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International Journal of Biological Macromolecules 70 (2014) 78–85

Contents lists available at ScienceDirect

International Journal of Biological Macromolecules

j ourna l ho me pa g e: www.elsev ier .com/ locate / i jb iomac

ovalent attachment of lipases on glyoxyl-agarose beads:pplication in fruit flavor and biodiesel synthesis

driano A. Mendesa,b,∗, Heizir F. de Castroc, Raquel L.C. Giordanob

Institute of Chemistry, Federal University of Alfenas, 37130-000 Alfenas, MG, BrazilDepartment of Chemical Engineering, Federal University of São Carlos, 13565-905 São Carlos, SP, BrazilDepartment of Chemical Engineering, Engineering School of Lorena, University of São Paulo, 12602-810, Lorena, SP, Brazil

r t i c l e i n f o

rticle history:eceived 30 December 2013ccepted 20 June 2014vailable online 27 June 2014

eywords:ipase immobilizationlyoxyl-agarose beadsster synthesis

a b s t r a c t

The aim of this work was to prepare biocatalysts to catalyze the synthesis of butyl butyrate by esterifica-tion reaction, and the synthesis of biodiesel by transesterification of palm and babassu oils with ethanol.Lipase preparations Lipolase® (TLL1) and Lipex® 100L (TLL2) from Thermomyces lanuginosus and LipaseAK from Pseudomonas fluorescens (PFL) were immobilized on glyoxyl-agarose beads prepared by activa-tion with glycidol (Gly) and epichlorohydrin (Epi). The influence of immobilization time, lipase sourceand activating agents on the catalytic activity of the biocatalysts were evaluated in both aqueous andorganic media. TLL1 immobilized on glyoxyl-agarose by 24 h of incubation resulted biocatalysts withhigh hydrolytic activity (varying from 1347.3 to 1470.0 IU/g of support) and thermal-stability, around300-fold more stable than crude TLL1 extract. The maximum load of immobilized TLL1 was around 20 mg

of protein/g of support. The biocatalyst prepared exhibited high activity and operational stability on thebutyl butyrate synthesis by esterification after five successive cycles of 24 h each (conversion around85–90%). Immobilized TLL1 and PFL were active in the synthesis of biodiesel by transesterification reac-tion. Maximum transesterification yield (≥98.5% after 48 h of reaction at 45 ◦C) was provided by usingpalm oil as feedstock.

© 2014 Elsevier B.V. All rights reserved.

. Introduction

Lipases (triacylglycerol hydrolases E.C. 3.1.1.3) are enzymes thatn vivo catalyze the hydrolysis of triacylglycerols to glycerol andree fatty acids. These enzymes, in vitro, also catalyze esterification,ransesterification and interesterification reactions in non-aqueousystems [1–3]. A typical feature of lipases is the so-called “interfa-ial activation”, although some lipases have been identified thato not undergo increased activity in the presence of hydropho-ic interfaces. A polypeptide chain called “lid” or “flap” covers itsctive site, making it inaccessible to solvent and substrates in manynstances (closed conformation). The lid undergoes conformationalhanges during the process of interfacial activation, allowing sub-

trate molecules access to the active site (open conformation) [4–6].

The limitations of the industrial use of lipases have beenainly due to their high cost, which may be overcome by proper

∗ Corresponding author at: Institute of Chemistry, Federal University of Alfenas,7130-000 Alfenas, MG, Brazil. Tel.: +55 35 3299 1399.

E-mail addresses: adriano.mendes@unifal-mg.edu.br,driano amendes@yahoo.com.br (A.A. Mendes).

ttp://dx.doi.org/10.1016/j.ijbiomac.2014.06.035141-8130/© 2014 Elsevier B.V. All rights reserved.

immobilization techniques on solid supports. The immobiliza-tion facilitates the recovery and further reuse of the biocatalyst,avoids enzyme aggregation and autolysis and increases flexibil-ity of reactor design. Furthermore, additional stabilization of theimmobilized enzyme three-dimensional structure may be achievedif an increase in the rigidification of the macromolecule structure ispromoted [7–10]. Lipases have been immobilized by different pro-tocols as physical adsorption on hydrophobic and ionic exchangesupports, covalent attachment on highly activated supports andencapsulation in inorganic and organic supports [11–21]. Althoughthere are several methods to immobilize enzymes, the immobiliza-tion/stabilization by multipoint covalent attachment on activatedsupports presents some practical advantages when compared withother immobilization methods: the immobilization process canbe controlled and different supports can be tested without dif-ficulty. Furthermore, the enzyme molecules become more rigid,and thus more resistant to conformational changes induced byheat and organic solvents than the corresponding unmodified ones

[7–9,12,15–20,22–24]. A great increase in the stability of severalenzymes after their multipoint covalent attachment on glyoxyl-supports has been widely reported [7–9,12,15,20,23]. The stabilityfactor for each enzyme depends on the enzyme structure, on the

A.A. Mendes et al. / International Journal of Biological Macromolecules 70 (2014) 78–85 79

Table 1Catalytic properties of crude lipase extracts used in the present work.

Lipase Source organism Designation Supplier Protein (mg/g) Activityc (IU/g) Specific activity(IU/mg protein)

t1/2 at 70 ◦C (min)

TLL1a T. lanuginosus Lipolase® Novozymes 17.9 3422.5 191.2 4.8TLL2a T. lanuginosus Lipex® 100L Novozymes 23.7 4161.7 175.6 5.4PFLb P. fluorescens AK Amano 20.5 4459.0 217.5 2.5

a

b

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rLgaaTebifFbpfiop[sb

vetAbSah[bb

icotm

2

2

f((ut6sba

Liquid lipase preparation.Solid powder lipase preparation.Activity measured on the olive oil emulsion hydrolysis (pH 8.0, 37 ◦C).

umber of aldehyde groups generated in the support and on themmobilization conditions [7–10,15–18,20,22–24].

