REVIEW ARTICLE
Structure, mechanism and function of prenyltransferases
Po-Huang Liang, Tzu-Ping Ko and Andrew H J. Wang
Institute of Biological Chemistry, Academia Sinica, Taipei, Taiwan
In this review, we summarize recent progress in studying
three main classes of prenyltransferases: (a) isoprenyl pyro-
phosphate synthases (IPPSs), which catalyze chain elonga-
tion of allylic pyrophosphate substrates via consecutive
condensation reactions with isopentenyl pyrophosphate
(IPP) to generate linear polymers with defined chain lengths;
(b) protein prenyltransferases, which catalyze the transfer of
an isoprenyl pyrophosphate (e.g. farnesyl pyrophosphate) to
a protein or a peptide; (c) prenyltransferases, which catalyze
the cyclization of isoprenyl pyrophosphates. The prenyl-
transferase products are widely distributed in nature and
serve a variety of important biological functions. The cata-
lytic mechanism deduced from the 3D structure and other
biochemical studies of these prenyltransferases as well as
how the protein functions are related to their reaction
mechanism and structure are discussed. In the IPPS reaction,
we focus on the mechanism that controls product chain
length and the reaction kinetics of IPP condensation in the
cis-type and trans-type enzymes. For protein prenyltrans-
ferases, the structures of Ras farnesyltransferase and Rab
geranylgeranyltransferase are used to elucidate the reaction
mechanism of this group of enzymes. For the enzymes
involved in cyclic terpene biosynthesis, the structures and
mechanisms of squalene cyclase, 5-epi-aristolochene syn-
thase, pentalenene synthase, and trichodiene synthase are
summarized.
Keywords: chain elongation; isoprenoid; lipid carrier;

20
and C
25
all-trans-polyprenyl pyro-
phosphates to make C
20
–C
20
and C
20
-C
25
ether-linked lipids
in archeon [9–11]. The C
40
product of octaprenyl pyro-
phosphate synthase (OPPS) constitutes the side chain of
ubiquinone in Escherichia coli [12–14]. Several cis-isoprenyl
Correspondence to P H. Liang, Institute of Biological Chemistry, Academia Sinica, Taipei 11529, Taiwan.
Fax: +886 2 2788 9759, Tel.: +886 2 2785 5696 ext. 6070, E-mail: or A.H J. Wang, Fax: +886 2788 2043,
E-mail:
Abbreviations: FPP, farnesyl pyrophosphate; GPP, geranyl pyrophosphate; GGPP, geranylgeranyl pyrophosphate; IPP, isopentenyl pyrophos-
phate; UPP, undecaprenyl pyrophosphate; IPPS, isoprenyl pyrophosphate synthase; UPPS, undecaprenyl pyrophosphate synthase; DDPPS,
dehydrodolichyl pyrophosphate synthase; PPPS, polyprenyl pyrophosphate synthase; GPPS, geranyl pyrophosphate synthase; FPPS, farnesyl
pyrophosphate synthase; GGPPS, geranylgeranyl pyrophosphate synthase; FGPPS, farnesylgeranyl pyrophosphate synthase; HexPPS, hexa-
prenyl pyrophosphate synthase; HepPPS, heptaprenyl pyrophosphate synthase; OPPS, octaprenyl pyrophosphate synthase; SPPS, solanesyl
pyrophosphate synthase; DPPS, decaprenyl pyrophosphate synthase; FTase, farnesyltransferases; GGTase, geranylgeranyltransferase.
Enzymes: UPPS from Escherichia coli (EC 2.5.1.31); DDPPS from yeast Saccharomyces cerevisiae (EC 2.5.1.31); PPPS from Arabidopsis thaliana
(EC 2.5.1.31); FPPS from E. coli (EC 2.5.1.10); GGPPS from yeast (EC 2.5.1.29); FGPPS from Aeropyrum pernix (EC 2.5.1.33); HexPPS from
Bacillus stearothermophilus and yeast (EC 2.5.1.30); HepPPS from Mycobacterium tuberculosis (EC 2.5.1.30); OPPS from E. coli (EC 2.5.1.11);

100
dolichols for
glycoprotein biosynthesis, a pathway similar to that of the
bacterial peptidoglycan synthesis [25,26]. An even longer
C
120
polymer was found as the final product of an isoprenyl
pyrophosphate synthase in plant Arabidopsis thaliana with
unknown function [27]. As the cis-prenyltransferases com-
monly synthesize long-chain products, a unique short-chain
cis,trans-FPP is made by Mycobacterium tuberculosis FPPS
which utilizes C
10
GPP and IPP to produce a FPP with a cis
double bond [28] (Fig. 1). This bacterium has a decaprenyl
pyrophosphate synthase (DPPS) to produce C
50
decaprenyl
pyrophosphate as a lipid carrier, which is one IPP unit
shorter than UPP found in other bacteria.
The products of these prenyltransferases have specific
chain lengths essential for their biological functions. An
intriguing question is how do they achieve product chain-
length specificity. In theory, restriction of the size of the
enzyme active site should play a major role in determining
the chain length of the final product. With the increasing
numbers of 3D structures available for prenyltransferases,
the mechanism of product chain-length determination has
begun to be elucidated [29]. The cis-type UPPS crystal
structures from Micrococcus luteus and E. coli have been

