Catalytic conversion of sugarcane bagasse to cellulosic ethanol: TiO2 coupled nanocellulose as an effective hydrolysis enhancer

Carbohydrate Polymers 136 (2016) 700–709

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Catalytic conversion of sugarcane bagasse to cellulosic ethanol: TiO2 coupled nanocellulose as an effective hydrolysis enhancer S. Anuradha Jabasingh a,∗ , D. Lalith b , M. Arun Prabhu b , Abubekker Yimam a , Taye Zewdu a a b

Process Engineering Division, School of Chemical and Bio Engineering, Addis Ababa Institute of Technology, Addis Ababa University, Ethiopia Chemical Engineering Department, Sathyabama University, Tamilnadu 600119, India

a r t i c l e

i n f o

Article history: Received 4 May 2015 Received in revised form 17 September 2015 Accepted 26 September 2015 Available online 30 September 2015 Keywords: Cellulosic ethanol Bagasse Titanium dioxide Nanocellulose Cellulase Saccharomyces cerevisiae

a b s t r a c t The present study deals with the production of cellulosic ethanol from bagasse using the synthesized TiO2 coupled nanocellulose (NC-TiO2 ) as catalyst. Aspergillus nidulans AJSU04 cellulase was used for the hydrolysis of bagasse. NC-TiO2 at various concentrations was added to bagasse in order to enhance the yield of reducing sugars. Complex interaction between cellulase, bagasse, NC-TiO2 and the reaction environment is thoroughly studied. A mathematical model was developed to describe the hydrolysis reaction. Ethanol production from enzymatically hydrolyzed sugarcane bagasse catalyzed with NC-TiO2 was carried out using Saccharomyces cerevisiae ATCC 20602. The glucose release rates and ethanol concentrations were determined. Ethanol produced was found to be strongly dependent on pretreatment given, hydrolysis and fermentation conditions. The study confirmed the promising accessibility of NC-TiO2 , for enhanced glucose production rates and improved ethanol yield. © 2015 Elsevier Ltd. All rights reserved.

Chemical compounds studied in this article: Ethyl alcohol (PubChem CID: 702) Titanium dioxide (PubChem CID: 26042) Na-carboxymethyl cellulose (PubChem CID: 6328154) d-Glucose anhydrous (PubChem CID: 5793) Sulphuric acid (PubChem CID: 1118) Sodium propionate (PubChem CID: 2723816) Citric acid (PubChem CID: 311) Sucrose (PubChem CID: 5988) Ammonium hydroxide (PubChem CID: 14923) Dinitrosalicylic acid (PubChem CID: 89779)

1. Introduction Fossil fuels, on which most life activities depend, are formed and deposited over millions of years. Limitation in their stock results in the crisis of depletion. In addition to this, carbon dioxide emissions during the combustion of fossil fuels are another source of major concern. Fuel ethanol is suggested as a sustainable fuel, which when blended with gasoline is reported to increase the octane number

∗ Corresponding author at: Process Engineering Division, E-104, School of Chemical and Bio Engineering, Addis Ababa Institute of Technology, Addis Ababa University, Ethiopia. E-mail address: [email protected] (S.A. Jabasingh). http://dx.doi.org/10.1016/j.carbpol.2015.09.098 0144-8617/© 2015 Elsevier Ltd. All rights reserved.

and provide oxygen to promote a complete combustion. Addition of ethanol to gasoline is found to reduce tailpipe emissions of CO and un-burnt hydrocarbons, which can contribute to improve the air quality. Ethanol is water-soluble and biodegradable, comparatively harmless to the environment, ground water and soil (Baeyens et al., 2015; Isenberg, 1999). Though, recent research shows the production of ethanol from various sources including wheat straw (Zhu et al., 2015), Kans grass (Kataria & Ghosh, 2011), cotton stalk (Binod et al., 2012), oil palm residues (Samsudin & Don, 2015), corn stover (Kim, Wi, Jung, Song, & Hyeun-Jong, 2015; Kim, Seo, Kim, & Han, 2015), low-cost agroindustrial by-products (Kelbert et al., 2015) and red seaweed Gelidium amansii (Kim, Wi, et al., 2015; Kim, Seo, et al., 2015), sugarcane bagasse remains the foremost feedstock (Nasidi, Agu, Deeni, & Walker, 2015; Wu et al., 2011).

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Cellulosic ethanol is a promising renewable fuel due to its carbon neutral characteristics, reduction of particulate and NOx emission (Wyman, 1999). This energy resource can be used as a substitute for petroleum-derived fuels (McMillan, 1997). Though, cellulosebased ethanol substantially decreases greenhouse gas emissions, the emission of pollutants, including VOCs and NOx may increase, due to the substitution of gasoline with cellulosic ethanol (Granda, Zhu, & Holtzapple, 2007). Nevertheless, cellulosic ethanol seems to be a promising choice from the perspectives of both net energy gain and overall emissions of contaminants (Von Blottnitz & Curran, 2007). The tight spot lies in the hinderance caused by incomplete conversion of cellulose to ethanol, due to the effect of its structural parameters and insolubility. There are many methods to hydrolyze cellulose into glucose, which is the fermentable sugar needed to produce cellulosic ethanol. Three major pretreatment processes are typically adopted to produce a variety of sugars suitable for ethanol production from lignocellulosic biomass. They are dilute acid hydrolysis, concentrated acid hydrolysis and enzymatic hydrolysis, of which, enzymatic hydrolysis is the most common (Kumar, Barrett, Delwiche, & Stroeve, 2009; Vallejos, Felissia, Kruyeniskia, & Area, 2015). This facet of a search for a better cellulolytic microorganism capable of efficient saccharification of lignocellulosic biomass, instigated the priority to be given for an already proven strain, A. nidulans AJSU04 cellulase (Anuradha Jabasingh, Varma, & Garre, 2014). In addition to cellulase being employed for hydrolysis, the cellulosic structure of bagasse, employed in this study, is modified and the effect of surface modification on the rate of enzymatic hydrolysis is investigated. A novel material, called nanocellulose (NC), was prepared using 80% sulfuric acid and modified with TiO2 . The resulting material NC-TiO2 , was studied as a possible glucose enhancer to increase the enzyme accessibility. Fermentable sugars produced by hydrolysis of cellulose using cellulase enzyme can easily be fermented into ethanol using yeast. Both separate hydrolysis and fermentation (SHF) and simultaneous saccharification and fermentation (SSF) processes are mostly accepted technologies for bioethanol production from lignocellulosic biomass. In SHF, hydrolysis and fermentation steps are carried out sequentially and separately, at optimal conditions of pH and temperature. In SSF, both steps are operated simultaneously in a single vessel, thus glucose released by the action of cellulases is converted directly to ethanol by fermenting microorganism and continuous removal of sugars from medium minimizes the feedback inhibition on enzyme activity (Singhania et al., 2014). Though, SSF process has shown to be superior to SHF in terms of overall ethanol yield, the differences between the optimal temperature for cellulase activity and yeast augmentation is an issue that needs to be solved for an efficient SSF as the optimal temperature for cellulase enzymes (about of 50 ◦ C) is higher than the tolerance range reported by yeast for industrial ethanol production (about 30–37 ◦ C) (Alfani, Gallifuoco, Saporosi, Spera, & Cantarella, 2000; Ruiz et al., 2012). This necessitates modulatory temperature conditions for the optimum performance of enzymatic and fermenting microorganism. Therefore, the present study emphasizes on separate SHF to achieve a maximum ethanol concentration. Furthermore, the emphasize was to observe the catalytic activity of the NC-TiO2 during hydrolysis, which could be better observed in SHF than in SSF. Economic obstacles, including the slow kinetics of enzymatic hydrolysis and the subsequent rate reduction during reaction progression, as well, hinder the industrial production of ethanol from biomass (Dasari & Berson, 2007; Dunaway, Dasari, Bennett, & Berson, 2010; Nidetzky & Steiner, 1993; Valjamae, Sild, Pettersson, & Johansson, 1998; Yang, Willies, & Wyman, 2006; Zhang, Xu, Xu, Yuan, & Guo, 2010). The properties of the lignocellulosic biomass such as crystallinity (Gharpuray, Lee, & Fan, 1983; Peitersen & Ross,

