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Keywords:

  • annexin;
  • calcium;
  • cellular interactions;
  • comparative genomics;
  • functional motifs;
  • molecular evolution;
  • profile hidden Markov model (pHMM);
  • signal transduction

Abstract

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Structural analyses
  5. III. Membrane-related functions
  6. IV. Enzyme-related functions
  7. V. Functional insights from proteome and transcriptome analyses
  8. VI. Future perspectives
  9. Acknowledgements
  10. References

Contents

 Summary695
I.Introduction695
II.Structural analyses696
III.Membrane-related functions702
IV.Enzyme-related functions703
V.Functional insights from proteome and transcriptome analyses704
VI.Future perspectives706
 Acknowledgements708
 References708

Summary

Annexins are an homologous, structurally related superfamily of proteins known to associate with membrane lipid and cytoskeletal components. Their involvement in membrane organization, vesicle trafficking and signaling is fundamental to cellular processes such as growth, differentiation, secretion and repair. Annexins exist in some prokaryotes and all eukaryotic phyla within which plant annexins represent a monophyletic clade of homologs descended from green algae. Genomic, proteomic and transcriptomic approaches have provided data on the diversity, cellular localization and expression patterns of different plant annexins. The availability of 35 complete plant genomes has enabled systematic comparative analysis to determine phylogenetic relationships, characterize structures and observe functional specificity between and within individual subfamilies. Short amino termini and selective erosion of the canonical type 2 calcium coordinating sites in domains 2 and 3 are typical of plant annexins. The convergent evolution of alternate functional motifs such as ‘KGD’, redox-sensitive Cys and hydrophobic Trp/Phe residues argues for their functional relevance and contribution to mechanistic diversity in plant annexins. This review examines recent findings and advances in plant annexin research with special focus on their structural diversity, cellular and molecular interactions and their potential integrated functions in the broader context of physiological responses.

I. Introduction

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Structural analyses
  5. III. Membrane-related functions
  6. IV. Enzyme-related functions
  7. V. Functional insights from proteome and transcriptome analyses
  8. VI. Future perspectives
  9. Acknowledgements
  10. References

Calcium is a universal signal in eukaryotic cells and is involved in a wide variety of plant responses and processes (White & Broadley, 2003; Hetherington & Brownlee, 2004; Hepler, 2005). Most of the protein targets for these calcium signals are conserved throughout the eukaryotes (Nagata et al., 2004). A review of calcium proteomics in plants suggests that there are at least 200 different targets of calcium in plant cells. The annexin family of proteins is an important subset of these sensors and may also participate in generation of the calcium signatures (Reddy & Reddy, 2004). In plants, three Ca2+-regulatory protein motifs have been well characterized – the EF-hand motif, the C2 domain and the annexin domain unique to this superfamily (Kopka et al., 1998). Both C2 and annexin domains have the common defined property of binding anionic phospholipids in a calcium-dependent manner, but this is neither a universal nor an essential property for these domains, which may interact with other targets, including each other, by diverse mechanisms (Jiménez et al., 2003; Creutz & Edwardson, 2009).

Annexins are found in most eukaryotic cells, including multiple paralogs in unicellular organisms and single gene copies in certain algae and yeast. Twelve principal annexin subfamilies extend throughout vertebrate animals (ANXA1–ANXA13), represented by 12 paralogous genes in humans, although the actual gene number differs among species from 10 to 21 as a result of additional selection pressures; for example, three to five copies of ANXA1 in amphibians, three copies of ANXA8 in humans, whole-genome duplication in teleost fishes, and ANXA3 and ANXA9 absent from birds. The earliest diverging members, ANXA13 and ANXA7, have their origins in invertebrates, collectively grouped in family ANXB which may harbor dozens of distinct annexin subfamilies as a consequence of phylum-specific radiation and adaptation. The ANXC family pertaining to fungi and various protozoa is similarly diverse. The plant annexin family ‘ANXD’ is monophyletic and, although members have < 45% amino acid (aa) identity with animal annexins, they do preserve the unique annexin fold secondary structure assembled into the characteristic tetrad of four homologous domains (Hofmann, 2004; Moss & Morgan, 2004) and are therefore expected to share some basic interaction mechanisms and fundamental physiological roles analogous to animal annexins. Plant annexin repeats 2 and 3 have generally lost their type 2 calcium-binding signature and most plant annexins have a very short N-terminal region. The functional motifs and post-translational modifications of these unique N-terminal regions generate functional diversity in animal annexins, whereas variation of the primary sequence in core domains must be primarily responsible for the functional divergence of plant annexins.

The concept of annexins as calcium-dependent, phosphatidylserine-binding proteins possessing distinct amino termini is based on original studies of select vertebrate models. Recent research has sought to explain the functional specificity of individual subfamilies by identifying much broader variability in annexin properties for binding to calcium, membrane lipids, cytoskeletal proteins and other molecular interactions that direct their involvement in signaling networks and membrane trafficking (Moss & Morgan, 2004; Morgan et al., 2004, 2006). The cellular processes affected and functional roles inferred have likewise become even more diverse and complex as novel annexins from invertebrates, plants, protozoa and even bacteria are being discovered. The analysis of completed genomes offers testimony to the repeated occurrence of whole-genome duplications in all branches of life. Such events have been especially rampant in the plant kingdom, and the consequences are visible in the differential amplification and reciprocal loss within gene families of individual lineages, notably angiosperms (Sémon & Wolfe, 2007). The ability to distinguish characteristics common to all annexins or related clades from those responsible for unique subfamily specificity requires a more comprehensive and precise view of the structure–function diversity within the entire superfamily, combined with a classification scheme that documents measurable changes in their relationships. Gene family phylogeny fulfils the necessary criteria for evaluating functional changes imposed by structural constraint within individual subfamilies that can be identified and traced from their evolutionary history (Morgan & Fernandez, 1997; Morgan et al., 2006). Beyond the primary subfamily classification, profile hidden Markov models (pHMM) and 3D protein models have become indispensible for predicting functionally relevant features that are worthy of structural scrutiny and empirical testing.

Over the years there have been many reviews focusing on plant annexins (Clark & Roux, 1995; Delmer & Potikha, 1997; Hofmann, 2004; Mortimer et al., 2008; Konopka-Postupolska et al., 2011; Laohavisit & Davies, 2009, 2011). However, in this review we highlight the potentially divergent functions of this gene family in plants based on the considerable sequence and expression data that have been generated from public sequencing projects and proteomic and transcriptomic analyses as well as annexin studies. The growing interest in plant annexins arises because they are increasingly implicated as key players in a variety of physiological processes. Their expression is regulated developmentally and by a number of different environmental stimuli such as abiotic and biotic stresses. Additionally, purified preparations of certain plant annexins exhibit different biochemical activities, revealing their potential to act directly as phosphodiesterases or peroxidases, calcium channels, or F-actin binding proteins.

Here we present the evolutionary relationships and conserved structural features of this protein family in plants and in this context discuss the potential physiological roles of plant annexins. We analyze results from individual annexin studies as well as plant proteomic and transcriptome studies to gain deeper insights into the physiological roles of this gene family. We also provide a valid basis for using the phylogenetic classification of plant annexins with data from structural analyses so that existing and future functional data can be more readily associated with specific molecular features of individual subfamilies. Although we are still only beginning to gain glimpses into the structure and function of plant annexins, it seems clear that not only do they share many of the exciting characteristics of their animal counterparts but they have also diverged and play highly specialized roles apparent only in plants.

II. Structural analyses

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Structural analyses
  5. III. Membrane-related functions
  6. IV. Enzyme-related functions
  7. V. Functional insights from proteome and transcriptome analyses
  8. VI. Future perspectives
  9. Acknowledgements
  10. References

The random discovery and characterization of isolated plant annexins has obviated their unified classification and nomenclature and precluded a comprehensive overview of their structural and functional relationships with other annexins. The high-throughput sequencing of > 35 complete plant genomes, together with transcriptomic and proteomic profiling, has now enabled a systematic analysis of an essentially complete family. Alignment of homologous sequences is the first major step toward comparative studies that include phylogenetic analysis to define individual gene subfamilies, pHMM models to identify functional constraints on structural evolution, and structural models that incorporate evolutionary and mechanistic information in a realistic context.

A representative family tree for major phyla of Embryophyta (land plants) rooted in Chlorophyta green algae (Timme et al., 2012) was reconstructed from the full paralogous repertoires of known plant genomes originating up to 450 million yr ago (Fig. 1). These included two monocots (Oryza sativa and Zea mays) and three dicots (Arabidopsis thaliana, Medicago truncatula and Nicotiana benthamiana) to encompass the disparate patterns of gen(om)e duplications for each lineage. The most extensive, but possibly still incomplete, transcriptome sequence data representative of the six annexins in gymnosperms came from Picea glauca (Pg; spruce), while Ceratopteris richardii (Cr) and Adiantum capillus-veneris transcripts described up to five Pteridophyta (fern) annexins and the six known annexins from Selaginella moellendorffii (Sm) represent Lycophyta (club mosses). The completed genome of Physcomitrella patens (Pp) contains seven unique moss annexin subfamilies in Bryophyta. Annexins have been identified in three distinct orders-genera of Chlorophyta green algae, Chaetosphaeridiales-Coleochaete (Co; = 4), Charales-Nitella (Nh; = 15+) and Klebsormidiales-Klebsormidium (Kf; = 1), as well as completed Prasinophyta genomes including the smallest free-living eukaryote Ostreococcus lucimarinus (Ol; = 1). Other outgroup taxa consisted of single annexins from the Haptophyta Prymnesium parvum (Ppar; = 3) and the Chromista Aureococcus anophagefferens (Aa; golden brown algae; = 3–6). The early divergence and lineage-specific amplification of algal, moss, clubmoss and fern annexins justified their letter suffix-based classification, whereas the 40 distinct subfamilies of seed plants identified by phylogenetic analysis allowed for a global, numeric classification scheme for known annexins pertaining to gymnosperms and angiosperms. The subfamily numbering scheme follows the evolutionary branching order with high confidence levels (node bootstrap values) and branch lengths that reflect the relative amounts of evolution (i.e. aa replacements per site) for each taxon.

image

Figure 1. Phylogenetic tree and proposed nomenclature for plant annexin subfamilies. Full-length annexin protein sequences were selected from plant species representing fully sequenced genomes for eight major phyla (see inset). Evolutionary releationships were inferred by maximum likelihood analysis of 100 bootstrap alignments (316 aa × 71 taxa) based on the JTT matrix model with gamma rate correction using RAxML (Stamatakis et al., 2008). All known angiosperm annexins (> 1000) were classifiable according to this nomenclature scheme using the corresponding subfamily profile hidden Markov model (pHMM) models, and boostrap values at the nodes provide confidence in the subfamily branching order. Transcriptomic data from gymnosperms, ferns and club mosses may still be incomplete, true moss and green alga complete genomes may not be fully representative, and sequence data on liverworts are lacking, so the numbering scheme should be regarded as tentative. Subfamily names correspond to the genus-species abbreviation (see text section II. Structural analyses) followed by the ANXD plant annexin family root symbol suffixed with a unique subfamily number and, optionally, an additional letter for paralogous genes confined to a single genus or species; for example, Physcomitrella patens ANXD2A–ANXD2G represent seven paralogous annexins, all distinct from the six paralogous genes ANXD2A–ANXD2F in Lycophyta Selaginella moellendorffii, because both are unique, basal lineages like the algal ANXD1A, B, C… paralogs. The amplification of dicot subfamilies is distinct for the 13-member annexin repertoire in Medicago, compared with the eight annexins in Arabidopsis thaliana and other Brassicaceae contributing three novel subfamilies ANXD25, D40 and D41, and the 12 annexins in Nicotiana benthamiana and other Solanaceae which have six additional, duplicated genes, as shown in the figure. Similarly for monocots, the 11 rice annexins are supplemented by three duplicated (paralogous) genes ANXD15, D19 and D26 shown for Zea mays and also found in the Sorghum genome. The domain architecture and putative functional motifs for each annexin subfamily are offset to the right, with green/red annexin domains having/lacking type II calcium-binding motifs, K, R or H for conserved ‘KGD’, ‘RGD’ or ‘HGD’ motifs, W for tryptophan and C for cysteine in the A-B or D-E loops.

