Eukaryotic and Bacterial Gene Clusters Related to an Alternative Pathway of Nonphosphorylated L-Rhamnose Metabolism*

The Entner-Doudoroff (ED) pathway is a classic central pathway of d-glucose metabolism in all three phylogenetic domains. On the other hand, Archaea and/or bacteria possess several modified versions of the ED pathway, in which nonphosphorylated intermediates are involved. Several fungi, including Pichia stipitis and Debaryomyces hansenii, possess an alternative pathway of l-rhamnose metabolism, which is different from the known bacterial pathway. Gene cluster related to this hypothetical pathway was identified by bioinformatic analysis using the metabolic enzymes involved in analogous sugar pathways to the ED pathway. Furthermore, the homologous gene cluster was found not only in many other fungi but also several bacteria, including Azotobacter vinelandii. Four putative metabolic genes, LRA1-4, were cloned, overexpressed in Escherichia coli, and purified. Substrate specificity and kinetic analysis revealed that nonphosphorylated intermediates related to l-rhamnose are significant active substrates for the purified LRA1-4 proteins. Furthermore, l-2-keto-3-deoxyrhamnonate was structurally identified as both reaction products of dehydration by LRA3 and aldol condensation by LRA4. These results suggested that the LRA1-4 genes encode l-rhamnose 1-dehydrogenase, l-rhamnono-γ-lactonase, l-rhamnonate dehydratase, and l-KDR aldolase, respectively, by which l-rhamnose is converted into pyruvate and l-lactaldehyde through analogous reaction steps to the ED pathway. There was no evolutionary relationship between l-KDR aldolases from fungi and bacteria.

Based on these insights, we here identified homologous gene clusters related to a novel L-rhamnose metabolism between eukaryotes and bacteria. Enzymatic and biochemical characterization revealed that the four metabolic genes encode L-rhamnose 1-dehydrogenase, L-rhamnono-␥-lactonase, L-rhamnonate dehydratase, and L-2-keto-3-deoxyrhamnonate (L-KDR) aldolase. The evolutionary relationship between sugar pathways analogous to ED pathways, including the alternative L-rhamnose pathway, is discussed by significant phylogenetic comparison of metabolic genes.

EXPERIMENTAL PROCEDURES
General Procedures-Basic recombinant DNA techniques were performed as described by Sambrook et al. (23). PCR was carried out using a PCR Thermal Cycler PERSONAL (Takara) for 30 cycles in 50 l of reaction mixture containing 1.25 units of Ex Taq DNA polymerase (Takara), appropriate primers (10 pmol), and template DNA under the following conditions: denaturation at 98°C for 10 s, annealing at 50°C for 30 s, and extension at 72°C for times calculated as an extension rate of 1 kbp/min. Protein concentrations were determined by the method of Lowry et al. (24) with bovine serum albumin as the standard. SDS-PAGE was performed as described by Laemmli (25). HPLC was performed using a Multi-Station LC-8020 model II system (TOSOH). Samples were applied at 35°C to an Aminex HPX-87H Organic Analysis column (300 ϫ 7.8 mm; Bio-Rad) linked to an RID-8020 refractive index detector (TOSOH) and eluted with 5 mM H 2 SO 4 at a flow rate of 0.6 ml/min. If necessary, 12% (w/v) trichloroacetic acid was added to samples (0.1 volume) to remove proteins. After filtration, 100 l of this solution was then analyzed. NMR spectra were recorded on a JEOL JNM-ECA600 FT NMR spectrometer (JEOL Ltd., Tokyo, Japan) operating at 600 MHz.
Microorganism Strains and Cell Growth-Pichia stipitis CBS 6054 was kindly provided by Dr. T. W. Jeffries (University of Wisconsin). Debaryomyces hansenii NBRC 0083 and Azotobacter vinelandii NBRC 102612 were purchased from the National Institute of Technology and Evaluation (Chiba, Japan). Two fungi were grown at 30°C in YPD medium (10 g of yeast extract, 20 g of peptone, and 20 g of glucose per liter) and in YNB medium (6.7 g of yeast nitrogen base without amino acid/liter) supplemented with 20 g/liter L-rhamnose. A. vinelandii was cultured at 30°C in medium (pH 7.6), containing 20.0 g of L-rhamnose, 0.5 g of yeast extract, 0.1 g of KH 2 PO 4 , 0.8 g of K 2 HPO 4 , 0.2 g of MgSO 4 ⅐H 2 O, 0.1 g of CaSO 4 ⅐2H 2 O, and trace amounts of FeCl 3 ⅐6H 2 O and NaMoO 4 ⅐2H 2 O per liter. Commonly, L-rhamnose was sterilized separately by filtration and added to each medium.
