Hindawi Publishing Corporation
Archaea
Volume 2013, Article ID 102972, 8 pages
http://dx.doi.org/10.1155/2013/102972
Research Article
Deposition of Biogenic Iron Minerals in a Methane
Oxidizing Microbial Mat
Christoph Wrede,1,2 Sebastian Kokoschka,1 Anne Dreier,1,3 Christina Heller,4,5
Joachim Reitner,3,4 and Michael Hoppert1,4
1 Institute of Microbiology and Genetics, Georg-August-University, Grisebachstr. 8, 37077 Go¨ttingen, Germany
2Hannover Medical School, Institute of Functional and Applied Anatomy, Carl-Neuberg-Str. 1, 30625 Hannover, Germany
3 Courant Centre Geobiology, Georg-August-University, Goldschmidtstr. 3, 37077 Go¨ttingen, Germany
4Geoscience Centre Go¨ttingen, Georg-August-University, Goldschmidtstr. 3, 37077 Go¨ttingen, Germany
5 Federal Institute for Geosciences and Natural Resources, Stilleweg 2, 30655 Hannover, Germany
Correspondence should be addressed to Christoph Wrede; cwrede@gwdg.de
Received 18 September 2012; Revised 13 April 2013; Accepted 20 April 2013
Academic Editor: Charles Cockell
Copyright © 2013 Christoph Wrede et al. This is an open access article distributed under the Creative Commons Attribution
License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly
cited.
The syntrophic community between anaerobic methanotrophic archaea and sulfate reducing bacteria forms thick, black layers
withinmulti-layeredmicrobialmats in chimney-like carbonate concretions ofmethane seeps located in the Black SeaCrimean shelf.
The microbial consortium conducts anaerobic oxidation of methane, which leads to the formation of mainly two biomineral by-
products, calcium carbonates and iron sulfides, building up these chimneys. Iron sulfides are generated by the microbial reduction
of oxidized sulfur compounds in the microbial mats. Here we show that sulfate reducing bacteria deposit biogenic iron sulfides
extra- and intracellularly, the latter in magnetosome-like chains. These chains appear to be stable after cell lysis and tend to attach
to cell debris within the microbial mat. The particles may be important nuclei for larger iron sulfide mineral aggregates.
1. Introduction
Frequently, biofilm formation in marine and freshwater
systems is accompanied by precipitation of minerals. These
minerals are also structurally integrative parts of the micro-
bial biofilm [1]. In most cases, mineral precipitates are
deposited in close contact to and in interaction with organic
macromolecules, that is, carbohydrates and/or proteins [2].
Formation of a biomineral in a microbial biofilm may be
detrimental to the organisms which is mainly due to the
enclosure of the living biomass by mineral precipitates. How-
ever, also positive effects, for example, when lithified precip-
itates provide a matrix or scaffold for the microbial biomass,
may be expected. It has also been considered that beneficial
effects predominate, for example, when biominerals act as
chemical filters or shield UV radiation [3]. It is known that, in
certain cases, biological macromolecules influence solubility
of minerals (e.g., by buffering the aqueous environment or
by chelating ions) and may direct the formation of a mineral
matrix in a more or less specific way. As a consequence, the
shape of biomineral deposits varies considerably at narrow
scales and seemingly similar environmental conditions [4].
Mineral deposits caused by the activity of microorgan-
isms are mostly based on either carbonates or silicates [4].
These mineral phases are regularly intermixed with other
organic or mineralic compounds (overviews in [2, 5]). A
special case of these organomineral precipitations is micro-
bialite formation during anaerobic oxidation of methane
(AOM). AOM is conducted by various groups of archaea in
a metabolic pathway reverting methanogenesis [6]. Mostly,
sulfate reducing bacteria (SRB) participate in AOM [7–9].
The role of SRB is still not fully understood, though is
generally accepted that, along with the oxidation of methane,
sulfate is reduced: CH
4
+ SO
4
2−
→ HCO
3
−
+ HS− + H
2
O
[8, 10]. As a result, carbonate phases (calcite and aragonite)
and iron sulfides are generated as byproducts of themetabolic
process.