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An alignment of 400 full-length plant annexin protein sequences was subjected to statistical evolutionary analysis by hmmer3 (Finn et al., 2011) to generate a pHMM sequence model (Fig. 2). Its visualization using logomat (Schuster-Böckler et al., 2004) shows the expected aa frequency distribution (relative aa letter size) and the probability of functional constraint at each site (total column height), based on aa abundance and the expected frequency of replacement catalogued in substitution matrices. This representation of plant annexin primary protein structure is combined with secondary structure analysis to facilitate inferences about potentially important sites based on their unusual conservation pattern and their location accessibility within the structure. This pHMM model thus represents a ‘typical consensus’ plant annexin that incorporates evolutionary information from 400 representatives and facilitates the direct comparison with an analogous pHMM model previously computed for vertebrate animal annexins (Moss & Morgan, 2004). The calcium-coordinating sites of the annexin domain highlighted by arrows in Fig. 2 establish the prominent conservation of this motif (GxGT…38 residues…E/D) in plant annexin domain 1, a general loss of this capacity in plant annexin domains 2 and 3, and moderate conservation in tetrad domain 4 (see Fig. 1 also). As membrane binding is cooperative with respect to phosphatidylserine content (Almeida et al., 2005) but probably also includes interactions with protein complexes therein, it is prudent to consider other ‘potentially functional’ sites of annexins, especially those in the interhelical loop regions disposed to external interactions. A string of approximately eight hydrophobic amino acids in the final α-helix E of all annexin domains includes a highly conserved Trp-80 in plant domain 1, similar to the terminal tryptophans in domains 1, 2 and 4 of the protist Giardia lamblia, but in contrast to a simple leucine in vertebrate annexins. The Trp location near inter-domain loops suggests a possible role in membrane lipid insertion, translocation or binding affinity, potentially more significant in the context of calcium-independent annexin domains. This property may be even more pronounced for the conserved Trp-27 in plant domain 1, by analogy with the well-studied Trp-187 in domain 3 of ANXA5 (Sopkova de Oliveira Santos et al., 2001). Likewise, the greater prominence of Cys-111 (vs Leu) and absence of Cys-313 in plants compared with animals could affect redox sensitivity, tetrad 3D stability and protein interactions. The single codon loss in repeat 3 of all plants beyond ANXD13 should also be expected to slightly alter the tetrad tertiary structure. Finally, it is important to remember that this particular model summarizes the primary and secondary structures of many diverse plant annexins; the functional specificity of individual subfamilies would be more precisely defined by the corresponding 41 models based only on orthologs from individual subfamilies.

image

Figure 2. Profile hidden Markov model (pHMM) of the plant annexin family. hmmer3 (Finn et al., 2011) and logomat (Schuster-Böckler et al., 2004) were used to predict and visualize the predicted amino acid profile of a composite plant annexin protein sequence, based on an alignment of 400 full-length members. hmmer performed evolutionary statistical analysis to predict the expected amino acid frequency distribution (shown by relative letter size), while the total column stack height measures the information content attributable to evolutionary constraint, and hence reflects the predicted functional importance of each site. Calcium-coordinating residue sites are marked by down arrows, conserved Trp-80, Arg-87, Cys-111 and Phe-190 common to plants are marked by stars, and other conserved sites of inferred functional significance (tallest site stacks) can be compared with the analogous model for vertebrate annexins (Moss & Morgan, 2004).

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While the pHMM model contains much structural and evolutionary information with which to infer functional significance, the sequence loops between α-helices A-B and D-E warrant special attention because they are known to be associated with membrane interaction of the convex molecular face. To the extent that repeat 1 calcium-coordinating residues are generally more highly conserved than those in repeat 4, while repeats 2 and 3 have largely lost this function, the question arises as to the identity and conservation of other amino acids in these strategic sites in different plant annexin subfamilies. Table 1 was therefore compiled to summarize the actual aa composition at these sites, and shows marked divergence among the many different subfamilies. This ranges from the additional calcium-binding capacity in combination with proximal Cys conservation in early diverging subfamilies ANXD1–D13, to complete loss of predicted calcium-binding capability in at least eight plant subfamiles. This disparity makes it clear that generalizations would be unjustified regarding the mechanism, affinity and capacity of different plant annexins to bind cell membranes, whereas individual plant annexins are likely to exhibit very distinct binding properties, subcellular localization kinetics and molecular interactions.

Table 1. The amino acid sequences of the external loops linking α-helices A-B and D-E in each of the four homologous annexin domains are displayed to highlight those residues capable of coordinating calcium ions (black reverse shading) as part of the discontiguous, type II calcium-binding site considered canonical for most annexinsThumbnail image of

Regarding the predicted, fairly well-conserved calcium-binding sites that many plant annexins have in the first or fourth repeat, a study by Lim et al. (1998) found that a substitution of specific acidic residues with an asparagine residue resulted in loss of binding activity only when the change is made in repeat 4, suggesting that this repeat is indeed important in binding calcium. There are Giardia annexin sequences lacking a classical conserved calcium-binding loop, which can bind calcium with a modified loop (Bauer et al., 1999).

A study by Dabitz et al. (2005) highlights some differences between the membrane-binding properties of plant and mammalian annexins. They found that 18% of an ANXD36 protein sample from bell pepper (Capsicum annuum) or cotton (Gossypium hirsutum) bound to PC/PS vesicles (3 : 1) in the absence of calcium, considerably higher levels of Ca2+-independent membrane binding than found for most mammalian annexins. A study by Hofmann et al. (2002) indicates that another unique feature of plant annexins is their tendency to oligomerize independent of calcium binding. These studies and our analyses make it clear that the calcium-binding and membrane-binding properties for any particular plant annexin cannot be taken for granted and thus need to be tested experimentally.

All tertiary structure models of plant annexins are based only on members of the ANXD36 subfamily (Hofmann et al., 2000, 2002; Hu et al., 2008). A consensus sequence for plant annexin was generated from the pHMM model and this was threaded by I-Tasser (Roy et al., 2010) through the known crystal structures from cotton (3brx.pdb), bell pepper (1dk5.pdb) and Arabidopsis thaliana (2q4c.pdb) ANXD36. The complete 3D structure of this consensus from 400 plant annexins was rendered using Chimera (Pettersen et al., 2004) in two ribbon modes to illustrate the α-helices and their interconnecting loops (Fig. 3).

image

Figure 3. Protein 3D models for plant annexins. (a) A plant protein consensus sequence based on an alignment of 300 annexins was generated from the HMM model (Fig. 2) and threaded by I-Tasser (Roy et al., 2010) onto the crystallographic coordinates for experimentally resolved plant annexin family ANXD36 structures from cotton (3brx.pdb), green pepper (1dk5.pdb) and thale cress (2q4c.pdb). The rendering by Chimera (Pettersen et al., 2004) is presented in ribbon style for α-helices with predicted calcium-binding sites in the external loops joining helices A-B and D-E in each of the four repeat domains. The loss of the functional residues in these sites in plant repeats 2 and 3 is shown by alternative basic Lys residues, and other significant, conserved residues such as Trp-27, Trp-80, Arg-87, Cys-111 and C-terminal KGD sequences are likewise identified and discussed in the text. The lower figures were based on (b) domain 2 of Pinus pinaster (maritime cluster pine) ANXD10 and (c) domain 4 of Fragaria vesca (woodland strawberry) ANXD39. The former protein has an unusually elevated cysteine content (18 Cys in 330 aa) with the second domain featuring an aromatic tryptophan and external cysteine in place of the lost calcium-binding site at the membrane interface. The latter strawberry domain 4 contains a conserved ‘KGD’ motif in the same loop as the intact calcium-binding ligands.

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The loss of calcium-coordinating residues in plant annexin repeats 2 and 3 contrasts with their elevated conservation in repeat 1, where they are also associated with conserved Trp-27 and Trp-80, which may further reinforce plant annexin binding to membrane lipids. The pHMM model shows the isolated conservation of redox-sensitive Cys, hydrophobic Trp/Phe residues and ‘KGD’ motifs in these strategic inter-helical loops in individual plant subfamilies, and it highlights their probable contribution to external interactions that may be independent of or associated with calcium. With regard to the conserved redox-sensitive Cys residues, a review by Hofmann (2004) emphasizes that this unique aspect of plant annexin structures may be important with regard to H2O2/reactive oxygen species (ROS) signaling and reduction of H2O2 during response to stress responses.

As previously mentioned, animal annexins have longer N-terminal regions that serve as sites for post-translational modifications, such as phosphorylation, which regulate annexin function. By contrast, most plant annexins have very short amino termini, and post-translational modifications and differentiation of specific functions may occur within the core repeats, especially repeats 2 and 3. For example, within core repeats of annexins, both evidence for phosphorylation of serine and threonine residues, and identification of a 14-3-3 protein-like motif were recently discussed by Konopka-Postupolska et al. (2011). Additionally, evidence for modification of the redox-sensitive Cys residues in the core repeats suggests that these Cys residues are indeed accessible and reactive.

Arabidopsis thaliana AnnAt1 can be subject to S-nitrosylation (Lindermayr et al., 2005) or S-glutathionylation (Konopka-Postupolska et al., 2009). S-glutathionylation of A. thaliana AnnAt1 alters its calcium-binding affinity. Another reversible, post-translational modification of Cys residues is S-acylation and palmitoylation which can induce association with specific membrane compartments and/or lipid microdomains and alter cellular localization (Zhao & Hardy, 2004; Saric et al., 2009; Aicart-Ramos et al., 2011; Wang et al., 2011). In addition to conserved Cys residues, certain annexins have additional Cys residues that could be redox-sensitive and play a role in differentiating the function of an individual annexin. Interestingly, there do appear to be exceptions where plant annexins have protein domains in addition to the core repeats that help distinguish their function. For example, there is a rice (Oryza sativa) annexin with an extended N-terminal region and there are two Ceratopteris fern annexins which have longer than usual C-terminal regions, and these regions could serve to provide a source for functional differences for these annexins.

Studies on the C2 domain superfamily have indicated that there are differences in membrane-binding aa residues found on the surface of these proteins that could account for functional and biochemical diversity within this protein family (Jiménez et al., 2003; Zhang & Aravind, 2010). The importance of surface-exposed basic and hydrophobic residues for phospholipid binding, especially in the calcium-independent C2 subfamilies, finds clear analogy with annexin subfamilies where highly conserved Arg residues in all annexin domain B-C loops determine the molecular conformation and membrane-binding affinity (Campos et al., 1999). An additional role for basic residues and conserved Trp residues adjacent to the type II calcium-binding sites in annexin A5 (Mo et al., 2003; Sopkova de Oliveira Santos et al., 2001) can also be extended to equivalent structures observed in plant annexin domains 1 (e.g. Trp-27 and surface-exposed lysines). The recognition that charged, aromatic and cysteine residues in plant annexins may similarly participate in multiple interactions with membranes, cytoskeleton and other proteins as well as their response to changes in ROS and pH represents an important mechanistic and conceptual advance in understanding annexin action. For example, there are differences in the number and position of lysine and histidine residues for A. thaliana annexins, some of which are on the surface in their predicted 3D structures. Lysines can participate in membrane binding but are also subject to reversible post-translational modifications such as acetylation, which occurs in plants and can affect protein conformation and regulate activities (Finkemeier et al., 2011; Wu et al., 2011). It is also well documented that histidine residues on a variety of proteins play an important role as pH sensors in eukaryotic cells (Srivastava et al., 2007). For example, aquaporin has a histidine residue that can be protonated, leading to conformation changes and regulation of its transport function (Fischer & Kaldenhoff, 2008; Ludewig & Dynowski, 2009). In a similar fashion, the function of individual annexins in response to pH might be regulated via the protonation status of its histidine residues and resultant protein conformation changes. Experimental validation would be needed to determine if there is a role(s) for certain conserved and nonconserved aa residues in regulating the structure and function of a particular annexin.

III. Membrane-related functions

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Structural analyses
  5. III. Membrane-related functions
  6. IV. Enzyme-related functions
  7. V. Functional insights from proteome and transcriptome analyses
  8. VI. Future perspectives
  9. Acknowledgements
  10. References

Annexins were first discovered in animal cells and were named for their ability to ‘annex’ or aggregate membranes, and based on this property were suggested to participate in membrane fusion. Not surprisingly, many suggested functions for annexins are based on their ability to interact with cellular membranes. In fact, there is evidence that they associate with many different plant membranes, including the plasma membrane, endoplasmic reticulum, tonoplast, nuclear envelope, and outer chloroplast membrane. The paradigm for annexins is that they bind membranes when cytosolic calcium concentrations increase in response to certain stimuli, but in recent years it has become clear that certain annexins can also bind membranes in a calcium-independent manner, as already discussed. It is also clear that annexins have amphipathic properties and some of them can insert into membranes as monomers or hexamers (Luecke et al., 1995; Ladokhin & Haigler, 2005). Animal annexins also participate in the formation of lipid microdomains and lipid microdomain signaling (Babiychuk & Draeger, 2000) and plant annexins may function in a similar manner (Konopka-Postupolska et al., 2011). The earliest suggested function for plant annexins was participation in Golgi-mediated secretion based on their localization and ability to bind F-actin, and there is much evidence to support their role in the secretory pathway (Konopka-Postupolska, 2007; Konopka-Postupolska et al., 2011).

