Regioselective Enzymatic Carboxylation of Bioactive (Poly)phenols

Article abstract of DOI:10.1002/adsc.201601046

In order to extend the applicability of the regioselective enzymatic carboxylation of phenols, the substrate scope of o-benzoic acid (de)carboxylases has been investigated towards complex molecules with an emphasis on flavouring agents and polyphenols possessing antioxidant properties. o-Hydroxycarboxylic acid products were obtained with perfect regioselectivity, in moderate to excellent yields. The applicability of this method was proven by the regioselective bio-carboxylation of resveratrol on a preparative scale with 95% yield. (Figure presented.).

Full text of DOI:10.1002/adsc.201601046

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DOI: 10.1002/adsc.201601046
Regioselective Enzymatic Carboxylation of Bioactive
(Poly)phenols
a
a
b
c
´
Katharina Plasch, Verena Resch, Julien Hitce, Jarosław Popłonski,
Kurt Faber,a,* and Silvia M. Gluecka,d,*
Department of Chemistry, Organic & Bioorganic Chemistry, University of Graz, Heinrichstrasse 28, A-8010 Graz, Austria
Fax : (+43)-(0)316-380-9840; e-mail: Kurt.Faber@Uni-Graz.at or Si.Glueck@Uni-Graz.at
LꢀOrꢁal Research & Innovation, 30 bis rue Maurice Berteaux, 95500 Le Thillay, France
Department of Chemistry, Wrocław University of Environmental and Life Sciences, ul. C. K. Norwida 25, 50-375
Wrocław, Poland
a
b
c
d
Austrian Centre of Industrial Biotechnology (ACIB) c/o University of Graz, Heinrichstrasse 28, A-8010 Graz, Austria
Received: September 20, 2016; Revised: November 21, 2016; Published online: && &&, 0000
Supporting information for this article is available on the WWW under http://dx.doi.org/10.1002/adsc.201601046.
Abstract: In order to extend the applicability of the
regioselective enzymatic carboxylation of phenols,
the substrate scope of o-benzoic acid (de)carboxy-
lases has been investigated towards complex mole-
cules with an emphasis on flavouring agents and
polyphenols possessing antioxidant properties. o-
Hydroxycarboxylic acid products were obtained
with perfect regioselectivity, in moderate to excel-
lent yields. The applicability of this method was
proven by the regioselective bio-carboxylation of
resveratrol on a preparative scale with 95% yield.
To date, a biocatalytic toolbox for the regioselective
ortho-,[1,2,8] para-[9] and b-carboxylation[10] of phenols
and hydroxystyrenes, respectively, has been estab-
lished, which employs decarboxylases acting in the
(reverse) carboxylation direction using bicarbonate as
CO2 source. In addition, electron-rich heteroaromat-
ics, such as pyrrole and indole were successfully car-
boxylated.[11] Enzymatic carboxylation of phenolic
substances is a common detoxification pathway in an
anaerobic environment[,12] and hence the correspond-
ing enzymes are expected to possess a relaxed sub-
strate tolerance, which is indeed true for the o-car-
boxylation of phenols[5,8f] and the b-carboxylation of
hydroxystyrenes.[10] However, the substrates reported
so far are predominantly small or medium-sized
phenol derivatives, which were converted at low sub-
strate loading, except for a recently reported study on
hydroxystilbenes and the naturally occurring polyphe-
nol resveratrol.[8g]
Keywords: ortho-benzoic acid decarboxylases; bio-
active (poly)phenols; biocatalysis; carboxylation;
regioselectivity
The carboxylation of (hetero)aromatic and phenolic
Based on our previous studies on the enzymatic
compounds is a convenient method to obtain aromat- ortho-carboxylation of phenols, we aimed to expand
ic carboxylic acids used as pharmaceuticals[1,2] (e.g., the scope of this method towards more complex
salicylic and m-aminosalicylic acid) as well as building (poly)phenolic substrates (Scheme 1, Figure 1). The
blocks for organic synthesis. The traditional chemical latter compounds are well known for their biological
(Kolbe–Schmitt) carboxylation process performed on activities such as antioxidant, anti-inflammatory and
an industrial scale requires high pressure and temper- antimicrobial properties, which promote them as
ature (~90 bar, 120–3008C) and often suffers from in- promising targets for the pharmaceutical industry as
complete regioselectivities resulting in product mix- well as for the cosmetic and food fields.[13,14,15] Enzy-
tures.[3] In order to circumvent these limitations, vari- matic carboxylation of these compounds would pro-
ous chemical CO2 fixation concepts using heterogene-
ous, (transition) metal and organic catalysts have
been established.[4] A novel attractive biocatalytic al-
ternative is the use of (de)carboxylases, which act at
ambient reaction conditions and show perfect regiose-
lectivities.[5,6,7]
Scheme 1. Enzymatic ortho-carboxylation of bioactive
(poly)ACTHNUTRGENUGpN henols.