2 Archaea
It is is known that AOM occurs worldwide in anoxic sed-
iments when methane and electrons acceptors are available
(e.g., [11]). The formation of large (several centimeters and
bigger) carbonate concretions depends on highmethane con-
centrations under hydrostatic pressure and on the presence
of sulfate [8, 9, 12]. In the anaerobic water column of the
Black Sea, huge carbonate concretions have been observed
at the Crimean shelf [8]. The carbonate buildups may be
considered as highly porous “fixed bed” bioreactors, allowing
the percolation of methane and the exchange of sea water.
The outer and inner surfaces of these carbonate buildups are
covered by complex microbial mats, primarily formed by the
organisms involved in AOM.
In previous investigations, distinct layers in these micro-
bial mats were discriminated. On the surface, exposed to
the sea water, a black layer consists mainly of aggregates
between methane-oxidizing archaea of the ANME-2 group
and sulfate reducing bacteria (SRB). SRB of this mat type
often exhibit intracytoplasmic magnetosome-like chains of
greigite precipitations [8, 13, 14]. Our results imply that
greigite magnetosomes are one sink for (otherwise toxic)
sulfides. These particles were found inside SRB but were also
present in the extracellular matrix of the biofilm.
2. Materials and Methods
Microbial mat samples were collected in 2001 during a cruise
with the Russian R/V Professor Logachev in the methane
seep area located in the GHOSTDABS field (Black Sea
north east the Crimean shelf). These samples have already
been subjected to extended geochemical and structural anal-
yses [9, 14]. Specific antibodies, directed against methyl-
coenzyme M reductase (MCR), the key enzyme of (reverse)
methanogenesis, were generated after purification of MCR
as essentially described according to [15] by immunization
of rabbits following established protocols (e. g., [16] and, the
references therein). Specificity of the antibodywas extensively
studied for methanogenic archaea and reverse methanogens
as already described [14, 16, 17].
For microscopic analyses, the samples were chemically
fixed in a 4.0% (v/v) aqueous formaldehyde solution (from
a 10%, w/v, stock solution, pH 8.0, freshly prepared from
paraformaldehyde) and stored in 100mM PBS (phosphate
buffered saline, pH 7.0) at 4∘C until further use. The material
was then washed several times in PBS and cut to small
fragments of about 200𝜇L volume. Samples were then chem-
ically fixed in a 0.5% (v/v) glutardialdehyde solution (in
100mM PBS) for 2 h. The samples were then processed as
described [18], for electron and light microscopy, and finally
cut in ultrathin or semithin sections of either 100–300 nm
or 1 𝜇m in thickness. Semithin sections were transferred,
with the aid of a transfer loop, on microscope slides, and
ultrathin sections were picked up with Formvar-coated grids.
For light microscopy, the sections were treated either with
an anti-MCR antibody or with the lectin concanavalin A
(ConA) coupled to fluorescent marker molecules (Sigma-
Aldrich, Deisenhofen, Germany) as described [18].The lectin
ConA IV coupled to Alexa Fluor 546 as fluorescence marker
(Molecular Probes, Eugene, OR, USA) was used in 1/1000
working concentration dilutions in PBS supplemented with
1mM CaCl
2
and MnCl
2
(lectin buffer). The sections were
mounted on glass slides by heat fixation at 60∘C for 15min
and then incubated for 30min at room temperature. After
this, the lectin dilution was soaked off, and the sections
were briefly rinsed in pure lectin buffer and covered with
coverslips. For immunofluorescence microscopy, the heat-
fixed semithin sections were incubated with the antiserum
(dil. 1/1000 with PBS, pH 7.5) for 2 h. The sections were
rinsed three times in 100𝜇L drops of PBS (supplementedwith
0.01% Tween 20) and incubated with a secondary goat anti-
rabbit antibody, coupled to Alexa Fluor 546 fluorescent dye
(Molecular Probes, Eugene, OR, USA), diluted 1 : 250 [14].
The rinsing steps were repeated. Fluorescence microscopy
was performed with an Axio Scope light microscope using
filter set 43 (BP: 545/25, FT 570, LP: 605/70) and the
AxioVision software package (Zeiss, Go¨ttingen, Germany).
