Tài liệu Báo cáo khóa học: Characterization of the products of the genes SNO1 and SNZ1 involved in pyridoxine synthesis in Saccharomyces cerevisiae pptx

Characterization of the products of the genes SNO1 and SNZ1 involved in pyridoxine synthesis in Saccharomyces cerevisiae Yi-Xin Dong, Shinji Sueda, Jun-Ichi Nikawa and Hiroki Kondo Department of Biochemical Engineering and Science, Kyushu Institute of Technology, Iizuka, Japan Genes SNO1 and SNZ1 are Saccharomyces cerevisiae homologues of PDX2 and PDX1 which participate in pyri- doxine synthesis in the fungus Cercospora nicotianae.In order to clarify their function, the two genes SNO1 and SNZ1 were expressed in Escherichia coli either individually or simultaneously and with or without a His-tag. When expressed simultaneously, the two protein products formed a complex and showed glutaminase activity. When purified to homogeneity, the complex exhibited a specific activity of 480 nmolÆmg )1 Æmin )1 as glutaminase, with a K m of 3.4 m M for glutamine. These values are comparable to those for other glutamine amidotransferases. In addition, the gluta- minase activity was impaired by 6-diazo-5-oxo- L -norleucine in a time- and dose-dependent manner and the enzyme was protected from deactivation by glutamine. These data sug- gest strongly that the complex of Sno1p and Snz1p is a glutamine amidotransferase with the former serving as the glutaminase, although the activity was barely detectable with Sno1p alone. The function of Snz1p and the amido acceptor for ammonia remain to be identified. Keywords: glutamine amidotransferase; pyridoxine biosyn- thesis; Saccharomyces cerevisiae; SNO1; SNZ1. Pyridoxal phosphate plays a crucial role in amino acid metabolism. Pyridoxine and its phosphate are the precur- sors of pyridoxal phosphate and the major forms of vitamin B6. Pyridoxine biosynthesis in Escherichia coli has been studied extensively but only recently has the whole synthetic pathway been finally established [1–3]. There are organisms such as budding yeast, Saccharomyces cerevisiae,which also synthesize pyridoxine but in a different pathway. This notion is based in part on an observation that the nitrogen of pyridoxine is derived from the amide group of glutamine in yeast [4], while glutamate is the source of the ring nitrogen in E. coli [5]. Recently, two independent groups identified pyroA and SOR1 (PDX1) as participating in pyridoxine synthesis in fungi Cercospora nicotianae and Aspergillus nidulans, respectively [6,7]. They are homologous genes and their homologues are distributed widely in various organ- isms, but nothing of their function is known except that SNZ1, the yeast homologue, works in the stationary phase of yeast cells together with SNO1 [8]. In addition to these observations, it was shown recently that a pentose or pentulose constitutes the skeleton of pyridoxine in yeast [9,10]. Herein, we report that Sno1p and Snz1p serve as a glutaminase to supply ammonia for the ring nitrogen of pyridoxine in yeast. Based on these and other lines of evidence, a putative synthetic pathway to pyridoxine is presented in the Discussion in which ribulose 5-phosphate and ammonia serve as the key starting or intermediary material. Experimental procedures Materials Inorganic salts and common organic chemicals including amino acids, nucleic bases and vitamins were obtained from commercial sources. Acetylpyridine adenine dinucleotide (APAD) and 6-diazo-5-oxo- L -norleucine (DON) were from Sigma (St. Louis, MO, USA). Glutamate dehydrogenase from bovine liver was obtained from Oriental Yeast (Tokyo, Japan). Reagents for genetic engineering such as restriction enzymes were purchased from Takara (Kyoto, Japan) and New England Biolabs (Beverly, MA, USA). Oligonucleo- tides were custom synthesized by Hokkaido Science (Sap- poro, Japan). Plasmid YEpM4 was a 2 lmDNA-based shuttle vector with gene LEU2 as the selectable marker [11]. Plasmids pET21a, pET21d (both ampicillin resistant), pET24a (kanamycin resistant) and His-bind columns were from Novagen (Madison, WI, USA). The TOPO TA cloning kit was the