As Ca2+-dependent membrane-binding proteins, one of the hallmark characteristics of many annexins is their ability to dynamically change their cellular localization during certain physiological responses. Thus, there are numerous reports describing the redistribution of annexins in plant cells in response to a particular environmental stimulus. For example, Thonat et al. (1997) described a change in the localization of annexins in Bryonia dioica internode parenchyma cells from the cytoplasm to the plasma membrane that occurred within 30 min of touch stimulation. An immunocytochemistry study in etiolated pea (Pisum sativum) shoots demonstrated that plant annexins can also redistribute in response to gravistimulation within 15 min (Clark et al., 2000). Another immunolocalization study showed that the distribution of a Mimosa annexin in pulvinus motor cells changes depending on the time of day, suggesting a role for this annexin in nyctinastic movement (Hoshino et al., 2004).

Breton et al. (2000) describe redistribution of wheat (Triticum aestivum) annexins in response to cold treatment. Cold treatment induces changes in cytoplasmic calcium concentrations, and in this study, increased concentrations of two annexins were found in the plasma membrane fraction 30 min after cold treatment. Interestingly, these two annexins can also associate with the plasma membrane in a Ca2+-independent manner, acting like intrinsic membrane proteins. Both proteins accumulated in response to cold in two cultivars differing in their cold hardiness. Other studies have found annexin expression induced by cold (Renaut et al., 2004; Winfield et al., 2010), and thus certain annexins may be important in the general response to low temperatures. The receptor for the low temperature signal in plants is still unknown and it is tempting to speculate that changes in the membrane that involve annexins might contribute to the perception of cold.

Certain animal annexins are suggested to function as a novel type of Ca2+ channel. Plant annexins that affect Ca2+ influx could influence cell function in several different ways, ranging from acting as sensors or receptors early in signal transduction pathways, to playing an important part in creating certain Ca2+ signatures, to acting in directed Golgi-mediated secretion. A review of crystallization of calcium oxalate in plants has even speculated that annexins may be involved in this process (Webb, 1999). Typically, channel proteins contain one to seven transmembrane domains that anchor the channel into the membrane and provide the structural basis for the channel pore, but annexins are hypothesized to insert into membranes as oligomers and in this way act as channels (Kourie & Wood, 2000). Ca2+ channel activity has been demonstrated for certain animal annexins in vitro (Caohuy et al., 1996; Kirilenko et al., 2002; Donahue et al., 2004; Wang et al., 2005), and there is evidence supporting in vivo channel activity for annexin A5 in response to H2O2 (Kubista et al., 1999). Annexins may also indirectly affect channel and other membrane protein activities by altering membrane properties upon binding to membranes or integral membrane proteins (Kaetzel et al., 1994; Peng et al., 2004; Watson et al., 2004).

The first evidence that annexins might also affect Ca2+ influx in plant cells came from a study by Hofmann et al. (2000), which showed that a Capsicum annexin had Ca2+ channel activity in vitro and that this activity was higher when compared with animal annexins. White et al. (2002) proposed that annexins might function as hyperpolarization-activated cation channels (HACCs) in root cells. Consistent with this hypothesis, and with the observation that ROS activate plasma membrane Ca2+ channels in plants (Mori & Schroeder, 2004), maize (Zea mays) annexins were shown to exhibit channel activity in oxidized membranes (Laohavisit et al., 2010). Arabidopsis thaliana AnnAt1 (ANXD36), which is expressed in root cells, was also reported to exhibit pH-dependent channel activity (Gorecka et al., 2007) and maize annexins exhibit active Ca2+ conductance in lipid bilayers at mildly acidic pH (Laohavisit et al., 2009). Based on these findings, annexins were suggested to allow for influx of Ca2+ during stress responses involving acidosis.

A recent seminal study found that A. thaliana AnnAt1 is indeed responsible for Ca2+ and K+ transport regulated by extracellular ROS in root hairs and root epidermal cells (Laohavisit et al., 2012). A key finding was that loss-of-function annAt1 mutant root cells lacked the OH-activated Ca2+- and K+-permeable conductance. Moreover, the AnnAt1-mediated transport was distinct from the HACC transport activity in root cells. This important study raises the question of whether the many roles attributed to AnnAt1 (and its orthologs in other plant species) in plant cells are all attributable to its functioning as a mediator of plasma membrane transport, or can it function in other ways? Ultimately, it will be important to determine if there are other plant annexins that function similarly to AnnAt1 and whether certain other plant annexins act as modulators of channel activity.

IV. Enzyme-related functions

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Structural analyses
  5. III. Membrane-related functions
  6. IV. Enzyme-related functions
  7. V. Functional insights from proteome and transcriptome analyses
  8. VI. Future perspectives
  9. Acknowledgements
  10. References

The first enzyme activity discovered for plant annexins was an ATPase activity from a maize annexin (McClung et al., 1993). Later, Calvert et al. (1996) found phospholipid-dependent nucleotide phosphodiesterase activity associated with several tomato (Lycopersicon esculentum) annexins, and Lim et al. (1998) showed that this tomato annexin (ANXD39) enzyme activity was calcium-independent.

Shin & Brown (1999) demonstrated that a cotton fiber annexin (ANXD36) had a higher GTPase than ATPase activity and was inhibited by calcium. By deletion mapping it was shown that GTP binding of this annexin was dependent on the fourth domain. There are two predicted GTP-binding sites in this repeat present in the cotton and other annexins, and one of these overlaps with the Ca2+-binding site, which could explain the inhibition by Ca2+ ions. Annexins lack the classical Walker-like ATP-binding motif typically found in ATPases, but certain animal annexins have been shown to bind and hydrolyze ATP/GTP and a consensus nucleotide-binding sequence has been proposed (Bandorowicz-Pikula et al., 2003). Even though the in vitro ATP/GTPase activities of plant annexins are 2–3 orders of magnitude higher than that of animal annexins (Bandorowicz-Pikula, 2003), it still remains to be determined if this in vitro activity reflects an in vivo property important for function. The best potential link of this activity to function is suggested in a study by Carroll et al. (1998), who demonstrated that annexins, Ca2+ and GTP modulate polysaccharide secretion in maize root cap protoplasts.

Consistent with the hypothesis that nucleotide binding is an important feature for plant annexins, Shang et al. (2009) have proposed that AnnAt1 and/or AnnAt4 (ANXD20), both of which associate with the plasma membrane, might function as a receptor for extracellular ATP. Considering that extracellular ATP signaling is mediated by plasma membrane NADPH oxidase (Demidchik et al., 2009) and AnnAt1 Ca2+ transport activity is activated by hydroxyl radicals generated as a downstream consequence of NADPH oxidase activity (Laohavisit et al., 2012), this is an attractive hypothesis. Interestingly, Rubio et al. (2009) found that root cells of the annAt1 mutant were impaired in their release of ATP into the extracellular matrix (ECM) and in Ca2+ influx during response to salt stress.

Another enzyme activity attributed to certain plant annexins is inherent peroxidase activity (Gorecka et al., 2005; Laohavisit et al., 2009), which was originally suggested for A. thaliana AnnAt1 based on complementation results of the oxy5 bacterial strain and sequence similarity with heme peroxidases (Gidrol et al., 1996). Because it is very difficult to experimentally discriminate between intrinsic activity and activation of a co-purifying enzyme, the peroxidase activity of plant annexins has remained a controversial topic. However, it is clear that certain annexins provide oxidative protection to cells as there are numerous cases where ectopic expression or overexpression of an annexin provided transgenic plants with abiotic stress resistance by reducing H2O2 and/or lipid peroxidation levels (Jami et al., 2008; Konopka-Postupolska et al., 2009; Divya et al., 2010; Zhang et al., 2011; Zhou et al., 2011). Recently, annexin A2 was shown to act as a redox regulatory protein during tumorogenesis in mammalian cells (Madureira et al., 2011). This study provided evidence that a cysteine residue found near the N-terminus of annexin A2 can be oxidized by H2O2, then reduced by thioredoxin, making the cysteine residue available for further redox cycles. It is clear that AnnAt1 is not a classical heme-binding peroxidase (Laohavisit et al., 2009; Konopka-Postulpolska et al., 2009), so a possible alternative mechanism to explain this in vitro activity might be one similar to that found for annexin A2, in which AnnAt1 interacts with another protein allowing the conserved cysteine residues to function in redox cycling. Consistent with this possibility, a complex of proteins purified from Brassica rapa floral buds that has peroxidase activity contains AnnBr1 and a peroxiredoxin (Clark et al., 2010). Additional studies will be needed to determine whether purified plant annexins alone have inherent peroxidase activity, and, more importantly, whether this property represents an in vivo function or whether annexins protect cells from ROS by an alternative mechanism. The many signaling roles of ROS in a variety of plant processes underscore the need to resolve the specific function of annexins in ROS-mediated signal transduction pathways.

Annexins might also activate other enzymes by interacting with them. A number of reports suggest that plant annexins could regulate certain plasma membrane enzyme complexes such as callose or cellulose synthases (Verma & Hong, 2001; Hofmann et al., 2003). Callose (β1[RIGHTWARDS ARROW]3 glucan polymer) is synthesized during new cell wall plate formation, during secondary wall synthesis in cotton fibers and during pollen tube elongation. Andrawis et al. (1993) found that callose synthase activity isolated from cotton fiber membranes was inhibited by calcium and restored by EDTA washes that also removed several proteins, including, predominately, a 34-kDa annexin (ANXD36). By doing ‘add back’ experiments the authors demonstrated that this annexin was the most likely candidate for an interaction with callose synthase. These in vitro results may be indicative of annexin association with callose synthase as a way of directing the specific localization of this enzyme. Interestingly, two cotton fiber annexins were shown to bind UDP-glucose, and Brown (1999) proposed that they might act as UDP-glucose transporters, thus affecting callose synthase activity by regulating substrate concentrations. Callose production is also important in wounding and defense responses where certain annexins are implicated to play a signaling role. If plant glucan polymerization by β-(1[RIGHTWARDS ARROW]3) callose synthase is actually regulated by annexins, this would corroborate evidence from oomycetes where an annexin activates (1[RIGHTWARDS ARROW]3)-β-D-glucan synthase (Bouzenzana et al., 2006) and mimick an analogous role of annexins in remodeling lipid domains and cytoskeletal proteins in animal cell membranes.

There may be a role for annexins in new cell wall plate formation following cell division, an event that requires callose synthase activity, vesicle-mediated secretion and endocytosis, all processes in which annexins might be expected to function (Dhonukshe et al., 2006). In support of this role, Proust et al. (1999) identified two tobacco (Nicotiana tabacum) annexins (ANXD36 and ANXD37) that exhibited tissue-specific and cell cycle-dependent expression patterns. Annexin epitopes were localized to intercellular junctions in a ring structure during the exponential growth phase of the cells. The authors interpreted this pattern as indicating a possible role for annexin in cell plate formation.

There is evidence that annexins may also have a role in the redox regulation of cellulose synthase function (Kurek et al., 2002). Structural data on a cotton annexin suggest the presence of an S3 cluster that could function as a redox reactive center (Hofmann et al., 2003). Several other plant annexins have amino acids conserved in positions needed to create the putative S3 cluster, including A. thaliana AnnAt1. Hofmann (2004) suggested two possible ways that this putative S3 cluster could affect cellulose synthase activity. It could mediate the reduction of intermolecular disulphide bonds resulting in monomerization and inactivation of cellulose synthase, or promote the oxidation of cysteine residues in zinc finger motifs leading to formation of intramolecular disulphide bonds and disabling dimerization of cellulose synthase. Our phylogenetic analysis of a broader range of plant annexins has identified extraordinarily high cysteine content in basal conifer annexins (e.g. 18 Cys in pine (Pinus pinaster) ANXD10). This clearly reflects a role in redox sensitivity and homeostasis but may also substitute for type 2 calcium-binding ligands in the loops posed for membrane interaction (see Fig. 3). Protein cysteines and other reducing agents are frequently cross-linked constituents of tree sap resins, with attributed roles in secretion and protection from microbes and fungi, analogous to the proposed role of human, cysteine-rich annexin A10 in gastrointestinal mucus (Morgan et al., 1999).

In animal cells, annexins appear to function as nuclear proteins. For example, annexin A2 has a nuclear localization signal (Boyko et al., 1994) and is associated with the nuclear matrix where it plays an enzymatic role in DNA replication (Vishwanatha & Kumble, 1993). Annexin A2 and other animal annexins have been found in the nucleus or in association with the nuclear envelope, often after signal-induced translocation (Mohiti et al., 1997; Kim et al., 2003; Liu & Vishwanatha, 2007). An early report documented that several pea annexins were immunolocalized in the nucleus and nuclear periphery and co-purified with nuclear isolates (Clark et al., 1998). Similarly, an alfalfa (Medicago sativa) annexin was immunolocalized in the nucleolus and nuclear periphery (Kovacs et al., 1998). In plant cells there is evidence that Ca2+ concentrations are regulated in the nucleoplasm (Pauly et al., 2000; Mazars et al., 2011), and calcium is implicated in the regulation of transcription, especially in biotic and abiotic stress responses (Galon et al., 2010; Mazars et al., 2010), so annexins could participate in regulating nuclear calcium signals or be targets of this nuclear calcium signal. Consistent with this hypothesis, altered expression of certain annexins results in changes in expression of stress-related genes (Jami et al., 2008; Huh et al., 2010) and a tobacco root annexin (ANXD25) has been suggested to play a role in regulating gene expression (Baucher et al., 2012). Membrane budding was recently suggested as a novel mechanism for nuclear export of mRNA (Montpetit & Weis, 2012) and this is one way in which annexins could regulate gene expression.

