Hindawi Publishing Corporation
Archaea
Volume 2013, Article ID 920241, 7 pages
http://dx.doi.org/10.1155/2013/920241
Research Article
Localization of Methyl-Coenzyme M Reductase as Metabolic
Marker for Diverse Methanogenic Archaea
Christoph Wrede,1,2 Ulrike Walbaum,1,3 Andrea Ducki,1,4
Iris Heieren,1 and Michael Hoppert1,5
1 Institute of Microbiology and Genetics, Georg-August-Universita¨t Go¨ttingen, Grisebachstraße 8, 37077 Go¨ttingen, Germany
2Hannover Medical School, Institute of Functional and Applied Anatomy, Carl-Neuberg-Straße 1, 30625 Hannover, Germany
3 Behavioral Ecology and Sociobiology Unit, German Primate Center, Kellnerweg 4, 37077 Go¨ttingen, Germany
4 School of Veterinary and Biomedical Sciences, Murdoch University, 90 South Street Murdoch, WA 6150, Australia
5 Courant Centre Geobiology, Georg-August-Universita¨t Go¨ttingen, Goldschmidtstraße 3, 37077 Go¨ttingen, Germany
Correspondence should be addressed to Michael Hoppert; mhopper@gwdg.de
Received 21 September 2012; Accepted 9 January 2013
Academic Editor: J. Reitner
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.
Methyl-Coenzyme M reductase (MCR) as key enzyme for methanogenesis as well as for anaerobic oxidation of methane
represents an important metabolic marker for both processes in microbial biofilms. Here, the potential of MCR-specific polyclonal
antibodies as metabolic marker in various methanogenic Archaea is shown. For standard growth conditions in laboratory culture,
the cytoplasmic localization of the enzyme in Methanothermobacter marburgensis, Methanothermobacter wolfei, Methanococcus
maripaludis, Methanosarcina mazei, and in anaerobically methane-oxidizing biofilms is demonstrated. Under growth limiting
conditions on nickel-depleted media, at low linear growth of cultures, a fraction of 50–70% of the enzyme was localized close
to the cytoplasmic membrane, which implies “facultative” membrane association of the enzyme.This feature may be also useful for
assessment of growth-limiting conditions in microbial biofilms.
1. Introduction
Methyl-coenzyme M reductase (MCR) is the key enzyme
of the final, methane-forming step in methanogenesis.
The enzyme catalyses the reductive cleavage of methyl-
coenzyme M (CoM-S-CH
3
) using coenzyme B (HS-CoB)
as reductant which results in the production of methane
and the heterodisulfide CoM-S-S-CoB. Though the involved
enzyme complexes as well as the reactants differ between
Methanosarcinales, Methanobacteriales, and other groups of
methanogens, the essential reaction steps are similar and
require several membrane-dependent steps (see [1, 2] for
review). The formation of methyl-coenzyme M is catalysed
by one subunit (MtrE) of a membrane-bound complex (the
N5-methyl-tetrahydromethanopterin:coenzyme M methyl-
transferase) and is coupled with energy conservation via
an electrochemical sodium potential across the cytoplasmic
membrane (see [3] for review). Regeneration of the reductant
HS-CoB is brought about by the enzyme heterodisulfide
reductase. For the regeneration of HS-CoB, reducing equiv-
alents are needed, provided by hydrogenases and/or dehy-
drogenases. The reducing equivalents are either guided via
a membrane-bound electron transport chain to the enzyme
or are directly transferred from the hydrogenase to the
heterodisulfide reductase. The reactions are also coupled to
chemiosmotic mechanisms, resulting in the generation of
ATP via a H+-potential [4–6]. Like MtrE, the heterodisulfide
reductase is a part of a membrane-bound complex. The
methyl-coenzyme M reductase reaction step itself is not
membrane-dependent. The enzyme has been purified from
the cytoplasmic fractions of methanogenic Archaea and
has been localized in the cytoplasm by immunoelectron
microscopy. The catalytic reaction does not depend on
the addition of membrane preparations [7–11]. A number
