A cold-loving crenarchaeon is a substantial part of a novel microbial
community in cold sulphidic marsh water
Marcus Koch1, Christian Rudolph1, Christine Moissl1 & Robert Huber2
1
Lehrstuhl für Mikrobiologie und Archaeenzentrum, Universität Regensburg, Regensburg, Germany; and 2Kommunale Berufsfachschule für BiologischTechnische Assistenten, Straubing, Germany
Correspondence: Robert Huber,
Kommunale Berufsfachschule für BiologischTechnische Assistenten, Stadtgraben 39,
D-94315 Straubing, Germany. Tel.: 149 9421
23811; fax: 149 9421 23884; e-mail:
[email protected]
Received 2 August 2005; revised 20 November
2005; accepted 21 November 2005.
First published online 31 January 2006.
doi:10.1111/j.1574-6941.2006.00088.x
Editor: Ralf Conrad
Keywords
Archaea; Crenarchaeota; Thiothrix;
sulphidic springs; string-of-pearls;
psychrophilic; microbial community.
Abstract
In this paper, we report the identification and first characterization of a novel,
cold-loving, prokaryotic community thriving among white-greenish ‘streamers’ in
the cold (c. 10 1C) sulphurous water of the marsh Sippenauer Moor near
Regensburg, Bavaria, Germany. It consists of the bacterial genus Thiothrix, the
bacterium ‘Sip100’ and one archaeal representative, forming together a unique
association structure with a distinct life cycle. Fluorescence in situ hybridization
studies have revealed that the archaeal member can be affiliated to the crenarchaeal
kingdom (‘Cre1’). This crenarchaeon was always observed attached to the bacterial
community member ‘Sip100’. Extended fluorescence in situ hybridization studies
showed that this crenarchaeon was not detected in a free-living form, raising the
idea of a probable host-dependent relationship. In line with our fluorescence in
situ hybridization studies, novel crenarchaeal 16S rRNA gene sequences were
identified in these samples. The design and application of a new in situ cultivation
method in the sulphurous water of the marsh allowed first insights into the
cohesion mechanisms, lifestyle and chronology of the microbes involved in this
prokaryotic community in nature. Our results suggest that hitherto unknown
Crenarchaeota thrive in cold sulphidic water and are a substantial part of a
synchronized microbial community.
Introduction
Archaea, one of the three domains of life, has long been
believed to consist almost exclusively of extremophilic microorganisms, inhabiting only specific environmental niches,
inhospitable to most Eucarya and Bacteria (Woese et al.,
1990; McInerney et al., 1997). The domain Archaea is divided
into the four kingdoms Crenarchaeota, Euryarchaeota, Korarchaeota and Nanoarchaeota (Huber et al., 2003). Within the
Crenarchaeota, all cultivated representatives originate almost
exclusively from geothermally heated habitats, such as submarine vents and terrestrial hot springs (Stetter, 1994, 1995).
Almost all are hyperthermophiles, requiring temperatures
above 80 1C for optimal growth (Stetter, 1996).
16S rRNA gene phylogenetic analysis, and its widespread
application in environmental microbiology (Olsen et al.,
1986; Pace et al., 1986), has revealed a multitude of new
phylogenetic lineages of, so far, uncultivated Crenarchaeota
in low- to moderate-temperature marine and terrestrial
environments (DeLong, 1998; Ochsenreiter et al., 2002;
Schrenk et al., 2003). Fluorescence in situ hybridization
FEMS Microbiol Ecol 57 (2006) 55–66
(FISH) with Crenarchaeota-specific probes has even allowed
some of these Crenarchaeota to be seen (Preston et al., 1996;
Großkopf et al., 1998; Karner et al., 2001; Pernthaler et al.,
2002). Within the tissue of the marine sponge Axinella
mexicana, rod-shaped cells were identified with a Crenarchaeota-targeted oligonucleotide probe (Preston et al., 1996).
This marine crenarchaeon, tentatively named ‘Cenarchaeum
symbiosum’, was maintained in stable association with its
host in laboratory aquaria for years at 10 1C (Preston et al.,
1996). Further studies have shown that two closely related
variants of ‘Cenarchaeum symbiosum’ were consistently
found in the majority of host individuals analysed (Schleper
et al., 1998). Although cold-adapted Crenarchaeota account
for up to 20–40% of the planktonic communities in some
marine geographical areas (DeLong et al., 1994; Massana
et al., 1997; Karner et al., 2001), their biology and function in
the natural environment remain largely unclear (Nicol et al.,
2003; Wuchter et al., 2003). Until now, none of these
Crenarchaeota has been obtained in pure culture (Ferguson
et al., 1984; Lee & Fuhrman, 1991; Stein et al., 1996; Vetriani
et al., 1998).
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M. Koch et al.
