Anaerobic biodegradation of lipids of the marine microalga Nannochloropsis salina

Organic Geochemistry 32 (2001) 795–808 www.elsevier.nl/locate/orggeochem

Anaerobic biodegradation of lipids of the marine microalga Nannochloropsis salina Vincent Grossi a,*, Peter Blokker b,1, Jaap S. Sinninghe Damste´ b a

Laboratoire d’Oce´anographie et de Bioge´ochimie, UMR CNRS 6535- Faculte´ des Sciences de Luminy, case 901- 13288 Marseille cedex 09, France b Netherlands Institute for Sea Research, Department of Marine Biogeochemistry and Toxicology, PO Box 59, 1790 AB Den Burg, The Netherlands Received 19 October 2000; accepted 19 March 2001 (returned to author for revision 17 January 2001)

Abstract In order to determine the susceptibility to anaerobic biodegradation of the different lipid biomarkers present in a marine microalga containing algaenan, portions of one large batch of cultured Nannochloropsis salina (Eustigmatophyceae) were incubated in anoxic sediment slurries for various times. After 442 days, all lipids studied [mono-, di-, and tri-unsaturated hydrocarbons, long-chain unsaturated alcohols and alkyl diols, phytol, sterols, saturated and (poly)unsaturated fatty acids] showed a significant decrease in concentration, which was accompanied by a strong production of sulfide and methane. However, the studied compounds showed a wide range of reactivity and different patterns and extent of degradation. Polyunsaturated fatty acids, phytol and triunsaturated hydrocarbons were the most labile compounds and showed initially rapid degradation rates, followed by a substantial reduction in degradation rate during the later stages of incubation. Long-chain alkyl diols and unsaturated alkenols, known to constitute the building blocks of the algaenan of N. salina, showed fluctuating concentrations with time clearly indicating their release from bound fractions in parallel with their degradation. Other lipids showed a continuous concentration decrease until the end of the incubation, with alkadienes and sterols being the most resistant compounds encountered. Besides providing an extended sequence of reactivity for lipids under anoxic conditions, the results demonstrate that the presence of resistant algaenan in the outer cell wall of microalgae does not protect the other lipids of the cell from anaerobic microbial degradation. # 2001 Elsevier Science Ltd. All rights reserved. Keywords: Lipid biomarkers; Nannochloropsis salina; Anaerobic biodegradation; Kinetics; Mixed microbial community; Recent sediments

1. Introduction Among the various classes of sedimentary organic components, lipids are widely used as biomarkers in geochemical studies for determining the source, transformation and fate of organic matter (e.g. Summons, 1993 and references therein), and specific palaeoenvironmental conditions (de Leeuw et al., 1995). This is mainly due to * Corresponding author. Tel.: +33-4-91-82-96-51; fax: +33-4-91-82-65-48. E-mail address: [email protected] mrs.fr (V. Grossi). 1 Present address: Department of Analytical and Applied Spectroscopy, Faculty of Sciences, Chemistry division, De Boelelaan 1083, 1081 HV Amsterdam, The Netherlands.

their better potential for preservation in marine sediments in comparison with carbohydrates and proteins (e.g. Harvey et al., 1995), and their more specific structures. Among lipids, however, a wide range of reactivity has been observed from laboratory incubations using individual compounds (e.g. Taylor et al., 1981; Sun et al., 1997; Grossi et al., 1998; 2000; Sun and Wakeham, 1998), or planktonic material (Afi et al., 1996; Harvey and Macko, 1997; Sun et al., 1997; Sun and Wakeham, 1998; Teece et al., 1998) and from field studies involving steady-state observations (e.g. Farrington et al., 1977; Haddad et al., 1992; Sun and Wakeham, 1994; 1999) or dynamic sedimentary processes (Canuel and Martens, 1996). It was further demonstrated that these rates are strongly influenced by the presence of oxygen; lipids being

0146-6380/01/$ - see front matter # 2001 Elsevier Science Ltd. All rights reserved. PII: S0146-6380(01)00040-7

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more slowly degraded under anoxia (Harvey and Macko, 1997; Sun et al., 1997; Sun and Wakeham, 1998; Teece et al., 1998). Under oxic conditions, degradation at different rates also affects the lipid distribution (Sinninghe Damste´ et al., 1997). However, whereas aerobic degradation of lipid biomarkers has been well documented, the impact of strict anaerobic bacterial degradation on marine sedimentary biolipids still needs to be investigated. Some species of green microalgae have been shown to contain biomacromolecules that exhibit a conspicuous resistance to bacterial degradation (Largeau and Derenne, 1993; Gelin et al., 1997a). These macromolecules, termed algaenans, often occur in trilaminar outer cell walls (TLS) and generally have a highly aliphatic structure (Gelin et al., 1999; Allard and Templier, 2000). Algaenan-containing TLS are known to escape diagenetic processes and thus are inferred to account for the bulk of the fossil matter in some organic matter-rich deposits (Largeau et al., 1990). However, knowledge of the influence of such resistant outer cell walls on the early diagenetic fate of other components of microalgae cells (i.e. neutral lipids) is rather limited (Afi et al., 1996). Indeed, while numerous studies have investigated the bacterial degradation of green microalgae, most of these studies considered only a few bulk parameters such as algal lysis, the chemical oxygen demand or the percentage of mineralization (reviewed by Afi et al., 1996). These reports did not indicate if the strains contained an algaenan-containing TLS. Afi et al. (1996) demonstrated that the presence of a TLS composed of algaenan in microalgae from the genus Chlorella does not protect the other lipids of the cells (hydrocarbons, fatty acids, triacylglycerols and chlorophyll) from aerobic bacterial degradation. However, the authors did not determine the susceptibility to anaerobic biodegradation of such classes of compounds. In this paper we address this question by simulating the biodegradation of an algaenan-containing microalga from the class Eustigmatophyceae, Nannochloropsis salina, in anoxic modern sediments. The microalga was incubated in the presence of a mixed marine bacterial/ archaeal community under strictly anaerobic conditions. This allowed us to assess and compare the kinetics of anaerobic biodegradation of a wide range of lipid biomarkers (classes and individual components) found in phytoplankton, including various (poly)unsaturated hydrocarbons, long-chain unsaturated alcohols and alkyl diols, phytol, sterols and fatty acids.