A Medicago truncatula annexin, MtAnn1 (ANXD23), whose expression is induced by Nod factors, associates with the nuclear periphery (de Carvalho-Niebel et al., 2002). MtAnn1 expression was simultaneously induced in both outer and inner cell layers of the root by Nod-factor treatment or rhizobial infection, and it can even be induced by Nod factors in the absence of infection, allowing it to be used as a marker for Nod-induced processes that happen pre-infection. MtAnn1 expression in the inner root cells is mainly in endodermal cells, where AnnAt2 (ANXD39) mRNA and protein have also been localized in A. thaliana seedlings (Clark et al., 2001, 2005). MtAnn1 expression in the outer cortical cells of the root was cytoplasmic and peri-nuclear, and de Carvalho-Niebel et al. (2002) suggest that the peri-nuclear localization could indicate its interaction with nuclear associated endoplasmic reticulum and/or cytoskeleton where MtAnn1 might be involved in changes in nuclear envelope that occur during cell cycle progression. In addition to functioning in the nodulation process, nuclear MtAnn1 was also found in dividing cells, in recently differentiated cells in the root apex and in lateral root primordia, and probably has key functions in these sites as well.

V. Functional insights from proteome and transcriptome analyses

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Structural analyses
  5. III. Membrane-related functions
  6. IV. Enzyme-related functions
  7. V. Functional insights from proteome and transcriptome analyses
  8. VI. Future perspectives
  9. Acknowledgements
  10. References

Annexin gene families from A. thaliana, Brassica juncea, rice and tomato have been well characterized (Clark et al., 2001; Cantero et al., 2006; Jami et al., 2009, 2011; Lu et al., 2012), while data on annexins from many other plant species are being generated from proteomic and transcriptome studies. Several proteomic studies have identified the association of certain plant annexins with purified plasma membrane preparations (Shi et al., 1995; Santoni et al., 1998; Alexandersson et al., 2004). Importantly, Santoni et al. (1998) found that membrane-localized AnnAt1 protein has two different molecular masses, 34 and 39 kDa. These two isoforms of AnnAt1 protein show different hydrophobicity properties, with the 34-kDa isoform of AnnAt1 behaving more like an integral membrane protein, similar to the reported membrane-binding properties for a wheat annexin (Breton et al., 2000).

Certain animal annexins are found extracellularly even though they lack signal peptides. A recent paper provides evidence for a novel mechanism by which annexin A2 is secreted. This study found that association of annexin A2 with lipid rafts targets it to the intralumenal vesicles of multivesicular endosomes that are then secreted into the ECM, releasing annexin A2 to associate with the cell surface (Valapala & Vishwanatha, 2011). Coincidently, Regente et al. (2012) found that plants may also utilize exosome-like vesicles for nonclassical protein secretion. Recent evidence suggests a role for A. thaliana AnnAt3 (ANXD16) in post-Golgi vacuolar transport (Scheuring et al., 2011). Specifically, this study found that separation of trans-Golgi network and multivesicular body marker proteins was prevented in the RNAi-mediated knock-down of AnnAt3 expression.

One proteomic study identified an annexin in plant cell wall fractions (Kwon et al., 2005). In agreement with these results, pea annexins have previously been localized in the ECM at the ultrastructural level (Clark et al., 1992). More recently, Laohavisit et al. (2009) provided evidence that annexins might function extracellularly to activate Ca2+-permeable channels in plant cells. Although plants lack integrins they do have RGD receptors as well as C2 domain proteins, both of which are able to bind the RGD motif (Simões et al., 2005). Thus, the ‘KGD’ motif found in many plant annexins, which is observed in various forms as [R/K/H]-G-[D/E], could be an extracellular or intracellular ligand able to interact with these or other unidentified proteins.

This is the case for A. thaliana AnnAt1, which has an RGD motif in domain 4 and has been found in the ECM. If AnnAt1 is an extracellular protein, its redox regulatory activity could be important in regulating growth, as ECM peroxidases play an important role in controlling cell wall extensibility during environmentally and hormonally induced growth changes (Lee & Lin, 1995). Interestingly, there are membrane vesicles in the ECM of plant cells, and these vesicles may play important physiological roles (Regente et al., 2008, 2009; Gonorazky et al., 2010, 2012).

Bassani et al. (2004) used suppression subtraction hybridization to analyze growth-related genes in the elongation zone of primary root tips in maize and identified annexin p35 (ANXD35) as being up-regulated in the region of accelerating elongation. Similarly, Birnbaum et al. (2003) found that messages for AnnAt1 and AnnAt2 were abundant especially in the elongation and root hair zone. AnnAt3 and AnnAt4 were generally expressed at much lower levels, but had their highest levels of expression in the epidermis. In a recent proteomic study in roots, Tan et al. (2011) found that higher AnnAt2 protein levels were induced in wild-type and agravitropic pin2 root apices under altered gravity conditions and that AnnAt2 was differentially expressed in wild-type and pin2 mutant root cap columella cells.

As previously discussed, early work implicated annexins in abiotic stress responses in plants (Gidrol et al., 1996; Kovacs et al., 1998). In agreement with these reports, results from proteomic studies have found that annexins are up-regulated by salinity in a wide variety of plant species (Zhang et al., 2012), and several transcriptome studies have found that expression of annexins is regulated by drought (Bianchi et al., 2002; Watkinson et al., 2003; Krugman et al., 2011) and by aluminum or heavy metal treatment (Repetto et al., 2003; Chandran et al., 2008; Hradilova et al., 2010). More evidence of the annexin-stress connection is that expression of all eight A. thaliana annexins is regulated by various abiotic stresses (Cantero et al., 2006).

In an important proteomic study, Lee et al. (2004) reported that salt treatment decreased levels of immunodetectable AnnAt1 and induced cytoplasmic AnnAt1 to move to the membrane fraction. Based on these results, the authors suggested that salt treatment may cause AnnAt1 to be targeted to the proteasome for degradation, an exciting suggestion that needs to be further investigated. Interestingly, annexin A2 is ubiquitinated (Lauvrak et al., 2005) and AnnAt2 was identified as a putative ubiquitinated protein (Maor et al., 2007), but thus far there is no evidence for AnnAt1 ubiquitination. More recently, ectopic expression of AnnBj1, the Brassica annexin ortholog of AnnAt1, was found to improve resistance to drought and other biotic and abiotic stresses in transgenic tobacco plants (Jami et al., 2008). In agreement, AnnAt1 loss-of-function mutants in short-day conditions are drought sensitive and have higher epidermal concentrations of H2O2, while AnnAt1 gain-of-function mutants are drought tolerant and have lower epidermal concentrations of H2O2 (Konopka-Postupolska et al., 2009). However, in long-day conditions AnnAt1 and AnnAt4 appear to be negative regulators of drought and salt stress responses (Huh et al., 2010). This study found that AnnAt1 and AnnAt4 interact with each other and that the double mutant annAt1annAt4 is more tolerant to drought and salt stress than the single mutants. Although it is known that plant annexins can oligomerize, this study provides the first evidence that plant annexins form heterodimers.

Many angiosperm plant species are able to form symbiotic associations with the fungi from the Glomeromycota called arbuscular mycorrhizas. Several studies have implicated annexins in the formation of arbuscular mycorrhizas. Manthey et al. (2004) found that the Medicago truncatula annexin, MtAnn2, is induced in arbuscular mycorrhizas, just as it is in root nodules. Another study used suppressive subtraction hybridization to analyze late-stage arbuscular mycorrhizal development in peas and found that annexin was one of the transcripts induced by mycorrhizas (Grunwald et al., 2004). These and related studies are discussed in an excellent review focusing on the role of annexins in nodulation and mycorrhization in Medicago (Talukdar et al., 2009).

Fungal pathogen attack in a number of different plant species regulates annexin expression (Dowd et al., 2004; Guilleroux & Osbourn, 2004; Zhao et al., 2009). Interestingly, Ramonell et al. (2002) found that A. thaliana AnnAt2 expression was down-regulated by chitin elicitation and hypothesized that the down-regulation of this and several other genes by chitin could potentially ameliorate the host defense response during fungal attack. Importantly, ectopic expression of an annexin in cotton provides tolerance to Fusarium oxysporum infection and drought (Zhang et al., 2011). Relevant to this discussion is the identification of annexins in the mycelial cell wall proteome of Phytophthora infestans (Meijer et al., 2006; Grenville-Briggs et al., 2010) and the finding that an Oomycete annexin that can activate callose synthase is present in plasma membrane lipid microdomains (Briolay et al., 2009).

Annexin expression is up-regulated by pathogenic bacterial and viral attacks (Xiao et al., 2001; Marathe et al., 2004; Thiel & Varrelmann, 2009). A tobacco annexin, NtAnn12 (ANXD25), is induced in BY2 cells by infection with two pathogenic bacteria, Pseudomonas syringae and Rhodococcus fascians, but not Agrobacterium tumefaciens or Escherichia coli (Vandeputte et al., 2007). The expression of this annexin is light- and auxin-dependent and is found mainly associated with the nucleus in cortex cells of the root maturation zone (Baucher et al., 2011). Clearly, the expression of specific annexins is induced by pathogenic bacterial and viral attack, but a specific role for annexins in pathogen defense has not yet been defined. Antiviral activity of annexins A2 and A6 has been documented (Kwak et al., 2011; Ma et al., 2012) but many other studies indicate that cell surface annexins can be subverted to facilitate viral entry into host cells.

The phloem sap from several species contains proteins involved in stress and defense responses, including antioxidant defense enzymes (Walz et al., 2002, 2004). Annexins were identified in the phloem sap from Ricinus communis and Brassica napus based on 2D gel electrophoresis and mass spectrometry (Barnes et al., 2004; Giavalisco et al., 2006). One way in which annexins could function in phloem is through its known interactions with the cytoskeleton, which plays such a central regulatory role in sieve elements and plasmodesmata. Alternatively, phloem sap annexins could regulate callose synthase, an enzyme that is needed to close sieve tubes after wounding.

Annexins have been implicated in plant defense mechanisms and their suggested participation in membrane repair (Monastyrskaya et al., 2009; Schapire et al., 2009; Draeger et al., 2011; Konopka-Postulpolska et al., 2011) could be one potential function for annexins during wound responses. Insect feeding and jasmonic acid induce differential annexin expression in a number of plant species (Schmidt et al., 2005; Puthoff & Smigocki, 2007; Yan et al., 2007; Ralph et al., 2008; Gfeller et al., 2011), indicating a likely role for annexins in plants’ wound responses to insect attack. Annexins have also been implicated in wounding responses in potato (Solanum tuberosum) tubers (Murphy et al., 2010; Urbany et al., 2012). Annexins are even found in latex, which has an important function in rubber (Hevea brasiliensis) plants’ defense against insect attack (Han et al., 2000). Nematode attack induces annexin expression (Klink et al., 2007) and an annexin secreted by nematodes was shown to play a significant role in nematode parasitism (Patel et al., 2010). This nematode annexin interacts with an A. thaliana oxidoreductase previously found to play an important role in susceptibility to downy mildew pathogens (Van Damme et al., 2008). Interestingly, the nematode annexin complements the annAt1 mutant stress seed germination phenotype, leading to the suggestion that it might mimic AnnAt1 function during infection.

Some studies have suggested novel and unexpected functions for annexins. For example, in their examination of the chloroplast transcription apparatus from mustard (Sinapis alba), Loschelder et al. (2004) isolated an annexin similar to A. thaliana AnnAt4 (both ANXD20) as a binding partner in the chloroplast-encoded RNA polymerase complex. Link (2003) suggested that this annexin might play a role in redox regulation of chloroplast transcription. Okamoto et al. (2004) found that annexin p35 was one of the major proteins present in egg cells and zygotes in maize. In this study, p35 annexin appeared to be egg-cell specific, which suggested that it could play a role in the fertilization-induced increase in secretion that is stimulated by an increase in cytoplasmic Ca2+ and associated with the production of a new cell wall surrounding the zygote. Another study comparing expression levels in tobacco egg cells and zygotes using subtractive hybridization found that the expression of a tobacco annexin was present in egg cells but was reduced in zygotes (Ning et al., 2006). An A. thaliana microarray study carried out by Honys & Twell (2004) compared expression levels of many different genes in pollen and found relatively high levels, and significant changes in expression, of transcripts for AnnAt1, AnnAt2 and AnnAt5 (ANXD2) at various stages of pollen development.

