Enzyme and Microbial Technology 29 (2001) 328 –334
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An endo--1,4-xylanase from Rhizopus oryzae: production, partial purification and biochemical characterization Ufuk Bakira,*, Sebnem Yavascaoglub, Ferda Guvencb, Aysegul Ersayinb a
Chemical Engineering Department, Middle East Technical University 06531 Ankara, Turkey b Biotechnology Department, Middle East Technical University 06531 Ankara, Turkey Received 31 August 2000; received in revised form 23 April 2001; accepted 22 May 2001
Abstract An endoxylanase (1,4--D-xylan xylanohydrolase, EC 3.2.1.8) was produced by Rhizopus oryzae fermentation. Different xylancontaining agricultural byproducts such as wheat straw, wheat stems, cotton bagasse, hazelnut shells, corn cobs and oat sawdust were used as the carbon source, while soybean bagasse was used as both the nitrogen and carbon source in the enzyme production medium. Partial steam hydrolysis of the agricultural byproducts increased the enzyme yield of the microorganism. The highest xylanase activity, 260 IU/ml fermentation medium, was obtained by using a medium containing 3% hydrolyzed corn cobs, 1% hydrolyzed soybean bagasse, 1% ammonium sulfate and 0.5% sodium chloride at 35°C, pH 5, 350 rpm and under aerobic conditions in a 2-1 fermenter. A maximal cellulose activity of 0.06 IU/ml was observed. The enzyme was partially purified from the culture medium by ammonium sulfate precipitation and cation exchange filtration. A 55-fold purification was achieved, with the purified xylanase having a specific activity of about 50 IU/mg protein. The molecular weight of the enzyme is about 22 kDa by SDS-PAGE. The optimal pH and temperature values of the enzyme were about 4.5 and 55°C, respectively. The enzyme obeys Michaelis-Menten kinetics with K m and V max values being 18.5 mg xylan/ml and 90 IU/mg protein, respectively. © 2001 Elsevier Science Inc. All rights reserved. Keywords: Rhizopus oryzae; Xylanase; Enzyme production; Purification and biochemical characterization
1. Introduction Lignocellulose is the most abundant and renewable biomass available on earth. It comprises three major group of polymers, cellulose (a linear homopolymer of -1,4-linked glucose units), hemicellulose (noncellulosic polymers including glucans, mannans, arabinans, galactans and xylans) and lignin (a complex polyphenol) [1]. Xylan, the major component of hemicellulose, is a heterogeneous polysaccharide consisting of -1,4-linked Dxylosyl residues on the backbone but also containing arabinose, glucuronic acid and arabinoglucuronic acids linked to the D-xylose backbone [2]. Due to xylan heterogeneity, the enzymatic hydrolysis of xylan requires different enzymatic activities. Two enzymes, -1,4-endo-xylanase (EC 3.2.1.8) and -xylosidase (EC 3.2.1.37), are responsible for hydrolysis of the main chain, the first attacking the internal main-chain xylosidic linkages * Corresponding author. Tel.: ⫹90-312-210-2619; fax: ⫹90-312-2101264. E-mail address:
[email protected] (U. Bakir).
and the second releasing xylosyl residues by endwise attack of xylooligosaccharides. These two enzymes are the major components of xylanolytic systems produced by biodegradative microorganisms such as Trichoderma, Aspergillus, Schizophyllum, Bacillus, Clostridium and Streptomyces species [3– 6]. However, for complete hydrolysis of the molecule, side-chain cleaving enzyme activities are also necessary. Xylanases have several different industrial applications including biodegradation of lignocellulose in animal feed, foods, and textiles, as well as biopulping in the paper and pulp industry [2]. Until now, no study on xylanases of any Rhizopus species has been appeared in the literature, except a recently published one in which 67 different Rhizopus strains were screened for the presence of xylanolytic activity [8]. Most of the Rhizopus strains, including three Rhizopus oryzae, produced xylanases by using either liquid or solid cultivation. We selected Rhizopus oryzae for xylanase production because of its GRAS status, which adds to the attractiveness of its enzymes, particularly in the food industry. Here we report the production of a -1,4-endo-xylanase from R.
