Phytochemistry 72 (2011) 2113–2123

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Phytochemistry

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Homology modeling of the three membrane proteins of the dhurrin metabolon:Catalytic sites, membrane surface association and protein–protein interactions

Kenneth Jensen a,b,c, Sarah Anne Osmani a,b,1, Thomas Hamann a,b,2, Peter Naur a,c,Birger Lindberg Møller a,b,c,⇑a Plant Biochemistry Laboratory, Department of Plant Biology and Biotechnology, University of Copenhagen, 40 Thorvaldsensvej, DK-1871 Frederiksberg C, Copenhagen, Denmarkb VKR Research Centre Pro-Active Plants, 40 Thorvaldsensvej, DK-1871 Frederiksberg C, Copenhagen, Denmarkc UNIK Center for Synthetic Biology, University of Copenhagen, 40 Thorvaldsensvej, DK-1871 Frederiksberg C, Copenhagen, Denmark

a r t i c l e i n f o

Article history:Received 25 February 2011Received in revised form 29 April 2011Available online 26 May 2011

Keywords:MetabolonDhurrinHomology modelingCytochrome P450NADPH-dependent cytochrome P450reductase

0031-9422/$ - see front matter � 2011 Elsevier Ltd. Adoi:10.1016/j.phytochem.2011.05.001

⇑ Corresponding author at: Plant Biochemistry LabBiology and Biotechnology, University of Copenhagen,Frederiksberg C, Copenhagen, Denmark. Tel.: +45 353

E-mail address: blm@life.ku.dk (B.L. Møller).1 Present address: ALK-Abello, 6–8 Bøge Allé, DK-292 Present address: Novo Nordisk A/S, Novo Nordisk Pa

a b s t r a c t

Formation of metabolons (macromolecular enzyme complexes) facilitates the channelling of substratesin biosynthetic pathways. Metabolon formation is a dynamic process in which transient structures med-iated by weak protein–protein interactions are formed. In Sorghum, the cyanogenic glucoside dhurrin isderived from L-tyrosine in a pathway involving the two cytochromes P450 (CYPs) CYP79A1 and CYP71E1,a glucosyltransferase (UGT85B1), and the redox partner NADPH-dependent cytochrome P450 reductase(CPR). Experimental evidence suggests that the enzymes of this pathway form a metabolon. Homologymodeling of the three membrane bound proteins was carried out using the Sybyl software and availablerelevant crystal structures. Residues involved in tight positioning of the substrates and intermediates inthe active sites of CYP79A1 and CYP71E1 were identified. In both CYPs, hydrophobic surface domainsclose to the N-terminal trans-membrane anchor and between the F0 and G helices were identified asinvolved in membrane anchoring. The proximal surface of both CYPs showed positively charged patchescomplementary to a negatively charged bulge on CPR carrying the FMN domain. A patch of surfaceexposed, positively charged amino acid residues positioned on the opposite face of the membrane anchorwas identified in CYP71E1 and might be involved in binding UGT85B1 via a hypervariable negativelycharged loop in this protein.

� 2011 Elsevier Ltd. All rights reserved.

1. Introduction

Co-localization of enzymes in macromolecular complexes cata-lyzing consecutive steps in a biosynthetic pathway is a central fea-ture of cell metabolism. Such complexes, also termed metabolons,are thought to improve catalytic efficiency and avoid metabolicinterference by channelling of intermediates between the activesites of co-localized enzymes (Jørgensen et al., 2005; Kurakin,2009; Srere, 1987; Sweetlove and Fernie, 2005; Thellier et al.,2006).

The molecular mechanisms governing metabolon formation re-main elusive, and direct evidence for the existence of metabolonscan be difficult to obtain experimentally when the associations be-tween the enzymes are dynamic, transient, with low affinity andfunction dependent (Jørgensen et al., 2005; Møller, 2010; Thellier

ll rights reserved.

oratory, Department of Plant40 Thorvaldsensvej, DK-187133352; fax: +45 35333333.

70 Hørsholm, Denmark.rk, DK-2760 Måløv, Denmark.

et al., 2006). The enzymes catalyzing the synthesis of the bio-activenatural product dhurrin in Sorghum bicolor are thought to be orga-nized as a metabolon (Kristensen et al., 2005; Møller and Conn,1980; Nielsen et al., 2008). In vitro experiments involving simulta-neous administration of dual radiolabeled intermediates to iso-lated microsomes accompanied by in vivo experiments usingtransgenic plants expressing fluorescent-labelled fusion proteinshave demonstrated that the biosynthesis of the L-tyrosine derivedcyanogenic glucoside dhurrin from S. bicolor is highly channelledand effectively prevents metabolic cross-talk (Kristensen et al.,2005; Møller and Conn, 1980; Nielsen et al., 2008; Tattersallet al., 2001). The dhurrin pathway involves two multifunctionalmembrane-anchored cytochromes P450 (CYPs), CYP79A1 andCYP71E1, a membrane bound NADPH–cytochrome P450 reductase(CPR) and a soluble UDP-glucosyltransferase (UGT85B1). In thefirst part of the pathway, CYP79A1 converts L-tyrosine to (Z)-p-hydroxyphenylacetaldoxime through a number of steps involvingtwo hydroxylation steps and unusual and labile intermediates(Koch et al., 1995; Sibbesen et al., 1995). The first step in the path-way is highly substrate specific, with L-tyrosine being the onlyknown substrate for the enzyme (Kahn et al., 1999; Møller andConn, 1980). [18O]-labelling experiments have demonstrated that

Table 1The quality of the structural models of CYP79A1, CYP71E1 and CPR1 assessed by theiroverall g-factors and ProQ scores.

CYP79A1 CYP71E1 SbCPR2b

Overall g-factor �0.05 �0,26 �0.15LGscore 3.186 1.889 4.420MaxSub 0.259 0.112 0.339

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the intermediates are held tightly in place in the active site ofCYP79A1 thereby preventing free rotation around the C–N singlebond of the N-hydroxytyrosine and N,N-dihydroxytyrosine inter-mediates (Halkier et al., 1991). In the second part of the pathway,CYP71E1 converts the oxime in a two step reaction into the cyano-hydrin p-hydroxymandelonitrile (Bak et al., 1998). This conversioninvolves a dehydration step followed by a classical monooxygen-ation reaction (Møller and Conn, 1980). Besides the tyrosine de-rived oxime, CYP71E1 has been shown to catalyze the conversionof a phenylalanine derived oxime, although at much lower rates.The CYP71E1 substrate specificity is thus less stringent comparedto CYP79A1 (Kahn et al., 1999). The cyanohydrin p-hydroxymand-elonitrile produced by CYP71E1 is labile at pH values above 6.0(Selmar et al., 1987). In vitro, the cyanohydrin dissociates into p-hydroxybenzaldehyde and toxic hydrogen cyanide. In vivo, dissoci-ation is avoided through rapid glucosylation by the soluble UDP-glucosyltransferase UGT85B1 (Hansen et al., 2003; Jones et al.,1999; Thorsøe et al., 2005). The organization of the biosyntheticpathway within a metabolon would serve to speed up conversionof the labile and highly reactive intermediates into the finalproduct.

