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The Crystal Structure of Arabidopsis thaliana Allene Oxide Cyclase: Insights into the Oxylipin Cyclization Reaction^[W]

^a Lehrstuhl für Biophysik, Ruhr-Universität Bochum, D-44780 Bochum, Germany
^b Lehrstuhl für Pflanzenphysiologie, Ruhr-Universität Bochum, D-44780 Bochum, Germany

² To whom correspondence should be addressed. E-mail eckhard.hofmann{at}bph.rub.de or florian.schaller{at}rub.de; fax 49-234-32-14748 or 49-234-32-14187.

	ABSTRACT

TOP ABSTRACT INTRODUCTION RESULTS DISCUSSION METHODS REFERENCES

 
We describe the crystallization and structure elucidation of Arabidopsis thaliana allene oxide cyclase 2 (AOC2), a key enzyme in the biosynthesis of jasmonates. In a coupled reaction with allene oxide synthase, AOC2 releases the first cyclic and biologically active metabolite, 12-oxo-phytodienoic acid (OPDA). AOC2 (AT3G25770) folds into an eight-stranded antiparallel ß-barrel with a C-terminal partial helical extension. The protein forms a hydrophobic binding cavity with two distinct polar patches. AOC2 is trimeric in crystals, in vitro and in planta. Based on the observed folding pattern, we assigned AOC2 as a low molecular weight member of the lipocalin family with enzymatic activity in plants. We determined the binding position of the competitive inhibitor vernolic acid (a substrate analog) in the binding pocket. Based on models for bound substrate 12,13-epoxy-9,11,15-octadecatrienoic acid and product OPDA, we propose a reaction scheme that explains the influence of the C15 double bond on reactivity. Reaction is promoted by anchimeric assistance through a conserved Glu residue. The transition state with a pentadienyl carbocation and an oxyanion is stabilized by a strongly bound water molecule and favorable – interactions with aromatic residues in the cavity. Stereoselectivity results from steric restrictions to the necessary substrate isomerizations imposed by the protein.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION RESULTS DISCUSSION METHODS REFERENCES

 
Since the initial discovery of methyl jasmonate (MeJA) as a secondary metabolite in essential oils of jasmine in 1962 (Demole et al., 1962), JAs have become accepted as a new class of plant hormone. In the early 1980s, their widespread occurrence throughout the plant kingdom (Meyer et al., 1984) and their growth-inhibitory (Dathe et al., 1981) and senescence-promoting activities (Ueda and Kato, 1980) were established. It has become increasingly clear, however, that biological activity is not limited to JA but extends to, and even differs among, its many metabolites and conjugates as well as its cyclopentenone precursors (Kramell et al., 1997; Stintzi et al., 2001). Because of the different biological function of each member of the group of JAs, the enzymes of JA biosynthesis and metabolism may thus have a regulatory function in controlling the activity and relative levels of different signaling molecules.

Research in recent years generally confirmed the Vick and Zimmerman pathway of JA biosynthesis (the octadecanoid pathway) and made considerable progress with respect to the biochemistry of the enzymes involved as well as the molecular organization and regulation of the pathway.

The early elucidation of the JA biosynthetic pathway and the demonstration of the growth-inhibiting and senescence-promoting activity were followed by the discovery that JAs are involved in plant defense reactions, which greatly stimulated the interest in these compounds as plant signaling molecules. A role in plant defense was first shown by Farmer and Ryan, who demonstrated the induction of proteinase inhibitors by MeJA and JA as part of the defense response against herbivorous insects (Farmer and Ryan, 1990; Farmer et al., 1991). JAs were then shown to be active inducers of antimicrobial phytoalexins by Gundlach et al. (1992), and subsequent work clearly established their defense gene–inducing activity for induced resistance against insect predators and pathogens (for a review, see Reymond and Farmer, 1998; Weiler et al., 1998; Blée, 2002). Such a role was unequivocally confirmed by the analysis of mutants compromised in either the synthesis or the perception of JA signals (for a review, see Schaller et al., 2005).

The pathway of JA biosynthesis is shown in Figure 1 . Biosynthesis is believed to start with the oxygenation of free -linolenic acid (LA), which is converted to (9Z,11E,15Z,13S)-13-hydroperoxy-9,11,15-octadecatrienoic acid [13(S)-HPOT] in a reaction catalyzed by 13-lipoxygenase. The presumed release of LA from membrane lipids through the action of a lipase may be triggered by local or systemic signals, such as oligogalacturonides, chitosan, systemin (Farmer and Ryan, 1992; Mueller et al., 1993; Narváez-Vásquez et al., 1999), or wounding (Conconi et al., 1996; Narváez-Vásquez et al., 1999). The 13(S)-HPOT serves as a substrate for several enzymes (Figure 1), such as divinylether synthase, lipoxygenase, hydroperoxide reductase, epoxyalcoholsynthase, peroxygenase, hydroperoxide lyase, or allene oxide synthase (AOS) (Blée and Joyard, 1996; Vick and Zimmerman, 1981, 1987). AOS converts 13(S)-HPOT to the unstable epoxide 12,13(S)-epoxy-9(Z),11,15(Z)-octadecatrienoic acid (12,13-EOT), which is cyclized by allene oxide cyclase (AOC) to the first cyclic and biologically active compound of the pathway, 12-oxo-phytodienoic acid (OPDA). Reduction of the 10,11-double bond by an NADPH-dependent OPDA reductase then yields 3-oxo-2(2'(Z)-pentenyl)-cyclopentane-1-octanoic acid (OPC-8:0), which is believed to undergo three cycles of ß-oxidation to yield the end product of the pathway [i.e., JA with (3R,7S)-configuration; (+)-7-iso-JA] (Vick and Zimmerman, 1984). The biosynthesis of JA appears to involve two different compartments: the conversion of LA to OPDA is localized in the chloroplasts (Vick and Zimmerman, 1987; Song et al., 1993; Laudert et al., 1997), while the reduction of OPDA to OPC-8:0 (Schaller and Weiler, 1997a, 1997b; Schaller et al., 2000; Strassner et al., 2002) and the three steps of ß-oxidation (i.e., conversion of OPC-8:0 to JA) occur in peroxisomes (Gerhardt, 1983; Vick and Zimmerman, 1984; Li et al., 2005). Like JA, OPDA may accumulate to substantial amounts in plant tissue (Stelmach et al., 1998). By contrast, OPC-8:0 occurs only in trace amounts.

