C � h C E a A b c I a A R R A A K P X � � E X 1 w p v 4 f p i P p 2 ( h 1 Process Biochemistry 51 (2016) 614–623 Contents lists available at ScienceDirect Process Biochemistry jo u r n al homep age: www.elsev ier .com/ locate /procbio o-immobilization and stabilization of xylanase, �-xylosidase and -l-arabinofuranosidase from Penicillium janczewskii for arabinoxylan ydrolysis ésar Rafael Fanchini Terrasana,∗, Lara Trobo-Masedaa, Sonia Moreno-Péreza, leonora Cano Carmonab, Benevides Costa Pesselac, José Manuel Guisana Departamento de Biocatálisis, Instituto de Catálisis y Petroleoquimica (ICP), Consejo Superior de Investigaciones Científicas (CSIC), Campus Universidad utónoma de Madrid (UAM), Cantoblanco, 28049 Madrid, Spain Biochemistry and Microbiology Department, Biosciences Institute, Univ. Estadual Paulista—UNESP, PO 199, 13506-900 Rio Claro, SP, Brazil Departamento de Biotecnología y Microbiología de Alimentos, Instituto de Investigación en Ciencias de los Alimentos (CIAL), Consejo Superior de nvestigaciones Científicas (CSIC), Campus Universidad Autónoma de Madrid (UAM), Cantoblanco, 28049 Madrid, Spain r t i c l e i n f o rticle history: eceived 30 November 2015 eceived in revised form 23 February 2016 ccepted 25 February 2016 vailable online 3 March 2016 eywords: enicillium janczewskii ylanase a b s t r a c t Differently activated agarose-based supports were evaluated for co-immobilization of a crude extract from Penicillium janczewskii containing xylanase, �-xylosidase and �-l-arabinofuranosidase activities. Adequately selecting support and immobilization conditions (8 h, using agarose with 10% crosslinking) increased enzyme levels substantially, mainly in relation to the xylanase (2-fold). A coating with dextran aldehyde MW 6000 Da, partially oxidized, covalently attached the enzymes to the support. Optimum activity was verified in the pH range 2–4, and at 50, 65 and 80 ◦C for the xylanase, �-l-arabinofuranosidase and �-xylosidase, respectively. The xylanase was highly thermostable retaining more than 70% of activity even after 24 h incubation at 60 and 70 ◦C; and at 80 ◦C its half-life was 1.7 h. The half-lives of the �- ◦ -xylosidase -l-arabinofuranosidase nzyme co-immobilization ylan hydrolysis xylosidase and �-l-arabinofuranosidase at 50 C were 2.3 and 3.8 h, respectively. The co-immobilization of the enzymes on a single support give raise to a functional multi-enzymatic biocatalyst acting in the complete hydrolysis of different and complex substrates such as oat spelt and wheat arabinoxylans, with xylose yield higher than 40%. The xylanase and the �-l-arabinofuranosidase presented high stability retaining 86.6 and 88.0% of activity after 10 reuse cycles. © 2016 Elsevier Ltd. All rights reserved. . Introduction Xylan is the second most abundant biopolymer in plant cell alls and the main hemicellulosic polysaccharide. It is com- osed of a �-(1 → 4) d-xylopyranosyl backbone substituted at arious degrees by side chain residues such as glucopyranosyl, -O-methyl-d-glucurono-pyranosyl, �-l-arabinofuranosyl, acetyl, eruloyl, and/or p-coumaroyl [1]. The precise composition of the olymer is strongly dependent on plant species and tissue. For nstance, hard wood xylans often have d-glucuronic acid attached ∗ Corresponding author at: Departamento de Biocatálisis, Instituto de Catálisis y etroleoquimica (ICP), Consejo Superior de Investigaciones Científicas (CSIC), Cam- us Universidad Autónoma de Madrid (UAM), Cantoblanco, Calle de Marie Curie, 8049 Madrid, Spain. E-mail addresses: cesarterrasan@hotmail.com, cesarterrasan@gmail.com C.R. Fanchini Terrasan). ttp://dx.doi.org/10.1016/j.procbio.2016.02.014 359-5113/© 2016 Elsevier Ltd. All rights reserved. to their backbone, whereas l-arabinose is the most common branch in cereal xylans [2]. Given the diversity of xylan structures, their complete and efficient hydrolysis involves the synergistic action of main chain degrading enzymes, including endo-�-1,4-xylanases (EC 3.2.1.8) and �-d-xylosidases (EC 3.2.1.37), and side chain cleav- ing enzymes, including �-l-arabinofuranosidases (EC 3.2.1.55), �-glucuronidases (EC 3.2.1.139), acetyl xylan esterase (EC 3.1.1.72), and feruloyl esterases (EC 3.1.1.73). Endo-�-1,4-xylanase and �-d- xylosidase are the main enzymes responsible for the degradation of the polymer: xylanases cleave the internal �-(1 → 4) bonds in the xylan backbone, liberating different chain-length-(substituted) xylooligosaccharides, and �-xylosidases are exoglycosidases that release xylose from the non-reducing ends of these xylooligosac- charides. �-xylosidases are critical for the systems since they carry the greatest work load in terms of number of glycosidic bonds cleaved, as well as in relieving product inhibition of xylanases [3]. Among other accessory enzymes, �-l-arabinofuranosidases are dx.doi.org/10.1016/j.procbio.2016.02.014 http://www.sciencedirect.com/science/journal/13595113 http://www.elsevier.com/locate/procbio http://crossmark.crossref.org/dialog/?doi=10.1016/j.procbio.2016.02.014&domain=pdf mailto:cesarterrasan@hotmail.com mailto:cesarterrasan@gmail.com dx.doi.org/10.1016/j.procbio.2016.02.014 cess B e a h i a t m [ e x c w c t l i M d i x c o e a l d e A i s x y s � c o A t 2 2 p p t d ( a ( a w l 2 2 e M m p C.R. Fanchini Terrasan et al. / Pro xo-type enzymes that catalyze the cleavage of the terminal �-l- rabinofuranosyl residues from arabinosylated substrates. Xylan ydrolysis is not aleatory, i.e., the degree of substitution in xylan nfluence the products of hydrolysis for xylanases [4]. In this sense, ccessory enzymes such as �-l-arabinofuranosidases are impor- ant since the removal of side-chain residues from xylan backbone ay have a synergistic effect with the other xylanolytic enzymes 5] and also the difference in substrate specificity among differ- nt xylanases has important implications in the deconstruction of ylan [4]. Enzyme immobilization poses as a possibility to improve the haracteristics of an enzyme in terms of stability and catalysis, as ell as for process improvement allowing the reuse of the bio- atalyst for many operational cycles [6]. Immobilization of more han one enzyme on the same support, however, is especially chal- enging, as it has to preserve the catalytic activity of all enzymes nvolved in the system and ideally improve their stability [7]. any xylanolytic enzymes have been individually immobilized by ifferent methods; and some studies have investigated the co- mmobilization of two xylanolytic enzymes [8]. In this sense, the ylanase and �-xylosidase from Talaromyces thermophilus were o-immobilized on chitosan and employed for the hydrolysis f oat spelt xylan, demonstrating the synergistic action of both nzymes by increasing the saccharification of the substrate [9]. In nother study, co-immobilization of recombinant xylanase and �- -arabinofuranosidase onglyoxyl agarose was evaluated through ifferent approaches in the hydrolysis of arabinoxylan [10]. The ffect of xylanase, �-xylosidase and �-l-arabinofuranosidase from spergillus oryzae in the decomposition of arabinoxylan was ver- fied using the soluble enzymes in the moromi mash during soy auce fermentation [11], nevertheless, co-immobilization of three ylanolytic enzymes acting cooperatively in the complete hydrol- sis of complex substrates has not been reported to date. This way, the aims of this work were to establish a protocol for imultaneous co-immobilization of the xylanase, �-xylosidase and -l-arabinofuranosidase from Penicillium janczewskii present in the rude extracellular extract, as well as improve the stabilization f the immobilized enzymes via post-immobilization techniques. fter that, the immobilized enzymes were biochemically charac- erized and evaluated in the hydrolysis of arabinoxylans. . Materials and methods .1. Materials Agarose with 4, 6 and 10% of cross-linking BCL were urchased from Agarose Bead Technologies (Madrid, Spain). -nitrophenyl �-d-xylopyranoside (pNPX), glycidol, potassium etraborate tetrahydrate, sodium borohydride, sodium perio- ate, ethylenediamine, glutaraldehyde, Leuconostocc spp. dextran MW 6000–100,000), polyethylenimine (PEI, MW 1300), oat spelt nd beechwood xylans were obtained from Sigma-Aldrich Co St. Louis, MO). d-xylose Assay Kit, xylose, p-nitrophenyl �-l- rabinofuranoside (pNPAra) and low viscosity wheat arabinoxylan ere from Megazyme (Wicklow, Ireland). All reagents were of ana- ytical grade. .2. Methods .2.1. Microorganism, enzyme production and preparation of nzyme extract P. janczewskii (CRM 1348) is deposited in The Central of icrobial Resources, CMR-UNESP, Brazil. The microorganism was aintained on Vogel solid medium [12] and liquid cultures were repared in the same medium with brewer’s spent grain as iochemistry 51 (2016) 614–623 615 substrate, under optimized conditions for xylanolytic enzymes pro- duction [13]. After cultivation, the mycelium was removed by vacuum filtration and the culture filtrate was centrifuged (10,000g, 4 ◦C, 15 min). The supernatant was dialyzed overnight against dis- tilled water, 0.025 M sodium acetate buffer pH 5.0 or 0.025 M sodium phosphate buffer pH 7.0 before immobilization. A sample of the supernatant was also treated with 0.01 M sodium periodate for 1.5 h in order to oxidize sugar moieties of the enzymes and then dialyzed against 0.025 M sodium phosphate buffer pH 7.0. 