Both CYPs involved in the catalysis of dhurrin synthesis areendoplasmic reticulum localized and classified as class II CYPsand require the transfer of electrons mediated by a membrane-an-chored CPR (Jensen and Møller, 2010; Werck-Reichhart and Feyer-eisen, 2000). Interactions between CYP and CPR have beenextensively studied using the flavocytochrome P450 BM3, a nativesoluble bacterial CYP-CPR fusion protein (Munro et al., 2007, 2002).Electron-transfer complex formation involving membrane-an-chored eukaryotic CYPs has been studied by chemical modifica-tions of key amino acid residues and by targeted mutagenesis(Hlavica et al., 2003). These studies have identified several surfaceexposed charged residues on the CYP and CPR proteins implied tobe important for efficient interaction. However, the CYP–CPR inter-action is expected to be somewhat transient. In the course of elec-tron transfer from NADPH through the FAD and FMN cofactors ofCPR to the heme of the CYP, CPR must undergo large conforma-tional changes. This serves to expose the FMN-cofactor otherwiseburrowed within the protein structure and thereby facilitatesCYP interaction. CPR has been proposed to be in equilibrium be-tween a compact and an open structure (Ellis et al., 2009). Theequilibrium is controlled by the redox state, coenzyme binding(Grunau et al., 2006; Gutierrez et al., 2003, 2002) and possibly byphosphorylation (Laursen et al., 2011). The structure of the com-pact conformation favors electron transfer from NADPH via theFAD cofactor to the FMN cofactor, whereas the open conformationleaves the FMN coenzyme solvent exposed and accessible to theheme of the CYP (Aigrain et al., 2009; Ellis et al., 2009; Hamdaneet al., 2009). Since CPR cycles between two distinct conformationsin the course of electron transfer, the interaction between CPR andCYPs is envisioned to be transient. Several crystal structures havebeen solved for mammalian microsomal class II CYP proteins(Bridges et al., 1998; Rowland et al., 2006; Sansen et al., 2007; Scottet al., 2003, 2004; Smith et al., 2007). In the field of plant CYPs, onlytwo crystal structures have been solved. Both of these were ob-tained with allene oxide synthases, peroxide-metabolising CYPsof the CYP74A subfamily (Lee et al., 2008; Li et al., 2008). Alleneoxide synthases differ from typical CYPs by not utilizing molecularoxygen as substrate and being independent of CPR. Based on mam-malian- and bacterial-derived CYP crystal structures, amino acidresidues forming the active site of a number of class II plant CYPshave been predicted by homology modeling (Lee et al., 2010;Rupasinghe et al., 2003; Rupasinghe and Schuler, 2006; Sawadaet al., 2002; Schoch et al., 2003; Thornton et al., 2010). None ofthese reports have enabled the characterization of the interactionsbetween plant CYPs and CPRs, the possible entry-exit routes of the

substrates, or possible interaction mechanisms in multi-enzymecomplexes.

Using homology modeling, we have constructed 3D structuralmodels of the three membrane bound proteins thought to interactas constituents of a dhurrin metabolon. Based on these models,predictions were made regarding amino acid residues involved inactive site formation and substrate docking, possible entry sitefor substrates and residues guiding proper interaction with the re-dox partner. The models demonstrate that the metabolon is verydynamic, requiring major structural conformation changes toachieve proper CYP–CPR interaction. It remains unclear whetherthe same CPR manages to interact with CYP79A1 as well asCYP71E1 or whether this interaction involves different CPR mole-cules. Likewise, the formation of the metabolon may be functiondependent with disassembly of the interaction between the CYPsresulting in formation of p-hydroxyphenylacetaldoxime as the fi-nal product (Møller, 2010).

2. Results

2.1. Evaluating the quality of structural models of CYP79A1, CYP71E1and SbCPR2b

Homology models of the S. bicolor proteins CYP79A1, CYP71E1and CPR2b were constructed in the Sybyl software suite using mul-tiple crystal structures of mammalian CYPs and CPR as templates(see Section 5 for details). The quality of the final models was as-sessed by calculation of their overall Procheck g-scores to evaluatethe distribution of phi and psi angles (Laskowski et al., 1993) andby calculating the LGscore and MaxSub using ProQ (Wallner andElofsson, 2003) to evaluate the quality of the protein models. Allcalculated values indicated a high likelihood of correct foldingand thus good quality of the models (Table 1).

2.2. Structural analysis of the structural models of CYP79A1 andCYP71E1

2.2.1. Secondary and tertiary structure is highly conserved in themodels

The final models of CYP79A1 and CYP71E1 contain the 12 majora-helices that form the conserved 3D fold of CYP proteins (Fig. 1).The conserved a-helices correspond to the helices that have previ-ously been assigned as helices A–L (Hasemann et al., 1995). Each ofthe helices F, K and J are divided into two helices F and F’, J and J0

and K and K0 in both models (Fig. 2). The structural model ofCYP71E1 contains an additional a-helix at the C-terminal end. Ofthe conserved b-strands in CYP proteins, six were present in themodels of CYP79A1 and CYP71E1. Together, the b-strands give riseto the two highly conserved b-sheets of the b-domain. One b-sheetis composed of b-strands 1-1, 1-2 (N terminal), 1-3 and 1-4 (fourthand first b strand of the four conserved K–K0 loop b-strands) andthe second of the two b-strands 2-1 and 2-2 (strands 2 and 3 ofthe four conserved K–K0 loop b-strands) (Hasemann et al., 1995).

CYP proteins are known to contain several conserved residuesinvolved in anchoring the heme group by H-bond formation tothe propionate side chains of the A and D rings of the porphyrin

Fig. 1. Overview of the modeled cytochrome P450 structures. The overall structure of CYP79A1 and CYP71E1 was colored blue to red from N-terminus to C-terminus. Theletters indicate the name of the helices. The positions of the N-terminal residue in the truncated structural models are shown by an asterisk. (For interpretation of thereferences to colur in this figure legend, the reader is referred to the web version of this article.)