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strates is the natural product resveratrol (7a) which
has fostered research and development towards cos-
metic, food, nutraceutical and pharmaceutical applica-
tions, due to its unique ability to modulate physiologi-
cal as well as pathological pathways.[13,19] In particular,
resveratrol (7a) exhibits antimicrobial[13c] and antioxi-
dant[13d] properties that are expected to be retained in
the more soluble carboxylated form 7b. Furthermore,
the g-resorcylic acid moiety found in carboxylation
products 7b–9b would bring about new biological ac-
tivities that are not displayed by the corresponding
parent compounds 7a–9a. For instance, g-resorcylic
acid derivatives bearing a lipophilic substituent are
known as thrombolytic[20] and as anti-inflammatory
agents through uncoupling of phosphorylation.[21] In
particular, g-resorcylic acids with an alkylaryl sub-
stituent in the 4-position were shown to reduce in-
flammation in vivo in a mice model.[22]
In order to examine the synthetic potential of the
method, the most promising substrate candidates
were subjected to preparative-scale carboxylation.
A set of suitable enzyme candidates which have
been shown to regioselectively catalyze the o-carbox-
ylation of phenolic compounds was applied: (i) 2,3-di-
hydroxybenzoic acid decarboxylase from Aspergillus
oryzae (2,3-DHBD_Ao),[8f,23] (ii) 2,6-dihydroxybenzoic
acid decarboxylases from Rhizobium sp. (2,6-
DHBD_Rs)[8f,24] and (iii) salicylic acid decarboxylase
from Trichosporon moniliiforme (SAD_Tm).[1,2,8e,f] To
simplify handling by avoiding protein purification, the
biocatalysts were employed as lyophilized whole-cell
preparations of recombinant overexpressed decarbox-
ylases in E. coli. Independent control experiments en-
Figure 1. Set of substrates (1a–9a) and corresponding car-
boxylated products (1b–9b and 9c) obtained from bio-car-
boxylation.
vide an efficient access to more polar derivatives with sured that the empty E. coli host was devoid of com-
enhanced water solubility thereby facilitating their peting carboxylase activities.
formulation and modulating their bioavailability. In
addition, the soft electron-withdrawing effect of the ed benzaldehydes, such as vanillin (1a) and p-
newly introduced carboxylate group should render hydroxybenzaldehyde (2a) were regioselectively car-
The results are summarized in Table 1. Hydroxylat-
AHCTUNGTRENNUNG
these products more stable towards autoxidation in boxylated by 2,3-DHBD_Ao with 40% and 62% con-
analogy to the beneficial effect of the eÀ-withdrawing version, respectively, whereas SAD_Tm was active
carbonyl group of green tea polyphenols.[16] Further- only on vanillin with 33% conversion (entries 1 and
more, phenolic acids are expected to impede light-in- 2). These activities were unexpected, because carbox-
duced degradation as demonstrated in the case of ylation represents an electrophilic aromatic substitu-
photolabile substances in food or feed, such as vita- tion, which is impeded by electron-withdrawing sub-
mins[17a] or nucleic acids.[17b]
stituents, such as an aldehyde.[8f] For instance, o- and
The substrate scope of this study ranges from the p-nitrophenol, o-hydroxybenzaldehyde (see the sup-
flavouring agent vanillin (1a) to secondary plant me- porting information of ref.[8f]) as well as isovanillin
tabolites, such as p-hydroxybenzaldehyde (2a), esters (10a, Supporting Information, Table S2, entry 1) were
of p-hydroxycinnamic (p-coumaric) (3a) and m-hy- unreactive with these enzymes.
