From the Institut für Mikrobiologie, Martin-Luther-Universität Halle-Wittenberg, Kurt-Mothes-Strasse 3, 06120 Halle, Germany
Received for publication, February 9, 2001, and in revised form, March 23, 2001
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The Gram-positive anaerobe
Eubacterium acidaminophilum contains at least two
tungsten-dependent enzymes: viologen-dependent formate dehydrogenase and aldehyde dehydrogenase.
185W-Labeled tungstate was taken up by this organism with a
maximum rate of 0.53 pmol min The function of tungsten as an essential trace element for some
archaea and bacteria has now been fully recognized (1-3). It is
incorporated into a pterin cofactor that is required for assembly and
function of enzymes such as acetylene hydratase, aldehyde ferredoxin
oxidoreductase, carboxylic acid reductase, formaldehyde dehydrogenase,
formate dehydrogenase, formylmethanofuran dehydrogenase, and
glyceraldehyde-3-phosphate ferredoxin oxidoreductase (3-10). Tungsten
is very similar in size and chemical behavior to molybdenum, and
therefore, tungsten can act as a specific antagonist of
molybdenum-containing enzymes. Tungsten-containing enzymes seem to
catalyze similar or even the same reactions as related molybdoenzymes
(1-3). In contrast to tungstate (11, 12), the uptake of molybdate into
cells has been well studied and is mediated by an ABC transporter
system (13-15). The molybdate-binding protein ModA has been identified
to bind specifically molybdate and tungstate (16), and the structures
of the ModA proteins from Escherichia coli and
Azotobacter vinelandii have been analyzed at 1.75 and 1.25 Å resolution, respectively (17-19). The specificity of ModA to bind
molybdate and tungstate, but not sulfate or other anions, is determined
mainly by the size of the binding pocket for molybdate/tungstate (17).
Because tungstate and molybdate have nearly identical sizes, ModA
cannot discriminate between these anions (16-19). At present, only a
little is known about a specific uptake mechanisms for tungstate into
microorganisms. Labeling studies were done for certain enzymes, and the
tungstate requirement for optimal growth was analyzed for some
organisms, most of them being (hyper)thermophilic (1-3).
The Gram-positive anaerobe Eubacterium acidaminophilum grows
at mesophilic temperatures, degrades amino acids by Stickland reactions, and requires the addition of selenite but not molybdate or
tungstate for growth (20). Two enzymes are involved in its metabolism:
formate dehydrogenase and aldehyde dehydrogenase, which are now
described to be specifically dependent on tungsten availability to
exhibit enzymatic activity. This correlates with a specific
incorporation into both enzymes (21, 22). This report focuses on
uptake experiments using resting cells of E. acidaminophilum
and radioactive labeled tungstate as well as on cloning and sequencing
of an ABC transporter. The extracytoplasmic binding protein of this
transporter, TupA, was overexpressed in E. coli, and its
anion binding characteristics show a specificity for tungstate. Thus,
this is the first report of a transport system specific for tungstate.
Materials--
All chemicals were obtained from commercial
sources unless otherwise specified.
Bacterial Strains and Plasmids--
E.
acidaminophilum (DSM 5388T) was grown anaerobically as
described (20). Glycine (50 mM) or the substrate
combination serine (10 mM), formate (40 mM),
betaine (60 mM) was used as growth substrate. The organism
was also grown without the addition of tungstate and molybdate as
described in this paper. E. coli XL1-Blue MRF' and E. coli SOLR were obtained from Stratagene (Heidelberg, Germany) and
used for cloning purposes and expression of proteins. They were grown
in Luria broth medium (23) or on agar plates containing 1.5% (w/v)
agar. 100 µg ml Formate Dehydrogenase and Aldehyde Dehydrogenase
Assay--
Formate dehydrogenase and aldehyde dehydrogenase were
measured by a standard procedure (12) at 34 °C under anaerobic
conditions by monitoring the substrate-dependent reduction
of benzyl viologen at 578 nm ( Uptake of [185W]Tungstate by E. acidaminophilum and
Labeling of Proteins--
[185W]Tungstate was
obtained from Amersham Pharmacia Biotech (Braunschweig, Germany); after
discontinuation of that service, the unlabeled salt was irradiated with
neutrons at the GKSS Forschungszentrum (Geesthacht, Germany) to obtain
[185W]tungstate (t1/2 = 75 d).
Na2WO4·2H2O (2.2 mg, 6.7 µmol)
was irradiated for 7 days to obtain a final radioactivity of
1.7·107 Bq and a specific radioactivity of
1.7·109 Bq/mmol.
For uptake experiments, cells (10 ml) were grown overnight with
serine/formate/betaine as substrates, harvested anaerobically at
2,000 × g, and washed twice with 10 ml of ice-cold 50 mM Na2HCO3 buffer (pH 7.5). Cells
were suspended in the same buffer, and 10 mM serine, 40 mM formate, 60 mM betaine, and 30 µg
ml
To label proteins of E. acidaminophilum with
[185W]tungstate, cells were grown overnight with glycine
as substrate in 3 liters of medium supplemented with
[185W]tungstate (4·105 Bq) and different
concentrations of unlabeled tungstate to give the final concentration.
Cells were harvested by centrifugation at 10,000 × g
and 15 min and were washed two times with potassium phosphate buffer
(50 mM, pH 7.5) and suspended in this buffer. After rupture
of the cells by two passages through a French pressure cell, cell
debris were removed by centrifugation at 20,000 × g for 20 min. To separate the labeled proteins, the supernatant was
desalted using a PD10 column (Amersham Pharmacia Biotech) and applied
to a Superdex-200 gel filtration column (Amersham Pharmacia Biotech).
The column had been preequilibrated with 50 mM potassium
phosphate buffer (pH 7.5) containing 150 mM KCl.
DNA Manipulations and Sequence Determination--
Routine DNA
techniques were performed as described by Sambrook et al.
