REVIEW www.rsc.org/npr | Natural Product Reports

Structural biology in plant natural product biosynthesis—architecture ofenzymes from monoterpenoid indole and tropane alkaloid biosynthesis†

Joachim Stockigt*a,b and Santosh Panjikarc

Received (in Cambridge, UK) 2nd August 2007First published as an Advance Article on the web 24th October 2007DOI: 10.1039/b711935f

Covering: 1997 to 2007

Several cDNAs of enzymes catalyzing biosynthetic pathways of plant-derived alkaloids have recentlybeen heterologously expressed, and the production of appropriate enzymes from ajmaline and tropanealkaloid biosynthesis in bacteria allows their crystallization. This review describes the architecture ofthese enzymes with and without their ligands.

1 Introduction2 Strictosidine synthase (STR1)2.1 Biological Pictet–Spengler reaction and the role of

STR1 for the entire indole alkaloid family2.2 Crystallization and structure determination of STR1,

and the “Auto-Rickshaw” software pipeline2.3 The 6-bladed 4-stranded b-propeller fold of STR1—the

first example from the plant kingdom2.4 Structures of STR1 in complex with its substrates and

product2.4 Substrate specificity of STR1 and mutagenesis studies2.6 Structure-based alignments and evolution of STR13 Strictosidine glucosidase (SG) and its role3.1 Optimization of crystallization of SG3.2 Overall 3D-structure of SG3.3 Site-directed mutagenesis and SG

Glu207Gln-strictosidine complex3.4 Recognition of the aglycone and glycone part of

strictosidine3.5 Substrate binding site of SG compared to those of

other plant glucosidases3.6 Prospects for the use of SG for alkaloid synthesis4 Polyneuridine aldehyde esterase (PNAE)4.1 PNAE—one of the most substrate-specific esterases?4.2 Inhibition, primary structure and site-directed

mutagenesis of PNAE4.3 The long route to PNAE crystals4.4 Homology modelling and structure of PNAE5 Vinorine synthase (VS)5.1 The role of VS in generating the ajmalan structure5.2 Inhibitors and mutants of VS5.3 Unusual crystallization conditions and instability of

VS crystals5.4 Structure elucidation of VS

aCollege of Pharmaceutical Sciences, Zijingang Campus, Zhejiang Univer-sity, 310058, Hangzhou, ChinabInstitute of Pharmacy, Johannes Gutenberg University Mainz, StaudingerWeg 5, D-55099, Mainz, GermanycEuropean Molecular Biology Laboratory Hamburg, Outstation DeutschesElektronen-Synchrotron, Notkestrasse 85, D-22603, Hamburg, Germany† In memory of Professor Pierre Potier.

5.5 Important amino acids and mechanistic aspects5.6 Significance of the vinorine synthase structure for the

BAHD enzyme family6 Raucaffricine glucosidase (RG), a side route of the

ajmaline pathway6.1 Crystallization attempts and the 3D-structure of RG7 Structure-based redesign of enzymes and applications

in chemo-enzymatic approaches8 Other structural examples from the alkaloid field9 Conclusions and future aspects of structural biology in

the alkaloid field10 Acknowledgements11 References

1 Introduction

Both their highly complex chemical structures and their pro-nounced pharmacological activities have made research on al-kaloids, including elucidation of their biosynthetic pathways, veryattractive for many decades. Major progress in the research intothe biosynthesis of plant monoterpenoid indole alkaloids has beenmade by investigations of single enzymes, partial biosyntheticroutes and entire pathways.1–3 This has been sporadically reviewedduring the last 10 years together with the genetics of alkaloidbiosynthesis.4,5

There are, however, only very few examples from the alkaloidfield, which deliver at present a coherent knowledge of biosyntheticroutes at the enzyme level. These include:

(a) The well investigated enzymatic steps connecting the Aspi-dosperma alkaloids tabersonine and vindoline, reactions occurringat the periphery of the basic Aspidosperma alkaloid skeleton.4,6–10

(b) The biosynthesis of camptothecin, our knowledge of which,despite some recent enzymatic work on the early steps,11 comeslargely from Hutchinson’s group, a long time ago.12,13

(c) The whole sequence between tryptamine plus secologaninand the heteroyohimbines ajmalicine and some of its isomers(Corynanthe alkaloids) was successfully investigated at the enzy-matic level a few decades ago.14–17

(d) Efficient alkaloid-producing cell suspension cultures ofthe Apocynaceae plant Catharanthus roseus, established at the

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Joachim Stockigt received a PhD in organic chemistry from Munster University (Germany) with Professor Burchard Franck. He hasworked with Professor Meinhart H. Zenk at the Faculty of Biology (Bochum University, Germany) and the Faculty of Pharmacy (MunichUniversity, Germany) and is currently Full Professor at the Institute of Pharmacy (Mainz University, Germany; College of PharmaceuticalSciences, Zhejiang University, Hangzhou, China). His research interests include natural products biosynthesis (phytochemistry, enzymology,molecular and structural biology).

Joachim Stockigt Santosh Panjikar

Santosh Panjikar earned a PhD in Biotechnology from Friedrich-Schiller University(Jena, Germany) and held a postdoctoral appointment at the EMBL HamburgOutstation with Dr Paul A. Tucker. He is currently Staff Scientist at the EMBLOutstation. His research interests focus on structure-based drug design, methoddevelopments in structural biology and synchrotron instrumentation.

beginning of the 1970s in Zenk’s laboratories, became the key forstudies of this plant.18,19

(e) Cell suspension cultures of the Indian medicinal plantRauvolfia serpentina Benth. ex Kurz from the same lab were alsothe major prerequisite for the isolation of some uncommon indolealkaloids20 and elucidation of the pathway for the formation ofthe antiarrhythmic ajmaline and structurally related alkaloids ofthe sarpagan- and ajmalan-type both by identification of manysingle enzymes21–23 and in part by in vivo NMR.24,25 The numberof enzyme-catalyzed reactions proven in that Rauvolfia cell systemamounts to more than 15, representing at the moment one of themost detailed investigations in alkaloid biosynthesis. The pathwaybetween tryptamine plus secologanin and ajmaline is illustrated inScheme 1.

Several of the cDNAs coding for enzymes of the above-mentioned ajmaline route have been detected, isolated and func-tionally expressed, mostly by using the “reverse genetic” approach,applying polymerase chain reaction (PCR) and heterologoussystems, especially Escherichia coli. For instance, most of thesoluble Rauvolfia enzymes are now functionally expressed andcharacterized in detail,23 especially with regard to their substratespecificity and their major kinetic data and general properties, suchas temperature and pH optimum, molecular size, and isoelectricpoints. For the first time, cloning has also provided amounts ofenzyme, in almost pure form, on the 20–50 mg scale by fusion-protein techniques. This is the amount necessary to developcrystallization conditions of these proteins and to elucidate theirthree-dimensional structure by X-ray analysis.

The following article summarizes in detail the achievementsin the elucidation of the architecture of five major enzymesof the ajmaline biosynthetic pathway in Rauvolfia,26 namelystrictosidine synthase (STR1), strictosidine-O-b-D-glucosidase(SG), polyneuridine aldehyde esterase (PNAE), vinorine synthase(VS) and raucaffricine glucosidase (RG), together with twotropinone reductases (TR-I and TR-II) from tropane alkaloidbiosynthesis.27,28 Other than a short general overview on 3D-

analysis of Rauvolfia enzymes,26 this is the first comprehensiveand detailed review on structural biology in the alkaloid field,providing a more direct insight into the mechanisms of alkaloidbiosynthesis.

2 Strictosidine synthase (STR1)

2.1 Biological Pictet–Spengler reaction and the role of STR1 forthe entire indole alkaloid family

Strictosidine synthase (STR1, EC 4.3.3.2), which catalyzes thestereoselective Pictet–Spengler reaction of tryptamine and secolo-ganin to form 3a(S)-strictosidine, has been described many timessince its original detection.29,30,31–34 It was the first enzyme fromalkaloid biosynthesis whose cDNA was functionally, heterolo-gously expressed.35–37 Later, the synthases from several plants otherthan Rauvolfia,35 such as Catharanthus38,39 and Ophiorhiza,40 werefunctionally expressed. The primary importance of STR1 is notonly its precursor role for the biosynthetic pathway of ajmaline, butalso because it initiates, in fact, all pathways leading to the entiremonoterpene indole alkaloid family. Some prominent membersare depicted in Fig. 1. Although STR1 is the crucial enzyme for allthe 2000 monoterpenoid indole alkaloids, little information aboutits reaction mechanism and the critical amino acids for enzymeactivity was known.

2.2 Crystallization and structure determination of STR1, and the“Auto-Rickshaw” software pipeline

Expression and crystallization of STR1 was straightforward andtypical of our crystallization of other Rauvolfia proteins, usingpQE-2-plasmid and M15 E. coli cell line. The hanging-dropvapour-diffusion technique was used routinely and was the mostefficient method in our hands. STR1 crystals were in spacegroup R3 with a hexagonal unit cell containing two moleculesof the enzyme. A set of selenium-labelled STR1s (4SeMet- and

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Scheme 1 Enzyme-catalysed biosynthesis of ajmaline in cell suspension culture of the medicinal plant Rauvolfia serpentina (L.) Benth. ex Kurz.Abbreviations are: STR1 = strictosidine synthase, SG = strictosidine glucosidase, SBE = sarpagan bridge enzyme, PNAE = polyneuridine aldehydeesterase, VS = vinorine synthase, VH = vinorine hydroxylase, CPR = cytochrome, P450 reductase, VR = vomilenine reductase, DHVR =dihydrovomilenine reductase, AAE = acetylajmalan esterase, NAMT = norajmalan methyltransferase, RG = raucaffricine glucosidine, VGT = vomilenineglucosyltransferase. Strictosidine aglycone has not been isolated. Enzymes shown in bold have been heterologously expressed, those marked with asteriskshave been crystallised and their 3D-structures determined recently in our laboratories. They are described in this review.

Fig. 1 Formation of strictosidine and its biosynthetic role as centralprecursor of various monoterpenoid indole alkaloids.

6SeMet-STR1) needed to be prepared in order to solve the enzymestructure, because no protein structures with significant sequencehomology to strictosidine synthase were available.41,42 The STR1

structure was finally solved using the multiple wavelength anoma-lous dispersion (MAD) approach together with software packageAuto-Rickshaw,43 and refined to 2.4 A.

Structures of all enzymes from the ajmaline biosyntheticpathway discussed in this article were determined using Auto-Rickshaw. In fact, early evaluation of the software pipeline wasperformed on several multiple wavelength diffraction (MAD)datasets from crystals of various proteins, including MAD dataof vinorine synthase (VS) and strictosidine synthase (STR1). Ata later stage, native structures or ligand complexes of the variousenzymes from the pathway were determined routinely using Auto-Rickshaw. Here we describe briefly the software pipeline whichprovides a means of rapid structure solution of proteins.

