The cellular language of myoinositol signaling - Wiley Online Library

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Tansley review The cellular language of myo-inositol signaling Author for correspondence: Glenda E. Gillaspy Tel: +1 540 231 1850 Email: [email protected]

Glenda E. Gillaspy Department of Biochemistry, Virginia Tech, Blacksburg, VA 24061, USA

Received: 20 September 2011 Accepted: 22 September 2011

Contents Summary

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I.

Introduction

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II.

Structural and terminology considerations: adding phosphates and fatty acids to myo-inositol creates a chemical and cellular language for signaling

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Hydrolysis of membrane PtdInsPs results in second messenger production

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III.

IV. What signals are linked to inositol signaling?

V. The controversial roles of DAG and Ins(1,4,5)P3 in the plant cell

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VI. A Multiplicity of enzymes that synthesize and metabolize Ins(1,4,5)P3

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VII. New connections of inositol signaling in plants

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VIII. Concluding remarks

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Acknowledgements

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References

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Summary New Phytologist (2011) 192: 823–839 doi: 10.1111/j.1469-8137.2011.03939.x

Key words: 5PTase, calcium, hormone receptor, inositol, inositol hexakisphosphate, InsP3 receptor, phosphatidylinositol, phosphatidylinositol(3)P.

2011 The Author New Phytologist 2011 New Phytologist Trust

The simple polyol, myo-inositol, is used as a building block of a cellular language that plays various roles in signal transduction. This review describes the terminology used to denote myo-inositol-containing molecules, with an emphasis on how phosphate and fatty acids are added to create second messengers used in signaling. Work in model systems has delineated the genes and enzymes required for synthesis and metabolism of many myo-inositol-containing molecules, with genetic mutants and measurement of second messengers playing key roles in developing our understanding. There is increasing evidence that molecules such as myo- inositol(1,4,5)trisphosphate and phosphatidylinositol(4,5)bisphosphate are synthesized in response to various signals plants encounter. In particular, the controversial role of myo-inositol(1,4,5)trisphosphate is addressed, accompanied by a discussion of the multiple enzymes that act to regulate this molecule. We are also beginning to understand new connections of myo-inositol signaling in plants. These recent discoveries include the novel roles of inositol phosphates in binding to plant hormone receptors and that of phosphatidylinositol(3)phosphate binding to pathogen effectors.

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I. Introduction All organisms require the ability to respond to their environment in order to adapt and survive. In response to extracellular signals, many organisms utilize receptors that respond to signals and convert them to intracellular second messengers (Berridge, 2009). Second messengers accumulate rapidly and transiently in response to external signals, a phenomena that allows signaling to be discrete and regulated. This review focuses on the role of the simple polyol, myo-inositol (Michell, 2007) (Fig. 1), and how it is used as a building block of a cellular language that plays various roles in plant signal transduction. The cellular language results because both lipids and phosphates can be added to myo-inositol and then removed (Michell, 2011), resulting in the production of new intracellular second messenger molecules that impart specific information to the cell. Although we do not currently understand the functional role of every isomer of each type of myo-inositol signaling molecule (Loewus, 1990), we do know much about how several of these molecules are synthesized and metabolized (Loewus & Murthy, 2000), and how the genes encoding such enzymes impact plant growth, development and physiology (Mueller-Roeber & Pical, 2002; Torabinejad & Gillaspy, 2006; Gillaspy, 2010). In addition, new approaches revealing genetic and biochemical interactions indicate that components of this pathway may be involved in other processes such as activation of plant hormone receptors (Tan et al., 2007; Tan & Zheng, 2009; Sheard et al., 2010) and plant pathogen signaling (Mosblech et al., 2011), which are detailed at the end of this review. Thus, this review seeks to provide a basic primer of information on myo-inositol synthesis and signaling, and to provide a forum for new ideas on this pathway in plants. At the outset, it is important to note that Fig. 2 contains a hypothetical model of the inositol synthesis and signaling pathway that may be useful for navigating this review.

II. Structural and terminology considerations: adding phosphates and fatty acids to myo-inositol creates a chemical and cellular language for signaling The terminology used in the inositol signaling pathway can be complex and possibly daunting for the uninitiated (Loewus, 1990). Besides the complexity of the terminology, there are also many different types of abbreviation styles used in the literature. As more genomic, proteomic and genetic screens identify interactions with this pathway, it is imperative to have a unifying set of abbreviations to facilitate understanding and description of the reactions within the pathway. To this end, this review uses the standard abbreviations suggested by the Journal of Biological Chemistry.

New Phytologist (2011) 192: 823–839 www.newphytologist.com

New Phytologist The first example of important terminology is the use of the latin prefix myo- to describe the most common inositol isomer in eukaryotic cells. The prefix myo- thus denotes an axial substitutent on carbon 2 of inositol while all other substituents of the ring are equatorial. It is the 2 position that provides the reference for subsequent assignment of stereoisomerism and enantiomerism. A seminal discussion of structure can be found in Parthasarathy & Eisenberg (1986). As myo-inositol constitutes the majority of inositol in the average eukaryotic cell, myo-inositol will be abbreviated as ‘inositol’ throughout this review. Of the eight other possible isomers, the D-chiro-, epi-, neo-, muco- and allo-isomers are probably minor in quantity (Michell, 2008). These other isomers of inositol have been found in specific organisms, such as the presence of D-chiro-inositol in insects (Hipps et al., 1972), humans (Larner, 2002; Carlomagno et al., 2011), pea (Pisum sativum; Lahuta & Dzik, 2011), buckwheat seeds (Fagopyrum esculentum; Horbowicz et al., 1998), soybean (Glycine max; Streeter, 1980), and Mesembryanthemum crystallinum (ice plant) (Mesembryanthemum crystallinum; Ishitani et al., 1996; Nelson et al., 1999). Further, lack of D-chiro-inositol has been linked to pathologies in humans such as polycystic ovary syndrome (Carlomagno et al., 2011; Galazis et al., 2011) and type 2 diabetes (Davis et al., 2000; Larner, 2002; Kang et al., 2006; Nissen et al., 2011), while in plants its presence is correlated with stress tolerance (Ishitani et al., 1996; Nelson et al., 1998). Thus it is theoretically possible that other isomers of inositol are used as precursors to the pool of inositol-containing signaling molecules in the cell, resulting in another layer of complexity in the cellular language of inositol signaling. However, when one considers the current data in the field, it seems likely that most of the inositol signaling in cells results from use of myo-inositol as a precursor. The second terminology issue in inositol signaling stems from the fact that inositol is used in a condensation reaction catalyzed by phosphatidylinositol synthase producing the phospholipid phosphatidylinositol (PtdIns). PtdIns contains diacylglycerol which is linked via a phosphate to the C1 carbon of the so-called headgroup of the phospholipid, the inositol ring (Fig. 1) (Nikawa & Yamashita, 1982; Robinson & Carman, 1982; Collin et al., 1999). The lipid nature of PtdIns and the membrane, Golgi or endoplasmic reticulum (ER) localization of PtdIns synthase (Robinson & Carman, 1982; Lofke et al., 2008) relegates PtdIns to cell membranes or vesicles. However, PtdIns is not very abundant overall, so it may not greatly contribute to membrane stability within cells. Some of the PtdIns within a cell can be phosphorylated sequentially by a group of specific PtdIns kinases, resulting in an even smaller pool of membrane-localized phosphoinositide phosphates (PtdInsPs) (Ischebeck & Heilmann, 2010) (Figs 1, 2). These PtdInsPs can have one, two or even three phosphates attached to the inositol ring, resulting in various isomers of


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Fig. 1 Structures of inositol, phosphatidylinositol phosphates and inositol phosphates. R1 and R2 denote fatty acyl chains. The arrow marks the C1 phosphate which is present in all phosphatidylinositol and phosphatidylinositol phosphates. Note that inositol(1,4,5)P3 also contains a C1 phosphate. Also note that the D isomers of inositol and inositol(1,4,5)trisphosphate are shown.

