Enantioselective Synthesis of Dilignol Model Compounds and Their Stereodiscrimination Study with a Dye-Decolorizing Peroxidase

Article abstract of DOI:10.1021/acs.orglett.7b00587

A four-step enantioselective approach was developed to synthesize anti (1R,2S)-1a and (1S,2R)-1b containing a β-O-4 linkage in good yields. A significant difference was observed for the apparent binding affinities of four stereospecific lignin model compounds with TcDyP by surface plasmon resonance, which was not translated into a significant difference in enzyme activities. The discrepancy may be attributed to the conformational change involving a loop widely present in DyPs upon H₂O₂ binding.

Full text of DOI:10.1021/acs.orglett.7b00587

Letter
pubs.acs.org/OrgLett
Enantioselective Synthesis of Dilignol Model Compounds and Their
Stereodiscrimination Study with a Dye-Decolorizing Peroxidase
Gaochao Huang,† Ruben Shrestha,† Kaimin Jia,† Brian V. Geisbrecht,‡ and Ping Li*,†
†Department of Chemistry and ‡Department of Biochemistry and Molecular Biophysics, Kansas State University, Manhattan, Kansas
66506, United States
S
* Supporting Information
ABSTRACT: A four-step enantioselective approach was
developed to synthesize anti (1R,2S)-1a and (1S,2R)-1b
containing a β-O-4 linkage in good yields. A significant
difference was observed for the apparent binding affinities of
four stereospecific lignin model compounds with TcDyP by
surface plasmon resonance, which was not translated into a
significant difference in enzyme activities. The discrepancy may be attributed to the conformational change involving a loop
widely present in DyPs upon H2O2 binding.
curvata using surface plasmon resonance (SPR) and enzyme
ye-decolorizing peroxidases (DyPs) are a new family of
Dheme peroxidases that catalyze H O -dependent oxida- activity assays.
2
2
tion of anthraquinone-based dyes under acidic conditions.1−3
They have been reported to degrade phenolic lignin model
compounds and wheat straw lignocellulose, though the
efficiency is low.4−7 Our interest in DyPs stems from their
potential applications in lignin degradation,8,9 a critical step in
converting lignocellulose biomass to biofuels.8,10−13 Lignin
biosynthesis produces a β-O-4 interunit linkage up to 50−65%
with α and β carbons exhibiting (R,R), (R,S), (S,R), and (S,S)
stereochemistry.11,14 While the lignin polymer is a racemic
mixture,15,16 studies have suggested that the localized stereo-
chemistry affects its enzymatic digestion.17−23 However,
stereochemical relationships between β-O-4 dilignols and
DyPs remained unknown due to limited access to these
stereospecific compounds in sufficient amounts.
While many endeavors had been pursued to synthesize
stereospecific lignin model compounds containing a β-O-4
linkage,21,24−31 significant progress was not made until recently
by Njiojob et al., who had successfully prepared the syn (1S,2S)-
dilignol using asymmetric epoxidation followed by kinetic
resolution.32 However, attempts to scale up this nine-step
reaction resulted in a low yield.33 Later, the same syn
compound was obtained by a five-step synthesis similar to
the method B (Scheme 1) described in this report.33 However,
they had to employ a Mitsunobu reaction to produce anti
(1R,2S)-dilignol,33 which resulted in an additional three steps
and is disadvantageous for reaction scale-up. Thus, we decided
to develop a short approach to synthesize anti dilignols
containing a β-O-4 linkage. Here, a four-step enantioselective
approach is reported to prepare anti (1R,2S)-1a and (1S,2R)-1b
(Scheme 1), achieving the goal of obtaining all four
stereoisomers in gram scales by controlling the conditions of
enolate formation in the presence of Evans chiral auxiliaries.