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1979; Zhu, O’Dwyer, Chang, Granda, & Holtzapple, 2010), accessibility (Asenjo, 1983), and reactivity (Drissen et al., 2007; Hong, Ye, & Zhang, 2007) affect the reaction rate. Other factors, including the loss in the activity of sorbed enzyme, due to deactivation (Converse, Matsuno, Tanaka, & Taniguchi, 1988; Howell & Mangat, 1978; Shen & Agblevor, 2008) and non-productive binding (Jalak & Valjamae, 2010) also contribute to the decline in the rate of enzymatic reaction. In this study, a mathematical model was developed to describe the variation of NC-TiO2 -B matrix during the hydrolysis. Howell and Mangat (1978) and Converse et al. (1988) also developed models to account for the slow kinetics of cellulose hydrolysis by considering activation of the adsorbed cellulase. Howell and Mangat (1978) proposed a model based on Michaelis–Menten kinetics, which is not appropriate for heterogeneous cellulose hydrolysis (Lynd, Weimer, van Zyl, & Pretorius, 2002). Converse et al. (1988) assumed direct proportionality of cellulase-substrate adsorption to the second order of substrate concentration, while it has not been validated by experiments. In this attempt, firstly, Langmuir adsorption, which has been broadly used and well accepted by researchers was employed to derive the hydrolysis rate equation (Kumar & Wyman, 2008; Kyriacou, Neufeld, & Mackenzie, 1988; Nidetzky, Steiner, & Claeyssens, 1994; Tu, Chandra, & Saddler, 2007) and secondly, model parameter values were determined by fitting the equations to experimental hydrolysis data. More significantly, this work investigated the possibility of improving biofuel production by the inclusion of TiO2 nano particles in the nanocellulose to produce TiO2 coupled nanocellulose, NC-TiO2 . In this context, this work aims for the intensification of hydrolysis, saccharification and fermentation processes combined with catalytic effect of NC-TiO, to attain robust ethanol production from sugarcane bagasse with S. cerevisiae ATCC 20602. 2. Materials and methods 2.1. Enzyme and reagents A. nidulans AJSU04 strain, employed previously in a solid-state fermenter was used for cellulase production (Anuradha Jabasingh et al., 2014). S. cerevisiae ATCC 20602, was procured from American Type Culture Collection, Manassas, VA, USA. Na-carboxy methyl cellulose (Na-CMC), microcrystalline cellulose (MCC) and titanium dioxide were purchased from Sigma Chemicals (Mumbai, India). Diluted solution of Na-CMC was prepared using sodium citrate buffer. Diluted solutions of d-glucose anhydrous (purity ≥99.0%), obtained from Merck (Mumbai, India) were used as standards. All other reagents were of analytical grade. 2.2. Lignocellulosic substrate Bagasse was obtained from local sugar mills in Tamilnadu. The substrate was washed with water at room temperature for 7 h, shredded and air dried. The initial composition of sugarcane bagasse was determined in %: cellulose, 42.7; hemi-cellulose, 25.4; lignin, 28.0; ash, 0.75% (Goering & Van Soest, 1970; Anuradha Jabasingh, 2011a). Alkaline pretreatment with NaOH was used to delignify bagasse (Jung et al., 2008). 10 tons dry weight bagasse was subjected to pretreatment with 8 L of 1 N NaOH solution in a digester for 8 h at 50 ◦ C. After treatment, the solution was filtered and solid fraction was suspended in hot boiling water for 10 min and subjected to drying in a tray drier. The solid fraction was dried and stored in air tight polythene bags. 2.3. Synthesis of TiO2 coupled nanocellulose About 20 mg of microcrystalline cellulose (MCC) powder was mixed with 150 mL of 80% (w/w) H2 SO4 using Evolution 6300

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Dissolution System at 4 ◦ C with 550 rpm agitation. A grayish-white material obtained after 30 min was precipitated by adding 100 mL of cold ethanol (−17 ◦ C) to allow efficient precipitation. The product was collected and centrifuged at 10 ◦ C thrice for 15 min at 3000 rpm(Heraeus Labofuge 200 Centrifuge). The product obtained is known as NC, nanocellulose. To this precipitate, 50 mL of TiO2 solution was added and subjected to sonication at 50% amplitude for 10 min. The product is collected, centrifuged at 5 ◦ C thrice for 20 min at 2000 rpm. The precipitate was collected and dialyzed for 12 h to ensure the neutral pH (Abushammala & Hashaikeh, 2011). The suspension obtained was further sonicated (Qsonica Q700 Sonicator) for 20 min (50% amplitude for 10 min, shut off for 5 min), re-started at 25% amplitude for 10 min followed by dialysis and freeze drying using Cole-Parmer Benchtop Freeze Dry System (Model: EW-03337-00) for 3 h. The material obtained was known as TiO2 coupled nanocellulose, NC-TiO2 (Hashaikeh & Abushammala, 2011). The MCC, NC and NC-TiO2 were tested for their efficacy to yield reducing sugars and their surface features analyzed using SEM. 2.4. Saccharification of substrate Solid-state fermentation (SsF) was performed in a solid-state fermenter employing pretreated bagasse and 0.005% NC-TiO2 with 60% moisture content, pH 5, at temperature 40 ◦ C for 11 days. The detailed description of the fermenter employed in the present study is provided elsewhere (Anuradha Jabasingh et al., 2014). A. nidulans AJSU04 spore suspension prepared by washing slant cultures with 5 mL sterilized water was transferred into sterilized minimal medium (mg L−1 ), KH2 PO4 , 0.75; NH4 NO3 , 1.5; thiamine HCl, 0.01; MgSO4 ·H2 O, 0.5; CaCl2 ·5H2 O, 0.05 at pH 5. The products of hydrolysis were assayed for cellulase production after 24 h. Extracts were drawn every 24 h by the addition of 10 mL distilled water to the solid bed, pressing, followed by filtering the extract through nylon cloth. The cellulase activity was analyzed in terms of FPU in accordance with the standard analytical methods (Ghose, 1987). One filter paper unit (FPU) of cellulase was defined as the amount of enzyme which produces 2.0 mg of reducing sugar from 50 mg of filter paper within 1 h. The enzyme activity of A. nidulans AJSU04 cellulase was 51.98 FPU/mL. A. nidulans AJSU04 cellulase (5 ␮L) was employed in the Lambda minifor laboratory bioreactor, 3 L containing 4800 mL of fermentation (minimal) medium for glucose production. 2.5. Inoculum preparation for ethanol production Yeast mold (YM) agar medium in %: yeast extract, 3; malt extract, 3; peptone, 5; dextrose, 10; agar, 20, was used to culture the yeast strain in sterile petriplates. Sterile citric acid is added to the sterile molten medium and cooled to 45–50 ◦ C. Spore suspension was prepared and transferred by washing slant cultures with 5 mL sterilized water into YM medium and inoculated on a rotary shaker for 3 days (Anuradha Jabasingh & Valli Nachiyar, 2012a). Sodium propionate, 0.01% is added to eliminate molds and permit the enumeration of S. cerevisiae ATCC 20602. Seed culture was prepared in a 1-L Erlenmeyer flask containing 200 mL YM growth medium supplemented with constituents in %: sucrose, 50; Na-CMC, 32; soy meal, 0.2; (NH4 )2 SO4 , 0.1; KH2 PO4 , 0.05; FeSO4 ·7H2 O, 0.003 using 1.0 mL frozen suspension followed by incubation at 30 ◦ C for 16 h at 200 rpm in an orbital shaker(Anuradha Jabasingh & Valli Nachiyar, 2012b). 2.6. Ethanol production 2% w/w concentrations of S. cerevisiae ATCC 20602 inoculum was transferred to CelliGen 310, a 2.5 L bioreactor containing 2 L

