High activity and selectivity immobilized lipase on Fe3O4 nanoparticles for banana flavour synthesis

Accepted Manuscript Title: High Activity and Selectivity Immobilized Lipase on Fe3 O4 Nanoparticles for Banana Flavour Synthesis Authors: Maria Sarno, Mariagrazia Iuliano, Massimiliano Polichetti, Paolo Ciambelli PII: DOI: Reference:

S1359-5113(16)31142-4 http://dx.doi.org/doi:10.1016/j.procbio.2017.02.004 PRBI 10935

To appear in:

Process Biochemistry

Received date: Accepted date:

22-12-2016 6-2-2017

Please cite this article as: Sarno Maria, Iuliano Mariagrazia, Polichetti Massimiliano, Ciambelli Paolo.High Activity and Selectivity Immobilized Lipase on Fe3O4 Nanoparticles for Banana Flavour Synthesis.Process Biochemistry http://dx.doi.org/10.1016/j.procbio.2017.02.004 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. 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.

High Activity and Selectivity Immobilized Lipase on Fe3O4 Nanoparticles for Banana Flavour Synthesis

Maria Sarno1,2*, Mariagrazia Iuliano1, Massimiliano Polichetti3, Paolo Ciambelli1,2

1

Department of Industrial Engineering,

2

NANO_MATES Research Centre,

3

Department of Physics “E.R. Caianiello”,

University of Salerno, via Giovanni Paolo II, 132 - 84084 Fisciano (SA), Italy

*

Corresponding authors: Tel.: +39 089 963460; fax: +39 089 964057; E-mail address:

[email protected] (M. Sarno).

Graphical abstract

Highlights  A novel Fe3O4 monodispersed nanoparticles anchoring TL lipase biocatalyst  Simple and green immobilization procedure  Physical adsorption, interfacial activation  Higher activity for the immobilized enzyme over the native counterpart  Less sensitivity to temperature and pH changes, long time stability  Banana flavor production at 100% selectivity

ABSTRACT Lipase (E.C.3.1.1.3) from Thermomyces lanuginosus (TL) was directly bonded, through multiple physical interactions, on citric acid functionalized monodispersed Fe3O4 nanoparticles (NPs) in presence of a small amount of hydrophobic functionalities. A very promising scalable synthetic approach ensuring high control and reproducibility of the results, and an easy and green immobilization procedure was chosen for NPs synthesis and lipase anchoring. The size and structure of magnetic nanoparticles were characterized by transmission electron microscopy (TEM) and X-ray diffraction (XRD). The samples at different degree of functionalization were analysed through thermogravimetric measurements. Lipase immobilization was further confirmed by enzymatic assay and Fourier transform infrared (FT-IR) spectra. Immobilized lipase showed a very high activity recovery up to 144% at pH=7 and 323% at pH=7.5 (activity of the immobilized enzyme compared to that of its free form). The enzyme, anchored to the Fe3O4 nanoparticles, to be easy recovered and reused, resulted more stable than the native counterpart and useful to produce banana flavour. The immobilized lipase results less sensitive to the temperature and pH, with the optimum temperature higher of 5 °C and optimum pH up shifted to 7.5 (free lipase optimum pH=7.0). After 120 days, free and immobilized lipases retained 64 % and 51% of their initial activity, respectively. Ester yield at 40°C for immobilized lipase reached 88% and 100% selectivity.

Keywords: Magnetic catalyst, green immobilization, high activity recovery, banana flavour synthesis

1.Introduction Enzymes are extremely important in the food industry to catalyse a large number of reactions. Lipases are the most widely used class of enzymes in biotechnology [1-10] and this for three main reasons: industrial preparations of many lipases were available in view of their applications in early industrial enzymology; they carry out reactions often in heterogeneous media showing the phenomenon of interfacial activation; they have broad substrate specificities, apart from hydrolysis lipases can also catalyze esterification, interesterification and transesterification [3]. They have various industrial applications: in the food, flavour industry, detergent; biocatalytic resolution of pharmaceuticals, esters and

amino acid derivatives; making of fine chemicals, agrochemicals; use as biosensor, bioremediation cosmetics and perfumery [1]. Free enzymes poor stability towards pH, temperature and time and their cost encourage the use of immobilizations to facilitate separation, recovery and enhance activity [11]. It is very difficult to predict the performance of the immobilized enzyme (e.g. predict effect of the immobilization protocol on one substrate based on the results with other substrate…), basically two central factors must be considered in the development of immobilized biocatalysts: the immobilization methods and the support [12-19]. Immobilization methods can be divided into chemical (covalent bond through ether, thio-ether, amide or carbamate bonds) and physical (characterized by weaker, monocovalent interactions such as hydrogen bonds, hydrophobic interactions, van der Waals forces, affinity binding, ionic binding, or mechanical containment of enzyme within the support) [12-15,20-27]. Covalent bonding is an irreversible enzyme immobilization method, in this case the activity recovery depends on the carrier and coupling materials and methods [12,16,17]. The physical, for which the stability control is more difficult, it can be affected by pH, temperature and ionic strength of the buffer conditions, is a reversible (permitting the support reusability even after enzyme deactivation), low cost and quite simple method, enzymes retain activity through relatively chemical-free enzyme binding [28]. Moreover, covalent and physical immobilization techniques have demonstrated able to successfully provide stable forms of enzymes [29]. Ideal support properties include inertness towards enzymes, biocompatibility, hydrophilicity, resistance to microbial attack and compression and accessibility at a low cost [12,30,31]. Supports can be classified as organic and inorganic according to their chemical composition. Important parameters are pore and/or particle size, establishing the total surface area that affects the capacity for enzymes binding. The immobilization of enzymes onto nanomaterials [11,32-36] is a topic of great interest. Indeed the reduction of

the enzyme support size can provide a larger surface area for the attachment, leading to higher loading. In particular, the large ratio between nanoparticles surface area and volume is not the only key aspect related to particle sizes. Nanoparticles offers lower diffusion limitation in solutions, unlike bulk materials they are mobile in a solution thanks to the Brownian motion, whereby enzyme attached to the nanoparticles are not immobilized [37]. On the other hand, according to the Stokes-Einstein equation the diffusivity of the nanoparticles is lower than that of the free enzyme, due to the larger size, setting these biocatalysts in a transition region between homogeneous and heterogeneous catalysis [37]. It has been found, that the catalyst mobility provides an explanation, among other considerations, for the high activities usually observed for enzymes attached to nanoparticles [38]. Among the inorganic systems to be used as support to immobilize enzymes, no toxic, cheap and biocompatible Fe3O4 magnetic nanoparticles are very promising [11,39-41]. They allow simple and quick collection just applying an external magnetic field avoiding difficult recovery, e.g. via centrifugation or filtration. The lipase from Thermomyces lanuginosus (TL) is a noticeably thermostable enzyme, initially employed in the food industry, the enzyme has found applications in many different industrial areas, from biodiesel production to fine chemicals [42-50]. It is able to catalyse hydrolysis and synthesis of esters and thus play a significant role in pharmaceutical, chemical, energy and food industry [42]. It is worth noticing that the limitations of the industrial use of TL lipase, as well as of other enzymes, have been mainly due to their high cost, which may be overcome by immobilization techniques on solid supports. In general, complex procedures and a large number of chemicals, even toxic, have been used for enzymes immobilization on Fe3O4 [11,51-54]. Moreover, enzymes can show reduced activity compared to the free counterpart after covalent immobilization [11,39,