Fig. 2 Inositol signaling pathway. Extracellular signals are perceived and phospholipase C (PLC) hydrolyzes substrate phosphatidylinositol (4,5)P2 (PtdInsP2) into the second messenger Ins(1,4,5)P3 and diacylglycerol (DAG). DAG can be phosphorylated by DAG kinases (DGKs) resulting in phosphatidic acid (PtdOH). PtdOH can also be made by phospholipase D (PLD) action on phosphatidylcholine (PC). Inositol (1,4,5) trisphosphate [Ins(1,4,5)P3] can stimulate intracellular Ca2+ release which triggers downstream biological responses. Termination of Inositol (1,4,5) trisphosphate [Ins(1,4,5)P3] signaling can occur by hydrolysis of second messengers by inositol polyphosphate 5-phosphatase (5PTase) enzymes. A putative pathway from Ins(1,4,5)P3 to InsP6 is shown and involves the inositol phosphate kinase (IPK) 1 and 2 enzymes. InsP6 has also been reported to stimulate intracellular Ca2+ release. Metabolism of Ins(1,4,5)P3 eventually results in free inositol (Ins), which can be used in a condensation reaction catalyzed by phosphatidylinositol synthase 1 (PIS1) and PIS2, resulting in phosphatidylinositol (PtdIns). PtdIns can be acted on by phosphatidylinositol(3)P kinase (PI3K), resulting in phosphatidylinositol(3)P (PtdIns3P). Phosphatidylinositol(4)P kinase action on PtdIns results in phosphatidylinositol(4)P (PtdIns4P), which can be further phosphorylated to phosphatidylinositol(4,5)P2 (PtdIns(4,5)P2). Note that there are multiple InsP3, InsP4, InsP5 and PtdInsP2 forms not shown. Shown also is the rate-limiting step in Ins synthesis, which is catalyzed by myo-inositol phosphate synthase (MIPS), which acts on glucose-6-phosphate (G6P). CDP, calcium-dependent protein.

PtdInsP, PtdInsP2 and PtdInsP3. The position and number of phosphates are extremely important, and it is important to note that each resulting unique PtdInsP probably conveys specific information to the cell. This is another example of diversity in the cellular language of inositolcontaining molecules which is used in communicating different types of signal (York et al., 2001). It is conventional to denote the position of phosphates on PtdInsPs with parentheses, and the final number of phos-


phates, apart from the C1 phosphate, at the end of the molecular name. For example, PtdIns(3,4,5)P3 contains three phosphates at the 3, 4 and 5 positions on the inositol ring, along with the C1-position phosphate. PtdIns(3,4,5)P3 has never been found in plant cells, even though a recombinant plant PtdInsP kinase can produce this molecule in vitro (Elge et al., 2001). There is ample evidence, however, that several PtdInsPs, such as PtdIns(4,5)P2, are present in plant cells and function as second messengers (Perera et al., 2002; Preuss et al., 2006; Ischebeck et al., 2008, 2011; Konig et al., 2008; Mosblech et al., 2008; Stenzel et al., 2008; Thole et al., 2008; Carland & Nelson, 2009; Naramoto et al., 2009; Vermeer et al., 2009; Whitley et al., 2009; Zhao et al., 2010; Hirano & Sato, 2011; Vollmer et al., 2011). Phosphate addition and removal on PtdIns and PtdIns(4)P probably drives some types of membrane and vesicular trafficking in plant cells, as suggested by the impact of mutations in various PtdIns kinases and PtdInsP phosphatases on root hair development, a tip growth process driven by membrane trafficking (Ischebeck et al., 2010). PtdInsPs can also be dephosphorylated by a group of specific phosphatases (PtdInsP or 5-phosphatases), resulting in a cycle of phosphate addition and removal (Gillaspy, 2010) (Fig. 2). Thus a molecule such as PtdIns(4,5)P2, which contains phosphates at both the C4 and C5 positions, can be dephosphorylated to PtdIns(4)P by a so-called PtdInsP inositol polyphosphate 5-phosphatase (5PTase) enzyme (Astle et al., 2007; Ooms et al., 2009). PtdIns(4)P can also be acted on by a PtdIns(4)P 5-kinase, resulting in synthesis of PtdIns(4,5)P2 (Ischebeck & Heilmann, 2010).

III. Hydrolysis of membrane PtdInsPs results in second messenger production Perhaps one of the most interesting functions of PtdInsPs is their hydrolysis by phospholipases in response to discrete signals. It is this signal-stimulated hydrolysis by phospholipases at the plasma membrane that allows for transmission of information to the interior of the cell. There are different types of phospholipases, but the most important for inositol signaling is phospholipase C (PLC) (EC 3.1.4.11) (Dowd & Gilroy, 2010). PLC enzymes are encoded by multiple genes in plants


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and can utilize different substrates such as PtdIns, PtdInsP and PtdIns(4,5)P2. The importance of PLC in cellular information transfer can be illustrated with discussion of a key reaction that this enzyme can catalyze. When PLC hydrolyzes PtdIns(4,5)P2 in the plasma membrane, a diacylglycerol (DAG) product is generated that remains associated with the membrane, and a water-soluble inositol (1,4,5) trisphosphate or Ins(1,4,5)P3 molecule is generated that can interact with other proteins or cellular components to propagate the transfer of information within the cell (Majerus, 1992; Berridge, 1993) (Fig. 2). This is a key event in PtdInsP signaling, and it provides movement of information from the membrane to the cytosol. Two signals that are known to stimulate these events in plants are the addition of ABA and the addition of salt (DeWald et al., 2001; Sanchez & Chua, 2001; Xiong et al., 2001; Burnette et al., 2003). After ABA has been given to Arabidopsis seedlings, one can measure a transient increase in Ins(1,4,5)P3 (Sanchez & Chua, 2001; Burnette et al., 2003). In addition to elevating Ins(1,4,5)P3, salt appears to also stimulate an increase in PtdIns(4,5)P2 which is later corrected by the cell (DeWald et al., 2001). Not all PLC enzymes hydrolyze PtdIns(4,5)P2 preferentially. In fact, some plant PLC enzymes may exclusively hydrolyze PtdIns(4)P or PtdIns (Munnik et al., 1998), which differs from the animal PLC enzymes that mostly hydrolyze PtdIns(4,5)P2. This key difference results in the possibility of a combination of second messengers being produced by the different plant PLC enzymes. For example, if PtdIns(4)P is used as a substrate by PLC, then production of DAG and Ins(1,4)P2 results. By contrast, If PtdIns is used as a substrate by a PLC enzyme, then DAG and Ins(1)P result. The Arabidopsis genome contains nine AtPLC genes (MuellerRoeber & Pical, 2002). In Arabidopsis, the PLC genes are differentially regulated during development and in response to various environmental stimuli, including cold, salt, nutrients, dehydration, and ABA (Dowd & Gilroy, 2010). In tomato (Solanum lycopersicum), expression of two PLC genes (SlPLC4 and SlPLC6) is increased in Cladosporium fulvumresistant tomato lines (Vossen et al., 2010). Together, these studies strongly suggest that transcriptional activation of the PLC gene family is important for adapting plants to both abiotic and biotic stress. Post-translational control of PLC enzymes may also be important for signaling. Application of the bacterial flagellin epitope elicitor flg22 has been shown via proteomics experiments to result in phosphorylation of PLC (Serna-Sanz et al., 2011). This suggests that some signals can modify PLC directly, although whether this modification results in different PLC activity is not known.

