©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
Proton-translocating Nicotinamide Nucleotide Transhydrogenase of Escherichia coli
INVOLVEMENT OF ASPARTATE 213 IN THE MEMBRANE-INTERCALATING DOMAIN OF THE SUBUNIT IN ENERGY TRANSDUCTION (*)

Mutsuo Yamaguchi , Youssef Hatefi (§)

From the (1)Division of Biochemistry, Department of Molecular and Experimental Medicine, The Scripps Research Institute, La Jolla, California 92037

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES

ABSTRACT

Mutations in the subunit of Escherichia coli proton-translocating nicotinamide nucleotide transhydrogenase of the conserved residue Asp-213 to Asn (D213N) and Ile (D213I) resulted in the loss, respectively, of about 70% and 90% NADPH 3-acetylpyridine adenine dinucleotide (AcPyAD) transhydrogenation and coupled proton translocation activities. However, the cyclic NADP(H)-dependent NADH AcPyAD transhydrogenase activities of the mutants were only 35% inhibited. The latter transhydrogenation, which is not coupled to proton translocation, occurs apparently via NADP under conditions that enzyme-NADP(H) complex is stabilized. Mutations D213N and D213I also resulted in decreases in apparent K for the NADPH AcPyAD and S (NADPH concentration needed for half-maximal activity) for the cyclic NADH AcPyAD transhydrogenation reactions, and in K , as determined by equilibrium binding studies on the purified wild-type and the D213I mutant enzymes. These results point to a structural role of Asp-213 in energy transduction and are discussed in relation to our previous suggestion that proton translocation coupled to NADPH NAD (or AcPyAD) transhydrogenation is driven mainly by the difference in the binding energies of NADPH and NADP.


INTRODUCTION

The energy-transducing nicotinamide nucleotide transhydrogenases constitute a class of membrane-bound enzymes which catalyze the reaction shown in Equation 1.

On-line formulae not verified for accuracy

The bovine enzyme is a homodimer of monomer M = 109,065 (Yamaguchi et al., 1988). The monomer is composed of 3 domains, a hydrophilic 430-residue-long NH-terminal domain that binds NAD(H), a hydrophobic 400-residue-long central domain that is largely membrane-intercalated and harbors the proton channel of the enzyme, and a hydrophilic 200-residue-long COOH-terminal domain that binds NADP(H) (Yamaguchi et al., 1988; Yamaguchi and Hatefi, 1991; Hatefi andYamaguchi, 1992). The enzyme exhibits half-of-the-sites reactivity. It binds 1 mol of NADH and 1 mol of NADPH per dimer (Yamaguchi and Hatefi, 1993), and its transhydrogenase activity is completely inhibited by covalent modification with 1 mol of [C]N,N`-dicyclohexylcarbodiimide (Phelps and Hatefi, 1984; Wakabayashi and Hatefi, 1987a) or 1 mol of [H]FSBA()(Phelps and Hatefi, 1985; Wakabayashi and Hatefi, 1987b) per dimer.

The transhydrogenases of Escherichia coli and Rhodobacter capsulatus are each composed of two unlike subunits, with M 53,000 and with M 48,000 (Clarke et al., 1986; Lever et al., 1991; Ahmad et al., 1992). The E. coli enzyme has been shown to be an heterotetramer (Hou et al., 1990). The transhydrogenase of Rhodospirillum rubrum chromatophores has 3 subunits, designated 1, 2, and for consistency with the earlier subunit designations of the E. coli enzyme (Yamaguchi and Hatefi, 1994; see also Williams et al.(1994)). The molecular masses of the R. rubrum transhydrogenase subunits are 1 = 40.3 kDa, 2 = 14.9 kDa, and = 47.8 kDa. Subunit 1 is water-soluble and easily detached from chromatophores. Together the two subunits of the E. coli or the three subunits of the R. rubrum enzyme (the Rb. capsulatus transhydrogenase has not been sequenced) exhibit the same tridomain hydropathy profile as the bovine enzyme (Yamaguchi and Hatefi, 1994). In addition, the predicted amino acid sequences of putative single-subunit transhydrogenases from Eimeria tenella (Kramer et al., 1993; Vermeulen et al., 1993) and Entamoeba histolytica (Yu and Samuelson, 1994) exhibit a similar hydropathy profile, except that they could be described as -tail-to--head spliced versions of the E. coli enzyme.

We have shown earlier that the bindings of NADPH and NADP, but not of NADH and NAD, alter the conformation of the bovine transhydrogenase in different ways and have argued on the basis of both theoretical considerations and experimental results that inside-to-outside proton translocation by the enzyme (reversal of Equation 1) is accomplished through substrate-induced conformation change of the protein and is paid for by the difference in the binding energies of substrates (NADPH + NAD) versus products (NADP + NADH). Because NADPH and NADP binding, but not NADH and NAD binding, results in detectable conformation changes of the bovine transhydrogenase, we further proposed that it is mainly the difference in the binding energies of NADPH and NADP that drives inside-to-outside proton translocation via protein conformation changes (Yamaguchi and Hatefi, 1989; Yamaguchi et al., 1990; Hatefi and Yamaguchi, 1992).