Substrates-All acid-sugars were prepared by hypoiodite-inmethanol oxidization (26) from the corresponding sugars as K ϩ salt (D-gluconate, D-galactonate, D-arabonate, L-arabonate, and D-xylonate) or Ba 2ϩ salt (L-rhamnonate, L-lyxonate, D-mannonate, L-mannonate, D-fuconate, and L-fuconate). The K ϩ salt was dissolved in an approximate amount of water, and the pH was adjusted to ϳ10.0 with 5 M NaOH. On the other hand, the Ba 2ϩ salt was dissolved by the addition of appropriate H 2 SO 4 , and the produced BaSO 4 was removed by centrifugation. The pH of the filtrate was adjusted as described above. The solution containing acid-sugar was applied to a column of a AG 1-X8 resin (200 -400 mesh, formate form) (Bio-Rad). The column was washed thoroughly with water and developed with 600 ml of a gradient of 0 -0.6 M formic acid. Fractions containing acidsugar (detected by HPLC) were combined and lyophilized to yield the corresponding lactone-sugar. Acid-sugar was obtained by base hydrolysis of the lactone-sugar, according to the method of Yew et al. (13). 1 Fig. 5C). Those of other acid-sugars were almost analogous to those reported by Yew et al. (13).
L-KDR was enzymatically synthesized from L-rhamnonate with AvLRA3 (L-rhamnonate dehydratase). The reaction mixture (100 ml) consisted of 50 mM sodium phosphate buffer (pH 7.0), 10 mM L-rhamnonate, 5 mM MgCl 2 , and 10% (v/v) glycerol. After the addition of 50 mg of AvLRA3, the mixture was left at 30°C overnight. The solution was filtered and then applied to a AG 1-X8 resin column followed by a 0 -0.6 M gradient of formic acid, and fractions containing L-KDR (detected by HPLC) were combined and lyophilized. L-and D-lactaldehyde were chemically synthesized from L-and D-threonine, respectively, according to the method of Huff and Rudney (27).
Enzyme Assays-All enzyme assays were performed at 30°C. L-Rhamnose 1-Dehydrogenase Assays-L-Rhamnose 1-dehydrogenase activity was assayed routinely in the direction of L-rhamnose oxidation by measuring the reduction of NAD(P) ϩ at 340 nm at 30°C using a Jasco spectrophotometer model V-550 (Japan Spectroscopic Co., Ltd., Tokyo, Japan). The standard assay mixture contained 10 mM L-rhamnose in 100 mM Tris-HCl (pH 9.0) buffer. The reaction was started by the addition of 10 mM NAD(P) ϩ solution (100 l) with a final reaction volume of 1 ml.
L-Rhamnono-␥-lactonase Assay-L-Rhamnono-␥-lactone hydrolase (lactonase) activity was measured by the modified method of Kondo et al. (28), which is based on the change in absorbance at 405 nm of the pH indicator p-nitrophenol caused by free acid formation from lactone-sugar. The reaction mixture contained 10 mM Pipes-NaOH (pH 6.4), 10 mM L-rhamnono-␥-lactone, and 0.25 mM p-nitrophenol. The substrate solution of lactone-sugar was freshly prepared immediately before the assay.
L-Rhamnonate Dehydratase Assay-L-Rhamnonate dehydration activity was monitored by the semicarbazide method (end point detection after 10 min) (29). The total reaction mixture of 400 l was incubated at 30°C in 50 mM sodium phosphate (pH 7.0) containing 10 mM L-rhamnonate, 5 mM MgCl 2 , 10% (v/v) glycerol, and a small volume of enzyme. After 10 min, the enzyme reaction was stopped by adding 100 l of 2 M HCl. To this solution, 300 l of semicarbazide solution consisting of 1.0% (w/v) semicarbazide hydrochloride and 1.5% (w/v) sodium acetate was then added, and the mixture was incubated at 30°C for 10 min. Finally, the reaction mixture was diluted with 500 l of distilled water, and its absorbance was measured at 250 nm (molar absorption coefficient ⑀ ϭ 571 M Ϫ1 ⅐cm Ϫ1 ). For the continuous assay, 50 mM sodium phosphate (pH 7.0), containing 10 mM L-rhamnonate, 5 mM MgCl 2 , 10% (v/v) glycerol, 1 unit of PsLRA4 (L-KDR aldolase, see below), and 1 unit of L-lactate dehydrogenase (LDH) (from bovine heart, Sigma) was used, and the reaction was started by the addition of 1 mM NADH solution (100 l) with a final reaction volume of 1 ml.
