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Applied and Environmental Microbiology, January 1999, p. 221-230, Vol. 65, No. 1
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
Jean-Francois
Rontani,1,*
Patricia C.
Bonin,1 and
John K.
Volkman2
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
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ABSTRACT |
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.
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INTRODUCTION |
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).
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MATERIALS AND METHODS |
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 
270°C in the presence of glycerol (20%
[vol/vol]). Bench cultures were made on solid tryptose blood agar
medium (43).
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 (
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.)
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 ml1 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.
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.
Structural assignments were based on interpretation of mass
spectrometric fragmentations and confirmed by comparison of retention times and mass spectra with those of authentic compounds. Quantitative determinations were based on integrator data that had been calibrated by using external standards.
Gas chromatography-electron impact mass spectrometry analyses were
carried out with an HP 5890 (series II plus) gas chromatograph
connected to an HP 5972 mass spectrometer (Hewlett-Packard). The
following operating conditions were used: a capillary column 30
m
by 0.25 mm (inner diameter) was coated with HP 5 (Hewlett-Packard),
and
the oven temperature was programmed from 60 to 130°C at 30°C
min
1 and then from 130 to 300°C at 4°C
min
1. The carrier gas (He) pressure was maintained at
1.05 × 10
5 Pa until the end of the temperature program and
then programmed
from 1.04 × 10
5 to 1.5 × 10
5 Pa at 0.04 × 10
5 Pa
min
1. 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.
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RESULTS AND DISCUSSION |
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.
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TABLE 1.
Metabolites detected during growth of the different
strains on 6,10,14-trimethylpentadecan-2-one (compound 1)
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The production of compounds 2 to 5 can be attributed to an oxidation
sequence involving the transformation of ketone 1 to
4,8,12-trimethyltridecan-1-ol acetate (compound 2).
Subsequent
hydrolysis of this ester affords
4,8,12-trimethyltridecan-1-ol
(compound 3), which can be metabolized
(after oxidation to the
corresponding acid 4) via a classical
-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).
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FIG. 1.
Proposed pathways for the production of isoprenoid wax
esters during the metabolism of 6,10,14-trimethylpentadecan-2-one. (The
processes for formation of esters 16 and 17 were omitted in order to
simplify the scheme.)
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If (
E)-4,8,12-trimethyltridec-2-enoic acid (compound 5)
constitutes a classical intermediate in the
-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 OsO
4 (
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 C
18 
-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
[C
nH
2n]
· + 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.
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FIG. 2.
Electron impact mass spectra of
(E)-4,8,12-trimethyltridecyl-4,8,12-trimethyltridec-3-enoate
(compound 14) (A),
4,8,12-trimethyltridecyl-4,8,12-trimethyltridecanoate (compound 11)
(B), and 6,10,14-trimethylpentadec-2-yl-4,8,12-trimethyl-tridecanoate
(compound 15) (C).
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Such wax esters had not been previously detected during studies of the
aerobic biodegradation of 6,10,14-trimethylpentadecan-2-one
(compound
1) (
41,
43). This can be explained by (i) the relatively
minor amounts of these compounds produced by
Acinetobacter
sp.
strain PHY9 and
Marinobacter sp. strain CAB (DSMZ 11874)
(Table
1) and (ii) the use of a gas chromatograph having a heated
splitless
injector during the earlier studies, which strongly
discriminates
against such high-molecular-weight compounds. Previous
attempts
to analyze phytyl esters by gas chromatography have, in most
cases,
not been successful. Gellerman et al. (
25) reported
that phytyl
esters decomposed when analyzed on OV-1 (methyl
silicone)-packed
columns, even though palmityl phytenate and esters of
phytanic
acid could be analyzed satisfactorily. Similar results were
obtained
by Withers and Nevenzel (
51).
The structures of these esters suggest that they are formed by
esterification between acids and alcohols produced during the
metabolism of ketone 1 by the different bacterial strains (Fig.