2. Experimental 2.1. Sampling and sediment characteristics Seawater and sediment were collected in Carteau Bay (Gulf of Fos, Mediterranean Sea) as described previously (Grossi et al., 1998). The sediment was main-

tained in isotherm bags (containing dry ice) during its transportation to the laboratory, where it was used immediately as a microbial inoculum. This sediment was selected for its shallow oxic layer (4 mm depth; Bonin et al., 1998) and its strong nitrate reduction, sulfate reduction and methanogenic activities, indicating the presence of an active, mixed population of anaerobic bacteria and archaea (Bonin et al., 1998; Grossi et al., 2000). The organic carbon content of this sediment varies with season and sediment depth from ca. 1.4–3.3%. 2.2. Algal substrate Axenic Nannochloropsis salina strain CCAP849/4 obtained from the Culture Collection of the Natural Environmental Research Council (UK) was grown axenically to the stationary phase at 16
C in 20 l batch culture with Si-free f/2 medium. Cells were harvested by centrifugation and kept frozen until preparation of the slurry. 2.3. Preparation of sediment slurries Microcosms were prepared in 60 ml serum bottles, to which 1 ml of a suspension of N. salina cells (corresponding to 24 mg of dry material) was added. The bottles were then transferred into an anaerobic chamber where all subsequent slurry preparation was performed under a N2:H2:CO2 (85:10:5, v/v/v) atmosphere. To each bottle, 25 ml of an aqueous medium and 5 ml of fresh sediment were added. The aqueous medium consisted of enriched (NH4Cl, Na2S and Na2S2O4) filtered seawater buffered to pH 7.2 (see Grossi et al., 2000 for details). A section, 2–8 cm below the surface, of a fresh sediment core was homogenised and used as the bacterial inoculum. Control cultures were performed in parallel either with autoclaved sediment (controls for abiotic reactions) or without algal cells (controls for microbial metabolism). Bottles were sealed with Teflon-coated butyl rubber stoppers before incubation at 20
C in the dark without agitation. Duplicate samples were analysed for each incubation time. Time courses of the concentration of inorganic sulfur species (H2S+HS+S2-; Cline, 1969) and methane (Marty, 1995) in the non-sterile cultures were monitored by spectrophotometry and gas chromatography (GC) respectively. 2.4. Lipid analysis The aqueous phase was extracted with dichloromethane (DCM) yielding an extract E1. The remaining sediment was ultrasonically extracted with methanol (MeOH) and DCM as described elsewhere (Grossi et al., 1998), and solvent-extractable lipids were partitioned into the DCM phase by the addition of water (extract E2). E1 and E2 were combined and saponified using 1 N KOH/MeOH (10% H2O) for 2 h; neutral lipids were

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then partitioned out of the basic solution, while acidic lipids were subsequently extracted with hexane after acidification of the residual water phase with 2 N HCl (pH <3). The neutral lipids were further separated into apolar and polar fractions by chromatography on an alumina column according to Kok et al. (2000). Extracted sediment residues were subsequently saponified for 4 h to release non-extractable, residual ‘‘ester-bound’’ lipids. These ‘‘ester-bound’’ neutral and acidic compounds were extracted as above. For individual component quantification, internal standards (9-tricosene, 1,12-dodecanediol and docosanoic acid for apolar, polar and acidic compounds respectively) were added and extracts were then derivatized before GC analyses. Alcohols were converted to trimethylsilyl ethers with a mixture of pyridine/BSTFA (1:1 v/v, 20 min at 60
C), whereas acids were esterified with 500 ml of a saturated solution of CH2N2 in ether (room temperature, evaporation under N2). Lipids were quantified by GC using a Hewlett Packard 5890 equipped with an on-column injector. Separations were performed with a fused silica capillary column (30 m0.32 mm i.d.) coated with CP-Sil 5CB (0.12 mm film thickness) with helium as the carrier gas. Both a flame ionization detector (FID) and a sulfur-selective flame photometric detector (FPD) were used simultaneously (split ratio FID:FPD=1:2). Samples were injected at 60
C and the oven temperature was programmed to

130
C at 20
C/min and then at 4
C/min to 310
C at which it was held for 10 min. Structural identifications were performed using a HP 5890 Series II Plus gas chromatograph coupled with a HP 5972 mass spectrometer operated at 70 eV with a mass range of m/z 50– 700. Similar conditions as for GC were used with the exception that a HP-5MS capillary column (30 m0.25 mm i.d., 0.25 mm film thickness) was employed.

3. Results and discussion 3.1. Lipid distribution in the initial slurries The major classes of lipids of N. salina strain CCAP849/4, obtained according to the analytical procedure described above, were total fatty acids (79%), sterols (9%), phytol (8%), alkyl diols (2%), alkenes (1%) and alkenols (1%) (relative percentages based on the sum of all quantified lipids). Within each class of compounds, the main individual components encountered and their relative distributions (Tables 1–4) are in good agreement with previous studies of the lipid composition of N. salina (Volkman et al., 1992; Gelin et al., 1997a). Although detected in N. salina, some minor components such as alkatetraenes (Gelin et al., 1997a), diunsaturated C32 alcohol (Volkman et al., 1992) or long-chain hydroxy-fatty acids (Gelin et al., 1997b) were

Table 1 Distributions, apparent first-order decay constants (k, year1), regression coefficients (r2, n=10), turn-over times (, days) and extent of degradation after 442 days of major alkenes during the anaerobic biodegradation of Nannochloropsis salina Alkenes

%/
Cx-y:i

C17:1 C19:1 C25:1 C27:1 C29:1
monoenes

16 11 8 42 23

C25:2 C27:2c C28:2 C29:2
dienes

8 58 28 6

c

C27:3 C28:3 C29:3
trienes
alk. a b c

%/
alk.

