Natural Communities of Novel Archaea and Bacteria Growing in Cold Sulfurous Springs with a String-of-Pearls-Like Morphology

Abstract

We report the identification of novel archaea living in close association with bacteria in the cold (approximately 10°C) sulfurous marsh water of the Sippenauer Moor near Regensburg, Bavaria, Germany. These microorganisms form a characteristic, macroscopically visible structure, morphologically comparable to a string of pearls. Tiny, whitish globules (the pearls; diameter, about 0.5 to 3.0 mm) are connected to each other by thin, white-colored threads. Fluorescent in situ hybridization (FISH) studies have revealed that the outer part of the pearls is mainly composed of bacteria, with a filamentous bacterium predominating. Internally, archaeal cocci are the predominant microorganisms, with up to 10⁷ cells estimated to be present in a single pearl. The archaea appear to be embedded in a polymer of unknown chemical composition. According to FISH and 16S rRNA gene sequence analysis, the archaea are affiliated with the euryarchaeal kingdom. The new euryarchaeal sequence represents a deep phylogenetic branch within the 16S rRNA tree and does not show extensive similarity to any cultivated archaea or to 16S rRNA gene sequences from environmental samples.

Ten years ago, the domain Archaea was considered to consist only of extremophiles such as methanogens, halophiles, and hyperthermophiles (49, 57). However, through the use of the 16S subunit rRNA as a molecular marker (43, 44), many new phylogenetic types within Archaea have recently been discovered in low- to moderate-temperature environments. New 16S rRNA gene sequences have been derived from ocean waters (14, 19, 38, 40), polar seas (41), marine sediments (29, 53, 54), highly saline brine sediments (17), freshwater lakes and sediments (12, 22, 36, 47), terrestrial soils (28, 51), rice roots (20, 30), and marine vertebrates and invertebrates (39, 45, 52), indicating a widespread distribution of Archaea.

Fluorescence in situ hybridization (FISH) using rRNA-targeted oligonucleotide probes is a widely used tool for visualizing microbial cells with a defined phylogenetic affiliation (2, 43). Many new bacterial groups have been detected by this method without the need for cultivation (2). This technique was also successfully applied in a few instances to identify phylogenetically predicted archaea in low- to moderate-temperature biotopes. FISH identified novel methanogens living as endosymbionts in close association with several different genera of protozoa (2, 18). Rod-shaped cells hybridized with a crenarchaeote-targeted oligonucleotide probe within the tissue of the marine sponge Axinella mexicana (45). This marine crenarchaeote, named Cenarchaeum symbiosum, was maintained in stable association with its host in laboratory aquaria for years at 10°C (45). Furthermore, archaeal cocci were identified by FISH on the rhizoplane of rice plants (20).

Recently, through the use of fluorochrome-labeled polyribonucleotide probes, group I and group II marine planktonic archaea in seawater to depths of 3,400 m have been successfully identified and quantified (15). Very recently, FISH studies showed the abundance of a marine microbial consortium consisting of archaea of the order Methanosarcinales and sulfate-reducing bacteria of the delta-proteobacteria. These microbial assemblages seem to mediate anaerobic oxidation of methane in gas-hydrate-rich sediments (6). None of the archaea in low- to moderate-temperature ecosystems, identified by in situ 16S rRNA gene sequence analysis or FISH, have been obtained in pure cultures so far. Therefore, the basic physiological and biochemical properties of these archaea and their function in the natural environment remain largely unclear. In contrast, a great variety of hyperthermophilic archaea have been isolated from sulfur-rich high-temperature environments (50). Many of these cultivated archaea are able to use oxidized or reduced sulfur compounds in their metabolic energy-yielding reactions (50).

We have chosen a sulfurous marsh as the main focus for detailed microbial investigations of novel archaea in nongeothermal environments. This marsh, the Sippenauer Moor, is located about 20 km south of Regensburg, Bavaria, Germany. The total size of the marsh is about 90,000 m². It has belonged to the Regensburgische Botanische Gesellschaft since 1911 and has been a designated environmentally protected area since 1939 (8, 55). About 20 springs arise in the Sippenauer Moor; some of them contain sulfide as a characteristic chemical compound. The waters of the springs form a sulfurous streamlet with a temperature of about 10°C. A specific feature of this streamlet is its high bioactivity, evident by the formation of different-colored large mats and streamers of microorganisms. The subject of the present study was to survey these large microbial assemblages for the existence of archaea by in situ hybridization and by in situ 16S rRNA gene sequence analysis. Here, we report for the first time on a specific microbial community comprising a novel group of archaea and bacteria. The organisms live in close association in this dynamic, low-temperature ecosystem and build up a characteristic, macroscopically visible structure.

