HPLC mapping of second generation ethanol production with lignocelluloses wastes and diluted sulfuric hydrolysis/HPLC mapeando a producao de etanol de segunda geracao utilizando residuos lignocelulosicos e hidrolise sulfurica diluida.
Concerns on the depletion of fossil fuel resources and climate changes attributed to C[O.sub.2] emissions give rise to strong global interest in renewable and carbon-neutral energy sources, as well as the production of chemical feedstock from vegetal sources (DOHERTY et al., 2011; ZHU; PAN, 2010). Biomass, one of the more important renewable energy sources, offers many advantages over petroleum-based fuels. The environment, economy and consumers are greatly benefitted when employing several types of biofuels. Moreover, they are biodegradable and contribute towards the planet's sustainability (DEMIRBAS et al., 2008, 2009, 2011).
The production of second generation ethanol from biomass is actually one method to reduce the consumption of crude oil and environmental pollution (BALAT et al., 2008, BALAT, 2011). Brazil is one of the most important producers of ethanol, especially from sugarcane (HARTEMINK, 2008). On the one hand, diluted acid hydrolysis is one of the most popular methods for converting celluloses into ethanol (KRISHNA et al., 2001; XIE et al., 2011; WEN et al., 2010). On the other hand, a great variety of degrading compounds are released during production (CARRASCO et al., 2010), most of which contain inhibitory activities which may affect the process and result in reduced yields and efficiency of biochemical conversion. Therefore, an efficient analytical approach is increasingly needed to quantify these compounds for a better understanding of their roles in the bioconversion process (XIE et al., 2011).
Great efforts have been taken to analyze the degradation products in biomass hydrolysate, with varying degrees of success. High performance liquid chromatography (HPLC) is a method frequently used in hydrolysis liquor analysis, although gas chromatography (GC), coupled to flame ionization or mass spectrometry detection, was also successful in identifying a large variety of organic degrading products (KARAGOZ et al., 2004; KLINKE et al., 2002).
The deployment of GC methodologies in quantitative studies has been impaired by inherent complexities of derived samples of unknown composition. Liquid chromatography (LC) methods, employing post-column UV or refractive index detection, have historically been jeopardized from incomplete resolution of its analysis. As a result, LC analysis of degrading products in hydrolysate samples has typically employed multiple chromatographic modes and several detection strategies, whereas the choice of utilization mainly depends on the analytical class (CHEN et al., 2006; 2009; LUO et al., 2002; PERSSON et al., 2002; ZEAMANN; BOBLETER, 1993).
Current assay demonstrates the production of second generation ethanol, utilizing several wood chips as biomass, for the bioconversion of cellulose into glucose; a sulfuric hydrolysis treatment was performed and the yeast Saccharomyces cerevisiae was used in the fermentation assays. An HPLC method employing the proton-exchange technique was used in the monitoring the process to map the compounds dispersed in the hydrolysate liquor.
Material and methods
All chemicals from analytical grade were purchased from commercial sources. Tests were performed in the Laboratory of the Department of Industrial Technology, Universidade Estadual de Santa Catarina (Udesc), and in the Laboratory of the Department of Chemistry, Universidade Regional de Joinville (Univille).
Samples were collected in wood transformation plants and furniture industries in the southern Brazilian states of Santa Catarina and Parana. The material analyzed hailed from twelve different wood species including: Hymenolobium petraeum, Tabebuia cassinoides, Myroxylon peruiferum, Nectandra lanceolata, Ocotea catharinensis, Cedrelinga catenaeformis, Cedrela fissilis Veil, Ocotea porosa, Laurus nobilis, Balfourodendron riedelianum, Pinus elliotti and Brosimum spp.
During hydrolysis and fermentation assays, at least one experimental condition was duplicated for each set of experiments, ensuring the consistence and accuracy of results. After collected, the samples were cataloged, packed in containers (5 kg) and stored for acclimatization in laboratory at 20[degrees]C during one week. Samples were then milled with a 0.75 mm screen centrifugal mill. Only samples that passed through a 0.6 mm sieve mesh were selected for tests (Tyler system). Klason lignin was determined following guidelines by the Technical Association of the Pulp and Paper Industry (TAPPI, 2002) T-222. Holocellulose content (cellulose + hemi-celluloses) was determined by (TAPPI, 2009) T-203. The determination of density was based on (TAPPI, 2001) T-258. Standards T-264 (TAPPI, 2007) and T-257 (TAPPI, 2012) were also consulted for the tests. Table 1 shows some characteristics of the samples studied.