View larger version (21K): [in this window] [in a new window] [as a PowerPoint slide]   Figure 1. The Metabolic Fate of Fatty Acid Hydroperoxides. Fatty acid hydroperoxides [only the 13(S)-hydroperoxy derivative of linolenic acid is shown] are channeled into seven distinct pathways for oxylipin biosynthesis. The committed steps are catalyzed by AOS for JA biosynthesis, divinylether synthase for the formation of divinyl ethers, lipoxygenase for the formation of keto polyunsaturated fatty acids (PUFAs), hydroperoxide reductase in the formation of hydroxy PUFAs, epoxyalcohol synthase and peroxygenase resulting in epoxy hydroxy PUFAs, and hydroperoxide lyase for the production of leaf aldehydes (green leaf volatiles). In the AOS branch, the Vick and Zimmerman pathway for JA biosynthesis is shown. 13-LOX, 13-lipoxygenase; OPR3, 12-oxophytodienoate reductase 3. Figure adapted from Feussner and Wasternack (2002) and Wasternack and Hause (2002).

An interesting aspect of the AOC reaction is the apparent competition between the spontaneous decomposition of its substrate, the unstable allene oxide, to form - and -ketols and racemic cis-OPDA, and the enzyme-catalyzed formation of optically pure (9S,13S)-OPDA [i.e., cis(+)-OPDA], the first pathway intermediate having the characteristic pentacyclic ring structure of JAs (Figure 1) and biological activity. The extremely short half-life of allene oxides (half-time <30 s in water; Brash et al., 1988) and the optical purity of natural OPDA suggest a tight coupling of the AOS and AOC reactions, possibly in a synthase-cyclase complex (Stenzel et al., 2003b). Coupling of the two reactions is also observed in vitro: AOC from potato (Solanum tuberosum) or recombinant Arabidopsis thaliana AOC2 in combination with recombinant Arabidopsis AOS resulted in the production of highly asymmetrical cis-OPDA consisting nearly exclusively of the (9S,13S)-enantiomer (Laudert et al., 1997). Recently, we demonstrated that a physical interaction of AOS and AOC is not essential for the cyclization reaction, but the yield of OPDA formation and optical purity of the product is diminished when separating both enzymes from each other (P. Zerbe, E.W. Weiler, and F. Schaller, unpublished data).

AOC has been cloned as a single-copy gene from tomato (Solanum lycopersicum; Ziegler et al., 2000) and barley (Hordeum vulgare; Maucher et al., 2004) and as a small gene family (AOC1-4) from Arabidopsis (Stenzel et al., 2003b). Inspection of the N termini of the cloned AOCs revealed the presence of transit peptides for plastid targeting, and localization in chloroplasts was confirmed immunohistochemically (Ziegler et al., 2000; Stenzel et al., 2003a, 2003b) and by transient expression of AOC1-4/green fluorescent protein fusions (F. Schaller and P. Zerbe, unpublished data).

To gain more information about the molecular mechanisms underlying the cyclization of oxylipins, we set out to elucidate the structure of an AOC from Arabidopsis. We decided to focus our biochemical and structural studies on AOC2 because this isozyme has been shown to be highly expressed in planta and was described as being the most active of the four Arabidopsis AOCs (Stenzel et al., 2003b).

	RESULTS

TOP ABSTRACT INTRODUCTION RESULTS DISCUSSION METHODS REFERENCES

 
Overexpression, Purification, and Crystallization of Arabidopsis AOC2
Because of relatively weak expression levels of published constructs (C. Wasternack, personal communication) we cloned differently truncated versions of AOC2 in pET 21b(+) (Novagen/Merck His6-tag, C-terminally) and pQE30 (Qiagen His6-tag, N-terminally) and analyzed expression levels of insoluble and soluble AOC2. Constructs beginning at base 132 showed the highest expression of soluble protein in both vectors and were subsequently used.

Using SDS-PAGE analysis, we observed a strong influence of the His6-tag location on the oligomerization state of the protein. While AOC2 with a C-terminal His6-tag runs as a monomer with an apparent mass of 22 kD, the N-terminally tagged protein shows a dominant band at a molecular mass of 60 kD, consistent with an SDS-stable trimer formation. The identity of the bands could be confirmed by immunoblot analyses using antibodies raised against AOC2 or the His6-tag.

A similar pattern to that of the N-terminally tagged protein has been observed in immunoblot analyses of native AOCs from plant extracts (P. Zerbe and F. Schaller, unpublished data). This clearly shows that the trimeric form is present in planta as well and is not an artifact due to addition of the N-terminal His6-tag. Addition of a C-terminal His6-tag seems to destabilize the trimers.

Overall Structure
The structure of Arabidopsis AOC2 has been determined by multiwavelength anomalous dispersion with selenomethionine (SeMet)-labeled protein crystals of the space group P2₁2₁2₁ and was refined to a resolution of 1.5 Å (SeMet-AOC2). Additionally, we solved the structure of the unlabeled protein in space group P2₁ at a resolution of 1.8 Å (WT-AOC2). In both space groups, AOC2 forms closely packed trimers. In space group P2₁, we find two copies of this AOC2 trimer, which superimpose with a root mean square deviation (RMSD) of 0.33 Å for all C atoms. Both align well with the single trimer found in the SeMet-AOC2 structure with RMSDs of 0.40 and 0.44 Å, respectively. Data collection and refinement statistics are presented in Table 1 .

View larger version (57K): [in this window] [in a new window] [as a PowerPoint slide]   Figure 2. Structure of the AOC2 Monomer. (A) Side view of the AOC2 monomer shown in ribbon representation, color changing from blue (N terminus) to red (C terminus). (B) Top view, rotated by 90° toward the viewer with respect to (A). The same representation is used as in (A). (C) Molecular surface representation of the proposed binding pocket. The orientation and color scheme are similar to (A). Also shown are the residues discussed in the text in stick representation. Water 75 is shown as a red sphere. (D) Topology diagram in the same color coding as in (A). ß-Strands are shown as arrows (with numbers indicating the first and last residues) and helices as small cylinders.

The barrel forms an elongated cavity, which is lined mostly by aromatic and hydrophobic residues and reaches 14 Å into the protein (Figure 2C). Three areas of the cavity surface are noteworthy. First, the conserved Glu-23 is positioned at the very bottom of the cavity, introducing a negative charge. Second, an additional polar patch is formed by Ser-31, Asn-25, and Asn-53 on one side of the cavity. These three residues together with the main chain nitrogen from Pro-32 coordinate a tightly bound water molecule that is found in all our AOC2 structures. Third, we found Cys-71 on the opposing wall of the cavity. These residues are strictly conserved among all AOC sequences in the EBI UNIRef100 database (Figure 3 ).

