Abstract The plant vacuole is a cellular compartment that is essential to plant development and growth. Often plant vacuoles accumulate specialized metabolites, also called secondary metabolites, which constitute functionally and chemically diverse compounds that exert in planta many essential functions and improve the plant’s fitness. These metabolites provide, for example, chemical defense against herbivorous and pathogens or chemical attractants (color and fragrance) to attract pollinators. The chemical composition of the vacuole is dynamic, and is altered during development and as a response to environmental changes. To some extent these alterations rely on vacuolar transporters, which import and export compounds into and out of the vacuole, respectively. During the past decade, significant progress was made in the identification and functional characterization of the transporters implicated in many aspects of plant specialized metabolism. Still, deciphering the molecular players underlying such processes remains a challenge for the future. In this review, we present a comprehensive summary of the most recent achievements in this field. Introduction It was in the middle of the 19th century that the word vacuole was first used to describe an apparently empty cellular organelle present in infusoria. Soon it became evident that despite this initial deceptive description, these fluid-filled organelles are involved in an incredibly diverse array of cellular functions. The vacuoles maintain cellular turgor and cellular homeostasis, and are involved in stomatal movement, cellular protection against UV light and in defense against herbivores and pathogens (Martinoia et al. 2012). The chemical composition of the vacuolar sap is plant, organ, tissue and cell specific, and it is altered upon development and environmental cues. The vacuolar membrane or tonoplast contains transporters and channels that are responsible for regulation of the fluxes of solutes and water between the cytoplasm and the vacuolar sap. The vacuole is often enriched in plant specialized metabolites (PSMs), formerly also called secondary metabolites. PSMs comprise > 200,000 compounds and, despite this huge chemical diversity, they can be structurally divided into three major groups: alkaloids, terpenoids and phenolic compounds, and some smaller classes such as cyanogenic glucosides, glucosinolates and betalains. PSMs are often accumulated at high levels in specific tissues or even specialized cells. For example, flavonoids accumulate in flowers and fruits, and contribute to the development of the diverse range of colors that we observe in nature. In many plants, saponins are synthesized and stored in underground organs such as the glycyrrhizin that accumulates in licorice (Glycyrrhiza glabra) roots (Moses et al. 2014) or the saponins of oats (Avena sativa; Kesselmeier and Urban 1983). In some cases, PSMs are translocated from the source to the sink organ, as is for example the case of cyanogenic glucosides in cassava (Manihot esculenta) that are synthesized in leaves and transported to the roots, or in bitter almond (Prunus dulcis) where they accumulate in fruits (Gleadow and Møller 2014). In contrast, Coptis japonica produces its alkaloid berberine in roots and it is subsequently translocated to the rhizome. Similarly, in tobacco (Nicotiana tabacum), nicotine is produced in roots and accumulates in leaves (for a review, see Shitan et al. 2014; Shitan 2016). In other circumstances, the synthesis of PSM intermediates and PSMs may occur in different cells. This is observed, for example, in the medical plant Madagascar periwinkle (Catharanthus roseus) where the synthesis of the terpenoid indole alkaloids (TIAs) has a sophisticated and complex spatial organization (for a review, see Courdavault et al. 2014). The vacuole is often considered the end-storage cellular compartment of many PSMs. However, vacuoles can also store inactive pools of PSMs, which when needed are secreted from the vacuole and converted into active compounds. For example, soybean (Glycine max) produces glucosylated isoflavones that are stored in the vacuoles of roots. When needed, these compounds are recruited from the vacuole by unknown mechanism(s) and are excreted to the cell wall, where the presence of a specific G. max β-glucosidase converts these compounds into their active aglycone form serving as chemoattractants for nitrogen-fixing bacteria (Suzuki et al. 2006). In the last years, several reviews with detailed information on the vacuolar transport mechanisms of primary and specialized metabolites have been published (Yazaki et al. 2008, Shitan and Yazaki 2013, Shitan 2016), focused in particular on the transport of flavonoids and alkaloids (Zhao and Dixon 2009, Petrussa et al. 2013, Shitan et al. 2014, Zhao 2015). This review will present the state of the art of the most important PSMs that accumulate in the vacuole. An overview of the most recent findings on the transporter-mediated mechanism responsible for the vacuolar accumulation of PSMs is also presented. We therefore update the library of the so far characterized vacuolar PSM transporters and also indicate a new direction of research on the vacuolar transport of PSMs. The Vacuolar Potpourri of Plant Specialized Metabolites: A Resource to Be Explored Phenylpropanoids Certain plant species accumulate lignin precursors, monolignol glucosides, such as coniferin and syringin in the vacuole. This led to the hypothesis that vacuoles might act both as a temporary and as a