1 Biodiesel production from crude Jatropha oil catalyzed by non- 2 commercial immobilized heterologous Rhizopus oryzae and Carica 3 papaya lipases 4 5 J. Rodrigues(a), A. Canet(b), I. Rivera(c), N.M. Osório(a), G. Sandoval(c), F. 6 Valero(b), S. Ferreira-Dias(a)* 7 (a) 8 Portugal; 9 (b) Instituto Superior de Agronomia, Universidade de Lisboa, LEAF, Lisbon, Departament d’Enginyeria Quimica, Biològica i Ambiental (EE), Universitat 10 Autònoma de Barcelona, Barcelona, Spain; 11 (c) 12 Jalisco (CIATEJ), Guadalajara, Jalisco, Mexico. n o i s r e v d Centro de Investigación y Asistencia en Tecnología y Diseño del Estado de 13 e t ep c c A 14 * 15 Suzana Ferreira-Dias 16 Instituto Superior de Agronomia, Tapada da Ajuda. 1349-017 Lisbon, Portugal 17 E-mail: suzanafdias@mail.telepac.pt Corresponding Author: 18 19 Abstract 20 The aim of this study was to evaluate the feasibility of biodiesel production by 21 transesterification of Jatropha oil with methanol, catalyzed by non-commercial 22 sn-1,3-regioselective lipases. Using these lipases, fatty acid methyl esters 23 (FAME) and monoacylglycerols are produced, avoiding the formation of glycerol This is the author's version of a work that was accepted for publication in Bioresource technology (Ed. Elsevier). Changes resulting from the publishing process, such as peer review, editing, corrections, structural formatting, and other quality control mechanisms may not be reflected in this document. Changes may 1 have been made to this work since it was submitted for publication. A definitive version was subsequently published in Rodrigues, J. et al. “Biodiesel production from crude Jatropha oil catalyzed by non-commercial immobilized heterologous Rhizopus oryzae and Carica papaya lipases” in Bioresource technology, vol. 213 (Aug. 2016), p. 88-95. DOI 10.1016/j.biortech.2016.03.011 24 as byproduct. Heterologous Rhizopus oryzae lipase (rROL) immobilized on 25 different synthetic resins and Carica papaya lipase (rCPL) immobilized on 26 Lewatit VP OC1600 were tested. Reactions were performed at 30ºC, with seven 27 stepwise methanol additions. 28 For all biocatalysts, 51-65 % FAME (theoretical maximum= 66%) was obtained 29 after 4 h transesterification. Stability tests were performed in 8 or 10 successive 30 4 h-batches, either with or without rehydration of the biocatalyst between each 31 two consecutive batches. Activity loss was much faster when biocatalysts were 32 rehydrated. For rROL, half-life times varied from 16 to 579 h. rROL on Lewatit 33 VPOC 1600 was more stable than for rCPL on the same support. n o i s r 34 e v d 35 Keywords: Biodiesel; Carica papaya lipase; Jatropha oil; Rhizopus oryzae 36 lipase; sn-1,3 regioselective lipase. 37 e t ep c c A 38 1. Introduction 39 Biofuels are a renewable alternative to fossil fuels that has lower greenhouse 40 gas emissions. Several biofuel crops can be grown locally (including in marginal 41 soils), helping countries to reduce their dependence on unstable foreign 42 sources of fossil fuels. These potential environmental and social advantages of 43 biofuels have led to some policy measures to support sustainable production. 44 For instance, the Renewable Energy Directive (European Directive, 45 2009/28/E.C, 2009) forces EU Member States to achieve a minimum target of 46 10 % renewable energy in all the energy used in the transport sector by 2020 2 47 and a 7 % limit on food crop based biofuels. The fact that more than 95% of 48 biodiesel production feedstocks come from edible oils, causes great concern 49 because of the competition with the food supply chain. Consequently, there is 50 now an increased interest in second generation biofuel crops, such as Jatropha 51 curcas L., whose high oil content (27-45 % dry basis) is not suitable for 52 consumption, because of the presence of toxic components (Makkar et al., 53 1998). 54 The most usual method in industry to transform oil into biodiesel is alkaline- 55 catalyzed transesterification. However, this method has some disadvantages: 56 uses large amounts of energy, the glycerol produced has low quality resulting in 57 difficult and high-cost recovery and purification; alkaline catalyst is inactivated 58 and removed by washing leading to the production of large amounts of alkaline 59 effluents that must be treated. In addition, the free fatty acids present in the oil 60 will form soaps by direct esterification with the catalyst (e.g. sodium hydroxide 61 or sodium methoxide) leading to a lower biodiesel yield. 62 Lipases (triacylglycerol acylhydrolases; EC 3.1.1.3) are enzymes that, besides 63 hydrolysis reaction, catalyze various synthetic reactions, including 64 transesterification, when in low water activity media (Casas et al., 2012; 65 Ferreira-Dias et al., 2013). The use of lipases as biocatalysts for biodiesel 66 production has become more appealing, since lipases can act in mild 67 temperature conditions, resulting in lower energy consumption, and with a wide 68 diversity of raw materials, such as waste oils and fats with high levels of free 69 fatty acids (FFA) and traces of water (Fan et al., 2012). Also, biodiesel recovery 70 is easier since no emulsions are formed, less unit operations are needed and 71 only small amounts of wastewater are produced. Furthermore, due to the high n o i s r e v d e t ep c c A 3 72 selectivity of lipases, side-reactions with the formation of undesirable products, 73 as well as soap formation occurring in alkaline-catalysis, are avoided, resulting 74 in easier and environmentally friendly separation and purification processes 75 (Juan et al., 2011). 