1). The
formation of these compounds implies the presence of a
very
active esterase system (especially in the case of
P. nautica [IP85/617] and
M. hydrocarbonoclasticus
[ATCC 49840]), since the
experimental conditions exclude a
chemical esterification. Indeed,
these esters were lacking in sterile
controls, and they were not
formed during the workup procedure. This
active esterase system
is probably the same as that involved during
hydrolysis of 4,8,12-trimethyltricecan-1-ol
acetate (compound 2) to
4,8,12-trimethyltridecan-1-ol (compound
3) and acetic acid. This
hypothesis is supported by the fact that
alcohol 3 constitutes the
alcohol moiety of most of the wax esters
identified (Table
1).
Although
Marinobacter sp. strain CAB (DSMZ 11874)
(
43),
P. nautica (IP85/617), and
M. hydrocarbonoclasticus (ATCC 49840)
are able to
grow on 6,10,14-trimethylpentadecan-2-one (compound
1) under
denitrifying conditions, they failed to produce isoprenoid
wax esters
under these conditions. This can be attributed to the
fact that (i) the
enzymatic oxidation of ketones by way of an
ester intermediate cannot
operate in the absence of oxygen, and
(ii) the esterases involved
during this process are generally
inducible enzymes (
9).
It is generally considered that wax esters constitute storage energy
components of marine bacteria (
3). Previous work
(
22)
has demonstrated that the amount of wax esters produced
increases
considerably in N-limited cultures under conditions of low
growth
rate, where carbon and energy are in excess. In the case of
P. nautica (IP85/617), the wax ester content increased by
approximately
20-fold in the N-limited culture compared with that in
the nonlimited
one (Fig.
3), while the
growth yield decreased slightly (9.5 ×
10
6 cells/mg
in the nonlimited culture and 3.3 × 10
6 cells/mg in
the N-limited culture). It is interesting to note
that the ammonium
concentration used in the N-limited culture
(0.1 mM) corresponds to
conditions often found in marine sediments
(
36). We suggest,
therefore, that the formation of isoprenoid
wax esters might be favored
in such environments when 6,10,14-trimethylpentadecan-2-one
or phytol
is available.
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FIG. 3.
Partial total ion chromatograms showing the wax ester
region of the extracts obtained after growth of P. nautica
on 6,10,14-trimethylpentadecan-2-one (compound 1) with ammonium ion
concentrations of 56 mM (A) and 0.1 mM (B). IS, internal standard
(triacontane).
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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).
Concurrently, the phytol can be metabolized via
(
E)-phytenic acid (compound 20) by two different
pathways (Fig.
4). The first
pathway
involves isomerization to (
Z)-phytenic acid (compound
19)
and subsequent alternating
-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).
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FIG. 4.
Proposed pathways for the production of isoprenoid wax
esters during the metabolism of phytol. (The processes for formation of
esters 28 and 29 were omitted in order to simplify the scheme.)
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The second pathway consists of a hydrogenation to
3,7,11,15-tetramethylhexadecanoic acid (phytanic acid)
(compound 21), followed
by
-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.
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FIG. 5.
Electron impact mass spectra of
(E)-3,7,11,15-tetramethylhexadec-2-enyl-3,7,11,15-tetramethylhexadecanoate
(phytylphytanate) (compound 24) (A),
(E,E)-3,7,11,15-tetramethylhexadec-2-enyl-3,7,11,15-tetramethylhexadec-2-enoate
(phytylphytenate) (compound 26) (B), and
3,7,11,15-tetramethylhexadec-2-enyl-4,8,12-trimethyltridecanoate
(compound 27) (C).
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Phytyl esters have been detected in a few recent sediments (
8,
15), but few data are available. Proposed sources have
included mosses (
18), bryophytes (
12,
25),
dinoflagellates
(
51), or zooplankton (
45), but in
most cases, no evidence
was presented to substantiate this. The results
obtained in the
present study show that the aerobic bacterial
degradation of phytol
might constitute another potential source of
these compounds in
sediments. It is generally considered that the major
proportion
of phytol released from chlorophyll
a is degraded
before it can
be incorporated into the anoxic region of the sediments
(
30).
