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Applied Microbiology and Biotechnology

, Volume 94, Issue 1, pp 1–10 | Cite as

On the evolutionary ecology of symbioses between chemosynthetic bacteria and bivalves

  • Guus Roeselers
  • Irene L. G. Newton
Open Access
Mini-Review

Abstract

Mutualistic associations between bacteria and eukaryotes occur ubiquitously in nature, forming the basis for key ecological and evolutionary innovations. Some of the most prominent examples of these symbioses are chemosynthetic bacteria and marine invertebrates living in the absence of sunlight at deep-sea hydrothermal vents and in sediments rich in reduced sulfur compounds. Here, chemosynthetic bacteria living in close association with their hosts convert CO2 or CH4 into organic compounds and provide the host with necessary nutrients. The dominant macrofauna of hydrothermal vent and cold seep ecosystems all depend on the metabolic activity of chemosynthetic bacteria, which accounts for almost all primary production in these complex ecosystems. Many of these enigmatic mutualistic associations are found within the molluscan class Bivalvia. Currently, chemosynthetic symbioses have been reported from five distinct bivalve families (Lucinidae, Mytilidae, Solemyidae, Thyasiridae, and Vesicomyidae). This brief review aims to provide an overview of the diverse physiological and genetic adaptations of symbiotic chemosynthetic bacteria and their bivalve hosts.

Keywords

Clams Hydrothermal vents Methane Sulfur Oxidizing bacterial symbionts Chemoautotrophy 

Introduction

The term “symbiosis” was coined to describe associations in which different species live closely together, in relationships ranging from mutualisms to parasitism. Symbiosis has played a major role in shaping the evolution and diversity of eukaryotic organisms, and it is now clear that eukaryotic organelles such as chloroplasts and mitochondria evolved via ancient endosymbiotic events (Margulis 1970). Chemosynthetic endosymbiosis is a nutritional strategy that loosely parallels photosynthesis in higher plants. Whereas chloroplasts convert light energy into chemical energy, in the form of ATP and NADPH, chemosynthetic endosymbionts harness chemical energy stored in reduced sulfur compounds or methane. In chemosynthetic symbioses, the host provides access to reduced compounds (e.g., hydrogen sulfide) and oxygen that the bacterium uses to drive the formation of fixed carbon from single carbon molecules (either CO2 or CH4). The symbionts support a substantial fraction, if not all, of the host’s nutrition (Cavanaugh et al. 2006). Although known to occur within several invertebrate groups, it is within the class Bivalvia that this nutritional strategy is most widespread, both phylogenetically and geographically. Here we aim to review the diverse genetic and functional characteristics of chemosynthetic symbiosis in bivalves, which reflect the evolutionary ecology of both hosts and symbionts within this important invertebrate group.

Host morphology

The unusual nutritional strategy used by these symbiotic bivalves is reflected in their atypical morphology. Symbiotic bivalves are characterized by rudimentary or absent digestive systems. The principal anatomical adaptations are reduction of labial palps, reduction in length and volume of intestine, reduction in stomach volume, disappearance of crystalline style and style sac, reduction in digestive tubule volume, disappearance of digestive gland secretory cells, and, in some cases, total disappearance of the digestive system (Boss and Turner 1980). Bivalves typically harbor their symbionts in typically large and conspicuous gills, often accounting for more than one third of the animal’s total soft tissue weight (Distel 1998). However, chemosynthetic symbioses within Bivalvia are excellent model systems for studying the evolution of bacteria–eukaryote interactions, as they display a range of intimacies with some symbionts being housed intracellularly within specialized gill cells called bacteriocytes (as in the bathymodiolin mussels and the vesicomyid and lucinid clams) or extracellularly within the subfilamentar tissue of the host gill (as in the thyasirid clams) (Dubilier et al. 2008). In some Idas and Adipicola species, the symbionts are epibiotic and are attached to the surface of the gill tissue (Southward 2008). The relevance of this intimacy to host function and metabolism is not currently known.

Host diversity

Solemyidae

Protobranch bivalves of the family Solemyidae occur ubiquitously throughout the world’s oceans (Zardus 2002). The Solemyidae contains ~25 species across two genera, Acharax and Solemya. The solemyid range in size from less than 30 to over 200 mm (Acharax bartschii) and form symbioses with thioautotrophic bacterial symbionts within their gill epithelial cells.

Solemyids live in “Y”-shaped burrows (Stewart and Cavanaugh 2006) (Fig. 1) and are capable of swimming by jet propulsion through flexure of the shell margins with a pulsing expulsion of water through the posterior aperture (Reid 1980). Several key studies of solemyid symbioses have focused on the Atlantic species Solemya velum, which inhabits soft coastal sediments from Florida to Nova Scotia (Vokes 1955) and its Pacific congener Solemya reidi. Each of these hosts associates with a single, unique symbiont phylotype, whose carbon fixation supports most if not all of the nutritional requirements of the adult host (Krueger et al. 1992). Symbiont DNA has been extracted from the host ovarian tissue (Krueger et al. 1996b), raising the hypothesis that the symbiont is transmitted maternally (vertically) between successive host generations via the egg (Krueger et al. 1996b). To our knowledge, all attempts to culture symbionts of S. velum (or other hosts) have failed. Nonetheless, the existence of a free-living form of the symbionts could explain the rare signals of horizontal transfers between hosts (Stewart et al. 2009a, Stewart and Cavanaugh 2009).
Fig. 1

Sediment cross section exposing the characteristic Y-shaped burrow dug by S. velum. Positioning itself at the triple junction of the Y, the bivalve alternates between actively pumping oxygenated water from the upper arms of the burrow through the mantle cavity and across the gills and accessing reduced sulfur compounds diffusing up from the anoxic zones below and pumped through a ventral incurrent opening in the mantle. Scale bar equals 2.5 cm

Lucinidae

The family Lucinidae is by far the most disparate and species-rich of the chemosynthetic hosts and occupies the greatest variety of habitats over a broad geographical range. Habitats occupied by the over 400 living Lucinidae species range from intertidal zones and offshore sediments, seagrass beds (Barnes and Hickman 1999; Johnson et al. 2002), mangrove muds (Schweimanns and Felbeck 1985; Lebata 2001), and sewage effluent sites (Herry et al. 1989). Some species are associated with cold seeps and mud volcanoes (Brissac et al. 2010; Callender and Powell 1997; Carney 1994), oxygen minimum zones (Cary et al. 1989; Oliver and Holmes 2006), and a single species (Bathyaustriella thionipta) from a hydrothermal vent site (Glover et al. 2004). The lucinid range from a few millimeters to 150 mm in size and harbor thioautotrophic symbionts within their gill epithelial cells (Frenkiel and Moueza 1995; Taylor and Glover 2000). The lucinid symbionts are thought to be laterally transmitted each generation, leading to the increased genetic diversity seen within hosts (Williams et al. 2004). Phylogenetic analyses places the Lucinidae symbionts in a γ-proteobacterial clade including Solemya and vestimentiferan tube worm symbionts (Cavanaugh et al. 2006) (Fig. 2).
Fig. 2