View larger version (104K): [in this window] [in a new window] [as a PowerPoint slide]   Figure 3. Sequence Alignment of Known AOCs. Representation of secondary structure elements and numbering is based on the SeMet-AOC2 structure. Asterisks mark residues modeled with alternate conformations. The following AOC2 sequences are listed (accession numbers in parentheses): Arabidopsis, AOC2_AT (Q9LS02), AOC1_AT (Q9LS03), AOC3_AT (Q9LS01), and AOC4_AT (Q93ZC5); Pisum sativum, AOC_PS (Q3LI84); Medicago truncatula, AOCa_MT (Q711Q9) and AOCb_MT (Q599T8); Oryza sativa, AOC_OS (Q8L6H4); Nicotiana tabacum, AOC_NT (Q711R1); Hordeum vulgare, AOC_HV (Q711R0); Zea mays, AOC_ZM (Q6RW09); Humulus lupulus, AOC1_HL (Q68IP7); AOC4_HL (Q68IP6); Bruguiera sexangula, MAN_BS (Q9AYT8); Lycopersicon esculentum, AOC_LE (Q9LEG5); S. tuberosum, AOC_ST (Q8H1X5); Physcomitrella patens, AOCa _PP (Q8GS38) and AOCb_PP (Q8H0N6). Figure was produced using the ESPript server (Gouet et al., 1999).

View larger version (63K): [in this window] [in a new window] [as a PowerPoint slide]   Figure 4. Trimeric Form of Arabidopsis AOC2. Ribbon representation of the AOC2 trimer. Residues 15 to 147 are shown in gray, and residues 148 to 188 are colored red. The view is along the trimer axis.

The lipocalin protein family (Pervaiz and Brew, 1985, 1987; Grzyb et al., 2006) is a large group of small extracellular proteins with great diversity at the sequence level but mostly with either three characteristic short conserved sequence motifs (SCR I-III; the kernel lipocalins) or else one conserved motif (SCR I; the outlier lipocalins). However, the crystal structures are highly conserved and comprise a single eight-stranded continuously hydrogen-bonded antiparallel ß-barrel with an integral ligand binding site (Figure 5 ). The structural basis for these SCRs is a clustering of conserved residues around a central Trp in SCR I at the beginning of ß-strand A, which stabilizes barrel closure. Several members of the lipocalin family bind to specific cell surface receptors or form complexes with soluble macromolecules. Initially, lipocalins were described as transport proteins; however, it has become clear that the lipocalins exhibit a multitude of functions, with roles in retinol transport, olfaction, and pheromone transport, prostaglandin biosynthesis, invertebrate cryptic coloration, and a role in the xanthophyll cycle. In addition to the central ß-barrel, the C-terminal part of the protein is usually appended by a characteristic -helix. One end of the barrel is closed by the N terminus that traverses the base of the barrel (Figure 5). At the opposite end, the ß-barrel is typically open to solvent and provides access to the cavity. These binding sites of the lipocalins may have very different shapes (e.g., with a ligand pocket like an extended cave with lobes, a ligand pocket deep within the hydrophobic core, with loops encapsulating the ligand, or a binding site with a wide, funnel-like opening to the solvent) (for a review, see Schlehuber and Skerra, 2005).

View larger version (45K): [in this window] [in a new window] [as a PowerPoint slide]   Figure 5. Structural Superposition of AOC2 and RBP. Stereo diagram of pig RBP (Protein Data Bank accession code 1AQB) with the bound ligand retinol superposed with AOC2. RBP is drawn as a yellow ribbon, and the bound retinol is shown in van der Waals representation in purple and red. AOC2 is shown as a gray ribbon. To illustrate the discussed strand exchange, N-terminal portions of both proteins are colored blue (residues preceding the first ß-strand in RBP and in the first ß-strand in AOC2). C-terminal stretches of the proteins are colored maroon (including the last ß-strand for RBP) and red (after the last residue in the barrel for AOC2). For clarity, the last 29 residues of RBP have been omitted, which include the conserved lipocalin helix mentioned in the text.

Stretches of superposed residues all fall within the ß-strands, therefore aligning the barrel walls well, while the loop regions show large differences. AOC2 does not show the classical C-terminal helix, and the barrel is not closed by the preceding sequence but by the extended C terminus. Most striking is the different position of the termini with respect to the barrel orientation. In the orientation shown in Figure 5, the origin of the closed ß-sheet is on the top for the lipocalins, whereas it is on the bottom side for AOC2. This topological difference could be the result of a strand exchange in which either the first or last ß-strand (strand A or strand H in lipocalin nomenclature) has been replaced by C- or N-terminal sequence stretches, respectively. Notably, the sequence DLVPFTNKLY (residues 47 to 56) from AOC2 could be automatically aligned to SCR I in pig RPB (accession number 1AQB). While we can observe a weak sequence similarity in this stretch, the conserved Gly and Trp (residues 22 and 24, respectively, in RBP) are missing in the AOC sequence. Interestingly, the region identified in AOC2 falls within the second ß-strand and superimposes spatially with the location of SCR I in ß-strand A of the lipocalins (Figure 5).

Recently, the structure of a coral AOS domain has been reported (Oldham et al., 2005). In soft corals, no AOC has been described so far, so an intriguing possibility would be that coral AOS catalyzes both epoxidation and cyclization reactions. Indeed, coral AOS has been found to have a catalase-type fold, which includes an eight-stranded antiparallel barrel. However, there has been no report that the barrel fulfils a catalytic role in cyclization in this case, and the interior is tightly packed with protein side chains, preventing a similar substrate binding as in lipocalins.

Substrate Specificity
To gain more information about the catalytic mechanism, we tested 13(S)-HPOT and 13(S)-hydroperoxy-octadecadienoic acid (13(S)-HPOD) as substrates for Arabidopsis AOC2 and AOS. We found that with 13(S)-HPOD as substrate, only negligible amounts of cyclic product were formed (data not shown). Since AOS makes use of linoleic acid and linolenic acid derivatives (Siedow, 1991; Song and Brash, 1991; Pan et al., 1995; Laudert et al., 1996), AOC2 is apparently incapable of cyclizing the corresponding epoxide, which was indicated before by studies of Ziegler et al. (1997) on AOC from maize seeds. Therefore, the double bond at position 15 seems to be essential for the enantioselective cyclization by AOC2. This is in agreement with a proposal for the chemical cyclization reaction, whereby the 15(Z) double bond facilitates the opening of the oxirane ring of the allene oxide, and the resulting C-13 carbocation is stabilized by the electrons of the 15(Z) double bond (Grechkin, 1994).

