Skip to main content

Optimization of upstream and downstream process parameters for cellulase-poor-thermo-solvent-stable xylanase production and extraction by Aspergillus tubingensis FDHN1



Xylanases are important members of the hemicellulolytic enzyme system. Xylanase plays a vital role in the hydrolysis of major hemicellulosic component xylan and converts it into xylooligosaccharides and ultimately yields xylose. Cellulase-lacking or cellulase-poor xylanase with high temperature and pH stability has gained special attention, especially in paper and pulp industries. Most of the available literature highlighted the fungal xylanase production by optimizing environmental and cultural parameters. However, the importance of enzyme recovery from fermented biomass still needs attention. In this study, upstream and downstream process parameters were studied for enhancing xylanase production and extraction by a newly isolated Aspergillus tubingensis FDHN1 under solid-state fermentation using low-cost agro-residues.


In the present study, A. tubingensis FDHN1 was used for the xylanase, with very low level of cellulase, production under solid-state fermentation (SSF). Among various agro-residues, sorghum straw enhanced the xylanase production. Under optimized upstream conditions, the highest xylanase production 2,449 ± 23 U/g was observed. Upon characterization, crude xylanase showed stability over a broad range of pH 3.0 to 8.0 up to 24 h. The temperature stability revealed the nature of the xylanase to be thermostable. Native polyacrylamide gel electrophoresis (native PAGE) and zymogram analysis revealed the multiple forms of the xylanase. Due to the many industrially important characteristics of the xylanases, the study was elaborated for optimizing the downstream process parameters such as volume of extractant, extraction time, temperature and agitation speed to recover maximum xylanase from fermented sorghum straw. The highest amount of xylanase (4,105 ± 22 U/g) was recovered using 0.05 M sodium citrate buffer (pH 6.5) at 12:1 (v/w) extractant/solid ratio, 90-min extraction time, 150-rpm agitation speed and 40°C. Finally, detailed bioprocess optimization shows an overall 6.66-fold enhancement in the xylanase yield.


The present study consolidates the importance of upstream and downstream process optimization for the overall enhancement in the xylanase production. The xylanase from A. tubingensis FDHN1 shows the stability at different pH and temperature, and it was also active in the presence of organic solvents. These properties of xylanase are very much important from an industrial application point of view.

Production, characterization and downstream processing of Aspergillus tubingensis FDHN1 xylanase


Hemicelluloses are the second most abundant renewable resources, only exceeded by cellulose. Xylan constitutes a major component of hemicellulose, which is a heteropolysaccharide, having a chain of β-1,4-linked xylopyranose residues. The complete hydrolysis of xylan requires the combined action of various enzymes such as endo β-1,4-xylanase (E.C., exoxylanase (β-D-xylan xylohydrolase), β-D-xylosidase (E.C., etc. [1,2]. Recently, xylanases have attracted considerable attention due to its application in many industrial processes such as enzymatic bleaching of paper pulp, juice clarification, extraction of plant oils, texture improvement in bakery, bioconversion of agricultural waste, bioscouring in textiles and improvement of animal feed digestibility [3-5]. Due to such immense industrial applications, xylanases have a worldwide market of around 200 million dollars.

Filamentous fungi are good producers of xylanase because they are capable of producing high levels of extracellular enzymes and can be cultivated very easily. On an industrial scale, xylanases are mainly produced by Aspergillus and Trichoderma sp. in solid-state fermentation (SSF). The research has indicated that xylanase production differs with the strains and is regulated by physiological, nutritional and biochemical nature of the microbes employed [6]. The fermentation profile of an organism that influences metabolism-mediated production includes pH, temperature, carbon and nitrogen source, metal ion requirement, incubation time, inoculum size, etc. The production of an industrial enzyme by optimizing these growth parameters is of prime importance because an improper optimization of these factors may result in the lower enzyme production. In general, no defined medium has been established for the best production of any metabolite because the genetic diversity present in different microbial sources causes each organism or strain to have its own special conditions for maximum product yield [7]. For the commercialization of xylanase production, it is necessary to identify a microorganism that produces high levels of xylanase using cheaper carbon sources and having novel properties. As lignocellulosic materials are abundantly found in nature in the form of agricultural and industrial residues, it can be exploited as a potential substrate for growing the organisms.

Apart from medium optimization, recovery of enzyme is also an important aspect of SSF technology. An optimized recovery process would generate concentrated enzyme extract, prevent enzyme loss, reduce the number of the downstream process step and thus gain better commercial viability. A wealth of information is available on medium optimization for xylanase production [8-11], but studies on xylanase recovery from fermented substrate are scarce [11]. Therefore, it is highly imperative to optimize production and extraction parameters which further facilitate economic design of the full-scale operation system for newly isolated microbial strains.

In the present study, we have highlighted the optimization of upstream process for xylanase production using newly isolated Aspergillus tubingensis FDHN1 under SSF. Produced xylanase was characterized for important properties such as pH, temperature and solvent tolerance. Looking towards the important features of this xylanase and thrust for its higher yield, downstream process parameters for xylanase recovery were also optimized.


Agro-residue preparation, chemicals and medium components

Different agro-residues such as caster straw, wheat straw, maize straw, rice straw, sorghum straw, barley straw, sugarcane bagasse, wood chips and groundnut shell were collected locally from agricultural farms. To remove the adhered surface dust particles, agro-residues were washed in distilled water, dried at 50°C and chopped into small pieces. These were ground in mixture to achieve average particle size of 0.5 to 2 mm.

Chemicals such as oat spelt xylan, carboxymethyl cellulose (CMC), bovine serum albumin (BSA) and dinitrosalicylic acid (DNSA) were purchased from HiMedia Laboratories Ltd (Mumbai, India). All the other chemicals, media, salts and reagents used were of analytical grade (Sigma-Aldrich, St. Louis, MO, USA; HiMedia, Mumbai, India; and Merck & Co., Inc., Whitehouse Station, NJ, USA).

