Báo cáo khoa học: Escherichia coli cyclopropane fatty acid synthase Mechanistic and site-directed mutagenetic studies potx

Escherichia coli
cyclopropane fatty acid synthase
Mechanistic and site-directed mutagenetic studies
Fabienne Courtois, Christine Gue
´
rard, Xavier Thomas and Olivier Ploux
Laboratoire de Chimie Organique Biologique, UMR7613 CNRS, Universite
´
Pierre et Marie Curie, Paris, France
Escherichia coli fatty acid cyclopropane synthase (CFAS)
was overproduced and purified as a His
6
-tagged protein.
This recombinant enzyme is as active as the native enzyme
with a K
m
of 90 l
M
for S-AdoMet and a specific activity of
5 · 10
)2
lmolÆmin
)1
Æmg
)1
. T he enzyme is devoid of organic
or metal cofactors and is unable to catalyze the wash-out of
the m ethyl protons of S-AdoMet to the solvent, data that do
not support the ylide mechanism. Inactivation of the enzyme
by 5,5¢-dithiobis-(2-nitrobenzoic acid) (DTNB), a pseudo
first-order process with a rate constant of 1.2

M
)1
Æs
)1
, is not
protected by substrates. Graphical analysis of t he inactiva-
tion by DTNB revealed that only one cyst eine is responsible
for the inactivation of the enzyme. The three strictly con-
served Cys residues among cyclopropane synthases, C139,
C176 and C354 of the E. coli enzyme, w ere mutated to
serine. The relative catalytic efficiency of t he mutants were
16% for C139S, 150% for C176S and 63% for C354S. The
three mutants were inactivated by DTNB at a rate com-
parable to the rate of inactivation of the His
6
-tagged wild-
type enzyme, indicating that the Cys responsible for the loss
of activity is not one of the conserved residues. Therefore,
none of the conserved Cys residues is essential for catalysis
and cannot be in volved in covalen t catalysis o r general
base catalysis. T he inactivation is p robably the result of
steric hindrance, a phenomenon irrelevant to catalysis. It is
very likely that E. coli CFAS operates via a carbocation
mechanism, but the base and nucleophile remain to be
identified.
Keywords: cyclopropane fatty acid s ynthase; hydrogen
isotope exchange; enzymatic reaction mechanism; site-
directed mutagenesis; chemical modification.
Cyclopropane synthases constitute an interesting class of
enzymes that catalyze the cyclopropanation of unsaturated

lipids in bacteria [1], plants [2,3] and parasites [4]. Escheris-
hia coli cyclopropane fatty acid synthase (CFAS) [5–9] and
its closely related homologs from Mycobacterium tuber-
culosis [10] are the best known representatives of this class
of enzymes. In E. coli, cyclopropanation is thought to be
involved in long-term survival of nongrowing cells and is
often associated with enviromental stresses [1]. In M. tuber-
culosis, cyclopropanation has recently been associated
with virulence and persistance of the pathogen [11]. Hence,
cyclopropane s ynthases m ight be good targets for new
antituberculous d rugs. Indeed, t uberculosis remains a major
cause of death in the world and there is a real need for new
drugs to combat strains of M. tuberculosis that are resistant
to existing drugs [12]. We have been interested in studying
CFAS from E. coli as a model for M. tuberculosis cyclo-
propane synthases, for which an in vitro assay is still lac king.
Our goal is to co ntribute to the elucidation of this intrigu-
ing enzymatic reaction, but also to discover inhibitors of
cyclopropane synthases that might be good leads to
antituberculous drugs [13].
This enzymatic cyclopropanation reaction proceeds by
transfer of a methylene group from the activated methyl
group of S-adenosyl-
L
-methionine (S-AdoMet) to the
(Z)-double bond of an unsaturated fatty acid chain,
resulting in the formation of a cyclopropane ring on the
alkyl chain (Scheme 1). Early in vivo studies [14–16] showed
that two of the three methyl protons of S-AdoMet are
retained in the product, although s ome exchange w as

observed under certain conditions [ 17,18], a nd that the
vinylic and allylic protons of the substrate are also retained
in the product. The stereochemistry is also retained; that is,
the (Z)-double bond gives a cis-cyclopropane [5], although
trans-cyclopropanes are also found in M. tuberculosis
mycolic acids [10]. Chiral methyl analysis was also conduc-
ted in v ivo using Lactobacillus plantarum cells, and showed
retention of t he stereochemistry of the reaction [19]. This
experimental observation is n ot in favor o f a carbenoid
species (see below), which would probably racemize.
Two types of reaction mechanism have b een proposed for
this fascinating reaction: a carbocation type and an ylide
type mechanism, schematically represented in Scheme 2.
Correspondence to O. Ploux, Laboratoire de Chimie Organique
Biologique – UMR CNRS 7613, Boıˆte 182, Tour 44–45, 4 P lace
Jussieu, F-75252 Paris cedex 05, France. Fax: +33 1 44 27 71 50,
Tel.: +33 1 44 27 55 11, E-m ail: ploux@ ccr.jussieu.fr
Abbreviations: BSA, bovine serum albumin; CFAS, cyclopropane
fatty acid synthase; DTNB, 5,5¢-dithiobis(2-nitrobenzoic acid);
EDTA, ethylenediaminetetraacetic acid; NEM, N-ethylmaleimide;
S-AdoMet, S-adenosyl-
L
-methionine; S-AdoHcy, S-adenosylhomo-
cysteine.
Enzymes: S-Adenosyl-
L
-homocysteine homocysteinylribohydrolase,
adenosylhomocysteine nuc leosidase (EC 3.2.2.9); S-adenosyl-
L
-