Substrate Binding Site and Biochemical Analysis of Mutated AOC2 Protein
In 1990, a competitive inhibitor of the AOC of maize [(+/–)-cis-12,13-epoxy-9(Z)-octadecenoic acid = vernolic acid] was described (Hamberg and Fahlstadius, 1990). Vernolic acid is a naturally occurring fatty acid enriched especially in the seed oil (50 to 90% [w/w] of the total fatty acids) of several Asteraceae genera (Gunstone, 1954; Badami and Patil, 1980) and Euphorbiaceae species (Kleinman et al., 1965; Spitzer et al., 1996) and is a substrate analog of 12,13-EOT only missing the C11 double bond. We tested vernolic acid for its inhibiting effects on Arabidopsis AOC2. In our coupled enzymatic assay, total activity was downregulated by 50%, with some loss of stereoselectivity (Figure 6 ).

View larger version (17K): [in this window] [in a new window] [as a PowerPoint slide]   Figure 6. Enzymatic Activity of AOC2 Mutants and AOC2 with the Inhibitor Vernolic Acid. For determination of enzymatic activity, 5 µg of AOS and 10 µg of AOC2 were mixed in 1 mL of 10 mM PPi buffer, pH 7.0. After addition of 100 µg of HPOT as substrate, samples were incubated at room temperature for 15 min. Synthesized OPDA was extracted with ethyl acetate and analyzed via gas chromatography–mass spectrometry (GC-MS) as described in Methods. Shown are the results of the coupled enzymatic assay for AOS alone (1), AOS and AOC2 (2), AOS and AOC2 in presence of 160 µM inhibitor vernolic acid (3), and the mutants F85L (4) and F85A (5). The results are presented as the relative amount of synthesized OPDA compared with the yield with AOC2. The gray bars show total synthesized OPDA, whereas the black and white bars represent the yield of (+)- and (–)-enantiomer, respectively. Error bars represent SD based on three independent triplicate measurements (1, 2, 4, and 5) or a duplicate measurement (3).

View larger version (37K): [in this window] [in a new window] [as a PowerPoint slide]   Figure 7. Structure of AOC2 with Bound Inhibitor Vernolic Acid. (A) In the stereoview, monomer B is shown in gray ribbon representation, vernolic acid (VA) as blue stick model, and Water75 as red sphere. Residues directly involved in interactions with either the inhibitor or Water75 are depicted in yellow stick representation and labeled. Phe-85 is colored red. Omit difference density for vernolic acid (see Methods) at 2.4 is drawn as red chicken wire. The view is rotated 90° around the vertical with respect to Figure 2A. (B) Ligplot sketch showing the molecular interactions between the protein and the bound inhibitor vernolic acid. Contacts to the conserved Water75 are shown in green dashed lines with distances, and hydrophobic contacts are represented by opposing red spoked arcs. The interaction of Glu-23 is represented as a yellow arc. Vernolic acid is shown in purple.

	DISCUSSION

TOP ABSTRACT INTRODUCTION RESULTS DISCUSSION METHODS REFERENCES

 
Comparison with Other Arabidopsis AOC Structures
During the preparation of this manuscript, the Center for Eukaryotic Structural Genomics consortium (Madison, WI) deposited the coordinates of ligand-free AOC2 (CESG-AOC2, entry 1Z8K) and ligand-free AOC1 (CESG-AOC1, entry 1ZVC) to the Protein Data Bank (G.E. Wesenberg, G.N. Phillips Jr., E. Bitto, C.A. Bingman, and S.T.M. Allard, unpublished data). Although there is no publication yet on these structures, it is interesting to compare our results with the available coordinates.

CESG-AOC2 has been solved using SeMet-labeled protein and was refined at 1.7-Å resolution in the same orthorhombic space group as our SeMet-AOC2 structure. Both models superimpose very well with an RMSD of only 0.31 Å for 173 C atoms. A stereoview of the superposition is shown in Figure 8 . Closer inspection does not reveal any significant structural differences; specifically, the trimerization and the tightly bound water molecule inside the cavity are also found in the CESG-AOC2 structure.

View larger version (42K): [in this window] [in a new window] [as a PowerPoint slide]   Figure 8. Structures of AOC2, CESG-AOC2, and CESG-AOC1 Shown is a stereoview of the C trace of AOC2 (in gray), CESG-AOC2 (accession code 1Z8K; in red), and CESG-AOC1 (accession code 1ZVC; in blue). The 10 residues of AOC2 that differ in the AOC1 sequence are depicted in yellow stick representation.

None of the changes seem to be of potential relevance for the ligand binding site. Indeed, we found no significant differences in either total activity, stereo, or substrate specificity for all four AOC isoforms (AOC1-4) in our activity assays (P. Zerbe, V. Effenberger, and F. Schaller, unpublished data). This is in contrast with the observation of Stenzel et al. (2003b), who reported AOC2 having the highest activity but without quantifying this.

A differential expression pattern of AOCs is clearly established. Upon wounding and application of jasmonic acid, expression of AOC2 is upregulated to a higher degree than that of AOC1 (Stenzel et al., 2003b), whereas during senescence, a stronger upregulation of expression has been observed for AOC1 than for AOC2 (He et al., 2002). The other two isoforms, AOC3 and AOC4, seem not to be as strongly upregulated or sometimes even downregulated. The small structural differences, which mainly involve a change in the surface potential pattern between AOC1 and AOC2, might therefore represent a subtle optimization for interactions of these proteins with different partners either in different cellular locations or physiological states of the plant. For a complete understanding of these regulatory processes, more data on the expression profiles for all four enzymes in planta are necessary. Meanwhile, the structure elucidation of AOC3, whose crystallization has already been indicated by the CESG in the structural genomics target database (http://targetdb.pdb.org/), might shed some more light on the structural variations between the isoforms.

Structural Relation to the Lipocalin Family
While a database search with the PROSITE lipocalin signature motif results in 113 hits from plants, there are to our knowledge only four examples of plant lipocalins explicitly assigned in the literature: the small temperature-induced lipocalin TIL (Charron et al., 2002), the two xanthophyll cycle enzymes violaxanthin deepoxidase (VDE) and zeaxanthin epoxidase (ZEP) (Burgos et al., 1998), and the recently identified chloroplast lipocalin CHL (Charron et al., 2005). The last three proteins are located in the chloroplast. While all four proteins share homology in the structurally conserved regions, especially in SCR I with the invariant Gly and Trp present, homology in SCR II and III is much weaker, suggesting that these plant proteins would fall in the class of outlier lipocalins (Burgos et al., 1998). The other hits produced by the automated search are either false positive hits or the classification as lipocalins has not yet been supported by analysis of functional data.