All eight A. thaliana annexins are present in seeds and germinating seedlings (Cantero et al., 2006), and several groups have published data pointing to high levels of annexin expression during seed development (Gallardo et al., 2003; Beilinson et al., 2005). Recently, it was shown that ectopic expression of a lotus (Nelumbo nucifera) annexin in A. thaliana provides seed germination vigor and thermotolerance during seed germination (Chu et al., 2012).

Certain annexins are differentially regulated during fruit development and ripening. Transcriptome analysis of ripening in strawberry fruit revealed that transcript levels of a strawberry (Fragaria ananassa) annexin dramatically increased at this time (Wilkinson et al., 1995; Wang et al., 2001). Gene expression analysis of strawberry achene and receptacle tissues during maturation revealed an annexin with a similar expression profile that was more highly expressed in receptacle tissue. This annexin could play a role in ROS detoxification that would be needed because ROS are generated in the receptacle tissue during the ripening process (Aharoni & O'Connell, 2002). It could also associate with Golgi to help mediate delivery of cell wall-loosening agents in the receptacle tissue during ripening. There is an annexin-ripening connection also in bell peppers (Capsicum annum), where two annexins are up-regulated during fruit development and ripening (Proust et al., 1996). Other fruit functions of annexins may be unrelated to ripening. For example, annexins are up-regulated in cherimoya (Annona cherimola) fruit after cold treatment (González-Agüero et al., 2011). The nine annexins in tomato fruit are hypothesized to play a role in the blossom-end rot believed to be caused by calcium deficiencies during cell elongation at the rapidly expanding tips of tomato fruit (Ho & White, 2005).

VI. Future perspectives

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Structural analyses
  5. III. Membrane-related functions
  6. IV. Enzyme-related functions
  7. V. Functional insights from proteome and transcriptome analyses
  8. VI. Future perspectives
  9. Acknowledgements
  10. References

Our current perspective on annexins in plant cells is that they are a diverse, multigene protein family that has been associated with a variety of processes and responses. A speculative model of plant annexin function highlights the different cellular locales and activities that have been discussed in this review (Fig. 4). Despite the great increase in information on plant annexins, there is still a general lack of specific details and direct experimental evidence for how annexins function in these processes and responses. Even with respect to their key biochemical feature, their interaction with calcium and membranes, there is still much that we do not understand. Structurally, the paradigm for plant annexins as calcium-dependent membrane-binding proteins clearly has exceptions. There appear to be some important similarities as well as important differences with animal annexins. While the independent emergence and strategic conservation of equivalent motifs in both kingdoms mark them as key functional determinants, the characterization of additional structure–function motifs unique to plant annexins is forthcoming. The verification of true ligand–receptor molecular interactions for plant annexins will be crucial to decipher basic mechanisms and active physiological processes. The application of bioinformatic analyses to the identification of regulatory motifs in plant annexin promoters, microRNAs, etc. could be especially useful for interpreting expression response studies.

image

Figure 4. Model illustrating potential functions of annexins in plant cells. Developmental or environmental signals can induce changes in calcium, pH and reactive oxygen species (ROS) which can result in structural and/or post-translational modifications of plant annexins. Specific annexins may function differently and in different cellular locales such as the extracellular matrix (ECM) or in association with different membranes or organelles.

Download figure to PowerPoint

In addition to their calcium- and membrane-binding activities, there are so many other reported activities for plant annexins that it is difficult to assign a specific function to any one annexin, leaving even annexin specialists with a myriad of unanswered questions. Can certain plant annexins act as receptors as they appear to do in animal cells, and, if so, is in vivo calcium transport activity important for this function? Which post-translational modifications are found on which plant annexins and what cellular activities do these modifications modulate? Are there examples where plant annexins or their polymorphic alleles function together to contribute to a phenotype controlled by quantitative trait loci? What are the specific protein–protein and network interactions for an individual plant annexin? Do plant annexins interact with annexins from fungal pathogens or nematodes to play roles in invasion or wound healing? Do plant annexins function in lipid microdomains and in membrane repair? Are all the annexins found in a particular plant species functionally relevant and are there levels of redundancy?

One answer that could impact all these questions may be that annexins are generally important in helping to maintain calcium homeostasis, especially in membrane domains. They may display isovariant dynamics in their cellular roles, a feature that has also been proposed to be characteristic of the actin multigene family. Isovariant dynamics can increase control and plasticity in plant signaling pathways by allowing a family of related signaling proteins to have distinct outputs during signal transduction via differences found in functional domains and protein–protein interactions (Meagher et al., 1999, 2008). In plant cells where multiple annexins are present, the isovariant dynamics model would predict a number of ways that different annexins could affect different outcomes in various signaling pathways. Each annexin could fill a slightly different niche in a signaling pathway, allowing fine-tuning of a response.

Whereas many of the cellular binding partners for various animal annexins have been well characterized, to date not much is known about the protein–protein interactions of plant annexins. Although methods for the computational analyses of plant gene networks are becoming available (Lee et al., 2010, 2011), determining experimentally the specific protein–protein and network interactions for individual plant annexins will be important to understanding the basis of their function.

The observation thus far that knocking out a single annexin in plants does not appear to cause lethality or result in an easily detectable phenotype under normal growth conditions suggests that annexins may function more as regulatory components of signaling pathways and membrane homeostasis or that there is a high level of redundancy in their function. Finally, the presence of either a single annexin or more than a dozen paralogs in different unicellular algae (e.g. Micromonas vs Nitella) suggests that their adaptive significance may vary considerably among different species.

In relating the evolutionary relationships of plant annexins with structural and physiological data available we have attempted to provide a bridge between structure, mechanism and function that can serve as a useful tool for plant annexin researchers. We also hope this review will inspire and spur further interest and new ideas regarding this family of proteins and their potential functions in plant cells.

Acknowledgements

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Structural analyses
  5. III. Membrane-related functions
  6. IV. Enzyme-related functions
  7. V. Functional insights from proteome and transcriptome analyses
  8. VI. Future perspectives
  9. Acknowledgements
  10. References

Grant BFU2007-67876 support for R.O.M. and M.P.F. came from the Spanish Ministry of Science and Innovation (MICINN), and UNOV-11-MA-14 came from the University of Oviedo, Spain. Support for S.J.R. and G.C. came from National Science Foundation Grant No. 0718890 and No. 1027514. We thank Drs Dorota Konopka-Postulpolska and Ann Kleinschmidt for helpful discussions on annexins.

