Lipid biomarker patterns reflect different formation environments of mussel- and tubeworm-dominated seep carbonates from the Gulf of Mexico (Atwater Valley and Green Canyon)

Accepted Manuscript Lipid biomarker patterns reflect different formation environments of mussel- and tubeworm-dominated seep carbonates from the Gulf of Mexico (Atwater Valley and Green Canyon)

Hongxiang Guan, Dong Feng, Daniel Birgel, Jörn Peckmann, Harry H. Roberts, Nengyou Wu, Duofu Chen PII: DOI: Reference:

S0009-2541(18)30590-4 https://doi.org/10.1016/j.chemgeo.2018.12.005 CHEMGE 18992

To appear in:

Chemical Geology

Received date: Revised date: Accepted date:

2 April 2018 7 June 2018 7 December 2018

Please cite this article as: Hongxiang Guan, Dong Feng, Daniel Birgel, Jörn Peckmann, Harry H. Roberts, Nengyou Wu, Duofu Chen , Lipid biomarker patterns reflect different formation environments of mussel- and tubeworm-dominated seep carbonates from the Gulf of Mexico (Atwater Valley and Green Canyon). Chemge (2018), https://doi.org/ 10.1016/j.chemgeo.2018.12.005

This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

ACCEPTED MANUSCRIPT

Lipid biomarker patterns reflect different formation environments of mussel- and tubeworm-dominated seep carbonates from the Gulf of Mexico (Atwater Valley and Green Canyon)

a,b

, Dong Feng

b,c*

, Daniel Birgel

d*

, Jörn Peckmann d, Harry H.

PT

Hongxiang Guan

Key Laboratory of Gas Hydrate, Guangzhou Institute of Energy Conversion, Chinese

SC

a

RI

Roberts e, Nengyou Wu f, Duofu Chen b,g

Academy of Sciences, Guangzhou 510640, China

Laboratory for Marine Mineral Resources, Qingdao National Laboratory for Marine

NU

b

c

MA

Science and Technology, Qingdao 266061, China

CAS Key Laboratory of Ocean and Marginal Sea Geology, South China Sea Institute

Institut für Geologie, Centrum für Erdsystemforschung und Nachhaltigkeit,

PT E

d

D

of Oceanology, Chinese Academy of Sciences, Guangzhou 510301, China

Universität Hamburg, 20146 Hamburg, Germany e

Coastal Studies Institute, Department of Oceanography and Coastal Sciences,

The Key Laboratory of Gas Hydrate, Ministry of Land and Resources, Qingdao

AC

f

CE

Louisiana State University, Baton Rouge, Louisiana 70803, USA

Institute of Marine Geology, Qingdao, 266071, China g

Shanghai Engineering Research Center of Hadal Science and Technology, College of

Marine Sciences, Shanghai Ocean University, Shanghai 201306, China

 Corresponding authors. E-mail addresses: [email protected] (D. Feng), [email protected] (D. Birgel). 1

ACCEPTED MANUSCRIPT Abstract Mussels and tubeworms thriving at many methane seeps typically live in symbiosis with chemosynthetic, chiefly methanotrophic or thiotrophic bacteria. It has been shown that the activities of chemosymbiotic animals can result in large differences in the sedimentary environments of their habitats. Here, we put forward the

PT

concept that such environmental variability can be archived in the lipid biomarker

RI

inventories of authigenic carbonates forming in different, locally confined

SC

environments at seeps, mussel beds and tubeworm bushes in this case. To test this hypothesis, lipid biomarker patterns of carbonates from mussel and tubeworm

NU

environments from two seep sites (Atwater Valley 340 and Green Canyon 852) of the Gulf of Mexico were analyzed. Previous work revealed stronger carbon isotope

MA

fractionation between the methane source and biomarkers of anaerobic methane oxidizing archaea-2 (ANME-2)/sulfate-reducing Desulfosarcina/Desulfococcus (DSS)

D

consortia than for ANME-1/DSS consortia, both performing anaerobic oxidation of

PT E

methane (AOM). Similar δ13Cmethane values were found at the mussel and tubeworm sites from the same seeps and the local microbial consortia also appear to be largely

CE

similar based on the observed AOM biomarker inventories. Yet, a large average offset of 32‰ between the δ13C values of molecular fossils of sulfate-reducing bacteria (SRB)

AC

involved in AOM was observed, with lower values typifying tubeworm carbonates than mussel carbonates. This pattern is interpreted to reflect local effects on isotope fractionation caused by the chemosymbiotic metazoans at mussel- and tubewormdominated sites. At tubeworm-dominated sites, the excess sulfate produced by thiotrophic symbionts of tubeworms and pumped down into the sediment results in persistent production of AOM-derived bicarbonate and the enrichment of

12

C in sub-

surface sediments. Interestingly, tubeworm carbonates also contain high amounts of 2

ACCEPTED MANUSCRIPT non-isoprenoidal dialkyl glycerol diethers (DAGEs) with extreme

13

C depletions,

representing compounds that derived from non-DSS cluster SRB. Most likely,

13

C-

depleted AOM-derived organic intermediates were used as carbon sources by the DAGE-producing non-DSS cluster SRB, possibly performing organoclastic sulfate reduction. Our study identifies significant variation in biomarker patterns between

PT

mussel and tubeworm carbonates at two seep sites in the Gulf of Mexico. Such

RI

variation allows to characterize different habitats at seeps, which are shaped by the

SC

interaction of chemosymbiotic seep metazoans and their symbionts with the local environment. Metazoan community composition apparently controls geobiological

MA

of chemosymbiosis into the rock record.

NU

interaction in seep ecosystems to a large degree, which may allow tracing of the effects

Keywords: Authigenic carbonates, Terminally-branched fatty acids, Dialkyl glycerol

AC

CE

PT E

D

diethers (DAGEs), Tubeworms, Mussels, Gulf of Mexico

3

ACCEPTED MANUSCRIPT 1. Introduction The Gulf of Mexico (GoM) is well known for its hydrocarbon seeps, many of which are typified by dense mussel beds and tubeworm aggregations, as well as exposed authigenic carbonate rocks (Fisher et al., 2007; Roberts et al., 2010a, b; Roberts and Feng, 2013). Macrofaunal communities flourish at deep-sea hydrocarbon

PT

seeps due to chemosynthetic, carbon-fixing bacterial symbionts (Jannasch and Nelson,

RI

1984; Madigan et al., 2002; Duperron et al., 2009; Rubin-Blum et al., 2017). These

SC

chemosynthetic symbionts use reduced compounds, such as methane and sulfide, to yield the energy required to fix carbon and provide a source of carbon to their

NU

metazoan host (Childress et al., 1986; Brooks et al., 1987; Fisher et al., 1987; Duperron et al., 2009, 2014; Duperron and Gros, 2016). The generation of sulfide at seeps is

MA

primarily the result of anaerobic oxidation of methane (AOM) coupled to sulfate reduction (Boetius et al., 2000; Joye et al., 2004; Levin, 2005; Jørgensen and Boetius,

D

2007), which promotes the precipitation of carbonates through an increase in alkalinity

PT E

of pore waters (Berner, 1980; Baker and Burns, 1985; Peckmann and Thiel, 2004). At least seven species of chemosymbiotic bathymodiolin mussels have been

CE

reported from GoM seep habitats (Cordes et al., 2009; Faure et al., 2015; Duperron and Gros, 2016). Large mussels, such as Bathymodiolus childressi, B. brooksi, B. heckerae,

AC

and B. boomerang, usually have one to four types of bacterial symbionts located inside their epithelial cells, whereas the smaller mussels can display a greater diversity of symbionts according to habitat characteristics and locations (Duperron et al., 2008; Laming et al., 2015). Among these seep-related mussels, B. childressi and B. brooksi are the best studied species. Bathymodiolus childressi harbors only methanotrophic bacterial endosymbionts (Fisher et al. 1987), whereas B. brooksi contains both methanotrophic and sulfide-oxidizing symbionts (Fisher et al., 1993; Duperron et al., 4

ACCEPTED MANUSCRIPT 2007). Of the two bacterial symbionts that co-occur in B. brooksi, the methanotrophic symbionts dominate (more than 80% of bacterial rRNA) and the relative abundances can vary in populations from different environments (Fisher et al., 1993; Duperron et al., 2007; Kellermann et al., 2012). Methane-oxidizing bacteria efficiently transform methane carbon into biomass carbon (Brown et al., 1964), and chemosymbiotic

PT

mussels with methanotrophic symbionts consequently take up mostly carbon from

RI

methane oxidation (Childress et al., 1986; Cary et al., 1988; Page et al., 1990).

SC

Vestimentiferan tubeworms lack a digestive system and rely on endosymbiotic, sulfide-oxidizing bacteria for their nutrition (Cavanaugh et al., 1981; Fisher, 1990;

NU

Julian et al., 1999; Cordes et al., 2009). Tubeworms not only take up sulfide from the sulfide-rich sediment surrounding the thin walled, posterior ends of tubes, but also

MA

release significant amounts of sulfate produced by their symbionts to the sediment (Cordes et al., 2005; Dattagupta et al., 2006, 2008). This input of sulfate significantly

D

impacts hydrocarbon-rich sediments, which are often limited in sulfate (Boetius, 2005;

PT E

Cordes et al., 2005; Dattagupta et al., 2006, 2008). In addition, strong variations in tissue δ13C values between larger adults and very small individuals have been observed,

CE

indicating that adult vestimentiferans take up dissolved inorganic carbon (DIC) from pore waters through their posterior ends, while small individuals incorporate inorganic

AC

carbon from the seawater across their plume (Kennicutt et al., 1992; Becker et al., 2011). Laboratory studies indicated that some tubeworms, for example L. luymesi, can also excrete hydrogen ions – byproducts of their sulfide-oxidizing symbionts – across their permeable posterior tube extensions (Dattagupta et al., 2006). It has been suggested that (1) vital effects during carbon uptake by mussels and tubeworms, (2) physical actions by mussels, and (3) the release of sulfate by tubeworms through their posterior ends can substantially impact the sediment 5

ACCEPTED MANUSCRIPT geochemistry of the respective habitats, resulting in changes in the isotopic composition of local pore water DIC pools (cf. Feng and Roberts, 2010; Feng et al., 2013). It was shown that tubeworm carbonates generally have lower δ13C values than mussel carbonates from the same site, which is in good accordance with the fact that tubeworms fuel extra subsurface methane oxidation through the release of sulfate into

PT

the sediment (Feng and Roberts, 2010; Feng et al., 2013). However, in many instances

RI

the differences in δ13C values between tubeworm and mussel carbonates were only within approximately 3‰.

SC

Apart from the seep macrofauna and the variability in δ13C values of seep

NU

carbonates, the distribution of seep-dwelling microbial communities should be monitored to classify seep environments. One way to achieve this goal is to analyze the

MA

molecular fossil inventory and the carbon isotopic composition of individual compounds. Because lipid biomarker patterns are diagnostic for varying environmental

D

conditions, including for example changing seepage intensities (e.g. Peckmann et al.,

PT E

2009; Haas et al., 2010; Birgel et al., 2011), it is to be expected that tubeworm and mussel carbonates archive subtle differences between the environments in which they

CE

had formed. Here, this approach is applied to authigenic carbonates from two sites of the northern GoM, Green Canyon 852 (GC852) and Atwater Valley 340 (AT340),

AC

representing both tubeworm and mussel carbonates that occur at both sites. New biomarker data are reported alongside mineralogical and stable isotope data published by Feng and Roberts (2010) and Feng et al. (2013). It is shown that such a comprehensive dataset allows to (1) tentatively characterize putative microbial abundance and diversity patterns at seeps and (2) constrain ecosystem dynamics.

6

ACCEPTED MANUSCRIPT 2. Geological settings and materials Two tubeworm carbonates (AT340-T and GC852-T) and two mussel carbonates (AT340-M and GC852-M), respectively, were retrieved from tubeworm- and musseldominated seeps of the AT340 and GC852 sites for a comparative study (Fig. 1). Both seep sites are located on the northern middle continental slope of the GoM (Fig. 1,

PT

Table 1). The study areas are characterized by abundant faults caused by salt diapirism.

RI

Dive observations indicate that the crestal areas of the sites are characterized by large

SC

authigenic carbonate outcrops (Roberts et al., 2010a, b). Detailed descriptions and more information on the two sites can be found in Roberts et al. (2010a). At times of

NU

sampling, dense bathymodiolin mussel beds were scattered among the carbonate blocks, and vestimentiferan tubeworms were widespread throughout the ridge crest

MA

area (Roberts et al., 2010a, b; Fig. 2). The carbonate samples analyzed in this study were collected during dives with the remotely operated vehicle (ROV) Jason II in 2007

D

(Table 1). They were exposed at the seafloor at the time of sampling. For the purpose

PT E

of this study, samples particularly rich in either mussels or tubeworms were selected (Fig. 2). Location, water depth, δ13Ccarb and δ18Ocarb values, and mineralogy of the

CE

studied carbonates are listed in Tables 1 and 2. A cross plot of δ13C and δ18O values of

AC

mussel and tubeworm carbonates is shown in Fig. 3.

3. Methods

The detailed protocol applied for preparation and extraction of lipids was described in Guan et al. (2013). Carbonates were crushed to small pieces, cleaned by distilled water and acetone, then freeze-dried, and grounded into powders. The powders were soxhlet-extracted for 72 h with a mixture of dichloromethane and methanol (9:1; v:v). Deuterated n-eicosane (C20D42), heptadecanoic acid, and tridecyl 7

ACCEPTED MANUSCRIPT alcohol were added as internal standards to each sample. The total lipid extracts (TLE) were dried with anhydrous sodium sulfate and saponified with 6% KOH (w/v) in methanol. The neutral fractions were extracted from the saponified TLE with hexane. To obtain carboxylic acids, the residuals were treated with 10% HCL to pH=2 and extracted using hexane. The neutral fractions were separated by column

PT

chromatography into three compound classes of (1) hydrocarbons (n-hexane), (2)

RI

aromatics (n-hexane/dichloromethane, 6:4; v:v), and (3) alcohols (methanol). Fatty

SC

acid methyl esters (FAMEs) and trimethylsiloxyl derivatives were obtained by subjecting the dried carboxylic acid and alcohol fractions to 14% BF3-methanol and

NU

bis(trimethylsilyl) trifluoroacetamide (BSTFA), respectively, heated at 60 °C for 2 h prior to GC-MS analysis. All fractions were analyzed using a Thermo Electron Trace

MA

GC-MS equipped with a 60-m DB-5 MS fused silica capillary column (0.32 mm i.d., 0.25 μm film thickness). Helium was supplied as the carrier gas at a flow rate of 1.2

D

ml/min. The following GC temperature program was used: initial temperature was 60 C, 2 min isothermal, from 60°C to 150 °C at 10 °C/min, from 150 to 320 oC at 4

PT E

o

o

C/min, 30 min isothermal for hydrocarbons and 45 min isothermal for carboxylic

CE

acids and alcohols. Compound-specific carbon isotope analysis was performed on a GV Isoprime system interfaced to a Hewlett-Packard 6890 gas chromatograph. The GC

AC

conditions were the same as those used in GC-MS analysis. The carbon isotope ratios are given as δ-values (δ13C in ‰) relative to the Vienna-PeeDee Belemnite (V-PDB) standard. FAMEs and trimethylsiloxyl derivatives were corrected for the addition of carbon during preparation. All samples were at least analyzed in duplicate. The standard deviation of compound-specific carbon isotope measurements was <0.8‰. For analysis of glycerol dialkyl glycerol tetraethers (GDGTs), an aliquot of the total extract was re-dissolved in hexane/isopropanol (99:1, v:v), before compounds 8

ACCEPTED MANUSCRIPT were

separated

and

identified

chromatography/atmospheric

pressure

with chemical

high

performance

ionization

mass

liquid

spectrometry

(HPLC/APCI-MS) at the Guangzhou Institute of Geochemistry, Chinese Academy of Sciences (GIG, CAS). Separation of GDGTs was achieved with an Altech Prevail Cyano Column (2.1 × 150 mm, 3 μm, Altech, USA) maintained at 30 °C. The

PT

conditions for HPLC/APCI-MS were set as described in Guan et al. (2016). The

RI

GDGTs were analyzed in the single ion monitoring (SIM) mode, and identified based

SC

on accurate mass, retention time, and diagnostic fragments (cf. Liu et al., 2012a). No internal standard was added for quantification of GDGTs. To determine the stable

NU

carbon isotopes of GDGT-cleaved biphytanes, the GDGTs were subjected to ether cleavage as described by Liu et al. (2012b). The ether-cleaved biphytanes were

MA

measured both by gas chromatography-mass spectrometry (GC-MS) and gas chromatography-isotope ratio mass spectrometry (GC-IRMS). Identification of

PT E

D

individual compounds was based on published mass spectra and GC retention times.

4. Results

CE

4.1. Lipid biomarker inventory

All AOM-related compounds are listed in Tables 3 and 4. The highest overall

AC

contents of lipid biomarkers were found for tubeworm carbonate GC852-T (20.8 µg/g rock), followed by tubeworm carbonate AT340-T (11.5 µg/g rock), and mussel carbonate AT340-M (10.6 µg/g rock); the lowest total contents were observed for mussel carbonate GC852-M (4.0 µg/g rock). The alcohol fraction of tubeworm carbonate GC852-T accounted for 69% of all lipid biomarkers, followed by tubeworm carbonate AT340-T, and mussel carbonate AT340-M (48% and 49%, respectively). In mussel carbonate GC852-M, alcohols only made up 17%, but carboxylic acids 9

ACCEPTED MANUSCRIPT accounted for 52% of the total lipid inventory. Hydrocarbons represented 10% to 14% of all lipids in carbonates AT340-T, GC852-T, and AT340-M, whereas in carbonate GC852-M hydrocarbons represented 31% of all lipids. Glycerol dialkyl glycerol tetraethers (GDGTs) were also present in all samples but were not quantified, since we did not add standards to the samples. The relative percentages of GDGTs can be found

PT

in Table S1. Apart from the compounds displayed in Tables 3 and 4, also hopanoids

RI

were found. The only hopanoids recognized were various C32-hopanols and hopanoic

SC

acids (Table S2). Due to their low specificity, they are not further discussed below.

NU

4.2. Oil-derived lipid biomarkers

Short-chain n-alkanes from n-C15 to n-C18 (summing up to 0.5-0.7 μg/g rock)

MA

were prominent in all samples, whereas long-chain n-alkanes from n-C23 to n-C31 represented less than 0.3 μg/g rock in mussel carbonate GC852-M and were close to

D

detection limit in tubeworm carbonates GC852-T and AT340-T as well as in mussel

PT E

carbonate AT340-M. Furthermore, an unresolved complex mixture (UCM) was present

CE

in all hydrocarbon fractions.

4.3. Archaeal lipid biomarkers

AC

Both tubeworm carbonates (AT340-T and GC852-T) and mussel carbonate AT 340-M contained high amounts of the irregular isoprenoid hydrocarbons crocetane (cr) and pentamethylicosane (PMI) with total contents varying from 0.49 μg/g rock to 1.19 μg/g rock (Fig. S1). Even more abundant were the isoprenoid glycerol diethers archaeol, sn2-hydroxyarchaeol, and di-hydroxyarchaeol with total contents ranging from 1.8 μg/g rock (AT340-T) to 5.43 μg/g rock (GC852-T). Crocetane coeluted with the regular isoprenoid phytane (ph), but dominated over phytane in samples GC852-T, 10

ACCEPTED MANUSCRIPT AT340-T, and AT340-M. Only in sample GC852-M, phytane dominated with a cr/ph ratio of 3:7 and was accompanied by only minor PMI (0.17 μg/g rock). The isoprenoidal diethers archaeol and sn2-hydroxyarchaeol were as well only present in minor amounts in the latter sample (Table 3). Other isoprenoids such as phytanic acid, phytanol, sn2-O- and sn3-O-phytanyl glycerol monoethers were found only in traces.

PT

The regular isoprenoid hydrocarbon pristane was identified with contents varying from

RI

0.1 μg/g rock to 0.8 μg/g rock. Whereas most isoprenoid biomarkers yielded δ13C

SC

values ranging from 137‰ to 104‰, indicating methane as major carbon source, pristane was the only isoprenoid with much higher δ13C values (30‰ to 28‰; Table

NU

3), agreeing with an oil-derived source. The combined cr/ph peak reflected a mixture of oil-derived, 13C-rich phytane and methane-derived, 13C-depleted crocetane, resulting

MA

in the significantly higher δ13C values of the cr/ph peak compared to other, almost exclusively methane-derived isoprenoids.