MATERIALS AND METHODS

Determination of environmental parameters.

The water temperature was measured using a GTH 1150 digital thermometer (Greisinger Electronic GmbH, Regenstauf, Germany). The pH was determined using pH indicator strips (Merck Eurolab GmbH, Darmstadt, Germany). Chemical analyses (Table 1) were performed at Blasy-Busse GmbH, Eching am Ammersee, Bavaria, Germany.

TABLE 1.

Chemical composition of the main sulfurous spring in the Sippenauer Moor

Component	Concn (mg/liter)
Cations
Ca2+	64
Mg2+	24
Na+	15
K+	2.6
NH4+	0.17
Anions
SO42−	27
CI−	13
NO3−	<1
NO2−	0.05
PO43−	<0.05
SO3−	<1
S2O32−	<0.5
S2−	1.2
Elements
O2	1.4
Mn	0.02
Ni	<0.01
As	<0.001
Al	0.033
Sb	<0.002
B	0.090
Cu	<0.005
Se	<0.002
Zn	0.011
Fe	<0.005

Collection and preparation of samples.

The different-colored microbial mats, filamentous streamers, and pearls from various sites of the streamlet within the Sippenauer Moor were all sampled. An aliquot of each sample was immediately fixed by the addition of formaldehyde (final concentration, 3% [wt/vol]) for in situ hybridization studies. The remaining part of each sample was preserved for DNA extraction (see below). All samples were transported to the laboratory in a refrigerated box at a temperature of about 4°C. Microscopic inspection of the original sample material was routinely carried out using a Zeiss (Oberkochen, Germany) standard phase-contrast microscope equipped with an oil immersion objective (magnification, ×100; numerical aperture, 1.3).

Oligonucleotide probes.

The following rRNA-targeted oligonucleotides were used: EUB338, ARCH915, EURY498, and CREN499. Oligonucleotides were synthesized and 5′ labeled with CY3 [indodicarbocyanine 3-1-O-(2-cyanoethyl)-(N,N-di-isopropyl)-phosphoramidite] or with rhodamine green [5(6)-carboxyrhodamine succinimidyl ester] from Metabion GmbH, Martinsried, Germany. All probe sequences and sources are given in Table 2.

TABLE 2.

Oligonucleotide probes used for FISH

Probe	Specificity	Probe sequence (5′-3′)	Target sitea (rRNA nucleotide positions)	Reference
EUB338	Bacteria	GCTGCCTCCCGTAGGAGT	16S (338–355)	1
ARCH915	Archaea	GTGCTCCCCCGCCAATTCCT	16S (915–934)	48
CREN499	Crenarchaeota	CCAGRCTTGCCCCCCGCT	16S (499–515)	10
EURY498	Euryarchaeota	CTTGCCCRGCCCTT	16S (498–510)	10

^aBased on Escherichia coli numbering (9). 

FISH.

In the laboratory, the formaldehyde-fixed part of each sample was transferred into 10 mM phosphate-buffered saline (pH 7.2) and allowed to stand for 1 h at room temperature. Afterward, the samples were gently squeezed onto a precleaned, gelatin-coated [0.1% gelatin, 0.01% KCr(SO₄)₂] microscope slide (Paul Marienfeld KG, Bad Mergentheim, Germany), dried at 45°C for 30 min, and dehydrated sequentially in 50, 80, and 99% (vol/vol) ethanol (3 min each). After desiccation, whole-cell hybridization was carried out as described elsewhere (10) by use of two labeled probes, either EUB338 combined with ARCH915 or CREN499 combined with EURY498, on a single spot. The hybridization solution contained 0.9 M NaCl, 100 mM Tris-HCl (pH 7.2), 20% (vol/vol) formamide, 0.005% (wt/vol) sodium dodecyl sulfate (SDS), and 50 ng of each probe. Hybridization was followed by a stringent washing step at 48°C for 15 min in washing buffer containing 100 mM Tris-HCl (pH 7.2), 0.18 M NaCl, and 0.005% (wt/vol) SDS. Afterward, the slides were rinsed with distilled water, dried, and mounted in Citifluor AF-1 (Citifluor Ltd., London, United Kingdom).

