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Applied and Environmental Microbiology, December 1999, p. 5484-5492, Vol. 65, No. 12
0099-2240/99/$04.00+0
Copyright © 1999, American Society for Microbiology. All rights reserved.
Biodegradation of Free Phytol by Bacterial
Communities Isolated from Marine Sediments under Aerobic and
Denitrifying Conditions
Jean-François
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 Marine Research,
Hobart, Tasmania 7001, Australia2
Received 30 July 1999/Accepted 4 October 1999
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ABSTRACT |
Biodegradation of (E)-phytol
[3,7,11,15-tetramethylhexadec-2(E)-en-1-ol] by two
bacterial communities isolated from recent marine sediments under
aerobic and denitrifying conditions was studied at 20°C. This
isoprenoid alcohol is metabolized efficiently by these two bacterial
communities via 6,10,14-trimethylpentadecan-2-one and
(E)-phytenic acid. The first step in both aerobic and
anaerobic bacterial degradation of (E)-phytol involves the
transient production of (E)-phytenal, which in turn can be
abiotically converted to 6,10,14-trimethylpentadecan-2-one. Most of the
isoprenoid metabolites identified in vitro could be detected in a fresh
sediment core collected at the same site as the sediments used for the
incubations. Since (E)-phytenal is less sensitive to
abiotic degradation at the temperature of the sediments (15°C), the
major part of (E)-phytol appeared to be biodegraded in situ
via (E)-phytenic acid. (Z)- and
(E)-phytenic acids are present in particularly large
quantities in the upper section of the core, and their concentrations
quickly decrease with depth in the core. This degradation (which takes place without significant production of phytanic acid) is attributed to
the involvement of alternating 
-decarboxymethylation and
-oxidation reaction sequences induced by denitrifiers. Despite the
low nitrate concentration of marine sediments, denitrifying bacteria
seem to play a significant role in the mineralization of
(E)-phytol.
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INTRODUCTION |
Acyclic isoprenoid compounds with 20 or fewer carbon atoms are relatively abundant and widespread components
of marine sediments (8, 50). These compounds, which are
often used as biological markers (50), originate mainly from
the phytyl side chain of chlorophyll a (14);
however, other, minor precursors, such as chlorophyll b and
bacteriochlorophyll a (19), tocopherols
(21), methyltrimethyltridecylchromans (31), and
wax esters (50), are also known. Although 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 (1), appreciable amounts of free
phytol can be detected in recent sediments (24, 41).
Many of the reactions thought to occur in sediments are known to
operate in biological systems. Although some early studies provided
information on the biodegradation of phytol (8, 9, 20), data
concerning these processes were very limited. Recently, however, there
has been a renewal of interest, and the biodegradation of phytol has
been studied under both aerobic (39, 44) and anaerobic
(24) conditions.
Rontani and Acquaviva (39) observed that different routes
can be involved in the aerobic bacterial metabolism of phytol, depending on the temperature. Indeed, the aerobic metabolism of phytol
by Acinetobacter sp. strain PHY9 occurs via a labile
intermediate, (E)-phytenal, which can be degraded
abiotically in seawater to 6,10,14-trimethylpentadecan-2-one. At
relatively high temperatures, this abiotic process is quite rapid and,
consequently, a large proportion of phytol is metabolized via this
C18 ketone. On the other hand, at low temperatures,
(E)-phytenal is much more stable abiotically and phytol is
biodegraded mainly via (E)-phytenic acid. However, in
anaerobic sediment slurries under sulfate-reducing conditions, phytol
was rapidly biodegraded by the mixed bacterial community to phytenes
via phytadiene intermediates (24).
There are many reports in the literature of denitrifying bacterial
strains able to grow on monoterpenes (for a review, see reference
28). Recently, we studied the production of
isoprenoid wax esters by some aerobic and denitrifying bacteria from
our laboratory collection (44). These bacteria metabolized
phytol aerobically via a pathway involving the formation of
(Z)-phytenic acid and subsequent alternating
-decarboxy-methylation and -oxidation reaction
sequences. Denitrifiers are aerobic facultative bacteria that can use
nitrate instead of oxygen as an electron acceptor. When oxygen is
depleted and anaerobic conditions become established, the bacteria can
make use of the same sequence of reactions since these reactions do not
require molecular oxygen (28).
The aim of the present work was to determine if such a pathway operates
in oxic and anoxic marine sediments. For this purpose, we studied the
biodegradation of free phytol by two bacterial communities isolated
from recent marine sediments under aerobic and denitrifying conditions.
Then, to compare our in vitro observations with naturally occurring
processes, the abundances of free isoprenoid compounds in recently
deposited sediments collected at the same site as the sediment used for
the in vitro inoculations were quantified.
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MATERIALS AND METHODS |
Sediment sampling and storage.