(23). Genomic DNA from E. acidaminophilum was isolated according to Saito and Miura (24). Enzymes were used according to the
recommendations of the manufacturer. Plasmid preparations were done
using the Qiagen (Hilden, Germany) kits as outlined in the
manufacturer's manual. Nucleotide sequences were determined by the
dideoxy chain termination method (25) using the Rhodamine Terminator
Cycle Sequencing Ready Reaction kit (PerkinElmer Life Sciences) and
analyzed using an Applied Biosystems PRISM 377 DNA sequencer. Sequence
information of large inserts were obtained using the GPSTM
genome priming system from New England Biolabs (Frankfurt, Germany) as
recommended by the manufacturer. Alternatively, the primer-walking method was used. The oligonucleotides were synthesized by Metabion (Martinsried, Germany). DNA fragments were labeled, and hybridizing bands were detected using the DIG DNA labeling and detection kit from
Roche Diagnostic according to the manufacturer's manual. A RNA Techniques--
E. acidaminophilum was grown in
medium containing glycine or serine/formate/betaine as substrates to an
absorbance of 0.9 and harvested by centrifugation at 4,000 × g and 4 °C for 10 min. Total RNA was isolated using the
RNeasy Mini Kit (Qiagen) with modifications. 10-25-ml cultures were
used, and the cells were lysed in 20 mg of lysozyme
ml
For RT-PCR experiments, purified RNA was treated with DNase RQ1
(Promega, Mannheim, Germany) following the instructions of the supplier
to obtain DNA-free RNA. For reverse transcription of mRNA, 1 µl
of purified RNA and 1 µl of hexanucleotide mixture (500 µg/ml) were
added to 10 µl of H2O, denatured for 10 min at 70 °C,
and cooled for 2 min on ice. Subsequently, 4 µl of first strand
buffer, 2 µl of 0.1 M dithiothreitol, 1 µl of 20 mM dNTP, and 0.7 µl of reverse transcriptase (Superscript
II, Life Technologies, Inc.) were added, and the mixture was incubated
for 10 min at room temperature and 1 h at 42 °C. The reaction
was terminated by incubation at 94 °C for 5 min. This reaction
mixture contained the synthesized cDNA, and 1 µl of it was used
as template in the following PCR. 100 pmol of forward and 100 of pmol
reverse primer, 1 µl of 20 mM dNTP, 1 µl of
Taq DNA polymerase (Quantum Appligene, Heidelberg), 5 µl
of Taq buffer, and 42 µl of H2O were mixed, and PCR was performed with 30 cycles: 30 s at 94 °C for
denaturing, 30 s of annealing at 42-72 °C (depending on the
primers used), and 1-3-min extension (depending on the distance
between both primers) at 72 °C.
Cloning of tupA into the pASK-IBA2 Vector--
To clone the
tupA coding sequence into the pASK-IBA2 expression vector
(IBA, Göttingen), the gene was amplified by PCR using Pwo DNA polymerase (Roche Diagnostics), primers PA3 and PA3r
(Table I), and chromosomal DNA from E. acidaminophilum as
template. The leader peptide of TupA including the conserved cysteine
residue was omitted and replaced by the OmpA leader peptide provided by the vector pASK-IBA2. Both primers contain a BsaI
restriction site and were cloned into the respective site of pASK-IBA2
resulting in the TupA fusion protein with an N-terminal OmpA signal
peptide and a C-terminal Strep-tagII. This plasmid,
pASK-tupA, was transformed into E. coli XL 1-Blue MRF'. The
insert of pASK-tupA was sequenced for confirmation.
Expression and Purification of TupA--
An E. coli
XL1-Blue clone harboring the plasmid pASK-tupA was inoculated into 3 ml
of LB medium containing 100 µg ml
E. coli cells overexpresing TupA were suspended in buffer A
(100 mM Tris (pH 8.0), 0.5 g of cells
ml TupA Protein Gel Shift Assay--
The
ligand-dependent gel shift assay developed by Rech et
al. (16) was used to analyze anion binding by TupA. Aliquots of purified TupA were incubated with the indicated concentration of anion
in binding buffer (50 mM potassium acetate (pH 5.0), 100 mM Tris (pH 8.0)) for 30 min on ice. Samples were mixed
with 0.25 volume of a sucrose solution (30% w/v) containing bromphenol blue and applied to a native 12% polyacrylamide gel. Protein was separated in a mini-Gel system (Biometra, Göttingen). The gel was
buffered with 50 mM Tris (pH 8.5), and the running buffer contained 0.1 M Tris and 0.1 M glycine.
Electrophoresis was done at 150 mV and 4 °C until bromphenol blue
left the gel.
Analytical Methods--
The concentration of proteins was
determined by the method of Bradford (27) with bovine serum albumin as
a standard. Whole cells were incubated in 0.1 M NaOH for 10 min at 95 °C and neutralized with 0.5 M HCl prior to
protein determination. The extinction coefficient for TupA was
determined to 24,460 mol Uptake of [185W]Tungstate by Growing Cells of E. acidaminophilum and Labeling of Proteins--
Omission of tungstate
and molybdate from the added trace element solution did not affect
growth of E. acidaminophilum over hundreds of generations in
a defined mineral medium on different substrates. After growth of
E. acidaminophilum in the presence of
10
However, a coelution of radioactivity and enzyme activity could already
be obtained for the 160-kDa protein with a
viologen-dependent formate dehydrogenase (31, 32) and for
the 74-kDa protein with a viologen-dependent acetaldehyde
dehydrogenase (32). Both enzymes were now independently purified to
homogeneity and contain tungsten but no
molybdenum.2,3
Tungstate and molybdate had quite different effects on both enzyme activities. In tungstate/molybdate-depleted cells of E. acidaminophilum, viologen-dependent acetaldehyde
dehydrogenase activity was no longer measurable, whereas
viologen-dependent formate dehydrogenase activity was still
present at 18% of its maximum value (100% = 1.1 unit
mg Uptake of [185W]Tungstate into Resting Cells of E. acidaminophilum--
Cells of E. acidaminophilum were grown
with serine/formate/betaine as carbon and energy source. Uptake
experiments using [185W]tungstate were carried out with 1 µM labeled tungstate and resting cells in carbonate
buffer and serine/formate/betaine as energy source. The highest uptake
rate of 0.53 pmol min Cloning of the tupABC Operon--
The genes encoding a tungstate
uptake system were identified in the downstream region of the genes
encoding the tungsten-containing formate dehydrogenase I and some
enzymes involved in pterin-cofactor biosynthesis of E. acidaminophilum (21) (Fig.
2A, depicted as 7.3-kb
EcoRI fragment). Sequence comparison and later biochemical analysis (see below) revealed that the three genes in the immediate downstream region of the mentioned 7.3-kb fragment encode an ABC uptake
system that seems to be specific for tungstate. According to the
results presented below, these genes have been named tup for
tungstate uptake (Fig.
2). To clone this EcoRI fragment downstream of formate
dehydrogenase I genes, a Southern blot analysis was performed using
probe A, which was deduced from the 3'-region of the 7.3-kb
EcoRI fragment (Fig. 2 and Table
I primers PprobeA). First, a
3-kb SacI fragment was identified which covers the 3'-part
of the 7.3-kb EcoRI fragment and the 5'-part of the
following 6.9-kb EcoRI fragment (Fig. 2).
