Coevolution of the Activity and Thermostability of an ?-Keto Ester Reductase for Better Synthesis of an (R)-α-Lipoic Acid Precursor

Article abstract of DOI:10.1002/cbic.201900693

In this work, we have identified a significantly improved variant (S131Y/Q252I) of the natural ?-keto ester reductase CpAR2 from Candida parapsilosis for efficiently manufacturing (R)-8-chloro-6-hydroxyoctanoic acid [(R)-ECHO] through co-evolution of activity and thermostability. The activity of the variant CpAR2_S131Y/Q252I towards the ?-keto ester ethyl 8-chloro-6-oxooctanoate was improved to 214 U mg^?1—from 120 U mg^?1 in the case of the wild-type enzyme (CpAR2_WT)—and the half-deactivating temperature (T₅₀, for 15 min incubation) was simultaneously increased by 2.3 °C in relation to that of CpAR2_WT. Consequently, only 2 g L^?1 of lyophilized E. coli cells harboring CpAR2_S131Y/Q252I and a glucose dehydrogenase (GDH) were required in order to achieve productivity similar to that obtained in our previous work, under optimized reaction conditions (530 g L^?1 d^?1). This result demonstrated a more economical and efficient process for the production of the key (R)-α-lipoic acid intermediate ethyl 8-chloro-6-oxooctanoate.

Full text of DOI:10.1002/cbic.201900693

Accepted Article
Title: Co-evolving the Activity and Thermostability of an ε-Ketoester
Reductase for Better Synthesis of (R)-α-Lipoic Acid Precursor
Authors: Yao Xu, Qi Chen, Zhi-Jun Zhang, Jian-He Xu, and Gao-Wei
Zheng
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ChemBioChem
10.1002/cbic.201900693
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Co-evolving the Activity and Thermostability of an ε-Ketoester
Reductase for Better Synthesis of (R)-α-Lipoic Acid Precursor
[
a]
[a]
[a]
[a]
[a]
Yao Xu, Qi Chen, Zhi-Jun Zhang, Jian-He Xu, and Gao-Wei Zheng*
Abstract: In this work, we identified a significantly improved variant S131Y/Q252I of natural ε-ketoester reductase CpAR2 from
Candida parapsilosis for efficient manufacturing (R)-ECHO [(R)-8-chloro-6-hydroxyoctanoate] by co-evolving the activity and
thermostability. The activity of variant CpAR2S131Y/Q252I towards ε-ketoester ethyl 8-chloro-6-oxooctanoate was improved to 214 U
mg−1 from 120 U mg−1 of the wild-type enzyme (CpAR2WT), and the half-deactivating temperature (T50, for 15 min incubation) was
simultaneously increased by 2.3°C compared with CpAR2WT. Consequently, only 2 g L−1 of lyophilized E. coli cells harboring
CpAR2S131Y/Q252I and glucose dehydrogenase (GDH) were required to achieve a similar productivity (530 g L-1 d-1) of our previous
work under the optimized reaction conditions. This result demonstrated a more economical and efficient process for the production of
the key (R)-α-lipoic acid intermediate.
Introduction
α-Lipoic acid is a kind of safe and potent chiral antioxidant that
can be used as dietary supplement and for treatment of diabetic
neuropathy[ and nephropathy, although only its R-enantiomer
1]
[2]
[
3]
shows the desired bioactivity . Nowadays, most of the products
available commercially are still produced and employed in a
form of racemic mixture, because the inactive S-enantiomer did
not show any obvious side effect,[4] and the price of the single R-
enantiomer is as much as five times that of the racemate.
Nevertheless, in order to reduce the metabolic burden of the
human body, it is highly desired to develop a green and efficient
method to asymmetrically synthesize the enantiomerically pure
Scheme 1. Chemoenzymatic route for the asymmetric synthesis of (R)-α-
Lipoic acid (LA).
