Extractive Fermentation of Xylanase from Aspergillus tamarii URM 4634 in a Bioreactor

Extractive Fermentation of Xylanase from Aspergillustamarii URM 4634 in a Bioreactor

Anna Carolina da Silva &

Alana Emília Soares de França Queiroz &

Talita Camila Evaristo dos Santos Nascimento &

Cristine Rodrigues & José Erick Galindo Gomes &Cristina Maria Souza-Motta &

Luciana Porto de Souza Vandenberghe &

Erika Valente de Medeiros & Keila Aparecida Moreira &

Polyanna Nunes Herculano

Received: 31 January 2014 /Accepted: 6 May 2014 /Published online: 22 May 2014# Springer Science+Business Media New York 2014

Abstract Of the many reported applications for xylanase, its use as a food supplement hasplayed an important role for monogastric animals, because it can improve the utilisation ofnutrients. The aim of this work was to produce xylanase by extractive fermentation in anaqueous two-phase system using Aspergillus tamarii URM 4634, increasing the scale ofproduction in a bioreactor, partially characterising the xylanase and evaluating its influenceon monogastric digestion in vitro. Through extractive fermentation in a bioreactor, xylanasewas obtained with an activity of 331.4 U mL−1 and 72 % yield. The xylanase was stable undervariable pH and temperature conditions, and it was optimally active at pH 3.6 and 90 °C.Xylanase activity potentiated the simulation of complete monogastric digestion by 6 %, andonly Mg2+ inhibited its activity. This process provides a system for efficient xylanaseproduction by A. tamarii URM 4634 that has great potential for industrial use.

Appl Biochem Biotechnol (2014) 173:1652–1666DOI 10.1007/s12010-014-0953-8

A. C. da Silva :A. E. Soares de França Queiroz : T. C. Evaristo dos Santos Nascimento : J. E. G. Gomes :E. Valente de Medeiros :K. A. Moreira (*)Unidade Acadêmica de Garanhuns, Universidade Federal Rural de Pernambuco, Av. Bom Pastor, s/n,Boa Vista, CEP 55.292-270 Garanhuns, PE, Brazile-mail: [email protected]

C. Rodrigues : L. Porto de Souza VandenbergheUniversidade Federal do Paraná/Centro Politécnico, Av. Coronel Francisco Heráclito dos Santos, 210-Jardimdas Américas, CEP 81531-970 Curitiba, PR, Brazil

C. M. Souza-MottaDepartamento de Micologia, Universidade Federal de Pernambuco, Av. Moraes Rego, s/n,CEP 50670-901 Recife, PE, Brazil

P. N. HerculanoUniversidade Federal Rural de Pernambuco, Rua Dom Manoel de Medeiros, s/n, Dois Irmãos,CEP 52171-900 Recife, PE, Brazil

Keywords Aqueous two-phase . Bioreactors . Filamentous fungi . Scale up . Xylanolyticcomplex .Monogastric digestion

Introduction

Hemicelluloses are hydrolysed by the cooperative action of a group of enzymes, and themain enzyme involved in the depolymerisation of xylan is endo-β-1,4-xylanase (1). Theβ-1,4-endoxylanases (1,4-β-D-xylan xylohydrolase, EC 3.2.1.8), which are produced byalgae, protozoa, mollusks, crustaceans, insects, the seeds of terrestrial plants, bacteria,and many fungal species belonging to the genus Aspergillus, are commonly found innature (2). However, the xylanase enzymes of filamentous fungi are potentiallyinteresting for industrial use because they are produced in greater amounts comparedwith those of yeast and bacteria (3). These enzymes can be used in monogastric feeds inorder to hydrolyse non-starch polysaccharides such as β-glucan and arabinoxylan thatare found in vegetables, thus increasing feed conversion, body weight gain, and the yieldof production (4, 5).

Of the total cost associated with enzyme production, approximately of 50–80 % isrelated to the downstream processing associated with extracting and purifying theenzyme. This high cost can be attributed to conventional methods of proteinspurification, which are performed in many steps with expensive reagents. Moreover, itis difficult to expand the process to an industrial scale. Therefore, it is necessary todevelop new methods of producing and extracting enzymes at a large scale with low cost(6, 7). There is increasing interest in innovative processes for the separation andpurification of biomolecules that are economically viable and preserve the biologicalactivity of proteins (8, 9).