Samples underwent a 2% ([H.sub.2]S[O.sub.4] v [v.sup.-1]) sulfuric hydrolysis and maintained during 2h in a water bath at 120 [+ or -] 5[degrees]C to partially remove the lignin fractions, so the yeast had an easier access to the cellulose. An additional step of partial delignification was performed with alkaline treatment 1.0% (NaOH v [v.sup.-1]) maintained at 120 [+ or -] 5[degrees]C during 30 min. (VASQUEZ et al., 2007, ZHANG et al., 2010) to correct treatment's pH from 2 to 5.5. The hydrolysis assay was performed in steps following procedures established by the National Renewable Energy Laboratory (NREL). The cellulose solid phase was separated with a hydraulic press and the content was filter by applying 2 ton pressure over an area of 200 [cm.sup.2] (MAEDA et al., 2011).
Strains of Saccharomyces cerevisiae were available from the microbiological culture collection of the Laboratory of Chemistry at Udesc. All materials were previously sterilized using a steam autoclave at 120 [+ or -] 5[degrees]C. For seed culture, strain was grown in an incubator by the agar"malt method consisting of malt (5 g [L.sup.-1]), yeast (5 g [L.sup.-1]) extract, peptone (5 g [L.sup.-1]), agar (20 g [L.sup.-1]) and distilled water (1L), supplemented with (1 g [L.sup.-1]) glucose in a flask. Prior to their use as inocula for fermentations, the culture was aerobically propagated utilizing 200 mL Erlenmeyer flasks. After incubation, the strains were grown overnight in a regulate climate environment (30[degrees]C), stirred at 200 rpm, and placed inside a shaking bath until the concentration reached approximately 3% (w [w.sup.-1] per liter). They were then separated by centrifugation, albeit monitored by optical density measurements (OD-600 nm, Agilent UV-visible Spectroscopy system).
Preparation of inocula
After hydrolysis, 250 g of each processed biomass were separately fermented in 200 mL Erlenmeyer flasks. Samples were inoculated with 3% of the microorganism colony formed and the solution was completed with 50 mL of distilled water. During the fermentation assay, samples were stored in anaerobic conditions (30[degrees]C) in a regulated climate environment during 8 hours, after which HPLC analysis was performed.
Analyses were performed with High Performance Liquid Chromatography (Merck-Hitachi) model (D-7000 IF), with refractive index (RI) detector and single column (Transgenomic ICE-ION/300). The analytical method employed was the proton-exchange technique, with ultra-pure water as mobile phase and 8.5 mM of sulfuric acid as eluent (isocratic).
Acquisition method: acid lacticbon 300; column type: RP18; pump A type: L-7100; solvent A: HAc 1%; solvent B: H2SO4 8.5 mM; solvent C: methanol; solvent D: can; method description: acid lactic determination using column (Transgenomic Ice-Ion 300), chromatography type: HPLC channel: 2, peak quantification: area, calculation method: EXT-STD. Additional parameters employed in HPLC analyses comprised injection volume 20 [micro]L; column-temperature 30[degrees]C. Samples were injected using an auto-sampler with injection volume of 0.25 [micro]L [min.sup.-1].
Calibration of HPLC was used to determine with precision the concentrations of analytics. The calibration curves were from series 4663 (glycerol), and series 4731, for the other target compounds. Figure 1 shows the calibration curve for ethanol.
y = (1/a). x
where: y = surface area, x = concentration of standard compounds is the straight slope, a = angular coefficient.
The Table 2 shows the retention times for the componds analyzed and the isocratic eluent used.
It must be underscored that the acceptability criteria for the acknowledgement of an individual component in validation studies employing high-purity reference samples, required retention times for a given analytic within [+ or -] 2% average for each respective standard used to construct the calibration curve for that analytic (CHENG et al., 2010).
