Terminal Alkenes from Acrylic Acid Derivatives via Non-Oxidative Enzymatic Decarboxylation by Ferulic Acid Decarboxylases

Article abstract of DOI:10.1002/cctc.201800643

Fungal ferulic acid decarboxylases (FDCs) belong to the UbiD-family of enzymes and catalyse the reversible (de)carboxylation of cinnamic acid derivatives through the use of a prenylated flavin cofactor. The latter is synthesised by the flavin prenyltransferase UbiX. Herein, we demonstrate the applicability of FDC/UbiX expressing cells for both isolated enzyme and whole-cell biocatalysis. FDCs exhibit high activity with total turnover numbers (TTN) of up to 55000 and turnover frequency (TOF) of up to 370 min^?1. Co-solvent compatibility studies revealed FDC's tolerance to some organic solvents up 20 % v/v. Using the in-vitro (de)carboxylase activity of holo-FDC as well as whole-cell biocatalysts, we performed a substrate profiling study of three FDCs, providing insights into structural determinants of activity. FDCs display broad substrate tolerance towards a wide range of acrylic acid derivatives bearing (hetero)cyclic or olefinic substituents at C3 affording conversions of up to >99 %. The synthetic utility of FDCs was demonstrated by a preparative-scale decarboxylation.

Full text of DOI:10.1002/cctc.201800643

Accepted Article
Title: Terminal Alkenes from Acrylic Acid Derivatives via Non-Oxidative
Enzymatic Decarboxylation by Ferulic Acid Decarboxylases
Authors: Godwin A. Aleku, Christoph Prause, Ruth T. Bradshaw-Allen,
Katharina Plasch, Silvia M. Glueck, Samuel S. Bailey, Karl A.
P. Payne, David A. Parker, Kurt Faber, and David Leys
This manuscript has been accepted after peer review and appears as an
Accepted Article online prior to editing, proofing, and formal publication
of the final Version of Record (VoR). This work is currently citable by
using the Digital Object Identifier (DOI) given below. The VoR will be
published online in Early View as soon as possible and may be different
to this Accepted Article as a result of editing. Readers should obtain
the VoR from the journal website shown below when it is published
to ensure accuracy of information. The authors are responsible for the
content of this Accepted Article.
To be cited as: ChemCatChem 10.1002/cctc.201800643
Link to VoR: http://dx.doi.org/10.1002/cctc.201800643
A Journal of
www.chemcatchem.org

ChemCatChem
10.1002/cctc.201800643
1
2
Terminal Alkenes from Acrylic Acid Derivatives via Non-Oxidative Enzymatic
Decarboxylation by FerulicAcid Decarboxylases
3
4
5
6
7
8
9
0
1
2
3
4
5
6
7
8
9
0
1
2
3
4
5
6
7
8
a
c
a
c
Dr. Godwin A. Aleku, Christoph Prause, Ruth T. Bradshaw-Allen, Katharina Plasch, Dr. Silvia
b,c
a
a
d
M. Glueck, Samuel S. Bailey, Dr. Karl A. P. Payne, Dr. David A. Parker, & Prof. Dr. Kurt
c
a
Faber, * & Prof. Dr. David Leys, *
a
Manchester Institute of Biotechnology, School of Chemistry, University of Manchester,
Manchester, UK. E-mail:David.Leys@manchester.ac.uk
b
1
1
1
1
1
1
1
1
1
1
2
2
2
2
2
2
2
2
2
Austrian Centreof IndustrialBiotechnology (ACIB), c/o
c
Department of Chemistry, University of Graz, Heinrichstrasse 28, 8010 Graz, Austria. E-mail:
Kurt.Faber@uni-graz.at
d
Innovation/Biodomain, Shell International Exploration and Production Inc., Westhollow
Technology Center, Houston, USA.
Abstract
Fungal ferulic acid decarboxylases (FDCs) belong to the UbiD-family of enzymes and catalyse
the reversible (de)carboxylation of cinnamic acid derivatives through the use of a prenylated
flavin cofactor. The latter is synthesised by the flavin prenyltransferase UbiX. Herein, we
demonstrate the applicability of FDC/UbiX expressing cells for both isolated enzyme and
whole-cell biocatalysis. FDCs exhibit high activity with total turnover numbers (TTN) of up to
-
1
55000 and turnover frequency (TOF) of up to 370 min . Co-solvent compatibility studies
revealed FDC’s tolerance to some organic solvents up 20% v/v. Using the in-vitro
(de)carboxylase activity of holo-FDC as well as whole-cell biocatalysts, we performed a
substrate profiling study of three FDCs, providing insights into structural determinants of
activity. FDCs display broad substrate tolerance towards a wide range of acrylic acid
derivatives bearing (hetero)cyclic or olefinic substituents at C3 affording conversions of up to
>99%. The syntheticutility of FDCs was demonstrated by a preparative-scaledecarboxylation.
2
3
3
3
3
9
0
1
2
3
Key words: Biocatalysis, Ferulic acid decarboxylase, Prenylated flavin, Decarboxylation;
Terminal alkenes
1
This article is protected by copyright. All rights reserved.

ChemCatChem
10.1002/cctc.201800643
3
4
Introduction
The production of organic building blocks from renewable carbon sources is a current trend
3
3
3
3
3
5
6
7
8
9
[
1–4]
in synthetic organic chemistry.
The major primary intermediates of traditional industrial-
scale synthesis are light alkenes such as ethylene, propylene and butadiene which are
produced from crude oil via steam-cracking, which has been described as the single most
[
5,6]
energy-demandingprocess in the petrochemicalindustry.
4
4
4
4
4
4
4
4
4
4
5
5
5
5
5
5
5
5
5
5
6
6
6
6
0
1
2
3
4
5
6
7
8
9
0
1
2
3
4
5
6
7
8
9
0
1
2
3
In view of the fact that biocatalytic transformations are operational under mild and
environmentally-friendly conditions and proceed with high chemo-, regio- and
[
7]
stereoselectivity, there is an increasing interest in expanding the scope and efficiency of
[
8–13]
enzymatic reactions.
investigated only recently.
Biological routes towards alkenes are rare and have been
[
14–20]
For instance, oxidative decarboxylation of (saturated) fatty
[
21–23]
[24]
acids by the P450 mono-oxygenase OleT
and the non-heme oxygenase UndA yields
[
25]
terminal alkenes on a small scale. In order to avoid the requirement for sophisticated and
sensitive electron-transfer proteins, redox-neutral decarboxylation of p-hydroxycinnamic
acids ('phenolic acids') derived from the breakdown of lignin catalysed by phenolic acid
[
7]
[26]
decarboxylases was investigated. The latter enzymes act via simple acid-base catalysis,
which requires the presence of a phenolic 'activating' group in the substrate, which severely
limits their applicability. Furthermore, the electron-rich p-hydroxystyrenes thus obtained are
not very stable and are prone to (spontaneous) oxidation and polymerisation.
Ferulic acid decarboxylases (FDCs) acting on 'non-phenolic' cinnamic acids are an intriguing
[
27–29]
new class of decarboxylases.
They are distinct members of the UbiD family of
decarboxylases and catalyse the non-oxidative decarboxylation of acrylic acid derivatives
[
30,31]
such as cinnamic, ferulic and sorbic acid yielding the corresponding terminal alkenes.
Recent structural and mechanistic studies revealed that these enzymes utilise a prenylated
[
32]
derivative of flavin (prFMN), a cofactor synthesised by UbiX. FDC-catalysed decarboxylation
of cinnamic acid derivatives mediated by prFMN is proposed to proceed via a 1,3-dipolar
[
27,28]
cycloaddition,
in which prFMN acts as 1,3-dipolar diene owing to its azomethine ylide
[
28,33–35]
character (Figure 1).
While this type of transformation - commonly referred to as
[
36,37]
'Huisgen-reaction'
this reaction are rare.
- is widely utilised in heterocyclic synthesis, enzymatic equivalents to
[
38–41]
6
6
6
4
5
6
Herein, we report on the broad substrate scope and high activity of three FDCs (Scheme 1).
Crucial reaction parameters such as co-solvent compatibility, temperature- and pH-optima of
these enzymes were investigated. Furthermore, we also performed a preparative-scale
2
This article is protected by copyright. All rights reserved.