SacI-digested chromosomal DNA from E. acidaminophilum was separated in a sucrose density gradient and
fractionated. Using probe A, a positive reacting fraction was
identified which contains fragments of 2-4 kb. Subsequently, these
fragments were ligated into pBSK Bluescript and transformed into
E. coli XL1-Blue MRF'. All transformants were washed from the agar plates and used to inoculate 100 ml of LB medium. After growth
overnight, the plasmid mixture was isolated from this culture and was
used directly as a template in a PCR that was done with primers
FDHlong and universal primer (Table I). A 3.1-kb fragment was amplified and partly sequenced to obtain sufficient sequence information from the so far unknown DNA region downstream of the EcoRI site to generate probe B (Fig. 2, Table I primers
PprobeB). With that probe a
The N-terminal amino acid sequence of TupA was similar to leader
peptides of bacterial lipoproteins (36). A conserved cysteine residue
at position 21 preceded by a glycine or alanine residue is
characteristic for such leader peptides, which are split at the
N-terminal side of the cysteine after transport across the cytoplasmic
membrane. Subsequently, the cysteine is modified in this class of
proteins to contain a fatty acid at the amino group and a diacyl
glycerol thioether at the thiol group. This lipid anchor is used to
attach the protein to the outside of the cytoplasmic membrane in
Gram-positive bacteria or mycoplasmas (37).
Expression and Purification of TupA--
If the TupA protein was
expressed in E. coli containing its native leader peptide,
90% of TupA was found in the cytoplasmic membrane of E. coli (data not shown), and only a very small amount of soluble but
inactive TupA was obtained. Thus, TupA was expressed as a C-terminal
Strep-tag fusion protein containing the OmpA leader peptide as outlined under "Experimental Procedures" resulting in a
higher yield (5 mg of TupA from 100 ml of E. coli culture) of soluble and active protein. Five bands were present in the purified
TupA preparation as revealed by analysis using native polyacrylamide
gel electrophoresis (Fig. 3A),
and no further purification/separation could be obtained (data not
shown). The N-terminal amino acid sequences of these protein bands
revealed that all bands represented only TupA, which differed slightly
by its N-terminal start (Fig. 3B). Although these TupA
protein species differed only by a few amino acids in length, their
size seems to vary between 34 and 38 kDa as judged by denaturing
SDS-polyacrylamide gel electrophoresis (data not shown). Most likely
these forms of TupA were processed differently by the signal peptidase
of E. coli. The native size of TupA as determined by gel
filtration eluted in one peak with a maximum at 38 kDa, indicating that
the expressed TupA was present as a monomer (data not shown).
Specificity of the TupA Protein for Binding Divalent
Oxyanions--
To examine the binding specificity of TupA, the native
polyacrylamide gel mobility assay as developed by Rech et
al. (16) was used. The anions sulfate, molybdate, tungstate,
phosphate, chlorate, chromate, vanadate, and selenate were tested and
supplied in a 90-fold molar excess of TupA (Fig. 3A). None
of these anions except tungstate caused a mobility shift of TupA,
indicating that TupA specifically binds tungstate. Binding specificity
was not altered even testing pH values of 5 and 8 in the binding buffer (data not shown). If molybdate and sulfate anions were added in a
higher than 1,000-fold molar excess to TupA, a mobility shift was
observed, but it was weaker than the shift induced by tungstate (data
not shown). 11 µM TupA was titrated with 0-30
µM tungstate to estimate the dissociation constant for
tungstate (Fig. 3C). An apparent Kd value
of ~0.5 µM was determined for tungstate binding using
Equation 1
TupA (286 amino acids, 30.9 kDa) exhibited no significant similarities
to the molybdate-binding proteins ModG, ModE, and Mop and only weak
similarities to ModA proteins and to other anion-binding proteins (Fig.
4 and data not shown) (14, 38). Highest
similarities were obtained to hypothetical proteins from
Methanobacterium thermoautotrophicum (accession no. G69162),
Haloferax volcanii (CAB42540), Campylobacter
jejuni (B81301), and Vibrio cholerae (A82188) (Fig. 4).
Therefore, these proteins are termed TupA-homologous proteins in this
publication. They are all included in operons encoding putative ABC
transporters and represent the substrate-binding component (Fig.
5). The TupA-homologous protein from
H. volcanii was identified during mutational analysis to be
essential for nitrate respiration (39). The Tup-homologous ABC
transporter from M. thermoautotrophicum has been suggested
in the data-base entry to be an ABC transporter specific for sulfate. A
phylogenetic tree was derived for TupA and the mentioned homologous
proteins as well as for binding proteins specific for other oxyanions
such as molybdate, sulfate, and phosphate. Additionally, other
hypothetical binding proteins obtained during similarity searches are
included. TupA and its homologs clustered together in a distinct
phylogenetic group compared with the other oxyanion-binding proteins.
Because the crystal structure of two ModA proteins is available, the
amino acids that bind molybdate have been identified to be Ser-12,
Ser-39, Ala-125, Val-152, and Tyr-170 in E. coli ModA and
Thr-9, Asn-10, Ser-37, Tyr-118, and Val-147 in A. vinelandii
ModA (Fig. 4) (17-19). TupA and its homologs show a conserved TTTS
motif close to the corresponding molybdate-binding amino acids Ser-12
(E. coli) and Thr-9/Asn-10 (A. vinelandii) and a
conserved threonine flanked by two glycine residues corresponding to
Ser-37 (E. coli) and Ser-39 (A. vinelandii) (Fig.
4). Ala-125 of E. coli ModA and Tyr-118 of A. vinelandii ModA are replaced in TupA and its homologs by the
positively charged Arg-135 present in the motif
SRGDXSGT (Fig. 4). Arg-135 and Asp-137 are
also conserved in the phosphate-binding protein of E. coli,
and Arg-135 is involved in phosphate binding and linked to Asp-137 by
salt bridges (40). Additionally, Val-147/152 in ModA is replaced by
glycine in the TupA group (Fig. 4).
TupB (228 amino acids, 24.5 kDa) exhibited highest similarities (36%
identity) to the permease protein MTH478 of the ABC transporter from
M. thermoautotrophicum, whose binding protein was similar to
TupA (Fig. 5). Nearly identical similarity values were obtained to the
permease components of the putative ABC transporters from C. jejuni (AL139078, 34% identity), V. cholerae
(AE004231, 33% identity), and H. volcanii (AJ238877, 32%
identity), which are located in the operons encoding the
TupA-homologous proteins (Fig. 5). Additionally, lower similarity
values were determined for permease components of other ABC
transporters including ModB from the molybdate-specific ABC transporter
(<20% similarity). Five transmembrane helices were postulated to be
present in TupB using the dense alignment surface method (41), which
indicates a location of this protein in the cytoplasmic membrane
(data not shown). The conserved C-terminal region
EAAX2GX9IXLP, which is generally present in permease components of ABC transporters was identified to be
RIGX2LGX8LXIR
in TupB and is, thus, only slightly conserved. This sequence was
suggested to constitute a recognition site for the ATP-binding protein
component (42).
TupC (214 amino acids, 23.6 kDa) was similar to a wide variety of
ATP-binding proteins of ABC transporters with values of up to 39%
identical amino acids calculated on the basis of the length of TupC.