The failure of standard techniques to isolate phylogenetically predicted prokaryotes is well known (Ward et al., 1990;
Barns et al., 1994; Amann et al., 1995), and may result from
their living in complex microbial communities (e.g. biofilms) and consequent host dependence (Schink, 1992; Paerl
& Pinckney, 1996;Abella et al., 1998; Fröstl & Overmann,
1998). Special partnerships between sulphate-reducing Bacteria and methanogenic Euryarchaeota have been described
for the anaerobic methane-oxidizing communities of gas
hydrate-rich, marine, anoxic sediments (Boetius et al., 2000)
and for the archaeal/bacterial string-of-pearls community
thriving in cold (10 1C) low-salt sulphurous spring water of
the marsh Sippenauer Moor (Rudolph et al., 2001) or in the
sulphurous streamlet Islinger Mühlbach (Rudolph et al.,
2004). The string-of-pearls community forms macroscopically visible globules, the ‘pearls’, containing microcolonies
of the nonmethanogenic SM1 euryarchaeon, which are
surrounded by mainly filamentous Bacteria (Rudolph et al.,
2001, 2004; Moissl et al., 2002, 2003). Single cells of the SM1
euryarchaeon produce surface appendages which are unique
among the prokaryotes studied so far, and represent a new
class of cell appendages of high complexity with a welldefined base-to-top organization, for which the name
‘hami’ has been proposed (Moissl et al., 2005).
During our FISH investigations of the string-of-pearls
community in the cold sulphurous marsh water of the
Sippenauer Moor near Regensburg, Bavaria, Germany (Rudolph et al., 2001), we also detected, very rarely, crenarchaeal cells, which were attached to rod-shaped Bacteria.
Advanced FISH studies during the last 4 years have shown
that these prokaryotes form a unique community, which
consists of a novel, cold-loving Crenarchaeum, tentatively
named ‘Cre1’, the bacterial genus Thiothrix and the bacterium ‘Sip100’. In this study, we report the identification,
in situ cultivation and life cycle of this particular microbial
community.
Materials and methods
Determination of environmental parameters
Water temperatures were measured using the flow and
temperature measurement tool Miniair 2 (Schildknecht
Messtechnik, Gossau, Switzerland). Oxygen concentrations
and conductivities were determined using the multifunction
measurement instrument MultiLineP4 (WTW, Weilheim,
Germany). Sulphide content and pH were determined as
described previously (Rudolph et al., 2004). Chemical
analyses of the marsh Sippenauer Moor and the streamlet
Islinger Mühlbach were performed at Blasy-Busse GmbH
(Eching am Ammersee, Germany) (Rudolph et al., 2001).
Collection and preparation of samples
In the Sippenauer Moor, c. 500 samples of different coloured
microbial mats, white-greenish coloured streamers, water
and sediment samples were taken at intervals of about 200 m
along the streamlet called Mineralbach emanating from the
spring area (Fig. 1). These samples were processed as
described previously (Rudolph et al., 2001). At the sampling
site of the Islinger Mühlbach, samples of greenish streamers
were taken using the same sampling procedure. A detailed
water analysis has been published recently (Rudolph et al.,
2004). Sampling from the new in situ cultivation system (socalled ‘agar noodles’) was performed by removing small
parts from the free-floating end of the noodles for subsequent FISH analyses. The slices were immediately fixed by
the addition of formaldehyde [final concentration, 3% weight
in volume (w/v)]. The inner part of the noodles was removed
3
4
0
25
50 75 100 125 m
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2
1
Fig. 1. Map of the protected area of the marsh
Sippenauer Moor near Regensburg, Bavaria,
Germany (N 48152.161 0 , E 11157.355 0 Warneke,
1992): 1, spring area with sulphidic springs (Str.1);
2, timber bridge at sampling site Str.6; 3, confluence of the Mineralbach and Feckinger Bach;
4, marsh area.
FEMS Microbiol Ecol 57 (2006) 55–66
57
Cold-loving crenarchaeon in cold sulphidic marsh water
using a sterile plastic tube as drill to reduce the amount of
agar. Alternatively, the surface of the noodle was scraped off
using a sterile razor blade. The material obtained was reduced
to small pieces using tweezers or a Silamat Plus dental shaker
(Vivadent Dental GmbH, Ellwangen, Germany), resuspended
in 0.5–1 mL of 1 phosphate-buffered saline (PBS), washed
twice in 1 PBS and used for FISH.