IV. What signals are linked to inositol signaling? Although the field of inositol signaling in plants has, in many ways, lagged behind that in animals, many studies over the


years have lead to identification of signals that perturb either PtdInsPs or InsPs in plants. Gravitropism is probably the best understood signal that utilizes inositol signaling in terms of the impact on InsP changes in plants, with sustained increases in Ins(1,4,5)P3 preceding growth in graviresponsive maize (Zea mays) and oat pulvini (Avena sativa; Perera et al., 1999, 2001, 2005). Other signals for which data exist include light (Morse et al., 1989, Shacklock et al., 1992), ABA (Lee et al., 1996; Sanchez & Chua, 2001; Burnette et al., 2003), salt stress (DeWald et al., 2001; Takahashi et al., 2001), gibberellic acid (Kashem et al., 2000), anoxia (Reggiani & Laoreti, 2000), cold (Smolenska-Sym & Kacperska, 1996), heat (Liu et al., 2006; Mishkind et al., 2009), drought (Perera et al., 2008; Khodakovskaya et al., 2010), and various plant pathogens and elicitors (Legendre et al., 1993; Mosblech et al., 2008). In some of these cases, Ins(1,4,5)P3 concentrations have been shown to increase transiently in response to the signal. This list of signals is by no means complete, and may only be limited by what investigators have been willing to test. Developmental transitions probably also require various inositol signaling molecules, as suggested by the fact that many mutants in genes that encode enzymes required for inositol signaling have developmental alterations (for a review, see Gillaspy, 2010 and also the section on 5PTases in the present review). This process has recently been referred to as ‘basal signaling’, which denotes the fact that the constant flux of on-going signaling is important for homeostasis (Boss et al., 2010). The special case for inositol signaling involvement in auxin and plant pathogen responses will be considered later within this review.

V. The controversial roles of DAG and Ins(1,4,5)P3 in the plant cell When PLC hydrolyzes PtdIns(4,5)P2 in response to signals, the result is the production of DAG and Ins(1,4,5)P3 (Berridge, 1993). These classic second messengers in animal cells have known, thoroughly examined functions. In particular, DAG activates protein kinase C near the membrane, and protein kinase C can phosphorylate many target substrate proteins in response (Newton, 2010). Even though several laboratories have identified plant PKC-like activities or genes (Park & Chae, 1990; Hayashida et al., 1993), no obvious, highly homologous protein kinase C protein or gene that would encode such a protein has ever been identified or purified from plants. Even more importantly, DAG has not been shown to activate any purified plant protein kinase. Thus, the canonical role of DAG as a protein kinase activator is controversial in plants. DAG has been demonstrated both to induce ion pumping in patchclamped guard cell protoplasts and to promote opening of intact stomata (Lee & Assmann, 1991), so it can perturb plant cell biology. Recently, it has been suggested that the plant calcium-dependent protein kinases (CDPKs) are


New Phytologist similar enough to PKC to function in an analogous role, as a maize CDPK-1 phosphorylates in vitro sequence motifs similar to those recognized by animal PKCs (Loog et al., 2000). A large family of CDPKs, some of them being activated by phospholipids, has been documented (Hrabak et al., 2003). Intriguingly, the activity of a shaker-like inward-rectifying K+ channel subunit gene called KAT1 has been recently shown to be regulated through calciumdependent phosphorylation by a Xenopus PKC (Sato et al., 2010). These results indicate that a kinase analogous to PKC might exist that may modulate KAT1 channel activity in plants. However, until a purified source of plant PKC-like enzyme is examined, the ability of DAG to activate protein kinases remains without substantial evidence to support this mode of action. One role for DAG in signaling, however, has been examined. Evidence exists that DAG produced from PLC hydrolysis of PtdIns(4,5)P2 can be phosphorylated, resulting in phosphatidic acid (PtdOH) production (Arisz et al., 2009) (Fig. 2). PtdOH is an abundant second messenger and has been linked to many diverse cellular processes, including the response to drought, cold, and pathogens (Testerink & Munnik, 2011). The DAG kinases (DGKs) from plants have been identified and seven genes encoding DGK have been found in Arabidopsis. Two of these DGKs (AtDGK2 and AtDGK7) have been examined (GomezMerino et al., 2004, 2005). Both genes encode active proteins that can catalyze phosphorylation of DAG species that are typically found in plants. Both are broadly expressed and each can be induced by an environmental stress. AtDGK2 is induced by exposure to low temperature (Gomez-Merino et al., 2004) and AtDGK7 is induced by wounding (Gomez-Merino et al., 2005), pointing to roles for these genes in specific response pathways. In addition, a rice (Oryza sativa) DGK gene, OsBIDK1 (Benzothiadiazoleinduced diacylglycerol kinase), has been studied that is transcriptionally regulated by the plant pathogen Magnaporthe grisea (Zhang et al., 2008). When overexpressed, this gene provides enhanced resistance against tobacco mosaic virus (TMV) and Phytophtora parasitica infection, indicating a role for DGKs in pathogen response pathways. PtdOH can also be produced via phospholipase D (PLD) (EC 3.1.4.4) hydrolysis of phosphatidylcholine (PtdCho), phosphatidylethanolamine (PtdEt), phosphatidylglycerol (PtdGly), and phosphatidylserine (PtdSer) to generate phosphatidic acid (PtdOH) and a free-head group (Pappan et al., 1998). This is a major pathway that produces second messenger PtdOH in plants. There are multiple PLD genes in plants (12 in Arabidopsis, 18 in poplar (Populus trichocarpa) and 11 in grape (Vitis vinifera)) and the encoded enzymes, like the PLC enzymes, have varying substrate specificities. These have been recently reviewed elsewhere (Zhang et al., 2010), so only a few facts that augment understanding of inositol signaling will be presented here.


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PLDs display different requirements for Ca2+, substrate vesicle composition and ⁄ or free fatty acids, and, interestingly, some require PtdIns(4,5)P2, a PLC substrate (Wang, 2005). Knock-out mutations in various PLD genes have confirmed the role of PLD isoforms in multiple developmental and physiological events (Zhang et al., 2010). Studies aimed at elucidating the relative contributions of DGK vs PLD to second messenger PtdOH production have yielded a picture in which the DGK dominates in certain pathways and PLD dominates in others. In addition, there are signaling conditions that utilize both routes of PtdOH production. For example, plant pathogen elicitors can induce a biphasic PtdOH response, in which the first phase involves the DGK pathway while the second phase involves PLD action (van der Luit et al., 2000; Arisz et al., 2009). Thus, DGK could provide an important cross-talk between PLC and PLD signaling. Another potential situation for coregulation of PLC and PLD signaling was revealed in studies on heat stress (Mishkind et al., 2009). In tobacco (Nicotiana tabacum) BY-2 cells, Arabidopsis seedlings and rice leaves, both PtdOH and PtdIns(4,5)P2 increase within minutes of heat stress, with PLD activity contributing to the PtdOH increase and a PtdInsP kinase contributing to the increase in PtdIns(4,5)P2 (Mishkind et al., 2009). Thus, it is important to keep in mind that a single signaling event may involve multiple activities and signaling molecules. How Ins(1,4,5)P3 functions in plant signaling is also controversial. In animal cells Ins(1,4,5)P3 binds to an intracellular tetrameric receptor (InsP3 receptor) on the ER that functions as an Ins(1,4,5)P3-gated calcium (Ca2+) channel (Berridge, 1993, 2005; Berridge et al., 1999). Binding of Ins(1,4,5)P3 to this receptor results in Ca2+ release from the ER, which can be measured in the cytoplasm with fluorescent dyes and imaging (Dupont et al., 2011). Ca2+ functions by binding to downstream targets such as calmodulin and other Ca2+-activated proteins, which in turn activates other molecules, ultimately resulting in biological responses such as stimulation of gene expression, secretion, cell enlargement, and cell division (Putney, 2009). There is ample evidence that plants use Ca2+ in signaling and Ca2+ increases in response to external stimuli have been well studied in plants (Trewavas & Knight, 1994; Trewavas, 1999). For example, guard cells undergo an initial increase in cytosolic Ca2+ concentration which occurs within minutes of ABA exposure (McAinsh et al., 2000; Allen et al., 2001; Webb et al., 2001). Further, this rapid Ca2+ increase is preceded by an increase in Ins(1,4,5)P3 (Lee et al., 1996), and is dependent on increased PLC activity in guard cells (Staxen et al., 1999) and in suspension cells (Meimoun et al., 2009). In addition, microinjection of caged Ins(1,4,5)P3 into guard cells has been shown to be sufficient for closure (Gilroy et al., 1990). Other studies in plant and algal cells also support a role for Ins(1,4,5)P3 in