Other ATP-binding subunits of ABC transporters are up to 120 amino acid
residues longer than TupC. The ATP-binding components of the ABC
transporter of C. jejuni and of V. cholerae, whose putative binding proteins and permease proteins are similar to
TupA and TupB, were also homologous to TupC with values of 34 and 32%
identity, respectively. The respective proteins from both archaea
H. volcanii and M. thermoautotrophicum showed
identity values below 28%. In TupC amino acid sequences were present,
which are similar to the Walker A (Gly-31 through Thr-38) motif and the
Walker B motif (Leu-149 through Asp-153) (43), and should be
responsible for nucleotide binding (data not shown). Near to the Walker
B motif a conserved linker peptide
(129LSGGETQRV137) typical for the ATP-binding
subunit of ABC transporters (42) was present in TupC. A conserved
histidine residue (His-187) was identified in TupC which might also be
involved in nucleotide binding (44).
Sequence Analysis of MoeA, MoeA-1, MoaA, and MoaC--
The
proteins encoded by moeA and moeA-1 exhibited
just 53% identity to each other and differed in size. They exhibit
similarities to many MoeA proteins from different sources present in
the data bases. However, the highest similarity values were obtained to MoeA proteins from hyperthermophilic archaea like Pyrococcus
abyssi and Archeoglobus fulgidus and lower values to
MoeA proteins from bacterial sources. MoeA is involved in the
biosynthesis of the molybdopterin cofactor. Hasona et al.
(45) suggest that MoeA participates in the synthesis of a
molybdenum-sulfur complex leading to formation of a thiomolybdate,
which is apparently the molybdenum donor in the production of the
molybdopterin cofactor. Additionally, evidence was provided that MoeA
might regulate the expression of formate hydrogen lyase and respiratory
nitrate reductase (46).
Sequence Analysis of MoaA and MoaC'--
MoaA and MoaC' are
similar to proteins involved in the formation of precursor Z, which is
an intermediate during molybdopterin cofactor biosynthesis (47, 48).
MoaA contains an N-terminal and a C-terminal cysteine-rich region. The
N-terminal motif (CXXXCYXC) has been also
identified in NifB and in PqqE, a protein involved in the biosynthesis
of PQQ (49). A mutation of these cysteine to serine residues leads to
an inactivation of MoaA (50). It is speculated that this motif is
necessary for the formation of iron-sulfur clusters (49). The function
of the C-terminal cysteine motif in MoaA
(CXXCX14C) is not known.
Transcriptional Analysis of the tupABC Gene
Region--
Transcription of the genes tupABC,
moeA, moeA-1, moaA, and
moaC was analyzed using RT-PCR because no signals were
detected by Northern blot analysis, probably due to a low transcription rate of these particular genes. RNA was isolated from cells grown in a
medium containing serine/formate/betaine as substrates. tupA and tupB were transcribed together on one transcript
(transcript I) (Fig. 2B). Stronger signals were obtained
when RT-PCR experiments were carried out with primers deduced from the
coding region of tupA compared with a situation where one
primer was deduced from tupA and one primer from
tupB (data not shown). A loop structure (111 kJ
mol All data presented here indicate that tungstate is taken up in
E. acidaminophilum by a high affinity and highly specific
transport system. For the first time such a specificity is described.
The deduced amino acid sequences exhibit similarities to ABC-type uptake permeases from different sources, particular to the sulfate, phosphate, and molybdate uptake transporters (SulT, PhoT, and MolT
family, TC 3.A.1.6-8) according to Saier (35).
Two different key metabolic enzymes formate dehydrogenase I and II (21,
31, 32) and aldehyde dehydrogenase (22, 32) of E. acidaminophilum contain tungsten. Thus, an uptake system specific
for tungstate is essential for this organism (20). It has to be very
efficient because the organism grew well and contained a reduced, but
still active, tungsten-dependent formate dehydrogenase
activity if no tungstate was added. Because E. acidaminophilum was cultivated over hundreds of generations
without tungsten supplementation, and no growth impairment was observed
(20, 32, and data not shown), it seems unlikely that tungstate was stored.
So far, no molybdenum-dependent enzyme has been detected in
E. acidaminophilum. Therefore, a specific uptake system for
tungstate, discriminating molybdate, would be reasonable. Otherwise, if
a classical molybdoenzyme, e.g. xanthine dehydrogenase,
would be present as in purinolytic anaerobes (17), E. acidaminophilum should contain a separate molybdate uptake system,
and additional principles must be exerted by the cell to incorporate
the proper metal ion into the respective enzyme. So far, no molybdate
specific ABC transporter has been identified in E. acidaminophilum using heterologous probes deduced from the
respective transporter genes from E. coli and
Arthrobacter nicotinovorans (data not shown). However, the
presence of two genes similar to moeA (moeA-1,
moeA) might be indicative of proteins responsible for a
selective incorporation of tungsten and molybdenum, respectively.
The ABC transporter described in this paper exhibits the typical
structure of ABC transporter systems (for a review, see Refs. 34 and
35). It includes a substrate-specific binding protein such as TupA,
which is localized in the periplasm of Gram-negative bacteria or
attached as a lipoprotein to the outer side of the cytoplasmic membrane
in Gram-positive bacteria; a permease (TupB), which is an integral
membrane protein that facilitates the transport across the membrane;
and a protein (TupC) attached to the cytoplasmic side of the membrane
that hydrolyzes ATP to provide energy for the transport.
TupA had a Kd value of 0.5 µM for
tungstate, indicating a high specificity. Molybdate and sulfate were
bound weakly by TupA when they were added in a more than 1,000-fold
molar excess, indicating that the Kd values for
these two anions are orders of magnitude higher than for tungstate or
that both substances contained tungstate as contaminant. Other anions
like vanadate and phosphate were not bound to TupA. Thus, TupA is able
to discriminate between tungstate and all other tested anions including
molybdate. Molybdate and tungstate are chemically very similar and have
the same ionic radii (cited in Ref. 1). In contrast to TupA, the molybdate-binding protein ModA of E. coli cannot
discriminate between molybdate and tungstate, and thus both anions
induce a mobility shift of ModA in native polyacrylamide gels (16). The Kd values were determined to be 3 µM
for molybdate and 7 µM for tungstate in ModA (16);
however, other authors reported a Kd value of 0.02 µM for molybdate, but giving no value for tungstate (51).
The Kd value for molybdate binding to the regulatory
protein ModE of E. coli was determined to be 0.8 µM (52).
In contrast to the ModA proteins investigated, the molybdate repressor
ModE (14, 51), and the small molybdate/tungstate-binding protein Mop of
Sporomusa ovata (53), TupA was able to discriminate between
the anions molybdate and tungstate. It is an interesting questions how
the protein accomplishes this specificity at the molecular level. A
discrimination by size seems unlikely because both anions are nearly
identical in size (1, 17). The pKa value of
tungstate is 4.7, the pKa of molybdate is 3.8 (1),
and thus, TupA might be able to take advantage of this difference.