The top layer of the sediment
was collected with a manual corer under 9 m of water. The cores
were maintained in bags (containing dry ice) during their
transportation to Marseille, where they were either used immediately as
bacterial inocula or stored at 
20°C until analysis of the free
isoprenoid compounds. Station 35 in Carteau Bay (Gulf of Fos,
Mediterranean Sea) was chosen as the site for this study of the
biodegradation of free phytol. Its suitability for such work was based
on (i) a minor variance in particle size distribution with depth (with
the percentage of particles <63 µm in diameter ranging from 89% at
2 cm to 95% at 10 cm [5]), suggesting minimal
variation of sedimentary conditions over the profile studied (see
reference 30); (ii) the relatively high chlorophyll
concentration (mostly of diatomaceous origin) at the water-sediment
interface; (iii) a strong dissimilative nitrate reduction activity
(5); and (iv) the significant amounts of phytol in the free
form (24, 41). The oxic layer of the sediment was 4 mm thick
(5), and the sedimentation rate in this area is
approximately 0.5 to 1 cm/year (22).
Bacterial isolation and culture.
Two bacterial communities
able to degrade free phytol were isolated from the coastal sediments
described above. One community was obtained under aerobic growth
conditions, and the other was isolated under anaerobic denitrifying
growth conditions in the presence of nitrate as an electron acceptor.
Portions (5 ml) of the upper 2 cm of the sediment samples were used to
inoculate 50-ml volumes of an enrichment medium consisting of
artificial seawater (3) supplemented with iron sulfate (0.1 mM), potassium phosphate (0.33 mM), and phytol (0.15 mM) (as the carbon
source). Aerobic enrichment cultures were incubated in 250-ml
Erlenmeyer flasks at 20°C on a reciprocal shaker. Denitrifying
cultures were incubated in 100-ml serum flasks containing 50 ml of the
above-described medium supplemented with nitrate (10 mM). These flasks
were sealed with rubber stoppers. Anaerobic conditions were obtained by
flushing nitrogen through the flasks for 30 min. The cultures were
magnetically stirred at 20°C. After two transfers of the cultures
into this medium, 1 ml of each enrichment culture was used to initiate
experiments. The aerobic and anaerobic (denitrifying) cultures used the
medium described above and were supplemented with sand (mainly composed of calcareous algal detritus of about 2 mm in diameter; 150 g liter
1) to support bacterial immobilization. Previous
studies have established that most benthic bacteria are not suspended
in interstitial water but are attached to sediment particles
(16). For each anaerobic experiment, two identical growth
media were inoculated: the first for the estimation of phytol
biodegradation and identification of metabolites, and the second for
monitoring nitrate reduction. Sterile control experiments were carried
out in parallel.
Treatment of bacterial cultures.
At the end of the growth
period, the aqueous and solid phases were separated. The aqueous phase
was continuously extracted with chloroform overnight, while the wet
solid phase was extracted ultrasonically with isopropanol-hexane (4:1,
vol/vol) (13). The chloroform and hexane extracts were
combined, dried over anhydrous Na2SO4,
filtered, and concentrated by rotary evaporation to yield the residual
substrate and the neutral metabolites. To recover acidic metabolites,
the isopropanol-water phase was filtered, evaporated under a vacuum,
acidified with HCl (pH 1), and then extracted with chloroform (three
times). Note that the isopropanol was removed by evaporation in order
to avoid the formation of isopropyl esters during acidification.
Treatment of sediments.
The wet sediment slices (2-cm
intervals) were extracted ultrasonically as described above for the
solid phase of the bacterial cultures. The combined neutral and acidic
extracts were dried over anhydrous Na2SO4,
filtered, concentrated by means of rotary evaporation, and then
chromatographed on a 13- by 0.6-cm (internal diameter) wet packed (in
hexane) glass column filled with silica gel (Kieselgel S plus 0.5%
H2O). Three fractions were eluted, one with hexane (100 ml), the second with dichloromethane (100 ml), and the third with
methanol (50 ml); fraction 1 (F1) contained hydrocarbons;
F2 contained alcohols, ketones, aldehydes, and other fairly
polar compounds; and F3 contained carboxylic acids and sugars.
Derivatization.
After evaporation of solvents,
F2 and F3 and the extracts obtained after
treatment of the bacterial cultures were taken up in 400 µl of a
mixture of pyridine and BSTFA
[bis(trimethylsilyl)trifluoroacetamide] (3:1, vol/vol) and
allowed to silylate at 50°C for 1 h. After evaporation to
dryness, the residue was dissolved in ethyl acetate and analyzed by gas
chromatography-mass spectrometry (GC-MS). Isophytol (a tertiary
alcohol) was only partially silylated by this procedure, but
quantitative silylation of this compound could be obtained via reaction
in a mixture of dimethyl sulfoxide and BSTFA (3:1, vol/vol) at 50°C overnight.
Identification and quantification of free isoprenoids.
Free
isoprenoids were identified by comparison of retention times and mass
spectra with those of standards and then quantified (calibration with
external standards) by electron impact GC-MS. For low concentrations,
or in the case of coelutions, quantification was assessed by selected
ion monitoring using the diagnostic ions at m/z
143 for silylated phytol and isophytol, m/z 58 for 6,10,14-trimethylpentadecan-2-one, and M-15 (loss of a methyl
group) for silylated dihydrophytol and phytanic, phytenic,
4,8,12-trimethyltridecanoic, and 5,9,13-trimethyltetradecanoic acids.
