Site-directed mutagenesis experiments on the putative deprotonation site of squalene-hopene cyclase from Alicyclobacillus acidocaldarius

Article abstract of DOI:10.1271/bbb.68.728

To provide insight into the catalytic mechanism for the final deprotonation reaction of squalene-hopene cyclase (SHC) from Alicyclobacillus acidocaldarius, mutagenesis experiments were conducted for the following ten residues: Thr41, Glu45, Glu93, Arg127,

Full text of DOI:10.1271/bbb.68.728

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Site-directed Mutagenesis Experiments on the
Putative Deprotonation Site of Squalene-hopene
Cyclase from Alicyclobacillus acidocaldarius
Tsutomu SATOa, Masanori KOUDAa & Tsutomu HOSHINOa
a Department of Applied Biological Chemistry, Faculty of Agriculture, and Graduate
School of Science and Technology, Niigata University
Published online: 22 May 2014.
To cite this article: Tsutomu SATO, Masanori KOUDA & Tsutomu HOSHINO (2004) Site-directed Mutagenesis Experiments
on the Putative Deprotonation Site of Squalene-hopene Cyclase from Alicyclobacillus acidocaldarius , Bioscience,
Biotechnology, and Biochemistry, 68:3, 728-738, DOI: 10.1271/bbb.68.728
To link to this article: http://dx.doi.org/10.1271/bbb.68.728
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Biosci. Biotechnol. Biochem., 68 (3), 728–738, 2004
Site-directed Mutagenesis Experiments on the Putative Deprotonation
Site of Squalene-hopene Cyclase from Alicyclobacillus acidocaldarius
y
Tsutomu SATO, Masanori KOUDA, and Tsutomu HOSHINO
Department of Applied Biological Chemistry, Faculty of Agriculture, and Graduate School of Science and Technology,
Niigata University, Ikarashi, Niigata 950-2181, Japan
Received November 18, 2003; Accepted December 4, 2003
To provide insight into the catalytic mechanism for
the final deprotonation reaction of squalene-hopene
cyclase (SHC) from Alicyclobacillus acidocaldarius,
mutagenesis experiments were conducted for the follow-
ing ten residues: Thr41, Glu45, Glu93, Arg127, Trp133,
Gln262, Pro263, Tyr267, Phe434 and Phe437. An X-ray
analysis of SHC has revealed that two types of water
molecules (‘‘front water’’ and ‘‘back waters’’) were
involved around the deprotonation site. The results of
these mutagenesis experiments allow us to propose the
functions of these residues. The two residues of Gln262
and Pro263 probably work to keep away the isopropyl
group of the hopanyl cation intermediate from the
‘‘front water molecule,’’ that is, to place the ‘‘front
water’’ in a favorable position, leading to the minimal
production of by-products, i.e., hopanol and hop-21(22)-
ene. The five residues of Thr41, Glu45, Glu93, Arg127
and Trp133, by which the hydrogen-bonded network
incorporating the ‘‘back waters’’ is constructed, in-
crease the polarization of the ‘‘front water’’ to facilitate
proton elimination from the isopropyl moiety of the
hopanyl cation, leading to the normal product, hop-
22(29)-ene. The three aromatic residues of Tyr267,
Phe434 and Phe437 are likely to play an important role
in guiding squalene from the enzyme surface to the
reaction cavity (substrate channeling) by the strong
affinity of their aromatic residues to the squalene
substrate.
that of 3 occurs by the addition of a water molecule to
final cation C-22 of 4 (path b), as shown in Scheme 1.
The ratio of product 2 to 3 is 84:16. Numerous triterpene
skeletons are formed by the biocyclization of common
substrate 1 or (3S)-2,3-oxidosqualene. The diversity of
triterpene skeletons found in nature originates from
many types of different termination reactions by
triterpene cyclases.1–5) Tetrahymanol from protozoa is
biosynthesized by water attack on the cation at C-21 to
the final cation having the 6/6/6/6/6-fused pentacyclic
skeleton. As for lanosterol from vertebrates and fungi,
the final protosterol cation (a 6/6/6/5-fused tetracyclic
ring system) undergoes backbone rearrangement by 1,2-
shifts of the hydride and methyl group and a subsequent
deprotonation reaction at C-9, leading to the introduc-
tion of a double bond at C8–C9 (lanostane numbering).
Cycloartenol from plants is formed via the protosterol
cation in a similar manner to that of lanosterol, but the
deprotonation reaction occurs from the methyl group of
C-19 after the backbone rearrangement (lanostane
numbering), resulting in the formation of a cyclopropane
ring.
In the past half a decade, there have been remarkable
advances in understanding the catalytic mechanisms by
A. acidocaldarius SHC.1,2) Site-directed mutants and
substrate analogs1,2) have enabled us to establish that
this polycyclization reaction consisted of eight reaction
steps, including two ring-expansion processes for the
formation of 6-membered C- and D-rings.6,7) The
polycyclization reaction is triggered by proton attack
on the terminal double bond of 1, the acidic proton being
donated by the DXDDTA motif.8) The transient carbo-
cations that are generated during the polycyclization
cascade are stabilized through cation-ꢀ interaction by
such aromatic amino acid residues as F365 and
F605.9,10) The steric bulk size of the amino acid residues
affects the stereochemical destiny during the polycyc-
lization process, which has been demonstrated by
mutation experiments on I261 and L607.11,12) However,
it remains uncertain as to which amino acids are
responsible for the final deprotonation reaction of 4 to
Key words: squalene; hopene; terpene cyclase; triter-
pene
The polycyclization of squalene 1 into pentacyclic
hopene 2 (hop-22(29)-ene) and hopanol 3 is one of the
most complicated biochemical reactions (Scheme 1)1–3)
and is catalyzed by squalene-hopene cyclases (SHCs)
[EC 5.4.99-] from prokaryotic species. The polycycliza-
tion reaction proceeds under precise enzymatic control
to form five rings and nine chiral centers. The formation
of 2 occurs by a proton elimination reaction from the
methyl group at C-29 of hopanyl cation 4 (path a), while
y
To whom correspondence should be addressed. Fax: +81-25-262-6854; E-mail: hoshitsu@agr.niigata-u.ac.jp
Abbreviations: SHC, squalene-hopene cyclase; GC, gas chromatography

Deprotonation Site of Squalene Cyclase
729
Scheme 1. Polycyclization Reaction of Squalene 1 into Hopene 2 and Hopanol 3 Catalyzed by Squalene-hopene Cyclase.
