The substrate tolerance of alcohol oxidases
Alcohols are a rich source of compounds from renewable sources, but they have to be activated in order to allow the modification of their carbon backbone. The latter can be achieved via oxidation to the corresponding aldehydes or ketones. As an alternative to (thermodynamically disfavoured) nicotinamide-dependent alcohol dehydrogenases, alcohol oxidases make use of molecular oxygen but their application is under-represented in synthetic biotransformations. In this review, the mechanism of copper-containing and flavoprotein alcohol oxidases is discussed in view of their ability to accept electronically activated or non-activated alcohols and their propensity towards over-oxidation of aldehydes yielding carboxylic acids. In order to facilitate the selection of the optimal enzyme for a given biocatalytic application, the substrate tolerance of alcohol oxidases is compiled and discussed: Substrates are classified into groups (non-activated prim- and sec-alcohols; activated allylic, cinnamic and benzylic alcohols; hydroxy acids; sugar alcohols; nucleotide alcohols; sterols) together with suitable alcohol oxidases, their microbial source, relative activities and (stereo)selectivities.
KeywordsOxidation Biocatalysis Alcohol oxidase Substrate tolerance Flavoprotein Cu-containing oxidase
Oxidation represents a fundamental reaction in nature (Hollmann et al. 2011; Turner 2011), and oxidases are a prominent subclass of redox enzymes, which use oxygen either as oxidant or as electron acceptor. This property made them particularly attractive for the production of chemicals (Vennestrom et al. 2010). In this context, the oxidation of alcohols is an important transformation in synthetic chemistry, which allows to introduce carbonyl groups, which represent excellent acceptors for C-, N-, O- and S-nucleophiles and thereby allows the extension of a given carbon backbone. Consequently, a large number of protocols has been developed, which depend on (i) transition metals in stoichiometric (e.g. Cr, Mn) or catalytic amounts (e.g. Ru, Fe), (ii) metal-free oxidations according to Swern or Pfitzner-Moffat (Pfitzner and Moffatt 1963; Omura and Swern 1978), (iii) molecular oxygen as oxidant (Tojo and Fernández 2006) and more recently, (iv) organocatalysts, such as TEMPO (Wertz and Studer 2013).
The over-oxidation of alcohols to carboxylic acids has been observed not only for choline oxidase but also for other flavoprotein oxidases, such as alditol oxidase (AldO), aryl alcohol oxidase (AAO), hydroxymethyl furfuryl oxidase (HMFO), hexose oxidase (HOX, Dbv29), isoamyl alcohol oxidase (IAO) or short- and long-chain alcohol oxidases (SCAOs, LCAOs). Labelling studies proved the existence of the aldehyde hydrate as intermediate (Van Hellemond et al. 2009), and for AAO, which naturally oxidises benzylic alcohols, NMR studies revealed that the gem-diol intermediate was favoured (Ferreira et al. 2010) (Scheme 3).
Structurally, most of the flavoprotein oxidases either belong to the glucose-methanol-choline (GMC) oxidase or the vanillyl alcohol oxidase (VAO) family. Both families have a flavin present in the active site where the binding domain and the binding mode of the flavin differ. In case of VAO, the flavin is covalently linked to a histidine, cysteine or tyrosine residue, while in the GMC family, the majority of the enzymes contain a dissociable flavin adenine dinucleotide (FAD) moiety. In P2O or CHO, a covalent linkage was found. The active sites and consequently the substrate scope of these enzymes show high variance (Fraaije et al. 1998a; Kiess et al. 1998; Leferink et al. 2008; Dijkman et al. 2013).
For Cu-containing alcohol oxidases, the oxidation stops at the aldehyde stage and over-oxidation was not observed (Monti et al. 2011).