The five transhydrogenases whose amino acid sequences are known contain regions of high sequence identity in the NAD(H) and especially the NADP(H) binding domains as well as in a 19-residue-long and a 58-residue-long stretch in the hydrophobic domain.()The 58-residue-long stretch is adjacent to the hydrophilic domain that binds NADP(H) (Fig. 1). Because such stretches of high sequence identity in the hydrophobic domains of related enzymes are rare, we felt that these membrane-intercalating regions may have functional significance with regard to the enzyme's proton translocation activity. In the entire 400-residue-long hydrophobic domains of the five sequenced transhydrogenases, there are only 4 conserved charged residues and 3 conserved histidine residues. In the 19-residue-long stretch of high sequence identity, there is a conserved histidine residue (E. coli His-91, see Fig. 1) which when mutated in the E. coli enzyme to Ser, Thr, or Cys results in 90% loss of transhydrogenation and proton pumping activities (Holmberg et al., 1994). As seen below, there is in the 58-residue-long stretch a conserved aspartic acid residue (E. coli Asp-213, see Fig. 1) whose mutation in the E. coli enzyme to Asn resulted in our hands in 70% activity loss (see also Holmberg et al.(1994)), but its mutation to Ile caused 90% loss of transhydrogenase and coupled proton translocation activities. Neither mutation greatly inhibited the NADP(H)-dependent cyclic NADH AcPyAD transhydrogenation activity of the enzyme, which involves both the NAD and the NADP binding sites, but is not coupled to proton translocation (see below). However, the D213I mutation increased the affinity of the enzyme for NADPH by 3-fold. The implications of these results with regard to energy transduction by the transhydrogenase will be discussed.


Figure 1: Locations in the subunit of E. coli transhydrogenase and compositions of the 19-residue-long and the 58-residue-long hydrophobic stretches of high sequence identity. The top rectangles depict that and the subunits of the E. coli transhydrogenase, with their hydrophilic NAD(H) and NADP(H) binding domains (unshaded) and their hydrophobic domains (shaded). The amino acid sequences of the 19- and the 58-residue-long stretches of high sequence identity (heavily shaded areas) are shown below the rectangles for the 5 sequenced transhydrogenases. The locations of His-91 (H91) and Asp-213 (D213) are marked.




EXPERIMENTAL PROCEDURES

Materials

NADPH, NAD, and ATP were obtained from Calbiochem; AcPyAD and NAD kinase were from Sigma; asolectin was from Associated Concentrates; Ultrogel AcA34 was from IBF Biotechnics; DEAE-Bio-Gel A agarose and Muta-Gene Phagemid In Vitro Mutagenesis Kit were from Bio-Rad; Sequenase version 1.0, nucleotide kit for DNA sequencing with 7-deaza-dGTP, and random-primed DNA labeling kit were from U. S. Biochemical Corp., and [4-H]NAD (1200 mCi/mmol) was from Amersham. Bacterial strains and plasmids were obtained from the following sources: E. coli strain MC4100 (F, araD139, (arg F-lac)U169, ptsF25, relA1, flb5301, rpsL 150.) from ATCC; pBluescript II KS(+) from Stratagene; pUC4K plasmid from Pharmacia Biotech; pLC10-19 from the Clarke and Carbon Collection, carrying the E. coli transhydrogenase gene (Clarke and Bragg, 1985), was supplied by Dr. B. Bachman (E. coli Genetic Stock Center, Yale University); pDC21, a derivative of pUC13, carrying the E. coli transhydrogenase gene (Clarke and Bragg, 1985), was the generous gift of Dr. P. D. Bragg, University of British Columbia; and pMAK705, whose replication is temperature-sensitive and carries the chloramphenicol resistance gene (Hamilton et al., 1989), was kindly supplied by Dr. S. R. Kushner, University of Georgia.

Construction of Transhydrogenase Deletion Mutant

Basic recombinant DNA techniques were performed according to Sambrook et al.,(1989). From pLC10-29, a 4.8-kilobase HindIII fragment carrying the transhydrogenase coding region was excised and inserted into the HindIII site of pBluescript II. By digestion of the constructed plasmid with HpaI, a 2.7-kilobase fragment, covering most of the transhydrogenase coding region, was cut out and replaced by the kanamycin resistance gene cartridge from pUC4K. Then the whole insert was excised with EcoRV and inserted into the HindIII site of pMAK705 (pMAKTK). As described by Hamilton et al.(1989), pMAKTK was integrated into the chromosomal DNA of E. coli strain MC4100, and then the transhydrogenase genes were replaced by the kanamycin resistance gene (strain MC4100 TH). The gene replacement was confirmed by Southern blotting (Southern, 1975).

Site-directed Mutagenesis

The transhydrogenase genes were removed from pDC21 by double digestion with BamHI and HindIII and ligated with pTZ18U digested with BamHI and HindIII. With this pTZ18U plasmid, site-directed mutagenesis was carried out to convert Asp-213 to Asn or Ile, using the reagents and protocol as outlined in the Bio-Rad Muta-Gene Mutagenesis Kit (Kunkel et al., 1987). The plasmid DNA was prepared from individual colonies, and the mutants were identified by double-strand DNA sequencing (Chen and Seeburg, 1985). After confirming the DNA sequence, each Tth111I-BssHII 450-base pair DNA fragment was excised and replaced with the counterpart of pDC21.

Enzyme Assays

Transhydrogenation from NADPH to AcPyAD was assayed spectrophotometrically at 375 nm at 37 °C in a reaction mixture containing 50 mM sodium phosphate (pH 7.0), 0.01% Brij 35, 10 µg of lysophosphatidylcholine, and 0.5 mM each of NADPH and AcPyAD. An extinction coefficient of 6.1 mM cm was used to calculate the rates. Transhydrogenation from NADH to AcPyAD in the presence of NADP or NADPH was measured spectrophotometrically at 375 nm at 37 °C in a reaction mixture containing 10 mM MES/KOH (pH 6.0), 10 µg of lysophosphatidylcholine, 0.2 mM NADP or NADPH, and 0.2 mM AcPyAD. The reaction was started by the addition of 20 µM NADH. Protein concentration was measured by the method of Peterson(1977). One unit of activity is defined as 1 µmol of AcPyAD reduced by NADPH per min.