of fermentation medium (YM growth medium supplemented with constituents). The fermenter system equipped with containers for base solution, feed medium, a collection reservoir, pumps, fermenter vessel with pH, dissolved oxygen and redox probes, and a nitrogen gas tank is operated in a batch mode. The fermenter was controlled at 30 ◦ C and pH 5.0. Aeration rate was set at 1.5 L/min (0.5 vessel volumes per minute). Agitation speed was maintained at 300 rpm. pH was controlled using a 29% NH4 OH base solution. Glucose obtained from the previous SSF, described in Section 2.4, was used as the feed medium. Dissolved oxygen and redox potential were monitored on a continuous basis and dissolved oxygen was controlled at 20% during the growth phase (Dawson & Boopathy, 2007; Kwan, Hwang, Lee, Ahn, & Park, 2007). Cell viability and cell concentration were monitored during the entire fermentation process (Gregg & Saddler, 1996). The anaerobic production phase was maintained using nitrogen gas (Palmqvist, Almeida, & HahnHagerdal, 1999). Ethanol production and glucose concentrations were measured off-line. After fermentation, the fermented mash was centrifuged for 10 min at 10,000 r/min (3 ◦ C) in a refrigerated Heraeus Labofuge 200 centrifuge and the supernatant was used for determination of bioethanol content, expressed in g/L (AOAC, 2000). 2.7. Analytical methods The residual glucose concentration during hydrolysis is measured by dinitrosalicylic acid method using UV–vis spectrophotometer (Hitachi Model: 100-40 Spectrophotometer) at max 540 nm (Miller, 1959). During ethanol fermentation, the glucose concentration was measured by high performance liquid chromatography (HPLC) using Agilent 1100 system (Palo Alto, CA). The sugars were separated on a 8 ␮m Hi-Plex H, 30 cm × 7.7 mm at room temperature and detected with a refractive index detector. Deionized water was used as mobile phase at a flow rate of 0.6 mL/min. Ethanol was quantified using Quantichrom TM Ethanol assay kit (DIET-500). The resulting bluish chromic product was analyzed at max 580 nm. Further quantification was carried out using gas chromatography (GC), equipped with flame ionization detector, and a column of Carbowax 20 M (Supelco, USA) at 90 ◦ C. Injector and detector temperature was maintained at 140 ◦ C. The surface morphology was determined by subjecting the samples to scanning electron microscopy (SEM) using Philips XL30 scanning electron microscope with electron acceleration voltage of 20 kV and probe current of 5 × 10−11 A after gold sputtering in a denton vacuum desk I for 1.5 min under a 200 m Torr Argon atmosphere and 30 mA current. 3. Results and discussion 3.1. Yield of reducing sugar using TiO2 coupled nanocellulose The effect of the yield of reducing sugar by employing MCC, NC and NC-TiO2 indicate the superiority of NC-TiO2 over MCC and NC. 1 ␮g of MCC, NC and NC-TiO2 were added to 20 mg of bagasse each layered in 250 mL sterilized Erlenmeyer flasks fitted with a threeway Claisen Adapter. 15 ␮L cellulase is added to each of the three Erlenmeyer flasks containing a combination of bagasse with MCC, bagasse with NC and bagasse with NC-TiO2 respectively. SsF was performed separately in three flasks (60% moisture content, pH 5, 40 ◦ C). The amount of glucose yielded during the course of fermentation was analyzed. It was observed that there was a maximum yield of reducing sugar at the end of 96 h, 64 h and 36 h in three flasks, in the order of the constituents – bagasse with MCC, bagasse with NC and bagasse with NC-TiO2 respectively. The yield of glucose in the three SsF vessels, with a combination of bagasse with

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Fig. 1. Surface morphology of microcrystalline cellulose (MCC) (a), nanocellulose (NC) (b), and TiO2 coupled nanocellulose, NC-TiO2 (c).

Fig. 2. Surface morphology of TiO2 coupled nanocellulose (NC-TiO2 ) subjected to hydrolysis for 0 min (a), 5 min (b), 20 min (c), 30 min (d), 50 min (e) and 70 min (f).

MCC, bagasse with NC and bagasse with NC-TiO2 were 221 ␮g/mL, 439 ␮g/mL and 612 ␮g/mL respectively. The reason for the acceleration in the glucose yield in ascending order through the course of MCC, NC and NC-TiO2 could be the complex networked structure,

created on MCC by concentrated sulfuric acid during the NC synthesis. Further, during the synthesis of NC-TiO2 , mixing with TiO2 suspension regenerates the cellulose chains around the TiO2 particles and thus affects the structure. In conclusion, acceleration of the

Fig. 3. Surface morphology of bagasse (a), TiO2 coupled nanocellulose (b), integrated NC-TiO2 -bagasse matrix before hydrolysis (c) and NC-TiO2 -bagasse matrix subjected to hydrolysis for 50 min (d).

