M. Matsumoto and Y. Tahara, Enhanced Esterification Activity and Thermostability…, Chem. Biochem. Eng. Q., 37 (4) 255–260 (2023)

Enhanced Esterification Activity and Thermostabilityof Imprinted Poly(Ethylene Glycol)-Lipase Complex
M. Matsumoto* and Y. Tahara
Department of Chemical Engineeringand Materials Science, Doshisha University,
Kyotanabe, Kyoto 610-0321, Japan

255

This work is licensed under aCreative Commons Attribution 4.0
International License

doi: https://doi.org/10.15255/CABEQ.2023.2205
Original scientific paper
Received: April 15, 2023
Accepted: September 19, 2023

Although the range of applications for enzymatic reactions in organic solvents is
rapidly expanding, this study focused on the enzymatic activity in the esterification oflauric acid with benzyl alcohol, and thermostability of lipase using poly(ethylene glycol)(PEG)-lipase complex and molecular imprinting techniques. The catalytic activity wasenhanced through molecular imprinting and the PEG-lipase complex. The imprinting operation was particularly effective for catalytic activity after forming the PEG-lipase complex. The kinetic analysis of the lipase-catalyzed esterification revealed that the increasein esterification rate with imprinted lipases was mainly due to the higher maximum rateachieved by the system. The thermostability of the lipases was significantly improved by
imprinting at all temperatures (50~70 °C). After forming a PEG-lipase complex, the imprinted lipase exhibited much higher reactivity and thermostability compared to the native lipase and the imprinted PEG-lipase complex.
Key words
lipase, imprinting, PEG-lipase complex, thermostability

Introduction
Enzymes typically function in aqueous environments in nature. However, as the application of
enzymes in chemical synthesis has become morepractical, they are increasingly employed in
non-aqueous media instead of aqueous media formany poorly water-soluble substrates1. Lipases, inparticular, are well-known enzymes that function in
non-aqueous media2. Depending on their source,certain lipases can remain active in hydrophobic organic solvents3. However, the disadvantage of enzymatic reactions in organic solvents is that the native
enzyme is insoluble in these solvents, and the reaction rates are affected by mass transfer of the substrate4. Therefore, enhancing enzymatic reaction
rates in organic solvents through simple methods
remains challenging from a practical standpoint.
Two approaches can be employed to address thischallenge: the design of a reaction apparatus, andactivation of enzyme functions. The former approach involves the use of microreactors5. Microreactors are characterized by a high specific surface
area, which often eliminates the effects of masstransfer. The latter approach involves enzyme activation through pretreatment of the enzyme with organic solvents or substrate analogs This method offers several advantages including easy preparation
Corresponding author: Tel/Fax +81-774-65-6655,
E-mail: mmatsumo@mail.doshisha.ac.jp*

and the absence of additives in the reaction system.
One method for enzyme activation involves pretreating an enzyme with an organic solvent before
its use in an enzymatic reaction in the organic solvent6. Another pretreatment method uses the molecular imprinting technique with a substrate analog,
which has been studied in our laboratory7–9. Thismethod entails the lyophilization of an enzyme
loaded with a substrate analog to form a complex,
followed by the removal of the substrate analog
through washing the organic solvent10. The conformation of the active site of the enzymes is retainedeven after the removal of the substrate analog10. Additionally, when lipases are used as the enzyme,
their active sites in the aqueous solution are coveredby a flexible region known as the “lid” of the lipase.
Interfacial activation by the organic solvent and amphiphiles causes an opening in this lid allowing theactive site to become accessible11. The imprinted
enzymes, especially lipases, have exhibited muchhigher activity compared to those non-imprinted.
Another method for activating an enzyme is theamphiphile-coating method, in which the enzyme’s
surface is covered by an amphiphile monolayerthat is soluble in most organic media12. The primaryreason for enzyme activation in this case is theincreased solubility of the amphiphile-enzyme complex in organic media. Amphiphiles such aspoly(ethylene glycol) (PEG)4,13, glycolipid12 and dialkyl glutamic ribitol amide14 have been reported

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M. Matsumoto and Y. Tahara, Enhanced Esterification Activity and Thermostability…, Chem. Biochem. Eng. Q., 37 (4) 255–260 (2023)

for coating lipase and a-chymotrypsin. Furthermore,
imprinted surfactant-coated lipases have been simultaneously prepared by combining imprinting
and surfactant coating,12,15 resulting in improved esterification activity and enantioselectivity. However, it remains quantitatively unclear which effect,surfactant or imprinting, exerts the most influenceon the reaction rate. Additionally, the surfactant
used in previous studies12,13,15 had to be synthesized,
making the use of this surfactant impractical. No
study has investigated the combined effects of imprinting and coating with commercially available
PEG on enzyme reaction systems including theirkinetics and thermostability.
In this study, we formed a PEG-lipase complex
using PEG as a coating material for the enzyme lipase and investigated the effect of imprinting and
PEG on the esterification reaction kinetics and thermostability using these enzyme catalysts.

