sesquiterpene synthases: passive catalysts or active players?€¦ · dedicated to these enzymes...

Dynamic Article LinksC<NPR

Cite this: Nat. Prod. Rep., 2012, 29, 60

www.rsc.org/npr REVIEW

Publ

ishe

d on

08

Nov

embe

r 20

11. D

ownl

oade

d on

28/

11/2

014

01:2

7:21

. View Article Online / Journal Homepage / Table of Contents for this issue

Sesquiterpene synthases: Passive catalysts or active players?

David J. Miller and Rudolf K. Allemann*

Received 29th July 2011

DOI: 10.1039/c1np00060h

Covering: up to July 2011

Sesquiterpene synthases catalyse the metal dependent turnover of farnesyl diphosphate to generate

a class of natural products characterised by an enormous diversity in structure, stereochemistry,

biological function and application. It has been proposed that these enzymes take a passive role in the

reactions they catalyse and that they serve mostly as stereochemical templates, within which the

reactions take place. Here, recent research into the structure and function of sesquiterpene synthases

and the mechanisms of the reactions that they catalyse will be reviewed to suggest that these fascinating

enzymes play multifaceted active roles in what are arguably the most complex biosynthetic reactions.

1 Introduction

2 Metal ion binding and diphosphate cleavage

3 Carbocationic reaction cascades

3.1 Site directed mutagenesis studies

3.2 Aza-analogues

3.3 Substrate analogues

4 Acid/base catalysis

5 Conclusions

6 Acknowledgments

7 References

1 Introduction

With tens of thousands of characterised members throughout all

forms of life, terpenoids are the largest and most structurally

diverse class of natural products.1,2 The chemically most

intriguing contribution to the generation of this diversity is made

by terpene synthases, enzymes that convert their linear isoprenyl

diphosphate substrates into terpenoid hydrocarbons in what are

perhaps the most complex biosynthetic reactions. They catalyse

the production of a multitude of compounds often containing

several stereocentres via carbocationic reaction cascades that

involve changes to the connectivity and hybridisation of up to

half the carbon atoms of the substrate.

Excellent reviews have been published on the biosynthesis of

monoterpenes,3–6 diterpenes5,7 and triterpenes;8–10 the focus here

will be on sesquiterpenes and particularly on sesquiterpene syn-

thases. Sesquiterpene synthases catalyse, within a shared three-

dimensional fold, the conversion of farnesyl diphosphate (FDP,

1a) into more than 300 different sesquiterpenes, all with the

School of Chemistry, Cardiff University, Main Building, Park Place,Cardiff, CF10 3AT, United Kingdom. E-mail: [email protected];Fax: (+41) 29-2087-4030

60 | Nat. Prod. Rep., 2012, 29, 60–71

shared formula C15H24. In the past, monographs have been

dedicated to these enzymes from experimental and theoretical

perspectives.3,11–13 We will be examining the role played by

sesquiterpene synthases in this masterpiece of combinatorial

chemistry from a mechanistic physical organic chemistry view-

point and will aim to describe the general principles shared by all

sesquiterpene synthases for catalysis.

The catalytic mechanisms of sesquiterpene synthases can be

divided into two phases; the metal ion dependent binding of the

substrate and cleavage of the C–O bond to generate an ion pair

between diphosphate and a farnesyl cation is followed by

a precise carbocationic reaction cascade to the final reaction

products. It has previously been suggested that after the initial

ionisation, the enzymes act as passive catalysts that simply

provide templates to chaperone the intermediates along the

reaction pathway. We will suggest a more active role for these

enzymes in catalysis, showing that they are intimately involved in

the chemistry at all stages of the reaction pathway.

The review will be divided into three sections: (i) investigation

of metal ion, substrate binding and generation of the initial

carbocation, (ii) analysis of the role of the enzyme in FDP

folding, cation stabilisation and coordination of the intermedi-

ates and transition states of the cyclisation cascade, and (iii)

study of the nature of putative acids and bases in the enzymes’

active sites.

2 Metal ion binding and diphosphate cleavage

The formation of sesquiterpenes is initiated by the divalent metal

(usually Mg2+) dependent cleavage of diphosphate (PPi) from

FDP; three metal ions are bound at the top of the active site and

coordinate diphosphate prior to cleavage. Metal coordination is

most easily visualised through structural analysis. At the time of

writing the protein databank (http://www.pdb.org) contains the

This journal is ª The Royal Society of Chemistry 2012

http://dx.doi.org/10.1039/c1np00060h






http://pubs.rsc.org/en/journals/journal/NP

http://pubs.rsc.org/en/journals/journal/NP?issueid=NP029001

Publ

ishe

d on

08

Nov

embe

r 20

11. D

ownl

oade

d on

28/

11/2

014

01:2

7:21

. View Article Online

crystal structures of seven distinct sesquiterpene synthases that

have been solved in forty two forms; both wild type and mutant

enzyme structures have been reported in unliganded forms and in

complex with various combinations of metal ions, diphosphate,

FDP, substrate analogues and/or organic cation mimics. All

sesquiterpene synthases contain the class I terpene synthase

a-helical fold, within which the reactions takes place; plant

sesquiterpene synthases also contain a catalytically silent class II

terpene synthase fold (Fig. 1).4 The most conserved motifs in the

class I fold are the ‘DDXXD/E’-motif, which is located on helix

D and binds Mg2+A and Mg2+C , and the ‘DTE/NSE’ motif on helix

H, which binds Mg2+B .4 The recently solved crystal structure of

(+)-d-cadinene (22) synthase from Gossypium arboreum (GA-

DCS) (Fig. 1) revealed that the DTE motif in this enzyme is

absent and instead takes the form of a second DDXXD motif

more reminiscent of the presumed ancestral isoprenyl

Fig. 1 Sketches of the three most recently solved X-ray single crystal stru

Streptomyces coelicolor A3 (EIZS) in complex with Mg2+3 , PPi and benzyltrim

synthase from Gossypium arboreum (GA-DCS) in complex with 3 Mg2+ and 2F

in yellow); C: aristolochene (11) synthase from Aspergillus terreus (AT-AS) in

David J: Miller

David Miller studied chemistry

at the University of Oxford

(1993). He obtained his PhD

(1997) in bioorganic chemistry

under the tutelage of Prof. Tim

Bugg at the University of

Southampton developing inhibi-

tors for enzymes involved in

peptidoglycan biosynthesis.

Post-doctoral positions followed

at the University of St. Andrews

and the University of Birming-

ham under Prof. David Gani and

Prof. Rudolf Allemann. He was

appointed as a research fellow in

the School of Chemistry at

Cardiff in 2007. His research interests include terpene biosynthesis,

synthetic biology and medicinal chemistry.


diphosphate synthases.14 (+)-Germacrene D (26) synthase (GDS)

from Solidago canadensis also has an altered metal binding motif,

although in this case, it is the first DDXXD motif that takes the

form of NDTYD.15 Nevertheless, the two metal ion binding sites

in all known sesquiterpene synthases are responsible for coor-

dination of three putative metal cations and it is these that

coordinate the diphosphate group of the substrate. The binding

of the three magnesium ions is highly ordered and triggers the

closure of the active site, in which the farnesyl chain adopts

a reactive conformation sheltered from water.4,16–26

The crystal structures of (+)-aristolochene (11) synthases from

Penicillium roquefortii (PR-AS)27 and Aspergillus terreus19 (AT-

AS) (Fig. 1) offer an opportunity to compare two separately

evolved sesquiterpene synthases that generate the same product.