The production ratio of 2:3 is ca. 84:16. The acyclic and flexible squalene molecule is cyclized through a discrete carbocation to form final
cation 4 (the hopanyl cation). Products 2 and 3 are formed by regiospecific deprotonation (path a) from the methyl group of C-29 of 4, derived
from Z-Me at C23 of 1, and hydroxylation (path b) at the C-22 cation of 4, respectively.14) The structure of hop-21(22)-ene 5 is also shown,
which occurs from the erroneous deprotonation reaction of H-21 of intermediate 4 (path c) by the mutated cyclases. Compound 5 is not produced
by the native cyclase, but is formed by the mutated cyclases.
form 2.
responsible for the deprotonation reaction is suggested
to be a water molecule (named ‘‘front water’’) whose
polarization can be enhanced by other types of water
(named ‘‘back waters’’) that construct the hydrogen-
bonding network associated with the seven residues of
T41, E45, E93, R127, W133, Q262 and Y267. The Q262
residue bears a further hydrogen bond with the ‘‘front
water’’. The strongly polarized ‘‘front water’’ can
therefore store the proton generated from Me-29 to
form 2, while 3 is produced if the ‘‘front water’’ adds as
hydoxyl to the C-22 cation of 4, instead of accepting the
proton (Fig. 1).15,16) In addition, the P263 and F437
residues are close to the ‘‘front water’’. To date, the
functions of the three residues of E45, F129 and F605,
which may be responsible for the deprotonation site,
have been evaluated.10,17,18) The ꢀ-electrons of F605
work to stabilize the transient carbocations at C-17 and
C-22 that are formed during the polycyclization process
through cation-ꢀ interaction.10) Residues E4518) and
F12917) play a crucial role in the catalysis. However, no
We have previously established, by using norsqualene
lacking methyl group at C-23, that the terminal
isopropylidene moiety of 1 was indispensable for
initiating the polycyclization reaction and for construct-
ing the five-membered E-ring.13) Incubation of deuter-
ated 1 having an E- or Z-trideuteriomethyl group at C-23
has revealed that proton elimination occurred exclu-
sively from the Z-methyl group at the terminal position
of 1, but not from the E-methyl group (Scheme 1).1,14) In
addition, we have suggested that the binding force for
the Z-methyl group to SHC was stronger than that for the
corresponding E-methyl group.13) Based on the X-ray
three-dimensional crystal structures of SHC, Wend et al.
have suggested the putative structure for the deprotona-
tion site involving some amino acids and water
molecules.2,15,16) According to their suggestion from
the results of an X-ray structural analysis, we have
illustrated the putative structure around the deprotona-
tion site as shown in Fig. 1. The catalytic base

730
T. SATO et al.
Fig. 1. Putative Structure for the Deprotonation Site of A. acidocaldarius SHC.
The locations of amino acid residues and water molecules are illustrated according to the X-ray crystal structure of SHC.16) Two types of water
molecules are depicted: the ‘‘front water’’ and ‘‘back waters’’ are represented by F and B, respectively. The ‘‘front water’’ and ‘‘back waters’’ are
associated with each other by hydrogen bonds, and the latter ones further construct the hydrogen-bonded network involving several amino acids.
The increased polarity of the ‘‘front water’’ can effectively behave as the catalytic base to form 2. The ‘‘front water’’ further forms the hydrogen
bond with Q262. The amino acid residues responsible for the formation of the hydrogen-bonded network including the ‘‘back waters’’ are as
follows: T41, E45, E93, R127, W133 and Y267. The P263 and F437 residues are close to the ‘‘front water’’. The F434 and C435 residues,
situated on the mobile loop, may behave as a gate that permits substrate passage. Y267 and F437 are aligned close to the substrate passage to
guide the substrate into the reaction cavity from the gate. This channel is shown with a dotted arrow.
results of mutagenesis experiments targeted on any other
of these described residues have been reported before.
Figure 1 also shows some amino acids responsible for
substrate entrance from the enzyme surface into the
active site cavity through a channel which contains a
narrow constriction formed by the four F166, V174,
F434 and C435 residues.15,16) It has been suggested that
the F434 and C435 residues on the mobile loop may also
behave as a gate that permits substrate passage.15,16)
Milla et al. have recently demonstrated that C435
actually has such a gate function by using a thiol-
modifying inhibitor.19) The function of F434 located at
the gate was also examined and is described here. The
locations of Y267 and F437 are also near the interface
between the putative deprotonation site and the substrate
gate (Fig. 1).
instrument. The reported chemical shifts of 1H- and 13C-
NMR (ppm) are relative to 7.28 and 128.0 ppm of the
solvent peak. MS spectra were recorded with a Jeol SX
102 mass spectrometer. GC analyses were carried out by
a Shimadzu GC-8A instrument with a DB-1 capillary
column (0:53 mm ꢀ 30 m, injection temperature of
290ꢁC, column temperature of 200–270ꢁC at 1ꢁC minꢂ1
,
and flow rate of carrier N2 of 0.37 kg/cm2). Specific
rotation values were measured at 25ꢁC by a Horiba
SEPA-300 instrument.
General methods. The detailed experimental protocol
has been reported in the previous papers20,21) for the
overexpression systems with the pET vector in E. coli
BL21 (DE3), DNA sequence analyses, preparation of
cell-free homogenates and enzyme purification methods.
