B a m A J a 1 b a A R R A A K N Z N B B 1 o c r e r s r S h 0 Colloids and Surfaces B: Biointerfaces 165 (2018) 150–157 Contents lists available at ScienceDirect Colloids and Surfaces B: Biointerfaces jo ur nal ho me p ag e: www.elsev ier .com/ locate /co lsur fb iocatalysts based on nanozeolite-enzyme complexes: Effects of lkoxysilane surface functionalization and biofuel production using icroalgae lipids feedstock driano de Vasconcellosa, Alex Henrique Millera, Donato A.G. Arandab, osé Geraldo Nerya,∗ Laboratory for Clean Energy Technology (LACET), Physics Department, São Paulo State University–UNESP, Campus de São José do Rio Preto, SP, 5054-000, Brazil Greentec Laboratory, School of Chemistry, Federal University of Rio de Janeiro, RJ, 21941-972, Brazil r t i c l e i n f o rticle history: eceived 20 September 2017 eceived in revised form 16 January 2018 ccepted 13 February 2018 vailable online 14 February 2018 eywords: anozeolite surface chemical modulation eolite-enzyme surface interaction on-edible lipid feedstocks iofuels iomass a b s t r a c t Nanozeolites with different crystallographic structures (Nano/TS1, Nano/GIS, Nano/LTA, Nano/BEA, Nano/X, and Nano-X/Ni), functionalized with (3-aminopropyl)trimethoxysilane (APTMS) and crosslinked with glutaraldehyde (GA), were studied as solid supports for Thermomyces lanuginosus lipase (TLL) immobilization. Physicochemical characterizations of the surface-functionalized nanozeolites and nanozeolite-enzyme complexes were performed using XRD, SEM, AFM, ATR-FTIR, and zeta potential measurements. The experimental enzymatic activity results indicated that the nanozeolitic supports functionalized with APTMS and GA immobilized larger amounts of enzymes and provided higher enzymatic activities, compared to unfunctionalized supports. Correlations were observed among the nanozeolite surface charges, the enzyme immobilization efficiencies, and the biocatalyst activities. The catalytic performance and reusability of these enzyme-nanozeolite complexes were evaluated in the ethanolysis transesterification of microalgae oil to fatty acid ethyl esters (FAEEs). TLL immobilized on the nanozeolite supports functionalized with APTMS and GA provided the most efficient biocatalysis, with FAEEs yields above 93% and stability during five reaction cycles. Lower FAEEs yields and poorer cat- alytic stability were found for nanozeolite-enzyme complexes prepared only by physical adsorption. The findings indicated the viability of designing highly efficient biocatalysts for biofuel production by means of chemical modulation of nanozeolite surfaces. The high biocatalyst catalytic efficiency observed in ethanolysis reactions using a lipid feedstock that does not compete with food production is an advantage that should encourage the industrial application of these biocatalysts. © 2018 Elsevier B.V. All rights reserved. . Introduction The production of liquid biofuels from biomass is one of the ptions available to help meet the increasing energy demands aused by the life styles in industrialized countries and by the apid economic growth of populous developing countries. This high nergy demand currently mainly relies on fossil fuels, but envi- onmental concerns and the depletion of fossil fuel resources have timulated the search for alternative renewable fuels that are envi- onmentally benign and sustainable. Biodiesel can play a major role ∗ Corresponding author at: Rua Cristóvão Colombo 2265, São José do Rio Preto, P, 15054-000, Brazil. E-mail address: nery@ibilce.unesp.br (J.G. Nery). ttps://doi.org/10.1016/j.colsurfb.2018.02.029 927-7765/© 2018 Elsevier B.V. All rights reserved. in this challenge of finding a viable alternative for the replacement of the fossil fuels used in the transportation sector [1,2]. The task of searching for new sources of clean and renewable energy is not straightforward and needs to consider various factors including the economic viability of non-edible biomass sources, the development of new catalysts, and the use of low cost process- ing technologies [3]. First generation biodiesels are produced from biomass feedstocks consisting of food and oil crops, using conven- tional chemical technologies [4,5], and a large proportion (95%) of biodiesel production still relies on edible oil sources [6]. The use of edible feedstocks has raised many ethical and economic ques- tions, due to its negative impacts on global food markets and food security [7]. Alternative biomass sources such as non-food crops [8], animal fats [9,10], and waste cooking oils [11,12] are employed in the production of second generation biodiesel, but the use of https://doi.org/10.1016/j.colsurfb.2018.02.029 http://www.sciencedirect.com/science/journal/09277765 http://www.elsevier.com/locate/colsurfb http://crossmark.crossref.org/dialog/?doi=10.1016/j.colsurfb.2018.02.029&domain=pdf mailto:nery@ibilce.unesp.br https://doi.org/10.1016/j.colsurfb.2018.02.029 A. de