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The Substitution of a Single Amino Acid Residue (Ser-116 → Asp) Alters NADP-containing Glucose-Fructose Oxidoreductase ofZymomonas mobilis into a Glucose Dehydrogenase with Dual Coenzyme Specificity

1997; Elsevier BV; Volume: 272; Issue: 20 Linguagem: Inglês

10.1074/jbc.272.20.13126

ISSN

1083-351X

Autores

Thomas Wiegert, Hermann Sahm, Georg A. Sprenger,

Tópico(s)

Biochemical and Molecular Research

Resumo

Glucose-fructose oxidoreductase (GFOR, EC1.1.1.99.-) from the Gram-negative bacterium Zymomonas mobilis contains the tightly bound cofactor NADP. Based on the revision of the gfo DNA sequence, the derived GFOR sequence was aligned with enzymes catalyzing reactions with similar substrates. A novel consensus motif (AGKHVXCEKP) for a class of dehydrogenases was detected. From secondary structure analysis the serine-116 residue of GFOR was predicted as part of a Rossmann-type dinucleotide binding fold. An engineered mutant protein (S116D) was purified and shown to have lost tight cofactor binding based on (a) altered tryptophan fluorescence; (b) lack of NADP liberation through perchloric acid treatment of the protein; and (c) lack of GFOR enzyme activity. The S116D mutant showed glucose dehydrogenase activity (3.6 ± 0.1 units/mg of protein) with both NADP and NAD as coenzymes (K mfor NADP, 153 ± 9 μm; for NAD, 375 ± 32 μm). The single site mutation therefore altered GFOR, which in the wild-type situation contains NADP as nondissociable redox cofactor reacting in a ping-pong type mechanism, to a dehydrogenase with dissociable NAD(P) as cosubstrate and a sequential reaction type. After prolonged preincubation of the S116D mutant protein with excess NADP (but not NAD), GFOR activity could be restored to 70 units/mg, one-third of wild-type activity, whereas glucose dehydrogenase activity decreased sharply. A second site mutant (S116D/K121A/K123Q/I124K) showed no GFOR activity even after preincubation with NADP, but it retained glucose dehydrogenase activity (4.2 ± 0.2 units/mg of protein). Glucose-fructose oxidoreductase (GFOR, EC1.1.1.99.-) from the Gram-negative bacterium Zymomonas mobilis contains the tightly bound cofactor NADP. Based on the revision of the gfo DNA sequence, the derived GFOR sequence was aligned with enzymes catalyzing reactions with similar substrates. A novel consensus motif (AGKHVXCEKP) for a class of dehydrogenases was detected. From secondary structure analysis the serine-116 residue of GFOR was predicted as part of a Rossmann-type dinucleotide binding fold. An engineered mutant protein (S116D) was purified and shown to have lost tight cofactor binding based on (a) altered tryptophan fluorescence; (b) lack of NADP liberation through perchloric acid treatment of the protein; and (c) lack of GFOR enzyme activity. The S116D mutant showed glucose dehydrogenase activity (3.6 ± 0.1 units/mg of protein) with both NADP and NAD as coenzymes (K mfor NADP, 153 ± 9 μm; for NAD, 375 ± 32 μm). The single site mutation therefore altered GFOR, which in the wild-type situation contains NADP as nondissociable redox cofactor reacting in a ping-pong type mechanism, to a dehydrogenase with dissociable NAD(P) as cosubstrate and a sequential reaction type. After prolonged preincubation of the S116D mutant protein with excess NADP (but not NAD), GFOR activity could be restored to 70 units/mg, one-third of wild-type activity, whereas glucose dehydrogenase activity decreased sharply. A second site mutant (S116D/K121A/K123Q/I124K) showed no GFOR activity even after preincubation with NADP, but it retained glucose dehydrogenase activity (4.2 ± 0.2 units/mg of protein). Glucose-fructose oxidoreductase (GFOR 1The abbreviations used are: GFOR, glucose-fructose oxidoreductase; kb, kilobase; MES, 4-morpholineethanesulfonic acid; HPLC, high performance liquid chromatography. ; EC 1.1.1.99.