Metabolons involving plant cytochrome P450s - Springer Link

Phytochem Rev (2006) 5:459–472 DOI 10.1007/s11101-006-9014-4

Metabolons involving plant cytochrome P450s Lyle Ralston Æ Oliver Yu

Received: 14 January 2006 / Accepted: 7 July 2006 / Published online: 28 October 2006 Springer Science+Business Media B.V. 2006

Abstract Arranging biological processes into ‘‘compartments’’ is a key feature of all eukaryotic cells. Through this mechanism, cells can drastically increase metabolic efficiency and manage complex cellular processes more efficiently, saving space and energy. Compartmentation at the molecular level is mediated by metabolons. A metabolon is an ordered protein complex of sequential metabolic enzymes and associated cellular structural elements. The sub-cellular organization of enzymes involved in the synthesis and storage of plant natural products appears to involve the anchoring of metabolons by cytochrome P450 monooxygenases (P450s) to specific domains of the endoplasmic reticulum (ER) membrane. This review focuses on the current evidence supporting the organization of metabolons around P450s on the surface of the ER. We outline direct and indirect experimental data that describes P450 enzymes in the phenylpropanoid, flavonoid, cyanogenic glucoside, and other L. Ralston Sigma-Aldrich Biotechnology, Life Science and High Technology Center, 2909 Laclede Avenue, St. Louis, MO 63103, USA O. Yu (&) Donald Danforth Plant Science Center, 975 North Warson Road, St. Louis, Missouri 63132, USA e-mail: [email protected]

biosynthetic pathways. We also discuss the limitations and future directions of metabolon research and the potential for application to metabolic engineering endeavors. Keywords Cytochrome P450 Æ Metabolon Æ Enzyme interaction Æ ER localization Æ Cytochrome P450 reductase Abbreviations 4CL 4-Coumarate: CoA ligase AFM Atomic force microscopy C4H Cinnamate 4-hydroxylase CHI Chalcone isomerase CHS Chalcone synthase CPR NADPH-cytochrome P450 reductase DFR Dihydroflavonol reductase ER Endoplasmic reticulum F3H Flavanone 3b-hydroxylase F3¢H Flavonoid 3¢-hydroxylase F3¢5¢H Flavonoid 3¢,5¢-hydroxylase FLIM Fluorescence lifetime imaging microscopy FRET Fluorescence energy resonance transfer I2¢H Isoflavone 2¢-hydroxylase IFS Isoflavone synthase IOMT Isoflavone O-methyltransferase P450 Cytochrome P450 monooxygenase PAL Phenylalanine ammonia-lyase

123

460

Introduction The ability to partition biological processes between discrete compartments is a major evolutionary advantage of eukaryotes. Metabolic processes in higher organisms are compartmentalized at multiple levels ranging from the macroorganizational level of a tissue or organ, to organization within different cell types, to subcellular organization at the level of the organelle, to molecular organization at the level of the multienzyme complex or metabolon. Compartmentalization at multiple levels allows an organism to manage various biological processes more precisely and with greater metabolic efficiency. The use of metabolons extends the coordination of metabolic compartmentalization to the molecular level. Paul Srere first defined a metabolon as a ‘‘supramolecular complex of sequential metabolic enzymes and cellular structural elements’’ (Srere 1985). Srere posited a number of characteristics as being inherent to metabolon structure and function: that component enzymes would channel substrates for increased catalytic efficiency; that flux through the metabolon would be regulated by the association and dissociation of component enzymes; that these component enzymes would form interactions with structural elements of the cell; and that assembly of the metabolon would be controlled, at least in part, through the coordinate regulation of the genes encoding its component enzymes. Such organization at the molecular level offers a number of potential benefits. The advantages of channeling intermediates through a metabolon include the ability to attain high local concentrations of pathway intermediates, bypass kinetic constraints resulting from diffusion of intermediates into the bulk solvent, decrease transition time of intermediates between the active sites of sequential enzymes, protect labile or reactive intermediates from the bulk solvent, and coordinate the metabolic flux through pathways with shared enzymes and/or intermediates. Classic examples of metabolons are found in both prokaryotes and eukaryotes (Srere 1987). Metabolons play well-established roles in the synthesis of amino acids, proteins, and nucleic acids (Srere 1987; Mendes et al. 1995). For

123

Phytochem Rev (2006) 5:459–472

example, the tryptophan synthase complex is a classical metabolon that forms an intra-molecular hydrophobic tunnel to channel indole between the active sites of the two subunits of the enzyme complex (Anderson et al. 1995; Miles 2001). In addition, several well-defined examples of metabolons exist in plants. Many of these examples come from central or primary metabolic pathways and include enzyme complexes of the Calvin– Benson cycle (Graciet et al. 2004; Winkel 2004), peroxisomal enzymes involved in the photorespiratory cycle (Reumann 2000), and enzymes involved in polyamine biosynthesis (Panicot et al. 2002). In many cases, metabolon formation appears to involve specific interactions between operationally soluble enzymes, cytoskeletal elements, or membrane proteins (Mendes et al. 1995; Srere 2000). Growing experimental evidence suggests that the enzymes of various natural product biosynthetic pathways in plants may be organized as metabolons on the cytoplasmic surface of the endoplasmic reticulum (ER). In these metabolons, cytochrome P450 monooxygenases (P450s) are hypothesized to serve as nucleation points that anchor the metabolon to a specific region within the cell (Winkel 2004; Jorgensen et al. 2005). The majority of plant P450s are tethered to the ER via a membrane-spanning amino-terminal domain (Chapple 1998) (Fig. 1). This localization provides a platform for potential interactions between proteins forming a given metabolon. The P450-anchored metabolons are suspected to associate with specific domains on ER membranes, thus forming ‘‘metabolic centers’’ (Galili et al. 1998). This phenomenon has been hypothesized to be facilitated by small accessory proteins that mediate interactions between component enzymes of a metabolon and cytoskeletal elements (Chuong et al. 2004; Graciet et al. 2004; Winkel 2004). Since cellular processes must constantly adjust in response to various environmental and physiological stimuli, dynamic formation and dissociation of metabolons may provide an additional level of metabolic regulation in biosynthetic pathways. This review focuses on the organization of metabolons around P450s in plant natural product pathways. Beyond a metabolic role, P450s are


461

Fig. 1 A plant cytochrome P450 localized to the cortical ER membrane. (A) Soybean isoflavone synthase (IFS, CYP93C1) was fused in-frame with a yellow fluorescent protein (EYFP) and transiently expressed in tobacco leaf epidermal cells. IFS was localized to the ER network (yellow color, Ralston and Yu, unpublished data). (B) As a

control, soybean chalcone isomerase (CHI) was fused inframe with a cyan fluorescent protein (ECFP) and transiently expressed in tobacco leaf epidermal cells. Soluble CHI was localized to the cytoplasm and nucleus (green color). The size bar is 10 lm

also thought to play a structural role in these complexes, providing a platform for their partners to establish and maintain protein–protein interactions. We will discuss the direct and indirect evidence supporting the hypothesis of P450-anchored metabolons in several plant natural product pathways.