Maximum likelihood tree illustrating the phylogenetic relationships among several thioautotrophic and methanoautotrophic symbionts of different bivalve hosts based on full-length 16S rRNA gene sequences selected from the ARB-SILVA SSU-Ref-108 database (http://www.arb-silva.de). Free-living bacteria are indicated with an asterisk. Agrobacterium tumefaciens [D14500], serving as an outgroup, was pruned from the ML tree. Bootstrap values ≥50% (1,000 replicates) are placed at the branch nodes

Vesicomyidae

The Vesicomyidae comprises at least 50 species occurring ubiquitously at hydrothermal vents, hydrocarbon seeps, and whale falls throughout the world’s oceans (Baco et al. 1999; Goffredi et al. 2003). They range in size from less than 5 mm to the extremely large species from vents and seeps that reach shell lengths up to 280 mm. Vesicomyidae retain only a rudimentary gut and rely instead on thioautotrophic gill endosymbionts for nutrition (Cavanaugh 1983).

Vesicomyid endosymbionts are found in host ovaries (Cary and Giovannoni 1993), and their genomes are highly reduced (Newton et al. 2007; Kuwahara et al. 2007) supporting the hypothesis that symbiont acquisition occurs via maternal inheritance in this group. Indeed, Vesicomyidae show a strong coupling of symbiont–host genetic variation (Peek et al. 1998). These symbioses are therefore considered useful models for studying bacteria–eukaryote coevolution. However, Stewart et al. (2009a, b) observed instances of decoupled symbiont and host mitochondrial phylogenies suggesting a mixed mode of symbiont transmission characterized by predominantly maternal transmission with periodical events of lateral symbiont acquisition.

Molecular analyses of symbionts from various species of Vesicomyidae from widely separated habitats show that they belong to one clade of γ-proteobacteria (Cavanaugh et al. 2006; Dubilier et al. 2008). These form a sister clade to the thioautotrophic symbionts from Bathymodiolus species, distinct from the symbiont group associated with shallow water families Lucinidae, Thyasiridae, and Solemyidae.

Mytilidae

Bathymodiolin mussels (family Mytilidae) are one of the most ecologically successful metazoans in the deep-sea; they are ubiquitous within these habitats, colonizing hydrothermal vents and cold seeps to wood falls, whale carcase, and oil impregnated muds from oil-drilling platform (Taylor and Glover 2010; Dubilier et al. 2008). The occurrence of bathymodiolin mussels in this broad range of environments may be attributed to their nutritional flexibility; they are capable of obtaining nutrition from filter feeding (Page et al. 1991) in addition to receiving nutrition from chemosynthetic symbionts (Dubilier et al. 1998; Fujiwara et al. 2010). The mussels can be very large; for example, Bathymodiolus boomerang reaches shell lengths of 370 mm and Gigantidas gladius to 316 mm (von Cosel and Marshall 2003). Sulfur and/or methane-oxidizing bacteria housed in their gill bacteriocytes cells have been reported for all studied species of bathymodiolins. All work done to date suggests that the bathymodiolin symbionts are acquired from the environment (Won et al. 2003)

Thyasiridae

Among the families of chemosymbiotic bivalves, the Thyasiridae stand out in several ways. Thyasiridae vary the most in their anatomical characters and in the extent of their nutritional reliance upon symbionts (Dufour 2005). Although Thyasiridae range in size from less than 1 to around 110 mm (Conchocele bisecta), most are less than 10 mm. They are globally distributed from polar to equatorial latitudes and live at all depths ranging from the intertidal to deepest trenches in the ocean (Taylor and Glover 2010).

Unlike the other chemosynthetic Bivalve families, only a fraction of Thyasiridae species harbor symbionts (Dufour 2005) which in most species are observed to be extracellular (Taylor and Glover 2010; Dufour 2005). This group may therefore be representative of early stages in the evolution of chemosynthetic bacterium–bivalve symbioses.

Symbiont transmission strategies

In order to continue to propagate these symbioses, each generation the host animals must be inoculated with the bacterial symbiont at some point during the life cycle, referred to as the transmission strategy for that organism. The transmission strategy utilized by a symbiosis has generally been inferred from phylogenetic evidence (lack or presence of codiversification) or the presence of bacterial DNA in the gametes of the host (Krueger et al. 1996b). Only for one bivalve group, bathymodiolin mussels, has the process of host colonization been experimentally tested. These experiments suggest that in these hosts, the symbionts are reacquired from the surrounding seawater after induced loss (Kadar et al. 2005; Won et al. 2003). The implications for host niche differentiation are quite clear: A host able to take up a genetically diverse pool of symbionts adapted to localized habitats may be at an advantage compared to those with static relationships. Similar mechanisms of niche adaptation have been suggested in other invertebrate symbioses, for instance, corals (Iglesias-Prieto et al. 2004).

Symbionts can be transmitted to the next host generation along a spectrum of transmission strategies: from strict vertical to strict lateral transmission. Vertically transmitted symbionts are passed down via the gametes (often intracellular). This transmission strategy imposes both a population bottleneck on the symbionts (reducing the symbiont genetic diversity available to the host and increasing the mutational load for the symbiont through genetic drift (Allen et al. 2009; Moran 1996)) as well as imposing codiversification (evidence often seen in phylogenetic analyses of host and symbiont). A vertical transmission strategy has been suggested for the Solemyidae and Vesicomyidae, based on phylogenetic and microscopic evidence. However, recent analyses suggest that vesicomyid bivalves are not strictly maternally transmitted (Stewart et al. 2009b) and that the symbiont of S. velum has a genome that contrast with those seen in vertically transmitted symbionts (Stewart et al. 2009a).

At the other end of the transmission strategy spectrum are those symbionts that are strictly laterally transmitted. These hosts must reacquire their symbionts each generation from a presumed free-living pool of bacteria. In contrast to the vertically transmitted symbioses, symbionts that are laterally transmitted are likely to be more genetically diverse within and between hosts, leading to the possibility of evolutionary conflicts. The bathymodiolin mussels and lucinid clams are thought to gain their symbiont through lateral transmission, culminating in the great diversity of symbionts we see in these hosts (Caro et al. 2007; Duperron et al. 2009). However, it is possible that the lateral transmission strategy comes with a substantial benefit; it would allow a host, during colonization of a new habitat, to sample bacteria from that environment, thereby selecting the organisms best suited for the symbiosis given the ecological parameters present.