For epifluorescence microscopy, an Olympus BX60 microscope (Olympus Optical Co. GmbH, Hamburg, Germany) was used with a Plan-Fluorite objective (UplanFl; magnification, ×100; numerical aperture, 1.3) and a 100-W mercury high-pressure bulb. Rhodamine green-labeled cells were visualized by use of filter U-MWB; CY3-marked cells were visualized by use of filter U-MNG. Micrographs were taken by use of an Olympus SC35 camera (Olympus Optical Co.) with Elitechrome 400 film (Kodak, Hemel Hempstead, United Kingdom) for color slides. Exposure times were calculated automatically.

Scanning electron microscopy.

For scanning electron microscopy, a pearl was gently pressed by a cover glass onto a microscope slide, which was then dipped into liquid nitrogen. The cover glass was removed immediately with a razor blade, and the specimens were fixed for 120 min at room temperature. The fixative was composed of cacodylate buffer (75 mM sodium cacodylate, 2 mM MgCl₂ [pH 7.0]) containing 2.5% (final concentration) glutaraldehyde. Further processing was carried out as previously described (25).

Extraction and purification of DNA.

DNA was extracted by the freeze-thaw lysis method (4). Archaeon-containing samples (identified by FISH) were concentrated by centrifugation. The pellets were resuspended in 500 μl of buffer A (100 mM NaCl, 500 mM Tris-HCl [pH 8.0], 1 mM Na₃C₆H₅O₇) and digested with 2 mg of proteinase K (Boehringer GmbH, Mannheim, Germany)/ml for 30 min at 37°C. After the addition of lysis buffer (100 mM NaCl, 200 mM Tris-HCl [pH 8.0], 4% SDS), the mixture was frozen (−80°C) and thawed (56°C) three times. The suspension was extracted with an equal volume of phenol (AquaPhenol, pH 7.5; Appligene, Illkirch, France) and then with phenol-chloroform-isoamyl alcohol (24:24:1, by volume). DNA was precipitated overnight at room temperature by the addition of 0.6 volume of isopropyl alcohol. After two wash steps with 70% (vol/vol) ethanol, the DNA was dried and resuspended in pure water (LiChrosolv; Merck). DNA was further purified on a 1% (wt/vol) low-melting-point agarose gel by electrophoresis at 70 V for 90 min. The ethidium bromide-stained DNA band was visualized under UV light and excised. DNA in the agarose slice was recovered using GeneClean II (Bio 101, Inc., Vista, Calif.) according to the manufacturer's instructions.

PCR amplification and cloning.

Extracted DNA was used as a template for the PCR amplification of archaeal 16S ribosomal DNA (rDNA) with the forward primer 344aF and the universal reverse primer 1406uR or 1512uR (Table 3). The 50-μl reaction cocktail contained (final concentrations) 5 to 50 ng of DNA, PCR buffer II (PE Applied Biosystems, Foster City, Calif.), 1.5 mM MgCl₂, 200 μM each deoxynucleotide triphosphate, 1 ng each of forward and reverse primers, and 2.5 U of AmpliTaq DNA polymerase (PE Applied Biosystems). Amplification was performed with 0.2-ml reaction tubes and a thermal cycler (model 9600; PE Applied Biosystems). The thermal profile used for amplification of the 16S rDNA sequences was as follows: 96°C for 90 s; 10 cycles of 96°C for 30 s, 60°C for 30 s, and 72°C for 1 min; 25 cycles of 94°C for 20 s, 60°C for 30 s, and 72°C for 1 min (plus a 2-s time increment for each further cycle); and a final incubation at 72°C for 10 min.

TABLE 3.

