Review: “Pyrophosphate and pyrophosphatases in plants, their involvement in stress responses and their possible relationship to secondary metabolism”

Accepted Manuscript Title: Review: “Pyrophosphate and pyrophosphatases in plants, their involvement in stress responses and their possible relationship to secondary metabolism” Authors: Francisca Morayna Guti´errez-Luna, Eric Edmundo Hern´andez-Dom´ınguez, Lili´an Gabriela Valencia-Turcotte, Rogelio Rodr´ıguez-Sotres PII: DOI: Reference:

S0168-9452(17)30608-8 https://doi.org/10.1016/j.plantsci.2017.10.016 PSL 9697

To appear in:

Plant Science

Received date: Revised date: Accepted date:

9-7-2017 19-10-2017 26-10-2017

Please cite this article as: Francisca Morayna Guti´errez-Luna, Eric Edmundo Hern´andez-Dom´ınguez, Lili´an Gabriela Valencia-Turcotte, Rogelio Rodr´ıguez-Sotres, Review: “Pyrophosphate and pyrophosphatases in plants, their involvement in stress responses and their possible relationship to secondary metabolism”, Plant Science https://doi.org/10.1016/j.plantsci.2017.10.016 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

Pyrophosphate and soluble Pyrophosphatases in plants

Gutiérrez-Luna et al., 2017

Review: “Pyrophosphate and pyrophosphatases in plants, their involvement

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in stress responses and their possible relationship to secondary metabolism”

By: Francisca Morayna Gutiérrez-Luna1,2, Eric Edmundo Hernández-Domínguez3,

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Lilián Gabriela Valencia-Turcotte1,4 y Rogelio Rodríguez-Sotres1,*

FACULTAD DE QUÍMICA, UNIVERSIDAD NACIONAL AUTÓNOMA DE

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1

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Author's details:

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Mexico City, Mexico.

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MÉXICO, Ave. Universidad 3000, Cd. Universitaria, Del. Coyoacán, P.C. 04510,

e-mail: [email protected].

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e-mail:[email protected]

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INSTITUTO DE ECOLOGÍA A.C., Carretera antigua a Coatepec 351, El Haya,

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Xalapa, P.C. 91070 Veracruz, Mexico; e-mail: [email protected]

*Corresponding Author, email: [email protected]

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Highlights By degrading pyrophosphate, inorganic pyrophosphatases promote growth.

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In plants, pyrophosphate can be an alternative source of energy, specially under stress.

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Plant genomes encode multiple isoforms of soluble and membrane-bound

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pyrophosphatases

The regulatory mechanisms of pyrophosphatases are still poorly understood.

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Pyrophosphatases may be relevant to the regulation of secondary metabolism

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Abstract.

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Pyrophosphate (PPi) is produced as byproduct of biosynthesis in the cytoplasm, nucleus, mitochondria

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and chloroplast, or in the tonoplast and Golgi by membrane-bound H+-pumping pyrophosphatases (PPv). Inorganic pyrophosphatases (E.C. 3.6.1.1; GO:0004427) impulse various biosynthetic reactions

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by recycling PPi and are essential to living cells. Soluble and membrane-bound enzymes of high

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specificity have evolved in different protein families and multiple pyrophosphatases are encoded in all plant genomes known to date. The soluble proteins are present in cytoplasm, extracellular space, inside

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chloroplasts, and perhaps inside mitochondria, nucleus or vacuoles. The cytoplasmic isoforms may compete for PPi with the PPv enzymes and how PPv and soluble activities are controlled is currently unknown, yet the cytoplasmic PPi concentration is high and fairly constant. Manipulation of the PPi metabolism impacts primary metabolism and vice versa, indicating a tight link between PPi levels and carbohydrate metabolism. These enzymes appear to play a role in germination, development and stress page 2 of 40

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adaptive responses. In addition, the transgenic overexpression of PPv has been used to enhance plant tolerance to abiotic stress, but the reasons behind this tolerance are not completely understood. Finally, the relationship of PPi to stress suggest a currently unexplored link between PPi and secondary

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metabolism.

Abbreviations

PPi, inorganic pyrophosphate; PPiase, inorganic pyrophosphate hydrolase (E.C. 3.6.1.1; GO:0004427);

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PPa, soluble inorganic pyrophosphatases of family I; PPa-ek, soluble inorganic pyrophosphate hydrolase of the family I, and eukaryotic type; PPa-pk, soluble inorganic pyrophosphate hydrolase of

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the family I, and prokaryotic type; PPv,membrane-bound, proton-pumping inorganic pyrophosphatase.

motif;

HAD,

haloacid

dehalogenase-related

protein

family;

pfp,

fructose-6-

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histidine

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PS2, phosphate starvation response 2 protein; DHH, family protein sharing the aspartic acid-histidine-

phosphate:pyrophosphate 1-phosphotransferase (EC 2.7.1.90); susy, sucrose synthase (EC 2.4.1.13);

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Ugppase, UDP-glucose pyrophosphorylase (EC 2.7.7.9). oppp, oxidative pentose phosphate pathway.

Keywords: inorganic pyrophosphate; inorganic pyrophosphatases; plant metabolism; subcellular

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localization; plant abiotic stress; plant stress responses; photosynthetic organisms.

