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Biochemical and Structural Analysis of Substrate Promiscuity in Plant Mg2+-Dependent O-Methyltransferases
J. Mol. Biol. (2008) 378, 154–164
doi:10.1016/j.jmb.2008.02.019
Available online at www.sciencedirect.com
Biochemical and Structural Analysis of Substrate Promiscuity in Plant Mg 2+ -Dependent O-Methyltransferases Jakub G. Kopycki 1 †, Daniel Rauh 1 †, Alexander A. Chumanevich 1 , Piotr Neumann 1 , Thomas Vogt 2 ⁎ and Milton T. Stubbs 1,3 ⁎ 1
Institut für Biotechnologie, Martin-Luther-Universität Halle-Wittenberg, Kurt-MothesStraße 3, D-06120 Halle (Saale), Germany 2
Leibniz-Institute of Plant Biochemistry, Department of Secondary Metabolism, Weinberg 3, D-06120 Halle (Saale), Germany 3
Mitteldeutsches Zentrum für Struktur und Dynamik der Proteine, Kurt-Mothes-Straße 3, D-06120 Halle (Saale), Germany
Plant S-adenosyl-L-methionine-dependent class I natural product Omethyltransferases (OMTs), related to animal catechol OMTs, are dependent on bivalent cations and strictly specific for the meta position of aromatic vicinal dihydroxy groups. While the primary activity of these class I enzymes is methylation of caffeoyl coenzyme A OMTs, a distinct subset is able to methylate a wider range of substrates, characterized by the promiscuous phenylpropanoid and flavonoid OMT. The observed broad substrate specificity resides in two regions: the N-terminus and a variable insertion loop near the C-terminus, which displays the lowest degree of sequence conservation between the two subfamilies. Structural and biochemical data, based on site-directed mutagenesis and domain exchange between the two enzyme types, present evidence that only small topological changes among otherwise highly conserved 3-D structures are sufficient to differentiate between an enzymatic generalist and an enzymatic specialist in plant natural product methylation. © 2008 Elsevier Ltd. All rights reserved.
Received 12 November 2007; received in revised form 5 February 2008; accepted 7 February 2008 Available online 20 February 2008 Edited by M. Guss
Keywords: crystal structure; substrate specificity; hybrid protein; loop exchange; plant secondary metabolism
Introduction
*Corresponding authors. M. T. Stubbs is to be contacted at Institut für Biotechnologie, Martin-Luther-Universität Halle-Wittenberg, Kurt-Mothes-Straße 3, D-06120 Halle (Saale), Germany. E-mail addresses: [email protected]; [email protected]. † J.G.K. and D.R. contributed equally to this work. Present address: D. Rauh, Chemical Genomics Centre of the Max Planck Society, Otto-Hahn-Straße 15, D-44227 Dortmund, Germany. Abbreviations used: OMT, O-methyltransferase; PFOMT, phenylpropanoid and flavonoid OMT; CoA, coenzyme A; CCoA, caffeoyl CoA; rOMT, rat catechol OMT; SAM, S-adenosyl-L-methionine; SAH, S-adenosyl-Lhomocysteine; SeMet, selenomethionine.
Methylation by S-adenosyl-L-methionine (SAM)dependent O-methyltransferases (OMTs) (EC 2.1.1) is a common modification in natural product biosynthesis. Site-specific O-methylation modulates the physiological properties and the chemical reactivity of phenolic hydroxy groups.1 Class I plant cationdependent OMTs (also known as caffeoyl coenzyme A OMTs or CCoAOMTs) constitute a group of low molecular mass (23–27 kDa) enzymes that play an important role in methylation of guaiacyl residues present in CCoA or 5-hydroxy feruloyl CoA, both precursors of monolignols.2 In angiosperms, members of a second class of metal-independent OMTs (caffeic acid OMTs) are able to methylate the monolignol precursors caffeic acid, caffeyl aldehyde, or caffeyl alcohol in the meta position.3–5 The action of
0022-2836/$ - see front matter © 2008 Elsevier Ltd. All rights reserved.
Substrate Promiscuity in Plant OMTs
both enzyme classes in a metabolic grid appears to be crucial for determining the structural integrity of the lignin polymer.6,7 Detailed structural information on the cationindependent enzymes is available, including the chalcone and isoflavone OMTs from alfalfa (Medicago sativa), a flavonoid OMT from Arabidopsis thaliana, as well as an OMT involved in methyl ester formation in Clarkia breweri.8–11 These enzymes are obligatory dimers, with each active site formed from residues of both monomeric polypeptide chains. Either general acid/base catalysis (in the case of phenolic ether formation) or proper substrate positioning (in the case of the methyl ester formation) is essential for efficient methyl transfer in the individual systems. Although a major goal of these investigations on the class II enzymes has been the tailored modification of methylation specificity, which would enable the engineering of biochemical pathways for the production of transgenic plants with altered lignin composition or alternative accumulation of diverse natural products, mutagenesis of individual amino acids has until now only resulted in lower substrate specificity.10 In contrast, less structural information is available for cation-dependent OMTs. The 2.0-Å-resolution structure of rat catechol OMT (rOMT)12 exhibits a typical Rossmann fold within the coenzyme-binding domain of the enzyme, suggesting that SAM methylases share a common structural motif.13 Methyl
155
Fig. 1. Structures of OMT substrates methylated by promiscuous CCoAOMT-like enzymes such as PFOMT from the ice plant M. crystallinum.18 1, quercetin; 2, quercetagetin; 3, caffeic acid; 4, CCoA; 5, caffeoyl glucose. The positions of the methyl group acceptor hydroxyl groups are marked; note the two methylation positions for quercetagetin.
transfer proceeds via an Sn2-like transition state, with one ionized hydroxyl group, supported by the Mg2+ ion, and a conserved lysine as a catalytic base, to attack the electron-deficient methyl group of SAM.14 In vertebrates, the enzyme is responsible for the inactivation of neurotransmitters such as dopamine in the brain and has therefore received particular attention as an attractive target to cure degenerate brain diseases.13,15 However, high concentrations of
Fig. 2. Overall structure of the PFOMT dimer in cartoon representation. Monomer A is in yellow, with the N-terminus in blue and the ‘insertion loop’ in orange; the secondary structure elements of PFOMT are shown for monomer B, which is color ramped from the N-terminus (blue) to the C-terminus (red). The positions of the active sites are indicated by the bound calcium ion (gray sphere) and the cofactor SAH (stick representation).
156 catechol OMT are also found in the liver, kidney, or placenta, where the enzyme inactivates potentially reactive or mutagenic phenolics such as quercetin.1 Recently, the 3-D structure of the CCoA-specific CCoAOMT from M. sativa was solved at a resolution of 2.4 Å.16 This enzyme, which almost exclusively methylates the CoA ester of caffeic acid, displays very low affinity for caffeic acid itself, with a direct impact on lignin polymer biosynthesis in gymnoand angiosperms. 7 The structure of the alfalfa CCoAOMT provides insights into the mode of action and specificity of this class of enzymes. While the overall fold and the active site of the CCoAOMT dimer resemble those of the animal catechol OMT, significant differences in the positioning of the Nterminus and intervening loops in the vicinity of the active site contribute to specificity for CCoA. In addition to the CCoA-specific enzymes, a second type of cation-dependent OMT from the ice plant Mesembryanthemum crystallinum [phenylpropanoid and flavonoid OMT (PFOMT)] with promiscuous substrate specificity has been described.17 Phylogenetic analyses indicate that plant cation-dependent
Substrate Promiscuity in Plant OMTs
OMTs can be divided into two subclades. The CCoAspecific cluster responsible for lignin biosynthesis exhibits a high sequence identity of over 80% at the amino acid level. The more divergent subcluster epitomized by PFOMT is found in several plant species as multiple sequences, which may be differentiated by substrate specificity or by differentially regulated expression (T.V., unpublished results). Functionally expressed in Escherichia coli, the promiscuous OMTs methylate a variety of natural products with vicinal dihydroxy groups in vitro that are not accepted by the lignin-specific enzyme, including caffeic acid and caffeoyl glucose17,18 (Fig. 1). In the ice plant, it has been established that the PFOMTcatalyzed methylation of flavonoids is induced upon ultraviolet irradiation to protect the plant against further damage.17 Further studies have shown that substrate specificity can be influenced by the nature of the cation,19 while the N-terminus of PFOMT (which, in the plant, is some 11 residues shorter than that derived from the gene sequence) plays a crucial role in determining the position specificity and kinetic properties of the enzyme.18
Fig. 3. Stereo view of the superposition of PFOMT (A monomer, colored as in Fig. 2) with (a) M. sativa CCoAOMT16 (green) and (b) rOMT12 (magenta). Coordinates of the cofactor SAH (red sticks) are from the present PFOMT structure, and positions of the bound cations are depicted by the colored spheres. CCoAOMT product 5-hydroxy feruloyl CoA (green) and rOMT inhibitor dinitrocatechol (magenta) from the respective structures are depicted as sticks. Note the difference in the ‘insertion loop,’ which supports the CoA moiety in the CCoAOMT structure, as well as the shorter loop between β6 and β7 and the alternative N-terminal arrangement of secondary structure elements in the plant enzymes compared to rOMT, which allows dimerization of the former.
Substrate Promiscuity in Plant OMTs
157
Here, we report the crystallization and 3-D structure determination of the cation-dependent PFOMT from the ice plant with promiscuous specificity at 1.4 Å resolution. Catalytic properties of N-terminally truncated variants and hybrids containing loops from the CCoA-specific alfalfa OMT demonstrate their significant influence on substrate specificity. Simultaneous replacement of two regions from the alfalfa into the ice plant enzyme converts the promiscuous enzyme to a CCoA-specific OMT, demonstrating the important contribution of substrate binding by the N-terminus and the loop domain.