9-Aminoacridine fluorescence was measured at 37 °C by an SLM photon-counting fluorescence spectrophotometer at the excitation wavelength of 420 nm and the emission wavelength of 500 nm. To the reaction mixture (2.0 ml), containing 10 mM Tricine/NaOH (pH 8.0), 2.5 mM MgSO, 0.25 mM dithiothreitol, and 0.2 mM EDTA, were added 20 µg of liposome-reconstituted transhydrogenase and 1 µM 9-aminoacridine followed by 0.2 mM NADPH. Then, 0.4 mM AcPyAD and 1 µM carbonyl cyanide p-trifluoromethoxyphenylhydrazone were added as indicated, and the fluorescence changes were monitored at 500 nm.

Culture of E. coli Cells and Preparation of Membranes

E. coli strain MC4100 TH was transformed with pDC21 or its mutated plasmids. Each single colony was inoculated into the LB medium containing ampicillin (100 µg/ml). Cells were grown until the late logarithmic phase, collected by centrifugation at 8,000 rpm for 10 min (Sorvall GSA rotor), and washed with 0.9% NaCl. The cells (wet weight, 14 g) were then suspended in 140 ml of 50 mM Tris-HCl (pH 7.8), containing 1 mM EDTA and 1 mM dithiothreitol (TED), and sonicated in a Branson sonifier at output 8 and 25% pulse for 10 min. Unbroken cells were removed by centrifugation at 8,000 rpm for 10 min, and membranes were collected by centrifugation at 39,000 rpm for 45 min (Beckman 42Ti rotor). Membranes were suspended in a small volume of TED and homogenized.

Purification of Enzymes

E. coli membranes (30 mg of protein) were suspended in 6 ml of TED, and 1.5 ml of 10% Triton X-100 was slowly added. After stirring for 15 min at room temperature, the sample was centrifuged at 48,000 rpm (Beckman 50Ti rotor) for 45 min. The supernatant (15 ml) was directly loaded on a DEAE Bio-Gel A agarose column (1.5 14 cm) equilibrated with TED, containing 0.125 M NaCl and 0.02% potassium cholate. The enzyme was eluted by a linear NaCl concentration gradient from 0.125 M to 0.5 M in TED, containing 0.02% potassium cholate. Approximately 1-ml fractions were collected. The active fractions eluted at 0.20 M to 0.23 M NaCl were combined and concentrated to 1.0 ml by a Centricon-30 concentrator and loaded on an Ultrogel AcA34 column (1.5 68 cm) equilibrated with TED, containing 0.02% potassium cholate. Active fractions were combined (10 ml) and concentrated to 700 µl by a Centricon-30 concentrator.

Reconstitution of the Purified Enzymes into Liposomes

Essentially the protocol developed by Sone et al.(1977) was followed. Asolectin (200 mg) and cholic acid (150 mg) were added to 10 ml of 10 mM Tricine/NaOH (pH 8.0), containing 5 mM dithiothreitol and 0.2 mM EDTA. After adjustment of pH to 8.0 with 1 M KOH, the mixture was sonicated to clarity. The purified enzyme (100 µg of protein) was added to 2 ml of the asolectin suspension, and the mixture was dialyzed against 1 liter of 10 mM Tricine/NaOH (pH 8.0), containing 2.5 mM MgSO, 0.25 mM dithiothreitol, and 0.2 mM EDTA for 20 h.

Preparation of [4-H]NADPH

[4-H]NADP was prepared by phosphorylation of [4-H]NAD (1200 mCi/mmol) in the presence of ATP and NAD kinase essentially according to Wang et al.(1954). [4-H]NAD and [4-H]NADP were separated on a Dowex 1 formate (200-400 mesh) column (0.7 6 cm). [4-H]NAD was eluted with 0.6 M formic acid, and [4-H]NADP was eluted with 1.2 M formic acid (Hatefi et al., 1980). The contents of the tubes containing [4-H]NADP were combined and lyophilized. [4-H]NADP was reduced in 0.5 ml of 10 mM sodium phosphate, pH 7.5, containing 4 mM MgCl, 7.5 mMD-isocitrate, and 120 µg of isocitrate dehydrogenase. The solution was incubated at room temperature and monitored spectrophotometrically for NADP reduction. After completion of the reaction (A/A = 2.3), the solution was heated in a boiling water bath for 2 min to inactivate and precipitate the enzyme. The resulting supernatant obtained after centrifugation was calibrated spectrophotometrically for [4-H]NADPH concentration and then used for binding studies. The extinction coefficient used for determining the concentration of NADPH was 6.22 mM cm at 340 nm.

Binding Experiments

The centrifugation method of Howlett et al.(1978) was followed for analyzing the binding of nucleotides to peptides. Proteins were mixed with different fixed concentrations of [H]NADPH in 110 µl of 10 mM sodium phosphate (pH 7.0), containing 0.5 mM dithiothreitol and 0.02% potassium cholate. Dextran T40 (2 mg/ml) was also added to provide density stabilization and prevent convective stirring of the tube contents during deceleration of the rotor. Samples were centrifuged at room temperature in a Beckman Airfuge at 30 p.s.i. Ten µl of the top layer before and after centrifugation was assayed for radioactivity, and the values were used to calculate total and free ligand concentrations, respectively. The difference in the concentration of radioactive ligand between the original solution and the supernatant after centrifugation was a direct measure of the amount of bound ligand. The data were treated according to the Scatchard equation for equilibrium binding (Segel, 1975) as before (Yamaguchi and Hatefi, 1993).

Gel Electrophoresis

Enzyme samples were incubated for 1 h at room temperature in 63 mM Tris-HCl, pH 6.8, containing 2% SDS, 5% -mercaptoethanol, 10% glycerol, and 0.002% bromphenol blue, and subjected to electrophoresis on 10% SDS-polyacrylamide slab gels (Laemmli, 1970). Gels were stained with Coomassie Blue and destained.