Quantifying target compounds
For the quantification of compounds, approximately 5 mL of hydrolysate fermented liquor from each sample was stored at 2[degrees]C during 30 min. until chromatography analysis. Samples were diluted (1:1 v v-1) with ultra-pure water and filtered with a 0.45-um Millipore membrane (VWR Scientific, Suwanee, GA, USA). Samples were then transferred to a vial (auto-sampler vial specific for chromatography) and placed in the machine carrousel.
HPLC analytical performance
The equipment mapped the chemical compounds dispersed in the hydrolysate liquor in a peak quantification area mode. The final concentration of ethanol after 8 hours of fermentation assay is given in g [L.sup.-1] [h.sup.-1]. Figures 2 and 3 and Tables 3 and 4 demonstrated the main results of current research.
Results and discussion
All target compounds remained within the calibration range. The identification of compounds was satisfactory and they appeared at peaks with good resolution between the analytics. Differences in resolutions among the compounds always respect the interval of [+ or -] 2% for retention times, as expected. The successfully mapped target compounds were glucose, fructose, lactic acid, acetic acid, glycerol and ethanol.
HPLC's capacity to separate the curves and still maintain linear peaks demonstrated its efficiency. Indeed, the temperature of the column remained equal to 30[degrees]C. In evaluating the target compounds, a total time of 40 min. was needed, using flow injection of 0.5 mL min.-1, which remained equal until the end of the analysis.
Glucose fractions which were not consumed by the microorganism during the initial 8h of the fermentation assay were detected using Cedrelinga catenaeformis (0.8356 g [L.sup.-1]) and Laurus nobilis (0.0148 g [L.sup.-1]). The above evidenced positive glucose consumption among the samples.
Moreover, the ability of the microorganism to survive in an environment fixed with pH 5.5 was also demonstrated.
Studying robust cellulosic ethanol production (SPORL), utilizing wood chips of pretreated lodgepole pine and an adapted strain of S. cerevisiae, Tian et al. (2010) produced ethanol ranging between 0.81 and 2.0 g [L.sup.-1] [h.sup.-1], over 4 and 24 hours in the fermentation assay, respectively, in the un-detoxified run.
When Brandberg et al. (2004) used wood chips and HPLC to analyze the hydrolysate liquor, the researchers proved that the more viable strains were able to consume nearly 2.0 g of glucose per gram of biomass during the first 8h in the fermentation assay, with ethanol production rates ranging between 0.1 and to 0.5 g [L.sup.-1] [h.sup.-1] during the same period.
According to Brandberg et al. (2004), even for the most metabolically active strains, the colony-forming capacity decreased by at least two orders of magnitude over the initial 8h. Actually the inhibitory effect of the diluted acid on biomass seemed to be directly linked to the reproductive ability of the microorganism, or biosynthesis, rather than to its catabolic activity.
It is highly relevant to emphasize that several researchers, such as Chen et al. (2009, 2010), Matias et al. (2011) and others, have spent efforts in substantial contributions towards HPLC analytical techniques by monitoring the compounds dispersed in hydrolysate liquor and thereby optimizing the process.
Fructose fractions were detected among all samples, with results varying between 0.012 g [L.sup.-1], using Cedrela fissilis Veil, and 0.15 g [L.sup.-1], using Balfourodendron riedelianum. The reminiscent presence of this carbohydrate indicates that fructose was consumed at a lower rate than glucose during the initial hours of the fermentation assay.
According to Costa et al. (2008), as a result of the catabolic activities of S. cerevisiae, lactic acid was produced during fermentations and consequently ethanol yields were reduced. In current research lactic acid production was detected in Ocotea porosa (0.293 g [L.sup.-1] [h.sup.-1]) and Balfourodendron riedelianum (l.644 g [L.sup.-1] [h.sup.-1]).
It should be underscored that in the evaluation of the production of ethanol on a large scale, all parameters must be completely free from competing source organisms, due to the fact that the whole production could be enormous advantageous, with the possibility to utilize antibiotics, although extra costs are required in the process.
A study conducted by Moreira et al. (2008) demonstrated that the production of microbial lactic acid was present in almost several fermentation tanks of sugarcane in ethanol conversion plants. The same author reported that lactic acid was produced, ranging between 6.84 and 3.48, Mol [L.sup.-1], thereupon contributing with significant diminutions in the total amounts of ethanol produced.
