Lipid Biomarker Patterns Reflect Nutritional Strategies of Seep-Dwelling Bivalves From the South China Sea

Introduction

Chemosymbiotic invertebrates from hydrothermal vents and methane seeps, including mussels and clams, gain nutrition through chemosynthetic bacterial symbionts (Childress et al., 1986; Brooks et al., 1987; Fisher et al., 1987; Duperron et al., 2009, 2014). Such symbionts use either reduced sulfur compounds like hydrogen sulfide or methane as electron donors, with oxygen as electron acceptor, providing the invertebrate hosts with nutrition (Petersen and Dubilier, 2010). The host animal provides a stable environment, electron donors, and electron acceptors for the symbionts (Petersen and Dubilier, 2010).

The first report of mussels living in symbiosis with thiotrophic bacteria was from a hydrothermal vent site (Rau and Hedges, 1979), followed by reports from methane seeps, where mussels were found to harbor abundant intracellular methanotrophic bacteria in their gills (Childress et al., 1986). To date, more than 10 species of chemosymbiotic bathymodiolin mussels and clams have been reported from methane seep habitats and whale falls (Cordes et al., 2009; Faure et al., 2015; Duperron and Gros, 2016; Feng et al., 2018; Lan et al., 2019; Xu et al., 2019). It has been demonstrated that some bathymodiolin mussels host either thiotrophic (Cavanaugh, 1983; Distel et al., 1988) or aerobic methanotrophic bacteria, others harbor both types of symbionts, a phenomenon referred to as dual symbiosis (Childress et al., 1986; Fisher et al., 1993; Fiala-Médioni et al., 2002; Cordes et al., 2009; Xu et al., 2019). Putative chemosymbiotic invertebrates were also common at ancient seeps (Kiel, 2010; Kiel and Peckmann, 2019; Kiel et al., 2021).

The phylogeny of bacterial symbionts varies with chemosymbiotic invertebrate species, habitat characteristics, and locations (Nelson and Fisher, 2000; Thornhill et al., 2008; Lorion et al., 2013; Laming et al., 2015; Szafranski et al., 2015; Duperron and Gros, 2016), but can also vary with symbiont populations in response to environmental change (Kádár et al., 2005; Bettencourt et al., 2008). For example different clades of symbionts were found for Escarpia laminata from Alaminos Canyon and the Florida Escarpment, both Gulf of Mexico (Nelson and Fisher, 2000; Thornhill et al., 2008). Furthermore, gradual loss and re-establishment of symbiosis in the gills of Bathymodiolus azoricus was observed when keeping the host in sulfide-free or sulfide-containing seawater, respectively (Kádár et al., 2005; Bettencourt et al., 2008). Fiala-Médioni et al. (2002) suggested the existence of dual symbiosis, providing the host with greater environmental tolerance and increased niche space. Unfortunately, symbiotic bacteria from hydrothermal vents and hydrocarbon seeps have not been cultured successfully to date (Franke et al., 2021). Symbiotic bacteria and their symbiotic associations between hosts and symbionts are typically studied with electron microscopy, enzymatic analyses, 16S rRNA work, as well as lipid biomarkers and their carbon stable isotope compositions (Duperron et al., 2009; Petersen and Dubilier, 2010; Feng et al., 2015).

The analysis of lipid biomarkers is not only a powerful tool to identify bacterial signatures in symbiotic invertebrates, but also to quantify the abundance of bacterial symbionts. Typically, the most abundant lipids are fatty acids. The fatty acid inventory of aerobic methanotrophic bacteria is characterized by the monounsaturated fatty acid (MUFA) C_16:1ω9, but especially by the group-specific MUFAs C_16:1ω8, and C_18:1ω8. In contrast, thiotrophic symbionts produce the ubiquitous MUFAs C_16:1ω7 and C_18:1ω7 (Jannasch, 1985; Nichols et al., 1985; Jahnke and Nichols, 1986; Bowman et al., 1991; Guezennec and Fiala-Medioni, 1996), which are common constituents of bacteria and eukaryotes, and are also produced or taken up by the host animals. Polyunsaturated fatty acids (PUFAs) n-C₁₈, n-C₂₀, and n-C₂₂ are most likely taken up and stored by the host after heterotrophic consumption (Fullarton et al., 1995; Pond et al., 2000) rather than produced by endosymbiotic bacteria (Dunstan et al., 1993; Kawashima and Ohnishi, 2003). Besides fatty acids, chemosymbiotic mussels contain bacteriohopanepolyols (aminotriol and aminotetrol), and various 4,4-dimethyl sterols and 4-methyl sterols, exclusively derived from symbiotic aerobic methanotrophic bacteria (Fang et al., 1993; Jahnke et al., 1995; Pond et al., 1998; Riou et al., 2010; Kellermann et al., 2012).

