Penicillium expansum lipase-coated magnetic Fe₃O₄–polymer hybrid hollow nanoparticles: a highly recoverable and magnetically separable catalyst for the synthesis of 1,3-dibutylurea

Abstract

Herein, amino-epoxy supports were innovatively imported onto magnetic nanoparticles (Fe₃O₄–polymer hybrid nanospheres) for immobilizing enzymes. This new support has a coating layer with dual functional groups (epoxy and amino-epoxy). Consequently, this support has great anionic exchange power and a high number of epoxy groups. The acquired immobilized Penicillium expansum lipase in combination with this heterofunctional support represents a novel class of heterogeneous catalyst towards the synthesis of 1,3-dibutylurea from ethylene carbonate and butylamine, which has not been very commonly catalyzed by enzymes. After optimization of the reaction conditions, the yield of 1,3-dibutylurea was 77% under solvent free conditions at 60 °C. Moreover, after completion of reaction, the catalyst was simply recovered by an external conventional magnet and recycled without significant loss in the catalytic activity (up to ten cycles).

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Introduction

Enzymes are versatile biocatalysts with high specificity, and their applications have been found in areas of biomaterials and organic synthesis.^1,2 However, the major drawbacks for their industrial applications are their low thermal and solvent stability as well as difficulty in recycling. Applications of lipase can be achieved more economically and efficiently by immobilization to enhance its activity, selectivity, and operational stability.^3–8 For example, the activity of lipase could be improved by adsorption of lipase on hydrophobic supports through the hydrophobic areas surrounding the active centre and thus stabilizing the open form of the lipase.⁹ Enantiomeric ratio may be changed from 1 to almost 100 simply by using different immobilized preparations in some cases.^10,11 Moreover, the stability of lipase could be enhanced through rigidification of the enzyme structure by multipoint covalent immobilization.^12,13 Therefore, a lot of fruitful efforts have been made on the preparation of lipases in immobilized forms, which involve a variety of both support materials and immobilization methods.^14–19

Nanophase materials have many advantages in immobilization due to their unique size and physical properties.^9,20,21 Lipases, immobilized onto nanoparticles, which have a high specific surface area and low diffusion resistance, possess much higher catalytic activity and stability despite their cumbersome preparation, painful separation and poor efficiency. Currently, using magnetic properties is a pinpoint protocol to solve these problems. With the rapid development of nanotechnology, magnetic nanoparticles for enzyme immobilization are now being studied extensively all over the world.^22–29 However, immobilization on nanoparticles has also some drawbacks: enzymes immobilized on nanoparticles can easily lose their activity due to the interaction with external interfaces or even with the enzyme molecules immobilized on other particles. This may be solved by covering the immobilized enzymes with polymers, which could provide a porous structure to stabilize the enzyme.³⁰

Nowadays, with the increasing use of green chemistry, magnetic chitosan (CS)–Fe₃O₄ nanoparticles with their potential application for enzyme immobilization have attracted extensive interest because of chitosan's excellent properties such as non-toxicity, biocompatibility, mucus-adhesion and biodegradation.^31–41 Zhi et al.⁴² prepared magnetic CS–Fe₃O₄ nanoparticles in situ with small pools of a water-in-oil microemulsion, which contains CS and the ferrous salt as micro-reactors, by adding a basic precipitant such as NaOH into the micro-emulsion. The defect of this method is poor productivity; in addition, it uses a lot of surfactant and co-surfactant compounds that are pernicious to the environment. Wu et al.⁴³ prepared magnetic CS–Fe₃O₄ nanoparticles by the covalent binding of CS and tripolyphosphate onto Fe₃O₄ nanoparticles by a one-step method. The problem of this method is that the magnetisation process is complicated and hard to control. Siódmiak et al.⁴⁴ prepared CS–poly[N-benzyl-2-(methacryloxy)-N,N-dimethylethanaminium bromide]-coated magnetic nanoparticles by co-precipitation method via the epichlorohydrin CS cross-linking reaction. The disadvantage of this method is that it uses some poisonous compounds, such as benzyl bromide, which is nocuous and not environmentally benign. Ding et al.⁴⁵ illustrated a successful and green synthetic route of magnetic hollow Fe₃O₄–polymer hybrid nanospheres (MHHNs) by the addition of Fe₃O₄ nanoparticles to an aqueous solution of polymer–monomer pairs composed of the cationic polymer, CS, and the anionic monomer, acrylic acid (AA), which is followed by polymerization of AA and selective crosslinking of CS at the end of the polymerization process. This innovative method with the highlights of simple operation, high performance, cost-effectiveness, and environmental friendliness, will usher in rosy prospects for magnetic nanospheres in the application of immobilized enzymes.

