Characterization of Bacterial Symbionts in Deep-Sea ...

Characterization of Bacterial Symbionts in Deep-Sea Fauna: Protocols for Sample Conditioning, Fluorescence In Situ Hybridization, and Image Analysis Se´bastien Duperron Abstract Symbioses with bacteria are key adaptations allowing various groups of metazoans to reach high biomasses at deep sea reducing habitats including hydrothermal vents and cold seeps. Characterizing these associations is challenging due to the constraints associated with work on deep-sea organisms. These include limited sample availability, impact of recovery procedures and shipment on sample quality, and general lack of environmental data. In this chapter, a standard procedure for sample processing at sea which can maximize sample use back in the laboratory is presented, with example protocols for sample fixation, fluorescence in situ hybridization (FISH)-based localization of symbionts in animal tissues, and estimation of their relative abundances in the case of multiple symbioses. Keywords: Deep sea, Fluorescence in situ hybridization, Sample conditioning, Symbiosis

1

Introduction The study of bacterial symbioses in animal hosts has enjoyed renewed interest in recent years, thanks to advances in culture-independent approaches with which information can be acquired without the need to isolate symbionts in pure cultures [1]. In the marine realm, symbioses are of prime importance to their hosts in various habitats and fundamental in maintaining the high biomass often recorded at deep-sea cold seeps and hydrothermal vents [2]. There, diverse metazoan groups including arthropods, annelids, and mollusks associate with different types of bacteria [3]. The most common associations involve sulfur-oxidizing Gamma- or Epsilonproteobacteria and methane-oxidizing Gammaproteobacteria, the latter being mainly reported in mytilid bivalves [4]. However, several recent studies employing molecular and microscopic approaches have revealed a broader diversity of bacterial taxa involved in symbiosis including methylotroph- and Cycloclasticus-related symbionts, member of the

T.J. McGenity et al. (eds.), Hydrocarbon and Lipid Microbiology Protocols, Springer Protocols Handbooks, DOI 10.1007/8623_2015_73, © Springer-Verlag Berlin Heidelberg 2015

Se´bastien Duperron

Bacteroidetes, as well as various unknown Gammaproteobacteria, some of which even occur within host cell nuclei [5–8]. Depending on the host taxa, symbionts can occur on the tegument surface (annelids, arthropods), in gills (mollusks), in the gut or digestive gland (arthropods, mollusks, annelids), or in dedicated organs, such as the trophosome in some annelids. Vertically transmitted symbionts also occur in the gonads, while horizontally acquired bacteria may occur free-living in the environment [9–12]. Gaining information regarding the identity, localization, and metabolism of bacterial symbionts is mandatory for understanding the symbiotic system. However, deep-sea biology is by nature sample limited due to constraints associated with sample collection, be they performed using submersibles or surface-operated devices such as box corers. Limited sample availability (not to mention rarity of certain species), pressure-related issues during recovery, sample processing onboard, and shipment must all be properly managed so that all necessary material is available back in the laboratory. Sampling and conditioning strategy must thus be planned carefully, particularly when no further visits to the deepsea site are anticipated. The aim of this chapter is to describe a standard procedure for sample processing at sea which can maximize sample use by permitting a wide array of techniques to be carried out in the laboratory with example protocols for the localization and relative abundance measurement of microbial symbionts provided. The first section is dedicated to study design, which summarizes important points to be considered prior to and during sampling at sea. Materials are summarized in the second section. Several protocols follow, detailing proper sample processing onboard with a variety of subsequent analyses in mind, appropriate means to facilitate sample shipment depending on constraints, and specific protocols for symbiont visualization and relative symbiont quantification in animal tissue, based on fluorescence in situ hybridization.

2

Study Design and Strategy Studying symbiosis can be summarized into a few objectives. The first is to identify the partners involved in symbiosis. This usually relies on gene- or (meta-) genome-sequencing approaches. The second is to localize symbionts in host tissues, which involves histological techniques and symbiont identification based on hybridization of specific molecular probes. The third is to understand the role of each partner in the relationship, the degree of interdependence, and their interplay in the context of their environment. This can involve various DNA- and RNA-based approaches, enzyme assays, proteomics, lipid characterization, and various more or less

Characterization of Bacterial Symbionts in Deep-Sea Fauna: Protocols. . .

quantitative methods. The fourth is to evaluate the degree of hostsymbiont coupling, by documenting the life cycle stages of each partner and in particular the mechanisms of symbiont transmission. The fifth integrates this knowledge in order to address the evolution of symbiotic systems. Designing a study on deep-sea symbiont-associated metazoans is not straightforward, and the whole procedure requires a thorough consideration in advance. Most sampling events are “one shot,” with little chance to return to the same sampling site. The sampling plan, including work on the seafloor, must therefore accommodate all the work envisioned. 2.1 The Importance of Background Environmental Data