Habitat

There are numerous environments in the biosphere where the biogeochemistry favors the colonization and emergence of chemosynthetic metabolisms. These sites are unified by the simultaneous availability of reduced compounds and molecular oxygen. Deep-sea hydrothermal vents were the first habitats in which chemosynthesis-driven primary production was shown to fuel large animal communities (Cavanaugh et al. 1981; Cavanaugh 1983). Hydrothermal vents result from geothermally driven circulation of seawater in the crust linked to magmatic activity at oceanic ridges. A few years after the discovery of the hydrothermal vent fauna, similar communities were discovered at cold seeps (Paull et al. 1984) where hydrocarbons and/or gas hydrates escape from the subsurface at continental margins. Later chemosymbiotic bivalves were found in a range of environments where sulfuric compounds originate from decaying organic matter as, such as in sea-grass beds (Cavanaugh 1983) and whale-falls (Bennett et al. 1994). Relevant to our discussions below, although each of these environments provides the biogeochemistry necessary for chemosynthetic metabolism, they differ in regards to the concentrations of particular reduced compounds and sources of nutrients available to the symbioses (Table 1). These geochemical differences may select for associations best specialized for a particular niche.
Table 1

Typical pore water concentrations of reduced compounds in different habitats

Habitat

Sulfur

Methane

References

Hydrothermal vents

3–40 mmol/kg

0.1–3.4 mmol/kg

Kelley et al. (2005); Cowen et al. (2002)

Cold seeps

0.57–19.43 mmol/kg

0.06–0.8 mmol/kg

Kelley et al. (2001); Barry et al. (1997)

Seagrass beds

5–35 μmol/kg

2–20 μmol/kg

Deborde et al. (2010); Calleja et al. (2007)

Symbiont diversity

It is striking that chemosynthesis has evolved in such phylogenetically diverse bivalve families ranging from the ancient Solemyidae, the bathymodiolin mussels to the heterodont families Thyasiridae, Lucinidae, and Vesicomyidae. In contrast to the remarkable phylogenetic divergence of the hosts involved in these symbioses, their chemosynthetic symbionts cluster all within a single bacterial class, the γ-proteobacteria, on the basis of 16S rRNA gene sequences (Distel and Cavanaugh 1994) (Fig. 2). Recent phylogenetic analyses of these symbionts revealed at least nine phylogenetically distinct clades within this bacterial class, most of which were interspersed with sequences from free-living bacteria (Dubilier et al. 2008). This evidence suggests that these symbioses have evolved multiple times, from different free-living bacterial ancestors.

The genetic and genomic diversity of within-host symbiont populations has been explored in a number of different chemosynthetic bivalves. The diversity of the population found within the host has the potential to affect host physiology, evolutionary adaptability, and define host niche. For example, in symbioses where a single, primary bacterial symbiont is known, polymorphisms can be observed with regards to symbiont genome number and bacterial size (as in the clam Codakia orbicularis) (Caro et al. 2007), and these differences may influence symbiont transcriptional response. A large amount of genetic diversity is expected in symbionts that are laterally transmitted (Stewart et al. 2009a, b; Dechaine et al. 2006), but even within primarily vertically transmitted symbioses, cryptic genetic diversity has been discovered within these presumed homogenous populations (Stewart and Cavanaugh 2009; Stewart et al. 2009b). The significance of this population level diversity is not yet known, but the implications for the evolution of the symbioses are large. Strict vertical transmission of symbionts is known to dramatically reduce genome content (Moran 2003). Indeed, the vertically transmitted vesicomyid symbionts have undergone dramatic genomic reductions, with the loss of many physiological capabilities (Kuwahara et al. 2007; Newton et al. 2007; Roeselers et al. 2010). If a symbiont population loses the ability to synthesize a particular nutrient, this can be particularly disadvantageous for the host. However, the existence of population level diversity within a host suggests avenues through with the bacterium might recuperate some function relevant to the symbiosis, perhaps thorough homologous recombination.

Within the bivalves, the bathymodiolin mussels are conspicuous for the huge diversity of symbionts found within an individual host’s tissues. Up to six bacterial symbionts have been identified in Idas species, with all symbionts capable of chemosynthesis (Duperron et al. 2008). More commonly, however, a dual symbiosis is seen in the mussels with a thioautotrophic symbiont co-occurring within the same cells as a methanoautotrophic symbiont (Duperron et al. 2006; Dechaine and Cavanaugh 2005). As detailed below, the levels of methane and sulfur in a particular environment may control the numbers of these distinct symbionts found within the host cells. This kind of symbiotic plasticity in combination with high dispersal ability, based on their larval shell morphology and small egg size (Le Pennec and Beninger 2000), may have allowed the bathymodiolin mussels to reach current cosmopolitan biogeographic distribution (Kyuno et al. 2009).

Functional genomics and ecology of chemosynthetic symbioses

The functionality of the chemosynthetic symbioses relies on both the availability of substrates for chemosynthetic metabolism and the existence of particular metabolic pathways in the symbiont to utilize those substrates. There is a link, therefore, between geochemistry and the type of symbiosis that can colonize a particular environment (Cordes et al. 2010). Hosts have adapted in a variety of ways to provide their symbionts with these essential substrates (for example, the behavioral extension of the foot to reach sulfur in thyasirid bivalves (Dufour and Felbeck 2003), but environmental concentrations are likely to dominate any ecological framework determining the localization and abundance of these symbioses.

Sulfur

For bivalves harboring thioautotrophic symbionts, the availability of sulfur (as well as a terminal electron acceptor such as oxygen) in their geochemical environment is an important variable in determining the presence of these symbioses (Van Dover 2000). Indeed, sulfur availability may define niche differentiation at cold seeps in Monterey Bay, where Calyptogena kilmeri inhabits areas of high porewater sulfide concentrations and Calyptogena pacifica beds surround the periphery of the C. kilmeri zone where sulfide concentrations are lower (Barry et al. 1997). Increased sulfide consumption rates, sulfide binding ability, and internal sulfide levels, as well as symbiont energy turnover, as measured by sulfide oxidation potential, sulfur metabolism enzymes, and symbiont densities suggest that C. pacifica, is physiologically better equipped for the uptake and transport of sulfide (Goffredi and Barry 2002).