final store of monolignols, depending on the metabolic stage of the plant or of the tissue (Whetten and Sederoff 1995). There has been great debate on the transport of lignin precursors. On the one hand it was shown that monolignol glucosides are transported into the vacuole by an ATP-binding cassette- (ABC) type specific transport mechanism (Miao and Liu 2010). It was suggested that the glucosylation of monolignols is a prerequisite for their selective import (Miao and Liu 2010). On the other hand, when woody plants were analyzed, the transport across the tonoplast was shown to be mainly driven by a proton (H+) gradient (Tsuyama et al. 2013). The apparent controversial results may reflect the differences in the analyzed plant tissues. In the first case, the isolated vacuolar membrane vesicles were prepared from Arabidopsis (Arabidopsis thaliana) leaves, whereas in the second case lignifying tissues such as of hybrid poplar (Populus sieboldii×Populus grandidentata) and pine (Pinus densiflora) were used. Furthermore, Arabidopsis vesicles are mainly derived from mesophyll cells where lignification is restricted to a number of cells at the leaf vein (Miao and Liu 2010). Despite the controversy, it was shown that Arabidopsis leaf vacuoles accumulate glucosylated lignin oligomers, and it was suggested that upon pathogen attack specific apoplastic β-glucosidases can mobilize these compounds to function as a rapid defense response (Dima et al. 2015). It is essential to identify the tonoplastic transporters involved in such processes and to determine whether the differences observed in the transport mechanisms are tissue or plant species dependent. Members of the Brassicaceae such as Arabidopsis or oilseed rape (Brassica napus) accumulate high levels of sinapate esters, namely sinapoyl malate in the leaves and sinapoyl choline (sinapine) in the seeds (Fraser and Chapple 2011). Sinapoyl malate has been suggested to act as a UV protectant in Arabidopsis (Fraser and Chapple 2011). Since the enzyme responsible for the conversion of sinapoyl glucose to sinapoyl malate, sinapoyl glucose:malate sinapoyltransferase, is localized in the vacuolar sap (Hause et al. 2002), sinapoyl glucose is likely to be transported into the vacuoles by an unknown mechanism. We can speculate that in Arabidopsis, similarly to monolignol glucosides (Miao and Liu, 2010), the mechanism might be mediated by ABC-like transporters. The subcellular localization of sinapine, the choline ester of sinapic acid, has not been elucidated so far, although it has been speculated that it may accumulate in vacuoles similarly to sinapoyl malate (Milkowski and Strack 2010). Sinapine accumulates up to 4% of the seed dry mass, and it has an important role during seed development as it may serve as the seed storage form of choline for the subsequent synthesis of phospholipids in developing seedlings (Bi et al. 2017). It also greatly contributes to anti-nutritive features, such as bitter and unpleasant flavor, for example in oilseed rape (Milkowski and Strack 2010). Various attempts have been made to produce plants with low sinapine content seeds. The majority of these studies focused on the inhibition of biosynthetic pathways (Milkowski and Strack 2010, Mittasch et al. 2013, Hettwer et al. 2016). However, it was shown that reprogramming sinapine metabolism in most cases produced modest reductions of these compounds (Mittasch et al. 2013) or induced strong metabolic modifications in late stages of seed development (Hettwer et al. 2016). From a biotechnological perspective, targeting transporters that are involved in such processes has the potential to offer new ways to control the accumulation of sinapate esters in a tissue-specific manner without compromising the whole-plant metabolism. Phenylpropanoids are also present in floral volatile organic compounds (Muhlemann et al. 2014). Volatile phenylpropanoids often undergo chemical modifications such as glycosylation. Recently, it was shown that in petal cells of Petunia×hybrida phenylpropanoid scent compounds are stored as glucosides in the vacuoles (Cna’ani et al. 2017). These authors propose that the glucosylated pool of flower scents and its vacuolar accumulation is dynamic and their recruitment from the vacuole occurs during peak scent emission. However, while it has been shown that a plasma membrane-localized ABC transporter is responsible for the release of the volatiles (Adebesin et al. 2017), it is unknown how floral scent compounds are imported and exported from the vacuole. Flavonoids: anthocyanins and proanthocyanins More than 600 anthocyanins have been identified in nature (He and Giusti 2010) and, despite this chemical diversity, the pathways involved in anthocyanin biosynthesis and their regulation are highly conserved in plants (Koes et al. 2005, Zhang et al. 2014). In this section, we will focus on some of the plant research fields where transport engineering may have a great impact in the accumulation of anthocyanins. Anthocyanin transport, from the site of synthesis to the site of storage, the vacuole, is a crucial process in anthocyanin metabolism. While several models for vacuolar sequestration of anthocyanins have been proposed (for a review, see Zhao 2015), we have still a limited knowledge of the transporters that are involved in such events. In the last years, several efforts have been made to enhance the anthocyanin content in fruit or to genetically engineer flower colors (Zhang et al. 2014). The anthocyanin-enriched tomato (Solanum lycopersicum) was