Apart from the lipids of endosymbiotic bacteria, lipid biomarkers and compound-specific carbon stable isotope compositions of free-living aerobic methanotrophic bacteria have been studied intensively (Summons et al., 1994; Jahnke et al., 1999; Schouten et al., 2000; Talbot et al., 2001; Cordova-Gonzalez et al., 2020). In contrast to the ubiquitous occurrence of Type I and X methanotrophs in diverse natural environments, including the marine realm, type II methanotrophs seem to be confined to terrestrial habitats (Knief, 2015). The identification of the different types of aerobic methanotrophs (Type I, X, and II) is commonly possible with biomarkers, since the different types of methanotrophs contain different biomarkers. For example, a high abundance of the MUFA C_16:1ω8 was found in Type I and X methanotrophs, whereas MUFA C_18:1ω8 tends to be predominant in Type II methanotrophs (Nichols et al., 1985; Bowman et al., 1991). On the other hand, 4-methyl steroids are highly specific biomarkers of Type I methanotrophs (e.g., Bouvier et al., 1976; Schouten et al., 2000). Information from lipid biomarkers can be complemented by their carbon stable isotope patterns, reflecting different carbon assimilation pathways of aerobic methanotrophs. Type-I and -X methanotrophs use the ribulose monophosphate (RuMP) pathway, whereas Type-II methanotrophs use the serine pathway (Anthony, 1982; Summons et al., 1994; Jahnke et al., 1999). The different assimilation pathways lead to distinct carbon isotopic fractionation (Jahnke et al., 1999; Cordova-Gonzalez et al., 2020). For example, Type I methanotrophs produce more highly ¹³C-depleted lipids relative to Type II methanotrophs (Summons et al., 1994; Jahnke et al., 1999; Cordova-Gonzalez et al., 2020). Moreover, the δ¹³C values of hopanoids and steroids synthesized by Type I methanotrophs are as much as 5–7‰ more negative than fatty acids, whereas in Type II methanotrophs, fatty acids are more depleted in ¹³C than hopanoids (Summons et al., 1994; Jahnke et al., 1999). Therefore, lipid biomarkers alongside compound-specific carbon stable isotopes are primed to become part of the standard toolkit to discriminate the different types of symbionts and evaluate the nutritional status and associations between symbionts and hosts in chemosymbiosis.

We studied seep bivalves collected from two active seep sites of the South China Sea (Figures 1, 2). The mussel Gigantidas platifrons was collected at Site F seep (also called Formosa ridge) at 1120 m water depth (Feng et al., 2015). One of the mussels and a clam retrieved from the Haima seep at 1390 m water depth were identified as new species and described as G. haimaensis and A. marissinica (Liang et al., 2017). The other mussel, Bathymodiolus aduloides, was collected at the Haima seep at 1390 m water depth. Transmission electron microscopy and 16S rRNA-encoding gene sequence analyses indicated that G. platifrons and G. haimaensis contain mainly aerobic methanotrophic bacteria as symbionts in their gills (Barry et al., 2002; Xu et al., 2019), whereas B. aduloides and A. marissinica were reported to live in symbiosis with thiotrophic bacteria (Feng et al., 2018; Lan et al., 2019). In this study, lipid biomarkers and compound-specific carbon stable isotope compositions were used to examine carbon assimilation pathways, nutritional status, and different host–symbiont associations in various environments, hosts, and symbionts of the four species of seep-dwelling bivalves from the South China Sea.

FIGURE 1

Figure 1. Map showing the location of the sampling sites. Site F and Haima are active methane seeps (highlighted by red circles); orange circles indicate inactive methane seeps on the northern and western continental slope of the South China Sea.

FIGURE 2

Figure 2. Seafloor images from Site F (a) and Haima (b–d) methane seeps and the four species of bivalves Gigantidas platifrons (a), Gigantidas haimaensis (b), Archivesica marissinica (c), and Bathymodiolus aduloides (d).

Materials and Methods

Sample Collection and Storage

Specimens of G. platifrons were collected in May 2018 using the ROV ROPOS during a cruise with the Research Vessel “Tan Kah Kee” to Site F. Specimens of G. haimaensis, A. marissinica, and B. aduloides were retrieved in April 2018 and May 2019 from the Haima seep using the ROV Haima (Table 1). At the sampling sites, dense G. platifrons, G. haimaensis, and A. marissinica clusters were observed, respectively, covering large areas of the seabed. Several B. aduloides were scattered among carbonate blocks. Bottom water temperatures at both Site F and Haima seep were 3.0 to 3.5°C. After sampling, bivalves were stored onboard at −80°C and shipped to the home laboratory and kept at −25°C. The bivalves were dissected into gill, foot, and remaining tissue, immediately freeze-dried, ground, homogenized and kept at −25°C until further analysis.

TABLE 1

Table 1. Information on the four studied seep bivalves.