product of Invitrogen (Carlsbad, CA, USA). Strains and media The S. cerevisiae strain used in this study was D373-1 (MATa, leu2, his3, trp1) [12]. The following medium was used to grow yeast: YPD [1% yeast extract, 2% polypep- tone, 2% glucose (v/v/v)] and synthetic medium [glucose, 20 g; (NH 4 ) 2 SO 4 ,1.02g;KH 2 PO 4 , 0.875 g; K 2 HPO 4 , 0.125 g; CaCl 2 ÆH 2 O, 0.02 g; NaCl, 0.01 g; MgSO 4 Æ7H 2 O, 0.05 g; CuSO 4 Æ5H 2 O, 40 lg; MnSO 4 ÆH 2 O, 400 lg; FeCl 3 Æ 6H 2 O, 200 lg; ZnSO 4 Æ7H 2 O, 400 lg; Na 2 MoO 4 Æ2H 2 O, Correspondence to H. Kondo, Department of Biochemical Engineering and Science, Kyushu Institute of Technology, Kawazu 680-4, Iizuka 820-8502, Japan. Fax: + 81 948 7801, Tel.: + 81 948 29 7814, E-mail: kondo@bse.kyutech.ac.jp Abbreviations: APAD, acetylpyridine adenine dinucleotide; APADH, reduced form of APAD; DON, 6-diazo-5-oxo- L -norleucine. (Received 7 October 2003, revised 2 December 2003, accepted 23 December 2003) Eur. J. Biochem. 271, 745–752 (2004) Ó FEBS 2004 doi:10.1111/j.1432-1033.2003.03973.x 200 lg; KI, 100 lg; H 3 BO 3 ,500lg; biotin, 2 lg; inositol, 10 mg; nicotinic acid, 0.2 mg and calcium pantothenate, 0.2 mg, per litre]. Where indicated, the following supple- ments were added to the synthetic media at a final concentration of 20 mgÆL )1 : histidine, leucine, tryptophan and pyridoxine. Solid media contained 2% agar. For each liquid culture experiment, yeast cells were shaken at 250 r.p.m. in 10 mL of medium at 30 °Cfortheperiodof time specified. LB medium [1% polypeptone, 0.5% NaCl and 0.5% yeast extract (v/v/v)] was used to grow E. coli. General procedures All of the gene manipulations of S. cerevisiae and E. coli were carried out using standard methods [13]. Chromo- somal DNA from S. cerevisiae was prepared according to the literature [14]. Yeast genomic DNA sequences were retrieved from a database by using the website (http:// genome-www.stanford.edu/Saccharomyces/). PCR was run on an Astec PC-700 (Tokyo, Japan) or Biometra T-gradient thermoblock (Gottingen, Germany). Sequencing of plas- mids was carried out by the dideoxy chain termination method on an automatic DNA sequencer model DSQ1000 from Shimadzu (Kyoto, Japan). Protein sequences were determined on an Applied Biosystems 491 A sequencer. Mutant construction The pyridoxine auxotrophic mutants of yeast were pro- duced by ethyl methanesulfonate mutagenesis [15]. In brief, cells were treated with 3% ethyl methanesulfonate for 50 min and then spread over the entire surface of synthetic medium plates supplemented with pyridoxine and the growth requirements, and the plates were incubated at 30 °C. The colonies that appeared on the supplemented plates were then transferred by replica plating first to a minimal plate without pyridoxine (–PN) and then to one supplemented with pyridoxine (+PN). The plates were scored for colonies that appeared on +PN media but not on the –PN media. This selection process was repeated several times to ensure that the colonies that were unable to grow in –PN medium are indeed pyridoxine auxotrophs. Transformation of yeast cells Pyridoxine auxotrophic mutants were transformed with a YEpM4-based genomic library [11] for complementation by the lithium acetate protocol [16]. Transformants were selected on synthetic media lacking pyridoxine and leucine. The plasmids were isolated from S. cerevisiae as described [14]. Construction of over-expression plasmids for the SNO1 and SNZ1 genes The coding regions of the SNO1 (672 bp) and SNZ1 (891 bp) genes were amplified by PCR in one step with the oligonucleotides shown in Table 1 as primers. Both of the genes were constructed either with or without a His-tag at the 3¢ terminus of the coding region. Thus, the stop codon of SNO1 and SNZ1 was replaced with bases coding for the sequence LEHHHHH as a C-terminal extension. The PCR was run as follows: After heating at 94 °C for 5 min, the following cycle was repeated 25 times; 94 °C, 1 min; 55 °C, 1min;70°C, 1–1.5 min, and finally heated at 72 °Cfor 5 min. Each PCR product was purified by agarose gel electrophoresis before ligation