Crystal structure determination both by isomorphous replace-ment and by anomalous scattering techniques is a multi-stepprocess in which each step, from substructure determination tomodel building and validation, requires certain decisions to bemade. These decisions comprise the choice of the crystallographiccomputer programs that are most suitable to perform the specifictasks and the optimal input parameters for each of these programs.The interpretability of the map depends to a large extent on thesuccess of the preceding steps and is generally limited by theresolution of the data and the quality of the phase information.Traditionally, each of the steps described was carried out by an ex-perienced crystallographer, whose skill manifested itself in finding

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the optimum, or at least a successful, path towards the completionof the structure determination. The automated crystal structuredetermination platform combines a number of macromolecularcrystallographic software packages with several decision-makingsteps. The entire process in the pipeline is fully automatic. Eachstep of the structure solution is governed by the decision makingmodule within the platform, which attempts to mimic the decisionsof an experienced crystallographer. The role of computer-codeddecision-makers is to choose the appropriate crystallographiccomputer programs and the required input parameters at eachstep of the structure determination. Various phasing protocols areencoded in the system. These are single anomalous diffraction(SAD), single isomorphous replacement with anomalous scatter-ing (SIRAS), multiple wavelength diffraction (MAD), standardmolecular replacement (MR), phased MR and combination ofMR and SAD. A large number of possible structure solutionpaths for each phasing protocol are encoded in the system, and theoptimal path is selected by the decision-makers as the structuresolution evolves.

Once the input parameters are given (number of amino acids,heavy atoms, molecules per asymmetric unit, probable space groupand phasing protocol) and X-ray data have been input to Auto-Rickshaw, no further user intervention is required. It proceeds stepby step through the structure solution using the decision makers.The Auto-Rickshaw server (http://www.embl-hamburg.de/Auto-Rickshaw) is available to EMBL-Hamburg beamline users, andthe server will be made more widely accessible in the immediatefuture.

2.3 The 6-bladed 4-stranded b-propeller fold of STR1—the firstexample from the plant kingdom

The overall structure of STR1 belongs to the six-bladed four-stranded b-propeller fold, where the blades are radially locatedaround a pseudo-six-fold symmetry axis. Each blade consists oftwisted four-stranded b-sheets. Although this particular fold hasbeen detected in different organisms several times, this is the firstexample from the plant kingdom. However, the other enzymeswith the six-bladed fold have a completely different function.44,45

The active site of STR1 is near to the symmetry axis, as shown bythe structure of the enzyme–tryptamine complex (Fig. 2).46 Thereare only three helices in the STR1 structure, of which two areconnected by an S–S bridge, which is essential for both the overallstructure and the shape of the catalytic centre.

2.4 Structures of STR1 in complex with its substrates andproduct

Tryptamine is located deep in the binding pocket, sandwichedbetween two hydrophobic residues (Tyr151 and Phe226) holdingthe molecule in the correct orientation for the Pictet–Spengler con-densation. In addition, Glu309, which is important for the STR1activity as proven by mutagenesis experiments, is coordinated withthe amino group of tryptamine. There are (primarily) hydrophobicamino acids lining the binding pocket, and both positively chargedand hydrophobic residues are located at the entrance to the bindingsite of STR1.

The complex structure of STR1 bound with secologanin exhibitsthe monoterpene in the same binding pocket (Fig. 3a). The

Fig. 2 Strictosidine synthase (STR1) from Rauvolfia in complex with itssubstrate tryptamine (N and C mark the N- and C-termini of the protein;the arrow points to the S–S bridge).

hydrophilic glucose unit of secologanin points out of the catalyticpocket towards the solvent. This complex is illustrated in Fig. 3b,which is rotated by 84◦ along the x-axis to show better theaccessibility of the hydrophilic part of the monoterpene to thesolvent. The aldehyde group of secologanin points towards Glu309and is in close proximity to the amino group of tryptamine(as observed in the previous structure), ready for the primarycondensation reaction.

The third complex was prepared by soaking crystals of STR1 ina solution containing strictosidine, and the structure illustrateshow the product of the STR1 reaction is accommodated inthe catalytic centre (Fig. 4). The location of the strictosidine isvery similar to that found for both substrates (see above) butis not totally identical, showing that the way the ligands areaccommodated is to some extent flexible before and after thereaction. The electron density gives additional information abouthow the product is shielded from the residues of the active centre.Such shielding might have important implications for the substratespecificity of STR1 and should help in structure-based rationaldesign of substrate acceptance.

2.4 Substrate specificity of STR1 and mutagenesis studies

As far as the substrate specificity of STR1 is concerned, severalstudies with both STRs from Catharanthus and Rauvolfia havebeen reported earlier.31,46–51 We have analyzed the substrate ac-ceptance of STR1 again and focused especially on explanationsof why particular tryptamine derivatives were not accepted bySTR1 based on the available structural information. Table 1highlights the comparison of substrate specificity of STR1,especially for tryptamines substituted differently at position 5.These tryptamines are not accepted in the case of 5-methyl- and5-methoxy-derivatives by the wild-type enzyme.52 The complexstructure demonstrates that the Val208 side-chain is shielding the

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Fig. 3 (a) Strictosidine synthase (STR1) in complex with its substratesecologanin. (b) Fig. 3a rotated by 84◦ around the x-axis.

5-position, preventing binding of 5-substituted indole bases. Re-placement of this particular Val by the smaller Ala clearly broadensthe substrate acceptance for 5-substituted tryptamines, deliveringnovel strictosidine analogues.52 Further structure-derived muta-tions based on this structure are expected to have significant impacton future enzymatic synthesis of new strictosidines structurallymodified at the tryptamine or secologanin parts.

Moreover, if the enzyme strictosidine glucosidase (SG), whichfollows STR1 in the biosynthesis of all the monoterpene alkaloids,can be modified in a similar way to that described for the synthase,future chemo-enzymatic approaches might be developed for thegeneration of novel alkaloid libraries with biologically importantcompounds.

2.6 Structure-based alignments and evolution of STR1

STR1 has no functional homologies to other six-bladed b-propeller folds. Also, a structurally based sequence alignment of

Fig. 4 3D-structure of STR1 in complex with its product strictosidine.

Table 1 Substrate acceptance of STR1 for differently substitutedtryptamines at positions 5 and 6 (n.d. = not detectable, data taken inpart from ref. 52)

Substrates Enzymes Km/mM (kcat/Km)/mM−1 s−1

Tryptamine Wild-type 0.072 147.92Val208Ala 0.219 246.99

5-Methyltryptamine Wild-type n.d. —Val208Ala 0.281 23.35

5-Methoxytryptamine Wild-type n.d. —Val208Ala 3.592 22.18

6-Methyltryptamine Wild-type 0.393 5.90Val208Ala 0.762 14.37

6-Methoxytryptamine Wild-type 0.962 5.53Val208Ala 0.307 54.27

5-Fluorotryptamine Wild-type 0.259 144.63Val208Ala 1.302 16.31

6-Fluorotryptamine Wild-type 0.136 171.84Val208Ala 0.356 38.29

5-Hydroxytryptamine Wild-type 2.255 249.29Val208Ala 0.844 21.47

these folds showed less than 16% sequence identity with most b-propellers such as diisopropylfluorophosphatase (DFPase)45 fromLoligo vulgaris, brain tumour NHL domain,53 serum paroxonase44

and low-density lipoprotein receptor YWTD domain. Analysis ofthe STR1 fold compared to the just-mentioned folds indicates thatthese structurally related proteins are also related from the pointof view of their evolution.46 Most probably they all evolved froman ancestral b-sheet gene. The ancestral structure might have beenmodified by e.g. deletions or insertions during evolution, leading toa differently shaped reaction pocket in order to adopt the differentfunctions of these propellers. These propellers keep a commonsequence homology which maintains the overall structural featuresof the protein fold. It is typical of these six-bladed b-propellersthat they are diverse in sequence and function and that they are ofdifferent phylogenetic origin.

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A similar reaction to that of STR1 found in the plant Alangiumlamarckii Thw. (Alangiaceae) is catalyzed by deacetylipecosidesynthase (DIS). This synthase catalyzes the condensation betweendopamine and secologanin, forming (1R)-deacetylipecoside whichhas the opposite configuration to strictosidine (at C-3).54 DISand STR1 are similar with respect to their reaction type (Pictet–Spengler condensation), their pH optimum, temperature optimumand their molecular size. Both enzymes exhibit high substratespecificity but they come from different plant species and families,and exhibit different substrate specificity and stereo-selectivity. Itcannot be excluded, however, that both synthases evolved from thesame ancestor but during evolution developed different substratespecificities leading to different alkaloid types. Cloning of DIS andcomparison of the 3D-structures of both enzymes could help tosupport this hypothesis.

3 Strictosidine glucosidase (SG) and its role

Similarly to STR1, strictosidine glucosidase (SG, EC 3.2.1.105)has been described several times from Catharanthus and fromRauvolfia.55–57 Here we refer mainly to the Rauvolfia enzymebecause it was the second protein from the ajmaline biosyntheticpathway to be crystallized. The general and major importance ofSG lies in its function to chemically “activate” strictosidine bydeglucosylation. The aglycone thus generated is highly unstableand reactive. It enters all the pathways to the various structuraltypes illustrated in Fig. 1, but the exact molecular stage atwhich it initiates all these pathways has not been clarified so far.Although glycosides are involved in many metabolic processessuch as regulation of plant hormone activity and biosynthesis58,59

and lignification,60 one of the main functions of glycosidases,especially in higher plants, is believed to be defence-related.61 Forexample, the generation by glycosidases of toxic products suchas cyanide or isothiocyanates from the appropriate glycosidesmakes their ecological significance as defensive agents immediatelyunderstandable.62 SG has also been discussed as being defence-related since its product(s) were shown to have antibiotic activity.56

However, in view of the important role that the aglycone ofstrictosidine plays as the biogenetic precursor of all of the 2000 orso monoterpenoid indole alkaloids, a synthesis-related functionof SG seems to be more important than its defence-related role.

Reasons for the pronounced substrate specificity and detailsof the reaction mechanism of this enzyme remained mostlyunknown until recently. Therefore, an efficient over-expression andpurification system for strictosidine glucosidase in Escherichia coliwas developed, followed by crystallization and preliminary X-rayanalysis of the enzyme.

3.1 Optimization of crystallization of SG

When our “routine” expression system (E. coli, M15 [pREP4])and purification method (fusion protein with N-terminal His6-tag), followed by removing the tag with dipeptidyl aminopeptidase(DAPase) and final enzyme purification by MonoQ ion exchangechromatography was used, about 10 mg of pure SG were obtainedfrom a 5 litre culture of E. coli. Such an amount of enzymeis acceptable for a large screening of crystallization conditions.More than 10 commercially available crystallization kits weretried. Many conditions resulted in formation of rather flat- or

needle-shaped crystals (Fig. 5a), which were not useful for X-ray experiments. However, in a large trial using a Cartesiansyn QUADTM Microsys crystallization robot with 600 nl sittingdrops, two conditions were found to produce rectangular crystalplates. These conditions gave, after extensive optimization withthe “hanging drop” method, improved crystals for X-ray mea-surements with a final resolution of 2.48 A (Fig. 5b).63

Fig. 5 (a) Needle-type crystals of SG not suitable for X-ray analysis.(b) A rectangular prism of SG (images are from ref. 26).