L-KDR Aldolase Assay-L-KDR aldolase activity in the aldol cleavage direction was continuously assayed in 50 mM sodium phosphate buffer (pH 7.0) containing 10 mM L-KDR, 0.5 mM MnCl 2 , and 1 unit of LDH. The reaction was started by the addition of 1 mM NADH solution (100 l) with a final reaction volume of 1 ml. For activity in the aldol condensation direction, the enzyme (ϳ0.5 mg/ml) was measured in 50 mM sodium phosphate buffer (pH 7.0) containing 0.5 mM MnCl 2 , 10 mM pyruvate, and 10 mM L-lactaldehyde.
To estimate the effect of metal ions on the activity of LRA4 proteins, aldol cleavage direction with L-KDR as a substrate was measured in the absence or presence of each metal chloride (0.5 mM). An assay in the presence of Zn 2ϩ , L-lactaldehyde dehydrogenase of P. stipitis 3 was used as a coupling enzyme instead of LDH, because of the significant inhibition of LDH activity (30).
Purification and Identification of Native L-Rhamnose 1-Dehydrogenase from P. stipitis-P. stipitis cells grown on L-rhamnose were harvested by centrifugation at 5,000 ϫ g for 10 min, washed with 50 mM Tris-HCl (pH 8.5) (referred to as Buffer A), and stored at Ϫ35°C until use. The washed cells were suspended in Buffer A, homogenized with an equal volume of glass beads (0.5 mm diameter, Sigma) for 30 min with appropriate intervals on ice using TORNADO Laboratory Power Mixer (AS ONE Co., Ltd., Osaka, Japan), and then centrifuged at 150,000 ϫ g for 1 h at 4°C to obtain cell-free extracts. All purification steps were performed below 4°C. All chromatography was carried out using an Á KTA purifier system (Amersham Biosciences) and/or BioAssist eZ system (TOSOH). The cellfree extract was loaded onto a column of HiPrep 16/10 Q FF (1.6 ϫ 10 cm, Amersham Biosciences) equilibrated with Buffer A and washed thoroughly with the same buffer. The column was developed with 300 ml of a linear gradient of 0 -0.5 M NaCl in Buffer A. Active fractions containing L-rhamnose 1-dehydrogenase were combined and concentrated by ultrafiltration with Centriplus YM-30 (Millipore) at 18,000 ϫ g for ϳ2 h. The enzyme solution was loaded onto a column of HiLoad 26/60 Superdex 200 pg (2.6 ϫ 60 cm; Amersham Biosciences) equilibrated with Buffer A. The active fractions were pooled, concentrated, and applied to a column of ceramic hydroxyapatite type I (1.6 ϫ 5 cm, Bio-Rad), equilibrated with Buffer A. The column was washed thoroughly with the same buffer and developed with 150 ml of linear gradient 0 -0.3 M potassium phosphate in Buffer A. Fractions with high enzymatic activity were combined, concentrated, and loaded onto a column of HiLoad 26/60 Superdex 200 pg (1.6 ϫ 60 cm, Amersham Biosciences), equilibrated with Buffer A. The active fractions were pooled and concentrated and run on native-PAGE with 6% (w/v) gel that was then soaked for 1 h at 30°C in a Zymogram staining solution (31) consisting of 100 mM Tris-HCl (pH 9.0), 10 mM L-rhamnose, 0.25 mM nitro blue tetrazolium, 0.06 mM phenazine methosulfate, and 15 mM NAD ϩ at 30°C for 30 min with gentle shaking. Dehydrogenase activity appeared as a dark band. The active band that appeared in staining was cut out and broken down in 50 mM Tris-HCl (pH 9.0) containing 1% (w/v) SDS. Protein was extracted from the gel by vigorous mixing overnight. A 4-fold volume of acetone chilled at Ϫ35°C was then added to the extracts. After cooling for 1 h at Ϫ35°C, the mixture was centrifuged at 39,120 ϫ g for 15 min at 4°C. The sample was dissolved in a small volume of SDS-PAGE sample buffer (500 mM Tris-HCl (pH 6.8), containing 5% (w/v) SDS, 10% (v/v) glycerol, 0.25% (w/v) bromphenol blue, and 5% (v/v) 2-mercaptoethanol) and separated by SDS-PAGE with 10% (w/v) gel. This procedure revealed that a polypeptide with a molecular mass of ϳ25 kDa corresponded to L-rhamnose 1-dehydrogenase (see lane 6 in Fig. 3A); therefore, this sample was used for internal peptide fragment analysis by a MALDI quadrupole ion trap mass spectrometer (MALDI-QIT-TOF MS) (AXIMA QIT, Shimadzu, Kyoto, Japan), according to the standard protocol (32).