methionine:unsaturated-ph ospholipid methyltransferase (cyclizing),
cyclopropane-fatty-acyl-phospholipid synt hase, cycloprop ane
synthase (EC 2.1.1.79).
Note: A website is available at />(Received 28 July 2004, revised 16 September 2004,
accepted 18 October 2004)
Eur. J. Biochem. 271, 4769–4778 (2004) Ó FEBS 2004 doi:10.1111/j.1432-1033.2004.04441.x
Even though the mechanism involving a carbocation
intermediate is often cited in the literature [1,10,20], the
other reasonable alternatives deserve c onsideration and in
particular the metal-assisted ylide mechanism [21]. H owever,
recent crystallographic data [22], inhibition and mechanistic
studies [13,18,23,24], and data reported in this study argue
in favor of the carbocation mechanism. On the basis of
chemical modification experiments [9], the involvement of a
cysteine residue in the catalysis has been invoked. Indeed, the
thiolate side chain could be either the base that is required for
abstraction of t he methyl proton, or could stabilize the
carbocation, if that intermediate were formed, or even
participate in a covalent catalysis (Scheme 2). Interestingly,
the three dimensional structure of three cyclopropane
synthases from M. tuberculosis [22] showed the presence of
two cyseines at, or near, the active site: C139 and C354
(E. coli CFAS numbering). Futhermore, these residues, as
well as C176 (E. coli numbering), are strictly conserved in all
cyclopropane synthases discovered so far [1].
We report here the purification of a His
6
-tagged CFAS
and its characterization. We also report exchange experi-
ments that are not in favor of the ylide mechanism. The role

of the conserved cysteines was studied using chemical
modification and site-directed m utagenesis. It was found
that the cysteines are not essential for catalysis.
Experimental procedures
General
E. coli strains JM109 and BL21(DE3) were from Promega
(Madison, WI, USA), and E. coli K12 was obtained from
the Institut Pasteur Collection (CIP; Paris, France). Plasmid
pET-24(+) was obtained from Novagen (Darmstadt,
Germany). Synthetic oligonucleotides were products of
Proligo (Paris, France) and were used without any fur-
ther purification. Chemicals were purchased from Sigma-
Aldrich (Saint Quentin, France) and were of the highest
purity available. S-[Methyl-
14
C]adenosyl-
L
-methionine
(60 m CiÆmmol
)1
)andS-[methyl-
3
H]adenosyl-
L
-methionine
(85 CiÆmmol
)1
or 15 CiÆmmol
)1
)werefromNewEngland

Nuclear (Boston, MA, USA). Restriction enzymes, Taq
polymerase, T4 DNA ligase and molecular biology kits
were either from Promega or f rom Roche (Meylan, France).
Culture medium components were purchased from Difco
Laboratories (Detroit, MI, USA). Chromatographic equip-
ment (GradiFrac) and column phases were from Amersham
Biosciences (Orsay, France). UV-visible spectra were
obtained on an Uvikon-930 Kontron spectrophotometer
(Munchen, Germany) or a Lambda-40 Perkin Elmer
apparatus (Norwalk, CT, USA). Scintillation counting
was run on a 1214 Rackbeta LKB Wallac radioactivity
counter (Per kin Elmer). Sonication was performed on a
VibraCell sonicator from Bioblock (Illkirch, France). SDS/
PAGE was run on a Bio-Rad Protean II system (Hercules,
CA, USA), using the conditions described by the manufac-
turer, and DNA electrophoresis on a Mupid apparatu s
(Eurogentec, Seraing, Belgium), in 40 m
M
Tris/acetate
buffer, pH 7.5, 1 m
M
EDTA. Centrifugations were run o n
a Sorval RF5plus centrifuge (DuPont, K endro, Cortaboeuf,
France).
1
Hand
13
C-NMR spectra were obtained on an AC
400 M Hz Bruker apparatus (Rheinstetten, Germany).
Plasmid construction and site-directed mutagenesis

The w ild-type histidine-tagged CFAS recombinant g ene
was obtained using PCR amplification of the cfa gene from
E. coli K12 genomic DNA. Briefly, E. coli K12 genomic
DNA was purified using the Wizard Genomic kit from
Promega, and the cfa gene was amplified using Taq DNA
polymerase (Promega) and the following two primers:
5¢-CGCGAATTCAGGAGGATTTTATGCACCACCA
CCACCACCACAGTTCATCGTGTATAGAAGAA-3¢
containing an EcoRI site, a ribosome binding site and a
His
6
-tag sequence, and 5¢-CGCAAGCTTTTAGCGAGC
CACTCGAAG-3¢ containing a Hin dIII site. The DNA
fragments was purified (PCR Preps, Promega), digested by
EcoRI and Hi ndIII, purified on agarose gel and ligated into
pET-24(+) previously cut by the same restriction enzymes.
After transformation in E. coli JM109, positive clones were
selected and the plasmid extracted and purified (Wizard
Plus Minipreps, Promega) for DNA sequencing ( ECSG,
Evry, France). Plasmid pET-24H6cfa, thus obtained, was
used for t ransformation in E. coli BL2 1(DE3) and t his
construction afforded efficient expression of the enzyme.
The mutated cfa genes, cfaC139S, cfaC176S and cfaC354S
(numbering corresponds to the wild-type s equence, that
is without counting the N-terminal His
6
-tag that has
R
1
S

R
2
CH
3
CH
3
CH
3
Enz-Nu
H
H
C
H
H
H
R
1
S
R
2
H
3
C
Ylide
R
1
S
R
2
CH

2
OR
Enz–Base
Enz–BaseH
Enz–Nu
Enz–Nu
A
B
Enz Metal
Metal
CH
2
Enz–Nu
Enz–Base
Enz–Base
Carbenoid
H
H
Scheme 2. Plausible reaction mechanisms for the catalyzed cyclopro-
panation. (A) The carbocation mechanism; (B) the ylide mechanism.
This mech anism would most probably require a metalloenzyme and
transfer of a carbenoid to the metal (details are not shown for clarity).
H
H
HH
O
A
OH
HO
S

NH
3
OOC
CH
3
O
A
OH
HO
S
NH
3
OOC
CFAS
S-Adenosyl-L-methionine
S-Adenosyl-L-homocysteine
+ H
Scheme 1. Reaction catalyzed by the cyclopropane synthases. In E. coli
CFAS the lipid substrate is an unsaturated ph ospholipid, while in
M. tuberculosis the unsat urated alkyl chain is probably bound to an
acyl carrier protein.
4770 F. Courtois et al.(Eur. J. Biochem. 271) Ó FEBS 2004
been engineered) of E. coli CFAS were constructed
using the QuikChange Site-Directed Mutagenesis Kit from
Stratagene (La Jolla, C A, USA). The following sets of
mutated primers (mutations are underlined) w ere u sed:
C139S: 5¢-CATGCAATATTCC
AGCGCTTACTGGAA
AG-3¢ and 5¢-CTTTCCAGTAAGCGC
TGGAATATTG