3.2 Overall 3D-structure of SG

From initial alignment studies SG unequivocally belongs to theglycosyl hydrolase (GH) family 1, which is grouped together with16 other families in clan GH-A, representing the biggest of the13 clans of glycosidases. There are only seven three-dimensionalstructures of glucosidases from family 1 known from eukaryoticsources, and six of these are of plant origin. The expression,purification, crystallization and preliminary X-ray analysis of SGhas recently been reported.63 In Fig. 6 the overall fold of SG

Fig. 6 “Tim barrel” overall fold of SG (2.48 A).

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is shown, representing the well known (b/a)8 barrel fold. Thestructure is built from 13 a-helices and 13 b-strands; the core ofthe structure consists of 8 parallel b-strands that form a b-barrel.This barrel is surrounded by eight helices and hosts the catalyticbinding site of SG for the substrate strictosidine.

3.3 Site-directed mutagenesis and SG Glu207Gln-strictosidinecomplex

Complete conservation of amino acid residues Glu207, Glu416and His161 is indicated by sequence alignment of SG withglucosidases of various origins. Glu207 is the proton donor thatassists nucleophilic attack of Glu416 at the anomeric carbon C-1.64

Hydrolysis of the glucoside bond is catalyzed in concert by bothglutamic acids.65 For SG no enzyme activity was obtained aftermutations Glu207Gln, Glu207Asp, Glu416Gln or Glu416Asp.66

This clearly points to the crucial role of both glutamates for thedeglucosylation of strictosidine. In the reaction catalyzed by SG,His161 also plays an important role. When it is replaced by Asnor Leu, enzyme activity is decreased to <0.1%. These results werethe basis for the assumption that the three mentioned residuesmust be involved in the reaction mechanism of strictosidinedeglucosylation. For this reason, they should be located near thebinding pocket or even belong directly to the catalytic centreof SG, which is, in fact, what is shown by the 3D-structure.The structure of the complex of inactive mutant SG-Glu207Glnwith strictosidine was generated (Fig. 7) to obtain additionalinformation on the role of other individual amino acids ofthe binding pocket.66 Since there was no detectable catalyticactivity of the Glu207Gln mutant, soaking was possible with thenatural substrate strictosidine and resulted in crystals exhibiting aresolution of 2.82 A.66

Fig. 7 Complex structure of inactive mutant SG-Glu207Gln withstrictosidine.

3.4 Recognition of the aglycone and glycone part of strictosidine

For both the glycone part and aglycone part of the substratemolecule, the electron density allowed detailed description of the

catalytic centre of SG in the substrate binding state. The bindingpocket is located at the top of the barrel. The base of the pocketcontains several charged residues whereas the gate to the pocketis hydrophobic in nature. The glycosidic part of strictosidine islocated deep in the catalytic pocket, and the indole part pointstowards the solvent even though this part of the substrate isstrongly hydrophobic. The aglycone part is surrounded by mostlyhydrophobic residues. The sugar unit of strictosidine interacts withmany hydrophilic amino acids. His161 is not a catalytic aminoacid—it is located too far away from the anomeric C-atom ofthe glucose unit—but it helps to bind strictosidine in the correctorientation for the deglucosylation process.

3.5 Substrate binding site of SG compared to those of other plantglucosidases

The structure of SG-Glu207Gln from R. serpentina in complexwith strictosidine can be compared with the structures of two otherplant glucosidase–substrate complexes, ZmGlu1-Glu191Asp fromZea mays in complex with DIMBOA-b-D-glucoside (PDB code1E56), and Dhr1-Glu189Asp from Sorghum bicolor in complexwith dhurrin, (PDB code 1V03) (Fig. 8).66–68 Ten out of the elevenamino acids that participate in the binding of the glucose unitare conserved in the sequences of the glucosidases in family 1.In some glucosidases only, Tyr481 is replaced by Phe, indicatingthe pronounced conservation which seems to keep the differentglycosylated compounds in the correct location for the hydrolysisreaction.

The aglycone binding site of family 1 members is much morevariable. The most striking difference in their active sites isobserved for the amino acid Trp388. This residue is conservedwithin the family 1 and its role in substrate recognition hasbeen discussed previously.69 Tryptophan shows in all known plantglucosidases its v-angle (the angle about the side chain Ca–Cbbond) of ∼60◦. That is obviously the typical conformation forplant glucosidases. In SG, this v-angle is changed to −180◦.Such a change in conformation results in more space withinthe binding pocket of SG. This additional space is needed toaccommodate the large substrate strictosidine, compared to thesmaller substrates of the other b-glucosidases. Moreover, Trp388forms a stacking interaction with the indole part of strictosidine.Due to the conformational change of Trp388, some amino acids(Gly386, Met275, Thr210 and Met297) can better interact withstrictosidine. Together with Phe221 and Trp388, all these aminoacids form the substrate binding site of SG for the indolic part ofthe substrate strictosidine.

The specific conformation of Trp388 of SG has been untilnow an exception amongst b-glucosidases. Structural comparisonwith fifteen glucosidases revealed three different conformationsfor this amino acid: (a) Trp points directly into the bindingpocket, which is found for glycosidases from bacteria and archaea;(b) Trp points in the direction of the N-terminus (which is theplant-typical conformation); (c) Trp points in the direction of theC-terminus, which is the SG Trp conformation.66 All plant-derivedglucosidases are folded in a similar way and the major differenceis the conformation of Trp388. If additional examples of this newTrp388 conformation are discovered in the future, b-glucosidasesmight be sub-divided based on this feature.

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Fig. 8 Comparison of the substrate complex of SG (a) with those of twoother b-glucosidases, from Zea mays (b) and from Sorghum bicolor (c),67,68

displaying the different conformation of tryptophan (W376) in SG. Thisillustration was partly taken from ref. 66 (hydrophobic residues in grey,hydrophilic in green, acidic residues (Glu, Asp) in red, positively chargedresidues in blue).

3.6 Prospects for the use of SG for alkaloid synthesis

A future demand for natural products, in this case of complexplant alkaloids with high structural diversity, will probably depend

on detailed biosynthetic knowledge, on rational molecular engi-neering of the enzymes, their synthetic application and metabolicengineering of the medicinal plants.70,71 Because of the special andmultiple role of the aglycone of strictosidine, the enzyme SG mayserve as an attractive candidate for such directed biosynthesisapproaches. Such approaches only make sense if the substratespecificity of SG can be modified. Because the three-dimensionalstructure of strictosidine synthase has become available, initialsteps were successful to engineer and to change the substrateacceptance of STR1.52 Generation of large alkaloid libraries bystructure-based enzyme re-design of STR1 and SG in combinationwith biomimetic approaches may therefore become possible in thenear future.47,51,52

4 Polyneuridine aldehyde esterase (PNAE)

4.1 PNAE—one of the most substrate-specific esterases?

Polyneuridine aldehyde (PNA) is the first alkaloid in the pathwayto ajmaline, exhibiting a sarpagan structure.72 The aldehydeis biosynthetically generated by the so-called sarpagan-bridgeenzyme (SBE),73 whose mechanism has not been elucidated so far.The esterase PNAE (EC 3.1.1.78) converts PNA after ester hydrol-ysis and decarboxylation into 16-epi-vellosimine.71,74 The substratespecificity of this enzyme has been investigated several times atvarious stages of enzyme purity,72,74,75 and was finally determinedwith the overexpressed homogenous His6-tag PNAE. From a seriesof 13 structurally different esters only the natural substrate (PNA)was hydrolyzed (Fig. 9).76–78 The esters included ten methyl esters,of which eight were monoterpenoid indole alkaloids and two moresimple methyl esters, and additionally three acetate esters. Theresults demonstrated an exceptionally high substrate specificity ofthe over-expressed enzyme and indicated that PNAE might be oneof the most substrate-specific esterases.

4.2 Inhibition, primary structure and site-directed mutagenesis ofPNAE

Inhibition studies with selective serine or cysteine inhibitorsand unselective histidine/cysteine inhibitors provided insight intoreactive residues of the enzyme (Table 2). They showed completeinhibition with diethyl pyrocarbonate, indicating the importanceof histidine for enzyme activity, and partial inhibition was ob-served with serine/cysteine inhibitors. However, these experimentsdid not allow PNAE to be defined as a serine or cysteinehydrolase. Site-directed mutagenesis was applied to get moredetailed information on particular amino acids that seemed likelyto be involved in the catalytic process of PNAE from a rigorousanalysis of the primary structure of the enzyme. Comparison of

Table 2 Inhibition of PNAE by serine, cysteine, and histidine inhibitors

Inhibitor Type of Inhibition Concentration Incubation time/min Relative inhibition (%)

AEBSF Selective Ser 4.0 mM 60 0E-64 Selective Cys 25 lM 60 0TPCK Ser-Cys 200 lM 60 12PMSF Ser-Cys 1.0 mM 60 20DEPC Unselective His 1.2 mM 60 100Hg2+ Unselective Cys 200 lM 60 100

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Fig. 9 Structurally different esters as putative substrates of PNAE. Only the natural substrate, polyneuridine aldehyde, displayed any activity.78

the overall amino acid sequence of PNAE showed relatively highidentity (up to 50%) to some putative lyases e.g. from Arabidopsisthaliana.79 A 40–43% identity was found for two well characterizedhydroxynitrile lyases (Hnls), one from Hevea brasiliensis (HbHnl)and one from Manihot esculenta (MeHnl)80–87 with known 3D-structure, demonstrating a relatively close relationship of PNAEto Hnls (Scheme 2). The above enzymes belong to the a/b-foldhydrolases, indicating that PNAE is a newly-detected member ofthat enzyme superfamily. The most important features of thatfamily are a catalytic triad formed by a nucleophilic residue, anacidic amino acid and a histidine, which appear in the same orderin PNAE (Ser87, Asp216, His244), and two additional stronglyrelated and conserved motifs. Indeed, mutations in PNAE showed

the presence of the catalytic triad, since replacement of each of thethree residues by alanine resulted in completely inactive mutants,76

classifying the enzyme as a novel a/b-fold hydrolase. Structuralanalysis of PNAE (see Section 4.4) supported that classification.

4.3 The long route to PNAE crystals

The key to the crystallization of an enzyme is its purity. Crystal-lization also depends on the availability of homogenous, ideallymonodisperse solutions of protein molecules with a uniform,rigid three-dimensional fold. Protein crystallization can be dividedinto two steps: coarse screening to identify initial crystallizationconditions and then optimization of these conditions to produce

Scheme 2 Alignment of the primary sequence of PNAE with hydroxynitrile lyases, showing the close relationship of the three enzymes, the conservedcatalytic triad (marked with asterisks) and two typical motifs (black on red) around His17 and Ser87.

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single, diffraction-quality crystals. Currently, there are no system-atic methods to ensure that ordered three-dimensional crystals willbe obtained.88 Sitting-drop and hanging-drop methods of vapourdiffusion are the most commonly used techniques. Microbatchcrystallization under oil is mainly used when these crystallizationmethods have failed.89 A recent development in protein crystal-lization has been the use of microfluidic systems for crystallizingproteins using the free-interface diffusion method on the nanolitrescale, but this has one major drawback in that it is often difficultto translate hits into higher volume solutions in order to growcrystals for diffraction experiments.