FISH and oligonucleotide probes
The streamer samples were either reduced to small pieces
using a Silamat Plus dental shaker or were gently squeezed
onto precleaned, gelatine-coated [0.1% gelatine, 0.01%
KCr(SO4)] microscope slides (Paul Marienfeld KG, Bad
Mergentheim, Germany; sample volume, 10–15 mL). After
desiccation, FISH was carried out as described recently
(Rudolph et al., 2001). All samples were stained with a
combination of three different Archaea-specific probes
(‘Arch-Mix’; Arch 344, Arch 915, Arch 1060 Moissl et al.,
2002) and a combination of three different Bacteria-specific
probes (‘Eub-Mix’ Rudolph et al., 2004). A kingdom-specific
characterization of the Archaea was carried out with the FISH
probes Cren499, Eury498 (Burggraf et al., 1994) and the new
crenarchaeal probe Cren457R (5 0 -TTG CCC CCC GCT TAT
TCS CCC G-30 ), designed after the PCR primer Cren457R
(Schleper et al., 1997). Archaeal 16S rRNA gene sequences
available in public databases (ARB program (Ludwig & Strunk,
1997); BLASTN, National Center for Biotechnology Information, http://www.ncbi.nlm.nih.gov/) were used to check the
specificity of the new probe for Crenarchaeota using the
‘Probe_Match’ function of the ARB program (Ludwig &
Strunk, 1997). The specificity of the probe was tested in several
FISH experiments with different genera of Euryarchaeota and
Crenarchaeota (Table 1). For further characterization of the
bacterium ‘Sip100’, different specific probes for Alpha-, Beta-,
Gamma- and Deltaproteobacteria (ALF968 (Neef, 1997);
Bet42a, GAM42a (Manz et al., 1992); DELTA495a/b/c Loy
et al., 2002) and for the Cytophaga/Flavobacterium/Bacteroides
(CFB) group (CFB319a/b Manz et al., 1996) were used. All
probes were labelled with different dyes [rhodamine green
(RG), carbocyanine (Cy3), Texas Red] and were synthesized
by Metabion (Martinsried, Germany). Following the FISH
procedure, 4 0 ,6-diamidino-2-phenylindole (DAPI) staining was
performed (Moissl et al., 2002). For microscopy, an Olympus
BX60 with the UV power supply unit U-RFL-T (Olympus,
Hamburg, Germany), with a set of filters (AHFanalysentechnik
AG, Tübingen, Germany; Table 2) for the different dyes, was
used. Photographs were taken using a Nikon Coolpix 990
(Nikon Corporation, Tokyo, Japan) digital camera.
Design of a new in situ cultivation system
In their biotope, the amount of the crenarchaeal/bacterial
community was variable and was dependent on environFEMS Microbiol Ecol 57 (2006) 55–66
Table 1. Verification of the newly designed hybridization probe
Cren457R with different archaeal genera
Probe
Organism
Cren457R
Eury498
Arch-Mix
Pyrobaculum islandicum
Thermoproteus tenax
Pyrodictium occultum
Sulfolobus acidocaldarius
Ignicoocus islandicus
Thermococcus celer
Methanococcus igneus
Pyrococcus furiosus
Archaeoglobus fulgidus
SM1 euryarchaeon
1
11
11
11
11
11
11
11
11
1
111
11
111
11
11
11
11
11
111
, no fluorescence in situ hybridization (FISH) signal; 1, satisfactory FISH
signal; 11, strong FISH signal; 111, very strong FISH signal.
Table 2. Filter sets used for fluorescence in situ hybridization (FISH)
Dye
Exciter
Long-pass
dichroic mirror
Band-elimination
filter
Cy3
DAPI
RG
TR
HQ 546/12
BP 360-370
HQ 480/40
D 585/10
Q 560 LP
DM 400
Q 505 LP
Q 595 LP
HQ 585/40
BA 420
HQ 527/30
HQ 645/75
Cy3, carbocyanine; DAPI, 4 0 ,6-diamidino-2-phenylindole; RG, rhodamine
green; TR, Texas Red.
2 cm
Fig. 2. Micrograph of the newly designed in situ cultivation system. The
red arrow points to the nylon thread, the white arrow to the ‘agar
noodle’.
mental factors. Therefore, it was of interest to develop an in
situ cultivation technique to obtain larger quantities of these
populations in a reproducible way. Different threads (polyethylene, polyamide, nylon, wool) and polyethylene nets
were used for cultivation attempts, according to Moissl et al.
(2003). However, as none of these materials led to a
successful adhesion and subsequent cultivation of the new
prokaryotic community, we designed another in situ cultivation system (Fig. 2) mainly based on a mixture of different
polysaccharides present in the biotope. For the preparation
of the in situ ‘agar noodles’ (length, 5 or 10 cm; diameter,
8 mm; Fig. 2) with integrated nylon threads, special epoxy
resin moulds were designed. The moulds were sterilized
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M. Koch et al.
with ethanol (70%) and prepared with the nylon threads,
which had been autoclaved in marsh water for 40 min at
121 1C. The agar noodles consisted of 1.8% prewashed agar,
ı-, k- and l-carrageenan (each 0.15%) and 0.15% alginate.