From a biocatalytic viewpoint, alcohol oxidases are a promising group of enzymes, because they are biochemically well characterised and a broad range of enzymes have been described (Whittaker 2003; Leferink et al. 2008; Dijkman et al. 2013) which were also employed in cascade reactions (Fuchs et al. 2012; Perez-Sanchez et al. 2013; Schrittwieser et al. 2011). Depending on their role in nature, substrates for alcohol oxidases vary to a great extent in terms of substrate size and/or polarity (Turner 2011). In fungi, extracellular alcohol oxidases produce hydrogen peroxide (needed for lignin degradation by peroxidases) by oxidation of cinnamyl alcohols (e.g. coniferyl, coumaryl and sinapyl alcohol). Furthermore, hydrogen peroxide acts as antibiotic in the rhizosphere to protect roots (Monti et al. 2011). As an alternative to alcohol oxidases, NAD(P)+-dependent alcohol dehydrogenases provide a well-investigated enzyme platform for the oxidation of prim- and sec-alcohol functionalities. Although these enzymes are more abundant than alcohol oxidases, the equilibrium for oxidation is strongly disfavored but can be overcome by NAD(P)+ recycling (Hollmann et al. 2011).
In the following, an overview on the current literature of alcohol oxidases is given, by focussing on their substrate tolerance to facilitate the choice of an appropriate enzyme for a given type of alcohol substrate.
Primary aliphatic alcohols
Primary aliphatic alcohols
SCAOb from P. pastoris (C1–C4), Hansenula sp. (C1–C5), C. boidinii, T. aurantiacus and A. terreus (C1–C2)
1-Alkanols C7–C14, C16
SCAOb from A. terreus (C7); LCAOb from A. terreus (C7–C14), A. thaliana (C12, C16) and C. tropicalis (C8, C10, C12, C14)
SCAO from C. boidinii
Menon et al. 1995
Diols and triols
SCAOb from T. aurantiacus and P. pastoris
LCAOb from C. tropicalis
Eirich et al. 2004
LCAOb from A. terreus, C. tropicalis and A. thaliana
AldOb from S. coelicolor
Van Hellemond et al. 2009
AldOb from S. coelicolor
Van Hellemond et al. 2009
AldOb from S. coelicolor
Van Hellemond et al. 2009
GOasea from Fusarium NRRL 2903
Klibanov et al. 1982
AldOb from S. coelicolor and A. cellulolyticus 11B
LCAOb from A. terreus (C12) and C. tropicalis (C12, C16)
CHOb from A. globiformis
SCAOb from P. pastoris
Siebum et al. 2006
AldOb from S. coelicolor
Van Hellemond et al. 2009
SCAOb from C. boidinii, P. pastoris and Hansenula sp.
Clark et al. 1995
SCAOb from T. aurantiacus; SAOc from A. terreus; IAOb from A. oryzae
SCAOb from A. terreus
Kumar and Goswami 2009
CHOb from A. globiformis
Gadda et al. 2004
Aliphatic alcohols with a chain length of one to seven C atoms were oxidised by short-chain alcohol oxidases (SCAOs) [EC 220.127.116.11] from Pichia pastoris, Hansenula sp. (Table 1, entry 1) and Aspergillus terreus (Table 1, entry 2) while methanol and ethanol were also converted by alcohol oxidases from Candida boidinii, Thermoascus aurantiacus and A. terreus (Table 1, entry 1) (Kato et al. 1976; Siebum et al. 2006; Menon et al. 1995; Couderc and Baratti 1980; Kumar and Goswami 2006; Ko et al. 2005; Perez-Sanchez et al. 2013). In general, the activity decreases with increasing chain length of the fatty alcohol, e.g. 1-pentanol shows 24 % relative activity compared to methanol (Ko et al. 2005). SCAO from Hansenula sp. was employed together with a C–C lyase in a cascade reaction, where short-chain alcohols (methanol, ethanol, 1-propanol and 1-pentanol) were oxidised with excellent conversion to the corresponding aldehydes, which were subjected to cross-acyloin condensation with benzoin generated in situ from benzaldehyde by benzaldehyde lyase to yield 2-hydroxyketones (Shanmuganathan et al. 2012; Perez-Sanchez et al. 2013). prim-Alcohols with a chain length of 7 to 16 carbon atoms were best oxidised by long-chain alcohol oxidases (LCAOs) [EC 18.104.22.168] from A. terreus, Candida tropicalis and Arabidopsis thaliana (Table 1, entry 2). Both, SCAOs and LCAOs, are flavoproteins located in fungal microsomes (Kemp et al. 1988; Eirich et al. 2004; Kumar and Goswami 2006, Cheng et al. 2004). Terminal alcohols bearing a polar functional group, such as α,ω-diols (Table 1, entries 5 and 6) and ω-carboxy fatty alcohols (Table 1, entry 12), with a long hydrocarbon backbone were also oxidised by long-chain alcohol oxidases (Kumar and Goswami 2006).
