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Structure and function of aldopentose catabolism enzymes involved in oxidative non-phosphorylative pathways

Biotechnology for Biofuels and Bioproducts volume 15, Article number: 147 (2022) Cite this article

Abstract

Platform chemicals and polymer precursors can be produced via enzymatic pathways starting from lignocellulosic waste materials. The hemicellulose fraction of lignocellulose contains aldopentose sugars, such as d-xylose and l-arabinose, which can be enzymatically converted into various biobased products by microbial non-phosphorylated oxidative pathways. The Weimberg and Dahms pathways convert pentose sugars into α-ketoglutarate, or pyruvate and glycolaldehyde, respectively, which then serve as precursors for further conversion into a wide range of industrial products. In this review, we summarize the known three-dimensional structures of the enzymes involved in oxidative non-phosphorylative pathways of pentose catabolism. Key structural features and reaction mechanisms of a diverse set of enzymes responsible for the catalytic steps in the reactions are analysed and discussed.

Background

Excessive exploitation of fossil resources and severe deterioration of the natural environment have become serious challenges for further development of human society, and has led to intense exploration for alternative sources of energy, raw materials and food [1]. Lignocellulose offers an alternative raw material that is abundant, cheap and renewable [2]. In particular utilization of various types of lignocellulosic waste, such as from agriculture, forestry, agroindustry and municipalities, as feedstock can promote resource efficiency and circular economic goals without affecting the food supply [3]. Industrial biotechnology offers sustainable approaches to converting lignocellulosic wastes to fuels, chemicals and materials. To be economically feasible, all fractions of lignocellulosics (the major components being cellulose, hemicellulose and lignin) need, however, to be considered. Concerning sugars, the most common monosaccharide found in plant biomasses is d-glucose, a 6-carbon (C6) hexose sugar, which is a structural component of cellulose [4]. The second most abundant sugar is a 5-carbon (C5) pentose sugar, d-xylose, which is especially abundant in xylan-rich hemicelluloses [5, 6]. l-arabinose, another type of C5 pentose sugar, is found in plant polysaccharides, hemicelluloses and pectin.

Efficient biotechnological use of lignocellulosic waste will require microorganisms (such as bacteria or yeast) that possess metabolic pathways for utilizing biomass-derived monosaccharides. Currently, most biotechnological processes rely on d-glucose as the primary carbon source, but utilization of abundant biomass-derived pentose sugars is also of importance [7, 8]. Three main microbial pathways for pentose sugar catabolism, including d-xylose, l-arabinose and d-arabinose, have been described [9]. In the first pathway, mainly found in bacteria, the conversion uses isomerases, kinases and epimerases to produce d-xylulose-5-phosphate, which then can be further utilized by the pentose phosphate pathway [10]. In the second pathway, commonly found in yeast and fungi, pentoses are metabolized by reductases, dehydrogenases and kinases to also produce d-xylulose-5-phosphate [11]. The third pathway of pentose sugar catabolism, found in archaea and bacteria, is an oxidative, non-phosphorylative pathway, originally discovered by Weimberg in Pseudomonas fragi [12]. Here, pentose sugars are converted through 2-dehydro-3-deoxy-aldopentonate to α-ketoglutarate, which is an intermediate in the citric acid cycle (Fig. 1). In the oxidative Dahms pathway [13], a pentose sugar is converted through 2-dehydro-3-deoxy-aldopentonate to glycolaldehyde and pyruvate (Fig. 1). The homologous pathway of Dahms pathway is also described in the catabolism of deoxyhexoses, for example, l-rhamnose and l-fucose [14, 15]. Furthermore, Weimberg/Dahms pathways are analogous to the non-phosphorylative Entner–Doudoroff (ED) pathway for hexose sugars. These non-phosphorylative metabolic pathways have the potential for efficient production of high-value bioproducts as an alternative to more traditional metabolic pathways, which consist of more steps and have more complex regulation that limits production yields and rates. Lately, Watanabe et al. found a third non-phosphorylative route of pentose metabolism in Herbaspirillum huttiense IAM 15032 [16] by analysis of gene clustering, in which the intermediate 2-dehydro-3-deoxy-aldopentonate is converted to glycolate and pyruvate via 5-hydroxy-2,4-dioxo-pentanonate. The gene cluster analysis has an increasing role in the discovery of new routes and novel enzymes.

Fig. 1
figure 1

a General scheme of oxidative non-phosphorylative pathways for catabolism of pentose sugars. Weimberg and Dahms pathway enzymes are shown in blue and green, respectively, and the collective upstream steps in the pathways are shown in grey. b Enzymes involved in Weimberg/Dahms catabolism of d-xylose

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In the Weimberg pathway (Fig. 1), a pentose sugar is first oxidized to a lactone by an aldopentose-1-dehydrogenase enzyme, followed by the lactone ring being opened to a sugar acid by a pentonolactonase enzyme. Two sequential dehydration steps then follow: in the first step, an aldopentonate dehydratase converts the sugar acid to 2-dehydro-3-deoxy-aldopentonate, which is then converted to 2,5-dioxopentanoate by another dehydratase. In the final reaction step of the Weimberg pathway, 2,5-dioxopentanoate is converted to α-ketoglutarate by α-ketoglutarate-semialdehyde dehydrogenase enzyme. In the Dahms pathway, the 2-dehydro-3-deoxy-aldopentonate is directly split into pyruvate and glycolaldehyde by an aldolase enzyme, while the upstream steps in the pathway are in common with the Weimberg pathway.

Recent studies have increasingly focused on metabolic engineering of the Weimberg/Dahms pathways for production of biobased compounds, i.e., glycolic acid, glutaric acid, ethylene glycol, 1,2,4-butanetriol and 1,4-butanediol, which can be used in the textile, food, pharmaceutical and chemical industries, among others [3, 8, 17,18,19,20]. In this review, we provide an overview of the structural characteristics of Weimberg/Dahms pathway enzymes and describe what is known of their catalytic mechanisms and enzyme families.

Aldopentose-1-dehydrogenases

The first step in the Weimberg/Dahms pathways is the oxidation of a pentose sugar to the corresponding lactone by aldopentose-1-dehydrogenases. In principle, l-arabinose is oxidized by l-arabinose dehydrogenase (E.C. 1.1.1.46 for the NAD- or 1.1.1.376 for the NAD/NADP-dependent forms), d-arabinose by d-arabinose dehydrogenase (E.C. 1.1.1.116 for the NAD- or 1.1.1.117 for the NADP-dependent forms) and d-xylose by d-xylose dehydrogenase (E.C. 1.1.1.175 for the NAD- and 1.1.1.179 for the NADP-dependent forms). However, many aldose-1-dehydrogenases have wide substrate specificity, and they can act on various aldopentoses as well as aldohexoses (E.C. 1.1.1.121). The general term aldose-1-dehydrogenase is used when enzymes utilize NAD or NADP as cofactors to oxidize aldoses to aldonolactones.

At present, there are only a few crystal structures of aldose-1-dehydrogenases with preference for pentose sugars available from the Protein Data Bank (PDB) (Table 1). These aldose-1-dehydrogenases belong to either the dehydrogenase/reductase superfamily or the Gfo/Idh/MocA superfamily.

Table 1 Available crystal structures of aldose-1-dehydrogenases acting on pentose sugars
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Dehydrogenase/reductase superfamily

The dehydrogenase/reductase superfamily is a very large and versatile superfamily of NAD(P)-dependent proteins with high sequence diversity and a wide range of catalytic activities [27]. The first solved protein structure of the family, described in 1976, was that from a horse liver alcohol dehydrogenase [28]. Based on polypeptide chain length, distinct sequence motifs, and structural comparisons, the dehydrogenase/reductase superfamily can be further divided into short-, medium- and long-chain dehydrogenase/reductase superfamilies (SDR, MDR and LDR, respectively) [29]. d-arabinose dehydrogenase from S. solfataricus [30] as well as l-arabinose dehydrogenase from the bacterial species R. leguminosarum [31], and a d-xylose dehydrogenase from the archaean species H. marismortui [32],belong to the MDR superfamily. Conversely, d-xylose dehydrogenase of the bacterial species C. crescentus [33] and an l-arabinose dehydrogenase of the archaean species H. volcanii [34] are reported to belong to the SDR superfamily based on sequence comparisons. No X-ray structures of pentose dehydrogenases belonging to the SDR superfamily have been published to date. However, there are some structures (unbound and ligand bound) of a deoxyhexose dehydrogenase of SDR family, the l-rhamnose-1-dehydrogenase from Azotobacter vinelandii [35]. SDR- and MDR-family pentose dehydrogenases belong to a branch of alcohol/polyol/sugar dehydrogenases along with the alcohol dehydrogenases, sorbitol dehydrogenases and aldohexose dehydrogenases [36].

A crystal structure of the d-arabinose dehydrogenase from S. solfataricus (hereafter referred to as Ss ADH; PDB code 2H6E) was determined in 2007 [21]. The year before, the X-ray structure of a glucose dehydrogenase, also from S. solfataricus (hereafter referred to as Ss GDH), had been solved in apo and complex forms (PDB codes 2CD9, 2CDA, 2CDB, and 2CDC). Ss GDH catalyzes the oxidation of glucose to gluconate in the non-phosphorylated ED pathway, but it is also able to act on d-galactose, d-xylose and l-arabinose [37]. Ss GDH shares only 19% sequence identity with Ss ADH, but the enzymes are structurally and functionally very similar.

D-arabinose dehydrogenase Ss ADH consists of 344 amino acids, and it folds into a catalytic domain (residues 1–154 and residues 292–344), and a nucleotide binding domain (residues 155–291) (Fig. 2a) [21]. MDR superfamily proteins typically have these two domains, where the N-terminal domain is responsible for substrate binding and the C-terminal domain with a Rossmann fold is responsible for nucleotide binding. The catalytic domain of Ss ADH has an N-terminal segment composed of two α-helices (α1, α2), four 310-helices (G1–G4), and nine β-strands (β1–β9). The C-terminal segment is composed of α-helix α9 (and a predicted additional α10) along with two β-strands (β17, β18). The core of the catalytic domain of Ss ADH is formed by five antiparallel β-strands (β4, β6–β9) and two parallel β-strands (β17, β18). The C-terminal nucleotide binding domain of Ss ADH has a classical Rossmann fold comprised of a six-stranded parallel β-sheet (β10–β16) surrounded by six α-helices (α3–α8). The overall fold is very similar to that of Ss GDH and other MDR dehydrogenases. The quaternary structure of Ss ADH is a homotetramer (Fig. 2b), which is a common quaternary structure of MDR enzymes, but monomers, dimers and trimers have also been observed among MDR family members [38]. The structurally related glucose dehydrogenase Ss GDH is a homotetramer [22].