However, the large scale application of these biocatalytic
processes are limited by low substrate loading, poor conversion,
or insufficient stereoselectivity and productivity. This is because
the bulky and special structure of ε-ketoester substrate ECOO,
(
R)-α-lipoic acid, instead of tedious and eco-unfriendly
diastereomeric resolution of the chemically synthesized
racemate, which is currently used in industry.[5]
Although numerous routes have been developed for the
synthesis of (R)-α-lipoic acid,[5,6] they are constrained by some
issues, including the maximum theoretical yield of 50% (kinetic
resolution), the complex synthetic process, and the generally
inadequate stereoselectivity (requiring the additional processes
to upgrade the optical purity of product). Biocatalytic asymmetric
reduction of ethyl 8-chloro-6-oxooctanoate (1, ECOO) developed
recently is a reliable, straightforward and easy to up-scale route
to the key precursor ethyl (R)-8-chloro-6-hydroxyoctanoate (2,
making it more difficult to be reduced by ketoreductases
compared with the well-studied α- or β-ketoesters.[8] This
dramatically decreases the probability of hitting an active
enzyme towards ECOO-like ε-ketoesters. Therefore, it is difficult
to create an industrially feasible biocatalyst for green synthesis
of (R)-ECHO, if considering the long distance between the ε-
position of the carbonyl group accepting the hydride from
NAD(P)H and the ester group positioned in the binding site of
ketoreductases.
(
R)-ECHO) for introduction of the chiral center into (R)-α-lipoic
Recently, we identified an unusual ε-ketoester reductase
CpAR2 from Candida parapsilosis.[9] Using CpAR2 as catalyst,
acid, which attracts widespread attention due to its advantages
of 100% theoretical yield and simple operation process for the
(
(
R)-ECHO with 99% ee was produced in a space-time yield
STY) of 530 g L−1 d−1 via the asymmetric reduction of ECOO,
(
R)-α-lipoic acid manufacture (Scheme 1).[7]
highlighting a particularly attractive and potential process for
enzymatic synthesis of (R)-α-lipoic acid precursor. However,
–
1
high biocatalyst loading (10
g
L
of dry E. coli cells
[
a]
Y. Xu, Q. Chen, Dr. Z. J. Zhang, Prof. J. H. Xu, Prof. G. W. Zheng
State Key Laboratory of Bioreactor Engineering, Shanghai
Collaborative Innovation Center for Biomanufacturing Technology,
East China University of Science and Technology, Shanghai
coexpressing CpAR2 and a glucose dehydrogenase BmGDH) is
required to achieve such high STY in the bioreduction process,
resulting in serious emulsification and tedious multi-step
downstream processing to obtain high product isolated yield. For
large scale application, a low biocatalyst loading is highly
desired. It not only reduces cost contribution also avoids post-
reaction processing issues (such as foaming or formation of
200237, China
E-mail: gaoweizheng@ecust.edu.cn
Supporting information for this article is given via a link at the end of
the document.
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ChemBioChem
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emulsions during the extraction of product from water using
organic solvents). Generally, the biocatalyst loading should be
less than 5 g L–1 for a green and economical chemical
process.[10]
Directed evolution has been proven to be an effective strategy
for improving the performance of enzymes by mimicking natural
evolution without the knowledge of protein structure-function
relationships.[11] To reduce the biocatalyst loading, and meet or
exceed the targets for the practical application, multiple enzyme
features are need to be evolved simultaneously. Due to the
excellent stereoselectivity (>99% ee) of CpAR2, we here intend
to improve its activity and thermal stability simultaneously. To
avoid the frequently encountered tradeoff between different
properties, we adopted a co-evolution strategy for identifying the
overall improved variants but excluding those mutants with bias
on any individual performance, either activity or stability, which is
important to create promising biocatalysts of industrial potential.
Figure 1. The locations of S131 and Q252 in the modelled structure of CpAR2.
Results and Discussion
To identify CpAR2 variants with improved activity and
thermostability, we first constructed a mutant library by error-
prone PCR[
12]
using pET28a-CpAR2 as a template, and
performed an initial screening of approximately 4,000 bacterial
clones with 1-4 mutated residues per clone on average. As a
result, two potential hot spots, S131 and Q252, for enhancing
both the activity and thermostability of CpAR2 were identified
through two rounds of epPCR-based random mutagenesis and
subsequent high-throughput screening based on 96-well plate
assay. A substrate-docking simulation also demonstrated that
Ser131 is a residue surrounding the active site of CpAR2, which
might interact with the ester group of the substrate ECOO (within
a distance of 6 Å) (Figure 1).
Figure 2. The specific activities of mutants at S131 position.