RESULTS

Free carboxyl groups in the membrane-intercalating domains of proton-translocating enzymes are generally considered as good candidates for participation in transmembrane proton translocation, witness Asp-85 and Asp-96 of bacteriorhodopsin (Rothschild, 1992) and Asp-61 of subunit c of E. coli ATP synthase complex or the corresponding Glu in ATP synthases from other species (Fillingame, 1990). As mentioned earlier, there is in the 400-residue-long hydrophobic domain of the transhydrogenase subunits a single conserved aspartic acid residue (E. coli Asp-213), which is located in a 58-residue-long segment of high sequence identity among the 5 sequenced transhydrogenases from diverse sources (Fig. 1). This 58-residue-long segment immediately precedes the extramembranous NADP(H) binding domain, and, for the reasons mentioned in the introduction, makes it a possible candidate for involvement in energy transduction and the enzyme's proton channel. Hence, it was of interest to investigate the role of the E. coli transhydrogenase Asp-213 in transhydrogenation and proton translocation by site-directed mutagenesis.

For this purpose, it was considered important to express the mutated forms of the enzyme in a strain of E. coli from which the wild-type transhydrogenase genes had been deleted (TH), so that no background wild-type activity, no matter how small, would complicate the interpretation of results. This was done as described under ``Experimental Procedures,'' and the deletion of transhydrogenase genes was confirmed by Southern blotting. Then, in this strain were expressed the wild-type enzyme as well as two mutants in which Asp-213 was mutated to Asn (D213N) and Ile (D213I).

Enzymatic Properties of the Mutated Enzymes

Fig. 2shows the transhydrogenase activities of E. coli membranes containing the expressed wild-type (A) and the D213N (B) and D213I (C) mutants. The transhydrogenase activity of membranes from the TH strain (D) is also shown. The minimal activity seen in trace D is probably due to another membrane-bound protein. Mutation of Asp-213 to Asn and Ile resulted, respectively, in a 72% and 91% loss of transhydrogenase activity (see Fig. 2legend). These data indicated that, even though Asp-213 does not appear to be an essential residue, its mutation (especially to Ile) results, nevertheless, in a dramatic loss of transhydrogenase activity. For further characterization of the mutants, all three expressed enzymes were purified from the E. coli membranes (Fig. 3).()The purified enzymes were then incorporated into liposomes and assayed for transhydrogenation-coupled proton translocation. As seen in Fig. 4, they were all capable of proton translocation upon initiation of NADPH AcPyAD transhydrogenation, and it was apparent that the proton translocation capability of the D213I mutant was less than those of the wild-type and the D213N mutant enzymes. The initial rates of 9-aminoacridine fluorescence quenching upon initiation of the reaction by addition of AcPyAD indicated that the rates of proton translocation by the D213N and the D213I mutant enzymes were, respectively, 25% and 9% of the proton translocation rate of the wild-type enzyme. These values agreed well with the relative transhydrogenation activities of the purified enzymes as given in the legend to Fig. 3. A more interesting difference between the wild-type and mutant transhydrogenases was, however, the effect the mutation of Asp-213 had on the apparent K values of the substrates. As seen in , the apparent K values for NADPH and AcPyAD, as determined from Lineweaver-Burk double reciprocal plots, were diminished in the mutant enzymes. This change was particularly noteworthy for the apparent K of the D213I mutant, which had decreased by about 3.5-fold as compared to that of the wild-type transhydrogenase.


Figure 2: Comparison of transhydrogenase activities of E. coli membranes containing expressed wild-type and mutant enzymes. The NADPH to AcPyAD transhydrogenase activities shown were each catalyzed by 28 µg of membrane protein. A, wild-type; B, D213N mutant; C, D213I mutant; D, MC 4100TH without transformation. Specific activities of A, B, and C, calculated from the initial rates of the progress curves shown, are 5.4, 1.52, and 0.48 µmol of AcPyAD reduced (minmg of protein), respectively. Assay conditions are described under ``Experimental Procedures.''




Figure 3: SDS-polyacrylamide gel electrophoretic patterns of purified wild-type and mutant transhydrogenases. SDS-polyacrylamide gel electrophoresis was carried out as described under ``Experimental Procedures.'' To each lane 3 µg of purified enzyme were added. A, wild-type enzyme; B, D213N mutant; C, D213I mutant. The specific activities of the purified enzymes A, B, and C were, respectively, 22.5, 5.14, and 2.20 µmol of AcPyAD reduced (minmg of protein).




Figure 4: Proton translocation by proteoliposomes of wild-type and mutant transhydrogenases. The reaction mixture (2.0 ml) contained 10 mM Tricine/NaOH, pH 8.0, 2.5 mM MgSO, 0.25 mM dithiothreitol, 0.2 mM EDTA, 20 µg of liposome-reconstituted transhydrogenase, 1 µM 9-aminoacridine, and 200 µM NADPH. Where indicated, 400 µM AcPyAD was added to start transhydrogenation and proton translocation, and 1 µM carbonyl cyanide p-trifluoromethoxyphenylhydrazone was added to uncouple. The fluorescence changes of 9-aminoacridine were monitored at 500 nm. Excitation was at 420 nm. A, wild-type enzyme; B, D213N mutant; C, D213I mutant.



