Regioselective para-Carboxylation of Catechols with a Prenylated Flavin Dependent Decarboxylase

Article abstract of DOI:10.1002/anie.201708091

The utilization of CO₂ as a carbon source for organic synthesis meets the urgent demand for more sustainability in the production of chemicals. Herein, we report on the enzyme-catalyzed para-carboxylation of catechols, employing 3,4-dihydroxybenzoic acid decarboxylases (AroY) that belong to the UbiD enzyme family. Crystal structures and accompanying solution data confirmed that AroY utilizes the recently discovered prenylated FMN (prFMN) cofactor, and requires oxidative maturation to form the catalytically competent prFMN^iminium species. This study reports on the in vitro reconstitution and activation of a prFMN-dependent enzyme that is capable of directly carboxylating aromatic catechol substrates under ambient conditions. A reaction mechanism for the reversible decarboxylation involving an intermediate with a single covalent bond between a quinoid adduct and cofactor is proposed, which is distinct from the mechanism of prFMN-associated 1,3-dipolar cycloadditions in related enzymes.

Full text of DOI:10.1002/anie.201708091

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Accepted Article
Title: Regioselective para-Carboxylation of Catechols by a Prenylated
Flavin Dependent Decarboxylase
Authors: Stefan Payer, Stephen A. Marshall, Natalie Bärland, Xiang
Sheng, Tamara Reiter, Andela Dordic, Georg Steinkellner,
Christiane Wuensch, Susann Kaltwasser, Karl Fisher,
Stephen E.J. Rigby, Peter Macheroux, Janet Vonck, Karl
Gruber, Kurt Faber, Fahmi Himo, David Leys, Tea Pavkov-
Keller, and Silvia M. Glueck
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To be cited as: Angew. Chem. Int. Ed. 10.1002/anie.201708091
Angew. Chem. 10.1002/ange.201708091
Link to VoR: http://dx.doi.org/10.1002/anie.201708091
http://dx.doi.org/10.1002/ange.201708091

10.1002/anie.201708091
Angewandte Chemie International Edition
COMMUNICATION
Regioselective para-Carboxylation of Catechols by a Prenylated
Flavin Dependent Decarboxylase
Stefan E. Payer,+ Stephen A. Marshall,+ Natalie Bärland, Xiang Sheng, Tamara Reiter, Andela Dordic,
Georg Steinkellner, Christiane Wuensch, Susann Kaltwasser, Karl Fisher, Stephen E. J. Rigby, Peter
Macheroux, Janet Vonck, Karl Gruber, Kurt Faber, Fahmi Himo, David Leys, Tea Pavkov-Keller,* Silvia
M. Glueck*
Abstract: The utilization of CO2 as carbon source for organic
syntheses meets the urgent demand of more sustainability in
chemical production. Here we report on the enzyme-catalyzed para-
carboxylation of catechols, employing 3,4-dihydroxybenzoic acid
decarboxylases (AroY) that belong to the UbiD enzyme family.
Crystal structures and accompanying solution data confirm that AroY
utilizes the recently discovered prenylated FMN (prFMN) cofactor,
and requires oxidative maturation to form the catalytically competent
prFMNiminium species. This study reports on the in vitro reconstitution
and activation of a prFMN-dependent enzyme that is capable of
directly carboxylating aromatic catechol substrates under ambient
conditions. A reaction mechanism for the reversible decarboxylation
involving a monocovalent quinoid-cofactor intermediate is proposed,
which is distinct from the prFMN associated 1,3 dipolar cycloaddition
mechanism in related enzymes.
dependent pyruvate decarboxylases)[4] and epoxides (epoxide
carboxylases from Xanthobacter sp.)[5] as well as of
heteroaromatics, such as pyrrole (pyrrole-2-carboxylate
decarboxylase from Bacillus megaterium)[6] and indoles (indole-
3-carboxylate decarboxylase from Arthrobacter nicotianae)[2e]
exhibited a narrow substrate specificity, promising results were
obtained in the biocarboxylation of phenols and styrenes. o-
Benzoic acid decarboxylases and phenolic acid decarboxylases
show a relaxed substrate specificity for the o-carboxylation of
phenols[7] and the β-carboxylation[8] of styrenes respectively,
whilst maintaining their exquisite regioselectivity.