Fig. 2. Sequence alignment of CYP79A1 and CYP71E1. The alignment was generated using the program TM-align (Zhang and Skolnick, 2005). Red tubes denote a-helices andgreen arrows b-sheets. Residues shown on a blue background are involved in heme anchoring and residues on a yellow background are within five angstroms of the dockedsubstrate (see Fig. 3). (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

K. Jensen et al. / Phytochemistry 72 (2011) 2113–2123 2115

system as well as to provide a fifth ligand for the heme iron(Rowland et al., 2006). These residues are conserved in thestructural models of CYP79A1 and CYP71E1 (Fig. 2).

2.2.2. Substrate docking and interactions in the model of CYP79A1CYP79A1 converts the amino acid L-tyrosine into p-hydrox-

yphenylacetaldoxime via the two intermediates N-hydroxytyro-sine and N,N-dihydroxytyrosine (Sibbesen et al., 1995). Thesubstrate of CYP79A1, L-tyrosine (Fig. 3A) and the first intermedi-ate N-hydroxytyrosine, were docked into the substrate bindingpocket above the heme plane. Docking of substrates and

intermediates was performed assuming a similar positioning ofthese during all three reaction steps. N,N-dihydroxytyrosine isunstable and its conversion into the final product of CYP79A1, p-hydroxyphenylacetaloxime is expected to be highly channelledwhile the likely transition state intermediates involved are keptin the same position as during the preceding steps. The substrateswere manually fitted into the substrate binding pocket as deter-mined by the physical constraints defined by the shape of thepocket and so that the L-tyrosine N-atom to be hydroxylated couldbe positioned at a favorable distance to the heme iron. The finaldocking resulted in a distance of 4.3 Ångstroms (Å) from the center

Fig. 3. Active site of CYP79A1 and CYP71E1 from Sorghum bicolor. Panels A and B show features of the substrate binding pocket of CYP79A1 and CYP71E1, respectively. Allamino acid residues within 5 Å of the docked substrate are shown in stick representation and dotted lines represent proposed hydrogen bonding between substrate andenzyme residues. Panels C and D show the substrate recognition sites 1–6 as defined by structural comparison with CYP101 (Gotoh, 1992). The substrates, L-tyrosine and p-hydroxyphenylacetaldoxime, are shown in magenta and the heme group in yellow. (For interpretation of the references to color in this figure legend, the reader is referred tothe web version of this article.)

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of the N-atom of the L-tyrosine or N-hydroxytyrosine amino groupto the center of the heme. Subsequently, positions of amino acidresidues proposing favorable H-bonding or hydrophobic interac-tions with L-tyrosine or N-hydroxytyrosine were considered. Thesubstrates of CYP79A1 were positioned so that the hydrophilic res-idues Ser144 (from the B–C loop) and Thr538 (from the C-terminal)positioned at the distal end of the pocket would engage in eitherdirect or water-mediated hydrogen bonding with the para-hydro-xy group of L-tyrosine or N-hydroxytyrosine (Fig. 3A). Dockingbased on these parameters positioned the carboxyl group and theNH2 group of the L-tyrosine and N-hydroxytyrosine substrates inclose association to residues Arg152 and Asn355 positioned inthe heme proximal end of the substrate binding pocket. This wouldallow for stabilizing interactions between the carboxy-group of thesubstrates and the positively charged side chain of Arg152, whileAsn355 can interact with the NH2 group of L-tyrosine or the N-OH function of N-hydroxytyrosine and N,N-dihydroxytyrosine byhydrogen bonding.

2.2.3. Substrate docking and interactions in the model of CYP71E1CYP71E1 converts p-hydroxyphenylacetaloxime into p-

hydroxymandelonitrile in two steps. The substrate p-hydrox-yphenylacetaloxime was docked into the substrate binding pocketso that the atoms involved in the initial dehydration reaction werepositioned above the heme and the p-hydroxy group of the aro-matic ring pointing towards the distal end of the pocket, whichwas displaced towards the propionate side chains of the heme(Fig. 3B). This positioned the p-hydroxyphenylacetaloxime sub-strate in an orientation similar to the position of L-tyrosine andN-hydroxytyrosine in the structural model of CYP79A1. Becauseof the smaller size of the substrate binding pocket in CYP71E1,caused mainly by the bulkiness of the two phenylalanine residuesPhe240 and Phe512 pointing into the cavity, the N-atom of theoxime function was positioned slightly closer to the heme iron incomparison to the positioning of the N-atom of L-tyrosine and

N-hydroxytyrosine in CYP79A1, resulting in a final distance of3.4 Å. The p-hydroxyphenylacetaloxime and p-hydroxyphenyl-acetonitrile substrates are tightly fitted into the substrate pocketwith the aromatic ring being surrounded by residues creating ahydrophobic environment. Ser125 from the B’-loop and Thr511from the C-terminal are positioned favorably for interaction withthe para-hydroxy group of the substrates in the distal end of thepocket, while Thr332 in the heme proximal end of the substratepocket is positioned to interact with the leaving group of p-hydrox-yphenylacetaloxime (Fig. 3B).

2.3. Membrane anchoring of CYP models

Eukaryotic CYP proteins are membrane bound (Werck-Reichhart and Feyereisen, 2000). Membrane anchoring is conferredby the presence of an N-terminal transmembrane spanning region.In the models of CYP79A1 and CYP71E1, the N-terminal aminoacids containing this membrane anchor were left out, because noappropriate template for modeling the membrane spanning seg-ments was available. Removal of the N-terminal anchor by proteo-lytic cleavage does not necessarily result in fully soluble CYPs(Cosme and Johnson, 2000). This demonstrates that ionic or hydro-phobic interactions with the membrane surface can be strongenough to conserve membrane association.

Analysis of the surface of the structural models of CYP79A1 andCYP71E1 identified surface exposed hydrophobic domains in the re-gion positioned near the N-terminal of CYP79A1 and CYP71E1 andfacing the membrane next to the attachment site of the N-terminalanchor. In CYP79A1, a hydrophobic tip is formed by hydrophobic res-idues at positions 72–78, 81–82 and 84–86 and in CYP71E1 byhydrophobic residues at positions 55–60, 62–65, 68 and 69–70.

