A biocatalytic method for the chemoselective aerobic oxidation of aldehydes to carboxylic acids

Article abstract of DOI:10.1039/c8gc01381k

Herein, we present a study on the oxidation of aldehydes to carboxylic acids using three recombinant aldehyde dehydrogenases (ALDHs). The ALDHs were used in purified form with a nicotinamide oxidase (NOx), which recycles the catalytic NAD⁺ at the expense of dioxygen (air at atmospheric pressure). The reaction was studied also with lyophilised whole cell as well as resting cell biocatalysts for more convenient practical application. The optimised biocatalytic oxidation runs in phosphate buffer at pH 8.5 and at 40 °C. From a set of sixty-one aliphatic, aryl-Aliphatic, benzylic, hetero-Aromatic and bicyclic aldehydes, fifty were converted with elevated yield (up to >99%). The exceptions were a few ortho-substituted benzaldehydes, bicyclic heteroaromatic aldehydes and 2-phenylpropanal. In all cases, the expected carboxylic acid was shown to be the only product (>99% chemoselectivity). Other oxidisable functionalities within the same molecule (e.g. hydroxyl, alkene, and heteroaromatic nitrogen or sulphur atoms) remained untouched. The reaction was scaled for the oxidation of 5-(hydroxymethyl)furfural (2 g), a bio-based starting material, to afford 5-(hydroxymethyl)furoic acid in 61% isolated yield. The new biocatalytic method avoids the use of toxic or unsafe oxidants, strong acids or bases, or undesired solvents. It shows applicability across a wide range of substrates, and retains perfect chemoselectivity. Alternative oxidisable groups were not converted, and other classical side-reactions (e.g. halogenation of unsaturated functionalities, Dakin-Type oxidation) did not occur. In comparison to other established enzymatic methods such as the use of oxidases (where the concomitant oxidation of alcohols and aldehydes is common), ALDHs offer greatly improved selectivity.

Full text of DOI:10.1039/c8gc01381k

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DOI: 10.1039/C8GC01381K
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ARTICLE
A biocatalytic method for the chemoselective aerobic oxidation of
aldehydes to carboxylic acids
Tanja Knaus a, Vasilis Tseliou a , Luke D. Humphreys b,c, Nigel S. Scrutton d and Francesco G. Mutti a,*
Received 00th January 20xx,
Accepted 00th January 20xx
Herein, we present a study on the oxidation of aldehydes to carboxylic acids using three recombinant aldehyde
dehydrogenases (ALDHs). The ALDHs were used in purified form with a nicotinamide oxidase (NOx), which recycles the
catalytic NAD+ at the expense of dioxygen (air at atmospheric pressure). The reaction was studied also with lyophilised whole
cell as well as resting cell biocatalysts for more convenient practical application. The optimised biocatalytic oxidation runs
in phosphate buffer at pH 8.5 and at 40 °C. From a set of sixty-one aliphatic, aryl-aliphatic, benzylic, hetero-aromatic and
bicyclic aldehydes, fifty were converted with elevated yield (up to >99%). The exceptions were a few ortho-substituted
benzaldehydes, bicyclic heteroaromatic aldehydes and 2-phenylpropanal. In all cases, the expected carboxylic acid was
shown to be the only product (>99% chemoselectivity). Other oxidisable functionalities within the same molecule (e.g.
hydroxyl, alkene, and heteroaromatic nitrogen or sulphur atoms) remained untouched. The reaction was scaled for the
oxidation of 5-(hydroxymethyl)furfural (2 g), a bio-based starting material, to afford 5-(hydroxymethyl)furoic acid in 61%
isolated yield. The new biocatalytic method avoids the use of toxic or unsafe oxidants, strong acids or bases, or undesired
solvents. It shows applicability across a wide range of substrates, and retains perfect chemoselectivity. Alternative oxidisable
groups were not converted, and other classical side-reactions (e.g. halogenation of unsaturated functionalities, Dakin-type
oxidation) did not occur. In comparison to other established enzymatic methods such as the use of oxidases (where the
concomitant oxidation of alcohols and aldehydes is common), ALDHs offer greatly improved selectivity.
DOI: 10.1039/x0xx00000x
www.rsc.org/
have been investigated using various metal salts or metal-
complexes in catalytic amounts (e.g. CuCl, AgNO3, Bi2O3, CoCl2,
pyridinium chlorochromate, VO(acac)2, Ni(dmp)2, CH3ReO3,
Introduction
Traditional methods for the oxidation of aldehydes to carboxylic
acids are no longer sustainable. In fact, they require
stoichiometric amounts (or more) of chemical reagents such as
toxic transition-metal oxidants (e.g. Cr or Ru oxides), or
different types of salts (e.g. permanganate, bromine,
hypochlorite, chlorite, perborate), or even Ag2O in combination
with sodium cyanide.1-13 Furthermore, the use of strong
oxidants in excess often produces side-products when multiple
functional groups are available for oxidation. A classic example
is the concomitant oxidation of an aldehyde group and the
halogenation of aliphatic or aromatic alkene groups within the
same molecule.9-11
Ag2O-IPr, Cu(acac)SIMes), and in the presence of stoichiometric
amounts of an oxidant (e.g. tert-BuOOH, H2O2, H5IO6) or
dioxygen.14-23 This second generation of methods provided
some improvements in terms of chemoselectivity, relatively
simple reaction procedures and complementary substrate
scope (e.g. depending on the method of choice and related
mechanism, substituted aromatic aldehydes bearing either
electron donating or electro withdrawing substituents can be
converted with high yields). The disadvantages in many cases
are that a considerable amount of metal-catalyst is still required
along with “ungreen” organic solvents as reaction medium.
Heterogeneous metal catalysts have also been developed for
the catalytic oxidation of aldehydes to carboxylic acids at the
expense of air, pure dioxygen or tert-BuOOH as oxidising agent.
Notable heterogeneous systems include: Au-CeO2 and Au-C
catalysts; polyoxometalates of Ni, Co, Mn and Cu; Co-
acetylacetonates on organically modified 3-aminopropyl silica;
Co(II)-substituted heteropolyacid supported on silica; Fe(III)-
hybrid polyoxometalate.24-27 These methods are currently
limited by either the relatively high loading of the
heterogeneous metal-catalyst (typically 5 mol%) or problems
related to long-term activity and recyclability or moderate
selectivity and substrate scope.