into pCR2.1-TOPO (Invi- trogen). After confirming the correct DNA sequence, the coding region of SNO1 and SNZ1 was excised from the plasmid and, where the His-tag was present, recloned into the NdeI/XhoI sites of pET21a or pET24a, respectively. Where a His-tag was absent, the coding region of both genes was recloned into the NcoI/BamHI sites of pET21d. The resulting recombinant plasmids are termed pSNO1H, pSNZ1H, pSNO1 and pSNZ1, respectively (Table 2). Protein production and purification The two genes were expressed in E. coli BL21(DE3) (Novagen) separately or simultaneously following trans- formation with one or two of the plasmids prepared above. Transformants were grown in 1 L of LB medium in the presence of ampicillin (50 lgÆmL )1 ), kanamycin (30 lgÆmL )1 ) or both, to late logarithmic phase, whereupon isopropyl thio-b- D -galactoside was added to 0.4 m M ,except for the expression of Sno1p (0.1 m M ). Eight hours after induction the cells were collected by centrifugation and washed with phosphate buffered saline. Subsequent steps of protein purification were carried out at 4 °C, unless otherwise stated. The complex of Sno1p with a His-tag and Snz1p without a tag. The washed cells were resuspended in 100 mL of 5m M imidazole, 0.5 M NaCl, 20 m M Tris/HCl, pH 8.0 Table 1. Sequences of oligonucleotides used as PCR primers. Symbols O, Z and H stand for Sno1p, Snz1p and His-tag, respectively. Underlined are the restriction enzyme sites. Primer Sequence P1O 5¢-ATA CCATGGACAAAACCCACAGTACAATG P1OH 5¢- CATATGCACAAAACCCACAGTAC P2O 5¢-TAT GGATCCTTAATTAGAAACAAACTGTCTGA TAAAC P2OH 5¢- CTCGAGATTAGAAACAAACTGTCTGATAAACC P1Z 5¢-ATA CCATGGCTGGAGAAGACTTTAAGATC P1ZH 5¢- CATATGACTGGAGACTTTAAGATC P2Z 5¢-TAT GGATCCTCACCACCCAATTTCGGAAAG P2ZH 5¢- CTCGAGCCACCCAATTTCGGAAAGT Table 2. Over-expression plasmids for Sno1p and Snz1p prepared in this study. Symbols O, Z and H stand for Sno1p, Snz1p and His-tag, respectively. Name of plasmid Primer VectorForward Backward pSNO1 P1O P2O pET21d pSNO1H P1OH P2OH pET24a pSNZ1 P1Z P2Z pET21d pSNZ1H P1ZH P2ZH pET24a 746 Y X. Dong et al.(Eur. J. Biochem. 271) Ó FEBS 2004 (buffer A) containing 5 m M glutamine, 0.4 m M phenyl- methanesulfonyl fluoride and 0.2 mL of dimethylsulfoxide. The suspension was sonicated and then centrifuged at 17 000 g for 30 min. The supernatant containing 120 mg of protein was filtered through a 0.45 lm filter and applied to a His-Bind affinity column (1 · 10 cm), pretreated with 50 m M NiSO 4 and equilibrated with buffer A. The column was washed successively with buffer A and buffer B (50 m M imidazole, 0.5 M NaCl, 20 m M Tris/HCl, pH 8.0) and protein was finally eluted with a gradient of 100–500 m M imidazole in buffer B. The protein-containing fractions were pooled and dialyzed against 35 m M potas- sium phosphate, 1 m M EDTA and 0.1 m M dithiothreitol, pH 7.5. The purity of the desired protein was greater than 95% at this stage and the yield was 25 mg from a 1 L culture. Sno1p and Snz1p with a His-tag were purified analogously with a yield of 30 mg each from 1 L of culture. Protein concentration was determined on the basis of the molar extinction coefficient of 15.9 and 12.3 m M )1 Æcm )1 at 280 nm for Sno1p and Snz1p, respectively, deduced from their amino acid compositions. Sno1p without a His-tag. The washed cells were resus- pended in 50 mL of 10 m M potassium phosphate buffer and 1m M EDTA, pH 7.0. The suspension was sonicated and then centrifuged at 17 000 g for 20 min. The pellet was taken up in 50 mL of the same buffer and 2 mL of Triton X-100 and shaken at room temperature for 30 min. The suspension was centrifuged (17 000 g) for 30 min and the supernatant discarded. This process was repeated twice more for the pellet. The washed pellet was mixed with 10 mL of 50 m M Tris/HCl containing 10 m M dithiothreitol and 8 M urea, pH 9.0, shaken at 37 °Cfor1hand centrifuged for 30 min. The supernatant was mixed with 30mLofthesamebufferwith6 M urea and dialyzed against the same buffer with 4 M urea. Dialysis was continued against the buffers containing 2, 1 and 0 M urea, successively. The