41,51-54], whereas physical adsorption can preserves its activity [55,56]. Here we report for the first time in food related applications, at the best of our knowledge, a direct bonding of Thermomyces lanuginosus (TL) lipase on citric acid modified Fe3O4 nanoparticles, which have been obtained through a “bottom up” approach [57] offering many advantages, such as results reproducibility and scale up easiness, returning surfactant coated hydrophobic monodispersed NPs. Particular care was devoted to the choice of the nanoparticles synthesis procedure and immobilization conditions. Indeed, our aim was to design a simple and efficient method for high activity enzymes immobilized on nanoparticles prepared by an easy controlled strategy. In particular, in an our previous paper [54] we have observed a reduced activity if compared with the native counterpart for our Fe3O4 nanoparticles activated via glutaraldehyde method to couple with TL lipase by a covalent bond. However, considering the high potential of interfacial activation [55] and taking advantage of the high hydrophobicity of our as prepared surfactant covered NPs we have performed TL activity tests after direct coordination with NPs surface obtaining, also in this case, unsatisfactory results °(e.g. low loading and thus low activity,..). On the other hand, it is well known that TL lipase, which isoelectric point is 4.4, can be easily linked on anionic support at its best activity pH=7 [58,59]. The nanoparticles surface was modified with citric acid (CA), through ligand exchange [60,61] and easily coupled with lipase through physical adsorption. Furthermore, taking advantage of our NPs synthetic approach, during immobilization interfacial activation can also occur [59,62] due to a small amount of residual oleic acid chains, indeed it is known that the ligand exchange process is not able to completely substitute functionality [63] and leading to higher enzymatic activity than that of the native counterpart. The immobilized enzyme was found to be more stable with temperature and showed higher stability towards pH and time. The experimental results evidence the formation of hydrogen bonds during lipase

immobilization suggesting that they not only affect stability [64] but the specificity of TL lipase can be modified according to support interaction [40,65]. In the present work, our magnetic nanoparticles immobilized TL lipase were tested for olive oil hydrolysis and used to synthesize banana flavour. Traditionally, banana flavour has been produced, often in a short supply, by extraction from natural sources. On the other hand, the fermentation is too expensive for commercial exploitation [66]. Transesterification reactions provides an economical alternative route to produce isoamyl acetate, that has a strong banana flavour and a lot of practical applications (e.g. increasingly demanded in food industry but also solvent for paints.. and used in rayon, dyes, penicillin extraction,..) via synthesis between isoamyl alcohol and vinyl acetate [67,68]. Lipases have been employed for isoamyl acetate synthesis in organic solvent [68,69]. However, the use of organic solvents can generate problems of separation, flammability and toxicity. The solvent-free systems have many advantage [70], however the ester was not been obtained in high yield [68]. It is worth noticing that this is the first experimental evidence of a solvent free isoamyl acetate production obtained using immobilized lipase on magnetite nanoparticles. In the banana flower synthesis the immobilized lipase showed very high activity and yield, with 100% selectivity and improved performance than the free counterpart.

2.Experimental 2.1 Preparation of magnetic nanoparticles Fe3O4 magnetic nanoparticles (Fe3O4@OA) synthesis was carried under nitrogen [57,71]. Ethanol and hexane were used as received. Benzyl ether (99%), oleic acid (OA), 1,2hexadecanediol (97%), (90%), iron(III) acetylacetonate (Fe(acac)3), were purchased from Aldrich Chemical Co.

For the synthesis 20 mL of benzyl ether, 2 mmol of Fe(acac)3, 10 mmol of 1,2hexadecanediol, 12 mmol of oleic acid were magnetically stirred from room temperature to 200 °C for 120 min and then to 285 °C for 60 min. Washing was obtained by centrifugation (7500 rpm, 30 min) in ethanol and then in an equal volume mixture of hexane and ethanol. 2.2 Functionalization procedure The magnetic nanoparticles modification with citric acid (CA), through ligand exchange [60,61], was performed at room temperature, a hexane dispersion of hydrophobic Fe3O4 nanoparticles (about 100 mg in 30 mL) was loaded into a 50 mL syringe and added in a vessel previously loaded with a solution 3 M of CA in water (30g in 30 mL). After 5 min a black precipitate was formed at the interface. A slow transfer of the Fe3O4 NPs into the CA solution started and completed in 24 h. Fe3O4 coated with citric acid, Fe3O4@CA in the following, were removed by magnetic field and washed with water several times [70]. It is worth noticing that the NPs CA functionalized can be easily dispersed in water showing a strongly hydrophilic behaviour.

2.3 Lipase Immobilization Then 10 mL of buffer solution (0.1 M phosphate buffer adjusted to pH 3.0, 4.0 and 7.0) containing 2 mg of lipase from Thermomyces lanuginosus (TL) (Sigma Aldrich, solution ≥100,000 U/g) were mixed with the above modified NPs. The mixtures were then shaken 200 rpm at 4 °C for 3 h. After completion of the reaction, the precipitate (Fe3O4@CA_L) was recovered under a magnetic field (the supernatant was removed with a Pasteur pipette, the absence of nanoparticles in the supernatant was then verified under magnetic field) and washed carefully with phosphate buffer for several times, to guarantee the absence of free enzyme, and directly used for the enzyme activity measurements. Furthermore, following the same procedure but using 10-20-40 mL of buffer solution (0.1 M phosphate buffer, pH

7.0) containing 2-4-8 mg of lipase from Thermomyces lanuginosus (TL) (Sigma Aldrich, solution ≥100,000 U/g) we prepare three samples to evaluate, at pH=7, the effect of lipase amount during lipase immobilization. 2.4 Lipase immobilization efficiency For the determination of the lipase protein amount in supernatant Bradford method (Bradford, 1976), using BSA as a protein standard for the calibration curve, was used. The immobilization efficiency of lipase onto the support was evaluated from the following equation ie = (Ci-Cf)V1 ⁄ CiV2, where: ie is the immobilization efficiency (%); Ci and Cf are the initial and final lipase concentrations (mg mL-1) in the supernatant after immobilization, respectively; and V1 and V2 are the solution volume (mL). All data in this formula are averages of duplicate experiments. 2.5 Lipase assay Olive oil emulsion 5.6 mL containing 1% (w/v) Polyvinyl alcohol (PVA) was used to test the enzymatic activities of immobilized and soluble lipase [58]. Soluble (0.2 mg/mL) or immobilized lipase (2 mg of lipase loaded on the nanoparticles) and 7 mL of the phosphate buffer (0.025 M, pH 7.0) were added to the emulsion. The hydrolysis reaction was carried out at 35-45-55-65 °C for 30 min and for 30-60-90-120 min at 40 °C, at pH 4, 5, 6, 7, 7.5, 8 and 8.5 at a temperature of 40°C. Both free and immobilized TL were stored for 120 days at 4 °C and their activity were determined after 30, 60 and 120 days. The amount of fatty acid liberated was measured by titration with 0.1 M KOH solution. The amount of lipase that liberates 1 μmol/min of oleic acid corresponds to one unit (1 U) of activity. As a reference test, titration with 0.1 M KOH solution was also performed for the as prepared nanoparticles after the washing step, no free carboxyl acids were found. The activity recovery (%) after immobilization was the ratio between the activity of immobilized lipase and the activity of its soluble form.