Lipid Extraction and Analysis

Lipids were extracted from the gill and foot tissues of G. platifrons, G. haimaensis, A. marissinica, and B. aduloides. The homogenate was extracted with the solvents dichloromethane (3×), methanol:dichloromethane (1:1, v:v) (3×), and methanol (3×) using an ultrasonic bath to obtain the total lipid extracts (TLE). An aliquot of the TLE was saponified after addition of internal standards (19-methylarachidic acid, and tridecyl alcohol) with 6% KOH (w/v) in methanol (2 h at 70°C). Unfortunately, the concentration of the 19-methylarachidic acid standard in the fatty acid fraction was too low in comparison with the target compounds, resulting in a potential error in the quantification of fatty acids. The absolute contents of fatty acids should consequently not be taken at face value. Therefore, the relative portions of carboxylic acids and alcohols of the overall compound inventory of the respective fractions are provided in Tables 2, 3 in addition to absolute contents. The neutral fractions were extracted from the saponified TLE with n-hexane. To obtain carboxylic acids, the residuals were treated with 10% HCl to pH = 2 and extracted using n-hexane. It needs to be stressed that bacteriohopanepolyols (BHPs) are commonly present in aerobic methanotrophic bacteria; however, we decided to focus on fatty acids and sterols and did not analyze BHPs in this study. The neutral fractions were silylated with bis(trimethylsilyl) trifluoroacetamide (BSTFA) for 3 h at 70°C, and the acid fractions were methylated with 14% BF₃-methanol in a screwcap vial (2 h at 60°C). To determine double bond positions of monounsaturated fatty acids, fatty acid methyl esters in n-hexane (50 μl) were treated with dimethyl disulfide (DMDS, 100 μ1) and iodine solution (500 μl; 6% w/v in diethyl ether), filled with argon and stored at 50°C for 48 h. After cooling, iodine was removed by shaking with drops of 5% (w:v) aqueous sodium thiosulfate (Nichols et al., 1986; Guan et al., 2021). The organic phase was removed and the aqueous phase was extracted with n-hexane to obtain the DMDS adducts. All fractions were analyzed with a Thermo Electron Trace 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 used as carrier gas at a flow rate of 1.2 ml/min. Compound-specific carbon stable isotope analysis was performed on a GV Isoprime system interfaced to a Hewlett-Packard 6890 gas chromatograph at the Guangzhou Institute of Geochemistry, Chinese Academy of Sciences (GIG, CAS). The same chromatographic column was used for GC-MS analysis. The following GC temperature program was applied for GC-MS and GC-IRMS: initial temperature was held at 60°C for 2 min, from 60°C to 160°C at a ramp of 10°C/min; from 160 to 320°C at a ramp of 1°C/min, and then held at 320°C for 30 min. The analytes were identified by their retention times and comparison with published mass spectra. The carbon isotopic composition is expressed in the standard δ notation in per mil (‰) relative to Vienna-PeeDee Belemnite (V-PDB) standard. The fatty acid methyl esters (FAMEs) and trimethylsiloxyl (TMS) derivatives were corrected for the addition of carbon during the derivatization. The fatty acids were measured as FAMEs and were corrected for the addition of an extra carbon due to the derivatization according to the following equation:

TABLE 2

Table 2. Contents and isotopic compositions of fatty acids (FAs) from bivalve tissues.

TABLE 3

Table 3. Contents and isotopic compositions of sterols, hopanoid, and n-alcohols from bivalve tissues.

$${\mathrm{\delta }}^{13}{C}_{\text{FA}}=[(C_{n+1}){\mathrm{\delta }}^{13}{C}_{\text{FAME}}-{\mathrm{\delta }}^{13}{C}_{\text{MEOH}}]/C{}_{\text{n}}$$

The δ¹³C_FA indicates the δ¹³C value of the fatty acid, C_n represents the number of carbons in the fatty acid, δ¹³C_FAME is the δ¹³C value of the FAME, and δ¹³C_MEOH is the δ¹³C value of the methanol used for the methylation reaction.

The sterols or n-alcohols were measured as TMS-derivatives and were corrected for the addition of three extra carbons according to the following equation:

$$\mathrm{\delta }{}^{13}C{}_{\text{alcohol}}=[(C{}_{n+3})\mathrm{\delta }{}^{13}C{}_{\mathrm{TMS}-\mathrm{derivatives}}-3\mathrm{\delta }{}^{13}C{}_{\text{TMS}}]/C{}_{\text{n}}$$

The δ¹³C_alcohol and C_n denotes the δ¹³C value and the number of carbons in the target biomarker, respectively; δ¹³C_TMS is the δ¹³C value of the TMS in BSTFA, and δ¹³C_{TMS–derivatives} is the δ¹³C value for the TMS-derivative.