In order to understand in detail the catalytic mech-
anism for the final deprotonation reaction, we carried out
mutagenesis experiments targeted on the ten residues,
T41, E45, E93, R127, W133, Q262, P263, Y267, F434
and F437, and functional analyses of the amino acid
residues are reported here.
Site-directed mutagenesis. All the site-directed muta-
genesis experiments were performed as described in the
previous papers.20,21) The following primers were used:
T41A, 50-pd[CTCCGCTTCCATGGCGACGTTGCTC-
AG]-30 (+Nco I); E45A, 50-pd[GAGGACGTACGCCG-
CTTCCATGGTGACGTTGCTCAG]-30 (+Nco I);
E93A, 50-pd[CACGACCATCGCGGCCTACGTCGC-
GC]-30 (ꢂBsiW I); R127Q, 50-pd[GAGTCGTCGCAA-
GTGTTCACGCGTATGTGGCTGG]-30 (+Mlu I);
W133A, 50-pd[GTGTTCACGCGTATGGCGCTGGCG-
Materials and Methods
1
Analytical methods. H-NMR and 13C-NMR spectra
were measured in C6D6 solutions by a Bruker DPX 400

Deprotonation Site of Squalene Cyclase
731
CTGG]-30 (+Mlu I); Q262G, 50-pd[CTGGGGCGG-
GATTGGGCCCCCTTGGTTTTACG]-30 (+Apa I);
Q262A, 50-pd[GGAGACGGCAGTTGGGGCGGGAT-
TGCGCCGCCTTGG]-30 (ꢂPvu II); P263G, 50-pd[GC-
TGGGGCGGAATTCAGGGGCCTTGGTTTTACG]-30
(+EcoR I); P263A, 50-pd[GCTGGGGCGGAATTCA-
GGCGCCTTGGTTTTAC]-30 (+EcoR I); Y267A, 50-
pd[GCCTTGGTTTGCCGCGCTCATAGCGCTCAAG-
ATTC]-30 (+Aor51 HI); F434A, 50-pd[CTTCGCCGA-
AGTCGCATGCCGGGATGTGGTTCGG]-30 (+Sph I);
and F437A, 50-pd[GGTCACTTCGCCGGCGTCGCAG-
AACGG]-30 (+Nae I). The bold letters designate the
altered bases, and the italic letters show the target
mutations. The underlined letters show the silent
mutations for easy screening of the desired mutants by
a restriction fragment analysis. The created or deleted
restriction sites are shown in parentheses. To ascertain
that the desired mutation had been carried out, the entire
region of all the inserted DNAs was sequenced.
were collected by centrifugation and washed twice with
a Tris-HCl buffer (pH 8.0, 50 mM). To the collected
pellets, 10 ml of a Tris buffer supplemented with 1%
Triton X-100 (w/v) was added. The bacterial cells were
subjected to ultrasonication at 4ꢁC and then centrifuged
to remove the cell debris to prepare a cell-free
homogenate. The pH value of the supernatant was
adjusted to 6.0 by adding 0.5 M citric acid. Five ml of the
cell-free extract thus prepared, which corresponds to ca.
0.5 mg of the homogeneously purified SHC, was used
for evaluating the product distribution. One mg of 1 was
separately incubated at 45ꢁC for 16 h. A hexane extract
was obtained from the reaction mixture. The detergent
was removed by passing the extract through a short SiO2
column [n-hexane:EtOAc = 100:20], and then the ex-
tract subjected to GC analyses to examine the product
distribution pattern and to quantify the product amounts.
The enzymatic products were identified by comparing
their GC-MS fragment patterns with those of authentic
samples that had previously been isolated by us.
Structural determination of product 5. Compound 5
had previously been isolated from the reaction mixture
of the mutated SHC of I261G (24% of all products).11)
However, we did not previously report the NMR data.
Complete NMR assignments of 5 were done by 2D-
NMR methods including COSY 45, HOHAHA, NO-
ESY, HMQC and HMBC. The spectroscopic data for
Enzyme assay. All the mutated SHCs were homoge-
neously purified according to the protocol described in
the previous paper.20,21) A mixed solution, which was
composed of 0.5 mM squalene, 0.2% Triton X-100 and
5 ꢃg of the homogeneously purified enzyme in a sodium
citrate buffer (60 mM, pH 6.0), was prepared in a final
volume of 5 ml for the enzyme reactions. Incubation was
done for 60 min at different temperatures (30, 35, 40, 45,
50, 55, 60, 65 or 70ꢁC) to examine the thermal stability
of each cyclase. To evaluate the kinetic parameters,
incubation was conducted for 60 min at 45ꢁC before
adding 15% methanolic KOH (6 ml) to terminate the
enzyme reaction. The lipophilic products (2, 3 and, in
some cases, 5) and unreacted 1 were extracted with n-
hexane (5 ml ꢀ 4) from each reaction mixture, and the
final products were quantified by GC analyses with a
DB-1 capillary column (30 m in length). The kinetic
values of Km and kcat were estimated from Lineweaver-
Burk plots.
