Hemisynthesis of deuteriated adenosylhopane and conversion into bacteriohopanetetrol by a cell-free system from Methylobacterium organophilum

Article abstract of DOI:10.1039/c4ob02560a

Adenosylhopane is a putative precursor of the widespread bacterial C₃₅ biohopanoids. A concise and flexible hemisynthesis of adenosylhopane has been developed including as key steps a cross metathesis between two olefins containing either the h

Full text of DOI:10.1039/c4ob02560a

Organic &
Biomolecular Chemistry
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Hemisynthesis of deuteriated adenosylhopane
and conversion into bacteriohopanetetrol by
a cell-free system from Methylobacterium
organophilum†
Cite this: Org. Biomol. Chem., 2015,
13, 3393
Wenjun Liu,a Anne Bodlennerb and Michel Rohmer*a
Adenosylhopane is a putative precursor of the widespread bacterial C35 biohopanoids. A concise and
flexible hemisynthesis of adenosylhopane has been developed including as key steps a cross metathesis
between two olefins containing either the hopane moiety or a protected adenosine derivative and a sub-
sequent diimide reduction of the resulting olefin. Reduction by deuteriated diimide allowed deuterium
labelling. This synthetic protocol represents a versatile tool to access to deuteriated composite bacterial
hopanoids required for biosynthetic studies. Deuteriated adenosylhopane was thus converted into
bacteriohopanetetrol by a crude cell-free system from Methylobacterium organophilum in the presence
of NADPH, showing for the first time the precursor to product relationship between these two bacterial
metabolites.
Received 8th December 2014,
Accepted 28th January 2015
DOI: 10.1039/c4ob02560a
www.rsc.org/obc
atom to the hopane moiety.6 The same configuration was also
found for 35-aminobacteriohopanetriol 6 via chemical corre-
Introduction
(22R)-Adenosylhopane 2 (Scheme 1) has been found in fairly lation with bacteriohopanetetrol.7 This side-chain configur-
large amounts in a few species of bacteria1a–e and in trace ation is consistent with the hypothesis deduced from the
amounts in many hopanoid producing bacteria.2 Adenosyl- labelling experiments: the additional C5 unit of elongated
hopane (and structurally tentatively related hopanoids) is one hopanoids originates from a D-ribose derivative.8,9 Adenosyl-
of the most common hopanoids in soils,3a–c and the presence hopane was consequently considered as a possible intermediate
of this biomarker in recent sediments is interpreted in terms in the biosynthesis of C35 hopanoids, leading to bacteriohopa-
of transport of soils into sediments.3b,4,5
netetrol 5 or aminobacteriohopanetriol 6 (Scheme 1).8–10 This
Adenosylhopane 2 is characterized by a unique carbon– hypothesis has been corroborated by the identification of the
carbon bond between C-30 of the hopane moiety and C-5′ of hpnG gene in Methylobacterium extorquens11 and Rhodopseudo-
adenosine. The commonly occurring configuration of adeno- monas palustris12 and the orf14 gene in Streptomyces coelico-
sylhopane was determined by circular dichroism and high- lor:13 their deletion results in the accumulation of
field NMR spectroscopy.1a Trace amounts of (22S)-adenosyl- adenosylhopane 2 and in the absence of the major hopanoids,
hopane as the minor isomer was also reported from Rhodopseu- bacteriohopanetetrol 5 or aminobacteriohopanetriol 6.
domonas acidophila.1b Chemical conversion of adenosylhopane
The elucidation of the biosynthesis of the side-chain of C35
2 into bacteriohopanetetrol 5 permitted the determination of hopanoids requires sufficient amounts of adenosylhopane for
the stereochemistry of all asymmetric centres of the side-chain future enzyme tests. Two distinct methods are available to
of bacteriohopanetetrol as 22R, 32R, 33R and 34S.1b Later the produce complex hopanoids: fermentation or chemical syn-
synthesis of the eight side chain diastereomers of bacterio- thesis. The amphiphilic character and the poor solubility in
hopanetetrol confirmed that the absolute configuration of the C5 most organic solvents of biohopanoids represent severe limit-
unit was identical to that of a D-ribitol linked via its C-5 carbon ations for their production by fermentation. Moreover, fermen-
tation is only appropriate for a few major compounds as many
complex hopanoids are produced by bacteria in only trace
amounts. In addition, isolation is until now only performed
on acetylated derivatives requiring a final deprotection to get
the native free metabolites, which is either cumbersome or
aUniversité de Strasbourg/CNRS, Institut Le Bel, 4 rue Blaise Pascal, F 67070
Strasbourg, Cedex, France. E-mail: mirohmer@unistra.fr
bUniversité de Strasbourg/CNRS, ECPM, 25 rue Becquerel, F 67087 Strasbourg,
Cedex 2, France
simply not available (e.g. for adenosylhopane or bacterio-
†Electronic supplementary information (ESI) available. See DOI: 10.1039/
hopanetetrol derivatives with carbamoyl groups). Chemical
c4ob02560a
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Scheme 1 Biogenetic scheme for the biosynthesis of C35 bacterial hopanoids. Identified genes in Methylobacterium extorquens, Rhodopseudomo-
nas palustris and Streptomyces coelicolor: (i) hpnH/orf15; (ii) hpnG/orf14; hpnO/orf18.10–13
synthesis, in contrast, allows an access to a broad range of required for the generation of the non-stabilized phosphor-
naturally occurring hopanoids in sufficient amounts. Extensive ane.18 Even the coupling strategy involving lithium (the smal-
work resulted in suitable syntheses of ribosylhopane, bacterio- lest cation) activated hopane moiety, which was very efficient
hopanetetrol and several other related structures.6,14–17 Some in ribosylhopane 3 synthesis, was disappointing to afford the
synthetic strategies are quite successful with excellent stereo- bulkier 35-O-benzyl ribosylhopane analogue.17 The second
chemistry control and good overall yields.
strategy, which involves a glycosidic coupling between a ribo-
No reliable synthesis is, however, available for adenosyl- sylhopane derivative and adenine in the presence of a Lewis
hopane. The initial attempts towards the synthesis of adenosyl- acid was also rather unsuccessful, not only because of the poor
hopane and related hopanoids were mainly based on two routes. solubility of adenine and the ribosylhopane derivative in aceto-
The first strategy, inspired from the hypothetical biogenetic nitrile, which promoted side reactions, but also because of the
biosynthetic pathway, relies on a coupling between a C30 triter- formation of both 35α and 35β isomers of adenosylhopane 2.14
pene derivative and an adenosine derivative. Wittig-type coup- Furthermore, the application of this methodology is also
ling between
a
triterpene phosphonium salt with an limited considering the number of synthetic steps.
This paper describes an efficient hemisynthesis of adeno-
adenosine-5′-aldehyde substrate led to the formation of adeno-
sylhopane 2 in a very low yield,14 probably due to the steric sylhopane including the possibility of deuterium labelling as
hindrance induced by both the adenosine moiety and the well as the conversion of deuterium labelled adenosylhopane
pentacyclic triterpene skeleton in addition to the instability of into bacteriohopanetetrol by a crude cell-free system form
the nucleoside-5′-aldehyde in the presence of the strong bases Methylobacterium organophilum, which represents the first
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direct and indisputable proof for a precursor to product different enough,21a initial attempts of cross metathesis were
relationship between adenosylhopane
panetetrol 5.
2
and bacterioho- performed with stoichiometric amounts. Reaction with alkene
8 and one equivalent of homohopene 10 in the presence of
Hoveyda–Grubbs catalyst (HII), did not, however, give the cross
coupling product 11 under reflux conditions (Table 1, entry 1).
This lack of reactivity for those two terminal alkenes probably
resulted from the high steric hindrance in the expected metalla-
cyclobutanyl transition state, together with an activation
Results and discussion
Hemisynthesis of adenosylhopane
The proposed strategy leading to adenosylhopane 2 is a conver-
gent hemisynthesis with an excellent stereochemical control
using commercially available starting materials. Moreover, in
order to meet the need for isotopic labelled hopanoids for bio-
synthetic investigations, this strategy allows also the introduc-
tion of deuterium into the hopanoid structure in the last but
one step. Our synthetic strategy relies on a cross-metathesis as
the key coupling step between a methylene elongated hopane
moiety 10 and a properly protected methylene elongated
5′-deoxy-5′-methyleneadenosine derivative 8 (Scheme 2). This
approach has the advantage of preserving the stereogenic
centres at C-22 on the hopane moiety and at C-4′ of the ribose
derivative and avoids the inconvenient glycosylation between
adenine and ribose,14 which must be regio- and diastereoselec-
tive. Metathesis19 seemed quite attractive because of its high
tolerance towards functional groups and bulky structures,20a,b
its mildness and impressive efficiency, and the possibility of
late labelling by deuteriation of the double bond. Cross meta-
thesis is known to be more difficult than dimerization, ring
closing or ring opening metathesis due to the lost advantages
from statistics, entropy and ring tension relief respectively. The
trick is to use alkenes having strong reactivity differences or an
excess of one counterpart.21a,b It was, however, encouraging to
note that Andrei and Wnuk succeeded in coupling a protected
5′-deoxy-5′-methyleneadenosine with N-Boc-protected six-
carbon amino acids bearing a terminal double bond,22 even if
the methylenetriterpene 10 (Scheme 2) in our synthesis pre-
sents a steric hindrance closer to the terminal double bond
than their cross-metathesis partner. Furthermore, both olefins
8 and 10 have the nice reactivity of terminal monosubstituted
olefins21 to counteract the steric hindrance present one bond
away from the olefin.
energy barrier too high to be overcome under classical thermic
conditions. Coupling product 11 was, however, isolated with
an up to 13% yield when the same reaction was performed
under microwave irradiation (Table 1, entry 2). Dimer 15
(Scheme 2) was also isolated in 43% yield, resulting from the
self-metathesis of the nucleoside substrate 8. Unreacted start-
ing materials 8 and 10, were also isolated (Table 1, entry 2).