During the process of screening, we also noticed that a double
mutant S131L/Q252I exhibited better thermostability than the
mutants S131L and S131Y (Table 1), suggesting that the
mutation of thermally labile residues (e.g., Gln, Asn, Asp, Cys)
on the surface of protein molecules is effective to increase
CpAR2 thermostability.[14]
To elucidate the reason behind the enhanced specific activity
of CpAR2 variants towards ECOO, we performed a kinetic
analysis as shown in Table 1. Specific activities of the purified
To further investigate the effect of all Ser131 mutations, we
performed site-directed mutagenesis to replace Ser131 of
CpAR2 with 19 other canonial amino acids, and assayed the
specific activity of all mutants. As shown in Figure 2, the
mutation of Ser131 into Tyr, Phe, Met or Leu significantly
improved the enzyme activity towards the ECOO. Among them,
the best mutant S131Y showed a nearly 2-fold higher specific
–
1
variants S131Y and S131Y/Q252I measured were 237 U mg
–
1
and 214 U mg , respectively, which are much higher than that
–
1
of the wild-type CpAR2 (120 U mg ). The kcat of variant S131Y
–
1
−1
activity (237 U mg ) compared with the wild-type enzyme (120
for ECOO reduction was up to 181 s , 2-fold higher than that of
–
1
−1
U mg ). Therefore, mutations of Ser131 seemed effective in
increasing the activity of CpAR2, even though Ser131 has no
direct interaction with the carbonyl group of the substrate.
Interestingly, unlike the general case, in which the mutation of
the residue near the active site into an amino acid with smaller
side chain group usually enhances the catalytic efficiency due to
the enlarged pocket and substrate channel.[13] Accordingly, the
mutations Ser131Tyr and Ser131Phe with larger side chain
containing a benzene ring exihibited the highest activity.
m
the wild-type enzyme (90.1 s ), and K decreased slightly,
resulting in an obviously increase in catalytic efficiency kcat/K
m
−
1
−1
−1
−1
(255 s mM vs 77 s mM ). The kinetic analysis revealed
that the S131 is a key residue for improve kcat of CpAR2 towards
ECOO, which contribute significantly to the increase of catalytic
efficiency. In addition, these mutations have no effect on the
stereoselectivity of CpAR2. All variants afforded the (R)-ECHO
with 99% ee.
Table 1. Properties of CpAR2 and its variants towards substrate ECOO.
1
50
5
Specific activity
k
cat
(s−1)
K
(mM)
m
k
cat/K
m
Half-life
(h)
T
Enzyme
WT
–1
(s mM−1)
−1
a
(U mg
)
(°C)
45.5
120
90.1
1.17
77
4.3
S131L
171
237
197
214
120 ± 6
181 ± 9
172 ± 7
140 ± 3
0.95 ± 0.19
0.71 ± 0.10
1.05 ± 0.10
0.65 ± 0.08
126
255
164
215
25.5
24.8
32.5
25.1
46.5
46.8
48.8
47.8
S131Y
S131L/Q252I
S131Y/Q252I
a
Half-life of wide-type CpAR2 and its mutants at 40°C.
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The specific activity of variant S131Y/Q252I was increased to
(Table 2, entries 2 and 3). In contrast, using the same amount
CpAR2WT/BmGDH cells as catalyst, only 68% substrate was
reduced within 22 h (Table 2, entry 1). Subsequently, a simple
reaction optimization was carried out. When 5% (v/v) DMSO
instead of 5% (v/v) EtOH was used as cosolvent and the
14 U mg–1 from 120 U mg . Moreover, the half-deactivating
–1
2
15
temperature (T50 ) was 2.3°C higher than the wild-type enzyme,
and the half-life of enzyme at 40°C were increased to more than
4 h from 4.3 h. To simplify the preparation of biocatalysts, we
2
o
o
coexpressed CpAR2S131Y/Q252I or CpAR2WT and glucose
a
reaction temperature was reduced to 35 C from 40 C, the same
amount substrate was completely converted within 4 h, providing
[
15]
dehydrogenase BmGDH for NADPH regeneration in an E. coli
cell according to previous reports.[9a] Then the whole cells of E.
−1
−1
a higher STY of 566 g L d , a similar productivity with our
[
9a]
coli
CpAR2WT/BmGDH,
CpAR2S131Y/BmGDH
and
previous work
but a 5-fold decrease in catalyst dosage.
