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
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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
C18 
-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.
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Grimalt et al. (23) detected relatively large amounts of
C17 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
C18 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
C18 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 C18 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 (F1), 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).
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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.
This work was supported by grants from the Centre National de la
Recherche Scientifique and the Elf Aquitaine Society (Research Groupment HYCAR 1123).
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Abstract
Introduction