ChemCatChem
10.1002/cctc.201800643
6
6
7
8
biotransformation and tested ScFDC in the (reverse) carboxylation of terminal alkenes
[
4,42–45]
utilisingKHCO orpressurized CO as C source.
1
3
2
6
7
7
7
9
0
1
2
Scheme 1. Enzymatic decarboxylation of ,-unsaturated carboxylic acids.
Results and Discussion
7
7
7
7
7
7
7
8
8
8
8
8
8
8
8
8
8
9
9
9
9
9
9
9
9
3
4
5
6
7
8
9
0
1
2
3
4
5
6
7
8
9
0
1
2
3
4
5
6
7
Optimisation of biotransformation conditions
In order to assess the biocatalytic potential of FDCs, three previously described
[
28]
representatives from Aspergillus niger (AnFDC), Saccharomyces cerevisae (ScFDC) and
Candida dubliniensis (CdFDC) were each co-expressed with the native E. coli UbiX in E. coli to
UbiX
UbiX
UbiX
produce the holo-enzymes AnFDC , ScFDC and CdFDC . In this system, the FDCs were
fused with a polyhistidine tag, whereas UbiX was co-expressed untagged to enable in vivo
production of prFMN, allowing for the purification of the prFMN-bound FDC to homogeneity
by Ni affinity chromatography.
UbiX
Using purified AnFDC as the catalyst, biotransformation conditions were optimised for the
decarboxylation of 20 mM 1a as a model reaction. The enzyme displayed a broad pH window
(pH 6.0 – 9.0) with highest conversions of >99% achieved at pH 7.5 (phosphate buffer) and pH
UbiX
8.0 (Tris-HCl buffer) (Supporting information Section S1.1). AnFDC
showed high activity
between 20 and 45 °C with highest rates obtained at 37 – 42 °C, however protein
precipitation was observed upon incubation at ≥37 °C for 1h. Hence, subsequent reactions
were performed at 30 °C. Under the optimised conditions, biotransformations were
performed with i) freshly purified enzyme preparations (snap-frozen or lyophilised), ii) E. coli
whole cells containing AnFDC either as fresh resting whole cells or in lyophilised form, and iii)
using fresh cell-free extract (snap-frozen or lyophilised). In all cases, conversions of >80%
were achieved highlighting the suitability of FDCs in isolated form or as whole cell biocatalyst.
Similarly, lyophilised whole-cell ScFDC showed a broad temperature optimum between 30 °C
and 45 °C, with a sharp drop beyond this value, while the pH-profile peaked at 6.0
(Supporting information Section S1.3). Monitoring ScFDC-catalysed decarboxylation of
(aromatic) ferulic and (non-aromatic) sorbic acid over time revealed a typical hyperbolic
decline of the substrate concentration, where ~90% conversion was reached within ~8 h, and
the reaction was complete after ~16 h (Supporting Information, Figure S5). Control reactions
3
This article is protected by copyright. All rights reserved.

ChemCatChem
10.1002/cctc.201800643
9
9
8
9
featuring all reaction conditions but containing E. coli whole cells harbouring an empty pET
vectorrevealed no conversion of 1a.
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
00
01
02
03
04
05
06
07
08
09
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
Substrate tolerance of FDCs
To highlight the synthetic utility of FDCs, the substrate scope of AnFDC, ScFDC and CdFDC was
investigated. An array of 60 different α,β-unsaturated carboxylic acids were tested in the
decarboxylation direction encompassing substituted cinnamic acids and heterocyclic analogs
thereof, as well as non-aromatic acrylic acid derivatives and α,β-acetylenic substrates
(Scheme 2 & Figure 1). Initially, isolated enzymes were used for the substrate profiling study
(Table 1). In addition, ScFDC was also applied as lyophilised whole cell preparation
(overexpressed in E. coli) to evaluate its applicability on preparative-scale for potential
industrial use. Overall, a broad set of substrates covering different structural motifs and
electronical properties were employed (Table1).
First, a range of cinnamic acid derivatives with various substituents at the p-position of the
aromatic moiety (1a-11a) were examined. Substrates bearing weakly electron-withdrawing
-
groups such as p-halogens (2a-4a) and weakly e -donating groups such as p-methyl (5a) were
-
well tolerated by the enzymes affording >84% conversion (Table 1, entries 2-5). Strong e -
donating groups such as p-NH 6a, p-OH 7a and p-OMe 8a were perfectly accepted by whole
2
cells (c = 86-99%, entries 6-8) while a drop in conversion was observed with purified enzymes
-
as catalyst (c = 61-80%). A strong e -withdrawing p-NO
2
group (10a) led to diminished
conversions (c = 18-50%, entry 10) using purified enzymes. Steric restriction seems to appear
with a larger p-Ph group (9a) which was only reasonably accepted by FDC from A. niger (c =
40%, entry 9). Complete loss of activity was observed with an even larger substituent (p-OPh,
48, Figure 1). Substrate 11a which carries two carboxyl groups was regioselectively
decarboxylated yielding 4-vinyl benzoic acid (11b) as sole product, albeit in low conversions
of up 8% (entry 11). Remarkably enough, in contrast to phenolic acid decarboxylases (PADs),
the confining requirement for an activating p-hydroxy group proved to be dispensable which
is in line with the proposed 1,3-dipolarcycloaddition mechanismof FDCs.
The influence of the substitution pattern at the aromatic ring on the enzyme’s performance
has been further evaluated applying mono- (o- or m-, for p- see above), di-, tri- and penta-
functionalised cinnamic acid derivatives. A NO -substituent in m-position was similarly
2
-
tolerated as the p-analogue (10a versus 15a, entries 10, 15) whereas a strong e -donating
group (such as OH) in m-position led to reduced reaction rates compared to the p-pendant
(7a versus 12a, entries 7, 12). Di-substitution in p- and m-position was well accepted (p-OH
4
This article is protected by copyright. All rights reserved.