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triggering physiological events (Blatt et al., 1990; Forster, 1990; Thiel et al., 1990; Tucker & Boss, 1996). Lastly, a Ca2+-sensing receptor (CAS) has been identified that may regulate the concentration of Ins(1,4,5)P3 via indirect means (Han et al., 2003). As CAS has been localized to the chloroplast, it has been suggested that CAS facilitates calcium release from this organelle (Nomura et al., 2008). Given the prominence of Ca2+ in plant signaling, it may seem confusing that the function of Ins(1,4,5)P3 as a bona fide second messenger is controversial in plants. Of interest is the fact that yeast probably does not use Ins(1,4,5)P3 to stimulate intracellular Ca2+ release. In fact yeast appears to use the Ins(1,4,5)P3 resulting from PLC-driven hydrolysis of PtdIns(4,5)P2 solely for the purpose of producing Ins(1,2,3,4,5,6)P6 or inositol hexakis phosphate (InsP6) (York et al., 1999). InsP6 is found in many organisms; in plants InsP6 is the primary storage form of phosphorus in seeds (Raboy & Bowen, 2006), although all plant cells probably contain much lower amounts of InsP6. In yeast and perhaps other organisms, InsP6 serves to regulate several nuclear processes, including mRNA export from the nucleus, chromatin remodeling, and regulation of teleomere length (Monserrate & York, 2010). Given the evolutionary relationship between yeast and plants, it would not be surprising if plants used Ins(1,4,5)P3 in a similar way, which has been suggested in the past (Munnik & Vermeer, 2010). One critical and missing piece of information that could resolve this controversy is a plant Ins(1,4,5)P3 receptor that might function as a calcium channel to effect calcium increases within the cell. Past attempts to identify such a channel suggested the vacuole rather than the ER as a likely site for such a channel (Brosnan & Sanders, 1993). However, nothing in any sequenced plant genome is predicted to encode an Ins(1,4,5)P3 receptor, even though a putative Ins(1,4,5)P3 receptor has been identified in the flagellar proteome of Chlamydomonas reinhardtii (Wheeler & Brownlee, 2008). Therefore, if such a gene product exists in plants, it must be very different from the animal counterpart in nucleic acid sequence. An excellent review on the existence of a plant Ins(1,4,5)P3 receptor has been written (Krinke et al., 2007). A skeptic must at least entertain the idea that Ins(1,4,5)P3 may play no role other than as a precursor to InsP6, a molecule that has also been shown to trigger intracellular calcium release after ABA addition (Lemtiri-Chlieh et al., 2003). Thus, the interplay between Ins(1,4,5)P3 signaling and synthesis of InsP6 is a distinct possibility in plants and merits consideration. Besides being generated through PLC action, InsPs can also be synthesized de novo through the sequential phosphorylation of inositol (Shi et al., 2005), the penultimate product being InsP6 (Raboy, 2003). These reactions are catalyzed by a group of specific inositol kinases (IPKs) that have been well studied in plants (Xia & Yang, 2005). If Ins(1,4,5)P3 were to act as an InsP6 precursor, it or one of its


New Phytologist breakdown products would need to be phosphorylated by one of the IPKs. Conversion of Ins(1,4,5)P3 to Ins(1,4,5,6)P4 can be catalyzed by the IPK2b gene product (StevensonPaulik et al., 2002; Xia et al., 2003) although this is probably a minor route of InsP6 synthesis in seeds as ipk2b mutants have only a 35% reduction in InsP6 (Stevenson-Paulik et al., 2005). The other possible route from second messenger Ins(1,4,5)P3 to InsP6 would require phosphorylation at the C3 position on the inositol ring. There is weak activity in plants that could catalyze this reaction (Josefsen et al., 2007). In addition, there is support for a route from Ins(1,4,5)P3 to InsP6 from transgenic plants and plant cells engineered to constitutively break down InsP3 or synthesize PtdInsP2. In both of these cases, changes in Ins(1,4,5)P3 are mirrored by changes in InsP6, and thus there is a correlation between these two molecules (Perera et al., 2002, 2008; Im et al., 2007). A skeptic would also have to wonder how InsP6 triggers intracellular calcium release. Indeed, no InsP6 receptor has been found in plants, or in any other organism. Therefore, a mechanism for InsP6 to fill InsP3’s role as an intracellular Ca2+-releasing signal is not readily apparent. Obviously InsP6 can impact many critical and key cellular events; for example, it has recently been discovered that InsP6 has the ability to bind and possibly regulate plant hormone receptors. However, InsP6 as a second messenger that controls Ca2+ release appears to lack the same critical key data as does InsP3. One possibility that needs to be considered is that there is no InsP-regulated calcium channel in plants, and an entirely different and uncharacterized mechanism allows both InsP3 and InsP6 to regulate Ca2+ release, either simultaneously, or in different signaling contexts. Given the redundancy of plant signaling mechanisms exemplified by the two routes to produce PtdOH, this last model seems the most reasonable explanation for the preponderance of data that exist concerning this issue. Biochemical and genomic data also lend support to the idea that Ins(1,4,5)P3 is a bona fide second messenger on its own in plants. Transient increases in Ins(1,4,5)P3 after a given stimulus have been measured by multiple groups. What we know about the kinetics of this Ins(1,4,5)P3 accumulation indicates that a pool of Ins(1,4,5)P3 accumulates for long enough to affect signaling pathways, as it does in animal cells. In effect, signal-generated Ins(1,4,5)P3 does not appear to be immediately phosphorylated to facilitate Ins(1,4,5)P3 production. One technique used to measure Ins(1,4,5)P3 alterations involves radiolabeling plant tissues or cells with 3H-myo-inositol, extracting the PtdInsPs and InsPs and then resolving them by HPLC using a radioisotope detector. Quantification of the radioactivity present in different PtdInsP and InsP peaks allows one to measure dynamic changes after a signal has been given (Stevenson-Paulik et al., 2006). Work using this technique has measured PtdIns(4,5)P2 and InsP3 changes after salt stimulation (DeWald et al., 2001). This technique has also been used to