76 The main reasons why lipases are not yet widely used in the industry are their 77 cost and longer reaction time compared with alkaline catalysts. An essential 78 strategy to lower the cost of the enzymatic process is the multiple reuse of the 79 biocatalyst or its use in continuous bioreactors, which can be achieved by using 80 immobilized enzymes. These biocatalysts must present both high 81 transesterification activity and operational stability. 82 Lipase denaturation and inhibition by methanol (or ethanol) is currently 83 observed during lipase-catalyzed transesterification. However, this problem can 84 be overcome by stepwise addition of the alcohol along the reaction (Canet et 85 al., 2014; Duarte et al., 2015; Kuo et al. 2015; Lotti et al., 2015; You et al 86 2013).Glycerol, the main byproduct of transesterification reaction, is one of the 87 constraints for lipase-catalyzed transesterification efficacy. It adsorbs onto 88 enzyme immobilization carriers, causing lipase deactivation and lowering the 89 process efficiency (Hama et al., 2011). The use of sn-1,3-regioselective lipases 90 to synthesize biodiesel and monoacylglycerols (MAG) simultaneously, avoiding 91 the generation of glycerol, could be a solution for this problem (Calero et al., 92 2015; Canet et al, 2014; Verdugo et al., 2010). The MAG obtained can be used 93 as emulsifiers in food, pharmaceutical and cosmetic industries. 94 In recent years, low-cost alternatives to commercial lipases have been 95 developed in order to reduce process costs. The non-commercial heterologous 96 Rhizopus oryzae lipase (rROL) has been produced and successfully used by n o i s r e v d e t ep c c A 4 97 our group as catalyst for lipid restructuring (Nunes et al., 2011, 2012a; 2012b; 98 Simões et al., 2014; Tecelão et al., 2012b), for the production of bile acids or 99 corticoesteroid derivatives for pharmaceuticals applications (Quintana et al., 100 2012, 2015) and also for biodiesel production (Bonet-Ragel et al., 2015; Canet 101 et al., 2014, 2016; Duarte et al., 2015). This recombinant lipase is a promising 102 new biocatalyst for biodiesel production that showed a 44-fold higher specific 103 activity compared to a commercially available lipase obtained directly from R. 104 oryzae, and a higher specificity towards the p-nitrophenol ester of long chain 105 length (Guillén et al, 2011). 106 Carica papaya lipase (CPL) is a naturally self-immobilized biocatalyst, since it is 107 attached to Carica papaya L. latex polymeric matrix. This low-cost biocatalyst 108 was also successfully used by us for enantioresolution (Rivera et al., 2013) and 109 the production of biopolymers (Sandoval et al., 2010), waxes (Quintana et al., 110 2011) and human milk fat substitutes (Tecelão et al., 2012a). Efforts have been 111 made to isolate CPL unsuccessfully. Heterologous expression of this protein 112 shows as an alternative to overcome this problem (Rivera et al., 2013). 113 This study aims to: (i) produce Jatropha biodiesel (FAME) and 114 monoacylglycerols (MAG), by transesterification of Jatropha crude oil with 115 methanol, in stirred batch reactor and solvent-free media, catalyzed by non- 116 commercial sn-1,3 regioselective lipases, from microbial and plant origins, 117 immobilized in different supports and (ii) select the best immobilized biocatalyst 118 for Jatropha biodiesel production in terms of activity and operational stability. 119 The non-commercial heterologous Rhizopus oryzae lipase (rROL), immobilized 120 on Lewatit VPOC 1600, IRA-96, LifetechTM ECR1030M, LifetechTM ECR8285M, n o i s r e v d e t ep c c A 5 121 LifetechTM AP1090M; and the recombinant Carica papaya lipase (rCPL) 122 immobilized on Lewatit VPOC 1600 were tested as biocatalysts. 123 124 2. Materials and Methods 125 2.1 Lipases 126 Two different non-commercial sn-1,3 regioselective lipases were tested: (i) the 127 heterologous Rhizopus oryzae lipase (rROL), produced by the Bioprocess 128 Engineering and Applied Biocatalysis group of the Universitat Autonoma de 129 Barcelona (UAB), Barcelona, Spain, and (ii) the heterologous Carica papaya 130 lipase (rCPL), produced by the group of Centro de Investigación y Asistencia en 131 Tecnología y Diseño del Estado de Jalisco (CIATEJ), Guadalajara, Mexico. 132 rROL was produced by over-expression of the corresponding gene in a mutant 133 strain of Pichia pastoris, according to Arnau et al. (2010) and Guillén et al. 134 (2011). The rROL used in this study presented a hydrolytic activity of 178.8 135 U/mg of total protein according to the methodology developed by Resina et al. 136 (2004). 