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Fig. 4. Concentration of glucose in the order of the composition of constituents, bagasse with MCC, bagasse with NC and bagasse with NC-TiO2 (a), effect of NC-TiO2 dosage on the reducing sugar yield (b), effect of initial cellulase concentration on the reducing sugar yield (c), and effect of hydrolysis time on the reducing sugar yield (d). Results are mean of triplicates in (a)–(d).

glucose yield is due to the enhanced hydrolysis of bagasse by NCTiO2 . Hence, the addition of NC-TiO2 with bagasse had a significant effect on the medium constituents both in terms of the duration of glucose yield and its quantity (Galbe & Zacchi, 2007; Hashaikeh & Abushammala, 2011). It was observed that the dispersed structure of NC-TiO2 was responsible for the significant enhancement in the overall enzymatic hydrolysis of feedstock supplemented with NCTiO2 in addition to a huge improvement in the initial hydrolysis rate. The yield of reducing sugar in the order of the composition of the constituents, bagasse with MCC, bagasse with NC and bagasse with NC-TiO2 are shown in Fig. 4a. Based on this study, there was a significant enhancement in the reducing sugar yield, effected by NC-TiO2 , which can be termed as a nanocatalyst, for it augments the hydrolyzing activity of the enzyme, cellulase. Henceforth, the effect of the NC-TiO2 dosage on the reducing sugar yield is of much interest. 3.2. Effect of NC-TiO2 dosage on production of glucose SsF studies were carried out using NC-TiO2 dosages (1–6 ␮g) incorporated into 20 mg of bagasse each layered in 250 mL Erlenmeyer flasks fitted with a three-way Claisen Adapter at conditions mentioned in Section 3.1. 15 ␮L of cellulase is added to each of the six Erlenmeyer flasks containing a combination of bagasse with NC-TiO2 . Concentration of glucose during the course of fermentation was analyzed. The effect of NC-TiO2 dosage on the reducing sugar yield is shown in Fig. 4b. It was found that the surface area available for the hydrolysis is limited for a specific dosage of NCTiO2 (Fan & Lee, 1983). The less hydrolysis of lignocellulosics at low concentration was due to the reduced surface area exposed to the access of cellulase. It is clear from Fig. 4b that the glucose yield increased from 620 ␮g/mL to 1563 ␮g/mL for the increase in the NC-TiO2 dosages from 1 ␮g to 6 ␮g. This is owed to the availability of more surface area and binding sites for complexation and hydrolysis. However, increasing the NC-TiO2 dosage above 6 ␮g had a very slight influence on hydrolysis. This effect may be due to the

interference caused by NC-TiO2 particles toward their access to bagasse. Hence, further addition of NC-TiO2 above 6 ␮g was considered to be economically unbefitting for the hydrolysis of bagasse. 3.2.1. Effect of cellulase concentration on production of glucose The initial concentration of cellulase provides an important driving force to overcome mass transfer resistance between aqueous cellulase enzyme and solid phases, which includes the integrated bagasse-NC-TiO2 structure (Fig. 4c). The initial cellulase concentrations were varied from 3 ␮g/mL to 20 ␮g/mL and added into 20 mg of bagasse each layered in 250 mL Erlenmeyer flasks fitted with a three-way Claisen Adapter. Studies were carried out using NC-TiO2 dosage of 6 ␮g integrated into bagasse in each flasks. SsF conditions were maintained as mentioned previously in Section 3.1. The glucose concentration was analyzed during the course of fermentation. The mass transfer of the cellulase enzyme is usually characterized by the external mass transfer (boundary layer diffusion) or by intraparticle diffusion or both for a solid–liquid hydrolysis. Mechanism of hydrolysis involves enzyme transport from the bulk solution through the liquid film to the surface of the integrated bagasse – NC-TiO2 exterior surface (Anuradha Jabasingh & Valli Nachiyar, 2012a). After which, the enzyme may be transported within the pores of NC-TiO2 or on the exterior surface of bagasse (Lagergren, 1898). Maximum glucose yield was obtained when, 12 ␮g/mL cellulase, was employed for the study. The glucose yield was less at a lower initial concentration of cellulase. As the cellulase concentrations were increased, beyond 12 ␮g/mL, it was observed that, due to the limited surface area available for a specific dosage of NC-TiO2 incorporated into bagasse for the excess cellulase, they remain unutilized and result in a more or less constant glucose yield. Another cause for this constant glucose yield may be due to the accumulation of the reducing sugars which tend to form a resistance to the mass transfer and hence hinder the access of cellulase toward the integrated bagasse-NC-TiO2 structure (NCTiO2 -B). After the imminence of this scenario, the hydrolysis rate was observed to be controlled by the rate of cellulase transport from

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the exterior regions to the interior sites of the integrated bagasse NC-TiO2 structure. For the same NC-TiO2 dosage, the hydrolysis yield could be improvised by enhancing the mass transfer. And this could be exercised by inculcating a stirring action in the SsF. But as far as the present investigations are concerned, cellulase concentration of 12 ␮g/mL is found to be optimum for the current context. 3.2.2. Effect of hydrolysis time on production of glucose The effect of hydrolysis time on the production of glucose using a combination of 6 ␮g NC-TiO2 , 20 mg bagasse and 12 ␮g/mL of cellulase was studied at the SsF conditions (Section 3.1). This study was carried out by varying the time of hydrolysis from 0 min to 120 min (Fig. 4d). The concentration of glucose was found to increase with an increase in the reaction time and attained equilibrium at 50 min for the optimum cellulase concentration employed. This behavior is attributed to the partial enzyme deactivation, product inhibition, chiefly caused by cellobiose and glucose. This might also could have occurred, possibly due to the recrystallization of amorphous cellulose (Segal, Creely, Martin, & Conrad, 1959). As known, the enzyme would be more stable in the presence of the substrate than in its absence. There might be, moreover, some loss in the enzyme activity during extended periods of hydrolysis. Amorphous cellulose is known to be more accessible to enzymatic action than crystalline cellulose. Lee, Shin, and Ryu (1982) and Bertran and Dale (1985) suggested that recrystallization of amorphous cellulose probably occurs during the enzymatic hydrolysis of cellulose. Such recrystallization of amorphous cellulose, if occurs, may join collectively in slowing down the enzymatic hydrolysis rate of cellulose. In the present scenario, as expected, NC-TiO2 integrated with bagasse showed significant improvement in the enzymatic hydrolysis rate as a result of high porosity and ample surface area of the networked, highly integrated and dispersed structure. In addition to the above observations, a significant increase in the initial hydrolysis rate could prove this material as promising boon to bioethanol production as it overcomes the hydrolysis inhibition by glucose and cellobiose. Hence, the optimum conditions for maximum glucose production were found to be an addition of cellulase at a concentration of 12 ␮g/mL to a combination of 6 ␮g NC-TiO2 and 20 mg of bagasse (NC-TiO2 -B) at hydrolysis time of 50 min.

possibly due to the penetration of cellulase into the matrix and the subsequent hydrolysis of the matrix due to infiltration (Fig. 3d) (Anuradha Jabasingh, 2011b). During hydrolysis, the surfaces of these pores are accumulated with cellulase, resulting in the yield of reducing sugar. Cellulase seems to get dilapidated out of the surface of NC-TiO2 -B during this scenario. SEM images clearly picturizes the entire scenario of NC-TiO2 -B hydrolysis. Variation in the surface texture is found to be generalized and very actual, as expected from the hydrolysis time. SEM images provide a qualitative confirmation of these results. Additionally, large localized deposits are found, exhibiting crevices and cracks. Distinction in the surface morphology of these large deposits can be expected due to hydrolysis and shrinking of the matrix (Knappert, Grethlein, & Converse, 1980). 3.4. Kinetic modeling of NC-TiO2 hydrolysis NC-TiO2 hydrolysis involves complex interaction between enzyme, substrate, and the reaction environment, and kinetic modeling is aimed at understanding the complete mechanism of these interactions. Glucose release was observed to decrease significantly as the reaction proceeds (Converse et al., 1988; Howell & Mangat, 1978). This model is based on Langmuir binding kinetics (Zhuoliang & Eric Berson, 2011) and investigates the inactivation of cellulose for the substrates employed in this study. Let us consider the simple rate equation for cellulose hydrolysis, as shown below. k1

E + S ⇔ ES → E + P

(1)

k−1

Eq. (1) describes the initial binding of cellulase and (NC-TiO2 -B) to form an active cellulase and NC-TiO2 -B complex followed by the breakdown of this complex to form product P and thereby releasing the cellulase from NC-TiO2 -B. Product inhibition is neglected in this model, due to the restriction in hydrolysis at the optimum conditions employed (Nidetzky & Steiner, 1993). The basic expression for the hydrolysis rate (␯) is given by

v=

dp K2 [ES] dt

(2)

Since the kinetics are described by the Langmuir model, the concentration of initial enzyme–substrate complex [ES] is expressed by Eq. (3):

3.3. Scanning electron microscopic images Surface morphology of microcrystalline cellulose (MCC), nanocellulose (NC) and TiO2 coupled nanocellulose (NC-TiO2 ) was observed using SEM. Microcrystalline cellulose was made up of more highly ordered crystalline region and less structured amorphous region (Fig. 1a). In the case of the nanocellulose, the degree of departure from crystallinity is invariable to form the less structured purely amorphous region with all degrees of order in between (Fig. 1b). Fig. 1c shows the TiO2 coupled nanocellulose (NC-TiO2 ), a highly amorphous surface created due to the increase in the surface area, possibly owing to the sonication at higher amplitude. These high surface areas were responsible for the exceptional assistance during the bagasse hydrolysis. Fig. 2a–f shows the morphology of NC-TiO2 , when subjected to hydrolysis for 0 min, 5 min, 20 min, 30 min, 50 min and 70 min respectively. At 50 min of hydrolysis, TiO2 coupled nanocellulose was found to be completely hydrolyzed. A comparison between the SEM images that were taken after 50 min and 70 min of hydrolysis showed, a very less significant change in the morphology. This indicated the maximum hydrolysis of NC-TiO2 , resulting in the yield of reducing sugar. Fig. 3a–d shows the surface morphology of bagasse, NC-TiO2 , NC-TiO2 -B before hydrolysis and NC-TiO2 -B subjected to hydrolysis for 50 min respectively. On subjecting NC-TiO2 -B to hydrolysis using the enzyme cellulase, a large number of pores were formed,

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[ES] =

[E]Amax [S] [E] + Kd

(3)

where Kd (␮g/mL) is the equilibrium constant of dissociation, for the binding mechanism considered and is given by k1 /k−1 . Amax is the maximum hydrolysed sites per unit substrate (␮g/mg). The mass balance for enzyme is given by Eq. (4): [E0 ] = [ES] + [E]

(4)

where (E0 ) in ␮g is the total cellulase concentration. On rearranging the above equation, we get an expression for [ES] as follows, Eq. (5): [E] = [E0 ] − [ES]

(5)

Now substituting for [E] in Eq. (3), we get the expression for [ES] as, [ES] =

{[E0 ] − [ES]}Amax [S] ([E0 ] − [ES]) + Kd

(6)

On rearranging and simplifying Eq. (6), we get Eq. (7), which has two roots for [ES]. [ES]2 − {[E0 ] + Kd + Amax + [S]}[ES] + [E0 ][S]Amax = 0

(7)

NC-TiO2 -B concentrations were kept near to the ground in the hydrolysis experiments performed, so that cellulase sorbed onto

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NC-TiO2 -B, [ES] was negligible compared to [E0 ]. Solving Eq. (7), for [ES], and considering the above assumption, we get, [ES] =

[E0 ][S]Amax [E0 ] + Kd + Amax [S]

(8)

Combining Eqs. (2) and (8), the hydrolysis rate for the enzymatic reaction will be:  = K2 [ES] =

K2 [E0 ][S]Amax [E0 ] + Kd + Amax [S]

(9)

Let us define the Michaelis–Mentens constant, Km as follows as in Eq. (10) Km =

Kd + [E0 ] Amax

(10)

Rearranging Eq. (9) and incorporating Km , into the rearranged Eq. (9), we get, the expression for  as follows: =

m [S] Km + [S]

(11)

where m is the maximum rate, defined as m = K2 [E0 ]

(12)

Also, the reaction rate is defined as the differential function of product with respect to time, henceforth, =

m [S] dp = Km + [S] dt

(13)

Since, m and Km are constants, the integration of the differential function of product with respect to time gives Eq. (14) t=

Km 1 [P] .[P] + . m [S] m

(14)

Rearranging the Eq. (14), by considering [S0 ], [S] and [P] as the initial concentration of NC-TiO2 -B, concentration of NC-TiO2 -B at any time t, and the final concentration of glucose respectively, we have, Eq. (15), as follows: Km [P] [P] t= . + m [S0 ] − [P] m

(15)

The Eq. (15) is rearranged as follows, and a plot of t/[P] versus 1/{[S0 ] − [P]} was found to give a straight line with a slope of Km /m and an intercept of 1/m (Fig. 5a) t = [P]

K  m

m

1 1 + m [S0 ] − [P]

(16)

The glucose formation rate,  in Eq. (13) at different time intervals is calculated by varying the initial substrate concentration [S] at different times in the hydrolysis reaction (Kyriacou et al., 1988; Nidetzky et al., 1994). The initial substrate concentration was varied from 1000 ␮g to 5000 ␮g of bagasse, supplemented with 250 ␮g to 1250 ␮g of NC-TiO2 . The first order kinetic plot shows the kinetics of substrate utilization, while maintaining the composition of NC-TiO2 -B substrate the integrated NC-TiO2 -B substrate as 5000 ␮g bagasse and 1250 ␮g of NC-TiO2 (Fig. 5a). The non linear regression of the kinetic plots for all the initial substrate utilization studies resulted in obtaining the values of Km and m . The values of m were found to increase from 1389 ␮g/mL min to 5128 ␮g/mL min and that of the values of Km decreased from 29 ␮g/mL to 10.3 ␮g/mL as the substrate concentration was increased from 1000 ␮g to 5000 ␮g of bagasse, supplemented with 250 ␮g to 1250 ␮g of NC-TiO2 . Our result is consistent with a similar results showing significant decrease in observed catalytic constants (Jalak & Valjamae, 2010; Zhuoliang & Eric Berson, 2011). This proves the efficiency of the biocatalyst integrated with bagasse that favored the affinity of cellulase for the substrate. Similar modeling method was

Fig. 5. Kinetic modeling of NC-TiO2 -B hydrolysis. Plot of t/[P] versus 1/{[S0 ] − [P]} (a), effect of time on the substrate and product concentration for an initial substrate concentration of 5000 ␮g/mL (b). Results are mean of triplicates in (b).

also applied to account for the hydrolysis of sigmacell and steamexploded wheat straw as a substrate, respectively (Zhang et al., 2010; Zhuoliang & Eric Berson, 2011). Fig. 5b shows the effect of time on the substrate and product concentration for an initial substrate concentration of 5000 ␮g/mL. It was observed that the initial substrate concentration of 5000 ␮g/mL was found to be completely depleted in about 50 min, while the glucose concentration reached a maximum of 1620 ␮g/mL for the optimum condition that are employed in the SsF.