GC-MS analyses were carried out with an HP 5890 Series II Plus
capillary gas chromatograph connected to an HP 5972 mass spectrometer (both from Hewlett-Packard). The following equipment and operating conditions were employed: a 30-m by 0.25-mm (internal diameter) capillary column coated with HP 5% phenyl methylsilicon
(Hewlett-Packard), an oven temperature programmed to increase from 60 to 130°C at 30°C min1 and then from 130 to 300°C at
4°C min1, a carrier gas (He) pressure maintained at
1.05 × 105 Pa until the end of the temperature
program and then programmed to increase from 1.05 × 105 to 1.5 × 105 Pa at 0.04 × 105 Pa min1, an injector (on column)
temperature of 50°C, an electron energy of 70 eV, a source
temperature of 170°C, and a cycle time of 1.5 s.
Standard compounds.
(E)-Phytol and isophytol were
purchased from Acros and Interchim, respectively.
6,10,14-Trimethylpentadecan-2-one was obtained by oxidation of phytol
with KMnO4 in acetone (11). (Z)- and (E)-Phytenic acids were synthesized in two steps: (i)
oxidation of a mixture of (Z)- and (E)-phytols
(Aldrich) with CrO3-pyridine in dry methylene chloride
(18), and (ii) oxidation of the resulting phytenals with
sodium chlorite (2). Phytanic acid and dihydrophytol were
obtained by hydrogenation of phytenic acids and phytol, respectively, in methanol with Pd-CaCO3 as a catalyst.
4,8,12-Trimethyltridecanoic acid was synthesized from isophytol by a
previously described procedure (38).
5,9,13-Trimethyltetradecanoic acid was produced from
6,10,14-trimethylpentadecan-2-one (32).
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RESULTS AND DISCUSSION |
Aerobic biodegradation of phytol.
The aerobic bacterial
community isolated from the sediments of Carteau Bay degrades
(E)-phytol very efficiently. We observed 96% degradation
after 10 days of incubation at 20°C, whereas extraction of sterile
controls yielded 93% recovery of the substrate. Several 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.
It was previously demonstrated that the first step of the aerobic
bacterial degradation of (
E)-phytol involves the transient
production of the corresponding aldehyde,
(
E)-3,7,11,15-tetramethylhexadec-2-enal
(phytenal) (compound
1 in Table
1) (
20). This labile compound
can be converted
abiotically in seawater to 6,10,14-trimethylpentadecan-2-one
(compound
2 in Table
1). The production of this ketone involves
addition of water
to the activated double bond of phytenal followed
by a retro-aldol
reaction (
37). In support of this view, we
detected ketone 2 and some of its metabolites after growth of
the aerobic bacterial
community on (
E)-phytol.
6,10,14-Trimethylpentadecan-2-one can be aerobically metabolized by two
different pathways (pathways II and III in Fig.
1).
The production of compounds 3 to 5 can be attributed to an oxidation
sequence involving the transformation
of ketone 2 to 4,8,12-trimethyltridecan-1-ol
acetate (compound 3)
(pathway II in Fig.
1). Subsequent hydrolysis
of this ester results in
4,8,12-trimethyltridecan-1-ol (compound
4), which can be metabolized
(after oxidation to the corresponding
acid [compound 5]) via
classical
-oxidation reaction sequences.
Such enzymatic oxidation of
ketones to esters by different bacteria
(which is analogous to
Baeyer-Villiger oxidation with peracids)
has been observed in other
studies (
7,
15,
33,
44). The
results obtained in the present
work with a bacterial community
isolated from marine sediments show
that microorganisms able to
carry out this sequence of reactions are
widely distributed in
the environment.
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FIG. 1.
Proposed pathways for the metabolism of
(E)-phytol by the aerobic bacterial community. The broken
arrows indicate abiotic processes.
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Due to the presence of traces of 5,9,13-trimethyltetradecanoic acid
(compound 6 in Table
1), we cannot exclude the possibility
of some
involvement of pathway III (Fig.
1) during the metabolism
of ketone 2. This pathway involves oxidation of the keto-terminal
methyl group of
the ketone and subsequent decarboxylation of the
resulting
C
18 
-keto acid to the 5,9,13-trimethyltetradecanoic
acid
(compound 6) (
20). Subsequently, the
-oxidation cycle
can
proceed only for one complete reaction sequence before a metabolic
blockage occurs (
-methyl branch). The assimilation of the resulting
3,7,11-trimethyldodecanoic acid (compound 7) requires the involvement
of an additional strategy, such as
-oxidation (
32),
-decarboxymethylation
(
10,
46), or
-oxidation
(
34). We were not able to find
any evidence of the presence
of branched
-dicarboxylic acids
in these experiments. Although it is
difficult to totally exclude
the possibility that some
-oxidation takes place, with the products
being rapidly assimilated,
we have not included the involvement
of
-oxidation processes during
the metabolism of (
E)-phytol by
our aerobic bacterial
community.
-Oxidation of acid 7 results
in
2-hydroxy-3,7,11-trimethyldodecanoic acid (compound 8), which
is then
converted to 2,6,10-trimethylundecanoic acid (compound
9) by
decarboxylation. The acid metabolite 9 thus formed may be
subsequently
totally metabolized via classical
-oxidation.