Each sample was measured at least in duplicate and the average values were used. Measurements were calibrated by several pulses of carbon dioxide with known δ¹³C value before and after each run. An n-alkane mixture (C₁₂ to C₃₅) of known isotopic composition was used to check instrument precision. All measured fractions and their compounds were diluted to concentrations of about 20 μg/ml. The injection volume was 1 to 2 μl. Generally, δ¹³C values of target compounds were only collected when peak heights were higher than one third of the reference carbon dioxide peaks (see Supplementary Figure S1). The standard deviation of the compound-specific carbon isotope measurement was <0.8‰.

Results

Fatty Acids

Generally, the net contents of fatty acids were much higher in gill than in foot tissues of G. platifrons, A. marissinica, and B. aduloides. Only for G. haimaensis, the contents of fatty acids were similar in gill and foot tissues. In foot tissues, PUFAs C₁₈, C₂₀, and C₂₂ were the most abundant fatty acids of G. platifrons, G. haimaensis, and B. aduloides with contents varying from 4 to 10 mg/g dry weight (dw). In contrast, both gill and foot tissues of A. marissinica were dominated by MUFAs. Among the MUFAs, C_16:1ω7 was the most abundant fatty acid, corresponding to 24% of total fatty acids in the gill tissues of G. platifrons, 30% of total fatty acids in B. aduloides, and 43% of total fatty acids in A. marissinica. However, PUFA C_22:2, accounting for 23% of total fatty acids, dominated in the gill tissue of G. haimaensis. The fatty acids C_16:1ω8 (12% and 9% of total fatty acids, respectively) and C_18:1ω8 (3% and 1% of total fatty acids, respectively) were only observed in the gill tissues of G. platifrons and G. haimaensis. Generally, the fatty acids C_16:1ω7, C_18:1ω7, and C_20:1ω7 were among the most abundant compounds of total fatty acids in gill and foot tissues of the four species, making up 32% and 22% for G. platifrons, 24% and 19% for G. haimaensis, and 58% and 28% for A. marissinica, and 44% and 17% for B. aduloides, respectively, with significantly higher relative abundances in gill tissues than in foot tissues. All relative percentages are referring to total fatty acids. Furthermore, C_16:1ω5 was found in the gill tissue of A. marissinica in trace amounts (<1% of total fatty acids), while C_16:1ω5 and C_18:1ω5 were present in both gill and foot tissues of B. aduloides (Figures 3, 4).

FIGURE 3

Figure 3. Partial gas chromatograms of fatty acids in gill tissues of (A) Gigantidas platifrons and (B) Bathymodiolus aduloides. The ω indicates the double bond position of monounsaturated fatty acids (MUFAs), the suffix c is for cis and t for trans, referring to stereoisomers of MUFAs. Double bond positions of polyunsaturated fatty acids (PUFAs) have not been determined.

FIGURE 4

Figure 4. Contents and carbon isotopic compositions of selected fatty acids and sterols for (A) Gigantidas platifrons and (B) Gigantidas haimaensis. Blue bars = contents of gill tissues; orange bars = contents of foot tissues; blue diamonds = δ¹³C values of biomarkers from gill tissues; orange diamonds = δ¹³C values of biomarkers from foot tissues.

The fatty acids from the gill and foot tissues of G. platifrons and G. haimaensis were strongly depleted in ¹³C, with δ¹³C values ranging from −75‰ to −59‰ and −72‰ to −54‰, respectively. Compared to G. platifrons and G. haimaensis, A. marissinica and B. aduloides yielded moderately negative δ¹³C values for fatty acids from −51‰ to −37‰ and −52‰ to −36‰, respectively (Table 2).

Lipid Biomarkers in the Alcohol Fractions

Various sterols were found in the gill and foot tissues of G. platifrons, G. haimaensis, A. marissinica, and B. aduloides (Table 3 and Figure 5). Even though the inventory of sterols was generally similar for G. platifrons and G. haimaensis, the contents of individual compounds varied significantly between the two species and types of tissues (Figure 4). The total content of sterols in the gill tissue of G. haimaensis was 60 mg/g dw, whereas the gill tissue content of sterols of G. platifrons was very low (2.6 mg/g dw). In B. aduloides, the sterol contents were even below 2 mg/g dw. Lanosterol (IX, see Figure 6 for structure) and cholesterol (I) were the most abundant sterols in the gill tissue of both G. haimaensis (16 mg/g dw) and G. platifrons (1.1 mg/g dw), representing 25% and 44% of the total alcohols, respectively. In B. aduloides, cholesterol (I) was the most abundant sterol, accounting for 46% and 53% of the total alcohols in gill and foot tissues, respectively, followed by lanost-8(9)-en-3β-ol (VIII), lanosterol, cholestanol, 24-methylenecholesterol (IV), and cholest-7-en-3β-ol (Table 3 and Figure 7). In the foot tissue, cholesterol (I) was most abundant in G. platifrons and G. haimaensis (0.4 mg/g dw and 12 mg/g dw, respectively), but was slightly more abundant in the gills (1.1 mg/g dw and 13 mg/g dw, respectively). Lanosterol, though, was not found in any of the foot tissue samples.

