Dodecenyl succinylated alginate as a novel material for encapsulation and hyperactivation of lipases

Carbohydrate Polymers 133 (2015) 194–202

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Carbohydrate Polymers journal homepage: www.elsevier.com/locate/carbpol

Dodecenyl succinylated alginate as a novel material for encapsulation and hyperactivation of lipases Mia Falkeborg a,b , Pattarapon Paitaid a,c , Allen Ndonwi Shu a , Bianca Pérez a , Zheng Guo a,∗ a b c

Department of Engineering, Aarhus University, Gustav Wieds Vej 10, DK-8000 Aarhus C, Denmark Center for Food Technology, Danish Technological Institute, Kongsvang Alle 29, DK-8000 Aarhus C, Denmark Department of Industrial Biotechnology, Faculty of Agro-Industry, Prince of Songkla University, Thailand

a r t i c l e

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Article history: Received 23 March 2015 Received in revised form 21 June 2015 Accepted 26 June 2015 Available online 8 July 2015 Keywords: Alginate Dodecenyl succinylated alginate Lipase Encapsulation Hyperactivation

a b s t r a c t Alginate was modified with dodecenyl succinic anhydride (SAC12) in an aqueous reaction medium at neutral pH. The highest degree of succinylation (33.9 ± 3.5%) was obtained after 4 h at 30 ◦ C, using four mole SAC12 per mol alginate monomer. Alginate was modified with succinic anhydride (SAC0) for comparison, and the structures and thermal properties of alg-SAC0 and alg-SAC12 were evaluated using FTIR, 1 H NMR, and DSC. Calcium-hydrogel beads were formed from native and modified alginates, in which lipases were encapsulated with a load of averagely 76 ␮g lipase per mg alginate, irrespective of the type of alginate. Lipases with a “lid”, which usually are dependent on interfacial activation, showed a 3-fold increase in specific activity toward water-soluble substrates when encapsulated in alg-SAC12, compared to the free lipase. Such hyperactivation was not observed for lipases independent of interfacial activation, or for lipases encapsulated in native alginate or alg-SAC0 hydrogels. © 2015 Elsevier Ltd. All rights reserved.

1. Introduction Alginate is a naturally occurring polysaccharide composed of ␣-l-guluronate and ␤-d-mannuronate arranged as linear homopolymeric and heteropolymeric blocks (Pawar & Edgar, 2012). Alginate is widely used for encapsulation purposes, commonly in the form of a hydrogel, which forms through coordination of divalent cations to the carboxylate groups in the guluronate moieties (Lee & Mooney, 2012). Calcium(II) is commonly used to form alginate hydrogels, and calcium–alginate hydrogels have been used to encapsulate probiotics (Dong et al., 2013), live cells for use in biomedicine (Lim & Sun, 1980), as well as enzymes for various technical applications (Won, Kim, Kima, Park, & Moon, 2005; Zhang Shang et al., 2014). Numerous derivatizations of alginate have been performed in order to tailor alginate toward specific applications (Pawar & Edgar, 2012; Yang, Xie, & He, 2011). Generally, the alginate polymer can be modified at the secondary hydroxyl groups at C2 and C3, and/or at the carboxyl groups. In this study, alginate was modified with succinic anhydrides (SAC0) and dodecenyl succinic anhydrides

∗ Corresponding author. Fax: +45 86 12 31 78. E-mail addresses: [email protected] (M. Falkeborg), [email protected] (P. Paitaid), [email protected] (A.N. Shu), [email protected] (B. Pérez), [email protected] (Z. Guo). http://dx.doi.org/10.1016/j.carbpol.2015.06.103 0144-8617/© 2015 Elsevier Ltd. All rights reserved.

(SAC12), forming succinylated alginates (alg-SAC0 and alg-SAC12), as illustrated in Fig. 1. Lipases (E.C. 3.1.1.3) are biocatalyst used in various industrial applications (Reis, Holmberg, Watzke, Leser, & Miller, 2009). Encapsulation of lipases can aid their reusability and protect the biocatalyst from external stresses. Lipases have successfully been encapsulated in alginate hydrogels by mixing the lipase with an aqueous solution of alginate, followed by drop-wise addition of the alginate–enzyme solution into an aqueous solution of calcium chloride (Toscano, Montero, Stoytcheva, Cervantes, & Gochev, 2014; Won et al., 2005; Zhang Shang et al., 2014). Lipases encapsulated in alginate hydrogels show good stability and good reusability, but generally suffer from lower activity compared to the free lipase, primarily due to mass transfer limitations (Toscano et al., 2014; Won et al., 2005). Most lipases have elements of secondary structure covering their active site (generally termed the “lid”), which opens in the presence of interfaces such as an oil–water interface or the surface of a micelle (Reis et al., 2009). The lipase is fully active only with the “lid” open, and lipases with a “lid”, such as the lipase from Thermomyces lanuginosus (TLL), show only low activity against water-soluble substrates. Studies have shown that encapsulating such lipases using hydrophobic supports can lead to hyperactivation of the lipase, as the hydrophobicity of the support enables the lipase to be encapsulated in an active “open” form (Bastida et al., 1998; Rodrigues, Ortiz, Berenguer-Murcia, Torres,

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Fig. 1. Reaction scheme of succinylation of alginate with SAC12 (left) or SAC0 (right).

