Present addresses: Anniet M. Laverman, Université P&M Curie Paris VI, Boite 123, 4 place Jussieu, 75252 Paris Cedex 05–France. Larry J. Forney, Department of Biological Sciences, University of Idaho, Moscow, ID 83844-3051, USA
Comparison of deep-sea sediment microbial communities in the Eastern Mediterranean
Article first published online: 15 APR 2008
Journal compilation © 2008 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd. No claim to original Dutch government works
FEMS Microbiology Ecology
Volume 64, Issue 3, pages 362–377, June 2008
How to Cite
Heijs, S. K., Laverman, A. M., Forney, L. J., Hardoim, P. R. and Van Elsas, J. D. (2008), Comparison of deep-sea sediment microbial communities in the Eastern Mediterranean. FEMS Microbiology Ecology, 64: 362–377. doi: 10.1111/j.1574-6941.2008.00463.x
Editor: Patricia Sobecky
- Issue published online: 15 APR 2008
- Article first published online: 15 APR 2008
- Received 31 May 2007; revised 23 December 2007; accepted 11 January 2008.First published online 15 April 2008.
- microbial community structure;
- molecular analysis
Bacterial and archaeal communities in sediments obtained from three geographically-distant mud volcanoes, a control site and a microbial mat in the Eastern Mediterranean deep-sea were characterized using direct 16S rRNA gene analyses. The data were thus in relation to the chemical characteristics of the (stratified) habitats to infer community structure–habitat relationships. The bacterial sequences in the different habitats were related to those of Actinobacteria, Bacilli, Chloroflexi, Alpha-, Beta-, Gamma-, Delta- and Epsilonproteobacteria and unclassified bacteria, including the JS1 group. The archaeal sequences found were affiliated with those of the Methanosarcinales, Thermoplasmales, Halobacteriales and Crenarchaea belonging to marine benthic group I and B, as well as MCG group archaea. In each sample, the communities were diverse and unique at the phylotype level. However, at higher taxonomic levels, similar groups were found in different sediments, and similar depth layers tended to contain similar communities. The sequences that dominated in all top layers (as well as in the mat) probably represented organisms involved in aerobic heterotrophy, sulfide-based chemoautotrophy and methanotrophy and/or methylotrophy. Sequences of organisms most likely involved in anaerobic methane oxidation, sulfate reduction and anaerobic heterotrophy were predominantly found in deeper layers. The data supported the notion of (1) uniqueness of each habitat at fine taxonomic levels, (2) stratification in depth and (3) conservation of function in the sediments.
In the past three decades, mud volcanoes, probably resulting from tectonic activity, have been discovered along the Eastern Mediterranean Ridge, which crosses the Eastern Mediterranean Sea (Limonov et al., 1996; Cronin et al., 1997; Woodside et al., 1998). Two areas, the Olimpi mud volcano area and the Anaximander Mountains area, have been extensively sampled and examined in two Dutch–French expeditions, MEDINAUT and MEDINETH, in 1998 and 1999. These expeditions revealed the occurrence of so-called cold seeps (MEDINAUT, MEDINETH Shipboard Scientific Parties et al., 2000), indicating the existence of chemotrophy-based microbial communities similar to those previously found in hydrothermal vent environments (Barry et al., 1996; Olu et al., 1996; Sibuet & Olu, 1998). Such communities, in the presence of suitable electron acceptors, may derive their energy mainly from the oxidation of methane and/or reduced sulfur compounds such as sulfides or even elemental sulfur (Gaill, 1993; Teske et al., 2002). The existence and nature of microbial communities in the seafloor habitats can be revealing with respect to our view of the functioning of the ecosystem Earth in ancient times, i.e. before and/or during the presumed transition from an atmosphere poor in molecular oxygen to one containing oxygen. In particular, the structure, diversity and functioning of these microbial communities represent important study objects.
Sulfide may be a key compound that is at the basis of life in the cold-seep environments at the Eastern Mediterranean mud volcanoes, and sulfide oxidation has been shown to take place, as evidenced from in-depth analyses of this process and organisms involved as revealed by lipid biomarkers (Peter & Shanks, 1992; Mizota & Maki, 1998). The process of sulfide oxidation is not confined to free-living organisms, as sulfide-oxidizing organisms have also been found as endosymbionts in the higher organisms present in these environments (Barry et al., 1996; Olu et al., 1996; Sibuet & Olu, 1998). On the other hand, methane can also be a key compound (electron donor and carbon source) supporting microbial life in the cold seeps (Pancost et al., 2000; Aloisi et al., 2002). The microbial lipid biomarkers in these environments were found to be strongly depleted in 13C-carbon, thus providing evidence for the occurrence of anaerobic methane oxidation (Pancost et al., 2000; Aloisi et al., 2002). In line with this contention was the finding of archaeal sequences that resembled those of organisms known to be involved in this process (Hinrichs et al., 1999; Boetius et al., 2000; Orphan et al., 2001, 2002; Aloisi et al., 2002; Michaelis et al., 2002; Heijs et al., 2005). In some studies, sulfate-reducing bacteria (SRB) were found to occur together with the methane oxidizers (Heijs et al., 2005, 2007), which is consistent with the hypothesis that anaerobic methane oxidation is often actually mechanistically coupled to sulfate reduction (Hoehler et al., 1994; Hansen et al., 1998; Boetius et al., 2000; Michaelis et al., 2002).