The product of self-metathesis of homohop-30-ene 10 was not
detected. Ruthenium catalyst Grubbs II (GII) promoted cross
metathesis between 8 and 10 in a lower yield, (Table 1, entry
4), suggesting a higher efficiency of HII over GII with those
olefins. Finally, the N,N-dimethylaminosulfonyl group contain-
ing Hoveyda–Grubbs-type complex catalyst Zhan-1B, was found
almost as active as HII for this cross metathesis reaction, and
similar yields were obtained for the coupling product 11 and
dimer 15 (Table 1, entries 2 and 3).
Attempts to obtain olefin 11 from secondary cross meta-
thesis between homohopene 10 and dimer 15 under micro-
wave irradiation at 75 °C did not lead to the formation of the
protected adenosylhopene 11, but allowed the recovery of the
two starting reagents. The inability of dimer 15 to undergo sec-
ondary cross metathesis in addition to its formation from 8
under our conditions but its absence in the cross metathesis
performed by Andrei and Wnuk with a type I olefin as cross
metathesis partner,22 suggest that it belongs to type II olefins
for cross metathesis reactions in Grubbs categorization of alke-
nes.21a Besides, the absence of the product of self-metathesis
of homohop-30-ene 10 suggests that it is a type III olefin in
spite of its terminal alkene.
Reacting two cross metathesis partners of different types
using feedstock stoichiometries as low as 1 : 1 normally allows
selective cross metathesis, but in our case the yield remained
low. Given these features, the crucial point to obtain higher
yield for coupling compound 11 was to maintain a low concen-
tration of adenosine derivative 8 compared to that of homo-
hopene 10, minimizing thus the amount of dimerization. A large
excess (10 eq.) of the less reactive homohopene 10 was thus
introduced, and, as expected, the yield of protected adenosyl-
hopene 11 rose significantly from 13% to 59% in the presence
of Zhan-1B catalyst under microwave irradiation (Table 1, entry
5). However, the cross metathesis yield still remained moder-
ate, probably due to the low reactivity of homohopene 10.
Increasing more the stoichiometry ratio was not possible for
solubility reasons.
Protected 5′-deoxy-5′-methyleneadenosine 8 (Scheme 2) was
obtained by a Wittig methylene elongation of aldehyde 7,
obtained in five steps from commercially available 2′,3′-O-iso-
propylideneadenosine by an adaptation of two published
methods (cf. ESI, Scheme S1†).23a,b In parallel, homohop-30-
ene 10 (Scheme 2), the triterpenic coupling partner, was syn-
thesized in high yield via a Wittig reaction from (22S)-hopan-
29-al 9, synthesized in four steps from hydroxyhopanone
extracted from Dammar resin,15,24 and presenting the C-22
configuration of most bacterial hopanoids.25
Cross metathesis
Cross metathesis reactions between homohop-30-ene 10 and
Under these conditions, the olefinic adenosine substrate 8
the 5′-methylene adenosine derivative 8 were tested under was fully consumed, resulting in the formation of 40% of
different conditions (Scheme 2 and Table 1). Given the high dimer 15. A prolonged reaction time failed to further increase
value of both olefins, and hoping that their reactivity are the yield in 11, supporting to a greater extent that dimer 15
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Scheme 2 Hemisynthesis of adenosylhopane 2. (i) Ph3PCH3Br, n-BuLi, DMSO–THF, 50 °C, 98%; (ii) Ph3PCH3Br, n-BuLi, THF, −40 °C, 75%; (iii) meta-
thesis; (iv) NH4OH, H2O, MeOH, CH2Cl2, 4 °C, 91%; (v) PADA, AcOH, Py, 90 °C, 73%; (vi) TFA, CHCl3–CH3OH, 0 °C, 92%; (vii) Ac2O, Py.
was indeed unable to undergo a secondary hetero-metathesis hindrance induced by the hopane ring system and the pseudo-
even with homohop-30-ene 10 in excess. The inability of com- axial position of the side chain, allowed the recycling of the
pound 10 to homodimerize, probably due to a strong steric large excess of olefin 10.
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Table 1 Cross-metathesis optimisation for the coupling of 10 and 8
Catalysta
Conditions
10/8 ratio
Products
1
2
3
4
5
6
7
8
H II
H II
Zhan-1B
G II
Zhan-1B
H II
40 °C, DCM, 24 hb
1/1
1/1
1/1
1/1
10/1
10/1
10/1
1/1
No reaction
MWc, 75 °C, DCM, 3 h
11 (13%), 15 (43%)
11 (12%), 15 (45%)
11 (6%), 15d
11 (59%), 15 (40%)
11 (51%), 15 (46%)
11 (52%), 15 (43%)
11 (6%), 15 (78%)
MW, 75 °C, DCM, 3 h
MW, 75 °C, DCM, 3 h
MW, 75 °C, DCM, 3 h
in sealed tubes, DCM, overnight
in sealed tubes, DCM, overnight
MW, 75 °C, perfluorobenzene, 3 hb
Zhan-1B
H II
a 10 mol% of catalyst were used. b Reaction under reflux at ambient pressure. c MW is abbreviation of microwave irradiation. d Yield was not
determined for 15.
The same reactions were performed in sealed tubes without labelled version, the acid promoted decarboxylation of potass-
microwave irradiation (Table 1, entries 6 and 7). A night long ium azodicarboxylate was chosen to generate diimide.31 Fur-
reaction time was required to reach a slightly lower cross meta- thermore, the acid catalysis may speed up the equilibration of
thesis yield, indicating that the highly beneficial effects of trans- and cis-diimide, favouring thus the hydrogen transfer
microwave irradiation did not only arise from the rapid from the cis-isomer to the alkene. Direct treatment of the
heating and pressure increase allowed in the microwave oven N-protected adenosylhopene 11 with potassium azodicarboxylate
(purely thermal/kinetic effect), but also from some specific and acetic acid, however, led to the partial loss of the 6′-N-
thermal microwave effect, such as wall effect and selective benzoyl group. This protecting group was therefore removed
heating of strongly microwave-absorbing heterogeneous cata- before diimide reduction using
a saturated methanolic
lysts in a less polar reaction medium. ammonia solution in dichloromethane to afford the N-depro-
Furthermore, a polar solvent (e.g. dichloromethane) is tected adenosylhopene 12. Diimide reduction of its double
required to enhance the heating effect of microwave. Although bond was then achieved by a continuous addition of potass-
perfluorobenzene was reported to be capable of increasing the ium azodicarboxylate and acetic acid for 36 h and afforded the
activity of cross metathesis catalysts,26 its apolar character expected protected adenosylhopane 13 in 73% yield. The reac-
probably led to less efficient heating by microwave, resulting tion required a large excess of diimide and a long reaction
in a lower yield of 11 in comparison to the one obtained with time, probably because of the steric hindrance around the
the use of dichloromethane (Table 1, entries 2 and 8).
double bond, resulting in a low reduction rate in comparison
As expected, the cross-metathesis product 11 presented pre- to the one of the disproportionation of diimide into nitrogen
dominantly the E configuration as shown by the larger vicinal and hydrazine, the major competing reaction consuming the
J30,31 coupling constant of the vinylic protons in the spectrum reducing agent. In order to minimize this disproportionation,
of the major E isomer (15.4 Hz) versus the smaller coupling potassium azodicarboxylate was added to the reaction mixture
constant in the spectrum of the minor Z isomer (11.0 Hz). in small portions. Acetic acid was added only when the nitro-
Based on the integration of the 32-H and 33-H signals of both gen evolution had ceased, keeping thus a low diimide concen-
protected (E)- and (Z)-adenosylhop-32-ene 11 in the 1H-NMR tration. Water was also avoided because it had been reported
spectrum, i.e. (E)-32-H at 4.74 ppm and (Z)-32-H at 5.03 ppm to be a powerful inhibitor of diimide reductions in aprotic sol-
and (E)-33-H at 4.96 ppm and (Z)-33-H at 4.85 ppm, the relative vents such as pyridine.31 Finally, acid catalysed deprotection of
amount of E isomer varied from 75% to 80%.
the acetonide of 13 with a methanolic TFA solution afforded
Reduction of the double bond of the protected adenosyl- adenosylhopane 2 in high yield. The reaction was conducted at
hopene 11 turned out to be more challenging than expected. 0 °C on a rotary evaporator to continuously remove 2,2-
Catalytic hydrogenation was disappointing even under pressure, dimethoxypropane and shift the equilibrium. Purification
and gave very low yields in the presence of Pd/C (<10%), of free adenosylhopane 2 also proved tricky, due to its amphi-
Adam’s catalyst (<10%)14,27 or no reaction with Wilkinson’s28 philic character and its insolubility in polar solvents such as
and Crabtree’s29 catalysts. Again, the steric hindrance around acetonitrile, methanol and water preventing purification by
the double bond probably allowed only restricted access or no reversed phase HPLC. Purification was indeed successfully
access at all for the different catalysts to the double bond. This achieved by gravity column chromatography on silica gel with
hindered access to the olefin was circumvented by the use of a polar ternary solvent (chloroform–methanol–ammonia).
the small sized diimide, a short-lived reagent that can be
Deuterium labelling of adenosylhopane
implicated in the reduction of nonpolar multiple bonds. In a
concerted mechanism, cis-diimide is converted into N2 via a Only minor changes were required to obtain the deuterium
six-centre transition state corresponding formally to the syn
labelled adenosylhopane 2-D (Scheme 3). Olefin 12 was first
addition of dihydrogen to the double bond.30a,b Taking into
treated with CH3O2H to avoid the presence of extra protons
account that this reaction had to be extended to a deuterium coming from the exchangeable hydrogens of the amino group
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Scheme 3 Hemisynthesis of (30,31-2H2)adenosylhopane 2-D. (i) MeOD; (ii) PADA, AcOD, Py, 90 °C and (iii) H2O, 60%, 2 steps; (iv) TFA, CHCl3–
CH3OH, 0 °C, 90%.