Fig. 2
figure 2

Crystal structure of d-arabinose dehydrogenase from S. solfataricus (PDB: 2H6E). a Overall fold is represented by a ribbon diagram. The catalytic domain is shown in green, where the N-terminal and C-terminal segments of the catalytic domain are shown in light and dark green, respectively. The nucleotide binding domain is shown in yellow. Zn2+ ion is shown as a purple sphere. The dashed line represents a missing part of the crystal structure due to weak electron density (here reconstructed by homology modelling). b Tetrameric quaternary structure generated by crystallographic symmetry operations. c Coordination geometry of the catalytic Zn2+ ion. d Coordination geometry of the structural Zn.2+ ion. (All structural figures in this article were prepared using Pymol [40] software)

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In the catalytic domain, Ss ADH has two Zn2+ ions, one being catalytic and the other structural. The catalytic Zn2+ ion is tetrahedrally bound by Cys41, His65, Asp150 and a water molecule (Fig. 2c). The Cys and His residues are well-conserved among MDR family members, but the third ligand can be a Glu or Cys residue. In Ss GDH, Cys69 and His66 residues, along with Glu67 from the β7-strand and a water molecule, are coordinated to the Zn2+ ion. In addition to the catalytic Zn2+ ion, there is the structural Zn2+ ion, which is bound to Asp94, Cys97, Cys100 and Cys108 in Ss ADH (Fig. 2d), similar to that in alcohol dehydrogenases. The structural Zn2+ ion stabilizes the loop, which protrudes from the catalytic domain. It has been observed that removal of the structural Zn2+ ion in yeast alcohol dehydrogenase results in decreased thermostability [39].

The active site of MDR proteins is located in a deep crevice between the two domains that accommodates the NAD(P) and substrate. At the moment, there is no available complex structure of Ss ADH or any other aldopentose-1-dehydrogenase of the dehydrogenase/reductase superfamily but the structure of Ss GDH complexed with d-glucose or d-xylose has been solved [22]. Based on structural similarities, Ss ADH most likely operates via the same reaction mechanism as that observed for Ss GDH. The aldose sugar in the pyranose form is oxidized to aldonolactone, when a hydride from C1 of aldose is transferred to C4 of the nicotinamide ring, which is facilitated by proton abstraction from the hydroxyl group at C1 of the sugar ring. Structures of Ss GDH complexed with d-glucose or d-xylose were observed in the chair conformation and found to exist in the β-anomer, which allows C1 of the pyranose ring to be close to C4 of nicotinamide ring. d-arabinose dehydrogenase converts sugars with opposite stereo configuration at C2 and C3 (2S,3R in d-arabinose) as compared to d-glucose dehydrogenase (2R, 3S in d-glucose, d-xylose and l-arabinose), and therefore, it has been proposed that the pyranose ring needs to be flipped by 180° in Ss ADH [21], but this proposal still needs to be confirmed by the structure of ADH complexed with d-arabinose. It is also unclear whether an aldose substrate interacts with the Zn2+ ion via coordinated water or whether the C1 hydroxyl group of the substrate can form a covalent bond with Zn2+, making it penta-coordinated.

Gfo/Idh/MocA superfamily

Proteins of the Gfo/Idh/MocA superfamily are NAD(P)-dependent oxidoreductases that act on a diverse set of substrates. The family name refers to glucose–fructose oxidoreductases, inositol 2-dehydrogenases, and the rhizopine catabolism protein MocA, but it also includes a growing number of other oxidoreductases [41]. The 6-phosphate glucose dehydrogenase from Leuconostoc mesenteroides was the first solved protein structure of the Gfo/Idh/MocA superfamily [42].

In 2019, the crystal structure of an l-arabinose dehydrogenase from the bacterial species A. brasiliensis (hereafter referred to as Ab ADH) was solved with and without the cofactor NADP (PDB codes 6JNK and 6JNJ) [23]. Very recently, its structure in a complex with l-arabinose (PDB code 7CGQ) was also published [24]. In addition, the crystal structure of a d-galactose-1-dehydrogenase from the bacterial species R. etli (hereafter referred to as Re GDH; PDB code 4EW6), has been solved by the Structural Genomics Research Consortium [25]. Re GDH is thought to be involved in galactose catabolism, but interestingly the orthologous enzyme from Rhizobium leguminosarum bv. trifolii was named l-arabinose/d-galactose-1-dehydrogenase due to its high catalytic efficiency toward l-arabinose, d-galactose and d-fucose [31]. The sequence identity of Re GDH with l-arabinose/d-galactose-1-dehydrogenase from R. leguminosarum is as high as 96%. The sequence identity between Ab ADH and Re GDH is 57%. The Gfo/Idh/MocA superfamily also contains an aldose–aldose oxidoreductase from C. crescentus (Cc AAOR), which was discovered through its sequence homology to the xylose dehydrogenases [43]. Cc AAOR has been shown to catalyze the concomitant oxidation and reduction of d-xylose and other aldose monosaccharides to the corresponding aldonolactones or alditols, respectively. Cc AAOR contains a tightly bound NADP cofactor, which is regenerated in this oxidation–reduction cycle. Several crystal structures of Cc AAOR have been solved including those complexed with d-xylose and d-glucose [26]. However, the sequence identity between Ab ADH and Cc AAOR is only 30%. Cc AAOR has been involved in co-production of d-xylonate and xylitol from d-xylose in Saccharomyces cerevisiae [44].

The L-arabinose dehydrogenase Ab ADH has the typical two-domain structure of Gfo/Idh/MocA family proteins (Fig. 3a). The N-terminal nucleotide-binding domain has a classical Rossmann fold with six-stranded parallel β-sheet (β1–β6) surrounded by five α-helices (kinked α1, α2, α3, α4 and α5) and one short 310-helix G1. The C-terminal domain has a two-layer α/β-sandwich fold consisting of an eight-stranded β-sheet (β7–β14) and eight helices (five α-helices, α6–α10, and three 310-helices, G2–G4). The C-terminal domain participates in oligomerization. Both Ab ADH and galactose dehydrogenase Re GDH shows structural similarity with Cc AAOR and glucose-6-phosphate dehydrogenases. Ab ADH and Re GDH exist as a homodimer, which is characteristic of Gfo/Idh/MocA proteins (Fig. 3b). Interestingly, Cc AAOR has a different dimerization mode compared with Ab ADH and Re GDH. In Cc AAOR, the β-sheets of two monomeric units are tightly packed together, and this dimerization creates an α–β–β–α sandwich-type structure at the dimeric interface.

Fig. 3
figure 3

Crystal structure of L-arabinose dehydrogenase from A. brasiliensis (PDB: 7CGQ). a Overall fold is represented by a ribbon diagram. The N-terminal domain is shown in yellow, and the C-terminal domain in green. b Quaternary structure is a homodimer. c NADP-binding site is located in the crevice between the two domains. d Amino acid residues interacting with l-arabinose. e Hydrogen-bonding network around the O1 hydroxyl of l-arabinose. The distance from C1 of l-arabinose to C4 of the nucleotide is also shown

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The ternary complex of Ab ADH with L-arabinose and NADP shows that the active site is located in the crevice formed between the two domains (Fig. 3c) [24]. NADP is bound by three loops (β1–α1, β2–β3 and β4–α3) and one α-helix (α3) of the N-terminal domain. The region binding the 2′-phosphate is highly positively charged in Ab ADH, where in some NAD-preferred Gfo/Idh/MocA family proteins, this region contains hydrophobic or negatively charged residues [45,46,47]. l-arabinose is observed to interact through hydrogen bonding with Lys91, His119, Glu147, Trp152, His153, Asp169, Asn173 and Trp231 (Fig. 3d). The oxidation mechanism of Gfo/Idh/MocA proteins is known to be involved in the transfer of hydride from the C1 carbon of the substrate to the C4 carbon of the nicotinamide ring, coupled with deprotonation of the O1 hydroxyl of the substrate by a catalytic base. In general, a tyrosine or histidine residue functions as the catalytic base. In Ab ADH, three residues Lys91, His119 and Asn173, are located close to the C1 hydroxyl group of l-arabinose (Fig. 3e). Site-directed mutagenesis studies of Ab ADH showed that the H119N mutant retained its activity, but the activity of the N173Y and N173H mutants were reduced by three orders of magnitude compared to the wild type [24]. These results suggested that Asn173 in Ab ADH plays an important role in catalysis.

Pentonolactonases

In the second step of the Weinberg/Dahms pathway, pentonolactone is hydrolyzed to pentoic acid. d-xylonolactone is catalyzed by xylonolactonase (E.C. 3.1.1.68), and l-arabinolactone by arabinolactonase (E.C. 3.1.1.15). This intramolecular ester bond hydrolysis step proceeds spontaneously but slowly at ambient temperature in vitro, although spontaneous lactone hydrolysis might proceed over a more reasonable timescale at elevated temperatures. The hyperthermophilic Sulfolobus solfataricus, for example, lacks an aldonolactonase gene in the pentose degradation gene cluster [30]. However, two genes encoding putative lactonases have been identified elsewhere in the S. solfataricus genome, but the involvement of these genes in pentose degradation is not yet clear. Structural information of pentonolactones is very limited and only very recently have crystal structures of xylonolactonase from C. crescentus complexed with d-xylose and 4-hydroxy-2-pyrrolidinone become available (PDB codes 7PLB, 7PLC and 7PLD) [48]. This enzyme, which is found in C. crescentus in the same operon as in the genes for the Cc XylB dehydrogenase and Cc XylD dehydratase enzymes, has been shown to improve formation of d-xylonic acid in in vitro enzyme cascade studies [49].

In addition to C. crescentus xylonolactonase [33], pentonolactonases have been characterized from Azospirillum brasilense [53] and Haloferax volcanii [54]. Based on sequence homology, pentonolactonases belong to the senescence marker protein (SMP30) superfamily, along with gluconolactonases, paraoxonases and luciferase regenerating enzymes. Available crystal structures of aldonolactonases belonging to the SMP30 superfamily are shown in Table 2. Gluconolactonase catalyses gluconolactone to gluconic acid [50], and SMP30 protein is reported to convert l-gulonate to l-gulonolactone in the vitamin C biosynthetic pathway, but also possess gluconolactonase activity [55]. In addition, SMP30 is sometimes called regucalcin due to its putative role in calcium homeostasis [56].