A shared active site contour for the two enzymes has been

revealed.19 A series of AT-AS structures were determined in the

ctures of sesquiterpene synthases. A: epi-isozizaene (17) synthase from

ethyl ammonium cation (BTAC, 51) (pdb 3KB9);23 B: (+)-d-cadinene (22)

-FDP (pdb 3G4F)14 (the catalytically silent class II-like domain is shown

complex with FDP (pdb 3BNX).21

Rudolf K: Allemann

Rudolf K Allemann studied

physics and chemistry at the

Eidgen€ossische Technische

Hochschule in Zurich (ETH-

Z), where he also received his

PhD after studies at Harvard

University and ETH-Z. He then

moved to the MRC-National

Institute for Medical Research

as a Postdoctoral Research

Fellow of the Royal Society

(London) and the Swiss

National Science Foundation.

After his habilitation at ETH-Z,

he took up positions at the

University of Birmingham,

ETH-Lausanne and Cardiff University, where he holds a Distin-

guished Research Professorship. The research of the Allemann

laboratory focuses on enzyme mechanism, catalysis and design.

Nat. Prod. Rep., 2012, 29, 60–71 | 61


Publ

ishe

d on

08

Nov

embe

r 20

11. D

ownl

oade

d on

28/

11/2

014

01:2

7:21


presence of the substrate and various fluorinated FDP

analogues.21 These crystals, which comprised the enzyme in

a tetrameric structure bound to Mg2+, PPi and FDP analogues in

various combinations provided a series of snapshots of the

enzyme in both open and closed form depending upon quater-

nary structure/crystal packing effects and the coordination state.

These structures led to the prediction of a sequence of binding

events for catalysis by AT-AS. Binding of the substrate to the

open form of the enzyme in the absence of metal ions is followed

by the coordination of Mg2+B and Mg2+C and active site closure;

the reactive conformation of FDP is formed after Mg2+A binds to

initiate the reaction cascade. Aristolochene (11), Mg2+A and Mg2+Crelease are accompanied by active site opening followed by

release of the third magnesium, Mg2+B , and diphosphate to

complete the reaction cycle.

Recently, a crystallographic analysis of epi-isozizaene (17)

synthase (EIZS) from Streptomyces coelicolor A3(2) (Fig. 1)

showed similar conformational changes upon binding of

ligands.23 The structure was solved in complex with three Mg2+

ions, PPi and benzyltrimethyl ammonium cation (BTAC, 51,

Fig. 2) as well as in complex with Hg2+; in addition, a structure of

the ligand-free EIZS with a mutation to the DDXXD motif was

also obtained. This work revealed a significant conformational

change of helix G to cap the active site upon metal binding. The

structure of trichodiene (20) synthase from Fusarium sporo-

trichioides (FS-TS) in complex with Mg2+, PPi and BTAC

showed similar conformational changes upon binding to metal

ions.22 In all these cases, it is the combination of binding to three

Mg2+ and PPi (FDP in the actual reaction) that triggers closure of

the active site, followed by the cleavage of the C–O bond in FDP.

Clearly, such a coordinated sequence of molecular events to

achieve the reactive conformation of the substrate must be

considered active enzyme participation.

Cation formation and carbocation cyclisation are of course

not separate but intimately linked events. For example, mutation

of the D100DSKD motif of FS-TS to E100DSKD resulted in an

increase in the active site volume and hence a more promiscuous

product spectrum indicating that metal ion binding and overall

active site contours are intimately linked.17 A site directed

mutagenesis (SDM) study aimed at investigating the role of the

NSE motif (typical of bacterial/fungal derived enzymes) in FS-

TS, where it was sequentially converted to a plant-like DTE

motif, showed a gradual reduction of product fidelity and cata-

lytic activity of the enzyme.20 Finally, Arg 304 of FS-TS forms

a hydrogen bond to PPi to stabilise the anion in the active site

and to ensure formation of the correct active site contour.18

These observations underline the notion that metal binding is

central to the fidelity of FS-TS catalysis.25,26,28

Recent work has shown that cation generation by sesquiter-

pene synthases need not necessarily precede the cyclisation

reactions, but the two processes often occur in a concerted

fashion. Interrogation of the mechanism of action of PR-AS

using 12,13-difluoro-farnesyl diphosphate (12,13-FF-FDP, 1n,

Fig. 4) showed that this compound was a competitive inhibitor of

PR-AS (vide infra for an explanation of the effects of fluoro

substituents).29 The fluorine atoms of 1n are distant from C1 and

should not inhibit formation of the farnesyl cation. Linear

difluoro-farnesene analogues were formed upon incubation of 1n

with the mutant enzyme F112A-PR-AS suggesting that in the

62 | Nat. Prod. Rep., 2012, 29, 60–71

absence of the phenyl ring of Phe 112 a farnesyl cation was

formed followed by proton loss to generate the farnesene prod-

ucts. This indicated that the role of Phe 112 in PR-AS is (together

with other bulky residues in the active site) to direct the distal

double bond of FDP into a reactive conformation to promote the

formation of germacrenyl cation (4) directly from FDP through

an SN2 like reaction and not via a farnesyl cation.29 This is

consistent with the observed inversion of configuration at C1

when (R)- and (S)-1-[2H]-FDP (1b and 1c, Fig. 3) were tested as

substrates of this enzyme.30 In further corroboration of the direct

displacement of PPi, 2-fluoro-farnesyl diphosphate (2F-FDP, 1k)

was turned over efficiently by PR-AS to a fluoro-germacrene A

analogue.31 Interestingly, 2F-FDP has been reported not to be

a substrate for tobacco 5-epi-aristolochene (12) synthase

(TEAS), an enzyme that also proceeds through the germacrenyl

cation (4), suggesting that a farnesyl cation is an intermediate in

the production of 12 and that the presence of fluorine prevents its

formation in this case. PR-AS plays a crucial active role in

determining the nature of the cation that is formed upon cleavage

of PPi – the catalysed reaction does not always proceed through

farnesyl cation and beyond for all sesquiterpene synthases.

Isomerisation of FDP to nerolidyl diphosphate (NDP 3) to

facilitate the production of products arising from a-bisabolyl (6),

cycloheptenyl (7), helminthogermacrenyl (8) or cis-humulenyl (9)

cations (Scheme 1) has been a difficult process to observe

experimentally. A study of the pre-steady state kinetics of FS-TS

revealed that formation of NDP is slow but once formed it reacts

onwards too quickly for NDP to be observed experimentally.32 A

recent study has revealed a single residue switch in the maize

terpene synthase 10 (TPS10) for NDP formation. TPS10

produces (mostly) a mixture of E-b-farnesene (30) and E-a-ber-

gamotene (21).33 The farnesene product arises from elimination

of H+ and PPi from FDP but the second product arises from

a downstream reaction after isomerisation to NDP. SDM of

TPS10 showed that Leu 356 controls the rate of isomerisation

from trans to cis farnesyl cations.33

3 Carbocationic reaction cascades

Following cleavage of PPi and the generation of the initial car-

bocation, sesquiterpene synthases steer the reaction through

often complex cyclisation cascades that contain additional car-

bocationic as well as neutral intermediates. In contrast to

promiscuous sesquiterpene synthases, such as g-humulene (27)

(GHS) and d-selinene (14) synthase (DSS) from Abies grandis34

or the recently discovered cadalane producing enzyme MtTPS5

from Medicago truncatula,35 high fidelity enzymes like AT- and

PR-AS or GA-DCS differentiate rigorously against alternative

reaction paths that will normally be of similar energies.