ChemCatChem
10.1002/cctc.201800643
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
61
62
63
64
65
and m-OMe, ferulic acid, 17a, c up to >99%; p- and m-OMe, 19a, c >99%, entries 17, 19) as
-
long as the m-substituent was not too e -pushing (p- and m-OH, caffeic acid, 18a) which led to
a significant drop in conversion (c = 33%, entry 18) correlating with the results from above.
The p-naphthyl derivatives (30a and 31a) which formally correspond to a p-/m-di-substitution
-
with weak e -donating groups were excellent substrates, which were quantitatively
decarboxylated (c >99%, entries 30, 31). The size as well as the electronic nature of the o-
substituents seem to play a crucial role which were well tolerated as long as they were small
(F, 20a, c = 82%, entry 20; F, 25a, c >81%, entry 25; Me, 16a, c >99%, entry 16). Sterically
more demanding methoxy- (21a, c = 36%; 22a, c = 31%, entries 21, 22) and nitro-groups (14a,
-
c up to 10%, entry 14) were less favoured which also applies to polar (strong e -donating) o-
substituents such as OH (13a, c up to 8%, entry 13) and led to a complete loss of activity in
case of two polar (o- and p-OH) groups (49, Figure 1). Tri-substituted compounds with
functional groups significantly larger than a F-atom were poor substrates (sinapic acid, 23a, c
= 3-15%, entry 23; 24a, c = 5-8%, entry 24).
The substrate profiling was further extended to α,β-unsaturated carboxylic acids containing
O-, S- and N-heteroaromatic systems at C3. The enzymes were excellent catalysts for the
decarboxylation of 2-furyl- (26a) and 2-thienyl acrylic acid (27a) furnishing the corresponding
UbiX
vinyl products in up to >99% conversion. AnFDC was also capable of decarboxylating the
imidazole-derivative 28a albeit with very low rate (c = 5%, entry 28), which is presumably
caused by the high degree of protonation (~90%/100%) at pH 6.0/7.5 creating a positive
charge. The bicyclic indole-derivative (29a) was reasonably well accepted (c up to 42%, entry
29).
In contrast to PADs which did not accept substitution (e.g. Me-group) at the α- or -carbon
atom to the carboxylate, FDCs showed a more relaxed behaviour tolerating small groups at
these positions (α-F, 32a, c up to >97%; α-Me, 33a, c =20-60%; -Me, 34a, c = 50-85%; entries
32-34), whereas bulky substituents led to a marked decrease (α-NHCOMe, 35a, c = 3-6%,
entry 35) or even loss of FDCactivity (α-Ph, 50, Figure 1).
In general, compounds lacking a C=C-spacer between the carboxylate and the aromatic
system were notconverted (compound 38– 44, Figure 1).
Conjugated 2,4-di-unsaturated acids 36a (sorbic acid) and 37a were excellent substrates,
which were quantitatively decarboxylated into the corresponding 1,3-dienes by whole cells
and only a minor decrease in rates were observed with isolated enzymes. The acceptance of
unsaturated substrates lacking an aromatic system by FDCs constitutes a valuable extension
of the substrate portfolio in the bio-decarboxylation.
5
This article is protected by copyright. All rights reserved.

ChemCatChem
10.1002/cctc.201800643
1
1
1
1
1
66
67
68
69
70
However, α,β-mono-unsaturated and 2,6-dienoic acids were unreactive, regardless of their
open-chain (52-54, 57) or cyclic structure (55, 56). Likewise, acetylenic substrates (58-60) and
symmetrical (E,E)-muconic acid (47) did not react. A switch of the C=C-bond configuration
from(E) to (Z) (51) resulted in substrate rejection (Figure 1).
1
1
71
72
Scheme 2. Substrates (1a – 37a) decarboxylated by FDCs and their corresponding products (1b – 37b).
1
1
73
74
Table 1. FDC-catalysed decarboxylation of acrylicacid derivatives (1a-37a).
Entry Substrates
Conversion [%]
UbiX
UbiX
UbiX
AnFDC
ScFDC
CdFDC
UbiX
[a]
purified ScFDC
E. coli whole cells
1
2
3
4
5
6
7
8
9
1a
2a
3a
4a
5a
6a
7a
8a
9a
>99
>99
n.d.
>99
>99
78
>99
88
>99
>99
>99
n.d.
>99
>99
86
96
84
n.d.
97
n.d.
98
>99
75
98
61
68
73
80
n.d.
40
n.d.
<5
>99
n.d.
n.d.
<5
6
This article is protected by copyright. All rights reserved.

ChemCatChem
10.1002/cctc.201800643
1
0
10a
11a
12a
13a
14a
15a
16a
17a
18a
19a
20a
21a
22a
23a
24a
25a
26a
27a
28a
29a
30a
31a
32a
33a
34a
35a
36a
37a
50
25
n.d.
n.d.
n.d.
n.d.
n.d.
n.d.
>99
>99
33
18
[
b]
[b]]
[b]
11
5
5
8
12
6
8
26
5
38
4
13
14
10
5
4
15
61
17
16
16
>99
35
>99
47
97
17
47
18
n.d.
n.d.
n.d.
n.d.
n.d.
15
n.d.
n.d.
n.d.
n.d.
n.d.
3
n.d.
n.d.
n.d.
n.d.
n.d.
6
19
>99
82
20
21
36
22
31
23
9
24
5
8
n.d.
n.d.
>99
>99
<1
6
25
91
94
81
26
>99
92
>99
87
>99
68
27
28
5
n.d.
14
n.d.
31
29
42
n.d.
n.d.
>99
n.d.
n.d.
n.d.
n.d.
>99
>99
30
>99
>99
97
>99
>99
58
>99
>99
47
31
32
33
22
60
20
34
85
77
50
35
6
<3
<3
36
95
80
88
37
99
90
87
-
1
1
1
1
1
1
1
75
76
77
78
79
80
Reaction conditions using purified enzymes: substrate (5 mM), purified enzyme (0.2 mg mL ), NaPi
buffer (100 mM, pH 7.5), 30 °C, 180 rpm, 18 h; conversion values were determined by GC-MS or HPLC
UbiX
analysis; [a] reaction conditions with E. coli whole cells: substrate (10 mM), ScFDC
E. coli whole cells
-
1
(30 mg mL ), NaPi buffer (100 mM, pH 6.0), 30 °C, 120 rpm, 18 h, 5% v/v DMSO (20% v/v DMSO for
31a and 48); n.d.= not determined; [b] decarboxylation occurred at the acrylic acid moiety furnishing
4-vinyl benzoic acid (11b) as soleproduct.
7
This article is protected by copyright. All rights reserved.

ChemCatChem
10.1002/cctc.201800643
1
81
1
1
82
83
Figure 1. Substrates rejected by FDC (conversion <1%), for standard conditions seeTable1.
1
1
1
1
1
1
1
1
1
1
1
1
84
85
86
87
88
89
90
91
92
93
94
95
The results from Scheme 2, Table 1 and Figure 1 reveal a clear substrate structure-activity
pattern of the FDCs enzymes:
i) Minimal substrate requirements consist of an acrylic acid moiety with an extended π-
system in the β-position, which is fulfilled by an aromatic system or a (minimal) second
conjugated C=Cbond.
ii) Compounds lacking an α,β-C=C bond, which is an essential requirement to undergo 1,3-
dipolarcycloaddition with the prFMN cofactor, are unreactive, as well as acetylenicanalogs.
iii) The (E) or (Z) configuration of the reactive C=Cbond seems to be critical.
iv) Sterically demandinggroups impedereaction rates.
v) Strongly electron-donatinggroups impede reaction rates.
Structural and mechanisticaspects
1
1
1
1
2
2
2
2
96
97
98
99
00
01
02
03
Azomethine ylides have been characterised as dipoles with pronounced nucleophilic
[
46]
character. Due to theirinherent reactivity, they are usually prepared in situ, for example by
[
47,48]
ring-opening of aziridines.
Initial cycloadduct formation in the reaction mechanism of
FDC is expected to proceed through interaction between the HOMO of prFMN and the
[
49]
substrate’s LUMO.
Thus, potential substrates must show a somewhat ambiguous
character: the α,β-unsaturated carboxylic acid molecule must be electrophilic enough to
allow cyloadduct formation with the nucleophilic cofactor in the first place. However, after
decarboxylation, the cycloadduct should dissociate easily into the olefinic decarboxylation
8
This article is protected by copyright. All rights reserved.