Table 2 Available crystal structures of aldonolactonases belonging to the SMP30 superfamily
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Xylonolactonase is folded into a beta-propeller consisting of six blades, each of which is formed by four beta-strands (Fig. 4a). The active site of the enzyme is located in the central cavity, where the divalent metal binding site exists at the bottom of the cavity. In the solved X. campestris gluconolactonase structure, the bound metal ion at the catalytic site was determined to be Ca2+ [50], but many studies have reported that Zn2+ would be responsible for gluconolactonase activity [51]. On the other hand, a very recent study of C. crescentus xylonolactonase shows that the enzyme binds only Fe2+ ion with high specificity and affinity, and the other divalent metal cations are suggested to assist non-enzymatic hydrolysis by stabilizing the short-lived bicyclic intermediate during isomerization of lactone [57]. The Fe2+ ion, located in the active site of the enzyme, is bound by three conserved residues: Glu18, Asn146 and Asp196 (Fig. 4b), but not by the highly conserved Asn101. Xylose is also coordinated to Fe2+ together with two water molecules, which complete the octahedral coordination.

Fig. 4
figure 4

Crystal structure of xylonolactonase from C. crescentus (PDB: 7PLB). a Overall fold is represented by a ribbon diagram. The Fe2+ ion is indicated by an orange sphere. b Coordination geometry of the Fe2+ ion

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The site of substrate entrance into the SMP30 lactonases is located at the top of the metal-binding site. Although the SMP30 proteins have a lid-like structure that partly covers the substrate entrance [51], no such lid exists in C. crescentus xylonolactonase [46] or X. campestris gluconolactonase [49]. The crystal structures of C. crescentus xylonolactonase complexed with d-xylose or the substrate analogue 4-hydroxy-2-pyrrolidinone reveal the binding mode of the substrates [48]. d-xylose was observed in xylopyranose form and the equatorial O1 was bound to the Fe2+ ion over a short distance of 2.0 Å. Similarly, the carbonyl oxygen of 4-hydroxy-2-pyrrolidinone was 1.9 Å away from the Fe2+ ion. The crystal structures of mouse SMP30 protein complexed with the substrate analogue xylitol (PDB code 4GNA) or the product analogues 1, 5-anhydro-d-glucitol (PDB code 4GN8) and d-glucose (PDB code 4GN9) are also available [52]. The divalent metal ion and polar residues in the active site pocket interact with hydroxyl groups of the substrates.

The quaternary structures of the published aldonolactonases are either monomeric or dimeric. C. crescentus xylonolactonase, human and mouse SMP30 proteins are reported to be monomers, but gluconolactonase from X. campestris has been shown to exist as a disulphide-bonded homodimer. One structural Ca2+ ion was found to be at the interface of the monomeric subunits in the dimer and was suggested to stabilize the dimeric form.

Aldopentonate dehydratases

In the third step of the Weimberg/Dahms pathways, an aldopentonate is converted to 2-dehydro-3-deoxy-aldopentonate by aldopentonate dehydratase. d-xylonate dehydratase (EC 4.2.1.82) converts d-xylonate to 2-dehydro-3-deoxy-d-xylonate, and l-arabinonate dehydratase (EC 4.2.1.25) converts l-arabinonate to 2-dehydro-3-deoxy-l-arabinonate by removing one water molecule. Currently, crystal structures of 2 homologous aldopentonate dehydratases, R. leguminosarum l-arabinonate dehydratase (hereafter referred to as Rl ADHT; (PDB codes 5J83, 5J84, 5J85) [58] and C. crescentus d-xylonate dehydratase (hereafter referred to as Cc XDHT; PDB code 5OYN) [59], have been solved. The 2 bacterial aldopentonate dehydratases have been shown to accept both pentonate and hexonate sugar acids as their substrates, being strictly stereospecific for the configuration of OH groups at C2 and C3 [60]. Rl ADHT is reported to have the highest catalytic efficiency (kcat/Km) for d-fuconate, followed by l-arabinonate and d-galactonate, while Cc XDHT prefers d-xylonate and d-gluconate. Metabolic engineering studies and in vitro enzyme cascade studies have indicated that the dehydratase reaction catalyzed by the Cc XDHT enzyme, which is a tetramer requiring a [2Fe–2S] cluster and Mg2+ ion for its activity [59,60,61], is a rate-limiting step in the Weimberg/Dahms pathways.

Rl ADHT and Cc XDHT enzymes belong to the iron–sulfur cluster-containing IlvD/EDD protein family, like many other aldopentonate dehydratases from bacterial species, such as A. brasiliense l-arabinonate dehydratase [53], E.coli D-xylonate dehydratase [67] and Pseudomonas putida d-xylonate dehydratase [68]. By contrast, aldopentonate dehydratases of the archaean species, such as d-arabinonate dehydratase from Sulfolobus solfataricus [30] and d-xylonate dehydratase from Haloferax volcanii [69], are typically reported to belong to the enolase superfamily, but no crystal structure of any aldopentonate dehydratase belonging to the enolase family is currently known. This may in part be due to difficulties in expressing the archaeal aldopentonate dehydratases in heterologous hosts like E.coli [49].

IlvD/EDD superfamily

The IlvD/EDD protein family consists of various dehydratases that are either involved in short-chain amino acid biosynthesis (IlvD refers to isoleucine/leucine/valine dehydrates), or in carbohydrate metabolic pathways (EDD refers to Entner–Doudoroff dehydratases). All these enzymes are thought to have an iron–sulfur cluster at their active site. The [Fe–S] clusters are evolutionarily ancient prosthetic groups found in many metabolically important enzymes, and are suggested to participate in electron transfer and iron–sulphur storage, catalysis, regulation of gene expression, and in oligomer formation [70].

The first crystal structure of the IlvD/EDD family was an apo-form of 6-phosphogluconate dehydratase from S. oneidensis (PDB code 2GP4) solved by the Structural Genomics Consortium Project [62]. The enzyme 6-phosphogluconate dehydratase is involved in the classical ED pathway of glucose. The first aldopentonate dehydratase crystal structure became available in 2017 when the apo-, holo- and variant S480A structures of Rl ADHT (PDB codes 5J83, 5J84, 5J85, respectively) were solved [58]. A year later, the crystal structure of Cc XDHT (PDB code 5OYN) was also published [59]. Recently, holo-forms of dihydroxy acid dehydratase from A. thaliana (PDB codes 5YM0 and 5ZE4) [63], M. tuberculosis (PDB code 6OVT) [65] and Synechocystis sp. (PDB code 6NTE) [66] were determined (Table 3).

Table 3 Available crystal structures of IlvD/EDD enzymes
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The overall structure of Rl ADHT, as well as all other IlvD/EDD enzymes, consists of two domains, the N-terminal αβα-sandwich domain and the C-terminal β-barrel domain (Fig. 5a). The core of the N-terminal domain is formed by a four-stranded parallel β-sheet flanked by α-helices, which is then further surrounded by additional secondary structures. The C-terminal domain consists of eight mixed β-strands. IlvD/EDD proteins exist as either homodimers or homotetramers, but the tetrameric structure can be also described as a dimer of dimers (Fig. 5b). Dimerization has a significant role in formation of the active site, which lies in the cavity between the two domains and at the interface of the monomeric units of the dimer. Dimerization restricts free access to the active site and protects the iron–sulfur cluster from oxidative damage, but transient access is achieved by a conformational shift of the N-terminal helix − loop − helix region as shown in the structure of the Rl ADHT S480A mutant [58] (Fig. 5c).

Fig. 5
figure 5

Crystal structure of Rl ADHT (PDB: 5J84). a Overall fold is represented by a ribbon diagram. The N-terminal domain is shown in yellow, and the C-terminal domain is shown in green. The Mg2+ ion is represented by a green sphere and the [2Fe–2S] cluster by orange–yellow spheres. b Overall structure of the homo-tetramer. c Conformational shift of the N-terminal helix − loop − helix region. The closed form (PDB: 5J84) is shown in yellow and the open form (PDB: 5J85) in cyan. d Active site of Rl ADHT (PDB: 5J84) with a docked l-Arabinonate

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Each monomer of the aldopentonate dehydratases contains an iron–sulfur [2Fe–2S] cluster and an Mg2+ ion in the active site. In Rl ADHT, a planar [2Fe–2S] cluster is bound to the N-terminal domain by three cysteine residues (Cys59, Cys127, and Cys200). All the published structures of IlvD/EDD enzymes show that [2Fe–2S] cluster is coordinated by three cysteines, and the fourth site is thought to be filled either by water or substrate. Mg2+ is octahedrally coordinated by a conserved Asp, two Glu and ɛ-carboxy-Lys residues, along with two water molecules. A Mg2+ ion most likely coordinates the sugar acid at its C1 carboxylate group to the active site. Based on solved crystal structures and site-directed mutagenesis studies, the reaction mechanism of aldopentonate dehydratases is suggested to begin by the abstraction of a proton from the C2 atom of aldopentonate by the alkoxide form of the conserved serine side chain (Ser480 in Rl ADHT and Ser490 in Cc XyDHT), which acts as a Lewis base [58]. A three-coordinated Fe atom acts as a Lewis acid and accepts an electron pair from the leaving hydroxyl group on the C3 of the substrate.

Enolase superfamily

Although no crystal structure of aldopentonate dehydratase belonging to the enolase superfamily is currently described, several crystal structures of aldohexonate and deoxyhexonate dehydratases—such as d-glucarate dehydratases [71, 72], l-rhamnonate dehydratase [73], l-fuconate dehydratase [74], galactarate dehydratase [75] and d-mannonate dehydratase [76]—are known. Based on variations in which amino-acid residues participate in catalysis, the enolase superfamily can be divided into the following seven subgroups: enolase, muconate lactonizing enzyme, mandelate racemase, d-glucarate dehydratase, d-mannonate dehydratase, β-methylaspartate ammonia lyase, and galactarate dehydratase [77]. Archeal aldopentonate dehydratases, such as from H. volcanii, and 2 dehydratases from S. solfataricus, belong to the mandelate racemase/muconate lactonizing enzyme family of the enolase superfamily along with d-gluconate/d-galactonate dehydratase from S. solfataricus (Ss GADHT) of the modified promiscuous ED pathway. Ss GADHT can convert d-gluconate and d-galactonate, but is unable to convert aldopentonates. The crystal structure of Ss GADHT is also currently unknown. Dehydratase from H. volcanii is reported to catalyze d-xylonate and d-gluconate [69], while Sso3124 S. solfataricus dehydratase [30] participates in d-arabinose degradation, and Sso2665 S. solfataricus dehydratase participates in degradation of d-xylonate and l-arabinonate [37].

Based on sequence comparison, it can be predicted that archaeal aldopentonate dehydratases contain a typical enolase fold with two domains: an N-terminal α + β capping domain and a C-terminal modified TIM-barrel domain, known as a (β/α)7β-barrel. Both H. volcanii and Sso3124 S. solfataricus dehydratases are characterized as homo-octamers, whereas Sso2665 S. solfataricus dehydratase is active as a tetramer [30, 69]. The N-terminal domain is primarily responsible for determining substrate specificity and the C-terminal domain is responsible for acid/base chemistry. The active site is located at the interface between the domains and contains a Mg2+ ion. The Mg2+ ion is suggested to play an essential role in the reaction mechanism as it stabilizes an enolate anion intermediate generated by the abstraction of a carboxylate α-proton by an active site base as the active site acid usually directs intermediates toward the product [77].