Identification of Putative Gene Cluster Related to Sugar Metabolism-Metabolic genes involved in several sugar pathways analogous to the ED pathway were used as a probe (supplemental Table SI). A preliminary homology search using the Fungal Genomic BLAST program distributed by the National Center for Biotechnology Information (NCBI) was carried out against the genome sequences of P. stipitis and D. hansenii using the probe protein sequences described above. The significant homologous candidate gene was further examined by estimating whether enzymes belong to the same protein family as probe proteins in the flanking region. Further analysis was carried out against the known genome sequences of microorganisms using amino acid sequences of PsLRA1-4 as a query sequence. In this study, the prefixes Ps (P. stipitis), Dh (D. hansenii), and Av (A. vinelandii) have been added to gene symbols or protein designations when required for clarity.
Functional Expression and Purification of His 6 -tagged Proteins-LRA1-4 genes from P. stipitis, D. hansenii, and A. vinelandii were amplified by PCR using primers containing appropriate restriction enzyme sites at the 5Ј-and 3Ј-ends and genome DNA as a template (supplemental Table S2). Each amplified DNA fragment was introduced into BamHI-PstI sites (for all others except PsLRA2 and PsLRA4 genes) or BamHI-SalI (for PsLRA2 and PsLRA4 genes) in pQE-80L (Qiagen), a plasmid vector for conferring N-terminal His 6 tag on expressed proteins. Escherichia coli DH5␣ harboring the expression plasmid for His 6 -tagged enzymes was grown at 37°C to a turbidity of 0.6 at 600 nm in Super broth medium (pH 7.0, 12 g of tryptone, 24 g of yeast extract, 5 ml of glycerol, 3.81 g of KH 2 PO 4 , and 12.5 g of K 2 HPO 4 per liter) containing 50 mg/liter ampicillin. After the addition of 1 mM of isopropyl-␤-D-thiogalactopyranoside, the culture was further grown for 6 h to induce the expression of His 6 -tagged protein. Cells were harvested and resuspended in Buffer X (50 mM sodium phosphate buffer (pH 7.0) containing 300 mM NaCl and 10 mM imidazole). The cells were then disrupted by sonication, and the solution was centrifuged. The supernatant was loaded onto a nickel-nitrilotriacetic acid Superflow column (Qiagen) equilibrated with Buffer X. The column was washed with Buffer Y (pH 7.0, Buffer X containing 10% (v/v) glycerol and 50 mM imidazole instead of 10 mM imidazole). The enzymes were then eluted with Buffer Z (pH 7.0, Buffer Y containing 250 mM imidazole instead of 50 mM imidazole). Five mM MgCl 2 (for AvLRA3, PsLRA4, and AvLRA4) and 10% (w/v) glycerol (for AvLRA3) were added to the buffer system.