In this paper, we introduce this kind of magnetic CS–Fe₃O₄ nanoparticles into the immobilization of lipases. MHHNs were activated by epoxy chloropropane. After that, Penicillium expansum lipase (PEL) was immobilized on this support by the covalent linkage between the epoxy groups on the support surface and the amino group on the enzymes surface to form the immobilized PEL magnetic Fe₃O₄–polymer hybrid nanospheres (MHHNs-lip). Recently, the production, purification, and application of PEL have been discussed in various reviews.^46,47 Immobilization on this kind of support follows a two-step mechanism:⁴⁸ (1) an ionic exchange with the amino groups in the support or hydrophobic adsorption is the first step in the immobilization process. (2) The protein, which has been previously adsorbed, is covalently attached to the epoxy groups present on the surface of the support. Finally, to complete the protein-support reaction, the epoxy groups can be easily blocked by reaction with very different thiol or amine compounds under mild conditions,⁴⁹ preventing further uncontrolled reactions between the support and the enzyme, which could decrease its stability. Obviously, the protocol is simple and allows the large-scale preparation of other enzymes, like Aspergillus niger lipase and Rhizopus chinensis satio lipase, which are still under investigation in our laboratory.

At the same time, we have investigated the application of MHHNs-lip for the synthesis of 1,3-disubstituted ureas, which are very essential intermediates in the preparation of pharmaceuticals and biochemicals. 1,3-Disubstituted ureas have extensive applications such as dyes, antioxidants, corrosion inhibitors, and intermediates for the preparation of pharmaceuticals and agricultural chemicals.^50–52 Recently, these compounds have attracted extensive interest because substituted ureas, which contain amino acid groups, have been used for brain cancer treatment and have been proven to have a marked inhibitory effect on the HIV protease enzyme.⁵² The currently studied alternative routes for the synthesis of ureas through phosgene substitutes are as follows: (a) the oxidative carbonylation of amines with carbon monoxide under various metal catalysts,^53–56 (b) the direct reaction of carbon dioxide with amines,^57,58 and (c) the use of some phosgene substitutes such as carbonates, which are less toxic and more stable.⁵⁹ Another strategy to synthesize 1,3-disubstituted urea is the composite of an amine and ethylene carbonate (EC) (Scheme 1). Although ethylene carbonate is a green reagent and solvent that is industrially obtained from an insertion reaction of carbon dioxide into epoxides,^60,61 some base catalysts such as NaOMe,⁶² CaO,⁶³ 1,5,7-triazabicyclo[4.4.0]dec-5-ene and thioureas⁶⁴ or Cs₂CO₃,⁶⁵ were used to catalyze this reaction. However, all of these catalysts operate under extreme conditions (above 90 °C) and are complicated to recycle. Moreover, most of these conventional catalysts contain metals, which are considered to be environmentally hazardous. Nowadays, with a universal awareness of environmental protection, eliminating the metal component involvement in catalyst design and avoiding volatile organic solvent usage in the process of chemical synthesis is more important and necessary for our future.

Scheme 1 Synthesis of 1,3-disubstituted urea from ethylene carbonate and amine.

In this study, we extended the scope of application of these nanoparticles in the synthesis of 1,3-disubstituted ureas. We report the immobilization of PEL-coated magnetic hollow Fe₃O₄–polymer hybrid nanospheres, which represents a novel class of heterogeneous catalyst for the synthesis of 1,3-dibutylurea from EC and butylamine. The main advantages behind using the immobilized PEL magnetic Fe₃O₄–polymer hybrid nanospheres (MHHNs-lip) as the catalyst are as follows: (1) compared to chemical catalysts, lipase is regarded as an optimized potential candidate for environmentally friendly reactions. (2) the synthesis of MHHNs-lip is simple and allows the large-scale preparation of nanospheres, which also follows an eco-friendly and non-toxic route. (3) The catalyst can be easily separated by an external magnet without using extra organic solvents and additional filtration steps. (4) The chemical reaction with the catalyst of MHHNs-lip can be carried out under mild conditions (60 °C and atmospheric pressure). All of these advantages are in agreement with the principles of green chemistry and sustainable development.