CpAR2S131Y/Q252I/BmGDH as biocatalysts were used to perform
the asymmetric reduction of ECOO.
These results indicated that a more efficient engineered ε-
ketoester reductase for the synthesis of (R)-α-lipoic acid
precursor was developed by cooperative directed evolution. The
green and economically viable bioreduction process has been
applied to (R)-α-lipoic acid precursor production in 2000-L scale
reactors by Suzhou Fushilai Pharmaceutical Co., Ltd (Suzhou,
China).
With these biocatalysts in hand, we further evaluated the
feasibility for their use in reduction of 110 g L−1 of ECOO at low
biocatalyst
loading.
Using
only
2
g
L−1
of
CpAR2S131Y/Q252I/BmGDH or CpAR2S131Y/BmGDH cells as
−
1
catalyst, 110 g L of ECOO was both completely converted
within 5 h, providing (R)-ECHO with a similar space-time yield
Table 2. Asymmetric reduction of ECOO with lyophilized E. coli cells of CpAR2WT/BmGDH and CpAR2S131Y/Q252I/BmGDH.
Substrate loading
Enzyme loading
Time
(h)
Conversion
(%)c
Yield
(%)
ee
(%)c
STY
(g L−1 d−1
)
Entry
Biocatalyst
−1
−1
(g L
)
(g L
)
1
2
3
4
CpAR2WT/BmGDHa
110
110
110
110
2
22
5
68
81
84
85
85
99
99
66
CpAR2S131Y/BmGDHa
CpAR2S131Y/Q252I/BmGDHa
CpAR2S131Y/Q252I/BmGDHb
2
2
2
>99
>99
>99
448
454
566
5
>99
>99
4
a
Reaction conditions: 0.5 M ECOO (110 g L−1), 1.5 equiv. glucose, 2.0 g L−1 lyophilized E. coli cells harboring pET28a-CpAR2/BmGDH (CpAR2WT: 17.7 kU/g
cells; CpAR2S131Y/Q252I: 32.1 kU/g cells; BmGDH: 3.5 kU/g cells), NADP+ (0.1 mM), 0.5 mL EtOH, and 9.5 mL Tris-HCl buffer (100 mM, pH 7.0), 40°C, with pH kept
2 3
at 7.0 by auto-titrating 1.0 M K CO .
b
Reaction conditions: 0.5 M ECOO (110 g L−1), 1.5 equiv. glucose, 2.0 g L−1 lyophilized E. coli cells harboring CpAR2S131Y/Q252I/BmGDH (CpAR2S131Y/Q252I: 32.1
kU/g cells; BmGDH: 3.5 kU/g cells), NADP+ (0.1 mM), 0.5 mL DMSO, and 9.5 mL Tris-HCl buffer (100 mM, pH 7.0), 35°C, with pH kept at 7.0 by auto-titrating 1.0
2 3
M K CO .
c
Conversion and ee were determined by GC analysis.
Conclusion
In summary, we performed the screening of a random mutation
library of CpAR2 for identifying an enzyme variant with high
activity and thermostability. It was demonstrated that mutations
at Ser131 and Gln252 contribute to the enhanced specific
activity and thermostability, respectively. Among saturation
mutants of S131, S131Y exhibited the highest specific activity
for ECOO reduction. Kinetic analysis showed that its increased
catalytic efficiency is largely attributable to the increased kcat
.
Subsequently, the beneficial mutants were combined, achieving
the best double mutant S131Y/Q252I. Its specific activity was
215 U mg , and the half-deactivating temperature (T50 ) was
Figure 3. Scheme and results for the co-operative directed evolution of
CpAR2’s activity and thermostability.
–
1
15
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2.3°C higher than the wild-type enzyme, resulting in 5-fold
agar plates containing kanamycin (50 g/mL). The resulting
decrease in the enzyme loading (only 2 g dry engineered cells
per liter) and obvious simplification of the post-reaction
processing which would otherwise be very difficult in the case of
high dosage of the interface active protein biocatalyst due to the
severe emulsification issue often encountered during the
extraction of product with organic solvents (Figure 3).
mutant plasmids were confirmed by DNA sequencing.