1
compound 5 (solid, hop-21(22)-ene) are as follows: H-
NMR (400.13 MHz, C6D6) ꢁH (ppm): 2.40 (1H, m, H-
20), 2.30 (1H, m, H-16), 2.25 (1H, m, H-20), 1.95 (3H,
bs, H-29), 1.84 (1H, bd, J ¼ 12:4 Hz, H-17), 1.78 (3H,
bs, H-30), 1.78 (2H, m, H-1 and H-16), 1.69 (1H, m, H-
19), 1.66 (1H, m, H-11), 1.63 (2H, m, H-2 and H-6),
1.62 (1H, m, H-7), 1.59 (2H, m, H-12), 1.52 (1H, m, H-
13), 1.51 (1H, m, H-3), 1.47 (1H, m, H-15), 1.42 (2H, m,
H-9 and H-11), 1.41 (2H, m, H-2 and H-6), 1.37 (1H, m,
H-7), 1.35 (1H, m, H-15), 1.28 (1H, m, H-3), 1.13 (1H,
m, H-19), 1.125 (3H, s, H-27), 1.10 (3H, s, H-26), 1.04
(3H, s, H-23), 0.994 (3H, s, H-24), 0.988 (3H, s, H-25),
0.89 (1H, m, H-1), 0.88 (1H, m, H-5), 0.84 (3H, s, H-
28). 13C-NMR (100.6 MHz, C6D6) ꢁC (ppm): 135.8 (C-
21), 120.6 (C-22), 56.5 (C-5), 56.3 (C-17), 50.8 (C-9),
48.4 (C-13), 44.6 (C-18), 42.4 (C-3), 42.2 (C-14), 41.7
(C-8), 40.6 (C-1), 39.5 (C-19), 37.7 (C-10), 33.62 (C-
23), 33.60 (C-7), 33.4 (C-4), 33.1 (C-15), 28.8 (C-20),
24.0 (C-12), 23.7 (C-16), 23.0 (C-30), 21.8 (C-24), 21.3
(C-11), 19.7 (C-29), 19.09 (C-6), 19.07 (C-2), 16.87 (C-
27), 16.84 (C-26), 16.1 (C-25). EIMS fragments m=z
(%): 410 (45) [Mþ], 395 (14), 367 (38), 341 (54), 231
(26), 218 (10), 205 (16), 203 (18), 191 (95), 189 (100),
Results and Discussion
To understand the catalytic mechanism for the
deprotonation reaction, the amino acid residues shown
in Fig. 1 were mutated. The following twelve mutants
were constructed: T41A, E45A, E93A, R127Q, W133A,
Q262G, Q262A, P263G, P263A, Y267A, F434A and
F437A. According to the location sites shown in Fig. 1,
these mutants are classified into three categories.
Category A: Q262G, Q262A, P263G and P263A, which
were located between 4 and the ‘‘front water’’. Category
B: T41A, E45A, E93A, R127Q and W133A, which were
positioned around the ‘‘back waters’’. Category C:
Y267A, F434A and F437A, which were situated near
the substrate channel.
161 (68). HREIMS for C30H50: calcd., 410.3913; found,
25
410.3900. ½ꢂꢃ
+29.1 (c ¼ 0:68, CHCl3), cf. +29.8
D
(c ¼ 0:3{0:6, CHCl3) in ref. 22.
Product distribution pattern. The E. coli transform-
ant, in which the mutated SHC was expressed, was
cultured at 30ꢁC for 20 h in a Luria-Bertani medium
(100 ml) containing 50 mg/l of ampicillin. The cells

732
T. SATO et al.
Product distribution pattern of mutated SHCs
ene (Scheme 1) by co-injection and comparing the GC-
MS fragment pattern with that of an authentic sample
which had previously been isolated by us from the
I261G mutant11) (see the Materials and Methods
section).
The product distribution patterns of all the mutated
SHCs are summarized in Table 1. All the mutants
produced hydroxylated 3 in a higher yield than that of
the wild-type (Table 1), indicating that the ‘‘front water’’
was used to add to the C-22 carbocation of 4 instead of
accepting the proton of Me-29 as the catalytic base
(Fig. 1). It is of particular interest that the Q262G,
G262A, P263G and P263A mutants (category A) pro-
duced 3 as the main product (60–73%) instead of 2.
These mutants (category A) also produced 5, although in
a small quantity, which can be produced by the proton
We have previously shown that point mutants, which
had been altered at active sites, afforded prematurely
cyclized products. Based upon the product distribution
patterns, the positions of numerous active residues
inside the central active cavity have been identified.1)
With a large amount of the cell-free homogenate from
E. coli encoding the mutated SHCs, 1 was incubated at
45ꢁC for 16 h, and the enzymic products were analyzed
by GC. As a typical example, the product distribution
patterns for the wild-type and P263G mutant are shown
in Fig. 2. The production of 3 by the P263G mutant was
significantly higher than that of the wild-type. Together
with 2 and 3, compound 5 was additionally produced by
the P263G mutant. No other aberrant cyclization product
was detected. Product 5 was identified to be hop-21(22)-
Fig. 2. Gas Chromatograms Showing the Product Distribution of the Wild-type and Mutated P263G SHCs.
A marked increase in hopanol 3 and new product 5 were found for the P263G mutant.
Table 1. Product Ratios (%) of the Wild-type and Mutant SHCs and Their Relative Activities (%)
Squalene 1 (1.00 mg) was incubated with 5 ml of a cell-free extract, the amount of which corresponds to a half mg of each purified enzyme, at pH
6.0, 45 ꢁC, and for 16 h (see the Materials and Methods section). Product ratios were estimated by GC analyses. Figures in parentheses show the
relative activity (%) of each mutated SHC, compared to that of the wild-type, which were determined by incubating 1 (1.00 mg) with 5 ꢃg of the
homogeneously purified SHC for 60 min. The incubation was done for a short time by using a small amount of the SHC, because a large amount of the
SHC and a prolonged incubation fully converted 1 into 2 and 3.
Hopene
(2)
Hopanol
(3)
SHC
Category*
5
Wild-type
84.1 (100)
15.9 (100)
—
Q262G
Q262A
P263G
P263A
(A)
(A)
(A)
(A)
22.4 (96)
35.9 (113)
20.1 (87)
21.1 (76)
71.8 (1753)
60.1 (1071)
73.4 (1502)
72.9 (1540)
5.8
4.0
6.5
6.0
T41A
E45A
(B)
(B)
(B)
(B)
(B)
82.0 (92)
61.3 (41)
67.4 (36)
68.2 (54)
76.4 (74)
18.0 (97)
38.7 (133)
32.6 (98)
31.8 (98)
23.6 (112)
—
—
—
—
—
E93A
R127Q
W133A
Y267A
F434A
F437A
(C)
(C)
(C)
75.3 (88)
77.9 (58)
67.5 (130)
24.7 (129)
22.1 (85)
32.5 (331)
—
—
—
* Category A–C was classified according to the location site (see Fig. 1).