Key ecosystem functions are likely to be better sustained under fluctuating environmental conditions if they are sufficiently redundant in a microbial community (Wilson & Botkin, 1990; Yachi & Loreau, 1999). Hence, in environments with fluctuating conditions, microbial diversity and community structure are crucial parameters that may relate to the expected stability of the local microbial processes. It is therefore important to provide community-level data next to data on local chemistries. The two datasets together can provide support for the notion of conservation of function of such systems. In addition, an assessment of a possible effect of site (geography) will shed light on the biogeographical aspects of the communities under study. Thus, the microbial communities in geographically separated sites, which are subjected to key chemical conditions that may be grossly similar between sites, may still be under slightly different selective pressures per site. Moreover, stochastic events of early colonization, migration and extinction may have differed. Hence different sites may encompass organisms that belong to similar ecophysiological groups as determined by the key processes they are involved in, yet are of different types as determined by their behavior in the light of other local conditions (Ward et al., 2006).
The objective of the current study was to determine the structures of the microbial communities as well as the chemistries of geographically separated mud volcanoes in the Eastern Mediterranean Sea in order to cross-compare these habitats with respect to microbial community structure and potential function. Next to the between-site comparisons, we were particularly interested in the stratification in the habitats under study.
Materials and methods
Sampling sites and sampling
Box cores of 50 cm diameter (resulting in sediment cores with an average depth of 35 cm) were taken at water depths between 1673 and 3342 m from a site near Urania brine lake (used as a control) and mud volcanoes in the Olimpi (O; containing two mud volcanoes, Napoli and Milano) and Anaximander Mountains (AM; containing mud volcanoes Kazan and Amsterdam) areas. This thus included four distinct locations, during the MEDINETH cruise in September 1999. From the centers of the box cores, subcores were taken with sterilized rectangular aluminum corers (length: 50 cm; width: 20 cm; depth: 10 cm). The subcores were divided into depth layers based on the sediment colors, which ranged from brown (oxidized) layers to black/dark-gray (strongly reduced) layers. Hence, we used, as the criterion in our sampling, the presumed prevailing presence or absence of molecular oxygen. Subsamples at depth intervals of c. 2 cm were thus aseptically taken from the sediment layers using cut-off sterile 1-mL syringes. For each layer, eight subsamples were pooled. The characteristics of all samples are shown in Table 1. In addition, microbial mat samples were collected from the Milano mud volcano during the MEDINAUT cruise in November/December 1998 (see Fig. 1). These samples were collected with the submersible Nautile using a titanium vacuum bottle. Sediment and microbial mat samples were placed in 12-mL sterile Greiner tubes, sealed and immediately frozen at −80 °C pending further analysis.
|Site||Site description||Sample name||Depth- layer (cm)||Water depth (m)||Position (N–E)||Genbank Accession No.|
|Urania||Near Urania brine lake, control site||Urania-1||0–10||3342||35°14.7′–21°28.9′||AY627426–AY627625|
|Napoli||Dome of Napoli mud volcano, a previously active cold seep.||Napoli-1||0–8||1910||33°43.6′–24°41.2′||AY592434–AY592798|
|Kazan||Kazan mud volcano, near active cold seep||Kazan-1*||0–6||1673||35°25.9′–30°33.7′||AY591932–AY592229|
|Amsterdam||Amsterdam mud volcano, nearby active seep||Amsterdam-1||0–14||1995||35°20.0′–30°16.0′||AY592230–AY592433|
|Milano||Milano mud volcano, microbial mat||Milano-1||5 μm filter†||1958||33°44.0′–24°46.7′||AY592799–AY592933|
|Milano-2||0.2 μm filter†|
The samples analyzed were from different depths [numbered 1 (top) through 3 or 4 (bottom)] of sediment cores from three cold seeps as well as one control site, as follows (Table 1):
- 1Napoli: The dome of the Napoli mud volcano. Based on the sulfidic smell of the sediment, the presence of a distinct layer with carbonate nodules, and the finding of elemental sulfur in the upper 20 cm of the core, this sample likely represented a cold seep (R. Haesse, pers. commun.). Four sediment depth layers, Napoli-1, -2, -3 and -4 (0–8, 8–18, 18–27 and 27–29 cm deep), were analyzed.
- 2Kazan: The top of the active (Werne et al., 2004; Heijs et al., 2005) Kazan mud volcano. Three depth layers were used, i.e. Kazan-1, -2 and -3 (respectively 0–6, 6–22 and 22–34 cm deep). Upon retrieval, this core showed strong degassing and had a strong sulfidic smell.
- 3Amsterdam: The top of the Amsterdam mud volcano dome. Two depth layers were used, i.e. Amsterdam-1 and -2 (respectively 0–14 and 14–31 cm deep). This core also showed degassing and a strong sulfuric smell (Table 1).
- 4Urania (control): A mound just outside Urania brine lake. This site was not an active cold seep and was therefore chosen as the control site. Two depths, Urania-1 and -2 (0–10 and 10–20 cm), were analyzed.
In addition, for reasons of comparison we analyzed two filter fractions, i.e. 5 μm (Milano-1) and 0.2 μm (Milano-2) of a microbial mat at the surface of the cold seep at the Milano mud volcano top, hereafter called Milano mat.
Pooled samples from each depth layer of the various sediments were used for DNA extraction. Each sample was thawed on ice and manually homogenized with a sterile spatula, after which c. 5 g (wet weight) of sediment was divided over 2-mL tubes, containing 0.5 g of a 1 : 1 mixture of zirconium and glass beads (Ø 0.1 mm).
In addition, filamentous mat material was thawed and a 20 mL portion (including adhering seawater) was sonicated three times at 0 °C for 3 min at 100 W in a sonicating water bath (Branson B1540, 100 W). Subsequently, the mixture was filtered through a 5 μm pore size filter and then over a 0.2 μm pore size filter to distinguish between particle- (or filament-) associated and free-living microorganisms in accordance with Crump et al. (1999). DNA was then extracted by a combination of enzymatic and mechanical treatments as previously described (Heijs et al., 2005, 2007).