of experiments, however, indicate that there is a certain
affinity of the enzyme to the membrane [12, 13]. MCR
2 Archaea
M
t m
b∗
M
t m
b
M
tw
o
M
cm
a
M
sf
r
M
sm
a
94
67
43
30
   20
(k
D
a)
M
t m
b-
N
i
M
t w
o-
N
i
M
sm
a-
N
i
Figure 1: Specificity of the polyclonal serum used for immunolo-
calization. The slots depict crude extracts of the organisms after
Western blotting of SDS gels and double-immunoperoxidase pre-
cipitation. All slots show the typical pattern of MCR. For most
organisms (except Ms ma and Mt mb), only the two larger of
the three MCR subunits are visible. When cells were grown on
nickel-depleted media, the respective slot is marked with -Ni. The
second slot (marked with an asterisk) shows a silver stained SDS-
polyacrylamide gel of the purified enzyme. Mt mb: Methanother-
mobacter marburgensis, Mt wo: Methanothermobacter wolfei, Mc
ma:Methanococcusmaripaludis,Ms fr:Methanosarcinamazei (DSM
3318, formerlyMethanosarcina frisia),Msma:Methanosarcinamazei
(DSM 3647).
of Methanothermobacter marburgensis was located at the
cytoplasmic membrane under nickel-depleted growth condi-
tions. Also electron microscopy of vesicle preparations from
Methanobacteriales andMethanosarcina showed that at least a
fraction of MCR is membrane-associated. From these data, it
was deduced that MCR might be part of a membrane-bound
multienzyme complex [14, 15].
For the reverse process, the anaerobic oxidation of
methane, a reverse operating methanogenic pathway has
been postulated, with anMCR structurally very similar to the
canonical enzyme [16–18]. In the postulated pathway, again,
membrane binding is not necessarily required. However, as
in methanogenesis, membrane association might also be of
advantage, since the samemembrane-dependent processes as
in methanogenesis are likely [17, 19].
In Methanobacterium thermoautotrophicum, two differ-
ent localizations of the MCR could already be shown [13]. In
our study, we show that these results are also true for other
methanogens, and we will discuss these results in view of
immunolocalization of the key enzyme MCR for studies in
environmental biofilms.
2. Materials and Methods
Methanothermobacter marburgensis (DSM 2133, formerly
Methanobacterium thermoautotrophicum, strain Marburg),
Methanothermobacter wolfei (DSM 2970, formerly Metha-
nobacterium wolfei), and Methanococcus maripaludis (DSM
2067) were grown autotrophically as described [20–23].
Methanosarcina mazei (DSM 3318, formerly Methanosarcina
Table 1: Partitioning of MCR as revealed by immunolocalization.
Organism
Approximate
doubling
time (h)
Concentration
of levulinic acid
in the medium
%markers
at the
membrane
Methanothermobacter
marburgensis
26 0.0 17
35 0.05 36
38 0.2 70
Methanothermobacter
wolfei
34 0.0 15
42 0.1 52
Methanosarcina
mazei (DSM 3647)
20 0.0 32
34 0.05 60
frisia) and Methanosarcina mazei (DSM 3647) were grown
heterotrophically [24, 25]. Nickel-limited media did not
contain nickel salts in trace element solutions and were
supplemented with up to 200mM levulinic acid (cf. Table 1).
For immunolocalization, cells were grown in batch cultures
at linear growth rates with approximate doubling times
between 25 and 45 h (Table 1). Cell disruption was performed
with a French pressure cell operated at 1,500 lb/in2 and
subsequent centrifugation by 15,000×g for 25min at 4∘C in
order to remove cell debris. The supernatant was used for
Western-blotting (see below). For protein purification, cells
of Methanothermobacter marburgensis were grown in 14 l-
fermenters with a doubling time of 2.9 h in the exponential
phase on mineral salt medium and continuous gassing with
H
2
/CO
2
(80%/20%, v/v) as described [20]. Purification of
MCR was performed according to [7]. The purified protein
(MCR, i.e. the isoform I of methyl-coenzyme M reductase,
Figure 1) was used for production of polyclonal antisera [26].