Deprotonation Site of Squalene Cyclase
733
elimination reaction of H-21 of intermediate 4 (path c in
Scheme 1), but 5 was not produced by the mutants of
categories B and C (Table 1).
decreased, while that of 3 was markedly increased at
lower temperatures by these mutations. The data in
Table 1 further support this result. The markedly
increased specific activities for all the products
(2 þ 3 þ 5) at low temperatures (Fig. 3 III) were thus
due to the significantly increased production of 3. We
have previously reported that the replacement of Phe365
and Phe605 by Tyr or Trp gave a marked increase in kcat
at lower temperatures. However, the mutants replaced
by aliphatic amino acids lacking ꢀ-electrons gave
premature cyclization products, which were generated
from the corresponding cationic intermediate(s), without
completion of the polycyclization reaction.9,10) Thus,
aromatic ꢀ-electrons play a crucial role in the accel-
eration of reaction velocity, resulting from the stabiliza-
tion of transient carbocations through a cation-ꢀ
interaction.9,10,23) However, the methyl group and hydro-
gen atom of the Ala and Gly mutants (Q262G, Q262A,
P263G and P263A) cannot stabilize a transient carbo-
cation(s), due to the lack of ꢀ-electrons. The residues of
Enzyme activities of mutated SHCs
The specific enzyme activities of mutant categories
A–C against the incubation temperature are illustrated in
Figs. 3–5, respectively. The specific activities for the
formation of 2, 3 or all the products (2 þ 3 þ 5) are
shown in I–III, respectively. The nine E45A, E93A,
R127Q, W133A, Q262G, Q262A, P263G, P263A and
F437A mutants had decreased activities in the high-
temperature range (Figs. 3–5). However, the optimal
temperature of the three T41A, Y267A and F434A
mutants remained unchanged. The kinetic data for
mutant categories A–C are shown in Tables 2–4,
respectively.
Category A
Figures 3 I and II show that the production of 2 was
Fig. 3. Specific Enzyme Activities of the Wild-type and Mutated SHCs in Category A against Incubation Temperature.
One mg of 1 was incubated with 5 ꢃg of each homogeneously purified SHC for 60 min at pH 6.0. The enzyme activities estimated by the
produced amounts of 2, 3, and all the products (2 þ 3 þ 5) are illustrated in panels I, II and III, respectively. The definition of symbols is as
follows:
wild-type,
Q262G,
Q262A, * P263G,
P263A.
Fig. 4. Specific Enzyme Activities of the Wild-type and Mutated SHCs in Category B against Incubation Temperature.
The incubation conditions were the same as those described for Fig. 3. The enzyme activities estimated by the amounts of 2, 3, and 2 þ 3 are
depicted in panels I, II and III, respectively.
wild-type,
T41A, * E45A,
E93A,
R127Q,
W133A.

734
T. SATO et al.
Fig. 5. Specific Enzyme Activities of the Wild-type and Mutated SHCs in Category C against Incubation Temperature.
The incubation conditions were the same as those described for Figs. 3 and 4. The enzyme activities estimated by the amounts of 2, 3, and
2 þ 3 are depicted in panels I, II and III, respectively.
wild-type,
Y267A,
F434A, * F437A.
Table 2. Kinetic Data and Optimal Temperatures for the Wild-type and Mutated SHCs in Category A
The kinetic values of Km and kcat were determined by incubating at 45ꢁC, pH 6.0 and for 60 min, during which, the purified proteins were not
denatured. These kinetic data were each estimated for the formation of 2, 3 and all the products (2 þ 3 þ ð5Þ).
Estimated
product
Km
kcat
(minꢂ1
)
Relative activity
(%)
Optimal temperature
for 60 min (ꢁC)
SHC
kcat=Km
(ꢃM)
2
3
1:62 ꢀ 10
1:62 ꢀ 10
1:62 ꢀ 10
51.8
9.8
3.19
0.60
3.79
100.0
100.0
100.0
60
60
60
Wild-type
2 þ 3
61.6
2
3
1:56 ꢀ 102
1:56 ꢀ 102
1:56 ꢀ 102
1:02 ꢀ 102
1:02 ꢀ 102
1:02 ꢀ 102
2:37 ꢀ 102
2:37 ꢀ 102
2:37 ꢀ 102
1:85 ꢀ 102
1:85 ꢀ 102
1:85 ꢀ 102
82.1
263.2
366.5
5:26 ꢀ 10ꢂ1
1.69
2.35
16.5
281
61.8
35–50
35–50
35–50
Q262G
Q262A
P263G
P263A
2 þ 3 þ 5
2
3
72.8
121.9
202.8
7.14
1.20
1.99
22.4
200
52.4
45–55
45–55
45–55
2 þ 3 þ 5
2
3
63.4
231.5
315.4
2:68 ꢀ 10ꢂ1
9:77 ꢀ 10ꢂ1
1.33
8.4
162.8
35.1
50
50
50
2 þ 3 þ 5
2
3
65.6
226.6
310.9
3:55 ꢀ 10ꢂ1
1.22
1.68
11.1
203.3
44.3
50
50
50
2 þ 3 þ 5
Q262 and P263 are located between 4 and the ‘‘front
water’’, as shown in Fig. 1.15,16) Table 2 shows that the
values of kcat and Km for the formation of all the
products (2 þ 3 þ 5) were higher than those of the wild-
type (kcat: 1.2–1.6-fold for 2, 12.4–26.9-fold for 3, 3.3–
6.0-fold for all the products; Km: 5.9–14.6-fold). This
finding indicates that the affinity of the cyclase to 1 was
less than the wild-type, despite the reaction velocity of
these mutants having been faster for the formation of all
the products. The decreased binding of 1 would have
occurred due to the decreased steric bulk size of the Ala
or Gly mutants, indicating the requirement of an
appropriate steric bulk size at positions 262 and 263.