Directing the reaction along a specific reaction coordinate is

achieved through the provision of an active site template largely

made up of aromatic and aliphatic residues, where the enzyme’s

active site surface contour aligns orbitals in productive spatial

orientations, and through the stabilisation of the mostly carbo-

cationic intermediates and transition states. The latter effect has

been studied using a variety of experimental approaches,

including SDM of aromatic amino acid residues, the use of aza-

analogues, ammonium containing species, which mimic carbo-

cationic intermediates and substrate analogues designed to



Scheme 1 Early steps in the formation of cyclic sesquiterpenoids from FDP (1a) catalysed by sesquiterpene synthases and some of the resulting

products discussed in the text. Loss of H+ and PPi from FDP (1a) or NDP (3) can also lead to the linear farnesenes.

Publ

ishe

d on

08

Nov

embe

r 20

11. D

ownl

oade

d on

28/

11/2

014

01:2

7:21


intercept the reaction at specific points or to examine the regio-

and/or stereochemistry of specific steps in the catalytic cycle.

3.1 Site directed mutagenesis studies

PR-AS contains four aromatic residues in its active site (Tyr 92,

Phe 112, Phe 178 and Trp 334) and the influence of these have

been analysed in a series of SDM experiments.36–40 Mutants of

Trp 334 produce varying amounts of (�)-germacrene A (10), the

putative intermediate of PR-AS catalytic activity.41 This result

was interpreted as evidence that Trp 334 stabilises the positive

charge on the eudesmane cation, the species that is formed upon

protonation of germacrene A (10).38 Replacement of Trp 334

with para-substituted phenylalanines of increasing electron-

withdrawing properties led to a progressive accumulation of 10


with a good correlation with the interaction energies of simple

cations, such as Na+, with substituted benzenes.42 These results

provide compelling evidence for the importance of Trp 334 in

cation stabilization during the energetically demanding trans-

formation of the neutral germacrene A to eudesmane cation in

PR-AS catalysis. Tyr 92 may, in part, play a similar role to that

played by Trp 334 since incubation of FDP with PR-AS-Y92V

gave a similar product profile to that observed for PR-AS-

W334V.39 In addition, the bulkiness of Tyr 92 clearly contributes

to the accurate folding of the substrate in the active site; a series

of replacements of Tyr 92 revealed that the amounts of aristo-

lochene (11) increased with the van der Waals volume of the side

chain. When the bulkiness of residue 92 was reduced, linear

farnesenes were observed as the main products.43 While these

results suggest that cyclisation of FDP might proceed through

Nat. Prod. Rep., 2012, 29, 60–71 | 63


Scheme 2 Bifurcation in sesquiterpenoid cyclisation pathways. Through

the alterations of active site residues, tobacco 5-epi-aristolochene syn-

thase (TEAS) andHysocyamus premnaspirodiene synthase (HPS) can be

tuned to transform the shared carbocationic intermediate 34 into either

5-epi-aristolochene (12) or premnaspirodiene (13).

Publ

ishe

d on

08

Nov

embe

r 20

11. D

ownl

oade

d on

28/

11/2

014

01:2

7:21


the farnesyl cation, experiments with a fluorinated substrate

analogue indicated that the formation of the germacryl cation

was indeed a concerted process due to optimal overlap of the

p-orbital of the C2,C3 double bond with the C1–O bond facili-

tated by Tyr 92 (together with i.a. Phe 112 and Phe 178) (vide

infra).29

In studies on the bacterial enzyme pentalenene (16) synthase

(PS), the roles of the active site residues Phe 76 and Phe 77 were

examined by separate replacement with alanine, resulting in

massive reductions of the catalytic activity in both cases.

Anomalous germacrene A production by these mutants might

have been considered to be due to alternate cation stabilisation

but instead was proposed, on energetic grounds, to be a derail-

ment of the cyclisation rather than perturbation of the stabilities

of carbocations by altered amino acid residues.16 Alteration of

Mg2+ binding motifs in FS-TS has also been observed to lead to

increased promiscuity in the product spectrum.20,28

EIZS has been the subject of SDM experiments, both of the

Mg2+ binding motifs44 and the aromatic portions of the active

site.23 In the former case, as with other sesquiterpene synthases,

altered products were formed, presumably due to altered coor-

dination geometries of the metal ions at the active site cleft.44

Alteration of the active site aromatic residues Phe 95, Phe 96 and

Phe 198 resulted in alternate products but the formation of these

could not be assigned purely to either perturbation of cation–p

stabilisation or a change of the active site contour and the role of

these residues remains ambiguous.23

TEAS and Hysocyamus premnaspirodiene (13) synthase

(HPS) are two evolutionarily related enzymes that share

a common carbocationic intermediate but catalyse different

rearrangements of this cation (Scheme 2). In an impressive

demonstration of artificial control of sesquiterpene synthase

activity, various amino acid residues in spheres of increasing

radii around the active site contours of these two enzymes were

identified and changed reciprocally to those of the other enzyme.

Thereby each enzyme was converted to the activity of the other,

indicating that close control over substrate folding and structural

dynamics are crucial to product specificity and demonstrating the

potential for modification of these enzyme activities for bespoke

synthetic biology purposes. The enzyme is clearly not an unin-

volved spectator as amino acid residues in and around the active

sites of TEAS and HPS determine which of two alkyl groups

migrates from a common carbcationic intermediate.45 It should,

however, be noted here that for several sesquiterpene synthases

from plants like amorpha-4,11-diene (18)46 or d-cadinene (22)

synthases (V. Gonzalez and R. K. Allemann, unpublished

results), such an exchange of activities through the replacement

of residues in or around the active sites were unsuccessful, in that

only a reduction of the activities were observed. Saturation

mutagenesis of GA-DCS using error-prone PCRwas only able to

produce a small number of clones that produced germacrene

D-4-ol instead of d-cadinene (22).47

Another successful example of man-made control of sesqui-

terpene cyclisation is the manipulation of the highly promiscuous

GHS from Abies grandis (52 known products34) towards much

higher fidelity production of some of its minor products; after

determination of residues in the two spheres around the active

site of GHS a computer algorithm was used to predict the

mutations required to alter the activity of GHS in the desired

64 | Nat. Prod. Rep., 2012, 29, 60–71

directions. With only five mutation GHS was converted from

a promiscuous sesquiterpene synthase into a variety of highly

active enzymes with reduced product spectra (Scheme 3).48

Recently a group of sesquiterpene synthases from the fungus

Coprinus cinereus have been examined in detail.49,50 Cop6

produces predominantly (�)-a-cuprenene (42). Cop4 is much

more promiscuous, with (+)-d-cadinene (22), b-copaene (43) and

b-cubebene (44) as major products, while Cop3 makes predom-

inantly a-muurolene (45) and (+)-germacrene A (32) (Scheme 4).

The binding pockets of Cop3 and Cop4 (as judged by homology

models based on the crystal structure of AT-AS) seem to be much

larger than that of Cop6, which might explain the greater

promiscuity. A SDM study looked at the exchange of the H1-

a loop between these sesquiterpene synthases, mirroring

a previous study with an a-farnesene synthase from an apple

species.51 The H1-a loop closes over the active site upon binding,

moving the NSE motif into the correct position for catalysis.

Various mutants altered the ability of the loop to close over the

active site leading to premature quenching of the cations and

hence production of germacrenes.

An interesting case of the flexibility of sesquiterpene synthase

catalysed chemistry has been provided by studies of the biosyn-

thesis of the trisnor-sesquiterpene geosmin (50). Although not

strictly a sesquiterpene, this compound is formed by a sesquiter-

pene synthase that eliminates acetone from an intermediate ses-

quiterpenoid alcohol to generate its product (Scheme 5).