solution was concentrated to one half the volume by dialysis against 50 m M Tris/HCl, 1 m M dithio- threitol and 0.1 m M EDTA, pH 9.0, containing 15% polyethylene glycol 20 000 and subjected to gel filtration chromatography on Superdex 200 (2.6 · 60 cm, Pharma- cia) to give 30 mg of virtually pure protein. Snz1p without a His-tag. Harvested cells were processed in a way identical to that for Sno1p up to the cell disruption step. The supernatant was subjected to DEAE- cellulose chromatography (2 · 10 cm, Whatman, Maid- stone, UK) and protein was eluted with a linear gradient of 0–500 m M NaCl in 50 m M potassium phosphate, pH 7.0. Snz1p was eluted at about 200 m M salt. The pooled fractions (25 mL) were concentrated and then subjected to gel filtration chromatography as described above to give 14 mg of the desired protein. Glutaminase assay The glutaminase activity of the complex of Sno1p with a His-tag and Snz1p was determined in two steps. Thus, glutamate formed was converted to 2-oxoglutarate by glutamate dehydrogenase with acetylpyridine adenine dinu- cleotide (APAD) as cosubstrate [17]. The glutaminase reaction was carried out in 0.3 mL of 50 m M Tris/HCl, pH 8.0, in the presence of 1–10 m M glutamine and 30 lgof the complex at 30 °C for 10 min. The sample was then boiledfor1minandkeptfrozenat)80 °C for subsequent analysis. To quantitate the amount of glutamate formed, 0.3 mL of the sample was incubated in 1 mL of 50 m M Tris/ HCl, pH 8.0, containing 1 m M EDTA, 0.5 m M APAD and 7 units of glutamate dehydrogenase at 37 °C for 90 min. After centrifugation (14 000 g) for 1 min, the absorbance of the supernatant was read at 363 nm. The absorbance of the reduced form of APAD (APADH) was linear over the 2–100 l M range of glutamate with a molar extinction coefficient of 8900 M )1 Æcm )1 [17]. Detection of glutamate Glutamate formed from glutamine by the complex of Sno1p with a His-tag and Snz1p, was further detected by TLC following dansylation. The reaction mixture (0.3 mL) contained 10 m M glutamine, 30 lgofthecomplexin 50 m M Tris/HCl, pH 8.0, and it was incubated at 30 °Cfor 10 min. Ten microliter aliquots of this solution together with 10 lLeachof10m M glutamine and glutamate were separately allowed to react with dansyl chloride under standard conditions [18]. Aliquots (1.5 lLeach)were spotted on silica gel 60 F 254 (Merck, Darmstadt, Germany) and developed in chloroform-t-amyl alcohol-glacial acetic acid (70 : 30 : 3, v/v/v). The dansylated products were visualized under ultraviolet light. Inhibition of Sno1p with DON Inhibition of the complex of Sno1p with a His-tag and Snz1p by 6-diazo-5-oxo-L-norleucine (DON) was studied according to the literature [19]. The reaction mixture contained 0.15 mg of the complex and various concentra- tions (1–10 m M )ofDONin1mLof50m M Tris/HCl, pH 8.0, and was incubated at 30 °C. Aliquots (200 lL) removed at a specified time were mixed with other ingredients for glutaminase assay and the remaining activity of the complex was determined as described above. Results Yeast pyridoxine auxotrophs Pyridoxine auxotrophs of S. cerevisiae were prepared by ethyl methanesulfonate mutagenesis. Several strains were tested including D373-1 and in all the cases auxotrophs showing clear pyridoxine requirements were obtained (data not shown). After several rounds of screening, around 20 mutants were established from D373-1. Among these was mutant K64 (Fig. 1). To identify the gene(s) affected by complementation, the mutant was transformed with a library of yeast chromosomal DNA. It was found that 4.6 and 5.1 kb overlapping fragments of chromosome XIII carrying SNO1 and SNZ1 were capable of complementing the defect of mutant K64 (Figs 1 and 2). They are homologues of PDX2 and PDX1 of fungus C. nicotianae, respectively [7,20], strongly suggesting that they are the genes responsible for the pyridoxine auxotrophy observed for that mutant. Ó FEBS 2004 Pyridoxine biosynthesis in yeast (Eur. J. Biochem. 