ChemCatChem
10.1002/cctc.201800643
2
2
2
2
2
2
2
2
2
2
2
2
2
2
2
2
2
2
04
05
06
07
08
09
10
11
12
13
14
15
16
17
18
19
20
21
product and cofactor, allowing a new catalytic cycle to initiate. This suggests that
decarboxylation itself (the loss of one EWG as CO ) is the crucial step that raises electron
2
density in the substrate-cofactor adduct, promoting it to undergo cyclo-elimination. Strongly
electron-deficient dipolarophiles are potent mechanistic inhibitors of FDC enzymes, which
[
35]
has been demonstrated experimentally. Additionally, the enzyme only accepted substrates
with an extended π-system conjugated to the acrylic acid moiety. This preference ensures
diffuse electron density in both cofactor and substrate, which allows enhanced matching
[
50–53]
orbital energy levels according to HSAB and FMO principles.
These considerations are in
excellentagreementwith the observed substrate preference of FDC enzymes.
An analysis of the AnFDC active site architecture provides a rationale for FDC tolerance to
1
cinnamic acid residues bearing small substituents (Figure 2a, R = F/Me) at the α-carbon to
2
the carboxylate (Figure 2). The orientation of the substrate in the active site positions R and
3
R substituents at a water filled cavity (Figure 2a), indicating that large groups can be
accommodated at the m- and p-positions of the aromatic ring. In contrast, the AnFdc1
1
structure highlights potential steric constraint with large R substituents and o-substitutions
4
of the aromatic ring (R ). These predictions are in excellent agreement with
biotransformation datapresented in Table 1.
2
22
2
2
2
2
2
2
2
2
2
23
24
25
26
27
28
29
30
31
Figure 2. Mechanism and substrate scope of ferulic acid decarboxylases (FDCs). a) Active site of
Aspergillus niger FDC (AnFDC) in complex with α-fluorocinnamic acid (PDB code 4ZAB). A transparent
surface reveals the solvent accessible surface on the re side of the prFMN that is complementary in
shape to the substrate. In addition, a water filled cavity is present near the cofactor ribityl moiety
(indicated by circle), providing ample space for m- and p-substitutions of the aromatic ring. Potential
1
steric constraint occurs with cinnamic acid derivatives bearing bulky substituents at the α-carbon (R )
4
to the carboxylate or o-substitutions of the aromatic ring (R ). b) a general mechanism proposed for
reversible decarboxylation of acrylic acid derivatives by prFMN in FDC enzymes via 1,3 dipolar
cycloaddition.
9
This article is protected by copyright. All rights reserved.

ChemCatChem
10.1002/cctc.201800643
2
2
2
2
2
2
2
2
2
2
2
2
2
2
2
2
2
2
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
Catalyticperformance
To assess the relative activity of the three FDCs, we determined the total turnover numbers
UbiX
for 7 representative substrates. AnFDC
displayed a high rate indicated by a turnover
-
1
frequency (TOF) of 370 min for cinnamic acid (1a). In general, the enzymes showed highest
activity towards acrylic acid derivatives bearing either an unactivated phenyl (1a) or naphthyl
group (31a, Table 2, entries 1, 7), affording a total turnover number of up to 55,000 for these
substrates. This activity value compares favourably with other industrially relevant enzymatic
[54,55]
UbiX
UbiX
reactions.
ScFDC and AnFDC displayed superior activity for the decarboxylation of
cinnamic acid (1a), however, the activity towards naphthylacrylic acid (31a) were comparable
UbiX
for the three enzymes. Although CdFDC
exhibited comparatively the lowest activity
towards cinnamic acid (1a) and 5a (p-Me-derivative, entry 2), it was the superior catalyst in
the decarboxylation of p-coumaric acid (7a), ferulic acid (17a) and the O-heterocyclic
derivative (26a). ScFDC displayed the highest tolerance to intensified reaction conditions,
showing high activity even at increased substrate loading of 1a up to 100 mM. However, at
>60 mM of 1a, decrease in reaction rate was observed, owing to substrate or product
inhibition.
Table 2. Comparison of FDCs catalytic activity forrepresentativesubstrates
Entry
Substrates
Total turnover number
UbiX
UbiX
UbiX
AnFDC
ScFDC
CdFDC
3
3
3
x 10
x 10
x 10
1
2
3
4
5
6
7
1
a
33.0 ± 4.0
11.0 ± 0.1
4.0 ± 0.8
4.2 ± 0.6
6.3 ± 0.8
6.3 ± 0.5
17.0 ±0.2
55.0 ± 0.5
11.0 ± 0.2
5.1 ± 0.4
5.2 ± 0.3
6.0 ± 1.0
11 ± 2.0
8.0 + 1.3
6 ± 0.5
5a
7a
7.5 ± 0.2
10.0 ± 0.4
11.0 ± 0.2
11.0 ± 1.0
15.0 ± 0.10
17a
26a
27a
31a
13.0 ± 0.2
-
1
2
2
2
2
50
51
52
53
Reaction conditions: substrate (20 – 1100 mM), purified enzyme (0.2 mg mL ), NaP buffer (200 mM,
i
pH 7.5), 30 °C, 180 rpm, 8 h. Total turnover numbers were calculated from conversions after 8 h
incubation. Reactions were run in triplicate and errors represent the standard deviation from the
mean.
2
2
54
55
1
0
This article is protected by copyright. All rights reserved.

ChemCatChem
10.1002/cctc.201800643
2
2
2
2
2
2
2
2
56
57
58
59
60
61
62
63
Co-solventcompatibility and upscaling
In order to overcome solubility problems of lipophilic substrates or products, the
compatibility of ScFDC with organic co-solvents was tested using 14 water-miscible and -
immiscible (co-)solvents at concentrations of 5%, 10% and 20% v/v (Figure 3). While water-
immiscible biphasic systems containing dichloromethane, chloroform or ethyl acetate led to
significant enzyme deactivation, water-miscible co-solvents were tolerated surprisingly well
at 5% v/v. MeOH, EtOH, 1,2-dimethoxyethane and DMF could be employed at 10% v/v and
DMSO was even compatible at20% v/v.
100
90
80
70
60
50
40
30
20
10
0
5
1
2
% v/v
0% v/v
0% v/v
Water-miscible
Water-immiscible
Organic co-solvents
Figure 3. Decarboxylation of sorbic acid (36a) by ScFDC in the presence of organic solvents. Reaction
conditions: NaPi (100 mM, pH 6.0), whole lyophilised cells of E. coli containing ScFDC (30 mg mL ),
substrate (10 mM), organic co-solvents (5 – 20% v/v), 30 °C, 120 rpm, 18 h. Conversions were
determined by calibrated RP-HPLC.
2
2
2
2
2
2
64
65
66
67
68
69
-
1
2
2
2
2
2
2
2
2
2
70
71
72
73
74
75
76
77
78
In order to prove the applicability of this method on preparative scale, the decarboxylation of
ferulic acid (17a) was performed. The substrate load was increased from 10 to 16.8 mM in 20
mL reaction volume. HPLC-analysis revealed incomplete conversion of the starting material
(48%). The product was isolated by extraction of the aqueous phase with EtOAc and was
purified by flash chromatography yielding 19 mg (38% yield) of 17b. Product identity and
purity were confirmed by NMRspectroscopy (see SupportingInformation).
1
1
This article is protected by copyright. All rights reserved.