Results and discussion

Characterization of immobilized lipase

Typical TEM micrographs of pure Fe₃O₄ and the MHHNs-lip nanoparticles are shown in Fig. 1. A salient feature of Fig. 1b–d is that these nanospheres have an intense dark circle within the shells of the spheres and dark spots on the surface of some spheres, which suggests that the Fe₃O₄ particles were coated by polymer acrylic acid (PAA), CS and PEL. The mean size of Fe₃O₄ particles was 30 ± 5 nm.

Fig. 1 TEM images of the (a) bare Fe₃O₄ nanoparticles and (b–d) MHHNs-lip.

The thermal stability of the samples was investigated by thermogravimetric analysis (TGA). The TGA curves for the particles of bare Fe₃O₄, PEL, MHHNs, and MHHNs-lip are shown in Fig. 2. As shown in Fig. 2, in all the samples, the weight loss within 200 °C was attributed completely to the loss of adsorbed water molecules.⁶⁶ MHHNs showed a higher weight loss than bare Fe₃O₄ because of the loss of the PAA and CS components (Fig. 2b). After immobilization of PEL, the weight loss shown in Fig. 2c increased again, which was due to the increased organic components on the MHHNs-lip. From this figure, the protein loading content was determined to be 56 mg g⁻¹, which was very close to the calculated value (the total amount of PEL protein assayed according to the Coomassie Brilliant Blue G-250 method was 58 mg g⁻¹).

Fig. 2 TGA of (a) bare Fe₃O₄, (b) MHHNs, (c) MHHNs-lip, and (d) Penicillium expansum.

Fig. 3 shows the FT-IR spectra of (a) Fe₃O₄ nanoparticles, (b) Penicillium expansum, and (c) MHHNs-lip. There was a strong peak observed at 598 cm⁻¹ in Fig. 3a, assigned to the characteristic absorbance of Fe₃O₄.⁶² In addition, the peaks at 1622 and 1362 cm⁻¹ were assigned to the absorbance of the asymmetric and symmetric stretching vibration of COO⁻ of the citric acid modifier, respectively, indicating the presence of Fe₃O₄ nanoparticles coated by the COO⁻ group.⁶⁷ The FT-IR spectra of free lipase contained a broad envelope at 3421 cm⁻¹ due to the overlap of the –NH stretching of the amide group in the proteins (Fig. 3b). The FT-IR absorption spectrum of lipase generally shows three major bands caused by peptide group vibrations in the range of 1800–1300 cm⁻¹.⁶⁸ The amide I band at 1605 cm⁻¹ is mainly due to C–O stretching vibrations, whereas free lipase illustrates a characteristic amide II band with a maximum at 1414 cm⁻¹ due to N–H bending with contribution from C–N stretching vibrations. The presence of the amide III band with a maximum at 1323 cm⁻¹ is due to N–H bending with C–Cα and C–N stretching vibrations. The absorption band at 2938 cm⁻¹ was due to the stretching vibrations of C–H. These significant bands were observed in the free lipase as well as MHHNs-lip (Fig. 3c), indicating that the lipase was successfully coated onto the surface of the Fe₃O₄ magnetic nanoparticles. The FT-IR spectrum of MHHNs-lip (Fig. 3c) shows that the amide I and II bands of the purified acidic lipase overlapped at 1647 cm⁻¹ with the absorption bands of the C=O groups in PAA and the ammonium groups in CS. This band became weaker, which may be due to the strong hydrogen bonding between Fe₃O₄ and lipase.⁶⁹ The characteristic peak for the Fe–O group of magnetite at 598 cm⁻¹ can be observed in Fig. 3c, which indicates the successful generation of MHHNs-lip particles.

Fig. 3 FT-IR spectra of: (a) bare Fe₃O₄, (b) Penicillium expansum, and (c) MHHNs-lip.

The magnetic properties of the magnetic nanoparticles were analyzed by VSM at room temperature. Fig. 4 shows the hysteresis loops of the samples. The saturation magnetization was found to be 38.79 emu g⁻¹ for MHHNs, which is less than that for the pure Fe₃O₄ nanoparticles (67.08 emu g⁻¹). This difference suggests that a large amount of chitosan and polymer of acrylic acid was coated onto the surface of the Fe₃O₄ nanoparticles. The lower saturation magnetization of MHHNs-lip (25.99 emu g⁻¹) compared with MHHNs suggests that PEL was immobilized successfully in the MHHNs. With large saturation magnetization, MHHNs-lip could be separated from the reaction medium rapidly and easily in a magnetic field. In addition, there is no hysteresis in the magnetization with both the remanence and coercivity being zero, suggesting that these magnetic nanoparticles are superparamagnetic. Moreover, when the external magnetic field is removed, the magnetic nanoparticles could be dispersed by gentle shaking. These magnetic properties are critical in the applications within the biomedical and bioengineering fields.