Hydrogen bond energy increases when the pKa values
of donor and acceptor become matched at the transition state (54). The
ligands binding tungstate should reflect this behavior exhibiting
stronger hydrogen bonds to the more basic tungstate and thus bind this
anion with higher affinity than molybdate and other oxyanions. At
least, the amino acids known to be involved in molybdate chelation by
ModA (17, 19) exhibited noticeable changes in the TupA of E. acidaminophilum and related proteins (Fig. 4). To identify the
molecular basis of the ability of TupA to discriminate tungstate and
other anions, the structure of TupA has to be analyzed. Experiments to
crystalize TupA are under way in our laboratory.
The ABC transporters exhibiting highest similarities to the
tupABC-encoded proteins were identified in archaea (H. volcanii, M. thermoautotrophicum) and Gram-negative
proteobacteria (V. cholerae, C. jejuni). TupA
from E. acidaminophilum and the four TupA-homologous proteins form a phylogenetic separate group compared with other anion-binding proteins (Fig. 5). These data suggest that all of these
organisms might have a specific tungstate uptake system and
consequently should have tungsten-containing cell components. Tungsten-dependent enzyme activities are known for M. thermoautotrophicum (1, 55), and it is likely that they should
also occur in H. volcanii because predominantly archaea are
known to contain tungstoenzymes (1, 3). To our knowledge, there is no
report that V. cholerae and C. jejuni contain
tungsten-dependent enzymes. However, both organisms can
grow anaerobically and might express tungstoproteins under anaerobic conditions.
1
mg
1 of protein at 36 °C. The uptake was
not affected by equimolar amounts of molybdate. The genes
tupABC coding for an ABC transporter specific for tungstate
were cloned in the downstream region of genes encoding a
tungsten-containing formate dehydrogenase. The substrate-binding
protein, TupA, of this putative transporter was overexpressed in
Escherichia coli, and its binding properties toward
oxyanions were determined by a native polyacrylamide gel retardation
assay. Only tungstate induced a shift of TupA mobility, suggesting that
only this anion was specifically bound by TupA. If molybdate and
sulfate were added in high molar excess (>1000-fold), they were also
slightly bound by TupA. The Kd value for tungstate
was determined to be 0.5 µM. The genes encoding the tungstate-specific ABC transporter exhibited highest similarities to
putative transporters from Methanobacterium
thermoautotrophicum, Haloferax volcanii, Vibrio
cholerae, and Campylobacter jejuni. These five
transporters represent a separate phylogenetic group of oxyanion ABC
transporters as evident from analysis of the deduced amino acid
sequences of the binding proteins. Downstream of the tupABC
genes, the genes moeA, moeA-1,
moaA, and a truncated moaC have been identified
by sequence comparison of the deduced amino acid sequences. They should
participate in the biosynthesis of the pterin cofactor that is present
in molybdenum- and tungsten-containing enzymes except nitrogenase.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
1 ampicillin was added, and
40 µg ml
1 5-bromo-4-chloro-3-indolyl
-D-galactopyranoside and 48 µg
ml
1 isopropyl
-D-thiogalactopyranoside if needed. The plasmid vector pBluescipt KS+ was obtained from Stratagene, the protein
expression vector pASK-IBA2 was from Institut für Bioanalytik
(Göttingen, Germany).
= 8.3 mM
1
cm
1) or methyl viologen at 600 nm (
= 13.1 mM
1
cm
1). To assay formate dehydrogenase, the
reaction mixture contained in a final volume of 1 ml, 50 mM
Tris buffer (pH 8.0), 5 mM benzyl or methyl viologen, 20 mM sodium formate, and 0.1-10 µl of extract. 1-5 µl
of 50 mM sodium dithionite was added to the reaction
mixture to obtain a light reduction of the viologen as an indication
for strict anaerobic conditions. Aldehyde dehydrogenase was measured under similar conditions (6) except that the reaction mixture contained
in a final volume of 1 ml, 50 mM Tris buffer (pH 8.5), 25 mM benzyl viologen, and 0.5 mM acetaldehyde.
Blanks were run with the same reaction mixture excluding the substrate.
The reactions were always started by the anaerobic addition of
substrate. 1 unit of enzyme activity was defined as the amount of
enzyme catalyzing the reduction of 2 µmol of viologen
min
1.
1 chloramphenicol were added to give a
final volume of 9.9 ml. Subsequently, cells were incubated at 30 °C,
and 100 µl of a 10 µM [185W]tungstate
solution was added. At different time intervals, 1-ml samples were
taken and collected on nitrocellulose filter discs (pore size 0.45 µm, Schleicher & Schüll, Dassel, Germany). The filter discs had
been preincubated in 50 mM Na2HCO3
buffer (pH 7.5) containing 1 mM tungstate and were washed
after cell collection with 20 ml of this buffer. Filter discs were
dried for 30 min at 60 °C, incubated in scintillation fluid
(Beckmann, München Germany), and the radioactivity was determined
in a Beckmann LS6500 scintillation counter with an efficiency of nearly
90%.
ZAP II
library was constructed using the
ZAP II-predigested EcoRI/CIAP-treated vector Kit (Stratagene). Genomic DNA of
E. acidaminophilum was digested with the restriction
endonuclease EcoRI and separated by centrifugation for
24 h at 200,000 × g using a sucrose density
gradient (10-40% (w/v) sucrose). DNA fragments of 5-10
kb1 were ligated into the
EcoRI site of the
ZAP II vector and subjected to in
vitro packaging according to the manufacturer. The resulting phage
particles represented a library of E. acidaminophilum DNA and were screened by plaque hybridization after infection of E. coli XL1-Blue MRF'. M13 ExAssist helper phage and E. coli SOLR were used for in vivo excision of the
pBluescript phagemid according to the instructions provided by Stratagene.
1 for 10 min. Northern
hybridizations were performed as described (26).
1
ampicillin and was grown overnight at 30 °C. 2 ml of this culture was transferred to 100 ml of LB medium containing 100 µg
ml
1 ampicillin. After the culture had reached
an absorbance of 0.5 at 550 nm, 10 µl of an anhydrotetracyclin
solution (2 mg ml
1 in dimethyl sulfoxide) was
added to induce expression of the binding protein. After 3 h at
30 °C the cells were harvested by centrifugation for 12 min at
4,000 × g and 4 °C and used directly for the
preparation of crude extract or stored at
20 °C.
1 buffer) and were lysed by sonication
using a GM 60 HD sonicator (Uni Eqip Laborgerätebau, Martinsried,
Germany) at intervals of 5 s until most cells were broken. After
each interval the samples were cooled on ice for 10 s. Cell debris
were removed by centrifugation at 25,000 × g and
4 °C for 5 min. The resulting supernatant was used directly or
stored at
20 °C. The supernatant containing TupA was applied to a
1-ml Strep-Tactin column (IBA) that had been equilibrated
previously with buffer A. The column was washed with 5 ml of buffer A
and subsequently, the TupA-binding protein was eluted in 3 ml of buffer
A containing 2.5 mM desthiobiotin.
1
cm
1 according to Gill and von Hippel
(28) and used to calculate protein concentration of the purified
protein. SDS-polyacrylamide gel electrophoresis was done with
the Laemmli buffer system as described (29). Proteins were blotted from
an SDS gel onto a polyvinylidene difluoride membrane according to
Towbin et al. (30) in 50 mM NaBO3
buffer (pH 9.0) containing 20% (v/v) methanol at 1.2 mA
cm
2 membrane for 1-2 h. For Edman
degradation, blotted proteins were cut from the polyvinylidene
difluoride membrane and analyzed in a 476A amino acid sequencer
(Applied Biosystems, Weiterstadt, Germany). Molybdenum and tungsten
were determined by ICP-MS with an Agilent 7500 Series ISP-MS (Agilent
Technologies, Waldbronn, Germany).
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
9 M
[185W]tungstate, labeled proteins were analyzed by gel
filtration. This revealed a strong radioactivity peak at 74 kDa and one
shoulder at about 160 kDa. If 10
8
M radioactive tungstate was present during growth, the
specific radioactivity eluting at about 160 kDa was nearly unchanged,
whereas the value of the radioactivity peak at 74 kDa increased 2-fold, and a new radioactivity peak appeared at about 40 kDa which became prominently labeled if higher concentrations of tungstate were provided
(data not shown). The simultaneous addition of equimolar amounts of
molybdate had a negligible effect on the observed labeling pattern
(data not shown), pointing to a high specificity for tungstate.
1) which was obtained after growth in the
presence of 10
7 M tungstate added
to the medium. The latter concentration was also optimal for aldehyde
dehydrogenase activity (0.56 unit mg
1).
Formate dehydrogenase activity was unaffected by higher tungstate concentrations added to the medium, whereas the aldehyde dehydrogenase activity declined by about 30% if 10
5
M tungstate was present during growth. In the absence of
tungstate, the addition of molybdate at physiological concentrations
(10
9 to 10
7
M) did not give rise to an aldehyde dehydrogenase activity.
However, if a high concentration of molybdate
(10
5 M) was added to the medium,
the specific activities of acetaldehyde dehydrogenase and formate
dehydrogenase increased to values of 14 and 62% of their maximum
activity, respectively. These activity levels corresponded to those
determined for a suboptimal tungstate supplementation by
10
9 M. Because molybdenum salts
are contaminated by tungsten (33), the induction of acetaldehyde
dehydrogenase and formate dehydrogenase activity by high molybdate
concentrations added to the medium might be caused by a contamination
by tungstate. However, it might also be possible that both
tungsten-containing enzymes incorporate molybdenum and form an active
enzyme when this metal is added in high concentrations to the medium.
At least, these physiological results suggested a strong preference for
tungstate compared with molybdate and a specific uptake of tungsten
even at quite low concentrations by growing cells.
1 mg of
protein
1 was obtained at 36 °C (Fig.
1). Significant lower values were obtained at room temperature (22 °C) and at 50 °C. This is in good agreement with the optimal growth temperature of E. acidaminophilum, which is between 30 and 35 °C (20). Uptake
experiments at 0 °C, performed as control, gave a strongly reduced
uptake rate of 0.1 pmol min
1 mg of
protein
1 (Fig. 1). Tungstate uptake was not
influenced by the presence of equimolar amounts of molybdate in the
uptake buffer (data not shown). No impairment of tungstate uptake was
observed if the ATPase inhibitor arsenate (5 mM) or the
uncoupler 2,4-dinitrophenol (2 mM) was present during the
uptake experiments by resting cells (data not shown).
View larger version (15K):
[in a new window]
Fig. 1.
Uptake of tungstate into E. acidaminophilum at different temperatures. Resting
cells of E. acidaminophilum were incubated with
[185W]tungstate as outlined under "Experimental
Procedures" at the indicated temperatures. At different time points
samples were withdrawn, and the radioactivity associated with the cells
was determined. , 0 °C;
, 15 °C;
, 22 °C;
,
36 °C;
, 50 °C.
ZAP-EcoRI library
of chromosomal DNA from E. acidaminophilum was screened. A
positive phage was isolated, and after in vivo excision
plasmid pKME5 containing a 6.9-kb EcoRI insert was obtained
(Fig. 2). On this 6.9-kb fragment seven open reading frames including a
truncated one were identified (Fig. 2A). The deduced amino
acid sequences of the genes tupABC located at the 5'-end of
this DNA fragment exhibited similarities to transporters of the
ABC-type uptake permeases (34, 35). TupA is similar to
substrate-binding lipoproteins (36), TupB to the permease part, and
TupC contains amino acid sequences similar to ATPases. The deduced
amino acid sequence of the other open reading frames exhibited
similarities to proteins involved in the biosynthesis of the
molybdopterin cofactor and were thus named: moeA,
moeA-1, moaA, and moaC' (Fig. 2). From
the last, a sequence of only 87 amino acids was encoded on the plasmid.
The genes tupA and tupB were identified by
Southern hybridization to be present as a single copy in the genome of
E. acidaminophilum (data not shown).
View larger version (8K):
[in a new window]
Fig. 2.
Organization of the sequenced tup
genes and the downstream region in E. acidaminophilum. A, the
restriction sites (EcoRI, SacI) and the probes
(probe A, probe B) relevant for cloning of the 6.87-kb EcoRI
fragment are shown. A putative termination signal is indicated.
B, putative transcripts were identified using RT-PCR and are
indicated.
Oligonucleotides used in this study
View larger version (69K):
[in a new window]
Fig. 3.
Anion binding to overexpressed TupA of
E. acidaminophilum. A, 11 µM TupA was incubated with a 90-fold excess of the
indicated anions and subjected to native polyacrylamide gel
electrophoresis (16). Separated proteins were stained with Serva
Brilliant Blue. Marker proteins are indicated at the left
side. Protein species indicated with a through e
were all identified to be TupA, which differed at its N terminus as
depicted in B. B, construction of the
overexpressed TupA protein and determined N-terminal sequences of the
five protein subspecies after heterologous expression and purification
of TupA by its Strep-tag. The designations a
through e correspond to those in A and
C. C, mobility shift assay with 11 µM TupA using different tungstate concentrations as
indicated.
where P0 is the protein concentration, and
A0 is the anion concentration with 50% of the
protein bound to the anion.
(Eq. 1)
View larger version (52K):
[in a new window]
Fig. 4.
Alignment of TupA from E. acidaminophilum with TupA-homologous proteins and with two
ModA proteins of known crystal structure. Two alignments are
combined in this figure. TupA was first aligned with ModA proteins and
subsequently with the TupA-homologous proteins from the indicated
organisms. Identical residues between TupA and ModA are
boxed; amino acids of ModA which are involved in
molybdate/tungstate binding are highlighted by gray boxes.
Identical residues in TupA and TupA-homologous proteins are printed in
bold. The salt bridge forming domain of phosphate-binding
protein from E. coli is shown below the respective sequence
of TupA and is boxed. Eco, ModA protein from E. coli (13); Avi, ModA protein from A. vinelandii (18);
Eac, TupA protein from E. acidaminophilum; Cje,
TupA-homologous protein from C. jejuni; Vch,
TupA-homologous protein from V. cholerae; Mth,
TupA-homologous protein from M. thermoautotrophicum; Hvo,
TupA-homologous protein from H. volcanii; Eco PBP,
phosphate-binding protein from E. coli (40).
View larger version (22K):
[in a new window]
Fig. 5.
Phylogenetic tree of TupA and other
anion-binding proteins from ABC transporters. ModA proteins: Hinf,
Hemophilus influenzae (accession no. P45323); Ecol, E. coli (P37329); Scar, Staphylococcus carnosus
(AAC83133); Cdif, Clostridium difficile (available from the
Sanger center); Rcap, Rhodobacter capsulatus (E36914); Avin,
A. vinelandii (ModA1; P37734; ModA2, 3891987); Hpyl,
Helicobacter pylori (AAD07541); Ani, A. nicotinovorans (CAA71776). Open reading frames similar to ModA:
Mkan, Orf2 M. kandleri (CAA67413). TupA-homologous
proteins: Mthe, M. thermoautotrophicum; Cjej, C. jejuni; Vcho, Vibrio cholerae; Hvol, H. volcanii; CysP: Ecol, E. coli (P16700). Sulfate-binding
protein: Ecol, E. coli (CAA26357). Phosphate-binding
protein: Ecol, E. coli (P06128); Spne, Streptococcus
pneumoniae (AAD22038). The tree was created using the program
ClustalW at the European Bioinformatics Institute. The bar
depicts 0.1 nucleotide substitution/site.
1) was identified in the sequence between
tupA and tupB, which might represent a
rho-independent transcriptional termination signal (Fig. 2).
Probably transcription started at a promoter structure upstream of
tupA, and one part of this transcript is terminated at the
loop structure between tupA and tupB, the other part runs through this structure and covers tupB. Thus, an
elevated transcription of tupA would be the result. By using
RT-PCR no common transcript of tupAB and tupC has
been identified. tupC was transcribed together with
moeA on one extra transcript (transcript II) and also
moeA-1, moaA, and moaC' were
transcribed together (transcript III). Probably the last transcript
extended into the unsequenced downstream region covering the complete
moaC gene. No difference in these mRNA pattern was
evident from RT-PCR data if RNA was isolated from cells grown with (1 µM) or without the addition of the anions tungstate and
molybdate to the medium (data not shown).
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
![]() |
ACKNOWLEDGEMENTS |
---|
We thank Karl Peter Ruecknagel for the N-terminal protein sequencing and Peter Planitz from Agilent Technologies (Waldbronn, Germany) for the inductively coupled plasma mass spectrometry analysis. We thank Ute Zindel and Kathrin Granderath for some FDH/AOR studies. We thank Roderick Brandsch (Freiburg) and Julia Vorholt (Marburg) for providing plasmids containing modAB of A. nicotinovorans and orf2 and orf3 of Methanopyrus kandleri, respectively. Thanks to Werner Klein from the GKSS in Geesthacht for the neutron irradiation of tungstate.
![]() |
FOOTNOTES |
---|
* This work was supported by grants from the Land Sachsen-Anhalt and the Fonds der Chemischen Industrie.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
The nucleotide sequence(s) reported in this paper has been submitted to the GenBankTM/EMBL Data Bank with accession number(s) AJ291988.
To whom correspondence should be addressed. Tel.: 49-345-552-6360;
Fax: 49-345-552-7010; E-mail: a.pich@mikrobiologie.uni-halle.de.
Published, JBC Papers in Press, April 5, 2001, DOI 10.1074/jbc.M101293200
2 A. Graentzdoerffer, A. Pich, and J. R. Andreesen, manuscript in preparation.
3 D. Rauh, A. Graentzdoerffer, A. Pich, and J.R. Andreesen, manuscript in preparation.
![]() |
ABBREVIATIONS |
---|
The abbreviations used are: kb, kilobase pair(s); RT-PCR, reverse transcription-polymerase chain reaction.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Kletzin, A., and Adams, M. W. (1996) FEMS Microbiol. Rev. 18, 5-63[CrossRef][Medline] [Order article via Infotrieve] |
2. | Ljungdahl, L. G. (1976) Trends Biochem. Sci. 1, 63-65 |
3. | Johnson, M. K., Rees, D. C., and Adams, M. W. W. (1996) Chem. Rev. 96, 2817-2839[CrossRef][Medline] [Order article via Infotrieve] |
4. | Chan, M. K., Mukund, S., Kletzin, A., Adams, M. W. W., and Rees, D. C. (1995) Science 267, 1463-1469[Medline] [Order article via Infotrieve] |
5. |
Mukund, S.,
and Adams, M. W. W.
(1995)
J. Biol. Chem.
270,
8389-8392 |
6. | White, H., and Simon, H. (1992) Arch. Microbiol. 158, 81-84[Medline] [Order article via Infotrieve] |
7. | Hu, Y., Faham, S., Roy, R., Adams, M. W. W., and Rees, D. C. (1999) J. Mol. Biol. 286, 899-914[CrossRef][Medline] [Order article via Infotrieve] |
8. |
Yamamoto, I.,
Saiki, T.,
Liu, S. M.,
and Ljungdahl, L. G.
(1983)
J. Biol. Chem.
258,
1826-1832 |
9. | Vorholt, J. A., Vaupel, M., and Thauer, R. K. (1997) Mol. Microbiol. 23, 1033-1042[CrossRef][Medline] [Order article via Infotrieve] |
10. |
Meckenstock, R. U.,
Krieger, R.,
Ensign, S.,
Kroneck, P. M.,
and Schink, B.
(1999)
Eur. J. Biochem.
264,
176-182 |
11. | Leonhardt, U., and Andreesen, J. R. (1977) Arch. Microbiol. 115, 277-284[Medline] [Order article via Infotrieve] |
12. | Wagner, R., and Andreesen, J. R. (1987) Arch. Microbiol. 147, 295-299 |
13. | Maupin-Furlow, J. A., Rosentel, J. K., Lee, J. H., Deppenmeier, U., Gunsalus, R. P., and Shanmugam, K. T. (1995) J. Bacteriol. 177, 4851-4856[Abstract] |
14. | Grunden, A. M., and Shanmugam, K. T. (1996) Arch. Microbiol. 168, 345-354[CrossRef] |
15. | Pau, R. N., Klipp, W., and Leimkühler, S. (1997) in Transition Metals in Microbial Metabolism (Winkelmann, G. , and Carrano, C. J., eds) , pp. 217-234, Harwood Academic Publishers, Amsterdam |
16. |
Rech, S.,
Wolin, C.,
and Gunsalus, R. P.