2-Dehydro-3-deoxy-aldopentonate dehydratases

In the penultimate reaction step of the Weimberg pathway, 2-dehydro-3-deoxy-aldopentonate is converted to 2,5-dioxopentanoate (often called α-ketoglutaric semialdehyde) by 2-dehydro-3-deoxy-aldopentonate dehydratase. The l-form intermediate of the pathway, 2-dehydro-3-deoxy-l-aldopentonate, is dehydrated by 2-dehydro-3-deoxy-l-aldopentonate dehydratase (EC 4.2.1.43), a.k.a. 2-dehydro-3-deoxy-l-arabinonate dehydratase, or 2-keto-3-deoxy-l-arabinonate dehydratase. The d-form intermediate, 2-dehydro-3-deoxy-d-aldopentonate, is converted by 2-dehydro-3-deoxy-d-aldopentonate dehydratase (EC 4.2.1.141), a.k.a. 2-dehydro-3-deoxy-d-arabinonate dehydratase, or 2-keto-3-deoxy-d-arabinonate dehydratase. This enzyme is also sometimes called 2-dehydro-3-deoxy-d-xylonate dehydratase due to its role in d-xylose degradation. However, the penultimate intermediate of the Weimberg pathway is the same for all d-aldopentoses, because only one stereocenter is left from the pentose sugar.

Interestingly, enzymes catalyzing different stereoisomers have remarkably different three-dimensional structures. Table 4 shows the available crystal structures of 2-dehydro-3-deoxy-aldopentonate dehydratases. The crystal structure of S. solfataricus 2-dehydro-3-deoxy-d-aldopentonate dehydratase (hereafter referred to as Ss DPDHT) has been solved with bound magnesium (PDB code 3BQB) and with calcium ions (PDB code 2Q1C), and also complexed with a substrate analogue 2-oxobutyrate (PDB code 2Q1A) and a product 2,5-dioxopentanoate (PDB code 2QID) [78]. Ss DPDHT belongs to the metal-dependent fumarylacetoacetate hydrolase (FAH) protein family. By contrast, 2-dehydro-3-deoxy-L-aldopentonate dehydratase from A. brasilense (hereafter referred as Ab LPDHT) belongs to dihydrodipicolinate synthase/N-acetylneuraminate lyase (DHDPS/NAL) protein family. In 2008, the crystal structure of Ab LPDHT was determined, both in its uncomplexed form (PDB code 3FKK) [79] and complexed with pyruvate (PDB code 3FKR) [80]. Recently, crystal structures of Ab LPDHT have been determined without a ligand (PDB code 7C0C) and complexed with two different substrate analogues: β-hydroxypyruvate (PDB code 7C0D) and 2-oxobutyrate (PDB code 7C0E) [81].

Table 4 Available crystal structures of 2-dehydro-3-deoxy-aldopentonate dehydratases
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FAH superfamily

The FAH superfamily is named after fumarylacetoacetate hydrolase, which was the first solved protein structure in the family [82]. The family contains a diverse set of enzymes that are essential for degrading complex carbon sources in various metabolic pathways in both prokaryotes and eukaryotes. Despite the enzymatic diversity, all FAH superfamily members contain a catalytic FAH-domain. Ss DPDHT has two domains: an N-terminal domain with a four-stranded antiparallel β-sheet flanked by two α-helices on either side, and a catalytic C-terminal FAH-domain with a mixed β-sandwich roll fold (Fig. 6a). The quaternary structure of Ss DPDHT is an oval-ring shaped homotetramer, composed of a dimer of dimers (Fig. 6b).

Fig. 6
figure 6

Crystal structure of Ss DPDHT (PDB: 3BQB). a Overall fold is represented by a ribbon diagram. The N- and C-terminal domains are shown in green and yellow, respectively. Mg.2+ ion shown as green sphere. b Overall structure of the homo-tetramer. c Crystal structure of Ss DPDHT complexed with a 2,5-dioxopentanoate product (PDB: 2Q1D)

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The active site pocket contains a catalytic Mg2+ ion that is coordinated by three conserved acidic residues (Glu143, Glu145, and Glu164 in Ss DPDHT). The metal ion is hexacoordinated by these three acidic residues together with one water molecule and two oxygen atoms of the ligand (Fig. 6c). The enzymatic mechanism for water elimination from 2-dehydro-3-deoxy-d-aldopentonate involves the catalytic metal ion and a catalytic dyad, Glu/Lys. Two putative reaction mechanisms for dehydration of 2-dehydro-3-deoxy-d-arabinonate have been proposed [78], which are differentiated by initial proton abstraction from either C3 or C5. In both mechanisms, it is suggested that the Glu114 subtracts a proton and Lys182 adds a proton to the C4 hydroxyl group to cause water elimination. The Mg2+ ion holds the substrate in place and the bidentate binding of the substrate might increase the acidity of the C3 proton.

DHDPS/NAL family

The DHDPS/NAL protein family is named after the first two solved archetypal structures: dihydrodipicolinate synthase (DHDPS) [83] and N-acetylneuraminate lyase (NAL) [84]. This is a large family of enzymes, such as 5-keto-4-deoxy-glucarate dehydratases and d-2-keto-deoxy-gluconate aldolases. DHDPS/NAL proteins have a common (α/β)8-barrel fold composed of a (β)8-barrel surrounded by eight α-helices, where the active site is at the C-terminal end of a central β-barrel. In addition, the structure contains three C-terminal α-helices (Fig. 7a). DHDPS/NAL proteins are tetramers composed of dimers of dimers (Fig. 7b). The wide interface area between the monomers suggests that the tetrameric structure is highly stable in solution.

Fig. 7
figure 7

Crystal structure of the monomer of Ab LPDHT (PDB: 7C0C). a Overall fold is represented by a ribbon diagram. The (α/β)8-barrel fold is shown in yellow; C-terminal α-helices are shown in light orange. b Overall structure of the homo-tetramer. c Active site of Ab LPDHT complexed with the substrate analogue β-hydroxypyruvate (PDB: 7C0D)

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The active site of the DHDPS/NAL enzyme contains a conserved lysine residue that forms a Schiff-base intermediate with the C2 carbon of an α-keto acid substrate, usually performing cleavage of C–C or C=C bonds typical of type I aldolases. The 2-dehydro-3-deoxy-l-aldopentonate dehydratase lacks the aldol cleavage functionality and performs only the dehydration reaction. In Ab LPDHT, Lys171 is located at the center of the barrel on β6 and mutation of conserved lysine to alanine completely inactivates the enzyme, suggesting Schiff-base formation also occurs with 2-dehydro-3-deoxy-l-arabinonate dehydratases [81]. Interestingly, 2-dehydro-3-deoxy-l-aldopentonate dehydratase seems to have a generally poor phylogenetic relationship with other DHDPS/NAL enzymes. It is thought that 2-dehydro-3-deoxy-l-aldopentonate dehydratase evolved from a common aldolase ancestor with a tyrosine residue having been replaced by Gln143 in Ab LPDHT. In addition, the substrate-binding motif in Ab LPDHT, which is significantly divergent and involved in acid/base catalysis in type 1 aldolases, has been replaced.

Based on crystallographic and mutagenesis studies, a reaction mechanism for 2-dehydro-3-deoxy-l-aldopentonate dehydratases has been proposed [81], where Glu173 and Glu200 acts as catalytic Brønsted bases for the C3 and C5 protons, respectively, of the Schiff-base intermediate. The C3 atom of hydroxypyruvate (and also the C3 atom from 2-oxobuturate) is about 3 Å away from the carboxylate group of Glu173 (Fig. 7c). Glu173 also interacts with a side-chain of Gln143 and the main-chain of Glu199. Notably, Glu173 is not conserved among any other DHDPD/NAL enzymes. In aldolases, the tyrosine residue typically acts as a Brønsted base catalyst, resulting in cleavage of the substrate.

Ketoglutarate-semialdehyde dehydrogenase

The last step of the Weimberg pathway is the conversion of 2,5-dioxopentanoate to α-ketoglutarate via ketoglutarate-semialdehyde dehydrogenase (E.C. 1.2.1.26). Ketoglutarate-semialdehyde dehydrogenase belongs to the aldehyde dehydrogenase (ALDH) family, which contains a variety of NAD(P)-dependent enzymes that catalyse the oxidation of aliphatic and/or aromatic aldehydes to their corresponding acids. ALDH proteins are found in all kingdoms and have multiple functions in cellular metabolism and defense systems [85]. In 1997, Liu et al. [86]. were the first to solve the structure of aldehyde dehydrogenase from Rattus norvegicus.

At present, structural information for ketoglutarate-semialdehyde dehydrogenases is very limited (Table 5) and only the crystal structure of ketoglutarate-semialdehyde dehydrogenase from A. brasiliensis (hereafter referred to as Ab KGSADH) has been solved with and without a cofactor (PDB codes 5X5T and 5X5U) [87].

Table 5 Available crystal structures of ketoglutarate-semialdehyde dehydrogenases
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The Ab KGSADH has a three-domain structure typical of ALDH proteins: 1) an N-terminal domain (Met1–Arg123 and Val145–Leu254), 2) an oligomerization domain (Val124–Pro144 and Tyr470–Val481), and 3) a C-terminal domain (Gly255–Pro469) (Fig. 8a). The N-terminal domain is responsible for nucleotide binding and is composed of seven α-helices (α1–α7) and nine β-strands (β1–β4 and β7–β11), the core of which has a five-stranded Rossmann-like fold (β7–β11). The C-terminal domain has a large β-sheet containing seven β-strands (β12–β18) surrounded by six α-helices (α8–α13) and two 310-helices (G1, G2). In addition, two 310-helices (G3, G4) and one short β-strand (β19) exist at the C-terminus of this domain. The oligomerization domain is composed of a three-stranded antiparallel β-sheet (β5, β6, and β20) that protrudes from the N-terminal domain. In the dimer, the last β-strand of the oligomerization domain interacts with the last β-strand of the C-terminal domain of another monomeric subunit. The quaternary structure of Ab KGSADH is a homotetramer, a dimer of dimers (Fig. 8b), which is a highly conserved architecture among ALDH proteins.

Fig. 8
figure 8

Crystal structure of the monomer of the ketoglutarate-semialdehyde dehydrogenase Ab KGSADH from Azospirillum brasilense (PDB: 5X5U). a Overall fold is represented by a ribbon diagram. The N-terminal domain, C-terminal domain and C-terminal oligomerization domains are shown in yellow, green, and purple, respectively. b Overall quaternary structure is a homotetramer

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Ab KGSADH prefers to utilize NAD as a cofactor [53]. The NAD binding pocket lies in a space between the N- and C-terminal domains. In NAD-dependent ALDHs, the ribose-ring binding site of an adenosine nucleotide is not large enough to accommodate the phosphorylated form of the ring, which accounts for the preference of NAD over NADP. Many NADP-dependent ALDHs have a serine residue instead of the Glu181 residue observed in Ab KGSADH.