Equilibrium Binding of [H]NADPH to the Wild-type and D213I Mutant Enzymes

Because the decreases in the substrate K values were accompanied by V decreases, it was important to see whether these K decreases were a reflection of k decreases, or there was in fact a change in the affinity of the enzyme for substrates. Hence, direct binding studies were done with the purified wild-type and the D213I mutant enzymes, using [H]NADPH. Binding studies with the D213N mutant and with AcPyAD were not done, because the K changes were not large enough to be reliably checked by equilibrium binding studies. The results of equilibrium binding of [H]NADPH to the purified wild-type (A) and the D213I mutant (B) transhydrogenases are shown in the Scatchard plots of Fig. 5. The K values derived from these plots were 19.7 µM for the wild-type transhydrogenase and 6.0 µM for the D213I mutant enzyme. It may be noted that in Fig. 5the abscissa intercepts (moles of [H]NADPH bound per mol of dimeric transhydrogenase) were n = 0.78 in panel A and n = 0.65 in panel B. Ideally, these values should be close to 1.0 for a dimeric, half-of-the-sites reactive enzyme. For the bovine transhydrogenase, we estimated n values of 1.02 for NADH and 0.92 for NADPH (Yamaguchi and Hatefi, 1993). However, we have noticed that the purified E. coli transhydrogenase has a greater tendency to aggregate at the concentrations used for binding studies than the purified bovine enzyme. We, therefore, repeated the experiments of Fig. 5in the presence of different low concentrations of lysophosphatidylcholine, which increase transhydrogenase activity when added to the assay mixture, but there was no improvement in the values of n.


Figure 5: Scatchard plots for binding of [H]NADPH to purified wild-type and D213I mutant transhydrogenases. Wild-type enzyme (A, 2.95 mg/ml) and D213I (B, 2.37 mg/ml) were mixed with different fixed concentrations of [H]NADPH (range 1.9-72.7 µM in A and 1.9-53.5 µM in B) in 110 µl of 10 mM sodium phosphate, pH 7.0, containing 0.02% potassium cholate and dextran T40 (2 mg/ml). The mixtures were centrifuged at 30 p.s.i. for 45 min. Before and after centrifugation, 10-µl aliquots of the upper layer of each mixture were removed, and their radioactivity was measured for calculation of the molar concentrations of free (L) and enzyme-bound (L) ligand. E is the molar concentration of the dimeric enzyme. The data were treated according to the Scatchard equation as described before (Yamaguchi and Hatefi, 1993). The constants obtained in these experiments were: K = 19.7 µM, n = 0.78 (A); K = 6.0 µM, n = 0.65 (B), where n is moles of NADPH bound per mol of dimeric transhydrogenase.



NADP(H)-dependent NADH AcPyAD Cyclic Transhydrogenation

The energy-transducing transhydrogenases of mitochondria and bacteria do not catalyze transhydrogenation from NADH to NAD or AcPyAD, but they do catalyze a slow transhydrogenation from NADPH to NADP or AcPyADP, which was shown in bovine submitochondrial particles to be energy-linked and accelerated by membrane energization (Hatefi et al., 1980; Phelps et al., 1980). However, Fisher and co-workers (Wu et al., 1981) discovered that in reconstituted proteoliposomes, where the normal NADPH NAD transhydrogenation becomes inhibited in the absence of an uncoupler to dissipate the membrane potential, the enzyme does catalyze a rapid and stoichiometric NADH AcPyAD transhydrogenation, which requires NADP(H) and is not coupled to proton translocation. Furthermore, they showed that whereas the stereospecificity of hydride ion transfer in the energy-transducing reaction is [4A]NAD(H) [4B]NADP(H), in the NADP(H)-dependent NADH NAD (or AcPyAD) reaction it is 4A 4A (Wu et al., 1981). A possible explanation offered was that creation of a membrane potential in proteoliposomes prevents the release of enzyme-bound NADP, which accepts a hydride ion equivalent from NADH. Then, the NAD so formed goes off the enzyme and is replaced by AcPyAD, which is reduced by the enzyme-bound NADPH (Wu et al., 1981; Fisher and Earle, 1982). Hutton et al.(1994) have shown recently that this cyclic NADP(H)-dependent NADH AcPyAD transhydrogenation can also be observed at pH 6.0 and low ionic strength with the detergent-solubilized enzyme not incorporated in liposomes, apparently because the enzyme-NADP(H) complex is much more stable under these conditions. In principle, therefore, the cyclic NADH AcPyAD transhydrogenation catalyzed via an enzyme-bound NADP(H) would be analogous to the [4B]NAD(H) [4B]NADP(H) transhydrogenation catalyzed by the so-called BB transhydrogenases. These enzymes are soluble flavoproteins in which the enzyme-bound flavin carries out the act of transferring a hydride ion equivalent from one nicotinamide dinucleotide to another as they bind interchangeably at the same site (Rydström et al., 1987; Lee and Ernster, 1989).

The cyclic transhydrogenase reaction was of interest to us, because it appeared to test the functionality of the catalytic sector of the enzyme without the constraints that an impaired proton channel might exert on the normal NADPH NAD transhydrogenation reaction. In other words, we could ask whether the inhibited D213I mutant was capable of catalyzing the NADP(H)-dependent NADH AcPyAD cyclic transhydrogenation reaction. As seen in Fig. 6A, the purified D213I mutant catalyzed a rapid cyclic transhydrogenation reaction upon successive additions of 20 µM NADH to a reaction mixture containing 0.2 mM each of NADPH and AcPyAD (see also Glavas, 1994). For comparison, the cyclic transhydrogenation reaction catalyzed by the purified, wild-type enzyme is also shown (Fig. 6B). The data of , derived from Lineweaver-Burk double-reciprocal plots, show that the rates of the cyclic transhydrogenation reactions catalyzed by the purified D213N and D213I mutated enzymes were somewhat slower at V than that catalyzed by the wild-type enzyme. However, comparison of the data of Tables I and II clearly indicate that, relative to the rates of the wild-type enzyme, the energy-transducing NADPH AcPyAD transhydrogenase activity of the D213I mutant enzyme was inhibited much more than its cyclic NADH AcPyAD reaction, which is not coupled to proton translocation. An obvious conclusion, therefore, is that the mutation of Asp-213 to Ile interferes with the proton translocation capability of the E. coli transhydrogenase, thereby inhibiting the coupled NADPH AcPyAD transhydrogenation much more than the cyclic NADH AcPyAD reaction. also shows that in the cyclic transhydrogenase reaction the apparent S (the concentration needed for half-maximal activity) of NADPH decreased in the mutated enzymes and that in the D213I mutant it is one order of magnitude smaller than in the wild-type enzyme. These results are in agreement with the apparent K data of and the K values derived from the equilibrium binding experiments of Fig. 5. Together, they suggest that mutations of Asp-213 to Asn and especially to Ile in the hydrophobic, membrane-intercalating domain of the enzyme are communicated to the NADP(H) binding site, altering the enzyme's affinity for NADPH. The implications of these results on the mechanism of energy transduction by the transhydrogenase enzyme are discussed below.