Crystal optimization aims to turn poor quality crystals intodiffraction-quality crystals that can be used for structure deter-mination. There are a variety of methods that can be used toimprove crystal quality, including crystal seeding. The nucleationevent in protein crystallization is a poorly understood process.In many crystallization experiments, it is not possible to reachsufficiently high levels of saturation for nucleus formation. In manycases, the introduction of a crystal or crystal seed stock at lowerlevels of saturation can facilitate nucleation and crystal growth.Micro-, macro-, heterogeneous and in situ seedings representdifferent seeding techniques. High-throughput crystallization andvisualization platforms have been widely established and are com-monly used by high-throughput structural genomics initiatives.90–93

Crystallization experiments can be monitored using an imagingrobot and the images can either be analyzed manually or usingautomatic crystal recognition systems. Such a kind of systemhas been installed at EMBL Hamburg, and the high-throughputcrystallization facility has been open to the general scientificcommunity.92 The facility covers every step in the crystallizationprocess from the preparation of crystallization cocktails for initialor customized screens to the setting up of hanging-drop vapour-diffusion experiments and their automatic imaging.

Obtaining useful PNAE crystals was our most tedious exampleof plant protein crystallization because it took in total four yearsto obtain good X-ray-diffracting crystals, although the entire timeperiod was not invested in this particular enzyme. The two majorobstacles were: (a) precipitation of PNAE in crystallization solu-tion and (b) the complete lack of crystal (nucleus) formation whennearly all of the commercially available crystallization kits wereapplied. Most of the typical crystallization procedures mentionedabove were used. In a total about 6000 crystallization conditionswere tried, with very few conditions resulting in crystal formation.Only “2-dimensional” crystals were obtained, which were uselessfor X-ray analysis. These conditions then were optimized in time-consuming trials by “hand-pipetting” of 4–8 ll hanging drops.From about 1000 drops, 10 crystals were finally detected, fromwhich the best gave a resolution of 2.0 A and allowed immediatestructure elucidation of the enzyme. Meanwhile, the success ratein crystallizing this particular enzyme increased and probably willalso become routine soon.

4.4 Homology modelling and structure of PNAE

The structure of PNAE was modelled76,77 based on the 3D-structure of hydroxynitrile lyase (EC 4.2.1.39) from Heveabrasiliensis, which was available at 1.9 A resolution (see Proteindatabank 1YAS) and which has an amino acid sequence identityto PNAE of 43%. Homology modelling gave a range of PNAE

models with the software MODELLER (version 4).94 The modelwith the best overall steric properties was chosen for comparisonwith the 3D-structure of PNAE generated from X-ray data.

The 3D-structure of His6-PNAE (resolution of 2.1 A) waselucidated by the “molecular replacement” method with thesoftware “Auto-Rickshaw”.43 The model of the salicylic acidbinding protein 2 (SABP2, pdb code: 1XKL) from Nicotianatabacum95 with identity of the amino acid sequence to PNAEof 54% was successfully used as search model for molecularreplacement. Sufficient electron density was not observed for theHis6-tag or for the first eight amino acids from the N-terminalend of the enzyme because of high flexibility. The fold of PNAEbelongs to the a/b-hydrolase family, which is one of the biggestand fastest growing enzyme families with about 4250 members.96,97

Based on the 3D-structure, PNAE indeed represents a novelmember of that family. The enzyme consists of eight b-strandsand eight a-helices (Fig. 10).

Fig. 10 Overall structure of PNAE highlighting the “cap-region”, whichis shown by the arrow.

The two anti-parallel strands and the three helices (see sec-ondary structure above, the straight line in Fig. 10) form a domainwhich is named the “cap region”. The connection of all thesestructural elements is illustrated by the 2D-topological diagram(Fig. 11). The three amino acids (Ser87, Asp216 and His244)that form the catalytic triad and are essential for enzyme activity,as shown by mutation experiments, are included in Fig. 11.77

The enzyme structure illustrates that these residues are locateddirectly in the reaction channel. Ser87 is positioned in the so-called “nucleophilic elbow” which is located between the b3 sheetand a3 helix. The narrow reaction channel connects the bindingsite with the surface of the enzyme, and the channel is flanked bypredominantly hydrophobic residues of the “cap-domain”. Thisdomain consists of a total of 71 amino acids from Asp116 toPhe187. In accord with the non-polar structure of the substratePNA, about 50% of the “cap” consists of hydrophobic residues.Most of the a/b-hydrolase enzymes exhibit this domain, which isinvolved in substrate recognition and interfacial activation.98 The“cap-region” has, in contrast to the core-region of these enzymes,

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Fig. 11 2D-topological diagram of PNAE illustrating the connection ofthe structural elements (“cap domain” at bottom).

a spectacular variability in topology, leading to the surprisingvariability in functions and to the substrate selectivity, which isespecially pronounced for PNAE.

Comparison of the overall structure of modelled PNAE withthe X-ray structure gave a root mean square deviation (rmsd)value of 0.916 A. For the “cap region” it was 1.045 A, and the“core region” exhibited a rmsd of 0.812 A. The “core region”,the conserved region in the a/b-hydrolase family, shows the lowerrmsd, whereas the flexible and variable “cap domain” differs morebetween the two models (rmsd of 1.045 A).

Comparison of the catalytic amino acids Ser87, Asp216 andHis244 shows only slight differences between the three X-ray

models of PNAE, HbHnl and the salicylic acid binding protein.The entrance to the channel is represented in Fig. 12 andshows clear differences between the modelled and the X-ray-derived structure of PNAE, indicating the necessity of the X-rayanalysis. There is a future need to generate appropriate substrate–enzyme complexes to understand more precisely the extraordinarysubstrate specificity of the enzyme and especially to get to knowdetails about the amino acids involved in the recognition of thesubstrate PNA.

5 Vinorine synthase (VS)

5.1 The role of VS in generating the ajmalan structure

On the pathway to the ajmalan basic skeleton, vinorine synthase(VS, EC 2.3.1.160) plays a central role by finalizing the synthesisof the six-ring ajmalan system. The enzyme converts the sarpagan-type alkaloid epivellosimine, which is the reaction product ofthe preceding enzyme PNAE (see Section 4), to the ajmalan-type in a coenzyme A-dependent reaction. VS has been identifiedfrom R. serpentina cell suspensions and its properties and werepreliminarily described a long time ago.99 Later, the cDNA of theenzyme was isolated and could successfully be expressed in E. coli.Again, the “reverse genetic approach” was the successful strategyto get the full length cDNA clone:100,101 after purification of VSfrom the plant cells, it was identified by SDS gel electrophoresisand partial sequencing. The primary structure of the encodedprotein and sequence alignment studies demonstrated that it was anovel enzyme. The pQE-2 vector and E. coli M-15 cell line servedfor good overexpression with mg amounts of enzyme per litre

Fig. 12 Geometry of the entrance of the reaction channel (A) derived from X-ray analysis and (B) from modelling of PNAE; and zoomed regions (C)and (D), respectively (images provided by Dr M. Hill).

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of bacterial suspension. The soluble enzyme was purified, as forthe previous proteins (Sections 2–4), by Ni-NTA chromatography,always an important step in getting homogenous enzyme prepara-tions for biochemical characterization as well as for crystallization.The functional identity of VS was proven by showing formation ofthe product of the reaction (the methoxylated indolenine alkaloidvinorine) by electron impact mass spectrometry.101

Sequence alignment studies supported classification of VSwithin the so-called BAHD enzyme superfamily.102 BAHD isthe abbreviated name from the first four enzymes of this familyisolated from plant species. Primarily this classification wasbased on two highly conserved motifs in that family, 362Asp-Phe-Gly-Trp-Gly366 and 160His-xxx-Asp164, respectively. As deducedfrom X-ray structures of chloramphenicol acetyltransferase anddihydrolipoamide acetyltransferase, these motifs participate inthe catalytic reaction of acetyl transfer.103,104 A catalytic triad(Lys-His-Asp) was discussed for the activity of arylamine N-acetyltransferase.105,106 The primary structure of VS containsseveral of these residues, suggesting an important function of theseparticular three amino acids.101

5.2 Inhibitors and mutants of VS

After expression of the VS cDNA in E. coli and its purificationto near homogeneity, inhibition studies gave the first clue as tothe catalytically active amino acids.101 Inhibitors acting on Ser,His and Cys such as chloromethyl ketones, the unselective His-directed diethyl pyrocarbonate, Hg2+-ion as a SH-group modifier,the Ser- and Cys-selective benzenesulfonylfluroide (AEBSF) andthe agmatine derivative E-64 were all tested. All these compoundsinhibited the VS activity.101 The results clearly indicated that Ser,His and Cys are essential for the VS-catalyzed reaction. Togetherwith results of sequence alignment studies, a series of mutationexperiments was performed101 and gave some detailed informationon the putative catalytic residues (see Table 3). Five conserved Serresidues were replaced by Ala and all the derived mutant enzymesstill showed catalytic activity. This made involvement of a Ser-His-Asp triad in the binding site unlikely. Also, when Cys89 andCys149 were mutated to Ala, the VS activity was not completely“knocked out”, making a Cys-His-Asp triad also unlikely. Butreplacement of His160 and Asp164 by Ala resulted in inactive

Table 3 Recombinant His6-tagged VS mutants and their relative activity(n.d. = not detectable, data are from ref. 101)

Enzyme Relative activity (%)

Wild-type 100S68A 100C89A 100S413A 100D360A 100S16A 71N293A 68D362A 35S29A 25S243A 17D32A 14C149A 10H160A n.d.D164A n.d.

enzyme mutants, which provided the first clear evidence for thefunctional and maybe exclusive role of the BAHD-typical domainHis-xxx-Asp in this particular enzyme family. The second motif,362Asp-Phe-Gly-Trp-Gly366(DFGWG), is completely conserved inthe BAHD enzymes. Mutation of the first Asp reduced the VSactivity by about one-third, a result matching with mutationof another BAHD enzyme involved in anthocyanin glucosidemalonylation.107 The 3D X-ray structure of VS discussed belowwill explain the significance of this motif. Based on sequencealignments, acyltransferases are divided into four evolutionarysequence clusters.108 Adopting this classification, VS falls intocluster C enzymes catalyzing esterification of hydroxyl groups ofmetabolically unrelated secondary metabolites.

From the above results only two probable catalytically activeamino acids were identified for VS; His160 and Asp164. Theirfunction was proven by the 3D-structure of VS (discussed inSection 5.4), after finding appropriate conditions of crystallizationof the recombinant enzyme in long crystallization trials.