Fungal isolation, identification and inoculum preparation

The novel cellulase-poor xylanase-producing fungal strain FDHN1 was isolated in our laboratory from active compost pit (approximately 10-cm depth). The strain was identified at the National Fungal Culture Collection of India (NFCCI), Pune, India, on the basis of morphological and molecular study. The strain was identified as A. tubingensis. Morphologically, the strain showed white velvety colonies which later turns to greyish-black on potato dextrose agar (PDA) plates. Under stereoscopic observation, the conidial heads were found to be globose, radiating, split into columns, greyish metallic in colour and measured up to 260 × 260 μm in size. The conidiophores were found to be hyaline to subhyaline which later turned into brown when old, smooth-walled and up to 625 × 5 μm in size. The vesicles were globose to subglobose, fertile all over and up to 39.0 × 45.5 μm in size. 18S rDNA gene sequence of fungus has been submitted to NCBI (Accession No. KF971693).

Spore inoculum was prepared from 5-day grown culture on PDA slants at 35°C. Suspension was prepared by lightly brushing the slant containing fungal mycelia with sterile distilled water to give a final spore count of approximately 1 × 106 spores/mL.

Xylanase production using agro-residues and upstream process optimization

Caster straw, wheat straw, maize straw, rice straw, sorghum straw, barley straw, groundnut shell, sugarcane bagasse and wood chips were used as substrates to produce xylanase under SSF. Erlenmeyer flasks (250 mL) containing 5 g of each substrates and 5 mL of mineral salt medium (g L−1: KH2PO4, 3.0; NaNO3, 2.5; MgSO4 · 7H2O, 1.0; CaCl2, 0.05; FeSO4 · 7H2O, 7.5; MnSO4, 2.5; ZnSO4 · 7H2O, 3.6; CoCl2 · 6H2O, 3.0 and 0.1% Tween-80 (v · v −1), pH 5.0) were autoclaved at 121°C for 15 min, cooled, inoculated with 1 mL of spore suspension and incubated at 35°C for 6 days.

SSF conditions were optimized by changing one factor at a time while keeping the other factors constant. To improve xylanase production, various upstream process parameters such as sorghum straw concentration (3.0 to 15 g), incubation period (2 to 8 days), substrate to moisture ratio (1:1 to 1:7 w/v), medium pH (3.0 to 9.0) and temperature (25 to 55°C) were optimized. Furthermore, effect of various carbon sources (glucose, fructose, sucrose, xylose, lactose, maltose and mannitol at 0.5% concentration), organic nitrogen sources (peptone, beef extract, urea, yeast extract, tryptone, malt extract, meat extract, gelatine and skimmed milk powder at 0.05 g ‘N’ equivalent in 50 mL medium) and inorganic nitrogen sources (ammonium hydrogen phosphate, ammonium dihydrogen phosphate ammonium sulphate, ammonium per sulphate, ammonium nitrate, ammonium chloride and sodium nitrate at 0.05 g ‘N’ equivalent in 50 mL medium) were evaluated for xylanase production. The xylanase production was also assessed in the presence of different metal additives (CuCl2, MgCl2, SnCl2, CaCl2, HgCl2, CoCl2, MnCl2, NiCl2 and FeCl3, 0.2%) and modulators (Tween-40, Tween-80, Triton X-100, glycine, glycerol, EDTA and SDS, 0.1%).

Enzyme assay and protein estimation

Xylanase activity was determined by adopting the modified method of Bailey et al. [12]. The reaction mixture containing 450 μL of oat spelt xylan (1%) prepared in 0.05 M citrate buffer (pH 5.0) and 50 μL of enzyme was incubated at 55°C for 10 min. The reaction was stopped by adding 2 mL of DNSA. The mixture was then heated in boiling water bath for 10 min and cooled down to room temperature, and then, 3 mL of water was added to the system. CMCase activity was assayed according to the method of Ghose [13]. The reaction system containing 1,000 μL of CMC (1%) dissolved in 0.05 M citrate buffer (pH 5.0) and 500 μL of enzyme was incubated at 60°C for 60 min. After incubation, 3 mL of DNSA was added to the system followed by 10-min incubation in a boiling water bath. The absorbance of the colour developed during xylanase and cellulase assay was measured at 540 nm as described by Miller [14]. One unit of xylanase/cellulase activity was defined as the amount of enzyme required to liberate 1 μmol of xylose/glucose equivalent per minute under the specified conditions. The unit ‘U/g’ denotes the xylanase/cellulase activity in international unit per gram dry substrate.

Soluble protein was determined by Folin's method using bovine serum albumin as standard [15].

Enzymatic properties of crude xylanase

The pH optima for xylanase activity was determined by measuring its activity in various buffers (0.05 M) such as sodium citrate (pH 3.0 to 6.0), sodium phosphate (pH 6.0 to 8.0) and glycine-NaOH (pH 8.0 to 10). The pH stability of xylanase was studied by incubating it in the above buffers for 24 h, and its residual activity was determined at a regular interval of 3 h.

The optimal temperature for xylanase activity was determined by assaying its activity at different temperatures (30 to 70°C). The thermostability of xylanase was determined at 30 to 70°C by measuring its activity at a regular interval of 30 min up to 120 min.

The solvent stability of xylanase was assessed by incubating the enzyme in the presence of various organic solvents (methanol, ethanol, propanol, butanol, acetone and benzene, at 10 to 30% concentration) for 30 min, and at the end of the reaction, residual activity was measured.

Native PAGE and xylan zymography

Native polyacrylamide gel electrophoresis (native PAGE) of the crude enzyme extract was performed at room temperature using 0.05 M citrate buffer (pH 5.0). The electrophoresed gel was stained with Coomassie Brilliant Blue G-250 for molecular weight determination [16]. A replica gel containing xylan (0.5%) was incubated at 50°C for 5 min in buffer and then stained in 0.1% Congo Red solution. After removal of dye using 1 M NaCl, gel was transferred to 5% acetic acid solution. Clear bands against a dark background indicated xylanase activity.

Enzyme extraction and downstream process optimization

Initially, the xylanase extraction was performed using 0.05 M citrate buffer (pH 5.0) at an extractant/solid ratio of 10:1 (v/w). Next, the suspension was agitated at 120 rpm for 60 min (35°C), centrifuged at 10,000×g for 15 min (4°C) and finally the clear supernatant was used as a crude enzyme source.