Vasconcellos et al. / Colloids and Surfaces B: Biointerfaces 165 (2018) 150–157 151 Table 1 Synthesis parameters of the nanozeolite materials. Sample Silica Source Aluminum or Titanium Source Cation Type Synthesis time (h) Temperature (◦C) Ref. Nano-TS1 TEOS TBOT TPA+ 24 100 [45] Nano-GIS TEOS AlIso TMA+ 312 100 [46] Nano-BEA Silica Fumed TEA TEA+ 120 140 [47] Nano-LTA Ludox AlIso TMA+ 24 60 [48] Nano-X Silica Fumed Sodium aluminate Na+ 48 60 [49] T numis T ). t l g b t t p d e c l a a c [ g i t l s h c t [ t b t t c o t [ t ( o t w f i s o t o a a i f i e t EOS = tetraethylorthosilicate; TBOT = tetrabutylorthotitanate; AlIso = alumi PA+ = tetrapropylammonium; Ludox = aqueoussilicasolution (Ludox HS-30, 30 wt% hese feedstocks has drawbacks related to the processing costs in arge-scale commercial operations [13,14]. The third generation of biodiesel proposes the use of microal- ae as an alternative biomass feedstock [15]. This is considered y experts in the energy field as a technically feasible alterna- ive capable of overcoming the main difficulties associated with he feedstocks employed in first and second generation biodiesel roduction [16]. Heterotrophic and photoautotrophic microalgae are viable can- idates for biodiesel production, due to their high photosynthetic fficiency, rapid formation of large amounts of biomass, high lipid ontents that can be equivalent to that of soybean, and faster arge-scale growth compared to other energy crops [17,18]. In ddition, some heterotrophic microalgae can be cultivated in the bsence of light, using a non-photosynthetic process, while others an be produced during fermentation of a reduced carbon source 19]. For practical and economic reasons, heterotrophic microal- ae have several advantages over photoautotrophic organisms, ncluding faster biomass accumulation under controlled condi- ions that decrease the likelihood of common problems affecting arge-scale microalgae cultivation, such as variations in light inten- ity and the available carbon source [20]. Another advantage of eterotrophic microalgae is that genetically modified strains are apable of producing lipid yields higher than 80%, while pho- oautotrophic cultures of native algae yield only 20–50% of lipids 21,22]. Although heterotrophic microalgae are incapable of cap- uring CO2 emissions, the overall heterotrophic process is cyclic, ecause the initial organic substrates are produced by photosyn- hetic plants [23]. There are four different catalytic routes in triacylglycerides ransesterification reactions: (i) base-catalyzed processes, (ii) acid- atalyzed processes, (iii) enzyme-catalyzed processes, and (iv) use f supercritical conditions. The advantages and disadvantages of hese different routes have been discussed in several review papers 24,25]. Currently, industrial biodiesel production mainly employs he homogeneous base-catalyzed process, using sodium hydroxide NaOH), potassium hydroxide (KOH), sodium methoxide (NaOCH3), r sodium ethoxide (NaOCH2CH3) as the catalyst [26]. However, here are several drawbacks associated with this process, especially hen low quality feedstocks with high contents of water and free atty acids (FFAs) are used. In this case, the use of base catalysts ncreases the energy cost and the likelihood of soap and emul- ion formation, which can lead to the generation of large amounts f wastewater during the processes of cleaning and separation of he glycerol (byproduct) and biodiesel (product) [26,27]. Hence, in rder to overcome these drawbacks, alternative and more sustain- ble catalytic routes for biodiesel production are being sought. The use of lipases as catalysts in biodiesel production is an ttractive option, due to their higher specificity for the transester- fication of triacylglycerides to fatty acid methyl esters (FAMEs) or atty acid ethyl esters (FAEEs), compared to the conventional chem- cal catalysts employed in industrial biodiesel production. Enzymes xhibit greater selectivity and higher catalytic activity in transes- erification reactions under mild operational conditions (25–60 ◦C), opropoxide; TMA+ = tetramethylammonium; TEA+ = tetraethylammonium, compared to homogeneous basic catalysts [28]. In addition, lipases can catalyze the transesterification of raw materials that contain high levels of FAAs and water, with decreased risk of soap and emulsion formation [28]. However, despite all the advantages of the enzymatic process, there are important issues that hinder its use in industrial scale processes for biodiesel production. These include: a) the high costs of enzymes, b) problems in separating the product from the reaction medium, c) difficulty in recover- ing and reusing enzymes, and d) deactivation of the active sites of enzymes by the substrates (short-chain alcohols such as methanol and ethanol) [29] and by the glycerol byproduct [30]. One way to overcome these obstacles is to immobilize the enzymes on solid carriers [31,32]. The selection of suitable solid supports needs to consider aspects including the overall enzymatic activity of the immobilized enzyme, the cost of the immobilization procedure, identification of the best operating conditions in order to avoid enzyme inactivation, and the ability to regenerate and reuse the enzyme-support complex [33]. Zeolites [34,35] and enzymes [36,37] are capable of catalyzing the transesterification of triacylglycerides into FAEEs or FAMEs. These different classes of materials have inherent characteristics that determine the yield of the desired final product under dif- ferent experimental conditions [38]. Nonetheless, the combination of these different classes of catalysts can result in zeolite-enzyme complexes that offer outstanding catalytic performance [38]. Nanozeolites are hydrophobic supports with high external sur- face areas, whose high dispersibility in both aqueous solutions and organic media allows better access of the enzymes to the substrate. This also acts to reduce the adsorption of glycerol molecules onto the biocatalyst microenvironment, and electrostatic and hydropho- bic interactions within the zeolite-enzyme system contribute to its stability. Several mechanisms have been proposed for the immobi- lization of lipase on zeolitic supports: a) electrostatic and acid-base linkages [38], b) formation of strong enzyme-support ionic inter- actions [39–41], and c) covalent bonding of lipases to the zeolite surface by functionalization of the zeolite surface with alkoxysi- lane, followed by crosslinking with glutaraldehyde [42]. Among the available immobilization methods, covalent attachment is the most effective in terms of enhancing uptake of the enzyme and its reten- tion on the solid support. The covalent bonding of enzymes to solid matrices can improve the feasibility of using enzymes in indus- trial applications, considering aspects such as denaturation of the enzyme by heat or organic solvents, pH control of the medium, stability during storage, and reduction of leaching [43]. Studies of biocatalysts based on nanozeolite-lipase complexes for the purpose of biodiesel production have not been widely reported in the literature. Transesterification of palm oil to FAEEs using lipases of Thermomyces lanuginosus (TLL) and Rhizomucor miehei (RML) immobilized on nanosized NaX zeolite (FAU) ion- exchanged with different transition metals has shown positive results. This has encouraged further synthetic and catalytic stud- ies of enzyme-nanozeolite complexes with supports derived from different zeolite structures, and their application in the catalytic conversion of different non-edible biomasses to biodiesel [44]. Nev- 1 urface e b n a w n i a m l L c e n r r t o a t 2 2 m w o ( t i ( p o w p f 2 G T a 0 [ B H p s f [ w H u s N n 2 u w 52 A. de Vasconcellos et al. / Colloids and S rtheless, careful analysis of the literature shows that there have een no systematic studies describing the chemical modulation of anozeolite support surfaces using chemical modifiers such as the lkoxysilane molecule (3-aminopropyl)trimethoxysilane (APTMS), ith crosslinking using glutaraldehyde (GA), and the effects on the anozeolite-enzyme complexes. In order to address this lack of nformation, the present research work was undertaken with the im of answering the following questions: a) What are the enzy- atic and catalytic performances of Thermomyces lanuginosus (TLL) ipases immobilized on different nanosized zeolites (TS1, GIS, BEA, TA, NaX, and NaX-Ni) previously functionalized with APTMS and rosslinked with GA? b) Are these biocatalysts effective in the trans- sterification of oil from the microalga Prototheca moriformis (a on-edible lipid feedstock source) into FAEEs using the ethanolysis oute? c) Do the functionalized biocatalysts provide better catalytic esults than unfunctionalized biocatalysts, especially in terms of he FAEEs yield and biocatalyst stability? The experimental results btained in the search to answer these questions are presented nd discussed here, with evaluation of the potential applications of hese biocatalysts in the biofuels industry. . Experimental section .1. Materials Flavorless high-oleic (>80%) oil extracted from the genetically odified heterotrophic algal strain Prototheca moriformis S2532 as obtained from Solazyme (Campinas, São Paulo, Brazil). The il consisted of triglycerides (95%), small amounts of diglycerides <5%), trace amounts of monoglycerides (<0.5%), and small quanti- ies of fatty acid (1%). Ethanol (99.8%), phosphate buffer, gum arabic, norganic salts, glutaraldehyde, (3-aminopropyl)trimethoxysilane H2N(CH2)3Si(OCH3)3) (APTMS), organic bases, and enzymes were urchased from Sigma-Aldrich and were used as received, with- ut further purification. The lipase enzyme employed in this work as Lipolase 100 L from T. lanuginosus. This enzyme consisted of urified 1,3-specific lipases (EC 3.1.1.3) produced by submerged ermentation of a genetically modified strain of Aspergillus oryzae. .2. Synthesis of nanozeolites The conditions used in the syntheses of the nanozeolites (TS1, IS, LTA, BEA, and NaX) prepared in this study are summarized in able 1. The final gel molar compositions of the nanozeolites were s follows: TS1 (0.36 TPAOH: 0.06TiO2: 1.00SiO2: 16.2H2O: 4 EtOH: .24 BuOH) [45]; GIS (1 Al2O3: 4.17 SiO2: 2.39 TMA2O: 253 H2O) 46]; BEA (0.36 TPAOH: 0.06TiO2: 1.00SiO2: 16.2H2O: 4 EtOH: 0.24 uOH) [47]; LTA (0.4 Na2O:1.9 Al2O3: 14.0 (TMA)2O: 11.9 SiO2:700 2O) [48]; NaX (5.5 Na2O:1.0 Al2O3:4.0 SiO2:190 H2O) [49]. In a typical synthesis procedure, an aluminosilicate gel was pre- ared by mixing together freshly prepared aluminate and silicate olutions (or titanate and silicate solutions, in the case of TS1), ollowing experimental procedures adapted from the literature 45–49]. The nanocrystals were cooled to room temperature and ere then recovered by centrifugation (Hitachi Koki himac CR22N igh-Speed Refrigerated Centrifuge), washed with deionized water ntil reaching pH <8, and dried at room temperature for 24 h. These amples were denoted U-Nano/TS1, U-Nano/GIS, U-Nano/LTA, U- ano/BEA, and U-Nano-X (where U stands for unfunctionalized anozeolite). .3. Ion exchange of the nanozeolitic supports Prior to enzyme immobilization, the U-Nano-X/Na material was sed in ion exchange experiments in which the sodium cations ere replaced with nickel ions. These experiments were only s B: Biointerfaces 165 (2018) 150–157 performed with U-Nano-X/Na, because some of the other nanoze- olites presented low aluminum contents (see the results of the ICP-AES analyses of U-Nano/TS1 and U-Nano/BEA),while others (U- Nano/LTA and U-Nano/GIS) did not retain their structures under the ion exchange conditions employed. The procedure was car- ried out as follows: 1 g of U-Nano-X/Na was transferred to a Teflon-lined digestion reactor (Parr Instruments), together with 30 mL of 0.5 mol L−1 NiSO4 solution. The ion exchange reaction was allowed to proceed for 72 h at 60 ◦C, followed by centrifuga- tion of the mixture at 13,400g, washing of the solid product three times with distilled water, and drying at room temperature. This ion-exchanged derivative was designated U-Nano-X/Ni. 2.4. Functionalization of the nanozeolitic supports 2.4.1. Functionalization of the nanozeolite surfaces with APTMS Prior to the functionalization, all the as-synthesized nanoze- olites (U-Nano/TS1, U-Nano/GIS, U-Nano/LTA, and U-Nano/BEA) were thermally treated in order to remove the organic templates used in their syntheses. The samples were heated in an air stream at 550 ◦C, using the following temperature ramps: 25–100 ◦C (1 h), 100–150 ◦C (30 min), 150–200 ◦C (30 min), and 200–550 ◦C (3 h). The samples were kept at the final temperature of 550 ◦C for 3 h. The functionalization reactions with APTMS were performed according to the method described by Plueddemann et al. [50] and updated by Li et al. [51]. The procedure involved three steps: (1) stir- ring a mixture of dichloromethane (30 mL) + APTMS (2 mL) + zeolite (1 g) for 16 h at room temperature; (2) centrifugation at 13,400xg and washing the solid samples three times with methanol to remove the unreacted APTMS; (3) drying the modified zeolite for 12 h at 60 ◦C under vacuum. These alkoxysilane functionalized nanozeolites were denoted N-Nano/TS1, N-Nano/GIS, N-Nano/LTA, N-Nano/BEA, N-Nano/X, and N-Nano-X/Ni (where N stands for the APTMS functionalized nanozeolite). 2.4.2. Crosslinking of the APTMS functionalized nanozeolites with GA The APTMS functionalized nanozeolites were crosslinked with glutaraldehyde (GA) as described in the literature [52]. A typical experiment was performed as follows: 1 g of the APTMS- nanozeolite was mixed with 30 mL of an aqueous 2.5 wt.% GA solution and stirred for 24 h at room temperature. The suspen- sion turned yellow immediately after mixing, then orange, and finally red. The GA-functionalized nanozeolite was collected by centrifugation (13,400g) and the solid product was washed three times with distilled water, dried at room temperature, and stored under vacuum in order to prevent oxidation. There covered sam- ples were denoted NG-Nano/TS1, NG-Nano/GIS, NG-Nano/LTA, NG-Nano/BEA, NG-Nano/X, and NG-Nano-X/Ni (where G stands for the APTMS-functionalized nanozeolite crosslinked with glu- taraldehyde). Experimental details of the lipase immobilization on the nanozeolitic supports and determination of the enzymatic hydrolytic activities of the immobilized enzymes (measured using a titrimetric method) are provided in the Supplementary materials section of this journal. The nanozeolite-enzyme complexes were named according to the nomenclatures of their preceding functionalized deriva- tives and the as-synthesized nanozeolites. The nanozeolite-enzyme complexes derived from T. lanuginosus lipase were designated U-Nano/TS1-TLL, U-Nano/GIS-TLL, U-Nano/LTA-TLL, U-Nano/BEA- TLL, U-Nano/X-TLL, and