“Recent advances on support materials for lipase immobilization and applicability as biocatalysts in inhibitors screening methods”-A review

Journal Pre-proof “Recent advances on support materials for lipase immobilization and applicability as biocatalysts in inhibitors screening methods”-A Review Jia Liu, Run-Tian Ma, Yan-Ping Shi PII:

S0003-2670(19)31439-4

DOI:

https://doi.org/10.1016/j.aca.2019.11.073

Reference:

ACA 237286

To appear in:

Analytica Chimica Acta

Received Date: 28 August 2019 Revised Date:

29 November 2019

Accepted Date: 30 November 2019

Please cite this article as: J. Liu, R.-T. Ma, Y.-P. Shi, “Recent advances on support materials for lipase immobilization and applicability as biocatalysts in inhibitors screening methods”-A Review, Analytica Chimica Acta, https://doi.org/10.1016/j.aca.2019.11.073. This is a PDF file of an article that has undergone enhancements after acceptance, such as the addition of a cover page and metadata, and formatting for readability, but it is not yet the definitive version of record. This version will undergo additional copyediting, typesetting and review before it is published in its final form, but we are providing this version to give early visibility of the article. Please note that, during the production process, errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain. © 2019 Elsevier B.V. All rights reserved.

“Recent advances on support materials for lipase immobilization and applicability as biocatalysts in inhibitors screening methods”-A Review Jia Liu a,b, Run-Tian Ma a∗, Yan-Ping Shi a∗∗ CAS Key Laboratory of Chemistry of Northwestern Plant Resources, Key Laboratory for Natural Medicine of Gansu Province, Lanzhou Institute of Chemical Physics, Chinese Academy of Sciences (CAS), Lanzhou 730000, P. R. China b University of Chinese Academy of Sciences, Beijing 100049, P. R. China a

∗

Corresponding author. E-mail address: [email protected] (R.-T. Ma). Corresponding author. E-mail address: [email protected] (Y.-P. Shi).

∗∗

1

“Recent advances on support materials for lipase immobilization and

2

applicability as biocatalysts in inhibitors screening methods”-A Review

3 4

Jia Liu a, b, Run-Tian Ma a∗, Yan-Ping Shi a∗∗

5 6 a

7

CAS Key Laboratory of Chemistry of Northwestern Plant Resources, Key Laboratory for Natural

8

Medicine of Gansu Province, Lanzhou Institute of Chemical Physics, Chinese Academy of Sciences

9

(CAS), Lanzhou 730000, P. R. China b

10

University of Chinese Academy of Sciences, Beijing 100049, P. R. China

11

∗

Corresponding authors. E-mail address: [email protected] (R.-T. Ma). Corresponding authors. E-mail address: [email protected] (Y.-P. Shi). 1

∗∗

12

ABSTRACT

13

With a substantial demand for new anti-obesity drugs for the treatment of obesity, screening lipase

14

inhibitors from natural products has become a popular approach toward drug discovery. Due to the

15

significant advantages of excellent reusability, stability and endurance in extreme pH and

16

temperature conditions, lipase immobilization has been employed as a promising strategy to screen

17

lipase inhibitors. Support is a key factor in the process of enzyme immobilization used to provide

18

excellent biocompatibility, stable physical and chemical properties and abundant binding sites for

19

enzymes. Thus, various supports, including nanofibers, polymeric monoliths, mesoporous materials,

20

nanomaterials, membrane and cellulose paper, are systematically introduced and discussed in this

21

review. Considering these supports, the application of the immobilization of lipase in screening

22

compounds from natural products is also comprehensively reviewed, and the outlook for future

23

research directions is described.

24

Keywords: immobilization support, immobilized lipase, lipase inhibitor screening, anti-obesity,

25

natural products.

26 27

1. Introduction

28

The prevalence of obesity, particularly among teenagers and children, has seen a dramatic

29

increase recently and has attracted considerable attention. In the United States, obesity is considered

30

the most common cause of death, surpassing smoking [1]. Approximately 3.40 million of adults die

31

annually due to health problems caused by obesity and excess weight [2]. Obesity occurs when the

32

energy intake is higher than energy consumption, and the patient is diagnosed as obese when the

33

value of the body mass index (BMI) exceeds 30. Obesity is also a serious epidemic that is associated

34

with the risk of complications, such as hyperlipidemia, arteriosclerosis, type 2 diabetes, 2

35

cardiovascular disease and hypertension [3, 4]. Therefore, there is an urgent need to take measures to

36

solve the problem of obesity. Changing the daily diet in combination with exercise has been a

37

relatively ineffective approach to losing weight. Therefore, several obesity treatments, such as

38

surgery and drug therapy, have become popular among obese patients. The drug Orlistat, which is a

39

lipase inhibitor, was approved by the Food and Drug Administration (FDA) for long-term clinical

40

treatment of obesity [5]. However, long-term use of this drug may cause adverse gastrointestinal

41

effects, such as diarrhea, fatty stool, and flatulence [5, 6]. Therefore, there is an urgent requirement to

42

explore additional lipase inhibitors that function similarly to Orlistat but are more safe. The

43

enormous variety of natural products provides a solid foundation for the discovery of the new drugs

44

and has inspired researchers to explore safer and effective lipase inhibitors from natural products [2,

45

7-10]. To date, various methods, including capillary electrophoresis (CE) [11-13], gas

46

chromatography-mass spectrometry (GC-MS) [14], high-performance liquid chromatograph-mass

47

spectrometry (HPLC-MS) [15-17] and ultra-performance liquid chromatography-mass spectrometry

48

(UPLC-MS) [18], have been extensively employed, due to their merits of limited sample

49

consumption and simple automation [11]. In these processes, the use of immobilized lipase for

50

inhibitor screening is superior to the use of free lipase due to its pH and heat stability, as first

51

reported in the literature in 1972 [19]. Since 1972, this technology has advanced continuously and

52

became widely employed in 2008. For example, Tao et al. [15] used hollow fibers as the support to

53

immobilize lipase and screened lipase inhibitors from lotus leaf. Wan et al. [17] used magnetic

54

nanoparticles for lipase immobilization for further screening of lipase inhibitors from Scutellaria

55

baicalensis extract. Wang et al. [20] immobilized lipase on halloysite nanotubes to selectively screen

56

for lipase inhibitors from Magnoliae cortex extract. Based on the reports in the literature in this field, 3

57

the advantages of immobilized lipase can be summarized as excellent reusability, good operational

58

stability, easy recycling, more convenient purification procedures, relatively low operational costs

59

and prolonged enzyme survival time [11, 21-23].

60

According to the literature [24], the interfacial activation of lipase is a key factor in lipase

61

immobilization. Generally, the active center of the lipase molecule is covered by a polypeptide chain

62

that is known as the lid [24]. In the presence of a hydrophobic surface, the lipase is absorbed on the

63

surface. The lid is open, and the active center is completely exposed [25]. Thus, the lipase is

64

immobilized on the supports. Based on this approach, various enzyme immobilization methods have

65

been developed and can be simply divided into physical methods and chemical methods [26]. As

66

shown in Fig. 1, in the physical methods, the enzyme is immobilized on the support via physical

67

absorption, such as by electrostatic and hydrophobic interactions [26]. On the contrary, the enzyme is

68

immobilized on the support via covalent interactions in the chemical methods [26]. The physical

69

methods specifically include adsorption and entrapment, whereas chemical methods include covalent

70

attachment and cross-linking. Physical adsorption is simple because there is no requirement for the

71

functionalization of the support. In addition, the conformation of the enzyme can be retained, and the

72

catalytic activity of the immobilized enzyme is relatively high. However, in physical adsorption,

73

enzyme leaching is a critical problem limiting the use of the immobilized enzyme in the different

74

reaction conditions [27]. To solve this problem, entrapment is employed to restrict the enzyme in the

75

polymer frameworks [28, 29]. Thus the operational stability is improved, enzymes leakage is reduced,

76

the enzyme conformations are maintained, and high catalytic activities are achieved [30]. However,

77

the large diffusion barrier that limits the passage of the enzymes cannot be avoided in this method.

78

For chemical methods, the enzyme is firmly immobilized on the chemically modified support 4

79

through covalent binding and cross-linking, effectively preventing enzyme leakage [23]. Additionally,

80

the modified support provides multipoint attachments for the enzyme, improving the operational

81

stability of the immobilized enzyme. Cross-linking is an improvement on the covalent attachment

82

because the enzyme is cross-linked to the support with the help of a cross-linker. By using this

83

method, the catalytic activity of the immobilized enzymes can be retained even under harsh reaction

84

conditions.

85 86

Fig. 1. The enzyme immobilization methods.

87

The support is a key factor in the enzyme immobilization process and has attracted much

88

attention [23, 31]. Generally, ideal supports for enzyme immobilization should have excellent

89

biocompatibility, stable physical and chemical properties and abundant binding sites for the enzyme

90

[26]. Accordingly, a number of supports were developed for lipase immobilization, with an intense

91

effort devoted to the advancement of the screening of lipase inhibitors.

92

This review describes various supports that are beneficial for lipase immobilization and the

93

application of lipase inhibitor screening. For the first time, the majority of the reports about the 5

94

screening of lipase inhibitors from natural products are summarized. Compared to the review

95

published by our group last year [32], this review has a greater focus on the various supports that are

96

specific to lipase immobilization and the application of the immobilized lipase. In contrast, the

97

previous review placed more emphasis on the easily separated support materials for the

98

immobilization of different enzymes. Furthermore, future perspectives of the potential

99

immobilization supports and lipase inhibitor screening based on the immobilization techniques are

100

also provided.

101

2. Immobilization supports

102

Biocatalysis of immobilized enzymes has been researched for decades because it meets the

103

demand of sustainable development [27, 33]. This approach also provides the theoretical basis for

104

various applications of immobilized enzymes. Therefore, enzyme inhibitor screening is a promising

105

development direction in recent applications of immobilized enzymes. In this process, enzyme

106

immobilization support plays a key role. Thus, the most commonly employed supports, including

107

nanofibers, polymeric monoliths, mesoporous materials, nanomaterials, membrane and cellulose

108

paper, are summarized and discussed below.

109

2.1 Nanofibers

110

Nanofibers are extensively employed as the supports for lipase immobilization because they

111

possess the necessary functional groups and have a uniform diameter, and ultrahigh surface area to

112

volume ratios; additionally, nanofibers can be easily separated from the reaction media [34].

113

Self-assembly, electrospinning, template synthesis and phase separation are commonly used to

114

prepare nanofibers. Electrospun nanofibers always have infinite length and favorable dispersion and

115

functional groups [35]. Thus, electrospinning is the most universal, simple and highly effective 6

116

method for nanofiber preparation [36, 37]. For example, Dogac et al. [38] prepared glutaraldehyde

117

(GA) cross-linked polyvinyl alcohol (PVA)/alginate and polyethylene oxide (PEO)/alginate

118

nanofibers to immobilize lipase. After 40-60 min maintenance at high temperatures, approximately

119

65-70% activity of the lipase immobilized in nanofibers was retained, while the free lipase lost all

120

activity. In another study, Candida rugosa lipase (CRL) was covalently immobilized onto electrospun

121

polyacrylonitrile (PAN) nanofibers with a diameter in the range of 150-300 nm and with large

122

surface area for CRL immobilization [34]. The loading capacity of CRL reached as high as 2.1%, and

123

the immobilized CRL retained a high activity of 81.3%. Işik et al. [39] immobilized lipase on

124

polyvinyl alcohol (PVA)/Zn2+ electrospun nanofbers that were prepared by embedding a

125

polymer/ionic metal composite in the hybrid fibers. The immobilized lipase nanofibers exhibited

126

excellent thermostability and reusability, and the immobilization of lipase on the PVA/Zn2+

127

electrospun nanofbers was demonstrated for the first time.