2.2.2. SDS-PAGE A sample containing 50 �g of protein prepared from the extra- cellular extract obtained under optimized conditions for xylanase production (medium with oat spelt xylan, pH 6.5, 7 days of cultiva- tion, 30 ◦C) was applied to SDS-PAGE performed in 8–18% (w/v) gradient gels, according to Laemmli [14]. The resolved protein bands were visualized after staining with 0.1% Coomassie bril- liant blue R-250 dissolved in methanol, acetic acid, and distilled water (4:1:5 v/v/v). Standard proteins (Sigma) were phosphorylase b (97 kDa), bovine serum albumin (66 kDa), ovalbumin (45 kDa), carbonic anhydrase (29 kDa), trypsin inhibitor (20 kDa), and �- lactalbumin (14.2 kDa). 2.2.3. Enzyme assays Xylanase activity was determined according to Bailey et al. [15] with 1% (w/v) beechwood xylan prepared in 0.05 M sodium acetate buffer pH 5.0 (before determining optimum pH) or pH 4.0 (after determining optimum pH) and appropriately diluted enzyme solution. Reducing sugars were quantified with DNS acid reagent [16]. �-xylosidase and �-l-arabinofuranosidase activities were determined in a reaction mixture containing, respectively, 3 mM pNPX and pNPAra prepared in 0.05 M sodium acetate buffer pH 5.0 (before determining optimum pH) or pH 4.0 (after deter- mining optimum pH) and appropriately diluted enzyme solution to 1 mL final volume. Reactions were stopped by adding 1 mL of a saturated potassium tetraborate solution and the absorbance was measured at 405 nm [17]. One unit of activity was defined as the amount of enzyme required to release 1 �mol of product equivalent per min in the assay conditions at 25 ◦C. 2.2.4. Preparation of support Monoaminoethyl-N-aminoethyl (MANAE)-agarose, was pre- pared as described elsewhere [18] and the glutaraldehyde-agarose support was prepared from MANAE agarose [19,20]. Briefly described, we used 10 g of MANAE in 20 mL of 0.5 or 15% (v/v) glu- taraldehyde solution prepared in 0.2 M phosphate buffer pH 7.0. The suspensions were kept under mild stirring at 25 ◦C for 1 and 15 h in the case of the supports activated with 0.5 or 15% (v/v) glutaraldehyde, respectively. This treatment permitted to fully modify the primary amino groups of the support with one or two glutaraldehyde molecules, respectively [19]. After that, the sup- ports were filtered and washed exhaustively with 0.025 M sodium phosphate buffer and then with distilled water. Glyoxyl-agarose was prepared with the maximal activation degree, as previously described [21]. Polyethylenimine (PEI) and dextran sulfate [22] and heterofunctional amino-glyoxyl and amino-epoxide [23,24] sup- ports were prepared as described elsewhere. The supports were initially prepared using agarose with 4% crosslinking. MANAE and 0.5% (w/v) glutaraldehyde supports were further prepared using agarose with 6 and 10% crosslinking. Acti- vated supports were stored at 4 ◦C and, before use, washed with incubation buffer according to immobilization condition. 2.2.5. Enzyme immobilization Immobilizations were performed by suspending 1:10 (w/v) the activated supports in the dialyzed/diluted enzyme solution. Buffers 6 cess B f p g s o w o t s s a 2 t w t c M e d 8 1 f d o s w u a t a d a a s 2 i p t a 2 c p e d 2 2 c p a a d a t 2 w i 16 C.R. Fanchini Terrasan et al. / Pro or each immobilization were: 0.025 M sodium phosphate buffer H 7.0 for MANAE, PEI, glutaraldehyde, amino-epoxide and amino- lyoxyl supports, 0.05 M sodium acetate buffer pH 5.0 for dextran ulfate support, 0.1 M sodium bicarbonate buffer pH 10.0 for gly- xyl support (enzyme solution dialyzed against distilled water as 2-fold diluted in the buffer). Immobilizations were carried ut under gentle agitation at 25 ◦C for 4 h or overnight incuba- ion. During immobilization, samples of the suspensions and the upernatants were withdrawn and enzyme activities were mea- ured. Proteins were measured during immobilization time-course nd immobilization for different periods. .2.6. Post-immobilization techniques Dextran-coated derivatives: a mass of 0.5 g of the uncoated glu- araldehyde (Glut) derivative (immobilization carried out by 8 h) as added to 5 mL of aldehyde-dextran suspensions, pH adjusted o 7.0. Dextrans with MW of 1500, 6000, 25,000 and 75,000 Da ompletely oxidized were initially evaluated [25]. Dextran with W of 6000 with 20 and 40% degree of oxidation were further valuated. The suspensions were gently agitated overnight, the erivative was then re-suspended in sodium borate buffer pH .5 or sodium phosphate buffer pH 7.0 and reduced by adding mg/mL sodium borohydride. The suspension was gently agitated or 30 min, washed abundantly with water and vacuum filtered. PEI-coated derivative: a mass of 0.5 g of the uncoated Glut erivative (immobilization carried out for 8 h) was added to 5 mL f 5% (w/v) PEI MW 1300 Da solution, pH was adjusted to 7.0. The uspension was gently agitated overnight, washed abundantly with ater and vacuum filtered. Glutaraldehyde cross-linked derivative: a mass of 1.0 g of the ncoated Glut derivative (immobilization carried out for 8 h) was dded to 10 mL of 0.5% (v/v) glutaraldehyde solution pH adjusted o 7.0. The suspensions were gently agitated for 30 min, washed bundantly with water and vacuum filtered. A mass of 0.5 g of this erivative was directly reduced with 1 mg/mL sodium borohydride nd 0.5 g was previously coated with dextran, as described above, nd then reduced with 1 mg/mL sodium borohydride. The suspen- ions were washed abundantly with water and vacuum filtered. .2.7. Immobilization parameters Immobilization yield was defined as the ratio between the activ- ties (or protein) in the supernatant compared to the activity (or rotein) in the control. Expressed activity was defined as the ratio of he activity in the final suspension after the immobilization process nd the initial enzyme activity. .2.8. Evaluation of the attachment between enzyme and support The glutaraldehyde agarose 10 BCL derivative (before and after oating with aldehyde dextran) was incubated in 0.005 M sodium hosphate buffer pH 7.0 with 0.5 M NaCl at 25 ◦C. After 1 h, nzyme activities were analyzed in the supernatant, as previously escribed. .2.9. Derivative characterization .2.9.1. Thermal stability. A mass of 0.1 g of the Glut derivative oated with dextran MW 6000 and 40% oxidation degree was sus- ended in 1.0 mL of 0.05 M acetate buffer pH 5.0 and incubated t 50 ◦C. In all cases, samples of the suspension were withdrawn t several intervals and the activity was assayed as previously escribed. Residual activity was calculated as the ratio between ctivity at a given time and the activity in the beginning of incuba- ion (regarded as 100%). .2.9.2. pH stability. A mass of 0.1 g of the Glut derivative coated ith dextran MW 6000 at 40% degree of oxidation was suspended n 1.0 mL of 0.05 M glycine HCl buffer pH 3.0, 0.05 M sodium acetate iochemistry 51 (2016) 614–623 buffer pH 4.0 and 5.0, and 0.05 M sodium phosphate buffer pH 7.0. The suspension was incubated at 50 ◦C and after 4 h residual activity was assayed. Initial activities before incubation were regarded as 100%. 2.2.9.3. Optima pH and temperature. Optimum pH was determined by assaying enzyme activities of the Glut derivative coated with dextran MW 6000 at 40% degree of oxidation at 25 ◦C at various pH from 2.0 to 8.0. The following buffers were utilized: 0.05 M glycine- HCl pH 2.0 and 3.0, 0.05 M sodium acetate pH 4.0 and 5.0, 0.05 M sodium phosphate pH 6.2 and 7.0, and 0.05 M Tris HCl pH 8.0. Optimum temperature was determined by assaying enzyme activities at temperatures ranging from 40 to 85 ◦C, with 5 ◦C inter- vals, in 0.05 M sodium acetate buffer pH 5.0. 2.2.10. Hydrolysis of arabinoxylans The hydrolysis of 0.5% (w/v) oat spelt xylan (arabinose residues ≤10%, glucose residues ≤ 15%, xylose residues ≥ 70%) and low vis- cosity wheat arabinoxylan (38/62 arabinose:xylose relation) were carried using the Glut derivative coated with dextran MW 6000 at 40% degree of oxidation. The reactions were carried out in 0.05 M sodium acetate buffer pH 4.0 at 40 ◦C. Collected samples were fil- tered and the adequately diluted supernatant was analyzed for xylose. 2.2.11. Reuse assay Successive hydrolysis cycles of 0.5% (w/v) wheat arabinoxylan prepared in 0.05 M sodium acetate buffer pH 4.0 were performed using the Glut derivative coated with dextran MW 6000 at 40% degree of oxidation, at 1:10 proportion (w/v). Each cycle was car- ried out at 40 ◦C for 1 h under magnetic stirring (300 rpm). At the end of the cycles, the derivative was filtered, washed with 0.05 M sodium acetate buffer pH 4.0 and new substrate was added for a new reaction round. Samples of the supernatant were withdrawn during the first, the fifth and the tenth cycles, filtered and analyzed for reducing sugars. After the fifth and the tenth cycles the deriva- tive was suspended in the washing buffer (1:10, w/v) and residual activities were measured, as previously described (activities before the first cycle were regarded as 100%). 