3,7,11-Trimethyldodecanoic
acid (compound 7) can also be assimilated
after
-decarboxymethylation
and
-oxidation reactions sequences.
The ability of microorganisms
to carry out the
-decarboxymethylation reaction sequence (Fig.
2) 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 (Fig.
2).
Traces of 6,10,14-trimethylpentadecan-2-ol (compound 10) were also
formed during the incubation (pathway V in Fig.
1), probably
by a
dehydrogenase (
35). The involvement of this "blind
alley"
pathway suggests that this process results from a nonspecific
enzyme activity that is not related specifically to phytol degradation
(
42).
Concurrently, (
E)-phytol is metabolized via
(
E)-3,7,11,15-tetramethylhexadec-2-enoic acid (phytenic
acid; compound 11) by
two different pathways (pathways I and IV in Fig.
1). Pathway
I involves isomerization to (
Z)-phytenic acid
(compound 12 in
Table
1) and subsequent
-decarboxymethylation and
-oxidation
reaction sequences. The involvement of such a mechanism
is supported
by the detection of significant amounts of
(
Z)-phytenic acid (compound
12), since activation of allylic
methyl groups via carboxylation
occurs only in the case of
Z
isomers (
10). The 5,9,13-trimethyltetradecanoic
acid
(compound 6) thus formed is then metabolized by the mechanisms
described above. Pathway IV, which was previously proposed by
Gillan et
al. (
20), consists of a hydrogenation to
3,7,11,15-tetramethylhexadecanoic
acid (phytanic acid) (compound 13)
followed by
-oxidation to
2-hydroxy-3,7,11,15-tetramethylhexadecanoic acid (compound 14),
which
is then converted to 2,6,10,14-tetramethylpentadecanoic
acid (pristanic
acid; compound 15) by decarboxylation. The pristanic
acid (compound 15)
thus formed can be subsequently metabolized
via classical
-oxidation
reactions.
We also detected several isoprenoid wax esters (compounds 16 to 19)
(Table
1) arising from the esterification of (
E)-phytol
with
some of its acidic metabolites (
44). It was previously
demonstrated that the amount of these esters (which constitute
energy
storage components of marine bacteria) increases considerably
in
N-limited cultures, in which the ammonium concentration corresponds
to
conditions often found in marine sediments (
44).
Anaerobic biodegradation of phytol under denitrifying
conditions.
The denitrifying bacterial community isolated from the
sediments of Carteau Bay efficiently degrades (E)-phytol
under anaerobic conditions. We observed 83% degradation after 30 days
of incubation at 20°C, while Grossi et al. (24) observed
80 to 95% degradation of (E)-phytol after 3 months of
incubation under sulfate-reducing conditions at 30°C. The kinetics of
nitrogen oxide production indicates that nitrate was consumed without a
transient accumulation of nitrite. The denitrifers appear to have
higher metabolic capacities, judging by the amounts of metabolites
accumulated at the end of the growth of these two communities. Thus,
Grossi et al. (24) detected relatively large quantities of
phytadienes and phytenes (41 to 74% of the amount of phytol that had
disappeared) after 3 months of incubation of phytol under
sulfate-reduction conditions, whereas the different metabolites
identified at the end of our experiment with the denitrifying community
(Table 1) represent only 2% of the degraded substrate. As aerobic
facultative bacteria, denitrifiers possess the powerful enzymatic
equipment of aerobic bacteria (4) and can use some of these
enzymes (which function without molecular oxygen) under anaerobic
conditions. The high metabolic capacities of these bacteria have been
recently confirmed by Harder and Probian (27). Indeed, these
authors showed that the denitrifying strain 72Chol mineralizes
cholesterol completely to carbon dioxide in the absence of oxygen,
whereas sulfate-reducing bacteria generally catalyze only the reduction
of the sterol double bond (51).
As under aerobic conditions, the first step of the anaerobic
degradation of (
E)-phytol involves the production of
(
E)-phytenal
(compound 1 in Table
1), which is then
abiotically partially
converted to
6,10,14-trimethylpentadecan-2-one (compound 2). Anaerobic
biodegradation of ketone 2 may involve either a carboxylation
reaction
(pathway II in Fig.
3) or hydration of
the enol forms
of this ketone (pathways III and IV in Fig.
3).
Anaerobic degradation
of acetone and higher ketones by different
denitrifying strains
of the genus
Pseudomonas involves an
initial carboxylation reaction
(
35). In the case of the
ketone 2, such a pathway produces 5,9,13-trimethyltetradecanoic
acid
(
6), which may be subsequently metabolized via alternating
-decarboxymethylation and
-oxidation reaction sequences as
described
above. A mechanism involving hydration of the enol form under
kinetic control was previously proposed for the metabolism of
6,10,14-trimethylpentadecan-2-one (compound 2) by the denitrifier
Marinobacter sp. strain CAB (
42). This pathway
produces 6,10,14-trimethylpentadecan-1,2-diol
(compound 20), which is
then metabolized to 5,9,13-trimethyltetradecanoic
acid (compound 6)
(pathway III in Fig.