128

Liu et al. [40] first reported the immobilization of Candida antarctica lipase B (CALB) on

129

poly(glycidyl methacrylate-co-methylacrylate)/feather polypeptide (P(GMA-coMA)/FP) nanofibrous

130

membrane that contained reactive epoxy groups and biocompatible FP (Fig. 2). This study revealed

131

that the nanofibrous membrane stabilized the enzyme conformation and improved the activity of the

132

immobilized enzyme. Under the treatment of 70°C for 3 h, the residual activity of the immobilized

133

lipase was approximately 38%. After 7 usages, the residual activity of the immobilized lipase was

134

approximately 62%. Further, the immobilized lipase also exhibited excellent endurance in organic

135

solvents, with approximately 75% of its activity preserved after storage in methanol for 12 h.

7

136 137

Fig. 2. Schematic illustration showing the detailed procedure for the preparation of electrospun

138

P(GMA-co-MA)/FP NFM immobilized with lipase [40]. [Reproduced by permission of the Copyright Clearance

139

Center, Elsevier].

140

2.2 Polymeric monoliths

141

Monoliths have been used as supports for lipase immobilization because of their simple

142

synthesis, good biocompatibility, hierarchically porous structure and a double-hole distribution [41,

143

42]. For example, Lathouder et al. [43] prepared three kinds of cordierite monoliths by using

144

carbonized sucrose, carbonized polyfurfurryl alcohol and carbon nanofibers, respectively. All of the

145

prepared monoliths were used to immobilize Candida antarctica lipase. Among the prepared

146

monolithic enzyme biocatalysts, carbon nanofiber-based monoliths exhibited the highest enzyme

147

loading capacity and the best storage stability. Samuel et al. [44] developed a lipase immobilized

148

poly(glycidyl

149

microreactor. The monolith microreactor was used to transesterificate castor oil triglycerides, and a

150

high conversion of 97% was realized. Xiao et al. [45] prepared a cellulose acetate monolith (CA-MN)

151

to immobilize lipase. Based on this, a continuous flow bioreactor was fabricated, and the catalytic

152

performance was tested. The results showed that the bioreactor could be continuously used for 200 h

153

without any loss of the catalytic activity. This study provided new ideas for the design and

methacrylate-co-ethylene

dimethacrylate

8

(poly(GMAco-EDMA))

monolith

154

application of a novel hierarchically structured monolith bioreactor. Sun et al. [46] synthesized an

155

acetoacetylated poly(vinyl alcohol) (AAPVA) monolith via the non-solvent-induced phase separation

156

(NIPS) technique, as shown in Fig. 3. This work represented the first example of a polymeric

157

monolith in which an acetoacetyl group was used for lipase immobilization, and a promising result

158

was obtained.

159 160

Fig. 3. The general fabrication process of AAPVA monolith via NIPS [46]. [Reproduced by permission of the Copyright Clearance Center, Springer].

161 162

2.3 Mesoporous materials

163

2.3.1 Mesoporous silica

164

Enzymes are extensively immobilized on mesoporous silica because of their large surface area,

165

well-organized pore geometry, confined pore size distributions and excellent thermal stability [47].

166

Furthermore, the surface of mesoporous silica supports can be chemically modified with various

167

functional groups to enhance the immobilization performance of the enzyme [48]. Recently, several

168

mesoporous silica materials have been employed as the supports for lipase immobilization. For

169

example, Jin et al. [49] synthesized three alkyl-functionalized mesoporous silica with different alkyls

170

(propyl, octyl and octadecyl) to immobilize lipase r27RCL. The result indicated that

171

octadecyl-functionalized mesoporous silica for immobilizing lipase r27RCL showed the highest

172

biocatalytic activity for the esterification reaction between ethanol and lauric acid. It was also found

173

that this silica could be reused at least 5 times without a significant activity loss. Ali et al. [50] used 9

174

amino-functionalized mesoporous silica nanoparticles (MSNPs) with a particle size of 200 nm and

175

pore size of 15-30 nm for the immobilization of CRL. Together, the specific channels in the

176

dendrimeric silica fibers combined with the function of GA provide favorable conditions for CRL

177

immobilization. The immobilized CRL maintained approximately 81% of the initial activity after

178

storage for 28 days, and 80% activity was retained after 8 reuses. In another study,

179

chitosan-mesoporous silica SBA-15 hybrid nanomaterials (CTS-SBA-15) with three-dimensional

180

(3D) structure were used to immobilize porcine pancreas lipase (PPL) with the help of GA [51]. The

181

immobilized PPL displayed improved stability and excellent reusability and enzymatic performance.

182

This study demonstrated that the CTS-SBA-15 material has great potential for use in enzyme

183

immobilization.

184

Zheng et al. [52] prepared the phenyl-modified ordered mesoporous silica (Ph-OMMs) with a

185

large pore size (>10 nm) for Burkholderia cepacia lipase (BCL) immobilization. According to the

186

morphology images presented in Fig. 4, the uniform cages were wrapped on the surface of the

187

hexagon-shaped OMMs and their mean diameter was 21 nm. Thus, the application of OMMs

188

introduced sufficient immobilization sites for the enzyme due to the large surface area of the OMMs,

189

tunable porosity and a functionalized pore wall. The as-prepared BCL@Ph-OMMs were successfully

190

used to catalyze the resolution of six secondary alcohols with high conversion (50%) and

191

enantioselectivity (≥99%). Specifically, the BCL@Ph-OMMs showed the best reusability among the

192

reported immobilized lipases for 50 continuous cycles, suggesting that BCL@Ph-OMMs may show

193

an extraordinary catalytic performance in industrial application. Based on the above, the pore

194

structures in mesoporous silica supports not only enhance the amount of the binding sites for the

195

lipase but also reduce the lipase leakage. Thus, lipase activity was greatly improved. 10

196 197

Fig. 4. Scanning electron microscopy (a-c) and transmission electron microscopy (d) images of OMMs [52]. [Reproduced by permission of the Copyright Clearance Center, Elsevier].

198 199

2.3.2 MOFs

200

Metal-organic frameworks (MOFs) are crystalline porous materials that are constructed by

201

interconnecting inorganic metal centers (metal ions or metal clusters) with bridging organic ligands

202

[53, 54]. Due to their extraordinary properties, such as giant porosity, a large surface area (even up to

203

6000 m2/g), tunable morphology and strong affinity for enzymes, MOFs have been extensively used

204

as promising supports for lipase immobilization [55, 56].

205

The first report of enzyme immobilization on MOFs was provided by Pisklak et al. [57].

206

Inspired by this work, the number of studies on the immobilization of lipases in MOFs has increased

207

gradually. For example, Samui et al. [58] developed an in situ method to synthesize CRL

208

immobilized on Zn-(NH2-BDC) MOFs. The immobilized CRL exhibited enhanced reusability and

209

thermal and pH stabilities. Bordbar et al. [59] used amino, trichlorotriazine amino and glutaraldehyde 11

210

amino groups to modify chromium terephthalate MIL-101 to obtain different supports for CRL

211

immobilization. For all of the MOFs supports, approximately 80-90% of the initial activities were

212

preserved after storage for 35 days, indicating an excellent storage stability of the immobilized CRL.

213

Wu et al. [60] compared the stability of the CRL/MOF composites and the free CRL at 80°C in

214

protein-denaturing solvents, as shown in Fig. 5. As a result, the immobilized CRL maintained 100%

215

of its activity in dimethyl sulfoxide, dimethyl formamide, methanol, and ethanol, whereas less than

216

20% of the initial activity was preserved for the free CRL exposed to the same conditions.

217 218

Fig. 5. Scheme of the green synthesis of enzyme-MOF composites exhibiting tolerance for denaturing solvents and

219

heat [60]. [Reproduced by permission of the Copyright Clearance Center, Springer].

220

Zeolitic imidazolate frameworks (ZIFs) are a type of MOFs with great prospects in enzyme

221

immobilization [61，62] because of their negligible cytotoxicity and outstanding chemical and

222

thermal stability [63]. Furthermore, ZIFs can be formed under mild conditions, solving the problem

223

of the relatively harsh reaction conditions required for in situ MOF synthesis [54]. Rafiei et al. [64]

224

developed the cobalt 2-methylimidazolate framework (ZIF-67) for immobilizing CRL by in situ

225

encapsulation. Adnan et al. [65] used a one-step biomineralization method to synthesize X-shaped

226

zeolite imidazolate framework-8 (ZIF-8) encapsulated RML. The immobilized RML was 12

227

successfully used as a biocatalyst for the transesterification of soybean oil to produce biodiesel. This

228

was the first report on the encapsulation of lipase within X-shaped ZIF-8 without changing the lipase

229

conformation, providing a novel strategy for the enzyme immobilization technology. Furthermore,

230

additional examples of the use of MOFs in lipase immobilization are given in Table 1. All of the

231

studies demonstrated that the application of MOFs in lipase immobilization is a worthwhile direction

232

for further exploration, even though this work is still in the initial stage.

233

Table 1

234

The applications of MOFs supports in lipase immobilization. Support Zeolite imidazolate framework-8 (ZIF-8)

Enzyme Rhizomucor miehei lipase

Functional reagent

Immobilization method

Application Biodiesel

Ref.

2-methylimidazole

Encapsulation

/

Encapsulation

Biocatalyst

[66]

Encapsulation

Biocatalyst

[67]

production

[65]

Pseudomonas ZIF-8

fluorescens lipase AK, Rhizomucor miehei

Sodium dodecyl ZIF-8

Candida rugose lipase

sulfate, bicinchoninic acid

Burkholderia cepacia

Cetyltrimethylammon

lipase

ium bromide

ZIF-67

Candida rugosa lipase

2-methylimidazole

Encapsulation

Zn-(NH2-BDC) MOFs

Candida rugosa lipase

/

Adsorption

UiO-66

Aspergillus niger lipase

Polydimethylsiloxane

Encapsulation

ZIF-8

Aspergillus niger lipase

ZIF-8

2-methylimidazole, zinc acetate

Adsorption

Encapsulation

Biodiesel production Biodiesel production Biocatalyst Biodiesel production

[68]

[64] [58] [69]

/

[70]

/

[71]

Physical

Magnetic-MOFs

Lipase

2-methylimidazole, H2BDC, H3BTC

adsorption, chemical binding, co-ordination bonding

13

MIL-100(Fe), HKUST-1

235

2.4 Nanomaterials

236

2.4.1 Nanotubes

Porcine pancreatic lipase

1,3,5-benzenetricarbo xylic acid

Encapsulation

Biocatalyst

[72]

237

A number of nanotubes, including carbon, halloysite and SnO2 nanotubes, have been employed

238

as supports for lipase immobilization. Nanotubes with a high surface area, easily functionalized

239

surface and opened lumens are favorable for enzyme immobilization [73]. A summary of the

240

nanotubes used as the supports in lipase immobilization is provided in Table 2.