built by combining the expression of R2R3MYB protein (Rosea 1) and bHLH protein (Delila) from snapdragon (Antirrhinum majus) (Butelli et al. 2008). Not only were biosynthetic and side chain modification genes in the anthocyanin pathway up-regulated, but also the expression of a likely vacuolar transporter of anthocyanin was induced (PAT/SlMTP77; Mathews et al. 2003). In a previous study, the overexpression of the ANTHOCYANIN MUTANT, which encodes a MYB transcription factor regulating anthocyanin production in tomato, also induced SlMTP77 (Mathews et al. 2003). Although tomato accumulates only small amounts of flavonoids in their peel and most cultivars do not produce anthocyanins in the fruit (Butelli et al. 2008), the previous results make SlMTP77 a good candidate for the vacuolar transport of flavonoids. Novel flower colors, particularly true-blue color in ornamental flowers such as rose, carnation or chrysanthemum, has been the aim of many researchers for many years. Designing of such flowers is far from being an easy task (Tanaka et al. 2010, Noda et al. 2017). In ornamental flowers, to date no vacuolar transporter has been directly implicated in the vacuolar accumulation of anthocyanins. From a biotechnological perspective, targeting such transporters might allow the design of novel strategies for improving or altering the levels and types of anthocyanins present in such flowers. It is known that different classes of anthocyanin transporters have substrate preferences according to their side chain modification. For example, in grapevine (Vitis vinifera), acylated and glucosylated anthocyanins are transported by different classes of proteins (Gomez et al. 2009, Francisco et al. 2013). Proanthocyanidins (PAs) or condensed tannins are polymers of flavan-3-ol units and are found in seed coats, leaves, fruit, flowers and bark in a variety of plant species (Dixon et al. 2005). In Arabidopsis, PAs are only produced in seeds, and the lack of PA pigmentation causes the transparent testa (tt) phenotype (Debeaujon et al. 2001). Apart from the transporters that are known to be involved in PA accumulation and that will be discussed later on in this review, other vacuolar proteins were shown to be indirectly involved in this process. AHA10 is a P-type H+-ATPase known to be involved in vacuolar PA accumulation in the seed coat (Baxter et al. 2005). Recently it was shown that AHA10 and TT13 are allelic (Appelhagen et al. 2015). TT13 was proposed to function as a proton pump that generates the driving force for TT12-mediated transport of PA precursors to the vacuole (Appelhagen et al. 2015). Another class of transporters that might also be involved in the transport of PAs into the vacuole are ABC transporters. Fu et al. (2017) characterized gene expression and flavonoid accumulation during the development of barrel medic (Medicago truncatula) seed coat. In macrosclereid cells, which are seed coat cells particularly enriched in flavonoids, several ABC transporters were involved in such a process. In particular, MRP14 (Medtr1g099280.1) is up-regulated at later stages of seed development. Still the exact mechanism of action of this transporter is unknown, and further biochemical studies need to be carried out. Saponins Saponins are a complex and chemically diverse group of triterpenoid or steroidal aglycones that are bound to oligosaccharides and have an important ecological role in plant defense against pathogens and herbivores (Sawai and Saito 2011, Moses et al. 2014). Despite their vacuolar localization (Mylona et al. 2008), so far no vacuolar saponin transporter has been identified. In a recent study, several transcriptionally active ABC-type transporters were nominated as candidates for the vacuolar storage of the saponin glycyrrhizin in licorice roots (Ramilowski et al. 2013). To investigate the possible co-regulation of these putative transporters with key enzymes of glycyrrhizin biosynthesis, namely β-amyrin synthase (bAS) and the Cyt P450 enzymes (CYP88D6 and CYP72A154), an unsupervised hierarchical clustering analysis was performed (Ramilowski et al. 2013), suggesting that a partial co-regulation exists. In a similar approach, RNA sequencing data from the medicinal plant Trillium govanianum identified several key genes involved in the complex steroidal saponin pathway, including ABC-type and multidrug and toxic compound extrusion (MATE) transporters (Singh et al. 2017). To unveil the mechanism of action of these transporters on the translocation of saponins into the vacuole, new studies need to be carried out. The genes involved in the synthesis of the saponin avenacins produced by oat are organized in a gene cluster. Interestingly, the final product is synthesized within the vacuole by the conjugation of des-acyl-avenacin A and N-methyl anthraniloyl-O-glucose. The fact that the transporters for both compounds are so far unknown indicates that in contrast to what was observed for the biosynthesis genes (Mugford et al. 2013), the transporter genes are not organized in a gene cluster. Glucosinolates In contrast to the major classes of PSMs, glucosinolates constitute a small yet diverse group of nitrogen- and sulfur-rich specialized metabolites that are mostly restricted to species of the Brassicales order (Grubb and Abel 2006, Halkier and Gershenzon 2006). Glucosinolate derivatives have an important role in plant defense against herbivores and pathogens, but also contribute, for example, to the flavor and aroma of cruciferous vegetables (Grubb and Abel 2006, Halkier and Gershenzon 2006). Although glucosinolates are non-toxic, highly toxic thiocyanates and related compounds are released when a cell is destroyed and glucosinolates come into contact with plant-specific hydrolases such as myrosinases (Grubb and Abel 2006, Halkier and Gershenzon 2006). This is a classic example of a plant defense binary system where cellular compartmentalization is essential to prevent the accumulation of deleterious, toxic compounds and to ensure that the plant response is released only upon pathogen attack. Arabidopsis accumulates glucosinolates in the vacuoles of the epidermis leaf margin side (Madsen et al. 2014) and in the vacuoles of the root cortex cells (Andersen et al. 2013). It is known that glucosinolates are transported between neighboring cells (Andersen et al. 2013, Madsen et al. 2014) or distant organs (Chen et al. 2001, Nour-Eldin et al. 2012) (for a review, see Jørgensen et al. 2015). In the proposed model for glucosinolates distribution in planta two plasma membrane-localized transporters from the nitrate/peptide (NTR/PTR) transporter family, AtGTR1 and AtGTR2, were shown to be involved in intra- and interglucosinolate leaf distribution and to be essential for glucosinolate accumulation in seeds (Nour-Eldin et al. 2012). The model also suggests the existence of tonoplastic transporters, importers and exporters that are responsible for storage and remobilization of glucosinolates, respectively (Jørgensen et al. 2015). However, so far the nature of these transporters is unknown. Cyanogenic glucosides Cyanogenic glucosides are PSMs widely distributed in plants. They are effective toxic compounds against herbivores, but may also exert a diversity of other biological functions (for reviews, see Gleadow and Møller 2014, Nielsen et al. 2016). Crops such as sorghum (Sorghum bicolor), cassava or almond are examples of cyanogenic plants (Gleadow and Møller 2014). This class of compounds are glucosides of amino acid-derived α-hydroxynitriles and, similarly to glucosinolates, are part of a plant’s defense binary system. Upon tissue disruption, the stored cyanogenic glucosides form, by the action of an endogenous β-glucosidases, an unstable compound that dissociates into hydrogen cyanide (Gleadow and Møller 2014). To avoid the release of hydrogen cyanide from intact tissue, cyanogenic glucosides and their degrading enzymes are stored in different cell compartments. The plant species S. bicolor accumulates dhurrin in the vacuoles (Saunders and Conn 1978) whereas dhurrinase is located in the chloroplasts (Thayer and Conn 1981). Genomic gene clusters for the biosynthesis of cyanogenic glucosides have been reported in several plant species including S. bicolor (Takos et al. 2011, Nützmann and Osbourn 2014). In a recent study, a tonoplastic transporter was identified in the cyanogenic glucoside dhurrin gene cluster (Darbani et al. 2016). Future research may help in understanding whether additional transporters are present in new gene clusters for cyanogenic glucosides relevant in cyanogenic crops. Recapitulation of Plant Vacuolar Transporters Several plant model systems of PSM metabolism and transport have emerged in the last years. Nicotiana tabacum and C. japonica can be considered plant models in the field of alkaloids; C. roseus for the study of terpenoid indole alkaloids; A. thaliana, V. vinifera and M. truncatula in the study of flavonoids; plant species from the Brassicales order for the investigation of glucosinolates; and Sorghum bicolor and Manihot esculenta for cyanogenic glucosides. To date, the vacuolar PSM transporters can be grouped into three families, the ABC transporters, the MATE and the nitrate/peptide family transporters (NPFs). So far the number of functionally characterized transporters remains limited; only one vacuolar ABC transporter, 10 vacuolar MATEs and one vacuolar NPF were unequivocally shown to be implicated in the transport of PSMs into/from the vacuole (Fig. 1). Fig. 1 View largeDownload slide Overview of the vacuolar transportome of plant specialized metabolites. Schematic representation of the vacuolar transporters of specialized metabolites in (A) Arabidopsis, (B) barrel medic, (C) tobacco, (D) Madagascar periwinkle, (E) Coptis japonica, (F) Sorghum and (G) grapevine. Only transporters that are functionally characterized are indicated. For Arabidopsis, we also include the proposed transport-mediated mechanism of flavonoids (Buer et al. 2007) and lignin precursors (Miao and Liu 2010). For Catharanthus roseus, it is also shown that terpenoid indole alkaloid end-products are actively taken up by a specific H+ antiport system into the vacuoles (Carqueijeiro et al. 2013). For tobacco, the induction of NtJAT1, NtJAT2, NtMATE1 and NtMATE2 by methyl jasmonate (MeJA) is indicated. Vac, vacuole. Fig. 1 View largeDownload slide Overview of the vacuolar transportome of plant specialized metabolites. Schematic representation of the vacuolar transporters of specialized metabolites in (A) Arabidopsis, (B) barrel medic, (C) tobacco, (D) Madagascar periwinkle, (E) Coptis japonica, (F) Sorghum and (G) grapevine. Only transporters that are functionally characterized are indicated. For Arabidopsis, we also include the proposed transport-mediated mechanism of flavonoids (Buer et al. 2007) and lignin precursors (Miao and Liu 2010). For Catharanthus roseus, it is also shown that terpenoid indole alkaloid end-products are actively taken up by a specific H+ antiport system into the vacuoles (Carqueijeiro et al. 2013). For