With the goal of improving the environmental footprint for the
selective oxidation of aldehydes, several alternative methods
a.Van’t Hoff Institute for Molecular Sciences, HIMS-Biocat, University of
Amsterdam, Science Park 904, 1098 XH, The Netherlands. E-mail: f.mutti@uva.nl.
b.GlaxoSmithKline Medicines Research Centre, Gunnel’s Wood Road, Stevenage,
SG1 2NY, UK.
c. Current address: Gilead Alberta ULC, 1021 Hayter Road NW, Edmonton, AB T6S
1A1, Canada.
d.Manchester Institute of Biotechnology, School of Chemistry, The University of
Manchester, 131 Princess Street, Manchester, M1 7DN, UK.
† Electronic Supplementary Information (ESI) available: A full list of the substrates
with related structures, detailed descriptions of the expression and purification of
the enzymes, full data sets and explanation of the biocatalytic reactions, NMR
spectra, GC analytical methods and data as well as GC traces. See
DOI: 10.1039/x0xx00000x
Despite the large variety of available metal-based catalytic
protocols, it was already shown more than seventy years ago
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that H2O2 can oxidise aldehydes to carboxylic acids without the compounds.45-48 Aldehyde dehydrogenases (EC 1.2.1), which
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need for a metal catalyst.28 However, the low reaction rate catalyse the oxidation of aldehydes via the nicotinamide
DOI: 10.1039/C8GC01381K
makes the oxidation with H2O2 alone synthetically non- coenzyme (NAD), have been applied only very recently in
applicable. Reaction rates were significantly enhanced by biocatalysis,49-51 although fundamental biochemical studies
combining H2O2 with additional organic reagents such as date back since early 1980s.52-54
.
stoichiometric trifluoroacetic anhydride (i.e. generating 2- Although marginally investigated in biocatalysis so far, aldehyde
hydroperoxyhexafluoro-2-propanol in situ).29 Unfortunately, dehydrogenases might become the first choice for oxidation of
even these types of method have disadvantages from a “green” aldehydes to carboxylic acids, if coupled with an efficient
chemistry perspective because of the requirement for system for the recycling of NAD+, because of their exquisite
halogenated solvents, the huge excess of carbonate salt and, chemoselectivity. Herein, we present a study on the
additionally,
a complicated reaction procedure. Similar optimisation of the chemoselective biocatalytic oxidation of
drawbacks apply with other related methods in which H2O2 is aldehydes to carboxylic acids catalysed by three recombinant
used in combination with excess of either urea and formic aldehyde dehydrogenases (ALDHs) originated from bovine lens
52
49, 53
acid,30 or formic acid alone,31 or monoperoxyphthalate and (ALDH-Bov),49,
Escherichia coli (ALDH-Ec)
and
urea. Variations using H2O2 with a catalytic amount of an Pseudomonas putida (PP-ALDH).50 For regeneration of the
organic molecule are also known. Benzeneseleninic acid33 or N- catalytic NAD+ coenzyme, the H2O forming NAD-oxidase from
methylpyrrolidin-2-one hydrotribromide (10 mol%),34 or Streptococcus mutans (NOx)55 was applied. The substrate scope
quaternary acidic salt (in aqueous/organic biphasic of the three ALDHs was elucidated with a wide range of
conditions)35 can be used as catalysts with H2O2 for aldehyde structurally diverse substrates.
oxidation. However, none of the systems based on H2O2 and
32
organic catalyst is of general applicability. For instance, the
H2O2/quaternary acid salt method developed by Noyori and
coworkers is efficient with straight-chain aliphatic aldehydes
and benzaldehydes, but chemoselectivity is mediocre with α-
branched aldehydes.35 Further research to develop a more
general metal-free method for the chemoselective oxidation of
aldehydes to carboxylic acids led others to consider the use of
alternative peroxides such as peroxymonosulphate (Oxone)36 or
even N-heterocyclic carbenes.37 On one hand, the oxidation
using Oxone requires toxic and high boiling DMF as solvent,38
Scheme 1 Oxidation of aldehydes catalysed by aldehyde dehydrogenases (ALDHs) using
complicating the isolation of low-boiling aliphatic products.
Furthermore, the Oxone oxidation method failed with para-
methoxy and hydroxy substituted aromatic aldehydes because
the related Dakin-type oxidation products (i.e. phenol esters)
were obtained. On the other hand, the N-heterocyclic carbene-
catalysed oxidation represents a valuable option for the
synthesis of high value α-chiral, α-halo carboxylic acids, but the
high loading of the sophisticated catalysts (10-20 mol%) makes
it economically unprofitable and unsustainable for a more
general oxidation of achiral aldehydes.
nicotinamide oxidase (NOx) for coenzyme recycling. The biocatalytic oxidation requires
dioxygen from air as ultimate oxidant. No side-product or waste is produced.
Results and discussion
Optimisation of the reaction conditions. We started our study
on the three ALDHs in highly purified form (Bov and Ec 5 µM; PP
20 µM) by determining the optimum temperature for the
biocatalytic oxidation of two model substrates (20 mM): a linear
aldehyde (1a, 2-methyl pentanal) and an aromatic aldehyde (2a
,
Biocatalytic methods for the oxidation of functional groups are
attracting interest because of the elevated (and often perfect)
chemo- and regio-selectivity, the mild reaction conditions
(ambient temperature and atmospheric pressure) in aqueous
environment and the possibility to use efficiently molecular
oxygen as benign oxidant.39 Interestingly, biocatalytic oxidation
of aldehydes to carboxylic acids in organic synthesis has mainly
been conducted using flavin or copper-dependent oxidases (EC
1.2.3), and with an emphasis for converting bio-derived
feedstock.40-44 Although aldehyde oxidases require only
dioxygen to accomplish the oxidation, the reaction is not fully
selective as also alcohol moieties are generally oxidised. Thus,
aldehyde oxidases (and even some alcohol oxidases) have been
benzaldehyde) in Tris/HCl buffer (50 mM, pH 9). The type of
buffer and pH initially applied was known to be suitable for the
biocatalytic reaction from our previous study.49 Nonetheless,
the influence of buffer has been assessed thoroughly in this
work. For regeneration of the NAD+ coenzyme (0.5 mM), the
H2O forming NAD-oxidase from Streptococcus mutans (NOx)
was applied (5 µM).55 DTT (100 µg mL-1) was also added in the
reaction mixture in order to prevent any possible oxidation of
the catalytic cysteine residue of the ALDHs. Indeed, addition of
DTT was mandatory for obtaining reproducible results using
different batches of the same enzyme (for details, see SI section
4). Figure 1 summarises the results for the temperature study
(for details see SI, Tables S3 and S4).