Deep-sea sciences are integrative. The symbiotic system must be considered in terms of interactions with its environment. Ideally, three main environmental aspects need to be investigated alongside the symbiotic system itself. First, the physicochemical environment must be characterized at a scale relevant to the target organisms, including – but not exclusive to – temperature, salinity, pH, oxygen content, reduced sulfur species, methane, and heavy metals. For certain compounds, direct in situ measurements are possible using sampler- or submersible-attached probes or analyzers [13, 14]. However, in other cases, samples of the habitat under examination, such as seawater or sediment, must be collected and characterized later. The second aspect is the characterization of potential freeliving forms of symbionts or closely related bacteria in the environment. This requires sampling habitats (e.g., seawater, reduced fluids, authigenic carbonates, biofilms, sediment, decaying substrates) and processing them for the same analyses intended for the partner organisms. The final aspect of environmental characterization deals with potential sources of locally available food, both for the metazoan hosts and their symbionts. This involves evaluating the composition and stable isotope ratios of potential sources, including dissolved and particulate organic matter, which require samples to be frozen as soon as possible upon recovery [15]. In concert, these data yield an informative contextual overview for the development of symbiotic systems and provide the necessary baseline and background data for the interpretation of ensuing results. Due to various constraints, many studies lack some or all of these environmental data.

2.2 The Issue of Replication and Optimal Sample Use

Several recent symbiosis studies have pointed toward significant intraspecific variability in the composition, abundances, and roles of symbionts at various scales [8, 16, 17]. Such variability, intrinsic to highly dynamic environments and subject to natural selection, reflects the adaptability and resilience of symbiotic organisms to their environment [18, 19]. As a critical ecological feature, it not only requires investigation but also emphasizes the need for replicate specimens. Replicates are also mandatory to gain minimal


statistical support for the patterns observed. These may be relatively easy to collect from large host aggregations (e.g., dense mussel beds), but may prove more challenging to obtain from low-density or infaunal species associated with certain seeps or with wood and whale falls, for example. When several complimentary approaches are to be applied on a limited number of samples, the best approach is to perform the whole suite of methods upon each specimen, having split individual tissues appropriately (see protocols below). In this way, interindividual replication is retained, and specific aspects of the host-symbiont association can be collated and compared directly using various methods. 2.3 The Impact of Recovery Methods

3 3.1

Deep-sea organisms suffer tremendous pressure loss during recovery, corresponding to 1 bar per 10 m during their ascent. This has considerable impact on their physiology, illustrated by various groups from which specimens collected deeper than ~1,500 m commonly arrive at the surface dying or dead. Pressure vessels which can restore, or ideally maintain, at-depth pressures have been developed, allowing ongoing experiments on live specimens once aboard [20–22]; recently, pressure-maintaining devices have been designed which permit both isobaric sampling and transfer into larger aquaria which permit manipulation [23]. When not available, as a minimum, the installation of specimens in insulated hermetic boxes at the seafloor will minimize temperature variations during recovery and maintain sample integrity by preventing flushing. Early developmental stages such as larvae and postlarvae are much more difficult to sample because of their small size, often lower densities of occurrence, and a general lack of knowledge regarding their environmental distribution. However, the recovery of pre-deployed colonization devices and sediment traps or actively deployed water pumps can be effective means for their collection [24–26].

Materials Sample Fixation

1. Oven is set at 60 C for sample drying; a hybridization oven can be used (see FISH protocol). 2. Caliper. 3. Petri dishes or dissection bowls. 4. Sterile tweezers and scalpels. 5. Cryotubes for storage in liquid nitrogen. 6. Parafilm. 7. Ethanol. 8. Formaldehyde 37%, histology grade, for example, SigmaAldrich cat: F1635.


9. Glutaraldehyde, electron microscopy grade, available at SigmaAldrich cat: G7526. 10. Seawater (SFS) filtered on a 0.22 μm filter. Syringe filtering is possible for low amounts; otherwise, use filtering units or disposable filters. 11. Sodium azide (NaN3) available at Sigma-Aldrich cat: S2002. Due to acute toxicity, this needs to be pre-weighted before a cruise: 0.13 g can be stored in an Eppendorf tube and will be added to 50 mL SFS. 12. Osmium tetroxide (OsO4) available at Sigma-Aldrich cat: 201030. 13. Uranyl acetate available at Electron Microscopy Sciences cat: 22400. This compound is radioactive and toxic and requires the use of appropriate procedures. 14. RNA-preserving buffer such as RNAlater® Stabilization solution, Ambion, cat: AM7021 (500 mL). 3.2 Fluorescence In Situ Hybridization

