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Previous Article | Next Article 
Applied and Environmental Microbiology, January 1999, p. 221-230, Vol. 65, No. 1
Laboratoire d'Océanographie et de
Biogéochimie, UMR 6535, Centre d'Océanologie de Marseille,
OSU, Campus de Luminy, 13288 Marseille,
France,1 and
CSIRO Division of
Marine Research, Hobart, Tasmania 7001, Australia2
Received 4 May 1998/Accepted 6 October 1998
This paper describes the production of isoprenoid wax esters during
the aerobic degradation of 6,10,14-trimethylpentadecan-2-one and phytol by four bacteria (Acinetobacter sp. strain PHY9,
Pseudomonas nautica [IP85/617],
Marinobacter sp. strain CAB [DSMZ 11874], and
Marinobacter hydrocarbonoclasticus [ATCC 49840])
isolated from the marine environment. Different pathways are
proposed to explain the formation of these compounds. In the case
of 6,10,14-trimethylpentadecan-2-one, these esters result from the
condensation of some acidic and alcoholic metabolites produced during
the biodegradation, while phytol constitutes the alcohol moiety of most
of the esters produced during growth on this isoprenoid alcohol. The
amount of these esters formed increased considerably in N-limited
cultures, in which the ammonium concentration corresponds to conditions
often found in marine sediments. This suggests that the bacterial
formation of isoprenoid wax esters might be favored in such
environments. Although conflicting evidence exists regarding the
stability of these esters in sediments, it seems likely that, under
some conditions, bacterial esterification can enhance the preservation
potential of labile compounds such as phytol.
The C20 isoprenoid
alcohol phytol usually occurs in an esterified form as the side chain
of chlorophyll a; it is generally considered to be the most
abundant acyclic isoprenoid compound in the biosphere (49).
In sediments, the fate of most of the deposited phytol, as with other
labile lipids, is remineralization to CO2. However, the
ester bond between phytol and the tetrapyrrolic macrocycle can resist
hydrolysis, as shown by the isolation of intact phytyl esters from
sediments several million years old (5). Small amounts of
free phytol are produced during early diagenesis in recent sediments
(30, 42), and there are numerous reports of
6,10,14-trimethylpentadecan-2-one in sediments (17, 29, 48),
which can be produced from free phytol, phytane, pristane, and
chlorophyll by various pathways (43).
Phytol and some of its metabolites have been identified in both the
alcohol and the fatty acid moieties of some naturally occurring wax
esters (28). Phytyl esters occur in higher plants (16), bryophytes (12, 25), mosses
(18), dinoflagellates (51) and marine zooplankton
(45). In bacteria of the genus Acinetobacter, wax
esters are generally considered to be energy storage components
(3, 22). Esters containing phytol have also been detected in
some marine (8) and lacustrine sediments (15),
but their origin has not been satisfactorily explained.
Some incubation experiments (10, 11) have suggested that the
esterification of phytol may be a significant process in microbially
active sediments. However, the bacterial production of isoprenoid wax
esters has not been reported. In this paper, we report the formation of
isoprenoid wax esters during aerobic growth of four marine
bacteria: Acinetobacter sp. strain PHY9, Pseudomonas
nautica (IP85/617), Marinobacter sp. strain CAB (DSMZ 11874), and Marinobacter hydrocarbonoclasticus (ATCC 49840)
when grown on free phytol and 6,10,14-trimethylpentadecan-2-one, which are two isoprenoid compounds widely distributed in marine sediments (43, 49).
Description of the strains.
Acinetobacter sp. strain
PHY9, P. nautica (IP85/617), M. hydrocarbonoclasticus (ATCC 49840), and Marinobacter
sp. strain CAB (DSMZ 11874) were used in this study. These strains had
been isolated in our laboratory from hydrocarbon-polluted marine
coastal sediments and foams collected from different sites in Golf of
Fos (Mediterranean Sea, France) and generally deposited in culture
collections. The identification and description of these strains can be
found elsewhere (7, 24, 39, 43). For our experiments, the
strains were maintained at Growth conditions.
The basic growth medium consisted of
autoclaved artificial seawater (6) (ASW) supplemented with
iron sulfate (0.1 mM), potassium phosphate (0.33 mM), and
6,10,14-trimethylpentadecan-2-one or phytol (3 mM) as the source of
carbon and energy ( Bacterial numeration.
Cultures at stationary phase were
fixed with formaldehyde to a final concentration of 2% and
refrigerated until needed. The samples were then diluted with filtered
(0.2-µm pore size) (Whatman, no. 7182002) ASW and gently sonicated in
a Branson 2200 ultrasonic bath for 5 min. The samples were then
vortexed for 15 s and incubated with DAPI
(4',6-diamidine-2'-phenylindole dihydrochloride) (Boehringer Mannheim)
to a final concentration of 2 µg ml Isolation and characterization of bacterial metabolites.