Experimental methods
Materials

Lipase from porcine pancreas (Type II, L3126,
Sigma-Aldrich (St. Louis, U.S.A.)) was utilized in
all our experiments. We employed poly(ethylene
glycol) (PEG) with an average molecular weight of
20,000. A PEG-lipase complex of 20,000 exhibits
higher lipase activity in ionic liquids compared toother molecular PEG weights13. Lauric acid andbenzyl alcohol served as substrates in the esterification. Cyclohexane and toluene were chosen as themedia for esterification, while hexane was used in
the pretreatment process and dried with 3 Å molecular sieves before use. Benzyl laurate, an esterification product, was used as a standard sample forquantification and was purchased from Tokyo Chem
Ind. (Tokyo, Japan). All other reagents were purchased from Wako Pure Chem. Ind. (Osaka, Japan),
except for the enzyme. All the reagents were of analytical grade and used without further purification.
Imprinting of lipase

The pretreatment of native lipase or PEG- lipase complex with octanoic acid as an imprint molecule followed a previously described method5–7.
Lipase (300 mg) was dissolved in 9 cm3 of a phosphate buffer solution at pH 7. Octanoic acid (3.5mmol) and Tween 80 (1000 mg) were dissolved in
10 cm3 of ethanol. To the enzyme solution, 1 cm3 ofoctanoic acid solution was added, and the mixed
solution was incubated for 30 min at 25 °C. The
mixed solution was then freeze-dried. The obtained
powder was washed with hexane to remove octanoic acid. The resulting lipases were dried in vacuo(Ulvac, GCD-051X, Chigasaki, Japan).

PEG-lipase complex

Our procedure for the preparation of the
PEG-lipase complexes closely followed the method
described in a previously published paper13. Lipase(10 mg) and PEG (16 mg) were dissolved in a phosphate buffer (5 mL, pH 7). Toluene (13 mL) was
added to the lipase solution and emulsified at 20,000rpm for 3 min using a homogenizer (Thermo Fisher
Scientific, Polytron PT 2500, Waltham, U. S. A.) inan ice-bath to prepare water-in-oil (w/o) emulsions.
The w/o emulsion was immediately frozen in liquid
nitrogen, followed by lyophilization for 24 h.
PEG-lipase complexes were obtained as white powders.
Esterification

To assess the effect of imprinting and the
PEG-lipase complex, lauric acid and benzyl alcohol
were used as substrates for esterification. The reactions were initiated by adding 1 mg of lipases­(native, imprinted or PEG-lipase complex) of an organic solution (cyclohexane or toluene) containing
both substrates (30 mmol dm–3 lauric acid and
30 mmol dm–3 benzyl alcohol) in a vial at 37 °C and1000 rpm (Eppendorf, Thermomixer Comfort,
Hamburg, Germany). Samples were taken every 30
minutes for up to 2 hours. The vial tubes were removed from the thermomixer, and immediately
centrifuged for 3 min at 10000 rpm (Eppendorf,
Centrifuge 5418). The benzyl laurate concentrationin the supernatant was measured using HPLC. Reaction rates were determined as the initial reaction
rate in the linear region during the first 90 minutesof the reaction. All experiments were performed intriplicate and demonstrated reproducibility with arelative standard deviation of 5 %.
Thermostability of lipase

The lipases were incubated in cyclohexane at
50~70 °C. Following the designated incubation period, the cyclohexane solution containing substrates,
lauric acid, and benzyl alcohol, was added and thereaction was carried out at 37 °C and 1000 rpm for
10 min in a thermomixer. Samples were collected tomeasure the initial esterification rate, and the residual activities were determined as the ratio of the initial esterification rates of the incubated lipases tothose of lipases without incubation.
AnalysisStandard benzyl laurate solutions were utilized
to prepare the calibration curves. The concentrations of the benzyl laurate in the cyclohexane solutions were determined by HPLC (Shimadzu, Prominence, Kyoto, Japan) with a Wakosil-II 5C18AR

M. Matsumoto and Y. Tahara, Enhanced Esterification Activity and Thermostability…, Chem. Biochem. Eng. Q., 37 (4) 255–260 (2023)(4.6 mm X 150 mm, Wako Pure Chem. Ind.) column and an eluent solution (ethanol: methanol: water = 15:5:3) as a mobile phase (0.6 cm3 min–1). Esters were detected using an UV detector at 255 nm(Shimadzu, SPD10AV). Retention time was 14 min.