Amplification of 16S rRNA gene regions, cloning and sequencing
The 16S rRNA gene regions were amplified using specific primers for bacterial and archaeal 16S rRNA genes. Bacterial 16S rRNA genes were amplified using the B8F (5′-AGAGTTTGATCMTGGCTCAG-3′) forward primer (Edwards et al., 1989) and the universal U1406R (5′-ACGGGCGGTGTGTRC-3′) reverse primer (Lane, 1991). Archaeal 16S rRNA genes were amplified with the A2F (5′-TTCCGGTTGATCCYGCCGGA-3′) forward primer (DeLong, 1992) in combination with the universal U1406R (5′-ACGGGCGGTGTGTRC-3′) reverse primer. For the Kazan sediment, PCR was performed using the same forward primer in combination with reverse primer A958-R (5′-YCCGGCGTTGAMTCCAATT-3′) (DeLong, 1992) when the total amount of PCR product obtained with primers A2F and U1406 was low (Heijs et al., 2007). PCR mixtures (25 μL) contained 10.2 mM Tris, 2.3 mM MgCl2, 50 mM KCl, 2% dimethyl sulfoxide (DMSO), 5 μg bovine serum albumin (BSA), 0.2 mM of each dNTP, 0.2 μM of each primer and 0.5 U of Taq DNA polymerase. The 16S rRNA genes were amplified in a Perkin-Elmer GeneAmp PCR System 9700 (Perkin-Elmer Applied Biosystems, Nieuwerkerk a/d IJsel, the Netherlands) using the following program: 95 °C for 5 min; 35 cycles of 94 °C for 1 min, 57.5 °C for 30 s, 72 °C for 4 min, with a final elongation step of 72 °C for 7 min.
PCR products were purified using QIAquick spin columns (Invitrogen BV, Groningen, the Netherlands) and subsequently cloned into the pGEM-T Easy vector system (Promega Benelux B.V., Leiden, the Netherlands) using Escherichia coli JM109 as the host. Inserts were amplified by colony PCR, using the pGEM-T specific primers T7 (5′-TAATACGACTCACTATAGGG-3′) and SP6 (5′-GATTTAGGTGACACTATAG-3′). PCR mixtures were as described above with the following PCR conditions: 94 °C for 5 min; 30 cycles of 94 °C for 1 min, 48 °C for 30 s, 72 °C for 4 min, with a final elongation step of 72 °C for 7 min. All clones with correct inserts from each 16S rRNA gene library were selected and partial 16S rRNA gene sequences were determined using Big Dye version-3 cycle sequencing reactions (Applied Biosystems, Foster City, CA) and an ABI3100 PRISM Genetic Analyzer (Applied Biosystems). Sequencing reactions used B8F, T7 or U515 (5′-GTGCCAGCMGCCGCGG-3′) forward primers for bacteria, and A2F, T7 or U515 forward primers for archaea. Partial sequences were manually edited in Chromas 1.45 (http://www.technelysium.com.au) and contig assemblies were done in bioedit (http://www.mbio.ncsu.edu/BioEdit/bioedit.html) according to Huang (1992), yielding a majority of sequences of ≥1000 base pairs.
Chimeric sequences were detected using the Check_Chimera tool available in the Ribosomal Database Project (http://rdp.cme.msu.edu/html/) and omitted from the dataset. Sequences were aligned using the arb software package fast aligner utility (Ludwig et al., 1998). Alignments were checked manually, using the secondary structure of the 16S rRNA gene molecule. Evolutionary distances of full-length sequences were calculated according to the Kimura two-parameter correction method (Kimura, 1980) after which neighbor-joining trees were constructed with 1000 bootstrap samplings using TreeconW (Van de Peer & De Wachter, 1994). Sequences with >97% similarity were considered to belong to the same phylotype. Hence, phylotype is here defined as a level of grouping that encompasses all sequences of at least 97% similarity to each other. Related 16S rRNA gene sequences were placed within taxa (between phylum and order) by determining the taxonomic class (using the NCBI taxonomy database) of the nearest GenBank relative of sequences that formed a phylogenetic group (class). Sequences that showed no relatedness with a known bacterial or archaeal phylogenetic group in the NCBI taxonomy database were listed as unclassified. Nearest relatives were obtained from the Genbank database using the basic local alignment search tool (blast) at the NCBI website (http://www.ncbi.nlm.nih.gov/). Alignments were made with the arb software package as mentioned before. Evolutionary distances were calculated for alignments of the comparable region of the 16S rRNA gene containing at least 800 nucleotides, as mentioned above.
Shannon–Weaver indices of diversity and CHAO1 and ACE estimators of richness were calculated for all samples on the basis of the phylotype distribution (97% similarity) using the dotur program (http://www.plantpath.wisc.edu/fac/joh/dotur.html). Similarity between the microbial communities in the selected mud volcanoes was determined using the Morisita–Horn index of similarity (Wolda, 1981) at both phylotype and phylogenetic group level. This similarity index is not affected by differences in sample size (Wolda, 1981). The Morisita–Horn indices of similarity at the group level were further analyzed by cluster analyses using past. The 16S rRNA gene sequences determined in this study were deposited in the Genbank sequence library (accession numbers in Table 1).