% Degraded

r2

k a

28

85 53 30 48 62 56

1.40 0.63 0.29 0.57 0.73 0.66

40

41 39 31 34 37

0.41 0.39 0.30 (0.52)b 0.33 (0.64)b 0.37

32

80 44 36 79

1.30 0.43 0.31 1.20

56

0.63 (1.10)a

78 9 13

(2.70) (1.40)a (0.52)b (1.30)a

(3.50)a (1.20)b (0.69)b (2.70)a

A more rapid decomposition rate was observed over the first 102 days of incubation. A more rapid decomposition rate was observed over the first 224 days of incubation. Present as two isomers.

0.80 0.86 0.85 0.98 0.94 0.99

 a

(0.92) (0.90)a (0.99)b (0.99)a

260 (140)a 580 (260)a 1300 (700)b 640 500 (290)a 550

0.93 0.85 0.92 (0.98)b 0.84 (0.97)b 0.96

890 940 1200 (700)b 1100 (570)b 990

(0.99)a (0.94)b (0.91)b (0.99)a

290 (110)a 850 (310)b 1200 (530)b 300 (130)a

0.92 (0.99)a

580 (330)a

0.78 0.77 0.88 0.91

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Table 2 Distributions, apparent first-order decay constants (k, year1), regression coefficients (r2, n=10), turn-over times (, days) and extent of degradation after 442 days of total, extractable and ‘‘ester-bound’’ phytol and major alkyl diols during the anaerobic biodegradation of Nannochloropsis salina Compounds

k

r2



84 92 36

1.60 (2.90)a 2.30 (4.10)a 0.29 (1.20)b

0.84 (0.99)a 0.83 (0.97)a 0.72 (0.98)b

230 (130)a 160 (90)a 1300 (320)b

22

70 53 87

0.90 0.44 1.70

0.97 0.53 0.96

410 830 220

5

73 44 57

0.93 0.46 0.91

0.83 0.66 0.41

390 790 400

54

54 38 87

0.53 0.26 0.71

0.83 0.41 0.94

690 1400 510

10

40 33 54

0.40 0.29 0.65

0.95 0.83 0.99

910 1300 560

9

63 54 100

0.63 0.48 1.00

0.52 0.51 0.35

580 760 360

58 43 86

0.63 0.33 1.60

0.90 0.52 0.94

580 1100 240

%/


Phytol Total Extractable OH-bound C30 diolsc Total Extractable OH-bound C31 diol Total Extractable OH-bound C32 diol Total Extractable OH-bound C32:1 diol Total Extractable OH-bound C36 diol Total Extractable OH-bound S diols Total Extractable OH-bound a b c

% Degraded

A more rapid decomposition rate was observed over the first 224 days of incubation. A more rapid decomposition rate was observed over the first 102 days of incubation. Present as two isomers.

Table 3 Distributions, apparent first-order decay constants (k, year1), regression coefficients (r2, n=10), turn-over times (, days) and extent of degradation after 442 days of total (extractable+‘‘ester-bound’’) sterols and major n-alkenols during the anaerobic biodegradation of Nannochloropsis salina Compoundsa

%/


% Degraded

k

r2



C27 C29;24ð28ÞE -ethyl+C295 -ethylb C295;24ð28ÞZ -ethyl S sterols

60 4+25b 11

31 22 20 27

0.29 0.18 0.18 0.25

0.90 0.94 0.86 0.93

1300 2000 2000 1500

n-C30:1 alcohol n-C32:1 alcohol
alkenols

47 53

40 57 49

0.43 0.69 0.56

0.93 0.84 0.88

850 530 650

5

a C275=cholest-5-en-3b-ol; C295,24(28)E-ethyl=24-ethylcholesta-5,24(28)E-dien-3b-ol; C295-ethyl=24-ethylcholest-5-en-3b-ol; C295,24(28)Z-ethyl=24-ethylcholesta-5,24(28)Z-dien-3b-ol. b Co-elution: the relative distribution was determined with MS data (Volkman et al., 1992)

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Table 4 Distributions, apparent first-order decay constants (k, year1), regression coefficients (r2, n=10), turn-over times (, days) and extent of degradation after 442 days of total (extractable+‘‘ester-bound’’) major fatty acids during the anaerobic biodegradation of Nannochloropsis salina Fatty acids

%/
Cx-y:i

C14:0 C16:0 C18:0
saturated

23 74 3

C16:1 C18:1
monounsat.

80 20

C16:2 C18:2 C18:3 C20:4 C20:5
PUFAs

2 4 2 14 79


acids a b

%/
acids

% Degraded

k

r2

20

58 64 72 63

0.70 0.80 1.10 0.78

0.91 0.94 0.92 0.94

520 460 340 470

31

67 68 67

0.90 0.94 0.91

0.99 0.91 0.99

410 390 400

49

74 61 87 92 91 89

1.00 0.73 1.30 1.80 1.60 1.60

76

1.10 (2.70)a

(2.70)a (1.80)a (8.80)b (8.60)b (8.60)b (8.10)b



(0.99)a (0.99)a (0.99)b (0.99)b (0.99)b (0.99)b

360 (140)1 500 (210)a 290 (42)b 210 (43)b 220 (42)b 240 (45)b

0.89 (0.98)a

340 (140)a

0.89 0.82 0.71 0.81 0.76 0.76

A more rapid decomposition rate was observed over the first 102 days of incubation. A more rapid decomposition rate was observed over the first 45 days of incubation.