-) is a homotetrameric enzyme from the Gram-negative bacterium Zymomonas mobiliswhich catalyzes the oxidation of glucose to gluconolactone and the reduction of fructose to sorbitol (Fig. 1) in a ping-pong type mechanism (1Zachariou M. Scopes R.K. J. Bacteriol. 1986; 167: 863-869Crossref PubMed Google Scholar, 2Hardman M.J. Scopes R.K. Eur. J. Biochem. 1988; 179: 203-209Crossref Scopus (50) Google Scholar). Sorbitol is used as a compatible solute by Z. mobilis to counteract the detrimental osmotic effects of high concentrations of sugars in the medium (3Loos H. Krämer R. Sahm H. Sprenger G.A. J. Bacteriol. 1994; 176: 7688-7693Crossref PubMed Google Scholar). The apparent physiological role of GFOR is the production of sorbitol from the two sugar moieties of sucrose, a natural carbon source of the bacterium which dwells in sugar-rich habitats (4Swings J. DeLey J. Bacteriol. Rev. 1977; 41: 1-46Crossref PubMed Google Scholar). The enzyme is synthesized as a precursor (pre-GFOR) with an NH2-terminal signal peptide of 52 amino acid residues (5Kanagasundaram V. Scopes R.K. J. Bacteriol. 1992; 174: 1439-1447Crossref PubMed Google Scholar) and is exported to the periplasm, at least partially, via the Sec pathway (6Wiegert T. Sahm H. Sprenger G.A. Arch. Microbiol. 1996; 166: 32-41Crossref PubMed Scopus (25) Google Scholar). The mature enzyme is located in the periplasm (7Loos H. Völler M. Rehr B. Stierhof Y.-D. Sahm H. Sprenger G.A. FEMS Microbiol. Lett. 1991; 84: 211-216Crossref Scopus (30) Google Scholar, 8Aldrich H.C. McDowell L. Barbosa M.de F.S. Yomano L.P. Scopes R.K. Ingram L.O. J. Bacteriol. 1992; 174: 4504-4508Crossref PubMed Google Scholar). Stoichiometrically, one molecule of the cofactor NADP is bound per monomer (1Zachariou M. Scopes R.K. J. Bacteriol. 1986; 167: 863-869Crossref PubMed Google Scholar). Pre-GFOR was shown to be enzymatically fully active and to bind NADP tightly (9Loos H. Sahm H. Sprenger G.A. FEMS Microbiol. Lett. 1993; 107: 293-298Crossref PubMed Scopus (26) Google Scholar). A deletion of 15 mainly hydrophobic amino acid residues (Δ32–46) in the signal peptide led to a cytosolic form of GFOR which could be purified and showed characteristics similar to the wild-type enzyme (6Wiegert T. Sahm H. Sprenger G.A. Arch. Microbiol. 1996; 166: 32-41Crossref PubMed Scopus (25) Google Scholar) in terms of reactivity and the mode of cofactor binding. A special feature of GFOR is the tight binding of the cofactor NADP(H). Treatment with perchloric acid removed the cofactor from the apoprotein (1Zachariou M. Scopes R.K. J. Bacteriol. 1986; 167: 863-869Crossref PubMed Google Scholar), suggesting that NADP(H) is bound in a noncovalent manner. During protein purification, the cofactor is not removed from the enzyme (1Zachariou M. Scopes R.K. J. Bacteriol. 1986; 167: 863-869Crossref PubMed Google Scholar,6Wiegert T. Sahm H. Sprenger G.A. Arch. Microbiol. 1996; 166: 32-41Crossref PubMed Scopus (25) Google Scholar). The published DNA sequence and the amino acid sequence derived thereof reveal that GFOR has little similarity to other enzymes (5Kanagasundaram V. Scopes R.K. J. Bacteriol. 