References

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Structural analyses
  5. III. Membrane-related functions
  6. IV. Enzyme-related functions
  7. V. Functional insights from proteome and transcriptome analyses
  8. VI. Future perspectives
  9. Acknowledgements
  10. References
  • Aharoni A, O'Connell AP. 2002. Gene expression analysis of strawberry achene and receptacle maturation using DNA microarrays. Journal of Experimental Botany 53: 20732087.
  • Aicart-Ramos C, Ana Valero R, Rodriguez-Crespo I. 2011. Protein palmitoylation and subcellular trafficking. Biochimica et Biophysica Acta-Biomembranes 1808: 29812994.
  • Alexandersson E, Saalbach G, Larsson C, Kjellbom P. 2004. Arabidopsis plasma membrane proteomics identifies components of transport, signal transduction and membrane trafficking. Plant Cell Physiology 45: 15431556.
  • Almeida PFF, Sohma H, Rasch KA, Wieser CM, Hinderliter A. 2005. Allosterism in membrane binding: a common motif of the annexins? Biochemistry 44: 1090510913.
  • Andrawis A, Solomon M, Delmer DP. 1993. Cotton fiber annexins – a potential role in the regulation of callose synthase. Plant Journal 3: 763772.
  • Babiychuk EB, Draeger A. 2000. Annexins in cell membrane dynamics: calcium-regulated association of lipid microdomains. Journal Cell Biology 150: 11131123.
  • Bandorowicz-Pikula J. 2003 The nucleotide face of annexins. In: Bandorowicz-Pikula J, ed. Annexins: biological importance and annexin-related pathologies. New York, NY, USA: Kluwer Academic Publishers, 234256.
  • Bandorowicz-Pikula J, Kirilenko A, van Deursen R, Golczak M, Kuhnel M, Lancelin JM, Pikula S, Buchet R. 2003. A putative consensus sequence for the nucleotide-binding site of annexin A6. Biochemistry 42: 91379146.
  • Barnes A, Bale J, Constantinidou C, Ashton P, Jones A, Pritchard J. 2004. Determining protein identity from sieve element sap in Ricinus communis L. by quadrupole time of flight (Q-TOF) mass spectrometry. Journal Experimental Botany 55: 14731481.
  • Bassani M, Neumann PM, Gepstein S. 2004. Differential expression profiles of growth-related genes in the elongation zone of maize primary roots. Plant Molecular Biology 56: 367380.
  • Baucher M, Oukouomi Lowe Y, Vandeputte OM, Mukoko Bopopi J, Moussawi J, Vermeersch M, Mol A, El Jaziri M, Homblé F, Pérez-Morga D. 2011. Ntann12 annexin expression is induced by auxin in tobacco roots. Journal of Experimental Botany 62: 40554065.
  • Baucher M, Pérez-Morga D, El Jaziri M. 2012. Insight into plant annexin function: from shoot to root signaling. Plant Signaling & Behavior 7: 15.
  • Bauer B, Engelbrecht S, Bakker-Grunwald T, Scholze H. 1999. Functional identification of α1-giardin as an annexin of Giardia lamblia. FEMS Microbiology Letters 173: 147153.
  • Beilinson V, Moskalenko OV, Ritchie RD, Nielsen NC. 2005. Differentially expressed genes during seed development in soybean. Physiologia Plantarum 123: 321330.
  • Bianchi MW, Damerval C, Vartanian N. 2002. Identification of proteins regulated by cross-talk between drought and hormone pathways in Arabidopsis wild-type and auxin-insensitive mutants, axr1 and axr2. Functional Plant Biology 29: 5561.
  • Birnbaum K, Shasha DE, Wang JY, Jung JW, Lambert GM, Galbraith DW, Benfey PN. 2003. A gene expression map of the Arabidopsis root. Science 302: 19561960.
  • Bouzenzana J, Pelosi L, Briolay A, Briolay J, Bulone V. 2006. Identification of the first Oomycete annexin as a (1,3)-β-D-glucan synthase activator. Molecular Microbiology 62: 552565.
  • Boyko V, Mudrak O, Svetlova M, Negishi Y, Ariga H, Tomilin N. 1994. A major cellular substrate for protein kinases, annexin II, is a DNA-binding protein. FEBS Letters 345: 139142.
  • Breton G, Vasquez-Tello A, Danyluk J, Sarhan F. 2000. Two novel intrinsic annexins accumulate in wheat membranes in response to low temperature. Plant Cell Physiology 41: 177184.
  • Briolay A, Bouzenzana J, Guichardant M, Deshayes C, Sindt N, Bessueille L, Bulone V. 2009. Cell wall polysaccharide synthases are located in detergent-resistant membrane microdomains in Oomycetes. Applied and Environmental Microbiology 75: 19381949.
  • Brown RM. 1999. Cellulose structure and biosynthesis. Pure Applied Chemistry 71: 767775.
  • Calvert CM, Gant SJ, Bowles DJ. 1996. Tomato annexins p34 and p35 bind to F-actin and display nucleotide phosphodiesterase activity inhibited by phospholipid binding. Plant Cell 8: 333342.
  • Campos B, Wang S, Retzinger GS, Kaetzel MA, Seaton BA, Karin NJ, Johnson JD, Dedman JR. 1999. Mutation of highly conserved arginine residues disrupts the structure and function of annexin V. Archives of Medical Research 30: 360367.
  • Cantero A, Barthakur S, Bushart T, Morgan RO, Fernandez MP, Chou S, Clark G, Roux SJ. 2006. Expression profiling of the Arabidopsis annexin gene family during abiotic stress, germination and de-etiolation. Plant Physiology and Biochemistry 44: 1324.
  • Caohuy H, Srivastava M, Pollard HB. 1996. Membrane fusion protein synexin (annexin VII) as a Ca2+/GTP sensor in exocytotic secretion. Proceedings of the National Academy of Sciences, USA 93: 1079710802.
  • Carroll AD, Moyen C, Van Kesteren P, Tooke F, Battey NH, Brownlee C. 1998. Ca2+, annexins, and GTP modulate exocytosis from maize root cap protoplasts. Plant Cell 10: 12671276.
  • de Carvalho-Niebel F, Timmers ACJ, Chabaud M, Defaux-Petras A, Barker DG. 2002. The Nod factor-elicited annexin MtAnn1 is preferentially localised at the nuclear periphery in symbiotically activated root tissues of Medicago truncatula. Plant Journal 32: 343352.
  • Chandran D, Sharopova N, Ivashuta S, Gantt JS, VandenBosch KA, Samac DA. 2008. Transcriptome profiling identified novel genes associated with aluminum toxicity, resistance and tolerance in Medicago truncatula. Planta 228: 151166.
  • Chu P, Chen H, Zhou Y, Li Y, Ding Y, Jiang L, Tsang EWT, Wu K, Huang S. 2012. Proteomic and functional analyses of Nelumbo nucifera annexins involved in seed thermotolerance and germination vigor. Planta 235: 12711288.
  • Clark G, Konopka-Postupolska D, Hennig J, Roux S. 2010. Is annexin 1 a multifunctional protein during stress responses? Plant Signaling & Behavior 5: 303307.
  • Clark GB, Dauwalder M, Roux SJ. 1992. Purification and immunolocalization of annexin–like protein in pea seedlings. Planta 187: 19.
  • Clark GB, Dauwalder M, Roux SJ. 1998. Immunological and biochemical evidence for nuclear localization of annexin in peas. Plant Physiology & Biochemistry 36: 621627.
  • Clark GB, Lee DW, Dauwalder M, Roux SJ. 2005. Immunolocalization and histochemical evidence for the association of two different Arabidopsis annexins with secretion during early seedling growth and development. Planta 220: 621631.
  • Clark GB, Rafati DS, Bolton RJ, Dauwalder M, Roux SJ. 2000. Redistribution of annexin in gravistimulated pea plumules. Plant Physiology & Biochemistry 38: 937947.
  • Clark GB, Roux SJ. 1995. Annexins of plant cells. Plant Physiology 109: 11331139.
  • Clark GB, Sessions A, Eastburn DJ, Roux SJ. 2001. Differential expression of members of the annexin multigene family in Arabidopsis. Plant Physiology 126: 10721084.
  • Creutz CE, Edwardson JM. 2009. Organization and synergistic binding of copine I and annexin A1 on supported lipid bilayers observed by atomic force microscopy. Biochimica Biophysica Acta 1788: 19501961.
  • Dabitz N, Hu NJ, Yusof AM, Tranter N, Winter A, Daley M, Zschornig O, Brisson A, Hofmann A. 2005. Structural determinants for plant annexin-membrane interactions. Biochemistry 44: 1629216300.
  • Van Damme M, Huibers RP, Elberse J, Van den Ackerveken G. 2008. Arabidopsis DMR6 encodes a putative 2OG-Fe(II) oxygenase that is defense-associated but required for susceptibility to downy mildew. Plant Journal 54: 785793.
  • Delmer DP, Potikha TS. 1997. Structures and functions of annexins in plants. Cell Molecular Life Science 53: 546553.
  • Demidchik V, Shang Z, Shin R, Thompson E, Rubio L, Laohavisit A, Mortimer J, Chivasa S, Slabas A, Glover B, et al. 2009. Plant extracellular ATP signaling by plasma membrane NADPH oxidase and Ca2+ channels. Plant Journal 58: 903913.
  • Dhonukshe P, Baluska F, Schlicht M, Hlavacka A, Samaj J, Friml J, Gadella TWJ. 2006. Endocytosis of cell surface material mediates cell plate formation during plant cytokinesis. Developmental Cell 10: 137150.
  • Divya K, Jami SK, Kirti PB. 2010. Constitutive expression of mustard annexin, AnnBj1 enhances abiotic stress tolerance and fiber quality in cotton under stress. Plant Molecular Biology 73: 293308.
  • Donahue TLH, Genetos DC, Jacobs CR, Donahue HJ, Yellowley CE. 2004. Annexin V disruption impairs mechanically induced calcium signaling in osteoblastic cells. Bone 35: 656663.
  • Dowd C, Wilson IW, McFadden H. 2004. Gene expression profile changes in cotton root and hypocotyl tissues in response to infection with Fusarium oxysporum f. sp. Vasinfectum. Molecular Plant-Microbe Interactions 17: 654667.
  • Draeger A, Monastyrskaya K, Babiychuk EB. 2011. Plasma membrane repair and cellular damage control: the annexin survival kit. Biochemical Pharmacology 81: 703712.
  • Finkemeier I, Laxa M, Miguet L, Howden AJM, Sweetlove LJ. 2011. Proteins of diverse function and subcellular location are lysine acetylated in Arabidopsis. Plant Physiology 155: 17791790.
  • Finn RD, Clements J, Eddy SR. 2011. HMMER web server: interactive sequence similarity searching. Nucleic Acids Research 39: W2937.
  • Fischer M, Kaldenhoff R. 2008. On the pH regulation of plant aquaporins. Journal of Biological Chemistry 283: 33889892.
  • Gallardo K, Le Signor C, Vandekerckhove J, Thompson RD, Burstin J. 2003. Proteomics of Medicago truncatula seed development establishes the time frame of diverse metabolic processes related to reserve accumulation. Plant Physiology 133: 664682.
  • Galon Y, Finkler A, Fromm H. 2010. Calcium-regulated transcription in plants. Molecular Plant 3: 653669.
  • Gfeller A, Baerenfaller K, Loscos J, Chetelat A, Baginsky S, Farmer EE. 2011. Jasmonate controls polypeptide patterning in undamaged tissue in wounded Arabidopsis leaves. Plant Physiology 156: 17971807.
  • Giavalisco P, Kapitza K, Kolasa A, Buhtz A, Kehr J. 2006. Towards the proteome of Brassica napus phloem sap. Proteomics 6: 896909.
  • Gidrol X, Sabelli PA, Fern YS, Kush AK. 1996. Annexin-like protein from Arabidopsis thaliana rescues delta oxyR mutant of Escherichia coli from H2O2 stress. Proceedings of the National Academy of Sciences, USA 93: 1126811273.
  • Gonorazky G, Laxalt AM, de la Canal L. 2010. Involvement of phospholipase C in the responses triggered by extracellular phosphatidylinositol 4-phosphate. Journal of Plant Physiology 167: 411415.
  • Gonorazky G, Laxalt A, Dekker H, Rep M, Munnik T, Testerink C, de la Canal L. 2012. Phosphatidylinositol 4-phosphate is associated to extracellular lipoproteic fractions and is detected in tomato apoplastic fluids. Plant Biology 14: 4149.
  • González-Agüero M, Cifuentes-Esquivel N, Ibañez-Carrasco F, Gudenschwager O, Campos-Vargas R, Defilippi BG. 2011. Identification and characterization of genes differentially expressed in cherimoya (Annona cherimola Mill) after exposure to chilling injury conditions. Journal of Agricultural and Food Chemistry 59: 1329513299.
  • Gorecka KM, Konopka-Postupolska D, Hennig J, Buchet R, Pikula S. 2005. Peroxidase activity of annexin 1 from Arabidopsis thaliana. Biochemistry Biophysics Research Communication 336: 868875.
  • Gorecka KM, Thouverey C, Buchet R, Pikula S. 2007. Potential role of annexin AnnAt1 from Arabidopsis thaliana in pH-mediated cellular response to environmental stimuli. Plant Cell Physiology 48: 792803.
  • Grenville-Briggs LJ, Avrova AO, Hay RJ, Bruce CR, Whisson SC, Van West P. 2010. Identification of appressorial and mycelial cell wall proteins and a survey of the membrane proteome of Phytophthora infestans. Fungal Biology 114: 702723.
  • Grunwald U, Nyamsuren O, Tarnasloukht M, Lapopin L, Becker A, Mann P, Gianinazzi-Pearson V, Krajinski F, Franken P. 2004. Identification of mycorrhiza-regulated genes with arbuscule development-related expression profile. Plant Molecular Biology 55: 553566.
  • Guilleroux M, Osbourn A. 2004. Gene expression during infection of wheat roots by the ‘take-all’ fungus Gaeumannomyces graminis. Molecular Plant Pathology 5: 203216.
  • Han KH, Shin DH, Yang J, Kim IJ, Oh SK, Chow KS. 2000. Genes expressed in the latex of Hevea brasiliensis. Tree Physiology 20: 503510.
  • Hepler PK. 2005. Calcium: a central regulator of plant growth and development. Plant Cell 17: 21422155.
  • Hetherington AM, Brownlee C. 2004. The generation of Ca2+ signals in plants. Annual Review of Plant Biology 55: 401427.
  • Ho LC, White PJ. 2005. A cellular hypothesis for the induction of blossom-end rot in tomato fruit. Annals of Botany 95: 571581.
  • Hofmann A. 2004. Annexins in the plant kingdom: perspectives and potentials. Annexins 1: 5161.
  • Hofmann A, Delmer DP, Wlodawer A. 2003. The crystal structure of annexin Gh1 from Gossypium hirsutum reveals an unusual S-3 cluster – implications for cellulose synthase complex formation and oxidative stress response. European Journal of Biochemistry 270: 25572564.