of different types of natural isoprenoids. A number of
enzymes catalyze cyclization of FPP to generate natural
products such as pentalenene (pentalenene synthase), 5-epi-
aristolochene (5-epi-aristolochene synthase) and trichodiene
(trichodiene synthase). Squalene synthase catalyzes the
cyclization of squalene which is synthesized by the coupling
of two FPP molecules. The crystal structures of these
enzymes have been solved and provide insights into the
catalytic mechanism of terpenoid cyclization [41–44].
This review summarizes these recent advances in the
structural and mechanistic studies of the above three
families of prenyltransferases with emphasis on IPPS, which
has essential biological functions (Table 1). A general
mechanism of product chain-length determination and the
reaction kinetics derived from a pre-steady-state kinetic
analysis for trans-IPPS and cis-IPPS are described.
CLASS I: ISOPRENYL PYROPHOSPHATE
SYNTHASES (IPPSs)
Structure and active site of
trans
-IPPS
Over the past decade, many trans-IPPSs have been purified
and their genes cloned [45,46]. The deduced amino-acid
sequences of these enzymes show amino-acid sequence
homology and two common DDxxD motifs [47], suggesting
that they evolved from the same origin (Fig. 2) [48,49].
These Asp-rich motifs were recognized from the 3D
structure [50] and site-directed mutagenesis studies [51–56]
to be involved in substrate binding and catalysis via
chelation with Mg

smaller k
cat
)[53].A10
4
)10
5
lower activity was observed
for the FPPS from Bacillus stearothermophilus when the first
Table 1.
7
The biological functions and three-dimensional structures of prenyltransferases presented in this review.
Prenyltransferases Biological functions 3D structure [ref]
trans-FPPS Precursor of steroids, cholesterol, sesquiterpenes, farnesylated proteins,
heme, and vitamin K12
[50,57]
trans-GGPPS Precursor of carotenoids, retinoids, diterpenes, geranylgeranylated
chlorophylls, and archaeal ether linked lipids
trans-GFPPS Archaeal ether linked lipids
trans-HexPPS Ubiquinone side chains
–DPPS
cis-UPPS Lipid carrier for peptidoglycan synthesis [30,31]
cis-DDPPS Lipid carrier for glycoprotein synthesis
Ras FTase Farnesylated Ras for signal transduction [37–39]
Rab GGTase Geranylgeranylated Rab for signal transduction [40]
Squalene cyclase Precursor of cholesterol [41]
epi-Aristolochene synthase Precursor of antifungal phytoalexin capsidol [42]
Pentalenene synthase Precursor of pentalenolactone antibiotics [43]
Trichodiene synthase Precursor of antibiotics and mycotoxins [44]
Fig. 2. Sequence alignment of trans-prenyl-
transferases. The sequence-related proteins

DDxxD motif of rat FPPS, substituting the first Asp with
Glu decreased k
cat
90-fold and increased IPP K
m
26-fold,
whereas GPP K
m
remained unchanged [51]. On the other
hand, mutation of the third Asp resulted in no change in the
kinetic parameters.
These results indicate that all the Asp residues in the two
DDxxD motifs except the last one in the second motif are
important for catalysis. In addition, the second motif is
essential for IPP binding. These results are consistent with
the cocrystal structure of avian FPP in complex with GPP
and IPP [57]. The structure of FPPS clearly shows that the
first DDxxD is bound to the allylic substrate GPP and the
second motif is the binding site of the homoallylic substrate
IPP. Site-directed mutagenesis of other amino acids around
the two DDxxD motifs was also performed to show their
effects in substrate binding and catalysis [54,56].
Amino-acid residues essential to product chain-length
determination of
trans
-prenyltransferases
In parallel with the site-directed mutagenesis studies, a
random chemical mutagenesis approach was used to select
FPPS mutants induced by NaNO
2

˚
from
the first Asp-rich motif, which is similar to the length of the
hydrocarbon moiety of FPP [59]. The chain length of the
product catalyzed by these mutants is inversely proportional
to the accessible surface volume of the substituted amino-
acid residue in the first DDxxD. Also in archaebacterial
GGPPS, mutation of Phe77, which is upstream from the
first DDxxD, led to a change in product [60]. For instance,
replacement of this large residue with the smaller Ser
resulted in the production of C
25
rather than C
20
.
There was a similar finding in avian FPPS, in which
replacement of aromatic Phe112 and Phe113 with smaller
amino acids resulted in the product specificity shifting from
C
15
(FPP) to C
20
(GGPP)
1
(F112A), C
25
geranylfarnesyl
pyrophosphate (F113S) and longer products (F112A/
F113S double mutant) [57]. These two residues are located
in the fifth and sixth position before the first DDxxD

from Methanobacterium thermoautotrophicum contains
phenylalanine and serine at the positions corresponding
to Phe112 and Phe113 in avian FPPS. From the X-ray
crystal structure and the sequence alignment of a variety of
polyprenyl pyrophosphate synthases (PPPSs), which gen-
erate C
15
,C
20
and larger products, the importance of
Phe112 and Phe113 in the mechanism of product chain-
length determination is evident. In conclusion, the large
amino-acid residue located before the first DDxxD motif
provides the ÔfloorÕ to block further elongation of the
product (Fig. 3B). Once this residue is replaced with a
smaller one, elongation can continue.
In addition to the 5th and 6th amino-acid residues, the
roles of the 8th and/or the 11th positions before the first
DDxxD in the control of product specificity of archaeal
GGPPS and FPPS were also examined [61]. The single
mutant (F77S, 5th amino acid residue before the first Asp-
rich motif), double mutant (L74G/F77G) and triple mutant
(I71G/74G/F77G) of GGPPS mainly produce C
25
,C
35
and
C
40
, respectively [61]. FPPS mutants display a similar