tobacco, the induction of NtJAT1, NtJAT2, NtMATE1 and NtMATE2 by methyl jasmonate (MeJA) is indicated. Vac, vacuole. Vacuolar transporters of alkaloids The vacuolar accumulation of alkaloids has been intensively studied, in particular in N. tabacum. So far this is the plant species where more vacuolar PSM transporters were characterized; four up to now. NtJAT1, NtJAT2, NtMATE1 and NtMATE2 all belonging to the MATE family are involved in nicotine transport into the vacuole (Morita et al. 2009, Shoji et al. 2009, Shitan et al. 2014). Shitan et al. (2014) and Shitan (2016) recently reviewed the state of the art of alkaloid transporters and also discussed the physiological role of alkaloids in planta. Coptis japonica is a medical plant grown in Asia that accumulates high amounts of the alkaloid berberine in the vacuole of rhizomes (Shitan et al. 2014). In C. japonica, a proton gradient-driven transporter(s) was proposed to transport berberine across the tonoplast (Otani et al. 2005). Takanashi et al. (2017) recently isolated CjMATE1 from cultured C. japonica cells. CjMATE1 is a tonoplast-localized MATE transporter that is preferentially expressed in rhizomes. When heterologously expressed in Saccharomyces cerevisiae, the berberine content was higher in yeast cells expressing CjMATE1 than in the control cells harboring the empty vector. These results strongly suggest that CjMATE1 acts as a vacuolar berberine uptake transporter. Catharanthus roseus has an elaborated metabolism producing >100 TIAs. In plants, these low abundant alkaloids provide a wide range of protection against pathogens and predators. Additionally, some of these compounds such as the bisindole alkaloids vinblastine and vincristine exhibit strong pharmacological activities such as anticancer properties (van der Heijden et al. 2004). The biosynthesis of TIAs involves complex multicellular compartmentation that comprises the phloem-associated parenchyma, the epidermis, the mesophyll, laticifers and idioblasts (for a review, see Courdavault et al. 2014). Despite the fact that some TIAs are accumulated in the vacuole and exported to the cytosol, only recently was a transporter that belongs to the NPF family identified as an exporter of the TIA precursor strictosidine (Payne et al. 2017). In this study, CrNPF2.9 was selected from transcriptomic expression data where TIA biosynthetic pathways genes and CrNPF2.9 were co-regulated. With virus-induced gene silencing, CrNPF2.9 loss of function in planta was observed. This approach increased the levels of strictosidine in the leaves together with a significant decrease of the end-products of the TIA pathway, vindoline and catharanthine. On the other hand, the heterologous expression of CrNPF2.9 in Xenopus laevis oocytes suggests that CrNPF2.9 is a high-affinity exporter of strictosidine out of the vacuole (Payne et al. 2017). The enzyme responsible for strictosidine synthesis, strictosidine synthase, is known to be vacuolar (Guirimand et al. 2010). This suggests that the precursors of strictosidine, tryptamine and secologanin must also be transported from the cytosol into the vacuole for strictosidine biosynthesis to occur; however, no transporter(s) have been identified yet. Vacuolar transporters of flavonoids To date, this is the class of PSMs where more tonoplastic transporters have been identified. In grapevine, VvAM1, VvAM3 and VvABCC1 are involved in vacuolar accumulation of acylated and glucosylated anthocyanins, respectively (Gomez et al. 2009, Francisco et al. 2013). VvMATE1 is also assigned as a putative PA transporter expressed during grapevine seed development (Pérez-Díaz et al. 2014). Medicago truncatula MATE1 is involved in flavonoid transport (Zhao and Dixon 2009), whereas MtMATE2 is involved in anthocyanin transport (Zhao et al. 2011). Genetic studies in maize provided evidence that ZmMRP3 acts as an anthocyanin transporter (Goodman et al. 2004). In Arabidopsis, two MATE transporters have been described to be involved in the compartmentalization of flavonoids into the vacuole. AtTT12 acts as a PA/H+-antiporter (Debeaujon et al. 2001, Marinova et al. 2007, Zhao and Dixon 2009), whereas FFT/DTX35 was characterized as a putative flavonoid transporter (Thompson et al. 2010). More recently, FFT/DTX35 was found to function in seed coats of nthe mutant banylus (ban) as an anthocyanin transporter (Kitamura et al. 2016). Since the immature seeds of the ban mutant accumulate anthocyanins instead of PAs, the authors screened for pigmentation phenotypes in a mutagenized population. The pale ban (pab1) seeds had reduced levels of cyanidin-3-O-glucoside (C3G) compared with the parental ban immature seeds, and no significant difference in the expression of anthocyanin biosynthetic genes was observed, although previous studies showed a feedback loop mechanism between biosynthesis and vacuolar transport (Goodman et al. 2004, Zhao and Dixon 2009). Furthermore, the levels of anthocyanins in young seedlings remained unchanged in the pab1ban mutant (Kitamura et al. 2016). All together, these results suggest that the true nature of FFT/DTX35 substrates remains to be elucidated since Arabidopsis seed coat does not contain C3G (Saito et al. 2013). Interestingly, FFT/DTX35 was also found to function in root hair and pollen tube elongation as a chloride channel essential for turgor regulation in Arabidopsis (Zhang et al. 2017). It remains to be clarified whether the FFT/DTX35 transporter exhibits multisubstrate specificity or whether changes in the membrane potential between the cytosol and the vacuole affect anthocyanin allocation. Functional homologs of AtTT12, MdMATE1 and MdMATE2, were identified in apple fruit (Malus domestica; Frank et al. 2011). In this study, complementation of the A. thaliana tt12-1 mutant could restore PA deposition in the seeds. Despite the fact that no biochemical data were presented, the results suggest that MdMATE1 and MdMATE2 are vacuolar flavonoid/H+-antiporters active in PA accumulation in apple fruit. The MdMYB1 transcription factor that regulates anthocyanin biosynthesis in red apples (Takos et al. 2006) was recently shown to regulate the expression of several genes including MdMATE-LIKE1 and a vacuolar MdABCB-LIKE27 transporter (Hu et al. 2016). It was also shown that AtPAP1, the homolog of apple MdMYB1, functions similarly to MdMYB1, binding to AtMATE (DTX35) and AtABCB27 (TAP2/ALS1) promoters, thereby directly mediating the transcriptional activation of these genes (Hu et al. 2016). AtABCB27 was described as a transporter involved in aluminum sequestration (Larsen et al. 2007). In many plant species, color is the association of anthocyanins with co-pigments namely flavonols and/or metals forming the so-called metalloanthocyanins (for a review, see Yoshida et al. 2009). The relationship between metal transporters and anthocyanin is intriguing. For example, in tulips (Tulipa gesneriana), the blue color was associated with the expression of the iron transporter gene TgVit1 (Momonoi et al. 2009). Despite the fact that TgVit1 is regulated differently compared with the structural anthocyanin biosynthesis genes, its expression is exclusive in blue-colored epidermal cells. In the Japanese morning glory (Ipomoea nil) where the vacuolar pH regulation has a great impact in flower color (Yoshida et al. 2005), the expression of the vacuolar Na+/H+ exchanger InNHX1 is associated with the colored cells of flowers. NHX1 mediates the vacuolar alkalinization, allowing the color change from pink to the heavenly blue anthocyanin pigment present in this species, likely to attract pollinators (Yamaguchi et al. 2001, Yoshida et al. 2005). Similarly, in petunia (Petunia hybrida), vacuolar acidification determines flower color (Faraco et al. 2014). Overall what all these studies indicate is that flower and fruit color mechanisms may activate vacuolar transporters (for uptake of flavonoids, metals or ions into the vacuoles) that are co-ordinated by the same transcriptions factors such as MdMYB1 (Hu et al. 2016) or PH4 (Quattrocchio et al. 2006). PH4 is a MYB transcription factor that was shown in petunia to interact physically with other transcription factors responsible for the transcriptional activation of a subset of structural anthocyanin genes (Quattrocchio et al. 2006). Interestingly, although in petunia our knowledge of the genetics and biochemistry of anthocyanin is well documented (Mol et al. 1998), we do not yet know much about the mechanisms and transporters behind the vacuolar storage of anthocyanins in this plant species. Sorghum bicolor vacuolar transporter of cyanogenic glucosides So far only one transporter was identified to mediate the vacuolar accumulation of cyanogenic glucosides. Darbani et al. (2016) identified SbMATE2 in the gene cluster for dhurrin biosynthesis. SbMATE2 was co-expressed with SbCYP79A1, encoding the first enzyme of the dhurrin biosynthetic pathway. SbMATE2 is tonoplast localized, as shown by its transient expression in N. benthamiana. The functional analysis of SbMATE2 was conducted in X. laevis oocytes where SbMATE2-expressing oocytes presented reduced dhurrin content in comparison with control oocytes. Interestingly, SbMATE2 was also able to transport structurally related aromatic cyanogenic glucosides such as prunasin and the diglucoside amygdalin, which are present in plant species such as those of the genus Prunus. Despite the fact that cyanogenic glucosides and glucosinolates are structurally related defense compounds, SbMATE2 shows no transport activity for indol-3-yl-methyl (Darbani et al. 2016). From Phylogeny to Function Genome and transcriptome data mining, gene clustering and phylogeny analysis allowed significant advances in the identification of vacuolar PSM transporters. However, we still have a long road in front of us. As we have already stated, the number of vacuolar PSM transporters that are characterized is limited. Phylogenetic analysis is a useful tool to select new candidates and, although with limitations, it can also assist in predicting the class of substrate–transporter affinities. We collected 60 MATE sequences which were vacuolar or putatively vacuolar localized, of known or as yet unknown function, and we performed a phylogenetic study (Fig. 2A). These proteins grouped into two clades: clades A and B. The first one can be divided into five subclades that we roughly classified as: subclade I (alkaloids and anthocyanins), subclade II (alkaloids), subclade III (proanthocyanidins) and subclade V (alkaloids and cyanogenic glucosides). Subclade IV is composed of four members all of unknown functions. Clade B (alkaloids) contains 15 members but so far only NtJAT1 has been functionally characterized (Morita et al. 2009). Altogether, this analysis suggests that vacuolar MATE transporters differentiated into transporters for different classes of specialized metabolites. For example, the characterized proteins of subclade III are clearly associated with PA metabolism. Thus, Solyc12g006360.2 and Peaxi162Scf00047g02333.1 are good candidates to be selected for functional studies on the accumulation of PAs or related compounds in tomato and petunia. Another point suggested by this phylogenetic analysis is the function of NtJAT2 sequence homologs: are they related to alkaloid transport or do they also take up other classes of substrates? It is tempting to speculate that the clustering of these proteins is related to their physiological role in planta. Still, different specialized metabolites can have identical physiological/ecological functions (Pichersky and Lewinsohn 2011). To provide the readers with an insight into tissue or development stage specificity of the vacuolar MATE transporters, we retrieved from Genenvestigator (Hruz et al. 2008) and from TomExpress (Zouine et al. 2017) gene expression data from Arabidopsis (see Supplementary Fig. S1) and tomato (see Supplementary Fig. S2). Tissue localization and physiological studies on these transporters are imperative for a full understanding of their role in planta. Fig. 2 View largeDownload slide Phylogenetic trees of vacuolar MATE and ABC transporters. The amino acid sequences of vacuolar (A) multidrug and toxic compound extrusion (MATE) and (B) ATP-binding cassette (ABC)-type C transporters were aligned with MUSCLE and subjected to phylogenetic analysis conducted in MEGA7 (Kumar et al. 2016) using the Maximum Likelihood method with 1,000 bootstraps. Clade numbers were arbitrarily assigned. Tomato (Solanum lycopersicum) and Petunia axilaris amino acid sequences were retrieved from the Solanaceae genomics database (https://solgenomics.net): Solyc01g094830, Solyc02g032660, Solyc02g080480, Solyc02g080490, Solyc03g025200, Solyc03g025230, Solyc03g118970, Solyc04g007540, Solyc04g009790, Solyc05g013450, Solyc05g013460, Solyc05g013470, Solyc06g036130, Solyc07g006730, Solyc07g006740, Solyc07g052380, Solyc10g007100, Solyc12g005850, Solyc12g006360, Solyc12g019320, Solyc00g283010.1, Solyc01g080640.2, Solyc03g007530.2, Solyc03g117540.2, Solyc05g014380.2, Solyc06g036490.1, Solyc07g065320.2, Solyc08g006880.2, Solyc08g081890.2, Solyc09g064440.2, Solyc09g075020.2, Solyc10g019270.1, Solyc10g024420.1, Solyc11g065710.1, Solyc12g036140.1, Solyc12g044820.1, Peaxi162Scf00007g00932.1, Peaxi162Scf00047g02333.1, Peaxi162Scf00057g00323.1, Peaxi162Scf00128g10012.1, Peaxi162Scf00262g01014.1, Peaxi162Scf00262g01015.1, Peaxi162Scf00565g00015.1, Peaxi162Scf00683g00247.1, Peaxi162Scf00692g00414.1, Peaxi162Scf00897g00113.1, Peaxi162Scf00932g00522.1, Peaxi162Scf00997g00111.1, Peaxi162Scf01340g00216.1, Peaxi162Scf00001g04810.1, Peaxi162Scf00043g00247.1, Peaxi162Scf00045g02134.1, Peaxi162Scf00061g00226.1, Peaxi162Scf00061g00228.1, Peaxi162Scf00074g01312.1, Peaxi162Scf00128g00825.1, Peaxi162Scf00177g00226.1, Peaxi162Scf00217g00622.1, Peaxi162Scf00217g00624.1, Peaxi162Scf00222g01019.1, Peaxi162Scf00285g00821.1, Peaxi162Scf00553g00540.1, Peaxi162Scf00618g00535.1, Peaxi162Scf00901g00110.1, Peaxi162Scf00904g00463.1, Peaxi162Scf01123g00025.1, Peaxi162Scf01123g00232.1. The barrel medic sequence was obtained from the Medicago truncatula (Mt) genome database (http://www.medicagogenome.org/). Sequence data from Arabidopsis thaliana (At), Catharanthus roseous (Cr), Coptis japonica (Cj), Glycine max (Gm), Malus domestica (Md), Medicago truncatula (Mt), Nicotiana tabacum (Nt), Sorghum bicolor (Sb), Vitis vinifera (Vv) and Zea mays (Zm) can be found under the following accession numbers: AtDTX35, NP_194294; AtDTX41/TT12, NP_191462; AtDTX16, NP_200058; AtDTX17, NP_177511; AtDX19/ALF5, NP_566730; AtDTX29, NP_189291; AtDTX30, NP_198619; AtDTX33, NP_175184; AtDTX40, NP_188806; CjMATE1, BAX73926; CrMATE1, AQM73450; CrMATE2, AQM73451; GmMATE10, KRH35833; MdMATE1, ADO22712; MdMATE2, NP_001280841; MdMATE-Like1, XP_008380016; MtMATE1, ACX37118; MtMATE2, ADV04045; NtJAT1, CAQ51477; NtJAT2, BAP40098; NtMATE1, BAF47751; NtMATE2, BAF47752; SlMTP77, NP_001234424; SbMATE2, XP_021303040; VvAM1, ACN88706; VvAM3, ACN91542; VvMATE1, XP_002282907; AtABCC1, NP_181013; AtABCC2, NP_001031116; AtABCC3, NP_187915; AtABCC4, NP_182301; AtABCC5, NP_171908; AtABCC6, NP_187916.3; AtABCC7, NP_187917; AtABCC8, Q8LGU1; AtABCC9, Q9M1C7; AtABCC10, NP_191473; AtABCC11, NP_174331; AtABCC12, Q9C8H0; AtABCC13, NP_001323940; AtABCC14, NP_191829; VvABCC1, AGC23330; and ZmMRP3, AAT37905. Fig. 2 View largeDownload slide Phylogenetic trees of vacuolar MATE and ABC transporters. The amino acid sequences of vacuolar (A) multidrug and toxic compound extrusion (MATE) and (B) ATP-binding cassette (ABC)-type C transporters were aligned with MUSCLE and subjected to phylogenetic analysis conducted in MEGA7 (Kumar et al. 2016) using the Maximum Likelihood method with 1,000 bootstraps. Clade numbers were arbitrarily assigned. Tomato (Solanum lycopersicum) and Petunia axilaris amino acid sequences were retrieved from the Solanaceae genomics database (https://solgenomics.net): Solyc01g094830, Solyc02g032660, Solyc02g080480, Solyc02g080490, Solyc03g025200, Solyc03g025230, Solyc03g118970, Solyc04g007540, Solyc04g009790, Solyc05g013450, Solyc05g013460, Solyc05g013470, Solyc06g036130, Solyc07g006730, Solyc07g006740, Solyc07g052380, Solyc10g007100, Solyc12g005850, Solyc12g006360, Solyc12g019320, Solyc00g283010.1, Solyc01g080640.2, Solyc03g007530.2, Solyc03g117540.2, Solyc05g014380.2, Solyc06g036490.1, Solyc07g065320.2, Solyc08g006880.2, Solyc08g081890.2, Solyc09g064440.2, Solyc09g075020.2, Solyc10g019270.1, Solyc10g024420.1, Solyc11g065710.1, Solyc12g036140.1, Solyc12g044820.1, Peaxi162Scf00007g00932.1, Peaxi162Scf00047g02333.1, Peaxi162Scf00057g00323.1, Peaxi162Scf00128g10012.1, Peaxi162Scf00262g01014.1, Peaxi162Scf00262g01015.1, Peaxi162Scf00565g00015.1, Peaxi162Scf00683g00247.1, Peaxi162Scf00692g00414.1, Peaxi162Scf00897g00113.1, Peaxi162Scf00932g00522.1, Peaxi162Scf00997g00111.1, Peaxi162Scf01340g00216.1, Peaxi162Scf00001g04810.1, Peaxi162Scf00043g00247.1, Peaxi162Scf00045g02134.1, Peaxi162Scf00061g00226.1, Peaxi162Scf00061g00228.1, Peaxi162Scf00074g01312.1, Peaxi162Scf00128g00825.1, Peaxi162Scf00177g00226.1, Peaxi162Scf00217g00622.1, Peaxi162Scf00217g00624.1, Peaxi162Scf00222g01019.1, Peaxi162Scf00285g00821.1, Peaxi162Scf00553g00540.1, Peaxi162Scf00618g00535.1, Peaxi162Scf00901g00110.1, Peaxi162Scf00904g00463.1, Peaxi162Scf01123g00025.1, Peaxi162Scf01123g00232.1. The barrel medic sequence was obtained from the Medicago truncatula (Mt) genome database (http://www.medicagogenome.org/). Sequence data from Arabidopsis thaliana (At), Catharanthus roseous (Cr), Coptis japonica (Cj), Glycine max (Gm), Malus domestica (Md), Medicago truncatula (Mt), Nicotiana tabacum (Nt), Sorghum bicolor (Sb), Vitis vinifera (Vv) and Zea mays (Zm) can be found under the following accession numbers: AtDTX35, NP_194294; AtDTX41/TT12, NP_191462; AtDTX16, NP_200058; AtDTX17, NP_177511; AtDX19/ALF5, NP_566730; AtDTX29, NP_189291; AtDTX30, NP_198619; AtDTX33, NP_175184; AtDTX40, NP_188806; CjMATE1, BAX73926; CrMATE1, AQM73450; CrMATE2, AQM73451; GmMATE10, KRH35833; MdMATE1, ADO22712; MdMATE2, NP_001280841; MdMATE-Like1, XP_008380016; MtMATE1, ACX37118; MtMATE2, ADV04045; NtJAT1, CAQ51477; NtJAT2, BAP40098; NtMATE1, BAF47751; NtMATE2, BAF47752; SlMTP77, NP_001234424; SbMATE2, XP_021303040; VvAM1, ACN88706; VvAM3, ACN91542; VvMATE1, XP_002282907; AtABCC1, NP_181013; AtABCC2, NP_001031116; AtABCC3, NP_187915; AtABCC4, NP_182301; AtABCC5, NP_171908; AtABCC6, NP_187916.3; AtABCC7, NP_187917; AtABCC8, Q8LGU1; AtABCC9, Q9M1C7; AtABCC10, NP_191473; AtABCC11, NP_174331; AtABCC12, Q9C8H0; AtABCC13, NP_001323940; AtABCC14, NP_191829; VvABCC1, AGC23330; and ZmMRP3, AAT37905. Many studies have shown that several ABC transporters recognize PSMs (for reviews, see Kang et al. 2011, Hwang et al. 2016). However, up to now the vacuolar-localized members that are functionally characterized are scarce. Genenvestigator (Hruz et al. 2008) and TomExpress (Zouine et al. 2017) platforms provide us with an overview of the expression of the vacuolar ABCCs in different plant tissues in Arabidopsis (see Supplementary Fig. S3) and tomato (see Supplementary Fig. S4). All Arabidopsis ABCC transporters predicted to be tonoplast localized (Jaquinod et al. 2007, Kang et al. 2011) were chosen to carry out this phylogenetic study. In total, we collected 50 sequences from several plant species, including Petunia axillaris and tomato (Fig. 2B). The maximum likelihood phylogenetic analysis revealed six clades. Based on the knowledge of the functions of the so far characterized AtABCC transporters (Kang et al. 2011), we suggest that clade I contains the best candidates involved in flavonoid transport. In plant species such as maize and grapevine, the involvement of ABCC transporters in the vacuolar accumulation of anthocyanins is known (Goodman et al. 2004, Francisco et al. 2013). Furthermore, the transport of flavonoids by ABCC transporters is likely also to occur in Arabidopsis (Buer et al. 2007). Conclusions and Future Outlook The vacuole is a dynamic cellular compartment that exerts many functions, including the accumulation of specialized metabolites. A comprehensive view of the vacuolar accumulation of specialized metabolites is hindered by an incomplete repertoire of transporters. Even so, significant progress in this field has been made in the past decade. To date, members of the ABC, MATE and NPF transporter families are functionally assigned as tonoplastic PSM transporters. As the vacuole temporarily or permanently stores PSMs, it must comprise importers and exporters. Nevertheless, the number of characterized vacuolar transporters with export activity is still very small. Strategies that help the identification of novel transporters have to include multidisciplinary approaches such as genome editing, bioinformatics, biochemistry and robust analytical methodologies. Many ABC and MATE transporters frequently show functional redundancy. Thus, structural information collected by crystallography studies could assist in accurately predictly the class of substrates of these transporters. However, so far, no plant vacuolar transporter has been crystallized. Recent developments in the fields of genomics and bioinformatics, where the number and quality of plant genome sequencing projects is rapidly increasing, could contribute to the study of non-model plant species such as medical or crop plants which are often enriched in PSMs. Lastly, an improved knowledge of plant transporters offers novel tools to develop successful strategies for effective metabolic engineering. We can exploit these tools to improve plant traits such as their resistance to biotic stresses or to improve the nutritional composition of crops. Supplementary Data Supplementary data are available at PCP online. Funding Results reported in this work were funded by the University of Zürich and the Swiss National Foundation (Switzerland) and Fundação para a Ciência e a Tecnologia of Ministério da Ciência, Tecnologia e do Ensino Superior (Portugal). 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Plant and Cell Physiology – Oxford University Press
Published: Feb 14, 2018
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