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\begin{document}

\title{Octadecyl and sulfonyl modification of diatomite synergistically
improved the immobilization efficiency of lipase and its application in
the synthesis of pine sterol esters}



\author[1]{Yifei Zhang}%
\author[1]{Wei Zhang}%
\author[1]{Guangzheng Ma}%
\author[1]{Binbin Nian}%
\author[1]{Yi Hu}%
\affil[1]{Nanjing Tech University}%


\vspace{-1em}



  
  \date{May 26, 2024}


\begingroup
\let\center\flushleft
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\selectlanguage{english}
\begin{abstract}
Phytosterols usually have to be esterified to various phytosterol esters
to avoid their disadvantages of unsatisfactory solubility and low
bioavailability. The enzymatic synthesis of phytosterol esters in
solvent-free system has advantages in terms of environmental
friendliness, sustainability, and selectivity. However, the limitation
of the low stability and recyclability of the lipase in the solvent-free
system, which often requires a relatively high temperature to induce the
viscosity, also increased the industrial production cost. In this
context, a low-cost material, namely diatomite, was employed as the
support in the immobilization of Candida rugosa lipase (CRL) due to its
multiple modification sites. The Fe3O4 was also then introduced to this
system for quick and simple separation via the magnetic field. Moreover,
to further enhance the immobilization efficiency of diatomite, a
modification strategy which involved the octadecyl and sulfonyl group
for regulating the hydrophobicity and interaction between the support
and lipase was successfully developed. The optimization of the ratio of
the modifiers suggested that the -SO3H/C18 (1:1.5) performed best with
an enzyme loading and enzyme activity of 84.8 mg\selectlanguage{ngerman}·g-1 and 54 U·g-1,
respectively. Compared with free CRL, the thermal and storage stability
of CRL@OSMD was significantly improved, which lays the foundation for
the catalytic synthesis of phytosterol esters in solvent-free systems.
Fortunately, a yield of 95.0\% was achieved after optimizing the
reaction conditions, and a yield of 70.0\% can still be maintained after
6 cycles.%
\end{abstract}\selectlanguage{ngerman}%



\sloppy


\textbf{Octadecyl and sulfonyl modification of diatomite synergistically
improved the immobilization efficiency of lipase and its application in
the synthesis of pine sterol esters}

Yifei Zhang, Wei Zhang, Guangzheng Ma, Binbin Nian* and Yi Hu*

\emph{(State Key Laboratory of Materials-Oriented Chemical Engineering,
School of Pharmaceutical Sciences, Nanjing Tech University, Nanjing,
China)}

\textbf{Abstract}

Phytosterols usually have to be esterified to various phytosterol esters
to avoid their disadvantages of unsatisfactory solubility and low
bioavailability. The enzymatic synthesis of phytosterol esters in
solvent-free system has advantages in terms of environmental
friendliness, sustainability, and selectivity. However, the limitation
of the low stability and recyclability of the lipase in the solvent-free
system, which often requires a relatively high temperature to induce the
viscosity, also increased the industrial production cost. In this
context, a low-cost material, namely diatomite, was employed as the
support in the immobilization of \emph{Candida rugosa} lipase (CRL) due
to its multiple modification sites. The
Fe\textsubscript{3}O\textsubscript{4} was also then introduced to this
system for quick and simple separation via the magnetic field. Moreover,
to further enhance the immobilization efficiency of diatomite, a
modification strategy which involved the octadecyl and sulfonyl group
for regulating the hydrophobicity and interaction between the support
and lipase was successfully developed. The optimization of the ratio of
the modifiers suggested that the
-SO\textsubscript{3}H/C\textsubscript{18} (1:1.5) performed best with an
enzyme loading and enzyme activity of 84.8 mg\selectlanguage{ngerman}·g\textsuperscript{-1} and
54 U·g\textsuperscript{-1}, respectively. Compared with free CRL, the
thermal and storage stability of CRL@OSMD was significantly improved,
which lays the foundation for the catalytic synthesis of phytosterol
esters in solvent-free systems. Fortunately, a yield of 95.0\% was
achieved after optimizing the reaction conditions, and a yield of 70.0\%
can still be maintained after 6 cycles.

\textbf{Keywords:} \emph{Candida rugosa} lipase, diatomite, octadecyl
and sulfonyl modification, immobilization, phytosterol esters

\textbf{Graphical abstract}

\textbf{Introduction}

Sterols are lipids that play a crucial role in all multicellular
eukaryotes. The distribution and synthesis of sterols varies among the
eukaryotic community, with animals primarily producing cholesterol
(C\textsubscript{27}), whereas fungi and plants synthesis sterols with
28 to 29 carbon atoms (C\textsubscript{28} and C\textsubscript{29}),
known as ergosterols and phytosterols{[}1, 2{]}. These differences
reflect the complex evolutionary history of sterol synthesis{[}3{]}.
Phytosterols, which include phytostanols and sterols, are similar in
structure and biological function to cholesterol{[}4, 5{]}. They are
primarily found in vegetable oils, nuts, fruits, grains, and other plant
products{[}6{]}. Phytosterols have been recognized for their various
pharmacological properties, including the potential to lower total and
low-density lipoprotein (LDL) cholesterol levels, thereby reducing the
risk of cardiovascular disease{[}7-9{]}. Other health-promoting effects
of phytosterols include anti-obesity{[}10{]}, anti-diabetic{[}9{]},
anti-microbial{[}11{]}, anti-inflammatory{[}12{]}, and immunomodulatory
effects{[}13{]}. Additionally, it has been strongly suggested that
phytosterols possess anti-cancer properties, as phytosterol-rich diets
may reduce the risk of cancer by 20\%{[}14, 15{]}. However, the high
melting point and low solubility in both water and oils have greatly
limited their further application in food, medicine, and other fields.
To overcome this problem, phytosterol esters are considered suitable
alternatives because they maintain all the excellent properties of
original phytosterol{[}16, 17{]}.

The chemical synthesis of phytosterol esters is industrially feasible,
but the high energy consumption, low selectivity of the reaction, and
the unavoidable by-products of dehydrated sterols limit its further
applications{[}18{]}. Enzymatic catalysis which is performed under mild
operating conditions, has high selectivity and fewer by-products and is
therefore attractive in this field{[}19{]}. In recent years, several
lipases, including Novozym\textsuperscript{@}435{[}20{]}, \emph{Candida
rugosa} lipase{[}21{]}, and others, have demonstrated their ability in
the synthesis of phytosterol esters. However, these reactions usually
were performed in organic solvents, which limited the space-time yield
and were harmful to the enzymes. To overcome the damage of organic
solvent, some environmentally friendly mediums, such as ionic
liquids{[}22{]}, and supercritical carbon dioxide{[}23{]} were applied
in biocatalysis. In this work, the esterification process is performed
in a solvent-free system. However, we just found that the solvent-free
system's high viscosity limited the substrates' mass-transfer effect,
which made the reaction time extremely long. It indicated that the
reaction can only be performed at a relatively higher temperature (50\selectlanguage{english}),
which may not be beneficial to the lipase.

In response to this challenge, the immobilization of enzymes that aimed
to improve stability and recyclability has attracted more and more
interest, and various immobilization methods and supports with low cost,
large specific surface area, and low diffusion limits of substrates,
have been developed in past decades. For instance, diatomite, which is
mainly composed of silicon dioxide (SiO\textsubscript{2}), was widely
used in the immobilization of various enzymes due to its low cost,
porous structure, low density, and chemical inertness. Chen et
al.{[}24{]} suggested that the half-lives of diatomite immobilized of
D-allulose 3-epimerase can be improved to 109 and 124 times than that of
free enzyme at 55\selectlanguage{english} and 60\selectlanguage{english}, respectively. Polyaniline-coated magnetic
diatomite can efficiently immobilize invertase, \selectlanguage{greek}β-\selectlanguage{english}galactosidase, and
trypsin{[}25{]}; \emph{Candida} sp. 99-125 immobilized on diatomite can
be used to produce biodiesel efficiently{[}26{]};
\emph{Burkholderia}lipase immobilized on diatomite can be applied to a
fixed-bed bioreactor and continuous biodiesel conversion{[}27{]}.
However, single immobilization support does not meet the needs of all
enzymes, therefore, for the immobilization of different enzymes,
different modifiers need to be designed to confer a more suitable
microenvironment to the support as well as the strength of interaction
with the enzyme, e.g., basso's study indicated that the octadecyl
functional group can change the hydrophobic microenvironment of lipase,
which can have a significant effect on the enzyme activity{[}28{]}, and
Singh modified the surface of silica nanoparticles with carboxylic acids
with different numbers of alkyl chains, and found that
silica-immobilized lipase modified by stearic acid with more
hydrophobicity has a higher activity{[}29{]}. Moreover, the production
of water during the esterification reactions will drive the reaction
toward hydrolysis and lead to enzyme inactivation via promoting the
aggregation of enzymes{[}30, 31{]}. Additionally, it will also
accelerate the leakage of lipase from the support during reutilization.

Therefore, to develop a low-cost, sustainable, and high-efficiency
enzymatic synthesis strategy for phytosterol esters, the solvent-free
system was applied in the present study. To overcome the challenge of
the low stability of lipase, a cheap inorganic porous material, namely
diatomite was applied as a support for the immobilization
of\emph{Candida rugosa} lipase (CRL). To further enhance its efficiency,
the magnetic nano-Fe\textsubscript{3}O\textsubscript{4} was first
attached to the surface of diatomite via a co-precipitation method.
Moreover, the octadecyl and sulfonyl groups were applied as modifiers of
the diatomite for regulating its hydrophobicity, flexibility, and
interaction and thus improve the efficiency of diatomite. Additionally,
the immobilized enzyme prepared by this method was used in the enzymatic
synthesis of phytosterol esters to test its stability and recyclability.
This study is anticipated to provide a green and efficient innovative
method for the enzymatic synthesis of phytosterol esters.