to yield 12-D. Monodeuteriated acetic acid CH3COO2H (98% hopane di-, tri- and tetraacetate 14a–c were compared with
isotope abundance) was adopted as deuterium source to gene- those of acetylated adenosylhopane isolated from bacteria.1a,b
rate dideuteriodiimide. Reduction of olefin 12-D was much All the analytical data were consistent with those previously
slower with deuteriodiimide than that with natural abundance described. The assignments of the 1H and 13C-NMR signals
diimide due to a primary deuterium isotope kinetic effect. were made with the help of 2D NMR (1H–1H COSY, HMBC and
This relatively slow reaction resulted in a decreased yield HSQC) and by comparison with those obtained on other
(60%), but allowed to recover unreacted starting material 12 hopanoids.16,32
1
after 36 h treatment. The H-NMR spectrum of the recovered
The structure of protected bisdeuteriated (31,32-2H2)adeno-
starting material 12 showed that the E/Z ratio decreased from sylhopane 13-D was confirmed by comparing its 1H- and
5 : 1 before diimide reduction to 1 : 1 after reduction, indicat- 13C-NMR spectra recorded in (2H5)pyridine with those of the
ing a faster hydrogenation rate of the E olefin. This phenom- corresponding natural abundance adenosylhopane derivative
enon has been frequently observed in diimide reductions and 18. Four different configurations at C-30 and C-31 were
2
is attributed to the more cluttered transition state for a Z observed after N2 H2 reduction. Given that diimide reacts
olefin than for an E olefin.30a
faster with the protected (E)-adenosylhopene 17, which is the
The deuterium isotope abundance of the protected adeno- dominant isomer, the major products resulting from syn
sylhopane 13-D was determined by mass spectrometry. In spite addition of deuteriated diimide to the E double bond should
of all precautions against water contamination the deuterium be protected (30R,31R-2H2)- and (30S,31S-2H2)adenosylhopane,
content was not maximal, and the reaction resulted in a and the (30R,31S-2H2)- and (30S,31R-2H2)adenosylhopane dia-
mixture of isotopomers: (30,31-2H2)-13-D (60%), (30-2H1)- and stereomers would be generated by the reduction of protected
(31-2H1)-13-D (31%) and natural abundance 13 (9%). This (Z)-adenosylhopene by diimide (cf. ESI and Scheme S2† for the
incomplete labelling is probably due to the large excess of discussion of the stereochemistry of those compounds and the
CH3COO2H (60 eq.) having only a 98% isotope abundance and interpretation of the NMR spectra).
to the primary deuterium kinetic effect, leading to a mixture of
2
2
N2 H2, N2 H H and N2H2. Deuterium isotope abundance can
be probably improved by using CH3COO2H with higher
isotope abundance. The obtained labelling was, however,
largely sufficient to perform the incorporation experiments.
Conversion of adenosylhopane into bacteriohopanetetrol by a
cell free system from Methylobacterium organophilum
Methylobacterium organophilum has been chosen for its sub-
stantial production of hopanoids (up to 13 mg g−1, dry
weight).33 It only synthesizes bacteriohopanetetrol derivatives,
a glycoside and two ethers involving a carbapseudopentose
Structure of adenosylhopane and of its bisdeuteriated
isotopomer
To confirm the structure, synthetic adenosylhopane 2 was moiety, free bacteriohopanetetrol being a minor hopanoid.
acetylated (Scheme 2). The spectroscopic data of adenosyl- This makes this bacterium a good candidate for incorporations
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Table 2 Incorporation of (30,32-2H2)adenosylhopane 2-D into bacterio-
hopanetetrol 5 by cell-free systems from Methylobacterium organo-
philum: 2H isotope abundances of fragments A and Ba
with a crude cell-free system, since dilution of the deuterium
labelled tetrol synthesized de novo from (2H)adenosylhopane
by the natural abundance endogenous tetrol synthesized
during the culture will be minimal. In addition, in order to
diminish this dilution, unbroken cells and large cell debris
containing most of the hopanoids were removed by centrifu-
gation, and the incubations were only performed with the
resulting supernatant.
Fragment A (m/z = 191)
Fragment B (m/z = 493)
Conditionsb m + 1/m (%) m + 2/m (%) m + 1/m (%) m + 2/m (%)
1
2
3
4
5
Referencec
20
19
21
24
19
4
5
4
2
28
27
36
29
28
3
6
47
19
6
1
A
B
C
D
4
According to the hypothetical biogenetic scheme
(Scheme 1), several steps are required for the conversion of
adenosylhopane into bacteriohopanetetrol via ribosylhopane:10
1d
2
6
a Bacteriohopanetetrol was analysed as tetraacetate by GC/MS. Isotopic
patterns of the fragments were measured according to the signal
intensities observed for m/z (reference signal, 100%), m/z + 1 and m/z +
2. The standard deviation of the signal intensities of both fragments
was calculated from the GC-MS analyses of seven samples of
peracetylated bacteriohopanetetrol 5 isolated from seven different cell
a
deadenylation into ribosylhopane, the hemiketal ring-
opening of ribosylhopane and a reduction. To obtain ribosyl-
hopane 3, the postulated precursor,10,13 two pathways have
been proposed for the deadenylation: either a direct hydrolysis
of adenosylhopane to ribosylhopane by a nucleosidase-like
enzyme12 or a phosphorolysis of the nucleotide hemiaminal
with the loss of adenine by a purine nucleoside phosphoryl-
ase-like enzyme, yielding ribosylhopane phosphate.11 In this
case, ribosylhopane 35-phosphate cannot be reduced and has
first to be converted by a phosphatase into ribosylhopane 3
prior reduction. The final step is in all cases the reduction of
the open aldehyde form of ribosylhopane by an NADPH-depen-
dent aldose reductase-type enzyme (Bodlenner and Rohmer,
unpublished results). For those reasons, the incubations were
performed in a phosphate buffer or in a phosphate free tri-
ethylamine buffer in the presence of NADPH. Adenosylhopane
was introduced as a THF solution. Although the use of this
batches.
doubling the standard deviation value. The intensities if signals of
fragments and have
2% deviation. b All the incorporation
experiments were performed at 30 °C for 24 h. A. Deuteriated
adenosylhopane with cell-free extract in phosphate
buffer. B. Deuteriated adenosylhopane with cell-free extract in TEA
buffer. C. Decomposed deuteriated adenosylhopane after 24 in
A confidence interval of 95% can also be calculated by
A
B
a
h
phosphate buffer and then incubated with the cell-free extract in
phosphate buffer. D. Deuteriated adenosylhopane with inactivated cell
free system in phosphate buffer. c Reference mass spectrum of natural
abundance bacteriohopanetetrol tetraacetate isolated from the
unbroken cells and cell debris pellets recovered after centrifugation
yielding the supernatant utilised for incubations. d High signal/noise
ratio.
2
solvent is not common in enzymatic processes, THF is a good indicating that the H labelling was strictly localized in the C5
solvent for this amphiphilic hopanoid and has the advantage side chain as expected (Table 2, entry 2). In order to minimize
of being water miscible. Fortunately, the presence of 10% THF the influence of the background noise, only the two most
in the incubation medium did not affect the enzymes activi- intense signals of the mass spectrum corresponding to frag-
ties. A lipid extraction followed by acetylation allowed isolation ments generated by ring C cleavage are shown and considered
of bacteriohopanetetrol tetraacetate and analysis of the deuter- for evaluating the conversion of deuteriated adenosylhopane
ium labelling by electron impact GC-MS (Fig. 1). Deuterium into bacteriohopanetetrol (Table 2).
incorporation was evaluated from the relative abundances of
Moreover, the ratio of bis- and monodeuteriated isotopo-
the (m + 1)/z and (m + 2)/z versus those of the m/z signals mers (d2/d1 = 1.9/1) of adenosylhopane was found unchanged
(Table 2). Evidence that adenosylhopane was efficiently con- in the labelled bacteriohopanetetrol (d2/d1 = 1.9/1), indicating
verted into BHT in a phosphate buffer was shown from intense that incubated (2H)adenosylhopane is the only deuterium
2H labelling for all ions containing the side chain [m/z 714 source and that there is no significant isotope effect in this
(M+), 699 (M+ − CH3), 654 (M+ − AcOH), 493 (fragment B, ring enzymatic conversion, which is logical as no reaction affects
C cleavage)], but not for those without the side-chain: [m/z 369 the deuterium labelled carbon atoms in this biosynthetic
(M+ − side chain), 191 (fragment A, ring C cleavage)] (Fig. 1),34 pathway.
Fig. 1 Fragmentation of the hopane skeleton by electron impact mass spectrometry.
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In order to confirm that the overall conversion of (2H)ade-
nosylhopane 2-D into bacteriohopanetetrol 5 is purely enzy-
Conclusion
matic, (2H)adenosylhopane 2-D was incubated with a cell-free A highly efficient synthesis of adenosylhopane and its deuter-
system inactivated by boiling. No deuterium labelling was ium labelled homologue has been achieved from the coupling
found in the bacteriohopanetetrol isolated from this assay of homohop-30-ene and a 5′-methylene adenosine derivative
(Table 2, entry 5).
via a cross-metathesis. This approach was found to be the only
A non-enzymatic conversion of (2H)adenosylhopane 2-D one able of linking the two highly hindered hopane and
into ribosylhopane 3 (or into ribosylhopane phosphate) was adenine counterparts to date, affording thus a total control of
also excluded since a 3 days preincubation of (2H)adenosyl- all the stereogenic centres of this large molecule. Moreover the
hopane in a phosphate buffer with 10% THF at 30 °C did not metathesis step allows further labelling through the reduction
lead to any detectable deuterium incorporation, even though a of the introduced double bond by diimide. The deuterium
partial degradation of adenosylhopane was observed on TLC. labelled adenosylhopane was thus obtained and incubated
Incubation of such a degraded sample resulted in no for- with a supernatant of a crude cell-free system of Methylobacter-
mation of 2H labelled bacteriohopanetetrol, indicating that ium organophilum yielding deuterium labelled bacteriohopane-
even the decomposition products of (2H)adenosylhopane tetrol as shown by EI-MS of its tetraacetate. This allowed for
cannot be converted into bacteriohopanetetrol (Table 2, the first time to demonstrate the substrate to product relation-
entry 4).
ship between adenosylhopane and bacteriohopanetetrol. This
The present conversion of deuterium labelled adenosyl- work opens also the way for the efficient synthesis of other bio-
hopane 2 into bacteriohopanetetrol 5 by a crude cell-free system hopanoids and for further enzymatic studies on isolated
of Methylobacterium organophilum in the presence of NADPH, enzymes required for the full elucidation of hopanoid
indicates that the nucleoside side chain of adenosylhopane 2 biosynthesis.
is the precursor of the D-ribitol side chain of bacterio-
hopanetetrol 5. This biotransformation requires at least two
reactions, depending on the nature of the nucleophile (phos-
phate or water) responsible of the cleavage of the glycosidic
Experimental
Reagents and solvents
bond with adenine loss. Hoping to solve that question, the
(2H)adenosylhopane incubations with a cell-free system of M.