encoding gene cloned from M. luteus is the first cis-
prenyltransferase gene identified, and the deduced amino-
acid sequence shows no sequence similarity to those of
trans-prenyltransferases [64]. The sequence comparison
with UPPS allows the identification of many cis-type
IPPSs in bacteria, plant and animals as a family [65].
Several regions of conserved sequences can be identified
(Fig. 4). Broadly grouped, they are (with the amino-acid
sequence shown in parentheses): region I (20–32), region II
(42–46), region III (66–88), region IV (142–154) and region
V (190–224). Most of the fully conserved amino acids are
involved in catalysis, substrate binding, or structural
interactions as revealed later by the crystal structures and
mutagenesis studies.
Unlike the trans-type enzymes, cis-prenyltransferases
lack the DDxxD motifs, although they require Mg
2+
for
activity. Earlier site-directed mutagenesis studies examined
the conserved Asp and Glu of E. coli UPPS and revealed
the importance of Asp26, Asp150 and Glu213 in substrate
binding and catalysis [66]. Replacement of Asp26 with Ala
results in 10
3
-fold smaller k
cat
without any significant
change in FPP and IPP K
m
values. Mutagenesis of Asp150

tunnel surrounded by two a helices (a2anda3) and four
b strands (bB, bA, bD, and bC)asshowninFig.5A.In
contrast with the symmetric structure of M. luteus UPPS,
two protein conformations (open and closed forms) are
seen in the two subunits of E. coli UPPS, implicating a
closed/open conformational change mechanism in sub-
strate binding and product release [31]. The difference
between the two conformers is mainly in the position of
the a3 helix. On the basis of site-directed mutagenesis
studies [31], the flexible loop with amino acids 72–83
connected to the a3 helix has been suggested to serve as a
hinge for the interconversion of two conformers. This
study also suggested a role for a Trp residue in the loop for
FPP binding and catalysis [69]. Fluorescent stopped-flow
technology and steady-state spectrophotometer have
recently been used to directly observe a Trp fluorescence
intensity change due to the change in protein conformation
during catalysis [70]. When Trp91, which is located in the
a3 helix, is mutated to Phe, the fluorescence quenching
upon addition of FPP is abolished, suggesting that the a3
helix moves toward the active site during substrate binding
[70]. Thus the change in UPPS conformation to a closed
form results in better interaction between the enzyme and
the substrates and intermediates. After the reaction, the
UPPS structure shifts to an open form for product release,
triggered by crowding of the prenyl chain of the product
because the large amino-acid residues seal the bottom of
the tunnel-shaped active site.
Ó FEBS 2002 Structure, mechanism and function of prenyltransferases (Eur. J. Biochem. 269) 3343
Mechanism of product chain-length determination

The single UPPS mutation, L137A, has converted UPPS
into DDPPS in terms of product specificity. V105 may also
play a role in blocking chain elongation, as its mutation
increases the proportion of C
60
,C
65
and C
70
in the absence
of Triton, but it is not as critical as L137.
All cis-prenyltransferases so far identified have products
of chain length at least C
55
, except a short-chain FPP-
synthesizing enzyme recently identified from M. tuberculo-
sis. When the amino-acid sequence of UPPS is compared
with that of the short-chain FPPS, the A69 and A143 in
E. coli UPPS correspond to the large L84 and V156 in
M. tuberculosis FPPS, respectively. Substitution of Leu for
A69 in E. coli UPPS indeed leads to production of a greater
amount of C
30
intermediate which is longer lived, suggesting
that this residue interferes with the chain elongation of the
C
30
product. From the structure, it is reasonable to assume
that A69 is located midway in FPP elongation to the C
55

Fig. 4. Alignment of the cis-type IPPSs. These
include, in turn, E. coli UPPS, yeast DDPPS
Rer2, Yeast DDPPS Srt1, M. tuberculosis FPs
Rv1086, M. tuberculosis DPPS Rv2361c,
Arabidopsis thaliana PPPS and human
DDPPS. The numbers and secondary-struc-
tural elements shown above the sequences
arefortheUPPSfromE. coli,basedon
PROCHECK
analysis of the crystal structure.
The green arrows denote the locations of
b-strands, and the cylinders in red, magenta
andcyanarefora helices, 3
10
helices and
turns, respectively. In the aligned sequences,
several conserved regions including (I) resi-
dues 20–32, (II) 42–46, (III) 66–88, (IV)
142–154, and (V) 190–224 can be identified,
which probably also have corresponding
secondary structures in common.
3344 P H. Liang et al.(Eur. J. Biochem. 269) Ó FEBS 2002
A tertiary carbocation on the C3 carbon of the IPP part is
proposed to form during the 1¢-4 condensation. The aza
analogues with nitrogen replacing the cationic carbon to
mimic the transition state have previously been demonstra-
ted to strongly inhibit enzymes in isoprenoid biosynthesis
where the carbocation transition state is presumably formed
during catalysis [73]. An aza analogue of the transition state,
3-azageranylgeranyl diphosphate, is also a potent inhibitor