not considered in our study due to their low concentrations. It should be noted that some of the lipids present in the microalga (phytol, sterols, fatty acids and some diols) were also detected in the sediment used as the source of the bacterial/archaeal inoculum; however, for each component, the contribution from the sediment always accounted for <3% of the amount detected in the slurries. At the start of the experiment, alkenes and alkenols appeared only as solvent-extractable compounds, whereas diols, phytol, fatty acids and sterols were present in both an extractable and a non-extractable ‘‘ester-bound’’ form. For these latter lipid classes, the proportion of extractable compounds relative to the total (extractable+‘‘ester-bound’’) was 63, 86, 91 and 95% respectively. 3.2. Microbial metabolisms Abundant production of sulfides and methane was observed during the first 102 days of incubation of the non-sterile slurries containing N. salina (Fig. 1). After day 102, the concentration of sulfides slightly increased until the end of the experiment, whereas methane was produced up to day 210 and then its concentration slowly decreased. The production of sulfides and methane was very low in the slurries incubated in the absence of the microalga (Fig. 1), which indicates that the addition of N. salina cells strongly stimulated anaerobic processes such as sulfate-reduction and methanogenesis. Since the anaerobic conditions employed were not in favour of denitrification processes (Grossi et al., 2000), it is likely that sulfate-reduction and methanogenesis were the major active microbial metabolisms during incubation

of the non-sterile slurries, although the involvement of other anaerobic metabolisms such as iron reduction (Lovley and Philipps, 1986) or fermentation cannot be excluded. 3.3. Anaerobic degradation of lipids The extent of anaerobic degradation of the different biomarkers of N. salina as well as their pattern of degradation varies strongly from one class of compounds to another. None of the biomarkers studied completely disappeared after 442 days of incubation (Fig. 2 and Tables 1–4). 3.3.1. Apolar compounds. The concentration of total (poly)unsaturated hydrocarbons (ranging from C17 to C29) showed a 56% decrease by the end of the experiment, with a more rapid degradation within the first 102 days (Fig. 2A). Alkenes grouped by degree of unsaturation showed varying degrees of degradation. Monounsaturated hydrocarbons exhibited the same decrease in concentration as total alkenes whereas, surprisingly, the summed alkadienes were more resistant (i.e. 63% remained after 442 days; Fig. 2A and Table 1). Alkatrienes showed the highest degree of degradation (ca. 80%) within the alkene class; triunsaturated alkenes accounted for only 15% of the total alkenes at the end of the incubation experiment compared with 32% at the start of the experiment. Individual alkenes showed a varying pattern of degradation from one compound to another (results not shown) and decreases in concentration ranged from 30 to 85% independent of their chain length,

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Fig. 1. Methane and sulfides production during the incubation of non-sterile anaerobic sediment slurries in the presence (assays) and absence (controls) of Nannochloropsis salina cells.

with the greatest variations observed for the mono- and the triunsaturated compounds (Table 1). Thus, the extent of degradation of unsaturated hydrocarbons seems to be related neither to their degree of unsaturation, nor to their chain-length. 3.3.2. Free and bound phytol Total phytol (extractable and residually ‘‘esterbound’’) showed a sharp decrease in concentration (ca. 85%) after 224 days and stabilised thereafter (Fig. 2B). Extractable phytol showed a similar pattern of loss as total phytol, whereas the concentration of residually ‘‘ester-bound’’ phytol only decreased by ca. 26% within the first 102 days after which it stabilised at ca. 70% of the initial concentration (Fig. 3A; Table 2). Consequently, the relative proportion of residually ‘‘ester-bound’’ phytol increased from 14 to 55% within 442 days. Since we did not observe an inverse relationship between the extractable and ‘‘ester-bound’’ phytols (Fig. 3A), the incorporation of extractable phytol into a non-extractable bound phase (via ester bonds) during the incubation seems unlikely. Algal-derived phytol occurs generally in an esterified form as the side chain of chlorophyll-a. The loss of phytol observed in our incubation experiments certainly involved the hydrolysis of the ester group and subsequent biodegradation of free phytol. This conclusion is based on the formation of various degradation products of free phytol during the incubations, i.e. isomeric phytadienes (mainly Z and E phyta-1,3-dienes) and phytenic acid. These degradation products, which were not detected in the corresponding abiotic controls, accounted for 65% of the amount of total phytol that was degraded after 45 days of incubation (76% of extractable phytol), 1% after 224 days, and were not detected in experiments of longer duration. Z and E isomers of phyta-1,3-dienes are important metabolic intermediates formed during the anaerobic biodegradation of free phytol under sul-

fate-reducing conditions (Grossi et al., 1998). The presence of phytenic acid in N. salina slurries is surprising since this compound was not detected in the earlier experiments (Grossi et al., 1998). However, the anaerobic biotransformation of phytol to phytenic acid (via phytenic aldehydes) has been recently demonstrated during the study of the biodegradation of free phytol by a denitrifying bacterial community (Rontani et al., 1999). Although we could not resolve the complete transformation pathways of phytol in our experiments, it seems that different diagenetic routes are involved. On the other hand, it was not possible to determine whether the hydrolysis of the ester group was biologically mediated because free phytol was not separated from its esterified form in the extractable fraction of the abiotic controls. Nevertheless, our results are in contrast with the scheme of degradation of the phytyl chain of chlorophyll-a in anoxic sediments proposed by Johns et al. (1980). These authors suggested that in the anoxic zone of a contemporary sediment esterified phytol was incorporated into a ‘‘bound’’ phase before being released, and that products of phytol degradation were not solvent-extractable but hydrolysable compounds ‘‘bound’’ to the residual sediment, which was not the case in our experiment. 3.3.3. Long-chain alkyl diols and alcohols The long-chain (C30–C36) alkyl diols present in the polar neutral fraction showed a much lower extent of degradation (58%) and a different pattern of degradation than phytol (Fig. 2B; Table 2). Their concentration decreased by 26% during the first 45 days and subsequently increased slightly until day 102, and decreased again at longer incubation times (Fig. 2B). The fluctuation in total diol concentration is even more pronounced if the extractable fraction is considered separately (Fig. 3C). Residual ‘‘ester-bound’’ diols continuously decreased in concentration throughout the incubation,

V. Grossi et al. / Organic Geochemistry 32 (2001) 795–808

801

Fig. 2. Bacterial degradation of: (A) alkenes, (B) alcohols, and (C) major fatty acids of Nannochloropsis salina in anaerobic sediment slurries. The values represent the sum of individual components belonging to these component classes (see Tables 1–4). Symbols represent the average values of duplicate samples (  6%). Open symbols are sterile controls (St.).