Reagents were purchased from Sigma-Aldrich, Acros, Alfa
Aesar or Carl Roth. Their purity was more than 98% in all
organophilum were performed using two different buffers.
Although, the conversion was significantly higher in the phos-
phate buffer than in the phosphate free triethylamine buffer
(Table 2, entries 2 and 3), there is no clear cut conclusion in
All solvents were distilled before use. Dry solvents used in
favour of one of the two proposed deadenylation mechanism.
A sufficient phosphate concentration originating from the cells
might indeed be still present in the triethylamine buffer to
achieve the phosphorolysis, and a slight inhibitory effect on
the putative aldose reductase of triethylamine/triethylammo-
nium cannot be excluded to explain the lower yields. In fact,
cases. Dowex 50 (H+) resin was purchased from Aldrich and
was activated right before use with a 10% HCl solution in
water.
moisture sensitive reactions were obtained as follows: THF was
freshly distilled from sodium benzophenone ketyl before use.
CH2Cl2 was distilled over P2O5. Pyridine was distilled over KOH
and then stored over molecular sieves (4 Å). DMSO was pur-
chased from Aldrich (sure sealed) and used directly.
ribosylhopane is effectively an intermediate in biohopanoid
biosynthesis. Deletion of the orf18 gene in a S. coelicolor
Purification techniques
The progress of reactions was monitored by analytical thin
layer chromatography using aluminum-coated Merck 60 F254
silica plates. UV visible spots were directly visualized under UV
light at λ = 254 nm. Hopanoids were revealed on TLC plate by
UV light (λ = 366 nm) after spraying with a 0.1% berberine
hydrochloride solution in EtOH.
Most products were purified by flash column chromato-
graphy using Merck 60 (230–400 mesh) silica gel and appropri-
ate eluents.35 Adenosylhopane 2 and 2-D were purified by
gravity column chromatography using Merck 60 (63–200 µm)
silica gel. Compounds in amounts below 10 mg were purified
via preparative thin layer chromatography using Merck 60 F254
silica gel plates (0.25 mm layer thickness).
strain (corresponding to the hpnO gene in M. extorquens and
R. palustris)11,12 allowed to identify for the first time in
the Δorf18 mutant ribosylhopane 3 (Scheme 1), postulated
twenty years ago as an intermediate in the biosynthesis of
C35 hopanoids.10,15 It is accumulated in the absence of
the putative transaminase encoded by the hpnO/orf18 gene
catalysing the reductive transamination of the free aldehyde
4 into aminobacteriohopanetriol 6.13 Decisive proofs will be
obtained later with tests performed with the isolated and
purified enzymes.
In the final step, a reduction converts the open free alde-
hyde 4 of ribosylhopane 3 into bacteriohopanetetrol 5, which
has been found to be NADPH-dependent (Bodlenner and
Rohmer, unpublished results). The reaction resembles the
reduction catalysed by an aldose reductase, but the gene
Spectroscopic analysis
encoding this enzyme is yet unknown in hopanoid producing 1H-NMR and 13C-NMR were performed on BRUKER Avance
1
bacteria.
600 spectrometer (600 MHz for H, 150 MHz for 13C), Bruker
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1
Biospin 500 spectrometer (500 MHz for H, 125 MHz for 13C)
M.p. = 203–205 °C.
and BRUKER Avance 300 spectrometer (300 MHz for 1H,
1H-NMR (300 MHz, C2HCl3): δ/ppm = 5.53 (1H, ddd, J30,31a
=
75 MHz for 13C). Most of the measurements were carried out 17.8 Hz, J30,31b = 9.5 Hz, J22,30 = 9.1 Hz, 30-H), 4.85 (1H, ddd,
using C2HCl3 as solvent and with CHCl3 (δ = 7.26 ppm) as J30,31a = 17.8 Hz, Jgem = 2.0 Hz, J22–31a = 0.5 Hz, 31-Ha), 4.85 (1H,
internal standard for H-NMR and C2H Cl3 (δ = 77.0 ppm) for dd, J30,31b = 9.5 Hz, Jgem = 2.0 Hz, 31-Hb), 2.21 (1H, ddq, J21,22
=
=
1
13C-NMR. NMR spectra of adenosylhopane 2 and 2-D were 9.7 Hz, J22,30 = 9.1 Hz, J22,29 = 6.5 Hz, 22-H), 1.8 (1H, tdd, J21,22
recorded in (2H5)pyridine as solvent and with (2H4)pyridine 9.7 Hz, J20,21 = J17,21 = 4.8 Hz, 21-H), 1.01 (3H, d, J22,29 = 6.5 H,
1
(δ = 8.73, 7.58 and 7.21 ppm) as internal standard for H-NMR 22R-Me), 0.960 (6H, s, 8β-Me and 14α-Me), 0.851 (3H, s, 4α-Me),
and (2H5)pyridine (δ = 149.9, 135.5 and 123.5 ppm) for 0.818 (3H, s, 10β-Me), 0.796 (3H, s, 4β-Me), 0.710 (3H, s,
13C-NMR. The following abbreviations represent the multi- 18α-Me).
plicity of the NMR signals: s, singlet; d, doublet; t, triplet; q,
13C-NMR (75 MHz, C2HCl3): δ/ppm = 145.3, 112.0, 56.1,
quadruplet; m, multiplet; dd, doublet of doublet; ddd, doublet 54.1, 50.4, 49.2, 45.0, 44.4, 42.2, 42.1, 41.9, 41.8, 41.7, 40.3,
of doublet of doublet; dddd, doublet of doublet of doublet of 37.4, 33.5, 33.4, 33.3, 33.2, 27.6, 24.0, 22.3, 22.0, 21.6, 20.9,
doublet; arom., aromatic protons or carbon atoms; br, broad.
1H- and 13C-NMR assignments of hopanoids were based on
18.7, 16.6, 15.9.
EI-MS (direct inlet, positive mode 70 eV): m/z = 424 (M+,
earlier assignments14,17,32 and were supported by additional 19%), 409 (M+–CH3, 5%), 369 (M+–side chain, 20%), 203 (ring
experiments including DEPT, HSQC and HMBC techniques. C cleavage, 100%), 191 (ring C cleavage, 88%).
13C shift values of the carbon atoms in the hopanoid skeleton
N-Benzoyl-33,34-O-isopropylidene-adenosylhopene (11) and
homodimer (15)
did not depend significantly on the nature of the side-chain
and remained virtually unchanged after introduction of new
carbon atoms or functional groups in the C5 side-chain.
Procedure A. Olefins 10 (156 mg, 0.37 mmol) and 8 (15 mg,
GC-MS spectra were acquired on a Thermo TSQ Quantum 0.037 mmol) and Ru-catalyst (10 mol%) were dissolved in dry
mass spectrometer connected to a Thermo Trace GC ultra gas DCM (3.6 mL) at ambient temperature under an atmosphere
chromatograph (PTV injector, HP-5 MS column, 30 m, of argon and the resulting solution was heated overnight at
0.25 mm internal diameter, 0.25 μm film thickness). Helium 75 °C (oil bath) in a pressure tube (Ace glass). Volatiles were
(constant flow 1 mL min−1) was used as carrier gas. The temp- evaporated and the residue was purified by flash chromato-
erature program for GC analysis was: 3 min at 55 °C, from graphy (DCM to DCM–MeOH 100 : 3) to afford compound 11
55 °C to 320 °C at 10 °C min−1 and 50 min at 320 °C.
Mass spectra were obtained by direct inlet on a Thermo 15 (6 mg, 43%) as a by-product.
(15 mg, 52%, E/Z ∼4 : 1) as a mixture of two isomers and dimer
TSQ Quantum mass spectrometer at 70 eV in the electron
Procedure B. Olefins 10 (210 mg, 0.49 mmol) and 8 (20 mg,
impact positive ionization mode (EIMS) over a mass range of 0.049 mmol) and Ru-catalyst (10 mol%) were dissolved in dry
50–800 Da (cycle time 0.5 s). The source temperature was set at CH2Cl2 (5 mL) in a sealed tube equipped with a Teflon-coated
230 °C.
ESI-HRMS mass spectra were carried out on a Bruker Micro- 3 h. The mixture was then evaporated to dryness in vacuo and
TOF spectrometer. purified by flash chromatography (CH2Cl2 to CH2Cl2–MeOH,
stirrer bar and heated under microwave irradiation at 75 °C for
Melting points were measured with a BIBBY SMP3 appar- 100 : 3) to afford compound 11 (23 mg, 59%, E/Z ∼4 : 1) and
atus and are corrected with benzophenone (m.p. 48 1.5 °C). dimer 15 (7.5 mg, 40%). Compound 11 was isolated as a col-
Optical rotations were measured on a Perkin Elmer 341 polari- ourless solid mixture of the E/Z isomers that were not separ-
meter at 20 2 °C and 589 nm using 10 cm length and 1 mL ated. Rf 0.47 (CH2Cl2–MeOH, 100 : 3).
volume cell.