In order to expand the biocatalytic carboxylation toolbox, we
searched for enzymes enabling the regiocomplementary p-
carboxylation of phenols. Most of the already characterized
enzymes either require an ATP-consuming activation
(phosphorylation) prior to carboxylation (phenylphosphate
carboxylases),[9] suffer from rapid loss of activity under aerobic
conditions, especially after purification (4-hydroxybenzoate-[10]
and 3,4-dihydroxybenzoate decarboxylases[2d,11]) or were not
extensively examined in vitro yet,[12] which limits their usability
Carboxylation reactions received considerable attention in
view of the use of CO2 as abundant C1-building block for
sustainable chemical production.[1] However, to date only a few
examples of CO2 fixation reactions have been realized on
industrial scale, mainly due to the high-energy input required for
substrate activation. In recent years, biocatalysts[2] have been
exploited as attractive alternatives to chemical methods[1,3] to
catalyze carboxylation reactions under mild, aqueous conditions.
Whereas the biocatalytic carboxylation of aldehydes (TPP-
for biotransformations. Based on
a literature survey and
preliminary activity screening of heterologously expressed
potential p-carboxylases (see Supporting Information), a 3,4-
dihydroxybenzoic acid decarboxylase form Enterobacter cloacae
(EcAroY) and Klebsiella pneumoniae (KpAroY, 89% identical)
were selected for further studies.[2d,13] Both enzymes belong to
the UbiD-family and are related to a ferulic acid decarboxylase
(Fdc1), which was shown to facilitate (de)carboxylation of
[*]
E-mail: Tea.Pavkov@uni-graz.at; Si.Glueck@uni-graz.at
S. E. Payer, Prof. Dr. K. Faber, Dr. S.M. Glueck
Department of Chemistry, Organic & Bioorganic Chemistry,
University of Graz, NAWI Graz, BioTechMed Graz
Heinrichstrasse 28/2, A-8010 Graz, Austria
S. A. Marshall, Dr. K. Fisher, Dr. S. E. J. Rigby, Prof. Dr. D. Leys
Manchester Institute of Biotechnology, University of Manchester
131 Princess Street, Manchester M1 7DN, United Kingdom
N. Bärland, Dr. S. Kaltwasser, Dr. J. Vonck
Max Planck Institute of Biophysics
cinnamic acids by
(prFMN) cofactor.[14]
a recently discovered prenylated flavin
Owing to the natural presence of the UbiD-associated
prenyltransferase UbiX in the E. coli expression host, lyophilized
whole cells displayed high decarboxylation activity in initial
screenings (Supporting Information). However, when purified
from E. coli, only little enzyme activity could be detected for
either of the two enzymes, despite additional co-expression with
the UbiD associated prenyltransferase UbiX to provide sufficient
prFMN in vivo. Upon in vitro reconstitution with reduced
prFMN,[14a,15] (Figure 1a) decarboxylation activity using 3,4-
dihydroxybenzoic acid (3,4-DHBA, 1) could be detected
following brief exposure to oxygen to generate the active
prFMNiminium form (Figure 1b). The lack of activity for
anaerobically reconstituted protein clearly demonstrates the
requirement of oxidative maturation of the prFMN cofactor. EPR-
and UV-Vis spectroscopy occasionally reveals the presence of a
radical semiquinone intermediate following in vitro reconstitution
and oxidation, reminiscent of the related E. coli UbiD[15] (see
Supporting Information). After reconstitution and maturation, the
enzyme activity has a half-life of only 5–8 minutes when stored
Max-Von-Laue Strasse 3, D-60438 Frankfurt am Main, Germany
Dr. X. Sheng, Prof. Dr. F. Himo
Department of Organic Chemistry, Arrhenius Laboratory,
Stockholm University
SE-10691 Stockholm, Sweden
Dr. A. Dordic, Dr. G. Steinkellner, Prof. Dr. K. Gruber, Dr. T.
Pavkov-Keller
Institute of Molecular Biosciences, University of Graz, NAWI
Graz, BioTechMed Graz
Humboldtstrasse 50, A-8010 Graz, Austria
Prof. Dr. P. Macheroux
Institute of Biochemistry, Graz University of Technology,
Petersgasse 12, A-8010 Graz, Austria
T. Reiter, Dr. A. Dordic, Dr. G. Steinkellner, Dr. C. Wuensch, Dr.
T. Pavkov-Keller, Dr. S.M. Glueck,
Austrian Centre of Industrial Biotechnology (ACIB)
These authors contributed equally.