Sediment pore water chemistry
Before analysis, subsamples from the different sediment depths were thawed and manually mixed. Water content and sediment density were determined from the loss of weight of 2 mL of sediment after extensive drying (60 °C, 72 h). The sediment subsamples were centrifuged at 0 °C at 16.000 g for 10 min. The supernatant was then collected with a 50-mL syringe and filtered over a 0.2 μm membrane filter into 10-mL Greiner tubes, and the filtrate was stored on ice. For the determination of levels of pore-water total organic carbon (TOC), , and , , Br− and Cl−, subsamples of pore water were stored at −20 °C. Aliquots of 2 mL for sulfide measurements were treated with 10 μL of ultrapure 1 M NaOH mL−1. Aliquots of 2 mL for measurements were treated with 10 μL ultrapure 1 M HCl mL−1 and stored at 4 °C. The pelleted material from each sediment layer (10–15 g) was oven dried (60 °C, 36 h) for subsequent analyses. The , , and concentrations were determined colorimetrically on a Nutrient Autoanalyzer-3 (Bran and Luebbe, De Meern, the Netherlands). , Br− and Cl− concentrations were determined by ion chromatography (Dionex DX-120 IC Waters, Milford, MA). TOC in the pore water was measured on a TOC analyzer (Shimadzu TOC-5050A, Shimadzu Benelux, Den Bosch, the Netherlands). Sediment total carbon and nitrogen were determined on dried sediment using an elemental analyzer (Carlo Erba NA1500, Milan, Italy). Pore-water chemistry measurements were carried out in duplicate, whereas sediment chemistry was performed on single samples.
Sediment parameters and chemistry of pore water
For all sediments, the average water contents and sediment densities indicated a porous, clay-like structure. Some striking features (Table 2, discussed below) characterize the sites as follows (detailed below):
|Parameter||Urania control sediment||Napoli mud volcano||Kazan mud volcano||Amsterdam mud volcano|
|1 0–10 cm||2 10–20 cm||1 0–8 cm||2 8–18 cm||3 18–27 cm||4 27–28 cm||1 0–6 cm||2 6–22 cm||3 22–34 cm||1 0–14 cm||2 14–31 cm|
|Water content (% v/v)||68||62||56||75||63||65||58||61||56||61||58|
|Sediment density (g cm−3)||2.83||2.49||2.49||2.35||2.66||2.76||2.37||2.50||2.30||2.58||2.56|
|Nitrogen content (%)||0.04||0.03||0.04||0.08||0.05||0.06||0.05||0.05||0.04||0.06||0.04|
|Total carbon content (%)||5.14||5.38||3.63||1.22||3.95||2.94||2.59||2.15||1.74||3.90||4.31|
|TOC (mg C L−1)||82.8||118.8||112.4||46.0||46.4||28.6||33.4||32.9||30.3||55.4||130.0|
|Ammonia (NH4) (μM)||18.4||26.8||40.6||80.9||115.7||217.2||30.0||42.7||158.5||152.5||50.6|
|Nitrite () (μM)||0.5||0.2||0.2||0.3||0.2||0.2||1.0||0.2||0.1||0.8||0.3|
|Nitrate () (μM)||10.3||2.2||1.7||1.2||1.8||1.6||15.7||2.0||1.1||2.2||2.3|
|Sulfate () (mM)||31.2||34.4||33.0||30.0||25.8||19.5||31.4||32.7||3.2||33.3||34.6|
|Sulfide (S2−) (mM)||B.D.||B.D.||B.D.||0.1||0.05||0.05||B.D.||6.2||2.8||N.D+||N.D++|
|Phosphate () (μM)||0.4||N.D.||0.9||2.7||6.8||8.1||0.5||0.3||0.2||0.8||0.3|
|Bromide (Br−) (μM)||922||940||1044||1081||651||458||937||921||691||944||1080|
|Chloride (Cl−) (mM)||622||624||858||1150||1742||2197||627||612||377||632||633|
- 1Urania: control site, high in carbon, evidence for nitrate reduction.
- 2Napoli: site with presumed sulfate reduction and possibly affected by gas hydrate or brine material in deeper layers.
- 3Kazan: site with strongest evidence for both nitrate and sulfate reduction.
- 4Amsterdam: high-carbon site.
Across all sediment layers, the average nitrogen contents were low (range 0.03–0.08%) and the total carbon contents high (range 1.22–5.38%). Hence, the carbon to nitrogen ratios were high in these samples, indicating that nitrogen might have been limiting throughout (Table 2). This contention was strengthened by a back-of-the-envelope comparison of molar C : N ratios to the critical value of 6.6 (Redfield ratio). In only two cases, namely Napoli-4 and Kazan-3, these values came to slightly above 10, whereas all other values were much higher. In all sediments, the pore water pH values were elevated (range 8.34–9.31). Furthermore, pore water (average) TOC levels were considerable and similar in the Napoli and Kazan sediments, and raised in the Amsterdam sediment. Ammonia constituted the largest fraction of inorganic nitrogen in the pore water of all sediments, while nitrate and nitrite levels were generally low. The ammonia levels showed clear increases with depth in Napoli and Kazan (no trend in Amsterdam), whereas the nitrite and nitrate levels decreased with depth in Kazan and the Urania control (no clear trend in Napoli and Amsterdam). Hence, the chemical data indicated, at least in Kazan, the occurrence of nitrate reduction. Furthermore, the sulfate concentrations in the top layers were comparable across all sediments (Table 2); these clearly decreased with depth in the Napoli and Kazan sediments (no trend in Amsterdam or Urania). These data indicate the occurrence of sulfate reduction in the lower layers of the Napoli and Kazan sediments (Table 2). Although sulfide measurements were obviously imprecise due to the problem of gas diffusion upon sampling, sulfide was mainly detected in the deeper sediment layers of all mud volcanoes, which corroborates the above statement.
Bromide was present in all sediments at levels ranging from 458 to 1081 μM, whereas the chloride levels ranged from 377 to 858 (average around 625) mM in most sediments. A striking example was formed by the Napoli sediments, in which the chloride levels were raised from about twofold (upper layer) to almost fourfold (deeper layer) the average levels found in all other samples. This observation hints at the occurrence of brine material or hydrates in deeper (unsampled) layers of the Napoli mud volcano. On the other hand, the decreasing chloride concentrations with depth in Kazan provide evidence for gas hydrate destabilization, which releases methane (and water) into the surrounding environment (De Lange & Brumsack, 1998).