U-Nano-X/Ni-TLL. The nanozeolite- enzyme complexes derived from the APTMS-functionalized supports were designated N-Nano/TS1-TLL, N-Nano/GIS-TLL, N-Nano/LTA-TLL, N-Nano/BEA-TLL, N-Nano/X-TLL, and N-Nano- X/Ni-TLL. The nanozeolite-enzyme complexes derived from the nanozeolitic supports crosslinked with glutaraldehyde were des- A. de Vasconcellos et al. / Colloids and Surfaces B: Biointerfaces 165 (2018) 150–157 153 i N 2 n u M r a C p n u t s e 6 E p t S p o m ( a t s ( T d g g F S Table 2 Chemical compositions of nanozeolitic supports obtained by ICP-OES. Sample SiO2 (%) Al2O3 (%) Na2O (%) TiO2(%) NiO(%) Nano-TS1 81.65 – <0.01 1.10 – Nano-GIS 42.73 16.60 1.27 – – Nano-LTA 37.42 24.98 11.69 – – Fig. 1. XRD patterns of the as made nanozeolites. gnated NG-Nano/TS1-TLL, NG-Nano/GIS-TLL, NG-Nano/LTA-TLL, G-Nano/BEA-TLL, NG-Nano/X-TLL,and NG-Nano-X/Ni-TLL. .5. Physicochemical characterization of the as-synthesized anozeolites and the nanozeolite-enzyme complexes The prepared materials described above were characterized sing XRD and SEM. The XRD analyses were performed with a iniFlex II instrument (Rigaku, Tokyo, Japan) equipped with a otating anode source with flat-plate Bragg-Brentano geometry and graphite monochromator, operating at 40 kV and 40 mA, with u K� radiation (wavelength = 1.5418 Å). The powder diffraction atterns were recorded in the 2� range from 3 to 50◦, with scan- ing at a goniometer rate of 2◦ min−1. SEM images were recorded sing an XL30 FEG instrument (FEI/Philips), with deposition of a hin coating of gold onto the samples prior to the analyses. FTIR pectra were acquired using a PerkinElmer Frontier FTIR spectrom- ter equipped with an ATR accessory. The samples were scanned 4 times between 4000 and 400 cm−1, at a resolution of 4 cm−1. lemental chemical analyses of the nanozeolitic supports were erformed using inductively coupled plasma atomic emission spec- roscopy (ICP-AES) at the Chemical Analysis Laboratory Facility of ao Paulo University. Atomic force microscopy (AFM) analyses were erformed at the Brazilian National Laboratory of Nanotechnol- gy (LNNano, Campinas), using a Dimension 3000 scanning probe icroscope (SPM) equipped with a NanoScopeIIIa SMP controller Digital Instruments Inc.). TESP tapping mode AFM images were cquired in ambient air using etched silicon probes. Zeta poten- ial measurements were performed with a Zetasizer Nano ZS90 ystem (Malvern Instruments, Worcestershire, U.K.), using 1–2% by weight) suspensions of the nanozeolites in deionized water. he samples were sonicated for 30 min before being transferred to isposable zeta potential cells for the measurements. Chromato- raphic analyses were performed with a PerkinElmer Clarus 580 as chromatograph equipped with a flame ionization detector (GC- ID). Experimental details of these analyses are provided in the upplementary material section of this journal. Nano-BEA 81.92 3.69 <0.03 – – Nano-X/Na 33.23 20.84 13.97 – – Nano-X/Ni 18.23 12.83 0.12 – 14.88 2.6. Syntheses of FAEEs The transesterification reactions of the high-oleic heterotrophic microalgae oil to biodiesel were performed with three different cat- alyst groups: (i) the as-synthesized nanozeolite-enzyme complexes (U-Nano/TS1-TLL, U-Nano/GIS-TLL, U-Nano/LTA-TLL, U-Nano/BEA- TLL, U-Nano/X-TLL, and U-Nano-X/Ni-TLL); (ii) the nanozeolite- enzyme complexes derived from the amino-functionalized sup- ports (N-Nano/TS1-TLL, N-Nano/GIS-TLL, N-Nano/LTA-TLL, N- Nano/BEA-TLL, N-Nano/X-TLL, and N-Nano-X/Ni-TLL); and (iii) the nanozeolite-enzyme complexes derived from the glutaraldehyde- activated supports (NG-Nano/TS1-TLL, NG-Nano/GIS-TLL, NG- Nano/LTA-TLL, NG-Nano/BEA-TLL, NG-Nano/X-TLL,and NG-Nano- X/Ni-TLL). Several experiments were performed in order to establish appropriate reaction conditions that would allow comparison among the performances of the three catalyst groups. Transesteri- fication reactions catalyzed by the different nanozeolite-enzyme complexes were performed at 40 ◦C, using an oil:ethanol ratio of 1:5, with slow addition of ethanol in order to avoid inactiva- tion of the catalysts. After a reaction time of 48 h, the product (biodiesel) and byproduct (glycerol) were separated by centrifu- gation at 13,400g. The progress of the transesterification reactions was followed using thin layer chromatography (TLC), according to the procedure described by Yang et al. [53]. At predetermined time intervals, a small volume (100 �L) of the reaction mixture was col- lected and mixed with 500 �L of hexane for 2 min. After separation by centrifugation, 3 �L of the upper layer was applied to a silica gel plate. A solution of hexane/ethyl acetate/acetic acid (90:10:1) was used as the developing solvent and iodine was used as the color reagent. 3. Results and discussion The XRD results (Fig. 1) revealed successful synthesis of Nano/TS1, Nano/GIS, Nano/LTA, Nano/BEA, Nano-X/Na, and Nano- X/Na ion-exchanged with nickel (Nano-X/Ni). In the case of Nano/TS1, most of the original Bragg reflections matched the XRD pattern reported for the MFI structure [45,54]. The XRD pattern of as-synthesized Nano/GIS with the Bragg reflections at 2� of 12.4◦, 21.7◦, and 27.2◦ corresponds to the typical reflections of a tetra- hedral GIS zeolite [46]. The XRD patterns of zeolites LTA and BEA (Fig. 1) indicated that the materials were obtained as pure crys- talline phases [47,48,55]. The XRD data for the Nano-X/Na zeolite were also indicative of a pure zeolite with faujasite topology, with- out the presence of organic templates or structure-directing agents (SDAs) [49,56]. However, ion exchange of the as-synthesized Nano- X/Na zeolite with nickel (producing Nano-X/Ni) resulted in a drastic change in the crystallographic structure (Fig. 1). The chemical compositions of the as-synthesized nanozeolites were determined by ICP-AES analyses and are summarized in Table 2. The SiO2: TiO2 ratio of 74.2 obtained for Nano/TS1 was in agreement with the results reported in previous studies [45,54]. The SiO2:Al2O3 ratios observed for the aluminosilicates Nano/BEA (22.2), Nano/GIS (2.57), Nano/LTA (1.49), and Nano/X (1.59) also agreed with the ratios reported elsewhere [46–48,56]. The high 154 A. de Vasconcellos et al. / Colloids and Surfaces B: Biointerfaces 165 (2018) 150–157 Fig. 2. Microscopy data for the U-Nano-BEA,SEM (a) and AFM (b). surfac S n a t e a I p s o a t T a a s A g p Fig. 3. The FTIR spectra of as made nanozeolite, amine-terminated iO2:Al2O3 ratio for Nano/BEA was indicative of the predomi- ance of Si O Si linkages, while the lower values for Nano/LTA nd Nano/X reflected higher amounts of aluminum in the crys- allographic structures, which had direct effects in the nickel ion xchange experiments. The SEM and AFM data for Nano/TS1, Nano/GIS, and Nano/LTA, re shown in the Supplementary materials section of this journal. n general TS1, GIS, and LTA nanozeolites presented spherical mor- hology, with sizes in the range 50–180 nm and. the homogeneous hapes and diameters of the nanocrystals were in accordance with ther results reported in the literature [45,46,48]. The morphology of Nano/BEA (Fig. 2) was characterized by gglomerates of nanocrystals, typical of nanozeolite beta [47], with he SEM and AFM images revealing crystals around 50 nm in size. he SEM and AFM images of Nano-X/Na (Supplementary materi- ls section) showed the presence of large nanoparticle aggregates pproximately 250 nm in size, composed of smaller particles with izes in the range 20–100 nm [49,56]. In contrast, the SEM and FM images for Nano-X/Ni showed the presence of crystal aggre- ates with no specific morphology, indicating that the ion exchange rocess damaged the original Nano/X structure. e nanozeolite and glutaraldehyde amino crosslinking nanozeolite. ATR-FTIR spectra of the as-synthesized and functionalized nanozeolites were measured in the range 400–4000 cm−1. Accord- ing to the literature, the infrared spectroscopy spectra for aluminosilicate zeolites can be divided into two distinct regions, corresponding to the skeletal IR spectrum(at 1600–500 cm−1) and the surface hydroxyl groups (at 4000–1500 cm−1) [57]. Analyses of the skeletal regions (1600–500 cm−1) of the ATR- FTIR spectra of the unfunctionalized U-Nano/TS1, U-Nano/GIS, U-Nano/LTA, U-Nano/BEA, and U-Nano-X/Na zeolites (see Supple- mentary materials section) showed that the absorption bands for the as-synthesized nanozeolite structures were in agreement with the values reported in the literature. However, for U-Nano-X/Ni, the absorption bands in the skeletal region (at 425, 552, 795, 912, and 1064 cm−1) were shifted from the positions expected for alu- minosilicates. This was a clear indication that the ion exchange treatment affected the original Nano-X/Na framework, corroborat- ing the XRD results. The ATR-FTIR spectra of the functionalized nanozeolites (Fig. 3) −1 revealed two absorption bands in the region 1350–1460 cm , which could be attributed to the bending of (CH) R CH3 and the methyl groups ( CH3) of glutaraldehyde [58]. Absorption bands at 1530 and 1750 cm−1 corresponded to the stretching of C O A. de Vasconcellos et al. / Colloids and Surfaces B: Biointerfaces 165 (2018) 150–157 155 urface nanozeolite and glutaraldehyde amino crosslinking nanozeolite. a b a t n m o b d b 3 m i s s z s t a ( N t t i i f ( t N N ( N l z i e w p e Table 3 The amount of Thermomyces lanuginosus lipase immobilized on the zeolitic supports, their enzymatic activity after immobilization and the transesterifi- cation reactions of microalgae oil to produce biodiesel. U-(unfunctionalized support), N-(APTMS funcionalized support), NG-(APTMS-Glutaraldehyde function- alized support). Nanozeolite Immobilization (%) Enzymatic activity (U/mg-support) Ethyl esters (%) U-Nano-TS1 31.9 15.6 12.9 N-Nano-TS1 97.6 48.0 56.6 NG-Nano-TS1 98.5 56.0 55.6 U-Nano-GIS 15.2 4.0 7.2 N-Nano-GIS 30.2 56.0 48.1 NG-Nano-GIS 79.1 28.0 60.3 U-Nano-LTA 18.4 8.0 7.7 N-Nano-LTA 97.1 68.0 89.8 NG-Nano-LTA 100 46.0 44.9 U-Nano-BEA 16.9 3.6 21.6 N-Nano-BEA 100 76.4 92.7 NG-Nano-BEA 100 68.0 67.7 U-Nano-X 18.3 2.0 17.8 N-Nano-X 76.6 74.0 94.6 NG-Nano-X 98.0 56.0 71.6 U-Nano-X/Ni 43.7 64.0 94.0 Fig. 4. Zeta potential for as made nanozeolite, amine-terminated s nd C N bonds, evidencing crosslinking of the GA group, while ands at 2950 and 2880 cm−1 corresponded to aldehyde C H and lkyl C H stretching vibrations, respectively. Since these absorp- ion bands were not observed in the spectra of the as-synthesized anozeolites, this provided strong evidence for the effective surface odification of the nanozeolite surfaces [52,59–62]. Absorption bands related to the APTMS, such as that for (C N) f the aminopropyl group, typically observed at wavelengths etween 1000 and 1200 cm−1, could not be unequivocally assigned, ue to the overlap of T O T bond absorption bands. Similarly, ands corresponding to stretching of amino group N H, at around 300 cm−1, overlapped with the strong absorption bands of water olecules and silanol groups in the region 3000–3600 cm−1 [63]. Electrostatic interaction plays a significant role in enzyme mmobilization. Therefore, in order to gain a better under- tanding of the role of organic modification of the nanozeolite urface and its influence on the overall immobilization process, eta potential determinations were performed for the as- ynthesized nanozeolites, the APTMS-functionalized nanozeolites, he APTMS/GA-functionalized nanozeolites, and the APTMS/Lipase nd APTMS/GA/Lipase complexes. Negative zeta potential values were found for U-Nano/TS1 −34.4 mV), U-Nano/GIS (−39.8 mV), U-Nano/LTA (−49.8 mV), U- ano/BEA (−25.1 mV), and U-Nano-X/Na (−38.2 mV) (Fig. 4). Since he Thermomyces lanuginosus enzyme is also negatively charged, he repulsive forces resulted in significantly smaller amounts of mmobilized enzymes, compared to the nanozeolites functional- zed with APTMS and GA (Table 3) [58,64]. The zeta potentials or the nanozeolites functionalized with APTMS and APTMS/GA Fig. 4) revealed that the surfaces of these zeolite were posi- ively charged: N-Nano/TS1 (19.6 mV), N-Nano/GIS (3.8 mV), N- ano/LTA (17.6 mV), N-Nano/BEA (24.1 mV), N-Nano/X (21.6 mV), -Nano/X/Ni (21.6 mV), NG-Nano/TS1(30.2 mV), NG-Nano/GIS 25.8 mV), NG-Nano/LTA (23.7 mV), NG-Nano/BEA (37.8 mV), G-Nano/X (28.2 mV), and NG-Nano-X/Ni (45.6 mV). After immobi- ization, all the nanozeolite-enzyme complexes presented negative eta potential values (Supplementary materials section). The pos- tively charged surfaces affected the amounts of immobilized nzyme, and it was also observed that the enzymatic activities ere higher for enzyme immobilized on the functionalized sup- orts (Table 3). A possible explanation for this enhancement of nzymatic activity could be related to the generation of spacer arms N-Nano-X/Ni 95.6 76.0 93.4 NG-Nano-X/Ni 97.7 78.0 93.0 between the enzyme and the supports, leading to an increase in the immobilized enzyme concentration promoted by the reactive organic chemical groups of the APTMS and GA molecules. According to the proposed mechanism, enzyme immobilization occurred at a greater distance from the support microenvironment, which could help to avoid distortions of the enzyme structure and enhance exposure of the active sites to the substrate, hence facil- itating enzyme/substrate contact and optimizing mass transfer in the system [65–70]. Systematic catalytic studies of the transesterification reac- tions of microalgae oil to produce biodiesel using the different nanozeolite-enzyme complexes were performed using the fol- lowing supports: a) unfunctionalized supports (U-Nano/TS1-TLL, U-Nano/GIS-TLL, U-Nano/LTA-TLL, U-Nano/BEA-TLL, U-Nano/X- TLL, and U-Nano-X/Ni-TLL); b) amine-terminated surface supports (N-Nano/TS1-TLL, N-Nano/GIS-TLL, N-Nano/LTA-TLL, N-Nano/BEA- TLL, N-Nano/X-TLL, and N-Nano-X/Ni-TLL); and c) glutaraldehyde crosslinked amine-terminated surface supports (NG-Nano/TS1- 156 A. de Vasconcellos et al. / Colloids and Surface F c T N t ( o T s l F 4 U c N N T l t m r e s w e t l ( c e s n b b t d w t p y p ig. 5. Turnover for the U-Nano-X/Ni-TLL,N-Nano-X/Ni-TLL and NG-Nano-X/Ni-TLL omplexes. LL, NG-Nano/GIS-TLL, NG-Nano/LTA-TLL, NG-Nano/BEA-TLL, NG- ano/X-TLL, and NG-Nano-X/Ni-TLL). The FAEEs yields of all he reactions were determined by gas chromatography analysis Table 3). The experimental reaction conditions were standardized in rder to compare the performances of the three catalyst groups. he FAEEs yields obtained for all the functionalized nanozeolitic upports were superior to those for the unfunctionalized zeo- ites (Table 3). For N-Nano/TS1-TLL and NG-Nano/TS1-TLL, the AEEs yields (56.6 and 55.6%, respectively) were approximately -fold higher than the yield obtained using the unfunctionalized -Nano/TS1-TLL support (12.9%) (Table 3). The FAEEs yields achieved with the nanozeolite-enzyme omplexes N-Nano/LTA-TLL (89.8%), N-Nano/BEA-TLL (92.7%),and -Nano/X-TLL (94.6%) were higher than obtainedwith the NG- ano/LTA-TLL (44.9%), NG-Nano/BEA-TLL (67.5%),and NG-Nano-X- LL (71.6%) complexes (Table 3). A possible explanation for the ower FAEEs yields observed for the NG supports was distortion of he enzyme structure caused by reticulation of the glutaraldehyde olecules with the enzyme amino groups, which consequently educed access of the substrate to the specific catalytic site of the nzyme [42,43]. One exception was observed among the Nano/GIS derivative upports. The FAEEs yield for the N-Nano/GIS-TLL complex (48.1%) as lower than for the NG-Nano/GIS-TLL complex (60.3%). Possible xplanations for this include low dispersibility of the biocatalyst in he reaction solution, or the low amount of the enzyme immobi- ized on this support (30.2%). High FAEEs yields were observed for the U-Nano-X/Ni-TLL 94.0%), N-Nano-X/Ni-TLL (93.4%), and NG-Nano-X/Ni-TLL (93.0%) omplexes (Table 3). A graphical representation of all the trans- sterification results is provided in the Supplementary materials ection. Reuse experiments were therefore performed using these anozeolite-enzyme complexes (Fig. 5). The important role played y nickel in enzyme immobilization and enzymatic activity has een reported previously [44,71] and was also observed here for he nanozeolite supports ion-exchanged with nickel. The FAEEs yield obtained with the U-Nano-X/Ni-TLL complex ecreased from 98 to 56.2% after five cycles of reuse in the reaction, hile the N-Nano-X/Ni-TLL complex showed a sharp decrease of he FAEEs yield from 93.4 to 26.7%. The NG-Nano-X/Ni-TLL complex resented the smallest decrease (from 93 to 85.1%) in the FAEEs ield. The highest decrease observed for N-Nano-X/Ni-TLL could in rinciple be attributed to the weak interactions between the APTMS s B: Biointerfaces 165 (2018) 150–157 amino groups and the enzyme, which could lead to the leaching or desorption of large amounts of the lipase from the solid during the successive reuses. The excellent performance shown by NG-Nano-X/Ni-TLL in the reuse experiments (Fig. 5) demonstrated the potential of the com- plex as a suitable biocatalyst for biofuel production reactions in long-term continuous systems, due to the fact that the immobilized enzyme was covalently bonded to the support, resulting in a more stable biocatalyst. The chemical functionalization of the Nano-X/Ni surfaces with APTMS and glutaraldehyde enhanced the hydropho- bicity of the biocatalyst, hence decreasing glycerol adsorption in the biocatalyst microenvironment and avoiding deactivation of the enzyme active sites [42,43]. 4. Conclusions • Nanozeolite surfaces chemically functionalized with APTMS and the glutaraldehyde crosslinking agent were efficient solid matri- ces for T. lanuginosus lipase immobilization; • Functionalization of the nanozeolite surfaces enhanced the amount of immobilized enzyme and increased the enzymatic activity, compared to unfunctionalized nanozeolitic supports; • Nanozeolite-enzyme complexes prepared with supports that had been previously chemically functionalized were more efficient biocatalysts in the transesterification reaction of microalgae oil to produce biodiesel, achieving high FAEEs yields; • The catalytic performances of biocatalysts prepared with unfunc- tionalized nanozeolitic supports were very poor, compared to the use of functionalized supports; • Although all the nanozeolite-enzyme complexes (U-Nanozeolite, N-Nanozeolite, and NG-Nanozeolite) were able to catalyze the transesterification of microalgae oil into biodiesel (FAEEs), the best catalytic performances were obtained with the complexes produced using nanozeolitic supports previously ion-exchanged with nickel (U-Nano-X/Ni-TLL, N-Nano-X/Ni-TLL, and NG-Nano- X/Ni-TLL). • In reuse experiments with the biocatalysts, the best results were obtained with NG-Nano-X/Ni-TLL, where the enzyme was cova- lently bonded to the nanozeolitic support. The NG-Nano-X/Ni-TLL complex showed excellent potential for use as a suitable bio- catalyst for biofuel production in long-term continuous system reactions. Acknowledgments Financial support for this ongoing research project is provided by the São Paulo State Research Foundation (FAPESP), in the form of a scientific award to JGN (grant #11/51851-5), and by CNPq (grants #465594/2014-0 and #406761/2013-2). AV (grant #11/10092-4) and AHM (grant #2016/24303-0) thank FAPESP for fellowships. We thank Dr. Marcia Cabrera for allowing us to use the zeta poten- tial instrument (FAPESP grant #2012/24259-0). We appreciate the assistance of Dr. Carlos A. O. Ramirez, Dr. Carlos A. Costa, and Dr. Evandro Lanzoni (Brazilian National Laboratory of Nanotechnology – LNNano) in the SEM-FEG and AFM microscopy experiments. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at https://doi.org/10.1016/j.colsurfb.2018.02. 029. 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