3). The hydration can also
take place on the enol
form of ketone 2 under thermodynamic control
(pathway IV in Fig.
3),
which leads to the production of 4,8,12-trimethyltridecanoic
acid
(compound 5), a compound that is easily assimilated via a
classical
-oxidation reaction sequence.
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FIG. 3.
Proposed pathways for the anaerobic metabolism of
(E)-phytol by the denitrifying bacterial community. The
broken arrows indicate abiotic processes.
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Reduction of ketone 2 to the corresponding alcohol, compound 10 (pathway VIII in Fig.
3), appears to be more intense under
anaerobic
conditions (Table
1). This result can be attributed
to the fact that a
nonspecific enzyme activity responsible for
this transformation
(
42) can act in this case on a larger amount
of substrate,
since 6,10,14-trimethylpentadecan-2-one (compound
2) is produced in
larger quantities under conditions of anaerobiosis
(Table
1).
The detection of significant amounts of (
Z)-phytenic acid
(compound 12 in Table
1) shows that the denitrifying bacterial
community metabolizes (
E)-phytol via alternating
-decarboxymethylation
and
-oxidation reaction sequences (pathway
I in Fig.
3). As expected,
this pathway (which is independent of
molecular oxygen ;[
28;])
is used by denitrifying
bacteria under anaerobic conditions. These
results are in good
agreement with the observed growth of
Pseudomonas citronellolis on 3,7-dimethyloctan-1-ol or citronellol under
anaerobic
conditions in the presence of nitrate (as an electron
acceptor)
(
26).
(
E)-Phytol can be anaerobically transformed to phytanic acid
(compound 13) by two pathways, one by way of
3,7,11,15-tetramethylhexadecan-1-ol
(dihydrophytol; compound 21)
(pathway VI in Fig.
3) and another
via (
E)-phytenal
(compound 1) and (
E)-phytenic acid (compound
11) (pathway V
in Fig.
3). Since dihydrophytol (compound 21) was
not among the
metabolites isolated and was not present among the
alcohol moieties of
the isoprenoid wax esters identified, we excluded
the possibility of
involvement of the first mechanism during the
anaerobic metabolism of
(
E)-phytol by the denitrifying
community.
3-Hydroxy-3,7,11,15-tetramethylhexadec-1-ene (isophytol) (compound 22)
was also detected at the end of the experiment (Table
1). The
production of this compound can be attributed to the
involvement of a
reversible enzyme-catalyzed allylic rearrangement
of
(
E)-phytol (pathway VII in Fig.
3) analogous to that
recently
proposed by Foss and Harder (
17) for the
transformation of linalool
to geraniol by the denitrifier
Thauera
linaloolentis.
We also detected some isoprenoid wax esters in the reaction products
(compounds 16 to 18) (Table
1). The production of these
wax esters
under denitrifying conditions contrasts with some of
our previous
results in which no wax esters were found when denitrifiers
from our
laboratory collection were grown anaerobically on
6,10,14-trimethylpentadecan-2-one
(compound 2) (
44).
However, during this previous work the bacteria
were not immobilized,
and it is generally believed that more of
the substrate is assimilated
when the bacteria are free (
29,
48).
Free isoprenoid compounds present in Carteau Bay sediments.
To
compare our laboratory observations with those of natural processes,
free isoprenoid compounds were quantified in 2-cm-long core sections to
a depth of 14 cm at the same site as that from which the sediment used
for bacterial incubations was obtained. GC-MS analyses showed that in
addition to (E)-phytol, 6,10,14-trimethylpentadecan-2-one (compound 2), dihydrophytol (compound 21), (Z)- and
(E)-phytenic (compounds 12 and 11), phytanic (compound 13),
4,8,12-trimethyltridecanoic (compound 5), and
5,9,13-trimethyltetradecanoic (compound 6) acids were present in the
core analyzed (Fig. 4).
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FIG. 4.
Depth profiles of the free isoprenoid compounds in
sediment core sections from Carteau Bay. 4,8,12-TMTD acid,
4,8,12-trimethyltridecanoic acid; 5,9,13-TMTD acid,
trimethyltetradecanoic acid; 6,10,14-TMPD-2-one,
6,10,14-trimethylpentadecan-2-one.
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|
(
Z) and (
E)-phytenic acids (compounds 12 and 11)
were present in particularly large quantities in the upper section of
the
core (Fig.
4). Silyl esters of these two isomers are readily
separated
by GC (Fig.
5) and have easily
distinguished electron impact mass
spectra (
43). Although
(
E)-phytenic acid (compound 11) has been
previously detected
in sediments (
6,
23), this is the first
report, to our
knowledge, of the presence of (
Z)-phytenic acid
(compound
12) in the marine environment. It is generally considered
that phytenic
acids do not accumulate in sediments because they
are readily degraded
or converted to phytanic acid (compound 13)
(
50). The
phytenic acid content declined rapidly with depth
of the core analyzed
(Fig.