271) 747 Site of mutation in K64 In order to identify the site of mutation, the two genes were amplified by PCR with the chromosomal DNA of mutant K64 as template. Sequencing of the mutated genes revealed that there was indeed a mutation on both of the genes. In SNO1, the 199th G from the 5¢ terminus of the open reading frame was deleted to result in a frame-shift and appearance of stop codons in the downstream. As a result, a protein as small as 70 residues is generated, a size too small for any protein to be functional as an enzyme (see below). In SNZ1, the 709th G was converted to A to result in replacement of Gly237 with Arg. It is noted that this residue and the surrounding regions are well conserved among the homo- logues of SNZ1 including pyroA and PDX1 [6,7], suggesting that the residue plays an important role. It should be emphasized that the dual mutation of the two genes was necessary to make yeast cells pyridoxine auxotrophic; cells were still viable in the absence of pyridoxine even when either one of the genes was disrupted separately (data not shown). This observation is consistent with those reported previously [21]. Presumably, their homologues SNO2, SNO3, SNZ2 and SNZ3 complement the defect. The relationship of all of these genes remains to be clarified. Expression and purification of Sno1p and Snz1p In light of the similarity in the amino acid sequence of Sno1p to that of the glutaminase subunit of imidazole glycerol phosphate synthase [22], Sno1p may possess the ability to hydrolyze glutamine. In addition, in reference to the observation that Sno1p and Snz1p act together in the stationary phase of yeast cells [8] and the result of two- hybrid analysis [23], they may form a complex. To assess these possibilities, we embarked on a characterization of the two proteins. Over-expression plasmids were prepared for both genes with or without a C-terminal His-tag (Table 2), as detailed under Experimental procedures. E. coli cells were trans- formed with one or two of the recombinant plasmids. The resulting cells over-expressed the desired protein(s) up to 15% of the total cellular proteins. When Sno1p was expressed alone, it was kept in the inclusion body, irrespective of the occurrence of a His-tag (data not shown). Whereas, Snz1p, expressed either alone or coexpressed with Sno1p, was found in the soluble fractions. Although purification procedures differed slightly from sample to sample, depending on the occurrence of a tag and the location of the protein in question, Sno1p, Snz1p and their complex were eventually purified to near homogeneity (Fig. 3). Typically, 30, 30 and 25 mg of protein were Fig. 1. Growth of S. cerevisiae strains D373-1, K64 and K64t. Yeast cells were treated with ethyl methanesulfonate as detailed under Experimental procedures. After several rounds of screening on media containing pyridoxine (+PN) and not containing pyridoxine (–PN), auxotrophic mutants were established, one of which was K64. K64t represents the transformant harboring plasmid pDYX11 (Fig. 2) and is capable of growing in –PN medium. Fig. 2. Partial map of S. cerevisiae chromosome XIII carrying SNO1 and SNZ1. Dark grey bars represent 4.6 and 5.1 kb DNA fragments capable of complementing the pyridoxine auxotrophy of mutant K64. Fig. 3. SDS/PAGE of purified Sno1p, Snz1p and their complex with or without a C-terminal His-tag. Lane 1, protein markers; lane 2, Sno1p without tag; lane 3, Sno1p with tag; lane 4, complex of Sno1p with tag and Snz1p without tag; lane 5, Snz1p without tag; lane 6, Snz1p with tag. About 10–20 lg of protein was loaded and stained with Coo- massie Brilliant blue. 748 Y X. Dong et al.(Eur. J. Biochem. 