Geosmin synthase is a bifunctional enzyme from Streptomyces

coelicolor52 that possesses functionally different C-and N-

terminal catalytic domains with significant sequence similarity to

pentalenene synthase. Cleavage of the protein into the two

separate domains gave two functional enzymes; the N-terminal

half was found to convert FDP to germacradienol (48) and



Publ

ishe

d on

08

Nov

embe

r 20

11. D

ownl

oade

d on

28/

11/2

014

01:2

7:21


(�)-germacrene D (25),52 while the C-terminal half is responsible

for the conversion of germacradienol to geosmin (50).53 A series

of mutant geosmin synthases with changed DDXXD and NSE

Mg2+ binding motifs provided insight into the mechanism; most

notably the S233A mutant produced wild-type products together

with 11% of isolepidozene (47), a neutral putative cyclopropyl

intermediate that is attacked by water under acid catalysis to give

germacradienol prior to conversion to geosmin.53

3.2 Aza-analogues

An alternative approach to SDM for the study of cation stabi-

lisation and conformational control in sesquiterpene synthase

catalysis is the use of synthetic analogues of the proposed

cationic intermediates. Nitrogen containing compounds that are

either quaternised or simply protonated in aqueous solution to

generate a cation, have been used in kinetic studies and as

structural analogues for single crystal X-ray analyses (for

examples of aza analogues see Fig. 2).

Scheme 3 Site directed mutagenesis of g-humulene synthase fromG. aroreum

the above sesquiterpene products.

Scheme 4 The major products generated by sesquiterpene synthases

Cop3 and Cop6 from Coprinus cinereus.

Fig. 2 Aza-analogues of sesquiterpenoid cationic intermediates.


In the crystal structure of wild-type EIZS, BTAC (51) was

observed to orient itself with aromatic residues.23 The crystal

structure of EIZS-F198A coordinated to three Mg2+ ions, PPi

and BTAC revealed a different orientation of the cation relative

to the wild type structure, implying that the resulting expansion

of the active site volume allowed other orientations of cation

binding and hence altered product specificity.23

A recent crystal structure of FS-TS-Y305F in complex with the

potent inhibitor 53 indicated that this aza-analogue bound to the

enzyme in a conformation that did not mimic the proposed

orientation of a-bisabolyl cation (6) during the catalytic

(GHS) has been used to alter the enzyme to a specific synthase for each of

Nat. Prod. Rep., 2012, 29, 60–71 | 65


Scheme 5 The mechanism for the biosynthesis of geosmin (50).

Publ

ishe

d on

08

Nov

embe

r 20

11. D

ownl

oade

d on

28/

11/2

014

01:2

7:21


cycle.17,54 Favourable interactions with PPi seemed to cause the

observed binding orientation rather than cation–p interactions

with aromatic residues, suggesting that sesquiterpene synthases

enforce cation location and formation through kinetic rather

than thermodynamic control.17

Use of BTAC in crystal structure analysis of mutant FS-TS

provided the first definitive proof that a cation can bind in the

active site of a sesquiterpene synthase since aza-bisabolyl cations

were of unknown protonation state in other known structures.22

An apparent ‘loosening’ of the coordination of PPi and Mg2+ in

FS-TS structures led to the interpretation that this leads to

greater versatility of PPi to act as a general base (vide infra) and

for the observed diversity of products for this enzyme.

Recently, aza-analogues designed to resemble eudesmane

cation have been prepared and tested as inhibitors of PR-AS. In

contrast to the observations discussed above for FS-TS, one of

these compounds (54, Fig. 2) proved to be a potent inhibitor of

PR-AS in both the presence and absence of PPi. This suggested

that this compound may be a genuine mimic of eudesmane cation

since no obvious favourable interactions with PPi were detec-

ted.55 The trigonal aza-analogues 55 and 56, designed to mimic

the likely transition state between (�)-germacrene A (10) and

eudesmane cation (33), inhibit PR-AS synergistically with PPi,

implying a role for PPi in carbocation formation, perhaps as the

active site acid.56

Scheme 6 Mechanisms for the biosynthesis of amorpha-4,11-diene (18).

3.3 Substrate analogues

Many of the finer points in the mechanism of formation of inter

alia (+)-aristolochene (11),30,57 trichodiene (20),32,58 d-cadinene

(22),59,60 epi-cubenol (24),61–64 pentalenene (16),65–67 and (�) and

(+)-germacrene D (25 and 26)68 were uncovered using deuter-

ated, tritiated and carbon-14 labelled analogues of FDP and

NDP. Much of this work has been reviewed previously.3,4,11

More recently, the catalytic mechanism of amorpha-4,11-diene

synthase from Artemisia annua (ADS) has been studied using

labelled FDP analogues. In contrast to the antimalarial

compound artemisinin, which is synthesised from amorpha-4,11-

diene, ADS has not been studied extensively from a mechanistic

point of view. Two parallel studies with (R)- and (S)-[1-2H]-FDP

66 | Nat. Prod. Rep., 2012, 29, 60–71

(1b and 1c) established a 1,3-shift of HSi on C1 of FDP to C7

implying a mechanism that proceeds via a-bisabolyl cation (6),

which is formed by 1,6 ring closure of NDP (Scheme 6).69,70

Interestingly, deuterium incorporation was observed upon

incubation in >75% 2H2O. Although the position of incorpora-

tion was not determined, the authors speculated that an inter-

mediate cation may be quenched and reprotonated as a side

reaction from the main catalytic pathway.70 This may hint at the

alternate reaction pathways to some of the minor products that

have been observed for this enzyme and present opportunities for

engineering altered reactivities into this enzyme.71 It will be

interesting to see how the enzyme promotes 1,6 ring closure of cis

farnesyl cation rather than the 1,10 ring closure catalysed by

cadalane producing sesquiterpene synthases.

Further to the SDM studies on the related enzymes TEAS and

HPS, deuterium labelling experiments were used to analyse the

stereochemistry and timing of protonation, rearrangement and

elimination steps for these two enzymes.72 Incubation with both

enantiomers of [8-2H]-FDP (1d and 1e), tris- and hexa-deuterated

FDP analogues at C12 and C13 (1f, 1g and 1h) and incubation of

unlabelled FDP in 2H2O gave results that were broadly in line

with mechanistic observations for the fungal aristolochene syn-

thases,57,72,73 namely that the proton lost upon formation of the

intermediate germacrene A is from C12; the intermediate is then

reprotonated from the Re face and HSi is lost from C8 to form

the final product.

A new trend in sesquiterpene synthase research is the hunt for

new activities using genome mining. Two presumptive sesqui-

terpene synthase genes from Streptomyces clavuligerus were

recently identified and codon optimised synthetic genes con-

structed and overexpressed in E. coli. The gene products were

identified as related sesquiterpene synthases producing the

cadalane family sesquiterpenoids (-)-d-cadinene (22) and (+)-T-

muurolol (23).74 Incubation of the enzymes with 1b and 1c and

analysis of the products by 1H and 2H NMR spectroscopy

revealed a [1,3]-hydride shift of HSi on C1 of FDP to C11 in the

reactions catalysed by both enzymes. This is in line with a similar

stereochemical prediction for the hydride shift observed for GA-

DCS.59 Further to this, a promiscuous cadalane producing

sesquiterpene synthase MtTPS5 from Medicago truncatula has

recently been identified.35 Incubations of unlabelled FDP in2H2O showed C15 proton/deuteron exchange, which was pre-

dicted through an alternative mechanism for cadalane produc-

tion that was first postulated many years ago but until this report



Publ

ishe

d on

08

Nov

embe

r 20

11. D

ownl

oade

d on

28/

11/2

014

01:2

7:21


had no experimental support.75 Some of the products appeared

to be generated via germacrene D, which after protonation at

C15 reacts to a variety of bicyclic products. Interestingly, the

alteration of GA-DCS by saturation mutagenesis led to an

enzyme that produced a close analogue of germacrene D,47

implying that only subtle differences in enzyme structure and

dynamics differentiate between production of germacrene D or

germacrene D-4-ol and d-cadinene from a common intermediate.