At
the end of the growth period, the contents of the flasks were acidified
with hydrochloric acid (pH 1) and continuously extracted with
chloroform overnight. The extracts thus obtained were then dried with
anhydrous Na2SO4, filtered, and concentrated by
means of rotary evaporation. After evaporation of the solvent, the
residues were taken up in ethyl acetate (1 ml per mg of residue) containing BSTFA
[N,O-bis(trimethylsilyl)trifluoroacetamide]
(100 µl), allowed to silylate at 40°C for 30 min, and analyzed by
gas chromatography-electron impact mass spectrometry.
0099-2240/99/$04.00+0
Copyright © 1999, American Society for Microbiology. All rights reserved.
Production of Wax Esters during Aerobic Growth of
Marine Bacteria on Isoprenoid Compounds
ABSTRACT
Top
Abstract
Introduction
Materials and methods
Results and discussion
References
INTRODUCTION
Top
Abstract
Introduction
Materials and methods
Results and discussion
References
MATERIALS AND METHODS
Top
Abstract
Introduction
Materials and methods
Results and discussion
References
270°C in the presence of glycerol (20%
[vol/vol]). Bench cultures were made on solid tryptose blood agar
medium (43).
medium). For experiments in which growth was N
limited, ammonium chloride was added to the medium to a
concentration of 0.1 mM instead of 60 mM. Aerobic cultures were
maintained in 250-ml Erlenmeyer flasks containing 50 ml of medium and
agitated on a reciprocal shaker at 96 rpm. For each experiment, two
flasks were inoculated: the first for estimation of substrate
degradation and identification of the metabolites and the second for
monitoring bacterial growth. Anaerobic growth experiments were
performed with 125-ml serum flasks containing 70 ml of medium
supplemented with KNO3 (20 mM). Anaerobic conditions were
obtained by flushing nitrogen through the flask for 1 h.
Experiments carried out in the presence of mercuric chloride (10 mg
liter
1) served as abiotic controls. (Autoclaved
sterilization was avoided, since phytol can be easily dehydrated during
such a treatment.)
1 in darkness
for 15 min before filtration on prestained (Irgalen black) membrane
filters (0.2-µm pore size) (Millipore, GTBP). Bacteria were counted
immediately with an epifluorescence microscope equipped with a mercury
lamp, on randomly selected fields, until 20 fields or about 500 bacteria were counted.
1 and then from 130 to 300°C at 4°C
min1. The carrier gas (He) pressure was maintained at
1.05 × 105 Pa until the end of the temperature program and
then programmed from 1.04 × 105 to 1.5 × 105 Pa at 0.04 × 105 Pa
min1. The injector (on column) temperature was 50°C,
the electron energy was 70 eV, the source temperature was 170°C, and
the cycle time was 1.5 s.
Chemicals. (E)-Phytol was isolated from a commercially available mixture of (Z and E)-phytol (Aldrich) by column chromatography on silica gel with n-hexane-ethyl acetate (19/1 [vol/vol]) as the eluant. 6,10,14-Trimethylpentadecan-2-one was produced by oxidation of phytol with KMnO4 in acetone (14). The syntheses of 6,10,14-trimethylpentadecan-2-ol, 4,8,12-trimethyltridecan-1-ol, 4,8,12-trimethyltridecanoic acid, 5,9,13-trimethyltetradecanoic acid, (Z)-3,7,11-trimethyldodec-2-enoic acid, phytenals, and (Z and E)-phytenic acids have been described previously (41, 43). Phytanic acid was obtained by hydrogenation of phytenic acids with Pd-CaCO3 as a catalyst. Pristanic acid was synthesized from dihydrophytol according to the method of Cason and Graham (14). The synthesis of 4,8-dimethylnonanoic acid required four steps: (i) hydrogenation of geranylacetone (Aldrich) in methanol with Pd-CaCO3 as a catalyst, (ii) oxidation of the resultant 6,10-dimethylundecan-2-one with perbenzoic acid in CH2Cl2 (Baeyer-Villiger reaction), (iii) alkaline hydrolysis of the resulting ester to 4,8-dimethylnonan-1-ol, and (iv) oxidation of this alcohol to the corresponding acid with chromic anhydride in acetic acid (35). Acetylation of 4,8,12-trimethyltridecan-1-ol with a mixture of pyridine and acetic anhydride (2:1) gave 4,8,12-trimethyltridecan-1-ol acetate. 4,8,12-Trimethyltridecan-4-olide had been previously isolated from phytadiene photooxidation products and characterized (27).