4), but this degradation took place
without any significant
production of phytanic acid (compound
13). The amount of phytanic acid
(compound 13) produced corresponds
to only 2% of the amount of
(
Z)- and (
E)-phytenic acids (compounds
11 and 12)
that had disappeared. Due to the presence of a large
proportion of the
Z isomer, the rapid degradation of phytenic
acid may be
attributed to the involvement of alternating
-decarboxymethylation
and
-oxidation reaction sequences induced by denitrifiers. This
is
in good agreement with the presence of 5,9,13-trimethyltetradecanoic
acid (compound 6) in the core and with the detection of small
amounts
of (
Z)-3,7,11-trimethyldodec-2-enoic acid in sediments
of
this area (
36a).
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FIG. 5.
Total ion chromatogram of the silylated fraction
(F3) obtained from the 0- to 2-cm core section. The
asterisks indicate silylated sugars. Question marks indicate
unidentified compounds. 4,8,12-TMTD acid, 4,8,12-trimethyltridecanoic
acid.
|
|
Grimalt et al. (
23) detected relatively large amounts of
C
17 acid 6 in Santa Olalla sediments and water particulate
matter
and theorized that the presence of this compound is indicative
of oxic decomposition processes. The production of this acid during
the
anaerobic biodegradation of phytol in our experiments does
not support
this
hypothesis.
6,10,14-Trimethylpentadecan-2-one (compound 2) was also detected in
this core (Fig.
4). This ketone can arise either via abiotic
degradation of the labile microbially produced (
E)-phytenal
(compound
1) or by hydrolysis of photochemically produced products of
the
phytyl chain of chlorophyll (
40), which are present in
the sediments
of Carteau Bay in relatively large quantities
(
41). The abiotic
degradation of phytenals to
C
18 ketone 2 is highly sensitive to
temperature
(
39); at the temperature of the sediments (annual
average,
15°C), the abiotic half-life of phytenals must be about
four times
longer than it is at 20°C. This explains the previous
detection of
this labile aldehyde in particulate matter (
37)
and sediment
samples (
23,
45). Consequently, only a small
proportion of
phytol must be biodegraded in situ via
6,10,14-trimethylpentadecan-2-one
(compound 2), and most of this ketone
detected in the sediments
probably results from the hydrolysis of
chlorophyll photoproducts.
The anaerobic biodegradation of
C
18 ketone 2 by the mechanisms
described above (Fig.
3) is
a likely source of the 4,8,12-trimethyltridecanoic
(compound 5) and
5,9,13-trimethyltetradecanoic (compound 6) acids
in the core
analyzed.
6,10,14-Trimethylpentadecan-2-ol (compound 10) is present
at trace levels in the sediments of Carteau Bay. Brooks et al.
(
9)
previously showed that a microbial population enriched
from a
lake sediment and grown on phytol under anaerobic conditions
produced
C
18 alcohol 10. These authors suggested that the
likely route
for the formation of this isoprenoid alcohol in Green
River shale
(
12) was via reduction of the corresponding
ketone produced
during phytol biodegradation. This hypothesis is well
supported
by the results obtained in the present study and also
probably
applies to the Santa Ollala sediments (
23).
Dihydrophytol (compound 21) is present in very small amounts in the
core (Fig.
4). This compound has often been considered
a hydrogenation
product of free phytol (
9,
49) and has been
proposed as a
marker for the presence of reducing conditions during
early diagenesis
(
13). The absence of dihydrophytol (compound
21) formation
during bacterial incubations with (
E)-phytol under
sulfate-reducing (
24) and denitrifying (present work)
conditions
strongly suggests that the presence of this compound in the
sediments
of Carteau Bay is more likely due to a direct input of lipids
from archaebacteria (
50) or to its production from the
chlorophyll
phytyl side chain during digestion of macrofauna
(
47) or copepods
(
36).
Isophytol (compound 22) was not detected in the sediments analyzed.
However, Brooks and Maxwell (
8) observed the formation
of this isoprenoid alcohol during incubation of
[U-
14C]phytol with sediment from Esthwaite water and
attributed its
formation to a rearrangement of phytol by an unknown
mechanism.
The reversible enzyme-catalyzed allylic rearrangement
observed
in the present work could account for this
isomerization.
In the hydrocarbon fractions (F
1), we failed to detect
significant amounts of isomeric phytenes and phytadienes, which were
the major metabolites identified during the incubation with
(
E)-phytol
under sulfate-reducing conditions
(
24). Owing to the complexity
of these fractions (Fig.
6), a search for these isoprenoid
hydrocarbons
required selected ion monitoring with the diagnostic ions
at
m/
z 82 (for phytadienes) and
m/
z 70 (for phytenes) (
24).
View larger version (15K):
[in this window]
[in a new window]
|
FIG. 6.