1992; 174: 1439-1447Crossref PubMed Google Scholar). This, together with the mode of tight cofactor binding, suggests a novel dinucleotide binding mode. Although diffracting crystals have been obtained from purified GFOR (10Loos H. Ermler U. Sprenger G.A. Sahm H. Protein Sci. 1994; 3: 2447-2449Crossref PubMed Scopus (9) Google Scholar), the three-dimensional structure of the enzyme had not been available while the present investigations were performed. Using site-directed mutagenesis we wanted to analyze the tight binding of cofactor to GFOR. Based on alignments and secondary structure predictions, we mutated amino acid residues that are likely to be involved in cofactor binding. As a striking result, a mutant GFOR with a single amino acid substitution (S116D) behaved as glucose dehydrogenase with dual coenzyme specificity for NADP and NAD. Z. mobilis strain ACM3963 (11Kirk L.A. Doelle H.W. Biotechnol. Lett. 1993; 15: 985-990Crossref Scopus (9) Google Scholar) and its recombinant derivatives were maintained and cultivated anaerobically as described previously (6Wiegert T. Sahm H. Sprenger G.A. Arch. Microbiol. 1996; 166: 32-41Crossref PubMed Scopus (25) Google Scholar). For protein purifications, cells were grown in complex medium with 10% glucose with isopropyl-1-thio-β-d-galactopyranoside (1 mm) to induce gfo expression from derivatives of plasmid pZY507 (CmR; Escherichia coli/Z.mobilis shuttle vector withlacI q/Ptac control; 6). E. coli cells were grown in LB medium with appropriate antibiotics (12Sambrook J. Fritsch E.F. Maniatis T. Molecular Cloning: A Laboratory Manual. 2nd Ed. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY1989Google Scholar). Construction of plasmids pZY470, pZY470Δ32–46, and pZY570 is described elsewhere (6Wiegert T. Sahm H. Sprenger G.A. Arch. Microbiol. 1996; 166: 32-41Crossref PubMed Scopus (25) Google Scholar). All manipulations of DNA, cloning, and transformation were done according to standard procedures (12Sambrook J. Fritsch E.F. Maniatis T. Molecular Cloning: A Laboratory Manual. 2nd Ed. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY1989Google Scholar, 13Hanahan D. J. Mol. Biol. 1983; 166: 557-580Crossref PubMed Scopus (8190) Google Scholar). DNA sequencing was performed by the chain termination method using dideoxynucleotides (14Sanger F. Nicklen S. Coulson A.R. Proc. Natl. Acad. Sci. U. S. A. 1977; 74: 5463-5467Crossref PubMed Scopus (52671) Google Scholar) in conjunction with T7 DNA polymerase (15Tabor S. Richardson C.C. Proc. Natl. Acad. Sci. U. S. A. 1987; 84: 4767-4771Crossref PubMed Scopus (1687) Google Scholar) using the nonradioactive A.L.F. fluorescence detection method according to the manufacturer's protocols (Pharmacia LKB, Freiburg, Germany). Site-directed mutagenesis was carried out on a derivative of M13mp18 (16Yanisch-Perron C. Vieira J. Messing J. Gene ( Amst. ). 1985; 33: 103-119Crossref PubMed Scopus (11465) Google Scholar) containing an internal 0.3 kb PstI/SphI fragment of the gfo gene according to a standard procedure (17Kunkel T.A. Proc. Natl. Acad. Sci. U. S. A. 1985; 82: 488-492Crossref PubMed Scopus (4900) Google Scholar) with oligonucleotidesI, 5′-GCTTTTTCAGCGTTACCATCGACCAAAGCTTCGAT-3′ (S116D); II, 5′-GACGCCATATTCAGCGGCAACTTTCTGAGCAGCTTCAGCGTTACCATCGACG-3′ (S116D/K121A/K123Q/I124K); and III, 5′-TAAAATCTGGTTAAGACCATATTTACCCAGACC-3′ (A95G); altered base triplets are underlined. In a second round of mutagenesis, an M13 clone with the exchange A95G was used as template together with oligonucleotides I and II to combine these mutations. Base exchanges were checked by DNA sequencing. After verification, the respective fragments were cloned into plasmid pZY470Δ32–46, sequenced again, introduced into the shuttle vector pZY507, and conjugated to Z. mobilis ACM3963 as described elsewhere (6Wiegert T. Sahm H. Sprenger G.A. Arch. Microbiol. 1996; 166: 32-41Crossref PubMed Scopus (25) Google Scholar). To avoid any problems with the export of recombinant GFOR proteins to the periplasm or related problems as stability and cofactor incorporation (6Wiegert T. Sahm H. Sprenger G.A. Arch. Microbiol. 1996; 166: 32-41Crossref PubMed Scopus (25) Google Scholar), respective mutations were introduced into a gfo allele that encodes a cytoplasmic form (GFORΔ32–46). This form had been shown earlier to bind NADP in the same tight manner as wild-type GFOR and was enzymatically fully active (6Wiegert T. Sahm H. Sprenger G.A. Arch. Microbiol. 