droxyphenylacetic acid (4a), as well as representatives
In order to convert substrates carrying a free car-
of flavonoids such as dihydrochalcones (5a, 6a) and boxylic acid group, where decarboxylation would be
hydroxystilbenes, such as resveratrol (7a) and resvera- favoured, the corresponding methyl esters were ap-
trol-like polyphenols (8a, 9a). Phloretin (6a) which is plied as masking groups. This strategy proved to be
abundantly present in apples is described to possess successful, as cinnamic and phenylacetic acid esters 3a
antioxidant properties and to act as a peroxynitrite and 4a were carboxylated by 2,3-DHBD_Ao and
scavenger and an inhibitor of lipid peroxidation.[13b,18] SAD_Tm with up to 66% conversion (entries 3 and
The most famous example among this group of sub- 4).
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Table 1. Regioselective enzymatic carboxylation of (poly)-
phenolic substrates.
complexity of the substrates containing two aromatic
moieties (entries 5–9).
Substrates 5a and 6a of the dihydrochalcone-type
family were regioselectively carboxylated at the most
electron-rich position[25] – even when it corresponds
to a sterically hindered position like in the phloroglu-
cinol moiety (6a, entry 6) – with moderate to excel-
lent conversions, regardless of the enzyme used (con-
version 57–97%, entries 5 and 6). The deactivating
(ÀM) effect of the electron-withdrawing carbonyl
group was compensated in the presence of three hy-
droxy groups exerting a +M effect (6a, entry 6),
whereas no carboxylation took place in presence of
a single (31a, 35a, Supporting Information, Table S2,
entries 22 and 26) or two (36a, Supporting Informa-
tion, Table S2, entry 27) OH moieties due to insuffi-
cient activation.
The well-known phytoalexin resveratrol (7a) and
derivatives (8a, 9a) were accepted as substrates by all
three enzymes (entries 7–9). Even though the steric
requirements of 7a and 8a are almost identical, the
conjugated system of 7a enabled additional electronic
activation towards carboxylation by the second aro-
matic moiety leading to considerably better conver-
sions (85–97%, respectively, entry 7) compared to the
non-conjugated (saturated) resveratrol derivative 8a
(45–77% conversion, entry 8). This is in line with the
results obtained with the well accepted oxyresveratrol
(9a), which underwent unique double carboxylation
on both resorcinol units, which are similarly electroni-
cally activated. Detailed investigation over time (see
the Supporting Information, Figure S7) revealed that
oxyresveratrol was first mono-carboxylated at the 4’
position to yield the mono-carboxylic acid 9b, which
was subsequently converted to diacid 9c with almost
full conversion by 2,6-DHBD_Rs and SAD_Tm (96%
and 97%, respectively, entry 9). Data from a recent
study[8g] showed that g-resorcylic acid decarboxylase
from Rhizobium radiobacter WU-0108 (95% se-
quence similarity to 2,6-DHBD_Rs, Supporting Infor-
mation, Figure S8) was also able to carboxylate re-
sveratrol (8a).
[a]
Reaction conditions: phosphate buffer (pH 8.5, 100 mM),
whole lyophilized cells of E. coli containing the corre-
sponding overexpressed enzyme (30 mgmLÀ1), substrate
(10 mM), KHCO3 (3M), 308C, 120 rpm, 24 h.
Conversions were determined by reversed-phase HPLC,
side products not detected (<2%).
Products 1b–9b and 9c were characterized by 1-D
(1H and 13C) and 2-D NMR (COSY, HSQC and
HMBC, see the Supporting Information) and HR-MS.
Various other classes of substrates (such as phenol
ether, heteroaromatics, coumarins, etc.), which were
not accepted by any of the three decarboxylases are
[b]
[c]
2,3-DHBD_Ao=2,3-dihydroxybenzoic acid decarboxy-
lase from Aspergillus oryzae, 2,6-DHBD_Rs=2,6-dihy-
droxybenzoic acid decarboxylase from Rhizobium sp.
and SAD_Tm=salicylic acid decarboxylase from Tricho-
sporon moniliiforme.
