- Research article
- Open Access
Facile, high efficiency immobilization of lipase enzyme on magnetic iron oxide nanoparticles via a biomimetic coating
© Ren et al; licensee BioMed Central Ltd. 2011
- Received: 20 January 2011
- Accepted: 8 June 2011
- Published: 8 June 2011
Immobilization of lipase on appropriate solid supports is one way to improve their stability and activity, and can be reused for large scale applications. A sample, cost- effective and high loading capacity method is still challenging.
A facile method of lipase immobilization was developed in this study, by the use of polydopamine coated magnetic nanoparticles (PD-MNPs). Under optimal conditions, 73.9% of the available lipase was immobilized on PD-MNPs, yielding a lipase loading capacity as high as 429 mg/g. Enzyme assays revealed that lipase immobilized on PD-MNPs displayed enhanced pH and thermal stability compared to free lipase. Furthermore, lipase immobilized on PD-MNPs was easily isolated from the reaction medium by magnetic separation and retained more than 70% of initial activity after 21 repeated cycles of enzyme reaction followed by magnetic separation.
Immobilization of enzyme onto magnetic iron oxide nanoparticles via poly-dopamine film is economical, facile and efficient.
- Immobilize Lipase
- Free Lipase
- Magnetic Iron Oxide Nanoparticles
Lipases (glycerol ester hydrolases E.C.22.214.171.124) are an important group of enzymes which have been widely used in the catalysis of different reactions [1, 2]. These enzymes have been applied in chemical and pharmaceutical industrial applications due to their catalytic activity in both hydrolytic and synthetic reactions. However, free lipases are easily inactivated and difficult to recover for reuse. Therefore, especially in large-scale applications, lipases are often immobilized on solid supports in order to facilitate recovery and improve operational stability under a wide variety of reaction conditions. Some lipase immobilization strategies involve the conjugation of lipases via covalent attachment, cross-linking, adsorption and entrapment onto hydrophobic or hydrophilic polymeric and inorganic matrixes [3–5].
In recent years, magnetic nanoparticles (MNPs) based on iron oxides, have attracted much interest thanks to their multifunctional properties, such as biocompatibility, superparamagnetism, small size and low toxicity . They have been applied in magnetic resonance imaging (MRI) , biosensors  and as anti-cancer drugs carriers . Due to their high specific surface area and easy separation from the reaction medium by the use of a magnet, they have been employed in enzymatic catalysis applications [10, 11]. Typical strategies for immobilizing lipase onto MNPs rely on surface grafting via low molecular weight linkers or polymers containing amino or epoxy functional groups to which lipases are reacted via covalent conjugation methods [1, 11]. Using such methods, the maximum reported loading capacity of lipase on nanoparticles is approximately 130 mg/g, using a complex methodology . One drawback of existing lipase immobilization technologies is that the activity of lipases decreases significantly upon immobilization due possibly to changes in enzyme secondary structure, or limited access of substrate to the active site of the surface bound enzyme . Thus, despite numerous reported approaches for immobilization of lipases on magnetic nanoparticles, there is still the need for simple, cost-effective and high loading capacity methods.
In this work, we present a facile, biomimetic approach to immobilize lipases onto iron oxide MNP surfaces modified with polydopamine, an in-situ formed coating inspired by the adhesive proteins secreted by marine mussels . The ortho-dihydroxyphenyl (catechol) functional group found in dopamine is also present in mussel adhesive proteins in the form of the amino acid DOPA, where it is highly adhesive to oxide surfaces [14, 15] and under alkaline conditions oxidizes to form quinone, a species that is reactive toward nucleophiles such as primary amines. Dopamine, containing both catechol and primary amine, was previously found to produce conformal coatings on surfaces by self-polymerization , which are further capable of immobilizing biomolecules . In the method described here, polydopamine serves as a conformal coating for the purposes of lipase immobilization onto MNPs. Our results demonstrate that PD-MNPs exhibit high efficiency for lipase immobilization under aqueous conditions, and the enzyme retains high activity after many cycles of magnetic separation and reuse.
Overall Strategy for Preparing Lipase Immobilized Iron Oxide MNPs
Synthesis and characterization of MNPs and PD-MNPs
Catechol derivatized polymers have been previously employed for grafting functional polymers onto surfaces of MNPs [16–18]. In the present work we took advantage of dopamine self-polymerization to form MNPs with polydopamine coatings. In-situ polymerization of dopamine represents a versatile method for modifying solid surfaces with adherent coatings for a variety of functional purposes . The reactions that occur in an alkaline dopamine solution are not unlike those that occur during melanin formation, beginning with oxidation of catechol to yield dopamine quinone, which in turn participates in a series of further intra- and intermolecular reactions that ultimately give rise to a high molecular weight heterogeneous polymer (polydopamine). Polydopamine formation occurs both in solution and as a conformal coating on surfaces. In the present case involving iron oxide MNPs, the polydopamine is likely bound to the surface and even nucleated by dopamine molecules that strongly interact with the oxide surface .