In this work, three lipase preparations from Pseudomonas fluo-escens (PFL) and Thermomyces lanuginosus (Lipolase® – TLL1 andipex® 100L – TLL2) were immobilized by covalent attachment onlyoxyl-agarose beads. The effect of the incubation time, activatinggent (glycidol or epiclorohydrin) and lipase source on the catalyticctivity and thermal-stability of the biocatalysts was investigated.he immobilization parameters were assayed in hydrolysis ofmulsified olive oil. The biocatalysts were also used to catalyzeutyl butyrate (pineapple flavor) synthesis by direct esterification

n heptane medium. Operational stability tests were performedor 5 successive cycles of 24 h each in butyl butyrate synthesis.atty acid ethyl ester (biodiesel) synthesis by transesterification ofabassu and palm oils with ethanol in solvent-free systems was alsoerformed. Recently, some studies have focused on the transesteri-cation of triglycerides with short-chain alcohols such as methanolr ethanol to produce biodiesel catalyzed by immobilized lipasesrepared by multipoint covalent attachment on glyoxyl-supports15–17]. However, the application of palm and babassu oils as feed-tocks in biodiesel production catalyzed by biocatalysts preparedy the above mentioned strategy has not been reported yet.

The immobilization of the lipases on glyoxyl-supports occursia nucleophilic attack of amine groups from Lys residues on thenzyme surfaces (pK of 10.5) to the aldehyde groups of the supportso form Schiff’s bases (C N double bonds) [7–9,12,16–18,22–24].fter immobilization, the biocatalysts prepared were then incu-ated in sodium borohydride solution for the reduction of thechiff’s bases, which transform them in stable covalent bonds,s well as change the reactive aldehyde groups to the inertydroxyl groups, an important step in the immobilization process7,12,16,24]. The representative scheme of preparation and immo-ilization of lipases on glyoxyl-supports, e.g. agarose beads, haseen previously reported [16].

Lipases from Pseudomonas fluorescens and Thermomyces lanug-nosus are able to form bimolecular aggregates at low enzymeoncentrations [20,25]. Thus, the covalent attachment of the lipasesn glyoxyl-agarose beads was carried out in the presence of Tri-on X-100 (0.15%, v/v) to obtain fully dispersed immobilized lipase

olecules oriented toward the reaction medium [16,19,20].

. Materials and methods

.1. Materials

Lipase preparations Lipolase® (TLL1) and Lipex® 100L (TLL2)rom Thermomyces lanuginosus were kindly donated by NovozymesAraucária, Brazil) and Lipase AK from Pseudomonas fluorescensPFL) was purchased from Amano Enzyme Inc. (Nagoya, Japan), andsed as received without further purification. The characteristics ofhese lipase preparations are presented in Table 1. Agarose beads

B-CL (SepharoseTM 6B-CL) was acquired from Amershan Bio-ciences (Uppsala, Sweden). Epichlorohydrin (Epi), glycidol (Gly),utyric acid, Triton X-100, Bradford reagent and bovine serumlbumin were purchased from Sigma–Aldrich Co. (St. Louis, USA).

Sodium borohydride, sodium periodate and anhydrous butanolwere obtained from Vetec (Sao Paulo, Brazil). Gum arabic wasacquired from Synth (São Paulo, Brazil). Olive oil (low acidity)from Carbonell (Spain) was purchased at a local market. Anhydrousethanol (minimum 99.5%, m/m) was supplied by Chromoline (SP,Brazil). Refined bleached palm oil was a kind gift from Agropalma(Belém, Brazil) and babassu oil kindly supplied by Pulcra (Jacareí,Brazil). The fatty acid composition of palm and babassu oils hasbeen previously reported [26].

2.2. Preparation of glyoxyl-agarose beads

Agarose beads were activated with glycidol and epichlorohydrinto produce glyoxyl-support. For glycidol activation, 10 g beads wereadded to a solution composed of 3 mL distilled water, 5 mL 1.7 MNaOH solution containing 0.15 g sodium borohydride (NaBH4). Fol-lowing this, 3.6 mL of glycidol was slowly added and the mixturekept at 0 ◦C for 15 h [27]. For epichlorohydrin activation, 10 g beadswere suspended in 100 mL 2 M NaOH solution containing 0.6 gNaBH4. Then, 10 mL of epichlorohydrin were slowly added and thesuspension was also kept at 0 ◦C for 15 h [28]. Glyceryl-agarose(prepared by activation with glycidol and epichlorohydrin) wassuspended in 60 mL Milli-Q water and then added 30 mL 100 mMsodium periodate solution to produce glyoxyl groups [27]. Thesuspensions were kept under slight stirring for 2 h at room temper-ature. Glyoxyl-agarose beads were thoroughly rinsed with Milli-Qwater and vacuum dried. The density of aldehyde groups for acti-vated agarose beads by glycidol was 95 �mol/g wet support [29].For the activation with epichlorohydrin, the densidy of aldehydeand epoxy groups was 102 and 6 �mol/g wet support, respectively[12].