In order to find the most suitable extractant for xylanase recovery from fermented sorghum straw, seven different solvents (0.05 M citrate buffer, pH 5.0; 0.05 M sodium citrate buffer, pH 6.5; double-distilled water, pH 7.0; 0.05 M phosphate buffer, pH 7.5; 0.05 M Tris-HCl buffer, pH 8.0; 0.1% Tween-80 and 1.0% NaCl) at an extractant/solid ratio of 10:1 (v/w) were used. After selecting extractant type, different recovery parameters such as extractant/solid ratio (4:1 to 16:1 (v/w)), extraction time (15 to 120 min), agitation speed (50 to 300 rpm) and extraction temperature (25 to 50°C) were also optimized.

After optimizing extraction conditions, the percentage efficiency of xylanase recovery was studied under un-optimized and optimized extraction conditions. The percentage efficiency of xylanase recovery was calculated as follows:

$$ \%\ \mathrm{Xylanase}\ \mathrm{recovery} = \frac{\mathrm{Xylanase}\ \mathrm{activity}\ \mathrm{extracted}}{\mathrm{Total}\ \mathrm{xylanase}\ \mathrm{activity}\ \mathrm{in}\ \mathrm{fermented}\ \mathrm{substrate}} $$

Effect of optimized upstream and downstream conditions on total xylanase yield

At the end of detailed upstream and downstream process optimization, additional set of experiment for total xylanase productivity was carried out under the optimized conditions and obtained results were compared with earlier-reported xylanase-producing Aspergillus strains.

Results and discussion

Selection of agro-residues for xylanase production

Due to its high cost, pure xylan is not affordable for large-scale production of xylanase. Utilization of agricultural residues for enzyme production may serve a dual purpose: on one side, it provides a valuable metabolite, and on the other side, it helps in solving environmental solid waste disposal problem. Therefore, various low-cost agricultural residues were explored for the xylanase production. After 6 days of cultivation, among the tested agro-residues, sorghum straw supported maximum xylanase activity (615.5 ± 19.21 U/g) followed by wheat straw, sugarcane bagasse and rice straw (Table 1). Till date studies on xylanase production using sorghum straw as a substrate under SSF are scarce. It is believed that the substrate from graminaceous plants containing arabinoxylans supports high xylanase production [17]. The high xylanase titre when sorghum straw was used may be attributed to the nature of hemicelluloses, presence of some activators, surface pore size and favourable degradability of the carbon source. Thus, sorghum straw was used as a substrate throughout the optimization of upstream and downstream process parameters for xylanase production by A. tubingensis FDHN1 under SSF. Previously, use of sorghum straw has been reported as a potent inducer for xylanase production from Thermomyces lanuginosus under SSF by Sonia et al. [18]. However, xylanase production under SSF using wheat bran has also been reported by Aspergillus niger [11], A. niger and Aspergillus niveus [19].

Table 1 Effect of various agro-residues on xylanase production from A. tubingensis FDHN1 under SSF

In the present study, a variation in xylanase production was observed with the change in sorghum straw concentration from 3.0 to 15 g. Maximum xylanase production (1,329 ± 29 U/g) with very poor cellulase activity (0.42 ± 0.02 U/g) was obtained using 9 g sorghum straw under SSF (Figure S1 in Additional file 1).

Evaluation of upstream bioprocess parameters for xylanase production

Effect of incubation period and substrate to moisture ratio

The fermentation time for the maximum enzyme production is dependent upon the organism type and cultural/environmental conditions. Xylanase production at different time intervals during SSF showed that it was initiated from the second day and reached the maximum (1,217 ± 14 U/g) on the fifth day with specific xylanase activity of 53.54 ± 1.88 U/mg (Figure 1). Xylanase activity was found to decrease sharply with prolonged incubation. This result suggested that the fermentation endpoint should be controlled carefully because non-specific proteases secreted by the fungus may degrade the synthesized xylanase. The decrease in xylanase production may also be due to the accumulation of the end products during fermentation. SSF has also been carried out by Pandya and Gupte [20] for 10 days to produce xylanase from A. tubingensis JP-1 using wheat straw. They have noticed optimum xylanase yield on the eighth day of incubation. However, an increase in the xylanase production was observed by Goyal et al. [21] up to 17 days of fermentation using various agro-residues under SSF by Trichoderma viride. The above results indicated that the fermentation time for enzyme production under SSF depends on the growth rate of the microorganism and its pattern for enzyme production. The strain A. tubingensis FDHN1 used in the present study showed relatively high xylanase production within a short incubation period which could be attractive for large-scale xylanase production.

Figure 1
figure 1

Effect of incubation time on the xylanase production. SSF was carried out using 9 g sorghum straw for respective days (2 to 8 days) at 35°C, and samples were analysed at a regular interval of 24 h.

Apart from incubation time, moisture content plays a major role during SSF in the regulation of microbial metabolism known as water activity [22]. In the present investigation, influence of moisture content was studied at different solid/liquid ratios of 1:1 to 1:7 (w/v). The highest xylanase production (1,659 ± 17 U/g) and specific xylanase activity (81.60 ± 1.18 U/mg) were obtained at 1:5 (w/v) moisture ratio after 5 days of incubation, while maximum cellulase production (2.03 ± 0.11 U/g) was achieved at 1:6 (w/v) moisture ratio. In some fungi, high xylanase production has been shown to be firmly associated with cellulase production [23], but A. tubingensis FDHN1 did not produce much cellulase at different time intervals and moisture ratios. At 1:6 and 1:7 (w/v) moisture ratios, the xylanase production decreased to 1,463 ± 6 and 1,397 ± 6 U/g, respectively (Table 2). The obtained results indicated that the deviations from the optimum level moisture content lead to reduced xylanase production. In SSF, moisture content is an important factor that determines the overall success of a process by altering the physical properties of the solid substratum. The lower xylanase production at higher moisture levels could attribute to the decrease in porosity, alteration in particle structure or lower oxygen transfer, whereas the lower moisture content leads to a reduction in the diffusion of the nutrients in the substrate, lower degree of swelling and higher water tension [24]. Previously, optimum moisture level of 1:5 (w/v) has been shown for the xylanase production after 10 days under SSF by Pandya and Gupte [20].