1. Confocal or epifluorescence microscope equipped with motorized stage for 3D acquisitions. 2. Histology molds, for example Peel-a-Way® disposable embedding molds available in different sizes. 3. Microtome. 4. Heating bench for warming slides. 5. Sealed slide-staining racks. 6. Hydrophobic PAP pen, available at Sigma-Aldrich cat: Z377821. 7. Hybridization oven. 8. SuperFrost Plus adhesive glass slides, available from Euromedex cat:71869. 9. Paraffin wax, available with different melting temperatures, an example being Sigma-Aldrich cat: 327212 with a Tmelting between 58 and 62 C. Wax can be removed using HistoClear available from Agar Scientific cat:AGR1345. For safety reasons, the use of xylene is not recommended. 10. Steedman’s wax, composed of polyethylene glycol distearate (Sigma-Aldrich cat: 305413) and 1-hexadecanol (Sigma-Aldrich cat: 258741) in a 9:1 weight ratio. Melting temperature is 37 C, but higher temperatures can be used when preparing the wax. Wax is soluble in pure ethanol. 11. Acryl resin, available at Sigma-Aldrich LR white resin cat: 62661. It requires the use of benzoyl peroxide as an initiator for polymerization cat: B5907. Wax is hydrophilic and does not need to be removed prior to hybridization.


Table 1 Composition of hybridization buffer depending on the formamide concentration employed. Concentrations of stock solutions are displayed, and volumes are given in mL for a final volume of ~6 mL, to be used for wetting tissue in hybridization chambers and for mixing with probes (see Sect. 4.3.2 steps 2 and 5) Formamide used

10%

20%

30%

40%

50%

60%

NaCl (5 M)

1.08

1.08

1.08

1.08

1.08

1.08

Tris-HCl (1 M)

0.12

0.12

0.12

0.12

0.12

0.12

Milli-Q

4.2

3.6

3

2.4

1.8

1.2

SDS (20%)

0.003

0.003

0.003

0.003

0.003

0.003

Formamide

0.6

1.2

1.8

2.4

3

3.6

Table 2 Composition of washing buffer depending on the formamide concentration employed during hybridization. Concentrations of stock solutions are displayed, and volumes are given in mL for a final volume of 100 mL, to be split between washing buffer 1 and 2 (see Sect. 4.3.2 step 7) Formamide used

10%

20%

30%

40%

50%

60%

NaCl (5 M)

9

4.3

2.04

0.92

0.36

0.08

Tris-HCl (1 M)

2

2

2

2

2

2

EDTA (0.5 M)

0

1

1

1

1

1

SDS (20%)

0.05

0.05

0.05

0.05

0.05

0.05

Milli-Q

89

93

95

96

96.5

97

12. Hybridization buffer containing 900 mM NaCl, 20 mM TrisHCl, 0.01% SDS, and 10–60% vol. formamide depending on the probe(s) used (Table 1). Prepare 6 mL for each hybridization chamber to be used (a chamber may contain more than a single slide). Unused buffer can be frozen. 13. Hybridization chambers can be 50 mL Falcon tubes or sealed slide-staining racks containing stuffed tissues wetted with 4 mL of hybridization buffer. They can be used for several hybridizations if still wet. 14. The washing buffer composition depends on formamide concentration used for hybridization. Table 2 indicates how to prepare 100 mL, to be split between the two consecutive washing steps 1 and 2 (see Sect. 4.3.2 step 7). Washing buffer can be stored at room temperature (RT) and be used for several slides. 15. Anti-fade mounting medium, for example, SlowFade with DAPI, available at Life Technologies cat: S36938. It can be


ordered with or without DAPI for counterstaining and is best stored frozen. 16. Custom DNA probes labeled with various fluorochromes in their 50 end can be ordered from various companies such as Eurogentec. Most commonly used fluorochromes are fluorescein isothiocyanate (FITC, green fluorescence) and cyanine dyes including Cy3 (orange fluorescence) and Cy5 (far-red fluorescence to be detected by cameras), but many more are available. Probes can be resuspended and diluted in sterile water. Prepare aliquots with final concentration of 50 ng/μL. Stocks and aliquots are stored frozen, and probes are light sensitive. 17. Install the SymbiontJ plugin into ImageJ by copying the source file to the “Plugins” folder of ImageJ (http://www.snv.jussieu. fr/~wboudier/softs/symbiontj.html).