2.6 Reaction setup for transesterification. Sampling and analysis Ester synthesis was carried out in a vessel containing isoamyl alcohol in vinyl acetate molar ratio 1:1 10 mL total volume. Vinyl acetate acted also as an acyl donor. Immobilized lipase was then added at 4 % (enzyme g per mL of reaction mixture). The reaction mixture and a control (solution with no enzyme) were placed in an orbital shaker. The reaction (200 rpm) was carried out at 30 °C for 7 h and for 4-6-8-10 h at 40°C. Effect of cycle number and operational stability on ester formation were also determined. All the experiments were performed in triplicate. Isoamyl acetate was analysed by GC-MS (Thermo Fischer Scientific) by using HP-5 capillary column (0.25 µm×0.25 mm×30 m) at 80 °C for 1 min and then raised to 180 °C for 5 min. Injector and detector temperatures were set 200 °C and 250 °C, respectively. Helium was used as the carrier gas. Isoamyl alcohol and isoamyl acetate retention times were 2.3 and 2.4 min, respectively. Before measurements the samples were separated under a magnetic field to remove the immobilized enzyme. The ester yield was determined as follows: % yield=weight of ester formed (g)/total weight of reaction mixture (g). 2.7 Characterization The characterization was obtained by: Transmission electron microscopy (TEM) (FEI Tecnai electron microscope operating at 200 KV) equipped with an EDX probe, for the measurement the nanoparticles was sonicated in hexane for 5 min; Thermogravimetric analysis (TG-DTG) (SDTQ 600 Analyzer (TA Instruments)) in flowing air at a 10 K/min heating rate; FT-IR spectra (Vertex 70 apparatus (Bruker Corporation)) by applying KBr technique; XRD measurements (Bruker D8 X-ray diffractometer using CuKα radiation). 2.8 Magnetic nanoparticles characterization Quantum Design PPMS 9T equipped with the Vibrating Sample Magnetometer (VSM) was used for the magnetic measurements in the temperature range 5-300 K. DC magnetic fields up to about 4 T maximum have been applied, paying attention to avoid any field

overshoot which could modify the initial samples magnetic state. The samples before each measurement were heated at 300 K for about 20 minutes to delete the effect of an eventual magnetic history. Demagnetisation cycles were performed on the superconducting magnet to reduced its trapped field to about 2 Oe. M vs H loops measurements have been performed at 5K and 300 K. Before measurements the samples were stabilized at the given temperature in zero magnetic field for about 20 minutes. The magnetization M is detected during applying the field up to a maximum value of Hmax and back to - Hmax and then again to Hmax and finally to zero. 3. Results and Discussion 3.1 Graphic representation When a black Fe3O4@OA – hexane solution (30 mL) was added to 30 mL of aqueous CA, the phase was separated showing Fe3O4@OA–hexane solution at the top and aqueous CA solution at the bottom. Fe3O4@OA was then precipitated and suspended in the interface (Figure 1a). At the interface the ligand exchange process took place and then Fe3O4 NPs gradually came to aqueous ligand solution (Figure 1b), which becomes increasingly yellow. This process was completed after 24 h. The pale yellow colour of CA-exchanged NPs solution indicates that the COO- group of CA binds towards the Fe3O4 atoms on the solid surface. The resulting CA-exchanged NPs solution has been maintained stable even after six months without aggregation at room temperature. The schematic representation of the surface modification of hydrophobic Fe3O4@OA, performed using CA as exchanging ligands, was shown in Figure 1c. The as synthesized nanoparticles surface, covered by oleic acid chains, results modified after ligand exchange with citric acid. In particular, citric acid chains together with small amount of residual oleic acid molecules, indeed it is known that the ligand exchange process is not able to completely substitute functionality, cover the surface. Finally, lipase was directly immobilized, without further modification on citric acid functionalized magnetite (Fe3O4@CA) through physical adsorption

(electrostatic interaction occurring between the negatively charged –COO- moieties of citric acid and positively charges on lipase happening in large pH range enclosing the pI of the enzyme [56] and hydrogen bonds between the –NH protein groups and the enriched carboxyl surface of NPs [39,72-74]) occurring together with interfacial activation due to residual surfactant chains [59, 62], see the discussion of the results in the sections below, while it seems difficult that covalent bonds occurs even if in literature it has been reported in similar conditions [39,72]. 3.2 Transmission electron microscopy (TEM) Analysis The morphological and structural characteristics of the NPs were determined by transmission electron microscopy (TEM) analysis. Highly uniform size nanoparticles were formed that, once deposited over a TEM grid, tend to self-organize in a hexagonal layer (Figure 2a, 2b, 2c). From statistical analysis of about 400 nanoparticles, the particle size distribution was obtained, indicating that the average diameter of inorganic core is d = 6.9 nm with σ = 1.7 nm. The corresponding electron diffraction pattern (insert in Figure 2a) confirms the ferrite nature of nanocrystals. Fe3O4@CA keeps its original organization in a monolayer, see Figure 2d. It is worth noticing that in this case, for the TEM sample preparation, the NPs were directly casted from the water solution on the TEM grid. 3.3 Thermogravimetric analysis (TG-DTG) In Figure 3a and 3b the TG-DTG analyses of free and immobilized lipase were reported, respectively. The occurrence of lipase in the sample after immobilization (Fe3O4@CA_L) is highlighted by the endset shift of the weight loss centred at about 230°C for the citric acid modified NPs after a treatment simulating lipase immobilization in the absence of enzyme (Fe3O4@CA_TS), compare Figure 3b and 3d, and by the weight loss in the range 442-619 °C (total weight loss equal to ~20 wt.%). To confirm the efficiency of the anchoring process Fe3O4@CA_L was further stirred in the same conditions of lipase immobilization and thermally analysed by TG. The maroon profile, in Figure 3b, of the

sample so obtained is almost superimposable to the Fe3O4@CA_L, green profile, indicating the efficiency of the anchoring process. Thermogravimetric analysis can help to understand what happened during the immobilization. In particular, the thermal stability of Fe3O4@OA was investigated by thermogravimetric analysis in flowing air (Figure 3c). In the same figure the TG-DTG profiles of oleic acid was also reported for comparison. The Fe3O4@OA TG curve shows: (i) a residual solvent slight weight loss of 7 wt.% at the initial stage; followed by (ii) a combined oxidation-decomposition of bounded oleic chains (20 wt. % total based, 18.6% solid based), occurring in this case mainly at higher temperature than free OA. More insight came from the TG analysis in air flow of Fe3O4@CA, shown in Figure 3d. The thermogravimetric profile of Fe3O4@CA is very similar to that of Fe3O4@OA. It is just after a stirring (sample was shaken 200 rpm at 4 °C for 3 h, simulating the lipase immobilization conditions) that in the TG-DTG profiles of Fe3O4@CA_TS the weight losses due to the lighter CA can be seen, while the loss in the range 254-425 °C (equal to 2.7 wt.% solid based) suggests the presence of residual OA on the NPs surface [63], indicating that during lipase immobilization interfacial activation can occur due to residual oleic acid chains [59,62]. 3.4 FT-IR spectroscopy analysis FT-IR spectra were acquired to analyse the nanoparticles capping with oleic acid and the functionalization at the end of the different steps. Fe3O4@OA spectrum, see Figure 4, showed the typical peaks at 645 and 590 cm-1 due to the 1 (Fe-O) and 2 (Fe-O) bonds [75]. In the spectrum of the coated nanoparticles the C=O stretch band of carboxyl group, at about 1710 cm-1 for pure oleic acid, was absent. In the Fe3O4@OA spectrum two new bands at 1531 and 1649 cm-1 appear due to the asymmetric as(COO-) and the symmetric s(COO-) stretch, indicating the bonding of the carboxylic acids on the surface of the nanoparticles [76]. To confirm the binding of citric acid on the surface of iron oxide nanoparticles, FTIR studies were performed. FTIR spectra of citrate coated iron oxide