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Table of Contents 1 Introduction .............................................................................................................................................4

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2 Inorganic pyrophosphatase activities in living cells. ..............................................................................4 2.1 DHH soluble pyrophosphatases. ....................................................................................................5 2.2 Haloacid dehalogenase related pyrophosphatases (PS2). ...............................................................5 2.3 Classic soluble pyrophosphatases of family I (PPa). .....................................................................6 2.3.1 Prokaryotic-type classic soluble pyrophosphatases of family I in plants. ..............................7

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2.3.2 Chloroplastic classic soluble pyrophosphatases of family I (PPa). ........................................8

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2.4 Membrane bound inorganic pyrophosphatases in plants. ..............................................................9

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3 Function and regulation of soluble inorganic pyrophosphatases from plants. ......................................10

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3.1 Expression and compartmentalization of inorganic pyrophosphatases. ........................................ 11

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3.2 Regulation of plant soluble inorganic pyrophosphatases by ligands. ...........................................12

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3.3 Regulation of plant soluble inorganic pyrophosphatases by covalent modification. ....................13 4 Role of pyrophosphatases in plants under stress and biotechnological applications. ...........................14

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4.1 Pyrophosphatases in relation to plant growth and development ...................................................14 4.2 Possible link between pyrophosphatases and stress responses in plants. ......................................15

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5 Conclusions and perspectives ...............................................................................................................21 Acknowledgements .................................................................................................................................21 Competing interests.................................................................................................................................22

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Abbreviations ..........................................................................................................................................22 References ...............................................................................................................................................22

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Introduction

All living cells generate pyrophosphate (PPi) as a byproduct of the biosynthesis of DNA, RNA, proteins, polysaccharides and membrane lipids [1] (Figure 1). As cells grow, PPi accumulation is

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prevented by the action of a number of proteins with inorganic diphosphate phosphohydrolase activity (E.C. 3.6.1.1; GO:0004427), commonly known as inorganic pyrophosphatases [2] (PPiase). All living cells studied to date contain one or more proteins with bona fide PPiase activity. Phosphohydrolases of low specificity such as alkaline phosphatases (EC 3.1.3.1), acid phosphatases (E.C: 3.1.3.2), or

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glucose-6-phosphatase (EC 3.1.3.9) [3–5] are able to hydrolyze PPi in vitro, but these are not

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considered true PPiase enzymes, as they are unlikely to play such role in vivo. The inorganic

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pyrophosphatases are required to have high specificity for inorganic PPi, because they have access in

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vivo to organic polyphosphates such ATP, ADP and NAD(P)+ which should not be hydrolyzed by these enzymes. Furthermore, their activity is probably tightly regulated, given the intricate complexity of

Inorganic pyrophosphatase activities in living cells.

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compartments (Fig. 1).

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plant metabolism which generates PPi through various biosynthetic pathways and in diverse

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The ability to catalyze the hydrolysis of pyrophosphate has evolved in nature several times in completely different protein families, and most but not all of these families are represented in plants.

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While previous reviews focusing on PPiase enzymes from different protein families have been published [1,2,6–8], these reviews deal with either the soluble [1,2,6] or membrane bound proteins [7,8]. Here an integrative overview of the features and properties of both soluble and membrane-bound PPiase proteins is provided, with focus on those representatives found in plants. page 5 of 40

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DHH soluble pyrophosphatases.

The Mn2+-dependent family II pyrophosphatases belong to a large superfamily of phosphohydrolases found in Bacteria, Archaea, and Eukarya (PFAM PF01368), sharing an Asp-His-His (DHH) motif.

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Some bacterial members of this protein family, including those present in pathogens, exhibit Mn2+dependent inorganic pyrophosphatase activity, with high specificity for PPi, and have been designated as the family II of inorganic pyrophosphate hydrolases [9]. Other orthologs of the same family found in other bacteria, yeast and animals exhibit exopolyphosphate hydrolase or phosphodiesterase activity

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[10,11]. Notably, none of the known plant proteins exhibit significant sequence similarity to the

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bacterial family II pyrophosphatases, the yeast exopolyphosphatase, the animal PRUNE protein, the

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CDC45 replication protein, or to the bacterial RecJ enzyme (from a BLAST search against NCBI's

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refseq [12] genomes database, on june 2017). A recent phylogenetic analysis of bacterial RecJ protein family suggest an ancient archeal early origin for these proteins [13]. Therefore, the plant ancestors

Haloacid dehalogenase related pyrophosphatases (PS2).

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seem to have lost this gene early in evolution.

Initially identified as highly overexpressed proteins in response to phosphate starvation in plants [14],

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the A. thaliana phosphate starvation response protein 2 (AtPS2) and its orthologs have similarity to the haloacid dehalogenase (HAD) superfamily. This is a functionally diverse protein family, initially

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associated to the hydrolytic cleavage of halogen-carbon bonds [15]. In plants, the Solanum lycopersicum protein (SlPS2, formerly LsPS2) has phosphatase activity at acid pH [14] and has been reported to exhibit okadaik acid-sensitive ser/thr protein phosphatase activity [16]. By contrast, the AtPS2 protein was found to be specific for PPi at alkaline pH [17]. This last page 6 of 40

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protein showed a basal Mg2+-independent activity, but could be activated by Mg2+ and was found active as tetramer [17]. Considered as cytoplasmic enzymes [18], the physiological role of these enzymes is still a mystery, because: (i) there are two or more isoforms in arabidopsis and tomato, and apparently

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also in most vascular plants (from sequence alignment to refseq [12]), and (ii) the analysis of their expression in tomato shows genes with tissue-specific expression patterns, not exclusive to the phosphate starvation response, but expressing also under phosphate sufficient conditions [18].