D

In both tubeworm and mussel carbonates the predominant GDGT was GDGT-2

PT E

(36% to 59% of all GDGTs), maximizing in mussel carbonate AT340-M (59%) and minimizing in GC852-M (36%), followed by GDGTs-1 and -3 in various percentages

CE

(Table S1). Only mussel carbonate GC852-M contained almost equal amounts of GDGT-0 and GDGT-2 (35% and 36% of all GDGTs, respectively). Crenarchaeol was

AC

either absent (AT340-T and GC852-M), or accounted for less than 3% of all GDGTs (Fig. 4). The acyclic, monocyclic, and bicyclic biphytanes, derived from ether cleavage of GDGTs, revealed pronounced 13C depletions with average δ13C values varying from 130‰ in sample AT340-T to 116‰ in GC852-M (Table S1).

4.4. Bacterial lipid biomarkers Various characteristic bacterial lipids were detected. Among those were 11

ACCEPTED MANUSCRIPT terminally-branched iso-/anteiso-C15:0-fatty acids (i-/ai-C15:0), i-/ai-C17:0, and n-C16:1ω5, which were found in tubeworm and mussel carbonates (Fig. S2). These compounds have been assigned to sulfate-reducing bacteria (SRB) in seep environments (e.g. Blumenberg et al., 2004; Table 4). Even though the inventory of fatty acids was similar, contents of individual compounds varied significantly between tubeworm and mussel

PT

carbonates. The total contents of i-/ai-C15:0, i-/ai-C17:0, and n-C16:1ω5 in tubeworm

RI

carbonates were as high as 0.80 μg/g rock, whereas the contents in the mussel carbonates were lower with <0.05 to 0.25 μg/g rock (Table 4). The i-/ai-C15:0- and i-/ai-

SC

C17:0-fatty acids yielded lower 13C values in tubeworm carbonate AT340-T (120‰ to

NU

107‰) than in mussel carbonate AT340-M (87‰ to 77‰), reflecting a significant offset in the δ13C values (‒32‰ on average) of these compounds (Table 4).

MA

Unfortunately, the contents of i-/ai-C15:0, i-/ai-C17:0, and n-C16:1ω5 in mussel carbonate GC852-M were too low to measure carbon stable isotope compositions. Even more

D

specific for SRB were the fatty acids n-C17:1ω6 and cycC17:0ω5,6, which were present in

PT E

minor amounts in tubeworm carbonates only; their δ13C values were not determined due to their low contents. Other short-chain n-fatty acids with C14 to C18 carbons

CE

showed only moderate 13C depletion (69‰ to 29‰; Table 4).

AC

The n-alcohols ranging from C14 to C16 and i-/ai-C15-alcohols were found in all samples, whereas n-C17:1-alcohol was found in all samples except for carbonate AT340-M (Table 4). Interestingly, i-/ai-C15-alcohols and n-C17:1-alcohol revealed extreme

13

C depletions from 120‰ to 102‰ in all samples. The saturated n-

alcohols C14 to C16 from tubeworm carbonates yielded similar δ13C values (114‰ to 93‰), whereas those from mussel carbonates were less 13C-depleted (63‰ to 38‰; Table 4). Besides the fatty acids diagnostic for SRB, bacterial, non-isoprenoid mono- and 12

ACCEPTED MANUSCRIPT dialkyl glycerol mono- and diethers (MAGEs and DAGEs) also assigned to SRB occurred in tubeworm and mussel carbonates (Fig. S3). Among the MAGEs, only sn-2 MAGE cycC17:0ω5,6 was found in tubeworm carbonates and in traces in mussel carbonate GC852-M. Considerably higher amounts of DAGEs, dominated by series 2 DAGEs (cf. Pancost et al., 2001a), were found in tubeworm carbonates with contents

PT

ranging from 2.7 μg/g rock to 7.4 μg/g rock for Green Canyon and Atwater Valley

RI

samples (Table 4, Fig. 5). DAGE contents were lower in mussel carbonates (from 0.1 μg/g rock to 0.3 μg/g rock) and were found to be dominated by series 1 DAGEs (Table

SC

4). All MAGEs and DAGEs revealed very negative δ13C values varying from 131‰

NU

to 101‰ (Table 4, Fig. 6).

MA

5. Discussion

5.1 AOM communities associated with mussel and tubeworm carbonates

D

At modern methane seeps, consortia of anaerobic methane oxidizing archaea

PT E

(ANME) and their sulfate reducing partner of the Desulfosarcina/Desulfococcus (DSS) cluster (ANME-1/DSS, ANME-2/DSS) commonly co-occur (Boetius et al., 2000;

CE

Orphan et al., 2002; Elvert et al., 2005). It has been demonstrated that ANME-1/DSS and ANME-2/DSS consortia have their own characteristic lipid biomarker

AC

compositions and isotopic fingerprints (cf. Blumenberg et al., 2004; Nauhaus et al., 2005; Niemann and Elvert, 2008; Knittel and Boetius, 2009; Rossel et al., 2011). A low ratio (1.1) of sn2-hydroxyarchaeol/archaeol is for example specific for the ANME1/DSS consortium, allowing to distinguish ANME-2/DSS from ANME-1/DSS consortia (Blumenberg et al., 2004; Elvert et al., 2005; Niemann and Elvert, 2008; Kellermann et al., 2016). Additionally, crocetane, which is abundant in ANME-2, but only present in minor contents or absent in ANME-1 (Niemann and Elvert, 2008 and reference therein), 13

ACCEPTED MANUSCRIPT is a good marker for ANME-2/DSS dominated systems (e.g. Blumenberg et al., 2004; Haas et al., 2010). Its absence is in accord with ANME-1/DSS dominated seep environments (Haas et al., 2010), and has been used, among other criteria, to identify the dominance of ANME-1 at some ancient seeps (cf. Peckmann et al., 2009; Natalicchio et al., 2015). PMI is abundant in both ANME-1 and ANME-2 and cannot

PT

be used to differentiate the AOM community. The sulfate-reducing partners (DSS) of

RI

ANME-2 can be discriminated from the DSS associated with ANME-1 by the presence

SC

of the fatty acids cycC17:0ω5,6, n-C17:1ω6, as well as the low abundance of ai-C15:0 relative to i-C15:0 (Niemann and Elvert, 2008 and reference therein).

NU

Less diagnostic lipid biomarkers that have been identified in the studied GoM samples including phytanyl-glycerol monoethers, phytanol, and phytanic acid.

MA

Although these compounds are of low source specificity, they all reveal extreme

13

C

depletions and, therefore, can be attributed to ANMEs as well. Most likely, phytanyl

D

monoethers are degradation products of archaeol and sn2-hydroxyarchaeol, as

PT E

indicated by a degradation scheme for GDGTs developed by Liu et al. (2016), which can also be applied to diether lipids. Pristane and phytane detected in the studied GoM

CE

samples, however, are unlikely to be derived from archaeols produced by ANME, because they are much less

13

C-depleted than the archaeols and their degradation

AC

products including phytanic acids and phytanyl monoethers. The presence of an UCM in the hydrocarbon fractions and the high δ13C values rather point to a crude oil source of pristane and phytane, as suggested for other modern oil-seep carbonates (e.g. Birgel et al., 2011). With regard to the mussel carbonate AT340-M and tubeworm carbonate GC852-T, the abundance of crocetane, sn2-hydroxyarchaeol/archaeol ratios >1.1, and the low content of ai-C15:0 relative to i-C15:0 indicate the predominance of ANME-2/DSS 14

ACCEPTED MANUSCRIPT consortia. Moreover, in both cases the carbonate mineralogy is aragonite, which is typically associated with ANME-2/DSS consortia (cf. Haas et al., 2010; Guan et al., 2013). The biomarker pattern of sample AT340-T also mostly reflects the dominance of ANME-2/DSS (presence of cycC17:0ω5,6 and n-C17:1ω6; anteiso-/iso-C15:0: 0.9; high content of crocetane); only the sn2-hydroxyarchaeol/archaeol ratio is <1, possibly

PT

reflecting a mixed community, which is also in accord with the composite

RI

aragonite/high magnesium calcite mineralogy of the sample. Finally, ANME-1/DSS

SC

consortia were apparently predominant (sn2-hydroxyarchaeol/archaeol ratio: 0.2; minor crocetane) during the formation of mussel carbonate GC852-M, agreeing with

NU

its dominantly high magnesium calcite mineralogy (cf. Haas et al., 2010; Guan et al., 2013).

MA

The GDGT patterns detected in the studied samples resemble patterns observed in other seep habitats with the dominance of GDGTs with 0-2 cyclopentane moieties (e.g.

D

Pancost et al., 2001b; Elvert et al., 2005; Birgel et al., 2008; Niemann and Elvert,

PT E

2008). ANME-1/DSS dominated seeps are typified by higher GDGT contents (Blumenberg et al., 2004; Niemann and Elvert, 2008; Rossel et al., 2011), but GDGTs

CE

are also found in ANME-2/DSS dominated environments with similar relative proportions among the different GDGTs (Zhang et al., 2003; Elvert et al., 2005;

AC

Chevalier et al., 2010, 2014; Feng et al., 2014; Guan et al., 2016). Elvert et al. (2005) reported a high abundance of AOM-related GDGTs with one or two cyclopentane rings in the sediments from Hydrate Ridge, reflecting an ANME-2c/DSS dominated system. Apparently GDGTs cannot be used to reliably differentiate between ANME-1/DSS and ANME-2/DSS consortia. This also seems to apply to seep carbonates, in most of which GDGT-2 is predominating. However, in few cases GDGT-0 has been found to be increased and this pattern was assigned to ANME-1 (Stadnitskaia et al., 2008; Feng et 15

ACCEPTED MANUSCRIPT al., 2014; Guan et al., 2016). The higher abundance of GDGT-0 in mussel carbonate GC852-M is best interpreted in the same way. Feng et al. (2014) found a deviation in the isotopic composition of GDGT-0 (measured as acyclic biphytane after ether cleavage) from the other cyclic biphytanes, which has been interpreted to reflect a contribution from methanogenic Euryarchaeota. Such an isotopic offset was not

PT

recorded in sample GC852-M and, therefore, an origin from methanogens is apparently