Results and Discussion PFOMT is a symmetric dimer of approximate dimensions 70 × 40 × 50 Å3 (Fig. 2). As expected, each monomer exhibits an α/β fold similar to the monomeric animal rOMT,13 with eight α-helices and seven β-strands, including the nucleotide-binding motif or Rossmann fold. In contrast to rOMT but in common with CCoAOMT,16 however, the first two helices of PFOMT show a distinctly different topological arrangement. The position of helix α1 in the plant enzymes is in part accomplished by a deletion between strands β6 and β7 with respect to the mammalian enzyme and contributes significantly to dimer formation (Fig. 3). Despite cocrystallization with a methyl group donor SAM and the substrate quercetin, only the resultant S-adenosyl-L-homocysteine (SAH) could be detected in the active center (Fig. 4a). Although no electron density could be observed for the catalytically important N-terminal residues, the positioning of the first visible amino acids indicates that the N-termini would be well placed to influence catalytic activity. The two active sites, demarcated by the presence of SAH and bound Ca2+ ions (Fig. 2), could act independently of one another; it is conceivable, however, that a mobile amino-terminus of one monomer might communicate with the catalytic apparatus of the second monomer, as has been observed recently for a dimeric bacterial OMT from Leptospira interrogans of unknown specificity.20 This is in contrast to the plant cation-independent OMTs, whose active site consists of residues from both monomers of the dimer, leading to the strong influence of catalytic properties of the corresponding second subunit.8 The catalytic apparatus and residues in the immediate vicinity are almost identical in each of the three metal-dependent enzymes, the only significant difference here being the replacement of rOMT Trp143, which partially covers the SAM moiety and forms a lower boundary to the substrate binding pocket, by Asp165. All attempts at soaking crystals or cocrystallization with a variety of substrates failed to yield any electron density to establish the binding modes within the active site of PFOMT (data not shown). Nevertheless, superposition of the active sites of PFOMT with ligand-bound structures of CCoAOMT and rOMT allows modeling of the binding of quercetin and quercetagetin, whose
Fig. 4. The active site of PFOMT. (a) Representative electron density (2Fo − Fc, contoured at 0.9σ) showing the cofactor SAH and the bound cation (gray sphere), interpreted as Ca2+. (b) Docking of the substrate quercetagetin in the PFOMT active site. Quercetagetin is methylated in two distinct positions (see Fig. 1), requiring opposing orientations of the substrate (yellow and pink). Both orientations make use of aromatic stacking interactions between the substrate and the side chains of Tyr51, Trp184, and Phe198 from the ‘insertion loop,’ positioning the pertinent hydroxyl groups in the vicinity of the SAM methyl donor and the active-site Mg2+ ion (gray sphere).
extended aromatic moieties would be sandwiched between the side chains of Tyr51, Trp184, and Phe198 (Fig. 4b). In silico docking experiments were carried out to obtain semiquantitative data for the relative specificities of the two plant enzymes (Table 1). Despite the absence of structural data for the amino-terminal residues for both proteins, the results are in general agreement with the observed specificities. For quercetin, a substrate-like orientation of the flavonoid could be determined in the active center of PFOMT with reasonable binding energies, whereas no enzy-
Substrate Promiscuity in Plant OMTs
158
Table 1. Results of docking of quercetin, caffeic acid, and feruloyl CoA into the active sites of PFOMT and CCoAOMT PFOMT Cluster rank
CCoAOMT
Lowest binding energy
Mean binding energy
Number in cluster
Cluster rank
Lowest binding energy
Mean binding energy
Number in cluster
Quercetin 1 2 3 4 5 5.1 6 7 8 9 10 11 12 13 14 15 16
− 7.93 − 7.51 − 7.00 − 6.93 − 6.84 − 6.84 − 6.83 − 6.75 − 6.68 − 6.61 − 6.54 − 6.52 − 6.49 − 6.38 − 6.34 − 6.17 − 5.82
− 6.81 − 6.72 − 6.56 − 6.32 − 6.49
53 41 23 15 13
− 6.45 − 6.31 − 6.37 − 6.37 − 6.08 − 6.25 − 6.18 − 6.32 − 6.34 − 5.80 − 5.78
8 8 14 5 10 44 7 3 1 3 2
1 2 3 4 5 6 7 8 · · · · 18 19 20 21 n.d.
− 8.18 − 7.94 − 7.73 − 7.73 − 7.48 − 7.21 − 7.17 − 7.16
−7.44 −7.47 −7.31 −7.03 −7.10 −6.92 −6.82 −7.16
− 6.40 − 6.37 − 6.35 − 6.02
−6.36 −6.37 −6.35 −6.02
67 63 16 32 25 3 15 1 · · · · 2 1 1 1
Caffeic acid 1 1.1 2 3 4 5 6 7 8 9
− 5.44 − 5.44 − 4.66 − 4.41 − 4.33 − 4.18 − 4.18 − 4.17 − 4.09 − 3.94
− 4.92
195
− 4.21 − 4.31 − 4.21 − 4.07 − 4.18 − 4.17 − 3.98 − 3.77
32 5 3 2 1 1 5 6
1 2 3 4 5 5.1 · · · 19
− 4.53 − 4.50 − 4.47 − 4.45 − 4.43 − 4.43
−4.23 −4.20 −4.22 −4.31 −4.09
16 13 11 3 83
+ 0.86
31
+ 1.22
20 · · · 2 1 1 1 1
5OH-feruloyl CoA 1 1.1 2 . . . 55 56 57 58 59
− 0.35 − 0.35 + 0.42
+ 2.65 + 2.92 + 2.96 + 3.08 + 3.25
+ 2.69 + 2.92 + 2.96 + 3.08 + 3.25
1 1.1 2 · · · 11 12 13 14 15
− 3.63
−3.63
− 2.85 − 2.85 − 2.64
−2.17
− 1.90 − 1.90 − 1.25 − 1.23 + 0.66
· · · 1 144
−2.16
−1.68 −1.90 −1.19 −1.23 +0.66
3 · · · 5 1 2 1 1
The highest ranking substrate-like binding modes are indicated in boldface, together with the corresponding cluster and binding energy (kcal mol− 1). n.d., not detected.
matically reasonable binding mode could be found for CCoAOMT. This was also the case for quercetagetin (data not shown). Caffeic acid and feruloyl CoA could be docked into both active sites with reasonable energies, although the in silico results clearly distinguish between the two enzymes. Caffeic acid docks to PFOMT favorably in terms of binding energy, cluster ranking, and cluster size; the results for CCoAOMT are less conclusive. This situation is reversed for the docking of feruloyl CoA, where only a small fraction of the binding poses for PFOMT show favorable (negative) energies. The most notable difference between the plant and animal enzymes is a large insertion loop between strand β5 and helix α8 (Fig. 3). The conformation of this insertion, which begins with a short α-helical turn, is supported by the N-terminal helix of the opposing monomer, so that the active-site cleft
of the dimeric plant enzymes appears deeper and more restricted (Fig. 4). This insertion loop provides a scaffold for the CoA moiety of CCoA in the CCoAOMT structure.16 Most interestingly, this loop in PFOMT exhibits a conformation markedly different from that in CCoAOMT (Fig. 3). Moreover, sequence analysis reveals that this region demonstrates the lowest sequence homology between CCoA-specific and promiscuous enzymes (30% identity for residues 185– 208, Fig. 5). Thus, it seems plausible that differences in this loop and in the N-termini of the two enzyme classes (which also exhibit low sequence conservation and are observed neither in the present structure nor in that of CCoAOMT16) could provide a clue to the promiscuous nature of PFOMT. Hybrid recombinant proteins were designed and expressed in E. coli and their substrate specificity was
Substrate Promiscuity in Plant OMTs
159
Fig. 5. Sequence alignment of promiscuous OMTs (from M. crystallinum and Stellaria longipes) and diverse, specific CCoAOMTs, together with the mammalian rOMT. Secondary structure elements of PFOMT are shown, colored as in Fig. 2 (monomer B). Boxed are the variable N-terminal residues (in blue) and the ‘insertion loop’ (in orange), which shows structural and sequence divergence between the promiscuous and specific enzymes.
characterized to investigate the influence of these two regions on substrate preference (Fig. 6). We have previously shown that the N-terminus of PFOMT plays a crucial role in determining the position specificity and kinetic properties of the enzyme.18 A new construct was made with a cleavable His tag in order to study the influence of the N-terminal His tag. Removal of the His tag from native PFOMT (ΔHisPFOMT, Table 2) resulted in a significant loss of catalytic efficiency toward the substrates caffeic acid and CCoA, largely due to a reduction in Km app, while turnover of the flavonoids was largely unaffected. In contrast, replacement of the PFOMT N-terminus with that of the specific CCoAOMT enzyme from alfalfa (N-term hybrid) resulted in a reduced catalytic efficiency toward flavonoids, an effect that is even more pronounced for the substrates caffeic acid, CCoA, and caffeoyl glucose. In all cases, a 10- to 100-fold increase in Km app reflects a reduction in substrate affinity, which is the major factor contributing to the strongly reduced catalytic efficiency kcat/Km app. As with PFOMT, removal of the His tag from the N-terminal hybrid (ΔHis–Nterm hybrid) resulted in a strong reduction in activity toward caffeic acid and CCoA; interestingly, the ΔHis–N-term hybrid exhibited an increased activity
toward quercetagetin, such that this enzyme has an almost exclusive flavonoid activity. Substitution of the insertion loop from the specific alfalfa CCoAOMT in the nonspecific PFOMT (loop hybrid) leads to a stark reduction in soluble protein expression (1 mg/ml instead of the usual 10 mg/ ml), with most of the protein depositing as insoluble inclusion bodies (data not shown). The loop hybrid exhibited a 10-fold increase in affinity for and activity against CCoA and a marked reduction in affinity for the flavonoids; that is, the loop-hybrid protein behaves more like CCoAOMT than PFOMT. On the other hand, replacement of the loop resulted in little change in Km app for caffeoyl glucose, although a considerable loss in activity against this substrate was observed. Despite the strongly reduced Km app for CCoA, the high affinity for caffeoyl glucose continues to distinguish the loop hybrid from the corresponding alfalfa protein. Variants in which both the N-terminal peptide and the insertion loop of PFOMT were replaced simultaneously by those of the corresponding CCoAOMT sequence (double hybrid) failed to increase the affinity for the CoA ester; instead, they led to a small reduction in affinity for all substrates. This is in contrast to the pronounced decrease in affinity ob-
160
Substrate Promiscuity in Plant OMTs
zymes. We have previously observed all these phenomena for variants of trypsin designed to behave like the closely related serine proteinase Factor Xa.22,23 Clearly, more experimental data are required to understand the complex mechanisms driving protein–ligand affinity and the evolution of substrate specificity. Results from directed evolution may yield important clues for future tailoring of enzymatic reactions.24,25
Conclusion and Summary
Fig. 6. (a) Cartoon illustrating hybrid enzymes of the promiscuous cation-dependent PFOMT and the M. sativa CCoAOMT. 1, PFOMT; 2, PFOMT after removal of the His tag using Factor Xa (ΔHis-PFOMT); 3, PFOMT with Nterminal domain (blue) from M. sativa CCoAOMT (N-term hybrid); 4, same as 3 after Factor Xa cleavage of His tag (ΔHis–N-term hybrid); 5, PFOMT with ‘insertion loop’ (orange) from M. sativa CCoAOMT (loop hybrid); 6, PFOMT with combined M. sativa N-terminal domain and ‘insertion loop’ (double hybrid); 7, M. sativa CCoAOMT. NheI indicates the restriction site used for the generation of the loop and double hybrids (see Materials and Methods) (b) SDS/PAGE showing purified recombinant cation-dependent OMT hybrids.