Screening of variants with improved activity and
thermostability: Library colonies were picked with sterile
toothpicks and inoculated into 200 μL LB medium containing 50
μg/mL kanamycin in 96-well plates, before incubated overnight
at 37°C. The resulting cultures were inoculated into 400 μL LB
medium containing kanamycin in new 96-well plates, and
incubated at 37°C for 1.5−2.0 h. The protein expression was
induced by adding 0.2 mM IPTG (final concentration) and then
the incubation was continued at 16°C for another 24 h. Cells
were lysed by adding 200 μL buffer containing lysozyme (750
mg/L) and DNase I (10 mg/L) and incubating at 37°C for 2 h.
The plates were centrifuged at 2395×g and 4°C for 10 min. The
supernatant was then subjected to activity and thermostability
assay as follows: After 50 μL properly diluted supernatants from
the deep-well plate were transferred to microtiter plates, the
reduction reaction was initiated by adding 150 μL reaction
mixture composed of 100 mM Tris-HCl buffer (pH 7.0), 0.2 mM
NADPH and 2 mM substrate. The activity of variants was
determined at 30°C by measuring the change of absorbance at
340 nm for 220 s using a microplate spectrophotometer (BioTek,
USA). Each of the supernatants (20 μL) from the deep-well
plates was transferred to 96-well PCR plates, heated at a
defined temperature for 15 min, and followed by cooling on ice
for 5 min. Each of the properly diluted supernatants (50 μL) from
the pre-incubated 96-well PCR plates were transferred to a
microtiter plate. Then the residual activity of each variant was
measured using the microplate spectrophotometer under the
standard assay condition. The variants with higher initial or
residual activity than the parental enzyme were chosen for
subsequent rescreening. The top mutants were confirmed by
DNA sequencing and activity / thermostability assay of the
purified enzymes.
Experimental Section
Materials: The enzyme substrate, ethyl 8-chloro-6-oxooctanoate
(
ECOO), was provided by Suzhou Fushilai Medicine & Chemical
Co., Ltd. (Jiangsu, China). All others chemicals and reagents
were purchased from commercial suppliers. E. coli BL21 (DE3)
was used as cloning and expression host. Cells were incubated
m
in Luria-Bertani (LB) medium. The melting temperature (T ) of
enzyme was measured using circular dichroism. The
oligonucleotide primers designed for mutagenesis were
synthesized by Sangon Biotech Co. (Shanghai, China).
Restriction enzymes were purchased from New England Biolabs
®
(
Beijing, China). PrimeSTAR HS (Premix), Recombinant Taq
DNA Polymerase (TaqTM) and T4 DNA Ligase were purchased
from TaKaRa (Dalian, China).
Random mutagenesis: The plasmid pET-28a containing
CpAR2-WT gene was used as the template for the construction
of random mutant library. Error-prone PCR was performed with
primers (Table S1) incorporating EcoRI and XhoI restriction sites.
The PCR mixture (50 μL) was composed of pET28a-CpAR2-WT
DNA (<500 ng), 10 mM Tris-HCl (pH 8.3), 50 mM KCl, 1.5 mM
2
MgCl , 0.2 mM each dNTPs mixture, 0.4 μM each of the two
primers, and 0.5 μL DNA polymerases (5 U/μL). The PCR
mixture was pre-incubated at 95°C for 3 min, then was cycled 30
times with the following program: 94°C for 10 s, 60°C for 30 s,
and 72°C for 90 s. The amplified PCR product was purified with
TRIpure Reagent Kit from Aidlab (Beijing, China), digested with
EcoRI and XhoI, and ligated into the EcoR I and Xho I sites of
pET-28a. The resultant recombinant plasmid was transformed
into chemical competent E. coli BL21 (DE3) and plated onto
Luria-Bertani (LB) agar plates containing 50 g/mL kanamycin,
and the plasmids extracted from the kanamycin-resistant
colonies were used to isolate CpAR2 mutants as the random
library.
Site-directed mutagenesis: MEGAWHOP method (Megaprimer
PCR of Whole Plasmid) was used to generate site-directed
mutant with corresponding primers (Table S1). MEGAWHOP
was carried out with a mixture (50 μL) containing 1 ng pET28a-
CpAR2-WT DNA, 0.25 μM each of the two degenerate primers
Expression and purification of proteins: The E. coli BL21
(
DE3) cells harboring the recombinant pET-28a(+) plasmids of
CpAR2-WT and its variants were cultivated overnight at 37°C in
mL LB medium containing 50 μg/mL kanamycin. The overnight
culture was then inoculated into 100 mL LB and cultivated at
7°C and 180 rpm until the culture’s optical density (OD600
4
3
)
reaching 0.6−0.8. The protein expression was induced at 16°C
with the addition of IPTG to a final concentration of 0.1 mM.