The active site tunnel exists in the inter-domain space, where two highly conserved catalytic Cys and Glu residues are found at the bottom of the pocket. In Ab KGSADH, the catalytic residues were concluded from site-directed mutagenesis experiments to be Cys287 and Glu253 [87]. During catalysis, the Cys residue first forms a tetrahedral intermediate with the carbonyl carbon of the aldehyde group, and the hydride ion is then transferred to NAD. The Glu residue acts as a general base and abstracts a proton from a water molecule, which then attacks the carbonyl carbon to again form a tetrahedral intermediate. The acidic product dissociates and the enzyme is then ready for a new cycle of catalysis [88].

2-Dehydro-3-deoxy-aldopentonate aldolase

In the Dahms pathway, the last step is catalyzed by 2-dehydro-3-deoxy-aldopentonate aldolase (a.k.a. 2-keto-3-deoxy-aldopentonate aldolase), which cleaves 2-dehydro-3-deoxy-aldopentonate into pyruvate and glycolaldehyde. Currently, no crystal structures of aldolases with strict specificity to 2-dehydro-3-deoxy-aldopentonate are described. However, several Entner–Doudoroff pathway-associated 2-dehydro-3-deoxy-gluconate aldolases, such as from S. solfataricus (hereafter referred to as Ss KDGA), S. acidocaldarius (hereafter referred to as Sa KDGA), S. tokodaii [37, 89], and E. coli (hereafter referred to as Ec KDGA) [17], have been reported to have some activity for aldopentonates (Table 6). Interestingly, these enzymes have been reported to have catalytic activity on both C4 epimers of hexose and pentose sugars as they cleave both 2-dehydro-3-deoxy-gluconate and 2-dehydro-3-deoxy-galactonate as well as both 2-dehydro-3-deoxy-l-aldopentonate and 2-dehydro-3-deoxy-d-aldopentonate [90]. Several crystal structures of Ss KDGA, Sa KDGA and Ec KDGA have been solved with and without ligands, such as pyruvate, 2-dehydro-3-deoxy-gluconate or 2-dehydro-3-deoxy-galactonate [89, 91,92,93].

Table 6 Available crystal structures of 2-dehydro-3-deoxy-d-gluconate aldolases with reported 2-dehydro-3-deoxy-aldopentonate activity
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The 2-dehydro-3-deoxy-aldopentonate aldolases belong to the DHDPS/NAL protein family along with the 2-dehydro-3-deoxy-l-aldopentonate dehydratases discussed earlier. However, compared with 2-dehydro-3-deoxy-l-aldopentonate dehydratase, 2-dehydro-3-deoxy-aldopentonate aldolase is expected to be a far more conventional C–C bond cleaving member of the DHDPS/NAL protein family as its active site contains the conserved catalytic residues lysine and tyrosine, and conserved substrate-recognizing residues (GXXG motif). Lysine is responsible for formation of a Schiff-base intermediate, and tyrosine is responsible for shuttling protons. Mutagenesis studies have shown that the catalytic triad of Tyr, Thr (or Ser) and Tyr residues act as a shuttle to transfer protons to and from active site that is needed for formation of a Schiff´s base and subsequent aldol condensation/cleavage [91]. In the 2-dehydro-3-deoxy-d-gluconate aldolase Ec KDGA, the triad is formed by Tyr145 above the Schiff´s base-forming Lys174, Ser56 from the GXXG motif, and Tyr119 from the adjacent monomer in the dimer [93].

2-Dehydro-3-deoxy-d-gluconate aldolases catalyse cleavage of 2-dehydro-3-deoxy-d-gluconate to pyruvate and glyceraldehyde, but they also possess a catalytic promiscuity that enables them to catalyse 2-dehydro-3-deoxy-d-galactose as well. The Schiff´s base complexes of Ss GDGA with C4 epimers of 2-dehydro-3-deoxy-d-glucose (D-KDG) and 2-dehydro-3-deoxy-d-galactose (D-KDGal) show that the active site of the enzyme is rigid, but the sugar substrates have conformational flexibility (Fig. 9a, b). The O4-hydroxyls of both substrates are in positions to interact with Tyr130, supporting the role of this tyrosine in proton abstraction. An alternative hydrogen-bonding network allows recognition of O5- and O6-hydroxyls even when they are in different orientations, which explains the substrate promiscuity of Ss GDGA [91]. The Ec KDGA–D-KDGal complex shows that its interactions with substrates are similar to those observed in the Ss KDGA–D-KDGal complex, but the O6–hydroxyl interacts directly with the backbone of Gly210 and Ala221 (Fig. 9c).

Fig. 9
figure 9

Active site of KDGA enzymes: a Ss KDGA complex with D-KDG (PDB: 1W3N); b Ss KDGA complex with D-KDGal (PDB: 1W3T); and c Ec KDGA complex with D-KDGal (PDB: 3NEV)

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Future prospects

The oxidative Weimberg and Dahms pathways provide five- and four-step enzymatic reactions, respectively, for converting pentose sugars to key glycolytic intermediates, such as α-ketoglutarate in the Weimberg pathway, and pyruvate and glycolaldehyde in the Dahms pathway, thus offering sustainable bioconversion of lignocellulose-derived aldopentoses to added-value products (Fig. 10). Weimberg/Dahms pathways are also independent of other carbohydrate assimilation pathways, which simplifies metabolic engineering efforts [99]. Oxidative aldopentose pathway enzymes have been utilized for production of d-xylonic acid or l-arabinoic acid from biomass-derived aldopentoses, either in E.coli or various yeasts [31, 44, 100]. Besides these pentonic acids, microbial synthesis of 1,2,4-butanetriol by the oxidative aldopentose pathway has been demonstrated [101]. A similar synthetic route, with a different last step reaction, of 3,4-dihydroxybutyric acid has also been established in E. coli [102]. At acidic condition, 3,4-dihydroxybutyric acid can be cyclized to 3-hydroxybutyrolactone (γHBL). Liu et al. [17, 103] have further reported biosynthesis of ethylene glycol from d-xylose by the Dahms pathway in E.coli, while Salusjärvi et al. [61] produced ethylene glycol and glycolic acid in S. cerevisiae yeast. This pathway of glycolic acid synthesis has also been used for PLGA copolymer (of glycolic acid and lactic acid) production in E. coli [104, 105]. In addition, Tai et al. [3] have applied partial Weimberg pathway for producing 1,4-butanediol in E. coli. The Weimberg pathway has been also utilized to yield mesaconic acid [106] as well as the glutaric acid via the α-ketoglutarate [107] in E. coli.

Fig. 10
figure 10

Added-value chemicals that can be produced from the intermediates or products of Weimberg/Dahms pathway. In the center of the scheme, the gray circle contains the common steps for Weimberg and Dahms pathways, the blue circle have the subsequent two steps in Weimberg pathway, and the green circle shows the last step of Dahms pathway. The added-value chemicals that can be derived from the pathways are shown. The yellow arrows correspond the enzymatic reaction and the purple arrow is the non-enzymatic reaction at acidic condition

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Metabolic engineering efforts have shown their potential in the production of valuable bioproducts from aldopentoses. However, incomplete biochemical and structural knowledge of the enzymes in these metabolic systems is still hampering these efforts. For example, it is evident that one of the bottlenecks in these oxidative pathways is the Fe–S cluster-containing aldopentonate dehydratase enzyme, which should be optimized in terms of expression and catalytic efficiency [49, 61, 99]. Three-dimensional enzyme structures and biochemical data are needed to provide a better understanding of structure–function relationships of the enzymes, including catalytic mechanisms, as well as to help protein and metabolic engineering efforts.

Protein engineering, including future novel artificial enzymes, provides a tremendous toolbox to broaden the potential applications of synthetic biochemical routes. The oxidative non-phosphorylative pathways of pentose catabolism have not yet been fully exploited, and the involving enzymes have not yet been subjected to protein engineering efforts. Particularly, the improvement of bottleneck enzyme, the aldopentonate dehydratase, will be beneficial to solve the drawbacks of utilizing these pathways in biosynthesis of chemicals from pentose sugars [99]. On the other hand, the optimization of some other related enzymes has been reported. Naturally existing diol dehydratase has been designed by rational protein engineering to have catalytic activity toward the artificial substrate 1,2,4-butanetriol, which led to enhanced production of 1,4-butanediol from xylose [108]. In addition to rational protein design, more random approaches have been used to modify the enzymes. Recently, dihydroxyacid dehydratase from S. solfataricus was successfully engineered to have tenfold higher activity for conversion of glycerate dehydration to pyruvate by iterative saturation mutagenesis [109].

The main aim of this review has been to provide a summary of the existing three-dimensional structures of enzymes involved in oxidative non-phosphorylative catabolic pathways for aldopentoses, as well as to highlight some missing links, including complex structures of enzymes, that may restrict understanding of enzymatic reactions. We believe that our structural review of Weimberg/Dahms pathway enzymes will be of value for researchers attempting to engineer more efficient pathways for producing biobased compounds from xylose and arabinose. Engineered enzyme pathways could be used in vivo or in vitro, or in hybrid systems. Ultimately, we hope that an increased range of biobased products can be developed to help reduce our dependence on oil by replacement with renewable energy sources.

Availability of data and materials

Not applicable.

Abbreviations

AAOR:

Aldose-aldose oxidoreductase

ADH:

Arabinose dehydrogenase

ADHT:

Arabinonate dehydratase

ALDH:

Aldehyde dehydrogenase

DHDPS:

Dihydrodipicolinate synthase

DPDHT:

2-Dehydro-3-deoxy-d-aldopentonate dehydratase

ED:

Entner–Doudoroff

EDD:

Entner–Doudoroff dehydratase

FAH:

Fumarylacetoacetate hydrolase

GADHT:

Gluconate/galactonate dehydratase

GDH:

Glucose dehydrogenase

IlvD:

Isoleucine/leucine/valine dehydrate

KDG:

2-Keto-3-deoxy-glucose

KDGA:

2-Keto-3-deoxy-gluconate aldolases

KDGal:

2-Keto-3-deoxy-galactose

KGSADH:

Ketoglutarate-semialdehyde dehydrogenase

LDR:

Long-chain dehydrogenase/reductase

LPDHT:

2-Dehydro-3-deoxy-l-aldopentonate dehydratase

MDR:

Medium-chain dehydrogenase/reductase

NAD:

Nicotinamide adenine dinucleotide

NADP:

Nicotinamide adenine dinucleotide phosphate

NAL:

N-acetylneuraminate lyase

PDB:

Protein data bank

PLGA:

Poly lactic-co-glycolic acid

SDR:

Short-chain dehydrogenase/reductase

SMP30:

Senescence marker protein 30

XDHT:

Xylonate dehydratase

References

  1. Sun X, Shen X, Jain R, Lin Y, Wang J, Sun J, et al. Synthesis of chemicals by metabolic engineering of microbes. Chem Soc Rev. 2015;44:3760–85.

    Article  CAS  Google Scholar 

  2. Kawaguchi H, Hasunuma T, Ogino C, Kondo A. Bioprocessing of bio-based chemicals produced from lignocellulosic feedstocks. Curr Opin Biotechnol. 2016;42:30–9.

    Article  CAS  Google Scholar 

  3. Tai Y-S, Xiong M, Jambunathan P, Wang J, Wang J, Stapleton C, et al. Engineering nonphosphorylative metabolism to generate lignocellulose-derived products. Nat Chem Biol. 2016;12:247–53.

    Article  CAS  Google Scholar 

  4. Klemm D, Heublein B, Fink H-P, Bohn A. Cellulose: fascinating biopolymer and sustainable raw material. Angew Chem Int Ed. 2005;44:3358–93.

    Article  CAS  Google Scholar 

  5. Biely P. Microbial xylanolytic systems. Trends Biotechnol. 1985;3:286–90.

    Article  CAS  Google Scholar 

  6. Webb SR, Lee H. Regulation of d-xylose utilization by hexoses in pentose-fermenting yeasts. Biotechnol Adv. 1990;8:685–97.

    Article  CAS  Google Scholar 

  7. Jeffries TW. Utilization of xylose by bacteria, yeasts, and fungi. Adv Biochem Eng Biotechnol. 1983;27:1–32.

    CAS  Google Scholar 

  8. Salusjärvi L, Havukainen S, Koivistoinen O, Toivari M. Biotechnological production of glycolic acid and ethylene glycol: current state and perspectives. Appl Microbiol Biotechnol. 2019;103:2525–35.

    Article  Google Scholar 

  9. Bräsen C, Esser D, Rauch B, Siebers B. Carbohydrate metabolism in archaea: current insights into unusual enzymes and pathways and their regulation. Microbiol Mol Biol Rev. 2014;78:89–175.

    Article  Google Scholar 

  10. Zhang M, Eddy C, Deanda K, Finkelstein M, Picataggio S. Metabolic engineering of a pentose metabolism pathway in ethanologenic Zymomonas mobilis. Science. 1995;267:240–3.

    Article  CAS  Google Scholar 

  11. Hahn-Hägerdal B, Jeppsson H, Skoog K, Prior BA. Biochemistry and physiology of xylose fermentation by yeasts. Enzyme Microb Technol. 1994;16:933–43.

    Article  Google Scholar 

  12. Weimberg R. Pentose oxidation by Pseudomonas fragi. J Biol Chem. 1961;236:629–35.

    Article  CAS  Google Scholar 

  13. Dahms AS. 3-Deoxy-d-pentulosonic acid aldolase and its role in a new pathway of d-xylose degradation. Biochem Biophys Res Commun. 1974;60:1433–9.

    Article  CAS  Google Scholar 

  14. Watanabe S, Saimura M, Makino K. Eukaryotic and bacterial gene clusters related to an alternative pathway of nonphosphorylated l-rhamnose metabolism. J Biol Chem. 2008;283:20372–82.

    Article  CAS  Google Scholar 

  15. Wolf J, Stark H, Fafenrot K, Albersmeier A, Pham TK, Müller KB, et al. A systems biology approach reveals major metabolic changes in the thermoacidophilic archaeon Sulfolobus solfataricus in response to the carbon source l-fucose versus d-glucose. Mol Microbiol. 2016;102:882–908.

    Article  CAS  Google Scholar 

  16. Watanabe S, Fukumori F, Nishiwaki H, Sakurai Y, Tajima K, Watanabe Y. Novel non-phosphorylative pathway of pentose metabolism from bacteria. Sci Rep. 2019;9:155.

    Article  Google Scholar 

  17. Liu H, Ramos KRM, Valdehuesa KNG, Nisola GM, Lee W-K, Chung W-J. Biosynthesis of ethylene glycol in Escherichia coli. Appl Microbiol Biotechnol. 2013;97:3409–17.

    Article  CAS  Google Scholar 

  18. Cao Y, Niu W, Guo J, Xian M, Liu H. Biotechnological production of 1,2,4-butanetriol: An efficient process to synthesize energetic material precursor from renewable biomass. Sci Rep. 2016;5:18149.

    Article  Google Scholar 

  19. Francois JM, Alkim C, Morin N. Engineering microbial pathways for production of bio-based chemicals from lignocellulosic sugars: current status and perspectives. Biotechnol Biofuels. 2020;13:118.

    Article  CAS  Google Scholar 

  20. McClintock MK, Wang J, Zhang K. Application of nonphosphorylative metabolism as an alternative for utilization of lignocellulosic biomass. Front Microbiol. 2017;8:2310.

    Article  Google Scholar 

  21. Brouns SJJ, Turnbull AP, Willemen HLDM, Akerboom J, van der Oost J. Crystal structure and biochemical properties of the d-arabinose dehydrogenase from Sulfolobus solfataricus. J Mol Biol. 2007;371:1249–60.

    Article  CAS  Google Scholar 

  22. Milburn CC, Lamble HJ, Theodossis A, Bull SD, Hough DW, Danson MJ, et al. The structural basis of substrate promiscuity in glucose dehydrogenase from the hyperthermophilic archaeon Sulfolobus solfataricus. J Biol Chem. 2006;281:14796–804.

    Article  CAS  Google Scholar 

  23. Watanabe Y, Iga C, Watanabe Y, Watanabe S. Structural insights into the catalytic and substrate recognition mechanisms of bacterial l -arabinose 1-dehydrogenase. FEBS Lett. 2019;593:1873-3468.13424.

    Google Scholar 

  24. Yoshiwara K, Watanabe S, Watanabe Y. Crystal structure of bacterial l-arabinose 1-dehydrogenase in complex with l-arabinose and NADP+. Biochem Biophys Res Commun. 2020;530:203–8.

    Article  CAS  Google Scholar 

  25. Eswaramoorthy S, Almo SC, Swaminathan S, New York Structure Genomics Research Consortium (NYSGRC). Crystal structure of D-galactose-1-dehydrogenase protein from Rhizobium etli. Protein Data Bank. 2012. https://www.rcsb.org/structure/4EW6

  26. Taberman H, Andberg M, Koivula A, Hakulinen N, Penttilä M, Rouvinen J, et al. Structure and function of Caulobacter crescentus aldose–aldose oxidoreductase. Biochemical Journal. 2015;472:297–307.

    Article  CAS  Google Scholar 

  27. Persson B, Hedlund J, Jörnvall H. Medium- and short-chain dehydrogenase/reductase gene and protein families: the MDR superfamily. Cell Mol Life Sci. 2008;65:3879–94.

    Article  CAS  Google Scholar 

  28. Eklund H, Nordström B, Zeppezauer E, Söderlund G, Ohlsson I, Boiwe T, et al. Three-dimensional structure of horse liver alcohol dehydrogenase at 2.4 Å resolution. J Mol Biol. 1976;102:27–59.

    Article  CAS  Google Scholar 

  29. Kavanagh KL, Jörnvall H, Persson B, Oppermann U. Medium- and short-chain dehydrogenase/reductase gene and protein families: the SDR superfamily: functional and structural diversity within a family of metabolic and regulatory enzymes. Cell Mol Life Sci. 2008;65:3895–906.

    Article  CAS  Google Scholar 

  30. Brouns SJJ, Walther J, Snijders APL, van de Werken HJG, Willemen HLDM, Worm P, et al. Identification of the missing links in prokaryotic pentose oxidation pathways. J Biol Chem. 2006;281:27378–88.

    Article  CAS  Google Scholar 

  31. Aro-Kärkkäinen N, Toivari M, Maaheimo H, Ylilauri M, Pentikäinen OT, Andberg M, et al. L-arabinose/d-galactose 1-dehydrogenase of Rhizobium leguminosarum bv. trifolii characterised and applied for bioconversion of l-arabinose to l-arabonate with Saccharomyces cerevisiae. Appl Microbiol Biotechnol. 2014;98:9653–65.

    Article  Google Scholar 

  32. Johnsen U, Schönheit P. Novel xylose dehydrogenase in the halophilic archaeon Haloarcula marismortui. J Bacteriol. 2004;186:6198–207.

    Article  CAS  Google Scholar 

  33. Stephens C, Christen B, Fuchs T, Sundaram V, Watanabe K, Jenal U. Genetic analysis of a novel pathway for d-Xylose metabolism in Caulobacter crescentus. J Bacteriol. 2007;189:2181–5.

    Article  CAS  Google Scholar 

  34. Johnsen U, Sutter J-M, Zaiß H, Schönheit P. l-Arabinose degradation pathway in the haloarchaeon Haloferax volcanii involves a novel type of L-arabinose dehydrogenase. Extremophiles. 2013;17:897–909.

    Article  CAS  Google Scholar 

  35. Yoshiwara K, Watanabe S, Watanabe Y. Crystal structure of l-rhamnose 1-dehydrogenase involved in the nonphosphorylative pathway of l-rhamnose metabolism in bacteria. FEBS Lett. 2021;595:637–46.

    Article  CAS  Google Scholar 

  36. Jörnvall H, von Bahr-Lindström H, Jany K-D, Ulmer W, Fröschle M. Extended superfamily of short alcohol-polyol-sugar dehydrogenases: structural similarities between glucose and ribitol dehydrogenases. FEBS Lett. 1984;165:190–6.

    Article  Google Scholar 

  37. Nunn CEM, Johnsen U, Schönheit P, Fuhrer T, Sauer U, Hough DW, et al. Metabolism of pentose sugars in the hyperthermophilic Archaea Sulfolobus solfataricus and Sulfolobus acidocaldarius. J Biol Chem. 2010;285:33701–9.

    Article  CAS  Google Scholar 

  38. Eklund H, Ramaswamy S. Medium- and short-chain dehydrogenase/reductase gene and protein families: three-dimensional structures of MDR alcohol dehydrogenases. Cell Mol Life Sci. 2008;65:3907–17.

    Article  CAS  Google Scholar 

  39. Magonet E, Hayen P, Delforge D, Delaive E, Remacle J. Importance of the structural zinc atom for the stability of yeast alcohol dehydrogenase. Biochem J. 1992;287:361–5.

