Plant Science 177 (2009) 239–244
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Review
Extracellular nucleotides: Ancient signaling molecules Greg Clark, Stanley J. Roux * Section of Molecular Cell and Developmental Biology, The University of Texas at Austin, Austin, TX 78712, USA
A R T I C L E I N F O
A B S T R A C T
Article history: Received 9 April 2009 Received in revised form 6 May 2009 Accepted 7 May 2009 Available online 15 May 2009
The ability of extracellular nucleotides to initiate diverse signaling responses in animal cells is well established, but is only just now being recognized in plants. Despite the newness of this field in the plant literature, there is reason to believe plant cells may have been using extracellular nucleotides to transduce environmental signals even before animal cells appeared on the biological scene. Recent evidence indicates that at least some green algae have purinoceptors that are subtly similar in structure to mammalian receptors, and that some algae employ extracellular ATP (eATP) as wound signals. Extracellular nucleotides can also induce superoxide and nitric oxide production in algae, two signaling intermediates that are also commonly used by other primitive plants. Another key molecular component of extracellular nucleotide signaling is the ectoapyrase enzyme, responsible for limiting the accumulation of nucleotides that are released during cell growth, wounding and pathogen attacks, and that can help terminate eATP signaling. The green alga Ostreococcus lucimarinus has several different enzymes that structurally qualify as apyrases, and one of these has a potential signal anchor, although none have been confirmed as ectoapyrases. As yet no plasma membrane-localized receptor for extracellular nucleotides has been identified in algae or in any other plant, and so it remains unclear whether the mechanisms by which algae and other plants respond to eATP and other extracellular nucleotides is similar to or has diverged from those used by animals. ß 2009 Elsevier Ireland Ltd. All rights reserved.
Keywords: ATP release Ectoapyrase Nitric oxide Ostreococcus Purinoceptors Reactive oxygen species
Contents 1. 2. 3. 4. 5. 6. 7.
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mechanisms of release of cellular nucleotides . . . . . . . . . . . . . . . . . . . . . . . . . . . . Purinoceptors – how early did they appear? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Signaling uses of extracellular nucleotides appeared early in evolution. . . . . . . . Amplification of the eATP signal occurs through ancient transduction pathways Ectoapyrases: ancient regulators of extracellular nucleotide levels . . . . . . . . . . . Summary and Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
1. Introduction During the last three years, over a thousand papers have been published on the topic of signaling responses initiated by extracellular nucleotides, such as extracellular ATP (eATP) and eADP. Over 98% of these papers were published on responses in animal cells, where the receptors for these changes, called P2 purinoceptors, are well known and characterized [1]. During the same period advances
* Corresponding author. Tel.: +1 512 471 4238; fax: +1 512 232 3402. E-mail address:
[email protected] (S.J. Roux). 0168-9452/$ – see front matter ß 2009 Elsevier Ireland Ltd. All rights reserved. doi:10.1016/j.plantsci.2009.05.004
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in documenting and understanding the roles of extracellular nucleotides in plant growth and development have been significant, and have included studies in algae, revealing that cells have used nucleotides that accumulate outside the cell membrane as signaling agents since early in biological history. This review will highlight some of the most recent evidence that eATP, eADP, and, likely, other extracellular nucleotides can serve as major regulators of diverse physiological responses in plants, and probably have done so for hundreds of millions of years. A more comprehensive coverage of the earlier (pre-2007) plant literature on eATP signaling can be found in [2]. Also, a broader and longer review on this topic, covering more of the animal literature has just been published [3].
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2. Mechanisms of release of cellular nucleotides The vast majority of nucleotides are synthesized inside the cell and must be released into the extracellular matrix (ECM) of cells before they can act as extracellular signaling agents. Remarkably, measurable levels of dissolved ATP near or above 1 nM can be found in the sea, much of it likely released from broken phytoplankton cells during their consumption by zooplankton [4], so it seems possible that primitive cells swam in an ocean of dilute ATP. Mechanisms for ATP release from cells are diverse, and several have been documented both in animals and plants. Wounding a cell, of course, would allow cytoplasmic ATP, typically near 1 mM [5], to escape into the ECM, and ATP levels above 40 mM have been measured in fluids at wound sites of Arabidopsis leaves [6]. Presumably the [eATP] immediately adjacent to the membrane at a cut site would initially be much higher. Mechanical stimulation (e.g., from touch, osmotic stress) also releases ATP from plant cells, just as it does from animal cells [6]. Multi-drug resistance (ABCB-type) transporters cause an efflux of ATP from animal cells [7], and there is evidence that the same happens in plant cells [8]. Another mode of delivery of cytoplasmic ATP into the ECM reported in animals is via secretion [9]. The contents enclosed in secretory vesicles typically contain 1 mM ATP, and this is released when these vesicles fuse with the plasma membrane [10]. As yet there are no published estimates of the [ATP] in secretory vesicles of plant cells, but the fact that significant eATP levels accumulate at the actively growing (and secreting) tip of root hairs [11] suggests that release of ATP may be correlated with secretion in plant cells, too. The luciferase reporter first used [11] to visualize the accumulation of ATP outside the tip of growing root hairs was expressed in bacteria as a luciferase:cellulose-binding peptide hybrid molecule, and then applied externally to plant roots, where it bound tightly to the surface of cell walls. Luciferase can be engineered into transgenic plants as an ecto-luciferase, which then can continuously report [eATP] in diverse tissues. Preliminary work [12] indicates that this may be another effective way to confirm the presence of significant levels of eATP surrounding plant cells, especially those engaged in active growth and/or secretion. 3. Purinoceptors – how early did they appear? There are two main classes of receptors in animal cells that bind extracellular nucleotides with a high affinity and initiate signal transduction chains, P2X and P2Y. P2X-type receptors (seven different types in mammals) are ion-channel linked and mediate the uptake of cations, including calcium, following activation [13]. P2Y-type receptors (eight different types in mammals) are G-protein linked, and they activate G-proteins when they bind extracellular nucleotides, which also results in an increase in cytoplasmic calcium [Ca2+]cyt. Plant cells respond to sub-mM levels of eATP with a rapid increase in [Ca2+]cyt [14], and, as a result of recent breakthrough studies in protoplasts [15], it is now clear that this ATP perception happens at the cell membrane not in the cell wall. Because no protein with close sequence similarity to any animal purinoceptor is coded for by the Arabidopsis genome, one could conclude that plants use very different receptors to initiate their responses to extracellular nucleotides. However, the same antagonists that block eATP responses in animal cells (e.g., pyridoxal-phosphate-6-azophenyl-20 , 40 -disulfonate (PPADS), and reactive blue 2) also block these responses in plant cells [6]. An advanced BLAST search was used to identify a candidate purinoceptor in the green alga Ostreococcus tauri [16], even though this candidate had only 28% sequence identity to animal
homologues. This candidate bound ATP with a high affinity and mediated ion transport changes, similar to P2X-type receptors in animals [16]. This specific receptor turned out to be localized internally instead of on the plasma membrane, but this result points to the possibility that other candidate proteins that are only subtly similar to animal receptors may yet be found in other plants, even multicellular ones. The high-affinity eATP-binding protein(s) that induced ion fluxes in protoplasts [15] could be very different from the animal and algal purinoceptors, but this remains for future studies to discover. 4. Signaling uses of extracellular nucleotides appeared early in evolution The macroalga Dasycladus vermicularis releases ATP when wounded [17], and this ATP could serve as a signaling agent to induce the production of nitric oxide (NO) and hydrogen peroxide (H2O2). These two agents help mediate the wound response in Dasycladus [18] (Fig. 1). Wound-released ATP in Arabidopsis leaves is known to induce superoxide (O2), which is rapidly converted to H2O2, and downstream wound-related changes in gene expression. Thus two groups have proposed that the increase in [Ca2+]cyt induced by the binding of eATP to its plasma membrane receptor could activate Respiratory burst oxidase homolog C NADPH oxidase activity in Arabidopsis, resulting in the increased production of O2 and, subsequently, H2O2 [6,15]. Overexpression of an ectoapyrase in Arabidopsis (which would lower the eATP at wound sites) diminishes O2 production in wounded leaves, and in Dasycladus, purinoceptor antagonists (such as PPADS) diminish NO and H2O2 production after the wound stimulus (Fig. 1). Taken together, these results suggest that plant cells have used eATP as an early signal to induce wound responses at least since Chlorophyta first appeared hundreds of millions of years ago. Some fungal elicitors induce an uptake of Ca2+ in plant cells [19], and this could be a mechanism for release of ATP from cells under attack from fungal pathogens [20]. Consistent with this hypothesis, the fungal elicitor chitin induces a rapid release of eATP and ROS production in Arabidopsis [11]. Also, a yeast fungal elicitor induces the release of ATP from hairy root cultures, and this ATP serves as a signal to induce H2O2 production [21]. This reactive oxygen species (ROS), in turn, is well documented to induce diverse plant defense responses, including a direct cytotoxic effect on pathogens, increased cross-linking of wall polymers that reinforce walls, and increased production of antimicrobial compounds, among others. It is not known how ancient the reactive oxygen species strategy of plant pathogen defense is, but the fossil record indicates that plants had already developed fungal defense mechanisms in the Devonian era a billion years ago [22]. The fungal elicitor fumonisin can induce the depletion of eATP from plant cells, and this depletion can serve as a signal to induce cell death [23]. Pathogen-induced cell death is a well characterized defense strategy of plants. The fact that non-hydrolysable nucleotides that can activate purinoceptors do not reverse the effects of eATP depletion [23] suggests that the eATP target in this response is probably not a purinoceptor, but is more likely an ATPrequiring enzyme, such as a protein kinase. The discovery of the eATP-requiring factor in the extracellular matrix that protects against cell death will represent a significant advance. 5. Amplification of the eATP signal occurs through ancient transduction pathways As indicated earlier, one of the first signaling changes that can be measured in plant cells that stimulated by extracellular nucleotides is an increase in [Ca2+]cyt. This signal amplification strategy must have arisen early in evolution, because virtually all photosynthetic
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Fig. 2. Schematic model of a proposed pathway for the induction of signaling changes by eATP. The model predicts that eATP can bind to a plasma membrane (PM)-localized receptor and induce the release of Ca2+ from internal stores, thus increasing the activity of a Ca2+-regulated NADPH oxidase and producing ROS in the ECM. An ROS like H2O2 can induce the opening of plasma membrane Ca2+ channels, further increasing the [Ca2+]cyt and turn on downstream responses including changes in the rate of transcription of nuclear genes. RBOHC, Respiratory burst oxidase homolog. Adapted from Demidchik et al. [15] by permission.
Fig. 1. Quantification of NO (A) and H2O2 (B) production, using the fluorescent indicators DAF-FM DA and H2DCFDA in Dasycladus verimularis. The graphs show that wounding or applied ATPgS (A and B) or applied ADPbS (A), but not AMPS, can induce the production of NO and H2O2. The inductive effects of ATPgS, ADPbS and wounding are blocked by the purinoceptor antagonist PPADS. The concentration of all nucleotides applied was 50 mM. DAF-FM DA, 4-amino-5-methylamino-20 ,70 difluorofluorescein diacetate; H2DCFDA 20 ,70 -dichlorodihydrofluorescein diacetate. Reproduced from Torres et al. [17] by permission.
eukaryotic cells tested, including algae, have at least some documented transduction pathways that involve changes in [Ca2+]cyt and activation of Ca2+ binding proteins like calmodulin and calmodulin-domain protein kinases (CDPKs) [24–26]. Although it is now clear that applied ATP can rapidly induce an increase in [Ca2+]cyt [14,15,27] it is not clear that this is mediated by a plasma-membrane localized ion-channel linked receptor. In contrast to the cation channel activity of mammalian P2X-type receptors, which is activated in milliseconds by eATP, there is a significant delay of at least 40 s after an eATP stimulus before channel activity in the plasma membrane can be detected [15]. This delay suggests there are probably precedent signaling steps before receptor activation induces a calcium influx across the membrane. It has been suggested that these steps could include a release of calcium from internal stores, the activation of calciumdependent NAPDH oxidase activity to generate O2 and H2O2 in the ECM, and, finally, the H2O2-induced opening of cation channels in the plasma membrane (Fig. 2) [15]. Discovery of the plasma
membrane-localized purinoceptor in plants, its enzyme activity (if any), and its cellular binding partners will be needed to resolve the sequence of signaling steps that link receptor activation to an increase in [Ca2+]cyt. Superoxide appears to be an intermediate signaling step induced by eATP [6,11,15,17], and, as indicated above, this ROS is already in the arsenal of transduction mechanisms of the earliest plants [17,18]. Diverse downstream responses that have survival benefit are induced by ROS, including wound-plug hardening in algae [28], heavy-metal stress responses in moss [29], induction of wound-response genes [6], and promotion of root and root hair growth in flowering plants [30,31]. That root hair growth positively responds to eATP has been widely reported [11,32,33]. Another early signaling intermediate induced by extracellular nucleotides is nitric oxide (NO), which, like Ca2+ and ROS, is currently used as a signal transducer by algae in the Chlorophyta, a phylum that first appeared close to a billion years ago. Extracellular nucleotides induce NO in unwounded D. vermicularis and Acetabularia acetabulum [17], mimicking a wound stimulus, as well as in tomato tissue culture and tobacco BY2 cells [34], in Salvia miltiorrhiza hairy roots [35], and in growing pollen tubes of Arabidopsis [36]. The threshold for all these responses is 50 mM nucleotides or higher, significantly greater than the 500 nM level needed to induce O2 production in Arabidopsis leaves [6]. As discussed in further detail later, it is premature to judge whether levels this high can be considered physiologically relevant. Just as Ca2+ and calcium-binding proteins may activate NADPH oxidase to increase O2 production, so, too, NO production can be stimulated by Ca2+ and calcium-binding proteins [37]. Although all the sources of NO production in plants are still not resolved [38], one source appears to be derived from nitrate reductase (NR) activity, because nia1nia2 NR mutants are suppressed in NO production and deficient in some NO-mediated responses [39]. Pollen from these mutants, as well as wild-type pollen treated with NO antagonists, fail to transduce the inhibitory effects of applied ATPgS on pollen germination and pollen tube growth [36], suggesting that NO mediation may be crucial for some responses induced by extracellular nucleotides.
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Given that there are multiple purinoceptors in animals, it would not be surprising if a similar situation exists in plants. This opens up the possibility that different receptors would have different thresholds for activation and would be connected to different downstream signaling pathways. For example, some might depend on downstream O2 and H2O2 mediation, others could be dependent more on NO mediation, and still others could utilize both agents [17,35] in linking eATP to an ultimate growth or development response. 6. Ectoapyrases: ancient regulators of extracellular nucleotide levels Extracellular nucleotide signaling pathways, like most other signaling pathways, require mechanisms to inactivate the signal. Diverse enzymes serve this function in animals, including, importantly, ectoapyrases. These enzymes are E-NTPDases (nucleoside triphosphate diphosphohydrolases) that can hydrolyse the alpha and gamma phosphates from all nucleotide di- and triphosphates. There are two types of apyrases, endoapyrases that function inside the cell in association with intracellular organelles and structures, and ectoapyrases that function outside the cell. Ectoapyrases play a key role in removing extracellular nucleotides in animal cells by hydrolyzing eATP and eADP signals to eAMP, which is unable to elicit responses [40]. Plants also have ectoapyrases that function extracellularly to regulate levels of extracellular nucleotides [2,3]. There are seven members of the apyrase gene family [41] in Arabidopsis thaliana. While the localization and function of AtAPYs three to seven are as yet unknown, there is biochemical and genetic evidence that AtAPY1 and AtAPY2, which are 87% identical [42], function extracellularly, maintaining low concentrations of eATP and eADP. These ectoapyrases appear to have overlapping functions, as there is no readily discernable phenotype for either of the single knockout mutants. However, when expression of AtAPY1 is suppressed using RNAi in the background of Atapy2 TDNA knockout mutant plants, the transgenic plants created are dwarf [43]. Polyclonal antibodies that block pollen APY1 and APY2 activity reduce the rate of eATP turnover in the pollen growth medium and inhibit pollen growth, providing additional evidence that these apyrases function extracellularly [43]. Mutants knocked out in both apyrases (apy1apy2) are sterile because their pollen does not germinate [43,44], but if this block is overcome by complementing apy1apy2 plants with a wild-type APY under the control of a pollen-specific promoter, the germinated pollen will successfully fertilize, and the resultant seeds will germinate, but the seedlings that emerge are extreme dwarfs [45]. Computer modeling of functional gene networks indicate that AtAPY1 and AtAPY2 share a common network of interacting genes (see http://www.functionalnet.org/aranet/). The same analysis predicts that the genes that interact with AtAPYs three to seven also form a common network, but one that is distinctly different from the network common to AtAPY1 and AtAPY2. The results of these analyses support the idea that these two groups of Arabidopsis ectoapyrases are functionally distinct. Typically, animal ectoapyrases have a Km value for ATP in the lower mM range, which corresponds to the levels of eATP measured in animal cells ranging from 20 mM to 150 mM [10]. The two Arabidopsis ectoapyrases, AtAPY1 and AtAPY2, have a Km for ATP of 26 mM and 30 mM, respectively [42]. The Km values for plant ectoapyrases are important to consider as there is not yet a published estimate of the [eATP] in the plant extracellular matrix. The Km of these enzymes may be predictive of the threshold concentration of extracellular nucleotides needed to stimulate a response if this concentration is controlled largely by ectoapyrases. There are certainly acid and alkaline phosphatases in the ECM, and
there may be other ecto-nucleotidases besides ectoapyrases in plants, but there is as yet insufficient information to judge the impact these enzymes play in controlling the levels of eATP. If extracellular nucleotide signaling does indeed occur in evolutionarily ancient plants like algae, as recent evidence suggests, then one would predict that algae would also have conserved ectoapyrases. The genome for the unicellular green alga, Ostreococcus lucimarinus, has recently been fully sequenced [46]. A search for apyrase-related sequences revealed the presence of three probable apyrase genes, designated OlAPY1 to 3, as judged by overall similarity with vascular plant apyrases and the presence of apyrase conserved regions. Phylogenetic analysis suggests that the OlAPYs are more closely related to the yeast apyrase than to the vascular plant apyrases (Fig. 3). Of the three Ostreococcus apyrases, OlAPY1 and 2 appear to be more closely related to the clade containing AtAPY1 and 2, while OlAPY3 is the most divergent of the plant apyrases. An alignment of the deduced amino acid sequences of the algal apyrases with Arabidopsis apyrases shows that OlAPY1 and 2 are most similar to AtAPY1 and 2 and that OlAPY3 is more similar to AtAPY3–7. There is also a fern EST from Ceratopteris richardii (Acc. No. BE641722) that appears to encode an apyrase based on the presence of ACRs and homology with Arabidopsis AtAPY1 and AtAPY2. OlAPY1 and 2 are possible candidates that may function in the extracellular matrix. Analyses of the predicted algal amino acid sequences using SignalP 3.0 [47] indicate that the only algal apyrase that has a predicted signal anchor is OlAPY2. However, not all ECM proteins have obvious signal peptides, and there are potato-specific apyrases that appear to function as soluble proteins in the ECM without evidence of the predicted signal peptide being cleaved [48]. Noteworthy, silencing the expression of potato apyrases by RNA interference leads to a decrease in tuber size, increase in tuber number per plant, and changes in tuber morphology [48]. Thus, apoplastic apyrases with no strong association to cell membranes can play an important role in regulating growth and development. The discovery of putative apyrase genes in Ostreococcus is exciting, but it underscores the paucity of available apyrase gene sequences from more ancient plant lineages. Increased sampling of these apyrase sequences will undoubtedly shed important light on the evolution of this gene family. The diversification of this family suggests that in addition to traditional functions regulating the concentration of extracellular nucleotides during signaling there are also likely new undiscovered functions for this gene family. Both our (Fig. 3) and others’ phylogenetic analyses show that endoapyrases and ectoapyrases are not separated into different clades. For example, in the wild soybean, Glycine soja, Gs52 and Gs50 are suggested to be an ecto- and endoapyrase, respectively [49,50]. Previous analyses have indicated that ectoapyrases in legumes have undergone increased rates of change resulting in specialization for functions during nodulation [51]. There are multiple lines of evidence that the Gs52 ecto-apyrase plays a critically important role in the early infection of wild soybean by Bradyrhizobium japonicum, as well as in the subsequent nodule primordium development and organogenesis [50]. Having a predicted signal peptide does not necessarily mean that an apyrase will function as an ectoapyrase. Several ectoapyrases from peas and potatoes have predicted signal peptides but have been suggested to function both as endoapyrases and ectoapyrases [52,53,54]. Sequence analysis alone may not clearly resolve whether an apyrase functions as an ecto- or endoapyrase, and it is likely that the type of signal sequence and the presence of other localization sequences will influence the location of a particular apyrase. Moreover, certain plant apyrases may function both inside and outside the cell. Two yeast endoapyrases play important roles in glycosylation by regulating
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Fig. 3. Unrooted phylogenetic tree of apyrase proteins, showing evolutionary relationships of algal, yeast, Arabidopsis, legume, and potato apyrases. Apyrase amino acid sequences were aligned with MUSCLE (v. 3.6, 1) [59], with gapped regions excluded. Maximum likelihood (ML) trees were inferred with RAxML using the PROTGAMMAJTTF amino acid substitution model. Accession numbers for apyrases are: PsAPY1 = AB071369; PsAPY2 = AB071370; GS50 = AF207687; GS52 = AF207688; StRROP1 = U58597; StAPY2 = AF535135; ScGDA1 = L19560; AtAPY1 = At3g04080; AtAPY2 = At5g18280; AtAPY3 = At1g14240; AtAPY4 = At1g14230; AtAPY5 = At1g14250; AtAPY6 = At2g02970; AtAPY7 = At4g19180; OlAPY1 = NC_009358 (OSTLU_36812); OlAPY2 = NC_009365 (OSTLU_3043); OlAPY3 = NC_009375 (OSTLU_29665). Note full length OlAPY2 was constructed by adding bp 307158–307185 and bp 308301–308558 from NC_009365 to OSTLU_3043 and full-length OlAPY3 was constructed by adding bp 74401–74735 from NC_009375 to OSTLU_29665.
nucleotide levels in the Golgi cisternae [55]. Plant endoapyrases can associate with a variety of intracellular organelles and structures, including mitochondria, nuclei and cytoskeleton/ endoplasmic reticulum. There they could play a role in regulating nucleotide concentrations associated with these intracellular locales, thus impacting important cellular processes such as mitochondrial adenylate metabolism, cytoskeletal mRNA transport and nuclear splicing [53,56,57]. 7. Summary and Conclusion Although signaling roles for extracellular nucleotides is a relatively new topic in plant biology, it is now clear that some of the earliest plants used these molecules to regulate many activities. How similar plant purinergic signaling is to that in animals, remains unclear. Pre-eminent among unanswered questions is the nature of the plasma membrane receptor that responds to released ATP and other nucleotides by initiating downstream signaling changes [15]. Current data indicate that the plant receptors will have distinctly different properties than their animal counterparts, so their discovery will mark a major milestone in understanding the evolution of purinergic signaling. Biochemical approaches (labeling membranes), mutant screening, and more advanced methods for identifying ATP-binding domains in proteins are all being used to attain this breakthrough. A second key question relates to the actual eATP concentrations that plant tissues experience in different real-life situa-
tions like wounding, pathogen attack, rapid growth, abiotic stress, etc. Knowing these parameters and the Km properties of ectoapyrases will facilitate a better evaluation of the role of these enzymes in maintaining [eATP] within physiologically supportive boundaries. It will also enable a more realistic evaluation of what doses of applied nucleotides can be considered physiologically meaningful, and which are likely to generate responses unlikely to occur in vivo. Luciferase-based methods have yielded valuable insights on these questions in animal cells, and have promise to be successful also in plant systems [11,12]. Meanwhile, we should recognize that we are only beginning to understand the signaling role of nucleotide-derived molecules in the ECM of plants. Extracellular pyridine signaling in plants has now been documented and likely plays a role in plant defense responses [58], so purinoceptors will likely emerge as only one sub-group of diverse receptors that respond to the various versions of modified purine and pyrimidine bases that can be released from plant cells. Acknowledgements We thank Andy Alverson for doing the apyrase alignments and creating the phylogenetic tree and for help interpreting the evolutionary relationships of the apyrases. Research of the authors was supported by a National Science Foundation grant to S.J.R. (IOS-0718890).
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