Extractive fermentation in an aqueous two-phase system is an excellent alternativemethod to decrease the total cost of enzyme production due its production characteristicsand the fact that the extraction occurs simultaneous in situ, removing steps from theenzyme purification process (10); the concept the in situ process involves integrating anextraction step into the first processing stage to simultaneously synthesis and remove theproduct. This not only ensures the primary recovery, but it also increases the rate ofproduct formation by minimizing final product inhibition during fermentation (11). Theapplication of extractive fermentation for the production of xylanase is particularlyinteresting because the production and simultaneous separation of the molecules in thissystem decreases the fermentative action of xylanase inhibitors such as proteases thatdegrade xylanase (12).

The aim of this work was to partially characterise the xylanase produced by Aspergillustamarii URM 4634 through extractive fermentation in a bioreactor.

Material and Methods

Microorganism

A. tamarii URM 4634 was obtained from the URM collection at the Department of Mycology,Federal University of Pernambuco, Brazil, and it was maintained in tubes containing potatodextrose agar. For sporulation, the fungus was grown in Petri dishes containing Czapek agarfor 5 days at 28 °C.

Appl Biochem Biotechnol (2014) 173:1652–1666 1653

Production in Flasks

Xylanase was produced using an extractive fermentation aqueous two-phase system (ATPS) in125 mL Erlenmeyer flasks containing 30 g total ATPS composed of polyethylene glycol,sodium citrate, citric acid, yeast extract (0.5 %), cassava bark (2 %), and water was added to all30 g. The molar mass of polyethylene glycol (MMPEG), concentration of sodium citrate (CC),concentration of PEG (CPEG) and pH of the system varied according to a 24 factorial design(Table 1). The inoculum was prepared at a concentration of 106 spores mL−1, and thefermentation was maintained at 28 °C and 120 rpm for up to 144 h.

The production kinetics of the microorganism was measured in Erlenmeyer flasks using theoptimum conditions for the production of xylanase. Aliquots were removed every 24 h until144 h of fermentation, and the extraction and the following production parameters wereevaluated: xylanase activity, total protein, pH, extraction yield, and partition coefficient.

Production in a Bioreactor

Extractive fermentation using ATPS was performed in a bioreactor (B.E. MARUBISHI®,Pathumthani, Thailand) with a total capacity of 10 L. The ATPS was composed of polyeth-ylene glycol, sodium citrate, citric acid, yeast extract (0.5 %), cassava bark (2 %), and water toachieve a total mass of 5,000 g. In this step, the best conditions for MMPEG, CC, and CPEG

were selected using a factorial design and carried out in shake flasks. The inoculum wasprepared at 106 spores mL−1, and the fermentation continued for 96 h. Aliquots were collectedevery 24 h to determination xylanase activity, total protein, partition coefficient, yield, and pH.Fermentation occurred at 28 °C and 150 rpm, and aeration was forced at 9 L min−1.

Determination of the Total Protein Content in the Enzyme Extract

The Bradford method was used to measure the total protein content (13). The absorbance ofthe solutions was measured at 595 nm using a spectrophotometer (Biochrom Libra S6®,Cambridge, UK).

Enzyme Assay

Xylanase activity was determined according to the method described by Bailey et al. (14), andthe free sugar released was analyzed by the dinitrosalicylic acid method, as described by Miller(15). One unit of enzymatic activity was defined as the amount of enzyme required to produce1 μmol of reducing sugars (measured as respective monosaccharide) per minute by hydrolys-ing the respective crude substrate under the specified assay conditions.

Table 1 Levels of factors used in the 24 factorial experimental design

Levels–coded values Low Central High

−1 0 +1

x1–MMPEG (g mol−1) 2,000 4,000 6,000

x2–CPEG (%) 15 20 25

x3–pH 6.0 7.0 8.0

x4–CC (%) 15 20 25

1654 Appl Biochem Biotechnol (2014) 173:1652–1666

Determination of Partition Coefficient and Yield

The partition coefficient for xylanase activity (kx) and partition coefficient for proteins (kp) inthe aqueous two-phase systems were defined using Eqs. 1 and 2, respectively:

Kx ¼ AT

ABð1Þ

Kp ¼ PT

PBð2Þ

Where AT and AB represent the equilibrium xylanase activities (per milliliter) in the top andbottom phases, and PT and PB represent the equilibrium concentration (milligrams permilliliter) of the proteins in the top and bottom phases, respectively (16, 17).

Xylanase yield was calculated using the following Eq. 3:

Yield 100%ð Þ 100

1þ 1

Vr*mathvariant ¼ }italic} > Kx

� � ð3Þ

Where Vr is the volume ratio of the top phase and the bottom phase (16, 17).

Effect of pH on Enzyme Activity and Stability

The optimum pH for the enzymatic activity of xylanase was determined usingdifferent buffers at 50 mM: glycine–HCl buffer (pH 2.4–3.6), acetate buffer(pH 3.6–5.0), citrate buffer (pH 5.0–18.0), Tris–HCl buffer (pH 6.0–8.0), and gly-cine–NaOH buffer (pH 8.0–10.0). The pH stability was determined by incubating thereaction mixture in the following buffers: glycine–HCl (pH 2.4–3.6), acetate (pH 3.6–5.0), citrate (pH 5.0–18.0), and Tris–HCl (pH 6.0–8.0). Aliquots were taken at 0, 60,120, and 180 min to determine the xylanase activity at each time point (18). Enzymeactivity was measured as described previously.

Effect of Temperature on Enzyme Activity and Stability

Optimum temperature was determined by measuring the xylanase activity of the crude extractat different temperatures in the range of 35–100 °C. To determine heat stability, the enzymewas subjected to temperatures of 30–90 °C. Aliquots were taken at 0, 60, 120, and 180 min todetermine their specific activities. Samples from each time point were subjected to analyticaldeterminations (18).

Effect of the Activation or Inhibition of Metal Ions and Inhibitors on Xylanase Activity

The following ions and inhibitors were tested at concentrations of 1 and 5 mM: ions, Cu2+,Mn2+, Na+, Fe2+, Zn2+, Mg2+, and Ca2+; inhibitors; ethylenediaminetetraacetic acid (EDTA);and 2-mercaptoethanol. The degree of inhibition or activation of xylanase was evaluated bysubjecting the enzyme extract to the ions and inhibitors for 1 h, followed by measurement ofxylanase activity (18).


Kinetic Parameters

For kinetic parameters, five different concentrations of beechwood xylan (4–12 g L−1) withconstant enzyme concentration were prepared. Assays were performed under standard assay.The kinetic parameters (apparent Km and Vmax) were calculated determined from aLineweaver–Burk plot.

Effect of Simulated Upper Monogastric Digestive Tract Conditions on Xylanase Activity

Simulations of monogastric digestion were determined in vitro, as described by Boyce andWalsh (19). Intestinal digestion of the test compound was simulated by adding trypsin and 1 %taurocholic acid for 4 h.

Polyacrylamide Gel Electrophoresis and Zymogram

Polyacrylamide gel electrophoresis (SDS-PAGE) was performed according to the method ofLaemmli (20), using gel concentrations of 4.9 % and 15.4 % to achieve separation. Sampleswere generated in the fermentation bioreactor in conditions that produced the best yield andfermentation at concentrations of 100 and 200 mg mL−1 protein. The molecular mass of thebands was determined using LabImage 1D software (Loccus Biotechnology, Brazil). Thezymogram was performed in a polyacrylamide gel (15.4 %), according to the methoddescribed by Mattéotti et al. (21).

Statistical Analysis

Statistica Version 7.0 (Statsoft, Tulsa, OK) software was used for graphical analyses of thedata. All experiments were performed in triplicate, and the results are expressed as the mean±standard deviation (SD). Effects of the variables and the significance of regression coefficientswere determined by Student's t test (p<0.05).

Results and Discussion

Fermentation Flasks

A. tamarii URM 4634 produced xylanase in all of the 24 factorial design assays. This specieswas reported to be a good producer of xylanase in other studies; however, no study cites theuse of extractive fermentation (22–24).

Table 2 shows the respective kx, kp, and yield values achieved by different variablecombinations. By analyzing the independent variables and using yield as the response variable,it is evident that CC and CPEG have a significant effect on xylanase production. Figure 1 showsthe effects of the independent variables and the effects of each combination. These datademonstrate that CC and CPEG have a negative effect, indicating that lower concentrationsincrease the fermentation yield.