4

Methods

4.1 Sample Processing: How to Make the Most of the Few Specimens Available?

To optimize sample use, the best option is to split each individual specimen in tissue subsamples for the various approaches that are planned. Dissection needs to be performed quickly after recovery, in a cold room and using sterile tweezers and scalpels, in large Petri dishes or ethanol-cleaned dissection bowls. Whenever size allows, specimens can be split along planes of symmetry. Figure 1 illustrates an example of how a bivalve mussel specimen can be dissected and split for different types of analyses. This allows the direct correlation of results from different analyses within each specimen so that genuine within-host processes can be elucidated (rather than inferred from multiple specimens) and ensures that data originates from a single species allowing intraspecific, interindividual variability to be described (Note 1). Photo-documentation of specimens under the dissecting microscope is important, and any morphometric measurements that are needed can be obtained at this stage (e.g., size parameters).

4.2 Sample Fixation Procedures

Sample fixation has to be performed as quickly as possible. For this, tubes should be clearly labeled and filled with appropriate fixatives before samples arrive onboard. Proper fixation requires the use of fragments small and thin enough to ensure good penetration of the fixatives, typically a 1:10 sample-to-fixative volume ratio.

4.2.1 Liquid Nitrogen Fixation and Alternatives

This fixation is suitable for samples to be used in nucleic acid (DNA, RNA)-, protein-, enzymatic activity-, and stable isotope-based analyses:


Fig. 1 An example of how a specimen of Bathymodiolus aff. boomerang can be split for various analyses. Adductor muscle (M) is classically used for host DNAbased identification. Gills (G) containing the symbionts are split to be used for symbiont and host DNA and RNA studies, for fluorescence in situ hybridization (FISH) in the anterior, median, and posterior regions (Gant, Gmed, Gpost), for electron microscopy, and for stable isotope analyses. The gonad (Go) is located in the dorsal region, but can extend laterally in the mantle and ventrally on the visceral mass. It can be split and used for histology including EM and FISH identification of potential symbionts and to investigate gamete formation. The digestive gland (Dg) and associated stomach and intestines are used for molecular and FISH identification of potential bacteria and are located dorsal to the anterior pedal retractor muscles. The foot (F) tissue is devoid of symbionts and can be used as a host-only reference for gene expression or stable isotope studies. The mantle tissue (Ma) is also used for reference. The image also features the commensal annelid Branchipolynoe seepensis (Bs), common in deep-sea mussels

1. In a cold room, dissect tissue of interest using sterile tweezers/ scalpels immediately upon recovery. 2. Place tissue in a cryotube and transfer immediately to liquid nitrogen. 3-. Store in liquid nitrogen or at 80 C.


Below are alternatives that are less efficient but still relevant when liquid nitrogen is not available or in places where shipment of frozen material back to the lab is problematic: 1. An alternative for DNA is to transfer tissue to pure ethanol, one volume of tissue in 10 volumes of ethanol. Samples can then be shipped at 4 C or even at room temperature for a short period of time. The cap can be sealed with Parafilm to prevent leakage. 2. An alternative for RNA is to transfer tissue to RNAlater™, one volume of tissue for 10 volumes of RNAlater. Samples can then be shipped at 4 C or even at room temperature for a short period of time. 3. An alternative for stable isotope analyses is to dry tissue samples directly in a chamber at 60 C for 3 days. The most common stable isotope analyses to be performed are those of carbon, nitrogen, and sulfur. Beware that a significant amount of material can be necessary for proper analyses, in particular for nitrogen and sulfur when these compounds are not abundant, and that it is not possible to provide general guidelines. This may require pooling samples prior to analysis. Once desiccated in appropriate tubes for transport and sealed prior to removing from the oven, these can be transported at room temperature. 4.2.2 Glutaraldehyde Fixation for Electron Microscopy

1. Tissue can be fixed in 2% paraformaldehyde and 2.5% EMgrade glutaraldehyde (or 2.5–4% EM-grade glutaraldehyde only) in 0.1 M phosphate-buffered saline (PBS), corrected to pH 7.4 (minimum 1 h, typically 4–16 h, 1 mm3 or less of tissue for optimal penetration at RT). Fixative-to-specimen volume ratio ought to be at least 20:1. 2. Rinse and store in sodium azide solution (0.13 g pre-weighted NaN3 in 50 mL SFS). Samples can be transported to the lab at 4 C or even at RT. 3. Transfer to 1% osmium tetroxide solution in 0.1 M phosphate buffer (pH 7.4, 45 min). Osmium is highly toxic and requires the use of an appropriate fume hood. This and the following steps are thus usually performed back to the lab. 4. Wash by aspiration, 4 times for 5 min in 0.1 M PBS. 5. Eventually stain using 2% aqueous uranyl acetate (2 h, RT, in the dark to avoid precipitation). 6. Transfer 3 times for 5 min each in increasing ethanol (30%, 50%, 75%, and 95%) in distilled water. 7. Store at 4 C or embed right away.