Scheme 7 Biosynthesis of epi-isozizaene (17).

Scheme 8 The biosynthesis of avermitilol (66). Virifloridol (67), ger-

macrene A (32) and germacrene B (63) are minor byproducts.

Fig. 3 Deuterated substrate analogues discussed in the text.

Another enzyme that produces germacrene D as a by-product

is geosmin synthase. Interrogation of the mechanism of this

enzyme using 1b and 1c and analysis of the products by GC-MS

established that HSi on C1 of FDP is the species that undergoes

[1,3]-hydride transfer in this case.76 Despite producing the same

enantiomer of germacrene D, (-)-GDS from Solidago canadensis

proceeds via [1,3]-migration of HRe on C1 of FDP indicating that

the two enzyme must convert FDP to (-)-germacrene D (25)

through different catalytic mechanisms.68,76 The mechanism of

the dominant geosmin production was established to result from

retro-Prins fragmentation of germacradienol by labelling studies.

Incubation of unlabelled FDP in 2H2O resulted in specific

incorporation of several deuterium atoms in geosmin and the

by-products although not in the (-)-germacrene D.77 The use of

tris-deuterated FDP at C13 (1h) resulted in trapped acetone

containing the label. This confirmed the retro-Prins fragmenta-

tion of 47 and the use of 1j as a substrate and analysis of the

resulting product demonstrated a predicted [1,2]-hydride shift

from C2 of FDP.78

Genome mining studies first identified EIZS from Strepto-

myces coelicolor. Incubation with 1b and 1c, [1-2H2]-FDP (1j)

and the trisdeutero-FDP analogues 1g and 1h established the

current understanding of the mechanism of this reaction

including the stereochemistry of the methyl migration in the

penultimate step (Scheme 7).44,79

Another genome mining study, this time of Streptomyces

avermitilis, identified a gene encoding a previously undiscovered

sesquiterpene synthase that produces the eponymous avermitilol

(66).80 The stereochemistry of the ring closures was elucidated

using 1b and 1c. The enzyme also produces small amounts of

(+)-germacrene A (32), germacrene B (63) and viridiflorol (67)

(Scheme 8).80

The product of presilphiperfolan-8b-ol (72) synthase from

Botrytis cinerea is a common precursor in the biosynthesis of


a series of toxins produced by this necrotic plant pathogen.81

With the use of deutero-FDP analogues it was established that

the reaction sequence proceeds via humulyl cation (5) and does

not require the intermediacy of NDP (3) (Scheme 9), although

this compound is an alternate substrate of this enzyme.

Fluorinated isoprenyl diphosphates have been used in the past

to study the mechanism of enzymes involved in terpene biosyn-

thesis,82–84 but only recently have they been extensively employed

for the analysis of sesquiterpene synthases. Their behaviour as

either inhibitors or substrates has revealed several subtle mech-

anistic details that may not otherwise have come to light. The

crystal structures of AT-AS and GA-DCS have now been solved

in the presence of several fluorinated sesquiterpene analogues

(vide supra).14,21 Fig. 4 illustrates the fluorinated FDP analogues

that have been used in this work and also shows several of the

fluorinated sesquiterpenoids produced with various sesquiter-

pene synthases.

Nat. Prod. Rep., 2012, 29, 60–71 | 67


Scheme 9 Mechanism for the biosynthesis of presilphiperfolan-8b-ol

(72).

Fig. 4 Fluorinated FDP analogues referred to in the text and some of

the germacrene A analogues resulting from turnover of these analogues

by sesquiterpene synthases.

Publ

ishe

d on

08

Nov

embe

r 20

11. D

ownl

oade

d on

28/

11/2

014

01:2

7:21


Fluorinated FDP analogues have been used to interrogate the

mechanism of the PR-AS; 12,13-FF-FDP (1n) was instrumental

in showing the role of Phe 112 (and other bulky active site resi-

dues) in aligning the distal double bond for attack at C1 during

PRAS catalysis (vide supra).29 In addition, 2F-FDP (1k),31 6F-

FDP(1l)85 and 14F-FDP (1o)85 have proved to be efficient

substrates of PR-AS, which converts these fluorinated FDPs to

analogues of (�)-germacrene A (73–75, Fig. 4). This provides

further evidence of the intermediacy of germacrene A in the

production of eremophilenyl sesquiterpenes such as (+)-aristo-

lochene (11) since the presence of fluorine on C2 or C6 prevents

the cyclisation of (�)-germacrene A by reduction of electron

68 | Nat. Prod. Rep., 2012, 29, 60–71

density on the C2,C3 or C6,C7 double bonds, respectively, and

also by preventing the build up of positive charge on C3 in the

eudesmane cation (34). In the case of fluorine on C14, there is

again a depletion of electron density in the C6,C7-double bond,

which reduces the p-basicity and so prevents protonation of 10.

It also shows that these enzymes can be utilised to produce

analogues of natural products that may be exploitable in the

future for medicinal or agrochemical uses. 6F-FDP (1l) is also an

efficient substrate of TEAS, giving the corresponding fluorinated

(+)-germacrene A analogue (76) as the sole product.86

The inclusion of TEAS amongst the so-called high-fidelity

sesquiterpene synthases has recently been challenged by the

observation of several minor products in its product pool

originating from both transoid and cisoid ring closures of

FDP.87 The amount of cisoid products increased dramatically

when the enzyme was incubated with the 2Z isomer of FDP.88

In addition, crystals of TEAS and a promiscuous mutant

(M4 TEAS) were soaked with 2F-FDP and 2-fluoro-2E-FDP to

provide snapshots of the binding modes of substrates in

conformations leading to both of these cyclisation pathways.89

Comparison of the X-ray structures showed that both substrates

were bound in the active site with the farnesyl chain in

a U-shaped conformation but with differing orientations

(inversions) of the first two isoprene units – hence showing

complete occupancy of the active site and also displaying

conformations consistent with the expected orientation that

would lead to their observed product pools. The cisoid substrate

analogue, however, was not bound in an active conformation

since the C–O bond was coplanar with the C2,C3 bond, in

contrast to the transoid substrate where the C–O bond was

perpendicular to this and hence oriented optimally for the

reaction. These results show that while the enzyme active site

surface contour is important for the control of product

outcome, it appears that an interplay exists between enzymatic

enforcement of substrate conformation and the intrinsic

conformational possibilities of the substrate; the use of a cisoid

C2,C3 double bond can produce dramatically altered products.

Interestingly, it was also observed that several of the mutations

in the M4 mutant TEAS cause little change to the contour of

the active site. Since this mutant is much more promiscuous

than the wild-type enzyme, this hints strongly at dynamic effects

playing an important part in the product outcome for this

enzyme. The electron density of the farnesyl chain in the crystal

structures of this mutant soaked with the two substrate

analogues showed discontinuities, perhaps indicating increased

motional flexibility of the substrate in the active site relative to

the wild type enzyme; this is fully consistent with the promis-

cuity observed for this mutant when incubated with FDP.

In agreement with results revealing active site plasticity in

GHS,48 the active site contour of PR-AS has been shown to be

highly plastic for binding FDP analogues. Three phenyl-FDP

analogues are potent competitive inhibitors of this enzyme.90

This reveals potential for synthetic biology applications, since it

may be possible to engineer a larger active site cavity to allow

productive binding of such bulky substrate analogues. It also

reveals that the active sites of these enzymes are not static and

rigid cavities but are able to adopt different shapes to accom-

modate bulkier ligands hinting at the dynamic role played by

these proteins during catalysis.