The loss of genes encoding the synthesis of dissimilatory nitrate reductase in the C. pacifica symbiont Candidatus Ruthia magnifica and retention of the pathway in the C. R. magnifica symbiont Candidatus Vesicomyosocious okutanii may reflect differences in host geochemical ecology. The C. V. okutanii specimen for the Candidatus V. okutanii genome project was collected off Hatsushima Island, in the Sagami Bay seep sites (Kuwahara et al. 2007). The vesicomyid hosts found at Sagami Bay protrude their feet deep within the sediment to access sulfide and may spend the majority of their time in anoxic conditions, where nitrate may be the more abundant oxidant (Masuzawa et al. 1992). As eukaryotic hosts must use oxygen as a terminal electron acceptor, the retention of the nitrate reductase in the seep clam symbionts may reduce competition for oxygen between host and symbiont (Newton et al. 2008).

Thioautotrophic symbionts are the most common type in Bivalvia and are found in bathymodiolin mussels as well as solemyid, thyasirid, lucinid, and vesicomyid clams (Cavanaugh et al. 2006). In laboratory-controlled experiments, reducing the availability of sulfur to the lucinid host induced physiological and morphological changes in the symbiont. The symbiont populations are greatly reduced, their shape and size altered, and the gill of the host was restructured (Caro et al. 2009). These changes can be interpreted as the presumed digestion of the symbiont, a process that is likely normal and ongoing at a lower rate when sulfur is readily available but accelerated when geochemical conditions are not favorable. There are bivalves that seem to be better equipped to handle periods of low sulfur availability. Among them are thyasirid and lucinid clams that can turn to particulate feeding to supplement their diet under sulfur starvation (Dufour and Felbeck 2006). Another adaptation to periods of sulfur starvation on the part of the symbioses is the reacquisition of symbionts after the populations are diminished. After experimentally induced symbiont loss, the reacquisition of symbionts was seen in Bathymodiolus azoricus (Kadar et al. 2005). However, it remains unclear whether or not the symbionts were repopulated from the standing intracellular pool of organisms or reacquired from the environment.

Methane

Mussels of the Bathymodiolus genus are the only known bivalve symbioses to harbor either or both sulfur and methane-oxidizing symbionts within their cells. These symbioses were first described in the late 1980s from organisms found at cold seeps (Cavanaugh et al. 1987), but subsequently, Bathymodiolus mussels have been found to have a wide biogeographic distribution, being found globally in environments such as cold seeps and hydrothermal vents (Dechaine and Cavanaugh 2005). Mussels that harbor methanoautotrophic symbionts exclusively include Bathymodiolus japonicus, Bathymodiolus platifrons and Bathymodiolus childressi (Childress et al. 1986; Fujiwara et al. 2000). In these hosts, the symbionts have been identified through 16S rRNA gene sequence analysis, morphological features typical of methanotrophs (such as stacked intracellular membranes), the incorporation of radiolabeled 14-C methane, and diagnostic enzymes.

The Bathymodiolus mussels harbor their symbionts, whether sulfur or methane oxidizing, primarily within epithelial cells in the gill tissue (Fiala-Médioni et al. 2002; Fisher et al. 1993), although these bacteria have been detected in other organs of the host as well (Streams et al. 1997). The location of the symbionts has led to speculation about how these hosts acquire their symbionts; it is hypothesized that the symbionts are endocytosed by the epithelial gill cells from the surrounding seawater (Won et al. 2003). The cytological observations support a lateral transmission strategy for these mussels. Additional evidence comes from the lack of congruence between host and symbiont phylogenies (DeChaine et al. 2005) and the fact that multiple host species can harbor the same symbiont phylotype and diverse phylotypes exist within a single host (Duperron et al. 2006; Won et al. 2003; DeChaine and Cavanaugh 2005).

It has long been observed that environmental concentrations of methane and sulfur predict the presence of the chemosynthetic symbioses (Barry et al. 1997; Goffredi and Barry 2002; Sibuet and Olu 1998); this is not surprising, as these symbioses require access to these reduced chemicals. Interestingly, however, environmental levels of these geochemicals may also control the relative numbers of methanoautotrophic and thioautotrophic symbionts in B. azoricus and Bathymodiolus heckerae (Salerno et al. 2005; Riou et al. 2008). The ability of the Bathymodiolus mussels to harbor both kinds of symbionts and utilize either reduced substrate as an energy source may be a significant advantage in colonizing vent, seep, and woodfall environments.

Electron acceptors and anoxic conditions

In order for the chemosynthetic symbioses to function, the symbionts need access to a terminal electron acceptor. In the process of sulfur or methane oxidation, electrons from these reduced substrates can be donated to oxygen (oxic conditions) or a molecule like nitrate (anoxic conditions). Some chemosynthetic bivalves (such as S. reidi and Lucinoma aequizonata) have shown evidence of nitrate respiration when under anoxic and/or oxic conditions (Hentschel and Felbeck 1995; Wilmot and Vetter 1992). The ability to utilize an alternative electron acceptor is a potentially significant trait for these symbioses as the environments in which they reside are often anoxic; indeed, the animals often bridge the oxic/anoxic interface to be able to acquire both reduced substrate and electron acceptor. The host animal, unable to utilize nitrate as a terminal electron acceptor, is likely to consume available oxygen for its own respiration under hypoxic conditions. More evidence for the ability of symbionts to respire nitrate comes from genomic sequencing of the vesicomyid clam symbionts. As mentioned earlier, the C. kilmeri symbiont, has a functional nitrate reductase while the closely related C. R. magnifica symbiont does not (Kuwahara et al. 2007; Newton et al. 2007, 2008). Evidence from comparative genomic analyses suggests that C. R. magnifica’s symbiont lost the ability to utilize nitrate—the nitrate respiration genes are now pseudogenes in this organism (Newton et al. 2008). Clearly the pressure to maintain functionality of the nitrate reductase in C. R. magnificas symbiont could not overcome the effects of genomic reduction and pseudogenization in this linage. The observation that the primarily vertically transmitted vesicomyid symbionts are occasionally laterally transmitted (Stewart et al. 2009a, b) opens up an interesting evolutionary scenario for their hosts—the loss nitrate respiration might be recouped through homologous recombination between divergent symbionts.

Conclusions and perspectives

Much remains to be learned about the biology of chemosymbiosis in bivalves. Continued genetic, physiological, and evolutionary explorations promise to further increase our understanding of these remarkable symbiotic associations.