  • Hofmann A, Proust J, Dorowski A, Schantz R, Huber R. 2000. Annexin 24 from Capsicum annuum: X-ray structure and biochemical characterization. Journal of Biological Chemistry 275: 80728082.
  • Hofmann A, Ruvinov S, Hess S, Schantz R, Delmer DP, Wlodawer A. 2002. Plant annexins form calcium-independent oligomers in solution. Protein Science 11: 20332040.
  • Honys D, Twell D. 2004. Transcriptome analysis of haploid male gametophyte development in Arabidopsis. Genome Biology 5: R85.
  • Hoshino D, Hayashi A, Temmei Y, Kanzawa N, Tsuchiya T. 2004. Biochemical and immunohistochemical characterization of Mimosa annexin. Planta 219: 867875.
  • Hradilova J, Rehulka P, Rehulkova H, Vrbova M, Griga M, Brzobohaty B. 2010. Comparative analysis of proteomic changes in contrasting flax cultivars upon cadmium exposure. Electrophoresis 31: 421431.
  • Hu NJ, Yusof AM, Winter A, Osman A, Reeve AK, Hofmann A. 2008. The crystal structure of calcium-bound annexin Gh1 from Gossypium hirsutum and its implications for membrane binding mechanisms of plant annexins. Journal of Biological Chemistry 283: 1831418322.
  • Huh SM, Noh EK, Kim HG, Jeon BW, Bae K, Hu HC, Kwak JM, Park OK. 2010. Arabidopsis annexins AnnAt1 and AnnAt4 interact with each other and regulate drought and salt stress responses. Plant and Cell Physiology 51: 14991514.
  • Jami SK, Clark GB, Ayele BT, Roux SJ, Kirti PB. 2011. Identification and characterization of annexin gene family in rice. Plant Cell Reports 31: 813825.
  • Jami SK, Clark GB, Turlapati SA, Handley CA, Roux SJ, Kirti PB. 2008. Ectopic expression of an annexin from Brassica juncea confers tolerance to abiotic and biotic stress treatments in transgenic tobacco. Plant Physiology and Biochemistry 46: 10191030.
  • Jami SK, Dalal A, Divya K, Kirti PB. 2009. Molecular cloning and characterization of five annexin genes from Indian mustard (Brassica juncea L. Czern and Coss). Plant Physiology and Biochemistry 47: 977990.
  • Jiménez JL, Smith GR, Contreras-Moreira B, Sgouros JG, Meunier FA, Bates PA, Schiavo G. 2003. Functional recycling of C2 domains throughout evolution: a comparative study of synaptotagmin, protein kinase C and phospholipase C by sequence, structural and modelling approaches. Journal of Molecular Biology 333: 621639.
  • Kaetzel MA, Chan HC, Dubinsky WP, Dedman JR, Nelson DJ. 1994. A role for annexin IV in epithelial-cell function – inhibition of calcium-activated chloride conductance. Journal of Biological Chemistry 269: 52975302.
  • Kim YS, Ko J, Kim IS, Jang SW, Sung HJ, Lee HJ, Lee SY, Kim Y, Na DS. 2003. PKC delta-dependent cleavage and nuclear translocation of annexin A1 by phorbol 12-myristate 13-acetate. European Journal of Biochemistry 270: 40894094.
  • Kirilenko A, Golczak M, Pikula S, Buchet R, Bandorowicz-Pikula J. 2002. GTP-induced membrane binding and ion channel activity of annexin VI: is annexin VI a GTP biosensor? Biophysics Journal 82: 27372745.
  • Klink VP, Overall CC, Alkharouf NW, MacDonald MH, Matthews BF. 2007. Laser capture microdissection (LCM) and comparative microarray expression analysis of syncytial cells isolated from incompatible and compatible soybean (Glycine max) roots infected by the soybean cyst nematode (Heterodera glycines). Planta 226: 13891409.
  • Konopka-Postupolska D. 2007. Annexins: putative linkers in dynamic membrane-cytoskeleton interactions in plant cells. Protoplasma 230: 203215.
  • Konopka-Postupolska D, Clark G, Goch G, Dębski J, Floras K, Cantero A, Fijolek B, Roux S, Hennig J. 2009. The role of annexin 1 in drought stress in Arabidopsis. Plant Physiology 150: 13941410.
  • Konopka-Postupolska D, Clark G, Hofmann A. 2011. Structure, function and membrane interactions of plant annexins: an update. Plant Science 181: 230241.
  • Kopka J, Pical C, Hetherington AM, Muller-Rober B. 1998. Ca2+/phopholipid-binding (C2) domain in multiple plant proteins: novel components of the calcium-sensing apparatus. Plant Molecular Biology 36: 627637.
  • Kourie JI, Wood HB. 2000. Biophysical and molecular properties of annexin-formed channels. Progress in Biophysics & Molecular Biology 73: 91134.
  • Kovacs I, Ayaydin F, Oberschall A, Ipacs I, Bottka S, Dudits D, Toth EC. 1998. Immunolocalization of a novel annexin-like protein encode by a stress and abscisic acid responsive gene in alfalfa. Plant Journal 15: 185197.
  • Krugman T, Peleg Z, Quansah L, Chague V, Korol AB, Nevo E, Saranga Y, Fait A, Chalhoub B, Fahima T. 2011. Alteration in expression of hormone-related genes in wild emmer wheat roots associated with drought adaptation mechanisms. Functional & Integrative Genomics 11: 565583.
  • Kubista H, Hawkins TE, Patel DR, Haigler HT, Moss SE. 1999. Annexin V mediates a peroxide-induced calcium influx in B cells. Current Biology 9: 14031406.
  • Kurek I, Kawagoe Y, Jacob-Wilk D, Doblin M, Delmer D. 2002. Dimerization of cotton fiber cellulose synthase catalytic subunits occurs via oxidation of the zinc-binding domains. Proceedings of the National Academy of Sciences, USA 99: 1110911114.
  • Kwak H, Park MW, Jeong S. 2011. Annexin A2 binds RNA and reduces the frameshifting efficiency of infectious bronchitis virus. PLoS ONE 6: e24067.
  • Kwon HK, Yokoyama R, Nishitani K. 2005. A proteomic approach to apoplastic proteins involved in cell wall regeneration in protoplasts of Arabidopsis suspension-cultured cells. Plant and Cell Physiology 46: 843857.
  • Ladokhin AS, Haigler HT. 2005. Reversible transition between the surface trimer and membrane-inserted monomer of annexin 12. Biochemistry 44: 34023409.
  • Laohavisit A, Brown AT, Cicuta P, Davies JM. 2010. Annexins: components of the calcium and reactive oxygen signaling network. Plant Physiology 152: 18241829.
  • Laohavisit A, Davies JM. 2009. Multifunctional annexins. Plant Science 177: 532539.
  • Laohavisit A, Davies JM. 2011. Annexins. New Phytologist 189: 4053.
  • Laohavisit A, Mortimer JC, Demidchik V, Coxon KM, Stancombe MA, Macpherson N, Brownlee C, Hofmann A, Webb AAR, Miedema H, et al. 2009. Zea mays annexins modulate cytosolic free Ca2+ and generate a Ca2+-permeable conductance. Plant Cell 21: 479493.
  • Laohavisit A, Shang Z, Rubio L, Cuin TA, Véry AA, Wang A, Mortimer JC, Macpherson N, Coxon KM, Battey NH, et al. 2012. Arabidopsis annexin1 mediates the radical-activated plasma membrane Ca2+- and K+-permeable conductance in root cells. Plant Cell 24: 15221533.
  • Lauvrak SU, Hollas H, Doskeland AP, Aukrust I, Flatmark T, Vedeler A. 2005. Ubiquitinated annexin A2 is enriched in the cytoskeleton fraction. FEBS Letters 579: 203206.
  • Lee I, Ambaru B, Thakkar P, Marcotte EM, Rhee SY. 2010. Rational association of genes with traits using a genome-scale gene network for Arabidopsis thaliana. Nature Biotechnology 28: 149156.
  • Lee S, Lee EJ, Yang EJ, Lee JE, Park AR, Song WH, Park OK. 2004. Proteomic identification of annexins, calcium-dependent membrane binding proteins that mediate osmotic stress and abscisic acid signal transduction in Arabidopsis. Plant Cell 16: 13781391.
  • Lee I, Seo YS, Coltrane D, Hwang S, Oh T, Marcotte EM, Ronald PC. 2011. Genetic dissection of the biotic stress response using a genome-scale gene network for rice. Proceedings of the National Academy of Sciences, USA 108: 1854818553.
  • Lee TM, Lin YH. 1995. Changes in soluble and cell wall-bound peroxidase activities with growth in anoxia-treated rice (Oryza sativa L.) coleoptiles and roots. Plant Science 106: 17.
  • Lim E-K, Roberts MR, Bowles DJ. 1998. Biochemical characterization of tomato annexin p35, independence of calcium binding and phosphatase activities. Journal of Biological Chemistry 273: 3492034925.
  • Lindermayr C, Saalbach G, Durner J. 2005. Proteomic identification of S-nitrosylated proteins in Arabidopsis. Plant Physiology 137: 921930.
  • Link G. 2003. Redox regulation of chloroplast transcription. Antioxidants and Redox Signaling 5: 7987.
  • Liu J, Vishwanatha JK. 2007. Regulation of nucleo-cytoplasmic shuttling of human annexin A2-a proposed mechanism. Molecular and Cellular Biochemistry 303: 211220.
  • Loschelder H, Homann A, Ogrzewalla K, Link G. 2004. Proteomics-based sequence analysis of plant gene expression – the chloroplast transcription apparatus. Phytochemistry 65: 17851793.
  • Lu Y, Ouyang B, Zhang J, Wang T, Lu C, Han Q, Zhao S, Ye Z, Li H. 2012. Genomic organization, phylogenetic comparison and expression profiles of annexin gene family in tomato (Solanum lycopersicum). Gene 499: 1424.
  • Ludewig U, Dynowski M. 2009. Plant aquaporin selectivity: where transport assays, computer simulations and physiology meet. Cellular and Molecular Life Sciences 66: 31613175.
  • Luecke H, Chang BT, Mailliard WS, Schlaepfer DD, Haigler HT. 1995. Crystal-structure of the annexin-XII hexamer and implications for bilayer insertion. Nature 378: 512515.
  • Ma H, Kien F, Manière M, Zhang Y, Lagarde N, San Tse K, Man Poon LL, Nal B. 2012. Human annexin A6 interacts with influenza A virus M2 protein and negatively modulates infection. Journal of Virology 86: 17891801.
  • Madureira PA, Hill R, Miller VA, Giacomantonio C, Lee PWK, Waisman DM. 2011. Annexin A2 is a novel cellular redox regulatory protein involved in tumorigenesis. Oncotarget 2: 10751093.
  • Manthey K, Krajinski F, Hohnjec N, Firnhaber C, Puhler A, Perlick AM, Kuster H. 2004. Transcriptome profiling in root nodules and arbuscular mycorrhiza identifies a collection of novel genes induced during Medicago truncatula root endosymbioses. Molecular Plant-Microbe Interactions 17: 10631077.
  • Maor R, Jones A, Nuhse TS, Studholme DJ, Peck SC, Shirasu K. 2007. Multidimensional protein identification technology (MudPIT) analysis of ubiquitinated proteins in plants. Molecular & Cellular Proteomics 6: 601610.
  • Marathe R, Guan Z, Anandalakshmi R, Zhao HY, Dinesh-Kumar SP. 2004. Study of Arabidopsis thaliana resistome in response to cucumber mosaic virus infection using whole genome microarray. Plant Molecular Biology 55: 501520.
  • Mazars C, Briere C, Bourque S, Thuleau P. 2011. Nuclear calcium signaling: an emerging topic in plants. Biochimie 93: 20682074.
  • Mazars C, Thuleau P, Lamotte O, Bourque S. 2010. Cross-talk between ROS and calcium in regulation of nuclear activities. Molecular Plant 3: 706718.
  • McClung AD, Carroll AD, Battey NH. 1993. Identification and characterization of ATPase activity associated with maize (Zea mays) annexins. Biochemistry Journal 303: 709712.
  • Meagher RB, Kandasamy MK, McKinney EC. 2008. Multicellular development and protein-protein interactions. Plant Signaling & Behavior 3: 333336.
  • Meagher RB, McKinney EC, Kandasamy MK. 1999. Isovariant dynamics expand and buffer the responses of complex systems: the diverse plant actin gene family. Plant Cell 11: 9951005.
  • Meijer HJG, van de Vondervoort PJI, Yin QY, de Koster CG, Klis FM, Govers F, de Groot PWJ. 2006. Identification of cell wall-associated proteins from Phytophthora ramorum. Molecular Plant-Microbe Interactions 19: 13481358.
  • Mo Y, Campos B, Mealy TR, Commodore L, Head JF, Dedman JR, Seaton BA. 2003. Interfacial basic cluster in annexin V couples phospholipid binding and trimer formation on membrane surfaces. Journal of Biological Chemistry 278: 24372443.
  • Mohiti J, Caswell AM, Walker JH. 1997. The nuclear location of annexin V in the human osteosarcoma cell line MG-63 depends on serum factors and tyrosine kinase signaling pathways. Experimental Cell Research 234: 98104.
  • Monastyrskaya K, Babiychuk EB, Draeger A. 2009. The annexins: spatial and temporal coordination of signaling events during cellular stress. Cellular and Molecular Life Sciences 66: 26232642.
  • Montpetit B, Weis K. 2012. An alternative route for nuclear mRNP export by membrane budding. Science 336: 809810.
  • Morgan RO, Fernandez MP. 1997. Distinct annexin subfamilies in plants and protists diverged prior to animal annexins and from a common ancestor. Journal of Molecular Evolution 44: 178188.
  • Morgan RO, Jenkins NA, Gilbert DJ, Copeland NG, Balsara BR, Testa JR, Fernandez MP. 1999. Novel human and mouse annexin A10 are linked to the genome duplications during early chordate evolution. Genomics 60: 4049.
  • Morgan RO, Martin-Almedina S, Garcia M, Jhoncon-Kooyip J, Fernandez MP. 2006. Deciphering function and mechanism of calcium-binding proteins from their evolutionary imprints. Biochimica Biophysics Acta 1763: 12381249.
  • Morgan RO, Martin-Almedina S, Iglesias JM, Gonzalez-Florez MI, Fernandez MP. 2004. Evolutionary perspective on annexin calcium-binding domains. Biochimica Biophysics Acta 1742: 133140.
  • Mori IC, Schroeder JI. 2004. Reactive oxygen species activation of plant Ca2+ channels. A signaling mechanism in polar growth, hormone transduction, stress signaling, and hypothetically mechanotransduction. Plant Physiology 135: 702708.
  • Mortimer JC, Laohavisit A, Macpherson N, Webb A, Brownlee C, Battey NH, Davies JM. 2008. Annexins: multifunctional components of growth and adaptation. Journal of Experimental Botany 59: 533544.