Overall, this analysis illustrated that clear differences existed with respect to sediment chemistries between sites as well as within samples from the same site (indicating stratification).
Diversity of microbial communities
A total of 699 bacterial and 479 archaeal 16S rRNA gene sequences were determined in this study (Table 3). Overall, the bacterial sequences represented 365 unique bacterial phylotypes (defined as a group of sequences sharing >97% similarity with each other) that were classified in 25 broader clusters of higher taxonomic ranking (i.e. at the taxonomic rank of class, hereafter designated as ‘phylogenetic group’). The archaeal 16S rRNA gene survey resulted in 136 phylotypes, which accounted for six archaeal phylogenetic groups (Table 3). Rarefaction analysis was carried out for each of the bacterial and archaeal 16S rRNA clone libraries of the sediment and microbial mat samples analyzed. This statistical analysis yielded the expected asymptotic accumulation curves for the distribution of the sequences over the phylotypes. On average, across the samples the curves indicated fairly high coverage levels for the archaea, but generally lower coverage for the bacteria (data not shown). This analysis was corroborated by the CHAO1 and ACE indicators of richness (Table 3), which suggested that the numbers of clones obtained indeed provided, in most cases, a fair representation of the dominant phylotypes per habitat for the archaea, whereas this was more limited for the bacteria.
|Domain||Phylogenetic group/ sequence statistics||Urania||Napoli||Kazan||Amsterdam||Milano|
|1 0–10 cm||2 10–20 cm||1 0–8 cm||2 8–18 cm||3 18–27 cm||4 27–28 cm||1 0–6 cm||2^ 6–22 cm||3 22–34 cm||1 0–14 cm||2 14–31 cm||WF1 5 μm||WF2 0.2 μm|
|Archaea||Marine Group I Archaea||54 (15)||–||19 (8)||6 (4)||3 (2)||–||46 (7)||3 (2)||–||5(3)||–||–||–|
|Crenarchaea related*||–||5 (3)||–||16 (9)||–||–||–||19 (12)||2 (2)||–||–||–||–|
|Halobacteriales related||–||25 (5)||–||2 (1)||3 (2)||3 (3)||1 (1)||3 (1)||2 (2)||–||–||–||1 (1)|
|Methanosarcinales related ANME-1/ANME-2||–||–||–||–||–||1 (1)||–||16 (2)†||45 (7)†||–||8 (3)†||31 (6)†||12 (2)†|
|Thermoplasmales related||–||–||7 (6)||14 (7)||23 (9)||6 (3)||–||1 (1)||2 (1)||23 (12)||18 (8)||–||1 (1)|
|Unclassified archaea‡||–||3 (1)||6 (4)||8 (3)||8 (5)||–||–||4 (2)||–||21 (5)||3 (3)||–||–|
|Total sequences§||54 (15)||33 (9)||32 (18)||46 (24)||37 (18)||10 (7)||47 (8)||46 (20)||51 (12)||49 (20)||29 (14)||31 (6)||14 (4)|
|Shannon–Weaver diversity||2.15 (± 0.3)||2.00 (± 0.3)||2.75 (± 0.28)||2.89 (± 0.23)||2.27 (± 0.29)||1.67 (± 0.58)||1.67 (± 0.19)||2.52 (± 0.3)||1.89 (± 0.35)||2.14 (± 0.27)||2.28 (± 0.36)||1.35 (± 0.35)||0.99 (± 0.46)|
|CHAO1 Richness||28 (18–70)||10 (10–15)||32 (21–73)||24 (21–39)||15 (13–30)||22 (10–75)||7 (ND)||37 (23–94)||49 (24–147)||15 (13–27)||26 (17–68)||10 (7–30)||5 (4–17)|
|ACE Richness||34 (21–84)||11 (10–14)||31 (22–67)||27 (24–31)||19 (14–40)||39 (11–235)||8 (7–17)||36 (24–79)||66 (27–246)||20 (15–45)||37 (20–100)||17 (8–84)||7 (4–30)|
|Bacteria||Acidobacteria||4 (4)||–||1 (1)||–||–||–||5 (4)||1 (1)||–||4 (4)||1 (1)||1 (1)||–|
|Actinobacteria||3 (2)||–||21 (11)||2 (2)||–||1 (1)||14 (12)||9 (7)||–||13 (8)||6 (4)||1 (1)||–|
|Alphaproteobacteria||11 (8)||28 (2)||3 (3)||1 (1)||–||11 (2)||–||2 (2)||–||–||1 (1)||7 (6)||1 (1)|
|Bacilli||2 (2)||10 (5)||–||2 (2)||10 (6)||1 (1)||–||–||–||–||–||–|
|Betaproteobacteria||1 (1)||13 (1)||–||–||2 (1)||11 (2)||–||–||–||–||3 (2)||–||–|
|CFB-group¶||1 (1)||–||–||1 (1)||2 (1)||3 (1)||–||–||–||1 (1)||–||4 (4)||–|
|Chloroflexi||13 (8)||6 (6)||13 (10)||25 (13)||25 (13)||14 (8)||5 (5)||17 (11)||17 (3)||13 (9)||12 (8)||3 (3)||–|
|Clostridia||–||–||–||–||–||–||1 (1)||1 (1)||–||–||–||–||–|
|Deltaproteobacteria||5 (3)||1 (1)||5 (5)||4 (4)||7 (5)||2 (1)||–||3 (3)||18 (7)||9 (8)||16 (10)||5 (5)||10 (3)|
|Epsilonproteobacteria||2 (2)||–||1 (1)||–||–||–||–||–||–||1 (1)||–||3 (1)||20 (8)|
|Gammaproteobacteria||2 (1)||–||2 (2)||3 (2)||–||–||4 (3)||3 (3)||–||8 (6)||4 (4)||19 (11)||11 (7)|
|Nitrospira||2 (2)||–||–||–||–||–||2 (2)||2 (1)||–||–||–||–||–|
|Candidate division OP-11||–||3 (1)||1 (1)||1 (1)||1 (1)||1 (1)||2 (2)||–||3 (3)||–||2 (1)||–||3 (3)|
|Planctomycetacaea||1 (1)||3 (3)||16 (7)||8 (8)||11 (10)||2 (2)||4 (3)||1 (1)||2 (2)||4 (3)||7 (7)||–||–|
|Unclassified∥||1 (1)||2 (2)||4 (4)||6 (4)||13 (6)||2 (2)||9 (6)||4 (3)||4 (3)||1 (1)||17 (4)||–||–|
|Total sequences§||48 (36)||66 (21)||67 (45)||51 (36)||63 (39)||57 (26)||47 (39)||43 (33)||44 (18)||54 (41)||71 (44)||43 (32)||45 (22)|
|Shannon–Weaver diversity||3.48 (± 0.21)||2.39 (± 0.34)||3.67 (± 0.23)||3.45 (± 0.22)||3.43 (± 0.24)||3.08 (± 0.27)||3.44 (± 0.22)||3.42 (± 0.23)||2.59 (± 0.33)||3.53 (± 0.21)||3.43 (± 0.26)||3.23 (± 0.28)||3.02 (± 0.26)|
|CHAO1 Richness||80 (52–153)||82 (42–215)||204 (107–454)||82 (53–163)||75 (53–131)||61 (42–115)||66 (45–123)||87 (52–185)||44 (27–103)||84 (55–162)||86 (60–153)||381 (197–767)||64 (38–152)|
|ACE Richness||93 (57–185)||95 (46–259)||274 (139–607)||91 (57–178)||122 (75–238)||111 (64–227)||85 (56–149)||93 (55–201)||65 (34–160)||90 (57–176)||90 (72–118)||229 (88–711)||65 (40–141)|