The reaction mechanism of the cis-prenyltransferases is
less well understood. The hydrolysis of the allylic substrate
by cis-prenyltransferases has not been observed. Analog-
ously to the two DDxxD motifs in the trans-prenyltrans-
ferases, site-directed mutagenesis studies of UPPS indicate
that Asp and Glu play a significant role in IPP binding and
Fig. 5. (A) Two orthogonal views of the ribbon representation of an E. coli UPPS dimer and (B) the proposed active site of the E. coli UPPS located in
a tunnel-like crevice surrounded by a2, a3, bD, bB, bA, and bC. (A) The top view is perpendicular to, and the bottom view is parallel to, the molecular
dyad axis. The seven a helices and six b strands are shown in red and green, respectively, for subunit A, and in magenta and cyan for subunit B, and
they are labelled separately. The blue arrows indicate locations of the active sites, each having a substrate-binding tunnel formed by helices a2, a3
and the central b-sheet. (B) The substrate site is located on the top of the hydrophobic tunnel with D26 and D213 playing a significant role in
substrate binding and catalysis. A69 is in the midle of FPP chain elongation to the product. The large amino acid L137 on the bottom of the tunnel
is essential for determination of product chain length by blocking further elongation of UPP.
Ó FEBS 2002 Structure, mechanism and function of prenyltransferases (Eur. J. Biochem. 269) 3345
activity probably through co-ordination with Mg
2+
.The
IPP condensation kinetics for UPPS, DDPPS (cis-type) and
OPPS (trans-type)hadbeenmeasuredforcomparisonof
IPP condensation catalyzed by cis-IPPS and trans-IPPS
[76–78]. These experiments were performed under the
enzyme single-turnover condition using a rapid-quench
instrument, because the slow release of the large hydropho-
bic product limits IPPS catalysis under steady-state condi-
tions. The IPP condensation rate constant in the UPPS
reaction is similar to that of OPPS (2.5 s
)1
vs. 2.0 s
)1
)

M
IPP, whereas the C
55
-UPP is synthesized with
5 l
M
FPP and 50 l
M
IPP. In contrast, OPPS produces no
intermediate products under these conditions [77]. Another
trans-prenyltransferase, SPPS from M. luteus, shows a
similar pattern of product distribution in which no inter-
mediate is displaced from the active site at high concentra-
tion of FPP [15]. As OPPS and UPPS have similar FPP and
IPP K
m
values, OPPS and SPPS (trans-type) apparently
have higher affinities for intermediates compared with
UPPS (cis-type). At a fixed concentration of allylic sub-
strate, the increased ratio of IPP to FPP results in the
synthesis of longer-chain products. As shown for a UPPS
from the Archaeon Sulfolobus acidocaldarius, the enzyme
mainly generates C
50
and C
55
when IPP is present in excess
of GGPP (or FPP), but decreasing the concentration of IPP
results in larger amounts of short-chain products [79]. This
could be due to the low affinity of IPP for the enzyme.

to C
55
)intheUPPS
reaction (Fig. 6). On the other hand, the rate for elongation
from C
40
to C
45
catalyzed by OPPS is 100 times slower than
for elongation from C
35
to C
40
(Fig. 6). The trans enzyme
seemstohaveamorerigidactivesitethanthecis-
prenyltransferase, as shown by the higher product specificity.
The most surprising observation on the product distri-
bution of the UPPS and OPPS reactions is that, when the
Fig. 6. Comparison of the kinetic pathways of UPPS and OPPS. OPPS
catalyzes elongation of C
15
to C
40
, resulting in all trans double bonds,
and UPPS catalyzes formation of the C
55
product containing newly
formed cis double bonds. The IPP condensation steps have rate con-
stants of 2 s
)1

Ras FTase is a Zn
2+
-dependent prenyltransferase contain-
ing a and b heterodimer, which catalyzes the farnesylation
on a C-terminal CaaX motif of the Ras protein. As shown
in Fig. 9B, the Zn
2+
-activated thiolate of Cys acts as a
nucelophile to attack the ionized farnesyl group. In the 3D
structure of a mammalian Ras FTase, both subunits are
largely composed of a helices (Fig. 7A) [37]. The a-2 to a-15
helices in the a subunit fold into a novel helical hairpin
structure, resulting in a crescent-shape domain that enve-
lopes part of the b subunit. On the other hand, the 12 helices
of the b subunit form an a–a barrel. Six additional helices
connect the inner core of helices and form the outside of the
helical barrel. A deep cleft surrounded by hydrophobic
amino acids in the center of the barrel is proposed as
the FPP-binding pocket. A single Zn
2+
ion is located at the
junction between the a-hydrophilic surface groove near the
subunit interface and the deep cleft in the b subunit. This
Zn
2+
ion is pentaco-ordinated by the Asp297 and Cys299
located in the N-terminal helix 11, His362 in helix 13 of the
b subunit, and a water molecule as well as a bidentate
2
ligand