1996; 166: 32-41Crossref PubMed Scopus (25) Google Scholar). Deletion mutagenesis was performed using a megaprimer method with two steps of polymerase chain reaction (18Ogel Z.B. McPherson M.J. Protein Eng. 1992; 5: 467-468Crossref PubMed Scopus (11) Google Scholar). In the first round of polymerase chain reaction, plasmid pZY470 DNA (6Wiegert T. Sahm H. Sprenger G.A. Arch. Microbiol. 1996; 166: 32-41Crossref PubMed Scopus (25) Google Scholar) served as a template with the loop-out primer IV 5′- ACGATCTTCCGGCATCGGGCGGATCATΔAATCCTTGTTTCTTTCTTAAC-3′ (Δ2–74) and the M13/pUC universe sequencing primer;Δ denotes the site of deletion. The resulting 222-base pair megaprimer was incubated with the M13/pUC universal and reverse sequencing primers (Boehringer Mannheim, Germany) together with a 1.5-kb PvuII fragment of pZY470 as template in the second polymerase chain reaction step. The 1.5-kb product was restricted withEcoRI plus HindIII, and the 0.3-kb fragment was ligated to plasmid pZY470 restricted withEcoRI/HindIII. Clones were checked for the desired deletion by restriction analysis and DNA sequencing. The resulting plasmid (pZY470Δ2–74) was restricted withEcoRI/HindIII, and the 0.3-kb fragment was cloned into pZY470/S116D or pZY470/S116D/K121A/K123Q/I124K to combine Δ2–74 with these amino acid substitutions. All gfo alleles were conjugated to Z. mobilis ACM3963 on plasmids derived from pZY507 as described elsewhere (6Wiegert T. Sahm H. Sprenger G.A. Arch. Microbiol. 1996; 166: 32-41Crossref PubMed Scopus (25) Google Scholar). Wild-type and mutant proteins (GFORΔ32–46, GFORΔ2–74, or derivatives) were purified from derivatives of the GFOR-defective Z. mobilis strain ACM3963 (11Kirk L.A. Doelle H.W. Biotechnol. Lett. 1993; 15: 985-990Crossref Scopus (9) Google Scholar) after growth in complex medium supplemented with isopropyl-1-thio-β-d-galactopyranoside (1 mm). GFOR was purified using a coupled anion-cation exchange chromatography as described elsewhere (6Wiegert T. Sahm H. Sprenger G.A. Arch. Microbiol. 1996; 166: 32-41Crossref PubMed Scopus (25) Google Scholar). A second cation exchange step was omitted, as apparent purity was already achieved in the first step (Fig. 2). As degradation was observed with some mutant proteins, NADP (0.5 mm) was added to all buffers during purification steps. Because of the lack of detectable GFOR activity in most of the mutant proteins, GFOR-containing fractions were identified by SDS-polyacrylamide gel electrophoresis and subsequent Western blot analysis (19Towbin H.T. Staehelin T. Gordon J. Proc. Natl. Acad. Sci. U. S. A. 1979; 76: 4350-4354Crossref PubMed Scopus (44923) Google Scholar). Fractions were pooled, equilibrated to 40 mm K-MES, 1 mmdithiothreitol, pH 6.4, by ultrafiltration, mixed with the same volume of glycerol (88%), and stored at −20 °C until further use. NH2-terminal peptide sequencing was performed by limited Edman degradation in conjunction with an Applied Biosystems Inc. 371 sequencer. Determination of the COOH-terminal amino acid was done with carboxypeptidase Y (Boehringer Mannheim) according to the manufacturer's protocol followed by reversed phase HPLC as described elsewhere (20Lindsroth P. Mopper K. Anal. Chem. 1979; 51: 1167-1174Crossref Google Scholar). Tightly bound NADP(H) was released from GFOR by acid denaturation and was subsequently detected by HPLC analysis. The enzymes were equilibrated to sodium phosphate buffer (200 mm, pH 5.0) by ultrafiltration and diluted to a final concentration of approximately 4 mg protein/ml. 60 μl of ice-cold perchloric acid (35%) was added to 60 μl of the enzyme solution, mixed, and kept on ice for 15 min. The samples were neutralized