These model compounds were studied for their stereo-
discrimination by the A-class TcDyP from Thermomonspora
As shown in Scheme 1, guaiacol was converted to acid 2 and
then reacted with oxazolidinone/thiazolidinethione 3 to yield
adduct 4 (Figure 1),28,34 which was screened for its efficiency in
diastereoselectivity by HPLC. A representative HPLC profile is
shown in Figure 2, and the results are summarized in Table 1.
For method A involving trimethylsilyl chloride (TMSCl) and
catalytic MgCl2,35,36 4a (X = O, R = R-Ph, R′ = H) was found
to be the best for diastereoselectivity to yield an anti-aldol
product. When 4a was reacted with vanillin, di-TMS-protected
aldol product was formed initially (see the NMR spectra in the
SI). However, the phenolic TMS was not stable and could be
easily cleaved to produce mono-TMS-protected 5a (X = O, R =
R-Ph, R′ = H). It has to be noted that isolation of the major
diastereomer in 5 by silica gel chromatography was difficult.
Therefore, without purification, 5 was directly treated with
trifluoroacetic acid (TFA) to afford compound 6, in which the
major diastereomer with anti-configuration was isolated in 75%
yields. To further optimize diastereoselectivity, effects of
organic bases and solvents were also screened. The results
are summarized in Table 1. It was concluded that Et3N and
CH2Cl2 were the best base and solvent, respectively. It was also
found that low temperature slightly improved diastereoselec-
tivity (entries 1 and 2 in Table 1).
Reductive cleavage of the auxiliary in compound 6 with
NaBH4 gave the desired stereoisomer (1R,2S)-1a or (1S,2R)-1b
depending on the configuration of phenyl group. The
enantiomeric excess (ee) of the final compounds was
determined to be >99% using chiral HPLC. Assignment of
the configurations in each stereoisomer was based on the
reported optical rotations, which were also consistent with
theoretical predictions.35,37 These results are summarized in
Received: February 26, 2017
© XXXX American Chemical Society
A
DOI: 10.1021/acs.orglett.7b00587
Org. Lett. XXXX, XXX, XXX−XXX

Organic Letters
Letter
a
Scheme 1. Enantioselective Synthesis of β-O-4 Dilignols
Figure 2. Representative HPLC profile of diastereoselectivity using
method A. The straight line indicates MeOH gradient. The peak at 4.8
min represents unreacted aldehyde.
a
Table 1. Optimization of Aldol Reactions Using Method A
b
entry
compd
base
Et3N
solvent
CH2Cl2
diastereoselectivity
1
4a
4a
4c
4e
4f
78:7:6:9 (rt)
80:12:4:4
2
Et3N
CH2Cl2
CH2Cl2
CH2Cl2
CH2Cl2
CH2Cl2
CH2Cl2
CH2Cl2
CH2Cl2
CH2Cl2
CH2Cl2
CH2Cl2
ClCH2CH2Cl
EtOAc
3
Et3N
78:4:16:2 (rt)
72:9:10:9 (rt)
4
Et3N
c
5
Et3N
57:27:16 (rt)
c
6
4g
4a
4a
4a
4a
4a
4a
4a
4a
4a
4a
4a
4a
Et3N
31:16:53 (rt)
7
BuNH2
PhNH2
i-Pr2NH
trace product
no product
73:15:4:8
8
9
d
10
11
12
13
14
15
16
17
18
DIPEA
no product
no product
no product
72:16:7:5
pyridine
e
DBU
Et3N
Et3N
Et3N
Et3N
Et3N
Et3N
76:7:8:9
THF
71:2:7:20
Et2O
64:6:14:16
57:6:6:31
toluene
CH3CN
a
Compounds 5 and 8 were used without purification.
no product
a
b
c
Unless indicated, all reactions were carried out at 0 °C.
Diastereoselectivity was determined by HPLC. See the SI for details.
Diastereomers were inseparable under these HPLC conditions.
d
e
Diisopropylethylamine. 1,8-Diazabicyclo[5.4.0]undec-7-ene.
Figure S1 and Table S1. Thus, the stereospecific anti-dilignols
were obtained in just four steps with an overall 42% yield,
which was a significant improvement over the other asymmetric
methods reported so far.32,33
Method B employing n-Bu2BOTf and DIPEA to form
enolates has been recently reported to synthesize syn-aldol 7e
(X = O, R = R-iPr, R′ = H) using chiral auxiliary 4e.33 However,
our screen on auxiliaries found that 4c had a slightly better
diastereoselectivity than 4e (see Figure S2 and Table S2). Thus,
4c was selected as the chiral auxiliary for subsequent
preparations. After the chiral auxiliary in 7 was cleaved by
NaBH4 to give 8, a subsequent hydrogenesis yielded (1R,2R)-
Figure 1. Structures of 4a−g containing Evans auxiliaries.
B
DOI: 10.1021/acs.orglett.7b00587
Org. Lett. XXXX, XXX, XXX−XXX

Organic Letters
Letter
1c or (1S,2S)-1d depending on the configuration of the benzyl
(Bn) group in the auxiliary.
summarized in Table 2, which are in the order of 1a > 1d > 1b
> 1c, consistent with the order of apparent binding affinities
(KD) obtained by SPR. However, one important difference was
found between SPR and enzyme activity assays. It was observed
that 1a bound TcDyP by 14- and 12-fold better than 1b and 1d,
respectively. Stereoisomer 1c displayed very weak binding with
TcDyP. Therefore, the significant difference observed in SPR
experiments was not reflected in enzyme activities toward these
stereoisomers, in which only 1.6-fold difference was observed
for the ones with the highest (1a) and lowest (1c) activities.
We recently determined the crystal structure of TcDyP,40
which revealed that a large loop consisting of residues 277−310
is present next to an access channel leading to the propionate
group on pyrrole ring C in the heme active site (Figure 4). The
Surface plasmon resonance (SPR) is a highly sensitive
technique to study noncovalent interactions of biomolecules,
especially protein−protein and protein−small molecule inter-
actions in real time.38 Purified TcDyP (Figure S3) was
immobilized on a CMD500 M sensor chip (Xantec Bioanalytics
GmbH) via amine-coupling chemistry.39 The synthesized
stereoisomers 1a−d at a series of concentrations were then
injected, and sensorgrams are shown in Figure S4. Steady-state
affinity analysis based on a 1:1 binding model (Figure S5) was
used to determine the apparent binding affinity (KD). The
results and parameters evaluating global fittings of the data are
summarized in Table 2. It was revealed that all stereoisomers
Table 2. SPR Analysis and Enzyme Assays with TcDyP
parameters
KD (mM)
1a (1R,2S) 1b (1S,2R) 1c (1R,2R) 1d (1S,2S)
c
SPR
0.166
35.6
1.51
232
2.37
163
ND
2.06
68.8
0.575
215
a
RUmax
Chi2 (RU2)
SA (mU/mg)
b
0.280
161
142
a
RUmax corresponds to the unconstrained value for the fitted data at
b
saturation, for which the theoretical values are 105 for TcDyP. The
fitting of the observed data to theoretical model is evaluated by Chi2.
c
ND: not determined.
had weak interactions with TcDyP. For stereoisomer 1c, its
affinity was too weak to be determined. Among the other three
stereoisomers, 1a appeared to bind the strongest with the
enzyme at submillimolar concentrations. The KD values of 1b
and 1d are at least 10-fold higher than that of 1a, suggesting
that stereochemistry at Cα and Cβ does affect the substrate
binding in the absence of H2O2. Such observations may shed
light on catalysis by DyPs.
We have previously reported that racemic mixtures of β-O-4
dilignol 1 can be degraded by TcDyP in the presence of H2O2.6
Based on the MS results, Cα−Cβ was proposed to be cleaved
during degradation.6 To investigate the stereochemical effects
of TcDyP, the four stereoisomers were individually incubated
with the enzyme in the presence of H2O2. Reaction rates were
determined on the basis of the disappearance of the peak at
13.2 min corresponding to the substrate (Figure 3). Specific
activities (SAs) of TcDyP toward the four stereoisomers are
Figure 4. Crystal structure of TcDyP (PDB 5JXU) in surface (left) and
cartoon (right) representations. The loop consisting of residues 277−
310 is colored in pink. The heme b, catalytic residues (D220, H312
and R327), W263, and L279 are shown in red, green, blue, and pink
sticks.
entrance of this channel in D-class DyP from Bjerkandera
adusta has been shown to be a substrate-binding site.41 The
same propionate group has also been demonstrated to provide
a direct electron transfer from the substrate to porphyrin radical
in heme-containing manganese and ascorbate peroxidases.42,43
Additionally, this 34-residue loop is close to W263, which has
been characterized as a surface-exposed substrate oxidation site
in TcDyP.40 The closest distance between the loop (L279) and
W263 is 6.1 Å. Moreover, this loop is widely present in DyPs,
although the size varies between classes. Deletion of the loop in
TcDyP has resulted in the loss of enzyme activity (data not
shown). Thus, it is highly possible that DyP enzymes undergo a
conformational change involving this large loop upon H2O2
binding. Such H2O2-induced conformational changes have been
reported for a number of other heme peroxidases including
cytochrome c peroxidase and horseradish peroxidase.44−48
Since our SPR experiments were carried out in the absence of
H2O2, the significant difference in KD with the four stereo-
isomers may not be translated into a significant difference in
enzyme activities toward them, though the overall order of
stereochemical preference is retained in both SPR and activity
assay experiments.
In summary, a 4-step enantioselective method has been
developed to prepare anti (1R,2S)-1a and (1S,2R)-1b in the
presence of TMSCl and catalytic MgCl2, achieving the goal of
synthesizing all four stereospecific β-O-4 dilignols on gram
scales by controlling the conditions of enolate formation. The
TcDyP displayed different stereodiscrimination against these
stereoisomers in the absence and presence of H2O2, which may
be attributed to the conformational change involving a loop
Figure 3. HPLC profiles of TcDyP incubating with 1a and H2O2 (276
nm). Peaks at 3.1, 12.0, 13.2, and 17.4−20.5 min correspond to
impurity from citrate buffer, internal standard, substrate 1a, and
degradation products, respectively. The inset shows the rate of 1a
degradation.
C
DOI: 10.1021/acs.orglett.7b00587
Org. Lett. XXXX, XXX, XXX−XXX

Organic Letters
Letter
(18) Gall, D. L.; Ralph, J.; Donohue, T. J.; Noguera, D. R. Environ.
Sci. Technol. 2014, 48, 12454−12463.
(19) Masai, E.; Ichimura, A.; Sato, Y.; Miyauchi, K.; Katayama, Y.;
Fukuda, M. J. Bacteriol. 2003, 185, 1768−1775.
(20) Katayama, T.; Tsutsui, J.; Tsueda, K.; Miki, T.; Yamada, Y.;
Sogo, M. J. Wood Sci. 2000, 46, 458−465.
widely present in DyP enzymes upon H2O2 binding. The
overall weak stereodiscrimination by DyPs may appear
advantageous in their applications as a biocatalyst for lignin
degradation, as they will not discriminate the source of biomass
that may have different localized stereochemistry.
(21) Hishiyama, S.; Otsuka, Y.; Nakamura, M.; Ohara, S.; Kajita, S.;
Masai, E.; Katayama, Y. Tetrahedron Lett. 2012, 53, 842−845.
(22) Sato, Y.; Moriuchi, H.; Hishiyama, S.; Otsuka, Y.; Oshima, K.;
Kasai, D.; Nakamura, M.; Ohara, S.; Katayama, Y.; Fukuda, M.; Masai,
E. Appl. Environ. Microbiol. 2009, 75, 5195−5201.
(23) Jonsson, L.; Karlsson, O.; Lundquist, K.; Nyman, P. O. FEBS
Lett. 1990, 276, 45−8.
ASSOCIATED CONTENT
■
S
* Supporting Information
The Supporting Information is available free of charge on the
ACS Publications website at DOI: 10.1021/acs.or-
glett.7b00587.
(24) Yue, F. X.; Lu, F. C.; Sun, R. C.; Ralph, J. Chem. - Eur. J. 2012,
18, 16402−16410.
Detailed experimental procedures and spectroscopic data
(PDF)
(25) Kawai, S.; Jensen, K. A.; Bao, W.; Hammel, K. E. Appl. Environ.
Microbiol. 1995, 61, 3407−3414.
(26) Forsythe, W. G.; Garrett, M. D.; Hardacre, C.; Nieuwenhuyzen,
M.; Sheldrake, G. N. Green Chem. 2013, 15, 3031−3038.
(27) Kishimoto, T.; Uraki, Y.; Ubukata, M. Org. Biomol. Chem. 2005,
3, 1067−1073.
AUTHOR INFORMATION
■
Corresponding Author
*E-mail: pli@ksu.edu.
ORCID
(28) Buendia, J.; Mottweiler, J.; Bolm, C. Chem. - Eur. J. 2011, 17,
13877−13882.
(29) Li, K. C.; Helm, R. F. Holzforschung 2000, 54, 597−603.
(30) Li, K. C.; Helm, R. F. J. Chem. Soc., Perkin Trans. 1 1996, 2425−
2426.
(31) Nakatsubo, F.; Sato, K.; Higuchi, T. Holzforschung 1975, 29,
165−168.
(32) Njiojob, C. N.; Rhinehart, J. L.; Bozell, J. J.; Long, B. K. J. Org.
Chem. 2015, 80, 1771−1780.
Ping Li: 0000-0003-3671-0269
Notes
The authors declare no competing financial interest.
ACKNOWLEDGMENTS
■
This work was supported by awards to P.L. from the pilot
project of NIH P30GM110761 and Johnson Cancer Research
Center. G.H. is supported by an NIH K-INBRE postdoctoral
fellowship under grant number P20GM103418.
(33) Njiojob, C. N.; Bozell, J. J.; Long, B. K.; Elder, T.; Key, R. E.;
Hartwig, W. T. Chem. - Eur. J. 2016, 22, 12506−12517.
(34) Prashad, M.; Kim, H. Y.; Har, D.; Repic, O.; Blacklock, T. J.
Tetrahedron Lett. 1998, 39, 9369−9372.
(35) Evans, D. A.; Tedrow, J. S.; Shaw, J. T.; Downey, C. W. J. Am.
Chem. Soc. 2002, 124, 392−393.
REFERENCES
■
(36) Evans, D. A.; Downey, C. W.; Shaw, J. T.; Tedrow, J. S. Org.
Lett. 2002, 4, 1127−1130.
(1) Strittmatter, E.; Plattner, D. A.; Piontek, K. Dye-Decolorizing
Peroxidase (DyP). In Encyclopedia of Inorganic and Bioinorganic
Chemistry; John Wiley and Sons: New York, 2011.
(2) Sugano, Y. Cell. Mol. Life Sci. 2009, 66, 1387−1403.
(3) Colpa, D. I.; Fraaije, M. W.; van Bloois, E. J. Ind. Microbiol.
Biotechnol. 2014, 41, 1−7.
(4) Ahmad, M.; Roberts, J. N.; Hardiman, E. M.; Singh, R.; Eltis, L.
D.; Bugg, T. D. H. Biochemistry 2011, 50, 5096−5107.
(5) Brown, M. E.; Barros, T.; Chang, M. C. Y. ACS Chem. Biol. 2012,
7, 2074−2081.
(6) Chen, C.; Shrestha, R.; Jia, K.; Gao, P. F.; Geisbrecht, B. V.;
Bossmann, S. H.; Shi, J. S.; Li, P. J. Biol. Chem. 2015, 290, 23447−
23463.
(37) Evans, D. A.; Rieger, D. L.; Bilodeau, M. T.; Urpi, F. J. Am.
Chem. Soc. 1991, 113, 1047−1049.
(38) Nguyen, H. H.; Park, J.; Kang, S.; Kim, M. Sensors 2015, 15,
10481−10510.
(39) Johnsson, B.; Lofas, S.; Lindquist, G. Anal. Biochem. 1991, 198,
268−277.
(40) Shrestha, R.; Chen, X. J.; Ramyar, K. X.; Hayati, Z.; Carlson, E.
A.; Bossmann, S. H.; Song, L. K.; Geisbrecht, B. V.; Li, P. ACS Catal.
2016, 6, 8036−8047.
(41) Yoshida, T.; Tsuge, H.; Hisabori, T.; Sugano, Y. FEBS Lett.
2012, 586, 4351−4356.
(42) Sharp, K. H.; Mewies, M.; Moody, P. C. E.; Raven, E. L. Nat.
Struct. Biol. 2003, 10, 303−307.
(7) Min, K.; Gong, G.; Woo, H. M.; Kim, Y.; Um, Y. Sci. Rep. 2015, 5,
8245.
(43) Sundaramoorthy, M.; Kishi, K.; Gold, M. H.; Poulos, T. L. J.
Biol. Chem. 1994, 269, 32759−32767.
(8) Brown, M. E.; Chang, M. C. Y. Curr. Opin. Chem. Biol. 2014, 19,
1−7.
(44) Miyazaki, C.; Takahashi, H. FEBS Lett. 2001, 509, 111−114.
(45) Wang, X.; Wang, L. K.; Wang, X.; Sun, F.; Wang, C. C. Biochem.
J. 2012, 441, 113−118.
(9) de Gonzalo, G.; Colpa, D. I.; Habib, M. H. M.; Fraaije, M. W. J.
Biotechnol. 2016, 236, 110−119.
(10) Paliwal, R.; Rawat, A. P.; Rawat, M.; Rai, J. P. N. Appl. Biochem.
Biotechnol. 2012, 167, 1865−1889.
(46) Hassler, K.; Rigler, P.; Blom, H.; Rigler, R.; Widengren, J.;
Lasser, T. Opt. Express 2007, 15, 5366−5375.
(11) Wong, D. W. S. Appl. Biochem. Biotechnol. 2009, 157, 174−209.
(12) ten Have, R.; Teunissen, P. J. Chem. Rev. 2001, 101, 3397−413.
(13) Bugg, T. D.; Ahmad, M.; Hardiman, E. M.; Singh, R. Curr. Opin.
Biotechnol. 2011, 22, 394−400.
(47) Wen, F.; Dong, Y. H.; Feng, L.; Wang, S.; Zhang, S. C.; Zhang,
X. R. Anal. Chem. 2011, 83, 1193−1196.
(48) Bonagura, C. A.; Bhaskar, B.; Shimizu, H.; Li, H. Y.;
Sundaramoorthy, M.; McRee, D. E.; Goodin, D. B.; Poulos, T. L.
Biochemistry 2003, 42, 5600−5608.
(14) Boerjan, W.; Ralph, J.; Baucher, M. Annu. Rev. Plant Biol. 2003,
54, 519−546.
(15) Akiyama, T.; Magara, K.; Matsumoto, Y.; Meshitsuka, G.; Ishizu,
A.; Lundquist, K. J. Wood Sci. 2000, 46, 414−415.
(16) Akiyama, T.; Sugimoto, T.; Matsumoto, Y.; Meshitsuka, G. J.
Wood Sci. 2002, 48, 210−215.
(17) Gall, D. L.; Kim, H.; Lu, F. C.; Donohue, T. J.; Noguera, D. R.;
Ralph, J. J. Biol. Chem. 2014, 289, 8656−8667.
D
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