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Fig. 3. Relative concentrations of: (A) extractable (E) and residual ‘‘ester-bound’’ (R) phytol, (B) total sterols, (C) total diols, and (D) total fatty acids during the incubation of Nannochloropsis salina in anaerobic sediment slurries. The proportion of the extractable versus residual ester-bound [E/(E+R)] fraction is also given and refers to the right scale of the plots. Symbols represent the average values of duplicate samples (  6%).

but showed a more rapid decrease between 45 and 102 days (Fig. 3C). Interestingly, this decrease in concentration coincided with the increase in concentration of extractable diols, which clearly indicates a release of diols from the residual ‘‘ester-bound’’ fraction to the extractable fraction. Since this inverse trend with time of extractable and non-extractable diols was not observed in the corresponding sterile controls (where ca. 90% of the original diols were recovered after 442 days), we conclude that the release of diols from the bound fraction was microbially mediated. Alkyl diols are building blocks of the algaenan present in different species of Nannochloropsis, where they occur essentially bound to extractable and non-extractable polar lipids through ester and (presumably) amide and/or sugar and/or sulfate linkages (Volkman et al., 1992; Gelin et al., 1997a). In the present study only diols bound with linkages non-resistant to base hydrolysis were considered. As indicated in Table 2, the rates of loss of the diols (total class and individual

components) after 442 days were systematically much lower for the extractable fraction than for the non-extractable fraction. This induced a significant increase in the relative proportion of extractable diols which accounted for ca. 90% of the total (extractable+non-extractable) ‘‘ester-bound’’ diols at the end of the incubation compared with ca. 60% at the beginning (Fig. 3C). Finally, individual diols showed the same pattern of loss as total diols, but their rates of loss appeared strongly variable, ranging from 40 to 73% (Table 2). Like the alkenes, the rate of loss of the different diols did not seem to be related to their chain length, whereas the presence of unsaturation in their carbon skeleton did not enhance their extent of degradation,(see data for C32 and C32:1 diols in Table 2). It is noteworthy that the oxidation of alkyl diols to alkyl keto-ols was not observed in these experiments. The pattern of loss of monounsaturated C30 and C32 alcohols, as well as their extent of degradation, is similar to those of alkyl diols (Fig. 2B and Table 3). Their con-

V. Grossi et al. / Organic Geochemistry 32 (2001) 795–808

centrations decreased (ca. 20% at day 45), subsequently increased slightly before decreasing again (albeit a little bit faster than alkyl diols) throughout the rest of the incubation experiment. Similar to alkyl diols, unsaturated alcohols present in Nannochloropsis cells have been reported to occur bound to extractable and non-extractable polar lipids, essentially through linkages resistant to base hydrolysis, and to a lesser extent through ester-linkages (Volkman et al., 1992; Gelin et al., 1997a). Although residual ester-bound alcohols could be obtained with our analytical procedure, they were never detected in the ‘‘ester-bound’’ fraction, indicating that ester-bound alkenols were neither present in the original slurries, nor incorporated into a non-extractable phase during the incubation. However, the increase in alkenols concentration between 45 and 102 days implies that some of these components were released from a bound fraction that was not analysable by the analytical procedure employed, i.e. extractable polar lipids resistant to base hydrolysis and/or the non-extractable algaenan biopolymers. 3.3.4. Sterols Four sterols, characteristic of N. salina (Volkman et al., 1992), were observed: cholest-5-en-3b-ol (cholesterol), 24ethylcholesta - 5,24(28)Z - dien - 3b - ol (isofucosterol), 24 ethylcholesta-5,24(28)E-dien-3b-ol (fucosterol) and 24ethylcholest - 5 - en- 3b - ol (24-ethylcholesterol) (Table 3). The latter two sterols co-elute and could not be quantified individually. The total sterol concentration showed a steady, but small, decrease during the incubation (Fig. 2B) resulting in the lowest rate of loss observed for the different classes of lipids, i.e. 27% (Tables 1–4). When individual sterols are considered, a difference in the rate of degradation is observed between cholesterol and the C29 sterols (Table 3); the concentration of the former showing an overall decrease of 31% compared to ca. 20% for the C29 sterols (Table 3). This difference is probably due to the presence of an ethyl substituent at C24 in the C29 sterols. Sterols were almost exclusively present as solvent-extractable compounds during the whole experiment as indicated by the proportion of extractable sterols (relative to total), which varied from 95% at the beginning of the incubation to 98% at the end (Fig. 3B). Thus, exchange between the extractable and the ‘‘ester-bound’’ fractions did not seem to occur for these compounds. Among steroidal components known to be intermediates in the degradation of sterols in anaerobic sediments (i.e. stanols and stanones; Taylor et al., 1981), only small amounts of 5a-cholestan-3b-ol (less than 1% of degraded cholesterol) were observed in the bacterial assays. We failed to detect any traces of steroidal ketones. 3.3.5. Fatty acids Microalgal fatty acids are defined as those present at T0 and exclude odd and branched chain bacterial acids