Scheme 10 The mechanism of formation of (S)-b-bisabolene (77) and

(S)-b-macrocarpene (19) catalysed by TPS6 and TPS11 from maize.

Scheme 11 Patchouli alcohol (80) and two of the minor hydrocarbon

products, b-patchoulene (81) and a-guaiane (82) produced by patchouli

alcohol synthase from Pogostemon cablin.

Publ

ishe

d on

08

Nov

embe

r 20

11. D

ownl

oade

d on

28/

11/2

014

01:2

7:21


4 Acid/base catalysis

Many sesquiterpene synthases require participation of acids and

bases during their catalytic cycle. For example, the two enan-

tiomers of germacrene A are thought to be neutral intermediates

in the pathways to (+)-aristolochene (11) (via 10) and 5-epi-

aristolochene (via 32) requiring an acid to regenerate a cation;

a base is then required in the final step of the reaction.11 Various

candidates have been proposed to act as active site acids/bases

including amino acid side chains, active site water molecules as

well as the diphosphate generated from the substrate itself.

Based on the proposal that Tyr 520 may be the active site acid

responsible for protonation of germacrene A in catalysis by

TEAS,91 it was suggested that the analogous function is fulfilled

by Tyr 92 in the active site of PR-AS.27 However, since

replacements of Tyr 92 in PR-AS lead to enzymes that still

produce significant quantities of (+)-aristolochene (11), Tyr 92

cannot be the active site acid.40 PPi has since been suggested to

act as the acid/base for AT-AS on the basis of crystallographic

analysis but there remains little other than structural evidence for

this.19 Meanwhile, a theoretical study proposed that instead of

being a reaction intermediate, (�)-germacrene A (10) could be

a by-product of PR-AS produced via a different catalytic

mechanism to the major product.92 Instead, it was proposed

based on gas-phase calculations that there may be a direct

hydride transfer between C12 and C6, or a proton transfer via an

intermediary water molecule. Such a mechanism would bypass

the neutral germacrene A and avoid an energetically expensive

sequence of cation quenching followed by cation regeneration.92

This attractive mechanistic proposal had to be rejected, however,

since in deuterium labelling experiments the stereochemistry of

protonation at C6 was inconsistent with either direct hydride

transfer or transfer by way of a water molecule.73 This labelling

study established that the protonation of 10 occurs on the same

face of the molecule as the later deprotonation event that

produces the final product, implying that the same species is

involved in both reaction steps.73 A similar study on TEAS

showed that protonation occurs on the same face of (+)-germa-

crene A in TEAS catalysis.72

The idea of tyrosine as an active-site acid in TEAS has proved

enticing for other sesquiterpene synthases also. Y295 has been

proposed as the active site base responsible for final deprotona-

tion in FS-TS catalysis – however, SDM of this residue ruled out

this possibility, since mutants had similar activity to the wild type

enzyme. Once more the PPi group most likely acts as the proton

donor/acceptor.20

Further enzymes where tyrosine has been investigated as an

active site acid are the (S)-b-macrocarpene (19) synthases TPS6

and TPS11 frommaize, which possess a tyrosine residue (Tyr 522

in both) that appears to be an evolutionarily conserved residue

(cf. Tyr 520 in TEAS).33 (S)-b-Macrocarpene was shown by

deuterium labelling studies to arise from protonation of a neutral

intermediate ((S)-b-bisabolene, 77) and SDM of Y522 sup-

pressed this protonation activity yielding b-bisabolene as

product (Scheme 10). Additionally, protonation of (S)-b-bisa-

bolene was found to be bothMg2+ and pH dependent, as has also

been reported for TEAS protonation of (+)-germacrene A (32).93

A similar pH sensitivity has been observed to affect the product

profile of Cop4 from Coprinus cinereus. This, however, appears


to be a protein folding effect since SDM experiments where H235

and N239 were altered to proline and leucine, respectively,

resulted in abolition of the pH dependence of product

formation - probably due to hydrogen bonding changes within

the enzyme.49 Whatever the specific identity of active site acids/

bases in various sesquiterpene synthases, it is clear that amino

acid residues are at least in part responsible for such processes in

some cases – notably TEAS and in TPS6 and TPS 11 from maize

and that the enzyme thereby plays a crucial active chemical role

in the catalysis.

Finally, a recent interrogation of patchouli alcohol (80,

Scheme 11) synthase from Pogostemon cablin using the singly

labelled [2-2H]-FDP 1j showed a surprising result. In addition to

the expected singly labelled patchouli alcohol product (�65% of

isolated alcoholic material), a doubly deuterated product was

also observed (�35% of isolated alcohols).94 From the positions

of deuteration it was concluded that an intermediate alkene

structure becomes protonated (or deuterated when 1j is used as

substrate) to give the final product. Minor hydrocarbon by-

products b-patchoulene (81) and a-guaiene (82) were depleted of

deuterium, indicating the source of the second deuteron.

Comparison of the amino acid sequence with other sesquiterpene

synthases and use of an active site model identified Leu 410 as an

active site residue that may undergo reorientation during the

catalytic cycle, effectively creating a second pocket in the active

site were the observed deuteration events occur.94 This may be

another example of active enzyme participation in catalysis.

Nat. Prod. Rep., 2012, 29, 60–71 | 69


Publ

ishe

d on

08

Nov

embe

r 20

11. D

ownl

oade

d on

28/

11/2

014

01:2

7:21


5 Conclusions

All of the work described here has helped unravel the general

principles that control the mode of action of sesquiterpene

synthases. Even though not the subject of this review, it is worth

noting that the same principles will also operate during catalysis

by mono- and diterpene synthases. Despite a lack of significant

sequence homology between sesquiterpene synthases in plants,

insects, bacteria and fungi, the chemistry that they facilitate

within the shared class I terpene synthase fold is the basis for

the generation of the enormous diversity in structure and

stereochemistry found in natural sesquiterpenes. The plasticity

of the sesquiterpene synthases appears to provide a framework

for the combinatorial production of many natural terpenoids

though subtle alterations in the composition of the active site

during evolution. The enzymes bind their substrates in reactive

conformations that take advantage of the chemical reactivity

inherent in FDP. This first binding step is central for the initi-

ation of the Mg2+-dependent expulsion of diphosphate and the

product profile. Perhaps not surprisingly, there is complexity in

the formation of the Michaelis complex of PR-AS21 and perhaps

all sesquiterpene synthases, in that after the initial molecular

recognition of FDP, coordination to Mg2+B reorients the

substrate diphosphate group. Subsequently, binding of Mg2+Ctriggers a more substantive conformational change leading to

active site closure. Finally, binding of Mg2+A completes the

formation of the trinuclear metal ion cluster and triggers further

structural changes to complete active site closure and to

generate the cyclisation-competent conformation of FDP. In the

Michaelis complex the three negative charges of the PPi leaving

group are fully neutralized to facilitate ionization and initiation

of the cyclisation cascade. It is likely that diphosphate stays

bound throughout the reaction thereby providing stability to

the carbocationic transition states and intermediates. The parts

of the active site that bind the isoprenoid tail of the substrate

are largely made up of aliphatic and aromatic amino acids to

allow additional stabilisation of the carbocations of the reaction

cascades through cation–p or charge–quadrupole interactions.

The specific orientation of the active site residues allows the

enzymes to channel the reaction along a specific pathway to

generate the desired outcome and at the same time strictly

controls water access.