Advances in single-cell methods that combine imaging techniques with metabolic analyses, such as nanometer-scale secondary ion mass spectrometry and Raman microscopy, permit the visualization of uptake and distribution of substrates from the environment as well as measuring the rates at which substrates are acquired (Lechene et al. 2007). Single-cell genomics in combination with methods that allow isolation of specific cells of interest from microscopic regions of tissue/cells/organisms such as laser capture microdissection will be particularly valuable for the characterization of novel chemosynthetic symbiosis and to identify the roles of the different bacteria within hosts harboring multiple symbionts.

The recent introduction of next generation sequencing methods in microbial ecology has offered a powerful alternative to traditional clone library-based and microarray assessments of microbial diversity. Using barcoded amplicon pyrosequencing, a single run can generate >1 Gb of sequence data, one can increase by orders of magnitude the ability to detect rare but potentially functionally important genetic variants in a given microbial ecosystem. This technology has recently been used to probe the rare genetic diversity in symbiont populations (Stewart and Cavanaugh 2009).

Similarly, advances in next-generation sequencing methods have revolutionized the application of metatranscriptome analyses, which until recently were limited to microarray experiments or analysis of mRNA-derived cDNA clone libraries. High-throughput sequencing of cDNA pools derived directly from microbial communities allows analysis of the (meta)transcription profiles and may provide a molecular framework for interpreting physiochemical measurements of metabolism in chemosynthetic symbionts (Stewart et al. 2011).

Chemosynthetic symbiotic bacteria have never been cultured and their hosts, especially those adapted to extreme deep-sea environments, are difficult to maintain in the laboratory. Nevertheless, despite the advantages of culture-independent detection methods, through the enrichment and isolation of bacterial symbiont species in pure culture, one would be able to fully characterize the physiology of these remarkable organisms. Surprisingly, alternative isolation efforts using, e.g., diffusion chambers with minimal amounts of host tissue or co-occurring microorganisms, or long-term incubation experiments have not yet been documented. It is possible that certain co-adapted intracellular symbionts with highly reduced genomes can never be cultivated in pure culture in vitro. However, novel culturing strategies targeting those chemosynthetic symbionts that show signs of temporal environmental stages deserve adequate investigation.

Notes

Acknowledgments

This work was funded by a Rubicon grant from the Netherlands Organisation for Scientific Research (NWO) to GR.

Open Access

This article is distributed under the terms of the Creative Commons Attribution Noncommercial License which permits any noncommercial use, distribution, and reproduction in any medium, provided the original author(s) and source are credited.