binding of FPP. The diphosphate moiety of the FPP
substrate is hydrogen-bonded with His248b,Arg291b,and
Tyr300b. Lys164a and Lys294b are also within hydrogen-
bonding distance of the diphosphate. Mutations of His248b,
Arg291b, Lys294b, Tyr300b, and Trp303b cause 3–15 times
increased FPP K
d
compared with the wild-type FTase [83],
consistent with the crystal structure. On the other hand,
replacement of Lys164a with Asn results in a markedly
decreased k
cat
value, suggesting that this residue plays a
catalyticrole[84].Fromthebinarystructure,Lys164may
interact with the diphosphate moiety of FPP and be
involved in the transfer of Cys thiol to C1 of the substrate.
As for the binding of the 5th CaaX peptide substrate,
the structure of FTase complexed with a-hydroxyfarnesyl-
Fig. 7. (A) Ribbon diagram of Ras FTase and (B) the molecular ruler mechanism for substrate specificity of FTase and GGTase. (A) The heterodimeric
enzyme consists of two subunits, a and b, colored in cyan–blue and yellow–green, respectively. Most of the secondary structures are helices, with the
a subunit comprising seven helical hairpins that surround the more compact b subunit. The peptide substrate, colored red, binds to a cleft between
the two subunits, and so does the substrate analogue, colored magenta. The active site is located in the b subunit. It contains a zinc ion, shown in
blue, which is bound by three residues Asp297, Cys299 and His362, shown in cyan, and also makes bonds with the substrate molecules. (B) When
FPP is replaced with GGPP in the active site of FTase, the thiolate nucleophile is further away from the electrophilic carbon next to the
pyrophosphate leaving group, thereby decreasing the enzyme activity. Similarly, FPP is a poor substrate for GGTase.
Ó FEBS 2002 Structure, mechanism and function of prenyltransferases (Eur. J. Biochem. 269) 3347
phosphonic acid and acetyl-Cys-Val-Ile-selenoMet-COOH
reveals that it binds in a cleft located in the subunit interface,
in agreement with the pervious apoenzyme structure [37].
The peptide is in contact with FPP and several amino acids

of the 3D structure of Rab GGTase [40]. The GGPP is
bound in the central cavity of the a–a barrel in the
b subunit with its diphosphate head group to the positively
charged cluster composed of Arg232b, Lys235b,and
L105a. The diphosphate is closed to the Zn
2+
,whichis
co-ordinated with Asp238b, Cys240 b and His290b in a
similar way to that observed in FTase, and an additional
His residue. The structure of Rab GGTase can be
superimposed on that of Ras FTase. One of the most
striking differences is that on the bottom of the GGTase
active-site cavity, Ser48b and Leu99b replace the more
bulky Trp102b and Tyr154b seen in FTase, thereby
enlarging the active site to accommodate GGPP. This is
consistent the molecular ruler mechanism (Fig. 7B) pro-
posed in FTase for substrate specificity.
Rab GGTase is unique and different from FTase and
type I GGTase in that it is able to prenylate Rab only in the
presence of Rab escort protein. It exclusively modifies
members of a single subfamily of Ras-related small GTPase,
the Rab proteins involved in the regulation of intracellular
vesicular transport in the biosynthetic secretory and exocy-
tic/endocytic pathways [86,87]. Upon binding with the
protein complex, the Rab GGTase transfers two GGPPs to
the two Cys residues of Rab C-terminal -CC, -CXC, -CCX,
or -CCXX motif [88]. As shown in the superimposition of
the GGTase and FTase structures, several residues in the
peptide-binding pockets are different, reflecting their pep-
tide substrate specificity.

[94]. The use of the more thiophilic Cd
2+
to increase the
thio affinity and lowering its pK
a
to display lower activity
suggests the direct participation of a metal ion in FTase
catalysis [95]. However, inclusion of high concentrations of
Mg
2+
in the reaction mixture increases enzyme activity
700-fold because excess Mg
2+
occupies a separate site,
facilitating departure of the diphosphate group [96].
Therefore, an associated character with partially positive
charge at C1 of FPP and partially negative charge
pyrophosphate oxygen as well as in the metal-co-ordinated
thiolate was proposed as the transition state of the FTase
reaction (see Fig. 9B).
CLASS III: TERPENOID CYCLASES
General structure and mechanism of terpenoid cyclase
Terpenoid cyclases such as squalene cyclase, pentalenene
synthase, 5-epi-aristolochene synthase, and trichodiene
synthase are responsible for the synthesis of cholesterol, a
hydrocarbon precursor of the pentalenolactone family of
antibiotics, a precursor of the antifungal phytoalexin
capsidiol, and the precursor of antibiotics and mycotoxins,
respectively. The last three enzymes catalyze the cyclization
of FPP involving: (a) ionization of FPP to an allylic cation

protruded into the active site to complex with Mg
2+
.
Together they facilitate the FPP carbocation formation as
shown in Fig. 9C (path a). Subsequent cyclization and
carbocation rearrangement processes use a His309 catalytic
residue in the active site [43].
5-epi-Aristolochene synthase
Tobacco 5-epi-aristolochene synthase adopts entirely an
a-helical structure with short loops and turns (Fig. 8B). The
structure folds into two domains: the N-terminal domain
aligns structurally with two glycosyl hydroxylases, and its
function is not known; the C-terminal domain shares the
common fold with FPPS. The active site of epi-aristolochene
synthase is located in the C-terminal domains containing two
Mg
2+
-binding sites. The Asp301 and Asp305 found in the
conserved DDxxD motif co-ordinate with a Mg
2+
[42]. The
Asp444, Thr448, Glu452 and a water molecule co-ordinate
Fig. 8. Ribbon representation of the structures of cyclic terpenoid-synthetic enzymes pentalenene synthase (A), 5-epi-aristolochene synthase (B),
trichodiene synthase (C) and squalene synthase (D). They are shown in different colors with the N-terminus and C-terminus labeled. All of these
enzymes have the terpenoid synthase fold and contain a helices as their major secondary-structure elements. The homologous helices that constitute
the active site are emphasized with broad ribbons.
Ó FEBS 2002 Structure, mechanism and function of prenyltransferases (Eur. J. Biochem. 269) 3349
with the other Mg
2+
.ThesetwoMg