Conversion corresponds to the doubly carboxylated
product 9c.
summarized
(Table S2).
in
the
Supporting
Information
In order to demonstrate the applicability of biocat-
alytic carboxylation on a preparative scale, substrates
3a, 4a and 6a–9a were subjected to up-scaling experi-
ments after optimization of the reaction conditions re-
garding the following parameters: (i) concentration of
[d]
Surprisingly, all enzyme candidates showed signifi- co-substrate bicarbonate, (ii) biocatalyst loading, (iii)
cantly enhanced conversion by further expanding the addition of organic co-solvents and (iv) substrate con-
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centration using resveratrol (7a) as model substrate sorcylic acid decarboxylase from Rhizobium radio-
and 2,6-DHBD_Rs as biocatalyst (for details see the bacter WU-0108 furnished 7b with only 26% yield.[8g]
Supporting Information). Optimal parameters to
In conclusion, our study shows that o-benzoic acid
reach complete conversion (ꢀ97%) within 18–24 h decarboxylases are able to convert large (poly)phe-
were identified as 2M bicarbonate concentration, nolic natural products possessing antioxidant activity,
2 mgmLÀ1 lyophilized whole-cell biocatalyst and 20% such as phloretin and resveratrol, with up to quantita-
(v/v) of methanol as water-miscible organic co-sol- tive conversion after optimization of the reaction con-
vent. For cinnamic and phenylacetic acid esters (3a, ditions. On complex substrates possessing more than
4a) moderate yields were achieved (32% and 43%, one potential carboxylation site, the relative electron-
respectively, Table 2, entries 1 and 2) employing 2,3- density of the aromatic moieties determines the regio-
DHBD_Ao as biocatalyst which is in accordance with selectivity. Surprisingly, various electron-withdrawing
the results obtained from the screening (Table 1, en- functional groups, such as carboxylic esters and alde-
tries 3 and 4). In the case of the dihydrochalcone-type hydes, were tolerated rather well. Overall, this study
substrate (6a) the isolated yield of 6b was 67% illustrates the versatility of bio-carboxylation as
(entry 3) with 2,3-DHBD_Ao. For the resveratrol-like a highly regioselective and synthetically useful biocat-
substrates (7a–9a) 2,6-DHBD_Rs was used as biocat- alytic alternative to the Kolbe–Schmitt reaction.
alyst since the latter gave the best results in the
screening experiments (Table 1). Whereas for saturat-
ed resveratrol (8a) and oxyresveratrol (9a) isolated Experimental Section
yields were moderate (45% and 66%, respectively, en-
tries 5 and 6), resveratrol (7a) was carboxylated to 7b General
in 95% isolated yield (entry 4). For comparison, g-re-
Substrates 1a, 2a, 6a and 8a as well as reference material of
1b and 2b were purchased from Sigma Aldrich, while 7a was
obtained from Acros Chemicals. p-Coumaric acid for the
Table 2. Up-scaling of various substrates.[a]
synthesis of substrate 4a was provided by Roche, substrate
8a was prepared by catalytic hydrogenation of resveratrol as
previously described[26] and provided by LꢀOrꢁal R&I. For
the synthesis of substrates 3a–5a and reference material for
product 5b see the Supporting Information. TLCs were run
on silica plates (Merck, silica gel 60, F254), for column chro-
matography silica gel 60 ꢃ (Merck) was used, compounds
were visualized using UV (254 and 365 nm) and by spraying
with cerium ammonium molybdate [5 g (CeSO4)2, 25 g
(NH4)6Mo7O24·4H2O, 50 mL concentrated H2SO4, 450 mL
H2O]. 2,3-Dihydroxybenzoic acid decarboxylase from As-
pergillus oryzae (2,3-DHBD_Ao), 2,6-dihydroxybenzoic acid
decarboxylase from Rhizobium species (2,6-DHBD_Rs) and
salicylic acid decarboxylase from Trichosporon moniliiforme
(SAD_Tm) were cloned and overexpressed as previously
described.[27]
General Procedure for Biotransformations
Lyophilized whole cells (30 mg E. coli cells, containing the
corresponding overexpressed enzyme) were rehydrated in
phosphate buffer (900 mL, pH 5.5, 100 mM) for 30 min. The
substrate [10 mM final concentration, dissolved in 100 mL or-
ganic co-solvent (DMSO, MeOH and H2O/MeCN 50:50, re-
spectively)] was added to the enzyme solution (1 mL final
volume) which was transferred into a glass vial containing
KHCO3 (3M) to give a final pH of 8.5. The vials were tight-
ly sealed with screw caps and were shaken for 18 h at 308C
with 120 rpm. After 18 h the reaction was stopped by taking
[a]
Reaction conditions: phosphate buffer (pH 8.5, 100 mM,
19 mL), whole lyophilized cells of E. coli containing the
corresponding overexpressed enzyme [2,3-DHBD_Ao
(651 mg) for substrates 3a, 4a and 6a; 2,6-DHBD_Rs
(651 mg) for substrates 7a–9a], substrate (50 mg, 10–
11 mM depending on the substrate) dissolved in MeOH
(1 mL), KHCO3 (3M), 308C, 400 rpm, 24 h.