by the stepwise addition of 180 μl of KHCO3(2 m), and insoluble material was spun down by centrifugation. An aliquot of the supernatant was submitted to HPLC on an octadecyl-silica gel column (Hypersil ODS, 5 μm, CS Chromatography Service, Langerwehe, Germany) that was eluted with a gradient of sodium phosphate (200 mm, pH 5.0), acetonitrile at a flow rate of 0.3 ml/min at 40 °C. Retention times of NADP(H) and NAD(H) were determined with standard solutions. Enzyme activities were determined at 30 °C in a thermostatted cuvette holder of a Shimadzu UV160 spectrophotometer by the measurement of acidification (formation of gluconic acid from glucose) according to a published method (1Zachariou M. Scopes R.K. J. Bacteriol. 1986; 167: 863-869Crossref PubMed Google Scholar). Gluconolactonase from Rhodotorula rubra was added in excess to the reaction mixture (7Loos H. Völler M. Rehr B. Stierhof Y.-D. Sahm H. Sprenger G.A. FEMS Microbiol. Lett. 1991; 84: 211-216Crossref Scopus (30) Google Scholar). Glucose dehydrogenase activity was measured by the increase of reduced NAD(P) at 340 nm. Reaction mixtures contained 5 μg/ml purified protein (GFOR or mutant protein), 5 units/ml gluconolactonase, glucose (400 mm in 40 mm K-MES buffer, pH 6.4), and 1 mm NAD(P). To determine K m for NAD(P), concentrations of coenzymes were varied from 2 to 1,000 μm. K m,k cat, and standard deviations thereof were calculated by the Enzfitter Program (Elsevier Biosoft, version 1.05). Protein was determined by a dye binding method (21Bradford M.M. Anal. Biochem. 1976; 72: 248-254Crossref PubMed Scopus (216428) Google Scholar). Fluorescence spectroscopy was performed with an Aminco-Bowman Series 2 Spectrometer at 20 °C. Prior to use, the enzyme solutions were equilibrated by ultrafiltration to sodium phosphate buffer (50 mm, pH 6.4). Excitation and emission slits were set to 4 nm. To minimize photodecomposition of the enzymes, the shutter of the exciting beam was kept closed until the measurement started. Fluorescence titrations were performed by the stepwise addition of 2.5–5 μl of NADPH to 2 ml of an enzyme solution with a concentration of 0.6 μm (tetramer). Dilution by the addition of NADPH was kept to a maximum of 2.5%. Controls were obtained following the same procedure without added enzyme. To minimize inner filter effects of nucleotide fluorescence, the excitation wavelength was set to 360 nm (22Basso L.A. Engel P.C. Walmsley A.R. Eur. J. Biochem. 1995; 234: 603-615Crossref PubMed Scopus (13) Google Scholar). Titration curves were fitted to a quadratic equation relating the fluorescence change to the coenzyme concentration for a second order binding process (23Birdsall B. King R.W. Wheeler M.R. Lewis C.A. Goode S.R. Dunlap B.D. Roberts G.C.K. Anal. Biochem. 1983; 132: 353-361Crossref PubMed Scopus (212) Google Scholar). (F−FL)(F∞−FL)−1=12[E]0Kd+[E]0+[S]0±(Kd+[E]0+[S]0)2−4[E]0[S]0Equation 1 where [E]0 and [S]0represent the initial enzyme and NADPH concentrations, respectively;F ∞ is the fluorescence intensity at saturating NADPH concentration; F L is the fluorescence intensity at a given NADPH concentration without enzyme; andF is the fluorescence intensity at a given NADPH concentration with enzyme. During former rounds of subcloning and site-directed mutagenesis (6Wiegert T. Sahm H. Sprenger G.A. Arch. Microbiol. 1996; 166: 32-41Crossref PubMed Scopus (25) Google Scholar) we had already encountered several deviations from the published DNA sequence of the gfo gene (5Kanagasundaram V. Scopes R.K. J. Bacteriol. 1992; 174: 1439-1447Crossref PubMed Google Scholar). This prompted us to sequence the complete gfo gene again. In a comparison with the former sequence (5Kanagasundaram V. Scopes R.K. J. Bacteriol. 