Total ion chromatogram of F1 obtained from
the 2- to 4-cm core section. The asterisk indicates the peaks
containing isomeric methyl-branched nonadecenes.
|
|
The results obtained in the present work clearly show that in the
sediment of Carteau Bay, the aerobic and anaerobic metabolism
of free
phytol involves a (
Z)-phytenic acid (compound 12)
intermediate
and subsequent alternating
-decarboxymethylation
and
-oxidation
reaction sequences. Although, it is generally
believed that the
denitrification rate contributes only 3 to 6% of the
total carbon
respiration in marine sediments (
25),
denitrifying bacteria
seem to play a significant role in the
mineralization of free
phytol. The ability of denitrifiers to grow on
isoprenoid structures
can be attributed to the fact that many of these
bacteria possess
the enzymatic equipment needed for the involvement of
alternating
-decarboxymethylation and
-oxidation reaction
sequences (
28),
a pathway avoiding
-methyl-branched
blockages and not requiring
the presence of molecular oxygen. These
very interesting results
led us to conclude that despite the low
nitrate concentration
of marine sediments, the role played by
denitrifiers in the anaerobic
mineralization of lipidic compounds must
not be neglected. These
aerobic facultative microorganisms appear to
possess higher metabolic
and adaptive capacities than sulfate-reducing
bacteria.
| |
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 (0)4 91 82 96 23. Fax: 33 (0)4 91 82 65 48. E-mail: rontani{at}com.univ-mrs.fr.
| |
REFERENCES |
| 1.
|
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
|
| 2.
|
Bal, B. S.,
W. E. Childers, Jr., and H. W. Pinnick.
1981.
Oxidation of ,-unsaturated aldehydes.
Tetrahedron
37:2091-2096.
|
| 3.
|
Baumann, P., and L. Baumann.
1981.
The gram negative eubacteria genera 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
|
| 4.
|
Blackburn, T. H.
1986.
Microbial processes of N- and C-cycles in marine sediments, p. 218-224.
In
F. Megusar, and M. Gantar (ed.), Perspectives in microbial ecology. Slovene Society for Microbiology, Ljubljana, Yugoslavia
|
| 5.
|
Bonin, P.,
P. Omnes, and A. Chalamet.
1998.
Simultaneous occurrence of denitrification and nitrate ammonification in sediments of the French Mediterranean coast.
Hydrobiologia
389:169-182.
|
| 6.
|
Boon, J. J.,
W. I. C. Rijpstra,
J. W. de Leeuw, and P. A. Schenck.
1975.
Phytenic acid in sediments.
Nature
258:414-416.
|
| 7.
|
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].
|
| 8.
|
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
|
| 9.
|
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.
|
| 10.
|
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].
|
| 11.
|
Cason, J., and D. W. Graham.
1965.
Isolation of isoprenoid acids from a California petroleum.
Tetrahedron
21:471-483.
|
| 12.
|
Cox, R. E.,
J. R. Maxwell,
R. G. Ackman, and S. N. Hooper.
1972.
The isolation of a series of acyclic isoprenoid alcohols from an ancient sediment: approaches to a study of the diagenesis and maturation of phytol, p. 263-276.
In
H. R. von Gaertner, and H. Wehner (ed.), Advances in organic geochemistry 1971. Pergamon Press, Oxford, United Kingdom
|
| 13.
|
de Leeuw, J. W.,
B. R. T. Simoneit,
J. J. Boon,
W. I. C. Rijpstra,
F. de Lange,
J. C. W. van der Leeden,
V. A. Correia,
A. L. Burlingame, and P. A. Schenck.
1977.
Phytol derived compounds in the geosphere, p. 61-79.
In
R. Campos, and J. Goni (ed.), Advances in organic geochemistry 1975. Enadimsa, Madrid, Spain
|
| 14.
|
Didyk, B. M.,
B. R. T. Simoneit,
S. C. Brassell, and G. Eglinton.
1978.
Organic geochemical indicators of palaeoenvironmental conditions of sedimentation.
Nature
272:216-222.
|
| 15.
|
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].
|
| 16.
|
Epstein, S. S., and J. Rossel.
1995.
Enumeration of sandy sediment bacteria: search for optimal protocol.
Mar. Ecol. Prog. Ser.
117:289-298.
|
| 17.
|
Foss, S., and J. Harder.
1997.
Microbial transformation of a tertiary allylalcohol: regioselective isomerisation of linalool to geraniol without nerol formation.
FEMS Microbiol. Lett.
149:71-75.
|
| 18.
|
Gellerman, J. L.,
W. H. Anderson, and H. Schlenk.
1975.
Synthesis and analysis of phytyl and phytenoyl wax esters.
Lipids
10:656-661[Medline].
|
| 19.
|
Gillan, F. T., and R. B. Johns.
1980.
Input and early diagenesis of chlorophylls in a temperate intertidal sediment.
Mar. Chem.
9:243-253.
|
| 20.
|
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].
|
| 21.
|
Goossens, H.,
J. W. de Leeuw,
P. A. Schenck, and S. C. Brassell.
1984.
Tocopherols as likely precursors of pristane in ancient sediments and crude oils.
Nature
312:440-442.
|
| 22.
|
Grenz, C.,
M.-N. Hermin,
D. Baudinet, and R. Daumas.
1990.
In situ biochemical and bacterial variation of sediments enriched with mussel biodeposits.
Hydrobiologia
207:153-160.
|
| 23.
|
Grimalt, J. O.,
I. Yruela,
C. Saiz-Jimenez,
J. Toja,
J. W. de Leeuw, and J. Albaiges.
1991.