fatty acid x-2 hydroxylase P450 BM3 (CYP102A1) from bacterium Bacillus megaterium, provide insight on the interaction between the P450 and CPR. The multifunctional P450s evolved through the fusion of the P450 and CPR enzymes. This type of P450 consists of two discrete domains that can be functionally separated genetically and proteolytically. The first 472 amino-terminal residues of P450 BM3 comprise the monooxygenase domain, while the remainder of the enzyme makes up the reductase domain (Munro et al. 2002). In P450 BM3, NADPH reduces a FAD flavin to a transient hydroquinone, and a flavin mononucleotide (FMN)-binding domain shuttles electrons between the FAD in the CPR domain and the heme in the P450 domain (Sevrioukova et al. 1999). The fusion provides a rare insight into the mechanism of the coupling of the two units and how protein–protein interactions facilitate the transfer of electrons (for reviews see (Munro et al. 2002)). Structural and biochemical data from P450 BM3 suggests that the rate of electron transfer between NADPH and FAD in the bipartate protein is much faster than the observed catalytic activity of the isolated monooxygenase domain (Munro et al. 1996). Two basic residues, Lys572 and Lys580, stabilize the hydroquinone-form of FAD and are essential for efficient coupling of the electron transfer chain (Sevrioukova et al. 1999; Murataliev and

Interactions between P450s and NADPH-cytochrome P450 reductases Given the central role of P450s in anchoring metabolons of plant natural product biosynthesis, the molecular architecture of the P450s provides a framework for organizing larger macromolecular complexes. All P450s rely on a heme co-factor to split molecular oxygen, inserting one oxygen atom to the substrate and reducing the other oxygen atom to water. To accomplish this reaction, a coupled, stepwise electron supply originating from NAD(P)H is indispensable (Denisov et al. 2005). The electron supply is provided by NADPH-cytochrome P450 reductase (CPR) (Paine et al. 2005). Depending on the type of P450, the CPR is either fused to the P450 to form a multifunctional protein or operates as a separate membrane-bound protein. Structural and functional studies of the multifunctional or ‘‘self-sufficient’’ P450s, such as the

123

462

Feyereisen 2000). Overall, for any P450 to be functional, a tightly coupled CPR is essential. In contrast to the multifunctional P450s, the typical eukaryotic P450 interacts with a separate membrane-bound CPR (Paine et al. 2005). The amino-terminal region of CPR (approximately 60 amino acids in length) is a hydrophobic membrane-spanning domain that anchors CPR to the cytoplasmic surface of the ER. Similar to the carboxyl-terminal region of the P450 BM3, the function of the membrane-bound CPR is to ferry electrons from NAD(P)H through FAD and FMN to the P450 (Murataliev et al. 2004). Compared to the vast number of P450 genes that exist in plant genomes, the number of CPR genes is much lower, and sequence homology is more highly conserved than in P450s (Koopmann and Hahlbrock 1997; Urban et al. 1997). This suggests that a limited number of CPRs would couple with a diverse array of P450 partners and, by extension, interact with a variety of metabolons. Interactions between CPR and P450s have been investigated mainly in mammalian systems. A rat CPR without the amino-terminal membrane domain has been crystallized (Wang et al. 1997). The enzyme is composed of three domains: an FAD-binding domain similar to ferredoxinNADP+ reductase, an FMN-binding domain similar to flavodoxins, and a linear linker that connects the other two domains. The linker domain is responsible for the positioning of the cofactors FMN and FAD in close proximity to the heme of the P450 to facilitate efficient electron transfer. Mutations in the linker region can cause significant differences in the relative positions of the two flavin domains and significantly reduce a P450’s catalytic activities (Hubbard et al. 2001). Similarly, removing the membrane anchor of a mammalian CPR will produce a soluble and enzymatically active CPR. However, the soluble CPR will be unable to support the activity of a P450 in vitro, due to the uncoupling of the electron transfer chain (Venkateswarlu et al. 1998). The direct interaction of CPR with P450s was visualized by atomic force microscopy (AFM) on a reconstituted phospholipid bilayer disk containing CPR and the P450 CYP2B4 (Bayburt et al. 1998; Bayburt and Sligar 2002). CYP2B4 was found to protrude above the lipid surface,

123


3.5 nm. This is thought to orient the CPR such that both the FMN- and FAD-binding domains lie close to the membrane surface, allowing a close interaction between FMN and the P450 heme (Wang et al. 1997). The lipid bilayer of the ER membrane may also contribute to interactions between CPR and P450 by altering the composition of the bilayers to provide a relatively stable support for the enzyme complexes when needed (Ingelman-Sundberg et al. 1981, 1983). More importantly, a group of basic residues, Arg45, Lys46, Lys47, and Lys48, which lies at the beginning of FMN-binding domain, may interact with the phospholipid head groups of the membrane and restrict the movement of CPR on the membrane surface (Wang et al. 1997; Zhao et al. 1999). Similarly, a second group of basic residues (Lys72, Lys74, Lys75, Arg78, Arg97, Lys100, His103, and Arg108) at the surface of the FMNbinding domain may provide an additional membrane-binding site. Surface residues on the P450 are also likely to contribute to the CPR–P450 interaction. Specific lysine and arginine residues on rat CYP1A1 are shown to be involved in the formation of an electron transfer complex with CPR (Shimizu et al. 1991). Similarly, mammalian CYP2B1 and CYP2B4 may also rely on complementary electrostatic charges to bind with CPR (Shen and Strobel 1993; Bridges et al. 1998). Site-directed mutagenesis of CYP2B4 identified a series of lysine and arginine residues on the proximal surface of the heme region that interact with CPR’s FMN region (Bridges et al. 1998). Moreover, chemical cross-linking studies indicate that several acidic residues on CPR are involved in the interaction with CYP1A1 (Nadler and Strobel 1991). Other evidence also suggests that the FMN-binding domain is the docking surface of the two enzymes, as with P450 BM3. Electrostatic potential measurement of the FMN-binding domain of a human CPR shows clusters of acidic residues that could form ion-pair interactions with a P450 (Shen and Strobel 1995). In fact, a decreased association (Kd 10–90 nM) between rabbit CPR and CYP2B4 was observed when the ionic strength of the buffer was increased 10-fold (Davydov et al. 2000). Mutations of the clusters located at the opposite side of the FMN domain significantly reduces


463

Arguably the strongest physical evidence for the existence of metabolic channels in a plant natural product pathway involves the interaction between the first two enzymes of the phenylpropanoid pathway (Fig. 2A). The phenylpropanoid path-

way exists in all higher plants and is a major consumer of energy and carbon fixed by photosynthesis (Werck-Reichhart and Feyereisen 2000). This pathway produces a myriad of structurally diverse phenolic compounds, including lignins, stilbenes, aurones, flavonoids, and isoflavonoids. These compounds serve as signal molecules, phytoalexins, anti-feedants, pigments, UV protectants, and cell wall components in plants. Since this pathway consumes large amounts of biochemical resources, phenylpropanoid metabolism is tightly regulated both transcriptionally and post-transcriptionally (Weisshaar and Jenkins 1998). Metabolic channeling between enzymes at the entry point of pathway is suggested to provide an additional level of regulation. The entry point enzyme of the phenylpropanoid pathway is phenylalanine ammonia-lyase (PAL). PAL commits phenylalanine to the phenylpropanoid pathway, deaminating phenylalanine and converting it into trans-cinnamate and ammonia. In all species examined so far, PAL is encoded by a multi-gene family (Dixon et al. 2002). Based on differential gene expression patterns, it has long been suspected that the