As AOC2 can be assigned structurally to the lipocalin fold, this would allow placement in one of several protein families showing this general folding pattern or even setting AOC2 aside as a new structural motif on its own. Considering the observed topology of the barrel and functional similarities, we propose to consider AOC2 as a distant member of the lipocalin family in plants. The most relevant differences are discussed below.

The different placement of the termini in the superposition is striking (Figure 5). The barrel organization can be described right- or left-handed in the following way: if the thumb points upwards in the direction of the first ß-strand, the bent fingers of the respective hand should indicate the sequential organization of the successive ß-strands. By this definition, all lipocalin family members known so far (to our knowledge, this extends to all structures assigned to the lipocalin fold) form a left-handed barrel, while the AOC2 calyx is right-handed. In both cases, possible ligand binding sites are inside the palm and are accessible from above. As mentioned before, N- or C-terminal strand insertion with concomitant strand removal on the opposite end would automatically convert right- to left-handed barrels and vice versa.

Along these lines, the first ß-strand in AOC2 (blue in Figure 5) could have been formed by the insertion of the longer N-terminal part of the classical lipocalins (also colored blue) into the barrel. This would push ß-strand H of the lipocalins (shown in maroon) out of the continuous ß-sheet. The resulting differences in the structural organization would reduce the central role of the tryptophane in SCR I, and a loss of similarity in the SCRs would be obvious. Indeed, the observed weak similarity to SCR I in the second ß-strand could be indicative of such an early process.

This divergence in structure from the classical fold is not unexpected. Investigation of the exon-intron structure of lipocalins clearly places the plant lipocalins together with Dictostelium lipocalins into a gene structure–related group distinct from lipocalins from bacteria, insects, or animals (Sanchez et al., 2003). Since the early acquisition of the ancestor gene, it has been duplicated and adapted to different tasks. While TIL still can be counted as a genuine lipocalin, VDE evolved to be catalytically active in deepoxidation and acquired additions to the central barrel structure. With a size of 350 amino acids, VDEs are much larger than the typical lipocalin monomer, which consists of 160 to 180 amino acids (Flower, 1994; Flower et al., 1995; Burgos et al., 1998). The same is true for ZEP, for which threading methods don't even predict a barrel structure. Therefore, the peculiar architecture of these proteins raised doubt as to whether they truly belong to the lipocalin family (Ganfornina et al., 2000; Salier, 2000).

Up to now, not enough representatives of plant lipocalin genes have been identified to decide whether the clade of plant lipocalins does indeed show a much higher variability as the better-characterized lipocalins from bacteria, insects, or animals.

On the other hand, one cannot exclude the possibility that we in fact do observe an example of convergent evolution. This is an ongoing debate given the very low sequence homology among lipocalins and especially for the outlier lipocalins, for which only sequence similarity in SCR I is detectable anyway (Grzyb et al., 2006).

Oligomerization of AOC2
In former investigations, AOC2 has been characterized to form dimers based on size exclusion chromatography (Ziegler et al., 1997), but several lines of evidence point toward a trimer as the physiological relevant oligomerization state of the protein. First, we can detect trimers of recombinant AOC2 or AOC2 from plant extracts in SDS page or immunoblots. Secondly, we observe very large buried surface areas upon trimerization that include three salt bridges involving the terminal carboxy group and several strong hydrogen bonds. This is in agreement with the loss of trimer formation for the construct with the C-terminal His-tag, which would likely interfere with the formation of the salt bridges. Taken together, this pattern is indicative of a stable protein–protein interaction. Importantly, residues involved in these interactions are predominantly conserved in all known AOC sequences. Additionally, while both wild-type AOC2 and the CESG-AOC1 from Arabidopsis crystallize in different space groups and have therefore a modified crystal packing organization, in both cases the same mode of trimerization is observed.

AOC from maize has been shown to have an apparent molecular mass of 47 kD in size exclusion chromatography and was discussed as a dimer (Ziegler et al., 1997). We found a similar behavior for AOC2 of Arabidopsis (data not shown), but based on our additional structural and biochemical data, we attribute this discrepancy of 25% to the calculated molecular weight of a trimer to an atypical retardation of this protein on the size exclusion matrix.

We therefore expect the trimeric form to be the physiological relevant oligomerization state for AOCs of Arabidopsis and maize. Nevertheless, this SDS-stable trimer formation seems not to be absolutely required, since both the C- and N-terminally tagged proteins show similar enzymatic activities. However, trimer formation might improve overall protein stability.

Active Site and Reaction Mechanism
Based on the structural similarity of AOC2 to the lipocalins, we propose that the active site is located inside the barrel cavity. While most lipocalins just bind small hydrophobic substrates in the interior, very few are known to be enzymatically active. One example is noteworthy: human ß-trace or prostaglandinsynthase-D catalyzes the conversion of prostaglandin H2 to prostaglandin D2. As in the case of AOC2, the substrate is unstable in water and catalysis in a protected environment seems to be favorable. Here, the active site Cys is located in the barrel interior (Nagata et al., 1991).

Indeed, in AOC2 we find the competitive inhibitor vernolic acid to be bound inside the barrel of one monomer. We don't observe any induced fit mechanism, as the protein scaffold superposes with an RMSD of 0.33 Å for all 174 C atoms with SeMet-AOC2. Even on the level of side chains only marginal differences are present both in comparison to SeMet-AOC2 and to the other two monomers without bound inhibitor.

Surprisingly, the carboxylic end of the inhibitor does not seem to be tightly bound to the protein, as there is no clear electron density for the first five carbon units. This is in contrast with the observation of Ziegler and coworkers that methylation of this group abolishes the inhibitory effect for AOC from maize (Ziegler et al., 1997). One possible explanation is the exchange of Leu-27 of AOC2 to Arg in maize AOC, which would be positioned to form a strong salt bridge to the inhibitor in this protein. Another possibility would be that the carboxylic moiety is involved in the interaction with AOS for both proteins. Because we don't have data on the influence of methylation on inhibition of AOC2, a decision between both cases is not possible yet. Work is in progress to obtain comparable data on both proteins with the different forms of the inhibitor.