Short-chain alcohol oxidase from several microorganisms (C. boidinii, Hansenula sp., P. pastoris and T. aurantiacus) was described to convert racemic branched alcohols (Table 1, entries 16–17) in an enantioselective fashion with conversions of 16–76 %, the non-reacted substrate enantiomers showed ees of up to 90 % for SCAO from C. boidinii (Clark et al. 1995). Isoamyl alcohol oxidase (IAO) [EC 1.1.3.x] from Aspergillus oryzae exhibits a narrow substrate range and prefers branched short-chain alcohols, such as 3-methyl-1-butanol (Table 1, entry 17) (Yamashita et al. 1999). Halogen-substituted alcohols, which were oxidised by SCAO, were used as molecular probes for mechanistic studies (Menon et al. 1995).
Saturated and unsaturated vic-1,2-diols were the substrates of choice for alditol oxidase [EC 22.214.171.124] from Streptomyces coelicolor and Acidothermus cellulolyticus (Table 1, entries 7–9, 11 and 15). This enzyme apparently prefers a glycol or 1,3-diol moiety. For rac-1-phenyl-1,2-ethanediol carrying a bulky aryl moiety, the (R)-enantiomer was preferentially oxidised by alditol oxidase (Table 1, entry 11) (Van Hellemond et al. 2009). Short (non-allylic) unsaturated alcohols lacking a second hydroxy group were completely (4-penten-1-ol) or partially (3-buten-1-ol) oxidised by short-chain alcohol oxidase from P. pastoris (Table 1, entry 14) (Siebum et al. 2006).
Another prominent enzyme of this group is choline oxidase from A. globiformis which oxidises choline and analogues, such as N,N-dimethylethanolamine, N-methylethanolamine, triethanolamine, diethanolamine and 3,3-dimethylbutan-1-ol (Table 1, entries 13 and 19) in a two-step oxidation to the corresponding carboxylic acid (Ikuta et al. 1977; Gadda et al. 2004).
Secondary aliphatic alcohols
Secondary aliphatic alcohols
SAOa from P. putida and P. vesicularis
2-Alkanols C3–C12, C16
SAOa from A. terreus (C3, C8, C12), P. putida (C3–C7) and P. vesicularis (C4–C8); SCAOb from T. aurantiacus (C3–C4), A. terreus (C8) and P. pastoris (C3); LCAOb from C. tropicalis (C10–C11, C16)
SAOa from P. putida (C5–C8), A. terreus (C8) and P. vesicularis (C6–C8)
SAOa from P. putida (C7–C9) and P. vesicularis (C7, C10)
SAOa from P. putida
Sakai et al. 1985
Cycloalkanols C6, C8
SAOa from P. vesicularis (C6) and A. terreus (C8)
SAOa from P. putida
Sakai et al. 1985
SAOa from P. vesicularis
Kawagoshi and Fujita 1997
For monomeric sec-alcohols, the relative activity of SAO from P. putida ranges between 5 and 30 % (compared to PVA). High activity for 2-octanol was found with the enzyme from P. vesicularis (83 % rel. activity), which also accepts cyclohexanol (42 % rel. activity). Its oxidised product (cyclohexanone) is used as a starting material for the synthesis of the polymer building block ε-caprolactam. sec-Alcohols bearing an additional OH group, such as 1,2-propanediol and 2,4-pentanediol, were also accepted as substrates (Table 2, entries 7 and 8); however, no details are reported about the regioselectivity of the oxidation (Sakai et al. 1985; Kawagoshi and Fujita 1997). Additionally, SCAO from T. aurantiacus, A. terreus and P. pastoris as well as LCAO from C. tropicalis showed broad activity on secondary alcohols (Table 2, entry 2) (Eirich et al. 2004; Kumar and Goswami 2009; Kjellander et al. 2013; Ko et al. 2005). Furthermore, 2-methyl-2-propanol was claimed to show 16 % relative activity with SCAO, but this tert-alcohol should be a non-substrate (Ko et al. 2005).