New Phytologist reliably and quantitatively measure steady-state changes in various inositol signaling mutants and transgenic plants (Gunesekera et al., 2007; Ercetin et al., 2008), illustrating that changes to Ins(1,4,5)P3 in plant cells can be measured. The second method used to measure Ins(1,4,5)P3 alterations is the so-called mass Ins(1,4,5)P3 assay and makes use of a partially purified bovine Ins(1,4,5)P3 receptor that is used along with 3H-Ins(1,4,5)P3 in competitive binding assays. This technique has been used more often than the radiolabeling assay, and is deemed to be reliable for various organisms. As this type of competition assay could have interference if other InsPs bind to the Ins(1,4,5)P3 receptor, it is worth describing in detail the specificity of the Ins(1,4,5)P3 receptor used. The Ins(1,4,5)P3 receptor most often used (adrenocortical) binds Ins(1,4,5)P3 c. 2400 times better than Ins(1,3,4,5)P4 and 500 times better than the next best interactor, Ins(1,3,4)P3 (Challiss et al., 1990). Thus, if endogenous plant InsPs of a unique nature were interfering with this assay they would have to be present at very high concentrations to interfere with Ins(1,4,5)P3 binding. While this possibility must be considered as a caveat, it does not warrant throwing out one of the most useful assays in the field of inositol signaling. One way to validate the Ins(1,4,5)P3 mass assay is to pretreat extracts with a mammalian 5PTase enzyme, which will hydrolyze Ins(1,4,5)P3 (Laxminarayan et al., 1994). Using this pretreatment it was found that 90% of the signal measured as Ins(1,4,5)P3 was removed from plant samples (Perera et al., 2001). Further, a separate study used both the radiolabeling and mass Ins(1,4,5)P3 assays on the same plant mutant samples and reported an increase in Ins(1,4,5)P3 in the mutant with both assays, although the radiolabeling indicated an overall larger fold increase in Ins(1,4,5)P3 (Ercetin et al., 2008). This result is similar to a comparison of both assays used in carbachol-treated animal cells in which Ins(1,4,5)P3 was noted to increase in both types of assay, although the radiolabeling showed a greater overall Ins(1,4,5)P3 increase (Baird et al., 1989; Lambert et al., 1992). It has been speculated that differences in mass levels of Ins(1,4,5)P3 and those from radiolabeling experiments may reflect the presence of a signaling pool of Ins(1,4,5)P3 that turns over faster and hence gets radiolabelled to a greater degree.

VI. A Multiplicity of enzymes that synthesize and metabolize Ins(1,4,5)P3 Plant genomes also provide evidence that Ins(1,4,5)P3 is a second messenger in plants. Specifically, plants appear to have expanded both the family of genes needed to produce Ins(1,4,5)P3 from PtdIns(4,5)P2 (the PLCs) and one class of genes needed to metabolize Ins(1,4,5)P3 from PtdIns(4,5)P2 (the 5PTases; EC 3.1.3.56) (Astle et al., 2007; Ooms et al., 2009). In the case of PLC genes,


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Arabidopsis contains nine PLC genes and mammals have 11, while yeast has only one gene capable of encoding PLC (Mueller-Roeber & Pical, 2002). In the case of the 5PTases, Arabidopsis has 15 genes and mammals have upward of 35 genes, while the yeast genome contains only four (Gillaspy, 2010). In addition, biochemical and genetic studies in yeast suggest that all four of the yeast 5PTases probably function to hydrolyze PtdInsPs (York et al., 2001), while some of the Arabidopsis 5PTases have been shown to preferentially impact Ins(1,4,5)P3 concentration when gene function is knocked out (Carland & Nelson, 2004; Ercetin & Gillaspy, 2004; Gunesekera et al., 2007; Ananieva et al., 2008). So, while this genome information is far from definitive, it does show expansion of these gene families in plants that may allow for specialization of Ins(1,4,5)P3 production and metabolism in plants. Once Ins(1,4,5)P3 is generated, it becomes quickly metabolized. The first step in Ins(1,4,5)P3 second messenger breakdown utilizes enzymes that dephosphorylate these molecules, often removing the 5-phosphate from the inositol ring (Erneux et al., 1998). All plant genomes queried contain two types of predicted 5PTases. Both types of plant 5PTases contain the highly conserved catalytic domain called the Exo_endo_phos domain, which confers the ability to hydrolyze the 5-phosphate from several different inositolcontaining substrates, including Ins(1,4,5)P3 and Ins (1,3,4,5)P4, and PtdIns(5)P, PtdIns(4,5)P2, PtdIns(3,5)P2 and PtdIns(3,4,5)P3 substrates (Erneux et al., 1998; York et al., 2001). The plant group A 5PTases consist entirely of this catalytic domain and have no other identified domains. The plant group B 5PTases, however, contain an additional large N-terminus containing five to six WD40 repeats (Zhong & Ye, 2004). WD40 repeats are found in a number of eukaryotic regulatory proteins (Li & Roberts, 2001; Higa & Zhang, 2007), where they form a stable b-propeller-like platform to which other proteins can bind either stably or reversibly. Their presence in plant 5PTases probably reflects the ability of group B 5PTases to participate in signaling protein complexes. Indeed, the WD40 domain of the At5PTase13 protein binds to the sucrose nonfermenting-like kinase 1.1 (SnRK1.1) and appears to protect SnRK1.1 from proteasomal degradation when sugars are in limited supply (Ananieva et al., 2008). As SnRK1.1 is an important energy ⁄ stress sensor that has been linked to lifespan determination in plants (Baena-Gonzalez et al., 2007; Coello et al., 2011), the regulation of SnRK1.1 by At5PTase13 may indicate a role for inositol signaling in these processes. The mechanism of At5PTase13 protection of SnRK1.1 probably precludes SnRK1.1 binding by an adaptor protein of the Cullin 4 ubiquitin-ligating (E3) enzyme complex called pleiotropic regulatory locus 1 (PRL1). PRL1 also contains a WD40 domain and mutation of PRL1 results in stabilization of SnRK1.1 (Lee et al., 2008) and ABA and stress phenotypes


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opposite in nature to At5PTase13 (Nemeth et al., 1998; Bhalerao et al., 1999), supporting an antagonistic role for these proteins in SnRK1.1 regulation (Ananieva & Gillaspy, 2009). It is not known if the other group B At5PTases (At5PTase12, At5PTase14 and Fragile fiber 3; FRA3) have protein-binding partners; however, the WD40 domains are different enough that it is possible that binding and protection of different proteasomal target proteins are a common function of these 5PTases (Ananieva et al., 2008). It is interesting to note that the only other type of nonplant organism that contains 5PTases with WD40 domains is a subgroup of filamentous fungi. This suggests that the presence of WD40 coding domains in 5PTase genes occurred before the diversification of plants and fungi c. 100 Mya. The substrate preferences of many of the 15 Arabidopsis 5PTases have been characterized (Berdy et al., 2001; Burnette et al., 2003; Ercetin & Gillaspy, 2004; Zhong & Ye, 2004; Zhong et al., 2004; Ercetin et al., 2008; Carland & Nelson, 2009; Kaye et al., 2011). When expressed as recombinant proteins and assayed in vitro, most of the Arabidopsis 5PTases will hydrolyze both InsP and PtdInsP substrates containing a 5-phosphate. The exceptions are At5PTase11 (Ercetin & Gillaspy, 2004) and At5PTase6 (Cotyledon vascular patterning; CVP2) (Carland & Nelson, 2009), which can only hydrolyze PtdInsP substrates, and At5PTase12 and At5PTase13, which can only hydrolyze Ins(1,4,5)P3 (Zhong & Ye, 2004). This restriction of substrates is also seen in certain animal 5PTases, so the ability to discriminate between water-soluble (i.e. InsP) and lipid-derived (i.e. PtdInsP) substrates appears to be conserved between animals and plants, although the basis for the discrimination is not currently known (Astle et al., 2007; Ooms et al., 2009). By examining plants containing either a gain or loss of function in specific 5PTases, it has been established that 5PTases are critical for plant development and signaling. The case of CVP2 ⁄ At5PTase6 exemplifies the critical connections among PtdInsPs, cell biology and development. Carland et al. isolated the cvp2 mutant as a result of its altered cotyledon vascular pattern (Carland et al., 1999). The cvp2 mutants contain elevated Ins(1,4,5)P3, are ABA hypersensitive, and contain a mutation in the At5PTase6 gene (At1g05470). At5PTase6 was subsequently shown to encode a lipid-specific 5PTase (Carland & Nelson, 2009). This indicates that the increase in Ins(1,4,5)P3 in cvp2 mutants is probably a result of decreased hydrolysis of PtdIns(4,5)P2 which is acted on and hydrolyzed by PLC, resulting in an Ins(1,4,5)P3 increase. The cvp2 gene interacts genetically with another vascular patterning gene product, scarface/vascular network defective3 (scf ⁄ van3), which contains a plextrin-homology domain that binds to PtdIns(4)P, and is required for intracellular trafficking of the Pinformed 1 (PIN1) protein (Feraru et al., 2011) that mediates auxin


New Phytologist efflux (Sieburth et al., 2006). Recently it was shown that a 5ptase6 ⁄ 5ptase7 double mutant (cvp2 ⁄ cvp2 like1 (cvl1)) phenocopies the scf ⁄ van3 mutant, and thus At5PTase6 and At5PTase7 are probably redundant in their function (Carland & Nelson, 2009). Together these data suggest that At5PTase6 and At5PTase7 hydrolyze PtdIns(4,5)P2 in developing leaves, and supply PtdIns(4)P to SCF ⁄ VAN3, which regulates vesicular trafficking of proteins important for vascular patterning. PIN1 is probably regulated by the At5PTase6, 7 ⁄ SCF ⁄ VAN3 network, thus establishing an important connection between inositol signaling and auxin transport. Other 5PTases that can impact development include FRA3, a WD40-containing 5PTase that when mutated results in altered fiber cells within the stem (Zhong et al., 2004). The fra3 mutant contains elevated Ins(1,4,5)P3 and PtdIns(4,5)P2, which is not surprising as FRA3 recombinant enzyme hydrolyzes both Ins(1,4,5)P3 and PtdIns(4,5)P2 (Zhong & Ye, 2004). Therefore, proper regulation of Ins(1,4,5)P3 and PtdIns(4,5)P2 concentrations is required for the development of fiber cells of plants. Regulation of inositol signaling may also be important for root hair initiation, as the Morphogenesis root hair 3 (mrh3) (5PTase5; At5g65090) mutants are altered in this process (Jones et al., 2006). Alterations in the Suppressor of actin mutation (SAC9) gene were found by Williams’ group to cause a constitutive stress response as a result of elevated InsP3 and PtdInsP2 concentrations (Williams et al., 2005). This mutant was also identified as being impaired in chloroplast ATP synthase activation ⁄ deactivation (Gong et al., 2006). The SAC proteins are not 5PTases, but can in some cases hydrolyze PtdInsP2 (Guo et al., 1999). As Ins(1,4,5)P3 is probably not an SAC9 substrate, it seems reasonable that the elevated Ins(1,4,5)P3 found in sac9 mutants results from elevated PtdIns(4,5)P2 which becomes hydrolyzed by PLC, similar to the scenario for cvp2 ⁄ 5ptase6 mutants. Several different developmental ⁄ signaling defects have been reported in At5PTase13 (At1g05630) mutants, including alterations in blue light responses and phototropin 1 signaling (Chen et al., 2008), nutrient ⁄ energy signaling (Ananieva et al., 2008), gravitropism, and vesicular trafficking required for PIN1 and PIN2 movements and subsequent polar auxin transport (Wang et al., 2009). Given that both CVP2 ⁄ At5PTase6 and At5PTase13 appear to impact PIN movement and polar auxin transport in the shoot and root, respectively, it is intriguing to note that both genes are present in a region of chromosome 1 that is speculated to have been duplicated during evolution. The resulting duplicated segment on chromosome 2 contains At5PTase7 and At5PTase14; thus, these genes probably represent duplicated genes that have at least partially redundant functions with At5PTase6 and At5PTase13, respectively. Interestingly, the phenotype of reduction of PIN movement can also be induced by a mutation in the rate-limiting


New Phytologist enzyme for inositol synthesis, the myo-inositol phosphate synthase (MIPS) (Chen & Xiong, 2010; Luo et al., 2011). In the Atmips1 mutant a reduction in inositol leads to a reduction in PtdIns (Donahue et al., 2010), and lower PtdIns is speculated to negatively impact vesicular trafficking of PIN2 (Chen & Xiong, 2010). This speculation seems reasonable given the involvement of PtdInsPs in several vesicular-trafficking-related events in plants. However, it is not known if Atmips1 mutants have differences in specific PtdInsPs, important information which could link these interesting phenotypes to a mechanism for altered PIN movement (Ischebeck et al., 2010). It is important to note that mutations in MIPS genes appear to be pleiotropic as growth changes are noted in potato (Solanum tuberosum) antisense MIPS plants (Keller et al.,1998) and both Atmips1 and Atmips2 mutants have changes in basal resistance to pathogens (Murphy et al., 2008). A recent study presented excellent evidence supporting a role for Ins(1,4,5)P3 and Ca2+ regulation of PIN movement and auxin transport in plants (Zhang et al., 2011). Zhang et al. isolated a suppressor of PIN1 overexpression that turns out to be an inositol 1-phosphate phosphatase called Fiery 1 (FRY1 ⁄ SAL1) (Xiong et al., 2001). The loss of FRY1 ⁄ SAL1 was associated with elevated Ins(1,4,5)P3 and Ca2+, and changes in PIN polarity and auxin transport within the root. The authors document changes in PIN trafficking in response to pharmacological inhibitors or activators which could alter Ins(1,4,5)P3 and ⁄ or Ca2+. This study went further in trying to determine the mechanism by which Ins(1,4,5)P3-mediated Ca2+ increase could impact PIN localization by examining the involvement of the pinoid kinase (PID). PID was previously shown to be negatively regulated by Ca2+ (Benjamins et al., 2003) and to phosphorylate PIN and impact PIN polar targeting and endocytic recycling (Benjamins et al., 2001; Friml et al., 2004; Michniewicz et al., 2007). Using a combination of genetic mutants and pharmacological inhibitors, the authors gained support for a model in which Ins(1,4,5)P3 mediates a Ca2+ increase that leads to inhibition of PID activity and subsequent changes in PIN polarity. It is important to note that this mechanism may not be exclusive, and thus Ca2+ could also impact PIN polarity and auxin transport in other ways, such as alteration of trafficking (Zhang et al., 2011). Reverse genetic strategies have also been helpful in delineating a role in growth regulation for the 5PTases. Ectopic expression of either 5PTase1 or 5PTase2 has been shown to decrease ABA signaling by increasing hydrolysis of Ins(1,4,5)P3 (Sanchez & Chua, 2001; Burnette et al., 2003). Loss-of-function mutants in these same genes result in ABA hypersensitivity in germinating seeds and increased seedling hypocotyl elongation in the dark, which is accompanied by an increase in Ins(1,4,5)P3 concentrations (Gunesekera et al., 2007). Overexpression of a human 5PTase exacerbates the second messenger changes seen with


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the plant enzymes and results in unique drought tolerance phenotypes (Khodakovskaya et al., 2010; Perera et al., 2008). Together the mutants and transgenic plants containing altered inositol signaling and the others described in this section illustrate that proper regulation of steady-state Ins(1,4,5)P3 and PtdIns(4,5)P2 concentrations is required for normal growth and development in plants. The fact that various 5PTase and SAC mutants are altered in vastly different developmental processes suggests that 5PTase and SAC proteins act in different subcellular compartments or within different tissues and/or organs within plants.

VII. New connections of inositol signaling in plants 1. Novel roles for InsPs in regulating plant hormone receptors Perhaps the most exciting recent development regarding inositol signaling is the potential involvement in regulating plant hormone receptors (Tan et al., 2007; Sheard et al., 2010). The discovery that higher InsPs, including InsP5 and InsP6, bind to specific plant hormone receptors and impact ligand binding suggests a role for InsPs in coincidence detection during hormone signaling. Specifically, these higher InsPs may be required, along with the hormone, in order for the receptor to become activated. Transport inhibitor response 1 (TIR1), the auxin receptor, is an F-box protein that binds auxin and mediates protein degradation of the Aux ⁄ IAA transcription repressors (Tan & Zheng, 2009). The crystal structure of the Arabidopsis TIR1 complex revealed that the leucine-rich repeat domain of TIR1 contains InsP6, and recognizes auxin and the Aux ⁄ IAA polypeptide substrate through a single surface pocket (Tan et al., 2007). Recently, the crystal structure of the jasmonic acid (JA) receptor revealed a similar binding pocket comprised of the F-box protein coronatine insensitive 1 (COI1), bound to transcriptional repressors known as jasmonate ZIM-domain (JAZ) proteins. This pocket also contains a higher InsP, namely InsP5 (Sheard et al., 2010). Remarkably, the InsP6 binding site on TIR1 and the InsP5 binding site on COI1 are strikingly similar (Sheard et al., 2010). This conservation relieves concerns that InsP6 or InsP5 binding to these receptors is nonspecific and attributable solely to the highly charged nature of these molecules. This is especially important given that the proteins were synthesized in a heterologous, nonplant host. The position of InsP6 in TIR1 and InsP5 in the JA receptor complex may reflect their status as co-factors. The recent JA receptor study explored this possibility using basic biochemical approaches to remove InsP5 from the receptor, followed by addition experiments to detect which InsPs were capable of restoring ligand (i.e. jasmonic acid-isoleucine (JA-ILE)) binding. The


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results indicate that Ins(1,2,4,5,6)P5 and Ins(1,4,5,6)P4 support JA-ILE ligand binding, but InsP6 and Ins(1,4,5)P3 do not (Sheard et al., 2010). Thus, auxin and JA signaling may differ in the higher InsP used in receptor binding. The functionality of InsP5 within the JA receptor complex has also been investigated, using the yeast two-hybrid system and Arabidopsis mutants with elevated InsP5 concentrations, by Mosblech et al. (2011). These investigators tested whether COI1 binding to its target, JAZ9, is influenced by InsP5. First, COI1 variants with changes in the putative InsP5 binding site were found to have reduced interaction with JAZ9 in yeast two-hybrid tests. In addition, the interaction of wild-type COI1 and JAZ9 is increased when a mutant yeast strain (Ives et al., 2000) is used that contains increased concentrations of InsP5 and reduced concentrations of InsP6. Next, the ability of these COI1 variants altered in the putative InsP5 binding site were tested for their ability to transduce JA responses in coi1 mutants (Feys et al., 1994). It was found that the COI1 variants displayed a reduced capability to rescue JAmediated root growth inhibition or silique development in Arabidopsis coi1 mutants. Lastly, Mosblech et al. used an Arabidopsis inositol phosphate kinase 1 (IPK1) mutant, which is defective in a 2-kinase activity, giving rise to elevated concentrations of Ins(1,3,4,5,6)P5 and reduced InsP6 (Stevenson-Paulik et al., 2005). These Arabidopsis ipk1-1 mutants have increased JA responses and defensive capabilities via COI1-mediated processes, as suggested by elevated wound-induced gene expression, defense against caterpillars or root growth inhibition by JA (Mosblech et al., 2011). Together, these data indicate that InsP5 contributes to COI1 function in plants. The subcellular implications for InsP5 and InsP6 binding to receptors are clear. As both TIR1 and COI complexes are located in the nucleus (Frankowski et al., 2009), if InsP6 and InsP5 bind to these receptors in vivo then they must also be present in the nucleus. This brings up the question of how these higher InsPs might get to the nucleus. Higher inositol phosphates are thought to be synthesized in the cytoplasm (Tsui & York, 2010). Given their charged nature, it is unlikely that they can cross the nuclear membrane. InsP6 could be actively transported, as an InsP6 ATPbinding cassette (ABC)-type transporter has been characterized and shown to transport InsP6 across the vacuolar membrane (Nagy et al., 2009). This particular transporter is not known to be located in the nuclear membrane, but other potential transporters could be. In addition, InsP6 and ⁄ or InsP5 could be synthesized in the nucleus, either de novo or after transport of a precursor into the nucleus. In particular, one inositol kinase, IPK2, has been shown to be present within the nucleus (Xia et al., 2003). This kinase can catalyze the phosphorylation of Ins(1,4,5)P3 to Ins(1,4,5,6)P4 and also converts it to Ins(1,3,4,5,6)P5. AtIPK2 also phosphorylates Ins(1,3,4,5)P4 to Ins(1,3,4,5,6)P5, and thus two routes of


New Phytologist InsP5 synthesis are possible with this nuclear kinase (Stevenson-Paulik et al., 2002; Xia et al., 2003). Of note, plants that overexpress AtIPK2 exhibit auxin-related phenotypes and are more resistant to exogenous auxin treatments, which would be consistent if these plants contained less InsP6 than is required for TIR1 function (Zhang et al., 2007). In yeast, the nuclear InsP synthesis pathway is extended by IPK1 which phosphorylates Ins(1,3,4,5,6)P5 to InsP6 (Ives et al., 2000). The plant IPK1 can complement the yeast mutant (Sweetman et al., 2006), suggesting that it may also function within the nucleus. In addition, the Arabidopsis IPK1 gene sequence contains a putative nuclear localization sequence. A last possible mechanism to provide higher InsPs to hormone receptors in the nucleus is the hydrolysis of PtdInsPs within the nuclear membrane. In particular, there are reports of a nuclear PLC signaling pathway in animals that can produce nuclear Ins(1,4,5)P3 (Divecha et al., 1991). If this exists in the plant nucleus, then Ins(1,4,5)P3 could be phosphorylated to InsP5 and InsP6 by the action of IPK2 and IPK1 within the nucleus. Regardless, future examination of nuclear InsPs and PtdInsPs is likely to be informative and to help sort out the pathways by which the auxin and JA receptor are regulated by their InsP co-factors. 2. Inositol signaling during plant–pathogen interactions A second exciting development involving inositol signaling is the connection between oomycete plant pathogens and PtdInsPs. Specifically, oomycete pathogens, like their bacterial counterparts, use pathogen-encoded effector proteins transferred into the cytoplasm of host cells to suppress host defenses (for a review, see Kale & Tyler, 2011). It has been recently established that several, but not all, oomycete and fungal effector proteins bind to monophosphorlyated PtdInsPs, with PtdIns(3)P emerging as a common binding partner (Kale et al., 2010). However, the portion of the effector protein that binds to PtdInsPs is a matter of debate (Gan et al., 2010; Ellis & Dodds, 2011). Kale et al. used both lipid blots and lipid pull-down assays to show that the N-terminal RXLR and dEER motifs that enable entry into host cells bind PtdIns(3)P. These authors also showed that the outer surface of the plant cell plasma membrane contains PtdIns(3)P and could participate in lipid raft-mediated endocytosis of the effector proteins (Kale et al., 2010). Further, they showed that blocking PtdIns(3)P binding to effectors can inhibit entry of certain effector proteins (Kale et al., 2010). By contrast, Yaeno et al. (2011) used NMR to identify a group of positively charged amino acids adjacent to the RXLR domain and have shown that this domain is responsible for PtdInsP binding. These authors used a combination of mutants and lipid blots to show that alterations that alter


New Phytologist positive charge in the RXLR adjacent region disrupt PtdInsP binding. In addition, these authors found that mutants incapable of binding to PtdInsPs have altered stability in plant cells and, importantly, reduced ability to suppress host cell death responses. In addition, co-expression of a PtdInsP 5-kinase 1 (PIP5K1) construct, which could phosphorylate endogenous PtdInsPs, also reduces the ability of an effector to suppress host cell death. Together, these data suggest that PtdInsP binding is required for the intracellular virulenceenhancing activity of the effector, possibly through internal membrane association facilitated by PtdInsP binding (Yaeno et al., 2011). However, these data do not completely rule out the possibility that PtdInsP impacts on the uptake of effectors. Both of these reports together illustrate the importance of understanding PtdInsPs in the infection process. Obviously, more work is needed before a concrete, mechanistic model for effector action can be articulated. It is important to note the possibility that different effectors with the RXLR motif may function in different ways. Given this new finding regarding the importance of PtdInsPs in infection, it should not be a surprise that powdery mildew, an oomycete, up-regulates one of the genes required for PtdIns synthase, the PtdIns synthase 1 gene (AtPIS1; At1G68000), 11-fold after infection (Chandran et al., 2010). This up-regulation may ensure that a robust pool of PtdIns is available after infection that could be phosphorylated to PtdInsP, which is needed for effector function. The up-regulation of AtPIS1 appears to be specific as the AtPIS2 gene is not up-regulated during powdery mildew infection (Chandran et al., 2010). Interestingly, AtPIS1 and AtPIS2 have been shown to have different substrate preferences for cytidinediphospho-DAG species differing in fatty acid composition, with PIS2 preferring unsaturated substrates in vitro (Lofke et al., 2008). In addition, overexpression of either gene yields different PtdIns species, with AtPIS1 overexpression resulting in lipids enriched in saturated or monounsaturated fatty acids (Lofke et al., 2008). This indicates that different AtPIS enzymes may preferentially produce different PtdIns species that could function in different signaling pathways. Limiting PtdIns synthesis may also have negative consequences for pathogens, as shown by recent results in mutants defective in the MIPS enzyme, which is rate limiting for inositol synthesis (Donahue et al., 2010). In Atmips1 mutants, an alteration of inositol synthesis leads to a reduction in PtdIns (Donahue et al., 2010), which is required by the inositol phosphorylceramide synthase enzyme that produces phytoceramide (Wang et al., 2008). The result is that Atmips1 mutants have elevated ceramide which leads to a cell death phenotype similar to that of lesion-mimic mutants (Donahue et al., 2010), which can prime defense response pathways. This connection between PtdIns, ceramide and cell death suggests that increasing


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PtdIns synthesis could be beneficial for pathogens in terms of reducing basal defense mechanisms. Other studies suggest that hydrolysis of specific PtdInsPs during plant–pathogen interactions is important for resistance. In a study using Pseudomonas syringae to infect plants, it was found that PtdInsP concentrations decreased in resistant plants, while DAG, PtdOH and InsP2 and InsP3 concentrations increased. These changes either did not occur or were much less pronounced when susceptible plants were infected (Andersson et al., 2006). This suggests that PtdInsP turnover is important during pathogen recognition. There is also genetic evidence that supports the role of PtdInsPs during pathogen recognition. Plants treated with the nonprotein amino acid b-aminobutyric acid (BABA) develop an enhanced capacity to resist an extraordinarily wide range of biotic and abiotic stresses (Ton et al., 2005). Intriguingly, BABA confers protection against certain pathogens via augmentation of basal defense responses, a phenomenon known as ‘priming’. A mutagenesis screen identified an ERlocalized phosphoinositide phosphatase, now called SAC6 (At5g66020), as required for BABA-induced pathogen resistance (Ton et al., 2005). The predicted AtSAC6 protein shares 50% amino acid identity with the SAC1 protein of yeast, and complements a yeast sac1 null mutant, indicating shared function between these genes (Despres et al., 2003). Although the substrate specificity of AtSAC6 has not been examined, the yeast SAC1 protein is known to hydrolyze PtdIns(3)P, PtdIns(4)P and PtdIns(3,5)P2 to PtdIns (Guo et al., 1999). In yeast, null mutations of the SAC1 gene result in enhanced accumulation of PtdIns(4)P and PtdIns(4,5)P2, which affects protein trafficking and organization of the actin cytoskeleton (Hama et al., 1999; Foti et al., 2001; Schorr et al., 2001). Based on these data, it has been speculated that the role of AtSAC6 in BABA-induced pathogen resistance is analogous to the role of SAC1 in yeast. Specifically, it has been suggested that PtdInsP hydrolysis may be required for vesicular trafficking and ⁄ or cytoskeleton rearrangements in the plant cell during infection by pathogens. These ultrastructural changes could direct secretory vesicles to the sites of pathogen attack, and mediate the delivery of the necessary molecules for callose production, allowing the plant to develop a faster and stronger activation of stress-specific defense mechanisms (Ton et al., 2005). InsPs have also been implicated in pathogen response pathways. A rapid accumulation of Ins(1,4,5)P3 has been observed in citrus infected with the fungus Alternaria alternata (Ortega & Perez, 2001), and treatment of rice (O. sativa) with salicyclic acid (SA) and JA has been reported to rapidly induce the expression of a PLC enzyme, which correlated with induced resistance against the blast fungus Magnaporthe grisea (Song & Goodman, 2002). Recently, wounding of plant leaves was shown to increase both JA and Ins(1,4,5)P3, while PtdIns(4,5)P2, PtdIns(4)P and PtdIns were transiently depleted. This effect is dependent on JA, as


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plants deficient in JA synthesis did not accumulate Ins(1,4,5)P3 in response to wounding (Mosblech et al., 2008). This suggests a role for PtdInsP signaling in mediating plant wound responses, and a possible role in mediating plant defense responses to herbivory. Given the role of InsP5 as a co-factor in the JA receptor, the location and metabolic fate of JA-stimulated Ins(1,4,5)P3 will be of future interest.

VIII. Concluding remarks Work over recent years has greatly expanded our understanding of the specific molecules involved in inositol synthesis and signaling and the gene products that regulate these molecules in plants. Physiological approaches, combined with analytical techniques, have delineated the signals that stimulate increases in inositol second messengers. Further, genetic mutants have provided tools with which to elucidate how individual genes function in the production or metabolism of inositol second messengers. Significant questions that remain are: How do Ins(1,4,5)P3 and ⁄ or InsP6 facilitate intracellular calcium release and stimulation of downstream pathways? How does the plant cell coordinate the function of multigene families of PtdInsP kinases, 5PTases and other enzymes to regulate inositol second messengers? How does the plant cell control the synthesis and transport of InsP5 and InsP6, which are necessary for plant hormone receptor binding? And lastly, do pathogens alter inositol synthesis and ⁄ or signaling pathways during infection?

Acknowledgements The members of the Gillaspy lab are acknowledged for their help with this review. The NSF is also gratefully acknowledged for supporting this work (MCB #0641954).

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