For comparison, phase contrast images were taken and were
digitally merged with fluorescence images.
For transmission electron microscopy, ultrathin sec-
tions obtained from five distinct samples were mounted
on Formvar-coated 300 mesh specimen grids. Immunolo-
calization with antibodies directed against MCR was per-
formed as described. Mounted sections were stained with
phosphotungstic acid (3%, w/v), if not stated otherwise [14].
Electron microscopy was performed in a Zeiss EM 902
transmission electron microscope (Zeiss, Oberkochen, Ger-
many), equipped with a eucentric goniometer stage. Images
were recorded with a 1 KB digital camera. Detection and
enhancement of colloidal gold markers in digitized electron
micrographs were performed as described [19]. Electron
energy loss spectroscopy (EELS) of iron-containing particles
was performed essentially as described in [20], with the aid
of the analysis V software package (Olympus-SIS, Mu¨nster,
Germany). Electron energy loss was measured between
655 eV and 751 eV. The L2 (720.6 eV) and L3 (708.0 eV)
edges, observed for pure iron minerals, were represented by
one broad peak at approximately 715 eV of the deposits in
embedded and ultrathin sectioned biofilms.
Goniometry was performed by tilting 300 nm sections
±60 degrees with 1-degree increments. Tomograms were
performedwith the EM3D2.0 software package (Department
of Neurobiology, Stanford University [21]).
3. Results and Discussion
Two layers of microbial mats retrieved from the the Black Sea
Crimean shelf have been identified as important for AOM:
the orange (or pink) layer and the black layer. The orange
layer consists of various cell morphotypes [8, 22]. Most of
them were identified as ANME-1 archaea. ANME-1 cells
are morphologically similar to the filamentous methanogens
Methanospirillum and Methanosaeta; these cells are covered
by a tight and very rigid protein sheath [23]. Sulfate reducing
bacteria were present in large clusters, but not in direct
contact with ANME-1 cells [8]. Visually, the black layer
could be clearly distinguished from other layers. In contrast
Archaea 3
20 𝜇m
(a)
20 𝜇m
(b)
20 𝜇m
(c)
Figure 1: Light micrographs of semithin sections after immunofluorescence staining with anti-MCR antibodies (merged fluorescence/phase
contrast images). (a) One large, cauliflower-like aggregate (lower right, the whole aggregate marked by three asterisks) surrounded by small
globular aggregates (arrows) in an unstained semithin section (phase contrast microscopy). (b) Periphery of a large aggregate.The cell density
at the periphery of the aggregate (arrows) is higher than in the central area (asterisks). (c) Small aggregates after fluorescence staining.
to the orange layer, the black layer consists of aggregates
formed by ANME-2 and SRB of the DSS group [8, 24, 25].
Immunofluorescence labelling of MCR, performed on resin
sections, marks the position of the MCR-expressing ANME-
2 inside large cauliflower-shaped aggregates, consisting of
thousands of cells visible in a section (Figure 1(a), asterisks,
Figure 1(b)). Similar labelling experiments have been also
performed on the same samples with antibodies directed
against the dissimilatory adenosine-5󸀠-phosphosulfate (APS)
reductase, along with the identification of the respective gene
in the microbial mat samples [14]. APS reductase is a key
enzyme of sulfate reduction and could be localized in the
magnetosome-bearing cell type (see below), identified as
SRB.