803

which were formed during incubation. These latter components never accounted for more than 5% of the total fatty acids present in the slurries suggesting a relatively low bacterial contribution to the fatty acids pool. Summed fatty acids showed a 50% decrease in concentration over the first 102 days of incubation and a slower rate of degradation thereafter (Fig. 2C), with ca. 20% of the total acids remaining at the end of incubation. Fatty acids grouped by unsaturation showed varied rates of degradation over time. Saturated and monounsaturated acids concentrations decreased similarly (ca. 65%) throughout the experiment, with a slightly more rapid decrease during the first 102 days (Fig. 2C). The concentration of polyunsaturated acids (PUFAs) decreased much more rapidly, with less than 40% of the original concentration remaining after only 45 days (Fig. 2C). At the end of the experiment, PUFAs showed an overall loss of ca. 90% and accounted for 15% of residual fatty acids compared with 49% in the original slurries. The concentrations of extractable and non-extractable ‘‘ester-bound’’ acids exhibited similar pattern of loss with a stronger decrease during the first 102 days and a steady but slower decrease afterwards (Fig. 3D). However, like sterols, the relative amount of residual ‘‘ester-bound’’ acids was very low ( < 10% of total) and stayed relatively stable over the 442-day incubation period (Fig. 3D) indicating that little exchange occurred between the extractable and the nonextractable pools of ‘‘ester-bound’’ acids. 3.4. Abiotic reactions The sterile slurries, incubated in parallel as controls for abiotic reactions, showed only a small decrease ( < 13%) in concentration of the different lipids over the time of incubation (Fig. 2). This demonstrated that: (i) changes in lipid concentration observed in the non-sterile incubations containing N. salina cells were microbially mediated, (ii) the abiotic degradation of lipids in our slurries was negligible, and (iii) sorption to mineral particles did not play a significant part in the preservation of lipids under these conditions (Teece et al., 1998). In the non-sterile incubations, where a strong production of sulfides was observed (Fig. 1), the possibility that sulfurisation reactions could have played a role in lipid sequestration (Sinninghe Damste´ and de Leeuw, 1990) was investigated by searching for the presence of organic sulfur compounds (OSC). The use of a sulfur specific detector could only reveal the formation of very small amounts of sulfur-containing steroids over the 442-day incubation. Gelin et al., (1998) recently demonstrated that the laboratory sulfurisation of N. salina results mainly in the formation of non-hydrolysable and non-extractable (poly)sulfide-bound long alkyl chains. To check for the presence of such chemical structures in our slurries, the final sediment residues were analysed by flash pyrolysis-

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GC–MS but no OSC could be detected in the pyrolysates. This indicated that the sulfurisation of lipids was not a major process in our experiments, despite the significant production of inorganic sulfur species (Fig. 1). This is not surprising since: (i) the conditions employed (i.e. 20
C, natural levels of sulfide, possibility for pyrite formation by reaction with iron) were closer to the conditions found in most modern sediments than the ones established by Gelin et al. (1998) (i.e. 50
C, high concentration of reactive sulfides and sulfur, absence of iron), and (ii) the time scales involved for observable sulfurisation of lipid biomarkers have been recently demonstrated to be 1–5 ka (Kok et al., 2000; Werne et al., 2000). In their study of the aerobic bacterial degradation of algaenan-containing microalgae, Afi et al. (1996) observed that up to 80% of lipid degradation after 4 months of incubation could be assigned to abiotic reactions, including those induced by autooxidative processes. The oxidation of lipids by molecular oxygen in our anaerobic slurries was not possible, but radical reactions could have been responsible for (part of) the degradation of unsaturated lipids, since hydroperoxides of phytoplanktonic origin (i.e. sterol photoproducts) have recently been detected in the anoxic sediment from Carteau Bay used for our bacterial inoculations (Rontani and Marchand, 2000). However, since some of these hydroperoxides resist autoclaving (J.-F. Rontani, 2000, unpubl. data), the low amount of lipids that were lost in the abiotic controls implied that the anaerobic oxidation of unsaturated components by hydroperoxides was negligible in our experiment. 3.5. Kinetics of degradation The decrease in concentration of lipids during the experiments was fitted using least square regression of concentration versus time to yield rate constants (k) using the simple first-order equation: GðtÞ ¼ G0 ekt , where GðtÞ ¼concentration of a component at time t, G0 ¼original concentration. This approach has been widely used to examine bulk and individual biochemical fractions during the degradative process (for review see Henrichs, 1993). For the present experiments, fitted experimental curves generally correspond well with the data (Tables 1–4) as shown by the correlation coefficients (r2). However, many (classes of) components (e.g. alkatrienes, phytol, PUFAs) showed an initial rapid rate, followed by a substantial reduction in rate over the later stages of incubation indicating that the reactivity of some lipids changes over time. This is in good agreement with the trends observed during the decomposition of lipid constituents (Canuel and Martens, 1996; Harvey and Macko, 1997; Teece et al., 1998) and of total organic carbon (Westrich and Berner, 1984; Middelburg, 1989) of natural phytoplanktonic communities.

For these component classes, the maximal extent of degradation was generally reached before the end of the experiment and a non-degradable fraction (GNR ; cf. Westrich and Berner, 1984) was evident in the experiments (Fig. 2). For those components whose concentrations did not stabilise over the 442-day incubation (e.g. alkadienes, sterols, diols, saturated acids), it may be that a longer incubation time would have resulted in further degradation. The fact that the reactivity of some lipids decreases over time may be due to varying degrees of availability to biological degradation processes (from readily biodegradable to refractory) of different matrices containing the same compounds. On the other hand, this may also reflect a concentration-dependent affinity to substrate degradation of microbial enzymes although, in this case, second-order kinetics ð1=C ¼ ktÞ should be more appropriate for calculating the rates of decay of labile lipids. The apparent first-order degradation constants of the different classes of compounds for the entire incubation period ranged from 0.25 year1 for sterols to 1.6 year1 for phytol (Tables 1–4), although some individual components (i.e. C29 sterols and C20 PUFAs) showed degradation constants slightly out of this range. The following decreasing order of reactivity was observed: phytol>PUFAs>alkatrienes>monounsaturated acids>saturated acids>monounsaturated alkenes  diols>alkenols>alkadienes>sterols. The range of reaction constants appeared much wider, however, when the rapid initial degradation was considered, with PUFAs showing a maximal decomposition rate constant of 8.1 year1 during the first 45 days. As discussed by various authors (Harvey et al., 1995; Canuel and Martens, 1996; Teece et al., 1998), the incubation time is an important parameter to consider when studying the reactivity of phytoplanktonic constituents that exhibit decreasing rates of degradation over time. Indeed, rates calculated over longer periods of time will integrate changes over a longer time window and thus yield lower average rate constants of degradation. In the present study, this was particularly evident for the most labile classes of compounds, i.e. phytol, alkatrienes and PUFAs, which rates determined over a period of ca. 100 days were respectively 1.8, 2.2 and 5.2 times greater than over the 442-day period. During this initial period of rapid degradation, PUFAs were degraded 3 times faster than phytol and alkatrienes (which showed similar degradation constants) and 9 to 32 times faster than the other classes of components, which showed the same sequence of decreasing reactivity as described above. This order of reactivity is generally consistent with those obtained by Harvey and Macko (1997) during the simulated degradation of different common microalgae (diatom, cyanobacteria) in anaerobic sea-water, although the overall degradation rates determined in the present study were systematically lower (for the same