Se´bastien Duperron 4.2.3 Tissue Fixation Using Formaldehyde

This procedure is employed to fix tissue samples for fluorescence in situ hybridization but can also be used for other histology staining techniques such as hematoxylin-eosin staining: 1. Dissect tissue of interest using sterile tweezers and scalpels immediately upon recovery, in a cold room. Volume of dissected tissue should not exceed 10% of the volume of fixative to be used. 2. Add 2–4% formaldehyde in sterile-filtered seawater (SFS can be syringe-filtered using a 0.22 μm filter). If working with hardshelled specimens, fixative penetration is markedly improved by preventing their closure (sever contracting musculature, or at least crack the shell, if dissection is impractical). Leave in the fridge (4 C) for 2–4 h, depending on volume of tissue (Note 2). 3. Remove fixative, rinse twice in SFS, and mix by inversion. 4. Dehydrate tissue in increasing ethanol series (50%, 70%, 80%, 20 min each). It is best to use SFS for diluting ethanol, at least on the first two steps. 5. Tissue samples can be stored in 80% ethanol at 4 C (Note 2) and can tolerate exposure to room temperature during transport if needed. Do not store at 80 C or below.

4.2.4 Conditioning and Shipment

Sample shipment to the lab is often a major (and risky) issue when working at sea. Nitrogen-fixed samples need to be transported on dry ice or in liquid nitrogen containers, shipment modes that are simply not available in some regions of the world. Transporters such as World Courier offer continuous monitoring of temperature and refill of nitrogen or dry ice but this comes at a cost. Liquid nitrogen is considered a hazardous substance and air shipment must fulfill stringent IATA guidelines, including the use of dry shippers. For this reason, many people prefer leaving frozen samples onboard until the ship stops at a readily accessible port from which samples can be transported easily. Given the risks associated with shipment of frozen material, it is good to duplicate samples and ship half under the best conditions, in dry ice for example, while the other half travels in an alternative fixative (see Sect. 4.2.1) at room temperature by courier. Note that various kits are also available for DNA or RNA extractions that require minimal material, usually a microcentrifuge. When possible, extraction onboard is a good approach, as it prevents the action of degrading enzymes during transport. Many countries, not only the final destination but also any country through which the samples transit, apply strict rules regulating the transport of animal samples. This often requires completing appropriate forms before shipment. Whatever the option chosen, sample shipment needs to be planned well in advance and discussed with local authorities and port agents.


4.3 Fluorescence In Situ Hybridization (FISH) on Sections of Animal Tissue 4.3.1 Tissue Embedding

Several types of wax can be used for tissue embedding. The choice depends on the size of tissue samples and the thickness of sections to be cut. Three classic examples are Steedman’s polyester wax, laboratory-grade paraffin wax, and, if semi-thin sectioning is needed, a hydrophilic methacrylate resin, such as LR white. Steedman’s wax melts around 37 C and dissolves in ethanol. It is appropriate for thicker sections (7–12 μm) and yields bright FISH signals, but usually the structures are less well preserved than with paraffin or methacrylate resin. Paraffin usually melts at Color - > Split channels.” Otherwise, you may work with images corresponding to each channel. 2. Select the area of interest using the “freehand selection” tool. 3. Select “Tool- > ROI Manager” in the menu “Analyze” and add object. 4. Crop the area of interest using the “Crop” function of the “Image” menu and then “Clear outside” in the “Edit” menu. This procedure may be applied to all images within a stack. If needed, set the background to black using “Edit - > Options - > color.” 5. Binarize images using “Image - > Adjust - > Threshold” either manually or automatically. This can be applied to a single or all images within a stack.


6. Numbers of black (value ¼ 0) and white (value ¼ 255) pixels/voxels can be obtained using the tool “Analyze - > Histogram - > List.” The output file can be saved. 7. When using several probes, results from individual channels can be compared to measure relative abundances. An automated procedure for counting dual symbionts, initially designed for methane- and sulfur-oxidizing symbionts of hydrothermal vent mussels, is also available as the ImageJ plugin SymbiontJ [16]. It automatically identifies bacterial signals, identifies thresholds for the channels, and computes the percentage of bacterial areas or volumes occupied by each of the two distinct symbionts: 1. You can directly use the RGB image without splitting channels. In this case, the red color will be used as a mask (anything outside is not considered true signal), so make sure to set the general bacterial probe signal to the red channel. 2. After cropping the area of interest, save it into a folder as a .TIFF file. 3. Start SymbiontJ (Plugins- > SymbiontJ) and select the folder in which you have saved the image. Several images can be stored within a single folder and will each be analyzed. 4. The resulting file summarizes the red, green, and blue areas in pixels/voxels and calculates the percentage of green and blue within the red area or volume. 5. Save the results as an Excel file.

5

Notes 1. Deep-sea metazoan hosts of bacterial symbionts often belong to poorly documented groups for which taxonomic data is scarce. Names are often inappropriate or messy, and many cryptic species are reported. Good examples are the deep-sea symbiont-bearing bathymodiolins for which most genera names actually correspond to polyphyletic groups [51]. It is thus important to back any morphology-based identification with molecular barcoding data for future comparison. 2. Over-fixation in formaldehyde or long-term storage in 96% ethanol is known to result in loss of FISH signal intensity. 3. Controls are of prime importance to confirm FISH signals. A given probe should always be tested associated to several fluorochromes. Positive (a general bacterial probe such as Eub338) and negative (a probe targeting no organism, such as the probe


Non338, antisense of Eub338) controls are mandatory. False positives that display signals even in the absence of a targeting probe are also problematic, but such signals can be identified as false by running RNAse-treated slides in parallel as further negative controls [12]. 4. A very common problem when working with bacterial symbionts within animal tissue is the low intensity of FISH signals compared to background tissue fluorescence. FISH signal intensities can be greatly improved by changing the approach to an improved version of FISH, such as CARD-FISH or DOPE-FISH [52, 53]. However, some simple steps can help optimize results of standard FISH. Regarding signal-to-noise ratios, as a rule of thumb, autofluorescence in animal tissues tends to decrease with increasing wavelength. For a given probe, signal-to-noise ratios are often improved when using Cy5 (infrared emission) compared to Cy3 (orange) and FITC (fluorescein, green). Long tissue exposure up to a few minutes to laser light may also decrease autofluorescence and result in higher signal-to-noise ratios. The use of confocal microscopes usually greatly improves the ratios. 5. Various alternative non-microscopy-based methods for the quantification of symbionts or their activity have been proposed including rRNA slot blot, qPCR using symbiont lineage-specific primer sets, and measurement of chemical compounds such as lipids produced by a partner, all with their own biases [42, 54, 55]. References 1. McFall-Ngai M, Hadfield MG, Bosch TCG et al (2013) Animals in a bacterial world, a new imperative for the life sciences. Proc Natl Acad Sci U S A 110:3229–3236 2. Tunnicliffe V, Juniper SK, Sibuet M (2003) Reducing environments of the deep-sea floor. In: Tyler PA (ed) Ecosyst. Elsevier, World Deep-Sea, pp 81–110 3. Dubilier N, Bergin C, Lott C (2008) Symbiotic diversity in marine animals: the art of harnessing chemosynthesis. Nat Rev Microbiol 6:725–740 4. Petersen JM, Dubilier N (2009) Methanotrophic symbioses in marine invertebrates. Env Microbiol Rep 1:319–335 5. Zielinski FU, Pernthaler A, Duperron S et al (2009) Widespread occurrence of an intranuclear bacterial parasite in vent and seep bathymodiolin mussels. Environ Microbiol 11:1150–1167

6. Duperron S (2010) The diversity of deep-sea mussels and their bacterial symbioses. In: Kiel S (ed) The vent and seep biota. Springer, Netherlands, pp 137–167 7. Raggi L, Schubotz F, Hinrichs K-U et al (2012) Bacterial symbionts of Bathymodiolus mussels and Escarpia tubeworms from Chapopote, an asphalt seep in the southern Gulf of Mexico. Environ Microbiol 15:1969–1987 8. Rodrigues CF, Cunha MR, Genio L, Duperron S (2013) A complex picture of associations between two host mussels and symbiotic bacteria in the Northeast Atlantic. Naturwissenschaften 100:21–31 9. Gros O, Liberge M, Heddi A et al (2003) Detection of the free-living forms of sulfideoxidizing gill endosymbionts in the lucinid habitat (Thalassia testudinum environment). Appl Environ Microbiol 69:6254–6257