1H-NMR (300 MHz, C2HCl3) of (E)-11 and (Z)-11 (subscripts
“E” and “Z” characterize respectively the 1H-NMR signals of
(E)-11 and (Z)-11, which are respectively present in a 4 : 1
(22S)-Homohop-30-ene (10)
A suspension of NaH (126 mg, 5 mmol) in DMSO (2.5 mL) was ratio): δ/ppm = 9.29 (1H, br. s, –NHCvO), 8.82 (1H, s, 2′-H),
stirred at 75 °C for 50 min. The light green suspension was 8.12 (0.2H, s, 8′-HZ), 8.10 (0.8H, s, 8′-HE), 8.04–8.01 (2H, m,
cooled down to room temperature, added into a solution of Ar–H), 7.61–7.47 (3H, m, Ar–H), 6.14 (1H, d, J34,35 = 1.9 Hz, 35-HZ
PPh3CH3Br (1.9 g, 5.4 mmol) in DMSO (27 mL) and stirred at and 35-HE), 5.55 (0.8H, dd, J33,34 = 6.2 Hz, J34,35 = 1.9 Hz,
room temperature for 20 min to afford a light green-grey sus- 34-HE), 5.51 (0.2H, dd, J33,34 = 6.1 Hz, J34,35 = 1.8 Hz, 34-HZ),
pension. The resulting mixture was then added into a solution 5.46 (0.8H, dd, J30,31 = 15.4 Hz, J22,30 = 8.9 Hz, 30-HE), 5.37
of aldehyde 9 (297 mg, 0.70 mmol) in THF (20 mL) and stirred (0.8H, dd, J30,31 = 15.4 Hz, J31,32 = 7.4 Hz, 31-HE), 5.28 (0.4H,
overnight at 50 °C. The reaction was quenched with water, and dd, J30,31 = 11.0 Hz, J31,32 = 8.7 Hz, 30-HZ and 31-HZ), 5.03
extracted three times with pentane. The combined organic (0.2H, dd, J31,32 = 8.7 Hz, J32,33 = 3.0 Hz, 32-HZ), 4.96 (0.8H, dd,
phases were dried over anhydrous Na2SO4, filtered through J33,34 = 6.2 Hz, J32,33 = 3.0 Hz, 33-HE), 4.85 (0.2H, dd, J33,34
=
cotton, and evaporated to dryness in vacuo to give an almost 6.0 Hz, J32,33 = 3.0 Hz, 33-HZ), 4.74 (0.8H, dd, J31,32 = 7.3 Hz,
pure crystalline product, which was further purified by flash J32,33 = 2.8 Hz, 32-HE), 2.71–2.65 (0.2H, m, 22-HZ), 2.17–2.11
chromatography (petroleum ether) to yield the terminal olefin (0.8H, m, 22-HE), 1.65 and 1.41 (6H, 2s, Me2C), 1.01 (0.6H, d,
10 (290 mg, 98%). Rf 0.67 (petroleum ether–EtOAc, 20 : 1).
J22,29 = 6.2 Hz, 22R-MeZ), 0.97 and 0.95 (1.2H, 2s, 8β-MeZ and
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14α-MeZ), 0.93 and 0.92 (4.8H, 2s, 8β-MeE and 14α-MeE), 0.89 J22,30 = 8.1 Hz, 30-HE), 5.36 (0.8H, dd, J30,31 = 15.3 Hz, J31,32
(2.4H, d, J22,29 = 6.4 Hz, 22R-MeE), 0.84 (18H, s, 4α-Me), 0.80 6.8 Hz, 31-HE), 5.31–5.29 (0.4H, m, 31-HZ and 30-HZ), 4.97
(0.2H, dd, J31,32 = 4.7 Hz, J32,33 = 2.9 Hz, 32-HZ), 4.94 (0.8H, dd,
=
(3H, s, 10β-Me), 0.781 (3H, s, 4β-Me), 0.64 (3H, s, 18α-Me).
13C-NMR of the major isomer (E)-11 (75 MHz, C2HCl3): J33,34 = 6.3 Hz, J32,33 = 3.1 Hz, 33-HE), 4.83 (0.2H, dd, J33,34 = 6.3
δ/ppm = 164.5, 152.6, 151.1, 149.6, 142.1, 133.6, 132.7, 128.7, Hz, J32,33 = 3.1 Hz, 33-HZ), 4.69 (0.8H, dd, J31,32 = 6.8 Hz,
127.8, 124.5, 123.5, 114.3, 91.2, 88.6, 84.8, 84.4, 56.0, 54.0, J32,33 = 2.9 Hz, 32-HE), 2.72–2.63 (0.2H, m, 22-HZ), 2.17–2.09
50.3, 49.1, 44.8, 44.3, 42.0, 41.8, 41.7, 41.5, 40.5, 40.2, 37.3, (0.8H, m, 22-HE), 1.63 (0.6H, s, Me2CZ), 1.61(2.4H, s, Me2CE),
33.4, 33.3, 33.2, 33.2, 27.5, 27.0, 25.4, 23.9, 21.9, 21.7, 21.5, 1.40 (3H, s, Me2C), 1.00 (0.6H, d, J22,29 = 6.4 Hz, 22R-MeZ), 0.96
20.9, 18.6, 16.5, 15.9, 15.8.
and 0.94 (1.2H, 2s, 8β-MeZ and 14α-MeZ), 0.93 (2.4H, s,
8β-MeE), 0.92 (2.4H, s, 14α-MeE), 0.89 (2.4H, d, J22,29 = 6.4 Hz,
22R-MeE), 0.834 (3H, s, 4α-Me), 0.799 (3H, s, 10β-Me), 0.779
(3H, s, 4β-Me), 0.640 (3H, s, 18α-Me).
+
HRMS (ESI): m/z [M + Na]+, calcd for C50H69N5NaO4
826.524, found 826.524.
Homodimer of 5′-methylene adenosine (15)
13C-NMR (75 MHz, C2HCl3) for (E)-12: δ/ppm = 155.5, 153.0,
Dimer 15 was isolated as a solid mixture of two E/Z isomers 149.4, 141.7, 139.8, 124.8, 120.3, 114.2, 90.8, 88.4, 85.0, 84.4,
(∼3 : 1). Rf = 0.16 (CH2Cl2–MeOH, 100 : 3).
56.1, 54.1, 50.4, 49.1, 44.9, 44.3, 42.1, 41.8, 41.7, 41.6, 40.5,
1H-NMR (300 MHz, C2HCl3) (subscripts “E” and “Z” charac- 40.3, 37.4, 33.5, 33.4, 33.3, 33.2, 27.5, 27.1, 25.4, 23.9, 22.1,
terize respectively the 1H-NMR signals of (E)-15 and (Z)-15, 22.0, 21.7, 21.6, 20.9, 18.7, 16.6, 15.9.
which are respectively present in a 3 : 1 ratio): δ/ppm = 9.24
HRMS (ESI): m/z [M + H]+, calcd for C43H65N5NaO3
+
(1.5H, br. s, –NHBzE), 9.11 (0.5H, br. s, –NHBzZ), 8.79 (0.5H, 722.498, found 722.497.
br. s, 2-HZ), 8.63 (1.5H, br. s, 2-HE), 8.10 (0.5H, s, 8-HZ),
33,34-O-Isopropylidene adenosylhopane (13)
To a stirred slurry of potassium azodicarboxylate (90 mg,
1.7 Hz, 1′-HE), 5.72 (1.5H, dd, J = 3.4, 1.5 Hz, 5′-HE), 5.63 (0.5H, 0.46 mmol) in pyridine (1 mL) in a two-neck round bottom
dd, J = 5.5, 1.3 Hz, 5′-HZ), 5.56 (0.5H, dd, J2′,3′ = 6.1 Hz, J1′,2′ flask equipped with a condenser under N2 atmosphere was
8.04–8.02 (4H, m, Ar-H), 7.95 (1.5H, s, 8-HE), 7.64–7.44 (6H, m,
Ar-H), 6.15 (0.5H, d, J1′,2′ = 1.5 Hz, 1′-HZ), 6.06 (1.5H, d, J1′,2′
=
=
1.5 Hz, 2′-HZ), 5.44 (1.5H, dd, J2′,3′ = 6.3 Hz, J1′,2′ = 1.7 Hz, added a solution of 12 (32 mg, 0.046 mmol) in pyridine
2′-HE), 5.14–5.11 (0.5H, m, 3′-HZ), 4.94 (1.5H, dd, J2′,3′ = 6.3, (3 mL). The mixture was heated to reflux, and a solution of
J3′,4′ = 3.3 Hz, 3′-HE), 4.69–4.67 (0.5H, m, 4′-HZ), 4.63 (1.5H, m, anhydrous acetic acid (0.030 mL, 0.55 mmol) in pyridine
4′-HE), 1.63 and 1.38 (3H, 2s, Me2CZ), 1.59 and 1.36 (9H, (0.1 mL) was carefully added dropwise. Each drop of acetic
2s, Me2CE).
acid solution was added after the end of N2 evolution. The
13C-NMR (75 MHz, C2HCl3): δ/ppm = 164.7E, 152.7Z, 152.4E, mixture was left under reflux until the yellow colour vanished.
150.9E, 149.8Z, 149.7E, 142.4Z, 142.3E, 133.5E, 133.4Z, 132.8Z, Another five portions of potassium azodicarboxylate and sub-
132.7E, 131.2Z, 130.0E, 128.9Z, 128.7E, 128.0E, 127.9Z, 123.6E, sequent acetic acid were necessary to increase the conversion.
114.6Z, 114.5E, 91.1Z, 90.7E, 87.3E, 85.4Z, 84.5E, 84.3Z, 84.0E, The reaction was quenched with water, and extracted with
83.5Z, 27.0E, 27.0Z, 25.3Z, 25.2E.
dichloromethane three times. The combined organic phases
were washed with brine and dried over anhydrous Na2SO4, fil-
tered through cotton, and evaporated to dryness in vacuo. The
crude product was further purified by flash chromatography
(CH2Cl2–MeOH 100 : 1 to 100 : 2) to yield the saturated product
13 (23 mg, 73%). Compound 13 was isolated as a colourless
HRMS (ESI): m/z [M + Na]+, calcd for C40H38N10NaO8
809.277, found 809.277.