RI

not the reason for the increase of the GDGT-0 relative proportion. Planktonic

SC

Thaumarchaeota can also be excluded as source of GDGT-0 in the studied samples, because of the absence or very minor contents of crenarchaeol, a compound produced

NU

by planktonic Thaumarchaeota (Jia et al., 2012; Schouten et al., 2013; Elling et al., 2017). The high proportion of GDGT-0 in mussel carbonate GC852-M may indeed be

MA

best interpreted as a signature of ANME-1 dominance, taking into account carbonate mineralogy (high magnesium calcite) and other lipids (see above). In contrast, the

D

predominance of GDGT-2 in the other, aragonite or mixed aragonite/high-magnesium

PT E

calcite samples agrees with the dominance of ANME-2/DSS consortia, except for sample AT 340-T, which might be mixed (see Table 2). To sum up, in some instances

CE

GDGTs may also be used to differentiate between ANME-1/DSS and ANME-2/DSS. Such an assessment should, however, not be based on the overall contents, but should

AC

rather look at the relative proportions of the different GDGTs. Another group of biomarkers occurring in both ANME-1/DSS and ANME-2/DSS dominated systems are DAGEs, which also have been suggested to allow for an assessment of the composition of AOM consortia. Abundant DAGEs have been first recognized in association with ANME-1/DSS dominated habitats (Aloisi et al., 2002; Blumenberg et al., 2004; Stadnitskaia et al., 2005; Bouloubassi et al., 2006). However, later on high amounts of DAGEs were also recognized in ANME-2/DSS dominated 16

ACCEPTED MANUSCRIPT systems independent of the prevalent carbonate mineralogy (Chevalier et al., 2010, 2011). Consequently, the presence or absence of DAGEs cannot be used to decide whether ANME-1/DSS or ANME-2/DSS consortia prevailed. Independent of that question, the two tubeworm carbonates were found to contain many different and more DAGEs (especially series 2) than mussel carbonates, which contain predominantly

PT

DAGE-C30, but also minor series 2 DAGEs in GC 852-M (see Table 4). The

RI

distribution of DAGEs is most likely not necessarily linked to the distribution of

SC

ANMEs and their adaptation to various ecological parameters (see chapter 5.3).

NU

5.2 Influence of the macrofauna on environmental isotope patterns As observed for many seep carbonates, lipid biomarker patterns in conjunction

MA

with carbonate microfabrics and mineralogy can be used to identify the dominant ANME consortia and the environmental conditions that prevailed at seep sites (e.g.

D

Peckmann et al., 2009). Moreover, the 13C values of archaeol and sn2-

PT E

hydroxyarchaeol can be used to calculate the isotopic composition of parent methane (Niemann and Elvert, 2008). This approach has been confirmed for carbonates from

CE

modern seep sites (Birgel et al., 2011; Himmler et al., 2015). For ANME-2, a  value of approximately 50‰ from methane to lipids has been found, for ANME-1 a  value

AC

of approximately 30‰. Based on the average δ13C values of the respective archaeal lipids, calculated δ13Cmethane values of 80‰ and 73‰ are obtained for the Green Canyon site based on the mussel carbonate and tubeworm carbonate, respectively, which are interpreted as ANME-1 and ANME-2 dominated systems, respectively. From the Atwater Valley samples, a δ13Cmethane value of 74‰ is calculated from the mussel carbonate, whereas the mixed ANME-2/ANME-1 consortium of the tubeworm carbonate (assuming a 50:50 proportion) results in a value of 78‰. For both sites, the 17

ACCEPTED MANUSCRIPT calculated δ13Cmethane values for tubeworm and mussel collections from the same site are very close to each other, with a variability of below 10‰. It has been demonstrated that tubeworms acquire sulfide produced in the sulfatemethane transition zone from the sediment through their posterior ends, providing their thiotrophic symbionts with sulfide. In return, tubeworms release sulfate and hydrogen

PT

ions into the sediment (Dattagupta et al., 2006). Cordes et al. (2005) modeled sulfide

RI

supply and requirements of adult L. luymesi and concluded that sulfide production

SC

mainly by microbial hydrocarbon degradation is sufficient to support moderate-sized tubeworm aggregations for hundreds of years, assuming that tubeworms release sulfate

NU

through their posterior ends. In addition, large adult tubeworms take up DIC from the sediment pore water through their posterior ends. This is confirmed by the fact that

MA

tubeworm tissue is typically characterized by pronounced

13

C depletion in seep

environments (Kennicutt et al., 1992; Roberts and Aharon, 1994; Becker et al., 2011).

D

Consequently, the activity of tubeworms intensifies sulfate-driven AOM by releasing

PT E

sulfate to the sediment, resulting in continued production of methane-derived DIC in sediments and the enrichment of

12

C within the sediment surrounding the posterior

CE

ends of tubeworms. Indeed, highly elevated concentrations of DIC in the pore water surrounding the permeable posterior tube have been documented (Joye et al., 2004).

AC

Feng and Roberts (2010) and Feng et al. (2013) already compared authigenic carbonates from mussel and tubeworm environments. These authors found that tubeworm carbonates were generally more

13

C-depleted than mussel carbonates from

the same site, which was in good accordance with the envisioned effect of tubeworms fueling extra subsurface methane oxidation through the release of sulfate into the sediment. In the course of this process, the persistent production of leads to strongly

13

13

C-depleted DIC

C-depleted SRB lipids, because the SRB of the AOM consortium 18

ACCEPTED MANUSCRIPT take up DIC (cf. Hinrichs and Boetius, 2002; Wegener et al., 2008 and references therein). Interestingly, in contrast to the relatively small differences between δ13Ccarbonate values of tubeworm and mussel lithologies, the differences between δ13Clipid values of compounds assigned to SRB is found to be particularly large (32‰ on average) when these two types of seep carbonates are compared, possibly

PT

suggesting that lipid biomarkers are more sensitive than carbonate isotope data to

RI

environmental variations, including variations caused by macrofaunal activity.

NU

metabolisms apart from sulfate-driven AOM.

SC

However, such variability may also reflect different microbial populations with other

5.3 Abundant DAGEs and their degradation products in tubeworm carbonates

MA

Apart from the characteristic lipid biomarkers of the ANME-1 and ANME-2 consortia discussed above, the most striking observation made on our samples are the

D

high amounts of 13C-depleted DAGEs. Although both tubeworm and mussel carbonates

PT E

contained DAGEs, tubeworm carbonates revealed much higher contents of this group of compounds (Table 4).

CE

The occurrence of 13C-depleted DAGEs in various AOM environments had been originally attributed to syntrophic SRB involved in AOM (e.g. Hinrichs et al., 2000;

AC

Pancost et al., 2001a), since some SRB in culture were known to produce DAGEs and MAGEs (Langworthy et al., 1983; Rütters et al., 2001). However, the SRBs living in syntrophy with the methane-oxidizing archaea were not found to produce DAGEs and MAGEs in culture. Moreover, for many seeps a large offset of δ13C values between DAGEs and the characteristic AOM-specific fatty acids of SRB has been reported, as for example for carbonates from the Nordic margin of the Norwegian Sea (Chevalier et al., 2010). Based on these observations, it was suggested that DAGEs may derive from 19

ACCEPTED MANUSCRIPT SRB species other than those involved in AOM (Chevalier et al., 2010). It was further suggested that the putative non-DSS cluster SRB grow on

13

C-depleted organic

intermediates (Neretin et al., 2007; Orcutt and Meile, 2008; Chevalier et al., 2010). During culture experiments with hydrocarbon-degrading sulfate reducers producing DAGEs, it was found that these SRB use hydrocarbons or fatty acids to build ether-

PT

bond membrane lipids (Grossi et al., 2015; Vinçon-Laugier et al., 2016). This

RI

observation may explain why the fatty acids of the SRB of the AOM consortia have

SC

similar alkyl chains like the seep DAGEs.

In the GoM samples studied here, the full range of series 1 and 2 compounds was

NU

found in tubeworm carbonates with series 2 DAGEs predominating, including the same alkyl chains like the fatty acids derived from SRB involved in AOM. In mussel

MA

carbonates, series 2 DAGEs were absent or occurred only in minor amounts, but series 1 DAGEs were present. DAGE-producing non-DSS cluster SRB may have grown on 13

D

C-depleted short-chain fatty acids or longer alkyl chains in tubeworm and mussel

PT E

dominated habitats. However, with regard to the environment in which mussel carbonates formed, non-DSS cluster SRB did apparently not use alkyl chains or fatty

CE

acids derived from the SRB involved in AOM, but rather used fatty acids of other, possibly heterotrophic bacteria. Because the fatty acids in the mussel carbonates

AC

showed different isotopic compositions than DAGEs, these fatty acids were apparently not used for DAGE biosynthesis. In tubeworm carbonates, the alcohols n-C14, n-C15, n-C16, i-/ai-C15, and n-C17:1 have δ13C values in the same range as i-/ai-C15-fatty acids. Since the latter are known to be products of SRB involved in AOM, the alcohols with similar

13

C depletions are

biosynthetic intermediates, by-products of DAGE biosynthesis using AOM lipids (e.g. Vinçon-Laugier et al., 2016), or degradation products as suggested by Liu et al. (2016) 20

ACCEPTED MANUSCRIPT for GDGT-derived compounds. However, in the mussel carbonate AT340-M, the difference in δ13C values between i-/ai-C15-alcohols and i-/ai-C15-fatty acids was 30‰, indicating that the strongly

13

C-depleted i-/ai-C15-alcohols originated from other

sources. The i-/ai-C15-alcohols are probably either degradation products of DAGEs or biosynthetic intermediates as indicated by (1) the same molecular structures as that of

PT

the ether-bound alkyl chains of DAGEs C30:0 (i-C15:0/ai-C15:0) and DAGEs C30:0 (ai-

RI

C15:0/ai-C15:0) and (2) the similar range of δ13C values of the corresponding DAGEs

SC

(Table 4 and Fig. 6).