241

Table 2

242

The applications of nanotubes supports in lipase immobilization. Support CNTs SWNTs MWNTs MWCNTs

MWCNTs

Enzyme

Functional

Immobilization

reagent

method

/

Adsorption

Yarrowia lipolytica lipase Pseudomonas cepacia lipase

BMIM-BF4

Candida rugosa lipase Burkholderia cepacia lipase Amano Lipase AK

Adsorption, chemical bonding

Application

Ref.

/

[74]

/

[75]

/

Adsorption

/

[76]

/

/

Biocatalyst

[77]

Cross-linking

Biocatalyst

[78]

DMF, NHS, EDC

mMWCNTs

Candida lipolytica lipase

/

Adsorption

Biocatalyst

[79]

Magnetic MWNTs

Yarrowia lipolytica lipase

EDC, NHS

Covalent binding

Biocatalyst

[80]

/

Covalent binding

mMWCNTs-PAMAM

m-MWCNTs-PAMAM

Peptide nanotubes SnO2 hollow nanotubes Halloysite clay nanotube Plasma-modified MWNTs

Burkholderia cepacia lipase Rhizomucor miehei lipase

Glutaraldehyd e, APTES, EDC, NHS

Candida rugosa lipase Lipase

Covalent bind

/ Glutaraldehyd e

Lipase

Chitosan

Lipozyme CALBL

/

14

Biodiesel production Biodiesel production

[81]

[82]

Encapsulation

/

[83]

Covalent bond

/

[84]

Antifouling

[85]

Biocatalyst

[86]

Electrostatic interaction Adsorption

Aminopropyl-grafted mesoporous silica

Candida sp. 99-125 lipase

APTES

Adsorption

Biocatalyst

[87]

Candida rugosa lipase

TEOS

/

/

[88]

Candida antarctica lipase B

Succinic acid

Biocatalyst

[89]

Biocatalysts

[90]

nanotubes CNTs–silica composites CheapTubes™ MWCNTs Functionalized MWCNTs

PDA@Co-MWCNTs

243

CAL-B

CRL

KMnO4, Na2SO3 Dopamine, CoCl2

Noncovalent binding Physical adsorption, covalent bonding Covalent binding

Nanobiocata lyst

[91]

2.4.1.1 Carbon nanotubes

244

Carbon nanotubes are extensively considered as versatile supports for lipase immobilization

245

because of their large surface area, small size, unique morphologies and thermal stability that

246

provides enhanced enzyme loading capacity [92]. For example, CALB was immobilized on

247

multiwalled carbon nanotubes (MWCNTs) and was used as a new heterogeneous nanobiocatalyst to

248

synthesize dicarboxylic acid esters. Compared to catalysts such as Novozyme-435, Amberlyst 14 and

249

CALB-CheapTubes™ MWCNTs, the CALB-immobilized MWCNTs had the shortest reaction time

250

[89]. Additionally, lipase was immobilized on carboxylated MWCNTs [77]. The resultant product

251

could be used 10 times to catalyze biodiesel production without any adverse effects on the lipase

252

activity. Li et al. [93] synthesized Ca3(PO4)2/CNT, Fe3(PO4)2/CNT and Cu3(PO4)2/CNT hybrid

253

nanotubes to immobilize BCL by adsorption and crystal encapsulation. All of the immobilized BCL

254

presented excellent reusability in the esterification reaction when exposed to organic medium,

255

indicating their great potential in industrial applications.

256

2.4.1.2 Halloysite nanotubes

257

Halloysite nanotubes (HNTs) constitute a special material with a hollow columnar structure and

15

258

nanoscale composition. HNTs have a positively charged inner surface composed of aluminum oxide

259

and a negatively charged silicon dioxide external surface composed of silicon dioxide [94, 95]. Thus,

260

the negatively charged enzymes can connect to the inner lumen of HNTs, and the positively charged

261

enzymes can adsorb on their external surface [96]. The unique structure of HNTs provides more

262

binding sites for enzyme and prevents nonspecific absorption. Thus, HNTs are an ideal support for

263

lipase immobilization. For example, Sun et al. [85] developed an efficient approach to immobilize

264

lipase on the chitosan-coated HNTs. In this case, the immobilized lipase maintained 90% of its initial

265

activity at 80°C, 70% of its initial activity at the pH of 9 and 85% of its activity after ten continuous

266

usages.

267

2.4.1.3 SnO2 hollow nanotubes

268

SnO2 hollow nanotubes are recognized as promising candidates for lipase immobilization due to

269

their large surface area and high porosity. As the reported in the literature, SnO2 hollow nanotubes

270

can obtain a high lipase loading value of 217 mg/g, immobilization yield of 93% and immobilization

271

efficiency of 89% [84]. The immobilized lipase had a half-life value of 4.5 h at 70°C and more than

272

91% of its initial activity was preserved even after 10 continuous usages. These results indicated that

273

the introduction of SnO2 hollow nanotubes can effectively reduce the activity loss of the immobilized

274

lipase upon exposure to a harsh environment. Thus, SnO2 hollow nanotubes are a promising direction

275

for future development as supports for the immobilization of various enzymes.

276

2.4.2 Magnetic nanoparticles

277

In recent decades, magnetic nanoparticles (MNPs) have emerged as versatile supports for lipase

278

immobilization due to their large surface-to-volume ratio, excellent physical and chemical stability,

279

low toxicity, tunable surface modification and easy separation [32, 97, 98]. Immobilization of lipase 16

280

on MNPs has been found to obtain high operational stability and retain good catalytic activity even

281

after several repeated usages [97, 98]. Among the various MNPs, such as Fe3O4, γ-Fe2O3, MgFe2O4,

282

MnFe2O4, CoFe2O4 and CoPt3 MNPs, the Fe3O4 MNPs have been mostly used for lipase

283

immobilization because of their excellent biocompatibility and nontoxicity [99]. However, bare

284

Fe3O4 MNPs always tend to aggregate due to the magnetic dipole-dipole attractions [100]. According

285

to the reports in the literature, functionalization of Fe3O4 MNPs can effectively improve their

286

dispersity and chemical stability [101]. Liu et al. from our group prepared Fe3O4/CS/GA NPs [102]

287

and chitosan-enriched magnetic composites (MCCs) [103] for α-glucosidase immobilization and

288

applied them for α-glucosidase inhibitor screening. Due to the application of magnetic supports,

289

multiple centrifugations that are used in conventional screening were avoided, and the separation

290

procedures were significantly simplified. The application of various modified Fe3O4 MNPs,

291

including polymer-modified MNPs, silica-coated MNPs, ionic liquids modified MNPs and

292

MOF-based MNPs, as the supports for lipase immobilization are discussed in detail below.

293

2.4.2.1 Polymer-modified MNPs

294

Fe3O4 MNPs are always modified with polymers because this provides more binding sites for

295

lipase immobilization through the various functional groups [104，105]. For example, Wu et al. [106]

296

prepared Fe3O4-chitosan NPs to immobilize lipase with the crosslinker GA. With the help of the

297

chitosan, hydroxyl, amino and carbonyl groups on the surface of the magnetic support, lipase

298

immobilization was promoted. Ren et al. [107] used polydopamine to modify magnetic nanoparticles

299

(PD-MNPs) in order to improve the dispersity of the bare Fe3O4 NPs and provide abundant hydroxyl

300

and amino groups for lipase immobilization. The immobilized lipase possessed excellent reusability,

301

enhanced pH and thermal stability. In addition, PAMAM dendrimer was also a common material 17

302

used for MNPs modification. Li et al. [108] prepared aminated MNPs grafted with melamine-GA

303

dendrimer-like polymers that provided more binding sites for lipase immobilization. The activity of

304

the immobilized lipase on the aminated MNPs grafted with melamine-GA dendrimer-like polymers

305

was three times higher than that of the lipase immobilized on the aminated MNPs. This study

306

demonstrated facile and efficient preparation of the biocatalyst that has great potential in industrial

307

application.

308

2.4.2.2 Silica-coated MNPs

309

Silica coating is the most commonly used material for the modification of MNPs due to its

310

enriched surface reactive groups, biocompatibility and water dispersibility. By using a common

311

sol-gel process, silica shells are developed on the surfaces of magnetic cores, producing silica-coated

312

MNPs (Fe3O4@SiO2) [109]. The formed silica shell protects the MNPs from aggregating and

313

oxidation, leading to their improved chemical stability [110, 111]. Through the hydrolysis reaction

314

between the silanol groups on the surface of Fe3O4@SiO2 NPs and the silane coupling agent, various

315

functional groups can be grafted on the surface of Fe3O4@SiO2 NPs to immobilize enzymes [112,

316

113]. For example, Fe3O4@SiO2 was functionalized with 3-aminopropyltriethoxysilane (APTES) to

317

provide more amino groups for Rhizopus oryzae lipase immobilization [111]. It was found that the

318

immobilized Rhizopus oryzae lipase retained 64% of its initial activity after 10 reusages. Tardioli et

319

al. [114] synthesized mono- and heterofunctionalized silica magnetic microparticles to immobilize

320

CALB. The as-prepared immobilized CALB was used to synthesize xylose fatty acid esters in the

321

tert-butyl alcohol medium. The results demonstrated that the magnetic biocatalyst displayed high

322

catalytic efficiency and excellent reusability.

323

2.4.2.3 Ionic liquids functionalized MNPs 18

324

Over the past decade, ionic liquids (ILs) have been found to be appropriate media for enzyme

325

catalysis [115, 116], and it was demonstrated that different lipases possess excellent activities and

326

high stabilities in ionic liquids. In recent years, ILs have become gradually more widely employed as

327

functional groups for the modification of MNPs. A number of studies have demonstrated that the

328

ILs-functionalized MNPs exhibit enhanced the electrostatic interactions, hydrophobic interactions

329

and hydrogen bonds between the support and the lipase, which prevent the leakage of lipase from the

330

support

331

acid)-imidazolium salt) functionalized MNPs were used to immobilize lipase [120]. This study was

332

the first use of ionic liquids as the cross-linker between the lipase and MNPs. Suo et al. [121]

333

immobilized PPL on the imidazole-based ionic liquid modified magnetic chitosan nanoparticles

334

(PPL-IL‑CS‑Fe3O4). Due to the presence of ILs, the PPL conformation was protected from damage.

335

Thus, the thermal stability of the immobilized PPL was significantly improved. In addition, 84.6% of

336

the initial activity of PPL‑IL‑CS‑Fe3O4 was retained even after 10 reuses, whereas 75.5% of the

337

initial activity for PPL‑CS‑Fe3O4 was retained. This result could also be attributed to the

338

introduction of ILs.

[117-119].

For

example,

[Cn(A)C4(-D)Im]X

(1-butyraldehyde-3-(carbonic

339

Huang et al. [122] immobilized PPL on magnetic nanocomposites by combining

340

Fe3O4@chitosan nanocomposites functionalized with ionic liquids with PPL (Fig. 6). The specific

341

activity of the immobilized PPL was 6.68 times higher than that of the free PPL. Approximately 91.5%

342

of the initial activity of the immobilized PPL was maintained even after 10 successive reuses. After

343

incubating in a urea solution for 1 h, the immobilized PPL retained 55.8% of its initial activity. This

344

study demonstrated that imidazole-based ionic liquids functionalized with Fe3O4@chitosan

345

nanocomposites can be used as promising supports for enzyme immobilization. 19

346 Fig. 6. Synthetic procedure of ionic liquids modified magnetic chitosan nanocomposites and its application in

347

PPL immobilization [122]. [Reproduced by permission of the Copyright Clearance Center, Elsevier].