Fig. 6. Time course of glucose hydrolysis and ethanol fermentation. Ethanol fermentation: 30 ◦ C; pH 5.0; 2% w/w concentrations of S. cerevisiae ATCC 20602.

S.A. Jabasingh et al. / Carbohydrate Polymers 136 (2016) 700–709

707

Table 1 Fermentation parameters for cellulosic ethanol production by S. cereviseae ATCC 20602 in CelliGen 310 bioreactor. Initial glucose concentration (g/L)

a

Maximum cell concentration (g/L)

*,a

Maximum cellulosic ethanol concentration (g/L)

a

Maximum time (h)

a Ethanol yield coefficient (g ethanol/g glucose)

a Biomass yield coefficient (g biomass/g glucose)

520 750 1250 1933 2400

153.9 226.5 394.8 620.5 1084.8

245.9 253.3 584.8 896.9 770.4

72 66 54 40 39

0.473 0.471 0.467 0.464 0.321

0.296 0.302 0.315 0.321 0.452

* a

Ethanol fermentation: 30 ◦ C; pH 5.0; 2% w/w concentrations of S. cereviseae ATCC 20602. At least three measurements were made for each condition and the data given were averages.

Fig. 7. GC–MS spectra of ethanol produced from glucose obtained by the hydrolysis of bagasse (a), integrated NC-TiO2 -B (b) and GC–MS spectra of standard ethanol (c).

3.5. Fermentation kinetics of ethanol production The initial glucose concentration obtained from integrated NC-TiO2 -B substrate had a pronounced effect on glucose hydrolysis and ethanol fermentation. Lower substrate concentrations were found to increase the yields, this may be because higher substrate concentrations may result in substrate inhibition, which

could result in lower yields, but this had a compromise on the maximum time required for the cellulosic ethanol yield. This effect of substrate inhibition was reported previously for ethanol fermen´ Nikolic, ´ Rakin, & Vukasinovic, ´ 2006). The glucose tation (Mojovic, feed at 1933 g/L from the previous fermenter (Lambda minifor laboratory Bioreactor, 3 L) was started at 3 h of elapsed fermentation time (EFT) (Tomás-Pejó, Negro, Sáez, & Ballesteros, 2012).

708

S.A. Jabasingh et al. / Carbohydrate Polymers 136 (2016) 700–709

Concentrations of glucose, ethanol and dry cell mass were continuously monitored by centrifuging the samples and collecting the supernatant and biomass. Biomass samples were dried at 60 ◦ C for 72 h. The cell concentration reached 1.52 × 106 cells mL−1 in 24 h, and cell viability remained above 92%. Time course of glucose consumption and ethanol production are shown in Fig. 6. Though cells were in an aerated fermentation condition, continuous monitoring of ethanol production denoted the production of a small quantity of ethanol (17 g/L) before the implementation of the anaerobic production phase using nitrogen gas. After 20 h, the aerobic, glucose-limited cultures were exposed to two different conditions that includes, a rapid oxygen depleting phase and a rapid glucose feed depleting phase (Kelbert et al., 2015). These scenarios were controlled and regulated by the redox potential measurement. Oxygen was replaced by nitrogen and sparged into the fermenter. The gas flow rate was maintained as 1.5 L/min. The medium feed was controlled using on-line redox potential measurements as an on-line input. The oxidation-reduction levels were maintained for the ethanol production. It was ensured that cells remain viable and healthy during the anaerobic ethanol production phase (Sassner, Galbe, & Zacchi, 2008). Cell viability was found to be 85% at 84 h of EFT. Throughout the fermentation, the culture broth pH was strictly maintained at 5.0. The percentage yield of ethanol, when the glucose feed was from bagasse alone was found to be 69.56% at 48 h, whereas, the glucose feed from integrated NC-TiO2 -B substrate yielded 89.69% of ethanol at the end of fermenter operation at 40 h. The cell viability in the latter case was found to be high as 92%, while the cell viability for the glucose obtained from integrated NC-TiO2 -B substrate was maintained at 88%. According to the results presented in Table 1, the maximum ethanol concentration, 896.9 g/L, and the maximum product yield on substrate (YP/S ), 0.464 g cellulosic ethanol per g glucose, were achieved for the initial glucose concentration in the mixture (1933 g/L). However, from the economic viewpoint, it is desired to attain as high ethanol concentrations as possible with the intention of decreasing the costs of ethanol distillation, which are considerable in the economical evaluation of the overall process. As presented in Fig. 6, concentration of glucose was constantly maintained low during the fermentation, since the produced glucose was simultaneously consumed by the yeast and converted to ethanol. The time course of the glucose depletion and ethanol production during the fermentation suggested that the consumption of glucose was in accordance with its consumption by yeasts. The progress of ethanol production showed that there was no shortage of fermentable sugars during the process. GC–mass spectrometry (MS) methods have also been applied to measure ethanol concentrations (Condor & Young, 1979). The GC–MS spectra of ethanol produced from glucose obtained by the hydrolysis of bagasse and ethanol produced from glucose obtained by the hydrolysis of integrated NC-TiO2 -B are shown in Fig. 7a and b. The GC–MS spectrum of ethanol standard is shown in Fig. 7c. The retention time for standard ethanol was 1.42 min and that for the ethanol produced from bagasse and bagasse integrated with integrated NC-TiO2 was 1.41 and 1.43 min respectively.

4. Conclusions In this study, NC-TiO2 integrated with bagasse is used as substrate for ethanol production. The present study aims at emphasizing the application of a catalyst to aid in the production of glucose, which in turn can be utilized for the production of cellulosic ethanol, a renewable fuel. NC-TiO2 matrix was developed for integration with bagasse. The production of this matrix was potentially cost-effective and energy efficient. NC-TiO2 -B substrate proved to be a potential feedstock for fuel ethanol production and could provide optional solutions for future cellulosic ethanol

industry. Though the enzymatic hydrolysis of bagasse gives a sustainable release of glucose, the incorporation of TiO2 coupled nanocellulose, (NC-TiO2 ), produced more amount of glucose due to the significant and enhanced surface area created, owing to its catalytic structure. The presented strategy would be further optimized and applied at an industrial capacity which could aid broader application in the global bioethanol market. Acknowledgements We thank Dr. Ing. Berhanu Assefa Demessie of AAiT for his support. The comments and recommendations of the anonymous reviewers and the editor Manuel A. Coimbra are greatly acknowledged. References Abushammala, H., & Hashaikeh, R. (2011). Enzymatic hydrolysis of cellulose and the use of TiO2 nanoparticles to open up the cellulose structure. Biomass and Bioenergy, 35, 3970–3975. Alfani, F., Gallifuoco, A., Saporosi, A., Spera, A., & Cantarella, M. (2000). Comparison of SHF and SSF processes for the bioconversion of steam-exploded wheat straw. Journal of Industrial Microbiology, 25, 184–192. Anuradha Jabasingh, S. (2011a). Response surface methodology for the evaluation and comparison of cellulase production by Aspergillus nidulans SU04 and Aspergillus nidulans MTCC344 cultivated on pretreated sugarcane bagasse. Chemical and Biochemical Engineering Quarterly, 25, 501–511. Anuradha Jabasingh, S. (2011b). Utilization of pretreated coir pith for the optimized bioproduction of cellulase by Aspergillus nidulans. International Biodeterioration and Biodegradation, 65, 1150–1160. Anuradha Jabasingh, S., & Valli Nachiyar, C. (2012a). Optimization of cellulase synthesis by RSM and evaluation of ethanol production from enzymatically hydrolyzed sugarcane bagasse using Saccharomyces cerevisiae. Journal of Scientific and Industrial Research, 71, 353–359. Anuradha Jabasingh, S., & Valli Nachiyar, C. (2012b). Immobilization of Aspergillus nidulans AJSU04 cellulase on modified activated carbon: Sorption and kinetic studies. Journal of Thermal Analysis and Calorimetry, 109, 193–202. Anuradha Jabasingh, S., Varma, S., & Garre, P. (2014). Production and purification of cellulase from Aspergillus nidulans AJSU04 under solid-state fermentation using coir pith. Chemical and Biochemical Engineering Quarterly, 28, 143–151. AOAC. (2000). International Official Methods of Analysis of AOAC International (17th ed.). Gaithersburg, MD, USA: Association of Analytical Communities. Asenjo, J. A. (1983). Maximizing the formation of glucose in the enzymatic-hydrolysis of insoluble cellulose. Biotechnology and Bioengineering, 25, 3185–3190. Baeyens, J., Kang, Q., Appels, L., Dewil, R., Lv, Y., & Tan, T. (2015). Challenges and opportunities in improving the production of bio-ethanol. Progress in Energy and Combustion Science, 47, 60–88. Bertran, M., & Dale, B. (1985). Enzymatic hydrolysis and recrystallization behavior of initially amorphous cellulose. Biotechnology and Bioengineering, 27, 177–181. Binod, P., Kuttiraja, M., Archana, M., Janu, K. U., Sindhu, R., Sukumaran, R. K., et al. (2012). High temperature pretreatment and hydrolysis of cotton stalk for producing sugars for bioethanol production. Fuel, 92, 340–345. Condor, J., & Young, C. (1979). Physicochemical Measurement by Gas Chromatography. Chichester, UK: John Wiley and Sons. Converse, A. O., Matsuno, R., Tanaka, M., & Taniguchi, M. (1988). A model of enzyme adsorption and hydrolysis of microcrystalline cellulose with slow deactivation of the adsorbed enzyme. Biotechnology and Bioengineering, 32, 38–45. Dasari, R. K., & Berson, R. E. (2007). The effect of particle size on hydrolysis reaction rates and rheological properties in cellulosic slurries. Applied Biochemistry and Biotechnology, 137–140, 289–299. Dawson, L., & Boopathy, R. (2007). Use of post-harvest sugarcane residue for ethanol production. Bioresource Technology, 98, 1695–1699. Drissen, R. E. T., Maas, R. H. W., Van Der Maarel, M. J. E. C., Kabel, M. A., Schols, H. A., Tramper, J., et al. (2007). A generic model for glucose production from various cellulose sources by a commercial cellulase complex. Biocatalysis Biotransformation, 25, 419–429. Dunaway, K. W., Dasari, R. K., Bennett, N. G., & Berson, R. E. (2010). Characterization of changes in viscosity and insoluble solids content during enzymatic saccharification of pretreated corn stover slurries. Bioresource Technology, 101, 3575–3582. Fan, L. T., & Lee, Y. H. (1983). Kinetic studies of enzymatic hydrolysis of insoluble cellulose: Derivation of a mechanistic kinetic model. Biotechnology and Bioengineering, 25, 2707–2733. Galbe, M., & Zacchi, G. (2007). Pretreatment of lignocellulosic materials for efficient bioethanol production. Advances in Biochemical Engineeering/Biotechnology, 108, 41–65. Gharpuray, M. M., Lee, Y. H., & Fan, L. T. (1983). Structural modification of lignocellulosics by pretreatments to enhance enzymatic-hydrolysis. Biotechnology and Bioengineering, 25, 157–172.

S.A. Jabasingh et al. / Carbohydrate Polymers 136 (2016) 700–709 Ghose, T. K. (1987). Measurement of cellulase activities. Pure and Applied Chemistry, 59, 257–268. Goering, H. K., & Van Soest, P. J. (1970). Forage Fiber Analysis, USDA Agriculture Handbook No. 379. Washington DC, USA: US Department of Agriculture, Agricultural Research Service, US Government Printing Office. Granda, C. B., Zhu, L., & Holtzapple, M. T. (2007). Sustainable liquid biofuels and their environmental impact. Environmental Progress, 26, 233–250. Gregg, D., & Saddler, J. N. (1996). A techno-economic assessment of the pretreatment and fractionation steps of a biomass-to-ethanol process. Applied Biochemistry and Biotechnology, 57–58, 711–727. Hashaikeh, R., & Abushammala, H. (2011). Acid mediated networked cellulose: Preparation and characterization. Carbohydrate Polymers, 83, 1088–1094. Hong, J., Ye, X. H., & Zhang, Y. H. P. (2007). Quantitative determination of cellulose accessibility to cellulase based on adsorption of a nonhydrolytic fusion protein containing CBM and GFP with its applications. Langmuir, 23, 12535–12540. Howell, J. A., & Mangat, M. (1978). Enzyme deactivation during cellulose hydrolysis. Biotechnology and Bioengineering, 20, 847–863. Isenberg, G. (1999). Assessment of automotive fuels. Journal of Power Sources, 84, 214–217. Jalak, J., & Valjamae, P. (2010). Mechanism of initial rapid rate retardation in cellobiohydrolase catalyzed cellulose hydrolysis. Biotechnology and Bioengineering, 106, 871–883. Jung, H., Yoon, H. G., Park, W., Choi, C., David, B. W., Shin, D. H., et al. (2008). Effect of sodium hydroxide treatment of bacterial cellulose on cellulase activity. Cellulose, 15, 465–471. Kataria, R., & Ghosh, S. (2011). Saccharification of Kans grass using enzyme mixture from Trichoderma reesei for bioethanol production. Bioresource Technology, 102, 9970–9975. Kelbert, M., Romaní, A., Coelho, E., Pereira, F. B., Teixeira, J. A., & Domingues, L. (2015). Lignocellulosic bioethanol production with revalorization of low-cost agroindustrial by-products as nutritional supplements. Industrial Crops and Products, 64, 16–24. Kim, H. M., Wi, S. G., Jung, S., Song, Y., & Hyeun-Jong, B. (2015). Efficient approach for bioethanol production from red seaweed Gelidium amansii. Bioresource Technology, 175, 128–134. Kim, I., Seo, Y. H., Kim, G. Y., & Han, J. I. (2015). Co-production of bioethanol and biodiesel from corn stover pretreated with nitric acid. Fuel, 143, 285–289. Knappert, D., Grethlein, H., & Converse, A. O. (1980). Partial acid hydrolysis of cellulosic materials as a pretreatment for enzymatic hydrolysis. Biotechnology and Bioengineering, 22, 1449–1463. Kumar, P., Barrett, D. M., Delwiche, M. J., & Stroeve, P. (2009). Methods for pretreatment of lignocellulosic biomass for efficient hydrolysis and biofuel production. Industrial and Engineering Chemistry Research, 48, 3713–3729. Kumar, R., & Wyman, C. E. (2008). An improved method to directly estimate cellulase adsorption on biomass solids. Enzyme and Microbial Technology, 42, 426–433. Kwan, N. B., Hwang, T. S., Lee, S. H., Ahn, D. H., & Park, D. H. (2007). Effect of electrochemical redox reaction on growth and metabolism of Saccharomyces cerevisiae as an environmental factor. Journal of Microbiology and Biotechnology, 17, 445–453. Kyriacou, A., Neufeld, R. J., & Mackenzie, C. R. (1988). Effect of physical parameters on the adsorption characteristics of fractionated Trichoderma reesei cellulase components. Enzyme and Microbial Technology, 10, 675–681. Lagergren, S. (1898). Zur Theorie der sogenannten Adsorption gelöster Stoffe Kungliga Svenska Vetenskapsakademiens. Handlingar, 24, 1–39. Lee, S. B., Shin, H. S., & Ryu, D. D. Y. (1982). Adsorption of cellulase on cellulose: Effect of physiochemical properties of cellulose on adsorption and rate of hydrolysis. Biotechnology and Bioengineering, 24, 2137–2153. Lynd, L. R., Weimer, P. J., van Zyl, W. H., & Pretorius, I. S. (2002). Microbial cellulose utilization: Fundamentals and biotechnology. Microbiology and Molecular Biology Reviews, 66, 506–577. McMillan, J. D. (1997). Bioethanol production: Status and prospects. Renewable Energy, 10, 295–302. Miller, G. L. (1959). Use of dinitrosalicylic acid reagent for determination of reducing sugar. Analytical Chemistry, 31, 426–428. ´ L., Nikolic, ´ S., Rakin, M., & Vukasinovic, ´ M. (2006). Production of Mojovic, bioethanol from corn meal hydrolyzates. Fuel, 85, 1750–1755. Nasidi, M., Agu, R., Deeni, Y., & Walker, G. (2015). Improved production of ethanol using bagasse from different sorghum cultivars. Biomass and Bioenergy, 72, 288–299.