The structure determination of the promiscuous metal-dependent OMT PFOMT reveals a dimeric protein with a high three-dimensional homology to the CCoA-specific enzyme OMT CCoAOMT. Based on the structures, hybrid proteins were designed through the exchange of N-terminal residues and of a loop region representing an insertion with respect to the mammalian catechol OMT. The resulting loop hybrids exhibited greater catalytic efficiency toward CCoA, while the N-terminal hybrids possessed lower affinity for nonflavonoid substrates. Although the overall structures of the PFOMT and CCoAOMT appear highly similar, our results show that combinations of two variable domains can afford novel enzymes of distinct kinetic properties. With the completion of several genome projects, it is apparent that plants contain several members of apparently redundant, but conserved CCoAOMT-like genes. The results presented here suggest that other members of this class may serve functions distinct to methylation of the single lignin precursor, CCoA.
Materials and Methods Crystallization of the recombinant full-length His-tagged protein
served toward caffeic acid, CCoA, and caffeoyl glucose for the N-terminal-alone hybrids. Based on the present results, the N-terminal residues of PFOMT would appear to be primarily responsible for the activity toward flavonoids, while the intermediate loop plays a role in the positioning and methylation of CCoA. While none of the hybrid variants created exhibit the desired specificity profile of CCoAOMT, the catalytic efficiency of the loop hybrid toward CCoA closely approaches this goal. The relaxed specificity of PFOMT appears to come from a novel combination of structural elements in the neighborhood of the active site, in particular through the use of a flexible N-terminal peptide. The modest change in specificity achieved through recombining surface loops is in line with other attempts to redesign enzymes by rational means.21 While we do not know the reasons for this for PFOMT at present, possible explanations could come from unpredictable changes in the scaffold conformation, changes in protein stability, and/or changes in the dynamic behavior of the hybrid en-
Recombinant PFOMT was produced and affinity purified as described previously.18 The protein was concentrated and rebuffered into 10 mM Tris/HCl, pH 7.5, centrifuged to remove any precipitant, and diluted to a concentration of 6 mg/ml with 10 mM Tris/HCl, pH 7.0. Crystals were obtained after optimization with 20% polyethylene glycol 4000 and 0.2 M CaCl2 in 100 mM Hepes/NaOH, pH 7.0, using a concentration of 3 mg/ml PFOMT, 250 μM MgCl2, 250 μM SAM, 25 μM quercetin, and 2.5% dimethyl sulfoxide through the sitting drop method. For the production of selenomethionine (SeMet)labeled protein for phase determination, bacteria were grown in the presence of SeMet (50 mg/l) medium as described previously26 with a yield of 10.5 mg/l of culture. SeMet crystals were obtained under the same conditions as wild type using a concentration of 2 mg/ml SeMet-PFOMT. Structure determination of PFOMT Crystals of the native PFOMT were measured at the Deutsches Elektronen-Synchrotron facility (BW6 Hamburg, Germany). Initial PFOMT crystals with the orthorhombic
Substrate Promiscuity in Plant OMTs
161
Table 2. Km app and kcat/Km app data obtained with recombinant PFOMT from M. crystallinum, CCoAOMT from M. sativa, and various constructs and hybrids as described in Fig. 6 Substrate Protein Km app (μM) PFOMT PFOMT-Xa N-term hybrid N-term hybrid-Xa Loop hybrid Double hybrid CCoAOMT kcat/Km app (s−1 M−1) PFOMT PFOMT-Xa N-term hybrid N-term hybrid-Xa Loop hybrid Double hybrid CCoAOMT
Quercetin
Quercetagetin
Caffeic acid
0.78 ± 0.26 1.00 ± 0.07 3.93 ± 0.42 4.74 ± 0.59 5.07 ± 0.50 6.97 ± 0.49 5.91 ± 0.71
0.51 ± 0.01 1.76 ± 0.22 1.57 ± 0.39 1.56 ± 0.62 0.96 ± 0.25 2.02 ± 0.24 3.63 ± 0.55
0.99 ± 0.07 8.61 ± 0.41 112 ± 10 N1000 2.81 ± 0.12 3.82 ± 0.93 56.2 ± 13.1
23,600 ± 7900 21,400 ± 1700 2880 ± 320 1920 ± 250 727 ± 77 328 ± 25 10,700 ± 1400
22,000 ± 400 17,200 ± 2300 8900 ± 2300 14,800 ± 6000 1510 ± 400 584 ± 71 8680 ± 1420
22,900 ± 1700 2310 ± 120 93.2 ± 11.5 n.d. 67.9 ± 3.1 20.4 ± 5.2 8.22 ± 2.44
space group C222 or C2221 (cell constants a = 51.2, b = 74.1, c = 117.0 Å, and one molecule per asymmetric unit) diffracted to 1.9 Å resolution. Unfortunately, these crystals showed signs of twinning and could not be used for further structural characterization. Slight modifications of the crystallization conditions resulted in new orthorhombic P212121 crystals with cell constants of a = 48.9, b = 71.8, c = 128.1 Å, and two molecules per asymmetric unit, that diffracted to 1.4 Å. Efforts to solve the structures by molecular replacement using the rOMT12 were unsuccessful; coordinates of the M. sativa CCoAOMT16 became available only later. Phase determination was performed by single-wavelength anomalous dispersion with data up to a resolution of 1.9 Å using CNS.27 Anomalous data for the SeMet crystals were collected at the peak wavelength of the selenium absorption (0.97905 Å), with reference data at the high remote wavelength of 0.95 Å. Model building was performed using the program O,28 coordinates for the SAH cofactor were obtained from the HIC-Up server,29 and the structures were refined using CNS. The final structure contains amino acid residues Gly13 to Thr237 for each monomer and has been refined to a resolution of 1.4 Å. The density clearly revealed the presence of SAH and one calcium ion per monomer (Fig. 4); in addition, a calcium ion was found between two crystallographically related molecules (involving the side chains of GluA106 and GluA109 and symmetry-related GluA204′ and GluA208′). Data collection and refinement statistics are given in Table 3. Visualization of the structure was performed using PyMOL.30 In silico docking experiments The in silico substrate docking experiments were carried out using AutoDock431 with the AutoDockTools' GUI‡. The starting positions and orientations of the substrates were modeled using the bound conformation of feruloyl CoA to CCoAOMT as reference [Protein Data Bank (PDB) code: 1sui].16 A Lamarckian Genetic Algorithm was used to search the active sites of both enzymes (PFOMT and M. sativa CCoAOMT) for alternative substrate conformations using a box size of 20 × 15 × 22 Å3 (36 × 28 × 22 Å3 for the bulkier CCoA). With the exception of CCoA, all ligand single bonds were free to rotate; for the former, all bonds ‡ http://mgltools.scripps.edu/
CCoA 2.86 ± 0.54 6.06 ± 1.97 103 ± 27 N1000 0.35 ± 0.10 0.62 ± 0.15 0.31 ± 0.09 12,400 ± 2400 2130 ± 740 105 ± 36 n.d. 27,900 ± 7800 8390 ± 2020 200,000 ± 60,800
Caffeoyl glucose 2.02 ± 0.15 1.72 ± 0.13 48.2 ± 8.8 49.1 ± 1.6 2.36 ± 0.42 3.07 ± 0.40 76.7 ± 11.2 15,700 ± 1200 16,400 ± 1300 233 ± 54 846 ± 34 145 ± 26 48.6 ± 6.5 289 ± 55
except for the C–C3 bond were fixed, as otherwise no meaningful results could be obtained (data not shown). Two hundred and fifty runs were conducted with a population size of 150 and 250,000 energy evaluations. A total of 27,000 generations were analyzed, with only one conformation surviving to the next generation. Three hundred local search iterations were carried out for each run. The resulting substrate conformations were clustered on the basis of their similarity with an rms tolerance of 2 Å. Docked poses were analyzed visually to detect ‘productive’ binding modes, that is, those in which the hydroxyl group to be methylated is in the vicinity of the SAH thiol group and the neighboring vicinal hydroxyl group approaches the metal ion. Docking results were ranked according to the calculated binding energies of the clusters and the rank of the cluster in which the first productive binding mode is found. Construction of the loop hybrid The first 396 bp of the loop-hybrid construct, containing an NheI restriction site at base pair 396, were amplified from the original PFOMT plasmid using Pfu DNA polymerase (Promega, Mannheim, Germany), introducing a BamHI restriction site at the beginning of the sequence with the primer (CGG GAT CCA ATG GAT TTT GCT GTG). A second 311-bp fragment, corresponding to the 3′ portion of the PFOMT gene with an altered sequence coding for the loop, was custom synthesized (Geneart, Regensburg, Germany). NheI and HindIII restriction sites were introduced at the beginning and the end of the synthesized sequence, respectively, with additional primers. The amplified fragments were gel purified and digested with NheI (New England Biolabs, Frankfurt, Germany) for 1 h at 37 °C. DNA cleavage reactions were monitored for purity by 1.2% agarose gel electrophoresis. Fragments were ligated using LigaFast Rapid DNA Ligation System (Promega). The ligated loop-hybrid gene was reamplified using Pfu DNA polymerase (Promega) as well as primers introducing a BamHI restriction site at the 5′-end and a stop codon followed by a HindIII restriction site (5′ CTC AAG CTT GTG TCA ATA AAG ACG CCT GCA G 3′) at the 3′-end for cloning into pQE30. The DNA was subsequently cut with BamHI and HindIII (Roche, Mannheim, Germany) and cloned into pQE30 expression vector (Qiagen, Hilden, Germany) using the abovementioned sites.
Substrate Promiscuity in Plant OMTs
162 Table 3. Crystallographic data, phasing, and refinement statistics for PFOMT Data set Data collection Wavelength (Å) Resolution (Å) Total reflections Unique reflections Completenessa (%) I/σ(I)a b Ra, sym Redundancy Space group Cell dimensions (Å) a b c Refinement Refinement reflections Rc Rfreed Protein atoms Ligand atoms Solvent molecules Ion atoms rmsd bond lengthse (Å) rmsd bond anglese (°) 〈B-factor〉 protein (Å2) chain A 〈B-factor〉 protein (Å2) chain B 〈B-factor〉 ligand (Å2) 〈B-factor〉 solvent (Å2) 〈B-factor〉 ion (Å2) Ramachandran plot (%) Most favored regions Additional allowed regions Generously allowed regions Disallowed regions
Native
Peak
High remote
1.05 0.97905 0.95 1.37 1.93 1.87 436,990 247,123 259,057 93,389 65,998 72,580 97.6 (92.9) 99.2 (87.1) 99.2 (88.8) 27.88 (2.33) 32.14 (10.46) 28.95 (3.71) 0.069 (0.392) 0.071 (0.190) 0.067 (0.355) 4.67 3.74 3.60 P212121 48.89 71.83 128.12
49.47 71.78 128.15
49.47 71.78 128.15
89,982 0.187 0.223 3577 52 481 3 0.009 1.260 17.22 17.00 14.28 28.90 15.29 92.6 6.3 1.0 0
a
Values in parentheses represent those for the highestresolution shell. b Rsym = ∑|Ih−〈Ih〉|/∑Ih, where 〈Ih〉 is the average intensity over symmetry-equivalent reflections. c R = ∑||Fobs|−|Fcalc||/∑|Fobs|. d Rfree is calculated as R, with 4.3% of the data (4041 reflections) excluded from the refinement. e The rmsd for bonds and angles are the rmsd from ideal values.
immediately prior to the desired N-terminal residue using the following forward primers: 5′ CGG GAT CCA TCG AGG GAA GGA TGG ATT TTG CTG TGA TGA 3′, for proteins containing the original PFOMT N-terminus, and 5′ CGG GAT CCA TCG AGG GAA GGA TGG CAA CCA ACG AAG ATC 3′, for proteins with the N-terminal sequence derived from the M. sativa CCoAOMT. cDNAs were amplified using Isis Proofreading DNA Polymerase (Q-Biogene, Heidelberg, Germany) and ligated into pQE9 vectors using the BamHI and HindIII restriction sites, introduced by the PCR primers. All resultant DNA constructs were confirmed by sequencing (Applied Biosystems, Darmstadt, Germany). Functional expression Plasmids were transformed into the E. coli strain M15 (Qiagen) harboring the plasmid pRep4. Expression was induced after 150 min by the addition of 1 mM isopropyl thio-β-D-galactoside. The bacteria from 400 ml of culture were harvested after 180 min as described previously.18 Incubation times for loop and double hybrids were extended to 6 h at 37 °C and then induced with 1 mM isopropyl thio-β-D-galactoside and transferred to 4 °C and grown for 48 h to increase the yield of the soluble protein. Crude bacterial extract was prepared by ultrasonication and centrifugation (30,000g, 4 °C, 10 min). The clear supernatant was affinity purified as described previously,18 concentrated into 50 mM Tris/HCl, pH 7.5, and 10% glycerol, and the protein was quantified based on the calculated extinction coefficient (ε = 18,730) at 280 nm (Protean Software Package, DNASTAR, Madison, WI, USA). The purification was monitored by SDS/PAGE. Dimerization of the variants was confirmed using gel filtration (GE Healthcare, Superdex 200 16/60) with the following standards: chymotrypsin (17 kDa), ovalbumin (45 kDa), bovine serum albumin (67 kDa), aldolase (158 kDa), and catalase (240 kDa). M. sativa CCoAOMT M. sativa CCoAOMT DNA in the pET15b vector was a kind gift from Dr. Joseph P. Noel (Howard Hughes Medical Institute and Salk Institute for Biological Studies, La Jolla, CA, USA). This plasmid was transformed into E. coli strain BL21DE3 (Novagen, Darmstadt, Germany), and the protein was expressed and purified as described for PFOMT hybrids.
N-terminal and double hybrids
Factor Xa cleavage
The DNA coding for the N-terminal hybrid was prepared by amplification of the PFOMT cDNA using Platinum PCR SuperMix High Fidelity (Invitrogen) and the 92-bp primer containing the sequence coding for the first 22 amino acids from the M. sativa CCoAOMT. The DNA was cloned into pQE9 (Qiagen) using BamHI and HindIII restriction sites as described for the loop hybrid. The double-hybrid DNA fragment was prepared analogously to the N-terminal hybrid using the cDNA coding for the loop hybrid as template and 92 bp primer.
Proteins containing the Factor Xa cleavage site were rebuffered into Factor Xa reaction buffer (20 mM Tris/ HCl, pH 6.5, 50 mM NaCl, and 1 mM CaCl2). The cleavage reactions were conducted in 400 μl volume. The amount of protein was adjusted to 0.25 mg/ml. Two units of Factor Xa protease were used per reaction (400 μl). The reactions were incubated at 37 °C for 24 h to achieve complete cleavage. After incubation, the protease was removed using Factor Xa removal resin (Qiagen). The reactions were purified by Talon affinity resin (BD Biosciences, Heidelberg, Germany). The cleaved protein that did not bind to the matrix and eluted with the flow through was concentrated and rebuffered to 50 mM Tris/HCl, pH 7.5, and 10% glycerol and checked for enzymatic activity. Cleavage and purity were confirmed by SDS/PAGE. Western blot analysis was also performed with His-tag-
OMTs containing a Factor Xa cleavage site To investigate the possible influence of the N-terminal His tag, a Factor Xa cleavage site (IEGR) was introduced
Substrate Promiscuity in Plant OMTs specific antibodies (Qiagen) in order to verify the absence of any His tag. Enzyme activity tests and kinetic data Enzyme assays were performed in a buffer containing 100 mM K2PO4, pH 7.5, and 10% glycerol with 10 μM substrate (dissolved in 30% dimethyl sulfoxide), 0.5–2 μg of total protein, and 400 μM SAM in a total volume of 50 μl. The assays were incubated at 30 °C for 60 to 3600 s (dependent on the protein and substrate tested) and then stopped by the addition of 20 μl of 7% trichloroacetic acid in 50% acetonitrile/water. The reaction products were analyzed by HPLC as described previously.17 Caffeoyl glucose was prepared as described previously from caffeic acid and UDP-glucose with the purified recombinant sinapic acid glucosyltransferase from Brassica napus.32 CCoA was prepared based on published methods.33,34 Caffeic acid was obtained from Serva (Heidelberg, Germany), quercetagetin was obtained from Extrasynthese (Genay, France), and quercetin was obtained from Roth (Karlsruhe, Germany). The reaction products were analyzed by reversed-phase liquid chromatography on a Nucleosil 5-μm C18 column (50 mm length × 4 mm inner diameter; Macherey and Nagel, Düren, Germany), as described previously.17 Compounds were analyzed with linear gradients from 10% B (100% acetonitrile) in A (1.5% aqueous phosphoric acid) to 70% B in A (for phenolics), from 5% B to 50% B in A (for free acids and CoA esters), from 5% B to 30% B (for glucose esters), and from 20% B in A to 80% B in A in 4 min (for flavonoids) at a flow rate of 1 ml min− 1. Detection of flavonoids, catechol, coumarins, and hydroxycinnamic acid esters was performed between 260 and 400 nm. Identification and quantification was achieved with reference compounds from our Institute collection. For Km determination of methyl group acceptors, acceptor concentrations were chosen between 2 and 20 μM, while SAM was kept constant at 1.5 mM. Km, Vmax, and kcat values were calculated by nonlinear curve fitting, assuming Michaelis–Menten steady state kinetics. All enzyme assays were recorded in triplicate for each experiment. PDB accession code Coordinates and structure factors have been deposited in the PDB with accession code 3C3Y.
Acknowledgements We thank Jean-Luc Ferrer (Grenoble) and Joseph P. Noel (San Diego) for providing the M. sativa CCoAOMT plasmid and information prior to publication and Norbert Sträter (Leipzig) for the use of his X-ray facilities at the initial stages of this project. Diffraction data were measured on the MPG/GBF beamline BW6 at Deutsches Elektronen-Synchrotron, where we gratefully acknowledge the assistance of Gleb Bourenkov and Hans Bartunik. This work was in part supported by grants VO719/5 and STU297/4 from the Deutsche Forschungsgemeinschaft as well as the Schwerpunktprogramm SPP1152 ‘Evolution metabolischer Diversität.’
163
References 1. Zhu, B. T., Ezell, E. L. & Liehr, J. G. (1994). Catechol-Omethyltransferase-catalyzed rapid O-methylation of mutagenic flavonoids. Metabolic inactivation as a possible reason for their lack of carcinogenicity in vivo. J. Biol. Chem. 269, 292–299. 2. Ye, Z. H., Kneusel, R. E., Matern, U. & Varner, J. E. (1994). An alternative methylation pathway in lignin biosynthesis in zinnia. Plant Cell, 6, 1427–1439. 3. Maury, S., Geoffroy, P. & Legrand, M. (1999). Tobacco Omethyltransferases involved in phenylpropanoid metabolism. The different caffeoyl coenzyme A/5-hydroxyferuloyl–coenzyme A 3/5-O-methyltransferase and caffeic acid/5-hydroxyferulic acid 3/5-O-methyltransferase classes have distinct substrate specificities and expression patterns. Plant Physiol. 121, 215–224. 4. Osakabe, K., Tsao, C. C., Li, L., Popko, J. L., Umezawa, T., Carraway, D. T. et al. (1999). Coniferyl aldehyde 5hydroxylation and methylation direct syringyl lignin biosynthesis in angiosperms. Proc. Natl Acad. Sci. USA, 96, 8955–8960. 5. Inoue, K., Sewalt, V. J. H., Murray Ballance, G., Ni, W., Sturzer, C. & Dixon, R. A. (1998). Developmental expression and substrate specificities of alfalfa caffeic acid 3-O-methyltransferase and caffeoyl coenzyme A 3-O-methyltransferase in relation to lignification. Plant Physiol. 117, 761–770. 6. Humphreys, J. M. & Chapple, C. (2002). Rewriting the lignin roadmap. Curr. Opin. Plant Biol. 5, 224–229. 7. Peter, G. & Neale, D. (2004). Molecular basis for the evolution of xylem lignification. Curr. Opin. Plant Biol. 7, 737–742. 8. Zubieta, C., He, X. Z., Dixon, R. A. & Noel, J. P. (2001). Structures of two natural product methyltransferases reveal the basis for substrate specificity in plant Omethyltransferases. Nat. Struct. Mol. Biol. 8, 271–279. 9. Zubieta, C., Kota, P., Ferrer, J. L., Dixon, R. A. & Noel, J. P. (2002). Structural basis for the modulation of lignin monomer methylation by caffeic acid/5-hydroxyferulic acid 3/5-O-methyltransferase. Plant Cell, 14, 1265–1277. 10. Zubieta, C., Ross, J. R., Koscheski, P., Yang, Y., Pichersky, E. & Noel, J. P. (2003). Structural basis for substrate recognition in the salicylic acid carboxyl methyltransferase family. Plant Cell, 15, 1704–1716. 11. Yang, H., Ahn, J. H., Ibrahim, R. K., Lee, S. & Lim, Y. (2004). The three-dimensional structure of Arabidopsis thaliana O-methyltransferase predicted by homologybased modelling. J. Mol. Graphics Modell. 23, 77–87. 12. Vidgren, J., Svensson, L. A. & Liljas, A. (1994). Crystal structure of catechol O-methyltransferase. Nature, 368, 354–358. 13. Mannisto, P. T. & Kaakkola, S. (1999). Catechol-Omethyltransferase (COMT): biochemistry, molecular biology, pharmacology, and clinical efficacy of the new selective COMT inhibitors. Pharmacol. Rev. 51, 593–628. 14. Schluckebier, G., O'Gara, M., Saenger, W. & Cheng, X. (1995). Universal catalytic domain structure of AdoMet-dependent methyltransferases. J. Mol. Biol. 247, 16–20. 15. Vidgren, J. (1998). X-ray crystallography of catechol O-methyltransferase: perspectives for target-based drug development. Adv. Pharmacol. 42, 328–330. 16. Ferrer, J. L., Zubieta, C., Dixon, R. A. & Noel, J. P. (2005). Crystal structures of alfalfa caffeoyl coenzyme A 3-O-methyltransferase. Plant Physiol. 137, 1009–1017. 17. Ibdah, M., Zhang, X. H., Schmidt, J. & Vogt, T. (2003).
