Molecular and Structural Perspectives on Cytochrome P450s in Plants

Molecular and Structural Perspectives on Cytochrome P450s in Plants

MARY A. SCHULER1 AND SANJEEWA G. RUPASINGHE

Departments of Cell and Developmental Biology, Biochemistry and Plant Biology, University of Illinois, Urbana, Illinois, USA

I. Biochemical Diversity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. General Considerations........................................................ B. Intracellular Locations and Electron-Transfer Partners.................. II. Molecular Diversity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Gene Nomenclature and Numbers .......................................... B. Conserved Gene Families ..................................................... C. Restricted Gene Families ...................................................... III. Structural Diversity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Structural Determinants in Classical P450s ................................ B. Structural Determinants in Nonclassical P450s............................ IV. Comparisons Between Oryza and Arabidopsis P450s . . . . . . . . . . . . . . . . . . . . . . . A. Phylogenetic Relationships Among Functionally Defined P450s ....... B. Structural Similarities in Highly Conserved P450s ........................ C. Structural Perspectives on Moderately Conserved P450s ................ V. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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ABSTRACT From a biochemical perspective, the ability to incorporate oxygen at very specific points in a substrate’s structure is essential to numerous synthetic and catabolic plant pathways that involve simple alkyl and aromatic hydroxylations or more complex 1

Corresponding author: E-mail: [email protected]

Advances in Botanical Research, Vol. 60 Copyright 2011, Elsevier Ltd. All rights reserved.

0065-2296/11 $35.00 DOI: 10.1016/B978-0-12-385851-1.00005-6

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epoxidations, aryl migrations, decarboxylations and carbon–carbon bond cleavages. Because of the restricted substrate specificities of many cytochrome P450 monooxygenases and the chemical versatility of the entire group of P450s existing in plants, significant interest exists in defining functions for those not yet characterized, in understanding catalytic-site constraints of those in biologically important pathways and in modifying them for crop improvement and biopharmaceutical production. From genomics and structural perspectives, these goals are especially challenging since P450 gene families have duplicated and diverged to unprecedented degrees as new plant pathways have evolved for the synthesis of defence toxins and other secondary metabolites. Phylogenetic comparisons based solely on primary sequences have facilitated grouping of the many, often divergent, sequences into families and subfamilies that can be compared within and between plant species. Within these individual groupings, mapping of variations that have accumulated in different regions of P450 proteins has shown that they can have drastically different effects on enzymatic functions with some affecting substrate recognition, others affecting interactions with electron-transfer partners and others not affecting activity at all. Bringing together biochemical, molecular and structural perspectives, it is now becoming possible to provide cohesive models of many functionally characterized P450s and to extend these models to related but uncharacterized P450s. Using examples taken from the sequenced Arabidopsis thaliana and Oryza sativa genomes and some functionally characterized P450s in other plant species, this review highlights their phylogenetic relationships and predicted structural similarities and differences that are likely or not likely to affect catalytic functions. Combinations of primary and tertiary structure analyses such as these can now allow researchers to better understand the evolutionary relationships among plant P450s in secondary metabolic pathways and assign tentative functions to those not yet functionally characterized. With the number of annotated plant P450 sequences exponentially increasing as genomes for medicinally important plants are being sequenced, these dual level assessments will become increasingly important for discriminating among the P450s needing to be functionally characterized.

I. BIOCHEMICAL DIVERSITY A. GENERAL CONSIDERATIONS

Data mining in plant genomes invariably results in the identification of cytochrome P450 monooxygenase (P450) genes whose structural and biochemical diversities perplex the uninitiated. Contributing to the extensive evolution of this gene superfamily is the enormous diversity of plant compounds that are synthesized or catabolized by the introduction of molecular oxygen into their structures. Examples of those synthesized include the numerous natural products that serve as defences against insect and vertebrate predators, fungal infections and bacterial infections (e.g. alkaloids, terpenoids, furanocoumarins, glucosinolates, cyanogenic glucosides; Croteau et al., 2000; Halkier and Gershenzon, 2006; Schuler, 2011),

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signalling molecules (e.g. jasmonic acid, brassinolide, gibberellins; Crozier et al., 2000), phenylpropanoids (e.g. flavonoids, anthocyanins, lignin monomers, isoflavonoids; Vogt, 2010), carotenoids (Dellapenna and Pogson, 2006), fatty acids and sterols (Pinot and Beisson, 2011). Examples of those catabolized include signalling molecules (e.g. abscisic acid, brassinolide; Crozier et al., 2000) and herbicides (Siminszky, 2006). Like most mammalian P450s, the vast majority of plant P450s mediating the aliphatic hydroxylations, aromatic hydroxylations, epoxidations, carboxylations and other oxygenations occurring on these substrates utilize molecular oxygen and electrons transferred from one or more transfer partners (Ortiz de Montellano, 2005; Sigel et al., 2007; Werck-Reichhart et al., 2006). A distinctly smaller (and more unusual) group of plant P450s mediate oxygen activations in the absence of molecular oxygen and/or electrontransfer partners and result in methylenedioxy bridge formation, phenolic couplings, carbon–carbon bond cleavages and the dimerization or oxidative rearrangement of carbon skeletons (Mizutani and Sato, 2010; Ortiz de Montellano and Nelson, 2011).

B. INTRACELLULAR LOCATIONS AND ELECTRON-TRANSFER PARTNERS

Most of the classical oxygen-requiring plant P450s are anchored in the endoplasmic reticulum (ER) with their N-terminal signal anchor sequence (SAD) buried in the lipid bilayer and their catalytic domain in the cytosol. These are equipped to utilize electrons transferred from NADPH via cytochrome P450 reductase (CPR) and, sometimes, NADH via cytochrome b5 (cyt b5) and cytochrome b5 reductase (Cb5R). Examples of many of these will be described below. Substantially fewer classical oxygen-requiring plant P450s are soluble in the chloroplast and equipped to utilize electrons transferred from NADPH via ferrodoxin (Fd) and ferrodoxin reductase (FdR). Examples of these include members of the CYP97 family that hydroxylate carotenoid ring structures (Dellapenna and Pogson, 2006). The more unusual nonclassical P450s, which fail to incorporate molecular oxygen into their final product or fail to use flavoproteins for dioxygen activation, are also situated in different cellular locales. Members of the CYP74 family (e.g. allene oxide synthase, hydroperoxide lyase, divinyl ether synthase) are localized in the chloroplast and are examples in which the standard P450 catalytic cycle is short-circuited by the direct activation of their hydroperoxide substrates (Brash, 2009; Stumpe and Feussner, 2006). Members of the CYP80 family (e.g. berbamunine synthase) are localized in the ER membrane and are examples in which oxygen present in one substrate

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molecule is coupled to carbon in another substrate utilizing the standard P450 catalytic cycle (Kraus and Kutchan, 1995; Mizutani and Sato, 2010). Complicating descriptions of interactions between plant P450s and their electron-transfer partners, the currently available sequences of higher plant genomes have shown that many contain multiple CPR Proteins (up to three in Populus trichocarpa (poplar) and Oryza sativa (rice)) and cyt b5 proteins (six in Arabidopsis thaliana (mouse ear’s cress)) and a few Cb5R Proteins (one in Arabidopsis, two in rice; Jensen and Møller, 2010; Paquette et al., 2009). Phylogenetic comparisons among the available plant CPR sequences have indicated that they fall into two distinct clusters with the CPR1 cluster restricted to dicots and the CPR2 cluster in both dicots and monocots (Jensen and Møller, 2010). The two A. thaliana CPR (designated ATR1, ATR2) are 63% identical to one another (Mizutani and Ohta, 1998; Urban et al., 1997) and representative of the CPR1 and CPR2 clusters (Jensen and Møller, 2010). Two of the three poplar CPR are 91% identical to one another (Ro et al., 2002) and representative of the CPR2 cluster; the third is 72% identical to the other two and representative of the CPR1 cluster. The three rice CPR are 62–79% identical to one another and representative of the CPR2 cluster. Heterologous expressions of the Arabidopsis and poplar CPR proteins in yeast have demonstrated that all are targeted to ER membranes with their cytosolic catalytic domains in orientations suitable for coupling with P450s (Ro et al., 2002; Urban et al., 1997). It is assumed that other plant CPR proteins are similarly localized even though the CPR sequences in current databases are most divergent in their N-terminal membrane-anchor sequences. Their extensive differences in this region have suggested that different CPR proteins associate with different sets of P450s in membrane-localized metabolons for the facilitated production of discrete components in individual pathways (Jensen and Møller, 2010). Support for this suggestion awaits reconstitution of membrane systems containing different P450 reductase and P450 components. Similar analysis of the Cb5R sequences has indicated that they are highly conserved with the two in rice being 85% identical to one another and 74–77% identical to the one in Arabidopsis. Analysis of the cyt b5 sequences has indicated that they are significantly less conserved with 35–67% identity among the five Arabidopsis cyt b5 members and many more divergent sequences in Oryza annotated as putative cyt b5 members. For these two groups of cyt b5 proteins, some of this variance may occur because of interactions that these proteins maintain for binding to other electron-transfer partners, such as fatty acid desaturases.

