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Autotrophy as a predominant mode of carbon fixation in anaerobic methane-oxidizing microbial communities
Associated Data
Abstract
The methane-rich, hydrothermally heated sediments of the Guaymas Basin are inhabited by thermophilic microorganisms, including anaerobic methane-oxidizing archaea (mainly ANME-1) and sulfate-reducing bacteria (e.g., HotSeep-1 cluster). We studied the microbial carbon flow in ANME-1/ HotSeep-1 enrichments in stable-isotope–probing experiments with and without methane. The relative incorporation of 13C from either dissolved inorganic carbon or methane into lipids revealed that methane-oxidizing archaea assimilated primarily inorganic carbon. This assimilation is strongly accelerated in the presence of methane. Experiments with simultaneous amendments of both 13C-labeled dissolved inorganic carbon and deuterated water provided further insights into production rates of individual lipids derived from members of the methane-oxidizing community as well as their carbon sources used for lipid biosynthesis. In the presence of methane, all prominent lipids carried a dual isotopic signal indicative of their origin from primarily autotrophic microbes. In the absence of methane, archaeal lipid production ceased and bacterial lipid production dropped by 90%; the lipids produced by the residual fraction of the metabolically active bacterial community predominantly carried a heterotrophic signal. Collectively our results strongly suggest that the studied ANME-1 archaea oxidize methane but assimilate inorganic carbon and should thus be classified as methane-oxidizing chemoorganoautotrophs.
Methane is an important greenhouse gas and the most abundant hydrocarbon in marine sediments. Its upward flux to the sediment–water interface is strongly reduced by sulfate-dependent anaerobic oxidation of methane (AOM) (1, 2). AOM is performed by syntrophic associations of anaerobic methane-oxidizing archaea (ANMEs) (3, 4) and their sulfate-reducing bacterial partners (SRBs) (mainly relatives of Desulfosarcina or Desulfobulbus) (5–8). The free energy yield of the AOM net reaction is one of the lowest known for catabolic reactions under environmental conditions (ΔG ranges from –20 to –40 kJ⋅mol–1; e.g., refs. 9, 10). Consequently, activity and biomass doubling times determined under optimized laboratory conditions range from 2 to 5 mo (11) and growth yields are extremely low, around 1% relative to oxidized methane (11, 12). The biomass of ANMEs and SRBs involved in AOM is usually strongly depleted in 13C. For instance, at methane seep locations, the δ13C values of specific bacterial fatty acids and archaeal ether lipids range from –60 to –100‰ and –70 to –130‰, respectively (e.g., refs. 13–17). Such low values have been interpreted as evidence for the incorporation of 13C-depleted methane into biomass (e.g., refs. 3, 4, 18).
One way to identify the carbon sources of microbial biomass is to perform stable–isotope–probing (SIP) experiments (19), followed by analysis of biomolecules such as membrane lipids (lipid-SIP hereafter). Application of lipid-SIP to cold seep sediments and microbial mats indicated that archaeal communities dominated by ANME-2 use both methane-derived carbon and inorganic carbon for lipid biosynthesis, whereas their bacterial partners assimilate only inorganic carbon to produce biomass and are thus complete autotrophs (20, 21). By contrast, Alperin and Hoehler (22) argued on the basis of isotopic considerations that autotrophic methanogens could equally be responsible for the observations of strongly 13C-depleted lipids and biomass at numerous cold seep sites.
Recently, dual SIP with simultaneous addition of deuterated water (D2O) and 13C-labeled inorganic carbon (13CDIC) was introduced as an assay for the quantification of rates of de novo lipid synthesis and inorganic carbon assimilation into lipids (23). Furthermore, via the relationship of incorporation of 13CDIC relative to D in microbial lipids, lipid biosynthesis by autotrophs and heterotrophs can be distinguished. To constrain mechanisms and patterns of carbon flow in microbial communities mediating AOM we performed a series of SIP experiments with combined 13CDIC and D2O as well as 13CH4, using enrichments of AOM-mediating microbial communities from the Guaymas Basin (cf. ref. 24). This approach enabled us to differentiate which lipids were produced in the presence and absence of methane and to quantify the roles of methane and inorganic carbon as carbon sources for microbes involved in AOM.