Fig. 4 Magnetic hysteresis curves of (a) bare Fe₃O₄, (b) MHHNs, and (c) MHHNs-lip.

Fig. 5 shows the XRD patterns of pure Fe₃O₄, MHHNs and MHHNs-lip. It is apparent that the diffraction pattern of our Fe₃O₄ nanoparticles is similar to the standard pattern for crystalline magnetite (Fig. 5a). The characteristic diffraction peaks marked by their indices (220), (311), (400), (422), (511), and (440) could be indexed to the inverse cubic spinel structure of Fe₃O₄ (JCPDS card no. 85-1436), and they were also observed for MHHNs (Fig. 5b) and MHHNs-lip (Fig. 5c). This reveals that the grafted and polymerized process, on the surface of Fe₃O₄ nanoparticles did not lead to their crystal phase change. The average crystallite size D was about 30.3 nm, which is in good agreement with that measured by TEM. The average crystallite size was obtained from the Sherrer equation D = Kλ/(β cos θ), where K is constant, λ is the X-ray wavelength, and β is the peak width of half-maximum. However, due the existence of the organic layer (CS and PAA) on the surface of the Fe₃O₄ nanoparticles in MHHNs (Fig. 5b), it may lead to a decrease in the intensity for the Fe₃O₄ specific peaks (Fig. 5a), which could also explain the decrease in intensity in Fig. 5c compared to Fig. 5b, because of the extra lipase layer on the surface of MHHNs.

Fig. 5 XRD pattern of (a) bare Fe₃O₄, (b) MHHNs, and (c) MHHNs-lip.

The preparation procedure is illustrated schematically in Scheme 2. We selected CS, which bears amino groups, as the cationic polymer and AA, which bears acid groups, as the anionic monomer. The CS and AA polymer–monomer pair and Fe₃O₄ nanoparticles stabilized by poly(vinyl alcohol) (PVA) were mixed and formed micelles loaded with Fe₃O₄ nanoparticles; the cores consist of the polyionic complexes of CS and AA (i.e. positively charged protonated CS chains and negatively charged dissociated AA) and the shells consist of protonated CS chains. Initiation of the polymerization of AA with potassium persulfate (K₂S₂O₈) followed by cross-linking of the shells with glutaraldehyde (GA) at the end of the polymerization process led to the formation of magnetic hollow Fe₃O₄–polymer hybrid nanospheres.⁴⁵

Scheme 2 Illustration of the preparation, activation of the epoxy groups and immobilization of the lipase.

Following the procedures described in the literature, it is hard to give an exact structure of the groups formed by the GA. However, whatever the exact structure of the GA on the support,^70,71 the final result is a support having a fairly hydrophobic moiety formed by the GA chains and free amino groups⁷² (the ratio⁷³ of aldehyde groups from GA to the amino groups from CS was 1 : 2). In the next activation procedure with epoxy chloropropane, the epoxy activation will proceed via the amino, hydroxyl⁶⁷ and carboxyl groups.⁷⁴ Therefore, the final support will be an amino-epoxy heterofunctional one (Scheme 3), which contributes a lot to promote the absorption via ionic exchange at low ionic strength.⁷⁵ However, due to the tendency of the open form of the lipases to become adsorbed versus the hydrophobic interface, even though immobilization is performed at low ionic strength, the enzyme will still be immobilized by both immobilization mechanisms: interfacial activation and ionic exchange.^76–78 After enzyme hydrophobic adsorption, the reactive groups of the enzyme near the support surface may produce some covalent bonding reactions, which may be produced after enzyme immobilization, and there is no guarantee that the enzyme will finally have any covalent attachment with the support.⁷⁵ Therefore, after completion of the immobilization, in order to protein-support reaction, aspartic acid was added to block the epoxy groups to prevent further uncontrolled reactions between the support and the enzyme that could decrease its stability.^48,49

Scheme 3 Illustration of the activation of the epoxy groups.