To compare the diversities of the prokaryotic communities, Shannon–Weaver diversity indices were calculated. These indices, with values generally above 3, indicated the existence of high bacterial diversities in all samples (Table 3). The highest diversities occurred in the sediment top layers, and there was a decrease in the lower layers. In contrast, the archaeal communities showed lower diversities, with indices ranging from 0.99 to 2.95, and there was no clear correlation of archaeal diversity with chemistry (defined by depth; Table 3). The archaeal diversity was highest in the Napoli sediment, and moderate for the Kazan sediment and Urania control. Archaeal diversity was lowest in the Milano mat (Table 3).
The similarities between, on the one hand, the bacterial and, on the other, the archaeal communities in the different sediment layers and the microbial mat were determined at both the phylotype and the phylogenetic group level. The Morisita–Horn indices of similarity of both communities were generally low (<0.4) at the phylotype level (data not shown). This indicated little overlap between both the bacterial and archaeal phylotypes across these samples, and revealed that each habitat contained its own unique community of bacteria and archaea. However, at the phylogenetic group level, the Morisita–Horn indices were, on several occasions, significantly raised. Clustering of the bacterial communities using these indices (Fig. 2) showed the appearance of basically four clusters: (bacterial) cluster I – the top layers of all three mud volcanoes sampled (plus the below-top layer Kazan-2), cluster II – four of six deeper mud volcano layers, cluster III – two Urania control site samples (and one deeper mud volcano layer), cluster IV – two microbial mat samples. Hence, the bacterial communities were grossly similar between comparable sediment depths/chemistries (here separated by top vs. lower layers) in all cold seeps, whereas the Urania control sediment communities (cluster III) were distinct and the Milano mat ones (cluster IV) even more distant. For archaea, this analysis showed clusterings akin to those of the bacterial communities, albeit with more exceptions (Fig. 2). In this case, three clusters plus one singleton could be distinguished, as follows. Archaeal cluster I – two of three mud volcano top layers plus the Urania control top layer, cluster II – five of six deeper sediment layers plus the Amsterdam top layer, singleton III – Urania deeper layer and cluster IV – the Milano mat samples plus Kazan-3. Hence, with the exception of one top layer and one deeper layer, again a clustering roughly along sediment sample type (top vs. lower layers) was observed.
Microbial community analysis
Phylogenetic trees constructed from the 16S rRNA gene sequences from this study and those of the nearest relatives from Genbank – obtained in 2005 – are shown in Figs 3 and 4. Because of the high numbers of bacterial and archaeal phylotypes found, only sequences belonging to phylotypes with >1 representative are shown. The majority of both the archaeal and the bacterial 16S rRNA gene sequences did not show high similarity (>97%) to sequences from cultivated species or sequences from clones present in the database. This indicated that the dominant members of the prokaryotic communities were comprised of novel, yet to be isolated, microorganisms.
In most sediment samples, 16S rRNA gene sequences related to those of Chloroflexi and Actinobacteria were present (Fig. 3a), and sequences affiliated with those of members of the Bacilli were found in the lower Napoli sediment layers as well as in the Urania control sediment (Fig. 3b, Table 3). These bacterial groups include many different types of heterotrophic organisms. Sequences related to those of the Alpha subgroup of the Proteobacteria, presumably encompassing numerous sulfate reducers, were found in the lowest sediment layers of all cold seep sediments, being most abundant in Kazan-3. They were also found in the Milano mat samples, mostly so in Milano-2 (Fig. 3b). Across the cold seep sediments, sequences affiliated with the Alphaproteobacteria (a group that is known to encompass key C1-metabolizers) were only found in the Napoli-4 sample, whereas they also occurred in both Urania samples (and in Milano-1). Sequences related to those of the Betaproteobacteria (a group that includes both baceria with highly versatile degradation capacities and chemolithotrophs such as ammonia oxidizers) were only observed in lower sediment layers, namely Napoli-3 and -4 and Amsterdam-2, whereas Urania-2 also contained such sequences (data not shown). Sequences related to Gammaproteobacteria (potentially including sulfur metabolizing prokaryotes as well as facultative heterotrophs) were found in most cold seep sediment layers, as well as in the Milano mat samples. Finally, sequences of Epsilonproteobacteria (including putative endosymbionts) were only detected in the Milano samples (Fig. 3b).