\textbf{2. Materials and Methods}

\textbf{2.1. Materials}

Pine sterol (91.0\% \selectlanguage{greek}β-\selectlanguage{english}sitosterol, 7.0\% campesterol, 0.5\% stigmasterol,
0.3\% brassicasterol, and 1.2\% others) was kindly donated from Jiangsu
Conat Biological Products Co., Ltd (Jiangsu, China); diatomite (200
mesh) was purchased from Aladdin (Shanghai, China);
FeCl\textsubscript{3}\selectlanguage{ngerman}·6H\textsubscript{2}O,
FeCl\textsubscript{2}·4H\textsubscript{2}O,
3-mercaptopropyltriethoxysilane (97\%), n-octadecyltriethoxysilane
(85\%), Oleic acid (99.0\%) were purchased from Sarn Chemical Technology
Co., Ltd (Shanghai, China).\emph{Candida rugosa} lipase (type VII) was
purchased from Sigma-Aldrich (St. Louis, MO, USA).

\textbf{2.2 Preparation and modification of magnetic diatomite}

4.1 g of FeCl\textsubscript{3}·6H\textsubscript{2}O and 2.6 g of
FeCl\textsubscript{2}·4H\textsubscript{2}O were dissolved in a quantity
of deionized water and mixed ultrasonically. After sufficient shaking,
6.0 g of diatomaceous earth was added and stirred at 80 °C for 30 min
using nitrogen protection. Add ammonia, adjust pH to 10, and continue
mixing for 1 h. Cooling slowly to room temperature and crystallize for 2
h. The precipitate obtained after magnetic field separation was
sequentially washed with deionized water and ethanol. After drying,
magnetic diatomite (MD) was obtained.

1.0 g of MD was added to 50.0 mL of toluene, sonicated, and mixed for
0.5 h. After that, 0.8-2.4 g of 3-mercaptopropyltriethoxysilane and
1.4-4.2 g of n-octadecyltriethoxysilane were added. Using nitrogen
protection, the reaction was heated to reflux for 12 hours. Then, the
diatomite was separated by a magnetic field, washed several times with
ethanol, and the product was dried under vacuum at 45°C to obtain
octadecyl and thiol co-bonded magnetic diatomite. The obtained octadecyl
and thiol co-bonded magnetic diatomite was added to a certain
concentration of H\textsubscript{2}O\textsubscript{2}and stirred at 40°C
for 24 h and then washed with water to remove the excess
H\textsubscript{2}O\textsubscript{2}. After drying under vacuum at 60°C,
octadecyl and sulfonyl co-bonded magnetic diatomite (OSMD) was obtained.

\textbf{2.3 Immobilization of \emph{Candida rugosa} lipase}

The aqueous lipase solution was centrifuged at 4025 xg for 10 min at 4°C
to remove the insoluble material to obtain a homogeneous semi-purified
lipase. The resulting supernatant was separated and used for subsequent
experiments.

As different supports have their suitable immobilization conditions, it
is reasonable to compare the immobilization effect of the supports on
this basis. 0.1 g of modified magnetic diatomite was added to 5 mL of
semi-purified CRL with different concentrations and different pH values
(6.0-8.0) and then stirred at 25-40°C for 4-12 h. After filtration under
reduced pressure, washing, and freeze drying, the immobilized enzyme
product was obtained. Enzyme loading and immobilization efficiency are
important parameters that define the immobilization process. Enzyme
loading is an important parameter to examine the loading capacity of the
support, and immobilization efficiency (immobilization yield) is usually
employed to define the percentage of the enzyme that is immobilized on
or in the support.

The BCA method was used to detect the protein content in the
supernatant. All the measurements were done in triplicate and the error
was less than 5\% in each case, and the following formula was used to
calculate CRL loading and immobilization efficiency.

\begin{equation}
\begin{matrix}\text{CRL\ loading}\left(\frac{\text{mg}}{g}\right)=\frac{C_{i}\times M_{i}-C_{f}\times V_{f}}{M_{f}}\#\left(1\right)\\
\end{matrix}\nonumber \\
 \end{equation}\begin{equation}
\begin{matrix}\text{Immobilization\ efficiency}\left(\%\right)=\frac{C_{i}\times M_{i}-C_{f}{\times V}_{f}}{C_{i}\times M_{i}}\ \times 100\#\left(2\right)\\
\end{matrix}\nonumber \\
 \end{equation}

where \(C_{i}\) (mg/g) is the initial protein concentration in
CRL before immobilization, \(C_{f}\) (mg/mL) is the final
protein concentration in supernatant and washing solution after
immobilization,\(M_{i}\) (mg) is the mass of CRL added to the
buffer solution,\(V_{f}\) (mL) is the total volume of the
supernatant and washing liquid, and \(M_{f}\) (g) is the mass
of the support.

\textbf{2.4} \textbf{Enzyme activity assay}

The expressed activity gives the enzyme activity expressed by the
immobilized enzyme itself and the specific activity shows the effect of
the immobilization process on the enzyme activity, these two parameters
together with the enzyme loading and the immobilization efficiency form
the parameters required to define the immobilization process{[}32{]}.

The p-nitrophenyl palmitate (p-NPP) assay has been widely used to
analyze lipase activity{[}33, 34{]}. One unit (1 U) is defined as the
amount of enzyme required to catalyze the hydrolysis of p-nitrophenyl
palmitate to produce 1 \selectlanguage{greek}μ\selectlanguage{english}mol of p-nitrophenol per minute at 40\selectlanguage{ngerman}°C and pH
7.0. All the measurements were done in triplicate and the error was less
than 5\% in each case.

The activity of CRL (immobilized CRL) and activity recovery were
calculated by the following equation:

\begin{equation}
\begin{matrix}\text{Expressed\ activity\ }\left({Ug}^{-1}\right)=\frac{A\times
10^{6}\times V_{t}}{\varepsilon\times t\times m_{1}}\#\left(3\right)\\
\end{matrix}\nonumber \\
 \end{equation}\begin{equation}
\begin{matrix}\text{Specific\ activity}\ \left({Ug}^{-1}\right)=\frac{A\times 10^{6}\times V_{t}}{\varepsilon\times t\times m_{2}}\#\left(4\right)\\
\end{matrix}\nonumber \\
 \end{equation}\begin{equation}
\begin{matrix}\text{Activity\ recovery\ }\left(\%\right)=\frac{\text{SA}_{1}}{\text{SA}_{0}}\times 100\#\left(5\right)\\
\end{matrix}\nonumber \\
 \end{equation}

Where \(A\) is the absorbance of the samples,
\(V_{t}\) (L) is the volume of the solution,
\(\varepsilon\)(L\selectlanguage{ngerman}·mol\textsuperscript{-1}·cm\textsuperscript{-1}) is
the molar extinction coefficient of p-nitrophenol, \(t\)
(min) is the reaction time, \(m_{1}\) (g) is the mass of
immobilized CRL and\(m_{2}\) (g) is the protein mass of
immobilized CRL,\(\text{SA}_{1}\)(U·g\textsuperscript{-1}) is the
specific activity of immobilization CRL,
\(\text{SA}_{0}\)(U·g\textsuperscript{-1}) is the specific activity of
free CRL.

The following formula was used to calculate relative activity and
residual activity.

\begin{equation}
\begin{matrix}\text{Relative\ activity\ }\left(\%\right)=\frac{A_{t}}{A_{\max}}\times 100\#\left(6\right)\\
\end{matrix}\nonumber \\
 \end{equation}\begin{equation}
\begin{matrix}\text{Residual\ activity\ }\left(\%\right)=\frac{A_{t}}{A_{i}}\times 100\#\left(7\right)\\
\end{matrix}\nonumber \\
 \end{equation}

Where \(A_{t}\) is the absorbance of the samples,
\(A_{\max}\) is the maximum absorbance, and \(A_{i}\)
is the initial absorbance value of the samples.

\textbf{2.5 Characterizations}

Scanning electron microscopy (SEM) was performed on a Hitachi FE-SEM
SU8200 instrument (Japan). Thermogravimetric analysis (TGA) was
performed on a TGA 550 instrument (TA Instruments, USA) at 20-800\selectlanguage{ngerman}°C
under N\textsubscript{2} atmosphere using a heating rate of 20°C/min.
The FTIR spectra of the MD and OSMD samples were measured using Nicolet
iS50 FT-IR (Thermo Scientific, USA). Spectra were taken from 4000 to 400
cm\selectlanguage{english}\textsuperscript{-1} and the resolution of the wavenumber was 2
cm\selectlanguage{english}\textsuperscript{-1}. XPS of raw diatomite and OSMD was carried out by
ESCALAB250Xi (Thermo Scientific, USA). The magnetization of the
immobilized enzyme was carried out by Lake Shore VSM 7307 (Lake Shore,
USA). N2 adsorption and desorption study was performed on the ASAP
2020Plus HD88 (Micromeritics, USA). Water contact angle measurement of
the samples was measured with a KRUSS drop shape analyzer DSA 100
instrument at 25degC for the support and the injecting syringe as well.

\textbf{2.6 Optimization of conditions for the synthesis of pine sterol
oleate}

Using the immobilized CRL as a catalyst, the process conditions such as
the amount of catalyst, the molar ratio of the substrate between pine
sterol and oleic acid, reaction time, and reaction temperature were
optimized for the synthesis of pine sterol oleate by direct
esterification of pine sterol with oleic acid. In a 15 mL reaction tube
on an automatic temperature-controlled magnetic stirrer, 1.0 mmol of
pine sterol, and 4.0-8.0 mmol of oleic acid were added as substrate,
catalyzed by the immobilized enzyme of 5.0-9.0
U*g\textsuperscript{-1}(relative to pine sterol mass), and the reaction
temperature was 35-55degC for 12-60 h.

\textbf{2.7 Gas Chromatography Analysis}

The quantitative analysis of pine sterol and sterol ester was performed
by gas chromatography (GC) with FID detector and dichloromethane as the
solvent, using a DB-5HT (0.1 \selectlanguage{greek}μ\selectlanguage{english}m, 0.25\selectlanguage{ngerman}×15 m) column with
N\textsubscript{2} as the carrier gas and a flow rate of 1.5 mL/min. The
inlet temperature and detector temperature were 350°C, and the initial
temperature of the column temperature chamber was 180°C, ramped up to
240°C at 10°C/min. The sample volume was 1 \selectlanguage{greek}μ\selectlanguage{english}L and the shunt ratio was
20:1.

\textbf{2.8 Substrate scope investigation}

To demonstrate the substrate range of CRL@OSMD, the esterification of
pine sterol with oleic, linoleic, linolenic acid, and mixed acid (oleic,
linoleic, and linolenic acids in equimolar ratios) was studied under
optimized reaction conditions.

\textbf{2.9} \textbf{Process scale-up and reusability studies}

To explore the scalability of the immobilized CRL-catalyzed pine sterol
ester synthesis process, the feed volume was increased up to 10 times
under optimized reaction conditions. The synthesis of pine sterol
easters was investigated for reusability under optimized conditions. At
the end of the reaction, the immobilized enzyme was filtered, separated
from the reaction system, and washed with 5 mL hexane 3 times. After
drying, the resulting immobilized CRL was used directly in the next
batch.

\textbf{3. Results and discussion}

\textbf{3.1 Characterization of modified diatomite}

The surface morphology and elemental analysis of OSMD and CRL were
characterized by SEM (\textbf{Figure 1} ). The Energy Dispersive
Spectroscopy (EDS) image (\textbf{Figure S1} ) showed that the OSMD
surface was rich in C, S, and Fe elements, which proved the success of
the surface modification. After immobilizing CRL, the proportions of S
and N elements increased significantly, indicating that the support
could load CRL efficiently (\textbf{Figure S2} ). The OSMD still
retained the original unique porous structure of diatomite after the
introduction of functional groups and
Fe\textsubscript{3}O\textsubscript{4}. Obviously, the surface of
immobilized CRL showed a rough morphology, indicating that CRL was
efficiently immobilized on the surface of the support.\selectlanguage{english}
\begin{figure}[H]
\begin{center}
\includegraphics[width=0.70\columnwidth]{figures/image2/image2}

\end{center}
\end{figure}

\textbf{Figure 1} . SEM images of OSMD (A, B, C) and CRL@OSMD (D, E, F)

\textbf{Figure 2. A} shows the FTIR spectra of the original diatomite,
MD, and OSMD. According to the FTIR diagram of the raw diatomite, the
absorption peaks of 3622.3 cm\textsuperscript{-1}, 3450.0
cm\textsuperscript{-1,} and 1639.7 cm\textsuperscript{-1} are formed by
the stretching vibration of -OH. The absorption peaks at 1025.2 and
792.4 cm\textsuperscript{-1} correspond to the stretching vibration of
Si-O-Si; the bending vibration of the Si-O bond forms the absorption
peak at 451.2 cm\textsuperscript{-1}. Instead, there are some new peaks
in the FTIR of modified diatomite. The absorption peaks at 2925.5
cm\textsuperscript{-1}, 2850.1 cm\textsuperscript{-1}, and 1463.1
cm\textsuperscript{-1} in the infrared spectrum of modified diatomite
correspond to the stretching vibration and bending vibration of the C-H
bond in -C\textsubscript{18}, which is consistent with the study of Li
et al{[}35{]}. The peak at around 1396.5 cm\textsuperscript{-1} and
1140.0 cm\textsuperscript{-1} is caused by the vibration of S=O in
-SO\textsubscript{3}H. The surface modification of the diatomite with
alkyl groups can be demonstrated by the Si-C associated peaks appearing
at 720.3 cm\textsuperscript{-1}, which is consistent with the work
reported by Xu et al{[}36{]}. All the above results indicated that the
diatomite was successfully modified.

The magnetization of OSMD and CRL@OSMD was measured using a Vibrating
Sample Magnetometer (VSM) at room temperature (\textbf{Figure 2. B} ).
The remanent magnetization and coercivity of OSMD and CRL@OSMD were
almost zero, indicating that the sample materials are soft magnetic
materials with superparamagnetic and outstanding magnetic
properties{[}37{]}. The specific saturation magnetization of OSMD and
CRL@OSMD were 12.2 emu\selectlanguage{ngerman}·g\textsuperscript{-1}and 12.1
emu·g\textsuperscript{-1}, respectively. There is almost no difference
between the two, which indicates that the immobilization process does
not affect the magnetic properties of the support. The exceptional
magnetic response helps to recover the immobilized enzyme under the
application of a magnetic field, providing valuable support for the
reuse of biocatalysts.

In addition, we employed the N\textsubscript{2} adsorption-desorption
isotherms to provide evidence for the successful immobilization of CRL
to the OSMD (\textbf{Figure 2. C} ). As shown in \textbf{Figure 2.D} ,
the overall pore size of CRL@OSMD was significantly decreased compared
to OSMD which may be attributed to that the CRL immobilized into the
pore of OSMD and thus decreased the pore size of it. Moreover, the
results showed that the Brunauer-Emmett-Teller (BET) surface area of
OSMD decreased from 53.6 m\textsuperscript{2}·g\textsuperscript{-1}to
27.2 m\textsuperscript{2}·g\textsuperscript{-1} after the immobilization
process, accompanied by a corresponding reduction in pore volume from
0.227 cm\textsuperscript{3}·g\textsuperscript{-1}to 0.176
cm\textsuperscript{3}·g\textsuperscript{-1} (\textbf{Table S1} ). These
findings indicate that a substantial portion of the enzyme occupies the
diatomite channels, thereby achieving the effective immobilization of
CRL on OSMD.

\textbf{Figure 2} . FTIR spectrum of diatomite before and after
modification (A), VSM plot (B), N\textsubscript{2} adsorption and
desorption isotherms of OSMD and CRL@OSMD (C), BJH-plot (D)

\textbf{Figure 3} compares the XPS spectra of OSMD, MD, and raw
diatomite. The peaks of 100.1 eV (Si 2p), 259.2 eV (O 1s), and 150.5 eV
(Si 2s) are formed by the diatomite, and the peak intensity is
significantly weakened after modification, which corroborates the change
in the surface groups of the diatomite. Relatively, the peak intensity
of C 1s (281.8 eV) was significantly enhanced after modification.
Despite the weak intensity, the peak at around 161.0 eV (S 2p) and 225.0
eV (S 2s) is caused by -SO\textsubscript{3}H. Compared to raw diatomite,
nanomagnetic Fe\textsubscript{3}O\textsubscript{4}increased the peaks
located at 710.5 eV and 719.9 eV{[}35{]}.\textbf{Table 1} shows that the
C peak intensity of modified diatomite is significantly enhanced, and
the content of C is increased from 15.8\% to 66.1\%. However, the
intensity of the O1s peak weakened, and the O content decreased from
61.8\% to 22.6\%. It is proved that the coating contains a large amount
of -C\textsubscript{18}, and -SO\textsubscript{3}H successfully modified
diatomite. In the high-resolution diagram of Si 2p, the electron
absorption peaks at 100.4 eV and 99.8 eV correspond to the Si-O bond and
Si-Si bond in diatomite{[}38{]}. Compared with unmodified diatomite and
MD, the absorption peak at 98.9 eV corresponds to the Si-C, which
demonstrates the surface modification of the diatomite with octadecyl
groups{[}39{]}.

\textbf{Hosted file}

\verb`image4.emf`  available at \url{https://authorea.com/users/697876/articles/685852-octadecyl-and-sulfonyl-modification-of-diatomite-synergistically-improved-the-immobilization-efficiency-of-lipase-and-its-application-in-the-synthesis-of-pine-sterol-esters}

\textbf{Figure 3} . XPS diagram of diatomite before and after
modification (A), XPS Si 2p spectra of Diatomite (B), MD (C), OSMD (D)

\textbf{Table 1} Relative atomic content of the surface before and after
modification\selectlanguage{english}
\begin{longtable}[]{@{}lllll@{}}
\toprule
& Si 2p (\%) & C 1s (\%) & O 1s (\%) & N 1s (\%)\tabularnewline
\midrule
\endhead
Diatomite & 21.8 & 15.8 & 61.8 & 0.7\tabularnewline
MD & 10.7 & 33.4 & 54.7 & 1.1\tabularnewline
OSMD & 11.3 & 66.1 & 22.5 & 0.1\tabularnewline
\bottomrule
\end{longtable}

As shown in the DTG results, MD reaches the point of maximum thermal
decomposition rate at 362.3\selectlanguage{ngerman}°C, which indicates the dehydration of
surface hydroxyl groups, which is consistent with the OSMD{[}40{]}. OSMD
shows another jump discontinuity at 561.8°C, which may be due to the
loss of grafted octadecyl and sulfonyl groups{[}41{]}. Throughout the
process, the overall mass loss of MD was 7.6\%, while the overall mass
loss of OSMD was 8.6\%, this indicated that diatomite was successfully
modified, while the support itself has excellent thermal stability.

The contact angle serves as a quantitative indicator of surface
wettability, which quantifies the angle formed between the interface of
the liquid or vapor and the solid surface. According to the
Young-According to the Young-Laplace equation, the morphology of a
droplet is influenced by the relative hydrophobic or hydrophilic
properties of the material's surface with respect to the liquid. A
larger contact angle corresponds to a greater repulsive force exerted by
the droplet on the surface, signifying a relatively higher degree of
hydrophobicity of the material's surface{[}24{]}. As depicted
in\textbf{Figure 3D (a)} , diatomaceous earth exhibits pronounced
hydrophilicity owing to the abundance of surface Si-OH groups{[}41{]}.
Conversely, the contact angle of OSMD increased to 133.5° following
additional octadecyl modification, indicative of its heightened
hydrophobicity, which can also support that the successful modification
of octadecyl.

\textbf{Figure 4} . TG of MD and OSMD (A), DTG of MD (B) and OSMD (C),
the contact angle of MD and OSMD (D)

\textbf{3.2 Optimization of the ratio of modifiers and surfactants}

Although different modification degrees of modifiers
(-SO\textsubscript{3}H and -C\textsubscript{18}) can result in distinct
properties of the carrier and thereby impact its immobilization
efficiency, quantitatively analyzing the modification degree can be
exceedingly challenging. Thus, to further investigate the effect of the
modification degree on the immobilization efficiency and to design a
more efficient immobilization carrier, we used the immobilization effect
of CRL as an indicator to optimize the modification degree. As shown
in\textbf{Table 2} , the results suggested that the enzyme loading and
activity recovery of modified diatomite modified with
-SO\textsubscript{3}H or -C\textsubscript{18} alone was considerably
lower than that of -SO\textsubscript{3}H/-C\textsubscript{18} (1:1)
group, which demonstrated that the -SO\textsubscript{3}H and
-C\textsubscript{18}groups may have a synergistic effect. It can be
attributed to several factors, on the one side, the long-chain
-C\textsubscript{18} provided a suitable hydrophobic microenvironment
for the lipase via an interfacial activation mechanism, and on the other
hand, the -SO\textsubscript{3}H groups changed the charge distribution
of the lipase surface, resulting in a stronger binding interaction
between lipase and carrier, which could reduce the enzyme leakage. In
addition, the ratio of the two modifiers had a significant effect on the
immobilization effect, and OSMD had the best immobilization effect with
an enzyme loading of 74.40 mg·g\textsuperscript{-1} and an activity
recovery of 63.35\% at the ratio of
-SO\textsubscript{3}H/-C\textsubscript{18}(1:1.5). The specific
mechanism needs to be further investigated to find out.

Given the high hydrophobicity of OSMD, the addition of surfactant
enhances the dispersion of the support in the phosphate buffer{[}42,
43{]}. In addition, the addition of surfactants can also prevent the
aggregation of lipase{[}44{]}. However, it also should be noted that the
surfactants may also play a negative role during immobilization due to
the inactivation of lipases. Hence, it becomes crucial to screen the
type and concentration of surfactant (\textbf{Figure S4} ). The results
showed that SDS has a detrimental effect on lipase activity. Nonionic
surfactants (Triton X-100, Tween 80) yielded favorable results but were
slightly inferior to the cationic surfactant (CTAB). Moreover, the
impact of different CTAB concentrations on immobilization efficacy was
assessed, revealing that at a concentration of 0.50 mM, CRL loading
reached 76.3 mg·g\textsuperscript{-1}, with an enzyme activity recovery
of 74.7\%. The presence of the cationic surfactant may alter the surface
charge of the lipase and enhance its electrostatic interactions with the
sulfonic acid groups while preventing lipase dimer formation and thus
beneficial to the immobilization processes{[}45, 46{]}.

\textbf{Table 2} . Effect of modifier ratio on the immobilization effect
of OSMD\selectlanguage{english}
\begin{longtable}[]{@{}lll@{}}
\toprule
\begin{minipage}[b]{0.32\columnwidth}\raggedright\strut
Modifier ratio\strut
\end{minipage} & \begin{minipage}[b]{0.32\columnwidth}\raggedright\strut
Enzyme loading (mg\selectlanguage{ngerman}·g\textsuperscript{-1})\strut
\end{minipage} & \begin{minipage}[b]{0.32\columnwidth}\raggedright\strut
Activity recovery (\%)\strut
\end{minipage}\tabularnewline
\midrule
\endhead
\begin{minipage}[t]{0.30\columnwidth}\raggedright\strut
-SO\textsubscript{3}H\strut
\end{minipage} & \begin{minipage}[t]{0.30\columnwidth}\raggedright\strut
32.62 \selectlanguage{ngerman}± 1.03\strut
\end{minipage} & \begin{minipage}[t]{0.30\columnwidth}\raggedright\strut
32.37 \selectlanguage{ngerman}± 1.68\strut
\end{minipage}\tabularnewline
\begin{minipage}[t]{0.30\columnwidth}\raggedright\strut
-SO\textsubscript{3}H/-C\textsubscript{18}(1:0.5)\strut
\end{minipage} & \begin{minipage}[t]{0.30\columnwidth}\raggedright\strut
56.77 \selectlanguage{ngerman}± 1.57\strut
\end{minipage} & \begin{minipage}[t]{0.30\columnwidth}\raggedright\strut
40.58 \selectlanguage{ngerman}± 0.79\strut
\end{minipage}\tabularnewline
\begin{minipage}[t]{0.30\columnwidth}\raggedright\strut
-SO\textsubscript{3}H/-C\textsubscript{18}(1:1.0)\strut
\end{minipage} & \begin{minipage}[t]{0.30\columnwidth}\raggedright\strut
63.14 \selectlanguage{ngerman}± 1.29\strut
\end{minipage} & \begin{minipage}[t]{0.30\columnwidth}\raggedright\strut
52.16 \selectlanguage{ngerman}± 1.34\strut
\end{minipage}\tabularnewline
\begin{minipage}[t]{0.30\columnwidth}\raggedright\strut
-SO\textsubscript{3}H/-C\textsubscript{18}(1:1.5)\strut
\end{minipage} & \begin{minipage}[t]{0.30\columnwidth}\raggedright\strut
74.40 \selectlanguage{ngerman}± 1.61\strut
\end{minipage} & \begin{minipage}[t]{0.30\columnwidth}\raggedright\strut
63.35 \selectlanguage{ngerman}± 1.48\strut
\end{minipage}\tabularnewline
\begin{minipage}[t]{0.30\columnwidth}\raggedright\strut
-SO\textsubscript{3}H/-C\textsubscript{18}(1:2.0)\strut
\end{minipage} & \begin{minipage}[t]{0.30\columnwidth}\raggedright\strut
65.95 \selectlanguage{ngerman}± 2.12\strut
\end{minipage} & \begin{minipage}[t]{0.30\columnwidth}\raggedright\strut
60.54 \selectlanguage{ngerman}± 1.61\strut
\end{minipage}\tabularnewline
\begin{minipage}[t]{0.30\columnwidth}\raggedright\strut
-C\textsubscript{18}\strut
\end{minipage} & \begin{minipage}[t]{0.30\columnwidth}\raggedright\strut
60.51 \selectlanguage{ngerman}± 0.66\strut
\end{minipage} & \begin{minipage}[t]{0.30\columnwidth}\raggedright\strut
54.19 \selectlanguage{ngerman}± 0.86\strut
\end{minipage}\tabularnewline
\begin{minipage}[t]{0.30\columnwidth}\raggedright\strut
MD\strut
\end{minipage} & \begin{minipage}[t]{0.30\columnwidth}\raggedright\strut
22.88 \selectlanguage{ngerman}± 0.68\strut
\end{minipage} & \begin{minipage}[t]{0.30\columnwidth}\raggedright\strut
24.57 \selectlanguage{ngerman}± 0.85\strut
\end{minipage}\tabularnewline
\bottomrule
\end{longtable}\selectlanguage{ngerman}

Conditions of the immobilization process: the reaction was carried out
in 0.1 mol·L\textsuperscript{-1} phosphate buffer (pH 7.0) with enzyme
addition of 100 mg·g\textsuperscript{-1} (mass of support) for 8 h at
30°C.

\textbf{3.3Optimization of\emph{Candida rugosa} lipase immobilization
conditions}

Lipase immobilization efficiency on hydrophobic supports is generally
influenced by enzyme concentration (enzyme addition), pH, temperature,
and reaction time{[}47{]}. In investigating the factor of enzyme
addition (\textbf{Figure 5. A} ), the results showed that the enzyme
loading increased with the enzyme addition. However, considering the
enzyme is relatively expensive, even though the enzyme loading and
activity of the CRL@OSMD were still slightly increased at more than 100
mg·g\textsuperscript{-1}\textsubscript{,} the optimum enzyme addition
was set to 100 mg·g\textsuperscript{-1}. As shown in\textbf{Figure 5. B}
, the higher temperature would increase the collision between enzyme
molecules and support, thus improving the immobilization rate, but too
high temperature will also affect the stability of enzymes in the buffer
solution. Taking the CRL loading and enzyme activity together as the
evaluation index, 30°C was finally selected as the optimal
immobilization temperature.

The pH of the buffer solution affects the surface charge of the CRL
(\textbf{Figure 5. C} ), which in turn affects the electrostatic
interaction of the enzyme protein with the sulfonic acid groups in the
support. The results showed that the immobilization efficiency and
enzyme activity were significantly increased at pH 6.5, indicating that
the amino group of CRL is more likely to interact electrostatically with
the sulfonic acid group after ion exchange under weakly acidic
conditions.

Finally, the time was optimized and the results showed that the longer
the immobilization process (\textbf{Figure 5.D} ), the higher the
loading of CRL@OSMD, but by 10 h later, the support was saturated.
Continued increase in time, on the contrary, led to a decrease in enzyme
activity.

Finally, immobilized CRL with a loading of 84.8 mg·g\textsuperscript{-1}
and enzyme activity of 54 U·g\textsuperscript{-1} was obtained after the
reaction at 30°C for 10 h in phosphate buffer at pH 6.5 with an enzyme
addition of 100 mg·g\textsuperscript{-1}\textsubscript{,}which is
significantly higher than that of some previous studies. For instance,
Cabrera et al. studied the immobilization of CALB in a series of
hydrophobic supports and found that only 30
mg·g\textsuperscript{-1}loading was obtained with octadecyl resin as the
support{[}48{]}. Similarly, Kurtovic et al. adsorbed CRL onto a highly
hydrophobic octadecyl methacrylate resin by interfacial activation and
only obtained an immobilized enzyme with a loading of 58.7
mg·g\textsuperscript{-1}{[}49{]}. Thus, in contrast to many existing
reports, OSMD exhibited its extreme advantages in enzyme loading.

\textbf{Figure 5.}Optimization of immobilization enzyme addition (A),
temperature (B), pH (C), and time (D) of CRL@OSMD

\textbf{3.4 Thestability study of immobilized CRL}

To investigate the effect of temperature on CRL@OSMD, the residual
enzyme activity of free CRL and CRL@OSMD at 50°C and 60°C were measured
with the p-NPP assay. As illustrated in \textbf{Figure S5} , as the
temperature increased, the residual activity of both free CRL and
CRL@OSMD exhibited a diminishing trend. However, it is noteworthy that
the relative activity of CRL@OSMD remained conspicuously higher than
that of free CRL. These findings suggested that under the experimental
conditions, both CRL@OSMD and free CRL retained their activities at
levels exceeding 90\% when subjected to a 50°C treatment for 1
h.\textbf{Figure S5} shows that the residual activity of free CRL and
CRL@OSMD decreased with increasing temperature, but the relative
activity of CRL@OSMD was significantly higher than that of free CRL. The
results indicated that both the activity of CRL@OSMD and free CRL can be
maintained above 90\% at 50°C treatment for 1 h. However, when the
temperature was increased to 60°C, a more pronounced decline in the
activity of free CRL was observed, which may be due to the irreversible
denaturation and inactivation of the secondary structure of CRL by the
higher temperature. In stark contrast, CRL@OSMD exhibited remarkable
resilience, maintaining its activity above 80\% even after 30 minutes of
exposure to the heightened temperature of 60°C. This observation serves
as compelling evidence that CRL@OSMD boasts superior thermal stability
compared to its free counterpart under the specified experimental
conditions.

To investigate the storage stability of the immobilized enzyme, free CRL
and immobilized CRL (CRL@OSMD) were stored at 4°C for one month and
residual enzyme activity was tested every 5 days at the respective
optimum pH and temperature and the initial enzyme activity was defined
as 100\%. As shown in\textbf{Figure S5} , the enzyme activity of free
CRL decreased rapidly to less than 50.0\% after 10 days, and only 11.3\%
residual enzyme activity can be maintained after 30 days. In contrast,
the storage stability of the OSMD immobilized enzyme was significantly
higher than free CRL. After 5 days of storage, 93.2\% of the initial
enzyme activity was still preserved, and this level of activity endured
above 50.0\% even after the 30-day storage duration. These results
unequivocally underscore the outstanding storage characteristics of the
immobilized enzyme, highlighting its impressive reusability and ability
to endure extended storage periods, making it well-suited to meet the
demands of production requirements.

When p-NPP was used as a model substrate to test the repetitive
hydrolysis activity of the immobilized enzyme, it was found that the
hydrolysis effect of CRL@OSMD could still reach 86.2\% after 10 times of
reuse (\textbf{Figure S6} ), which indicated that the immobilized enzyme
had good operational stability and repetitive use performance.

\textbf{3.5} \textbf{Synthesis of pine sterol oleate with OSMD@CRL in a
solvent-free system}

Oleic acid, as a common monounsaturated \selectlanguage{greek}ω-\selectlanguage{english}9 fatty acid, works
synergistically with phytosterols to regulate blood lipid levels and
effectively reduce hypercholesterolemia and cardiovascular
disease{[}50{]}. In addition, our previous studies also found that oleic
acid is one of the most efficient substrates for the esterification
reaction due to its low melting point and its high solubility of
phytosterols{[}50{]} (\textbf{Figure 6} ). With excess oleic acid as a
solvent, the additional costs and environmental concerns associated with
the use of organic solvents can be avoided{[}51, 52{]}. Optimization of
solvent-free esterification involves obtaining high conversions while
avoiding excess reagents and catalysts and saving energy{[}53{]}.
However, solvent-free reactions present specific challenges given the
drastic changes that can occur in the reaction medium during the
reaction processes{[}54{]}. Specific studies are needed to determine the
optimal amounts of reagents and catalysts and the optimal temperatures
under these conditions, where the thermodynamic and kinetic aspects
converge toward high conversion{[}50{]}.

Most enzymatic esterification studies consider the molar ratio of
reagents, biocatalyst addition, reaction time, and temperature as the
main variables that determine the reaction yield{[}50{]}. Moreover,
there are two possible strategies to change the esterification
equilibrium: (1) use an excess amount of one of the reagents or (2)
remove one of the product mediators (water) from the reaction{[}55{]}.
As the relatively low cost of oleic acid and it can act as the solvent
of the solvent-free system, the ratio of oleic acid: pine sterol was
selected from 1:4 to 1:8. As shown in the results of \textbf{Figure 6.
A} , the yield can only reach 53.4\% and 65.7\% at ratios of 1:4 and 1:5
which can largely be attributed to the incomplete dissolution of pine
sterol in the reaction medium. Nonetheless, it's worth noting that
excessive quantities of oleic acid can also exert a detrimental
influence on esterification efficiency. This is likely attributable to a
reduction in the relative concentration of pine sterol within the oleic
acid medium, leading to a diminished likelihood of substrate access to
the enzyme's active center. The optimal esterification yield was
achieved when the molar ratio of pine sterol to oleic acid was
maintained at 1:6. This ratio takes into consideration the intricate
dynamics introduced by the reaction medium, resulting in the highest
efficiency.

The amount of biocatalyst in the reaction is limited by the dispersing
capacity of the stirring system and the filtration capacity of the
system{[}56-58{]}. The amount of enzyme addition in the system does not
affect the final yield of the ester in equilibrium, but it does affect
the reaction rate. Reaction product-induced lipase inhibition may occur
under specific circumstances of the reaction{[}55, 59{]}. As shown in
the results of \textbf{Figure 6. B} , the lower yield of the reaction at
an enzyme addition of 5.0 U\selectlanguage{ngerman}·g\textsuperscript{-1} (relative to the mass
of pine sterol) is most likely due to a decrease in the rate of the
reaction, as well as product inhibition. The efficiency of the
esterification reaction remained essentially the same at enzyme
additions above 8.0 U·g\textsuperscript{-1}, which was finally chosen
given the high cost of biocatalysts.

The temperature has a positive effect on the energy of the reagents,
favoring the effective number of collisions leading to product
formation{[}60-62{]}. However, collateral effects may occur - high
temperatures may cause conformational changes in the enzyme, leading to
an increase (or loss) of enzyme catalytic activity{[}61, 63{]}. Thus,
incremental increases in reaction kinetics due to increased temperatures
may be offset by reductions in the catalytic activity of
lipases{[}64{]}. Temperature also affects the solubility of the reagents
and viscosity reduction, which means that there are considerable changes
in the reaction medium, and these changes affect the apparent
equilibrium position because only dissolved reactants are involved in
the thermodynamic process, and only dissolved substrates can be
contacted by the enzyme{[}65{]}. The optimal temperature for enzymatic
esterification should facilitate proper diffusion of the reagents in the
medium, thus maintaining the performance of the biocatalyst. As shown
in\textbf{Figure 6. C} , the low yield at 35°C was attributed to the
poor solubilization of the pine sterols and the high mass transfer
resistance of the medium due to the high viscosity of oleic acid at a
low temperature, which resulted in the diffusion of substrates to the
active site of CRL@OSMD became more difficult. In addition, the
temperature above 55°C may also have a negative effect on the
esterification yield due to the denature of CRL@OSMD. Taking into
account the catalytic activity of the immobilized enzyme and the
efficiency of the esterification reaction, 50°C was finally selected as
the optimum reaction temperature. Reaction time affects the efficiency
of product production and reduces the number of times the biocatalyst
can be recycled, and reaching reaction equilibrium cannot always be
pursued to maximize the benefits of immobilized enzyme-catalyzed
esterification. The esterification reaction yield can reach 95.0\% at 48
h (\textbf{Figure 6.D} ), and further increasing the reaction time did
not have a significant effect on the esterification yield. Moreover,
only a conversion of 91.1\% of phytosterols can be obtained even though
the dry air was introduced to regulate water activity at 72 h{[}66{]},
which may increase the production cost. Fortunately, in this study, we
found that our modified diatomite with long-chain alkyls, namely OSMD,
can prevent the water molecules from adhering to the surface and thus
beneficial to the reaction without dry air treatment.

\textbf{Figure 6.} Optimization of molar ratio (A), enzyme addition (B),
temperature (C) and time (D) of the esterification reaction

\textbf{3.} \textbf{6Substrate scope investigation of immobilized CRL}

To investigate the substrate scope of the immobilized lipase (CRL@OSMD),
several medium or long-chain fatty acids, such as linoleic acid (C18:2),
linolenic acid (C18:3), lauric acid (C12:0), and decanoic acid (C10:0)
were selected as the acyl donors. As summarized in \textbf{Table 3} ,
both saturated and unsaturated fatty acids can be used as the acyl
donors of the esterification of pine sterol with phytosterol conversions
at close to or above 90\%. However, as the high melting point of the
long-chain (above C12) saturated fatty acids are not beneficial to the
solubility of pine sterol and the transfer mass effect of the reactions,
only lauric acid and decanoic acid were selected as the model of
saturated fatty acid with a yield of 92.33 ± 1.29\% and 90.45 ± 1.70\%.
To further promote the industrial production processes, edible oil,
which contains oleic acid, linoleic acid, and linolenic acid was also
selected as the substrate in the synthesis of pine sterol esters, and
the results showed that the yield can reach as high as 94.14 ± 1.37\%.
This indicates that our novel-designed immobilized enzyme (CRL@OSMD) can
catalyze the esterification reaction of pine sterols with excellent
substrate applicability, which can be adapted to the needs of processing
and production.

\textbf{Table 3} . Substrate scope investigation\selectlanguage{english}
\begin{longtable}[]{@{}ll@{}}
\toprule
\begin{minipage}[b]{0.48\columnwidth}\raggedright\strut
\textbf{Fatty Acid}\strut
\end{minipage} & \begin{minipage}[b]{0.48\columnwidth}\raggedright\strut
\textbf{Conversion rate} \textbf{(\%)}\strut
\end{minipage}\tabularnewline
\midrule
\endhead
\begin{minipage}[t]{0.47\columnwidth}\raggedright\strut
Oleic acid (C18:1)\textsuperscript{b}\strut
\end{minipage} & \begin{minipage}[t]{0.47\columnwidth}\raggedright\strut
95.01 \selectlanguage{ngerman}± 0.87\strut
\end{minipage}\tabularnewline
\begin{minipage}[t]{0.47\columnwidth}\raggedright\strut
Linoleic acid (C18:2)\strut
\end{minipage} & \begin{minipage}[t]{0.47\columnwidth}\raggedright\strut
94.50 \selectlanguage{ngerman}± 0.93\strut
\end{minipage}\tabularnewline
\begin{minipage}[t]{0.47\columnwidth}\raggedright\strut
Linolenic acid (C18:3)\strut
\end{minipage} & \begin{minipage}[t]{0.47\columnwidth}\raggedright\strut
93.80 \selectlanguage{ngerman}± 0.49\strut
\end{minipage}\tabularnewline
\begin{minipage}[t]{0.47\columnwidth}\raggedright\strut
Lauric acid (C12:0)\strut
\end{minipage} & \begin{minipage}[t]{0.47\columnwidth}\raggedright\strut
90.45 \selectlanguage{ngerman}± 1.70\strut
\end{minipage}\tabularnewline
\begin{minipage}[t]{0.47\columnwidth}\raggedright\strut
Decanoic acid (C10:0)\strut
\end{minipage} & \begin{minipage}[t]{0.47\columnwidth}\raggedright\strut
92.33 \selectlanguage{ngerman}± 1.29\strut
\end{minipage}\tabularnewline
\begin{minipage}[t]{0.47\columnwidth}\raggedright\strut
Mixed acids\strut
\end{minipage} & \begin{minipage}[t]{0.47\columnwidth}\raggedright\strut
94.14 \selectlanguage{ngerman}± 1.37\strut
\end{minipage}\tabularnewline
\bottomrule
\end{longtable}\selectlanguage{ngerman}

\textsuperscript{a} Reaction conditions: 1 mmol pine sterol, 6 mmol
fatty acid, 8 U/g (based on the mass of pine sterol) CRL@OSMD, 50°C, 48
h.

\textsuperscript{b} The first number in parentheses represents the
number of carbon atoms of the fatty acid, and the second number
represents the number of carbon-carbon double bonds of the fatty acid.

\textsuperscript{c} Mixing oleic, linoleic, and linolenic acids in
equimolar ratios to simulate edible oils.

\textbf{3.7 Process scale-up andreusability studies}

To explore the scalability of the pine sterol ester synthesis process
developed above, the amount of substrate was scaled up to 10 times, and
a final esterification of 94.3\% of pine sterol ester was obtained,
demonstrating the scalable production potential of the new process for
the synthesis of pine sterol esters catalyzed by the immobilized enzymes
in the study. Moreover, the reusability of the catalyst was investigated
under the optimized conditions, and the results are shown
in\textbf{Figure S6} . It can be seen that the esterification yield
decreased with the increase of cycles, and could still be maintained
above 70.9\% after 6 cycles. The decrease in esterification yield is
most likely due to the unavoidable loss of the immobilized enzyme during
the separation from the reaction system, as well as denaturation and
inactivation of the enzyme during the reaction. For instance, Zhang et
al. suggested that only 82.0\% activity of CRL immobilized with a hollow
cubic carbon can be maintained in the second cycle{[}67{]}. However, in
this study, our novel-designed OSMD immobilization can maintain the
residual activity as high as 90.6\% in two cycles, which suggests that
the novel-designed OSMD in this study is a good candidate in lipase
immobilization and has great potential for application in further food
biomanufacturing.

\textbf{4. Conclusion}

A low-cost and designable material, namely diatomite, was investigated
in the immobilization of CRL and then used for the synthesis of various
pine sterol esters. To further improve its efficiency, sulfonyl and
octadecyl were employed as modifiers to regulate the hydrophobicity and
the binding interaction between support and lipase. In addition, the
Fe\textsubscript{3}O\textsubscript{4} was also introduced to this system
for quick and simple separation via the magnetic field. Interestingly,
the enzyme loading and activity recovery of the single modification
diatomite via octadecyl or sulfonyl group was lower than that of two
functional groups modification diatomite, namely OSMD. It indicated that
there may be a synergistic effect between these two modifiers, the
octadecyl modification endowed a suitable hydrophobicity of diatomite,
and the sulfonyl group changed the interaction between lipase and
support which enhanced the enzyme loading of CRL. The optimization of
the ratio of the modifiers suggested that the
-SO\textsubscript{3}H/C\textsubscript{18} (1:1.5) performed best with an
enzyme loading and enzyme activity of 84.8 mg·g\textsuperscript{-1} and
54 U·g\textsuperscript{-1}, respectively. Compared to free CRL, the
thermal and storage stability of CRL@OSMD was significantly improved,
which lays the foundation for the catalytic synthesis of phytosterol
esters in a solvent-free system. The molar ratio of pine sterol and
oleic acid, enzyme addition, reaction temperature, and time in the
esterification reaction were optimized, and finally the reaction was
carried out at a molar ratio of pine sterol to oleic acid of 1:6, an
enzyme addition of 8.0 U·g\textsuperscript{-1} (concerning the mass of
pine sterol), and at a temperature of 50°C for 48 h, which could
ultimately achieve an esterification yield of 95.0\%. After cycling the
reaction 6 times, the esterification yield still reached more than
70.0\%. This study shows that OSMD materials are good candidates for
lipase immobilization and have great potential for application in
further food biomanufacturing with low-cost and high efficiency.

\textbf{ACKNOWLEDGEMENT}

This research was financially supported by the National key R\&D program
of China (No.2021YFC2103800), National Natural Science Foundation of
China (No. 22178174), Jiangsu Synergetic Innovation Center for Advanced
Bio-Manufacture (XTC2206).

\textbf{AUTHOR INFORMATION}

Corresponding author:

Yi Hu, E-mail:\emph{huyi@njtech.edu.cn}; Binbin Nian,
E-mail:\emph{bbnian@njtech.edu.cn}.

\textbf{Notes}

The authors declare no competing financial interests.

\textbf{References:}

{[}1{]} Lagarda, M., García-Llatas, G., Farré, R., Analysis of
phytosterols in foods. \emph{Journal of pharmaceutical and biomedical
analysis} 2006, \emph{41} , 1486-1496.

{[}2{]} Weete, J. D., Abril, M., Blackwell, M., Phylogenetic
distribution of fungal sterols. \emph{PloS one} 2010, \emph{5} , e10899.

{[}3{]} Michellod, D., Bien, T., Birgel, D., Violette, M.\emph{, et al.}
, De novo phytosterol synthesis in animals. \emph{Science}
2023,\emph{380} , 520-526.

