Organic matter sources in an enclosed coastal inlet assessed using lipid biomarkers and stable isotopes

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Organic Geochemistry Organic Geochemistry 39 (2008) 689–710 www.elsevier.com/locate/orggeochem

Organic matter sources in an enclosed coastal inlet assessed using lipid biomarkers and stable isotopes John K. Volkman a,*, Andrew T. Revill a, Daniel G. Holdsworth a, David Fredericks b,1 a

CSIRO Marine and Atmospheric Research, GPO Box 1538, Hobart 7001, Tasmania, Australia b Geoscience Australia, GPO Box 378, Canberra 2601, ACT, Australia

Received 14 June 2007; received in revised form 15 February 2008; accepted 22 February 2008 Available online 29 February 2008

Abstract The sources of organic matter (OM) in surface sediments from Wilson Inlet, an enclosed and seasonally barred inlet in Western Australia, have been determined using a combination of lipid biomarkers (including fatty acids, sterols, phytol, long chain alcohols, alkyl diols, tetrahymanol and C32 hopanol) and stable carbon and nitrogen isotope data for bulk OM and d13C data for n-alkanols. The organic composition of sediments from the inflowing rivers was used as a proxy for terrestrial inputs of higher plant OM, since these sediments contained only small amounts of carbon from microalgae (mostly diatoms), bacteria and cyanobacteria. The relative proportions of ‘‘marine” and ‘‘terrestrial” OM in the sediments could be estimated using the stable carbon isotope signature of the bulk organic carbon. The sediments from Wilson Inlet showed similar lipid compositions and isotope values, and the sterol and fatty acid distributions showed the complexity typical of marine ecosystems. Relative contributions from algal (phytoplankton and microphytobenthos), bacterial, terrestrial plants and seagrass sources were estimated using biomarker/organic carbon ratios in the different source terms. Microalgae and bacteria are the most important source of OM within the inlet (over 80% in most sediments) and most of the algal material originates from the water column, except in sandy shallow sediments at the eastern end of the inlet where a contribution from microphytobenthos can be discerned. The distributions, abundances and d13C isotope values of long chain n-alkanols proved to be useful in quantifying the contributions from terrestrial plants and the seagrass Ruppia megacarpa. The latter is a relatively small contributor (< 10%) to the OM in the sediments, despite the presence of large seagrass beds around the edges of the inlet and the presumed importance of seagrass as a source of carbon in the foodwebs of the inlet. Nonetheless, seagrass is a significant contributor of the ‘‘higher plant” marker 24-ethylcholesterol (sitosterol) in the sediments. Crown Copyright Ó 2008 Published by Elsevier Ltd. All rights reserved.

1. Introduction *

Corresponding author. Tel.: +61 3 62325281; fax: +61 3 62325090. E-mail address: [email protected] (J.K. Volkman). 1 Present address: Water Environment and Sanitation, 7 Fox Place, Lyneham, Canberra 2602, ACT, Australia.

A goal of many organic geochemical studies is to determine the sources and relative proportions of OM in the samples being analysed. The use of stable isotope data to infer C and N sources in marine sediments has a long history (e.g. Parker, 1964; Parker

0146-6380/$ - see front matter Crown Copyright Ó 2008 Published by Elsevier Ltd. All rights reserved. doi:10.1016/j.orggeochem.2008.02.014

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J.K. Volkman et al. / Organic Geochemistry 39 (2008) 689–710

et al., 1972; Fry et al., 1977; Anderson et al., 1992; Andrews et al., 1998; Galimov, 2006), although the differentiation tends to be limited to rather broad categories such as ‘‘terrestrial” or ‘‘marine” or different types of plants (C3, C4 and CAM). Many isotope studies use simple mixing of two end members (e.g. marine and terrestrial; Schultz and Calder, 1976), but this is a simplification since many more sources often need to be considered. Quantitative estimates become uncertain when multiple sources are involved or where the differences in d13C values are relatively small. For a discussion of some of the pitfalls in binary mixing schemes involving d13C and C/N values, see Perdue and Koprivnjak (2007). Specific organic compounds (biomarkers or signature lipids) can also be used to infer the contribution of organic carbon from different sources (e.g. Volkman et al., 1980; Canuel et al., 1995; Boschker et al., 1999; Dachs et al., 1999; Mead et al., 2005; Xu et al., 2007 and references therein), but the quantitative aspects of the approach are still being refined. Ideally, if we knew the ratio of the biomarker concentration to that of organic carbon in the different sources then it would be relatively straightforward to determine the contribution of carbon from each source from measurements of the biomarker content in the sediment. However, this approach has a number of potential pitfalls including: 1. Unique biomarkers may not be known for each possible source (Volkman et al., 1998); some compounds occur only in certain species [e.g. highly branched isoprenoid (HBI) alkenes in relatively few diatoms; dinosterol in some dinoflagellates; etc.], whereas others are widely distributed (e.g. cholesterol in animals and some phytoplankton, sitosterol in terrestrial plants). 2. The biomarker/Corg ratio, C and N content and stable isotope values in a plant, alga, animal or bacterium can vary due to inter-species differences or effects due to seasonality or environmental conditions etc. (e.g. Fourqurean et al., 1997; Papadimitriou et al., 2006). 3. Like all organic compounds, biomarkers degrade in sediments and the rate varies depending on the chemical structure and on the particular characteristics of the microbial community (e.g. Canuel and Martens, 1996). This can actually be used to advantage in some cases since the presence of labile lipids such as phospholipids or polyunsaturated fatty acids (PUFAs) clearly indicates the

presence of ‘‘fresh” OM, which may be present in living cells (e.g. Rajendran and Nagatomo, 1999). A more advanced approach uses the d13C values of individual compounds determined from isotope ratio monitoring gas chromatography-mass spectrometry (IRM GC–MS) as indicators of carbon source (e.g. Rieley et al., 1991; Bull et al., 1999; Jaffe´ et al., 2001; Oakes et al., 2005). This approach is attractive since the isotope data provide additional evidence supporting the proposed origins of the biomarkers. Unfortunately, baseline data on the d13C signatures of biomarkers in the contributing organisms are still limited and these values can also show significant variation as a result of environmental, physiological and genetic differences (e.g. Boon and Bunn, 1994; Buskey et al., 1999; Riebesell et al., 2000). In this study, we have identified possible sources of OM in Wilson Inlet, a bar-built estuary in Western Australia (Humphries et al., 1982; Lukatelich et al., 1987; Twomey and Thompson, 2001) and estimated their relative contributions to sediments collected at different sites of increasing distance from riverine input (Fig. 1). Potential sources include: pelagic phytoplankton, including diatoms and dinoflagellates; phototrophic, sulfate-reducing and other bacteria; aquatic animals, including zooplankton and benthic fauna; macroalgae; seagrass (Ruppia megacarpa Mason) and its epiphytes; terrigenous OM mainly of higher plant origin; and microphytobenthos (benthic microalgae and cyanobacteria). Given the range of potential contributors, we studied a range of biomarkers including FAs, sterols, long chain alcohols, triterpenoid alcohols and phytol. Only a limited set of sediment and source samples was available, so we were constrained to working mainly with literature data on the biomarker compositions of many of these sources. However, we needed to analyze the seagrass and its detritus since literature data are limited (e.g. Maurer and Parker, 1967; Attaway et al., 1970, 1971; Hernandez et al., 2001; Xu et al., 2006), and no data for alcohols were available for this Ruppia species. Others have used hydrocarbons to successfully assign terrestrial and seagrass sources in sediments (e.g. Jaffe´ et al., 2001) and Ruppia maritima does contain long chain n-alkanes (Attaway et al., 1970), but hydrocarbons were relatively minor constituents of the sediments and so were not utilized as source indicators since

J.K. Volkman et al. / Organic Geochemistry 39 (2008) 689–710

NT

691

QLD

WA SA NSW VIC

TAS

33

35 18

12 14

Fig. 1. Map of Wilson Inlet showing sampling locations and inflowing rivers.

more abundant lipids were available. We also analyzed sediments from the inflowing rivers as a proxy for the composition of terrestrial higher plant OM entering the estuary rather than carry out an exhaustive study of the many possible plant sources in this environment. This approach assumes that phytoplankton and seagrass contributions to the river sediments are relatively minor. From the calculations we have developed a general approach to semi-quantitative estimation of multiple carbon sources that should be applicable to other coastal settings. 2. Study area Wilson Inlet is a shallow estuarine ecosystem situated on the remote SW coast of Western Australia (117° 250 E, 34° 200 S; Fig. 1) near the town of Denmark (Lukatelich et al., 1987; Fredericks et al., 1999). It covers an area of about 48 km2 and is about 14 km long by up to 4 km wide. The maximum water depth is about 5 m, but the mean depth is only 1.8 m, so that a high proportion of the sediments receives sufficient light for benthic photosynthesis, although this is attenuated by the tanninstained water. The inlet is seasonally open to the sea and is generally closed from January–February to July–August. When the bar is breached, about 35% of the water in the inlet is lost to the sea within a few days. When the inlet is closed to the sea, the main source of nutrients is likely to be from the Denmark, Hay and Sleeman rivers (Twomey and

Thompson, 2001); sediment samples from each of these were obtained for analysis. Another possible source is the Cuppup drain, but this was not sampled. Runoff can occur throughout the year, but it is usually greatest in winter. The catchment covers some 2263 km2, of which 89% is drained by the Denmark and Hay rivers (Lukatelich et al., 1987). The headwaters of these rivers are mostly flat and swampy (producing highly coloured waters), but change to deeply incised valleys dissecting a lateritic landscape. The Denmark River catchment is mostly vegetated with jarrah forest (Eucalyptus marginata), and with less than one quarter of the land cleared for sheep and wool production, and with some cereal crops. In contrast, over half of the Hay River catchment is cleared (Lukatelich et al., 1987). Wilson Inlet was chosen as one of the focus catchments of Australia’s National Eutrophication Management Program (NEMP) because it was on the verge of eutrophication. Ruppia growth had expanded dramatically and constituted about 90% of the plant biomass in the estuary, impeding boat traffic and often rotting in the shallows (Lukatelich et al., 1987). Of particular interest is the role of bottom sediments and benthic processes as a source of nutrients (Fredericks et al., 1999). The water quality, nutrient status and ecology of Wilson Inlet have been the subject of previous investigations (e.g. Lukatelich et al., 1987; Hodgkin and Clark, 1988; Carruthers et al., 1997; Twomey et al., 1998). The likely importance of the microphytobenthos in

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Table 1 Stable isotope and carbon contents of sediment and seagrass samples (analysed in duplicate)a %N

d13C ‰

%C

C/Nb

Wilson Inlet sediments WI-12 2.8 WI-14 3.5 WI-18 3.2 WI-33 4.0 WI-35 5.1 SGsed 4.3

0.21 0.03 1.1 1.0 1.2 0.21

21.6 21.3 20.7 20.9 22.1 19.6

1.5 0.6 8.0 5.7 8.0 1.6

7.4 18.7 7.6 5.4 6.6 7.6

River sediments DR 2.3 SR 1.8 HR 0.5

0.11 0.17 0.09

26.8 26.0 26.8

2.0 2.5 2.1

17.3 14.7 26.7

Fresh seagrass SG1 5.8

2.2

14.0

22.2

10.1

Dead seagrass SG3 nmc

nmc

20.4

38.8

nmc

Sample

d15N ‰

a

Data are mean of at least 2 analyses and in most cases 4 (apart from SG3 where only one sample was analysed). b C/N values are by weight. c Not measured.

nutrient uptake has recently been assessed by Fredericks et al. (1999) using benthic chambers. The samples are listed in Table 1. The number of sediments analysed was not sufficient to characterise the OM sources in all parts of the inlet, but was sufficient to discern the major features of the system and to test the usefulness of the combined biomarker-isotope approach. 3. Methods 3.1. Sediment analysis Surface sediments collected from Wilson Inlet (designated WI-##) and influent rivers (HR, DR, SR) were provided by the Australian Geological Survey Organization (AGSO, now Geoscience Australia) as frozen samples in glass jars. Sample locations are shown in Fig. 1. A portion of each sample was extracted at the CSIRO Hobart laboratories using a CHCl3/MeOH/water mixture (1:2:0.8 v/v/v) according to a modified version of the Bligh and Dyer (1959) procedure. Approximately 15 g wet sediment was extracted. An aliquot of the extract (50%) was taken for saponification with 3 ml of 5% KOH in MeOH/water (80:20, v/v). The test tubes were vortex-mixed and heated at 80 °C for 2 h. Neutrals were extracted with hexane and the solution was then brought to pH 2 for

extraction of the FA fractions. Methyl esters (FAMEs) were formed by treating the FA fraction with MeOH/HCl at 80 °C for 2 h and extracted into hexane/chloroform (4:1, v/v). The neutral fractions were treated with bis(trimethylsilyl)trifluoroacetamide (BSTFA) to convert hydroxylated compounds such as sterols and alcohols to the TMSi-ethers. C23 n-alkanoic acid and n-C22 alkane were used as internal standards for the FAMEs and neutral lipid fractions, respectively. 3.2. GC GC was performed with a Varian 3400 GC equipped with a 50 m  0.32 id HP 1 column using a temperature programmable injector and flame ionisation detection. Samples were injected at 45 °C. The oven temperature was ramped to 140 °C at 30 °C min1 and to 310 °C at 3 °C min1 (held 10 min). Compounds were identified using relative retention time, co-injection with authentic standards and with previously identified compounds, and GC–MS data. Concentrations were calculated using peak areas referenced to internal standards. 3.3. GC–MS GC–MS analyses of FAME and neutral lipid fractions was performed with a Fisons MD 800 mass spectrometer fitted with on-column injector set at 45 °C. Injections were made into a retention gap attached to a 50 m  0.32 mm id HP 5 column (0.17 lm film thickness) using He as carrier gas. The same temperature ramp as for the GC analysis was used. Typical MS operating conditions were: 70 eV; transfer line 310 °C, source temperature 250 °C, 0.8 scan s1 and mass range 40–650 Da. 3.4. Stable C and N isotope analysis Sediment samples were dried (24 h, 50 °C), ground and weighed into tin cups (Elemental Microanalysis Ltd., UK). Samples for C analysis were decalcified using sulfurous acid according to the method of Verardo et al. (1990). Duplicates of each sample were analysed for N and C contents (both as dry wt.%) and d15N and d13C values using a Carlo Erba NA1500 CNS analyser interfaced via a Conflo II to a Finnigan MAT Delta S mass spectrometer operating in continuous flow mode. Combustion and oxidation were achieved at 1090 °C and

J.K. Volkman et al. / Organic Geochemistry 39 (2008) 689–710

reduction at 650 °C. Where necessary, as a result of high C content, the C signal was quantitatively diluted with He. Samples were analysed at least in duplicate (Table 1). Percent organic carbon and nitrogen were determined by comparison of instrument response (area) calibrated using standards with known C and N content. Stable isotope ratio values were determined using laboratory standards calibrated with NBS-22 oil for C and IAEA-N3 for N. Results are presented in standard d notation:   Rsample d ð‰Þ ¼  1  100% Rstandard where R = 15N/14N or 13C/12C. The standard for N is air and for C Vienna Pee Dee Belemnite (VPDB). Instrument reproducibility was 0.2‰ for C and 0.3‰ for N; sample reproducibility varied between 0‰ and 1‰. 3.5. Compound specific stable isotope analysis Compound specific isotope ratio mass spectrometry was performed with selected sediment extracts using a Hewlett Packard 5890 series II gas chromatograph coupled via a Finnigan MAT GC combustion interface to the isotope ratio mass spectrometer. The chromatograph was equipped with a 60 m J&W DB-1, 0.32 mm id column with He as carrier gas. Samples were injected on-column via a cold on-column injector (‘‘Duck Bill”, Hewlett Packard). The initial oven temperature of 40 °C was maintained for 1 min followed by a 30 °C min1 ramp rate up to 120 °C followed by a 4 °C min1 ramp rate up to 315 °C (held 15 min). Samples were run in duplicate with coinjected C16 and C24 deuterated n-alkanes of known isotopic composition; the average of the two analyses is reported. C isotope ratio values of derivatized components were corrected for the number of carbons added during derivatization (3 for TMSi-ethers and 1 for FAMEs). 3.6. Analysis of seagrass Samples of Ruppia megacarpa Mason were collected by T. Carruthers in August 1996. They were obtained from 20  20 cm quadrants from a site on the north side of the middle of the estuary. The water depth at this site was 1–1.5 m. Samples were stored in plastic bags and delivered on dry ice to Hobart on July 2, 1997. Sediment from this site was also collected and designated SGsed. Seagrass

693

was suspended in filtered seawater to remove any sediment, shells and other extraneous matter. Any obviously old or decayed seagrass blades were removed. The remaining seagrass was separated into two fractions. One half (SG1) was cleaned of epiphytes by suspending it in beakers containing salt water and dilute acid and physically wiping blades with glass fibre filter. The other was untreated (SG2; data not shown). These samples and a third of decayed leaves (SG3) (all ca. 1 g) were then cut into small pieces and ground while wet with clean laboratory sand. Each was extracted using the same procedure as for the sediments. 4. Results 4.1. Organic carbon and nitrogen contents C and N contents (as dry wt.%) are presented in Table 1. Samples were analysed in duplicate. Sediment WI-14 was a coarse sandy sediment and had low N and C contents, leading to imprecision in the measured values and C/N ratio. N content of river sediments was generally low, in the range 0.09–0.17%. Values for WI-12 and WI-14 were also low (0.21 and 0.03, respectively) but, in the more marine-influenced samples WI-18, WI-33 and WI35, the N content was in the range 1.01.2% (Table 1). A similar trend was found for the C contents, with comparatively low values for the river sediments and much higher values for the more marine sediments, with WI-18 having the highest content (8.0%). These carbon values are at the high end of the range normally found for marine sediments, but the C/N values are typical (Table 1). Note the much higher C/N values for the river sediments and WI-14, reflecting the much higher proportion of OM derived from terrestrial higher plants (Table 1). There was a distinct difference in the C content and d13C values of the fresh and decayed seagrass material (Table 1). The C content of fresh seagrass was 38.8% compared with only 22.2% for the decayed material. Qu et al. (2006) reported an intermediate value of 34.3% for Ruppia growing in Lake Illawarra, eastern Australia. 4.2. Stable isotopes 4.2.1. Carbon The mean d13C value for the fresh seagrass samples (–14.0‰) is in the middle of the range reported