Apart from residues near the N-terminal membrane anchor, theregion between helices F and G has also been implicated in mem-brane association. In the crystal structure of human CYP1A2, thisregion consists of two amphipatic helix segments which are

K. Jensen et al. / Phytochemistry 72 (2011) 2113–2123 2117

thought to be partly embedded in the membrane (Sansen et al.,2007). These segments are modeled as loops in CYP79A1 andCYP71E1, and their hydrophobicity is most pronounced inCYP79A1 (Fig. 1A and B). The loops are localized between a-helicesF’ and G and in both enzymes contain hydrophobic and nonpolarresidues, for CYP79A1 Leu 272, Pro273, Trp274 and Leu275 andfor CYP71E1 Ala249, Ala250, Leu253, Ala254.

2.4. Proposed substrate access to the active site in the structuralmodels of CYP79A1 and CYP71E1

CYP79A1 and CYP71E1 were modeled using crystal structurecoordinates of CYP proteins crystallized in their closed conforma-tions. In these structures, the substrates are observed to be entirelyburied inside the CYP protein with no apparent entry or exit chan-nel. Accordingly, in the structural models of CYP79A1 andCYP71E1, the substrates are buried inside the protein. Substrate ac-cess is most likely to occur when the enzyme is in an open confor-mation. Opening of the enzyme is envisioned to be analogous towhat happens in CYP2B4 (Scott et al., 2003) where displacementof helices F, F0 and G leads to formation of a cleft near the mem-brane surface on the side facing away from the CPR interactionsurface.

2.5. Modeling and membrane anchoring/positioning of SbCPR2b

Activity of CYPs is dependent on electron transfer from a mem-brane bound CPR. Homology modeling of SbCPR2b was based on

Fig. 4. Structural ribbon model of N-terminally truncated CPR from Sorghum bicolor. (A) Scontact to the membrane is facilitated by hydrophobic patches in the FAD/NADPH domembrane (dashed line) is 50 Å. (B) Vertical orientation of CPR onto the membrane. TheSbCPR2b over the membrane is 57 Å. Panels C and D: CPR undergoes conformational chanhave been proposed, a swinging model (Panel C) in which the FAD/NADPH domain (gre(Panel D). The conformational changes are illustrated with a red arrow. The linker domaicolored in yellow, orange and blue, respectively. The position of the N-terminal of the truthe lower part of the figure represents the membrane plane. (For interpretation of the refarticle.)

the solved crystal structures of rat, yeast and human CPR as de-scribed in Section 5. The spatial orientation of CPR on the mem-brane is not yet clear and two different orientations have beenproposed; a horizontal and a vertical orientation. Wang et al.(1997) suggested a horizontal orientation in which both the FMNand FAD/NADPH binding domains are partly embedded in themembrane surface, which would facilitate electron transfer fromthe FMN binding domain to the CYP heme. Horizontal membranedocking of SbCPR2b was accomplished by superimposing themembrane interacting regions previously proposed for the rat liverCPR onto the model structure of SbCPR2b (Fig. 4A) (Wang et al.,1997). This identified a limited number of hydrophobic aminoacids in SbCPR2b (274–305, 546–565 and 583–587) which arepositioned on the surface of the model but did not give rise tohydrophobic patches suitable for membrane interaction, conse-quently showing no clear linkage between the hydrophobic aminoacids and a horizontal positioning of SbCPR2b on the membrane.An analysis of the surface hydrophobicity of the horizontal posi-tioned SbCPR2b model did not identify other hydrophobic patchesto support a horizontal orientation of CPR on the membrane. In thehorizontal orientation, CPR has a vertical height of 5.0 nm. In con-trast, single molecule height measurements on CPR in nanodiscsshowed that the molecule had a vertical height of 5.6 nm, suggest-ing a vertical orientation of CPR, in which the orientation on themembrane is controlled solely by the N-terminal membrane an-chor (Ellis et al., 2009) and illustrated in Fig. 4B. The spatial androtational freedom of the catalytic domains in the vertical orientedmodel of SbCPR2b would ease the rearrangements of the catalytic

bCPR2b positioned in a horizontal orientation on the membrane. In this orientation,main and by the N-terminal membrane anchor. The height of SbCPR2b over themembrane association is solely controlled by the membrane anchor. The height ofges during electron transfer to CYPs. Two different types of conformational changesen) is displaced and a rotating model, in which the FMN-domain is rotated (blue)n is colored in red and the hinge in yellow. The FMN, FAD and NADPH cofactors arencated protein is shown by an asterisk. All modeled structures are positioned so thaterences to color in this figure legend, the reader is referred to the web version of this

2118 K. Jensen et al. / Phytochemistry 72 (2011) 2113–2123

domains compared to a horizontal positioning of SbCPR2b. Giventhe lack of constraints on rearrangements of the catalytic domainsand the importance of these rearrangements for completing thecatalytic cycle, the vertical orientation would describe the mostfavorable model.

2.6. CYP79A1/71E1-SbCPR2b interactions and membrane positioning

Electron transfer between CPR and CYP proteins proceeds byinteraction of the FMN domain of CPR with the proximal surfaceof the CYP. Interaction of CYPs with redox partners has been shownto involve ionic interactions between positively charged residuesof the CYP protein with negatively charged residues on the redoxpartner proteins (Bridges et al., 1998; Hasemann et al., 1995). Stud-ies of CPR show that negatively charged residues surrounding theFMN cofactor are involved in the interaction (Shen and Strobel,1993; Wang et al., 1997; Zhao et al., 1999). The negatively chargedresidues are divided into three clusters and in the primary struc-ture of SbCPR2b, cluster 1 corresponds to the amino acid residuesAsp231-Asp232-Asp233, cluster 2 to Glu237-Asp238-Asp239 andcluster 3 to Asp163-Glu165-Asp168. In the structural model ofSbCPR2b as well as in the published CPR crystal structures(Hubbard et al., 2001; Lamb et al., 2006; Wang et al., 1997; Zhaoet al., 1999), all residues proposed to engage in CYP interactionsare positioned on the side of the SbCPR2b harboring the FMN bind-ing domain. In the structures, this side is directly facing the FADbinding domain of CPR. As expected, analysis of the surface ofthe structural model of SbCPR2b showed that not all clusters weresurface exposed. Only cluster 2 and residue Asp168 from cluster 3were freely available for CYP interaction without rearrangement ofthe SbCPR2b structure. Accordingly, CYP binding requires a confor-mational change of the SbCPR2b protein to expose the FMN bind-ing domain for CYP interaction. Two different models to exposethe FMN domain have been suggested; a swinging model and arotation model (Ellis et al., 2009; Laursen et al., 2011). The swing-ing model is based on a study utilizing a combination of NMR andSAXS that documented that such a major conformational change ofCPR does indeed take place and that CPR is thought to exist in equi-librium between a compact and an open conformation (Ellis et al.,2009). The CPR hinge documented to mediate this flexibility corre-sponds to the amino acid residues 254–268 in the SbCPR2b proteinstudied here. Accordingly, during the conformation change fromthe closed to the open form, the FMN domain is envisioned to re-tain its position on the membrane surface whereas the FAD/