Wax esters were prepared from alcohols and acids by the procedure described by Gellerman et al. (25).Determination of double-bond position in monounsaturated acid metabolites. The method used to determine double-bond position involved the formation of diols by stereospecific oxidation of double bonds with OsO4 in pyridine-dioxane (1:8 [vol/vol]) (33) and subsequent analyses by gas chromatography-electron impact mass spectrometry of the silylated {dimethyl sulfoxide-BSTFA [5:1 (vol/vol)] for 12 h at 60°C} diols. The double-bond position was obtained from the mass fragmentation patterns of these derivatized compounds.
| |
RESULTS AND DISCUSSION |
|---|
|
Top
Abstract
Introduction
Materials and methods
Results and discussion
References
|
|---|
Metabolism of 6,10,14-trimethylpentadecan-2-one.
Like
Marinobacter sp. strain CAB (DSMZ 11874)
(43) and Acinetobacter sp. strain PHY9
(41), P. nautica (IP85/617) and M. hydrocarbonoclasticus (ATCC 49840) were able to grow
on 6,10,14-trimethylpentadecan-2-one (compound 1) as the sole carbon
and energy source under aerobic conditions. After 10 days of growth,
the extent of degradation ranged from 50 to 95%, depending on the
strain used. Several isoprenoid metabolites were detected (Table
1). These compounds (which were not found
in sterile controls) were formally identified by comparison of their
retention times and mass spectra with those of reference compounds.
|
-oxidation sequence (Fig. 1). Such
enzymatic oxidation of ketones to esters by microorganisms has been
reported (9, 20, 37). This process is analogous to
Baeyer-Villiger oxidation with peracids and seems to operate for each
of the four strains studied (Table 1).
|
-oxidation sequence, the
presence of (Z and
E)-4,8,12-trimethyltridec-3-enoic acids (compounds 6 and 7)
is more surprising. The position of the double bonds of these acids was
determined by gas chromatography-electron impact mass spectrometry
after oxidation with OsO4 (33) and silylation.
Subsequent hydration of these curious ,
-unsaturated acids affords
4-hydroxy-4,8,12-trimethyltridecanoic acid, which lactonizes easily to 4,8,12-trimethyltridecan-4-olide (compound 8) (Fig. 1).
Small amounts of 6,10,14-trimethylpentadecan-2-ol (compound 9) were
also formed in the experiments, probably by a dehydrogenase (38). The involvement of this "blind alley" pathway
suggests that this process results from nonspecific enzyme activity
that is not related specifically to
6,10,14-trimethylpentadecan-2-one (compound 1)
degradation (32).
As previously described (41, 43), Acinetobacter
sp. strain PHY9 and Marinobacter sp. strain CAB (DSMZ 11874)
also produce 5,9,13-trimethyltetradecanoic acid (compound 10) after
oxidation of the keto-terminal methyl group of the ketone 1 and
subsequent decarboxylation of the resulting C18 
-keto acid.
In addition to these different metabolites, we also detected several
isoprenoid wax esters (compounds 11 to 17) (Table 1). Electron impact
mass spectra of the most abundant esters are given in Fig.
2. Fragment ions of the general formula
[RC(OH)==OH]+ formed after rearrangement of
two hydrogen atoms (34) allow an easy characterization
of the acid moiety of these esters, whereas the alcohol moiety
generally gives
[CnH2n]· + fragment
ions (1). In contrast to compound 15, compounds 11 and 14 give notable molecular ions (M+ ·) (Fig. 2). The lack of
M+ · in the electron impact mass spectrum of compound 15 can be attributed to the presence of a secondary carbon in the position relative to the saturated oxygen atom, which strongly favored
the cleavage of the molecule and thus decreased the abundance of the
molecular ion. Identification of compounds 12 to 14, which were not
synthesized, was based on the characterization of the respective
isoprenoid alcohols and acids obtained after alkaline hydrolysis.
|
|
Metabolism of phytol.
All four strains were able to grow on
phytol as the sole source of carbon and energy under aerobic
conditions, but this compound appeared to be a poorer substrate for
these organisms than 6,10,14-trimethylpentadecan-2-one. This relative
biological recalcitrance can be attributed to the presence of a methyl
group on the carbon 3 of phytol, which prevents classical
-oxidation, requiring an additional strategy, such as -oxidation
(26) or -alkyl group removal (-decarboxymethylation) (13, 46), to allow oxidation to proceed.