ChemCatChem
10.1002/cctc.201800643
2
2
2
2
2
2
2
2
2
2
2
2
2
2
2
2
2
79
80
81
82
83
84
85
86
87
88
89
90
91
92
93
94
95
Carboxylationexperiments
Converting the decarboxylation of acrylic acid derivatives into the reverse carboxylation
[54-56]
reaction has been demonstrated for o-(de)carboxylases,
phenolic acid
indole-3-carboxylate
3-Methoxy-4-hydroxystyrene (17b), 1,3-pentadiene (36b) and two
[26]
[59–61]
(de)carboxylases
and
pyrrole-2-carboxylate
or
[
62]
(de)carboxylases.
further 1,3-dienes (61, 62), which fulfil the minimal substrate requirements for ScFDC, were
subjected to carboxylation with ScFDC using elevated concentrations of bicarbonate (0.5 – 3
M), as well as pressurized CO
2
(30 bar) as CO -source (Scheme 3). Using varying amounts of
2
-
1
ScFDC preparation (30 – 50 mg mL lyophilised cells) and DMSO (5 – 20% v/v) as co-solvent,
no formation of the desired products was observed after18 h.
-
1
Enhanced biocatalyst loading (100 mg whole cells mL ) and CO (30 bar) produced small
2
amounts of 17a from 17b within 18 h (c <1%). Although this might be taken as proof-of-
principle for the carboxylation of alkenes with ScFDC, the reaction was plagued by
decomposition of (sensitive) vinylphenol 1b, dimerization of structurally similar 4-vinylphenol
[63]
(7b) has been reported. Using 1,3-pentadiene (36b), isoprene (61) or myrcene (62) did not
resultin any formation of carboxylation productusingKHCO
3
.
2
96
2
2
2
3
3
3
3
3
97
98
99
00
01
02
03
04
Scheme 3. Substrates tested in the carboxylation direction with ScFDC (conversions <1%). The arrow
indicates the expected carboxylation site. Reaction conditions using KHCO3: NaPi (100 mM, pH 5.5),
-
1
whole lyophilised cells of E. coli containing ScFDC (30 – 50 mg mL ), 10 mM substrate (17b, 36b, 61,
62), KHCO3 (0.5 – 3 M), 30 °C, 120 rpm, 18 – 20 h, 5 – 20% v/v DMSO or DME. Reaction conditions-
using CO2 (gas): NaPi (250 mM, pH 7.5), whole lyophilised cells of E. coli containing ScFDC (100 mg mL
1
), 10 mM substrate (17b), 30 bar CO2, 30 °C, 50 rpm, 18 h, 5% v/v DMSO. Conversions were
determined by calibrated RP-HPLC.
3
3
3
3
3
3
3
05
06
07
08
09
10
11
Experimental Section
General
Commercially available chemicals and reagents of the highest purity were purchased from
Sigma-Aldrich (Poole, Dorset, UK) unless stated otherwise. Compounds 2a, 48, 51, 36a, 34b,
27a, 6a, 38, 61 were donated by BASF (Ludwigshafen, Germany); 16a was obtained from
abcr; 5b, 52, 59, 62 were purchased from Fluka; 8a and 8b were sourced from Lancaster and
7a and 7b were purchased from Alfa Aesar (Karlsruhe, Germany), while 58 was purchased
1
2
This article is protected by copyright. All rights reserved.

ChemCatChem
10.1002/cctc.201800643
3
3
3
12
13
14
from Acros Organics (Geel, Belgium). HPLC solvents were obtained from Sigma-Aldrich (Poole,
Dorset, UK) or ROMIL (Waterbeach, Cambridge, UK) and GC gases from BOC gases (Guildford,
UK).
3
3
3
3
3
3
3
3
3
3
3
3
15
16
17
18
19
20
21
22
23
24
25
26
Production and preparation of biocatalysts
UbiX
UbiX
UbiX
Cloning, expression and purification of AnFDC , ScFDC and CdFDC were performed as
[26,30]
previously described.
The purified enzymes were either snap-frozen or stored at -80 °C
until when needed or lyophilised and stored at -20 °C. For the preparation of the whole cell
biocatalysts, cultivation was performed in 500 mL LB broth medium with kanamycin (30
-
1
-1
μgꢀmL ) and ampicillin (50 μgꢀmL ). Cultures were initially incubated at 37 °C with shaking at
200 rpm. At an optical density (OD600) between 0.6 and 0.8, isopropyl β-D-1-
thiogalactopyranoside (IPTG) was added to a final concentration of 0.3 mM to induce protein
expression and MnCl to the final concentration of 1 mM was added. Incubation was
2
continued at 20 °C and 250 rpm for 18 h. Cells were then harvested by centrifugation and
suspended in sodium phosphate buffer (100 mM, pH 7.5). The harvested cells were used as
fresh restingcells orlyophilised preparation.
3
3
27
28
General procedure for isolated enzyme decarboxylation
UbiX
3
3
3
3
3
3
3
3
3
3
3
3
3
3
3
3
3
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
For FDC -catalysed decarboxylation reaction using purified enzyme preparation, a 500 µL
reaction mixture contained carboxylic acid substrate (5 mM), 2 – 10% (v/v) DMSO, purified
UbiX
-1
FDC (0.2 mg mL ) in sodium phosphate buffer (100 mM, pH 7.5). Reaction mixtures in 2
mL tightly-closed glass vials were incubated at 30 °C with 180 rpm shaking for 18 h, after
which the enzyme was inactivated by the addition of an equal volume of MeCN and
vigorously mixed. The reaction mixtures were centrifuged (4 °C, 2,831 rcf, 5 min); the clear
supernatant was filtered and analysed by reverse phase HPLC. Where analysis of
biotransformation was performed on the GC-MS, an equal volume of EtOAc (containing a
known concentration of an internal standard where necessary) was added to
biotransformation mixture, vigorously mixed, centrifuged and the organic layer was extracted
twice. The aqueous layer was then acidified to a pH of ~2 and further extracted with EtOAc
with centrifugation (4 °C, 2,831 rcf, 5 min) to improve the separation of phases. The organic
layers were combined and dried over anhydrous MgSO
MS.
4
and samples were analysed by GC-
General procedure for whole-cell decarboxylation
Lyophilised whole cells of E. coli (30 mg) containing overexpressed ScFDC were rehydrated for
1
3
This article is protected by copyright. All rights reserved.