The finding that the location of the epoxy group in the 12,13-position of vernolic acid is essential for binding to AOC from maize (Ziegler et al., 1997) is consistent with the observed binding position in our structure. Any shift of the epoxy group would inhibit the formation of the hydrogen bond to the conserved water molecule.

To allow a discussion of possible reaction mechanisms, we tentatively modeled 12,13-EOT (substrate) and OPDA (product) molecules into the pocket in the same binding mode as vernolic acid (Figure 9A ). Based on the resulting arrangement of protein side chains around the bound substrate, we postulate the following reaction mechanism for the cyclization reaction inside the protein (Figure 10 ).

View larger version (48K): [in this window] [in a new window] [as a PowerPoint slide]   Figure 9. Substrate and Product Modeled into the Binding Pocket of AOC2 and Comparison with PGHS-1. (A) Shown are 12,13-EOT and OPDA as green and blue sticks, respectively. The protein is shown in a similar representation as in Figure 7. Also drawn is the molecular protein surface colored according to atom types. Possible hydrogen bonds between substrate and protein are shown as dashed green lines. (B) View into the cyclooxygenase active site of PGHS-1 from sheep (Ovis aris) (accession number 1DIY) with bound substrate arachidonic acid (AA). A similar representation is chosen as compared with (A). Also shown are the residues discussed in the text.

View larger version (17K): [in this window] [in a new window] [as a PowerPoint slide]   Figure 10. Schematic Overview of the Proposed Cyclization Reaction. The AOC2 catalyzed cyclization reaction involves the opening of the oxirane ring and a subsequent conrotary pericyclic ring closure (see text for details). The fatty acid moiety of 12,13-EOT [(CH2)7COOH] is labeled as R, and the schematic binding pocket of AOC2 is displayed in gray.

The proposed reaction scheme assumes as a final step a classical conrotary pericyclic ring closure, according to the Woodward-Hoffmann rules (Figure 10C). The absolute stereoselectivity of the reaction follows from the steric restrictions by the protein environment on the formation of the cis intermediate. Phe-85, Val-45, and Tyr-105 especially seem to form a greasy slide, which at the same time facilitates and restricts the conformational change of the hydrocarbon tail. Indeed, the F85L and F85A mutants show a reduced reactivity and stereoselectivity of the reaction. The latter effect would be explained by the removal of some of the steric restrictions on the trans-cis rotation around C10-C11. A rotation in the opposite direction would automatically result in the opposite stereoisomer of the product.

Our proposed mechanism explains the importance of the C15 double bond for reactivity along the lines of the proposal of Grechkin (1994) by the mechanism of anchimeric assistance. It also rationalizes the impact of the protein environment on the stereoselectivity of the cyclization reaction. It is clear by the packing in the cavity that changes in the position of the epoxide group will affect binding affinities of possible substrates.

A similar biosynthesis of cyclic hormones can be found in mammals during formation of prostanoids, which are structurally similar to JAs. One of the enzymes involved is the Lipocalin-type prostaglandin D synthase, which has been shown to be an example for an enzymatically active lipocalin (Urade and Hayaishi, 2000). The enzyme responsible for the first committed step in prostanoid catalysis from the linear precursor is prostaglandin endoperoxide synthase (PGHS). Due to its medical relevance as target for anti-inflammatory drugs like aspirin, it has been very well characterized. PGHS catalyzes both the cyclization of arachidonic acid to prostaglandin G2 and the successive peroxidation of PGG2 to prostaglandin H2. While PGHS shows a completely different overall architecture than AOC, the functional homology makes a comparison of the active sites worthwhile.

PGHS is a dimeric monotopic membrane protein with a molecular mass of 70 kD (monomer), which is located in the endoplasmic reticulum and nuclear envelopes. Besides an N-terminal EGF domain, the protein is mainly -helical and has two distinct active sites for the two catalyzed reaction steps (Picot et al., 1994; for a review, see Garavito et al., 2002). In the cyclooxygenase active site, arachidonic acid binds with the carboxylic end, forming salt bridges to a conserved Arg residue. The rest of the molecule is tightly packed into an L-shaped cavity lined mainly with hydrophobic and aromatic residues, as found in the AOC2 binding pocket (Figure 9B). The catalytically active Tyr is positioned at the kink of the cavity for abstraction of the 13proS hydrogen, the initiating step of the reaction. Interestingly, the only other polar residue is situated on the opposite side of the polycarbon chain, similar to the position of Cys-71 in the case of AOC2. This has been shown to be critical in determining the stereochemistry of the product (Schneider et al., 2002). It has been found that the packing at the -end of the substrate molecule is very important for maintaining the catalytic selectivity and activity (Rowlinson et al., 1999; Malkowski et al., 2001). While increasing the size of residues in this region can abolish activity due to steric inhibition of binding, mutations increasing the volume of the binding pocket mostly diminish activity, as optimal positioning of substrate is not enforced. It has been proposed that the carboxylic end stays fixed during catalysis in PGHS, while the -end has to move closer to allow for the substantial conformational changes necessary for ring closure (Garavito et al., 2002). In the case of AOC2 from Arabidopsis, a movement of the carboxyl end of the substrate along our proposed reaction scheme seems to be more likely. While the relative flexibility of the carboxy group of vernolic acid might not represent the situation for the substrate 12,13-EOT in planta, there are no positively charged residues appropriately placed to form such a strong salt linkage as observed in the case of PGHS.

Here, we find two completely different structures showing the same binding principles. A hydrophobic mold binds the substrate exactly in the right position to a single catalytically critical polar site without apparent induced fit during binding. While the exact reaction schemes are very different, the hydrophobic mold in both cases also ensures the stereospecificity of the reaction.

As the comparison with PGHS shows, residues in the vicinity of the substrate binding site are probably highly optimized to control the reaction in the cavity. Structural and functional analyses of additional point mutants will help to clarify the individual influence of the key amino acids in the binding pocket on the proposed mechanism and the stereoselectivity of the reaction.

	METHODS

TOP ABSTRACT INTRODUCTION RESULTS DISCUSSION METHODS REFERENCES

 
Protein Expression and Purification
A truncated version of AOC2 from Arabidopsis thaliana has been cloned into the vector pQE30 (Qiagen) and pET21b (Novagen/Merck) to yield a fusion protein (20.2 kD) in which the first 77 amino acids (the predicted transit signal) have been replaced by a His6-tag (sequence MRGSHHHHHHRS) or in which the His6-tag has been coupled with the C terminus of the truncated protein. The resulting constructs were transformed into Escherichia coli strain M15. Cells were grown in 2YT media and induced at an OD of 0.5 to 0.6 with addition of 0.2 mM isopropylthio-ß-galactoside and 0.2% arabinose, respectively. E. coli with the pQE30 construct were harvested after an induction time of 5 h at 37°C and 220 rpm. Bacteria with the pET21b construct were induced for a further 30 to 40 h at 25°C. Cells were broken by ultrasonication, and the fusion protein was purified by affinity chromatography (Ni-NTA; Qiagen) and concentrated with Centricon concentrators (5000 D cutoff; Millipore). The yield of purified protein exceeded 10 mg/L culture.

SeMet-labeled N-terminally His6-tagged AOC2 was produced by growth of M15 cells in M9 minimal medium supplemented with SeMet as described (Van Duyne et al., 1993). The mutated proteins of AOC2 were expressed in the pQE vector under the same conditions as the native AOC2.

AOS (pQE30-AOS; Laudert et al., 1996) was expressed and purified in accordance with Oh and Murofushi (2002). Protein expression was induced at an OD₆₀₀ of 0.5 by addition of isopropylthio-ß-galactoside to a final concentration of 0.4 mM and further incubation at 16°C and 150 rpm overnight. Cells were broken by ultrasonication, and recombinant AOS was purified via Ni-NTA agarose chromatography (Qiagen) by adding Triton X-100 up to a final concentration of 0.1% (v/v) to the recommended buffers.

Generation of Mutant Proteins of AOC2
The generation of AOC2 mutants was done via PCR according to the sequential PCR step method of the current protocols in molecular biology. Phe at position 85 was exchanged to Ala (primers: 5'-GAAAGGTGAAAGAGCCGAAGCTACTTATA-3'and 5'-TATAAGTAGCTTCGGCTCTTTCACCTTTC-3') and Leu (primers: 5'-GAAAGGTGAAAGACTCGAAGCTACTTATA-3'and 5'-TATAAGTAGCTTCGAGTCTTTCACCTTTC-3'). The N- and C-terminal primers were 5'-TATGGATCCCCAAGCAAAGTTCAAGAACTG-3' (BamHI restriction site underlined) and 5'-TATGTCGACTTAGTTGGTATAGTTACTTATAAC-3' (SalI restriction site underlined). The mutated cDNA was cloned afterwards into the expression vector pQE30, and the protein was expressed under standard conditions.

Gel Electrophoresis and Protein Immunoblotting
Denaturing gel electrophoresis was performed according to Laemmli (1970). The discontinuous systems consisted of 4% stacking gels and 12.5% resolving gels. Protein blotting onto nitrocellulose was done electrophoretically overnight (4°C, 64 mA) as described by Towbin et al. (1979). Immunodetection followed standard procedures (Parets-Soler et al., 1990) with goat anti-rabbit immunoglobin-conjugated alkaline phosphatase and the enzyme substrates 4-nitrotetrazolium blue and 5-bromo-4-chloro-3-indolyl phosphate.

Determination of Total Enzymatic Activity via Chiral Capillary GC-MS
For activity analysis, 5 to 20 µg of purified AOS and AOCs (weight-based ratio 1:2) were added to 1 mL of 10 mM PP_i buffer, pH 7.0. The reaction was started by addition of 100 µg of HPOT. After incubation for 15 min at room temperature, the reaction was stopped by acidifying with 1 M HCl to pH 3.0. The synthesized OPDA was extracted twice with 2 volumes of ethyl acetate and taken to dryness. For GC-MS analysis, 1 nmol of ²[H]₅-OPDA was added before extraction as an internal standard. Dried samples were incubated in 250 µL of 0.1 M KOH for 45 min at room temperature to convert the synthesized OPDA into trans-configuration. After a second extraction step, OPDA was dissolved in methanol and methylated with ethereal diazomethane (Hamberg and Fahlstadius, 1990). The dried fractions were then redissolved in 50 to 100 µL of chloroform, and 1 µL of the sample was injected into a Varian GC 3400 gas chromatograph in splitless mode in direct connection to a MAT Magnum ion trap mass spectrometer (Finnigan) using a chemical ionization mode with methanol as reactant gas. Separations of the (+)- and the (–)-enantiomer of OPDA were made on a ß-Dex120 column (30 m x 0.25 mm x 0.15 µm stationary-phase thickness) coated with 20% permethyl-ß-cyclodextrin in SPB-35 (Supelco) with He carrier gas at 1 mL/min (gas prepressure of 80 kPa) using the following temperature program: injector temperature of 220°C, 1 min isothermally at 50°C, with 10°C/min up to 190°C, 75 min isothermally at 190°C, with 3°C/min up to 220°C, 15 min isothermally at 220°C, and transfer line temperature 220°C. For details, see Laudert et al. (1997) and Schaller et al. (1998). Quantitation of AOC enzymatic activity was based on (+)-OPDA-to-(–)-OPDA ratios derived from integration of total ion current traces.

Crystallization
Initial crystallization screening for AOC2 was performed using vapor diffusion against the Wizard I and II kits from Emerald BioSystems. Protein crystals were found with condition WI-28, which was subsequently optimized. Using 14 to 16% PEG-8000, 6% tert-butanol, and 0.1 M MES, pH 5.0 to 6.0, as reservoir solution in either hanging- or sitting-drop setups (4 mg/mL protein concentration), we were able to grow crystals (space group P2₁) up to a size of 250 x 150 x 50 µm, which diffract to 1.7-Å resolution.

AOC2-SeMet crystals (space group P2₁2₁2₁) were grown from reservoir solutions containing 10% PEG-3350, 200 mM NaCl, and 100 mM phosphate-citrate, pH 4.2, with a protein concentration of 13.5 mg/mL. They typically reach a size of 250 x 200 x 100 µm and diffract to better than 1.3-Å resolution. Crystals grew at 18°C in 1 week.

For soaking experiments, AOC2-SeMet crystals were transferred for 1 to 2 d into appropriate mother liquor saturated with inhibitor.

X-Ray Diffraction Data Collection
Data collection was performed at 100K on flash-frozen crystals at the Swiss Light Source (Villigen, Switzerland) at beamline PXII using a MAR CCD detector. Cryoprotection was achieved by briefly soaking the crystals in paraffin oil or in mother liquor supplemented with glycerol. For SeMet-AOC2 crystals, MAD data at the Se edge were collected at the European Synchroton Radiation Facility (Grenoble, France) on beamline ID13.1 at 100K using a MAR CCD detector. Data of inhibitor-soaked SeMet-AOC2 crystals were collected in house at 100K on a Bruker FR591 rotating anode with Montel multilayer optics and a MAR345dtb detector system.