Lipase immobilization onto PD-MNPs
After formation, the polydopamine coating contains residual quinones that are reactive toward nucleophilic groups, affording covalent immobilization of polymers and biomolecules through Michael addition and/or Schiff base formation [13, 15]. In the present case, similar reactions between the active quinone groups at the surface of the PD-MNPs and the amino or thiol groups of the lipase enzyme result in lipase immobilization onto the surfaces of the PD-MNPs during the lipase immobilization step (Figure 1). The reaction between PD-MNPs and lipase was spontaneous and rapid, resulting in precipitation of lipase-containing MNPs particles soon after mixing. Analysis of free protein concentration in solution during lipase immobilization at 4°C showed little change after 2 hours, further confirming the completion of the immobilization reaction.
XPS analysis of the lipase immobilized PD-MNPs (Figure 2C) showed increases in the N1s peak (399.5 eV) and the calculated N/C ratio (0.139) compared to PD-MNPs, reflecting the higher nitrogen content of lipase compared to polydopamine. Little change in aggregate appearance and morphology was observed upon immobilization of lipase onto PD-MNPs, although SEM and TEM analysis (Figure 3C) revealed further agglomeration (average size 150 - 250 nm). It is not known whether the additional aggregation occurred during lipase immobilization or was an artifact of EM sample preparation.
Effect of preparation conditions on lipase immobilization efficiency and activity of lipase immobilized PD-MNP
Bound Lipase (mg)
Specific Activity (U/mg lipase)
3.16 ± 0.52
9.75 ± 2.28
3.09 ± 0.16
6.96 ± 0.49
18.19 ± 2.90
2.62 ± 0.13
7.81 ± 0.62
21.08 ± 2.49
2.70 ± 0.17
8.26 ± 0.48
28.81 ± 3.08
3.49 ± 0.17
9.17 ± 0.58
59.00 ± 3.39
6.44 ± 0.38
9.37 ± 0.55
62.50 ± 3.04
6.67 ± 0.37
9.46 ± 0.57
64.89 ± 2.69
6.86 ± 0.40
4.29 ± 0.22
37.65 ± 2.52
8.78 ± 0.33
118.76 ± 4.13
11.88 ± 0.41
Effect of temperature and pH on free and immobilized lipase activity
Magnetic Isolation and Reuse
In this work, we described a facile method to immobilize lipase onto magnetic nanoparticles through an adhesive polydopamine film. Our immobilization experiments show that the PD-MNPs exhibit high lipase loading capacity due to high surface area and strong adhesive interactions between lipase and polydopamine. In addition, the immobilized lipase shows high specific activity and favorable thermal and pH stability compared to free lipase. Importantly, the lipase immobilized PD-MNPs display good reusability as well as the convenience to be magnetically recovered. These results confirm that immobilization of enzyme onto magnetic iron oxide nanoparticles by poly-dopamine film, is economical, facile and efficient.
Candida rugosa lipase type VII (E.C.126.96.36.199), ferric chloride hexahydrate (FeCl3·6H2O), ferrous chloride tetrahydrate (FeCl2·4H2O), dopamine hydrochloride, tributyrin, gum acacia, tris buffer and sodium hydroxide were purchased from Sigma-Aldrich (USA). Ultrapure water (resistivity = 18.2 MΩ, pH 6.82) was used in all experiments and obtained from a NANOpure Infinity@ system from Barnstead/Thermolyne Corporation (Dubuque, IA).
Synthesis and surface modification of MNPs
Figure 1 presents the synthesis and modification of MNPs. MNPs were prepared by the conventional co-precipitation method  with some modifications. Briefly, 0.5 mmol FeCl2 and 1.0 mmol FeCl3 were dissolved in 50 ml ultrapure water under nitrogen at room temperature, then the pH of the solution was adjusted to 10.0 using 6.0 M NaOH under vigorous stirring. After stirring for 2 hours, the magnetite precipitates were separated and washed several times with ultrapure water by magnetic decantation. The precipitate was dispersed in 50 ml Tris buffer (10 mM, pH 8.5) under ultrasonication for 15 min, after which large precipitates were removed. Dopamine hydrochloride (125 mg, 2.5 mg/ml) was added to the remaining MNPs suspension with vigorous stirring, and the pH of the solution was kept at 8.5 by addition of 10 mM NaOH. After 3 hours, the PD-MNPs was collected by magnetic decantation and washed 5 times with ultrapure water, and finally re-dispersed in ultrapure water to 2 mg PD-MNPs per milliliter solution by ultrasonication for 15 min.