Results and Discussion
Activity and Microbial Composition of the Guaymas Basin Enrichments.
We incubated replicates of hot seep sediments naturally enriched in moderately thermophilic ANME-1 dominated communities at suitable growth conditions (200 kPa CH4, 37 °C, and artificial seawater medium for sulfate reducers) (25). Samples were amended either with 13CDIC [resulting 13C fraction was 9.6% of dissolved inorganic carbon (DIC)] and deuterated water (3% deuterium in total water) or with 13CH4 (15.9% 13C of methane carbon) in a time series of 10, 17, and 24 d, with and without methane headspace and in killed controls (Table 1). The incubations with methane showed a strong increase of sulfide production in response to stimulation of sulfate reduction. Incubations with 13CH4 revealed methane oxidation rates (SI Text, Eq. S1) that were roughly 85% (3.4 μmol·d–1·gdm–1) of the sulfate reduction rate (Table 1). In the absence of methane, sulfate reduction decreased by ∼90% (Table 1). Sequencing of 16S rRNA and fluorescence in situ hybridization revealed a dominance of aggregate-forming ANME-1 archaea and partner bacteria from the HotSeep-1 cluster (Fig. S1 and Table S1).
Table 1.
Overview of conditions in incubation experiments
| Incubation time | Substrate combination | δ13CDIC, ‰ | δDH2O, ‰ | H2S production, µmol⋅d–1⋅gdm–1 |
| Control | ||||
| t0, no label | CH4 | NA | NA | NA |
| t0, + CH4 | CH4; D2O; H13CO3– | NA | 244,000 | NA |
| t0, + 13CH4 | 13CH4 (∼16,000‰) | NA | NA | NA |
| t24, killed | CH4; D2O; H13CO3–; ZnCl2 | 8,500 | 199,000 | 0.4 |
| + CH4 | ||||
| t10, + CH4 | 8,400 | 196,000 | ||
| t17, + CH4 | CH4; D2O; H13CO3– | 8,200 | 206,000 | 6.0 ± 0.4 |
| t24, + CH4 | 8,000 | 206,000 | ||
| t24, + 13CH4 | 13CH4 | 950 | NA | 4.0 |
| w/o CH4 | ||||
| t24, w/o CH4 | D2O; H13CO3– | 8,700 | 210,000 | 0.7 |
Duration of the incubation, substrate combinations (DH2O and 13CDIC; 13CCH4), labeling strength (δ13CDIC and δ13CCH4 of the headspace; δDH2O of the aqueous media), and the sulfide production rate of all analyzed samples incubated at 37 °C are shown. CO2 = 50 kPa; CH4 = 200 kPa; N2 = 200 kPa. + CH4, presence of methane headspace; killed, killed control (ZnCl2, final concentration 2% wt/vol); NA, not analyzed; w/o, without.
Microbial Lipid Distribution and Natural Isotopic Compositions.
Concentrations and relative abundances of lipids did not change significantly during the incubations, an observation that is consistent with low growth rates reported for this microbial community (doubling time of 77 d at 37 °C) (24). Ether cleavage of archaeal lipids in the total lipid extract released mainly acyclic biphytane, phytane, and bicyclic biphytane (Fig. S2 A and C and Table S2). High relative amounts of biphytanes are derived from glycerol dibiphytanyl glycerol tetraethers (GDGTs), which have been routinely found in methane-rich sediments and cold seeps dominated by ANME-1 (26–28), whereas phytanes derived from the cleavage of archaeols are less specific and likely produced by all ANMEs (e.g., refs. 16, 17, 26). However, because archaeal 16S rRNA gene libraries and microscopic analysis of this enrichment revealed dominance of ANME-1 archaea, this specific group is most likely the main source of both phytanes and biphytanes.