2.2.12. Analytical methods Protein concentration was determined with the modified Brad- ford’s method, with bovine serum albumin as standard [26]. Reducing sugars were quantified with DNS acid reagent, with xylose as standard [16]. Xylose was quantified using the enzymatic d-xylose Assay Kit. The first reaction involves the conversion of the �- to the �-anomeric form of d-xylose catalyzed by xylose mutaro- tase. The �-d-xylose was then oxidized by NAD+ to d-xylonic acid in the presence of �-xylose dehydrogenase. The amount of NADH formed in the reaction is stoichiometric with the amount of d-xylose. NADH was measured by the increase in absorbance at 340 nm (� = 6300 M−1 cm−1). Samples of hydrolyzed oat spelt and wheat arabinoxylans were adequately diluted and analyzed according to the supplier instruction, in duplicate, and expressed as mean value. Xylose yield was calculated using 0.88 as the con- version factor of pentose to equivalent xylan, as below: Xylose yield (%) = xylose released (g) × 0.88 initial xylan (g) × 100 3. Results and discussion 3.1. Enzyme immobilization P. janczewskii was isolated from soil of the Brazilian Rainfor- est [27] and characterized as an excellent producer of xylanolytic C.R. Fanchini Terrasan et al. / Process Biochemistry 51 (2016) 614–623 617 Table 1 Co-immobilization of xylanase, �-xylosidase and �-l-arabinofuranosidase from P. janczewskii on different agarose-based supports. Derivative Condition/strategy Xylanase �-xylosidase �-l-arabinofuranosidase Yield (%) Expressed activity (%) Yield (%) Expressed activity (%) Yield(%) Expressed activity (%) MANAE pH 7, 4 h 53.4 20.0 100 58.5 100 18.0 pH 8.5, overnight – – – – – – pH 7, 24 h, sodium periodate treatment 100 167.5 100 73.3 100 55.1 Glutaraldehyde 0.5% pH 7, 4 h 63.2 20.8 100 76.2 100 22.4 Glutaraldehyde 10% pH 7, 4 h 100 1.2 100 100 100 74 PEI 1300 0,5% pH 7, overnight 100 6.8 0 0 30 28.9 Enzyme activities were assayed in 0.05 M sodium acetate buffer pH 5.0 at 25 ◦C. Initial loads: xylanase—64 U; �-xylosidase—23 mU; �-l-arabinofuranosidase—34 mU; proteins—5.3 mg. Supports were prepared using agarose with 4% crosslinking. Table 2 Co-immobilization of xylanase, �-xylosidase and �-l-arabinofuranosidase from P. janczewskii on different agarose beads MANAE and Glut supports. Derivative Agarose beads (% CL) Expressed activity (%) Xylanase �-xylosidase �-l-arabinofuranosidase MANAE 4 20.0 58.5 18.0 6 14.6 54.6 14.2 10 14.0 54.0 14.0 Glut 4 20.8 76.2 22.4 6 23.1 82.5 25.1 10 26.1 100 42.5 Immobilization was carried out in 0.025 M sodium phosphate buffer pH 7.0 for 4 h at 25 ◦C. Initial loads: xylanase—64 U; �-xylosidase—23 mU; �-l-arabinofuranosidase—34 m Glut—0.5% (v/v) glutaraldehyde activated support. Fig. 1. SDS-PAGE (8–18%) of the crude extracellular extract from P. janczewskii. Lane 1: standard proteins, phosphorylase b (94 kDa), bovine serum albumin (67 kDa), o l [ e � o m t t t o x b d � l ( valbumin (43 kDa), carbonic anhydrase (30 kDa), trypsin inhibitor (20 kDa) and �- actalbumin (14.4 kDa); Lane 2: crude extracellular extract (50 �g). Modified from 28] . nzymes in the absence of cellulases [13]. Production of xylanase, -xylosidase and �-l-arabinofuranosidase has been previously ptimized [28,29] but other enzymes from the xylanolytic complex ay also be produced by this fungal strain since SDS-PAGE revealed he presence of several extracellular proteins (Fig. 1) [28]. Due to heir potential application in many bioprocesses, several activa- ion protocols, techniques and strategies were evaluated in order to ptimize the concomitant immobilization of the crude extracellular ylanase, �-xylosidase and �-l-arabinofuranosidase on agarose- ased supports, which have been widely used for immobilizing ifferent enzymes [30]. In general, good results were observed in relation to the -xylosidase immobilization, although the xylanase and the �- -arabinofuranosidase were hardly immobilized on the supports Table 1). Good balance for the immobilization of the three enzymes 25 ◦C. Enzyme activities were assayed in 0.05 M sodium acetate buffer pH 5.0 at U; proteins—5.3 mg. CL—crosslinking. MANAE—monoaminoethyl-N-aminoethyl, was obtained with ionic immobilization on MANAE agarose. The use of pH 8.5 and more prolonged immobilization period did not improve the activities on this support. When the enzymes were previously treated with sodium periodate to oxidize sac- charide moieties of the proteins, the three enzymes were easily attached to the anionic MANAE support. This fact indicate that the presence of carbohydrates may be blocking the access of the enzymes to the reactive groups of the support or the formed aldehyde can readily react with these groups forming imine or Schiff bases. The removal of glycosylation was good in terms of immobilization; nevertheless, the enzymes became unstable even during mild conditions of enzymatic assays (not shown). The use of MANAE agarose cross-linked with 0.5% (w/v) glutaraldehyde also resulted in good enzyme immobilization. When this sup- port was activated with higher glutaraldehyde concentration (10%, w/v), which results in the formation of glutaraldehyde dimers, improved immobilization yield of the xylanase was observed, probable consequence of the higher glutaraldehyde concentration, which allowed more attachments between enzyme and support. Nevertheless, these attachments may have been excessive, caus- ing distortion in the xylanase structure resulting in the very low expressed activity. Expressed activities of the �-xylosidase and �-l-arabinofuranosidase increased by increasing glutaraldehyde concentration that may be related to the fact that these enzymes usually have higher MW than the xylanases or correspond to mul- timeric enzymes [31,32]. In this case, the higher glutaraldehyde concentration provided more attachments stabilizing the structure of the immobilized enzyme or provided enough attachments to sta- bilize all enzyme subunits, thus resulting in the higher expressed activity. Immobilization of enzymes on glutaraldehyde-activated supports has been largely used on supports previously activated with amine groups. Immobilization is promoted through a two- step mechanism: in a first step the enzyme is adsorbed on the support via an anionic exchange mechanism and then, the covalent immobilization occurs [20,33]. The enzymes could not be immobilized on PEI (Table 1), dextran sulfate and glyoxyl activated supports (not shown). In the case of glyoxyl support, the enzymes probably present low stability in pH 618 C.R. Fanchini Terrasan et al. / Process Biochemistry 51 (2016) 614–623 Table 3 Co-immobilization of xylanase, �-xylosidase and �-l-arabinofuranosidase from P. janczewskii for different periods on Glut agarose 10% crosslinking. Incubation (h) Protein yield (%) Xylanase �-xylosidase �-l-arabinofuranosidase Yield (%) Expressed activity (%) Yield (%) Expressed activity (%) Yield (%) Expressed activity (%) 4 78.1 79.8 26.1 94.3 100 85.4 42.5 8 83.1 80.1 41.8 100 105.6 89.8 38.3 16 84.3 81.7 11.2 100 61.5 98.2 23.5 Immobilization was carried out in 0.025 M sodium phosphate buffer pH 7 at 25 ◦C. Enzyme activities were assayed in 0.05 M sodium acetate buffer pH 5.0 at 25 ◦C. Initial loads: xylanase—64 U; �-xylosidase—23 mU; �-l-arabinofuranosidase—34 mU; proteins—5.3 mg. Table 4 Post-immobilization strategies evaluated on co-immobilized xylanase, �-xylosidase and �-l-arabinofuranosidase from P. janczewskii on Glut agarose 10% crosslinking. Step Residual activity (%)a First coating Sodium borohydride reduction Second coating Sodium borohydride reduction Xylanase �-xylosidase �-l-arabinofuranosidase PEI − − − 0 0 0 Dextran + − − 66.9 57.0 55.5 Glutaraldehyde − − − 39.6 21.7 47.8 + − − 33.0 7.7 25.9 − Dextran + 15.8 7.4 10.9 Immobilization was carried out in 0.025 M sodium phosphate buffer pH 7 at 25 ◦C. Enzyme activities were assayed in 0.05 M sodium acetate buffer pH 5.0 at 25 ◦C. Step present (+) or absent (−). a In relation to the expressed activities of the derivative prepared by 8 h incubation (Table 3). S u p er n at an t ac ti vi ty o r p ro te in ( % ) 0 20 40 60 80 100 Time (h) 0 5 10 15 Fig. 2. Time-course of total protein, xylanase, �-xylosidase and �- arabinofuranosidase from P. janczewskii co-immobilization on Glut agarose 10% crosslinking. Immobilization was carried out in 0.025 M sodium phosphate buffer pH 7.0 at 25 ◦C. Enzymatic activities were assayed in 0.05 M sodium acetate buffer pH 5.0 at 25 ◦C. Initial protein (5.3 mg) and activities (xylanase—64 U; �-xylosidase—23 mU; �-l-arabinofuranosidase—34 mU) in the supernatant were r � 1 [ e r i i c 6 a i e f a t R es id u al a ct iv it y (% ) 0 20 40 60 80 100 Time (h) 0 2 864 Fig. 3. Thermostability of xylanase, �-xylosidase and �-l-arabinofuranosidase from P. janczewskii co-immobilized on MANAE-Glut derivative coated with MW 6000 and 40,000 dextrans (completely oxidized). Incubation was carried in 0.05 M sodium phosphate buffer pH 7.0 at 50 ◦C. Enzymatic activities were assayed in 0.05 M sodium acetate buffer pH 5.0 at 25 ◦C. Initial activities were regarded as 100%. Full sym- bols: derivative coated with dextran MW 6000. Empty symbols: derivative coated Based on previous observations, immobilization on the Glut sup- egarded as 100%. Relative (%) protein (*), and xylanase (�), �-xylosidase (�) and -l-arabinofuranosidase (�) activities. 