It has been shown that, at higher levels of MMPEG, lower concentrations of polymer arerequired for the formation of phases (25). For PEG 6000, it was possible to reduce theconcentration without affecting the formation phase. Moreover, changes in polymer concen-tration can change the partition coefficient of a molecule. For example, Schmidt et al. (26)


reported a 3.2-fold decrease in the k of α-amylase due to increasing CPEG using a PEG/phosphate system, whereas in a system using PEG 4000/phosphate, increasing the concentra-tion of PEG from 14 % to 20 % increased the partition coefficient of the enzyme 5-fold. Forxylanase produced by A. tamarii URM 4634, the concentration of the polymer did notsignificantly affect the partition.

Ooi et al. (27) studied the production of lipase by Burkholderia pseudomallei usingextractive fermentation, varying the CPEG levels from 5 to 20 % (w/w) and observed thatcell growth was retarded in media containing elevated CPEG. Increased CPEG can raise theviscosity of the medium, reducing oxygen transfer and affecting cell growth (28).

Regarding salt concentrations, Kuboi et al. (29) reported that high concentrations are toxicto microbial cells. This toxicity may be related to osmotic pressure due to dehydration of thecells in hypertonic medium. For this reason, salts are often considered inappropriate forcomposition of the lower phase system. However, studies by Ooi et al. (27) show that a

Table 2 Matrix of the 24 factorial with the results of the production xylanase in ATPS using extractivefermentation by A. tamarii URM 4634

Experimental factors

Run MMPEGa CPEG

b pH CCc Kpd KX

e Yf

1 2,000 15 6.0 15 8.48 0.19 95.2

2 6,000 15 6.0 15 7.20 0.14 96.5

3 2,000 25 6.0 15 5.85 0.12 89.8

4 6,000 25 6.0 15 1.05 1.89 32.7

5 2,000 15 8.0 15 1.41 0.12 89.3

6 6,000 15 8.0 15 3.08 0.16 86.2

7 2,000 25 8.0 15 0.91 0.55 71.8

8 6,000 25 8.0 15 0.90 0.77 64.4

9 2,000 15 6.0 25 1.72 3.21 35.6

10 6,000 15 6.0 25 1.28 0.15 94.0

11 2,000 25 6.0 25 2.57 2.98 25.1

12 6,000 25 6.0 25 0.49 2.27 36.4

13 2,000 15 8.0 25 3.47 1.00 80.0

14 6,000 15 8.0 25 2.01 3.98 20.0

15 2,000 25 8.0 25 1.04 1.51 49.9

16 6,000 25 8.0 25 5.96 2.93 32.9

17 4,000 20 7.0 20 10.37 0.47 62.9

18 4,000 20 7.0 20 9.90 0.55 62.5

19 4,000 20 7.0 20 10.38 0.89 62.7

20 4,000 20 7.0 20 8.21 0.66 64.2

a PEG molar mass (grams per mole)b PEG concentration (%)c Citrate concentration (%)d Partition coefficient of the proteine Partition coefficient of xylanasef Yield of the fermentation


system consisting of PEG/sodium citrate (5 % w/w) was adequate for the production of theenzyme with lipase activity and promoted similar cell growth to that observed with PEG/dextran. For the production of xylanase by A. tamariiURM 4634, low salt concentrations werebetter, but the CC was superior to that reported by Ooi et al. for the production of lipase (27)without damaging the growth of the fungus and the activity of the resulting xylanase.

The amount ofMMPEG used did not have a significant effect. PEG 6000 was included in theassay with increased yield (Run 2, Table 2), so it was used in subsequent stages of fermen-tation. Based on the results obtained by analyzing the 24 factorial design, a new factorialdesign was prepared using sodium citrate and PEG concentrations lower than those previouslytested (Table 3).

Figure 2 demonstrates the effect caused by CPEG and Cc. CPEG has a significant positiveeffect, and CC showed no significant effect on fermentation efficiency. Therefore, the systemconsisting of 14 % PEG 6000 and 14 % sodium citrate was chosen for subsequent stages offermentation.