0141-0229/01/$ – see front matter © 2001 Elsevier Science Inc. All rights reserved. PII: S 0 1 4 1 - 0 2 2 9 ( 0 1 ) 0 0 3 7 9 - 9
U. Bakir et al / Enzyme and Microbial Technology 29 (2001) 328 –334
oryzae ATTC 9363 by using low-cost lignocellulosic waste products. In addition we report partial purification and biochemical characterization of the enzyme.
2. Materials and methods 2.1. Organism and growth conditions R. oryzae strain ATCC 9363 was maintained on a medium containing 0.2% ammonium sulfate, 0.065% monobasic potasium phosphate, 0.025% magnesium sulfate, 0.005% zinc sulfate, 0.5% glucose and 1.5% agar. The plates were incubated at 30°C for three to four days for spore production and then stored at 4°C until use. 2.2. Fermentation studies Fermentations were performed in either shake flasks or in a 2-1 Virtis fermentor. Two different media, a minimal salt and a rich medium, were used. The salt medium was as above less glucose and agar but including 1% xylan. The rich medium contained 1% tryptone, 0.5% sodium chloride, 0.5% yeast extract and 1% xylan. Agricultural byproducts were chopped and dried at 100°C overnight and either directly added in the range of 1–3% to the rich medium or after steam hydrolysis at 121°C for 30 min. To start 100-ml shake flask and 1-l fermentor cultures, 106–107 spores and a 100-ml overnight shake culture were used as inocula, respectively. Daily samples were taken from the fermentation broth to check xylanase production until xylanase activities started to decrease, generally after five to eight days.
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2.4. Xylanase assay Endo--1,4-xylanase activity was assayed by measuring the release of reducing sugars from xylan by the SomogyiNelson method using a xylose standard [10]. To compare the enzyme yield of the microorganism with literature values, the dinitrosalicylic acid method was also used once to determine xylanase activity of the fermentation medium under optimal fermentation conditions, since these two procedures give different activity values [11–13]. The reaction was started with the addition of one volume of appropriately diluted enzyme solution to ten volumes of 1% birchwood xylan solution in 100 mM citrate buffer, pH 4.8, previously equilibrated to 40°C. Samples were taken at regular intervals and the reactions were immediately stopped by adding Nelson’s alkaline copper reagent to determine the initial reaction rate. One unit (IU) of xylanase activity is defined as the enzyme necessary to catalyze the production of 1 mol of xylose equivalent per min at 40°C and pH 4.8. 2.5. Cellulase assay Cellulase activity was measured by the release of reducing sugars from cellulose by the Somogyi-Nelson method using a glucose standard [10]. The substrate was 50 mg Whatman No. 1 filter paper in 2 ml of 0.1 M citrate buffer, pH 4.8. An appropriately diluted 1.5-ml enzyme solution was mixed with the substrate solution previously equilibrated to 40°C. Samples were taken and reducing sugar concentrations were determined. One unit (IU) of cellulase activity is defined as the enzyme necessary to catalyze the production of 1 mol of glucose equivalent per min at 40°C and pH 4.8. 2.6. Protein content
2.3. Growth rate determination Growth rate measurements by weighing the mycelium were impossible due to the difficulty of separating it from the compact structure formed with the insoluble materials present in the fermentation medium. Therefore a correlation was determined between the mass of extracted intracellular protein and cell mass by using fermentations performed in rich medium, which does not contain any insoluble material. To this end, multiple fermentations in rich media were performed in shake flasks. Two of them were harvested each day by centrifugation at 10,000 ⫻ g and the mycelium was dried at 45°C until a constant weight was reached. The dried mycelium was frozen and crushed in the presence of liquid nitrogen in a mortar and pestle. The broken cells were suspended in 100 mM citrate buffer, pH 4.8, and the released protein concentration was determined by using a modified Lowry’s procedure [9]. The cell mass produced in fermentation medium containing insoluble materials was calculated based upon the released intracellular protein quantity.
Protein concentration was measured by using a modified Lowry method with bovine serum albumin as the standard [9]. 2.7. Purification All purifications were carried out at 4°C unless otherwise stated. After harvesting the R. oryzae mycelia by centrifugation at 10,000 ⫻ g for 10 min, the supernatant was used as crude xylanase extract. Protein precipitated between 40 and 80% ammonium sulfate saturation was dissolved in 50 mM citrate buffer at pH 4.8 and dialyzed immediately against the same buffer overnight. After equilibrating the membrane pH to 4.8 by using 30 ml buffer, sample was loaded onto a strong cation exchange membrane (Sartorius S5F) at pH 4.8. The membrane was washed with 50 ml buffer to remove unbound proteins, and then the bound proteins were eluted with 1 M KCl solution. Finally the eluted proteins were immediately dialyzed against 50 mM citrate buffer at pH 4.8.
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2.8. Biochemical characterization 2.8.1. Analytical gel electrophoresis Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) was performed as described by Laemmli et al. [14] by using 4% stacking and 12% separating polyacrylamide gel in the presence of 0.1% SDS and run on a Bio-Rad Mini Protean II DUAL SLAB cell according to the manufacturer’s instructions. Mr markers of 15, 25, 35, 50, 75 and 100 kDa (Sigma M 0671) were used. Silver staining was employed for Mr estimation, while for activity check of the band presumed to be xylanase, KCl staining [15] was used. 2.8.2. KCl activity measurement After SDS-PAGE was performed, the nature of the xylanase band was verified by testing xylanase activity. The presumed xylanase band was cut from the KCl-stained SDS-PAGE gel. After removal of SDS, protein was renatured following total denaturation [15]. Afterwards xylanase activity was measured by using a semiquantitative gel diffusion assay [16]. Enzyme solutions were injected into 4-mm diameter holes punctured in a 0.2% RBB (Remazol Brilliant Blue) xylan-containing agar plate. Diffusion zones formed around the holes verified xylanase activity. 2.8.3. Kinetic studies Initial reaction rates of birchwood xylan hydrolysis were determined at different substrate concentrations ranging from 2.5 to 25 mg xylan/ml. Higher xylan concentrations could not be used due to low xylan solubility. Reaction rate vs. substrate concentration curve was plotted to determine whether the enzyme obeys Michaelis-Menten kinetics, and constants were determined from a Lineweaver-Burk plot. 2.8.4. Effects of pH and temperature on xylanase activity Xylanase activity was assayed using 50 mM citrate buffer, pH 3.5–5.5, at 30 – 60°C using 25 mg/ml xylan. During pH- and temperature-activity experiments, the assay temperature and pH were kept constant, respectively, at 40°C and 4.8.
Fig. 1. Xylanase production by R. oryzae in rich and minimal salt media. Fermentations were carried out in an incubator shaker at 30°C and 175 rpm. Minimal salt medium contained 1% xylan, 0.2% ammonium sulfate, 0.065% potassium phosphate (monobasic), 0.025% magnesium sulfate, and 0.005% zinc sulfate, while the rich medium contained 1% xylan, 1% tryptone, 0.5% sodium chloride and 0.5% yeast extract (F: rich medium, }: salt medium).
was produced on rich medium, almost three times more than that produced on the minimal salt medium. Following this, in an attempt to investigate the relationship between production of xylanase activity and use of a carbon source as inducer, the xylan in the rich medium was replaced by 2% wheat straw, wheat stem or soybean bagasse as the only carbon source. Each agricultural byproduct was either untreated or treated by steam hydrolysis as explained in section 2.2. R. oryzae seemed to use agricultural byproducts inefficiently unless they were steam-hydrolyzed, as maximal enzyme concentrations were 3–7 times higher with this pretreatment (Fig. 2). After observing the positive effect of pretreatment, all the other agricultural byproducts, including corn cobs, hazelnut shells, cotton bagasse and oak sawdust, were used after steam hydrolysis. As observed from Table 1, all the agricultural byproducts gave better xylanase yields than pure xylan. In the best case, use of corn cobs increased xylanase yield approximately seven times.
3. Results and discussion 3.1. Production of R. oryzae xylanase After observing the xylanolytic activity of the strain on an agar medium containing xylan as the only carbon source and congo red as the indicator [17], production studies started in an incubator shaker with a minimal salt medium and a rich medium containing 1% xylan, the only carbon source, as inducer. As observed in Fig. 1, the use of minimal salt medium was not as successful as use of the rich medium. After a six-day fermentation, 0.39 IU/ml xylanase
Fig. 2. The effect of thermal pretreatment of agricultural byproducts on the xylanase production by R. oryzae. The fermentations were carried out in an incubator shaker at 30°C and 175 rpm in a media containing 1% tryptone, 0.5% sodium chloride and 0.5% yeast extract and one of the carbon sources (⫻: hydrolyzed wheat straw, 䊐: hydrolyzed soybean, E: hydrolyzed wheat stem, ‚: unhydrolyzed wheat straw, {: unhydrolyzed soybean, 兩: unhydrolyzed wheat stem).
U. Bakir et al / Enzyme and Microbial Technology 29 (2001) 328 –334 Table 1 Effect of carbon source on xylanase production by R. oryzae* Carbon source Xylan Hazelnut shells Wheat stems Soybean bagasse Wheat straws Cotton bagasse Oak sawdust Corn cobs
Maximum Xylanase activity (IU/ml)
Ferm. Period** (day)
0.39 0.42 0.70 0.74 1.5 1.26 1.26 2.81
6 5 5 5 6 5 6 5
* The fermentation medium contained 1% tryptone, 0.5% yeast extract, 0.5% NaCl, and 2% of a carbon source, except 1% xylan. Carbon sources were added after thermal pretreatment. The fermentations were carried out in an incubator shaker at 30°C and 175 rpm. ** Fermentation period at which maximum enzyme titers were observed.
Maximal xylanase activities were achieved after a five-day fermentation for all cases except wheat straw and oak sawdust, which took six days. Gomes and Purkarthofer (1993) also reported corn cob powder as the best carbon source for xylanase production from a thermotolerant fungus, Thermomyces lanuginosus [18]. Considering both its high protein content and the nearly twofold increase in enzyme yield when soybean bagasse was used as the only carbon source (Table 1), the effect of this substrate as a nitrogen source was tested. To this purpose, the previously used nitrogen sources, yeast extract and tryptone, were replaced by 1% soybean bagasse one at a time. As observed from Table 2, when soybean bagasse was added without omitting any of the nitrogen sources, the maximal enzyme activity remained constant. But when either yeast extract or both yeast extract and tryptone were omitted, 7 and 38% increases in enzyme yields, respectively, were observed. However, omitting both of the simpler nitrogen sources increased the fermentation period from five to eight days. After corn cobs and soybean bagasse were recognized as the best carbon and nitrogen sources, the effects of changing their concentrations were tested. For this purpose, corn cobs and soybean bagasse concentrations were increased to 3% and 2%, respectively, one at a time. The increase in the corn cob concentration increased maximal enzyme concentration about 80%; however, the increase in soybean bagasse concentration led to an increase of 26%, probably due to higher production levels of the proteolytic enzymes or the necessity of nitrogen limitation for high enzyme production. Finally, 1% of the inorganic nitrogen sources, ammonium sulfate, potassium nitrate, or sodium nitrate were included in addition to 1% soybean bagasse, causing enzyme concentrations to increase almost sixfold for the first two salts and fivefold for the third. The fermentation period necessary to obtain maximal yields decreased to five days with ammonium sulfate and to six days with the other two salts.
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Table 2 Effect of medium composition, temperature and pH on xylanase production by R. oryzae Medium compositiona,b
Xylanase act. (IU/ml)
Ferm. Periodc (day)
2% C ⫹ 0.5% YE ⫹ 1% T 2% C ⫹ 1% S ⫹ 0.5% YE ⫹ 1% T 2% C ⫹ 1% S ⫹ 1% T 2% C ⫹ 1% S 3% C ⫹ 1% S 3% C ⫹ 2% S 3% C ⫹ 1% S ⫹ 1% (NH4)2 SO4 3% C ⫹ 1% S ⫹ 1% KNO3 3% C ⫹ 1% S ⫹ 1% NaNO3 Temperature (°C)d 30 35 37 pHe 4 5 6
4.10 4.15 4.39 5.64 7.38 5.15 23.3 22.2 20.2
5 5 5 8 8 6 5 6 6
2.55 4.14 3.15
5 5 5
7.53 20.5 17.4
5 5 6
a C: corn cobs; S: soybean; YE: yeast extract; T: tryptone. All the media contains 0.5% NaCl. b Fermentations were performed in shake flasks at 35°C and 175 rpm. c Fermentation period at which maximum enzyme titers were observed. d Fermentations were performed in shake-flasks at 175 rpm in media containing 2% corn cobs ⫹ 1% S ⫹ 1% tryptone ⫹ 0.5% NaCl. e Fermentations were performed in a 2-1 fermentor at 35°C in media containing 2% corn cobs ⫹ 1% S ⫹ 0.5% NaCl.
Corn cob concentrations higher than 3% were not used due to the increase in medium viscosity. Therefore, the best enzyme producing medium was 3% corn cobs, 1% soybean bagasse, 0.5% NaCl, and 1% ammonium sulfate. After studying the effects of fermentation composition on enzyme yield, the effect of temperature was investigated in shake-flask cultures. Three different temperatures, 30, 35 and 37°C, were tested and the best enzyme yields were obtained at 35°C (Table 2). To determine the effect of pH on enzyme production, a new set of fermentations were performed in a 2-1 fermentor. As observed in Table 2, although best enzyme yields were obtained at pH 5, these were not much different from those at pH 6. Optimization studies were completed by performing the fermentation in the 2-1 fermentor under the best conditions determined in the previous experiments, using a medium containing 3% corn cobs, 1% soybean bagasse, 0.5% NaCl and 1% ammonium sulfate at 35°C, pH 5, and 350 rpm under aerobic conditions. Highest xylanase activity was 32 IU/ml (Fig. 3). Cellulase production, with a maximal activity of 0.06 IU/ml, quite low compared with xylanase activity, occurred during the fermentation. Xylanase and cellulase production and microbial growth rate profiles are given in Fig. 3. Maximal xylanase activity was reached after five days of fermentation. Mycelial dry weight decreased after
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Fig. 3. Xylanase, cellulase production and microbial growth rate profiles during R. oryzae fermentation in a medium containing 1% soybean bagasse, 3% corn cobs, 0.5% sodium chloride and 1% ammonium sulfate at 35°C and 350 rpm with aeration in a 2-1 fermentor. Enzyme activities were measured by Somogyi-Nelson method (}: xylanase activity, Œ: dry cell weight, ■: cellulase activity ⫻ 100). Fig. 4. SDS-PAGE of xylanase preparations. Lane 1: Mr markers, Lane 2–3: crude enzyme, Lane 4 –5: after ammonium sulfate precipitation, Lanes 6 –7: after ion-exchange filter. Samples containing 5–10 g of protein were loaded into wells. Protein bands were stained with silver stain.
an increase during the first three days. This decrease might be an artifact related to increased cell wall strength with age during fermentation, since dry weight was correlated with intracellular protein content released after cell disruption as explained in Section 2.3. In all our experiments, the Somogyi-Nelson method was used to determine xylanase activity. However, the DNSA assay is generally preferred in the literature to test reducing sugar concentrations for xylanase activity measurements; it gives seven to ten times more activity than the SomogyiNelson method [12,13]. To compare the enzyme yield of R. oryzae with the literature, xylanase activity of the last fermentation broth was also measured by using DNSA method and maximal activity was 260 IU/ml. Veluz and coworkers [8] reported 306, 372 and 508 IU/ml xylanase activities for R. oryzae P17, P1 and P3 strains. However comparing these values with our data is impossible since they did not measure initial reaction rates as we did [8]. On the other hand, there are many reports on xylanase production from different fungi. For example, highest xylanase activities obtained from Penicillium canescens, Trichoderma viride and Aspergillus tamarii were 300, 112 and 77 IU/ml, when grown on wheat straw, barley husk and corn cobs, sequentially [19 –21]. Since the assay conditions and enzyme units used in these studies are similar, these values can be compared with the maximal xylanase yield of R. oryzae, 260 IU/ml.
3.2. Purification The supernatant obtained by centrifugation of the fiveday old culture was used as a crude extract, and partial purification was performed by ammonium sulfate precipitation (40 –75%) and cation-exchange filtration (Sartorius S5). Specific activity increased from 0.93 IU/mg to approximately 50 IU/mg and nearly 55-fold purification was achieved with 10% yield (Table 3). 3.3. Biochemical characterization The molecular weight and purity of the partially purified enzyme were evaluated with SDS-PAGE. After electrophoresis, the gels were silver-stained. Several minor protein bands were also observed. The xylanase activity of the band at 22 kDa was checked on a parallel gel stained with KCl after SDS removal. The presence of 50% of the original xylanase activity verified the molecular weight of the enzyme as about 22 kDa (Fig. 4). Initial reaction rates were determined at different substrate concentrations ranged from 2.5 to 25 mg birchwood xylan/ml. Due to low xylan solubility, concentrations higher than 25 mg/ml could not be used. Rate vs. xylan concen-
Table 3 Purification of xylanase from R. oryzae culture supernatant
Crude extract 40–75% (NH4)2SO4 fraction Cation-exchange filter
Xylanase activity (IU/ml)
Total activity (IU)
Protein Conc. (mg/ml)
Total protein (mg)
Spec. activity (IU/mg)
Yield, %
Purification fold
4.40 6.00 4.98
2200 270 214
4.75 2.60 0.10
2375 117 4
0.93 2.31 49.8
100 12.3 9.73
1.00 2.48 53.6
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Table 4 Some characteristics of low molecular weight xylanases from bacteria and fungi
Fig. 5. Lineweaver-Burk plot for the partially purified R. oryzae xylanase. Xylanase activity was measured at 40°C and pH 4.8.
tration data showed that Michaelis-Menten kinetics were obeyed, with V max and K m values using a Lineweaver-Burk plot as 90 IU/mg protein and 18.5 mg xylan/ml, respectively (Fig. 5). The pH dependence of xylanase activity was determined at 40°C by varying the reaction mixture pH in 50 mM citrate buffer from 3.5 to 5.5. The highest activity occurred at pH 4.5 (Fig. 6). Enzyme activity in 25 mg xylan/ml and 50 mM citrate buffer at pH 4.8 was measured at temperatures from 30°C to 60°C; the maximal activity was 55°C (Fig. 6). An interesting physicochemical property of fungal and bacterial xylanases seems to be the strong relationship between their molecular weight and isoelectric point values. Many xylanases fall into a pattern of high molecular weight/ low pI value (Family 10) or low molecular weight/high pI value (Family 11) [2]. The properties of the R. oryzae xylanase are given together with properties of some other low molecular weight xylanases (Table 4). They have quite similar molecular weights, optimal pHs and temperature values.
Fig. 6. Effect of temperature and pH on the activity of R. oryzae xylanase. Assay temperature was kept constant at 40°C during pH-activity experiments while pH was kept constant at 4.8 during temperature-activity experiments (F: pH, }: temperature).
Xylanase source
Mol. wt. (kDa)
Opt. pH
Opt. temp. (°C)
Reference
Trichoderma harzianum Trichoderma viride Streptomyces sp No 3137 Trichoderma lignorum Bacillus sp. W2 Rhizopus oryzae
20 22 25 20 22.5 22
5.0 5.0 5.0–6.0 6.5 6.0 4.5
50 53 60–65 — 65 55
6 22 23 24 25 This study
In conclusion, R. oryzae produces a low molecular weight endoxylanase from low-cost lignocellulosic byproducts at reasonable rates. Since R. oryzae is a GRAS microorganism, the xylanase produced can be used in the food industry as well as in other industrial areas. Acknowledgements This project was supported by Middle East Technical University (AFP-97-07-02-02) and the State Planning Organization of Turkey (AFP-06-02DPT.98K 122770). Aysegul Ersayın and Sebnem Yavascaoglu were supported by the Graduate School of Natural and Applied Sciences of METU and TUBITAK, respectively. We would like to thank Professor Peter J. Reilly for his help to edit the manuscript.
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