Protein purity and specificity of the antisera was tested by
SDS polyacrylamide gel electrophoresis andWestern blotting
[27–29] and by immunolocalization control experiments (see
below, [30]). Protein assays were performed according to [31].
Samples of an environmental methane-oxidizing biofilms
were obtained and processed as described [32, 33]. Microbial
mat samples were collected in 2001 during a cruise with the
Russian R/V “Professor Logachev” from the methane seep
area located on the NW’ Shelf region (Crimean Shelf) in
the Black Sea. Material for transmission electron microscopy
and immunofluorescence analyses was chemically fixed in a
4.0% (w/v) formaldehyde solution and kept at 4∘C in 100mM
PBS (phosphate-buffered saline, pH 7.0). The samples were
washed several times in PBS and fixed in 0.3% (v/v) solution
of glutardialdehyde and 0.5% (w/v) formaldehyde in PBS for
2 h at 4∘C.The samples were then washed three times in PBS
supplemented with 10mM glycin. See below for subsequent
dehydration and resin embedding.
Active cultures were chemically fixed anaerobically by
adding 0.2% (v/v) solution of glutardialdehyde and 0.3%
(w/v) formaldehyde to the active culture under anaerobic
conditions. After incubation for 2 h at 4∘C, the culture was
centrifuged three times for 10min at 9.000×g and resus-
pended in PBS supplemented with 10mM glycin. Molten
agar (2%, w/v, 50∘C) was added to an equal volume of the
Archaea 3
400 nm
(a)
400 nm
(b)
200 nm
(c)
200 nm
(d)
Figure 2: Ultrathin sections ofMethanosarcina mazei (DSM 3318, formerlyMethanosarcina frisia (a, b) and Methanococcus maripaludis (c,
d) grown on media without nickel depletion. Immunolabeled cells (b, d; a, c are negative controls) show cytoplasmic localization of MCR.
The inset in (b) shows an enlarged area of the image before (open star symbol) and after processing (closed star) of the gold marker.
resuspended pellet. After mixing thoroughly, the sample was
allowed to solidify.
Subsequently, biofilm samples and agar-embedded cul-
ture samples were dehydrated. For dehydration, an ascending
methanol series was used [30]: 15% (v/v), 30% for 15min,
50%, 75% for 30min, 90%, and 100% for 1 h.The temperature
was successively lowered down to −35∘C (steps: 15%, 30% at
0∘C, 50% at −20∘C, and all other steps at −35∘C). Samples
were then incubated in Lowicryl K4M resin dilutions in
methanol (1 : 3, 1 : 2, and 3 : 1; Lowicryl resin obtained from
Electron Microscopy Sciences, Hatfield, PA, USA), followed
by incubation in pure resin for 1 h per step and then overnight.
The blocks were transferred to small gelatin capsules (Plano,
Wetzlar, Germany) containing pure resin and were polymer-
ized for at least 48 h at −35∘C and 3 d at room temperature
under UV light.
Ultrathin sections of trimmed specimens (80–90 nm)
were cut with glass knives in a Reichert Jung FC 4 ultrami-
crotome (Leica Microsystems, Wetzlar, Germany). Sections
were transferred onto Formvar-coated grids [30].
For TEM immunocytochemistry, grids were placed with
sections facing downwards, for 30min on drops of 3% (w/v)
bovine serum albumin (BSA) in PBS, then for 2 h on anti-
MCR antibodies (1mg protein/mL; dilution in PBS). Negative
controls were performed by incubation of the grids on PBS
without the antibody.