Fatty acids (FAs) are composed of saturated even-numbered, saturated, and unsaturated terminally or subterminally branched FAs (iso, i; and anteiso, ai); monounsaturated C16 and C18 FAs; and isoprenoidal FAs (Fig. S2 B and C and Table S2). This distribution differs significantly from those found at cold seeps (14, 17, 26), which is consistent with the distinct bacterial members in our enrichment. The unusual methyl branched and unsaturated FAs iC16:1ω6 (Fig. S3) and iC18:1ω6 have to our knowledge not been detected at cold seeps, but are known constituents of some sulfate-reducing bacteria (mostly in Desulfovibrio species) (29, 30). Phylogenetic as well as catalyzed reporter deposition fluorescence in situ hybridization (CARD-FISH) analysis of the enrichment indicated that the dominant sulfate reducers belong to the HotSeep-1 cluster (Table S1 and Fig. S1), which has been repeatedly found at Guaymas Basin but nowhere else yet.
Both, archaeal isoprenoids and bacterial FAs displayed relatively low natural δ13C values indicative of AOM activity (13–17); phytanes and biphytanes ranged from –18 to –47‰ and bacterial FAs from –21 to –50‰, whereas δD values ranged from –147 to –286‰ for archaeal isoprenoids and from –86 to –232‰ for FAs (Table S2). This large range is similar to earlier observations of δD values in anoxic marine sediments (31).
Assimilation of Stable Isotope Label into Microbial Lipids.
Although changes in lipid concentrations were not detectable, the incorporation of D and 13C from labeled substrates indicated freshly synthesized lipids (Fig. 1 A and B, respectively, and Table S3). The incorporation of both heavy isotopes in microbial lipids relative to time zero (Δδ) was most pronounced for incubations with methane, and Δδ increased fairly linearly with time (Fig. 1 A and B). The extent of label uptake differed between individual compounds, with the highest change in ΔδD and Δδ13C found for archaeal phytane and bacterial lipids such as iC16:1ω6, iC18:1ω6, and C18:1ω7 FAs (Table S3). For instance, in the 24-d incubation with methane, changes in phytane’s δ-values were remarkably higher than in biphytanes (Fig. 1 A and B and Table S3). The ΔδD of bacterial C14–C18 FAs was high (∼ +12,000‰) and similar to that of phytane, whereas Δδ13C was significantly lower in the FAs (∼ +380‰; Fig. 1 A and B and Table S3). In the killed control, no enrichment of D and 13C in lipids was detected (Table S2). In incubations without methane, archaeal lipids showed only traces of D-label uptake but none of 13C; likewise the uptake into bacterial lipids was strongly reduced compared with the incubations with methane (Fig. 1 A and B and Table S3). This observation confirms the central role of methane as an energy source in the examined AOM system.
Results of lipid stable-isotope probing. (A and B) Isotopic shifts of (A) hydrogen and (B) carbon, expressed as weighted-average Δδ of total fatty acids (FA, light gray bars), archaeols as phytanes (Phy, dark gray bars), and glycerol dibiphytanyl glycerol tetraethers (GDGTs) as biphytanes (BPs, black bars) during the experiment. (C) Production of lipids (prodlipid, green bars) and assimilation of inorganic carbon (assimIC, blue bars) into bacterial (Left) and archaeal (Right) lipids. Time series experiments in C are presented as average values (n = 3, error bar = SD). The ratio of assimIC to prodlipid (Ra/p) gives a measure for the dominance of a heterotrophic (Ra/p < 0.3) or autotrophic (Ra/p ~ 1) mode of carbon fixation. In the absence of methane, Ra/p values are not presented because of low values detected for archaeal inorganic carbon lipid assimilation. The symbol “+” indicates incubations with 13CDIC ∼ 9.6% 13C, D ∼ 3%, and unlabeled methane. Incubations with13CH4 contained ∼15.9% 13C.