After cultivation for 24 h, the cells were collected by
centrifugation. For preparing the lyophilized cells, a portion of
the cells was directly freeze-dried for 72 h. For protein
purification, another portion of the cells were washed and
resuspended in buffer A (20 mM sodium phosphate buffer, pH
7.4, 500 mM NaCl, 10 mM imidazole). The cell suspension was
disrupted by ultrasonication, and centrifugated (13,000×g, 4°C,
(
Table S1), 0.2 mM each dNTPs and 25 μL PrimeSTAR HS
4
0 min). Then the resulting supernatant was filtered through a
DNA Polymerase (1.25 U/25 μL). The PCR mixture was pre-
incubated at 98°C for 3 min, and then subjected to thermal
cycles (98°C for 10 s, 55°C for 5 s and 72°C for 6 min, 30
cycles). The resulting PCR product was digested with 1 μL DpnI
0.22 μm filter and then loaded into a standard Ni-NTA affinity
column which was pre-equilibrated with ice chilled buffer A. The
column was subsequently washed by elution buffer with an
increasing gradient of imidazole from 10 to 500 mM, using the
mixture of buffer A and buffer B (20 mM sodium phosphate
(
37°C for 2 h). The cloned plasmids were then transformed into
chemically competent E. coli BL21 (DE3) cells and plated on LB
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buffer, pH 7.4, 500 mM NaCl, 500 mM imidazole). The fractions
containing the target protein was collected and concentrated by
ultrafiltration using Ultra-4 centrifugal filter with 10 kDa cut off
For the half-life determination, each of the purified enzymes (1.0
mg/mL) was incubated at 40°C. Aliquots were taken at specific
time intervals and cooled on ice before activity assay, followed
by measuring the residual activity using the standard activity
assay method described above. The initial activity detected
before incubation was normalized as 100%. The half-lives were
determined according to first-order exponential decay function.
(
Millipore, USA) and then washed three times by buffer C (25
mM Tris-HCl, 150 mM NaCl, 1 mM DTT). The purity of target
protein was checked by SDS-PAGE (Figure S1). The purified
protein was stored at −80°C for further use. Protein
concentration of the purified enzyme was estimated by detecting
the absorbance at 280 nm using
a
NanoDrop 2000c
Steroselectivity of CpAR2 for substrate ECOO: The reaction
mixture consisted of 20 mM substrate, 20 U of purified CpAR2,
20 U of BmGDH, 30 mM glucose, 0.5 mM NADP+ in 1 mL of
Tris-HCl buffer (pH 7.0), with shaking at 30°C for 12 h. Then
each of the reaction mixture was extracted with ethyl acetate
and the conversion of substrate was determined by GC analysis.
The product was acetylated before GC analysis. An appropriate
amount of acetic anhydride, 4-dimethylaminopyridine (DMAP)
and pyridine was added to the ethyl acetate solution containing
the product, and incubated at 30°C for 12 h. The mixture was
then diluted properly with ethyl acetate for GC analysis.
spectrophotometer (Thermo Scientific) and taking into account
the calculated extinction coefficients with the ExPASy
ProtParam Tool.
Enzyme assays: The reductase activity was assayed at 30°C
by monitoring the decrease in the absorbance of NADPH at 340
nm within
UV1800). The standard assay mixture (1 mL) was composed of
70 μL Tris-HCl buffer (100 mM, pH 7.0), 10 μL ethyl 8-chloro-6-
1 min on a UV-spectrophotometer (Shimadzu,
9
oxooctanoate (800 mM), 10 μL NADPH (10 mM), and 10 μL
enzyme with an appropriate concentration. One unit of enzyme
activity (U) was defined as the amount of enzyme that can
catalyze the oxidation of 1 μmol NADPH per minute under above
conditions.