In addition to these large cauliflower-shaped aggregates,
also a smaller globular-shaped aggregate type of 5–20 labeled
ANME-2 cells was identified (Figure 1(c)). This aggregate
type is located in the surrounding (Figure 1(a), arrows) and in
the center of the cauliflower-shaped features. All aggregates
are separated by areas of low cell densities. It has to be
noted that not all cells visible in the depicted sections are
labelled, since markers do only bind to cells with their
cytoplasm exposed to the section surface. In the large type
of aggregates, cells are not completely randomly distributed;
higher cell densities are observable at the periphery, sepa-
rated in irregular lobes (Figure 1(b), arrows). Both types of
aggregates consist of ANME-2/SRB consortia [8, 14, 22]. The
electron micrographs in Figure 2 show ANME-2 (immuno-
gold labelled) and SRB (nearly unlabeled) in a globular
aggregate, surrounded by multiple layers of extracellular
material. The gaps between the large aggregates are filled
with EPS [18]. These gaps show a distinct lectin labelling
(Figure 3(a), arrows), in contrast to the unlabeled extracel-
lular surrounding of the ANME-2/SRB consortia. Various
morphotypes of prokaryotic cells could be detected in these
empty spaces (Figure 3(b), cf. [14]), including thin filaments
of several 𝜇m in length and 200 nm in diameter (arrows
in Figure 3(b)). Some filaments still contain a dark stained
4 Archaea
2 𝜇m
(a)
0.5 𝜇m
(b)
Figure 2: Immunoelectron microscopy. (a) Electron microscopy of a typical small aggregate (cf. Figure 1(c)), consisting of ANME-2/SRB.
Several single cells are surrounded by a thick multilayered mucilage. (b) Detail of the aggregate as depicted in (a). The MCR expressing
ANME-2 cells are labelled with small gold dots (black arrows point to some dots); the SRB (upper right cell) show a low background labelling
(black arrows).
20 𝜇m
(a)
5𝜇m
(b)
Figure 3: Appearance of gaps between large aggregates. (a) Overview: large patches outside aggregates show intense fluorescence (arrows)
after staining with fluorescently labeled ConA lectin. (b) Electron micrograph of a 300 nm thick section from the aggregate periphery with
intact cells (left) and cell debris embedded in EPS (right). Arrows point to long filaments.
matrix, putatively cytoplasmic contents. Shallowly stained
filaments likely represent empty cell envelopes (Figures 4(b),
5(a), and 5(b)).
Within these large, cauliflower-shaped aggregates, SRB
exhibit peculiar cytological features. Apart from occasionally
observed intracytoplasmic membranes, intracellular magne-
tosome-like particles, arranged in straight rows and com-
posed of greigite, were observed frequently [8, 14]. Here, we
show that these chains appear to be stable after cell death
and cell lysis, but exhibit structural modifications. In intact
cells, chains are arranged in straight rows, mostly as paral-
lel pairs (Figure 4(b), inset, Figure 4(c)). Figure 4(c) shows
magnetosome chains in unstained sections; that is, the cells,
though still present and morphologically intact, are invisible
here. Basically, three variants of magnetosome-like chains
could be observed. These variants may represent three stages
of development. In a (putatively) early stage, chains appear
to be absent (Figure 4(a)). During aggregate development,
the organisms deposit intracellular chains (Figures 4(b) and
4(c)). Finally, the organisms get lysed, and in stained sections,
just the magnetosomes and some cell debris are still present;
the free chains still mark the positions of the SRB in the
aggregate (Figure 4(d)); distances between these deposits are
similar to the distances between magnetosome chains in
Archaea 5
2.5 𝜇m
0.5 𝜇m
(a)
10𝜇m
0.5 𝜇m
∗
∗
∗
∗
∗
(b)
0.5 𝜇m 2𝜇m
(c)
10𝜇m
200 nm
(d)
Figure 4: Aggregates in different stages of magnetosome-like chain formation. (a) ANME-2/SRB aggregate without visible precipitates. (b)
Periphery of a large aggregate (asterisk), with gap between aggregates (arrows). Cells with multiple magnetosome-like chains (inset; see
also (c)). Note cell debris, mainly consisting of envelopes from filamentous cells (arrows), outside the aggregates. (c) Periphery of an intact
aggregate as depicted in (b) (unstained section; cells are invisible), showing the position of straight magnetosome chains inside cells. (d)
ANME-2/SRB aggregate after cell lysis (cells are absent in spite of staining, compare (c)), with magnetosome chains still in place (some chains
are marked by arrows). The inset shows a single chain. The dotted line marks the border of the aggregate (right of the dotted line). Some
chains are found outside the area of the aggregate.
neighboured intact cells (Figure 4(c)). Chains of the same
size are also intermixed with the cell debris in the gaps
between the ANME-2/SRB aggregates; here they appear to
be attached to cell envelopes of filamentous morphotypes
(Figure 5(a)). Mostly, the chains exhibit curves and wrinkles,
perhaps due to the loss of their intracellular scaffolding struc-
ture (cf. [26]), though also straight chains are observable.