Consequently, aerobic bacterial degradation processes,
which must
play an important role in this disappearance, could produce
significant
amounts of phytyl esters in the water column and in the
oxic zone
of the
sediments.
During incubation of [U-
14C]phytol in sediments, Brooks
and Maxwell (
10) observed the formation of phytyl
esters and suggested
that esterification of the introduced phytol
is brought about
by enzymatic processes, possibly microbial. Our
results strongly
support this hypothesis and are also in good agreement
with those
of Volkman and Maxwell (
49), who said in their
review of acyclic
isoprenoid compounds that esterification may be a
significant
process in microbially active
sediments.
Phytyl esters have been reported in cultures of a few marine microalgae
(
21,
51), and small amounts are also present in
senescent cultures of the green microalga
Tetraselmis chui
(
48a).
These limited examples suggest that microalgae are
not likely
to be major sources of phytyl or other wax esters in
sediments.
However, there are a variety of other sources, as discussed
previously,
and amounts can be large in some zooplankton. Zooplankton
are
a major source of wax esters in sedimenting particles in marine
ecosystems (
50), and even though phytyl esters have yet to
be
reported, one might still expect significant contributions to
some
sediments. In addition, our results suggest that they are
also formed
in sediments by bacterial action and that this process
might be
enhanced at low ammonium concentrations. Despite these
observations, phytylphytenate and phytylphytanate are generally
present
in sediments at relatively low concentrations. We believe
that
this is probably due to their rapid hydrolysis during early
diagenesis. Indeed, de Leeuw et al. (
18) analyzed a lake
sediment
which received inputs of the moss
Fontinalis
antipyretica (in
which phytylphytenate is a major wax ester) and
failed to detect
this ester. They concluded that a rapid hydrolysis of
the ester
occurred in the detritus, resulting in its absence in the
underlying
sediment. However, it is interesting to note that, despite
the
occurrence of these hydrolytic processes, Cranwell
(
15) detected
phytylphytenate in lacustrine sediments up
to 40,000 years old.
Similarly, esters of sterols are thought to be
more stable in
sediments than the corresponding free compounds
(
19), so that,
clearly under some conditions, esterification
can enhance the
preservation potential of labile compounds such as
phytol.
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 |
*
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.
| |
REFERENCES |
| 1.
|
Aasen, A. J.,
H. H. Hofstetter,
B. T. R. Iyengar, and R. T. Holman.
1971.
Identification and analysis of wax esters by mass spectrometry.
Lipids
6:502-507.
|
| 2.
|
Ahlquist, L.,
G. Bergström, and C. Liljenberg.
1977.
Acyclic diterpene alcohols: occurrence and synthesis of geranylcitronellol, phytol and geranylgeraniol.
Prog. Chem. Fats Other Lipids
16:231-255.
|
| 3.
|
Alvarez, H. M.,
O. H. Pucci, and A. Steinbüchel.
1997.
Lipid storage compounds in marine bacteria.
Appl. Microbiol. Biotechnol.
47:132-139.
|
| 4.
|
Bacchin, P.,
A. Robertiello, and A. Viglia.
1974.
Identification of n-decane oxidation products in Corynebacterium cultures by combined gas chromatography-mass spectrometry.
Appl. Microbiol.
28:737-741[Medline].
|
| 5.
|
Baker, E. W., and G. D. Smith.
1974.
Pleistocene changes in chlorophyll pigments, p. 649-660.
In
B. Tissot, and F. Bienner (ed.), Advances in organic geochemistry 1973. Editions Technip, Paris, France.
|
| 6.
|
Baumann, P., and L. Baumann.
1981.
The gram negative eubacteria genus Protobacterium, Beneckae, Alteromonas, Pseudomonas and Alcaligenes, p. 1302-1330.
In
P. S. Mortimer, et al. (ed.), The prokaryotes: a handbook on habitats, isolation and identification of bacteria. Springer-Verlag, Berlin, Germany.
|
| 7.
|
Bonin, P.,
M. Gilewicz, and J.-C. Bertrand.
1987.