bicyclic eudesmane carbocation intermediate. The following
rearrangement of the carbocation is facilitated by the
electron-rich side chain of Trp273 to generate the final
product aristolochene.
Trichodiene synthase
The recently solved 3D structure of trichodiene synthase
(Fig. 8C) reveals 17 a helices, six of which (C, D, G, H, I,
and J) form the hydrophobic active-site cleft [44]. An Asp-
rich D
100
DSKD motif, which is important for catalytic
activity as suggested by the mutagenesis studies [98], is
located in the active site at the C-terminus of helix D.
Three Mg
2+
ions were found in the electron map of the
enzyme–pyrophosphate complex. Among them, two metal
ions in the active-site region co-ordinate with Asp100 (the
first residue of the conserved Asp-rich motif). The
conserved Asp residues are also found in aristolochene
synthase and FPPS. The other Mg
2+
is located in the site
of N225, S229, and E233 on helix H, which are in a
consensus sequence conserved among all known terpene
synthase sequences. The metal ions are bound to the
enzyme in the presence of pyrophosphate, and the
pyrophosphate (also probably FPP) triggers a protein
conformational change, in which D101 breaks a salt link
with R62 and forms a new salt bridge with R304 after

2+
. The deprotonated IPP or a Zn
2+
-activated thiolate
anion of a protein acts as a nucleophile to attack the carbocation at C1
resulting in condensation with departure of the pyrophosphate. In the
cyclase reaction, the nearby C¼C bond donates electrons for ring
closure. The paths a, b, and c denote the routes of electron migration
from the C¼C bond to C1 with the release of pyrophosphate catalyzed
by pentalenene synthase, epi-aristolochene synthase, and trichodiene
synthase, respectively.
3350 P H. Liang et al.(Eur. J. Biochem. 269) Ó FEBS 2002
(Fig. 8D) [41]. Domain 1 is an a6–a6 barrel of two
concentric rings of parallel a helices, which is similar to the
fold of FPPSs and glucanases [99]. Domain 2 forms an a–a
barrel and inserts into domain 1. The active site is located
in a central cavity, with two amino acids in a DxDD motif,
Asp376 and Asp377, on the top of the cavity [41].
Mutagenesis of these two Asp residues abolishes enzyme
activity, indicating their importance in catalysis [100].
Based on the structure, Asp376 is the catalytic acid to
protonate the first C¼C double bond, resulting in a
carbocation for the initiation of ring cyclization. A water
molecule acts as a base to receive a proton on the other
end of the squalene molecule, which is polarized by the
other water in a hydrogen-bonding network formed by
Glu45 and Gln262. A constriction formed by Phe166,
Val174, Phe434, and Cys435 in a mobile loop may serve as
a gate for substrate passage.
CONCLUSION

available, the general mechanism for product chain-length
determination of the cis and trans forms of IPPS has
gradually been understood. As revealed by the 3D
structures, the size of the active-site cavity apparently
controls the chain length of the final product of the IPPS
reaction. Several large amino acids form a floor to seal the
bottom of the active site and block further elongation of
the product. Site-directed substitution of key large amino
acids with smaller ones extends the size of the active site,
thereby allowing the formation of a longer product and
vice versa. The substrate specificities of protein FTase and
GGTase are controlled by the size of the substrates. The
larger GGPP is more distant from the Zn
2+
catalytic site
when binding on FTase and hence produces lower activity.
The accurately tailored active sites of FPP cyclases
(pentalenene synthase, 5-epi-aristolochene synthase, and
trichodiene synthase) guarantee formation of the correct
cyclization products.
A large number of isoprenoids have significant biological
functions in nature. The structural and mechanistic infor-
mation available so far provides an initial step to under-
standing the functions of many other prenyltransferases and
a basis for designing enzyme agonists and antagonists to
treat various diseases. However, the structures and even
functions of many isoprenoid-synthetic enzymes, as well as
the proteins catalyzing and undergoing prenylation, are still
not known. Further studies on the structure, mechanism
and function of prenyltransferases are therefore required.

isoprenoid biosynthetic pathway. Curr. Opin. Chem. Biol. 1, 570–
578.
8. Ogura, K. & Koyama, T. (1998) Enzymatic aspects of isoprenoid
chain elongation. Chem. Rev. 98, 1263–1276.
9. Ohnuma, S I., Suzuki, M. & Nishino, T. (1994) Archaebacterial
ether-linked lipid biosynthetic gene. Expression cloning,
sequencing and characterization of geranylgeranyl-diphosphate
synthase. J. Biol. Chem. 269, 14792–14797.
10. Chen, A. & Poulter, C.D. (1994) Isolation and characterization
of idsA: the gene for the short chain isoprenyl diphosphate syn-
thase from Methanolbacterium thermoautotrophicum. Arch. Bio-
chem. Biophys. 314, 399–404.
11. Tachibana, A., Yano, Y.Y., Otani, S., Nomura, N., Sako, Y. &
Taniguchi, M. (2000) Novel prenyltransferase gene encoding
farnesylgeranyl diphosphate synthase from a hyperthermophilic
archeon, Aeropyrum pernix. Eur. J. Biochem. 267, 321–328.
12. Asai, K I., Fujisaki, S., Nishimura, Y., Nishino, T., Okada, K.,
Nakagawa, T., Kawamukai, M. & Matsuda, H. (1994) The
identification of Escherichia coli ispB (cel ) gene encoding the
octaprenyl diphosphate synthase. Biochem. Biophys. Res. Com-
mun. 202, 340–345.
13. Okada, K., Suzuki, K., Kamiya, Y., Zhu, X., Fujisaki, S.,
Nishimura, Y., Nishino, T., Nakagawa, T., Kawamukai, M. &
Matsuda, H. (1996) Polyprenyl diphosphate synthase essentially
defines the length of the side chain of ubiquinone. Biochim.
Biophys. Acta 1302, 217–223.
14. Okada, K., Minehira, M., Zhu, X., Suzuki, K., Nakagawa, T.,
Matsuda, H. & Kawamukai, M. (1997) The ispB gene encoding
octaprenyl diphosphate synthase is essential for growth of
Escherichia coli. J. Bacteriol. 179, 3058–3060.

