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J Biol Chem, Vol. 274, Issue 42, 29772-29778, October 15, 1999

Rubredoxin from the Green Sulfur Bacterium Chlorobium tepidum Functions as an Electron Acceptor for Pyruvate Ferredoxin Oxidoreductase^*

Ki-Seok Yoon^§, Russ Hille^¶, Craig Hemann^¶, and F. Robert Tabita

From the  Department of Microbiology and the Plant Biotechnology Center and the ^¶ Department of Medical Biochemistry, The Ohio State University, Columbus, Ohio 43210-1292

	ABSTRACT

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Rubredoxin (Rd) from the moderately thermophilic green sulfur bacterium Chlorobium tepidum was found to function as an electron acceptor for pyruvate ferredoxin oxidoreductase (PFOR). This enzyme, which catalyzes the conversion of pyruvate to acetyl-CoA and CO₂, exhibited an absolute dependence upon the presence of Rd. However, Rd was incapable of participating in the pyruvate synthase or CO₂ fixation reaction of C. tepidum PFOR, for which two different reduced ferredoxins are employed as electron donors. These results suggest a specific functional role for Rd in pyruvate oxidation and provide the initial indication that the two important physiological reactions catalyzed by PFOR/pyruvate synthase are dependent on different electron carriers in the cell. The UV-visible spectrum of oxidized Rd, with a monomer molecular weight of 6500, gave a molar absorption coefficient at 492 nm of 6.89 mM¹ cm¹ with an A₄₉₂/A₂₈₀ ratio of 0.343 and contained one iron atom/molecule. Further spectroscopic studies indicated that the CD spectrum of oxidized C. tepidum Rd exhibited a unique absorption maximum at 385 nm and a shoulder at 420 nm. The EPR spectrum of oxidized Rd also exhibited unusual anisotropic resonances at g = 9.675 and g = 4.322, which is composed of a narrow central feature with broader shoulders to high and low field. The midpoint reduction potential of C. tepidum Rd was determined to be 87 mV, which is the most electronegative value reported for Rd from any source.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Chlorobium tepidum is a moderately thermophilic, anoxygenic green sulfur photosynthetic bacterium (1) capable of obtaining cell carbon through the action of two reduced ferredoxin (Fd)¹-linked CO₂ fixation reactions, much like other specialized prokaryotes. The enzymes that catalyze these reactions, pyruvate synthase (PS) and -ketoglutarate synthase, are key components of the reductive tricarboxylic acid pathway of CO₂ fixation (2-7). PS (Equation 1) is classically thought to also catalyze a pyruvate ferredoxin/flavodoxin oxidoreductase (PFOR) reaction, in which pyruvate is oxidized to acetyl-CoA and CO₂, essentially the reverse of the PS reaction (Equation 2).

(Eq. 1)

(Eq. 2)

In the PFOR reaction, coenzyme A and thiamine diphosphate (ThDP) are necessary coenzymes, with oxidized Fd or flavodoxin previously shown to be necessary electron acceptors (Equation 2) (8-13). PFOR is thus distinct from the well characterized pyruvate dehydrogenase multienzyme complex, which catalyzes coenzyme A- and NAD-dependent oxidation of pyruvate (Equation 3) (14, 15). PFOR also differs from pyruvate decarboxylase, which catalyzes the ThDP-dependent conversion of pyruvate to acetaldehyde and CO₂ (Equation 4) (16, 17).

(Eq. 3)

(Eq. 4)

To date, the physiologically important CO₂ fixation (or PS) reaction has only been demonstrated for one purified PFOR/PS enzyme, the protein from Hydrogenbacter thermophilus (7, 12);however, the reaction was initially discovered in another green sulfur bacterium, C. limicola forma thiosulfatophilum (3, 18). The PS carboxylation reaction must overcome a large negative potential energy barrier, which is facilitated by the use of Fd, one of the most electronegative electron carriers known in cellular metabolism (19, 20). Thus, PS catalyzes a carboxylase reaction, which distinguishes it from pyruvate dehydrogenase multienzyme complex, pyruvate decarboxylase, and perhaps even some sources of PFOR.