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A novel Mg++-dependent O-methyltransferase in the phenylpropanoid metabolism of Mesembryanthemum crystallinum. J. Biol. Chem. 278, 43961–43972. Vogt, T. (2004). Regiospecificity and kinetic properties of a plant natural product O-methyl-transferase are determined by its N-terminal domain. FEBS Lett. 561, 159–162. Lukacin, R., Matern, U., Specker, S. & Vogt, T. (2004). Cations modulate the substrate specificity of bifunctional class I O-methyltransferase from Ammi majus. FEBS Lett. 577, 367–370. Hou, X. W., Wang, Y. L., Zhou, Z. W., Bao, S. L., Lin, Y. J. & Gong, W. M. (2007). Crystal structure of SAM-dependent O-methyltransferase from pathogenic bacterium Leptospira interrogans. J. Struct. Biol. 159, 523–528. Kohara, A., Nakajima, C., Yoshida, S. & Muranaka, T. (2007). Characterisation and engineering of glycosyltransferases responsible for steroid saponin biosynthesis in Solanaceous plants. Phytochemistry, 68, 478–486. Reyda, S., Sohn, C., Klebe, G., Rall, K., Ullmann, D., Jakubke, H.-D. & Stubbs, M. T. (2003). Reconstructing the binding site of factor Xa in trypsin reveals ligandinduced structural plasticity. J. Mol. Biol. 325, 963–977. Rauh, D., Klebe, G. & Stubbs, M. T. (2004). Understanding protein–ligand interactions: the price of protein flexibility. J. Mol. Biol. 335, 1325–1341. Aharoni, A., Gaidukov, L., Khersonsky, O., McQ Gould, S., Roodveldt, C. & Tawfik, D. S. (2005). The ‘evolvability’ of promiscuous protein functions. Nat. Genet. 37, 73–76. Williams, G. J., Zhang, C. & Thorson, J. S. (2007). Expanding the promiscuity of a natural product glycosyltransferase by directed evolution. Nat. Chem. Biol. 3, 657–662.
Substrate Promiscuity in Plant OMTs 26. Doublie, S. (1997). Preparation of selenomethionyl proteins for phase determination. Methods Enzymol. 276, 523–530. 27. Brünger, A., Adams, P., Clore, G., DeLano, W., Gros, P., Grosse-Kunstleve, R. et al. (1998). Crystallography and NMR system: a new software suite for macromolecular structure determination. Acta Crystallogr., Sect. D: Biol. Crystallogr. 54, 905–921. 28. Jones, T. A., Zou, J. Y. & Cowan, S. W. (1991). Improved methods for building protein models in electron density maps and the location of errors in these models. Acta Crystallogr., Sect. A: Found. Crystallogr. 47, 110–119. 29. Kleywegt, G. J. & Jones, T. A. (1998). Databases in protein crystallography. Acta Crystallogr., Sect. D: Biol. Crystallogr. 54, 1119–1131. 30. DeLano, W. (2002). The PyMOL Molecular Graphics System, DeLano Scientific, Palo Alto, CA. 31. Morris, G. M., Goodsell, D. S., Halliday, R. S., Huey, R., Hart, W. E., Belew, R. K. & Olson, A. J. (1998). Automated docking using a Lamarckian Genetic Algorithm and empirical binding free energy function. J. Comput. Chem. 19, 1639–1662. 32. Milkowski, C., Baumert, A. & Strack, D. (2000). Cloning and heterologous expression of a rape cDNA encoding UDP-glucose:sinapate glucosyltransferase. Planta, 211, 883–886. 33. Strack, D., Keller, H. & Weissenböck, G. (1987). Enzymatic synthesis of hydroxycinnamic acid esters of sugar acids and hydroaromatic acids by protein preparations of Raphanus sativus. J. Plant Physiol. 131, 61–73. 34. Stöckigt, J. & Zenk, M. H. (1975). Chemical synthesis and properties of hydroxycinnamoyl coenzyme A derivatives. Z. Naturforsch., C: Biosci. 30, 352–358.
doi:10.1016/j.jmb.2008.02.019
Available online at www.sciencedirect.com
Biochemical and Structural Analysis of Substrate Promiscuity in Plant Mg 2+ -Dependent O-Methyltransferases Jakub G. Kopycki 1 †, Daniel Rauh 1 †, Alexander A. Chumanevich 1 , Piotr Neumann 1 , Thomas Vogt 2 ⁎ and Milton T. Stubbs 1,3 ⁎ 1
Institut für Biotechnologie, Martin-Luther-Universität Halle-Wittenberg, Kurt-MothesStraße 3, D-06120 Halle (Saale), Germany 2
Leibniz-Institute of Plant Biochemistry, Department of Secondary Metabolism, Weinberg 3, D-06120 Halle (Saale), Germany 3
Mitteldeutsches Zentrum für Struktur und Dynamik der Proteine, Kurt-Mothes-Straße 3, D-06120 Halle (Saale), Germany
Plant S-adenosyl-L-methionine-dependent class I natural product Omethyltransferases (OMTs), related to animal catechol OMTs, are dependent on bivalent cations and strictly specific for the meta position of aromatic vicinal dihydroxy groups. While the primary activity of these class I enzymes is methylation of caffeoyl coenzyme A OMTs, a distinct subset is able to methylate a wider range of substrates, characterized by the promiscuous phenylpropanoid and flavonoid OMT. The observed broad substrate specificity resides in two regions: the N-terminus and a variable insertion loop near the C-terminus, which displays the lowest degree of sequence conservation between the two subfamilies. Structural and biochemical data, based on site-directed mutagenesis and domain exchange between the two enzyme types, present evidence that only small topological changes among otherwise highly conserved 3-D structures are sufficient to differentiate between an enzymatic generalist and an enzymatic specialist in plant natural product methylation. © 2008 Elsevier Ltd. All rights reserved.
Received 12 November 2007; received in revised form 5 February 2008; accepted 7 February 2008 Available online 20 February 2008 Edited by M. Guss
Keywords: crystal structure; substrate specificity; hybrid protein; loop exchange; plant secondary metabolism
Introduction
*Corresponding authors. M. T. Stubbs is to be contacted at Institut für Biotechnologie, Martin-Luther-Universität Halle-Wittenberg, Kurt-Mothes-Straße 3, D-06120 Halle (Saale), Germany. E-mail addresses: [email protected]; [email protected]. † J.G.K. and D.R. contributed equally to this work. Present address: D. Rauh, Chemical Genomics Centre of the Max Planck Society, Otto-Hahn-Straße 15, D-44227 Dortmund, Germany. Abbreviations used: OMT, O-methyltransferase; PFOMT, phenylpropanoid and flavonoid OMT; CoA, coenzyme A; CCoA, caffeoyl CoA; rOMT, rat catechol OMT; SAM, S-adenosyl-L-methionine; SAH, S-adenosyl-Lhomocysteine; SeMet, selenomethionine.
Methylation by S-adenosyl-L-methionine (SAM)dependent O-methyltransferases (OMTs) (EC 2.1.1) is a common modification in natural product biosynthesis. Site-specific O-methylation modulates the physiological properties and the chemical reactivity of phenolic hydroxy groups.1 Class I plant cationdependent OMTs (also known as caffeoyl coenzyme A OMTs or CCoAOMTs) constitute a group of low molecular mass (23–27 kDa) enzymes that play an important role in methylation of guaiacyl residues present in CCoA or 5-hydroxy feruloyl CoA, both precursors of monolignols.2 In angiosperms, members of a second class of metal-independent OMTs (caffeic acid OMTs) are able to methylate the monolignol precursors caffeic acid, caffeyl aldehyde, or caffeyl alcohol in the meta position.3–5 The action of
0022-2836/$ - see front matter © 2008 Elsevier Ltd. All rights reserved.
Substrate Promiscuity in Plant OMTs
both enzyme classes in a metabolic grid appears to be crucial for determining the structural integrity of the lignin polymer.6,7 Detailed structural information on the cationindependent enzymes is available, including the chalcone and isoflavone OMTs from alfalfa (Medicago sativa), a flavonoid OMT from Arabidopsis thaliana, as well as an OMT involved in methyl ester formation in Clarkia breweri.8–11 These enzymes are obligatory dimers, with each active site formed from residues of both monomeric polypeptide chains. Either general acid/base catalysis (in the case of phenolic ether formation) or proper substrate positioning (in the case of the methyl ester formation) is essential for efficient methyl transfer in the individual systems. Although a major goal of these investigations on the class II enzymes has been the tailored modification of methylation specificity, which would enable the engineering of biochemical pathways for the production of transgenic plants with altered lignin composition or alternative accumulation of diverse natural products, mutagenesis of individual amino acids has until now only resulted in lower substrate specificity.10 In contrast, less structural information is available for cation-dependent OMTs. The 2.0-Å-resolution structure of rat catechol OMT (rOMT)12 exhibits a typical Rossmann fold within the coenzyme-binding domain of the enzyme, suggesting that SAM methylases share a common structural motif.13 Methyl
155
Fig. 1. Structures of OMT substrates methylated by promiscuous CCoAOMT-like enzymes such as PFOMT from the ice plant M. crystallinum.18 1, quercetin; 2, quercetagetin; 3, caffeic acid; 4, CCoA; 5, caffeoyl glucose. The positions of the methyl group acceptor hydroxyl groups are marked; note the two methylation positions for quercetagetin.
transfer proceeds via an Sn2-like transition state, with one ionized hydroxyl group, supported by the Mg2+ ion, and a conserved lysine as a catalytic base, to attack the electron-deficient methyl group of SAM.14 In vertebrates, the enzyme is responsible for the inactivation of neurotransmitters such as dopamine in the brain and has therefore received particular attention as an attractive target to cure degenerate brain diseases.13,15 However, high concentrations of
Fig. 2. Overall structure of the PFOMT dimer in cartoon representation. Monomer A is in yellow, with the N-terminus in blue and the ‘insertion loop’ in orange; the secondary structure elements of PFOMT are shown for monomer B, which is color ramped from the N-terminus (blue) to the C-terminus (red). The positions of the active sites are indicated by the bound calcium ion (gray sphere) and the cofactor SAH (stick representation).