5.3 Unusual crystallization conditions and instability of VScrystals

“Beginner’s luck is definitely a factor in finding conditions forcrystallizing a protein the first time. This is partly becausebeginners are more willing to try new conditions and will oftendo naıve things to the sample, thus finding novel conditions forcrystal growth. This is also because no one can predict the properconditions for crystallizing a new protein”.109

In this light, VS represents a good example of successfully usingrather uncommon crystallization conditions. Similarly to the otherenzymes described in this article, on the one hand the purificationprocedure of VS after over-expression in E. coli was crucial forobtaining the first crystals, but on the other hand, removal of theHis6-tag by DAPase was the second requirement to succeed. Eachexperiment to crystallize the His-tagged enzyme resulted in pre-cipitation or in undetectable nucleation of the enzyme. After Ni-NTA chromatography, two additional chromatographic steps (ionexchange on MonoQ followed by Sephacryl S-100) were essentialand gave highly pure VS as judged by SDS gel chromatography andCoomassie staining. Also in this case the “hanging drop” methodwas used and initially small, clustered crystals were observedon applying “Crystal Screen” and “Crystal Screen2” kits fromHampton Research110,111 at 22 ◦C and using ammonium sulfate asthe precipitant in the presence of polyethylene glycol (PEG) 400.Optimum crystallization conditions were found after systematicchanges of precipitant concentrations, types of PEG, buffers, pH,temperature and enzyme concentrations. VS crystals obtainedonly at the relatively high temperature of 32 ◦C and low proteinconcentration of 2 mg ml−1, proved to be the best ones for X-raymeasurements. These conditions are exceptional, because such acombination of temperature and enzyme concentration was notfound in the Biological Macromolecule Crystallization Database(BMCD),112 and was just based on trying unusual conditions.The well diffracting VS crystals proved to be very sensitive totemperature changes. In contrast to all other crystals mentionedin this article, it was not possible to transport VS crystals forsynchrotron measurements without careful freezing procedures,and such procedures were always essential.

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5.4 Structure elucidation of VS

Up to the time when this review was written, there were nocrystal structures known in the BAHD enzyme family whichcould be helpful for 3D-structure elucidation of VS. By inhibitionof methionine biosynthesis,113 the recombinant E. coli produced(in the presence of selenomethionine) SeMet-VS, which waspurified and crystallized similarly to the wild-type VS. The “multi-wavelength anomalous diffraction” (MAD) approach togetherwith the diffraction data of the wild-type VS (2.6 A resolution)allowed for the first time determination of the VS structure, forwhich the selenium sites were taken as marker residues to placethe amino acid side chains into the appropriate electron densities.After several rounds of refinement, the final three-dimensionalstructure of VS was obtained.114–116 The structure contains 14 b-strands (b1–b14) and 13 a-helices (a1–a13) and consists of twodomains, A and B, of about the same size. The two domains areconnected by a long loop. Both have a similar backbone fold, buttheir topology is different. Between the domains a solvent channelis formed, running through the entire VS molecule (Fig. 13).The His-xxx-Asp motif is located at the interface of the twodomains. His160 is situated directly in the centre of the solventchannel and is accessible from both ends of the channel. Suchan arrangement allows both reaction partners, 16-epi-vellosimineand acetyl-CoA, to approach the active site. Kinetic data obtainedwith a partially purified VS preparation from Rauvolfia cells hadearlier suggested a ternary complex between both ligands and theenzyme with independent binding of the ligands.99 The mechanism

Fig. 13 Image of VS illustrating the two-domain structure (A and B), thereaction channel and the catalytic His in the binding pocket.

of the reaction is illustrated in Scheme 3. The 3D-structure of VSstrongly supports this proposal.

5.5 Important amino acids and mechanistic aspects

The mutagenesis studies indicated the indispensability of His160for VS activity, and the 3D-structure of the enzyme confirmsthe functional importance of this residue. It is this amino acidwhich is located directly in the centre of the solvent channel(Fig. 14). This is in agreement with the above-mentioned kineticdata, that the two substrates may approach the binding pocketfrom different directions (from which direction still needs to beanalyzed). Based on His160, the acetyl-transfer reaction wouldoccur as illustrated in Scheme 3. The amino acid acts as a base,taking off the proton of the hydroxy group of 17-deacetylvinorine.The 17-oxygen will then attack the carbonyl carbon of acetyl-CoA, resulting in its acetylation and release of CoA. The reactionprobably proceeds without formation of an acetylated enzymeintermediate. The second conserved residue (Asp164) is alsolocated directly at the binding site and belongs to the essentialHis-xxx-Asp motif. Asp164 has no catalytic function. Its sidechain points away from His160, not allowing hydrogen-bondformation between the residues. Such an arrangement excludesan amino acid dyad in the catalytic process as found for e.g.human carnitine acetyltransferase.117 Asp164 forms a salt bridgeto Arg279, which is also a conserved residue in the BAHD family.This interaction of the two residues appears to be of structural, notof catalytic, importance. Exchange of Arg279 against Ala resultsin loss of enzyme activity. Most probably it is the geometry ofthe binding pocket which is significantly changed when the saltbridge is interrupted. It must be demonstrated in future whetherthe importance of the His-xxx-Asp motif in other enzymes ofthe BAHD family is based on the same reasons as shown forVS. The DFGWG motif completely conserved in that family isnot localized in the binding pocket of the enzyme. In contrast, itis far away from the region of catalysis, indicating its structuralimportance. It is not involved in substrate binding and mightmaintain the conformation of the enzyme structure. Mutationof the Asp residue to Ala therefore caused decrease of VS activity(Table 4). Such mutation also resulted in complete loss of reactivityof another BAHD enzyme participating in anthocyanin glucosidemalonylation.107 A more detailed insight into the binding pocket ofVS requires more structural information. In particular, the crystalstructure of vinorine synthase with substrate or product boundshould provide much deeper understanding of the catalytic processin this enzyme family.

Scheme 3 Proposed reaction mechanism of VS; scheme from ref. 116.

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Fig. 14 Surface representation, reaction channel and the localization ofthe catalytic residue His160 (in red) in VS (images kindly provided byDr M. Hill).

Very recently, structural and mutational studies on anthocyaninmalonyltransferase were reported.118 This enzyme is also a BAHDmember and now represents the second example where the 3D-structure has been solved. The complex of this malonyltransferasewith its ligand malonyl-CoA together with a series of site-directedmutations allowed the acyl-acceptor binding site to be identified.These results together with those on VS are very helpful for theunderstanding in more detail of the diversity of the acyl-acceptorspecificity within the BAHD family.118

5.6 Significance of the vinorine synthase structure for the BAHDenzyme family

The BAHD enzymes belong to a constantly growing family withan increasing number of functionally expressed members. At theDNA level it is suggested that in two organisms alone ∼180 genesoccur that might code for BAHD acyltransferases; in Oryza sativa(rice) 119 genes of this family have been identified, but none havebeen investigated for their biochemical function. The Arabidopsisthaliana genome delivered 64 genes of the family, but very fewhave been functionally described so far. The (Z)-3-hexen-1-olO-acetyltransferase (producing the acetylated hexenol)119 oranother anthocyanidin 5-O-glucoside-O-malonyltransferase120 are

recent examples. Rosmarinic acid synthase is one of the mostrecent members of the BAHD family detected by “reversegenetics”.121 Prediction of the reactions catalyzed or the substratesaccepted by the enzymes, based on sequence alignments, isdifficult.122 But mutation experiments, such as those describedabove, combined with sequence comparison might help to elu-cidate the functionalities of other members of the BAHD family.

The structure of VS could also be helpful in solving otherstructures by molecular replacement approaches, or at least couldprovide opportunities for homology modelling. This is particularlyimportant for those family members for which the cDNAs havefunctionally been cloned and which take part in the biosynthe-sis of important plant natural products. Prominent examplesare salutaridinol acetyltransferase in morphine biosynthesis,123

deacetylvindoline acetyltransferase124 on the way to vinblastine,and the acyltransferases acting on the skeleton of taxol,125 just tomention BAHD members involved in alkaloid biosynthesis.

6 Raucaffricine glucosidase (RG), a side route of theajmaline pathway

As a part of a metabolic network, the ajmaline biosyntheticpathway has at several intermediate stages side routes, of whichthe formation of raucaffricine is one of the most important.Raucaffricine is the glucoside of the intermediate vomilenine. Thisglucoalkaloid has previously been detected as the main alkaloidof Rauvolfia cell cultures, exceeding significantly the amountsin differentiated Rauvolfia cells.126 The enzyme, raucaffricineglucosidase (RG, EC 3.2.1.125), converts the glucoside back tovomilenine. It has been identified, characterized and its cDNA hasbeen cloned in E. coli.127,128 RG represents the second glucoside-hydrolyzing enzyme in Rauvolfia alkaloid biosynthesis, the firstbeing SG. The primary structure and the substrate specificity ofthe two glucosidases are different; RG accepts strictosidine, thesubstrate of SG, but SG does not hydrolyze raucaffricine.127,128 Tocompare the two glucosidases, it was important to determine the3D-structure of RG in order to evaluate substrate recognition andbetter understand the deglucosylation process. Over-expressionyielded enough pure RG enzyme for crystallization experimentswhen the routine strategy, applied for the other Rauvolfia enzymes,was followed.

6.1 Crystallization attempts and the 3D-structure of RG

His6-tag RG was crystallized by the hanging-drop vapour diffusiontechnique using similar conditions as for strictosidine glucosidase.Crystals reached maximum dimensions of about 0.2 × 0.15 ×0.05 mm. The crystals belong to space group I222 and diffract to2.30 A.129

The structure of RG was determined by the molecular replace-ment method (unpublished data). The structure of Rauvolfia SGserved as a search model, because RG shares a sequence identitywith SG of 56%. RG belongs to the family 1 of the glycosylhydrolases. Crystal packing showed two enzyme molecules foreach crystallographic asymmetric unit, but the enzyme is active asthe monomer.127

The refined model of RG consists of 13 a-helices and 13 b-strands. RG adopts the expected topology of a single (b/a)8 barrelfold (TIM barrel) (Fig. 15). The barrel hosts a binding site for the

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natural substrate raucaffricine. The groove leading to the catalyticcentre is formed mainly by irregular loops between the secondarystructures on top of the enzyme. Similarly to SG, RG contains thecatalytic residues E186 and E420. The most striking difference isat the catalytic centre of the RG and SG. An identical position ofTrp392 but a different orientation of the tryptophan in the twoglucosidases helps the protein to recognize its substrate.

Fig. 15 The (b/a)8 barrel fold of RG, illustrating the binding pocket andthe catalytic acids E186 and E420.

7 Structure-based redesign of enzymes andapplications in chemo-enzymatic approaches

The enzymes described in this review are of relatively highsubstrate specificity. This is a great disadvantage in terms of usingthem in the future as biocatalysts e.g. for the enzymatic synthesisof structurally novel alkaloids. For instance, the best substrate forSTR1 is tryptamine, and benzene-ring-substituted tryptaminesgenerally react at less than 10% of the rate for tryptamine. TheSTR1 mutant Val208Ala, however, converts 5-methyltryptamine

and 5-methoxytryptamine into the corresponding novel substi-tuted strictosidines, indicating the importance of knowing the 3D-structure of STR1 for redesigning its enzyme activity.51,52 Basedon such information, a more systematic mutation approach will inthe future show whether a further expanded substitution patternat the indole moiety can be achieved.

A biomimetic approach which uses strictosidine (or derivatives),strictosidine glucosidase and an excess of primary amines (with ob-viously any residues “R” followed by reduction) (Scheme 4), resultsin formation of novel N-analogous heteroyohimbine alkaloids.47,52

After producing STR1 mutants (allowing synthesis of a range ofnovel strictosidines) and optimizing the second enzyme strictosi-dine glucosidase by mutation, such a chemo-enzymatic approachmay be an excellent tool for the generation of new alkaloid librariescontaining thousands of alkaloids. Knowledge of biosynthesiscombined with structural biology might be an excellent tool in fu-ture to develop such combinatorial enzyme-mediated approaches.