271) Ó FEBS 2004 obtained from a 1 L culture for Sno1p, Snz1p and the complex, respectively. Identity of each protein was con- firmed by N-terminal sequencing; the sequences were correct at least to the 9th cycle from the N-terminus including the initiating Met: MHKTHSTMS for Sno1p and MTGEDFKIKS for Snz1p. Properties of the complex of Sno1p and Snz1p When Sno1p with a His-tag and Snz1p without a tag were coexpressed and then applied to a His-Bind affinity column, not only Sno1p but also Snz1p bound to the column and was eluted simultaneously by imidazole, strongly suggesting that they form a complex. The ratio of the two proteins, assessed by SDS/PAGE, seemed to be equimolar, although a more rigorous assessment is necessary to prove this assertion. The glutaminase activity of this complex was assessed in two ways. First, the glutamate formed was oxidized to 2-oxoglutarate by APAD and glutamate dehydrogenase, and the generated APADH was determined spectroscopically. Second, the glutamate formed was dansy- lated and analyzed by TLC (Fig. 4). These experiments consistently pointed to the formation of glutamate, proving unambiguously that the complex did indeed hydrolyze glutamine efficiently. Kinetics of the glutaminase reaction The kinetics of glutamine hydrolysis mediated by the complex of Sno1p with a His-tag and Snz1p was determined at 30 °C. The reaction rate was proportional to protein concentration over a 5–50 lg range and it was constant up to at least 1 h (data not shown). The reaction followed the simple Michaelis–Menten formulation as illustrated in Fig. 5. The specific activity (V max )andK m for glutamine obtained from analysis of these data were 0.48 lmolÆ min )1 Æmg )1 and 3.4 m M , respectively. These values seem to be reasonable for a glutaminase, though the specific activity varies drastically from enzyme to enzyme and depends on whether a proper synthetase partner and cosubstrate are present or not. For example, the V max and K m values of imidazole glycerol phosphate synthase in the absence of substrate are 0.084 lmolÆmin )1 Æmg )1 and 4.8 m M , respectively [22]. The V max was enhanced 39-fold and K m lowered 20-fold in the presence of substrate N 1 -[(5¢- phosphoribulosyl)formimino]-5-aminoimidazole 4-carbox- amide ribonucleotide. It may be reasonable therefore to assume that once the unknown substrate or ligand for Snz1p was added, the glutaminase activity could have been even higher. It should be noted that the enzyme activity is gradually lost over time with a half life of 2–3 days at 4 °C. Several additives such as ATP were tested as a stabilizer, but thus far we have not been successful in stabilizing the enzyme activity. Incidentally, no glutaminase activity was found for Snz1p alone. The function of Sno1p The data shown above revealed that the complex of Sno1p and Snz1p is a glutamine amidotransferase, with the former serving as the glutamine-hydrolyzing machinery. This latter notion is based solely on the sequence homology of Sno1p with the glutaminase subunit of imidazole glycerol phos- phate synthase. To test this hypothesis directly, Sno1p and Snz1p were separately expressed, purified and characterized as detailed under Experimental procedures. As described above, Sno1p expressed in E. coli resided in the inclusion body and hence use of Triton X-100 and urea was necessary for its solubilization. Even after renaturation, however, Sno1p exhibited no glutaminase activity, nor did the addition of 0.5 to 3-fold Snz1p have any effect. It was only when Sno1p and Snz1p were partially denatured together and then renatured, that glutaminase activity was observed. Thus, the two proteins were mixed in 8 M urea in various Fig. 4. TLC analysis of glutamate formed from glutamine mediated by the complex of Sno1p with a His-tag and Snz1p. Glutamate, glutamine and the reaction mixture were dansylated separately and aliquots (2.5 nmol each) were applied on silica gel 60 F 254 and developed in chloroform-t-amyl alcohol-glacial acetic acid (70 : 30 : 3, v/v/v). The dansylated (DANS) products were visualized under ultraviolet light. Fig. 5. Michaelis–Menten kinetics for glutamine hydrolysis mediated by the complex of Sno1p with a His-tag and Snz1p. The reaction was carried out in 1 mL of 50 m M Tris/HCl, pH 8.0, in the presence of 1–10 m M glutamine and 50 lg of the complex, at 30 °C for 10 min. The sample was then boiled for 1 min and a 0.3 mL aliquot was incubated in 1 mL of 50 m M Tris/HCl, pH 8.0, containing 1 m M EDTA, 0.5 m M APAD and 7 units of glutamate dehydrogenase at 37 °C for 90 min. After centrifugation for 1 min, the absorbance of the supernatant was read at 363 nm for APADH. The curve drawn is a theoretical one based on 0.48 lmolÆmin )1 Æmg )1 for V max and 3.4 m M for K m for glutamine. Ó FEBS 2004 Pyridoxine biosynthesis in yeast (Eur. J. Biochem. 