On the basis of the two-step mechanism of enzyme immobilization,^67,79 the adsorption of the enzymes on the support surface was the first step. In the second step, with enzymes absorbed onto the surface of the support successfully, the high concentration of reactive groups in the PEL and the epoxy groups in the support could make a rapid “intramolecular” covalent reaction (Scheme 4). It has been reported that the probability of the formation of multiple covalent linkages between PEL and epoxy could be enhanced, leading to an increase in the stability of the PEL immobilized microspheres.^67,80 The immobilization of the enzymes inside the hollow MHHNs may increase the stability by preventing any intermolecular process and by preventing the enzyme from interacting with external interfaces.⁸¹ Therefore, the cavities in the cellulose microspheres could provide not only a space for storage of the enzyme but also a wall shell to protect its structure and nature.

Scheme 4 Illustration of the immobilization of lipase.

Screening of the biocatalyst for 1,3-dibutylurea synthesis

The catalytic activity of lipase from various natural sources has been directly evaluated in the transesterification of EC with butylamine. All the available lipases were screened based on their activities for the production of 1,3-dibutylurea. As shown in Table 1, PEL showed high catalytic performance for the production of 1,3-dibutylurea, while other lipases revealed little activity. Based on these results, we selected PEL as the lipase for immobilization.

Table 1 Screening the lipase sources for 1,3-dibutylurea synthesis. Conditions: catalyst: 0.7 g; EC: 40 mmol; butylamine: 80 mmol; temperature: 60 °C; time: 24 h

Entry	Catalyst	Yield (%)
1	Penicillium expansum	69
2	Penicillium neutral expannsum	37
3	Aspergillus niger	28
4	Aspergillus oryzae	15
5	Rhizopus chinensis satio	9
6	Porcine pancreatic	6
7	Novozym 435	13
8	Neural proteolytic enzyme	5
9	Acidic proteolytic enzyme	3
10	Alkaline proteolytic enzyme	4

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As seen in Table 2, when bare Fe₃O₄ was used in the reaction medium, the yield of reaction was 6% in 24 h. Using bare MHHNs in the reaction at room temperature gave a 4% yield. In the experiment using MHHNs-lip gave 77% yield. This value is higher than that using the same lipase in its free form, where the yields ranged from 69% to 77%. It seems that after immobilization, there was an enhancement of the enzymes activity, as shown by Dalla-Vecchia in esterification reactions of carboxylic acids with n-pentanol in organic media by Rhyzopus oryzae lipase.⁸² However, the activity of the immobilized lipase decreased when PEL was immobilized onto conventional epoxy supports (epoxy Sepabeads). This different effect on the enzyme activity of the immobilization when using the different supports (although both of them have epoxy groups as the reactive group) may be related to different enzyme orientations of the enzyme on the support. This indicates that the enzyme likely reacts with the support via different areas (the areas of the protein surface that have more hydrophobic residues or areas of the protein that have more negative charge) with concomitant different effects on enzyme activity.⁴⁸

Table 2 Synthesis of 1,3-dibutylurea. Conditions: catalyst: 0.7 g; EC: 40 mmol; butylamine: 80 mmol; temperature: 60 °C; time: 24 h

Entry	Catalyst	Yield (%)
1	PEL	69
2	Fe3O4	6
3	MHHNs	4
4	MHHNs-lip	77
5	Epoxy sepabeads-lip	70

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The temperature influence on the transesterification process has also been investigated. As seen from Table 3, the temperature increase is favourable for the production of 1,3-dibutylurea, since the enzyme activity increases with an increase in temperature. The yield of 1,3-dibutylurea increased gradually with the temperature till a maximum at 60 °C. However, a further increase in temperature (up to 70 °C) led to a slight decrease in the yield of 1,3-dibutylurea. Nevertheless, we adopted 60 °C for this work, which is in agreement with the our earlier reports for the appropriate reaction temperature of this lipase.⁸³

Table 3 Influence of reaction temperature on the synthesis of 1,3-dibutylurea. Conditions: MHHNs-lip catalyst: 0.7 g; EC: 40 mmol; butylamine: 80 mmol; time: 24 h

Temperature (°C)	Yield (%)
20	9
30	23
40	35
50	52
60	77
70	76

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As shown in Table 4, the influence of incubation time on the transesterification process was also investigated. It was apparent that increasing the reaction time from 0 to 24 h is more advantageous for the reaction process, giving rise to improved yields. When increasing the reaction time further from 24 h to 36 h, no apparent change on the yield of 1,3-dibutylurea yield can be seen, indicating the appropriate reaction time was 24 h.