156 catechol OMT are also found in the liver, kidney, or placenta, where the enzyme inactivates potentially reactive or mutagenic phenolics such as quercetin.1 Recently, the 3-D structure of the CCoA-specific CCoAOMT from M. sativa was solved at a resolution of 2.4 Å.16 This enzyme, which almost exclusively methylates the CoA ester of caffeic acid, displays very low affinity for caffeic acid itself, with a direct impact on lignin polymer biosynthesis in gymnoand angiosperms. 7 The structure of the alfalfa CCoAOMT provides insights into the mode of action and specificity of this class of enzymes. While the overall fold and the active site of the CCoAOMT dimer resemble those of the animal catechol OMT, significant differences in the positioning of the Nterminus and intervening loops in the vicinity of the active site contribute to specificity for CCoA. In addition to the CCoA-specific enzymes, a second type of cation-dependent OMT from the ice plant Mesembryanthemum crystallinum [phenylpropanoid and flavonoid OMT (PFOMT)] with promiscuous substrate specificity has been described.17 Phylogenetic analyses indicate that plant cation-dependent
Substrate Promiscuity in Plant OMTs
OMTs can be divided into two subclades. The CCoAspecific cluster responsible for lignin biosynthesis exhibits a high sequence identity of over 80% at the amino acid level. The more divergent subcluster epitomized by PFOMT is found in several plant species as multiple sequences, which may be differentiated by substrate specificity or by differentially regulated expression (T.V., unpublished results). Functionally expressed in Escherichia coli, the promiscuous OMTs methylate a variety of natural products with vicinal dihydroxy groups in vitro that are not accepted by the lignin-specific enzyme, including caffeic acid and caffeoyl glucose17,18 (Fig. 1). In the ice plant, it has been established that the PFOMTcatalyzed methylation of flavonoids is induced upon ultraviolet irradiation to protect the plant against further damage.17 Further studies have shown that substrate specificity can be influenced by the nature of the cation,19 while the N-terminus of PFOMT (which, in the plant, is some 11 residues shorter than that derived from the gene sequence) plays a crucial role in determining the position specificity and kinetic properties of the enzyme.18
Fig. 3. Stereo view of the superposition of PFOMT (A monomer, colored as in Fig. 2) with (a) M. sativa CCoAOMT16 (green) and (b) rOMT12 (magenta). Coordinates of the cofactor SAH (red sticks) are from the present PFOMT structure, and positions of the bound cations are depicted by the colored spheres. CCoAOMT product 5-hydroxy feruloyl CoA (green) and rOMT inhibitor dinitrocatechol (magenta) from the respective structures are depicted as sticks. Note the difference in the ‘insertion loop,’ which supports the CoA moiety in the CCoAOMT structure, as well as the shorter loop between β6 and β7 and the alternative N-terminal arrangement of secondary structure elements in the plant enzymes compared to rOMT, which allows dimerization of the former.
Substrate Promiscuity in Plant OMTs
157
Here, we report the crystallization and 3-D structure determination of the cation-dependent PFOMT from the ice plant with promiscuous specificity at 1.4 Å resolution. Catalytic properties of N-terminally truncated variants and hybrids containing loops from the CCoA-specific alfalfa OMT demonstrate their significant influence on substrate specificity. Simultaneous replacement of two regions from the alfalfa into the ice plant enzyme converts the promiscuous enzyme to a CCoA-specific OMT, demonstrating the important contribution of substrate binding by the N-terminus and the loop domain.
Results and Discussion PFOMT is a symmetric dimer of approximate dimensions 70 × 40 × 50 Å3 (Fig. 2). As expected, each monomer exhibits an α/β fold similar to the monomeric animal rOMT,13 with eight α-helices and seven β-strands, including the nucleotide-binding motif or Rossmann fold. In contrast to rOMT but in common with CCoAOMT,16 however, the first two helices of PFOMT show a distinctly different topological arrangement. The position of helix α1 in the plant enzymes is in part accomplished by a deletion between strands β6 and β7 with respect to the mammalian enzyme and contributes significantly to dimer formation (Fig. 3). Despite cocrystallization with a methyl group donor SAM and the substrate quercetin, only the resultant S-adenosyl-L-homocysteine (SAH) could be detected in the active center (Fig. 4a). Although no electron density could be observed for the catalytically important N-terminal residues, the positioning of the first visible amino acids indicates that the N-termini would be well placed to influence catalytic activity. The two active sites, demarcated by the presence of SAH and bound Ca2+ ions (Fig. 2), could act independently of one another; it is conceivable, however, that a mobile amino-terminus of one monomer might communicate with the catalytic apparatus of the second monomer, as has been observed recently for a dimeric bacterial OMT from Leptospira interrogans of unknown specificity.20 This is in contrast to the plant cation-independent OMTs, whose active site consists of residues from both monomers of the dimer, leading to the strong influence of catalytic properties of the corresponding second subunit.8 The catalytic apparatus and residues in the immediate vicinity are almost identical in each of the three metal-dependent enzymes, the only significant difference here being the replacement of rOMT Trp143, which partially covers the SAM moiety and forms a lower boundary to the substrate binding pocket, by Asp165. All attempts at soaking crystals or cocrystallization with a variety of substrates failed to yield any electron density to establish the binding modes within the active site of PFOMT (data not shown). Nevertheless, superposition of the active sites of PFOMT with ligand-bound structures of CCoAOMT and rOMT allows modeling of the binding of quercetin and quercetagetin, whose
Fig. 4. The active site of PFOMT. (a) Representative electron density (2Fo − Fc, contoured at 0.9σ) showing the cofactor SAH and the bound cation (gray sphere), interpreted as Ca2+. (b) Docking of the substrate quercetagetin in the PFOMT active site. Quercetagetin is methylated in two distinct positions (see Fig. 1), requiring opposing orientations of the substrate (yellow and pink). Both orientations make use of aromatic stacking interactions between the substrate and the side chains of Tyr51, Trp184, and Phe198 from the ‘insertion loop,’ positioning the pertinent hydroxyl groups in the vicinity of the SAM methyl donor and the active-site Mg2+ ion (gray sphere).
extended aromatic moieties would be sandwiched between the side chains of Tyr51, Trp184, and Phe198 (Fig. 4b). In silico docking experiments were carried out to obtain semiquantitative data for the relative specificities of the two plant enzymes (Table 1). Despite the absence of structural data for the amino-terminal residues for both proteins, the results are in general agreement with the observed specificities. For quercetin, a substrate-like orientation of the flavonoid could be determined in the active center of PFOMT with reasonable binding energies, whereas no enzy-
Substrate Promiscuity in Plant OMTs
158
Table 1. Results of docking of quercetin, caffeic acid, and feruloyl CoA into the active sites of PFOMT and CCoAOMT PFOMT Cluster rank
CCoAOMT
Lowest binding energy
Mean binding energy
Number in cluster
Cluster rank
Lowest binding energy
Mean binding energy
Number in cluster
Quercetin 1 2 3 4 5 5.1 6 7 8 9 10 11 12 13 14 15 16
− 7.93 − 7.51 − 7.00 − 6.93 − 6.84 − 6.84 − 6.83 − 6.75 − 6.68 − 6.61 − 6.54 − 6.52 − 6.49 − 6.38 − 6.34 − 6.17 − 5.82
− 6.81 − 6.72 − 6.56 − 6.32 − 6.49
53 41 23 15 13
− 6.45 − 6.31 − 6.37 − 6.37 − 6.08 − 6.25 − 6.18 − 6.32 − 6.34 − 5.80 − 5.78
8 8 14 5 10 44 7 3 1 3 2
1 2 3 4 5 6 7 8 · · · · 18 19 20 21 n.d.
− 8.18 − 7.94 − 7.73 − 7.73 − 7.48 − 7.21 − 7.17 − 7.16
−7.44 −7.47 −7.31 −7.03 −7.10 −6.92 −6.82 −7.16
− 6.40 − 6.37 − 6.35 − 6.02
−6.36 −6.37 −6.35 −6.02
67 63 16 32 25 3 15 1 · · · · 2 1 1 1
Caffeic acid 1 1.1 2 3 4 5 6 7 8 9
− 5.44 − 5.44 − 4.66 − 4.41 − 4.33 − 4.18 − 4.18 − 4.17 − 4.09 − 3.94
− 4.92
195
− 4.21 − 4.31 − 4.21 − 4.07 − 4.18 − 4.17 − 3.98 − 3.77
32 5 3 2 1 1 5 6
1 2 3 4 5 5.1 · · · 19
− 4.53 − 4.50 − 4.47 − 4.45 − 4.43 − 4.43
−4.23 −4.20 −4.22 −4.31 −4.09
16 13 11 3 83
+ 0.86
31
+ 1.22
20 · · · 2 1 1 1 1
5OH-feruloyl CoA 1 1.1 2 . . . 55 56 57 58 59
− 0.35 − 0.35 + 0.42
+ 2.65 + 2.92 + 2.96 + 3.08 + 3.25
+ 2.69 + 2.92 + 2.96 + 3.08 + 3.25
1 1.1 2 · · · 11 12 13 14 15
− 3.63
−3.63
− 2.85 − 2.85 − 2.64
−2.17
− 1.90 − 1.90 − 1.25 − 1.23 + 0.66
· · · 1 144
−2.16
−1.68 −1.90 −1.19 −1.23 +0.66
3 · · · 5 1 2 1 1
The highest ranking substrate-like binding modes are indicated in boldface, together with the corresponding cluster and binding energy (kcal mol− 1). n.d., not detected.
matically reasonable binding mode could be found for CCoAOMT. This was also the case for quercetagetin (data not shown). Caffeic acid and feruloyl CoA could be docked into both active sites with reasonable energies, although the in silico results clearly distinguish between the two enzymes. Caffeic acid docks to PFOMT favorably in terms of binding energy, cluster ranking, and cluster size; the results for CCoAOMT are less conclusive. This situation is reversed for the docking of feruloyl CoA, where only a small fraction of the binding poses for PFOMT show favorable (negative) energies. The most notable difference between the plant and animal enzymes is a large insertion loop between strand β5 and helix α8 (Fig. 3). The conformation of this insertion, which begins with a short α-helical turn, is supported by the N-terminal helix of the opposing monomer, so that the active-site cleft
of the dimeric plant enzymes appears deeper and more restricted (Fig. 4). This insertion loop provides a scaffold for the CoA moiety of CCoA in the CCoAOMT structure.16 Most interestingly, this loop in PFOMT exhibits a conformation markedly different from that in CCoAOMT (Fig. 3). Moreover, sequence analysis reveals that this region demonstrates the lowest sequence homology between CCoA-specific and promiscuous enzymes (30% identity for residues 185– 208, Fig. 5). Thus, it seems plausible that differences in this loop and in the N-termini of the two enzyme classes (which also exhibit low sequence conservation and are observed neither in the present structure nor in that of CCoAOMT16) could provide a clue to the promiscuous nature of PFOMT. Hybrid recombinant proteins were designed and expressed in E. coli and their substrate specificity was
Substrate Promiscuity in Plant OMTs
159
Fig. 5. Sequence alignment of promiscuous OMTs (from M. crystallinum and Stellaria longipes) and diverse, specific CCoAOMTs, together with the mammalian rOMT. Secondary structure elements of PFOMT are shown, colored as in Fig. 2 (monomer B). Boxed are the variable N-terminal residues (in blue) and the ‘insertion loop’ (in orange), which shows structural and sequence divergence between the promiscuous and specific enzymes.