(green) and citric acid (blue) are shown in Figure 4. Citric acid shows three bands at 1705, 1740, and 1755 cm-1, due to C=O vibration band of COOH groups present in citric acid [77]. When citric acid adsorbs on the NPs surface C=O bands at 1705 and 1755 cm-1 disappeared and two partially overlapped new bands appeared at about 1660 and 1626 cm-1 [77-79]. It indicates that C-O single bond appears due to a complexation with the Fe3O4 surface [77]. The band at about 1420 cm-1 can be assigned to the asymmetric stretching of CO form COOH group [78]. It is worth noting that the typical bands due to the oleic acid bonding on the surface of the nanoparticles [80] are not clearly visible, on the other hand they could be hidden under the bands between 1720 cm-1 and 1535 cm-1 of Fe3O4@CA_TS. Figure 4 shows the FT-IR spectrum of the lipase-bound nanoparticles, too. After the immobilization of lipase, the sample showed the characteristic bands of both lipase and Fe3O4 (profile in lime). Moreover, the FT-IR profile of Fe3O4@CA_L_stirred is practically superimposable to that of Fe3O4@CA_L in agreement with thermogravimetric analysis. In particular, when the spectra of Fe3O4@CA_stirred and Fe3O4@CA_L were compared the vibrational bands series in the regions over 1000 cm-1 and at about 1700 cm-1 changed due to enzyme-citric acid complex formation [72]. Between 1500 and 1750 cm-1 in the spectrum of the free lipase the typical vibrational bands due to the in-plane N-H bending vibration of amide II at 1657 cm-1 and amide I band due to two carbonyl stretching vibration around 1543 cm-1 [72,73], can be seen. Amide I band is also responsible of enzyme secondary structure. After lipase immobilization the spectrum is dominated by a broad band, due to the CA-NPs vibrational bands and the typical lipase bands also due to the interaction with the nano-support. In particular, the vibrational bands of amide I and amide II are no longer distinguishable, indicating the possibility of hydrogen bonds formation [72,73] and a protein conformational change, probably co-responsible of higher activity observed in the following. 3.5 XRD studies on Fe3O4 NPs

Figure 5 shows the XRD profile of Fe3O4 NPs. The magnetite typical peaks at 30.1° (220), 35.6° (311), 43.1° (400), 53.4° (422), 57.2° (511) and 62.5° (440) can be seen [71]. 3.6 Effect of pH, concentration of lipase and immobilization time on the activity of immobilized lipase The effect of pH on the immobilization efficiency has been evaluated at pH 3 (far before isoelectric point (pI) of TL), at pH 4 (near the TL pI equal to 4.4) and at pH 7 (the optimum activity pH for free lipase [58]). The immobilization efficiency %, as defined in the experimental section, has been found ~60% at pH 3 and about 74 %, error bar ±1.8, at pH 4 and 7, indicating that a larger amount of enzyme can be immobilized, on the hydrophilic NPs functionalized surface, in the range of pH in which the polar interactions are not favoured (indeed the ionic and polar interactions are favoured at pH lower than pI) [39,72-74]. Therefore, hydrogen bonds, which are established between –NH protein groups and the enriched carboxyl surface of nanoparticles, prevail in the interactions between our functionalized NPs and TL. As can be seen in Table 1 the activity recovery % (the activity recovery (%) after immobilization was the ratio between the activity of immobilized lipase and the activity of its soluble form), measured at pH 7 results higher for Fe3O4@CA_L as high as the immobilization pH, this can be due to the abrupt change in pH and conformational modifications of lipase structure induced by pH. As it is expected, bearing in mind the free lipase behaviour, the recovery activity at smaller pH, see Table 1, results lower. The best lipase amount for immobilization was analysed at pH 7 on 100 mg of Fe3O4 nanoparticles. The dependence of lipase loading with the immobilization efficiency and the activity recovery (olive oil emulsion was used as substrate for incubation in a phosphate buffer (pH 7)) is shown in Figure 6. From this figure, it can be seen that the immobilization efficiency and the activity recovery were decreased when the lipase loading rose from 2 to 8 mg, probably due to diffusion limitations caused by the high activity of the catalyst

(concentration and pH gradients in the biocatalyst microenvironment) [81,82] A lipase loading of 2 mg was thus chosen for the lipase immobilization under the assay conditions. The immobilization efficiency and activity recovery of immobilized lipase was also studied at two immobilizing time (Figure 7). It was found that, under coupling time increase from 1 to 3 h, the percentage of immobilized lipase increased, due to residual active carboxyl groups still available on the surface of NPs for immobilization. After 3 hours immobilization the activity recovery results increased, likely due to improved conformation flexibility to interact with the substrate molecules [83]. From Figure 6 and 7 it is also evident that the activity recovery of immobilized lipase results higher (until 144% at pH=7 and 40 °C) than that of the native counterpart suggesting that the specificity of TL lipase was modified according to the support interaction [40,65]. This can be attributed to: (i) modifications of lipase conformational structure; (ii) enzyme stabilization after the immobilization on high surface area NPs and confinement, resulting in higher availability of the active sites for the reaction media [65]; and (iii) even more interfacial activation enhancing enzyme activity [59,62]. 3.7 Effect of pH and temperature on the activity of immobilized lipase The relative activity for different temperatures between 35 and 65 °C was determined, and the results given in Figure 8. The optimum temperature for free lipase was 40 °C, while immobilized lipase showed the highest hydrolytic activity at 45 °C. Moreover, in the temperature range 45-65 °C the lipase immobilized on Fe3O4 results less sensitive to the temperature change, probably due to a support inducing stability. The hydrolytic activity of both free and immobilized lipase was tested at different pH values between 4 and 8 to find the optimal conditions, see Figure 9. It can be observed an higher pH tolerability for immobilized lipase (pH range 6-8.5), the optimum pH for immobilized lipase results up shifted to 7.5 (free lipase optimum pH=7.0) [58]. These results are probably due to multipoint non-covalent interaction of lipase and support which

preserve the activity [39]. It is worth noticing that at pH=7.5 the recovery activity of the immobilized lipase was 323% of that showed by its native counterpart. 3.8 Thermal and storage stability of immobilized lipase Time-dependent thermal stability of free and immobilized lipase was determined at 40 °C as shown in Figure 10. The immobilized enzyme was found to be more thermostable than the free enzyme, it retained 88% of its activity after 2 h (free enzyme retained 74% of its activity). Moreover, the activity of immobilized lipase keeps constant after 60 min. The effect of operational and storage stability on the activity of free ad immobilized TL was investigated as shown in Figure11. Both free and immobilized TL were stored for 120 days at 4 °C and their activity were determined after 30, 60 and 120 days using the standard activity assay. The immobilized TL retains 80% of its initial activity after 30 days of storage. Under the same conditions, free enzyme lost 40% of its initial activity during 30 days of storage, as enzymes are not stable in solution and their activity is gradually reduced [50,79]. After 120 days, free and immobilized lipases retained 51% and 64 % of their initial activity, respectively, indicating that the electrostatic interaction and even more the hydrogen bonds are a good way to stabilize the enzyme activity, also if compared with previous results obtained for TL cross-linked with glutaraldehyde [50]. 3.9 Effect of immobilized lipase on transesterification reaction Ester yields obtained at 30 °C after 7 h of reaction were 46 % and 61% for free and immobilized lipase respectively. The ester yields for free and immobilized lipases were also determined at 40 °C and different times. As expected enzyme activity results higher at 40 °C, furthermore it can be observed (Figure 12), in particular for 6 h and 8 h of reaction, a significantly higher activity vs ester formation for immobilized lipase, in agreement with the results of the lipase assay tests. The ester yield for immobilized TL increases under time until 8 h as for free lipase. It is worth noticing that the selectivity for immobilized

lipase was 100%, whereas acetaldehyde formation was also observed by using free lipase catalyst. To explore the very key aspect of the enzyme recycling and reuse, the reusability of the immobilized enzyme for the ester formation was investigated. Activity of the immobilized lipase was measured seven times over a period of 2 weeks as shown in Figure 13. Before each cycle the immobilized lipase was washed with sodium phosphate buffer solution (0.1 M, pH 7), after reactions easily separated form product by magnet and finally stored at 4 °C in the same buffer for further use. The immobilized lipase activity was maintained almost intact after 3 cycles, it remains above 75% at the 5° cycle and at 44% of its initial activity after seven successive reuse, showing an excellent reusability even higher than that shown by TL anchored on Fe3O4 by covalent bonds [58]. The effect of operational and storage stability on ester formation for free ad immobilized TL was investigated. Both free and immobilized TL were stored for 60 days at 4 °C and their activity were determined after 30, 45 and 60 days (see Figure 14). The immobilized TL retains 75% of its initial activity at the third working cycle and after 60 days of storage, showing also in the production of aroma banana performance better than those exhibited by the free lipase. 3.10