FIGURE 5

Figure 5. Gas chromatograms of alcohols in gill tissues of (A) Gigantidas platifrons and (B) Gigantidas haimaensis. Roman numerals indicate n-alcohols; ? denotes presently unknown; Cont. = contaminant.

FIGURE 6

Figure 6. Structures of sterols found in soft tissues of seep bivalves in the South China Sea.

FIGURE 7

Figure 7. Contents and carbon isotopic compositions of selected fatty acids and sterols for (A) Archivesica marissinica and (B) Bathymodiolus aduloides. Blue bars = contents of gill tissues; orange bars = contents of foot tissues; blue diamonds = δ¹³C values of biomarkers from gill tissues; orange diamonds = δ¹³C values of biomarkers from foot tissues.

Other sterols, such as nor-lanosterol (X), cholestanol (II), cholesta-5,24-dien-3β-ol (III), cholest-7-en-3β-ol (V), 4α-methylcholesta-8(14),24-dien-3β-ol (VI), and 4α-methylcholesta-8(14)-en-3β-ol (VII), were as well more prominent in the gill tissues than in foot tissues of G. platifrons and G. haimaensis. In B. aduloides, the composition and contents of sterols in the foot tissue are similar to gill tissue. The single hopanoid, found only in traces in the gills of G. platifrons, was diplopterol (Figure 5). Minute amounts of cholesterol (I), cholestanol (II), and stigmast-5-en-3β-ol (XI) were found in A. marissinica. The total sterols accounted for 17% and 6% of all alcohols in the gill and foot tissues, respectively, with relatively higher contents in the foot than in the gill.

The n-alcohols C_22:0, C_28:0, and C_30:0 were found in gill and foot tissues of G. haimaensis, A. marissinica, and B. aduloides. The gill and foot tissues revealed similar ¹³C depletion for sterols, with δ¹³C values varying from −84‰ to −78‰ and −80‰ to −74‰ for G. platifrons and G. haimaensis, respectively. For B. aduloides, cholesterol, cholestanol, cholest-7-en-3β-ol, and the 4,4-dimethyl sterols (lanosterol and lanost-8(9)-en-3β-ol) in both gill and foot tissues yielded δ¹³C values from −46‰ to −40‰, whereas the δ¹³C values of 24-methylenecholesterol in gill and foot tissues ranged from −32‰ to −30‰.

Discussion

Lipid Biomarker Signatures of Aerobic Methanotrophic Symbionts and Carbon Assimilation of Gigantidas platifrons and Gigantidas haimaensis

Both G. platifrons (Site F) and G. haimaensis (Haima seeps) were referred to as B. platifrons previously (Feng et al., 2015, 2018) and were reported to host methanotrophs (Duperron et al., 2009; Feng et al., 2015; Assié et al., 2016; Xu et al., 2019). Xu et al. (2019) analyzed the microbial 16S rRNA genes in gill tissue of G. haimaensis collected from the Haima seep site, and found that the bacteria were dominated by three phylotypes of symbiotic Gammaproteobacteria and three phylotypes of symbiotic Epsilonproteobacteria. The most abundant gammaproteobacterial phylotype of G. haimaensis clustered close to methanotrophic Gammaproteobacteria of other Gigantidas species and were assigned to Type I aerobic methanotrophic bacteria (Xu et al., 2019). Given Type II methanotrophs have never been reported in the marine realm (Knief, 2015), the symbionts harbored in G. platifrons are likely to be Type I methanotrophs too. Type I methanotrophs are the most prominent symbionts in the gills of both G. platifrons and G. haimaensis, which is supported by characteristic fatty acid inventories, including cis and trans isomers of C_16:1 with double bonds at ω6, ω7, ω8, and ω9 (Bowman et al., 1993) and a predominance of C_16:1ω8 fatty acid over C_18:1ω8 fatty acid (cf. Nichols et al., 1985; Bowman et al., 1991; Niemann et al., 2006; Riou et al., 2010).