Data were processed using the XDS package (Kabsch, 1993). Statistics of the different data sets are shown in Table 1.

Structure Solution and Refinement
SeMet-AOC2
Self-rotation functions indicated the presence of a threefold noncrystallographic symmetry. Search for the Se substructure and initial phasing were done with the programs of the SHELX package (Sheldrick and Schneider, 1997) using the hkl2map interface (Pape and Schneider, 2004) with data to 2-Å resolution. SHELXD identified four major peaks, two of which in retrospect correspond to two copies of SeMet42. The other two are a double peak corresponding to the third copy of SeMet42. This solution showed correlation coefficients of 41 for all data and 32 for weak data. Phasing with SHELXE using data to 3 Å clearly identified the correct hand and already resulted in a well-interpretable map. Fifty-eight cycles of density modification using an estimated solvent content of 0.4 resulted in density contrast of 0.73 (0.38), connectivity of 0.95 (0.91), and a pseudo-free correlation coefficient of 83.2 (63.3) (values for the wrong hand in parentheses).

Density modification (using the threefold symmetry), phase extension, and auto building in RESOLVE (Terwilliger, 2000) could assign 387 of the expected 564 (3 x 188) residues, including side chains. An additional 44 residues were placed without side chains.

For refinement, the peak wavelength data set was used. Five percent of the data were randomly assigned to the R_free test set and omitted from refinement.

The resulting model was further improved in several cycles of refinement, automated water placement, and manual rebuilding using programs of the CCP4 package (Collaborative Computational Project, Number 4, 1994) and ARP/wARP (Cohen et al., 2004).

The final model contains three protein monomers composed of residues 15 to 188. The first 14 residues, including the His6-tag, could not be placed due to missing density. Based on clear density, a total of 19 residues had to be modeled with alternate conformations. Additionally, three glycerol and 556 water molecules were included. Part of the experimental electron density map after RESOLVE with the final model is shown in Supplemental Figure 2 online.

WT-AOC2
The structure of WT-AOC2 was solved using the refined coordinates of SeMet-AOC2 after removal of solvent and alternate conformations and the mutation of SeMet to Met as a search model. Two trimers were found in the asymmetric unit of the monoclinic unit cell. After several rounds of refinement in REFMAC (Murshudov et al., 1997) and manual rebuilding and water picking in COOT (Emsley and Cowtan, 2004), the final model consists of 1036 amino acids, five glycerol molecules, two sodium ions, and 660 water molecules. Gln-102 had to be modeled in alternate conformations in five of the six copies of AOC2 in the asymmetric unit. Weak NCS restraints were used throughout the refinement.

SeMet-AOC2 Inhibitor
For analysis of the inhibitor data, one copy of the 2BRJ model without alternate conformations and solvent molecules was placed in the unit cell. After several rounds of refinement and water picking, extended residual density in monomer B was fitted with a model of the inhibitor vernolic acid. The initial atomic coordinates and the necessary geometry definitions were created using the PRODRG server (Schuettelkopf and van Aalten, 2004). At the end of the refinement, the occupancy of the inhibitor was set to 0.25, 0.5, 0.75, and 1 and the structure refined using identical protocols. With full occupancy, the best R-factors were reached. While we observed clear density for the majority of the inhibitor, the carboxylic end up to C5 did not show clear density due to disorder (Figure 7A). The shape of the resulting difference density together with the higher flexibility of the inhibitor in comparison to the surrounding protein atoms also points to different minor binding modes of the inhibitor (possibly also the second enantiomer), which were not modeled.

Elongated residual density in monomer C was judged too weak to safely allow positioning of the inhibitor. In monomer A, the entrance to the binding site is completely blocked by Leu-41 from a crystallographically related molecule. Therefore, binding of inhibitor during soaking in this monomer is prohibited. Weak NCS restraints were used in the initial stages of the refinement but were released after inclusion of the inhibitor.

For preparation of the omit maps, vernolic acid was removed from the final model and random positional shifts (average RMSD 0.12 Å) were applied to the remaining coordinates to remove model bias. After another round of refinement with REFMAC, an mFo-DFc difference omit map was calculated.

Figure Preparation
Figures were generated using Pymol (http://www.pymol.org). To allow coloring of residues by conservation, an alignment of 22 different AOCs found with a National Center for Biotechnology Information BLAST search in the UNIREF100 database was analyzed using the ESPript server (Gouet et al., 1999). In the resulting coordinate file, the similarity score has been assigned to the B-factor column, ranging from 100 (fully conserved) to 0 (no conservation). This range was assigned to the color range red to blue for figure generation.

Superposition of molecules was either performed in COOT (Emsley and Cowtan, 2004) or using the SuperPose server (Maiti et al., 2004).

Accession Numbers
Coordinates and structure factors can be found in the Protein Data Bank under accession numbers 2BRJ, 2GIN, and 2DIO for the AOC2-SeMet, WT-AOC2, and AOC2-SeMet inhibitor structure, respectively.

Supplemental Data
The following materials are available in the online version of this article.

Supplemental Figure 1. Conservation of the Trimer Interface.

Supplemental Figure 2. MAD-Solvent-Flattened Electron Density of AOC2.

	Acknowledgments

 
We thank C. Kötting for helpful discussions, K. Diederichs and D.R. Madden for comments on the manuscript, and E.W. Weiler for continuing support and interest in the work. Part of this work was performed at the Swiss Light Source, Beamline X10SA, Paul Scherrer Institute (Villigen, Switzerland) and at the European Synchroton Radiation Facility, Beamline ID13.1 (Grenoble, France). We would especially like to acknowledge the help of the respective beamline staff and of our colleagues from the Max Planck Institutes for Medical Research (Heidelberg) and Molecular Physiology (Dortmund) during data collection. We also acknowledge financial support from the Deutsche Forschungsgemeinschaft by Grants SFB480-C6 (E.H.), HO2600 (E.H.), and SCHA939 (F.S.).

	Footnotes

 
¹ These authors contributed equally to this work.

The authors responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantcell.org) are: Eckhard Hofmann (eckhard.hofmann{at}bph.rub.de) and Florian Schaller (florian.schaller{at}rub.de).

^[W] Online version contains Web-only data.

www.plantcell.org/cgi/doi/10.1105/tpc.106.043984

Received May 10, 2006; Revision received August 4, 2006. accepted October 17, 2006.

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