Finally, general acid and base catalysis is involved in

terpene synthase reactions. There has so far been little

evidence that the enzymes provide amino acids to act as acids

and bases and, by exclusion, it is now assumed that in many

cases the bound diphosphate ion helps move protons. The

selectivities of the protonation and deprotonation reactions

depend on the correct orientation of acid and bases with in the

active sites.

Sesquiterpene synthases play an active controlling role in

catalysis; they provide a fascinating example of how enzymes

have evolved to take advantage of the chemical properties

inherent in their substrates to generate a biologically desirable

outcome with remarkable efficiency and specificity. (Sesqui)-

terpene synthases, the chemistry they facilitate, their biology

and their applications will no doubt continue to astound us

through a combination of structural complexity and chemical

simplicity.

70 | Nat. Prod. Rep., 2012, 29, 60–71

6 Acknowledgments

We are grateful for the financial support of the Biotechnology

and Biological Sciences Research Council (grants 6/B17177; BB/

G003572/1 and BB/H01683X/1), the Engineering and Physical

Sciences Research Council (grant EP/D06958/1) and Cardiff

University. We thank Dr Juan A. Faraldos, Dr Sabrina Touchet,

DrMahmoud Akhtar, Dr RobertMart and Dr E. Joel Loveridge

for critical comments on the manuscript and past and present

members of the terpene cyclase group for stimulating

discussions.

7 References

1 J. D. Connolly and R. A. Hill, Dictionary of Terpenoids, Chapmanand Hall, London, 1991.

2 J. S. Glasby, Encyclopedia of Terpenoids, Wiley, Chichester, 1982.3 J. Degenhardt, T. G. Kollner and J. Gershenzon, Phytochemistry,2009, 70, 1621.

4 D. W. Christianson, Chem. Rev., 2006, 106, 3412.5 E. M. Davis and R. Croteau, Biosynthesis: Aromatic Polyketides,Isoprenoids, Alkaloids, Springer, Berlin Heidelberg, 2000, vol. 209,pp. 53–95.

6 E. M. Davis, in Comprehensive Natural Products II, ed. L. Manderand H.-W. Liu, Elsevier, 2010, vol. 1, pp. 585–608.

7 T. Toyomasu and T. Sassa, in Comprehensive Natural Products II, ed.L. Mander and H.-W. Liu, Elsevier, 2010, vol. 1, pp. 643–672.

8 T. Kushiro and Y. Ebizuka, in Comprehensive Natural Products II, ed.L. Mander and H.-W. Liu, Elsevier, 2010, vol. 1, pp. 673–708.

9 D. R. Phillips, J. M. Rasbery, B. Bartel and S. P. T. Matsuda, Curr.Opin. Plant Biol., 2006, 9, 305.

10 R. Xu, G. C. Fazio and S. P. T. Matsuda, Phytochemistry, 2004, 65,261.

11 D. Cane, Chem. Rev., 1990, 90, 1089.12 D. J. Tantillo, Nat. Prod. Rep., 2011, 28, 1035.13 J. Chappell and R. M. Coates, in Comprehensive Natural Products II,

ed. M. L. and H.-W. Liu, Elsevier, 2010, vol. 1, pp. 609–641.14 H. A. Gennadios, V. Gonzalez, L. Di Costanzo, A. A. Li, F. L. Yu,

D. J. Miller, R. K. Allemann and D. W. Christianson,Biochemistry, 2009, 48, 6175.

15 I. Prosser, I. G. Altug, A. L. Phillips, W. A. Konig, H. J. Bouwmeesterand M. H. Beale, Arch. Biochem. Biophys., 2004, 432, 136.

16 M. Seemann, G. Z. Zhai, J. W. de Kraker, C. M. Paschall,D. W. Christianson and D. E. Cane, J. Am. Chem. Soc., 2002, 124,7681.

17 L. S. Vedula, M. J. Rynkiewicz, H. J. Pyun, R. M. Coates, D. E. Caneand D. W. Christianson, Biochemistry, 2005, 44, 6153.

18 L. S. Vedula, D. E. Cane and D.W. Christianson, Biochemistry, 2005,44, 12719.

19 E. Y. Shishova, L. Di Costanzo, D. E. Cane and D. W. Christianson,Biochemistry, 2007, 46, 1941.

20 L. S. Vedula, J. Y. Jiang, T. Zakharian, D. E. Cane andD. W. Christianson, Arch. Biochem. Biophys., 2008, 469, 184.

21 E. Y. Shishova, F. L. Yu, D. J. Miller, J. A. Faraldos, Y. X. Zhao,R. M. Coates, R. K. Allemann, D. E. Cane andD. W. Christianson, J. Biol. Chem., 2008, 283, 15431.

22 L. S. Vedula, Y. X. Zhao, R. M. Coates, T. Koyama, D. E. Cane andD. W. Christianson, Arch. Biochem. Biophys., 2007, 466, 260.

23 J. A. Aaron, X. Lin, D. E. Cane and D. W. Christianson,Biochemistry, 2010, 49, 1787.

24 C. A. Lesburg, G. Z. Zhai, D. E. Cane and D. W. Christianson,Science, 1997, 277, 1820.

25 M. J. Rynkiewicz, D. E. Cane and D. W. Christianson, Biochemistry,2002, 41, 1732.

26 M. J. Rynkiewicz, D. E. Cane and D. W. Christianson, Proc. Natl.Acad. Sci. U. S. A., 2001, 98, 13543.

27 J. M. Caruthers, I. Kang, M. J. Rynkiewicz, D. E. Cane andD. W. Christianson, J. Biol. Chem., 2000, 275, 25533.

28 D. E. Cane, Q. Xue and J. E. VanEpp, J. Am. Chem. Soc., 1996, 118,8499.

29 F. L. Yu, D. J. Miller and R. K. Allemann, Chem. Commun., 2007,4155.



Publ

ishe

d on

08

Nov

embe

r 20

11. D

ownl

oade

d on

28/

11/2

014

01:2

7:21


30 D. E. Cane, P. C. Prabhakaran, E. J. Salaski, P. H. M. Harrison,H. Noguchi and B. J. Rawlings, J. Am. Chem. Soc., 1989, 111, 8914.

31 D. J. Miller, F. L. Yu and R. K. Allemann, ChemBioChem, 2007, 8,1819.

32 D. E. Cane, H. T. Chiu, P. H. Liang and K. S. Anderson,Biochemistry, 1997, 36, 8332.

33 T. G. Kollner, J. Gershenzon and J. Degenhardt, Phytochemistry,2009, 70, 1139.

34 C. L. Steele, J. Crock, J. Bohlmann and R. Croteau, J. Biol. Chem.,1998, 273, 2078.

35 S. Garms, T. G. Kollner and W. Boland, J. Org. Chem. 75, p. 5590.36 S. Forcat and R. K. Allemann, Org. Biomol. Chem., 2006, 4, 2563.37 S. Forcat and R. K. Allemann, Chem. Commun., 2004, 2094.38 A. Deligeorgopoulou, S. E. Taylor, S. Forcat and R. K. Allemann,

Chem. Commun., 2003, 2162.39 M. J. Calvert, S. E. Taylor and R. K. Allemann, Chem. Commun.,

2002, 2384.40 B. Felicetti and D. E. Cane, J. Am. Chem. Soc., 2004, 126, 7212.41 M. J. Calvert, P. R. Ashton and R. K. Allemann, J. Am. Chem. Soc.,

2002, 124, 11636.42 J. A. Faraldos, A. K. Antonczak, V. G. Gonz�alez, R. Fullerton,

E. M. Tippmann and R. K. Allemann, J. Am. Chem. Soc., 2011,133, 13906.

43 A. Deligeorgopoulou and R. K. Allemann, Biochemistry, 2003, 42,7741.

44 X. Lin and D. E. Cane, J. Am. Chem. Soc., 2009, 131, 6332.45 B. T. Greenhagen, P. E. O’Maille, J. P. Noel and J. Chappell, Proc.