During the course of studying the carboxylation enzymes of the reductive tricarboxylic acid cycle of C. tepidum, a Rd was isolated and purified to homogeneity from photoautotrophically grown cells. Rd is a small nonheme iron protein and is an important electron transfer component of many bacteria, especially anaerobic organisms. Rd contains a single iron atom coordinated to four cysteine residues in a tetrahedral arrangement (21) with the redox-active site located at the surface of the protein, indicating a directed outer sphere electron transfer mechanism. There are many specific examples where Rd has been shown to participate in a variety of metabolic reactions. For example, Rd is proposed to be the primary electron carrier for the acetogenic CO dehydrogenase of several anaerobes (22) and is an intermediary electron carrier in the NAD(P)H-dependent nitrate reduction system in Clostridium perfringens (23). Similarly, in Pseudomonas oleovorans, a soluble Rd reductase and a membrane-bound -hydroxylase are coupled to the hydroxylation of aliphatic hydrocarbons, functionalized hydrocarbons, and various aromatic compounds (24, 25). In Azotobacter vinelandii (26) and in Rhizobium leguminosarum (27), the genes coding for proteins with sequences very similar to Rd have recently been recognized as part of hydrogenase gene clusters, which may indicate the involvement of Rd in hydrogen oxidation or reduction (28). The Rd of Desulfovibrio gigas has also shown to function as a redox coupling protein between NADH oxidoreductase and Rd oxidoreductase, allowing this organism to increase ATP production via the degradation of internal polyglucose in the presence of oxygen (21, 29). Despite these disparate studies, the actual physiological function of Rd is still unclear, especially in organisms that may obtain much of their organic carbon from CO₂ assimilation, such as green photosynthetic bacteria (30-33). Certainly, from the foregoing, Rd might have very important catabolic and also anabolic functions in different organisms. In this study, it is demonstrated that Rd from C. tepidum exhibits unique characteristics; most interestingly, it serves as a high potential electron acceptor for C. tepidum PFOR, resulting in the conversion of pyruvate to acetyl-CoA + CO_2. However, Rd has thus far not been shown to participate in the PS reaction of this protein, suggesting a potential specific physiological role in pyruvate oxidation.

	EXPERIMENTAL PROCEDURES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Bacterium and Growth Conditions-- C. tepidum strain TLS was grown photoautotrophically in 20-liter Carboy bottles according to a modification of a method described previously (34). The cells were harvested anaerobically, using continuous centrifugation at 35,000 × g at 4 °C under a flow of nitrogen gas. The harvested cells were washed twice with anaerobic 50 mM phosphate buffer, pH 7.0, containing 50 mM -mercaptoethanol. The cells were then resuspended in the same buffer (1 g of wet weight cells/4 ml of buffer), after which the cell suspension was disrupted by two passes through a French pressure cell at 10,000 psi under a flow of argon gas. Cell debris and unbroken cells were removed by centrifugation (20,000 × g, 10 min); the supernatant fraction was then centrifuged at 150,000 × g for 1.5 h. The supernatant fraction from this step was used for the purification of all soluble proteins, including Rd, two Fds, and PFOR. The high speed precipitate fraction was used for the isolation of membrane-bound reaction center particles.