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J.K. Volkman et al. / Organic Geochemistry 39 (2008) 689–710

for seagrasses, although values as enriched as –6‰ have been reported for Thalassia and Syringodium (Hemminga and Mateo, 1996). There can be significant differences between species and there is a general trend for seagrass to become more 13C depleted with increasing latitude (Hemminga and Mateo, 1996). Environmental influences such as source carbon, irradiance and temperature can also produce significant variation in the isotope value of aquatic species (France and Holmquist, 1997; Macleod and Barton, 1998; Lepoint et al., 2004; Papadimitriou et al., 2006), so caution should be applied when using isotope values to calculate OM sources. However, for practical purposes the differences are only likely to be a few ‰ for a particular species of seagrass in a given environment (Hemminga and Mateo, 1996). The d13C value for the dead seagrass leaves was surprisingly light (–20.4‰), perhaps due to loss of isotopically heavy biochemical constituents such as sugars on degradation. This appears to be a novel finding. In general terms, the d13C values of sediments within the inlet (–20.7‰ to –22.1‰) suggest mixed terrestrial and microalgal sources. The sediment under the seagrass sample (SGsed), where we might expect the seagrass isotope signature to be more evident, does show the most enriched value (–19.6‰), but even this is significantly different from that of the fresh seagrass, suggesting that seagrass is not the dominant direct source of organic carbon in this sediment. Rather, the value is consistent with significant contributions from microalgal OM, with some contribution from terrestrial plant material (d13C values assumed to be ca. –27‰ based on the values obtained for river sediments and literature values). Detached seagrass blades can easily be transported by currents, winds and tides to the shoreline and occasionally long distances from their source. They are also known to physically degrade quite rapidly, as shown through litter bag experiments (Fourqurean and Schrlau, 2003), so it is perhaps not too surprising to see such a small direct input. It is likely that much of the seagrass-derived OM in sediments in seagrass meadows derives from the roots and rhizomes, but these were specifically removed from our sediment samples. However, such sources could be important in palaeo-sediments containing buried seagrass remains (Xu et al., 2007). 4.2.2. Nitrogen isotopes The d15N values were quite variable, indicating multiple OM sources. The river sediments had val-

ues in the range 0.5–2.3‰, indicating a predominant contribution from terrestrial sources. Sediments from the inlet had values from 2.8‰ to 5.1‰; the latter is typical of marine microalgae, although a range of values can be found, depending on the N source and the extent to which production utilises regenerated N (Waser et al., 1998). The fresh seagrass had the highest value (5.8‰; Table 1). 4.3. FA compositions FAs provide information about a wide variety of OM sources since they are present in all organisms. They occur in different forms (e.g. wax esters, triacylglycerols, glycolipids, phospholipids etc), but these rapidly break down in sediments to produce free FAs. We analysed total FAs after saponification so that no information was obtained about the form in which the FAs occurred. 4.3.1. FAs in seagrass Total FA contents after saponification are given in Table 2. Typical GC traces are shown in Fig. 2. The major FAs in the fresh seagrass were 16:0, 18:3(n3) and 18:2(n6), in order of decreasing abundance (Table 2; Fig. 2). Such distributions are commonly found in other seagrasses such as Zostera (e.g. Johns et al., 1978), and the results are similar to a previous analysis of FAs of Ruppia (Maurer and Parker, 1967). In the decayed leaves, there is a substantial loss of the more polyunsaturated FAs, such as 18:3(n3), leading to a relative enhancement in 18:2(n6). Long chain (>C20) saturated FAs, often abundant in the epicuticular waxes of terrestrial higher plants, were not abundant in any of the seagrass samples. The chloroplast FA, 16:1(n13)t, comprised 1.5% of the FAs in the fresh seagrass, but only 0.4% in the dead leaves (Table 2). 4.3.2. FAs in river sediments The distributions in DR, SR and HR surface sediments were generally similar (Table 2). However, some significant differences are apparent on closer examination of the data. For example, each contains a higher proportion of branched FAs (more pronounced in HR and SR) and a lower proportion of 18:1(n7) than the inlet sediments, reflecting the different microbial communities present. The FAs in the Denmark River sediment (DR) show some unusual features, with high abundances of 15:0 and anteiso 17:0. The latter is derived from bacteria, but the origins of such high levels of 15:0 are not

J.K. Volkman et al. / Organic Geochemistry 39 (2008) 689–710

695

Table 2 FA compositions of sediment and seagrass (SG1 and SG3) samples (lg g1 dry weight of sample) Sample

WI-12

WI-14

WI-18

WI-33

WI-35

SGsed

DR

SR

HR

SG1

SG3

Saturates 12:0 14:0 i14:0 i15:0 a15:0 15:0 16:0 i16:0 10Me16:0 i17:0 a17:0 17:0 i17:1 18:0 19:0 20:0 22:0 24:0 26:0 28:0

0.5 9.5 1.2 4.1 7.9 5.9 37 6.8 3.1 0.3 2.6 2.2 0.3 3.7 0.9 0.8 0.4 1.2 0.9 0.4

0.2 5.7 0.5 1.9 3.5 6.7 36 3.6 2.4 2.0 2.7 1.7 1.2 2.6 0.4 0.4 0.1 0.2 0.2 0.1

0.9 14.8 2.2 8.2 14.4 8.8 55 14 6.8 2.0 3.3 2.8 1.0 3.9 0.3 1.1 0.5 1.1 1.0 0.9

0.9 12.2 2.1 8.6 13.5 5.7 47 8.1 5.0 2.2 2.4 2.7 0.9 5.8 1.2 2.6 1.8 4.6 4.8 3.6

0.9 13.9 2.3 9.1 14.9 7.8 53 1.0 7.1 2.2 3.1 3.0 1.0 4.9 0.9 1.5 0.7 2.0 1.9 1.7

tra 3.7 0.5 1.6 2.7 1.7 16 1.8 1.1 0.6 0.9 0.9 0.4 1.7 0.2 0.3 0.1 tra ndb ndb

0.6 12.6 2.1 7.2 9.9 23 35 3.9 4.9 4.2 6.7 2.5 1.1 3.4 1.5 1.3 1.2 1.4 1.5 1.3

0.3 4.6 1.0 3.7 4.2 1.4 19 0.3 1.0 0.8 1.0 0.7 0.6 3.1 0.5 1.6 1.5 2.1 1.7 0.6

0.4 4.2 0.9 3.2 4.5 1.9 16 0.4 1.2 0.7 1.5 0.8 0.5 3.1 1.5 0.7 0.5 0.6 1.0 0.5

26.9 72.9 ndb 9.6 tr 19 1454 ndb ndb 2.4 3.6 21 13 103 tra 11 5.7 12 7.1 tra

3.2 25.2 ndb 5.2 3.9 10 603 ndb ndb 4.1 2.3 9.0 1.0 44 tra 7.7 2.1 2.4 tra ndb

Monounsaturates 14:1 15:1 16:1(n7)c 16:1(n5)c 16:1(n13)t 18:1(n9)c 18:1(n7)c 18:1(n5) br19:1

0.3 ndb 41 1.5 0.3 5.8 17 0.2 0.0

0.1 ndb 47 1.0 0.4 5.6 8.8 0.1 0.2

0.5 ndb 66 2.9 0.8 7.3 28 0.4 0.4

0.4 ndb 42 1.7 0.6 6.9 25 0.4 0.4

0.5 ndb 57 3.2 0.6 7.6 29 tra tra

tra ndb 13 0.7 0.2 3.3 5.7 0.1 0.1

0.3 3.4 24 3.6 0.2 5.7 8.8 ndb tra

0.1 ndb 7.5 2.2 0.1 3.1 4.2 ndb tra

0.4 ndb 6.5 1.3 0.0 4.7 3.6 ndb tra

tra ndb 140 1.7 71 115 75 ndb tra

ndb ndb 52 3.9 8.0 49 38 ndb tra

Polyunsaturates C16 PUFA 18:3(n6) 18:4(n3) 18:2(n6) 18:3(n3) 20:4(n6) 20:4(n3) 20:5(n3) 22:5(n6) 22:5(n3) 22:6(n3)