Grids were then washed by incubation (two times 5min)
on drops of PBS containing 0.05% (v/v) Tween 20 and by
incubation (5min) on PBS without Tween, followed by a 1 h
incubation step on the secondary antibody (goat anti-rabbit
IgG-10 nm gold conjugate; British Biocell International Ltd.,
Cardiff, UK). The secondary antibody was used in a 1 : 80
dilution (in the same solutions as used for the respective
primary antibodies). Again, two 5min washing steps with
PBS containing 0.05% (v/v) Tween 20, and one step with PBS
was performed, followed by washing for 10 s in distilled water
for desalting. Poststaining was performed with 4% (w/v)
uranyl acetate solution for 3min. All steps were conducted
at room temperature.
Electron micrographs were taken, at calibrated mag-
nifications, with a Philips EM 301 transmission electron
microscope (Philips, Eindhoven, The Netherlands) operated
in the conventional bright field mode and a Jeol JEM 1011
(Jeol, Eching,Germany) equippedwith aGatanOrius SC1000
4 Archaea
400 nm
(a)
400 nm
(b)
400 nm
(c)
300 nm
(d)
300 nm
(e)
300 nm
(f)
400 nm
(g)
300 nm
(h)
300 nm
(i)
400 nm
(j)
Figure 3: Localization of MCR in Methanosarcina mazei (DSM 3647; a, b, c),Methanothermobacter marburgensis (d–g), andMethanother-
mobacter wolfei (h, i, j). On media without nickel depletion, the markers are localized in the cytoplasm (b, e, i). On nickel-depleted media
with levulinic acid (c, f: 0.05M; g 0.2M; j: 0.1M levulinic acid) a tendency to membrane localization is obvious (a, d, h are negative controls).
CCD camera (Gatan, Munich, Germany). For enhancement
of gold particles of 5 nm in diameter, images were processed
as follows. High-pass filtering was applied to suppress low
spatial frequencies, that is, large image components with
smooth contrast gradients. After readjustment of the contrast
level by stretching the intensity histogram, the image was
transformed by thresholding at a gray level of 100. In the black
and white image, particles larger than 7 nm and smaller than
4 nm (original size) were eliminated. The resulting images
depict 5 nm gold particles in pure black and white contrast.
Archaea 5
300 nm
(a)
300 nm
(b)
Figure 4: Localization of MCR in predominant morphotypes of microbial mats conducting AOM. Filamentous ANME-1 in the pink layer
of the AOM community (a) and coccoid ANME-2 in the black layer (b) exhibit dense cytosolic MCR labelling.
Particle sizes were enlarged by a factor of 2.5 to allow an
easier identification. When these images were merged with
the original, a higher final contrast was gained (Figure 2(b),
inset). Image processing was performed with the NIH Image
software (National Institute of Health; see also [30, 34]). For
statistical analysis, 20 randomly selected cells were counted.
Gold markers in a range of 25 nm (original size) inside
or outside the cytoplasmic membrane were referred to as
“membrane associated”; all other markers inside the cell were
referred to as cytoplasmic.
3. Results and Discussion
The apparent subunit molecular weights (𝛼: 65 kDa, 𝛽:
49 kDa, and 𝛾: 38 kDa) of the purified methyl-coenzyme
M reductase correspond to the subunit molecular weights
of the MCR I isoenzyme from Methanothermobacter mar-
burgensis [35]. This isoform is predominant when H
2
and
CO
2
supply is low and growth limiting [11]. Western blotting
of crude extracts, prepared from cells that were also used
for immunolocalization, revealed that the antiserum detects
protein bands from a variety methanogens. The apparent
sizes of these bands correspond to the expected molecular
weights of the 𝛼 and 𝛽 subunit of MCR. In some cases,
also the smallest (𝛾) subunit is visible (Figure 1). The same
pattern has already been shown for MCR extracts obtained
from anaerobic methane-oxidizing microbial mats [36]. The
𝛾 subunit of MCR is known to be less immunogenic and
produces a weaker or no signal (Figure 1; [8, 15]). In some
of the Western blots, a second band in the range of 67 kDa
may account for the presence of the MCR isoenzyme II
(MRT) in minor amounts. This enzyme exhibits a slightly
smaller 𝛼 subunit and a 𝛾 subunit of 33 kDa [35]. MRT
was isolated from Methanothermobacter marburgensis [35]
and from various otherMethanobacteriales andMethanococ-
cales, but as yet not from Methanosarcinaceae [35, 37–40].