8 Other structural examples from the alkaloid field

Despite the fact that alkaloids have been an excellent sourceof biologically active compounds, e.g. anti-infectious drugs,130

only limited efforts have been focussed on understanding themode of action of alkaloids and their interactions with hostproteins or enzymes from other organisms at the molecular level.These include crystallographic studies of tropinone reductase,phospholipase, transcriptional repressor, acetylcholine esterase,calmodulin and tyrosine kinase and their complexes with variousalkaloids, listed in Table 4.

Below the only other structural examples from alkaloid biosyn-thesis, the tropinone reductases, are discussed in some detail.

Tropinone reductase (TR) comprises a branching point in thebiosynthetic pathway of tropane alkaloids which includes suchmedicinally important compounds as hyoscyamine, scopolamineand cocaine (Scheme 5). All the tropane-alkaloid-producing plantsspecies so far examined have two TR activities and their amino acidsequence is known from Datura stramonium,131,132 Hyoscyamusniger,133 Atropa belladonna134,135 and Solanum tuberosum.136 Inthese species, there are two types of TR, TR-I and TR-II. These

Scheme 4 A biomimetic approach to indole alkaloid diversity based on rational enzyme design (STR1, strictosidine synthase; SG, strictosidineglucosidase; X, various substituents).

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Table 4 3D-structures of enzyme–alkaloid complexes from the PDB data bank

PDBcode Protein Complex with alkaloid

2AE2 Tropinone reductase-II Pseudotropine142

1IPF Tropinone reductase-II Tropinone143

1ZR8 Phospholipase Ajmaline144

1JUM Transcriptional repressor Berberine145

1VOT Acetylcholine-esterase Huperzine A146

1ZGB Acetylcholine-esterase (R)-Tacrine-C10H20-hupyridone147

1DX6 Acetylcholine-esterase Galanthamine148

1H22 Acetylcholine-esterase (S)-Hupyridone-C10H20-(S)-hupyridone149

1XA5 Calmodulin A bis-indole alkaloid150

Scheme 5 Biosynthetic connection of tropane alkaloids through the intermediate tropinone (ODC, ornithine decarboxylase; PMT, putrescinemethyltransferase; TR-I, tropinone reductase I; TR-II, tropinone reductase II).

TRs share 64% sequence identity and belong to the short-chaindehydrogenase/reductase (SDR) family. TRs catalyze NADPH-dependent reductions of the 3-carbonyl group of their commonsubstrate, tropinone, to hydroxy groups with different diastere-omeric configurations: TR-I (EC 1.1.1.206) produces tropine(3a-hydroxytropane), and TR-II (EC 1.1.1.236) produces pseu-dotropine (W-tropine, 3b-hydroxytropane). These enzymes havedifferent Km values for tropinone and its analogues but have similarKm values for NADPH, and both catalyze transfer of the pro-Shydrogen atom of NADPH to tropinone.137

This indicates that the two TR enzymes have different bindingsites for tropinone but have similar ones for NADPH, and thattheir different stereospecificities result from the different bindingmodes of tropinone. Both TRs from Datura stramonium have beencloned and expressed in E. coli, and purified and crystallizedusing hanging drop vapour diffusion techniques with sodium

citrate and 2-methyl-2,4-pentenediol as buffer and precipitantrespectively.138,139 The structure of each TR was solved using theisomorphous replacement method.140 The two structures are al-most indistinguishable from each other in both subunit folding andtheir association into dimers. Both TR subunits consist of a coredomain that includes most of the polypeptide and a small lobe thatprotrudes from the core. In the centre of the core domain is a seven-stranded parallel b-sheet, flanked on each side by three a-helices,which constitutes the Rossmann-fold topology. This core structureis highly conserved among the SDR family members, despiterelatively low residue identity between these enzymes (∼30%).141

The structure of TR-I was also determined in the presence ofNADP+. The cofactor was found to be located at the bottom ofthe cleft between the core domain and the small lobe. The carbox-amide group of the nicotinamide ring is anchored by the main-chain nitrogen and oxygen atoms of Ile204 and the side-chain

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oxygen of Thr206. This tight binding of the carboxamide groupto the protein directs the B-face of the nicotinamide ring towardthe void of the cleft, consistent with the observed specificity forthe pro-S hydride transfer of both TRs.137

TR-II catalyzes the one step chemical reaction via the followingstates:

(1) TR-II (E) + NADPH (S1) + tropinone (S2) ↔(2) ES1S2 ↔(3) EP1P2 ↔(4) TR-II + NADP+(P1)+ W-tropinone (P2),where the states are: (1) E, the free enzyme; (2) ES1S2, the enzyme

with the substrates [NADPH (S1) and tropinone (S2)] bound priorto reaction initiation; (3) EP1P2, the enzyme with the products[NADP+ (P1) and W-tropine (P2)] bound just after the reactionis completed; and (4) the free enzyme again after releasing theproducts.

The crystallographic structures of TR-II have been determinedin an unliganded form,140 as the complex with NADPH,141 as thecomplex with NADPH and tropine,141 and as the complex withNADP+ and W-tropine.142 These structures provide a great dealof insight into each state of the reaction in the biosynthesis oftropane alkaloids (Fig. 16).

Fig. 16 (a) Complex structure of tropinone reductase II (TR-II) withtropinone in black and NADPH in blue+grey. (b) Complex structure oftropinone reductase I (TR-I) with NADPH illustrated in blue+grey.

9 Conclusions and future aspects of structuralbiology in the alkaloid field

Elucidation of the three-dimensional structures of several of theRauvolfia enzymes has extensively broadened our knowledge on

details of the enzymatic biosynthesis of monoterpenoid indolealkaloids, particularly of the pathway to the antiarrhythmicajmaline. For five major enzymes, the fold-families, their overallstructures and substrate binding sites are now known, includingidentification of amino acids of catalytic and structural impor-tance. This will facilitate elucidation of the corresponding enzymemechanisms. For enzymes of more general significance, such asstrictosidine synthase, crystal structures of substrates and productcomplexes have been analyzed; this allows understanding of ligandrecognition and permitted the first successful rational enzymeengineering for the synthesis of novel alkaloids. Structure-guidedcombinatorial approaches with redesigned enzyme mutants to-gether with biomimetic strategies will help to generate largealkaloid libraries with hopefully important and novel biologicalactivities.52

10 Acknowledgements

We thank Mrs Yang Liuqing for kind help in preparing thediagrams for this article. The described research was continuouslysupported by the Deutsche Forschungsgemeinschaft (Bonn, Bad-Godesberg, Germany), the Fonds der Chemischen Industrie(Frankfurt/Main, Germany) in cooperation with the Bundesmin-isterium fur Bildung und Forschung (BMBF, Bonn, Germany), theDLR Office Bonn of BMBF, and the Deutscher AkademischerAustauschdienst (DAAD), Bonn, Germany. The work was alsosupported by European Community Access to Research Infras-tructure Action of the Improving Human Potential Programmeto the European Molecular Biology Laboratory (EMBL) at Ham-burg Outstation (contract No. HPRI-CT-1999-100017) and theFP6 Programme (No. RII3/CT/2004/5060008), and the BerlinerElektronenspeicherring-Gesellschaft fur Synchrotronstrahlung(Berlin, Germany). We also acknowledge staff members at theDORIS storage ring (DESY, Hamburg, Germany), Prof. Lottspe-ich’s team at the Max-Planck-Institute of Biochemistry (MPI,Martinsried, Germany) for protein sequencing, Prof. H. Michel’sgroup at MPI (Frankfurt/Main, Germany) and Prof. T. Kutchan(St Louis, USA) for introduction into structural and molecularbiology techniques, respectively, and the ESRF beamline (Greno-ble, France) for technical help. Present co-workers (in alphabeticalorder: Dr Leif Barleben, Dr Marco Hill, Petra Kercmar, ElkeLoris, Kerstin Oelrich, and Dr Martin Ruppert) and former co-workers of various nationalities are very much appreciated for theirenthusiastic performing of chemical, enzymatic and crystallizationexperiments. Both Dr E. Mattern-Dogru and Dr Xueyan Ma orig-inally initiated structural biology research in our group. We thankDr F. Leeper (Cambridge, UK) and Dr P. Tucker (EMBL Ham-burg, Germany) for advice and help in correcting the manuscript.

11 References

1 E. J. Saxton, Nat. Prod. Rep., 1997, 14, 559–590.2 J. Leonhard, Nat. Prod. Rep., 1999, 16, 319–338.3 J. Stoeckigt, ‘Biosynthesis in Rauvolfia serpentina—Modern Aspects

of an Old Medicinal Plant’, in: The Alkaloids, ed. G. A. Cordell,Academic Press, San Diego, California, vol. 47, pp. 115–172.

4 S. O’Connor and J. J. Maresh, Nat. Prod. Rep., 2006, 23, 532–547.5 T. M. Kutchan, Recent Adv. Phytochem., 2002, 36, 163–178.6 V. De Luca, Recent Adv. Phytochem., 2003, 37, 181–202.7 T. Hashimoto and Y. Yamada, Curr. Opin. Biotechnol., 2003, 14, 163–

168.8 J. Murata and V. De Luca, Plant J., 2005, 44, 581–594.

1398 | Nat. Prod. Rep., 2007, 24, 1382–1400 This journal is © The Royal Society of Chemistry 2007

Dow

nloa

ded

on 0

1 N

ovem

ber

2011

Publ

ishe

d on

24

Oct

ober

200

7 on

http

://pu

bs.r

sc.o

rg |

doi:1

0.10

39/B

7119

35F

View Online

9 F. Vazquez-Flota, E. De Carolis, A. Alarco and V. De Luca, PlantMol. Biol., 1997, 34, 935–948.

10 B. St. Pierre, P. Laflamme, A. Alarco and V. De Luca, Plant J., 1998,14, 703–713.

11 Y. Yamazaki, A. Urano, H. Sudo, M. Kitajima, H. Takayama, M.Yamazaki, N. Aimi and K. Saito, Phytochemistry, 2003, 62, 461–470.

12 A. H. Heckendorf and C. R. Hutchinson, Tetrahedron Lett., 1977, 48,4153–4154.

13 C. R. Hutchinson, A. H. Heckendorf, J. L. Straughn, P. E. Daddonaand D. E. Cane, J. Am. Chem. Soc., 1979, 101, 3358–3369.

14 A. I. Scott and S.-L. Lee, J. Am. Chem. Soc., 1975, 97, 6906–6908.15 J. Stoeckigt, J. Treimer and M. H. Zenk, FEBS Lett., 1976, 70, 267–

270.16 J. Stoeckigt, T. Hemscheidt, G. Hoefle, P. Heinstein and V. Formacek,

Biochemistry, 1983, 22, 3448–3452.17 J. Stoeckigt, ‘The Biosynthesis of Heteroyohimbine-Type Alkaloids’,

in: Indole and biogenetically related alkaloids, ed. J. D. Philipson andM. H. Zenk, Academic Press, London/New York, 1980, pp. 113–141.