271) 749 molar ratios (0.2 : 4 of Sno1p : Snz1p). The urea concen- tration was lowered gradually to 6, 4, 2, and 1 M by dialysis. Finally, the enzyme solution was dialyzed against buffer without urea, and glutaminase activity was determined. The mixtures of Sno1p and Snz1p with a molar ratio of the former greater than 0.5 exhibited partial activity; at a maximum of 5% of that of the intact complex described above. This suggests that although Sno1p and Snz1p associate spontaneously, renaturation of either one or both of the proteins was incomplete and the complex formation is slow and/or incomplete under the conditions employed. Inhibition of the glutaminase activity by DON Glutamine amidotransferases are inhibited irreversibly by 6-diazo-5-oxo- L -norleucine (DON) [24]. Hence, the suscep- tibility of the complex of Sno1p and Snz1p to DON was studied under conditions detailed under Experimental procedures. As shown in Fig. 6, DON deactivated the enzyme in a time- and dose-dependent manner. Typically, the half-life of deactivation was 10 min at 10 m M DON at 30 °C. It is noted that the inhibition did not go to completion even at the highest concentration of DON employed but halted at a point where about half of the enzyme activity is lost. The reason for this phenomenon is not known but a similar observation was made for phosphoribosylpyrophos- phate amidotransferase [24]. Glutamine was effective in protecting the enzyme from inactivation and its effect was again dose-dependent, suggesting that inhibition by DON occurs at the active site or the glutamine-binding site of the enzyme (Sno1p). Although DON inhibition of Sno1p was not pursued further, it is worth pointing out that the cysteine serving as the key catalytic residue and modified covalently by DON in other amidotransferases is also conserved in Sno1p at position 100. Discussion As described above, the gene products of SNO1 and SNZ1 serve as a glutamine amidotransferase, which is needed to supply ammonia as a source of the ring nitrogen of pyridoxine [4]. Although Sno1p alone does not exhibit detectable glutaminase activity, it seems certain that it is responsible for the hydrolysis of glutamine mediated by the complex with Snz1p. For example, the amino acid sequence of Sno1p has 40% identity to that of the glutaminase subunit of yeast imidazole glycerol phosphate synthase [22]. In addition, the key catalytic residues required for glutamine hydrolysis by glutamine amidotransferases including imi- dazole glycerol phosphate synthase, i.e. Cys100, His203 and Glu205 (numbering based on Sno1p), are conserved in Sno1p as well. Presumably, Sno1p hydrolyzes glutamine by the same mechanism as those of other glutamine amido- transferases such as imidazole glycerol phosphate synthase and carbamoyl-phosphate synthetase, whose three-dimen- sional structures are available [25,26]. In light of the function of these glutamine amidotrans- ferases, Snz1p may be a synthetase that is mediating the coupling of ammonia released from glutamine with an unknown acceptor substrate. In reactions of many amido- transferases, ammonia is delivered from the site of forma- tion to the site of coupling through a tunnel that is present in the synthetase subunit [25,26]. It is hence tempting to assume that there is such a tunnel in Snz1p as well, although this notion awaits experimental verification of the structure Fig. 6. Inhibition of the glutaminase activity of the complex of Sno1p with a His-tag and Snz1p by 6-diazo-5-oxo- L -norleucine (DON). The reaction mixture contained 150 lgofthecomplexandno(s), 1 (e), 5 (h,) and 10 m M (n) DON in the absence (open symbols) and presence (j)of10 m M glutamine in 1 mL of 50 m M Tris/HCl, pH 8.0, at 30 °C. Aliquots (200 lL)werewithdrawnatspecifiedtimesandtheremaining activity of the complex was determined. Scheme 1. Putative synthetic pathway to pyridoxine in yeast. Ammonia released from glutamine by Sno1p condenses with an unknown acceptor (?) or a derivative of ribulose 5-phosphate. This process is mediated by Snz1p but the subsequent steps remain obscure. 