22. Collin, M.D. & Jones, D. (1981) Distribution of isoprenoid
quinone structural types in bacteria and their taxonomic
implication. Microbiol. Rev. 45, 316–354.
23. Robyt, J. (1998) In Essentials of Carbohydrate Chemistry Chapter
10, pp. 305–318. Springer-Verlag, New York.
24. Sutherland, I.W. (1977) In Surface Carbohydrates of the Proka-
rotic Cell,p.27.AcademicPress,London.
25. Nishikawa, S. & Nakano, A. (1993) Identification of a gene
required for membrane protein retention in the early secretory
pathway. Proc. Natl. Acad. Sci. USA 90, 8179–8183.
26. Sato, M., Sato, K., Nishikawa, S I., Hirata, A., Kato, J I. &
Nakano, A. (1999) The yeast RER2 gene, identified by
endoplasmic reticulum protein localization mutations, encodes
cis-prenyltransferase, a key enzyme in dolichol synthesis. Mol.
Cell. Biol. 19, 471–483.
27. Oh, S.K., Han, K.H., Ryu, S.B. & Kang, H. (2000) Molecular
cloning, expression, and functional analysis of a cis-pre-
nyltransferase from Arabidopsis thaliana. J. Biol. Chem. 275,
18482–18488.
28. Schulbach, M.C., Brennan, P.J. & Crick, D.C. (2000) Identifi-
cation of a short (C
15
) chain Z-isoprenyl diphosphate synthase
and a homologous long (C
50
) chain isoprenyl diphosphate syn-
thase in Mycobacterium tuberculosis. J. Biol. Chem. 275, 22876–
22881.
29. Wang, K.C. & Ohnuma, S I. (2000) Isoprenyl diphosphate
synthases. Biochim. Biophys. Acta 1529, 33–48.

38. Long, S.B., Casey, P.J. & Beese, L.S. (1998) Cocrystal structure
of protein farnesyltransferase complexed with a farnesyl dipho-
sphate substrate. Biochemistry 37, 9612–9618.
39. Strickland, C.L., Windsor, W.T., Syto, R., Wang, L., Bond, R.,
Wu, Z., Schwart, J., Le, H.V., Beese, L.S. & Weber, P.C. (1998)
Crystal structure of farnesyl protein transferase complexed with a
CaaX peptide and farnesyl diphosphate analogue. Biochemistry
37, 16601–16611.
40. Zhang, H., Seabra, M. & Deisenhofer, J. (2000) Crystal structure
of Rab geranylgeranyltransferase at 2.0 A
˚
resolution. Structure 8,
241–251.
41. Wendt, K.U., Poralla, K. & Schulz, G.E. (1997) Structure and
function of a squalene cyclase. Science 275, 1811–1815.
42. Starks, C.M., Back, K., Chappel, J. & Noel, J.P. (1997) Struc-
tural basis for cyclic terpene biosynthesis by tobacco 5-epi-aris-
tolochene synthase. Science 275, 1815–1820.
43. Lesburg, C.A., Zhai, G., Cane, D.E. & Christianson, D.W.
(1997) Crystal structure of pentalenene synthase: mechanistic
insights on terpenoid cyclization reactions in biology. Science
275, 1820–18224.
44. Rynkiewicz, M.J., Cane, D.E. & Christianson, D.W. (2001)
Structure of trichodiene synthase from Fusarium sporotrichioides
provides mechanistic inferences on the terpene cyclization cas-
cade. Proc. Natl. Acad. Sci. USA 98, 13543–13548.
45. Ogura, K., Toyama, T. & Sagami, H. (1997) Polyprenyl
diphopshate synthases. In Subcellular Biochemistry (Bittman, R.,
ed.), Vol. 28, pp. 57–88. Plenum, New York.
46. Wang, K. & Ohnuma, S. (1999) Enzymatic aspects of isoprenoid

served prenyltransferase domain I and II. Proc. Natl. Acad. Sci.
U.S.A. 91, 3044–3048.
54. Koyama, T., Tajima, M., Nishino, T. & Ogura, K. (1995) Sig-
nificance of Phe220 and Gln-221 in the catalytic mechanism of
farnesyl diphosphate synthase of Bacillus stearothermophilus.
Biochem. Biophys. Res. Commun. 212, 681–686.
55. Koyama, T., Tajima, M., Sano, H., Doi, T., Koike-Takeshita,
A., Obata, S., Nishino, T. & Ogura, K. (1996) Identification of
significant residues in the substrate binding site of Bacillus
stearothermophilus farnesyl diphosphate synthase. Biochemistry
35, 9533–9538.
56. Koyama, T., Obata, S., Saito, K., Takeshita-Koike, A. & Ogura,
K. (1994) Structural and functional roles of the cysteine residues
of Bacillus stearothermophilus farnesyl pyrophosphate synthase.
Biochemistry 33, 12644–12648.
57. Tarshis, L.C., Proteau, P.J., Kellogg, B.A., Sacchettini, J.C. &
Poulter, C.D. (1996) Regulation of product chain length by iso-
prenyl diphosphate synthase. Proc. Natl. Acad. Sci. USA 93,
15018–15023.
58. Ohnuma, S I., Narita, K., Nakazawa, T., Hemmi, H., Hallberg,
A M., Koyama, T., Ogura, K. & Nishito, T. (1996) Conversion
from farnesyl diphosphate synthase to geranylgeranyl diphos-
phate synthase by random chemical mutagenesis. J. Biol. Chem.
271, 10087–10095.
59. Ohnuma, S I., Narita, K., Nakazawa, T., Ishida, C., Takeuchi,
Y., Ohto, C. & Nishito, T. (1996) A role of the amino acid residue
located on the fifth position before the first Aspartate-rich motif
of farnesyl diphosphate synthase on determination of the final
product. J. Biol. Chem. 271, 30748–30754.
60. Ohnuma, S I., Hirooka, K., Hemmi, H., Ishida, C., Ohto, C. &