Figure 6: NADPH-dependent cyclic NADH AcPyAD transhydrogenation catalyzed by purified wild-type and D213I mutated transhydrogenases. The reaction mixtures contained 10 mM MES/KOH, pH 6.0, 10 µg of lysophosphatidylcholine, 0.2 mM NADPH, 0.2 mM AcPyAD, 2.8 µg of purified D213I (A) or 2.3 µg of purified wild-type (B) transhydrogenase. After 1 min, 20 µM NADH was added to start the reaction followed by an additional 20 µM NADH where indicated by arrows. Reduction of AcPyAD was monitored at 375 nm.




DISCUSSION

It has been shown here that the mutation in E. coli of the only conserved dicarboxylic acid residue in the hydrophobic domain of the subunit of nicotinamide nucleotide transhydrogenase (Asp-213) to Asn (D213N) or Ile (D213I) diminishes in parallel the hydride ion transfer (NADPH AcPyAD) and the transmembrane proton translocation activities of the enzyme. In the case of the D213I mutant, these activities, as assayed with the purified and liposome-reconstituted enzymes, were 90% lower than the activities of the wild-type transhydrogenase. However, another activity of the D213I mutant, namely, the NADP(H)-dependent cyclic NADH AcPyAD transhydrogenation, which is not coupled to proton translocation, was only about 35% inhibited. As seen in Fig. 1, Asp-213 is located in a hydrophobic stretch of 58 highly conserved amino acid residues, which is immediately adjacent to the COOH-terminal extramembranous domain of the subunit that binds NADP(H). In the hydrophobic domain of the subunit, there is a second, much shorter stretch (19 residues) of high sequence conservation (Fig. 1). This segment contains a conserved His (His-91) whose mutation to Ser, Thr, or Cys also results in 90% loss of NADPH AcPyAD hydride ion transfer and coupled proton translocation activities (Holmberg et al., 1994). Because such regions of high sequence identity are rare in the hydrophobic domains of functionally related proteins, it is possible that in the folded structure of the transhydrogenase these regions of high sequence identity are in close apposition and are together involved as membrane-intercalated helices in the proton translocation function of the enzyme. Whether His-91 and Asp-213 are within these membrane-intercalated helices or in the extramembranous loops that connect the helices is not known. Nor do the catalytic properties of the mutant enzymes cited above indicate whether these amino acid residues play a chemical or a structural role in the coupled reactions of the transhydrogenase. Such distinctions are difficult to make, especially when one considers the recent results of Sonar et al.(1994) on bacteriorhodopsin. These authors have shown that in a Y57D mutant of bacteriorhodopsin the deprotonation of the retinal Schiff base and formation of the M intermediate is blocked. Nevertheless, the Y57D mutant is capable of appreciable proton translocation activity, apparently via a redirected proton pathway. Therefore, in the E. coli transhydrogenase, the incomplete loss of proton translocation activity upon mutations of His-91 and Asp-213 does not necessarily mean that one or both of these residues are not directly involved in proton translocation in the wild-type enzyme.

It has also been shown here that, in addition to 90% inhibition of NADPH AcPyAD transhydrogenation and coupled proton translocation activities, the D213I mutation lowers the apparent K by 3.5-fold in the above reaction (assayed at pH 7.0) and by 10-fold in the cyclic NADH AcPyAD transhydrogenation reaction (assayed at pH 6.0 and low ionic strength). In agreement with these results, it was shown by equilibrium binding experiments (at pH 7.0) that in the D213I mutant enzyme K is 3-fold lower than in the wild-type transhydrogenase. These results indicate, therefore, that Asp-213 or a mutation of this residue influences the affinity of the nearby NADP(H) binding site for its substrate. We have discussed elsewhere that the energetically uphill (inside-to-outside) proton translocation that is coupled to NADPH NAD transhydrogenation (reversal of Equation 1) is driven by the difference in the binding energies of the reactants (NADPH and NAD) and the products (NADP and NADH). This means that the enzyme must undergo a conformation change coupled to hydride ion transfer from NADPH to NAD. Because substrate binding by the bovine transhydrogenase is random, we were able to show that the bindings of NADPH and NADP, but not of NADH and NAD, change the conformation of the enzyme in different ways. We, therefore, proposed that the difference in the binding energies of NADPH and NADP is the principal force that via coupled conformation changes of the protein results in vectorial proton release and uptake across the membrane (for elegant discussions of the utilization of binding energy to drive coupled vectorial processes, including proton translocation, see Jencks(1975, 1989)). Accordingly, if we now assume that the hydrophobic stretches of high sequence identity depicted in Fig. 1are involved in the coupled conformation changes of the transhydrogenase, we could then rationalize why a point mutation in these hydrophobic stretches might alter the affinity of the enzyme for NADPH. In other words, it is possible that the D213I mutation causes a structural change in the hydrophobic domain of the subunit that has a reciprocal effect on the binding energy of NADPH. This structural change could restrict proton translocation and inhibit the coupled transhydrogenation reaction.