100 mL of the reaction mixture and diluting it in 900 mL of
H2O/MeCN/trifluoroacetic acid (TFA, 50:50:3) to precipi-
tate the enzyme, which was removed by centrifugation
(10 min, 14000 rpm). The resulting supernatant was directly
used for measurements on a reversed-phase HPLC system
using H2O/MeCN (0.1% TFA) as eluent (gradient: MeCN/
H2O 5%–100%) on an achiral C18 column (Phenomenex
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Luna, C18 100 ꢃ, 250ꢄ4.6 mm, 5 mm). Anisole was used as
an internal standard. All screening experiments were carried
out at least in triplicate.
a MeCN/acetic acid (2%) gradient (0–16 min 20–65%, 16–
22 min 65–20%). Retention times for all substrates (1a--9a)
and products (1b–9b and 9c) are listed in the Supporting In-
formation (Table S1).
NMR and HR-MS analysis: NMR spectra were recorded
with a Bruker AVANCE III 300 MHz spectrometer using
a 5 mm BBO probe at 300 K. Chemical shifts d are ex-
pressed in ppm, coupling constants J are given in Hz. HR-
MS analysis was performed on an Agilent 1260 Infinity
system coupled with an Agilent 6230 TOF LC/MS instru-
ment with an APCI-ionisation in positive mode.
Preparative-Scale Biotransformation of Resveratrol
(7a)
Lyophilized whole cells (651 mg E. coli cells, containing the
corresponding expressed enzyme) were rehydrated in phos-
phate buffer (19 mL, pH 5.5, 100 mM) for 30 min. Substrate
7a (50 mg in 1 mL MeOH) was added to the enzyme solu-
tion and this mixture was transferred into a glass reaction
vessel containing KHCO3 (6.5 g, 3M) to give a final pH of
8.5. The vessel was tightly closed with a rubber stopper
(fixed with a clamp to avoid loss of CO2) and was allowed
to react for 24 h at 308C with 400 rpm shaking. Overall,
150 mg of resveratrol were carboxylated in triplicate parallel
experiments with 50 mg substrate each, which were com-
bined before work-up. After 24 h the reaction was stopped
by slow addition of HCl (6M) until a pH of around 1–2 was
reached (caution: CO2 formation). The quenched reaction
mixture was saturated with NaCl and extracted with EtOAc
(4ꢄ, 10 mL) and the combined organic layers were dried
with Na2SO4. EtOAc was removed under reduced pressure
and remaining solids were purified using silica gel column
chromatography. Silica gel was deactivated with aqueous
ammonia solution, which was added to the solvent mixture
used for packing of the column [CH2Cl2/MeOH 90:10 with
2% v/v of aqueous ammonia solution (30%)]. After column
chromatography, solvents were removed under reduced
pressure and the residue (containing the ammonium salt of
7b) was redissolved in EtOAc (10 mL) containing HCl (6M,
1 mL) and water (10 mL). The acidified aqueous phase was
re-extracted with EtOAc (3ꢄ10 mL), the combined organic
phases were dried over Na2SO4 and the organic solvent was
removed under reduced pressure to give free acid 7b as
a yellow solid; yield: 169 mg (94%).
3a: 1H NMR (300 MHz, DMSO-d6): d=10.02 (s, 1H),
7.54–7.59 (m, 3H), 6.80 (s, 1H), 6.78 (s, 1H), 6.40 (d, J=
16.0 Hz, 1H), 3.69 (s, 3H);[28] 13C NMR (75 MHz, DMSO-
d6): d=167.1, 159.9, 144.8, 130.3, 125.1, 115.8, 113.9, 51.2.[29]
3b: 1H NMR (300 MHz, DMSO-d6): d=7.57 (d, J=
7.8 Hz, 1H), 7.53 (s, 1H), 6.81 (s, 1H), 6.78 (t, J=1.9 Hz,
1H), 6.39 (d, J=16.0 Hz, 1H), 3.69 (s, 3H); 13C NMR
(75 MHz, DMSO-d6): d=175.8, 167.1, 159.9, 144.8, 132.6,
130.4, 125.1, 115.8, 115.0, 113.9, 45.6.[30]
4a: 1H NMR (300 MHz, DMSO-d6): d=9.37 (bs, OH),
7.09 (t, J=7.09 Hz 1H), 6.66 (s, 2H), 6.63–6.66 (m, 1H),
3.60 (s, 3H), 3.56 (s, 2H);[31] 13C NMR (75 MHz, DMSO-d6):
d=171.6, 157.3, 135.6, 129.3, 119.9, 116.2, 113.8, 51.7, 40.2.[32]
4b: 1H NMR (300 MHz, DMSO-d6): d=7.51 (d, J=
7.7 Hz, 1H), 6.52 (s, 1H), 6.48 (dd, J=7.9, 1.2 Hz, 1H), 3.19
(s, 3H), (CH2 signal obscured by water peak 3.43 ppm);
13C NMR (75 MHz, DMSO-d6): d=172.1, 163.3, 162.3,
142.1, 129.3, 117.7, 117.5, 116.5, 67.3, 45.1.[33]
5a: 1H NMR (300 MHz, CDCl3) d=7.92–7.99 (m, 2H),
7.53–7.59 (m, 1H), 7.41–7.49 (m, 2H), 7.16 (t, J=7.8 Hz,
1H), 6.81 (d, J=7.6 Hz, 1H), 6.74–6.77 (m, 1H), 6.70 (dd,
J=8.0, 2.0 Hz, 1H), 3.30 (dd, J=8.4, 6.9 Hz, 2H), 3.02 (t,
J=7.7 Hz, 2H);[34] 13C NMR (75 MHz, CDCl3) d=199.9,
156.0, 143.2, 136.8, 133.4, 129.9, 128.8, 128.2, 120.8, 115.6,
113.3, 40.4, 30.1.
5b: 1H NMR (300 MHz, CDCl3) d=7.99–7.94 (m, 2H),
7.82 (d, J=8.1 Hz, 1H), 7.58 (ddd, J=7.4, 3.9, 1.3 Hz, 1H),
7.50–7.43 (m, 2H), 6.87 (s, 1H), 6.82 (dd, J=8.0, 1.2 Hz,
1H), 3.32 (t, J=7.5 Hz, 2H), 3.07 (t, J=7.6 Hz, 2H);
13C NMR (75 MHz, CDCl3) d=204.3, 168.4, 162.4, 145.3,
141.1, 133.4, 131.1, 128.8, 128.2, 120.1, 117.1, 117.0, 112.8,
30.3, 22.8. HR-MS (ESI+): m/z=269.081932, calculated for
C16H14O4 [M+H]+: 269.081739.
Preparative-scale experiments with 3a, 4a, 6a, 8a and 9a
were performed analogously with 50 mg substrate each.
Analytics
HPLC analysis: HPLC/UV experiments were performed on
an HPLC Agilent 1260 Infinity system with a diode array
detector and a reversed phase Phenomenex Luna column
C18 (100 ꢃ, 250ꢄ4.6 mm, 5 mm, column temperature 248C).
Conversions were determined by comparison with calibra-
tion curves for products and substrates prepared with au-
thentic reference material. All compounds were spectropho-
tometrically detected at 254, 280 and 310 nm, respectively.
Method A was run over 22 min with H2O/TFA (0.1%) as
the mobile phase (flow rate 1 mLminÀ1) and a MeCN/TFA
(0.1%) gradient (0–2 min 5%, 2–15 min 5–100%, 15–17 min
100%, 17–22 min 100–5%). Method B was run over 27 min
with H2O/TFA (0.1%) as the mobile phase (flow rate
1 mLminÀ1) and a MeCN/TFA (0.1%) gradient (0–5 min
5%, 5–20 min 5–100%, 20–22 100%, 22–27 min 100–5%).