References

  1. Allen JM, Light JE, Perotti MA, Braig HR, Reed DL (2009) Mutational meltdown in primary endosymbionts: selection limits Muller’s ratchet. PLoS One 4:e4969CrossRefGoogle Scholar
  2. Baco AR, Smith CR, Peek AS, Roderick GK, Vrijenhoek RC (1999) The phylogenetic relationships of whale-fall vesicomyid clams based on mitochondrial COI DNA sequences. Mar Ecol Prog Ser 182:137–147CrossRefGoogle Scholar
  3. Barnes PAG, Hickman CS (1999) Lucinid bivalves and marine angiosperms: a search for causal relationships. In: Walker DI, Wells FE (eds) The seagrass flora and fauna of Rottnest Island, Western Australia. Western Australian Museum, Perth, pp 215–238Google Scholar
  4. Barry JP, Kochevar RE, Baxter CH (1997) The influence of pore-water chemistry and physiology on the distribution of vesicomyid clams at cold seeps in Monterey Bay: implications for patterns of chemosynthetic community organization. Limnol Oceanogr 42:318–328CrossRefGoogle Scholar
  5. Bennett BA, Smith CR, Glaser B, Maybaum HL (1994) Faunal community structure of a chemoautotrophic assemblage on whale bones in the deep northeast Pacific Ocean. Mar Ecol Prog Ser 108:205–223CrossRefGoogle Scholar
  6. Boss KJ, Turner RD (1980) The giant white clam from the Galapagos Rift, Calyptogena magnifica n. sp. (Bivalvia; Vesicomyidae). Malacologia 20:161–194Google Scholar
  7. Brissac T, Rodrigues CF, Gros O, Duperron S (2010) Characterization of bacterial symbioses in Myrtea sp. (Bivalvia: Lucinidae) and Thyasira sp. (Bivalvia: Thyasiridae) from a cold seep in the Eastern Mediterranean. Mar Ecol-Evol Persp 32:198–210Google Scholar
  8. Calleja ML, Marbà N, Duarte CM (2007) The relationship between seagrass (Posidonia oceanica) decline and sulfide porewater concentration in carbonate sediments. Estuar Coast Shelf Sci 73:583–588CrossRefGoogle Scholar
  9. Callender WR, Powell EN (1997) Autochthonous death assemblages from chemoautotrophic communities at petroleum seeps: palaeoproduction, energy flow, and implications for the fossil record. Hist Biol 12:165–198CrossRefGoogle Scholar
  10. Carney RS (1994) Consideration of the oasis analogy for chemosynthetic communities at Gulf-of-Mexico hydrocarbon vents. Geo-Mar Lett 14:149–159CrossRefGoogle Scholar
  11. Caro A, Gros O, Got P, De Wit R, Troussellier M (2007) Characterization of the population of the sulfur-oxidizing symbiont of Codakia orbicularis (Bivalvia, Lucinidae) by single-cell analyses. Appl Environ Microb 73:2101–2109CrossRefGoogle Scholar
  12. Caro A, Got P, Bouvy M, Troussellier M, Gros O (2009) Effects of long-term starvation on a host bivalve (Codakia orbicularis, Lucinidae) and its symbiont population. Appl Environ Microb 75:3304–3313CrossRefGoogle Scholar
  13. Cary SC, Giovannoni SJ (1993) Transovarial inheritance of endosymbiotic bacteria in clams inhabiting deep-sea hydrothermal vents and cold seeps. Proc Natl Acad Sci U S A 90:5695–5699CrossRefGoogle Scholar
  14. Cary SC, Vetter RD, Felbeck H (1989) Habitat characterization and nutritional strategies of the endosymbiont-bearing bivalve Lucinoma aequizonata. Mar Ecol Prog Ser 55:31–45CrossRefGoogle Scholar
  15. Cavanaugh CM (1983) Symbiotic Chemoautotrophic bacteria in marine-invertebrates from sulfide-rich habitats. Nature 302:58–61CrossRefGoogle Scholar
  16. Cavanaugh CM, Gardiner SL, Jones ML, Jannasch HW, Waterbury JB (1981) Prokaryotic cells in the hydrothermal vent tube worm Riftia pachyptila Jones—possible chemoautotrophic symbionts. Science 213:340–342CrossRefGoogle Scholar
  17. Cavanaugh CM, Levering PR, Maki JS, Mitchell R, Lidstrom ME (1987) Symbiosis of methylotrophic bacteria and deep-sea mussels. Nature 325:346–348CrossRefGoogle Scholar
  18. Cavanaugh CM, McKiness ZP, Newton ILG, Stewart FJ (2006) Marine chemosynthetic symbioses. In: Dworkin M, Falkow S, Rosenberg E, Schleifer KH, Stackebrandt E (eds) The prokaryotes, 3rd edn. Springer, New YorkGoogle Scholar
  19. Childress JJ, Fisher CR, Brooks JM, Kennicutt MC, Bidigare R, Anderson AE (1986) A methanotrophic marine molluscan (Bivalvia, Mytilidae) symbiosis: mussels fueled by gas. Science 233:1306–1308CrossRefGoogle Scholar
  20. Cordes EE, Cunha MR, Galeron J, Mora C, Olu-Le, Roy K, Sibuet M, Van Gaever S, Vanreusel A, Levin LA (2010) The influence of geological, geochemical, and biogenic habitat heterogeneity on seep biodiversity. Mar Ecol-Evol Persp 31:51–65Google Scholar
  21. Cowen JP, Wen XY, Popp BN (2002) Methane in aging hydrothermal plumes. Geochim Cosmochim Acta 66(20):3563–3571CrossRefGoogle Scholar
  22. Deborde J, Anschutz P, Guerin F, Poirier D, Marty D, Boucher G, Thouzeau G, Canton M, Abril G (2010) Methane sources, sinks, and atmospheric fluxes in a temperate tidal lagoon (Arcachon Bay, France). Estuar Coast Shelf Sci 89:256–266CrossRefGoogle Scholar
  23. DeChaine EG, Cavanaugh CM (2005) Symbioses of methanotrophs and deep-sea mussels (Mytilidae: Bathymodiolinae). In: Overmann J (ed) Molecular basis of symbiosis. Springer, New York, pp 227–249Google Scholar
  24. DeChaine EG, Bates AE, Shank TM, Cavanaugh CM (2006) Off-axis symbiosis found: characterization and biogeography of bacterial symbionts of Bathymodiolus mussels from Lost City hydrothermal vents. Environ Microbiol 8:1902–1912CrossRefGoogle Scholar
  25. Distel DL (1998) Evolution of chemoautotrophic endosymbioses in bivalves. BioScience 48:277–286CrossRefGoogle Scholar
  26. Distel DL, Cavanaugh CM (1994) Independent phylogenetic origins of methanotrophic and chemoautotrophic bacterial endosymbioses in marine bivalves. J Bact 176:1932–1938Google Scholar
  27. Dubilier N, Windoffer R, Giere O (1998) Ultrastructure and stable carbon isotope composition of the hydrothermal vent mussels Bathymodiolus brevior and B. sp. affinis brevior from the North Fiji Basin, western Pacific. Mar Ecol Prog Ser 165:187–193CrossRefGoogle Scholar
  28. Dubilier N, Bergin C, Lott C (2008) Symbiotic diversity in marine animals: the art of harnessing chemosynthesis. Nat Rev Microbiol 6:725–740CrossRefGoogle Scholar
  29. Dufour SC (2005) Gill anatomy and the evolution of symbiosis in the bivalve family Thyasiridae. Bio Bull 208:200–212CrossRefGoogle Scholar
  30. Dufour SC, Felbeck H (2003) Sulphide mining by the superextensile foot of symbiotic thyasirid bivalves. Nature 426:65–67CrossRefGoogle Scholar
  31. Dufour SC, Felbeck H (2006) Symbiont abundance in thyasirids (Bivalvia) is related to particulate food and sulphide availability. Mar Ecol Prog Ser 320:185–194CrossRefGoogle Scholar
  32. Duperron S, Bergin C, Zielinski F, Blazejak A, Pernthaler A, McKiness ZP, DeChaine E, Cavanaugh CM, Dubilier N (2006) A dual symbiosis shared by two mussel species, Bathymodiolus azoricus and Bathymodiolus puteoserpentis (Bivalvia: Mytilidae), from hydrothermal vents along the northern Mid-Atlantic Ridge. Environ Microbiol 8:1441–1447CrossRefGoogle Scholar
  33. Duperron S, Halary S, Lorion J, Sibuet M, Gaill F (2008) Unexpected co-occurrence of six bacterial symbionts in the gills of the cold seep mussel Idas sp. (Bivalvia: Mytilidae). Environ Microbiol 10:433–445CrossRefGoogle Scholar
  34. Duperron S, Lorion J, Samadi S, Gros O, Gaill F (2009) Symbioses between deep-sea mussels (Mytilidae: Bathymodiolinae) and chemosynthetic bacteria: diversity, function and evolution. CR Biol 332:298–310CrossRefGoogle Scholar
  35. Fiala-Médioni A, McKiness ZM, Dando PD, Boulegue JB, Mariotti AM, Alayse-Danet A, Robinson J, Cavanaugh CM (2002) Ultrastructural, biochemical, and immunological characterization of two populations of the mytilid mussel Bathymodiolus azoricus from the Mid-Atlantic Ridge: evidence for a dual symbiosis. Mar Biol 141:1035–1043CrossRefGoogle Scholar
  36. Fisher CR, Brooks JM, Vodenichar JS, Zande JM, Childress JJ, Burke RA (1993) The co-occurrence of methanotrophic and chemoautotrophic sulfur-oxidizing bacterial symbionts in a deep-sea mussel. Mar Ecol-Evol Persp 14:277–289Google Scholar
  37. Frenkiel L, Moueza M (1995) Gill ultrastructure and symbiotic bacteria in Codakia orbicularis (Bivalvia, Lucinidae). Zoomorphology 115:51–61CrossRefGoogle Scholar
  38. Fujiwara Y, Takai K, Uematsu K, Tsuchida S, Hunt JC, Hashimoto J (2000) Phylogenetic characterization of endosymbionts in three hydrothermal vent mussels: influence on host distributions. Mar Ecol Prog Ser 208:147–155CrossRefGoogle Scholar
  39. Fujiwara Y, Kawato M, Noda C, Kinoshita G, Yamanaka T, Fujita Y, Uematsu K, Miyazaki JI (2010) Extracellular and mixotrophic symbiosis in the whale-fall mussel Adipicola pacifica: a trend in evolution from extra- to intracellular symbiosis. PLoS One. doi: 10.1371/journal.pone.0011808
  40. Glover EA, Taylor JD, Rowden AA (2004) Bathyaustriella thionipta, a new lucinid bivalve from a hydrothermal vent on the Kermadec Ridge, New Zealand and its relationship to shallow-water taxa (Bivalvia: Lucinidae). J Mollus Stud 70:283–295CrossRefGoogle Scholar
  41. Goffredi SK, Barry JP (2002) Species-specific variation in sulfide physiology between closely related Vesicomyid clams. Mar Ecol Prog Ser 225:227–238CrossRefGoogle Scholar
  42. Goffredi SK, Hurtado LA, Hallam S, Vrijenhoek RC (2003) Evolutionary relationships of deep-sea vent and seep clams (Mollusca: Vesicomyidae) of the ‘pacifica/lepta’ species complex. Mar Biol 142:311–320Google Scholar
  43. Hentschel U, Felbeck H (1995) Nitrate respiration in chemoautotrophic symbionts of the bivalve Lucinoma aequizonata is not regulated by oxygen. Appl Environ Microb 61:1630–1633Google Scholar
  44. Herry A, Diouris M, Pennec M (1989) Chemoautotrophic symbionts and translocation of fixed carbon from bacteria to host tissues in the littoral bivalve Loripes lucinalis (Lucinidae). Mar Biol 101:305–312CrossRefGoogle Scholar
  45. Iglesias-Prieto R, Beltran VH, LaJeunesse TC, Reyes-Bonilla H, Thome PE (2004) Different algal symbionts explain the vertical distribution of dominant reef corals in the eastern Pacific. Proc R Soc Lond B Biol Sci 271:1757–1763CrossRefGoogle Scholar
  46. Johnson MA, Fernandez C, Pergent G (2002) The ecological importance of an invertebrate chemoautotrophic symbiosis to phanerogam seagrass beds. B Mar Sci 71:1343–1351Google Scholar
  47. Kadar E, Bettencourt R, Costa V, Santos RS, Lobo-Da-Cunha A, Dando P (2005) Experimentally induced endosymbiont loss and re-acquirement in the hydrothermal vent bivalve Bathymodiolus azoricus. J Exp Mar Biol Ecol 318:99–110CrossRefGoogle Scholar
  48. Kelley DS, Karson JA, Blackman DK, Früh-Green GL, Butterfield DA, Lilley MD, Olson EJ, Schrenk MO, Roe KK, Lebon GT, Rivizzigno P, and the AT3–60 Shipboard Party (2001) An off-axis hydrothermal vent field near the Mid-Atlantic Ridge at 30°N. Nature 412:145–149CrossRefGoogle Scholar
  49. Kelley DS, Karson JA, Früh-Green GL, Yoerger DR, Shank TM, Butterfield DA, Hayes JM, Schrenk MO, Olson EJ, Proskurowski G, Jakuba M, Bradley A, Larson B, Ludwig K, Glickson D, Buckman K, Bradley AS, Brazelton WJ, Roe K, Elend MJ, Delacour A, Bernasconi SM, Lilley MD, Baross JA, Summons RE, Sylva SP (2005) A serpentinite-hosted ecosystem: the Lost City hydrothermal field. Science 307:1428–1434CrossRefGoogle Scholar
  50. Krueger DM, Gallager SM, Cavanaugh CM (1992) Suspension feeding on phytoplankton by Solemya velum, a symbiont-containing clam. Mar Ecol Prog Ser 86:145–151CrossRefGoogle Scholar
  51. Krueger DM, Gustafson RG, Cavanaugh CM (1996a) Vertical transmission of chemoautotrophic symbionts in the bivalve Solemya velum (Bivalvia: Protobranchia). Bio Bull 190:195–202CrossRefGoogle Scholar
  52. Kuwahara H, Yoshida T, Takaki Y, Shimamura S, Nishi S, Harada M, Matsuyama K, Takishita K, Kawato M, Uematsu K, Fujiwara Y, Sato T, Kato C, Kitagawa M, Kato I, Maruyama T (2007) Reduced genome of the thioautotrophic intracellular symbiont in a deep-sea clam, Calyptogena okutanii. Curr Biol 17(10):881–886CrossRefGoogle Scholar
  53. Kyuno A, Shintaku M, Fujita Y, Matsumoto H, Utsumi M, Watanabe H, Fujiwara Y, Miyazaki JI (2009) Dispersal and differentiation of deep-sea mussels of the genus Bathymodiolus (Mytilidae, Bathymodiolinae). J Mar Biol. doi: 10.1155/2009/625672
  54. Le Pennec M, Beninger PG (2000) Reproductive characteristics and strategies of reducing-system bivalves. Comp Biochem Physiol A Mol Integr Physiol 126:1–16CrossRefGoogle Scholar
  55. Lebata MJHL (2001) Oxygen, sulphide and nutrient uptake of the mangrove mud clam Anodontia edentula (Family: Lucinidae). Mar Pollut Bull 42:1133–1138CrossRefGoogle Scholar
  56. Lechene CP, Luyten Y, McMahon G, Distel DL (2007) Quantitative imaging of nitrogen fixation by individual bacteria within animal cells. Science 317:1563–1566CrossRefGoogle Scholar
  57. Margulis, L., (1970) Origin of Eukaryotic Cells (Yale University Press, New Haven, Connecticut)Google Scholar