Another finding of interest, especially in the case of the D213I mutation, is that the enzyme's cyclic NADH AcPyAD transhydrogenase activity is much less inhibited than its normal, energy-coupled NADPH AcPyAD transhydrogenation reaction. The inhibition of the latter reaction, as discussed above, is consistent with the possibility that the D213I mutation restricts proton translocation and as a consequence inhibits the coupled NADPH AcPyAD transhydrogenation. However, this does not explain why the cyclic NADH AcPyAD reaction should not be equally inhibited as well. The explanation, we feel, lies in the difference between the normal and the cyclic transhydrogenation reactions, more precisely between the role of NADP(H) in each. In the energy-coupled NADPH NAD transhydrogenation reaction, enzyme-bound NADPH and NADP are in equilibrium with their counterparts in the medium. As a result, the enzyme experiences their different binding energies with each turnover and undergoes the conformation changes that drive proton translocation. By contrast, the cyclic NADH NAD transhydrogenation occurs while NADP(H) is enzyme-bound. Hence, the enzyme does not experience the different binding energies of NADPH and NADP and does not undergo the conformation changes that drive proton translocation. This important difference can, therefore, explain why the cyclic NADH AcPyAD transhydrogenation is not coupled to proton translocation and why restriction of the latter process by the D213I mutation does not have a comparable inhibitory effect on the cyclic transhydrogenation reaction.

  
Table: Kinetic parameters of wild-type and mutant enzymes for the NADPH AcPyAD transhydrogenation reaction

The apparent K and V values were derived from Lineweaver-Burk double reciprocal plots.


  
Table: Effect of NADPH concentration on the cyclic NADH AcPyAD transhydrogenation catalyzed by the purified wild-type and mutant enzymes

The apparent S (NADPH concentration required for half-maximal activity) and V values were derived from Lineweaver-Burk double reciprocal plots.



FOOTNOTES

*
This work was supported by United States Public Health Service Grant GM24887. Computer facilities were supported by United States Public Health Service Grant M01 RR0033 for the General Clinical Research Center. Synthesis of oligonucleotides was supported in part by the Sam and Rose Stein Charitable Trust. This is Publication 9137-MEM from The Scripps Research Institute, La Jolla, CA. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
To whom correspondence and reprint requests should be addressed. Tel.: 619-554-8092; Fax: 619-554-6838.

The abbreviations used are: FSBA, 5`-p-fluorosulfonylbenzoyladenosine; AcPyAD, 3-acetylpyridine adenine dinucleotide; AcPyADP, 3-acetylpyridine adenine dinucleotide phosphate; MES, 2-(N-morpholino)ethanesulfonic acid; Tricine, N-[2-hydroxy-1,1-bis(hydroxymethyl)-ethyl]glycine.

Following are the degrees of sequence identity in the 5 sequenced transhydrogenases for: 100 residues around the NAD(H) binding site, 26%; 100 residues around the NADP(H) binding site, 45%; the 19-residue-long hydrophobic segment, 58%; and the 58-residue-long hydrophobic segment, 65.5%.

The identical purification procedures of the wild-type and the mutant enzymes suggest that no major structural alteration had taken place as a result of D213N and D213I mutations.


ACKNOWLEDGEMENTS

We thank Drs. Takao Yagi and Akemi Matsuno-Yagi for fruitful discussions.