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compound classes). As discussed above, this may be due to the extended incubation time of our experiments. However, a major difference between our results and those of Harvey and Macko (1997) was observed for phytol, which appeared as one of the most reactive compound in our study whereas, conversely, its degradation rate was one of the lowest observed by Harvey and Macko (1997). Differences in degradation rates may be due to varied bacterial/archaeal communities from one study to the other and/or to differences between the cellular matrix of the (micro)algae. The reactivity of phytol may be further influenced by the activity of autocatalytic enzymes known to play an important role in chlorophyll defunctionalisation (Spooner et al., 1994). The anaerobic degradation rates of the different classes of fatty acids encountered in N. salina are in agreement with the general view that unsaturation is an important parameter in regulating the lability of many lipids. Indeed, polyunsaturated fatty acids were degraded much faster than monounsaturated fatty acids, which in their turn were degraded faster than saturated acids (Table 4). This feature seems to be, however, essentially verified when anaerobic conditions are prevailing since other studies in which contrasting redox conditions were considered, have demonstrated that saturated and unsaturated acids of planktonic origin are degraded at the same rate in the presence of oxygen, whereas unsaturated acids are preferentially degraded under anoxic conditions (Harvey and Macko, 1997; Sun et al., 1997). As observed in the present study, the higher degradation rate of unsaturated fatty acids under anoxic conditions may result in an enrichment of the sediments in saturated fatty acids compared with the composition in fatty acids of their precursor organism in the overlying water column (e.g. Farrington et al., 1977; Wakeham and Beier, 1991; Harvey and Macko, 1997). Surprisingly, unsaturation did not play a similar role in lipid lability for other classes of compounds that exhibited different degrees of unsaturation. Within the alkene class, alkatrienes were the most labile compounds, but diunsaturated alkenes appeared much more resistant to anaerobic degradation than monounsaturated hydrocarbons (Table 1). Also, the monounsaturated C32 diol was not degraded faster than its saturated homologue (Table 2). The low degradation rates observed for alkadienes (and to a lesser extent monounsaturated alkenes) likely result from their specific chemical structure, especially the position and the geometry of the double bonds (which are presently not known). Alkenes are generally readily degraded under both oxic and anoxic conditions (Harvey and Macko, 1997; Grossi et al., 1998; 2000). Differences in the potential for preservation have been noted, however, between several linear alkenes detected in the Black Sea (Wakeham et al., 1991). Moreover, in their study of the biodegradation of the marine coccolithophorid Emiliania huxleyi under sulfate-reducing and

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methanogenic conditions, Teece et al. (1998) observed decay constants of C31 dienes very close to the ones observed in the present study. Information concerning the reactivity of alkyl diols during early diagenesis is rather limited. At first sight, it seems that these compounds are not highly reactive in spite of the presence of hydroxyl groups and unsaturation (Sun and Wakeham, 1994). This is confirmed by the relatively low degradation constants of the different diols observed in the present study (Table 2). Data from the Madeira Abyssal Plain turbidites indicate that over periods exceeding 5 ka, diols are better preserved than sterols under oxic conditions and that no difference in degradation rates occurs between the various isomers (Sinninghe Damste´ et al., 1997). However, recently, we observed that following their deposition at the surface of coastal sediments, the C32 diols of N. salina can be degraded at rates similar to those of the sterols of the same microalga (V. Grossi, 2000, unpubl. data). The different isomers of diols present in our incubation experiments showed varying degradation constants, which did not seem to be dependent on the chain-length of the compound, nor to its degree of unsaturation (Table 2). It is likely that the release of diols from a bound pool observed during the incubation have influenced the overall degradation rate constants of these compounds and thus enhanced variations between isomers. Acyclic long-chain (>C22) unsaturated alcohols are rarely found in sediments (Mudge and Norris, 1997) and no information concerning their reactivity is presently available. Our results suggest that, whenever present in anoxic marine sediments, (i) long-chain alkyl diols and alkenols are degraded at similar rates, (ii) these compounds can be more rapidly degraded than sterols. The sterols of N. salina were the more recalcitrant lipids in the anaerobic slurries. Although many studies have demonstrated rapid rates for sterols degraded under oxic conditions (Sinninghe Damste´ et al., 1997; Sun and Wakeham, 1998), these compounds are considered to be much less reactive in the absence of oxygen. Sun and Wakeham (1998) observed that 14C-cholesterol degraded ca. 3 times faster under oxic conditions than under anoxic conditions at a simulated sediment-water interface, whereas Harvey and Macko (1997) observed a 13fold decrease in the degradation rate of sterols between oxic and anoxic conditions during simulated sedimentation experiments. Among individual sterols selective preservation is generally considered important, especially in strictly anaerobic sediments (Taylor et al., 1981). This was confirmed in the present study since cholesterol appeared significantly more reactive than the C29 sterols. For the extended incubation period, the turnover times (defined as the reciprocal of the rate constants) of the different lipid classes of N. salina ranged from ca. 210 to 2000 days (Tables 1–4). These values are in good agreement with those reported by Teece et al. (1998) for