applied for the two-step oxidation of alcohols to carboxylic
ALDH-Bov converted >90% of 1a to 1b after 2 h reaction time at
all three temperatures tested (Figure 1 A). The initial activity,
however, was higher at 40 °C. The higher reaction rate at
42, 44
acids.41,
In contrast, synthetic applications of
dehydrogenases during the past two decades mainly involved
the reversible oxidation/reduction of alcohols to carbonyl
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temperatures moderately above room temperature was also time. Thus, it appeared that the choice of the temperature for
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DOI: 10.1039/C8GC01381K
observed for the oxidation of 2a. In fact, the analytical yield of performing the biocatalytic oxidation should be a compromise
2a into 2b already exceeded 90% after 90 min at 40 °C and 50 between activity and stability of the tested ALDHs. This
°C. In contrast, only 33% analytical yield into 2b was obtained observation is in agreement with our thermostability data (see
after 90 min at 30 °C, and 4 h were required to reach a maximum next paragraph) and the data reported for ALDH-Ec in
of 88% (Figure 1 D). ALDH-Ec also operated well at 30 °C and 40 literature.53 PP-ALDH showed similar activity for the oxidation
°C, having a slight better catalytic performance at 40 °C (Figure of 1a at 40 and 50 °C, with a slight preference for 50 °C (Figure
1 B and E). In this case, 1b and 2b were obtained with >90% 1 C, >90% analytical yield after 180 min). At 30 °C, only 48%
analytical yield in less than 90 min. Increasing the temperature analytical yield into 1b was obtained. An interesting first
to 50 °C, ALDH-Ec produced more than 90% of 1b only just after indication of the different substrate acceptance of the ALDHs
90 min, whereas no more than 68% analytical yield of 2b could was revealed already at this stage, since PP-ALDH prefers to
be obtained under the same reaction conditions. Observing in oxidise aliphatic aldehydes over aromatic ones. Indeed, only a
Figure E the progress of the analytical yield of 2a over time maximum of 10-13% of 2b was formed during the bio-oxidation,
revealed that 64% of 2b was obtained already within 15 min, independent of the temperature applied (Figure 1 F).
but the reactions did not proceed significantly further after that
Figure 1 Temperature profile for the conversion of 1a (panel A to C) and 2a (panel D to F) by ALDH-Bov, ALDH-Ec and PP-ALDH. Experimental conditions: 1 mL final volume
in Eppendorf tubes, buffer = 50 mM Tris/HCl pH 9, 180 rpm on orbital shaker, [1a] or [2a] = 20 mM, [NAD+] = 0.5 mM, [ALDH-Bov] = 5 µM for 1a and 10 µM for 2a, [ALDH-
Ec] = 5 µM, [PP-ALDH] = 20 µM, [NOx] = 5 µM, [DTT] = 100 µg/mL. Extraction with EtOAc under acidic conditions using toluene as IS and derivatisation of acids to the
methyl esters with (trimethylsilyl)diazomethane. Conversions were measured by GC-FID (DB-1701).
In a second step, we investigated the influence of the pH and yield values at 40 °C were compared for the reactions using
type of buffer used for the oxidation of 1a and 2a. Analytical Tris/HCl buffer at pH 7, 8 and 9, KPi buffer pH 8 and Gly/NaOH
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buffer at pH 9 (all buffers at 50 mM concentration) and the essential cysteine residue in the active site of the ALDHs (pKa
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results are summarized in figure 2 and the SI (tables S5 and S6). 8.6),52 as well as the required nucleophilicity of the attacking
DOI: 10.1039/C8GC01381K
In general, all three enzymes are poorly active at pH 7, leading water molecule in the last step of the mechanism.54 In fact, the
to maximum analytical yields of 14-40% for 1a and 15-30% for thiol group of the cysteine residue must be deprotonated in
2a after 4h. The origin for the low activity at neutral pH might order to perform the first nucleophilic attack on the aldehyde
have been related to the protonation state of the catalytic substrate.
Figure 2 pH and buffer profile for the conversion of 1a (panel A to C) and 2a (panel D to F) by ALDH-Bov, ALDH-Ec and PP-ALDH. Experimental conditions: 1 mL final volume
in Eppendorf tubes, buffer strength = 50 mM (Gly/NaOH buffer = 50 mM glycine titrated with NaOH to pH 9), T = 40 °C, 180 rpm on orbital shaker, [1a] or [2a] = 20 mM,
[NAD+] = 0.5 mM, [ALDH-Bov] = 5 µM for 1a and 10 µM for 2a, [ALDH-Ec] = 5 µM, [PP-ALDH] = 20 µM, [NOx] = 5 µM, [DTT] = 100 µg/mL. Extraction with EtOAc under acidic
conditions using toluene as IS and derivatization of acids to the methylester with (trimethylsilyl)diazomethane. Conversions were measured by GC-FID (DB-1701).
Then, after the hydride shift from the first thiohemiacetal (>90% analytical yield after 60 min for 1a and 2a; Figure 2 A and
intermediate to the NAD+, a polarized water molecule is D). Moreover, even at pH 9, the enzyme was found to perform
necessary to hydrolyse the second thioester intermediate. As better in Tris/HCl (>90% after 60 and 90 min for 1a and 2a) than
both steps are enabled by Brønsted base catalysis, a basic pH in Gly/NaOH (70% and 60% for 1a and 2a after 60 and 90 min,
buffer should accelerate the reaction.
respectively).
Previous biochemical studies indicated that ALDH-Bov is ALDH-Ec was described as a sensitive enzyme towards pH
inhibited in Tris/HCl buffer.52 However, the use of Tris/HCl changes, reaching a maximum of activity at pH 8. Increasing or
buffer did not influence the outcome of our biocatalytic decreasing the pH by one unit led to 70% loss of activity.53
reactions since the same increase of the analytical yield over Indeed, using Gly/NaOH buffer at pH 9 (as reported in
53
time was observed using both Tris/HCl and KPi buffer at pH 8 literature), the analytical yields did not exceed 40-45% for
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both substrates tested. However, when we applied Tris/HCl established formate dehydrogenase (FDH)/formVaiteew ArrteiccleyOclnilninge
DOI: 10.1039/C8GC01381K
buffer at the same pH, the analytical yield increased over 90% system. Formation of aldehydes 1a and 2a was not detected at
after 60 and 30 min for the oxidation of 1a and 2a, respectively. all.
The results for the oxidations of 1a and 2a in Tris/HCl were
comparable with the results in KPi buffer at pH 8 (Figure 2 B and Thermostability study on the ALDHs and NOx
E).