REFERENCES
  1. Ahmad, S., Glavas, N. A., and Bragg, P. D. (1992) Eur. J. Biochem.207, 733-739 [Abstract]
  2. Chen, E. Y., and Seeburg, P. H. (1985) DNA (NY)4, 165-170 [Medline] [Order article via Infotrieve]
  3. Clarke, D. M., and Bragg, P. D. (1985) J. Bacteriol.162, 367-373 [Medline] [Order article via Infotrieve]
  4. Clarke, D. M., Loo, T. W., Gillam, S., and Bragg, P. D. (1986) Eur. J. Biochem.158, 647-653 [Abstract]
  5. Fillingame, R. H. (1990) in The Bacteria (Krulwich, T. A., ed) Vol. XII, pp. 345-391, Academic Press, New York
  6. Fisher, R. R., and Earle, S. R. (1982) in The Pyridine Nucleotide Coenzymes (Everse, J., Anderson, B., and You, K.-S., eds) pp. 279-324, Academic Press, New York
  7. Glavas, N. A. (1994) Structural and Functional Studies of the Pyridine Nucleotide Transhydrogenase of the Escherichia coli, Ph.D. dissertation, University of British Columbia
  8. Hamilton, C. M., Aldea, M., Washburn, B. K., Babitzke, P., and Kushner, S. R. (1989) J. Bacteriol.171, 4617-4622 [Medline] [Order article via Infotrieve]
  9. Hatefi, Y., and Yamaguchi, M. (1992) in Molecular Mechanisms in Bioenergetics (Ernster, L., ed) pp. 265-281, Elsevier Science Publishers B.V., Amsterdam
  10. Hatefi, Y., Phelps, D. C., and Galante, Y. M. (1980) J. Biol. Chem.255, 9526-9529 [Abstract/Free Full Text]
  11. Holmberg, E., Olausson, T., Hultman, T., Rydström, J., Ahmad, S., Glavas, N. A., and Bragg, P. D. (1994) Biochemistry33, 7691-7700 [Medline] [Order article via Infotrieve]
  12. Hou, C., Potier, M., and Bragg, P. D. (1990) Biochim. Biophys. Acta1018, 61-66 [Medline] [Order article via Infotrieve]
  13. Howlett, G. J., Yeh, E., and Schachman, H. K. (1978) Arch. Biochem. Biophys.190, 809-819
  14. Hutton, M., Day, J. M., Bizouarn, T., and Jackson, J. B. (1994) Eur. J. Biochem.219, 1041-1051 [Abstract]
  15. Jencks, W. P. (1975) Adv. Enzymol. Relat. Areas Mol. Biol.43, 219-410 [Medline] [Order article via Infotrieve]
  16. Jencks, W. P. (1989) Methods Enzymol.171, 145-164 [Medline] [Order article via Infotrieve]
  17. Kramer, R. A., Tomchak, L. A., McAndrew, S. J., Becker, K., Hug, D., Pasamontes, L., and Hümbelin, M. (1993) Mol. Biochem. Parasitol.60, 327-332 [CrossRef][Medline] [Order article via Infotrieve]
  18. Kunkel, T. A., Roberts, J. D., and Zakour, R. A. (1987) Methods Enzymol.154, 367-382 [Medline] [Order article via Infotrieve]
  19. Laemmli, U. K. (1970) Nature227, 680-685 [Medline] [Order article via Infotrieve]
  20. Lee, C.-P., and Ernster, L. (1989) Biochim. Biophys. Acta1000, 371-376 [Medline] [Order article via Infotrieve]
  21. Lever, T. M., Palmer, T., Cunningham, I. J., Cotton, N. P. J., and Jackson, J. B. (1991) Eur. J. Biochem.197, 247-255 [Abstract]
  22. Peterson, G. L. (1977) Anal. Biochem.83, 346-356 [Medline] [Order article via Infotrieve]
  23. Phelps, D. C., and Hatefi, Y. (1984) Biochemistry23, 6340-6344 [Medline] [Order article via Infotrieve]
  24. Phelps, D. C., and Hatefi, Y. (1985) Biochemistry24, 3503-3507 [Medline] [Order article via Infotrieve]
  25. Phelps, D. C., Galante, Y. M., and Hatefi, Y. (1980) J. Biol. Chem.255, 9647-9652 [Abstract/Free Full Text]
  26. Rothschild, K. J. (1992) J. Bioenerg. Biomembr.24, 147-167 [Medline] [Order article via Infotrieve]
  27. Rydström, J., Persson, B., and Carlenor, E. (1987) in Pyridine Nucleotide Coenzymes: Chemical, Biochemical and Medical Aspects (Dolphin, D., Poulson, R., and Avramovic, O., eds) pp. 433-460, Academic Press, New York
  28. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989) in Molecular Cloning: a Laboratory Manual, 2nd Ed, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY
  29. Segel, I. H. (1975) in Enzyme Kinetics: Behavior and Analysis of Rapid Equilibrium and Steady-State Enzyme Systems, pp. 218-220, John Wiley and Sons, New York
  30. Sonar, S., Marti, T., Rath, P., Fischer, W., Coleman, M., Nilsson, A., Khorana, H. G., and Rothschild, K. J. (1994) J. Biol. Chem.269, 28851-28858 [Abstract/Free Full Text]
  31. Sone, N., Yoshida, M., Hirata, H., and Kagawa, Y. (1977) J. Biochem. (Tokyo)81, 519-528 [Abstract]
  32. Southern, E. M. (1975) J. Mol. Biol.98, 503-517 [Medline] [Order article via Infotrieve]
  33. Vermeulen, A. N., Kok, J. J., Van den Boogaart, P., Dijkema, R., and Claessens, J. A. J. (1993) FEMS Microbiol. Lett.110, 223-230 [Medline] [Order article via Infotrieve]
  34. Wakabayashi, S., and Hatefi, Y. (1987a) Biochem. Int.15, 915-924 [Medline] [Order article via Infotrieve]
  35. Wakabayashi, S., and Hatefi, Y. (1987b) Biochem. Int.15, 667-675 [Medline] [Order article via Infotrieve]
  36. Wang, T. P., Kaplan, N. O., and Stolzenbach, F. E. (1954) J. Biol. Chem.211, 465-472 [Free Full Text]
  37. Williams, R., Cotton, N. P. J., Thomas, C. M., and Jackson, J. B. (1994) Microbiology140, 1595-1604 [Abstract]
  38. Wu, L. N. Y., Earle, S. R., and Fisher, R. R. (1981) J. Biol. Chem.256, 7401-7408 [Abstract/Free Full Text]
  39. Yamaguchi, M., and Hatefi, Y. (1989) Biochemistry28, 6050-6056 [Medline] [Order article via Infotrieve]
  40. Yamaguchi, M., and Hatefi, Y. (1991) J. Biol. Chem.266, 5728-5735 [Abstract/Free Full Text]
  41. Yamaguchi, M., and Hatefi, Y. (1993) J. Biol. Chem.268, 17871-17877 [Abstract/Free Full Text]
  42. Yamaguchi, M., and Hatefi, Y. (1994) J. Bioenerg. Biomembr.26, 435-446 [Medline] [Order article via Infotrieve]
  43. Yamaguchi, M., Hatefi, Y., Trach, K., and Hoch, J. A. (1988) J. Biol. Chem.263, 2761-2767 [Abstract/Free Full Text]
  44. Yamaguchi, M., Wakabayashi, S., and Hatefi, Y. (1990) Biochemistry29, 4136-4143 [Medline] [Order article via Infotrieve]
  45. Yu, Y., and Samuelson, J. (1994) Mol. Biochem. Parasitol.68, 323-328 [Medline] [Order article via Infotrieve]

©1995 by The American Society for Biochemistry and Molecular Biology, Inc.