1.6 0.8 1.8 3.3 3.4 11 0.4 18 0.2 1.1 0.4

0.7 0.9 2.0 2.3 1.7 7.5 0.1 15 0.6 1.2 2.6

3.0 0.8 1.3 4.1 3.4 17 tra 29 1.0 0.8 2.5

3.0 0.4 5.8 1.2 1.8 12 tra 18 1.1 2.9 0.7

3.0 0.8 1.0 1.8 3.6 15 tra 24 1.0 1.5 3.2

0.4 0.4 2.5 1.3 2.4 3.2 0.2 6.3 0.2 0.2 1.1

2.0 0.7 1.6 3.3 3.0 5.3 ndb 11 0.7 1.2 2.4

1.3 0.2 0.3 3.1 3.4 1.1 ndb 1.0 ndb ndb 0.7

1.3 0.2 0.3 1.9 1.0 1.1 ndb 1.4 ndb ndb 0.5

230 0.6 34 830 1240 17 ndb 80 ndb ndb 15

8.3 0.0 7.8 640 470 7.8 ndb 14 ndb ndb tra

Total

200

170

312

258

285

75

202

78

68

4600

2020

a b

Trace. Not detected.

known. FAs common in diatoms, such as 16:1(n7) and 20:5(n3), are very much less abundant in the river sediments than in those from the inlet, but their presence is good evidence for some contribution from microalgae. Long chain (C22C28) FA abundances were comparable for DR and SR (5.4 and 5.9 lg g1), but HR had much lower amounts

(2.6 lg g1). Both higher and lower contents were found in sediments from the inlet (Table 2). The SR sediment also contains a higher proportion of C18 PUFA, indicating a greater contribution from higher plants or freshwater green algae to the fatty acids, but this was still subordinate to that from diatoms and bacteria.

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J.K. Volkman et al. / Organic Geochemistry 39 (2008) 689–710 100

16:0

Rel. Intensity

16:1

WI-33 18:1

i15:0 a15:0

20:4(n-6)

14:0

20:5(n-3) 18:0

15:0

20:0

26:0

24:0

22:0

28:0

0 100

16:0

Rel. Intensity

IS

Hay River 18:2(n-3) 16:1

18:1

14:0

18:0

18:2(n-6)

28:0

26:0

24:0

22:0

20:0

0 100

16:0

Rel. Intensity

18:3 (n-3)

Seagrass

14:0

16:1 16:3

18:2(n-6) 18:1 18:0

IS 20:5 22:0

20:0

24:0

0 100

150

200

250

300

350

400

450

500

Time (minutes)

Fig. 2. Partial total ion chromatograms of total FAs as methyl esters in sediment from site WI-33 and Hay River and the seagrass Ruppia megacarpa. FAs are denoted by number of carbons: number of double bonds. Quantitative data for these and other samples are shown in Table 2.

4.3.3. FAs in Wilson Inlet sediments All of the Wilson Inlet sediments contain similar FA distributions (Fig. 2; Table 2). Saturated FAs are dominated by 16:0 (1721%), monounsaturated FAs by 16:1(n7) (1627%) and PUFAs by 20:5(n3) (69%). These 3 are the major FAs in diatoms (e.g. Volkman et al., 1980; Dunstan et al., 1994) and the distributions are characteristic of these microalgae. The data are consistent with the view that diatoms are a significant source of OM in the sediments. Similar distributions have been found in intertidal sediments in which diatoms are a major source of the lipids (Volkman and Johns, 1977). Note that HBI alkenes, found in a few specific diatoms (e.g. Volkman et al., 1994), were not observed (unpublished data), but the sterol distributions (see below) also support a significant diatom source of OM. The relatively low abundance of 22:6(n3) FA indicates only minor contributions from dinoflagellates and marine animals. Similarly, the low proportion of C18 PUFAs indicates minor contributions from green algae. 4.4. Sterol distributions Sterols provide a good indication of eukaryotic sources of OM since, with very few exceptions, they

are not synthesised by bacteria. Some cyanobacteria have been reported to produce very small amounts of sterols such as cholesterol and 24-ethylcholesterol, although evidence for sterol biosynthesis by cyanobacteria is equivocal (Volkman, 1986, 2005). Sterols are considered to be more stable than FAs in sediments and usually occur predominantly in the free (non-esterified) form. Note that the absolute concentration of sterols varies widely between algal groups, so relative amounts may give a false impression of the relative contributions of OM from different species. Quantitative data expressed on a dry weight basis are given in Table 3. Representative GC traces are shown in Figs. 3 and 4. 4.4.1. Sterols in seagrass The seagrass had a simple distribution dominated by the so-called ‘‘higher plant” sterol 24-ethylcholesterol (sitosterol), with smaller amounts of 24-methylcholesterol (campesterol) and 24-ethylcholesta-5,22E-dien-3b-ol (stigmasterol). Trivial sterol names can be used here since the stereochemistry of the alkyl group at C-24 is a, but this need not be the case in the sediments where both isomers could occur. In this respect the sterols are the same as those found in a previous analysis of Ruppia

Table 3 Concentration (in lg g1 dry wt.) of selected neutral lipids in sediment and seagrass samples Neutral lipid

a b c

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21

Inlet

Sediments

SGsed

WI-12

WI-14

WI-18

WI-33

WI-35

20 0.18 tra 1.9 0.32 8.1 0.79 2.2 0.61 0.15 1.1 2.0 0.08 1.2 2.8 0.65 2.4 0.80 2.5 1.2 0.47 1.1 2.5

22 0.05 tra 1.1 0.20 4.0 0.61 1.9 0.51 tra 0.63 0.99 0.19 0.72 1.4 0.27 2.2 0.66 0.98 0.28 tra 0.25 1.6

30 0.10 tra 3.2 0.90 8.9 1.7 3.1 1.6 tra 1.6 3.0 0.77 1.8 5.0 1.3 4.1 1.5 5.4 4.6 tra 3.0 5.8

23 0.45 0.22 2.7 1.0 8.2 2.7 3.9 1.6 0.73 1.9 2.9 0.80 2.8 6.6 2.3 5.1 2.0 9.6 5.4 1.5 2.7 6.6

30 tra tra 3.3 1.2 12 2.9 4.3 2.8 0.25 2.4 4.2 0.53 2.8 7.8 3.0 5.9 2.3 11 6.6 tra 3.6 6.8

8.2 ndb ndb 0.80 0.15 3.9 0.74 1.1 0.68 tra 0.64 0.96 0.27 0.28 1.1 0.30 2.0 0.25 0.79 0.23 ndb 0.21 1.0

River

Sediments

DR

SR

HR

SG1

Seagrass SG3

25 ndb ndb 0.53 0.12 6.1 1.8 4.5 2.5 1.8 0.82 4.0 1.5 0.54 3.9 1.3 18 6.0 3.4 ndb 0.15 0.60 22

9.8 ndb ndb 0.44 tra 5.5 2.0 4.2 3.4 2.5 0.71 3.6 1.6 0.72 5.9 1.4 24 7.1 1.9 ndb tra 0.55 17

8.0 ndb ndb 0.49 0.12 4.6 1.8 4.0 3.1 1.9 0.72 2.6 0.81 0.75 3.5 0.58 11 4.1 1.5 ndb 0.30 0.21 11

1450 ndb ndb 2.0 tra 28 0.89 20 tra ndb 8.0 76 tra ndb 130 8.5 470 18 1.7 ndb ndb ndb 120

130 ndb ndb 1.0 ndb 14 ndb 6.1 tra ndb 2.7 90 ndb ndb 140 trt 400 9.6 3.0 ndb ndb ndb 68

J.K. Volkman et al. / Organic Geochemistry 39 (2008) 689–710

Phytol 24-Nor-cholesta-5,22E-dien-3b-ol 24-Nor-5a-cholest-22E-en-3b-ol Cholesta-5,22E-dien-3b-ol 5a-Cholest-22E-en-3b-ol Cholesterol 5a-Cholestanol 24-Methylcholesta-5,22E-dien-3b-ol 24-Methyl-5a-cholest-22E-en-3b-ol Ergosterol 24-Methylenecholesterol 24-Methylcholesterol 24-Methyl-5a-cholestan-3b-ol 23,24-Dimethylcholesta-5,22E-dien-3b-ol 24-Ethylcholesta-5,22E-dien-3b-ol 24-Ethyl-5a-cholest-22E-en-3b-ol 24-Ethylcholesterol 24-Ethyl-5a-cholestanolc Dinosterol C30 alkyl-1,15-diol Tetrahymanol C32 17b,21b-hopanol C20-C26 even n-alkanols

GC peak

Trace constituent. Not detected. Includes a contribution from fucosterol (up to ca. half of concentration indicated).

697

698

J.K. Volkman et al. / Organic Geochemistry 39 (2008) 689–710 50

x2 WI-12 Total Neutrals –TMS Total Ion Chromatogram

14 5 16 18

7

Relative Intensity

11 3 % 13 10

19

6 21

15 20

17

8

4 9

12

1 2

0 2250

2300

2350

2400

2450

2500

2550

2600

2650

2700

2750

2800

2850

2900

2950

3000

3050

3100

3150

3200

3250

3300

3350

3400

3450

Scan number

Fig. 3. Partial total ion chromatogram of total neutrals in sediment from site WI-12 in Wilson Inlet. Note that the major peak is off scale by a factor of 2. Major sterols and alcohols are labelled with numerals identified in Table 3 together with quantitative data for all samples.

(Attaway et al., 1971) and other flowering plants (Volkman, 1986), including seagrasses such as Zostera (Volkman et al., 1981) and Thalassia (Jaffe´ et al., 2001), but variations in the ratio of these sterols have been noted between species. Note that pentacyclic triterpenoid alcohols were not detected. Caution must be applied when using sterol distributions to distinguish between terrestrial plant and seagrass contributions, unless isotope measurements of the individual compounds are available. Such data clearly provide an elegant means for confirming sources of the different compounds, and hence of the OM (Jaffe´ et al., 2001), as we show below for the alcohol distributions. Very small amounts of microalgal sterols are also present in these seagrass samples, indicating the presence of some epiphyte material, even in the well washed samples. Extensive fouling of Ruppia with diatoms embedded in a polysaccharide slime has been reported for Wilson Inlet by Carruthers et al. (1997). These authors also reported cyanobacteria and green algae in some epiphyte samples.

4.4.2. Sterols in river sediments The three river sediments contain a mixture of sterols. The major one is 24-ethylcholesterol which, together with 24-methylcholesterol and 24-ethylcholesta-5,22E-dien-3b-ol, is almost certainly derived from terrestrial plants. Sediment from the Hay River had the lowest proportion of these plantderived sterols. The river sediments also contain algal-derived sterols such as 24-methylcholesta5,22E-dien-3b-ol (a common constituent of diatoms and haptophyte microalgae; Barrett et al., 1995) and cholesterol from animals and microalgae. Saturated sterols (stanols) are present, which indicates that some double bond reduction occurs in the sediments. However, the stenol/stanol values are still quite high and therefore not indicative of substantial degradation. Small amounts of the faecal sterol coprostanol were found in SR and HR, but the concentrations are consistent with natural sources and do not indicate significant contamination with human faecal material (e.g. Nichols and Leeming, 1991; Leeming et al., 1997).

J.K. Volkman et al. / Organic Geochemistry 39 (2008) 689–710 100

699 dinosterol

Rel. Intensity

WI-33 cholesterol

C29Δ 5,22

C30Diol

C28Δ 5,22 C22

C27 Δ 5,22

C26

C24

0 C29Δ 5

100 Rel. Intensity

Denmark River C26

cholesterol

C24

C22

C29Δ 0

0 C29Δ 5

100 Rel. Intensity

Seagrass IS C29Δ 5,22 cholesterol

C28Δ 5

0 300

325

350

375

400

425

450

475

500

525

550

575

600

625

Scan number

Fig. 4. Capillary gas chromatograms showing distributions of fatty alcohols and sterols in sediment from site WI-33 and Denmark river compared to the total neutrals from the seagrass Ruppia megacarpa. Retention times in the latter are slightly different due to a change in GC conditions and column.

4.4.3. Sterols in Wilson Inlet sediments All contain a complex mixture of sterols indicative of mixed marine-terrigenous sources. The distributions are quite similar, with the same complement of sterols in all the sediments analyzed. The major sterol in each case is cholesterol (15.529.8% of total sterols) which is commonly associated with animalderived OM although some microalgae do contain this sterol. A range of algal-derived sterols is present, including 24-methylcholesta-5,22E-dien-3b-ol and 23,24-dimethylcholesta-5,22E-dien-3b-ol. The latter is less abundant in the river sediments, reflecting differences in microalgal communities between these sites. The dinoflagellate marker dinosterol is particularly abundant in sediments from the marine end of the inlet, where it represents 18.1% and 16.8% of the sterols in WI-33 and WI-35, respectively. 4.5. Other neutral lipids

roalgae), since this class of microalgae is the main source of these compounds identified to date (Volkman et al., 1992). Diatoms of the genus Proboscia have been shown to synthesize C28 and C30 nalkyl-1,14-diols (Sinninghe Damste´ et al., 2003), but these were only present in trace amounts in the Wilson Inlet sediments. The C30 diol was not detected in the river samples (Table 3). All the Wilson Inlet total neutrals fractions contained a suite of compounds identified as 1-O-alkylglyceryl-2,3-diols from their characteristic spectra and base peak at m/z 205 for the TMSi-ethers (Fig. 5). Note that 1,3-diols give a base peak at m/z 218, but these were only present in very minor amounts. They showed a range of saturated nand branched (mainly iso- and anteiso-) alkyl chain lengths from C14 to C19, with only minor amounts of unsaturated components. They occurred in both the Wilson Inlet and river sediments (Fig. 5).

The total neutrals fractions from the sediments also contained significant amounts of other hydroxylated compounds, including the C30 n-alkyl-1,15diol (Fig. 3; Table 3). This may be derived from eustigmatophytes (a class of mainly planktonic mic-

4.5.1. Acyclic and cyclic alcohols Coastal sediments commonly contain a range of acyclic and cyclic alcohols such as phytol (derived from chlorophyll-a; chlorophyll-a/phytol = 3), long chain (C20C32) n-alkanols showing a strong even

700

J.K. Volkman et al. / Organic Geochemistry 39 (2008) 689–710

100

16:0

WI-33 Neutrals –TMS m /z 205 Glyceryl ether 2,3-diols CH 2OR CHOTMSi

Relative Intensity

CH 2OTMSi

10Me-16:0?

i15:0 a15:0

18:0

l17:0 17:0 a17:0

15:0 16:1

14:0

18:1

l16:0 l18:0

i14:0 0 1300

1350

1400

1450

1500

1550

1600

1650

1700

1750

1800

1850

1900

1950

2000

2050

2100

2150

2200

2250

Scan number

Fig. 5. Distribution of 1-O-alkyl-glyceryl-2,3-diols in Wilson Inlet sediment WI-33 determined from the characteristic m/z 205 base peak of the TMSi-ethers. The alkyl group is designated by R and shown by the shorthand nomenclature of number of carbon atoms:number of double bonds. Branched chains are shown as i for iso and a for anteiso.

predominance (derived from the waxes of terrestrial higher plants and found in seagrasses; Johns et al., 1980; Volkman et al., 1999) and pentacyclic triterpenoids such as tetrahymanol (derived from protozoa; ten Haven et al., 1989) and C32 b,b-hopanol (derived from bacteria and cyanobacteria; Dastillung et al., 1980; Summons et al., 1999). Phytol was detected in all the sediments, but it is rapidly degraded and the amount measured includes both free and esterified phytol. In surface sediments such as these, it is highly likely that most of the measured phytol is still esterified to chlorophyll-a. The C32 b,b-hopanol was detected in all the sediments, but the abundance of tetrahymanol was more variable, with some sediments containing barely detectable amounts. Similar distributions have been reported for other marine sediments (e.g. ten Haven et al., 1989; Volkman et al., 1987). Neither compound was present in the seagrass (Table 3). 4.5.2. n-Alkanols in seagrass The three seagrass samples contained very similar distributions of C20C28 n-alkanols, dominated by

C22 and C24, with the former predominating (Fig. 6). Zostera muelleri contains a similar distribution, with C22 predominating although C20 is more abundant in that species than in Ruppia (Johns et al., 1980; Volkman et al., 1981). Note that all seagrasses analysed to date contain these long chain alkanols (Johns et al., 1980; Volkman et al., 1981; Nichols and Johns, 1985; Jaffe´ et al., 2001), so there is the potential to incorrectly assign such long chain alcohols in coastal marine sediments to a terrestrial origin rather than from seagrasses (Jaffe´ et al., 2001). Jaffe´ et al. (2001) reported that the seagrass Thalassia contains C22C32 alcohols, maximizing at 28:0 when analysed with epiphytes removed, but the distribution was dominated by 22:0 when epiphytes were present. We analysed Ruppia with and without epiphytes and the distributions were almost identical (unpublished data). Moreover, we analysed epiphytes from two samples of seagrass and these contained only minor amounts of long chain alcohols (more than an order of magnitude less abundant per dry weight than in the seagrass) and the

J.K. Volkman et al. / Organic Geochemistry 39 (2008) 689–710 60

C20

C22

C24

701

C26

Relative Abundance

50 40 30 20 10 0

WI-12

WI-14

WI-18

WI-33

WI-35

Wilson Inlet Sediments

DR

SR

HR

River sediments

Live SG

Dead SG

Seagrass

Fig. 6. Distributions (%) of long chain (C20, C22, C24 and C26) alcohols in sediments and seagrass samples from Wilson Inlet. Note the high relative abundance of 22:0 alcohol in the seagrass samples and elevated levels in several of the sediments.

distributions were similar to that of the seagrass, probably due to some inclusion of seagrass fragments. We thus conclude that an epiphyte source for the 22:0 alcohol in Ruppia is highly unlikely. 4.5.3. n-Alkanols in river sediments The three river sediments contain significant amounts of long chain n-alkanols; these showed a predominance of the C26 and only moderate amounts of C22 (Fig. 6). Such distributions are typical of those in the waxes of terrestrial plants (Kolattukudy, 1970), and so can be used as a signature for the higher plant wax input. 4.5.4. n-Alkanols in Wilson Inlet sediments Long chain n-alkanols are present in all of the Wilson Inlet sediments and the total concentrations are surprisingly similar, but much lower than in the river sediments (Table 3). The distributions show a strong predominance of even chain lengths, with a maximum at C26, except for WI-33 where C22 is more abundant (Fig. 6). Indeed, in all but WI-14 there is an enhanced abundance of C22 compared to that expected if all of the n-alkanols were derived from terrestrial higher plant waxes. This variability and the different chain length distributions between seagrass and the river sediments suggest to us that the ratio of C22 to C26 n-alkanol might be used to distinguish between seagrass and terrestrial plantderived n-alkanols in this environment. This proxy may not be useful in other environments since Jaffe´ et al. (2001) have suggested that seagrass epiphytes might be a source of the 22:0 alcohol in Florida Shelf sediments and Cranwell (1981) suggested an

association of it with ‘‘decomposer organisms”. Possible algal sources are discussed by Volkman et al. (1998). The alcohol data provide evidence for a contribution from seagrasses to the sedimentary OM, although the contribution is relatively minor compared with algal and terrestrial plant sources. Indeed, even in the sediment obtained from within the seagrass beds (SGSed), the seagrass long-chain n-alkanol signature is no more prominent than at the other sites. Seagrass does not appear to be a significant source of OM in sediments at site WI-14, even though this site is closest to the seagrass beds. These sandy sediments experience a lot of winnowing by waves and currents which removes seagrass detritus. 5. Discussion 5.1. Phytoplankton or microphytobenthos? The presence of phytol and PUFAs such as 20:4(n6), 20:5(n3) and 22:6(n3) in the sediments indicates that there is a contribution from relatively fresh (i.e. undegraded) OM since such FAs are rapidly decomposed after cells lyse. It is not immediately clear, however, whether this fresh algal matter is derived primarily from the water column (i.e. phytoplankton) or from the benthic populations of microalgae and cyanobacteria (microphytobenthos). Benthic mats of Oscillatoria have been reported from the Inlet (Lukatelich et al., 1987), with chlorophyll-a content ranging from 2.8 to 45.8 lg g1 (sediment dry weight). All the sediment

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J.K. Volkman et al. / Organic Geochemistry 39 (2008) 689–710

extracts were highly coloured due to the presence of chlorophylls and carotenoids. Preliminary investigations of the pigments isolated from the sediment using HPLC indicated the presence of intact chlorophylls and carotenoids likely to be associated with living algal cells (unpublished data). It is possible to get an approximate estimate of the contribution of the microphytobenthos to the OM in the sediments if chlorophyll/carbon ratio values in contributing sources are known. Benthic mats were not sampled, but previous research indicates that a value of 40 is typical of benthic diatom mats (e.g. de Jonge, 1980; Santos et al., 1997). Note, however, that it can vary considerably, with values as low as 10.2 and as high as 153.9 occasionally recorded (de Jonge, 1980). If we assume that all the phytol in the sediments is associated with intact chlorophyll-a (which sets a maximum for the chlorophyll-a content and thus is likely to be an overestimate since some phytol occurs in the free form and at least some of the chlorophyll is derived from the water column), then the inferred chlorophyll-a content falls in the range 59.191.2 lg g1. This is only slightly higher than the maximum of 45.8 lg g1 reported by Lukatelich et al. (1987) for sediments sampled in April 1983. Converting these to carbon content indicates a range of 2.43.6 mg of living algal carbon in the sediments per gram dry weight. For WI-12 this suggests that about 15% of the measured organic carbon might be associated with living biomass, but for samples WI-18, WI-33 and WI-35 the inferred proportions are only 4.3%, 4.0% and 4.6%, respectively. Even if the highest reported C/Chl value in the literature is used (153.9; de Jonge, 1980) in the calculation this suggests a maximum contribution of living microphytobenthos in the sediments of 15%. However, values in excess of 100 in microalgae are only associated with very low growth rate and high C/N values (e.g. Goldman, 1980), neither of which seems appropriate for our samples. Additional algal organic carbon in the sediment must either be derived from the water column or from dead microphytobenthos. The inlet experiences spring blooms, which appear to be increasing in frequency (Twomey et al., 1998; Twomey and Thompson, 2001). Chlorophyll values are typically low in spring, autumn and winter, typically <5 lg Chl-a L1, but can rise to around 50 lg Chl-a L1 in spring in the middle of the inlet (Twomey and Thompson, 2001), so contributions from the water column could be large. The phytoplankton in the

estuary shows a diversity typical of ecosystems having a Mediterranean-type climate, with blooms of diatoms (especially Chaetoceros), dinoflagellates and cryptophytes, with smaller numbers of chlorophytes, cyanobacteria, chrysophytes and other classes (Twomey et al., 1998). This diversity is matched by the complexity of the sterol distributions (Table 3, Fig. 3). All sediments show moderate amounts of the typical diatom sterol 24-methylcholesta5,22E-dien-3b-ol (although this may also be derived from cryptophytes; Volkman, 1986) and 24-methylenecholesterol (Table 3). The dinoflagellate marker dinosterol is the most abundant sterol in WI-33 and is only slightly less abundant than cholesterol in WI35 (Table 3), reflecting the high abundance of dinoflagellate populations near the marine end of the inlet. Note that dinoflagellates are not typically associated with microphytobenthos populations, which usually show a predominance of diatoms (Barranguet et al., 1997). C30 alkyl diols possibly from eustigmatophytes and the C32 b,b-hopanol derived from cyanobacteria provide further evidence of diverse algal populations. The relatively high abundance of the two sterols 24-methylcholest-5-en-3b-ol (campesterol if the Me group at C-24 is a) and 24-ethylcholesta-5,22Edien-3b-ol (stigmasterol if the Et group at C-24 is a) is only partly explained by their presence in seagrass and terrestrial plants. Indeed, ‘‘stigmasterol” is more abundant than ‘‘sitosterol” in most of the inlet sediments. A significant fraction must be derived from microalgae, with ‘‘campesterol” being derived from some diatoms, dinoflagellates and green algae, while ‘‘stigmasterol” occurs in some diatoms, green algae and chrysophytes (Volkman, 1986). Haptophytes have not been reported in the estuary (Twomey et al., 1998), in agreement with the very minor amounts of long chain alkenones in the sediments. Note that the proportion of stanols is not particularly high for marine sediments, but clearly there has been some sterol diagenesis, including reduction of stenols to stanols. 5.2. Bacterial contributions An obvious feature of the FA distributions is the high proportion of branched FAs indicative of bacterial biomass (Table 2). C15 and C17 iso- and anteiso- FAs predominate in this group. Anteiso-isomers are more abundant than the iso ones in all sediments. Small amounts of even C14 and C16 isoFAs, also of bacterial origin, are also present.

J.K. Volkman et al. / Organic Geochemistry 39 (2008) 689–710

Similar distributions have been reported for many marine sediments (e.g. Volkman and Johns, 1977; Volkman et al., 1980). It is noteworthy that 18:1(n7) (cis-vaccenic acid), common in many bacteria, greatly predominates over 18:1(n9) (oleic acid), more commonly associated with green algae and higher plants (Volkman and Johns, 1977). Given these high abundances of bacterially derived branched acids, it is highly likely that a significant proportion of the 16:0 and 16:1(n7) acids are also derived from bacteria, as hypothesised for some intertidal sediments (Volkman et al., 1980), and not solely from diatoms. Taken together, the data point to substantial bacterial reworking of the OM deposited in the sediment. The alkyl distributions of the 1-O-alkyl-glyceryl2,3-diols are very reminiscent of the FA distributions in bacteria. A major component eluting after 16:0 was tentatively assigned as a C17 alkyl chain (10-methylhexadecanoic) which, if confirmed, would strongly link the distributions to those found in certain bacteria such as the Actinomycetes (Cranwell, 1973), or sulfate-reducing bacteria of the genus Desulfobacter (e.g. Oude Elferink et al., 1998). Smallwood and Wolff (2000) speculated that glyceryl ethers in Arabian Sea cores may have had an origin from bacteria or, more likely, from benthic macrofauna. No information was provided on the alkyl distribution to compare with our results. Since we analysed the samples after saponification, information on the identity of constituents esterified to the 2- and 3-positions on the glycerol was lost. If both are FAs then the compounds would be alkyldiacylglycerols, as detected in sediment trap material off Peru (Wakeham et al., 1983), or it may be that a more polar group is esterified in C-3 of the glycerol. Further studies are needed to confirm whether the 1-O-alkyl-glyceryl-2,3-diols are of bacterial origin or not.