348 349

2.4.2.4 MOFs modified MNPs

350

Combining the merits of MNPs and MOFs, the use of MOFs-modified MNPs has been reported

351

as a novel support for lipase immobilization. For example, Wang et al. [123] combined

352

carboxyl-functionalized Fe3O4 nanorods with MIL-100(Fe) to prepare Fe3O4@MIL-100(Fe) in a

353

simple and environmentally friendly manner. Then, the prepared magnetic support was employed to

354

immobilize CRL. Approximately 60% of the initial activity for the immobilized CRL was retained

355

even after ten reaction cycles. Sargazi et al. [124] synthesized Ta-MOF@Fe3O4 to immobilize

356

Bacillus licheniformis Km12 lipase for the first time. After incubating at 50°C for 3 h, the

357

immobilized Km12 lipase activity was unchanged, while 91% activity of the free Km12 lipase was

358

retained. Additionally, applications of magnetic supports in lipase immobilization are listed in Table

359

3.

360

Table 3

361

The applications of magnetic supports in lipase immobilization. Support Fe3O4@SiO2 NPs

Enzyme Candida antarctica lipase B

Functional

Immobilization

reagent

method

GPTMS,TEOS

Covalent attachment

20

Application

Ref.

Biodiesel

[125]

Fe3O4-chitosan NPs Fe3O4@MIL-100(Fe )

Porcine pancreatic lipase

Chitosan,

Cross-linking

/

[106]

Covalent bonding

/

[123]

/

[119]

Covalent linkages

Biocatalyst

[114]

Covalent conjugation

Biosensor

[105]

APTES

Covalent bonding

/

[111]

Imidazole

Covalent binding

/

[120]

/

[122]

/

[126]

/

[108]

Glutaraldehyde EDTA-2Na,

Candida rugosa lipase

H3btc,EDC,NH S 3-chloropropyl

IM/BF4-Fe3O4@CA

Porcine pancreatic

trimethoxysila

Electrostatic

lipase

ne, methyl

adsorption

imidazole Silica magnetic microparticles Polydopamine-Fe3O4 NPs Amino-functionalize d Fe3O4 NPs [Cn(A)C4(-D)Im]X Fe3O4 NPs

Candida antarctica lipase B

Candida rugosa lipase Rhizopus oryzae lipase Candida rugosa lipase

APTES Dopamine hydrochloride

Chitosan,1(3IL-Fe3O4@Chitosan nanocomposites

Porcine pancreatic lipase

Amino

Covalent

propyl)

crosslinking

-imidaz ole Poly(carboxybetaine methacrylate)-Fe3O4 NPs

Porcine pancreatic lipase

Dendrimer-polymers -aminated Fe3O4 NPs

362

CBMA, APS, MBAA, TEMED APTES,

Burkholderia cepacia lipase

Crosslinking

glutaraldehyde, melamine

Covalent bonding

2.5 Membrane

363

Membranes are promising supports for enzyme immobilization because they can promote the

364

application of enzymes in membrane bioreactors, enzymatic reactors and biosensors [127]. The

365

membrane surface is typically modified with functional groups for lipase immobilization. The

366

covalent attachment of the lipase on the membrane surface is more popular than physical absorption

367

because it can increase the stability and reusability of the lipase [128]. For example, Aghababaie et al. 21

368

[127] reported that the aminated poly acrylonitrile membranes (MNP@PAN) were activated by

369

glutaraldehyde (GA) and trichlorotriazine (TCT), resulting in TCT-MNP@PAN and GA-MNP@PAN

370

membranes, respectively. Then, CRL was covalently immobilized on the prepared nanocomposite

371

membrane. The results showed that the activity of GA-MNP@PAN and TCT-MNP@PAN

372

membranes were approximately 50% and 31%, respectively, higher than that of GA-activated PAN

373

membrane. Li et al. [129] prepared a functionalized PAN membrane for the immobilization of CALB.

374

As shown in Fig. 7, polyethyleneimine (PEI) was introduced on the surface of the membrane with

375

the nitrile-click chemistry. Then, the prepared PAN-PEI was treated with sodium alginate (SA) and

376

post treated by CaCl2. The resultant PAN-PEI-SA-CaCl2 was used for lipase immobilization and

377

acted as a catalyst for biodiesel production. After the PAN-PEI-SA-CaCl2 was successively used for

378

20 times, only 11% of the biodiesel yield was lost.

379 380

Fig. 7. Process of enzyme immobilization [129]. [Reproduced by permission of the Copyright Clearance Center, Elsevier].

381 382

2.6 Cellulose paper

383

Paper materials have the advantages of low cost, portability, commercial availability,

384

hydrophilicity, environmental friendliness and ease of handling. Therefore, paper is expected to serve 22

385

as a novel support for lipase immobilization [130]. Through the reaction between the C-OH on the

386

cellulose paper and the silane coupling agent, various functional groups can be introduced into

387

cellulose paper for lipase immobilization. For example, Koga et al. [131] prepared cellulose paper

388

with methacryloxy groups to immobilize lipase. Then, the catalytic performance of the immobilized

389

lipase was evaluated by a transesterification reaction between 1-phenylethanol and vinyl acetate to

390

produce 1-phenylethylacetate. Due to the hyperactivation of the methacryloxy groups towards lipase

391

and the unique structure of the cellulose paper, the immobilized lipase exhibited excellent reusability

392

and high catalytic activity. In another study [132], nuclease p1 was immobilized on paper cellulose.

393

The immobilized nuclease p1 showed a tolerance for a wide range of pH and temperature variations.

394

Based on the above, cellulose paper is a relatively green support for lipase immobilization and can be

395

applied to the immobilization of other enzymes. For example, Liu et al. [130] from our group applied

396

chitosan-modified cellulose paper for α-Glu immobilization. After 10 successive cycles, 71.0% of the

397

initial activity was preserved, and the immobilized α-Glu exhibited favorable temperature and pH

398

stability. Lawrence et al. [133] immobilized GOx on a cellulose paper disk via adsorption. Tyagi et al.

399

[134] prepared cellulose filter paper grafted with glycidyl methacrylate (GMA) for urease

400

immobilization. All of the above examples demonstrated that cellulose paper is a promising support

401

for enzyme immobilization.

402

3. Lipase inhibitors screening based on immobilized enzyme

403

Lipase inhibitors can be used as anti-obesity drugs to inhibit the absorption of fats, thus

404

achieving weight loss. However, the conventional strategies for lipase inhibitor screening have

405

several shortcomings, such as high time consumption and labor intensity and low efficiency [135,

406

136]. In recent years, researchers have preferred to screen lipase inhibitors rapidly from natural 23

407

products combined with the immobilized lipase due to the easy manipulation and automation of this

408

approach. Although lipase inhibitor screening technology based on immobilization enzyme strategy

409

is still in the early stages, a batch of compounds was surprisingly successfully screened out. A basis

410

for further research and development of the new anti-obesity drugs has gradually been established.

411

Accordingly, a system for the screening and identification potential lipase inhibitors from natural

412

products via immobilization enzyme technology was developed, as shown in Fig. 8.

24

413 414

Fig. 8. Schedule for screening and identifying potential lipase inhibitors from natural products.

415

For example, Zhu et al. [137] prepared carboxylated magnetic nanoparticles for covalent PPL

416

immobilization and used them to identify the lipase inhibitory activities of two compounds,

417

(−)-epigallocatechin gallate (EGCG) and (−)-epigallocatechin (EGC), isolated from Oolong tea. The

418

specific screening procedures were as follows. The immobilized PPL magnetic nanoparticles, in the 25

419

presence or absence of the samples, were mixed with the substrates (p-NPP). After incubating with

420

the enzyme buffer, the supernatants were injected into HPLC for the analysis of the product (p-NP).

421

By comparing the chromatographic peak area before and after the addition of the samples, the

422

inhibition ratio was calculated according to the reduction of the peak area of p-NP. The results

423

indicated that the IC50 values of EGCG was 55.00 ± 0.50 µM, demonstrating that EGCG possessed a

424

remarkable lipase inhibitory effect. Further, EGC exhibited no inhibitory against lipase even at the

425

high concentration. This work was the first report on the magnetically separable immobilized lipase

426

used for the screen lipase inhibitors, demonstrating that this is a facile and rapid screening method.

427

Wan et al. [17] used lipase-immobilized magnetic nanoparticles (LMNPs) as a solid extraction

428

absorbent. First, they used the chemical coprecipitation method to synthesize amino-functionalized

429

Fe3O4 MNPs. Then, the Stöber method was applied to form an SiO2 layer on the surface of the MNPs.

430

After that, the Fe3O4@SiO2 was modified to graft carboxyl groups for the further lipase

431

immobilization. By incubating LMNPs with Scutellaria baicalensis extract, LMNP-ligand

432

complexes were formed. The nonspecifically bounded compounds were removed by a Tris-HCl

433

buffer, and the specifically bounded ligands were eluted and analyzed by HPLC-MS/MS. Three

434

ligands of lipase were identified as baicalin, wogonin and oroxylin A, and their half maximal

435

inhibitory concentration (IC50) values were calculated as 229.22 ± 12.67, 153.71 ± 9.21 and 56.07 ±

436

4.90 µM, respectively. To further explore the IC50 difference of the compounds, the molecular

437

docking technique was used to simulate the binding mode between the lipase and the ligands. In this

438

experiment, oroxylin A exhibited the best affinity for PPL among the three lipase inhibitors, and this

439

result successfully explained why oroxylin A had the best inhibitory activity.

440

In another study, Zhu et al. [18] immobilized PL on the amino-functionalized MNPs (PL-MNPs) 26

441

and employed it as an absorbent in combination with UPLC-MS for screening lipase inhibitors from

442

Oolong tea. A suspension of PL-MNPs was incubated with Oolong tea extract and then washed three

443

times with an NH4OAc solution to remove the nonspecifically absorbed compounds. Subsequently,

444

the eluent of the mixture was filtrated and analyzed by UPLC-MS/MS. By comparing the

445

chromatograms of the eluent and the Oolong tea extract solutions, three PL inhibitors were found and

446

identified as EGCG, (-)-gallocatechin-3-O-gallate (GCG) and (-)-epicatechin-3-O-gallate (ECG). In

447

addition, Wang et al. [15] reported a novel strategy called hollow fiber-based affinity selection

448

(HF-AS) and developed it for lipase inhibitor screening from the total flavonoids of lotus leaf. The

449

detailed processes of screening active compounds are illustrated in Fig. 9. Briefly, lipase was

450

absorbed onto the hollow fibers immersed in the lipase solution. Then, the lotus leaf extract was

451

incubated with the immobilized lipase for a period of time. Subsequently, the specifically bounded

452

compounds were dissociated and analyzed by HPLC-MS. Through the proposed HF-AS approach,

453

three

454

quercetin-3-O- β-D-glucuronide and kaempferol-3-O-β-D-glucuronide, were screened out. The results

455

showed that the HF-AS strategy can be a rapid and convenient approach for lipase inhibitor

456

screening from natural product resources.

active

compounds,

quercetin-3-O-β-D-arabinopyranosyl-(1→2)-β-D-galactopyranoside,

27

457 458 459

Fig. 9. Schematic diagram of the proposed hollow fibers based affinity selection method [15]. [Reproduced by permission of the Copyright Clearance Center, Elsevier].