In current study, fractions of acetic acid were also registered among the chromatograms, albeit in small quantities, without ethanol production being affected.
In a previous research, glycerol reduced nutrient effects, thereby reducing hydrolysis and ethanol yields (TENGBORG et al., 2001). Indeed Stenberg et al. (1998), studying biomass-to-ethanol conversions, reported that reductions in cellulose conversion and glycerol accumulation were also observed, with increased recirculation of the process flow.
This compound normally appears as a byproduct of fermentations when utilizing yeasts, as result of the catabolic activities of the microorganism, together with carbonic gas (C[O.sub.2]), certain alcohols, and pyruvic and succinic acids. However, from the quantitative viewpoint, only the most important component is glycerol. However, glycerol fractions were only detected in Ocotea porosa 0.23 g [L.sup.-1].
Cellulosic ethanol was produced with varying results: Myroxylon peruferum (1.69 g [L.sup.-1] [h.sup.-1]); Tabebuia cassiniodes (1.11 g [L.sup.-1] [h.sup.-1]); Balfourodendron riedelianum (1.60 g [L.sup.-1] [h.sup.-1]); Cedrela fssilis Vell. (0.93 g [L.sup.-1] [h.sup.-1]); Cedrelinga catenaeformis (1.91 g [L.sup.-1] [h.sup.-1]); Brosimum (0.77 g [L.sup.-1] [h.sup.-1]); Ocotea catharinensis (0.75 g [L.sup.-1] [h.sup.-1]); Pinus elliotti (1.03 g [L.sup.-1] [h.sup.-1]); Ocotea porosa (1.59 g [L.sup.-1] [h.sup.-1]); Laurus nobilis (1.59 g [L.sup.-1] [h.sup.-1]); Hymenolobium petraeum (1.71 g [L.sup.-1] [h.sup.-1]); and Nectandra lanceolata (0.95 g [L.sup.-1] [h.sup.-1]).
Zhu et al. (2011) found similar results when studying Eucalyptus sludge, sulfuric hydrolysis and enzymatic saccharifications and registered ethanol concentrations ranging between 1.88 and 36.42 g [L.sup.-1], during the initial 120h of the fermentation assay.
Current analysis showed that the highest quantities of ethanol were produced by Cedrelinga catenaeformis, Ocotea porosa, Balfourodendron riedelianum and Laurus nobilis. These samples actually revealed a potential for their employment at high scale biomass in ethanol plants.
From the technical viewpoint, current assay successfully demonstrated the possibility to produce second generation ethanol, utilizing diluted sulfuric hydrolysis and several lignocellulosic materials. In the case of ethanol productivity, the softwoods showed similar productivity to hardwoods. Research also contributes towards the reduction of these raw materials by employing them as fuel. These promising results demonstrated that bioconversion was efficient even without the need of detoxifications or nutrient supplementation.
Current investigation validated the bioconversion of cellulose into ethanol by monitoring the hydrolysate liquor with subsequent fermentation. The ethanol productivity ranged between 1.91 and 0.75 g [L.sup.-1] [h.sup.-1] after 8h of fermentation assay. The yeast showed resistance and positive carbohydrates consumption, coupled to low biosynthesis of degrading compounds. The HPLC proton-exchange technique proved to be a quick, sensitive and precise method to analyze the hydrolysate liquor even after fermented, therefore demonstrating good resolution among the analytics. Furthermore, the risk of degradation of the target compounds under analysis seemed to be low when such technique is used. Calibrations and recoveries for all standard compounds were satisfactory, despite the complex matrix content of the hydrolysate.
BALAT, M. Production of ethanol from lignocellulosic materials via the biochemical pathway: A review. Energy Conversion and Management, v. 52, n. 2, p. 858-887, 2011.
BALAT, M.; BALAT, H.; CAHIDE, O. Progress in ethanol processing. Progress in Energy and Combustion Science, v. 34, n. 5, p. 551-573, 2008.