& Fernandez-Lafuente, 2013; Wilson et al., 2006). For example, Bastida et al. (1998) showed that lipases from Rubus niveus, Rhizomucor miehei, and Humicola lanuginosa were 6-, 7-, and 20-fold more active toward water-soluble substrates when encapsulated in octyl-agarose gels, compared to the corresponding free lipases. Such hyperactivation of lipases towards water-soluble substrates is particularly advantageous in medical applications, where lipasecatalyzed resolution of drugs (water soluble or partially soluble), e.g., menthol (Bai, Guo, Liu, & Sun, 2006), is an important area of lipase application. The present work is based on our recent findings with alginatebased oligomers/polymers, obtained upon depolymerization or modification, which demonstrated interesting properties as ingredients in lipid encapsulation and formulation (Falkeborg & Guo, 2015; Falkeborg et al., 2014). We hypothesize that modification of alginate with hydrophilic SAC0 and hydrophobic SAC12 may not only change the chemical structure, but also alter the polymer packing and chemical properties of alginate, and create possible interactions with proteins/enzymes. Hence, the properties and activities of lipases encapsulated in these modified alginates were examined in this study. Two lipases, Candida antartica lipase B (CALB) and T. lanuginosus lipase (TLL), were selected as respective representatives of non-interfacial- and interfacial active lipases. This work was aimed to exploit the application potentials of algSAC12 as a novel biomimicking material for use in biotechnological areas. 2. Materials and methods 2.1. Materials Sodium alginate (Grindsted® Alginate FD 170) was provided by DuPont, Brabrand, Denmark. This alginate originated from brown algae, and had an average molecular weight of 100 kDa.

The ratio of ␣-L-guluronate units to ␤-d-mannuronate units was 40–60. Lipase B from C. antartica (CALB) and lipase from T. lanuginosus (TLL) were obtained from Novozymes Denmark in liquid formulations containing 25% propylene glycol. The formulations contained 30 mg lipase/mL as determined from the bicinchoninic acid (BCA) protein assay (Section 2.8), and were used as received. Succinic anhydride (SAC0), 2-dodecen-1-yl succinic anhydride (SAC12), tris(hydroxymethyl) aminomethane (Tris; ≥99.8%), p-nitrophenylbutyrate (p-NPB; 98%), cupric acetate–pyridine reagent, and calcium chloride (CaCl2 ), were from Sigma–Aldrich Co., Ltd, Denmark.

2.2. Alginate succinylation Alginate was succinylated with SAC0 and SAC12 with inspiration from previously reported methods (Le-Tien, Millette, Mateescu, & Lacroix, 2004; Rao, Prakasham, Rao, & Yadav, 2008). Small-scale alginate succinylations for process optimization were performed in 20 mL capped vials, thermostated using circulating water and stirred with magnetic bars at 300 rpm. Sodium alginate (100 mg, corresponding to 0.51 mmol monomeric units) was dissolved in 10 mL pure water, followed by addition of 2.02 mmol SAC0 or SAC12. The pH was adjusted to 7.0 ± 0.3 using 1% sodium hydroxide (NaOH) and an automated pH-meter (inoLab pH 7110, Wissenschaftlich-Technische Werkstätten GmbH, Weilheim, Germany). After 4 h, the succinylated products (alg-SAC0 and alg-SAC12) were precipitated using 35 mL ethanol and isolated by centrifugation at 4000 rpm for 10 min at 20 ◦ C. Excess SAC0 and SAC12, and water, were removed by washing four times with 50 mL acetone. The reaction velocities were followed by determining the degree of succinylation (DS) after varying reaction times (1–8 h). Optimization toward a high DS was performed with SAC12 by varying the reaction temperature (25, 30, 40, and 50 ◦ C) and the mole

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excess of SAC12 with respect to alginate monomeric units (1–4, and 5-times mole excess). Larger-scale preparations of alg-SAC0 and alg-SAC12 were performed at optimum conditions in 250 mL screw-capped flasks at 30 ◦ C in 4 h with magnetic stirring at 400 rpm. Alginate (750 mg) was dissolved in 75 mL H2 O, followed by addition of 15.2 mmol SAC0 or SAC12 (substrate molar ratio 1:4). The pH was adjusted to 7.0 ± 0.3 using 3% NaOH. The succinylated alginates were precipitated in 300 mL ethanol and isolated by vacuum filtration; washed four times with 200 mL acetone; and finally dried at room temperature for 48 h. The structures and thermal properties of alg-SAC0 and alg-SAC12 were analyzed by 1 H nuclear magnetic resonance (NMR), Fourier transformed infrared spectrometry (FTIR), and differential scanning calorimetry (DSC) (Section 2.4). 2.3. Degree of succinylation (DS)