Denitrification by a marine bacterium, Pseudomonas nautica strain 617.
Ann. Inst. Pasteur Microbiol.
138:371-383[Medline].
|
| 8.
|
Boon, J. J., and J. W. de Leeuw.
1979.
The analysis of wax esters, very long mid-chain ketones and sterol ethers isolated from Walvis Bay diatomaceous ooze.
Mar. Chem.
7:117-132.
|
| 9.
|
Britton, L. N.,
J. M. Brand, and A. J. Markovetz.
1974.
Source of oxygen in the conversion of 2-tridecanone to undecyl acetate by Pseudomonas cepacia and Nocardia sp.
Biochim. Biophys. Acta
369:45-49[Medline].
|
| 10.
|
Brooks, P. W., and J. R. Maxwell.
1974.
Early stage fate of phytol in a recently-deposited lacustrine sediment, p. 977-991.
In
B. Tissot, and F. Bienner (ed.), Advances in organic geochemistry 1973. Editions Technip, Paris, France.
|
| 11.
|
Brooks, P. W.,
J. R. Maxwell, and R. L. Patience.
1978.
Stereochemical relationships between phytol and phytanic acid, dihydrophytol and C18 ketone in recent sediments.
Geochim. Cosmochim. Acta
42:1175-1180.
|
| 12.
|
Buchanan, M. S.,
T. Hashimoto, and Y. Asakawa.
1996.
Phytyl esters and phaeophytins from the hornwort Megaceros flagellaris.
Phytochemistry
41:1373-1376.
|
| 13.
|
Cantwell, S. G.,
E. P. Lau,
D. S. Watt, and R. R. Fall.
1978.
Biodegradation of acyclic isoprenoids by Pseudomonas species.
J. Bacteriol.
135:324-333[Abstract/Free Full Text].
|
| 14.
|
Cason, J., and D. W. Graham.
1965.
Isolation of isoprenoid acids from a California petroleum.
Tetrahedron
21:471-483.
|
| 15.
|
Cranwell, P. A.
1986.
Esters of acyclic and polycyclic isoprenoid alcohols: biochemical markers in lacustrine sediments.
Org. Geochem.
10:891-896.
|
| 16.
|
Csupor, L.
1971.
Das Phytol in vergilbten Blättern.
Planta Med.
19:37-41.
|
| 17.
|
de Leeuw, J. W.,
V. A. Correia, and P. A. Schenck.
1974.
On the decomposition of phytol under simulated geological conditions and in top layer of natural sediments, p. 993-1004.
In
B. Tissot, and F. Bienner (ed.), Advances in organic geochemistry 1973. Editions Technip, Paris, France.
|
| 18.
|
de Leeuw, J. W.,
W. I. C. Rijpstra,
J. J. Boon,
F. de Lange, and P. A. Schenck.
1977.
The relationship between lipids from Fontinalis antipyretica, its detritus and the underlying sediment: the fate of wax esters and sterol esters, p. 141-147.
In
H. L. Golterman (ed.), Interactions between sediments and freshwater. Junk, The Hague, The Netherlands.
|
| 19.
|
de Leeuw, J. W.,
W. I. C. Rijpstra,
P. A. Schenck, and J. K. Volkman.
1983.
Free, esterified and residual bound sterols in Black Sea Unit I sediments.
Geochim. Cosmochim. Acta
47:455-465.
|
| 20.
|
Donoghue, N. A.,
D. B. Norris, and P. W. Trudgill.
1976.
The purification and properties of cyclohexanone oxygenase from Nocardia globurela CL1 and Acinetobacter NCIB 9871.
Eur. J. Biochem.
63:175-192[Medline].
|
| 21.
|
Findlay, J. A., and A. D. Patil.
1984.
Antibacterial constituents of the diatom Navicula delognei.
J. Nat. Prod.
47:815-818[Medline].
|
| 22.
|
Fixter, L. M.,
M. N. Nagi,
J. G. McCormack, and C. A. Fewson.
1986.
Structure, distribution and function of wax esters in Acinetobacter calcoaceticus.
J. Gen. Microbiol.
132:3147-3157.
|
| 23.
|
Forney, F. W., and A. J. Markovetz.
1970.
Subterminal oxidation of aliphatic hydrocarbons.
J. Bacteriol.
102:281-282[Abstract/Free Full Text].
|
| 24.
|
Gauthier, M. J.,
B. Lafay,
R. Christen,
L. Fernandez,
M. Acquaviva,
P. Bonin, and J.-C. Bertrand.
1992.
Marinobacter hydrocarbonoclasticus gen. nov., sp. nov., a new, extremely halotolerant, hydrocarbon-degrading marine bacterium.
Int. J. Syst. Bacteriol.
42:568-576[Abstract/Free Full Text].
|
| 25.
|
Gellerman, J. L.,
W. H. Anderson, and H. Schlenk.
1975.
Synthesis and analysis of phytyl and phytenoyl wax esters.
Lipids
10:656-661[Medline].
|
| 26.
|
Gillan, F. T.,
P. D. Nichols,
R. B. Johns, and H. J. Bavor.
1983.
Phytol degradation by marine bacteria.
Appl. Environ. Microbiol.
45:1423-1428[Abstract/Free Full Text].
|
| 27.
|
Grossi, V.
1996.
Formation et devenir des phytadiènes dans le milieu marin: implication de ces hydrocarbures dans les processus de dégradation des chlorophylles à chaîne phytyle. Ph.D. thesis.
University of Aix-Marseille II, Marseille, France.
|
| 28.
|
Hansen, R. P.
1980.
Phytol: its metabolic products and their distribution. A review.
N. Z. J. Sci.
23:259-275.
|
| 29.
|
Ikan, R.,
M. J. Baedecker, and I. R. Kaplan.
1973.
C18-isoprenoid ketone in recent marine sediment.
Nature
244:154-155.
|
| 30.
|
Johns, R. B.,
F. T. Gillan, and J. K. Volkman.
1980.
Early diagenesis of phytyl esters in a contemporary temperate intertidal sediment.
Geochim. Cosmochim. Acta
44:183-188.
|
| 31.
|
Kaneshiro, T.,
L. K. Nakamura,
J. J. Nicholson, and M. O. Bagby.
1996.
Oleyl oleate and homologous wax esters synthesized coordinately from oleic acid by Acinetobacter and coryneform strains.
Curr. Microbiol.
32:336-342[Medline].
|
| 32.
|
Klug, M. J., and A. J. Markovetz.
1971.
Utilization of aliphatic hydrocarbons by micro-organisms.
Adv. Microbiol. Physiol.
5:1-43[Medline].
|
| 33.
|
McCloskey, J. A., and M. J. McClelland.
1965.
Mass spectra of O-isopropylidene derivatives of unsaturated fatty esters.
J. Am. Chem. Soc.
87:5090-5093.
|
| 34.
|
McLafferty, F. W.
1980.
Mass spectra of ethyl and higher esters, p. 203-205.
In
N. J. Turro (ed.), Interpretation of mass spectra. University Science Books, Mill Valley, Calif.
|
| 35.
|
Mize, C. E.,
J. Avigan,
D. Steinberg,
R. C. Pittman,
H. M. Fales, and G. W. A. Milne.
1969.
A major pathway for the mammalian oxidative degradation of phytanic acid.
Biochim. Biophys. Acta
176:720-739[Medline].
|
| 36.
|
Omnes, P.
1996.
Mise en évidence de la voie de réduction dissimilatrice du nitrate en ammonium. Application in situ aux systèmes particulaires et sédimentaires en Méditerranée. Ph.D. thesis.
University of Aix-Marseille II, Marseille, France.
|
| 37.
|
Patrick, M. A., and P. R. Dugan.
1974.
Influence of hydrocarbons and derivatives on the polar lipid fatty acids of an Acinetobacter isolate.
J. Bacteriol.
119:76-81[Abstract/Free Full Text].
|
| 38.
|
Platen, H., and B. Schink.
1989.
Anaerobic degradation of acetone and higher ketones via carboxylation by newly isolated denitrifying bacteria.
J. Gen. Microbiol.
135:883-891[Abstract/Free Full Text].
|
| 39.
|
Rambeloarisoa, E.,
J.-F. Rontani,
G. Giusti,
Z. Duvnjak, and J.-C. Bertrand.
1984.
Degradation of crude oil by a mixed culture of bacteria isolated from surface foams.
Mar. Biol.
83:69-81.
|
| 40.
|
Rontani, J.-F.,
I. Combe, and P. J.-P. Giral.
1990.
Abiotic degradation of free phytol in the water column: a new pathway for the production of acyclic isoprenoids in the marine environment.
Geochim. Cosmochim. Acta
54:1307-1313.
|
| 41.
|
Rontani, J.-F., and M. Acquaviva.
1993.
The aerobic bacterial metabolism of phytol in seawater: temperature dependence of an abiotic step and its consequences.
Chemosphere
26:1513-1525.
|
| 42.
|
Rontani, J.-F.,
D. Raphel, and P. Cuny.
1996.
Early diagenesis of intact and photooxidized chlorophyll phytyl chain in a recent temperate sediment.
Org. Geochem.
24:825-832.
|
| 43.
|
Rontani, J.-F.,
M. J. Gilewicz,
V. Michotey,
T. L. Zheng,
P. C. Bonin, and J.-C. Bertrand.
1997.
Aerobic and anaerobic metabolism of 6,10,14-trimethylpentadecan-2-one by a denitrifying bacterium isolated from marine sediments.
Appl. Environ. Microbiol.
63:636-643[Abstract].
|
| 44.
|
Russell, N. J., and J. K. Volkman.
1980.
The effect of growth temperature on wax ester composition in the psychrophilic bacterium Micrococcus cryophilis ATCC 15174.
J. Gen. Microbiol.
118:131-141.
|
| 45.
|
Sargent, J. R., and S. Falk-Petersen.
1981.
Ecological investigations on the zooplankton community in Balsfjorden, Northern Norway.
Mar. Biol.
62:131-137.
|
| 46.
|
Seubert, W.
1960.
Degradation of isoprenoid compounds by microorganisms. I. Isolation and characterization of an isoprenoid-degrading bacterium, Pseudomonas citronellolis n. sp.
J. Bacteriol.
79:426-434[Free Full Text].
|
| 47.
|
Shum, A. C., and A. J. Markovetz.
1974.
Specificity and induction of undecyl acetate esterase from Pseudomonas cepacia grown on 2-tridecanone.
J. Bacteriol.
118:890-897[Abstract/Free Full Text].
|
| 48.
|
Simoneit, B. R. T., and A. L. Burlingame.
1974.
Ketones derived from the oxidative degradation of Green River formation oil shale kerogen, p. 191-201.
In
B. Tissot, and F. Bienner (ed.), Advances in organic geochemistry 1973. Editions Technip, Paris, France.
|
| 48a.
| Volkmann, J. K. Unpublished results.
|
| 49.
|
Volkman, J. K., and J. R. Maxwell.
1986.
Acyclic isoprenoids as biological markers, p. 1-46.
In
R. B. Johns (ed.), Biological markers in the sedimentary record. Elsevier, Amsterdam, The Netherlands.
|
| 50.
|
Wakeham, S. G.,
J. W. Farrington, and J. K. Volkman.
1983.
Fatty acids, wax esters, triacylglycerols and alkyldiacylglycerols associated with particles collected in sediment traps in the Peru upwelling, p. 185-197.
In
M. Bjorøy, et al. (ed.), Advances in organic geochemistry 1981. John Wiley and Sons, Chichester, United Kingdom.
|
| 51.
|
Withers, N. W., and J. C. Nevenzel.
1977.
Phytyl esters in a marine dinoflagellate.
Lipids
12:989-993.
|
Applied and Environmental Microbiology, January 1999, p. 221-230, Vol. 65, No. 1
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Copyright © 1999, American Society for Microbiology. All rights reserved.
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Abstract
Introduction