Table 4 Influence of incubation time on the 1,3-dibutylurea synthesis. Conditions: MHHNs-lip catalyst: 0.7 g; EC: 40 mmol; butylamine: 80 mmol; temperature: 60 °C

Incubation time (h)	Yield (%)
1	22
3	34
6	46
12	58
24	77
36	77

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The recyclability and storage stability of lipase

To make the process more economical, it is necessary to study the recyclability of the immobilized lipase, which is regarded as the most important advantage of immobilized enzymes when compared to the free enzyme. After completion of the reaction, the catalyst in the free PEL catalyst reaction was recovered by centrifugation of the resulting suspension and washed using acetone. The recovery free PEL obtained was dried at 45 °C under reduced pressure overnight (at 1 Torr for 24 h), and then used for the next reaction. At the same time, the catalyst in the MHHNs-lip catalysed reaction was magnetically separated from the reaction medium directly (Fig. 6) and run in another reaction vessel under the same conditions. The residual activity of both catalysts during their reuse is presented in Fig. 7, which shows that MHHNs-lip has a better recyclability compared to free PEL. This suggests that the lipase immobilized on MHHNs may display a higher stability than the original lipase. The activity of the first batch was taken to be 100%. The MHHNs-lip exhibited a better reusability, and it retained over 94.8% of its residual activity after being used 10 times. Moreover, there was no further loss in activity after these cycles. Fig. 8 shows the FT-IR spectra of the MHHNs-lip before and after the catalyst was used 10 times in the reaction medium. As seen in Fig. 8, no change in the FT-IR spectra of the catalyst was observed after 10 cycles. Moreover, according to elemental analysis, the organic content in the catalyst after 10 cycles was found to be C = 35.5%, Fe = 14.6%, and N = 39.9%, and no considerable change was seen in the CHN data after recycling.

Fig. 6 Magnetic separation of catalyst by an external magnet.

Fig. 7 Reusability of catalyst.

Fig. 8 FT-IR spectra of the MHHNs-lip before and after the catalyst was used 10 times.

The storage stability study of free lipase and MHHNs-lip was investigated and the immobilized lipase, which retains its original catalytic activity, was found to be appreciably stable for a period of 60 days (Fig. 9). The immobilized enzyme provided excellent yield up to 77% of desired product, whereas commercially available lipase provided 69% yield after a period of 60 days.

Fig. 9 The storage stability of the immobilized lipase.

Thermal stability of the immobilized PEL

The thermal stability of the immobilized PEL is one of the most important criteria when dealing with its possible applications. Fig. 10 shows the relative activity of the free and the immobilized PEL incubated in a phosphate buffer at 80 °C for 12 h. As shown in Fig. 10, the relative activity of the immobilized PEL was much higher than that of free PEL. Clearly, the immobilized PEL was more stable than free PEL at relatively high temperatures, indicating a better thermal stability. The increase in the thermal stability of immobilized PEL was caused by its stabilization through the weak intermolecular forces and the prevention of autolysis of the PEL.⁶⁷ Therefore, the thermal stability of the PEL was significantly improved through its immobilization on the MHHNs. In addition, no protein could be observed from the immobilization of PEL (MHHNs-lip) even after a period of 48 h under magnetic stirring, which indicates that the mechanical stability of MHHNs-lip is also high.

Fig. 10 Relative activity of the free and the immobilized PEL incubated in 10 mM phosphate buffer (pH 7.0) at 80 °C for 12 h.

Conclusion

We have developed a novel class of heterogeneous biocatalyst, which is particularly attractive in the environmentally friendly synthesis of 1,3-dibutylurea. PEL was immobilized successfully in the hollow structure of activated MHHNs. The immobilized PEL exhibited high effective activity and thermal stability as well as good reusability. Moreover, the cellulose magnetic microspheres loaded with PEL could be conveniently and easily magnetically separated from reaction solution, leading to recovery of the catalyst. The immobilization carriers prepared from CS and acrylic acid solution in aqueous solvent via a simple, easily prepared, and environmental friendly process will have wide applications in the field of biocatalysis.