V − V control × 100 Vcontrol

2.6. Hydrogel bead characterization The morphologies of the hydrogel beads were observed at room temperature in a Leica M165FC stereomicroscope with a PLANAPO 1,0x objective. The size distributions of the hydrogel beads were determined by digital inspection of images of 30 randomly selected beads from each batch, using a digital single-lens reflex camera. The diameter of each bead was determined at two angles and averaged. The beads were grouped according to size (±50 ␮m) and the number of beads in each group was counted.

2.7. Encapsulation of lipases

The DS was determined as reported previously (Falkeborg & Guo, 2015), shortly as follows. Succinylated alginates (100 mg) were dispersed in 10 mL 2.5 M HCl in isopropanol (IPA) and shaken for 10 min. Additional 20 mL IPA was added and the acidified products were centrifuged at 4000 rpm at 17 ◦ C for 10 min. The precipitated products were washed 3 times with 40 mL IPA, dispersed in 30 mL H2 O, and titrated with 0.1 M NaOH using phenolphthalein as an indicator. Alginate (100 mg) was acidified and titrated correspondingly as a control. The DS was quantified according to Eq. (1), where V is the titration volume of succinylated alginate, and Vcontrol is the titration volume of alginate. The DS was determined as the average of duplicate titrations. DS = 0.5 ×

the beads were determined immediately after preparation (Section 2.6).

(1)

2.4. Characterization of succinylated alginates by 1 H NMR, FTIR, and DSC 1 H NMR spectra were recorded in deuterium oxide at room tem-

perature on a Bruker Avance III 400 spectrometer. FTIR spectra were recorded in absorbance mode in the 4000–650 cm−1 region at a resolution of 4 cm−1 , using a Qinterline QFAflex spectrometer equipped with a deuterium triglycine sulfate detector. The samples were mounted in their pure solid form in a Pike attenuated total reflectance (ATR) device at 25 ◦ C, and the spectra were ratioed against a single-beam spectrum of the clean ATR crystal. DSC measurements were performed in a Pyris 6 DSC from PerkinElmer, using a sample mass of 8–12 mg in an aluminum pan. The temperature was increased from 20 ◦ C to 350 ◦ C at a rate of 5 ◦ C/min under nitrogen flowing at 20 mL/min. 2.5. Hydrogel bead formation Hydrogel beads were formed from the native and succinylated alginates with inspiration from previously reported methods (Betigeri & Neau, 2002; Won et al., 2005; Zhang Shang et al., 2014). Alginate, alg-SAC0, or alg-SAC12, was dissolved in 50 mM Tris–HCl buffer (pH 7.3) to 10 g/L and stirred overnight at 300 rpm at room temperature for complete hydration. Portions of 0.5 mL of this alginate solution were added drop wise (approximately 0.25–0.50 mL/min) from a plastic syringe through a 27-gauge needle (0.4 × 20 mm), from a distance of approximately 5 cm, into 3 mL 1 M CaCl2 at 4 ◦ C. The alginate beads were left in the CaCl2 solution for 20–30 min to cure. The CaCl2 solution was then carefully drained off using a thin-tipped pipette, and the alginate beads were washed twice by dispersing them in 3 mL 50 mM Tris–HCl buffer (pH 7.3) for 2 min under mild magnetic stirring (∼100 rpm), and draining off the washing buffer. The size distributions and morphologies of

Lipases (CALB and TLL) were encapsulated in alginate hydrogel beads by mixing the alginate solution (2 mL) with the lipase solution (0.125 mL) at room temperature for 10 min, before forming the beads by drop wise addition into the 1 M CaCl2 solution, as described in Section 2.5. The loaded alginate hydrogel beads were characterized as described in Section 2.6, and the encapsulation efficiency was measured by determining the protein content in the recovered CaCl2 fraction and in the wash fractions (Section 2.8). The activities and reusability of the encapsulated lipases were determined as described in Section 2.9 and Section 2.10, respectively.

2.8. Encapsulation efficiency Protein contents were determined using the Thermo Scientific PierceTM BCA Protein Assay Kit enhanced protocol, suitable for detecting protein concentrations in the range 5–250 ␮g/mL. A standard curve prepared from solutions of bovine serum albumin in 50 mM Tris–HCl buffer (pH 7.3) was used to determine the contents of lipase in the original formulation and in the residual CaCl2 solutions and wash fractions. The protein contents were determined as averages of three samples with duplicate spectrophotometric measurements.