460

Wang et al. [20] prepared lipase-immobilized halloysite nanotubes and used them as a medium

461

to screen lipase inhibitors from natural products for the first time. The immobilized lipase was

462

employed to screen potential compounds with anti-obesity activity from the Magnoliae cortex. The

463

lipase-immobilized halloysite nanotubes were incubated with Magnoliae cortex extract solution at

464

room temperature. After 2 h of incubation, the specifically bounded ligands were released by

465

washing with acetonitrile and then were centrifuged for 5 min. According to the results of the

466

HPLC-MS analysis of the supernatants, four compounds, magnotriol A, magnaldehyde A,

467

magnaldehyde D and magnaldehyde B, were isolated and identified as potential lipase inhibitors. In

468

addition, the values of the binding degree of the four compounds were calculated as 4.5%, 5.0%, 7.0%

469

and 7.4%, respectively. Considering the highest binding degree of magnaldehyde B, molecular

470

docking was further performed. The obtained result indicated that magnaldehyde B binds with 28

471

several main amino acids located at the catalytic site of the pancreatic lipase, demonstrating that it is

472

the most promising lipase inhibitor in Magnoliae cortex. The above mentioned reports not only

473

comprehensively described the whole screening process of the lipase inhibitors from natural products

474

but also demonstrated the feasibility and great potential of the future development of lipase inhibitor

475

screening technology based on enzyme immobilization.

476

4. Conclusions and future perspectives

477

This review provides an overview of the supports used for lipase immobilization and the

478

application of immobilized lipase to the screening of lipase inhibitors from natural products.

479

Different supports that possess various advantages for lipase immobilization were summarized and

480

discussed. Nanofibers provide large specific surface area for lipase immobilization and greatly

481

improve the immobilization efficiency and reaction catalytic efficiency. Polymer monoliths have

482

been widely used as the supports for lipase immobilization because of their high chemical stability,

483

good biocompatibility and ease of modification with different functional groups. Mesoporous silica

484

is a promising support for the immobilization of lipase because of its large surface areas,

485

well-organized pore geometry, confined pore size distributions, excellent thermal stability and easily

486

modified surface. Nanotubes are another kind of lipase immobilization support that have attracted

487

research interest due to their merits of favorable surface area, small size, unique morphologies, and

488

excellent mechanical and thermal stability. Additionally, MOFs have been employed as suitable

489

lipase immobilization supports due to their ultrahigh surface area to volume ratios, giant porosity,

490

tunable morphology and appropriate pore size. Furthermore, MNPs can be used to overcome the

491

difficulty of filtration and centrifugal separation methods in the conventional screening process. To

492

increase the amount of the binding sites on MNPs, polymers, silica, ionic liquids and MOFs have 29

493

been coated on their surfaces. Thus, functionalized MNPs are potential supports for efficient lipase

494

immobilization. Membranes are an ideal choice for lipase immobilization because they can enhance

495

the stability and reusability of the lipase. Cellulose paper is expected to serve as a novel lipase

496

immobilization support due to its extraordinary advantages of portability, hydrophilicity, low cost,

497

commercial availability, environmental friendliness and ease of handling. In our opinion,

498

MOFs-modified MNPs may be regarded as the main direction for the further development of lipase

499

immobilization supports, as they integrate the merits of giant porosity, simple operation, tunable

500

morphology and ultrahigh surface area to volume ratios. Nevertheless, recyclable supports that have

501

potential for use in high-throughput screening need to be developed. New functional materials, such

502

as silica aerogels with large surface area to volume ratios, may be introduced as a potential support

503

for lipase immobilization. Furthermore, new immobilization techniques that are simpler, faster and

504

more effective are also expected.

505

Screening lipase inhibitors from natural products is one of the main approaches for anti-obesity

506

drug discovery. Scutellaria baicalensis, Oolong tea, lotus leaf and Magnoliae cortex have been found

507

to possess lipase inhibitory activities, as reviewed in Section 3. These studies demonstrated the

508

feasibility of the screening of lipase inhibitors based on the immobilized lipase strategy and inspired

509

more research focused on the screening from natural products. In our opinion, lipase inhibitor

510

screening based on immobilized-lipase can be implemented in screening active compounds with

511

lipase inhibition, rather than only focusing on screening inhibitors from natural products.

512

Furthermore, the screening range of plants can be broadened and should not be restricted to the

513

plants that have been reported to possess lipase inhibitory activities. Finally, mass spectrometry with

514

more advanced instruments should be comprehensively utilized in the process of screening lipase 30

515

inhibitors to obtain more accurate screening results.

516 517

Declarations of interests

518

The authors declare that they have no known competing financial interests or personal relationships

519

that could have appeared to influence the work reported in this paper.

520 521

Acknowledgements

522

This work was financially supported by the National Natural Science Foundation of China (Nos.

523

21775153, 21804135 and 21974145), and the Scholar Program of West Light Project of the Chinese

524

Academy of Sciences.

31

525

References

526

[1] Y.G. Shi, P. Burn, Lipid metabolic enzymes: Emerging drug targets for the treatment of obesity, Nat. Rev. Drug

527

Discov. 3 (2004) 695-710.

528

[2] T.G. Chen, Y.Y. Li, L.W. Zhang, Nine different chemical species and action mechanisms of pancreatic lipase

529

ligands screened out from Forsythia suspensa leaves all at one time, Molecules 22 (2017) 795-806.

530

[3] P.G. Kopelman, Obesity as a medical problem, Nature 404 (2000) 635-645.

531

[4] K.L. Canning, R.E. Brown, V.K. Jamnik, J.L. Kuk, Relationship between obesity and obesity-related

532

morbidities weakens with aging, J. Gerontol. A-Biol. 69 (2014) 87-92.

533

[5] A.M. Heck, J.A. Yanovski, K.A. Calis, Orlistat, a new lipase inhibitor for the management of obesity,

534

Pharmacotherapy 20 (2000) 270-279.

535

[6] D. Chaudhari, C. Crisostomo, C. Ganote, G. Youngberg, Acute oxalate nephropathy associated with orlistat: a

536

case report with a review of the literature, Case Rep. Nephrol. 2013 (2013) 124604-124607.

537

[7] C. Fu, Y. Jiang, J. Guo, Z. Su, Natural products with anti-obesity effects and different mechanisms of action, J.

538

Agric. Food Chem. 64 (2016) 9571-9585.

539

[8] R.B. Birari, K.K. Bhutani, Pancreatic lipase inhibitors from natural sources: unexplored potential, Drug Discov.

540

Today 12 (2007) 879-889.

541

[9] X.D. Hou, G.B. Ge, Z.M. Weng, Z.R. Dai, Y.H. Leng, L.L. Ding, L.L. Jin, Y. Yu, Y.F. Cao, J. Hou, Natural

542

constituents from Cortex Mori Radicis as new pancreatic lipase inhibitors, Bioorg. Chem. 80 (2018) 577-584.

543

[10] Y. Fu, J. Luo, J. Qin, M. Yang, Screening techniques for the identification of bioactive compounds in natural

544

products, J. Pharm. Biomed. Anal. 168 (2019) 189-200.

545

[11] Z.M. Tang, J.W. Kang, Enzyme inhibitor screening by capillary electrophoresis with an on-column

546

immobilized enzyme microreactor created by an ionic binding technique, Anal. Chem. 78 (2006) 2514-2520. 32

547

[12] X.W. Ji, F.G. Ye, P.T. Lin, S.L. Zhao, Immobilized capillary adenosine deaminase microreactor for inhibitor

548

screening in natural extracts by capillary electrophoresis, Talanta 82 (2010) 1170-1174.

549

[13] O. Hodek, T. Krizek, P. Coufal, H. Ryslava, Online screening of alpha-amylase inhibitors by capillary

550

electrophoresis, Anal. Bioanal. Chem. 410 (2018) 4213-4218.

551

[14] M. Wang, D. Gu, H. Li, Q. Wang, J. Kang, T. Chu, H. Guo, Y. Yang, J. Tian, Rapid prediction and

552

identification of lipase inhibitors in volatile oil from Pinus massoniana L. needles, Phytochemistry 141 (2017)

553

114-120.

554

[15] Y. Tao, Y.F. Zhang, Y. Wang, Y.Y. Cheng, Hollow fiber based affinity selection combined with high

555

performance liquid chromatography-mass spectroscopy for rapid screening lipase inhibitors from lotus leaf, Anal.

556

Chim. Acta 785 (2013) 75-81.

557

[16] Y. Tao, Z. Chen, Y. Zhang, Y. Wang, Y. Cheng, Immobilized magnetic beads based multi-target affinity

558

selection coupled with high performance liquid chromatography-mass spectrometry for screening anti-diabetic

559

compounds from a Chinese medicine "Tang-Zhi-Qing", J. Pharm. Biomed. Anal. 78-79 (2013) 190-201.

560

[17] L.H. Wan, X.L. Jiang, Y.M. Liu, J.J. Hu, J. Liang, X. Liao, Screening of lipase inhibitors from Scutellaria

561

baicalensis extract using lipase immobilized on magnetic nanoparticles and study on the inhibitory mechanism,

562

Anal. Bioanal. Chem. 408 (2016) 2275-2283.

563

[18] Y.T. Zhu, X.Y. Ren, L. Yuan, Y.M. Liu, J. Liang, X. Liao, Fast identification of lipase inhibitors in oolong tea

564

by using lipase functionalised Fe3O4 magnetic nanoparticles coupled with UPLC-MS/MS, Food Chem. 173 (2015)

565

521-526.

566

[19] T. Ogiso, M. Sugiura, Y. Kato, Studies on bile-sensitive lipase. X. Preparation and properties of carrier-bound

567

mucor lipase, Chem. Pharm. Bull. 20 (1972) 2542-2550.

568

[20] H.B. Wang, X.P. Zhao, S.F. Wang, S. Tao, N. Ai, Y. Wang, Fabrication of enzyme-immobilized halloysite 33

569

nanotubes for affinity enrichment of lipase inhibitors from complex mixtures, J. Chromatogr. A 1392 (2015) 20-27.

570

[21] P. Adlercreutz, Immobilisation and application of lipases in organic media, Chem. Soc. Rev. 42 (2013)

571

6406-6436.

572

[22] C.J. Gray, M.J. Weissenborn, C.E. Eyers, S.L. Flitsch, Enzymatic reactions on immobilised substrates, Chem.

573

Soc. Rev. 42 (2013) 6378-6405.

574

[23] U. Hanefeld, L. Gardossi, E. Magner, Understanding enzyme immobilisation, Chem. Soc. Rev. 38 (2009)

575

453-468.

576

[24] A.M. Brzozowski, U. Derewenda, Z.S. Derewenda, A model for interfacial activation in lipases from the

577

structure of a fungal lipase-inhibitor complex, Letters to Nature 351 (1991) 491-494.

578

[25] E.A. Manoel, J.C. Dos Santos, D.M. Freire, N. Rueda, R. Fernandez-Lafuente, Immobilization of lipases on

579

hydrophobic supports involves the open form of the enzyme, Enzyme Microb. Tech. 71 (2015) 53-57.