The hydrogenotrophic methanogens investigated here were
grown in batch culture with limited substrate availability.
Thus, it may be expected that for those methanogens that
contain, in analogy to Methanothermobacter marburgensis,
two methyl-coenzyme M reductases, the MCR isoenzyme I
is dominant [11].
Immunolocalization of MCR from Methanosarcina
mazei (DSM 3318, formerly Methanosarcina frisia;
Figure 2(b)), Methanococcus maripaludis (Figure 2(d)),
Methanosarcina mazei (DSM 3647; Figure 3(b)), Methano-
thermobacter marburgensis (Figure 3(e)), and Methano-
thermobacter wolfei (Figure 3(i)) grown on media without
depletion of nickel show that MCR antigens are distributed
throughout the whole cell. For the autotrophically growing
Methanothermococcus thermolithotrophicus (DSM 2095,
formerly Methanococcus thermolithotrophicus) and for
Methanolobus tindarius (DSM 2278), cytoplasmic local-
ization ofMCR has already been shown [33].Thoughmost of
the organisms tested did not grow on nickel-depleted media,
Methanosarcina mazei (DSM 3647), Methanothermobacter
marburgensis, and Methanothermobacter wolfei grew on
media without the trace element nickel and after addition
of up to 0.2M levulinic acid (final concentration) to the
respective standard growth medium. Levulinic acid inhibits
the biosynthesis of the nickel tetrapyrrole cofactor F430.
This reduces, in addition to nickel-limitation, the expression
of MCR [13, 41]. Under these conditions, the distribution
of the enzyme changed; a higher amount of MCR is now
located at the membrane. Table 1 summarizes the results
obtained after immunolocalization. Statistical errors of
individual cells counted were around 20% for all counts.
Thus, the values show a clear trend, but not an exactly
reproducible value. Cellular redistribution of MCR markers
was most pronounced inMethanothermobacter marburgensis
(Figures 3(f) and 3(g)). The organismmay be considered as a
reference for our experiments, since similar results have been
6 Archaea
described previously [13]. AlsoM. wolfei andM.mazei (DSM
3647) showed redistribution of the gold marker (Figures
3(c) and 3(j)). Though the effect is less obvious than in M.
marburgensis; 50–60% of the markers could be located at the
membrane.
Samples taken from different layers of the multilayered
anaerobically methane-oxidizing microbial mats [32, 36]
show distinct mophotypes of methane oxidizing Archaea
[33]: ANME-1 Archaea are filamentous organisms, related to
Methanomicrobiales and are dominating in the pink-coloured
layer of the microbial mat, whereas the ANME-2 Archaea of
the outermost black layer are related to Methanosarcinales
[42, 43]. Immunolocalization of MCR showed for both
morphotypes intensive cytosolic labelling (Figures 4(a) and
4(b)). In this respect, the expression of MCR may not be
limited in the environmental biofilm.