The thermostability (melting temperature Tm) of the three
The preferred buffer conditions for the aldehyde oxidation ALDHs as well as NOx was determined by differential scanning
catalysed by PP-ALDH were Tris/HCl and KPi at pH 8 (>90% after fluorometry. Tm values were determined in Tris/HCl buffers (50
120 min for 1a and 2a, Figure 1 C and F). Whilst similar analytical mM, pH 8, 8.5 and 9), KPi buffers (50 mM, pH 8 and 8.5) and
yields were obtained for 1a at pH 9 (>90% after 180 min), only Gly/NaOH buffer (50 mM, pH 9). Additionally, we measured the
10% analytical yield was measured for 2a. In the case of PP- Tm values of the three ALDHs in presence as well as absence of
ALDH, we investigated also the oxidation at pH 8.5 in Tris/HCl as NAD+ and DTT, since stabilising effect of coenzyme is expected.
well as KPi buffer, which demonstrated the preference for KPi The Tm values of ALDH-Bov, ALDH-EC and PP-ALDH in KPi (50
buffer for the oxidation of substrate 2a
.
mM, pH 8.5) were 47 °C, 44 °C and 42 °C (for details, see SI
For the continuation of our study, we finally decided to use KPi section 14). The Tm value of NOx was 51 °C. These data are in
buffer at pH 8.5. In fact, tris(hydroxymethyl)aminomethane (i.e. agreement with the temperature profile of the biocatalytic
Tris buffer) contains a primary amine moiety that is amenable reactions reported in Fig. 1 and Fig. 2.
for condensation with some aldehydes in aqueous solution as
also observed by other authors.56, 57 Consequently, the possible Elucidation of the substrate scope. A panel of sixty-one
interaction between Tris and some aldehyde substrates might structurally diverse substrates was tested with the three ALDHs
influence negatively the efficiency of the biocatalytic oxidation. under the optimised reaction conditions. The substrates
On the other hand, the buffer capacity of KPi buffer is maximum investigated can be divided into four main structural groups: (1)
in the range of pH between 6.5 to 8. Thus, KPi buffer is not aliphatic (linear and cyclic) aldehydes, (2) aryl-alkyl aldehydes
commonly applied above pH 8. However, in our specific cases, (e.g. cinnamic/hydrocinnamic motif), (3) aromatic aldehydes
KPi at pH 8.5 is applicable, because the oxidation of the (e.g. substituted benzaldehydes) and (4) heteroaromatic
aldehyde generates only the carboxylic acid product. Thus, only aldehydes (i.e. mono- and bicyclic). Moreover, we assessed the
buffering towards the production of Brønsted acid species is potential chemospecificity of the ALDHs for the oxidation of the
required. In fact, under our optimised reaction conditions, no aldehyde moiety to the carboxylic moiety by testing additional
significant variation of pH was observed between the start and substrates that contain further oxidisable groups such as
the end of the reaction. The temperature was instead set to 40 alcohols. Racemic phenyl-2-propanal was also tested. The
°C for the continuation of the study, to avoid any possible partial progress of the reaction was monitored after 4 and 24 h. In
evaporation of volatile aldehydes and to retain enzyme stability. general, we did not observe any significant differences in the
An interesting aspect is the possible enantiodiscrimination of analytical yield after 4 and 24 h reaction time. Either the
the three ALDHs towards racemic substrate 1a, which contains reaction proceeded to high analytical yield within 4 h or did not
a stereogenic centre in α-position to the aldehyde moiety. Our proceed to a significant extent at all. The results of the substrate
data clearly shows that all three ALDHs can efficiently oxidise screening are summarized in Table 1 to 4 (for the full dataset
both (S)-1a and (R)-1a, although the oxidation of (R)-1a is including the blank measurements see SI, Tables S9 and S10).
slightly kinetically favoured. However, when the oxidation Comparing the results of the biocatalytic reactions (i.e. in
reaches completion (for details, see SI section 9), product 1b is presence of ALDH) with the blank experiments (i.e. same
obtained in racemic form. Thus, significant spontaneous conditions but without addition of enzyme), we could detect a
chemical racemisation of substrate 1a does not occur within the minor spontaneous aerobic oxidation of some aldehydes to the
reaction time (1 h).
related carboxylic acids under aerobic mixing in aqueous buffer
Finally, the irreversibility of the biocatalytic oxidation catalysed at pH 8.5. This observation is in agreement with a recent
by the ALDHs under our reaction conditions was demonstrated publication by Zhang et al., in which the spontaneous oxidation
by running the reverse reaction using carboxylic acids 1b and 2b of some aldehydes at 5-10 mM concentration was reported in
(20 mM) as substrates (see SI section 13). The reduced aqueous environment at 1 bar of pure dioxygen pressure and
nicotinamide coenzyme (1 mM) was recycled via the 37 °C without any additional catalyst.58
Table 1 Biocatalytic aerobic oxidation of aliphatic/cyclic aldehydes to carboxylic acids catalysed by ALDHs.
substrate
ALDH
Bov
Ec
yield [%]
67[a]
substrate
ALDH
Bov
Ec
yield [%]
69[a]
substrate
ALDH
Bov
Ec
yield [%]
70[a]
O
O
O
76[a]
92
62[a]
3a
PP
Bov
Ec
72[a]
>99
>99
6a
PP
Bov
Ec
64[a]
76[a]
76[a]
10a[b]
PP
Bov
Ec
61[a]
75[a]
69[a]
O
O
O
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4a
5a
PP
Bov
Ec
>99
>99
7a[b]
8a
PP
Bov
Ec
70[a]
10[a]
92[a]
94[a]
60[a]
11a
12a
PP
67[a]
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O
O
O
O
DOI: 10.1039/C8GC01381K
Bov
Ec
77
78
>99
PP
>99
>99[a]
PP
PP
80
O
Bov
Bov
Bov
>99
O
Ec
>99[a]
Ec
54[a]
55[a]
Ec
>99
>99
1a
PP
>99[a]
9a[b]
PP
13a[b]
PP
If not specified, reaction time was 4 h; [a] reaction time was 24 h.
[b] High analytical yield was observed without the addition of enzyme. Conversion of blank reactions (24 h, average of two samples): 7a, 20%; 9a, 15%; 13a, 10%. For full
data set, see SI section 8.
Experimental conditions: 1 mL Eppendorf tubes in KPi buffer (50 mM, pH 8.5), T = 40 °C, 180 rpm on orbital shaker, [substrate] = 20 mM, [ALDH] = 10 µM, [NOx] = 5 µM,
[DTT] = 100 µg/mL, [NAD+] = 0.5 mM. Extraction with EtOAc under acidic conditions using toluene as IS and derivatization of acids to the methylester with
(trimethylsilyl)diazomethane. Analytical yields were measured by GC-FID (DB-1701).