Interestingly, the n-C14, n-C15, and n-C16-alcohols in mussel carbonates revealed 13

C enrichment compared to i-/ai-C15-alcohols, pointing again to different

NU

significant

sources, which is further supported by the absence of DAGEs with n-C14 to n-C16 alkyl

MA

chains in mussel carbonates. Although n-C14, n-C15, and n-C16-alcohols are ubiquitous, non-diagnostic lipids sourced by various bacteria, their strong 13C depletion, along with

D

the 13C depletion of the n-C17:1-alcohol, accounts for a derivation from non-DSS cluster

PT E

SRB in case of the tubeworm carbonates. These compounds probably represent breakdown products or biosynthetic intermediates of DAGEs, given the significant

CE

amounts of series 2 DAGEs and their similar δ13C signatures in tubeworm carbonates. Liu et al. (2016) suggested that hydrolysis of different ether bonds could result in

AC

discrete series of hydroxyl derivatives, and oxidation of each terminal hydroxyl functional group may generate related carboxyl products. Accordingly, the n-C14, n-C15, n-C16, i-/ai-C15, and n-C17:1-alcohols in tubeworm carbonates were most likely hydrolysis products of DAGEs. However, if a degradation scenario as suggested by Liu et al. (2016) is valid in this case, we would have expected to find MAGEs too, which is only partially the case. For that reason, it cannot be excluded that some of these compounds are rather biosynthetic intermediates. 21

ACCEPTED MANUSCRIPT Indeed, various SRB species other than members of the DSS cluster have been reported to co-occur with methanotrophic archaea at marine seeps (Orphan et al., 2002; Knittel et al., 2003; Nauhaus et al., 2005). In culture experiments, increased sulfate reduction rates have been attributed to the enhanced growth of non-DSS cluster SRB induced by the addition of hydrogen or formate (Nauhaus et al., 2005; Wegener et al., 13

C-depleted molecular fossils of SRB of

PT

2015). In the present study, the extremely

RI

tubeworm carbonates are argued to reflect excess sulfate delivered to the sediment by

SC

tubeworms, which intensified sulfate-driven AOM and this process in turn favored

NU

organoclastic sulfate reduction by DAGE-producing non-DSS cluster SRB.

6. Conclusions

MA

Authigenic carbonates from mussel and tubeworm environments collected from AT340 and GC852 seep sites of the Gulf of Mexico revealed varying contents of 13C-

D

depleted archaeal and bacterial biomarkers, reflecting their formation as a result of

PT E

anaerobic oxidation of methane under varying conditions. Individual biomarker patterns revealed differences in the microbial consortia with changing abundances of

CE

ANME-1 to ANME-2 archaea, consuming methane typified by similar δ13C values at all seeps, and reflected by the dominance of authigenic aragonite in the case of ANME-

AC

2 dominated systems and by the dominance of calcite in the case of ANME-1 dominated systems. The large difference in δ13C values (‒32‰ on average) of terminal branched fatty acids between tubeworm carbonates and mussel carbonates is interpreted to reflect the impact of the distinct modes of chemosymbiosis on the local geochemical conditions. The persistent release of sulfate from the permeable posterior tube extensions of tubeworms promotes the production of bicarbonate in the sediment and results in the enrichment of 12C in the local carbon pool. In comparison to mussel 22

ACCEPTED MANUSCRIPT carbonates, the much higher amounts of

13

C-depleted DAGEs and their hydrolysis

products in tubeworm carbonates are interpreted to reflect input from organoclastic, non-DSS cluster SRB, which contribute to an increased sulfate reduction induced by the supply of sulfate by tubeworms. The differences in biomarker patterns between mussel and tubeworm carbonates reflect distinct types of chemosymbiosis and may be

RI

PT

used to trace the effects of chemosymbiosis into the rock record.

SC

Acknowledgments

The authors thank BOEM and NOAA for their support of the Gulf of Mexico deep-

NU

sea dives. We express our sincere appreciation to the crews of the ROV Jason II and R/V Ron Brown for their professionalism in sampling. We thank Y. Tian and J. He

MA

(GIG, CAS) for sample preparation and technical assistance. The research was partially supported by Qingdao National Laboratory for Marine Science and Technology (Grant:

D

QNLM2016ORP0210) and the NSF of China (Grants: 41473080, 41761134084, and

PT E

41730528). We thank the journal editor H.L. Dong, V. Grossi, and one anonymous

AC

CE

reviewer for constructive comments, which considerably improved the manuscript.

23

ACCEPTED MANUSCRIPT References Aloisi, G., Bouloubassi, I., Heijs, S.K., Pancost, R.D., Pierre, C., Sinninghe Damsté, J.S., Gottschal, J.C., Forney, L.J., Rouchy, J.-M., 2002. CH4-consuming microorganisms and the formation of carbonate crusts at cold seeps. Earth Planet. Sci. Lett. 203, 195–203.

PT

Baker, P.A., Burns, S.J., 1985. Occurrence and formation of dolomite in organic-rich

RI

continental-margin sediments. AAPG Bull. 69, 1917–1930.

SC

Becker, E.L., Macko, S.A., Lee, R.W., Fisher, C.R., 2011. Stable isotopes provide new insights into vestimentiferan physiological ecology at Gulf of Mexico cold seeps.

NU

Naturwissenschaften 98, 169–174.

Berner, R.A., 1980. Early Diagenesis—A Theoretical Approach. Princeton University

MA

Press, Princeton.

Birgel, D., Elvert, M., Han, X., Peckmann, J., 2008. 13C-depleted biphytanic diacids as

D

tracers of past anaerobic oxidation of methane. Org. Geochem. 39, 152–156.

PT E

Birgel, D., Feng, D., Roberts, H.H., Peckmann, J., 2011. Changing redox conditions at cold seeps as revealed by authigenic carbonates from Alaminos Canyon, northern

CE

Gulf of Mexico. Chem. Geol. 285, 82–96. Blumenberg, M., Seifert, R., Reitner, J., Pape, T., Michaelis, W., 2004. Membrane lipid

AC

patterns typify distinct anaerobic methanotrophic consortia. P. Natl. Acad. Sci. USA 101, 11111–11116. Boetius, A., 2005. Microfauna-macrofauna interaction in the seafloor: lessons from the tubeworm. PLOS Biol. 3, 375–378. Boetius, A., Ravenschlag, K., Schubert, C.J., Rickert, D., Widdel, F., Gieseke, A., Amann, R., Jørgensen, B.B., Witte, U., Pfannkuche, O., 2000. A marine microbial consortium apparently mediating anaerobic oxidation of methane. Nature 407, 24

ACCEPTED MANUSCRIPT 623–626. Bouloubassi, I., Aloisi, G., Pancost, R.D., Hopmans, E., Pierre, C., Sinninghe Damsté, J.S., 2006. Archaeal and bacterial lipids in authigenic carbonate crusts from eastern Mediterranean mud volcanoes. Org. Geochem. 37, 484–500. Brooks, J.M., Kennicutt, M.C., Fisher, C.R., Macko, S.A., Cole, K., Childress, J.J.,

PT

Bidigare, R., Vetter, R.D., 1987. Deep-sea hydrocarbon seep communities:

RI

evidence for energy and nutritional carbon sources. Science 238, 1138–1142.

SC

Brown, L.R., Strawinski, R.J., McCleskey, C.S., 1964. The isolation and characterization of Methanomonas methanooxidans Brown and Strawinski. Can. J.

NU

Microbiol. 10, 791–799.

Cary, S.C., Fisher, C.R., Felbeck, H., 1988. Mussel growth supported by methane as

MA

sole carbon and energy source. Science 240, 78–80. Cavanaugh, C.M., Gardiner, S.L., Jones, M.L., Jannasch, H.W., Waterbury, J.B., 1981.

D

Prokaryotic cells in the hydrothermal vent tube worm Riftia pachyptila Jones:

PT E

Possible chemoautotrophic symbionts. Science 213, 340–342. Chevalier, N., Bouloubassi, I., Birgel, D., Crémière, A., Taphanel, M.-H., Pierre, C.,

CE

2011. Authigenic carbonates at cold seeps in the Marmara Sea (Turkey): A lipid biomarker and stable carbon and oxygen isotope investigation. Mar. Geol. 288,

AC

112–121.

Chevalier, N., Bouloubassi, I., Stadnitskaia, A., Taphanel, M.H., Lorre, A., Sinninghe Damsté, J.S., Pierre, C., 2010. Distributions and carbon isotopic compositions of lipid biomarkers in authigenic carbonate crusts from the Nordic Margin (Norwegian Sea). Org. Geochem. 41, 885–890. Chevalier, N., Bouloubassi, I., Stadnitskaia, A., Taphanel, M.H., Sinninghe Damsté, J.S., 2014. Lipid biomarkers for anaerobic oxidation of methane and sulphate 25

ACCEPTED MANUSCRIPT reduction in cold seep sediments of Nyegga pockmarks (Norwegian margin): discrepancies in contents and carbon isotope signatures. Geo-Mar. Lett. 34, 269– 280. Childress, J.J., Fisher, C.R., Brooks, J.M., Kennicutt, M.C., Bidigare, R., Anderson, A.E., 1986. A methanotrophic marine molluscan (Bivalvia: Mytilidae) symbiosis:

PT

Mussels fueled by gas. Science 233, 1306–1308.

RI

Cordes, E.E., Arthur, M.A., Shea, K., Arvidson, R.S., Fisher, C.R., 2005. Modeling the

SC

mutualistic interactions between tubeworms and microbial consortia. PLOS Biol. 3, 497–506.

NU

Cordes, E.E., Bergquist, D.C., Fisher, C.R., 2009. Macro-ecology of Gulf of Mexico cold seeps. Annu. Rev. Mar. Sci. 1, 143–168.

MA

Dattagupta, S., Arthur, M.A., Fisher, C.R., 2008. Modification of sediment geochemistry by the hydrocarbon seep tubeworm Lamellibrachia luymesi: A

PT E

2298–2315.

D

combined empirical and modeling approach. Geochim. Cosmochim. Acta 72,

Dattagupta, S., Miles, L.L., Barnabei, M.S., Fisher, C.R., 2006. The hydrocarbon seep

CE

tubeworm Lamellibrachia luymesi primarily eliminates sulfate and hydrogen ions across its roots to conserve energy and ensure sulfide supply. J. Exp. Biol. 209,

AC

3795–805.