  • Moss SE, Morgan RO. 2004. The annexins. Genome Biology 5: 219.
  • Murphy JP, Kong F, Pinto DM, Wang-Pruski G. 2010. Relative quantitative proteomic analysis reveals wound response proteins correlated with after-cooking darkening. Proteomics 10: 42584269.
  • Nagata T, Iizumi S, Satoh K, Ooka H, Kawai J, Carninci P, Hayashizaki Y, Otomo Y, Murakami K, Matsubara K, et al. 2004. Comparative analysis of plant and animal calcium signal transduction element using plant full-length cDNA data. Molecular Biology Evolution 21: 18551870.
  • Ning J, Peng XB, Qu LH, Xin HP, Yan TT, Sun MX. 2006. Differential gene expression in egg cells and zygotes suggests that the transcriptome is restructed before the first zygotic division in tobacco. FEBS Letters 580: 17471752.
  • Okamoto T, Higuchi K, Shinkawa T, Isobe T, Lorz H, Koshiba T, Kranz E. 2004. Identification of major proteins in maize egg cells. Plant Cell Physiology 45: 14061412.
  • Patel N, Hamamouch N, Li C, Hewezi T, Hussey RS, Baum TJ, Mitchum MG, Davis EL. 2010. A nematode effector protein similar to annexins in host plants. Journal of Experimental Botany 61: 235248.
  • Pauly N, Knight MR, Thuleau P, van der Luit AH, Moreau M, Trewavas AJ, Ranjeva R, Mazars C. 2000. Cell signalling – control of free calcium in plant cell nuclei. Nature 405: 754755.
  • Peng S, Publicover NG, Airey JA, Hall JE, Haigler HT, Jiang D, Chen SRW, Sutko JL. 2004. Diffusion of single cardiac ryanodine receptors in lipid bilayers is decreased by annexin 12. Biophysics Journal 86: 145151.
  • Pettersen EF, Goddard TD, Huang CC, Couch GS, Greenblatt DM, Meng EC, Ferrin TE. 2004. UCSF chimera – a visualization system for exploratory research and analysis. Journal of Computational Chemistry 25: 16051612.
  • Proust J, Houlne G, Schantz M-L, Schantz R. 1996. Characterization and gene expression of an annexin during fruit development in Capsicum annum. FEBS Letters 383: 208212.
  • Proust J, Houlne G, Schantz M-L, Shen W-H, Schantz R. 1999. Regulation of biosynthesis and cellular localization of Sp32 annexins in tobacco BY2 cells. Plant Molecular Biology 39: 361372.
  • Puthoff DP, Smigocki AC. 2007. Insect feeding-induced differential expression of Beta vulgaris root genes and their regulation by defense-associated signals. Plant Cell Reports 26: 7184.
  • Ralph SG, Chun HJE, Cooper D, Kirkpatrick R, Kolosova N, Gunter L, Tuskan GA, Douglas CJ, Holt RA, Jones SJM, et al. 2008. Analysis of 4,664 high-quality sequence-finished poplar full-length cDNA clones and their utility for the discovery of genes responding to insect feeding. BMC Genomics 9: 57.
  • Ramonell KM, Zhang B, Ewing RM, Chen Y, Xu D, Stacey G, Somerville S. 2002. Microarray analysis of chitin elicitation in Arabidopsis thaliana. Molecular Plant Pathology 3: 301311.
  • Reddy VS, Reddy ASN. 2004. Proteomics of calcium-signaling components in plants. Phytochemistry 65: 17451776.
  • Regente M, Corti-Monzon G, Maldonado AM, Pinedo M, Jorrin J, de la Canal L. 2009. Vesicular fractions of sunflower apoplastic fluids are associated with potential exosome marker proteins. FEBS Letters 583: 33633366.
  • Regente M, Monzon GC, de la Canal L. 2008. Phospholipids are present in extracellular fluids of imbibing sunflower seeds and are modulated by hormonal treatments. Journal of Experimental Botany 59: 553562.
  • Regente M, Pinedo M, Elizalde M, de la Canal L. 2012. Apoplastic exosome-like vesicles: a new way of protein secretion in plants? Plant Signaling & Behavior 7: 13.
  • Renaut J, Lutts S, Hausman L, Hoffmann JF. 2004. Responses of poplar to chilling temperatures: proteomic and physiological aspects. Plant Biology 6: 8190.
  • Repetto O, Bestel-Corre G, Dumas-Gaudot E, Berta G, Gianinazzi-Pearson V, Gianinazzi S. 2003. Targeted proteomics to identify cadmium-induced protein modifications in Glomus mosseae-inoculated pea roots. New Phytologist 157: 555567.
  • Roy A, Kucukural A, Zhang Y. 2010. I-TASSER: a unified platform for automated protein structure and function prediction. Nature Protocols 5: 725738.
  • Rubio L, Laohavisit A, Mortimer JC, Dark A, Davies JM. 2009. Salt stress signalling involves ATP release and Arabidopsis annexin 1. Comparative Biochemistry and Physiology, Part A: Molecular & Integrative Physiology 153: S193S194.
  • Santoni V, Rouquie D, Doumas P, Mansion M, Boutry M, Degand H, Dupree P, Packman L, Sherrier J, Prime T, et al. 1998. Use of a proteome strategy for tagging proteins present at the plasma membrane. Plant Journal 16: 633641.
  • Saric M, Vahrmann A, Niebur D, Kluempers V, Hehl AB, Scholze H. 2009. Dual acylation accounts for the localization of alpha 19-giardin in the ventral flagellum pair of Giardia lamblia. Eukaryotic Cell 8: 15671574.
  • Schapire AL, Valpuesta V, Botella MA. 2009. Plasma membrane repair in plants. Trends in Plant Science 14: 645652.
  • Scheuring D, Viotti C, Krueger F, Kuenzl F, Sturm S, Bubeck J, Hillmer S, Frigerio L, Robinson DG, Pimpl P, et al. 2011. Multivesicular bodies mature from the trans-Golgi network/early endosome in Arabidopsis. Plant Cell 23: 34633481.
  • Schmidt DD, Voelckel C, Hartl M, Schmidt S, Baldwin IT. 2005. Specificity in ecological interactions. Attack from the same Lepidopteran herbivore results in species-specific transcriptional responses in two Solanaceous host plants. Plant Physiology 138: 17631773.
  • Schuster-Böckler B, Schultz J, Rahmann S. 2004. HMM Logos for visualization of protein families. BMC Bioinformatics 5: 7.
  • Sémon M, Wolfe KH. 2007. Consequences of genome duplication. Current Opinion in Genetics & Development 17: 505512.
  • Shang Z, Laohavisit A, Davies JM. 2009. Extracellular ATP activates an Arabidopsis plasma membrane Ca2+-permeable conductance. Plant Signaling & Behavior 4: 13.
  • Shi JR, Dixon RA, Gonzales RA, Kjellbom P, Bhattacharyya MK. 1995. Identification of cDNA clones encoding valosin-containing protein and other plant plasma membrane-associated proteins by a general immunoscreening strategy. Proceedings of the National Academy of Sciences, USA 92: 44574461.
  • Shin H, Brown M. 1999. GTPase activity and biochemical characterization of a recombinant cotton fiber annexin. Plant Physiology 119: 925934.
  • Simões I, Mueller EC, Otto A, Bur D, Cheung AY, Faro C, Pires E. 2005. Molecular analysis of the interaction between cardosin A and phospholipase Dα. Identification of RGD/KGE sequences as binding motifs for C2 domains. FEBS Journal 272: 57865798.
  • Sopkova de Oliveira Santos J, Vincent M, Tabaries S, Chevalier A, Kerboeuf D, Russo-Marie F, Lewit-Bentley A, Gallay J. 2001. Annexin A5 D226K structure and dynamics: identification of a molecular switch for the large-scale conformational change of domain III. FEBS Letters 493: 122128.
  • Srivastava J, Barber DL, Jacobson MP. 2007. Intracellular pH sensors: Design principles and functional significance. Physiology 22: 3039.
  • Stamatakis A, Hoover P, Rougemont J. 2008. A rapid bootstrap algorithm for the RAxML Web servers. Systematic Biology 57: 758771.
  • Talukdar T, Gorecka KM, de Carvalho-Niebel F, Downie TA, Cullimore J, Pikula S. 2009. Annexins – calcium- and membrane-binding proteins in the plant kingdom Potential role in nodulation and mycorrhization in Medicago truncatula. Acta Biochimica Polonica 56: 199210.
  • Tan C, Wang H, Zhang Y, Qi B, Xu GX, Zheng HQ. 2011. A proteomic approach to analyzing responses of Arabidopsis thaliana root cells to different gravitational conditions using an agravitropic mutant, pin2 and its wild type. Proteome Science 9: 72.
  • Thiel H, Varrelmann M. 2009. Identification of beet necrotic yellow vein virus P25 pathogenicity factor-interacting sugar beet proteins that represent putative virus targets or components of plant resistance. Molecular Plant-Microbe Interactions 22: 9991010.
  • Thonat C, Mathieu C, Crevecoeur M, Penel C, Gaspar T, Boyer N. 1997. Effects of a mechanical stimulation on localization of annexin-like proteins in Bryonia dioica internodes. Plant Physiology 114: 981988.
  • Timme RE, Bachvaroff TR, Delwiche CF. 2012. Broad phylogenomic sampling and the sister lineage of land plants. PLoS ONE 7: e29696.
  • Urbany C, Colby T, Stich B, Schmidt L, Schmidt J, Gebhardt C. 2012. Analysis of natural variation of the potato tuber proteome reveals novel candidate genes for tuber bruising. Journal of Proteome Research 11: 703716.
  • Valapala M, Vishwanatha JK. 2011. Lipid raft endocytosis and exosomal transport facilitate extracellular trafficking of Annexin A2. Journal of Biological Chemistry 286: 3091130925.
  • Vandeputte O, Lowe YO, Burssens S, van Raemdonck D, Hutin D, Boniver D, Geelen D, El Jaziri M, Baucher M. 2007. The tobacco Ntann12 gene, encoding an annexin, is induced upon Rhodoccocus fascians infection and during leafy gall development. Molecular Plant Pathology 8: 185194.
  • Verma DPS, Hong ZL. 2001. Plant callose synthase complexes. Plant Molecular Biology 47: 693701.
  • Vishwanatha JK, Kumble S. 1993. Involvement of annexin II in DNA replication: evidence from cell-free extracts of Xenopus eggs. Journal Cell Science 105: 533540.
  • Walz C, Giavalisco P, Schad M, Juenger M, Klose J, Kehr J. 2004. Proteomics of curcurbit phloem exudate reveals a network of defence proteins. Phytochemistry 65: 17951804.
  • Walz C, Juenger M, Schad M, Kehr J. 2002. Evidence for the presence and activity of a complete antioxidant defence system in mature sieve tubes. Plant Journal 31: 189197.
  • Wang Q, Sun JL, Bao L, Lian JP, Zhao HX. 2011. Twenty putative palmitoyl-acyl transferase genes with distinct expression patterns in Arabidopsis thaliana. African Journal of Biotechnology 10: 1057510584.
  • Wang W, Xu JP, Kirsch T. 2005. Annexin V and terminal differentiation of growth plate chondrocytes. Experimental Cell Research 305: 156165.
  • Wang GL, Yang HY, Xia R, Fang HJ, Jing SX. 2001. Cloning and sequencing the full-length cDNA of annexin from strawberry fruit. Acta Botanica Sinica 43: 874876.
  • Watkinson JI, Sioson AA, Vasquez-Robinet C, Shukla M, Kumar D, Ellis M, Heath LS, Ramakrishnan N, Chevone B, Watson LT, et al. 2003. Photosynthetic acclimation is reflected in specific patterns of gene expression in drought-stressed loblolly pine. Plant Physiology 133: 17021716.
  • Watson WD, Srivastava M, Leighton X, Glasman M, Faraday M, Fossam LH, Pollard HB, Verma A. 2004. Annexin 7 mobilizes calcium from endoplasmic reticulum stores in brain. Biochimica Biophysics Acta 1742: 151160.
  • Webb MA. 1999. Cell-mediated crystallization of calcium oxalate in plants. Plant Cell 11: 751761.
  • White PJ, Bowen HC, Demidchik V, Nichols C, Davies JA. 2002. Genes for calcium-permeable channels in the plasma membrane of plant root cells. Biochimica Biophysica Acta-Biomembranes 1564: 299309.
  • White PJ, Broadley MR. 2003. Calcium in plants. Annals of Botany 92: 487511.
  • Wilkinson JQ, Lanahan MB, Conner TW, Klee HJ. 1995. Identification of mRNAs with enhanced expression in ripening strawberry fruit using polymerase chain reaction differential display. Plant Molecular Biology 27: 10971108.
  • Winfield MO, Lu CG, Wilson ID, Coghill JA, Edwards KJ. 2010. Plant responses to cold: transcriptome analysis of wheat. Plant Biotechnology Journal 8: 749771.
  • Wu X, Oh M-H, Schwarz EM, Larue CT, Sivaguru M, Imai BS, Yau PM, Ort DR, Huber SC. 2011. Lysine acetylation is a widespread protein modification for diverse proteins in Arabidopsis. Plant Physiology 155: 17691778.
  • Xiao FM, Tang XY, Zhou JM. 2001. Expression of 35S:Pto globally activates defense-related genes in tomato plants. Plant Physiology 126: 16371645.
  • Yan Y, Stolz S, Chételat A, Reymond P, Pagni M, Dubugnon L, Farmer EE. 2007. A downstream mediator in the growth repression limb of the jasmonate pathway. Plant Cell 19: 24702483.
  • Zhang D, Aravind L. 2010. Identification of novel families and classification of the C2 domain superfamily elucidate the origin and evolution of membrane targeting activities in eukaryotes. Gene 469: 1830.
  • Zhang H, Han B, Wang T, Chen SX, Li HY, Zhang YH, Dai SJ. 2012. Mechanisms of plant salt response: insights from proteomics. Journal of Proteome Research 11: 4967.
  • Zhang YG, Wang QH, Zhang X, Liu XL, Wang P, Hou YX. 2011. Cloning and characterization of an annexin gene from Cynanchum komarovii that enhances tolerance to drought and Fusarium oxysporum in transgenic cotton. Journal of Plant Biology 54: 303313.
  • Zhao H, Hardy RW. 2004. Long-chain saturated fatty acids induce annexin II translocation to detergent-resistant membranes. Biochemical Journal 381: 463469.
  • Zhou L, Duan J, Wang X-M, Zhang H-M, Duan M-X, Liu JY. 2011. Characterization of a novel annexin gene from cotton (Gossypium hirsutum cv CRI 35) and antioxidative role of its recombinant protein. Journal of Integrative Plant Biology 53: 347357.
  • Zhao JW, Buchwaldt L, Rimmer SR, Sharpe A, McGregor L, Bekkaoui D, Hegedus D. 2009. Patterns of differential gene expression in Brassica napus cultivars infected with Sclerotinia sclerotiorum. Molecular Plant Pathology 10: 635649.