Magnetic characterization

Figure 15 shows the Fe3O4 magnetic hysteresis loops (M vs H) measured at 300 K. The inserts in the figures show, in a smaller range to highlight the hysteresis, magnetization loops at 300 K and 5 K for the three NPs. The Hc values at the two temperatures agree with the particles size distribution measured for our NPs [85]. In particular, the particles are close to the superparamagnetic regime [86]. The saturation magnetization (Ms) of oleic acid, citric acid and citric acid_Lipase coated Fe3O4 was found to be 70.29, 71.72 and 68.64 emu/g, respectively, likely due to the different organic content. The three samples

Ms values were lower than that shown by bulk magnetite (Ms= 89.0 emu/g), mainly due to the size and surface effect [61]. 4. Conclusion Highly uniform size NPs that, once deposited over a TEM grid, due to oleic acid chains tend, to self-organize in a hexagonal layer, with an average diameter of 6.9 nm and 1.7 nm standard deviation, were prepare by a promising synthetic approach. A new magnetic lipase catalyst, utilizing Fe3O4 monodispersed nanoparticles as support, was developed. The enzyme was immobilized through a simple and green procedure. The very high enzymatic activity and stability showed by our bio-catalyst, obtained through a one-step immobilization process, was likely due to the multiple interaction consisting in electrostatic forces, hydrogen bonds and interfacial activation. The immobilized TL retains 80% of its initial activity during 30 days of storage, while free lipase in the same time, due to a gradual activity reduction in no-stable solution lost 40% of its initial activity. The immobilized TL retains 75% of its initial activity after 60 days of storage at the third working cycle of aroma banana production, showing better performance than those exhibited by the free lipase. After 120 days immobilized lipases retained 64 % of its initial activity indicating that the anchoring of lipase on our functionalized nanoparticles is a good way to stabilize the enzyme activity, also if compared with previous results obtained for chemical immobilized TL. The saturation magnetization (Ms) of oleic acid, citric acid and citric acid_Lipase coated Fe3O4 was found to be 70.29, 71.72 and 68.64 emu/g, respectively and therefore substantially unmodified by the coating. References.

[1] Hasan F, Shah AA, Hameed A. Industrial applications of microbial lipases. Enzyme Microb Tech 2006;27:235-251.

[2] Jaeger KE, Eggert T. Lipases for biotechnology. Curr Opin Biotech 2002;13:390397. [3] Kapoor M, Gupta MN. Lipase promiscuity and its biochemical applications. Process Biochem 2012;47:555-569. [4] Jayani RS, Saxena S, Gupta R. Microbial pectinolytic enzymes: A review. Process Biochem 2005;40:2931–2944. [5] Alkorta I, Garbisu C, Llama MJ, Serra JL. Industrial applications of pectic enzymes: a review. Process Biochem 1998;33:21-28. [6] Rodriguesa RC, Lafuenteb RF. Lipase from Rhizomucor miehei as an industrial biocatalyst in chemical process. J Mol Catal B-Enzym 2010;64:1–22. [7] Sharmaa R, Chistib Y, Banerjeea UC. Production, purification, characterization, and applications of lipases. Biotechnol Adv 2001;19:627–662. [8] Jaeger KE, Reetz MT. Microbial lipases form versatile tools for Biotechnology. Trends Biotechnol 1998;16:396-403. [9] Pandey A, Benjamin S, Soccol CR, Nigam P, Krieger N, Soccol VT. The realm of microbial lipases in biotechnology. Biotechnol Appl Bioc 1999;29:119–131. [10] Sheldon RA, Van Pelt S. Enzyme immobilisation in biocatalysis: Why, what and how. Chem Soc Rev 2013;42:6223-6235. [11] Cao M, Li Z, Wang J, Ge W, Yue T, Li R, Colvin VL, Yu WW. Food related applications of magnetic iron oxide nanoparticles: Enzyme immobilization, protein purification, and food analysis. Trends Food Sci Tech 2012;27:47-56. [12] Mohamada NR, Marzukia NHC, Buanga NA, Huyopb F, Wahab RA. An overview of technologies for immobilization of enzymes and surface analysis techniques for immobilized enzymes. Biotechnol Biotechnol Equip 2015;29:205-220. [13] Ruedaa N, Dos Santos JCS, Ortiz C, Barbosa O, Lafuente RF, Torres R. Chemical amination of lipases improves their immobilization on octyl-glyoxyl agarose beads. Catal Today 2015;259:107-118. [14] Suescun A, Rueda N, Dos Santos JCS, Castillo JJ, Ortiz C, Torres R, Barbosa O, Lafuente RF. Immobilization of lipases on glyoxyl–octyl supports:Improved stability and reactivation strategies. Process Biochem 2015;50:1211-1217. [15] L de Albuquerque T, Rueda N, Dos Santos JCS, Barbosa O, Ortiz C, Binay B, Özdemir E, Gonçalves LRB, Lafuente RF. Easy stabilization of interfacially activated lipases usingheterofunctional divinyl sulfone activated-octyl agarose beads. Modulation

of the immobilized enzymes by altering their nano environment. Process Biochem 2016;51:865–874. [16] Dos Santos JCS, Rueda N, Torres R, Barbosa O, Gonçalves LRB, Lafuente RF. Evaluation of divinylsulfone activated agarose to immobilize lipases and to tune their catalytic properties. Process Biochem 2015;50:918–927. [17] Garcia-Galan C, Dos Santos JCS, Barbosa O, Torres R, Pereira EB, Corberan VC, Gonçalves LRB, Lafuente RF. Tuning of Lecitase features via solid-phase chemical modification: Effect of the immobilization protocol. Process Biochem 2014;49:604– 616. [18] De Souza TC, De Fonseca TS, Da Costa JA, Rocha MVP, De Mattos MC, Lafuente RF, Gonçalves LRB, Dos Santos JCS. Cashew apple bagasse as a support for the immobilization of lipase B from Candida antarctica: Application to the chemoenzymatic production of (R)-Indanol. J Mol Catal B-Enzym 2016;130:58-69. [19] Rueda N, Albuquerque TL, Cabrero RB, Lopez LF, Torres R, Ortiz C, Dos Santos JCS, Barbosa O, Lafuente RF. Reversible Immobilization of Lipases on Heterofunctional Octyl-Amino Agarose Beads Prevents Enzyme Desorption. Molecules 2016;21:646. [20] Manoel EA, Dos Santos JCS, Freire DMG, Rueda N, Lafuente RF. Immobilization of lipases on hydrophobic supports involves the open form of the enzyme. Enzyme Microb Technol 2015;71:53-57. [21] Schmid RD, Verger R. Lipases: Interfacial Enzymes with Attractive Applications. Angew. Chem. Int. Ed. 1998;37:1608-1633. [22] Verger R. ‘Interfacial activation’ of lipases: facts and artifacts. Trends Biotechnol 1997;15:32-38. [23] Cambillau C, Longhi S, Nicolas A, Martinez C. Acyl glycerol hydrolases: inhibitors, interface and catalysis. Curr Opin Struct Biol 1996;6:449-455. [24] Brzozowski AM, Derewenda U, Derewenda ZS, Dodson GG, Lawson DM, Turkenburg JP, Bjorkling F, Huge-Jensen B, Patkar SA, Thim L. A model for interfacial activation in lipases from the structure of a fungal lipase-inhibitor complex. Nature 1991;351:491-494. [25] Kim KK, Song HK, Shin DH, Hwang KY, Suh SW. The crystal structure of a triacylglycerol lipase from Pseudomonas cepacia reveals a highly open conformation in the absence of a bound inhibitor. Structure 1997;5:173-185.

[26] Jaeger KE, Ransac S, Koch HB, Ferrato F, Dijkstra BW. Topological characterization and modeling of the 3D structure of lipase from Pseudomonas aeruginosa. FEBS Lett 1993;332:143-149. [27] Cygler M, Schrag JD. Structure and conformational flexibility of Candida rugosa lipase. Biochim Biophys Acta 1999;1441:205-14. [28] Hanefeld U, Gardossi L, Magner E. Understanding enzyme immobilisation. Chem Soc Rev 2009;38:453-468. [29] Murty VR, Bhat J, Muniswaran PKA. Hydrolysis of Oils by Using Immobilized Lipase Enzyme: A Review. Biotechnol Bioprocess Eng 2002;7:57-66. [30] Rodrigues RC, Hernandez K, Barbosa O, Rueda N, Galan CG, Dos Santos JCS, Murcia AB, Lafuente RF. Immobilization of Proteins in Poly-Styrene-Divinylbenzene Matrices: Functional Properties and Applications. Curr Org Chem 2015;19:1707-1718. [31] Evelin A. Manoel, Marcela F.P. Ribeiro, Jose C.S. Dos Santos, Maria Alice Z. Coelho, Alessandro B.C. Simas, Roberto Fernandez-Lafuente, Denise M.G. Freire. Accurel MP 1000 as a support for the immobilization of lipase from Burkholderia cepacia: Application to the kinetic resolution of myo-inositol derivatives. Process Biochem 2015;50:1557-1564. [32] Kim BC, Nair S, Kim J, Kwak JH, Grate JW, Kim SH, Gu MB. Preparation of biocatalytic nanofibers with high activity and stability via enzyme aggregate coating on polymer nanofibers. Nanotechnology 2005;16:S382-S388. [33] Jia H, Zhu G, Vugrinovich B, Kataphinan W, Reneker DH, Wang P. EnzymeCarrying polymeric nanofibers prepared via electrospinning for use as unique biocatalysts. Biotechnol Progr 2002;18:1027-1032. [34] Cao Y, Wen L, Svec F, Tan T, Lv Y. Magnetic AuNP@Fe3O4 nanoparticles as reusable carriers for reversible enzyme immobilization. Chem Eng J 2016;286:272-281. [35] Wang J, Meng G, Tao K, Feng M, Zhao X, Li Z, Xu H, Xia D, Lu JR. Immobilization of lipases on alkyl silane modified magnetic nanoparticles: effect of alkyl chain length on enzyme activity. PLoS ONE 2012;7:e43478. [36] Dos Santos JCS, Barbosa O, Ortiz C, Murcia AB, Rodrigues RC, Lafuente RF. Importance of the Support Properties for Immobilization or Purification of Enzymes. Chem Cat Chem 2015;7:2413-2432. [37] Wang P. Nanoscale Biocatalyst Systems. Current Opinion in Biotechnology 2006;17:574–579.

[38] Jia H, Zhu G, Wang P. Catalytic behaviors of enzymes attached to nanoparticles: the effect of particle mobility. Biotechnol Bioeng 2003;84:406-14. [39] Sahoo B, Sahu SK, Bhattacharya D, Dhara D, Pramanika P. A novel approach for efficient immobilization and stabilization of papain on magnetic gold nanocomposites. Colloid Surface B 2013;101:280-289. [40] Jiang Y, Guo C, Xia H, Mahmood I, Liu C, Liu H. Magnetic nanoparticles supported ionic liquids for lipase immobilization: Enzyme activity in catalyzing esterification. J Mol Catal B-Enzym 2009; 58:103-109. [41] Hosseinipour SL, Khiabani MS, Hamishehkar H, Salehi R. Enhanced stability and catalytic activity of immobilized a-amylase on modified Fe3O4 nanoparticles for potential application in food industries. J Nanopart. Res 2015;17:382. [42] Lafuente RF. Lipase from Thermomyces lanuginosus: Uses and prospects as an industrial biocatalyst. J Mol Catal B-Enzym 2010; 62:197-212. [43] Ferrer M, Soliveri J, Plou FJ, Cortes NL, Duarte DR, Christensen M, Patino JLC, Ballesteros A. Synthesis of sugar esters in solvent mixtures by lipases from Thermomyces lanuginosus and Candida antarctica B, and their antimicrobial properties. Enzyme Microb Tech 2005;36:391-398. [44] Mogensen JE, Sehgal P, Otzen DE. Activation, Inhibition, and Destabilization of Thermomyces lanuginosus Lipase by Detergents. Biochemistry 2005;44:1719-1730. [45] Lage FAP, Bassi JJ, Corradini MCC, Todero LM, Luiz JHH, Mendes AA. Preparation of a biocatalyst via physical adsorption of lipase from Thermomyces lanuginosus on hydrophobic support to catalyze biolubricant synthesis by esterification reaction in a solvent-free system. Enzyme Microb Tech 2016;84:56-67. [46] Paludo N, Alves JS, Altmann C, Ayub MAZ, Lafuente RF, Rodrigues RC. The combined use of ultrasound and molecular sieves improves the synthesis of ethyl butyrate catalyzed by immobilized Thermomyces lanuginosus lipase. Ultrason Sonochem 2015;22:89-94. [47] Martins AB, Da Silva AM, Schein MF, Galan CG, Ayub MAZ, Lafuente RF, Rodrigues RC. Comparison of the performance of commercial immobilized lipases in the synthesis of different flavor esters. J Mol Catal B-Enzym 2014;105:18-25. [48] Martins AB, Friedrich JLR, Rodrigues RC. Optimized butyl butyrate synthesis catalyzed by Thermomyces lanuginosus lipase. Biotechnol Progr 2013;29:1416-1421. [49] Martins AB, Friedrich JLR, Cavalheiro JC, Galan CG, Barbosa O, Ayub MAZ, Lafuente RF, Rodrigues RC. Improved production of butyl butyrate with lipase from

Thermomyces lanuginosus immobilized on styrene-divinylbenzene beads. Bioresour Technol 2013;134:417-422. [50] Ondul E, Dizge N, Albayrak N. Immobilization of Candida antarctica A and Thermomyces lanuginosus lipases on cotton terry cloth fibrils using polyethyleneimine. Colloids Surf B 2012;95:109-114. [51] Hu B, Pan J, Yu HL, Liu JW, Xu JH. Immobilization of Serratia marcescens lipase onto amino-functionalized magnetic nanoparticles for repeated use in enzymatic synthesis of Diltiazem intermediate. Process Biochem 2009;44:1019-1024. [52] Xie W, Ma N. Enzymatic transesterification of soybean oil by using immobilized lipase on magnetic nano-particles. Biomass Bioenerg 2010;34:890-896. [53] Jordan J, Kumar CSSR, Theegala C. Preparation and characterization of cellulasebound magnetite nanoparticles. J Mol Catal B-Enzym 2011;68:139-146. [54] Sarno M, Paciello L, Cirillo C, Parascandola P , Ciambelli P. Improvement of the Lipase Immobilization Procedure on Monodispersed Fe3O4 Magnetic Nanoparticles. Chem Eng Trans 2016;49:121-124. [55] Lee DG, Ponvel KM, Kim M, Hwang S, Ahn IS, Lee CH. Immobilization of lipase on hydrophobic nano-sized magnetite particles. J Mol Catal B-Enzym 2009;57:62-66. [56] Bahrami A, Hejazi P. Electrostatic immobilization of pectinase on negatively charged AOT-Fe3O4 nanoparticles. J Mol Catal B-Enzym 2013;93:1-7. [57] Altavilla C, Sarno M, Ciambelli P. Synthesis of Ordered Layers of Monodisperse CoFe2O4 Nanoparticles for Catalyzed Growth of Carbon Nanotubes on Silicon Substrate. Chem Mater 2009;21:4851–4858. [58] Xie W, Ma N. Immobilized lipase on Fe3O4 nanoparticles as biocatalyst for biodiesel production. Energ Fuels 2009;23:1347-1353. [59] Quilles JCJ, Brito RR, Borges JP, Aragon CC,Lorente GF, Bocchini-Martins DA, Gomes E, Da Silva R, Boscolo M, Guisan JM. Modulation of the activity and selectivity of the immobilized lipases by surfactants and solvents. Biochem Eng J 2015;93:274– 280. [60] Krishna SH, Divakar S, Prapulla SG, Karanth NG. Enzymatic synthesis of isoamyl acetate using immobilized lipase from Rhizomucormiehei. J Biotechnol 2001;87:193201. [61] Kumari A, Mahapatra P, Garlapati VK, Banerjee R, Dasgupta S. Lipase Mediated Isoamyl Acetate Synthesis in Solvent-Free System Using Vinyl Acetate as Acyl Donor. Food Technol Biotechnol 2009;47:13–18.

[62] Cui J, Zhao Y, Liu R, Zhong C, Jia S. Surfactant-activated lipase hybridnanoflowers with enhanced enzymatic performance. Sci Rep 2016;6:27928. [63] Herranz F, Salinas B, Groult H, Pellico J, Lechuga-Vieco AV, Bhavesh R, Cabello JR. Superparamagnetic Nanoparticles for Atherosclerosis Imaging. Nanomaterials 2014;4:408-438 [64] Klapiszewski Ł, Wysokowski M, Norman M, Kołodziejczak-Radzimska A, Moszyński D, Ehrlich H, Maciejewski H, Stelling AL, Jesionowski T. Chitin-Lignin Material as a Novel Matrix for Enzyme Immobilization. Mar. Drugs 2015;13:24242446. [65] Santos JC, Bueno T, Molgero PC, Rós D, Castro HFD. Lipase-catalyzed synthesis of butyl esters by direct esterification in solvent-free system. J Chem Technol Biot 2007;82:956-961. [66] Romero MD, Calvoa L, Alba C, Daneshfarb A, Ghaziaskarb HS. Enzymatic synthesis of isoamyl acetate with immobilized Candida antarctica lipase in n-hexane. Enzyme Microb Tech 2005;37:42-48. [67] Krishna SH, Divakar S, Prapulla SG, Karanth NG. Enzymatic synthesis of isoamyl acetate using immobilized lipase from Rhizomucormiehei. J Biotechnol 2001;87:193201. [68] Kumari A, Mahapatra P, Garlapati VK, Banerjee R, Dasgupta S. Lipase Mediated Isoamyl Acetate Synthesis in Solvent-Free System Using Vinyl Acetate as Acyl Donor. Food Technol Biotechnol 2009;47:13–18. [69] Sarno M, Cirillo C, Ponticorvo E, Ciambelli P. Synthesis and Characterization of FLG/Fe3O4 Nanohybrid Supercapacitor. Chem Eng Trans 2015;43:727-732. [70] Vo DQ, Kim EJ, Kim S. Surface modification of hydrophobic nanocrystals using short-chain carboxylic acids. J Colloid Interf Sci 2009;337:75-80. [71] Sun S, Zeng H, Robinson DB, Raoux S, Rice PM, Wang SX, Li G. Monodisperse MFe2O4(M = Fe, Co, Mn) Nanoparticles. J Am Chem Soc 2004;126:273-279. [72] Atacan K, Özacar M. Characterization and immobilization of trypsin on tannic acid modified Fe3O4 nanoparticles. Colloids Surf B 2015;128:227-236. [73] Atacan K, Çakıroğlu B, Özacar M. Improvement of the stability and activity of immobilized trypsin on modified Fe3O4 magnetic nanoparticles for hydrolysis of bovine serum albumin and its application in the bovine milk. Food Chem 2016;212:460468.

[74] Agostinelli E, Belli F, Tempera G, Mura A, Floris G, Toniolo L, Vavasori A, Fabris S, Momo F, Stevanato R. Polyketone polymer: a new support for direct enzyme immobilization. J Biotechnol 2007;127:670-678. [75] Klokkenburg M, Hilhorst J, Erne B.H. Surface analysis of magnetite nanoparticles in cyclohexane solutions of oleic acid and oleylamine. Vib Spectrosc 2007;43:243-248. [76] Zhang L, He R, Gu HC. Oleic acid coating on the monodisperse magnetite nanoparticles. Appl Surf Sci 2006;253:2611-2617. [77] Srivastava S, Awasthi R, Gajbhiye NS, Agarwal V, Singh A, Yadav A, Gupta RK. Innovative synthesis of citrate-coated superparamagnetic Fe3O4 nanoparticles and its preliminary applications. J Colloid Interf Sci 2011;359:104–111. [78] Singh D, Gautam RK, Kumar R, Shukla BK, Shankar V, Krishna V. Citric acid coated magnetic nanoparticles:Synthesis,characterization and application in removal of Cd(II) ions from aqueous solution. J Water Process Eng 2014;4:233-241. [79] Nigam S, Barick KC, Bahadur D. Development of citrate-stabilizer Fe3O4 nanoparticles: conjugation and release of doxorubicin for therapeutic applications. J Magn Magn Mater 2011;323:237-243. [80] Sarno M, Ponticorvo E, Cirillo C. High surface area monodispersed Fe3O4 nanoparticles alone and on physical exfoliated graphite for improved supercapacitors J Phys Chem Solids 2016;99:138–147. [81] Hernandez K, Galan CG, Lafuente RF. Simple and efficient immobilization of lipase B from Candida antarctica on porous styrene-divinylbenzene beads. Enzyme Microb Technol 2011;49:72-78. [82] Lage FAP, Bassi JJ, Corradini MCC, Todero LM, Luiz JHH, Mendes AA. Preparation of a biocatalyst via physical adsorption of lipase from Thermomyces lanuginosus on hydrophobic support to catalyze biolubricant synthesis by esterification reaction in a solvent-free system. Enzyme Microb Technol 2016;84:56-67 [83] Huang SH, Liao MH, Chen DH. Direct Binding and Characterization of Lipase onto Magnetic Nanoparticles. Biotechnol Prog 2003;19:1095-1100. [84] Arıca MY, Bayramoglu G. Reversible immobilization of tyrosinase ontopolyethyleneimine-grafted and Cu(II) chelated poly(HEMA-co-GMA) reactivemembranes, J Mol Catal B Enzym 2004;27:255–265. [85] Chesne K, Trevino M, Cai Y, Hancock JM, Smith SJ, Harrison RG. Particle size effects on the magnetic behavior of 5 to 11 nm Fe3O4 nanoparticles coated with oleic acid. JPCS 2014;521:012004.