Euryarchaeal, Methanosarcinales-related, 16S rRNA gene sequences were found in Kazan-2 and-3, and Amsterdam-2 sediments, as well as in the Milano mat samples (Fig. 4a and Table 3). Most of the sequences (110) obtained from these samples were related to ANME-2 archaea; less abundant (seven) ANME-1-related sequences were only found in the Amsterdam-2 sample. These ANME-1- and ANME-2- related populations might be involved in the anaerobic oxidation of methane (AOM) (Hinrichs et al., 1999; Orphan et al., 2002). The remaining euryarchaeal sequences found in the Napoli, Kazan and Amsterdam sediments were all related to those of organisms belonging to the Thermoplasmales and Halobacteriales, archaeal groups that include thermophiles, acidophiles and aerobic halophiles. The Halobacteriales-related sequences were most abundant in the lowest Urania sediment layer, Urania-2 (Fig. 4a).
Fairly abundant crenarchaeal sequences affiliated with those of marine benthic Group I archaea (Fig. 4b and Table 2) were detected in all sediment top layers; such sequences are broadly found in marine sediments across the world. The remaining abundant crenarchaeal sequences formed two distinct groups (Fig. 4b). One group consisted of sequences from Urania-2, Amsterdam-1 and all Napoli sediment layers. This group of sequences belongs to a sequence group denominated by marine benthic group B (MBG-B), which is synonymous with the deep-sea archaeal group DSAG (Teske, 2006). The second group of Crenarchaea was only found in the Kazan-2, Urania-2 and Napoli-2 samples (Fig. 4b). This group of sequences is related to those of the miscellaneous archeal group MCG (Teske, 2006). Related sequences have been observed in various environments ranging from freshwater sediments (Hershberger et al., 1996) to deep-sea subfloor samples from the North Atlantic and Sea of Ochotsk (Teske et al., 2002; Inagaki et al., 2003) as well as a mudflow on a deep-sea mud volcano off the coast of Guyana (Madrid et al., 2001).
The results of this study provide insight into the identities of dominant members of the microbial communities at geographically separated deep-sea mud volcanoes in the Eastern Mediterranean Sea. The data indicated the existence of diverse microbial communities in the cold seep sediment (as well as the microbial mat) samples, which were unique for each site studied, as illustrated by the low similarity of phylotypes between the different samples. The Shannon-Weaver diversity index values indicated, in a general sense, that the sediment top layer samples contained more diverse microbial communities than the deeper sediment layers.
Different metabolic processes likely dominated the different sediment layers studied, resulting in different bacterial and archaeal communities. For two cold seeps, namely Napoli and Kazan, we obtained chemical evidence for nitrate and sulfate reductions in the profile (Table 2). Although methane was not measured, it is likely that methane oxidation was also prominent in the Kazan and Amsterdam layers (MEDINAUT, MEDINETH Shipboard Scientific Parties et al., 2000; Werne et al., 2004). Therefore, and in spite of the pitfalls that accompany the inference of function based on phylogeny (Holmes et al., 1995; Chandler et al., 1997; Costello & Lidstrom, 1999; Gray & Head, 2001; Kato & Nogi, 2001; Torsvik & Øvreås, 2002; Purdy et al., 2003), the results obtained provided evidence for the hypothesis that the microbial communities in the Eastern Mediterranean mud volcano cold seep sediments were involved in methane- (and, presumably, sulfide-) based trophic processes.
In the upper layers of mud volcano sediment samples, the most abundant 16S rRNA gene sequences were related to phylogenetic groups that include putative aerobic chemoautotrophs, methanotrophs, methylotrophs as well as aerobic and facultatively anaerobic heterotrophs (Fuerst, 1995; Hugenholtz et al., 1998; Barns et al., 1999). Sequences related to those of sediment top layer sequences from Urania and the mud volcanoes studied have been obtained previously from hydrocarbon-rich sediments or hydrocarbon-metabolizing communities (Phelps et al., 1998; Reed et al., 2002; Teske et al., 2002). Sequences related to those of aerobic methylotrophs (Doronina et al., 2000), chemoautotrophs capable of utilizing reduced sulfur compounds (Eisen et al., 1992; Schweitzer et al., 2001; Smirnov et al., 2001) and sulfide-oxidizing bacteria such as Beggiatoa and Thiobacillus sp. were predominant in the microbial mat. The archaeal sequences from the top layers of mud volcanoes, Urania sediment, and the Milano microbial mat samples were related to sequences that have been previously found in deep-sea environments (Vetriani et al., 1999) or in association with marine plankton (Fuhrman & Davis, 1997). Such sequences are thought to be representative of aerobic heterotrophs or autotrophs (Massana et al., 1997; Ouverney & Fuhrman, 2000; DeLong, 2003). Thus, we hypothesized that oxic sulfide- or methane-dependent chemotrophy was important in the microbial mat, whereas these processes probably played a minor role in the mud volcano sediment top layers. In addition, there is no evidence that these processes occur to a large scale in the Urania sediment.