[+]
Supporting information for this article is given via a link at the end
of the document.
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10.1002/anie.201708091
Angewandte Chemie International Edition
COMMUNICATION
under aerobic conditions but remained undiminished for at least
14 hours when rendered anaerobic again (Figure 1c).
more akin to that observed for UbiD, compared to the more
closed conformation observed for Fdc1 (see Supporting
Information). This open conformation is observed for all
crystallographically independent AroY monomers, and in the 4.6
Å cryo-EM solution structure of apo-EcAroY (Figure 2a).
The active site of AroY is clearly defined by the prenylated
isoalloxazine moiety of the cofactor, and the presence of key
conserved residues (Figure 2b). It is situated near the hinge
point of the prFMN-binding domain motion, and at the cleft
between the oligomerization and the prFMN-binding domains.
Two water molecules are clearly defined in the active site: one
hydrogen atom bonded to His327 and Lys363, the other to
Lys363, His436 and Arg188. We hypothesize that the two water
molecules mimic the two hydroxyl groups at the m- and p-
positions of the substrate protocatechuic acid (1). Given the
relative rigidity of the substrate, we superimposed a catechol
moiety on the two water molecules where the carboxylate moiety
is positioned in close proximity of the prFMN iminium group.
While this has some similarity to the Fdc1:substrate complex
structures (PDB-ID: 4ZA7),[14a] the exact relative position of the
substrate carboxylate and the prFMN N5–C1’ iminium linkage is
distinct. In Fdc1, the substrate α-carbon adjacent to the carboxyl
group is located directly above the prFMN C1’ atom, while in the
case of AroY, the α-carbon is located above the isoalloxazine
N5 (see Supporting Information).
Figure 1. a) Established in vitro reconstitution of AroY with prFMN generated
by UbiX from FMN and DMAP and subsequent aerial oxidation. b) AroY
requires oxygen for activation; activity was measured using 150 µM 3,4-DHBA
(1) and is relative to the highest activity detected. c) Decay of oxidized KpAroY
and EcAroY when kept aerobic or anaerobic. Measured with 150 µM 1. t
8.1 ± 1.3 minutes for KpAroY and 5.3 ± 1.2 minutes for EcAroY.
=
½
Amino acid exchanges within the putative catechol-binding
motif (Arg188, His327, Lys363 to Ala, and His436 to Lys or Thr),
led to a complete loss of activity. The same effect was observed
upon exchange of Arg181 or Glu289 located near the
carboxylate group of the substrate to Ala. The analogous Glu282
in Fdc1 is proposed to be required for the donation of a proton to
the covalent intermediate,[14a,16] which hints at a similar role for
Glu289 as a catalytic acid in AroY.
A
comparison between KpAroY and EcAroY crystal
structures revealed only small differences in accordance to the
similar activities observed in solution experiments. The AroY
monomer consists of an N-terminal prFMN-binding domain
(residues 1–339), an oligomerization domain (residues 340–475)
and a C-terminal α-helix (residues 476–495) (Figure 2a). A
comparison with the previously reported structures of the fungal
Fdc1 and E. coli UbiD[14] shows that AroY structures adopt an
“open” conformation where the position of the prFMN domain is
Figure 2. AroY structure and suggested mechanism based on calculations a) Hexameric quaternary structure of EcAroY: dimer pairs are shown in a ribbon
representation coloured in green and blue. The cryo-EM envelope is shown as a grey translucent surface. In circle is a detailed view of the EcAroY monomeric
structure showing prFMN binding domain (blue), oligomerization domain (green) and C-terminal helix (red). b) Optimized structure of the active site model
employed in the computational study. Atoms marked with asterisks are fixed during the geometry optimization. prFMN cofactor is shown in green and the
substrate in salmon ball and sticks presentation. Distances are given in Å. For clarity, only polar hydrogen atoms and the hydrogen atoms on the substrate are
shown. c) Reaction mechanism suggested on the basis of the calculations.