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II. MOLECULAR DIVERSITY A. GENE NOMENCLATURE AND NUMBERS

Acknowledging that the number of genes in the P450 superfamily is continually increasing as the more and more genome sequences are completed, a universal nomenclature system has evolved to categorize these proteins based on their phylogenetic relationships with other monooxygenases. In this system, families share greater than 40% amino acid identity and are designated with numbers (CYP1, CYP2, etc.), subfamilies share greater than 55% amino acid identity and are designated with alphabetical characters (A, B, C, etc.) and individual sequences are designated with a second set of numbers (CYP1A1, CYP1A2, CYP1A3, etc.; Nelson et al., 1993, 1996). In cases where allelic variants of individual loci are known, these are designated with yet another set of numbers (v1, v2, etc.). In sequenced higher plant genomes where P450 annotations have been completed, the complement of full-length P450 genes includes 142 in Carica papaya (papaya), 225 in Bracypodium distachyon (model wild grass), 245 in A. thaliana, 270 in Lycopersicon esculentum (tomato), 310 in P. trichocarpa (poplar), 316 in Vitis vinifera (grape), 334 in O. sativa (rice), 337 in Glycine max (soybean) and 399 in Solanum tuberosum (potato; Nelson, 2009; Nelson and Werck-Reichhart, 2011; Nelson et al., 2004, 2008; Paquette et al., 2000, 2009). Additional P450 annotations are in progress for the sequenced Zea mays, Sorghum bicolor and Jatropha curcas genomes. In sequenced lower plant genomes where P450 annotations are finished, the full-length P450 gene complement includes 71 in Phycomistrella patens (moss) and 227 in Selanginella moellendorffii (lycopod moss; Nelson and Werck-Reichhart, 2011). As in the Arabidopsis genome (Paquette et al., 2000; Werck-Reichhart et al., 2002), many of these full-length P450 genes are organized in long tandem arrays of related P450s from the same subfamily and, sometimes, are interspersed with P450s from more divergent families and subfamilies. In addition to these full-length genes, numerous non-functional P450 pseudogenes (e.g. prematurely truncated coding sequences, exon fragments) are scattered throughout these plant genomes and tandem arrays of functional genes. Comparisons of the gene structures in individual P450 families and subfamilies have highlighted the enormous complexity of the evolutionary process resulting in this large P450 superfamily. Together with the clustering of many subfamilies in tandem gene arrays, commonalities in their intron positions have provided evidence of the numerous gene duplications that have led to many of the current arrays of P450 subfamily genes. Divergences in their intron positions when different P450 families and, sometimes, P450

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subfamilies are compared have provided evidence of more complicated genetic restructurings that have led to the current high degree of organizational divergence among its members. Between these extremes, there exist examples in which intron positions of genes in one subfamily are shared with one or more other subfamilies as though current phylogenetic nomenclatures placing them in different subfamilies are but a timepoint in the long evolutionary process that is continually changing these genes. Coupled with phylogenetic comparisons, these structural intermediates in genome evolution have begun to unravel the various reiterations and deletions that have contributed to P450 genomic diversity. B. CONSERVED GENE FAMILIES

P450 gene families conserved in higher plants include CYP51 mediating obtusifol 14a-demethylation, CYP74 mediating oxylipin synthesis (producing jasmonic acid) and catabolism (producing C6-volatiles), CYP86 and CYP94 mediating fatty acid hydroxylations, CYP85 and CYP90 mediating brassinosteroid syntheses, and CYP97 mediating carotenoid hydroxylations (Mizutani and Ohta, 2010; Nelson and Werck-Reichhart, 2011; Nelson et al., 2008). P450 gene subfamilies existing in these genomes include CYP73A mediating cinnamic acid hydroxylation in early phenylpropanoid synthesis; CYP84A and CYP98A mediating various hydroxylations in lignin synthesis; CYP75B mediating 30 -hydroxylation in flavonoid synthesis; CYP88A and CYP701A mediating multiple conversions in gibberellin synthesis; CYP77A, CYP703A and CYP704B mediating fatty acid hydroxylations; CYP710A mediating sterol C22-desaturations; CYP711A mediating strictolactone synthesis; CYP707A mediating abscisic acid catabolism; CYP734A mediating brassinosteroid catabolism; and CYP735A mediating cytokinin catabolism. Other conserved subfamilies include CYP78A, which mediates an unspecified conversion in organ development, as well as CYP72A, CYP77B, CYP87A, CYP704A, whose functions have not yet been defined (Mizutani and Ohta, 2010; Nelson and Werck-Reichhart, 2011; Nelson et al., 2008). C. RESTRICTED GENE FAMILIES

P450 genes conserved in more restricted groups of higher plants vary depending on the groups of plants compared, their chemical constituents and their levels of functional analyses. These first two points are apparent when one considers the complexities of plant secondary metabolic pathways. In comparing two related species synthesizing similar metabolites, one is likely to find P450s within the same subfamily, and, in comparing distant species

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synthesizing quite different metabolites, one is likely to find little overlap in their P450 complements. Examples of the selective nature of some subfamilies include members of the CYP720B subfamily in Picea (spruce) and Pinea (pine) genuses that are involved in producing the diterpene olefins, alcohols and resin acids used for insect defence (Zulak and Bohlmann, 2010). Others are members of the CYP725A subfamily in the Taxus (yew) species that are involved in producing taxadiene and its derivative paclitaxel used for herbivore defence (Kaspera and Croteau, 2006) and members of multiple CYP80 and CYP719 subfamilies in Coptis, Papaver and Eschscholzia species that are involved in benzylisolquinoline alkaloid synthesis (Facchini, 2001; Ziegler et al., 2009). Others are members of the CYP71C subfamily in Z. mays (maize) that synthesizes benzoxazinoids (e.g. DIMBOA), which are broken down to toxic aglycons upon herbivore damage (Frey et al., 2009; Glawischnig et al., 1999). Complicating the association of this subfamily with benzoxazoid production, the CYP71C subfamily exists in rice, which is not known to synthesize benzoxazinoids, and not in any other plant genome sequenced to date. The fact that benzoxazinoids are produced in several other monocot grasses and dicots (Schullehner et al., 2008) has suggested that this particular set of biosynthetic activities has evolved multiple times using different subfamilies in the very large CYP71 family or completely different P450 families. Sorting out the many origins of this biochemical pathway awaits future genome sequencings and functional characterizations. The third point made concerning the restrictedness of some P450 subfamilies is less apparent and reflective of the many ways in which P450 catalytic sites can acquire new activities and discard old ones. These include cases where variations in one or just a few catalytic-site residues allow for the repositioning of the same substrate (e.g. limonene hydroxylations) or the acceptance of a new substrate and cases where completely different P450s have convergently evolved the ability to mediate the same reactions (e.g. benzoxazinoid synthesis). Some examples, which highlight the complexities of phylogenetic comparisons without detailed biochemical information, exist in the CYP79 and CYP83 subfamilies of papaya and Arabidopsis. Papaya, which produces benzylglucosinolates and not indole glucosinolates (Bennett et al., 1997), contains several CYP79B and CYP83B members closely related to Arabidopsis CYP79B and CYP83B members that mediate the conversion of tryptophan to indole glucosinolates (Nafisi et al., 2006; Sønderby et al., 2010) and lacks members related to Arabidopsis CYP79A members that mediate conversion of phenylalanine to benzylglucosinolates (Wittstock and Halkier, 2000). It would seem, therefore, that production of benzylglucosinolates in papaya has necessitated the subtle evolution of the catalytic

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site in one of the papaya CYP79B subfamily members allowing it to utilize phenylalanine in place or tryptophan or the more drastic evolution of an entirely different P450 family converging on this activity. Differentiating between these possibilities awaits biochemical characterizations of the papaya CYP79B and CYP83B proteins. Additional examples of these complexities exist in the CYP734A and CYP72C subfamilies mediating brassinolide inactivations (Nomura and Bishop, 2006; Thornton et al., 2010). The CYP734A proteins, which are encoded in all higher plant genomes (Nelson and Werck-Reichhart, 2011; Nelson et al., 2008), catabolize brassinolide via 26-hydroxylation. The CYP72C proteins, which are encoded by one gene in Arabidopsis and two genes in papaya and are absent in other sequenced plant genomes, catabolize brassinolide by hydroxylation at another site that has not yet been specified. Compared at the level of their primary sequences, these proteins share low enough identity that they fall in different families while still modifying the same substrate. Other examples of divergent P450 families mediating similar reactions on related substrates exist in the six Arabidopsis families (CYP77, CYP86, CYP94, CYP703, CYP704, CYP709) responsible for hydroxylations at terminal and internal points on fatty acids (Kandel et al., 2006; Pinot and Beisson, 2011). Together, these examples of convergence and divergence in catalytic activities highlight the evolutionary plasticities of this large group of monooxygenases that have enabled the development of many novel restricted activities while still maintaining ancient and, presumably, essential activities.

III. STRUCTURAL DIVERSITY A. STRUCTURAL DETERMINANTS IN CLASSICAL P450S

With several examples of different P450 subfamily members mediating reactions at different positions on the same substrates (e.g. limonene hydroxylations, fatty acid hydroxylations), three-dimensional perspectives on their catalytic-site structures are now beginning to provide insight into the number and types of amino acid variations needed to change the position of hydroxylation and/or the range of substrates. They are also beginning to provide insight into convergently evolved catalytic sites capable of accommodating the same substrate(s). Our ability to predict the structure of these catalytic sites has been greatly enabled by the availability of many mammalian P450 crystal structures defined with and without their respective substrates (Johnson and Stout, 2005; Poulos and Johnson, 2005). Studies of these have identified extensive conservation at the secondary and tertiary structure

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levels that have allowed individual P450 monooxygenases to maintain a conserved P450-fold even though their primary sequences may share as little as 23% identity (Poulos and Johnson, 2005). Within the core structure formed by 11 a-helices (labelled A–K) and four b-sheets (labelled 1–4), catalytic sites buried in each of these proteins contain six comparatively small regions (termed substrate recognition sites SRS1-6) that control substrate access, binding and catalysis (Poulos and Johnson, 2005; Poulos and Meharenna, 2007). Among these, SRS1 corresponds to the loop region between the B- and C-helices positioned over the catalytically essential haem, SRS2 and SRS3 correspond to the F- and G-helices contributing to formation of the substrate access channel, SRS4 corresponds to the I-helix extending over the haem, SRS5 and SRS6 correspond to the amino-terminus of b-sheet 1-4 and b-turn at the end of b-sheet 4 that both protrude into the catalytic site. By separating plant P450s into two classes depending on whether they are membrane-anchored (ER-localized) or soluble (chloroplast-localized; Baudry et al., 2006; Rupasinghe and Schuler, 2006), it is now possible to align their sequences with mammalian P450 templates (which are also typically membrane-bound and localized in the ER) or bacterial P450 templates (which are soluble) and predict catalytic-site structures with reasonable accuracy. Alignments at the level of primary sequence that are needed to initiate development of predictive structures have indicated that, despite their varying lengths and low sequence identities, the many P450 sequences in the Arabidopsis genome have no length variations in SRS regions closest to the haem (SRS4, SRS5, SRS6) and relatively few length variations in more distal SRS regions contributing to the substrate access channel (SRS1, SRS2, SRS3; Rupasinghe and Schuler, 2006). Instead, the length variations occur in a number of a-helices and b-sheets contributing to the overall P450-fold, external surface loops and the FG-loop (i.e. between the F- and G-helices), which is involved in defining substrate access and, probably, interactions with the ER membrane. Assessed at the level of their predicted structures, this means that the positioning of substrates in the catalytic site is actually defined by the small number of SRS side chains in or near the catalytic site. Examples of plant P450s known to have different reactivities on the same substrate due to side chain variations in SRS regions include Mentha spicata (spearmint) CYP71D18 and Mentha piperita (peppermint) CYP71D15 sequences that differentiate between C6- and C3-hydroxylations on limonene, respectively, based on a single Phe363Ile switch in SRS5 (Schalk and Croteau, 2000). Recently added to this subfamily is the Perilla frutescens (perilla) CYP71D174 sequence that has 63–68% identity to the spearmint and peppermint sequences and hydroxylates at the C7-, C3- and C6-positions (Mau

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et al., 2010). Comparisons among predicted structures that we have built for these three proteins indicate that these enzymes show extremely high conservation in most catalytic-site residues and that the spearmint and perilla sequences share Phe363 important for C6-hydroxylation. Overlays of these proteins suggest that the loss of regioselectivity seen for perilla CYP71D174 is probably due to the replacement of Ser297 (SRS4) in spearmint CYP71D18 with Ala277 in perilla CYP71D174. Dockings of limonene in the predicted CYP71D18 and CYP71D174 catalytic sites suggest that this single change allows the limonene molecule to bind in different orientations. The slight increase in catalytic-site volume due to the replacement of Ser297 with Ala277 in perilla CYP71D174 positions limonene in a horizontal orientation relative to the haem and brings the C7, C6 and C3 positions closer to the reactive oxygen. The more constricted catalytic site in CYP71D18 allows limonene to bind in a vertical orientation and positions only the C6 atom close to the reactive oxygen. Similarly, site-directed mutagenesis has demonstrated that substrate contacts in Heliothus tuberosus CYP73A1 (artichoke 4-cinnamic acid hydroxylase in phenylpropanoid synthesis) include Asn302 in SRS4 (I-helix) and Ile371 in SRS5 (loop between the K-helix and b1-4 strand; Schoch et al., 2003). Others influencing substrate positioning and activity include Lys484 in SRS6 (b-turn at the end of b-sheet 4), Ala306-Ala307 in SRS4 (I-helix) and Pro372 in SRS5 (N-terminus of b1-4 strand; Schalk et al., 1999; Schoch et al., 2003). Site-directed mutagenesis in Nicotiana tabacum CYP71D20 (tobacco 5-epiaristolochene 1,3-dihydroxylase in sesquiterpene phytoalexin synthesis) has indicated that Ser368 in SRS5 controls its overall activity towards 5-epiaristolochene: its mutagenesis to Ala368 or Thr368 decreases activity and mutagenesis to Val368 compromises capsidiol formation without affecting overall activity (Takahashi et al., 2005). Another residue critical for double hydroxylation of this substrate is Ile486 in SRS6: its mutagenesis blocks C3-hydroxylation on the 1b-hydroxy epiaristolochene intermediate. Site-directed mutagenesis in the related Hyoscyamus muticus CYP71D55 (henbane premnaspirodiene oxygenase in sesquiterpene phytoalexin synthesis) has indicated that Val366 in SRS5 and Val480, Val482 and Ala484 in SRS6 (aligning with Ser482, Ile484 and Ile486 in tobacco CYP71D20) control catalytic formation of solavetivone by affecting catalytic-site geometry rather than directly interacting with its substrate (Takahashi et al., 2007). Further comparisons between these proteins suggest that several others of their 91 amino acid differences account for their observed regio- and stereospecific differences. In addition, site-directed mutagenesis in Vicia sativa (vetch) CYP94A2 (o-hydroxylase in fatty acid synthesis) has indicated that Phe494 in SRS6

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affects positioning of short-chain fatty acids within its catalytic site: its mutagenesis to smaller Leu, Val, or Ala residues progressively increases the levels of in-chain and o-1 hydroxylations due to reductions in the steric constraints imposed by the larger Phe side chain (Kahn et al., 2001). Overlays of the predicted structures for the related Arabidopsis CYP94B1 and CYP94C1 subfamily members have indicated that they stabilize their oxygenated fatty acid substrates (e.g. 9,10-epoxystearic acid, 9,10-dihydroxystearic acid) with polar side chains from SRS2 and SRS5 (Rupasinghe and Schuler, 2006). These same side chains are conserved in the CYP94A subfamily members of vetch and tobacco and absent from several Arabidopsis CYP86A subfamily members that hydroxylate only non-oxygenated fatty acids (Rupasinghe and Schuler, 2006; Rupasinghe et al., 2007). Arabidopsis CYP86A1 that can hydroxylate these substrates is predicted to stabilize their binding with Arg49, a non-SRS residue in the A-helix. Overlays of the vetch CYP94A1, tobacco CYP94A5 and Arabidopsis CYP94C1 with their respective fatty acid substrates show many other catalytic-site side chains that directly align despite the more limited identities between these CYP94 subfamilies (Rupasinghe and Schuler, 2006). More comprehensive mappings of the positions affecting substrate recognition in other fatty acid hydroxylases are included in Hlavica and Lehnerer (2010). Finally, site-directed mutagenesis in Glycyrrhiza echinata (licorice) CYP93C2 (2-hydroxyisoflavonone synthase in isoflavonoid synthesis) has indicated that the unusual aryl migration occurring in its substrate is controlled by Ser310 in SRS4, which replaces the conserved oxygen-activating Thr in the I-helix of many P450s, and by Leu371 and Lys375 in SRS5 (Sawada et al., 2002). Site-directed mutagenesis in Gerbera hybrida (gerber daisy) CYP75B15 (flavonoid 30 -hydroxylase in flavonoid/anthocyanin synthesis) has indicated that Thr487 in SRS6 controls the position of aromatic ring hydroxylation: its conversion to Ser487 allows both 30 - and 50 -hydroxylations to occur (Seitz et al., 2007). In addition to this long list of SRS variations affecting plant P450 catalytic activities, a few non-SRS variations have been shown to affect substrate range. Examples here include variations between the three Triticum aestivum (wheat) CYP98A subfamily members mediating 3-hydroxylations on phenolic rings. CYP98A11 and CYP98A12, which are capable of modifying r-coumaroyltyramine, contain an additional amino acid (Cys52) at the N-terminus of their A-helices that orients the preceding Arg51 for interaction with the carboxylate group on shikimate and quinic esters and the phenolic hydroxyl of tyramine (Morant et al., 2007). CYP98A10, which cannot hydroxylate r-coumaroyltyramine, lacks Cys52 and orients Arg51 in a way

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that prevents it from interacting with this potential substrate. Introduction of the missing Cys52 into CYP98A10 enhances metabolism of the shikimate and quinic esters and allows for metabolism of r-coumaroyltyramine. B. STRUCTURAL DETERMINANTS IN NONCLASSICAL P450S

The recent availabilities of a crystal structure for the atypical A. thaliana and Parthenium argentatum (guayule) CYP74A subfamily members (allene oxide synthases in oxylipin synthesis; Lee et al., 2008; Li et al., 2008) have delineated residues in these chloroplast-localized P450s that affect haem orientation and substrate positioning. Among the most significant differences distinguishing this group of nonclassical soluble P450s from classical ERlocalized P450s is an insertion of nine residues upstream of the haem-binding Cys471 that reorganizes the surface regions typically interacting with electron-transfer partners. Other differences in Arabidopsis CYP74A1 reposition the kink in the I-helix so that Asn321 is over the haem and substitutes Ile328 for the catalytically important Thr in the I-helix of other P450s. As a result of these structural changes, CYP74 proteins are able to short-circuit the normal P450 catalytic cycle using hydroperoxides as oxygen donors rather than the dioxygen used by classical P450s (Brash, 2009; Lee et al., 2008). Even with these structural and catalytic cycle differences, residues in SRS regions affect activity. Site-directed mutagenesis of Phe137 (SRS1) in Arabidopsis CYP74A1 has shown it to be important for stabilization of the radical intermediates leading to allene oxide formation: its replacement with Leu allowed it mediate hydroperoxide cleavage (Lee et al., 2008). Similarly, mutagenesis of Phe92 in rice CYP74A5, which aligns with Phe137 in Arabidopsis, also converted it from an allene oxide synthase to a hydroperoxide lyase (Lee et al., 2008).

IV. COMPARISONS BETWEEN ORYZA AND ARABIDOPSIS P450S A. PHYLOGENETIC RELATIONSHIPS AMONG FUNCTIONALLY DEFINED P450S

With the number of annotated plant P450s exponentially increasing as new genomes are sequenced, the remainder of this review is focused on more manageable comparisons of Arabidopsis and Oryza P450 functions and their predicted structures. In the 7 years since the P450 families in these two species were first phylogenetically analysed (Nelson et al., 2004), functions have been assigned to 64 Arabidopsis P450s and 22 Oryza P450s. Comparisons between

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the P450 subfamilies in which one or more of members have been functionally defined in Arabidopsis or Oryza are listed in Table I. Primary references for the Arabidopsis P450s initially presented in Schuler et al. (2006) are updated in Table II. Primary references for the Oryza P450s are summarized for the first time in Table III. Comparisons among the 33 families presented in Table I highlight five types of phylogenetic relationships. First, several P450 families are highly conserved (i.e. one subfamily) and maintain low copy numbers in both species. Examples here include many with essential roles in plant growth and development such as CYP73A, CYP84A and CYP98A in phenylpropanoid synthesis, CYP85A in brassinosteroid synthesis, CYP88A and CYP701A in gibberellin synthesis, CYP703A in fatty acid synthesis, CYP707A in abscisic acid inactivation, CYP710A in sterol synthesis, CYP734A in brassinolide inactivation and CYP735A in cytokinin synthesis. Second, several P450 families contain multiple conserved subfamilies that closely align in both species. Examples here include CYP97 with three subfamilies in carotenoid synthesis, CYP77 with two subfamilies in fatty acid synthesis and CYP704 with two subfamilies in fatty acid synthesis. Three closely aligning subfamilies also exist in the CYP90 family mediating brassinosteroid synthesis but, in this instance, a fourth subfamily (CYP90C) in Arabidopsis is absent from Oryza and possibly replaced by one of the duplicated CYP90A sequences in Oryza or by an even more divergent sequence. Three closely aligning subfamilies also exist in the CYP94 family mediating fatty acid hydroxylations and, in this instance, a fourth subfamily (CYP94E) is present in Oryza and absent from Arabidopsis. Third, several P450 families contain one highly conserved subfamily and other more divergent subfamilies (i.e. nonoverlapping subfamilies occur in these species). Examples here include the conserved CYP74A subfamily in jasmonate synthesis and the divergent CYP74B subfamily (Arabidopsis) and CYP74E and CYP74F subfamilies (Oryza) in C6-volatile synthesis. Other examples include the conserved CYP86A and CYP86B subfamilies in fatty acid synthesis and the divergent CYP86C subfamily (Arabidopsis) and CYP86E subfamilies (Oryza) as well as the CYP51, CYP72, CYP75, CYP78, CYP79 families that each contains one highly conserved subfamily and one (or more) divergent subfamily in one or the other of these species. Fourth, several P450 families have proliferated and diverged so considerably (i.e. numerous genes in nonoverlapping subfamilies). Examples here include CYP76 with one Oryza member involved in diterpenoid phytoalexin synthesis, CYP81 with one Oryza member in herbicide metabolism and several Arabidopsis members in glucosinolate synthesis, CYP93 with one Oryza member in flavone synthesis, CYP96 with several Arabidopsis members in fatty acid synthesis, CYP714 with one Oryza

TABLE I Arabidopsis and Oryza P450 Families with One or More Functions Defined Oryza 51G1 51G3

Function in Oryza – –

Arabidopsis 51G1

Function in Arabidopsis obtusifol 14a-demethylase

51H1 51H3–9 71A12 71A13

– – – conversion of indole-3-acetaldoxime to indole-3-acetonitrile – – – – conversion of cysteine indole 3-acetonitrile and dihydrocamalexic acid to camalexin – –

71A14–16 71A18–26 71A28 71B2–14 71B15

71Ca 71E4–6 71Ka 71P1 71Q–Za 71AA–ACa 71AD1 71AF1 71AG1 71AK1–2 72A17 72A18 72A19–25

71B16–29 71B31–38 – – – tryptamine 5-hydroxylase – – – – – – – pelargonic acid (o-1) hydroxylase –

Modifications on/products Sterols

Camalexin

Camalexin

Serotonin

72A7–11 72A13–15

– –

Herbicide detoxification

72A32–35

–

73A38–40 74A4 74A5

– – allene oxide synthase

74E1 74E2 74F1 75A11 75B3 75B11

9-/13-hydroperoxide lyase 9-/13-hydroperoxide lyase – – flavonoid 30 -hydroxylase –

76H4–11 76M7 76K–Na 76P–Qa 76U1 76V1 77A18

– ent-cassadiene C11a-hydroxylase – – – – –

72C1 73A5 74A1

degradation of brassinosteroids 4-cinnamic acid hydroxylase allene oxide synthase

74B2

13-hydroperoxide lyase

Brassinosteroids Core phenylpropanoids Jasmonic acid Jasmonic acid C6-Volatiles

75B1

flavonoid 30 -hydroxylase

Flavonoids/anthocyanins

76C1–7 76G1

– – Diterpenoid phytoalexins

77A4 77A6

77B2 78A11

– plastochron 1 mutant

78A12–17 78D1 79A7

– –

79A9–11

–

77A7 77A9 77B1 78A5 78A6–10 79A2

epoxidase and o-hydroxylase on C18 fatty acids in-chain hydroxylase on 16-hydroxypalmitate – – – synthesis of mobile signalling molecule –

Fatty acids

conversion of phenylalanine to oxime

Benzylglucosinolates

Fatty acids

(continues)

TABLE I Oryza

Function in Oryza

Arabidopsis 79B2 79B3 79C1 79C2 79F1 79F2

81A5 81A6 81A7–8

Function in Arabidopsis conversion of tryptophan and analogs to oximes conversion of tryptophan to oxime – – synthesis of short and long chain aliphatic glucosinolates synthesis of long chain aliphatic glucosinolates

– bentazon and sulfonylurea metabolism –

Modifications on/products Indole glucosinolates Indole glucosinolates Aliphatic glucosinolates Aliphatic glucosinolates Herbicide metabolism

81D1–11 81F1

81F4 81G1 81H1 81K1–2

– conversion of l3M to 4-hydroxy-l3M, 1-hydroxy l3M conversion of l3M to 4-hydroxy-l3M, 1-hydroxy l3M conversion of l3M to 4-hydroxy-l3M, 1-hydroxy l3M conversion of l3M to 1-hydroxy l3M – – –

82C2 82C3 82C4

hydroxylase for 8-methoxypsoralen – hydroxylase for 8-methoxypsoralen

81F2 81F3

81L–Na 81P1

(continued )

– –

Glucosinolates Glucosinolates Glucosinolates Glucosinolates

82F1 82G1 83A1 84A5–7

–

85A1

6-oxidase for 6-deoxycastasterone and other steroids

83B1 84A1 84A4 85A1 85A2

86A9–11

–

86A1 86A2 86A4 86A7 86A8

86B3

–

86B1 86B2 86C1–4

86E1 88A5

– ent-kaurenoic acid oxidase

90A3–4

–

90A19

–

88A3 88A4 90A1

– degradation of C20 geranyllinalool and C15 nerolidol oxidation of methionine-derived oximes oxidation of indole-3-acetaldoxime 5-hydroxylase for coniferaldehyde/ coniferyl alcohol/ferulic acid – 6-oxidase for 6-deoxycastasterone and other steroids 6-oxidase for 6-deoxycastasterone and other steroids o-hydroxylase on C12–C18 fatty acids o-hydroxylase on C12–C18 fatty acids o-hydroxylase on C12–C18 fatty acids o-hydroxylase on C12–C18 fatty acids o-hydroxylase on C12–C18 fatty acids o-hydroxylase on C22–C24 fatty acids – – ent-kaurenoic acid oxidase ent-kaurenoic acid oxidase 23a-hydroxylase for 6-oxocathasterone/cathasterone

Homoterpene volatiles Aliphatic glucosinolates Indole glucosinolates Lignin monomers Brassinosteroids Brassinosteroids Fatty acids Fatty acids Fatty acids Fatty acids Fatty acids Fatty acids

Gibberellins Brassinosteroids

(continues)

TABLE I Oryza 90B2

Function in Oryza 22a-hydroxylase for campesterol

Arabidopsis 90B1 90C1

90D2

90D3

conversion of teasterone and 6-deoxoteasterone to 3-dehydroteasterone and 3-dehydro 6-deoxoteasterone –

93F1 93G1 93G2 94B4–5

– – flavanone 2-hydroxylase –

90D1

–

94B1

o-hydroxylase on saturated and oxygenated fatty acids – o-hydroxylase on saturated and oxygenated fatty acids o-hydroxylase on saturated and oxygenated fatty acids – –

94C2–4

–

94C1

94D4–7 94D9–13 94D15 94E1–3

– – – –

94D1 94D2 96A1–3 96A4 96A5 96A7–13 96A15

–

Function in Arabidopsis 22a-hydroxylase for campesterol/ campestanol/6-deoxocampestanol 23a-hydroxylase for multiple brassinosteroids 23a-hydroxylase for multiple brassinosteroids

93D1

94B2 94B3

96B2–10

(continued )

– o-hydroxylase for saturated C12–C14 fatty acids and oleic acid – – mid-chain hydroxylase for alkanes and secondary alcohols

Modifications on/products Brassinosteroids Brassinosteroids Brassinosteroids

Flavones Fatty acids Fatty acids Fatty acids

Fatty acids

Epidermal waxes

96D1–2 96E1 97A4 97B4 97C2 98A4

– – b-ring carotene hydroxylase – e-ring carotene hydroxylase –

98A18

–

99A2 99A3 701A6 701A7–9 701A19 703A3

– diterpene oxidase ent-kaurene oxidase – – –

704A3–8 704B2 707A5 707A6 707A37

– o-hydroxylase on C16–C18 fatty acids ABA 80 -hydroxylase ABA 80 -hydroxylase –

710A5–8

–

98A8 98A9

b-ring carotene hydroxylase b-ring carotene hydroxylase e-ring carotene hydroxylase 30 -hydroxylase for p-coumaroylshikimic/quinic acids triferuloylspermidine hydroxylase triferuloylspermidine hydroxylase

701A3

ent-kaurene oxidase

703A2

in-chain hydroxylase on C10–C14 fatty acids – in-chain hydroxylase on C16–C18 fatty acids ABA 80 -hydroxylase ABA 80 -hydroxylase ABA 80 -hydroxylase ABA 80 -hydroxylase 22-desaturase for b-sitosterol 22-desaturase for 24-epicampesterol and b-sitosterol – 22-desaturase for b-sitosterol –

97A3 97B3 97C1 98A3

704A1–2 704B1 707A1 707A2 707A3 707A4 710A1 710A2 710A3 710A4 714A1–2

714B1 714C1–3 714D1

Carotenoids Carotenoids Carotenoids Lignin monomers Phenolamides Phenolamides Momilactones Gibberellins Fatty acids Fatty acids ABA inactivation ABA inactivation ABA inactivation ABA inactivation Sterols Sterols Sterols

– – epoxidase on non-13-hydroxylated GAs (continues)

TABLE I Oryza

Function in Oryza

Arabidopsis 724A1

724B1

(continued ) Function in Arabidopsis

Modifications on/products

–

734A2

22a-hydroxylase for brassinosteroid precursors –

734A1

26-hydroxylase for brassinolide and castasterone

Brassinolide inactivation

734A4–6 735A3

– –

735A1

Cytokinins

735A4

–

735A2

trans-hydroxylase for isopentenyladenine phosphates trans-hydroxylase for isopentenyladenine phosphates

a

Brassinosteroids

Individual members of these multigene subfamilies are not listed.

Cytokinins

TABLE II Arabidopsis thaliana P450s Functionally Defined P450 51G1 71A13 71B15 72C1 73A5 74A1 74B2 75B1 77A4 77A6 79A2 79B2 79B3 79F1 79F2 81F1

Activity Obtusifoliol 14a-demethylase Conversion of indole-3-acetaldoxime to indole-3-acetonitrile Conversion of cysteine indole 3-acetonitrile and dihydrocamalexic acid to camalexin Degradation of brassinosteroids Cinnamic acid 4-hydroxylase (t-CAH) Allene oxide synthase (AOS) Hydroperoxide lyase (HPL) 30 -Hydroxylase for narigenin, dihydrokaempferol (F30 H) Epoxidase and o-hydroxylase on C18 fatty acids In-chain hydroxylase on 16-hydroxypalmitate Conversion of phenylalanine to oxime Conversion of tryptophan and analogs to oximes Conversion of tryptophan to oxime Mono- to hexahomomethionine in synthesis of short and long chain aliphatic glucosinolates Long chain penta- and hexahomomethionine in synthesis of long chain aliphatic glucosinolates Conversion of indol-3-ylmethylglucosinolate to 4-hydroxy-l3M and 1-hydroxy-l3M

Pathway

References

Sterols Camalexin

Kushiro et al. (2001), Kim et al. (2005b) Nafisi et al. (2006)

Camalexin Brassinosteroid inactivation Phenylpropanoids Oxylipins Oxylipins Phenylpropanoids

Zhou et al. (1999), Schuhegger et al. (2006), Bottcher et al. (2009) Nakamura et al. (2005), Takahashi et al. (2005) Urban et al. (1997), Mizutani et al. (1997) Laudert et al. (1996) Bate et al. (1998) Schoenbohm et al. (2000)

Fatty acids

Sauveplane et al. (2009)

Fatty acids Benzylglucosinolates Indole glucosinolates

Li-Beisson et al. (2009) Wittstock and Halkier (2000) Hull et al. (2000), Mikkelsen et al. (2000)

Indole glucosinolates Aliphatic glucosinolates

Hull et al. (2000) Hansen et al. (2001), Reintanz et al. (2001), Chen et al. (2003)

Aliphatic glucosinolates

Reintanz et al. (2001), Chen et al. (2003)

Glucosinolates

Pfalz et al. (2011) (continues)

TABLE II

(continued )

P450

Activity

81F2

83A1

Conversion of indol-3-ylmethylglucosinolate to 4-hydroxy-l3M and 1-hydroxy-l3M Conversion of indol-3-ylmethylglucosinolate to 4-hydroxy-l3M and 1-hydroxy-l3M Conversion of indol-3-ylmethylglucosinolate to 1-hydroxy-l3M Hydroxylase for 8-methoxypsoralen Hydroxylase for 8-methoxypsoralen Oxidative degradation of C20 geranyllinalool and C15 nerolidol Oxidation of methionine-derived oximes

Aliphatic glucosinolates

83B1

Oxidation of indole-3-acetaldoxime

Indole glucosinolates

84A1

5-Hydroxylase for coniferaldehyde, coniferyl alcohol and ferulic acid (F5H) C6-Oxidase for 6-deoxycastasterone and other steroids C6-Oxidase for 6-deoxycastasterone and other steroids o-Hydroxylase for saturated and unsaturated C12 to C18 fatty acids o-Hydroxylase for saturated and unsaturated C12 to C18 fatty acids o-Hydroxylase for saturated and unsaturated C12 to C18 fatty acids o-Hydroxylase for lauric acid

Phenylpropanoids

81F3 81F4 82C2 82C4 82G1

85A1 85A2 86A1 86A2 86A4 86A7 86A8

o-Hydroxylase for saturated and unsaturated C12 to C18 fatty acids

Pathway

References

Glucosinolates

Bednarek et al. (2009), Pfalz et al. (2009)

Glucosinolates

Pfalz et al. (2011)

Glucosinolates

Pfalz et al. (2011)

Homoterpene volatiles

Kruse et al. (2008) Kruse et al. (2008) Lee et al. (2010)

Brassinosteroids Brassinosteroids Fatty acids Fatty acids Fatty acids Fatty acids Fatty acids

Hemm et al. (2003), Bak and Feyereisen (2001), Naur et al. (2003) Bak et al. (2001), Bak and Feyereisen (2001), Naur et al. (2003) Meyer et al. (1996), Ruegger et al. (1999), Humphreys et al. (1999) Shimada et al. (2001), Shimada et al. (2003) Shimada et al. (2003), Nomura et al. (2005), Kim et al. (2005c) Benveniste et al. (1998), Rupasinghe et al. (2007), Hofer et al. (2008) Duan and Schuler (2005), Rupasinghe et al. (2007) Duan and Schuler (2005), Rupasinghe et al. (2007), Li-Beisson et al. (2009) Duan and Schuler (2005), Rupasinghe et al. (2007) Wellesen et al. (2001), Rupasinghe et al. (2007)

Fatty acids Gibberellins Gibberellins Brassinosteroids

Compagnon et al. (2009) Helliwell et al. (2001) Helliwell et al. (2001) Szekeres et al. (1996)

Brassinosteroids

Choe et al. (1998), Fujita et al. (2006)

Brassinosteroids

Kim et al. (2005a), Ohnishi et al. (2006)

Brassinosteroids

Kim et al. (2005a), Ohnishi et al. (2006)

Fatty acids

Benveniste et al. (2006)

Fatty acids

Benveniste et al. (2006)

Fatty acids

Benveniste et al. (2006), Kandel et al. (2007)

Fatty acids

Benveniste et al. (2006)

Epidermal waxes

Greer et al. (2007)

97A3

o-Hydroxylase for C22–C24 fatty acids Multifunctional ent-kaurenoic acid oxidase Multifunctional ent-kaurenoic acid oxidase 23a-Hydroxylase for 6-oxo-cathasterone and cathasterone 22a-Hydroxylase for campesterol, campestanol and 6-oxo-campestanol 23a-Hydroxylase for multiple brassinosteroids 23a-Hydroxylase for multiple brassinosteroids o-Hydroxylase for saturated and oxygenated fatty acids o-Hydroxylase for saturated and oxygenated fatty acids o-Hydroxylase and in-chain hydroxylase for saturated C12 and unsaturated C18 fatty acids and 9, 10 epoxystearic acid o-Hydroxylase for saturated C12, C14 fatty acids and oleic acid Mid-chain hydroxylase for alkanes and secondary alcohols b-Ring carotene hydroxylase

Carotenoids

97B3 97C1

b-Ring carotene hydroxylase e-Ring carotene hydroxylase

Carotenoids Carotenoids

98A3

Phenylpropanoids

98A8 98A9 701A3

30 -Hydroxylase for p-coumaryl shikimic/ quinic acids (C30 H) Hydroxylase on triferuloylspermidine Hydroxylase on triferuloylspermidine Multifunctional ent-kaurene oxidase

Kim and DellaPenna (2006), Tian et al. (2004) Kim et al. (2009) Kim and DellaPenna (2006), Tian et al. (2004) Schoch et al. (2001)

Phenolamides Phenolamides Gibberellins

703A2

In-chain hydroxylase for C10–C14 fatty acids

Fatty acids

86B1 88A3 88A4 90A1 90B1 90C1 90D1 94B1 94B3 94C1 96A4 96A15

Matsuno et al. (2009) Matsuno et al. (2009) Helliwell et al. (1998, 1999), Morrone et al. (2010) Morant et al. (2007) (continues)

TABLE II P450

(continued )

Activity

Pathway

References

704B1 707A1 707A2 707A3 707A4 710A1

In-chain hydroxylase for C16–C18 fatty acids 80 -Hydroxylase for ABA 80 -Hydroxylase for ABA 80 -Hydroxylase for ABA 80 -Hydroxylase for ABA C-22 desaturase for b-sitosterol

Fatty acids ABA inactivation ABA inactivation ABA inactivation ABA inactivation Sterols

710A2

C-22 desaturase on 24-epicampesterol and b-sitosterol C-22 desaturase for b-sitosterol 26-Hydroxylase for brassinolide and castasterone trans-Hydroxylase for isopentenyladenine phosphates trans-Hydroxylase for isopentenyladenine phosphates

Sterols

Dobritsa et al. (2009) Saito et al. (2004), Kushiro et al. (2004) Saito et al. (2004), Kushiro et al. (2004) Saito et al. (2004), Kushiro et al. (2004) Saito et al. (2004), Kushiro et al. (2004) Morikawa et al. (2006), Arnqvist et al. (2008) Morikawa et al. (2006)

Sterols Brassinolide inactivation

Arnqvist et al. (2008) Neff et al. (1999), Turk et al. (2003)

Cytokinins

Takei et al. (2004)

Cytokinins

Takei et al. (2004)

710A4 734A1 735A1 735A2

TABLE III Oryza sativa P450s Functionally Defined CYP

Activity/induction

Pathway

71P1

Tryptamine 5-hydroxylase

Serotonin

72A18 74A5 74E1 74E2 75B3 76M7 81A6 85A1 88A5 90B2 90D2

Herbicide detoxification Jasmonic acid Oxylipins Oxylipins Flavonoids Diterpenoid phytoalexins Herbicide detoxification Brassinosteroids Gibberellins Brassinosteroids Brassinosteroids

93G2 97A4 97C2 99A3

Peralogonic acid (o-1) hydroxylase Allene oxide synthase 9-/13-Hydroperoxide lyase 9-/13-Hydroperoxide lyase Flavonoid 30 -hydroxylase ent-Cassadiene C11a-hydroxylase Bentazon and sulfonylurea metabolism C6-Oxidase for 6-deoxocastasterone and other steroids ent-Kaurenoic acid oxidase 22a-Hydroxylase for campesterol Conversion of teasterone and 6-deoxoteasterone to 3-dehydroteasterone and 3-dehydro 6-deoxoteasterone Flavanone 2-hydroxylase b-Ring carotene hydroxylase e-Ring carotene hydroxylase Multifunctional diterpene oxidase

Flavones Carotenoids Carotenoids Momilactones

701A6 704B2 707A5 707A6 714D1 724B1

ent-Kaurene oxidase o-Hydroxylase on C16–C18 fatty acids ABA 80 -hydroxylase ABA 80 -hydroxylase Epoxidase on non-13-hydroxylated GAs 22a-Hydroxylase for brassinosteroid precursors

Gibberellins Fatty acids ABA inactivation ABA inactivation Gibberellin inactivation Brassinosteroids

Reference Fujiwara et al. (2010) Park et al. (2011) Imaishi and Matumoto (2007) Mei et al. (2006) Kuroda et al. (2005) Kuroda et al. (2005) Shih et al. (2008) Swaminathan et al. (2009) Pan et al. (2006) Hong et al. (2002) Sakamoto et al. (2004) Sakamoto et al. (2006) Hong et al. (2003) Du et al. (2010) Quinlan et al. (2007) Quinlan et al. (2007) Shimura et al. (2007) Wang et al. (2011) Ko et al. (2008) Li et al. (2010) Yang and Choi (2006) Saika et al. (2007) Zhu et al. (2006) Tanabe et al. (2005) Sakamoto et al. (2006)

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member in gibberellin metabolism and the enormous CYP71 family with several Arabidopsis members in camalexin synthesis and one Oryza member in serotonin synthesis. Although substantially smaller in size than these other families, the CYP724 family might also be placed in this same category since its single Oryza sequence (CYP724B1) mediating brassinosteroid synthesis is annotated in a different subfamily than the single Arabidopsis CYP724A sequence. Fifth, some P450 families have diverged so considerably that they have no close relatives in the other species. Examples of these, which may mediate dicot- or monocot-specific functions, include CYP82 in Arabidopsis with one member mediating production of terpene volatiles, CYP83 in Arabidopsis mediating glucosinolate synthesis and CYP99 in Oryza mediating momilactone synthesis. B. STRUCTURAL SIMILARITIES IN HIGHLY CONSERVED P450S

With functions defined for some of these highly conserved and more divergent P450 families, it becomes possible to examine the extent of catalytic-site evolution in P450s mediating various functions and provide three-dimensional perspectives on the amino acids conserved for maintenance of essential functions versus variant for acquisition of new functions. These comparisons, which are done by overlaying the predicted structures for different groups of proteins, indicate that there are often more conservations in catalytic-site residues than is evident in primary sequence alignments. The first set of examples for this are in the CYP98A subfamily mediating lignin synthesis where alignments of the SRS regions in Arabidopsis CYP98A3 and Oryza CYP98A4 and CYP98A18 (65– 75% overall identity) show absolutely conserved SRS5 (0/10 differences), highly conserved SRS4 (2–3/19 differences) and SRS1 (3/14 differences), less conserved SRS2 (3/7 differences), SRS3 (3/8 differences) and SRS6 (3–5/9 differences) regions. Overlays of predicted structures for Arabidopsis CYP98A3 and Oryza CYP98A4 (Fig. 1) indicate that, despite significant divergence in some SRS regions, there is extremely high conservation in most side chains contacting the r-coumaroyl shikimic acid substrate. The change of His95 (SRS1) in Arabidopsis CYP98A3 to Pro99 in Oryza CYP98A sequences is a dicot– monocot difference that occurs in many but not all sequenced CYP98A proteins. Based on our predicted structures, this difference is likely to affect substrate access and/or positioning of the shikimic/quinic acid ‘tails’ on their substrates but not substantially affect substrate range. The change of Phe239 (FG-loop between SRS2 and SRS3) in Arabidopsis CYP98A3 to Tyr243 in Oryza CYP98A4 sequences is not predicted to change substrate binding. With these many side chain conservations, these models allow us to predict with near certainty that rice CYP98A4 and CYP98A18 mediate the

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289

F239/Y243

H95/P99

CYP98A3/CYP98A4

Fig. 1. Predicted structures for CYP98A proteins. The docking mode for rcoumaroyl shikimate (elemental colours in ball-and-stick format) in the Arabidopsis ˚ of this CYP98A3 predicted structure is shown with substrate contacts within 4.5 A substrate. Overlaid with this is the Oryza CYP98A4 predicted structure with identical side chains shown in green and variable side chains H95/P99 (SRS1) and F239/Y243 (loop between SRS2 and SRS3) shown in elemental colours in stick format. The haem in the floor of the catalytic site is shown in space-filling format.

same 30 -hydroxylation on r-coumaroylshikimic and quinic acids as documented for Arabidopsis CYP98A3. With their substantial divergence from Arabidopsis CYP98A3 in their overall identity (50%) and even more in their SRS regions, it is not surprising that the divergent Arabidopsis CYP98A8 and CYP98A9 mediate hydroxylations on triferuloylspermidine (Matsuno et al., 2009) rather than r-coumaroylshikimic/quinic acids as in the case of other CYP98A proteins. Overlays of these indicate that the CYP98A8 and CYP98A9 catalytic sites are most substantially affected by changes in distal regions contributing to formation of the substrate access channel and interacting with tails on their substrates. The second set of examples for catalytic-site conservation are in the CYP84A subfamily mediating lignin production where alignments of Arabidopsis and Oryza CYP84A sequences (51–70% overall identity) show highly conserved SRS regions. Of the three rice CYP84A sequences, two are more closely related to Arabidopsis CYP84A1 with one difference in SRS5 (rice CYP84A5 only), two differences in SRS1, SRS4 (rice CYP84A6 only) and

290

MARY A. SCHULER AND SANJEEWA G. RUPASINGHE

SRS6, four differences in SRS3. And, several SRS regions (SRS2, SRS4 in CYP84A5, SRS5 in CYP84A6) show no variations between these two Arabidopsis and two Oryza proteins. With just a few SRS differences occurring in Arabidopsis CYP84A1 and CYP84A4 sequences, Oryza CYP84A7 is substantially diverged from the other two Oryza CYP84A sequences with two to seven differences in SRS1, SRS2, SRS3 and SRS5, five differences and a three amino acid insertion in SRS4; unexpectedly, its SRS6 is identical to Arabidopsis CYP84A1. Overlays of predicted structures for Arabidopsis CYP84A1, CYP84A4 and Oryza CYP84A5, CYP84A6 (Fig. 2) show extensive catalytic-site identity than is evident from primary sequence alignments. All SRS side chains predicted to contact the ferulic acid substrate are identical, except for the bulky Phe405 in SRS5 of CYP84A4 that is predicted to slightly reorient this substrate without affecting its hydroxylation position (Fig. 2). Compared to these four predicted structures, the catalytic site in Oryza CYP84A7 has evolved to the point that it is not predicted to bind ferulic acid or any other lignin precursor.

I384/F405/I389/I371

CYP84A1/CYP84A4/CYP84A5/CYP84A6

Fig. 2. Predicted structures for CYP84A proteins. The docking mode for ferulic acid (elemental colours in ball-and-stick format) in the Arabidopsis CYP84A1 pre˚ of this substrate. dicted structure is shown with substrate contacts within 4.5 A Overlaid with this are the predicted structures for Arabidopsis CYP84A4 as well as Oryza CYP84A5 and CYP84A6 with identical side chains shown in green and the variable side chain I384/F405/I389/I371 (SRS5 residues designated as CYP84A1/A4/ A5/A6) that slightly reorients this substrate in the CYP84A4 catalytic site. The haem in the floor of the catalytic site is shown in space-filling format.

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291

In a third set of examples for catalytic-site conservations, the overall identities among the five Arabidopsis CYP86A proteins (61–87%) and three Oryza CYP86A proteins (63–90%) compare with overall identities of 56–90% for all eight proteins. Comparisons among the Arabidopsis CYP86A proteins indicate that all five are moderately variable in SRS1, SRS4 and SRS5, more variable in SRS2 and SRS3 and identical in SRS6. The three Oryza CYP86A proteins follow this same pattern of variation but with less divergence than some of the Arabidopsis CYP86A proteins. In contrast to their variabilities in other regions, all eight of these CYP86A proteins are identical in SRS6. Biochemical analyses have indicated that all five Arabidopsis CYP86A proteins mediate hydroxylations on fatty acids in the C12 to C18 range (Benveniste et al., 1998; Duan and Schuler, 2005; Li-Beisson et al., 2009; Rupasinghe et al., 2007; Wellesen et al., 2001) and overlays of their predicted structures (Rupasinghe et al., 2007) show that this is due highly conserved substrate contacts. Of the 32 predicted contact residues within 4.5A of oleic acid (C18:1), 22 are conserved in all five proteins and 10 are variable in one or more. Most of these conserved residues, including the nine absolutely conserved residues in SRS6, make up the hydrophobic core of the fatty acid binding site associated with the acyl chain. Variations that do occur are in the distal regions of the substrate access channel (b1–2 and b1–4 regions (nonSRS), b1–5 strand (SRS1) and b1–3 strand (SRS5)) contacting the carboxy group on short-chain fatty acids. Overlays of the eight predicted Arabidopsis and Oryza CYP86A structures (not shown) show absolute conservation near the o-carbon binding site and some variations in the carboxylate-binding regions. In a fourth set of examples, Arabidopsis CYP86B1 and CYP86B2 and the Oryza CYP86B3 share significant overall identity (62–79%) with absolute conservation in SRS4 (20 amino acids) and SRS5 (9 amino acids), high conservation in SRS2 (0–2/7 differences) and SRS6 (1/7 differences), moderate conservation in SRS1 (5–6/23 differences) and somewhat less conservation in SRS3 (1–4/8 differences). Homology modelling and overlays of these three CYP86B proteins show a highly conserved and unusually large catalytic site. Examination of the residues in the predicted catalytic site shows that, of 22 residues predicted to constitute the substrate-binding cavity, only three residues in its distal region are not conserved. C. STRUCTURAL PERSPECTIVES ON MODERATELY CONSERVED P450S

In a fifth set of examples that can begin to detail the catalytic-site conservations and divergences leading to new activities, Arabidopsis CYP90B1 and Oryza CYP90B2, which are known to mediate 22a-hydroxylation on

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campesterol, campestanol and 6-deoxocampestanol (Choe et al., 1998; Fujita et al., 2006; Sakamoto et al., 2006), are closely related in their overall sequences (66% identity) and even more closely related in their SRS regions. Among these, SRS5 has no differences, SRS3 and SRS6 have one difference, SRS2 and SRS4 have two differences and SRS1 has three differences. Overlays of the predicted CYP90B1 and CYP90B2 structures (Fig. 3B) show that only two of these changes exist in side chains predicted to contact their campesterol substrate: Asn82 and Pro87 (both in SRS1) in Arabidopsis CYP90B1 are changed to Ser99 and Arg104 in Oryza. Similar comparisons of Oryza CYP724B, another brassinosteroid 22a-hydroxylase (Sakamoto et al., 2006), and Arabidopsis CYP724A1 (48% overall identity) indicate that these two members of the CYP724 family are substantially more divergent in all SRS regions except SRS5 (1/10 differences) and SRS6 (1/7 differences). These changes are predicted to substantially affect catalytic-site dimensions and cause the two CYP724 proteins to have different catalytic activities. In contrast and despite their phylogenetic placements in different

A

B

L200/A214

L468/P488 T294/L303

N82/S99 P87/R104

R199/T213 D196/E210

N79/H84

CYP724A1/CYP724B1

C

CYP90B1/CYP90B2

F498/499 E97 H84 M318 Q96

CYP90B1/CYP90B2/CYP724B1

Fig. 3.

(Continued)

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D

R235/254/246

F501/502/483

I238/259/249

T202/221/213

L83/100/85 P506/507/488 V499/500/481 L505/506/487

E199/218/210 E314/314/299 L309/309/294

S317/317/302

T88/105/90 L306/306/296 L304/304/294

V381/385/368

V90/107/92 L96/112/98

S91/108/93 Y75/92/77

T315/315/300

G310/310/297

H385/389/372

D93/110/95

CYP90B1/CYP90B2/CYP724B1

Fig. 3. Predicted structures for CYP90 and CYP724 proteins. (A) The docking mode for campesterol (magenta in ball-and-stick format) in the Oryza CYP724B1 ˚ of this substrate. predicted structure is shown with substrate contacts within 4.5 A Overlaid with this is the predicted structure for Arabidopsis CYP724A1 with identical side chains shown in green and the many variable side chains shown in elemental colours. (B) The docking mode for campesterol (orange in ball-and-stick format) in the Arabidopsis CYP90B1 predicted structure is shown with substrate contacts within ˚ of this substrate. Overlaid with this is the predicted structure for Oryza 4.5 A CYP90B2 with identical side chains shown in green and the variable side chains N82/S99 and P87/R104 (SRS1) shown in elemental colours. (C) The docking modes for campesterol in the Oryza CYP724B1 (magenta) and the Arabidopsis CYP90B1 and Oryza CYP90B2 (orange) predicted structures are shown with substrate contacts ˚ of this substrate, identical side chains shown in green and variable side within 3.0 A chains in elemental colours. Side chains predicted to cause campesterol to bind in different orientations correspond to H84, Q96, E97 (all in SRS1) in Oryza CYP724B1 versus M318/318 (SRS4) and F498/499 (SRS6) in Arabidopsis CYP90B1 and Oryza CYP90B2. (D) The docking modes for campesterol in the Oryza CYP724B1 (magenta) and the Arabidopsis CYP90B1 and Oryza CYP90B2 (orange) predicted structures ˚ of the substrate (in either orientation) are shown with identical residues within 4.5 A in green. Variable side chains shown in panel (C) are not included.

P450 families, comparisons of the two CYP90B proteins with Oryza CYP724B1 (28–29% overall identity) show that they have many contiguous identities throughout the lengths of their coding sequences, significant conservations in SRS3 (5/8 identities) and SRS5 (7/10 identities), moderate conservation in SRS1 (10/22 identities) and very low conservations in SRS2

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(2/7 identities), SRS4 (6/19 identities) and SRS6 (3/7 identities). Overlays of CYP90B1, CYP90B2 and CYP724B1 (Fig. 3C and D) indicate that many regions of their catalytic sites are highly conserved (green side chains in Fig. 3D). Differences in their catalytic sites that are likely to explain the alternate binding modes of campesterol in CYP90B proteins (orange in Fig. 3A) versus the CYP724B protein (magenta in Fig. 3A) occur in SRS1 (His84, Gln96, Glu97 in CYP724B1), SRS4 (Met318 in CYP90B proteins), and SRS6 (Phe498/Phe499 in CYP90B proteins) and are shown in Fig. 3C. These, in fact, correspond to three of the regions having lower conservations in our CYP90B1–CYP90B2–CYP724B1 alignment and provide evidence that the ability of all three proteins to hydroxylate this particular brassinosteroid in the same way has evolved from a common ancestral sequence by sequence variations in relatively constrained regions. With our previous text indicating that the CYP90C subfamily is absent in Oryza, comparisons of Arabidopsis CYP90C1 with the remaining unassigned Oryza CYP90A and CYP90D subfamily members indicate that these sequences are only 34–43% identical with no extensive conservation in the SRS regions. Previous suggestions in the literature that the missing rice CYP90C1 activity is mediated by one of the eight rice CYP92A subfamily members can be excluded based on the fact that all six SRS regions are highly diverged in these two subfamilies. In the sixth set of examples, comparisons of less closely related P450s whose functions have been defined, such as Oryza CYP81A6 in herbicide catabolism (Pan et al., 2006) and Arabidopsis CYP81F subfamily members in glucosinolate production (Bednarek et al., 2009; Pfalz et al., 2009), have indicated that the differing activities of these proteins arise from substantial subfamily divergence in all SRS except SRS6. Yet, as their phylogenetic classifications reflect, the remainder of the CYP81A and CYP81F proteins maintain significant (40–42%) overall identity between these subfamilies (Fig. 4). Stepping beyond phylogenetic comparisons, our structural predictions highlight several sets of P450s that are more similar in their catalytic sites than their overall sequence identities might reflect. When viewed from this three-dimensional perspective, extensive conservation in predicted substrate contact residues can provide suggestions for putative substrates for functionally uncharacterized Arabidopsis and Oryza P450s (e.g. CYP84A5 and CYP84A6 in Fig. 2) as well as information on variations in substrate positioning (e.g. CYP98A4 in Fig. 1, CYP84A4 in Fig. 2, CYP90B1 vs. CYP724B1 in Fig. 3C) that may result in different modifications or specific activities among even the most closely related P450s in these species. Contrasting with this, substantial divergence in predicted substrate contact

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CYP724A1 (A) CYP724B1 (R) CYP90B1 (A) CYP90B2 (R) CYP72C1 (A) CYP734A1 (A) CYP734A2 (R) CYP734A6 (R) CYP734A5 (R) CYP734A4 (R) CYP85A1 (A) CYP85A2 (A) CYP85A1 (R) CYP90A1 (A) CYP90A3 (R) CYP90C1 (A) CYP90D1 (A) CYP90D2 (R) CYP90D3 (R)

Fig. 4.

Phylogenetic relationships of CYP90 and CYP724 sequences.

residues (e.g. CYP724A and CYP724B proteins in Fig. 3A) can discriminate between distantly related proteins in the same subfamily and exclude the possibility that they are capable of metabolizing similar substrates. While computationally intensive, these perspectives argue that the time is now right for structural comparisons within individual P450 subfamilies to determine whether various members of some P450 subfamilies mediate similar or dissimilar reactions. With this predictive information and targeted biochemical analyses, it will become increasingly apparent which P450 subfamilies are especially prone to duplications and neofunctionalizations leading to the evolution of new catalytic activities and, potentially, the production of new plant defence compounds. It will also be more apparent which P450 subfamilies have maintained the levels of catalytic-site conservations needed for retention of various essential activities.

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V. CONCLUSIONS Phylogenetic comparisons among the vast collection of plant P450s have helped enormously to subdivide this large and diverse gene family into tractable groups. But, because variations in different external and internal regions of P450 molecules can have dramatically different effects on the catalytic activities of P450s, the substrate specificities of individual enzymes do not necessarily map to their phylogenetic classifications. Some within highly conserved subfamilies retain highly conserved catalytic sites and common functions while some vary in just enough catalytic-site side chains that they have developed new functions. Some within less conserved subfamilies have so few conserved catalytic-site residues that they handle dramatically different substrates. Some within divergent families and subfamilies have maintained enough commonality in their catalytic-site residues that is it clear that they evolved from a common ancestor in ways that have allowed them to modify common substrates even while positioning these differently in their catalytic sites. Other catalytic sites are so divergent that it is not possible to piece together their evolutionary relationships except by comparing their overall sequences and their genomic organizations. Given the growing number of plant P450s that are being annotated and current limits in the number of monooxygenases that can be functionally characterized in various heterologous expression systems (Duan and Schuler, 2006), it would seem that a cohesive mixture of phylogenetic and structural analysis can substantially advance our understanding of this botanically important group of enzymes. With information on the SRS variations in representative dicot and monocot P450s now accessible from genome sequencing and protein structure predictions, it will soon become possible to define the limits on the evolution of essential activities and the catalytic-site variations not likely to affect activities. With high-throughput substrate prediction programs being developed for use with P450 molecular models, it will also become possible to assign function to the novel monooxygenases now being identified in medicinal and other plant genomes.

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