CATG-3¢; C176S: 5¢-GGATATTGGC
AGCGGCTGGG
GCGGACTGGC-3¢ and 5¢-GCCAGTCCGCCCCAGCC
GC
TGCCAATATCC-3¢; C354S: 5¢-CTGAATGCCTCT
GCAGGTGCTTTCCGCGCC-3¢ and 5¢-GGCGCGCGG
AAAGCACCTGCA
GAGGCATTCAG-3¢. Plasmid pET-
24H6cfa was used as the template. Transformants
were selected and the plas mids were extracted, purified
(Wizard P lus miniprep kit from Promega) and se quenced
(Eurogentec) to ens ure integrity and the presence of the
desired mutation. In the case of the C354S mutant,
the mutation was confirmed by digestion with PstIasthe
mutation introduces a new restriction site. Each mutated
plasmid was then transformed into competent E. coli
BL21(DE3) for protein expression.
Protein assay
Protein concentrations were determined using the colori-
metric assay described by Bradford [25] and supplied by
Bio-Rad.
Phospholipids preparation
Unsaturated E. coli K12 phospholipids were prepared
according to Cronan [ 6] and as originally described by
Ames [26]. Once purified the phospholipids were stored as a
chloroform solution at )20 °C. Aqueous solution s of
phospholipids were prepared by evaporating the chloroform
and resuspending the phospholipids in 20 m
M
potassium

phosphate buffer, pH 7.4 at the desired concentration
(% 20 mgÆmL
)1
). Phospholipids were assayed using the
ferric hydroxamate method, as described previously [27],
and using tripalmitin standards for calibration. Phospho-
lipid solutions were sonicated f or 30 s for dispersion prior to
use as substrates. Cyclopropanated phospholipids were
extracted, using t he same protocol, from isopropyl thio-b-
D
-galactoside (IPTG)-induced E. coli BL21(DE3)/pET-
24H6cfa cells.
CFAS purification
An overnight preculture [10 mL Luria–Bertani (LB)
medium, 50 lgÆmL
)1
kanamycin] of E. coli BL21(DE3)/
pET-24H6cfawasusedtoinoculate800mLofLBmedium
supplemented with 5 0 lgÆmL
)1
kanamycin. The culture was
shaken (180 r.p.m., 37 °C), and when the absorbance a t
600 nm reached a v alue of 0.7, IPTG was added at a final
concentration o f 100 l
M
. T he culture was then shaken
overnight at 37 °C. The cells were collecte d by centrifuga-
tion (4000 g, 15 min), washed (0.1
M
potassium phosphate

buffer, pH 7.4), centrifuged (4000 g, 15 min) and kept at
)20 °C until use. The cell paste was resuspended in 40 m L
of 20 m
M
potassium phosphate buffer, p H 7.4, and the
suspension was sonicated on ice (5 min, with 1 min cooling
period every minute). After centrifugation (15 000 g,
20 min), the supernatent was loaded directly on a nickel
affinity column (Chelating Sepharose, Amersham Bio-
science; 1.6 cm i.d., 5 cm long, 10 mL) prepared as recom-
mended by the manufacturer and equilibrated with buffer A
(20 m
M
potassium phosphate buffer, pH 7.4, 0.5
M
NaCl).
The column was successively washed with 30 mL of buffer
A and 30 mL of buffer A containing 5% (v/v) of buffer B
(20 m
M
potassium phosphate buffer, pH 7.4, 0.5
M
NaCl,
1.0
M
imidazole). T he proteins were eluted by a linear
gradient starting from 5% (v/v) of buffer B to 40% (v/v) of
buffer B, in buffer A . The column was run at a flow rate of
1mLÆmin
)1

and 7 mL fractions were collected. The pres-
ence of proteins was detected using the Bradford assay and
the purity of i ndividual fractions was analyzed by SDS/
PAGE. Fractions containing pure CFAS were pooled and
desalted on PD-10 columns (Amersham Bioscience) equili-
brated with buffer A. Highly concentrated enzyme solu-
tions were obtained by ammonium sulfate precipitation as
follows. Solid ammonium sulfate was added at 0 °Ctothe
enzyme solution, up to 40% saturation, and the precipitated
protein was recovered by centrifugation (10 min at
12 000 g). The pellet was then dissolved in the minimum
volume of 20 m
M
potassium pho sphate buffer , pH 7.4,
50% (v/v) glycerol, and the enzyme solution was stored at
)20 °C. The mutant proteins, which all carry an N -terminal
His
6
-tag, were purified as des cribed for the His
6
-tagged
wild-type enzyme.
Biochemical characterization
N-terminal protein sequencing of the enzyme, t ransfered
onto a polyvinyliden e fluoride membrane, was obtained at
the Plateau Technique d’An alyse et de Microsequenc¸ age
des Prote
`
ines (Institut Pasteur). For the determination of
the metal content, several samples (18 nmol each) of

purified CFAS were lyophilized and s ubjected to metal
analysis (Zn, Ni, Co, Fe, Cu) using ICP-AES methodology
(Service Central d’Analyse; CNRS, Vernaison, France).
The metal content, in each case, represented less than 1%
of what was expected for 1 mol of metal per mol of
enzyme.
CFAS assay
CFAS activity was assayed as described previously [7] with
slight modifications. The assay consisted in 1.0 mgÆmL
)1
phopholipids, 0.5 mgÆmL
)1
bovine serum albumin (BSA),
10% (v/v) glycerol, 2 m
M
dithiothreitol (dithiotheithol),
0.75 m
M
S-AdoMet, either
14
C-labelled (specific radioac-
tivity of 5.0 mC iÆmmol
)1
)or
3
H-labelled (specific radioac-
tivity of 1.5 mCiÆmmol
)1
), 2 lgCFAS,in20m
M

potassium
phosphate buffer, pH 7.4, in a final volume of 100 lL. The
reaction was initiated by addition of the enzyme and
incubated at 37 °C for 20 min. The reaction was stopped by
adding 1 mL 10% (v/v) trichloroacetic acid, a nd the solu-
tion was filtered over glass fi ber filters (Whatman GF/c,
Middlesex, USA; 25 mm). The filters, adapted on a
filtration device (Millipore, Billerica, M A, USA; 1225
model), were washed three times with 1 mL 10% (v/v)
trichloroacetic acid, three times with 1 mL H
2
O, oven-dried
(60 °C, 20 min) and finally counted for radioactivity in
5 mL o f s cintillation cocktail (Optiphase, Wallac). The
activity measured under these conditions was linear with
Ó FEBS 2004 Cyclopropane synthase reaction mechanism (Eur. J. Biochem. 271) 4771
time over a period of 2 0 min and linear with enzyme
concentration up to 0.1 mgÆmL
)1
of protein (data not
shown). One unit of CFAS is defined as the amount of
enzyme that transforms 1 lmol of substrate per min. The
kinetic parameters of His
6
-tagged wild-type and mutant
CFAS were determined by measuring the activity
(as described above for the e nzyme a ssay) at different
concentrations of S-AdoMet. D ata were a nalyzed using
nonlinear regression analysis run on
KALEIDAGRAPH

soft-
ware, to fit to M ichaelis–Menten kinetics. S-Adenosyl-
homocysteine nucleosidase was purified from an
overproducing strain (E. coli BL21(DE3)/pEXH6MTAN),
generously given by K. Cornell and M. Riscoe (VAMC,
Portland, OR, USA), and assayed as described previously
[28].
pH profile
CFAS activity was determined as described above in the
following buffers: 4-morpolinoethanesulfonic acid ( pH 5.5–
7.0), 2 -(4-(2-hydoxyethyl)-1-piperazine) ethanesulfonic a cid
(pH 7.0–8.5) and 3-(tris(hydroxymethyl)methylmino)-1-
proanesulfonicacid (pH 7.7–9.5) all at a concentration of
150 m
M
. The pH was a djusted by a dding aqueous HCl or
aqueous NaOH. The activity vs. pH p rofile was bell-shaped
and the data points were fitted to Eqn (1):
V ¼ V
max
=ð1 þ 10
pHÀpKa1
þ 10
pKa2ÀpH
Þ Eqn ð1Þ
using a nonlinear regression analysis supported by
KALEIDAGRAPH
software (Synergy Software, Reading, PA,
USA).
Exchange experiments

A sample consisting of 2 lg (45 pmol) CFAS, 2 m
M
dithiotheithol, 0.5 mgÆmL
)1
BSA, 10% (v/v) glycerol,
20 m
M
potassium ph osphate buffer, pH 7.4, and 1 m
M
[methyl-
3
H]S-AdoMet (13 mCiÆmmol
)1
) was incubated at
37 °C for 3 h. A control sample that did not contain the
enzyme was run at the same time. The reaction was stopped
by dilution with 1 mL water and immediate f reezing in
liquid nitrogen. Water was then lyophilized, re covered and
counted for r adioactivity in 4 mL of scintillation liquid. For
the incorporation of deuterium from D
2
O, the experiment
was run directly in the NMR tube (500 lL, total volume).
The sample c onsised of 2 lg (45 pmol) CFAS, 2 m
M
dithiotheithol, 0.5 mgÆmL
)1
BSA, 10% (v/v) glycerol,
20 m
M

potassium phosphate buffer, pD 7.4 (corrected),
and 1 m
M
S-AdoMet. In order to minimize the H
2
O
concentration the buffer was exchanged in D
2
Oand
lyophilized prior t o u se. A control sample that did not
contain the enzyme was run at the same time. The samples
were incubated f or 4 h and were a nalyzed by
1
H-NMR. The
signal at 3.11 p.p.m., w hich corresponds to the methyl
group of the natural diastereoisomer of S-AdoMet [(S,S)
configuration], was quantified and compared to an authen-
ticsampleofcommercialS-AdoMet. The methyl group of
para-toluenesulfonate, present in the commercial sample of
S-AdoMet, was used as an internal standard. No modifi-
cation of the signal was ob served. A
13
C-NMR (
1
H
decoupled) s pectrum was also recorded to see if a ny
exchange on the methyl g roup had occured, becau se
deuterium incorporation would shift the signal and would
give a scalar coupling.
Thiol titration by 5,5¢-dithiobis-(2-nitrobenzoic acid)

(DTNB)
Thiol titrations were run a s described by Riddles et al.[29].
Breifly, for titration i n d enaturing c onditions, 1.9 nmol
(84 lg, 2.4 l
M
final concentration) of purified His
6
-tagged
wild-type CFAS w ere added t o a solution (800 lL final
volume) c ontaining 6.0
M
guanidine hydrochloride,
0.31 m
M
5,5¢-dithiobis-(2-nitrobenzo ic acid) (DTNB),
0.1
M
potassium phosphate, pH 7.3, 1 m
M
EDTA at
20 °C. The exposed thiols were titrated by measuring the
change in absorbance at 412 nm (e ¼ 13 700 cm
)1
Æ
M
)1
).
For titration under n ative conditions the same protocol was
applied except that the guanidine hydrochloride was not
added. Thiols were titrated by measuring the change in

absorbance at 412 nm (e ¼ 14 150 cm
)1
Æ
M
)1
).
Inactivation by DTNB
His
6
-tagged wild-type and mutant CFAS pr oteins were
treated with various concentration of DTNB in 0.1
M
potassium phosphate, pH 7.3, 1 m
M
EDTA, at 20 °C.
Aliquots of 30 lL were transferred, at different time points,
to a CFAS assay mixture (final volume of 100 lL)
containing, 1.0 mgÆmL
)1
phopholipids, 0.5 mgÆmL
)1
BSA,
0.76 m
M
[methyl-
3
H]S-AdoMet at a final specific radio-
activity of 25 mCiÆmmol
)1
,5m

M
reduced glutathione to
quench t he inactivation, in 20 m
M
potassium phosphate
buffer, pH 7.4. The mixture was incu bated at 37 °Cfor
15 min. The reaction was stopped by adding 1 m L 10%
(w/v) trichloroacetic acid and treated as described above
for radioactivity counting.
Protection from inactivation by DTNB
His
6
-tagged wi ld-type C FAS was incubated with 2 m
M
DTNB in presence of 1 mg ÆmL
)1
phospholipids or
380 l
M
S-AdoMet, in 0.1
M
potassium phosphate,
pH 7.3, at 20 °C. Residual activity was measured as
described for the inactivation experiments (see above).
Tsou plot
Two identical samples w ere p repared as f ollows. His
6
-
tagged wild-type CFAS (1.9 nmol; 84 lg, 2.4 l
M

final
concentration) was added to a solution (800 lL final
volume) containing 0.1
M
potassium phosphate, pH 7.3,
1m
M
EDTA, 0.31 m
M
DTNB. Addition o f the enzyme was
performed at the same time in both samples and the
absorbance at 412 nm (e ¼ 14 150 cm
)1
Æ
M
)1
) was followed
against time, using one sample. Th e residual activity was
followed against time using the second samp le. Both
incubations were run at 20 °C. Determination of residual
activity was carried out as described a bove for the DTNB
inactivation experiments. Data obtained at the same time
points, that is the number of titrated thiols and the residual
activity, were used in the graphical representation described
by Tsou [30].
4772 F. Courtois et al.(Eur. J. Biochem. 271) Ó FEBS 2004
Results
Cloning, expression and purification of CFAS
E. coli CFAS was expre ssed as an N-terminal His
6

-tagged
recombinant protein, in order to simplify the purification
protocol [9]. The recombinant gene, containing an engin-
eered ribosome binding site [31] and a His
6
-tag was
constructed using PCR-based recombinant technology
and cloned into a pET-24(+) vector. A C-terminal tagged
protein was also constructed but the protein was expressed
as an insoluble and inactive polypeptide. The N-terminal
His
6
-tagged construct, pET-24H6cfa, whose DNA sequence
was verified, was used th roughout this study. Overexpres-
sion in E. coli BL21 (DE3)/pET-24H6cfa was optimized by
varying the usual parameters, that is IPTG concentration
(from 40 l
M
to 1 m
M
), t emperature ( 20 °C, 30 °Cand
37 °C), and incubation time after induction (from 3 h to
15 h). Our best results were obtained using the following
conditions: 100 l
M
IPTG, 37 °C and overnight incubation.
The His
6
-taggedCFASwaspurifiedintwosteps(Fig.1).
An affinity nickel chromatography was followed by a

necessary desalting s tep by g el filtration because high
imidazole concentration inhibits the enzyme activity. Start-
ing from 0.8 L of culture (100 mg of protein in the crude
extract with a specific activity of 0.9 · 10
)2
UÆmg
)1
), 5 m g
of pure protein (Fig. 1) was obtained with a specific activity
of 5.0 · 10
)2
UÆmg
)1
, a value comparable to previously
reported data [9,24]. The yield of this purification is 28%,
and the purification fold is 5.5. This simple protocol is fast
enough (a few hours in total) to keep this labile enzyme
active. The enzyme was stored best in 20 m
M
phosphate
buffer, pH 7.4 containing 50% (v/v) glycerol, at )20 °C.
Assay and characterization
The recombinant CFAS was assayed as described by
Cronan and coworkers w ith slight modifications [7]. We
found that addition of 0.5 mgÆmL
)1
BSA, 2 m
M
dithiothei-
thol and 10% (v/v) g lycerol substantially stabilized the

enzyme activity during the assay. Typically, after 60 min
incubation the activity of a sample containing the additives
was twice over that of a c ontrol sample. Addition of
S-AdoHcy nucleosidase as suggested previously [7] to
hydrolyze the product, a competitive inhibitor [6,13], was
not necessary in our assay b ecause the concentration of
S-AdoHcy reached was too low to cause inhibition. The
effect of ph ospholipid concentration was also checked and
we found a biphasic curve as already observed, with a
saturation at 1 mgÆmL
)1
phospholipid [6]. Using this assay
we measured a K
m
of 90 ± 5 l
M
for S-AdoMet and a k
cat
of 2.2 ± 0.1 min
)1
, values in close agreement t o t hose
reported for the native enzyme [7]. Therefore, the presence
of the His
6
-tag does not perturb the catalytic activity.
N-terminal sequencing showed no contaminants and was
in agreement with the predicted seq uence. The UV-visible
spectrum of the protein did not show any absorption over
300 nm and thus no organic cofactor could be detected.
Search for usual metals found in proteins (Zn, Ni, Cu, Co,

Fe) was unsuccessful. We therefore concluded that CFAS
has no cofactor, a result in agreement with the three
dimensional structure obtained for the M. tuberculosis
cyclopropane synthases [22].
The effect of pH on the activity of CFAS, under saturating
conditions, is shown in Fig. 2. The profile is b ell-shaped with
a maximum around pH 7.5. Fitting the data to a simple
model using Eqn (1) (with two ionisable groups involved in
catalysis) gave a pK
a1
of 6.8 and a pK
a2
of 8.7.
Exchange experiments
Exchange of the methyl p roton of S-AdoMet catalyzed by
CFAS was tested by measuring the wash-out of tritium
from the methyl group to the solvent water. Incubation of
1
10
100
5678910
V (mU/mg)
pH
pK
a1
=6.8
pK
a2
=8.7
Fig. 2. pH profile for CFAS activity. His

6
-tagged wild-type CFAS
activity was measured at different pH values, using a series of buffers
(see Experiental procedures). Each data point represents the average of
two independent experiments with less than 5% deviation to the mean.
The data points were fitted to Eqn (1), for estimation of the two pK
a
.
Ordinates are plotted on a log scale.
Fig. 1. SDS/PAGE analysis of the purification of His
6
-tagged wild-type
and E. coli CFAS mutants. From left to right: lane 1, C139S; lane 2 ,
C176S; lane 3, C354S; lane 4, wild-typeCFAS;lane5,molecularmass
markers (from top to bottom, 66 kDa, 45 kDa, 36 kDa, 29 kDa,
24 k Da, 20.1 kDa).
Ó FEBS 2004 Cyclopropane synthase reaction mechanism (Eur. J. Biochem. 271) 4773
the enzyme in the presence of [methyl-
3
H]S-AdoMet but
without unsaturated phospholipids, for 3 h gave no more
counts in the water fraction than a control sample
containing no enzyme. The detection limit of this experi-
ment was e stimated at 0.2% exchange (i.e. that an exchange
of 0.2% or more would have been easily detected). Addition
of cyclopropanated phospholipids in the reaction mixture
that could trigger a conformational change upon binding
did not enhance this exchange reaction. The reverse
experiment, incorporation o f solvent protons into the
substrate, which should be faster than the wash-out as no

intramolecular kinetic isotope effect on th e abstraction
should occur, was tested using the same conditions but in
the presence of unlabeled S-AdoMet and deuteriated buffer.
The reaction was followed by
1
Hand
13
C-NMR, and again
no incorporation of deuterium could be detected. Therefore,
under our conditions, CFAS is unable t o c atalyze the
exchange of the methyl protons of S-AdoMet, a result that
does not support the ylide mechanism.
Thiol titration by DTNB
The amino acid sequence of wild-type E. coli CFAS
predicts eight cysteines [9]. The total thiol content of the
purified enzyme was spectrophotometrically titrated using
DTNB, in denaturing conditions using standard protocols
[29]. A ratio of 7 .5 ± 0.3 mol of free thiols p er mol of
CFAS monomer was f ound, consistent with eight free
cysteines in the E. coli wild-type CFAS. In native condi-
tions (Fig. 3), six thiols per monomer were titrated in one
hour with triphasic kinetics. Three cysteines reacted within
4 min, two more cysteines reacted more slowly within
40min,andonecysteinereactedinathirdveryslowphase.
If the enzyme was left longer under these conditions (for
two further hours), the absorbance at 412 nm finally
reached a value compatible with eight free cysteines.
Therefore E. coli CFAS contains three classes of free
cysteines, three fast reacting thiols (exposed), two slowly
reacting thiols (less accessible) and three buried cysteines

that react extremely slowly. The upward curvature of the
trace in Fig. 3, after 40 min (a reproducible phenomenon),
is probably due to a partial unfolding o f the protein,
exposing the buried cysteines, which consequently react
faster. It i s not clear i f the protein unfolds because of
multiple chemical modifications or if it is simply due to the
long incubation time.
CFAS inactivation by DTNB
As already reported by Cronan and coworkers [9], we found
that CFAS could be inactivated by thio l-directed reagents
such as DTNB and N-ethylmaleimide (NEM). Kinetic
analysis of the inactivation process by DTNB is shown i n
Fig. 4. The inactivation follows a pseudo first-order kinetics
with no saturation and w ith a second-order r ate constant of
1.2
M
)1
Æs
)1
, a low but not unprecedented value [32]. Similar
analysis using NEM showed that the inactivation occurred
similarly with a rate constant of 2.4
M
)1
Æs
)1
(not shown).
The inactivation process was not significantly slowed down
in the presence of a saturating concentration of S-AdoMet
(0.38 m

M
) o r i n the presence of 1 mgÆmL
)1
unsaturated
phospholipids ( Table 1). This suggests t hat the cys teine
residue responsible for the inactivation is not located in the
active site.
For the graphical analysis of t he inactivation, His
6
-tagged
wild-type CFAS (2.4 l
M
) was treated with excess DTNB
(0.31 m
M
) at pH 7.3 and the residual activity together with
the number of modified sulfhydryls per CFAS were
determined at the same time points. The data were analyzed
graphically as described by Tsou (Fig. 5) using t he following
Eqn (2) [30]:
(a)
1=i
¼ðp þ s À mÞ=p ð2Þ
where m is the number of modified cysteines per monomeric
CFAS, s t he number o f fast r eacting cysteines that are
nonessential, a the fraction of remaining activity when m
residues have been modified, p is the number of cysteines
reacting slower than the s group, and i is the number of
essential r esidues for catalytic a ctivity, as defined by Tsou.
Note that the i class belongs to the p class. The graph shown

in Fig. 5 c onfirms the presence o f three classes of free
cysteines in the enzyme. First, three cysteines react quickly,
with no loss of activiy, then two more cysteines react with
concomitant loss of e nzyme activity, and finally the buried
cysteines react. The portion of the graph where the activity
is lost perfectly fits to a straight line when i ¼ 1(the
correlation coefficient i s 0 .99). Wh en the same data are
plotted with i ¼ 2ori¼ 3 the data points clearly deviate
from linearity. Therefore the data of Fig. 5 are most
consistent with one cysteine, chemical modifi cation of which
leads to inactivation. Futhermore the plot a llows the
estimation of s and p, as the o rdinate in tercept is (p+s)/
p ¼ 2.1, the abscissa intercept is p+s ¼ 5 and the slope is
)1/p ¼ –0.44. Thus p ¼ 2.3 and s ¼ 2.7, values in close
agreement with the numbers deduced from Fig. 3 (where
p ¼ 2ands¼ 3).
0
1
2
3
4
5
6
7
0 102030405060
Number of reacting thiol
per monomer (mol/mol)
Time (min)
Fig. 3. Kinetics of DTNB titration of E. coli His
6

-tagged wild-type
CFAS in nondenaturing conditions. CFAS (2.4 l
M
) was titrated with
0.31 m
M
DTNB at 2 0 °Cin0.1
M
potassium phosphate bu ffer,
pH 7.3, containing 1 m
M
EDTA. The absorbance at 412 nm was
recorded against time.
4774 F. Courtois et al.(Eur. J. Biochem. 271) Ó FEBS 2004
Characterization of C139S, C176S and C354S mutant
proteins
Alignment o f the sequence of a ll cyclopropane syntha-
ses known s o far shows t hat among the e ight cysteine
residues of the E. coli CFAS only three are strictly
conserved: C139, C176 and C 354 [1]. W e thus prepared
three corresponding Cys fi Ser mutants for analysis. This
particular re placement was chosen because it is isosteric, but
the O H group is much less acidic and much less nucleophilic
0.00
0.05
0.10
0.15
0.0 0.5 1.0 1.5 2.0 2.5
k
obs

(min
-1
)
[DTNB] (mM)
10
100
0 2 4 6 8 10 12 14 16
Residual activity (%)
Time (min)
Fig. 4. Inactivation of His
6
-tagged wild-type E. coli CFAS by DTNB.
Top: His
6
-tagged wild-type CFAS was incubated in the presence of
DTNB, 0 m
M
(d), 0.5 m
M
(h), 1 m
M
(r), 1.5 m
M
(s), 2 m
M
(j), at
20 °Cin0.1
M
potassium phosphate buffer, pH 7.3, containing 1 m
M

EDTA. Aliquots were withdrawn at different time points and the
residual activity was m easured (se e Experim antal proced ures for
details). Each data point represents the average of two independant
experiments, with less than 5% deviation from the mean. Error bars
are omitted for clarity. The observed first-order rate constants, k
obs
,
were calculated by fitting the data points to simple exponential decays.
Bottom: observed rate con stants, k
obs
, corrected for the slow inacti-
vation in the absence of DTNB, were plotted aga inst DTNB concen -
tration. Data were fitted to a straight line.
Table 1. Summary of the inactivation r ate constant in the presence of
DTNB. All experiments were run under the following conditions:
2m
M
DTNB in 0.1
M
potassium p hosphate buffer, pH 7.3, 1 m
M
EDTA at 20 °C. A protectant was sometimes added as indicated.
Enzyme Protectant k
obs
(min
)1
)
His
6
-tagged

wild-type CFAS
No protection 0.13
0.38 m
M
S-AdoMet 0.10
1mgÆmL
)1
Phospholipids 0.15
C139S CFAS No protection 0.14
C176S CFAS No protection 0.11
C354S CFAS No protection 0.12
0.0
0.2
0.4
0.6
0.8
1.0
1.2
1.4
012345678
(Fraction of resudual activity)
1/i
Number of titrated thiol per monomer
Fig. 5. Tsou plot for the inactivation of His
6
-tagged wild-type CFAS by
DTNB. CFAS (1.9 nmol, 2.4 l
M
) was incubat ed with 0.31 m
M

DTNB
in 0.1
M
potassium buffer, pH 7.3, 1 m
M
EDTA, a t 20 °C f or 60 min.
The absorbance at 412 nm (e ¼ 14 150 cm
)1
Æ
M
)1
) and the activity
were monitored at the same time (see Experimental procedures). The
fraction of residual activity, a
1/i
,i¼ 1(d), i ¼ 2(s), i ¼ 3(n)was
plotted against the n umber of modified thiols. The data for i ¼ 1were
fitted to a straight line. Each data point represents the average of tw o
independent experiments. Error bars are not shown for clarity. See text
for details of the analysis.
Table 2. Kinetic parameters for His
6
-tagged wild-type and muta nt
CFAS.
Enzyme
K
m
for
S-AdoMet
(l

M
)
k
cat
(min
)1
)
k
cat
/K
m
(min
)1
Æm
M
)1
)
Relative catalytic
efficiency (%)
Wild type 90 2.2 24.8 100
C139S 88 0.3 3.9 16
C176S 73 2.7 37.9 150
C354S 105 1.6 15.7 63
Ó FEBS 2004 Cyclopropane synthase reaction mechanism (Eur. J. Biochem. 271) 4775
than the t hiol group. All mutant genes were obtained by
PCR amplification using two sets of mutated primers, and
the Stratagene technology. The desired mutations were
verified by DNA sequencing and the mutated proteins were
expressed and purified as described for the His
6

-tagged wild-
type enzyme (Fig. 1). Table 2 summarizes the kinetic
parameters for individual mutants and His
6
-tagged wild-
type enzyme. It is clear that all m utant are active, the slowest
being C139S (16% relative catalytic efficiency). Therefore
none of the conserved cysteines is essential for the activity.
The three mutants w ere inactivated by DTNB at a rate
similar to that m easured for the H is
6
-tagged wild-type
enzyme under the same conditions (Table 1). Thus, the
cysteine residue that is responsible for the inactivation is not
one of the conserved cysteines, a result compatible with the
fact that no protection was observed by the substrates.
Discussion
The cyclopropyl group is present in number of natural
products and its unusual properties have always stimulated
chemists and biochemists [33,34]. Biosynthesis of this
structural element follows diverse schemes, but the direct
methylenation of double bonds, catalyzed by cyclopropane
synthases, is one of the most interesting. Most intriguing is
the c hemical m echanism by which this class o f c losely
related e nzymes effects the cyclopropanation. The carboca-
tion mechanism, first described by Lederer [20], is chemic-
ally sound and can also be applied to other methyl
transferases found in mycobacteria that are homologous
with cyclopropane synthases, and which catalyze modifica-
tions of unsaturated lipids, such as the formation of

a-methylketo- or a-methylhydoxy- fatty acids [35,36].
However, it quickly appeared that addition of a s ulfur
ylide, derived from S-AdoMet, to the double bond of the
fatty acid could b e another plausible alternate r eaction
mechanism [14,17,18,21]. The two mechanisms differ from
one another not only in the order of making and breaking
bonds, but also in the type of intermediate formed. Progre ss
has recently been achieved with cloning and purification
of the E. coli enzyme [9], as well as solving the three
dimensional structure of M. tuberculosis enzymes [22], and
reports of some mechanistic experiments [23,24].
We report here the purification and characterization of a
recombinant CFAS bearing an N-terminal His
6
-tag. The
use of nickel affinity chromatography allowed rapid
preparation o f pure enzyme in s ubstantial amounts. A
similar successful strategy was recently followed by Liu and
coworkers [24].
As the ylide mechansim would be most likely to i nvolve
carbenoid transfer to a metal [21], we searc hed for metals in
the enzyme. No cofactors, organic or metallic, were found, in
accordance with structural data obtained for the M. tuber-
culosis enzymes(forwhichnoin v itro catalytic a ctivity has
ever been reported). The reaction mechanism mu st therefore
rely solely on side chain functional groups, and thus only
acid-base or nucleophilic catalysis must operate.
The pH profile of the activity, in saturating conditions,
revealed two ionisable g roups important for catalysis: a first
pK

a1
of 6.8 and a second pK
a2
of 8.7. Interpretation of pH
effects are most difficult, but it is interesting to note that a
carbonate (pK
a
¼ 6.4), bound in the a ctive s ite of the
M. tu berculosis enzymes, has been suggested to be the base
necessary to abstract the methyl proton [22]. It is then
tempting to attribute the pK
a1
¼ 6.8, detected by kinetic
means to the carbonate, that was proposed to be the base on
structural grounds. Alternatively, this pK
a
could b e attrib-
uted to a His r esidue, such as His266 (His 167 in the
M. tu berculosis sequence), which lies in the active site and
could participate in a proton relay. Futh er mutagenetic
experiments are in progress to clarify this point.
One of the best ways to prove the ylide mechanism would
be to show that the enzyme is able to catalyze the exchange
of the methyl protons of S-AdoMet in the absence of other
substrates. Numbers of enzymatic r eaction mechanisms
have been supported on this type of experimental grounds
[37]. However, under our conditions we did not observe
such an exchange, even in the presence of the cycloprop-
anated product that could mimic the second substrate and
hence trigger a conformational change, a strategy that was

successful in the citrate synthase case [37]. Therefore, our
data do not support the ylide mechanism. Of course, one
cannot exclude the possibility that the abstraction is
promoted by a monoprotic base that exchanges its proton
with the s olvent very slowly. However, Buist and coworkers
reported feeding experiments, using deuteriated methionine
and L. plantarum cells, which were then interpreted by
invoking an exchange of the methyl protons on the
carbocation intermediate ( Scheme 2A) but not on an ylide
species [17,18]. Such a fast exchange (33% exchange) would
probably require a polyprotic base and a reversible forma-
tion of th e cyclopropane ring. However, these experiments
were conducted using whole cells and were dependent on
growth conditions, and thus need to be confirmed on the
isolated enzyme.
We have also addressed the role, in c atalysis, of the
cysteines of the E. coli enzyme. It has been suggested in the
literature that a cysteine could b e important for catalysis [9].
Indeed, a thiolate could e ither abstract a proton on the
methyl group or stabilize the carbocation, or even form a
covalent adduct (the base or the nucleophile in Scheme 2).
The alignment of the seq uence of all cyclopropane synthases
known so far [1], shows that only three cysteines among the
eight cysteine residues of the E. coli enzyme are conserved:
C139, C176 and C354. In the three dimensional structure of
the homologous M. tuberculosis enzymes [22], C35 which
corresponds to C139 of E. col i CFAS shares a hydrogen
bond, by its N-H, to a carbonate in the active site. C269 of
the M. tuberculosis enzyme, which corresponds to C354 in
the E. coli enzyme, was found to be located near the active

site. The third conserved c ysteine, C72 in the M. tuberculosis
enzyme, corresponding to C176 in the E. coli enzyme, is
located near the S-AdoMet binding site but far from the
active site. T herefore, it was reasonable to postulate an
important role in catalysis for these conserved residues.
Cronan and coworkers reported very briefly in a re view [1]
that mutation of Cys176 and Cys354 to Ala, in the E. coli
enzyme, gave active mutants. No experimental data were
reported and the third conserved cysteine, Cys139, was not
mutated. We thus decided to re-examine this issue in detail.
Our results show that, in the E. coli enzyme, all of the eight
cysteines are free and that they can be classified into three
classes: three fast reacting cysteines, two cysteines re acting at
a moderate rate, and three cysteines reacting very slowly.
4776 F. Courtois et al.(Eur. J. Biochem. 271) Ó FEBS 2004
Futhermore, using the graphical analysis developed by Tsou
[30], we s how here that only one cysteine of the t wo
moderately reacting residues, is responsible for the inactiva-
tion of CFAS by DTNB. Thus, five cysteines react with
thiol-directed reagent in 40 min under our conditions, but
only one modification leads to inactivation. The three
mutated enzymes, C139S, C176S and C354S, w ere p repared
by site-directed mutagenesis, and were found to be active, the
slowest (16% active) being C139S, primarily affected on its
catalytic constant. If any of these cysteines were invo lved in
base catalysis or nucleophilic catalysis, the corresponding
serine mutant should have been at least a hundred to a
thousand times less efficient than the wild-type enzyme. A
dramatic drop in activity is usually observed in CysfiSer
mutants of enzymes known to use the thiolate group as a

base (racemases) or as a nucleophile (methyl transferases)
[38–40]. The fact that inactivation by DTNB of the His
6
-
tagged wild-type enzyme was not protected by substrates,
and that the three C ysfiSer mutants prepared in this report
are inactivated by DTNB at the same rate as the His
6
-tagged
wild-type enzyme, shows that the cysteine responsible for the
inactivation cannot be C139, C176 or C354. There are five
other cysteine residues in the E. coli enzyme, and it is not
possible at t he moment to attribute the residue t hat is
responsible for the inactivation. Furthermore, this chemical
inactivation does not seem to be related to catalysis.
In conclusion, the findings reported here do not support
the ylide m echanistic proposal but further s upport the
carbocation mechanism. Furthermore, it is shown here that
the conserved cysteines of E. coli CFAS are not directly
involved in catalysis and that the inactivation observed after
chemical modification o f another cysteine probably comes
from steric hindrance, that is no t relevent to catalysis. The
base and the nu cleophile supposedly involved in the r eaction
mechanism are very likely other residues o r f unctional
groups, e.g. E239 or the active site carbonate. F uther
mutagenesis experiments are underway to explore these
hypotheses.
Acknowledgements
We wish to thank Thierry Drujon and Diane Delaroche for technical
assistance in preparing E. coli pho pholipids a nd in constructing

mutants of E. coli cyclop ropane fatty acid s ynthase. We are grateful
to Sabin e Cornet for initial work on cloning the cfa ge ne. We are
indebted to Dr Kenneth A. Cornell and Dr Michael Riscoe for the
generous gift of plasmid p EXH6MTAN and for advice in preparing
S-AdoHcy nucleosidase. This work was supported in part by the ACI
program of the ÔMiniste
`
re de lÕEducation Nationale de la Recherche et
de la Technologie’, grant number 0693.
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