1992; 174: 1439-1447Crossref PubMed Google Scholar), several frameshifts were observed which resulted in a deviating amino acid sequence of GFOR comprising only 433 residues (instead of 439). According to our DNA sequencing, the COOH-terminal residue should be a tyrosine. We subjected purified GFOR to a carboxypeptidase Y treatment and found that, indeed, tyrosine appeared as the first residue (data not shown). Using the corrected gfo sequence and the derived amino acid sequence, we performed similarity searches with the HUSAR package provided by the European Molecular Biology Laboratory (EMBL, Heidelberg) in all accessible data bases using the BLAST data base search program. Several amino acid sequences, in some cases open reading frames with putative enzyme functions, were found which showed significant similarities to the NH2-terminal half of GFOR (Fig. 3); similar findings, using the uncorrected GFOR sequence, have been reported by others (24Rossbach S. Kulpa D.A. Rossbach U. DeBruijn F.J. Mol. & Gen. Genet. 1994; 245: 11-24Crossref PubMed Scopus (56) Google Scholar). All of these proteins are NAD(P)-dependent dehydrogenases displaying a possible fingerprint motif of a classical Rossmann fold (25Wierenga R.K. Terpstra P. Hol W.G.J. J. Mol. Biol. 1986; 187: 101-107Crossref PubMed Scopus (996) Google Scholar) at their immediate NH2 termini. In the GFOR sequence, a possible fingerprint motif could also be recognized, although it was preceded by the signal sequence of 52 amino acid residues and a proline-rich sequence of approximately 30 amino acid residues (Fig. 3). GFOR displays the characteristic fingerprint of a NADP binding Rossmann fold,i.e. the sequence Gly-X-Gly-X-X-Ala with alanine at position 95 and the absence of a negatively charged amino acid residue (Asp or Glu), which is typically found as the last conserved residue of the fingerprint sequence in NAD-binding βαβ folds (25Wierenga R.K. Terpstra P. Hol W.G.J. J. Mol. Biol. 1986; 187: 101-107Crossref PubMed Scopus (996) Google Scholar, 26Hanukoglu I. Gutfinger T. Eur. J. Biochem. 1989; 180: 479-484Crossref PubMed Scopus (173) Google Scholar). From the sequence alignment in Fig. 3, a highly conserved motif with the consensus AGKHVXCEKP became apparent; this box is found around amino acid residues 170–185 of GFOR. A data base search for this motif returned exclusively the sequences listed in Fig. 3. All of the listed enzymes, with the exception of biliverdin reductase, are known, or can be expected, to react with substrates that are structurally similar to glucose. Therefore this motif may constitute a putative fingerprint for a novel class of sugar dehydrogenases. A secondary structure prediction of GFOR was performed using the PHD program (27Rost B. Sander C. J. Mol. Biol. 1993; 232: 584-599Crossref PubMed Scopus (2647) Google Scholar), based on the multiple alignment shown in Fig. 3. The amino-terminal half of GFOR, according to this prediction, consists of six β-folds interspaced by α-helical elements, resembling the structure of Rossmann-type NAD(P) binding sites (28Rossmann M.G. Liljas A. Brändén C.-I. Banaszak L.J. Boyer P.D. The Enzymes. 3rd Ed. 11. Academic Press, New York1975: 61-102Google Scholar). Taken together with the possible fingerprint motif for NAD(P) binding sites, it could be predicted that GFOR binds its cofactor NADP in a domain comprising the NH2-terminal half (approximately amino acid residues 80–250) in a βαβ dinucleotide binding fold, resembling the Rossmann fold of dehydrogenases. Only a few amino acid residues are highly conserved in Rossmann folds. These are 10–11 amino acid residues, termed the fingerprint sequence of βαβ dinucleotide binding folds, in the region of the first and second β-sheet (βa and βb) and the interspacing pyrophosphate binding α-helix (αb). From sequence and structural data and from mutational analyses, it has been established that the fingerprint regions of binding sites for NAD or NADP differ to some extent and that these differences play a key role in determining the coenzyme specificity (29Scrutton N.S. Berry A. Perham R.N. Nature. 