Perturbation around these residues may have resulted in
closer contact of the ‘‘front water’’ to cationic inter-
mediate 4 and thus led to the significantly higher
production of 3. The slightly accelerated reaction
velocity for the formation of 2 at lower temperatures,
and the unusual formation of 5, which was produced by
the deprotonation reaction from H-21 (path c), may also
be explained in terms of the closer orientation of the
‘‘front water’’ to 4. As an alternative explanation,
perturbation around the ‘‘front water’’ may have
promoted the formation of 5, because the deprotonation
reaction from H-21 for the formation of 5 would be
more preferable to that from H-29 for 2 (a Zaitsev rule).
We thus propose that residues Q262 and P263 worked to
situate the catalytic base of the ‘‘front water’’ at the
appropriate position. The favorable placement of the
‘‘front water’’ would be effective for suppressing the
production of by-products 3 and 5. Indeed, the Q262G
and P263G mutants having the smaller side chain
produced 3 and 5 in higher yields than the Q262A and
P263A mutants (Table 1). We have previously reported

Deprotonation Site of Squalene Cyclase
735
Table 3. Kinetic Data and Optimal Temperatures for the Wild-type and Mutated SHCs in Category B
The incubation conditions were the same as those described for Table 2. These kinetic data were each estimated for the formation of 2, 3 and all the
products (2 þ 3).
Estimated
product
Km
kcat
(minꢂ1
)
Relative activity
(%)
Optimal temperature
for 60 min (ꢁC)
SHC
kcat=Km
(ꢃM)
2
3
1:62 ꢀ 10
1:62 ꢀ 10
1:62 ꢀ 10
51.8
9.8
3.19
6:05 ꢀ 10ꢂ1
3.79
100.0
100.0
100.0
60
60
60
Wild-type
2 þ 3
61.6
2
3
1:67 ꢀ 10
1:67 ꢀ 10
1:67 ꢀ 10
44.1
9.8
2.64
5:87 ꢀ 10ꢂ1
3.23
82.8
97.0
85.2
60
60
60
T41A
2 þ 3
53.9
2
3
2:13 ꢀ 10
2:13 ꢀ 10
2:13 ꢀ 10
20.7
13.3
34.0
9:72 ꢀ 10ꢂ1
6:24 ꢀ 10ꢂ1
1.60
30.5
103
42.2
50
50
50
E45A
2 þ 3
2
3
1:89 ꢀ 10
1:89 ꢀ 10
1:89 ꢀ 10
21.2
10.4
31.6
1.12
5:50 ꢀ 10ꢂ1
1.67
35.1
90.9
44.1
55
55
55
E93A
2 þ 3
2
3
2:55 ꢀ 10
2:55 ꢀ 10
2:55 ꢀ 10
26.9
12.8
39.7
1.05
5:02 ꢀ 10ꢂ1
1.56
32.9
83.0
41.2
50
50
50
R127Q
W133A
2 þ 3
2
3
1:69 ꢀ 10
1:69 ꢀ 10
1:69 ꢀ 10
37.3
11.5
48.8
2.21
6:80 ꢀ 10ꢂ1
2.89
69.3
112.4
76.3
55
55
55
2 þ 3
Table 4. Kinetic Data and Optimal Temperatures for the Wild-type and Mutant SHCs in Category C
The incubation conditions were the same as those described for Table 2 and 3. These kinetic data were each estimated for the formation of 2, 3 and
all the products (2 þ 3).
Estimated
product
Km
kcat
(minꢂ1
)
Relative activity
(%)
Optimal temperature
for 60 min (ꢁC)
SHC
kcat=Km
(ꢃM)
2
3
1:62 ꢀ 10
1:62 ꢀ 10
1:62 ꢀ 10
51.8
9.8
3.19
6:05 ꢀ 10ꢂ1
3.79
100.0
100.0
100.0
60
60
60
Wild-type
2 þ 3
61.6
2
3
1:97 ꢀ 102
1:97 ꢀ 102
1:97 ꢀ 102
38.5
12.6
51.1
1:95 ꢀ 10ꢂ1
6:40 ꢀ 10ꢂ2
2:59 ꢀ 10ꢂ1
3:96 ꢀ 10ꢂ1
1:13 ꢀ 10ꢂ1
5:09 ꢀ 10ꢂ1
6:44 ꢀ 10ꢂ1
3:10 ꢀ 10ꢂ1
9:54 ꢀ 10ꢂ1
6.1
10.6
6.8
50–60
50–60
50–60
Y267A
F434A
F437A
2 þ 3
2
3
8:16 ꢀ 10
8:16 ꢀ 10
8:16 ꢀ 10
32.3
9.2
12.4
18.7
13.4
60
60
60
2 þ 3
41.5
2
3
9:55 ꢀ 10
9:55 ꢀ 10
9:55 ꢀ 10
61.5
29.6
91.1
20.2
51.7
25.1
45
45
45
2 þ 3
the enzymic products from the mutants of Phe605A,10)
but 5 could not be isolated, possibly due to its low
production. We reexamined the enzymic products from
the mutants of Phe605A; GC-MS analyses clearly
showed the involvement of 5 in the reaction mixtures,
although in a small amount (3.8%). As found with the
P263A and P263G mutants, the less bulky size at the
605 position would also have induced perturbation
around the ‘‘front water’’, resulting in the high produc-
tion of 310) and 5. As shown in Fig. 1, the terminal amide
group of Q262 forms a hydrogen bond with the ‘‘front
water’’. The substitution of Gln with Ala or Gly could
also disrupt the hydrogen bond, thus leading to higher
production of 3 and 5 due to perturbation around the
‘‘front water’’.
Category B
As shown in Table 1, 2 and 3 were produced, although
there was no detectable amount of 5, by the category B
mutants (T41A, E45A, E93A, R127Q and W133A). The
production of 2 was less throughout the entire temper-
ature range examined (Fig. 4 I) than that by the wild-
type, but that of 3 was almost the same as that of the
wild-type, being slightly higher in the low-temperature
ranges (Fig. 4 II). The specific activities for the
formation of (2 þ 3) were smaller than those of the

736
T. SATO et al.
wild-type throughout the entire temperature range
examined (Fig. 4 III). The kinetic analyses revealed
that kcat for the production of 2 was lower (40–85%), but
that for the production of 3 was slightly higher (106–
135%) than by the wild-type (Table 3). These mutations
would have disrupted the hydrogen-bonding network
constructed for the ‘‘back waters’’, thus lowering the
polarity of the ‘‘front water’’ and leading to a slower
reaction for the deprotonation of H-29 (path a) to form 2.