Interestingly, a soluble inorganic pyrophosphatase activity purified from etiolated maize leaves [19],

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presented a phosphatase activity at pH 6.2, with Zn2+ non-essential activation, but also Mg2+-dependent

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pyrophosphatase activity at pH 9. These activities could not be separated by chromatography [19].

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Perhaps, the PS2 enzyme class exhibits dual activity, or alternatively, the A. thaliana and tomato

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orthologs differ in specificity and metal requirements; clearly, this is still an open question. In addition, HAD bacterial proteins with PPi-specific hydrolytic activity have been reported [20] and

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their essential role during DNA polymerization in vivo has been demonstrated [21], however, the A.

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thaliana and bacterial HAD pyrophosphatase orthologs share only 25% identity and 38% similarity (∂-

Classic soluble pyrophosphatases of family I (PPa).

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2.3

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BLAST alignment [22]).

With high specificity for PPi [23], these enzymes are widely distributed among Prokaryotes (PPa-pk)

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and Eukaryotes (PPa-ek). They exhibit high activity against PPi at alkaline pH, with Mg2+ as their essential and physiological activator [23], and in many cases Ca2+ inhibits their activity (see below, section ), although the protein from Leishmania major was reported to be activated by Ca2+ [24]. The prototype of the PPa-pk is the homohexameric PPiase from E. coli [25], while there is a well page 7 of 40

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characterized homodimeric PPa-ek prototype in Saccharomyces cerevisiae [26]; yet, the recently reported protein from Trypanosoma brucei brucei has a novel tetrameric structure [27]. In plants, there are both PPa-pk and PPa-ek orthologs, but these proteins function as monomers [28,29]. All the

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members with solved structure show a five-fold αβ imperfect barrel at the catalytic core, but differ significantly in their N- and C-terminal extensions [2,6,23,27,30,31].

2.3.1 Prokaryotic-type classic soluble pyrophosphatases of family I in plants.

Six soluble PPa proteins are encoded in A. thaliana genome (designated as AtPPa1 to AtPPa6). AtPPa1

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to AtPPa5 are known to be cytoplasmic proteins [32,33], and there is evidence confirming the presence

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of PPa proteins in the cytoplasm from potato tubers [34]. The AtPPa1 and AtPP4 isoforms were cloned,

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expressed, purified and characterized as recombinant proteins [29]. These isoforms were shown to be

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Mg2+-dependent, and Mn2+ was a poor activator, while other divalent cations tested were inhibitors.

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Both enzymes were highly specific for PPi but displayed a complex kinetics [29]. All of these

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cytoplasmic AtPPa isoforms share more sequence similarity to the PPa-pk proteins [28,35] and a minimum of three different isoforms seem to be present in all terrestrial plants [33]. There are many

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reports on the kinetics and molecular properties of pyrophosphatases directly purified from plant

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tissues, but many of them were performed at the time when genetic information was scarce and in most cases the data cannot be associated to a particular protein. In addition, there is a significant variation in

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the molecular mass reported for the subunit, the assigned oligomeric state (when given) and in the relative affinities for PPi and Mg2+ [19,36–38]. Fortunately, a few reports present enough information to identify the protein as belonging to the PPa-pk group [39–41], and the data presented are consistent with the properties found for the Arabidopsis enzymes. In this sense, AtPPa1 and AtPPa4 enzymes

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appear to be good representatives of the enzymes of this class in plants; yet, there is still no satisfactory explanation to the need for several active isoforms of PPa-pk in plants, because their kinetic properties are very similar [29], more than one has constitutive expression [35], and because there are examples of

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unicellular photosynthetic Eukaryotes lacking a cytoplasmic protein of this class [28,33].

2.3.2 Chloroplastic classic soluble pyrophosphatases of family I (PPa).

In contrast to the PPa-pk proteins encoded in A. thaliana genome, AtPPa6 appears to be an exclusively chloroplastic enzyme [33,42]. It is the most abundant [36,43] and is the only one truly essential for

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plant survival [32], although no attempt has been reported to produce a multiple null mutant lacking all

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of the PPa-pk isoforms. The plant chloroplastic PPa protein shares higher sequence similarity with PPa-

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ek proteins and despite being essential, neither the kinetics, nor the structure of the pure chloroplastic

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PPa-ek from A. thaliana (AtPPa6) have been studied in detail. Orthologs from other species have been

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purified from plant tissues and characterized [36,38,44,45], and their properties are consistent with

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those of bona fide PPiase enzyme, but contamination of these preparations by PPa-pk isoforms cannot be ruled out. A recent work [28] reported the cloning and expression the two encoded PPa proteins

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from the photosynthetic Eukaryote Chlamydomonas reinhardtii. In this work, the authors report also

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the isolation of the chloroplastic protein from spinach, the protein from the cyanelle of the photosynthetic microalgae Cyanophora paradoxa and the organellar PPa proteins from other

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photosynthetic Eukaryotes. In all of the studied organisms, these authors found most of the activity associated with organelles, and this activity was Mg2+-dependent, presented high affinity for PPi, and the protein identity was confirmed by mass spectrometry or in some cases by Edman degradation [28]. The obtention of a pure recombinant AtPPa6 mature peptide would help to confirm the plant PPa-ek

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enzyme properties, and would allow a more in depth comparison of the kinetic and regulatory properties of PPa-ek and PPa-pk proteins, but this work is still pending. In addition, the plant chloroplasts are considered descendants of endosymbiotic cyanobacteria and a