ChemCatChem
10.1002/cctc.201800643
3
3
3
3
3
3
3
3
3
3
3
46
47
48
49
50
51
52
53
54
55
56
30 min at 30 °C with 120 rpm shakingin phosphate buffer (950 μL, 100 mM, pH 6.0) in 1.5 mL
plastic Eppendorf tubes. Substrates were supplied by adding 50 μL of 200 mM stock solution
in DMSO to achieve a substrate concentration of 10 mM in 1 mL of total reaction volume,
followed by incubation for 18 h at 30 °C with shaking in horizontal position at 120 rpm under
exclusion of light. For substrates showing limited solubility (48, 31a), lyophilised cells were
suspended in buffer (800 μL) and after rehydration, pure DMSO (150 μL) was supplemented
followed by addition of a substrate stock (50 μL) and incubation. After given reaction time,
samples were centrifuged at 14,000 rpm for 10 min and supernatant (100 μL) was diluted
with 900 μL of H O/MeCN/trifluoroacetic acid (TFA, 50:50:3) to precipitate residual protein.
2
The diluted sample was centrifuged again, followed by analysis with HPLC. All reactions were
performed in triplicate plus negative control withoutlyophilised cells.
3
3
3
3
3
3
3
3
3
57
58
59
60
61
62
63
64
65
Co-solventstudies
Stock solutions of 36a (200 mM) were prepared in MeCN, acetone, 1,4-dioxane, MeOH, EtOH,
i-PrOH, t-BuOH, DME, DMF, DMSO, THF, DCM, chloroform and EtOAc. Lyophilised cells were
rehydrated in 800, 900 or 950 μL phosphate buffer (100 mM, pH 6.0). 50 μL of the
corresponding stock solution was added to the mixture and pure co-solvent was added to
achieve a reaction volume of 1 mL, followed by incubation. For water-miscible co-solvents,
sample workup and analysis was performed as described above. For immiscible solvents,
partial evaporation of the organic layer was observed and therefore, only the aqueous phases
were analysed usingHPLC.
3
3
3
3
3
3
3
66
67
68
69
70
71
72
General procedure for carboxylation using KHCO3
Lyophilised cells (30 – 50 mg) were rehydrated in phosphate buffer (800 – 950 μL, 100 mM,
pH 5.5). Pure co-solvent (0 – 150 μL) followed by substrate stock (50 μL; 17b, 36b and 61 200
mM in DMSO or 62 200 mM in DME) was added to achieve a reaction volume of 1 mL,
followed by transfer of the mixture into a screw-neck glass vial containing KHCO (0.5 – 3 M).
3
The vessels were swiftly closed to avoid the loss of emerging CO gas and were incubated for
2
18 – 20 h.
3
3
3
3
3
73
74
75
76
77
General procedure for carboxylation using pressurized CO
2
Lyophilised cells (300 mg) were rehydrated in phosphate buffer (2850 μL, 250 mM, pH 7.5).
150 μL of a 200 mM stock solution of 17b in DMSO was added and the mixture was
transferred into a steel pressure vessel equipped with a stirring bar. The reaction mixture was
pressurized with technical CO gas (30 bar) and was stirred (50 rpm) at 30 °C for 18 h.
2
1
4
This article is protected by copyright. All rights reserved.

ChemCatChem
10.1002/cctc.201800643
3
3
3
3
3
3
3
3
3
3
3
3
3
3
78
79
80
81
82
83
84
85
86
87
88
89
90
91
Preparative scalebiotransformation
560 mg lyophilised cells were rehydrated in a plastic vial (50 mL) with phosphate buffer (19
mL, 100 mM, pH 6.0). 17a (65.1 mg, 0.34 mmol) and hydroquinone (6.5 mg, 0.06 mmol,
radical scavenger to inhibit product decomposition) were dissolved in MeOH (1 mL) and
added to the mixture. The vessel was wrapped in aluminum foil to ensure protection from
light and was incubated for 24 h with shaking at 120 rpm at 30 °C. Solids were separated by
centrifugation at 4000 rpm at 4 °C for 20 min. The supernatant (100 μL) was diluted with
H
2
O/MeCN/TFA and was subjected to HPLC analysis, which revealed incomplete turnover of
the starting material (48% conversion). The remainingliquid was extracted with EtOAc (4 × 20
mL). The combined organic phases were dried with Na SO and filtered. After evaporation, a
2
4
mixture of off-white solids and dark yellow oil was obtained. The oil was diluted with DCM
and purified by flash column chromatography (silica gel Merck 60, DCM), giving 19 mg (0.13
mmol, 38% yield) of spectroscopically pure 17b as colorless oil with adistinctclove-likeodor.
1
17b: H NMR (300 MHz, DMSO-d ): δ = 9.09 (s, 1H), 7.04 (d, J = 1.9 Hz, 1H), 6.85 (dd, J = 8.1,
6
3
3
3
92
93
94
1.9 Hz, 1H), 6.72 (d, J = 8.1 Hz, 1H), 6.60 (dd, J = 17.6, 10.9 Hz, 1H), 5.63 (dd, J = 17.6, 1.1 Hz,
1
3
1H), 5.05 (dd, J = 10.9, 1.0 Hz, 1H), 3.78 (s, 3H); C NMR (75 MHz, DMSO-d ): δ = 147.69,
6
6
2,63]
146.71, 136.71, 128.79, 119.53, 115.36, 110.95, 109.57, 55.56.
3
3
3
3
3
4
4
4
4
4
4
4
4
4
4
4
4
95
96
97
98
99
00
01
02
03
04
05
06
07
08
09
10
11
Analyses of whole-cell biotransformations
HPLC analysis: HPLC/UV experiments were performed on a HPLC Agilent 1260 Infinity system
with a diode array detector and a reversed-phase Phenomenex Luna C18 column (100 Å, 250
× 4.6 mm, particle size 5 μm, column temperature 24 °C). All compounds were
spectrophotometrically detected at 220, 254, 263, 280 and 310 nm, respectively. Method
-
1
was run over 22 min with H O/TFA (0.1%) as the mobile phase (flow rate 1 mL min ) and a
2
MeCN/TFA (0.1%) gradient (0-2 min 5%, 2-15 min 5-100%, 15-17 min 100%, 17-22 min 100-
5%). Conversions were determined by comparison with calibration curves for products and
substrates prepared with authentic reference material. Due to the instability of the
decarboxylation products, concentrations were determined indirectly via the reduction of
substrate peaks.
Headspace GC-MS analysis: To verify the formation of volatile decarboxylation products not
detectable on HPLC, reactions were performed in glass vials capped with rubber septa.
Volatiles were analysed directly with an Agilent 7697A headspace sampler (oven temp. 80 °C,
loop temp. 90 °C, transfer line temp. 100 °C, vial equilibration time 2 min, vial pressurization
15 psi). In addition, a 10 μL syringe (Agilent syringe FN 26/50/cone) was pre-heated (10 min,
80 °C) to prevent condensation prior to injection. From headspace of reaction vials 9 μL were
1
5
This article is protected by copyright. All rights reserved.