In addition to fatty acids, 4,4-dimethyl sterols (lanosterol and nor-lanosterol), 4-methyl sterols (4α-methylcholesta-8(14),24-dien-3β-ol and 4α-methylcholesta-8(14)-en-3β-ol), and desmethyl sterols (cholesterol, cholestanol, cholesta-5,24-dien-3β-ol, and cholest-7-en-3β-ol) were observed in soft tissues of G. platifrons and G. haimaensis. The absolute and relative contents of sterols are generally different, for example lanosterol accounted for 15% and 25% of the total alcohols in gills of G. platifrons and G. haimaensis, respectively. However, the overall inventory of sterols of G. platifrons and G. haimaensis is similar (Table 3). Cholesterol is the most abundant sterol in the gill of G. platifrons (44%), whereas in G. haimaensis lanosterol dominates (25%; Figure 4). Cholesterol and other desmethyl sterols are common in eukaryotes, but lanosterol-like 4,4-dimethyl sterols and 4-methyl sterols are only present in free-living (Jahnke and Nichols, 1986; Cordova-Gonzalez et al., 2020) and endosymbiotic aerobic methanotrophs among bacteria (Jahnke et al., 1995; Kellermann et al., 2012). Both 4,4-dimethyl sterols and 4-methyl sterols are demethylation products of lanosterol in aerobic methanotrophic bacteria (Methylococcus capsulatus; Bouvier et al., 1976; Summons et al., 1994). Based on similar ¹³C-depletions and the co-occurrence of 4-methyl sterol intermediates and desmethylated cholesterol in soft tissues other than gills, cholesterol was supposed to be synthesized by the animal host using bacterial 4-methyl sterols as diet (Jahnke et al., 1995). For G. platifrons and G. haimaensis, 4-methyl sterols and cholesterol occurred in both gill and foot tissues and yielded similar δ¹³C values, indicating that 4-methyl sterols are most likely biosynthetic intermediates and represent metabolic precursors in cholesterol biosynthesis. The high abundance of 4,4-dimethyl sterols and 4-methyl sterols agrees with the presence of Type I methanotrophs, which are the only bacteria known to synthesize these source-specific compounds in noteworthy amounts (Bird et al., 1971; Schouten et al., 2000; Wei et al., 2016; Cordova-Gonzalez et al., 2020). In culture, Methylococcus capsulatus biosynthesizes fatty acids with significant ¹³C-depletion compared to methane, with Δ values ranging from −25‰ to −17‰ (Jahnke et al., 1999), whereas the fatty acids in the gills of type I methanotroph-containing mussels were more ¹³C-depleted by ca. −16‰ (Jahnke et al., 1995). At site F and Haima seeps, available δ¹³C values of methane approximately correspond to −60‰ (Feng et al., 2015; Fang et al., 2019). In the gills of G. platifrons and G. haimaensis, the symbiont-specific C_16:1ω8 fatty acid and most fatty acids yielded δ¹³C values from −75‰ to −72‰ and −75‰ to −65‰, respectively, which is consistent with fractionation patterns of fatty acids in type I/X methanotrophs and mussels that live in symbiosis with type I methanotrophs (Jahnke et al., 1995; Kellermann et al., 2012). Further, both 4,4-dimethyl sterols and 4-methyl sterols were more ¹³C-depleted than fatty acids, with an average Δδ¹³C_{sterols–fatty acids} between −13‰ and −8‰, similar to values reported from laboratory cultures (Summons et al., 1994; Jahnke et al., 1999; Cordova-Gonzalez et al., 2020). As for fatty acids, the isotopic pattern of sterols confirms a predominance of Type I and/or X methanotrophs (cf. Cordova-Gonzalez et al., 2020).

Although δ¹³C values of fatty acids and sterols were uniform for G. platifrons and G. haimaensis, contents of the various lipids varied in the two mussels. While the same fatty acid inventory was found in gill and foot tissues of both mussels, contents of fatty acids were much higher in G. platifrons (see Figure 4), whereas both 4,4-dimethyl and 4-methyl sterols were more abundant in G. haimaensis. The strong variation in the membrane lipid composition, especially the ratio of fatty acids over sterols, is most likely caused by variation of oxygen levels in the environment. Such an effect was shown by Jahnke and Nichols (1986) for Methylococcus capsulatus grown under varying oxygen tensions, revealing that sterols (4-methyl and 4,4-dimethyl sterols) and phospholipids were most abundant in cells grown with 0.5 and 1.1% oxygen. As oxygen was reduced, the amount of MUFA C_16:1 decreased but the 4,4-dimethyl sterols were predominantly produced at very low oxygen tensions (0.1%). Except for type X methanotrophs, oxygen level affecting membrane lipid composition was also reported for Methylobacterium organophilum (Type II methanotroph; Patt and Handon, 1978). Bathymodiolin mussels acquire symbionts horizontally and may take up various types and different amounts of symbionts in response to varying environmental conditions (Won et al., 2003; Kádár et al., 2005; Franke et al., 2021), which may also affect the biosynthesis of membrane lipids. In the present study, G. platifrons yielded significantly higher contents of MUFA C_16:1 than G. haimaensis, which may reflect a higher oxygen level for G. platifrons. Although no oxygen concentrations have been reported for the study sites, applying these observations to the biomarker patterns in G. haimaensis suggests that G. haimaensis most likely lived at a low level of dissolved oxygen.