Natl. Acad. Sci. U. S. A., 2006, 103, 9826.46 M. K. Julsing, PhD thesis, University of Groningen, 1975.47 Y. Yoshikuni, V. J. J. Martin, T. E. Ferrin and J. D. Keasling, Chem.

Biol., 2006, 13, 91.48 Y. Yoshikuni, T. E. Ferrin and J. D. Keasling, Nature, 2006, 440,

1078.49 F. Lopez-Gallego, G. T. Wawrzyn and C. Schmidt-Dannert, Appl.

Environ. Microbiol., 2010, 76, 7723.50 F. Lopez-Gallego, S. A. Agger, D. Abate-Pella, M. D. Distefano and

C. Schmidt-Dannert, ChemBioChem, 2010, 11, 1093.51 S. Green, C. J. Squire, N. J. Nieuwenhuizen, E. N. Baker and

W. Laing, J. Biol. Chem., 2009, 284, 8652.52 D. E. Cane and R.M.Watt, Proc. Natl. Acad. Sci. U. S. A., 2003, 100,

1547.53 J. Y. Jiang, X. F. He and D. E. Cane, Nat. Chem. Biol., 2007, 3, 711.54 D. E. Cane, G. H. Yang, R.M. Coates, H. J. Pyun and T.M. Hohn, J.

Org. Chem., 1992, 57, 3454.55 J. A. Faraldos, B. Kariuki and R. K. Allemann, J. Org. Chem., 2010,

75, 1119.56 J. A. Faraldos and R. K. Allemann, Org. Lett., 2011, 13, 1202.57 D. E. Cane, P. C. Prabhakaran, J. S. Oliver and D. B. McIlwaine, J.

Am. Chem. Soc., 1990, 112, 3209.58 D. E. Cane and Q. Xue, J. Am. Chem. Soc., 1996, 118, 1563.59 C. R. Benedict, J. L. Lu, D. W. Pettigrew, J. G. Liu, R. D. Stipanovic

and H. J. Williams, Plant Physiol., 2001, 125, 1754.60 I. Alchanati, J. A. A. Patel, J. G. Liu, C. R. Benedict,

R. D. Stipanovic, A. A. Bell, Y. X. Cui and C. W. Magill,Phytochemistry, 1998, 47, 961.

61 K. Nabeta,M. Fujita, K. Komuro, K. Katayama and T. Takasawa, J.Chem. Soc., Perkin Trans. 1, 1997, 2065.

62 K. Nabeta, K. Kigure, M. Fujita, T. Nagoya, T. Ishikawa,H. Okuyama and T. Takasawa, J. Chem. Soc., Perkin Trans. 1,1995, 1935.


63 D. E. Cane, M. Tandon and P. C. Prabhakaran, J. Am. Chem. Soc.,1993, 115, 8103.

64 D. E. Cane and M. Tandon, Tetrahedron Lett., 1994, 35,5355.

65 D. E. Cane and S. W. Weiner, Can. J. Chem., 1994, 72, 118.66 D. E. Cane, J. S. Oliver, P. H. M. Harrison, C. Abell, B. R. Hubbard,

C. T. Kane and R. Lattman, J. Am. Chem. Soc., 1990, 112, 4513.67 P. H. M. Harrison, J. S. Oliver and D. E. Cane, J. Am. Chem. Soc.,

1988, 110, 5922.68 C. O. Schmidt, H. J. Bouwmeester, S. Franke and W. A. Konig,

Chirality, 1999, 11, 353.69 S. Picaud, P. Mercke, X. F. He, O. Sterner, M. Brodelius, D. E. Cane

and P. E. Brodelius, Arch. Biochem. Biophys., 2006, 448, 150.70 S. H. Kim, K. Heo, Y. J. Chang, S. H. Park, S. K. Rhee and

S. U. Kim, J. Nat. Prod., 2006, 69, 758.71 P. E. O’Maille, J. Chappell and J. P. Noel, Anal. Biochem., 2004, 335,

210.72 D. J. Schenk, C. M. Starks, K. R. Manna, J. Chappell, J. P. Noel and

R. M. Coates, Arch. Biochem. Biophys., 2006, 448, 31.73 D. J. Miller, J. L. Gao, D. G. Truhlar, N. J. Young, V. Gonzalez and

R. K. Allemann, Org. Biomol. Chem., 2008, 6, 2346.74 Y. Hu, W. K. W. Chou, R. Hopson and D. E. Cane, Chem. Biol.,

2011, 18, 32.75 D. Arigoni, Pure Appl. Chem., 1975, 41, 219.76 X. F. He and D. E. Cane, J. Am. Chem. Soc., 2004, 126, 2678.77 J. Y. Jiang, X. F. He and D. E. Cane, J. Am. Chem. Soc., 2006, 128,

8128.78 J. Y. Jiang and D. E. Cane, J. Am. Chem. Soc., 2008, 130, 428.79 X. Lin, R. Hopson and D. E. Cane, J. Am. Chem. Soc., 2006, 128,

6022.80 W. K.W. Chou, I. Fanizza, T. Uchiyama, M. Komatsu, H. Ikeda and

D. E. Cane, J. Am. Chem. Soc., 2010, 132, 8850.81 C. M. Wang, R. Hopson, X. Lin and D. E. Cane, J. Am. Chem. Soc.,

2009, 131, 8360.82 C. D. Poulter, J. C. Argyle and E. A. Mash, J. Biol. Chem., 1978, 253,

7227.83 R. Croteau, Arch. Biochem. Biophys., 1986, 251, 777.84 D. E. Cane, G. H. Yang, Q. Xue and J. H. Shim, Biochemistry, 1995,

34, 2471.85 D. J. Miller, F. L. Yu, D. W. Knight and R. K. Allemann, Org.

Biomol. Chem., 2009, 7, 962.86 J. A. Faraldos, Y. X. Zhao, P. E. O’Maille, J. P. Noel and

R. M. Coates, ChemBioChem, 2007, 8, 1826.87 P. E. O’Maille, J. Chappell and J. P. Noel, Arch. Biochem. Biophys.,

2006, 448, 73.88 J. A. Faraldos, P. E. O’Maille, N. Dellas, J. P. Noel and R.M. Coates,

J. Am. Chem. Soc., 2010, 132, 4281.89 J. P. Noel, N. Dellas, J. A. Faraldos, M. Zhao, B. A. Hess,

L. Smentek, R. M. Coates and P. E. O’Maille, ACS Chem. Biol.,2010, 5, 377.

90 D. J. Miller, F. L. Yu, N. J. Young and R. K. Allemann,Org. Biomol.Chem., 2007, 5, 3287.

91 K. A. Rising, C. M. Starks, J. P. Noel and J. Chappell, J. Am. Chem.Soc., 2000, 122, 1861.

92 R. K. Allemann, N. J. Young, S. H. Ma, D. G. Truhlar and J. L. Gao,J. Am. Chem. Soc., 2007, 129, 13008.

93 P. E. O’Maille, B. Greenhagen, J. Chappell, Y. Zhao, R. M. Coatesand J. P. Noel, in Terpnet 2003, Lexington, Kentucky USA, 2003.

94 J. A. Faraldos, S. Q. Wu, J. Chappell and R.M. Coates, J. Am. Chem.Soc., 2010, 132, 2998.

Nat. Prod. Rep., 2012, 29, 60–71 | 71


sesquiterpene synthases: passive catalysts or active players?€¦ · dedicated to these enzymes...

Documents