Fig. 5. Possible interactions between CYP79A1/CYP71E1 and CPR. The cytochromes P450electrostatic potential (blue: positively charged surfaces; red: negatively charged surfaacidic residues on CPR. CPR is displayed as a cyan cartoon representation with the acidirepresentation is used for the FMN cofactor. The distance shown is the shortest distancereferences to color in this figure legend, the reader is referred to the web version of thi

NADPH domain moves away from the membrane (Fig. 4C). In con-trast, the rotating model describes a horizontal rotation of the FMNdomain around the flexible hinge resulting in exposure of the FMNbinding domain to the solvent (Fig. 4D) (Hamdane et al., 2009).Surface analysis of the exposed FMN domain of SbCPR2b revealedthat all residues in the three clusters 1, 2 and 3, will be surface ex-posed in the open conformation of CPR and properly positioned tointeract with positive residues at the proximal side of the CYP. Inaddition, Glu203 was also found to be in a favorable position forCYP interaction.

The CPR interacting surface of CYP proteins is proposed to be onthe proximal side of the protein where the heme is closest to thesurface (Williams et al., 2000). Analysis of the proximal surfaceof the two CYP models shows patches of positively charged resi-dues positioned to form a ring around the heme. Four patches ofpositively charged residues on each of the CYP79A1 and CYP71E1models were identified. Patch 1 consists of residues of the C-helix(K164, K165 and R168 in CYP79A1 and R148, R151 and R152 inCYP71E1). Patch 2 of residues of the B-C loop (R125, R129, R130,CYP79A1 and R109, K113, H115, CYP71E1) and a highly conservedarginine from the cys-pocket (CYP79A1 R490 and CYP71E1 R465).Patch 3 consists of residues of the K-helix and a residue just beforethe Cys-pocket (79A1 residues K398, K403, K407 and R482 andCYP71E1 residues K376, K381, K385 and H455). Patch 4 consistsof residues of the C-D loop (patch 3) (CYP79A1 Arg179 and His180 and CYP71E1 Arg163 and Arg165) (Supplementary Fig. 1Aand B).

Due to the lack of templates for modeling SbCPR2b in an openconformation, docking of CYP79A1/71E1 and SbCPR2b was doneusing only the FMN domain of SbCPR2b. The manual docking wasperformed to maximize the potential interactions between theidentified charged patches on the protein surfaces as well as tominimize the distance from the FMN cofactor of SbCPR2b to thecenter of the heme (Fig. 5). Both CYPs and SbCPR2b are membranebound, and while the overall membrane position of both proteinswas kept, some flexibility in the position of the CYP proteins wasallowed in accordance with the above proposed CYP membranepositioning.

The proposed interacting surfaces of either of the CYP proteinmodels and the SbCPR2b protein model were found to have com-plementary surfaces, with a bulge on the exposed SbCPR2b surfaceFMN domain fitting into a positively charged surface cavity onCYP79A1/71E1 (Supplementary Fig. 1). This would position theheme center�12 Å from the FMN cofactor (Fig. 5A and B). Studying

(Panel A: CYP79A1, Panel B: CYP71E1) are illustrated as surface models showing theces), clearly demonstrating the positively charged patch proposed to interact withc residues important for CYP interaction colored orange and labeled. A yellow stick

between the dimethyl group of FMN and the heme iron. (For interpretation of thes article.)

K. Jensen et al. / Phytochemistry 72 (2011) 2113–2123 2119

the distribution of positively and negatively charged patches of theCYPs and CPR respectively shows that the SbCPR2b cluster 1 and 2residues are positioned close to CYP patch 1 and 4. SbCPR2b cluster3 is facing CYP patch 3 and the SbCPR2b residue Glu203 and cluster3 residue Asp163 are facing CYP patch 2 (Fig. 5A and B). In theresulting structural model, the angle of the CYP heme plane in rela-tion to the membrane is �50�. This compares well with the valuediscussed by Bayburt and Sligar (2002).

3. Discussion

3.1. Substrate interactions

The specificity of the CYP79A1 and CYP71E1 proteins in the S.bicolor dhurrin biosynthetic pathway has been extensively studiedbiochemically (Kahn et al., 1999). Here we used homology model-ing to propose specific amino acid residues responsible for sub-strate binding and for defining the substrate specificity of thetwo proteins. The homology modeling showed that the substratepockets of both proteins were small and severely restricted thenumber of possible overall orientations and positions of the twoin vivo substrates L-tyrosine and p-hydroxyphenylacetaldoximewithin the active site of CYP79A1 and CYP71E1, respectively.

CYP79A1 is not able to metabolize phenylalanine (Kahn et al.,1999; Møller and Conn, 1980), demonstrating the importance ofthe p-hydroxy group for substrate recognition. Two residues inthe distal part of the substrate pocket Ser144 from the B-C loopand Thr538 from the C-terminal in CYP79A1 were found in a posi-tion to present hydrogen-bonding possibilities for the p-hydroxygroup of the substrate (Fig. 3). Site directed mutagenesis to ex-change these two amino acid residues with e.g. alanine residuesmight alter the substrate specificity to include phenylalanine. Inother CYP proteins, Ser and Thr residues from the B–C loop posi-tioned at the distal end of the substrate pocket are also proposedto interact with substrates either directly or mediated by the pres-ence of water molecules (Sansen et al., 2007).

The structural model of CYP79A1 proposes that residues Arg152and Asn355 interact with the carboxylate and amino group of theL-tyrosine substrate. The only substrate known to be metabolizedby CYP79A1 is L-tyrosine. In addition, 18O labelling studies haveshown that the hydroxy group introduced during the conversionof L-tyrosine to N-hydroxytyrosine is removed in the CYP79A1 cat-alyzed conversion of N,N-dihydroxytyrosine into p-hydrox-yphenylacetaldoxime (Halkier et al., 1991). This is in agreementwith the tight substrate specificity of CYP79A1 and the small sub-strate binding pocket elucidated by structural modeling but alsodictates that specific and strong interactions prevents free rotationof the C-N bond following the first N-hydroxylation step maintain-ing the CH-N-OH function in a rigid position during CYP79A1 catal-ysis. The residues Arg152 and Asn355 offer positive electrostaticcharges to position the carboxylate ion and by establishing hydro-gen bonds, maintain the N-hydroxy groups into a fixed position asrequired for specific loss of the hydroxyl group introduced in thefirst N-hydroxylation reaction during the final conversion of N,N-dihydroxytyrosine into p-hydroxyphenylacetaldoxime. Likewise,an internal stabilizing interaction between the substrate carboxyl-ate and hydroxy group could add to a locked and rigid conforma-tion around the C–N bond.