703

5.3. Terrigenous input Terrestrial higher plant matter in sediments is usually indicated by the presence of long-chain even FAs from 22:0 to 30:0, since these are abundant in plant waxes (e.g. Kolattukudy, 1970). However, these FAs collectively represent only 0.3–5.6% of the total FAs in the Wilson Inlet sediments studied (Table 2), and one might infer incorrectly from this that higher plants are not major contributors. The use of these FAs as unique markers for terrestrial plants, particularly when present in relatively small amounts, is questionable since, as we and others have shown, small amounts occur in seagrasses and in some microorganisms (see Volkman et al., 1980; Volkman et al., 1998). For example, Ruppia contains small amounts of 22:0, 24:0 and 26:0 FAs (Table 2). Sterol and long-chain alcohol distributions are often used to infer contributions from terrestrial higher plants. In the river sediments, almost all the sterols present are derived from higher plants and the distributions of C28 D5 and C29 D5 and D5,22 sterols are quite similar, with sitosterol (C29 D5) by far the most abundant. However, in the marine-influenced inlet sediments, there are clearly significant contributions from non-terrestrial sources to the abundance of campesterol (C28 D5) and stigmasterol (C29 D5,22), since they are relatively more abundant than in the river sediments, and a small amount of ‘‘sitosterol” may also have an algal source. The calculation of terrestrial higher plant OM should be relatively straightforward if biomarkers specific to this source are known, together with their ratio to organic carbon in the source organism. In Table 4, we provide calculations based on the absolute abundance of the C26 n-alkanol. The ratio of it to TOC content in the 3 river sediments varies (Table 4), but the range is surprisingly small given

Table 4 Concentrations of terrestrial biomarker 26:0 n-alkanol in Wilson Inlet sediments and inferred contribution of organic carbon (Corg) from terrestrial higher plants (based on mean biomarker/Corg ratio in river sediments as a proxy for terrestrial plant matter entering the inlet)a Corg% (dry wt.) Corg mg g1 26:0 n-alkanol (lg g1) lg 26:0 per mg Corg Inferred terrestrial Corg mg g1 % Corg that is terrestrial

WI-12

WI-14

WI-18

WI-33

WI-35

DR

SR

HR

1.5 15 0.8 0.05 2.5 17

0.6 6 0.6 0.10 1.9 33

8.0 80 2.0 0.025 6.2 8

5.7 57 1.6 0.028 5.0 9

8.0 80 2.4 0.03 7.4 10

1.6 16 8.7 0.54

2.0 20 6.1 0.305

2.5 25 5.1 0.204

a The 26:0 alcohol has a very minor contribution from seagrass which has been ignored; its abundance was converted to terrestrial Corg based on the mean of the 3 river sediment samples, but note that there is significant variation in the 26:0/Corg ratio in the sediments.

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have used the 22:0 n-alkanol, but this has both seagrass and terrestrial sources. By assuming that the ratio of 26:0 n-alkanol to 22:0 n-alkanol in terrestrial plants is 0.62 (based on the average of the 3 river sediments which provide an average of plant input over time), we can estimate the amount of 22:0 n-alkanol in the sediments that is due to seagrass. This value is then converted into an amount of seagrass per gram of sediment using the data for fresh seagrass SG1 (67.1 lg of 22:0 n-alkanol per g of seagrass carbon) or decayed seagrass SG3 (35.3 lg of 22:0 n-alkanol per g of seagrass carbon). Note that these values may not be applicable to other seagrass samples. At this point we can use either the value for the fresh seagrass of 22.1% carbon content (Table 1), which seems a little low, or the higher value of 38.8% for the dead seagrass. Finally these values are compared with the total carbon in the sediment (mean values from Table 1) to calculate the percentage of organic carbon that is due to seagrass. Data calculated in this way for both fresh and decayed seagrass are shown in Table 5. Clearly, such an approach involving subtracting a terrestrial component to calculate the seagrassderived 22:0 value is prone to error so the derived values should be treated with caution. What is inter-

the possible variation in biomarker content in the many higher plants in the catchment. We have used a mean value in the calculations and assumed that all the OM in the river sediments is of terrestrial origin, which is a slight overestimate. This calculation indicates low contributions of organic carbon from terrestrial higher plants to WI-18, WI-33 and WI-35 (511%), moderate contributions to WI-12 (1519%), but a large contribution at site WI-14 (3435%). Sites WI-12 and WI-14 are closest to the Hay and Sleeman rivers (Fig. 1) and there appears to be a general gradient from E to W in terrestrial OM content. 5.4. Estimation of OM contribution from seagrass using n-alkanol distributions In the absence of a unique neutral lipid biomarker for seagrass, we have used the absolute concentrations of the n-alkanols in the sediments to calculate an approximate contribution (Table 5). We assumed that all the 26:0 n-alkanol in the sediments is derived from terrestrial sources (see previous section). This introduces a very small error since the seagrass does contain trace amounts of this n-alkanol (Table 3). As a marker for seagrass we

Table 5 Summary of inferred OM sources in Wilson Inlet sediments determined from long chain alcohol abundances and d13C data

WI-12 WI-14 WI-18 WI-33 WI-35 SGSed DR SR HR

Mean measured Corg % dry wt.

Seagrassa (if fresh) % of Corg

Seagrassa (if dead) % of Corg

Terrestrialb,c % of Corg

‘‘P + B”d (if fresh sg) % of Corg

‘‘P + B”d (if dead sg) % of Corg

Estimated d13Ce (if fresh sg) ‰

Estimated d13Cf (if dead sg) ‰

Measured d13Cg ‰

1.5 0.56 8.0 5.7 8.0 7.7 2.0 2.5 2.1

5.2 1.0 1.6 9.0 1.4 0.5 0.0 0.0 0.0

15 3.1 4.6 26 4.1 1.5 0.0 0.0 0.0

17 33 8 9 10 1.4 97 83 97

78 66 90 82 89 98 3 17 3

68 64 87 63 86 97 3 17 3

21.7 22.9 21.4 20.7 21.5 21.2

21.0 22.6 21.2 19.7 21.3 21.1 26.8 26.0 26.8

21.6 21.3 20.7 20.9 22.1 19.6 26.8 26.0 26.8

a Seagrass-derived (sg) Corg as dry wt.% determined from 22:0 alcohol content (after correction for a small terrestrial plant contribution); seagrass-derived organic matter is assumed to be zero in the river sediments. b Terrestrial-derived (i.e. of higher plant origin) Corg in river samples as determined from d13C values. c Terrestrial-derived Corg in WI samples as determined from 26:0 alcohol content using a mean value for 26:0/Corg ratio in river sediments. d P + B-derived (i.e. mainly phytoplankton plus bacteria) Corg as determined by difference assuming either live or dead seagrass contribution. e Estimated d13C in sediment based on fresh seagrass content and following values: fresh seagrass –14‰, terrestrial: 27‰, phytoplankton –21‰. f Estimated d13C in sediment based on dead seagrass content and following values: dead seagrass –20.4‰, terrestrial: 27‰, phytoplankton –21‰. g Measured d13C values are the mean of at least two measurements and in most cases four measurements.

J.K. Volkman et al. / Organic Geochemistry 39 (2008) 689–710

esting is, however, that the calculated seagrass contribution varies by a factor of 3 depending on whether one assumes that the seagrass is fresh or decayed. A priori, both forms of seagrass might contribute to a sediment in varying proportions, depending on proximity to the seagrass bed. A summary of the inferred OM sources (terrestrial, seagrass and marine algae by difference) based on the abundance of n-alkanols is shown in Table 5. This suggests that dead seagrass comprises <10% of the TOC in most sediments, except for WI-33. While such a low contribution might be surprising at first glance, similar inferences have been drawn by Boschker et al. (2000) in their stable isotope studies of a Zostera seagrass bed. Moreover, these authors were able to show from compound specific analysis of the bacterial FAs that the bacteria in the sediments made little use of seagrass-derived material. This is not to imply that seagrass OM is not an important component of the food web, which has been amply demonstrated in many environments using the distinctive enriched carbon signal of seagrass (e.g. Buskey et al., 1999). Macrophytes such as seagrass are also an important substrate for epiphytes, which can also be a major food source (e.g. France, 1996). Also, much of the carbon in seagrass meadows is below ground and this could be an important source of carbon in some palaeodeposits. The contribution of terrestrial higher plantderived OM is about 10% in the western end of the inlet and about 19% at WI-12. The remaining 7288% organic carbon in these sediments is primarily of algal and bacterial origin. Such high values are consistent with the low C/N values, which are typical of living microalgae and very much less than the values of 2030 found in vascular plants. As a further check on these estimates we can calculate the expected d13C value of the sedimentary OM from a linear addition of the three main source terms. When reasonable values are chosen (i.e. 14‰ for fresh seagrass, 27‰ for terrestrial