Among the set of 61 substrates investigated (20 mM), to ten carbon atoms (6-12a), the ALDHs could oxidise the
spontaneous oxidation afforded analytical yields in the range of substrates with analytical yields ranging from 54% to 94%. A
1-5% in 21 cases, and in the range of 6-10% in 7 cases. general trend was not revealed as the best performing ALDH
Spontaneous oxidation slightly above 10% was detected for depended on the particular substrate. The interesting
substrates 2a, 7a, 9a and 21a. Aliphatic and aryl aliphatic exceptions were the bio-oxidation of 6a and 8a, which both
aldehydes (see SI Table S9 for data and Table 1, 2 for structures) contain a carbon-carbon double bond conjugated to the
were the substrates to be affected generally by spontaneous aldehyde moiety. 6a is preferentially converted by ALDH-Ec
oxidation, with the exception of 16a. Instead, the occurrence of (92% analytical yield), whereas ALDH-Bov and PP-ALDH
spontaneous oxidation with benzaldehyde derivatives afforded 69% and 64% analytical yield, respectively. 8a was also
depended on the type of substituent on the phenyl ring (see SI well accepted by ALDH-Ec and PP-ALDH (92 and 94% analytical
Table S9 and S10 for data and Table 3 for structures). In yield, respectively), but not by ALDH-Bov (10% analytical yield).
contrast, other homoaromatic (bicyclic) and heteroaromatic This feature was also observed in our previous work, in which
aldehydes did not show any spontaneous oxidation (see SI Table we exploited the higher activity of ALDH-Bov towards the
S10 for data and Table 4 for structures).
saturated aldehyde rather than the unsaturated one in a dual-
Table 1 summarizes the results for the oxidation of enzyme cascade process.49 The same effect is also evident in the
aliphatic/cyclic aldehydes. The shortest chain aldehyde tested case of other structurally related substrates (Table 2): the α,β-
was butanal (3a), whose biocatalytic oxidation resulted in the unsaturated aldehyde cinnamaldehyde (15a) and its saturated
formation of 67-76% analytical yield of acid 3b after 24 h. counterpart hydrocinnamaldehyde (14a) (Table 2). In fact, the
However, the actual productivity of the system might be analytical yield for the saturated substrate 14a (83, 85, 79% for
improved as a fraction of volatile aldehyde 3a may have Bov, Ec, and PP) was always higher than the analytical yield of
evaporated during the course of the reaction with the current the unsaturated one 15a (6, 25, 5% for Bov, Ec, and PP) with all
experimental set-up. Indeed, homologous aldehydes but less three ALDHs.59 This discrimination between saturated and
volatile such as 4a, 5a and 1a (5 and 6 carbon-atom chains) gave unsaturated aldehydes did not occur when testing substrates
quantitative analytical yield with all three ALDHs tested. The possessing the unsaturation moiety in further ω-positions. In
same applied for cyclohexancarbaldehyde 13a. The capability of this case, respectively saturated and unsaturated substrates 4a
converting short chain aliphatic aldehydes by ALDHs is of and 5a were converted equally (>99%). The substituted
interest. In fact, a number of catalytic organic and metal-organic cinnamaldehyde 16a, which bears an additional methoxy group
reagents are not applicable for the oxidation of these substrates in meta and a hydroxyl group in para position was converted to
because of the need for medium-high boiling solvents for the low extend (4-5% max with ALDH-Bov and Ec). It might be that,
reaction, which complicates product isolation, or lack of activity. in this case, steric hindrances become already a limiting factor.
12, 19, 36 In the case of medium and long chain aldehydes from six
Table 2 Biocatalytic aerobic oxidation of aromatic aldehydes (cinnamic/hydrocinnamic motif) to carboxylic acids catalysed by ALDHs. The reaction time was 24 h.
substrate
ALDH
Bov
yield [%]
83
substrate
ALDH
Bov
yield [%]
6
substrate
ALDH
Bov
yield [%]
4
O
O
O
Ec
85
79
Ec
25
Ec
5
HO
O
14a
PP
15a
PP
5[a]
16a
PP
<1
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[a] The blank measurement without the addition of enzyme gives the same analytical yield.
View Article Online
DOI: 10.1039/C8GC01381K
Experimental conditions: 1 mL Eppendorf tubes in KPi buffer (50 mM, pH 8.5), T = 40 °C, 180 rpm on orbital shaker, [substrate] = 20 mM, [ALDH] = 10 µM, [NOx] = 5 µM,
[DTT] = 100 µg/mL, [NAD+] = 0.5 mM. Extraction with EtOAc under acidic conditions using toluene as IS and derivatization of acids to the methylester with
(trimethylsilyl)diazomethane. Analytical yields were measured by GC-FID (DB-1701).
Furthermore, a wide panel of 32 benzaldehyde derivatives was ortho- substituted benzaldehyde derivatives, as the bulkier
tested (Table 3). Chemoselective oxidations to carboxylic acids methoxy (37a) and acetyl (46a) groups in the ortho position
starting from benzaldehyde derivatives bearing electro- gave only 10% and 2% analytical yield for ALDH-Bov,
donating substituents such as hydroxy and methoxy, are respectively. Di- or tri-substituted derivatives with at least one
problematic by chemocatalytic methods because of the substituent in the ortho position, like 20a, 28a and 40a, were
frequent occurrence of competitive Dakin-type oxidation to not accepted by any of the ALDHs used in this study. Table 4
phenols (and their oligomerisation). 12, 21, 36 On the contrary, we demonstrates the broad applicability of the ALDHs since all the
found that all the biocatalytic oxidations of benzaldehyde monocyclic six-membered and five-membered ring
derivatives catalysed by ALDHs were perfectly chemoselective heteroaromatic compounds tested were converted
as the expected carboxylic acids were the only products. As quantitatively by at least one of the enzymes (48-54a).
suggested above from the pH and temperature studies, PP- Interestingly, even some bicyclic heteroaromatic compounds,
ALDH preferentially oxidises aliphatic aldehydes rather than such as 56a and 57a, were converted (51-67%), whereas 55a
aromatic aldehydes. Among the benzaldehyde derivatives and 59a were converted to a lesser extent (11 and 3%). It is
tested, quantitative analytical yield was only observed for the important to note that the oxidation of these substrates
oxidation of benzaldehyde 2a. For all other benzaldehyde proceeded with perfect chemoselectivity because sulphur
derivatives bearing a substituent in meta or para position, there heteroatoms (52-54a
was at least one ALDH enzyme that was capable of catalysing 59a), or additional hydroxyl groups (51a) were not oxidised at
the reaction above 90% analytical yield (18-19a 22-23a 26-27a all by the ALDHs. However, it is known that all these functional
30-31a 32-33a 35-36a 38-39a 47a). Moreover, the oxidation groups can become oxidised when chemical catalysts and/or
, 56a), nitrogen heteroatoms (48-49a, 57a,
,
,
,
,
,
,
,
of meta and para di-substituted aromatic aldehydes worked reagents are used for oxidation of aldehydes.2, 3 Furthermore,
well with analytical yields from 91-99% for 41-44a and 57% for the perfect chemoselectivity for the biocatalytic oxidation of
45a. In contrast, the oxidation of aldehydes possessing furan-2-carbaldehydes
(50-51a
)
and
benzofuran-2-
substituents in ortho position were generally more challenging. carbaldehyde (55a) is also noteworthy as formation of Dakin-
It seems that electron donating substituents in the ortho type by-products is known using, among others, peroxides as
position still favour the reaction (51% for 17a and >99% 34a chemical oxidising agent.36 In addition, ALDH-Bov converted
with ALDH-Bov), whereas analytical yields were moderate or bulkier bicyclic homoaromatic aldehydes such as 58a and 60a
poor with electron withdrawing groups in the ortho position with elevated analytical yields (>99% and >90% respectively)
(11% for 25a, 15% for 29a). The exceptions are the fluoro- and again perfect chemoselectivity. In contrast, strong chemical
substituted aromatic benzaldehyde derivatives 21a and 24a
,
oxidising agents such as pyridinium chlorochromate/periodate
which were fully converted by ALDH-Bov and ALDH-Ec. It might fail in the oxidation of naphtaldehydes such as 58a and 60a
.
also be that steric effects can play a role in the bio-oxidation of
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Table 3 Biocatalytic aerobic oxidation of aromatic aldehydes (substituted benzaldehydes) to carboxylic acids catalysed by ALDHs.
View Article Online
DOI: 10.1039/C8GC01381K
substrate
ALDH
Bov
yield [%]
>99
substrate
ALDH
Bov
yield [%]
>99
substrate
ALDH
yield [%]
O
O
O
Bov
Ec
>99
Ec
>99
Ec
>99
49[a]
O
Cl
2a[c]
PP
>99[a]
51
27a
Cl
PP
7[a]
38a
PP
n.d.
>99
O
O
O
Bov
Bov
n.d.
Bov
Ec
28[a]
Ec
n.d.
Ec
12[a]
Cl
O
17a
PP
1[a] [b]
>99
28a
PP
n.d.
15[a]
39a
PP
<1[a] [b]
2[a
O
O
O
Bov
Bov
Bov
Ec
25
Ec
8
Ec
n.d.
O
O
Br
18a
PP
2[a] [b]
>99
29a
PP
7[b]
40a
PP
n.d.
95[a]
O
O
Bov
Bov
97[a]
Bov
O
1[a]
O
Ec
>99
Ec
>99
Ec
O
Br
19a
PP
3[a] [b]
2[b]
30a
PP
14
41a
PP
n.d.
91
O
Bov
Bov
>99
Bov
O
O
O
Ec
2[b]
Ec
>99[a]
Ec
8[a]
Br
O
20a
PP
1[b]
31a
PP
8
42a
PP
4[a] [b]
>99[a]
O
O
O
Bov
>99
Bov
94
Bov
16[a]
O
Ec
>99
Ec
93
Ec
CN
F
Cl
21a[c]
PP
16
32a
PP
92
98
43a
PP
3[a]
O
O
O
Bov
>99
Bov
Bov
>99[a]
HO
Ec
>99
Ec
96
Ec
3[a]
F
NC
Cl
22a
PP
13
33a
34a
PP
96[a]
>99
44a
PP
2[a] [b]
57[a]
O
O
O
Bov
>99
Bov
Bov
2[a]
Ec
<1[a]
HO
Ec
>99
Ec
F
OH
O
O
23a
PP
17
PP
<1[b]
>99
45a
PP
n.d.
O
Bov
>99
Bov
Bov
2[a] [b]
F
O
F
Ec
>99
Ec
20
Ec
2[a] [b]
OH
O
24a
25a
PP
3[a] [b]
11[a]
35a
PP
1[a] [b]
96[a]
46a
47a
PP
2[a] [b]
>99[a]
O
O
O
Bov
Bov
Bov
Ec
11[a]
Ec
6[a]
Ec
>99
18
F3C
HO
Cl
PP
5[a] [b]
>99[a]
36a
PP
2[a]
10
PP
O
Experimental: 1 mL in KPi buffer (50 mM, pH
8.5), T = 40 °C, 180 rpm, [substrate] = 20 mM,
[ALDH] = 10 µM, [NOx] = 5 µM, [DTT] = 100
µg/mL, [NAD+] = 0.5 mM. Analytical yields were
measured by GC-FID as methyl ester.
O
Bov
Bov
Ec
>99
Ec
2[a]
O
Cl
26a
PP
20
37a
PP
1[a] [b]
If not specified, reaction time was 4 h; [a] reaction time was 24 h.
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[b] The blank measurement without the addition of enzyme gives the same analytical yield; [c] High analytical yield observed without the addition of enzyme. Conversion
View Article Online
DOI: 10.1039/C8GC01381K
of blank reactions (24 h, average of two samples): 21a, 12%; 2a, 13%. For full data set, see SI section 8.
Table 4 Biocatalytic aerobic oxidation of mono- and bicyclic heteroaromatic aldehydes, bicyclic aromatic aldehydes and profen aldehydes to carboxylic acids catalysed by ALDHs.
substrate
ALDH
Bov
yield [%]
40[a]
substrate
ALDH
Bov
yield [%]
99
substrate
ALDH
Bov
yield [%]
>99[a]
O
O
S
O
Ec
>99
Ec
>99
Ec
17[a]
Br
Br
N
48a
PP
13[a]
>99
53a
PP
4
58a
59a
PP
n.m.
3[a]
Bov
Bov
>99
Bov
O
O
S
Cl
n.m.[a]
N
N
Ec
>99
Ec
>99
Ec
H
O
49a
PP
4
54a
PP
3
PP
n.m.[a]
94[a]
O
Bov
>99
Bov
11
Bov
O
O
O
Ec
>99
Ec
8[a]
Ec
3
O
50a
PP
>99
90
55a
PP
1[a]
60a
61a
PP
2[a] [b]
25[c]
Bov
Bov
51[a]
Bov
O
O
HO
O
Ec
91[a]
Ec
8[a]
Ec
27[c]
25[c]
S
O
51a
PP
Bov
Ec
25]
>99
>99
6
56a
PP
Bov
Ec
n.m.[a]
67
PP
O
O
S
N
<1
52a
PP
57a
PP
n.m.
If not specified, reaction time was 4 h, [a] reaction time was 24 h
[b] The blank measurement without the addition of substrate gives the same analytical yield
[c] For ALDH-Bov and ALDH-Ec, racemic product was obtained. PP-ALDH afforded the product in slightly enantioenriched form (ee 16%, (R)).
Experimental conditions: 1 mL Eppendorf tubes in KPi buffer (50 mM, pH 8.5), T = 40 °C, 180 rpm on orbital shaker, [substrate] = 20 mM, [ALDH] = 10 µM, [NOx] = 5 µM,
[DTT] = 100 µg/mL, [NAD+] = 0.5 mM. Extraction with EtOAc under acidic conditions using toluene as IS and derivatization of acids to the methylester with
(trimethylsilyl)diazomethane. Analytical yields were measured by GC-FID (DB-1701).
Finally, racemic 2-phenylpropanal (61a) was tested for the on the oxidation of two heterocyclic compounds with
oxidation with the three ALDHs, affording the carboxylic acid lyophilised whole cells containing overexpressed ALDH (E. coli
product 61b in 25-27% analytical yield and in racemic (ALDH- cells devoid of expressed ALDH were also tested as negative
Bov, ALDH-Ec) or slightly enantioenriched form (PP-ALDH, ee control
reactions).
We
chose
5-hydroxymethyl-2-
16% R). furancarbaldehyde (51a) as substrate because it is a starting
Overall, these results highlight the perfect chemoselectivity of material derivable from renewable fructose. Furthermore, the
the oxidation catalysed by aldehyde dehydrogenases.18 This related oxidation product 5-hydroxymethyl-2-furancarboxylic
level of chemospecificity in the oxidation of the aldehyde
acid (HMFCA, 51b) is a versatile building block for the synthesis
moiety is unrivalled by any chemical or other enzymatic method of polyesters as well as an interleukin inhibitor.60, 61 In addition,
(i.e. oxidase enzymes).
it was reported that HMFCA possesses antitumor activity. 40 As
second test substrate thiophene-2-carbaldehyde (54a) was
Biocatalytic oxidation using E. coli/ALDH whole cells. For chosen as its chemoselective oxidation is challenging, due to the
practical applications, the use of whole cells rather than purified fact that both, sulphur heteroatom and carbon-carbon double
enzymes is the preferred solution because time consuming bonds are prone to oxidation by conventional chemical
purification steps are omitted. At this stage, we decided to focus oxidants.62, 63 Additionally, 54b was quantitatively recovered
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after bio-oxidation catalysed by ALDH-Bov and ALDH-Ec on study was conducted following the sameViewsyArstticelemOanltinice
DOI: 10.1039/C8GC01381K
analytical scales and further extraction of the reaction mixture. methodology as for the biocatalytic oxidation with lyophilised
Before scaling-up the biocatalytic oxidation, we carried out a cells. In general, we observed a correlation between the yields
preliminary set of experiments at analytical scale. The bio- for the oxidation of substrate 51a using either resting cells
oxidation (substrate 20 mM, reaction volume 1 mL) was (Table S16) or lyophilised cells (Table S14). In contrast, few
performed using lyophilised E. coli overexpressing the ALDHs minor discrepancies were observed for the oxidation of
(10 mg mL-1) in presence and absence of NOx (0-5 µM) and NAD+ substrate 54a (Table S17 and Table S15, respectively). The major
(0-0.5 mM) in order to assess the influence of the exogenous discrepancy was revealed for the oxidation of 54a catalysed by
recycling system and NAD coenzyme in a whole cell ALDH-Ec in absence of NOX and NAD+. Resting cells afforded
environment (SI, tables S12 and S13).
higher yields (89-90%) than lyophilised cells (26-38%). This
According to the results for the biocatalytic oxidation of 51a in difference may be attributed to the fact that a significant level
analytical scale using purified ALDHs (Table 4), the substrate of endogenous NAD+ is available to sustain the reaction in
must be converted to the product 51b with a maximum of 91% resting cells, whereas it is mainly degraded during lyophilisation
analytical yield. In that case, no starting material was detected procedures.
after the reaction with purified ALDHs. Using lyophilised E. coli
cells overexpressing the ALDHs, the aldehyde substrate 51a was Preparative scale biocatalytic oxidation. A preparative scale
converted again to a similar level (ca. 80% analytical yield, see bio-oxidation of 51a (>500 mg) using lyophilised whole cells was
SI Table S12). Surprisingly, the same conversion of 80% from 51a conducted without the addition of NAD+ and recycling enzyme
to 51b was also obtained using lyophilised E. coli cells/ALDH but NOx, resulting in 74% and 75% analytical yield for ALDH-Bov and
without the addition of NAD+ and NOx. For substrate 51a, also ALDH-Ec, respectively. After extraction and solvent
a considerable level of oxidation to the carboxylic acid (36%) evaporation, the isolated yields of 51b were 63% and 56% (i.e.
was observed when E. coli whole cells devoid of any 356 and 315 mg). We further increased the scale of the reaction
overexpressed ALDHs were tested.
up to 2 g of substrate using ALDH-Ec lyophilised whole cells; that
Conversely, the bio-oxidation of 54a with lyophilized cells resulted in 71% analytical yield and 61% isolated yield of 51b
overexpressing ALDH-Ec required the mandatory addition of (1.37 g). Thus, applicability of the ALDHs in chemical synthesis
either NAD+ or NOx in order to reach elevated analytical yields. and potentially in large scale can be demonstrated.
In fact, a drop in the analytical yield of 54a was observed when
both NOx and NAD+ were excluded from the reaction (26-38%).
In any case, the analytical yield was slightly lower using ALDH-
Conclusions
Ec as lyophilised whole cells biocatalyst as the maximum
analytical yield with addition of either NAD+ or NOx was 89%
(see SI, Table S13) In comparison, the same oxidation using the
purified ALDH-EC or ALDH-Bov gave >99% analytical yield (Table
4). The reason for that might be the probable interaction
between the reactive aldehyde moiety of the substrate with
abundant nucleophilic groups of the whole cell environment,
hence reducing free substrate availability in solution.
Interestingly, the bio-oxidation of 54a using ALDH-Bov required
mandatory addition of both NOX and NAD+ to achieve elevated
analytical yield (87%). Without NAD+ or NOx the analytical yield
using ALDH-Bov fell to 42-46%. Absence of both, coenzyme and
recycling enzyme led to only 15% analytical yield. Thus, it seems
that the need for addition of coenzyme to lyophilized whole
cells systems depends from the particular overexpressed ALDH
and the substrate converted.
In the following step, lyophilized whole cells (ALDH-Bov and Ec)
were tested for the oxidation at increased substrate
concentrations of 51a (20-100 mM) either with (table S14) or
without (table S15) addition of NAD+ and NOx. Already at 40
mM substrate concentration, the analytical yield dropped down
from >80% to 35% (addition of NOx and NAD+) or 5-18%
(without coenzyme and recycling enzyme). Thus, 20 mM
substrate concentration was kept for the preparative scale
oxidation.
The oxidation of aldehydes to carboxylic acids is a relatively
simple organic chemical transformation, but a “green” and
general method for performing this reaction with perfect
chemoselectivity (>99%) and on a wide range of structurally
diverse substrates is needed. Some chemical methods offer
elevated efficiencies in terms of yields, selectivities and
substrate acceptance but these are accompanied by one or
more of the following drawbacks: i) a requirement for
increasingly unsustainable transition metal-catalysts; ii) the
need for organic oxidants in stoichiometric amounts (or even
large excess); iii) the need for strong acidic or basic conditions;
iv) the requirement for elevated temperatures or pressures.
Thus, there is still a need for new methods of aldehyde
oxidation which are both chemically efficient, chemoselective
and green.
In this work, we have performed the oxidation of aldehydes
using three recombinant aldehyde dehydrogenases that
originated from bacteria and bovine lens. The enzymes were
applied in highly purified form in combination with a
nicotinamide oxidase for the efficient recycling of NAD+ at the
expense of molecular oxygen from air. The biocatalytic
oxidation was optimised in terms of temperature, pH and type
of buffer. The final bio-oxidation runs in aqueous phosphate
buffer, under mild reaction conditions (40 °C and atmospheric
pressure) and consumes only dioxygen from air as benign
oxidant. Sixty-one structurally diverse aldehydes (aliphatic, aryl
aliphatic, benzylic-, hetero-aromatic and bicyclic aldehydes)
were tested. With the exception of only a few ortho-substituted
In the final set of experiments, we tested both substrates 51a
and 54a for the biocatalytic oxidation using resting E. coli cells
(i.e. wet cells) overexpressing either ALDH-Bov or ALDH-Ec. The
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benzaldehydes and two bicyclic heteroaromatic aldehydes, all Preparative biocatalytic oxidation of 51a
the other substrates were converted with elevated yield (up to
View Article Online
DOI: 10.1039/C8GC01381K
A total volume of 200 mL consisted of KPi buffer (50 mM, pH
8.5), lyophilised whole cells of E. coli overexpressing ALDH-Bov
or ALDH-Ec (2 g, 10 mg mL-1, resuspended in buffer), DTT (20
mg, final concentration 100 µg mL-1) and 51a (504 mg, 4 mmol
for ALDH-Bov and 502 mg, 3.98 mmol for ALDH-Ec; final
concentration 20 mM). Biotransformations were performed in
500 mL glass bottles, incubated at 40 °C in an orbital shaker.
After 21 h, a sample was taken and, after derivatisation, it was
analysed by GC. The reaction mixture was acidified to pH 2–4
via addition of HCl (3 M). NaCl was added (36 g) to enhance the
extraction of the acid. The aqueous reaction mixture was then
extracted with methyl tert-butyl ether (3 × 70 mL). Phase
separation was facilitated by centrifugation. The organic
fractions were combined and dried over MgSO4. After filtration
and evaporation of the solvent, the product was obtained in a
pure form. Column chromatography was not required. The
>99%). In all cases, the chemoselectivity was perfect: no other
product was detected except the expected carboxylic acid.
Oxidisable functionalities such as the hydroxyl moiety, alkene
groups, aryl groups, and sulphur as well as nitrogen
heteroatoms remained untouched. The bio-oxidation was also
investigated with lyophilised whole cell as well as resting E. coli
cell biocatalysts for a more convenient practical application.
Interestingly, supplementation of NAD+ and NOx recycling
enzyme can be omitted in some cases as the microbial host
produces sufficient amount of coenzyme, which can be recycled
by endogenous enzymes. The reaction was upscaled to the
oxidation
of
bio-based
starting
material
5-
(hydroxymethyl)furfural (2 g) to afford 5-(hydroxymethyl)furoic
acid in 61% isolated yield.
We conclude that aldehyde dehydrogenases have the potential
to become the first choice for chemoselective oxidation of
aldehydes into carboxylic groups. This biocatalytic method is
particularly attractive for the oxidation of aldehyde moieties
within molecules possessing further oxidisable groups. Future
research will focus on improving the enzymes tolerance to
substrate concentration and long-term stability in order to
enable even broader application of these enzymes.
1
authenticity of the product was confirmed by H-NMR (ESI,
paragraph 11). Upscaling of 2 g (15.86 mmol) 51a was
performed similar to the reaction described above using
lyophilized E.coli cells overexpressing AldDH-Ec. The reaction
was performed in 2 L Erlenmeyer flasks, covered with
aluminium foil.
Conflicts of interest
The authors declare to have no competing interests, or other
interests that might be perceived to influence the results
and/or discussion reported in this article.
Experimental
The expression and purification of the enzymes, influence of
DTT, temperature and pH studies, calculation of response
factors, the full dataset for the substrate scope using lyophilised
cells and resting cells, thermostability studies as well as the
analytical methods can be found in the ESI.
Acknowledgements
General procedure for the biocatalytic oxidation of aldehydes in
analytical scale
This project received funding from the European Research
Council (ERC) under the European Union’s Horizon 2020
research and innovation programme (grant agreement no.
638271, BioSusAmin). Dutch funding from the NWO Sector Plan
for Physics and Chemistry is also acknowledged.
Unless otherwise stated, standard reactions (1.0 mL, in
Eppendorf tubes) were performed in KPi buffer (50 mM
KH2PO4/K2HPO4, pH 8.5), containing ALDH purified (10 µM), NOx
purified (5 µM), NAD+ (0.5 mM) and DTT (100 µg/mL). The
reactions were started by the addition of substrate (1-61a, 20
mM final concentration), added pure or as DMSO stock
solutions (2 M, 1 M or 0.5 M, depending on their solubility). The
reactions were shaken at 40 °C at 170 rpm in an orbital shaker
for 4 and 24 h. Reactions were stopped by the extraction with
EtOAc (2 × 500 µL) under acidic conditions (addition of 100 µL
HCl 3N). The extracts were dried using anhydrous MgSO4 and
derivatised to the methyl ester as described below. Analytical
yields were measured by GC-FID.
Notes and references
1. G. Tojo and M. Fernández, Oxidation of Primary Alcohols to
Carboxylic Acids, chapter 7: Oxidation of Alcohols to Carboxylic
Acids via Isolated Aldehydes, Springer, New York, 2007.
2. M. Hudlicky, Oxidations in Organic Chemistry, American
Chemical Sociecty, Washington DC, 1990.
3. A. H. Hainess, Methods for the Oxidation of Organic
Compounds, Academic, New York, 1988.
4. I. Heilbron, E. R. H. Jones and F. Sondheimer, Journal of the
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Derivatisation of carboxylic acids to methyl esters
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TOC
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Aldehyde dehydrogenases enable the chemospecific oxidation of structurally
diverse aldehydes to carboxylic acids in aqueous environment and using only
air.
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