1990; 343: 38-43Crossref PubMed Scopus (646) Google Scholar, 30Baker P.J. Britton K.L. Rice D.W. Rob A. Stillman T.J. J. Mol. Biol. 1992; 228: 662-671Crossref PubMed Scopus (156) Google Scholar, 31Levy R.L. Vought V.E. Yin X. Adams M.J. Arch. Biochem. Biophys. 1996; 326: 145-151Crossref PubMed Scopus (36) Google Scholar). To analyze the mode of NADP binding in GFOR and to assess the involvement of a possible Rossmann fold, we intended to weaken the interaction of GFOR with its cofactor NADP by engineering an NAD binding motif using site-directed mutagenesis. The conserved Ala residue in NADP binding sites is known to induce a hydrogen bond pattern that differs from that of NAD binding sites, where Gly occupies this position (30Baker P.J. Britton K.L. Rice D.W. Rob A. Stillman T.J. J. Mol. Biol. 1992; 228: 662-671Crossref PubMed Scopus (156) Google Scholar, 32Mittl P.R.E. Berry A. Scrutton N.S. Perham R.N. Schulz G.E. Protein Sci. 1994; 3: 1504-1514Crossref PubMed Scopus (48) Google Scholar). To assay this for GFOR, we changed Ala-95 of GFOR to Gly (A95G; mutant A, Fig.4). Negatively charged amino acid residues (Glu or Asp) are invariably found at the end of the second β-sheet of NAD binding sites. The oxygen atoms of the side chain carboxyl group form hydrogen bonds to the 2′- and 3′-OH groups of the adenine ribose moiety of NAD. In contrast, NADP binding sites usually contain an uncharged amino acid residue at this position, which is followed immediately by a positively charged residue in many cases (29Scrutton N.S. Berry A. Perham R.N. Nature. 1990; 343: 38-43Crossref PubMed Scopus (646) Google Scholar, 33Adams M.J. Ellis G.H. Gover S. Naylor C.E. Phillips C. Structure. 1994; 2: 385-393Abstract Full Text Full Text PDF PubMed Scopus (110) Google Scholar). Our secondary structure predictions suggested that Ser-116 at the end of the putative βB in GFOR might be replaced by Asp (S116D; mutant B) to lower the affinity for NADP and to combine mutations A and B (A95G/S116D; mutant C; Fig.4), to complete disruption via the H bonding pattern. Site-directed mutageneses were performed as described under "Materials and Methods." The resulting mutant alleles were introduced and expressed in the GFOR-defective strain Z. mobilis ACM3963 (6Wiegert T. Sahm H. Sprenger G.A. Arch. Microbiol. 1996; 166: 32-41Crossref PubMed Scopus (25) Google Scholar, 11Kirk L.A. Doelle H.W. Biotechnol. Lett. 1993; 15: 985-990Crossref Scopus (9) Google Scholar). These and additional mutant proteins (see below) are listed in Fig. 4. Mutant proteins were purified to apparent homogeneity as judged by SDS-polyacrylamide gel electrophoresis (Fig.2). Only mutant A (A95G) showed GFOR activity comparable to the GFORΔ32–46 wild-type enzyme (Table I). No GFOR activity could be detected by the standard photometric GFOR enzyme assay (without addition of NADP) with mutants B (S116D) and C (A95G/S116D). The loss of GFOR activity could be due to the loss of the cofactor NADP(H). Proteins, therefore, were analyzed by fluorescence spectroscopy. Fluorescence at 450 nm is a sensitive proof of the presence of reduced NAD(P) (34Roskoski R. in Pyridine Nucleotide Coenzymes.in: Dolphin D. Poulson R. Avramovic O. Part B. John Wiley and Sons, New York1987: 173-188Google Scholar). In addition, shifts in tryptophan fluorescence at 330 nm may reveal conformational differences between wild-type and mutant GFOR (35Schmid F.X. Creighton T.E. Protein Structure: A Practical Approach. IRL Press, Oxford, U. K.1989: 251-285Google Scholar). GFORΔ32–46, as