Acinetobacter sp. strain PHY9 failed to produce detectable
amounts of isoprenoid wax esters during growth on this substrate. This
is in good agreement with the results described above, since this
organism has the best cell yield and the lowest production of wax
esters when grown on 6,10,14-trimethylpentadecan-2-one (Table 1). The
first step of the bacterial degradation of phytol involves the
transient production of the corresponding aldehyde
(E)-3,7,11,15-tetramethylhexadec-2-enal (phytenal) (compound
18). This labile compound can be converted quickly and abiotically in
seawater to 6,10,14-trimethylpentadecan-2-one (compound 1)
(40). The production of this ketone involves addition of
water to the activated double bond of phytenal followed by a
retro-aldol reaction. In support of this view, we detected ketone 1 and
some of its metabolites (compounds 3, 4, and 11 [described above])
after growth of the three strains on phytol (Table
2).
|
-decarboxymethylation and
-oxidation sequences. The ability of microorganisms to carry out
-decarboxymethylation was originally established by Seubert
(46). The net effect of this process is to replace a methyl
substituent (which prevents -oxidation) with a carbonyl oxygen
(13). In the case of P. nautica (IP85/617) and
Marinobacter sp. strain CAB (DSMZ 11874), the involvement of
such a mechanism is supported by the detection of only the Z
isomer of 3,7,11-trimethyldodec-2-enoic acid (compound 23) (Table 2).
Activation of allylic methyl groups via carboxylation occurs only in
the case of the Z isomers (13).
|
-oxidation to
2-hydroxy-3,7,11,15-tetramethylhexadecanoic acid, which is then
converted to 2,6,10,14-tetramethylpentadecanoic acid (pristanic acid)
(compound 22) by decarboxylation. The pristanic acid (compound
22) thus formed may be subsequently metabolized via classical
-oxidation reactions. This pathway, which was previously proposed by
Gillan et al. (26) during a study of the aerobic degradation
of phytol by bacteria isolated from surface sediments, operates in the
case of the three strains studied (Table 2).
After 10 days of growth, we also detected several isoprenoid wax esters
(compounds 24 to 29 in Table 2) arising from the esterification of
phytol with its acidic metabolites (Fig. 4). The electron impact mass
spectra of some of these compounds are given in Fig.
5. The mass spectrometric
characterization of the acid moiety of phytyl esters is not easy, since
the double bond of phytol considerably decreases the intensity of the
double hydrogen atom rearrangement. In contrast, the fragment ion at
m/z 278 is of great diagnostic value for the
characterization of the phytyl alcohol chain. The presence of a notable
molecular peak only in the electron impact mass spectrum of compound 26 (Fig. 5) seems to indicate that the abundance of M· + of
these wax esters increases with increased unsaturation. Esterification activity in these cultures was not confined to
4,8,12-trimethyltridecan-1-ol (compound 3) and
6,10,14-trimethylpentadecan-2-ol (compound 9); phytol appeared also
to be an excellent substrate. This can be attributed to the well-known
overlapping substrate selectivity of esterases involved during the
enzymatic oxidation of ketones by way of an ester intermediate
(47). The ability of phytol to give acyl esters is supported
by the detection of phytyl esters in several plants (2, 12, 18,
51), which must also possess active esterase systems.
|
Conclusions. The production of linear wax esters in bacteria is well known (44) and has been demonstrated during the growth of bacteria on n-alkanes (4) and oleic acid (3, 22, 31). However, to our knowledge, this study is the first report of the production of wax esters when marine bacteria are grown on isoprenoid substrates. This property seems to be a characteristic of bacteria able to oxidize methyl ketones by way of an ester intermediate (9, 23).
There is a demonstrable need to identify bacterial metabolites that have sufficient structural specificity to act as biological markers for microbial degradation in the aquatic environment. Consequently, our results suggest that it would be useful to search for isoprenoid esters, such as compounds 11 to 17 and 27 to 29 (Tables 1 and 2, respectively), in marine sediments and particulate matter samples.| |
ACKNOWLEDGMENTS |
|---|
This work was supported by grants from the Centre National de la Recherche Scientifique and the Elf Aquitaine Society (Research Groupment HYCAR 1123).
| |
FOOTNOTES |
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* Corresponding author. Mailing address: Laboratoire d'Océanographie et de Biogéochimie, UMR 6535, Centre d'Océanologie de Marseille, OSU, Campus de Luminy, case 901, 13288 Marseille, France. Phone: 33 4 91 82 96 23. Fax: 33 4 91 82 65 48. E-mail: rontani{at}com.univ-mrs.fr.
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Top
Abstract
Introduction
Materials and methods
Results and discussion
References
|
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