2.9. Assay of lipase activity The activities of the lipases in free and encapsulated forms were determined as the rate of hydrolysis of water-soluble p-NPB at 30 ◦ C, with inspiration from the methods of Won et al. (2005). A substrate solution containing 4 mM p-NPB in 50 mM Tris–HCl buffer with 12% acetonitrile was prepared and incubated for 2 h at 30 ◦ C before each analysis. The incubation with acetonitrile ensured that the substrate p-NPB was fully solubilized and not present in micellar forms. The substrate solution was filtered through a 0.45 ␮m membrane and combined with either free lipase (11 ␮L liquid formulation, corresponding to 330 ␮g lipase) or portions of alginate beads (prepared from 0.5 mL alginate–lipase solution). Samples of 25 ␮L were withdrawn every 1 min and dissolved in 975 ␮L 50 mM Tris–HCl buffer (pH 7.3), and the absorbance at 400 nm was determined immediately in a UV–visible spectrophotometer (Pharmacia LKB, Utrospec II). The lipase activity was determined as the rate of formation of p-nitrophenol, determined as the initial increase in absorbance at 400 nm, using the extinction coefficient of pnitrophenol (18,000 M−1 cm−1 )(Zhang & Vanetten, 1991). Activity was expressed in units of ␮M p-nitrophenol/min, and specific enzymatic activity was expressed as enzymatic activity per mg protein encapsulated in the beads.

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2.10. Assay of lipase reusability To evaluate the reusability of the encapsulated lipase (TLL), an assay based on hydrolysis of olive oil was employed. The rate of hydrolysis of olive oil was determined in a two-phase system at 37 ◦ C based on the colorimetric determination of liberated fatty acids (Kwon & Rhee, 1986). Isooctane (1 mL) containing 10% (w/v) olive oil was combined with 1 mL 50 mM Tris–HCl buffer (pH 7.0) and either 100 mg TLL-loaded alginate hydrogel beads, 100 mg TLLloaded alg-SAC12 hydrogel beads, or free TLL corresponding to the amount encapsulated in 100 mg hydrogel beads (5.5 ␮L), and the mixture was shaken at 1200 rpm for 15 min. Aliquots of 0.1 mL were withdrawn periodically from the upper layer and mixed with 0.9 mL isooctane and 0.4 mL cupric acetate-pyridine reagent, followed by determination of the absorbance at 715 nm. A standard curve for this colorimetric assay was prepared using oleic acid (R2 = 0.9965). One unit lipase activity was defined as the amount of enzyme required to liberate one ␮mol free acid per minute. The TLL-loaded hydrogel beads were retrieved from the reaction mixture and washed three times with isooctane to remove residual substrate and product, before being used for a new round of reaction. The reusability was examined for up to 10 cycles. 2.11. Statistics Data processing including regression analyses were performed in Microsoft Excel 2013. All measurements were conducted with two-to-four repeats, and the results are reported as means ± standard deviations. One-way analysis of variance (oneway ANOVA) were performed using Microsoft Excel Analysis Toolpak (2010) to identify differences between groups. 3. Results and discussion 3.1. Alginate succinylation The modifications of alginate with SAC0 and SAC12 were performed in aqueous reaction media at pH neutral conditions and low temperature. No catalysts were needed due to the high reactivity of the anhydride substrates. A series of reactions with varying substrate ratio, reaction temperature, and reaction time, were performed to evaluate the influence of these parameters on the degree of succinylation of alginate. Fig. 2A shows the time course of the succinylation reactions, using SAC0 and SAC12 in 4-times mole excess at 30 ◦ C. For both reactions, no significant increase (p > 0.3) in the degree of succinylation was observed after 4 h of reaction. The degree of succinylation was higher for alg-SAC12 compared to SAC0. This observation could be related to the water solubility of the substrates and their stability in water. Anhydrides are unstable in water and lose reactivity upon reaction with water (ring opening). SAC0 can be dissolved readily in water, and the higher susceptibility of this anhydride towards water hydrolysis could have caused the lower degree of succinylation compared to SAC12, which had a much lower solubility in water and higher resistance to hydrolysis. The influence of temperature was examined for the succinylation of alginate with 4-times mole excess of SAC12 (Fig. 2B). Changes in reaction temperature affects the degree of succinylation in contradicting ways, as has also been reported for alkylsuccinylations of starch (Bhosale & Singhal, 2006; Song, He, Ruan, & Chen, 2006). Lower reaction temperature slows the reaction rate and decreases the substrates’ solubility in water. In principle, higher temperature increases the reaction velocity kinetically; however, it also leads to faster anhydride ring opening, which decreases the reactivity of the anhydride. Thirty Celsius degrees was found to be the optimum temperature for succinylation of alginate in this