580

[26] M. Hartmann, X. Kostrov, Immobilization of enzymes on porous silicas-benefits and challenges, Chem. Soc.

581

Rev. 42 (2013) 6277-6289.

582

[27] M. Hartmann, D. Jung, Biocatalysis with enzymes immobilized on mesoporous hosts: the status quo and future

583

trends, J. Mater. Chem. 20 (2010) 844-857.

584

[28] S.M. Safaryan, A.V. Yakovlev, E.A. Pidko, A.V. Vinogradov, V.V. Vinogradov, Reversible sol-gel-sol medium

585

for enzymatic optical biosensors, J Mater. Chem. B 5 (2017) 85-91.

586

[29] R.L. Souza, E.L.P. Faria, R.T. Figueiredo, S. Mettedi, O.A.A. Santos, A.S. Lima, C.M.F. Soares, Protic ionic

587

liquid applied to enhance the immobilization of lipase in sol-gel matrices, J. Therm. Anal. Calorim. 128 (2017)

588

833-840.

589

[30] R.A. Sheldon, Enzyme immobilization: The quest for optimum performance, Adv. Synth. Catal. 349 (2007)

590

1289-1307. 34

591

[31] A.A. Khan, M.A. Alzohairy, Recent advances and applications of immobilized enzyme technologies: A review,

592

Curr. Res. J. Biol. Sci. 5 (2010) 565–575

593

[32] D.M. Liu, J. Chen, Y.P. Shi, Advances on methods and easy separated support materials for enzymes

594

immobilization, TrAC-Trend. Anal. Chem. 102 (2018) 332-342.

595

[33] L.Q. Cao, Immobilized enzymes: Science or art. Curr. Opin. Chem. Biol. 9 (2005) 217-226.

596

[34] S.F. Li, J.P. Chen, W.T. Wu, Electrospun polyacrylonitrile nanofibrous membranes for lipase immobilization, J.

597

Mol. Catal. B-Enzym. 47 (2007) 117-124.

598

[35] S.A. Ansari, Q. Husain, Potential applications of enzymes immobilized on/in nano materials: A review,

599

Biotechnol. Adv. 30 (2012) 512-523.

600

[36] F.E. Ahmed, B.S. Lalia, R. Hashaikeh, A review on electrospinning for membrane fabrication: Challenges and

601

applications, Desalination 356 (2015) 15-30.

602

[37] Z.G. Wang, L.S. Wan, Z.M. Liu, X.J. Huang, Z.K. Xu, Enzyme immobilization on electrospun polymer

603

nanofibers: An overview, J. Mol. Catal. B-Enzym. 56 (2009) 189-195.

604

[38] Y. Ispirli Dogac, I. Deveci, B. Mercimek, M. Teke, A comparative study for lipase immobilization onto

605

alginate based composite electrospun nanofibers with effective and enhanced stability, Int. J. Biol. Macromol. 96

606

(2017) 302-311.

607

[39] C. Isik, G. Arabaci, Y. Ispirli Dogac, I. Deveci, M. Teke, Synthesis and characterization of electrospun

608

PVA/Zn2+ metal composite nanofibers for lipase immobilization with effective thermal, pH stabilities and

609

reusability, Mat. Sci. Eng. C-Mater. Biol. Appl. 99 (2019) 1226-1235.

610

[40] X. Liu, Y. Fang, X. Yang, Y. Li, C. Wang, Electrospun epoxy-based nanofibrous membrane containing

611

biocompatible feather polypeptide for highly stable and active covalent immobilization of lipase, Colloid. Surface.

612

B 166 (2018) 277-285. 35

613

[41] F.A. Hasan, P. Xiao, R.K. Singh, P.A. Webley, Zeolite monoliths with hierarchical designed pore network

614

structure: Synthesis and performance, Chem. Eng. J. 223 (2013) 48-58.

615

[42] J. Ma, L. Zhang, Z. Liang, W. Zhang, Y. Zhang, Monolith-based immobilized enzyme reactors: Recent

616

developments and applications for proteome analysis, J. Sep. Sci. 30 (2007) 3050-3059.

617

[43] K.M. de Lathouder, T. Marques Fló, F. Kapteijn, J.A. Moulijn, A novel structured bioreactor: Development of

618

a monolithic stirrer reactor with immobilized lipase, Catal. Today 105 (2005) 443-447.

619

[44] S.M. Mugo, K. Ayton, Lipase immobilized methacrylate polymer monolith microreactor for lipid

620

transformations and online analytics, J. Am. Oil Chem. Soc. 90 (2012) 65-72.

621

[45] Y. Xiao, M. Zheng, Z. Liu, J. Shi, F. Huang, X. Luo, Constructing a continuous flow bioreactor based on a

622

hierarchically porous cellulose monolith for ultrafast and nonstop enzymatic esterification/transesterification, ACS

623

Sustain. Chem. Eng. 7 (2018) 2056-2063.

624

[46] X. Sun, Y. Xin, X. Wang, H. Uyama, Functionalized acetoacetylated poly(vinyl alcohol) monoliths for enzyme

625

immobilization: a phase separation method, Colloid Polym. Sci. 295 (2017) 1827-1833.

626

[47] E. Magner, Immobilisation of enzymes on mesoporous silicate materials, Chem. Soc. Rev. 42 (2013)

627

6213-6222.

628

[48] S. Hudson, J. Cooney, E. Magner, Proteins in mesoporous silicates, Angew. Chem. Int. Edit. 47 (2008)

629

8582-8594.

630

[49] W.B. Jin, Y. Xu, X.W. Yu, Improved catalytic performance of lipase under non-aqueous conditions by

631

entrapment into alkyl-functionalized mesoporous silica, New J. Chem. 43 (2019) 364-370.

632

[50] Z. Ali, L. Tian, P.P. Zhao, B.L. Zhang, N. Ali, M. Khan, Q.Y. Zhang, Immobilization of lipase on mesoporous

633

silica nanoparticles with hierarchical fibrous pore, J. Mol. Catal. B-Enzym. 134 (2016) 129-135.

634

[51] X.R. Xiang, H.B. Suo, C. Xu, Y. Hu, Covalent immobilization of lipase onto chitosan-mesoporous silica 36

635

hybrid nanomaterials by carboxyl functionalized ionic liquids as the coupling agent, Colloid. Surface. B 165 (2018)

636

262-269.

637

[52] M.M. Zheng, X. Xiang, S. Wang, J. Shi, Q.C. Deng, F.H. Huang, R.H. Cong, Lipase immobilized in ordered

638

mesoporous silica: A powerful biocatalyst for ultrafast kinetic resolution of racemic secondary alcohols, Process

639

Biochem. 53 (2017) 102-108.

640

[53] P. Falcaro, R. Ricco, C.M. Doherty, K. Liang, A.J. Hill, M.J. Styles, MOF positioning technology and device

641

fabrication, Chem. Soc. Rev. 43 (2014) 5513-5560.

642

[54] E. Gkaniatsou, C. Sicard, R. Ricoux, J.P. Mahy, N. Steunou, C. Serre, Metal-organic frameworks: a novel host

643

platform for enzymatic catalysis and detection, Mater. Horiz. 4 (2017) 55-63.

644

[55] J. Mehta, N. Bhardwaj, S.K. Bhardwaj, K.H. Kim, A. Deep, Recent advances in enzyme immobilization

645

techniques: metal-organic frameworks as novel substrates, Coordin. Chem. Rev. 322 (2016) 30-40.

646

[56] H.M. He, H.B. Han, H. Shi, Y.Y. Tian, F.X. Sun, Y. Song, Q.S. Li, G.S. Zhu, Construction of thermophilic

647

lipase-embedded metal organic frameworks via biomimetic mineralization: A biocatalyst for ester hydrolysis and

648

kinetic resolution, ACS Appl. Mater. Inter. 8 (2016) 24517-24524.

649

[57] T.J. Pisklak, M. Macias, D.H. Coutinho, R.S. Huang, K.J. Balkus, Hybrid materials for immobilization of

650

MP-11 catalyst, Top. Catal. 38 (2006) 269-278.

651

[58] A. Samui, A.R. Chowdhuri, S.K. Sahu, Lipase immobilized metal-organic frameworks as remarkably

652

biocatalyst for ester hydrolysis: A one step approach for lipase immobilization, ChemistrySelect 4 (2019)

653

3745-3751.

654

[59] A. Zare, A.K. Bordbar, F. Jafarian, S. Tangestaninejad, Candida rugosa lipase immobilization on various

655

chemically modified Chromium terephthalate MIL-101, J. Mol. Liq. 254 (2018) 137-144.

656

[60] X. Wu, C. Yang, J. Ge, Green synthesis of enzyme/metal-organic framework composites with high stability in 37

657

protein denaturing solvents, Bioresources and Bioprocessing 4 (2017) 24.

658

[61] K. Liang, C.J. Coghlan, S.G. Bell, C. Doonan, P. Falcaro, Enzyme encapsulation in zeolitic imidazolate

659

frameworks: a comparison between controlled co-precipitation and biomimetic mineralisation, Chem. Commun. 52

660

(2016) 473-476.

661

[62] L.Z. Cheong, Y.Y. Wei, H.B. Wang, Z.Y. Wang, X.R. Su, C. Shen, Facile fabrication of a stable and recyclable

662

lipase@amine-functionalized ZIF-8 nanoparticles for esters hydrolysis and transesterification, J. Nanopart. Res. 19

663

(2017) 280-291.

664

[63] K.S. Park, Z. Ni, A.P. Cote, J.Y. Choi, R.D. Huang, F.J. Uribe-Romo, H.K. Chae, M. O'Keeffe, O.M. Yaghi,

665

Exceptional chemical and thermal stability of zeolitic imidazolate frameworks, Proc. Natl. Acad. Sci. USA 103

666

(2006) 10186-10191.

667

[64] S. Rafiei, S. Tangestaninejad, P. Horcajada, M. Moghadam, V. Mirkhani, I. Mohammadpoor-Baltork, R.

668

Kardanpour, F. Zadehahmadi, Efficient biodiesel production using a lipase@ZIF-67 nanobioreactor, Chem. Eng. J.

669

334 (2018) 1233-1241.

670

[65] M. Adnan, K. Li, L. Xu, Y.J. Yan, X-Shaped ZIF-8 for immobilization Rhizomucor miehei lipase via

671

encapsulation and its application toward biodiesel production, Catalysts 8 (2018) 96-110.

672

[66] F. Pitzalis, C. Carucci, M. Naseri, L. Fotouhi, E. Magner, A. Salis, Lipase encapsulation onto ZIF-8: A

673

comparison between biocatalysts obtained at low and high zinc/2-methylimidazole molar ratio in aqueous medium,

674

ChemCatChem 10 (2018) 1578-1585.

675

[67] L. Qi, Z.G. Luo, X.X. Lu, Biomimetic mineralization inducing lipase-metal-organic framework

676

nanocomposite for pickering interfacial biocatalytic system, ACS Sustain. Chem. Eng. 7 (2019) 7127-7139.

677

[68] M. Adnan, K. Li, J.H. Wang, L. Xu, Y.J. Yan, Hierarchical ZIF-8 toward immobilizing Burkholderia cepacia

678

lipase for application in biodiesel preparation, Int. J. Mol. Sci. 19 (2018) 1424-1438. 38

679

[69] Y.L. Hu, L.M. Dai, D.H. Liu, W. Du, Rationally designing hydrophobic UiO-66 support for the enhanced

680

enzymatic performance of immobilized lipase, Green Chem. 20 (2018) 4500-4506.

681

[70] S.S. Nadar, V.K. Rathod, Encapsulation of lipase within metal-organic framework (MOF) with enhanced

682

activity intensified under ultrasound, Enzyme Microb. Tech. 108 (2018) 11-20.

683

[71] S.S. Nadar, V.K. Rathod, Magnetic-metal organic framework (magnetic-MOF): A novel platform for enzyme

684

immobilization and nanozyme applications, Int. J. Biol. Macromol. 120 (2018) 2293-2302.

685

[72] N. Nobakht, M.A. Faramarzi, A. Shafiee, M. Khoobi, E. Rafiee, Polyoxometalate-metal organic

686

framework-lipase: An efficient green catalyst for synthesis of benzyl cinnamate by enzymatic esterification of

687

cinnamic acid, Int. J. Biol. Macromol. 113 (2018) 8-19.

688

[73] S. Gupta, C.N. Murthy, C.R. Prabha, Recent advances in carbon nanotube based electrochemical biosensors,

689

Int. J. Biol. Macromol. 108 (2018) 687-703.

690

[74] W. Feng, X.C. Sun, P.J. Ji, Activation mechanism of Yarrowia lipolytica lipase immobilized on carbon

691

nanotubes, Soft Matter 8 (2012) 7143-7150.

692

[75] H.K. Lee, J.K. Lee, M.J. Kim, C.J. Lee, Immobilization of lipase on single walled carbon nanotubes in ionic

693

liquid, B. Kor. Chem. Soc. 31 (2010) 650-652.

694

[76] N.Z. Prlainovic, D.I. Bezbradica, Z.D. Knezevic-Jugovic, S.I. Stevanovic, M.L.A. Ivic, P.S. Uskokovic, D.Z.

695

Mijin, Adsorption of lipase from Candida rugosa on multi walled carbon nanotubes, J. Ind. Eng. Chem. 19 (2013)

696

279-285.

697

[77] C.X. Ke, X. Li, S.S. Huang, L. Xu, Y.J. Yan, Enhancing enzyme activity and enantioselectivity of Burkholderia

698

cepacia lipase via immobilization on modified multi-walled carbon nanotubes, Rsc. Adv. 4 (2014) 57810-57818.

699

[78] Akash Deep a, Amit L. Sharma b, Parveen Kumar a, Lipase immobilized carbon nanotubes for conversion of

700

Jatropha oil to fatty acid methyl esters, Biomass Bioenerg. 81 (2015) 83-87. 39

701

[79] M.M. Zheng, S. Wang, X. Xiang, J. Shi, J. Huang, Q.C. Deng, F.H. Huang, J.Y. Xiao, Facile preparation of

702

magnetic

703

1,3-dioleoyl-2-palmitoylglycerol-rich human milk fat substitutes, Food Chem. 228 (2017) 476-483.

704

[80] H.S. Tan, W. Feng, P.J. Ji, Lipase immobilized on magnetic multi-walled carbon nanotubes, Bioresour. Technol.

705

115 (2012) 172-176.

706

[81] Y.L. Fan, F. Su, K. Li, C.X. Ke, Y.J. Yan, Carbon nanotube filled with magnetic iron oxide and modified with

707

polyamidoamine dendrimers for immobilizing lipase toward application in biodiesel production, Sci. Rep-UK 7

708

(2017) 45643-45656.

709

[82] Y. Fan, G. Wu, F. Su, K. Li, L. Xu, X. Han, Y. Yan, Lipase oriented-immobilized on dendrimer-coated

710

magnetic multi-walled carbon nanotubes toward catalyzing biodiesel production from waste vegetable oil, Fuel 178

711

(2016) 172-178.

712

[83] L.T. Yu, I.A. Banerjee, X.Y. Gao, N. Nuraje, H. Matsui, Fabrication and application of enzyme-incorporated

713

peptide nanotubes, Bioconjugate Chem. 16 (2005) 1484-1487.

714

[84] M.Z. Anwar, D.J. Kim, A. Kumar, S.K.S. Patel, S. Otari, P. Mardina, J.H. Jeong, J.H. Sohn, J.H. Kim, J.T. Park,

715

J.K. Lee, SnO2 hollow nanotubes: a novel and efficient support matrix for enzyme immobilization, Sci. Rep. 7

716

(2017) 15333-15344.

717

[85] J. Sun, R. Yendluri, K. Liu, Y. Guo, Y. Lvov, X. Yan, Enzyme-immobilized clay nanotube-chitosan membranes

718

with sustainable biocatalytic activities, Phys. Chem. Chem. Phys. 19 (2016) 562-567.

719

[86] X. Cao, R. Zhang, W.M. Tan, C. Wei, J. Wang, Z.M. Liu, K.Q. Chen, P.K. Ouyang, Plasma treatment of

720

multi-walled carbon nanotubes for lipase immobilization, Korean J. Chem. Eng. 33 (2016) 1653-1658.

721

[87] W. Bai, Y.J. Yang, X. Tao, J.F. Chen, T.W. Tan, Immobilization of lipase on aminopropyl-grafted mesoporous

722

silica nanotubes for the resolution of (R, S)-1-phenylethanol, J. Mol. Catal. B-Enzym. 76 (2012) 82-88.

carbon

nanotubes-immobilized

lipase

40

for

highly

efficient

synthesis

of

723

[88] S.H. Lee, T.T.N. Doan, K. Won, S.H. Ha, Y.M. Koo, Immobilization of lipase within carbon nanotube-silica

724

composites for non-aqueous reaction systems, J. Mol. Catal. B-Enzym. 62 (2010) 169-172.

725

[89] A. Szelwicka, S. Boncel, S. Jurczyk, A. Chrobok, Exceptionally active and reusable nanobiocatalyst

726

comprising lipase non-covalently immobilized on multi-wall carbon nanotubes for the synthesis of diester

727

plasticizers, Appl. Catal. A-Gen. 574 (2019) 41-47.

728

[90] M.C. Bourkaib, Y. Guiavarc’h, I. Chevalot, S. Delaunay, J. Gleize, J. Ghanbaja, F. Valsaque, N. Berrada, A.

729

Desforges, B. Vigolo, Non-covalent and covalent immobilization of Candida antarctica lipase B on chemically

730

modified multiwalled carbon nanotubes for a green acylation process in supercritical CO2, Catal. Today (2019)

731

https://doi.org/10.1016/j.cattod.2019.08.046.

732

[91] S. Asmat, A.H. Anwer, Q. Husain, Immobilization of lipase onto novel constructed polydopamine grafted

733

multiwalled carbon nanotube impregnated with magnetic cobalt and its application in synthesis of fruit flavours, Int.

734

J. Biol. Macromol. 140 (2019) 484-495.

735

[92] M. Shim, N.W.S. Kam, R.J. Chen, Y.M. Li, H.J. Dai, Functionalization of carbon nanotubes for

736

biocompatibility and biomolecular recognition, Nano. Lett. 2 (2002) 285-288.

737

[93] K. Li, J. Wang, Y. He, M.A. Abdulrazaq, Y. Yan, Carbon nanotube-lipase hybrid nanoflowers with enhanced

738

enzyme activity and enantioselectivity, J. Biotechnol. 281 (2018) 87-98.

739

[94] Y. Lvov, W. Wang, L. Zhang, R. Fakhrullin, Halloysite clay nanotubes for loading and sustained release of

740

functional compounds, Adv. Mater. 28 (2016) 1227-1250.

741

[95] C. Chao, J.D. Liu, J.T. Wang, Y.W. Zhang, B. Zhang, Y.T. Zhang, X. Xiang, R.F. Chen, Surface modification of

742

halloysite nanotubes with dopamine for enzyme immobilization, ACS Appl. Mater. Inter. 5 (2013) 10559-10564.

743

[96] D.G.S. Shchukin, G. B. Price, R. R. Lvov, Y. M., Halloysite nanotubes as biomimetic nanoreactors, Small 1

744

(2005) 510-513. 41

745

[97] J. Xu, J. Sun, Y. Wang, J. Sheng, F. Wang, M. Sun, Application of iron magnetic nanoparticles in protein

746

immobilization, Molecules 19 (2014) 11465-11486.

747

[98] Y. Hu, S. Mignani, J.P. Majoral, M.W. Shen, X.Y. Shi, Construction of iron oxide nanoparticle-based hybrid

748

platforms for tumor imaging and therapy, Chem. Soc. Rev. 47 (2018) 1874-1900.

749

[99] J. Liu, Z. Sun, Y. Deng, Y. Zou, C. Li, X. Guo, L. Xiong, Y. Gao, F. Li, D. Zhao, Highly water-dispersible

750

biocompatible magnetite particles with low cytotoxicity stabilized by citrate groups, Angew. Chem. Int. Edit. 48

751

(2009) 5875-5879.

752

[100] X.Z. Zang, W.L. Xie, Immobilized lipase on core-shell structured Fe3O4-MCM-41 nanocomoposites as a

753

magnetically recyclable biocatalyst for interesterification of soybean oil and lard, Food Chem. 194 (2016)

754

1283-1292.

755

[101] J.M. Rosenholm, J.X. Zhang, W. Sun, H.C. Gu, Large-pore mesoporous silica-coated magnetite core-shell

756

nanocomposites and their relevance for biomedical applications, Micropor. Mesopor. Mat. 145 (2011) 14-20.

757

[102] D.M. Liu, J. Chen, Y.P. Shi, Screening of enzyme inhibitors from traditional Chinese medicine by magnetic

758

immobilized alpha-glucosidase coupled with capillary electrophoresis, Talanta 164 (2017) 548-555.

759

[103] D.M. Liu, J. Chen, Y.P. Shi, Alpha-Glucosidase immobilization on chitosan-enriched magnetic composites

760

for enzyme inhibitors screening, Int. J. Biol. Macromol. 105 (2017) 308-316.

761

[104] Y.F. Wang, F. Xu, L. Zhang, X.L. Wei, One-pot solvothermal synthesis of Fe3O4-PEI composite and its

762

further modification with Au nanoparticles, J. Nanopart. Res. 15 (2013) 1338-1349.

763

[105] M. Martin, P. Salazar, R. Villalonga, S. Campuzano, J.M. Pingarron, J.L. Gonzalez-Mora, Preparation of

764

core-shell Fe3O4@poly(dopamine) magnetic nanoparticles for biosensor construction, J. Mater. Chem. B 2 (2014)

765

739-746.

766

[106] Y. Liu, S.Y. Jia, Q.A. Wu, J.Y. Ran, W. Zhang, S.H. Wu, Studies of Fe3O4-chitosan nanoparticles prepared by 42

767

co-precipitation under the magnetic field for lipase immobilization, Catal. Commun. 12 (2011) 717-720.

768

[107] Y.H. Ren, J.G. Rivera, L.H. He, H. Kulkarni, D.K. Lee, P.B. Messersmith, Facile, high efficiency

769

immobilization of lipase enzyme on magnetic iron oxide nanoparticles via a biomimetic coating, Bmc. Biotechnol.

770

11 (2011) 63-71.