Fig. 2 Schematic diagram of the phenylpropanoid (A) and dhurrin pathways (B). The P450 enzymes are in red italics. Enzyme abbreviations are listed in the text, except

for CHR, chalcone reductase; STS, stilbene synthase; HID, 2-hydroxyisoflavanone dehydratase; and IFR, isoflavone reductase

P450 activity, suggesting that the P450 may partially ‘‘engulf’’ CPR at the FMN domain to bring the heme and FMN in closer contact (Zhao et al. 1999). In spite of the above evidence, the proposed metabolon model has to reconcile with the fact that, at least in mammalian cells, P450s are present in a 10–25-fold molar excess over CPR (Estabrook et al. 1971; Peterson et al. 1976). This ratio dictates that any association of CPR with a given P450 has to be transient and dynamic to allow multiple P450s from different pathways to have access to this essential partner. Such interactions in plant cells are likely to be more complicated due to the higher number of P450 genes found in plant genomes.

Entry into the phenylpropanoid pathway: the CYP73A-related metabolon

123

464

various PAL isoforms are activated by different environmental signals to divert phenylalanine into different branches of the phenylpropanoid pathway (Kao et al. 2002). The second enzyme of the phenylpropanoid pathway is the P450 cinnamate 4-hydroxylase (CYP73A, C4H). C4H adds a hydroxyl group at the C4 (para) position of trans-cinnamate to form p-coumarate (Russell and Conn 1967; Schoch et al. 2003). C4H is encoded by a single copy gene in Arabidopsis and occurs in only one or two copies in other species studied to date (Plant Cytochrome P450 Database; http://www.drnelson.utmem.edu/P450dbplant.html). C4H was the first plant P450 to be functionally characterized, and it is among the most extensively studied of the plant P450s (Chapple 1998). PAL exists as an operationally soluble homotetramer with no obvious membrane localization domains (Wanner et al. 1995); however, a number of reports describe the association of PAL with endomembranes in plant cells. In 1975, based on sub-cellular fractionation and microsome enzyme activity assays, Czichi and Kindl reported that PAL and C4H may assemble on microsomal membranes as an enzyme complex (Czichi and Kindl 1975). While C4H activity is located exclusively in the microsomal fraction of plant cells and PAL activity resides in the soluble fraction of plant cell extracts, they noted that both PAL and C4H activities co-localized to the microsomal fraction. Substrate feeding experiments suggested that cinnamate produced from [3H]-phenylalanine by PAL is preferred over exogenously added [14C]-cinnamate by C4H. Hence, the majority of the C4H product, p-coumarate, was derived from radiolabeled phenylalanine (Czichi and Kindl 1977). Further feeding assays carried out by others demonstrated that disrupting the microsomal structure, which may disrupt the interaction of the two enzymes, led to the increased conversion of exogenously added cinnamate conversion to p-coumarate (Hrazdina and Wagner 1985). Taken together, these earlier experiments provide strong indirect evidence that PAL and C4H are co-localized on the cytoplasmic surface of the ER and that cinnamate is channeled directly to the active site of C4H from PAL.

123


Rick Dixon’s lab has produced a good deal of evidence for the existence of a tobacco PAL-C4H metabolon. Rasmussen et al. demonstrated that cinnamate produced from [3H]-phenylalanine does not equilibrate with exogenously added [14C]-cinnamate, indicating channeling between PAL and C4H (Rasmussen and Dixon 1999). Further, sub-cellular fractionation and protein gel blot analysis demonstrated that tobacco PAL1 is localized to both the soluble and microsomal protein fractions, whereas the PAL2 isozyme is strictly localized to the soluble fraction. More recently, Achinine et al. provided more evidence of direct enzyme interactions between PAL and C4H. They used a combination of biochemical and microscopic techniques to study the localization of PAL1, PAL2, and C4H (Achnine et al. 2004). Using these techniques, they verified the sub-cellular localization of PAL1 and PAL2. Moreover, they showed that over-expression of C4H resulted in the ER localization of both PAL1 and PAL2. This suggests that C4H itself may be responsible for the membrane association of PAL. Surprisingly, co-expression of unlabeled PAL1 with PAL2-GFP resulted in a shift of fluorescence from ER membranes to the cytoplasm in C4H over-expressing plants, whereas coexpression of unlabeled PAL2 with PAL1-GFP did not affect PAL1-GFP localization. This indicates that PAL1 has a higher affinity for its membrane localization site than PAL2. Fluorescence energy resonance transfer (FRET) studies based on dual-labeling immuno-fluorescence showed the PAL and C4H are located within ˚ of each other on the membrane. In addi100 A tion, negative photo-bleaching results may suggest that the connection between PAL and C4H is fluid (Achnine et al. 2004). In whole, there is extensive compelling evidence suggesting that C4H anchors an ER-localized metabolon at the entry point of the phenylpropanoid pathway. However, at least one example can be found in the literature disputing the existence of a metabolon between PAL and C4H. Ro and Douglas argue that enzyme kinetics and biochemical coupling of PAL and C4H are enough to drive carbon flux into the phenylpropanoid pathway (Ro and Douglas 2004). They reconstituted the entry point of the pathway by


expressing PAL, C4H, and CPR in yeast. They then fed the engineered yeast [3H]-phenylalanine and [14C]-cinnamate simultaneously. In this system, inhibition of C4H by P450 inhibitors indeed reduced PAL activity, demonstrating biochemical coupling. However, unlike experiments using plant microsome fractions, the two radiolabeled substrates showed equal accessibility to C4H, indicating that no channeling occurs between PAL and C4H. Earlier analysis using transgenic plants supported this biochemical coupling theory: when the C4H gene was silenced in tobacco, PAL activity was significantly reduced while PAL silencing did not disturb C4H activity. This data provides further evidence of a feedback loop at the entry point of the phenylpropanoid pathway (Blount et al. 2000). Clearly, biochemical coupling is part of the regulation mechanism between PAL and C4H. Yet the physical interactions between these enzymes inside plant cells should not be excluded based on experiments in yeast. Cytoskeletal elements and accessory proteins critical for interactions between PAL and C4H may well be lacking in a heterologous host. It is possible that at the entry point of the phenylpropanoid pathway, both regulation mechanisms co-exist inside plant cells.

Flavonoid biosynthesis: CYP75B-related metabolon While the majority of the carbon flux into the phenylpropanoid pathway is directed towards lignin production (discussed below), the second greatest carbon sink from this pathway is flavonoid biosynthesis. Flavonoid compounds are produced by the coordinate action of a series of operationally soluble and membrane-bound cytoplasmic enzymes (Fig. 2A) (Winkel-Shirley 2001a). Soluble enzymes of the flavonoid branch pathway include 4-coumarate: CoA ligase (4CL), chalcone synthase (CHS), chalcone isomerase (CHI), flavanone 3b-hydroxylase (F3H), and dihydroflavonol reductase (DFR). A number of P450s also participate in this branch of the pathway. Flavonoid 3¢-hydroxylase (CYP75B, F3¢H) modifies the B-ring of the flavanone backbone at the C3¢ position. Another P450, flavonoid 3¢,5¢-

465

hydroxylase (CYP75A, F3¢5¢H), is found in a limited number of species and modifies the B-ring at the C5¢ position (Winkel-Shirley 2001b). There is strong evidence suggesting that the operationally soluble enzymes of the flavonoid branch pathway participate in metabolons. In a yeast two-hybrid assay, Burbulis and WinkelShirley demonstrated that CHS, CHI, and DFR interact with one another in an orientationdependent manner (Burbulis and Winkel-Shirley 1999). Affinity chromatography and immunoprecipitation assays also demonstrated interactions between CHS, CHI, and F3H in lysates from Arabidopsis seedlings (Burbulis and WinkelShirley 1999). Although three-dimensional structures are available for CHS and CHI (Ferrer et al. 1999; Jez et al. 2000), no structural model of this proposed macromolecular complex has been published. However, mechanistic studies of CHS and CHI suggest the need for metabolic channeling in this pathway. The non-enzymatic cyclization of chalcones into flavanones readily occurs in solution but yields an enantiomeric mix of biologically inactive and active isomers. Channeling between CHS and CHI would prevent the formation of mixed isomers (Jez et al. 2002). Interestingly, the catalytic efficiency of CHI (kcat/ Km = 109 M–1 min–1) approaches the diffusion limit, so one might ask why CHI would need metabolites to be directed towards it. One possible answer is that the moderately lipophilic nature of chalcones and flavanones may require channeling to limit the potential for sequestration in cellular membranes. Moreover, in legumes, channeling through the metabolon may ‘‘slow’’ the activity of CHI and protect the chalcone pool from complete conversion into naringenin (Jez et al. 2002). There is no direct evidence that the complex formed by CHS, CHI, F3H, and DFR interacts with P450s. However, if these enzyme do form such an interaction, the most likely candidate is F3¢H (or perhaps F3¢5¢H, if it is expressed). The complex could also involve the upstream P450 C4H, thus forming a metabolon that encompasses the majority of the enzymes of the general phenylpropanoid and flavonoid branch pathways (Winkel 2004; Jorgensen et al. 2005). There is interesting indirect evidence suggesting that CHS/

123

466

CHI-related metabolons are indeed localized to the ER. First, CHI contains a motif that indicates the enzyme might be modified post-translationally by a fatty acid moiety with a thiol-sensitive linkage. This would provide a mechanism for the membrane localization of this soluble enzyme (Burbulis and Winkel-Shirley 1999). Earlier reports indicate that in the hypocotyl epidermis of buckwheat (Fagopyrum esculentum), CHS activity and protein were detected at the cytoplasmic face of the rough ER, based on linear sucrose density gradient fractionation, immuno-blots, and immuno-gold analysis (Hrazdina et al. 1987). Finally, membrane localization of CHS, and colocalization of CHS and CHI were observed at the ER and tonoplast in Arabidopsis root cells (Saslowsky and Winkel-Shirley 2001). The antibodies detected an electron-dense region with membrane structures. In the Arabidopsis tt7 mutant in which the P450 F3¢H is deleted, the electron-dense regions containing these two enzymes were not detected. However, CHS and CHI were still found to co-localize to the ER and tonoplast in the tt7 mutant, suggesting that other proteins may function in recruiting the soluble flavonoid pathway enzymes to membranes (Saslowsky and Winkel-Shirley 2001). Interestingly, recent data suggests that flavonoids and CHS and CHI proteins are also localized to nucleus of Arabidopsis (Saslowsky et al. 2005). However, the implications of this discovery are not yet clear. Collectively, the evidence amassed over the past 30 years suggests the existence of metabolons involving enzymes of the general phenylpropanoid and flavonoid branch pathways that are anchored to the cytoplasmic surface of the ER by P450s.

Isoflavonoid biosynthesis: CYP93C-related metabolons In legumes, the majority of the naringenin produced by CHS is converted to isoflavonoids by the P450 isoflavone synthase (CYP93C, IFS) (Yu et al. 2003). Isoflavonoids are major defense compounds for legumes. While isoflavonoids are constitutively produced at low levels, their biosynthesis is drastically induced upon pathogen

123


infection. During the defense response, isoflavonoids are further metabolized into more complex and more potent phytoalexins by a set of legumespecific enzymes (Dixon et al. 2002). In the Medicago family, the first such modification is the methylation of the 4¢-hydroxy on the B-ring of the isoflavones. This step is mediated by an operationally soluble isoflavone-O-methyltransferase (IOMT). Liu et al. observed that IOMT from alfalfa (Medicago sativa) was translocated to the ER upon elicitor treatment, co-localizing with IFS (Liu and Dixon 2001). They also noted that feeding studies in alfalfa cell suspension cultures suggested channeling between IFS and the IOMT. Radiolabel from [3H]-liquiritigenin was preferentially incorporated into formononetin, the product of IOMT, as well as other downstream isoflavonoids. However, radiolabel from daidzein, the in vitro substrate of IOMT, was not incorporated into downstream isoflavonoids. This data suggests that the methyltransferase is closely associated with IFS, facilitating rapid methylation and stabilization of the product of IFS. This association was also used to explain the observed differences in IOMT regio-specificity between in vivo and in vitro assays (Liu and Dixon 2001). However, different types of IOMTs have been cloned, indicating that these differences in regio-specificity may instead be caused by the activity of different IOMT enzymes (Akashi et al. 2000, 2003). Liu et al. provided a model suggesting that intermediates of the isoflavonoid pathway could rapidly flow from one enzyme center anchored by IFS to next enzyme center in a metabolon anchored by another P450, isoflavone 2¢-hydroxylase (CYP81E, I2¢H), and eventually enter into the vacuole (Liu and Dixon 2001). This process is more complicated in other legume species. In these species, the product of I2¢H is prenylated by plastid-associated prenyltransferases to form pterocarpans and furanocoumarins before entering the vacuole (Dhillon and Brown 1976). This must require the shuttling of compounds between different organelles for the metabolon model to hold true. The association of IFS with upstream enzymes such as CHI and CHS can only be implied. Several groups have shown that efforts to engineer isoflavonoid synthesis in non-legume plants by


introducing IFS resulted in a disproportionate decrease in flavonoid production, which cannot be explained by a shift in metabolic flux alone (Yu et al. 2000; Liu et al. 2002). In transgenic Arabidopsis expressing IFS, the production of genistein can be increased 3-fold by irradiation with UV light, but the ratio of genistein in total flavonoid levels actually decreased. This suggests that the pool of naringenin substrate is not equally accessible to the flavonoid and isoflavonoid branches of the pathway (Yu et al. 2000). Similar experiments using transgenic tobacco and maize cell cultures demonstrated similar ‘‘compartmentalization’’ of the naringenin substrate (Yu et al. 2000). Additionally, the treatment of tobacco leaves with UV light to increase naringenin levels resulted in elevated flavonol levels but failed to raise anthocyanin or isoflavone levels. This data suggests that flux through the phenylpropanoid pathway is tightly channeled to flavonol production in tobacco leaves, and IFS has difficulty accessing this substrate. To delineate enzyme interactions between IFS and endogenous flavonoid pathway genes, a CHI from alfalfa, shown to be an legume-specific isomerase by its ability to convert isoliquiritigenin into liquiritigenin, was transformed into Arabidopsis and then genetically crossed into a transgenic Arabidopsis line carrying the IFS gene (Liu et al. 2002). The combination of genes enhanced genistein accumulation moderately. However, flavonol accumulation was significantly reduced compared to transgenic plants carrying only the legume-specific CHI. The disproportionate reduction of flavonol biosynthesis caused by the presence of IFS further confirmed that the flux of substrate is preferentially channeled towards endogenous flavonoid biosynthesis (Liu et al. 2002). One possible explanation offered for this outcome is that the membrane-localized IFS might interfere with the assembly of metabolic complexes dedicated to flavonoid biosynthesis, which may be anchored to the ER by the P450 F3¢H in Arabidopsis (Yu et al. 2000). When the IFS and CHI were introduced into a tt6/tt3 double mutant background, genistein accumulation was enhanced by up to 30-fold as compared to plants expressing IFS alone. This mutant has structural defects in both F3¢H and DFR genes and is thus

467

blocked in flavonol and anthocyanin production. Again, this suggests that the bottleneck for isoflavone production in Arabidopsis is competition for flavanones between IFS and endogenous flavonol biosynthetic enzymes (Liu et al. 2002). This brings us to an interesting topic regarding the regulation of the competing branch pathways. In legumes, where the flavonoid and isoflavonoid pathways co-exist, the flux of flavanone intermediates should be differentially divided into the two pathways in response to abiotic and biotic stresses. Plants can activate the expression of a subset of genes that divert the precursor phenylalanine into specific products of the pathway using a number of relatively well-defined transcription factors (Yu et al. 2003). However, metabolons could play a major role in controlling flux through the pathway. In fact, two functionally different CHI exist in legumes (Shimada et al. 2003; Ralston et al. 2005): the type I CHI exists in all higher plants and converts tretrahydroxychalcones into the flavanone naringenin, while the legume specific type II CHI converts both tetrahydroxy-chalcones and trihydroxy-chalcones into naringenin and liquiritigenin, respectively. The two types of CHI are differentially expressed, and only the type II CHI is induced by defense signals (Ralston et al. 2005). It’s possible that the type II CHI, together with other isoflavone biosynthetic enzymes may form their own metabolic channels, independent of the flavonoid channels described in Arabidopsis (Fig. 3) (Winkel 2004; Yu and McGonigle 2005).

Cyanogenic glucoside biosynthesis: CYP79A/ CYP71E-related metabolons and others The biosynthesis of cyanogenic glucosides in sorghum adds another dimension to the involvement of P450s in metabolons. Dhurrin is a type of cyanogenic glucoside that is produced from the amino acid tyrosine by two P450s, CYP79A1 and CYP71E1, and the UDP-glucosyltransferase UGT85B1 (Fig. 2B) (Celenza 2001; Morant et al. 2003; Jorgensen et al. 2005). Feeding assays using the radioisotopically labeled substrate tyrosine and the intermediate Z-p-hydroxyphenylacetal doxime demonstrated clearly that the two P450s are

123

468

Fig. 3 Proposed models of the flavonoid (A) and isoflavonoid (B) metabolons. The enzyme abbreviations are the same as in Fig. 2, except for UFGT, UDP-glucose: anthocyanidin-3-O-glucosyl trasnsferase; LDOX, leucoanthocyanidin deoxygenase

highly channeled, with the majority of the radiolabel in dhurrin coming from tyrosine (Moller and Conn 1980). Epitope tagging and confocal laser scanning microscopy revealed that this metabolon is located in a distinct domain of the ER (Nielsen and Moller 2005). From an evolutionary point of view, the coupling of the three enzymes likely arose out of the need to prevent the highly labile and toxic p-hydroxymandelonitrile intermediate from releasing cyanide into the cytoplasm. Genetic engineering has provided further support for the existence of P450 anchored metabolons. When the entire cyanogenic glucoside pathway was introduced into Arabidopsis by expressing all three enzymes, the production of dhurrin had virtually no effect on the metabolic profiles of the host plants (Kristensen et al. 2005). This toxic product was effectively contained and sequestered in vacuoles. However, when only the two P450s were expressed in Arabidopsis, without the metabolon partner UGT85B1, plants showed a variety of toxicity-related phenotypes. Interestingly, when introduced separately into Arabidopsis, the first enzyme CYP79A1 was able to establish highly efficient interactions with down-

123


stream glucosinolate-producing enzymes, and altered the overall glucosinolate profile of Arabidopsis (Bak et al. 1999). With the rapid advances in the understanding of cellular and biochemical processes, metabolons have been invoked in other metabolic pathways. For example, the lignin biosynthetic enzymes have previously been proposed to form a metabolic grid, in which the flux could proceed in a relatively random manner (Humphreys and Chapple 2002; Boudet et al. 2003), however, recent molecular analysis, particularly the new enzyme kinetic and substrate specificity observations suggested that many of the routes through the initially proposed grid are unlikely to function in vivo, thus suggesting a more or less linear metabolic pathway. Dixon et al. proposed an interesting model suggesting that the linearity of enzyme reactions is the result of metabolic channeling among the key lignin enzymes (Dixon et al. 2001). The metabolons are thought to prevent the intermediates from coming into contact with competing enzymes that might lead to the development of a less efficient metabolic grid. However, direct evidence of enzyme interactions in this pathway is scarce. While the existence of metabolons has been postulated in other plant natural product pathways, including the alkaloid and terpenoid pathways (Chappell 1995; Memelink 2004), investigators have yet to produce compelling evidence of this phemonenon.

Conclusion and perspectives A considerable amount of evidence argues for the formation of metabolic P450-anchored enzyme complexes in plant natural product pathways. However, unlike the more conventional concept of the stable multienzyme complexes that are focused on completing one particular task more efficiently, the P450-related metabolons probably take on broader roles in regulating complicated pathways and even metabolic grids. Such functions require these metabolons to be able to accommodate the constantly changing environment both inside and outside of the cell. Therefore, most of these metabolons are likely to be very dynamic and transient. The assembly and


disassociation of enzymes within these complexes is expected to occur rapidly in response to internal and external stimuli. As a consequence, it has been very difficult to isolate and study these complexes. Traditional biochemical assays, including isotope dilution, substrate feeding, and sub-cellular fractionation, while valuable, are limited in their ability to provide direct evidence of enzyme interactions. Co-immunoprecipitation, protein cross-linking and affinity purifications are best suited for stable interactions (Blancaflor and Gilroy 2000); and yeast two-hybrid assays are not applicable for membrane bound P450s. Therefore, newer technologies, such as multiphoton confocal co-localization (Blancaflor and Gilroy 2000), FRET and fluorescence lifetime imaging microscopy (FLIM) analysis (Wallrabe and Periasamy 2005), split ubiquitin assays (Thaminy et al. 2004), protein fragment complementation assays (Subramaniam et al. 2001), and surface plasmon resonance assays (Magee et al. 2002) may provide direct and in vivo enzyme interaction analysis. If one or more plant P450 structures can be resolved by X-ray crystallography, it will certainly provide tremendous help in understanding of nature of these protein–protein interactions, since many of the soluble partners of P450s have already been resolved structurally. This will allow directed mutagenesis to be carried out to evaluate the interactions at the interface of these proteins. However, the crystallization of membrane-bound proteins is notoriously difficult and may require intensive labor and extensive modifications of the P450. In addition, direct visualization based on AFM (Janovjak et al. 2005) or cryo-EM (Chiu et al. 2005) is also possible if the enzyme complexes can be functionally reconstituted or isolated. Understanding the structure and function of the P450-related metabolons clearly is a considerable challenge. However, these complexes very likely provide another layer of cellular regulation mechanisms and play important roles in plant growth and development. The information obtained on metabolons will also guide new metabolic engineering efforts and allow the plant community to harvest the desired products with more certainty and higher efficiency.

469 Acknowledgements We would like to thank Dr. Joe Jez of the Danforth Center for the critical review of this manuscript and for many thoughtful discussions on the topic. The research in Dr. Yu’s lab is supported by the National Science Foundation (MCB0519634), United States Department of Agriculture (CSREES: 2005-05190), and Missouri Soybean Merchandising Council (02-242).

References Achnine L, Blancaflor EB, Rasmussen S, Dixon RA (2004) Colocalization of L-phenylalanine ammonialyase and cinnamate 4-hydroxylase for metabolic channeling in phenylpropanoid biosynthesis. Plant Cell 16:3098–3109 Akashi T, Sawada Y, Aoki T, Ayabe S (2000) New scheme of the biosynthesis of formononetin involving 2,7,4¢-trihydroxyisoflavanone but not daidzein as the methyl acceptor. Biosci Biotechnol Biochem 64:2276– 2279 Akashi T, Sawada Y, Shimada N, Sakurai N, Aoki T, Ayabe S (2003) cDNA cloning and biochemical characterization of S-adenosyl-L-methionine: 2,7,4¢trihydroxyisoflavanone 4¢-O-methyltransferase, a critical enzyme of the legume isoflavonoid phytoalexin pathway. Plant Cell Physiol 44:103–112 Anderson KS, Kim AY, Quillen JM, Sayers E, Yang XJ, Miles EW (1995) Kinetic characterization of channel impaired mutants of tryptophan synthase. J Biol Chem 270:29,936–29,944 Bak S, Olsen CE, Petersen BL, Moller BL, Halkier BA (1999) Metabolic engineering of p-hydroxybenzylglucosinolate in Arabidopsis by expression of the cyanogenic CYP79A1 from Sorghum bicolor. Plant J 20:663–671 Bayburt TH, Carlson JW, Sligar SG (1998) Reconstitution and imaging of a membrane protein in a nanometersize phospholipid bilayer. J Struct Biol 123:37–44 Bayburt TH, Sligar SG (2002) Single-molecule height measurements on microsomal cytochrome P450 in nanometer-scale phospholipid bilayer disks. Proc Natl Acad Sci USA 99:6725–6730 Blancaflor EB, Gilroy S (2000) Plant cell biology in the new millennium: new tools and new insights. Am J Bot 87:1547–1560 Blount JW, Korth KL, Masoud SA, Rasmussen S, Lamb C, Dixon RA (2000) Altering expression of cinnamic acid 4-hydroxylase in transgenic plants provides evidence for a feedback loop at the entry point into the phenylpropanoid pathway. Plant Physiol 122:107–116 Boudet AM, Kajita S, Grima-Pettenati J, Goffner D (2003) Lignins and lignocellulosics: a better control of synthesis for new and improved uses. Trends Plant Sci 8:576–581 Bridges A, Gruenke L, Chang YT, Vakser IA, Loew G, Waskell L (1998) Identification of the binding site on cytochrome P450 2B4 for cytochrome b5 and cytochrome P450 reductase. J Biol Chem 273:17,036– 17,049

123

470 Burbulis IE, Winkel-Shirley B (1999) Interactions among enzymes of the Arabidopsis flavonoid biosynthetic pathway. Proc Natl Acad Sci USA 96:12,929–12,934 Celenza JL (2001) Metabolism of tyrosine and tryptophan – new genes for old pathways. Curr Opin Plant Biol 4:234–240 Chappell J (1995) Biochemistry and molecular biology of the isoprenoid biosynthetic pathway in plants. Ann Rev Plant Physiol Mol Biol 46:521–547 Chapple C (1998) Molecular genetics analysis of plant cytochrome P450-dependent monooxygenases. Ann Rev Plant Physiol Mol Biol 49:311–343 Chiu W, Baker ML, Jiang W, Dougherty M, Schmid MF (2005) Electron cryomicroscopy of biological machines at subnanometer resolution. Structure 13:363–372 Chuong SD, Good AG, Taylor GJ, Freeman MC, Moorhead GB, Muench DG (2004) Large-scale identification of tubulin-binding proteins provides insight on subcellular trafficking, metabolic channeling, and signaling in plant cells. Mol Cell Proteomics 3:970–983 Czichi U, Kindl H (1975) Formation of p-coumaric acid and o-coumaric acid from L-phenylalanine by microsomal membrane fractions from potato: evidence of membrane-bound enzyme complexes. Planta 125:115–125 Czichi U, Kindl H (1977) Phenylalanine ammonia lyase and cinamic acid hydroxylases as assembled consecutive enzymes on microsomal membranes of cucumber cotyledons: cooperation and subcellular distribution. Planta 134:133–143 Davydov DR, Kariakin AA, Petushkova NA, Peterson JA (2000) Association of cytochromes P450 with their reductases: opposite sign of the electrostatic interactions in P450BM-3 as compared with the microsomal 2B4 system. Biochemistry 39:6489–6497 Denisov IG, Makris TM, Sligar SG, Schlichting I (2005) Structure and chemistry of cytochrome P450. Chem Rev 105:2253–2277 Dhillon DS, Brown SA (1976) Localization, purification, and characterization of dimethylallylpyrophosphate: umbelliferone dimethylallythransferase from Ruta graveolens. Arch Biochem Biophys 177:74–83 Dixon RA, Achnine L, Kota P, Liu C-J, Reddy MSS, Wang LJ (2002) The phenylpropanoid pathway and plant defense – a genomics perspective. Mol Plant Pathol 3:371–390 Dixon RA, Chen F, Guo D, Parvathi K (2001) The biosynthesis of monolignols: a ‘‘metabolic grid’’, or independent pathways to guaiacyl and syringyl units? Phytochemistry 57:1069–1084 Estabrook RW, Hildebrandt AG, Baron J, Netter KJ, Leibman K (1971) A new spectral intermediate associated with cytochrome P-450 function in liver microsomes. Biochem Biophys Res Commun 42:132–139 Ferrer JL, Jez JM, Bowman ME, Dixon RA, Noel JP (1999) Structure of chalcone synthase and the molecular basis of plant polyketide biosynthesis. Nat Struct Biol 6:775–784 Galili G, Sengupta-Gopalan C, Ceriotti A (1998) The endoplasmic reticulum of plant cells and its role in protein maturation and biogenesis of oil bodies. Plant Mol Biol 38:1–29

123

Phytochem Rev (2006) 5:459–472 Graciet E, Lebreton S, Gontero B (2004) Emergence of new regulatory mechanisms in the Benson–Calvin pathway via protein–protein interactions: a glyceraldehyde-3-phosphate dehydrogenase/CP12/phosphoribulokinase complex. J Exp Bot 55:1245–1254 Hrazdina G, Wagner GJ (1985) Metabolic pathways as enzyme complexes: evidence for the synthesis of phenylpropanoids and flavonoids on membrane associated enzyme complexes. Arch Biochem Biophys 237:88–100 Hrazdina G, Zobel AM, Hoch HC (1987) Biochemical, immunological, and immunocytochemical evidence for the association of chalcone synthase with endoplasmic reticulum membranes. Proc Natl Acad Sci USA 84:8966–8970 Hubbard PA, Shen AL, Paschke R, Kasper CB, Kim JJ (2001) NADPH-cytochrome P450 oxidoreductase: structural basis for hydride and electron transfer. J Biol Chem 276:29,163–29,170 Humphreys JM, Chapple C (2002) Rewriting the lignin roadmap. Curr Opin Plant Biol 5:224–229 Ingelman-Sundberg M, Blanck J, Smettan G, Ruckpaul K (1983) Reduction of cytochrome P-450 LM2 by NADPH in reconstituted phospholipid vesicles is dependent on membrane charge. Eur J Biochem 134:157–162 Ingelman-Sundberg M, Haaparanta T, Rydstrom J (1981) Membrane charge as effector of cytochrome P-450LM2 catalyzed reactions in reconstituted liposomes. Biochemistry 20:4100–4106 Janovjak H, Kedrov A, Cisneros DA, Sapra KT, Struckmeier J, Muller DJ (2005) Imaging and detecting molecular interactions of single transmembrane proteins. Neurobiol Aging 88:1423–1428 Jez JM, Bowman ME, Dixon RA, Noel JP (2000) Structure and mechanism of the evolutionarily unique plant enzyme chalcone isomerase. Nat Struct Biol 7:786– 791 Jez JM, Bowman ME, Noel JP (2002) Role of hydrogen bonds in the reaction mechanism of chalcone isomerase. Biochemistry 41:5168–5176 Jorgensen K, Rasmussen AV, Morant M, Nielsen AH, Bjarnholt N, Zagrobelny M, Bak S, Moller BL (2005) Metabolon formation and metabolic channeling in the biosynthesis of plant natural products. Curr Opin Plant Biol 8:280–291 Kao YY, Harding SA, Tsai CJ (2002) Differential expression of two distinct phenylalanine ammonialyase genes in condensed tannin-accumulating and lignifying cells of quaking aspen. Plant Physiol 130:756–760 Koopmann E, Hahlbrock K (1997) Differentially regulated NADPH: cytochrome p450 oxidoreductases in parsely. Proc Natl Acad Sci USA 94:14,954– 14,959 Kristensen C, Morant M, Olsen CE, Ekstrom CT, Galbraith DW, Moller BL, Bak S (2005) Metabolic engineering of dhurrin in transgenic Arabidopsis plants with marginal inadvertent effects on the metabolome and transcriptome. Proc Natl Acad Sci USA 102:1779–1784

Phytochem Rev (2006) 5:459–472 Liu CJ, Blount JW, Steele CL, Dixon RA (2002) Bottlenecks for metabolic engineering of isoflavone glycoconjugates in Arabidopsis. Proc Natl Acad Sci USA 99:14,578–14,583 Liu CJ, Dixon RA (2001) Elicitor-induced association of isoflavone O-methyltransferase with endomembranes prevents the formation and 7-O-methylation of daidzein during isoflavonoid phytoalexin biosynthesis. Plant Cell 13:2643–2658 Magee T, Pirinen N, Adler J, Pagakis SN, Parmryd I (2002) Lipid rafts: cell surface platforms for T cell signaling. Biol Res 35:127–131 Memelink J (2004) Putting the opium in poppy to sleep. Nat Biotechnol 22:1526–1527 Mendes P, Douglas KB, Welch GR (1995). Metabolic channelling in organized enzyme systems: experiments and models. In: Brindle KM (ed) Enzymology in vivo, vol 11. JAI Press Inc., Greenwich, CN, pp 1–19 Miles EW (2001) Tryptophan synthase: a multienzyme complex with an intramolecular tunnel. Chem Rec 1:140–151 Moller BL, Conn EE (1980) The biosynthesis of cyanogenic glucosides in higher plants. Channeling of intermediates in dhurrin biosynthesis by a microsomal system from Sorghum bicolor (linn) Moench. J Biol Chem 255:3049–3056 Morant M, Bak S, Moller BL, Werck-Reichhart D (2003) Plant cytochromes P450: tools for pharmacology, plant protection and phytoremediation. Curr Opin Biotechnol 14:151–162 Munro AW, Daff S, Coggins JR, Lindsay JG, Chapman SK (1996) Probing electron transfer in flavocytochrome P-450 BM3 and its component domains. Eur J Biochem 239:403–409 Munro AW, Leys DG, McLean KJ, Marshall KR, Ost TW, Daff S, Miles CS, Chapman SK, Lysek DA, Moser CC, Page CC, Dutton PL (2002) P450 BM3: the very model of a modern flavocytochrome. Trends Biochem Sci 27:250–257 Murataliev MB, Feyereisen R (2000) Functional interactions in cytochrome P450BM3. Evidence that NADP(H) binding controls redox potentials of the flavin cofactors. Biochemistry 39:12,699–12,707 Murataliev MB, Feyereisen R, Walker FA (2004) Electron transfer by diflavin reductases. Biochim Biophys Acta 1698:1–26 Nadler SG, Strobel HW (1991) Identification and characterization of an NADPH-cytochrome P450 reductase derived peptide involved in binding to cytochrome P450. Arch Biochem Biophys 290:277–284 Nielsen KA, Moller BL (2005) Cytochrome P450 in plants. In: Ortiz de Montellano PR (ed) Cytochrome P450: structure, mechanism, and biochemistry. Kluwer Academic/Plenum Publishers, New York, pp 553–583 Paine MJ, Scrutton NS, Munro AW, Gutierrez A, Roberts GC, Wolf CR (2005) Electron transfer partners of cytochrome P450. In: Ortiz de Montellano PR (ed) Cytochrome P450: structure, mechanism, and biochemistry. Kluwer Academic/Plenum Publishers, New York, pp 115–148

471 Panicot M, Minguet EG, Ferrando A, Alcazar R, Blazquez MA, Carbonell J, Altabella T, Koncz C, Tiburcio AF (2002) A polyamine metabolon involving aminopropyl transferase complexes in Arabidopsis. Plant Cell 14:2539–2551 Peterson JA, Ebel RE, O’Keeffe DH, Matsubara T, Estabrook RW (1976) Temperature dependence of cytochrome P-450 reduction. A model for NADPHcytochrome P-450 reductase: cytochrome P-450 interaction. J Biol Chem 251:4010–4016 Ralston L, Subramanian S, Matsuno M, Yu O (2005) Partial reconstruction of flavonoid and isoflavonoid biosynthesis in yeast using soybean Type I and Type II chalcone isomerases. Plant Physiol 137: 1375–1388 Rasmussen S, Dixon RA (1999) Transgene-mediated and elicitor-induced perturbation of metabolic channeling at the entry point into the phenylpropanoid pathway. Plant Cell 11:1537–1552 Reumann S (2000) The structural properties of plant peroxisomes and their metabolic significance. Biol Chem 381:639–648 Ro DK, Douglas CJ (2004) Reconstitution of the entry point of plant phenylpropanoid metabolism in yeast (Saccharomyces cerevisiae): implications for control of metabolic flux into the phenylpropanoid pathway. J Biol Chem 279:2600–2607 Russell DW, Conn EE (1967) The cinnamic acid 4hydroxylase of pea seedlings. Arch Biochem Biophys 122:256–258 Saslowsky D, Winkel-Shirley B (2001) Localization of flavonoid enzymes in Arabidopsis roots. Plant J 27:37– 48 Saslowsky DE, Warek U, Winkel BS (2005) Nuclear localization of flavonoid enzymes in Arabidopsis. J Biol Chem 280:23,735–23,740 Schoch GA, Attias R, Le Ret M, Werck-Reichhart D (2003) Key substrate recognition residues in the active site of a plant cytochrome P450, CYP73A1. Homology guided site-directed mutagenesis. Eur J Biochem 270:3684–3695 Sevrioukova IF, Li H, Zhang H, Peterson JA, Poulos TL (1999) Structure of a cytochrome P450-redox partner electron-transfer complex. Proc Natl Acad Sci USA 96:1863–1868 Shen S, Strobel HW (1993) Role of lysine and arginine residues of cytochrome P450 in the interaction between cytochrome P4502B1 and NADPH-cytochrome P450 reductase. Arch Biochem Biophys 304:257–265 Shen S, Strobel HW (1995) Functional assessment of specific amino acid residues of cytochrome P4501A1 using anti-peptide antibodies. Arch Biochem Biophys 320:162–169 Shimada N, Aoki T, Sato S, Nakamura Y, Tabata S, Ayabe S (2003) A cluster of genes encodes the two types of chalcone isomerase involved in the biosynthesis of general flavonoids and legume-specific 5deoxy(iso)flavonoids in Lotus japonicus. Plant Physiol 131:941–951

123

472 Shimizu T, Tateishi T, Hatano M, Fujii-Kuriyama Y (1991) Probing the role of lysines and arginines in the catalytic function of cytochrome P450d by site-directed mutagenesis. Interaction with NADPH-cytochrome P450 reductase. J Biol Chem 266:3372–3375 Srere PA (1985) The metabolon. Trends Biochem Sci 10:109–110 Srere PA (1987) Complexes of sequential metabolic enzymes. Annu Rev Biochem 56:89–124 Srere PA (2000) Macromolecular interactions: tracing the roots. Trends Biochem Sci 25:150–153 Subramaniam R, Desveaux D, Spickler C, Michnick SW, Brisson N (2001) Direct visualization of protein interactions in plant cells. Nat Biotechnol 19:769–772 Thaminy S, Miller J, Stagljar I (2004) The split-ubiquitin membrane-based yeast two-hybrid system. Methods Mol Biol 261:297–312 Urban P, Mignotte C, Kazmaier M, Delorme F, Pompon D (1997) Cloning, yeast expression, and characterization of the coupling of two distantly related Arabidopsis thaliana NADPH-cytochrome P450 reductases with P450 CYP73A5. J Biol Chem 272:19,176–19,186 Venkateswarlu K, Lamb DC, Kelly DE, Manning NJ, Kelly SL (1998) The N-terminal membrane domain of yeast NADPH-cytochrome P450 (CYP) oxidoreductase is not required for catalytic activity in sterol biosynthesis or in reconstitution of CYP activity. J Biol Chem 273:4492–4496 Wallrabe H, Periasamy A (2005) Imaging protein molecules using FRET and FLIM microscopy. Curr Opin Biotechnol 16:19–27 Wang M, Roberts DL, Paschke R, Shea TM, Masters BS, Kim JJ (1997) Three-dimensional structure of NADPH-cytochrome P450 reductase: prototype for FMN- and FAD-containing enzymes. Proc Natl Acad Sci USA 94:8411–8416

123

Phytochem Rev (2006) 5:459–472 Wanner LA, Li G, Ware D, Somssich IE, Davis KR (1995) The phenylalanine ammonia-lyase gene family in Arabidopsis thaliana. Plant Mol Biol 27:327–338 Weisshaar B, Jenkins GI (1998) Phenylpropanoid biosynthesis and its regulation. Curr Opin Plant Biol 1:251– 257 Werck-Reichhart D, Feyereisen R (2000) Cytochromes P450: a success story. Genome Biol 1:3003.1–3003.9 Winkel-Shirley B (2001a) Flavonoid biosynthesis. A colorful model for genetics, biochemistry, cell biology, and biotechnology. Plant Physiol 126:485–493 Winkel-Shirley B (2001b) It takes a garden. How work on diverse plant species has contributed to an understanding of flavonoid metabolism. Plant Physiol 127:1399–1404 Winkel BSJ (2004) Metabolic channeling in plants. Ann Rev Plant Physiol Mol Biol 55:85–107 Yu O, Jung W, Shi J, Croes RA, Fader GM, McGonigle B, Odell JT (2000) Production of the isoflavones genistein and daidzein in non-legume dicot and monocot tissues. Plant Physiol 124:781–794 Yu O, McGonigle B (2005) Metabolic engineering of isoflavone biosynthesis. Adv Agron 86:147–190 Yu O, Shi J, Hession AO, Maxwell CA, McGonigle B, Odell JT (2003) Metabolic engineering to increase isoflavone biosynthesis in soybean seed. Phytochemistry 63:753–763 Zhao Q, Modi S, Smith G, Paine M, McDonagh PD, Wolf CR, Tew D, Lian LY, Roberts GC, Driessen HP (1999) Crystal structure of the FMN-binding domain of human cytochrome P450 reductase at 1.93 A resolution. Protein Sci 8:298–306