BRANDBERG, T.; FRANZEN, C. J.; GUSTAFSSON, L. The fermentation performance of nine strains of Saccharomyces cerevisiae in batch and fed-batch cultures in dilute-acid wood hydrolysate. Journal of Bioscience and Bioengineering, v. 98, n. 2, p. 122-125, 2004.
CARRASCO, C.; BAUDEL, H. M.; SENDELIUS, J.; MODIG, T.; ROSLANDER, C.; GALBE, M.; HAHNHAGERDAL, B.; ZACCHI, G.; LIDEN, G. SO2catalyzed steam pretreatment and fermentation of enzymatically hydrolyzed sugarcane bagasse. Enzyme Microbiology and Technology, v. 46, n. 2, p. 64-73, 2010.
CHEN, S. F.; MOWERY, R. A.; CASTLEBERRY, V. A.; VAN WALSUM, G. P.; CHAMBLISS, C. K. High-performance liquid chromatography method for simultaneous determination of aliphatic acid, aromatic acid and neutral degradation products in biomass pretreatment hydrolysates. Journal of Chromatography A, v. 1104, n. 1-2, p. 54-61, 2006.
CHEN, Z.; JIN, X.; WANG, Q.; LIN, Y.; GAN, L. Confirmation and determination of sugars in soft drink products by IEC with ESI-MS. Chromatographia, v. 69, n. 7-8, p. 761-764, 2009.
CHENG, C.; CHEN, C.-S.; HSIEH, P.-H. On-line desalting and carbohydrate analysis for immobilized enzyme hydrolysis of waste cellulosic biomass by column-switching high-performance liquid chromatography. Journal of Chromatography A, v. 1217, n. 14, p. 2104-2110, 2010.
COSTA, V. M.; BASSO, T. O.; ANGELONI, L. H. P.; OETTERER, M.; BASSO, L. C. Production of acetic acid, ethanol and optical isomers of lactic acid by Lactobacillus strains isolated from industrial ethanol fermentations. Ciencia e Agrotecnologia, v. 32, n. 2, p. 503-509, 2008.
DEMIRBAS, A. Biofuels sources, biofuel policy, biofuel economy and global biofuel projections. Energy Conversion Management, v. 49, n. 8, p. 2106-2116, 2008.
DEMIRBAS, M. F.; BALAT, M.; BALAT, H. Potential contribution of biomass to the sustainable energy development. Energy Conversion and Management, v. 50, n. 7, p. 1746-176, 2009.
DEMIRBAS, M. F.; BALAT, M.; BALAT, H. Biowastes-to-biofuels. Energy Conversion and Management, v. 52, n. 4, p. 1815-1828, 2011.
DOHERTY, W. O. S.; MOUSAVIOUN, P.; FELLOWS, C. M. Review: Value-adding to cellulosic ethanol: Lignin polymers. Industrial Crops and Products, v. 33, n. 2, p. 259-276, 2011.
HARTEMINK, A. E. Sugarcane for ethanol: soil and environmental issues. Advances in Agronomy, v. 99, p. 125-182, 2008.
KARAGOZ, S.; BHASKAR, T.; MUTO, A.; SAKATA, Y. Effect of Rb and Cs carbonates for production of phenols from liquefaction of wood biomass. Fuel, v. 83, n. 17-18, p. 2293-2299, 2004.
KLINKE, H. B.; AHRING, B. K.; SCHMIDT, A. S.; THOMSEN, A. B. Characterization of degradation products from alkaline wet oxidation of wheat straw. Bioresource Technology, v. 82, n. 1, p. 15-26, 2002.
KRISHNA, S. H.; REDDY, T. J.; CHOWDARY, G. V. Simultaneous saccharification and fermentation of lignocellulosic wastes to ethanol using a thermotolerant yeast. Bioresource Technology, v. 77, n. 2, p. 193-196, 2001.
LUO, C.; BRINK, D. L.; BLANCH, H. W. Identification of Potential Inhibitors in Conversion of Poplar Hydrolyzate to Ethanol. Biomass and Bioenergy, v. 22, n. 2, p. 125-138, 2002.
MAEDA, R. N.; SERPA, V. I.; ROCHA, V. A. L.; MESQUITA, R. A. A.; ANNA, L. M. M. S.; DE CASTRO, A. M.; DRIEMEIER, C. E.; PEREIRA, N.; POLIKARPOV, I. Enzymatic hydrolysis of pretreated sugar cane bagasse using Penicillium funiculosum and Ttichoderma harzianum cellulases. Process Biochemistry, v. 46, n. 5, p. 1196-1201, 2011.
MATIAS, J.; JERONIMO, G.; LUIS, R.; BARRENA, R. A. Analysis of sugars by liquid chromatography-mass spectrometry in Jerusalem artichoke tubers for ethanol production optimization. Biomass and Bioenergy, v. 35, n. 5, p. 2006-2012, 2011.
MOREIRA, A. L.; ALMEIDA, W. S.; SCABBIA, R. J. A.; TEIXEIRA, R. R. P. Dosagem de acido latico na producao de etanol a partir da cana-de-acucar. Biologico, v. 70, n. 1, p. 35-42, 2008.
PERSSON, P.; ANDERS SON, J.; GORTON, L.; LARSSON, S.; NILVEBRANT, N.-O.; JONSSON, L. J. Effect of different forms of alkali treatment on specific fermentation inhibitors and on the fermentability of lignocellulose hydrolysates for production of fuel ethanol. Journal of Agricultural and Food Chemistry, v. 50, n. 19, p. 5318-5325, 2002.
STENBERG, K.; TENGBORG, C.; GALBE, M.; ZACCHI, G.; PALMQVIST, E.; HAHN-HAGERDAL, B. Recycling of process streams in ethanol production from softwoods based on enzymatic hydrolysis. Applied Biochemistry Biotechnology, v. 70, n. 72, p. 697-708, 1998.
TAPPI-Technical Association of the Pulp and Paper Industry. TAPPI T-222 om-6. Acid insoluble lignin in wood and pulp. Atlanta: Tappi, 2002. (Tappi Test Methods).
TAPPI-Technical Association of the Pulp and Paper Industry. TAPPI T-264 om-97. Preparation of wood for chemical analysis. Atlanta: Tappi, 2007. (Tappi Test Methods).
TAPPI-Technical Association of the Pulp and Paper Industry. TAPPI T-257 om-2. Sampling and preparation wood for analysis. Atlanta: Tappi, 2012. (Tappi Test Methods).
TAPPI-Technical Association of the Pulp and Paper Industry. TAPPI T-203 om-93. Alpha-, beta- and gamma-cellulose in pulp. Atlanta: Tappi, 2009. (Tappi Test Methods).
TAPPI-Technical Association of the Pulp and Paper Industry. TAPPI T-258 om-11. Basic density and moisture content of pulpwood. Atlanta: Tappi, 2001. (Tappi Test Methods).
TENGBORG, C.; GALBE, M.; ZACCHI, G. Reduced inhibition of enzymatic hydrolysis of steam-pretreated softwood. Enzyme and Microbial Technology, v. 28, n. 9-10, p. 835-844, 2001.
TIAN, S.; LUO, X. L.; YANG, X. S.; ZHU, J. Y. Robust cellulosic ethanol production from SPORL-pretreated lodgepole pine using an adapted strain Saccharomyces cerevisiae without detoxification. Bioresource Technology, v. 101, n. 22, p. 8678-8685, 2010.
VASQUEZ, M. P.; SILVA, J. N. C.; SOUZA JR., M. B.; PEREIRA JR., N. Enzymatic hydrolysis optimization to ethanol production by simultaneous saccharification and fermentation. Applied Biochemistry and Biotechnology, v. 137, n. 140, p. 141-154, 2007.
WEN, F.; SUN, J.; ZHAO, H. Yeast surface display of trifunctional mini-cellulosomes for simultaneous saccharification and fermentation of cellulose to ethanol. Applied Environment Microbiology, v. 76, n. 4, p. 1251-1260, 2010.
XIE, R.; TU, M.; WU, Y.; ADHIKARI, S. Improvement in HPLC separation of acetic acid and levulinic acid in the profiling of biomass hydrolysate. Bioresource
Technology, v. 102, n. 7, p. 4938-4942, 2011. ZEAMANN, A. J.; BOBLETER, O. Separation of biomass degradation products by capillary electrophoresis. Advanced Thermochemisty Biomass Conversion, v. 2, p. 953-965, 1993.
ZHANG, J.; DENG, H.; LIN, L.; SUN, Y.; LIU, C. P. S. Isolation and characterization of wheat straw lignin with a formic acid process. Bioresource Technology, v. 101, n. 7, p. 2311-2316, 2010.
ZHU, J. Y.; PAN, X. J. Woody biomass pretreatment for cellulosic ethanol production: technology and energy consumption evaluation. Bioresource Technology, v. 101, n. 13, p. 4992-5002, 2010.
ZHU, Z.-S.; LI, X.-H.; ZHENG, Q.-M.; ZHANG, Z.; YU, Y.; WANG, J.-F.; LIANG, S.-Z.; ZHU, M.-J. Bioconversion of a mixture of paper sludge and extraction liquor from water pre-hydrolysis of Eucaliptus chips to ethanol using separate hydrolysis and fermentation. Bioresources, v. 6, n. 4, p. 5012-5026, 2011.
Received on April 26, 2013.
Accepted on April 4, 2014.
License information: This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.
Diogo Jose Horst (1) *, Rodolfo Reinaldo Hermes Petter (2), Rogerio de Almeida Vieira (3) and Theodoro Marcel Wagner (4)
(1) Departamento de Engenharia de Producao, Universidade Tecnologica Federal do Parana, Av. Monteiro Lobato, s/n, km 4, 84016-210, Ponta Grossa, Parana, Brazil. (2) Departamento de Engenharia de Producao, Universidade Federal do Rio Grande do Sul, Porto Alegre, Rio Grande do Sul, Brazil. (3) Departamento de Engenharia Mecanica, Universidade Federal de Juiz de Fora, Juiz de Fora, Minas Gerais, Brazil. (4) Departamento de Engenharia Quimica, Universidade da Regiao de Joinville, Joinville, Santa Catarina, Brazil. *Author for correspondence. E-mail: email@example.com
Table 1. Physical and chemical properties of the wood species analyzed. Hard/ Name of Species Chemical (%) Softwood Cellulose Hemicellulose Lignin Hardwood Hymenolobium petraeum 42.2 27.2 28.4 Hardwood Myroxylon peruiferum 41.1 25.4 27.3 Hardwood Tabebuia cassinoides 44.2 29.4 25.6 Softwood Nectandra lanceolata 45.4 30.1 23.6 Hardwood Ocotea catharinensis 44.7 27.5 27.7 Hardwood Cedrelinga 40.6 29.5 27.2 catenaeformis Hardwood Cedrela fissilis Vell. 40.4 28.1 29.7 Hardwood Ocotea porosa 43.8 26.9 30.2 Softwood Laurus nobilis 46.7 32.4 20.1 Softwood Balfourodendron 45.1 26.6 22.2 riedelianum Softwood Pinus elliotti 45.3 30.5 22.9 Hardwood Brosimum spp. 44.1 26.5 26.2 Hard/ Physical Softwood (g [cm.sup.-3]) Density Hardwood 0.67 Hardwood 0.61 Hardwood 0.99 Softwood 0.50 Hardwood 0.62 Hardwood 0.50 Hardwood 0.47 Hardwood 0.66 Softwood 0.44 Softwood 0.69 Softwood 0.48 Hardwood 0.54 Table 2. Retention times (RT) and isocratic eluent. Compounds Isocratic eluent ([H.sub.2]S[O.sub.4]) Retention Time (RT) Mobile phase identification (min.) (mM) Glucose 14.93 8.0 Fructose 16.12 8.0 Lactic-acid 20.74 8.0 Glycerol 21.70 8.0 Acetic acid 24.00 8.0 Ethanol 34.04 8.0 Table 3. Quantitative results of mapped standard compounds. Wood species A-Ethanol (min.) (a) (g [L.sup.-1] [h.sup.-1]) (b) 1 Hymenolobium petraeum 35.85 1.71 2 Myroxylon peruferum 33.88 1.69 3 Tabebuia cassiniodes 33.87 1.11 4 Nectandra lanceolata 33.85 0.95 5 Ochotea chatarinensis 34.87 0.75 6 Cedrelinga catenaeformis 33.84 1.91 Wood species B-Fructose (min.) (a) (g [L.sup.-1] [h.sup.-1]) (b) 1 Hymenolobium petraeum *n.d *n.d 2 Myroxylon peruferum *n.d *n.d 3 Tabebuia cassiniodes *n.d *n.d 4 Nectandra lanceolata 16.05 0.03 5 Ochotea chatarinensis *n.d *n.d 6 Cedrelinga catenaeformis *n.d *n.d Wood species C-Glycerol (min.) (a) (g [L.sup.-1] [h.sup.-1]) (b) 1 Hymenolobium petraeum *n.d *n.d 2 Myroxylon peruferum *n.d *n.d 3 Tabebuia cassiniodes *n.d *n.d 4 Nectandra lanceolata *n.d *n.d 5 Ochotea chatarinensis *n.d *n.d 6 Cedrelinga catenaeformis *n.d *n.d Wood species D-Acetic acid (min.) (a) (g [L.sup.-1] [h.sup.-1]) (b) 1 Hymenolobium petraeum 23.93 *n.d 2 Myroxylon peruferum 23.94 *n.d 3 Tabebuia cassiniodes 23.90 *n.d 4 Nectandra lanceolata 23.91 *n.d 5 Ochotea chatarinensis 23.94 *n.d 6 Cedrelinga catenaeformis 23.91 *n.d Wood species F-Lactic acid (min.) (a) (g [L.sup.-1] [h.sup.-1]) (b) 1 Hymenolobium petraeum 20.82 0.89 2 Myroxylon peruferum 20.83 0.68 3 Tabebuia cassiniodes 20.84 0.99 4 Nectandra lanceolata 20.81 0.58 5 Ochotea chatarinensis 20.83 0.77 6 Cedrelinga catenaeformis 20.81 0.53 (a) (min.): retention times, (b)(g [h.sup.-1] [L.sup.-1]): concentration, * n.d: not detected. Table 4. Quantitative results of mapped standard compounds. Wood species A-Ethanol (min.) (a) (g [L.sup.-1] [h.sup.-1]) (b) 7 Cedrelafissilis Veil 33.88 0.93 8 Ocotea porosa 33.83 1.59 9 Laurus nobilis 33.88 1.59 10 Balfourodendron riedelianum 34.01 1.60 11 Pinus elliotti 33.85 1.03 12 Brosimum 33.84 0.77 Wood species B-Fructose (min.) (a) (g [L.sup.-1] [h.sup.-1]) (b) 7 Cedrelafissilis Veil 16.11 0.01 8 Ocotea porosa *n.d *n.d 9 Laurus nobilis 16.08 0.02 10 Balfourodendron riedelianum 15.90 0.15 11 Pinus elliotti *n.d *n.d 12 Brosimum 16.17 0.02 Wood species C-Glycerol (min.) (a) (g [L.sup.-1] [h.sup.-1]) (b) 7 Cedrelafissilis Veil *n.d *n.d 8 Ocotea porosa 21.79 0.23 9 Laurus nobilis *n.d *n.d 10 Balfourodendron riedelianum 21.78 *n.d 11 Pinus elliotti *n.d *n.d 12 Brosimum *n.d *n.d Wood species D-Acetic acid (min.) (a) (g [L.sup.-1] [h.sup.-1]) (b) 7 Cedrelafissilis Veil 23.95 *n.d 8 Ocotea porosa 23.93 *n.d 9 Laurus nobilis 23.93 *n.d 10 Balfourodendron riedelianum 23.83 *n.d 11 Pinus elliotti 23.91 *n.d 12 Brosimum 23.92 *n.d Wood species F-Lactic acid (min.) (a) (g [L.sup.-1] [h.sup.-1]) (b) 7 Cedrelafissilis Veil 20.84 0.75 8 Ocotea porosa 20.82 0.29 9 Laurus nobilis 20.83 1.38 10 Balfourodendron riedelianum 20.64 1.64 11 Pinus elliotti 20.81 0.69 12 Brosimum 20.81 0.84 (a) (min.): retention times, (b) (g [h.sup.-1] [L.sup.-1]): concentration, *n.d: not detected.