Duperron, S., Gaudron, S.M. Lemaitre, N., Bayon, G., 2014. A microbiological and biogeochemical investigation of the cold seep tubeworm Escarpia southwardae (Annelida: Siboglinidae): Symbiosis and trace element composition of the tube. Deep-Sea Res. I 90, 105–114. Duperron, S., Gros, O., 2016. Colwellia and sulfur-oxidizing bacteria: An unusual dual symbiosis in a Terua mussel (Mytilidae: Bathymodiolinae) from whale falls in the 26

ACCEPTED MANUSCRIPT Antilles arc. Deep-Sea Res. I 115, 112–122. Duperron, S., Halary, S., Lorion, J., Sibuet, M., Gaill, F., 2008. Unexpected cooccurence of six bacterial symbionts in the gill of the cold seep mussel Idas sp. (Bivalvia: Mytilidae). Environ. Microbiol. 10, 433–445. Duperron, S., Lorion, J., Samadi, S., Gros, O., Gaill, F., 2009. Symbioses between

PT

deep-sea mussels (Mytilidae: Bathymodiolinae) and chemosynthetic bacteria:

RI

diversity, function and evolution. C. R. Biol. 332, 298–310.

SC

Duperron, S., Sibuet, M., MacGregor, B.J., Kuypers, M.M.M., Fisher, C.R., Dubilier, N., 2007. Diversity, relative abundance and metabolic potential of bacterial

NU

endosymbionts in three Bathymodiolus mussel species from cold seeps in the Gulf of Mexico. Environ. Microbiol. 9, 1423–38.

MA

Elling, F.J., Könneke, M., Nicol, G.W., Stiegelmeier, M., Bayer, B., Spieck, E., de laTorre, J.R., Becker, K.W., Thomm, M., Prosser, J.I., Herndl, G.J., Schleper, C.,

D

Hinrichs, K.-U., 2017. Chemotaxonomic characterisation of the thaumarchaeal

PT E

lipidome. Environ. Microbiol. 19, 2681–2700. Elvert, M., Hopmans, E.C., Treude, T., Boetius, A., Suess, E., 2005. Spatial variations

CE

of methanotrophic consortia at cold methane seeps: implications from a highresolution molecular and isotopic approach. Geobiology 3, 195–209.

AC

Faure, B., Schaeffer, S.W., Fisher, C.R., 2015. Species distribution and population connectivity of deep-sea mussels at hydrocarbon seeps in the Gulf of Mexico. PLOS One 10, e0118460. Feng, D, Cordes, E.E., Roberts, H.H., Fisher, C.R., 2013. A comparative study of authigenic carbonates from mussel and tubeworm environments: Implications for discriminating the effects of tubeworms. Deep-Sea Res. I 75, 110–118. Feng, D., Birgel, D., Peckmann, J., Roberts, H.H., Joye, S.B., Sassen, R., Liu, X.L., 27

ACCEPTED MANUSCRIPT Hinrichs, K-U., Chen, D., 2014. Time integrated variation of sources of fluids and seepage dynamics archived in authigenic carbonates from Gulf of Mexico gas hydrate seafloor observatory. Chem. Geol. 385, 129–139. Feng, D., Roberts, H.H., 2010. Initial results of comparing cold-seep carbonates from mussel- and tubeworm-associated environments at Atwater Valley lease block 340,

PT

northern Gulf of Mexico. Deep-Sea Res. II 57, 2030–2039.

RI

Fisher, C., Roberts, H., Cordes, E., Bernard, B., 2007. Cold seeps and associated

SC

communities of the Gulf of Mexico. Oceanography 20, 118–129. Fisher, C.R., 1990. Chemoautotrophic and methanotrophic symbioses in marine

NU

invertebrates. Rev. Aquat. 2, 399–436.

Fisher, C.R., 1993. Oxidation of methane by deep-sea mytilids in the Gulf of Mexico.

MA

In Biochemistry of Global Change: Radiatively Active Trace Gases. Oremland, R.S. (ed.). New York, NY, USA: Chapman and Hall, pp. 606–618.

D

Fisher, C.R., Childress, J.J., Oremland, R.S., Bidigare, R.R., 1987. The importance of

PT E

methane and thiosulfate in the metabolism of the bacterial symbionts of two deepsea mussels. Mar. Biol. 96, 59–71.

CE

Grossi, V., Mollex, D., Vinçon-Laugier, A., Hakil, F., Pacton, M., Cravo-Laureau, C., 2015. Mono- and dialkyl glycerol ether lipids in anaerobic bacteria: biosynthetic

AC

insights from the mesophilic sulfate reducer Desulfatibacillum alkenivorans PF2803T. Appl. Environ. Microbiol. 81, 3157–3168. Guan, H., Feng, D., Wu, N., Chen, D., 2016. Methane seepage intensities traced by biomarker patterns in authigenic carbonates from the South China Sea. Org. Geochem. 91, 109–119. Guan, H., Sun, Y., Zhu, X., Mao, S., Feng, D., Wu, N., Chen, D., 2013. Factors controlling the types of microbial consortia in cold-seep environments: A 28

ACCEPTED MANUSCRIPT molecular and isotopic investigation of authigenic carbonates from the South China Sea. Chem. Geol. 354, 55–64. Haas, A., Peckmann, J., Elvert, M., Sahling, H., Bohrmann, G., 2010. Patterns of carbonate authigenesis at the Kouilou pockmarks on the Congo deep-sea fan. Mar. Geol. 268, 129–136.

Formation of seep carbonates along the Makran convergent margin,

RI

2015.

PT

Himmler, T., Birgel, D., Bayon, G., Pape, T., Ge, L., Bohrmann, G., Peckmann, J.,

SC

northern Arabian Sea and a molecular and isotopic approach to constrain the carbon isotopic composition of parent methane. Chem. Geol. 415, 102–117.

NU

Hinrichs, K.-U., Boetius, A., 2002. The anaerobic oxidation of methane: new insights in microbial ecology and biogeochemistry. In: Wefer, G., Billett, D., Hebbeln, D.,

MA

Jørgensen, B.B., Schlüter, M., van Weering, T.C.E. (Eds.), Ocean Margin Systems. Springer, pp. 457–477.

D

Hinrichs, K.-U., Summons, R.E., Orphan, V., Sylva, S.P., Hayes, J.M., 2000. Molecular

PT E

and isotopic analysis of anaerobic methane-oxidizing communities in marine sediments. Org. Geochem. 31, 1685–1701.

CE

Jannasch, H.W., Nelson, D.C., 1984. Recent progress in the microbiology of hydrothermal vents, in: M.J. Klug, C.A. Reddy (Eds.), Current Perspectives in

AC

Microbial Ecology, American Society for Microbiology, Washington, DC, 1984, pp. 170–175.

Jia, G., Zhang, J., Chen, J., Peng, P., Zhang, C.L., 2012. Subsurface water temperatures recorded by archaeal tetraether lipids in the South China Sea. Org. Geochem. 50, 68–77. Jørgensen, B.B., Boetius, A., 2007. Feast and famine—microbial life in the deep-sea bed. Nat. Rev. Microbiol. 5, 770–781. 29

ACCEPTED MANUSCRIPT Joye, S.B., Boetius, A., Orcutt, B.N., Montoya, J.P., Schulz, H.N., Erickson, M.J., Lugo, S.K., 2004. The anaerobic oxidation of methane and sulfate reduction in sediments from Gulf of Mexico cold seeps. Chem. Geol. 205, 219–238. Julian, D., Gaill, F., Wood, E., Arp, A., Fisher, C., 1999. Roots as a site of hydrogen sulfide uptake in the hydrocarbon seep vestimentiferan Lamellibrachia sp. J. Exp.

PT

Biol. 202, 2245–2257.

N.,

Hinrichs,

K.-U.,

2012.

Symbiont–host

relationships

in

SC

Dubilier,

RI

Kellermann, M.Y., Schubotz, F., Elvert, M., Lipp, J.S. Birgel, D., Prieto-Mollar, X.,

chemosynthetic mussels: A comprehensive lipid biomarker study. Org. Geochem.

NU

43, 112–124.

Kellermann, M.Y., Yoshinaga, M.Y., Wegener, G., Krukenberg, V., Hinrichs, K.-U.,

MA

2016. Tracing the production and fate of individual archaeal intact polar lipids using stable isotope probing. Org. Geochem. 95, 13–20.

D

Kennicutt II, M.C., Burke, R.A., Jr, MacDonald, I.R., Brooks, J.M., Denoux, G.J.,

PT E

Macko, S.A. 1992. Stable isotope partitioning in seep and vent organisms: chemical and ecological significance. Chem. Geol. 101, 293–310.

CE

Knittel, K., Boetius, A., 2009. The anaerobic oxidation of methane—progress with an unknown process. Annu. Rev. Microbiol. 63, 311–334.

AC

Knittel, K., Boetius, A., Lemke, A., Eilers, H., Lochte, K., Pfannkuche, O., Linke, P., Amann, R., 2003. Activity, distribution, and diversity of sulfate reducers and other bacteria in sediments above gas hydrate (Cascadia margin, Oregon). Geomicrobiol. J. 20, 269–294. Laming, S.R., Szafranski, K.M., Rodrigues, C.F., Gaudron, S.M., Cunha, M.R., Hilário A., Bris, N.L., Duperron, S., 2015. Fickle or faithful: the roles of host and environmental

context

in

determining 30

symbiont

composition

in

two

ACCEPTED MANUSCRIPT bathymodioline mussels. PLOS One 10, e0144307. Langworthy, T.A., Holzer, G., Zeikus, J.G., Tornabene, T.G., 1983. Iso- and anteisobranched

glycerol

diethers

of

the

thermophilic

anaerobe

Thermodesulfotobacterium commune. Syst. Appl. Microbiol. 4, 1–17. Levin, L.A., 2005. Ecology of cold seep sediments: Interactions of fauna with flow,

PT

chemistry and microbes. Oceanogr. Mar. Biol. 43, 1–46.

RI

Liu, X.-L., Birgel, D., Elling, F.J., Sutton, P.A., Lipp, J.S., Zhu, R., Zhang, C.,

SC

Könneke, M., Peckmann, J., Rowland, S.J., Summons R.E., Hinrichs K.-U., 2016. From ether to acid: A plausible degradation pathway of glycerol dialkyl glycerol

NU

tetraethers. Geochim. Cosmochim. Acta 183,138–152.

Liu, X.-L., Lipp, J.S., Schröder, J.M., Summons, R.E., Hinrichs, K.-U., 2012b.

MA

Isoprenoid glycerol dialkanol diethers: A series of novel archaeal lipids in marine sediments. Org. Geochem. 43, 50–55.