Immobilization of lipase
Lipase immobilization was carried out by adding the suspension of PD-MNPs in ultrapure water to a buffered enzyme solution. An aqueous solution of lipase (2 mg/ml) was prepared by dissolving the lyophilized enzyme in sodium phosphate buffer (10 mM, pH 7.0) solution. A fresh solution of the previously described PD-MNPs (5 ml, 2 mg/ml) was added to the lipase solution (5 ml, 2 mg/ml) at 4°C. Precipitation was observed immediately as the two solutions mixed. After shaking at 180 rpm for 3 h (at 4°C), the lipase-loaded precipitates were collected and washed 3 times with ultrapure water and stored at 4°C prior to use. The protein concentration in the lipase solution before and after immobilization was determined using the Bradford method . The difference in protein concentration was used to calculate the loading of lipase onto the PD-MNPs.
Characterization of MNPs and PD-MNPs
XPS spectra were obtained using an Omicron ESCALAB (Omicron, Taunusstein, Germany) with a monochromatic Al Kα (1486.8 eV) 300-W X-ray source, a flood gun to counter charging effects, and ultrahigh vacuum (~10-9 torr). The takeoff angle was fixed at 45°. Substrates were mounted on sample studs by means of double-sided adhesive tape. All binding energies were calibrated using the C1s peak (284.5 eV). Scanning electron microscopy (SEM) and transmission electron microscopy (TEM) images were acquired on a Hitachi HD2300 electron microscope (Hitachi, Japan) operated at 200 kV. TEM specimens were prepared by casting drops of dilute dispersion of nanoparticles aqueous solution on 200-mesh carbon coated copper grids (Ted Pella).
Activity assay of free and immobilized lipase
The enzymatic activities of free and immobilized lipase were measured by titration of the organic acid that results from the hydrolysis of the tributyrin ester . Briefly, the activity of the free lipase was assayed by adding 0.5 ml of free lipase (5 mg/ml, w/v) in phosphate buffer (10 mM, pH 7.0), 20 ml of a tributyrin solution consisting of 1% tributyrin as the substrate with 1% gum acacia as emulsifier. After 20 min of incubation at 37°C with shaking at 150 rpm, the reaction was stopped by the addition of 20 ml of ethanol. Immediately, the mixture was titrated against 50 mM NaOH in ethanol solution using phenolphthalein as indicator. The immobilized lipase activity was determined as described above by adding a known amount of lipase immobilized PD-MNPs. The lipase loaded PD-MNPs were sonicated for 10 min at 4°C before combining with substrate solution. After incubation of the mixture at 37°C for 20 min with shaking at 200 rpm, the lipase PD-MNPs were separated by a magnet, and the supernatant titrated using an NaOH in ethanol solution. One unit of lipase activity was defined as the amount of lipase which liberated 1 μmol of free acid per minute under the assay conditions. All activity measurements were carried out at least three times and the experimental error was less than 3%.
Effect of temperature and pH
The effect of temperature on the free and immobilized enzyme activity was determined by pre-incubating in phosphate buffer (10 mM, pH 7.0) at temperatures ranging from 20°C to 80°C for 1 h, followed by measurement of the residual enzyme activity at 37°C as described above. The effect of pH on the activity of the free and immobilized enzyme was determined by pre-incubating at room temperature in phosphate buffer (10 mM) at pH ranging from 4 to 9, followed by determination of enzymatic activity at pH 7.0 as described in activity assay section. The residual activity of the immobilized lipase was normalized to the initial value determined at 37°C, pH 7.0 (the initial activity was defined as 100%).
Recovery and reuse of immobilized lipase
The stability of immobilized lipase under conditions of repeated magnetic isolation and reuse was studied under the same conditions as described in activity assay section. After each enzyme run, the lipase containing PD-MNPs were magnetically isolated and washed twice with hexane and ultrapure water to remove any remaining substrate and product species before the next experiment. The residual activity of the immobilized lipase after each cycle was normalized to the initial value (the initial activity was defined as 100%).
This research was partially supported by grant R37 DE 014193 from the National Institutes of Health.
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