Purification of Rd-- All purification procedures were carried out at room temperature under anaerobic conditions using a Coy anaerobic chamber under an atmosphere of 95% nitrogen and 5% hydrogen. All buffers were repeatedly degassed and flushed with argon and were maintained under a positive pressure of argon; for overnight storage, the buffers were stirred continuously with a magnetic stir bar in the anaerobic chamber. With the exception of the last two chromatographic steps, buffers contained sodium dithionite (1 mM) and dithiothreitol (1 mM) to protect against trace oxygen contamination. Protein purification was performed in the Coy anaerobic chamber using the Biological Autoprogram System from Bio-Rad. The supernatant following ultracentrifugation was loaded onto a DEAE-Sepharose fast flow column (2.5 × 15 cm), pre-equilibrated with 20 mM Tris-HCl, pH 8.0, 1 mM dithiothreitol, 1 mM sodium dithionite, 2 mM MgCl₂, and 0.1 mM ThDP. The column was washed with 100 ml of the same buffer containing 0.1 M NaCl at a flow rate of 7 ml/min. A stepwise gradient method was used that consisted of 600 ml of buffer containing a gradient of 0.1-0.3 M NaCl, 200 ml of 0.3-0.5 M NaCl, and 100 ml of 0.5-2.0 M NaCl. PFOR, Rd, and Fds were eluted around 0.2, 0.3, and 0.4 M NaCl, respectively, from this first column. The Rd fraction, eluting at approximately 0.32 to 0.34 M NaCl, was diluted 2-fold with nonsalt buffer and then applied onto a Q-Sepharose high performance column (1.6 × 10 cm) pre-equilibrated with wash buffer. After sample application, the column was washed with wash buffer and then eluted isocratically using buffer containing 0.25 M NaCl, followed by a gradient of buffer containing 0.25-0.4 M NaCl at a flow rate of 3 ml/min. The total gradient volume was 180 ml. The Rd-containing fractions from this column were then directly loaded onto a hydroxyapatite column (1.0 × 15 cm) pre-equilibrated with 1 mM potassium phosphate, pH 6.8, containing 1 mM dithiothreitol. At this stage, the Rd-containing fraction was concentrated using Utrafree-15, PBCC 5,000 NMWL (Millipore). Then the Rd was loaded onto a Superose-12 column (1.6 × 50 cm) pre-equilibrated with 20 mM Tris-HCl, pH 8.0, containing 0.2 M NaCl and eluted at a flow rate of 1.0 ml/min. PFOR and two different Fds were also purified to homogeneity via several column chromatographic steps under anaerobic conditions.²

EPR, CD, and UV-visible Spectra of Rd-- EPR spectra were obtained with a Brüker instruments, Inc. ER 300 spectrometer equipped with ER 035M NMR gaussmeter and a Hewlett-Packard 5352B microwave frequency counter. Double integration of the EPR spectra was performed with the ESP 300 Software package from Brüker Instruments. Air-oxidized Rd was injected into quartz EPR tubes and frozen by plunging the tubes into liquid nitrogen. Samples of reduced Rd were prepared under anaerobic conditions with either an enzymatic protocol using the activity of PFOR or by chemical means after the addition of sodium dithionite. The reduced liquid samples were collected directly into EPR tubes and sealed with a rubber stopper; the samples were then frozen at specific times after mixing in a dry ice/acetone bath by plunging the tube into liquid nitrogen. EPR spectra were measured at 4.7-20 K using 10 mW of microwave power under liquid helium. CD spectra of oxidized or reduced Rd were measured at room temperature on an AVIV 62DS Spectropolarimeter. CD data were collected at 1-mm intervals using averaging times of 1 s/m, depending on the signal to noise ratio. UV-visible spectra were measured with a Beckman DU®-70 spectrophotometer.

General Properties of C. tepidum Rd-- The midpoint reduction potential value of the C. tepidum Rd was determined by cyclic voltammetry at a glassy carbon electrode at 25 °C (35). The scan rate was 20 mV/s over the potential range 0 to 600 mV (versus the Ag/AgCl electrode). All values were referenced to the standard hydrogen electrode. The N-terminal amino acid sequence and amino acid analyses of Rd were determined using standard procedures. The purity of Rd was established by SDS-PAGE using a discontinuous Tris-Tricine buffer system (36); samples were heated at 75 °C for 1 h. The apparent molecular weight of Rd was estimated by calibration with a commercial kit of small molecular weight markers from Sigma. Protein concentrations were routinely estimated by established procedures (37). The iron content of the protein was determined by the o-phenanthroline procedure (38). The molecular weight of Rd was determined by gel filtration using a Superose-12 HR (10/30) column calibrated with -amylase (M_r = 200,000), bovine serum albumin (M_r = 67,000), carbonic anhydrase (M_r = 29,000), cytochrome c (M_r = 12,400), and aprotinin (M_r = 6,500). In all cases, 50 mM Tris-HCl, pH 8.0, containing either 0.1 or 1.0 M NaCl, was used as the eluate buffer.

Enzyme Assays-- PFOR activity was routinely determined spectrophotometrically by following the pyruvate-dependent reduction of methyl viologen in anaerobic cuvettes under nitrogen gas at 30 °C (12). The standard assay mixture contained 5 mM pyruvate, 0.5 mM coenzyme A, 2 mM MgCl₂, 0.1 mM ThDP, and 2 mM methyl viologen in 50 mM phosphate buffer, pH 7.8. PFOR activity was expressed as units/mg of protein, where 1 unit is equivalent to the reduction of 1 µmol of methyl viologen per min at 578 nm (_{578 nm} = 9.8 mM¹ cm¹). Acetyl-CoA, formed as a result of pyruvate oxidation catalyzed by Rd-dependent PFOR activity, was measured using a coupled assay with malate dehydrogenase and citrate synthase (39). The amount of pyruvate consumed after the PFOR reaction was also determined with a lactate dehydrogenase coupled assay (40). The amounts of pyruvate and acetyl-CoA oxidized and produced, respectively, were standardized using commercial preparations of pyruvate and acetyl-CoA, as above. PS activity was determined using a radiometric assay (2, 3). Reduced Fd and Rd were employed in reaction mixtures to reductively carboxylate acetyl-CoA; isolated spinach chloroplast fragments (41) or C. tepidum membrane-bound reaction center particles were used to photoreduce these electron carriers (42). Citrate synthase and malate dehydrogenase were obtained from Roche Molecular Biochemicals, and lactate dehydrogenase, acetyl-CoA, NAD, NADH, coenzyme A, and pyruvate were purchased from Sigma.

	RESULTS

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

General Properties of C. tepidum Rd-- After purification, the Rd preparation was oxidized and pink-red in color; it eluted from DEAE-Sepharose columns at salt concentrations between 0.32 and 0.34 M NaCl or about 20-25 mS/cm conductivity. This column effectively separated other important soluble proteins, including PFOR, two Fds, and others. Furthermore, the hydroxyapatite column was crucial to separate rubredoxin from another rubredoxin-like molecule that was found to compromise the spectral and other molecular properties obtained. Rd was reduced and blanched in a reaction mixture containing PFOR; similar results were obtained after incubation with sodium dithionite. When reoxidized in air, the pink-red color returned after a few minutes. The UV-visible absorption spectra of oxidized Rd showed several peaks at 280, 370, 492, 570, and 774 nm (Fig. 1A); at 492 nm, the absorption coefficient was 6.89 mM¹ cm¹, calculated using a molecular weight of 6,500. C. tepidum Rd differed from other bacterial Rds by the presence of at least two absorption bands in the 300-400 nm region, a property that may be unique to only green photosynthetic bacteria (32, 33). The A₄₉₂/A₂₈₀ ratio was 0.343. A molecular weight of 6,500 ± 1,000 was obtained from gel filtration experiments in the presence of 1.0 M NaCl. In the presence of 0.1 M NaCl, the molecular weight increased to 13,000 ± 1,000, indicating that the protein exists as a monomer at high ionic strength. The amino acid composition of C. tepidum Rd also yielded an approximate molecular weight of 6,500 (data not shown), which is typical of other Rds (30, 31, 43-45). Electrophoresis of the purified Rd gave rise to a single protein band (Fig. 1B), which migrated with the dye front after SDS-PAGE using a discontinuous Tris-Tricine buffer system (36). This behavior is typical of acidic proteins such as Rd or Fd even in the presence of 20% acrylamide (46). PFOR was shown to have a homodimeric structure comprised of two 125 ± 1 kDa subunits, as determined by SDS-PAGE (Fig. 1C), with a native molecular weight of 250,000 ± 10,000, as determined by gel filtration. The N-terminal amino acid sequence of the C. tepidum Rd was determined to be MQKWVCVPCGY-DPAD-, which indicated high similarity with other previously isolated Rds of anaerobic bacteria (30, 31, 43, 44). The midpoint reduction potential value of C. tepidum Rd, 87 mV (Fig. 2), is more negative than other sources of Rd (31, 33, 45, 47-49), perhaps indicating that the C. tepidum Rd has more negatively charged amino acids at neutral pH. The reduction potential of Rd from the hyperthermophilic archaeon Pyrococcus furiosus is highly temperature-dependent, changing from about 0 mV at 25 °C to about 160 mV at 90 °C at pH 8.0 (47). Thus, the in vivo reduction potential of Rd from C. tepidum, with an optimum growth temperature of around 48 °C, must be even more negative than the value determined in vitro (87 mV at 25 °C).

View larger version (38K): [in this window] [in a new window]   Fig. 1.   A, UV-visible absorption spectra, including inset, of C. tepidum Rd in 50 mM Tris-HCl, pH 8.0. The solid line refers to air oxidized protein, and the dotted line was obtained after reduction in the presence of sodium dithionite. B, SDS-PAGE of purified Rd from C. tepidum. Lane 1, standard marker proteins (Mr = 17-2.5 kDa); lane 2, Rd treated with 2% SDS and heated at 75 °C for 1 h in the presence of -mercaptoethanol. C, SDS-PAGE of purified C. tepidum PFOR. Lane 1, standard marker protein (Mr = 36-205 kDa); lane 2, purified PFOR.

View larger version (15K): [in this window] [in a new window]   Fig. 2.   Cyclic voltammogram of C. tepidum Rd. Bulk solution voltammetry of 300 µM Rd in 0.1 M NaCl, 25 mM potassium phosphate buffer, pH 7.5 at 25 °C; scan rate 20 mV/s. MgCl2 (50 mM) was added to promote and stabilize the response.

EPR and CD Spectra of C. tepidum Rd-- The near UV-visible CD spectra of oxidized C. tepidum Rd closely resembled spectra obtained previously (31, 45, 50), with the exception of the absorption maximum at 385 nm and the shoulder at 420 nm (Fig. 3A). The optical absorption and CD spectra of reduced and oxidized Rd at wavelengths from 300 to 600 nm arise from optically active charge-transfer transitions in the iron-sulfur chromophore, which are presumably sensitive to changes in conformation. The CD features above 320 nm were lost when the protein was reduced (Fig. 3A). The far UV-CD spectrum of C. tepidum Rd showed a weak negative band near 225 nm, a strong positive transition between 200 and 215 nm, and also a strong negative band between 185 and 190 nm (Fig. 3B), which usually signifies an antiparallel -sheet environment. This property is very similar to Rd from P. furiosus (50).

View larger version (18K): [in this window] [in a new window]   Fig. 3.   Circular dichroism spectra of isolated C. tepidum Rd (in 50 mM potassium phosphate buffer, pH 7.2) at 25 °C in the near UV-visible (A) and far UV (B). The solid line refers to air oxidized protein, and the dotted line indicates the spectrum obtained after dithionite reduction.

Reduced Rd was produced when air-oxidized Rd was added to a complete PFOR assay system under anaerobic conditions; this occurred in the complete absence of artificial electron acceptors such as methyl viologen. Under these conditions, PFOR apparently uses oxidized Rd as an electron acceptor. This was manifested by a change from the pink-red color observed upon air oxidation and the appearance of a strong EPR signal (Fig. 4A). Reduced Rd was not EPR-active, as expected (Fig. 4B). Oxidized Rd from C. tepidum exhibited a principal resonance at g = ~4.3, which is composed of a narrow sharp component, at g = 4.322 with broad shoulders to high and low field; in addition, a smaller feature at g = 9.675 was observed. The EPR features at g = ~4.3 and g = ~9.7 regions were similar to states previously observed for other Rds (31, 45, 48, 49, 51) and have been assigned to excited and ground state transitions, respectively. EPR spectral resonance signals at g = ~4.3 and g = ~9.7 were temperature-dependent, increasing in amplitude with decreasing temperature below 16.0 K (Fig. 5, A and B). These results differ from other Rds where the g = ~4.3 region decreases with temperature (31, 45, 51). The relative change in intensity of the temperature-dependent EPR signal at the g = ~9.7 region was over 2.5-fold more affected than the g = ~4.3 region under 10 K (Fig. 5C). Thus, it is conceivable that future studies below the 4.7 K limit of the presently available instrumentation might indicate that the g = ~9.7 region may be increased while the g = ~4.3 region may be reduced.

View larger version (28K): [in this window] [in a new window]   Fig. 4.   EPR spectra of C. tepidum Rd. A, magnetic field (50-750 mT) of aerobically purified Rd; (B) Magnetic field (50 to 750 mT) of aerobically purified Rd reduced by pyruvate- and coenzyme A-dependent PFOR activity at 30 °C for 15 min. The 1-ml reaction mixture contained 5 mM pyruvate, 0.1 mM ThDP, 2.0 mM MgCl2, 1.0 mM coenzyme A, 167 nM PFOR, and 800 µM Rd in 50 mM Tris-HCl, pH 7.8, under anaerobic conditions. The EPR parameters were: center field, 415 mT; microwave power, 10 milliwatts; modulation amplitude, 0.505 mT (A) and 0.201 (B); the receiver gain, 1 × 104 for A and 5 × 103 for B; microwave frequency, 9.45 GHz; modulation frequency, 100 kHz.

View larger version (17K): [in this window] [in a new window]   Fig. 5.   Temperature-dependent EPR spectral resonances of C. tepidum Rd. A, magnetic field (149-164 mT) of the aerobically purified Rd at 4.7-16.0 K. B, magnetic field (62-78 mT) of aerobically purified Rd at 4.7-16.0 K. Each feature was shown at the same value of intensity. The EPR parameters were: center field, 156 mT (A) and 70 mT (B); microwave power, 10 milliwatts; modulation amplitude, 0.505 mT (A) and 0.201 mT (B); the receiver gain, 2 × 103 at 4.7 K and 5 × 103 at 15.0 K (A) and 1 × 104 (B); microwave frequency, 9.45 GHz; modulation frequency, 100 kHz. C, temperature-dependent relative intensity of EPR spectra at the g = ~9.7 region () and g = ~4.3 region ().

C. tepidum PFOR-dependent Reduction of C. tepidum Rd-- As a result of the EPR results, which indicated that PFOR activity caused reduction of oxidized Rd, additional experiments were undertaken to characterize the specificity of this interaction. The catalytic activity of PFOR was coupled to the time-dependent and pyruvate/coenzyme A-dependent change of the absorption spectra of Rd (Fig. 6A). Rd was not reduced if pyruvate, coenzyme A, or PFOR were omitted from the reaction mixture, suggesting that full PFOR activity was necessary to reduce oxidized Rd. During this reaction, the amounts of acetyl-CoA produced, as well as the amount of pyruvate consumed, were also determined (Fig. 6B). The rate of acetyl-CoA formation from pyruvate depended on the amounts of added Rd supplied. Moreover, the specificity of Rd was suggested by substituting either of two different Fd molecules isolated from this organism² for an equivalent amount of Rd (approximately 90 µM for each electron donor), resulting in 30- and 64-fold less activity, respectively (results not shown). In this catabolic reaction, the iron-sulfur clusters of PFOR release electrons to Rd, resulting in the formation of acetyl-CoA (52, 53). In the presence of excess oxidized Rd, this catalytic reaction was continuous. However, after the iron-sulfur clusters of PFOR are reduced in the absence of Rd or some other electron acceptor, this catalytic reaction eventually stops and acetyl-CoA can no longer be generated. As expected, no acetyl-CoA was produced if pyruvate, coenzyme A, or PFOR were omitted (Table I). If Rd was omitted or if it was heated at 100 °C for 1 h, virtually all pyruvate-dependent acetyl-CoA formation was lost. Rd did not blanch, however, when boiled at 100 °C over 1 h. Only after over 1.5~2 h at 100 °C did the pink-red color of Rd disappear, as determined by the visible absorption spectrum at 492 nm (data not shown). Thus, Rd from this moderate thermophile possesses high thermostability. Because it is known that P. furiosus PFOR catalyzes the conversion of pyruvate to acetaldehyde (53) via a coenzyme A-dependent pyruvate decarboxylase-like reaction (Equation 4), it is conceivable that the approximate 2-fold greater amount of pyruvate consumed compared with acetyl-CoA formed might be due to ancillary reactions of this type. However, we were unable to detect a pyruvate decarboxylation reaction catalyzed by C. tepidum PFOR; this and other unique properties of C. tepidum PFOR/PS will be described at a later date.²

View larger version (24K): [in this window] [in a new window]   Fig. 6.   A, pyruvate- and coenzyme A-dependent spectral changes of C. tepidum Rd catalyzed by PFOR. The 1-ml reaction mixture contained 5 mM pyruvate, 0.1 mM ThDP, 2.0 mM MgCl2, 1.0 mM coenzyme A, 42 nM PFOR, and 90 µM Rd in 50 mM Tris-HCl, pH 7.8, under anaerobic conditions. The reaction was initiated by the addition of coenzyme A to the reaction mixture under anaerobic conditions. Interval spectra were shown every 30 s at 30 °C. B, acetyl-CoA production and pyruvate consumption by Rd-dependent PFOR activity. The 1-ml reaction mixtures contained 0.5 mM pyruvate, 0.5 mM coenzyme A, 0.1 mM ThDP, 1.0 mM MgCl2, and 63 nM PFOR in 50 mM Tris-HCl, pH 7.8, under anaerobic conditions. Acetyl-CoA was determined using a coupled enzymatic reaction with malate dehydrogenase/citrate synthase. The consumption of pyruvate was determined using lactate dehydrogenase.

Finally, Rd was found to function specifically in the PFOR reaction of the bifunctional C. tepidum PFOR/PS protein and not the PS reaction. A CO₂ fixation assay for PS activity was optimized for this enzyme, based on a previously employed method for measuring the reductive carboxylation of acetyl-CoA (2, 3). It was apparent that two different reduced Fd proteins isolated from this organism² were much more effective than Rd in supporting PS activity (Table II). It was also apparent that there was some specificity in the Fd that best supported PS activity.

View this table: [in this window] [in a new window]   Table II Pyruvate synthase reaction of C. tepidum PFOR/PS The complete system contained 50 µg PFOR, heated spinach chloroplast fragments (~1.5 mg of protein) and approximately 200 µg of either Rd or Fd from the indicated source, all in 50 mM phosphate buffer, pH 7.0, containing 1.0 mM acetyl-CoA, 2.0 mM MgCl2, 0.1 mM ThDP, 10 mM -mercaptoethanol, and 5 mM [14C]-NaHCO3 (2 µCi of H14CO3) in a 1-ml reaction mixture. The reaction was initiated with bicarbonate after preincubation of the reaction mixture for 30 min and was run at 48 °C for 30 min under a nitrogen atmosphere at a light intensity of 10,000 lux. The reaction was then quenched by the addition of propionic acid, and the acid stable pyruvate was counted by liquid scintillation spectrometry.

	DISCUSSION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

During the course of investigating the enzymology of reductive tricarboxylic acid cycle enzymes of C. tepidum, several low molecular weight electron transfer proteins were isolated, some of which were associated with CO₂ fixation and light-dependent energy generation. Rd was of particular interest because it was shown to be required as an electron acceptor for PFOR activity. Further work indicated that this Rd was highly stable because little change in the absorption spectrum at 492 nm was noted even after exposure to boiling temperatures for 1 h or less. Because virtually all pyruvate-dependent acetyl-CoA formation was lost under the same conditions, it would appear that this catalytic function might serve as a convenient measure of thermal stability independent of any bleaching of its redox center. The major EPR signals resembled those obtained for other Rds. The signal at g = 4.322 was much more prominent than the signal at g = 9.675; in addition, the sharp EPR signal at g = 4.322 was different from other Rds, which usually exhibit broader signals at g = 4.1-5.6 (31, 45, 49, 51). The rhombicity of the S = 5/2 system is approximately E/D = 0.31. The g value is consistent with a  = 1/2 subspectrum for the g = 9.7 region and with a  = 3/2 subspectrum for the g = 4.3 region. Because the EPR signals of Rd disappeared and the molecule was completely reduced upon the addition of the complete PFOR reaction mixture, it would appear that a major function of Rd was as an electron acceptor for PFOR. This was supported by the Rd-dependent oxidation of pyruvate to acetyl-CoA and was further buttressed by the UV-visible spectral changes, which are coenzyme A-dependent, when Rd was reduced via the activity of PFOR. Under the assay conditions described, oxidized Rd appeared to be the favored electron acceptor for the PFOR reaction, because its specificity was 30- and 64-fold greater, respectively, than two different Fds² purified from this organism. The specificity for PFOR in this reaction was further indicated by the fact that as little as 167 nM PFOR was capable of completely reducing 800 µM of Rd within 15 min at 25 °C. Therefore, the results obtained indicate a potentially important role for this small protein in controlling anabolic and catabolic metabolism at the level of PFOR/PS. However, future genetic and physiological experiments must be performed to establish the actual role for Rd in C. tepidum. In addition, the same enzyme that catalyzes PFOR activity (Equation 2) unequivocally catalyzes the PS reaction, basically the CO₂ assimilation or anabolic reaction where acetyl-CoA is reductively carboxylated to form pyruvate (Equation 1). However, only reduced Fd was employed for the PS reaction because Rd was shown to be virtually unable to support CO₂ fixation under the conditions reported.

PFOR is known to contain redox sensitive Fe-S clusters, as does the enzyme from C. tepidum.² Therefore, in the presence of pyruvate and coenzyme A under anaerobic conditions, the enzyme is reduced from the oxidized form (9-11, 52). As a result of PFOR activity, acetyl-CoA is produced from pyruvate, with the reduced 4Fe-4S cluster of the enzyme reoxidized by the electron acceptor. As for PFOR and Rd of C. tepidum, it is not yet clear how reduced Rd may be reoxidized in the cell; however, it is feasible that an NAD-dependent hydrogenase or a Rd/Fd oxidoreductase may be involved to balance the reduction potentials in the cell. In vitro, with only Rd and PFOR present, the reduced iron-sulfur clusters at the active site of PFOR are catalytically more important than when the centers are oxidized, resulting in a shift of the apparent midpoint potential toward the positive direction, thus allowing Rd to function as an electron acceptor. In addition, because the reduction potential of P. furiosus Rd becomes more electronegative as the temperature increases (47), the actual reduction potential of Rd from C. tepidum, which grows at a temperature of 48 °C, may be even more electronegative than the value determined at 25 °C or at the optimum temperature for C. tepidum PFOR activity (58 °C).

It is clear that this phototrophic green sulfur bacterium metabolizes both pyruvate and acetate and that the formation of a key intermediate like acetyl-CoA via PFOR must be an important regulatory step. The fact that Rd may serve as an electron acceptor for PFOR could thus be very important as an appreciation of the metabolic control profile of C. tepidum develops. Several efforts have been made to assign specific functions to redox carriers such as Rd, Fd, and flavodoxin or other proteins such as cytochrome c₅₅₀ and cytochrome c₅₅₃, all of which have recently been purified from C. tepidum.³ C. tepidum is an organism for which a functional genetic system exists (34); thus it should be feasible to elucidate the physiological role of Rd and other electron carriers, especially since the genomic sequence of this organism has recently reached the completion stage. Such studies will nicely complement biochemical studies with purified proteins, e.g. Rd and PFOR. Certainly, the fact that Rd participates in the PFOR reaction but is not efficiently used in the PS reaction of this bifunctional protein under the reported conditions suggests that PFOR/PS catalyzes two fundamentally different reactions, with different catalytic mechanisms, for diverse physiological purposes.

	ACKNOWLEDGEMENT

We express our appreciation to Dr. Chulhwan Kim for determining the reduction potential of Rd by cyclic voltammetry.

	FOOTNOTES

^* This work was supported by Department of Energy Grant DE-FG02-91ER20033 (to F. R. T.) and National Institutes of Health Grant GM52322 (to R. H.).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.

^§ Supported by an Ohio State University postdoctoral fellowship during the early portion of this study.

To whom correspondence should be addressed: Dept. of Microbiology, Ohio State University, 484 West 12^th Ave., Columbus, OH 43210-1292. Tel.: 614-292-4297; Fax: 614-292-6337; E-mail: tabita.1@osu.edu.

² K.-S. Yoon and F. R. Tabita, manuscript in preparation.

³ K. S. Yoon and F. R. Tabita, unpublished results.

	ABBREVIATIONS

The abbreviations used are: Fd, ferredoxin; PS, pyruvate synthase; PFOR, pyruvate ferredoxin/flavodoxin oxidoreductase; ThDP, thiamine diphosphate; Rd, rubredoxin; PAGE, polyacrylamide gel electrophoresis; Tricine, N-[2-hydroxy-1,1-bis(hydroxymethyl)ethyl]glycine; mT, millitesla.

	REFERENCES

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