Metabolic Genes Related to Fungal L-Rhamnose Metabolism-
Phylogenetic analysis of the equivalent reaction step catalyzing metabolic enzymes is helpful to estimate the evolutionary relationship between sugar pathways analogous to the ED pathway. Classification of these metabolic enzymes based on the "Cluster of Orthologous Groups of proteins" (COG) is summarized in Fig. 1. On the other hand, metabolic genes often form a single gene cluster on the genomes of bacteria and Archaea (supplemental Fig. S1). Based on these insights, we searched for significant homologs to the known metabolic genes on the genomes of microorganisms. We particularly focused on fungal P. stipitis and D. hansenii, because it is believed that there is an alternative pathway for L-rhamnose metabolism in these microorganisms (33,34). The schematic conversion of L-rhamnose is different from that of the known bacterial pathway, depending on protein products of the rhaDABSRT operon (35) (Fig. 2, A and B), and completely analogous to nonphosphorylative sugar pathways (type I), although no gene encoding four metabolic enzymes has been identified so far. As a result, we identified a potential gene cluster related to the sugar metabolic pathway on the genomes of two fungi consisting of LRA1-4 genes. Further bioinformatic analysis revealed that the (partially) homologous gene cluster (together with other putative sugar-related genes) was found on the genomes of not only many fungi (supplemental Fig. S2A) but also several bacteria, including A. vinelandii ( Fig. 2E and supplemental Fig. S2B). A. vinelandii can grow on L-rhamnose as a sole carbon source (data not shown), and interestingly, AvLRA4 is not related to the fungal LRA4(s) (see below in detail); therefore, we expressed LRA1-4 proteins of P. stipitis, D. hansenii, and/or A. vinelandii in E. coli cells. Although several expression levels and/or solubility problems were often found, we were successful in homogeneously purifying PsLRA1, DhLRA1, AvLRA1, DhLRA2, AvLRA3, PsLRA4, and AvLRA4 as His 6 -tagged enzymes (see Fig. 3B, Fig. 4A, Fig. 5A, and Fig. 6A).
L-Rhamnose 1-Dehydrogenase-LRA1 was homologous to D-glucose 1-dehydrogenase of Bacillus megaterium (E11, see Ref. 36) and D-xylose 1-dehydrogenase of Caulobacter crescentus (E66, Ref. 11), which belong to the short-chain dehydrogenases/reductases family (COG1028): 35 and 29% of identity with PsLRA1, respectively. To preliminarily estimate the physiological role of the gene cluster, we attempted to biochemically characterize a dehydrogenase with L-rhamnose in P. stipitis. When compared with nutrient medium (0.056 unit/mg protein), approximately 6-fold higher activity of NAD ϩ -dependent dehydrogenation with L-rhamnose was found in the cell-free extract from P. stipitis cells grown on L-rhamnose as a sole carbon source (0.33 unit/mg protein), and appeared as a single active band in zymogram-staining analysis (data not shown). L-Rhamnose 1-dehydrogenase was (partially) purified by four chromatographic steps (Fig. 3A) and showed the NAD ϩ -dependent specific activity of 22 Table S1), which are referred to as En in the text.

Novel L-Rhamnose Metabolic Pathway
result of purification is summarized in Table 1. No NADP ϩ -dependent activity was found in the purified enzyme. MALDI-QIT-TOF MS analysis revealed that the enzyme is identical to a putative D-glucose 1-dehydrogenase of P. stipitis CBS6054 (ABN68405), which corresponds to the LRA1 gene involved in the gene cluster with 56% of sequence coverage (supplemental Fig. S3 and supplemental Table S3). These results indicated that the LRA1 gene encodes NAD(P) ϩ -dependent L-rhamnose 1-dehydrogenase (EC 1.1.1.173) ( Fig. 2A) and that the remaining LRA2ϳ3 genes may also be related to L-rhamnose metabolism.
Among 16 sugars (listed in Table 2), significant sugar dehydrogenase activities in PsLAR1, DhLRA1, and AvLAR1 were commonly observed with L-rhamnose, L-lyxose, L-mannose, and L-fucose, indicating that the same configuration at C-2, C-3, and C-4 may be important for activity. The catalytic efficiency (k cat /K m ) value with L-rhamnose was much higher between active substrates because of the lowest values of K m and the highest values of k cat , in the order of L-rhamnose Ͼ L-lyxoseϾ L-mannose Ͼ L-fucose (Fig. 3C and Table 2). The most different property between three LRA1 enzymes was coenzyme specificity. As expected from the native enzyme, both PsLRA1 and DhLRA1 were strictly NAD ϩ -dependent enzymes, and their activity in the presence of NADP ϩ (using L-rhamnose as a substrate) was less than 1% that in the presence of NAD ϩ . On the other hand, significant preference for NADP ϩ over NAD ϩ was found in AvLRA1; the k cat /K m value with L-rhamnose in the presence of NADP ϩ (2140 min Ϫ1 ⅐mM Ϫ1 ) was 2.5-fold higher than in the presence of NADP ϩ (856 min Ϫ1 ⅐mM Ϫ1 ), mainly because of the higher k cat value. When compared with other short-chain dehydrogenases/reductase enzymes, essential amino acid residues for coenzyme binding (Gly-X 3 -Gly-X-(Gly/Ala)) and catalytic triad (Ser-Tyr-Lys) were conserved in L-rhamnose 1-dehydrogenases: Gly 14 -Gly-Val-Thr-Gly-Ile-Gly 20    the significant modifications of amino acid residues that directly interact with 2Ј-and 3Ј-functional groups in the ribosyl moiety of NAD(P) ϩ (Fig. 3D), confirming the different coenzyme specificity between fungal and bacterial LAR1 enzymes.