Characterization of Bacterial Symbionts in Deep-Sea Fauna: Protocols. . . 10. Harmer TL, Rotjan RD, Nussbaumer AD et al (2008) Free-living tube worm endosymbionts found at deep-sea vents. Appl Environ Microbiol 74:3895–3898 11. Petersen JM, Wentrup C, Verna C et al (2012) Origins and evolutionary flexibility of chemosynthetic symbionts from Deep-Sea animals. Biol Bull 223:123–137 12. Szafranski KM, Gaudron SM, Duperron S (2014) Direct evidence for maternal inheritance of bacterial symbionts in small deep-sea clams (Bivalvia: Vesicomyidae). Naturwissenschaften 101:373–383 13. Vuillemin R, Le Roux D, Dorval P et al (2009) CHEMINI: A new in situ CHEmical MINIaturized analyzer. Deep Sea Res Part Oceanogr Res Pap 56:1391–1399 14. Wankel SD, Huang Y, Gupta M et al (2013) Characterizing the distribution of methane sources and cycling in the deep sea via in situ stable isotope analysis. Environ Sci Technol 47:1478–1486 15. Carlier A, Ritt B, Rodrigues CF et al (2010) Heterogeneous energetic pathways and carbon sources on deep eastern Mediterranean cold seep communities. Mar Biol 157:2545–2565 16. Halary S, Riou V, Gaill F et al (2008) 3D FISH for the quantification of methane- and sulphuroxidising endosymbionts in bacteriocytes of the hydrothermal vent mussel Bathymodiolus azoricus. ISME J 2:284–292 17. Lorion J, Halary S, do Nascimento J et al (2012) Evolutionary history of Idas sp. Med, (Bivalvia: Mytilidae), a cold seep mussel bearing multiple symbionts. Cah Biol Mar 53:77–87 18. Stewart FJ, Young CR, Cavanaugh CM (2008) Lateral symbiont acquisition in a maternally transmitted chemosynthetic clam endosymbiosis. Mol Biol Evol 25:673–683 19. Decker C, Olu K, Arnaud-Haond S, Duperron S (2013) Physical proximity may promote lateral acquisition of bacterial symbionts in vesicomyid clams. PLoS One 8(7):e64830 20. Shillito B, Jollivet D, Sarradin PM et al (2001) Temperature resistance of Hesiolyra bergi, a polychaetous annelid living on vent smoker walls. Mar Ecol Prog Ser 216:141–149 21. Shillito B, Gaill F, Ravaux J (2014) The ipocamp pressure incubator for deep-sea fauna. J Mar Sci Technol Taiwan 22:97–102 22. Pradillon F, Shillito B, Chervin JC et al (2004) Pressure vessels for in vivo studies of deep-sea fauna. High Press Res 24:237–246 23. Shillito B, Hamel G, Duchi C et al (2008) Live capture of megafauna from 2300 m depth,

using a newly designed Pressurized Recovery Device. Deep-Sea Res Part -Oceanogr Res Pap 55:881–889 24. Mullineaux LS, Mills SW, Sweetman AK et al (2005) Vertical, lateral and temporal structure in larval distributions at hydrothermal vents. Mar Ecol Prog Ser 293:1–16 25. Beaulieu SE, Mullineaux LS, Adams DK, Mills SW (2009) Comparison of a sediment trap and plankton pump for time-series sampling of larvae near deep-sea hydrothermal vents. Limnol Oceanogr Methods 7:235–248 26. Gaudron SM, Pradillon F, Pailleret M et al (2010) Colonization of organic substrates deployed in deep-sea reducing habitats by symbiotic species and associated fauna. Mar Env Res 70:1–12 27. Gros O, Maurin LC (2008) Easy flat embedding of oriented samples in hydrophilic resin (LR White) under controlled atmosphere: application allowing both nucleic acid hybridizations (CARD-FISH) and ultrastructural observations. Acta Histochem 110:427–431 28. Verna C, Ramette A, Wiklund H et al (2010) High symbiont diversity in the bone-eating worm Osedax mucofloris from shallow whalefalls in the North Atlantic. Environ Microbiol 12:2355–2370 29. Duperron S, De Beer D, Zbinden M et al (2009) Molecular characterization of bacteria associated with the trophosome and the tube of Lamellibrachia sp., a siboglinid annelid from cold seeps in the eastern Mediterranean. FEMS Microbiol Ecol 69:395–409 30. Zimmermann J, Lott C, Weber M et al (2014) Dual symbiosis with co-occurring sulfuroxidizing symbionts in vestimentiferan tubeworms from a Mediterranean hydrothermal vent. Environ Microbiol 16:3638–3656 31. Lo¨sekann T, Robador A, Niemann H et al (2008) Endosymbioses between bacteria and deep-sea siboglinid tubeworms from an Arctic Cold Seep (Haakon Mosby Mud Volcano, Barents Sea). Environ Microbiol 10:3237–3254 32. Nussbaumer AD, Fisher CR, Bright M (2006) Horizontal endosymbiont transmission in hydrothermal vent tubeworms. Nature 441:345–348 33. Goffredi SK, Orphan VJ, Rouse GW et al (2005) Evolutionary innovation: a bone-eating marine symbiosis. Environ Microbiol 7:1369–1378 34. Polz MF, Cavanaugh CM (1995) Unique dominance of one bacterial phylotype at a