Anaerobic respiration with sulfate, involving methane or even other carbonaceous compounds, appeared to be a major process in the deeper mud volcano sediment layers, as evidenced by the finding of sequences of bacteria capable of sulfate reduction, of archaea that were potentially involved in AOM and of heterotrophs. The archaea in these samples were related to the ANME-2 and ANME-1 phylogenetic groups. Both groups have been found together in the microbial communities of deep-sea sediments from different areas of the Pacific Ocean and the Gulf of Mexico (Hinrichs et al., 1999; Boetius et al., 2000; Orphan et al., 2001; Valentine, 2002), whereas ANME-1 alone was found to be the dominant group in the Black Sea (Michaelis et al., 2002). In the Eastern Mediterranean, both groups were found, with ANME-2 as most abundant archaea, dominating in samples from the Kazan and Milano mud volcanoes, and ANME-1 predominant in the sediment of the Amsterdam mud volcano. Therefore, AOM in deep-sea cold seep sediments in the Eastern Mediterranean Sea may be carried out by communities dominated by either ANME-2 or ANME-1 archaea (Knittel et al., 2005). These results are comparable to those obtained in previously investigated deep-sea AOM communities in sediments at cold seeps and/or hydrothermal vents (Hinrichs et al., 1999; Orphan et al., 2002; Teske et al., 2002; Knittel et al., 2005).
The inferred metabolic processes for the communities found were supported by the chemistry data. The moderate to high total dissolved carbon concentrations may have constituted an ample supply of carbon for either (chemo-)autotrophic or heterotrophic microbial communities. The degradation of organic matter was supported by the increase of ammonia with depth at two of the three mud volcano sites. We have no explanation for the apparent absence of this process from the Amsterdam mud volcano, but this discrepancy could be explained by advective fluid flow from deep sediment layers, albeit not supported by the bromide and chloride profiles (Schulz, 2000). Although speculative, alternative energy-yielding processes at this site might include anammox or the site might, for reasons unknown, be biologically stalled. Detailed chemistry measurements in the Kazan mud volcano sediments have shown the existence of a largely oxic zone in the uppermost 10 cm of the sediment (Werne et al., 2004). In combination with the high total (2.6%) and organic carbon concentrations (0.6%) found (Werne et al., 2004, Table 2), this is consistent with the occurrence of organisms capable of aerobic heterotrophic growth. Given the presence of nitrate and the high concentrations of total carbon in the sediment top layers, aerobic heterotrophic degradation of organic matter indeed seems to be the most important process in all the sediment top layers examined.
Sulfate reduction (either coupled or not to AOM) and sulfide oxidation were likely metabolic processes in the sediments studied. Given the sometimes abundant presence of Deltaproteobacteria, indicative of SRB, and sulfide in the deeper sediment layers of all three mud volcanoes (but not of the Urania control site), sulfate reduction appeared to be an important process in the mud volcano deeper layers. Supportive of this were the decreasing sulfate concentrations in the sediment layers of the Napoli and Kazan mud volcanoes with depth. However, in the Amsterdam mud volcano, high sulfate concentrations were measured concomitantly with the detection of relatively abundant Deltaproteobacterial sequences related to known SRB. This unexpected result may relate to either advective fluid flow or enhanced mixing of the Amsterdam samples, resulting in the blurring of the stratification of chemistry, microbial processes and communities.
We surmised that populations in the intermediate and deeper layers of the mud volcano sediments, related to Chloroflexi, Bacilli, Thermoplasmales and novel Crenarchaea, were capable of anaerobic (e.g. fermentative) metabolism. This is consistent with the likely absence of oxygen in these layers (Table 2; Werne et al., 2004). The presence of archaea associated with AOM in the deeper, anoxic sediment layers of the Kazan mud volcano was consistent with published data on Kazan pore-water chemistry, notably the typical methane and sulfate depth profiles (indicating sulfate/methane transition zones; Table 3), as well as pore-water carbon isotope measurements (low δ13C values in total carbon; see Werne et al., 2004). Our data on chloride concentrations provide evidence for gas hydrate destabilization, which would release methane (and water) into the surrounding environment (De Lange & Brumsack, 1998) and support AOM. This could explain why chloride concentrations decreased with depth in the Kazan mud volcano sediments (Table 2).
The microbial communities in the Urania sediment samples were different from those of the mud volcano sediments as well as the mat (Fig. 2), and this may relate to the fact that in the Urania sediment there was no evidence for the presence of a cold seep. As a consequence, the local microbial communities most likely depend on organic matter input from the water column by sedimentation. This is consistent with the high abundance of putatively heterotrophic prokaryotes in communities in the Urania sediment layers.
We showed that at the phylogenetic group level (taxonomic rank of ‘class’), similar bacterial communities were present in comparable habitats as well as sediment layers. The identities of the abundant sequences from these communities indicated that specific physiological functions, most notably sulfate reduction, aerobic and anaerobic methanotrophy, aerobic sulfide oxidation, and aerobic and anaerobic heterotrophy, were most likely present among diverse microbial communities in each of the mud volcano sediments studied. This is consistent with the notion that microbial communities in geographically different sites where similar energy-yielding processes reign are often composed of many different species that can perform similar metabolic functions (Yachi & Loreau, 1999).
Samples used in this study were obtained during the French–Dutch Medinaut and Medineth expeditions, which are integrated geological, geochemical and biological studies of mud volcanism and fluid seepage in the Eastern Mediterranean Sea. We thank Maria Schneider and Mayee Wong of the University of Idaho (Moscow, USA) for their support with DNA sequencing and Stephen Bent (Univ. of Idaho, Moscow, USA) for help with the Morisita–Horn similarity calculations. Financial support was provided by the funding organizations IFREMER and NWO-ALW (project-grant 809.63.013).
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