18 M. H. Zenk, H. El-Shagi, H. Arens, J. Stoeckigt, E. W. Weiler and B.Deus, ‘Formation of the Indole Alkaloids Serpentine and Ajmalicinein Cell Suspension Cultures of Catharautus roseus’, in: Plant TissueCulture and its Bio-technological Application, ed. W. Barz, E. Reinhardand M. H. Zenk, Springer Verlag, Berlin/Heidelberg/New York,1977, pp. 27–43.

19 J. Stoeckigt and H. J. Soll, Planta Med., 1980, 40, 22–30.20 J. Stoeckigt, P. Obitz, H. Falkenhagen, R. Lutterbach and S. Endress,

Plant Cell, Tissue Organ Culture, 1995, 43, 97–109.21 R. B. Herbert, ‘The Biosynthesis of Terpenoid Indole Alkaloids’,

in: Monoterpenoid Indole Alkaloids, The Chemistry of HeterocyclicCompounds, Vol. 25 (supplement), ed. J. E. Saxton, John Wiley andSons, Chichester/New York/Brisbane/Toronto/Singapore, 1994, pp.1–13.

22 J. Stoeckigt, ‘Alkaloid Metabolism in Plant Cell Culture’, in NaturalProduct Analysis, ed. P. Schreier, M. Herderich, H. U. Humpf andW. Schwab, Vieweg, Braunschweig/Wiesbaden, Germany, 1998, pp.313–325.

23 M. Ruppert, X. Y. Ma and J. Stoeckigt, Curr. Org. Chem., 2005, 9,1431–1444.

24 C. Hinse, Y. Sheludko, A. Provenzani and J. Stoeckigt, J. Am. Chem.Soc., 2001, 123, 5118–5119.

25 C. Hinse, C. Richter, D. Moskau, A. Provenzani and J. Stoeckigt,Bioorg. Med. Chem., 2003, 11, 3913–3919.

26 J. Stoeckigt, S. Panjikar, M. Ruppert, C. Barleben, X. Y. Ma, E. Lorisand M. Hill, Phytochem. Rev., 2007, 6, 15–34.

27 E. Leete, Planta Med., 1990, 56, 339–352.28 T. Hashimoto and Y. Yamada, Annu. Rev. Plant Physiol. Plant Mol.

Biol., 1994, 45, 257–285.29 J. Stoeckigt and M. H. Zenk, J. Chem. Soc., Chem. Commun., 1977,

646–648.30 J. Stoeckigt and M. H. Zenk, FEBS Lett., 1977, 79, 233–237.31 J. F. Treimer and M. H. Zenk, Eur. J. Biochem., 1979, 10, 225–233.32 A. de Waal, A. H. Meijer and R. Verpoorte, Biochem. J., 1995, 306,

571–580.33 L. H. Stevens, C. Giroud, J. M. E. Pennings and R. Verpoorte,

Phytochemistry, 1993, 33, 99–106.34 H. Wang, R. Chen, M. Chen, M. Sun and Z. H. Liao, Acta Bot.

Boreali Occident. Sin., 2006, 26, 900–905.35 T. M. Kutchan, N. Hampp, F. Lottspeich, K. Beyreuther and M. H.

Zenk, FEBS Lett., 1988, 237, 40–44.36 T. M. Kutchan, FEBS Lett., 1989, 257, 127–130.37 T. M. Kutchan, H. Dittrich, D. Bracher and M. H. Zenk, Tetrahedron,

1991, 47, 5945–5954.38 T. D. McKnight, C. A. Roessner, R. Devagupta, A. I. Scott and C. L.

Nessler, Nucleic Acids Res., 1990, 18, 4939.39 C. Canel, M. I. Lopes-Cardoso, S. Whitmer, L. von der Fits, G.

Pasquali, R. von der Heijden, J. H. Hoge and R. Verpoorte, PlantMol. Biol., 1999, 39, 1299–1310.

40 Y. Yamazaki, H. Sudo, M. Yamazaki, N. Aimi and K. Saito, PlantCell Physiol., 2003, 44, 395–403.

41 X. Y. Ma, J. Koepke, G. Fritzsch, R. Diem, T. M. Kutchan, H. Micheland J. Stoeckigt, Biochim. Biophys. Acta, 2004, 1702, 121–124.

42 J. Koepke, X. Y. Ma, G. Fritzsch, H. Michel and J. Stoeckigt, ActaCrystallogr., Sect. D, 2005, 61, 690–693.

43 S. Panjikar, V. Parthasarathy, V. S. Lamzin, M. S. Weiss and P. A.Tucker, Acta Crystallogr., Sect. D, 2005, 61, 449–457.

44 M. Harel, A. Ahoroni, L. Gaidukov, B. Brumshtein, O. Khersonsky,R. Meged, H. Dvir, R. B. G. Ravelli, A. McCarthy, L. Toker, L.Silman, J. L. Sussman and D. S. Tawfik, Nat. Struct. Mol. Biol., 2004,11, 412–419.

45 E. I. Scharff, J. Koepke, G. Fritzsch, C. Luecke and H. Rueterjans,Structure, 2001, 9, 493–502.

46 X. Y. Ma, S. Panjikar, J. Koepke, E. Loris and J. Stoeckigt, Plant Cell,2006, 18, 907–920.

47 J. Stoeckigt, M. Ruppert, ‘Strictosidine: The biosynthetic key tomonoterpenoid indole alkaloids’, in: Comprehensive Natural ProductsChemistry: Amino Acids, Peptides, Porphyrins and Alkaloids, ed.D. H. R. Barton, K. Nakanishi, O. Meth-Cohn and J. W. Kelly,Elsevier, Amsterdam, 1999, vol. 4, pp. 109–138.

48 N. Hampp and M. H. Zenk, Phytochemistry, 1988, 27, 3811–3815.49 L. H. Stevens, G. Giroud, E. J. M. Pennings and R. Verpoorte,

Phytochemistry, 1993, 35, 353–360.50 E. McCoy, M. C. Galan and S. E. O’Connor, Bioorg. Med. Chem.

Lett., 2006, 16, 2475–2478.51 S. Chen, M. C. Galan, C. Coltharp and S. E. O’Connor, Chem. Biol.,

2006, 13, 1137–1141.52 E. Loris, S. Panjikar, L. Barleben, M. Unger, H. Schubel and J.

Stoeckigt, Chem. Biol., 2007, 14, 979–985.53 T. A. Edwards, B. D. Wilkinson, R. P. Wharton and A. K. Aggarwal,

Genes Dev., 2003, 17, 2508–2513.54 W. De-Eknamkul, N. Suttipanta and T. M. Kutchan, Phytochemistry,

2000, 55, 177–181.55 T. Hemscheidt and M. H. Zenk, FEBS Lett., 1980, 110, 187–

191.56 A. Geerlings, J. I. Memelink, R. von der Heiden and R. Verpoorte,

J. Biol. Chem., 2000, 275, 3051–3056.57 I. Gerasimenko, Y. Sheludkov, X. Y. Ma and J. Stoeckigt,

Eur. J. Biochem., 2002, 269(2), 204–2213.58 G. Haberer and J. J. Kieber, Plant Physiol., 2002, 128, 354–362.59 K. Ljung, A. Oestin, C. Lioussanne and G. Sandberg, Plant Physiol.,

2001, 125, 464–475.60 D. P. Dharmawardhana, B. E. Ellis and J. E. Carlson, Plant Physiol.,

1995, 107, 331–339.61 M. Zagrobelny, S. Bak, A. V. Rasmussen, B. Jorgensen, C. M.

Naumann and B. L. Moller, Phytochemistry, 2004, 65, 293–306.62 J. E. Poulton, Plant Physiol., 1990, 94, 401–405.63 L. Barleben, X. Y. Ma, J. Koepke, G. Peng, H. Michel and J. Stoeckigt,

Biochim. Biophys. Acta, 2005, 1747, 89–92.64 J. D. McCarter and S. G. Withers, Curr. Opin. Struct. Biol., 1994, 4,

885–892.65 J. Sanz-Aparicio, J. A. Martinez-Ripoll, M. Lequerica and J. L.

Polaina, J. Mol. Biol., 1998, 275, 491–502.66 L. Barleben, S. Panjikar, M. Ruppert, J. Koepke and J. Stoeckigt, Plant

Cell, 2007, DOI: 10.1105/tpc.106.045682.67 L. Verdoucq, M. Czjzek, J. Moriniere, D. R. Bevan and A. Esen,

J. Biol. Chem., 2003, 278, 25055–25062.68 L. Verdoucq, J. Moriniere, D. R. Bevan, A. Esen, A. Vasella, B.

Henrissat and M. Czjzek, J. Biol. Chem., 2004, 279, 31796–31803.69 M. Czjzek, M. Cicek, V. Zamboni, D. R. Bevan and A. Esen, Proc.

Natl. Acad. Sci. U. S. A., 2000, 97, 13555–13560.70 M. H. Zenk, ‘Chasing the Enzymes of Alkaloid Biosynthesis’, in

Organic Reactivity: Physical and Biological Aspects, ed. B. T. Golding,R. J. Griffin and H. Maskill, The Royal Society of Chemistry,Cambridge, UK, 1995, pp. 89–109.

71 T. M. Kutchan, Plant Cell, 1995, 7, 1059–1070.72 A. Pfitzner and J. Stoeckigt, Planta Med., 1983a, 48, 221–227.73 D. Schmidt and J. Stoeckigt, Planta Med., 1995, 61, 254–258.74 A. Pfitzner and J. Stoeckigt, J. Chem. Soc., Chem. Commun., 1983,

459–460.75 H. Warzecha and J. Stoeckigt, ‘New Enzymes of Indole Alkaloid

Biosynthesis’, in: Proceedings of the 3rd International Symposium onFrontiers in Protein Chemistry and Biotechnology, ed. Z. Wang andW. Li, Changchun Publishing House, China, 1999, pp. 38–43.

76 E. Dogru, H. Warzecha, F. Seibel, S. Haebel, F. Lottspeich and J.Stoeckigt, Eur. J. Biochem., 2000, 267, 1397–1406.

77 E. Mattern-Dogru, X. Y. Ma, J. Hartmann, H. Decker and J.Stoeckigt, Eur. J. Biochem., 2002, 269, 2889–2896.

78 E. Mattern-Dogru, Molecular Analysis of Polyneuridine AldehydeEsterase by Functional Expression: Site-directed Mutagenesis andHomology-Modelling, Der Andere Verlag, Osnabrueck, 2002, pp. 1–116.

This journal is © The Royal Society of Chemistry 2007 Nat. Prod. Rep., 2007, 24, 1382–1400 | 1399

Dow

nloa

ded

on 0

1 N

ovem

ber

2011

Publ

ishe

d on

24

Oct

ober

200

7 on

http

://pu

bs.r

sc.o

rg |

doi:1

0.10

39/B

7119

35F

View Online

79 The Arabidopsis Initiative, Nature, 2000, 408, 796–815.80 M. Hasslacher, M. Schall, M. Hayn, H. Griengl, S. D. Kohlwein and

H. Schwab, J. Biol. Chem., 1996, 271, 5884–5891.81 M. Hasslacher, C. Kratky, Z. H. Griengl, H. Schwab and S. D.

Kohlwein, Proteins, 1997, 27, 438–449.82 U. Hanefeld, A. J. J. Straathof and J. J. Heijnen, Biochim. Biophys.

