APPLIED AND ENVIRONMENTAL MICROBIOLOGY, Dec. 1999, p. 5484–5492 0099-2240/99/$04.00⫹0 Copyright © 1999, American Society for Microbiology. All Rights Reserved.
Vol. 65, No. 12
Biodegradation of Free Phytol by Bacterial Communities Isolated from Marine Sediments under Aerobic and Denitrifying Conditions JEAN-FRANC ¸ OIS RONTANI,1* PATRICIA C. BONIN,1
AND
JOHN K. VOLKMAN2
Laboratoire d’Oce´anographie et de Bioge´ochimie (UMR 6535), Centre d’Oce´anologie 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
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. 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 -decarboxymethylation 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.
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
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 m in diameter ranging from 89% at 2 cm to 95% at 10 cm [5]), suggesting minimal variation of
* Corresponding author. Mailing address: Laboratoire d’Oce´anographie et de Bioge´ochimie (UMR 6535), Centre d’Oce´anologie 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:
[email protected]. 5484
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TABLE 1. Metabolites detected during growth of the two bacterial communities isolated from Carteau Bay sediments Metabolite
Amt (‰) detected duringa:
Code
Aerobic incubationb
Anaerobic incubationc
1
2.31
0.40
12 and 11
0.08(E/Z ⫽ 3)
8.34(E/Z ⫽ 3.1)
13
0.02
0.82
22
2.10
Traced
10
2
0.15
3
0.32
0.70
Trace
4
0.02
5
0.12
Trace
6
Trace
0.44
16
Trace
1.82
17
0.66
1.78
18
0.25
0.60
19
1.66
a
Based on the amount of degraded substrate (accuracy estimated to be ⫾0.02‰). Biodegradation percentage, 96; 10-day incubation. c Biodegradation percentage, 83; 30-day incubation. d Trace, amount was ⬍0.02%. b
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 con-
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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.
sisting 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 l 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 min⫺1 and then from 130 to 300°C at 4°C min⫺1, 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 min⫺1, 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.
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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). 5,9,13Trimethyltetradecanoic acid was produced from 6,10,14-trimethylpentadecan-2one (32).
RESULTS AND DISCUSSION
FIG. 2. -Decarboxymethylation reaction sequence. COSCoA, acyl CoA thioester. 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
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)-3,7,11,15tetramethylhexadec-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.
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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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.
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. 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 5,9,13trimethyltetradecanoic 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), -decarboxy-
methylation (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)-3,7,11,15tetramethylhexadec-2-enoic acid (phytenic acid; compound 11) by two different pathways (pathways I and IV in Fig. 1). Path-
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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.
way 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
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.
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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-trimethylpentadecan2-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. 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-
APPL. ENVIRON. MICROBIOL.
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 3,7,11,15tetramethylhexadecan-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). (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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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). The results obtained in the present work clearly show that in the sediment of Carteau Bay, the aerobic and anaerobic me-
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tabolism 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 sulfatereducing 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). 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. 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. 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. 16. Epstein, S. S., and J. Rossel. 1995. Enumeration of sandy sediment bacteria:
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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. 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. 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. 27. Harder, J., and C. Probian. 1997. Anaerobic mineralization of cholesterol by a novel type of denitrifying bacterium. Arch. Microbiol. 167:269–274. 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. 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. 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. 34. Pirnik, M. P. 1977. Microbial oxidation of methyl branched alkanes. Crit. Rev. Microbiol. 5:413–422.
APPL. ENVIRON. MICROBIOL. 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. 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. 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. 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. 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. 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. 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.