D

Liu, X.-L., Summons, R.E., Hinrichs, K.-U., 2012a. Extending the known range of

PT E

glycerol ether lipids in the environment: structural assignments based on tandem mass spectral fragmentation patterns. Rapid Commun. Mass Sp. 26, 2295–2302.

CE

Madigan, M.T., Martinko, J.M., Parker, J., 2002. Brock Biology of Microorganisms, Pearson Education.

AC

Natalicchio, M., Peckmann, J., Birgel, D., Kiel, S., 2015. Seep deposits from northern Istria, Croatia: a first glimpse into the Eocene seep fauna of the Tethys region. Geol. Mag. 152, 444–459. Nauhaus, K., Treude, T., Boetius, A., Krüger, M., 2005. Environmental regulation of the anaerobic oxidation of methane: a comparison of ANME-1 and ANME-II communities. Environ. Microbiol. 7, 98–106. Neretin, L.N., Abed, R.M., Schippers, A., Schubert, C.J., Kohls, K., Kuypers, M.M., 31

ACCEPTED MANUSCRIPT 2007. Inorganic carbon fixation by sulfate-reducing bacteria in the Black Sea water column. Environ. Microbiol. 9, 3019–3024. Niemann, H., Elvert, M., 2008. Diagnostic lipid biomarker and stable carbon isotope signatures of microbial communities mediating the anaerobic oxidation of methane with sulphate. Org. Geochem. 39, 1668–1677.

PT

Orcutt, B., Meile, C., 2008. Constraints on mechanisms and rates of anaerobic

RI

oxidation of methane by microbial consortia: process-based modeling of ANME-2

SC

archaea and sulfate reducing bacteria interactions. Biogeosciences 5, 1587–1599. Orphan, V.J., House, C.H., Hinrichs, K.-U., McKeegan, K.D., DeLong, E.F., 2002.

NU

Multiple archaeal groups mediate methane oxidation in anoxic cold seep sediments. P. Natl. Acad. Sci. USA 99, 7663–7668.

MA

Page, H.M., Fisher, C.R., Childress, J.J., 1990. Role of filter-feeding in the nutritional biology of a deep-sea mussel with methanotrophic symbionts. Mar. Biol. 104,

D

251–257.

PT E

Pancost, R.D., Bouloubassi, I., Aloisi, G., Sinninghe Damsté, J.S., MEDINAUT scientific party, 2001a. Three series of non isoprenoidal dialkyl glycerol diethers

CE

in cold seeps carbonate crusts. Org. Geochem. 32, 695–707. Pancost, R.D., Hopmans, E.C., Sinninghe Damsté, J.S., MEDINAUT scientific party,

AC

2001b. Archaeal lipids in Mediterranean Cold Seeps: Molecular proxies for anaerobic methane oxidation. Geochim. Cosmochim. Acta 65, 1611–1627. Peckmann, J., Birgel, D., Kiel, S., 2009. Molecular fossils reveal fluid composition and flow intensity at a Cretaceous seep. Geology 37, 847–850. Peckmann, J., Thiel, V., 2004. Carbon cycling at ancient methane-seeps. Chem. Geol. 205, 443–467. Roberts, H.H., Aharon, P., 1994. Hydrocarbon-derived carbonate buildups of the 32

ACCEPTED MANUSCRIPT northern Gulf of Mexico continental slope: a review of submersible investigations. Geo-Mar. Lett. 14, 135–148. Roberts, H.H., Feng, D., 2013. Carbonate precipitation at Gulf of Mexico hydrocarbon seeps: An overview, in Abrams M, Aminzadeh, F, Berge T, Connolly D, O’Brien G. (eds.) Hydrocarbon Seepage: From Source to Surface: SEG/AAPG Special

PT

Publication, 43–61.

RI

Roberts, H.H., Feng, D., Joye, S.B., 2010b. Cold-seep carbonates of the middle and

SC

lower continental slope, northern Gulf of Mexico. Deep-Sea Res. II 57, 2040– 2054.

NU

Roberts, H.H., Shedd, W., Hunt Jr, J., 2010a. Dive site geology: DSV ALVIN (2006) and ROV JASON II (2007) dives to the middle-lower continental slope, northern

MA

Gulf of Mexico. Deep-Sea Res. II 57, 1837–1858. Rossel, P.E., Elvert, M., Ramette, A., Boetius, A., Hinrichs, K.-U., 2011. Factors

D

controlling the distribution of anaerobic methanotrophic communities in marine

PT E

environments: evidence from intact polar membrane lipids. Geochim. Cosmochim. Acta 75, 164–184.

CE

Rubin-Blum, M., Paul Antony, C., Borowski, C., Sayavedra, L., Pape, T., Sahling, H., Bohrmann, G., Kleiner, M., Redmond, M.C., Valentine, D.L., Dubilier, N., 2017.

AC

Short-chain alkanes fuel mussel and sponge Cycloclasticus symbionts from deepsea gas and oil seeps. Nat. Microbiol. 2, 17093. Rütters, H., Sass, H., Cypionka, H., Rullkötter, J., 2001. Monoalkylether phospholipids in the sulfate-reducing bacteria Desulfosarcina variabilis and Desulforhabdus amnigenus. Arch. Microbiol. 176, 435–442. Schouten, S., Hopmans, E.C., Sinninghe Damsté, J.S., 2013. The organic geochemistry of glycerol dialkyl glycerol tetraether lipids: A review. Org. Geochem. 54 19–61. 33

ACCEPTED MANUSCRIPT Stadnitskaia, A., Muyzer, G., Abbas, B., Coolen, M.J.L., Hopmans, E.C., Bass, M., van Weering, T.C.E., Ivanov, M.K., Poludetkina, E., Sinninghe Damsté, J.S., 2005. Biomarker and 16S rDNA evidence for anaerobic oxidation of methane and related carbonate precipitation in deep-sea mud volcanoes of the Sorokin Trough, Black Sea. Mar. Geol. 217, 67–96.

PT

Stadnitskaia, A., Nadezhkin, D., Abbas, B., Blinova, V., Ivanov, M.K., Sinninghe

RI

Damsté, J.S., 2008. Carbonate formation by anaerobic oxidation of methane:

SC

Evidence from lipid biomarker and fossil 16S rDNA. Geochim. Cosmochim. Acta 72, 1824–1836.

NU

Vinçon-Laugier, A., Grossi, V., Pacton, M., Escarguel, G., Cravo-Laureau, C., 2016. The alkyl glycerol ether lipid composition of heterotrophic sulfate reducing

MA

bacteria strongly depends on growth substrate. Org. Geochem. 98, 141–154. Wegener, G., Krukenberg, V., Riedel, D., Tegetmeyer, H.E., Boetius, A., 2015.

D

Intercellular wiring enables electron transfer between methanotrophic archaea and

PT E

bacteria. Nature 526, 587–590.

Wegener, G., Niemann, H., Elvert, M., Hinrichs, K.-U., Boetius, A., 2008. Assimilation

CE

of methane and inorganic carbon by microbial communities mediating the anaerobic oxidation of methane. Environ. Microbiol. 10, 2287–2298.

AC

Zhang, C., Pancost, R.D., Sassen, R., Qian, Y., Macko, S.A., 2003. Archaeal lipid biomarkers and isotopic evidence of anaerobic methane oxidation associated with gas hydrates in the Gulf of Mexico. Org. Geochem. 34, 827–836.

34

ACCEPTED MANUSCRIPT Figure captions Fig. 1. Bathymetry map showing the two seep sites from which samples were analyzed in this study (see Table 1 for details). Fig. 2. Authigenic carbonates are abundant in well-developed mussel beds (a) and tubeworm bushes (b) at site AT340. Images are ~ 1-2 m across.

PT

Fig. 3. Plot of δ13C and δ18O values of seep carbonates of mussel (AT340-M and

SC

Roberts et al. (2010b) and Feng et al. (2013).

RI

GC852-M) and tubeworm (AT340-T and GC852-T) carbonates. Data are from

Fig. 4. Relative proportions of archaeal biomarkers in mussel (AT340-M and GC852-

NU

M) and tubeworm (AT340-T and GC852-T) carbonates. (a) Isoprenoidal glycerol diethers, normalized to archaeol and (b) glycerol dialkyl glycerol tetraethers

MA

(GDGTs; in % of total GDGTs).

Fig. 5. Contents of non-isoprenoid dialkyl glycerol diethers (DAGEs) and n-C14, n-C15,

D

n-C16, i-/ai-C15 and n-C17:1-alcohols in mussel (AT340-M and GC852-M) and

PT E

tubeworm (AT340-T and GC852-T) carbonates. ω-cycC17:0: ω-cyclohexyl C17; cycC17:0ω5,6: cyclopropyl C17.

CE

Fig. 6. The isotopic compositions of sulfate-reducing bacteria derived fatty acids, DAGEs, and n-C14, n-C15, n-C16, i-/ai-C15, and n-C17:1-alcohols in mussel (AT340-

AC

M and GC852-M) and tubeworm (AT340-T and GC852-T) carbonates. ω-cycC17:0: ω-cyclohexyl C17; cycC17:0ω5,6: cyclopropyl C17. Fig. S1. Partial gas chromatograms (FID) of hydrocarbon fractions from tubeworm (AT340-T, (a)) and mussel (AT340-M, (b)) carbonates; Istd: internal standard; Cr: crocetane; PMI: pentamethylicosane; Pr: pristane. Gray dots: n-alkanes. Fig. S2. Partial gas chromatograms (FID) of carboxylic acid fractions from tubeworm (AT340-T, (a)) and mussel (AT340-M, (b)) carbonates; Istd: internal standard; Ph: 35

ACCEPTED MANUSCRIPT phytanic acid; ββ-32-acid: 17β(H),21β(H)-32-hopanoic acid. White dots: n-fatty acids. Fig. S3. Partial gas chromatogram (FID) of the alcohol fraction of tubeworm carbonate GC852-T, showing the distributions of DAGEs. Please refer to the Appendix for

RI

PT

DAGE structures from Series 1 (1 a-f) and Series 2 (2 a-d); THM: tetrahymanol.

SC

Table captions

Table 1. Cruise information, geographical coordinates, and water depth of the seep

NU

sites GC852 and AT340 of the Gulf of Mexico.

Table 2. Carbonate contents, mineral compositions, and carbon and oxygen stable

T and GC852-T) carbonates.