Figures 5(b) and 5(c) show particles of increased size, from
30 nm to up to 100 nm in diameter. The particles appear
to be attached to the filamentous morphotype (Figure 5(c)).
Larger agglomerations of these particles are found near
0.5 𝜇m sized microcrystals (arrows in Figure 5(d)), similar
to particles forming pyrite framboids (cf. [7]). It is unclear
if the greigite magnetosomes contribute, in the end, to
the formation of these crystals and/or framboidal pyrite
(e.g., [7, 27, 28]), but the contribution of free magnetosome
chains to the iron sulfide minerals in the black layer appears
to be obvious. In particular, the “reactive” surfaces of prokary-
otic cell envelopes are involved in binding and accumulation
of these particles. Figure 6 summarizes our observations and
proposes a schematic sequence of the observed features.
Active consortia may not contain magnetosome-like chains
in the beginning (a), but in most of the aggregates, well-
developed chains are present ((b) and (c)). The involvement
of thesemagnetosomes in chemotaxis appears to be doubtful,
since the SRB are immotile during all stages of biofilm
development. It may be speculated that the particles are,
in this case, an intracellular “dead end” storage granule,
accumulating iron sulfides as waste product from sulfate
reduction (cf. [29]). It has to be expected that not all reduced
6 Archaea
1𝜇m
(a)
0
50
100
710 720 730
(eV)
In
te
ns
ity
0.5 𝜇m
(b)
1𝜇m
(c)
1𝜇m
(d)
Figure 5: Extracellular magnetosome-like features. (a) Extracellular magnetosome chains (short arrow) bound to filamentous cell envelopes.
Occasionally, also intact organisms, filled with dark cytoplasm, are visible (long arrow). (b) Particles (arrows), double in size ofmagnetosomes
close or attached to filaments. The EELS spectrum (upper left inset) shows the energy loss at the Fe L2/L3 edges. The lower right insets
show a tomogram of a small section (encircled) as depicted in the lower left inset (blue: cell envelope, red: particles). The small dots
represent randomly distributed colloidal gold particles (no markers) necessary for image alignment of the tilted sections. (c) Overview image
showing aggregates of the particles adjacent to filamentous envelopes. (d) Aggregates of particles (arrowheads) and typical microcrystals from
framboidal pyrite (arrows).
sulfur compounds end up inside cells and, as known from
other sulfate reducing bacteria, sulfides are also deposited
outside cells. However, intracellular deposition may be also
a rapid way to keep the concentration of sulfides as low
as possible. It is obvious that both syntrophic partners die
and lyse (possibly all cells at the same time) leaving the
magnetosome chains as still visible remains (d) inside the
aggregates. Free chains (e) migrate, by diffusion in the matrix
outside the aggregate bind to a specific type of cell envelope
(f), and lose their chain-like appearance and regular size (g).
Conflict of Interests
The authors declare that they have no conflict of interests.
Archaea 7
Active ANME/SRB
Formation of 
greigite 
magnetosomes
Cell death 
and lysis
Free chains
Interaction
with 
envelopes
Particle growth 
(a)
(b)
(c)
(d)
(e)
(f)
(g)
SRB
ANME
Figure 6: Turnover of magnetosome-like chains in the black layer. See Section 3 for further explanation.
Acknowledgments
The authors thank the crew of the R/V Professor Logachev,
the Hamburg research group of Professor W. Michaelis, and
the Jago-Submersible Team (J. Schauer and K. Hissmann)
for collaboration during the sampling cruise for deep sea
sediments. The authors thank also Jo¨rn Peckmann (RCOM-
Bremen) for the helpful discussions. Part of this study
received financial support by the Bundesministerium fu¨r
Bildung und Forschung (BMBF-GEOTECHNOLOGIEN-
Program GHOSTDABS 03G0559A) and the Deutsche For-
schungsgemeinschaft (DFG-Research Unit 571—Geobiology
of Organo—and Biofilms, DFG Grants Re 665/31-1 and
Ho 1830/2-1). Anne Dreier is supported by a fellowship of
the Studienstiftung des Deutschen Volkes. The authors also
acknowledge the support by the Open Access Publication
Funds of theGo¨ttingenUniversity.This is a Courant Research
Centre Geobiology publication.
References
[1] R. V. Burne and L. S.Moore, “Microbialites: organosedimentary
deposits of benthic microbial communities,” Palaios, vol. 2, no.
3, pp. 241–254, 1987.
[2] A. W. Decho, “Microbial biofilms in intertidal systems: an
overview,”Continental Shelf Research, vol. 20, no. 10-11, pp. 1257–
1273, 2000.
[3] V. R. Phoenix and K. O. Konhauser, “Benefits of bacterial
biomineralization,” Geobiology, vol. 6, no. 3, pp. 303–308, 2008.
[4] D. S. S. Lim, B. E. Laval, G. Slater et al., “Limnology of Pavilion
Lake, B. C., Canada: characterization of a microbialite forming
environment,” Fundamental andApplied Limnology, vol. 173, no.
4, pp. 329–351, 2009.
[5] P. U. P. A. Gilbert, M. Abrecht, and B. H. Frazer, “The organic-
mineral interface in biominerals,” Reviews in Mineralogy and
Geochemistry, vol. 59, pp. 157–185, 2005.
[6] S. J. Hallam, N. Putnam, C.M. Preston et al., “Reversemethano-
genesis: testing the hypothesis with environmental genomics,”
Science, vol. 305, no. 5689, pp. 1457–1462, 2004.
[7] J. Peckmann, A. Reimer, U. Luth et al., “Methane-derived
carbonates and authigenic pyrite from the northwestern Black
Sea,”Marine Geology, vol. 177, no. 1-2, pp. 129–150, 2001.
[8] J. Reitner, J. Peckmann, M. Blumenberg, W. Michaelis, A.
Reimer, and V.Thiel, “Concretionary methane-seep carbonates
and associated microbial communities in Black Sea sediments,”
Palaeogeography, Palaeoclimatology, Palaeoecology, vol. 227, no.
1–3, pp. 18–30, 2005.
[9] J. Reitner, J. Peckmann, A. Reimer, G. Schumann, and V.
Thiel, “Methane-derived carbonate build-ups and associated
microbial communities at cold seeps on the lowerCrimean shelf
(Black Sea),” Facies, vol. 51, no. 1–4, pp. 66–79, 2005.
[10] J. T. Milucka, T. G. Ferdelman, L. Polereck et al., “Zero-valent
sulphur is a key intermediate in marine methane oxidation,”
Nature, vol. 491, no. 7425, pp. 541–546, 2012.
[11] K. U. Hinrichs and A. Boetius, “The anaerobic oxidation of
methane: new insights in microbial ecology and biogeochem-
istry,” inOceanMargin Systems, G. Wefer, D. Billet, D. Hebbeln,
B. B. Jørgensen, M. Schlu¨ter, and T. vanWeering, Eds., pp. 457–
477, Springer, Heidelberg, Germany, 2002.
[12] W. Michaelis, R. Seifert, K. Nauhaus et al., “Microbial reefs in
the black sea fueled by anaerobic oxidation ofmethane,” Science,
vol. 297, no. 5583, pp. 1013–1015, 2002.
[13] C. T. Lefe`vre, N. Menguy, F. Abreu et al., “A cultured greigite-
producing magnetotactic bacterium in a novel group of sulfate-
reducing bacteria,” Science, vol. 334, no. 6063, pp. 1720–1723,
2011.
8 Archaea
[14] C. Wrede, V. Krukenberg, A. Dreier, J. Reitner, C. Heller,
and M. Hoppert, “Detection of metabolic key enzymes of
methane turnover processes in cold seep microbial biofilms,”
Geomicrobiology Journal, vol. 30, no. 3, pp. 214–227, 2013.
[15] M. Hoppert and F. Mayer, “Electron microscopy of native and
artificial methylreductase high-molecular-weight complexes in
strain Go 1 and Methanococcus voltae,” FEBS Letters, vol. 267,
no. 1, pp. 33–37, 1990.
[16] I. J. Braks, M. Hoppert, S. Roge, and F. Mayer, “Structural
aspects and immunolocalization of the F420-reducing and non-
F420-reducing hydrogenases,” Journal of Bacteriology, vol. 176,
no. 24, pp. 7677–7687, 1994.
[17] M. Kru¨ger, A.Meyerdierks, F. O. Glo¨ckner et al., “A conspicuous
nickel protein in microbial mats that oxidize methane anaero-
bically,” Nature, vol. 426, no. 6968, pp. 878–881, 2003.
[18] C. Wrede, C. Heller, J. Reitner, and M. Hoppert, “Correlative
light/electron microscopy for the investigation of microbial
mats from Black Sea Cold Seeps,” Journal of Microbiology
Methods, vol. 73, no. 2, pp. 85–91, 2008.
[19] M.Ka¨mper, S. Vetterkind, R. Berker, andM.Hoppert, “Methods
for in situ detection and characterization of extracellular poly-
mers in biofilms by electron microscopy,” Journal of Microbio-
logical Methods, vol. 57, no. 1, pp. 55–64, 2004.
[20] R. Bauer, “Electron spectroscopic imaging: an advanced tech-
nique for imaging and analysis in transmission electron
microscopy,”Methods inMicrobiology, vol. 20, pp. 113–146, 1988.
[21] D. Ress, M. L. Harlow, M. Schwarz, R. M. Marshall, and U.
J. McMahan, “Automatic acquisition of fiducial markers and
alignment of images in tilt series for electron tomography,”
Journal of Electron Microscopy, vol. 48, no. 3, pp. 277–287, 1999.
[22] C. Heller, M. Hoppert, and J. Reitner, “Immunological localiza-
tion of coenzyme M reductase in anaerobic methane-oxidizing
archaea of ANME 1 and ANME 2 type,” Geomicrobiology
Journal, vol. 25, no. 3-4, pp. 149–156, 2008.
[23] T. J. Beveridge, G. D. Sprott, and P. Whippey, “Ultrastructure,
inferred porosity, and gram-staining character of Methanospir-
illum hungatei filament termini describe a unique cell perme-
ability for this archaeobacterium,” Journal of Bacteriology, vol.
173, no. 1, pp. 130–140, 1991.
[24] A. Boetius, K. Ravenschlag, C. J. Schubert et al., “A marine
microbial consortium apparently mediating anaerobic oxida-
tion methane,” Nature, vol. 407, no. 6804, pp. 623–626, 2000.
[25] L. Schreiber, T. Holler, K. Knittel, A. Meyerdierks, and R.
Amann, “Identification of the dominant sulfate-reducing bacte-
rial partner of anaerobic methanotrophs of the ANME-2 clade,”
Environmental Microbiology, vol. 12, no. 8, pp. 2327–2340, 2010.
[26] A. Scheffel, M. Gruska, D. Faivre, A. Linaroudis, J. M. Plitzko,
and D. Schu¨ler, “An acidic protein aligns magnetosomes along
a filamentous structure in magnetotactic bacteria,” Nature, vol.
440, no. 7080, pp. 110–114, 2006.
[27] J. Schieber, “Sedimentary pyrite: a window into the microbial
past,” Geology, vol. 30, no. 6, pp. 531–534, 2002.
[28] L. C.W.MacLean, T. Tyliszczak, P. U. P. A. Gilbert et al., “A high-
resolution chemical and structural study of framboidal pyrite
formedwithin a low-temperature bacterial biofilm,”Geobiology,
vol. 6, no. 5, pp. 471–480, 2008.
[29] M. Po´sfai, B. M. Moskowitz, B. Arato´ et al., “Properties
of intracellular magnetite crystals produced by Desulfovibrio
magneticus strain RS-1,” Earth and Planetary Science Letters, vol.
249, no. 3-4, pp. 444–455, 2006.