The components were dissolved in filtered marsh water and
autoclaved for 20 min at 121 1C. The polymer mixture was
cooled to about 60 1C and then poured into the moulds.
After 30 min at room temperature, the moulds were opened
and the solidified agar noodles were stored in autoclaved
marsh water until use. All preparation steps were performed
under a laminar flow to prevent contamination.
The agar noodles were fixed with the nylon threads in the
streamlet at site Str.6 of the Sippenauer Moor (Fig. 1),
thereby floating between white-greenish streamers for about
5–7 weeks. Sampling was performed about once or twice a
week by removing small parts from the free-floating end of
the noodles for subsequent FISH analyses (see ‘Collection
and preparation of samples’).
Extraction of DNA
All equipment was treated with 5% HClO4 to remove DNA
impurities and washed with double-distilled water. Cell lysis
and bulk DNA extraction were performed as described previously (Rudolph et al., 2001). Alternatively, DNA was isolated
with the DNeasys Tissue Kit (Qiagen GmbH, Hilden, Germany) according to the manufacturer’s instructions.
Purification of DNA
Humic acids were removed by a modified method (Kuske
et al., 1998) using Sephadex G200-120/polyvinylpolypyrrolidone (PVPP)-filled centrifugal devices (Nanoseps MF
Centrifugal Devices 0.2 mm; Pall Corporation, Ann Arbor,
MI). The preparation of PVPP was performed as described
by (Holben et al., 1988), but using a cellulose filter (Schleicher & Schuell, Nr. 595, Dassel, Germany) instead of MIRACLOTH (Holben et al., 1988). The following purification
steps were performed in a laminar flow to prevent contamination. Sephadex G200–120 (0.7 mL) was mixed with
20 mg of washed PVPP. Sterile TE buffer (EDTA, 0.001 M;
Tris/HCl, 0.001 M; pH 8.0) was added until all resin was
wetted out, and equilibrated overnight at 4 1C. After dilu-
tion with sterile TE buffer, 600 mL was pipetted onto a
Nanosep centrifugal device and packed to about 500 mL by
centrifugation at 800 g for 15 min. The flow-through was
discarded. Premade columns could be stored at 4 1C for
several days. For purification, up to 200 ng or 100 mL of
humic acid-contaminated brownish coloured DNA was
carefully loaded onto a single column and centrifuged for
15 min at 800 g. DNA elution was performed by a further
centrifugation step with 50–100 mL of TE buffer. The DNA
solution obtained became colourless after centrifugation.
The DNA concentration was estimated using an ethidium
bromide plate and different concentrations of lambda DNA
by intensity comparison.
PCR amplification and cloning
Purified DNA was used as template for the PCR amplification of archaeal 16S rRNA gene sequences (Table 3) with
different Archaea-specific primer combinations (Table 4).
PCR was performed as described previously (Rudolph et al.,
2001). PCR products were purified using MontageTM PCR
centrifugal filter devices (Millipore Corporation, Bedford,
MA), as recommended by the manufacturer. In order to
amplify exclusively Crenarchaeota-specific sequences in
these purified PCR products, they were utilized for subsequent nested PCR with Archaea- and Crenarchaeota-specific
primer combinations (Tables 3 and 4). The sequence of the
primer 457R was also converted to the forward primer 457F.
The thermal PCR profile for the amplification of the 16S
rDNA sequences consisted of 30 cycles at 95 1C for 60 s,
50 1C for 60 s and 72 1C for 120 s. After purification (see
above), the PCR products were cloned using the original TA
Clonings Kit (Invitrogen BV, Breda, the Netherlands) or the
Qiagen PCR Cloning Kit (Qiagen GmbH), according to the
manufacturer’s instructions.
Restriction fragment length polymorphism,
sequencing of rRNA gene clones and
phylogenetic analyses
Restriction fragment length polymorphism (RFLP) and
sequencing were carried out as described previously (Rudolph et al., 2001). Clones with unique RFLP patterns were
Table 3. Primers used for 16S rRNA gene amplification
Primer
Specificity
Primer sequence (5 0 –3 0 )
Target site (rRNA nucleotide positions)
Reference
8aF
28aF
344aF
1119aR
1406uR
457R
9bF
Archaea
Crenarchaeota
Archaea
Archaea
Universal
Crenarchaeota
Bacteria
TCY GGT TGA TCC TGC C
AAT CCG GTT GAT CCT GCC GGA CC
CGG GGY GCA SCA GGC GCG AA
GGY RSG GGT CTC GCT CGT T
ACG GGC GGT GTG TRC AA
TTG CCC CCC GCT TAT TCS CCC G
GRG TTT GAT CCT GGC TCA G
8–23
28–51
345–364
1101–1119
1406–1390
489–510
9–27
Burggraf et al. (1992)
Schleper et al. (1997)
Burggraf et al. (1997)
Burggraf et al. (1997)
Lane (1991)
Schleper et al. (1997)
Burggraf et al. (1992)
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FEMS Microbiol Ecol 57 (2006) 55–66
59
Cold-loving crenarchaeon in cold sulphidic marsh water
Table 4. Primer combinations for PCR and nested PCR
PCR primer
combination
Nested PCR primer
combination
Cloned PCR
products
8aF-1119aR
8aF-1406uR
8aF-457R
8aF-1119aR
28aF-457R
28aF-1119aR
457F-934aR
457F-1060aR
28aF-457R
28aF-1119aR
457F-934R
457F-1060R
457F-1119aR
—
344aF-1119aR
344aF-1406uR
chosen for sequencing. Phylogenetic analyses were performed as described by Rudolph et al. (2001).