+
(E)-33,34-O-Isopropylidene adenosylhop-30-ene ((E)-12) and
(Z)-33,34-O-isopropylidene adenosylhop-30-ene ((Z)-12)
To a mixture of compounds (E)-11 and (Z)-11 (150 mg, solid. Rf = 0.38 (CH2Cl2–MeOH, 100 : 5).
0.19 mmol) in CH2Cl2 (5 mL) was added a solution of
ammonia in MeOH (7N, 20 mL) at 0 °C, and the mixture was
M.p. = 256–257 °C. [α]2D0 = +34 (c 0.35, CHCl3).
1H-NMR (C2HCl3, 300 MHz): δ/ppm = 8.35 (1H, s, 2′-H),
stirred at 5 °C for 5 days. The volatiles were evaporated and the 7.89 (1H, s, 8′-H), 6.03 (1H, dd, J34,35 = 2.4 Hz, 35-H), 5.85 (2H,
residue was purified via flash chromatography (CH2Cl2 to s, –NH2), 5.51 (1H, J33,34 = 6.5 Hz, J34,35 = 2.4 Hz, 34-H), 4.80
CH2Cl2–MeOH, 100 : 2) to give a mixture of (E)-12 and (Z)-12 (1H, dd, J33,34 = 6.5 Hz, J32,33 = 3.5 Hz, 33-H), 4.15 (1H, ddd,
(120 mg, 0.17 mmol, 91%) as a colourless solid, which was J31a,32 = 7.4 Hz, J31b,32 = 6.5 Hz, J32,33 = 3.5 Hz, 32-H), 1.60 and
used in the next step without further purification. Rf = 0.36 1.38 (2 × 3H, 2s, Me2C), 0.928 (3H, s, 8β-Me), 0.921 (3H, S,
(CH2Cl2–MeOH, 100 : 5).
14α-Me), 0.83 (3H, d, J = 6.2 Hz, 22-Me), 0.834 (3H, s, 4α-Me),
1H-NMR (300 MHz, C2HCl3) for (E)-12 and (Z)-12 (subscripts 0.800 (3H, s, 10β-Me), 0.779 (3H, s, 4β-Me), 0.656 (3H, s,
“E” and “Z” characterize respectively the 1H-NMR signals of 18α-Me).
(E)-12 and (Z)-12, which are respectively present in a 4 : 1
13C-NMR data (C2HCl3, 75 MHz): δ/ppm = 155.4 (C-6′),
ratio): δ/ppm = 8.36 (0.2H, s, 2′-HZ), 8.35 (0.8H, s, 2′-HE), 7.91 153.0 (C-2′), 139.8 (C-8′), 149.4 (C-4′), 120.3 (C-5′), 114.4
(0.2H, s, 8′-HZ), 7.89 (0.8H, s, 8′-HE), 6.07 (1H, d, J34,35 = 1.8 Hz, (CMe2), 90.5 (C-35), 87.7 (C-32), 84.3 (C-34), 84.0 (C-33), 56.1
35-HZ and 35-HE), 5.90 (2H, br. s, –NH2), 5.54 (0.8H, dd, (C-5), 54.4 (C-17), 50.4 (C-9), 49.3 (C-13), 45.7 (C-21), 44.3
J33,34 = 6.2 Hz, J34,35 = 1.9 Hz, 34-HE), 5.50 (0.2H, dd, J33,34
=
(C-18), 42.1 (C-3), 41.8 (C-14), 41.7 (C-8), 41.5 (C-19), 40.3 (C-1),
6.2 Hz, J34,35 = 1.9 Hz, 34-HZ), 5.45 (0.8H, dd, J30,31 = 15.3 Hz, 37.4 (C-10), 36.4 (C-22), 33.6 (C-15), 33.4 (C-23), 33.2 and 33.1
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(C-4 and C-7), 31.4 (C-30), 30.0 (C-31), 27.5 (C-20), 23.9 (C-12), (rotary evaporator). When starting material could no more be
22.7 (C-16), 21.6 (C-24), 20.9 (C-11), 20.0 (C-29), 18.7 (C-2 and detected by TLC, the reaction mixture was evaporated to
C-6), 16.6 and 16.5 (C-26 and C-27), 15.9 and 15.8 (C-25 and dryness and the residue was purified by column chromato-
C-28).
HRMS (ESI): m/z [M + H]+, calcd for C43H67N5NaO3
724.514, found 724.512.
graphy over silica gel (63–200 µm) (CHCl3–MeOH–NH4OH,
100 : 2 : 0.5 to 100 : 5 : 0.5 to 100 : 7.5 : 0.5) to afford adenosyl-
hopane 2 as a colourless solid (5.3 mg, 8 µmol 90%). Rf = 0.09
(CH2Cl2–MeOH 100 : 5). [α]2D0 = +34 (c 0.3, THF).
+
[30,31-2H2]-33,34-O-Isopropylidene adenosylhopane (13-D)
1H-NMR ((2H5)pyridine, 600 MHz): δ (ppm) = 8.77 (1H, s,
Compound 12 (51 mg, 0.073 mmol) was dissolved in a mixture 2′-H), 8.61 (1H, s, 8′-H), 8.29 (2H, s, –NH2), 7.74 (1H, d, J34,OH
=
of CH3O2H–C2HCl3 (1 : 2) under an atmosphere of N2 for H-2H 5.5 Hz, 34-OH), 7.04 (1H, d, J33,OH = 5.8 Hz, 33-OH), 6.72 (1H,
exchange. After 5 min standing, the solvents were carefully d, J34,35 = 4.4 Hz, 35-H), 5.43 (1H, pseudo q, J = 4.7 Hz, 34-H),
evaporated under vacuum. The same procedure was repeated 4.77 (1H, pseudo q, J = 5.2 Hz, 33-H), 4.56 (1H, pseudo dt, J =
three times, and the N-deuteriated intermediate 12-D was care- 8.4 Hz, J = 4.9 Hz, 32-H), 2.13 (1H, m, 31-Ha), 1.96 (1H, m,
fully dried overnight under vacuum in the presence of P2O5. 31-Hb), 1.79 (2H, m, 30-Ha and 20-Ha), 1.73 (1H, m, 21-H),
Compound 13-D was prepared following the procedure 0.994 (3H, d, J = 6.3 Hz), 0.959 (3H, s, 8β-Me), 0.948 (3H, s,
described for the synthesis of natural abundance 13 using pot- 14α-Me), 0.882 (3H, s, 4α-Me), 0.819 (3H, s, 4β-Me), 0.815 (3H,
assium azodicarboxylate (850 mg, 43.8 mmol) and CH3COO2H s, 10β-Me), 0.665 (3H, s, 18α-Me).
(98% isotope abundance, 3.3 mL, 56.9 mmol) to generate deu-
13C-NMR ((2H5)pyridine, 150 MHz): δ (ppm) = 157.6 (C-6′),
teriated diimide in situ. Compound 13-D (31 mg, 60%) was iso- 153.8 (C-2′), 150.6 (C-4′), 140.0 (C-8′), 121.3 (C-5′), 90.0 (C-35),
lated as a colourless solid with unreacted starting material 12 85.2 (C-32), 75.24 and 75.16 (C-33 and C-34), 54.6 (C-17), 56.4
(8.6 mg, 17%) by TLC (CH2Cl2–MeOH, 100 : 5, Rf = 0.38). Deu- (C-5), 50.7 (C-9), 49.6 (C-13), 46.4 (C-21), 44.5 (C-18), 42.3(C-3),
terium content of 13-D was determined from HRMS (ESI) spec- 42.0 (C-14), 41.9 (C-8), 41.8 (C-19), 40.5 (C-1), 37.6 (C-10), 36.9
trum by evaluating the relative intensities of the m/z 724, 725 (C-22), 34.0 (C-15), 33.6 (C-7 and C-23), 33.4 (C-4), 32.3 (C-30),
and 726 peaks corresponding to each isotopomer: natural 31.0 (C-31), 27.9 (C-20), 24.2 (C-12), 23.0 (C-16), 21.8 (C-24),
abundance adenosylhopane (8%), monodeuteriated adenosyl- 21.2 (C-11), 20.4 (C-29), 19.0 (C-2 and C-6), 16.8 and 16.7 (C-26
hopane (32%) and bisdeuteriated adenosylhopane (60%), tris- and C-27), 16.1 and 16.0 (C-25 and C-28).
HRMS (ESI): m/z [M + H]+, calcd for C40H64N5O3 662.501,
+
deuteriated adenosylhopane (0%).
1H-NMR (C2HCl3, 600 MHz): δ/ppm = 8.35 (1H, s, 2′-H), found 662.500.
7.90 (1H, s, 8′-H), 6.03 (1H, d, J34,35 = 2.4 Hz, 35-H), 5.92 (2H, s,
(30,31-2H2)Adenosylhopane (2-D)
–NH2), 5.51 (1H, ddd, J33,34 = 6.4 Hz, J34,35 = 2.4 Hz, J32,34
=
1.2 Hz, 34-H), 4.80 (1H, ddd, J33,34 = 6.4 Hz, J32,33 = 3.5 Hz, Deprotection of 13-D (9.3 mg, 13.2 µmol) by TFA (3 mL) in
J31,33 = 1.2 Hz, 33-H), 4.15 (1H, dd for the major isotopomer, CHCl3–MeOH (2 : 1, 7 mL) following procedure D afforded deu-
J31,32 = 7.2 Hz, J32,33 = 3.5 Hz, 32-H), 1.60 and 1.38 (2 × 3H, 2s, teriated adenosylhopane 2-D (7.7 mg, 11.6 µmol, 90%). Rf =
Me2C), 0.927 (3H, s, 8β-Me), 0.920 (3H, S, 14α-Me), 0.833 (3H, 0.09 (DCM–MeOH 100 : 5). Deuterium content of 2-D was cal-
s, 4α-Me), 0.832 (3H, d, J = 6.3 Hz, 22-CH3), 0.798 (3H, s, culated from HRMS (ES) spectrum: natural abundance adeno-
10β-Me), 0.778 (3H, s, 4β-Me), 0.655 (3H, s, 18α-Me).
sylhopane 8%; monodeuteriated adenosylhopane 32%;
13C-NMR data (C2HCl3, 150 MHz): δ/ppm = 155.5 (C-6′), bisdeuteriated adenosylhopane 60%; trisdeuteriated adenosyl-
152.9 (C-2′), 139.8 (C-8′), 149.4 (C-4′), 120.3 (C-5′), 114.4 hopane 0%.