ChemCatChem
10.1002/cctc.201800643
4
4
4
4
4
4
4
4
4
4
4
4
4
4
4
4
4
4
4
4
4
4
4
4
4
4
4
4
4
4
4
4
4
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
injected split-less (for analysis of compound 36b and 37b). For separation and detection, an
Agilent 7890A GC machine (oven temp. 50 °C) with a HP-5ms capillary column (30 m × 0.25
mm × 0.25 μm; stationary phase: bonded and cross-linked 5% phenyl-methylpolysiloxane)
equipped with a 5975C mass-selective detector (electron impact ionisation, 70 eV;
quadrupole mass selection)usingheliumas carriergas was used.
NMR analysis: NMR spectra were recorded with a Bruker AVANCE III 300 MHz spectrometer
using a 5 mm BBO probe at 300 K. Chemical shifts (δ) are expressed in ppm, coupling
constants (J) are given in Hz.
Analysis of purified enzyme biotransformations
GC-MS analysis was performed on an Agilent 5977A Series GC/MSD System with an Agilent
7890B Series GC coupled to Mass Selective Detector. Analysis was performed using GC/MSD
MassHunter Data Acquisition and ChemStation Data Analysis. A 30 m DB-WAX column with
0.25 mm inner diameter and 0.25 μm film thickness (Agilent, Santa Clara, CA, USA) was used.
Analysis method: Inlet temperature: 240 C, detector temperature: 250 C, MS source 230 C,
-
1
-1
heliumflow:1.2mL min ;oven temperature40 – 240 C, 15 C min .
Reverse phase HPLC was performed on an Agilent system (Santa Clara, CA, USA) equipped
with a G1379A degasser, G1312A binary pump, a G1367A well plate autosampler unit, a
G1316A temperature controlled column compartment and a G1315C diode array detector.
Columns used include: Kinetex C18; 250 mm length, 4.6 mm diameter, 5 μm particle size
(Phenomenex, Macclesfield, Cheshire, UK) and Syncronis; C18; 250 mm length, 4.6 mm
diameter, 5μm particle size (Thermo Scientific;Waltham, MA USA).
Substrates standards and product markers, and the resulting biotransformation products
were analysed by reverse phase chiral HPLC using isocratic methods with different solvent
ratios of MeCN and H O, with 0.1% TFA as additive. The flow rate was maintained at 1 mL
2
-1
min and elutes were detected by the UV detector at a wavelength of 245 nm (except for
pyrrole which was monitored at 210 nm). To account for the variation in UV response
between the starting material and the product, relative response factors were experimentally
determined. Correction factors were calculated from the ratio of the slopes of standard
curves plotted for varying concentrations of both the acid and the corresponding alkene at a
UV detection wavelength of 245 nm.
Conclusion
1
6
This article is protected by copyright. All rights reserved.

ChemCatChem
10.1002/cctc.201800643
4
4
4
4
4
4
4
4
4
45
46
47
48
49
50
51
52
53
In summary, we elucidated the substrate scope and high activity of FDCs as reversible
(de)carboxylation catalysts. The enzymes displayed broad substrate tolerance towards a
variety of phenylacrylic acids and heteroaromatic analogs thereof, as well as non-aromatic
2,4-dienoic acids. The minimum structural requirement for substrate acceptance is a non-
aromatic or (hetero)aromatic conjugated π-system linked to C3. The observed substrate-
activity pattern is in agreement with the proposed 1,3-dipolar cycloaddition mechanism.
Steric requirements and the (E/Z)-configuration of the acrylic C=C bond had a strong impact
on reaction rates. Attempts to reverse the reaction into the carboxylation direction in
presence of bicarbonate orpressurized CO were unsuccessful.
2
4
4
4
4
4
54
55
56
57
58
AssociatedContent
Data on pH study, co-solvent compatibility study, analytical protocols including HPLC, GC-MS
analyses and representative traces of biotransformation products are available in the
Supporting Information.
4
4
59
60
Author Information
CorrespondingAuthors
4
4
4
61
62
63
David Leys, Manchester Institute of Biotechnology, School of Chemistry, University of
Manchester, Manchester, 131 Princess Street, Manchester, M1 7DN, UK., e-mail
<David.Leys@manchester.ac.uk>
4
4
4
64
65
66
Kurt Faber, Department of Chemistry, University of Graz, Heinrichstrasse 28, A-8010 Graz,
Austria; e-mail <Kurt.Faber@Uni-Graz.at>, phone: +43-316-380-5332, fax: +43- 316-380-
9840.
4
67
Acknowledgements
4
4
4
4
68
69
70
71
G. A. A. received funding by grant BB/KO17802/1 from the Biotechnology and Biological
Sciences Research Council (BBSRC). S. S. B. was funded by BBSRC iCASE studentship. K. A. P. P.
received funding from the European Research Council grant 695013. DL is grateful to the
Royal Society forWolfson MeritAward.
4
4
4
72
73
74
Funding by the Austrian Science Fund (FWF, project I 1637-N19 and project P 26863-N19) is
gratefully acknowledged. This was also supported by the Austrian BMWFJ, BMVIT, SFG,
StandortagenturTirol and ZIT through the Austrian FFG-COMET-FundingProgram.
4
75
BASF SE (Ludwigshafen) is thanked forthe donation of specialty chemicals.
1
7
This article is protected by copyright. All rights reserved.

ChemCatChem
10.1002/cctc.201800643
4
4
76
77
Conflictof Interests
The authors declare thatthey have no conflictof interestwith the contents of this article
4
4
4
4
78
79
80
81
References
4
82
4
4
4
4
4
4
4
4
4
4
4
4
4
4
4
4
4
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
83
84
85
86
87
88
89
90
91
92
93
94
95
96
97
98
99
00
01
02
03
04
05
06
07
08
09
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
[1] R. Tinikul, P. Chenprakhon, S. Maenpuen, P. Chaiyen, Biotechnol. J. 2018, DOI
10.1002/biot.201700632.
[2] R. Kourist, J.-K. Guterl, K. Miyamoto, V. Sieber, ChemCatChem 2014, 6, 689–701.
[3] K. Fesko, M. Gruber‐Khadjawi, ChemCatChem 2013, 5, 1248–1272.
[4] S. M. Glueck, S. Gümüs, W. M. F. Fabian, K. Faber, Chem. Soc. Rev. 2010, 39, 313–328.
[5] T. Ren, M. Patel, K. Blok, Energy 2006, 31, 425-451.
[6] M. Tuomaala, M. Hurme, A.-M. Leino, Appl. Therm. Eng. 2010, 30, 45–52.
[7] D. H. Jung, W. Choi, K. Y. Choi, E. Jung, H. Yun, R. J. Kazlauskas, B.-G. Kim, Appl. Microbiol.
Biotechnol. 2013, 97, 1501–1511.
[8] G. A. Aleku, et al., ACS Catal. 2016, 6, 3880–3889, "see Supporting Information".
[9] G. A. Aleku, S. P. France, H. Man, J. Mangas-Sanchez, S. L. Montgomery, M. Sharma, F. Leipold, S.
Hussain, G. Grogan, N. J. Turner, Nat. Chem. 2017, 9, 961–969.
[10] G. A. Aleku, J. Mangas-Sanchez, J. Citoler, S. P. France, S. L. Montgomery, R. S. Heath, M . P.
Thompson, N. J. Turner, ChemCatChem 2018, 10, 515–519.
[11] S. P. France, et al., Angew. Chem. Int. Ed. 2017, 56, 15589–15593, "see Supporting Information".
[12] B. M. Nestl, S. C. Hammer, B. A. Nebel, B. Hauer, Angew. Chem. Int. Ed. 2014, 53, 3070–3095.
[13] A. J. L. Wood, et al., Angew. Chem. Int. Ed.. 2017, 56, 14498–14501, "see Supporting
Information".
[14] S. Noda, Y. Kawai, T. Tanaka, A. Kondo, Bioresour. Technol. 2015, 180, 59–65.
[15] M. Takemoto, K. Achiwa, Tetrahedron Lett.1999, 40, 6595-6598.
[16] J. T. Claypool, D. R. Raman, L. R. Jarboe, D. R. Nielsen, J. Ind. Microbiol. Biotechnol. 2014, 41,
1211–1216.
[17] R. Fujiwara, S. Noda, T. Tanaka, A. Kondo, J. Biosci. Bioeng. 2016, 122, 730–735.
[18] R. McKenna, D. R. Nielsen, Metab. Eng. 2011, 13, 544–554.
[19] R. McKenna, B. Thompson, S. Pugh, D. R. Nielsen, Microb. Cell Fact. 2014, 13, 123.
[20] J. M. Salgado, R. Rodríguez-Solana, J. A. Curiel, B. de Las Rivas, R. Muñoz, J. M. Domínguez,
Enzyme Microb. Technol. 2014, 58–59, 22–28.
[21] A. Dennig, M. Kuhn, S. Tassoti, A. Thiessenhusen, S. Gilch, T. Bülter, T. Haas, M. Hall, K. Faber,
Angew. Chem. Int. Ed. 2015, 54, 8819–8822.
[22] M. A. Rude, T. S. Baron, S. Brubaker, M. Alibhai, S. B. Del Cardayre, A. Schirmer, Appl. Environ.
Microbiol. 2011, 77, 1718–1727.
[23] J. L. Grant, C. H. Hsieh, T. M. Makris, J. Am. Chem. Soc. 2015, 137, 4940–4943.
[24] Z. Rui, X. Li, X. Zhu, J. Liu, B. Domigan, I. Barr, J. H. D. Cate, W. Zhang, Proc. Natl. Acad. Sci. U. S. A.
2014, 111, 18237–18242.
[25] N. A. Herman, W. Zhang, Curr. Opin. Chem. Biol. 2016, 35, 22–28.
[26] C. Wuensch, et al., Adv. Synth. Catal. 2015, 357, 1909–1918, "see Supporting Information".
[27] M. Stratford, A. Plumridge, D. B. Archer, Appl. Environ. Microbiol. 2007, 73, 6534–6542.
[28] K. A. P. Payne, et al., Nature 2015, 522, 497–501, "see Supporting Information".
[29] S. A. Marshall, K. Fisher, A. N. Cheallaigh, M. D. White, K. A. P. Payne, D. A. Parker, S. E. J. Rigby,
D. Leys, J. Biol. Chem. 2017, 292, 4623-4637.
[30] G. A. Aleku, B. Nowicka, N. J. Turner, ChemCatChem 2018, 10, 124–135.
[31] S. A. Marshall, K. A. P. Payne, D. Leys, Arch. Biochem. Biophys. 2017, 632, 209–221.
[32] M. D. White, et al., Nature 2015, 522, 502–506, "see Supporting Information".
1
8
This article is protected by copyright. All rights reserved.

ChemCatChem
10.1002/cctc.201800643
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
5
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
61
62
63
64
65
66
67
68
[33] K. L. Ferguson, N. Arunrattanamook, E. N. G. Marsh, Biochemistry 2016, 55, 2857–2863.
[34] F. Lin, K. L. Ferguson, D. R. Boyer, X. N. Lin, E. N. G. Marsh, ACS Chem. Biol. 2015, 10, 1137–1144.
[35] K. L. Ferguson, J. D. Eschweiler, B. T. Ruotolo, E. N. G. Marsh, J. Am. Chem. Soc. 2017, 139,
10972–10975.
[36] H. C. Kolb, M. G. Finn, K. B. Sharpless, Angew. Chem. Int. Ed. 2001, 40, 2004–2021.
[37] R. Huisgen, Proc. Chem. Soc. 1961, 0, 357–396.
[38] D. Leys, N. S. Scrutton, Curr. Opin. Struct. Biol. 2016, 41, 19–26.
[39] V. Piano, B. A. Palfey, A. Mattevi, Trends Biochem. Sci. 2017, 42, 457–469.
[40] C. T. Walsh, T. A. Wencewicz, Nat. Prod. Rep. 2013, 30, 175–200.
[41] M. Baunach, C. Hertweck, Angew. Chem. Int. Ed. 2015, 54, 12550–12552.
[42] K. Plasch, V. Resch, J. Hitce, J. Popłoński, K. Faber, S. M. Glueck, Adv. Synth. Catal. 2017, 359,
959–965.
[43] C. Wuensch, N. Schmidt, J. Gross, B. Grischek, S. M. Glueck, K. Faber, J. Biotechnol. 2013, 168,
264–270.
[44] C. Wuensch, S. M. Glueck, J. Gross, D. Koszelewski, M. Schober, K. Faber, Org. Lett. 2012, 14,
1974–1977.
[45] S. E. Payer, et al., Angew. Chem. Int. Ed. 2017, 56, 13893–13897, "see Supporting Information".
[46] P. Pérez, L. R. Domingo, M. J. Aurell, R. Contreras, Tetrahedron 2003, 59, 3117–3125.
[47] A. L. Cardoso, T. M. V. D. Pinho e Melo, Eur. J. Org. Chem. 2012, 2012, 6479–6501.
[48] P. Dauban, G. Malik, Angew. Chem. Int. Ed. 2009, 48, 9026–9029.
[49] H. Pellissier, Tetrahedron 2007, 16, 3235–3285.
[40] F. Méndez, J. Tamariz, P. Geerlings, J. Phys. Chem. A 1998, 102, 6292–6296.
[51] D. H. Ess, G. O. Jones, K. N. Houk, Adv. Synth. Catal. 2006, 348, 2337–2361.
[52] K. N. Houk, Acc. Chem. Res. 1975, 8, 361–369.
[53] P. K. Chattaraj, J. Phys. Chem. A 2001, 105, 511–513.
[54] T. A. Rogers, A. S. Bommarius, Chem. Eng. Sci. 2010, 65, 2118–2124.
[55] H. Kohls, F. Steffen-Munsberg, M. Höhne, Curr. Opin. Chem. Biol. 2014, 19, 180–192.
[56] C. Wuensch, J. Gross, G. Steinkellner, A. Lyskowski, K. Gruber, S. M. Glueck, K. Faber, RSC Adv.
2014, 4, 9673–9679.
[57] Y. Ishii, Y. Narimatsu, Y. Iwasaki, N. Arai, K. Kino, K. Kirimura, Biochem. Biophys. Res. Commun.
2004, 324, 611–620.
[58] T. Matsui, T. Yoshida, T. Yoshimura, T. Nagasawa, Appl. Microbiol. Biotechnol. 2006, 73, 95–102.
[59] M. Wieser, N. Fujii, T. Yoshida, T. Nagasawa, Eur. J. Biochem. 1998, 257, 495–499.
[60] M. Wieser, T. Yoshida, T. Nagasawa, J. Mol. Catal. B Enzym. 2001, 11, 179–184.
[61] M. Wieser, T. Yoshida, T. Nagasawa, Tetrahedron Lett. 1998, 39, 4309–4310.
[62] T. Yoshida, K. Fujita, T. Nagasawa, Biosci. Biotechnol. Biochem. 2002, 66, 2388–2394.
[63] R. Bernini, E. Mincione, M. Barontini, G. Provenzano, L. Setti, Tetrahedron 2007, 63, 9663–9667.
[64] E. Zago, E. Dubreucq, J. Lecomte, P. Villeneuve, F. Fine, H. Fulcrand, C. Aouf, New J. Chem. 2016,
40, 7701–7710.
[65] P. Terpinc, T. Polak, N. Šegatin, A. Hanzlowsky, N. P. Ulrih, H. Abramovič, Food Chem. 2011, 128,
62–69.
5
5
5
5
5
5
5
5
5
69
70
71
72
73
74
75
76
77
1
9
This article is protected by copyright. All rights reserved.

ChemCatChem
10.1002/cctc.201800643
5
5
5
5
5
5
5
78
79
80
81
82
83
84
Table of Contentgraphics
5
85
2
0
This article is protected by copyright. All rights reserved.