0, a required condition for optimal immobilization in this support 21]. In the case of heterofunctional amino-glyoxyl and amino- poxide supports (not shown) the enzymes were not immobilized, emaining in the supernatant even after pH increase or long-term ncubation. Considering the previous results, a comparison between the mmobilization on MANAE and 0.5% (w/v) glutaraldehyde (further alled Glut agarose) prepared with different agarose beads, i.e., 4, and 10% of crosslinking, was carried out (Table 2). The expressed ctivity gradually increased by increasing the agarose crosslink- ng in the Glut derivative, whereas in the MANAE derivative the xpressed activity decreased by increasing agarose crosslinking rom 4 to 6 or 10%. The low crosslinking level in the 4 BCL garose results in larger pores, which contain higher concentra- ion of reactive groups for the co-immobilization of the enzymes with dextran MW 40,000. Residual (%) xylanase (�), �-xylosidase (�) and �-l- arabinofuranosidase (�) activities. in the MANAE support. The higher quantity of weak ionic inter- actions were not enough to distort enzyme structure resulting in the higher expressed activities. In opposition, the lower degree of derivatization with glutaraldehyde and the consequent fewer covalent bonds in the 10 BCL agarose resulted in less distortion to enzyme structures and consequent higher expressed activities. The Glut derivative using 10% cross-linked agarose was that which resulted in the highest expressed activity levels, corresponding to 26.1, 100 and 42.5% for the xylanase, �-xylosidase and �-l- arabinofuranosidase, respectively. port was followed by measuring the enzyme activities and protein in the supernatant during 16 h (Fig. 2). The enzymes were quickly immobilized on the support, i.e., after 30 min most of the enzyme C.R. Fanchini Terrasan et al. / Process Biochemistry 51 (2016) 614–623 619 R es id u al a ct iv it y (% ) 0 20 40 60 80 100 pH 3.0 pH 4.0 pH 5.0 pH 7.0 Fig. 4. Stability in different pH of the xylanase from P. janczewskii immobilized on Glut derivative coated with dextran MW 6000 at different degrees of oxidation. Incubation was carried for 4 h in 0.05 M glycine HCl buffer pH 3.0, 0.05 M sodium acetate buffer pH 4.0 and 5.0, and 0.05 M sodium phosphate buffer pH 7.0, at 50 ◦C. Enzymatic activity was assayed in 0.05 M sodium acetate buffer pH 5.0 at 25 ◦C. Ini- tial activities before incubation were regarded as 100%. Residual (%) activity in the derivatives coated with dextran at 20 (black), 40 (white), and 100% (grey) degree of oxidation. Table 5 Xylanase, �-xylosidase and �-l-arabinofuranosidase from P. janczewskii on Glut derivative coated with dextrans of different MW and degrees of oxidation. MW (Da) DO (%) Expressed activity (%) Xylanase �-xylosidase �-l-arabinofuranosidase 6000 100 53.5 41.7 26.4 40,000 100 66.9 57.0 55.5 100,000 100 8.8 10.2 24.3 6000 20 35 31.8 14.8 40 58.8 58.7 60.4 E M a i x t a t i a b i i t t ( f e s a c g 8 b h R el at iv e ac ti vi ty ( % ) 0 20 40 60 80 100 (a) (b) pH 321 87654 R el at iv e ac ti vi ty ( % ) 20 40 60 80 100 Temperature ( οC) 40 50 60 70 80 90 Fig. 5. Influence of pH (a) and temperature (b) on the activity of xylanase, �- xylosidase and �-l-arabinofuranosidase from P. janczewskii co-immobilized on Glut derivative coated with dextran MW 6000 at 40% degree of oxidation. (a) Enzymatic activities were assayed at 25 ◦C in 0.05 M glycine-HCl buffer pH 2.0 and 3.0, 0.05 M sodium acetate buffer pH 4.0 and 5.0, 0.05 M sodium phosphate buffer pH 6.2 and before, and 33.0, 7.7 and 25.9% after borohydride reduction, for nzyme activities were assayed in 0.05 M sodium acetate buffer pH 5.0 at 25 ◦C. W—molecular weight; DO—degree of oxidation. ctivities had disappeared from the supernatant. After this period, mmobilization proceeded at lower rate, with exception of the ylanase whose activity remained in the supernatant (≈20%) for he subsequent period. Immobilization of proteins also proceeded t lower rate and after 16 h, 20% of the proteins still remained in he supernatant. In sequence, three derivatives were prepared by carrying out mmobilization for 4, 8 and 16 h in order to verify the enzyme ctivities in the derivatives obtained after these three immo- ilization periods (Table 3). As previously observed in Fig. 2, mmobilization yield in relation to protein and enzymatic activities ncreased by increasing the immobilization period, nevertheless he best result in terms of expressed activities was obtained with he derivative prepared by proceeding immobilization during 8 h Table 3). These results indicated that enzyme–support reaction or more prolonged periods may be causing distortion of the nzyme structures, resulting in activity loss. Thus, the selected upport for the co-immobilization of the xylanase, �-xylosidase nd �-l-arabinofuranosidase from P. janczewskii was agarose 10% rosslinking activated with MANAE cross-linked with 0.5% (v/v) lutaraldehyde and by carrying out immobilization at pH 7.0 for h. In order to obtain covalent bonds from Schiff base formed etween protein lysines and aldehyde groups of glutaralde- yde, the support was incubated in pH 8.5 and reduced with 7.0 and 0.05 M Tris-HCl buffer pH 8.0. (b) Enzymatic activities were assayed 0.05 M sodium acetate buffer pH 4.0. Relative (%) xylanase (�), �-xylosidase (�) and �-l- arabinofuranosidase (�) activities. sodium borohydride. The pH increase and reduction in itself negatively influenced the enzymatic activities, i.e., after the pro- cedure, xylanase and �-xylosidase activities were 50 and 90% reduced, respectively, and no �-l-arabinofuranosidase activity was detected on the support. Due to these results, the non-reduced Glut derivative was submitted to post-immobilization techniques (Table 4null, i.e., coating with PEI or dextran-aldehyde (and fur- ther reduced), and it was also submitted to a second round of cross-linking with glutaraldehyde (further reduced or not). Coating with the ionic polymer PEI had a strong negative influence and the three enzyme activities were totally depleted from the biocatalyst. Dextran-aldehyde coating and the maintenance of pH 7.0 during borohydride reduction resulted in residual activity corresponded to 66.9, 57.0 and 55.5% of those previously observed for the xylanase, �-xylosidase and �-arabinofuranosidase, respectively. When the derivative was submitted to a second round of glutaraldehyde crosslinking it presented residual activity of 39.6, 21.7 and 47.8% the xylanase, �-xylosidase and �-l-arabinofuranosidase, respec- tively. If this latter derivative was coated with dextran-aldehyde and then reduced, the residual activity was even lower, corre- 620 C.R. Fanchini Terrasan et al. / Process Biochemistry 51 (2016) 614–623 R es id u al a ct iv it y (% ) 0 20 40 60 80 100 Time (h) 20 864 Fig. 6. Thermostability of xylanase, �-xylosidase and �-l-arabinofuranosidase from P. janczewskii co-immobilized on Glut derivative coated with dextran MW 6000 at 40% degree of oxidation. Incubation was carried out in 0.05 M glycine HCl buffer pH 3.0 at 50 ◦C for the �-xylosidase and �-l-arabinofuranosidase, and at 80 ◦C for the xylanase. Enzymatic activities were assayed in 0.05 M sodium acetate buffer pH 4.0 at 25 ◦C. Initial enzyme activities were regarded as 100%. Residual (%) xylanase (�), �-xylosidase (�) and �-l-arabinofuranosidase (�) activities. Table 6 Hydrolysis of wheat and oat spelt arabinoxylans by multienzymatic derivative of xylanolytic enzymes from P. janczewskii. Time (h) Xylose yield (%) WAX OSX 0.25 0.8 1.2 0.5 1.4 1.9 1 2.3 2.9 2 5.0 5.8 4 9.2 9.5 8 16.9 15.1 24 28.9 26.8 48 41.0 37.4 72 43.5 42.8 Hydrolysis was carried out by incubating 0.05 g of Glut derivative coated with dextran MW 6000 at 40% degree of oxidation with 10 mL of 0.5% (w/v) substrate pre- pared in 0.05 M sodium acetate buffer pH 4.0 at 40 ◦C. WAX—wheat arabinoxylan; O s � w t v e p a p a a a t r t v v s m X yl o se ( m g m l -1 ) 0 1 2 3 Time (h) 100 20 30 40 50 60 70 80 Fig. 7. Hydrolysis of wheat and oat spelt arabinoxylans by multienzymatic deriva- tive of xylanolytic enzymes from P. janczewskii. Hydrolysis was carried out by incubating 0.05 g of Glut derivative coated with dextran MW 6000 at 40% degree of oxidation with 10 mL of 0.5% (w/v) substrate prepared in 0.05 M sodium acetate ◦ When the stability of the xylanase in the derivatives coated with SX—oat spelt xylan. ponding to 15.8, 7.4 and 10.9% for the xylanase, �-xylosidase and -l-arabinofuranosidase, respectively. When the Glut derivative coated with dextran (and reduced) as incubated in 0.005 M sodium phosphate buffer pH 7.0 con- aining high salt concentration (0.5 M), no enzyme activity was erified in the supernatant after 1 h incubation, indicating that the nzymes were covalently immobilized in the support. The pro- osed mechanism of enzyme immobilization on glutaraldehyde as bifunctional support involves initial immobilization occurring by hysical adsorption due to the presence of primary amino groups nd then the covalent immobilization rapidly occurs through the ldehyde portion of glutaraldehyde [34]. The use of neutral pH nd completely oxidized aldehyde-dextran for coating the deriva- ive surface not only preserved the enzyme activities during the eduction but may also provide rigidification of the enzyme struc- ures due to stronger attachment of the enzymes to the support ia covalent bonds. It may have the additional advantage of pre- enting subunit dissociation of enzymes [35], specially considering ome xylanolytic enzymes, mainly �-xylosidases, which may be ultimeric enzymes as cited by [31]. buffer pH 4.0 at 40 C. Xylose released (mg/mL) from oat spelt (�) and wheat (©) arabinoxylans. In the subsequent step, dextrans of different MW (6000 40,000 and 100,000 Da; 100% degree of oxidation) were evaluated for coat- ing the Glut derivative in order to improve enzyme activity and stabilization (Table 5). Coating the derivative with dextran MW 100,000 negatively influenced the immobilized enzymes and their activities were greatly reduced. Coating with dextran MW 6000 resulted in similar xylanase and �-xylosidase activities and half of the �-l-arabinofuranosidase activity was observed in relation to the derivative coated with dextran MW 40,000. A previous stability study, however, demonstrated that the xylanase was more ther- mostable in the derivative coated with dextran MW 6000 than with dextran MW 40,000 (as further presented). In this sense, derivatives coated with dextran MW 6000 at 20 and 40% degree of oxidation (DO) were prepared and the expressed activities were compared to those from the derivative coated with the completely oxidized dex- tran. Among them, the lowest enzyme levels were observed in the derivative coated with dextran at 20% DO. Similar levels of xylanase and �-xylosidase were observed in the derivatives coated with dex- trans at 40 and 100% DO, while the �-l-arabinofuranosidase activity was substantially higher in the derivative coated with dextran at 40% DO (Table 5). 3.2. Derivative characterization After selecting the most adequate support and optimiz- ing immobilization conditions, the stability of the immobilized enzymes in the Glut derivative coated with dextrans MW 6000 and 40,000 Da (completely oxidized) was initially evaluated (Fig. 3). The immobilized xylanase was highly stabilized, retaining 80% of activ- ity in the case of the derivative coated with dextran MW 6000, and around 60% in the derivative coated with dextran MW 40,000. Sim- ilar activity was observed even after 24 h incubation (shown up to 8 h). The �-xylosidase coated with dextran MW 6000 was only a little more stable than that coated with dextran 40,000; while for the �-l-arabinofuranosidase no differences were observed. dextran MW 6000 at different DO was evaluated at different pH and 50 ◦C (Fig. 4), it was observed that coating the biocatalyst sur- face with this polymer stabilized the enzymes in all pH. Among the cess Biochemistry 51 (2016) 614–623 621 d d e w t w W l t a s h m i I o a l p o p g r e e c b y c p � a a o o a t t x r o t � t t � b r t a a 3 h a d a h o p R es id u al a ct iv it y (% ) 0 20 40 60 80 100 (a) (b) Initia l a ctiv ity 5th cy cle 10th cy cle R ed u ci n g s u g ar s (% ) 0 20 40 60 80 100 Time (min.) 200 40 60 Fig. 8. Operational stability (a) and product release (b) from multienzymatic deriva- tive of xylanolytic enzymes from P. janczewskii. Successive hydrolysis 1 h-cycles were carried out by incubating 0.05 g of Glut derivative coated with dextran MW 6000 at 40% degree of oxidation with 10 mL of 0.5% wheat arabinoxylan prepared in 0.05 M sodium acetate buffer pH 4.0 at 40 ◦C. (a) Enzymatic activities were assayed in 0.05 M sodium acetate buffer pH 4.0 at 25 ◦C. Enzymatic activities before the first cycle were regarded as 100%. Residual (%) xylanase (black), �-xylosidase (ligh grey) and �-l-arabinofuranosidase (dark grey) activities after five and ten consecutive C.R. Fanchini Terrasan et al. / Pro extrans oxidized at different degrees, the derivative coated with extran at 20% DO presented the lowest stability, intermediate lev- ls were verified with dextran at 100% DO and the highest stability as observed with the 40% DO, especially in the pH range from 3.0 o 5.0, in which more than 85% of the activity was retained. Dextran coating as a post-immobilization technique has been idely used in enzyme technology for many different enzymes. hen dextran at low degree of oxidation is used only some cova- ent attachments are formed between enzyme and the polymer, and he residual sugar chains remain attached rendering non-natural nd large sugar moiety for the enzyme, i.e., the chemical glyco- ylation of a protein [36]. On the other hand, when the dextran is ighly oxidized, it results in the formation of many covalent attach- ents between enzyme and support rigidifying enzyme structure, mproving stabilization with consequent thermostabilization [25]. n this sense, some stabilization was achieved by glycosylation f the enzymes as observed for the xylanase from P. janczewskii, lthough it was not stabilized in a wide pH range. Additional cova- ent attachments rendered more stability to the enzyme at different H, but when excessive it results in loss of activity as previously bserved in Table 5. Thus, the best results were obtained with artially oxidized dextran, which corresponds to a mix of both lycosylation and an intermediate level of covalent attachments, endering the highest stabilization to the co-immobilized enzymes. Considering the stability of the immobilized xylanase in differ- nt pH and also that the derivative coated with dextran at 40% DO xpressed good levels of the three enzyme activities, the derivative oated with dextran MW 6000 at 40% DO was the most promising, eing further characterized and applied for arabinoxylans hydrol- sis. When the enzymatic reactions were carried out at different onditions, the immobilized xylanase and �-l-arabinofuranosidase resented optimum activity in pH between 2.0 and 4.0, and the -xylosidase in pH 3.0 (Fig. 5a). The immobilized xylanase, �-l- rabinofuranosidase and �-xylosidase presented optimum activity t 50, 65 and 80 ◦C, respectively (Fig. 5b). A shift in the pH for ptimum activity is observed since the free xylanase presented ptimum activity at pH 5.0 and both free �-xylosidase and �-l- rabinofuranosidase were optimally active at pH 4.0. The optimum emperature of the xylanase was similar to that observed for he free-enzyme, while for the �-l-arabinofuranosidase and �- ylosidase the optimum temperature was increased by 5 ◦C in elation to those previously observed for the free enzymes [28,29]. The xylanase was highly thermostable retaining 83.1 and 70% f activity even after 24 h incubation at 60 and 70 ◦C, respec- ively (not shown). At 80 ◦C, its half-life was 1.7 h (Fig. 6). The -xylosidase and the �-l-arabinofuranosidase were very stable up o 40 ◦C, retaining 53 and 100% of the activity after 24 h incuba- ion (not shown). At 50 ◦C, the half-lives of the �-xylosidase and -l-arabinofuranosidase were 2.3 and 3.8 h (Fig. 6). The immo- ilized �-l-arabinofuranosidase is therefore 23-fold stabilized in elation to its free counterpart [29]. The higher thermostability of he xylanase may be associated to its possible monomeric structure nd lower MW that resulted in more attachments to the support nd a more adequate coating by the dextran. .3. Hydrolysis of arabinoxylans The synergistic action of xylanolytic enzymes, divided into omeosynergy, occurring between main-chain cleaving enzymes, nd heterosynergy, occurring between main-chain cleaving and ebranching enzymes, has been reported by using free- [37] and t a lesser extent by using immobilized enzymes [8]. In this ydrolysis study a low viscosity wheat arabinoxylan (WAX) and at spelt xylan (OSX) were used. WAX is a highly arabinosylated olymer containing 41% arabinose and 59% xylose residues, accord- cycles. (b) Reducing sugars (%) released from the first (�), fifth (�) and tenth (�) cycles. ing to the supplier. OSX holds a complex structure containing 81.4% xylose, 9.7% arabinose, 3.4% glucose, 1.2% galactose and 4.3% uronic acid residues [38]. The presence of substituents intensely limits the action of endo-xylanase, hampering the complete degra- dation of the polymers into their monomers. The functionality of the multienzymatic derivative was evaluated by hydrolyz- ing these substrates in pH 4.0 at 40 ◦C during 72 h (Fig. 7). As previously observed, at these conditions the xylanase and the �-l-arabinofuranosidase were very active, and the �-xylosidase operate with 65% of its maximum activity; at this temperature the enzymes do not operate with maximum activity, nevertheless, they present high stability to operate for prolonged cycles. The prepared biocatalyst was active and the similar hydrolysis profiles revealed that they equally degraded both substrates, independent of struc- ture and composition. After 8 h, the reaction velocity is reduced probably due to the inhibitory effect of xylose or arabinose on enzyme activities since the enzymes were stable in the reaction conditions. The maximum xylose yield verified after 72 h corre- sponded to 43.5 and 42.8% for WAX and OSX, respectively (Table 6). The application of a biocatalyst in industry requires the sta- bility of immobilized enzymes through many operational cycles. After consecutive cycles of wheat arabinoxylan hydrolysis, the 6 cess B x i � c a r c 4 x s a i t m d e b h t s c t i o o d f i s l R [ [ [ [ [ [ [ [ [ [ [ [ [ [ [ [ [ [ [ [ [ [ [ [ [ 22 C.R. Fanchini Terrasan et al. / Pro ylanase and the �-l-arabinofuranosidase were very stable retain- ng 86.6 and 88.0% of the activity even after 10 reuse cycles. The -xylosidase activity decreased to 78.7 and 47% after five and ten ycles, respectively (Fig. 8a). The product release decreased to 80 nd 60% after five and ten cycles, respectively, that may be mainly elated to the reduction in the �-xylosidase activity during the ycles (Fig. 8b). . Conclusions The simultaneous co-immobilization of the crude xylanase, �- ylosidase and �-l-arabinofuranosidase from P. janczewskii was not uch a simple procedure, but by adequately selecting the support nd by optimizing parameters such as agarose cross-linking and mmobilization period, the amount of each enzyme in the deriva- ive can be considerably increased, allowing the preparation of a ultienzymatic biocatalyst acting cooperatively in the complete egradation of complex substrates. By selecting conditions, the xpressed activity of the xylanase, the most difficult enzyme to e immobilized, in the glutaraldehyde derivative can be two-fold igher than initially verified. By optimizing the dextran coating, he activity and the stability of the enzymes could also be sub- tantially increased. The mildly activated glutaraldehyde derivative oated with low molecular weight partially oxidized dextran is he most promising, presenting interesting properties, and offer- ng advantages over the use of free enzymes. It is possible to btain a very stable derivative, with improved temperature, pH and perational stability. Furthermore, the use of this multi-enzymatic erivative leads to the direct formation of xylose monosaccharide rom different arabinoxylans, therefore representing an encourag- ng alternative for degradation of hemicelluloses since it can be uccessively reused thereby reducing the need for new enzyme oads and consequently reducing the process cost. eferences [1] B. Saha, Hemicellulose bioconversion, J. Ind. Microbiol. Biotechnol. 30 (2003) 279–291, http://dx.doi.org/10.1007/s10295-003-0049-x. [2] H.V. Scheller, P. Ulvskov, Hemicelluloses, Annu. Rev. Plant Biol. 61 (2010) 263–289, http://dx.doi.org/10.1146/annurev-arplant-042809-112315. [3] D.B. Jordan, K. Wagschal, Properties and applications of microbial�-d-xylosidases featuring the catalytically efficient enzyme from Selenomonas ruminantium, Appl. Microbiol. Biotechnol. 86 (2010) 1647–1658, http://dx.doi.org/10.1007/s00253-010-2538-y. [4] D. Dodd, I.K.O. Cann, Enzymatic deconstruction of xylan for biofuel production, Global Change Biol. Bioenergy 1 (2009) 2–17, http://dx.doi.org/ 10.1111/j.1757-1707.2009.01004.x. [5] A.K.M.S. Rahman, N. Sugitani, M. Hatsu, K. Takamizawa, A role of xylanase, alpha-l-arabinofuranosidase, and xylosidase in xylan degradation, Can. J. Microbiol. 49 (2003) 58–64, http://dx.doi.org/10.1139/w02-114. [6] R.A. Sheldon, S. van Pelt, Enzyme immobilisation in biocatalysis: why, what and how, Chem. Soc. Rev. 42 (2013) 6223–6235, http://dx.doi.org/10.1039/ c3cs60075k. [7] L. Betancor, H. Luckarift, Co-immobilized coupled enzyme systems in biotechnology, Biotechnol. Genet. Eng. Rev. 27 (2010) 95–114, http://dx.doi. org/10.1080/02648725.2010.10648146. [8] C.R.F. Terrasan, E.P. Cipolatti, L.T.A. Souza, R.O. Henriques, S. Moreno-Perez, W.G. Morais Junior, A.O. Chioma, J.M. Guisan, B.C. Pessela, Immobilization of plant cell wall degrading enzymes, in: R.N. Silva (Ed.), Mycol. Curr. Futur. Dev., Bentham Science Publishers B.V., 2015, pp. 276–315, http://dx.doi.org/10. 2174/97816810807411150101. [9] M. Guerfali, I. Maalej, A. Gargouri, H. Belghith, Catalytic properties of the immobilized Talaromyces thermophilus �-xylosidase and its use for xylose and xylooligosaccharides production, J. Mol. Catal. B Enzym. 57 (2009) 242–249, http://dx.doi.org/10.1016/j.molcatb.2008.09.011. 10] A.R.L. Damásio, B.C. Pessela, T.M. da Silva, L.H.S. Guimarães, J.A. Jorge, J.M. Guisan, et al., Co-immobilization of fungal endo-xylanase and �-l-arabinofuranosidase in glyoxyl agarose for improved hydrolysis of arabinoxylan, J. Biochem. 154 (2013) 275–280, http://dx.doi.org/10.1093/jb/ mvt053. 11] T. Hashimoto, Y. Nakata, Synergistic degradation of arabinoxylan with �-l-arabinofuranosidase, xylanase and �-xylosidase from soy sauce koji mold, Aspergillus oryzae, in high salt condition, J. Biosci. Bioeng. 95 (2003) 164–169, http://dx.doi.org/10.1016/S1389-1723(03)80123-8. [ iochemistry 51 (2016) 614–623 12] H.J. Vogel, A convenient growth medium for neurospora crassa, Microb. Genet. Bull. 13 (1956) 42–47. 13] C.R.F. Terrasan, B. Temer, M.C.T. Duarte, E.C. Carmona, Production of xylanolytic enzymes by Penicillium janczewskii, Bioresour. Technol. 101 (2010) 4139–4143, http://dx.doi.org/10.1016/j.biortech.2010.01.011. 14] U.K. Laemmli, Cleavage of structural proteins during the assembly of the head of bacteriophage T4, Nature 227 (1970) 680–685, http://dx.doi.org/10.1038/ 227680a0. 15] M.J. Bailey, P. Biely, K. Poutanen, Interlaboratory testing of methods for assay of xylanase activity, J. Biotechnol. 23 (1992) 257–270, http://dx.doi.org/10. 1016/0168-1656(92)90074-J. 16] G.L. Miller, Use of dinitrosalicylic acid reagent for determination of reducing sugar, Anal. Chem. 31 (1959) 426–428, http://dx.doi.org/10.1021/ ac60147a030. 17] H. Kersters-Hilderson, M. Claeyssens, E. Van Doorslaer, E. Saman, C.K. De Bruyne, Complex Carbohydrates Part D, vol. 83, Elsevier, 1982, http://dx.doi. org/10.1016/0076-6879(82)83062-0. 18] R. Fernandez-Lafuente, C.M. Rosell, V. Rodriguez, C. Santana, G. Soler, A. Bastida, et al., Preparation of activated supports containing low pK amino groups. A new tool for protein immobilization via the carboxyl coupling method, Enzyme Microb. Technol. 15 (1993) 546–550, http://dx.doi.org/10. 1016/0141-0229(93)90016-U. 19] L. Betancor, F. López-Gallego, N. Alonso-Morales, G. Dellamora, C. Mateo, R. Fernandez-Lafuente, et al., Glutaraldehyde in protein immobilization, in: J. Guisan (Ed.), Immobil. Enzym. Cells SE—5, vol. 22, Humana Press, 2006, pp. 57–64, http://dx.doi.org/10.1007/978-1-59745-053-9 5. 20] F. López-Gallego, L. Betancor, C. Mateo, A. Hidalgo, N. Alonso-Morales, G. Dellamora-Ortiz, et al., Enzyme stabilization by glutaraldehyde crosslinking of adsorbed proteins on aminated supports, J. Biotechnol. 119 (2005) 70–75, http://dx.doi.org/10.1016/j.jbiotec.2005.05.021. 21] C. Mateo, J.M. Palomo, M. Fuentes, L. Betancor, V. Grazu, F. López-Gallego, et al., Glyoxyl agarose: a fully inert and hydrophilic support for immobilization and high stabilization of proteins, Enzyme Microb. Technol. 39 (2006) 274–280, http://dx.doi.org/10.1016/j.enzmictec.2005.10.014. 22] C. Mateo, B.C. Pessela, M. Fuentes, R. Torres, C. Ortiz, F. López-Gallego, et al., Very strong but reversible immobilization of enzymes on supports coated with ionic polymers, in: J. Guisan (Ed.), Immobil. Enzym. Cells SE—18, vol. 22, Humana Press, 2006, pp. 205–216, http://dx.doi.org/10.1007/978-1-59745- 053-9 18. 23] C. Mateo, R. Torres, G. Fernández-Lorente, C. Ortiz, M. Fuentes, A. Hidalgo, et al., Epoxy-amino groups: a new tool for improved immobilization of proteins by the epoxy method, Biomacromolecules 4 (2003) 772–777, http:// dx.doi.org/10.1021/bm0257661. 24] J.M. Bolivar, C. Mateo, V. Grazu, A.V. Carrascosa, B.C. Pessela, J.M. Guisan, Heterofunctional supports for the one-step purification, immobilization and stabilization of large multimeric enzymes: amino-glyoxyl versus amino-epoxy supports, Process Biochem. 45 (2010) 1692–1698, http://dx.doi. org/10.1016/j.procbio.2010.07.001. 25] L. Betancor, F. López-Gallego, A. Hidalgo, N. Alonso-Morales, M. Fuentes, R. Fernández-Lafuente, et al., Prevention of interfacial inactivation of enzymes by coating the enzyme surface with dextran-aldehyde, J. Biotechnol. 110 (2004) 201–207, http://dx.doi.org/10.1016/j.jbiotec.2004.02.003. 26] J.J. Sedmak, S.E. Grossberg, A rapid, sensitive, and versatile assay for protein using Coomassie brilliant blue G250, Anal. Biochem. 79 (1977) 544–552, http://dx.doi.org/10.1016/0003-2697(77)90428-6. 27] S.M. Tauk-Tornisielo, A. Garlipp, M. Ruegger, D.S. Attili, E. Malagutti, Soilborne filamentous fungi in Brazil, J. Basic Microbiol. 45 (2005) 72–82, http://dx.doi. org/10.1002/jobm.200410418. 28] C.R.F. Terrasan, B. Temer, C. Sarto, F.G. Silva Junior, E.C. Carmona, Xylanase and �-xylosidase from Penicillium janczewskii: production, physico-chemical properties, and application of the crude extract to pulp biobleaching, BioResources 8 (2013) 1292–1305. 29] B. Temer, C.R.F. Terrasan, E.C. Carmona, �-l-arabinofuranosidase from Penicillium janczewskii: production with brewers spent grain and orange waste, Afr. J. Biotechnol. 13 (2014) 1796–1806, http://dx.doi.org/10.5897/ AJB2013.13361. 30] R.A. Sheldon, Enzyme Immobilization: the quest for optimum performance, Adv. Synth. Catal. 349 (2007) 1289–1307, http://dx.doi.org/10.1002/adsc. 200700082. 31] A. Knob, C.R.F. Terrasan, E.C. Carmona, �-xylosidases from filamentous fungi: an overview, World J. Microbiol. Biotechnol. 26 (2010) 389–407, http://dx.doi. org/10.1007/s11274-009-0190-4. 32] B.C. Saha, �-l-arabinofuranosidases, Biotechnol. Adv. 18 (2000) 403–423, http://dx.doi.org/10.1016/S0734-9750(00) 00044-6. 33] O. Barbosa, C. Ortiz, A. Berenguer-Murcia, R. Torres, R.C. Rodrigues, R. Fernandez-Lafuente, Glutaraldehyde in bio-catalysts design: a useful crosslinker and a versatile tool in enzyme immobilization, RSC Adv. 4 (2014) 1583–1600, http://dx.doi.org/10.1039/C3RA45991H. 34] L. Betancor, F. López-Gallego, A. Hidalgo, N. Alonso-Morales, G. Dellamora-Ortiz, J.M. Guisán, et al., Preparation of a very stable immobilized biocatalyst of glucose oxidase from Aspergillus niger, J. Biotechnol. 121 (2006) 284–289, http://dx.doi.org/10.1016/j.jbiotec.2005.07.014. 35] R. Fernandez-Lafuente, Stabilization of multimeric enzymes: strategies to prevent subunit dissociation, Enzyme Microb. Technol. 45 (2009) 405–418, http://dx.doi.org/10.1016/j.enzmictec.2009.08.009. dx.doi.org/10.1007/s10295-003-0049-x dx.doi.org/10.1007/s10295-003-0049-x dx.doi.org/10.1007/s10295-003-0049-x dx.doi.org/10.1007/s10295-003-0049-x dx.doi.org/10.1007/s10295-003-0049-x dx.doi.org/10.1007/s10295-003-0049-x dx.doi.org/10.1007/s10295-003-0049-x dx.doi.org/10.1007/s10295-003-0049-x dx.doi.org/10.1007/s10295-003-0049-x dx.doi.org/10.1007/s10295-003-0049-x dx.doi.org/10.1146/annurev-arplant-042809-112315 dx.doi.org/10.1146/annurev-arplant-042809-112315 dx.doi.org/10.1146/annurev-arplant-042809-112315 dx.doi.org/10.1146/annurev-arplant-042809-112315 dx.doi.org/10.1146/annurev-arplant-042809-112315 dx.doi.org/10.1146/annurev-arplant-042809-112315 dx.doi.org/10.1146/annurev-arplant-042809-112315 dx.doi.org/10.1146/annurev-arplant-042809-112315 dx.doi.org/10.1146/annurev-arplant-042809-112315 dx.doi.org/10.1146/annurev-arplant-042809-112315 dx.doi.org/10.1007/s00253-010-2538-y dx.doi.org/10.1007/s00253-010-2538-y dx.doi.org/10.1007/s00253-010-2538-y dx.doi.org/10.1007/s00253-010-2538-y dx.doi.org/10.1007/s00253-010-2538-y dx.doi.org/10.1007/s00253-010-2538-y dx.doi.org/10.1007/s00253-010-2538-y dx.doi.org/10.1007/s00253-010-2538-y dx.doi.org/10.1007/s00253-010-2538-y dx.doi.org/10.1007/s00253-010-2538-y dx.doi.org/10.1111/j.1757-1707.2009.01004.x dx.doi.org/10.1111/j.1757-1707.2009.01004.x dx.doi.org/10.1111/j.1757-1707.2009.01004.x dx.doi.org/10.1111/j.1757-1707.2009.01004.x dx.doi.org/10.1111/j.1757-1707.2009.01004.x dx.doi.org/10.1111/j.1757-1707.2009.01004.x dx.doi.org/10.1111/j.1757-1707.2009.01004.x dx.doi.org/10.1111/j.1757-1707.2009.01004.x dx.doi.org/10.1111/j.1757-1707.2009.01004.x dx.doi.org/10.1111/j.1757-1707.2009.01004.x dx.doi.org/10.1111/j.1757-1707.2009.01004.x dx.doi.org/10.1111/j.1757-1707.2009.01004.x dx.doi.org/10.1139/w02-114 dx.doi.org/10.1139/w02-114 dx.doi.org/10.1139/w02-114 dx.doi.org/10.1139/w02-114 dx.doi.org/10.1139/w02-114 dx.doi.org/10.1139/w02-114 dx.doi.org/10.1139/w02-114 dx.doi.org/10.1139/w02-114 dx.doi.org/10.1039/c3cs60075k dx.doi.org/10.1039/c3cs60075k dx.doi.org/10.1039/c3cs60075k dx.doi.org/10.1039/c3cs60075k dx.doi.org/10.1039/c3cs60075k dx.doi.org/10.1039/c3cs60075k dx.doi.org/10.1039/c3cs60075k dx.doi.org/10.1080/02648725.2010.10648146 dx.doi.org/10.1080/02648725.2010.10648146 dx.doi.org/10.1080/02648725.2010.10648146 dx.doi.org/10.1080/02648725.2010.10648146 dx.doi.org/10.1080/02648725.2010.10648146 dx.doi.org/10.1080/02648725.2010.10648146 dx.doi.org/10.1080/02648725.2010.10648146 dx.doi.org/10.1080/02648725.2010.10648146 dx.doi.org/10.1080/02648725.2010.10648146 dx.doi.org/10.2174/97816810807411150101 dx.doi.org/10.2174/97816810807411150101 dx.doi.org/10.2174/97816810807411150101 dx.doi.org/10.2174/97816810807411150101 dx.doi.org/10.2174/97816810807411150101 dx.doi.org/10.2174/97816810807411150101 dx.doi.org/10.2174/97816810807411150101 dx.doi.org/10.1016/j.molcatb.2008.09.011 dx.doi.org/10.1016/j.molcatb.2008.09.011 dx.doi.org/10.1016/j.molcatb.2008.09.011 dx.doi.org/10.1016/j.molcatb.2008.09.011 dx.doi.org/10.1016/j.molcatb.2008.09.011 dx.doi.org/10.1016/j.molcatb.2008.09.011 dx.doi.org/10.1016/j.molcatb.2008.09.011 dx.doi.org/10.1016/j.molcatb.2008.09.011 dx.doi.org/10.1016/j.molcatb.2008.09.011 dx.doi.org/10.1016/j.molcatb.2008.09.011 dx.doi.org/10.1016/j.molcatb.2008.09.011 dx.doi.org/10.1093/jb/mvt053 dx.doi.org/10.1093/jb/mvt053 dx.doi.org/10.1093/jb/mvt053 dx.doi.org/10.1093/jb/mvt053 dx.doi.org/10.1093/jb/mvt053 dx.doi.org/10.1093/jb/mvt053 dx.doi.org/10.1093/jb/mvt053 dx.doi.org/10.1093/jb/mvt053 dx.doi.org/10.1016/S1389-1723(03)80123-8 dx.doi.org/10.1016/S1389-1723(03)80123-8 dx.doi.org/10.1016/S1389-1723(03)80123-8 dx.doi.org/10.1016/S1389-1723(03)80123-8 dx.doi.org/10.1016/S1389-1723(03)80123-8 dx.doi.org/10.1016/S1389-1723(03)80123-8 dx.doi.org/10.1016/S1389-1723(03)80123-8 dx.doi.org/10.1016/S1389-1723(03)80123-8 dx.doi.org/10.1016/S1389-1723(03)80123-8 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0060 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0060 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0060 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0060 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0060 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0060 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0060 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0060 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0060 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0060 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0060 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0060 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0060 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0060 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0060 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0060 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0060 dx.doi.org/10.1016/j.biortech.2010.01.011 dx.doi.org/10.1016/j.biortech.2010.01.011 dx.doi.org/10.1016/j.biortech.2010.01.011 dx.doi.org/10.1016/j.biortech.2010.01.011 dx.doi.org/10.1016/j.biortech.2010.01.011 dx.doi.org/10.1016/j.biortech.2010.01.011 dx.doi.org/10.1016/j.biortech.2010.01.011 dx.doi.org/10.1016/j.biortech.2010.01.011 dx.doi.org/10.1016/j.biortech.2010.01.011 dx.doi.org/10.1016/j.biortech.2010.01.011 dx.doi.org/10.1016/j.biortech.2010.01.011 dx.doi.org/10.1038/227680a0 dx.doi.org/10.1038/227680a0 dx.doi.org/10.1038/227680a0 dx.doi.org/10.1038/227680a0 dx.doi.org/10.1038/227680a0 dx.doi.org/10.1038/227680a0 dx.doi.org/10.1038/227680a0 dx.doi.org/10.1016/0168-1656(92)90074-J dx.doi.org/10.1016/0168-1656(92)90074-J dx.doi.org/10.1016/0168-1656(92)90074-J dx.doi.org/10.1016/0168-1656(92)90074-J dx.doi.org/10.1016/0168-1656(92)90074-J dx.doi.org/10.1016/0168-1656(92)90074-J dx.doi.org/10.1016/0168-1656(92)90074-J dx.doi.org/10.1016/0168-1656(92)90074-J dx.doi.org/10.1016/0168-1656(92)90074-J dx.doi.org/10.1021/ac60147a030 dx.doi.org/10.1021/ac60147a030 dx.doi.org/10.1021/ac60147a030 dx.doi.org/10.1021/ac60147a030 dx.doi.org/10.1021/ac60147a030 dx.doi.org/10.1021/ac60147a030 dx.doi.org/10.1021/ac60147a030 dx.doi.org/10.1016/0076-6879(82)83062-0 dx.doi.org/10.1016/0076-6879(82)83062-0 dx.doi.org/10.1016/0076-6879(82)83062-0 dx.doi.org/10.1016/0076-6879(82)83062-0 dx.doi.org/10.1016/0076-6879(82)83062-0 dx.doi.org/10.1016/0076-6879(82)83062-0 dx.doi.org/10.1016/0076-6879(82)83062-0 dx.doi.org/10.1016/0076-6879(82)83062-0 dx.doi.org/10.1016/0076-6879(82)83062-0 dx.doi.org/10.1016/0141-0229(93)90016-U dx.doi.org/10.1016/0141-0229(93)90016-U dx.doi.org/10.1016/0141-0229(93)90016-U dx.doi.org/10.1016/0141-0229(93)90016-U dx.doi.org/10.1016/0141-0229(93)90016-U dx.doi.org/10.1016/0141-0229(93)90016-U dx.doi.org/10.1016/0141-0229(93)90016-U dx.doi.org/10.1016/0141-0229(93)90016-U dx.doi.org/10.1016/0141-0229(93)90016-U dx.doi.org/10.1007/978-1-59745-053-9_5 dx.doi.org/10.1007/978-1-59745-053-9_5 dx.doi.org/10.1007/978-1-59745-053-9_5 dx.doi.org/10.1007/978-1-59745-053-9_5 dx.doi.org/10.1007/978-1-59745-053-9_5 dx.doi.org/10.1007/978-1-59745-053-9_5 dx.doi.org/10.1007/978-1-59745-053-9_5 dx.doi.org/10.1007/978-1-59745-053-9_5 dx.doi.org/10.1007/978-1-59745-053-9_5 dx.doi.org/10.1007/978-1-59745-053-9_5 dx.doi.org/10.1007/978-1-59745-053-9_5 dx.doi.org/10.1007/978-1-59745-053-9_5 dx.doi.org/10.1016/j.jbiotec.2005.05.021 dx.doi.org/10.1016/j.jbiotec.2005.05.021 dx.doi.org/10.1016/j.jbiotec.2005.05.021 dx.doi.org/10.1016/j.jbiotec.2005.05.021 dx.doi.org/10.1016/j.jbiotec.2005.05.021 dx.doi.org/10.1016/j.jbiotec.2005.05.021 dx.doi.org/10.1016/j.jbiotec.2005.05.021 dx.doi.org/10.1016/j.jbiotec.2005.05.021 dx.doi.org/10.1016/j.jbiotec.2005.05.021 dx.doi.org/10.1016/j.jbiotec.2005.05.021 dx.doi.org/10.1016/j.jbiotec.2005.05.021 dx.doi.org/10.1016/j.enzmictec.2005.10.014 dx.doi.org/10.1016/j.enzmictec.2005.10.014 dx.doi.org/10.1016/j.enzmictec.2005.10.014 dx.doi.org/10.1016/j.enzmictec.2005.10.014 dx.doi.org/10.1016/j.enzmictec.2005.10.014 dx.doi.org/10.1016/j.enzmictec.2005.10.014 dx.doi.org/10.1016/j.enzmictec.2005.10.014 dx.doi.org/10.1016/j.enzmictec.2005.10.014 dx.doi.org/10.1016/j.enzmictec.2005.10.014 dx.doi.org/10.1016/j.enzmictec.2005.10.014 dx.doi.org/10.1016/j.enzmictec.2005.10.014 dx.doi.org/10.1007/978-1-59745-053-9_18 dx.doi.org/10.1007/978-1-59745-053-9_18 dx.doi.org/10.1007/978-1-59745-053-9_18 dx.doi.org/10.1007/978-1-59745-053-9_18 dx.doi.org/10.1007/978-1-59745-053-9_18 dx.doi.org/10.1007/978-1-59745-053-9_18 dx.doi.org/10.1007/978-1-59745-053-9_18 dx.doi.org/10.1007/978-1-59745-053-9_18 dx.doi.org/10.1007/978-1-59745-053-9_18 dx.doi.org/10.1007/978-1-59745-053-9_18 dx.doi.org/10.1007/978-1-59745-053-9_18 dx.doi.org/10.1007/978-1-59745-053-9_18 dx.doi.org/10.1021/bm0257661 dx.doi.org/10.1021/bm0257661 dx.doi.org/10.1021/bm0257661 dx.doi.org/10.1021/bm0257661 dx.doi.org/10.1021/bm0257661 dx.doi.org/10.1021/bm0257661 dx.doi.org/10.1021/bm0257661 dx.doi.org/10.1016/j.procbio.2010.07.001 dx.doi.org/10.1016/j.procbio.2010.07.001 dx.doi.org/10.1016/j.procbio.2010.07.001 dx.doi.org/10.1016/j.procbio.2010.07.001 dx.doi.org/10.1016/j.procbio.2010.07.001 dx.doi.org/10.1016/j.procbio.2010.07.001 dx.doi.org/10.1016/j.procbio.2010.07.001 dx.doi.org/10.1016/j.procbio.2010.07.001 dx.doi.org/10.1016/j.procbio.2010.07.001 dx.doi.org/10.1016/j.procbio.2010.07.001 dx.doi.org/10.1016/j.procbio.2010.07.001 dx.doi.org/10.1016/j.jbiotec.2004.02.003 dx.doi.org/10.1016/j.jbiotec.2004.02.003 dx.doi.org/10.1016/j.jbiotec.2004.02.003 dx.doi.org/10.1016/j.jbiotec.2004.02.003 dx.doi.org/10.1016/j.jbiotec.2004.02.003 dx.doi.org/10.1016/j.jbiotec.2004.02.003 dx.doi.org/10.1016/j.jbiotec.2004.02.003 dx.doi.org/10.1016/j.jbiotec.2004.02.003 dx.doi.org/10.1016/j.jbiotec.2004.02.003 dx.doi.org/10.1016/j.jbiotec.2004.02.003 dx.doi.org/10.1016/j.jbiotec.2004.02.003 dx.doi.org/10.1016/0003-2697(77)90428-6 dx.doi.org/10.1016/0003-2697(77)90428-6 dx.doi.org/10.1016/0003-2697(77)90428-6 dx.doi.org/10.1016/0003-2697(77)90428-6 dx.doi.org/10.1016/0003-2697(77)90428-6 dx.doi.org/10.1016/0003-2697(77)90428-6 dx.doi.org/10.1016/0003-2697(77)90428-6 dx.doi.org/10.1016/0003-2697(77)90428-6 dx.doi.org/10.1016/0003-2697(77)90428-6 dx.doi.org/10.1002/jobm.200410418 dx.doi.org/10.1002/jobm.200410418 dx.doi.org/10.1002/jobm.200410418 dx.doi.org/10.1002/jobm.200410418 dx.doi.org/10.1002/jobm.200410418 dx.doi.org/10.1002/jobm.200410418 dx.doi.org/10.1002/jobm.200410418 dx.doi.org/10.1002/jobm.200410418 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 http://refhub.elsevier.com/S1359-5113(16)30026-5/sbref0140 dx.doi.org/10.5897/AJB2013.13361 dx.doi.org/10.5897/AJB2013.13361 dx.doi.org/10.5897/AJB2013.13361 dx.doi.org/10.5897/AJB2013.13361 dx.doi.org/10.5897/AJB2013.13361 dx.doi.org/10.5897/AJB2013.13361 dx.doi.org/10.5897/AJB2013.13361 dx.doi.org/10.5897/AJB2013.13361 dx.doi.org/10.1002/adsc.200700082 dx.doi.org/10.1002/adsc.200700082 dx.doi.org/10.1002/adsc.200700082 dx.doi.org/10.1002/adsc.200700082 dx.doi.org/10.1002/adsc.200700082 dx.doi.org/10.1002/adsc.200700082 dx.doi.org/10.1002/adsc.200700082 dx.doi.org/10.1002/adsc.200700082 dx.doi.org/10.1007/s11274-009-0190-4 dx.doi.org/10.1007/s11274-009-0190-4 dx.doi.org/10.1007/s11274-009-0190-4 dx.doi.org/10.1007/s11274-009-0190-4 dx.doi.org/10.1007/s11274-009-0190-4 dx.doi.org/10.1007/s11274-009-0190-4 dx.doi.org/10.1007/s11274-009-0190-4 dx.doi.org/10.1007/s11274-009-0190-4 dx.doi.org/10.1007/s11274-009-0190-4 dx.doi.org/10.1007/s11274-009-0190-4 dx.doi.org/10.1016/S0734-9750(00) 00044-6 dx.doi.org/10.1016/S0734-9750(00) 00044-6 dx.doi.org/10.1016/S0734-9750(00) 00044-6 dx.doi.org/10.1016/S0734-9750(00) 00044-6 dx.doi.org/10.1016/S0734-9750(00) 00044-6 dx.doi.org/10.1016/S0734-9750(00) 00044-6 dx.doi.org/10.1016/S0734-9750(00) 00044-6 dx.doi.org/10.1016/S0734-9750(00) 00044-6 dx.doi.org/10.1016/S0734-9750(00) 00044-6 dx.doi.org/10.1039/C3RA45991H dx.doi.org/10.1039/C3RA45991H dx.doi.org/10.1039/C3RA45991H dx.doi.org/10.1039/C3RA45991H dx.doi.org/10.1039/C3RA45991H dx.doi.org/10.1039/C3RA45991H dx.doi.org/10.1039/C3RA45991H dx.doi.org/10.1016/j.jbiotec.2005.07.014 dx.doi.org/10.1016/j.jbiotec.2005.07.014 dx.doi.org/10.1016/j.jbiotec.2005.07.014 dx.doi.org/10.1016/j.jbiotec.2005.07.014 dx.doi.org/10.1016/j.jbiotec.2005.07.014 dx.doi.org/10.1016/j.jbiotec.2005.07.014 dx.doi.org/10.1016/j.jbiotec.2005.07.014 dx.doi.org/10.1016/j.jbiotec.2005.07.014 dx.doi.org/10.1016/j.jbiotec.2005.07.014 dx.doi.org/10.1016/j.jbiotec.2005.07.014 dx.doi.org/10.1016/j.jbiotec.2005.07.014 dx.doi.org/10.1016/j.enzmictec.2009.08.009 dx.doi.org/10.1016/j.enzmictec.2009.08.009 dx.doi.org/10.1016/j.enzmictec.2009.08.009 dx.doi.org/10.1016/j.enzmictec.2009.08.009 dx.doi.org/10.1016/j.enzmictec.2009.08.009 dx.doi.org/10.1016/j.enzmictec.2009.08.009 dx.doi.org/10.1016/j.enzmictec.2009.08.009 dx.doi.org/10.1016/j.enzmictec.2009.08.009 dx.doi.org/10.1016/j.enzmictec.2009.08.009 dx.doi.org/10.1016/j.enzmictec.2009.08.009 dx.doi.org/10.1016/j.enzmictec.2009.08.009 cess B [ [ C.R. Fanchini Terrasan et al. / Pro 36] M.L.E. Gutarra, O. Romero, O. Abian, F.A.G. Torres, D.M.G. Freire, A.M. Castro, et al., Enzyme surface glycosylation in the solid phase: improved activity and selectivity of Candida antarctica Lipase B, ChemCatChem 3 (2011) 1902–1910, http://dx.doi.org/10.1002/cctc.201100211. 37] J.S. Van Dyk, B.I. Pletschke, A review of lignocellulose bioconversion using enzymatic hydrolysis and synergistic cooperation between enzymes–factors [ iochemistry 51 (2016) 614–623 623 affecting enzymes, conversion and synergy, Biotechnol. Adv. 30 (2012) 1458–1480, http://dx.doi.org/10.1016/j.biotechadv.2012.03.002. 38] F.J.M. Kormelink, a. GJ. Voragen, Degradation of different [(glucurono) arabino]xylans by a combination of purified xylan-degrading enzymes, Appl. Microbiol. Biotechnol. 38 (1993) 688–695, http://dx.doi.org/10.1007/ BF00182811. dx.doi.org/10.1002/cctc.201100211 dx.doi.org/10.1002/cctc.201100211 dx.doi.org/10.1002/cctc.201100211 dx.doi.org/10.1002/cctc.201100211 dx.doi.org/10.1002/cctc.201100211 dx.doi.org/10.1002/cctc.201100211 dx.doi.org/10.1002/cctc.201100211 dx.doi.org/10.1002/cctc.201100211 dx.doi.org/10.1016/j.biotechadv.2012.03.002 dx.doi.org/10.1016/j.biotechadv.2012.03.002 dx.doi.org/10.1016/j.biotechadv.2012.03.002 dx.doi.org/10.1016/j.biotechadv.2012.03.002 dx.doi.org/10.1016/j.biotechadv.2012.03.002 dx.doi.org/10.1016/j.biotechadv.2012.03.002 dx.doi.org/10.1016/j.biotechadv.2012.03.002 dx.doi.org/10.1016/j.biotechadv.2012.03.002 dx.doi.org/10.1016/j.biotechadv.2012.03.002 dx.doi.org/10.1016/j.biotechadv.2012.03.002 dx.doi.org/10.1016/j.biotechadv.2012.03.002 dx.doi.org/10.1007/BF00182811 dx.doi.org/10.1007/BF00182811 dx.doi.org/10.1007/BF00182811 dx.doi.org/10.1007/BF00182811 dx.doi.org/10.1007/BF00182811 dx.doi.org/10.1007/BF00182811 dx.doi.org/10.1007/BF00182811 Co-immobilization and stabilization of xylanase, β-xylosidase and α-l-arabinofuranosidase from Penicillium janczewskii for... 1 Introduction 2 Materials and methods 2.1 Materials 2.2 Methods 2.2.1 Microorganism, enzyme production and preparation of enzyme extract 2.2.2 SDS-PAGE 2.2.3 Enzyme assays 2.2.4 Preparation of support 2.2.5 Enzyme immobilization 2.2.6 Post-immobilization techniques 2.2.7 Immobilization parameters 2.2.8 Evaluation of the attachment between enzyme and support 2.2.9 Derivative characterization 2.2.9.1 Thermal stability 2.2.9.2 pH stability 2.2.9.3 Optima pH and temperature 2.2.10 Hydrolysis of arabinoxylans 2.2.11 Reuse assay 2.2.12 Analytical methods 3 Results and discussion 3.1 Enzyme immobilization 3.2 Derivative characterization 3.3 Hydrolysis of arabinoxylans 4 Conclusions References