In CYP71E1, residues Ser125 and Thr511 are placed in the sameregion as Ser144/Thr538 of CYP79A1, and would appear to fulfillthe same role, i.e. positioning the p-hydroxy group of p-hydrox-yphenylacetaldeoxime in a favorable spatial orientation for cata-lytic conversion into p-hydroxymandelonitrile. However,CYP71E1 has slightly broader substrate specificity than CYP79A1as it metabolizes both p-hydroxyphenylacetaldeoxime and

phenylacetaldeoxime, derived from tyrosine and phenylalanine,respectively (Kahn et al., 1999). The lack of a p-hydroxy groupand subsequently a non-optimal spatial orientation of phen-ylacetaldeoxime for catalytic processing correspond well withexperimental results showing a lower catalytic conversion of phen-ylacetaldeoxime, making it a suboptimal substrate for CYP71E1compared to p-hydroxyphenylacetaldeoxime (Kahn et al., 1999).Substrate specificity in CYP71E1 is probably conferred by a combi-nation of the restricted size of the substrate binding pocket mainlycaused by the presence of two phenyalanines (Phe240 and Phe512)which impose conformational restrictions on the substrate andthe energetically favorable electrostatic interactions between thearomatic rings of Phe240/512 and of (p-hydroxy)phenylace-taldeoxime.

Six substrate recognition sites termed SRS1–6 have been shownto be important determinants for substrate recognition in CYPs(Gotoh, 1992). These SRSs were identified in CYP79A1 andCYP71E1 by structural alignment with CYP101 (PDB code 2CPP).Whereas the aromatic ring of the substrate is contacted by residuesfrom SRS2, 5 and 6, SRS1 and 4 is primarily responsible for shapingthe environment around the end of the substrate that is convertedby the enzymes, i.e. the amino acid group for CYP79A1 and theoxime group for CYP71E1 (Fig. 3C and D). Four CYPs (CYP73A5,CYP84A1, CYP75B1 and CYP98A3) active in the phenylpropanoidpathway in Arabidopsis thaliana have previously been modeled,and substrate docking showed a remarkable conservation in regardto substrate orientation and substrate interactions with the SRS re-gions within the different CYPs (Rupasinghe et al., 2003). The sub-strates all consisted of a reactive aromatic region, which was foundin proximity to the SRS4, 5 and 6 regions, whereas the non-reactivetail region was localized next to the SRS1 and 2 regions. Despitethat the aromatic ring system in tyrosine and its derived oximeis the non-reactive region of the substrate, the interactions be-tween substrate and SRS regions are very similar in the A. thalianaCYPs compared to interactions in CYP79A1/71E1, A difference isthe spatial orientation of the substrate, which in CYP79A1 andCYP71E1 is positioned over heme rings II and III and perpendicularto SRS4 (helix I) and not parallel to SRS4 as seen for the phenyl-propanoid CYPs. Instead, CYP79A1/CYP71E1 substrate bindingmodes appears to be more comparable to the bacterial CYP102,in which palmitoleic acid is positioned perpendicular to SRS4 (Liand Poulos, 1997). Given the differences in substrate sizes andhydroxylation specificities of the different CYPs, further in-depthcomparisons of substrate binding were not feasible.

3.2. Membrane positioning and CPR interaction

Membrane insertion of CYP proteins holds important implica-tions for their functional properties, both with respect to theirinteraction with other proteins such as CPR and for potential sub-strate availability. Measurements based on tryptophane fluores-cence and antibody interactions have suggested that somehydrophobic loop sequences of CYP proteins are membrane associ-ated and buried in the lipid bilayer of the membrane to varying de-grees dependent on each particular CYP protein (Ozalp et al., 2006).The depth of insertion into the lipid bilayer is thought to influencesubstrate entry/product exit at the protein-membrane interface(Bayburt and Sligar, 2002; Ozalp et al., 2006) and to guide properinteraction with CPR or potential interactions with other CYP pro-teins. The solved crystal structure of the mammalian CYP5C2showed that a hydrophobic tip protrudes around the site of attach-ment of the N-terminal anchor. This suggests that this tip wasembedded into the lipid bilayer (Williams et al., 2000). The major-ity of the amino acids forming this protrusion on the foldedCYP5C2 protein were found in the region just downstream of themembrane anchor and before the A-helix. In the proposed

2120 K. Jensen et al. / Phytochemistry 72 (2011) 2113–2123

structural models of CYP79A1 and the CYP71E1 presented in thisstudy, a hydrophobic tip protruding from the surface was formedby residues positioned in corresponding regions of the primaryamino acid sequence. A hydrophobic patch suitable for membraneinteraction was also identified in the F–G loop, most clearly in thestructural model of CYP79A1. In accordance with this observation,membrane interaction of mammalian microsomal CYP proteins hasalso been suggested to involve hydrophobic residues in the F–Gloop region (Sansen et al., 2007; Williams et al., 2000).

Modeling and Trp fluorescence quenching studies of the mam-malian CYP2C5 and CYP2C2 proteins have suggested amino acidsin the b1 sheet region as well as residues in the b-2-2 strand tobe membrane associated (Ozalp et al., 2006; Williams et al.,2000). The proposed CYP79A1/CYP71E1 dockings onto/into themembrane positioned the residues of the b1 sheet region at themembrane surface (Fig. 1) and tilting the CYP protein to a heme an-gle of �50� brought the b-2-2 strand (shown in orange in Fig. 1)into contact with the membrane surface. The 50� angle betweenthe heme plane and the membrane is within the range of reportedangles discussed by Bayburt and Sligar (2002).

The CPR interacting surface on CYP proteins have been studiedby point mutational studies. A patch of positively charged residuescorresponding to the CYP models patch 1 in the C-helix has beenidentified to be important for CPR interactions (Bridges et al.,1998; Shimizu et al., 1991). In the identified patch 2, the conservedarginine from the cys-pocket has been shown experimentally to beimportant (Bridges et al., 1998; Shen and Strobel, 1993; Shimizuet al., 1991) as has the C–D loop arginine from patch 3 (Bridgeset al., 1998). Studies of mammalian CPR and CYP crystal structuressuggest rearrangement of the CPR domains for a productive inter-action between the CPR FMN domain and the CYP heme to occur(Williams et al., 2000; Zhao et al., 1999). This has recently beenshown to occur by structural changes centered on the hinge regionresulting in a major conformational change of the overall CPRstructure without interfering with the 3D structure of the individ-ual CPR domains (Ellis et al., 2009; Laursen et al., 2011). Such amechanism is in accordance with our model where the twopatches of negatively charged residues cluster 1 and 3 are posi-tioned on the side of the SbCPR2b FMN domain facing towardsthe FAD domain. Opening of the CPR protein allowed by flexibilityin the hinge region connecting the domains would expose this sur-face and allow for CYP interaction.

3.3. Substrate channelling and metabolon formation

The crystal structure of one of the templates used in this study,CYP2B4, has been solved both in a closed (PDB: 1SUO) (Scott et al.,2004) and in an open conformation (PDB: 1PO5) (Scott et al., 2003).Comparing the open CYP2B4 crystal with the CYP79A1 andCYP71E1 models, a similar opening would occur by displacementof the B loop and the F–G loop. The B-loop runs from the N-termi-nal end of the I-helix into the protein and displacement would re-sult in an opening facing downwards towards the membrane(Fig. 1). Entry of hydrophobic substrates is likely to occur throughan entry facing directly towards the lipid bilayer. The substrate forCYP79A1 is L-tyrosine. Tyrosine is an amphoteric and hydrophilicmolecule, an unusual characteristic for a CYP substrate. In contrast,p-hydroxyphenylacetaloxime, the substrate of CYP71E1, has theclassical hydrophobic properties of a CYP substrate. The entryroute for the CYP79A1 substrate is thus likely to be either directlyfrom the cytosol or along the charged phospholipid heads at thetop of the lipid bilayer. The proposed cytosolic entry site is onthe surface of CYP79A1 opposite the SbCPR2b interacting surfaceand would allow substrate entry simultaneous with SbCPR2binteraction.

From the models of CYP79A1 and CYP71E1, no clear indicationsof interacting domains facilitating channelled product transferfrom CYP79A1 to CYP71E1 were obtained. Both CYPs are boundto the cell membrane by a transmembrane hydrophobic N termi-nus. In several studies, the transmembrane domain has beenshown to mediate CYP oligomerization (Ozalp et al., 2005;Subramanian et al., 2010; Szczesna-Skorupa et al., 2003). Thesuccessful crosslinking of microsomal CYPs using a crosslinking re-agent aimed at charged residues primarily exposed to the aqueousenvironment, indicated that the transmembrane domain was mostlikely not the sole determinant of CYP interaction (Alston et al.,1991). The function of the transmembrane domain in mediatingprotein–protein interaction could be to properly direct andposition CYPs in specific lipid environments (‘‘rafts’’) within themembrane, thereby facilitating CYP-CYP and CPR-CYP interactions.The CYP distribution in the endoplasmic reticulum is still debated,ranging from non-uniformly distributed to semiorganized zonaldifferences within the endoplasmic reticulum (Jungermann,1995; Matsuura et al., 1978; Oinonen and Lindros, 1998). Thetransmembrane domains could easily be envisioned to influenceCYP distribution in the endoplasmic reticulum and thereby medi-ate the formation of hotspots for CYP-CYP interaction. Based onthese observations, it would be interesting to modify or replacethe transmembrane domains of CYP79A1 and CYP71E1 to furtherinvestigate their significance for metabolon formation.

Modeling is further complicated because it is unknown whetherthe same or different CPR molecules are required to mediate elec-tron donation to CYP79A1 and CYP71E1. In both cases it needs tobe resolved how the large conformational changes of the CPR mol-ecule in the course of its catalytic action as electron donor to CYPsmay interfere with or favor shuttling of electrons from the sameCPR molecule to CYP79A1 and CYP71E1. Reconstitution of theCYP79A1, CYP71E1 and SbCPR2b in nanodiscs followed by bindingstudies with UGT85B1 as monitored by surface plasmon resonancespectroscopy and atomic force microscopy may provide answers tothese issues (Borch and Hamann, 2009). Some metabolons are nowknown to be function dependent structures (Brandina et al., 2006;Sweetlove and Fernie, 2005; Thellier et al., 2004) where proteinassembly is controlled e.g. by the presence of intermediates orinhibitors. CYP71E1 is known to lose activity when isolated inthe absence of a strong reductant (Møller and Conn, 1980). Thismay be related to major structural changes of CYP71E1 and wouldresult in p-hydroxyphenylacetaldoxime as being the final productformed. A shift between cyanogenic glucoside and oxime forma-tion would enable the plant to alternate between producing insectand fungal repellants, respectively (Møller, 2010). It remains to beshown whether such factors control establishment and assemblyof a dhurrin metabolon. CYP79 proteins are also involved in thesynthesis of glucosinolates, again catalyzing conversion of parentamino acids to the corresponding oximes (Halkier and Gershenzon,2006). In glucosinolate producing plants the oximes may be usedfor synthesis of additional secondary metabolites like camalexin(Bottcher et al., 2009). A function dependent metabolon would ex-plain how CYP79 proteins could be involved in directed synthesisof numerous different types of compounds even within the sameplant species.

3.4. Putative interactions between CYP71E1 and UGT85B1 derivedfrom the structural models

Surface charge and hydrophobicity of the models of CYP79A1and CYP71E1 were analysed to identify regions of potential impor-tance for interaction between the two CYP proteins or domainssuitable to provide a binding region for recruitment of the solubleUGT85B1 glucosyltransferase required for the final step of dhurrinsynthesis (Jones et al., 1999). The interaction between CYP71E1

K. Jensen et al. / Phytochemistry 72 (2011) 2113–2123 2121

and UGT85B1 is an important step in product channelling becausea rapid glucosylation of the labile p-hydroxymandelonitrileproduct of CYP71E1 is necessary to avoid its dissociation intop-hydroxybenzaldehyde and hydrogen cyanide. A patch of posi-tively charged surface exposed amino acid residues (Lys88,Arg92, His95, Arg99, Arg450, Arg460, Lys531) was observed onthe opposite face of CYP71E1 compared to the attachment sitefor CPR. Complementary to this positively charged patch, a patchof negatively charged residue was found to be located on the C-ter-minal end of the B loop in UGT85B1 (Osmani et al., 2009; Thorsøeet al., 2005). This loop is hypervariable and usually longer inUGT85s than in other UGTs and has, based on the latter two char-acteristics, previously been suggested as a domain involved inCYP71E1 interaction (Thorsøe et al., 2005). The modeling studyprovides further support to the validity of this suggestion.

4. Conclusion

In conclusion, we have generated structural models of the threemembrane anchored proteins of the dhurrin metabolon. Thesemodels will provide a structural basis both for expanding ourknowledge of the individual proteins and elucidating how theyinteract. Given the significant advantages associated with metabo-lon formation, such as improved kinetics and catalytic efficiency(Møller, 2010), a greater understanding of the factors involved inmetabolon formation is very interesting in a biotechnologicalperspective.

5. Experimental

5.1. Homology modeling

Protein models were built using the Orchestra protein modelingcomponent of the Sybyl software from Tripos. Secondary structurepredictions of query sequences were performed using Jpred http://www.compbio.dundee.ac.uk/~www-jpred/submit.html. Structuralalignments of query and template sequences were edited manuallyand entered into Orchestra for modeling. Structural models werevisualized using PYMOL (www.pymol.org). APBS was used for cal-culation and visualization of the electrostatic potential (Bakeret al., 2001).

5.2. Structural modeling of CYP79A1

CYP79A1 was modeled using the coordinates for the solvedcrystal structures of the human CYP proteins 2B4 (1SUO) (Scottet al., 2004), 1A2 (2HI4) (Sansen et al., 2007), 2A13 (2P85) (Smithet al., 2007), and 2D6 (2F9Q) (Rowland et al., 2006). A structuralalignment of CYP79A1 against the four template sequences wasachieved by a BLAST search followed by manual editing. The aminoacid sequence identity of CYP79A1 was 21.7% to 2B4, 24.1% to 1A2,22.2% to 2A13 and 22.6% to 2D6. The model was built including thecoordinates of the heme group from the crystal structure 2HI4 dur-ing modeling. From the structural alignment, Orchestra built amodel that included 387 of the 558 amino acid residues inCYP79A1. Addition of loops identified by a loop search in Orchestraresulted in a model with 445 residues and six missing regions, onecorresponding to the N-terminal. The remaining five missing re-gions (residues 117–124, 138–154, 312–322, 330–333 and 536–549) were modeled in Sybyl by a database search suggesting struc-tures for the regions and guided by overlay with the CYPs used astemplates for modeling. The final model included 489 residuesleaving out only the N-terminal residues 1–69 from the structuralmodel. The energy of the model structure was optimized by severalrounds of energy minimisations in Sybyl, monitored manually to

avoid major rearrangements of the structure. Bond lengths and an-gles were verified and optimized by consultation of Ramachandranmaps followed by energy minimisations. In the final structuralmodel, no residues were in disallowed regions and only 11 (2.6%)were in generously allowed regions.

5.3. Structural modeling of CYP71E1

CYP71E1 was modeled using the coordinates for the solvedcrystal structures of the human CYP proteins 2B4 (1SUO) (Scottet al., 2004), 1A2 (2HI4) (Sansen et al., 2007), 2C9 (1R9O) (Westeret al., 2004), and 2D6 (2F9Q) (Rowland et al., 2006). FollowingBLAST searches and manual optimization of the structural align-ment, the amino acid sequence identity of CYP71E1 was 26.3% to1SUO, 24.9% to 2HI4, 21.9% to 1R9O and 23.3% to2F9Q. The modelwas built including the coordinates of the heme group from thestructure for 2HI4 during the modeling. From the structural align-ment, a model was built in Orchestra including 366 of the 531 ami-no acid residues in CYP71E1. Addition of loops identified by a loopsearch in Orchestra resulted in a structural model encompassing458 residues and three missing regions of which two correspondedto the N- and C terminal residues 1–47 and 529–531, respectively,and were not included in the final model. The third missing se-quence region contained the amino acid residues 115–137. To beable to include this loop in the structural model of CYP71E1, the re-gion containing the amino acid residues 114–148 was remodeledin Orchestra by alignment to the crystal template 1SUO. The en-ergy of the model was optimized by several rounds of energy min-imizations in Sybyl, monitored manually to avoid majorrearrangements of the structure. Bond lengths and angles wereverified and optimized by consultation of Ramachandran maps fol-lowed by energy minimizations. The final model included 481 ofthe 531 residues of CYP71E1. No amino acid residues were in dis-allowed regions and only five (1.2%) were in generously allowedregions.

5.4. Substrate docking into models of CYP79A1 and CYP71E1

Substrates were docked manually into the models of CYP79A1and CYP71E1. The substrates were docked into the substrate pock-ets above the heme so that the atom to be hydroxylated was posi-tioned 4 Å above the center of the heme. Other aspects taken intoconsideration during substrate docking were the available spaceand shape of the substrate pocket and the potential for formationof favorable interactions between the substrates and side chainsof the amino acids lining the substrate pockets.

5.5. Structural modeling of SbCPR2b

SbCPR2b was modeled using the coordinates for the crystalstructures of the reductases RnCPR (1AMO) (Wang et al., 1997),RnCPR W677X (1JA0) (Hubbard et al., 2001), HsCPR (1B1C) (Zhaoet al., 1999) and ScCPR (2BF4) (Lamb et al., 2006) sharing 38.9%,38.4%, 39.6% and 33.5% amino acid sequence identity, respectively,when structurally aligned. Modeling was performed includingcoordinates for the cofactors FAD, FMN and NAD from the structureof 1AMO. A structural model of SbCPR2b including 489 residueswas built in Orchestra. Additional loop searches in orchestra re-sulted in a model with 626 residues and two gaps correspondingto the N-terminal residues 1–63 and to the residues 463–481.The amino acid residues 461–481 were modeled following data-base searches in Sybyl. Bond lengths and angles of all amino acidresidues in the model were verified and optimized by consultationof Ramachandran maps followed by energy minimizations. The fi-nal model included 643 of the 706 amino acid residues of SbCPR2bleaving out residues 1–63. Nine (1.6%) residues were in disallowed

2122 K. Jensen et al. / Phytochemistry 72 (2011) 2113–2123

regions and only eight (1.4%) were in the generously allowedregions.

Acknowledgements

The research was supported by the Villum Foundation researchcentre ‘‘Pro-Active Plants’’, by the UNIK Center for Synthetic Biol-ogy funded by the Danish Ministry for Science, Technology andInnovation and by a research grant from the Danish ResearchCouncil for Independent Research/Technology and Production Sci-ences. Kenneth Jensen and Thomas Hamann were supported by aPh.D. fellowship from the University of Copenhagen. Peter Naurwas supported by the Carlsberg Foundation.

Appendix A. Supplementary data

Supplementary data associated with this article can be found, inthe online version, at doi:10.1016/j.phytochem.2011.05.001.

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