The nucleophilic property of the ‘‘front water’’ to the C-
22 cation (path b) for the formation of 3 was slightly
increased. This may suggest that the correct position of
the ‘‘front water’’ could have been slightly disordered
due to breakage of the hydrogen bond between these
amino acid residues and the ‘‘back waters’’ by a single
amino acid substitution. The difference in Km for the
T41A, E45A, E93A, R127Q and W133A mutants was
minor (1.0–1.5-fold) in comparison with that of the
wild-type. This finding suggests that the affinity of 1 to
the cyclase was not affected by these mutations
(Table 3); that is, the interactive contact of substrate 1
with these amino acid residues was little. The relative
activity, kcat=Km, for the formation of 2 was significantly
decreased by the mutations. This finding further supports
the idea that the hydrogen-bonding network, which had
been constructed by assembling the residues of E45,
E93, R127 and W133 including the ‘‘back waters’’,
strongly worked to polarize the ‘‘front water’’, leading to
the higher production of 2 than of 3. There was little
difference in the relative activity (kcat=Km) for the
formation of 3, suggesting that the position of the ‘‘front
water’’ remained unchanged or only slightly altered by
these site-directed mutations. There was also little
difference in the kinetic values and specific activities
with incubation temperature (Table 3 and Fig. 4) for the
T41A and W133A mutants when compared to the
figures for other mutants E45A, E93A and R127Q. This
finding indicates that polarization of the ‘‘front water’’
could have been archived mainly by the three E45, E93
and R127 residues.
Y267A had an exceptionally large Km value, probably
due to the location of Y267 in the proximity of the
substrate passage. A higher Km value was also observed
with the F434 mutant situated at the substrate entrance.
In respect of the specific activity against the incubation
temperature, mutated F437A showed a different profile
from that of the F434A or Y267A mutant; at lower
temperatures, the production of 2 and 3 was each higher
than that of the wild-type (Fig. 5 I–III). As shown in
Table 4, the increase in kcat for the production of 3 (3-
fold) was higher than that of 2 (1.2-fold). This profile is
characteristic of category A (Fig. 3 I–III). The higher
values of Km and kcat with F437A at lower temperatures
also supports the classification of F437 into category A,
but not into category B, suggesting that the F437 residue
is likely to have been situated near the ‘‘front water’’,
and not near the ‘‘back waters’’. However, the increase
in the formation of 3 (3-fold) was significantly smaller
than that of the category A mutants (12–27-fold)
(compare Tables 2 and 4). In addition, the formation
of product 5, characteristic of category A, was not
detected by the F437A mutant. These findings suggest
that F437 was located at three interfaces among the
‘‘front water’’, ‘‘back waters’’ and substrate channel,
which is in a good agreement with the inference from
the X-ray analysis (Fig. 1).
We have previously reported the kinetic data for the
F129A mutant:17) Km 1:85 ꢀ 10 ꢃM and kcat 13.7 minꢂ1
for the formation of 2; Km 1:85 ꢀ 10 ꢃM and kcat
2.5 minꢂ1 for that of 3. This Km value is nearly equal to
that for the wild-type, while kcat is significantly lower.
F129 is not involved in the hydrogen-bonded network
with the ‘‘back waters’’ and is located around residues
Y267 and W133 as shown in Fig. 1. The hydrogen bond
could be disrupted around Y267 and W133 due to the
decreased bulk size by mutating Phe into Ala, leading to
a slower reaction rate as shown for category B. Thus, no
detectable amount of 5 and no increased amount of 3
was observed for the mutant F129A.
In conclusion, the results of the site-directed muta-
genesis experiments described here allowed us to
identify the amino acid residues that play a crucial role
in the final deprotonation reaction leading to 2. Sub-
stitution of Gln with Ala at the 262 position disrupted a
hydrogen bond of the side chain of Gln with the ‘‘front
water’’, thus enabling the disordered ‘‘front water’’ to
approach final cation 4. This could lead to the formation
of a larger quantity of 3 than 2. Pro263 did not form a
hydrogen bond with the ‘‘front water’’, but the Ala and
Gly mutants produced 3 in a significantly large amount.
This would have been due to perturbation of the correct
position of the ‘‘front water’’ induced by the decreased
bulk size of Ala and Gly. The F437A mutant also
showed a high production of 3, but the increase was less
than that by P263, suggesting that Pro263 was closer to
the ‘‘front water’’ than in F437. Several amino acids are
responsible for the hydrogen-bonded network with the
Category C
The F434 residue is located at the substrate gate
(Fig. 1). Compared to the wild-type, the Ala mutant had
significantly lower affinity (5-fold in Km), although
difference in reaction velocity was small (1/1.6 fold in
kcat), as shown in Table 4. This may suggest an
important role of the aromatic residue for capturing
substrate 1 and for guiding 1 into the reaction cavity.
The Km value of Y267A for the formation of 2, 3 or
(2 þ 3) was also each significantly decreased by 12-fold.
The Y267 residue participates in the hydrogen-bonding
network with the ‘‘back water’’ (Fig. 1); thus, Y267 may
be classified into category B. The lower of kcat value for
the formation of 2 is characteristic of category B.
Indeed, kcat decreased by 1/1.3 fold with a small change.
The difference in Km for category B was little by the
point mutations, as shown in Table 3. However, mutated

Deprotonation Site of Squalene Cyclase
Soc. Chem. Commun., 291–301 (2002).
737
‘‘back waters’’, which form further hydrogen bonds with
the ‘‘front water’’, thus increasing the polarity of the
‘‘front water’’ to give rise to a strong catalytic base for
the ‘‘front water’’. The disruption of the hydrogen-
bonded network responsible for the ‘‘back waters’’ led to
reduced polarity of the ‘‘front water’’ (a weakened
catalytic base), thus giving less activity for the formation
of 2. The residues located at the substrate entrance or
passage were also examined. The F434A mutant
resulted in significantly decreased binding to the
substrate. The Y267 and F437 residues were aligned
near the passage. The mutation of these residues into Ala
also decreased the binding of the substrate with the
cyclase. The Phe166 aromatic residue was also located
at the substrate gate (Fig. 1). Aromatic residues may
play an important role in capturing a hydrophobic
substrate and guiding it into the reaction cavity. To
validate this assumption, further mutation experiments
are necessary with the residues F166, V174, V440 and
so on (see Fig. 1).
A variety of quenching sites for the last cation
generated during the polycyclization cascade leads to
structural diversity of the triterpene skeleton. At the
present time, the amino acid residue(s) responsible for
final reaction step (quenching of the final carbocation,
e.g., the deprotonation site or hydroxylation site) of the
triterpene cyclase family is still unknown, except for the
SHC. If detailed information can be collected for the
final reaction step, it will be possible to create many
‘‘unnatural natural’’ triterpenes in future. We have
demonstrated that the steric bulk size of the crucial
residues, which are in close contact with the folded and/
or extended substrate, significantly affected the folding
conformation of squalene, leading to novel triter-
penes.11,12) Very interesting reports have recently ap-
peared that squalene analogs (truncated C29 analogs and
an elongated C35 analog) were successfully cyclized to
give novel carbocyclic skeletons by using A. acido-
caldarius squalene-hopene cyclase24,25) and that the
natural indole diterpene, petromindole, was synthesized
in a single step from the indole derivative of the
geranylgeranyl isoprenoid, 3-(!-oxidogeranylgeranyl)-
indole, by using Arabidopsis thaliana lupeol synthase (a
plant oxidosqualene cyclase).26) The combined use of
altered triterpene cyclases and substrate analogs will
also be promising for creating further unnatural natural
compounds and for one-pot syntheses of natural terpe-
noids.
2) Wendt, K. U., Schulz, G. E., Corey, E. J., and Liu, D. R.,
Enzyme mechanisms for polycyclic triterpene formation.
Angew. Chem. Int. Ed., 39, 2812–2833 (2000).
3) Abe, I., Rohmer, M., and Prestwich, G. D., Enzymatic
cyclization of squalene and oxidosqualene to sterols and
triterpenes. Chem. Rev., 93, 2189–2206 (1993).
4) Segura, M. J. R., Jackson, B. E., and Matsuda, S. P. T.,
Mutagenesis approaches deduce structure-function rela-
tionship in terpene synthases. Nat. Prod. Rep., 20, 304–
317 (2003).
5) Kushiro, T., Shibuya, M., and Ebizuka, Y., Cryptic
regiospecificity in deprotonation step of triterpene bio-
synthesis catalyzed by new members of lupeol synthase.
Tetrahedron Lett., 40, 5553–5556 (1999).
6) Hoshino, T., Kouda, M., Abe, T., and Ohashi, S., New
cyclization mechanism for squalene: a ring expansion
step for the five-membered C-ring intermediate in
hopene biosynthesis. Biosci. Biotechnol. Biochem., 63,
2038–2041 (1999).
7) Sato, T., Abe, T., and Hoshino, T., On the cyclization
mechanism of squalene: a ring expansion process of the
five-membered D-ring intermediate. J. Chem. Soc.
Chem. Commun., 2617–2618 (1998).
8) Sato, T., and Hoshino, T., Functional analysis of the
DXDDTA motif in squalene-hopene cyclase by site-
directed mutagenesis experiments: initiation site of the
polycyclization reaction and stabilization site of the
carbocation intermediate of the initially cyclized A-ring.
Biosci. Biotechnol. Biochem., 63, 2189–2198 (1999).
9) Hoshino, T., and Sato, T., Functional analysis of
phenylalanine 365 in hopene synthase, a conserved
amino acid in the families of squalene and oxidosqualene
cyclases. J. Chem. Soc. Chem. Commun., 2005–2006
(1999).
10) Hoshino, T., Kouda, M., Abe, T., and Sato, T., Func-
tional analysis of Phe605, a conserved aromatic amino
acid in squalene-hopene cyclase. J. Chem. Soc. Chem.
Commun., 1485–1486 (2000).
11) Hoshino, T., Abe, T., and Kouda, M., Unnatural natural
triterpenes produced by altering isoleucine into alanine
at position 261 in hopene synthase and the importance of
having the appropriate bulk size at this position for
directing the stereochemical destiny during the polycyc-
lization cascade. J. Chem. Soc. Chem. Commun., 441–
442 (2000).
12) Sato, T., Sasahara, S., Yamakami, T., and Hoshino, T.,
Functional analyses of Tyr420 and Leu607 of Alicyclo-
bacillus acidocaldarius squalene-hopene cyclase. Neo-
achillapentaene, a novel triterpene with the 1,5,6-
trimethylcyclohexene moiety produced through a folding
of the constrained boat structure. Biosci. Biotechnol.
Biochem., 66, 1660–1670 (2002).
13) Hoshino, T., and Kondo, T., The cyclization mechanism
of squalene in hopene biosynthesis: the terminal methyl
groups are critical to the correct folding of this substrate
both for the formation of the five-membered E-ring and
for the initiation of the polycyclization reaction. J.
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Acknowledgments
This work was supported by grant-aid to T. H. (Nos.
13306009 and 15380081) and to T. S. (No. 15780083).
14) Hoshino, T., Nakano, S., Kondo, T., and Miyoshi, A.,
Report in preparation. Preliminary results have been
reported at Annual Meeting Japan Society for Biosci.
Biotechnol. and Agrochem., April 1998, Nagoya, Ab-
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