characterized to investigate the influence of these two regions on substrate preference (Fig. 6). We have previously shown that the N-terminus of PFOMT plays a crucial role in determining the position specificity and kinetic properties of the enzyme.18 A new construct was made with a cleavable His tag in order to study the influence of the N-terminal His tag. Removal of the His tag from native PFOMT (ΔHisPFOMT, Table 2) resulted in a significant loss of catalytic efficiency toward the substrates caffeic acid and CCoA, largely due to a reduction in Km app, while turnover of the flavonoids was largely unaffected. In contrast, replacement of the PFOMT N-terminus with that of the specific CCoAOMT enzyme from alfalfa (N-term hybrid) resulted in a reduced catalytic efficiency toward flavonoids, an effect that is even more pronounced for the substrates caffeic acid, CCoA, and caffeoyl glucose. In all cases, a 10- to 100-fold increase in Km app reflects a reduction in substrate affinity, which is the major factor contributing to the strongly reduced catalytic efficiency kcat/Km app. As with PFOMT, removal of the His tag from the N-terminal hybrid (ΔHis–Nterm hybrid) resulted in a strong reduction in activity toward caffeic acid and CCoA; interestingly, the ΔHis–N-term hybrid exhibited an increased activity
toward quercetagetin, such that this enzyme has an almost exclusive flavonoid activity. Substitution of the insertion loop from the specific alfalfa CCoAOMT in the nonspecific PFOMT (loop hybrid) leads to a stark reduction in soluble protein expression (1 mg/ml instead of the usual 10 mg/ ml), with most of the protein depositing as insoluble inclusion bodies (data not shown). The loop hybrid exhibited a 10-fold increase in affinity for and activity against CCoA and a marked reduction in affinity for the flavonoids; that is, the loop-hybrid protein behaves more like CCoAOMT than PFOMT. On the other hand, replacement of the loop resulted in little change in Km app for caffeoyl glucose, although a considerable loss in activity against this substrate was observed. Despite the strongly reduced Km app for CCoA, the high affinity for caffeoyl glucose continues to distinguish the loop hybrid from the corresponding alfalfa protein. Variants in which both the N-terminal peptide and the insertion loop of PFOMT were replaced simultaneously by those of the corresponding CCoAOMT sequence (double hybrid) failed to increase the affinity for the CoA ester; instead, they led to a small reduction in affinity for all substrates. This is in contrast to the pronounced decrease in affinity ob-
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Substrate Promiscuity in Plant OMTs
zymes. We have previously observed all these phenomena for variants of trypsin designed to behave like the closely related serine proteinase Factor Xa.22,23 Clearly, more experimental data are required to understand the complex mechanisms driving protein–ligand affinity and the evolution of substrate specificity. Results from directed evolution may yield important clues for future tailoring of enzymatic reactions.24,25
Conclusion and Summary
Fig. 6. (a) Cartoon illustrating hybrid enzymes of the promiscuous cation-dependent PFOMT and the M. sativa CCoAOMT. 1, PFOMT; 2, PFOMT after removal of the His tag using Factor Xa (ΔHis-PFOMT); 3, PFOMT with Nterminal domain (blue) from M. sativa CCoAOMT (N-term hybrid); 4, same as 3 after Factor Xa cleavage of His tag (ΔHis–N-term hybrid); 5, PFOMT with ‘insertion loop’ (orange) from M. sativa CCoAOMT (loop hybrid); 6, PFOMT with combined M. sativa N-terminal domain and ‘insertion loop’ (double hybrid); 7, M. sativa CCoAOMT. NheI indicates the restriction site used for the generation of the loop and double hybrids (see Materials and Methods) (b) SDS/PAGE showing purified recombinant cation-dependent OMT hybrids.
The structure determination of the promiscuous metal-dependent OMT PFOMT reveals a dimeric protein with a high three-dimensional homology to the CCoA-specific enzyme OMT CCoAOMT. Based on the structures, hybrid proteins were designed through the exchange of N-terminal residues and of a loop region representing an insertion with respect to the mammalian catechol OMT. The resulting loop hybrids exhibited greater catalytic efficiency toward CCoA, while the N-terminal hybrids possessed lower affinity for nonflavonoid substrates. Although the overall structures of the PFOMT and CCoAOMT appear highly similar, our results show that combinations of two variable domains can afford novel enzymes of distinct kinetic properties. With the completion of several genome projects, it is apparent that plants contain several members of apparently redundant, but conserved CCoAOMT-like genes. The results presented here suggest that other members of this class may serve functions distinct to methylation of the single lignin precursor, CCoA.
Materials and Methods Crystallization of the recombinant full-length His-tagged protein
served toward caffeic acid, CCoA, and caffeoyl glucose for the N-terminal-alone hybrids. Based on the present results, the N-terminal residues of PFOMT would appear to be primarily responsible for the activity toward flavonoids, while the intermediate loop plays a role in the positioning and methylation of CCoA. While none of the hybrid variants created exhibit the desired specificity profile of CCoAOMT, the catalytic efficiency of the loop hybrid toward CCoA closely approaches this goal. The relaxed specificity of PFOMT appears to come from a novel combination of structural elements in the neighborhood of the active site, in particular through the use of a flexible N-terminal peptide. The modest change in specificity achieved through recombining surface loops is in line with other attempts to redesign enzymes by rational means.21 While we do not know the reasons for this for PFOMT at present, possible explanations could come from unpredictable changes in the scaffold conformation, changes in protein stability, and/or changes in the dynamic behavior of the hybrid en-
Recombinant PFOMT was produced and affinity purified as described previously.18 The protein was concentrated and rebuffered into 10 mM Tris/HCl, pH 7.5, centrifuged to remove any precipitant, and diluted to a concentration of 6 mg/ml with 10 mM Tris/HCl, pH 7.0. Crystals were obtained after optimization with 20% polyethylene glycol 4000 and 0.2 M CaCl2 in 100 mM Hepes/NaOH, pH 7.0, using a concentration of 3 mg/ml PFOMT, 250 μM MgCl2, 250 μM SAM, 25 μM quercetin, and 2.5% dimethyl sulfoxide through the sitting drop method. For the production of selenomethionine (SeMet)labeled protein for phase determination, bacteria were grown in the presence of SeMet (50 mg/l) medium as described previously26 with a yield of 10.5 mg/l of culture. SeMet crystals were obtained under the same conditions as wild type using a concentration of 2 mg/ml SeMet-PFOMT. Structure determination of PFOMT Crystals of the native PFOMT were measured at the Deutsches Elektronen-Synchrotron facility (BW6 Hamburg, Germany). Initial PFOMT crystals with the orthorhombic
Substrate Promiscuity in Plant OMTs
161
Table 2. Km app and kcat/Km app data obtained with recombinant PFOMT from M. crystallinum, CCoAOMT from M. sativa, and various constructs and hybrids as described in Fig. 6 Substrate Protein Km app (μM) PFOMT PFOMT-Xa N-term hybrid N-term hybrid-Xa Loop hybrid Double hybrid CCoAOMT kcat/Km app (s−1 M−1) PFOMT PFOMT-Xa N-term hybrid N-term hybrid-Xa Loop hybrid Double hybrid CCoAOMT
Quercetin
Quercetagetin
Caffeic acid
0.78 ± 0.26 1.00 ± 0.07 3.93 ± 0.42 4.74 ± 0.59 5.07 ± 0.50 6.97 ± 0.49 5.91 ± 0.71
0.51 ± 0.01 1.76 ± 0.22 1.57 ± 0.39 1.56 ± 0.62 0.96 ± 0.25 2.02 ± 0.24 3.63 ± 0.55
0.99 ± 0.07 8.61 ± 0.41 112 ± 10 N1000 2.81 ± 0.12 3.82 ± 0.93 56.2 ± 13.1
23,600 ± 7900 21,400 ± 1700 2880 ± 320 1920 ± 250 727 ± 77 328 ± 25 10,700 ± 1400
22,000 ± 400 17,200 ± 2300 8900 ± 2300 14,800 ± 6000 1510 ± 400 584 ± 71 8680 ± 1420
22,900 ± 1700 2310 ± 120 93.2 ± 11.5 n.d. 67.9 ± 3.1 20.4 ± 5.2 8.22 ± 2.44
space group C222 or C2221 (cell constants a = 51.2, b = 74.1, c = 117.0 Å, and one molecule per asymmetric unit) diffracted to 1.9 Å resolution. Unfortunately, these crystals showed signs of twinning and could not be used for further structural characterization. Slight modifications of the crystallization conditions resulted in new orthorhombic P212121 crystals with cell constants of a = 48.9, b = 71.8, c = 128.1 Å, and two molecules per asymmetric unit, that diffracted to 1.4 Å. Efforts to solve the structures by molecular replacement using the rOMT12 were unsuccessful; coordinates of the M. sativa CCoAOMT16 became available only later. Phase determination was performed by single-wavelength anomalous dispersion with data up to a resolution of 1.9 Å using CNS.27 Anomalous data for the SeMet crystals were collected at the peak wavelength of the selenium absorption (0.97905 Å), with reference data at the high remote wavelength of 0.95 Å. Model building was performed using the program O,28 coordinates for the SAH cofactor were obtained from the HIC-Up server,29 and the structures were refined using CNS. The final structure contains amino acid residues Gly13 to Thr237 for each monomer and has been refined to a resolution of 1.4 Å. The density clearly revealed the presence of SAH and one calcium ion per monomer (Fig. 4); in addition, a calcium ion was found between two crystallographically related molecules (involving the side chains of GluA106 and GluA109 and symmetry-related GluA204′ and GluA208′). Data collection and refinement statistics are given in Table 3. Visualization of the structure was performed using PyMOL.30 In silico docking experiments The in silico substrate docking experiments were carried out using AutoDock431 with the AutoDockTools' GUI‡. The starting positions and orientations of the substrates were modeled using the bound conformation of feruloyl CoA to CCoAOMT as reference [Protein Data Bank (PDB) code: 1sui].16 A Lamarckian Genetic Algorithm was used to search the active sites of both enzymes (PFOMT and M. sativa CCoAOMT) for alternative substrate conformations using a box size of 20 × 15 × 22 Å3 (36 × 28 × 22 Å3 for the bulkier CCoA). With the exception of CCoA, all ligand single bonds were free to rotate; for the former, all bonds ‡ http://mgltools.scripps.edu/
CCoA 2.86 ± 0.54 6.06 ± 1.97 103 ± 27 N1000 0.35 ± 0.10 0.62 ± 0.15 0.31 ± 0.09 12,400 ± 2400 2130 ± 740 105 ± 36 n.d. 27,900 ± 7800 8390 ± 2020 200,000 ± 60,800
Caffeoyl glucose 2.02 ± 0.15 1.72 ± 0.13 48.2 ± 8.8 49.1 ± 1.6 2.36 ± 0.42 3.07 ± 0.40 76.7 ± 11.2 15,700 ± 1200 16,400 ± 1300 233 ± 54 846 ± 34 145 ± 26 48.6 ± 6.5 289 ± 55
except for the C–C3 bond were fixed, as otherwise no meaningful results could be obtained (data not shown). Two hundred and fifty runs were conducted with a population size of 150 and 250,000 energy evaluations. A total of 27,000 generations were analyzed, with only one conformation surviving to the next generation. Three hundred local search iterations were carried out for each run. The resulting substrate conformations were clustered on the basis of their similarity with an rms tolerance of 2 Å. Docked poses were analyzed visually to detect ‘productive’ binding modes, that is, those in which the hydroxyl group to be methylated is in the vicinity of the SAH thiol group and the neighboring vicinal hydroxyl group approaches the metal ion. Docking results were ranked according to the calculated binding energies of the clusters and the rank of the cluster in which the first productive binding mode is found. Construction of the loop hybrid The first 396 bp of the loop-hybrid construct, containing an NheI restriction site at base pair 396, were amplified from the original PFOMT plasmid using Pfu DNA polymerase (Promega, Mannheim, Germany), introducing a BamHI restriction site at the beginning of the sequence with the primer (CGG GAT CCA ATG GAT TTT GCT GTG). A second 311-bp fragment, corresponding to the 3′ portion of the PFOMT gene with an altered sequence coding for the loop, was custom synthesized (Geneart, Regensburg, Germany). NheI and HindIII restriction sites were introduced at the beginning and the end of the synthesized sequence, respectively, with additional primers. The amplified fragments were gel purified and digested with NheI (New England Biolabs, Frankfurt, Germany) for 1 h at 37 °C. DNA cleavage reactions were monitored for purity by 1.2% agarose gel electrophoresis. Fragments were ligated using LigaFast Rapid DNA Ligation System (Promega). The ligated loop-hybrid gene was reamplified using Pfu DNA polymerase (Promega) as well as primers introducing a BamHI restriction site at the 5′-end and a stop codon followed by a HindIII restriction site (5′ CTC AAG CTT GTG TCA ATA AAG ACG CCT GCA G 3′) at the 3′-end for cloning into pQE30. The DNA was subsequently cut with BamHI and HindIII (Roche, Mannheim, Germany) and cloned into pQE30 expression vector (Qiagen, Hilden, Germany) using the abovementioned sites.
Substrate Promiscuity in Plant OMTs
162 Table 3. Crystallographic data, phasing, and refinement statistics for PFOMT Data set Data collection Wavelength (Å) Resolution (Å) Total reflections Unique reflections Completenessa (%) I/σ(I)a b Ra, sym Redundancy Space group Cell dimensions (Å) a b c Refinement Refinement reflections Rc Rfreed Protein atoms Ligand atoms Solvent molecules Ion atoms rmsd bond lengthse (Å) rmsd bond anglese (°) 〈B-factor〉 protein (Å2) chain A 〈B-factor〉 protein (Å2) chain B 〈B-factor〉 ligand (Å2) 〈B-factor〉 solvent (Å2) 〈B-factor〉 ion (Å2) Ramachandran plot (%) Most favored regions Additional allowed regions Generously allowed regions Disallowed regions
Native
Peak
High remote
1.05 0.97905 0.95 1.37 1.93 1.87 436,990 247,123 259,057 93,389 65,998 72,580 97.6 (92.9) 99.2 (87.1) 99.2 (88.8) 27.88 (2.33) 32.14 (10.46) 28.95 (3.71) 0.069 (0.392) 0.071 (0.190) 0.067 (0.355) 4.67 3.74 3.60 P212121 48.89 71.83 128.12
49.47 71.78 128.15
49.47 71.78 128.15
89,982 0.187 0.223 3577 52 481 3 0.009 1.260 17.22 17.00 14.28 28.90 15.29 92.6 6.3 1.0 0
a
Values in parentheses represent those for the highestresolution shell. b Rsym = ∑|Ih−〈Ih〉|/∑Ih, where 〈Ih〉 is the average intensity over symmetry-equivalent reflections. c R = ∑||Fobs|−|Fcalc||/∑|Fobs|. d Rfree is calculated as R, with 4.3% of the data (4041 reflections) excluded from the refinement. e The rmsd for bonds and angles are the rmsd from ideal values.
immediately prior to the desired N-terminal residue using the following forward primers: 5′ CGG GAT CCA TCG AGG GAA GGA TGG ATT TTG CTG TGA TGA 3′, for proteins containing the original PFOMT N-terminus, and 5′ CGG GAT CCA TCG AGG GAA GGA TGG CAA CCA ACG AAG ATC 3′, for proteins with the N-terminal sequence derived from the M. sativa CCoAOMT. cDNAs were amplified using Isis Proofreading DNA Polymerase (Q-Biogene, Heidelberg, Germany) and ligated into pQE9 vectors using the BamHI and HindIII restriction sites, introduced by the PCR primers. All resultant DNA constructs were confirmed by sequencing (Applied Biosystems, Darmstadt, Germany). Functional expression Plasmids were transformed into the E. coli strain M15 (Qiagen) harboring the plasmid pRep4. Expression was induced after 150 min by the addition of 1 mM isopropyl thio-β-D-galactoside. The bacteria from 400 ml of culture were harvested after 180 min as described previously.18 Incubation times for loop and double hybrids were extended to 6 h at 37 °C and then induced with 1 mM isopropyl thio-β-D-galactoside and transferred to 4 °C and grown for 48 h to increase the yield of the soluble protein. Crude bacterial extract was prepared by ultrasonication and centrifugation (30,000g, 4 °C, 10 min). The clear supernatant was affinity purified as described previously,18 concentrated into 50 mM Tris/HCl, pH 7.5, and 10% glycerol, and the protein was quantified based on the calculated extinction coefficient (ε = 18,730) at 280 nm (Protean Software Package, DNASTAR, Madison, WI, USA). The purification was monitored by SDS/PAGE. Dimerization of the variants was confirmed using gel filtration (GE Healthcare, Superdex 200 16/60) with the following standards: chymotrypsin (17 kDa), ovalbumin (45 kDa), bovine serum albumin (67 kDa), aldolase (158 kDa), and catalase (240 kDa). M. sativa CCoAOMT M. sativa CCoAOMT DNA in the pET15b vector was a kind gift from Dr. Joseph P. Noel (Howard Hughes Medical Institute and Salk Institute for Biological Studies, La Jolla, CA, USA). This plasmid was transformed into E. coli strain BL21DE3 (Novagen, Darmstadt, Germany), and the protein was expressed and purified as described for PFOMT hybrids.
N-terminal and double hybrids
Factor Xa cleavage
The DNA coding for the N-terminal hybrid was prepared by amplification of the PFOMT cDNA using Platinum PCR SuperMix High Fidelity (Invitrogen) and the 92-bp primer containing the sequence coding for the first 22 amino acids from the M. sativa CCoAOMT. The DNA was cloned into pQE9 (Qiagen) using BamHI and HindIII restriction sites as described for the loop hybrid. The double-hybrid DNA fragment was prepared analogously to the N-terminal hybrid using the cDNA coding for the loop hybrid as template and 92 bp primer.
Proteins containing the Factor Xa cleavage site were rebuffered into Factor Xa reaction buffer (20 mM Tris/ HCl, pH 6.5, 50 mM NaCl, and 1 mM CaCl2). The cleavage reactions were conducted in 400 μl volume. The amount of protein was adjusted to 0.25 mg/ml. Two units of Factor Xa protease were used per reaction (400 μl). The reactions were incubated at 37 °C for 24 h to achieve complete cleavage. After incubation, the protease was removed using Factor Xa removal resin (Qiagen). The reactions were purified by Talon affinity resin (BD Biosciences, Heidelberg, Germany). The cleaved protein that did not bind to the matrix and eluted with the flow through was concentrated and rebuffered to 50 mM Tris/HCl, pH 7.5, and 10% glycerol and checked for enzymatic activity. Cleavage and purity were confirmed by SDS/PAGE. Western blot analysis was also performed with His-tag-
OMTs containing a Factor Xa cleavage site To investigate the possible influence of the N-terminal His tag, a Factor Xa cleavage site (IEGR) was introduced
Substrate Promiscuity in Plant OMTs specific antibodies (Qiagen) in order to verify the absence of any His tag. Enzyme activity tests and kinetic data Enzyme assays were performed in a buffer containing 100 mM K2PO4, pH 7.5, and 10% glycerol with 10 μM substrate (dissolved in 30% dimethyl sulfoxide), 0.5–2 μg of total protein, and 400 μM SAM in a total volume of 50 μl. The assays were incubated at 30 °C for 60 to 3600 s (dependent on the protein and substrate tested) and then stopped by the addition of 20 μl of 7% trichloroacetic acid in 50% acetonitrile/water. The reaction products were analyzed by HPLC as described previously.17 Caffeoyl glucose was prepared as described previously from caffeic acid and UDP-glucose with the purified recombinant sinapic acid glucosyltransferase from Brassica napus.32 CCoA was prepared based on published methods.33,34 Caffeic acid was obtained from Serva (Heidelberg, Germany), quercetagetin was obtained from Extrasynthese (Genay, France), and quercetin was obtained from Roth (Karlsruhe, Germany). The reaction products were analyzed by reversed-phase liquid chromatography on a Nucleosil 5-μm C18 column (50 mm length × 4 mm inner diameter; Macherey and Nagel, Düren, Germany), as described previously.17 Compounds were analyzed with linear gradients from 10% B (100% acetonitrile) in A (1.5% aqueous phosphoric acid) to 70% B in A (for phenolics), from 5% B to 50% B in A (for free acids and CoA esters), from 5% B to 30% B (for glucose esters), and from 20% B in A to 80% B in A in 4 min (for flavonoids) at a flow rate of 1 ml min− 1. Detection of flavonoids, catechol, coumarins, and hydroxycinnamic acid esters was performed between 260 and 400 nm. Identification and quantification was achieved with reference compounds from our Institute collection. For Km determination of methyl group acceptors, acceptor concentrations were chosen between 2 and 20 μM, while SAM was kept constant at 1.5 mM. Km, Vmax, and kcat values were calculated by nonlinear curve fitting, assuming Michaelis–Menten steady state kinetics. All enzyme assays were recorded in triplicate for each experiment. PDB accession code Coordinates and structure factors have been deposited in the PDB with accession code 3C3Y.
Acknowledgements We thank Jean-Luc Ferrer (Grenoble) and Joseph P. Noel (San Diego) for providing the M. sativa CCoAOMT plasmid and information prior to publication and Norbert Sträter (Leipzig) for the use of his X-ray facilities at the initial stages of this project. Diffraction data were measured on the MPG/GBF beamline BW6 at Deutsches Elektronen-Synchrotron, where we gratefully acknowledge the assistance of Gleb Bourenkov and Hans Bartunik. This work was in part supported by grants VO719/5 and STU297/4 from the Deutsche Forschungsgemeinschaft as well as the Schwerpunktprogramm SPP1152 ‘Evolution metabolischer Diversität.’
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