From the Institut für Biologie,
Humboldt-Universität zu Berlin, Chausseestrasse 117, 10115 Berlin, Germany, the § Swammerdam Institute for Life
Sciences, Department of Biochemistry, University of Amsterdam, Plantage
Muidergracht 12, NL-1018 TV Amsterdam, The Netherlands, and the
¶ Instituto de Catalisis, CSIC, Campus Universidad
Autonoma-Cantoblanco, 28049 Madrid, Spain
Received for publication, October 26, 2000, and in revised form, February 6, 2001
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ABSTRACT |
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Previous genetic studies have revealed a
multicomponent signal transduction chain, consisting of an
H2 sensor, a histidine protein kinase, and a response
regulator, which controls hydrogenase gene transcription in the
proteobacterium Ralstonia eutropha. In this study, we
isolated the H2 sensor and demonstrated that the purified
protein forms a complex with the histidine protein kinase. Biochemical
and spectroscopic analysis revealed that the H2 sensor is a
cytoplasmic [NiFe]-hydrogenase with unique features. The
H2-oxidizing activity was 2 orders of magnitude lower than that of standard hydrogenases and insensitive to oxygen, carbon monoxide, and acetylene. Interestingly, only H2 production
but no HD formation was detected in the
D2/H+ exchange assay. Fourier transform
infrared data showed an active site similar to that of standard
[NiFe]-hydrogenases. It is suggested that the protein environment
accounts for a restricted gas diffusion and for the typical kinetic
parameters of the H2 sensor. EPR analysis demonstrated that
the [4Fe-4S] clusters within the small subunit were not reduced under
hydrogen even in the presence of dithionite. Optical spectra revealed
the presence of a novel, redox-active, n = 2 chromophore that is reduced by H2. The possible involvement of this chromophore in signal transduction is discussed.
The detection of physiologically important gases by organisms is
mediated by biological sensors that convert the molecular signal into a
cellular response. Sensors for O2, CO, and NO have been
described, and the signaling mechanism is the subject of current
research (1-3). One of the best studied examples is the two-component
FixL-FixJ system of Rhizobium meliloti and
Bradyrhizobium japonicum. In this case the presence of
O2 is detected by a heme-containing histidine protein
kinase (4). The heme group in FixL binds the oxygen molecule that
induces a transition of the ferrous iron from high spin to low spin.
This triggers the inactivation of the kinase domain of FixL. The
release of O2, at low O2 tensions, restores the
S = 2 state of the heme iron, which in turn leads to activation of
the kinase by autophosphorylation. Subsequent phosphoryl transfer
to the response activator FixJ finally stimulates gene transcription
(5).
In the facultative chemolithotrophic proteobacterium Ralstonia
eutropha (formerly Alcaligenes eutrophus), the
oxidation of molecular hydrogen is catalyzed by two
[NiFe]-hydrogenases, a membrane-bound
(MBH)1 and a cytoplasmic
NAD-reducing hydrogenase (SH) (6, 7). The structural genes of both
[NiFe]-hydrogenases together with sets of accessory genes are grouped
in the MBH and SH operons, which are induced in the presence of
molecular hydrogen (8, 9). Hydrogenase gene transcription is controlled
by a multicomponent regulatory system consisting of the proteins HoxB,
HoxC, HoxJ, and HoxA, which are encoded in the MBH operon (8-10). HoxJ
and HoxA share typical features of a bacterial two-component regulatory system that recognizes and responds to various environmental stimuli (9, 11). Our studies showed that HoxJ displays autokinase activity (9)
and communicates with the activator
HoxA,2 a member of the NtrC
family of response regulators (12). HoxA, the final target of the
H2-sensing signal transduction chain, binds to the upstream
region of the hydrogenase promoters and activates open complex
formation by Genetic studies revealed that recognition of H2 requires in
addition to HoxA and HoxJ the protein HoxBC (9). Proteins similar to
HoxBC, designated HupUV, have been identified in Rhodobacter capsulatus and B. japonicum (14, 15). HoxBC-like
proteins show typical features of a [NiFe]-hydrogenase (16). Although HoxBC is essential for lithoautotrophic growth of R. eutropha (9), it cannot compensate for the loss of the MBH and the
SH. This observation points to a regulatory rather than an
energy-yielding function of the HoxBC protein (16). The low level of
expression combined with an extremely low activity allowed only
preliminary biochemical analysis of HoxBC in crude extracts (17).
Attempts to express a functional HoxBC protein in Escherichia
coli were unsuccessful. This prompted us to develop a homologous
overexpression system in R. eutropha (16) and to use it
successfully for the purification of HoxBC, later named regulatory
hydrogenase (RH).
Biochemical and spectroscopic analysis of the homogenous RH uncovered
unique enzymatic features that are clearly distinct from the properties
of standard hydrogenases. The data suggest that the RH shows a common
[NiFe] active site but displays significant changes in the protein
environment. In order to study the mechanism of H2 signal
transduction in more depth, we began to establish an in
vitro system, using purified components. First data show that the
RH forms a specific complex with the sensor kinase HoxJ, supporting the
notion that the RH is a direct component of the signal transduction chain.
Cell Growth--
R. eutropha strain HF371, a
derivative of R. eutropha H16, harboring plasmid pGE378, was
used for protein purification (16). Cells were heterotrophically grown
in a mineral medium in a 10-liter Braun Biostat fermentor (Braun,
Melsungen, Germany) at 30 °C under hydrogenase derepressing
conditions. At an OD436 of 11 the cells were harvested, washed in 50 mM potassium phosphate buffer,
pH 7.0 (K-PO4 buffer), and stored frozen in liquid nitrogen.
RH Purification--
Cells (83 g, wet weight) were resuspended
in 30 ml of K-PO4 buffer containing 0.1 mM
phenylmethylsulfonyl fluoride. Cells were disrupted by two passages
through a chilled Amicon French press cell at 1100 pounds/square inch
(75.8 bar). Soluble extracts were prepared by high speed centrifugation
(100 000 × g, 60 min, 4 °C). The resulting
supernatant was degassed and saturated with hydrogen. The extract was
kept under an atmosphere of 100% H2 and subsequently
incubated for 10 min at 65 °C. After the heat treatment the sample
was chilled on ice. All further purification steps were carried out
under air. The denatured proteins were removed by centrifugation (13 000 × g, 20 min, 4 °C), and the supernatant was
fractionated by addition of
(NH4)2SO4 to give a final
concentration of 1 M. The precipitated proteins were
removed by centrifugation (13 000 × g, 20 min,
4 °C), and the clear supernatant was directly applied to a POROS
20ET column (Applied Biosystems; ethyl ether; 10 × 100 mm),
preequilibrated with K-PO4 buffer containing 1 M (NH4)2SO4 at a flow
rate of 40 ml/min (BioCAD Perfusion Chromatography Workstation). The
column was washed with 2 bed volumes of K-PO4 buffer
containing 1 M
(NH4)2SO4. The protein was eluted
with K-PO4 buffer containing 0.4 M
(NH4)2SO4, and fractions of 4 ml
were collected. The active fractions of several column runs were
combined, concentrated, and dialyzed against K-PO4 buffer.
The RH was further purified on a POROS 20HQ column (Applied Biosystems;
quarternized polyethyleneimine; 4.6 × 100 mm) preequilibrated
with K-PO4 buffer. The eluent was pooled, concentrated
(Centriprep-10; Amicon), and directly frozen in liquid N2.
Protein concentrations were determined according to the methods of
Lowry et al. (18).
Immunological Procedures--
Proteins were resolved by
electrophoresis in 12% polyacrylamide/SDS gels and transferred to
Protran BA85 nitrocellulose membranes (Schleicher & Schüll). HoxC
was detected with anti-HoxC serum, diluted 1:1000, and an
alkaline-phosphatase-labeled goat anti-rabbit IgG (Dianova, Hamburg, Germany).
Complex Formation Assay--
The histidine protein kinase HoxJ
was overproduced in E. coli and purified as a
polyhistidine-tagged protein, His6-HoxJ, by metal chelate
affinity chromatography (9). Purified His6-HoxJ and RH were
mixed and subsequently applied to a native 4-15% polyacrylamide gel.
Native gel electrophoresis was carried out as described previously (19). Complex formation was either monitored by in-gel hydrogenase activity staining (19) or by protein staining using Coomassie Blue.
Metal Analysis--
Nickel and iron were determined with a
Hitachi 180-80 polarized Zeeman atomic absorption spectrophotometer
against a standard series. Samples were made devoid of extraneous metal
ions by passage over a Chelex-100 column (Bio-Rad).
Activity Measurements--
Hydrogen uptake activity was measured
amperometrically at 30 °C in a cell (2.15 ml) with 50 mM
Tris-HCl, pH 8.0, using a Clark-type electrode (YSI 5331) according to
Coremans et al. (20). As O2 did not affect the
activity, no efforts were made to remove air. Hydrogen, in the form of
H2-saturated water, was added to final concentrations
varying from 36 to 100 µM. As electron acceptor either
benzyl viologen (4.2 mM, Em = -359 mV)
or methylene blue (4 mM,
Em, pH 7 = +11 mV) were used. The
measured specific activities were plotted against the H2
concentration. The dependence was simulated using the program Leonora
by Cornish-Bowden, assuming Michaelis-Menten kinetics (21). Protein
concentrations in the assay were typically 2.5-5 nM RH
(
D+/H+ exchange activity was measured in a
stirred membrane leak chamber fitted to a mass spectrometer (Masstorr
200 DX quadrupole, VG Quadrupoles Ltd.). Two different assays were
used. In the first assay 10 ml of Mes/Mops/Tris buffer solution (ionic
strength 90 mM; pH 6.5) was saturated with 20%
D2 and 80% argon, and 1 µmol of sodium dithionite was
added to eliminate residual oxygen. The reaction was started by the
addition of RH ( EPR Spectroscopy--
X-band (9.4 GHz) spectra with a 100 kHz
field modulation frequency were recorded on a Bruker ECS106 EPR
spectrometer equipped with an Oxford Instruments ESR900 helium flow
cryostat with an ITC4 temperature controller. The magnetic field was
calibrated with an AEG Magnetic Field Meter. The frequency was measured
with a Hewlett-Packard 5350B Microwave Frequency Counter. Illumination of the samples was performed by shining white light (Osram Halogen Bellaphot, 150 watts) via a light guide through the irradiation grid of
the Bruker ER 4102 ST cavity. Spectra were simulated according to the
formulas published by Beinert and Albracht (24).
FTIR Spectroscopy--
Fourier transform infrared (FTIR) spectra
were taken on a Bio-Rad FTS 60A spectrometer equipped with an MCT
detector. Spectra were recorded at room temperature with a resolution
of 2 cm Ultraviolet/Visible Spectroscopy--
Optical spectra were taken
on an Aminco DW-2000 spectrophotometer interfaced with an IBM computer.
Purification of the RH Protein--
To avoid interferences with
the dominant activities of the MBH and the SH, we started the
purification of the RH protein with mutant R. eutropha
HF371, in which the MBH and SH genes had been deleted by mutation.
After cell disruption and high speed centrifugation, the soluble
extract was incubated at 65 °C for 10 min under an atmosphere of
H2 (100%). The heat treatment was necessary prior to high
resolution hydrophobic interaction chromatography to provide an
effective and rapid isolation of the RH. Purification to apparent homogeneity was achieved by subsequent anion exchange chromatography. The total procedure is summarized in Table
I. Beginning with 83 g of cells (wet
weight), 10.9 mg of RH was obtained. The specific activity of the
preparation was 0.94 units/mg of protein with methylene blue as the
electron acceptor. The H2 concentration in the assay was
57.8 µM. The protein was purified 26-fold with a yield of
6%.
Biochemical Properties--
Two protein bands occurred after
denaturing the RH by SDS-PAGE corresponding to molecular masses of 37 and 55 kDa, respectively (Fig.
1A). These values are in good
agreement with those predicted from the nucleotide sequence of
hoxB (36.5 kDa) and hoxC (52.4 kDa). The identity
of the purified protein was confirmed by immunoblot analysis, using an
antibody raised against the HoxC subunit of the RH (Fig.
1B). Analysis of the enzyme on a Superdex G-200 (Amersham Pharmacia Biotech) revealed a single peak corresponding to a molecular mass of ~165 kDa (data not shown) indicating that the RH was purified as a tetramer with an
The oxidation of H2 by the purified RH turned out to be
O2-insensitive. The level of activity was the same in
aerobic and anaerobic buffers. Moreover, the rate of H2
oxidation determined with methylene blue as the electron acceptor did
not show the typical lag phase that is found with most as isolated
[NiFe]-hydrogenases. This observation is consistent with the result
obtained with soluble extract (17). The Km for
H2 was 25 ± 5 µM, and the calculated
specific activity at Vmax conditions was
1.2 ± 0.2 units/mg of protein. The activity of the RH remained
constant over a broad pH range between 5 and 10 irrespective of the
used buffers (potassium acetate, K-PO4, and Tris-HCl; 50 mM each), whereas most hydrogenases show a distinct pH
optimum. In contrast to the H2 uptake activity, the
production of H2 by the RH was pH-dependent.
Highest H2 evolution rates (0.8 units/mg of protein) were
obtained at pH 4.0 with benzyl viologen as electron donor. Acetylene
has been shown to be a competitive inhibitor for several hydrogenases
(25, 26). Incubation of the RH with C2H2 did not affect the RH activity (data not shown).
Storage of the purified RH at 4 °C under air or an atmosphere of
100% O2 resulted in a loss of 50% of the
H2-dependent methylene blue-reducing activity
within 48 h. Replacement of the air atmosphere by 100% argon or
N2 caused a decrease of 20% of the activity within the
same period. Addition of metal ions (Fe3+,
Ni2+, Mn2+, Mg2+, and
Zn2+) or glycerol (20%) or addition of KCl up to 0.5 M did not affect the stability of the RH. The supply of
dithionite or ferricyanide under anoxic conditions also did not
contribute to the stability of the RH. Moreover, storage of the
isolated RH under an atmosphere of 100% H2 inactivated the
RH rapidly; 50% of its activity disappeared within 12 h. The
H2 sensitivity contrasts with data obtained with the
soluble extract, which showed constant RH activity over a period of
24 h under comparable conditions. In all cases inactivation of the
RH was irreversible.
D+/H+ Exchange Activity--
The
D+/H+ exchange assay with the RH yielded only
H2 production but no HD formation (Fig.
2A). The initial rate of
H2 production at pH 6.5 was 2.1 ± 0.1 units/mg of
protein. This behavior is distinct from that of other
[NiFe]-hydrogenases, which show higher initial rates of HD production
than of H2 production. When the exchange activity assay was
measured in deuterated water saturated with H2 some HD
production was detected with the RH, although the rate of
D2 evolution was definitively higher (Fig. 2B).
The initial rate of D2 production at pD 6.5 was 1.3 ± 0.2 units/mg of protein, whereas the initial rate of HD production was
0.5 ± 0.1 units/mg of protein. The pH optimum of the
D+/H+ exchange activity of the RH was at pH 5.5 (data not shown).
EPR Spectroscopy--
Preliminary studies of the RH in crude cell
extracts prohibited a study of the EPR properties of its Fe-S clusters
(17). The purified enzyme now allowed this approach. The as
isolated RH showed no EPR signals at temperatures between 4.2 and 100 K. Also after addition of the oxidizing agent DCIP
(dichlorophenolindophenol, Em = +230 mV), no signal
occurred. Upon reduction of the RH (15 min under 100% H2
at room temperature in 50 mM Tris-HCl, pH 8.0), a rhombic
EPR signal with g values at 2.19, 2.13, and 2.01 appeared (Fig.
3, trace A). The
double-integrated intensity of the signal amounted to a spin
concentration equal to 69% of the nickel concentration.
The EPR signal is very similar to the well studied Nia-C*
signal observed in standard hydrogenases (e.g. from
Desulfovibrio gigas and Allochromatium vinosum),
and it is due to a paramagnetic state of the active site nickel in the
3+ state (27). A typical feature of enzyme in the Nia-C*
state is its light sensitivity at cryogenic temperatures, yielding the
so-called Nia-L* signal as a result of the
photodissociation of a hydrogen (28). A model for this
photodissociation has been described by Happe et al. (27).
In the case of the RH the Nia-C* signal also showed this
light-sensitive behavior. Upon illumination at 30 K a spectrum
(Nia-L*: gxyz = 2.045, 2.09, and 2.24), only
slightly different from the Nia-L* signal of standard
hydrogenases, appeared (Fig. 3, trace B). The small
difference concerns the position of the gz (2.24 in the RH as
compared with 2.28/2.30 in standard [NiFe] hydrogenases). This points
to a small structural difference around the active site nickel. Upon
warming of the sample to 200 K for 15 min in the dark a third,
transient, spectrum came up with g values at 2.047, 2.069, and 2.30 (Fig. 3, trace C). Only after several hours at 200 K the
sample returned to the Nia-C* state.
Contrary to observations in standard [NiFe]-hydrogenases no signal of
a [3Fe-4S]+ cluster could be observed in the oxidized
protein, not even after treatment with excess DCIP. This is in
agreement with the presence of three [4Fe-4S] clusters as predicted
from the amino acid sequence data. When the protein was treated with
100% H2, however, no signals due to reduced cubanes were
detectable, not even if 20 mM dithionite was added. None of
the nickel signals (Nia-C*, Nia-L*, or the transient signal) showed any spin coupling due to a reduced proximal [4Fe-4S] cluster (Fig. 3). This indicates that this cluster was in
the oxidized, diamagnetic state in the RH under H2. In
standard [NiFe]-hydrogenases the proximal cluster is usually reduced
under 100% H2. The interaction of the nickel with the
reduced proximal cluster is observed as a clear 2-fold splitting of the
Nia-C* signal at 4.5 K. At low temperatures it was also
possible to completely saturate the Nia-C* signal at high
microwave power (260 milliwatts), which is again indicative of an
oxidized proximal cluster (28, 29). Reduction of the RH with dithionite
in the presence or absence of low potential electron acceptors (methyl
viologen and benzyl viologen) under 100% H2 did not evoke
any signal of a reduced Fe-S cluster. Also inspection of the integrated
EPR signals did not uncover any broad signal due to reduced Fe-S
clusters as can be seen in the right-hand panel in Fig. 3
for the Nia-L* signal.
FTIR Spectroscopy--
The FTIR measurements on purified RH
confirmed the presence of only two redox states described earlier to be
present in the RH from soluble extracts (17). Untreated protein showed
a spectrum (Fig. 4A) with two
small bands (2082 and 2071 cm
When the gas phase was changed from 100% H2 to 100% CO
(equilibration time 60 min) a mixture of the Nia-C* and
Nia-S state was observed (Fig. 4B). The spectrum
clearly showed that it was not possible for exogenous CO to bind to the
active site of the RH since no extra peak around 2060 cm UV-Visible Absorption Spectroscopy--
UV-visible spectra of
oxidized and reduced RH showed differences in absorption between the
two species. Incubation of the RH under 100% H2 resulted
in an increase in absorption in the 250-280 and 300-400 nm spectral
regions (Fig. 5). The difference spectrum
of reduced minus oxidized RH showed a large peak at 251 nm and a
smaller one at 342 nm with an apparent shoulder at 305 nm. The
calculated Complex Formation--
To elucidate the nature of the interaction
between the RH and the signal transduction chain, purified kinase HoxJ
and the RH were mixed, and the sample was subjected to native PAGE
(Fig. 6). In one experiment the gel was
resolved by protein staining (Fig. 6A), and in the parallel
experiment a hydrogenase in-gel activity staining with phenazine
methosulfate as electron acceptor was performed (Fig. 6B).
Incubation of the RH with increasing amounts of HoxJ led to a bandshift
indicating the formation of a high molecular weight complex (Fig.
6A, lanes 2-4). A bandshift was not observed in the control
containing the RH and an excess of bovine serum albumin (Fig. 6,
A and B, lane 5). The in-gel assay (Fig.
6B) demonstrated that the hydrogenase activity of the RH was
maintained at high level upon complex formation (Fig. 6B, lanes
3 and 4). Exposure to H2 resulted in
considerable loss of hydrogenase activity (Fig. 6B, lane 6),
which is consistent with the observed instability of the RH in the
presence of H2.
Genetic and biochemical studies uncovered a signal transduction
chain, which directs H2-dependent gene
activation in R. eutropha. This signal transduction chain
consists of the transcription activator HoxA, the histidine protein
kinase HoxJ, and the H2 sensor RH. The RH is absolutely
necessary for the recognition of dihydrogen suggesting its primary role
in signal reception (9). Sequence alignment revealed that the RH
contains typical signatures of [NiFe]-hydrogenases (16), and a
preliminary EPR and FTIR study showed an active site similar to that of
prototypic [NiFe]-hydrogenases (17). Characterization of the purified
RH achieved in this study confirmed some biochemical features that are
compatible to those of standard [NiFe]-hydrogenases. On the other
hand, some characteristics were uncovered that are obviously uniquely
assigned to the subgroup of H2-sensing proteins (16).
Unlike standard [NiFe]-hydrogenases, which usually have
H2 uptake activities of about 200-300 units/mg of protein,
the RH displayed a specific activity at Vmax of
only 1.2 ± 0.2 units/mg protein. Moreover, in the air-oxidized
state the RH showed no lag phase, suggesting that it does not require a
reductive activation step before the protein is enzymatically active.
Interestingly, the activity of the RH was not inhibited by
O2, CO, or C2H2. Most hydrogenases
are sensitive to these gases with the exception of the SH of R. eutropha (33). In this case a modified catalytic center probably
excludes the binding of CO and O2 (34). Although the EPR
and FTIR spectra of the [NiFe] site of the RH resemble those of
standard [NiFe]-hydrogenases, the active site of the RH exhibits some
important redox differences. Only the Nia-S and
Nia-C* states are attainable, and CO cannot bind to the
active enzyme. This indicates that the nickel site (where in standard
[NiFe]-hydrogenases CO binds and where H2 is proposed to
react under turnover conditions (27)) is altered such that it cannot
react with CO or H2. This would restrict the reaction with
H2 to the iron site resulting in the Nia-C*
state only. The very low activity of the RH is in line with this idea. The D2/H+ exchange data suggest that
D2 diffusion to and from the active site is severely
restricted resulting in a molecular cage effect (35). The formed HD
then reacts again to form H2, before diffusion of HD from
the enzyme to the bulk occurs. In the H2/D+
exchange, the formed D2 escapes slower than HD allowing
some HD detection. The gas channel detected in the x-ray structures of
[NiFe]-hydrogenases (36, 37) points right to the nickel site. Changes
in the amino acid composition of this channel close to the nickel site,
e.g. the presence of more bulky residues, could explain both
the redox and the exchange properties. The kinetic behavior of the RH
in the D2/H+ activity assay is in agreement
with the low activity of the RH in the other assays.
The described EPR and FTIR data on the purified RH do not differ from
those presented earlier for the protein in crude extracts, so
purification does not change these properties. A previously unobserved
state occurred when the Nia-L* state was warmed up to 200 K. A transient state was then observed with g values at 2.047, 2.069, and 2.30. This points to changes induced in the vicinity of the nickel
site. As yet, we do not understand the nature of these changes.
Another typical feature of the light sensitivity in the RH is that all
conversions are much slower than in several other hydrogenases tested
in this laboratory using the same experimental set up (e.g. Allochromatium vinosum, Methanococcus
thermoautotrophicum, and Wollinella succinogenes). The
Nia-C* to Nia-L* conversion in membrane-bound hydrogenase (MBH) of A. vinosum is completed within 5 min,
whereas in the RH it took about 15 min. The difference in the reverse reaction was even more pronounced. After 2 h at 200 K the RH was still in the transient dark state, whereas the MBH of A. vinosum requires only 10-15 min at 200 K to return completely to
the Nia-C* state. This slow photolysis and the extremely
slow annealing might be due to a less spacious, obstructed active site.
It was shown that it was impossible to reduce the three [4Fe-4S]
clusters (predicted to be present from sequence data), although highly
reductive conditions were applied (100% H2 with benzyl viologen, methyl viologen and/or 20 mM dithionite). Also no
splitting of the Nia-C* or Nia-L* signals at
4.5 K by a reduced proximal cluster was observed.
Our current model of signal transduction in the RH is as follows;
H2 binds to the active site (presumably at the iron site (27)) and causes a formal oxidation of the nickel ion from the 2+ to
the 3+ state. The released electron is transferred to the Fe-S
clusters. However, no spectroscopic evidence for a reduced Fe-S cluster
was found, and no other S = 1/2 EPR signal was detected. Since the
RH is apparently functional as an Transmission of the H2-induced changes in the RH to the
histidine protein kinase HoxJ proceeds via direct protein-protein interaction as shown by complex formation. The N-terminal part of HoxJ,
the so-called input domain, is the most likely region for the
signal-accepting site. Sequence comparison revealed that this domain is
a member of the PAS domain superfamily, which is found in a wide
variety of regulatory systems involved in the sensing of light, oxygen,
or redox potential (38- 40). Several PAS domain proteins mediate signal
transmission by the way of an associated cofactor (40), like the FAD in
the aerotaxis signal transducer Aer of E. coli (41). Such a
two-electron cofactor in HoxJ might be a good candidate to be reduced
by the yet unidentified cofactor in the RH. In this scenario, electron
flow from the RH to the histidine kinase should induce a conformational
switch to modulate the activity of the HoxJ transmitter domain and
thereby affect the autophosphorylation activity of HoxJ. To resolve
such a mechanism we intend to block electron transport within the RH by
site-directed mutagenesis. Attractive targets will be the ligands of
the three FeS clusters and the nonmetal cofactor of the RH.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
54 RNA polymerase (8, 13).
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
2
2). Benzyl
viologen-dependent H2 evolution was determined
amperometrically at 30 °C. The reaction mixture contained 50 mM acetate buffer, 1 mM benzyl viologen, and 3 mM sodium dithionite.