Fig. 2. Degree of alginate succinylation at varying (A) reaction times, (B) reaction temperature (for reaction with SAC12), and (C) molar ratio of alginate monomeric units and SAC12.

study, which is comparable to the values reported for succinylation of starch (Bhosale & Singhal, 2006; Song et al., 2006). The effect of increasing the dosage of SAC12 was studied (Fig. 2C), and as expected, the degree of succinylation increased with increasing amounts of SAC12. The degree of succinylation leveled off at four-time mole excess, and no significant (p = 0.235) increase in the degree of succinylation was observed by increasing the mole ratio further. This phenomenon could be related to an effect of steric hindrance, in that, due to the more bulky structure of the alkyl substitution, not every monomeric unit in alginate could be substituted. The highest degrees of alginate succinylation were obtained using 4-times mole excess of SAC0 and SAC12, after four hours of reaction at 30 ◦ C. Under these conditions, the degree of succinylation in alg-SAC0 and alg-SAC12 were 24.5 ± 2.7% and 33.9 ± 3.5%, respectively. This interprets to that, on average, every second monomeric unit of alginate were modified with one SAC0 moiety,

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and two out of three monomeric unit of alginate were modified with one SAC12 moiety, for alg-SAC0 and alg-SAC12, respectively. 3.2. Structure identification The succinylations of alginate were structurally verified using NMR and FTIR. All 1 H NMR spectra of native and succinylated alginates showed a series of multiplets in the 3.0–5.5 ppm region corresponding to the protons in the alginate backbone (H C OH and H C O C). The 1 H NMR spectrum of alg-SAC12 displayed two additional signals in the upfield region: a multiplet at 1-4-1.1 ppm and another multiplet at 0.9–0.7 ppm associated to the protons of the secondary and primary alkyl group from the dodecenyl substituents, respectively. Furthermore, the spectrum of alg-SAC0 showed a peak at 2.4 ppm characteristic of the H 2 C COOH originating from the modification with SAC0 (1 H NMR data are not shown). FTIR spectra of native and succinylated alginates are presented in Fig. 3. Distinct bands assignable to the alkane moieties of the substitutions were detected at 2925 cm−1 and 2854 cm−1 in alg-SAC12, and at 2938 cm−1 and 2914 cm−1 in alg-SAC0. In the spectrum of succinylated alginates, the C O asymmetric stretching vibrations of the carboxylates in the alginate backbone, the free acids in the substitutions, and the ester bonds between alginate and succinic anhydride, are not well separated, but can be identified at 1596 cm−1 , 1649 cm−1 , and 1728 cm−1 for alg-SAC12; and at 1601 cm−1 , 1669 cm−1 , and 1740 cm−1 for alg-SAC0, respectively. The band of the carboxylate asymmetric stretching vibrations (1603 cm−1 ) is expectedly also present in the native alginate. In all spectra, two bands at ∼1080 cm−1 and ∼1030 cm−1 occur. These are assigned to the C O and C C stretching vibrations of the pyranose rings in alginate (Falkeborg et al., 2014). The bands at ∼945 cm−1 in each spectra are assigned to the 1 → 4 glycosidic linkages in the alginate (Chandia, Matsuhiro, & Vasquez, 2001). The alginates all have the characteristic broad absorbance band in the region 3200–3500 cm−1 , assignable to the hydroxyl stretching vibrations. The intensity of this band is higher in the succinylated alginates, as it is contributed from the O H stretching vibration of the free carboxylic acids (Yadav, 2005). 1H

3.3. Thermal properties The thermal properties of the native and modified alginates were analyzed by DSC and the results are presented in Fig. 4. All alginates underwent a significant endothermic phase transition, which for native alginate occurred at 149.6 ◦ C, preceded by a small endothermic event at 143.2 ◦ C. For alg-SAC0, the phase transition occurred over a broader temperature range and peaked at a slightly lower temperature (145.6◦ ). For alg-SAC12, the endothermic phase transition occurred at a significantly lower temperature, 123.3 ◦ C, which indicates that this polymer had lower interchain interaction forces and a lower degree of molecular packing ordering. Most likely, this is due to the introduced dodecenyl succinyl groups, which disturb the hydrogen-bonding network originally occurring in alginate. A small endothermic peak is observed at 59.7 ◦ C in the thermogram of alg-SAC0, which is not observed for native alginate or alg-SAC12. This peak is probably related to the loss of water, which shows that the drying process using acetone was less successful for alg-SAC0 compared to alg-SAC12, possibly as a consequence of a higher water-binding capacity of alg-SAC0 compared to alg-SAC12. The alginates showed a broad exothermic peak at 226.4 ◦ C, 227.0 ◦ C, and 220.9 ◦ C, for alginate, alg-SAC12, and alg-SAC0, respectively. In agreement with related studies (Abulateefeh, Khanfar, Al Bakain, & Taha, 2014; Soares, Santos, Chierice, & Cavalheiro, 2004), we attribute these exothermic peaks to the

decomposition of the polymer through reactions such as decarboxylation and oxidation. In agreement with this, the alginate samples appeared black and “charred” after the analysis, and an expansion of the samples at high temperatures led to breaking of the aluminum pans containing the alginates at ∼250 ◦ C. 3.4. Characterization of alginate hydrogel beads The native and succinylated alginates all formed beads upon contact with the CaCl2 solution. The beads appeared transparent for the native alginate and alg-SAC0, and turbid/white for algSAC12. Approximately 70–90 hydrogel beads were formed per batch (0.5 mL), which is comparable to, or slightly higher than values previously reported (Taqieddin & Amiji, 2004). The size distributions of loaded and unloaded hydrogel beads are presented in Fig.5. Beads loaded with CALB were similar to beads loaded with TLL, and only the size distributions of beads loaded with TLL are presented. Beads formed from native alginate had a broader size distribution compared to beads formed from alg-SAC0 and algSAC12. No significant difference (p = 0.281) was observed between the sizes of beads formed from native alginate and alg-SAC12, whereas beads formed from alg-SAC0 were significantly smaller (p < 0.001). For alginate and alg-SAC0, the bead size increased when TLL was encapsulated in the hydrogel matrix. The average bead size increased from 2.30 mm (unloaded) to 2.64 mm (loaded) for native alginate (p = 0.0019), and from 1.77 mm (unloaded) to 2.00 mm (loaded) for alg-SAC0 (p < 0.001). Oppositely, for alg-SAC12, no significant (p = 0.488) change in average bead size was observed between unloaded (2.38 mm) and loaded (2.33 mm) beads, yet, a portion of smaller hydrogel beads were obtained when loaded with TLL. These results indicate that the mechanism of encapsulation of lipases may be different for alg-SAC12 compared to alg-SAC0 and native alginate. The sizes of the alginate hydrogel beads are comparable to values reported in literature (Toscano et al., 2014; Won et al., 2005). Generally, the bead sizes can be varied by using different syringe tips during preparation, as has been shown by Taqieddin and Amiji (2004) and Won et al. (2005), and automated methods for formation of alginate hydrogel beads have also been developed (Liu, Kost, Yan, & Spiro, 2012). The morphologies of the beads were observed using optical stereomicroscopy, and respective images of all hydrogel beads, unloaded and loaded, are presented in Fig. 6. The beads formed from native alginate were irregular in shape, which correlates well with their broad size distribution. The beads formed from alg-SAC0 and alg-SAC12 were more spherical and had smoother surfaces. The unloaded beads formed from alg-SAC0 showed protrusions, probably originating from the syringe tip used in the preparation process. The beads formed from alg-SAC12 did not change their shape upon loading with TLL, but remained spherical and turbid. 3.5. Lipase encapsulation efficiency The efficiencies of the native and succinylated alginates for encapsulation of lipases were determined (Fig. 7). On average, 82.5 ± 6.3 ␮g CALB and 70.1 ± 3.9 ␮g TLL could be encapsulated per mg alginate carrier material. This corresponds to 38% and 44% of the initial amount of lipase added, for CALB and TLL, respectively. The main loss of enzyme during the encapsulation process probably occurred during the initial gelling period, when the hydrogel network had not yet fully formed (Betigeri & Neau, 2002). Due to variations in the encapsulation procedure, such as amount of lipase initially added and its ratio to the amount of carrier material, actual loadings, and not encapsulation efficiencies in percentages, should be used when comparing the loading capacities to values reported in literature. Thus, when using similar starting concentrations as in this study, Won et al. (2005) encapsulated 75–85 ␮g lipase per mg

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Fig. 3. FTIR spectra of native alginate, alg-SAC12, and alg-SAC0.

Fig. 4. DSC thermograms of alginate, alg-SAC12, and alg-SAC0.

Fig. 5. Size distributions of hydrogels beads, unloaded (top panels) or loaded with TLL (lower panel).

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Fig. 6. Microscopy images of hydrogel beads unloaded (top panels) or loaded with TLL (lower panel). Scale bars (in red in the lower right corner) are 200 ␮m.

Fig. 7. Lipase load (␮g lipase/mg alginate) of TLL and CALB in alginate, alg-SAC0, and alg-SAC12 hydrogel beads.

alginate; and Betigeri & Neau (2002) encapsulated 56–66 ␮g per mg alginate, which is comparable to the loading capacities obtained in this study. A slightly higher encapsulation efficiency was observed for CALB compared to TLL (p = 0.096, p = 0.004, and p = 0.009, for alginate, alg-SAC0, and alg-SAC12, respectively). This trend could be explained by differences in interactions between the lipase and the alginate, which may have been stronger for CALB compared to TLL. This was not examined further in this study. Alg-SAC12 was found capable of encapsulating more CALB (91.3 ± 6.1 ␮g/mg) compared to native alginate (79.3 ± 4.4 ␮g/mg; p = 0.006) or algSAC0 (76.9 ± 3.6 ␮g/mg; p = 0.001); this trend was, however, not observed for encapsulation of TLL, which was encapsulated by all alginates to a similar extent (p = 0.213).

3.6. Specific lipase activity and hyperactivation The water-soluble p-NPB was used as substrate for determining lipase activity. To obtain this substrate in soluble form, and not in micellar form, the substrate solution was incubated with acetonitrile for 2 h prior to analysis. The very low specific activity of free TLL toward this substrate solution, compared to its activity towards the substrate solution before incubation (not shown), confirms that the substrate p-NPB was in fully soluble form. During the

Fig. 8. Specific lipase activity (␮M/min/mg lipase) of free and encapsulated CALB (A) and TLL (B).

incubation time, less than 0.3% of the substrate was hydrolyzed to p-nitrophenol due to autohydrolysis. Unloaded hydrogel beads did not catalyze the hydrolysis of pNPB (data not shown). Free CALB had a specific activity against soluble p-NPB of 303.9 ± 23.3 ␮M/min/mg lipase. When encapsulated, the specific activity of CALB was significantly (p < 0.001) decreased to an average of 74.6 ± 11.8 ␮M/min/mg lipase. The specific activity of CALB did not differ (p = 0.891) depending on the type of alginate hydrogel in which it was encapsulated (Fig. 8A). As CALB do not have a hydrophobic “lid” in its structure, it is independent of interfacial activation. The hydrophobic moieties introduced in the alginate through modification with SAC12 were hence not expected

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Fig. 9. Reusability of TLL encapsulated in modified (alg-SAC12) and native alginate hydrogel beads per 100 mg hydrogel beads. *The activity of free TLL is presented per corresponding amount of free enzyme (5.5 ␮L liquid formulation).

to influence the specific activity of this lipase, as was also experimentally observed (Fig. 8A). The decrease in specific activity when CALB was encapsulated in the alginate hydrogels might be due to mass transfer limitations, which possibly could be eliminated by making smaller beads (Won et al., 2005). Physical or ionic interactions between the alginate polymer and the enzyme has also been suggested to decrease the specific activity of enzymes encapsulated in alginate hydrogels (Betigeri & Neau, 2002). These parameters were not evaluated in this study, but the studies with CALB confirm that the type of alginate, succinylated or native, did not alter the mass transfer limitations or deactivating interactions, as CALB had the same specific activity regardless of the type of alginate carrier. TLL is dependent on interfacial activation for high activity, and free TLL showed low specific activity against the water-soluble pNPB (75.8 ± 4.7 ␮M/min/mg lipase). When encapsulated in native alginate or alg-SAC0 hydrogels, the specific activity of TLL increased to an average of 118.0 ± 8.1 ␮M/min/mg lipase, with no significant difference between the two carriers (p = 0.490). However, more than 3-fold increase in specific activity was observed when TLL was encapsulated in alg-SAC12, compared to the specific activity of free TLL. This hyperactivation of TLL when encapsulated in algSAC12 hydrogel beads is proposedly related to the hydrophobic moieties in alg-SAC12. The TLL recognizes the hydrophobic moieties as interfaces, and the hydrophobic pocket of the lipase, formed by the area surrounding the active site and the internal part of the “lid”, is exposed, and may adsorb to the hydrophobic parts of the carrier material (Rodrigues et al., 2013). Hence, TLL might exist in an active form with the “lid” open when it is encapsulated in algSAC12 hydrogel, leading to the observed higher specific activity of the encapsulated lipase. The mass transfer limitations observed for CALB are expectedly also decreasing the activity of TLL, as the TLL encapsulated in the center of the beads expectedly is less active. This consideration makes the observed 3-fold increase in specific activity even more pronounced. The effects of mass transfer limitations could be eliminated by making smaller beads. Several other improvements to enzyme encapsulation in alginate hydrogels have been reported in literature, including coating of the alginate beads with chitosan and/or freeze-drying (Taqieddin & Amiji, 2004), or crosslinking with gluteraldehyde (Fadnavis, Sheelu, Kumar, Bhalerao, & Deshpande, 2003). Such processes can expectedly also be applied to hydrogel beads formed from alg-SAC12, thus combining the