    Article  CAS  Google Scholar 

  40. Schrödinger L, DeLano W. PyMOL. 2020. http://www.pymol.org/pymol.

  41. Taberman H, Parkkinen T, Rouvinen J. Structural and functional features of the NAD(P) dependent Gfo/Idh/MocA protein family oxidoreductases: Gfo/Idh/MocA protein family. Protein Sci. 2016;25:778–86.

    Article  CAS  Google Scholar 

  42. Rowland P, Basak AK, Gover S, Levy HR, Adams MJ. The three–dimensional structure of glucose 6–phosphate dehydrogenase from Leuconostoc mesenteroides refined at 2.0 Å resolution. Structure. 1994;2:1073–87.

    Article  CAS  Google Scholar 

  43. Andberg M, Maaheimo H, Kumpula E-P, Boer H, Toivari M, Penttilä M, et al. Characterization of a unique Caulobacter crescentus aldose-aldose oxidoreductase having dual activities. Appl Microbiol Biotechnol. 2015. https://doi.org/10.1007/s00253-015-7011-5.

    Article  Google Scholar 

  44. Wiebe MG, Nygård Y, Oja M, Andberg M, Ruohonen L, Koivula A, et al. A novel aldose-aldose oxidoreductase for co-production of d-xylonate and xylitol from d-xylose with Saccharomyces cerevisiae. Appl Microbiol Biotechnol. 2015;99:9439–47.

    Article  CAS  Google Scholar 

  45. Thoden JB, Holden HM. Structural and functional studies of WlbA: A dehydrogenase involved in the biosynthesis of 2,3-diacetamido-2,3-dideoxy-d-mannuronic acid. Biochemistry. 2010;49:7939–48.

    Article  CAS  Google Scholar 

  46. van Straaten KE, Zheng H, Palmer DRJ, Sanders DAR. Structural investigation of myo-inositol dehydrogenase from Bacillus subtilis : implications for catalytic mechanism and inositol dehydrogenase subfamily classification. Biochem J. 2010;432:237–47.

    Article  Google Scholar 

  47. Liu QP, Sulzenbacher G, Yuan H, Bennett EP, Pietz G, Saunders K, et al. Bacterial glycosidases for the production of universal red blood cells. Nat Biotechnol. 2007;25:454–64.

    Article  CAS  Google Scholar 

  48. Pääkkönen J, Hakulinen N, Andberg M, Koivula A, Rouvinen J. Three-dimensional structure of xylonolactonase from Caulobacter crescentus : a mononuclear iron enzyme of the 6-bladed β-propeller hydrolase family. Protein Sci. 2021;31:371–83.

    Article  Google Scholar 

  49. Boer H, Andberg M, Pylkkänen R, Maaheimo H, Koivula A. In vitro reconstitution and characterisation of the oxidative d-xylose pathway for production of organic acids and alcohols. AMB Expr. 2019;9:48.

    Article  Google Scholar 

  50. Chen C-N, Chin K-H, Wang AH-J, Chou S-H. The first crystal structure of gluconolactonase important in the glucose secondary metabolic pathways. J Mol Biol. 2008;384:604–14.

    Article  CAS  Google Scholar 

  51. Chakraborti S, Bahnson BJ. Crystal structure of human senescence marker protein 30: insights linking structural enzymatic, and physiological functions. Biochemistry. 2010;49:3436–44.

    Article  CAS  Google Scholar 

  52. Aizawa S, Senda M, Harada A, Maruyama N, Ishida T, Aigaki T, et al. Structural basis of the γ-lactone-ring formation in ascorbic acid biosynthesis by the senescence marker protein-30/gluconolactonase. PLoS ONE. 2013;8:e53706.

    Article  CAS  Google Scholar 

  53. Watanabe S, Kodaki T, Makino K. A novel α-ketoglutaric semialdehyde dehydrogenase. J Biol Chem. 2006;281:28876–88.

    Article  CAS  Google Scholar 

  54. Sutter J-M, Johnsen U, Schönheit P. Characterization of a pentonolactonase involved in d-xylose and l-arabinose catabolism in the haloarchaeon Haloferax volcanii. FEMS Microbiol Lett. 2017;364:fnx140.

    Article  Google Scholar 

  55. Kondo Y, Inai Y, Sato Y, Handa S, Kubo S, Shimokado K, et al. Senescence marker protein 30 functions as gluconolactonase in l-ascorbic acid biosynthesis, and its knockout mice are prone to scurvy. Proc Natl Acad Sci. 2006;103:5723–8.

    Article  CAS  Google Scholar 

  56. Shimokawa N, Yamaguchi M. Molecular cloning and sequencing of the cDNA coding for a calcium-binding protein regucalcin from rat liver. FEBS Lett. 1993;327:251–5.

    Article  CAS  Google Scholar 

  57. Pääkkönen J, Penttinen L, Andberg M, Koivula A, Hakulinen N, Rouvinen J, et al. Xylonolactonase from Caulobacter crescentus is a mononuclear nonheme iron hydrolase. Biochemistry. 2021;60:3046–9.

    Article  Google Scholar 

  58. Rahman MM, Andberg M, Thangaraj SK, Parkkinen T, Penttilä M, Jänis J, et al. The crystal structure of a bacterial l-arabinonate dehydratase contains a [2Fe-2S] cluster. ACS Chem Biol. 2017;12:1919–27.

    Article  CAS  Google Scholar 

  59. Rahman MM, Andberg M, Koivula A, Rouvinen J, Hakulinen N. The crystal structure of D-xylonate dehydratase reveals functional features of enzymes from the Ilv/ED dehydratase family. Sci Rep. 2018;8:865.

    Article  Google Scholar 

  60. Andberg M, Aro-Kärkkäinen N, Carlson P, Oja M, Bozonnet S, Toivari M, et al. Characterization and mutagenesis of two novel iron–sulphur cluster pentonate dehydratases. Appl Microbiol Biotechnol. 2016;100:7549–63.

    Article  CAS  Google Scholar 

  61. Salusjärvi L, Toivari M, Vehkomäki M-L, Koivistoinen O, Mojzita D, Niemelä K, et al. Production of ethylene glycol or glycolic acid from d-xylose in Saccharomyces cerevisiae. Appl Microbiol Biotechnol. 2017;101:8151–63.

    Article  Google Scholar 

  62. Schormann N, Symersky J, Southeast Collaboratory for Structural Genomics (SECSG). Structure of [FeS]cluster-free Apo Form of 6-Phosphogluconate Dehydratase from Shewanella oneidensis. Protein Data Bank. 2006. https://www.rcsb.org/structure/2GP4

  63. Yan Y, Liu Q, Zang X, Yuan S, Bat-Erdene U, Nguyen C, et al. Resistance-gene-directed discovery of a natural-product herbicide with a new mode of action. Nature. 2018;559:415–8.

    Article  CAS  Google Scholar 

  64. Zang X, Huang WX, Cheng R, Wu L, Zhou JH, Tang Y, et al. The crystal structure of DHAD. Protein Data Bank. 2018. https://www.rcsb.org/structure/5YM0

  65. Bashiri G, Grove TL, Hegde SS, Lagautriere T, Gerfen GJ, Almo SC, et al. The active site of the mycobacterium tuberculosis branched-chain amino acid biosynthesis enzyme dihydroxyacid dehydratase contains a 2Fe–2S cluster. J Biol Chem. 2019;294:13158–70.

    Article  CAS  Google Scholar 

  66. Zhang P, MacTavish BS, Yang G, Chen M, Roh J, Newsome KR, et al. Cyanobacterial dihydroxyacid dehydratases are a promising growth inhibition target. ACS Chem Biol. 2020;15:2281–8.

    Article  CAS  Google Scholar 

  67. Jiang Y, Liu W, Cheng T, Cao Y, Zhang R, Xian M. Characterization of d-xylonate dehydratase YjhG from Escherichia coli. Bioengineered. 2015;6:227–32.

    Article  CAS  Google Scholar 

  68. Bator I, Wittgens A, Rosenau F, Tiso T, Blank LM. Comparison of three xylose pathways in Pseudomonas putida KT2440 for the synthesis of valuable products. Front Bioeng Biotechnol. 2020;7:480.

    Article  Google Scholar 

  69. Johnsen U, Dambeck M, Zaiss H, Fuhrer T, Soppa J, Sauer U, et al. d-Xylose degradation pathway in the halophilic archaeon haloferax volcanii. J Biol Chem. 2009;284:27290–303.

    Article  CAS  Google Scholar 

  70. Johnson DC, Dean DR, Smith AD, Johnson MK. Structure, function, and formation of biological iron-sulfur clusters. Annu Rev Biochem. 2005;74:247–81.

    Article  CAS  Google Scholar 

  71. Gulick AM, Palmer DRJ, Babbitt PC, Gerlt JA, Rayment I. Evolution of enzymatic activities in the enolase superfamily: crystal structure of d-glucarate dehydratase from Pseudomonas putida. Biochemistry. 1998;37:14358–68.

    Article  CAS  Google Scholar 

  72. Gulick AM, Hubbard BK, Gerlt JA, Rayment I. Evolution of enzymatic activities in the enolase superfamily: crystallographic and mutagenesis studies of the reaction catalyzed by d-glucarate dehydratase from Escherichia coli. Biochemistry. 2000;39:4590–602.

    Article  CAS  Google Scholar 

  73. Rakus JF, Fedorov AA, Fedorov EV, Glasner ME, Hubbard BK, Delli JD, et al. Evolution of enzymatic activities in the enolase superfamily: l-rhamnonate dehydratase. Biochemistry. 2008;47:9944–54.

    Article  CAS  Google Scholar 

  74. Yew WS, Fedorov AA, Fedorov EV, Rakus JF, Pierce RW, Almo SC, et al. Evolution of enzymatic activities in the enolase superfamily: l-fuconate dehydratase from Xanthomonas campestris. Biochemistry. 2006;45:14582–97.

    Article  CAS  Google Scholar 

  75. Yew WS, Fedorov AA, Fedorov EV, Almo SC, Gerlt JA. Evolution of enzymatic activities in the enolase superfamily: l-talarate/galactarate dehydratase from Salmonella typhimurium LT2. Biochemistry. 2007;46:9564–77.

    Article  CAS  Google Scholar 

  76. Qiu X, Tao Y, Zhu Y, Yuan Y, Zhang Y, Liu H, et al. Structural insights into decreased enzymatic activity induced by an insert sequence in mannonate dehydratase from gram negative bacterium. J Struct Biol. 2012;180:327–34.

    Article  CAS  Google Scholar 

  77. Gerlt JA, Babbitt PC, Jacobson MP, Almo SC. Divergent evolution in enolase superfamily: strategies for assigning functions. J Biol Chem. 2012;287:29–34.

    Article  CAS  Google Scholar 

  78. Brouns SJJ, Barends TRM, Worm P, Akerboom J, Turnbull AP, Salmon L, et al. Structural insight into substrate binding and catalysis of a novel 2-Keto-3-deoxy-d-arabinonate dehydratase illustrates common mechanistic features of the FAH superfamily. J Mol Biol. 2008;379:357–71.