(CMe2), 90.5 (C-35), 87.63 and 87.60 (C-32, β-shift induced by
deuterium at C-31), 84.3 (C-34), 84.0 (C-33), 56.1 (C-5), 54.3 2′-H), 8.60 (1H, s, 8′-H), 8.28 (2H, s, –NH2), 7.73 (1H, d, J34,OH
1H-NMR ((2H5)pyridine, 600 MHz): δ (ppm) = 8.72 (1H, s,
=
(C-17), 50.4 (C-9), 49.2 (C-13), 45.7 (C-21), 44.3 (C-18), 42.1 5.1 Hz, 34-OH), 7.03 (1H, d, J33,OH = 5.6 Hz, 33-OH), 6.72 (1H,
(C-3), 41.8 (C-14), 41.6 (C-8), 41.5 (C-19), 40.3 (C-1), 37.4 (C-10), d, J34,35 = 4.4 Hz, 35-H), 5.43 (1H, pseudo q, J = 4.3 Hz, 34-H),
36.35, 36.28 and 36.25 (C-22, β-shifts induced by deuterium 4.77 (1H, pseudo q, J = 5.0 Hz, 33-H), 4.57–4.54 (1H, m, 32-H),
at C-30), 33.6 (C-15), 33.4 (C-23), 33.2 and 33.1 (C-4 and 2.11 (1H, dd, J31a, 32 = 8.3 Hz, J30b,31a = 4.6 Hz, 31-Ha), 1.93
C-7), missing signal (C-30 bearing a deuterium), missing (1H, pseudo t, J31b,32 = J30a,31b = 5.0 Hz, 31-Hb), 1.82–1.71
signal (C-31 bearing a deuterium), 27.5 (C-20), 23.9 (C-12), (2.7H, m, 30-Ha, 20-Ha and 21-H of isotopomer resulting from
22.7 (C-16), 21.6 (C-24), 20.9 (C-11), 20.0 (C-29), 18.7 (C-2 deuteriation of major isomer E-12), 1.75–1.72 (1H, m, 21-H),
and C-6), 16.6 and 16.5 (C-26 and C-27), 15.9 and 15.8 (C-25 0.993 (3H, d, J = 6.3 Hz), 0.961 (3H, s, 8β-Me), 0.949 (3H, s,
and C-28).
14α-Me), 0.883 (3H, s, 4α-Me), 0.820 (3H, s, 4β-Me), 0.816 (3H,
s, 10β-Me), 0.667 (3H, s, 18α-Me).
HRMS (ESI): m/z [M + H]+, calcd for C43H65D2N5NaO3
+
726.526, found 726.523.
13C-NMR ((2H5)pyridine, 150 MHz): δ (ppm) = 157.6 (C-6′),
153.8 (C-2′), 150.6 (C-4′), 140.0 (C-8′), 121.3 (C-5′), 90.0 (C-35),
85.20 and 85.15 (C-32, β-shift induced by deuterium at C-31),
Adenosylhopane (2)
A solution of 13 (6.3 mg, 9 µmol) and TFA (2 mL) in CHCl3– 75.21 and 75.15 (C-33 and C-34), 54.6 (C-17), 56.4 (C-5), 50.7
MeOH (2 : 1, 5 mL) in a round bottomed flask was shaken (C-9), 49.6 (C-13), 46.4 (C-21), 44.5 (C-18), 42.3(C-3), 42.0
under reduced pressure (450–500 mbar) at 0 °C for 2 h30 (C-14), 41.9 (C-8), 41.8 (C-19), 40.5 (C-1), 37.6 (C-10), 36.90,
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36.86 and 36.81 (C-22, β-shifts induced by deuterium at C-30), Culture of M. organophilum and preparation of the cell-free
33.9 (C-15), 33.6 (C-7 and C-23), 33.4 (C-4), missing signal at system
ca. 32.3 (C-30 bearing a deuterium), missing signal at ca. 31.0
Methylobacterium organophilum DSM 760 was grown on a
(C-31 bearing a deuterium), 27.9 (C-20), 24.2 (C-12), 23.0
modified medium of Hestrin and Schramm36 (6 × 500 mL,
(C-16), 21.8 (C-24), 21.2 (C-11), 20.3 (C-29), 19.0 (C-2 and C-6),
yeast extract 5 g L−1, bactopeptone 5 g L−1, glucose 1 g L−1
,
16.8 and 16.7 (C-26 and C-27), 16.1 and 16.0 (C-25 and C-28).
HRMS (ESI): m/z [M + H]+, calcd for C40H622H2N5O3
664.513, found 664.513.
anhydrous citric acid 1.04 g L−1, Na2HPO4 2.7 g L−1, pH = 6.8)
in 2 L Erlenmeyer flasks on a rotatory shaker (200 rpm) at
30 °C. Optical densities of the bacterial cultures were
measured at 595 nm on a Genesys 10UV spectrophotometer
using 1 cm in length and 1 mL volume cuvettes. The cells were
harvested before the end of the exponential growth phase (OD
∼ 0.6) by centrifugation (8 000 rpm, 10 min), and then washed
with buffer either a sodium phosphate buffer (50 mM
NaH2PO4–Na2HPO4, 0.5 mM, MgCl2, 1 mM dithiothreitol, pH
7.5) or a triethylamine buffer (0.1 M triethylamine, 0.5 mM
MgCl2, 1 mM dithiothreitol, pH 7.5). After centrifugation (8
000 rpm, 10 min), the cell pellet (3 g) was suspended in the
buffer (15 mL) and disrupted by sonication at 0 °C in a
melting ice bath using a Branson SONIFIER B-30 (8 × 40 s with
3 min pause at 0 °C, 60% pulsed duty cycle, output control: 6).
The crude cell-free system was centrifuged (10 000 rpm,
10 min). The pellet was freeze-dried to recover bacteriohopane-
tetrol of natural abundance, and the supernatant was directly
used for the incubation deuterium labelled adenosylhopane.
+
Adenosylhopane acetates (14a–c)
Adenosylhopane 2 (1.8 mg) was acetylated overnight at room
temperature with a mixture of acetic anhydride and pyridine
(0.3 mL, v/v, 1/2). Reagents were removed under vacuum. The
resulting residue was purified by preparative thin layer chromato-
graphy (CH2Cl2–MeOH, 100 : 5) to afford adenosylhopane
diacetate 14a (0.4 mg, Rf = 0.30), triacetate 14b (0.6 mg, Rf =
0.38)1a,b and tetraacetate 14c (0.8 mg, Rf = 0.66).
Adenosylhopane diacetate (14a). 1H-NMR (300 MHz,
C2HCl3): δ/ppm = 8.30 and 7.95 (2 × 1H, 2s, 2′-H and 8′-H),
6.68 (2H, br. s, –NH2), 6.14 (1H, d, J = 5.5 Hz, 35-H), 5.87 (1H,
dd, J = 5.7, 5.5 Hz, 34-H), 5.38 (1H, dd, J = 5.5, 4.5 Hz, 33-H),
4.15 (1H, td, J = 8.4, 4.3 Hz, 32-H), 2.14 (3H, s, CH3COO–), 2.06
(3H, s, CH3COO–), 0.94 (9H, m, 8β-Me, 14α-Me and 22R-Me),
0.84 (3H, s, 4α-Me), 0.81 (3H, s, 10β-Me), 0.79 (3H, s, 4β-Me),
0.68 (3H, s, 18α-Me).
Conversion of deuteriated adenosylhopane into
bacteriohopanetetrol by a crude cell-free system from
M. organophilum
MS (EI, direct inlet, positive mode 70 eV): m/z = 746 (M+,
9%), 686 (M+ − AcOH, 6%), 626 (M+ − 2AcOH, 24%), 595 (6%),
538 (15%), 491 (5%), 389 (17%), 367 (M+ − CH2CO − side
chain, 9%), 191 (ring C cleavage, 10%), 136 ([adenine + H]+,
100%).
A solution of deuteriated adenosylhopane 2-D (200 μg) in THF
(500 µL) was added to the supernatant (5 mL) described above
in a glass screw cap vial. After addition of a solution of NADPH
(1.5 mg) in water (100 µL), the sample was incubated at 30 °C
by shaking (200 rpm). An additional portion of the cofactor
solution (100 µL) was added after 4 h incubation and the
sample was incubated for another 20 h. After lyophilisation,
the residue was directly acetylated overnight at room tempera-
ture with Ac2O–pyridine (1 : 2 v/v, 3.9 mL). The reaction
mixture was filtered over cotton and washed with toluene (3 ×
2 mL) and CH2Cl2 (3 × 2 mL). The combined filtrates were con-
centrated in vacuo, and the residue was subjected to prepara-
tive TLC (cyclohexane–ethyl acetate, 3 : 7) to give an apolar
fraction (Rf > 0.7) containing diploptene, diplopterol and
acetylated bacteriohopanetetrol. This less polar fraction was
further separated by TLC (cyclohexane–ethyl acetate, 5 : 1),
Adenosylhopane triaacetate (14b). 1H-NMR (300 MHz,
C2HCl3): δ/ppm = 8.87 (1H, s, NHCOCH3), 8.69 and 8.06
(2 × 1H, 2s, 2′-H and 8′-H), 6.14 (2H, d, J = 5.5 Hz, 35-H), 5.94
(1H, dd, J = J = 5.5 Hz, 34-H), 5.42 (1H, dd, J = 5.5, 4.5 Hz,
33-H), 4.17 (1H, td, J = 8.4, 4.3 Hz, 32-H), 2.60 (3H, s,
CH3CONH–), 2.15 (3H, s, CH3COO–), 2.06 (3H, s, CH3COO–),
0.94 (9H, m, 8β-Me, 14α-Me and 22R-Me), 0.84 (3H, s, 4α-Me),
0.81 (3H, s, 10β-Me), 0.79 (3H, s, 4β-Me), 0.68 (3H, s, 18α-Me).