prim- and sec-Allylic alcohols
Small allylic alcohol was oxidised poorly by galactose oxidase (Table 3, entries 1 and 2), which prefers large analogues, such as cinnamyl alcohol (Table 3, entry 4). A mutant of galactose oxidase from Fusarium sp. oxidised cinnamyl alcohol with full conversion (Sun et al. 2002; Fuchs et al. 2012). In contrast to galactose oxidase, which does not accept sec-allylic alcohols, cholesterol oxidase from Rhodococcus erythropolis converted sterically demanding secondary allylic alcohols in a complete stereo- and enantioselective fashion with conversions up to 70 % and high to excellent ees. For methyl-substituted bicyclic substrates (Table 3, entries 7 and 8), the relative (cis) position of the hydroxyl group with respect to the methyl group were mandatory to be accepted and non-activated (non-allylic) hydroxy groups were unreactive. Even comparably small monocyclic substrates could be converted (Dieth et al. 1995). Aryl alcohol oxidase exhibits a broad substrate scope and accepts phenyl substituted allylic alcohols such as coniferyl and cinnamyl alcohol (Table 3, entries 4 and 5), as well as slim counterparts, such as 2,4-hexadien-1-ol (Table 3, entry 3), which shows that this enzyme does not necessarily need a cyclic structure, but only a conjugated system (Ferreira et al. 2005; Romero et al. 2009). 5-Hydroxymethylfurfural oxidase exhibited a similar behaviour and appears to be a promising candidate for the oxidation of allylic alcohols, as it showed excellent acceptance of cinnamyl alcohol (Table 3, entry 4) and 2,4-hexadien-1-ol (Table 3, entry 3) (Dijkman and Fraaije 2014). With cinnamyl alcohol and its p-methoxy derivative, AAO shows over-oxidation and forms the corresponding acids (Table 3, entry 4) (Guillen et al. 1992).
In the case of benzyl alcohol, two more AAOs (from A. terreus and Pleurotus ostreatus) showed activity, as well as SCAO from C. boidinii and T. aurantiacus, SAO from P. vesicularis and HMFO from Methylovorus sp. (Table 4, entry 2) (Kumar and Rapheal 2011). Although not visible on a non-chiral substrate, AAO acts in a stereoselective fashion by removing the pro-R hydride as shown by deuterium experiments (Hernandez-Ortega et al. 2012b). Various substituents on the aromatic ring system are freely tolerated: Although wild-type galactose oxidase from Fusarium has a broad substrate scope for benzylic prim-alcohols, the activity was considerably increased by mutations. For instance, all regioisomers of pyridine methanol were transformed by a R330K, Q406T-mutant of galactose oxidase, which showed up to 2000-fold enhanced activity towards 2-pyridine methanol compared to the canonical d-galactose (Sun et al. 2002). Meta- and para-substituted substrates (3-F, 3-Br, 3-Cl, 3-NO2, 4-F, 4-Cl, 4-Br, 4-I, 4-NO2, 4-OMe, 4-SMe, 4-Me, 4-CF3) (Table 4, entries 4, 6, 7, 17, 18, 20–24) were converted with up to 20-fold variation of relative rates (Whittaker and Whittaker 2001).
Secondary aryl alcohols undergo kinetic resolution with partly excellent ees using an (R)-selective mutant of galactose oxidase from Fusarium sp. created by directed evolution (Table 4, entries 28–40) (Escalettes and Turner 2008). The same group also reported a rare example of the successful recognition of an atropisomeric pair of enantiomers possessing axial chirality (Table 4, entry 41) (Yuan et al. 2010). Furthermore, an engineered variant of HMFO was able to oxidise phenylethanol in a stereoselective fashion (Dijkman et al. 2015).
Methoxy groups (Table 4, entry 6) were accepted independently from the position on the ring with comparable activities relative to unsubstituted benzyl alcohol, whereas para-substituted analogues reacted more than fivefold faster with aryl alcohol oxidase. Furthermore, dimethoxy benzyl alcohols (Table 4, entries 8 and 9) were converted by aryl alcohol oxidase with high activity (Hernandez-Ortega et al. 2011; Hernandez-Ortega et al. 2012a). In particular, 3,4-dimethoxybenzyl alcohol (veratryl alcohol, Table 4, entry 9) was converted with 326 % activity, while the 2,4-substituted pendant (Table 4, entry 8) was accepted with 178 % activity relative to benzyl alcohol (Guillen et al. 1992). Sterically demanding 3,4,5-trimethoxybenzyl alcohol (Table 4, entry 10) was converted slowly. Besides methoxy groups, also hydroxy groups, combinations thereof and even a meta-substituted phenoxy group were accepted (Table 4, entries 12–16). The hydroxy substrates (Table 4, entries 12 and 13) were poorly converted compared to the 3-phenoxybenzyl alcohol (Table 4, entry 16) which was well accepted (Guillen et al. 1992). Additionally, the name-giving enzyme for the VAO family, vanillyl alcohol oxidase (VAO) [EC126.96.36.199] acts on 4-hydroxy-3-methoxybenzyl alcohol (vanillyl alcohol, Table 4, entry 15) (de Jong et al. 1992; Van den Heuvel et al. 1998; Fraaije et al. 1998b; Van Den Heuvel et al. 2000; Van den Heuvel et al. 2001a; Van den Heuvel et al. 2001b). While the enzyme seems to accept bulky substituents, e.g. bearing a phenoxy group, additional methoxy or especially hydroxy groups (Table 4, entries 12–16) cause unfavourable interactions in the active site. The aryl alcohol oxidase from P. eryngii also acts on 4-hydroxy-substituted α-aryl alcohols (Table 4, entry 13) (Guillen et al. 1992). Piperonyl alcohol (1,3-benzodioxole-5-methanol, Table 4, entry 11), a building block in epinephrine synthesis, was oxidised with full conversion by galactose oxidase from Fusarium sp. (Fuchs et al. 2012). A broad range of chloro- and fluoro-substituted aryl alcohols were accepted by both aryl alcohol oxidase and galactose oxidase (Table 4, entries 20 and 21) (Guillen et al. 1992; Whittaker and Whittaker 2001; Romero et al. 2009). The only exception being meta-chlorobenzyl alcohol, which was not converted at all. A substrate which is sterically demanding and well accepted by AAO is 2-naphthalene methanol (Table 4, entry 26). It showed a relative activity of 746 % compared to the monocyclic substrate analogue (Table 4, entry 2). In conclusion, the position of substituents and their polarity seem to play a crucial role in substrate acceptance. The recently characterised 5-hydroxymethylfurfural oxidase from Methylovorus sp. MP688 showed a broad substrate acceptance of various furfuryl alcohols (Table 4, entry 42), but it also showed activity on benzylic alcohols with substituents in para-position (Table 4, entries 3 and 5) and vanillyl alcohol (Table 4, entry 15) (Dijkman and Fraaije 2014). In view of the growing importance of furan derivatives, such as hydroxymethyl furfural, which can easily be obtained via double elimination of H2O from hexoses or pentoses and hence constitute a promising C source for organic synthesis (Schwartz et al. 2014), HMFO has a considerable potential to be used in large-scale applications. In a recent study, site-directed mutagenesis allowed to boost the activity of HMFO on 5-formyl-2-furancarboxylic acid leading to improved yields of 2,5-furandicarboxylic acid, which is a promising monomer for polyester production from renewable resources (Dijkman et al. 2015).
Due to the presence of numerous hydroxy groups, carbohydrates are usually bound in the active site of proteins via a tight hydrogen-bonding network, which is not possible for lipophilic mono-alcohols or diols. Consequently, one might surmise, that alcohol oxidases acting on lipophilic (mono) alcohols would not accept polar carbohydrates, and vice versa. However, comparison of Tables 1 and 3 shows that many sugar alcohol oxidases are also surprisingly active on small non-polar alcohols, in particular galactose oxidase and alditol oxidase.
The relative reactivity of hydroxy groups in sugars can be associated with different subgroups of alcohol oxidases, most of which possess a strong regio-preference for a specific hydroxyl group, which is exemplified on a schematic hexose (Scheme 12). With its hemiacetal structure, the anomeric OH is most reactive, which can be oxidised by glucose oxidase (GOX), hexose oxidase (HOX) and oligosaccharide oxidases forming the corresponding sugar lactone. Next, the terminal prim-OH is sterically least hindered among the non-activated hydroxy groups; it can be selectively oxidised by GOase to yield the aldehyde; no over-oxidation to the acid is observed in this case. Due to small steric and electronic differences, internal secondary hydroxy groups show very similar reactivities, they are oxidised by P2O with mixed regioselectivities with a prevalence of C2 > C3 yielding ketoses. C3-Oxidation products are only formed on 2-deoxy and methylated sugars.
Furthermore, also glucooligosaccharide oxidase (GOO) [EC 1.1.3.x] from various sources oxidised d-glucose and its oligomers at C1 (Huang et al. 2005). Lactose oxidase (LAO) [EC 1.1.3.x] from Microdochium nivale displayed a similar substrate preference. Cellobiose (Table 7, entry 18) with 100 % relative activity was the preferred substrate, whereas di-sugars as d-maltose (84 % rel. activity) and d-lactose (52 % rel. activity) were also well accepted (Table 7, entries 9,10). Furthermore, the monosugars d-glucose (69 % rel. activity) and d-galactose (31 % rel. activity) were both oxidised at C1 (Xu et al. 2001) (Table 7, entries 1 and 2). Moreover, Pezzotti and Therisod synthesised aldonic acids starting with C6 sugars (d-galactose, d-xylose, d-mannose and 2-deoxy-d-glucose) employing glucose oxidase for the oxidation of the C1 hydroxy group (2006). HOX [EC 188.8.131.52] from Chondrus crispus is an enzyme with a fairly broad substrate scope for the oxidation of sugars at C1. Hexose oxidase accepted d-xylose, d-arabinose and d-glucose containing di-sugars, like d-lactose and d-cellobiose (Table 7, entries 4, 5, 10 and 18) (Poulsen and Hostrup 1998; Savary et al. 2001; Rand et al. 2006).
(ii) The sterically least hindered prim-OH group of sugars can be selectively oxidised by copper-containing galactose oxidase (Scheme 12). Relative activities were measured in relation to the reactivity of the C6-hydroxy group of d-galactose as the canonical substrate. The most prominent galactose oxidase from Fusarium converted d-galactose containing substrates d-lactose (10 % conv.), lactitol (20 % conv.), lactobionic acid and the synthetic disaccharide and laxativum d-lactulose completely (Table 7, entries 8, 10 and 17) (Siebum et al. 2006). For substrate acceptance of GOase, the axial position of the C4 position is crucial. The di-sugars d-melibiose, d-raffinose and d-stachyose were good substrates for galactose oxidase (83 % rel. activity for d-melibiose, up to 161 % rel. activity for d-stachyose) (Table 7, entries 14–16) (Mendonca and Zancan 1987). For d-fructose (Table 7, entry 7), a GOase mutant from Fusarium seems to be an appropriate biocatalyst (Deacon et al. 2004). Recently, a FAD-containing hexose oxidase was discovered. The so-called Dbv29 oxidised a glycopeptide at C6 to the corresponding carboxylic acid in a two-step reaction (Li et al. 2007; Liu et al. 2011).
(iii) d-Glucose (Table 7, entry 1) was also oxidised by the flavoenzyme pyranose oxidase (P2O) [EC 184.108.40.206] (Giffhorn 2000), which was obtained from several fungi (Peniophora sp., Trametes sp., Tricholoma matsutake and Gloeophyllum sepiarium). It oxidises hydroxyl groups on the C2 position, but also oxidation at C3 can occur (Scheme 12) (Kujawa et al. 2006). The process based on C2 oxidation of d-glucose followed by catalytic hydrogenation yielding d-fructose is known as ‘Cetus process’, which was also utilised for the synthesis of d-tagatose (Geigert et al. 1983; Freimund et al. 1996). d-Galactose was a rather poor substrate for pyranose oxidase from P. gigantea (Table 7, entry 2) (Freimund et al. 1998; Cook and Thygesen 2003; Bastian et al. 2005). Furthermore, the configuration on C4 played an important role in substrate acceptance. d-Allose (94 % overall yield), d-xylose (100 % overall yield) and d-mannose (only moderate rel. activity of 23 %) were all oxidised by pyranose oxidase originating from several microorganisms (Table 7, entries 3, 4 and 6) (Danneel et al. 1993; Freimund et al. 1998; Takakura and Kuwata 2003; Bannwarth et al. 2006; Machida and Nakanishi 1984). Pyranose oxidase accepted the di-sugars d-trehalose (54 % rel. activity), d-gentiobiose (1 % conversion) and d-maltose (8–56 % rel. activity) as substrates (Table 7, entries 9, 13 and 19) (Danneel et al. 1993; Freimund et al. 1998; Takakura and Kuwata 2003). Moreover, P2O was used as a biocatalyst for the C2 oxidation of disaccharides to obtain 2-keto-aldopyranose intermediates (Leitner et al. 2001) and the di-sugar d-sucrose (Table 7, entry 11) was fully converted by P2O in a multistep process (Seto et al. 2008).
Deoxy sugars were often employed in kinetic studies to investigate the catalytic mechanism of enzymes. 1-, 2-, 3- and 6-deoxy-d-glucose and 2-deoxy-d-galactose (Table 7, entries 20–24) were used for this purpose showing full conversions. The enzymes exhibited their expected regioselectivity. For pyranose oxidase, activity was observed for 2-deoxy-d-glucose (52 % rel. activity) for oxidation at C3 (Table 7, entry 21). 1-Deoxy-d-glucose (Table 7, entry 20) was converted by pyranose oxidase (8 % rel. conversion P2O from Phanerochaete gigantea, 22 % from Trametes versicolor and 69 % from T. matsutake). The substrate 3-deoxy-d-glucose was almost as good for pyranose oxidase as the natural one, but 6-deoxy-d-glucose showed significantly diminished relative conversion rate of 15 % (Table 7, entries 22 and 23). Glucose oxidase also shows activity for 2-deoxy-d-glucose and 6-deoxy-d-glucose (Table 7, entries 21 and 23). Galactose oxidase showed 74 % relative activity for 2-deoxy-d-galactose (Table 7, entry 24) (Danneel et al. 1993; Freimund et al. 1998; Takakura and Kuwata 2003; Leskovac et al. 2005; Siebum et al. 2006; Masuda-Nishimura et al. 1999).
In addition, various sugar derivatives were tested: 4-O-Methylated sugars were accepted by pyranose oxidase and galactose oxidase (Schoevaart and Kieboom 2004). With pyranose oxidase, oxidation occurred at C3. Phenyl- and hexyl-glucosides were well accepted, but underwent a glycosyl transfer reaction forming a disaccharide (Table 7, entry 28). These bulky substrates indicate that the size of the active site is not a limiting factor. Nitro sugars were tested with pyranose oxidase, and glycosyl transfer occurred yielding a 4:1 ratio of 1-6 vs. 1-3 di-sugar at C2 position in 15 % overall yield. At C4 position, a 2:1 mixture of 1-6 vs. 1-3 di-sugar was obtained in 24 % yield. α-d-Glucosyl fluoride (Table 7, entry 31) was a moderate substrate for pyranose oxidase from P. gigantea (40 % yield, Danneel et al. 1993; Freimund et al. 1998). Pyranose oxidase also converted the unnaturall-sugar l-sorbose completely (Table 7 entry 34). Mono- and poly-fluorinated galactose analogues were oxidised by galactose oxidase (Table 7, entry 33) (Ioannou et al. 2011), and also hydroxyacetone derivatives represented excellent substrates. Dihydroxyacetone (Table 7, entry 36) was also oxidised at a fair rate by glycerol oxidase (GlycOx) from Aspergillus japonicus (59 % rel. activity) (Uwajima and Terada 1980). Furthermore, galactose oxidase was active on guaran, a galactomannan (Table 7, entry 38) (47 % rel. activity) (Mendonca and Zancan 1987). This enzyme was also applied for the oxidation of the nucleotide sugars uridine 5′-diphospho-α-d-galactose and uridine 5′-diphospho-N-acetyl-α-d-galactosamine (Table 7, entry 39) for subsequent biotinylation (Bülter et al. 2001; Namdjou et al. 2007).
Sugar alcohols and amino sugar alcohols
Sugar alcohols and amino sugars
Cofactor presence, substrate scope and propensity for over-oxidation of alcohol oxidases
Substrate (major activities)
Alditol oxidase (AldOx)
Primary alcohols, sugar alcohols
Aryl alcohol oxidase (AAO)
Benzylic alcohols, allylic alcohols
Chitooligosaccharide oxidase (ChitO)
Cholesterol oxidase (ChOx)
Sterols, allylic alcohols
Choline Oxidase (CHO)
Galactose oxidase (GOase)
Benzylic alcohols, sugars
Glucooligosaccharide oxidase (GOO)
Glucose oxidase (GOX)
Glycerol Oxidase (GlycOx)
Glycerol 3-phosphate oxidase (GPO)
Glycolate oxidase (GlyO)
Hexose oxidase (HOX)
Hydroxymethylfurfural oxidase (HMFO)
Benzylic alcohols, allylic alcohols
(S)-2-Hydroxy acid oxidase (HAOX)
Isoamyl alcohol oxidase (IAO)
Branched aliphatic alcohols
L-lactate oxidase (LLO)
Lactose oxidase (LAO)
Long-chain alcohol oxidase (LCAO)
Secondary alcohol oxidase (SAO)
Secondary aliphatic alcohols
Short-chain alcohol oxidase (SCAO)
Pyranose oxidase (P2O)
Vanillyl alcohol oxidase (VAO)
Summary and outlook
The broad substrate scope coupled with high regio- and stereoselectivity makes alcohol oxidases a fantastic tool for the oxidation of primary and secondary alcohols using molecular oxygen as an alternative to traditional chemical methods. Owing to their mechanism, copper-depending oxidases selectively yield aldehydes from primary alcohols, while over-oxidation to furnish carboxylic acids may take place to a varying degree with flavin-depending oxidases. For a broad range of alcohols—non-activated prim- and sec-alcohols, activated allylic, cinnamic and benzylic alcohols, hydroxy acids, hydroxy steroids, carbohydrates and derivatives thereof—alcohol oxidases are available from various microbial sources, which are reviewed with respect to their substrate tolerance to facilitate the choice of the optimal enzyme for a given alcohol substrate.
Funding by the Austrian Science Fund (FWF), within the DK Molecular Enzymology (project W9), an Erwin-Schrödinger fellowship (J3466) and the Austrian BMWFW, BMVIT, SFG, Standortagentur Tirol, Government of Lower Austria and ZIT through the Austrian FFG-COMET-Funding Program is gratefully acknowledged.
Conflict of interest
The authors declare that they have no competing interests.
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