Experimental

Chemical reagents

Crude lipase from Penicillium expansum (PEL; 5000 U mg⁻¹ solid), Penicillium neutral expannsum (PNEL; 10 000 U mg⁻¹ solid), Aspergillus niger (ANL; 120 000 U mg⁻¹ solid), Rhizopus chinensis satio (RCSL; 10 000 U mg⁻¹ solid), Aspergillus oryzae (AOL; 30 000 U mg⁻¹ solid), Porcine pancreatic (PPL; 135 U mg⁻¹ solid), neural proteolytic enzyme (50 000 U mg⁻¹ solid), acidic proteolytic enzyme (60 000 U mg⁻¹ solid), alkaline proteolytic enzyme (40 000 U mg⁻¹ solid) were kindly donated by Shenzhen Leveking Bio-engineering Co. Ltd., China. These enzymes were produced by spraying the concentrated supernatant from fermentation with the addition of a certain amount of starch as a thickening agent. Immobilized C. antarctica lipase B (Novozyme 435; EC 3.1.1.3) was donated by Novozymes Investment Co. Ltd (China). Epoxy Sepabeads, EC, butylamine, Fe₃O₄ nanoparticles and all other chemicals were purchased from the Aladdin reagent company, China and were used without further purification. The water used in the study was prepared using a Milli-Q Water Purification System (Millipore, Bedford, MA, USA).

Procedure for the preparation of magnetic hollow Fe₃O₄–polymer hybrid nanospheres

The procedure for the preparation of magnetic hollow Fe₃O₄–polymer hybrid nanospheres was carried out as previously reported.⁴⁵ 10 mL of a PVA-stabilized magnetite nanoparticle suspension (≈1 × 10¹⁸ Fe₃O₄ particles mL⁻¹) was added to 50 mL of an aqueous solution of CS–AA, which consisted of 0.5 g of purified CS (Mw = 200 kDa, degree of deacetylation = 90%) and 0.22 g of AA with a stoichiometric molar ratio of 1 : 1.1 (glucosamine unit–acid). Polymerization was then initiated by K₂S₂O₈ at 70 °C under a stream of nitrogen. As the reaction system appeared opalescent, the reaction was allowed to proceed for another 100 minutes at 50–60 °C. Afterwards, 0.5 mL of GA, a bifunctional cross-linker, was added to the reaction system at 40 °C to crosslink the chitosan selectively. The resultant suspension was separated by an external magnet and washed using acetone. The amino bands of the shells were cross-linked with glutaraldehyde, in a ratio of glucosamine unit in CS to aldehyde unit in GA of 2. The amine moieties were quantified by measuring the color obtained by the reaction of the primary amino groups on the support and an excess of picrylsulphonic acid solution.^84,85 However, in order to immobilize the lipase, we needed to activate the MHHNs with epoxy chloropropane.

Activation of the magnetic hollow Fe₃O₄–polymer hybrid nanospheres

MHHNs (4 g) was added to a solution of 12 mL of epoxy chloropropane in 40 mL of DMSO; then, 40 mL of 2 N NaOH was slowly dripped into the reaction mixture system. The mixture was then allowed to stand for 6 h at room temperature; subsequently, the resulting solution was removed by magnetic separation. The activated MHHNs were obtained after washing with acetone and distilled water until the eluate was neutral.⁶⁷ The epoxy groups in MHHNs were determined by a reaction between the oxirane ring and sodium thiosulfate, followed by titration with 0.01 mol L⁻¹ hydrochloride acid to neutralize any OH⁻ released in the reaction.⁸⁶ The amount of epoxy groups in the activated MHHNs was estimated to be 1.68 mmol g⁻¹ (dry). The epoxide group readily reacts with the enzyme. Under relatively mild conditions, these groups in the cellulose support may react with both amino groups and carboxyl group in the enzyme.

Preparation of the immobilized PEL

The immobilization was carried out as follows. About 1.0 g of the activated magnetic carriers was mixed with 1.0 g of lipase in 25 mL of 10 mM sodium phosphate buffer at pH 7.0 to avoid diffusional problems that could alter the apparent properties of the immobilized enzymes under continuous and gentle stirring. The mixture was shaken at room temperature for 24 h at 250 rpm. Then, the remaining epoxy groups were blocked by incubation with 3 M aspartic acid at pH 8.5 for 24 h.⁸⁷ The supernatant was removed by magnetic separation, and the MHHNs-lip was washed with deionized water, respectively, to remove the noncovalently coupled enzymes on the support. The MHHNs-lip was stored at 4 °C. The total amount of PEL protein was assayed according to the Coomassie Brilliant Blue G-250 method with bovine serum albumin (BSA) used as the standard.⁸⁸

Measurement of enzyme activity

Soluble PEL Activity. The hydrolytic activity of the PEL in an aqueous solution was determined according to the method described in the literature⁸⁹ with a slight modification. The PEL solution (∼0.23 U ml⁻¹, based on the activity in catalyzing the hydrolysis of p-nitrophenyl palmitate (pNPP) at 37 °C in 50 mM phosphate buffer, pH 6.0) was obtained as the supernatant after dissolution of 1.5 g of the enzyme powders in 10.0 ml phosphate buffer (50 mM, pH 8.0) followed by centrifugation. A substrate mixture was prepared by mixing 1 ml of 15.0 mM pNPP in isopropanol and 9 ml of a 50 mM buffer solution (phosphate buffer, pH 5.0–8.0; Tris–HCl buffer, pH 7.0–9.0; barbiturate–HCl buffer, pH 7.5–10.0) containing 0.1% (w/v) arabic gum and 0.4% (w/v) Triton X-100. A cuvette containing 2.4 ml of this substrate mixture was placed in a Pharmacia Biotech Ultraspec 2000UV/V spectrophotometer equipped with a thermostatic cell, for pre-equilibration at 37 °C, and the reaction started by the addition of 0.1 ml of the PEL solution. The variation in the absorbance at 410 nm of the assay against a blank without enzyme was monitored for 2–5 min. The reaction rate was calculated from the slope of absorbance versus the time curve using the molar extinction coefficient for p-nitrophenol (pNP), obtained as measured above.

Immobilized PEL Activity. The enzyme activity of immobilized PEL was measured in the same manner as in the determination of the soluble PEL activity described in the previous section, except that the reaction mixture was continuously stirred during the reaction. The activity yield remaining after immobilization was defined as follow:and the immobilization yield was calculated bywhere A is the total activity of enzyme added in the initial immobilization solution, B is the activity of the residual enzyme in the immobilization and washing solution after the immobilization procedure, and C is the activity of the immobilized PEL. The maximum activity was defined as 100%, and the relative activity refers to the percentage that an enzyme activity accounts for the highest one. The concluded activity yield and immobilization yield were 55.6% and 44.8%, respectively.

Characterization of free and immobilized lipase

Transmission electron microscopy (TEM) was performed using a JEM-2010FEF field-emission electron microscope operating at 200 kV equipped with an EDAX Phoenix EDS analyzer. The thermogravimetric analysis (TGA) was carried out using a Q-series 600 analyzer. About 8–10 mg of samples were placed in a ceramic crucible and the analysis programmed from 30 to 800 °C with a 10 °C min⁻¹ rise in temperature, under a 99.99% pure nitrogen atmosphere with flow rate of 100 mL min⁻¹. The reference run was carried out with an empty sample crucible pan and the results were recorded accordingly. The immobilized biocatalysts and free lipase were investigated for their native confirmation using FT-IR analysis (Bruker. Vertex 70). The magnetic properties of Fe₃O₄, MHHNs and MHHNs-lip were determined using a vibrating sample magnetometer (Lake Shore 7410 VSM). X-ray diffraction of the magnetic microspheres was carried out on the X-ray diffractometer (Rigaku D/MAX-2400 XRD with Ni-filtered Cu Ka radiation). The XRD patterns were recorded in the region of 2θ from 10 to 80 °C. Samples were ground into powders and dried in a vacuum oven at 60 °C for 48 h before characterization.

Procedure for the synthesis of 1,3-dibutylurea from EC and butylamine

The reaction was performed in a 25 mL conical flask. After 40 mmol of EC, 80 mmol of butylamine and the lipase catalyst was charged into the reactor, the mixtures were incubated for 24 h under stirring at temperatures in the range of 60 °C with an agitation speed of 180 rpm. At the end of the reaction, the catalyst was separated by an external magnet and washed using acetone. The recyclable immobilized lipase obtained was dried at 45 °C under reduced pressure overnight (at 1 Torr for 24 h), and it was then used for the next reaction. After separating the catalyst, the reactor was cooled to room temperature and the reaction mixture dispersed in 50 mL of water. Then, the resulting mixture was stirred for 1 h at room temperature. The solid formed was filtered off and washed with water. The product yield was determined from the weight of the solid. The product was characterized by gas chromatography, gas chromatography-mass spectrometry and ¹H NMR spectroscopy and comparison with authentic samples was made whenever possible.

Acknowledgements

We gratefully acknowledge the financial support of the National Natural Science Foundation of China (no. 20972120, 21202037).

Notes and references

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