GC analysis: The conversion of substrates and enantiomeric
excess of products were determined by GC analysis using a
Shimadzu GC-2014 gas chromatography, equipped with a CP-
Chirasil-Dex CB column (25 m × 0.25 mm × 0.39 mm, Agilent
Technologies), a capillary column (30 m length, 0.25 μm film
Kinetic properties: The kinetic parameters of the purified
CpAR2 were determined by measuring the activity at the
different substrate concentrations (0.1−10 mM) and a fixed
NADPH concentration. All data were handled by using the
software OriginPro 9.0 to adjust to the Michaelis-Menton model
thickness, HP-5MS, Agilent Technologies) and
a
flame
ionization detector (FID). Injection and detection temperatures
were set at 280°C, while the oven temperature was fixed at
160°C (Figure S4).
(
Figure S2).
Homology modeling: A homology model of CpAR2 was built
Thermostability: The thermal stability of CpAR2 was also
based on the crystal structure of a methylglyoxal reductase
GRE2 from Saccharomyces cerevisiae (PDB entry: 4PVD),
−
1
[16]
examined by incubating the purified enzymes (1.0 mg mL ) at
varied temperatures (44−49°C), and the residual activity was
assayed as described above.
which shares 40.5% sequence identity with CpAR2. The three-
dimensional structure model of CpAR2 was generated using
Modeller 9.11 based on the crystal structure of a yeast
methylglyoxal / isovaleraldehyde reductase Gre2 (with 40.5%
identity to CpAR2) in complex with NADPH (PDB ID: 4PVD).
AutoDock Vina was used for the docking of substrate ECOO into
the model structure. All figures of protein models were prepared
with PyMOL.[17]
15
The half-deactivating temperature (T50 ) was determined by
measuring the residual activity after incubation at a gradient
15
temperatures for 15 min. The T50 was estimated as temperature
at which heat treatment for 15 min reduced the initial activity by
50%. The purified enzymes were diluted to 1.0 mg/mL and 20 μL
each of the purified enzyme was distributed into 8 thin-walled
PCR tubes, which were then subjected to incubation at a
temperature gradient using a thermocycler (Bio-Rad, USA). After
incubation, the PCR tubes were cooled on ice immediately.
Then, the residual activities of the enzymes were measured with
the standard protocol (Figure S3).
Biocatalytic asymmetric reduction of ECOO: A biocatalytic
reaction 10-mL scale was conducted to monitor the reaction
time-courses of CpAR2 wild type and its variants. The reaction
−
1
mixture was composed of substrate ECOO (0.5 M, 110 g L ),
1.5 equiv. of glucose, 2 g L−1 of lyophilized E. coli cells co-
expressing CpAR2 and BmGDH (CpAR2WT: 17.7 U/mg cells;
CpAR2S131Y/Q252I: 32.1 U/mg cells; BmGDH: 3.5 U/mg cells), 5%
m
The melting temperature (T ) of CpAR2 were determined via
circular dichroism (CD) spectroscopy. The concentration of initial
protein samples was diluted to 0.3 mg/mL with 100 mM
potassium phosphate buffer (pH 7.0). The CD spectra of the
diluted samples were individually detected in a 0.2 cm-path-
length cuvette using a Chirasan circular dichroism spectrograph
+
ethanol (v/v) or 5% DMSO (v/v) and 0.1 mM NADP in Tris-HCl
(100 mM, pH 7.0). The mixture was agitated at 30°C-40°C and
500 rpm, and the pH of reaction system was maintained at 7.0
(
Applied Photophysics, UK) by scanning at 180-260 nm (1
nm/step) under increasing temperatures (35−65°C, 2°C/step,
°C/min). Then the protein CD spectra measured as a function
of temperature were analyzed using Global 3 analysis software
Applied Photophysics, UK) to obtain the calculated T values.
2 3
by auto-titrating 1.0 M K CO solution. Aliquots of sample (200
μL each) were taken at pre-determined times. The product in the
sample was extracted into 200 μL ethyl acetate and the resultant
2
organic phase was dried over anhydrous Na
2 4
SO for the analysis
(
m
of substrate conversion and product ee by GC.
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Entry for the Table of Contents (Please choose one layout)
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Yao Xu, Qi Chen, Zhi-Jun Zhang, Jian-
He Xu, and Gao-Wei Zheng*
Page No. 1 – Page No. 6
Title
Co-evolving
the
Activity
and
Thermostability of an ε-Ketoester
Reductase for Better Synthesis of (R)-
α-Lipoic Acid Precursor
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