hyperactivating property of this matrix material with improved encapsulation properties. 3.7. Lipase reusability The ability to reuse the lipase is the main advantage of lipase encapsulation, and accordingly, the reusability of TLL encapsulated in alginate and alg-SAC12 hydrogel beads were evaluated. The results are presented in Fig. 9. Free TLL had an activity of 6.2 U/5.5 ␮L and could not be reused due to its liquid nature. Encapsulation in native alginate led to a decrease in hydrolytic activity compared to the free TLL (3.7 U/100 mg in the first round), probably related to mass transfer limitations as discussed in Section 3.7. Comparatively, TLL encapsulated in alg-SAC12 hydrogel beads had an activity of 7.8 U/100 mg beads in the first round, and hence, as observed previously, encapsulation in this matrix led to a hyperactivation of the TLL. A significant decrease in lipase activity was observed after the first round of hydrolysis, and 65% of the original lipase activity was recovered after the first round of reuse for both types of alginate hydrogel beads. This observation might be ascribed to the deactivation of the portion of the lipase located on the hydrogel bead surface due to washing with organic solvent. The activity of TLL encapsulated in alg-SAC12 was stabilizing after four cycles of reuse, and no further activity was lost upon further reuse. Oppositely, the residual activity after four cycles of TLL encapsulated in native alginate was only 16%, and after eight cycles, the alginate hydrogel beads were completely dissolved in the reaction mixture and could no longer be recovered for additional rounds of reuse. Hence, alg-SAC12 as a carrier might provide increased protection against solvent-induced lipase deactivation compared to native alginate. After 10 cycles of reuse, the TLL encapsulated in alg-SAC12 was able to retain about 50% of its original activity, indicating the potential of alg-SAC12 in industrial applications. 4. Conclusion In this study, alginate was chemically modified with dodecenyl succinic anhydrides, and the resulting alg-SAC12 was capable of encapsulating lipase in a hyperactivated form. The modification procedure took place in an aqueous reaction medium at low temperature and neutral pH, and required no catalysts or

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organic solvents, which is an environmentally benign approach. The hydrophobically modified alginate could hyperactivate lipases due to the presence of hydrophobic moieties, and more than 3fold higher activity towards water-soluble substrate was observed for encapsulated lipase, compared to free lipase. The loading capacity of the modified alginate was not decreased compared to native alginate. The properties of alg-SAC12 thus combines the good hydrogel-forming and encapsulation properties of native alginate, with the hyperactivating properties usually observed for hydrophobic supports, when encapsulating interfacial-dependent lipases. The reusability of lipase encapsulated in alg-SAC12 was markedly enhanced compared to the reusability of lipase encapsulated in native alginate, which lost its activity completely after eight cycles of reuse, while lipase encapsulated in alg-SAC12 retained 50% of its activity after 10 cycles. The encapsulation properties of algSAC12 can expectedly be further improved by mechanisms already in use for native alginate, such as coating or cross-linking reactions. The hydrophobically modified alginate represents a new functional material with potential applications in the encapsulation of proteins/enzymes for use in biological or biotechnological areas, and this biomimicking material deserves further exploration. Acknowledgements This work was partly funded by “Innovation Consortium Multicaps” in collaboration with Danish Technological Institute, DuPont Denmark, Marine Bioproducts, and University of Southern Denmark. The authors thank Anna Malgorzata Jurkiewicz for the support with optical stereo microscopy analysis. References Abulateefeh, S. R., Khanfar, M. A., Al Bakain, R. Z., & Taha, M. O. (2014). Synthesis and characterization of new derivatives of alginic acid and evaluation of their iron(III)-crosslinked beads as potential controlled release matrices. Pharmaceutical Development and Technology, 19(7), 856–867. Bai, S., Guo, Z., Liu, W., & Sun, Y. (2006). Resolution of (±)-menthol by immobilized Candida rugosa lipase on super paramagnetic nano particles. Food Chemistry, 96(1), 1–7. Bastida, A., Sabuquillo, P., Armisen, P., Fernandez-Lafuente, R., Huguet, J., & Guisan, J. M. (1998). Single step purification, immobilization, and hyperactivation of lipases via interfacial adsorption on strongly hydrophobic supports. Biotechnology and Bioengineering, 58(5), 486–493. Betigeri, S. S., & Neau, S. H. (2002). Immobilization of lipase using hydrophilic polymers in the form of hydrogel beads. Biomaterials, 23(17), 3627–3636. Bhosale, R., & Singhal, R. (2006). Process optimization for the synthesis of octenyl succinyl derivative of waxy corn and amaranth starches. Carbohydrate Polymers, 66(4), 521–527. Chandia, N. P., Matsuhiro, B., & Vasquez, A. E. (2001). Alginic acids in Lessonia trabeculata: characterization by formic acid hydrolysis and FT-IR spectroscopy. Carbohydrate Polymers, 46(1), 81–87. Dong, Q. Y., Chen, M. Y., Xin, Y., Qin, X. Y., Cheng, Z., Shi, L. E., & Tang, Z. X. (2013). Alginate-based and protein-based materials for probiotics encapsulation: a review. International Journal of Food Science and Technology, 48(7), 1339–1351.

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