Acta, 1999, 1432, 185–193.83 K. Gruber, M. Gugganig, U. G. Wagner and C. Kratky, Biol. Chem.,

1999, 380, 993–1000.84 J. Hughes, F. J. P. Carvalho and M. A. Hughes, Arch. Biochem.

Biophys., 1994, 311, 496–502.85 H. Wajant and K. Pfizenmaier, J. Biol. Chem., 1996, 271, 25830–

25834.86 H. Lauble, S. Foerster, B. Miehlich, H. Wajant and F. Effenberger,

Acta Crystallogr., Sect. D, 2001, 57, 194–200.87 H. Lauble, B. Miehlich, S. Foerster, C. Koblar, H. Wajant and F.

Effenberger, Protein Sci., 2002, 11, 65–71.88 N. E. Chayen, Methods Mol. Biol., 2007, 363, 175–190.89 N. E. Chayen, Structure, 1997, 5, 1269–1274.90 I. M. Berry, O. Dym, R. M. Esnouf, K. Harlos, R. Meged, A. Perrakis,

J. L. Sussman, T. S. Walter, J. Wilson and A. Messerschmidt, ActaCrystallogr., Sect. D, 2006, 62, 1137–1149.

91 B. Hazes and L. Price, Acta Crystallogr., Sect. D, 2005, 61, 1165–1171.

92 J. Mueller-Dieckmann, Acta Crystallogr., Sect. D, 2006, 62, 1446–1452.

93 L. J. De Lucas, D. Hamrick, L. Cosenza, L. Nagy, D. McCombs, T.Bray, A. Chait, B. Stoops, A. Belgovskiy, W. W. Wilson, M. Parhamand N. Chernov, Prog. Biophys. Mol. Biol., 2005, 88, 285–309.

94 A. Sali and T. Blundell, J. Mol. Biol., 1993, 234, 779–815.95 F. Forouhar, Y. Yang, D. Kumar, Y. Chen and E. Fridman, Proc. Natl.

Acad. Sci. U. S. A., 2005, 102, 1773–1778.96 Esther database, 02.2007: T. Hotelier, L. Renault, X. Cousin, V. Negre,

P. Marchot and A. Chatonnet, Nucleic Acids Res., 2004, 32, D145–D147.

97 N. Siew, H. K. Saini and D. Fischer, FEBS Lett., 2005, 579, 3175–3182.

98 Y. Cajal, A. Svendsen, J. De Bolos, S. A. Patkar and M. A. Alsina,Biochimie, 2000, 82, 1053–1061.

99 A. Pfitzner, L. Polz and J. Stoeckigt, Z. Naturforsch., C: Biosci., 1986,41, 103–114.

100 I. Gerasimenko, X. Y. Ma, Y. Sheludko, R. Mentele, F. Lottspeich andJ. Stoeckigt, Bioorg. Med. Chem., 2004, 12, 2781–2786.

101 A. Bayer, X. Y. Ma and J. Stoeckigt, Bioorg. Med. Chem., 2004, 12,2787–2795.

102 B. St. Pierre and V. De Luca, ‘Evolution of actyltransferase genes:origin and diversification of the BAHD superfamily of acyltrans-ferases involved in secondary metabolism’, in: Recent Advances inPhytochemistry: Evolution of Metabolic Pathways, ed. J. T. Romeo,R. Ibrahim, L. Varin and V. De Luca, Elsevier Science, 2000, vol. 34,pp. 285–315.

103 A. G. Leslie, Mol. Biol., 1990, 213, 167–186.104 T. A. Keating, C. G. Marshall, C. T. Walsh and A. E. Keating, Nat.

Struct. Biol., 2002, 9, 522–526.105 J. C. Sinclair, J. Sandy, R. Delgoda, E. Sim and M. E. M. Noble, Nat.

Struct. Biol., 2000, 7, 560–564.106 A. Upton, N. Johnson, J. Sandy and E. Sim, Trends Pharmacol. Sci.,

2001, 22, 140–146.107 H. Suzuki, T. Nakayama and T. Nishino, Biochemistry, 2003, 42,

1764–1771.108 L. Hoffmann, S. Maury, F. Martz, P. Geoffroy and M. Legrand, J. Biol.

Chem., 2003, 278, 95–103.109 D. E. McRee, Practical Protein Crystallography (2nd edn), Academic

Press, San Diego, California, 1999, p. 9.110 J. Jansarik and S.-H. Kim, J. Appl. Crystallogr., 1991, 24, 409–

411.111 R. Cudney, S. Patel, K. Weisgraber, Y. Newhouse and A. McPherson,

Acta Crystallogr., Sect. D, 1994, 50, 414–423.112 G. L. Gilliland, M. Tung, D. M. Blakeslee and J. Ladner, Acta

Crystallogr., Sect. D, 1994, 50, 408–413.113 G. D. Van Duyne, R. F. Standaert, P. A. Karplus, S. L. Schreiber and

J. Clardy, J. Mol. Biol., 1993, 229, 105–124.114 X. Y. Ma, J. Koepke, A. Bayer, V. Linhard, G. Fritzsch, B. Zhang,

H. Michel and J. Stoeckigt, Biochim. Biophys. Acta, 2004, 1701, 129–132.

115 X. Y. Ma, J. Koepke, A. Bayer, G. Fritzsch, H. Michel and J. Stoeckigt,Acta Crystallogr., Sect. D, 2005, 61, 694–696.

116 X. Y. Ma, J. Koepke, S. Panjikar, G. Fritzsch and J. Stoeckigt, J. Biol.Chem., 2005, 280, 13576–13583.

117 D. Wu, L. Govindasamy, W. Lian, Y. Gu, T. Kukar, M. Agbandje-McKenna and R. McKenna, J. Biol. Chem., 2003, 278, 13159–13165.

118 H. Unno, F. Ichimaida, H. Suzuki, S. Takahashi, Y. Tanaka, A. Saito,T. Nishino, M. Kusunoki and T. Nakayama, J. Biol. Chem., 2007, 282,15812–15822.

119 J. C. D’Auria, E. Pichersky, A. Schaub, A. Hansel and J. Gershenzon,Plant J., 2007, 49, 194–207.

120 J. C. D’Auria, M. Reichelt, K. Luck, A. Svatos and J. Gershenzon,FEBS Lett., 2007, 581, 872–878.

121 A. Berger, J. Meinhard and M. Petersen, Planta, 2006, 224, 1503–1510.122 J. Beekwilder, M. Alvarez-Huerta, E. Neef, F. Verstappen, H.

Bouwmeester and A. Aharoni, Plant Physiol., 2004, 135, 1865–1878.123 T. Grothe, R. Lenz and T. M. Kutchan, J. Biol. Chem., 2001, 276,

30717–30723.124 B. St-Pierre, P. Laflamme, A. M. Alarco and V. De Luca, Plant J.,

1998, 14, 703–713.125 K. Walker, R. Long and R. Croteau, Proc. Natl. Acad. Sci. U. S. A.,

2002, 99, 9166–9171.126 H. Schuebel and J. Stoeckigt, Plant Cell Rep., 1984, 3, 72–74.127 H. Warzecha, P. Obitz and J. Stoeckigt, Phytochemistry, 1999, 50,

1099–1109.128 H. Warzecha, I. Gerasimenko, T. M. Kutchan and J. Stoeckigt,

Phytochemistry, 2000, 54, 657–666.129 M. Ruppert, S. Panjikar, L. Barleben and J. Stoeckigt, Acta Crystal-

logr., Sect. F, 2006, 62, 257–260.130 C. Mahidol, S. Ruchirawat, H. Prawat, S. Pisutjaroenpong, S. Eng-

prasert, P. Chumsri, T. Tengchaisri, S. Sirisinha and P. Picha, PureAppl. Chem., 1998, 70, 2065–2072.

131 K. Nakajima, T. Hashimoto and Y. Yamada, Proc. Natl. Acad. Sci.U. S. A., 1993, 90, 9591–9595.

132 K. Nakajima and T. Hashimoto, Plant Cell Physiol., 1999, 40, 1099–1107.

133 K. Nakajima, T. Hashimoto and Y. Yamada, Plant Physiol., 1993,103, 1465–1466.

134 U. Richter, G. Rothe, A. K. Fabian, B. Rahfeld and B. Draeger, J. Exp.Bot., 2005, 56, 645–652.

135 B. Draeger, Phytochemistry, 2006, 67, 327–337.136 R. Keiner, H. Kaiser, K. Nakajima, T. Hashimoto and B. Draeger,

Plant Mol. Biol., 2002, 48, 299–308.137 T. Hashimoto, K. Nakajima, G. Ongena and Y. Yamada, Plant

Physiol., 1992, 100, 836–845.138 K. Nakajima, T. Hashimoto and Y. Yamada, J. Biol. Chem., 1994,

269, 11695–11698.139 A. Yamashita, K. Nakajima, H. Kato, T. Hashimoto, Y. Yamada and

J. Oda, Acta Crystallogr., Sect. D, 1998, 54, 1405–1407.140 K. Nakajima, A. Yamashita, H. Akama, T. Nakatsu, H. Kato, T.

Hashimoto, J. Oda and Y. Yamada, Proc. Natl. Acad. Sci. U. S. A.,1998, 95, 4876–4881.

141 H. Joernvall, B. Persson, M. Krook, S. Atrian, R. Gonzalez-Duarte,J. Jeffey and D. Ghosh, Biochemistry, 1995, 34, 6003–6013.

142 A. Yamashita, H. Kato, S. Wakatsuki, T. Tomizaki, T. Nakatsu, K.Nakajima, T. Hashimoto, Y. Yamada and J. Oda, Biochemistry, 1999,38, 7630–7637.

143 A. Yamashita, M. Endo, T. Higashi, T. Nakatsu, Y. Yamada, J. Odaand H. Kato, Biochemistry, 2003, 42, 5566–5573.

144 M. Mahendra, N. Singh, P. Kaur, S. Sharma and T. P. Singh, to bepublished.

145 M. A. Schumacher, M. C. Miller, S. Grkovic, M. H. Brown, R. A.Skurray and R. G. Brennan, Science, 2001, 294, 2158–2163.

146 M. L. Raves, M. Harel, Y. P. Pang, I. Silman, A. P. Kozikowski andJ. L. Sussman, Nat. Struct. Biol., 1997, 4, 57–63.

147 H. Haviv, D. M. Wong, H. M. Greenblatt, P. R. Carlier, Y. P. Pang, I.Silman and J. L. Sussman, J. Am. Chem. Soc., 2005, 127, 11029–11036.

148 H. M. Greenblatt, G. Kryger, T. Lewis, I. Silman and J. L. Sussman,FEBS Lett., 1999, 463, 321–326.

149 D. M. Wong, H. M. Greenblatt, H. Dvir, P. R. Carlier, Y. F. Han,Y. P. Pang, I. Silman and J. L. Sussman, J. Am. Chem. Soc., 2003, 125,363–373.

150 I. Horvath, V. Harmat, A. Perczel, V. Palfi, L. Nyitray, A. Nagy, E.Hlavanda, G. Naray-Szabo and J. Ovadi, J. Biol. Chem., 2005, 280,8266–8274.

1400 | Nat. Prod. Rep., 2007, 24, 1382–1400 This journal is © The Royal Society of Chemistry 2007

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