2.3. Immobilization procedure

Immobilization of lipase preparations on glyoxyl-agarose wascarried out by adding 1.0 g of support to 9 mL of a solution atpH 10.05 (buffer sodium bicarbonate 100 mM) containing 0.15%(v/v) of Triton X-100 and 5 mg protein. The suspensions were keptunder mild stirring in an orbital shaker during at different time ofincubation varying from 4 to 72 h at room temperature. The immo-bilization was followed by measuring the hydrolytic activity andprotein concentration in the supernatant solution. The immobi-lization of TLL1 on glyoxyl-agarose was further tested by offeringdifferent loadings of protein (5.0, 10.0, 30.0, 60.0 and 80 mg/g ofsupport) to determine the support saturation enzymatic load. Afterthe enzyme immobilization step, 1.0 mg/mL sodium borohydridewas added to the immobilization suspension and kept under agita-

tion during 30 min at 25 ◦C. After this, the biocatalysts were filtered(Whatman filter paper 41) and thoroughly rinsed with 200 mMbuffer sodium phosphate pH 7.0 and finally washed thoroughlywith distilled and Milli-Q water.

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.4. Determination of hydrolytic activity

Hydrolytic activities of crude extract and immobilized lipaseere assayed using olive oil emulsion as substrate, according to

he methodology described by Soares et al. [30], with slight mod-fications. The substrate was prepared by mixing 50 g of olive oil

ith 150 g of gum Arabic solution (3%, m/v). The reaction mixtureontaining 5 mL of the emulsion, 5 mL of 100 mM buffer sodiumhosphate at pH 8.0 and soluble (0.2 mL, 0.5 mg/mL) or immobilized

ipase (0.1 g) was incubated for 5 min at 37 ◦C under continuous agi-ation in an orbital shaker (200 rpm). The reaction was stopped bydding 10 mL of commercial ethanol (minimum 95.0%, m/m). Theiberated fatty acids were titrated with 20 mM sodium hydroxideolution in the presence of phenolphthalein as indicator. The reac-ion blanks were made by adding immobilized biocatalysts afterhe ethanol. One international unit (IU) of activity was defined ashe amount of enzyme that is necessary to liberate 1 �mol of freeatty acid per minute under the conditions described above [21].

.5. Determination of protein

Protein was determined according to the methodologyescribed by Bradford [31] using bovine serum albumin astandard. All solutions were prepared using Milli-Q water.

.6. Immobilization parameters

Immobilization yield percentage (YI) was calculated by mea-uring the units of lipase (determined by the activity on oliveil hydrolysis) immobilized on glyoxyl-agarose. Specific activ-ty (SA) was calculated as the hydrolytic activity of biocatalystser milligram of immobilized protein (IU/mgIP). Recovery activityercentage (RA) was calculated after determining the hydrolyticctivity of immobilized enzyme and comparing with the numberf enzyme units that disappeared from the supernatant.

.7. Thermal stability

Soluble (0.1 mL) and immobilized lipase (0.1 g) were incubatedn the presence of 1 mL buffer sodium phosphate pH 8.0 (100 mM)t 70 ◦C for different time intervals [16,17]. Samples were taken inetermined interval times and immediately cooled in ice bath forhe interruption of inactivation reaction. The half-life time (t1/2)as determined by applying the exponential non-linear decayodel proposed by Sadana and Henley [32]. The experimental dataere analyzed using Origin Pro software, version 8.0 (OriginLaborporation, Northampton, USA). Stabilization factors (SF) werebtained as the ratio between the half-lives of the immobilizediocatalysts and the crude lipase extracts, as shown in Eq. (1).

=timmob1/2

tcrude1/2

(1)

.8. Butyl butyrate synthesis

Immobilized biocatalysts prepared by offering 5 mg protein perram of support (1.0 g) were incubated in 10 mL of heptane con-aining butanol and butyric acid at molar ratio 1:1 (100 mM of eacheactant). The reactions were performed at 37 ◦C for 24 h of incuba-ion under continuous stirring in an orbital shaker at 200 rpm [21].he ester conversion percentage was determined by measurements

f the concentration of residual butyric acid in the reaction mix-ure, as shown in Eq. (2). Samples were withdrawn (1 mL), dilutedn 10 mL of an ethanol/acetone 1:1 (v/v) mixture and titrated withaOH solution (20 mM) using phenolphthalein as indicator. The

ogical Macromolecules 70 (2014) 78–85

reaction blanks (substrate incubated in the presence of activatedagarose beads by Gly and Epi) were performed and no conversionwas observed at 24 h of incubation.

Conversion (%) =(

Ainitial − Afinal

Ainitial

)× 100 (2)

where Ainitial is the initial concentration and Afinal is the final con-centration of butyric acid in the reaction medium (mM).

2.9. Operational stability tests

Operational stability tests of immobilized TLL1 on glyoxyl-agarose prepared by activating with epoxyde agents (Gly and Epi)were performed in the synthesis of butyl butyrate in batch reactorsincubated at 37 ◦C, under the experimental conditions describedabove (Section 2.8). At the end of each cycle (five batch reactionsof 24 h each), immobilized TLL1 was then removed from the reac-tion medium and rinsed with ice heptane in excess to remove anysubstrate and product retained in the microenvironment of the bio-catalyst. After, the biocatalyst was introduced into a fresh medium.The reactions were periodically monitored by assessing the residualbutyric acid concentration.

2.10. Biodiesel synthesis

The reactions were performed in closed flasks with working vol-ume of 100 mL containing 20 g substrate consisting of babassu orpalm oils and anhydrous ethanol in solvent-free systems at fixedmolar ratio oil to alcohol 1:9 and 1:18, respectively, accordingmethodology previously reported [16,17,21]. The mixtures wereincubated with immobilized lipase at proportion of 2 mg immobi-lized protein per gram of oil. The experiments were carried out at45 ◦C by a maximum period of 48 h under continuous stirring in anorbital shaker at 250 rpm. For the time course studies, an aliquot ofreaction medium was taken at different time intervals and dilutedin hexane for determining fatty acid ethyl esters (biodiesel) by gaschromatograph (GC) analysis.

2.11. GC analysis

Fatty acid ethyl esters from babassu and palm oils were analyzedby gas chromatograph using a Varian CG 3800 model (Varian, Inc.Corporate Headquarters, Palo Alto, CA, USA) equipped with flameionization detector and 5% DEGS on Chromosorb WHP 80/100 mesh(6 ft, 2.0 mm ID) in stainless steel packed column (Restek, FrankelCommerce of Analytic Instruments Ltd., SP, Brazil). Nitrogen wasused as the carrier gas with a flow rate of 25 mL/min. Tempera-ture programming was performed. The column temperature waskept at 90 ◦C for 3 min, heated to 120 ◦C at 25 ◦C/min and keptconstant for 10 min. Then, the temperature was programmed at25 ◦C/min to 170 ◦C and kept constant for 15 min. The temperaturesof the injector and detector were set at 250 ◦C. Data collection andanalyses were performed using the software Galaxie Chromatog-raphy Data System version 1.9. Calibration curves were built fromstandard ethyl esters (ethyl caproate, caprate, laurate, myristate,palmitate, stearate, oleate, and linoleate) by using hexanol as inter-

nal standard. The transesterification yield was calculated by takinginto account the mass of ester content obtained by GC analysis andthe total theoretical ester mass based on the reaction molar ratio[33].

A.A. Mendes et al. / International Journal of Biological Macromolecules 70 (2014) 78–85 81

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ig. 1. Immobilization course of TLL1 (A), TLL2 (B) and PFL (C) on agarose beads puffer sodium bicarbonate pH 10.05 containing 0.15% (v/v) of Triton X-100 at room tonditions. The assays were conducted with biocatalysts prepared by offering 5 pro

. Results and discussion

.1. Influence of activation protocol, lipase source, and incubationime on the catalytic properties of the biocatalysts prepared

Fig. 1 shows the immobilization course of microbial lipases TLL1A), TLL2 (B) and PFL (C) on glyoxyl-agarose beads prepared byctivating with Gly and Epi. Firstly, it was verified the stabilityf the lipases under the immobilization conditions: incubation atH 10.05 buffer sodium bicarbonate by a maximum period of 72 h

n the presence of 0.15% (v/v) of Triton X-100. Under these con-itions, TLL1 and TLL2 were fully active and PFL retained around0% of its initial activity after 72 h of incubation. The progressf the immobilization of lipases was determined by measuringhe activity disappeared in the supernatant. According to Fig. 1,he immobilization of the lipases by offering protein loading of

mg/g of support was achieved at first 12–14 h of incubation.fter this period, a strong effect on the enzyme immobilizationas not observed. Maximum immobilization yield around 80% was

bserved in the immobilization of TLL2 on glyoxyl-agarose beads,ollowed by PFL (around 60%) and TLL1 (around 25–30%). Althoughoth TLL1 and TLL2 are lipase preparations from T. lanuginosus, the

atter presented immobilization yield around 3-fold higher thanLL1. The different performance of these two lipase preparationsould be attributed to different additives and preserving agentsresent in their formulations which may hindered the immobi-

ization of lipases onto solid supports [34] or that TLL2 may be variation of TLL1 obtained by genetic modification which has aumber higher of Lys residues in its structure than TLL1 that haseven Lys residues [3], or changes in the glycosilation of the enzymeurface.

The catalytic properties of the biocatalysts prepared by immo-ilizing lipases on glyoxyl-agarose beads are shown in Table 2.ccording to results, the activating agent did not exhert strong

nfluence on the catalytic properties of the biocatalysts. Theseesults could be attributed to similar density of reactive groupsn the support surface, as previously described in “Preparation oflyoxyl-agarose beads” (see Section 2.2). Furthermore, the acti-ation by both epoxyde agents (Gly and Epi) generates shortpace-arms with only two carbon atoms, providing aldehyderoups with similar reactivity [16]. These results are in agreement

ith previous studies reported for the immobilization of TLL1 and

FL on glyoxyl-supports such as Toyopearl AF-amino-650 M resinnd hybrid hydrogels chitosan/alginate chemically modified with,4,6 trinitrobenzesulfonic acid (TNBS) prepared by activation with

sly activated by Gly (full square) and Epi (open square) by incubating at 100 mMrature. Lipase extracts control (open circle) incubated under the same experimentalg/g of support.

these agents [16,17]. Conversely, a strong influence of the time ofimmobilization and lipase source on the catalytic properties of thebiocatalysts prepared was observed, i.e. hydrolytic activity, specificactivity, recovered activity percentage and thermal-stability. Max-imum hydrolytic activity varying from 230 to 250 IU/g of supportwas found for the biocatalysts prepared by immobilizing TLL1 andTLL2 on glyoxyl-agarose beads by 24 h of incubation. On the otherhand, the immobilization of PFL on glyoxyl-agarose beads renderedbiocatalysts with the lowest hydrolytic activity (around 50–60 IU/gof support).

Although TLL1 and TLL2 have presented similar hydrolytic activ-ity values, TLL1 rendered biocatalysts with lower immobilizationyield (by a 3-fold factor in relation to TLL2), as shown in Fig. 1. Forthis reason, the specific activity and recovered activity percentagevalues for TLL1 biocatalysts were higher than TLL2 (Table 2). It canbe seen in Table 1 that the specific activity values for TLL1 and TLL2extracts were 191.2 and 175.6 IU/mg protein, respectively. Afterimmobilization step, a drastic reduction of specific activity for TLL2by a 3-fold factor in relation to crude extract was observed (52.3and 54.2 IU/mgIP), while immobilized TLL1 by 4–24 h of incubationexhibited similar values in relation to the crude extract. Similarly,recovered activity percentage (ratio between apparent hydrolyticactivity and number of enzyme units theoretically immobilized)for the biocatalysts prepared by immobilizing TLL1 was drasticallyhigher than those ones prepared with TLL2 (recovered activity per-centage <40%). This means that, in this case, the immobilization ofTLL2 on glyoxyl-agarose beads prepared by activating with bothepoxyde agents strongly distorted its three-dimensional structure.

From data in Table 2, it is possible to verify that the immo-bilization of PFL on glyoxyl-agarose beads also distorted itsthree-dimensional structure rendering biocatalysts with the low-est catalytic activity. Crude PFL extract has specific activity of217.5 IU/mg protein, however after the immobilization on glyoxyl-agarose beads its specific activity was drastically reduced by a11-fold factor (around 20.0 IU/mgIP). Similarly, strong reduction ofrecovered activity percentage was also observed. These results arealso in accordance with the immobilization of the lipase by covalentattachment on highly activated supports such as hybrid hydrogelepoxy-chitosan/alginate and glyoxyl-Toyopearl AF-amino-650 Mresin [17,19].

The increase of immobilization time reduced the catalytic activ-

ity of biocatalysts prepared by immobilizing TLL1 and TLL2 asexpected, however no reduction was observed for PFL as shown inTable 2. Multipoint covalent attachment between enzyme and sup-port occurs in two steps. First, some bonds are quickly formed. Since

82 A.A. Mendes et al. / International Journal of Biological Macromolecules 70 (2014) 78–85

Table 2Catalytic properties of the biocatalysts prepared by covalent attachment on glyoxyl-agarose beads (protein loading – 5 mg/g of support).

Lipase (source) Activating agent Time (h) HAa (IU/g) SAb (IU/mgIP) RAc (%) t1/2d (h) SFe

TLL1

Gly

4 182.5 192.1 100 12.1 151.012 247.9 193.7 100 19.4 242.024 246.5 183.9 95.4 23.4 291.048 220.8 167.2 87.7 26.3 303.072 209.9 155.5 81.8 19.4 277.0

Epi

4 175.8 193.2 100 11.5 144.012 211.7 194.2 100 18.6 232.024 227.3 181.9 96.0 22.1 276.048 216.3 172.8 89.0 23.6 286.072 208.8 154.8 81.5 21.7 271.0

PFL

Gly24 66.0 18.7 8.60 0.33 7.8048 64.6 19.3 8.33 0.31 7.4072 58.8 17.3 8.40 0.29 6.90

Epi24 59.0 18.2 8.30 0.31 7.4048 58.8 19.6 8.43 0.37 8.7072 52.9 16.8 6.90 0.35 8.40

TLL2

Gly24 253.0 52.3 36.1 0.55 6.1048 235.2 49.2 33.5 0.60 6.7072 211.7 44.1 30.1 0.67 7.50

Epi24 244.0 54.2 34.7 0.58 6.5048 231.1 52.1 32.8 0.58 6.4072 217.0 48.5 30.8 0.81 9.00

a Hydrolytic activity.b Specific activity.c Recovered activity.

tsatg[si

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d Half-life.e Stabilization factor.

he support and the enzyme molecules are not complementarytructures, after the first bonds the enzyme loses their flexibilitynd the formation of new bonds may require longer immobiliza-ion times to allow a correct alignment between the nucleophilicroups of the enzyme and aldehyde groups on the support surface7–9,12,16–18,20,22–24]. This effect can also distort the enzymetructure and reduce its catalytic activity, however an increase ints thermal-stability may be expected [7,8,12,23].

Indeed, the results summarized in Table 2 indicated that anncrease of the immobilization time enhanced the thermal-stabilityor the biocatalysts prepared by immobilizing TLL1. At 4 h of incu-ation, TLL1 was around 150-fold more stable than crude extracthat presented half-life (t1/2) of 4.8 min at 70 ◦C, but increasing themmobilization time for 24–48 h a maximum stabilization around00-fold was reached. However, no influence of immobilizationime on the thermal-stability for PFL and TLL2 biocatalysts wasbserved (around 10-fold more stables than crude lipase extracts).imilarly, half-life (t1/2) values for TLL1 biocatalysts varied from 12o 26 h, whereas for TLL2 and PFL was observed a maximum valuef 0.9 h (Table 2). At 70 ◦C, crude lipase extracts from TLL2 and PFLxhibited half-lives of 5.4 and 2.5 min, respectively (see Table 1).hese results suggest that an intense multipoint covalent attach-ent occurred only for the biocatalysts prepared by immobilizing

LL1. Conversely, the results indicated that the immobilizationf PFL and TLL2 on glyoxyl-agarose beads was not successfullychieved by an intense multipoint covalent attachment. Fig. 2isplays the inactivation profiles of the biocatalysts prepared by

mmobilizing TLL1 at different incubation times on agarose beadsreviously activated with glycidol (Fig. 2A) and epichlorohydrinFig. 2B). The biocatalysts exhibited similar inactivation profiles,.e. after 28 h of incubation at 70 ◦C the biocatalysts prepared bymmobilizing TLL1 on glyoxyl-agarose at 24–48 h retained around5% of their initial activities.

Results from Table 2 showed that the multipoint covalentttachment of TLL1 on glyoxyl-agarose beads for 24 h of incuba-ion was enough to prepare highly active and stable biocatalysts.ence, the immobilization of lipases on glyoxyl-agarose performed

at 24 h of incubation was then selected for further studies concern-ing the determination of maximum immobilized protein amountand synthesis of esters in organic medium such as butyl butyrate(pineapple flavor) by direct esterification in heptane medium, andsynthesis of fatty acid ethyl esters (biodiesel) by transesterificationof vegetable oils in solvent-free systems.

3.2. Influence of protein loading on the catalytic properties ofimmobilized TLL1

The effect of protein loading on the catalytic properties ofimmobilized TLL1 on glyoxyl-agarose beads activated by bothagents (Gly and Epi) was investigated, as shown in Table 3. Theamount of immobilized TLL1 increased greatly with the offeredprotein loading and the concentration of immobilized proteinreached a maximum value around 20 mg/g of support by offer-ing 80 mg protein per gram of support. The hydrolytic activityalso increased proportionally to the immobilized protein amount,as expected. At highest protein loading offered (80 mg/g of sup-port), maximum hydrolytic activity for immobilized TLL1 variedfrom 1350 to 1470 IU/g of support by activation with glycidol andepichlorohydrin, respectively. A drastic reduction of the recov-ered activity percentage was observed due to possible reduction ofthe porous effective diameter caused by steric hindrances, whichreduces the diffusion of the substrate molecules (oil droplets) andproducts [16,17,19]. The maximum capacity of glyoxyl-agarosebeads to immobilize TLL1 was higher than other glyoxyl-supportstested in previous studies such as chemically modified hybridchitosan/alginate hydrogels and Toyopearl AF-amino-650 M resin[16,17].

3.3. Butyl butyrate synthesis and operational stability assays

The catalytic activity of immobilized biocatalysts prepared bycovalent attachment of lipases on glyoxyl-agarose was also inves-tigated in the synthesis of butyl butyrate (pineapple flavor) inheptane medium. Maximum ester conversion for immobilized TLL1

A.A. Mendes et al. / International Journal of Biological Macromolecules 70 (2014) 78–85 83

0 5 10 15 20 25 300,0

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ivity

Time (h)

0 5 10 15 20 25 300,0

0,2

0,4

0,6

0,8

1,0(B)

Res

idua

l act

ivity

Time (h)

F covalent attachment on agarose beads previously activated by Gly (A) and Epi (B) atd cle), 48 h (full square) and 72 h (full triangle). Inactivation tests were carried out at 70 ◦Cb

aEtahseaHeaoplvgw

tiTaftrc

543210

20

40

60

80

100

Con

vers

ion

(%)

Cycles

Fig. 3. Operational stability of immobilized TLL1 on agarose beads activated by Gly

TI

ig. 2. Thermal stability of TLL1 extract (open square) and immobilized lipase byifferent time of immobilization: 4 h (full circle), 12 h (open triangle), 24 h (open ciry incubating at 100 mM buffer sodium phosphate pH 8.0.

nd TLL2 on glyoxyl-agarose prepared by activating with Gly andpi was around 85–88% after 24 h of reaction. Butyl butyrate syn-hesis by immobilized PFL presented also similar conversion – 89nd 85% for activation with Gly and Epi, respectively. Although PFLas exhibited lower catalytic activity in aqueous reaction (emul-ified olive oil hydrolysis) than TLL1 and TLL2, both biocatalystsxhibited similar catalytic activity in esterification reaction, ingreement with previous studies carried out in our lab [16,19].owever, the biocatalysts prepared by immobilizing PFL and TLL2xhibited around 2- to 3-fold more immobilized enzyme than TLL1,s shown in Fig. 1. Based on these results, the direct esterificationf butyric acid and butanol catalyzed by TLL1 biocatalysts is moreromising from the industrial point of view because is required a

owest immobilized enzyme amount to exhibit similar ester con-ersion. Hence, the biocatalysts prepared by immobilizing TLL1 onlyoxyl-agarose beads activated by different protocols (Gly and Epi)ere then selected for operational stability tests.

The most important advantage of immobilization procedure ishe repeated use of the biocatalysts prepared. The catalyst reusabil-ty was carried out to determine the stability of the immobilizedLL1 on glyoxyl-agarose beads previously activated by both Glynd Epi. In this set of experiments, crude TLL1 extract was excluded

rom the study since it is very difficult to separate it from the reac-ion medium for reuse, while the immobilized enzyme can be easilyecovered and recycled for its reuse. According to Fig. 3, the bio-atalysts retained around 95% of their initial activities after five

(white) and Epi (light gray) in butyl butyrate synthesis. The reactions were per-formed in heptane medium at molar ratio butanol:butyric acid 1:1 (100 mM of eachreactant), 10% m/v biocatalysts, 37 ◦C under continuous stirring in an orbital shaker(200 rpm) for 24 h of reaction.

able 3nfluence of protein loading on the immobilization parameters of immobilized TLL1 on glyoxyl-agarose beads.

Activating agent Protein loading (mg/g of support) Parameters

HA (IU/g) IP (mg/g) SA (IU/mgIP) RA (%)

Gly 5 246.5 1.34 183.9 95.410 703.4 4.14 169.9 67.630 1246.6 12.7 98.2 39.160 1390.4 17.9 77.7 30.980 1347.3 21.2 63.5 25.4

Epi 5 227.3 1.25 181.9 96.010 560.6 3.89 144.1 57.430 1113.9 12.1 92.1 36.660 1335.5 17.0 78.5 31.380 1470.0 19.4 75.8 30.2

84 A.A. Mendes et al. / International Journal of Biological Macromolecules 70 (2014) 78–85

Gly-TLL1 Ep i-TLL 1 Gly-PF L Epi-PF L0

20

40

60

80

100(A)

Tran

sest

erifi

catio

n yi

eld

(%)

Biocatal ystsGly-T LL1 Epi-T LL1 Gly-P FL Epi-P FL

0

20

40

60

80

100(B)

Tran

sest

erifi

catio

n yi

eld

(%)

Biocatalysts

F obilizer 0 rpm

sbwer

3t

fAnciofintfb1htra

dAnrtwt9aoG9H

ig. 4. Transesterification yield for babassu (A) and palm (B) oils catalyzed by immeactions were performed at 40 ◦C under continuous stirring in an orbital shaker (25

uccessive cycles of 24 h each. Based on these results, the immo-ilization of TLL1 on glyoxyl-agarose beads prepared by activatingith epoxyde agents (Gly and Epi) seems to be an excellent strat-

gy to prepare highly stable and active biocatalysts in esterificationeaction.

.4. Production of fatty acid ethyl esters (biodiesel) byransesterification of vegetable oils in solvent-free systems

Biodiesel synthesis by transesterification reaction was per-ormed using immobilized TLL1 and PFL on glyoxyl-agarose beads.lthough the immobilization of PFL on glyoxyl-agarose beads didot result highly thermal-stable biocatalysts, it was also used toatalyze biodiesel synthesis because previous studies carried outn our lab report its high catalytic activity in transesterificationf vegetable oils [16,21]. In this set of experiments, transesteri-cation reactions performed by using crude lipase extracts wereot carried out because immobilized lipases have been consis-ently more active [21]. Babassu and palm oils were employed aseedstocks under conditions previously established, i.e. ethanol toabassu oil molar ratio of 9:1 and ethanol to palm molar ratio of8:1 [16,17,21]. Biodiesel synthesis by transesterification reactionas been performed in an excess alcohol to shift the equilibriumo the products side [35]. Additionally, some studies previouslyeported show that both TLL1 and PFL exhibit high ethanol toler-nce [16,17,21,36,37].

As can be seen in Fig. 4, the activating agent and lipase sourceid not exhibit strong influence on the transesterification yields.lthough the difference among the transesterification yields wasot considerable, a higher transesterification yield was found foreactions performed with palm oil. At 24 h of reaction, transes-erification yields for babassu oil varied from 75 to 85% (Fig. 4A),hereas for palm oil was above 90% (Fig. 4B). After 48 h of reac-

ion, maximum transesterification yield for babassu oil varied from0 to 94%, however the transesterification of palm oil reachedlmost full conversion (≥98.5%). According to specifications rec-

mmended by the Brazilian National Agency of Petroleum, Naturalas and Biofuels (ANP), a minimum transesterification yield of6.5% is required to be used as biofuel (Method EN 140103) [38].ence, only fatty acid ethyl esters (biodiesel) from palm oil are

d TLL1 and PFL on agarose beads activated by Gly (white) and Epi (light gray). The) for 24 h (white bars) and 48 h (light gray bars) of reaction in solvent-free systems.

in accordance with specifications recommended by the ANP forbiofuels.

From this point of view, immobilized TLL1 and PFL presentedsimilar apparent activities in transesterification reactions of Brazil-ian vegetable oils. However, TLL1 immobilized on glyoxyl-agarosebeads may be more attractive option for biotransformation reac-tions due to its high catalytic activity and thermal-stability in bothaqueous and organic media.

4. Conclusions

Agarose beads showed to be a very promising support for lipaseimmobilization. This support produced biocatalysts with high cat-alytic activity in both aqueous and organic media. Among them,immobilized TLL1 on glyoxyl-agarose exhibited high hydrolyticactivity and thermal-stability, around 300-fold more stable thanthe lipase crude extract after 24–48 h of incubation. These resultssuggested that the immobilization of TLL1 on glyoxyl-agarosebeads proceeded via multipoint covalent attachment. The bio-catalysts prepared were highly active in synthesis of butylbutyrate. In esterification reaction, immobilized TLL1 exhibitedhigh operational stability after five successive cycles. The high-est transesterification yield was reached for palm oil after 48 hof reaction, in which provided biodiesel samples in accordancewith specifications recommended by the ANP to be used as bio-fuel. Although the biocatalysts prepared have been highly active inesterification and transesterification reactions, immobilized TLL1was the most active and stable biocatalyst in both aqueous andorganic media. Multipoint covalent attachment of TLL1 on glyoxyl-agarose may be interesting from industrial point of view forapplication in esterification and transesterification reactions.

Acknowledgments

The authors gratefully acknowledge to FAPESP (Project

04/14593-4), CNPq and CAPES (Brazil) for financial supportAgropalma (Belém, PA, Brazil) for the donation of refined palm oiland Novozymes S.A. (Araucária, PR, Brazil) for the donation of lipasepreparations TLL1 and TLL2 used in this work.

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A.A. Mendes et al. / International Journal

eferences

[1] A.A. Mendes, P.C. Oliveira, H.F. Castro, J. Mol. Catal. B: Enzym. 78 (2012)119–134.

[2] D. Sharma, B. Sharma, A.K. Shukla, Biotechnology 10 (2011) 23–40.[3] R. Fernandez-Lafuente, J. Mol. Catal. B: Enzym. 62 (2010) 197–212.[4] L. Sarda, P. Desnuelle, Biochim. Biophys. Acta 30 (1951) 513–521.[5] R. Verger, Trends Biotechnol. 15 (1997) 32–38.[6] P. Reis, K. Holmberg, H. Watzke, M.E. Leser, R. Miller, Adv. Colloid Interface Sci.

147–148 (2009) 237–250.[7] C. Mateo, O. Abian, M. Bernedo, F. Cuenca, M. Fuentes, G. Fernández-Lorente,

J.M. Palomo, V. Grazú, B.C.C. Pessela, C. Giacomini, G. Irazoqui, A. Villarino, K.Ovsejevi, F. Batista-Viera, R. Fernández-Lafuente, J.M. Guisán, Enzyme Microb.Technol. 37 (2005) 456–462.

[8] C. Mateo, J.M. Palomo, M. Fuentes, L. Betancor, V. Grazú, F. López-Gallego, B.C.C.Pessela, A. Hidalgo, G. Fernández-Lorente, R. Fernández-Lafuente, J.M. Guisán,Enzyme Microb. Technol. 39 (2006) 274–280.

