Phytochem Rev DOI 10.1007/s11101-017-9520-6
Aromatic amino acid aminotransferases in plants Minmin Wang . Hiroshi A. Maeda
Received: 2 March 2017 / Accepted: 20 June 2017 Ó Springer Science+Business Media B.V. 2017
Abstracts Aromatic amino acid aminotransferases (AAA-ATs) catalyze the reversible transamination reactions of proteinogenic and non-proteinogenic aromatic amino acids to corresponding keto acids and vice versa. The products of plant AAA-ATs serve as key precursors of many primary and secondary metabolites that are crucial for both plant and human metabolism and physiology. In most microbes, Ltyrosine (Tyr) and L-phenylalanine (Phe) aminotransferases (Tyr and Phe-ATs) catalyze the final steps of Phe and Tyr biosynthesis. On the other hand, plants use different pathways to synthesize Tyr and Phe via arogenate, in which prephenate-specific aminotransferases (PPA-ATs) catalyze the committed step in the plastids. Plant Tyr and Phe-ATs, unlike microbial counterparts, often prefer the reverse reactions and metabolize Tyr and Phe to their respective aromatic keto acids, which serve as precursors of various plant natural products (e.g. benzenoid volatiles, tocochromanols, plastoquinone, and tropane and benzylisoquinoline alkaloids). Unlike plastidic PPA-ATs, plant Tyr/Phe-ATs are localized outside of the plastids, have broad substrate specificity, and interlink Tyr and Phe Electronic supplementary material The online version of this article (doi:10.1007/s11101-017-9520-6) contains supplementary material, which is available to authorized users. M. Wang H. A. Maeda (&) Department of Botany, University of Wisconsin-Madison, Madison, WI, USA e-mail:
[email protected]
metabolism. L-Tryptophan (Trp) aminotransferases (Trp-ATs) are involved in biosynthesis of the plant hormone auxin. Although significant advancement has been made on biochemical, molecular, and genetic characterizations of plant AAA-ATs, there are still many critical knowledge gaps, which are highlighted in the current review. Keywords Aromatic amino acids Aminotransferase Transaminase Amino acid biosynthesis Plant natural products Auxin biosynthesis Abbreviations AAA Aromatic amino acid ADT Arogenate dehydratase ADH Arogenate dehydrogenase Asp L-Aspartate Asp-AT Aspartate aminotransferase BCAT Branch-chain amino acid aminotransferase L-DOPA 3,4-Dihydroxyphenylalanine Glu L-Glutamate HPP 4-Hydroxyphenylpyruvic acid HPPD HPP dioxygenase IAA Indole-3-acetic acid IPA Indole-3-pyruvate Met L-Methionine PDT Prephenate dehydratase PDH Prephenate dehydrogenase
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Phe Phe-AT PLP PPA-AT PPY Trp Trp-AT Tyr Tyr-AT VAS1
L-Phenylalanine
Phe aminotransferase Pyridoxal-50 -phosphate Prephenate aminotransferase Phenylpyruvic acid L-Tryptophan Trp aminotransferase L-Tyrosine Tyr aminotransferase REVERSAL OF SAV3 PHENOTYPE 1
Introduction Aromatic amino acid aminotransferases (AAA-ATs) refer to transaminases (EC 2.6.1, Fig. 1a) of proteinogenic aromatic amino acids (AAAs), L-phenylalanine (Phe), L-tyrosine (Tyr), and L-tryptophan (Trp) (Fig. 1b), as well as those of non-proteinogenic AAAs, such as L-3,4-dihydroxyphenylalanine (L-DOPA). This review also considers aminotransferases that utilize prephenate and arogenate, which are key precursors of Tyr and Phe biosynthesis (Fig. 1b). Although the official naming of transaminases should follow amino donor—keto acceptor aminotransferase (e.g. alanine—glyoxylate aminotransferase), different nomenclatures have been used for AAA-ATs in different studies. This review categorizes different AAA-AT types based on their preferred AAA substrate (e.g. Trp-ATs; Fig. 1b), while acknowledging gene/enzyme names that were used in original articles. Prephenate aminotransferase is abbreviated as PPAAT rather than PAT to avoid confusion with phosphoribosylanthranilate transferase (PAT) catalyzing the second step of Trp biosynthesis (Rose et al. 1992). In Escherichia coli and Saccharomyces cerevisiae, AAA-ATs (e.g. EcTyrB and ScARO8) are responsible for the final step of Phe and Tyr biosynthesis (Gelfand and Steinberg 1977; Urrestarazu et al. 1998). On the other hand, plants use different pathways to synthesize Phe and Tyr via arogenate (Jensen 1986; Tzin and Galili 2010; Maeda and Dudareva 2012), which is produced by prephenate aminotransferases (PPAATs) (Graindorge et al. 2010; Dal Cin et al. 2011; Maeda et al. 2011) (Fig. 1c). Plant AAA-ATs are instead often involved in the metabolism of AAAs to the plant hormone auxin and a diverse array of plant natural products, such as phenylpropanoids and
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Fig. 1 AAA-ATs involved in AAA biosynthesis and metabo- c lism in plants. a The overall mechanism of a PLP-dependent aminotransferase reaction. An amino group (pink highlight) of an amino acid substrate (e.g. Phe) is transferred to a PLP-bound enzyme (top), generating a PMP-bound enzyme (bottom) and releasing a keto acid product (e.g. PPY) in the first half reaction. In the second half reaction, a keto acid substrate (e.g. oxaloacetate) accepts the amino group to be converted to an amino acid product (e.g. Asp), while regenerating the PLP enzyme. b Amino acid and keto acid substrates of the major aminotransferases discussed in this study. As denoted in the box, black arrows indicate the deamination reactions of an amino acid substrate, whereas gray arrows are the transamination reactions of a keto acid substrate. Most aminotransferases catalyze reversible reactions but usually have a preferred reaction direction. c AAA-ATs catalyze key biochemical steps of AAA biosynthesis and metabolism. In plants, AAAs are mainly synthesized within the plastids from chorismate, the product of the shikimate pathway, though cytosolic pathways also exist for Phe and Tyr biosynthesis in some plants. AAAs are exported from the plastids and used for protein synthesis and also metabolized to diverse plant natural products and hormones (green arrows and letters). Dotted arrows denote multiple enzymatic steps. Preferred directions of AAA-AT catalyzed reactions are indicated by large arrowheads. ACC, 1-aminocyclopropane-1-carboxylic acid, ADT, arogenate dehydratase; ADH, arogenate dehydrogenase; Asp, L-aspartate; cCM, cytosolic chorismate mutase; pCM, plastidic chorismate mutase; L-DOPA, 3,4-dihydroxyphenylalanine; Glu, L-glutamate; HPP, 4-hydroxyphenylpyruvic acid; HPPD, HPP dioxygenase; IAA, indole-3-acetic acid; IPA, indole-3-pyruvate; aKG, a-ketoglutarate; Met, L-methionine; OAA, oxaloacetate; PDT, prephenate dehydratase; PDH, prephenate dehydrogenase; Phe, L-phenylalanine; Phe-AT, Phe aminotransferase; PLP, pyridoxal-50 -phosphate; PMP, pyridoxamine-50 -phosphate; PPA-AT, prephenate aminotransferase; PPY, phenylpyruvic acid; Trp, L-tryptophan; Trp-AT, Trp aminotransferase; Tyr, L-tyrosine; Tyr-AT, Tyr aminotransferase; SAM, S-adenosyl methionine; VAS1, REVERSAL OF SAV3 PHENOTYPE 1. (Color figure online)
alkaloids (Facchini 2001; Vogt 2010; Aniszewski 2015) (Fig. 1c). AAA-derived plant natural products play crucial roles for the fitness of various plant species, as antioxidants, attractants, and defense compounds (Rice-Evans et al. 1997; Maeda and DellaPenna 2007; Møller 2010; Falk and Munne´Bosch 2010). These compounds are also of industrial and pharmaceutical interest to human society as medicines (e.g. isoquinoline and tropane alkaloids), nutrients (e.g. tocochromanols collectively known as vitamin E), and natural pigments (e.g. betalains and anthocyanins) (Kutchan 1995; Bridle and Timberlake 1997; Robins 1998; Hunter and Cahoon 2007; Gandı´aHerrero and Garcı´a-Carmona 2013; Beaudoin and Facchini 2014; Aniszewski 2015) (Fig. 1c). Plant
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AAA-ATs have been investigated for more than fifty years since AAA-AT activity was first detected in a shoot extract of common bean (Phaseolus vulgaris) (Gamborg and Wetter 1963). Despite their importance in both plant primary and specialized metabolism, a large body of literature on plant AAA-ATs has not been reviewed previously. This article summarizes previous biochemical examinations of AAA-AT activity from various plant tissues as well as more recent biochemical, molecular, and genetic characterizations of plant AAA-AT enzymes, while highlighting remaining knowledge gaps on plant AAA-ATs.