(1996)
J. Biol. Chem.
271,
2557-2562 |
17. | Hu, Y., Rech, S., Gunsalus, R. P., and Rees, D. C. (1997) Nature 4, 703-707 |
18. | Lawson, D. M., Williams, C. E., White, D. J., Choay, A. P., Mitchenall, L. A., and Pau, R. N. (1997) J. Chem. Soc. Dalton Trans. 3981-3984 |
19. | Lawson, D. M., Williams, C. E., Mitchenall, L. A., and Pau, R. N. (1998) Structure 15, 1529-1539 |
20. | Zindel, U., Freudenberg, W., Rieth, M., Andreesen, J. R., Schnell, J., and Widdel, F. (1988) Arch. Microbiol. 150, 254-266 |
21. | Graentzdoerffer, A. (2000) Formiat-Stoffwechsel in Eubacterium acidaminophilum: Molekulare und biochemische Charakterisierung der Wolfram- und Selen-haltigen Formiat-Dehydrogenasen sowie einer Eisen-Hydrogenase. Ph.D. thesis , University of Halle |
22. | Rauh, D. (2000) Isolierung: Charakterisierung und molekulare Analyse der wolframhaltigen Aldehyd-Oxidoreduktase aus Eubacterium acidaminophilum. Diploma thesis , University of Halle |
23. | Sambrook, J., Fritsch, E. F., and Maniatis, T. (1998) Molecular Cloning: A Laboratory Manual , 2nd Ed. , Cold Spring Harbor Laboratory, Cold Spring Harbor, NY |
24. | Saito, H., and Miura, K. I. (1963) Biochim. Biophys. Acta 72, 619-620[CrossRef] |
25. | Sanger, F., Nicklen, S., and Coulson, A. R. (1977) Proc. Natl. Acad. Sci. U. S. A. 74, 5463-5467[Abstract] |
26. | Graentzdoerffer, A., Pich, A., and Andreesen, J. R. (2001) Arch. Microbiol. 175, 8-18[CrossRef][Medline] [Order article via Infotrieve] |
27. | Bradford, M. M. (1976) Anal. Biochem. 72, 248-254[CrossRef][Medline] [Order article via Infotrieve] |
28. | Gill, S. C., and von Hippel, P. H. (1989) Anal. Biochem. 182, 319-326[Medline] [Order article via Infotrieve] |
29. | Laemmli, U. K. (1970) Nature 277, 680-685 |
30. | Towbin, H., Staehelin, T., and Gordon, J. (1992) Bio/Technology 24, 145-149[Medline] [Order article via Infotrieve] |
31. | Meyer, M., Granderath, K., and Andreesen, J. R. (1995) Eur. J. Biochem. 234, 184-191[Abstract] |
32. | Granderath, K. (1993) Charakterisierung der Formiat-Dehydrogenase und Aldehyd-Dehydrogenase als Wolframhaltige Proteine von Eubacterium acidaminophilum. Ph.D. thesis , University of Göttingen |
33. | Herzig, A. J., and Briggs, J. Z. (1968) in Encyclopedia of the Chemical Elements (Hampel, C. A., ed) , pp. 412-422, Reinhold Book Corp., New York |
34. | Nikaido, H., and Hall, J. A. (1998) Methods Enzymol. 292, 3-20[Medline] [Order article via Infotrieve] |
35. |
Saier, M. H., Jr.
(2000)
Microbiology
146,
1775-1795 |
36. |
Sutcliffe, I. C.,
and Russel, R. R. B.
(1995)
J. Bacteriol.
177,
1123-1128 |
37. | Chambaud, I., Wroblewski, H., and Blanchard, A. (1999) Trends Microbiol. 7, 493-499[CrossRef][Medline] [Order article via Infotrieve] |
38. | Williams, C. E., White, D. J., Delarbre, L., Mitchenall, L. A., Pau, R. N., and Lawson, D. M (1999) Acta Crystallogr. Sect. D Biol. Crystallogr. 55, 1356-1358[CrossRef][Medline] [Order article via Infotrieve] |
39. |
Wanner, C.,
and Soppa, J.
(1999)
Genetics
152,
1417-1428 |
40. | Luecke, H., and Quiocho, F. A. (1990) Nature 347, 402-406[CrossRef][Medline] [Order article via Infotrieve] |
41. | Cserzo, M., Wallin, E., Simon, I., von Heinje, G., and Elofson, A. (1997) Prot. Eng. 10, 673-676[Abstract] |
42. |
Mourez, M.,
Hofnung, M.,
and Dassa, E.
(1997)
EMBO J.
16,
3066-3077 |
43. | Walker, J. E., Saraste, M., Runswick, M. J., and Gay, N. J. (1982) EMBO J. 1, 945-951[Medline] [Order article via Infotrieve] |
44. | Hung, L. W., Wang, I. X., Nikaido, K., Liu, P. Q., Ames, G. F., and Kim, S. H. (1998) Nature 396, 703-707[CrossRef][Medline] [Order article via Infotrieve] |
45. |
Hasona, A.,
Ray, R. M.,
and Shanmugam, K. T.
(1998)
J. Bacteriol.
180,
1466-1472 |
46. | Hasona, A., Self, W. T., Ray, R. M., and Shanmugam, K. T. (1998) FEMS Microbiol. Lett. 169, 111-116[CrossRef][Medline] [Order article via Infotrieve] |
47. | Rivers, S. L., McNairn, E., Blasco, F., Giordano, G., and Boxer, D. H. (1993) Mol. Microbiol. 8, 1071-1081[Medline] [Order article via Infotrieve] |
48. |
Allen, R. M.,
Roll, J. T.,
Rangaraj, P.,
Shah, V. K.,
Roberts, G. P.,
and Ludden, P. W.
(1999)
J. Biol. Chem.
274,
15869-15874 |
49. | Menendez, C., Siebert, D., and Brandsch, R. (1996) FEBS Lett. 391, 101-103[CrossRef][Medline] [Order article via Infotrieve] |
50. | Menendez, C., Igloi, G., Henninger, H., and Brandsch, R. (1995) Arch. Microbiol. 164, 142-151[CrossRef][Medline] [Order article via Infotrieve] |
51. | Imperial, J., Hadi, M., and Amy, N. K. (1998) Biochim. Biophys. Acta 1370, 337-346[Medline] [Order article via Infotrieve] |
52. | Anderson, L. A., Palmer, T., Price, N. C., Bornemann, S., Boxer, D. H., and Pau, R. N. (1997) Eur. J. Biochem. 246, 119-126[Abstract] |
53. | Wagner, U. G., Stupperich, E., and Kratky, C. (2000) Structure 8, 1127-1136[CrossRef][Medline] [Order article via Infotrieve] |
54. | Shan, S., Loh, S., and Herschlag, D. (1996) Science 272, 97-101[Abstract] |
55. | Hochheimer, A., Schmitz, R. A., Thauer, R. K., and Hedderich, R. (1995) Eur. J. Biochem. 234, 910-920[Abstract] |