the control with tightly bound NADP, was preincubated with glucose to reduce bound NADP. When fluoresence was excited at 295 nm, only weak tryptophan fluorescence was observed, with a distinct peak of NADPH fluorescence at 450 nm. Fluorescence spectra with the mutant enzyme B (GFOR-deficient) at the same conditions showed clearly enhanced tryptophan fluorescence compared with the wild-type enzyme, with the same emission maximum at 330 nm (Fig. 5). However, no NADPH fluorescence at 450 nm could be measured. Mutant A showed the same fluorescence emission spectra as the wild-type GFORΔ32–46, and mutant C behaved similarly to mutant B (data not shown).Table IGlucose-fructose oxidoreductase and glucose dehydrogenase activitiesGFOR activityGlcDH activity−NADP+NADPμmol/min/mgμmol/min/mgGFORΔ32–46144 ± 7136 ± 10NDMutant A154 ± 4159 ± 3NDMutant BND43 ± 23.6 ± 0.1Mutant CND41 ± 13.1 ± 0.3Mutant DNDND4.2 ± 0.2Mutant ENDND4.4 ± 0.1GFORΔ2–74NDNDNDMutant FNDND2.8 ± 0.1Mutant GNDND3.6 ± 0.1Specific GFOR and glucose dehydrogenase (GlcDH) activities of GFORΔ32–46, GFORΔ2–74, and mutants thereof (compare with Fig. 4) are shown. Mean values and deviations were obtained from two independently performed assays. ND, not detectable (ΔE/min for the GFOR and glucose dehydrogenase test with 5 μg/ml enzyme <0.01/min). GFOR activities were determined with the standard photometric test without supplemented NADP (−NADP). In addition, GFOR activities were determined after preincubation of 5 mg/ml each of GFORΔ32–46, GFORΔ2–74, or mutants thereof with 20 mmNADP in 40 mm K-MES, pH 6.4, for 5 min, at 25 °C (+NADP). The final concentration of enzyme was 5 μg/ml and 20 μm NADP in the assay mix. Open table in a new tab Specific GFOR and glucose dehydrogenase (GlcDH) activities of GFORΔ32–46, GFORΔ2–74, and mutants thereof (compare with Fig. 4) are shown. Mean values and deviations were obtained from two independently performed assays. ND, not detectable (ΔE/min for the GFOR and glucose dehydrogenase test with 5 μg/ml enzyme <0.01/min). GFOR activities were determined with the standard photometric test without supplemented NADP (−NADP). In addition, GFOR activities were determined after preincubation of 5 mg/ml each of GFORΔ32–46, GFORΔ2–74, or mutants thereof with 20 mmNADP in 40 mm K-MES, pH 6.4, for 5 min, at 25 °C (+NADP). The final concentration of enzyme was 5 μg/ml and 20 μm NADP in the assay mix. To examine whether the differences in intensity of tryptophan fluorescence between wild-type GFOR and mutant B reflected differences in the native conformation, the respective enzymes were denaturated with 6 m guanidinium hydrochloride. In the denatured state, wild-type GFOR and mutant B showed the same intensity of tryptophan fluorescence and the NADPH fluorescence at 450 nm of wild-type GFOR vanished (Fig. 5). These results indicate that mutant B did not contain the tightly bound cofactor NADP(H). Moreover, in wild-type GFOR a quenching of tryptophan fluorescence energy occurred, most likely by a direct transfer of fluorescence energy from tryptophan residues to the 1,4-dihydronicotineamide ring of NADPH which does not absorb light at a wavelength of 295 nm. To release any bound cofactor (oxidized or reduced forms of NADP or NAD), protein from wild-type GFORΔ32–46, mutant A, and mutant B was denatured by perchloric acid, and the supernatants were analyzed for NAD(P) by HPLC. NADP was detected with GFORΔ32–46 and from mutant protein A. No NADP appeared from mutant B. NAD was not detected from any protein (data not shown). We deduce that a single amino acid residue exchange (S116D) is sufficient to destroy tight cofactor binding to GFOR. The enzymatic assay for GFOR activity is usually performed without NAD

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