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reasonable expectation would be to find in this organelle an inherited member of a PPa-pk subfamily, instead of a member of the PPa-ek subfamily, formerly associated to higher Eukaryotes [2,6]. Furthermore, the genomes of microalgae from order Mamiellales (Viridiplantae) did only encode for the putatively chloroplastic PPa-ek isoform [33], making unclear the origin and diversification of this

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protein family in plants. Some efforts to group the amino acid sequences of PPa proteins in

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photosynthetic Eukaryotes have been made [2,28,33], but a robust and properly rooted phylogenetic

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analysis of this protein family in plants is still missing and could provide better hypotheses related to:

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when did the PPa-ek protein first appear? how did this protein subfamily diversify? was there a PPa-pk protein ancestor in primitive Eukaryotes and was lost in Amoebozoa (amoebas) and Opisthokonta

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(fungi, animals, and other unicellular Eukaryotes with posterior flagellum) after their separation from

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Archaeplastidia (Green Plants, Rhodophytas and Glaucophytas)? or were PPa-pk proteins acquired by

Membrane bound inorganic pyrophosphatases in plants.

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2.4

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photosynthetic organisms later on?

In addition to the soluble forms of inorganic pyrophosphatases, plants have membrane-bound enzymes

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with inorganic diphosphate hydrolase activity (PPv) [46,47]. The PPv isoenzymes localize to the tonoplast (Fig. 1, 10) [46], Golgi apparatus [48], and plasma membrane [49]. These Mg2+-dependent enzymes can couple the PPi hydrolysis to translocation of protons across the membrane and sustain secondary active transport. Such activity may play a role in nitrate accumulation [50], and may be page 10 of 40

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involved in the vectorial movement of auxins from the roots to the leaves [51]. From subcellular fractionation, activity measurements and enzyme kinetics, a membrane-bound pyrophosphatase activity was found in the chloroplast (Fig. 1,17?) [52], but its gene has not been identified and the protein was

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not purified. Possibly, the chloroplast enzyme is one of the PPv isoforms [53], since the transport of plant proteins from the Golgi to the chloroplast has been shown to occur [54] and there is proteomic evidence of a PPv-like protein in chromoplast membranes [55,56]; however, evidence is still inconclusive.

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A recent report has demonstrated a role for the PPv from A. thaliana (AVP1) as a PPi synthase [57],

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and apparently this activity is important for the maintenance of PPi homeostasis in the phloem, favors

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the flow of photoassimilates to the root, and it is necessary for normal growth. Using immunogold and

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epifluorescence, the A. thaliana PPv AVP1 was found in the xylem vessels of the primary vein of leaves, in the sieve elements (Ses), and in companion cells (CCcs) [49].

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Given the relevance of H+-pumping pyrophosphatases in plant metabolism, the role of PPi as an

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alternative source of energy in plants (see below, section ) is a reality, at least in relation to membrane

Function and regulation of soluble inorganic pyrophosphatases from plants.

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polarization and secondary active transport.

As shown in figure 1, in plant cells, PPi is produced in several cell compartments, but PPiase activity

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has only been demonstrated in the cytoplasm (Fig. 1-9), the tonoplast and Golgi (Fig. 1-11), and the chloroplast (Fig. 1-16). In addition, the tonoplast membrane-bound enzyme and the soluble cytoplasmic isoforms could compete for the substrate. In fact, the simultaneous unregulated operation of the PPv, the vacuolar proton-pumping ATPase and the PPa-pk plant isoforms would consume the page 11 of 40

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celular ATP cytoplasmic pools making the cell unviable. Unfortunately, as discussed in this section the mechanisms that regulate all these proteins are not well understood.

Expression and compartmentalization of inorganic pyrophosphatases.

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3.1

While PPi hydrolysis is essential to support growth in living cells [32,58–61], the several PPa and PPv isoforms present in plants complicate the analysis of their expression and regulation under normal development or under stress; and only recently this aspect has received some attention [35,62]. Many of the cytoplasmic PPa-pk isoforms show constitutive expression [29,35,62], but 90% of the extractable

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soluble Mg2+-dependent PPiase activity was found associated to the chloroplast [43], and in addition,

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chromatographic fractionation [29, 33] and gel zymograms [35] have revealed a complex and changing

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activity profile [33, 35]. A detailed analysis of the promoter regions of the genes encoding the PPa-pk

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isoforms using the classic functional promoter dissection together with the search for miRNA acting of

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the PPa-pk transcripts would most probably lead to better understanding of how the expression of these

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genes is regulated.

In a recent work [33], the authors provided evidence for a non-overlapping localization of A. thaliana

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PPa-pk isoforms, but there is still limited information to link this differential distribution to specific

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pathways or functions. The same work also found the AtPPa4 isoform in the apoplast, and again the function of this PPa-pk isoform within this compartment is currently unknown, as it is the trafficking

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mechanism that carries it out there [33]. Certainly, there is still pending research in relation to the mechanisms controlling the trafficking of AtPPa4, and the interactions behind the uneven distribution of PPa-ek isoforms. Novel proteomic approaches and yeast 2 hybrid experiments could help to identify the protein partners of these enzymes and should help to answer such questions. page 12 of 40

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Regulation of plant soluble inorganic pyrophosphatases by ligands.

At the activity level, little is known about the regulation of PPa-pk plant isoforms, but the high levels of PPi in plant cytoplasm suggest a form of control of their activity. Possible mechanisms include pH

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changes, Mg2+ levels, Ca2+ levels, the high concentration of phosphate and their association to other proteins within defined subcytoplasmic domains. Calcium has been shown to be an inhibitor of the potato cytosolic enzymes [29,63], but the Ca2+ concentration required to inhibit the enzyme is above 10 µM, and the physiological impact of this inhibition is unclear. The concentration of free Mg2+ and Ca2+

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has also been proposed to be a regulatory factor for PPv enzymes [46].

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Certainly, the drastic change in Mg2+ concentration associated to the operation of photosynthesis is

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known to be a major factor in the control of the Calvin cycle in chloroplast, and this change have some

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effect on the cytoplasmic Mg2+ concentration [64]. Mg2+ is the known physiological activator for both PPa and PPv, and these changes in its concentration are very likely to modulate the activity of the

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chloroplastic and/or cytoplasmic pyrophosphatases in photosynthetic tissue. However, current evidence

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is not enough for predicting the amplitude of PPa and PPv activity changes associated to Mg 2+

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variations along the light/dark cycle. Clearly further research is required to fully understand the role of this divalent ion in the regulation of PPiase enzymes.

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The effect of orthophosphate as inhibitor cannot be studied with the very sensitive standard assay, because added Pi interferes with the quantification of the Pi produced by PPi hydrolysis. A different

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assay would be needed to tackle this question, but since Pi in the plant cytoplasm can reach a concentration two or times higher than PPi [65], this factor deserves consideration. Other metabolites may also influence PPa and PPv activity, but currently, based on in vitro evidence, only L-malate has been proposed to play a physiological role in the regulation of the PPa-ek isoform [44] and this page 13 of 40

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observation has not been confirmed.

3.3

Regulation of plant soluble inorganic pyrophosphatases by covalent modification.

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Covalent modification may be an additional form a regulation for these enzymes, and in Papaver rhoeas pollen-specific PPa proteins were found to be inhibited by phosphorylation in vivo during incompatible pollen rejection at the flower style [66]. Recently, the phosphorylation of these proteins was found to enhance Ca2+ and hydrogen peroxide inhibition in vitro, and this has been proposed as a regulatory mechanism relevant to pollen rejection [41]. Furthermore, A. thaliana proteins AtPPa1,

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AtPPa3 and AtPPa5 were found phosphorylated in a phosphoproteomic analysis of the plant response

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to iron deficiency [67], although no further analysis of that observation was presented. In accordance,

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Molecular Dynamics simulations of modeled AtPPa1 and AtPPa4 proteins suggested a significant

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influence of an N-terminal segment over the active site conformation [30], the segment involved seems

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to include the putative phosphorylation site in AtPPa1. Evidence of regulation by in vitro

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phosphorylation has also been reported for PPa-pk enzymes in bacteria [68] and for the PPa-ek in animal tissues [69]. Clearly, more work is needed to establish if phosphorylation is a general form of

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regulation for PPa enzymes and how this modification relates to the physiology of the plant.

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In addition, regulation by redox state has been shown to take place in a the PPa-ek enzyme from the cattle parasite tick Rhipicephalus microplus [70]. Redox regulation has been shown to play a central

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role in the regulation of chloroplasts enzymes [71], and is at this organelle where the PPa-ek isoform is located. The maturation proteolytic signal in the N-terminal peptide of the AtPPa6 protein is currently unknown, and only one cysteine is expected to remain in the mature enzyme. Nevertheless, it seems worth testing if this protein is sensitive to regulation through changes in its redox state.

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Role of pyrophosphatases in plants under stress and biotechnological applications.

4.1

Pyrophosphatases in relation to plant growth and development

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In contrast to animal tissues, PPi accumulates in the cytoplasm of plant cells [43] and contrary to ATP, its concentration remains high under stress conditions causing energy-deficit [72,73]. Alterations of PPi metabolism by transgenesis reduces plant growth, apparently through changes in the metabolism of carbohydrates, which affect carbon partitioning between source and sink organs [74–76]. For example,

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transgenic expression of the E. coli PPa protein in the phloem tobacco, reduced cytosolic PPi pools,

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induced accumulation of soluble sugars in older leaves and increased starch accumulation in all the

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leaves, resulting in stunted growth [74]. This last observation suggests a possible role for PPi on the

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phloem, and is consistent with: (i) the extracellular localization of AtPPa4 [33], (ii) data demonstrating a role for PPv in the phloem [57], and (iii) the unexpected ability of a transgenic yeast invertase to

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compensate the negative effects of the bacterial PPiase, when both proteins are overexpressed together

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in the phloem [77].

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On the other hand, the fugu5 mutant with reduced expression of the AVP1 protein (tonoplast PPv) presents an increased level of PPi and has defects in seed development and germination. The fugu5

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phenotype can be rescued by complementation with the yeast PPa-ek under the control of the A. thaliana AVP1 promoter [61]. The authors provided evidence indicative of gluconeogenesis inhibition

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by PPi, rather than impairment of vacuole acidification [78]. According to this last evidence, soluble PPiases would not be active in the Arabidopsis' seed, but RNAi silencing the expression of AtPPa1 and AtPPa4 resulted in a higher accumulation of seed storage lipids [79]. A reasonable explanation to these opposing observations could lie in the regulatory mechanisms of cytoplasmic PPa-pk enzymes, which page 15 of 40

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may limit the access of AtPPa-pk isoforms to cytoplasmic PPi pool and favor its use by AtPPv. This mechanisms may still operate in the fugu5 mutant, but could fail to control a heterologous protein, such

the PPi-pk proteins in the seed, through a still unidentified link.

4.2

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as the yeast PPa-ek protein. Alternatively, the fugu5 mutant might also show a reduced expression of

Possible link between pyrophosphatases and stress responses in plants.