Sedimentary lipid biogeochemistry of a hypereutrophic alkaline lagoon.
Geochim. Cosmochim. Acta
55:2555-2577.
|
| 24.
|
Grossi, V.,
A. Hirschler,
D. Raphel,
J.-F. Rontani,
J. W. de Leeuw, and J.-C. Bertrand.
1998.
Biotransformation pathways of phytol in recent anoxic sediments.
Org. Geochem.
29:845-861.
|
| 25.
|
Hansen, L. S., and T. H. Blackburn.
1991.
Aerobic and anaerobic mineralization of organic material in marine sediment microcosms.
Mar. Ecol. Prog. Ser.
75:283-291.
|
| 26.
|
Harder, J., and C. Probian.
1995.
Microbial degradation of monoterpenes in the absence of molecular oxygen.
Appl. Environ. Microbiol.
61:3804-3808[Abstract].
|
| 27.
|
Harder, J., and C. Probian.
1997.
Anaerobic mineralization of cholesterol by a novel type of denitrifying bacterium.
Arch. Microbiol.
167:269-274[Medline].
|
| 28.
|
Hylemon, P. B., and J. Harder.
1999.
Biotransformation of monoterpenes, bile acids, and other isoprenoids in anaerobic ecosystems.
FEMS Microbiol. Rev.
22:475-488.
|
| 29.
|
Iriberri, J.,
M. Unanue,
B. Ayo,
I. Barcina, and L. Egea.
1990.
Bacterial production and growth rate estimation from [3H]thymidine incorporation for attached and free-living bacteria in aquatic systems.
Appl. Environ. Microbiol.
56:483-487[Abstract/Free Full Text].
|
| 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.
|
Li, M. W.,
S. R. Larter,
P. Taylor,
D. M. Jones,
B. Bowler, and M. Bjoroy.
1995.
Biomarkers or not biomarkers a new hypothesis for the origin of pristane involving derivation from methyltrimethyltridecylchromans (MTTCs) formed during diagenesis from chlorophyll and alkylphenols.
Org. Geochem.
23:159-167.
|
| 32.
|
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].
|
| 33.
|
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].
|
| 34.
|
Pirnik, M. P.
1977.
Microbial oxidation of methyl branched alkanes.
Crit. Rev. Microbiol.
5:413-422.
|
| 35.
|
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].
|
| 36.
|
Prahl, F. G.,
G. Eglinton,
E. D. S. Corner, and S. C. M. O'Hara.
1984.
Copepod fecal pellets as a source of dihydrophytol in marine sediments.
Science
224:1235-1237[Abstract/Free Full Text].
|
| 36a.
| Rontani, J.-F. Unpublished data.
|
| 37.
|
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.
|
| 38.
|
Rontani, J.-F.,
G. Baillet, and C. Aubert.
1991.
Production of acyclic isoprenoid compounds during the photodegradation of chlorophyll a in seawater.
J. Photochem. Photobiol. A Chem.
59:369-377.
|
| 39.
|
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.
|
| 40.
|
Rontani, J.-F., and V. Grossi.
1995.
Abiotic degradation of the intact and photooxidized chlorophyll phytyl chain under simulated geological conditions.
Org. Geochem.
23:355-366.
|
| 41.
|
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.
|
| 42.
|
Rontani, J.-F.,
M. J. Gilewicz,
V. D. 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].
|
| 43.
|
Rontani, J.-F.
1998.
Electron ionization mass spectrometric determination of double bond position in monounsaturated ,- and , -isomeric isoprenoid acids.
Rapid Commun. Mass Spectrom.
12:1-7.
|
| 44.
|
Rontani, J.-F.,
P. C. Bonin, and J. K. Volkman.
1999.
Production of wax esters during aerobic growth of marine bacteria on isoprenoid compounds.
Appl. Environ. Microbiol.
65:221-230[Abstract/Free Full Text].
|
| 45.
|
Rowland, S. J., and J. R. Maxwell.
1990.
Phytenic aldehydes in a freshwater sediment.
Org. Geochem.
15:457-460.
|
| 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.
|
Sun, M.-Y.,
S. G. Wakeham,
R. C. Aller, and C. Lee.
1998.
Impact of seasonal hypoxia on diagenesis of phytol and its derivatives in Long Island Sound.
Mar. Chem.
62:157-173.
|
| 48.
|
van Loosdrecht, M. C. M.,
J. Lyklema,
W. Norde, and A. J. B. Zehnder.
1990.
Influence of interfaces on microbial activity.
Microbiol. Rev.
54:75-87[Abstract/Free Full Text].
|
| 49.
|
van Vleet, E. S., and J. G. Quinn.
1979.
Early diagenesis of fatty acids and isoprenoid alcohols in estuarine and coastal sediments.
Geochim. Cosmochim. Acta
43:289-303.
|
| 50.
|
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
|
| 51.
|
Wakeham, S. G., and J. A. Beier.
1991.
Fatty acid and sterol biomarkers as indicators of particulate matter source and alteration processes in the Black Sea.
Deep-Sea Res.
38(Suppl. 2):S943-S968.
|
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Abstract
Introduction