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plants and 21‰ for phytoplankton) the correspondence between the calculated and measured values is surprising good (Table 5). Note that for a three end member system like this, the stable carbon isotope data on their own would not be sufficient to estimate the relative contributions. Indeed, most measured isotope values are very similar to the calculated values even though the biomarker data suggest quite different proportions of the three source terms. Indeed, the d13C values do not provide sufficient power to distinguish between a fresh or dead seagrass input (Table 5). Note also that the calculated d13C values for the WI samples are relatively insensitive to a 1‰ change in the chosen d13C values for ‘‘algal+bacterial” and ‘‘terrestrial plant” end members, but in the river sediments this can lead to major changes in the inferred contributions. For example, if terrestrial plants are assumed to have a d13C value of 28‰ instead of 27‰, the contribution from algae + bacteria in DR and HR increases from 3% to 17%, with a concomitant decrease in the terrestrial component. 5.5. Validation of source assignments from d13C values of individual alkanols A key requirement of the calculations using nalkanol abundances is that all of the 26:0 alcohol is assumed to be terrestrial and that seagrass contributes significantly to the 22:0 alcohol. We have checked this by determining d13C values for long chain alkanols in selected sediments (DR, WI-18 and WI-33) and the two seagrass samples (Table 6). The dataset is more limited due to instrumental difficulties in obtaining accurate values for some of the sediment samples where the amounts of alkanols were small and co-elution with other compounds was a problem. The values for the alcohols in the fresh and decayed seagrass samples (SG1 and SG3) matched quite well despite differences in the bulk isotope values (Table 1). These values are approximately 12‰ heavier than for alcohols in

Table 6 d13C values of long chain alcohols in selected sediment and seagrass samples Alkanol

DR

WI-18

WI-33

20:0 22:0 24:0 26:0 28:0 30:0

28.9 ± 1.4 33.6 ± 1.0 33.5 ± 0.2 33.7 ± 0.1 34.2 ± 0.5 34.7 ± 1.8

28.8 ± 0.7 27.9 ± 0.4 27.1 ± 0.6 32.9 ± 0.6 32.7 ± 0.6

28.7 ± 1.4 28.7 ± 1.5 31.8 ± 3.0

SG1

SG3

21.5 ± 0.3 23.3 ± 0.7 22.5 ± 1.5

23.2 ± 0.1 20.4 ± 0.8

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Table 7 Predicted values for 24-ethylcholesterol (sitosterol) content in Wilson Inlet sediments from terrestrial plants and seagrass Total Corg in sediment (dry wt.%)a Calculated Corg from terrestrial plantsa Calculated Corg from fresh seagrassa Calculated Corg from dead seagrassa Expected sitosterol from terrestrial plantsb Expected sitosterol if seagrass is freshc Expected sitosterol if seagrass is deadd Measured sitosterol % Sitosterol of terrestrial plant origin

WI-12

WI-14

WI-18

WI-33

WI-35

1.5 0.26 0.08 0.22 0.58 0.80 4.8 2.4 24

0.56 0.19 0.00 0.01 0.42 0.03 0.18 2.2 19

8.0 0.63 0.13 0.37 1.4 1.3 7.8 4.1 34

5.7 0.52 0.51 1.5 1.2 5.2 31.1 5.1 23

8.0 0.77 0.11 0.32 1.7 1.2 6.8 5.9 29

a

Calculated Corg values derived from Table 4. Sitosterol from terrestrial plant input calculated from a mean value of 7.9 lg sitosterol per Corg (as dry wt.%) in the three river sediments. c Sitosterol from fresh seagrass calculated from 10.3 lg sitosterol per Corg (as dry wt.%) in fresh seagrass (from data in Tables 1 and 4). d Sitosterol from dead seagrass calculated from 21.2 lg sitosterol per Corg (as dry wt.%) in fresh seagrass (from data in Tables 1 and 4). b

the river sediment DR, indicating that alkanol d13C values can be used to distinguish between seagrass and terrestrial plant contributions. In both WI-18 and WI-33, the 26:0 alkanol shows a d13C value close to that found in the river sediment DR, confirming that the major source of this alcohol in all sediments is terrestrial higher plants. We were unable to obtain a value for the 26:0 alcohol in the seagrass samples due to its low abundance, but we would expect it to be about 22‰ (i.e. much heavier than in the sediments). Moreover, the d13C values for the 22:0 alcohol in the WI-18 and WI33 sediments is intermediate between the values for the seagrass and river sediment, providing clear evidence for a significant seagrass origin. For WI33, the value for the seagrass contribution is calculated to be 52% from simple isotopic balance, which is not very different from that of 43% calculated from absolute values of 26:0 and 22:0 alcohols as discussed above. 5.6. Origins of 24-ethylcholesterol in WI sediments The fact that the d13C values for the alkanols are essentially the same in both fresh and decayed seagrass (as would be expected) means that these values cannot be used to determine whether the seagrass contribution in the sediments is from fresh or decayed material. Our calculation that 26% of the measured organic carbon in the sediment might be derived from decayed leaves in WI-33 seems very high, even though this site is closest to extensive seagrass beds. As a check, we calculated how much of the 24-ethylcholesterol (‘‘sitosterol”) in the sediments might be contributed from fresh or dead sea-

grass leaves using data in Tables 1 and 3. For a dead seagrass contribution of 26% of the total Corg in the sediment we would expect the sitosterol content in the sediments to be about 31 lg g1 from this source alone (Table 7). This is almost six times the measured value of 5.1 lg g1. However, if the seagrass contribution is from fresh seagrass we calculate an expected sitosterol content of 5.2 lg g1, which is only slightly greater than the measured value of 5.1 lg g1 and is consistent with the view that in this sediment most of the seagrass-derived OM is fresh and most of the sitosterol is seagrass-derived. A similar calculation for the other sediments suggests that the calculated contribution from dead seagrass leads to sitosterol values greater than those found in the sediment (Table 7). Note that in these sediments, only 1934% of the sitosterol is derived from terrestrial higher plants. While such calculations can only be approximate they do lend support to the view that non-terrestrial plant sources can be a significant source of 24-ethylcholesterol in sediments (e.g. Volkman, 1986). 6. Conclusions Lipid and stable isotope data provide a consistent picture of the OM sources in surface sediments of Wilson Inlet. In the river sediments there is a strong contribution from terrestrial plants, with a small input from microalgae, mostly diatoms and dinoflagellates, and a relatively minor input from bacteria. Cyanobacteria are a minor source judging from the low content of C32 b,b-hopanol. In Wilson Inlet itself, the sediments show similar lipid compositions and isotope values, suggesting at first sight

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that the gradients in OM sources within the inlet are not large. However, a more detailed analysis of the data indicates a spatial trend east to west for 13C, C/ N and some of the lipid biomarkers, with eastern basin sediments showing a more terrestrial signal consistent with their closer proximity to river inputs. Microalgae are the most important source of OM within the inlet and most of this is inferred to originate from the water column rather than the benthos. The terrestrial plant contribution of carbon is relatively small in most sediments (511%), but can be significant (up to 35%) in the fine-grained sediments nearer the inflowing rivers. Perhaps surprisingly, the seagrass does not appear to make a significant contribution to the OM preserved in the sediments (<10%), although it is obviously an important component of the OM cycle. It may be that much of this material is deposited along the shoreline and effectively remineralized there. The study demonstrates the benefit of combining quantitative data for lipid biomarkers with stable isotope data, and the need to adequately characterise potential OM sources. Provided that specific biomarkers can be identified for the major contributing organisms and that the ratios of these biomarkers to carbon are known for each source, then the amount of organic carbon from each can be independently calculated. The sum can then be compared with the actual TOC measured and cross checked against source assignments apportioned from stable carbon isotope data or other biomarkers. The study has highlighted that additional sources need to be considered for some lipids (e.g. long chain alcohols from seagrass, 24-ethylcholesterol from seagrass and microalgae) but, even so, corrections for these contributions can be estimated. The difficulty lies in defining a single biomarker/Corg ratio for each source when multiple species are involved. In this case, analysis of river sediments provided a reasonable proxy of the mixed higher plant contribution, and direct analysis of seagrass could be used, but phytoplankton species are likely to show a wide range of values and their contribution had to be calculated by difference. Moreover, the approach will only work where diagenesis has not significantly changed the biomarker/Corg ratio or, if this has occurred, then the diagenetic effects have to be modelled. Stable carbon isotopes alone could not be used to apportion sources of the organic carbon in this multi-component system, but they do provide an objective test as to whether or not the sum of isotope values determined from the calculated propor-

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tions provides an isotope value consistent with that measured. Stable isotope values of individual biomarkers proved very useful in determining whether or not a biomarker had mixed sources and could be used to determine the relative proportions. Acknowledgements The work was commissioned by the Australian Geological Survey Organisation (AGSO; now Geoscience Australia) as part of that organisation’s contribution to the National Eutrophication Management Program (NEMP), which was a joint initiative of the Land and Water Resources Research and Development Corporation (LWRRDC) and the Murray-Darling Basin Commission (MDBC). We are grateful to T. Carruthers for collecting samples of seagrass. B. Dudley provided helpful comments on an earlier version of the manuscript. We thank L. Bell for the artwork and two reviewers for their helpful comments which significantly improved the manuscript. Associate Editor—P. Farrimond

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