137 rCPL, expressed extracellularly in Pichia pastoris, was produced by submerged 138 fermentation in a rich medium at 30°C. Then, the culture broth was centrifuged 139 and the supernatant was tested for lipase activity. Immobilized rCPL presents a 140 hydrolytic activity of 30 U/mg of total protein. n o i s r e v d e t ep c c A 141 142 2.2 Carriers 6 143 rCPL was immobilized by adsorption on Lewatit VPOC 1600. rROL was 144 immobilized by adsorption on different carriers: (i) Lewatit VP OC 1600, donated 145 by Lanxess, Germany; (ii) LifetechTM ECR1030M and (iii) LifetechTM 146 ECR1090M, gifts from Purolite. Also, rROL was immobilized by covalent binding 147 on: (iv) LifetechTM ECR8285M, kindly donated by Purolite, Wales, U.K; and (v) 148 Amberlite IRA 96, from Rhom and Haas, Philadelphia, USA. The main physical 149 and chemical properties of these five lipase immobilization carriers are 150 presented in Table 1. 151 n o i s r 152 2.3 Jatropha oil extraction and characterization 153 Jatropha curcas L. seeds were collected from healthy and ripened fruits 154 harvested in central Mozambique, in Sofala province (19º56’S; 34º24’E). The 155 whole seeds (not dehulled) were crushed with a hammer and the fraction 156 smaller than 2 mm diameter was mechanically extracted in a screw press, Täby 157 Press type 20 (Skeppsta Maskin AB, Sweden), as previously described 158 (Rodrigues et al., 2016). 159 The acidity (% of free fatty acids, FFA) of Jatropha oil was 3.7 %, and was 160 determined by titration, according to ISO standard 660:2009. This oil has 41.1 161 % of oleic acid, 38.8 % of linoleic acid and 11.6 % of palmitic acid, as major 162 fatty acids (Rodrigues et al., 2015). e v d e t ep c c A 163 164 2.4 Methods 165 2.4.1 rROL and rCPL immobilization on Lewatit VP OC 1600 by adsorption 7 166 rROL immobilization on Lewatit VP OC 1600 was carried out, as previously 167 described by Tecelão et al. (2012b), by mixing the lipase powder with the carrier 168 in 50 mL of 0.1 M phosphate buffer solution, at room temperature for 18 hours. 169 The ratio lipase powder:support had been previously optimized for rROL 170 (Tecelão et al., 2012b) and corresponds to 0.25 g of rROL (85.1 ± 10.5 mg of 171 protein) per gram of Lewatit VP OC 1600. The beads were recovered by 172 vacuum filtration and incubated, under gentle stirring with 25 mL of 2.5 % (v/v) 173 glutaraldehyde solution, for 2 h at room temperature. The liquid phase was 174 filtered and collected in order to determine protein content and evaluate 175 immobilization yield. The immobilized lipase was rinsed twice with 50 mL of 176 immobilization buffer solution, in order to remove the free enzyme and the liquid 177 phase was collected for subsequent analysis. Beads were dried under reduced 178 pressure for 10 minutes, transferred into a suitable container and kept 179 refrigerated at 5 ºC until use. 180 rCPL was immobilized by direct adsorption on Lewatit VP OC 1600 (Sigma- 181 Aldrich, Mexico) at 4°C, using 22 mg of total protein per g of support, without 182 any subsequent treatment with glutaraldehyde. One of the reasons to select this 183 hydrophobic support is because the natural support of papaya latex is also 184 hydrophobic and can be used for lipase selective adsorption. n o i s r e v d e t ep c c A 185 186 2.4.2 rROL immobilization on LifetechTM AP1090M and LifetechTM 187 ECR1030M by adsorption 8 188 The immobilization of rROL on Purolite ECR resins was performed based on 189 the Purolite Application Guide - Purolite ECR Enzyme Immobilization 190 Procedures, with some modifications. 191 For lipase immobilization on LifetechTM AP1090M and LifetechTM ECR1030M 192 macroporous styrenes, the resin was previously equilibrated by washing it with 193 phosphate buffer solution (pH = 7, 0.05 M) and then filtered. A resin/buffer ratio 194 of 1/1 v/v was used. 195 The lipase (0.25 g of rROL per gram of wet resin) was dissolved in buffer 196 solution in a ratio of 1/4 (w/v) resin/buffer. Then, Purolite ECR resins were 197 added to the lipase solution and the mixture was gently stirred with the resin for 198 24 hours at room temperature. After, the liquid phase was filtered and collected, 199 in order to determine the protein content in the liquid and evaluate the 200 immobilization yield. The immobilized enzyme was washed with the 201 immobilization buffer (ratio resin/buffer of 1/1, w/v), the immobilized lipase was 202 filtered under reduced pressure and kept refrigerated at 5 ºC. n o i s r e v d e t ep c c A 203 204 2.4.3. rROL immobilization on LifetechTM ECR8285M epoxy acrylate by 205 covalent binding 206 The resin equilibration was carried out following the same procedure described 207 for LifetechTM AP1090M and LifetechTM ECR1030M resins. rROL was also 208 dissolved in immobilization buffer in a ratio of 0.25 g of rROL per gram of wet 209 resin and the ratio resin/buffer of 1/4 (w/v) was also used. 9 210 The mixture of lipase solution with the Purolite LifetechTM ECR8285M resin was 211 placed under gentle stirring at room temperature for 18 hours. After, the stirring 212 was stopped and the solution was left static for another 20 h. Then, the liquid 213 phase was filtered and collected for subsequent analysis, and the immobilized 214 lipase was washed once with buffer. The immobilized enzyme was kept 215 refrigerated at 5 ºC. 216 217 2.4.4 rROL immobilization on AmberliteTM IRA 96 by covalent binding 218 The methodology used for immobilizing rROL on anion exchange resin 219 Amberlite™ IRA96 is based on the method described by Wang et al. (2010) 220 with some modifications, as follows: 5 g of AmberiteTM IRA 96 were added to 50 221 mL of deionized water and put under gentle stirring for 30 minutes at 50 ºC. The 222 support was washed three times with 25 mL of NaOH aqueous solution 1 M, 223 alternating with 25 mL of HCl aqueous solution 1 M. The anion exchange resin 224 was then equilibrated by immersion in 100 mL of sodium phosphate buffer 225 solution 0.2 M (pH = 7.5). After, AmberliteTM IRA 96 was mixed together with 226 rROL dissolved in 10 mL of sodium phosphate buffer solution 0.2 M (pH = 7.5), 227 at room temperature and under magnetic stirring for 4 h. The ratio lipase 228 powder: support used was also 0.25 g of rROL per gram of AmberliteTM IRA 96. 229 After, particles were filtered under reduced pressure and then brought into 230 contact with 0.5 % (v/v) glutaraldehyde aqueous solution using 25 mL of 231 glutaraldehyde solution per gram of support. The immobilized lipase was rinsed 232 three times with 15 mL of immobilization phosphate buffer solution. Beads were 233 dried in a desiccator, transferred into a suitable container and stored at 5 ºC. n o i s r e v d e t ep c c A 10 234 235 2.4.5 Protein assay 236 The method described by Bradford (1976) was used to determine the total 237 amount of protein immobilized on the resins, using bovine serum albumin from 238 Sigma-Aldrich, Saint Louis, USA, as a standard. The immobilization yield was 239 defined as the difference between protein amount in the initial lipase solution 240 (before the immobilization support was added), and the residual protein present 241 in the supernatant after immobilization (as well as in the subsequent washing 242 solutions), divided by the protein content in the initial lipase solution. n o i s r 243 e v d 244 2.4.5. Time-course transesterification reactions catalyzed by rROL and 245 CPL 246 Transesterification reactions were carried out in 25 mL cylindrical glass reactors 247 for 48 h, at 30°C, and under magnetic stirring. Reaction conditions, were the 248 same as previously optimized by Canet et al. (2014), for biodiesel production 249 from olive oil by rROL immobilized in octadecyl-Sepabeads: 4% (w/w) water 250 content in the reaction medium, substrate molar ratio (methanol:Jatropha oil) of 251 3:1 and seven methanol additions. A load of 5% (w/w) of biocatalyst in relation 252 to the amount of Jatropha oil (10 g) was used. Samples were taken before 253 every methanol addition (after 0, 30, 60, 90, 120, 150 and 180 min) and at the 254 end of the reaction, and stored at -18°C for subsequent analyses. e t ep c c A 255 256 2.4.6 Operational Stability Tests 11 257 The operational stability of the immobilized rROL on different resins or rCPL on 258 Lewatit VPOC 1600 was evaluated during consecutive 4 h batches, carried out 259 under the same reaction conditions of time-course transesterification 260 experiments (c.f. 2.4.5.). At the term of each batch, the biocatalyst was removed 261 from the reaction medium by vacuum filtration. After, it was (i) immediately 262 added into fresh reaction medium and reutilized in the next batch (total of 10 263 batches) or (ii) rehydrated with 10 mL of 0.1 M sodium phosphate buffer 264 solution (pH 7.0), filtered under reduced pressure, added into fresh medium and 265 used in the subsequent batch (total of 8 batches). Samples were collected at 266 the end of each batch and stored at -18 ° C until further analysis. 267 It was considered that each biocatalyst has 100 % of its original activity, at the 268 end of the first batch. In order to describe the deactivation kinetics of 269 biocatalysts, each experimental point (FAME yield), at the end of each batch n, 270 was converted into the fraction of the original activity, i.e. its residual activity. 271 The residual activity (Ares, %) after each reuse was calculated as the ratio 272 between FAME yield of batch n, divided by FAME yield observed in the first 273 batch, and multiplied by 100. The fit of lipase deactivation models to the 274 experimental data was performed using “solver”, a tool included in Microsoft 275 Excel for Windows, by minimizing the sum of squares of errors between the 276 experimental data and those estimated by the respective model. The following 277 deactivation models, first order exponential decay (eq. 1) and two-component 278 first order exponential decay (eq 2), were tested: n o i s r e v d e t ep c c A 279 280 (Eq. 1) = 12 281 282 Ares = a e-k1n + b e-k2n 283 where, k, k1 and k2 are deactivation coefficients (n-1). 284 The kinetic constants were obtained by non-linear regression analysis for the 285 tested models. Also, the operational half-life of the biocatalyst (t1/2), i.e. the time 286 after which the activity of the biocatalyst is reduced to 50 %, was estimated by 287 the deactivation model fitted to the experimental results. (Eq. 2) 288 n o i s r 289 2.4.7 Analysis of reaction products 290 With the purpose of monitoring the transesterification reaction kinetics, the 291 determination of MAG, diacylglycerols (DAG), triacylglycerols (TAG) and FAME 292 contents was carried out for each sample, based on the European standard EN 293 14105: 2011, with some modifications. This European standard refers only to 294 the detection of trace amounts of glycerol, MAG, DAG and TAG in purified 295 biodiesel (FAME). Therefore, it was necessary to adapt the methodology to be 296 able to follow the transesterification kinetics. 297 The preparation of the samples was carried out according to Faustino et al. 298 (2015). Samples were derivatized with N-methyl-N-trimethylsilyl-tri- 299 fluoroacetamide (MSTFA) to convert the -OH groups to -OSi (Me)3 groups 300 The sample analysis was performed on a GC Agilent Technologies 7820A, 301 equipped with an on-column injector and a flame ionization detector. The 302 capillary column used for sample analysis was a J & W DB - 5HT (15 m x 0.32 e v d e t ep c c A 13 303 mm x 0.10 mm). The main operating conditions of the equipment were the 304 same used by Faustino et al. (2015) in a study about the production of human 305 milk fat substitutes, by acidolysis of tripalmitin with camelina oil FFA, catalyzed 306 by rROL. All compounds with retention times equal or higher than 25 min were 307 considered as TAG; DAG and MAG were assumed as the compounds with 308 retention times between 22 and 25 min or 17.8 and 21 min, respectively. FAME 309 presented retention times between 11 and 17 min. 310 Calibration curves for methyl oleate (retention time of 12.7 min) and triolein 311 (retention time of 35.2 min) were established, in order to quantify each group of 312 compounds (%, w/w), using mononodecanoin as internal standard (Mono C19; 313 retention time of 19.4 min).The masses of partial acylglycerols (MAG and DAG) 314 were calculated using the equations from the European standard EN 315 14105:2011. FAME yield (%) was defined as the ratio between the amount of 316 methyl esters formed and the total amount of fatty acids (free and esterified in 317 MAG, DAG and TAG) in the oil at the beginning of the reaction. 318 n o i s r e v d e t ep c c A 319 3. Results and Discussion 320 3.1 Immobilization yield 321 rROL and rCPL were immobilized on synthetic resins by adsorption, since it is 322 an economic and easy immobilization technique that maintains lipase activity 323 and specificity. rROL was also immobilized on ECR8285 M and AmberiteTM IRA 324 96 by covalent binding, which is considered one of the most efficient technique 325 for enzyme immobilization, due to the formation of chemically stable covalent 14 326 linkages between the different functional groups of the lipases and the active 327 functionalities of the carrier. 328 Table 2 shows the immobilization results in terms of yield and amount of 329 immobilized protein. The highest immobilization yield was achieved with rCPL in 330 Lewatit VPOC 1600 (98 %), which was higher than the value observed for rROL 331 in the same support (77.2 %). It is worthy to notice that the amount of 332 immobilized rCPL protein is much lower than that of rROL immobilized in 333 Lewatit VP OC 1600. 334 With respect to rROL, the immobilization yields were similar for Lewatit VPOC 335 1600, ECR1030M and ECR8285M (77.2-79.5 %) and slightly lower for 336 AP1090M (70.1 %). The lowest immobilization yield was observed for IRA-96. 337 The immobilization method used (adsorption or covalent binding) seems not to 338 affect the immobilization yield, evaluated in terms of immobilized protein. 339 n o i s r e v d e t ep c c A 340 3.2. Time-course of the transesterification reactions catalyzed by rROL 341 and rCPL 342 The results obtained after 48 h batch transterification reactions with methanol 343 and catalyzed by rROL immobilized on Lewatit VP OC 1600, IRA-96, 344 ECR1030M, ECR8285M, AP1090M or rCPL on Lewatit VP OC 1600 are 345 presented in Fig. 1. 346 The highest methyl ester production rates were observed in the beginning of the 347 transesterification reactions. After the second methanol addition, reaction 348 progress was slower and quasi-equilibrium was attained in less than 4 h for all 15 349 the biocatalysts tested. No glycerol was detected along the reactions. Thus, 350 acyl migration was not produced. 351 The maximum percentage of FAME (%, w/w) in the reaction medium obtained 352 with rROL immobilized on Lewatit VPOC 1600, IRA-96, ECR1030M, 353 ECR8285M or AP1090M and rCPL immobilized on Lewatit VPOC 1600, was 354 64.5, 63.8, 64.8, 60.6, 60.4 and 51.7 %, respectively. These results are very 355 close to the theoretical maximum FAME production, which is 66 (mol-%) for sn- 356 1,3 regioselective lipases. They do not directly reflect the amounts of 357 immobilized protein in the supports. In fact, for rROL, similar FAME production 358 was observed when this lipase was immobilized in Lewatit VP OC 1600, IRA-96 359 or ECR1030M, while IRA-96 showed the lowest protein load (Table 2). With 360 rCPL, 51.7 % FAME was obtained, in spite of the low amount of immobilized 361 protein (21.6 mg/g Lewatit VP OC 1600). Probably, a higher rCPL load in the 362 support would increase FAME yield. 363 More important than the amount of immobilized protein is the catalytic activity of 364 this protein. Also, deactivation and/or steric hindrance on the lipase 365 conformation occurring during immobilization, as well as internal diffusion 366 effects during the reaction may be responsible for the different results observed. 367 The maximum amount of MAG varied from 1.5 % with rCPL immobilized in 368 Lewatit VP OC 1600 to 27.9 % in rROL immobilized on Lewatit VP OC 1600. 369 In a study carried out by Canet et al. (2014), rROL immobilized in octadecyl- 370 Sepabeads was successfully used as catalyst for the transesterification of virgin 371 olive oil with methanol. Reaction conditions were the same as described in the 372 present study and a 50.3 % FAME yield was achieved in 3 hour reaction. This n o i s r e v d e t ep c c A 16 373 value is similar to that obtained with rCPL in our study. Also, Duarte et al. 374 (2015) produced biodiesel from yeast oil and olive oil using the same rROL 375 immobilized in Relizyme OD403 (polymethacrylate) as catalyst, in a solvent 376 system, with stepwise methanol addition. However, a lower FAME yield (40.6%) 377 was obtained with yeast oil as substrate, when comparing with olive oil (Canet 378 et al., 2014; Duarte et al., 2015) and Jatropha oil, in our study. 379 The transesterification of jatropha oil with methanol has been also carried out by 380 non-regioselective lipases, namely Burkholderia cepacia lipase immobilized on 381 modified attapulgite (You et al., 2013) and free recombinant Candida rugosa 382 lipase isozymes (Kuo et al., 2015), also using stepwise methanol addition. 383 When Burkholderia cepacia lipase was used, 94 % of biodiesel yield was 384 attained after 24 h reaction at 35 ºC (You et al., 2013). With C. rugosa lipase 385 isozymes, a maximum of 95.3 % FAME yield was obtained after 48h reaction at 386 37 ºC (Kuo et al., 2015). 387 In fact, in the studies on the production of biodiesel from jatropha oil, using non- 388 regioselective lipases as catalysts for the transesterification reaction, either in 389 solvent or solvent-free systems, FAME yields between 75 and 98 % have been 390 attained after 24 to 90 h reaction times (Juan et al., 2011) . 391 In our study, reaction equilibrium was attained after 4 h transesterification with 392 all the biocatalysts tested. This result, together with the absence of free glycerol 393 in the reaction medium, is highly beneficial in terms of industrial scale-up of the 394 process and reaction implementation in continuous bioreactors. n o i s r e v d e t ep c c A 395 396 3.4 Operational stability of the tested lipases 17 397 The short reaction time needed to attain equilibrium, together with the high 398 FAME yields obtained with all biocatalysts tested are very interesting results. 399 However, a high operational stability of the biocatalyst is also a key-factor for 400 industrial implementation of the process. 401 Thus, the operational stability of rROL and rCPL immobilized in different 402 supports was assayed in 10 or 8 consecutive batches, as previously described 403 (c.f. 2.4.6). The duration of each batch (4 h) was selected from the results 404 obtained in the 48 h time-course transesterification. Since biocatalyst 405 inactivation by dehydration is currently described (Nunes et al., 2012b; Tecelão 406 et al., 2012b), lipases were rehydrated between batches. 407 Residual activities along reuses, deactivation models fitted to the experimental 408 data and estimated half-life times for the biocatalysts tested, are presented in 409 Fig. 2 and Table 3. Lipase stability was found to be dependent on the 410 characteristics of immobilization matrices. The behaviour of rROL in ECR8285M 411 and rehydrated rROL in Lewatit VP OC 1600 can be described by a two 412 component first-order decay model. The inactivation profiles of rROL in 413 AP1090M, ECR1030M or IRA-96 and rCPL in Lewatit VP OC 1600, could be 414 well described by a first-order deactivation model. 415 The best results, in terms of operational stability, were observed when the 416 biocatalysts were reused without rehydration. The highest half-life value was 417 estimated for rROL immobilized in ECR1030M (579 h), followed by IRA-96 (381 418 h), AP1090M (270 h) and Lewatit VP OC 1600 (113 h). For rROL in ECR8285M 419 and rCPL in Lewatit VP OC 1600, the stability was much lower, with half-lives of 420 16 and 27.3 h, respectively. The lower stability exhibited by rCPL immobilized in n o i s r e v d e t ep c c A 18 421 Lewatit VP OC 1600, compared to that of rROL in the same support, may be 422 explained by different protocols followed for immobilization. The treatment with 423 glutaraldehyde after rROL adsorption will promote the formation of stable 424 crosslink between the lipase and the matrix, as well as intermolecular bonds 425 between enzyme molecules, hindering the leakage of enzyme molecules along 426 operation (Tecelão et al., 2012b). Also, the inhibitory effect of methanol for rCPL 427 may be stronger than for rROL. When lipases were rehydrated between reuses, 428 a dramatic loss of activity was observed, probably due to the leaching of 429 enzyme molecules during hydration. Also, the presence of water molecules in 430 the support will increase its hydrophilicity, promoting methanol diffusion inside 431 the support, with the risk of reaching inhibitory concentrations at the 432 microenvironment of the enzyme. 433 When rROL immobilized in Lewatit VP OC 1600 was used as catalyst for the 434 production of (i) human milk fat substitutes (HMFS) by acidolysis of tripalmitin 435 with oleic acid, in solvent-free media, an increase in operational stability was 436 observed when the biocatalyst was rehydrated between reuses (t1/2 increased 437 from 64 to 195 h) (Tecelão et al., 2012b). Similar behaviour was observed with 438 the same immobilized rROL used for the production of low calorie TAG: an 439 increase in t1/2 from 49 to 234 h was observed after rehydration (Nunes et al., 440 2012b). It worth to notice that in these studies carried out by Tecelão et al. 441 (2012b) and Nunes et al. (2012b), no water was added to the reaction medium. 442 In the present study, the addition of 4% water to the reaction medium shows to 443 be sufficient to maintain rROL activity. Conversely, the rehydration of rROL 444 immobilized in Eupergit C, also used for the production of low calorie TAG in n o i s r e v d e t ep c c A 19 445 solvent-free media, resulted in a decrease in operational stability of the 446 biocatalyst (Nunes et al., 2011; 2012 b). 447 The operational stability of rROL in octadecyl sepabeads used by Canet et al. 448 (2014) in the transesterification of olive oil was evaluated after 2 and 21 h of 449 reaction time. No significant differences were observed in methyl esters yield 450 (%) when this biocatalyst was reused. As in the experiments without biocatalyst 451 rehydration between batches of the present study, methanol was not washed 452 out between batches. In view of that, methanol did not seem to inactivate rROL 453 in octadecyl-Sepabeads under the considered conditions. However, the same 454 lipase immobilized in Relizyme OD403 lost 30 % of its activity after 6 455 reutilizations of 4 h each in transesterification of yeast oil or olive oil (Duarte et 456 al., 2015). 457 Luna et al (2014) used a low-cost multipurpose additive for the food industry 458 (Biolipase-R, from Biocon-Spain), containing Rhizopus oryzae lipase, as 459 catalyst for the transesterification reaction of sunflower oil with ethanol. When 460 this enzyme preparation was covalently immobilized on amorphous 461 AlPO4/sepiolite support with the p-hydroxybenzaldehyde linker, a conversion of 462 84.3 % of TAG into a blend of fatty acid ethyl esters (FAEE), MAG and DAG 463 was obtained, after 2h reaction at 30 ºC, using a molar ratio oil/ethanol of 1:6. 464 This biocatalyst did not show a significant loss of its initial catalytic activity for 465 more than five successive reuses of 2 h each. 466 The rROL used in the present study showed similar or even higher stability in 467 presence of methanol than Biolipase-R in presence of ethanol, which is an 468 alcohol with lower deactivation effect on lipases. n o i s r e v d e t ep c c A 20 469 470 4. Conclusions 471 The non-commercial recombinant sn-1,3 regioselective lipases rROL and rCPL 472 are promising catalysts for the production of biodiesel and MAG from crude 473 Jatropha oil. Transesterification was rather fast and equilibrium was reached 474 after 4 h-reaction with high FAME yields, varying from 51.7 to 64.8 %, which is 475 very close to the theoretical maximum for sn-1,3 regioselective lipases (66%). 476 All biocatalysts were active during 10 consecutive batches without rehydration. 477 However, when lipases were rehydrated between each two consecutive 478 batches, the loss of activity was much faster. rROL in Lewatit VPOC 1600 479 showed to be more stable than rCPL in the same support. n o i s r e v d e t ep 480 c c A 481 Acknowledgements 482 This work was supported by the national funding of FCT – Fundação para a 483 Ciência e a Tecnologia, Portugal, to the research unit LEAF 484 (UID/AGR/04129/2013); by CONACYT (Mexico) project CB-2014-01-237737 485 and BIOCATEM network; and by the project CTQ2013-42391-R of the Spanish 486 Ministry of Economy and Competitiveness. 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A 610 comprehensive study of reaction parameters in the enzymatic production of 611 novel biofuels integrating glycerol into their composition. Biores. Technol. 612 101, 6657-6662. 26 613 34. Wang, Y., Shen, X., Li, Z., Li, W., Wang, F., Nie, X., Jiang, J., 2010. 614 Immobilized recombinant Rhizopus oryzae lipase for the production of 615 biodiesel in solvent free system. J. Mol. Catal. B: Enzym. 67, 45-51. 616 35. You, Q., Yin, X., Zhao, Y., Zhang, Y., 2013. Biodiesel production from 617 jatropha oil catalyzed by immobilized Burkholderia cepacia lipase on 618 modified attapulgite. Bioresource Technology. 148, 202–207. 619 n o i s r e v d e t ep c c A 27 620 Figure captions 621 622 Figure 1. Evolution of fatty acid methyl ester (FAME), monoacylglycerol (MAG), 623 diacylglycerol (DAG) and triacylglycerol (TAG) concentrations in the reaction 624 medium, during the 48 hour transesterification reaction catalysed by rROL 625 immobilized in different supports or rCPL in Lewatit® VP OC 1600. 626 627 Figure 2. Operational stability of rROL immobilized in different supports or CPL 628 in Lewatit® VP OC 1600, with and without rehydration of the biocatalyst 629 between each consecutive 4-h batch, when transesterification of Jatropha oil 630 with methanol was performed. 633 e v d e t ep 631 632 n o i s r c c A 28 Table 1. Main physical and chemical properties of synthetic resins tested for rROL or rCPL immobilization. Carrier LifetechTM ECR8285M LifetechTM AP1090M LifetechTM ECR1030M Method of Functional immobilization Group Covalent Epoxy Polymer structure Epoxy/butyl methacrylate Macroporous styrene A DVB-crosslinked 1600 methacrylate d e t Adsorption p e cc DVB/ methacrylate Lewatit® VP OC binding Pore range Diameter (mm) (A) n o si r e v None Particle size 0.30 - 0.71 400 - 500 0.30 - 0.71 900- 1100 Styren/divinylbenzene Adsorption None 0.30 - 0.71 250 Adsorption None 0.315 - 1.0 150 96 copolymer Covalent amine: binding at least 85 % 29 Appearance White, spherical, porous beads. Tan, opaque, Tertiary Amberlite™ IRA- Structure/ 0.55 - 0.750 - spherical beads Table 2- Immobilization yields (± STD) and amounts of immobilized protein for rROL immobilized in different supports and rCPL immobilized in Lewatit VP OC 1600. Biocatalyst Immobilization yield (%) Immobilized protein (mg/g support) rROL in Lewatit VPOC 1600 77.4 ± 4.1 rROL in Amberlite IRA-96 58.8 ± 1.0 e t ep rROL in Lifetech ECR1030M rROL in Lifetech ECR8285M c c A rROL in Lifetech AP1090M rCPL in Lewatit VP OC 1600 e v d n o i s r 65.7 50.0 79.5± 1.5 67.6 78.4 ± 8.3 66.7 70.1± 7.8 59.6 98.0 ±1.2. 21.6 30 Table 3. Deactivation model equations fitted to the experimental data and the estimated half-lives for rROL immobilized in different supports and rCPL in Lewatit® VP OC 1600 (n= batch number; 1 batch= 4 h). Biocatalyst rROL in Lifetech rROL in Lifetech rROL in Lifetech TM TM TM Deactivation Model Model Equation 0.61n Non rehydrated Two component first-order Ares = 0.06e Rehydrated Two component first-order Ares = 22178.04e Non rehydrated First-order Rehydrated First-order Non rehydrated First-order Rehydrated First-order Non rehydrated First-order ECR8285M p e c rROL in Amberlite™ IRA-96 Rehydrated c A Non rehydrated rROL in Lewatit® VP OC 1600 d e t −0.21n + 115.20 e −5.64n n o si −0.12n + 23.55 e r e v −0.17n 15.6 −0.002n 579.0 Ares = 66.79 e −0.73n Ares = 203.92 e −0.006n Ares = 294.02 e First-order Ares = 80.91 e 6.6 −0.017n 113.0 -1.65n Rehydrated Two component first-order Non rehydrated First-order Ares = 90.26 e Rehydrated First-order Ares = 710.83 e 31 380.7 −1.07n First-order CPL in Lewatit® VP OC 1600 7.7 Ares = 88.51 e -0.10n 4.7 270.0 Ares = 97.17 e Ares = 8.14 e 16.0 −0.008n Ares = 85.76 e AP1090M ECR1030M Half life time (h) + 475.3 e −0.087n −1.96n 5.8 27.3 5.4 rROL - ECR1030M 10 20 30 100 90 80 70 60 50 40 30 20 10 0 40 0 10 20 Time (h) MAG DAG TAG FAME MAG 100 90 80 70 60 50 40 30 20 10 0 10 20 DAG 30 d e pt 100 90 80 70 60 50 40 30 20 10 0 e c c A 40 0 10 20 Time (h) MAG DAG TAG TAG 0 10 n o si FAME r e v 30 100 90 80 70 60 50 40 30 20 10 0 40 rROL - Lewatit VPOC 1600 [Compund] % (w/w) [Compound] %(w/w) rROL - IRA 96 0 30 Time (h) 40 20 MAG DAG MAG DAG TAG Fig. 1 32 TAG FAME 0 10 20 30 Time (h) FAME 40 100 90 80 70 60 50 40 30 20 10 0 Time (h) FAME 30 Time (h) Carica papaya - Lewatit VPOC1600 [Compound] % (w/w) 0 rROL - ECR8285 [Compound] % (w/w) 100 90 80 70 60 50 40 30 20 10 0 [Compound] (%, w/w) [Compound] % (w/w) rROL - AP1090M MAG DAG TAG FAME 40 rROL - ECR8285M rROL - ECR 1030M rROL-AP1090M 2 3 4 5 6 7 8 9 90 2 3 4 5 Rehydrated lipase Non rehydrated lipase rROL - IRA-96 7 8 9 d e pt 100 100 90 80 70 60 50 40 30 20 10 0 90 80 e c c A 70 60 50 40 2 3 4 5 6 7 8 9 10 Non rehydrated lipase 30 0 2 3 4 5 6 1 r e v 20 Batch number Rehydrated lipase 40 7 8 2 3 4 9 10 Non rehydrated lipase Fig. 2 33 6 7 8 9 10 Non rehydrated lipase Carica papaya - Lewatit VPOC1600 100 90 80 70 60 50 40 30 20 10 0 1 2 3 4 5 6 7 8 Batch number Batch number Rehydrated lipase 5 Batch number Rehydrated lipase 30 1 50 n o si 10 1 60 0 10 Non rehydrated lipase rROL - Lewatit VPOC 1600 Residual activity (%) Residual Activity (%) 6 Batch number Batch number Rehydrated lipase 70 10 1 10 80 20 Residual activity (%) 1 100 90 80 70 60 50 40 30 20 10 0 Residual activity (%) 100 90 80 70 60 50 40 30 20 10 0 Residual activity (%) Residual activity (%) 100 Rehydrated lipase Non rehydrated lipase 9 10