MS (EI, direct inlet, positive mode 70 eV): m/z = 788 (M+,
2%), 610 (M+ − N,N-acetyladenine, 4%), 595 (ring C cleavage,
2%), 491 (1%), 389 (6%), 367 (M+ − CH2CO − side chain, 4%),
191 (ring C cleavage, 11%), 178 ([N-acetyladenine + H]+, 100%).
Adenosylhopane tetraacetate (14c). 1H-NMR (300 MHz,
C2HCl3): δ/ppm = 8.97 and 8.21 (2 × 1H, 2s, 2′-H and 8′-H),
6.18 (2H, d, J = 5.5 Hz, 35-H), 5.96 (1H, dd, J = J = 5.5 Hz, 34-
H), 5.42 (1H, dd, J = 5.5, 4.5 Hz, 33-H), 4.21–4.17 (1H, m, 32-
H), 2.37 (6H, s, 2 × CH3CONH–), 2.16 (3H, s, CH3COO–), 2.09
(3H, s, CH3COO–), 0.94 (6H, br. s, 8β-Me and 14α-Me), 0.94
(3H, d, J = 6.1 Hz, 22R-Me), 0.84 (3H, s, 4α-Me), 0.81 (3H, s,
10β-Me), 0.79 (3H, s, 4β-Me), 0.69 (3H, s, 18α-Me).
yielding pure bacteriohopanetetrol tetraacetate (0.16 < Rf
<
0.20), which was analysed by GC-EIMS.
Isolation of natural abundance bacteriohopanetetrol from
M. organophilum or from cell-free system unbroken cells and
cell debris pellet
MS (EI, direct inlet, positive mode 70 eV): m/z = 830 (M+, The freeze-dried material (unbroken cells and large cell debris)
0.3%), 788 (M+ − CH2CO), 662 (1%), 610 (M+ − CH2CO − N,N- recovered after the centrifugation yielding the supernatant
diacetyladenine, 8%), 595 (ring C cleavage − CH2CO, 5%), 491 used for the incubations was extracted with CHCl3–CH3OH
(3%), 389 (16%), 367 (M+ − CH2CO − side chain, 8%), 191 (2 : 1, v/v) three times under reflux. The combined extracts
(ring C cleavage, 20%), 178 (100%).
were brought to dryness, acetylated with Ac2O–pyridine (1 : 2,
3404 | Org. Biomol. Chem., 2015, 13, 3393–3405
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Paper
v/v) and the excess of reagents was evaporated in vacuo. The 13 W. Liu, E. Sakr, P. Schaeffer, H. M. Talbot, J. Donisi,
residue was separated by preparative TLC (silica gel, cyclo-
T. Härtner, E. Kannenberg, E. Takano and M. Rohmer,
hexane–ethyl acetate, 3 : 7) to give an apolar fraction (Rf > 0.7)
ChemBioChem, 2014, 15, 2156.
containing diploptene, diplopterol, and acetylated bacterioho- 14 P. Stampf, PhD Thesis, Université de Haute Alsace, Mul-
panetetrol. This mixture was further separated by preparative house, France, 1992.
TLC (silica gel, cyclohexane–ethyl acetate, 5 : 1), yielding pure 15 T. Duvold and M. Rohmer, Tetrahedron, 1999, 55,
bacteriohopanetetrol tetraacetate (0.16 < Rf < 0.20).
9847.
16 W. D. Pan, PhD Thesis, Université de Paris VI, Paris,
France, 2005.
17 W. D. Pan, C. Sung, Y. M. Zhang, G. Y. Liang, P. Sinaÿ and
S. P. Vincent, Chem. – Eur. J., 2007, 13, 1471.
18 S. F. Wnuk and M. J. Robins, Can. J. Chem., 1991, 69,
334.
19 R. H. Grubbs, in Handbook of metathesis, Wiley-VCH, Wein-
heim, Germany, 2003.
20 (a) T. M. Trnka and R. H. Grubbs, Acc. Chem. Res., 2001, 34,
18; (b) A. B. Flynn and W. W. Ogilvie, Chem. Rev., 2007, 107,
4698.
Acknowledgements
This work was supported by a grant from the Foundation
“Frontier Research in Chemistry” (Strasbourg, France). WL
acknowledges financial support from the China Scholarship
Council and the French embassy in Beijing. The authors
express their gratitude to E. Motsch and Dr P. Schaeffer for
EIMS and GC-EIMS measurements, Dr L. Allouche, B. Vincent
and J.D. Sauer for NMR measurements and R. Carrière and Dr
H. Nierengarten for ES-HRMS measurements.
21 (a) A. K. Chatterjee, T. L. Choi, D. P. Sanders and
R. H. Grubbs, J. Am. Chem. Soc., 2003, 125, 11360;
(b) S. J. Connon and S. Blechert, Angew. Chem., Int. Ed.,
2003, 42, 1900.
References
1 (a) S. Neunlist and M. Rohmer, Biochem. J., 1985, 228, 769;
(b) S. Neunlist, P. Bisseret and M. Rohmer, Eur. J. Biochem.,
1988, 171, 245; (c) M. Seemann, P. Bisseret, J. P. Tritz,
A. B. Hooper and M. Rohmer, Tetrahedron Lett., 1999, 40,
1681; (d) J. M. Bravo, M. Perzl, T. Härtner, E. L. Kannenberg
and M. Rohmer, Eur. J. Biochem., 2001, 268, 1323;
(e) M. Eickhoff, D. Birgel, H. M. Talbot, J. Peckman and
A. Kappler, Geobiology, 2013, 11, 268.
22 D. Andrei and S. F. Wnuk, Org. Lett., 2006, 8, 5093.
23 (a) R. S. Ranganathan, G. H. Jones and J. G. Moffatt, J. Org.
Chem., 1974, 39, 290; (b) S. Eppacher, P. K. Bhardwaj,
B. Bernet, J. L. B. Gala, T. Knopfel and A. Vasella, Helv.
Chim. Acta, 2004, 87, 2969.
24 W. J. Dunstan, H. Fazakerley, T. G. Halsall and
E. R. H. Jones, Croat. Chem. Acta, 1957, 29, 173.
25 A. Ensminger, PhD Thesis, Université Louis Pasteur, Stras-
bourg, France, 1974.
26 C. Samoljowicz, M. Bienek, A. Zarecki, R. Kadyrov and
K. Grela, J. Chem. Soc., Chem. Commun., 2008, 6282.
27 V. Voorhees and R. Adams, J. Am. Chem. Soc., 1922, 44,
1397.
28 J. A. Osborn, F. H. Jardine, J. F. Young and G. Wilkinson,
J. Chem. Soc. A, 1966, 1711.
2 H. M. Talbot, M. Rohmer and P. Farrimond, Rapid
Commun. Mass Spectrom., 2007, 21, 880.
3 (a) M. P. Cooke, H. M. Talbot and P. Farrimond, Org.
Geochem., 2008, 39, 1347; (b) M. P. Cooke, H. M. Talbot and
T. Wagner, Org. Geochem., 2008, 39, 965; (c) Y. Xu,
M. P. Cooke, H. M. Talbot and M. J. Simpson, Org.
Geochem., 2008, 40, 79.
4 A. D. Selver, H. M. Talbot, O. Gustafsson, S. Boult and
B. E. van Dongen, Org. Geochem., 2012, 51, 63.
29 R. Crabtree, Acc. Chem. Res., 1979, 12, 331.
30 (a) C. E. Miller, J. Chem. Educ., 1965, 42, 254;
(b) J. L. G. Spears and J. S. Hutchinson, J. Chem. Phys.,
1988, 88, 240.
31 J. W. Hamersma and E. I. Snyder, J. Org. Chem., 1965, 30,
3985.
32 J. Toulouse, PhD Thesis, Université de Strasbourg, Stras-
bourg, France, 2011.
33 J. M. Renoux and M. Rohmer, Eur. J. Biochem., 1985, 151,
405.
5 T. Wagner, W. Kallweit, H. M. Talbot, G. Mollenhauer,
A. Boom and T. Zabel, Org. Geochem., 2014, 67, 85.
6 P. Bisseret and M. Rohmer, J. Org. Chem., 1989, 54, 2958.
7 S. Neunlist and M. Rohmer, J. Chem. Soc., Chem. Commun.,
1988, 830.
8 G. Flesch and M. Rohmer, Eur. J. Biochem., 1988, 175, 405.
9 M. Rohmer, B. Sutter and H. Sahm, J. Chem. Soc., Chem.
Commun., 1989, 1471.
10 M. Rohmer, Pure Appl. Chem., 1993, 65, 1293.
11 A. S. Bradley, A. Pearson, J. P. Sáenz and C. J. Marx, Org.
Geochem., 2010, 41, 1075.
12 P. V. Welander, D. M. Doughty, C. H. Wu, S. Mehay,
R. Summons and D. K. Newman, Geobiology, 2012, 10, 163.
34 H. Budzikiewicz, J. M. Wilson and C. Djerassi, J. Am. Chem.
Soc., 1963, 85, 3688.
35 W. C. Still, M. Kahn and A. Mitra, J. Org. Chem., 1978, 43,
2923.
36 S. Hestrin and M. Schramm, Biochem. J., 1954, 58, 345.
This journal is © The Royal Society of Chemistry 2015
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