771

[108] K. Li, J.H. Wang, Y.J. He, G.L. Cui, M.A. Abdulrazaq, Y.J. Yan, Enhancing enzyme activity and

772

enantioselectivity of Burkholderia cepacia lipase via immobilization on melamine-glutaraldehyde dendrimer

773

modified magnetic nanoparticles, Chem. Eng. J. 351 (2018) 258-268.

774

[109] Y. Deng, D. Qi, C. Deng, X. Zhang, D. Zhao, Superparamagnetic high-magnetization microspheres with an

775

Fe3O4@SiO2 core and perpendicularly aligned mesoporous SiO2 shell for removal of microcystins, J. Am. Chem.

776

Soc. 130 (2008) 28-29.

777

[110] M. Bilal, Y.P. Zhao, T. Rasheed, H.M.N. Iqbal, Magnetic nanoparticles as versatile carriers for enzymes

778

immobilization: A review, Int. J. Biol. Macromol. 120 (2018) 2530-2544.

779

[111] K. Pashangeh, M. Akhond, H.R. Karbalaei-Heidari, G. Absalan, Biochemical characterization and stability

780

assessment of Rhizopus oryzae lipase covalently immobilized on amino-functionalized magnetic nanoparticles, Int.

781

J. Biol. Macromol. 105 (2017) 300-307.

782

[112] W.W. Huan, Y.X. Yang, B. Wu, H.M. Yuan, Y.N. Zhang, X.N. Liu, Degradation of 2,4-DCP by the

783

immobilized laccase on the carrier of Fe3O4@SiO2-NH2, Chinese J. Struc. Chem. 30 (2012) 2849-2860.

784

[113] Z.X. Cai, Y. Wei, M. Wu, Y.L. Guo, Y.P. Xie, R. Tao, R.Q. Li, P.G. Wang, A.Q. Ma, H.B. Zhang, Lipase

785

immobilized on layer-by-layer polysaccharide-coated Fe3O4@SiO2 microspheres as a reusable biocatalyst for the

786

production of structured lipids, ACS Sustain. Chem. Eng. 7 (2019) 6685-6695.

787

[114] L.N. de Lima, G.N.A. Vieira, W. Kopp, P.W. Tardioli, R.L.C. Giordano, Mono- and heterofunctionalized

788

silica magnetic microparticles (SMMPs) as new carriers for immobilization of lipases, J. Mol. Catal. B-Enzym. 133 43

789

(2016) S491-S499.

790

[115] T. Itoh, Ionic liquids as tool to improve enzymatic organic synthesis, Chem. Rev. 117 (2017) 10567-10607.

791

[116] J.L. Kaar, A.M. Jesionowski, J.A. Berberich, R. Moulton, A.J. Russell, Impact of ionic liquid physical

792

properties on lipase activity and stability, J. Am. Chem. Soc. 125 (2003) 4125-4131.

793

[117] J. Qin, X.Q. Zou, S.T. Lv, Q.Z. Jin, X.G. Wang, Influence of ionic liquids on lipase activity and stability in

794

alcoholysis reactions, Rsc. Adv. 6 (2016) 87703-87709.

795

[118] R. Patel, M. Kumari, A.B. Khan, Recent advances in the applications of ionic liquids in protein stability and

796

activity: A review, Appl. Biochem. Biotech. 172 (2014) 3701-3720.

797

[119] J.J. Xia, B. Zou, G. B. Zhu, P. Wei, Z.J. Liu, Quick separation and enzymatic performance improvement of

798

lipase by ionic liquid-modified Fe3O4 carrier immobilization, Bioproc. Biosyst. Eng. 41 (2018) 739-748.

799

[120] Y.Y. Jiang, C. Guo, H.S. Gao, H.S. Xia, I. Mahmood, C.Z. Liu, H.Z. Liu, Lipase immobilization on ionic

800

liquid-modified magnetic nanoparticles: Ionic liquids controlled esters hydrolysis at oil-water interface, Aiche. J. 58

801

(2012) 1203-1211.

802

[121] H. Suo, Z. Gao, L. Xu, C. Xu, D. Yu, X. Xiang, H. Huang, Y. Hu, Synthesis of functional ionic liquid

803

modified magnetic chitosan nanoparticles for porcine pancreatic lipase immobilization, Mat. Sci. Eng. C-Mater. 96

804

(2019) 356-364.

805

[122] H.B. Suo, L.L. Xu, C. Xu, H.Y. Chen, D.H. Yu, Z. Gao, H. Huang, Y. Hu, Enhancement of catalytic

806

performance of porcine pancreatic lipase immobilized on functional ionic liquid modified Fe3O4-chitosan

807

nanocomposites, Int. J. Biol. Macromol. 119 (2018) 624-632.

808

[123] J. Wang, G. Zhao, F. Yu, Facile preparation of Fe3O4@MOF core-shell microspheres for lipase

809

immobilization, J. Taiwan Inst. Chem. 69 (2016) 139-145.

810

[124] G. Sargazi, D. Afzali, A.K. Ebrahimi, A. Badoei-Dalfard, S. Malekabadi, Z. Karami, Ultrasound assisted 44

811

reverse micelle efficient synthesis of new Ta-MOF@Fe3O4 core/shell nanostructures as a novel candidate for lipase

812

immobilization, Mater. Sci. Eng. C-Mater. Biol. Appl. 93 (2018) 768-775.

813

[125] M.R. Mehrasbi, J. Mohammadi, M. Peyda, M. Mohammadi, Covalent immobilization of Candida antarctica

814

lipase on core-shell magnetic nanoparticles for production of biodiesel from waste cooking oil, Renew. Energ. 101

815

(2017) 593-602.

816

[126] H.S. Qi, Y. Du, G.N. Hu, L. Zhang, Poly(carboxybetaine methacrylate)-functionalized magnetic composite

817

particles: A biofriendly support for lipase immobilization, Int. J. Biol. Macromol. 107 (2018) 2660-2666.

818

[127] M. Aghababaie, M. Beheshti, A.-K. Bordbar, A. Razmjou, Novel approaches to immobilize Candida rugosa

819

lipase on nanocomposite membranes prepared by covalent attachment of magnetic nanoparticles on poly

820

acrylonitrile membrane, Rsc. Adv. 8 (2018) 4561-4570.

821

[128] P.C. Chen, Z. Ma, X.Y. Zhu, D.J. Chen, X.J. Huang, Fabrication and optimization of a lipase immobilized

822

enzymatic membrane bioreactor based on polysulfone gradient-pore hollow fiber membrane, Catalysts 9 (2019)

823

495-506.

824

[129] Y. Li, H. Wang, J. Lu, A. Chu, L. Zhang, Z. Ding, S. Xu, Z. Gu, G. Shi, Preparation of immobilized lipase by

825

modified polyacrylonitrile hollow membrane using nitrile-click chemistry, Bioresour. Technol. 274 (2019) 9-17.

826

[130] D.M. Liu, J. Chen, Y.P. Shi, alpha-Glucosidase immobilization on chitosan-modified cellulose filter paper:

827

Preparation, property and application, Int. J. Biol. Macromol. 122 (2019) 298-305.

828

[131] H. Koga, T. Kitaoka, A. Isogai, Paper-immobilized enzyme as a green microstructured catalyst, J. Mater.

829

Chem. 22 (2012) 11591-11597.

830

[132] L.E. Shu, Z.X. Tang, Y. Yi, J.S. Chen, H. Wang, W.Y. Xiong, G.Q. Ying, Study of immobilization of nuclease

831

P1 on paper cellulose, Biotechnol. Biotechnol. Equip. 24 (2010) 1997-2003.

832

[133] C.S.K. Lawrence, S.N. Tan, C.Z. Floresca, A "green" cellulose paper based glucose amperometric biosensor, 45

833

Sensor Actuat. B-Chem. 193 (2014) 536-541.

834

[134] C. Tyagi, L.K. Tomar, H. Singh, Surface modification of cellulose filter paper by glycidyl methacrylate

835

grafting for biomolecule immobilization: Influence of grafting parameters and urease immobilization, J. Appl.

836

Polym. Sci. 111 (2009) 1381-1390.

837

[135] H. Sugiyama, Y. Akazome, T. Shoji, A. Yamaguchi, M. Yasue, T. Kanda, Y. Ohtake, Oligomeric procyanidins

838

in apple polyphenol are main active components for inhibition of pancreatic lipase and triglyceride absorption, J.

839

Agric. Food Chem. 55 (2007) 4604-4609.

840

[136] M. Nakai, Y. Fukui, S. Asami, Y. Toyoda-Ono, T. Iwashita, H. Shibata, T. Mitsunaga, F. Hashimoto, Y. Kiso,

841

Inhibitory effects of Oolong tea polyphenols on pancreatic lipase in vitro, J. Agric. Food Chem. 53 (2005)

842

4593-4598.

843

[137] Y.T. Zhu, X.Y. Ren, Y.M. Liu, Y. Wei, L.S. Qing, X. Liao, Covalent immobilization of porcine pancreatic

844

lipase on carboxyl-activated magnetic nanoparticles: Characterization and application for enzymatic inhibition

845

assays, Mater. Sci. Eng. C-Mater. Biol. Appl. 38 (2014) 278-285.

46

Highlights •

Immobilized enzyme strategy has been currently recognized as a powerful tool in the field of inhibitors screening.

•

A variety of materials have been systematically introduced and employed as the supports for lipase immobilization.

•

Lipase inhibitors screening based on the immobilized enzyme technology has been detailed presented.

•

Future perspectives of novel supports for immobilizing lipase and screening techniques for inhibitors are stated as well.

Jia Liu, current M.A. student at the Lanzhou Institute of Chemical Physics, Chinese Academy of Sciences (CAS). She obtained her B.E. degree in 2017, at the HeFei University of Technology. Her research interests are mainly centred on enzyme inhibitor screening, especially the application of immobilized enzyme technology coupled with capillary electrophoresis for enzyme inhibitor screening.

Run-Tian Ma received her Ph.D. from Lanzhou Institute of Chemical Physics, Chinese Academy of Sciences (CAS) in 2016. She is currently an associate research fellow in Lanzhou Institute of Chemical Physics, Chinese Academy of Sciences (CAS). Her research interests are mainly on sample pretreatment technology, especially molecular imprinting technology and solid-phase microextraction.

Yan-Ping Shi was a Full Professor, Ph.D. Supervisor, and Former Deputy Director of the Lanzhou Institute of Chemical Physics (LICP), Chinese Academy of Sciences (CAS), P. R. China. Shi was received his Ph.D. and M.Sc. degrees in organic chemistry from Lanzhou University, China in 1996 and 1992, respectively. He completed periods as a post-doctoral research associate at University of Puerto Rico, USA in 2001 and at Lanzhou University in 1998. He has been awarded by the hundred talent program of the Chinese Academy of Sciences for 2000. He obtained a permanent position in LICP, CAS, as a Research Professor during 2001 to now and was the vice Director of LICP, CAS during 2012 to 2018. Prof. Shi has authored over 400 scientific publications. He was honored with two 2nd Prizes for the Advancement of Science & Technology by the National Education Department of China in 2005 and 1999, respectively, as well as three 2nd Prizes for the Natural Sciences by Gansu of China in 2019, 2016, 2007, respectively. His research interests cover the analysis chemistry, especially focus on sample pretreatment technology and chromatographic analysis.

Declaration of interests The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper. ☐The authors declare the following financial interests/personal relationships which may be considered as potential competing interests: