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.


Introduction
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).
In a related fashion, alcohol oxidases convert primary and secondary alcohols to aldehydes and ketones, respectively. During this reaction, molecular oxygen is reduced to hydrogen peroxide. In order to avoid enzyme deactivation, a catalase is usually employed, particularly on preparative scale. For screening purposes, a spectrophotometric assay based on horse radish peroxidase (HRP) together with a suitable artificial electron acceptor, such as 2,2′ -azino-bis(3ethylbenzthiazoline-6-sulfonic acid) (ABTS) may be employed (Scheme 1). The ABTS-radical generated shifts its absorption maximum (Baron et al. 1994;Uwajima and Terada 1980).
Although cofactor-lacking oxidases are reported (Fetzner and Steiner 2010), commonly used alcohol oxidases depend on flavin (Macheroux et al. 2011;Dijkman et al. 2013) or a Electronic supplementary material The online version of this article (doi:10.1007/s00253-015-6699-6) contains supplementary material, which is available to authorized users. metal (Cu) as a cofactor (Whittaker 2003), which mediates the electron transfer. In flavoprotein oxidases, the oxidation proceeds via two half reactions, where the alcohol is first oxidised by a two-electron transfer during the reductive half reaction, yielding reduced flavin. The oxidised flavin is regenerated by a stepwise single-electron transfer via the oxidative half reaction, which requires triplet oxygen, as it is a spinforbidden reaction. Hence, di-oxygen acts as single-electron acceptor and forms superoxide (O 2 −· ), stabilised by a positively charged histidine residue (Dijkman et al. 2013;Wongnate et al. 2014). Another single-electron transfer yields a covalent hydroperoxy flavin intermediate, which eliminates hydrogen peroxide and re-forms oxidised flavin (Scheme 2) (Gadda 2012). The highly unstable C 4 a-hydroperoxyflavin intermediate has only been detected for pyranose oxidase (P2O) (Mattevi 2006;Chaiyen et al. 2012;Wongnate and Chaiyen 2013).
The oxidation of primary alcohols catalysed by flavoprotein oxidases does not necessarily stop at the aldehyde stage, but may further proceed to the corresponding carboxylic acid. This second oxidation step is mechanistically less investigated, but it is obvious that the actual substrate is the aldehyde hydrate (gem-diol), rather than its carbonyl form, because hydride abstraction from the former yields a doubly resonance-stabilised oxocarbenium cation, which upon expulsion of H + furnishes the carboxylic acid. In contrast, hydride abstraction from the carbonyl form would lead to a highly unstable (hypothetical) acylium cation, which would be quenched by a water molecule (Scheme 3).
This mechanism for over-oxidation has been proposed for choline oxidase (CHO), whose natural role is the formation of N-trimethylammonium glycine ('betaine') from choline via the aldehyde hydrate through a two-step oxidation (Scheme 4) (Rungsrisuriyachai and Gadda 2008).
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 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  (Fraaije et al. 1998a;Kiess et al. 1998;Leferink et al. 2008;Dijkman et al. 2013).
Another redox cofactor found in alcohol oxidases, such as galactose oxidase (GOase), is the transition metal copper, whose role in catalysis is well described in several reviews (Ridge et al. 2008;Guengerich 2013). Since only a single copper(I) ion is found in the active site, it seems surprising that a two-electron transfer can occur. Detailed investigations revealed that the latter proceeds via two consecutive singleelectron transfer steps. Thus, abstraction of the first electron by Cu 2+ yields Cu + , which transfers its electron onto a tyrosine residue, which forms a transient radical anion (Monti et al. 2011). The latter is stabilised by a rare covalent thioether bridge with an adjacent cysteine (Ito et al. 1991). The second electron transfer yields a Cu + -tyrosine radical. From this, oxygen accepts two electrons (Wang 1998;Whittaker 2003) (Scheme 5). GOase from Fusarium NRRL 2903 is the most prominent member of Cu-containing alcohol oxidases and belongs to the family of radical copper oxidases, a family with a wide phylogenetic distribution and broad range of functions. The crystal structure of the enzyme revealed that a mononuclear copper ion is centred in a distorted pyramid structure, which is coordinated by two tyrosine residues (Tyr272 and Tyr495) and two histidine side chains (His496 and His581) (Whittaker and Whittaker 2001).
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.

Secondary aliphatic alcohols
Racemic secondary aliphatic alcohols are interesting substrates, because enantioselectivities in kinetic resolution are usually much higher than with prim-alcohols bearing a stereogenic centre. In contrast to prim-alcohols, which may undergo over-oxidation to carboxylic acids, the oxidation products derived from sec-alcohols are solely ketones (Scheme 7). Compared to prim-alcohol oxidases, enzymes acting on secondary alcohols are less abundant, but several enzymes were found to be highly active (  (Sakai et al. 1985;Kawagoshi and Fujita 1997), and it was discovered that one non-heme Fe 2+ species is present in the enzyme. To date, it remains unclear whether the iron species serves as a cofactor like the copper in galactose oxidase or if it does not participate in catalysis at all.
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).

Activated alcohols Allylic alcohols
In contrast to saturated (non-activated) aliphatic alcohols, allylic and benzylic alcohols are much easier to oxidise, because radicals and carbene ions occurring as intermediates are resonance stabilised (see Electronic Supplementary Material, Scheme S1). Owing to their high intrinsic reactivity, allylic alcohols are easily oxidised by a broad range of alcohol oxidases, such as copper-containing galactose oxidase (GOase Table 3) (Guillen et al. 1992;Dieth et al. 1995;Sun et al. 2002;Dijkman and Fraaije 2014).
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 parasubstituted 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, 3,4dimethoxybenzyl 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 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 (1,3benzodioxole-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 H 2 O 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).

α-Hydroxy acids
Owing to the negative charge of α-hydroxy acids at neutral pH, the latter are oxidised by a subgroup of flavoprotein oxidases, which are specific for this type of polar substrate and furnish the corresponding α-ketoacids (Scheme 10). On a first glimpse, this transformation appears to have little value, because it goes in hand with the destruction of a chiral centre. However, αhydroxy acids are usually more easily accessible than the corresponding sensitive α-keto-analogues, which are prone to decarboxylation; this transformation is of practical value, and in addition, racemic α-hydroxy acids undergo kinetic resolution with a preference for the (S)-enantiomer (Turner 2011).

Sterols
The bioactivity of steroids strongly depends on their substitutional pattern, which is dominated by secondary hydroxy groups in αor β-positions, which upon oxidation furnish keto-steroids. This transformation can be achieved in a regio-and stereoselective fashion by alcohol oxidases. Owing to the spacious molecular framework, it is conceivable that alcohol oxidases acting on steroids have a strong preference for large substrates and are generally not ideally suited for small alcohols (Scheme 11).
Cholesterol oxidase (ChOx) [EC 1.1.3.6] found in Streptomyces hygroscopicus, Rhodococcus and Brevibacterium sterolicum is the enzyme of choice for the oxidation of the secondary alcohol function at C 3 , which leads to rare ketosteroids (Table 6). From a biochemical point of view, it is remarkable that cholesterol oxidases are strictly FAD containing, although they belong to two different families: Cholesterol oxidase from Streptomyces is a member of the GMC oxidase family, whereas B. sterolicum ChOx belongs to the VAO family. Remarkably, most cholesterol oxidases are bifunctional enzymes (Pollegioni et al. 1999;Gadda et al. 1997;Pollegioni et al. 2009;Vrielink and Ghisla 2009), as they not only oxidise the alcohol functionality at C 3 yielding 5-cholesten-3-one but also mediate the isomerisation of the C 5 -C 6 double bond of the latter into conjugation with the newly formed keto-function by assistance of an active-site glutamate residue to furnish the corresponding 4-en-3-one, as demonstrated in detail with ChOx from B. sterolicum (Kass and Sampson 1995) (Scheme 11). The enzyme exhibited a surprisingly broad substrate scope, and a variant from R. erythropolis even lacks enantiospecificity at the C 3 position (Dieth et al. 1995;Biellmann 2001). For the enzyme from Rhodococcus sp., moderate activities (relative to the natural substrate cholesterol) on β-sitosterol (80 % rel. activity) and stigmasterol (78 % rel. activity) were found by Wang et al. (2008) (Table 6, entries 6 and 7). Furthermore, the enzyme was active on cholestanol, 7dehydrocholesterol and dehydroepiandrosterone (15-37 % rel. activity) (Table 6, entries 2, 4 and 8), and 5 % relative activity was found on 5α-androstane-3α,17β-diol (Table 6, entry 11) (Labaree et al. 1997;Toyama et al. 2002;Wang et al. 2008, Scheme 10 Enzymatic oxidation of hydroxy acids by hydroxy acid oxidases  Fujishiro et al. 2002;Xiang and Sampson 2004). Moreover, cholesterol oxidase from B. sterolicum was employed for the oxidation of 7α-and 7β-hydroxycholesterol (90 % conv.) ( Table 6, entry 3) in a chemoenzymatic multistep synthesis (Alexander and Fisher 1995).

Sugar-related alcohols Sugars
Although sugars constitute the most abundant group of renewable compounds/materials (Straathof 2014), their polyhydroxy structure imposes several unsolved problems in view of their utility as starting materials in organic synthesis: (i) they possess only a single type of functional group-the hydroxy group, and (ii) there are too many of them with similar reactivity (Scheme 12). This causes a selectivity problem, which is usually circumvented by tedious and inefficient protection-deprotection chemistry. (iii) Furthermore, except for the anomeric carbon, the carbon framework is inaccessible to C-C extension/modification, because the [CH-OH] moiety cannot be directly accessed without prior activation of the hydroxy group. In this context, regioselective oxidation of OH groups in sugars at the expense of O 2 offers an elegant method to introduce a carbonyl group, which is an ideal acceptor for C nucleophiles in C-C bond forming reactions.
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 C 2 >C 3 yielding ketoses. C 3 -Oxidation products are only formed on 2-deoxy and methylated sugars.
(i) The most reactive anomeric hydroxy group in sugars can be selectively oxidised by a range of well-studied oxidases (Scheme 12): D-Glucose (Table 7, entry 1) is the natural substrate of the flavoenzyme GOX [EC 1.1.3.4], well studied from Aspergillus niger, which displayed a very narrow substrate spectrum and oxidises glucose at the C 1 position (Nakamura and Ogura 1968). Furthermore, chitooligosaccharide oxidase (ChitO) [EC 1.1.3.x] from Fusarium graminearum catalyses the oxidation of C 1 of D-glucose. The catalytic activity was improved by mutation (Heuts et al. 2007a), and the wild-type and mutant enzymes also accepted cellulose degradation products like cellobiose, cellotriose and cellotetraose (Table 7, entry 18). Mutants of chitooligosaccharide oxidase also accepted Dlactose and D-maltose besides the before mentioned D-glucose oligomers (Table 7, entries 9 and 10) (Heuts et al. 2007a). Variants obtained by further mutagenesis studies showed a switch in the preference for the oligosugar preference as well as improved activities on D-lactose, D-maltose and D-glucose (Ferrari et al. 2015).
(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 C 6 -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 C 4 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 C 6 to the corresponding carboxylic acid in a twostep reaction (Li et al. 2007;Liu et al. 2011).

Sugar alcohols and amino sugar alcohols
Several enzymes were reported to oxidise sugar alcohols to the corresponding aldoses, and in case of flavoprotein oxidases, aldonic acids were obtained via over-oxidation. FAD-containing alditol oxidase (AldO) [EC 1.1.3.41] has shown a broad acceptance for sugar alcohols: AldO from Streptomyces sp. and thermophilic A. cellulolyticus acted on several D-and even L-sugar alcohols (Table 8) and oxidised them to the corresponding aldoses or even further to carboxylic acids. D-Galactitol, D-xylitol, D-sorbitol, D-mannitol, L-threitol and prochiral glycerol (Table 8, entries 1-5, 9) were tested as substrates in kinetic studies (Heuts et al. 2007b;Forneris et al. 2008; Van Hellemond e t a l . 2 0 0 9 ; M u r o o k a a n d Ya m a s h i t a 2 0 0 1 ; Drueckhammer et al. 1991;Yamashita et al. 2000). Glycerol was oxidised to L-glyceraldehyde as a building block for a follow-up aldolase reaction in a multienzyme cascade (Franke et al. 2003). The latter is also oxidised by the Cu-containing glycerol oxidase which exhibited excellent activity towards glycerol, which was selected as a name-giving substrate (Uwajima and Terada 1980;Uwajima et al. 1984). The building block dihydroxyacetone phosphate (DHAP), which is a popular C donor in asymmetric aldol reactions, can be obtained using flavoprotein glycerol 3-phosphate oxidase (GPO) [EC 1.1.3.21] for the oxidation of L-glycerol 3-phosphate (Table 8, entry 10) at the sec-OH (Babich et al. 2011). Furthermore, also copper-containing galactose oxidase from Fusarium exhibited a broad acceptance of sugar alcohols without acid formation (Table 8, entries 1, 2, 5, 6 and 8).

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 overoxidation 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.