709

Nidetzky, B., & Steiner, W. (1993). A new approach for modeling cellulase cellulose adsorption and the kinetics of the enzymatic-hydrolysis of microcrystalline cellulose. Biotechnology and Bioengineering, 42, 469–479. Nidetzky, B., Steiner, W., & Claeyssens, M. (1994). Cellulose hydrolysis by the cellulases from Trichoderma reesei: Adsorptions of two cellobiohydrolases, two endocellulases and their core proteins on filter-paper and their relation to hydrolysis. Biochemical Journal, 303, 817–823. Palmqvist, E., Almeida, J. S., & Hahn-Hagerdal, B. (1999). Influence of furfural on anaerobic glycolytic kinetics of Saccharomyces cerevisiae in batch culture. Biotechnology and Bioengineering, 62, 447–454. Peitersen, N., & Ross, E. W. (1979). Mathematical-model for enzymatic-hydrolysis and fermentation of cellulose by Trichoderma. Biotechnology and Bioengineering, 21, 997–1017. Ruiz, H. A., Silva, D. P., Ruzene, D. S., Lima, L. F., Vicente, A. A., & Teixeira, J. A. (2012). Bioethanol production from hydrothermal pretreated wheat straw by a flocculating Saccharomyces cerevisiae strain – Effect of process conditions. Fuel, 95, 528–536. Samsudin, M. D. M., & Don, M. M. (2015). Assessment of bioethanol yield by S. cerevisiae grown on oil palm residues: Monte Carlo simulation and sensitivity analysis. Bioresource Technology, 175, 417–423. Sassner, P., Galbe, M., & Zacchi, G. (2008). Techno-economic evaluation of bioethanol production from three different lignocellulosic materials. Biomass and Bioenergy, 32, 422–430. Segal, L., Creely, J. J., Martin, A. E., Jr., & Conrad, C. M. (1959). An empirical method for estimating the degree of crystallinity of native cellulose using the X-ray diffraction. Textile Research Journal, 29, 786–794. Shen, J., & Agblevor, F. A. (2008). Kinetics of enzymatic hydrolysis of steam-exploded cotton gin waste. Chemical Engineering Communication, 195, 1107–1121. Singhania, R. R., Saini, J. K., Saini, R., Adsul, M., Mathur, A., Gupta, R., et al. (2014). Bioethanol production from wheat straw via enzymatic route employing Penicillium janthinellum cellulases. Bioresource Technology, 169, 490–495. Tomás-Pejó, E., Negro, M. J., Sáez, F., & Ballesteros, M. (2012). Effect of nutrient addition on preinoculum growth of S. cerevisiae for application in SSF processes. Biomass and Bioenergy, 45, 168–174. Tu, M. B., Chandra, R. P., & Saddler, J. N. (2007). Evaluating the distribution of cellulases and the recycling of free cellulases during the hydrolysis of lignocellulosic substrates. Biotechnology Progress, 23, 398–406. Valjamae, P., Sild, V., Pettersson, G., & Johansson, G. (1998). The initial kinetics of hydrolysis by cellobiohydrolases I and II is consistent with a cellulose surface-erosion model. European Journal of Biochemistry, 253, 469–475. Vallejos, M. E., Felissia, F. E., Kruyeniskia, J., & Area, M. C. (2015). Kinetic study of the extraction of hemicellulosic carbohydrates from sugarcane bagasse by hot water treatment. Industrial Crops and Products, 67, 1–6. Von Blottnitz, H., & Curran, M. A. (2007). A review of assessments conducted on bioethanol as a transportation fuel from a net energy, greenhouse gas, and environmental life cycle perspective. Journal of Cleaner Production, 15, 607– 619. Wu, L., Li, Y., Arakane, M., Ike, M., Wada, M., Terajima, Y., et al. (2011). Efficient conversion of sugarcane stalks into ethanol employing low temperature alkali pretreatment method. Bioresource Technology, 102, 11183–11188. Wyman, C. E. (1999). Biomass ethanol: Technical progress opportunities, and commercial challenges. Annual Review of Energy and the Environment, 24, 189–226. Yang, B., Willies, D. M., & Wyman, C. E. (2006). Changes in the enzymatic hydrolysis rate of avicel cellulose with conversion. Biotechnology and Bioengineering, 94, 1122–1128. Zhang, Y., Xu, J. L., Xu, H. J., Yuan, Z. H., & Guo, Y. (2010). Cellulase deactivation based kinetic modeling of enzymatic hydrolysis of steam-exploded wheat straw. Bioresource Technology, 101, 8261–8266. Zhu, L., O’Dwyer, J. P., Chang, V. S., Granda, C. B., & Holtzapple, M. T. (2010). Multiple linear regression model for predicting biomass digestibility from structural features. Bioresource Technology, 101, 4971–4979. Zhu, S., Huang, W., Huang, W., Wang, K., Chen, Q., & Wu, Y. (2015). Coproduction of xylose, lignosulfonate and ethanol from wheat straw. Bioresource Technology, 185, 234–239. Zhuoliang, Ye., & Eric Berson, R. (2011). Kinetic modeling of cellulose hydrolysis with first order inactivation of adsorbed cellulase. Bioresource Technology, 102, 11194–11199.