[9] C. Mateo, J.M. Palomo, G. Fernández-Lorente, J.M. Guisán, R. Fernández-Lafuente, Enzyme Microb. Technol. 40 (2007) 1451–1463.

10] R.C. Rodrigues, C. Ortiz, A. Berenguer-Múrcia, R. Torres, R. Fernández-Lafuente,Chem. Soc. Rev. 42 (2013) 6290–6307.

11] A.I.S. Brígida, A.D.T. Pinheiro, A.L.O. Ferreira, G.A.S. Pinto, L.R.B. Gonc alves, Appl.Biochem. Biotechnol. 137–140 (2007) 67–80.

12] H. Tümtürk, N. Karaca, G. Demirel, F. S ahin, Int. J. Biol. Macromol. 40 (2007)281–285.

13] A.A. Mendes, D.S. Rodrigues, M. Filice, R. Fernández-Lafuente, J.M. Guisán, J.M.Palomo, Tetrahedron 64 (2008) 10721–10727.

14] D.S. Rodrigues, A.A. Mendes, M. Filice, R. Fernández-Lafuente, J.M. Guisán, J.M.Palomo, J. Mol. Catal. B: Enzym. 58 (2009) 36–40.

15] R.C. Rodrigues, B.C.C. Pessela, G. Volpato, R. Fernández-Lafuente, J.M. Guisán,M.A.Z. Ayub, Process Biochem. 45 (2010) 1268–1273.

16] A.A. Mendes, R.C. Giordano, R.L.C. Giordano, H.F. Castro, J. Mol. Catal. B: Enzym.68 (2011) 109–115.

17] A.A. Mendes, H.F. Castro, D.S. Rodrigues, W.S. Adriano, P.W. Tardioli, E.J. Mam-marella, R.C. Giordano, R.L.C. Giordano, J. Ind. Microbiol. Biotechnol. 38 (2011)1055–1066.

18] J.A. Silva, G.P. Macedo, D.S. Rodrigues, R.L.C. Giordano, L.R.B. Gonc alves,Biochem. Eng. J. 60 (2012) 16–24.

[

[

ogical Macromolecules 70 (2014) 78–85 85

19] A.A. Mendes, H.F. Castro, G.S.S. Andrade, P.W. Tardioli, R.L.C. Giordano, React.Funct. Polym. 73 (2013) 160–167.

20] L.N. Lima, C.C. Aragon, C. Mateo, J.M. Palomo, R.L.C. Giordano, P.W. Tardioli, J.M.Guisán, G. Fernández-Lorente, Process Biochem. 48 (2013) 118–123.

21] A.A. Mendes, P.C. Oliveira, A.M. Vélez, R.C. Giordano, R.L.C. Giordano, H.F. Castro,Int. J. Biol. Macromol. 50 (2012) 503–511.

22] W.S. Adriano, D.B. Mendonc a, D.S. Rodrigues, E.J. Mammarella, R.L.C. Giordano,Biomacromolecules 9 (2008) 2170–2179.

23] J. Pedroche, M.M. Yust, C. Mateo, R. Fernández-Lafuente, J. Girón-Calle, M. Alaiz,J. Vioque, J.M. Guisán, F. Millán, Enzyme Microb. Technol. 40 (2007) 1160–1166.

24] A. Manrich, C.M.A. Galvão, C.D.F. Jesus, R.C. Giordano, R.L.C. Giordano, Int. J. Biol.Macromol. 43 (2008) 54–61.

25] J.M. Palomo, M. Fuentes, G. Fernández-Lorente, C. Mateo, J.M. Guisán, R.Fernández-Lafuente, Biomacromolecules 4 (2003) 1–6.

26] K.C. Santos, D.M.J. Cassimiro, M.H.M. Avelar, D.B. Hirata, H.F. Castro, R.Fernández-Lafuente, A.A. Mendes, Ind. Crops Prod. 49 (2013) 462–470.

27] J.M. Guisán, Enzyme Microb. Technol. 10 (1988) 375–382.28] M.M. Beppu, E.J. Arruda, R.S. Vieira, N.N. Santos, J. Membr. Sci. 240 (2004)

227–235.29] A.A. Mendes, L. Freitas, A.K.F. Carvalho, P.C. Oliveira, H.F. Castro, Enzyme Res.

2011 (2011) 1–8.30] C.M.F. Soares, H.F. Castro, G.M. Zanin, F.F. Moraes, Appl. Biochem. Biotechnol.

77/79 (1999) 745–757.31] M.M. Bradford, Anal. Biochem. 72 (1976) 248–254.32] A. Sadana, J.P. Henley, Biotechnol. Bioeng. 30 (1987) 717–723.33] D. Urioste, M.B.A. Castro, F.C. Biaggio, H.F. Castro, Quím. Nova 31 (2008)

407–412.34] N. López, R. Pérez, F. Vásquez, F. Valero, A. Sánchez, J. Chem. Technol. Biotechnol.

77 (2002) 175–182.35] J.M. Bernal, P. Lozano, E. García-Verdugo, M.I. Burguete, G. Sánchez-Gómez,

G. López-López, M. Pucheault, M. Vaultier, S.V. Luis, Molecules 17 (2012)8696–8719.

36] A. Salis, M. Pinna, M. Monduzzi, V. Solinas, J. Mol. Catal. B: Enzym. 54 (2008)

19–26.

37] A. Salis, M.S. Bhattacharyya, M. Monduzzi, V. Solinas, J. Mol. Catal. B: Enzym. 57(2009) 262–269.

38] J.S. de Sousa, E.A. Cavalcanti-Oliveira, D.A.G. Aranda, J. Mol. Catal. B: Enzym. 65(2010) 133–137.