2
2) to a final
concentration of 0.12 µM. Masses 1-6 were scanned at 1 atomic mass unit/s. In the second assay the buffer solution was in
99.9% D2O (Aldrich) and saturated with H2. A
control experiment was done in D2/D2O in order
to evaluate the HD production catalyzed by the protein due to
contaminant H+. This effect was subtracted to the
H2/D2O assay. The pD of the assay mixtures was
measured with a glass electrode calibrated with pH standards in
H2O. It was taken into account that pD = pH + 0.41 (22, 23). All experiments were done at 30 °C.
1. Typically, averages of 1524 spectra were taken against proper blanks. Enzyme samples (10 µl) were
loaded into a gas-tight transmission cell (CaF2, 56 µm
path length). The spectra were corrected for the base line using a
spline function provided by the Bio-Rad software.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
Purification of the R. eutropha regulatory hydrogenase
2
2 structure.
Atomic absorption spectroscopy showed an average metal content of 11.2 iron/nickel. After Chelex treatment this ratio decreased to 7.6 iron/nickel. The activity after the Chelex-100 column was 75% of the
initial activity.
View larger version (39K):
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Fig. 1.
Purification of the RH. A,
Coomassie Blue staining; B, immunoblot analysis of protein
samples from various purification steps after separation by SDS-PAGE.
The subunits HoxB and HoxC of the RH are indicated by an arrow.
Lane 1, soluble extract; lane 2, supernatant
after treatment at 65 °C for 10 min; lane 3, supernatant
of 25% (NH4)2SO4 precipitation;
lane 4, hydrophobic interaction; lane 5, anion
exchange chromatography.
View larger version (12K):
[in a new window]
Fig. 2.
D+/H+ exchange
activity of the RH. A, D2/H+
exchange kinetics catalyzed by the RH from R. eutropha in
Mes/Mops/Tris buffer solution (ionic strength 90 mM, pH
6.5) saturated with 20% D2 and 80% argon. B,
H2/D+ exchange kinetics measured in
Mes/Mops/Tris buffer solution in D2O (ionic strength 90 mM, pD 6.5) saturated with 100% H2. The
arrow marks the addition of the RH to a final concentration
of 0.12 µM. Masses 2-4 were recorded.
View larger version (15K):
[in a new window]
Fig. 3.
EPR spectra at 4.5 K of the RH under 100%
H2. Left-hand panel: A, the
Nia-C* state. B, after illumination at 30 K
(Nia-L* state). C, after 15 min at 200 K in the
dark a transient state was observed. Relative gains for A,
B, and C are 1.6, 1, and 1. Right-hand
panel: the light-induced Nia-L* state is shown over a
much wider field sweep (400 millitesla), either as the direct
spectrum (E) or as the first integral (D).
Trace D shows that there are no other paramagnets with broad
signals hidden in the base line of the first derivative. All spectra
were recorded with a microwave power incident to the cavity of 0.26 microwatts.
1) and one large
band (1943 cm
1) in the 2150-1850
cm
1 spectral region. This EPR-silent state of
the active RH resembles the Nia-S state of standard
[NiFe]-hydrogenases. Maximal reduction, already obtained after a few
minutes under 100% H2 at room temperature, yielded the
Nia-C* state (Fig. 4C) as identified previously
in other [NiFe]-hydrogenases (30, 31). This state showed a CO stretch
vibration at 1960 cm
1. The two bands at 2082 and 2071 cm
1, which did not shift, are
ascribed to the symmetrical and antisymmetrical coupled vibrations of
two cyanides bound to iron in the active site (17). It was not possible
to reduce further this state by adding excess dithionite (20 mM, spectrum not shown).
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Fig. 4.
FTIR spectra of the RH. Trace
A shows the RH in the oxidized state. After reduction under
H2 the RH ends up in the reduced state (trace
C). If the gas phase was then exchanged for CO, a mixture of
oxidized and reduced RH was observed (trace B). A similar
spectrum was obtained by flushing with argon.
1 could be seen. Such a band from added CO
is observed in the A. vinosum and D. gigas
enzyme3 (32). A similar
change was observed by replacing H2 with argon (results not
shown). Upon complete oxidation with excess DCIP (2 mM) the
sample returned to the Nia-S state.
251 was 11.96 mM
1 cm
1
based on protein concentration. Similarly the
342 was
calculated to be 5.36 mM
1
cm
1. The protein concentration used (0.64 mg/ml) was such that the absorption at 280 nm was about 1.0. At this
intensity the detector is still sensitive enough to pick up reliable
differences in the UV, meaning that these are not due to mismatching in
this region.
View larger version (14K):
[in a new window]
Fig. 5.
Difference spectrum of reduced minus oxidized
RH. Aerobic RH was diluted to a concentration of 0.64 mg/ml and
divided over two cuvettes. One cuvette was put under 100%
H2 and measured against the aerobic sample in the reference
cuvette at 2 nm resolution.
View larger version (46K):
[in a new window]
Fig. 6.
Complex formation of the RH and the histidine
protein kinase HoxJ. Purified RH, HoxJ, and a mixture of RH and
HoxJ, preincubated for 10 min, were applied to native PAGE.
A, Coomassie staining; B,
H2-dependent PMS reduction of native gels.
Lane 1, 50 pmol of RH; lane 2, 50 pmol of RH and
10 pmol of HoxJ; lane 3, 50 pmol of RH and 50 pmol of HoxJ;
lane 4, 50 pmol of RH and 250 pmol of HoxJ; lane
5, 50 pmol of RH and 250 pmol of bovine serum albumin; lane
6, 50 pmol of RH and 50 pmol of HoxJ preincubated for 30 min under
H2; lane 7, 50 pmol of HoxJ.
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
2
2
tetramer, the possibility exists that two unpaired spins released by
the two [Ni-Fe] sites in the tetramer are united in a yet undetected, diamagnetic prosthetic group. Hence UV-visible spectroscopy was applied. Much to our surprise reduction of the RH by H2
resulted in an increase in absorption with clear maxima at 251 and 342 nm. We tentatively conclude that this increase is caused by the reduction of a two electron accepting cofactor, shared by the two dimer
(
) molecules in the RH (
2
2). The
exact identity of this cofactor is currently under investigation. The
position of the 342 nm band and its approximate molecular absorption
coefficient (5.4 mM
1
cm
1) resemble those of NADH.
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ACKNOWLEDGEMENT |
---|
We thank A. Strack for excellent technical assistance.
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FOOTNOTES |
---|
* This work was supported by the The Netherlands Organization for Scientific Research, the Deutsche Forschungsgemeinschaft, 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.
This paper is dedicated to Prof. Ernst-G. Jäger on the occasion of his 65th birthday on May 5, 2001.
To whom correspondence should be addressed. Tel.: 49 30 20 93 81 01; Fax: 49 30 20 93 81 02; E-mail:
baerbel.friedrich@rz.hu-berlin.de.
Published, JBC Papers in Press, February 16, 2001, DOI 10.1074/jbc.M009802200
2 M. Forgber, O. Lenz, E. Schwartz, and B. Friedrich, unpublished results.
3 De Lacey, A.L., and Fernandez, V.M., unpublished results.
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ABBREVIATIONS |
---|
The abbreviations used are: MBH, membrane-bound hydrogenase; DCIP, dichlorophenolindophenol; FTIR, Fourier transform infrared; RH, regulatory hydrogenase; SH, soluble hydrogenase; PAGE, polyacrylamide gel electrophoresis; Mes, 4-morpholineethanesulfonic acid; Mops, 4-morpholinepropanesulfonic acid.
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REFERENCES |
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1. | Ignarro, L. J., Degnan, J. N., Baricos, W. H., Kadowitz, P. J., and Wolin, M. S. (1982) Biochim. Biophys. Acta 718, 49-59[Medline] [Order article via Infotrieve] |
2. |
Shelver, D.,
Kerby, R. L.,
He, Y.,
and Roberts, G. P.
(1997)
Proc. Natl. Acad. Sci. U. S. A.
94,
11216-11220 |
3. | Gilles-Gonzalez, M. A., Ditta, G. S., and Helinski, D. R. (1991) Nature 350, 170-172[CrossRef][Medline] [Order article via Infotrieve] |
4. | Gilles-Gonzalez, M. A., Gonzalez, G., and Perutz, M. F. (1995) Biochemistry 34, 232-6[Medline] [Order article via Infotrieve] |
5. |
Gong, W.,
Hao, B.,
Mansy, S. S.,
Gonzalez, G.,
Gilles-Gonzalez, M. A.,
and Chan, M. K.
(1998)
Proc. Natl. Acad. Sci. U. S. A.
95,
15177-15182 |
6. | Schink, B., and Schlegel, H. G. (1979) Biochim. Biophys. Acta 567, 315-324[Medline] [Order article via Infotrieve] |
7. | Schneider, K., and Schlegel, H. G. (1976) Biochim. Biophys. Acta 452, 66-80[Medline] [Order article via Infotrieve] |
8. | Schwartz, E., Gerischer, U., and Friedrich, B. (1998) J. Bacteriol. 180, 3197-3204[Abstract] |
9. |
Lenz, O.,
and Friedrich, B.
(1998)
Proc. Natl. Acad. Sci. U. S. A.
95,
12474-12479 |
10. | Lenz, O., Strack, A., Tran-Betcke, A., and Friedrich, B. (1997) J. Bacteriol. 179, 1655-1663[Abstract] |
11. | Stock, J. B., Surette, M. G., Levit, M., and Park, P. (1995) in Two-component Signal Transduction (Hoch, J. A. , and Silhavy, T. J., eds) , pp. 25-51, American Society for Microbiology, Washington, D. C. |
12. | Eberz, G., and Friedrich, B. (1991) J. Bacteriol. 173, 1845-54[Medline] [Order article via Infotrieve] |
13. | Zimmer, D., Schwartz, E., Tran-Betcke, A., Gewinner, P., and Friedrich, B. (1995) J. Bacteriol. 177, 2373-80[Abstract] |
14. | Elsen, S., Colbeau, A., Chabert, J., and Vignais, P. M. (1996) J. Bacteriol. 178, 5174-5181[Abstract] |
15. | Black, L. K., Fu, C., and Maier, R. J. (1994) J. Bacteriol. 176, 7102-7106[Abstract] |
16. |
Kleihues, L.,
Lenz, O.,
Bernhard, M.,
Buhrke, T.,
and Friedrich, B.
(2000)
J. Bacteriol.
182,
2716-2724 |
17. | Pierik, A., Schmelz, M., Lenz, O., Friedrich, B., and Albracht, S. P. (1998) FEBS Lett. 438, 231-5[CrossRef][Medline] [Order article via Infotrieve] |
18. |
Lowry, O. H.,
Rosebrough, N. J.,
Farr, A. L.,
and Randall, R. J.
(1951)
J. Biol. Chem.
193,
265-275 |
19. | Bernhard, M., Schwartz, E., Rietdorf, J., and Friedrich, B. (1996) J. Bacteriol. 178, 4522-4529[Abstract] |
20. | Coremans, J. M. C. C., Van der Zwaan, J. W., and Albracht, S. P. J. (1989) Biochim. Biophys. Acta 997, 256-267 |
21. | Cornish-Bowden, A. (1995) Analysis of Enzyme Kinetic Data , pp. 133-189, Oxford University Press, New York |
22. | Covington, A. K., Paabo, M., Robinson, R. A., and Bates, R. G. (1968) Anal. Chem. 40, 700-706 |
23. | Quinn, D. M., and Sutton, L. D. (1991) in Enzyme Mechanism from Isotope Effects (Cook, P. F., ed) , pp. 73-126, CRC, Boca Raton, FL |
24. | Beinert, H., and Albracht, S. P. (1982) Biochim. Biophys. Acta 683, 245-277[Medline] [Order article via Infotrieve] |
25. | Hyman, M. R., and Arp, D. J. (1987) Biochemistry 26, 6447-54 |
26. | Zorin, N. A., Dimon, B., Gagnon, J., Gaillard, J., Carrier, P., and Vignais, P. M. (1996) Eur. J. Biochem. 241, 675-681[Abstract] |
27. |
Happe, R. P.,
Roseboom, W.,
and Albracht, S. P.
(1999)
Eur. J. Biochem.
259,
602-608 |
28. | Van der Zwaan, J. W., Albracht, S. P. J., Fontijn, R. D., and Mul, P. (1987) Eur. J. Biochem. 169, 377-384[Abstract] |
29. | Teixeira, M., Fauque, G., Moura, I., Lespinat, P. A., Berlier, Y., Prickril, B., Peck, H. D., Jr., Xavier, A. V., LeGall, J., and Moura, J. J. G. (1987) Eur. J. Biochem. 167, 47-58[Abstract] |
30. | Bagley, K. A., Duin, E. C., Roseboom, W., Albracht, S. P., and Woodruff, W. H. (1995) Biochemistry 34, 5527-5535[Medline] [Order article via Infotrieve] |
31. | De Lacey, A. L., Hatchikian, E. C., Volbeda, A., Frey, M., Fontecilla-Camps, J. C., and Fernandez, V. M. (1997) J. Am. Chem. Soc. 119, 7181-7189[CrossRef] |
32. | Bagley, K. A., Van Garderen, C. J., Chen, M., Duin, E. C., Albracht, S. P., and Woodruff, W. H. (1994) Biochemistry 33, 9229-9236[Medline] [Order article via Infotrieve] |
33. | Schneider, K., Cammack, R., Schlegel, H. G., and Hall, D. O. (1979) Biochim. Biophys. Acta 578, 445-461[Medline] [Order article via Infotrieve] |
34. | Happe, R. P., Roseboom, W., Egert, G., Friedrich, C. G., Massanz, C., Friedrich, B., and Albracht, S. P. (2000) FEBS Lett. 466, 259-263[CrossRef][Medline] [Order article via Infotrieve] |
35. | Krasna, A. I. (1979) Enzyme Microb. Technol. 1, 165-172 |
36. | Volbeda, A, Charon, M. H., Piras, C., Hatchikian, E. C., Frey, M., and Fontecilla-Camps, J. C. (1995) Nature 373, 580-587[CrossRef][Medline] [Order article via Infotrieve] |
37. | Montet, Y., Amara, P., Volbeda, A., Vernede, X., Hatchikian, E. C., Field, M. J., Frey, M., and Fontecilla-Camps, J. C. (1997) Nat. Struct. Biol. 4, 523-527[Medline] [Order article via Infotrieve] |
38. | Ponting, C. P., and Aravind, L. (1997) Curr. Biol. 7, 674-7 |
39. | Zhulin, I. B., Taylor, B. L., and Dixon, R. (1997) Trends Biochem. Sci. 22, 331-333[CrossRef][Medline] [Order article via Infotrieve] |
40. |
Taylor, B. L.,
and Zhulin, I. B.
(1999)
Microbiol. Mol. Biol. Rev.
63,
479-506 |
41. |
Bibikov, S. I.,
Barnes, L. A.,
Gitin, Y.,
and Parkinson, J. S.
(2000)
Proc. Natl. Acad. Sci. U. S. A.
97,
5830-5835 |