diphosphate synthase. J. Biol. Chem. 276, 28459–28464.
69. Fujikura, K., Zhang, Y W., Yoshizki, T. & Koyama, T. (2000)
Significance of Asn-77 and Trp-78 in the catalytic function of
undecaprenyl diphosphate synthase of Micrococcus luteus B-P
26. J. Biochem. (Tokyo) 128, 917–922.
70. Chen, Y.H., Chen, A.P C., Chen, C.T., Wang, A.H J. & Liang,
P.H. (2002) Probing the conformational change of Escherichia
coli undecaprenyl pyrophosphate synthase during catalysis using
an inhibitor and tryptophan mutants. J. Biol. Chem. 277, 7369–
7376.
71. Poulter, C.D. & Rilling, H.C. (1976) Prenyltransferase: the
mechanism of the reaction. Biochemistry 15, 1079–1083.
72. Poulter, C.D. & Rilling, H.C. (1978) The prenyl transfer reaction.
Enzymatic and mechanistic studies of the 1¢-4 coupling reaction
in the terpene biosynthetic pathway. Acc. Chem. Res. 11, 307–
311.
73. Rahier, A., Taton, M. & Benveniste, P. (1990) Inhibition of sterol
biosynthesis enzymes in vitro by analogues of high-energy car-
bocationic intermediates. Biochem. Soc. Trans. 18, 48–52.
74. Steiger, A., Pyun, H J. & Coates, R.M. (1992) Synthesis and
characterization of aza analog inhibitors of squalene and
geranylgeranyl diphosphate synthases. J. Org. Chem. 57, 3444–
3449.
75. Sagami, H., Korenaga, T., Ogura, K., Steiger, A., Pyun, H.J. &
Coates, R.M. (1992) Studies on geranylgeranyl diphosphate
synthase from rat liver: specific inhibition by 3-azageranylgeranyl
diphosphate. Arch. Biochem. Biophys. 297, 314–320.
76. Pan, J.J., Chiou, S.T. & Liang, P.H. (2000) Product distribution
and pre-steady-state kinetic analysis of Escherichia coli
undecaprenyl pyrophosphate synthase reaction. Biochemistry 39,

1383–1390.
85. Yokoyama, K., Zimmerman, K., Scholten, J. & Gelb, M.H.
(1997) Differential prenyl pyrophosphate binding to mammalian
protein geranylgeranyltransferase-I and protein farnesyl-
transferase and its consequence on the specificity of protein
Prenylation. J. Biol. Chem. 272, 3944–3952.
86. Pfeffer, S.R. (1994) Rab GTPases: master regulators of mem-
brane trafficking. Curr. Opin. Cell Biol. 6, 522–526.
Ó FEBS 2002 Structure, mechanism and function of prenyltransferases (Eur. J. Biochem. 269) 3353
87. Novick, P. & Zerial, M. (1997) The diversity of Rab proteins in
vesicle transport. Curr. Opin. Cell Biol. 9, 496–504.
88. Farnsworth, C.C., Seabra, M.C., Ericsson, L.H., Gelb, M.H. &
Glomset, J.A. (1994) Rab geranylgeranyl transferase catalyzes
the geranylgeranylation of adjacent cysteines in the samll
GTPases Rab1A, Rab3A, and Rab5A. Proc. Natl. Acad. Sci.
USA 91, 11963–11967.
89. Furfine, E., Leban, J., Landavazo, A., Moomaw, J. & Casey, P.
(1995) Protein farnesyltransferase: kinetics of farnesyl
pyrophosphate binding and product release. Biochemistry 34,
6857–6862.
90. Dolence, J.M., Cassidy, P.B., Mathis, J.R. & Poulter, C.D. (1995)
Yeast protein farnesyltransferase: steady-state kinetic studies of
substrate binding. Biochemistry 34, 16687–16694.
91. Pompliano, D., Schaber, M., Mosser, S., Omer, C., Shafer, J. &
Gibbs, J. (1993) Isoprenoid diphosphate utilization by
recombinant human farnesyl: protein transferase: interactive
binding between substrates and a preferred kinetic pathway.
Biochemistry 32, 8341–8347.
92. Huang, C.C., Hightower, K.E. & Fierke, C.A. (2000) Mechan-
istic studies of rat protein farnesyltransferase indicate an associ-

hopene cyclase. Eur. J. Biochem. 242, 51–55.
3354 P H. Liang et al.(Eur. J. Biochem. 269) Ó FEBS 2002