Multi-scale structural and chemical analysis of sugarcane bagasse in the process of sequential acid–base pretreatment and ethanol production by Scheffersomyces shehatae and Saccharomyces cerevisiae
© Chandel et al.; licensee BioMed Central Ltd. 2014
Received: 9 September 2013
Accepted: 4 February 2014
Published: 16 April 2014
Heavy usage of gasoline, burgeoning fuel prices, and environmental issues have paved the way for the exploration of cellulosic ethanol. Cellulosic ethanol production technologies are emerging and require continued technological advancements. One of the most challenging issues is the pretreatment of lignocellulosic biomass for the desired sugars yields after enzymatic hydrolysis. We hypothesized that consecutive dilute sulfuric acid-dilute sodium hydroxide pretreatment would overcome the native recalcitrance of sugarcane bagasse (SB) by enhancing cellulase accessibility of the embedded cellulosic microfibrils.
SB hemicellulosic hydrolysate after concentration by vacuum evaporation and detoxification showed 30.89 g/l xylose along with other products (0.32 g/l glucose, 2.31 g/l arabinose, and 1.26 g/l acetic acid). The recovered cellulignin was subsequently delignified by sodium hydroxide mediated pretreatment. The acid–base pretreated material released 48.50 g/l total reducing sugars (0.91 g sugars/g cellulose amount in SB) after enzymatic hydrolysis. Ultra-structural mapping of acid–base pretreated and enzyme hydrolyzed SB by microscopic analysis (scanning electron microcopy (SEM), transmitted light microscopy (TLM), and spectroscopic analysis (X-ray diffraction (XRD), Fourier transform infrared (FTIR) spectroscopy, Fourier transform near-infrared (FT-NIR) spectroscopy, and nuclear magnetic resonance (NMR) spectroscopy) elucidated the molecular changes in hemicellulose, cellulose, and lignin components of bagasse. The detoxified hemicellulosic hydrolysate was fermented by Scheffersomyces shehatae (syn. Candida shehatae UFMG HM 52.2) and resulted in 9.11 g/l ethanol production (yield 0.38 g/g) after 48 hours of fermentation. Enzymatic hydrolysate when fermented by Saccharomyces cerevisiae 174 revealed 8.13 g/l ethanol (yield 0.22 g/g) after 72 hours of fermentation.
Multi-scale structural studies of SB after sequential acid–base pretreatment and enzymatic hydrolysis showed marked changes in hemicellulose and lignin removal at molecular level. The cellulosic material showed high saccharification efficiency after enzymatic hydrolysis. Hemicellulosic and cellulosic hydrolysates revealed moderate ethanol production by S. shehatae and S. cerevisiae under batch fermentation conditions.
KeywordsSugarcane bagasse Sequential acid–base pretreatment Enzymatic hydrolysis Structural analysis Bioethanol Yeasts
Harnessing the carbohydrates from lignocellulosic biomass into bioethanol is not only a ‘nice idea’ but an ‘important necessity’ owing to the increased energy demand globally, safe environment, and sustainable employment . Among the most feasible second generation feedstock for bioethanol production, sugarcane bagasse (SB), a fibrous residue generated during the extraction of cane juice in mills, is an excellent raw commodity due to large abundance, non-competitiveness with food or feed requirement, easy transportation, and rich in accessible carbohydrates . The fullest exploitation of SB in the integrated biorefineries setting (1G + 2G ethanol production in one unit) may provide a unique breakthrough for the commercialization of biofuels .
The key driver for the successful conversion of SB into ethanol is the selection of efficient pretreatment technology followed by maximal sugars recovery coupled with ethanol production with desired yield and productivities [2, 4, 5]. Pretreatment of lignocellulose is an inevitable process for the commercial deployment of cellulosic biofuels. Nevertheless, there are only a few robust pretreatment technologies available which efficiently enable the cellulosic fraction of the cell wall for enzymatic conversion into monomeric sugars [5, 6]. Sequential acid–base pretreatment has shown efficient removal of hemicellulose and lignin from SB eventually releasing high amounts of fermentable sugars upon enzymatic hydrolysis [7–9]. Apart from acid–base pretreatments, several pretreatment methods have been studied in the past which selectively remove either hemicellulose or lignin from the SB matrix. The choice of effective pretreatment methods depends upon the minimum degradation of carbohydrate, ability to enhance the surface area of cellulosic substrates, and minimizing the production of inhibitors and toxic products [1, 5].
SB is consisted of crystalline cellulose nanofibrils embedded in an amorphous matrix of cross-linked lignin and hemicelluloses that impair enzyme and microbial accessibility . Structural changes in the cellular components of SB elucidating the hemicellulose degradation and removal of lignin after sequential acid–base pretreatment are very important for the investigation of changes in the cell wall at molecular level . Structural mapping of lignocellulosic materials after various thermochemical pretreatments provide important insights at molecular level to facilitate the mechanistic action of catalysts used for pretreatment [10–13]. Microbial conversion of hemicellulose and cellulose derived sugars into ethanol with optimum yields and productivities are the key parameters to obtain cheap ethanol in biorefineries . For the conversion of xylose, Scheffersomyces shehatae (syn. Candida shehatae) is the preferred choice [14, 15]. On the other hand, Saccharomyces cerevisiae is the most employed microorganism for ethanol production from cellulosic derived sugars with high productivities .
In the present study, we attempted to pretreat the SB by sequential acid–base followed by enzymatic saccharification using commercial enzymes. The hemicellulosic hydrolysate after vacuum concentration and detoxification (30.89 g/l xylose) and cellulosic hydrolysate (48.50 g/l glucose) were fermented by S. shehatae UFMG HM 52.2 and S. cerevisiae 174 to ethanol under batch cultivation conditions. Further, efforts were made to unveil the structure of native, sulfuric acid pretreated, sodium hydroxide delignified, and enzyme-digested SB by using scanning electron microscopy (SEM), transmitted light microscopy (TLM), Fourier transform infrared (FTIR) spectroscopy, Fourier transform near-infrared (FT-NIR) spectroscopy, micro-Raman spectroscopy, and solid-state nuclear magnetic resonance (NMR) spectroscopy.
Results and discussion
Native SB used in this study was found to contain cellulose (39.52%), hemicellulose (25.63%), total lignin (30.36%), ash (1.44%), and extractives (2.90%). These values are within the range found in other studies. The composition of SB varies with variety, origin, cultivation type of sugarcane, and the analytical method used for the characterization . For instance, Rocha et al.  observed 45.5% cellulose, 27% hemicellulose, 21.1% lignin, and 2.2% ash in SB. Rabelo et al.  observed 38.4% cellulose, 23.2% hemicellulose, 25% lignin, and 1.5% ash in SB.
Dilute sulfuric acid treatment
Chemical composition of sugarcane bagasse (SB) native hemicellulose hydrolysate, vacuum concentrated, and detoxified (overliming plus activated charcoal)
Concentrated hydrolysate (3x)
6 × 10-3
5.93 × 10-4
2.85 × 10-3
1 × 10-3
After the dilute acid hydrolysis, 92.78% hemicellulose was removed, which in turn increased the amount of lignin (45.09%), cellulose (48.95%), and ash (3.2%) in the substrate, showing the efficiency of this pretreatment. Extractives were found in negligible amounts. Canilha et al.  reported 3.7% hemicellulose in the substrate after the dilute sulfuric acid pretreatment (2.5% w/v H2SO4, 150°C, 30 minutes) from the dilute sulfuric acid pretreated SB. Rezende et al.  recovered 51.2% cellulose, 29.5% lignin, and 7.8% hemicellulose in the dilute sulfuric acid pretreated SB (1% H2SO4, 120°C, 40 minutes).
Vacuum evaporation and detoxification of dilute acid hydrolysate
In order to increase the concentration of sugars in the hydrolysate, native hemicellulosic hydrolysate was concentrated by vacuum evaporation at 70°C. The hydrolysate after concentration was found to contain an efficient amount of sugars (37.44 g/l xylose, 2.73 g/l arabinose, and 0.39 g/l glucose) and other undesired products (Table 1). It is essential to eliminate these inhibitors from the hydrolysate before fermentation to obtain the desired ethanol yield and productivities . Sequential detoxification strategy (CaO mediated overliming followed by activated charcoal treatment) has shown an effective removal of inhibitors from the hydrolysates without the loss of sugars (reduction of approximately 96% of furfural and 90% of HMF comparing crude with treated hydrolysate). However, the removal of acetic acid was negligible (Table 1). Activated charcoal treatment effectively eliminates the furfurals and HMF from dilute acid hydrolysate of SB under the process conditions employed in this study. These results are in good accordance with the previous study on detoxification of SB acid hydrolysate by overliming and charcoal .
Sodium hydroxide treatment
The recovered cellulignin after dilute acid hydrolysis was delignified by sodium hydroxide pretreatment (1.0% w/v NaOH, 120°C, 60 minutes). These conditions showed the maximal removal of lignin (55.65%) from the substrate after a response surface optimization (L9 Taguchi orthogonal design of experiments) for delignification . It is necessary to remove the lignin from the substrate to obtain the desired sugars recovery from cellulignin after enzymatic saccharification. Lignin significantly aids the biomass recalcitrance eventually strongly impairing the cellulase amenability towards the substrate.
Sodium hydroxide mediated pretreatment disrupts the SB cell wall by solubilization of the remaining hemicellulose and lignin. Sodium hydroxide mechanistically cleavages the alpha-aryl ester bonds from its polyphenolic monomers along with weakening of hydrogen bonds, which in turn promotes the swelling of cellulose . Sequential acid–base pretreatment of SB has been found effective for the removal of hemicellulose and lignin from SB increasing the amenability of cellulases towards cellulose [7, 8]. NaOH mediated delignification of cellulignin (1% w/v NaOH, 120°C, 1 hour of pretreatment time) showed 76.5% cellulose, 20.0% lignin, and 3.50% ash in the substrate. Rezende et al.  reported 88% lignin removal which showed 84.7% cellulose and 3.3% hemicellulose in acid-alkali pretreated SB (2% w/v NaOH, 120°C, 40 minutes) . Recently, Rocha et al.  observed 91.2% lignin removal from cellulignin of SB under the alkali pretreatment conditions (1% w/v NaOH, 100°C, 1 hour).
The acid–base pretreated SB, the co-called SB-cellulose, was enzymatically saccharified to obtain clean sugar stream (glucose solution). A maximum of 48.50 g/l glucose (0.91 g/g SB-cellulose) was obtained from the acid–base pretreated SB after 96 hours of enzymatic hydrolysis (15 FPU/g, 20 CBU/g of enzyme loading). Acid pretreated bagasse (cellulignin) showed only 22.75 g/l sugars recovery proving the requirement of alkali mediated delignification.
Apart from the efficient pretreatment of the substrate, appropriate enzyme loading and hydrolytic conditions are necessary to obtain the maximum sugars recovery from the substrate [7, 22]. Sequential acid–base pretreatment efficiently removes the hemicellulose and lignin from the substrate simultaneously promoting the swelling of cellulose. Further, Tween 20 was also added in the hydrolytic reaction as surfactant for the enhancement of specific surface area for better enzyme action. Surfactants generally enhance the surface area of lignocellulosic substrates to improve the extent of enzymatic hydrolysis. Non-ionic surfactant-like Tween 20 is more effective due to its adsorption on hydrophobic surfaces mainly composed of lignin fragments .
It is difficult to compare the hydrolytic efficiency of acid–base pretreated SB with the existing reports due to the different conditions employed in the studies. For example, Giese et al.  performed sequential acid–base pretreatment of SB (100 mg H2SO4/g of SB, 121°C, 20 minutes) plus alkali (1% w/v NaOH, 100°C, 1 hour). The acid–base pretreated SB was further enzymatically hydrolyzed (cellulase AF28918 300 IU/ml, cellobiase 80 IU/ml, Tween 80 0.15 g/g, and pretreated SB 5% (w/v)), which led to sugars recovery (0.89 g total reducing sugars (TRS)/g). Rezende et al.  reported 72% cellulose conversion from consecutive acid (1% v/v H2SO4, 120°C, 40 minutes) and base (1% w/v NaOH, 120°C, 40 minutes) pretreated SB after enzymatic hydrolysis (25 FPU of Accelerase 1500 (DuPont, Wilmington, DE, USA) and 50 IU of β-glucanase from Novozym 188 (Sigma Aldrich, USA).
Scanning electron microscopy (SEM)
Transmitted light microscopy (TLM)
X-ray diffraction (XRD)
Earlier, Rezende et al.  observed the CrI of raw bagasse (48.7 ± 2.5%), corresponding to a cellulose amount of 35.2%. A liner increment in sample crystallinity was observed with the cellulose amount as the sample was treated with 1% H2SO4, 0.25%, or 0.5% NaOH, corresponding to cellulose percentages of 51%, 66%, and 68%, respectively. Sindhu et al.  reported the increased CrI (67.83%) of dilute sulfuric acid plus formic acid pretreated SB samples. Velmurugan and Muthukumar  also observed the CrI (66%) of sodium hydroxide pretreated bagasse which was further increased (up to 70.7%) after sono-assisted pretreatment as compared to native SB (50%). The CrI of enzyme digested samples (66.98%) was not found to be increased. Furthermore, additional peaks were more visible in the spectra of enzyme digested bagasse due to the presence of SiO2 in enzyme hydrolyzed bagasse samples. This is an intriguing finding, probably due to the high beta-glucosidase concentration in the commercial enzymatic preparations and complexity of the SB cellulose chemical composition and morphology. Earlier, Binod et al.  reported the CrI of native SB (53.44%), microwave-alkali pretreatment (65.29%), and enzyme hydrolyzed microwave-alkali pretreated bagasse (58.58%).
Fourier transform infrared (FTIR) spectroscopy
The region between the wavelengths of 1,450 to 1,300 cm-1 was omitted. According to literature, this region exhibits a high molecular coupling, which makes the area quite complex, involving a superposition of several modes of vibration of the lignin and carbohydrates . A band around 1,458 cm-1 is reported to be a deformation of lignin CH2 and CH3, and 1,604 cm-1 is reported to be stretching of the C = C and C = O lignin aromatic ring. The band around 1,515 cm-1 is because of the C = C stretching of the aromatic ring in lignin [33, 34]. A band around 1,733 cm-1 is characteristic of C = O stretching of unconjugated hemicellulose. The peak around 2,850 cm-1 is reported due to the symmetric stretch of CH and CH2, while the peak at 2,918 cm-1 is due to asymmetrical stretching of CH2 and CH. Both denote the characteristics of cellulose . The region between 3,800 and 3,000 cm-1 covers the related crystalline structure of cellulose. This region represents the sum of the vibration of valence bands of the hydrogen bond of the OH group and the bands of intra-molecular and intermolecular hydrogen bonds .
Figure 4b shows the FTIR spectrum of the region 1,300 to 750 cm-1 of native SB, sulfuric acid pretreated, NaOH-pretreated cellulignin, and enzymatic digested SB. The region between 1,100 and 1,000 cm-1 clearly shows two peaks after acid hydrolysis (red line), indicating the removal of hemicellulose. The removal of hemicellulose is also evident in the region of 1,247 cm1. Alkaline hydrolysis (blue line) seems to be affected due to the removal of lignin moieties. After the enzymatic hydrolysis (green line), the peaks between 1,200 and 1,000 cm-1 are accentuated, which demonstrates the hydrolysis of cellulose.
Figure 4c shows the FTIR spectrum of the region 1,800 to 1,400 cm-1 of native SB, cellulignin, alkaline hydrolyzed cellulignin, and enzymatic digested material. The region of 1,733 cm-1 is affected after acid hydrolysis (red line), which indicates the decrease in hemicellulose content. The regions 1,604 cm-1, 1,515 cm-1, and 1,458 cm-1 relating to the lignin macromolecule have noticeable changes after alkaline hydrolysis (blue line). These changes are good evidence of effective lignin degradation after the alkali mediated delignification process.
Figure 4d shows the FTIR spectrum of the region 3,900 to 2,700 cm-1 of native SB, acid hydrolyzed bagasse, alkaline hydrolyzed cellulignin, and enzyme hydrolyzed substrate. Bands around 2,918 cm-1 and 2,850 cm-1 do not seem to change during acid hydrolysis (red line) which is related to the cellulose. This indicates that the major effect of treatment with dilute acid is the removal of hemicellulose. However, this region is affected during alkaline hydrolysis (blue line) and enzymatic hydrolysis (green line). The changes in the two local maxima of 2,918 cm-1 and 2,850 cm-1 indicate that chemical treatment has also affected the cellulose chain. The increase in line width and asymmetry of the curves in the range of 3,800 to 3,000 cm-1 in the course of the treatments indicates disturbances in the crystalline structure of cellulose. These changes are strong evidence of intra-molecular hydrogen bonding disruption in cellulose .
Fourier transform near-infrared (FT-NIR) spectroscopy
Regarding structural changes in hemicellulose after sulfuric acid pretreatment, the region around 5,808 cm-1 is the first of the CH stretching band harmonic (overtone CH stretch) attributed to variations of hemicellulose [40, 41]. The minimum local amplitude variation in this region indicates the removal of hemicellulose content . The range of the 6,000 to 5,920 cm-1 band is attributed to the first harmonic of the stretching vibration of aromatic CH (CH stretching vibration of aromatics) and is responsible for variation of lignin content. The change in local minimum amplitude in this region indicates degradation in units of lignin macromolecules. Figure 5b,c shows the second derivative of the absorption spectra for the FT-NIR range from 5,500 to 4,000 cm-1 of native, dilute sulfuric acid pretreated, sodium hydroxide pretreated, and enzyme digested SB. The regions around 4,813, 4,285, and 5,208 cm-1 are combinations of the first harmonic of the CH stretching, and the region around 4,405 cm-1 is the combination of the first harmonic of CO stretching in polysaccharides [40, 41]. According to Figure 5b,c, it is possible to observe changes in the content of polysaccharides. The steps in which the samples are subjected to hydrolysis result in a relative increase in local minimum amplitude, indicating the increase of cellulose in acid pretreated and delignified samples.
Solid-state 13C nuclear magnetic resonance (NMR) spectroscopy
Kinetic parameters for ethanol production from detoxified sulfuric acid hydrolysate after detoxification and enzymatic hydrolysates by Scheffersomyces shehatae UFMG HM 52.2 and Saccharomyces cerevisiae 174, respectively
S. shehatae a
S. cerevisiae b
Initial sugars (gs/l)
Sugar consumed (%)
Ethanol produced (gp/l)
Ethanol yield (gp/gs)
Ethanol productivity (gp/l/h)
Biomass produced (gx/l)
Biomass yield (gx/gs)
Biomass productivity (gx/l/h)
The spectrum of the sulfuric acid treated sample is shown in Figure 7b. The intensity decrease of signals 1 and 17 as well as the improved spectral resolution in the 50 to 120 ppm region indicates the almost complete removal of hemicellulose in this sample. The spectrum of the samples pretreated by sodium hydroxide is shown in Figure 7c. The intensity decrease of signals *, 2, 11, 12, 13, 14, and 15 show the reduction of the lignin to cellulose fraction in the bagasse sample after the sodium hydroxide treatment. Moreover, in agreement with the Raman results, the remaining lignin signals in the spectra of Figure 7c show that lignin is not completely removed by these treatments.
Figure 7d shows the spectrum of the solid fraction obtained after enzyme hydrolyze of the sodium hydroxide and sulfuric acid treated sample. Assuming that the relative amount of lignin is maintained during the enzyme hydrolysis, the increase of lignin signals, lines *, 2, 11, 12, 13, 14, and 15, relative to cellulose ones, lines 3, 4, 5, 6, 7, 8, and 10, is associated to the removal of cellulose. Information that can be obtained from Figure 7 is the increase in the crystallinity of the cellulose after the enzymatic treatment. As already mentioned, signals at 84 and 88 ppm are due to the amorphous and crystalline cellulose, respectively. Spectra of Figure 7c,d show the change in the relative intensity of signal 7 and 8 as well as 3 and 4, which might suggests an increase in crystallinity of cellulose upon enzymatic hydrolysis, showing the preference for removal of amorphous cellulose. However, it is worth pointing out that there is indeed a lignin peak in the region of the C4 signal which may compromise the calculation of the ratio between the crystalline and amorphous signal in the spectrum of the enzymatic treated sample. Zhao et al.  observed that the ordered structure of crystalline cellulose was not found to be disrupted after the hydrolysis of cellulose. They found that the peak ratio of C4 (79 to 86 ppm)/C4 (86 to 92 ppm) to calculate the cellulose crystallinity remains the same after the hydrolysis reaction. The relative ratio of amorphous cellulose and crystalline cellulose was almost similar prior to hydrolysis.
Fermentation of acid hydrolysate byScheffersomyces shehatae
Fermentation of enzyme hydrolysate bySaccharomyces cerevisiae
Plant cell walls are a useful source of renewable energy. For the biochemical ethanol production from lignocellulosics, it is essential to overcome the complex, rigid, and recalcitrant characteristics of the plant cell wall. Chemical pretreatment encompassing sequential acid–base pretreatment of SB separates hemicellulose and lignin, and increases the accessibility of cellulose to cellulase enzyme mediated action to convert into glucose. The microscopic (SEM and TLM) and spectroscopic techniques (FTIR, FT-NIR, Raman, NMR, and XRD) used in this work provided in-depth structural investigation of chemical changes at the molecular level during sequential pretreatment and enzymatic digestion.
Dilute sulfuric acid pretreatment significantly removed hemicellulose (10.9 g/l xylose) in addition to lignin relocalization. Cellulignin was further delignified by dilute sodium hydroxide pretreatment, which efficiently removed lignin from the substrate eventually increasing the cellulose fraction in the substrate. Acid–base pretreated substrate showed efficient enzymatic action toward the depolymerization of cellulose into glucose (0.91 g sugars/g pretreated bagasse). Detoxified hemicellulosic hydrolysate, when fermented by S. shehatae UFMG HM 52.2, showed ethanol production of 9.11 g/l (yield 0.38 g/g). The cellulosic hydrolysate showed ethanol production of 8.13 g/l (yield 0.22 g/g) by S. cerevisiae 174. Both microorganisms showed a moderate ethanol yield which needs further investigation. Both microorganisms are native and showed average ethanol production potential from SB hydrolysates. System metabolic engineering-based approaches, improvements in media formulation, and modified fermentation methods could provide the desired ethanol yields from these microorganisms growing on lignocellulose hydrolysates.
Material and methods
Preparation of raw substrate
The raw substrate, SB, was acquired from Usina Vale do Rosário (Morro Agudo, São Paulo, Brazil). During preliminary processing, the SB was air-dried and knife-milled (model number MA 680; Marconi Equipamentos, Piracicaba, São Paulo, Brazil) to pass through with a 20-mesh sieve. The finely milled SB was washed under running tap water to remove the dust and dried at 45°C for further experiments.
Dilute acid hydrolysis
The dilute acid hydrolysis of SB was carried out in a Parr reactor 4848 (Moline, IL, USA) with a capacity of 5 l. For the hydrolysis of the SB, H2SO4 (98% purity) was used as a catalyst in a ratio of 100 mg of acid/g of SB, at 121°C for 20 minutes, using a ratio of 1/10 between the bagasse mass and the volume of acid solution. After the reaction, the solid material (cellulignin) was recovered by filtration using muslin cloth. The hydrolysate obtained was maintained at 4°C. Cellulignin was washed with running tap water until neutral pH and dried at 45°C.
Detoxification of SB hemicellulosic hydrolysate
The SB hemicellulosic hydrolysate was vacuum concentrated at 70°C in a concentrator and further detoxified by sequential calcium oxide-activated charcoal pretreatment according to Alves et al. . The hydrolysate was finally filtered under vacuum and then autoclaved under 0.5 atm (110°C) for 15 minutes.
Dilute sodium hydroxide pretreatment
Cellulignin obtained after dilute acid hydrolysis was subsequently pretreated by sodium hydroxide mediated delignification. It was carried out in a Parr reactor 4848 of 5 l capacity. Sodium hydroxide (1% m/v) was used as a catalyst in a ratio of 1/10 between the cellulignin mass and the volume of alkali solution, at 121°C for 1 hour. After the reaction, the solid material (cellulose) was recovered by filtration using muslin cloth. The recovered solid residue was washed with running tap water until neutral pH and dried at 45°C.
Enzymatic hydrolysis of acid–base pretreated bagasse was performed in a 250 ml Erlenmeyer flask containing 7.5 g d.wt. of acid–base pretreated bagasse and 100 ml of citrate buffer (50 mM, pH 4.8). Substrates with buffer were pre-incubated at room temperature for 90 minutes. The substrate soaked in citrate buffer was supplemented with cellulase loadings (15 FPU/g of the dry substrate from Celluclast 1.5 L and 20 IU/g of β-glucosidase from Novozym 188). Surfactant (Tween 20) was also added (0.10 g/g substrate) in the hydrolysis experiment. Enzymatic hydrolysis was performed at 50°C at 150 rpm in an incubator shaker (Innova 4000; New Brunswick Scientific, Enfield, CT, USA). The enzymes were purchased from Sigma Aldrich (St Louis, MO, USA). The enzymatic hydrolysis was performed for a period of time up to 96 hours. Samples were collected after every 24 hours, centrifuged, and analyzed to determine the sugars released.
The chemical composition of the solid material (raw bagasse, cellulignin, and cellulosic pulp (1% m/v NaOH, 120°C, 90 minutes)) was analyzed by a methodology validated by Gouveia et al.. The determination of constituent concentrations in dilute acid hydrolysate and enzymatic hydrolysates were verified by HPLC. The content of glucose, xylose, arabinose, formic acid, and acetic acid were verified in chromatograph Shimadzu LC-10AD (Kyoto, Japan) with a column equipped with Aminex HPX-87H (300 × 7.8 mm; Bio-Rad, Hercules, CA, USA), coupled to a refractive index detector (RID-6A), and 0.01 N sulfuric acid as an eluent at a flow rate of 0.6 ml/min, column temperature of 45°C, and injected volume of 20 μl. The samples were previously filtered through a Sep-Pak C18 filter (Sigma Aldrich, USA). The determination of furfural and HMF was obtained in chromatograph Shimadzu LC 10AD with column HP-RP18 (200 × 4.6 mm), coupled to an ultraviolet detector SPD-10A UV–VIS in a wavelength of 276 nm, with eluent acetonitrile/water (1/8), and 1% of acetic acid. The used flow was 0.8 ml/min, the column temperature was 25°C, and the volume injected was 20 μl. All the samples were filtered in membrane Minisart 0.22 μm (Sartorius, Epsom, UK) before the readings. TRS in enzyme hydrolysates were estimated by using spectrophotometer (Beckman DU-640B; Beckman Coulter, Brea, CA, USA) following the dinitrosalicylic acid (DNS) method of Miller . All the experiments were carried out in triplicates. The values are the mean of three replicates.
S. shehatae UFMG HM 52.2 was isolated from a sample of rotting wood grown in xylan medium (1% xylan, 0.67% yeast nitrogen base, 0.02% chloramphenicol, and pH 5.0 ± 0.2), collected in a private natural reserve, Bello and Kerida, located in Rio Grande de Cima, Nova Friburgo, Rio de Janeiro, Brazil. The strains were maintained on yeast extract peptone dextrose (YPD) plates and stored at 4°C. S. shehatae UFMG HM 52.2 was grown in 150 ml Erlenmeyer flasks containing 50 ml of seed medium (30 g/l of xylose), 20 g/l of peptone, and 10 g/l of yeast extract in an orbital incubator shaker at 30°C, 200 rpm. Synthetic medium consisted of commercial xylose (37 g/l), and the other medium ingredients were the same as the hydrolysate supplemented medium. For the fermentation of S. shehatae UFMG HM 52.2, the medium was composed of the hydrolysates supplemented with yeast extract (3.0 g/l), malt extract (3.0 g/l), and ammonium sulfate (5.0 g/l), as described by Parekh et al. . Flasks were maintained in a rotator shaker at 30°C and 200 rpm for 72 hours. Samples were collected at 0 hours, 6 hours, 14 hours, 24 hours, and 48 hours to determine the residual sugars and ethanol and biomass production.
S. cerevisiae 174 was isolated from the Atibaia River, São Paulo state, Brazil. It was grown in 150 ml Erlenmeyer flasks containing 50 ml of seed medium (30 g/l of glucose, 5 g/l of peptone, 3 g/l of yeast extract, and 0.25 g/l diammonium hydrogen phosphate) in an orbital incubator shaker at 30°C, shaken at 100 rpm. Following 24 hours of growth, fermented broth was centrifuged and S. cerevisiae was prepared corresponding to 1.0 g/l cells (d.wt.). Inoculums were aseptically transferred into enzymatic hydrolysates (50 ml) supplemented with medium ingredients.
Scanning electron microscope (SEM)
The SEM analysis of native, dilute sulfuric acid pretreated, dilute sodium hydroxide pretreated, and enzymatically hydrolyzed SB was performed as described by Kristensen et al. . Briefly, native, acid–base pretreated, and enzymatically hydrolyzed SB were distributed on a 12 mm glass coverslip coated with poly-L-lysine (Sigma Diagnostics, São Paulo, Brazil). The dried sections were mounted on aluminum stubs, sputter-coated (JEOL JFC-1600) with a gold layer, and used for scanning. The prepared samples were scanned and imaged using Hitachi S520 SEM (Tokyo, Japan).
Transmitted light microscopy (TLM)
TLM analysis of native, dilute sulfuric acid pretreated, dilute sodium hydroxide pretreated, and enzyme digested SB samples was performed to reveal the changes in surface morphology in response to transmitted light. After the light passes through the samples, the image of the specimen goes through the objective lens and to the oculars where the enlarged image is viewed. The samples were mounted on aluminum stubs and the light was passed through a condenser to focus it on the samples to obtain very high illumination with the microscope (Axioskop 40; Zeiss, Oberkochen, Germany) and camera (Axiocam ICC 3; Zeiss). All the images were captured through 25, 50, and 400× magnifications.
X-ray diffraction (XRD)
Fourier transform infrared (FTIR) spectroscopy
FTIR spectroscopic analysis of native, dilute sulfuric acid pretreated, dilute sodium hydroxide pretreated, and enzyme digested SB samples was performed to detect the changes in functional groups. Samples were milled in agate cups for 1.5 hours with 400 rpm (PM 400; RETSCHW, Haan, Germany), followed by passing through a 100-mesh sieve (A Bronzinox, São Paulo, Brazil) with an aperture of 150 μm. The pellets were prepared by mixing 300 mg of spectroscopic grade KBr with 3 mg of sample in an agate mortar. Each sample was then submitted to 10 tons for 3 minutes in a hydraulic press (Atlas 25T; Specac, Swedesboro, NJ, USA). The spectra were collected in the range 4,000 to 400 cm-1 with a resolution of 4 cm-1 and 72 scans per sample on a VERTEX 70 spectrometer (Bruker Optics, Ettlingen, Germany).
Fourier transform near-infrared (FT-NIR) spectroscopy
The FT-NIR spectroscopy of native, dilute sulfuric acid pretreated, dilute sodium hydroxide pretreated, and enzyme digested SB samples was performed with a spectrometer FT-NIR multi-purpose analyzer (MPA) from Bruker Optics. The measurements were used to diffuse reflectance, which was analyzed via an integrating macro sample sphere, the diameter of the measured area was 15 mm, and 32 scans per sample were performed with a resolution of 4 cm–1 covering a range from 13,000 to 3,500 cm–1. Second derivative spectra were calculated with 21 smoothing points after unit vector normalization. All calculations were conducted with OPUS version 6.5 software.
The Raman spectra of native and sequential acid–base pretreated SB samples (dilute sulfuric acid pretreatment of bagasse followed by dilute sodium hydroxide pretreatment) were carried out using a Bruker Optics RFS 100 instrument and Nd:YAG laser operating at 1,064 nm. The instrument with 4 cm-1 of spectral resolution was equipped with a germanium detector cooled with liquid nitrogen and coupled to a RamanScopeIII microscope system. Good signal-to-noise ratios were obtained with 600 scans, using a range of laser power between 50 and 200 mW.
Solid-state 13C nuclear magnetic resonance (NMR) spectroscopy
Solid-state 13C NMR spectroscopy of native, dilute sulfuric acid pretreated, dilute sodium hydroxide pretreated, and enzyme digested SB samples was performed using a Varian Inova spectrometer (Eugene, OR, USA) at 13C and 1H frequencies of 88.02 and 350.50 MHz, respectively . A Varian 7 mm magic angle spinning (MAS) double resonance probe head was used. Spinning frequencies of 4.5 kHz were controlled by a Varian pneumatic system that ensures a rotation stability of approximately 2 Hz. Radio frequency ramped cross-polarizations under magic angle spinning (CPMAS) combined with total suppression of spinning sidebands (TOSS) and heteronuclear 1H decoupling (CPMAS-TOSS) was used to acquire the 13C spectra. Typical π/2 pulse lengths of 4.0 μs (13C) and 4.5 μs (1H), cross-polarization time of 1 ms, acquisition time of 30 ms, and recycle delays of 2 seconds were used in all NMR experiments.
Cross polarization under magic angle spinning and total suppression of spinning sidebands
Filter paper unit
Fourier transform infrared
Fourier transform near-infrared
High performance liquid chromatography
Magic angle spinning
YAG: Neodymium-doped yttrium aluminum garnet
Nuclear magnetic resonance
Refractive index detector
Scanning electron microscopy
Transmitted light microscopy
Total suppression of spinning sidebands
Total reducing sugars
Yeast extract peptone dextrose.
The authors are grateful to Fundação de Amparo à Pesquisa do Estado de São Paulo (FAPESP) Research Program on Bioenergy (BIOEN; process numbers: 2008/57926-4 and 2010/11258-0) and Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq) for financial support. FAFA gratefully acknowledges Coordenação de Aperfeiçoamento de Pessoal de Nível Superior (CAPES). VA, MJVB, LNR, and CAR acknowledge Fundação de Amparo à Pesquisa de Minas Gerais (FAPEMIG), CNPq, and CAPES for financial support. ODB and ERA are grateful to FAPESP and CNPq (process number: 159341/2011-6) for financial support. The authors are thankful to Ms Juliana RG Reis for technical assistance. The authors would also like to thank Dr Durval Rodrigues Jr, Engineering School of Lorena (EEL), São Paulo, Brazil, and Dr Rogerio Rhein, Universidade Estadual Paulista (UNESP), Guaratingueta, São Paulo, Brazil, for SEM and TLM analysis, respectively.
- Dale BE, Ong RG: Energy, wealth, and human development: why and how biomass pretreatment research must improve. Biotechnol Prog. 2012, 28: 893-898. 10.1002/btpr.1575.View ArticleGoogle Scholar
- Chandel AK, da Silva SS, Carvalho W, Singh OV: Sugarcane bagasse and leaves: foreseeable biomass of biofuel and bioproducts. J Chem Technol Biotechnol. 2012, 87: 11-20. 10.1002/jctb.2742.View ArticleGoogle Scholar
- Dias MOS, Junqueira TL, Cavalett O, Cunha MP, Jesus CDF, Rossell CEV, Filho RM, Bonomi A: Integrated versus stand-alone second generation ethanol production from sugarcane bagasse and trash. Bioresour Technol. 2012, 103: 152-161. 10.1016/j.biortech.2011.09.120.View ArticleGoogle Scholar
- Himmel ME, Ding SY, Johnson DK, Adney WS, Nimlos MR, Brady JW, Foust TD: Biomass recalcitrance: engineering plants and enzymes for biofuels production. Science. 2007, 315: 804-807. 10.1126/science.1137016.View ArticleGoogle Scholar
- Canilha L, Chandel AK, Milessi TSS, Antunes FAF, Freitas WLC, Felipe MGA, da Silva SS: Bioconversion of sugarcane biomass into ethanol: An overview about composition, pretreatment methods, detoxification of hydrolysates, enzymatic saccharification and ethanol fermentation. J Biomed Biotechnol. 2012, doi: 10.1155/2012/989572Google Scholar
- Agbor VB, Cicek N, Sparling R, Berlin A, Levin DB: Biomass pretreatment: fundamentals toward application. Biotechnol Adv. 2011, 29: 675-685. 10.1016/j.biotechadv.2011.05.005.View ArticleGoogle Scholar
- Rezende CA, de Lima MA, Maziero P, de Azevedo ER, Garcia W, Polikarpov I: Chemical and morphological characterization of sugarcane bagasse submitted to a delignification process for enhanced enzymatic digestibility. Biotechnol Biofuels. 2011, 11: 4-54.Google Scholar
- Giese EC, Pierozzi M, Dussán KJ, Chandel AK, da Silva SS: Enzymatic saccharification of acid-alkali pretreated sugarcane bagasse using commercial enzymatic preparations. J Chem Technol Biotechnol. 2012, 88: 1266-1272.View ArticleGoogle Scholar
- Chandel AK, Antunes FAF, Freitas WLC, da Silva SS: Sequential acid–base pretreatment of sugarcane bagasse: a facile method for the sugars recovery after enzymatic hydrolysis. J Bioproc Eng Bioref. 2013, 2: 1-9. 10.1166/jbeb.2013.1041.View ArticleGoogle Scholar
- Chandel AK, Antunes FAF, Anjos V, Bell MJV, Rodrigues LN, Singh OV, Rosa CA, Pagnocca FC, da Silva SS: Ultra-structural mapping of sugarcane bagasse by oxalic acid fiber expansion (OAFEX) and ethanol production by Candida shehatae and Saccharomyces cerevisiae. Biotechnol Biofuels. 2013, 6: 4-10.1186/1754-6834-6-4.View ArticleGoogle Scholar
- Singh S, Simmons BA, Vogel KP: Visualization of biomass solubilization and cellulose regeneration during ionic liquid pretreatment of switchgrass. Biotechnol Bioeng. 2009, 104: 68-75. 10.1002/bit.22386.View ArticleGoogle Scholar
- Chundawat SPS, Donohoe BS, Sousa LD, Elder T, Agarwal UP, Lu FC, Ralph J, Himmel ME, Balan V, Dale BE: Multi-scale visualization and characterization of lignocellulosic plant cell wall deconstruction during thermochemical pretreatment. Ener Environ Sci. 2011, 4: 973-984. 10.1039/c0ee00574f.View ArticleGoogle Scholar
- Hansen MAT, Hidayat BJ, Mogensen KK, Jeppesen MD, Jørgensen B, Johansen KS, Thygesen LG: Enzyme affinity to cell types in wheat straw (Triticum aestivum L.) before and after hydrothermal pretreatment. Biotechnol Biofuels. 2013, 6: 54-10.1186/1754-6834-6-54.View ArticleGoogle Scholar
- Cadete RM, Melo MA, Dussán KJ, Rodrigues RC, da Silva SS, Zilli JE, Vital MJ, Gomes FC, Lachance MA, Rosa CA: Diversity and physiological characterization of D-xylose-fermenting yeasts isolated from the Brazilian Amazonian forest. PLoS One. 2012, 7: e43135-10.1371/journal.pone.0043135.View ArticleGoogle Scholar
- Urbina H, Blackwell M: Multi-locus phylogenetic study of the Scheffersomyces yeast clade and characterization of the N-terminal region of xylose reductase gene. PLoS One. 2012, 7: e39128-10.1371/journal.pone.0039128.View ArticleGoogle Scholar
- Martín C, Galve M, Wahlbom F, Hagerdal BH, Jonsson LJ: Ethanol production from enzymatic hydrolysates of sugarcane bagasse using recombinant xylose–utilizing Saccharomyces cerevisiae. Enzyme Microbiol Technol. 2012, 31: 274-282.View ArticleGoogle Scholar
- Rocha GJM, Martin C, Soares IB, Souto-Maior AM, Baudel HM, de Abreu CAM: Dilute mixed-acid pretreatment of sugarcane bagasse for ethanol production. Biomass Bioener. 2011, 35: 663-670. 10.1016/j.biombioe.2010.10.018.View ArticleGoogle Scholar
- Rabelo SC, Carrere H, Filho RM, Costa AC: Production of bioethanol, methane and heat from sugarcane bagasse in a biorefinery concept. Bioresour Technol. 2011, 102: 7887-7895. 10.1016/j.biortech.2011.05.081.View ArticleGoogle Scholar
- Chandel AK, Kapoor RK, Singh AK, Kuhad RC: Detoxification of sugarcane bagasse hydrolysate improves ethanol production by Candida shehatae NCIM 3501. Bioresour Technology. 2007, 98: 1947-1950. 10.1016/j.biortech.2006.07.047.View ArticleGoogle Scholar
- Canilha L, Santos VTO, Rocha GJM, Silva JBA, Giulietti M, da Silva SS, Felipe MGA, Ferraz AL, Milagres AMF, Carvalho W: A study on the pretreatment of a sugarcane bagasse sample with dilute sulfuric acid. J Ind Microbiol Biotechnol. 2011, 38: 1467-1475. 10.1007/s10295-010-0931-2.View ArticleGoogle Scholar
- Rocha GJM, Martín C, da Silva VF, Gomez EO, Goncalves AR: Mass balance of pilot-scale pretreatment of sugarcane bagasse by steam explosion followed by alkaline delignification. Bioresour Technol. 2012, 111: 447-452.View ArticleGoogle Scholar
- Santos VTO, Esteves PJ, Milagres AMF, Carvalho W: Characterization of commercial cellulases and their use in the saccharification of a sugarcane bagasse sample pretreated with dilute sulfuric acid. J Ind Microbiol Biotechnol. 2011, 38: 1089-1098. 10.1007/s10295-010-0888-1.View ArticleGoogle Scholar
- Martín C, Rocha GJM, Santos JRA, Wanderley MCA, Gouveia ER: Enzyme loading dependence of cellulose hydrolysis of sugarcane bagasse. Quim Nova. 2012, 35: 1927-1930. 10.1590/S0100-40422012001000007.View ArticleGoogle Scholar
- Shogren RL, Peterson SC, Evans KO, Kenar JA, Kenar JA: Preparation and characterization of cellulose gels from corn cobs. Carb Polym. 2011, 86: 1351-1357. 10.1016/j.carbpol.2011.06.035.View ArticleGoogle Scholar
- FitzPatrick MA: PhD thesis. Characterization and processing of lignocellulosic biomass in ionic liquids. 2011, ON: Queen’s UniversityGoogle Scholar
- Segal L, Creely JJ, Martin AE, Conrad CM: An empirical method for estimating the degree of crystallinity of native cellulose using the X-ray diffractometer. Text Res J. 1962, 29: 786-794.View ArticleGoogle Scholar
- Park S, Baker JO, Himmel ME, Parilla PA, Jonhson DK: Cellulose crystallinity index: measurement techniques and their impact on interpreting cellulase performance. Biotechnol Biofuels. 2010, 3: 1-10. 10.1186/1754-6834-3-1.View ArticleGoogle Scholar
- Sindhu R, Binod P, Satyanagalakshmi K, Janu KU, Sajna KV, Kurien N, Sukumaran RK, Pandey A: Formic acid as a potential pretreatment agent for the conversion of sugarcane bagasse to bioethanol. Appl Biochem Biotechnol. 2010, 162: 2313-2323. 10.1007/s12010-010-9004-2.View ArticleGoogle Scholar
- Velmurugan R, Muthukumar K: Utilization of sugarcane bagasse for bioethanol production: sono-assisted acid hydrolysis approach. Bioresour Technol. 2011, 102: 7119-7123. 10.1016/j.biortech.2011.04.045.View ArticleGoogle Scholar
- Binod P, Satyanagalakshmi K, Sindhu R, Janu KU, Sukumaran RK, Pandey A: Short duration microwave assisted pretreatment enhances the enzymatic saccharification and fermentable sugar yield from sugarcane bagasse. Ren Ener. 2012, 37: 109-116. 10.1016/j.renene.2011.06.007.View ArticleGoogle Scholar
- Pandey KK, Pitman AJ: FTIR studies of the changes in wood chemistry following decay by brown-rot and white-rot fungi. Int Biodet Biodeg. 2003, 52: 151-160. 10.1016/S0964-8305(03)00052-0.View ArticleGoogle Scholar
- Oh SY, Yoo D, Shin Y, Kim HC, Kim HY, Chung YS, Park WH, Youk JH: Crystalline structure analysis of cellulose treated with sodium hydroxide and carbon dioxide by means of X-ray diffraction and FTIR spectroscopy. Carbohydr Res. 2005, 340: 2376-2391. 10.1016/j.carres.2005.08.007.View ArticleGoogle Scholar
- Colom X, Carrillo F, Nogués F, Garriga P: Structural analysis of photodegraded wood by means of FTIR spectroscopy. Polym Deg Stab. 2003, 80: 543-549. 10.1016/S0141-3910(03)00051-X.View ArticleGoogle Scholar
- Pandey KK: Study of the effect of photo-irradiation on the surface chemistry of wood. Polym Deg Stab. 2005, 90: 9-20. 10.1016/j.polymdegradstab.2005.02.009.View ArticleGoogle Scholar
- Pandey KK: A study of chemical structure of soft and hardwood and wood polymers by FTIR spectroscopy. J Appl Polym Sci. 1999, 12: 1969-1975.View ArticleGoogle Scholar
- Ivanova NV, Korolok EV: IR spectrum of cellulose. J Appl Spec. 1989, 51: 847-851. 10.1007/BF00659967.View ArticleGoogle Scholar
- Hinterstoisser B, Salmén L: Two-dimensional step-scan FTIR: a tool to unravel the OH-valency-range of the spectrum of Cellulose I. Cellulose. 1999, 6: 251-263.View ArticleGoogle Scholar
- Cao Y, Huimin T: Structural characterization of cellulose with enzymatic treatment. J Mol Str. 2004, 705: 189-193. 10.1016/j.molstruc.2004.07.010.View ArticleGoogle Scholar
- Stuart BH: Infrared Spectroscopy: Fundamentals and Applications. 2004, Chichester: Wiley-Blackwell, 224-View ArticleGoogle Scholar
- Krongtaew C, Meesner K, Ters T, Fackler K: Qualitative NIR and pretreatment. Bioresour Technol. 2010, 5: 2063-2080.Google Scholar
- Belini UL, Hein PRG, Filho MT, Rodrigues JC, Chaix G: NIR for bagasse content of MDF. Bioresour Technol. 2011, 6: 1816-1829.Google Scholar
- Agarwal UP, Ralph SA: FT-Raman spectroscopy of wood: identifying contributions of lignin and carbohydrate polymers in the spectrum of black spruce (Picea mariana). Appl Spec. 1997, 51: 1648-1655. 10.1366/0003702971939316.View ArticleGoogle Scholar
- Wiley JH, Atalla RH: Band assignments in the Raman spectra of celluloses. Carb Res. 1987, 160: 113-129.View ArticleGoogle Scholar
- Wickholm K, Larsson PT, Iversen T: Assignment of non-crystalline forms in cellulose I by CP/MAS C-13 NMR spectroscopy. Carb Res. 1998, 312: 123-129. 10.1016/S0008-6215(98)00236-5.View ArticleGoogle Scholar
- Templeton DW, Scarlata CJ, Sluiter JB, Wolfrum EJ: Compositional analysis of lignocellulosic feedstocks. 2. Method uncertainties. J Agr Food Chem. 2010, 58: 9054-9062. 10.1021/jf100807b.View ArticleGoogle Scholar
- Hallac BB, Sannigrahi P, Pu Y, Ray M, Murphy RJ, Ragauskas AJ: Biomass characterization of Buddleja davidii: a potential feedstock for biofuel production. J Agr Food Chem. 2009, 57: 1275-1281. 10.1021/jf8030277.View ArticleGoogle Scholar
- Foston MB, Hubbell CA, Ragauskas AJ: Cellulose isolation methodology for NMR analysis of cellulose ultrastructure. Materials. 2011, 4: 1985-2002. 10.3390/ma4111985.View ArticleGoogle Scholar
- Zhao H, Kwak JH, Zhang ZC, Brown HM, Arey BW, Holladay JE: Studying cellulose fiber structure by SEM, XRD, NMR, and acid hydrolysis. Carb Polym. 2007, 68: 235-241. 10.1016/j.carbpol.2006.12.013.View ArticleGoogle Scholar
- Sanchez G, Pilcher L, Roslander C, Modig T, Galbe M, Liden G: Dilute-acid hydrolysis for fermentation of the Bolivian straw material Paja brava. Bioresour Technol. 2004, 93: 249-256. 10.1016/j.biortech.2003.11.003.View ArticleGoogle Scholar
- Sreenath HK, Jeffries TW: Production of ethanol from wood hydrolyzate by yeasts. Bioresour Technol. 2000, 72: 253-260. 10.1016/S0960-8524(99)00113-3.View ArticleGoogle Scholar
- Chandel AK, Narasu ML, Chandrasekhar G, Manikeyam A, Rao LV: Use of Saccharum spontaneum (wild sugarcane) as biomaterial for cell immobilization and modulated ethanol production by thermotolerant Saccharomyces cerevisiae VS3. Bioresour Technol. 2009, 100: 2404-2410. 10.1016/j.biortech.2008.11.014.View ArticleGoogle Scholar
- Alves LA, Felipe MGA, Silva JBA, da Silva SS, Prata AMR: Pretreatment of sugarcane bagasse hemicellulose hydrolysate for xylitol production by Candida guilliermondii. Appl Biochem Biotechnol. 1998, 70/72: 89-98. 10.1007/BF02920126.View ArticleGoogle Scholar
- Gouveia ER, Nascimento RT, Maior AMS, Rocha JM: Validação de metodologia para a caracterização química de bagaço de cana-de-açúcar. Quim Nova. 2009, 32: 1500-1503. 10.1590/S0100-40422009000600026.View ArticleGoogle Scholar
- Miller GL: Use of dinitrosalicylic acid reagent for determination of reducing sugar. Anal Chem. 1959, 31: 426-428. 10.1021/ac60147a030.View ArticleGoogle Scholar
- Parekh SR, Yu S, Wayman M: Adaptation of Candida shehatae and Pichia stipitis to wood hydrolysates for increased ethanol production. Aspen Bib. 1986, 25: 300-3004.Google Scholar
- Kristensen JB, Thygesen LG, Felby C, Jørgensen H, Elder T: Cell-wall structural changes in wheat straw pretreated for bioethanol production. Biotechnol Biofuels. 2008, 1/5: 1-9.Google Scholar
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