Oligonucleotide primers used for PCR and sequencing

Primer	Specificity	Primer sequence (5′-3′)	Target sitea (rRNA nucleotide positions)	Reference
344aF	Archaea	CGGGGYGCASCAGGCGCGAA	16S (344–363)	11
1119aR	Archaea	GGYRSGGGTCTCGCTCGTT	16S (1101–1119)	11
1406uR	Universal	ACGGGCGGTGTGTRCAA	16S (1390–1406)	31
1512uR	Universal	ACGGHTACCTTGTTACGACTT	16S (1492–1512)	31

^aBased on E. coli numbering (9). 

Amplified PCR products were purified with Microcon 100 microconcentrators (Amicon Inc., Beverly, Mass.) as recommended by the manufacturer. The purified PCR products were cloned using a TOPO TA cloning kit (Invitrogen, San Diego, Calif.) in accordance with the manufacturer's instructions. The presence of inserts of the expected sizes was analyzed by direct PCR screening of 30 to 40 transformants without plasmid extraction. A small part of each colony was used for a PCR with the plasmid-specific primers M13F(−40) and M13R. The sizes of the inserts were checked by electrophoresis on a 1% (wt/vol) agarose gel.

RFLP and sequencing of rRNA gene clones.

For restriction fragment length polymorphism (RFLP) analyses, the PCR products were digested with a mixture of four different restriction endonucleases. Ten microliters of each PCR product was mixed with 1 μl of buffer REact 1 (10×) and 0.25 μl (10 U/μl) each of the enzymes AluI, HhaI, HinfI, and RsaI (Life Technologies, Paisley, United Kingdom). All of these enzymes cleave at specific sites within 4- or 5-bp recognition sequences with different GC contents. The preparations were incubated at 37°C for 3 h and then electrophoresed on a 3% (wt/vol) agarose 1000 gel (Life Technologies) for 90 min at 85 V using a 25-bp ladder as a DNA size standard (Life Technologies). The ethidium bromide-stained DNA bands were visualized under UV light, and the fingerprints were compared. Representative transformants were selected on the basis of the 16S rRNA gene fingerprint patterns. The corresponding plasmid DNA was extracted using QIAprep Spin Miniprep kits (Qiagen GmbH, Hilden, Germany) as recommended by the manufacturer. Sequencing of the 16S rDNA inserts was performed at the Department of Biology (University of Regensburg, Regensburg, Germany) with an ABI 310 capillary sequencer (PE Applied Biosystems) and a Big Dye Terminator Cycle Sequencing Ready Reaction kit (PE Applied Biosystems). The sequences were amplified with the archaeal primer 1119aR (Table 3) and the plasmid-specific primers M13F(−40) and M13R.

Phylogenetic analyses.

For phylogenetic analyses, an alignment of approximately 10,500 homologous full and partial sequences (>1,300 nucleotides) available in public databases (ARB Project [33, 34]) was used. The new 16S rRNA gene sequence (1,085 nucleotides) was integrated in the 16S rRNA alignment using the corresponding automated tools of the ARB software package (34, 35). The resulting alignment was checked manually and corrected if necessary. For tree reconstruction, maximum-parsimony, distance-matrix (Jukes-Cantor correction), and maximum-likelihood (fastDNAml) methods were applied as implemented in the ARB software package. The sequence was submitted to the CHECK_CHIMERA program of the Ribosomal Database Project (37) to detect possible chimeric artifacts.

Nucleotide sequence accession number.

The 16S rRNA gene sequence of archaeal sequence clone SM1 was deposited in the EMBL nucleotide sequence database under accession number AJ296315.

RESULTS

Chemical and physical analyses.

Since January 1998, temperature and pH measurements have been obtained periodically at different sites of the selected streamlet in the Sippenauer Moor (Fig. 1). A nearly constant water temperature of 10°C (±2°C) and a constant pH of 6.5 have been determined. Chemical analyses of water from the main sulfurous spring have revealed a sulfide concentration of 1.2 mg/liter and a low oxygen concentration of 1.4 mg/liter. The chemical composition is listed in more detail in Table 1. The low salt content is typical for a freshwater environment.

FIG. 1.

Topographic map of the location of the Sippenauer Moor in Bavaria, Germany. The topographic map was generated using the Online Map Creation program (Geomar, Kiel, Germany; http://www.aquarius.geomar.de).

FISH studies.

