From the Institute of Biophysics, Biological Research Center of the Hungarian Academy of Sciences, P. O. Box Szeged H-6701, Hungary
Received for publication, January 21, 2003
, and in revised form, March 25, 2003.
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ABSTRACT |
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INTRODUCTION |
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It is known that hydrogenase needs to be activated under anaerobic conditions (4, 5, 6). During activation and the enzyme cycle, several stable intermediates have been determined by electron paramagnetic resonance (EPR)1 and Fourier transform infrared (FTIR) spectroscopy (7, 8), ranging from the fully oxidized Form A through oxygen-free Form Siab, and from Form B to the active enzyme (Form C and Form R). These forms have not yet been satisfactorily assigned to the corresponding theoretical enzyme forms of hydrogen splitting.
Although the enzymatic activity of hydrogenase is determined routinely, a number of contradictory results have been published. Despite the many features described in the hydrogenase reaction, the activity of this class of enzymes has not yet been thoroughly explained (5, 9, 10, 11, 12). An autocatalytic kinetic step was recently proposed in hydrogenase activation or during the hydrogenase kinetic cycle.2,3
The ability of nitrogen laser flash-reduced methyl or benzyl viologen to initiate a redox transition of a redox protein has been demonstrated previously (14). Here we report fast kinetic measurements of the hydrogenase enzyme reaction involving the use of an excimer laser flash-reduced redox dye. The results are evaluated on the assumption of an autocatalytic reaction in the hydrogenase kinetic cycle.
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EXPERIMENTAL PROCEDURES |
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Protein PurificationThe wet cell paste of T.
roseopersicina was treated with 90% cold (20 °C) acetone, and
the pellet was dried and stored at4 °C. For purification, 15 g of
pellet was dissolved in distilled water at 50 °C. The solution was
centrifuged at 12,500 x g for 1 h, and the supernatant was
used. The first purification step comprised batch chromatography with Whatman
DEAE DE-52 in 20 mM Tris-HCl, pH 7.5. The hydrogenase was washed
off with 450 mM NaCl. The next step was butyl-Sepharose column
chromatography in 1 mM Tris-HCl at pH 7.5 with 10% ammonium
sulfate, using the Amersham Biosciences FPLC system. Hydrogenase eluted at
0% ammonium sulfate. Two Q-Sepharose Fast Flow columns were used for
further purification steps using different buffer systems (20 mM
Tris-HCl, pH 7.5, and 50 mM MES, pH 5.5). In both cases, the
hydrogenase eluted in the interval 350500 mM NaCl. Final
purification of the hydrogenase was achieved by 9% native preparative
polyacrylamide gel electrophoresis. The hydrogenase band was excised, and the
electrophoretically pure enzyme was collected on a small Q-Sepharose
column.
Sample PreparationControl samples containing 10 mM oxidized methyl viologen in 20 mM Tris-HCl, pH 7.5, were incubated overnight under anaerobic conditions. Before measurement, the sample was poured into a 0.1-mm quartz cell under anaerobic conditions.
Samples containing 5 mg/ml hydrogenase (50 µM) in 20
mM Tris-HCl (pH 7.5) buffer were incubated overnight in an anaerobe
box (Coy Laboratories) containing 10% of hydrogen and 90% of nitrogen.
After incubation, the hydrogenase was in activated form (Form C) as revealed
by the EPR spectrum. The sample was poured into a 0.1-mm quartz cell and
oxidized to Form B by adding a negligible volume of concentrated oxidized
methyl viologen to a final concentration of 10 mM under anaerobic
conditions. The cell was sealed with a quartz cover plate, which maintained
anaerobic conditions for several hours.
Kinetic MeasurementsThe nanosecond flash photolysis setup involved an excimer laser, a controlled-temperature sample holder, a Xenon lamp, a detector, and a DSA sampling oscilloscope connected to an IBM personal computer.
The data collection, evaluation, and model fitting were performed with a specifically modified SPSERV program.4 The program was modified so it would be able to supervise the DSA sampling oscilloscope and collect and process the data.
The geometry of excitation was pseudo front face, and the analyzed volume was completely irradiated by the excitation beam from the excimer laser at 307 nm. The frequency of laser flashes was 10 Hz. The samples were slowly rotated so that each laser flash came into contact with a fresh portion of the sample. After each measurement, a new cylinder of the cell was selected to avoid saturation of the sample (for an explanation of the saturation, see "Results and Discussion"). In every experiment, the temperature of the cell was maintained at 25 °C.
The wavelength of the measuring light beam for the methyl viologen (control) sample was varied in the range 360680 nm in 20-nm steps. The bandwidth was 10 nm. 100 flashes were averaged and collected for each wavelength.
The wavelength of the measuring light beam for hydrogenase samples was varied in the range 350600 nm, and kinetic measurements were made at indicated wavelengths. The bandwidth was 30 nm. Data collected from 1000 individual flashes were averaged at each wavelength.
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RESULTS AND DISCUSSION |
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In the control samples (containing no hydrogenase) the concentration of reduced methyl viologen produced after one excimer laser flash was 1.5 µM. The activation spectrum reconstructed from the signal amplitude of the samples containing only methyl viologen is in excellent agreement with the absorption spectrum of reduced methyl viologen (Fig. 1).
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The correspondence of the activation spectrum reconstructed from the signal amplitude demonstrates that methyl viologen reduction did occur. This confirms our previous findings with the nitrogen laser (14).
In the case of hydrogenase, it is more difficult to demonstrate a redox transition as compared with flavocytochrome c552. Hydrogenase is not only a redox protein but also a catalyst that produces hydrogen in this reaction. Because the volume of the sample is limited and hydrogen cannot leave it, after a while the back reaction of the hydrogenase should be taken into account. Thus, upon prolonged irradiation the forward and back reactions of the hydrogenase will come into equilibrium, and, therefore, no further net methyl viologen oxidation will be observed. Rotation of the sample holder and irradiation of a fresh portion of the sample can partially resolve this problem; but, after the entire surface of the cell has been irradiated, the whole sample becomes saturated with hydrogen gas, and no further disappearance of the reduced dye can be observed despite the presence of hydrogenase in the sample. Hence, one sample is suitable for only a limited number of experiments.
The results of fast kinetic measurements on the hydrogenase-methyl viologen reaction at different wavelengths are presented in Fig. 2. The rate of change of the absorption of hydrogenase samples on reaction with methyl viologen is very characteristic. At every wavelength where reduced methyl viologen absorbs, a fast initial increase can be observed, followed by damped oscillatory behavior of the kinetic trace. The spectral dependence of the initial amplitude follows the methyl viologen absorption spectrum (there is no observable initial increase in Fig. 2 at 450- and 500-nm excitation, where the reduced methyl viologen absorption spectrum exhibits a minimum). This is exactly the same phenomenon as observed in hydrogenase-free samples.
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The measured rate of change of OD for a hydrogenase/methyl viologen
sample can be fitted with the sum of a fast and a slow decay curve
(A1·exp(t/
1) and
A2·exp(t/
2)) and two damped
sinusoidal time functions
(A3·sin(
(tt01))
·exp(/
3) and
A4·sin(
(tt02))
·exp(t/
4))
(Fig. 3).
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The amplitude of the fast decay curve (A1) reflects the methyl
viologen spectrum, and the value for this exponential
(
1) remains constant throughout the wavelength range measured.
The reduced methyl viologen concentration decreases as the hydrogenase takes
over the electrons from the reduced methyl viologen. We therefore assign this
process to the disappearance of reduced methyl viologen by the transfer of
electrons to hydrogenase. The rate constant for the electron transfer from the
reduced methyl viologen to the hydrogenase (calculated according to Ref.
14) is k =
1.33(5)·108 M1
s1. This value is of the same order of magnitude
as the 109 M1 s1
found for Chromatium vinosum hydrogenase by cyclic voltammetry
(17).
We have not been able to assign the second decay to any process. This decay
can be visualized at some wavelengths as a slight decrease in the kinetic
trace with time. This is clearly pronounced at 350 nm in
Fig. 2, where the damped
oscillations are not around the zero value. The dependence of the amplitude
(A2) on the wavelength demonstrates no clear-cut tendency. The
value for this exponential (
2) changes from 15,000 µs
at 350 nm to 400 µs at 600 nm, but the fitting error for the high values
was high (>50%). This change in A2 and
2
indicates that the concentration of a corresponding reaction partner is
changing during the measurement. Because we collected 1000 flashes with a
frequency of 10 Hz in order to obtain a reasonable signal-to-noise ratio, a
slow process can develop during the data collection (100 s), and a hydrogenase
component, the substrate (the reduced viologen dye) or the product (the
hydrogen gas) can diffuse over the cell and interfere with the following
measurement. We therefore assign this process as a methodological
artifact.
The kinetic characteristics of the hydrogenase reaction, obtained by conventional activity measurements, led us to the conclusion that there should be an autocatalytic reaction step in the hydrogenase cycle or during the activation process.2,3 The autocatalytic interaction between two enzyme forms results in a conformational change of the enzyme, thereby activating it. We know from experimental results that such conformational changes are possible and that different conformations of hydrogenase exist. Following the changes in behavior of the hydrogenase under SDS-PAGE conditions, we were able to distinguish between three different states of the enzyme. The first conformer is present under aerobic conditions. Incubation of the enzyme under anaerobic conditions changes it to a different conformer, whereas incubation under hydrogen and methyl viologen transforms it into a third conformational state (18, 19, 20). We do not know, however, if these conformational changes are associated with an autocatalytic reaction. It is very interesting to note, however, that, similarly to prion proteins, the hydrogenases are partially resistant to proteolytic digestion (20).
The autocatalytic behavior of an enzyme reaction leads to an oscillating
concentration of enzyme products contributing to the autocatalytic step. On
the assumption of a simple autocatalytic reaction for hydrogenase activation,
as shown in Reaction 1 (the
autocatalytic step is between E2 and E3, as indicated by
the curved back arrow), the changes in concentration of E2 and
E3 would be reflected by a damped oscillation with
2 =
k1·k0·E1, if
k1·k0·E1
« k22 and if we can neglect second-order
terms (21,
22). E4 is the
active form of the enzyme that is able to catalyze hydrogen evolution or
hydrogen splitting. The reaction cycle closes back to E2 (recent
model) or E3, the autocatalytic step being included in the enzyme
cycle or left only in the activation process.
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k0 describes an intraenzyme reaction (probably a conformation change or charge redistribution), while k1 is the autocatalytic rate constant. It should be noted that, at the start of the experiment, the hydrogenase is in the E1 form, and this is not the oxidized form (Form A) of the enzyme (see sample preparation). The two oscillating concentrations observed in our experiment would correspond to E2 and E3 in the above model. The spectral dependence of the initial amplitudes of both of the damped sine curves can be described as the difference in absorption of the spectra of reduced and oxidized hydrogenase, which has a maximum at around 440 nm, though the fitting errors are rather large (Fig. 4). The spectral dependence of the amplitude of the second oscillation is the mirror image of the first. This implies that the oscillation occurs between an oxidized and a reduced form of hydrogenase.
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An alternative explanation is that, while the first oscillation proceeds from a reduced hydrogenase, the second oscillation originates from the reduced dye. As a consequence, it is possible that an enzyme form (if any) binding the reduced dye is responsible for this spectral component, or, alternatively, the methyl viologen plays the role of the autocatalytic partner2. To be able to select between these two possibilities, the rate of changes of the spectra of the hydrogenase-methyl viologen samples should be measured in smaller wavelength steps. Unfortunately, the saturation of the hydrogenase samples by hydrogen and the limited volume did not allow us to obtain more precise wavelength information by using the same hydrogenase sample.
The frequencies (0.9 (1) ·103/s and 1.04 (9)
·103/s) of the two damped oscillations are very close
(within the fitting error); only the phases differ. This is in accordance with
the above autocatalytic model, where the two oscillations should have the same
frequency. The
k1·k0·E1
value calculated from the average of this fit is roughly
6·106 s2. For the conditions
discussed above,
k0·k1·E1
« k22, which puts k2
at > 3·103 s1. In the
knowledge of the overall concentration of the enzyme and on the assumption
that the enzyme is in the E1 form at the beginning of the
experiment, k0·k1
1011 M1 s2.
The rate of the autocatalytic process should be higher than or at least the same order of magnitude as that of the reaction with the real substrate (reduced methyl viologen in our case); otherwise, we would not be able to observe the autocatalytic behavior of the reaction because it would be very fast. A comparison of the reaction rate constant of the reduced methyl viologen binding, calculated from the fast decay of the reduced dye, puts k1 at around or higher than 108 M1 s1. As a result, k0 < 103 s1.
In summary, the ability to use flash-reduced methyl viologen as a light-induced trigger in transient kinetic phenomena associated with the intermolecular electron transfer of hydrogenase has been demonstrated. The possibility of fitting the kinetics with a hydrogenase autocatalytic process has been demonstrated, and the kinetic constants of the autocatalytic reaction and the binding of methyl viologen or its release from hydrogenase have been determined. Limits of the kinetic constants relating to the intramolecular (intraenzyme) reactions have also been set.
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FOOTNOTES |
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To whom correspondence should be addressed: Temesvári krt. 62, P. O.
Box 521, Szeged H-6701, Hungary. Tel.: 36-62-599-605; Fax: 36-62-433-133;
E-mail:
csaba{at}nucleus.szbk.u-szeged.hu.
1 The abbreviations used are: EPR electron paramagnetic resonance; MES,
4-morpholineethanesulfonic acid.
2 J. Ösz and Cs. Bagyinka, submitted for publication.
3 Bagyinka, Cs., Ösz, J., and Kovács, K. L. (2000) Poster
presented at the 6th International Conference on the Molecular Biology of
Hydrogenases, Potsdam, Germany (August 510, 2000).
4 Copyrighted in 2000 by Csaba Bagyinka.
5 Cs. Bagyinka and M. J. Maroney, unpublished data.
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REFERENCES |
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