When assessing production kinetics, we observed that yield decreased over time, and ahigher yield was obtained within 48 h (Fig. 3). The pH changed little over time, varyingbetween 6.17 and 6.2. Up to 72 h, the partition coefficient of the protein was maintained above1 (Fig. 4), indicating that these molecules were in phase with the higher concentration of

Fig. 1 Main effects (% citrate, % PEG,MMPEG, and pH) of the response variable and fermentation efficiency infactorial design 24

Table 3 Levels of factors used in the 22 factorial experimental design

Levels–coded values Low Central High

−1 0 +1

x1–CPEG (%) 12 13 14

x2–CC (%) 12 13 14


polymer (PEG-rich phase). In contrast, the kX value remained was kept below 1during this time (k=0.2 at 48 h of fermentation), indicating that the enzyme wasextracted in the phase rich in sodium citrate. Similar results have been described foralkaline phosphatise (30), where the enzyme partitioned into the salt-rich phase in asystem consisting of 10 % PEG 6000 and sodium citrate 12.9 %, with k similar to thexylanase produced by A. tamarii URM 4634 (k=0.26). However, after 96 h, there wasa reversal of values, and the xylanase remained in the opposite phase compared withother proteins.

Bioreactor Fermentation

In the bioreactor, enzymes behave similarly to those produced in shake flasks, with 72 % yield;however, the enzyme remained in the lower phase throughout the process in the bioreactor andshowed higher xylanase activity than in flasks (141.0 U mL−1 in flasks and 331.4 U mL−1 inbioreactor). Studies show xylanase production by other filamentous fungi such as Aspergillusniger US368 (14.5 U mL−1) (31), Aspergillus candidus (100.23 U mL−1) (32), and Aspergillusfumigatus SK1 (418.70 U g−1) (33). The high activity of xylanase produced by A. fumigatusSK1 might be explained due to the solid fermentation employed at work (33) in contrast to thesubmerged fermentation used in the production of xylanase by A. tamarii URM 4634 andother works cited above, because the solid fermentation may provide a near natural habitat offilamentous fungus to enhance protein secretion (34). About extractive fermentation, Penicil-lium janthinellum produced xylanase with 160.7 U mL−1 of activity in the top phase of thesystem (35).

Although the production of xylanase by extractive fermentation in ATPS has not beenreported, this type of fermentation is used for the production of other enzymes, such asendoglucanase (11), alkaline phosphatise (30), and lipase (27, 32). These studies reportedincreasing the production scale using extractive fermentation in a bioreactor (32), and the

Fig. 2 Main effects (% citrate and % PEG) of the response variable and fermentation efficiency in factorialdesign 22


reported k and fermentation yield values were better than those obtained in shake flasks, likelydue to better homogeneity in the agitation of the medium and forced aeration.

These factors allow for better control of the fermentation environment and more efficientnutrient transport, especially in regard to oxygen. These results suggest that agitation plays acrucial role in the extractive fermentation, especially in the presence of high viscosity. This isdue to the influence of agitation on the rate of oxygen transfer, the concentration of dissolvedoxygen, and the production and recovery of products (36).

The yield obtained by extractive fermentation in bioreactors was considerably higher(72 %) compared with other purification methods, including submerged fermentation in shakeflasks (in which the yield reached 17.5 %), ammonium sulfate precipitation, and

Fig. 3 Curve of the fermentation yield over time

Fig. 4 Curve of the partition coefficient of the protein (filled diamond), xylanase (filled square), and pH curve(filled circle)


chromatography (37), and with similar methodology, other authors achieved an 87 % yield(38). For xylanase produced by Aspergillus nidulans KK-99, a yield of 74.6 % was obtainedby precipitation with ammonium sulphate (38), but the application of other phases of enzymepurification would diminish yield. Other corroborating data were reported by Khanum and Pal(39), who purified xylanase from A. niger DFR-5 in three steps, obtaining a yield of 81.92 %in the first stage, but only 38.9 % by the end of the process.

Characterisation of the Xylanase Produced in a Bioreactor

According to our partial enzyme characterisation, the best conditions for the production ofxylanase were achieved in a bioreactor. Regarding optimal pH, the highest activities wereobserved in the acidic pH range, and its optimum and lowest pH values were 3.6 in sodiumacetate buffer and pH 7.5 in Tris-HCl (61.4 %), respectively (Fig. 5). The fungal xylanasesreported in the literature are optimally active at an acidic (pH 4.5 to 6.5) or neutral pH (5,40–44), but, unlike our results, the xylanase produced by Chaetomium sp. CQ31 had higheractivity at pH 7.5 (37).

pH stability of the enzyme is essential for enzymatic characterisation for commercial use.The xylanase produced by A. tamarii URM 4634 was stable at a range of pH values andremained active at all range of pH tested, even after 180 minutes; the residual activity washigher than 40 % (Fig. 6) and 97.18 % at the lowest and optimum pHs, respectively. Thus, thestability of xylanase in a wide pH range is an important feature that demonstrates the potentialof this method for industrial applications. The xylanase produced by Arthrobacter sp. MTCC5214 was stable over a smaller range of pH values (7.0 to 8.0) (39), while the enzymeproduced by Thermomyces lanuginosus retained over 95 % of its activity between pH 5.5and 9.5 (45). Furthermore, the xylanase of A. nidulans was stable over a pH range of 3.5 to10.0 after 4 h of incubation (46), and the xylanase produced by A. niger DFR-5 lost activitybetween pH 4.0 to 6.5 (39).

Fig. 5 Curve of the optimum pH of the xylanase produced by A. tamarii URM 4634. Glycine–HCl buffer(pH 2.4, 3.0, and 3.6) (filled diamond), acetate buffer–Na (pH 3.6, 4.0, 4.5, and 5.0) (filled circle), Na citratebuffer (pH 5.0, 5.5, 6.0, 7.0 and 8.0) (filled triangle), Tris–HCl (pH 6.0, 7.0, 7.5, and 8, 0) (filled square), andglycine–NaOH (pH 8.0, 9.0, and 10.0) (multiplication symbol)


The effect of temperature on xylanase activity was positive, as activity increased withincreasing temperature, up to 90 °C, where the highest xylanase was observed. Xylanaseactivity was reduced at 100 °C (Fig. 7). For most of fungal xylanases, the optimum activity isbetween temperatures of 45 and 60 °C (38, 44, 47–50). Our results corroborated the findings ofJiang et al. (37), because they obtained xylanase produced by A. tamarii URM 4634 alsodisplayed optimal activity above 60 °C.

The enzyme was stable at a range of temperatures, retaining over 50 % activity after180 min (Fig. 7). In contrast, the activity of the xylanase produced by Chaetomium sp. wascompletely abolished after a 30-min reaction at 180 °C (37). The thermostability of xylanaseinvolves factors related to its thermodynamic stability, and its kinetics depend on molecularinteractions, such as hydrogen bonding, electrostatic and hydrophobic interactions, disulfidebonds, and the existence of metals that can promote a different conformational structure of theenzyme that displays greater structural efficiency that reduces entropy in the unfolding andrelease of tension (50, 51).

The xylanase produced in our study was not inhibited in the presence of Cu2+, Mn2+, Fe2+,Ca2+ (1 mM), and 2-mercaptoethanol (Tables 4 and 5), but Mg2+ (5 mM) inhibited enzymeactivity by 65 %; at 1 mM, the inhibition was ten times lower. Other ions and EDTA inhibitedthe enzyme between 1.8 and 12.7 %. These values were similar to those reported by Jiang et al.(37), where xylanase activity was inhibited in the presence of Cu2+, EDTA, and 2-mercaptoethanol at 1 mM. These results were positive, as animal feed often requires mineralsupplementation, which would not inhibit the activity of the xylanase produced by A. tamariiURM 4634.

The kinetic constants (Km) and (Vmax) of xylanase produced by A. tamarii URM 4634 were0.464 g L−1 and 63.39 U mL−1, respectively. When compared with other fungal species Km, itis noted that the xylanase produced by Aspergillus tamari URM 4634 can be considered morespecific for substrate utilized once the value Km is lower: Chaetomium sp. CQ31 (0.06 g L−1)(37), A. niger GH10 (2.43 g L -1) (52), A. niger US368 (1.03 g mL -1) (31), and A. niger DSM1957 (25 g L−1) (53). The Vmax value fit xylanse from Aspergillus tamari URM 4634

Fig. 6 pH-stability curve of the xylanase produced by A. tamarii URM 4634 after 180 min. Glycine–HCl buffer(pH 2.4, 3.0 and 3.6) (filled diamond), Na–acetate buffer (pH 3.6, 4.0, 4.5, and 5.0) (filled square), Na-citratebuffer (pH 5.0, 5.5, 6.0, 7.0 and 8.0) (filled triangle), and Tris-HCl (pH 6.0, 7.0, 7.5, and 8.0) (multiplicationsymbol)


(63.39 U mL−1) was higher than that obtained for A. niger GH10 (53.7 μmol mL−1) (52) butlower than that obtained for A. niger DSM 1957 (5,000 μmol mg−1) (53).

The xylanase produced in this study behaved positively in the presence of digestiveenzymes from monogastric animals in vitro. The lowest enzyme activity was approximately65 % in gastric conditions (pepsin+HCl), and its activity was above 70 % in response tointestinal activity (pancreatin and bile extract+trypsin+taurocholic acid). When completedigestion was simulated, the xylanase activity was 10 %. These results suggest that theenzyme retains its activity or is potentiated in the presence of digestive enzymes frommonogastric animals, and in vivo, the presence of digestive components can improve theperformance of the enzyme (19).

Fig. 7 Curve of the optimum temperature of xylanase produced by A. tamarii URM 4634 (filled square). Curvetemperature stability of the xylanase produced by A. tamarii URM 4634 after 180 min (filled triangle)

Table 4 Inhibition of xylanase in presence of metal ions and inhibitors

Substances Residual activity (1 mM) Inhibition (%) Residual activity (5 mM) Inhibition (%)

Controla 100.0±0.23 – 100.0±0.23 –

Cu2+ 103.4±0.24 0 104.7±0.65 0

Mn2+ 155.5±1.53 0 156.6±0.37 0

Na+ 85.7±0.67 14.3 91.8±0.56 8.2

Fe2+ 100.2±0.33 0 98.2±0.13 1.8

Zn2+ 96.2±0.87 3.8 87.3±0.38 12.7

Mg2+ 93.5±0.03 6.5 34.6±1.25 65.4

Ca2+ 101.1±0.45 0 93.1±0.61 6.9

EDTA 94.3±0.57 5.7 90.9±0.62 9.1

2-Mercaptoethanol 100.0±0.77 0 127.0±0.33 0

a Data are shown as residual activity (%)±SD. The control was conducted to test the xylanase activity normally,without addition of ions or inhibitory substances


SDS-PAGE analysis (Fig. 8a) was used to visualize bands contained in the enzyme extract,and these bands displayed molecular weights ranging between 13.53 and 71.46 kDa. The areasof hydrolysis observed in the zymogram (Fig. 8b) indicate that the A. tamarii URM 4634produced only an extracellular xylanase with a molecular weight of approximately 19.08 kDa.Generally, xylanases with molecular weight below 30 kDa are classified within a family of 11of glycosyl hydrolases with a high isoelectric point (54). This would explain why the xylanasehad better activity at an acidic pH, as its isoelectric point, which must be high, would presentless solubility and interaction with the middle, which hinders its activity. This finding is similarto that of other groups who have identified xylanases with weights of 17.5 and 23.0, 21 and26 kDa (31, 55).

Table 5 Effect of monogastric digestion in vitro on activity of xylanase produced by A. tamarii URM 4634

Assay Total activity (U mL−1) Control (U mL−1) Residual activity (%)

1 13.66±1.11 20.96±0.199 65±3.85

2 15.50±1.19 20.96±0.199 74±2.95

3 16.30±0.55 20.96±0.199 78±2.65

4 13.68 ± 0.76 12.96±0.92 106±0.92

Data are shown as residual activity (%)±SD

Fig. 8 a Electrophoretic profile of the enzyme produced by A. tamarii URM 4634 during this phase of the saltextractive fermentation; (1) column with protein standards and (2) column with sample. b Zymogram demon-strating xylanase activity


Conclusion

Extractive fermentation with ATPS is an effective method to extract the enzyme, resulting inhigh yield (72 %) that was more than doubled after scaling-up. The xylanase produced byA. tamariiURM 4634 has great potential for use in industrial feeds for monogastric animals, asit presents good xylanase activity and stability in a wide range of pH and temperaturecharacteristics, which is very important for industrial manipulation. Furthermore, the enzymeactivity did not change in response to in vitro simulated digestion (106 %). Therefore, thisenzyme can be used to supplement feeds for monogastric animals, improving the availabilityof metabolic energy and increasing their food production.

Acknowledgments The authors thank FACEPE, CNPq, and CAPES (PROCAD/NF 2009 N. 091/2010) byfunding and encouraging research.

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