Method C was run over 27 min with H2O/TFA (0.1%) as the
mobile phase (flow rate 1 mLminÀ1) and a MeCN/TFA
(0.1%) gradient (0–2 min 5%, 2–15 min 5–60%, 15–21.5 min
60%, 21.5–23.5 min 60–100%, 23.5–27 min 100–5%).
Method D was run over 22 min with MeOH/acetic acid
(2%) as the mobile phase (flow rate 1 mLminÀ1) and
1
6b: H NMR (300 MHz, DMSO-d6): d=7.0 (d, J=8.4 Hz,
2H), 6.7 (d, J=8.4 Hz, 2H), 5.5 (s, 1H), 3.3 (s, 2H), 2.73–
2.78 (m, 2H); 13C NMR (75 MHz, DMSO-d6): d=204.1,
170.3, 169.5, 168.1, 155.3, 131.7, 129.2, 115.1, 103.1, 96.0,
92.6, 45.0, 29.4; HR-MS (ESI+): m/z=317.066416, calculated
for C16H14O7 [M+H]+: 317.066676.
7b: 1H NMR (300 MHz, DMSO-d6): d=7.45 (d, J=
8.6 Hz, 2H), 7.19 (d, J=16.4 Hz, 1H), 6.87 (d, J=16.4 Hz,
1H), 6.78 (d, J=8.6 Hz, 2H), 6.55 (s, 2H), broad signal
from OH-moieties overlapping with signals from 7.45–6.87);
13C NMR (75 MHz, DMSO-d6): d=172.9, 161.4, 158.3,
144.3, 131.9, 129.0, 128.0, 124.6, 116.0, 104.9, 100.8; HR-MS
(ESI+): m/z=271.061226, calculated for C15H12O5 [M+H]+:
271.061197.
8b: 1H NMR (300 MHz, DMSO-d6): d=6.99 (d, J=
8.5 Hz, 2H), 6.65 (d, J=8.4 Hz, 2H), 6.23 (s, 2H), 2.69–2.71
(m, 4H); 13C NMR (75 MHz, DMSO-d6): d=172.8, 161.0,
155.8, 150.3, 131.7, 129.6, 115.4, 107.6, 100.0, 38.8, 35.7; HR-
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MS (ESI+): m/z=275.09158, calculated for C15H14O5 [M+
H]+: 275.09140.
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9b: 1H NMR (300 MHz, DMSO-d6): d=7.37 (d, J=
8.5 Hz, 1H), 6.87 (d, J=8.9 Hz, 1H), 6.80 (t, J=8.2 Hz,
1H), 6.41 (s, 2H), 6.33 (d, J=2.4 Hz, 1H), 6.27–6.23 (m,
1H). 13C NMR (75 MHz, DMSO-d6): d=172.9, 161.2, 160.8,
158.8, 145.9, 129.7, 126.4, 125.5, 115.0, 107.4, 104.0, 103.8,
103.7, 102.6.
9c: 1H NMR (300 MHz, DMSO-d6): d=7.58 (d, J=
8.7 Hz, 1H), 7.35 (d, J=16.5 Hz, 1H), 6.95 (d, J=16.5 Hz,
1H), 6.51 (s, 2H), 6.30 (t, J=9.2 Hz, 1H); 13C NMR
(75 MHz, DMSO-d6): d=174.5, 172.5, 162.5, 161.2, 160.8,
145.3, 132.0, 126.9, 124.1, 114.6, 107.3, 104.7, 103.0, 100.3;
HR-MS (ESI+): m/z=333.060595, calculated for C16H14O8
[M+H]+: 333.060494.
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Acknowledgements
Funding by the Austrian Science Fund (FWF, project I 1637-
N19) is gratefully acknowledged. This work has been sup-
ported by the Austrian BMWFJ, BMVIT, SFG, Standortagen-
tur Tirol and ZIT through the Austrian FFG-COMET-Fund-
ing Program. The authors would like to thank LꢀOrꢁal for
a financial contribution. Klaus Zangger and his team and
Martin Mittelbach and co-workers (Graz) are cordially
thanked for their support in NMR and HR-MS analysis, re-
ˇ
spectively. Vladimir Kren (Prague) is thanked for providing
substrates.
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