Incubation with 13CH4 (24 d) resulted in relatively low 13C assimilation into lipids. The Δδ13C values of around 30‰ for both bacterial and archaeal lipids (Table S3) corresponded to about 1/12th and 1/4th of the Δδ13C values for the respective lipid pools in the parallel experiment with 13CDIC but unlabeled methane. We assign this slight positive shift in δ13C values to indirect uptake via assimilation of inorganic carbon from the 13C-enriched DIC pool resulting from 13CH4 oxidation (after 24 d δ13CDIC ∼ +950‰; Table 1). This indicates that not only SRBs, but also ANME-1 archaea assimilate predominantly inorganic carbon. Our findings are in agreement with the result from Treude and coworkers (12), who detected a predominance of 14CO2 over 14CH4 incorporation within a microbial mat known to be dominated by ANME-1/SRB consortia. However, this result differs from a previous labeling study with cold seep sediments by Wegener et al. (21), in which methane contributed up to 50% during archaeal lipid carbon fixation for mixed ANME-1/ANME-2 and ANME-2–dominated communities.
Lipid Production, Inorganic Carbon Assimilation, and Methane-Dependent Carbon Fixation.
We converted the incorporation of D and 13CDIC in microbial lipids into rates of lipid production (prodlipid) and inorganic carbon assimilation (assimIC), respectively. The ratio of assimIC to prodlipid (Ra/p) indicates the dominant mode of microbial carbon fixation, with Ra/p values ≤ 0.3 indicative of heterotrophic metabolism and Ra/p values of close to 1 signaling autotrophic metabolism (cf. ref. 23) (Materials and Methods). Intermediate Ra/p values are interpreted to represent a mixture of auto- and heterotrophically produced lipids, e.g., via production of a single compound by multiple sources (23).
In the presence of methane as an energy source the values of archaeal prodlipid reached 10 ± 3 µg lipid⋅gdm–1⋅y–1, indicating archaeal cell growth (Fig. 1C and Table S4). The assimIC value was nearly identical and thus indicated strictly autotrophic carbon fixation by the archaeal community (Ra/p values of 1.03 ± 0.05; Fig. 1C). In the absence of methane, the values of archaeal prodlipid and assimIC were marginal (Table S4), demonstrating that the presence of methane is mandatory to sustain lipid production and thus the archaeal community.
In all incubations with methane, prodlipid of phytane exceeded that of biphytanes by at least one order of magnitude (Fig. 2A, Fig. S4, and Table S4), whereas Ra/p values of around 1 for both phytane and biphytanes indicated a strictly autotrophic carbon metabolism of these lipids’ microbial producers. Detection of assimIC in biphytane was hindered by the large background of fossil core-GDGTs in the total lipid extract (Fig. S5), which reduces the SIP sensitivity and prevents the use of biphytane-derived Ra/p values as a gauge for distinguishing hetero- or autotrophic producers of lipids (Table S4). To increase the sensitivity of the SIP experiment, we performed a similar isotopic assay on biphytanes from intact polar GDGTs purified from the total lipid extract by preparative liquid chromatography (cf. ref. 32) and obtained an Ra/p value of 1.2 in support of autotrophic production (254 and 217 ng lipid⋅gdm–1⋅y–1 for assimIC and prodlipid, respectively).
Carbon assimilation and lipid production of microbial lipids. Compared are carbon assimilation (assimIC) and production (prodlipid) of bacterial fatty acids (diamonds) and archaeal ether lipid-derived isoprenoids (circles) in the (A) presence and (B) absence of methane after 24 d of incubation (see Fig. S4 for more details).
As observed for archaeal lipids, the prodlipid and assimIC values of bulk bacterial FAs were highest in the presence of methane (24 ± 2 and 16 ± 1 µg lipid⋅gdm–1⋅y–1, respectively; Fig. 1C and Table S4), resulting in an Ra/p value of 0.70 ± 0.01. Without methane, both prodlipid and assimIC of bulk bacterial FAs were strongly reduced; the resulting Ra/p value of 0.2 signaled predominant production by heterotrophic microbes (Fig. 1C and Table S4). This contrast emphasizes the importance of bacterial CO2 assimilation in connection with methane oxidation.