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10.1002/anie.201708091
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Based on the available structural, mutational and kinetic data
as well as DFT calculations (see Supporting information) we
propose a reaction mechanism involving a quinoid intermediate
(Figure 2c). It has chemical similarity to that proposed for the
phenolic acid decarboxylases (PAD) [8a] and is different from the
1,3 dipolar cycloaddition mechanism proposed for Fdc1[14a,16–18]
The calculations, employing a large model of the active site
consisting of 283 atoms (Figure 2b), suggest that the generation
of a cycloadduct is unlikely in the case of AroY, as it requires
formation of a very strained intermediate (Figure S29). The latter
is not required in the case of the cinnamic acid-like substrates of
Fdc1, for which previous calculations validate the proposed 1,3
dipolar cycloaddition mechanism to the exocyclic alkene moiety.
Whereas the carboxylation of simple phenols was not successful
(non-substrates are shown in Table S7), EcAroY is capable of
catalyzing the regioselective p-carboxylation of catechol (4) at
the expense of either bicarbonate or pressurized CO2 (30 bar).
Apart from 4, EcAroY also accepted pyrogallol (5) as substrate
and carboxylation with bicarbonate predominantly occurred in
para-position to the central hydroxy group to give 2.
Carboxylation in vicinity to the peripheral hydroxy groups
occurred only to a minor extent to give 3. Since the catechol
scaffold appears to be crucial for substrate acceptance, the
carboxylation of substituted catechols 6–8 with bicarbonate was
tested (Scheme 1b). Small electron withdrawing (3-F, 6) to
medium-sized electron donating (3-OMe, 7; 3-Me, 8)
substituents in position 3 were accepted and carboxylation
occurred exclusively in position 5, i.e. in para-position to the
central hydroxy group, which was confirmed by NMR-
spectroscopy (see Supporting Information).
[16b,17b]
Instead, the mechanism involving a quinoid intermediate
is calculated to have feasible energy barriers, which are
significantly lower as compared to the 1,3-cycloaddition
mechanism. The calculated energy profile and optimized
structures of all intermediates and transition states along the
reaction pathway are given in Figure S30–S32.
In conclusion, our data contributes to a better understanding
of the ATP-independent para-carboxylation of phenolic
substrates. Crystal structures and in vitro reconstitution data
unambiguously demonstrate that prFMN is employed as cofactor,
and that oxidative maturation is required for activity. The exact
mechanism of oxidative maturation, and the cause of the
observed oxygen sensitivity remain unclear at this stage. Due to
the preference of AroY towards catechols over simple phenols, a
second hydroxy group seems to be mandatory in order to
provide ideal positioning of the substrate (H-bond between OH
and His327, Figure 2b) in the active site of the enzyme, the latter
further enhancing the nucleophilicity of the (catechol) substrates
to facilitate the nucleophilic addition step onto prFMN. Electron
withdrawing and -donating groups are allowed in the m-position
to the carboxylation site whereas substitution in the o-position
was not tolerated due to steric hindrances in the active site.
In view of synthetic applicability, the substrate scope of
EcAroY was investigated with different protocatechuic acid
derivatives (Scheme 1a, 1–3). EcAroY exhibited high
decarboxylation activity on 3,4-dihydroxybenzoic acid (1, >99%
after 1 h) (Scheme 1a) and gallic acid (2), which proceeded at a
slightly slower rate compared to 1 (ca. 83% conv. after one hour).
For its isomer 2,3,4-trihydroxybenzoic acid (3) however, no
reaction was observed. Highly similar kinetic parameters for the
decarboxylation of 1 with purified and in vitro reconstituted AroY
enzymes were determined for both AroY enzymes (KpAroY
app
app
app
vmax = 4.7 ± 0.4 s–1, Km = 96 ± 23 µM. EcAroY vmax = 4.6
app
± 0.4 s–1, Km = 61 ± 17 µM). These parameters are reported
as apparent values, given the very low prevalence of species
that are likely inactive (i.e. FMN bound or radical prFMN bound
enzyme).
In the context of the wider UbiD-family, the metal assisted-
binding and associated oxidative maturation of prFMN is
common to all biochemically and structurally characterized
enzymes (the fungal Fdc1, E. coli UbiD and AroY). The
substrate binding specificity is distinct for each of these, but in all
cases appears largely governed by residues derived from the
oligomerization domain that are involved in binding to the non-
carboxylate substrate moiety. The carboxylate group on the
other hand is bound near the conserved Glu-Arg-Glu/Asp triad of
ionic residues located near the prFMN N5=C1’ iminium. A key
difference is the relative position of the oligomerization and
prFMN domain, and by extension the substrate binding and the
carboxylate binding motifs. These are considerably closer in the
fungal Fdc1 structure as opposed to the bacterial UbiD and AroY.
A putative domain motion might allow the UbiD/AroY enzymes to
adopt a more Fdc1-like conformation, but this has not been
directly observed. While the quinoid-based mechanism
proposed for AroY suggests an alternative to 1,3-dipolar
cycloaddition in the case of aromatic substrates, it does not
explain how (de)carboxylation of non-phenolic substrates[19] is
achieved. A better understanding of the UbiD-enzyme family will
require further study of these and additional family members.[20]
O
CO2
HO
HO
HO
HO
EcAroY
OH
R
R
CO2
a) Decarboxylation of protocatechuic acids, conv. after 1 h (grey) and 24 h (black) in %:
O
O
OH
O
HO
HO
HO
HO
OH
OH
OH
>99
>99
<1
<1
>83
>99
HO
HO
1
2
3
OH
b) Carboxylation of catechols, conv. after 24 h in %:
15
3
13
2
9
16
(6)[a]
HO
HO
HO
HO
HO
HO
HO
HO
HO
HO
4
5
6
7
8
CH3
O
OH
F
CH3
carboxylation
decarboxylation
Scheme 1. EcAroY substrate screening with lyophilized E. coli whole cells
containing the heterologously expressed decarboxylase. Potassium
bicarbonate (3 M) was used as CO2 source in carboxylation assays;
[a]
pressurized CO2 (30 bar) was used for carboxylation.
To force the reaction equilibrium towards the carboxylation
of suitable substrates, we subjected phenolic substrates 4–7 to a
3 M potassium bicarbonate buffer as CO2-source (Scheme 1b).
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10.1002/anie.201708091
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Acknowledgements
S.E.P. and S.M.G. thank the Austrian FWF for financial funding
within project no. 26863-N19. This work has been supported by
the Austrian BMWFJ, BMVIT, SFG, Standortagentur Tirol and
ZIT through the Austrian FFG-COMET-Funding Program This
work was also supported by the BBSRC grant BB/K017802/1
and a BBSRC iCASE studentship to S.A.M.; D.L. is a Royal
Society Wolfson Merit Award holder.
Matthew Cliff (University of Manchester) performed deuterium-
exchange NMR experiments. Klaus Zangger (University of Graz)
measured NMR spectra. Katharina Plasch aided in the use of
CO2 pressure reactors. We thank Deryck Mills for assistance
with cryo-EM. We acknowledge the assistance given by the use
of the Manchester Protein Structure Facility as well as the DESY
(PETRAIII, Hamburg, Germany), Diamond Light Source (Didcot,
UK) (proposal number MX12788), Elettra (Trieste, Italy) and
ESRF (Grenoble, France) for provision of synchrotron-radiation
facilities and support during data collection.
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Keywords: Biocatalysis • Carboxylation • Catechols • prenylated
FMN
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10.1002/anie.201708091
Angewandte Chemie International Edition
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Biocatalytic CO2-fixation: 3,4-
dihydroxybenzoic acid
Stefan E. Payer,+ Stephen A. Marshall,+
Natalie Bärland, Xiang Sheng, Tamara
Reiter, Andela Dordic, Georg
Steinkellner, Christiane Wuensch,
Susann Kaltwasser, Karl Fisher,
Stephen E. J. Rigby, Peter Macheroux,
Janet Vonck, Karl Gruber, Kurt Faber,
Fahmi Himo, David Leys, Tea Pavkov-
Keller,* Silvia M. Glueck*
decarboxylases are shown to
catalyze the regioselective para-
carboxylation onto the aromatic
core of catechols under ambient
conditions. The enzymes depend on
the recently discovered prenylated
FMN cofactor for catalysis, which
is proposed to proceed via a
monocovalent quinoid-cofactor
intermediate.
Page No. – Page No.
Regioselective para-Carboxylation of
Catechols by a Prenylated Flavin
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