In the initial studies, samples from the different-colored microbial mats and streamer-like cell masses were used for in situ hybridization studies with domain-specific bacterial and archaeal hybridization probes. In almost all samples, bacteria with different morphologies were the main constituents. Filament-forming bacteria were very frequently the dominant morphotype. Very rarely, archaeal cocci (about 0.01% of the total microbial population) could be identified randomly in the samples.

During our investigations of the Sippenauer Moor, we detected macroscopically visible microbial cell assemblages. They exhibited a characteristic morphology, comparable to a string of pearls. Tiny, whitish individual globules (the pearls) connected by a thin white thread were floating in the current of the sulfurous water. Approximately 2 to 15 pearls were arranged in a relatively linear series. Each string of pearls was attached at one end to a rigid material (e.g., branches, algae, leaves, or stones) (Fig. 2). The total length of this unusual structure was up to 15 cm; the diameter of the pearls varied between 0.5 and 3.0 mm (Fig. 2). In contrast to the other samples studied (see above), the ratio between archaea and bacteria was dramatically different within the pearls (Fig. 3). FISH revealed that the outer part of a pearl was composed mainly of bacteria and dominated by a filamentous morphotype. Internally, in contrast, archaeal cocci were the predominant microorganisms, with up to 10⁷ archaeal cells estimated to be present in a single pearl (Fig. 3). The use of kingdom-specific hybridization probes showed that these archaeal cocci belonged to the Euryarchaeota. When single pearls enlarged (up to 8 mm in diameter), the concentration of the archaea decreased. A population of morphologically diverse bacteria could be identified by FISH at this stage. Additionally, the pearls collapsed easily when sampled, and gas bubbles of unknown chemical composition were released. This finding indicates that gas had accumulated inside the pearls.

FIG. 2.

Photograph of archaeal and bacterial communities growing in a morphologically characteristic structure (the strings of pearls) in the sulfurous water of the Sippenauer Moor. Orange arrows point to single pearls.

FIG. 3.

FISH of a part of a pearl. (A) Phase-contrast micrograph. (B) Epifluorescence micrograph. Dual hybridization was done with a rhodamine green-labeled archaeal probe (ARCH915) and a CY3-labeled bacterial probe (EUB338). The archaeal cocci stain green; the bacteria stain red.

The growth of microorganisms in a string-of-pearls-like morphology was observed for more than 1 year at different sulfurous sites of the streamlet within the Sippenauer Moor. Periodically performed FISH studies showed that the smaller pearls (a diameter of up to 3 mm) especially revealed the characteristic archaeal-bacterial composition.

16S rRNA gene sequence analysis.

The phylogenetic position of the archaea inside a pearl was further investigated by 16S rRNA gene sequence analysis. The PCR products of the archaeal 16S rRNA genes were cloned, and a total of 21 randomly selected clones were screened by RFLP analysis. All clones showed the same restriction pattern, indicating that the archaeal component inside a pearl consisted of a single phylotype. The 16S rRNA gene sequences of four clones were determined (about 1,100 bp) and showed an identical base composition. Comparative analysis of the 16S rRNA gene sequences showed that the new sequence clustered within the euryarchaeal kingdom (Fig. 4), in agreement with the results of the FISH studies. The sequence branched deeply in the phylogenetic tree, with no close phylogenetic relationship to either environmentally derived sequences or any cultivated euryarchaeal sequences (Fig. 4). The branching point was supported by all three tree construction methods (not shown). The definitive confirmation of a single archaeal phylotype by use of a sequence-specific FISH probe awaits further study.

FIG. 4.

16S rRNA gene-based phylogenetic tree showing the position of the archaeal sequence clone (SM1) derived from a single pearl. The topology of the tree is based on the results of a maximum-parsimony analysis (ARB software package [34, 35]). Reference sequences were chosen to represent the broadest diversity of archaea. The scale bar shows a 10% estimated difference in nucleotide sequence positions.

Structural studies.

Phase-contrast microscopy was used to investigate the structural arrangement of the archaeal cocci within the pearls without chemical treatment. Large cell assemblages of cocci could be identified when a pearl was gently squeezed onto a microscope slide by use of a cover glass. The cells appeared to be grouped in a three-dimensional arrangement at defined distances from each other. This typical orientation of the archaeal cocci was observed for all pearls investigated throughout the year.