DISCUSSION
Evolutionary Relationships between the L-Rhamnose and Other Sugar Metabolic Pathways-There is significant phylogenetic mosaicism between the metabolic enzymes involved in analogous sugar pathways to the ED pathway (Fig. 1), strongly indicating that these pathways did not evolve from a single ancestral pathway. In fact, the combination of these protein families in the alternative L-rhamnose pathway is completely different from the known sugar pathways. On the other hand, the numbers of protein families belonging to these metabolic enzymes are limited, suggesting that analogous sugar pathways to the ED pathway evolved by the combination of a limited number of ancestral enzymes.
In archaeal S. solfataricus, both D-glucose and D-galactose are metabolized through the npED pathway (3)(4)(5). Furthermore, the E. coli mutant able to grow on L-lyxose possesses a mutated L-rhamnulokinase, which phosphorylates L-xylulose as efficiently as L-rhamulose, by which L-lyxose is metabolized through the L-rhamnose pathway via L-xylulose and L-xylulose 1-phosphate (46) (Fig. 2A). This indicates that different sugars, in particular with structural similarity, are metabolized through a (inherent) single pathway. Although P. stipitis, D. hansenii, and A. vinelandii also cannot grow on L-lyxose (data not shown), there is concomitant activity with the related intermediates in LRA1, LRA3, and LRA4; therefore, (small) modification of substrate specificities in LRA1-4, in particular LRA2, incapable of utilizing L-lyxono-␥-lactone, may lead to a novel ability to grow on L-lyxose.
Convergent Evolution of Metabolic Pathways-Despite the same L-rhamnose pathway, L-KDR aldolases of fungi and bacteria belong to different protein families, as well as three metabolic enzymes involved in the D-xylose pathway of bacteria (E66 -E68) and Archaea (E71-E73): convergent evolution of the same pathway in different domains. On the other hand, we previously identified and characterized ␣-ketoglutaric semialdehyde dehydrogenases involved in L-arabinose and hexaric acid pathways (type II) of bacteria (E43, E46, E51, and E56). They are commonly members of the aldehyde dehydrogenase superfamily (COG1012), but classified into three subgroups in the phylogenetic tree, indicating the independent acquisition of substrate specificity for ␣-ketoglutaric semialdehyde rather than divergence from a common ancestor (8,21). Similar phenomena are found in bacterial D-glucose 1-dehydrogenases (E8, E9 and E11) and L-2-keto-3-deoxyarabonate dehydratases (E50, E55, and E60). This suggests that even the same pathway in the same domain evolved independently, at least partially.
Over 20 years after the discovery of L-rhamnose degradation in fungi (33,34), the metabolic enzymes and intermediates involved in the pathway can now be understood at the molecular level. Gerlt and co-workers (13,40,41) have identified several novel acid-sugar dehydratases by an unique approach, in which the following three steps are carried out: 1) selection of a putative mandelate racemase enzymes (COG0148) on the bacterial genomes; 2) purification of the recombinant enzyme; and 3) estimation of substrate specificity using 52 chemically synthesized acid-sugars. Although they also mention that the acidsugar dehydratase gene is often clustered with other genes probably involved in the sugar metabolic pathway, this approach may be applicable only for a pathway containing mandelate racemase-type acid-sugar dehydratase. We previously revealed that L-arabonate dehydratase (E49) belongs to the dihydroxyacid dehydratase (ILVD) family (COG0129), another type of acid-sugar dehydratase, together with 6-phosphogluconate dehydratase (E3) (14). Therefore, the combination of significant insights about the metabolic gene illustrated in Fig. 1 and the approach by Gerlt and co-workers (13,40,41) will also be helpful to identify unknown sugar pathways analogous to ED and npED pathways containing either type of acidsugar dehydratases, even if the phenotype of the microorganism is not available.