The bacterial lipids with the highest production in the presence of methane were iC16:1ω6, iC16:0, C16:0, iC18:1ω6, and C18:1ω7. Most of these lipids, including iC16:1ω6 and iC18:1ω6, which are both specific to HotSeep-1 phylotype, and C18:1ω7, show comparable prodlipid and assimIC values that are indicative of predominant production by autotrophs (Figs. 2A and and3,3, Fig. S4, and Table S4). The production intensity of these lipids does not correspond to the distribution of FA concentrations (Fig. S5). This indicates that the lipid distribution is a cumulative record affected by the temporally varying rates of production and degradation of individual lipids, whereas the prodlipid records the metabolic activity of those community members stimulated by the culture conditions. In the absence of methane, production of all lipids decreased, and Ra/p values dropped toward heterotrophic signatures. This decline was most pronounced in lipids with autotrophic signatures in the presence of methane (e.g., C18:1ω7, iC16:1ω6, and iC18:1ω6; Figs. 2B and and3,3, Fig. S4, and Table S4). We cannot resolve to what degree the drastic change in both production and carbon metabolism expressed in dual isotopic signatures of individual lipids is due to metabolic adaptation or simply to the shutoff of SRBs affiliated with AOM. The permanently low Ra/p values observed for some i- and ai-FAs, specifically iC15:0 and iC17:0 (Fig. 3 and Table S4), are best explained by other, less active, heterotrophic bacterial community members not involved in AOM, such as relatives of the genus Anaerolinea within subphylum I of Chloroflexi, which are abundant in the 16S rRNA gene library from this Guaymas Basin enrichment (Table S1). Fermenters usually show low heterotrophic CO2 fixation (33).
Relative increase in individual lipid production in incubations with methane compared with incubations without methane. Increases are plotted against the ratio of carbon assimilation to lipid production (Ra/p) as an indicator for auto- or heterotrophic source organisms of the lipid. Archaeal ether lipid-derived hydrocarbons (circles) and bacterial fatty acids (diamonds) were produced in either the presence (red symbols) or the absence (white symbols) of methane. Ra/p values in the absence of methane were not determined for iC16:1ω6, C16:1ω7, and iC18:1ω6 and for BP(0) and BP(1) due to assimIC values close to the detection limit. Shaded bars indicate ranges primarily interpreted to reflect heterotrophy (brown) or autotrophy (blue).
We demonstrated that both archaea and bacteria mediating AOM in hydrothermal sediments in the Guaymas Basin assimilate preferentially inorganic carbon. This was unexpected for the archaeal community members. Our results indicate that the archaea do not assimilate methane but instead act as autotrophic methane oxidizers. The carbon assimilation of many Euryarchaeota, in particular methanogens, is performed via the reductive acetyl-CoA pathway (34). This pathway combines two one-carbon-atom moieties of the oxidation state of +II (equivalent of carbon monoxide) and –III (methyl group) to form acetyl-CoA. It was speculated that ANME archaea shuttle methyl groups from the acetyl-CoA pathway into their assimilatory system (21, 35). We have shown that in ANME-1 archaea from the Guaymas Basin growing at 37 °C, methane oxidation is decoupled from the assimilatory system. Hence these ANME-1 archaea qualify as chemoorganoautotrophs. The lacking transfer of methane carbon into the assimilatory system may appear counterintuitive. However, autotrophic carbon fixation during growth on organic substrates has been previously demonstrated (36, 37). Particularly under the strongly negative redox potentials in sulfidic environments, microorganisms are able to fix inorganic carbon with relatively low input of energy, using the acetyl-CoA pathway (38). This is especially true for methane-oxidizing archaea, which demand a sink for the four electron pairs per molecule methane oxidized. Our findings strongly suggest that in ANME-1, as in many methanogens, lipid biosynthesis is linked to autotrophic carbon fixation. Because methane utilization appears to be limited to the dissimilatory pathway, the term methanotrophy should be used with caution, as “trophy” involves the assimilation of a compound.