Biochemical properties of plant AAA-AT activity and enzymes AAA-ATs are pyridoxal-50 -phosphate (PLP)-dependent enzymes (Mehta et al. 1993; Jensen and Gu 1996), which catalyze transamination reactions between an amino acid donor and a keto acid acceptor substrates (Fig. 1a) utilizing the ‘‘ping-pong bi-bi’’ reaction mechanism (Cleland 1963; John 1995; Toney 2014). In the first half reaction, an amino acid substrate (e.g. Phe) reacts with a PLP-bound enzyme to generate a pyridoxamine-50 -phosphate (PMP) enzyme and release a keto acid product (e.g. phenylpyruvate, PPY, Fig. 1a). In the second half reaction, a keto acid substrate (e.g. oxaloacetate) reacts with the PMP enzyme, regenerating the PLP enzyme and releasing an amino acid product (e.g. Asp, Fig. 1a). The transamination activity of plant AAA-AT is abolished when PLP is depleted (Koshiba et al. 1993; Simpson et al. 1997) or in the presence of carbidopa, an inhibitor of PLP-dependent enzymes (Hirata et al. 2012). Crystal structures of microbial and mammalian AAA-ATs have been obtained, including Tyr-ATs of Trypanosoma cruzi, mouse, and human (Blankenfeldt et al. 1999; Karlberg et al. 2008; Mehere et al. 2010) as well as kynurenine aminotransferases of human, mouse and yeast (Rossi et al. 2004, 2008; Wogulis et al. 2008; Han et al. 2009, 2011). Overall structures of these AAA-ATs are similar with one another and also to other class I aminotransferases (e.g. Asp-ATs) (Blankenfeldt et al. 1999; Ko et al. 1999). AAA-ATs form homodimers and each monomer subunit consists of a small domain and a large domain, with its active site located between the two domains and at the subunit interface. Substrate binding induces
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movement of the small domain from the open form to close the active site (Blankenfeldt et al. 1999; Ko et al. 1999; Kamitori et al. 1990; Jensen and Gu 1996). The enzyme-PLP Schiff base is formed at the lysine residue in the active sites (Fig. 1a), like in other PLPdependent enzymes (John 1995; Toney 2005). These structural analyses combined with site-directed mutagenesis further identified six active site residues that can introduce AAA-AT activity in E. coli Asp-AT (i.e. AspC) (Onuffer and Kirsch 1995). In contrast, limited information is available for structures of plant AAAATs and for the role of active site residues, especially ones involved in amino and keto acid substrate specificities. Prephenate aminotransferase (PPA-AT, EC 2.6.1.78 or 79) In most microbes, the Phe and Tyr biosynthetic pathways diverge at prephenate, which is converted by prephenate dehydratase and dehydrogenase (PDT and PDH) to (PPY) and 4-hydroxyphenylpyruvate) (Cotton and Gibson 1965; Davidson and Hudson 1987; Mannhaupt et al. 1989; Euverink et al. 1995; Zhang et al. 1998) followed by transamination to Phe and Tyr, respectively (Gelfand and Steinberg 1977; Urrestarazu et al. 1998; Bentley 1990). In contrast, most plants first transaminate prephenate by prephenate aminotransferase (PPA-AT, EC 2.6.1.78 or 79) into arogenate, which is then converted to Phe and Tyr by arogenate dehydratase and dehydrogenase (ADT and ADH), respectively (Fig. 1c, Jensen 1986; Rippert and Matringe 2002; Cho et al. 2007; Tzin and Galili 2010; Maeda and Dudareva 2012). In vascular plants, up to 30% of total deposited carbon can pass through the shikimate and Phe pathways for synthesizing abundant phenylpropanoid compounds such as lignin and tannins (Weiss 1986; Benner et al. 1987; Razal et al. 1996). Thus, plant PPA-ATs catalyze one of the major transamination reactions in plants. Euglena gracilis, which acquired chloroplasts through secondary symbiosis of green algae (Bhattacharya and Medlin 1998), has four PPA-AT activities that can be separated by chromatography and all have comparable activity towards prephenate, HPP, and PPY (Byng et al. 1981). PPA-AT activity was also detected from various plants, Nicotiana sylvestris, Sorghum bicolor, Vigna radiata (mung bean) and A.
Tissue
Cell culture
Seedling
Cell culture
Nicotiana sylvestris
Sorghum bicolor
Anchusa officinalis
Seedling
Seedling
Seedling
Seedling
Cell culture
Phaseolus vulgaris
Vigna radiata
Pisum sativum
Vigna radiata
Anchusa officinalis
AAA-AT activity
Seedling
Vigna radiata
PPA-AT activity
Plant species
–
–
75
29
–
–
Tyr-AT3: Tyr 0.45 [a-KG]
Tyr-AT2: Tyr 5 [aKG]
Tyr-AT1: Tyr 20 [aKG]
–
a-KG (100), KDP (36), OAA (23), GLO (10), PRY (5) [Tyr]
–
–
–
OAA (100), a-KG (86), KDP (81), PYR (67), GLO (14) [Tyr] OAA (100), PYR (60), KDP (49), a-KG (35), GLO (12) [Tyr]
– HPP (100), PPY (51), PPA (24) [Asp]
8.8–9.4
9.6
9
–
8.5
–
PYR (100), OAA (92), a-KG (61), GLO (30) [D,L-Trp]
–
D,L-Ala (100), D-Trp (34), D,L-Met (31), D,L-Leu (30), D,L-Trp (30), L-Trp (11) [a-KG]
8.5
–
PYR (100), 4MOP (89), OAA (82), 3MOP (72), 3MOB (52), GLO (8), HPY (4) [Trp]
Trp 0.33 [aKG]
Lys (100), Leu (86), Met (75), Arg (74), Ala (70), Phe (57), Tyr (46), Trp (35), Asp (33), His (29), Val (22), Cys (1) [a-KG]
–
–
–
–
–
–
–
–
9
8.5
PPA 0.08 [Asp]
Plastid [S]
–
–
Subcellular localization [method]b
8–8.5
8.25
–
pH optimum
a-KG 0.8 [Phe]
PPA (100), HPP (19), PPY (10), IPA (7) [Asp]
PPA (100), OAA (78), HPP (10), pyruvate (10) [Glu] PPA 0.07 [Glu]
–
a-KG (100), PPA (63), OAA (50), PYR (29) [Glu]; a-KG (100), OAA (76), PPA (49) [Asp]
–
Km (mM)
PPA (100), HPP (14), PPY (7) [Asp]; PPA (100), HPP (15), PPY (4) [Glu]
Preferred substrate (% of the best)a
PYR (100), a-KG (76) [Phe]
–
AGN 0.8 [aKG]
–
–
Km (mM)
Keto acid substrate [co-substrate used]
Tyr 0.27, Phe 0.4, Trp 1.7 [PYR]
Gln (100), Asp (58), Lys (53), Glu (49), Arg (39), Phe (36), Tyr (29), Met (26), Leu (21), Trp (11), Ile (9), Asn (8), His (6) [PYR]
–
Glu (100), Asp (18) [PPA]
–
–c
Preferred substrate (% of the best)a
Amino acid substrate [co-substrate used]
84
13
53
29
17
41
–
–
5 to 8
Purification (fold)
Table 1 Biochemical properties of purified native plant AAA-AT activity
Ellis and DeEknamkul (1987)
Rubin and Jensen (1979)
Matheron and Moore (1973)
Truelsen (1972)
Gamborg and Wetter (1963), Gamborg (1965)
De-Eknamkul and Ellis (1988)
Siehl et al. (1986a, b)
Bonner and Jensen (1985)
Rubin and Jensen (1979)
References
Phytochem Rev
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Seedling
Coleoptile
Seedling
Pisum sativum
Zea mays
Vigna radiata Trp (100), Tyr (83), Phe (75), Arg (35), Ala (25), Lys (25), Leu (10), Asp (6), Asn (5), His (4), Val (3) [a-KG]
–
27
33,000
–
24
Phe 0.07, Tyr 0.08, Trp 0.095 [a-KG]
–
–
–
–
Plastid extract converted DTrp better than L-Trp into IAA by [sevenfold. –
Km (mM)
Preferred substrate (% of the best)a
Amino acid substrate [co-substrate used]
4
1508
Purification (fold)
–
PYR 0.24, OAA 0.25, aKG 0.65, [Trp]
OAA (100), PYR (91), a-KG (78), HPP (31), HPY (7), GLO (0.8) [Trp]
Km (mM)
D-Trp-AT: PYR (100), GLO (26), a-KG (12), OAA (2.6) [Trp]
–
–
L-Trp-AT1: a-KG (100), OAA (23), PYR (14), GLO (4) [Trp] L-Trp-AT2: a-KG (100), OAA (15), PYR (15), GLO (2.9) [Trp]
–
PYR (100), OAA (93), a-KG (54), PPY (47) [D-Trp]
Preferred substrate (% of the best)a
Keto acid substrate [co-substrate used]
–
8–9
8–9
8–9
–
pH optimum
–
–
–
–
Plastid [S]
Subcellular localization [method]b
Simpson et al. (1997)
Koshiba et al. (1993)
McQueen-Mason and Hamilton (1989)
References
c
b
a
‘–’ Indicates that data is not available
Subcellular localization was analyzed by subcellular fractionation [S] or localization of a GFP-fusion protein [G]
Percent activity for each substrate is shown in parentheses relative to the highest activity
Ala, alanine; Asp, aspartate; Glu, glutamate; GLO, glyoxylate; HPP, 4-hydroxyphenylpyruvic acid; IPA, indole-3-pyruvate; a- KG, a-ketoglutarate; Met, Methionine; 4MTOB, 2-oxo-4-methylthiobutyric acid; OAA, oxaloacetate; Phe, phenylalanine; PPA, prephenate; PPY, phenylpyruvic acid; PYR, pyruvate; Trp, tryptophan; Tyr, tyrosine
Aromatic amino acid and corresponding keto acid substrates are indicated in bold
Tissue
Plant species
Table 1 continued
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officinalis), which all show high specificity towards prephenate, with low to undetectable activity towards other aromatic keto acid substrates (i.e. PPY and HPP) (Rubin and Jensen 1979; Bonner and Jensen 1985; Siehl et al. 1986a; De-Eknamkul and Ellis 1988) (Table 1). These plant PPA-AT activities prefer acidic amino donors, Asp and Glu, and favor the direction of prephenate transamination to arogenate (Siehl et al. 1986a), though the reverse reaction, the arogenate deamination (Fig. 1b), was rarely tested likely due to limited availability of arogenate. PPA-AT is usually not feedback inhibited, such as by Phe or Tyr, with an exception of A.officinalis PPA-AT activity that was inhibited by 3,4-dihydroxyphenyllactic acid, the intermediate of rosmarinic acid biosynthesis (De-Eknamkul and Ellis 1988). Genes encoding PPA-ATs were identified from Arabidopsis (At2g22250), Petunia hybrida, and tomato (Solanum lycopersicum) through biochemical purification of PPA-AT activity and gene co-expression analyses (Graindorge et al. 2010; Dal Cin et al. 2011; Maeda et al. 2011) (Table 2). Like PPA-AT activity detected from plant tissues, PPA-AT recombinant enzymes specifically use prephenate, but not PPY or HPP, with Asp and Glu amino donors (Graindorge et al. 2010; Dal Cin et al. 2011; Maeda et al. 2011) and also prefer prephenate transamination to arogenate (Maeda et al. 2011). Plant PPA-AT enzymes also have comparable Asp-AT activity to PPA-AT activity and are able to transaminate aketoglutarate and oxaloacetate, besides prephenate (Rubin and Jensen 1979; Bonner and Jensen 1985; Siehl et al. 1986a; De-Eknamkul and Ellis 1988; Graindorge et al. 2010; Dal Cin et al. 2011; Maeda et al. 2011), consistent with in silico docking of Asp and Glu substrate on a structure model of maritime pine (Pinus pinaster Ait.) PPA-AT ortholog (de la Torre et al. 2009). Thus, plant PPA-ATs specifically utilize acidic dicarboxylic acid substrates (i.e. prephenate, a-ketoglutarate, and oxaloacetate keto acid acceptors, Asp and Glu amino donors). Another unique feature of plant PPA-AT activity was their high temperature stability up to 70°C, which was used to purify the PPA-AT activity (Bonner and Jensen 1987; Siehl et al. 1986a; De-Eknamkul and Ellis 1988) and was also observed in the recombinant enzymes of maritime pine, Arabidopsis, and tomato PPA-ATs (de la Torre et al. 2007; Dal Cin et al. 2011; Dornfeld et al. 2014).
To identify amino acid residues responsible for the acidic dicarboxylic acid substrate specificity of PPAATs, phylogeny-guided biochemical characterization of PPA-AT orthologs was conducted from a wide range of plants and microbes (Dornfeld et al. 2014). Combined with PPA-AT structure model generated based on a crystal structure of microbial Asp-AT, two active site residues, Lys169 and Thr84 (numbering based on Arabidopsis PPA-AT), were found to be highly conserved among functional PPA-AT orthologs truly having PPA-AT activity. Simultaneous mutations of these two residues completely abolished PPA-AT (and Asp-AT) activity but instead introduced a new AAA-AT activity that efficiently transaminates HPP into Tyr using neutral amino donors (e.g. alanine, Trp) (Dornfeld et al. 2014). Although actual crystal structures of plant PPA-ATs remain to be determined, these active site residues are likely involved in direct recognition of dicarboxylic acidic substrates of PPAATs (i.e., prephenate keto acid substrate and Asp or Glu amino donor). These results suggest that Lys169 and Thr84 likely played a key role in the evolution of PPA-AT enzymes and hence the arogenate pathways of Tyr and Phe biosynthesis that operate in plants and some microbes today. Tyrosine and Phenylalanine aminotransferases (Tyr-ATs and Phe-ATs, EC 2.6.1.5) In plants, owing to the presence of the major Tyr and Phe pathway mediated by PPA-ATs (Fig. 1c), Tyr and Phe-ATs have taken diverse roles in utilizing Tyr and Phe to synthesize a variety of plant specialized metabolites, while some provide alternative microbial-type Phe and Tyr biosynthesis pathways (Fig. 1c). The first AAA-AT activity partially purified from mung bean shoots showed activity towards Phe, Tyr and Trp, with the lowest Km towards Tyr (Gamborg and Wetter 1963) (Table 1), which was later found to be active towards other amino acids as well (e.g. lysine, glutamine) when pyruvate was used as keto acceptor (Gamborg 1965). Aminotransferase activity that deaminates both AAA and aliphatic amino acids was also detected in shoot tip of pea (Pisum sativum) seedlings (Matheron and Moore 1973) and mung bean (Vigna radiata) seedlings (Truelsen 1972). Tyr-AT activity that deaminates Tyr into HPP using aketoglutarate keto acceptor was detected and partially purified in opium poppy (Papaver somniferum) and
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123
ADM67557 (PhPPA-AT)
ADQ00383 (SlPAT)
Petunia hybrida
Solanum lycopersicum
ADC45389 (CmArAT1)
ADC33123 (PsTyrAT)
XP_011457732 (RyAAAT3)
Cucumis melo
Papaver somniferum
Rosa x hybrida
Tyr-AT and Phe-AT enzymes
At2g22250 (AtPPA-AT)
Accession number (name)
Arabidopsis thaliana
PPA-AT enzymes
Plant species
Glu 5.6 [OAA]; Asp 12 [aKG]
Glu 36 [OAA];
Tyr 1.8, Phe 6.3, Trp 7.8 [a-KG] Phe 0.7 [OAA]; Phe 1.5 [a-KG]
Tyr 0.13, Trp 0.05, Phe 0.01 [a-KG]
Vmax/Km (mM s-1)a:
Phe 0.89 [a-KG]; Phe 0.56 [OAA]
26-fold higher activity towards Tyr than other amino acids [PPY]
*Fourfold higher activity towards Phe than Tyr [a-KG]
–
AGN 2.1 [aKG]
AGN 3.1 [a-KG]
–
Asp 3.2, Glu 3.3 [PPA];
Glu 2.7, Asp 2.8 [PPA]
Glu 1.7, Asp 1.3 [PPA]
Glu 1.4, Asp 1.3 [PPA];
–
–
Asp 5.4 [a-KG]
Glu 1.5, Asp 2.2 [PPA];
Glu 63, Asp 13 [PPA];
PPA 0.36,
a-KG 25, PPA 13 [Asp];
a-KG 0.28 [Asp]; PPA 0.10, OAA 0.32 [Glu]
PPA 115, OAA 47 [Glu]
–
a-KG 0.63, PYR 0.14, OAA 0.01 [Tyr]
–
a-KG 0.35, PYR 2.5, OAA 56 [Tyr]
1.5-fold higher activity towards HPP than aKG [Phe]
PPA 0.08,
a-KG 70 [Asp];
OAA 12 [Glu]
PPA 142,
PPA 0.48,
OAA 4.9 [Glu]
a-KG 1.0 [Asp];
PPA 20 [Asp]; PPA 28,
PPA 0.32,
OAA 8.2 [Glu]
PPA 0.68,
a-KG 34,
OAA 21, PPA 13 [Glu]
PPA 0.2 [Asp] a-KG 0.69 [Asp];
PPA 0.1 [Glu];
PPA 168 [Glu]
OAA 0.025 [Glu]
PPA 118 [Asp];
PPA 6,786 [Glu]
PPA 0.014,
a-KG 0.2 [Asp];
a-KG 325 [Asp]; OAA 8,000.
PPA 0.013,
PPA 2,154,
Km (mM)
kcat/ Km (mM-1 s-1)a
kcat/Km (mM-1 s-1)a Km (mM)
Keto acid substrate [co-substrate used]
Amino acid substrate [cosubstrate used]
Table 2 Biochemical properties of purified native and recombinant plant AAA-ATs
Phe ? PPYf
–
PPY [Tyr] ? Phe [HPP]f
–
PPA [Glu/ Asp] ? AGN [aKG/OAA]e
PPA [Glu/ Asp] ? AGN [aKG/OAA]e
–
–c
Favored reaction direction
9
8.5
–
–
–
–
–
–
pH optimum
–
–
–
Plastid [G]
Plastid [P]
Plastid [P]
Plastid [P][S]d
Subcellular localization [method]b
Hirata et al. (2012)
Lee and Facchini (2011)
Gonda et al. (2010), Yoo et al. (2013)
Dal Cin et al. (2011)
Maeda et al. (2011)
Maeda et al. (2011)
Dal Cin et al. (2011)
Graindorge et al. (2010)
References
Phytochem Rev
Accession number (name)
AHA62827 (PhPPY-AT)
AHN10104 (AbArAT4)
XP_004250335 (SlGTK)
NP_001143647 (ZmGTK)
AGK24944 (EsAroAT1)
At5g53970 (AtTAT1)
At5g36160 (AtTAT2)
Plant species
Petunia hybrida
Atropa belladonna
Solanum lycopersicum
Zea mays
Ephedra sinica
Arabidopsis thaliana
Arabidopsis thaliana
Table 2 continued
PPY 1.5 [Tyr];
HPP 0.5, OAA 0.5, aKG 5.2, PYR 19 [Phe]; PPY 0.5 [Tyr]
PPY 0.97 [Tyr]; PPY 0.46 [Glu]; a-KG 0.50, HPP 0.09 [Phe] HPP 87, a-KG 0.26, OAA 0.21, PYR 0.008 [Phe]; PPY 0.35 [Tyr]
Tyr 1.9, Glu 11 [PPY]; Phe 4.3 [HPP]; Phe 14 [aKG] Phe 1.4 [a-KG];
Tyr 0.77, Glu 0.04 [PPY];
Phe 0.04 [HPP];
Tyr 0.03 [PPY]
Met 0.32, Tyr 0.23, Phe 0.08 [a-KG]
Glu 1.4 [HPP] Tyr 2.9, Met 1.7, Phe 6.7 [a-KG]
4MTOB 0.51, a-KG 0.07, PPY 0.04, OAA 0.02 [Tyr]
HPP 1.4, 4MTOB 1.2, PPY 0.5 [Glu];
–
HPP 0.4, 4MTOB 0.8, PPY 1.3 [Glu]
4MTOB 0.7, PPY 3.4, aKG 7.6, OAA 43 [Tyr];
a-KG 1.2 [Phe]
HPP 0.22 [Glu];
PPY 0.13,
HPP 10.1 [Glu]
PPY 0.46, 4MTOB 1.3,
PPY 54, 4MTOB 16, HPP 2.5 [Glu]
Tyr 0.19, Phe 0.84 [a-KG];
PPY 3.1, a-KG 3.3, 4MTOB 3.5, [Tyr];
a-KG 15.1, PPY 9.9, 4MTOB 5.5 [Tyr];
Tyr 0.2, Met 24, Phe 6.9 [aKG]
Tyr 282, Phe 4.5, Met 1.1 [a-KG]
–
a-KG 0.42 [Tyr]
a-KG 2.94 [Tyr]
HPP [Glu] ? Tyr [a-KG]
8.5
8
8.5–9
Tyr [a-KG] ? HPP [Glu]
–
8
HPP 0.3 [Phe];
HPP 0.14 [Phe]
Tyr ? HPPf
9
Tyr [a-KG] ? HPP [Glu]
IPA 0.04, HPP 0.1, PPY 0.12 [Glu];
a-KG 41 [Tyr]; PPY 7.2, HPP 6.1, IPA 4.5 [Glu]; a-KG 0.8 [Tyr]
–
–
–
–
pH optimum
–
–
Phe [HPP] ? PPY [Tyr]
PPY [Tyr] ? Phe [HPP]
Favored reaction direction
% of the best: 4MTOB (100), PPY (10) [Gln]
Tyr 0.18 [a-KG]
Tyr 0.9, Phe 8.7, Trp 5.4 [a-KG]
Tyr 0.37 [4MTOB];
Tyr 0.26 [PYR]; Tyr 0.27 [OAA]; Tyr 0.7, Glu 9.4 [PPY];
Gln 0.07 [4MTOB]
% of the best: 4MTOB (100), PPY (81), 4MOP (15) [Gln]
a-KG 1.8, HPP 7.9 [Phe];
Tyr 7.72 [a-KG]
Tyr 0.16, Glu 0.09 [PPY];
Tyr 1.0 [PYR];
Tyr 7.4 [4MTOB]; Tyr 3.0 [OAA];
Tyr 36, Phe 0.97, Trp 0.67 [aKG];
Gln 1.73 [4MTOB]
Gln 0.22 [4MTOB]
Tyr 4.1 [PPY];
Phe 1.3 [a-KG];
Gln 0.71 [4MTOB]
Phe 2.1, Trp 20 [HPP];
Phe 15, Trp 0.2 [HPP];
Phe 0.04 [a-KG]
Km (mM)
kcat/ Km (mM-1 s-1)a
Km (mM)
kcat/Km (mM-1 s-1)a
PPY 2.6 [Glu];
Keto acid substrate [co-substrate used]
Amino acid substrate [cosubstrate used]
Cytosol [G]
–
Cytosol [G]
–
–
Cytosol [G]
Cytosol [G]
–
Cytosol [G]
Subcellular localization [method]b
Wang et al. (2016)
Prabhu and Hudson (2010)
Wang et al. (2016)
Riewe et al. (2012)
Kilpatrick et al. (2016)
Ellens et al. (2015)
Ellens et al. (2015)
Bedewitz et al. (2014)
Yoo et al. (2013)
References
Phytochem Rev
123
123
At1g80360 (ATVAS1)
Zheng et al. (2013)
Arabidopsis thaliana
Cytosol [G]
Met 0.63, Phe 2.6 [IPA]
Met 0.05,
Trp 0.3, Tyr 4.7, Phe 9.4 [a-KG]
Phe 0.002[IPA]
Phe 0.01 [a-KG]
Trp 2.7, Tyr 1.1
Vmax/Km (mM s-1):
IPA 0.6 [Met]
–
IPA
–
Favored reaction direction
9
pH optimum
–
[Met] ? Trp[4MTOB]
Subcellular localization [method]b
–
Tao et al. (2008)
References
f
e
d
c
b
a
A preferred reaction direction was examined by comparing specific activities
The best co-substrate are shown in brackets when substrate specificity was analyzed
At2g22250 was identified as aspartate aminotransferase and recognized in the plastid fraction by immunoblotting (de la Torre et al. 2006)
‘–’ Indicates that data is not available
Subcellular localization was analyzed by subcellular fractionation [S] or localization of a GFP-fusion protein [G]
When catalytic efficiency was not reported, Vmax/Km is indicated instead
Ala, alanine; Asp, aspartate; Glu, glutamate; GLO, glyoxylate; HPP, 4-hydroxyphenylpyruvic acid; IPA, indole-3-pyruvate; a- KG, a-ketoglutarate; Met, Methionine; 4MTOB, 2-oxo-4-methylthiobutyric acid; OAA, oxaloacetate; Phe, phenylalanine; PPA, prephenate; PPY, phenylpyruvic acid; PYR, pyruvate; Trp, tryptophan; Tyr, tyrosine
IPA, 0.07 [Met]
–
Km (mM)
kcat/ Km (mM-1 s-1)a
kcat/Km (mM-1 s-1)a Km (mM)
Keto acid substrate [cosubstrate used]
Amino acid substrate [cosubstrate used]
Aromatic amino acid and corresponding keto acid substrates are indicated in bold
At1g70560 (AtTAA1)
Accession number (name)
Arabidopsis thaliana
Trp-AT enzymes
Plant species
Table 2 continued
Phytochem Rev
Phytochem Rev
was proposed to catalyze the initial step of the benzylisoquinoline alkaloid biosynthesis from Tyr (Jindra et al. 1966; Rueffer and Zenk 1987). To search for AAA-AT(s) involved in the final step of Tyr and Phe biosynthesis (as the major Tyr/Phe biosynthetic pathway was unresolved at that time), Rubin and Jensen (1979) partially purified aminotransferase activity from mung bean that transaminates HPP, but not PPY, using a glutamate (Glu) amino donor, though the reverse reaction (Tyr deamination) was not tested. Tyr-AT activities from Anchusa officinalis were chromatographically separated into four distinct peaks, three of which were specific to Tyr with \30% activity towards Phe (Ellis and De-Eknamkul 1987), while the fourth peak showed PPA-AT activity (De-Eknamkul and Ellis 1988). All four purified Tyr-AT activities of A. officinalis were inhibited by 3,4-dihydroxyphenyllactate, the intermediate of rosmarinic acid biosynthesis, and two of them were also inhibited by rosmarinic acid itself (Ellis and De-Eknamkul 1987). These early studies detected AAA-AT activities utilizing Tyr and Phe substrates from various plants, which often exhibit broad substrate specificity when other amino acids and keto acid substrates were tested. More recently, genes and enzymes responsible for these Tyr- and Phe-ATs have been identified and their recombinant enzymes have been biochemically characterized (Table 2). A recombinant AAA-AT of melon (Cucumis melo, CmArAT1) has higher activity towards Phe than Tyr, though Trp-AT activity and the reverse reactions were not tested (Gonda et al. 2010). Tyr-AT from opium poppy (PsTryAT) exhibits a much higher catalytic efficiency towards Tyr than Phe or Trp (Lee and Facchini 2011). Detailed substrate specificity analyses of AAA-AT from deadly nightshade (Atropa belladonna, Ab-ArAT4) using various amino donors and keto acceptors demonstrated that Ab-ArAT4 prefers Phe deamination to PPY using HPP as the best keto acceptor (Bedewitz et al. 2014). AAAAT from rose (Rosa hybrida var. Yves Piaget, RyAAAT3) shows higher specific activity with Phe than Tyr, Trp, or other amino acid substrates, and also favors Phe deamination than the reverse reaction, PPY to Phe, by *tenfold (Hirata et al. 2012). AAA-ATs from the gymnosperm Ephedra sinica (EsAroAT1 and EsAroAT2) have activities towards both Tyr and Phe, with EsAroAT1 having 3 to 7-fold higher specific activity than EsAroAT2 (Kilpatrick et al. 2016).
EsAroAT1 shows the highest catalytic efficiency toward Tyr deamination to HPP using a-ketoglutarate as the best keto acceptor, followed by 2-oxo-4methylthiobutyric acid, the keto acid of methionine (Met) (Kilpatrick et al. 2016). Substrate specificity analyses of AAA-AT from petunia (Petunia hybrida, PhPPY-AT) and melon (C. melo, CmArAT1) showed that both of them had the highest catalytic efficiency in transaminating PPY to synthesize Phe while using Tyr as the best amino donor (Yoo et al. 2013), which is the same substrate combination but the opposite direction to Ab-ArAT4 (Bedewitz et al. 2014). Arabidopsis thaliana has at least two Tyr-ATs encoded by AtTAT1 (At5g53970) and AtTAT2 (At5g36160) (Prabhu and Hudson 2010; Riewe et al. 2012; Wang et al. 2016). AtTAT1 strongly prefers Tyr over Phe, Trp, or other amino acid donors and also favors Tyr deamination to HPP rather than HPP transamination to Tyr (Riewe et al. 2012; Wang et al. 2016). The Tyr deamination reaction of AtTAT1 was best coupled with a-ketoglutarate followed by PPY and 2-oxo-4-methylthiobutyric acid (Wang et al. 2016). Unlike AtTAT1, AtTAT2 favors HPP transamination to synthesize Tyr and has a much broader substrate specificity using both AAAs and aliphatic amino acids (Wang et al. 2016). AtTAT2 generally exhibits much lower activity than AtTAT1 (by *40-fold) and it is possible that AtTAT2 utilizes yet to be identified non-proteinogenic amino acid substrates. Taken together, many AAA-ATs that utilize Tyr and Phe (thus categorized as Tyr and Phe-ATs) often have broad substrate specificity, though some are relatively specific to Tyr and/or Phe or not examined extensively for different substrate specificity. Interestingly, when different combinations of amino donors and keto acceptors were rigorously tested for substrate specificity, the deamination of Tyr to HPP was often coupled to the PPY to Phe synthesis (i.e. PhPPA-AT, CmArAT1, AtTAT1), or vice versa (AbArAT4) (Fig. 1c). Tryptophan aminotransferases (Trp-AT, EC 2.6.1.27 or 99) Trp-AT activity was detected from many plant tissues but mostly as a side activity of Tyr/Phe-AT or other aliphatic amino acid aminotransferase activity (Table 1) (Gamborg 1965; Truelsen 1972; Matheron
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and Moore 1973). Trp-AT activity was also detected from a peroxisomal fraction of spinach (Spinacia oleracea) leaf extract, but prefers serine and alanine over Trp by *tenfold (Noguchi and Hayashi 1980). Three Trp-AT activities were purified from maize (Zea mays) coleoptiles: two of them converted L-Trp to indole-3-pyruvic acid (IPA) using a-ketoglutarate, while the remaining one was specific to D-Trp using pyruvate as its keto acceptor (Koshiba et al. 1993). Trp-AT activity partially purified from P. sativum seedlings also preferred D-Trp over L-Trp (Matheron and Moore 1973; McQueen-Mason and Hamilton 1989), whereas that from crude extract of wheat seedlings favored L-Trp over D-Trp (Rekoslavskaya et al. 1999). The partially purified AAA-AT activity from mung bean seedlings exhibited the highest activity towards L-Trp among other amino acids, had little activity toward D-Trp, and was slightly inhibited by a-naphthaleneacetic acid, an analog of auxin indole-3-acetic acid (IAA), but not by IAA itself (Simpson et al. 1997). Thus, depending on tissues analyzed, Trp-AT activity can convert either L-Trp or D-Trp or both to IPA, a precursor of plant hormone auxin, IAA (Vanneste and Friml 2009). D-Amino acid aminotransferase (At5g57850), which belongs to class IV aminotransferases, was identified from Arabidopsis and showed activity towards a wide range of D-amino acids including DTrp (Funakoshi et al. 2008); however, its physiological role remained to be tested. Arabidopsis mutants, shade avoidance 3 (sav3), weak ethylene insensitive 8 (wei8), and transport inhibitor response 2 (tir2) were independently found to be defective in the Trp-AT enzyme encoded by TRYPTOPHAN AMINOTRANSFERASE OF ARABIDOPSIS 1 (TAA1, At1g70560) (Tao et al. 2008; Stepanova et al. 2008; Yamada et al. 2009). TAA1 deaminates L-Trp into IPA and prefers Trp over Phe or Tyr as an amino donor mainly due to its low Km towards L-Trp (0.3 mM vs. 5 and 9 mM for Tyr and Phe, respectively) (Tao et al. 2008). X-ray crystal structure analysis of Arabidopsis TAA1 (PDB: 3BWN), the only structure of plant AAA-ATs to date, and in silico docking analysis further supported that IPA and L-Trp serve as preferred ligands of TAA1 among Phe, Tyr, histidine, and D-Trp (Tao et al. 2008). The reverse reaction, IPA transamination to Trp, has not been tested. One of the sav3 suppressors, vas1 (reversal of sav3 phenotype 1), encoded an aminotransferase that efficiently transaminates IPA back to
123
Trp and is incapable of deaminating Trp into IPA (Zheng et al. 2013). Interestingly, among various amino donors tested, VAS1 most efficiently utilized Met followed by valine, isoleucine, Tyr, and leucine (Zheng et al. 2013). Therefore, TAA1 deaminates Trp into IPA, whereas VAS1 reverts IPA back to Trp and at the same time converts Met to 2-oxo-4-methylthiobutyric acid in Arabidopsis (Tao et al. 2008; Zheng et al. 2013, Fig. 1c). TAA1 related proteins (TARs) (Stepanova et al. 2008; Yoshikawa et al. 2014), which are closely related to TAA1 (see Sect. 3), also likely contribute to Trp deamination to IPA, though their biochemical properties have not been examined. Other AAA-ATs L-3,4-Dihydroxyphenylalanine
(L-DOPA) is synthesized from Tyr by an additional ring hydroxylation reaction and serves as a key precursor of benzylisoquinoline alkaloids and betalain pigments (Gandı´aHerrero and Garcı´a-Carmona 2013; Beaudoin and Facchini 2014). Plant DOPA-AT (EC 2.6.1.49) activity was first detected in leaf crude extract of opium poppy (Jindra et al. 1966) and also from the cell cultures of isoquinoline alkaloid-containing plants (Rueffer and Zenk 1987). In many studies investigating AAA-ATs, however, L-DOPA (a non-proteinogenic AAA) was not commonly tested as a substrate. Genes and enzymes responsible for DOPA-AT activity and their physiological significance are currently unknown in plants. Kynurenine is an intermediate of Trp catabolism towards the generation of neuroactive compounds, kynurenate, 3-hydroxykynurenine, and quinolinate, in the mammalian brain (Schwarcz et al. 2012). Kynurenine and downstream metabolites are toxic to animals at high concentrations but are sometimes accumulated in various fungi (McGary et al. 2013). Kynurenine aminotransferase (EC 2.6.1.7) activity that converts kynurenine to kynurenate has been reported in the kingdoms of fungi and animals (Wogulis et al. 2008; Han et al. 2010), but not in plants so far. Human kynurenine aminotransferase I is identical to glutamine transaminase K (EC 2.6.1.64), involved in the Met salvage pathway (Cooper 2004), and exhibits broad substrate specificity utilizing Gln, Met, Leu, kynurenine, Tyr, and Phe as amino donors, and aketoglutarate, PPY, and 2-oxo-4-methylthiobutyrate as keto acceptors (Cooper et al. 2008). Similarly,
Phytochem Rev
glutamine transaminase K enzymes of tomato and maize transaminate 2-oxo-4-methylthiobutyric acid and, to a lesser extent PPY, into Met and Phe, respectively, specifically using glutamine as the amino donor (Table 2) (Ellens et al. 2015). Thus, plant glutamine transaminase K enzymes also catalyze the final step of the Met salvage pathway (Ellens et al. 2015), also known as the Yang cycle (Miyazaki and Yang 1987; Sauter et al. 2013) (Fig. 1c). Other aminotransferases that utilize non-AAA substrates appear to have residual activity towards some AAAs as well. When Asp-AT activity was purified from P. vulgaris seedlings and tested for 19 proteinogenic amino acid donors, it exhibited aminotransferase activity towards Phe, Tyr, and Trp, though was less than 10% of that towards Asp (Forest and Wightman 1972, 1973). Also, rose RyAAAT1 and RyAAAT2 that are closely-related to plant Asp-ATs and alanine aminotransferases, respectively, showed residual Phe-AT activity (Phe transamination to PPY), though much weaker than RyAAAT3 (Hirata et al. 2012). Although AAAs and their corresponding keto acids were not always tested for other aminotransferases, moonlight activity of other aminotransferases having broad substrate specificity could also contribute to total AAA-AT activity detected in plants.
Phylogenetic relationships of AAA-ATs AAA-ATs belong to class I aminotransferases, which also include Asp-ATs and alanine aminotransferases (Jensen and Gu 1996). The identifications of representative AAA-AT enzymes and genes from different plant species (as described above) now allow us to examine how plant AAA-ATs are related with each other and to AAA-ATs from other kingdoms. Here we constructed a phylogenetic tree of previously characterized plant AAA-ATs together with closely related Arabidopsis sequences as well as characterized class I aminotransferases from animals and microbes (Fig. 2). Available crystal structure information (as indicated in the tree) was incorporated using PROMALS3D to improve the alignment of divergent sequences (Pei et al. 2008). Branch-chain amino acid aminotransferases (BCATs) from class IV were used as an outgroup (Table 3, Diebold et al. 2002; Schuster et al. 2006; Knill et al. 2008; Binder 2010; Kochevenko et al. 2012; La¨chler et al. 2015).
Tyr- and Phe-ATs from angiosperm form a tight monophyletic clade (Fig. 2). Both Tyr-ATs of Arabidopsis (AtTAT1 and AtTAT2) belong to this clade, so do all five homologs of AbArATs from deadly nightshade (Bedewitz et al. 2014). The Tyr/Phe-AT clade of angiosperm is sister to the clade containing carbon-sulfur (C–S) lyases, which are uniquely present in the Brassicaceae family (Wang et al. 2016) and do not have Tyr-AT activity (Jones et al. 2003; Mikkelsen et al. 2004). Some of the C–S lyases (i.e. CORI3, SUR1) were shown to be involved in glucosinolates metabolism (Mikkelsen et al. 2004; Klein and Sattely 2017), but the function and potential Tyr-AT activity of the remaining proteins within the C–S lyase clade are yet to be examined. Gymnosperm AAA-ATs (e.g. EsAroAT1 and 2) form basal branches of the Tyr/Phe-AT and C–S lyase clades (Fig. 2). Plant Tyr/Phe-ATs are much more closely related to those of animals (HcTAT, MmTAT) than of fungi (ScARO8 and 9) and bacteria (EcTyrB, KpTyrB), the latter is sister to ubiquitous class Ia Asp-ATs found in all organisms (Schultz and Coruzzi 1995; Schultz et al. 1998, Fig. 2). In agreement with the acidic substrate specificity of plant PPA-AT enzymes (see Sect. 2.3), plant PPA-ATs belong to class Ib Asp-ATs (Dornfeld et al. 2014), which also specifically utilize acidic, but not neutral, amino donors and are distantly related to class Ia AspATs (Kuramitsu et al. 1990; Nobe et al. 1998; Nakai et al. 1999). Interestingly, class Ib Asp-ATs from Chlorobi/Bacteroidetes show much higher PPA-AT activity and sequence similarity to plant PPA-ATs than those of cyanobacteria, suggesting that a common ancestor of plants and algae obtained a PPA-AT enzyme through a novel horizontal gene transfer event (Dornfeld et al. 2014). Cyanobacteria (e.g. Synechocystis sp. PCC6803) instead possess an additional class IV aminotransferase that belongs to BCATs but efficiently catalyzes the PPA-AT reaction (Graindorge et al. 2014), representing convergent evolution of PPA-AT enzymes in distinct classes of aminotransferases (i.e. class I and IV). Like other AAA-ATs, Trp-ATs of Arabidopsis (TAA1 and TARs) and monocots (ZmTAR1, ZmVT2, OsTAR2) form a well-supported clade (Fig. 2). As expected, maize and tomato glutamine transaminase K enzymes and their Arabidopsis orthologs (At1g77670) (Ellens et al. 2015) are sister to mammalian and yeast kynurenine aminotransferase I/glutamine
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Phytochem Rev
123
Phytochem Rev b Fig. 2 Maximum likelihood phylogenetic tree of characterized
aminotransferases with AAA-AT activity and their Arabidopsis homologs. Alignment was performed by structure-guided multiple sequence alignment using PROMALS3D (Pei et al. 2008). The PDB IDs of the crystal structures used in the analysis are indicated in red letters. The original alignment is found in Supplemental Data. Analysis was performed under default setting in MEGA 6 (Tamura et al. 2013) with partial deletion site coverage set to 50%. While each plant AAA-AT type formed a well-supported monophyletic clade (green boxes), the relationship among different AAA-AT types was not well resolved. Percentage values of replicate trees in bootstrap test (1000 replicates) are shown next to the branches (value less than 50 are not shown) with branch length indicating the number of substitutions per site. Class IV BCATs were used as an outgroup. Names of characterized enzyme are listed after NCBI or AGI accession number and also found in Table 3. Asp-AT aspartate aminotransferase, BCAT branched-chain amino acid aminotransferase, C–S lyase carbon–sulfur lyase, DAPAT diaminopimelate aminotransferase, KAT/GTK Kynurenine aminotransferase/glutamine transaminase K, Phe-AT Phe aminotransferase, PPA-AT prephenate aminotransferase, TrpAT Trp aminotransferase, Tyr-AT Tyr aminotransferase. Aa, Alicyclobacillus acidocaldarius; Ab, Atropa belladonna; Am, Antirrhinum majus; As, Aspergillus sp; At, Arabidopsis thaliana; Ba, Bacillus anthracis; Bc, Bacillus cereus; Ct, Chlorobium tepidum; Cm, Cucumis melo; Dd, Dickeya dadantii; Ec, Escherichia coli; Es, Ephedra sinica; Hs, Homo sapiens; Kp, Klebsiella pneumoniae; Ll Lactococcus lactis; Mf, Malassezia furfur; Mm, Mus musculus; Mt, Mycobacterium tuberculosis; Os, Oryza sativa; Ne, Nitrosomonas europaea; Ph, Petunia hybrid; Pp, Pinus pinaster; Ps, Papaver somniferum; Rs_Rhodobacter sphaeroides; Rh, Rosa hybrida; Sa, Streptomyces avermitilis; Sb, Streptomyces bingchenggensis; Sc, Saccharomyces cerevisiae; Sm, Sinorhizobium meliloti; Ssp, Synechocystis sp PCC6803; Ssc, Synechococcus sp. CC9605; Tc, Trypanosoma cruzi; Tt, Thermus thermophiles; Um, Ustilago maydis; Zm, Zea mays. (Color figure online)
transaminase K (Cooper et al. 2008; Wogulis et al. 2008; Han et al. 2009). While all plant AAA-ATs belong to class I aminotransferases and are closely related with each other, the relative relationships among different AAA-AT types were not confidently resolved using different alignment and phylogenetic methods (not shown). Nevertheless, each AAA-AT type forms a well-supported monophyletic clade, allowing us to predict different types of plant AAAATs based on gene and protein sequences.
Subcellular localizations of plant AAA-ATs Plastids are autonomous in producing all three AAAs, Phe, Tyr, and Trp (Fig. 1c) (Bickel et al. 1978; Bagge and Larsson 1986), and enzymes of the shikimate and
AAA pathways localize in the plastids (Jensen 1986; Tzin and Galili 2010; Maeda and Dudareva 2012), though some isoforms (e.g. chorismate mutase, Eberhard et al. 1996b) were found outside of the plastids. ADH and ADT enzymes involved in the final steps of plant Tyr and Phe biosynthesis, respectively (Fig. 1c), are mainly localized in the plastids based on subcellular fractionation and fluorescence protein-tagged protein localization (Jung et al. 1986; Rippert et al. 2009; Bross et al. 2017). Similar approaches as well as immunoblotting and chloroplast import assays also revealed that Trp biosynthetic enzymes (i.e. anthranilate synthase, phosphoribosylanthranilate transferase, phosphoribosylanthranilate isomerase, and Trp synthase) are plastidic in various plants (Last et al. 1991; Bohlmann et al. 1995; Zhao and Last 1996; Kriechbaumer et al. 2008). The plastid synthesized AAAs are then exported from the plastids and further utilized for the synthesis of proteins, hormones, and natural products in the cytosol and other compartments (Fig. 1c). Consistent with the involvement of PPA-ATs in the biosynthesis of Phe and Tyr in plants, PPA-ATs are localized in the plastids. Subcellular fractionation of sorghum leaves showed that PPA-AT activity is mainly detected in the plastid fraction (Siehl et al. 1986b), so is the PPA-AT activity in petunia flower and soybean leaf tissues (Yoo et al. 2013; Schenck et al. 2015). Arabidopsis PPA-AT (At2g22250) was enriched in the plastid fraction detected by immunoblotting (de la Torre et al. 2006). GFP-fused tomato PPA-AT was also targeted to the plastids, when expressed in the protoplasts of Nicotiana benthamiana (Dal Cin et al. 2011). In contrast to PPA-ATs, petunia PhPPY-AT is localized outside of the plastids, based on the localization of GFP-fused PhPPY-AT in Arabidopsis protoplasts as well as subcellular fractionation of PPY-AT activity from flower extracts (Yoo et al. 2013). Given that ADT homologs having some PDT activity from petunia and Arabidopsis are found in the plastids (Cho et al. 2007; Maeda et al. 2010), PPY, the product of PDT, is likely exported from the plastids to the cytosol, where PPY-AT converts PPY into Phe (Fig. 1c). Thus, the plastidic PDT and cytosolic PPYAT enzymes likely provide an alternative cytosolic pathway of Phe biosynthesis via PPY (Yoo et al. 2013). A plastid envelop-localized AAA transporter, which has transport activity towards Phe, Tyr and to a
123
Phytochem Rev Table 3 List of aminotransferase sequences used in the phylogenetic analysis of Fig. 2 Enzyme name
Plant species (common name)
Accession number (PDB ID)
References
Ab-ArAT4
Atropa belladonna (deadly nightshade)
KC954706
Bedewitz et al. (2014)
AmPPA-AT
Antirrhinum majus (snapdragon)
AII23743
Dornfeld et al. (2014)
AtPPA-AT
Arabidopsis thaliana
AT2G22250.2
Graindorge et al. (2010), Dal Cin et al. (2011), Maeda et al. (2011)
AtTAA1
Arabidopsis thaliana
AT1G70560.1 (3BWN)
Stepanova et al. (2008), Tao et al. (2008)
AtTAR1
Arabidopsis thaliana
AT1G23320.1
Stepanova et al. (2008)
Plant AAA-ATs
AtTAR2
Arabidopsis thaliana
AT4G24670.1
Stepanova et al. (2008), Won et al. (2011)
AtTAT1
Arabidopsis thaliana
AT5G53970.1
Riewe et al. (2012), Wang et al. (2016)
AtTAT2
Arabidopsis thaliana
AT5G36160.1
Prabhu and Hudson (2010), Wang et al. (2016)
AtVAS1
Arabidopsis thaliana
AT1G80360.1
Zheng et al. (2013)
EsAroAT1
Ephedra sinica
AGK24944
Kilpatrick et al. (2016)
EsAroAT2
Ephedra sinica
AGK24945
Kilpatrick et al. (2016)
CmArAT1 OsTAR2
Cucumis melo (melon) Oryza sativa (rice)
ADC45389 BAS70606
Gonda et al. (2010) Yoshikawa et al. (2014)
PhPPA-AT
Petunia hybrida (petunia)
ADM67557
Maeda et al. (2011)
PhPPY-AT
Petunia hybrida (petunia)
KF511589
Yoo et al. (2013)
PpPPA-AT
Pinus pinaster (maritime pine)
CAF31327
de la Torre et al. (2007)
PsTyrAT
Papaver somniferum (opium poppy)
ADC33123
Lee and Facchini (2011)
RyAAAT3
Rosa x damascene (rose)
XP_011457732
Hirata et al. (2012)
SlGTK
Solanum lycopersicum (tomato)
XP_004250335
Ellens et al. (2015)
SlPAT
Solanum lycopersicum (tomato)
ADQ00383
Dal Cin et al. (2011)
ZmGTK
Zea mays (maize)
NP_001143647
Ellens et al. (2015)
ZmTAR1
Zea mays (maize)
ACG56678
Chourey et al. (2010)
ZmVT2
Zea mays (maize)
DAA34789
Phillips et al. (2011)
Mammalian AAA-ATs HsKAT-I
Homo sapiens (human)
NP_004050 (1W7L)
Rossi et al. (2004), Han et al. (2010)
HsKAT-II
Homo sapiens (human)
NP_872603 (2QLR)
Rossi et al. (2008), Han et al. (2010)
HsTAT MmKAT-III
Homo sapiens (human) Mus musculus (mouse)
P17735 (3DYD) AAQ15190 (2ZJG)
Rettenmeier et al. (1990) Han et al. (2009)
MmKAT-IV
Mus musculus (mouse)
P05202
Han et al. (2011)
MmTAT
Mus musculus (mouse)
NP_666326 (3PDX)
Mehere et al. (2010)
Microbial enzymes with AAA-AT activities AaAAA-AT
Alicyclobacillus acidocaldarius
ZP_03494134
Dornfeld et al. (2014)
AsTdiD
Aspergillus sp.
ABU51605
Balibar et al. (2007)
BaBCAT2
Bacillus anthracis
AF527044
Berger et al. (2003)
BcBCAT2
Bacillus cereus
AF527043
Berger et al. (2003)
CtPPA-AT
Chlorobium tepidum
NP_661859
Dornfeld et al. (2014)
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Phytochem Rev Table 3 continued Enzyme name
Plant species (common name)
Accession number (PDB ID)
References
Dornfeld et al. (2014)
DdAspAT
Dickeya dadantii
ACS85258
EcAspC
Escherichia coli
EG10096 (1AAM)
Gelfand and Steinberg (1977), Almo et al. (1994)
EcIlvE
Escherichia coli
EG10497 (1A3G)
Gelfand and Steinberg (1977), Okada et al. (1997)
EcTyrB
Escherichia coli
EG11040 (3TAT)
Gelfand and Steinberg (1977), Ko et al. (1999)
KpTyrB
Klebsiella pneumoniae
AF074934
Heilbronn et al. (1999)
LlAraT
Lactococcus lactis
AAF06954
Rijnen et al. (1999)
MfTam1
Malassezia furfur
JX453496
Preuss et al. (2013)
MtPPA-AT/ DAPAT NePPA-AT/ DAPAT
Mycobacterium tuberculosis
O50434
Graindorge et al. (2014)
Nitrosomonas europaea
Q82S89
Graindorge et al. (2014)
SaPPA-AT/ DAPAT
Streptomyces avermitilis
Q82IK5
Graindorge et al. (2014)
SbAAA-AT
Streptomyces bingchenggensis Saccharomyces cerevisiae (yeast)
ADI11720
Dornfeld et al. (2014)
YGL202 W
Urrestarazu et al. (1998)
ScARO8 ScARO9
Saccharomyces cerevisiae (yeast)
YHR137 W
Urrestarazu et al. (1998)
ScKAT
Saccharomyces cerevisiae (yeast) Synechococcus sp. CC9605
NP_012475 (3B46)
Wogulis et al. (2008)
P54691
Graindorge et al. (2014)
SspPPA-AT/ PPA-AT
Synechocystis sp PCC6803
AGF52553
Dornfeld et al. (2014)
SspPPA-AT/ BCAT TcTAT
Synechocystis sp PCC6803
Q3AJX2
Graindorge et al. (2014)
Trypanosoma cruzi
AAA02975 (1BW0)
Nowickia et al. (2001)
TtPPA-AT
Thermus thermophilus
YP_143312
Dornfeld et al. (2014)
UmTam1
Ustilago maydis
XP_011387757
Zuther et al. (2008)
UmTam2
Ustilago maydis
XP_011389975
Zuther et al. (2008)
SscPPA-AT/ BCAT
Arabidopsis C–S lyase, Asp-AT, BCAT homologs CORI3
Arabidopsis thaliana
AT4G23600.1
Jones et al. (2003)
SUR1
Arabidopsis thaliana
AT2G20610.1
Mikkelsen et al. (2004), Klein and Sattely (2017)
–a
Arabidopsis thaliana
AT4G23590.1
Uncharacterized
b
–
Arabidopsis thaliana
AT2G24850.1
Uncharacterized
–
Arabidopsis thaliana
AT4G28410.1
Uncharacterized
–
Arabidopsis thaliana
AT4G28420.2
Uncharacterized
–
Arabidopsis thaliana
AT1G77670.1
Uncharacterized
BCAT1
Arabidopsis thaliana
AT1G10060.2
Diebold et al. (2002)
BCAT2
Arabidopsis thaliana
AT1G10070.1
Diebold et al. (2002)
BCAT3 BCAT4
Arabidopsis thaliana Arabidopsis thaliana
AT3G49680.1 AT3G19710.1
Diebold et al. (2002), Knill et al. (2008) Schuster et al. (2006)
BCAT5
Arabidopsis thaliana
AT5G65780.1
Diebold et al. (2002)
BCAT6
Arabidopsis thaliana
AT1G50110.1
Diebold et al. (2002), La¨chler et al. (2015)
BCAT7
Arabidopsis thaliana
AT1G50090.1c
Binder (2010)
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Phytochem Rev Table 3 continued Enzyme name
Plant species (common name)
Accession number (PDB ID)
References
AspAT1
Arabidopsis thaliana
AT4G31990.3
Schultz and Coruzzi (1995), Schultz et al. (1998), Wilkie and Warren (1998)
AspAT2
Arabidopsis thaliana
AT5G19550.1
Schultz and Coruzzi (1995), Schultz et al. (1998), Wilkie and Warren (1998)
AspAT3
Arabidopsis thaliana
AT5G11520.1
Schultz and Coruzzi (1995), Schultz et al. (1998)
AspAT4
Arabidopsis thaliana
AT1G62800.2
Schultz and Coruzzi (1995)
AspAT aspartate aminotransferase, AraT aromatic amino acid aminotransferase, BCAT branched-chain amino acid aminotransferase, C–S lyase carbon-sulfur lyase, DAPAT diaminopimelate aminotransferase, KAT/GTK Kynurenine aminotransferase/glutamine transaminase K, Phe-AT Phe aminotransferase, PPA-AT prephenate aminotransferase, Tam Trp amino transferase, Trp-AT Trp aminotransferase, Tyr-AT Tyr aminotransferase a
Not designated yet as they have not been characterized
b
Co-expressed with other AAA network genes
c
Not expressed (likely pseudogene)
lesser extent Trp, has been recently identified in petunia (Widhalm et al. 2015). However, PPY was not tested as a substrate and a plastid transporter(s) responsible for the PPY export is currently unknown. Arabidopsis AtTAT1 and AtTAT2 localize outside the plastids (Wang et al. 2016). Based on the lack of N-terminal plastid transit peptide, other Phe-/TyrATs, such as Ab-ArAT4 of deadly nightshade (Bedewitz et al. 2014), are also likely to be extra-plastidic. Subcellular fractionation of Arabidopsis leaf protein extracts, however, detected substantial Tyr-AT activity also within the plastid fraction (Wang et al. 2016), suggesting that there is an unknown aminotransferase(s) having Tyr-AT activity in the plastids as well. Interestingly, many plant Phe-/Tyr-ATs, despite their localization outside the plastids, have the pH optima at the basic region in the vicinity of 8 and even 9 (Tables 1, 2), although its physiological reason is unknown. HPP dioxygenase (HPPD), which converts HPP into homogentisate, the precursor of tocochromanols and plastoquinone (Norris et al. 1998, Fig. 1c), localizes in the cytosol in carrot (Daucus carota) (Garcia et al. 1997) and Arabidopsis (Wang et al. 2016). Homogentisate is also an intermediate of the Tyr degradation pathway in plants (Dixon and Edwards 2006) and the subsequent enzyme, homogentisate dioxygenase (HGO), also localizes to the cytosol in soybean (Stacey et al. 2016). Thus, homogentisate is mostly synthesized in the cytosol (Fig. 1c). Given that
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enzymes directly involved in tocochromanol and plastoquinone biosynthesis are plastidic (Soll and Schultz 1980; Soll et al. 1985; Schulze-Siebert et al. 1987; Joyard et al. 2009; Yang et al. 2011; Mehrshahi et al. 2014), homogentisate should be transported into the plastids with an unknown transporter(s). In spinach leaves, however, HPPD activity was enriched in the plastid fraction (Fiedler et al. 1982). Also, maize HPPD is mainly localized in the plastids (Siehl et al. 2014). On the other hand, soybean HPPD enzyme is dual localized in the cytosol and plastids due to alternative splicing of the N-terminal transit peptide (Siehl et al. 2014). Thus, depending on HPPD localization, the identity and the role of the plastidic Tyr-AT activity needs to be further investigated in different plants to clarify relative contributions of cytosolic vs. plastidic supply of HPP. Some legumes such as soybean have, besides the plastidic ADHs, a cytosolic PDH enzyme(s) that converts prephenate into HPP (Rubin and Jensen 1979; Schenck et al. 2015; 2017). Also, all plants have at least one cytosolic isoform of chorismate mutases, which convert chorismate into prephenate (Fig. 1c, Benesova and Bode 1992; Eberhard et al. 1996a, b; Colquhoun et al. 2010; Westfall et al. 2014). Although most Tyr-AT activity and enzymes prefer Tyr deamination to HPP, AtTAT2 favors HPP transamination to synthesize Tyr (Prabhu and Hudson 2010; Wang et al. 2016). Thus, some of the AtTAT2 orthologs from legumes may efficiently transaminate HPP into Tyr
Phytochem Rev
and, together with cytosolic CM and PDH enzymes, allow legumes to synthesize Tyr within the cytosol. Subcellular fractionation studies showed that LTrp-AT activity is detected in all fractions (i.e. plastid, cytosol, mitochondria) of P. sativum etiolated seedlings, while D-Trp-AT activity was enriched in the plastid fraction of dark-grown P. sativum seedlings (McQueen-Mason and Hamilton 1989). The L-Trp-AT activity detected in spinach leaves was enriched in the peroxisome fraction but had tenfold higher serineglyoxylate aminotransferase activity (Noguchi and Hayashi 1980) that are likely involved in photorespiration (Liepman and Olsen 2001). Maize ZmTAR1 fused with GFP and expressed in tobacco leaves mainly colocalized with an ER luminal marker (Kriechbaumer et al. 2015). Also, cytosolic and microsomal fractions of maize coleoptile extracts accounted for *90% of total activity converting Trp to IAA (Kriechbaumer et al. 2015). Arabidopsis VAS1, which converts IPA back to Trp while deaminating Met (Fig. 1c), was also shown to localize outside of the plastids through GFP localization (Zheng et al. 2013). Early subcellular fractionation studies also showed that the two enzymes involved in ethylene biosynthesis, S-adenosyl methionine synthase, and 1-aminocyclopropane-1-carboxylic acid synthase, are cytosolic (Boller et al. 1979; Wallsgrove et al. 1983). Thus, L-Trp-ATs and other enzymes involved in auxin and ethylene biosynthesis are localized outside of the plastids (Fig. 1c), though their exact localizations (e.g. cytosolic or microsomal) remain to be examined.
In planta functions of AAA-ATs To discuss physiological functions of different AAAATs in various plant species, this section summarizes expression profiles and genetic characterizations of AAA-ATs in different plants. Gene expression and activity profiles of AAA-ATs in plants All three AAA-AT activity (i.e. Phe-AT, Tyr-AT, and Trp-AT) were found to peak together in light-grown P. vulgaris shoots, cotyledons, and roots at 4, 6, and 8-day-old, respectively (Forest and Wightman 1972). Taylor and Wightman (1987) also found that Phe-AT
activity of P. vulgaris shoots and roots peaks at 10 days after germination and decreases thereafter. The highest Tyr-AT and Phe-AT activities were together observed after 9-day growth of organ-forming rice callus culture (Kishor 1989). These results suggest that AAA-AT activities generally increase during early seedling development likely to support active synthesis of proteins required for growth. In A. officinalis cell culture, Tyr-AT activity, especially of Tyr-AT1 isoform, increased together with the accumulation of rosmarinic acid during the cell growth (De-Eknamkul and Ellis 1987). In the cell suspension and hairy root cultures of rosmarinic acidproducing Lamiaceae plants (Orthosiphon aristatus, Lithospermum erythrohizon, and Salvia miltiorrhiza), Tyr-AT activity increased upon different elicitors treatments (i.e. methyl jasmonate, yeast extract or silver ion), which enhance the production of rosmarinic acid (Sumaryono et al. 1991; Mizukami et al. 1993; Yan et al. 2006). An AtTAT1 homolog of red sage (Salvia miltiorrhiza, also known as danshen, a Chinese medicinal plant) is also upregulated upon treatments with salicylic acid, methyl jasmonate, abscisic acid, and UV-B (Huang et al. 2008). These results are consistent with the involvement of Tyr-AT in rosmarinic acid biosynthesis (Petersen and Simmonds 2003). Spatial and temporal expressions of Tyr-AT and Phe-AT genes highly correlate with the production of downstream specialized metabolites derived from Tyr and Phe. The expression of melon AAA-AT, CmArAT1, increases during the ripening stage of climacteric cultivars, when biosynthesis of Phe-derived volatile esters becomes active (Gonda et al. 2010). Petunia PhPPY-AT expression increases towards the end of the day in flowers when Phe-derived benzenoid volatile production is elevated (Yoo et al. 2013). EsAroAT1 and EsAroAT2 of E. sinica are expressed in young stems, where ephedrine alkaloids accumulate (Kilpatrick et al. 2016). Similarly, ArAT4 from deadly nightshade is co-expressed with tropane alkaloid biosynthetic genes especially in secondary roots where high levels of alkaloids accumulate (Bedewitz et al. 2014). Arabidopsis AtTAT1 is upregulated upon various abiotic stress conditions and coronatine treatment (Holla¨nder-Czytko et al. 2005; Less and Galili 2008) and is most strongly co-expressed with HPPD (Wang et al. 2016), which together are required for tocochromanol and plastoquinone biosynthesis.
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Because Tyr- and Phe-AT enzymes are usually not very efficient (Tables 1, 2) and likely rate-limiting enzymes of biosynthetic pathways of specialized metabolites, transcriptional regulations of these Tyrand Phe-AT genes appear to be the major mechanism to efficiently turn on and off the utilization of the Tyr and Phe precursors to their downstream pathways (Less and Galili 2008, 2009). The relationship between Trp-AT activity and auxin biosynthesis has been investigated in various plant species. L-Trp-AT activity and IAA content rapidly increase together and peak at 12 h after root excision in the epicotyl of pea seedlings (Minh et al. 1984). In tobacco cell culture, on the other hand, LTrp-AT activity continues to increase during maturation, whereas IAA production peaks around 20 days (El Bahr et al. 1987). The expression of Arabidopsis TAA1 as well as IAA level are both higher in the shade than under normal light (Tao et al. 2008) and also higher at 28 °C than 20 °C (Franklin et al. 2011). Expressions of OsTAR1 and OsTAR2, rice orthologs of Arabidopsis TAR1, increase during rice grain development together with the large increase of IAA production (Abu-Zaitoon et al. 2012). Thus, IAA production is in general positively correlated with TrpAT expression, consistent with the major role of the Trp-AT-mediated, IPA-dependent pathway in auxin biosynthesis (Zhao 2012, 2014). D-Trp-AT activity increases during seedling growth of wheat, maize, and pea (McQueen-Mason and Hamilton 1989; Koshiba et al. 1993; Rekoslavskaya et al. 1999). On the other hand, the gene expression of D-amino acid aminotransferase (At5g57850) that exhibits some D-Trp-AT activity (Funakoshi et al. 2008) is enriched in root quiescent center (Nawy et al. 2005) and overall evenly expressed throughout the development and tissues of Arabidopsis (eFP browser, Winter et al. 2007). Thus, At5g57850 is unlikely to be responsible for the D-Trp-AT activity detected in various seedlings. A D-Trp conjugate, N-malonyl-Dtryptophan, accumulates under germinating and stressed conditions in multiple plant species, and is considered to be the source of D-Trp (Zenk and Scherf 1963, 1964; Robinson 1976; Rekoslavskaya 1986, 1988; Gamburg et al. 1993). Furthermore, Trp racemase (EC 5.1.1.10) activities interconverting DTrp and L-Trp were also detected in seedlings of wheat and P. sativum, and tobacco cell culture (Miura and Mills 1971; Law 1987; Rekoslavskaya et al. 1999).
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However, it is still unclear how D-Trp is synthesized in plants and how much D-Trp contributes to the production of IPA-derived metabolites including auxin. It will be also important to analyze endogenous D-Trp levels especially during early seedling development when D-Trp-AT activity is detectable and the demand of L-Trp for protein and auxin biosynthesis is especially high. PPA-AT is co-expressed with many genes involved in the shikimate pathway (e.g. 3-deoxy-7-phosphoheptulonate synthase), Phe biosynthesis (e.g. ADT), and the phenylpropanoid pathways (e.g. Phe ammonia-lyase, PAL) in Arabidopsis (Maeda et al. 2011). Expressions of PPA-AT and PAL are co-regulated by Myb8 transcription factor in maritime pine (CravenBartle et al. 2013). Also, PPA-AT expression is increased after anthesis and at night in petunia flower, when biosynthesis of benzenoid and phenylpropanoid volatiles is induced (Maeda et al. 2011). Heterologous expression of Arabidopsis MYB12 or petunia ODORANT1 transcription factor in tomato fruits also led to induction of PPA-AT as well as other shikimate, Phe and phenylpropanoid pathway genes (Dal Cin et al. 2011; Zhang et al. 2015). Thus, the coordinated transcriptional regulation of PPA-AT together with its upstream and downstream genes is important for biosynthesis of Phe and Phe-derived natural products. Genetic characterization of AAA-ATs in plants In vivo functions of AAA-ATs have been investigated through genetic suppression of various AAA-AT genes in planta. RNA interference (RNAi) suppression of PPA-AT led to decreased Phe and increased prephenate levels in petunia flowers, providing genetic evidence that PPA-AT is responsible for Phe biosynthesis via the arogenate intermediate in this tissue (Maeda et al. 2011). PPA-AT down-regulation in Nicotiana benthamiana via virus-induced gene silencing resulted in lower lignin content and dwarf phenotype, though the levels of Phe was unexpectedly increased and asparagine was decreased (de la Torre et al. 2014). Given that PPA-AT can efficiently transaminate a-ketoglutarate and oxaloacetate, besides prephenate, in vitro (Rubin and Jensen 1979; Bonner and Jensen 1985; Siehl et al. 1986a; DeEknamkul and Ellis 1988; Graindorge et al. 2010; Dal Cin et al. 2011; Maeda et al. 2011), the conversion between a-ketoglutarate/oxaloacetate and Glu/Asp
Phytochem Rev
(i.e. Asp-AT activity) that is coupled to prephenate transamination may have significant impacts on overall amino acid metabolism in the plastids (Fig. 1c). Since significant carbon flux goes through Phe biosynthesis and metabolism to phenylpropanoids (e.g. lignin) in plants, it is important to understand how ammonia released in the cytosol by PAL in most plants (Zhang and Liu 2015) or Tyr ammonia-lyase, TAL, in grasses (Barros et al. 2016) will be recycled back to the prephenate transamination reaction catalyzed by PPA-AT in the plastids (Razal et al. 1996). Genetic characterizations of Tyr/Phe-ATs are overall consistent with their biochemical characterizations and further support that directionally preferred reactions of Tyr/Phe-ATs observed in vitro (Sect. 2) play physiological roles in planta by providing precursors for the synthesis of various downstream specialized metabolites. Virus-induced gene silencing of PsTyrAT in opium poppy resulted in at least 80% transcript reduction and almost 50% reduction of total benzylisoquinoline alkaloids (Lee and Facchini 2011), suggesting that PsTyrAT is involved in the initial step of benzylisoquinoline alkaloid biosynthesis from Tyr (Jindra et al. 1966; Rueffer and Zenk 1987), though some redundancy may still exist. RNAi suppression of PhPPY-AT in petunia flowers led to decline in the levels of Phe and benzenoid volatile compounds, suggesting that the cytosolic PPY-AT enzyme is involved in the conversion of PPY into Phe and Phederived compounds (Yoo et al. 2013). Similarly, RNAi silencing of Ab-ArAT4 by *60% in deadly nightshade led to more than 50% reduction in tropane alkaloids, together with 90% reduction in phenyllactic acid, suggesting that the main role of the Ab-ArAT4 is to deaminate Phe into PPY in the cytosol for tropane biosynthesis via phenyllactic acid (Bedewitz et al. 2014). Transient RNAi silencing of RyAAAT3 in rose protoplasts resulted in reduced 2-phenylethanol level, suggesting that PPY synthesized from Phe by PyAAAT3 is used for the formation of 2-phenylethanol volatile (Hirata et al. 2012, 2016). T-DNA knockout lines of Arabidopsis AtTAT1 show higher Tyr and lower tocopherol levels than wild type (Riewe et al. 2012), providing genetic evidence that AtTAT1 plays the major role in converting Tyr to HPP for tocopherol biosynthesis. Consistently, bleaching phenotypes of Arabidopsis and duckweed (Lemna paucicostata) treated by cinmethylin and 5-benzyloxymethyl-1,2-isoxazolines herbicides,
which inhibit AtTAT1, can be rescued by HPP supplementation (Grossmann et al. 2012). Forward genetics studies identified Trp-ATs as essential components in auxin biosynthesis in various plants (Stepanova et al. 2008; Tao et al. 2008; Won et al. 2011; Yoshikawa et al. 2014). Arabidopsis TAA1 knock-out mutants do not exhibit rapid accumulation of IAA in response to the shade and hence lack shade avoidance response (Tao et al. 2008). Under normal light conditions, the taa1 single mutant still has wildtype level of IAA, but IAA level decreased in the taa1 tar2 double mutants (Stepanova et al. 2008). Also, the taa1 tar2 double and taa1 tar1 tar2 triple mutants showed retarded plant growth and auxin-related defects in embryo development, respectively (Stepanova et al. 2008). Transient overexpression of ZmTAR1 in tobacco enhanced IAA accumulation, whereas knock-out mutants of maize ZmVT2 and rice OsTAR2 reduced IAA content and led to abnormal auxin-related phenotypes (Chourey et al. 2010; Phillips et al. 2011; Yoshikawa et al. 2014). These genetic data support the major and redundant roles of closely-related TAA1 and TARs (Fig. 2) in catalyzing the committed step of the IPA-dependent auxin biosynthetic pathway in plants (Zhao 2012, 2014). Knockout mutants of VAS1 showed higher IPA and IAA levels than wild type under both normal light and shade, while VAS1 overexpression led to reduced seed set and IAA accumulation under shade, consistent with the preference of VAS1 for IPA transamination to Trp (Zheng et al. 2013). Expression patterns of bglucuronidase (GUS) reporters fused with the VAS1 and auxin-responsive DR5 promoters showed that VAS1 expression coincides with auxin accumulation. The levels of 1-aminocyclopropane-1-carboxylic acid, the key intermediate of ethylene from Met (Fig. 1c), were also increased in the vas1 mutant, in agreement with efficient deamination of Met into 2-oxo-4methylthiobutyric acid by VAS1 (Zheng et al. 2013). The opposite activity to transaminate 2-oxo-4methylthiobutyric acid to Met in the Yang cycle (Fig. 1c) is instead catalyzed by glutamine transaminase K orthologs (Ellens et al. 2015) and possibly cytosolic BCATs (e.g. Arabidopsis AtBCAT4 and 6, Schuster et al. 2006; La¨chler et al. 2015). Notably, several other AAA-ATs, besides VAS1, also showed comparable activities towards Met or 2-oxo-4methylthiobutyric acid to aromatic amino or keto acid substrates (Matheron and Moore 1973; Kilpatrick
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Phytochem Rev
et al. 2016; Wang et al. 2016), though their physiological roles linking AAA and Met metabolism have not been investigated. These results revealed key roles of VAS1 in coordinating the production of two classical plant hormones, auxin and ethylene (Zheng et al. 2013, Fig. 1c).
Conclusions and open questions In summary, plant AAA-ATs play crucial roles in both biosynthesis and metabolism of AAAs. In plants, PPA-ATs are responsible for transamination of prephenate to arogenate, the common precursor of Tyr and Phe biosynthesis, within the plastids (Fig. 1c). Plant PPA-ATs so far characterized belong to the class Ib Asp-AT clade, but the role of redundant PPA-AT enzymes needs to be further investigated, some of which may be derived from side activity of other aminotransferases, such as weak PPA-AT activity of AtTAT2 (Wang et al. 2016) or orthologs of cyanobacteria class IV aminotransferases having PPA-AT activity (Graindorge et al. 2014). Considering the significant carbon flow passing through the PPA-ATcatalyzed reaction in vascular plants, it is also important to investigate the physiological significance of Asp-AT activity coupled with PPA-AT activity in planta. A potential knockout mutant of AtPPA-AT was reported among 130 Arabidopsis mutants defective in female gametophyte development (maternal effect embryo arrest, mee17) (Pagnussat et al. 2005), but still requires genetic rescue or independent alleles to verify the in vivo function of plant PPA-ATs. Plant Tyr/Phe-ATs, which generally have broad substrate specificity, are responsible for producing precursors of various specialized metabolites derived from Phe, Tyr, PPY, or HPP in various plants. Interestingly, Tyr deamination to HPP is often efficiently coupled with PPY transamination to Phe or vice versa (Fig. 1c) and could be a common feature of AtTAT1 orthologs (Fig. 2), though its physiological significance remains to be examined. Also, contributions of redundant AAA-ATs or other classes of aminotransferases having moonlight Tyr/Phe-AT activity remain to be investigated. Trp-ATs are involved in the production of auxin and ethylene through the fine balancing of Trp deamination (catalyzed by TAA1 and likely TARs) and IPA transamination to Trp (catalyzed by VAS1),
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while the latter is coupled to deamination of Met, the precursor of ethylene (Fig. 1c). It remains to be explored, however, whether Trp-ATs is also involved in the production of IPA-derived metabolites besides auxin, such as indoleacetaldehyde, tryptophol, and their derivatives, detected in various plants (Helianthus annus, P. sativum, Cucumis sativus and Euglena gracilis) (Rajagopal 1967; Rayle and Purves 1967; Rajagopal 1968; Lacan et al. 1984, 1985). Also, the genes and enzymes responsible for D-Trp-AT activity, detected from many seedlings, have not been determined nor their contributions to the production IPA-derived metabolites including auxin in different plants, tissues, and developmental stages. Unexpected couplings of Phe and Tyr metabolism, as well as Trp and Met transamination reactions, suggest that aminotransferases could mediate ‘‘metabolic interlock’’ (Jensen 1969) or ‘‘cross-pathway regulation’’ (Guyer et al. 1995) of different amino acid pathways at the enzyme levels. Thus, rigorous testing of substrate specificity of both amino donors and keto acceptors can potentially identify novel metabolic links between different amino acid metabolic pathways. Detailed characterization of various AAA-ATs guided by their phylogenetic relationships, as was employed to identify key residues of PPA-AT substrate specificity (Dornfeld et al. 2014), can potentially identify residues responsible for the substrate specificity of other AAAATs. Also, structural analyses of plant AAA-ATs will be necessary to uncover molecular mechanisms underlying the extensive diversification of plant AAA-ATs (Fig. 2). Understanding of plant AAA-ATs at the molecular level will allow us to better predict functionality of aminotransferases having different AAA-AT activities and also to redesign their substrate specificity and metabolic function in planta. Acknowledgements This work was supported by the IOS1354971 grant from the US National Science Foundation and the Agriculture and Food Research Initiative competitive Grant (2015-67013-22955) from the USDA National Institute of Food and Agriculture to H.A.M.
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