{[}4{]} Piironen, V., Lindsay, D. G., Miettinen, T. A., Toivo, J.,
Lampi, A. M., Plant sterols: biosynthesis, biological function and their
importance to human nutrition. \emph{Journal of the Science of Food and
Agriculture} 2000, \emph{80} , 939-966.

{[}5{]} Salehi-Sahlabadi, A., Varkaneh, H. K., Shahdadian, F., Ghaedi,
E.\emph{, et al.} , Effects of Phytosterols supplementation on blood
glucose, glycosylated hemoglobin (HbA1c) and insulin levels in humans: a
systematic review and meta-analysis of randomized controlled
trials.\emph{Journal of Diabetes \& Metabolic Disorders} 2020, \emph{19}
, 625-632.

{[}6{]} Moreau, R. A., Whitaker, B. D., Hicks, K. B., Phytosterols,
phytostanols, and their conjugates in foods: structural diversity,
quantitative analysis, and health-promoting uses. \emph{Progress in
lipid research} 2002, \emph{41} , 457-500.

{[}7{]} Bacchetti, T., Masciangelo, S., Bicchiega, V., Bertoli, E.,
Ferretti, G., Phytosterols, phytostanols and their esters: from natural
to functional foods. \emph{Mediterranean Journal of Nutrition and
Metabolism} 2011, \emph{4} , 165-172.

{[}8{]} Dash, R., Mitra, S., Ali, M. C., Oktaviani, D. F.\emph{, et al.}
, Phytosterols: Targeting neuroinflammation in
neurodegeneration.\emph{Current Pharmaceutical Design} 2021, \emph{27} ,
383-401.

{[}9{]} Hannan, M. A., Sohag, A. A. M., Dash, R., Haque, M. N.\emph{, et
al.} , Phytosterols of marine algae: Insights into the potential health
benefits and molecular pharmacology. \emph{Phytomedicine} 2020,\emph{69}
, 153201.

{[}10{]} Kury\selectlanguage{polish}ł\selectlanguage{english}owicz, A., C\selectlanguage{polish}ą\selectlanguage{english}ka\selectlanguage{polish}ł\selectlanguage{english}a-Jakimowicz, M., Puzianowska-Ku\selectlanguage{polish}ź\selectlanguage{english}nicka,
M., Targeting abdominal obesity and its complications with dietary
phytoestrogens. \emph{Nutrients} 2020, \emph{12} , 582.

{[}11{]} Bur\selectlanguage{polish}č\selectlanguage{english}ov\selectlanguage{ngerman}á, Z., Kreps, F., Greifová, M., Jablonský, M.\emph{, et
al.} , Antibacterial and antifungal activity of phytosterols and methyl
dehydroabietate of Norway spruce bark extracts. \emph{Journal of
biotechnology} 2018, \emph{282} , 18-24.

{[}12{]} Chen, L., Deng, H., Cui, H., Fang, J.\emph{, et al.} ,
Inflammatory responses and inflammation-associated diseases in
organs.\emph{Oncotarget} 2018, \emph{9} , 7204.

{[}13{]} Scolaro, B., Andrade, L. F. d., Castro, I. A., Cardiovascular
disease prevention: The earlier the better? A review of plant sterol
metabolism and implications of childhood
supplementation.\emph{International Journal of Molecular Sciences} 2019,
\emph{21} , 128.

{[}14{]} Ramprasath, V. R., Awad, A. B., Role of phytosterols in cancer
prevention and treatment. \emph{Journal of AOAC International}
2015,\emph{98} , 735-738.

{[}15{]} Suttiarporn, P., Chumpolsri, W., Mahatheeranont, S.,
Luangkamin, S.\emph{, et al.} , Structures of phytosterols and
triterpenoids with potential anti-cancer activity in bran of black
non-glutinous rice. \emph{Nutrients} 2015, \emph{7} , 1672-1687.

{[}16{]} Yang, Y., He, W., Jia, C., Ma, Y.\emph{, et al.} , Efficient
synthesis of phytosteryl esters using the Lewis acidic ionic
liquid.\emph{Journal of Molecular Catalysis A: Chemical} 2012,
\emph{357} , 39-43.

{[}17{]} Tan, Z., Le, K., Moghadasian, M., Shahidi, F., Enzymatic
synthesis of phytosteryl docosahexaneates and evaluation of their
anti-atherogenic effects in apo-E deficient mice. \emph{Food
chemistry}2012, \emph{134} , 2097-2104.

{[}18{]} Shi, H., Li, S., Liu, J., Wang, S.\emph{, et al.} , Green and
efficient synthesis of pine sterol oleate catalyzed by
SO\textsubscript{3}H-functionalized ionic liquid. \emph{Journal of
Chemical Technology \& Biotechnology} 2022, \emph{97} , 3083-3090.

{[}19{]} Lortie, R., Enzyme catalyzed esterification.\emph{Biotechnology
Advances} 1997, \emph{15} , 1-15.

{[}20{]} Maldonado, R. R., Burkert, J. F. M., Mazutti, M. A., Maugeri,
F., Rodrigues, M. I., Evaluation of lipase production by Geotrichum
candidum in shaken flasks and bench-scale stirred bioreactor using
different impellers. \emph{Biocatalysis and Agricultural
Biotechnology}2012, \emph{1} , 147-151.

{[}21{]} Zheng, M.-M., Lu, Y., Dong, L., Guo, P.-M.\emph{, et al.} ,
Immobilization of \emph{Candida rugosa} lipase on hydrophobic/strong
cation-exchange functional silica particles for biocatalytic synthesis
of phytosterol esters. \emph{Bioresource Technology} 2012, \emph{115} ,
141-146.

{[}22{]} Zeng, C., Qi, S., Li, Z., Luo, R.\emph{, et al.} , Enzymatic
synthesis of phytosterol esters catalyzed by \emph{Candida rugosa}lipase
in water-in-{[}Bmim{]}PF\textsubscript{6} microemulsion.\emph{Bioprocess
and biosystems engineering} 2015, \emph{38} , 939-946.

{[}23{]} Hu, L., Llibin, S., Li, J., Qi, L.\emph{, et al.} ,
Lipase-catalyzed transesterification of soybean oil and phytosterol in
supercritical CO\textsubscript{2}. \emph{Bioprocess and biosystems
engineering} 2015, \emph{38} , 2343-2347.

{[}24{]} Chen, B., Jia, Y., Zhang, M., Li, X.\emph{, et al.} , Facile
modification of sepiolite and its application in superhydrophobic
coatings. \emph{Applied Clay Science} 2019, \emph{174} , 1-9.

{[}25{]} Cabrera, M. P., da Fonseca, T. F., de Souza, R. V. B., de
Assis, C. R. D.\emph{, et al.} , Polyaniline-coated magnetic diatomite
nanoparticles as a matrix for immobilizing enzymes. \emph{Applied
Surface Science} 2018, \emph{457} , 21-29.

{[}26{]} Zhao, K., Cao, X., Di, Q., Wang, M.\emph{, et al.} , Synthesis,
characterization and optimization of a two-step immobilized
lipase.\emph{Renewable energy} 2017, \emph{103} , 383-387.

{[}27{]} Tran, D.-T., Chen, C.-L., Chang, J.-S., Continuous biodiesel
conversion via enzymatic transesterification catalyzed by immobilized
Burkholderia lipase in a packed-bed bioreactor. \emph{Applied
Energy}2016, \emph{168} , 340-350.

{[}28{]} Basso, A., Hesseler, M., Serban, S., Hydrophobic
microenvironment optimization for efficient immobilization of lipases on
octadecyl functionalised resins. \emph{Tetrahedron} 2016, \emph{72} ,
7323-7328.

{[}29{]} Singh, A. K., Mukhopadhyay, M., Immobilization of lipase on
carboxylic acid-modified silica nanoparticles for olive oil
glycerolysis. \emph{Bioprocess and biosystems engineering}
2018,\emph{41} , 115-127.

{[}30{]} Fu, D., Li, C., Lu, J., ur Rahman, A., Tan, T., Relationship
between thermal inactivation and conformational change of \emph{Yarrowia
lipolytica} lipase and the effect of additives on enzyme
stability.\emph{Journal of Molecular Catalysis B: Enzymatic} 2010,
\emph{66} , 136-141.

{[}31{]} Hilterhaus, L., Thum, O., Liese, A., Reactor concept for
lipase-catalyzed solvent-free conversion of highly viscous reactants
forming two-phase systems. \emph{Organic Process Research \&
Development} 2008, \emph{12} , 618-625.

{[}32{]} Boudrant, J., Woodley, J. M., Fernandez-Lafuente, R.,
Parameters necessary to define an immobilized enzyme
preparation.\emph{Process Biochemistry} 2020, \emph{90} , 66-80.

{[}33{]} Guo, J., Chen, C.-P., Wang, S.-G., Huang, X.-J., A convenient
test for lipase activity in aqueous-based solutions. \emph{Enzyme and
Microbial Technology} 2015, \emph{71} , 8-12.

{[}34{]} Pencreac'h, G., Baratti, J. C., Activity of Pseudomonas cepacia
lipase in organic media is greatly enhanced after immobilization on a
polypropylene support. \emph{Applied Microbiology and
Biotechnology}1997, \emph{47} , 630--635.

{[}35{]} Li, Y., Li, B., Zhao, X., Tian, N., Zhang, J., Totally
waterborne, nonfluorinated, mechanically robust, and self-healing
superhydrophobic coatings for actual anti-icing. \emph{ACS applied
materials \& interfaces} 2018, \emph{10} , 39391-39399.

{[}36{]} Xu, F., Li, D., Chen, W., Gao, S., Formation of hydrophobic
silica coatings on stones for conservation of historic
sculptures.\emph{Chinese Journal of Chemistry} 2010, \emph{28} ,
1487-1490.

{[}37{]} Zhang, W., Zhang, Y., Lu, Z., Nian, B.\emph{, et al.} ,
Enhanced stability and catalytic performance of laccase immobilized on
magnetic graphene oxide modified with ionic liquids. \emph{Journal of
Environmental Management} 2023, \emph{346} , 118975.

{[}38{]} Feng, K., Hung, G.-Y., Liu, J., Li, M.\emph{, et al.} ,
Fabrication of high performance superhydrophobic coatings by
spray-coating of polysiloxane modified halloysite
nanotubes.\emph{Chemical Engineering Journal} 2018, \emph{331} ,
744-754.

{[}39{]} Wu, S., Wang, C., Jin, Y., Zhou, G.\emph{, et al.} , Green
synthesis of reusable super-paramagnetic diatomite for aqueous nickel
(II) removal. \emph{Journal of Colloid and Interface Science}
2021,\emph{582} , 1179-1190.

{[}40{]} Peng, X., Yuan, Z., Zhao, H., Wang, H., Wang, X., Preparation
and mechanism of hydrophobic modified diatomite coatings for oil-water
separation. \emph{Separation and Purification Technology}
2022,\emph{288} , 120708.

{[}41{]} Zhang, Y., Zhang, J., Wang, A., From Maya blue to biomimetic
pigments: durable biomimetic pigments with self-cleaning
property.\emph{Journal of Materials Chemistry A} 2016, \emph{4} ,
901-907.

{[}42{]} Fernández\selectlanguage{english}-Lorente, G., Palomo, J. M., Fuentes, M., Mateo,
C.\emph{, et al.} , Self\selectlanguage{english}-assembly of Pseudomonas fluorescens lipase into
bimolecular aggregates dramatically affects functional
properties.\emph{Biotechnology and bioengineering} 2003, \emph{82} ,
232-237.

{[}43{]} Martins, A. B., Friedrich, J. L., Cavalheiro, J. C.,
Garcia-Galan, C.\emph{, et al.} , Improved production of butyl butyrate
with lipase from Thermomyces lanuginosus immobilized on
styrene--divinylbenzene beads. \emph{Bioresource technology}
2013,\emph{134} , 417-422.

{[}44{]} Wiemann, L. O., Nieguth, R., Eckstein, M., Naumann, M.\emph{,
et al.} , Composite particles of novozyme 435 and silicone: advancing
technical applicability of macroporous enzyme
carriers.\emph{ChemCatChem} 2009, \emph{1} , 455-462.

{[}45{]} dos Santos, J. C., Garcia-Galan, C., Rodrigues, R. C., de
Sant'Ana, H. B.\emph{, et al.} , Stabilizing hyperactivated lecitase
structures through physical treatment with ionic polymers. \emph{Process
Biochemistry} 2014, \emph{49} , 1511-1515.

{[}46{]} Santos, J. C. S. d., Barbosa, O., Ortiz, C., Berenguer\selectlanguage{english}-Murcia,
A.\emph{, et al.} , Importance of the support properties for
immobilization or purification of enzymes. \emph{ChemCatChem}
2015,\emph{7} , 2413-2432.

{[}47{]} Rodrigues, R. C., Virgen-Ortiz, J. J., Dos Santos, J. C.,
Berenguer-Murcia, A.\emph{, et al.} , Immobilization of lipases on
hydrophobic supports: immobilization mechanism, advantages, problems,
and solutions. \emph{Biotechnology Advances} 2019, \emph{37} , 746-770.

{[}48{]} Verdasco-Martin, C. M., Garcia-Verdugo, E., Porcar, R.,
Fernandez-Lafuente, R., Otero, C., Selective synthesis of partial
glycerides of conjugated linoleic acids via modulation of the catalytic
properties of lipases by immobilization on different supports.\emph{Food
Chemistry} 2018, \emph{245} , 39-46.

{[}49{]} Kurtovic, I., Nalder, T. D., Cleaver, H., Marshall, S. N.,
Immobilisation of Candida rugosa lipase on a highly hydrophobic support:
A stable immobilised lipase suitable for non-aqueous
synthesis.\emph{Biotechnology Reports} 2020, \emph{28} , e00535.

{[}50{]} Sousa, R. R., Silva, A. S. A., Fernandez-Lafuente, R.,
Ferreira-Leitao, V. S., Solvent-free esterifications mediated by
immobilized lipases: A review from thermodynamic and kinetic
perspectives. \emph{Catalysis Science \& Technology} 2021, \emph{11} ,
5696-5711.

{[}51{]} Hari Krishna, S., Karanth, N., Lipases and lipase-catalyzed
esterification reactions in nonaqueous media. \emph{Catalysis
Reviews}2002, \emph{44} , 499-591.

{[}52{]} Dossat, V., Combes, D., Marty, A., Lipase-catalysed
transesterification of high oleic sunflower oil. \emph{Enzyme and
Microbial Technology} 2002, \emph{30} , 90-94.

{[}53{]} Ghamgui, H., Karra-Chaabouni, M., Gargouri, Y., 1-Butyl oleate
synthesis by immobilized lipase from Rhizopus oryzae: a comparative
study between n-hexane and solvent-free system. \emph{Enzyme and
Microbial Technology} 2004, \emph{35} , 355-363.

{[}54{]} Sandoval, G., Condoret, J., Monsan, P., Marty, A.,
Esterification by immobilized lipase in solvent\selectlanguage{english}-free media: Kinetic and
thermodynamic arguments. \emph{Biotechnology and bioengineering}
2002,\emph{78} , 313-320.

{[}55{]} Castillo, E., Torres-Gavilan, A., Sandoval, G., Marty, A.,
Thermodynamical methods for the optimization of lipase-catalyzed
reactions. \emph{Lipases and Phospholipases: Methods and Protocols}2012,
383-400.

{[}56{]} Adlercreutz, P., Immobilisation and application of lipases in
organic media. \emph{Chemical Society Reviews} 2013, \emph{42} ,
6406-6436.

{[}57{]} Goldberg, M., Thomas, D., Legoy, M.-D., Water activity as a key
parameter of synthesis reactions: the example of lipase in biphasic
(liquid/solid) media. \emph{Enzyme and microbial technology}
1990,\emph{12} , 976-981.

{[}58{]} Colombie, S., Tweddell, R. J., Condoret, J. S., Marty, A.,
Water activity control: A way to improve the efficiency of continuous
lipase esterification. \emph{Biotechnology and bioengineering}
1998,\emph{60} , 362-368.

{[}59{]} Lopresto, C. G., Calabro, V., Woodley, J. M., Tufvesson, P.,
Kinetic study on the enzymatic esterification of octanoic acid and
hexanol by immobilized \emph{Candida antarctica} lipase B. \emph{Journal
of Molecular Catalysis B: Enzymatic} 2014, \emph{110} , 64-71.

{[}60{]} Vadgama, R. N., Odaneth, A. A., Lali, A. M., Green synthesis of
isopropyl myristate in novel single phase medium Part I: Batch
optimization studies. \emph{Biotechnology Reports} 2015, \emph{8} ,
133-137.

{[}61{]} Kuperkar, V. V., Lade, V. G., Prakash, A., Rathod, V. K.,
Synthesis of isobutyl propionate using immobilized lipase in a solvent
free system: optimization and kinetic studies. \emph{Journal of
Molecular Catalysis B: Enzymatic} 2014, \emph{99} , 143-149.

{[}62{]} Scillipoti, J., Nioi, C., Marty, A., Camy, S., Condoret, J.-S.,
Prediction of conversion at equilibrium for lipase esterification in
two-phase systems. \emph{Biochemical engineering journal}
2017,\emph{117} , 162-171.

{[}63{]} Martins, A. B., Schein, M. F., Friedrich, J. L.,
Fernandez-Lafuente, R.\emph{, et al.} , Ultrasound-assisted butyl
acetate synthesis catalyzed by Novozym 435: Enhanced activity and
operational stability. \emph{Ultrasonics sonochemistry} 2013, \emph{20}
, 1155-1160.

{[}64{]} Romero, M., Calvo, L., Alba, C., Daneshfar, A., Ghaziaskar, H.,
Enzymatic synthesis of isoamyl acetate with immobilized \emph{Candida
antarctica} lipase in n-hexane. \emph{Enzyme and microbial
technology}2005, \emph{37} , 42-48.

{[}65{]} Khan, N. R., Rathod, V. K., Enzyme catalyzed synthesis of
cosmetic esters and its intensification: A review. \emph{Process
Biochemistry} 2015, \emph{50} , 1793-1806.

{[}66{]} Cui, C., Guan, N., Xing, C., Chen, B., Tan, T., Immobilization
of \emph{Yarrowia lipolytica} lipase Ylip2 for the biocatalytic
synthesis of phytosterol ester in a water activity controlled
reactor.\emph{Colloids and Surfaces B: Biointerfaces} 2016, \emph{146} ,
490-497.

{[}67{]} Zhang, S., Hou, H., Zhao, B., Zhou, Q.\emph{, et al.} , Hollow
mesoporous carbon-based enzyme nanoreactor for the confined and
interfacial biocatalytic synthesis of phytosterol esters. \emph{Journal
of Agricultural and Food Chemistry} 2023, \emph{71} , 2014-2025.

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