The unexpected chemoorganoautotrophic nature of ANME-1 demands a reevaluation of isotopic signatures of archaeal lipid biomarkers in marine sediments with active AOM communities. Strongly 13C-depleted archaeal lipids are established as important proxies for tracking the activity of methane-oxidizing archaea in such settings (e.g., refs. 3, 13–18); this isotopic depletion has been attributed to the direct transfer of 13C-depleted methane carbon into lipid biosynthesis. As suggested recently (22), the 13C-depleted lipids could equally be products of autotrophic methanogens that use 13C-depleted CO2. Although lipid products of CO2-assimilating ANME archaea could probably not be distinguished easily from those of such putative methanogens on the basis of their isotopic composition. However, our study identified close relatives of ubiquitous cold-seep archaea, in which CO2 assimilation is strictly coupled to methane oxidation rather than methanogenesis. The much larger range of isotopic compositions in putative lipids of methane-oxidizing archaea compared with methane at cold seeps (39) could be the result of a large variation of δ-values of CO2 in conjunction with a more widespread distribution of ANME archaea that predominantly assimilate CO2.
Materials and Methods
Sample Collection, Genetic Analyses, and Stable Isotope-Labeling Experiments.
Sediments derived from the gas-rich, Beggiatoa-covered hydrothermal site in the Guaymas Basin (27°00.437 N, 111°24.548 W) were retrieved during the R/V Atlantis cruise AT 15–56 (Alvin dive 4,570, 2009). Samples were diluted 1:1 with anoxic artificial seawater medium supplemented with trace elements and vitamins for sulfate-reducing bacteria (25) and incubated at 37 °C with a methane headspace at 100 kPa until further processing (24). After 90 d of preincubation, DNA was extracted from a replicate sediment batch incubated following established protocols. rRNA (16S) was amplified, cloned, and sequenced. The phylogenetic affiliation was inferred with the ARB software package (SI Text). For the SIP experiments, enrichments were equally distributed into 256-mL culture vials [∼4 g dry mass (gdm) per vial], filled up with seawater medium as described above. Samples were amended with labeled substrates (D2O, 13CDIC, 13CH4) in different combinations and incubated at 37 °C for 0, 10, 17, and 24 d (t0, t10, t17, t24) with either CH4 or N2 atmosphere at 200 kPa + 50 kPa CO2. The pH in all incubations was 7.2. Samples amended with 3% D2O and 9.6% 13CDIC had δDH2O values of ∼ +200,000‰ vs. Vienna Standard Mean Ocean Water (VSMOW) and δ13CDIC values of ∼ +8,400‰ vs. Vienna PeeDee Belemnite (VPDB) (VPDB = 9.6% 13C). The carbon isotopic composition of the methane headspace in the 13CH4 (15.9% 13C) experiments had δ13C values of ∼ +16,000‰ vs. VPDB (detailed information in Table 1). Killed controls were sterilized with ZnCl2 [2% (wt/vol) final concentration] before label addition. Sulfide production was constantly monitored using a spectrophotometric method described by Cord-Ruwisch (40). For cell hybridization, a sediment aliquot was fixed (in formaldehyde; 30 g·L–1, final concentration, at room temperature) and blotted on a membrane filter and CARD-FISH was performed with established probes for ANME-1 and ANME-2. Subsequently samples were stained with DAPI and cells and/or microbial phyla were identified with fluorescence microscopy.
Medium Isotopic Compositions.
Hydrogen isotopic composition of the incubation medium was determined using cavity ring-down laser spectroscopy (Liquid Water Isotope Analyzer DLT-100; Los Gatos Research) after dilution with pure water (1:100). The δ-values of DIC and CH4 were measured from the headspace of acidified medium by gas chromatography coupled to isotope-ratio mass spectrometry (irMS) (VG Optima, Fisons; and Trace GC ultra + DeltaPlus XP isotope-ratio MS, ThermoFinnigan, respectively).
Lipid Extraction and Sample Treatment.
Lipids were extracted from enrichment slurries (∼4 gdm), using a modified Bligh and Dyer protocol (41). Bacterial membrane-derived FAs were retrieved by saponification and analyzed as methyl esters after derivatization with BF3 and MeOH. Double-bond positions of FAs were identified by examining mass spectra of their pyrrolidide derivatives (42) (Fig. S3). Ether-bound archaeal isoprenoids were released using boron tribromide treatment to cleave ethers, followed by reduction of the resulting alkyl bromides with superhydride (Aldrich) (43). The products were purified on a silica gel column. FAs and isoprenoidal hydrocarbons were quantified by gas chromatography coupled to a flame ionization detector (GC-FID) (ThermoFinnigan), using squalane as an injection standard.
The analysis of isoprenoidal hydrocarbons from intact polar lipids was performed after purification by an orthogonal preparative high-performance liquid chromatography method in normal and reversed mode (cf. ref. 32) (SI Text). Subsequently, intact polar tetraether fractions were subjected to ether cleavage as described above.
Sample Analyses: GC-FID, GC-MS, and GC-irMS.
Lipids were quantified by gas chromatography coupled to a FID (Trace GC Ultra; Thermo Scientific). Carbon and hydrogen isotopic composition was determined using GC-irMS at least in duplicate measurements (Trace GC Ultra coupled to a GC-IsoLink/ConFlow IV interface and a Delta V Plus irMS; all from Thermo Scientific). Compounds were oxidized in a combustion reactor at 940 °C, and molecular hydrogen was produced from eluting lipids in a pyrolysis reactor at 1,420 °C. The analytical error was <0.5‰ and <10‰ for nonlabeled δ13C and δD values, respectively. δ13C and δD values were corrected for additional carbon and hydrogen introduced during derivatization.
Calculations of Total Lipid Production Rate (Prodlipid), Assimilation Rate of Inorganic Carbon (AssimIC), and the AssimIC/Prodlipid Ratio (Ra/p).
The stable hydrogen and carbon isotope values are expressed in the δ-notation in per mill (‰) as deviation of the isotope ratio from the reference standards. Prodlipid and assimIC are calculated by multiplying lipid concentration (conclipid) by the increase of the fraction of either D (ΔFDlipid) or 13C (ΔF13Clipid) relative to the fraction of nonlabeled samples, divided by the fraction of D (FDmedium) and 13C (F13Cmedium) in the incubated medium (Eqs. 1 and 2) (23):
The fractions of FD and F13C in individual lipids and incubation medium are calculated from the isotope ratios FD = RD/H/(RD/H + 1); F13C = R13C/12C/(R13C/12C + 1), where R derives from the δ-notations (Eqs. S2 and S3 and SI Text). Both prodlipid and assimIC are expressed in µg lipid⋅gdm–1⋅y–1.
Acknowledgments
We thank the shipboard scientific crew and pilots of the R/V Atlantis and Research Submersible Alvin. We thank Jenny Wendt and Arne Leider for technical support and Jan Hoffmann for the water isotopic composition analysis. This study was supported by the Deutsche Forschungsgemeinschaft (through the Research Center/Excellence Cluster MARUM–Center for Marine Environmental Sciences and the graduate school GLOMAR-Globale Change in the Marine Realm), the European Research Council under the European Union's Seventh Framework Programme–“Ideas” Specific Programme, ERC grant agreement No. 247153 (to K.-U.H.), the Alexander von Humboldt Foundation (to M.Y.Y.), the Gottfried Wilhelm Leibniz Prize awarded to Antje Boetius (to G.W.), and the Max Planck Society (to G.W., T.H. and K.K.). Sample acquisition was supported by Grant NSF OCE-0647633 to Andreas Teske.
Footnotes
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
Data deposition: The nucleotide sequences reported in this paper have been deposited at the European Molecular Biology Laboratory, GenBank, and the DNA Data Base of Japan (DDBJ) [accession nos. FR682479–FR682487 and HE817766–HE817767 (archaeal 16S rRNA genes); and FR682618–FR682643 (bacterial 16S rRNA genes)].
This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1208795109/-/DCSupplemental.