  58. Masuzawa T, Handa N, Kitagawa H, Kusakabe M (1992) Sulfate reduction using methane in sediments beneath a bathyal “cold seep” giant clam community off Hatsushima Island, Sagami Bay, Japan. Earth Planet Sci Lett 110:39–50CrossRefGoogle Scholar
  59. Moran NA (1996) Accelerated evolution and Muller’s rachet in endosymbiotic bacteria. Proc Natl Acad Sci U S A 93:2873–2878CrossRefGoogle Scholar
  60. Moran NA (2003) Tracing the evolution of gene loss in obligate bacterial symbionts. Curr Opin Microbiol 6:512–518CrossRefGoogle Scholar
  61. Newton ILG, Woyke T, Auchtung TA, Dilly GF, Dutton RJ, Fisher MC, Fontanez KM, Lau E, Stewart FJ, Richardson PM, Barry KW, Saunders E, Detter JC, Wu D, Eisen JA, Cavanaugh CM (2007) The Calyptogena magnifica chemoautotrophic symbiont genome. Science 315:998–1000CrossRefGoogle Scholar
  62. Newton ILG, Girguis PR, Cavanaugh CM (2008) Comparative genomics of vesicomyid clam (Bivalvia: Mollusca) chemosynthetic symbionts. BMC Genomics 9:585CrossRefGoogle Scholar
  63. Oliver PG, Holmes AM (2006) A new species of Lucinoma (Bivalvia: Lucinoidea) from the oxygen minimum zone of the Oman margin, Arabian Sea. J Conchol 39:63–77Google Scholar
  64. Page HM, Fiala-Medioni A, Fisher CR, Childress JJ (1991) Experimental evidence for filter-feeding by the hydrothermal vent mussel, Bathymodiolus thermophilus. Deep Sea Res Part 1 Oceanogr Res Pap 38:1455–1461CrossRefGoogle Scholar
  65. Paull CK, Hecker B, Commeau R, Freeman-Lynde RP, Neumann C, Corso WP, Golubic S, Hook JE, Sikes E, Curray J (1984) Biological communities at the Florida escarpment resemble hydrothermal vent taxa. Science 226:965–967CrossRefGoogle Scholar
  66. Peek AS, Feldman RA, Lutz RA, Vrijenhoek RC (1998) (1998) Cospeciation of chemoautotrophic bacteria and deep-sea clams. Proc Natl Acad Sci U S A 95:9962–9966CrossRefGoogle Scholar
  67. Reid RGB (1980) Aspects of the biology of a gutless species of Solemya (Bivalvia: Protobranchia). Can J Zool 58:386–393CrossRefGoogle Scholar
  68. Riou V, Halary S, Duperron S, Bouillon S, Elskens M, Bettencourt R, Santos RS, Dehairs F, Colaço A (2008) Influence of CH4 and H2S availability on symbiont distribution, carbon assimilation and transfer in the dual symbiotic vent mussel Bathymodiolus azoricus. Biogeosciences 5:1681–1691CrossRefGoogle Scholar
  69. Roeselers G, Newton ILG, Woyke T, Auchtung TA, Dilly GF, Dutton RJ, Fisher MC, Fontanez KM, Lau E, Stewart FJ, Richardson PM, Barry KW, Saunders E, Detter JC, Wu D, Eisen JA, Cavanaugh CM (2010) Complete genome sequence of Candidatus Ruthia magnifica. Stand Genomic Sci 3:163–173CrossRefGoogle Scholar
  70. Salerno JL, Macko SA, Hallam SJ, Bright M, Won YJ, McKiness Z, Van Dover CL (2005) Characterization of symbiont populations in life-history stages of mussels from chemosynthetic environments. Bio Bull 208:145–155CrossRefGoogle Scholar
  71. Schweimanns M, Felbeck H (1985) Significance of the occurrence of chemoautotrophic bacterial endosymbionts in lucinid clams from Bermuda. Mar Ecol Progr Ser 24:113–120CrossRefGoogle Scholar
  72. Sibuet M, Olu K (1998) Biogeography, biodiversity and fluid dependence of deep-sea cold-seep communities at active and passive margins. Deep Sea Res Part 2 Top Stud Oceanogr 45:517–567CrossRefGoogle Scholar
  73. Southward EC (2008) The morphology of bacterial symbioses in the gills of mussels of the genera Adipicola and Idas (Bivalvia: Mytilidae). J Shellfish Res 27:139–146CrossRefGoogle Scholar
  74. Stewart FJ, Cavanaugh CM (2006) Bacterial endosymbioses in Solemya (Mollusca: Bivalvia)—model systems for studies of symbiont–host adaptation. Antonie Van Leeuwenhoek 90:343–360CrossRefGoogle Scholar
  75. Stewart FJ, Cavanaugh CM (2009) Pyrosequencing analysis of endosymbiont population structure: co-occurrence of divergent symbiont lineages in a single vesicomyid host clam. Environ Microbiol 11:2136–2147CrossRefGoogle Scholar
  76. Stewart FJ, Baik AHY, Cavanaugh CM (2009a) Genetic subdivision of chemosynthetic endosymbionts of Solemya velum along the southern New England coast. Appl Environ Microb 75:6005–6007CrossRefGoogle Scholar
  77. Stewart FJ, Young CR, Cavanaugh CM (2009b) Evidence for homologous recombination in intracellular chemosynthetic clam symbionts. Mol Biol Evol 26:1391–1404CrossRefGoogle Scholar
  78. Stewart FJ, Dmytrenko O, DeLong EF, Cavanaugh CM (2011) Metatranscriptomic analysis of sulfur oxidation genes in the endosymbiont of Solemya velum. Front Microbiol 2:134Google Scholar
  79. Streams ME, Fisher CR, FialaMedioni A (1997) Methanotrophic symbiont location and fate of carbon incorporated from methane in a hydrocarbon seep mussel. Mar Biol 129:465–476CrossRefGoogle Scholar
  80. Taylor JD, Glover EA (2000) Functional anatomy, chemosymbiosis and evolution of the Lucinidae. In: The evolutionary biology of the Bivalvia. Geological Society Special Publications. Geological Society, London, pp 207–225Google Scholar
  81. Taylor JD, Glover EA (2010) Chemosymbiotic bivalves. In: Landman NH, Jones DS (eds) The vent and seep biota. Springer, The Netherlands, pp 107–135CrossRefGoogle Scholar
  82. Van Dover CL (2000) The ecology of deep-sea hydrothermal vents. Princeton University Press, PrincetonGoogle Scholar
  83. Vokes HE (1955) Notes on tertiary and recent Solemyacidae. J Paleontol 29:534–545Google Scholar
  84. von Cosel R, Marshall BA (2003) Two new species of large mussels (Bivalvia: Mytilidae) from active submarine volcanoes and a cold seep off the eastern North Island of New Zealand, with description of a new genus. Nautilus 117:31–46Google Scholar
  85. Williams ST, Taylor JD, Glover EA (2004) Molecular phylogeny of the Lucinoidea (Bivalvia): non-monophyly and separate acquisition of bacterial chemosymbiosis. J Mollus Stud 70:187–202CrossRefGoogle Scholar
  86. Wilmot DB, Vetter RD (1992) Oxygen- and nitrogen-dependent sulfur metabolism in the thiotrophic clam Solemya reidi. Biol Bull 182:444–453CrossRefGoogle Scholar
  87. Won YJ, Hallam SJ, O'Mullan GD, Pan IL, Buck KR, Vrijenhoek RC (2003) Environmental acquisition of thiotrophic endosymbionts by deep-sea mussels of the genus Bathymodiolus. Appl Environ Microb 69:6785–6792CrossRefGoogle Scholar
  88. Zardus JD (2002) Protobranch bivalves. Advances Mar Biol 42:1–65CrossRefGoogle Scholar

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© The Author(s) 2011

Authors and Affiliations

  1. 1.Microbiology and Systems Biology GroupTNOZeistThe Netherlands
  2. 2.Department of BiologyIndiana UniversityBloomingtonUSA

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