MA

isotope compositions of mussel (AT340-M and GC852-M) and tubeworm (AT340-

D

Table 3. Contents and stable carbon isotopic composition of archaeal and oil-seepage

PT E

derived biomarkers in mussel (AT340-M and GC852-M) and tubeworm (AT340-T and GC852-T) carbonates.

CE

Table 4. Contents and stable carbon isotopic composition of bacterial biomarkers in mussel (AT340-M and GC852-M) and tubeworm (AT340-T and GC852-T)

AC

carbonates.

Table S1. The distribution of GDGTs and δ13C values of ether-cleaved biphytanes. Table S2. Contents and stable carbon isotopic composition of hopanoids in mussel (AT340-M and GC852-M) and tubeworm (AT340-T and GC852-T) carbonates.

36

ACCEPTED MANUSCRIPT Table 1

Year-Dive number-Sample ID

Lat. Mean

Lon. Mean

Maximum water depth (m)

2007-Jason II 270-AT340-T

N27.6453° W88.3640° 2216

2007-Jason II 270-AT340-M N27.1063° W91.1661° 1450

PT

2007-Jason II 273-GC852-T

AC

CE

PT E

D

MA

NU

SC

RI

2007-Jason II 273-GC852-M

37

ACCEPTED MANUSCRIPT

Table 2 ANME Sample ID

Mineral composition a

Carb. Cont. (%)

Isotopic composition a

Qtz

LMC

mol%

HMC

mol%

Arag.

Protodol./

(%)

(%)

Mg

(%)

Mg

(%)

dolo. (%)

54

13

39

7

I R

ANMEAT340-T

1/ANME-

86

2 AT340-M

GC852-T

GC852-M

ANME-2

ANME-2

ANME-1

90

2

3

78

7

4

77

7

T P

mol%

SC

Mg

30

U N

1

T P E

D E 98

A M 11

‒60.8 to ‒51.8 ‒58.4 to

97

92

δ13C (‰)

‒46.8

4

50

2

50

‒53.7 to ‒44.2 ‒42.9 to ‒36.4

δ18O (‰)

Comments

3.3 to 5.1

Matrix

3.6 to 4.6

Matrix

3.1 to 4.8

Matrix

3.6 to 3.8

Matrix

Carb. Cont., Carbonate content; Qtz, Quartz; HMC, High-Mg Calcite; LMC, Low-Mg Calcite; Arag, Aragonite; Protodol/Dolo,

C C

Protodolomite/Dolomite. a

A

Data were originated from Roberts et al. (2010b).

38

ACCEPTED MANUSCRIPT

Table 3 Sample ID

AT340-T

AT340-M

GC852-T

Content (μg/g

δ13C (‰)

Content (μg/g

δ13C (‰)

Content (μg/g

rock)

V-PDB

rock)

V-PDB

rock)

Crocetanea

0.41

−106

0.38

−99

PMI

0.40

−131

0.11

−119

Phytanoic acid

0.08

n.d.

0.06

Phytanol

0.08

−132

0.11

0.09

−131

δ13C (‰)

V-PDB

rock)

V-PDB

0.76

−90

0.14

−50

0.43

−127

0.03

−103

n.d.

0.08

n.d.

n.d.

n.d.

−127

0.11

−129

0.01

−111

0.07

−131

0.06

−126

0.02

−104

<0.01

n.d.

<0.01

n.d.

<0.01

n.d.

<0.01

n.d.

glycerolether

C C

A

D E

T P E

sn2-O-phytanyl-

sn3-O-phytanyl-

T P

Content (μg/g

N A

M

δ13C (‰)

I R

C S U

Archaeal lipids

glycerolether

GC852-M

Archaeol

1.01

−130

1.42

−127

2.26

−128

0.08

−122

sn2-hydroxyarchaeol

0.81

−137

1.72

−133

2.80

−135

0.01

n.d.

39

ACCEPTED MANUSCRIPT

di-hydroxyarchaeol

n.d.

n.d.

Average δ13C value* Sum AOM

0.06

−128

−134

−127

0.37

−124

−123

2.88

3.93

6.87

0.8

1.2

1.2

C S U

/archaeol

Pristane

0.14

‒28

0.15

n-C16

0.21

‒32

0.2

n-C17

0.23

‒33

n-C18

0.14

N A

Sum

0.72

n.d., not detected. a

C A

M

n.d. −110

0.29

0.2

‒30

0.26

‒30

0.13

‒29

‒30

0.18

n.d.

0.16

‒30

0.21

‒30

0.20

‒32

0.17

‒29

0.15

‒30

0.17

n.d.

0.13

‒30

D E

PT

E C ‒30

T P

I R

sn2-hydroxyarchaeol

Oil-derived lipids

n.d.

0.71

0.81

coelution with phytane.

* Crocetane/phytane is excluded.

40

0.59

ACCEPTED MANUSCRIPT

Table 4

Sample ID

AT340-T

δ13C (‰)

(μg/g rock)

V-PDB

(μg/g rock)

V-PDB

Content

δ13C (‰)

(μg/g rock)

V-PDB

(μg/g rock)

V-PDB

−79

0.27

−110

<0.01

n.d.

−77

0.21

−107

<0.01

n.d.

n.d.

0.02

n.d.

<0.01

n.d.

0.04

−87

0.08

−111

<0.01

n.d.

n.d.

n.d.

n.d.

0.02

n.d.

n.d.

n.d.

n.d.

n.d.

n.d.

0.01

n.d.

n.d.

n.d.

n.d.

n.d.

n.d.

0.02

n.d.

n.d.

n.d.

0.33

−111

0.11

anteiso-C15-fatty acid

0.29

−109

0.08

iso-C17-fatty acid

0.04

n.d.

anteiso-C17-fatty acid

0.11

−120

n-C16:1ω5-fatty acid

0.03

n-C17:1ω6-fatty acid

C A

Average δ13C value

Content

I R

GC852-M

δ13C (‰)

iso-C15-fatty acid

anteiso-/iso-C15:0 ratio

δ13C (‰)

GC852-T

Content

0.01 0.02

PT

D E

E C

0.9

0.02

Content

SC

U N

Bacterial biomarkers (SRB, DSS cluster)

cycC17:0ω5,6-fatty acid

T P

AT340-M

A M

0.7

0.8

−113

−81

41

1.2 −109

ACCEPTED MANUSCRIPT

Sum SRB

0.83

0.25

0.63

<0.05

T P

Bacterial biomarkers (non-DSS cluster SRB)

I R

Monoalkyl glycerol ethers(MAGEs) Sn-2 MAGE cycC17:0ω5,6

−126

0.03

n.d.

C S U

n.d.

Dialkyl glycerol ethers(DAGEs)

N A

Series 1 DAGEs DAGE-C29:0(i-C14:0/ai-C15:0)a

0.07

−122

DAGE-C29:0(n-C14:0/i-C15:0)a

0.03

−112

DAGE-C29:0(n-C14:0/ai-C15:0)a

0.05

n.d.

DAGE-C30:0(i-C15:0/i-C15:0)a

0.05

DAGE-C30:0(i-C15:0/ai-C15:0)a DAGE-C30:0(ai-C15:0/ai-C15:0)a

M

0.04

−117

<0.01

n.d.

n.d.

0.18

−119

n.d.

n.d.

n.d.

0.16

−112

n.d.

n.d.

n.d.

n.d.

0.21

−115

n.d.

n.d.

−117

0.05

n.d.

0.07

−105

<0.01

n.d.

C A

−131

0.05

−125

0.09

−126

<0.01

−131

0.35

−123

0.21

−122

0.52

−119

0.02

−111

0.89

−120

n.d.

n.d.

2.38

−115

0.03

n.d.

0.12

0.03

PT

D E

E C

n.d.

Series 2 DAGEs DAGE-C31:0(n-

42

ACCEPTED MANUSCRIPT

C14:0/cycC17:0ω5,6)a DAGE-C32:0(n0.29

−122

n.d.

n.d.

1.14

0.54

−123

n.d.

n.d.

1.54

0.37

−115

n.d.

n.d.

C15:0/cycC17:0ω5,6) +C32:1 DAGE-C33:0(n-

DAGE-C34:0(ω-

N A

cycC17:0/cycC17:0ω5,6)a Average δ13C value of DAGEs Sum DAGEs

−120

D E

2.76

0.34

M

PT

Δδ13CDAGEs-Fa

−7‰

I R

C S U

C16:0/cycC17:0ω5,6)a+C33:1

1.06

−124

−117

0.01

n.d.

−112

0.01

n.d.

−111

0.02

n.d.

T P

a

−115 7.35

−43‰

−121 0.10

−6‰

n.d.

E C

0.32

C A

−56

0.23

−36

0.15

−69

0.06

n.d.

n-C15-fatty acid

0.12

−46

0.07

−34

0.07

n.d.

0.02

n.d.

n-C16-fatty acid

2.36

−34

2.78

−30

2.76

−33

1.67

−29

Other bacterial biomarkers n-C14-fatty acid

43

ACCEPTED MANUSCRIPT

n-C18-fatty acid

0.32

−30

0.43

−35

0.35

−32

0.15

−29

n-C14-alcohol

0.10

−96

0.07

−38

0.11

−108

0.02

−48

iso-C15-alcohol

0.02

−118

0.02

−102

0.02

−115

0.01

n.d.

anteiso-C15-alcohol

0.06

−119

0.04

−116

0.05

−120

0.02

−32

n-C15-alcohol

0.03

−97

0.03

−40

−114

0.02

−63

n-C16-alcohol

0.08

−93

0.11

−62

0.11

−108

0.03

−48

n-C17:1-alcohol

0.15

−116

n.d.

0.26

−115

0.03

−101

n-C18-alcohol

0.04

−33

0.07

0.03

−45

0.06

−28

Sum

3.60

n.d.

M

T P E

n.d., not detected. a

D E 3.85

N A

C S U

−23

I R

0.05

3.96

T P

2.09

From Pancost et al. (2001b). ω-cycC17:0, ω-cyclohexyl C17; cycC17:0ω5,6, cyclopropyl C17. C32:0/C32:1 and C33:0/C33:1 were in co-elution,

C C

determined by their mass spectra on GC–MS.

A

44

ACCEPTED MANUSCRIPT

AC

CE

PT E

D

MA

NU

SC

RI

PT

Appendix

45

Figure 1

Figure 2

Figure 3

Figure 4

Figure 5

Figure 6

Figure 7

Figure 8

Figure 9