According to all available biochemical data, membrane
association of MCR is not necessary for functioning. How-
ever, localization of the soluble enzyme in vicinity to the
membrane is favourable for the whole pathway, in particular
when the enzyme production is limited. Membrane binding
becomes obvious, as already stated before [13], when growth
conditions limit the synthesis of MCR. Putatively, the diffu-
sion paths of the reactants are shorter and the final step of
methanogenesis is more effective. We could show that this
feature is not restricted toM. marburgensis, but appears to be
true for the related M. wolfei as well as the phylogenetically
distant M. mazei and may also be expected for the related
methane-oxidizing Archaea. Thus the enzyme may be an
interesting example for a “facultative” membrane association
of proteins, with a certain capability of membrane bind-
ing, but without a specific membrane-dependent metabolic
mechanism [44]. For environmental processes, location of
the enzyme at the membrane may be an indirect indicator
for the physiological status, in our case for nickel-limited
conditions and reduced cofactor biosynthesis. According
to this assumption, this does not appear to be the case
for ANME-1 and ANME-2, prominent in AOM-performing
microbial mats from the Black Sea (cf. Figures 4(a) and
4(b)). The images show dense cytoplasmic localization of the
marker, accounting for nonlimited production of the enzyme.
Acknowledgments
Part of this work was supported by the Deutsche Forschungs-
gemeinschaft (Grant Ho 1830/2-1).We also acknowledge sup-
port by the Open Access Publication Funds of the Go¨ttingen
University. This is a Courant Research Centre Geobiology
publication.
References
[1] R. K. Thauer, “Biochemistry of methanogenesis: a tribute to
Marjory Stephenson,” Microbiology, vol. 144, no. 9, pp. 2377–
2406, 1998.
[2] R. K. Thauer, A. K. Kaster, H. Seedorf, W. Buckel, and R.
Hedderich, “Methanogenic archaea: ecologically relevant dif-
ferences in energy conservation,” Nature Reviews Microbiology,
vol. 6, no. 8, pp. 579–591, 2008.
[3] G. Gottschalk and R. K. Thauer, “The Na+-translocating
methyltransferase complex from methanogenic archaea,”
Biochimica et Biophysica Acta, vol. 1505, no. 1, pp. 28–36, 2001.
[4] E. Stupperich, A. Juza, M. Hoppert, and F. Mayer, “Cloning,
sequencing and immunological characterization of the
corrinoid-containing subunit of the N5-methyltetrahydro-
methanopterin:coenzyme-M methyltransferase from Metha-
nobacterium thermoautotrophicum,” European Journal of Bio-
chemistry, vol. 217, no. 1, pp. 115–121, 1993.
[5] U. Deppenmeier, “Redox-driven proton translocation in
methanogenic archaea,” Cellular and Molecular Life Sciences,
vol. 59, no. 9, pp. 1513–1533, 2002.
[6] A. Stojanowic, G. J. Mander, E. C. Duin, and R. Hedderich,
“Physiological role of the F420-non-reducing hydrogenase
(MvH) from Methanothermobacter marburgensis,” Archives of
Microbiology, vol. 180, no. 3, pp. 194–203, 2003.
[7] R. P. Hausinger, W. H. Orme-Johnson, and C. Walsh, “Nickel
tetrapyrrole cofactor F430: comparison of the forms bound
to methyl coenzyme M reductase and protein free in cells
ofMethanobacterium thermoautotrophicum ΔH,” Biochemistry,
vol. 23, no. 5, pp. 801–804, 1984.
[8] I. Thomas, H.-C. Dubourguier, G. Prensier, P. Debeire, and G.
Albagnac, “Purification of component C from Methanosarcina
mazei and immunolocalization in Methanosarcinaceae,”
Archives of Microbiology, vol. 148, no. 3, pp. 193–201, 1987.
[9] J. Ellermann, R. Hedderich, R. Bo¨cher, and R. K. Thauer,
“The final step in methane formation. Investigations with
highly purified methyl-CoM reductase (component C) from
Methanobacterium thermoautotrophicum (strain Marburg),”
European Journal of Biochemistry, vol. 172, no. 3, pp. 669–677,
1988.
[10] J. Ellermann, S. Rospert, R. K.Thauer et al., “Methyl-coenzyme-
M reductase from Methanobacterium thermoautotrophicum
(strain Marburg)-purity, activity and novel inhibitors,” Euro-
pean Journal of Biochemistry, vol. 184, no. 1, pp. 63–68, 1989.
[11] L. G. Bonacker, S. Baudner, and R. K. Thauer, “Differen-
tial expression of the two methyl-coenzyme M reductases
in Methanobacterium thermoautotrophicum as determined
immunochemically via isoenzyme-speficic antisera,” European
Journal of Biochemistry, vol. 206, no. 1, pp. 87–92, 1992.
[12] R. Ossmer, T. Mund, and P. L. Hartzell, “Immunocytochemical
localization of component C of the methylreductase system in
Methanococcus voltae andMethanobacterium thermoautotroph-
icum,” Proceedings of the National Academy of Sciences of the
United States of America, vol. 83, no. 16, pp. 5789–5792, 1986.
[13] H. C. Aldrich, D. B. Beimborn, M. Bokranz, and P. Scho¨nheit,
“Immunocytochemical localization of methyl-coenzyme
M reductase in Methanobacterium thermoautotrophicum,”
Archives of Microbiology, vol. 147, no. 2, pp. 190–194, 1987.
[14] F. Mayer, M. Rohde, M. Salzmann, A. Jussofie, and G.
Gottschalk, “The methanoreductosome: a high-molecular-
weight enzyme complex in the methanogenic bacterium strain
Go¨1 that contains components of the methylreductase system,”
Journal of Bacteriology, vol. 170, no. 4, pp. 1438–1444, 1988.
[15] M. Hoppert and F. Mayer, “Electron microscopy of native and
artificial methylreductase high-molecular-weight complexes in
strainGo¨ 1 andMethanococcus voltae,” FEBS Letters, vol. 267, no.
1, pp. 33–37, 1990.
[16] 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.
Archaea 7
[17] 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.
[18] S. Shima, M. Kru¨ger, T. Weinert et al., “Structure of a methyl-
coenzyme M reductase from Black Sea mats that oxidize
methane anaerobically,” Nature, vol. 481, no. 7379, pp. 98–101,
2011.
[19] A. Meyerdierks, M. Kube, I. Kostadinov et al., “Metagenome
and mRNA expression analyses of anaerobic methanotrophic
archaea of the ANME-1 group,” Environmental Microbiology,
vol. 12, no. 2, pp. 422–439, 2010.
[20] P. Scho¨nheit, J. Moll, and R. K. Thauer, “Growth parameters
(K(s), 𝜇(max), Y(s)) of Methanobacterium thermoautotroph-
icum,” Archives of Microbiology, vol. 127, no. 1, pp. 59–65, 1980.
[21] H. Huber, M. Thomm, H. Konig, G. Thies, and K. O. Stet-
ter, “Methanococcus thermolithotrophicus, a novel thermophilic
lithotrophicmethanogen,”Archives ofMicrobiology, vol. 132, no.
1, pp. 47–50, 1982.
[22] N. Belay, R. Sparling, and L. Daniels, “Dinitrogen fixation by
a thermophilic methanogenic bacterium,” Nature, vol. 312, no.
5991, pp. 286–288, 1984.
[23] J. Winter, C. Lerp, and H. P. Zabel, “Methanobacterium wolfei,
sp. nov., a new tungsten-requiring, thermophilic, autotrophic
methanogen,” Systematic and Applied Microbiology, vol. 5, no.
4, pp. 457–466, 1984.
[24] H. Hippe, D. Caspari, K. Fiebig, and G. Gottschalk, “Utilization
of trimethylamine and other N methyl compounds for growth
and methane formation by Methanosarcina barkeri,” Proceed-
ings of the National Academy of Sciences of the United States of
America, vol. 76, no. 1, pp. 494–498, 1979.
[25] H. Ko¨nig and K. O. Stetter, “Isolation and characterization
of Methanolobus tindarius, sp. nov., a coccoid methanogen
growing only on methanol and methylamines,” Zentralblatt fu¨r
Bakteriologie Mikrobiologie und Hygiene, vol. 3, no. 4, pp. 478–
490, 1982.
[26] E. Harlow and D. Lane, Antibodies-A Laboratory Manual, Cold
Spring Harbor Laboratory, New York, NY, USA, 1988.
[27] U. K. Laemmli, “Cleavage of structural proteins during the
assembly of the head of bacteriophage T4,” Nature, vol. 227, no.
5259, pp. 680–685, 1970.
[28] H. Towbin, T. Staehelin, and J. Gordon, “Electrophoretic trans-
fer of proteins frompolyacrylamide gels to nitrocellulose sheets:
procedure and some applications,” Proceedings of the National
Academy of Sciences of the United States of America, vol. 76, no.
9, pp. 4350–4354, 1979.
[29] H. Blum, H. Beier, and J. H. Gross, “Improved silver staining
of plant proteins, RNA and DNA in polyacrylamide gels,”
Electrophoresis, vol. 8, no. 2, pp. 93–99, 1987.
[30] M. Hoppert and A. Holzenburg, Electron Microscopy in Micro-
biology, Bios Scientific, Oxford, UK, 1998.
[31] M. M. Bradford, “A rapid and sensitive method for the quanti-
tation of microgram quantities of protein utilizing the principle
of protein dye binding,”Analytical Biochemistry, vol. 72, no. 1-2,
pp. 248–254, 1976.
[32] 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.
[33] 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.
[34] 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.
[35] L. G. Bonacker, S. Baudner, E. Mo¨rschel, R. Bo¨cher, and
R. K. Thauer, “Properties of the two isoenzymes of methyl-
coenzymeM reductase inMethanobacterium thermoautotroph-
icum,” European Journal of Biochemistry, vol. 217, no. 2, pp. 587–
595, 1993.
[36] 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.
[37] S. Rospert, D. Linder, J. Ellermann, and R. K. Thauer,
“Two genetically distinct methyl-coenzyme M reductases in
Methanobacterium thermoautotrophicum strain Marburg and
delta H,” European Journal of Biochemistry, vol. 194, no. 3, pp.
871–877, 1990.
[38] A. Lehmacher and H. P. Klenk, “Characterization and phy-
logeny ofmcrII, a gene cluster encoding an isoenzyme ofmethyl
coenzyme M reductase from hyperthermophilic Methanother-
mus fervidus,” Molecular and General Genetics, vol. 243, no. 2,
pp. 198–206, 1994.
[39] C. J. Bult, O. White, G. J. Olsen et al., “Complete genome
sequence of the Methanogenic archaeon, Methanococcus jan-
naschii,” Science, vol. 273, no. 5278, pp. 1058–1073, 1996.
[40] E. Springer, M. S. Sachs, C. R. Woese, and D. R. Boone,
“Partial gene sequences for the A subunit of methyl-coenzyme
M reductase (mcrI) as a phylogenetic tool for the family
Methanosarcinaceae,” International Journal of Systematic Bacte-
riology, vol. 45, no. 3, pp. 554–559, 1995.
[41] R. Jaenchen, H. H. Gilles, and R. K. Thauer, “Inhibition of
factor F430 synthesis by levulinic acid in Methanobacterium
thermoautotrophicum,” FEMS Microbiology Letters, vol. 12, no.
2, pp. 167–170, 1981.
[42] K. U. Hinrichs, J. M. Hayes, S. P. Sylva, P. G. Brewert, and
E. F. DeLong, “Methane-consuming archaebacteria in marine
sediments,” Nature, vol. 398, no. 6730, pp. 802–805, 1999.
[43] V. J. Orphan, C. H. House, K. U. Hinrichs, K. D. McKeegan,
and E. F. DeLong, “Methane-consuming archaea revealed by
directly coupled isotopic and phylogenetic analysis,” Science,
vol. 293, no. 5529, pp. 484–487, 2001.
[44] M. Hoppert and F. Mayer, “Principles of macromolecular
organization and cell function in bacteria and archaea,” Cell
Biochemistry and Biophysics, vol. 31, no. 3, pp. 247–284, 1999.