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alkadienes (1300–2500 days), alkenones (220–1200 days) and alkyl alkenoates (160–630 days) after similar incubation periods (415–450 days) under sulfate reducing and methanogenic conditions. Shorter turn-over times were reported by Harvey et al. (1995) for total lipids (130–160 days) and phytoplanktonic POC (120–150 days) and by Otsuki and Hanya (1972) for total algal carbon (110–200 days) after incubation under anoxic conditions for 180 and 40 days, respectively. Harvey and Macko (1997) reported turnover times of 27–350 days for individual lipids after 180 days of anaerobic laboratory incubation whereas, during their field study in the anoxic sediments of Cape Lookout Bight (NC, USA), Canuel and Martens (1996) observed turn-over times of 23–500 days for freshly deposited (31–144 days) lipids. As discussed above, extended incubation times lead often to lower average degradation rates and thus longer turnover times to be obtained. For those lipids of N. salina that showed a more rapid decomposition rate during the first 102 days of incubation (i.e. PUFAs, phytol, and some mono- and triunsaturated hydrocarbons), the turnover times calculated over this short period ranged from 42 to 260 days (Tables 1, 2 and 4), which is in full agreement with the values reported in the literature discussed above. Beside the period of time considered for degradation, it is likely that differences in the chemical structure, in microbial assemblages/activities, and in resistance of distinct (micro)algae to bacterial attack may have influenced the turnover times of the lipids of N. salina in comparison to previous studies. This seems especially true for the compounds that exhibited lower and unique degradation rates over the 442 days of incubation (e.g. alkadienes, sterols). In the natural environment, such diagenetic parameters may be further influenced by the type of depositional environment encountered [e.g. temperature, sediment accumulation rates, bioturbation, redox conditions, water depth; see Sun and Wakeham (1994) and Canuel and Martens (1996)]. 3.6. Susceptibility of neutral lipids of algaenancontaining microalgae to microbial degradation Although the biodegradation of N. salina under aerobic conditions was not investigated in the present study, it was previously demonstrated that neutral lipids of an algaenan-containing green microalga (Chlorella emersonii) can be readily degraded in the presence of oxygen (Afi et al., 1996). Thus, in spite of the selective preservation of algaenan-based TLS during fossilization, it is obvious that such outer cell walls do not protect other components of the cell from microbial degradation under either oxic or anoxic conditions. To determine whether the presence of an algaenancontaining outer cell wall could influence the role of oxygen in the degradation of the neutral lipids of

(micro)algal cell, we have calculated (using least square regression) the apparent oxic degradation first-order rate constants of alkenes and fatty acids present in C. vulgaris and C. emersonii from the data of Afi et al. (1996). These constants (k=6.7–7.1 year1 and 21–25 year1 for alkenes and FA, respectively) are 6–9 times greater than those determined in the present study for similar components. This is consistent with the results of Harvey and Macko (1997) who determined that similar lipids present in phytoplankton species that do not contain algaenan (diatom and cyanobacteria) were degraded 4–5.5 times faster in the presence of oxygen compared with anoxic conditions. Therefore, the potential of (bio)degradation/preservation of neutral lipids under both oxic and anoxic conditions does not seem to be significantly different between algaenan-containing green microalgae and other phytoplankton species.

4. Conclusions The long-term incubation of the algaenan-containing microalga N. salina in sediment slurries under anaerobic conditions demonstrated that the lipid biomarkers present in the cells can be significantly, albeit not entirely, (bio)degraded after 442 days. During the study of the bacterial degradation of (micro)algal lipids, these compounds very often reach a threshold concentration below which no further degradation occurs. This nondegradable fraction composed of otherwise labile lipids is thus likely to contribute to the fraction of organic matter which is preserved in marine sediments (GNR ; cf. Westrich and Berner, 1984) together with naturally refractory material such as algaenan. Although the concentration of all components decreased by the end of the incubation, the lipids known to constitute the building blocks of the algaenan of N. salina (i.e. long-chain alkyl diols and unsaturated alkenols) showed fluctuating concentrations with time. This is most likely due to their release from bound fractions, a process already suggested by Blokker et al. (2000) when comparing extant and fossil algaenans, in parallel with their degradation. This contrasted with the continuous decrease in concentration observed for the other compounds studied (i.e. alkenes, phytol, sterols and fatty acids). The release of bound alkyl diols and alkenols suggests that complex lipidic structures present in phytoplankton can be significantly modified in modern anaerobic sediments. By comparing the reactivity of different classes of components present in N. salina, this work provides an extended sequence of reactivity for lipids under anoxic conditions. Beside confirming the strong reactivity of polyunsaturated compounds (PUFAs and alkatrienes), the results further demonstrate that in anoxic sediments: (i) the presence of unsaturation and the chain-length are

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not always good predictors of lipid reactivity and, (ii) phytol can be degraded much faster than linear unsaturated alcohols whereas sterols appeared far less reactive than acyclic alcohols. The wide range of decay rates observed for the numerous lipid constituents of N. salina reinforces the idea that bulk parameters used to approximate the degradation of major biochemical classes may be misleading as they integrate varying rates for individual compounds. On the other hand, this further highlights the complexity of understanding the reactivity of sedimentary lipids at the molecular level, as many parameters may influence their diagenetic fate (e.g. source organism, molecular structure, microbial assemblages, redox conditions).

Acknowledgements This work was supported by grants from the Netherlands Geosciences Foundation (NWO, French-Dutch collaboration 1999 No. 0001/717-704) and from the Centre National de la Recherche Scientifique (CNRS) and Elf Society (GDR Hycar No. 1123). We are grateful to Dr. A. Hirschler for help with microcosms preparation and sulfide measurements, and to Dr. D. Marty for methane measurements. We thank Dr. E.A. Canuel and Dr. M. Teece for their constructive reviews and the associate editor Dr. J.K. Volkman for his helpful comments and suggestions. NIOZ publication 3533. Associate Editor — J.K. Volkman

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