750 Y X. Dong et al.(Eur. J. Biochem. 271) Ó FEBS 2004 by means of X-ray crystallography. Incidentally, the amino acid sequence of Snz1p does not show homology to any other known proteins and hence its three-dimensional structure could be unique. In addition, once the amido- acceptor substrate of Snz1p is identified, its addition to the reaction system will enhance the glutaminase activity of Sno1p significantly. In light of the fact that a ketopentose seems to be a component of the skeleton of pyridoxine in yeast and related organisms (see below), dihydroxyacetone phosphate, glyceraldehyde 3-phosphate or related com- pounds are probable candidates for the coupling partner. These possibilities are presently under scrutiny in this laboratory. Recently, it was shown that a ketopentose is one of the starting materials for pyridoxine in yeast and the initial form of vitamin B6 produced is 2¢-hydroxypyridoxine [9,10]. Our unpublished observation supports these findings; the gene RKI1 coding for ribose 5-phosphate ketol-isomerase (Rki1p), which interconverts ribose 5-phosphate and ribu- lose 5-phosphate, dictates somehow pyridoxine synthesis in yeast. In this light, ribulose 5-phosphate may be the more probable candidate for the starting material, as its structure fits the skeleton from positions 2¢ to 4¢ of 2¢-hydroxypyri- doxine neatly. Hence, the possibility that ribulose 5-phos- phate serves as the direct ammonia-acceptor was addressed. It was found that this compound considerably inhibits the glutaminase activity of the complex of Sno1p and Snz1p in a competitive fashion; the activity decreased to 70% at 8 m M . Ribose 5-phosphate was as equally effective as ribulose 5-phosphate but dihydroxyacetone phosphate was without effect. These data seem to suggest that, although it does interact with the glutaminase complex, ribulose 5-phosphate is not the direct acceptor of ammonia. Based on these arguments, the following scheme is proposed for the synthetic route to pyridoxine in yeast and related organisms (Scheme 1). Ammonia released from glutamine undergoes condensation with either a derivative of ribulose 5-phosphate or an unknown acceptor with a three-carbon unit to give an aminated intermediate. These two components eventually undergo coupling and cycliza- tion to form 2¢-hydroxypyridoxine. If this scheme holds, the synthetic pathways to pyridoxine of yeast and related organisms are remarkably similar to that of E. coli, as both utilize a ketopentose or its derivative as the starting material. Although it seems certain that 3-amino-1-phosphohydroxy- propan-2-one, a key component in the E. coli pathway and generated by the product of gene pdxA, is not involved, compounds related to it structurally could be the coupling partner of the ketopentose in yeast. One characteristic feature of the genes SNO1 and SNZ1 is that they act simultaneously in the stationary phase of yeast cells [8]. They are assumed to participate in the metabolism of nucleotides. In this context, the assertion that pyridoxine may be the precursor of the pyrimidine moiety of thiamin in yeast is intriguing [27]. In fungi, pyroA and SOR1 (PDX1), homologues of yeast SNZ1, play a defensive role against reactive oxygen species such as singlet oxygen [6,7]. Taken together, pyridoxine synthesis in various organisms ranging from Bacillus to plants seems to play a broader physiological role than simply supplying the cofactor that is essential for amino acid metabolism. 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Dong et al.(Eur. J. Biochem. 271) Ó FEBS 2004 . Characterization of the products of the genes SNO1 and SNZ1 involved in pyridoxine synthesis in Saccharomyces cerevisiae Yi-Xin Dong, Shinji Sueda,. to the 9th cycle from the N-terminus including the initiating Met: MHKTHSTMS for Sno1p and MTGEDFKIKS for Snz1p. Properties of the complex of Sno1p and Snz1p When