[86] Brollo MEF, Ruiz RL, Muraca D, Figueroa SJA, Pirota KR, Knobel M. Compact Ag@Fe3O4 Core-shell Nanoparticles by Means of Single-step Thermal Decomposition Reaction. Sci Rep 2014;4:6839.

a

b

c

H

In the case of chemical bond

NH2

O HO

O

OH O OH

OH

Oleic Acid Citric Acid Lipase from Thermomyces Lanuginousus in closed form; Lipase from Thermomyces Lanuginousus; in open form Figure 1 Photographs of Fe3O4@OA NPs suspended at the CA aqueous solution interface (a) and precipitated in aqueous CA under magnetic field (b). Schematic representation of the of ligand exchange process of Fe3O4@OA with CA and Lipase immobilization on the surface of Fe3O4@CA (c).

b

a

d

100 nm

c

200 nm

200 nm

100 nm

Figure 2 TEM images of Fe3O4@OA nanoparticles at different magnifications (a, b, c), TEM of Fe3O4@CA (d)

100

L

0.5

Deriv. Weight (%/min)

Weight (%)

80 60

a

40 20 0 60

260 460 Temperature (°C)

100

660

––––––– Fe3O4@CA_L ––––––– Fe3O4@CA_L_stirred

90

0.0

Universal V4.5A TA Instruments

3

Deriv. Weight (%/min)

80 Weight (%)

70 60 50

b

40 30 20 10 0 60

260 460 Temperature (°C)

120

660

-1

Universal V4.5A TA Instruments

100

Deriv. Weight (%/min)

––––––– Fe3O4@OA ––––––– OA 35

Weight (%)

80 60

c

40

15

20 230

120

430

Temperature (°C) ––––––– ––––––– –––––––

100

630

-5 Universal V4.5A TA Instruments

FE3O4@CA 19 CA Fe3O4@CA_TS

Weight (%)

80 60

9

d

40

Deriv. Weight (%/min)

0 30

20 0 30

230

430 Temperature (°C)

630

-1 Universal V4.5A TA Instruments

Figure 3. TG-DTG analyses of: lipase solution (Sigma Aldrich) (a); Fe3O4@CA_L and Fe3O4@CA_L after stirring (b); Fe3O4@OA and OA (c) and Fe3O4@CA and CA (d).

Figure 4 FT-IR spectra in the range of wavenumber 2000-400 cm-1 of nanoparticles synthesized in the presence of surfactants (Fe3O4@OA), nanoparticles after the ligand exchange (Fe3O4@CA_TS), nanoparticles after lipase immobilization (Fe3O4@CA_L and Fe3O4@CA_L_stirred), free citric acid, oleic acid and lipase.

311

0

220 400

Intensity a.u.

4@OA

440 511

422

F3O4@OA

80

100

20

40

60

2 ° Figure 5 XRD spectra of F3O4@OA.

80

120

80

80

40

40

Activity Recovery (%)

Immobilization Efficiency (%)

120

Activity Recovery (%) Immobilization Efficiency (%)

0 2,5

5,0

7,5

0 10,0

Amounts of lipase (mg)

Figure 6. Effect of lipase amounts on the immobilization efficiency and activity recovery. Immobilization conditions: coupling temperature, 4 °C; coupling time, 3 h. Reaction conditions: reaction temperature, 40 °C; reaction time, 30 min.

120

80

80

40

40

Activity Recovery (%)

Immobilization Efficiency (%)

120

Activity Recovery (%) Immobilization Efficiency (%)

0

0 60

80

100

120

140

160

180

time (min)

Figure 7. Effect of coupling time on the immobilization efficiency and activity recovery. Immobilization conditions: coupling temperature, 4 °C; lipase amount, 2 mg. Reaction conditions: reaction temperature, 40 °C; reaction time, 30 min.

Free Lipase Immobilized Lipase

Relative activity (%)

100

80

60

40

20 35

40

45

50

55

60

65

Temperature (°C)

Figure 8. Effect of temperature on hydrolytic activities of the free and immobilized lipases. Immobilization conditions: coupling temperature, 4 °C; coupling time: 3h; lipase amount, 2 mg. Reaction conditions: reaction time, 30 min. The maxima were defined as 100% activity.

Relative activity (%)

100

Free Lipase Immobilized Lipase

80 60 40 20 0 4

5

6

7

8

9

pH

Figure 9. Effect of pH on hydrolytic activities of the free and immobilized lipases. Immobilization conditions: coupling temperature, 4 °C; coupling time: 3h; lipase amount, 2 mg. Reaction conditions: reaction time, 30 min; reaction temperature 40°C. The maxima were defined as 100% activity.

Free Lipase Immobilized Lipase

Relative activity (%)

100 80 60 40 20 0 20

40

60

80

100

120

time (min)

Figure 10. Effect of time on hydrolytic activities of the free and immobilized lipases. Immobilization conditions: coupling temperature, 4 °C; coupling time: 3h; lipase amount, 2 mg. Reaction conditions: reaction temperature, 40 °C. The maxima were defined as 100% activity.

Relative activity (%)

100 Free Lipase Immobilized Lipase

80 60 40 20 0 0

20

40

60

80

100

120

storage time (days)

Figure 11. Effect of storage time on hydrolytic activities of the free and immobilized lipases. Immobilization conditions: coupling temperature, 4 °C; coupling time: 3h; lipase amount, 2 mg. Reaction conditions: reaction time 30 min; reaction temperature, 40 °C. The maxima were defined as 100% activity.

100

Yield (%)

80

60

40 Free Lipase Immobilized Lipase

20

0 4

6

8

10

time (h)

Figure 12. Ester yield of the free and immobilized lipases at different time. Immobilization conditions: coupling temperature, 4 °C; coupling time: 3h; lipase amount, 2 mg. Reaction conditions: reaction temperature, 40 °C.

Immobilized Lipase

Relative activity (%)

100

80

60

40

20 0

1

2

3

4

5

6

7

8

Cycle Number

Figure 13. Effect of cycle number on ester formation of immobilized lipases. Immobilization conditions: coupling temperature, 4 °C; coupling time: 3h; lipase amount, 2 mg. Reaction conditions: reaction time 8 h; reaction temperature, 40 °C. The maxima were defined as 100% activity.

Relative activity (%)

100 80 60 40 Free Lipase Immobilized Lipase

20 0 0

20

40

60

storage time (days)

Figure 14. Effect of storage time on ester formation of the free and immobilized lipases. Immobilization conditions: coupling temperature, 4 °C; coupling time: 3h; lipase amount, 2 mg. Reaction conditions: reaction time 8 h min; reaction temperature, 40 °C. The maxima were defined as 100% activity.

80

300 K

Fe3O4@OA

60 40

-20

5K

40

300 K

20

20

M (emu/g)

0

M (emu/g)

M (emu/g)

20

40

0

-20 -40

0 -20 -40

-1000

-500

0

500

1000

-1500 -1000 -500

0

500

1000 1500

H (Oe)

H (Oe)

-40 -60

a -80 -90000

-60000

-30000

0 H (Oe)

30000

60000

90000

80

300 K

Fe3O4@CA_stirred 60 40

-20

5K

20

M (emu/g)

0

40

300 K

20

M (emu/g)

M (emu/g)

20

40

0 -20 -40

0 -20 -40

-1000

-500

-40

0

500

1000

-1500 -1000 -500

H (Oe)

0

500 1000 1500

H (Oe)

-60

b -80 -90000

80

-60000

-30000

0 H (Oe)

30000

60000

90000

300 K

Fe3O4@CA_L_stirred

60 40 40

-20

0 -20 -40 -1000

-40

5K

20

M (emu/g)

0

20

M (emu/g)

M (emu/g)

20

40

300 K

0 -20 -40

-500

0

500

1000

-1500 -1000 -500

H (Oe)

0

500 1000 1500

H (Oe)

-60

c -80 -90000

-60000

-30000

0 H (Oe)

30000

60000

90000

Figure 15. Magnetic hysteresis loop at T=300 K, for the sample Fe3O4@OA (a), Fe3O4@CA_stirred (b) and Fe3O4@CA_L_stirred (c).

Table 1 Immobilization efficiency (%) and activity recovery (%) at different pH

T=40°C pH

Immobilization efficiency %

Activity Recovery % pH 3

3

60.2

65.3

4

74,1

7

73,9

Activity Recovery % pH 4

Activity Recovery % pH 7 80,9

52.4

121,9 144,4