In addition to their main role in the recycling of PPi, plant PPa and PPv proteins may have additional roles in the adaptive responses to stress [61,80,81]. Based on a number of metabolic changes in planta

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under phosphate deficiency, a previous work [82] has proposed a plant adaptive response to this stress

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condition which makes use of PPi as an energy source, involving an alternative form of glycolysis. This

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response may be in operation under stress conditions leading to an energy deficit, such as anoxia or

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phosphate deficiency. However, this and other stress responses are accompanied by a reduction, or even

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a halt, in plant growth [83], which should limit biosynthesis and its associated production of

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pyrophosphate [84,73]. Since, unlike ATP levels, the PPi pools are fairly stable under stress [73,85,86], the consumption rate should be equal to its production, and the pyrophosphatase activity should be

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modulated to match the drop in PPi generation. In fact, phosphate starvation was shown to associate with changes in transcriptional expression and activity patterns of PPa proteins in common bean [35],

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the AtPPa4 protein was found phosphorylated in response to iron deficiency in A. thaliana [67], and

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transgenic expression of a bacterial PPa-pk in potato reduced plant survival under anoxia [87]. In this scenario, PPi could then be a relevant energy source only if an alternative source of pyrophosphate is available under energy deficit. In this sense, PPi synthesis has been observed by pea isolated mitochondria [88], and the existence of a H+-pumping pyrophosphatase in the mitochondrial inner membrane has been proposed [89], unfortunately, in none of these cases the corresponding protein has page 16 of 40

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been identified. On the other hand, overexpression of a PPv (AVP1) in A. thaliana, tomato and other plant species enhanced their tolerance to limiting phosphorous conditions [81,90], and salt stress or drought [91–93].

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Thus, the transgenic expression of PPv proteins have been proposed as a way to enhance crop production [90,91,94], although care should be taken, because the overexpression of PPv in the rice developing seed increases the grain chalkiness, an undesirable trait [95]. In addition, PPv has been implicated in auxin transport and organ development [51], but this role has been questioned recently

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[96] because avp1-1 (GABI-kat 05D04) used in this study has a second alteration in a neighbor locus

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encoding the ARFGEG GNOM protein, which is involved in the establishment of root basal polarity

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and participates in the correct trafficking of PIN auxin transporters [97]. Clearly the mechanism behind

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the beneficial effects of PPv overexpression deserves additional research. From the biotechnological point of view, tracking possible alterations to the relative growth rates of aerial parts and of the root

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system, as well as changes in crop yields, caused by the overexpression of PPv requires further

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consideration.

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One condition where experimental evidence points to a central role for PPi cytoplasmic pools is hypoxia. Hypoxia is a stressful condition affecting plants under flooding and when soils are too rich in

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organic matter, and it is prevalent in species adapted to grow in marshes, riversides and estuaries, such as rice, mangroves, and the Montezuma bald cypress, among others. Extreme hypoxia becomes anoxia

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and stops ATP provision through mitochondrial respiration, plant tissues have developed a modified form of glycolysis which involves three possible bypasses to ATP consumption [84] and may take advantage of the use of PPi as energy source [73]. Three PPi-dependent mechanisms are likely to be central to this adaptive response, and these have recently been reviewed in great detail [98 and page 17 of 40

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references therein]. Briefly, these include: (i) The preferential acidification of the vacuole by the PPv instead of the V-ATPase (Fig 1.). (ii) The sucrose mobilization via sucrose synthase plus UDPG pyrophosphorylase (see Fig 2.)[84], and (iii) the use of alternative enzymes to reduce ATP

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consumption, mainly PPi-dependent phosphofructotransferase (Fig 3)[98], PEP phosphatase [84] and pyruvate orthophosphate dikinase (EC 2.7.9.1) [73, 84,98].

A second stress condition where the link to PPi is evident is salinity, because the Na negative effects in plant metabolism can be alleviated by the compartmentation of this cation in the vacuole in a secondary

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active exchange with protons, using either the vacuolar H+-ATPase or the PPv as primary pump [99].

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The biotechnological use of this knowledge has already been mentioned, was proposed at the beginning

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of the century [91,100] and the mechanisms behind the regulation of this response have recently been

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reviewed [99]. Recent metabolomic data from euryhaline seagrass Cymodocea nodosa [101] and proteomic evidence from rice seedling germinating under salt stress [102] are consistent with a role for

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tolerance to salt stress in plants.

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PPi metabolism, and PPv and PPa enzymes in the biochemical and physiological mechanisms of

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In addition, multiple stress conditions are known to induce changes in the secondary metabolism [103]. The production of a number of secondary metabolites in plants has a role in protection against

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oxidative, osmotic, ionic and biotic stress, and it has been proposed as an energy overflow protection mechanism, when plant growth is limited by the stress condition, but the photosynthetic apparatus is

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still under daily illumination [103,104]. The biosynthetic pathways involved do require energy in the form of ATP and NADPH, but an energy deficit may accompany some stress conditions, specially phosphorous deficiency or anoxia, known to cause ATP levels to drop substantially [73,84,86]. Energy deficit could become critical at the sink tissues because normal glycolysis would be impaired by ATP page 18 of 40

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limitations. The glycolytic bypasses proposed and already mentioned [82,84,98] would become relevant, and in the present review, a novel oxidative pentose phosphate pathway (oppp) operational mode is also proposed (Fig. 2), which could help to metabolize sucrose and produce NADPH in non-

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photosynthetic plant cells, using PPi to reduce ATP consumption. As for other oppp operational modes previously proposed [71], the operational mode described here would not be exclusive. Thus, the fraction of the total metabolic flow going through the pathways depicted in figure 2, at any time, would depend on the prevailing conditions. In the scheme (Fig. 2), sucrose could be degraded by sucrose synthase (Fig. 2, susy) and the energy in the glycosidic bond would be conserved in the UDP-glucose

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bonds. UDP-glucose pyrophosphorylase (Fig. 2, Ugppase) could convert UDP-glucose into glucose-

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phosphate and one ATP equivalent (as UTP), which could serve to phosphorylate the fructose. After

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conversion into fructose-6-phosphate, the oppp pathway could oxidize the hexoses, provide fructose-

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1,6-biphosphate to serve as PPi source through the PPi-dependent phosphofructotransferase (Fig. 2,

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pfp), while the remaining fructose-6P could be channeled into oppp again.

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Erythose-4P could remain in the plastid and along with additional pentose-P could be used to build up the regenerative phase of oppp and eventually support the operation of the shikimate pathway, provide

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more triose-P for glycolisis and respiration and also for terpene biosynthesis, required to support

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carotenoids and ABA biosynthesis in the plastids (Fig. 2), or sterols in the cytoplasm (shown in Fig. 1). Terpene biosynthesis does require ATP, but also preserves part of the energy as PPi, which under

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energy deficit could support the acidification of the vacuole [84,90] (Fig. 1). A careful analysis of the basic stoichiometry is shown in figure 3 and supports complete degradation of 2 sucrose (24 carbons, no phosphate) and 2

inorganic phosphate (2-phosphates) using only only 2 ATP (introducing 2

phosphates in the sugar products). These sugars would be converted into 2 ribulose-5-phosphate (10 page 19 of 40

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carbons, 2 phosphates), 2 erythrose-4-phosphate (8 carbons, 2 phosphates), 12 NADPH (and 12 protons), and 6 CO2 (6 carbons). Here two triose-P can produce one fructose-1,6-P2, and two ATP are consumed to convert one fructose-6P (produced directly by the regenerative segment of oppp) into

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fructose-1,6-P2. ATP could be produced from PPi by the combined action of the vacuolar protonpumping ATPase and PPv, however, under stress many biosynthetic reactions slow down and the supply of PPi is reduced. Under this scenario, an active consumption of PPi would deplete the PPi pools, which is inconsistent with experimental observations [85–87]. A novel source of PPi might exist,

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but it is yet to be discovered.

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The scheme in figure 2 is speculative, but is supported by the following observations:

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A) All the enzymes and pathways considered are known to be expressed in plant tissues [84,71].

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B) The oppp is known to present several operational modes in plants, and biochemical and genetic

D

evidence is consistent with the expression of active decarboxylating segments of this pathway in both

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cytoplasm and plastids, for a wide range of tissues from plants of different families [71]. C) Metabolomics evidence from cell suspensions under phosphorous limitation indicates an increased

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carbon flow through the oppp under the stress, in comparison to normal conditions [105], and oppp has

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been found to protect plants against oxidative stress, which is inherent to energy deficit [106]. D) The activity of sucrose synthase (Fig. 2, susy) [84], pyrophosphosphate:glucose-6-phosphate 1-

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fosfotransferase (Fig. 2, pfp) [84], and UDP-glucose pyrophosphorylase (Fig. 2, Ugppase) [107] are known to increase under some stress conditions, including those causing energy-deficit. D) Mutations of the NDB1 burst NADPH dehydrogenase in arabidopsis resulted in an elevation of NADPH/NADP+ ratios, and this in turn, resulted in enhanced expression of many genes related to plant page 20 of 40

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defense responses [108]. E) There is evidence of important roles for the shikimate [109] and the phenylpropanoids [110]

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biosynthetic pathways under stress conditions. F) Overexpression of a plasma membrane isoform of geranylgeranyl pyrophosphate synthase in sweet potato increased carotenoid accumulation and afforded protection against hydric and oxidative stress [111]; and terpene biosynthesis is known to produce up to 8 mol of PPi per mol of triterpene [112]. In this sense, the chromoplast of Capsicum annuum is known to have a very active PPiase [45], and the

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regulation of its expression and activity has not been studied in relation to pigment accumulation

N

during fruit development under neither normal nor stressful conditions, but recent proteomic evidence

A

indicates changes in the content of the PPa-ek protein during the ripening of tomato fruit [56]. Thus, a

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detailed study of this activity and its role on fruit development seem relevant, at least in Solanales.

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One very interesting prediction of the scheme in figure 2 is the need for a tight regulation of oppp

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intermediates branching to support the synthesis of purines and cytokininis, or shikimate and aromatic amino acids. Under ATP sufficient conditions, purines and cytokinins biosynthesis should occur

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without limitations to stimulate and support normal plant growth, and the shikimate pathway should also operate to provide aromatic amino acids, some cell wall precursors and other metabolites. Under

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stress, particularly under energy deficit, the synthesis of purines would be limited and the production of

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shikimate and phenylpropanoids would consume NADPH and would help to protect the cell against NADP+ exhaustion and free radical accumulation. A number of approaches can be used to explore this prediction experimentally and whatever the outcome, the data should improve our understanding of plant physiology under stress.

page 21 of 40

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Conclusions and perspectives

Despite an important research effort, many aspects of the PPi metabolism in plants are still a mystery. Among other problems, the significant diversification and subspecialization of the PPi hydrolases

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makes difficult to explore their individual roles and regulatory mechanisms, particularly at the protein level. In addition, many studies in the literature tend to focus on the PPv and overlook the PPa proteins, or the other way around. To complicate the issue, several other enzymes may catalyze the unspecific hydrolysis of PPi in vitro, and interfere with classic biochemical approaches. Surely, modern

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approaches of genetic manipulation could be used to tag individual PPa and PPv proteins in vivo and

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study their synthesis, trafficking, activity regulation, and degradation in planta, under normal and

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stressful conditions. These studies would benefit from an integral approach, taking into account both,

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the membrane-bound and soluble PPiases in the plant cell, and should be backed up by a more comprehensive biochemical and molecular characterization of the plant proteins with PPiase activity,

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belonging to the different families.

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Finally, secondary metabolism is also known to be the subject of regulation under stress [103] and

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some metabolic pathways, such as terpene biosynthesis, can produce a large quantity of PPi [112]. Therefore, PPi could be an alternative energy source under stress if secondary metabolism can maintain

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the cytoplasmic pools of PPi.

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Acknowledgements This work was done in relation to projects funded by PAPIIT-DGAPA-UNAM: IN216815, SC-DGTICUNAM/LANCAD: SC151IR47 and PAIP-FQ-UNAM: 5000-9122. We thank the anonymous reviewers for their helpful comments. page 22 of 40

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Competing interests

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All authors declare having no competing interests.

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e1002323.

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Legend to Figures

Figure 1. Main pathways of pyrophosphate production and degradation in plants. Processes or

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reactions shown are: (1,2) DNA and RNA polymerases, (3?) nuclear soluble pyrophosphatase, (4?) Pi and/or PPi (5?) diffusion into cytoplasm, (6?) mitochondrial PPi synthesis, (7?) mitochondrial

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pyrophosphatase, (8?) mitochondrial PPi transport to the cytoplasm, (9) cytosolic soluble pyrophosphatases, (10) tonoplast-bound H+-pyrophosphatase (also in Golgi and plasma membrane, not

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shown), (11) tonoplast-bound H+-ATPase, (12) PPi production by protein and sterol biosynthesis in the cytoplasm, (13) PPi produced by sucrose synthesis in cytoplasm, (14) thylakoidal ATP synthase, (15) pyrophosphate produced by DNA, RNA, protein, starch and lipid synthesis in chloroplast stroma, (16) chloroplastic soluble inorganic pyrophosphatase, (17?) chloroplastic pyrophosphate synthesis. page 36 of 40

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Numbered with a question mark superindex represent proposals with incomplete experimental support.

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Figure 2. Possible sucrose degradation pathway with reduced ATP requirements, which might take place in non-photosynthetic tissues of plant cells under energy-deficit. The steps shown with gray arrows represent reactions likely to exhibit limited flow at low ATP level. Processes or reactions

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depicted include 4 transport systems at the inner plastid envelope: 1, phosphoenolpyruvate:phosphate translocator; 2, pyruvate:H+ symporter; 3, xylulose-5-phosphate:phosphate translocator ; 4, glucose-6phosphate:phosphate translocator. Here, translocators 3 and 4 may carry the indicated exchange alone or in combination with the triosephosphate:phosphate translocator (not shown). Individual enzymes depicted are: ald, aldolase (EC 4.1.2.13); fk, fructokinase (EC 2.7.1.4); pfp, fructose-6-phosphate:

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pyrophosphate 1-phosphotransferase (EC 2.7.1.90); pk, pyruvate kinase (EC 2.7.1.40); pgi,

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phophoglucoisomerase (EC 5.3.1.9); pgm, phosphoglucomutase (EC 5.4.2.2);

nk, nucleotide

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diphosphate kinase (EC 2.7.4.6); rppik, ribose-5P, pyrophosphokinase (EC 2.7.6.1); susy, sucrose

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synthase (EC 2.4.1.13); Ugppase, UDP-glucose pyrophosphorylase (EC 2.7.7.9). Pathway segments

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shown (condensed into a single arrow) are: oppp, oxidative pentose phosphate pathway, subdivided

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respectively in cdoppp and pdoppp, the cytoplasmic and the plastidial segment of the decarboxylating oxidative pentose phosphate pathway; and roppp, the regenerative segment of oxidative pentose

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phosphate pathway (mostly plastidial, but some steps can also occur in cytosol). Metabolites

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abbreviations are: CKs, citokynins; pentose-P, xilulose-5P and/or ribulose-5P and/or ribose-5P; pyr, pyruvate; prpp; phosphoribosyl pyrophosphate; triose-P, glyceraldehyde-3P and/or dihydroxyacetone-P.

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Notes: †The biosynthetic pathway of terpene derivatives produces up to 8 mols of pyrophosphate/mol of lipid.

ǂ→

Phenylalanine should be exported to cytoplasm to serve as substrate to phenylalanine

ammonium liase, the first enzyme of phenylpropanoid pathway.

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Figure 3. Basic stoichiometry of the low ATP requiring sucrose degradation segment shown in figure 2. The steps where PPi is consumed or produced are circled, the final carbon products are enclosed in a rectangle and the net ATP consumption is enclosed in an hexagon. Abbreviations are the

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same as in figure 2, but some enzymes steps are made explicit here: gpdh, glucose-6-phosphate dehydrogenase (EC 1.1.1.49); gnpdh, phosphogluconate dehydrogenase (NADP+-dependent,

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decarboxylating; EC 1.1.1.44); pfk-1, ATP:fructose-6-phosphate 1-phosphotransferase (EC 2.7.1.11);

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gluconolatonase

(EC

3.1.1.17)

is

also

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