257

(a)

Results and discussion
Effect of pretreatment on esterification

The effect of pretreatment (imprinting and
PEG-lipase complex) on the esterification of thelauric acid with benzyl alcohol was examined. Fig.
1 shows the initial esterification rates in cyclohexane and toluene. In a previous paper16 it was reported that the initial esterification rates exhibited a linear relationship with solvent hydrophobicity (log P)
in porcine pancreatic lipase-catalyzed esterificationof oleic acid with ethanol. Therefore, cyclohexane,
possessing a higher log P value than toluene, yielded a larger initial reaction rate than toluene, except
in the case of imprinted lipase. As described in previous papers7–9, imprinting was effective for activating the lipase in toluene, but had little effect in cyclohexane. Conversely, the PEG-lipase complex
substantially increased the initial reaction rate incyclohexane, although it had minimal effect in toluene. This suggests that PEG-lipase complexes were
not effectively formed in toluene. Therefore, we
prepared lipases by combining imprinting and
PEG-lipase complex. As shown in Fig. 1, the activity of the lipase imprinted after forming a PEG-lipase complex was clearly higher than that of the(b)

F i g . 2 – Effects of substrate concentrations on initial rate ofesterification. (a) native lipase and, (b) PEG + imprinted lipase. Solid lines were calculated using Eq. (1).

complexes of imprinted lipase with PEG. The degree of increase in activity exceeded the combined
activity of the imprinted lipase and the PEG-lipase
complex alone, indicating a synergistic effect. The
lipase imprinted after the PEG-lipase complex
demonstrated a 4.1-fold and 7.0-fold increase in activity in cyclohexane and toluene compared to thenative lipase. In subsequent experiments, cyclohexane was used as the reaction solvent, which gavehigher reaction rates and lower toxicity17.
Effect of PEG-lipase complex and imprinting onesterification kinetics

F i g . 1 – Effect of imprinting and PEG-lipase complex on initial esterification rate in cyclohexane and toluene. Imprinted +PEG denotes that lipase was imprinted and formed a complex
with PEG. PEG + Imprinted denotes that lipase was a complex
with PEG and imprinted. Lauric acid concentration = benzyl
alcohol concentration = 30 mmol dm–3.

Fig. 2 shows the effect of substrate concentrations on the initial rate of esterification of both native and treated lipases. No substrate inhibitions
were observed. The kinetic rate equation for esterification by lipase can be written using the PingPong Bi-Bi mechanism without substrate inhibition
as shown in Eq. 118,19.
Vm [ LA ][ BA ]V=
(1)
K m,LA [ BA ] + K m,BA [ LA ] + [ LA ][ BA ]

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M. Matsumoto and Y. Tahara, Enhanced Esterification Activity and Thermostability…, Chem. Biochem. Eng. Q., 37 (4) 255–260 (2023)

Ta b l e 1 – Kinetic parameters for esterification reaction
Lipases

(a)

Vm
Km,LA
Km,BA
[mol m–3 h–1] [mol m–3] [mol m–3]

Native0.801±0.036 3.21±0.70 3.06±0.69

PEG complex

2.04±0.16

Imprinted

0.780±0.032 2.89±0.61 2.88±0.61

Imprinted/PEG complex

5.59±0.96

20.3±5.8 45.4±11.9

PEG complex/Imprinted

7.22±1.12

5.27±2.20 61.3±14.2

8.31±1.75 10.3±2.0

Lauric acid concentration = benzyl alcohol concentration =30mmol dm–3

where Vm is the maximum rate, [LA] and [BA] are
the concentrations of lauric acid and benzyl alcohol,
and Km,LA and Km,BA are the Michaelis constants oflauric acid and benzyl alcohol, respectively. Table 1
lists the kinetic parameters determined using a nonlinear least-squares method integrated into Sigma
Plot 14.0. In Fig. 2, the solid lines calculated withthese kinetic parameters agree well with the experimental data. In previous reports7–9, imprinting led toa decrease in Michaelis constants, Km,LA and Km,BA,
without changing the maximum rate Vm. However,in our present system, the Michaelis constants were
not significantly different from those in the native
system, despite the decrease in Michaelis constants.
On the other hand, the PEG-lipase complex increased the Michaelis constants, suggesting that thePEG coating on the lipase’s surface probably inhibited the interaction between the substrate and thelipase. The PEG-lipase complex significantly increased the maximum rate. This increase in the
maximum rate can be attributed to the following
factors: The net effect of the PEG molecules shifts
the equilibrium between the closed and open formsof the lid in lipases from Candida rugosa toward
the open form20. The preferable conformation
change of the lipase (lid open) was initially inducedby the contact with the PEG molecule, and imprinting subsequently facilitated substrate accessibility
to the active site.