Apart from the symbiont-specific fatty acids, the MUFAs C₂₀ and C₂₂, as well as PUFAs C₂₀ and C₂₂ of the animal host (Conway and McDowell-Capuzzo, 1991; Pond et al., 1998) were present in both G. platifrons and G. haimaensis. In chemotrophic symbioses, some bivalves are highly dependent on the metabolic intermediates transferred by the symbionts, which leads to a similar range of δ¹³C values for the lipids produced by hosts and symbionts (Jahnke et al., 1995; Pond et al., 1998; Kellermann et al., 2012). With respect to the seep-dwelling bivalves from the South China Sea, most symbiont- and host-specific fatty acids of G. platifrons and G. haimaensis yielded δ¹³C values from −75‰ to −68‰ and −72‰ to −60‰, respectively, indicating that the host obtained most of its nutrition either directly from the symbionts or consumes symbiont-derived carbon.

Nutritional Signatures in Archivesica marissinica and Bathymodiolus aduloides

Membrane lipids of free-living sulfide-oxidizing bacteria are usually characterized by prominent ω7 fatty acids with either C_16:1ω7 or C_18:1ω7 predominating (Jannasch, 1985; Guezennec and Fiala-Medioni, 1996; Arning et al., 2008). These compounds were also found to be abundant in thiotrophic bacteria living in mussels (Pond et al., 1998; Kellermann et al., 2012) and mussels cultured in the presence of hydrogen sulfide (Riou et al., 2010). The high abundance of MUFAs C_16:1ω7 and C_18:1ω7 and the absence of 4-methyl sterols in A. marissinica (yielding cholesterol, cholestanol, and stigmast-5-en-3β-ol instead) agrees with thiotrophic bacteria as only endosymbionts.

For B. aduloides, in addition to the abundant MUFAs C_16:1ω7 and C_18:1ω7, a range of sterols, including cholesterol, cholestanol, 24-methylenecholesterol, cholest-7-en-3β-ol, lanost-8(9)-en-3β-ol, and lanosterol, were found in both gill and foot tissues. It has been demonstrated that some mussels are capable of synthesizing sterols (Teshima and Kanazawa, 1974; Teshima and Patterson, 1981). Sterols in soft tissues of symbiont-containing mussels may originate from its symbionts, diets from ingested plant and algal material, or synthesis by the animal host. For B. aduloides, a derivation of sterols except for 24-methylenecholesterol from filter-feeding is unlikely, because sterols synthesized by plants and algae are characterized by high abundance of sterols alkylated at C-24 containing either a methyl group or ethyl group (Jarzebski and Popov, 1985; Volkman, 2003, 2005). Furthermore, such photosynthetically derived carbon reveals δ¹³C values around −30‰ to −20‰ (Conway and McDowell-Capuzzo, 1991; Kellermann et al., 2012), which is inconsistent with the carbon stable isotopic pattern of the B. aduloides sterols (from −46‰ to −40‰). Feng et al. (2018) reported δ¹³C_biomass values (−37‰ to −33‰) in soft tissues of B. aduloides from Site F of the South China Sea and concluded that the mussels host thiotrophic bacteria. However, the thiotrophic symbionts are not a potential source candidate for sterols, since no sulfide-oxidizing bacteria have been shown to synthesize sterols to date (McCaffrey et al., 1989; Conway and McDowell-Capuzzo, 1991). Given the absence of ω8 fatty acids, a derivation of 4,4-dimethyl sterols (lanosterol and lanost-8(9)-en-3β-ol) from aerobic methanotrophic bacteria is highly unlikely as well.

Apart from the bacterial domain, eukaryotes including animals are known to produce lanosterol. Using ¹⁴C-labeling in cultures revealed that the oyster Crassostrea virginica incorporated acetate to form squalene, 4,4-dimethylsterols, and 4-methylsterols, which can also convert lanosterol to cholesterol (Teshima and Patterson, 1981). These results suggest that the lanosterol in B. aduloides was most likely synthesized by the animal host. However, the lanost-8(9)-en-3β-ol as an intermediate has never been reported in aerobic methanotrophs and symbiont-containing invertebrates before, but it was found to co-occur with lanosterol in hyperlipidemic serum and rat livers (Gray et al., 1969; Scallen et al., 1971). As one of the intermediates in the conversion from lanosterol to cholesterol, the production of lanost-8(9)-en-3β-ol may occur as the first step when the side chain of lanosterol is reduced (Gray et al., 1969; Scallen et al., 1971). Although uncertainties are present, the absence of lanost-8(9)-en-3β-ol in aerobic methanotrophs is most likely caused by differences in biosynthetic steps from lanosterol to cholesterol between aerobic methanotrophs and animals.

In addition to nutrients supplied by the endosymbionts, B. aduloides complemented its diet by filter-feeding on photosynthetically derived carbon, as indicated by the presence of ¹³C-enriched 24-methylenecholesterol (δ¹³C values from −32‰ to −30‰) in both gill and foot tissues. Since consumers are generally ¹³C-enriched by only 0.4‰ to 1‰ compared to their diet (McCutchan et al., 2003), these δ¹³C values suggest higher plants or marine phytoplankton as sources (Goericke et al., 1994). Similar observations were made for the Pacific-Antarctic Ridge and the Gulf of Mexico. At the Pacific-Antarctic Ridge, a chemosynthesis-based community was observed to progressively shift its nutritional lifestyle to filter-feeding when diffuse hydrothermal venting vanished (Stecher et al., 2002). Reduction in symbiont abundances and changes in energy sources induced by decrease of hydrogen sulfide concentration were also reflected by very low contents of symbiont-specific lipids and relatively ¹³C-enriched cholesterol (−22‰) in Bathymodiolus cf. thermophilus from seeps of the Gulf of Mexico (Kellermann et al., 2012). Although both A. marissinica and B. aduloides contain thiotrophic bacteria, there are major differences when it comes to their symbionts. A. marissinica transmits the symbionts from parent to offspring (Ohishi et al., 2016; Lan et al., 2019), whereas B. aduloides acquires symbionts from the ambient environments for each new generation (Franke et al., 2021). Such differences in symbiont clades and amounts may lead to variations in the biomarker inventories of both hosts and symbionts. The above findings including our study confirm that lipid biomarkers and their isotope signatures are excellent tracers to decode varying environmental conditions and carbon and energy pathways in chemosynthesis-based ecosystems.

Conclusion

Four species of seep bivalves collected from Site F (Gigantidas platifrons) and Haima seeps (Gigantidas haimaensis, Bathymodiolus aduloides, and Archivesica marissinica) in the South China Sea were analyzed for lipid biomarkers and compound-specific carbon stable isotope patterns. Gigantidas platifrons and Gigantidas haimaensis were demonstrated to live in symbioses with methanotrophic bacterial symbionts as revealed by abundant ¹³C-depleted lanosterol, nor-lanosterol, 4-methylsterols, and monounsaturated fatty acids C_16:1ω9 and C_16:1ω8 extracted from gill tissues. The relatively higher abundance of C_16:1ω8 relative to C_18:1ω8 and the occurrence of 4,4-dimethylsterols and 4-methylsterols agree with an assignment to type I methanotrophs. The much higher sterol contents and the less abundant fatty acids in G. haimaensis compared to G. platifrons probably reflect low oxygen levels during the growth of G. haimaensis. The gill tissues of both B. aduloides and A. marissinica contained significant amounts of C_16:1ω7 and C_18:1ω7 fatty acids, and their nutritional status was found to be different from G. platifrons and G. haimaensis. A. marissinica apparently contains only thiotrophic bacteria, indicated by the lack of ω8 fatty acids, 4,4-dimethylsterols, and 4-methylsterols. Based on the biomarker patterns, B. aduloides is also suggested to contain thiotrophic bacteria, but the animal host complemented its diet through filter-feeding as revealed by relatively ¹³C-enriched 24-methylenecholesterol. The different types of 4-methyl sterols between the thiotroph-containing mussel (B. aduloides) and the methanotroph-containing mussels (G. platifrons and G. haimaensis) are likely caused by different biosynthetic steps from lanosterol to cholesterol between animal hosts and aerobic methanotrophs. Furthermore, the host-specific polyunsaturated fatty acids C₂₀ and C₂₂ in all four bivalve species yielded δ¹³C values close to those of symbiont-specific fatty acids, suggesting that the hosts acquired most of their nutrition from the symbionts.

Data Availability Statement

The original contributions presented in the study are included in the article/Supplementary Material, further inquiries can be directed to the corresponding authors.

Ethics Statement

Ethical review and approval was not required for the animal study because invertebrate studies at seeps are common.

Author Contributions

HG, DF, and JT collected the samples. HG did the experiments. HG, DB, and JP analyzed the biomarker data. All authors contributed to the preparation of the manuscript.

Funding

This research was supported by NSF of China (Grant Number 91958105) and the Shandong Special Fund of Pilot National Laboratory for Marine Science and Technology (Qingdao; Grant Number 2021QNLM020002).

Conflict of Interest

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Publisher’s Note

All claims expressed in this article are solely those of the authors and do not necessarily represent those of their affiliated organizations, or those of the publisher, the editors and the reviewers. Any product that may be evaluated in this article, or claim that may be made by its manufacturer, is not guaranteed or endorsed by the publisher.

Acknowledgments

We express our sincere appreciation to the crews of the Haima, ROPOS, Haiyang-06, and “Tan Kah Kee” Cruises for their professionalism in sampling. We thank S. Gao (GIG, CAS) for sample preparation and technical assistance. The manuscript benefited from insightful comments of the reviewers.

Supplementary Material

The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fmars.2022.831286/full#supplementary-material

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