    Article  CAS  Google Scholar 

  79. Shimada N, Mikami B. Structure of L-2-keto-3-deoxyarabonate dehydratase. Protein Data Bank. 2010. https://www.rcsb.org/structure/3FKK

  80. Shimada N, Mikami B. Structure of L-2-keto-3-deoxyarabonate dehydratase complex with pyruvate. Protein Data Bank. 2010. https://www.rcsb.org/structure/3FKR

  81. Watanabe S, Watanabe Y, Nobuchi R, Ono A. Biochemical and structural characterization of L-2-Keto-3-deoxyarabinonate dehydratase: a unique catalytic mechanism in the class i aldolase protein superfamily. Biochemistry. 2020;59:2962–73.

    Article  CAS  Google Scholar 

  82. Timm DE, Mueller HA, Bhanumoorthy P, Harp JM, Bunick GJ. Crystal structure and mechanism of a carbon–carbon bond hydrolase. Structure. 1999;7:1023–33.

    Article  CAS  Google Scholar 

  83. Mirwaldt C, Korndorfer I, Huber R. The crystal structure of dihydrodipicolinate synthase from Escherichia coliat 2.5 Å resolution. J Mol Biol. 1995;246:227–39.

    Article  CAS  Google Scholar 

  84. Izard T, Lawrence MC, Malby RL, Lilley GG, Colman PM. The three-dimensional structure of N -acetylneuraminate lyase from Escherichia coli. Structure. 1994;2:361–9.

    Article  CAS  Google Scholar 

  85. Hempel J, Nicholas H, Lindahl R. Aldehyde dehydrogenases: widespread structural and functional diversity within a shared framework. Protein Sci. 1993;2:1890–900.

    Article  CAS  Google Scholar 

  86. Liu Z-J, Sun Y-J, Rose J, Chung Y-J, Hsiao C-D, Chang W-R, et al. The first structure of an aldehyde dehydrogenase reveals novel interactions between NAD and the Rossmann fold. Nat Struct Biol. 1997;4:317–26.

    Article  CAS  Google Scholar 

  87. Son HF, Park S, Yoo TH, Jung GY, Kim K-J. Structural insights into the production of 3-hydroxypropionic acid by aldehyde dehydrogenase from Azospirillum brasilense. Sci Rep. 2017;7:46005.

    Article  CAS  Google Scholar 

  88. Hempel J, Perozich J, Chapman T, Rose J, Boesch JS, Liu Z-J, et al. Aldehyde dehydrogenase catalytic mechanism. In: Weiner H, Maser E, Crabb DW, Lindahl R, editors., et al., Enzymology and molecular biology of carbonyl metabolism 7. Houston: Gulf Professional Publishing; 2021. p. 53–9.

    Google Scholar 

  89. Wolterink-van Loo S, van Eerde A, Siemerink MAJ, Akerboom J, Dijkstra BW, van der Oost J. Biochemical and structural exploration of the catalytic capacity of Sulfolobus KDG aldolases. Biochem J. 2007;403:421–30.

    Article  CAS  Google Scholar 

  90. Archer RM, Royer SF, Mahy W, Winn CL, Danson MJ, Bull SD. Syntheses of 2-Keto-3-deoxy-d-xylonate and 2-Keto-3-deoxy-L-arabinonate as stereochemical probes for demonstrating the metabolic promiscuity of Sulfolobus solfataricus towards d-Xylose and L-arabinose. Chem Eur J. 2013;19:2895–902.

    Article  CAS  Google Scholar 

  91. Theodossis A, Walden H, Westwick EJ, Connaris H, Lamble HJ, Hough DW, et al. The structural basis for substrate promiscuity in 2-Keto-3-deoxygluconate aldolase from the entner-doudoroff pathway in Sulfolobus solfataricus. J Biol Chem. 2004;279:43886–92.

    Article  CAS  Google Scholar 

  92. Manicka S, Peleg Y, Unger T, Albeck S, Dym O, Greenblatt HM, et al. Crystal structure of YagE, a putative DHDPS-like protein from Escherichia coli K12. Proteins. 2008;71:2102–8.

    Article  CAS  Google Scholar 

  93. Bhaskar V, Kumar M, Manicka S, Tripathi S, Venkatraman A, Krishnaswamy S. Identification of biochemical and putative biological role of a xenolog from Escherichia coli using structural analysis. Proteins. 2011;79:1132–42.

    Article  CAS  Google Scholar 

  94. Zaitsev V, Johnsen U, Reher M, Ortjohann M, Taylor GL, Danson MJ, et al. Insights into the substrate specificity of archaeal entner-doudoroff aldolases: the structures of Picrophilus torridus 2-Keto-3-deoxygluconate aldolase and Sulfolobus solfataricus 2-Keto-3-deoxy-6-phosphogluconate aldolase in complex with 2-Keto-3-deoxy-6-phosphogluconate. Biochemistry. 2018;57:3797–806.

    Article  CAS  Google Scholar 

  95. Manoj Kumar P, Baskar V, Manicka S, Krishnaswamy S. Crystal structure of YagE, a KDG aldolase protein, in complex with aldol condensed product of pyruvate and glyoxal. Protein Data Bank. 2014. https://www.rcsb.org/structure/4OE7

  96. Manoj Kumar P, Bhaskar V, Manicka S, Krishnaswamy, S. Crystal structure of YagE, a KDG aldolase protein in complex with 2-Keto-3-deoxy gluconate. Protein Data Bank. 2014. https://www.rcsb.org/structure/4ONV

  97. Manoj Kumar P, Bhaskar V, Manicka S, Krishnaswamy S. Crystal structure of 0.5M urea unfolded YagE, a KDG aldolase protein in complex with Pyruvate. Protein Data Bank. 2014. https://www.rcsb.org/structure/4U4M

  98. Manoj Kumar P, Baskar V, Manicka S, Krishnaswamy S. Crystal Structure of YagE, a KDG aldolase protein in complex with Magnesium cation coordinated L-glyceraldehyde. Protein Data Bank. 2014. https://www.rcsb.org/structure/4PTN

  99. Valdehuesa KNG, Ramos KRM, Nisola GM, Bañares AB, Cabulong RB, Lee W-K, et al. Everyone loves an underdog: metabolic engineering of the xylose oxidative pathway in recombinant microorganisms. Appl Microbiol Biotechnol. 2018;102:7703–16.

    Article  CAS  Google Scholar 

  100. Mehtiö T, Toivari M, Wiebe MG, Harlin A, Penttilä M, Koivula A. Production and applications of carbohydrate-derived sugar acids as generic biobased chemicals. Crit Rev Biotechnol. 2016;36:904–16.

    Article  Google Scholar 

  101. Niu W, Molefe MN, Frost JW. Microbial synthesis of the energetic material precursor 1,2,4-butanetriol. J Am Chem Soc. 2003;125:12998–9.

    Article  CAS  Google Scholar 

  102. Wang J, Shen X, Jain R, Wang J, Yuan Q, Yan Y. Establishing a novel biosynthetic pathway for the production of 3,4-dihydroxybutyric acid from xylose in Escherichia coli. Metab Eng. 2017;41:39–45.

    Article  CAS  Google Scholar 

  103. Liu H, Lu T. Autonomous production of 1,4-butanediol via a de novo biosynthesis pathway in engineered Escherichia coli. Metab Eng. 2015;29:135–41.

    Article  CAS  Google Scholar 

  104. Choi SY, Park SJ, Kim WJ, Yang JE, Lee H, Shin J, et al. One-step fermentative production of poly(lactate-co-glycolate) from carbohydrates in Escherichia coli. Nat Biotechnol. 2016;34:435–40.

    Article  CAS  Google Scholar 

  105. Choi SY, Kim WJ, Yu SJ, Park SJ, Im SG, Lee SY. Engineering the xylose-catabolizing Dahms pathway for production of poly(d-lactate-co-glycolate) and poly(d-lactate-co-glycolate-co-d -2-hydroxybutyrate) in Escherichia coli. Microb Biotechnol. 2017;10:1353–64.

    Article  CAS  Google Scholar 

  106. Bai W, Tai Y-S, Wang J, Wang J, Jambunathan P, Fox KJ, et al. Engineering nonphosphorylative metabolism to synthesize mesaconate from lignocellulosic sugars in Escherichia coli. Metab Eng. 2016;38:285–92.

    Article  CAS  Google Scholar 

  107. Wang J, Shen X, Lin Y, Chen Z, Yang Y, Yuan Q, et al. Investigation of the synergetic effect of xylose metabolic pathways on the production of glutaric acid. ACS Synth Biol. 2018;7:24–9.

    Article  CAS  Google Scholar 

  108. Wang J, Jain R, Shen X, Sun X, Cheng M, Liao JC, et al. Rational engineering of diol dehydratase enables 1,4-butanediol biosynthesis from xylose. Metab Eng. 2017;40:148–56.

    Article  CAS  Google Scholar 

  109. Wang J, Qu G, Xie L, Gao C, Jiang Y, Sun Z, et al. Engineering of a thermophilic dihydroxy-acid dehydratase to enhance its dehydration ability on glycerate to pyruvate and its application in in vitro synthetic enzymatic biosystems. Authorea. 2021. https://doi.org/10.22541/au.162601310.07045038/v1.

    Article  Google Scholar 

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Acknowledgements

We acknowledge the Academy of Finland for financial support.

Funding

This work was supported by the Academy of Finland (Decision 322619).

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  1. Department of Chemistry, University of Eastern Finland, 111, 80101, Joensuu, Finland

    Yaxin Ren, Veikko Eronen & Nina Hakulinen

  2. VTT Technical Research Centre of Finland Ltd, Espoo, Finland

    Martina Blomster Andberg & Anu Koivula

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  1. Yaxin Ren
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YR, VE, and NH conceptualized, implemented, and wrote the review together. NH was responsible of funding acquisition and supervision. MBA, and AK reviewed and edited the manuscript. All authors read and approved the final manuscript.

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Correspondence to Nina Hakulinen.

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Ren, Y., Eronen, V., Blomster Andberg, M. et al. Structure and function of aldopentose catabolism enzymes involved in oxidative non-phosphorylative pathways. Biotechnol Biofuels 15, 147 (2022). https://doi.org/10.1186/s13068-022-02252-5

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  • DOI: https://doi.org/10.1186/s13068-022-02252-5

Keywords

  • Aldopentose
  • Non-phosphorylative pathways
  • Pentose catabolism
  • Aldose-1-dehydrogenase
  • Lactonase
  • Sugar acid dehydratase
  • Ketoglutarate-semialdehyde dehydrogenase
  • Aldolase

Biotechnology for Biofuels and Bioproducts

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