(b)

(c)

Effect of PEG-lipase complex and imprinting on
thermostability of lipase

Fig. 3 shows the time-courses of the residual
activities of the lipases in cyclohexane at 70 °C. In
all the lipases, enzyme activity declined rapidly at
20 min and then remained constant. Similar behaviors were observed at other temperatures (60 and 50°C), indicating that enzyme inactivation conforms
to a first-order reversible rate equation. The residual
activity at, is determined by the following equation
which is based on reversible first-order deactivation
kinetics21.
at - a¥(2)= exp {-( kd + kr ) t}
1- a¥

F i g . 3 – Thermostability of treated lipases at (a) 70 °C, (b)60 °C and (c) 50 °C. Solid lines were calculated using Eq. (2).
Imprinted + PEG denotes that lipase was imprinted and formed
a complex with PEG. PEG + Imprinted denotes that lipase wasa complex with PEG and imprinted.

where kd and kr are lipase deactivation and refolding
rate constants, respectively, a∞ is the residual activity at equilibrium and t is the elapsed time.

M. Matsumoto and Y. Tahara, Enhanced Esterification Activity and Thermostability…, Chem. Biochem. Eng. Q., 37 (4) 255–260 (2023)

259

Ta b l e 2 – Parameters for deactivation kinetics
70 °C

NativePEG-complex

Imprinted

Im+PEG*PEG+Im**a∞0.177±0.004

0.292±0.008

0.586±0.011

0.434±0.018

0.586±0.012

kd /min

0.276±0.006

0.458±0.040

0.0982±0.0087

0.140±0.016

0.0927±0.0092

kr/min0.0594±0.0023

0.189±0.021

0.139±0.016

0.107±0.017

0.131±0.017

a∞0.439±0.010

0.453±0.013

0.670±0.009

0.610±0.008

0.739±0.010

kd /min

0.181±0.014

0.181±0.019

0.0570±0.0046

0.0603±0.0032

0.0386±0.0039

kr/min0.141±0.014

0.150±0.021

0.116±0.012

0.0944±0.067

0.109±0.014

a∞0.519±0.019

0.565±0.015

0.793±0.009

0.676±0.007

0.817±0.012

kd /min

0.0286±0.0023

0.0264±0.0016

0.0179±0.0018

0.0368±0.0019

0.0167±0.0026

kr/min0.0309±0.0040

0.0269±0.0028

0.0687±0.0092

0.0768±0.0053

0.0757±0.0152

60 °C

50 °C

* Im + PEG denotes that lipase was imprinted and formed a complex with PEG.
** PEG + Im denotes that lipase was a complex with PEG and imprinted.

Three parameters, kd, kr and a∞ were calculated
for the data in Fig. 3 using Sigma Plot 14.0. The
parameter values obtained at each temperature arelisted in Table 2. In Fig. 3, the solid lines calculatedusing Eq. (2), closely approximate the experimental
data, affirming the justifiability of employing Eq. 2.
Table 2 reveals that the thermostability of both thenative lipase and the PEG-lipase complex is significantly enhanced by imprinting at all temperatures.
On the other hand, the PEG-lipase complex did not
produce a significant improvement in thermostability. The improved thermostability due to imprinting
indicates the importance of retaining the imprinted
structure for thermostability, and the subsequent
PEG modification after imprinting is presumed toslightly impair the retention of the imprinted structure. Our prior studies on imprinting for transesterification in cyclohexane and ionic liquids haveshown slight stabilization of lipase, although theextent was not notably large7,8. This result suggeststhat carboxylic acid was not the ideal substrate analog in the transesterification reaction, and that theester should have been considered as the imprinting
molecule. Stabilization of lipase by imprinting not
only significantly decreased the deactivation rate
constant, but also led to a very high retention of activity after 90 minutes. Although longer-term stability will be investigated in the future, our finding
indicate that that imprinting lipase-catalyzed esterification significantly improves its thermostability.

Conclusions
The effects of imprinting and the PEG-lipase
complex on catalytic activities and thermostabilityof lipase were investigated for esterification in or-

ganic solvents. Both imprinting and the PEG-lipase
complex enhanced the enzyme’s activity. The activity of the imprinted PEG-lipase complex increased
5.9-fold in cyclohexane and 27-fold in toluene whencompared to the native lipase, respectively. Kinetic
analysis of lipase-catalyzed esterification revealed
that while the PEG-lipase complex and imprinting
had a minor effect on Michaelis constants, the increase in the esterification rate primarily resulted
from an enhanced maximum rate. Furthermore, imprinting significantly improved the thermostabilityof the lipases at every temperature. In contrast, thePEG-lipase complex did not significantly improve
thermostability. After forming a PEG-lipase complex, the imprinted lipase exhibited significantly
improved reactivity and thermostability compared
to both the native lipase and the imprinted PEG-lipase complex. These results hold promising prospects for practical applications in the future.
Acknowledgments
We thank Toshihide Shirai for his experimental
assistance.
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