Scanning electron microscopy was performed on individual pearls fixed on a cover glass. An overview of the inner part of a pearl is shown in Fig. 5A. The overwhelming majority of the organisms were cocci; a small number of rod-shaped cells also were present. An enlarged view of part of Fig. 5A revealed that the cells were spherical and had a diameter of about 0.6 μm (Fig. 5B). Single cocci were embedded in fibrous material (most likely representing a polymeric substance) with an as-yet-undetermined chemical composition (Fig. 5B).

FIG. 5.

Scanning electron micrographs of a single pearl from the Sippenauer Moor. (A) Overview of the inner part of a pearl, showing large numbers of small cocci. (B) Detail of the inset in panel A. Single round cocci are embedded in a fibrous matrix.

DISCUSSION

Characteristic, macroscopically visible structures, morphologically comparable to strings of pearls, were detected in the cold (approximately 10°C) sulfurous marsh water of the Sippenauer Moor. The combination of environmental observations and data from phase-contrast microscopy, scanning electron microscopy, and FISH has enabled initial interpretation of the microbial architecture of these unusual structures. So far, two groups of microorganisms which contribute significantly to the structures have been identified. One group is represented by filamentous bacteria. They seem to play an important role in the formation of the white threads connecting single pearls, and they also participate in the formation of the outer, whitish part of the pearls. The whitish appearance may be due to the presence of elemental sulfur, formed by the oxidation of sulfide during growth of the filamentous organisms (5). The interior of the pearls is dominated by a novel group of archaea, which form microcolonies within the filamentous bacteria. Interestingly, the archaeal cells of a microcolony are encased in a polymeric matrix with an as-yet-undetermined chemical composition (Fig. 5B), in which they are arranged three-dimensionally at defined distances from each other. So far, it is not known whether the polymer is bacterial or archaeal in origin. However, because of the specific archaeal cell orientation, the polymer most likely is produced by the archaea during growth. The biological function of the polymer is not known yet, but it could play an important role in the formation and maintenance of the globular structure of the pearls in nature. The formation of different polymeric substances by various bacteria in natural biofilms has been observed frequently but has not previously been reported for members of the archaeal domain in nature (13, 21).

In a pearl, the archaea and bacteria form a stable association over an extended period of time; this association makes specific interactions between the two partners possible (7). These defined communities are observed especially in smaller pearls, with a diameter of 0.5 to 3 mm. At this stage, a perfect milieu for the horizontal transfer of genetic material between members of two different domains is present, as recently inferred by DNA sequence comparisons between hyperthermophilic archaea and bacteria (16, 42, 56).

When single pearls enlarge (a diameter of up to 8 mm), the interior becomes colonized by a variety of morphologically diverse bacteria, while the number of archaea decreases. Additionally, the pearls collapse easily when sampled, and gas bubbles of unknown chemical composition are released. These observations indicate that the structure-stabilizing polymer has been digested and that gas has been formed as a degradation product. At this stage, the encased archaea can be released from the polymeric matrix.

The newly discovered strings of pearls provide a significant opportunity for further studies, which may lead to interesting physiological and ecological insights. They represent a hot spot for a novel archaeal group and are permanently present in the sulfurous springs of the Sippenauer Moor. Therefore, they are easily accessible for investigation in the natural biotope as well as in the laboratory. A first insight into the metabolic properties of the archaea within the pearls could be derived directly from natural samples, e.g., by the combination of FISH and microautoradiography (3, 32). The internal milieu could be characterized by use of different microelectrodes or micro-optodes (23, 46). The basis for a deeper understanding of the new archaea in the pearl microenvironment is the cultivation and isolation of these organisms. Isolation attempts will be preferentially performed by a newly developed isolation strategy based on the use of an “optical tweezer trap” (24, 26, 27).

An interesting question concerns the relative occurrence, distribution, and abundance of strings of pearls in diverse sulfide-containing ecosystems. Recent field studies have revealed that the growth of microorganisms in this characteristic morphology is not restricted to the sulfurous freshwater springs of the Sippenauer Moor. We observed similar structures in other sulfurous freshwater ecosystems located at different sites in Bavaria, Germany. Interestingly, strings of pearls were also detected in a sulfurous seawater streamlet near the coast of Dalyan, Turkey, indicating that the growth of microorganisms in a strings of pearls-like morphology may be a widely distributed form of life.