Se´bastien Duperron Mid-Atlantic Ridge hydrothermal vent site. Proc Nat Acad Sci USA 92:7232–7236 35. Petersen JM, Ramette A, Lott C et al (2010) Dual symbiosis of the vent shrimp Rimicaris exoculata with filamentous gamma- and epsilonproteobacteria at four Mid-Atlantic Ridge hydrothermal vent fields. Environ Microbiol 12:2204–2218 36. Duperron S, Pottier M-A, Leger N et al (2013) A tale of two chitons: is habitat specialisation linked to distinct associated bacterial communities? FEMS Microbiol Ecol 83:552–567 37. Rodrigues CF, Duperron S (2011) Distinct symbiont lineages in three thyasirid species (Bivalvia: Thyasiridae) form the eastern Atlantic and Mediterranean Sea. Naturwissenschaften 98:281–287 38. Distel DL, Beaudoin DJ, Morrill W (2002) Coexistence of multiple proteobacterial endosymbionts in the gills of the wood-boring bivalve Lyrodus pedicellatus (Bivalvia: Teredinidae). Appl Environ Microbiol 2002:6292–6299 39. Wentrup C, Wendeberg A, Schimak M et al (2014) Forever competent: deep-sea bivalves are colonized by their chemosynthetic symbionts throughout their lifetime. Environ Microbiol 16:3699–3713 40. Duperron S, Rodrigues CF, Leger N et al (2012) Diversity of symbioses between chemosynthetic bacteria and metazoans at the Guiness cold seep site (Gulf of Guinea, West Africa). MicrobiologyOpen 1:467–480 41. Duperron S, Nadalig T, Caprais JC et al (2005) Dual symbiosis in a Bathymodiolus mussel from a methane seep on the Gabon continental margin (South East Atlantic): 16S rRNA phylogeny and distribution of the symbionts in the gills. Appl Environ Microbiol 71:1694–1700 42. Duperron S, Sibuet M, MacGregor BJ et al (2007) Diversity, relative abundance, and metabolic potential of bacterial endosymbionts in three Bathymodiolus mussels (Bivalvia: Mytilidae) from cold seeps in the Gulf of Mexico. Env Microbiol 9:1423–1438 43. Distel DL, Cavanaugh CM (1994) Independent phylogenetic origins of methanotrophic and chemoautotrophic bacterial endosymbioses in marine bivalves. J Bacteriol 176:1932–1938 44. Distel DL, Lee HKW, Cavanaugh CM (1995) Intracellular coexistence of methano- and thioautotrophic bacteria in a hydrothermal vent mussel. Proc Natl Acad Sci USA 92:9598–9602

45. Duperron S, Bergin C, Zielinski F et al (2006) A dual symbiosis shared by two mussel species, Bathymodiolus azoricus and B. puteoserpentis (Bivalvia: Mytilidae), from hydrothermal vents along the northern Mid-Atlantic Ridge. Environ Microbiol 8:1441–1447 46. Duperron S, Halary S, Lorion J et al (2008) Unexpected co-occurrence of 6 bacterial symbionts in the gill of the cold seep mussel Idas sp. (Bivalvia: Mytilidae). Environ Microbiol 10:433–445 47. Urakawa H, Dubilier N, Fujiwara Y et al (2005) Hydrothermal vent gastropods from the same family (Provannidae) harbour ε- and γ-proteobacterial endosymbionts. Environ Microbiol 7:750–754 48. Bates AE, Harmer TL, Roeselers G, Cavanaugh CM (2011) Phylogenetic characterization of episymbiotic bacteria hosted by a hydrothermal vent limpet (Lepetodrilidae, Vetigastropoda). Biol Bull 220:118–127 49. Cavanaugh CM, Levering PR, Maki JS et al (1987) Symbiosis of methylotrophic bacteria and deep-sea mussels. Nature 325:346–347 50. Fiala-Me´dioni A, McKiness ZP, Dando P et al (2002) Ultrastructural, biochemical and immunological characterisation of two populations of the Mytilid mussel Bathymodiolus azoricus from the Mid Atlantic Ridge: evidence for a dual symbiosis. Mar Biol 141:1035–1043 51. Lorion J, Buge B, Cruaud C, Samadi S (2010) New insights into diversity and evolution of deep-sea Mytilidae (Mollusca: Bivalvia). Mol Phyl Evol 57:71–83 52. Pernthaler A, Pernthaler J, Amann R (2002) Fluorescence in situ hybridization and catalysed reporter deposition for the identification of marine Bacteria. Appl Environ Microbiol 68:3094–3101 53. Stoecker K, Dorninger C, Daims H, Wagner M (2010) Double labeling of oligonucleotide probes for fluorescence in situ hybridization (DOPE-FISH) improves signal intensity and increases rRNA accessibility. Appl Environ Microbiol 76:922–926 54. Pond DW, Bell MV, Dixon DR et al (1998) Stable-carbon-isotope composition of fatty acids in hydrothermal vent mussels containing methanotrophic and thiotrophic bacterial endosymbionts. Appl Environ Microbiol 64:370–375 55. Guezi H, Boutet I, Andersen AC et al (2014) Comparative analysis of symbiont ratios and gene expression in natural populations of two Bathymodiolus mussel species. Symbiosis 63:19–29