and Blackbourn 1993).
Cytoskeleton proteins provide a mechanism for vesicle
movement, but they are not able to support fusion. Membrane fusion is an energetically unfavourable process and
is accomplished by a special set of proteins called
SNAREs (Rothman 1994). Special sets of complementary
SNAREs from different membranes interact, which results in docking and permits membrane fusion (Hughson
1999, Mellman and Warren 2000, Chen and Scheller
2001). This is accomplished first by the juxtaposition of
interacting organelles and then merging of the external
leaflets of the two apposing bilayers (hemifusion). In the
next step, merging of the two internal lipid leaflets occurs,
thereby completing the full fusion and establishing direct
continuity between the two previously separate compartments. Because of their ability to bind calcium and membrane lipids, annexins have been proposed to function as
CAPS (Ca2-dependent activator of protein secretion) in
Ca2-dependent exocytosis (Zorec and Tester 1992, Thiel
et al. 1994, Carroll et al. 1998). They can mediate hemifusion, thus enhancing the possibility of further full-fusion
events governed by SNARE proteins. In tight junction of
epithelial cells, annexin A2 was shown to promote stable
hemifusion, anchoring the cytoplasmic leaflets of secretory vesicles to each other or to the plasma membrane
(D. Lee et al. 2004). Strong support for this hypothesis
has come from experiments with heterologous expression
of mammalian annexins in yeast cells that do not express
endogenous proteins from this family. Annexins interacted
in a specific manner with the cellular secretory machinery
(Creutz et al. 1992) despite the lack of an endogenous
proteinaceous target, which strongly suggested that this
effect was due to interactions of annexins with membrane
elements of the yeast exocytosis machinery.
212
D. Konopka-Postupolska: Annexins in regulation of membrane–cytoskeleton dynamics
Further perspectives
Despite nearly 30 years of investigation, we are still far
from a comprehensive description of annexin function in
cell processes. Still much more is suggested than has been
established and much more remains to be done. Like
actin, annexins are so similar that they can easily substitute for each other, but this does not exclude the possibility that particular proteins are also specialised in specific
functions. Recently recognised active interplay between
membranes and the cytoskeleton makes annexins particularly interesting as they may function as a linker and modulate communication and cooperation between those two
cellular compartments. Their fusogenic activity also allows them to participate in secretion, which is particularly
important for plant cell growth. Understanding of annexin-mediated signal transduction could allow manipulation of plant stress response and in the future help in
obtaining more resistant crops.
Acknowledgments
The work in our laboratory is partially supported by Grant nr. 2P06A
007 29 from the Polish Ministry of Education and Science.
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