Nucleotide sequence accession number
The archaeal 16S rRNA gene sequences were deposited in
the European Molecular Biology Laboratory (EMBL)
nucleotide sequence database. The accession numbers are
AM055699–AM055710, as shown in Fig. 6.
Results
10 µm
Fig. 3. Fluorescence in situ hybridization (FISH) of white-greenish streamers from the Sippenauer Moor, sampling site Str.6 (Fig. 1). Epifluorescence micrograph. Hybridization was performed with a rhodamine
green-labelled, Archaea-specific probe mix (Moissl et al. 2003) and a
carbocyanine-labelled, Bacteria-specific probe mix (Rudolph et al. 2004).
The newly discovered Crenarchaeota stains green (red arrows), the
Bacteria appear orange/red (white arrows). The green arrow shows the
Thiothrix filament.
Chemical and physical analyses
Since March 2001, temperature and pH measurements have
been performed periodically at site Str.6 of the Mineralbach
in the marsh Sippenauer Moor (Fig. 1) (Bresinsky, 1991,
1999, 2001), located about 90 m downstream of the main
spring Str.1 (Fig. 1) (Rudolph et al., 2001, 2004; Moissl et al.,
2002, 2003). The same analyses were performed at the
sampling site Islinger Mühlbach, about 10 m downstream
of the drill hole (Rudolph et al., 2004). Some characteristic
and site-specific chemical and physical parameters are
shown in Table 5; a detailed chemical composition of both
springs has been described by (Rudolph et al., 2004).
FISH studies
In our initial FISH studies, all samples were hybridized with
domain-specific bacterial and archaeal probes. In almost all
samples, Bacteria with different morphologies were the main
constituents. Filament-forming Bacteria were very frequently the dominant morphotype in mat and streamer
samples. In addition, archaeal cocci were identified in these
samples. They could be attributed to the already described
SM1 euryarchaeon (Rudolph et al., 2001) using the specific
SMARCH714 probe (Moissl et al., 2003).
During FISH investigations of the main streamlet of
Sippenauer Moor, a novel, characteristically shaped microbial cell assemblage was detected at sampling site Str.6
within the streamlet of Sippenauer Moor (Figs 1 and 3). It
FEMS Microbiol Ecol 57 (2006) 55–66
consisted of two types of the Bacteria and one crenarchaeal
representative. The growth of this community was reliably
observed for more than 4 years at Str.6, analysing samples by
FISH, freshly collected twice a week. FISH studies showed
that this community was only found within free-floating
white-greenish streamers, but not in microbial mats, water
samples or sediments.
Within the community, filamentous Bacteria represented
the dominant morphotype, often showing sulphur granules
within the cells. These microorganisms were classified as
Thiothrix sp. using a Thiothrix-specific FISH probe (TN1
Wagner et al., 1994). The second partner, tentatively named
‘Sip100’, was also a filament-forming bacterium, which
attached perpendicular to the Thiothrix filaments. For
further characterization of ‘Sip100’, FISH with different
specific probes for the Proteobacteria (ALF968, Neef, 1997;
Bet42a, GAM42a, Manz et al., 1992; DELTA495a/b/c, Loy
et al., 2002) and CFB (CFB319a/b Manz et al., 1996) groups,
which are often main constituents in such biotopes, was
performed. However, none of these probes produced a
positive FISH signal. The third organism in the community
was a small coccus/bacillus (diameter about 0.5–0.7 mm),
which was attached to ‘Sip100’ and gave a positive hybridization signal with a crenarchaeal FISH probe (Cren499
Burggraf et al., 1994). This novel crenarchaeon is tentatively
named ‘Cre1’.
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Probe design
To verify the phylogenetic affiliation of the new organism
‘Cre1’ to the Crenarchaeota, a second Crenarchaeota-specific
FISH probe was designed on the basis of primer Cren457R
(Schleper et al., 1997) using the ‘Probe_Design’ function of
the ARB program (Ludwig & Strunk, 1997). The specificity of
the probe was tested in several FISH experiments with
different genera of Euryarchaeota and Crenarchaeota (Table
1). To obtain optimal hybridization signals, varying formamide concentrations were tested (Olsen et al., 1986; Amann
et al., 1995). The strongest hybridization signals were
obtained using 100–150 ng mL1 probe per spot, 0.01%
sodium dodecylsulphate (SDS) and 25% formamide in the
hybridization buffer. This new probe was successfully applied in FISH experiments with original samples containing
the new crenarchaeal/bacterial community. The results are
shown in Table 6.
Table 5. Chemical and physical parameters of the two sampling sites
Str.6 (Sippenauer Moor; Fig. 1) and Islinger Mühlbach
Sampling site
Sippenauer Moor
0
Site location
Temperature ( 1C)
pH
Conductivity (mS cm1)
Sulphide (mg L1)
Oxygen (mg L1)
Flow speed (m s1)
Sulphate (mg L1)
Sulphite, nitrate (mg L1)
Nitrite (mg L1)
Ferrous iron (mg L1)
N 48152.161
E 11157.355 0
8.9 (wintertime)
12.6 (summertime)
6–6.5
520
0.1–0.2
3.0
0.06–0.16
o 200
o 10
o2
1
Islinger Mühlbach
N 48159.140 0
E 12107.632 0
9.5 (wintertime)
11.5 (summertime)
6–6.5
600
0.1
1.3
n.d.
n.d.
n.d.
n.d.
n.d.
n.d., not determined.
Table 6. Hybridization signals of the newly discovered crenarchaeon
(‘Cre1’) using different hybridization probes
Probe
Hybridization signal of
the new crenarchaeon
Reference
Arch-Mix
Arch344
Arch915
Arch1060
Cren499
Cren457R
Eury498
Aqui542
111
111
11
1
11
Moissl et al. (2003)
Moissl et al. (2003)
Stahl & Amann (1991)
Moissl et al. (2003)
Burggraf et al. (1994)
This work
Burggraf et al. (1994)
Huber (1998)
, no fluorescence in situ hybridization (FISH) signal; 1, satisfactory FISH
signal; 11, strong FISH signal; 111, very strong FISH signal.
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The life cycle of the new crenarchaeal/bacterial
community
The use of the newly designed in situ cultivation system
provided the basis to shed light on the life cycle of the
specific crenarchaeal/bacterial community in time and
space. After about 1 week of incubation of the in situ
cultivation system in the streamlet of the Sippenauer Moor,
it became whitish coloured as a result of the growth of
filamentous Bacteria belonging to the genus Thiothrix (Fig.
4a, green arrow). By phase contrast microscopy, it became
evident that Thiothrix was sticking to the agar noodles as a
result of its distal holdfast material (Larkin & Nelson, 1987).
After 2 weeks of incubation, the first rod-shaped Bacteria,
designated ‘Sip100’, attached to the Thiothrix filaments (Fig.
4b, green arrows) in a characteristic perpendicular orientation (Fig. 4b, white arrows). Three to five days later, the
number of ‘Sip100’ Bacteria had increased. Afterwards, they
divided and formed short filaments with up to five cells (Fig.
4c and d, white arrows). After 3–3.5 weeks of incubation, the
first Crenarchaeota ‘Cre1’ attached to the short ‘Sip100’
filaments (Fig. 4e, red arrows). Within another week, the
Crenarchaeota proliferated enormously, and almost all rodshaped ‘Sip100’ cells became infested with one to ten
crenarchaeal cells (Fig. 4f, red arrows). During multiplication, the almost coccoid-shaped Crenarchaeota (+ =
0.5–0.7 mm) increased by up to 3–5 mm in length into rods,
which remained attached to the ‘Sip100’ cells at a characteristic 901angle. Afterwards, the rods fragmented at the free
end into up to four coccoid ‘daughter’ cells simultaneously.
Thereby, the most distal crenarchaeal cells were obviously
released and possibly attached again to the same or another
cell of ‘Sip100’ (Fig. 5). After 4–5 weeks of incubation, the
number of Crenarchaeota and ‘Sip100’ decreased almost to
zero within only 1–2 days and the agar noodles became
brownish as a result of the growth of various algae. At this
point, the experiment was stopped. The whole cultivation
experiment was repeated four times over a period of about
18 months with reproducible results.
Phylogenetic analyses
In a FISH-independent study, the crenarchaeal 16S rRNA
gene pool of white-greenish streamers from sampling site
Str.6 at Mineralbach of Sippenauer Moor was analysed. The
DNA showed a brownish colour after isolation as a result of
contamination with humic acids, which inhibited PCR
amplifications. A modified, fast and reliable DNA purification method was developed, and archaeal and crenarchaeal
PCR products were obtained with all primer combinations
tested, and four products were cloned (Table 4).
Based on RFLP analyses, 26 clones were sequenced
(421–1082 bp) in total and all sequences belonged to the
Crenarchaeota. None of the sequences was judged to be
FEMS Microbiol Ecol 57 (2006) 55–66
61
Cold-loving crenarchaeon in cold sulphidic marsh water
(a)
(c)
5 µm
5 µm
(b)
(d)
5 µm
5 µm
(e)
(f)
5 µm
5 µm
Fig. 4. Growth cycle of the novel crenarchaeal/bacterial community at Str.6, analysed by fluorescence in situ hybridization (FISH). (a–f) Epifluorescence
micrographs. Hybridization was performed with a rhodamine green-labelled, Archaea-specific probe mix and Carbocyanine-labelled, Bacteria-specific probes.
The new Crenarchaeota stains green (red arrows), the bacterium ‘Sip100’ appears red (white arrows) and the Thiothrix cells stain orange (green arrows).
chimeric (CHECK-CHIMERA; Ribosomal Database Project
Maidak et al., 2000). Phylogenetic analysis showed that all
sequences were spread in ‘marine group I’ and the ‘freshwater and marine benthic group’ (Vetriani et al., 1998,
1999). Depending on the primer combination, between 26
and 51% of the sequences belonged to one single, dominating phylotype (clone Str.6_6/K12) clustering within marine
group I. The phylogenetic position of all clone sequences
FEMS Microbiol Ecol 57 (2006) 55–66
was verified by three different tree reconstruction methods.
The maximum parsimony phylogenetic tree is shown in
Fig. 6.
Phylogenetic analysis of a white-greenish streamer from
sampling site Str.6, performed 1 year later, showed the same
dominance of the identical crenarchaeal sequence Str.6_6/
K12 (52% of all clones analysed). The other derived 16S
rRNA gene sequences clustered together in the freshwater
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62
M. Koch et al.
about 10–15 m in length within the Mineralbach at site Str.6
(Fig. 1). However, the community has been reliably detected
in this small region for about 4 years. The association is very
low in number (up to one crenarchaeon vs 104–106 freeliving Bacteria in total) and often appears in clusters with up
to 100 crenarchaeal cells.
(a)
Distribution of the new community in another
sulphidic spring
5 µm
(b)
For ecological studies, samples from a sulphidic spring at the
Islinger Mühlbach (Rudolph et al., 2004) were taken and
analysed via FISH using Bacteria-, Archaea- and Crenarchaeota-specific probes. FISH studies revealed that most
samples consisted almost exclusively of Bacteria, with filamentous morphotypes predominating. In addition, a very
small number of rod-shaped Crenarchaeota could be detected, which were attached to Bacteria at a characteristic
901 angle. These Bacteria were again associated perpendicular to filamentous Bacteria of the genus Thiothrix, which
showed sulphur granules inside the cells. This prokaryotic
cell association is highly reminiscent of the crenarchaeal/
bacterial community from Str.6 of Sippenauer Moor.
Discussion
5 µm
Fig. 5. Fluorescence in situ hybridization (FISH) and 4 0 ,6-diamidino-2phenylindole (DAPI) staining of the novel crenarchaeal/bacterial community at Str.6. (a) Epifluorescence micrograph. Hybridization was performed
with a rhodamine green-labelled, Archaea-specific probe mix and carbocyanine-labelled, Bacteria-specific probes. The new Crenarchaeota stains
green (red arrows), the ‘Sip100’ cells appear red (white arrows) and the
Thiothrix filaments stain green (green arrows). (b) DAPI stain; the fragmented Crenarchaeota are marked with red arrows, the ‘Sip100’ cells
with white arrows and the Thiothrix filaments with green arrows.
and marine benthic group with no close phylogenetic
relationship to already published crenarchaeal sequences.
Distribution of the new crenarchaeal/bacterial
community within Sippenauer Moor
From about 500 samples of different coloured streamers,
mats, water and sediment taken from Mineralbach at intervals of about 200 m, the crenarchaeal/bacterial community
was detected via FISH in only a few samples, composed of
white-greenish streamer material. Interestingly, the growth
of the community is obviously limited to a small region of
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A unique, characteristically formed prokaryotic community
was detected in the cold (c. 10 1C) sulphurous marsh water
of the Sippenauer Moor (Bresinsky, 1991, 1999, 2001). The
combination of environmental observations, the design and
use of a new in situ cultivation technique, data from phase
contrast microscopy and FISH has enabled an initial interpretation to be made of the microbial architecture and
growth behaviour of this unusual microbial community.
So far, three groups of microorganisms have been identified, which contribute significantly to the association structure. One group is represented by filamentous Bacteria
belonging to the genus Thiothrix, which seem to play the
initial role in the formation of this unique microbial
community. The whitish coating of the new in situ cultivation system may be the result of the presence of elemental
sulphur, formed by the oxidation of sulphide during the
growth of this filamentous organism (Bland & Staley, 1978).
The second group is represented again by a bacterium
tentatively named ‘Sip100’, which attaches nearly perpendicular to the Thiothrix filaments and can grow out to short
filaments. A more detailed phylogenetic affiliation of
‘Sip100’ to the Proteobacteria or to the CFB group via FISH
with specific probes was not successful. ‘Sip100’ either does
not belong to these groups, has sequence variations in the
regions of probe binding or inaccessible target sites as a
result of the higher order structure of the ribosome (Amann
et al., 1995; Fuchs et al., 1998; Wagner et al., 2003). A novel
crenarchaeon represents the third group present in this
FEMS Microbiol Ecol 57 (2006) 55–66
63
Cold-loving crenarchaeon in cold sulphidic marsh water
Fig. 6. Phylogenetic tree based on 16S rRNA
gene sequences derived from DNA extracted
from white-greenish streamers from sampling
site Str.6. The tree was constructed using the
maximum parsimony method. The scale bar
indicates 10 estimated substitutions per 100
nucleotides.
specific community, verified by the use of the Crenarchaeota-specific FISH probe Cren499 and the newly designed
probe Cren457R. These cells attach to the ‘Sip100’ filaments
as small cocci (+ = 0.5–0.7 mm), elongate to rods of 2–5 mm
in length (901 angle to ‘Sip100’) and fragment at the free end
into up to three to four daughter cells, which may attach
again to the same or another ‘Sip100’ bacterium. Free-living
Crenarchaeota were not found in the biotope. At the
moment, the biological relationship between Thiothrix sp.,
‘Sip100’ and the crenarchaeon remains unclear; however, a
variety of possibilities, such as symbiosis or parasitism, can
be envisaged (Stolp & Starr, 1963; Schink, 1992; Abella et al.,
1998).
During comparative studies of cold, sulphidic springs, we
discovered a second biotope (Islinger Mühlbach Rudolph
et al., 2004) harbouring morphologically very similar microbial associations between Crenarchaeota and two groups of
FEMS Microbiol Ecol 57 (2006) 55–66
rod-shaped filamentous Bacteria (including Thiothrix sp.).
Based on the morphology and structure of this prokaryotic
community, it can be assumed that the Crenarchaeota from
both biotopes are phylogenetically closely related. This is an
interesting result considering the geographical and spatial
separation of the two sampling sites (Rudolph et al., 2004).
The occurrence of Crenarchaeota was proven with the
FISH-independent method of environmental 16S rRNA
gene sequence analysis. This study represents the first report
of crenarchaeal 16S rRNA gene sequences occurring in the
cold, sulphidic water of the Sippenauer Moor. Using a
modified DNA purification method, humic acids could be
removed rapidly and effectively and PCR products were
obtained. Using different primer combinations, the most
commonly occurring RFLP pattern from all cloning attempts showed the same 16S rRNA gene sequence clustering
within the Crenarchaeota. The sequence could be classified
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64
within ‘marine group I’ (Vetriani et al., 1998, 1999), which
contains 16S rRNA gene sequences from marine biotopes
(DeLong et al., 1994; Preston et al., 1996; Massana et al.,
1997; Maarel et al., 1998), as well as sequences from freshwater sediments (McGregor et al., 1997). In addition, we
obtained many more different crenarchaeal 16S rRNA gene
sequences which all belonged to the ‘freshwater and marine
benthic group’ (Vetriani et al., 1998, 1999). This group
predominantly contains representatives from freshwater
sediments (Hershberger et al., 1996) and paddy fields
(Großkopf et al., 1998).
Our investigations show that a novel, cold-loving crenarchaeon thrives in the cold, sulphurous water of the
Sippenauer Moor and the Islinger Mühlbach, and lives in
close association with two specific members of the bacterial
domain. As a result of the constant water temperature of
Mineralbach throughout the year, this observation provides
the strong evidence that this crenarchaeon, whose closest
cultivated relatives are thermophiles or hyperthermophiles,
grows at low temperatures. Actively dividing Crenarchaeota
cells could be found in white-greenish streamers as well as
on the new in situ cultivation system. The results from the
new in situ cultivation system reported in this paper are the
prerequisite for culture attempts employing enrichment
cultures and ‘optical tweezers’ as described recently (Huber
et al., 1995; Huber, 1999).
The newly discovered crenarchaeal/bacterial community
provides a significant opportunity for further studies, which
may lead to the identification of interesting physiological
and ecological insights in such close microbial relationships.
The community represents a hot spot for a novel crenarchaeon, which is permanently present in the sulphurous
streamlet of the Sippenauer Moor.
Acknowledgements
We are indebted to the Government of Bavaria (Germany)
for a sampling permit. Financial support from the Deutsche
Forschungsgemeinschaft (Hur 711/2) is grateful acknowledged.
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