Table 2 Evaluation of physiological parameters for xylanase production from A. tubingensis FDHN1 under SSF

Effect of pH and temperature

The enzyme production by microbial strains strongly depends on the initial pH of the medium as it influences many enzymatic processes and transport of various components across the cell membrane [25]. In the present study, xylanase production was noticed in the wide range of pH 3.0 to 9.0 (Table 2) and high xylanase production (2,012 ± 18 U/g) with poor cellulase activity (1.64 ± 0.12 U/g) was achieved at pH 6.0. However, substantial xylanase production was also observed at pH 5.0 (1,606 ± 7 U/g), 7.0 (1,924 ± 15 U/g), 8.0 (1,745 ± 13 U/g) and 9.0 (1,565 ± 19 U/g). These results indicated that A. tubingensis FDHN1 was not only grown successfully in acidic and alkaline pH but also generated considerable xylanase titre. However, xylanase production by A. tubingensis JP-1 has been observed to decrease drastically under acidic and alkaline pH ranges [20]. Liao et al. [26] have reported higher xylanase production by Penicillium oxalicum GZ-2 at pH 5.0. The highest xylanase has been produced by A. niveus RS2 at a neutral to alkaline pH than at an acidic and high alkaline pH [27].

In the present study, optimum xylanase production (1,998 ± 12 U/g) and specific activity (71.35 ± 2.97 U/mg) were noticed at 40°C. At the same temperature, cellulase activity was increased to 2.87 ± 0.16 U/g. The xylanase and cellulase activity data suggested that the temperature is an important parameter which regulates the enzyme production by A. tubingensis FDHN1. At higher temperatures, xylanase production declined sharply and it was 50% at 55°C (Table 2). Although, the physiological changes induced by high temperature during enzyme production are not completely understood. It has been suggested that at higher temperatures, microorganisms may synthesize only a reduced number of proteins essential for growth and other physiological processes [8]. It has been shown in a previous study that A. tubingensis JP-1 was unable to produce a considerable xylanase yield above 30°C [20]. The results of incubation time, pH and temperature study indicate that A. tubingensis FDHN1 has far better xylanase production ability across the pH and temperature range within short fermentation time as compared to earlier reported A. tubingensis JP-1.

Influence of carbon and organic/inorganic nitrogen sources

Carbon sources are very important for the growth and metabolic processes of microorganisms. The xylanase and cellulase production in the presence of different carbon sources are shown in Table 3. Addition of 0.5% xylose in the fermentation medium enhanced the xylanase production (2,267 ± 12 U/g) with low cellulase activity (1.85 ± 0.09 U/g). However, fructose, mannitol, lactose, maltose, glucose and sucrose were affected adversely to the xylanase production. The level of cellulase production was reached to 3.68 ± 0.07 U/g in the presence of glucose, but the xylanase production was affected negatively. Xylanase and cellulase are known to be inducible in fungi and are affected by the nature of the substrate used in the study [23]. To enhance the xylanase production, xylose was selected as the best carbon source and its concentration in medium was varied from 0.1 to 1.1%. The results indicated that 0.3% xylose gave high xylanase yield (2,349 ± 17 U/g) (Figure S2 in Additional file 1). It has been suggested that xylose is not only a carbon source but also an effective inducer for the xylanase production [28]. In the present study, xylose concentration above 0.3% reduced the xylanase production gradually. Such carbon source concentration-dependent enzyme production regulation has been noticed in different xylanase-producing microbial strains by various research groups [29,30].

Table 3 Effect of carbon sources on xylanase production from A. tubingensis FDHN1 under SSF

Formation of extracellular enzymes is greatly influenced by the availability of nitrogen source as it is the ultimate precursor of protein synthesis. Furthermore, nitrogen source can significantly affect the pH of the medium during the course of fermentation, and hence, the enzyme activity and stability may get influenced. Among the different organic nitrogen sources tested, gelatine was found to support the maximum xylanase production (2,453 ± 11 U/g). In this study, all the tested nitrogen sources enhanced the xylanase production as compared to control (Table 4). During the study with different inorganic nitrogen sources, sodium nitrate induced the xylanase production (2,589 ± 22 U/g). The xylanase production from Trichoderma viride in the presence of NaNO3 as an inorganic nitrogen source has also been reported by Goyal et al. [21]. In the present study, combination of gelatine and sodium nitrate further improved the xylanase (2,697 ± 18 U/g) and cellulase (3.86 ± 0.06 U/g) yield.

Table 4 Effect of nitrogen sources on xylanase production from A. tubingensis FDHN1 under SSF

Influence of metal additives and modulators

The choice of an appropriate additive is of great importance for the successful production of xylanase. Among tested metal compounds, MgCl2 exhibited the clear stimulation in xylanase yield (2,446 ± 27 U/g) followed by CuCl2, CaCl2, MnCl2 and FeCl3. However, HgCl2, CoCl2, SnCl2 and NiCl2 responded negatively towards xylanase production (Table S1 in Additional file 1). Similarly, addition of various modulators to the culture medium exerts a range of effects on the enzyme secretion. In the presence of Tween-80, the levels of cellulase and xylanase activities were increased to 4.21 ± 0.13 and 2,449 ± 23 U/g, respectively, which could be due to its favourable effect on cell permeability and thus it affects the secretion of certain proteins [31]. In addition, Tween-40, glycine and glycerol also showed its stimulatory effects on the xylanase synthesis.

Enzymatic properties of crude xylanase

Effect of pH on the activity and stability

Xylanase activity was studied in the wide range of pH 3.0 to 10 using different buffers to find out the best pH for the enzyme. The crude enzyme extract exhibited the maximum xylanase activity at pH 5.0 and retained more than 80% of its activity between pH 3. to 7.0 (Figure 2A). The xylanase activity was found to be affected slightly by variation in the pH outside its optimum pH. However, it decreased drastically at pH 9.0 and 10. Betini et al. [19] have measured xylanase activity at different pH 4.0 to 8.0. They have observed the optimum pH 5.0 to 5.5 for A. niveus, pH 5.5 to 6.0 for A. niger and pH 5.0 for Aspergillus ochraceus xylanase activities from crude enzyme extracts. During pH stability studies, A. tubingensis FDHN1 xylanase showed high stability towards a broad range of pH 3.0 to 9.0 (Figure 2B). The enzyme retained its 50% activity (t 50) after 9, 15 and 15 h of incubation in the acidic range pH 3.0, 4.0 and 5.0, respectively. At pH 6.0 and 7.0, xylanase showed good stability even after 24 h. However, at pH 7.0, 8.0 and 9.0, the half xylanase activities were noticed after 15, 9 and 6 h, respectively. These results indicate that xylanase from A. tubingensis FDHN1 is highly active and stable within a broad pH range as compared to the reported xylanases from several other Aspergillus sp. [19,32].

Figure 2
figure 2

Optimal pH (A) and pH stability (B) of the xylanase from A. tubingensis FDHN1. The influence of pH on xylanase activity was determined using 0.05 M sodium citrate (pH 3.0 to 6.0), sodium phosphate (pH 6.0 to 8.0) and glycine-NaOH (pH 8.0 to 10) buffers. The pH stability was studied at a regular interval of 3 up to 24 h.

Effect of temperature on the activity and stability

The effect of different temperatures on xylanase activity is shown in Figure 3A. The highest xylanase activity was observed at 50°C. In the present study, approximately 30 and 70% reduction in the xylanase activity were observed at 55 and 60°C, respectively. Enzyme activity reduced drastically with further increase in the temperature. The present xylanase exhibited more than 70% thermal stability at temperature range of 35 to 55°C after 60 min (Figure 3B), and it showed maximum stability at 45°C by retaining 85% residual activity after 120 min. The xylanase t 50 was observed after 60 and 90 min at 30 and 35°C, respectively. At 40, 45 and 50°C, t 50 was noticed after 195, 180 and 155 min, respectively. The xylanase retained 51, 53 and 50% activity even at temperature 55°C (90 min), 60°C (60 min) and 65°C (50 min), respectively. These results show that the activity and stability of xylanase at various temperatures were high as compared to the existing Aspergillus sp. xylanases [19,32,33].

Figure 3
figure 3

Optimal temperature (A) and thermal stability (B) of the xylanase from A. tubingensis FDHN1. Xylanase activity was measured at different temperatures to determine its optimal range. For the determination of thermal stability, the residual activity of the xylanase was measured after 30, 60, 90 and 120 min pre-incubation at different temperatures.

Effect of organic solvents on the activity

Published studies of the mechanism of adaptation of enzymes for functioning into the organic solvent are relatively few. In the present study, effects of various organic solvents and its concentration on the xylanase activity were examined (Figure S3 in Additional file 1). The xylanase retained more than 85% of its initial activity with all the solvents at 10% concentration except benzene. In the presence of 30% acetone, butanol and propanol, xylanase retained more than 70% of the initial activity after 30 min. This result is interesting with an aim of using the present xylanase in an application requiring the incubation of the enzyme with organic solvents. The mechanism of organic solvent tolerance has been investigated by Ogino et al. [34] in a Pseudomonas aeruginosa PST-01 protease by site-directed and random mutagenesis. They have reported that the disulfide bonds and amino acid residues located on the surface of the molecule play an important role in the organic solvent stability of the enzymes. Gupta et al. [35] have proposed that the presence of hydrophobic clusters on the protein surface and disulfide bonds was responsible for the solvent-stable nature of enzymes. Previously, Ines et al. [36] have studied the effect of various solvents up to 40% concentration on the activity of xylanase produced by Talaromyces thermophilus and observed maximum activity in the presence of 5% ethanol. However, reduced xylanase activity has been observed in the presence of methanol when its concentration was increased from 20 to 40%.

Native PAGE and xylan zymography

The crude enzyme from A. tubingensis FDHN1 was analysed by native PAGE and zymography. Figure 4 shows the presence of three clear zones upon xylan zymography, suggesting the presence of at least three xylanases. The approximate molecular weight of three xylanases, namely, Xyl-D1, Xyl-D2 and Xyl-D3 was found to be approximately 94, 30 and 18.60 kDa, respectively. This study for the first time highlights multiplicity of the xylanase from A. tubingensis. It is believed that the protein modification (e.g. post-translational cleavage) such as glycosylation, proteolysis or both could be responsible for the genesis of multiple enzymes [37]. Apart from post-translational modifications, multiplicity might also be due to the differential mRNA processing, post-secretional modification and the presence of different alleles of the same gene [3]. In 2010, two thermostable xylanases with a molecular weight of 25.2 and 30 kDa were identified by Sharma et al. from Malbranchea flava which were active under the alkaline conditions [38].

Figure 4
figure 4

Native PAGE and zymogram analysis of crude enzyme extract from A. tubingensis FDHN1. (A) Molecular weight marker, (B) crude enzyme extract, and (C) xylan zymography.

Thus, cellulase-poor xylanase from A. tubingensis FDHN1 seems a novel enzyme, being active and stable at broad pH and temperature ranges and in the presence of organic solvents. The multiplicity is also an interesting feature of this xylanase. Due to such desirable properties of xylanase, the present study was elaborated to achieve its better recovery from fermented sorghum straw by optimizing downstream process parameters.

Evaluation of downstream process parameters for maximum xylanase recovery

Selection of an appropriate extraction solvent

Recovery of the enzyme from solid support is another important aspect of SSF. An ideal solvent extracts the enzyme selectively and completely in minimum contact time. During initial solvent screening experiments, 0.05 M sodium citrate buffer (pH 6.5) was found suitable for efficient xylanase recovery (3,573 ± 22 U/g) with cellulase activity of 4.02 ± 0.08 U/g from the fermented sorghum straw. Extractions with all other solvents gave lower enzyme recoveries (Table 5). Interestingly, in this study, the xylanase was highly active and stable within the acidic range of pH and its optimum recovery was also found in the presence of an acidic extractant. This could be due to the fact that the use of suitable buffer solutions as a solvent helps in maintaining the pH-dependent enzyme stability. Earlier, distilled water and sodium citrate buffer 50 mM, pH 6.5 were used by Pal and Khanum [11] and Sonia et al. [18] as a solvent for the effective recovery of xylanase from cultivated biomass, respectively. Fernendez-Lahore et al. [39] have pointed that the hydrophobic or hydrophilic nature of fungal mycelia, ionic bonds, hydrogen bonds and van der Waal forces determines the efficiency of extractant for enzyme recovery.

Table 5 Effect of various extraction solvents on xylanase recovery from the fermented sorghum straw

Effect of extractant volume and extraction time

In this study, volumes of extractant and extraction time were optimized to improve the xylanase recovery. An extractant/solid ratio was kept in the range of 4:1 to 16:1 (v/w) while extraction time was varied from 15 to 120 min. During the study, the best xylanase extraction (3,769 ± 22 U/g) was obtained at an extractant/solid ratio of 12:1 (v/w) and the extraction was further improved (3,951 ± 17 U/g) for an extraction time of 90 min. Such increase in the xylanase extraction could be due to the maximum solubilization of the enzyme from the fermented sorghum straw (Figure 5A,B). At 12:1 (v/w) extractant/solid ratio, the cellulase activity was 4.89 ± 0.14 U/g which was reached to 5.17 ± 0.11 U/g at 90 min extraction. The improved xylanase recovery under the above optimized conditions is extremely important because concentrated crude extract ease downstream processes mainly the purification step, reducing the time and cost of enzyme recovery. The use of small volumes of solvent for xylanase recovery greatly reduces the energy requirements, equipment size and pollution, but the volumes that are too low leads to unsatisfactory recoveries, since a significant fraction of the xylanase remains in the fermented solid [40]. Previous studies have shown that 10 to 11 mL g−1 [11] and 17 mL g−1 [41] solvent/substrate ratio effectively recovers maximum xylanase that was produced by Aspergillus niger DFR-5 and Trichoderma longibrachiatum, respectively.

Figure 5
figure 5

Optimization of xylanase recovery parameters using 0.05 M sodium citrate buffer pH 6.5. (A) Xylanase recovery was performed using different volumes of extractant at 120 rpm for 60 min at 37°C, (B) xylanase recovery was performed at different time intervals using 12:1 (v/w) extractant/solid ratio at 120 rpm and 37°C, (C) xylanase recovery was performed at different agitation rates using 12:1 (v/w) extractant/solid ratio for 90 min at 37°C, and (D) xylanase recovery was performed at different temperatures using 12:1 (v/w) extractant/solid ratio at 150 rpm for 90 min.

Effect of agitation speed and extraction temperature

The xylanase extraction was also studied at different agitation speed and temperature values. During the study, 150 rpm was found suitable for the xylanase recovery (4,067 ± 27 U/g) which was further improved at 40°C (4,105 ± 22 U/g). The maximum cellulase activity under above optimized conditions was 5.36 ± 0.17 U/g (Figure 5C,D). In the present study, xylanase was most stable at 45°C and its optimum recovery was observed at 40°C suggesting that the enzyme is optimally active at moderately high temperatures. Ghildyal et al. [40] have reported that the prolonged higher rate of agitation causes a loss of enzyme activity. The maximum xylanase extraction has been observed at the highest rate of agitation (200 rpm) with an extraction time of 60 min at 37°C by Pal and Khanum [11]. However, 25°C and 200 rpm agitation speed for xylanase recovery has also been reported by Azin et al. [41].

Effect of un-optimized and optimized recovery conditions on xylanase recovery efficiency

After studying the detailed downstream process parameters, the efficiency of un-optimized and optimized recovery conditions for xylanase recovery were studied at different time intervals. The results indicated that xylanase recovery efficiency was 89.4% under the optimized conditions after 90 min, whereas it was only 53.3% under un-optimized conditions after 60 min (Figure S4 in Additional file 1). After 90 min, the recovery efficiency remained almost constant indicating that the maximum xylanase from the fermented sorghum straw was leached into the extraction solvent during this period. In the present study, the maximum xylanase recovery within a short time indicated the importance of detailed downstream process optimization.

The existing novel strain of A. tubingensis FDHN1 has been found to show ability to produce higher cellulase-poor xylanase yield under controlled optimized conditions. The overall productivity of xylanase by A. tubingensis FDHN1 has been compared with the existing xylanases produced by different Aspergillus sp. under SSF (Table 6).

Table 6 Comparison of A. tubingensis FDHN1 with different Aspergillus sp. for xylanase yield and productivity


The present study has revealed the potential of a newly isolated A. tubingensis FDHN1 to produce xylanase under SSF. The prominent induction of xylanase has been observed in the presence of the least highlighted agricultural residue sorghum straw. The optimization of upstream and downstream process parameters enhanced the xylanase yield from 615.5 ± 19.21 to 4,105 ± 22 U/g, suggesting the importance of the detailed bioprocess profiling. Apart from that cellulase-poor nature, high temperature, pH and solvent stability are suitable features for its application in paper, juice, feed improvement and in bioconversion of lignocelluloses to the solvents. The multiplicity in xylanase from A. tubingensis FDHN1 has been shown for the first time in the present study. Further scaling-up and purification of the xylanase still deserve more attention to reach the commercial feasibility and to find the novel properties of xylanase from this novel strain of A. tubingensis.



solid-state fermentation


potato dextrose agar

w/v :

weight by volume




xylanase/cellulase activity as unit per gram dry substrate


specific xylanase activity per milligram of protein

Native PAGE:

native polyacrylamide gel electrophoresis

v/w :

volume by weight




  1. Tseng MJ, Yap MN, Ratanakhanokchai K, Kyu KL, Chen ST (2002) Purification and characterization of two cellulase-free xylanases from an alkaliphilic Bacillus firmus. Enzyme Microb Technol 30:590–595

    Article  CAS  Google Scholar 

  2. Collins T, Gerday C, Fellre G (2005) Xylanases, xylanase families and extremophilic xylanases. FEMS Microbiol Rev 29:3–23

    Article  CAS  Google Scholar 

  3. Polizeli MLTM, Rizzatti ACS, Monti R, Terenzi HF, Jorge JA, Amori DS (2005) Xylanase from fungi: properties and industrial applications. Appl Microbiol Biotechnol 67:577–591

    Article  CAS  Google Scholar 

  4. Butt MS, Tahir-Nadeem M, Ahmad Z, Sultan MT (2008) Xylanases and their applications in baking industry. Food Technol Biotechnol 46:22–31

    CAS  Google Scholar 

  5. Nagar S, Mittal A, Kumar D, Kumar L, Gupta VK (2012) Immobilization of xylanase on glutaraldehyde activated aluminium oxide pellets for increasing digestibility of poultry feed. Process Biochem 47:1402–1410

    Article  CAS  Google Scholar 

  6. Coughlan MP, Hazlewood GP (1993) β-1, 4-D xylan-degrading enzyme systems: biochemistry, molecular biology and applications. Appl Biochem 17:259–289

    CAS  Google Scholar 

  7. Rao CS, Sathish T, Laxmi MM, Laxmi GS, Rao RS, Prakasham RS (2008) Modeling and optimization of fermentation factors for enhancement of alkaline protease production by isolated Bacillus circulans using feed-forward neural network and genetic algorithm. J Appl Microbiol 104:889–898

    Article  CAS  Google Scholar 

  8. Gawande PV, Kamat MY (1999) Production of Aspergillus xylanase by lignocellulosic waste fermentation and its application. J Appl Microbiol 87:511–519

    Article  CAS  Google Scholar 

  9. Park Y, Kang S, Lee J, Hong S, Kim S (2002) Xylanase production in solid state fermentation by Aspergillus niger mutant using statistical experimental designs. Appl Microbiol Biotechnol 58:761–766

    Article  CAS  Google Scholar 

  10. Maciel GM, Vandenberghe LPS, Haminiuk CWI, Fendrich RC, Bianca BED, Brandalize TQS, Pandey A, Soccol CR (2008) Xylanase production by Aspergillus niger LPB 326 in solid state fermentation using experimental designs. Food Technol Biotechnol 46(2):183–189

    CAS  Google Scholar 

  11. Pal A, Khanum F (2010) Production and extraction optimization of xylanase from Aspergillus niger DFR5 through solid state fermentation. Bioresour Technol 101:7563–7569

    Article  CAS  Google Scholar 

  12. Bailey MJ, Biely P, Poutanen K (1992) Interlaboratory testing of methods for assay of xylanase activity. J Biotechnol 23:257–270

    Article  CAS  Google Scholar 

  13. Ghose TK (1987) Measurement of cellulase activities. Pure Appl Chem 59:257–268

    CAS  Google Scholar 

  14. Miller LG (1959) Use of dinitrosalicylic acid reagent for determination of reducing sugars. Anal Chem 31:426–428

    Article  CAS  Google Scholar 

  15. Lowry OH, Rosebrough NJ, Farr AL, Randall RJ (1951) Protein measurement with Folin phenol reagent. J Biol Chem 31:426–428

    Google Scholar 

  16. Laemmli UK (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227(5259):680–685

    Article  CAS  Google Scholar 

  17. Puls J, Suhuseil J (1993) Chemistry of hemicelluloses: relationship between hemicellulose structure and enzyme required for hydrolysis, in hemicellulose and hemicellulases. Portland Press, London, pp 103–127

    Google Scholar 

  18. Sonia KG, Chadha BS, Saini HS (2005) Sorghum straw for xylanase hyper production by Thermomyces lanuginosus (D2W3) under solid state fermentation. Bioresour Technol 96:1561–1569

    Article  CAS  Google Scholar 

  19. Betini JHA, Michelin M, Peixoto-Nogueira SC, Jorge JA, Terenzi HF, Polizeli MLTM (2009) Xylanase from Aspergillus niger, Aspergillus niveus and Aspergillus ochraceus produced under solid-state fermentation and their application in cellulose pulp bleaching. Biprocess Biosyst Eng 32:819–824

    Article  CAS  Google Scholar 

  20. Pandya JJ, Gupte A (2012) Production of xylanase under solid state fermentation by Aspergillus tubingensis JP-1 and its application. Bioprocess Biosyst Eng 35:769–779

    Article  CAS  Google Scholar 

  21. Goyal M, Kalra KL, Sareen VK, Soni G (2008) Xylanase production with xylan rich lignocellulosic wastes by a local soil isolate of Trichoderma viride. Braz J Microbiol 39:535–541

    Article  Google Scholar 

  22. Biswas R, Sahai V, Mishra S, Bisaria VS (2010) Bioprocess strategies for enhanced production of xylanase by Melanocarpus albomyces IITD3A on agro-residual extract. J Biosci Bioeng 110(6):702–708

    Article  CAS  Google Scholar 

  23. Kang SW, Park YS, Lee JS, Hong SI, Kim SW (2004) Production of cellulases and hemicellulases by Aspergillus niger KK2 from lignocellulosic biomass. Bioresour Technol 91:153–156

    Article  CAS  Google Scholar 

  24. Hasseltine CW (1972) Solid state fermentations. Biotechnol Bioeng 14:517–532

    Article  Google Scholar 

  25. Prakasham RS, SubbaRao C, Rao RS, Rajesham S, Sarma PN (2005) Optimization of alkaline protease production by Bacillus sp. using Taguchi methodology. Appl Biochem Biotechnol 120:133–144

    Article  CAS  Google Scholar 

  26. Liao H, Xu C, Tan S, Wei Z, Ling N, Yu G, Raza W, Zhang R, Xu QSY (2012) Production and characterization of acidophilic xylanolytic enzymes from Penicillium oxalicum GZ-2. Bioresour Technol 123:117–124

    Article  CAS  Google Scholar 

  27. Sudan R, Bajaj BK (2007) Production and biochemical characterization of xylanase from an alkalitolerant novel species Aspergillus niveus RS2. World J Microbiol Biotechnol 23:491–500

    Article  CAS  Google Scholar 

  28. Priem B, Dobberstein J, Emies CC (1991) Production of β-1,4- xylanase in continuous culture by Aureobasidium pullulans CBS 58475. Biotechnol Lett 13:149–154

    Article  CAS  Google Scholar 

  29. Lakshmi GS, Rao CS, Rao RS, Hobbs PJ, Prakasham RS (2009) Enhanced production of xylanase by newly isolated Aspergillus terreus under solid state fermentation using palm industrial waste: a statistical optimization. Biochem Eng J 48:51–57

    Article  CAS  Google Scholar 

  30. Oliveira LA, Porto ALF, Tambourgi EB (2006) Production of xylanase and protease by Penicillium janthinellum CRC 87-M- 115 from different agricultural wastes. Bioresour Technol 97:862–867

    Article  CAS  Google Scholar 

  31. Bakri Y, Mohammed J, Mohammed IEA (2008) Improvement of xylanase production by Cochliobolus sativus in submerged culture. Food Tehnol Biotechnol 46(1):116–118

    CAS  Google Scholar 

  32. Delabona PS, Pirota RDPB, Codima CA, Tremacoldi CR, Rodrigues A, Farinas CS (2013) Effect of initial moisture content on two rainforest Aspergillus strains cultivated on agro-industrial residues: biomass-degrading enzymes production and characterization. Ind Crop Prod 42:236–242

    Article  CAS  Google Scholar 

  33. Lu FX, Lu M, Lu ZX, Bie XM, Zhao HZ, Wang Y (2008) Purification and characterization of xylanase from Aspergillus ficcum AF-98. Biresour Technol 99:5938–5941

    Article  CAS  Google Scholar 

  34. Ogino H, Uchiho T, Yokoo J, Kobayashi R, Ichise R, Ishikawa H (2001) Role of intermolecular disulfide bonds of the organic solvent-stable PST-01 protease in its organic solvent stability. Appl Environ Microbiol 67:942–947

    Article  CAS  Google Scholar 

  35. Gupta A, Ray S, Kapoor S, Khare SK (2008) Solvent-stable Pseudomonas aeruginosa PseA protease gene: identification, molecular characterization, phylogenetic and bioinformatic analysis to study reasons for solvent stability. J Mol Microbiol Biotechnol 15:234–243

    Article  CAS  Google Scholar 

  36. Ines MA, Mohamed G, Ines BBR, Ali G, Hafedh B (2012) The effect of Talaromyces thermophilus cellulase-free xylanase and commercial laccase on lignocellulosic components during the bleaching of kraft pulp. Int Biodet Biodeg 75:43–48

    Article  CAS  Google Scholar 

  37. Li XL, Ljungdahl LG (1994) Cloning, sequencing and regulation of a xylanase gene from the fungus Aureobasidium pullulans Y-2311-1. Appl Environ Microbiol 60:3161

    Google Scholar 

  38. Sharma M, Chadha BS, Saini HS (2010) Purification and characterization of two thermostable xylanases from Malbranchea flava under alkaline conditions. Bioresour Technol 101:8834–8842

    Article  CAS  Google Scholar 

  39. Fernendez-Lahore HM, Fraile ER, Cascone O (1998) Acid protease recovery from a solid state fermentation system. J Biotechnol 62:83–93

    Article  Google Scholar 

  40. Ghildyal NP, Ramakrishna M, Lonsane BK, Karanth NG (1991) Efficient and simple extraction of mouldy bran in a pulsed column extractor for recovery of amyloglucosidase in concentrated form. Process Biochem 26:235–241

    Article  CAS  Google Scholar 

  41. Azin M, Moravej R, Zareh D (2007) Production of xylanase by Trichoderma longibrachiatum on a mixture of wheat bran and wheat straw: optimization of culture conditions by Taguchi method. Enzyme Microb Technol 40:801–805

    Article  CAS  Google Scholar 

Download references


One of the authors (DNA) gratefully acknowledges the financial assistance from the Department of Science and Technology (DST), Ministry of Science and Technology, Govt. of India, for the award of Junior Research Fellowship under the INSPIRE Program during the course of the investigation. The authors would like to thank Mr. Mahesh Bhatt, Gujarat Arts & Science College, Ahmedabad, for the language editing.

Author information

Authors and Affiliations


Corresponding author

Correspondence to Nikhil S Bhatt.

Additional information

Competing interests

The authors declare that they have no competing interests.

Authors' contributions

DNA designed experiments, carried out the laboratory work and wrote the manuscript. NSB provided supervision and research direction of the experimental work, as well as editing the manuscript. HAM critically observed the results and finalized the manuscript draft. All authors contributed intellectually via scientific discussions during the study and read and approved the final manuscript.

Authors' information

DNA is registered as a Ph.D. Fellow and is also DST-INSPIRE-JRF at P. G. Department of Microbiology, Gujarat Vidyapeeth, Sadra- 382 320, Gujarat, India. NSB is a Professor at P. G. Department of Microbiology, Gujarat Vidyapeeth, Sadra- 382 320, Gujarat, India. HAM is a Reader at the Department of Life Sciences, School of Sciences, Gujarat University, Ahmadabad, Gujarat, India.

Additional file

Additional file 1: Table S1 and Figures S1 to S4.

Table S1. Effect of metals and modulators on xylanase production from A. tubingensis FDHN1 under SSF. Figure S1. Effect of sorghum straw concentration (3 to 15 g) on xylanase production. SSF was carried out by moistening sorghum straw with mineral salt medium pH 5.0 at 35°C for 6 days. Figure S2. Effect of xylose concentration (0.1 to 1.1%) on xylanase production. SSF was carried using 9 g sorghum straw under optimized physiological conditions (at 5 days incubation, 1:5 (w:v) substrate: moisture ratio, pH 6.0 and 40°C). Figure S3. Effect of organic solvents (10 to 30%) on the xylanase activity. The crude xylanase was incubated in the presence of various organic solvents concentration for 30 min and at the end of reaction residual activities were measured. Figure S4. Comparison of the un-optimized and optimized extraction conditions on the xylanase recovery efficiency (%). Optimized extraction conditions were extractant/ solid ratio 12: 1 (v:w), 150 rpm agitation speed and 40°C. The data presented are the mean values of three replicates with the standard deviations.

Rights and permissions

Open Access  This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made.

The images or other third party material in this article are included in the article’s Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

To view a copy of this licence, visit

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Adhyaru, D.N., Bhatt, N.S. & Modi, H.A. Optimization of upstream and downstream process parameters for cellulase-poor-thermo-solvent-stable xylanase production and extraction by Aspergillus tubingensis FDHN1. Bioresour. Bioprocess. 2, 3 (2015).

Download citation

  • Received:

  • Accepted:

  • Published:

  • DOI: