journal of biomedicine and biotechnologydownloads.hindawi.com/journals/focusissues/748976.pdfthe...

Biofuels

Journal of Biomedicine and Biotechnology

Biofuels

Journal of Biomedicine and Biotechnology

Biofuels

Copyright © 2011 Hindawi Publishing Corporation. All rights reserved.

This is a focus issue published in volume 2011 of “Journal of Biomedicine and Biotechnology.” All articles are open access articlesdistributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in anymedium, provided the original work is properly cited.

Editorial BoardThe editorial board of the journal is organized into sections that correspond to

the subject areas covered by the journal.

Agricultural Biotechnology

Guihua H. Bai, USAChristopher P. Chanway, CanadaRavindra N. Chibbar, CanadaIan Godwin, Australia

Hari B. Krishnan, USACarol A. Mallory-Smith, USADennis P. Murr, CanadaRodomiro Ortiz, Mexico

B. C. Saha, USAMariam B. Sticklen, USAChiu-Chung Young, Taiwan

Animal Biotechnology

E. S. Chang, USAHans H. Cheng, USABhanu P. Chowdhary, USANoelle E. Cockett, USAPeter Dovc, SloveniaScott C. Fahrenkrug, USADorian J. Garrick, USA

Thomas A. Hoagland, USATosso Leeb, SwitzerlandJames D. Murray, USAAnita M. Oberbauer, USAJorge A. Piedrahita, USADaniel Pomp, USAKent M. Reed, USA

Lawrence Reynolds, USALawrence B. Schook, USAMari A. Smits, The NetherlandsLeon Spicer, USAJ. Verstegen, USAMatthew B. Wheeler, USAKenneth L. White, USA

Biochemistry

Robert Blumenthal, USADavid Ronald Brown, UKSaulius Butenas, USAVittorio Calabrese, ItalyF. Castellino, USARoberta Chiaraluce, ItalyD. M. Clarke, CanadaFrancesca Cutruzzola, ItalyPaul W. Doetsch, USA

Hicham Fenniri, CanadaNick V. Grishin, USAJ. Guy Guillemette, CanadaPaul W. Huber, USAChen-Hsiung Hung, TaiwanMichael Kalafatis, USAB. E. Kemp, AustraliaPhillip E. Klebba, USAWen-Hwa Lee, USA

Richard D. Ludescher, USAGeorge Makhatadze, USALeonid Medved, USASusan A. Rotenberg, USAJason Shearer, USAAndrei Surguchov, USAJohn B. Vincent, USAYujun George Zheng, USA

Bioinformatics

T. Akutsu, JapanMiguel A. Andrade, GermanyMark Y. Borodovsky, USARita Casadio, ItalyArtem Cherkasov, CanadaDavid Corne, UKSorin Draghici, USA

Stavros J. Hamodrakas, GreecePaul Harrison, USAGeorge Karypis, USAJack A. Leunissen, The NetherlandsGuohui Lin, CanadaSatoru Miyano, JapanZoran Obradovic, USA

Florencio Pazos, SpainZhirong Sun, ChinaYing Xu, USAA. Zelikovsky, USAAlbert Zomaya, Australia

Biophysics

Miguel Castanho, PortugalP. Bryant Chase, USAKuo-Chen Chou, USARizwan Khan, India

Ali A. Khraibi, Saudi ArabiaRumiana Koynova, USASerdar Kuyucak, AustraliaJianjie Ma, USA

S. B. Petersen, DenmarkPeter Schuck, USAClaudio M. Soares, Portugal

Cell Biology

Omar Benzakour, FranceSanford I. Bernstein, USAPhillip I. Bird, AustraliaEric Bouhassira, USAMohamed Boutjdir, USAChung-Liang Chien, TaiwanRichard Gomer, USAPaul J. Higgins, USAPavel Hozak, Czech Republic

Xudong Huang, USAAnton M. Jetten, USASeamus J. Martin, IrelandManuela Martins-Green, USAShoichiro Ono, USAGeorge Perry, USAM. Piacentini, ItalyGeorge E. Plopper, USALawrence Rothblum, USA

Michael Sheetz, USAJames L. Sherley, USAG. S. Stein, USARichard Tucker, USAThomas van Groen, USAAndre Van Wijnen, USASteve Winder, UKChuanyue Wu, USABin-Xian Zhang, USA

Genetics

Adewale Adeyinka, USAClaude Bagnis, FranceJ. Birchler, USASusan Blanton, USABarry J. Byrne, USAR. Chakraborty, USADomenico Coviello, ItalySarah H. Elsea, USACelina Janion, Poland

J. Spencer Johnston, USAM. Ilyas Kamboh, USAFeige Kaplan, CanadaManfred Kayser, The NetherlandsBrynn Levy, USAXiao Jiang Li, USAThomas Liehr, GermanyJames M. Mason, USAMohammed Rachidi, France

Raj S. Ramesar, South AfricaElliot D. Rosen, USADharambir K. Sanghera, USAMichael Schmid, GermanyMarkus Schuelke, GermanyWolfgang Arthur Schulz, GermanyJorge Sequeiros, PortugalMouldy Sioud, NorwayRongjia Zhou, China

Genomics

Vladimir Bajic, Saudi ArabiaMargit Burmeister, USASettara Chandrasekharappa, USAYataro Daigo, JapanJ. Spencer Johnston, USA

Vladimir Larionov, USAThomas Lufkin, SingaporeJoakim Lundeberg, SwedenJohn L. McGregor, FranceJohn V. Moran, USA

Yasushi Okazaki, JapanGopi K. Podila, USAMomiao Xiong, USA

Immunology

Hassan Alizadeh, USAPeter Bretscher, CanadaRobert E. Cone, USATerry L. Delovitch, CanadaAnthony L. DeVico, USANick Di Girolamo, AustraliaDon Mark Estes, USASoldano Ferrone, USAJeffrey A. Frelinger, USAJohn Robert Gordon, Canada

James D. Gorham, USASilvia Gregori, ItalyThomas Griffith, USAYoung S. Hahn, USADorothy E. Lewis, USABradley W. McIntyre, USAR. Mosley, USAMarija Mostarica-Stojkovic, SerbiaHans Konrad Muller, AustraliaAli Ouaissi, France

Kanury V. S. Rao, IndiaYair Reisner, IsraelHarry W. Schroeder, USAWilhelm Schwaeble, UKNilabh Shastri, USAYufang Shi, ChinaPiet Stinissen, BelgiumHannes Stockinger, AustriaJ. W. Tervaert, The NetherlandsGraham R. Wallace, UK

Microbial Biotechnology

Jozef Anne, BelgiumYoav Bashan, MexicoMarco Bazzicalupo, ItalyNico Boon, BelgiumLuca Simone Cocolin, Italy

Peter Coloe, AustraliaDaniele Daffonchio, ItalyHan de Winde, The NetherlandsYanhe Ma, ChinaBernd H. A. Rehm, New Zealand

Angela Sessitsch, AustriaEffie Tsakalidou, GreeceJ. Wiegel, USA

Microbiology

D. Beighton, UKSteven R. Blanke, USAStanley Brul, The NetherlandsIsaac K. O. Cann, USAPeter Dimroth, SwitzerlandStephen K. Farrand, USAAlain Filloux, UK

Gad Frankel, UKRoy Gross, GermanyHans-Peter Klenk, GermanyTanya Parish, UKGopi K. Podila, USAFrederick D. Quinn, USADidier A. Raoult, France

Isabel Sa-Correia, PortugalP. L. C. Small, USALori Snyder, UKMichael Thomm, GermanyH. C. van der Mei, The NetherlandsSchwan William, USA

Molecular Biology

Rudi Beyaert, BelgiumMichael Bustin, USADouglas Cyr, USAK. Iatrou, GreeceLokesh Joshi, IrelandDavid W. Litchfield, Canada

Noel F. Lowndes, IrelandWuyuan Lu, USAPatrick Matthias, SwitzerlandJohn L. McGregor, FranceS. L. Mowbray, SwedenElena Orlova, UK

Yeon-Kyun Shin, USAWilliam S. Trimble, CanadaLisa Wiesmuller, GermanyMasamitsu Yamaguchi, Japan

Oncology

Colin Cooper, UKF. M. J. Debruyne, The NetherlandsNathan Ames Ellis, USADominic Fan, USAGary E. Gallick, USADaila S. Gridley, USAXin-yuan Guan, Hong KongAnne Hamburger, USAManoor Prakash Hande, SingaporeBeric Henderson, Australia

Steve B. Jiang, USADaehee Kang, Republic of KoreaAbdul R. Khokhar, USARakesh Kumar, USAMacus Tien Kuo, USAEric W. Lam, UKSue-Hwa Lin, USAKapil Mehta, USAOrhan Nalcioglu, USAVincent C. O. Njar, USA

P. J. Oefner, GermanyAllal Ouhtit, USAFrank Pajonk, USAWaldemar Priebe, USAF. C. Schmitt, PortugalSonshin Takao, JapanAna Maria Tari, USAHenk G. Van Der Poel, The NetherlandsHaodong Xu, USADavid J. Yang, USA

Pharmacology

Abdel A. Abdel-Rahman, USAM. Badr, USAStelvio M. Bandiera, CanadaRonald E. Baynes, USAR. Keith Campbell, USAHak-Kim Chan, AustraliaMichael D. Coleman, UKJ. Descotes, FranceDobromir Dobrev, Germany

Ayman El-Kadi, CanadaJeffrey Hughes, USAKazim Husain, USAFarhad Kamali, UKMichael Kassiou, AustraliaJoseph J. McArdle, USAMark J. McKeage, New ZealandDaniel T. Monaghan, USAT. Narahashi, USA

Kennerly S. Patrick, USAVickram Ramkumar, USAMichael J. Spinella, USAQuadiri Timour, FranceTodd W. Vanderah, USAVal J. Watts, USADavid J. Waxman, USA

Plant Biotechnology

P. L. Bhalla, AustraliaJ. R. Botella, AustraliaElvira Gonzalez De Mejia, USAH. M. Haggman, Finland

Liwen Jiang, Hong KongPulugurtha Bharadwaja Kirti, IndiaYong Pyo Lim, Republic of KoreaGopi K. Podila, USA

Ralf Reski, GermanySudhir Kumar Sopory, India

Toxicology

M. Aschner, USAMichael L. Cunningham, USALaurence D. Fechter, USAHartmut Jaeschke, USA

Youmin James Kang, USAM. Firoze Khan, USAPascal Kintz, FranceR. S. Tjeerdema, USA

Kenneth Turteltaub, USABrad Upham, USA

Virology

Nafees Ahmad, USAEdouard Cantin, USAEllen Collisson, USAKevin M. Coombs, CanadaNorbert K. Herzog, USATom Hobman, CanadaShahid Jameel, India

Fred Kibenge, CanadaFenyong Liu, USAEric Rassart, CanadaGerald G. Schumann, GermanyY.-C. Sung, Republic of KoreaGregory Tannock, Australia

Ralf Wagner, GermanyJianguo Wu, ChinaDecheng Yang, CanadaJiing-Kuan Yee, USAXueping Zhou, ChinaWen-Quan Zou, USA

Contents

Continuous Production of Lipase-Catalyzed Biodiesel in a Packed-Bed Reactor: Optimization andEnzyme Reuse Study, Hsiao-Ching Chen, Hen-Yi Ju, Tsung-Ta Wu, Yung-Chuan Liu, Chih-Chen Lee,Cheng Chang, Yi-Lin Chung, and Chwen-Jen ShiehVolume 2011, Article ID 950725, 6 pages

Performance and Emission Characteristics of Diesel Engine Fueled with Ethanol-Diesel Blends inDifferent Altitude Regions, Jilin Lei, Yuhua Bi, and Lizhong ShenVolume 2011, Article ID 417421, 10 pages

Biomethane Production as an Alternative Bioenergy Source from Codigesters Treating Municipal Sludgeand Organic Fraction of Municipal Solid Wastes, M. Evren Ersahin, Cigdem Yangin Gomec,R. Kaan Dereli, Osman Arikan, and Izzet OzturkVolume 2011, Article ID 953065, 8 pages

In Situ Biodiesel Production from Fast-Growing and High Oil Content Chlorella pyrenoidosa in RiceStraw Hydrolysate, Penglin Li, Xiaoling Miao, Rongxiu Li, and Jianjiang ZhongVolume 2011, Article ID 141207, 8 pages

Molecular Breeding of Advanced Microorganisms for Biofuel Production, Hiroshi Sakuragi,Kouichi Kuroda, and Mitsuyoshi UedaVolume 2011, Article ID 416931, 11 pages

Rapeseed Oil Monoester of Ethylene Glycol Monomethyl Ether as a New Biodiesel, Jiang Dayong,Wang Xuanjun, Liu Shuguang, and Guo HejunVolume 2011, Article ID 293161, 8 pages

Utilization of Biodiesel By-Products for Biogas Production, Nina Kolesarova, Miroslav Hutnan,Igor Bodık, and Viera SpalkovaVolume 2011, Article ID 126798, 15 pages

A Simple and Fast Method for the Production and Characterization of Methylic and Ethylic Biodieselsfrom Tucum Oil via an Alkaline Route, Marcelo Firmino de Oliveira, Andressa Tironi Vieira,Antonio Carlos Ferreira Batista, Hugo de Souza Rodrigues, and Nelson Ramos StradiottoVolume 2011, Article ID 238474, 4 pages

Hindawi Publishing CorporationJournal of Biomedicine and BiotechnologyVolume 2011, Article ID 950725, 6 pagesdoi:10.1155/2011/950725

Research Article

Continuous Production of Lipase-Catalyzed Biodiesel ina Packed-Bed Reactor: Optimization and Enzyme Reuse Study

Hsiao-Ching Chen,1 Hen-Yi Ju,1 Tsung-Ta Wu,1 Yung-Chuan Liu,2 Chih-Chen Lee,2, 3

Cheng Chang,4 Yi-Lin Chung,4 and Chwen-Jen Shieh4

1 Department and Graduate Program of Bioindustry Technology, Dayeh University, Chang-Hua 515, Taiwan2 Department of Chemical Engineering, National Chung Hsing University, Taichung 402, Taiwan3 Derlin Biotech Corporation Ltd., Yun-Lin 640, Taiwan4 Biotechnology Center, National Chung Hsing University, Taichung 402, Taiwan

Correspondence should be addressed to Chwen-Jen Shieh, [email protected]

Received 29 June 2010; Accepted 2 September 2010

Academic Editor: Nico Boon

Copyright © 2011 Hsiao-Ching Chen et al. This is an open access article distributed under the Creative Commons AttributionLicense, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properlycited.

An optimal continuous production of biodiesel by methanolysis of soybean oil in a packed-bed reactor was developed usingimmobilized lipase (Novozym 435) as a catalyst in a tert-butanol solvent system. Response surface methodology (RSM) and Box-Behnken design were employed to evaluate the effects of reaction temperature, flow rate, and substrate molar ratio on the molarconversion of biodiesel. The results showed that flow rate and temperature have significant effects on the percentage of molarconversion. On the basis of ridge max analysis, the optimum conditions were as follows: flow rate 0.1 mL/min, temperature 52.1◦C,and substrate molar ratio 1 : 4. The predicted and experimental values of molar conversion were 83.31± 2.07% and 82.81± .98%,respectively. Furthermore, the continuous process over 30 days showed no appreciable decrease in the molar conversion. The paperdemonstrates the applicability of using immobilized lipase and a packed-bed reactor for continuous biodiesel synthesis.

1. Introduction

Biodiesel, also known as fatty acid methyl esters (FAME),has attracted great attention in recent years because of thedepletion of fossil fuels, increased crude oil price, and itsenvironmental benefits. It is mainly synthesized by transes-terifying triglycerides from soybean, rapeseed, and palm oilswith methanol [1]. The advantages of biodiesel over dieselfuel are its renewability, biodegradability [2], lower emissionof toxic compounds [3], and higher combustion efficiency[4]. However, the industrial scale production of biodieselis limited due to the undesirable byproducts, the recoveryof glycerol, the removal of inorganic salts and water, thetreatment of wastewater, and the requirement of high energy.Many research works related to enzymatic catalysts have beencarried out to overcome these drawbacks [5–7].

Lipases classified as hydrolysis E.C. 3.1.1.3 have bothhydrolytic and synthetic activity. They can catalyze trans-esterification of triglycerides as well as esterification of free

fatty acids to produce biodiesel. Compared to chemicalcatalysis, the lipase-catalyzed synthesis of biodiesel is morecompatible with variations in the quality of raw materials. Itrequires only simple purification steps and conducts undermoderate reaction conditions. The main obstacle for com-mercialization of the lipase-catalyzed process is the cost ofthe enzyme [7, 8]. The cost of Novozym 435 lipase is approx.1000 USD per kg [7]. The alkaline catalyst (NaOH) is approx.0.62 USD per kg [9]. To compensate for the high cost of theenzyme, an effective way is to extend the operational life ofthe lipase, which would significantly increase productivityof a given amount of enzyme and lower the biodieselproduction price. This can be achieved by using immobilizedenzymes. In the batch system, Novozym 435, an immobilizedCandida antarctica lipase B on acrylic resin, could be repeat-edly used 5−50 cycles under their optimal reaction conditions[10–12]. Li et al. [13] reported that the enzyme activity ofNovozym 435 showed no obvious loss after being used for200 cycles. Furthermore, Royon et al. used 13.5% cotton

2 Journal of Biomedicine and Biotechnology

oil and 54% methanol with Novozym 435 in tret-butanolsolvent system to produce biodiesel at 50◦C reaction temper-ature and 24 hours reaction time with 97% yield. Chang etal. reported that canola oil and methanol (molar ratio 1 : 3.5)was employed to biosynthesize biodiesel with 42% Novozym435 at reaction temperature of 38◦C and reaction time of 12.4hours in hexane system. Therefore, we chose Novozym 435 asthe biocatalyst in a pack-bed reactor system in this study.

For industrial scale production, the use of immobilizedenzyme with packed-bed reactor is more cost effective thanthe batch operation [14]. The immobilized lipase is packedinto a column and the reaction mixture is continuouslypumped through the column. The enzyme can be reusedwithout a prior separation. Besides, the advantages of usinga packed-bed reactor also include continuous removal ofglycerol and excess alcohol, effective reuse of enzymes, andthe protection of enzyme particles from mechanical shearstress [14–16]. However, long-term operation of a packed-bed reactor is still a challenge. In a solvent-free system, thepacked-bed reactor can be operated for 3−7 days withoutdecreasing ester yield [17, 18]. The stepwise addition ofmethanol prevented the deactivation of the lipase, whichallowed a constant enzyme activity during 100 days ofoperation [19]. Furthermore, Chen and Wu [20] regeneratedlipase by periodically washing with tert-butanol, and thelipase maintained its activity for 70 days. The stability ofimmobilized lipase also can be improved using tert-butanolas the solvent in the continuous packed-bed reactor. Theenzyme activity was maintained through 5 and 20 days byHalim et al. [21] and Royon et al. [15], respectively.

In this study, we developed a continuous packed-bedreactor, which was packed with Novozym 435 lipase, toproduce biodiesel from soybean oil with methanol. tert-butanol was used as the solvent, because it is an ideal mediumthat enhances miscibility of methanol with vegetable oilsas well as being regenerating agent of lipase [10, 13, 15,20]. Statistical experiment design and RSM analysis wereemployed to investigate the affinities between the reactionvariables (reaction temperature, flow rate, and substratemolar ratio) and response (molar conversion percentage),and to obtain the optimal conditions for continuous packed-bed reactor operation. This paper provides a feasible modelfor long-term operation of packed-bed reactor and shedslight on industrial scale production of biodiesel with enzy-matic catalysts.

2. Materials and Methods

2.1. Materials. Immobilized lipase (triacylglycerol hydrolase,EC 3.1.1.3; Novozym 435) from Candida antarctica, sup-ported on acrylic resin beads, was purchased from NovoNordisk Bioindustrials, Inc. (Bagsvaerd, Denmark). Accord-ing to the commercial production manual, its catalyticactivity was 10,000 PLU/g (propyl laurate units per gram),and it contained 1%-2% (w/w) moisture. Soybean oil waspurchased from Taisun Enterprise Co., Ltd. (Chang-Hua,Taiwan). Methanol and tert-butanol were purchased fromKatayama Chemical Co., Ltd. Tributyrin was purchased fromSigma Chemical Co. (St. Louis, MO, USA). Molecular sieves

(4 A) were purchased from Davison Chemical (Baltimore,MD, USA), and n-hexane was obtained from Merck Chem-ical Co. (Darmstadt, Germany). All other chemicals wereanalytical reagent grade.

2.2. Experimental Design. A 3-level-3-factor Box-Behnkendesign with three replicates at the center was employedin this study, requiring 15 experiments [22, 23]. Thevariables and their levels selected for the study of biodieselsynthesis were reaction temperature (40◦C−60◦C), flow rate(0.1−0.5 mL/min), and substrate molar ratio (1 : 3−1 : 5). Allexperiments in this paper were performed in a 32.5% tert-butanol system with a packed-bed reactor. Table 1 presentsthe independent factor (Xi) levels and the experimentaldesign in terms of coded and uncoded values. To avoid bias,15 runs were performed in a fully random order.

2.3. Enzymatic Transesterification in a Continuous Reactor.All materials were dehydrated by molecular sieves (4 A)for 24 hours. Soybean oil (0.05 mole), tert-butanol (32.5%,based on the weight of soybean oil), and different molarratios of methanol were well mixed in a feeding flask. Thetransesterification reaction was carried out in a packed-bedreactor consisting of a stainless steel tube 25 cm in lengthand with a 0.46 cm inner diameter. The mixture was pumpedthrough the continuous reactor (a packed-bed column with1.7 g Novozym 435 lipase) at the designed conditions. Thesystem was placed into the temperature-controlled chamberto prevent temperature gradients.

2.4. Determination of Biodiesel Yield. The biodiesel for-mation was determined by injecting a 1 μL aliquot insplitless mode into a gas chromatograph (Hewlett Packard7890, Avondale, PA, USA) equipped with a flame-ionizationdetector (FID) and an MXT-65TG-fused silica capillarycolumn (30 m × 0.25 mm i.d.; film thickness 1 μm; Restek,Bellefonte, PA, USA). Injector and FID temperatures wereset at 300◦C and 320◦C, respectively. The oven temperaturewas maintained at 195◦C for 10 minutes. Nitrogen was usedas the carrier gas. The percentage of molar conversion wasdefined as (mmol of biodiesel/3 mmol of initial soybean oil)× 100% and was estimated using peak area integrated by on-line software Hewlett Packard 6890 Series II Chem Station(Hewlett Packard 6890, Avondale, PA, USA).

2.5. Statistical Analysis. The experimental data (Table 1)were analyzed by the response surface regression (RSREG)procedure of SAS software to fit the following second-orderpolynomial equation

Y = βk0 +3∑

i=1

βkixi +3∑

i=1

βkiixi2 +

2∑

i=1

3∑

j=i+1

βki jxix j , (1)

where Y is the response (percent molar conversion), βk0,βki, βkii, and βki j are constant coefficients, and xi is theuncoded independent variable. The ridge max option wasused to compute the estimated ridge of maximum responsefor increasing radii from the center of the original design.

Journal of Biomedicine and Biotechnology 3

0.60.50.40.30.20.10

Flow rate (mL/min)

0

20

40

60

80

100

Mol

arco

nver

sion

(%)

Figure 1: The effects of flow rate on the conversion of soybean oil tobiodiesel. Reaction conditions: temperature 50◦C, substrate molarratio 1 : 4, and 1.7 g enzyme.

3. Results and Discussion

3.1. Effect of Reaction Parameters. Reaction temperature andmolar substrates ratio are crucial parameters affecting theyield of enzymatic synthesis of biodiesel [6]. Many researches[10, 24, 25] showed that lipase such as Novozym 435deactivated at a methanol-to-oil ratio more than 3. Thedeactivation was resulted from the contact between lipaseand insoluble methanol in the reaction mixture. However,the molar conversion increased with the increase of methanolto soybean oil ratio and did not decrease even at the ratioabove 4 : 1 (data not shown). The addition of tert-butanol tothe mixture enhanced miscibility of methanol with vegetableoils and therefore protected lipase from deactivation [10,26]. Tert-butanol also enhanced mass transfer by reducingviscosity of oils [10]. Thus, the lipase maintained highactivity even at high methanol concentration.

Another important parameter affecting the conversionefficiency in a packed-bed reactor is substrate flow rate.Figure 1 showed the conversion decreased with increasingflow rate. The highest conversion (81%) was obtained at aflow rate of 0.1 mL/min. The increase in substrate flow ratecaused reduction in the residence time of substrate in thepacked-bed reactor, which resulted in poor contact betweenlipases and substrates.

The choice of temperature, flow rate, and substrate molarratio range (Table 1) needs to be examined precisely in Box-Behnken design. The optimal conditions of synthesis can befound inside the experimental region through the analyses ofstatistics and contour plots.

3.2. Model Fitting. The major objective of this study isthe development and evaluation of a statistical approachto better understand the affinity between the variables ofa continuous lipase-catalyzed transesterification reactionin continuously packed-bed reactor. On the basis of thisconcept, the large-scale and continuous process can beoptimized to obtain greater efficiency and to decrease human

resource requirements. Compared with one-factor-at-a-timedesign, the RSM employed in this study was more efficient inreducing the number of experimental runs and time requiredto determine the optimal parameters for optimal biodieselproduction.

The RSREG procedure was employed to fit the second-order polynomial (1) to the experimental data (Table 2) witha coefficient of determination, R2 = 0.985. Among the varioustreatments, the highest molar conversion (81.23 ± 1.49%)was treatment no. 2 (0.1 mL/min, 60◦C, and substrate molarratio 1 : 4), and the lowest molar conversion (27.53± 1.76%)was treatment no. 3 (0.5 mL/min, 40◦C, substrate molar ratio1 : 4). Moreover, treatments no. 2, no. 11, and no. 12 showedthat the optimal flow rate was 0.1 mL/min, resulting inapproximately 80% molar conversion. From the SAS outputof RSREG, the second-order polynomial (2) obtained wasgiven below:

Y = 7.188 + 1.613x1 − 163.456x2 + 13.633x3

− 0.023x21 + 1.236x1x2 + 0.292x2

2

+ 0.222x1x3 + 1.125x2x3 − 2.708x23 .

(2)

The analysis of variance (ANOVA) data were presentedin Table 2. The results indicated that the second-orderpolynomial model was an adequate representation of theactual relationship between the response percentage andthe significant variables with very small P-value (.0006).Furthermore, the overall effect of the three synthesis variableson the molar conversion of biodiesel was further analyzed bya joint test (Table 3). These results revealed that the flow rateand temperature were important parameters, which have astatistically significant overall effect (P < .01) on the responsemolar conversion of biodiesel.

3.3. Mutual Effect of Parameters. The relationships betweenreaction factors and response could be better understoodby examining the planned series of contour plots generatedfrom the predicted model. Figures 2(a)−2(c) represented thesame range of temperature (40◦C−60◦C) and substrate molarratio (1 : 3−1 : 5). Overall, the three contour plots presenteda similar behavior in that predicted molar conversionincreased. At a flow rate of 0.1 mL/min, the maximummolar conversion obtained was up to 80% (Figure 2(a))whereas the yield was significantly reduced at 0.5 mL/min(Figure 2(c)). Thus, the most suitable flow rate would bearound 0.1 mL/min.

3.4. Obtaining Optimal Synthesis Conditions. The optimalpoint of synthesis was determined by ridge max analy-sis, which approximates the estimated ridge of maximumresponse for increasing radii from the center of originaldesign. Table 4 indicated that molar conversion increasedwith a reduced flow rate, which is in agreement withour previous study [23]. The ridge max analysis indicatesthat maximum molar conversion was 83.31 ± 2.07% at0.1 mL/min, 52.1◦C, with a 1 : 4 substrate molarratios.


Table 1: 3-level-3-factor Box-Behnken design of experiments.

Factors Predicted

Treatmenta Temperature ( ◦C) Flow rate (mL/min) Molar ratio (alcohol/oil) Yield (%) Yieldc (%)

no. X1 X2 X3 Y1± SD Y2

1 1b (60) 1 (0.5) 0 (4) 55.00 ± 2.53 58.04

2 1 (60) −1 (0.1) 0 (4) 81.23 ± 1.49 82.26

3 −1 (40) 1 (0.5) 0 (4) 27.53 ± 1.76 26.50

4 −1 (40) −1 (0.1) 0 (4) 70.53 ± 1.69 67.49

5 1 (60) 0 (0.3) 1 (5) 72.40 ± 1.69 71.65

6 1 (60) 0 (0.3) −1 (3) 63.65 ± 1.49 60.34

7 −1 (40) 0 (0.3) 1 (5) 39.72 ± 0.93 43.03

8 −1 (40) 0 (0.3) −1 (3) 41.90 ± 0.09 42.65

9 0 (50) 1 (0.5) 1 (5) 56.94 ± 1.09 54.66

10 0 (50) 1 (0.5) −1 (3) 41.14 ± 0.41 41.42

11 0 (50) −1 (0.1) 1 (5) 80.14 ± 2.32 79.87

12 0 (50) −1 (0.1) −1 (3) 79.13 ± 3.88 81.42

13 0 (50) 0 (0.3) 0 (4) 63.98 ± 1.26 63.98

14 0 (50) 0 (0.3) 0 (4) 63.22 ± 0.35 63.98

15 0 (50) 0 (0.3) 0 (4) 64.73 ± 0.71 63.98aThe treatments were run in random order.bThe values (−1), (0), and (1) are coded levels.cCalculated from the refined model.

807570

807570

80 807570

80757065

6055504540

Temperature (◦C)

3

3.5

4

4.5

5

Subs

trat

em

olar

rati

o(a

lcoh

ol/o

il)

(a) Flow rate 0.1 mL/min

70

65

65

65

60

60

60

555045

555045

555045

7065

60555045

6055504540

Temperature (◦C)

3

3.5

4

4.5

5

Subs

trat

em

olar

rati

o(a

lcoh

ol/o

il)

(b) Flow rate 0.3 mL/min

4540

40

40

35

35

25

30

30

30

3530

35

35

6055504540

Temperature (◦C)

3

3.5

4

4.5

5

Subs

trat

em

olar

rati

o(a

lcoh

ol/o

il)

(c) Flow rate 0.5 mL/min

Figure 2: Contour plots of molar conversion of continuous synthesized fatty acid methyl esters. Reaction conditions: constant flow rate with1.7 g enzyme. Numbers inside the plots indicate the molar conversion at given reaction conditions.

Table 2: Analysis of variance for synthesis variables pertaining tothe response of percent molar conversion.

Source df Sum of squares Prob > F

Model 9 0.21783 0.0006

Linear 3 0.18552 <0.0001

Quadratic 3 0.01473 0.0314

Crossproduct 3 0.01757 0.0221

Pure error 2 0.00003

Total error 5 0.00355

R2 0.985

3.5. Model Verification. The biocatalysis of such estersby lipase-catalysis reactions under milder conditions hasbecome a current industrial interest. Many reactions were

Table 3: Analysis of variance for the joint test.

Factor df Sum of squares Prob > Fa

Temperature (X1) 4 1388.879994 0.0010

Flow rate (X2) 4 2269.962598 0.0003

Molar ratio (X3) 4 166.191998 0.0898b

aProb > F: level of significance.bNot significant at P > .01.

conducted by immobilized lipase in a continuous packed-bed bioreactor for minimizing labors and overhead costsin the industry [27]. An optimized enzymatic catalysis ofbiodiesel production with higher yield at reduced cost inthe optimal condition would be more appealing to the con-sumers and benefit to the manufacturers. The validity of the


Table 4: Estimated ridge of maximum response for variable percent molar conversion.

Uncoded factor values

Coded Estimated responseStandard error

X1 X2 X3

radius (% corporation) ( ◦C) (mL/min) (methanol/oil)

0.0 63.98 1.92 50.00 0.30 4.00

0.1 65.97 1.92 50.53 0.28 4.01

0.2 67.91 1.90 50.98 0.27 4.03

0.3 69.82 1.87 51.34 0.25 4.03

0.4 71.71 1.84 51.63 0.23 4.04

0.5 73.59 1.81 51.84 0.21 4.04

0.6 75.49 1.79 51.98 0.19 4.04

0.7 77.39 1.80 52.07 0.17 4.03

0.8 79.33 1.84 52.12 0.15 4.02

0.9 81.30 1.92 52.12 0.13 4.01

1.0 83.31 2.07 52.09 0.10 4.00

302520151050

Reaction time (day)

0

20

40

60

80

100

Mol

arco

nver

sion

Figure 3: The effects of operating time on the conversion ofsoybean oil to biodiesel. Reaction conditions: temperature 52.1◦C,flow rate 0.1 mL/min, substrate molar ratio 1 : 4, and 1.7 g enzyme.

predicted model was examined by conducting experimentsat the suggested optimum synthesis conditions. The valuepredicted by ridge max analysis was 83.31 ± 2.07%, and theactual value was 82.81 ± 0.98%.

3.6. Enzyme Reuse. The operational stability of the con-tinuous packed-bed reactor was shown in Figure 3. Theimmobilized lipase showed excellent stability under itsoptimal conditions. The ridge max analysis suggested theoptimal reaction conditions were flow rate of 0.1 mL/min,temperature of 52.1◦C, and a methanol-to-oil ratio of 1 : 4. Ahigh molar conversion (80%) was obtained and the immobi-lized lipase was operated over 30 days without losing catalyticactivity. Watanabe et al. [19] achieved a high conversion(90%) over 100 days in the stepwise methanolysis withthree columns of continuous packed-bed reactor system;however, they required a large amount of immobilized lipase(10 g). Chen and Wu [20] studied the methanolysis reactionwith a continuous stirred tank reactor and obtained a 70%conversion over 70 days. The drawback is the requirement for

periodical regeneration of the lipase by tert-butanol washing.Halim et al. [21] reported a conversion of 79%-80% over5 days by methanolysis of waste palm oils in a tert-butanolmediated packed-bed system. These results, applied to large-scale biodiesel production, will increase costs associated withreactor design and poor operational stability. The packed-bed reactor proposed in this paper not only extended theusability of the lipase but also improved the conversionefficiency in the continuous one-step operation system.

4. Conclusions

We developed an optimal packed-bed reactor for continuousproduction of biodiesel from soybean oil in a tert-butanolsolvent system. RSM and 3-factor-3-level Box-Behnkendesign were employed for optimization of transesterificationof soybean oil. The ridge max analysis indicates thatmaximum molar conversion was 83.31% at 0.1 mL/min,52.1◦C, with a 1 : 4 substrate molar ratio. The continuouspacked-bed reactor can operate more than 30 days withoutan appreciable loss in substrate conversion. This systemdemonstrates the potential for industrial scale production ofbiodiesel by using a packed-bed reactor with a tert-butanolsolvent system.

References

[1] I. Kralova and J. Sjoblom, “Biofuels-renewable energy sources:a review,” Journal of Dispersion Science and Technology, vol. 31,no. 3, pp. 409–425, 2010.

[2] H. K. Speidel, R. L. Lightner, and I. Ahmed, “Biodegradabilityof new engineered fuels compared to conventional petroleumfuels and alternative fuels in current use,” Applied Biochemistryand Biotechnology A, vol. 84−86, no. 1−9, pp. 879–897, 2000.

[3] USEPA, “A comprehensive analysis of biodiesel impacts onexhaust emissions,” Draft Technical Report, 2002.

[4] A. Demirbas, “Fuel properties and calculation of higherheating values of vegetable oils,” Fuel, vol. 77, no. 9-10, pp.1117–1120, 1998.


[5] A. Bajaj, P. Lohan, P. N. Jha, and R. Mehrotra, “Biodieselproduction through lipase catalyzed transesterification: anoverview,” Journal of Molecular Catalysis B: Enzymatic, vol. 62,no. 1, pp. 9–14, 2010.

[6] M. Szczesna Antczak, A. Kubiak, T. Antczak, and S. Bielecki,“Enzymatic biodiesel synthesis—key factors affecting effi-ciency of the process,” Renewable Energy, vol. 34, no. 5, pp.1185–1194, 2009.

[7] L. Fjerbaek, K. V. Christensen, and B. Norddahl, “A reviewof the current state of biodiesel production using enzymatictransesterification,” Biotechnology and Bioengineering, vol.102, no. 5, pp. 1298–1315, 2009.

[8] W. Du, W. Li, T. Sun, X. Chen, and D. Liu, “Perspectivesfor biotechnological production of biodiesel and impacts,”Applied Microbiology and Biotechnology, vol. 79, no. 3, pp. 331–337, 2008.

[9] M. J. Haas, A. J. McAloon, W. C. Yee, and T. A. Foglia,“A process model to estimate biodiesel production costs,”Bioresource Technology, vol. 97, no. 4, pp. 671–678, 2006.

[10] M. M. R. Talukder, J. C. Wu, T. B. van Nguyen, N. M. Fen,and Y. L. S. Melissa, “Novozym 435 for production of biodieselfrom unrefined palm oil: comparison of methanolysis meth-ods,” Journal of Molecular Catalysis B: Enzymatic, vol. 60, no.3-4, pp. 106–112, 2009.

[11] M. M. R. Talukder, J. C. Wu, N. M. Fen, and Y. L. S. Melissa,“Two-step lipase catalysis for production of biodiesel,” Bio-chemical Engineering Journal, vol. 49, no. 2, pp. 207–212, 2010.

[12] D. Yu, L. Tian, H. Wu et al., “Ultrasonic irradiation withvibration for biodiesel production from soybean oil byNovozym 435,” Process Biochemistry, vol. 45, no. 4, pp. 519–525, 2010.

[13] L. Li, W. Du, D. Liu, L. Wang, and Z. Li, “Lipase-catalyzedtransesterification of rapeseed oils for biodiesel productionwith a novel organic solvent as the reaction medium,” Journalof Molecular Catalysis B: Enzymatic, vol. 43, no. 1−4, pp. 58–62,2006.

[14] P. M. Nielsen, J. Brask, and L. Fjerbaek, “Enzymatic biodieselproduction: technical and economical considerations,” Euro-pean Journal of Lipid Science and Technology, vol. 110, no. 8,pp. 692–700, 2008.

[15] D. Royon, M. Daz, G. Ellenrieder, and S. Locatelli, “Enzymaticproduction of biodiesel from cotton seed oil using t-butanol asa solvent,” Bioresource Technology, vol. 98, no. 3, pp. 648–653,2007.

[16] A. Robles-Medina, P. A. Gonzalez-Moreno, L. Esteban-Cerdan, and E. Molina-Grima, “Biocatalysis: towards evergreener biodiesel production,” Biotechnology Advances, vol. 27,no. 4, pp. 398–408, 2009.

[17] C. Chang, J.-H. Chen, C.-M. J. Chang, T.-T. Wu, andC.-J. Shieh, “Optimization of lipase-catalyzed biodiesel byisopropanolysis in a continuous packed-bed reactor usingresponse surface methodology,” New Biotechnology, vol. 26,no. 3-4, pp. 187–192, 2009.

[18] N. Ognjanovic, D. Bezbradica, and Z. Knezevic-Jugovic,“Enzymatic conversion of sunflower oil to biodiesel in asolvent-free system: process optimization and the immobi-lized system stability,” Bioresource Technology, vol. 100, no. 21,pp. 5146–5154, 2009.

[19] Y. Watanabe, Y. Shimada, A. Sugihara, and Y. Tominaga,“Enzymatic conversion of waste edible oil to biodiesel fuel ina fixed-bed bioreactor,” Journal of the American Oil Chemists’Society, vol. 78, no. 7, pp. 703–707, 2001.

[20] J.-W. Chen and W.-T. Wu, “Regeneration of immobilizedCandida antarctica lipase for transesterification,” Journal ofBioscience and Bioengineering, vol. 95, no. 5, pp. 466–469,2003.

[21] S. F. A. Halim, A. H. Kamaruddin, and W. J. N. Fernando,“Continuous biosynthesis of biodiesel from waste cookingpalm oil in a packed bed reactor: optimization using responsesurface methodology (RSM) and mass transfer studies,”Bioresource Technology, vol. 100, no. 2, pp. 710–716, 2009.

[22] SAS, SAS User Guide, SAS Institute, Campus Drive Cary, NC,USA, 1990.

[23] S.-F. Chang, S.-W. Chang, Y.-H. Yen, and C.-J. Shieh, “Opti-mum immobilization of Candida rugosa lipase on Celite byRSM,” Applied Clay Science, vol. 37, no. 1-2, pp. 67–73, 2007.

[24] S. Tamalampudi, M. R. Talukder, S. Hama, T. Numata, A.Kondo, and H. Fukuda, “Enzymatic production of biodieselfrom Jatropha oil: a comparative study of immobilized-wholecell and commercial lipases as a biocatalyst,” BiochemicalEngineering Journal, vol. 39, no. 1, pp. 185–189, 2008.

[25] Y. Shimada, Y. Watanabe, A. Sugihara, and Y. Tominaga,“Enzymatic alcoholysis for biodiesel fuel production andapplication of the reaction to oil processing,” Journal ofMolecular Catalysis. B Enzymatic, vol. 17, no. 3-5, pp. 133–142,2002.

[26] W. Du, D. Liu, L. Li, and L. Dai, “Mechanism explorationduring lipase-mediated methanolysis of renewable oils forbiodiesel production in a tert-butanol system,” BiotechnologyProgress, vol. 23, no. 5, pp. 1087–1090, 2007.

[27] N. S. Nielsen, T. Yang, X. Xu, and C. Jacobsen, “Productionand oxidative stability of a human milk fat substitute producedfrom lard by enzyme technology in a pilot packed-bedreactor,” Food Chemistry, vol. 94, no. 1, pp. 53–60, 2006.


Research Article

Performance and Emission Characteristics of Diesel EngineFueled with Ethanol-Diesel Blends in Different Altitude Regions

Jilin Lei, Yuhua Bi, and Lizhong Shen

Faculty of Transportation Engineering, Kunming University of Science and Technology, Kunming 650224, China

Correspondence should be addressed to Lizhong Shen, [email protected]

Received 11 June 2010; Revised 5 October 2010; Accepted 8 November 2010

Academic Editor: Andrei Surguchov

Copyright © 2011 Jilin Lei et al. This is an open access article distributed under the Creative Commons Attribution License, whichpermits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

In order to investigate the effects ethanol-diesel blends and altitude on the performance and emissions of diesel engine, thecomparative experiments were carried out on the bench of turbo-charged diesel engine fueled with pure diesel (as prototype)and ethanol-diesel blends (E10, E15, E20 and E30) under different atmospheric pressures (81 kPa, 90 kPa and 100 kPa). Theexperimental results indicate that the equivalent brake-specific fuel consumption (BSFC) of ethanol-diesel blends are betterthan that of diesel under different atmospheric pressures and that the equivalent BSFC gets great improvement with the rise ofatmospheric pressure when the atmospheric pressure is lower than 90 kPa. At 81 kPa, both HC and CO emissions rise greatly withthe increasing engine speeds and loads and addition of ethanol, while at 90 kPa and 100 kPa their effects on HC and CO emissionsare slightest. The changes of atmospheric pressure and mix proportion of ethanol have no obvious effect on NOx emissions. Smokeemissions decrease obviously with the increasing percentage of ethanol in blends, especially atmospheric pressure below 90 kPa.

1. Introduction

Recently, diesel engine has received considerable attentionbecause of its high heat efficiency and low emission; however,with the stringent emission standard and limited petroleumreserve, alternative fuels for diesel engine have been used.As a renewable and oxygen-containing biofuel, ethanol is aprospective fuel for vehicle, which can be blended with dieselor be injected into cylinder directly. There are many studieson the application of ethanol on diesel engine, which focuson the three aspects: application techniques of ethanol ondiesel engine, fuel properties of ethanol-diesel blends, andeffects on the combustion and emission characteristics ofethanol-diesel blends [1–6].

Because ethanol is polar molecule and its solubility indiesel is prone to be affected by temperature and watercontent, high percentage addition of ethanol to diesel isdifficult, especially under low temperature (below about10◦C). In order to mix ethanol and diesel, an emulsifier orcosolvent should be added. Many literatures indicated thataromatic hydrocarbon, middle distillate, and wax content ofdiesel are important factors of its blend with ethanol [1, 2].Presently, the application techniques of ethanol on diesel

engine can be divided into the following four classes: (1)ethanol-diesel blend by high-pressure pump [3], (2) ethanolfumigation to the intake air charge by using carburetionor manifold injection, which is associated with limits tothe amount of ethanol due to the incipience of engineknock at high loads, and prevention of flame quenchingand misfire at low loads [3–6], (3) dual injection systemrequiring an extra high-pressure injection system and arelated major design change of the cylinder head [6, 7], and(4) blends of ethanol and diesel fuel by using an emulsifier orcosolvent to mix the two fuels for preventing their separation,requiring no technical modifications on the engine side[6, 8, 9].

The physical and chemical characteristics of ethanol-diesel blends are very important to its application on dieselengine. The stability, density, viscosity, surface tension,specific heat, heat value, and cetane number of blendshave great impact on the injection, atomization, ignition,and combustion properties, as well as cold start, power,fuel consumption, and emission characteristics of engine.Additionally, the poking and leakage of conventional tank,fuel pipe, and sealing part can be rendered. More stringentdemands are necessary for the mixture, transportation,


storage, and usage of fuel because of low flash point ofethanol-diesel blends [9–13].

The cetane number is an important fuel propertyfor diesel engines. It has an influence on engine startability, emissions, peak cylinder pressure, and combustionnoise. According to research carried out by Li et al. [12],each 10-vol% ethanol added to the diesel fuel, results ina 7.1-unit reduction in cetane number of the resultingblend. References [8, 14, 15] indicated that the additionof ethanol resulted in increased ignition delay, reducedcombustion duration, high maximum pressure rates, andslightly decreased gas temperature because of its low cetanenumber and high/low heat value. With the addition of cetanenumber improver, the combustion properties can reach thelevel of prototype at middle-high load.

Without modification, the ethanol-diesel blendsdecreased the power of diesel engine and increased thebrake-specific fuel consumption; however, the performanceof prototype can be rehabilitated after adjusting the fueldelivery and injection timing of engine [16–18]. Reference[19] showed no significant power reduction in the engineoperation on different blends of ethanol-diesel (up to 20%)at a 5% level of significance. Brake-specific fuel consumptionincreased by up to 9% as compared to diesel alone. Theexhaust gas temperature and lubricating oil temperatureswere lower with operations on ethanol-diesel blends ascompared to operation on diesel.

Ethanol-diesel blends can reduce the smoke and PMemissions of diesel engine. The higher this reduction is,the higher the percentage of ethanol is in the blends. Thereason is that the oxygen content in blends can promote thecombination of fuel and oxygen, even in fuel-rich region[16, 20–22]. The NOx emissions remained the same orvery slightly reduced with the use of the ethanol-diesel fuelblends with respect to those of the diesel; however, theNOx emissions can be reduced by other techniques, suchas EGR and SCR. The hydrocarbons (HCs) emissions wereincreased with the use of ethanol-diesel blends. The higherthis increase is, the higher the percentage of ethanol in theblend, however, the HC emissions of blends can still meet theemission standards due to low HC emissions of diesel engine.References [12, 20] showed that the CO emissions of ethanol-diesel blends were increased at low load and were decreasedat high load. Additionally, the CO2 emissions were decreaseddue to the low C/H ratio of ethanol-diesel blends.

The irregular emissions of diesel engine were also affectedby the addition of ethanol. Cheung et al. [23] reported thatthe unburned ethanol and acetaldehyde increased when a4-cyclinder direct-injection diesel engine was fueled withethanol-diesel blends, but formaldehyde, ethene, ethyne,1,3-butadiene, and BTX (benzene, toluene, and xylene) ingeneral decreased, especially at high engine load. A dieseloxidation catalyst (DOC) is found to reduce significantlymost of the pollutants, including the air toxics. Song et al.[24] showed that the content of 16 kinds of PAHs and DNAdamage level decreased in exhaust of E5 compare with thatof diesel.

The atmospheric pressure and air density can affect thecombustion process of engine, so the power performance,

Table 1: Engine Configuration.

Type In-line, 4 cylinders

Bore× stroke (mm) 100× 105

Displacement (L) 3.298

Combustion chamber ω-type direct injection

Induction system Turbocharged and intercooler

Compression ratio 17.5 : 1

Rated power (kW/(r·min−1)) 73/3200

Maximum torque (N·m/(r·min−1)) 245/2200

fuel consumption, and emission characteristics of engine willbe different when the engine was run at different altitudes.So far, the application researches of ethanol-diesel blendswere almost carried out at low altitude. Therefore, in orderto investigate the effects of ethanol-diesel blends on theperformance and emissions of diesel engine under differentatmospheric pressures, the comparative experiments weredone between the engine fueled with pure diesel (asprototype) and ethanol-diesel blends at different altitudes[25–27].


2.1. Test Engine. The test engine was a 3.298 L, direct-injection, turbocharged diesel engine. The relevant char-acteristic of detailed engine configuration was given inTable 1. During experiment the engine was tested withoutany modification.

2.2. Emission Test Apparatus and the Realization of DifferentAtmospheric Pressures. The emission test devices includedan AC electric dynamometer (AVL AFA Drive 250/4–8),an exhaust analyzer (AVL CEB), a fuel consumption meter(AVL 733), and a smoke meter (AVL 415). The altitude oftest bench is 1912 m, and the local atmospheric pressure is81 kPa. The relative humidity is 40 ∼ 60%, and temperatureranges from 18◦C to 21◦C.

The different atmospheric pressures were produced byan engine condition system (AVL ACS1300/300), which canautomatically controls the atmospheric pressures and inletgas temperatures. The inlet of turbocharger compressor wasconnected to the pressure output of engine condition system,and the pressure sensor and temperature sensor were used.When the c was 81 kPa, the exhaust back pressure wasset at local environmental pressure. When the atmosphericpressure was 90 kPa or 100 kPa, the back pressure of enginewas adjusted to inlet pressure [17, 18].

2.3. Blend of Ethanol and Diesel. A hydraulic vibrationemulsification device was developed, which was installedon the high-pressure pump of diesel engine. The ethanoland diesel were delivered to the emulsification device bytwo fuel delivery systems. The emulsified ethanol/dieselwas injected into the cylinder by pump and injector. Theemulsification device can provide different proportions ofethanol and diesel without modifying engine and stopping


80 90 100

D100 E10E15 E20E30

210

215

220

225

230

235

240

Atmospheric pressure (kPa)

be/(

g/(k

W·h

))

(a) 2200 r/min 230 N·m

240

245

250

255

260

265

270

275

80 90 100

D100 E10E15 E20E30


be/(

g/(k

W·h

))

(b) 3200 r/min 190 N·m

Figure 1: Effects of different atmospheric pressure and mix proportion on equivalent BSFC.

D10

0E

10E

20E

30D

100

D10

0

D10

0

D10

0

D10

0

D10

0

D10

0

D10

0

E10

E10 E10 E

10

E10 E10 E10

E10

E20

E20

E20

E20

E20 E20

E20E20

E30

E30

E30

E30

E30

E30

E30

E30

0

100

200

300

400

500

600

50 140 180

10−6

Ttq/(N·m)

81 (kPa)90 (kPa)100 (kPa)

ϕ(C

O)

Figure 2: Comparison of HC emission of different atmospheric pressure and mix proportion at speed 1400 r/min.

engine. The emulsification device can use the 95% ethanolwithout any emulsifier and surfactant. The test diesel is0# diesel [5].

3. Results and Discussions

3.1. Analysis of Engine Performance. The low heat value (Qi)of ethanol is lower than that of diesel, so it is necessary toconsider the effect of heat value when making comparison ofbrake-specific fuel consumption (BSFC) and then to refer toequivalent BSFC (be), defined as be = BSFC∗Qie/Qid. Qie andQid are the low heat value of ethanol-diesel blends and diesel,respectively. Figure 1 illustrated the comparison of equivalentBSFC under three atmospheric pressures.

It can be seen that be of ethanol-diesel blends are lowerthan those of diesel. The ethanol is an oxygenated fuelwith lower surface tension and boiling point, so the fast

vaporization of ethanol can promote the spray performanceand the formation of mixture gas, which is good for thepremix and diffused combustion. Additionally, the higheroxygen content of ethanol can increase the excess air ratioand improve the heat efficiency. On the other hand, thedecrease of be was not proportioned to the addition ofethanol. Compared to diesel, E10 reduced be by 1.0 ∼ 2.6%,while E15 by 1.8 ∼ 3.0%, E20 by 2.6 ∼ 2.7%, and E30 by1.4 ∼ 2.1%. The results indicated that E15 and E20 hadbetter performance than E10 and E30 because E10 has lowerproportion of ethanol and E30 maybe has bad emulsification.

It can be seen that be of both ethanol-diesel blendsand diesel are decreased with the increase of atmosphericpressure. The reduction of be was great when atmosphericpressure changed from 81 kPa to 90 kPa, while the reductionwas slight when atmospheric pressure changed from 90 kPato 100 kPa.


D10

0

D10

0

D10

0

D10

0

D10

0

D10

0

D10

0

D10

0

D10

0

E10

E10

E10

E10 E

10

E10

E10E

10E10 E20

E20 E20

E20

E20

E20E

20

E20

E20 E30

E30 E

30

E30

E30E30

E30

E30

E30

0

100

200

300

400

50010−6

60 160

Ttq/(N·m)

230

81 (kPa)90 (kPa)100 (kPa)

ϕ(C

O)


D10

0 D10

0

D10

0

D10

0

D10

0

D10

0D10

0

D10

0

E10

E10

E10 E10E

10

E10

E10

E10

E10

E20

E20

E20

E20

E20

E20E

20E20

E20

E30

E30

E30

E30

E30 E

30

E30

E30

E30

0

100

200

300

400

50 140

10−6

Ttq/(N·m)

190

81 (kPa)90 (kPa)100 (kPa)

ϕ(C

O)


D10

0

D10

0

D10

0

D10

0

D10

0

D10

0

D10

0

D10

0

D10

0

E10

E10

E10

E10

E10

E10

E10

E10

E10

E20

E20

E20

E20

E20

E20

E20

E20

E20

E30

E30

E30

E30

E30

E30

E30

E30

E30

10−6

0

200

400

600

800

1000

1200

1400

50 140 180

Ttq/(N·m)

81 (kPa)90 (kPa)100 (kPa)

ϕ(C

O)

Figure 5: Comparison of CO emission of different atmospheric pressure and mix proportion at speed 1400 r/min.


D10

0

D10

0

D10

0

E10

E10E10

E10

E10

E10 E10

E10

E10 E

20E20

E20E

20

E20

E20

E20

E20

E20 E30

E30

E30 E30

E30E

30

E30

E30

E30

D10

0

D10

0

D10

0

D10

0

D10

0

D10

0

60 160

Ttq/(N·m)

230

10−6

0

200

400

600

800

1000

81 (kPa)90 (kPa)100 (kPa)

ϕ(C

O)


D10

0E

10E

20E

30D

100

E10

E20 E

30D

100

E10 E20 E30

D10

0 E10 E20 E30

D10

0E

10E

20E

30D

100

E10 E20 E

30

D10

0E

10E

20 E30

D10

0E

10E

20E

30D

100

E10

E20

E30

50 140

Ttq/(N·m)

190

10−6

0

200

400

600

800

81 (kPa)90 (kPa)100 (kPa)

ϕ(C

O)


50 140 180

Ttq/(N·m)

D10

0E

10 E20

E30

D10

0E

10 E20 E30

D10

0E

10 E20 E30

D10

0E

10 E20

E30

D10

0E

10 E20 E

30D

100

E10 E

20 E30 D10

0E

10 E20 E30

D10

0 E10 E20 E30

D10

0E

10E

20 E30

0

200

400

600

800

1000

120010−6

81 (kPa)90 (kPa)100 (kPa)

ϕ(N

Ox)

Figure 8: Comparison of NOx emission of different atmospheric pressure and mix proportion at speed1400 r/min.


D10

0

D10

0

D10

0

D10

0

D10

0

D10

0

E10

E10

E10

E10

E10

E10

E20

E20

E20

E20 E20

E20E

30 E30

E30

D10

0

D10

0

D10

0

E10

E10

E10

E20

E20

E20

E30 E30

E30

E30E

30E30

0

300

600

900

1200

150010−6

60 160

Ttq/(N·m)

230

81 (kPa)90 (kPa)100 (kPa)

ϕ(N

Ox)

Figure 9: Comparison of NOx emission of different atmospheric pressure and mix proportion at speed 2200 r/min.

D10

0

D10

0

D10

0

E10 E10

E10E20 E20

E20E

30

D10

0E

10E

20 E30 D10

0E

10E

20 E30 D10

0E

10E

20 E30

D10

0E

10E

20 E30 D

100

E10

E20

E30 D10

0E

10E

20E

30

E30

E30

50 140

Ttq/(N·m)

190

10−6

0

200

400

600

800

1000

1200

1400

1600

81 (kPa)90 (kPa)100 (kPa)

ϕ(N

Ox)

Figure 10: Comparison of NOx emission of different atmospheric pressure and mix proportion at speed 3200 r/min.

3.2. Emission Characteristics of HC. The HC emissions ofdiesel-ethanol blends under three atmospheric pressureswere shown in Figures 2, 3, and 4. It can be seen that theHC emissions under different atmospheric pressures showsignificant divergences when the mix proportions, enginespeeds, and loads change. With increasing speeds and loads,the effect of atmospheric pressure on HC emission was notsignificant. At 2200 r/min and 81 kPa, the mix proportionshad great effects on the HC emissions, especially at light load(50 N·m), which rendered the increase by 47% ∼ 293%. Theincrease of HC emissions of E30 was great. The HC emissionincreased with the increasing percentage of ethanol inblends; however, the HC emissions of ethanol-diesel blendsnearly reached the level of prototype at 3200 r/min.

Because the ethanol has higher latent heat of vaporiza-tion, which reduces the gas temperature and promotes thechilling of cylinder wall, the HC emission rises evidentlywith the increasing content of ethanol at low speed andload of engine. When engine speeds and loads go up, thetemperature of gas and combustion chamber wall increases,which accelerates the formation of mixture gas and promotesthe combustion of fuel, so the increasing blends of ethanolhas litter influence on the HC emissions at higher enginespeed and load. Thus, HC emission had slight increase andreached the level of diesel-fueled engine at some engineloads. Due to its higher latent heat of vaporization andlower cetane number, higher proportion of ethanol reducesthe gas temperature and retards the ignition delay, which


0

0.5

1

1.5

2

2.5

3

80 90 100

SF/F

SN

D100E10

E20E30


(a) 1400 r/min 140 N·m

0

0.5

1

1.5

2

2.5

3

SF/F

SN

80 90 100

D100E10

E20E30


(b) 1400 r/min 180 N·m

Figure 11: Comparison of smoke of different atmospheric pressure and mix proportion at speed 1400 r/min.

0

0.5

1

1.5

2

2.5

3

80 90 100

D100E10

E20E30

S F/F

SN


(a) 2200 r/min 160 N·m

0

0.5

1

1.5

2

2.5

3

80 90 100

D100E10

E20E30

S F/F

SN


(b) 2200 r/min 230 N·m


results in the significant rise of HC emissions of E30 at lowerspeed and load. Additionally, the limited emulsifiable abilityof mixture device at higher proportion of ethanol may beanother reason. Based on the above analysis, it can be saidthat HC emissions of ethanol-diesel blends are depended onthe engine speed, load, and the mix proportion of ethanol.

3.3. Emission Characteristics of CO. The CO emissions ofethanol-diesel blends under three atmospheric pressureswere shown in Figures 5, 6, and 7. At 2200 r/min andlow load (50 N·m), E10, E20, and E30 augmented theCO emissions by 20% ∼ 250%, 33% ∼ 301%, and 35% ∼210%, respectively. With increasing engine speed and engineload, atmospheric pressure had litter influence on the COemission. At low and middle loads, the higher proportion

of ethanol increased the CO emission slightly. At full load,CO emissions of ethanol-diesel blends were lower thanthose of pure diesel, especially at 81 kPa. The experimentalresults indicated that the ethanol-diesel blends would notdeteriorate the CO emissions except for 2200 r/min and lowload.

The addition of ethanol causes the reduction of gastemperature, which restrains the oxidation of CO, so COemission goes up at low load. With the increase of enginespeed and load, the increase of gas temperature, walltemperature, and oxygen content of ethanol promote theoxidation condition of CO, which decreases the negativeeffect of addition of ethanol. At full load, the excess airratio is comparatively low, so the increasing proportion ofethanol decreases the CO emission greatly. With the increase


0

0.5

1

1.5

2

2.5

3

80 90 100

D100E10

E20E30

S F/F

SN


(a) 3200 r/min 140 N·m

80 90 100

D100E10

E20E30

0

0.5

1

1.5

2

2.5

3

S F/F

SN


(b) 3200 r/min 190 N·m


of atmospheric pressure, the excess air ratio increases and theeffect of ethanol is weakened, so the influence of atmosphericpressure on the CO emission is slight. Based on the aboveanalysis, it can be said that CO emissions of ethanol-dieselblends are depended on the engine speed, load, and the mixproportion of ethanol.

3.4. Emission Characteristics of NOx. Figures 8, 9, and10 showed the NOx emissions of ethanol-diesel blendsunder three atmospheric pressures. At different atmosphericpressures and mix proportions, the NOx emissions showedthe similar trend. The ethanol-diesel blends reduced the NOxemission at most modes. At 1400 and 2200 r/min and lowload, the slight increase of NOx emission for E30 should berendered by the bad emulsification at higher mix proportion.The increasing oxygen content can promote the formation ofNOx; however, the maximum gas temperature is the mostimportant factor of NOx formation, so the decreased gastemperature caused by higher latent heat of vaporization ofethanol can reduce the NOx emission.

3.5. Emission Characteristics of Smoke. Figures 11, 12, and13 showed the smoke emissions of ethanol-diesel blendsunder three atmospheric pressures at full load. At differentatmospheric pressures, the smoke emissions of ethanol-diesel blends had similar tendency as those of diesel. Thesmoke emissions of both blends and diesel were decreasedwith the increasing atmospheric pressures. Compared withpure diesel, E10, E20, and E30 reduced the smoke emissionsby 18% ∼ 26%, 36% ∼ 47%, and 50% ∼ 63%, respectively,at 81 kPa, by 18% ∼ 19%, 40% ∼ 38%, and 63% ∼ 59%,respectively at 90 kPa, and by 17% ∼ 19%, 34% ∼ 42%, and58% ∼ 62%, respectively, at 100 kPa. It showed that highermix proportion of ethanol resulted in lower smoke emissionat the same atmospheric pressure and load. At 2200 r/min

when atmospheric pressure ranged from 81 kPa to 90 kPa thesmoke emissions of E10, E20, and E30 were reduced by 39%,43%, and 55%, respectively. However, when atmosphericpressure ranged from 90 kPa to 100 kPa, the smoke emissionsof E10, E20, and E30 were reduced by 14%, 6%, and 4%,respectively. It can be seen that atmospheric pressure hassignificant effect on the smoke emission when atmosphericpressure is lower than 90 kPa. The influence is weakenedwhen it is above 90 kPa.

The oxygen atom is usually connected to carbon atom inoxygenated fuel, and it is difficult to break the bond, whichrestrains the formation of aromatic hydrocarbon and blackcarbon, so the oxygen content of ethanol can provide oxygenatom in the fuel-rich region and inhibit the formation ofsmoke, especially at heavy load. At heavy load, the excessair ratio is low, so the oxygen content of ethanol can showgreatly positive effect on the smoke emission. On the otherhand, ethanol has lower carbon and sulfur percentage, littlearomatic hydrocarbon, and lower surface tension and boilingpoint, which can promote the spray and combustion char-acteristics of ethanol-diesel blends and restrain the smokeemission.

4. Conclusions

(1) The power performance of engine fueled withethanol-diesel blends can meet the demand ofprototype after adjusting the fuel delivery. Withincreasing atmospheric pressure, the equivalent spe-cific fuel consumption of both mixtures and purediesel showed the same trend of decrease. Whenthe atmospheric pressure is lower than 90 kPa, theequivalent specific fuel consumption is significantlyimproved with the rise of atmospheric pressure; andthe improvement is weakened when atmosphericpressure is above 90 kPa.


(2) At 81 kPa, the HC emission rises greatly with thedecrease of speed and load and the increase of ethanolcontent, especially at low load. The increasing mixproportion of ethanol has little influence on the HCemission when atmospheric pressure ranges from90 kPa to 100 kPa.

(3) At 81 kPa, the CO emission rises greatly with thedecrease of speed and the increase of ethanol content,especially at low load. At 90 kPa and 100 kPa, the COemission increases slightly with the increasing mixproportion at low and middle load, while the COemission is reduced at heavy load.

(4) Atmospheric pressure and mix proportion have noobvious influence on NOx emission. Under mostworking conditions, NOx emission of ethanol-dieselblends has a slight drop compared to that of diesel.

(5) The smoke emission drops obviously with increasingatmospheric pressure. Furthermore, the higher mixproportion of ethanol results in the lower smokeemission. Atmospheric pressure has significant effecton the smoke emission when it is lower than 90 kPa.The influence is weakened when it is above 90 kPa.

Acknowledgment

This work was supported by the National Natural ScienceFoundation of China (Grant no. 50766001).

References

[1] C. D. Rakopoulos, K. A. Antonopoulos, D. C. Rakopoulos, andD. T. Hountalas, “Multi-zone modeling of combustion andemissions formation in DI diesel engine operating on ethanol-diesel fuel blends,” Energy Conversion and Management, vol.49, no. 4, pp. 625–643, 2008.

[2] A. C. Hansen, Q. Zhang, and P. W. L. Lyne, “Ethanol-dieselfuel blends—a review,” Bioresource Technology, vol. 96, no. 3,pp. 277–285, 2005.

[3] L. Z. Shen, W. S. Yan, Y. H. Bi, and J. L. Lei, “Performance com-parison of ethanol/diesel blends mixed in different methods ofdiesel engine,” Journal of Combustion Science and Technology,vol. 13, no. 5, pp. 389–392, 2007.

[4] M. Abu-Qudais, O. Haddad, and M. Qudaisat, “Effect of alco-hol fumigation on diesel engine performance and emissions,”Energy Conversion and Management, vol. 41, no. 4, pp. 389–399, 2000.

[5] E. A. Ajav, B. Singh, and T. K. Bhattacharya, “Performanceof a stationary diesel engine using vapourized ethanol assupplementary fuel,” Biomass and Bioenergy, vol. 15, no. 6, pp.493–502, 1998.

[6] P. Satge de Caro, Z. Mouloungui, G. Vaitilingom, and J. C.Berge, “Interest of combining an additive with diesel-ethanolblends for use in diesel engines,” Fuel, vol. 80, no. 4, pp. 565–574, 2001.

[7] N. Noguchi, H. Terao, and C. Sakata, “Performance improve-ment by control of flow rates and diesel injection timing ondual-fuel engine with ethanol,” Bioresource Technology, vol. 56,no. 1, pp. 35–39, 1996.

[8] H. Chen, S. J. Shuai, and J. X. Wang, “Study on combustioncharacteristics and PM emission of diesel engines using ester-ethanol-diesel blended fuels,” Proceedings of the CombustionInstitute, vol. 31, pp. 2981–2989, 2007.

[9] E. W. De Menezes, R. da Silva, R. Cataluna, and R. J. C.Ortega, “Effect of ethers and ether/ethanol additives on thephysicochemical properties of diesel fuel and on engine tests,”Fuel, vol. 85, no. 5-6, pp. 815–822, 2006.

[10] S. G. Poulopoulos, D. P. Samaras, and C. J. Philippopoulos,“Regulated and unregulated emissions from an internalcombustion engine operating on ethanol-containing fuels,”Atmospheric Environment, vol. 35, no. 26, pp. 4399–4406,2001.

[11] X. Lu, J. Yang, W. Zhang, and H. Zhen, “Effect of cetanenumber improver on heat release rate and emissions of highspeed diesel engine fueled with ethanol-diesel blend fuel,”Fuel, vol. 83, no. 14-15, pp. 2013–2020, 2004.

[12] D. Li, H. Zhen, X. Lu, W. Zhang, and J. Yang, “Physico-chemical properties of ethanol-diesel blend fuel and its effecton performance and emissions of diesel engines,” RenewableEnergy, vol. 30, no. 6, pp. 967–976, 2005.

[13] B. Q. He, S. J. Shuai, J. X. Wang, and H. He, “The effectof ethanol blended diesel fuels on emissions from a dieselengine,” Atmospheric Environment, vol. 37, no. 35, pp. 4965–4971, 2003.

[14] D. C. Rakopoulos, C. D. Rakopoulos, E. G. Giakoumis, R. G.Papagiannakis, and D. C. Kyritsis, “Experimental-stochasticinvestigation of the combustion cyclic variability in HSDIdiesel engine using ethanol-diesel fuel blends,” Fuel, vol. 87,no. 8-9, pp. 1478–1491, 2008.

[15] C. D. Rakopoulos, K. A. Antonopoulos, and D. C. Rakopoulos,“Experimental heat release analysis and emissions of a HSDIdiesel engine fueled with ethanol-diesel fuel blends,” Energy,vol. 32, no. 10, pp. 1791–1808, 2007.

[16] D. C. Rakopoulos, C. D. Rakopoulos, E. C. Kakaras, and E.G. Giakoumis, “Effects of ethanol-diesel fuel blends on theperformance and exhaust emissions of heavy duty DI dieselengine,” Energy Conversion and Management, vol. 49, no. 11,pp. 3155–3162, 2008.

[17] N. Noguchi, H. Terao, and C. Sakata, “Performance improve-ment by control of flow rates and diesel injection timing ondual-fuel engine with ethanol,” Bioresource Technology, vol. 56,no. 1, pp. 35–39, 1996.

[18] C. Sayin and M. Canakci, “Effects of injection timing onthe engine performance and exhaust emissions of a dual-fueldiesel engine,” Energy Conversion and Management, vol. 50,no. 1, pp. 203–213, 2009.

[19] E. A. Ajav, B. Singh, and T. K. Bhattacharya, “Experimentalstudy of some performance parameters of a constant speedstationary diesel engine using ethanol-diesel blends as fuel,”Biomass and Bioenergy, vol. 17, no. 4, pp. 357–365, 1999.

[20] H. Chen, J. Wang, S. Shuai, and W. Chen, “Study of oxygenatedbiomass fuel blends on a diesel engine,” Fuel, vol. 87, no. 15-16, pp. 3462–3468, 2008.

[21] X. Shi, Y. Yu, H. He, S. Shuai, J. Wang, and R. Li, “Emissioncharacteristics using methyl soyate-ethanol-diesel fuel blendson a diesel engine,” Fuel, vol. 84, no. 12-13, pp. 1543–1549,2005.

[22] M. Lapuerta, O. Armas, and J. M. Herreros, “Emissions from adiesel-bioethanol blend in an automotive diesel engine,” Fuel,vol. 87, no. 1, pp. 25–31, 2008.

[23] C. S. Cheung, Y. Di, and Z. Huang, “Experimental investi-gation of regulated and unregulated emissions from a dieselengine fueled with ultralow-sulfur diesel fuel blended with


ethanol and dodecanol,” Atmospheric Environment, vol. 42, no.39, pp. 8843–8851, 2008.

[24] C. L. Song, Y. C. Zhou, R. J. Huang et al., “Influence ofethanol-diesel blended fuels on diesel exhaust emissions andmutagenic and genotoxic activities of particulate extracts,”Journal of Hazardous Materials, vol. 149, no. 2, pp. 355–363,2007.

[25] L. Shen, Y. Shen, and W. Yan, “Combustion process of dieselengines at regions with different altitudes,” SAE Paper, 950857,1995.

[26] L. Shen, Y. Shen, and W. Yan, “Dimensionless analysis of theproperties of diesel engines at different pressures,” SAE Paper,952064, 1995.

[27] P. Benjumea, J. Agudelo, and A. Agudelo, “Effect of altitudeand palm oil biodiesel fuelling on the performance andcombustion characteristics of a HSDI diesel engine,” Fuel, vol.88, no. 4, pp. 725–731, 2009.


Research Article

Biomethane Production as an Alternative BioenergySource from Codigesters Treating Municipal Sludge andOrganic Fraction of Municipal Solid Wastes

M. Evren Ersahin, Cigdem Yangin Gomec, R. Kaan Dereli, Osman Arikan,and Izzet Ozturk

Department of Environmental Engineering, Istanbul Technical University, Maslak, 34469 Istanbul, Turkey

Correspondence should be addressed to Cigdem Yangin Gomec, [email protected]

Received 27 July 2010; Revised 5 October 2010; Accepted 30 November 2010

Academic Editor: R. S. Tjeerdema

Copyright © 2011 M. Evren Ersahin et al. This is an open access article distributed under the Creative Commons AttributionLicense, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properlycited.

Energy recovery potential of a mesophilic co-digester treating OFMSW and primary sludge at an integrated biomethanization plantwas investigated based on feasibility study results. Since landfilling is still the main solid waste disposal method in Turkey, landscarcity will become one of the most important obstacles. Restrictions for biodegradable waste disposal to sanitary landfills in EULandfill Directive and uncontrolled long-term contamination with gas emissions and leachate necessitate alternative managementstrategies due to rapid increase in MSW production. Moreover, since energy contribution from renewable resources will berequired more in the future with increasing oil prices and dwindling supplies of conventional energy sources, the significanceof biogas as a renewable fuel has been increased in the last decade. Results indicated that almost 93% of annual total costcan be recovered if 100% renewable energy subsidy is implemented. Besides, considering the potential revenue when replacingtransport fuels, about 26 heavy good vehicles or 549 cars may be powered per year by the biogas produced from the proposedbiomethanization plant (PE = 100,000; XPS = 61 g TS/PE·day; XSS-OFMSW = 50 g TS/PE·day).

1. Introduction

Primary energy sources are classified as fossil and renewableenergy sources and nuclear fuels. Globally, most energy isprovided from the fossil energy sources whereas nuclearpower plants contribute to only a very small percentage.Therefore, the dependence of the global economy on fossil-derived fuels, coupled with political instability in oil-pro-ducing countries, has pushed petroleum prices near all-timehighs. On the other hand, although energy supply fromrenewable energy sources has been improved significantly, itscontribution is quite limited. Water, sun, wind, geothermalheat, tides, and biomass are reported as the renewableenergy sources. Since demand for energy is expected to in-crease more than 50% by 2025, there is an ongoing searchto develop sustainable, affordable, environmentally soundenergy from renewable sources [1, 2]. Biofuels derived from

plant-based feedstocks, such as corn and sugarcane, are con-sidered renewable and are an environmentally clean energysource, and they have potential to significantly decrease fossilfuel consumption. Bioethanol and biodiesel can be usedin the form of a gasoline/diesel blend [2]. Besides, biogaswhich is produced at most biological treatment plants hasbeen considered as one of the most important renewableenergy sources. Anaerobic biotechnology has been reportedas a sustainable alternative to current disposal strategiesbecause the volume of the organic waste is reduced andstabilized, a residue (compost) that can be used for soilconditioning is produced, and energy in the form of methaneis recovered [3]. Methane production can be estimated fromchemical oxygen demand (COD) or ultimate biochemicaloxygen demand (BODL) stabilization based on the factthat 1 kg COD destroyed produces 0.35 m3 CH4 at stan-dard temperature and pressure (STP). Although methane


generation is the focus, anaerobic biotechnology may alsoconvert organic-rich waste streams into hydrogen or ethanol.Temperature, pH, alkalinity, hydraulic and organic loadingrates, the presence of toxic and inhibitory substances, type ofsubstrate, and total solids (TS)/volatile solids (VS) contentare the factors affecting the amount of gas produced. Themethane content of biogas is also very important, becauseit imparts fuel value to the gas. It was reported that bio-gas, produced during anaerobic digestion of biodegradableorganic solids, typically contains about 60–65% methane,which is a valuable resource and can be used to offset partof the energy needs for in-plant use. The produced bio-gas can replace natural gas, coal, and electricity. Moreover,the residues resulting from anaerobic digestion are rich innutrients (N and P) and can be land applied which alsosignificantly reduce use of chemical fertilizer. In addition, thetreated effluent can be recycled as process water for in-plantuse [2, 4].

In the countries like Turkey where landfilling is still themain solid waste management method, land scarcity willbecome one of the most important obstacles for municipalsolid wastes (MSW) disposal. It was reported that ca.60× 103 tonnes of MSW are produced per day in Turkey,which includes almost 50% biodegradable materials. Forinstance in Istanbul, almost 45% of the produced MSW isin organic nature whereas 34% of it may be recovered and21% is composed of miscellaneous materials. Thus, MSWshould be considered as the “valuable products” instead ofthe “wastes” [5]. In the framework of the Kyoto agreements,which Turkey has already signed, the production of methanefrom wastes should be encouraged, for example, by subsi-dizing electricity from biogas with the net added price of asmuch as C0.1 per kWh. Since current biomethane potentialof the produced OFMSW in Turkey is about 109 m3 CH4 peryear considering 50% biomethanization of 5× 106 ton/yearbiodegradable waste as volatile solids (VSs), the biometh-anization of OFMSW can become a feasible option byapplying subsidies to electricity production from wastes [6].In year 2000, the law of “Renewable Energies,” which statedthe rules for the subsidization of the power supplied bybiogas facilities, became effective in Europe [1]. It wasreported that around the world, the potential for biogasproduction is so large that it could replace about 20–30% oftotal natural gas consumption [4]. Moreover, the restrictionsfor biodegradable waste disposal in EU Landfill Directive[7] and uncontrolled long-term contamination with gasemissions and leachate necessitate alternative sustainableand integrated management strategies due to rapid increasein the production of MSW, that is, anaerobic biologicaltreatment, either in anaerobic digesters or in landfill biore-actors [8, 9]. It was reported that most wastes with ahigh-solid content, for example, municipal sludge (primarysolids and waste-activated sludge (WAS)), animal manure,food wastes, and agricultural residues contain a significantamount of biodegradable organic carbon. These organic-rich wastes and residues are ideal feedstocks for renewableenergy (methane, hydrogen, or butanol) generation throughnonoxidative metabolism (anaerobic fermentation) [10].Codigestion of the organic fraction of municipal solid

wastes (OFMSW) with other cosubstrates is reported as anattractive alternative for sustainable management of differentseparate waste streams produced at large amounts. Sincecodigestion concept involves the treatment of several typesof waste in a single treatment facility, mixing of several wastetypes has positive effects both on the anaerobic digestionprocess itself and on the treatment economy which showsitself with an increase in methane yield and in processstability. The most appropriate cosubstrate alternatives forcodigestion with OFMSW are reported as the sewage sludgewhich provides codigestion application to be realised inexisting digesters at the treatment plants without greatinvestments, a situation which makes this alternative moreappealing. Moreover, theoretical CH4 content for primarysludge is reported as about 63% [3, 4].

The aim of this study is to evaluate energy recoverypotential if mesophilic (30–35◦C) codigestion of OFMSWwith primary sludge is applied at a full-scale municipalwastewater treatment plant (WWTP) for 100,000 populationequivalent (PE) (Figure 1). Here, the treatment plant iscomposed of an activated sludge system (C, N, and Premoval) with presedimentation. The produced primarysludge will be codigested with the mixed wastes originatingfrom the restaurants and canteens and from the supermar-kets (greengrocery part) where both waste streams werethe source sorted organic fraction of the municipal solidwastes (SS-OFMSWs). In the scope of this study, feasibility,biogas production and its potential revenue when replacingtransport fuels (i.e., diesel or gasoline) were investigated forthe proposed integrated biomethanization plant.

2. The Mass Balance Calculations

In order to calculate the fundamental data for the processdesign and to establish the mass balance, first of all theproduced primary sludge and the OFMSW should bedetermined. The typical raw primary sludge production andits total solids (TSs) content from a municipal wastewatertreatment plant are given in the ranges of 0.92–2.20 L/PE·dayand 2–8%, respectively [11]. If raw primary sludge produc-tion and its TS content are assumed as 1.5 L/PE·day and4%, respectively, primary sludge amount (XPS) is calculatedas about 61 g/PE·day (density ∼= 1010 kg/m3). TS contentand VS/TS ratio for the SS-OFMSW from the restaurantsand canteens are given in the range of 21.4–27.4% and 91–99%, respectively. On the other hand, TS content and VS/TSratio for the SS-OFMSW from the greengrocery parts of thesupermarkets are given in the range of 5.4–13.3% and 78–92%, respectively.

The OFMSW production from municipal sources isabout 0.25 kg/PE·day. If the mixed SS-OFMSW productionfrom the restaurants and canteens and from the greengroceryparts of the supermarkets is assumed as 20% of this amount,SS-OFMSW amount (XSS-OFMSW) per capita is calculated asabout 50 g/PE·day (TS = 12%; VS/TS = 90%; density ∼=1015 kg/m3). If it is assumed that 20% of dual (separate)sorted (DS) OFMSW originating from households (onlykitchen wastes) is added, XSS-OFMSW amount will increase to


Fine screen

Pumpingstation

Grit removal Pre-sedimentation Anaerobic

Biological N, P removalInternal recycle

Anoxic Aerobic2nd sedimentation Disinfection

Dischar.

WAS recycle

WASPrimary sludge

SS-OFMSWMagnet

Metals

Grinder Pulper

Inerts

Light (floated)Part

Mid-phase

Anaerobicdigestion

Energyrecovery

Biogas

Sludgestorage

Mixer

Flotation thickener

Mechanicaldewatering

Cake

Separatedewatering

Liquid manureFiltrate (except irrigation time)

Figure 1: Process scheme of the municipal WWTP and the integrated biomethanization plant.

about 100 g/PE·day (i.e., 0.25× 0.2 + 0.05 = 0.1 kg/PE·day).The amount of the XSS-OFMSW would increase if DS-OFMSWaddition is more than 20%. In the light of the abovegiven data, the flowrates of raw primary sludge and SS-OFMSW as the influents of the pulper (Figure 1) can becalculated as about 150 and 41 m3 per day, respectively,for the 100,000 PE plant. The effluent of the pulper iscomposed of the supernatant and the bottom phases whichcan be calculated as about 3 and 4 m3 per day, respectively.Therefore, the flowrate of the mid-phase (Figure 1) of thepulper (which is the influent of the anaerobic codigesterwhen primary sludge is given to the effluent stream of thegrinder) is about 184 m3/day with a TS content of about5.5%. The results of the above calculations are summarizedand explained in Table 1.

3. Biomethane Recovery asa Potential Energy Source

Since daily methane production is calculated as about2220 m3 (Table 1), total annual savings based on electric-ity and heat energies can be determined as about C263,350using (1) as follows:

Melectricity = 2220 m3/d× 0.35× 10 kWh/m3 CH4

× C0.07/kWh× 365 d/yr ≈ C198, 525/year,

Mheat = 2220 m3/d× 0.40× 10 kWh/m3 CH4

× C0.02/kWh× 365 d/yr ≈ C64, 825/year.(1)

However, since it is assumed that 85% of this energy can besold in the market, net energy income is expected to beca. C223,850/yr. Here, energy potential per m3 of methaneis approximately 33,810 kJ which corresponds to about10 kWh. Unit energy selling prices are assumed accordingto Turkey’s current market conditions, that is, C0.07 for

electricity and C0.02 for heat per kWh, respectively. If theabove calculations are repeated according to XSS-OFMSW =100 g/PE·day (XSS-OFMSW = 10 t/day for PE = 100,000), dailymethane production is calculated as about 2980 m3 whichcorresponds to net energy income of about C300,500. Hence,results indicated that energy savings that can be recoveredannually from the proposed biomethanization plant wouldbe in the range of C2.24–3.00 per capita by the codigestion ofthe primary sludge and SS-OFMSW. This range may increaseto C4.48–6.00/PE·year if 100% renewable energy subsidy isimplemented. It was observed that over the past few years, thenumber of biogas facilities has been continuously rising inEurope, especially after implementing even higher subsidies.For example, in Germany about 1500 biogas facilities werein use. From these 1500 installed facilities, about 720 MWelectric power production is reported [1].

4. Total Cost of the Biomethanization Plant

Pretreatment of the feedstock is generally essential toenhance their digestibility and bioenergy generation poten-tial because it was reported that one major challenge to thefeedstocks is their slow digestibility and the rate-limitinghydrolysis step. In order to accelerate digestion, variouspretreatment methods are applied such as mechanical, ther-mal, chemical, or biological which help solubilize particulatematter. Pretreatment would become a standard practice forall high-solids organic wastes and residues in the comingyears [10]. In the scope of this study, a preliminary treatmentunit (PTU) is included for the pretreatment of the SS-OFMSW before the mesophilic anaerobic codigester. Thisunit consists of a magnet by which the metals are separated, agrinder, and a pulper. The primary sludge is not subjected tomagnet and grinder but may be given to the effluent streamof the grinder when needed. In order to calculate total projectcost including investment and operation and maintenance(O&M) costs for the proposed integrated biomethanizationplant, the cost for PTU (magnet, grinder, and pulper)


Table 1: Summary of the mass balance for the proposed integrated biomethanization plant (PE = 100,000; XPS = 61 g TS/PE·day; XSS-OFMSW)= 50 g TS/PE·day) (values in brackets show the case for XSS-OFMSW = 100 gr/PE·day).

Waste streams TS (%) VS/TS (%) Density (kg/m3) Total solids (t/day) Flowrate (m3/day)

Pulper

Inlet (PS) 4.0 80 1010 6 150 (150)

Inlet (SS-OFMSW) 12 (15) 90 (85) 1015–1020 5 (10) 41 (66)

Supernatant phase 15 — 1015 ∼0.50 (∼1.0) 3(1) (5.5)

Bottom phase 12 — 1010 ∼0.50 (∼1.0) 4(2) (8)

Outlet (PS+SS-OFMSW) 5.5(3) (7) 90 (85) — 10(4) (14.2) 184(5) (202.5)

Codigester

Inlet (PS + SS-OFMSW) 5.5(3) (7) 90 (85) — 10(4) (14.2) 184(5) (202.5)

Inert solids — 10 (15) — 1.0 (2.13) —

Excess sludge (Px) — — — 0.34(6,7) (0.45) —

Outlet (digested sludge) — 83(8) (75) — 5.84(8) (8.615) 4.84(7,8) (6.485) —

Solids converted into CH4 — — — 4.16(7,9) (5.585) ∼2220(10) (∼2980)(1)

If 10% of VS of OFMSW is wasted from the pulper with supernatant, then 0.10 ×[(41 × 103× 0.12 × 0.90)/(0.15× 1015)] = 3.(2)If 10% of inorganic materials in OFMSW is wasted from the bottom of the pulper, then 0.10 ×[(41 × 103× 0.12)/(0.12× 1010)] = 4.(3)0.055 = [(150 × 0.04 + 41 × 0.12) − (3 × 0.15 + 4 × 0.12)/184].(4)10 = (184× 0.055).(5)184 = [150 + 41−(3 + 4)].(6)If VS/TS = 90%, = 0.05, VS Removal Yobs = 50%, and 1 g VS = 1.5 g COD, then Px = [(10 × 0.90) × 0.50 × 1.5 × 0.05] = 0.34.(7)As volatile solids.(8)5.84 = [(10 × 0.10) + 9 × (1−0.50) + 0.34]; 4.84 = (9 × 0.50 + 0.34); 4.84/5.84 = 0.83.(9)4.16 = [(10 × 0.90)−4.84].(10)Net CH4 recovery = 90%; 1 g VS = 1.5 g COD; CH4 Production/1 kg CODdest. = 0.395 L (at 35◦C), then QCH4 = (4160× 0.90 × 1.5 × 0.395) ∼= 2220.

should be determined. For this purpose, primary compo-nents of the investment cost which were already determinedfor a biomethanization plant treating OFMSW and theprimary sludge for a PE = 200,000 capacity were takeninto consideration [12]. According to the study, the primaryinvestment cost components were construction includingpreconstruction (engineering and consultancy) and civilworks as ∼ C2.5× 106, equipment (mechanical, electrical,instrumentation, piping, and mounting) as ∼ C1.65× 106,and thus giving a total investment cost of about C4.15× 106.On the other hand, annual O&M cost without PTU wascalculated ca. C300,000/year [12]. If investment cost for PTUis assumed as 25% of total investment cost (for PE = 200,000plant) and if it is assumed that O&M cost (for PE = 200,000plant) would increase by 20% by the addition of PTU, annualtotal cost is calculated as C4.82/PE·yr for the proposedbiomethanization plant (PE = 100,000). Cost estimationresults are summarized in Table 2, and change in annualcost values for the plants serving 100,000–1,000,000 PE ispresented in Figure 2. Consequently, when compared withthe annual energy savings (C4.48/PE·yr), almost 93% ofannual total cost (C4.82/PE·year) can be recovered fromthe proposed integrated biomethanization plant (for PE =100,000; XPS = 61 g/PE·day; XSS-OFMSW = 50 g/PE·day) in thecase of implementing 100% renewable energy subsidy with apayback period of 13.2 years.

Unfortunately, integrated biomethanization plants wouldnot be much feasible without public subsidy at the currentsituation. Subsidies have undeniable effect on the improve-ment of renewable energy production. It is considered thatenergy subsidies not only cover operating costs of companiesbut also encourage access to the modern energy sources and

6

5

4

3

2

1

0100000 500000 1000000

Population equivalent (PE)

InvestmentO and MTotal

R2 = 0,9563y = −0,567 ln(x)+ 11,477

y = −0,335 ln(x)+ 6,5903R2 = 0,9413

y = −0,232 ln(x)+ 4,8865R2 = 0,9743

An

nu

alco

st(E

uro

/PE·y

ear)

Figure 2: Change in annual cost values for 100,000–1,000,000 PE.

help to reduce energy import dependency especially in devel-oping countries such as Turkey. The gate fee (or tip-ping fee)for MSW disposal also has significant effect on the decreaseof the payback period although no subsidy is implementedfor energy. Municipalities currently pay approximately 15Euro/tonne landfill gate fee for the MSW and municipalsludge in Turkey. However, this fee needs to be increased inorder to meet EU limitations. Comparative costs of landfillin different EU Member States have indicated that gatefee values might fluctuate in the range of 6–220 Euro pertonne of waste excluding tax [13]. Regarding the proposedbiomethanization plant, if approximately 40 Euro gate fee isassumed per tonne of waste, the payback period is calculatedto be 3.2 years in the case of only gate fee implementation.


Table 2: Summary of total cost estimated for the proposed bio-methanization plant for 100,000 PE (adapted from Ozturk et al.[12]).

Cost components Unit Value

Investment

Construction works C (×106) 1.78(1,2)

Equipment C (×106) 1.33(3,4)

Total investment C (×106) 3.11(5)

Total investment C/PE 31(6)

Annual investment C/yr (×106) 0.266(7)

Annual investment C/PE·yr 2.66(8)

O&M

Annual O&M C/yr (×106) 0.216(9)

Annual O&M C/PE·yr 2.16(10)

Total

Annual total C/PE·yr 4.82(11)

(1)If investment cost of PTU is 25% of total investment cost determined

for PE = 200,000 and if 45% of total investment cost is assumed as theconstruction work, then (2.5× 106 + 0.25× 4.15× 106× 0.45) = 2.97× 106.(2)Since the values were determined for PE = 200,000, and values werecorrected for PE = 100,000 by 60%, then 0.60× 2.97× 106 = 1.78× 106.(3)If investment cost of PTU is 25% of total investment cost determinedfor PE = 200,000 and if 55% of the total investment cost is assumed as theequipment work, then (1.65× 106 + 0.25× 4.15× 106× 0.55) = 2.22× 106.(4)Since the values were determined for PE = 200,000, and values werecorrected for PE = 100,000 by 60%, then 0.60× 2.22× 106 = 1.33× 106.(5)3.11× 106 = (1.78 + 1.33) × 106.(6)For PE = 100,000, (3.11× 106÷ 100000) ∼= 31.(7)If interest + depreciation + amortization, lifetime, and CRF equal 6%, 30years, and 0.07265 for preconstruction and 6%, 15 years, and 0.10296 forequipment works, then 1.78× 106× 0.07265 + 1.33× 106× 0.10296= 0.266.(8)For PE = 100,000, (0.266× 106÷ 100000) = 2.66.(9)O&M cost for PE = 200,000 would increase by 20% with PTU,and if values were corrected for PE = 100,000 by 60%, then [0.60×(300,000× 1.2)] = 216,000 = 0.216× 106.(10)For PE = 100,000, (0.216× 106÷ 100000) = 2.16.(11)4.82 = 2.66 + 2.16.

However, if public subsidy (as 100%) is also implementedtogether with this gate fee, the payback period will shortento 2.6 years indicating the positive effects of the incentives onsystem feasibility.

5. Utilization of Biogas as Transport Fuel

There are different ways of using the produced biogas fromthe biomethanization plants. These are (i) direct steamgeneration, (ii) direct combustion, for example, by internalcombustion engines (ICE), (iii) simultaneous electricity anduseful heat generation, that is, cogeneration at combinedheat and power (CHP) plants, (iv) pumping into centralnatural gas grid after pretreatment, and (v) usage forpowering vehicles after pretreatment. Typical fuel value isreported to be 22,400 kJ per 1 m3 normal biogas. Biogascan be used either for the production of heat only or forthe generation of electric power. When current is obtained,normally heat is produced in parallel by the combinedheat and power (CHP) generation plants. Here, biogasis converted to electricity using on-site power generation

equipment at which heat is recovered in the form of hot wateror steam for heating applications [1]. Although boilers andother direct combustion applications are by far the cheapestand easiest use options for energy utilization, CHP is oneof the most common forms of energy recycling. Turkey hasa wide-spread natural gas grid typically used as domesticand industrial power sources, and biogas generated fromanaerobic digestion of wastes can also be distributed viathis network for community use. However, it was reportedthat using biogas to power vehicles had the lowest carbonfootprint, followed by the use of biogas on-site in a CHPplant. Pumping gas into the grid was the next most efficient[14]. Due to increased use of fossil fuels, atmospheric carbondioxide which hastens the global warming crisis has alsoincreased [2]. The EU Directive on the promotion of theuse of energy from renewable sources sets ambitious targetsfor all Member States, such that the EU will reach a 20%share of energy from renewable sources by 2020 and a10% share of renewable energy specifically in the transportsector [15]. Besides, key goals of the “An EU Strategy forBiofuels” is to promote the use of biofuel in transport withthe aim of reducing the environmental impact of fossil fuelsto define a minimum share of biofuel to be sold on themarket of each Member State by 2010 [16]. Thus, use ofbiogas produced from the wastes as the fuel (gasoline and/ordiesel) for transport has increased extensively in recent years.By this way, significant decreases in air emissions from thevehicles and noise pollution would be achieved, dependencyon other countries to supply petroleum would cease, biogaswastage at the treatment plants via gas flares would beprevented, and a new income source as well as economicbenefit would be provided. It was reported that biogas hasbeen used as vehicle fuel in around 14 cities in Europe,among them the leading two countries are Sweden andSwitzerland. For example, in Stockholm, the annual biogasproduction of about 1.5× 106 m3 could power about 425automobiles and 3 trucks, which was obtained from themunicipal WWTP serving 600,000 PE [4, 17, 18]. However,some modifications are required on the vehicle and enginetechnologies. For example, gasoline and diesel vehicles needto be retrofitted for natural gas unless new vehicles areused. Biofuels need to be processed to consistent standardsfor optimal performance in ICE. Optimization of vehiclesincludes fine-tuning control systems and engine designs torun on varying blends for maximum fuel efficiency andminimum emissions across the full range of potential blendmixes. Since the Member States of EU must ensure that theminimum share of biofuels sold on their markets is 2% by 31December 2005 at the latest, and 5.75% by December 2010,most of the automobile industries guarantee its usage safely[16, 19]. Biogas should also comply some quality standardsin order to be used as the fuel for transport purposes. Notonly does biogas consist mainly of methane and CO2, butit also contains several impurities [1]. It was reported thatbiogas has to be upgraded to natural gas quality for use invehicles designed to function on natural gas. This calls forremoval of particulates, carbon dioxide, hydrogen sulfide,moisture, and any other contaminants present in the gas toincrease the methane content to over 95% (by volume). The


Table 3: Transport fuel substitutes of the produced biomethane (PE= 100,000; XPS = 61 g TS/PE·day; XSS-OFMSW) = 50 g TS/PE·day) (adaptedfrom Murphy et al. [21]).

Statement Value Unit

Replacement of diesel in heavy good vehicles (operating at 6 mpg on diesel 0.052 km/MJ)

Heavy good vehicle operating on CH4-enriched biogas, 90% efficiency of diesel 0.047 km/MJ

Potential travel distance powered by biogas: (2340 m3/day × 330(1) day/year × 35.9(2)

MJ/m3× 0,047 km/MJ)1,302,933 km/yr

Number of heavy good vehicles powered (50,000 km per year) 26 —

Diesel substituted of 1 m3 enriched biogas: [2340 × 330 × 35.9 × (0.047/0.052)]÷ (40.7(3,4)) 615,636 (∼0.8)(5) L/yr (L/m3)

Replacement of gasoline in cars (operating at 40 mpg on gasoline 0.439 km/MJ)

Car operating on CH4-enriched biogas, 90% efficiency of gasoline 0.396 km/MJ

Potential travel distance powered by biogas: (2340 m3/day × 330(1) day/year × 35.9(2)

MJ/m3× 0.396 km/MJ)10,977,904 km/yr

Number of cars powered (20,000 km per year) 549 —

Gasoline substituted of 1 m3 enriched biogas: [2340 × 330 × 35.9 × (0.396/0.439)]÷(32.23(3,4))

775,880 (∼1.0)(5) L/yr (L/m3)

(1)Annual operation period of 330 days is assumed.

(2)Energy value of 1 m3 enriched biogas (95% CH4) is 35.9 MJ.(3)Energy value of oil 47.89 GJ/T; densities of diesel and gasoline are 850 and 673 kg/m3, respectively.(4)Energy values of diesel and gasoline are 40.7 MJ/L (47.89 × 0.850) and 32.23 MJ/L (47.89× 0.673), respectively.(5)Since annual CH4-enriched biogas production is 2340 m3/d× 330 d/yr∼= 772,200 m3/yr, (615,636÷ 772,200)∼= 0.8 L/m3 for diesel and (775,880÷ 772,200)∼= 1.0 L/m3 for gasoline.

carbon dioxide present in biogas dilutes the fuel value of thegas and has to be removed. Hydrogen sulfide can increasecorrosion in the presence of water and moisture can causeclogging at lower temperatures. For this purpose, the biogasshould be purified by a suitable method, for example, bywater scrubbing. [4]. Then this purified biogas can be storedunder 200–250 bar pressure at biogas stations and can be soldafterwards. Since clean biogas has the same characteristicsas natural gas, the existing natural gas infrastructure cansupport the use of biogas. Generally biogas is mixed withnatural gas, and blending with natural gas can also enhancethe distribution of biogas even at relatively small quantities.Moreover, CO2, NOx, and particulate emissions are reportedat much lower rates compared to other fuel types; for exam-ple, CO, hydrocarbon, NOx, CO2, and particulate emissionsfrom heavy good vehicles powered by biogas are almost 2.5-,1.14-, 0.6-, 5-, and 7-folds lower than the heavy good vehiclespowered by diesel, respectively. It is also reported thatbenzene (which is a known human carcinogen) emissionsare eliminated [4]. Tilche and Galatola [20] reported thatthe use of biomethane, in particular as biofuel, has beenshown to be able to produce large greenhouse gas savings.Methane amounts for the proposed integrated biometh-anization plant (at 35◦C and 1 atm) were calculated as2220 m3/day (XSS-OFMSW= 50 g TS/PE·day) and 2980 m3/day(XSS-OFMSW = 100 g TS/PE·day). CH4-enriched biogas (95%CH4) equivalents of the above values can be calculatedas 2340 and 3140 m3/day, respectively. Since 1 m3 enrichedbiogas replaces about 0.8 L of diesel or 1.0 L of gasolineaccording to Table 3, annual amounts in diesel or gasolinesubstitutes are about 615,636 or 775,880 L, respectively, for2340 m3/day CH4-enriched biogas. Moreover, results showedthat about 26 or 549 (XSS-OFMSW = 50 g TS/PE·day) and 35or 737 (XSS-OFMSW=100 g TS/PE·day) heavy good vehicles

and cars could be powered by the produced biogas from theproposed integrated biomethanization plant (PE = 100,000)instead of diesel or gasoline, respectively. The productioncost and the selling price of 1 m3 biogas are reportedin the ranges of C0.2–0.75 and C0.66–1.1, respectively[17]. Transport fuel susbstituted values determined forthe proposed biomethanization plant are summarized inTable 3.

6. Conclusion

The significance of renewable alternative fuels has increasedin the last decade due to dwindling supplies of conventionalenergy sources and rising oil prices. Since natural gas andelectricity costs have been increasing day by day, the shareof renewable energy in total global energy consumptionincreases and energy recovery from biogas becomes attrac-tive. Biogas is reported as one of the most importantrenewable energy sources, and biogas production is mainlybased on anaerobic digestion of organic wastes. Turkey’s oilimport bills likely to grow and put increasing pressure on theeconomy. Turkey has the highest gasoline and diesel prices inthe world, and there is a high motivation and public accep-tance in alternative and relatively cheaper fuels. Only about0.2% of current energy power in Turkey is obtained frombiogas and solid wastes. Since the current energy productionis heavily dependant on the imported natural gas and coalin Turkey, renewable energy recovery by anaerobic digestionof organic wastes (biomethanization) has been gaining vitalimportance in recent years. Towards the EU membershipof Turkey, waste minimization and recovery strategy willbe based on this approach especially in metropolitan cities.Codigestion of OFMSW with other cosubstrates is reportedas an attractive alternative for sustainable management of


different separate waste streams produced at large amounts.Although, the most appropriate cosubstrate alternatives forcodigestion with OFMSW are reported as the sewage sludge,animal slurries, garden wastes, fruit wastes, and so forth,they may be also used as alternative feedstocks. In thisstudy, an integrated biomethanization plant is proposed atwhich primary sludge is codigested with SS-OFMSW (mixedwastes originating from the restaurants and canteens andfrom the greengrocery parts of the supermarkets). Resultsof this study indicated that almost 93% of annual totalcost can be recovered if 100% renewable energy subsidy isimplemented with a payback period of 13.2 years. However,if approximately 40 Euro gate fee is also implemented pertonne of waste with public subsidy, the payback period wouldshorten to 2.6 years indicating the positive effects of theincentives on system feasibility. Since landfilling is still themain solid waste disposal method in Turkey, other benefits ofthe proposed biomethanization plant would be the reductionof landfill volume requirement and the protection of thegroundwater.

In accordance with the EU, the authoritatives in Turkeyare aware of that serious precautions should be taken interms of sustainable environmental protection such as usageof alternative fuels for transport. For example, urban airpollution is often a key driver of alternative fuels fortransport in developing countries due to its magnitude andvisibility and the degree of public exposure to the problem.In this scope, if transport fuel substitute of the producedbiogas is calculated, about 26 heavy good vehicles using dieselor 549 cars using gasoline may be powered per year (PE =100,000; XPS = 61 g TS/PE·day; XSS-OFMSW=50 g TS/PE·day).The number of heavy good vehicles or cars might increaseup to 35 or 737 in the case of XSS-OFMSW= 100 g TS/PE·day.However, the biogas has to be upgraded for use in vehicles,and some modifications are required on the vehicle andengine technologies.

It is known for certain that number of such proposedbiogas plants will increase in Turkey following the gov-ernment incentive to promote their installations. How-ever, the related regulations regarding energy productivity,environment, Kyoto agreements, clean ecoproduction, andrenewable energy should be immediately completed with thesupply of the required infrastructure for investment.

Acknowledgment

The authors gratefully acknowledge The Scientific and Tech-nological Research Council of Turkey (TUBITAK) (Projectno. 105G024) for the financial support.

References

[1] D. Deublein and A. Steinhauser, Biogas from Waste and Renew-able Resources: An Introduction, Wiley-VCH, Weinheim, Ger-many, 2008.

[2] S. K. Khanal, “Bioenergy generation from residues of biofuelindustries,” in Anaerobic Biotechnology for Bioenergy Pro-duction: Principles and Applications, S. K. Khanal, Ed.,

pp. 161–188, Wiley-Blackwell, Oxford, UK; John Wiley &Sons, New York, NY, USA, 2008.

[3] H. Hartmann, I. Angelidaki, and B. K. Ahring, “Co-digestionof the organic fraction of municipal waste with other wastetypes,” in Biomethanization of the Organic Fraction of Munic-ipal Solid Wastes, J. Mata-Alvarez, Ed., pp. 181–199, IWA,London, UK, 2002.

[4] S. Harikishan, “Biogas processing and utilization as an energysource,” in Anaerobic Biotechnology for Bioenergy Production:Principles and Applications, S. K. Khanal, Ed., pp. 267–291,Wiley-Blackwell, Oxford, UK; John Wiley & Sons, New York,NY, USA, 2008.

[5] I. Ozturk, A. Bir, A. F. Aydin et al., “Investigation of bioenergyrecovery technologies (biomethanisation) by integrated treat-ment of municipal wastewaters and organic fraction of solidwastes,” Final Report 105G024, The Scientific and TechnicalResearch Council of Turkey (TUBITAK), 2009.

[6] R. K. Dereli, M. E. Ersahin, C. Y. Gomec, I. Ozturk, and O.Ozdemir, “Co-digestion of the organic fraction of municipalsolid waste with primary sludge at a municipal wastewatertreatment plant in Turkey,” Waste Management and Research,vol. 28, no. 5, pp. 404–410, 2010.

[7] EU Council Directive, EU Council Directive on the Landfill ofWaste, 99/31/EC, 1999.

[8] M. Balat and H. Balat, “Biogas as a renewable energy sourceareview,” Energy Sources A, vol. 31, no. 14, pp. 1280–1293, 2009.

[9] F. G. Pohland, A. B. Al-Yousfi, and D. R. Reinhard, “Anaerobicdigestion of organic solid waste in bioreactor landfills,” inBiomethanization of the Organic Fraction of Municipal SolidWastes, J. Mata-Alvarez, Ed., pp. 303–315, IWA, London, UK,2002.

[10] S. Harikishan, “Pretreatment of high-solids wastes/residues toenhance bioenergy recovery,” in Anaerobic Biotechnology forBioenergy Production: Principles and Applications, S. K. Khanal,Ed., pp. 247–265, Wiley-Blackwell, Oxford, UK; John Wiley &Sons, New York, NY, USA, 2008.

[11] L. Spinosa and P. A. Vesilind, Sludge into Biosolids Processing,Disposal, Utilization, IWA, London, UK, 2001.

[12] I. Ozturk, R. K. Dereli, A. Ozabali, K. Ericyel, and I. Karakaya,“The feasibility report of Central Biomethanization Plantof Stock Farmer Union in Suluova (Amasya) at the StockOrganised Industrial District,” Istanbul, 2009.

[13] D. Hogg, “Costs for municipal waste management in the EU,”Final Report, Directorate General Environment, EuropeanCommission, Eunomia Research & Consulting, Bristol, UK,2001.

[14] Letsrecycle.com, “Research raises questions over best use ofbiogas,” London, UK, 2010, http://www.letsrecycle.com/do/ecco.py/view item?listid=37&listcatid=5477&listitemid=5634&section=composting.

[15] EU Council Directive, Directive 2009/28/EC of The EuropeanParliament and of The Council of 23 April 2009 on thepromotion of the use of energy from renewable sources andamending and subsequently repealing Directives 2001/77/ECand 2003/30/EC, 2009/28/EC, 2009.

[16] An EU Strategy for Biofuels {SEC(2006)142}, COM(2006) 34final, 2006.

[17] G. Landahl and C. Plombin, “Trendsetter: biogas in Europeanvehicles,” in Resource Recovery and Reuse in Organic SolidWaste Management, P. Lens and B. Hamelers, Eds., pp. 411–422, IWA, London, UK, 2004.

[18] G. Boyle, “An overview of alternative transport fuels indeveloping countries: drivers, status, and factors influencing


market deployment,” in Hydrogen Fuel Cells and Alternativesin the Transport Sector: Issues for Developing Countries, UnitedNations University International Conference, November 2005.

[19] “Biofuels for Transportation Global Potential and Implica-tions for Sustainable Agriculture and Energy in the 21stCentury,” Worldwatch Institute, Washington, DC, USA, 2006,http://www.worldwatch.org/system/files/EBF038.pdf.

[20] A. Tilche and M. Galatola, “The potential of bio-methane asbio-fuel/bio-energy for reducing greenhouse gas emissions: aqualitative assessment for Europe in a life cycle perspective,”Water Science and Technology, vol. 57, no. 11, pp. 1683–1692,2008.

[21] J. D. Murphy, E. McKeogh, and G. Kiely, “Technical/eco-nomic/environmental analysis of biogas utilisation,” AppliedEnergy, vol. 77, no. 4, pp. 407–427, 2004.


Research Article

In Situ Biodiesel Production from Fast-Growing and High OilContent Chlorella pyrenoidosa in Rice Straw Hydrolysate

Penglin Li,1, 2 Xiaoling Miao,1, 2 Rongxiu Li,1, 2 and Jianjiang Zhong1, 2

1 Key Laboratory of MOE for Microbial Metabolism and School of Life Science & Biotechnology, Shanghai Jiao Tong University,Shanghai 200240, China

2 Biomass Energy Research Center, Shanghai Jiao Tong University, Shanghai 200240, China

Correspondence should be addressed to Xiaoling Miao, [email protected]

Received 14 October 2010; Accepted 22 December 2010

Academic Editor: Rodomiro Ortiz

Copyright © 2011 Penglin Li et al. This is an open access article distributed under the Creative Commons Attribution License,which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Rice straw hydrolysate was used as lignocellulose-based carbon source for Chlorella pyrenoidosa cultivation and the feasibilityof in situ biodiesel production was investigated. 13.7 g/L sugar was obtained by enzymatic hydrolyzation of rice straw. Chlorellapyrenoidosa showed a rapid growth in the rice straw hydrolysate medium, the maximum biomass concentration of 2.83 g/L wasobtained in only 48 hours. The lipid content of the cells reached as high as 56.3%. In situ transesterification was performed forbiodiesel production. The optimized condition was 1 g algal powder, 6 mL n-hexane, and 4 mL methanol with 0.5 M sulfuric acidat the temperature of 90◦C in 2-hour reaction time, under which over 99% methyl ester content and about 95% biodiesel yieldwere obtained. The results suggested that the method has great potential in the production of biofuels with lignocellulose as analternative carbon source for microalgae cultivation.

1. Introduction

As a renewable, biodegradable, and environmentally friendlyfuel, biodiesel has received more attention in the past fewyears. Biodiesel refers to fatty acids methyl esters (FAMEs)from vegetable oils or animal fats with high stability; lowwater and volatiles content; a low amount of polymerscontaining sulfur and nitrogen elements. Current sources ofcommercial biodiesel are primarily soybean oil, rapeseed oil,palm oil, corn oil, waste cooking oil, and animal fat, butthese sources of biodiesel cannot meet even a slight fractionof present demand for fuels [1]. However, a major criticismagainst large-scale fuel production from agricultural crops, isthat it will consume vast swaths of farmland, native habitatsand drive up food prices [2]. For example, meeting only halfthe existing US transport fuel by biodiesel would requireunsustainably 54% and 24% of the US cropping land usingcoconut and oil palm, respectively [1].

Microalgae have been considered as one of the mostpromising feedstocks for biodiesel production due to theirshort cell cycle (within 24 hours), high oil content (20%–50% normally and highest exceeding 80%), strong adaptive

capacity to environment (high salinity, heavy metal ion,toxicants, high CO2 concentration, etc.) and no occupationfor cropping area [1]. However, price of algae-based biodieselis still much higher than normal diesel with the priceof US$1.25/lb and US$0.43/lb, respectively, [3, 4], whichencumbers large-scale manufacturing and applications ofalgae-based biodiesel. Some attempts have been made toreduce the cost such as using cheap carbon sources forhigh oil microalgae production. For example, Jerusalemartichoke [5], cassava starch [6], and sugar cane juice [7]were introduced as new carbon supplies for microalgaecultivation and algae-based biodiesel production. The resultsshowed that hydrolysate medium could provide an exceeding15.2% to 42.3% biomass production leading to an 8.8% to27.7% increase in algal oil production based on the carbonsource. Then good quality biodiesel, whose FAME contentreached as high as 93%, was produced from algal biomass[8]. These studies suggested that microalgae could be ableto utilize such resources for high oil accumulation andbetter biodiesel production. However, such materials, mainlyagricultural crops, have been also studied for bioethanolproduction for years [9, 10]. The conflict between food and


fuel still exists either for biodiesel or bioethanol. For furtherfeasible study and sustainability, lignocellulose material hasaroused people’s interest for recent years [11]. Since theworld’s annual cereal production has been over 2 billiontons for years [12], straw, making up of 50% weight ofcereal crops, is one of the largest lignocellulose resourcesas an agricultural byproduct. Additionally, biodegradablesolid waste fraction from municipal refuse provides anotheralternative large source for discarded lignocellulose dueto target for minimizing landfill use in many countries[13]. Based on these sources, microalgae cultivation couldbe coupled with wastes recovery, management, and reuse.Therefore biofuels derived from lignocellulose, especiallystraw and other nonfood feedstock offer a better option forfood, energy, and environmental concerns [2].

The current algae-based biodiesel is mainly producedby conventional route: extraction of the lipids from themicroalgal biomass followed by its conversion to FAMEs andglycerol [14–16]. However, such method is time consuming,costly, and difficult to be implemented in algae’s crushingstep because of the rigid cell walls [17]. Recently in situtransesterification method, in which the oil-bearing materialcontacts with alcohol directly instead of reacting with pre-extracted oil, has received serious attention for algae-basedbiodiesel production. As a method for biodiesel production,not only FAME content but also conversion efficiency fromalgal oil to biodiesel are imperative as quality and quantitystandards in the process of transesterification [18]. However,current papers provide limited information about bothquality and quantity studies of in situ transesterificationfrom microalgal biomass. Johnson and Wen [17] reported a63.5% of the FAME content using direct transesterificationof Schizochytrium limacinum biomass with 100% biodieselyield; Ehimen et al. [19] obtained a high-quality biodieselwith almost 93% FAME content from Chlorella strains, butno data was provided on yield from algal oil to biodiesel.Therefore, further research is needed to develop a feasiblemethod for high yield and high quality in situ biodieselproduction from microalgae.

In this paper, we studied the feasibility of rice straw asthe raw material of carbon source for Chlorella pyrenoidosa’scultivation and the effects of solvent, methanol volume,temperature and reaction time on FAME content andbiodiesel yield were also investigated in the process of in situtransesterification from algal biomass.

2. Methods

2.1. Preparation of Cellulose Enzymatic Hydrolysate. Asreported by many papers, the barrier to production andrecovery of valuable materials from lignocellulose was itsresistance to degradation due to cross-linking between thepolysaccharides (cellulose and hemicellulose) and the ligninvia ester and ether linkages [20]. To utilize lignocelluloseas a raw material, a pretreatment is needed [21]. Ricestraw purchased from local market was selected as thelignocellulose material for hydrolysation as rice is one ofthree largest grain types produced in China. The strawwas previously cut into 10-cm pieces and then pretreated

in a two-step method according to our published paper[22]. First, 50 g dried rice straw was mixed with 1L 1%trifluoroacetate at 95◦C for 5 hours. Then the samplesafter desiccation filled in a steel reaction kettle reacted with60% alcohol at 210◦C for 4 hours. After pretreatment,a commercially available cellulase, Accellerase 1000 fromGenencor, was used for rice straw hydrolyzation. Accordingto our pre-experiment and the enzyme protocol [23], 20 g/Lpretreated rice straw and 20 mL/L enzyme were mixed withwater (pH 5.0) in an Erlenmeyer flask. The reaction was doneon the shaker (150 rpm) for 24 hours at a temperature of50◦C. Then the enzyme was deactivated (heating) and thesolution was filtered to obtain rice straw hydrolysate (RSH).

2.2. Algae Strain and Cultivation Condition. The greenmicroalga Chlorella pyrenoidosa was purchased from thealgae strain laboratory of Aquatic Organism Research Insti-tute of Chinese Academy of Science. A modified BG11medium was used as the basic medium which contains100 mg NaNO3, 74.9 mg MgSO4·7H2O, 30.5 mg KH2PO4,27.18 mg CaCl2, 20 mg Na2CO3, 8.9 mg C6H5O7Na3·2H2O,6 mg ferric ammonium citrate, 1.04 mg Na2·EDTA, 2.86 mgboric acid, 0.222 mg ZnSO4·7H2O, 0.079 mg CuSO4·5H2O,1.81 mg MnCl2·4H2O, 0.39 mg Na2MoO4, and 0.049 mgCo(NO3)2·6H2O every liter. Glucose and RSH were used ascarbon sources. 10 g/L glucose were selected for cultivation[15] and the same amount of glucose was used in RSHmedium by adding appropriate volumes of RSH solution tothe basic medium to achieve the concentration. The mediumwas adjusted to pH 7.5 and autoclaved; alga (OD600 = 0.5)was inoculated into an Erlenmeyer flask (250 mL, containing50 mL medium) and cultivated on the shaker (110 rpm) ata temperature of 25 ± 1◦C with continuous illumination atintensities of 40 µmol/(m2s).

2.3. In Situ Transesterification for Biodiesel Production. Adirect methanolysis of algal biomass was introduced forin situ biodiesel production due to its higher yield andtime and cost saving. Algal powder, methanol, solvent,and 0.5 M sulfuric acid were reacted in a 20 mL-cylindricaltetrafluoroethylene reaction vessel and there was a steel shelloutside the reactor for seal and high-temperature protection.With 1 g algal powder and 0.5 M sulfuric acid, differentamounts of solvent (2.0 mL to 8.0 mL), methanol (0.5 mL to10.0 mL), temperature (20◦C to 110◦C), and reaction time(0.25 hour to 4 hours) were studied. After the reaction,2.0 mL water was added to the mixture. Samples werethen cooled down to room temperature, and centrifugedat 8,228×g for 10 min. Three layers formed which werecomposed of a water phase for water and alcohol, a solidphase for algae residue, and an organic phase for solventand biodiesel, respectively. The organic layer was collectedand evaporated at 60◦C to constant weight for analysis. Thebiodiesel yield from algal biomass was calculated by (1):

Biodiesel yield (%)

= Biodiesel mass(g)

algae mass(g)× oil content (%)

× 100%.(1)


2.4. Dry Cell Weight and Reducing Sugar Concentration. Forbiomass measurement, 5 mL culture medium was trans-ferred to a preweighed centrifuge tube and centrifuged at4,500×g for 5 min. After rinsing three times, the samplewas lyophilized to constant weight and warmed up to roomtemperature in a desiccator, then the dry cell weight wasdetermined.

The content of reducing sugars in medium was deter-mined by the 3,5-dinitrosalicylic acid method [22]. Thereducing sugar concentration was measured using an ST360biosensor. The sugar components of the hydrolysate wereanalyzed by HPLC. The enzymatic hydrolysis yield wascalculated by

Enzymatic hydrolysis yield (%)

= Reducing sugar(g)× 0.9

Carbohydrate(g) × 100%.

(2)

2.5. Lipid Content. Lipid in the algal cells was extractedaccording to a modified method from Xu et al. [14]. 0.2 gdry algae were triturated in the liquid nitrogen for cellfragmentation. Then the algae were blended with 3 mL chlo-roform/methanol (2 : 1), shaken for 20 min and centrifuged(4,500×g) for 10 min. The supernatant was collected in apre-weighed tube, and the same process was repeated twice.All the supernatant was collected together, evaporated anddried to constant weight at 60◦C, and finally weighed tocalculate the total lipid content.

2.6. Composition Analysis of Biodiesel. The composition ofthe biodiesel produced from in situ transesterification of algalbiomass was analyzed by GC-linked mass spectrometry (GC-MS) equipped with a DB-5MS column (30 m× 0.25 mm IDDF = 0.25 µm) and with a flow rate of 1.0 mL/min.


3.1. Enzymatic Hydrolysis of Rice Straw. To obtain a higherglucose concentration for algae cultivation, time of enzy-matic hydrolysis was modified. 20 g/L pretreated rice strawmixed with 20 mL/L cellulase was hydrolyzed for 24 hoursat 50◦C. According to our pre-experiments, a dosage of1mL enzyme to 1 g straw was essential for fast hydrolyzation(within one day) and other conditions were followed by theprotocol from the enzyme. The time course of hydrolyzationwas shown in Figure 1. The reducing sugar concentrationincreased notably in the first 8 hours, reaching over 12 g/L(Figure 1). Then its concentration remained almost constantbetween 12 to 14 g/L in the following time. The highest sugarconcentration (about 13.7 g/L) appeared in the 20th hour(Figure 1). The enzymatic hydrolysis yield was about 61.7%calculated by (1). This data was close to a 67% yield ofwheat straw hydrolyzation pretreated by an aqueous glycerolmethod at the same temperature [24]. The HPLC analysis ofthe hydrolysate showed that the glucose took up over 98% ofthe total reducing sugar and the rest was mainly xylose. Othercompounds such as acetic acid and glycerol was also detectedin the hydrolysate, but their concentration was very low

0

2

4

6

8

10

12

14

16

0 5 10 15 20 25

Red

uci

ng

suga

r(g

/L)

Time (hours)

Figure 1: Time course of rice straw hydrolyzation. Reactioncondition: 20 g pretreated rice straw and 20 mL enzyme in 1 Lvolume on the shaker (150 rpm) at 50◦C Error bars = mean ±standard deviation.

0

2

4

6

8

10

12

0

0.5

1

1.5

2

2.5

3

0 12 24 36 48 60

Res

idu

alsu

gar

(g/L

)

Bio

mas

sco

nce

ntr

atio

n(g

/L)

Time (hours)

Figure 2: Cell growth and sugar consumption of Chlorellapyrenoidosa in the glucose medium and the rice straw hydrolysate(RSH) medium. Biomass concentration (�) and residual sugarconcentration (•) in the glucose medium; biomass concentration(�

) and residual sugar concentration (©) in the RSH medium.Error bars = mean ± standard deviation.

(<0.05 g/L). This suggests that higher percentage of glucosecould be obtained from rice straw through our presentmethod. For further algae cultivation, the hydrolysate wasprepared in a 20-hour hydrolysis.

3.2. Shake-Flask Cultivation of C. pyrenoidosa with Rice StrawHydrolysate. To improve C. pyrenoidosa’s utilization of ricestraw hydrolysate in the presence of probable inhibitors,algae adaptation had been done in the RSH medium forgenerations. After adaptation, comparison experiments inshake-flask cultivation using glucose medium and RSHmedium were conducted. The effects of glucose and RSHmedium on the growth and sugar consumption wereshown in Figure 2. The algae showed rapid growth in RSHmedium; the maximum biomass concentration of 2.83 g/Lwas obtained in only 48 hours. The lipid content of the cellsreached as high as 56.3% (Table 1). The biomass productivity


Table 1: The effects of different carbon sources on cell growth andlipid accumulation of Chlorella pyrenoidosa.

Carbon source

Glucose Rice straw hydrolysate

Maximum biomassconcentration (g/L)

0.92± 0.01 2.83± 0.04

Lipid content (%, w/w) 50.3± 0.9 56.3± 1.4

Biomass productivity(g/L/day)

0.37± 0.00 1.10± 0.02

Lipid productivity(g/L/day)

0.19± 0.00 0.62± 0.01

and lipid productivity of the cells growing in RSH mediumwere about 3.0 times and 3.3 times higher than that of thecells growing in the glucose medium, respectively (Table 1).This is probably because the C. pyrenoidosa could fully utilizecarbon sources in the RSH medium. Larson and Rees [26]reported that lipid accumulation could be triggered in alow nitrogen medium (0.1 g/L in our medium). Under sucha condition, cell division ceases while respiration does notdecrease, so lipid could be accumulated for energy storageif cells absorb additional carbon sources. Almost all thesugars in the RSH medium had been used up after 48hours, while about two-thirds of the glucose in the glucosemedium was not consumed (Figure 2). The maximumbiomass concentration of the cells growing in the glucosemedium was only 0.92 g/L after 60 hours (Table 1). Anotherpossible reason for these phenomena reported by manypapers [8, 25] was that there were probably some beneficialgrowth factors for cell growth and oil accumulation in thehydrolysate system. Compared with other research, the C.pyrenoidosa growing in RSH medium also showed higherbiomass productivity and higher lipid content (Table 2).As reported, different microalgal biomass concentrationswere obtained at relatively longer culture period in otherresearch such as 3.92 g/L [14], 7.2 g/L [6], 4.26 g/L [25], and5.1 g/L [8] in 6 day, 10 days, 125 hours, and 120 hours,respectively (Table 2). Our present research suggests thathigher biomass and higher lipid content microalgal cellscould be obtained in a shorter culture period (48 hours) byusing RSH medium, which would have a better applicationin industrial production.

3.3. In Situ Transesterification for Biodiesel Production

3.3.1. Effect of Solvent. Generally, methanol plays the rolesof both reactant and extractant in in situ transesterification,but it was proved to be a poor solvent for oil extraction [27].This would probably lead to low conversion efficiency fromalgal biomass to biodiesel by in situ transesterification. Oneof the possible solutions to this problem is to introduce agood oil extraction solvent such as n-hexane or chloroformin the process of in situ biodiesel production. Accordingto our previous experiments, adding these two kinds ofsolvents to the in situ reaction system could lead to a higherbiodiesel yield. Therefore, these two kinds of widely used oilextraction solvent, chloroform, and n-hexane, were selected

Table 2: Comparison of maximum biomass concentration,biomass productivity, and lipid content of Chlorella strains growingin different hydrolysate medium.

Hydrolysatematerial

Maximumbiomass

concentration(g/L)

Biomassproductivity

(g/L/day)

Lipid content(%, w/w)

Rice straw 2.83 1.10 56.3

Cornpowdera 3.92 0.65 55.3

cassavastarchb 7.20 0.72 28.9

Cassavac 4.26 0.82 50.2

Sweetsorghumd 5.10 1.02 53.3

aXu et al. [14]; bWei et al. [6]; cLu et al. [25]; dGao et al. [8].

for further in situ transesterification studies. Reaction wascarried out at 90◦C with 1 g algal powder, 4 mL methanoland 0.5 M sulfuric acid. Solvent volume was set at 2, 4,6, and 8 mL, respectively. The effects of different volumeof chloroform and n-hexane on biodiesel yield and FAMEcontent were shown in Figure 3. All the samples’ FAMEcontent maintained over 98% suggesting that high-qualitybiodiesel could be prepared by our in situ transesterificationmethod. For solvent volume from 2 mL to 6 mL, biodieselproduction increased with the increase of solvent amountfor both groups and approximately 10% higher biodieselyield was obtained in n-hexane group compared with thatin chloroform group. While at solvent volume of 8 mL, chlo-roform showed a better result with a more 5% conversionratio. The highest biodiesel yield was obtained under theconditions of 6.0 mL n-hexane and 8.0 mL chloroform wherethe conversion efficiency from algal biomass to biodiesel was94.3% and 95.7%, respectively. Although highest yield andFAME content were almost equal to each other at 6.0 mL n-hexane, and 8.0 mL chloroform conditions, the colour of thebiodiesel was different. A light yellow sample was made withno solid residue in the n-hexane group. In contrast, therewere some solid residues in the chloroform group where thesamples’ color was brown. As reported by Kulkarni et al. [28]and Goff et al. [29], oxidization of fatty acid and formation ofsulfones or sultones from sulfuric acid are two of the possiblereasons for dark products in biodiesel production. Thatprotein has a better solubility in higher polarity solvent suchas chloroform might be one of the reasons to solid residuein the samples. Dark products could have a smaller cetanenumber representing a lower power take off and efficiency ofbiodiesel and the additional residue in the biodiesel could beharmful for engine’s lifespan. This suggested that in in situtransesterification, nonpolarity solvents should be a betterchoice than polarity ones. Therefore, 6 mL n-hexane waschosen for following experiments considering both biodieselyield and quality levels.

3.3.2. Effect of Methanol Volume. Apart from the positiveeffect of n-hexane on in situ transesterification, the amount


90

91

92

93

94

95

96

97

98

99

100

0

10

20

30

40

50

60

70

80

90

100

110

2 4 6 8

FAM

Eco

nte

nt

(%)

Bio

dies

elyi

eld

(%)

Solvent volume (mL)

Figure 3: Effects of chloroform and n-hexane volume on thebiodiesel yield and FAME content by in situ transesterification ofC. pyrenoidosa. Reaction condition: 1 g algal powder with 4 mLmethanol and 0.5 M sulfuric acid at 90◦C. Solvent volume: 2, 4,6, and 8 mL. Biodiesel yield (�) and FAME content (©) in thechloroform group, biodiesel yield (�) and FAME content (�) inthe n-hexane group. Error bars = mean ± standard deviation.

of methanol is one of the most important factor affectingquality and quantity of biodiesel production. As supportedby many papers [18, 30], an exceeding amount of methanolis essential for in situ biodiesel production. In order to knowthe appropriate volume for algae’s in situ transesterification,a series of methanol volume (0.5, 1, 2, 3, 4, 6, 8, 10 mL) wasselected for biodiesel production with 1 g algal powder, 6 mLn-hexane and 0.5 M sulfuric acid at 90◦C. The methanolvolume from 0.5 to 10 mL represented a reacting alcohol tooil molar ratio from about 21 : 1 to 413 : 1 (calculated witha methanol density of 0.792 g/mL). The results were shownin Figure 4. Biodiesel yield was enhanced from 69.4% to94.3% when increasing methanol volume from 0.5 mL to4 mL and the FAME content remained at over 98% level.For further increase of methanol amount, there was nosignificant difference on the biodiesel yield (about 95%) andFAME content (>98%). Moreover, the excess of methanolquantities also slows down the separation process [15].Therefore, 4 mL methanol was suggested to be a suitablevolume.

Miao and Wu [15] reported a 63% biodiesel yieldfrom algal oil transesterification using an optimized 56 : 1methanol to oil molar ratio. In our experiments, whenmethanol to oil molar ratio was 41 : 1 (1.0 mL methanol to1 g algae), about 78.4% biodiesel yield from algal biomasswas obtained, which was 24% higher than a 63% biodieselproduction. Such situation could be attributed to introduc-tion of n-hexane as an effective solvent in the reaction. Thehighest biodiesel yield of 94.7% was obtained under theoptimized methanol to oil molar ratio of about 165 : 1 (4 mLmethanol to 1 g algae). Adequate amount of methanol isrequired for high yield of in situ biodiesel production because

92

93

94

95

96

97

98

99

100

0

10

20

30

40

50

60

70

80

90

100

0.5 1 2 3 4 6 8 10

FAM

Eco

nte

nt

(%)

Bio

dies

elyi

eld

(%)

Methanol volume (mL)

Figure 4: Effects of methanol volume on the biodiesel yield (�) andFAME content (�) by in situ transesterification of C. pyrenoidosa.Reaction condition: 1 g algal powder with 6 mL n-hexane and 0.5 Msulfuric acid at 90◦C. Methanol volume: 0.5, 1, 2, 3, 4, 6, 8, and10 mL. Error bars = mean ± standard deviation.

methanol plays the role of both reactant and substance tosubmerge algal powder [19, 31]. These results suggestedthat in situ transesterification for biodiesel production frommicroalgal biomass could not only save the time and cost, butalso have high conversion efficiency.

For the comparison of in situ transesterification ofalgal biomass, Johnson and Wen [17] reported an about100% yield crude biodiesel using a proportion of 3.4 mLmethanol/g algae from Schizochytrium limacinum cells.However, the FAME content of their biodiesel was only63.5%, while it was about 98% in our samples under a 4 mLmethanol condition. Ehimen et al. [19] prepared the algae-based biodiesel from Chlorella strains that contained 93%FAME. However, the optimized methanol to oil molar ratiothey used was about two times as high as that of our results(315 : 1 compared with 165 : 1). This was probably due tothe introduction of solvent in our in situ transesterificationsystem. The addition of n-hexane could help oil extractionso that half of the methanol could be saved. The resultssuggested that 4 mL methanol to 1 g algal powder was areasonable ratio for in situ biodiesel production, but forspecific strains, the suitable reaction condition should bemodified.

3.3.3. Effect of Temperature and Reaction Time. Temperatureplays an important role in the acid-catalyzed synthesisof biodiesel. To investigate the influence of temperatureand reaction time, 1 g algal powder, 6 mL n-hexane, 4 mLmethanol, and 0.5 M sulfuric acid were mixed for in situtransesterification. Reaction temperature was set as 20◦C(room temperature), 50◦C, 70◦C, 90◦C, and 110◦C. Foreach temperature, we chose 0.25 hour, 0.5 hour, 1 hour,2 hours, and 4 hours as sampling time. The data wasshown in Figure 5. Our data agreed with the conclusion thathigher temperature could produce biodiesel with high FAMEcontent in much shorter time [32]. Higher yield of biodieselwas produced with higher temperature, especially in the firsthour of the reaction. Interestingly, we found that there was


Table 3: Components of biodiesel from Chlorella pyrenidosabiomass under the conditions of 1 g algal powder, 6 mL n-hexane,and 4 mL methanol with 0.5 M sulfuric acid at 90◦C.

Components Molecular formula Content (%)

10-Octadecenoic acidmethyl ester

C19H36O2 70.18± 3.07

9,12-Octadecadienoicacid methyl ester

C19H34O2 14.53± 5.28

Hexadecanoic acidmethyl ester

C17H34O2 10.48± 1.36

Octadecanoic acidmethyl ester

C19H38O2 3.21± 1.24

Methyl tetradecanoate C15H30O2 0.68± 0.07

9-hexadecenoic acidmethyl ester (z)

C17H32O2 0.31± 0.01

Eicosanoic acid methylester

C21H42O2 0.24± 0.06

Total 99.63± 0.36

no distinction on the FAME content (>98%) of samples from0.25 hour to 4 hours, which suggests that the key point ofin situ transesterification would be how to efficiently extractalgal oil from biomass rather than transesterification itself.The largest amount of biodiesel is obtained at both 90◦C and110◦C with about 95% yield in 2 hours. At 20◦C, 50◦C, and70◦C conditions, the yield was lower and still increasing after4 hours (Figure 5). The reason for more biodiesel productionat higher temperature (90◦C, 110◦C) is that high temperaturecould lead to a fast transesterification rate. Moreover, hightemperature may also improve cell disruption in the in situtransesterification system so that oil in the algal biomasscould contact the reactants more easily, resulting in a higherbiodiesel yield. It was found that there was a slight decreaseof biodiesel from 2 hours to 4 hours at 110◦C (Figure 5).This is probably caused by that high temperature and longreaction time could burn some of the oil [15]. Based onthe quality and quantity standard of biodiesel, the optimizedtemperature and the reaction time for in situ biodieselproduction from microalgal biomass were 90◦C and 2 hours.

3.3.4. Composition of In Situ Biodiesel. The main FAMEcomponents of biodiesel produced from microalgal biomass(1 g) under the optimized conditions of 6 mL n-hexane,4 mL methanol, and 0.5 M sulfuric acid at 90◦C with 2hours reaction time was shown in Table 3. The total FAMEcontent was over 99%. There were seven FAMEs derivatizedin the biodiesel, and the most abundant composition wasoctadecenoic acid methyl ester with the content of 70.18%.Octadecenoic acid methyl ester, octadecadienoic acid methylester, hexadecanoic acid methyl ester and octadecanoic acidmethyl ester are the dominating acid methyl esters, and thetotal content of these four FAMEs was over 98%. This couldresult in the high quality of the biodiesel.

3.4. Possible Methods for Further Utilization of Residue andByproducts. Chlorella pyrenoidosa was able to accumulatehigh content of lipids in the cells with fast growth in the

0

10

20

30

40

50

60

70

80

90

100

0 0.5 1 1.5 2 2.5 3 3.5 4

Bio

dies

elyi

eld

(%)

Time (hours)

20◦C50◦C70◦C

90◦C110◦C

Figure 5: Effects of temperature and reaction time on the biodieselyield by in situ transesterification of C. pyrenoidosa. Reactioncondition: 1 g algal powder with 6 mL n-hexane, 4 mL methanoland 0.5 M sulfuric acid. Temperature: 20◦C, 50◦C, 70◦C, 90◦C, and110◦C.

rice straw hydrolysate medium. Apart from the cellulosepart, other parts of straw could also be used for energyproduction. Kaparaju et al. [33] reported that the liquidhemicellulose fraction from straw pretreatment could beused for biohydrogen production by dark fermentationand the effluents from biohydrogen process could alsobe converted to methane. Thus, the processes would beenergetically most efficient compared to production ofmonofuel from straws in such cases. The present results ofin situ transesterification demonstrated that approximately95% algal oil was converted to high-quality biodiesel. Thissuggested that in situ biodiesel production from microalgalbiomass would be an alternative way for the conversionaltwo-step method. In addition to the biodiesel, glyceroland algae waste, as two main byproducts in algae-basedbiodiesel production, can be made for different uses.Crude glycerol can be transformed into many high valuechemical compounds such as 1,3-propanediol [34], poly-3-hydroxybutyrate [35] and D-glyceric acid production [36].For algae residue treatment, Sialve et al. [37] suggested thatmethane production from algae waste after lipid recovery isviable especially for higher lipid content (>40%). All of theabove methods can be used to make the best and sustainableuse of raw materials in the whole process from straw to algae-based biodiesel. Based on these studies, the idea of biodieselproduction from microalgae by using lignocellulose materialmight be attractive and promising.

4. Conclusions

The study investigated the feasibility of using lignocellulosematerial, rice straw, as carbon sources for microalgae


cultivation. Enzymatic hydrolyzation of pretreated ricestraw produced glucose available for microalgae cultivation.Chlorella pyrenoidosa showed a fast growth rate and a highlipid content growing in the hydrolysate medium. Goodquality and high yield of biodiesel could be obtained fromalgal biomass by adding n-hexane in in situ transesterifi-cation system. This method for in situ biodiesel productionshowed a better performance than the traditional two-stepmethod. The strategy provides a combined idea of recoveringand reusing discarded lignocellulose and utilizing fibrousbiomass for biodiesel production. Further research shouldbe focused on biorefinery of lignocellulose and algal biomassand cost analysis.

Acknowledgments

This work was financially supported by Shanghai MunicipalCommittee of Science and Technology (08DZ1204400). Itwas also supported by the 863 Key Program (2007AA100503)and by China National Offshore Oil Corporation.

References

[1] Y. Chisti, “Biodiesel from microalgae,” Biotechnology Advances,vol. 25, no. 3, pp. 294–306, 2007.

[2] V. Patil, K. -Q. Tran, and H. R. Giselrød, “Towards sustainableproduction of biofuels from microalgae,” International Journalof Molecular Sciences, vol. 9, no. 7, pp. 1188–1195, 2008.

[3] J. O. B. Carioca, J. J. Hiluy Filho, M. R. L. V. Leal, and F. S.Macambira, “The hard choice for alternative biofuels to dieselin Brazil,” Biotechnology Advances, vol. 27, no. 6, pp. 1043–1050, 2009.

[4] EIA, “Gasoline and diesel fuel update,” March 2010,http://tonto.eia.doe.gov/oog/info/gdu/gasdiesel.asp.

[5] Y. Cheng, W. Zhou, C. Gao, K. Lan, Y. Gao, and Q. Wu,“Biodiesel production from Jerusalem artichoke (HelianthusTuberosus L.) tuber by heterotrophic microalgae Chlorella pro-tothecoides,” Journal of Chemical Technology and Biotechnology,vol. 84, no. 5, pp. 777–781, 2009.

[6] A. Wei, X. Zhang, D. Wei, G. U. Chen, Q. Wu, and S. T.Yang, “Effects of cassava starch hydrolysate on cell growthand lipid accumulation of the heterotrophic microalgaeChlorella protothecoides,” Journal of Industrial Microbiologyand Biotechnology, vol. 36, no. 11, pp. 1383–1389, 2009.

[7] Y. Cheng, Y. Lu, C. Gao, and Q. Wu, “Alga-Based biodiesel pro-duction and optimization using sugar cane as the feedstock,”Energy and Fuels, vol. 23, no. 8, pp. 4166–4173, 2009.

[8] C. Gao, Y. Zhai, Y. Ding, and Q. Wu, “Application of sweetsorghum for biodiesel production by heterotrophic microalgaChlorella protothecoides,” Applied Energy, vol. 87, no. 3, pp.756–761, 2010.

[9] A. Margaritis and P. Bajpai, “Continuous ethanol productionfrom Jerusalem artichoke tubers. II. Use of immobilized cellsof Kluyveromyces marxianus,” Biotechnology and Bioengineer-ing, vol. 24, no. 7, pp. 1483–1493, 1982.

[10] C. Martın and A. B. Thomsen, “Wet oxidation pretreatment oflignocellulosic residues of sugarcane, rice, cassava and peanutsfor ethanol production,” Journal of Chemical Technology andBiotechnology, vol. 82, no. 2, pp. 174–181, 2007.

[11] C. E. Wyman, “Potential synergies and challenges in refiningcellulosic biomass to fuels, chemicals, and power,” Biotechnol-ogy Progress, vol. 19, no. 2, pp. 254–262, 2003.

[12] FAO, “Global cereal production brief,” 2009, http://www.fao.org/docrep/012/ai484e/ai484e04.htm.

[13] A. Li, B. Antizar-Ladislao, and M. Khraisheh, “Bioconversionof municipal solid waste to glucose for bio-ethanol produc-tion,” Bioprocess and Biosystems Engineering, vol. 30, no. 3, pp.189–196, 2007.

[14] H. Xu, X. Miao, and Q. Wu, “High quality biodieselproduction from a microalga Chlorella protothecoides byheterotrophic growth in fermenters,” Journal of Biotechnology,vol. 126, no. 4, pp. 499–507, 2006.

[15] X. Miao and Q. Wu, “Biodiesel production from heterotrophicmicroalgal oil,” Bioresource Technology, vol. 97, no. 6, pp. 841–846, 2006.

[16] K. Vijayaraghavan and K. Hemanathan, “Biodiesel productionfrom freshwater algae,” Energy and Fuels, vol. 23, no. 11, pp.5448–5453, 2009.

[17] M. B. Johnson and Z. Wen, “Production of biodiesel fuelfrom the microalga schizochytrium limacinum by directtransesterification of algal biomass,” Energy and Fuels, vol. 23,no. 10, pp. 5179–5183, 2009.

[18] S. Siler-Marinkovic and A. Tomasevic, “Transesterification ofsunflower oil in situ,” Fuel, vol. 77, no. 12, pp. 1389–1391,1998.

[19] E. A. Ehimen, Z. F. Sun, and C. G. Carrington, “Variablesaffecting the in situ transesterification of microalgae lipids,”Fuel, vol. 89, no. 3, pp. 677–684, 2010.

[20] G. Y. S. Mtui, “Recent advances in pretreatment of lignocellu-losic wastes and production of value added products,” AfricanJournal of Biotechnology, vol. 8, no. 8, pp. 1398–1415, 2009.

[21] B. C. Saha, L. B. Iten, M. A. Cotta, and Y. V. Wu, “Dilute acidpretreatment, enzymatic saccharification and fermentation ofwheat straw to ethanol,” Process Biochemistry, vol. 40, no. 12,pp. 3693–3700, 2005.

[22] D. Dong, J. Sun, F. Huang, Q. Gao, Y. Wang, and R.Li, “Using trifluoroacetic acid to pretreat lignocellulosicbiomass,” Biomass and Bioenergy, vol. 33, no. 12, pp. 1719–1723, 2009.

[23] Genencor, “AccelleraseTM 1000,” 2007, http://www.genencor.com/wps/wcm/connect/d23ac9804fa2d4cdaec6be4895e32-24e/ACCELLERASE+1000+prod+info+sheet.pdf?MOD=AJPERES%CACHEID=d23ac9804fa2d4cdaec6be4895e3224e.

[24] F. Sun and H. Chen, “Enhanced enzymatic hydrolysis ofwheat straw by aqueous glycerol pretreatment,” BioresourceTechnology, vol. 99, no. 14, pp. 6156–6161, 2008.

[25] Y. Lu, Y. Zhai, M. Liu, and Q. Wu, “Biodiesel productionfrom algal oil using cassava (Manihot esculenta Crantz) asfeedstock,” Journal of Applied Phycology, vol. 22, no. 5, pp. 573–578, 2010.

[26] T. R. Larson and T. A. V. Rees, “Changes in cell compositionand lipid metabolism mediated by sodium and nitrogenavailability in the marine diatom Phaeodactylum tricornutum(Bacillariophyceae),” Journal of Phycology, vol. 32, no. 3, pp.388–393, 1996.

[27] G. Kildiran, S. O. Yucel, and S. Turkay, “In-situ alcoholysis ofsoybean oil,” Journal of the American Oil Chemists’ Society, vol.73, no. 2, pp. 225–232, 1996.

[28] A. S. Kulkarni, R. R. Khotpal, R. L. Mehta, and H. A. Bhakare,“Studies in sulfation and sulfonation of some semi-dryingoils,” PaintIndia, vol. 41, pp. 41–42, 1991.

[29] M. J. Goff, N. S. Bauer, S. Lopes, W. R. Sutterlin, and G. J.Suppes, “Acid-catalyzed alcoholysis of soybean oil,” Journal ofthe American Oil Chemists’ Society, vol. 81, no. 4, pp. 415–420,2004.


[30] J. Qian, F. Wang, S. Liu, and Z. Yun, “In situ alkalinetransesterification of cottonseed oil for production of biodieseland nontoxic cottonseed meal,” Bioresource Technology, vol.99, no. 18, pp. 9009–9012, 2008.

[31] S. Ozgul-Yucel and S. Turkay, “Variables affecting the yieldsof methyl esters derived from in situ esterification of rice branoil,” Journal of the American Oil Chemists’ Society, vol. 79, no.6, pp. 611–614, 2002.

[32] X. Miao, R. Li, and H. Yao, “Effective acid-catalyzed transes-terification for biodiesel production,” Energy Conversion andManagement, vol. 50, no. 10, pp. 2680–2684, 2009.

[33] P. Kaparaju, M. Serrano, A. B. Thomsen, P. Kongjan, and I.Angelidaki, “Bioethanol, biohydrogen and biogas productionfrom wheat straw in a biorefinery concept,” BioresourceTechnology, vol. 100, no. 9, pp. 2562–2568, 2009.

[34] P. A. Selembo, J. M. Perez, W. A. Lloyd, and B. E. Logan,“Enhanced hydrogen and 1,3-propanediol production fromglycerol by fermentation using mixed cultures,” Biotechnologyand Bioengineering, vol. 104, no. 6, pp. 1098–1106, 2009.

[35] C. Zhu, C. T. Nomura, J. A. Perrotta, A. J. Stipanovic,and J. P. Nakas, “Production and characterization of poly-3-hydroxybutyrate from biodiesel-glycerol by Burkholderiacepacia ATCC 17759,” Biotechnology Progress, vol. 26, no. 2,pp. 424–430, 2010.

[36] H. Habe, T. Fukuoka, D. Kitamoto, and K. Sakaki, “Biotech-nological production of d-glyceric acid and its application,”Applied Microbiology and Biotechnology, vol. 84, no. 3, pp. 445–452, 2009.

[37] B. Sialve, N. Bernet, and O. Bernard, “Anaerobic digestion ofmicroalgae as a necessary step to make microalgal biodieselsustainable,” Biotechnology Advances, vol. 27, no. 4, pp. 409–416, 2009.


Review Article

Molecular Breeding of Advanced Microorganisms forBiofuel Production

Hiroshi Sakuragi, Kouichi Kuroda, and Mitsuyoshi Ueda

Division of Applied Life Sciences, Graduate School of Agriculture, Kyoto University, Kitashirakawa-oiwake-cho,Sakyo-ku, Kyoto 606-8502, Japan

Correspondence should be addressed to Mitsuyoshi Ueda, [email protected]

Received 1 September 2010; Revised 29 November 2010; Accepted 1 December 2010

Academic Editor: Effie Tsakalidou

Copyright © 2011 Hiroshi Sakuragi et al. This is an open access article distributed under the Creative Commons AttributionLicense, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properlycited.

Large amounts of fossil fuels are consumed every day in spite of increasing environmental problems. To preserve the environmentand construct a sustainable society, the use of biofuels derived from different kinds of biomass is being practiced worldwide.Although bioethanol has been largely produced, it commonly requires food crops such as corn and sugar cane as substrates. Todevelop a sustainable energy supply, cellulosic biomass should be used for bioethanol production instead of grain biomass. For thispurpose, cell surface engineering technology is a very promising method. In biobutanol and biodiesel production, engineered hostfermentation has attracted much attention; however, this method has many limitations such as low productivity and low solventtolerance of microorganisms. Despite these problems, biofuels such as bioethanol, biobutanol, and biodiesel are potential energysources that can help establish a sustainable society.

1. Introduction

In the 20th and 21st century, especially in the last severaldecades, our lifestyle has dramatically changed and becamemuch more comfortable owing to the developments insciences and technology. However, the advantages affordedby such progress caused many environmental problems suchas air pollution, climate changes, and global warming. Globalwarming is mainly caused by excess and accumulating green-house gases such as CO2, NOX, and SO2. The major cause ofincreasing CO2 concentration in the atmosphere is excessiveconsumption of fossil fuels, for example, petroleum, coal,and natural gases. Energy and some products such as plasticsare derived from oil. However, their production causesemission of several greenhouse gasses, such as CO2, in theatmosphere. Moreover, excessive consumption of fossil fuelcauses not only global warming but also global economicalproblems. Paradigm-changing new ideas and technologiesare required to overcome these problems.

Utilization of biomass energy can potentially reducethe emission of greenhouse gases. This is because throughphotosynthesis, CO2 released from the combustion of

biomass energy is recycled without affecting the overall CO2

balance in the atmosphere. Therefore, the introduction ofbiofuels is considered a promising approach to reduce thedependence on fossil resources. Some examples of biofuelsare bioethanol, biobutanol, and biodiesel. Bioethanol fer-mentation is one of the largest-scale microbial processesusing sugars or polysaccharides that can be depolymerizedto a fermentable sugar [1]. Biomass sources such as grains,however, are also used as food, and thus the increasingdemand for bioethanol has led to a global increase in theprices of food crops. Therefore, inedible cellulosic biomassresources should be used for the production of bioethanol(the so-called second-generation bioethanol).

Biobutanol has been attracting more attention thanother biofuels, especially after the announcement by BP andDuPont. These companies started to finance the develop-ment of modernized biofuel production plants supportedby research and development [2, 3]. Biobutanol is producedby the fermentation process of Clostridium species, whichnaturally possess metabolic pathways for the conversion ofsugar into solvents such as acetone, butanol, and ethanol.Although the process of biofuel conversion by clostridia


SSCF process

CBP process

Physicochemicalpretreatment

Enzyme production + saccharification+ pentose and hexose cofermentation

Saccharification + pentose andhexose cofermentation

Enzyme productionBiomass

SSCF: simultaneous saccharification andCBP: consolidated bioprocessing

cofermentation

Figure 1: Illustration of CBP (consolidated bioprocessing) system.

is known (acetone-butanol-ethanol [ABE] fermentation),the mechanism to regulate the metabolic fluxes in theseorganisms is still obscure. For the industrial-level productionof butanol, clostridia fermentation has been widely studied,including pretreatment of substrates, techniques of butanolrecovery, and induction of solventogenesis. In addition, theintroduction of butanol synthetic pathways into hosts withclostridial metabolic pathways or keto-acid pathways has alsobeen investigated.

Biodiesel is also expected to be a substitute for petro-leum-based diesel fuel. The most common process for bio-diesel production is extraction of oils from vegetable oilfeedstocks, such as palm tree, soybean, and rapeseed, andconverting the oil to biodiesel by transesterification oftriacylglycerols (TAGs) with methanol and ethanol [4].Fatty acid methyl esters (FAMEs) and fatty acid ethylesters (FAEEs) are synthesized by this reaction. However,the cost and energy requirement of this process and thesubsequent separation are high, and the availability of cheapvegetable oil feedstocks is also limited [5]. To overcomethese limitations, several studies have focused on enzymatictransesterification using lipase with whole cell biocatalyststechnology and biodiesel production from microalgae. Inaddition, engineered host fermentation by using Escherichiacoli and Saccharomyces cerevisiae containing heterologousgenes has also been shown to be involved in the productionof FAEEs.

The major problems in the production of biofuel includelow and nonspecific productivity and high cost. However,advances in genetic manipulation and metabolic engineeringcan offer solutions to these problems.

2. Bioethanol Production

Bioethanol is a carbon-neutral fuel derived from biomassfeedstocks such as grain and wood. The yeast S. cerevisiaeproduces ethanol in batch fermentation, with CO2 and smallamounts of methanol, glycerol, and others as byproducts.Modern industrial yeast strains produce up to 20% (v/v)of ethanol, by using glucose derived from starch, owing totheir increased inhibitor resistance, ethanol tolerance, andethanol-specific productivity. When ethanol is used as a fuel,the azeotropic mixture of 95.57 wt% ethanol and 4.43 wt%water could be used as car fuel. The large amount of water in

the mixture, however, causes phase separation between waterand gasoline [3]. Most of the raw materials currently used forbioethanol production are grain biomass such as corn grainand sugar cane; however, future limitation in the supply ofthese materials is inevitable because they are also used as foodcrops. Therefore, lignocellulosic biomasses are consideredattractive raw materials for bioethanol production. To con-struct an energy-saving and sustainable society, developmentof technologies to utilize lignocellulosic biomasses is stronglyrequired.

Lignocellulosic biomass is composed of cellulose (40–50%), hemicellulose (25–35%), and lignin (15–20%) [6].Cellulose is the most abundant biomacromolecule on earthand is embedded with hemicelluloses and lignin in the plantcell wall matrix. It consists of a long straight chain formedby β-1,4 glucosidic linkage of d-glucose, and containsmicrofibrils with a crystalline stable structure formed by astrong hydrogen bond between molecular chains. Celluloseis difficult to convert into the amorphous state and easilyaccumulates in the environment. Hemicelluloses are nonho-mogeneously distributed in the plant cell wall, and are boundto cellulose by hydrogen bonds and to lignin by ether, ester,and glycoside bonds. They include xylan, mannan, xyloglu-can, and glucomannan. Lignin is hydrophobic and has a 3-dimensional structure, in which phenylpropane monomersrandomly polymerize, which strengthens the physical prop-erties of cell wall polysaccharides. For efficient utilization ofcellulose and hemicelluloses, their lignin content should beremoved because lignin covers the polysaccharides used asmaterials for bioethanol production and prevents cellulosicbiomass from being hydrolyzed.

2.1. Consolidated Bioprocessing (CBP) and Cell SurfaceEngineering Technology. There are 4 major bioprocesses inbioethanol production from cellulosic biomass: productionof cellulases and hemicellulases, hydrolytic degradation ofcellulose and hemicelluloses, C6 sugar fermentation, and C5sugar fermentation. The consolidated bioprocessing (CBP)is the system by which these 4 bioprocesses are allowed tooccur in a single fermenter (Figure 1). The simultaneoussaccharification and cofermentation (SSCF) is the system bywhich saccharification and fermentation are simultaneouslyperformed. This is the primarily existing method of ethanolfermentation, but simplification and cost reduction of the


process are highly desired. Whereas SSCF needs the processof enzyme production, CBP can simultaneously also produceenzymes in a single fermenter. CBP can allow more efficientproduction of bioethanol in a smaller fermenter. Therefore,CBP has an advantage in terms of reduced cost of productionand equipment. To carry out CBP, however, it is essential thata single and advanced species of microorganism is used inthe production of saccharification enzymes, saccharificationof biomasses, and fermentation of saccharified sugar [7].Thus, yeasts having saccharification ability are ideal forethanol production with CBP. Because yeasts can performethanol fermentation of sugars but not saccharification, anengineered yeast can be utilized as a microorganism for CBP.

The cell surface engineering technology is a very usefulstrategy for molecular breeding of yeasts as CBP microor-ganisms. Using this technology, functional proteins canbe displayed on the cell surface of the microorganism.Therefore, cell surface engineering technology can provideintact cells with new functions, and construct various armingcells with novel functions [8–11]. The cell surface is afunctional interface between the inside and the outside of thecell. S. cerevisiae has the rigid cell wall that is about 200 nmthick, mainly composed of mannoproteins and β-linkedglucans, and lies outside the plasma membrane. In yeasts,several proteins on the cell surface have secretion signalpeptides at the N-terminal and glycosylphosphatidylinositol(GPI) anchors at the C-terminal, which play important rolesin surface production and are essential for cell viability. Uponcompletion of protein synthesis, the secreted proteins aretranslocated into the lumen of the endoplasmic reticulum(ER), and transported from the ER to the Golgi apparatusand then to the plasma membrane in membrane-enclosedvesicles. GPI-anchored proteins are further transported tothe outside of the plasma membrane through the secre-tory pathway, released from the plasma membrane bya phosphatidylinositol-specific phospholipase C (PI-PLC),and transferred to the outermost surface of the cell wall(Figure 2) [10]. α-Agglutinin, which is encoded by AGα1and interacts with the binding subunit of the agglutinincomplex of a-type cells, is one of the GPI-anchored cellsurface proteins. α-Agglutinin is composed of a secretionsignal region, an active region, an anchoring region richin serine and threonine, and a GPI anchor-attachmentsignal (Figure 3(a)). Using this molecular information ofcell wall localization mechanism of proteins, it has becamepossible to display target heterologous proteins on the yeastcell surface by genetic engineering techniques (Figure 3(b))[11]. Furthermore, cell surface engineering is an innovativemolecular tool by which the function of a displayed proteincan be analyzed on intact cells. Because of the lack of need forprotein purification and concentration, the mutated proteinscan be analyzed by treating the cells as microparticles coveredwith proteins [12]. Therefore, cell surface engineering tech-nology enables the construction of various biocatalysts withpotential for industrial utilization.

2.2. Ethanol Production by Using Cell Surface Engineering.Starch is the most conveniently used biomass materialand consists of 2 types of macromolecules, amylose and

ER

Golgi

Exocytosis

Secretory vesicleProtein folding quality

control system

GPI anchorCleavage by PI-PLC andcovalent linkage with glucan

Nucleus

Cell wallCell membrane

Figure 2: Mechanism of cell surface display of proteins by cellsurface engineering.

Secretion signal sequenceGPI anchor attachment

signal sequence

Active region Ser/Thr rich region-

(a)

Protein to be displayedencoding gene

Promoter

Secretion signal sequence

Protein to be displayed C-terminal half of α-agglutinin

GPI anchor

GPI anchor attachmentsignal sequence

3half of α-agglutinin

(b)

Figure 3: Molecular structure of α-agglutinin (a) and moleculardesign of cell surface-displayed enzyme (b).

amylopectin. When starch is converted into glucose byAspergillus, yeasts assimilate glucose and ferment ethanolbecause they cannot directly utilize starch. The cell surfacedisplay system is very useful in bioethanol production fromstarch by CBP. When various amylases are displayed onthe yeast cell surface, the constructed yeasts acquire theability to directly utilize starch as the sole carbon sourcein ethanol production. Arming yeast cells with displayedglucoamylase were cultivated in aerobic condition withsoluble starch as the sole carbon source, and were grown [8].Yeasts codisplaying glucoamylase and α-amylase grew faster


with starch as the sole carbon source than yeasts displayingonly glucoamylase [13]. These results show that the cellsurface display of amylolytic enzymes in yeasts through cellsurface engineering can integrate the multiple conventionalprocesses using Aspergillus into a single process using arming(cell-surface-engineered) yeast cells. This shows why cellsurface engineering technology can realize the constructionof a CBP system. In addition, the concentration of glucosein the fermentation medium is always kept zero becauseglucose produced by starch degradation on the cell surfaceis quickly imported into the yeast cell. Therefore, bacterialcontamination can be prevented during ethanol productionby using surface-engineered yeast cells.

Cellulosic biomass is a more abundant renewableresource than grain biomass and is attracting much attentionas a cheap and available material that does not competewith food supply. Cellulose is a major component of cellu-losic biomass and a high-molecular-weight polysaccharide.Degradation of cellulose requires many enzymes and is moredifficult than that of starch because of the existence ofcrystalline and amorphous regions in cellulose. Three typesof cellulolytic enzymes are required for cellulose degradation:endoglucanases (EGs), cellobiohydrolases (CBHs), and β-glucosidase (BGL). The endo-exo synergism of EGs andCBHs efficiently degrades the cellulose chain into solublecellobiose and cello-oligosaccharides. Yeasts, however, donot have these enzymes and cannot degrade cellulose. Toendow yeast cells with the ability to degrade cellulose,EG II and CBH II from Trichoderma reesei and BGL1from Aspergillus aculeatus were displayed on the yeast cellsurface by cell surface engineering (Figure 4). When EG II,CBH II, and BGL1 were simultaneously codisplayed on theyeast cell surface, the sum of the numbers of individualdisplayed enzymes in cells displaying 3 enzymes is largerthan that of cells displaying a single enzyme, although thenumbers of the individual enzymes displayed on the cellsurface were considered to be decreased by the simultaneousdisplay of 3 enzymes. When the surface-engineered yeast cellswere suspended and cultivated in anaerobic condition withphosphoric-acid-swollen cellulose as amorphous celluloseafter aerobic cultivation, the yeast strain could directlyproduce ethanol from amorphous cellulose [14].

β-1,4-Xylan is a major component of hemicelluloses anda complex polysaccharide consisting of a backbone of β-1,4-linked xylopyranoside. To produce ethanol efficiently fromcellulosic biomass, bioconversion of xylan is also required.Endo-β-xylanase hydrolyzes xylan to xylooligosaccharides,and β-xylosidase subsequently hydrolyzes xylooligosaccha-rides to d-xylose. These enzymes need to be displayed onthe yeast cell surface because S. cerevisiae cannot degradexylan to xylose. Using the cell surface display system, xylanaseII (XYN II) from T. reesei and β-xylosidase (XylA) from A.oryzae were displayed on the yeast cell surface [15]. Theseenzymes displayed on the yeast cell surface showed activities,and the yeast cells codisplaying XYN II and XylA hydrolyzexylan to xylose subsequently. For further conversion of thegenerated xylose into ethanol, yeast cells should be endowedwith the ability to assimilate xylose. The ability to convertxylose into xylulose needs to be furnished in yeasts because

yeasts can intrinsically assimilate xylulose, which is theisomer of xylose. Microorganisms use 2 pathways for theconversion of xylose into xylulose. One pathway involvesxylose reductase (XR) and xylitol dehydrogenase (XDH),whereas the other pathway involves xylose isomerase (XI).An example of a yeast strain endowed with a relatively highxylose fermentation ability has been recently reported, butthe pathway involving XI has not been utilized until now[16]. Focusing on the pathway involving XR and XDH, axylose-utilizing yeast strain was constructed by the codisplayof XYN II and XylA, the intracellular production of XR andXDH from Pichia stipitis, and the enhanced expression ofthe xylulokinase (XK) gene from S. cerevisiae [15]. Whenethanol production from xylan by using the constructedyeast was anaerobically examined after aerobic cultivation,it was confirmed that xylan was simultaneously saccharifiedand fermented.

2.3. Electrical Current Production from Ethanol. These days,the production of electrical current from biofuels, especiallyethanol, is also very attractive. Biofuel cells are energyconversion devices like traditional fuel cells that convertthe chemical energy of fuels into electricity. The featureof biofuel cells is energy conversion through the use ofbiological catalysts at the anode and/or the cathode. Withtwo electrode interfaces, biocatalyzed oxidation of organicsubstances has a way to convert chemical energy to electricalenergy, and the chemical energy of various biological sub-stances such as biofuels may be transformed into electricalcurrent by enzymes and microorganisms in biofuel cellssystem [17, 18]. Direct electron transfer (DET) to or fromenzymes was important for producing an electric potentialbetween the anode and cathode. However, the most ofenzymes cannot transfer electrons directly. In enzymaticbiofuel cells, enzyme-modified electrodes are powered byethanol using the anode where quinohemoprotein-alcoholdehydrogenase (QH-ADH) is immobilized and the cathodewhere alcohol oxidase (AOx) and microperoxidase (MP-8)are immobilized. QH-ADH transfers electrons directly to thecarbon-based electrode and MP-8 accepts electrons directlyfrom same type of electrodes. Although the biofuel cells hasweakness of long-term operation, they can convert commonbiological substances such as ethanol to an electrical currentwithout redox mediators [19].

3. Biobutanol Production

Biofuels are produced from renewable biomass by microbes,and butanol has more significant advantages as biofuels thanethanol. First, butanol can be used either in its pure form orin a mixture with gasoline at any concentration. Ethanol canbe mixed only up to 85%. Second, combustion of butanoldoes not require modification of existing car engines becausebutanol is less soluble in water than ethanol. Third, the vaporpressure of n-butanol (4 mm Hg at 20◦C) is approximately11 times less than that of ethanol (45 mm Hg at 20◦C),and n-butanol is safer than ethanol owing to lower vaporpressure. Fourth, butanol is less corrosive than ethanol; thus,


(Synergistic reaction)

(Sequential reaction)

Armingyeast

Cellulose

Cellobiose

Cellooligosaccharide

Glucose

Ethanol

Ethanol

Glucose

Cellobiohydrolase (CBH)

C-terminal half of α-agglutinin

OOH

OHOH

OH

HOHOHO

OH

OO

β-Glucosidase (BGL)

Endoglucanase (EG)

HO

HO

HO

HO OH

OH

OH

OH

OO

O OO O

OO

HO

HO

OH

OH

HO

HO

HO

HO OH

OH

OH

OH

OO

O OO O

OO

HO

HO

OH

OH

HO

HO

HO

HO OH

OH

OH

OH

OO

O OO O

OO

HO

HO

OH

OH

HO

HO

HO

HO OH

OH

OH

OH

OO

O OO O

OO

HO

HO

OH

OH

Figure 4: Cellulose-assimilating yeast by codisplaying of cellulolytic enzymes.

Table 1: Comparison of butanol with ethanol and gasoline.

FuelCaloric value

(MJ/L)Air-fuel ratio

Research octanenumber

Butanol 29.2 11.2 96

Ethanol 21.2 3 129

Gasoline 32.5 14.6 91–99

it will not cause damage to existing infrastructures, forexample, tanks, pipelines, pumps, and filling stations. Fifth,the energy content of butanol is about 40% higher than thatof ethanol. Table 1 shows the comparison of 3 characteristicsof the important biofuels with those of gasoline [20, 21].The less desirable characteristic of butanol is its researchoctane number (i.e., 96), which is lower than that of ethanol.The octane number generally increases with the number ofdouble bonds and methyl branches of molecules. Amongbranched C4 and C5 alcohols that are also consideredpotential gasoline alternatives, isobutanol is also currentlyunder investigation as one of new biofuel targets. Isobutanolhas very similar characteristics to n-butanol, although it hasa higher octane number [22].

3.1. n-Butanol Production by Clostridia Fermentation. It iswell known that n-butanol can be produced by clostridia

fermentation. Clostridia consist of a diverse group ofanaerobic, spore-forming, and gram-positive bacteria thatinclude notable pathogens or industrial significant microor-ganisms. Clostridium acetobutylicum was found to produceacetone/butanol/ethanol at a ratio of 3 : 6 : 1. This process iscalled ABE fermentation [2, 3]. Corn starch was used as asubstrate at facilities in the UK and France, while rice starchwas used at facilities in India [23]. The bacterial productionof butanol and acetone by using the ABE fermentationprocess was valuable in the production of the lacquer solventbutylacetate and in the development of the synthetic rubberindustry. However, bacterial production has declined withthe advancement in the petrochemical industry, which canproduce acetone and butanol at low costs. Biofuel productionis currently increasingly being practiced worldwide, andresearch and development into microbial butanol produc-tion is again becoming actively pursued.

Clostridia such as C. acetobutylicum, C. beijerinckii, andC. saccharoperbutylacetonicum were demonstrated to havevery similar metabolic pathways (Figure 5) [24–26]. WhenC. acetobutylicum conducts fermentation, it produces 3major classes of products: solvents (acetone, ethanol, and n-butanol), organic acids (acetic acid, lactic acid, and butyricacid), and gases (carbon dioxide and hydrogen). Using starchor sugar, it first carries out acid fermentation, such asbutyrate and acetate in the exponential growth phase, andsolvent fermentation at the end of the exponential growth


phase. In the solventogenic phase, the excreted acids aretaken up again and converted to n-butanol and ethanol. Asa result, more n-butanol is produced than ethanol, and morebutyrate is produced than acetate [27, 28]. The change inculture condition has been shown to be required for theinduction of n-butanol production. Acetate and butyrateproduced during the exponential growth phase reduce theculture pH, and the acidic condition at the end of theexponential growth phase is widely regarded to trigger theshift in metabolism resulting from solvent production [23].Another factor shown to trigger n-butanol production isthe regulator protein SpoOA-P, which is responsible forsporulation as well as induction of genes involved in n-butanol production [29, 30]. The induction mechanisms ofsolventogenesis and sporulation in C. acetobutylicum havesimilar characteristics; therefore, more studies on the keyfactors for these processes are required to further clarify thesolvent, especially in n-butanol production.

The traditional clostridial fermentation of butanol hasseveral limitations, especially the high cost of substrates. Tosolve such problems, biomasses like cassava, wheat straw, andliquefied corn starch, and not pure sugar such as glucose, areused as substrates in ABE fermentation of clostridia. A previ-ous study reported that addition of ammonium acetate to thecassava medium significantly facilitates solvent productionfrom cassava fermented by C. acetobutylicum EA 2018 [31].When C. beijerinckii P260 fermented wheat straw hydrolysatein the batch culture, the total solvent productivity and yieldwere superior to the control fermentation of glucose, andproductivity was improved by 214% [32]. Liquefied cornstarch is a promising industrial substrate and has beenused for successful ABE fermentation. Batch fermentation ofliquefied corn starch by using C. beijerinckii BA101 resultedin the production of total solvent that is almost equivalentto that of glucose. In addition, when solvent was recoveredfrom a fed-batch reactor by gas stripping, more total solventwas produced and much more liquefied corn starch wasconsumed. Its consumption was improved by 487% ofcontrol [33]. Gas stripping is a simple and useful recoverytechnique that can be integrated with ABE fermentation andsimultaneously recovers solvent during ABE fermentation.This technique has the additional advantage of not requiringa membrane, because membrane-based recovery systems cancause fouling and clogging, and is significant for keepingbutanol concentration in the fermentation reactor below thethreshold of butanol toxicity to the culture [25]. Moreover,other recovery techniques, such as liquid-liquid extraction[34], perstraction [35], and pervaporation [36], are available.However, their recovery levels of butanol do not satisfyindustrial-level requirements.

The butanol production pathway from acetyl-CoA tobutanol in C. acetobutylicum is illustrated in Figure 5. Thepathway from acetyl-CoA to n-butanol requires 8 enzymes:acetyl-CoA acetyltransferase (thiolase; THL), β-hydroxy-butyryl-CoA dehydrogenase (HBD), 3-hydroxybutyryl-CoAdehydratase (crotonase; CRT), butyryl-CoA dehydrogenase(BCD), electron transfer flavoprotein A (ETFA) and B(ETFB), aldehyde dehydrogenase (ADHE), and aldehyde-alcohol dehydrogenase (ADHE1). THL plays an important

β-Hydroxybutyryl-CoA

1

5

Crotonyl-CoA

Glucose

Butyryl-CoAButyrate

Acetoacetyl-CoA Acetone

Acetate Ethanol

EMP pathway

NADH

NAD+

2 NADH

Butyraldehyde

2

3

4

5

NADH

NAD+

NADH

NAD+

NADH NAD+

2 NAD+

H2O

2 Pyruvate

2 Acetyl-CoA

n-Butanol

Figure 5: The metabolic pathways of C. acetobutylicum. Numbersrefer to the enzymes: 1: acetyl-CoA acetyltransferase (thiolase;THL), 2: β-hydroxybutyryl-CoA dehydrogenase (HBD), 3: 3-hydroxybutyryl-CoA dehydratase (crotonase; CRT), 4: butyryl-CoAdehydrogenase (BCD), electron transfer flavoprotein A (ETFA) andB (ETFB), 5: aldehyde dehydrogenase (ADHE)/aldehyde-alcoholdehydrogenase (ADHE1).

role in the production of both acids and solvents. BCDinteracts with ETFA and ETFB in its redox reaction. The openreading frames of bcd, etfB, and etfA are located betweencrt and hbd. Clustered genes encoding CRT, BCD, ETFB,ETFA, and HBD are transcribed as 1 transcriptional unitand form an operon [37]. ADHE and ADHE1 catalyze theconversion of butyryl-CoA to butyraldehyde and butyralde-hyde to butanol accompanied by oxidation of NADH. Thegenome sequence of C. acetobutylicum has been revealed,which allows us to have a considerable understanding ofits metabolic pathways, cellular regulation, and genetics[38]. Using genome and metabolic information, metabolicengineering of C. acetobutylicum has been shown to increasesolvent production and solvent tolerance.

An antisense RNA (asRNA) strategy has been shown toimprove the selectivity for butanol production. Althoughonly asRNA against ctfB (the second CoA transferase gene


in the polycistronic aad-ctfA-ctfB message) in C. aceto-butylicum drastically decreased the acetone and butanol lev-els compared to control, asRNA against ctfB combined withoverexpression of the alcohol-aldehyde dehydrogenase gene(aad) resulted in an increase of butanol/acetone ratio. Theconcentration of butanol produced by the mutant strain wasimproved by about 230% of control strain, and the highestever reported in C. acetobutylicum. This demonstrated thatasRNA against ctfB degraded the entire sol operon (aad-ctfA-ctfB) transcript. Indeed, the butanol/acetone ratio ofthe mutant strain was more than twice as much as thatof the control strain [39]. The method of asRNA strategywas a successful one in increasing the selectivity of butanolproduction.

It is known that clostridial cellular metabolism ceaseswhen solvent concentration reaches 20 g/L. Thus, one of thecrucial problems of ABE fermentation is the tolerance ofclostridia to solvent toxicity; however, clostridial tolerancemechanism has not been elucidated because of the increasingnumber of genes involved in solvent tolerance. Based on thereports that clostridia preferentially grow under conditionsof butanol stress, the elements of its genomic library wereidentified. Overexpression of groESL, the class I heat shockprotein gene, resulted in increased final solvent concentra-tions [40]. DnaK, the protein product of the dnaK gene, mayenhance the tolerance of C. acetobutylicum to solvent becauseit is induced during the onset of solventogenesis. Moreover,a cluster of heat shock genes in the dnaK gene region of C.acetobutylicum, including grpE, dnaK, and dnaJ, and a newheat shock gene encoding an unknown heat shock proteinhas been identified [41].

The butanol toxicity limit of the wild-type strain is13 g/L [23]. The final concentration of strain PJC4BK andstrain PJC4BK(pTAAD) surpassed that of the wild typewithout any selection for butanol tolerance. Strain PJC4BKis prepared by the inactivation of butyrate kinase (buk) inC. acetobutylicum, and strain PJC4BK(pTAAD) is preparedby the overproduction of alcohol aldehyde dehydrogenase(aad). This suggests that butanol production is not triggeredby, or directly related to butanol concentration or tolerancelimits [42]. It is also clear that butanol production is notlimited by alcohol aldehyde dehydrogenase activity underthis experiment’s fermentation conditions because strainPJ4BK(pTAAD) produced as much butanol as PJC4BK.Strain SolRH, which is prepared by the inactivation ofsolH in C. acetobutylicum, had a higher rate of glucoseutilization and produced higher solvent compared to thewild type. The gene product of solH is a putative repressorof solvent formation gene. Strain SolRH(pTAAD) producedeven higher concentrations of solvents than strain SolRH[43]. These reports demonstrate that an increased solventtolerance is related to increased butanol production withoutany selection for solvent tolerance.

3.2. Metabolic Engineering for Butanol Production

3.2.1. Application of the Clostridial Metabolic Pathway.Although metabolic engineering of clostridia has beendeveloped for many years, this is not ideal for butanol

production because of the relative lack of genetic toolsto regulate the organisms’ metabolism, anaerobicity, slowgrowth, weak butanol tolerance, and byproduct production,such as acetone and ethanol. Therefore, metabolic engineer-ing of organisms has been essential for the production ofbutanol with the rapidly expanding genomic informationand molecular biology techniques [44]. The genes of C.acetobutylicum ATCC 824 (thl, hbd, crt, bcd, etfA, etfB, andadhE2) involved in the biosynthesis of n-butanol in clostridiawere cloned into, and expressed in E. coli. Under aerobicconditions, the engineered strain produced n-butanol, andmore n-butanol was produced by deleting the pathwayscompeting with n-butanol production when the bacteria wasgrown in rich media [21]. Similarly, when different genesof C. acetobutylicum ATCC 824 (thl, hbd, crt, bcd-etfB-etfA,and adhe1 or adhe) were cloned into and expressed in E.coli, the engineered strain with adhe produced 4 times asmuch n-butanol as the engineered strain with adhe1 [2].Although the activity of the product of the bcd gene wasnot detected in E. coli in the previous study [37], this studyshowed this activity and its requirement for etfA and etfBcoexpression. Without any selection for butanol tolerance orengineering for increased toxicity thresholds, E. coli exhibitedtolerance up to 1.5% of butanol [45], which is competitivewith clostridia. The amount of butanol produced using E.coli was lower than clostridia, but E. coli still has potential asa host for high butanol production.

S. cerevisiae has been considered another ideal engineeredhost for butanol production. It has inherent tolerance tosolvents due to its extensive use in industrial production ofethanol, and is able to grow in aerobic conditions, unlikeclostridia. In addition, it is thought to produce more CoAand NADH, which are required for the clostridial metabolicpathway of butanol production because it is a eukaryoticorganism. When genes encoding related enzymes chosenfrom C. beijerinckii, E. coli, and Ralstonia eutropha, andothers were cloned into S. cerevisiae, the engineered strainproduced n-butanol [46]. However, this concentration ofbutanol is still much less than that of butanol producedby clostridia and the most recently engineered E. colistrains. Butanol toxicity is not the limiting factor of butanolproduction because S. cerevisiae can tolerate up to 2%butanol [47].

Lactic acid bacteria are used for several biotechnologicalapplications, and methods for their manipulation havebeen well developed. Lactobacillus brevis naturally has thehighest tolerance of butanol (3.0% of butanol) among severalorganisms [47]; thus, it was also subjected to the basis ofmetabolic engineering to produce butanol. Recombinant L.brevis strains were able to produce n-butanol on glucosemedium [48].

3.2.2. Application of Keto-Acid Pathways. When clostridialmetabolic pathways are introduced into other organisms forbutanol production, expression of heterogeneous genes maycause the imbalance of electron flow and metabolic pathway,and the accumulation of the heterogeneous metabolites mayelicit the cytotoxicity of engineered hosts. For the pro-duction of greater amounts of butanol, synthetic pathways


1

2

3

4

5

Isobutyraldehyde

Isobutanol

Glucose

2 NAD+ 2 NADH

CO2

NAD(P)+

NAD(P)+

2-Acetolacetate

2-Ketoisovalerate

2,3-Dihydroxy-isovalerate

2 Pyruvate

NADH (NADPH)

NADH (NADPH)

Figure 6: Synthetic pathway of isobutanol through keto-acidpathway. Numbers refer to the enzymes; 1: acetolactate synthase(AlsS), 2: acetohydroxy acid isomeroreductase (IlvC), 3: dihydroxy-acid dehydratase (IlvD), 4: 2-ketoacid decarboxylase (KDC), 5:alcohol dehydrogenase (ADH).

introduced into the heterologous host could be suitableto the native metabolic machinery. There is a metabolicengineering approach using E. coli to produce longer-chainalcohols, including n-propanol, isobutanol, n-butanol, 2-methyl-1-butanol, 3-methyl-1-butanol, and 2-phenylethanolfrom glucose. This strategy utilizes the host’s highly activeamino acid biosynthetic pathway and converts its 2-keto-acid intermediates to higher alcohols [49]. To divert themetabolic intermediates from amino acid biosynthesis path-ways to higher alcohols, only 2 non-native steps for 2-keto-acid degradation were introduced into E. coli. First isthe conversion of 2-keto-acid into aldehydes by 2-keto-aciddecarboxylases (KDCs), and second is the conversion ofaldehydes into alcohols by alcohol dehydrogenases (ADHs).Moreover, when specific 2-keto-acids were added to theculture, the E. coli strain with overexpression of KDC andADH produced the specific and corresponding alcohols.These results suggest that increasing the flow to the 2-keto-acids could develop the productivity and specificity of thealcohols [50].

As the other pathway of n-butanol production, the 2-keto-acid degradation pathway can also be used. The keto-acid precursor of n-butanol is 2-ketovalerate. It is a raremetabolite in the cell causing the synthesis of norvaline,which is an unnatural amino acid, and is produced througha minor side reaction of the leucine biosynthesis pathway.

CH2-OOC-R1

CH-OOC-R2

CH2-OOC-R3

3ROH

R1-COO-R

R2-COO-R

R3-COO-R

CH2-OH

CH-OH

CH2-OH

Glyceride Alcohol Esters Glycerol

+ +

Figure 7: Biodiesel production by transesterification of triglyc-erides with alcohol.

Then, increasing the upstream precursors of 2-ketovalerateis required to produce n-butanol through this pathway.The upstream precursors are threonine and its deaminationproduct, 2-ketobutyrate [50, 51]. To increase the intracellularlevels of threonine and 2-ketovalerate, thrABC, ilvA, andleuABCD were overexpressed in the E. coli strain withcompeting pathways deleted. The product of thrABC gene isinvolved in threonine production. The product of ilvA geneis threonine dehydrogenase, which catalyzes the reactionof threonine to 2-ketobutyrate. The protein product ofleuABCD gene catalyzes the conversion of 2-ketobutyrate to2-ketovalerate. The engineered strain produced 0.9 g/L n-butanol in the initial shake flask experiments without muchoptimization [50].

The 2-keto-acid degradation pathway was also used forisobutanol production. Isobutanol can be produced from2-ketoisovalerate, an intermediate in valine biosynthesis.When alsS, ilvC, and ilvD were overexpressed in the E. colistrain, the carbon source was converted to 2-ketoisovalerate,and 2-ketoisovalerate was converted to isobutanol usingKDC and ADH (Figure 6). The genes of the competingpathways were deleted, which caused an increase in thelevel of pyruvate available for the synthetic isobutanolpathway. The engineered strain produced 20 g/L isobutanol,and its yield is 86% of the theoretical maximum [49, 50].The previous report that similarly used this pathway forisobutanol production showed the effects of various ADHs[52]. The high production of isobutanol demonstrates thatthis pathway is one of promisings for industrial production.Additionally, isobutanol is one of the advanced biofuels liken-butanol, and has a higher octane number than n-butanol.

4. Biodiesel Production

Biodiesel is a monoalkyl ester of fatty acids from vegetableoils. These oils consist of triglycerides in which 3 fatty acidmolecules are esterified with a molecule of glycerol. Whenbiodiesel is made, triglycerides are reacted with short-chainalcohols, primarily methanol and ethanol. This reaction isknown as transesterification or alcoholysis. Transesterifica-tion produces monoalkyl ester of fatty acids and glycerol(Figure 7) [4]. However, this reaction is not sustainablebecause the cost and energy of this process is higher, andcheaper vegetable oil feedstocks for biodiesel are scarce.Therefore, there are 3 major investigations in biodieselproduction: the use of lipase with whole cell biocatalyststechnology, biodiesel production from microalgae, and theuse of a metabolically engineered strain of E. coli that canproduce FAEE.


Whereas transesterification using alkali-catalysis iswidely utilized in many countries for biodiesel production,enzymatic transesterification using lipase has been attractingmuch attention because of reducing processes in biodieselproduction and an easy separation of the glycerol byproduct.However, the high cost of lipase production is the mainobstacle to this lipase reaction. The use of whole cellbiocatalysts immobilized within biomass support particlescan have a potential for reducing the cost [53].

Biodiesel was believed to be produced from microalgae.Unlike other oil feedstocks, microalgae grow extremelyrapidly, and their oil productivity greatly exceeds that ofthe best-producing oil crop. These characteristics makemicroalgae a desired organism for biodiesel production [54].

To produce FAEE from E. coli, the ethanol pathwayfrom Zymomonas mobilis pyruvate decarboxylase (pdc) andalcohol dehydrogenase (adhB), and the unspecific acyltrans-ferase (atfA) from Acinetobacter baylyi, are required. Ethanolproduction was combined with esterification of ethanol withthe acyl moieties of coenzyme A thioesters from fatty acidsin the supplied glucose and oleic acid. The engineered strainproduced 1.28 g/L of FAEE under aerobic conditions in thepresence of glucose and oleic acid by fed-batch fermentation[55]. In addition, there are other studies on engineered E.coli with the ability to produce FAEE. The engineered E.coli strain produced FAEE directly from glucose and ethanol,from only glucose, and from glucose and xylan [56]. Thisstrategy of synthetic biology may improve industrial effortsto produce a variety of diesel-type fuels.

In the same way, heterologous production of the bifunc-tional wax ester synthase/acyl-coenzyme A:diacylglycerolacyltransferase (WS/DGAT) from A. calcoaceticus ADP1 in S.cerevisiae resulted in the production of FAEEs and fatty acidisoamyl ester [57].Other studies have also been performedin oleaginous yeasts such as Rhodosporidium toruloides toproduce microbial lipids [58, 59].

5. Conclusion

To realize a sustainable society, the use of biofuels isneeded to reduce the dependence on fossil fuels as energysources. One of the most ideal systems to effectively producebioethanol is the CBP system. Yeast cell surface engineeringtechnology is a very promising method for the constructionof CBP. In future studies, cell surface engineering willbe a more valuable molecular tool for the degradationof cellulosic materials and the production of all biofuelsincluding bioethanol, biobutanol, and biodiesel at highefficiency and low cost.

Moreover, metabolic engineering and synthetic biologyare important tools for the construction of engineered hostfermentation systems, such as biobutanol and biodieselproduction, although further improvements are requiredbecause of their low productivity and low tolerance ofproducts. In addition, the combination of cell surfaceengineering and metabolic engineering will create moreadvantageous microorganisms for the lower-cost and lessintensive production of biofuels.

References

[1] A. L. Demain, “Biosolutions to the energy problem,” Journalof Industrial Microbiology and Biotechnology, vol. 36, no. 3, pp.319–332, 2009.

[2] M. Inui, M. Suda, S. Kimura et al., “Expression of Clostridiumacetobutylicum butanol synthetic genes in Escherichia coli,”Applied Microbiology and Biotechnology, vol. 77, no. 6, pp.1305–1316, 2008.

[3] D. Antoni, V. V. Zverlov, and W. H. Schwarz, “Biofuels frommicrobes,” Applied Microbiology and Biotechnology, vol. 77, no.1, pp. 23–35, 2007.

[4] F. Ma and M. A. Hanna, “Biodiesel production: a review,”Bioresource Technology, vol. 70, no. 1, pp. 1–15, 1999.

[5] M. R. Connor and J. C. Liao, “Microbial production ofadvanced transportation fuels in non-natural hosts,” CurrentOpinion in Biotechnology, vol. 20, no. 3, pp. 307–315, 2009.

[6] K. A. Gray, L. Zhao, and M. Emptage, “Bioethanol,” CurrentOpinion in Chemical Biology, vol. 10, no. 2, pp. 141–146, 2006.

[7] L. R. Lynd, W. H. Van Zyl, J. E. McBride, and M. Laser,“Consolidated bioprocessing of cellulosic biomass: an update,”Current Opinion in Biotechnology, vol. 16, no. 5, pp. 577–583,2005.

[8] T. Murai, M. Ueda, M. Yamamura et al., “Construction ofa starch-utilizing yeast by cell surface engineering,” Appliedand Environmental Microbiology, vol. 63, no. 4, pp. 1362–1366,1997.

[9] M. Ueda and A. Tanaka, “Genetic immobilization of proteinson the yeast cell surface,” Biotechnology Advances, vol. 18, no.2, pp. 121–140, 2000.

[10] M. Ueda and A. Tanaka, “Cell surface engineering of yeast:construction of arming yeast with biocatalyst,” Journal ofBioscience and Bioengineering, vol. 90, no. 2, pp. 125–136,2000.

[11] A. Kondo and M. Ueda, “Yeast cell-surface display—applications of molecular display,” Applied Microbiology andBiotechnology, vol. 64, no. 1, pp. 28–40, 2004.

[12] M. Ueda, “Future direction of molecular display by yeast-cellsurface engineering,” Journal of Molecular Catalysis B, vol. 28,no. 4–6, pp. 139–143, 2004.

[13] T. Murai, M. Ueda, Y. Shibasaki et al., “Development of anarming yeast strain for efficient utilization of starch by co-display of sequential amylolytic enzymes on the cell surface,”Applied Microbiology and Biotechnology, vol. 51, no. 1, pp. 65–70, 1999.

[14] Y. Fujita, J. Ito, M. Ueda, H. Fukuda, and A. Kondo, “Syn-ergistic saccharification, and direct fermentation to ethanol,of amorphous cellulose by use of an engineered yeast straincodisplaying three types of cellulolytic enzyme,” Applied andEnvironmental Microbiology, vol. 70, no. 2, pp. 1207–1212,2004.

[15] S. Katahira, Y. Fujita, A. Mizuike, H. Fukuda, and A. Kondo,“Construction of a xylan-fermenting yeast strain throughcodisplay of xylanolytic enzymes on the surface of xylose-utilizing Saccharomyces cerevisiae cells,” Applied and Environ-mental Microbiology, vol. 70, no. 9, pp. 5407–5414, 2004.

[16] M. Kuyper, H. R. Harhangi, A. K. Stave et al., “High-levelfunctional expression of a fungal xylose isomerase: the keyto efficient ethanolic fermentation of xylose by Saccharomycescerevisiae?” FEMS Yeast Research, vol. 4, no. 1, pp. 69–78, 2003.

[17] A. Ramanavicius and A. Ramanaviciene, “Hemoproteins indesign of biofuel cells,” Fuel Cells, vol. 9, no. 1, pp. 25–36, 2009.

[18] D. Bhatnagar, S. Xu, C. Fischer, R. L. Arechederra, and S. D.Minteer, “Mitochondrial biofuel cells: expanding fuel diversity


to amino acids,” Physical Chemistry Chemical Physics, vol. 13,no. 1, pp. 86–92, 2011.

[19] A. Ramanavicius, A. Kausaite, and A. Ramanaviciene, “Enzy-matic biofuel cell based on anode and cathode powered byethanol,” Biosensors and Bioelectronics, vol. 24, no. 4, pp. 761–766, 2008.

[20] P. Durre, “Biobutanol: an attractive biofuel,” BiotechnologyJournal, vol. 2, no. 12, pp. 1525–1534, 2007.

[21] S. Atsumi, A. F. Cann, M. R. Connor et al., “Metabolicengineering of Escherichia coli for 1-butanol production,”Metabolic Engineering, vol. 10, no. 6, pp. 305–311, 2008.

[22] S. K. Lee, H. Chou, T. S. Ham, T. S. Lee, and J. D. Keasling,“Metabolic engineering of microorganisms for biofuels pro-duction: from bugs to synthetic biology to fuels,” CurrentOpinion in Biotechnology, vol. 19, no. 6, pp. 556–563, 2008.

[23] D. T. Jones and D. R. Woods, “Acetone-butanol fermentationrevisited,” Microbiological Reviews, vol. 50, no. 4, pp. 484–524,1986.

[24] Y. Jiang, C. Xu, F. Dong, Y. Yang, W. Jiang, and S. Yang,“Disruption of the acetoacetate decarboxylase gene in solvent-producing Clostridium acetobutylicum increases the butanolratio,” Metabolic Engineering, vol. 11, no. 4-5, pp. 284–291,2009.

[25] T. C. Ezeji, N. Qureshi, and H. P. Blaschek, “Acetone butanolethanol (ABE) production from concentrated substrate:reduction in substrate inhibition by fed-batch technique andproduct inhibition by gas stripping,” Applied Microbiology andBiotechnology, vol. 63, no. 6, pp. 653–658, 2004.

[26] V. H. Thang, K. Kanda, and G. Kobayashi, “Productionof acetone-butanol-ethanol (ABE) in direct fermentationof cassava by Clostridium saccharoperbutylacetonicum N1-4,”Applied Biochemistry and Biotechnology, pp. 1–14, 2009.

[27] P. Durre, “Fermentative butanol production: bulk chemicaland biofuel,” Annals of the New York Academy of Sciences, vol.1125, pp. 353–362, 2008.

[28] Y. N. Zheng, L. Z. Li, MO. Xian et al., “Problems withthe microbial production of butanol,” Journal of IndustrialMicrobiology and Biotechnology, vol. 36, no. 9, pp. 1127–1138,2009.

[29] A. Ravagnani, K. C. B. Jennert, E. Steiner et al., “Spo0A directlycontrols the switch from acid to solvent production in solvent-forming clostridia,” Molecular Microbiology, vol. 37, no. 5, pp.1172–1185, 2000.

[30] P. Durre and C. Hollergschwandner, “Initiation of endosporeformation in Clostridium acetobutylicum,” Anaerobe, vol. 10,no. 2, pp. 69–74, 2004.

[31] Y. Gu, S. Hu, J. Chen et al., “Ammonium acetate enhancessolvent production by Clostridium acetobutylicum EA 2018using cassava as a fermentation medium,” Journal of IndustrialMicrobiology and Biotechnology, vol. 36, no. 9, pp. 1225–1232,2009.

[32] N. Qureshi, B. C. Saha, and M. A. Cotta, “Butanol productionfrom wheat straw hydrolysate using Clostridium beijerinckii,”Bioprocess and Biosystems Engineering, vol. 30, no. 6, pp. 419–427, 2007.

[33] T. C. Ezeji, N. Qureshi, and H. P. Blaschek, “Production ofacetone butanol (AB) from liquefied corn starch, a commercialsubstrate, using Clostridium beijerinckii coupled with productrecovery by gas stripping,” Journal of Industrial Microbiologyand Biotechnology, vol. 34, no. 12, pp. 771–777, 2007.

[34] P. J. Evans and H. Y. Wang, “Enhancement of butanol forma-tion by Clostridium acetobutylicum in the presence of decanol-oleyl alcohol mixed extractants,” Applied and EnvironmentalMicrobiology, vol. 54, no. 7, pp. 1662–1667, 1988.

[35] T. C. Ezeji, N. Qureshi, and H. P. Blaschek, “Bioproductionof butanol from biomass: from genes to bioreactors,” CurrentOpinion in Biotechnology, vol. 18, no. 3, pp. 220–227, 2007.

[36] T. C. Ezeji, N. Qureshi, and H. P. Blaschek, “Butanol fermen-tation research: upstream and downstream manipulations,”Chemical Record, vol. 4, no. 5, pp. 305–314, 2004.

[37] Z. L. Boynton, G. N. Bennett, and F. B. Rudolph, “Cloning,sequencing, and expression of clustered genes encodingβ-hydroxybutyryl-coenzyme A (CoA) dehydrogenase, cro-tonase, and butyryl-CoA dehydrogenase from Clostridiumacetobutylicum ATCC 824,” Journal of Bacteriology, vol. 178,no. 11, pp. 3015–3024, 1996.

[38] J. Nolling, G. Breton, M. V. Omelchenko et al., “Genomesequence and comparative analysis of the solvent-producingbacterium Clostridium acetobutylicum,” Journal of Bacteriol-ogy, vol. 183, no. 16, pp. 4823–4838, 2001.

[39] S. B. Tummala, S. G. Junne, and E. T. Papoutsakis, “AntisenseRNA downregulation of coenzyme A transferase combinedwith alcohol-aldehyde dehydrogenase overexpression leadsto predominantly alcohologenic Clostridium acetobutylicumfermentations,” Journal of Bacteriology, vol. 185, no. 12, pp.3644–3653, 2003.

[40] C. A. Tomas, N. E. Welker, and E. T. Papoutsakis, “Over-expression of groESL in Clostridium acetobutylicum resultsin increased solvent production and tolerance, prolongedmetabolism, and changes in the cell’s transcriptional pro-gram,” Applied and Environmental Microbiology, vol. 69, no.8, pp. 4951–4965, 2003.

[41] F. Narberhaus, K. Giebeler, and H. Bahl, “Molecular character-ization of the dnaK gene region of Clostridium acetobutylicum,including grpE, dnaJ, and a new heat shock gene,” Journal ofBacteriology, vol. 174, no. 10, pp. 3290–3299, 1992.

[42] L. M. Harris, R. P. Desai, N. E. Welker, and E. T. Papoutsakis,“Characterization of recombinant strains of the Clostridiumacetobutylicum butyrate kinase inactivation mutant: Needfor new phenomenological models for solventogenesis andbutanol inhibition?” Biotechnology and Bioengineering, vol. 67,no. 1, pp. 1–11, 2000.

[43] L. M. Harris, L. Blank, R. P. Desai, N. E. Welker, and E. T.Papoutsakis, “Fermentation characterization and flux analysisof recombinant strains of Clostridium acetobutylicum with aninactivated soIR gene,” Journal of Industrial Microbiology andBiotechnology, vol. 27, no. 5, pp. 322–328, 2001.

[44] H. Huang, H. Liu, and Y. R. Gan, “Genetic modification ofcritical enzymes and involved genes in butanol biosynthesisfrom biomass,” Biotechnology Advances, vol. 28, no. 5, pp. 651–657, 2010.

[45] T. Ezeji, C. Milne, N. D. Price, and H. P. Blaschek, “Achieve-ments and perspectives to overcome the poor solvent resis-tance in acetone and butanol-producing microorganisms,”Applied Microbiology and Biotechnology, vol. 85, no. 6, pp.1697–1712, 2010.

[46] E. J. Steen, R. Chan, N. Prasad et al., “Metabolic engineeringof Saccharomyces cerevisiae for the production of n-butanol,”Microbial Cell Factories, vol. 7, Article ID 36, 2008.

[47] E. P. Knoshaug and M. Zhang, “Butanol tolerance ina selection of microorganisms,” Applied Biochemistry andBiotechnology, vol. 153, no. 1–3, pp. 13–20, 2009.

[48] O. V. Berezina, N. V. Zakharova, A. Brandt, S. V. Yarotsky, W.H. Schwarz, and V. V. Zverlov, “Reconstructing the clostridialn-butanol metabolic pathway in Lactobacillus brevis,” AppliedMicrobiology and Biotechnology, vol. 87, no. 2, pp. 635–646,2010.


[49] S. Atsumi, T. Hanai, and J. C. Liao, “Non-fermentativepathways for synthesis of branched-chain higher alcohols asbiofuels,” Nature, vol. 451, no. 7174, pp. 86–89, 2008.

[50] S. Atsumi and J. C. Liao, “Metabolic engineering for advancedbiofuels production from Escherichia coli,” Current Opinion inBiotechnology, vol. 19, no. 5, pp. 414–419, 2008.

[51] C. R. Shen and J. C. Liao, “Metabolic engineering ofEscherichia coli for 1-butanol and 1-propanol production viathe keto-acid pathways,” Metabolic Engineering, vol. 10, no. 6,pp. 312–320, 2008.

[52] S. Atsumi, T. Y. Wu, E. M. Eckl, S. D. Hawkins, T. Buelter,and J. C. Liao, “Engineering the isobutanol biosyntheticpathway in Escherichia coli by comparison of three aldehydereductase/alcohol dehydrogenase genes,” Applied Microbiologyand Biotechnology, vol. 85, no. 3, pp. 651–657, 2010.

[53] H. Fukuda, A. Kondo, and H. Noda, “Biodiesel fuel produc-tion by transesterification of oils,” Journal of Bioscience andBioengineering, vol. 92, no. 5, pp. 405–416, 2001.

[54] Y. Chisti, “Biodiesel from microalgae,” Biotechnology Advances,vol. 25, no. 3, pp. 294–306, 2007.

[55] R. Kalscheuer, T. Stolting, and A. Steinbuchel, “Microdiesel:Escherichia coli engineered for fuel production,” Microbiology,vol. 152, no. 9, pp. 2529–2536, 2006.

[56] E. J. Steen, Y. Kang, G. Bokinsky et al., “Microbial productionof fatty-acid-derived fuels and chemicals from plant biomass,”Nature, vol. 463, no. 7280, pp. 559–562, 2010.

[57] R. Kalscheuer, H. Luftmann, and A. Steinbuchel, “Synthesisof novel lipids in Saccharomyces cerevisiae by heterologousexpression of an unspecific bacterial acyltransferase,” Appliedand Environmental Microbiology, vol. 70, no. 12, pp. 7119–7125, 2004.

[58] Q. Li, W. Du, and D. Liu, “Perspectives of microbial oils forbiodiesel production,” Applied Microbiology and Biotechnol-ogy, vol. 80, no. 5, pp. 749–756, 2008.

[59] X. Zhao, C. Hu, S. Wu, H. Shen, and Z. K. Zhao, “Lipidproduction by Rhodosporidium toruloides Y4 using differentsubstrate feeding strategies,” Journal of Industrial Microbiologyand Biotechnology. In press.


Research Article

Rapeseed Oil Monoester of Ethylene Glycol Monomethyl Ether asa New Biodiesel

Jiang Dayong, Wang Xuanjun, Liu Shuguang, and Guo Hejun

Xi’an Research Institute of High Technology, Xi’an, Shaanxi 710025, China

Correspondence should be addressed to Jiang Dayong, [email protected]

Received 19 July 2010; Revised 8 December 2010; Accepted 3 January 2011

Academic Editor: R. S. Tjeerdema

Copyright © 2011 Jiang Dayong et al. This is an open access article distributed under the Creative Commons Attribution License,which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

A novel biodiesel named rapeseed oil monoester of ethylene glycol monomethyl ether is developed. This fuel has one more estergroup than the traditional biodiesel. The fuel was synthesized and structurally identified through FT-IR and P1PH NMR analyses.Engine test results show that when a tested diesel engine is fueled with this biodiesel in place of 0# diesel fuel, engine-out smokeemissions can be decreased by 25.0%–75.0%, CO emissions can be reduced by 50.0%, and unburned HC emissions are lessenedsignificantly. However, NOx emissions generally do not change noticeably. In the area of combustion performance, both enginein-cylinder pressure and its changing rate with crankshaft angle are increased to some extent. Rapeseed oil monoester of ethyleneglycol monomethyl ether has a much higher cetane number and shorter ignition delay, leading to autoignition 1.1◦CA earlierthan diesel fuel during engine operation. Because of certain amount of oxygen contained in the new biodiesel, the engine thermalefficiency is improved 13.5%–20.4% when fueled with the biodiesel compared with diesel fuel.

1. Introduction

In recent years, growing awareness of the complete depletionof petroleum oil in the near future and serious atmosphericpollution caused by automobile industry has inspired muchresearch for clean alternative fuels to substitute for fossil fuels[1–3]. One of the most promising alternative energy sourcesis biodiesel. Biodiesel contains significantly less sulfur andnitrogen, which makes the fuel more environment-friendlythan petroleum fuels. Because it is renewable and availableworldwide, it has a bright future for practical application.A traditional biodiesel used to be the methyl ester ofvegetable oil, which is prepared through transesterificationof vegetable oils such as rapeseed oil, peanut oil, and soybeanoil with methanol. Many studies have revealed that suchbiodiesel, containing a certain amount of oxygen, can leadto remarkable reduction in diesel engine exhaust emissions[4–6]. Thus, it has been called a green fuel for diesel engine.However, because it has only one ester group (i.e., twooxygen atoms, existing in each monoester molecule), theoxygen content in conventional biodiesel is comparativelylow. Consequently, the reduction in smoke emissions is notas significant as anticipated used in diesel engines when

mixed with diesel fuel. Experiments have shown that thereduction rate in engine-out smoke emissions is correlatedwith the oxygen content of the fuel. To enhance the effect oftraditional biodiesel in reducing engine-out smoke forma-tion, the introduction of other ether group into its moleculehas been attempted. Rapes are largely planted in China,and rapeseed oil is a vegetable oil extensively used for food.Therefore, a new type of biodiesel, rapeseed oil monoesterof ethylene glycol monomethyl ether, is synthesized in thispaper. This study focuses on its performance in reducingengine-out exhaust emissions and combustion.

2. Experimental Section

2.1. Preparation and Structure Analysis. The new rapeseedoil monoester was synthesized using a commercial refinedrapeseed oil and ethylene glycol monomethyl ether as reac-tants. Initially, the selected rapeseed oil was treated throughextraction with ethanol as solvent at a temperature of 90◦Cto remove tiny amounts of organic fatty acid. It was thenpurified under vacuum condition. FT-IR analysis verifiedthat there was no vibration (–OH) absorption peak in the


721.41

1097.55

1160.02

1375.86

1458.57

1743.42

2852.42

2921.93

3006.46

50556065707580859095

100105110115120125130

4000 3500 3000 2500 2000 1500 1000 500

Wavenumbers (cm−1)

Refl

ecta

nce

(%)

Figure 1: IR spectrum of rapeseed oil after pretreatment.

Table 1: Specification of the used chemicals.

ChemicalsDensity/kg·m−3

Boilingpoint /◦C

Molecularweight

Standard

Rapeseed oil 909–914 — — primes state

EthyleneglycolMonomethylether

965.0 134 76.10analyticallypure

Ethanolabsolute

789–791 78 46.07analyticallypure

Potassa 950 — 23.5chemicallypure

region of 3200–3600 cm−1, indicating that there was littlefatty acid and ethanol in the treated rapeseed oil (Figure 1,Table 1).

The subsequent transesterification reaction was carriedout in a flask with the acid-free rapeseed oil of 600 mLand ethylene glycol monomethyl ether of 210 mL at atemperature of 65◦C using 0.6% KOH as catalys,. Uponcompletion of the reaction which lasted for approximately2 h, the crude product was first neutralized with diluted HClsolution and separated from the water phase. Subsequently,it was purified in a vacuum to remove ethylene glycolmonomethyl ether left over in the ester phase. Finally, it wasdried using CaCl2 agent [7].

Gel permeation chromatography (GPC) can help simul-taneously detect diglyceride, triglyceride, glycerol, and fattyacid content of methyl ester in the process of transesterifi-cation reaction with a refractive index detector. The maincomponent of rapeseed oil is fatty acid glycerin ester, whosenumber average molecular weight is approximately 1426or so. This is consistent with the known composition ofrapeseed oil. After transesterification, the peak of fatty acidester (Mw 1426) had almost disappeared, which means fattyacid glycerides have been transformed into fatty acid etherester (biodiesel) thoroughly. Correspondingly, the peaks ofnumber average molecular weight of 564 and 338 haveappeared (Figure 2, Table 2).

Table 2: GPC Data of rapeseed oil monoester.

Peak Mn MP RT/minArea/mVsecs

Peakarea/%

Rapeseedoil

I 2713 2649 23.363 4041 0.55

II 1426 1382 24.921 696199 95.25

III 1005 1029 25.622 9767 1.34

IV 835 893 25.931 9626 1.32

V 568 573 26.731 7390 1.01

VI 342 350 27.364 3920 0.54

RapeseedoilMonoester

I 1105 1038 25.604 176060 16.29

II 564 525 26.858 365764 33.83

III 338 337 27.405 539238 49.88

Table 3: Specification of DI diesel engine utilized.

Parameter Magnitude Parameter Magnitude

Bore 100 mm Rated speed 2300 rpm

Stroke 115 mm Rated power 11 kw

Connectingrod length

190 cmCombustion

chamberω shape

Displacement 0.903 LCompression

ratio18

2.2. Engine Test. A single cylinder, four-stroke, water-cooled,DI diesel engine was adapted to complete the determinationof exhaust emissions and combustion performances. Thetechnical parameters of the engine are presented in Table 3.An AVL DiSmoke 4000 smoke opacity indicator was used torecord smoke number in extinction coefficient, and an on-line exhaust emission analyzer was utilized to examine NOx,CO, and HC emitted. An angle calibration apparatus and akistler pressure transducer were used to pick up crankshaftangle and in-cylinder pressure. A CS20000 Data Gatheringand Analyzing System was utilized to process data [8].

For comparison study, a 0# diesel fuel meeting Chinanational technical specification was utilized and named B0.B0 was mixed with the rapeseed oil (B100) monoester ina proportion of 1 : 1 by volume to investigate effect ofthe mixture (B50) on engine-out exhaust emissions andcombustion performance [9–12].

The engine tests were carried out under the followingconditions: ambient temperature of 23◦C, humidity of 71%,and engine water-cooling temperature of 70◦C. In theexperiment, when the engine achieved stable operation at afixed steady state, of the necessary determinations were madeaccording to certain well-defined procedures.


3.1. Chemical Structure. Figure 3 and Table 4 list the mainabsorption frequencies displayed in IR spectrum obtainedfor the new rapeseed oil monoester. No absorption peaksabove 3100 cm−1 was found, implying that there is nohydroxyl group (–OH) in the molecules of the synthesizedproduct. So, it is easily confirmed that the product is an ester


2649

1029

1382

893

573

350

−505

1015202530

20 21 22 23 24 25 26 27 28 29

(min)

MV

(a) rapeseed oil

1038

525

337

−505

1015202530

20 21 22 23 24 25 26 27 28 29

(min)

MV

(b) rapeseed oil monoester

Figure 2: GPC spectrum of rapeseed oil and rapeseed oil monoester.

3461

.41

3005

.88

2925

.71

2864

.56

1740

.36

1461

.59

1376

.49

1244

.91

1174

.62

1130

.69

1038

.43

8633

7

7229

0

30

40

50

60

70

80

4000 3500 3000 2500 2000 1500 1000 500

Wavenumbers (cm−1)

Tran

smit

ten

ce(%

)

Figure 3: IR spectrum of rapeseed oil monoester.

7.27

4

5.36

5.34

25.

324

4.24

24.

226

4.21

3.60

73.

591

3.57

63.

391

2.36

52.

342.

315

2.01

81.

999

1.64

81.

626

1.60

31.

31.

255

1.22

0.9

0.88

0.85

8

0

1

1.81

22.

98

2.66

2.24

2.45

27.7

3.04

0.21

7 6 5 4 3 2 1 0

ppm (t1)

Figure 4: P1PH NMR spectrum of rapeseed oil monoester.

involving an ether group. Figure 4 and Table 5 illustrateits P1PH NMR data collected. Chemical shift 5.359 ppmbelonged to the protons attaching to the C=C group inthe rapeseed oil monoester molecules, and chemical shiftof 4.228, 3.596, and 3.395 ppm, respectively, belonged tothe protons existing in the group –COOCH2CH2OCH3 inthe order from left to right. The peaks occurring in thechemical shift region below 2.771 ppm all were attributed toprotons in the fatty groups (–R) and other non-fatty estercompounds. Therefore, the chemical structure of the new

Table 4: Data of FT-IR spectrum of rapeseed oil monoester.

Frequency/cm−1 Group attribution Vibration type Strength

2925.71 –CHB3B, –CHB2B νBasB s

2854.56 –CHB3B, –CHB2B νBsB s

1740.36 C=O ν s

1461.59 –CHB2B δ m

1376.49 –CHB3B δ m

1244.91 C–O–C νBasB m

1174.62 C–O–C νBsB m

722.90 (CH2) n(n > 4) – w

prepared rapeseed oil monoester was easily confirmed asRCOOCH2CH2OCH3.

3.2. Exhaust Emissions. Two types of engine operation modesrunning at 1400 and 2000 rpm, respectively, were selectedto study the exhaust emissions changes under differentbrake mean effective pressures (BMEP). Figure 5 displaysthe effects of the new rapeseed oil monoester on engine-out smoke emissions at 1400 and 2000 rpm, respectively.Clearly, a considerable decrease in smoke emissions hasbeen achieved in the experiment. When the engine burntB100 under partial loads at 1400 rpm, there was a relativereduction in smoke emissions by 25.0%–56.5% within thetested partial load range compared with B0. A decreaseof 25.9%–75.0% was also obtained for B50. At 2000 rpm,reductions of 43.5%–72.2% and of 25.0%–38.8% werereached, respectively, for B100 and B50 within the testedpartial load range. As an oxygenated fuel containing morethan 10% oxygen, the molecule of biodiesel does not containaromatics. In addition, its hydrocarbon ratio (C/H) is farless than that of saturated hydrocarbons. Oxygen capacitycan be improved through the issue of local hypoxia andsmoke mainly generated in the diffusion combustion. Thus,the accession of biodiesel can increase premixed combustion,and decrease diffusion combustion, especially in areas of highfuel concentration, reducing the oxygen fuel combustion,and thereby reducing the soot emissions.

Figure 6 reveals that CO emissions reduced noticeably atthe speed of 1400 and 2000 rpm. At 1400 rpm, CO emissionswere lessened by 25.0%–33.3% when the tested engineburned B100; a slightly larger decline occurred for B50


0

0.2

0.4

0.6

0.8

1

0.2 0.4 0.6 0.8

BMEP (MPa)

Ext

inct

ion

coeffi

cien

t(1

/m)

B0B50B100

(a) 1400 rpm

0

0.2

0.4

0.6

0.8

1

1.2

0.2 0.4 0.6 0.8

BMEP (MPa)E

xtin

ctio

nco

effici

ent

(1/m

)

B0B50B100

(b) 2000 rpm

Figure 5: Effect of the rapeseed oil monoester on smoke emissions.

0

0.1

0.2

0.3

0.2 0.4 0.6 0.8

BMEP (MPa)

CO

(%(v

))

B0B50B100

(a) 1400 rpm

0

0.1

0.2

0.3

0.2 0.4 0.6 0.8

BMEP (MPa)

CO

(%(v

))

B0B50B100

(b) 2000 rpm

Figure 6: Effect of the rapeseed oil monoester on CO emissions.


0

400

800

1200

1600

2000

0.2 0.4 0.6 0.8

BMEP (MPa)

NO

x(p

pm)

B0B50B100

(a) 1400 rpm

0

300

600

900

1200

1500

0.2 0.4 0.6 0.8

BMEP (MPa)

NO

x(p

pm)

B0B50B100

(b) 2000 rpm

Figure 7: Effect of the rapeseed oil monoester on NOx emissions.

Table 5: P1PH NMR data of rapeseed oil monoester.

Chemicalshift/ppm

Proton peaksplitting

Couplingconstant/Hz

Peakarea/proton

number

5.359 quartet 6.4 1.001/—

4.228 triplet 3.6 0.78/2

3.596 triplet 3.6 0.83/2

3.395 singlet — 0.99/3

<2.771 more — more

with B0. Similar to the changing patterns at 2000 rpm, COemissions were lowered by 16.0%–50.0% when the enginecombusted B100; a slightly larger decline also emerged forB50 with diesel fuel. Because of the low calorific valueof biodiesel at the high load and the low temperature incylinder, combustion is not sufficient. Another explanationfor the increasing CO emissions may be that oil-rich areasmay be formed when the combustion begins to deteriorate,causing the CO emissions to increase gradually.

Figure 7 exhibits the results of NOx under differentBMEP at 1400 and 2000 rpm, respectively. Clearly, NOxemissions did not change noticeably in general. This isdue to more adequate oxygen of the mixture at the smallload for diesel engines, and the lower temperature in thecombustion chamber causing lesser NOx emissions andlower gas temperature. Biodiesel has high cetane numberand short ignition delay; however, there is a trend toreduce NOx emissions. With increased combustion, the

maximum temperature also increases in the combustionchamber, which is a major factor facilitating an increase inthe production and emission of NOx. As a result, the NOx ofbiodiesel is slightly less than that of diesel at 2200 rpm.

The effect of the rapeseed oil monoester on unburnedHC emissions at 1400 and 2000 rpm were also investigatedin the experiment. HC emissions at levels from 8–13 ppmwhen the engine was fueled with the diesel fuel droppedto 1–3 ppm when fueled with the blend. However, whenthe engine burned B100, unburned HC increased to thesame levels equal to that of the pure diesel fuel. The changefluctuated with BMEP increase. This may be attributed to thelower heating value of biodiesel when mixed with diesel fuel.The ignition delay was decreased, resulting in the reductionof heat in the combustion process. Thus, the temperature ofgas starts to drop increasingly, causing the quenching layerto thicken in chamber. Because the fuel of quenching layerdoes not vaporize, a greater amount of unburned HC doesnot participate in the combustion as the quenching layerthickens. When the amount of unburned HC becomes morethan the amount of decreased part of HC due to the extraoxygen from oxygenated fuels, HC emissions increase.

3.3. Combustion Performances. Figure 8 displays the testresults of in-cylinder pressure when the tested diesel engineburnt three fuels at 1400 and 2000 rpm. Under BMEP of0.14 MPa and speed of 1400 rpm engine in-cylinder pressurewas noticeably enhanced. This occurred when the engine wasfueled with B100 in place of the reference diesel fuel. Thein-cylinder peak pressure was raised by approximately 2.0%


0

1

2

3

4

5

6

7

8

9

P(M

Pa)

−20 −10 0 10 20 30 40

Crank angle (◦CA)

(a) BMEP = 0.14 MPa (1400 rpm)

0

1

2

3

4

5

6

7

8

9

10

P(M

Pa)

−20 −10 0 10 20 30 40

Crank angle (◦CA)

B0B50B100

(b) BMEP = 0.70 MPa (2000 rpm)

Figure 8: Diesel engine in-cylinder pressure when burning differentfuels.

when burning B100. The peak pressure was also observed torise when combusting B50. The engine ignition delay clearlybecame shorter when the biodiesel was substituted with B0.The new biodiesel spontaneously ignited at approximately1.1◦CA earlier than B0. Under BMEP of 0.70 MPa andspeed of 2000 rpm, these remarkable changes also occuredobviously, as shown distinctly in the figure. The engineignition delay was 1.5◦CA shorter than that of diesel fuel.In-cylinder peak pressure also increased by approximately6.9% when fueled with the new biodiesel. This is becauseB100 with the high oxygen content has a greater amount ofpremixed combustion than B0, which means its heat releasedis so concentrated that the maximum cylinder pressurerises.

Figure 9 exhibits the change rates of engine in-cylinderpressure with the crank angle at 1400 and 2000 rpm, respec-tively. The two graphs clearly show that in-cylinder pressurechange rate increased slightly under the engine running

−0.6

−0.4

−0.2

0

0.2

0.4

0.6

0.8

1

1.2

Pre

ssu

reri

se(k

J/◦ C

A)

−20 −10 0 10 20 30 40

Crank angle (◦CA)

(a) BMEP = 0.14 MPa (1400 rpm)

−0.6

−0.4

−0.2

0

0.2

0.4

0.6

0.8

1

1.2

1.4

Pre

ssu

reri

se(J

/◦C

A)

−20 −10 0 10 20 30 40

Crank angle (◦CA)

B0B50B100

(b) BMEP = 0.70 MPa (2000 rpm)

Figure 9: Diesel engine in-cylinder pressure changing rate whenburning different fuels.

modes (a) and (b), when the diesel engine combusted B100.This means that the combustion rate of the new biodieselin diesel engine increased and its combustion duration wasshorter than that of diesel fuel. The two figures also distinctlyreveal that the new biodiesel began to burn earlier than thediesel fuel.

Figure 10 demonstrates the heat release rates of the dieselengine when it was fueled with the biodiesel, diesel fuel, anda mixture of both. Under two modes, the heat release rateincreased when the biodiesel replaced the reference dieselfuel during the experiment. At the mode (a), no changewas observed in the heat release rate. This indicates thatthe combustion velocity of the new rapeseed oil monoesterin diesel engine combustion chamber was much higherthan diesel fuel, likely because the oxygen contained in thenew biodiesel is able to accelerate the engine combustion.The curves in the two figures also clearly show that thenew biodiesel began to burn earlier than diesel fuel. Thisimplies that the cetane number of the new biodiesel ishigher than that of diesel fuel with CN45 to CN 50


−20

0

20

40

60

80

100

120

140

160

180

Hea

tre

leas

era

te(J

/◦C

A)

−20 −10 0 10 20 30 40

Crank angle (◦CA)

(a) BMEP = 0.14 MPa (1400 rpm)

−20

0

20

40

60

80

100

120

140

160

180

Hea

tre

leas

era

te(k

J/◦ C

A)

−20 −10 0 10 20 30 40

Crank angle (◦CA)

B0B50B100

(b) BMEP = 0.70 MPa (2000 rpm)

Figure 10: Diesel engine heat release rate when burning differentfuels.

currently used in China. This higher cetane number is likelybecause the introduced ether group in the biodiesel is moreeasily oxidized. The biodiesel also releases more heat at agiven time, raising the temperature, thereby accelerates fueloxidation. The cetane number of the new biodiesel is beingfurther determined in the laboratory.

Figure 11 demonstrates the improved engine thermalefficiency when biodiesel and the biodiesel-diesel blendreplaced the reference diesel fuel in the experiment. Asignificant increase of 13.5%–20.4% in engine thermalefficiency was observed when the tested engine burnt thenew biodiesel in place of diesel fuel at 1400 rpm. A similarresult was attained at 2000 rpm. This may be explained bythe fact that the new biodiesel contains a certain amount ofoxygen, which promotes a more complete combustion of thebiodiesel than diesel fuel.

4. Conclusions

(1) A novel biodiesel named rapeseed oil monoester ofethylene glycol monomethyl ether, which contains

0

10

20

30

40

0 0.2 0.4 0.6 0.8

BMEP (MPa)

η(%

)

(a) 1400 rpm

0

5

10

15

20

25

30

35

40

0 0.2 0.4 0.6 0.8

BMEP (MPa)

η(%

)

B0B50B100

(b) 2000 rpm

Figure 11: Engine brake thermal efficiency when burning differentfuels.

more oxygen than traditional rapeseed oil alkyl ester,has been prepared and structurally identified.

(2) When diesel engine is fueled with this rapeseed oilmonoester and its mixture with diesel fuel in theproportion of 1 : 1 by volume, engine-out exhaustemissions such as smoke and CO can be substantiallyreduced under partial load modes. However, NOxemissions do not change significantly in general.

(3) When diesel engine is fueled with the new biodiesel inplace of diesel fuel, both engine in-cylinder pressureand its changing rate with crankshaft angle increaseto some extent. Biodiesel has a much higher cetanenumber and shorter ignition delay than diesel fuel,and therefore ignites earlier during diesel engineoperation.

(4) The combustion of the new rapeseed oil monoestercan lead to a slightly higher heat release rate thandiesel fuel. Utilization of the new biodiesel canremarkably improve engine brake thermal efficiencybecause of its specific oxygen content.


Acknowledgment

This research was supported by the National Natural ScienceFoundation of China (Grant no. 50976125).

References

[1] R. Crooles, “New findings on combustion behavior of oxy-genated synthetic diesel fuels,” Biomass and Bioenergy, vol. 30,pp. 461–468, 2006.

[2] M. Pugazhvadivu and K. Jeyachandran, “Experimental studiesof the impact of CETANERTM on diesel combustion andemission,” Renewable Energy, vol. 30, pp. 2189–2202, 2005.

[3] G. Kaplan, R. Arslan, and A. Surmen, “Modeling the effectsof oxygenated fuels and split injections on DI diesel engineperformance and emission,” Energy Sources, vol. A28, pp. 751–755, 2006.

[4] N. Usta, E. Ozturk, O. Can, and E. Conkur, “Emissioncharacteristics of a navistar 7.3 L turbo diesel fueled withblends of oxygenates and diesel,” Energy Conversion andManagement, vol. 46, pp. 741–755, 2005.

[5] M. Lapuerta, O. Armas, and R. Ballesteros, “Oxygenated fuelsfor particulate emissions reduction in heavy duty DI dieselengines with common rail fuel injection,” Fernandez Fuel, vol.84, pp. 773–780, 2005.

[6] M. Cetinkaya and F. Karaosmanoglu, “The effect of oxy-genated fuels on emissions from a modern heavy-duty dieselengine,” Energy & Fuels, vol. 19, pp. 645–652, 2007.

[7] D. Y. C. Leung, X. Wu, and M. K. H. Leung, “A reviewon biodiesel production using catalyzed transesterification,”Applied Energy, vol. 87, no. 4, pp. 1083–1095, 2010.

[8] J. Feng, Structure Analysis and Identification of Organic Com-pound, Defense Industry, Beijing, China, 2003.

[9] G. Fontaras, G. Karavalakis, M. Kousoulidou et al., “Effectsof low concentration biodiesel blends application on modernpassenger cars. Part 2: impact on carbonyl compound emis-sions,” Environmental Pollution, vol. 158, pp. 2496–2504, 2010.

[10] X. Wang, Z. Huang, O. A. Kuti, W. Zhang, and K. Nishida,“Experimental and analytical study on biodiesel and dieselspray characteristics under ultra-high injection pressure,”International Journal of Heat and Fluid Flow, vol. 31, no. 4, pp.659–666, 2010.

[11] I. M. Atadashi, M. K. Aroua, and A. A. Aziz, “High qualitybiodiesel and its diesel engine application: a review,” Renew-able and Sustainable Energy Reviews, vol. 14, pp. 1999–2009,2010.

[12] K. T. Tan, M. M. Gui, K. T. Lee, and A. R. Mohamed, “Anoptimized study of methanol and ethanol in supercriticalalcohol technology for biodiesel production,” Journal ofSupercritical Fluids, vol. 53, no. 1–3, pp. 82–87, 2010.


Review Article

Utilization of Biodiesel By-Products for Biogas Production

Nina Kolesarova, Miroslav Hutnan, Igor Bodık, and Viera Spalkova

Institute of Chemical and Environmental Engineering, Faculty of Chemical and Food Technology,Slovak University of Technology, Radlinskeho 9, 812 37 Bratislava, Slovakia

Correspondence should be addressed to Nina Kolesarova, [email protected]

Received 22 August 2010; Revised 22 November 2010; Accepted 7 January 2011

Academic Editor: Leon Spicer

Copyright © 2011 Nina Kolesarova et al. This is an open access article distributed under the Creative Commons AttributionLicense, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properlycited.

This contribution reviews the possibility of using the by-products from biodiesel production as substrates for anaerobic digestionand production of biogas. The process of biodiesel production is predominantly carried out by catalyzed transesterification. Besidesdesired methylesters, this reaction provides also few other products, including crude glycerol, oil-pressed cakes, and washing water.Crude glycerol or g-phase is heavier separate liquid phase, composed mainly by glycerol. A couple of studies have demonstratedthe possibility of biogas production, using g-phase as a single substrate, and it has also shown a great potential as a cosubstrate byanaerobic treatment of different types of organic waste or energy crops. Oil cakes or oil meals are solid residues obtained after oilextraction from the seeds. Another possible by-product is the washing water from raw biodiesel purification, which is an oily andsoapy liquid. All of these materials have been suggested as feasible substrates for anaerobic degradation, although some issues andinhibitory factors have to be considered.

1. Introduction

Renewable energy sources and biofuels, including biodiesel,have been gaining increasing attention recently as a replace-ment for fossil fuels [1]. However, their implementation inthe general market depends on making these fuels morecompetitive. A convenient way to lower the costs of biofuels isto use the by-products as a potential source of energy, ratherthen treat them as waste.

Biodiesel is a prominent candidate as alternative dieselfuel. It is offering few advantages compared to conventionaldiesel, including the status of renewable energy sourceand lower emissions. Advances against petroleum dieselfuel are represented by the terms of sulfur content, flashpoint, content of aromatic substances, and biodegradability[1].

With approximately 245 processing plants and annualproduction of about 9 million tons, European Unionhas had the leading position in both production andconsumption of biodiesel [2]. These plants are mainlylocated in Germany, Italy, Austria, France, and Sweden.Production of biodiesel has been expanding rapidly also onthe other continents, mainly in the USA and developing

countries, such as India, Brazil, Argentina, Malaysia, andFiji.

As a primary feedstock, vegetable oils, animal fats, orwaste cooking oils can be used for the production ofbiodiesel. In Europe, rapeseed oil is predominantly used,while, in the world extent, highest quantities of biodiesel areproduced from soya oil [3].

The process of biodiesel production is usually carriedout by catalyzed transesterification with alcohol, most likelymethanol (Figure 1). A catalyst is usually involved to improvethe reaction rate and yield [3]. Alkalies (sodium hydrox-ide, potassium hydroxide, carbonates, and correspondingsodium and potassium alkoxides), acids (sulfuric acid,sulfonic acid or hydrochloric acid), or enzymes can be usedto catalyze the reaction. Base-catalyzed transesterificationis much faster than the acid-catalyzed one (base catalyzedtransesterif ication is basically finished within one hour) andis most often used commercially [4–6].

Besides the desired methylesters this reaction providesalso few other products. Isolation of oil from the oil seedplants by pressing and extraction provides oil cakes or oilmeal as a by-product. In the reaction of transesterification,triglycerides are converted into glycerol and methylesters


Table 1: Analysis results of macroelements, carbon and nitrogen in crude glycerol from different feedstocks (BDL indicates values that arebelow the detection limit for the corresponding analytical method) [25].

Feed stocks Ida Gold Mustard Pac Gold Mustard Rapeseed Canola Soybean Crambe Waste vegetable oils

Calcium, ppm 11.7 ± 2.9 23.0 ± 1.0 24.0 ± 1.7 19.7 ± 1.5 11.0 ± 0 163.3 ± 11.6 BDL

Potassium, ppm BDL BDL BDL BDL BDL 216.7 ± 15.3 BDL

Magnesium, ppm 3.9 ± 1.0 6.6 ± 0.4 4 ± 0.3 5.4 ± 0.4 6.7 ± 0.2 126.7 ± 5.8 0.4 ± 0

Phosphorus, ppm 25.3 ± 1.2 48 ± 2.0 65 ± 2.0 58.7 ± 6.8 53.0 ± 4.6 136.7 ± 57.7 12.0 ± 1.5

Sulfur, ppm 21.0 ± 2.9 16.0 ± 1.4 21.0 ± 1.0 14.0 ± 1.5 BDL 128.0 ± 7.6 19.0 ± 1.8

Sodium, % wt 1.17 ± 0.15 1.23 ± 0.12 1.06 ± 0.07 1.07 ± 0.12 1.2 ± 0.1 1.10 ± 0.10 1.40 ± 0.16

Carbon, % wt 24.0 ± 0.00 24.3 ± 0.58 25.3 ± 0.58 26.3 ± 0.58 26.0 ± 1 24.0 ± 0.00 37.7 ± 0.58

Nitrogen, % wt 0.04 ± 0.02 0.04 ± 0.01 0.05 ± 0.01 0.05 ± 0.01 0.04 ± 0.03 0.06 ± 0.02 0.12 ± 0.01

R1

R1

R1

R2

R2

R3

O

O

O

O

O

O

OO

O

+ +CH3OH

KOH

OCH3

OCH3

OCH3

HO

OH

OH

Triglyceride Biodiesel

Glycerol

Figure 1: Reaction of biodiesel production by base-catalyzedtransesterification with methanol.

(biodiesel) are separated from the heavier glycerol phase(crude glycerine) by settling. To remove the impurities,soaps, short chain fatty acids and excess methanol, crudebiodiesel is subsequently washed generating the washingwater as another potential by-product.

Although the present production of biodiesel is mainlycarried out by homogeneous base-catalyzed transesterifi-cation, implementation of alternative approaches has beenincreasingly studied [7–24]. Homogeneous base catalysis hasfew disadvantages, such as the high requirement for thepurity of oil. The high consumption of energy and costlyseparation of the homogeneous catalyst from the reactionmixture have also called for development of new catalysts.

Different alternative techniques have been applied forbiodiesel production [7–11]. The most promising onesinclude employing heterogeneous catalyst, lipase catalyst orsupercritical alcohol [12].

Catalysis using solid heterogeneous catalysts runs slowerthan homogeneouscatalysis, however it can be integratedwith continuous processing technologies. A great varietyof catalysts in catalytic transesterification of vegetable oilshave been used recently, including zeolites, hydrotalcites,oxides, and so forth, [13–17]. Utilization of heterogeneouscatalysts provides few advantages over the homogeneousbase catalysis, including mainly the easier purification ofmethylesters from glycerol and impurities. Also, the contentof free fatty acids and water in the raw material does notaffect the reaction [12].

Enzyme catalyzed transesterification using lipases forbiodiesel production is also increasingly studied [18–20].

Similarly to heterogeneous catalysis, it also provides asolution of avoiding difficult recovery of glycerol andmethylesters purification. Although this technology offersan attractive alternative, the industrial application has beenslow due to feasibility aspects and some technical challenges,resulting from the low solubility of methanol and glycerol inbiodiesel and high cost of lipases as catalyst.

Transesterification with supercritical methanol providesseveral advantages, compared to traditional methods [21–24]. The reaction is fast, in addition no catalyst is neededand therefore the separating process of the catalyst andsaponified products becomes unnecessary. Generation ofwashing water can also be avoided. However the highpressure and temperature (239–385◦C) is required, whichleads to high energy consumption and production costs.

Commonly the most important by-products frombiodiesel production are pressed cakes from oil extraction,crude glycerol and washing water [5, 7, 18]. The nature ofthese products is highly dependent on the character of rawmaterial and processing technique, although generally theypresent suitable substrates for anaerobic digestion with theproduction of biogas.

2. Crude Glycerol

Crude glycerol (g-phase) is heavier separate liquid phase, co-mposed mainly by glycerol. In general for every 100 kilo-grams of biodiesel about 10 kilograms of g-phase is pro-duced.

Crude glycerol generated by homogeneous base-cata-lyzed transesterification contains approximately 50–60% ofglycerol, 12–16% of alkalies especially in the form of alkalisoaps and hydroxides, 15–18% of methyl esters, 8–12% ofmethanol, 2-3% of water and further components [25, 26].Tables 1 and 2 summarize the characteristics of g-phasesbased on the source of oil used for the production ofbiodiesel. Analytical results from the macroelement screen-ing tests are listed in Table 1. Crude glycerol contains a vari-ety of elements, such as calcium, magnesium, phosphorus orsulfur, originating from the primary oil. Larger quantities ofsodium or potassium are also contained, coming from thecatalyst.

Table 2 shows the content of protein, fat, ash, carbohy-drates in percents and caloric value for kg. G-phase is mostlycomposed of carbohydrates, represented by glycerol. The ash


Table 2: Food nutrient analysis for crude glycerol samples [25].

Feed stocks Ida Gold Mustard Pac Gold Mustard Rapeseed Canola Soybean Crambe Waste vegetable oils

Fats, % 2.03 1.11 9.74 13.1 7.98 8.08 60.1

Carbohydrates, % 82.8 83.8 75.5 75.2 76.2 78.6 26.9

Protein, % 0.14 0.18 0.07 0.06 0.05 0.44 0.23

Calories, kJ/kg 14.6 14.5 16.3 17.5 15.8 16.3 27.2

Ash, % 2.8 1.9 0.7 0.65 2.73 0.25 5.5

contained in crude glycerol is mainly sodium or potassiumfrom the catalyst.

Considering that the processing technology of biodieselproduction affects the characteristics of by-products, thenew technologies and modern catalysts can be expected toinfluence the composition and utilization of crude glycerol.For example, g-phase originating from biodiesel productionusing rapeseed oil with heterogeneous catalyst is limpid andcolorless, containing at least 98% of glycerin and neither ash,nor inorganic compounds were detected in it [27].

As the biodiesel production is increasing exponentially,the crude glycerol generated in this process has also beengenerated in a large quantity. Despite the wide applicationsof pure glycerol in pharmaceutical, food and cosmeticindustries, the refining of crude glycerol to a high purityis too expensive, especially for small and medium biodieselproducers [28]. The investments for the construction andstartup operation of crude glycerol purification facility makeaccording to Singhabhandhu [29] roughly 65 million Euros(facilities with production capacities of 1.4–2 ML/y). Weber[30] mentions 27% of the capital investment costs goingto construction of the technical glycerin facility in a 12 MLbiodiesel refinery (Aschach, Austria).

To improve the economic feasibility of biodiesel industry,new alternate ways of utilization of g-phase have beenstudied recently. Possibilities such as combustion, coburning,composting, animal feeding, thermochemical conversionsand biological conversion have been applied for crudeglycerol processing [31–46].

One of the possible applications is utilization of g-phase as carbon and energy source for microbial growth inindustrial microbiology. Microbial conversion of glycerol tovarious compounds has been investigated recently, with par-ticular focus on the production of 1,3-propanediol [47–50],which has been considered as a main product of glycerol fer-mentation [28]. 1,3-propanediol presents several interestingapplications, it can be used as a monomer for polycondensa-tions to produce plastics with special properties, (polyesters,polyethers and polyurethanes) [51–54] as a monomer forcyclic compounds, as a polyglycol-type lubricant [55] andit also may serve as a solvent [56]. The biotechnologi-cal production of 1,3-propanediol from glycerol has beendemonstrated for several bacteria, Klebsiella pneumoniae,Clostridium butyricum and Citrobakter freundii have beenmost commonly used in the studies [47–50, 57–59].

Besides the production of 1,3-propanediol, glycerol canalso be used as a carbon source to obtain other valuablemicrobial products, such as recombinant proteins and

enzymes, microbial lipids (single-cell oils), medicinal drugs,antibiotics and fine chemicals. Bioconversion of g-phaseinto chemicals, such as dihydroxyacetone, 1,2-propanediol,ethanol, hydrogen,citric acid, propionic acid, polyglycerols,succinate, have been also increasingly studied recently[60–77].

Another option offers biological production of methanefrom crude glycerol using anaerobic sludge [78–88]. Besidesthe production of methane, the advantages include lownutrient requirements, energy savings, generation of lowquantities of sludge and excellent waste stabilization. Glyc-erol is a readily digestible substance, which can be easilystored over a long period. High energy content in g-phasemakes it an interesting substrate for anaerobic digestion aswell, since it offers high production of biogas in smallerreactor volumes. A great variety of microorganisms is able touse this substrate as a carbon source for the growth underanaerobic conditions, such as Citrobacter freundii, Kleb-siella pneumoniae, Clostridium pasteurianum, Clostridiumbutyricum, Enterobacter agglomerans, Enterobacter aerogenesor Lactobacillus reuteri [28, 34, 58, 67]. The production ofbiogas through anaerobic digestion offers significant advan-tages over other forms of crude glycerol treatment. It requireslower investments and simpler operational conditions com-pared to more sophisticated preprocessing technologies,which makes it ideal for local applications. Less biomasssludge is produced in comparison to aerobic treatmenttechnologies. The digestate is an improved fertilizer in termsof both its availability to plants and its rheology. A source ofcarbon neutral energy is produced in the form of biogas.

3. Anaerobic Digestion of Crude Glycerol

Considering anaerobic treatment of crude glycerol, potentialof its main component glycerol has been well-known for alonger period [89–91]. Digestion of pure glycerol has beeninvestigated both as a primary substrate [89, 90], and asan intermediate product of anaerobic degradation of fats[91]. Biodegradation have been carried out using either purecultures of microorganisms [90] or sludge composed ofmixed cultures from wastewater treatment plant [89].

Few studies focused on biogas production from g-phase [78–88] have also been realized recently. Anaerobictreatment of g-phase as a single substrate [78–81] was carriedout as well as coprocessing of crude glycerol with differentsubstrates [82–88].

Mesophilic anaerobic digestion of crude glycerol wasstudied in work Lopez et al. (2009). The substrate was


previously treated in two different ways: (1) acidificationwith phosphoric acid and centrifugation (so-called acidifiedglycerol) or (2) acidification followed by distillation (so-called distilled glycerol) [78]. Either granular sludge fromanaerobic reactor treating brewery wastewater or nongranu-lar sludge from anaerobic reactor treating urban wastewaterwas used for inoculation of batch laboratory-scale reactors,having the working volume of one liter. The variationsin the methane production were studied, considering thedifferent ways of substrate pretreatment and different typesof sludge. The use of the combination of granular sludgewith acidified glycerol was found to be the best option foranaerobic treatment of glycerol [78]. The organic loadingrates for each substrate and sludge type were in the rangeof 0.92–2.0 kg/m3·d (COD). Organic loading rate (ORL)is presented as the weight of organic matter per dayapplied over a specific volume of reactor. The parameterCOD (chemical oxygen demand) represents indirectly theamount of organic compounds in the sample. It is ameasure of the oxygen needed to degrade organic matter.A decrease in specific methane production was observedwhen the ORL was increased further. Considering thebiomass production and cell maintenance null, 0.382 m3 ofmethane are theoretically produced per kilogram of removedCOD. Experimentally, the effectiveness of the process ineach case was: 76% using granular sludge-acidified glycerol,75% using nongranular sludge-acidified glycerol and 93%with granular sludge-distilled glycerol (0.292; 0.288 and0.356 m3/kg COD removed, resp.). Besides the methaneproduction coefficient, the removed COD percentage is alsoimportant in order to determine biodegradability. This wasfound to be around 100% using granular sludge-acidifiedglycerol, 75% with nongranular sludge-acidified glycerol and85% using granular sludge-distilled glycerol.

Crude glycerol was processed in anaerobic laboratory-mixed reactor under mesophilic conditions for several mothsby Bodık et al. [79]. The anaerobic reactor achieved stableoperation at the volume loading of 4 kg/m3·d with biogasproduction ca0.980 m3/L of dosed g-phase. The maximalreached volumetric loading was 8–10 kg/m3·d, but the load-ing was considered to be very sensitive and unstable, becauseit caused decrease of the specific methane production andincrease of concentrations of volatile fatty acids (VFA) anddissolved COD. Very effective transformation of g-phase intobiogas was measured (more than 95%) which gives very goodassumptions for posttreatment of sludge water. The concen-tration of dissolved inorganic substances increased duringthe monitored period very slowly but continuously from1.3 g/L up to 15 g/L. Higher concentrations of dissolved saltscould cause inhibition of anaerobic degradation, however nosignificant influence was observed during this experiment.

In the work Hutnan et al. (2009) results of crude glyceroltreatment in the laboratory-mixed reactor (with effectivevolume of 4 liters) and in the laboratory UASB (upflowanaerobic sludge blanket) reactors (volume of 3.7 liters) aredescribed [80]. From this work resulted that the operationof mesophilic anaerobic degradation of crude glycerol asthe only organic substrate is feasible, however the processoperation is very sensible to organic overloading of reactor.

The laboratory-mixed reactor achieved stable operation atORL of 4 kg/m3·d (COD). The specific production of biogasachieved ca0.980 m3/L of glycerol added. The laboratoryUASB reactor with granulated biomass achieved stableoperation at ORL of 6.5 kg/m3·d and the specific biogasproduction was ca0.840 m3/L of glycerol added. Inoculationof the UASB reactor with suspended biomass showed thatthis type of sludge is not suitable for this purpose becauseof sludge flotation during the reactor operation.

Yang et al. (2008) examined biodegradation of glycerol-containing synthetic wastes using a fixed-bed laboratorybioreactor packed with polyurethane under mesophilic andthermophilic anaerobic conditions [81]. Better performancewas obtained from the reactor under the thermophilic con-ditions. When increasing the ORL from 0.25 to 1 kg/m3·d,the COD removal efficiency was decreasing under mesophilicconditions, however under thermophilic conditions higherCOD removal was achieved corresponding to higher load-ing rate. After 516 days of reactor operation, the bedmaterials under the thermophilic reactors were removedto measure the quantity of attached biomass and formicroscopic observation. The polyurethane immobilizationcarrier retained more biomass than did the liquid phase ofreactor. About 95% of the microbes were maintained onthe fixed-bed. The immobilized microorganisms present inthe thermophilic reactor were primarily Methanobacteriumsp., Methanosarcina sp., Bacillus sp., Clostridium sp., Desulfo-tomaculum sp. and Ruminococcus.

Feasibility of utilization of crude glycerol as a cosubstratehas been proven for example in the work of Fountoulakis(2009). The effects of g-phase on the performance ofanaerobic reactor treating different types of organic waste(organic fraction of municipal solid waste, mixture ofolive mill wastewater and slaughterhouse wastewater) wereexamined, in order to enhance methane production andincrease the yield of hydrogen [82]. Digestion was carriedout in a single-stage reactor with a working volume of3 liters, inoculated with anaerobic sludge from municipalsewage treatment plant and the share of crude glycerolmade 1% (v/v) of the dose. The supplementation of thefeed with crude glycerol had a significant positive effectand the methane production rate in both cases increasedclose to the theoretical values given total biodegradationof glycerol. Addition of g-phase to a reactor treating theorganic fraction of municipal solid waste resulted in theincrease of methane production to 2.094 L/d, comparedto 1.400 L/d. An enhanced methane production was alsoobserved when a mixture of olive mill wastewater andslaughterhouse wastewater was supplemented with crudeglycerol. Specifically, by adding 1% of g-phase to the feed,the methane production rate increased from 0.479 L/d to1.210 L/d. Stable concentration levels of COD indicated thatCOD attributed to glycerol in the feed was totally digestedin the reactor. The estimated yield of methane generatedfrom the digestion of glycerol was in both cases almostreaching the value of theoretical methane production. Thetheoretical methane production from digestion of glycerolwas estimated at 0.751 L/d (using the Buswell formula andthe ideal gas law).


Fountoulakis et al. studied also the feasibility of addingcrude glycerol to the anaerobic digesters treating sewagesludge in wastewater treatment plants [83]. Both batch andcontinuous experiments were carried out at 35◦C. It wasobserved that glycerol addition up to 1% (v/v) in the feedincreased methane production in the reactor above theexpected theoretical value, as it was totally digested andfurthermore enhanced the growth of active biomass in thesystem. On the other hand, any further increase of glycerolcaused a high imbalance in the anaerobic digestion process.The reactor treating the sewage sludge produced 1.106 ±0.036 L/d of methane before the addition of glycerol and2.353 ± 0.094 L/d after the addition of glycerol (1% in thefeed). The extra glycerol-COD added to the feed did nothave a negative effect on reactor performance, but seemedto increase the active biomass (volatile solids) concentrationin the system. Also, the kinetic experiments have shown thatglycerol biodegradation took place significantly faster thanpropionate (which is an intermediate product) biodegrada-tion, and it was therefore suggested that the glycerol overloadin the reactors increased propionate concentration.

Ma et al. studied the improvement of anaerobic treat-ment of potato processing wastewater in a laboratory UASBreactor by codigestion with crude glycerol [84]. Influenceof three types of glycerol was tested: pure glycerol, crudeglycerol and high conductivity glycerol. All 3 types ofglycerol are generated as a by-product by the productionof biodiesel. They are obtained by different processingtechnologies and thus their characteristics differ. Supplementof pure glycerol of 2 mL/L of potato processing wastewaterresulted in increase of the specific biogas production by0.740 m3/L of glycerol added. High COD removal efficiencies(around 85%) were obtained. Moreover, a better in-reactorbiomass yield (surplus of active biomass in the reactor) wasobserved for the UASB reactor supplemented with so-calledpure glycerol (0.012 g VS (volatile solids) per gram of CODremoved) compared to the reactor without added glycerol(0.002 g VS per gram of COD removed), which suggests apositive effect of glycerol on the sludge blanket growth.

Alvarez et al. carried out a laboratory study, aimed atmaximizing methane production by anaerobic codigestion ofthree agroindustrial wastes: crude glycerol, pig manure andtuna fish waste [85]. Experiments were performed by batch(discontinuous) assays and 500 mL reactors were operatedunder the temperature of 35◦C. Different blends composedby various percentages of these substrates were fed intothe reactors. Compositions of these blends were specifiedusing linear programming optimization method to find mostsuitable ratios of cosubstrates which would achieve highestbiodegradation potential or highest methane productionrate. The highest biodegradation potential (methane pro-duction of 0.321 m3/kg COD) was reached with a mixturecomposed of 84% pig manure, 5% fish waste and 11%biodiesel waste, while the highest methane production rate(16.4 L/kg·d (COD)) was obtained by a mixture containing88% pig manure, 4% fish waste and 8% biodiesel waste.Mixture composed of 84% pig manure, 5% fish waste and11% biodiesel waste and mixture of 79% pig manure, 5%fish waste and 16% biodiesel waste have also achieved very

high methane production rates (14.4 L/kg·d (COD) and12.8 L/kg·d (COD) resp.) compared to the control sampleusing pig manure substrate (8.3 L/kg·d (COD)).

Anaerobic codigestion of crude glycerol in the reactorsprocessing maize, maize silage and pig manure as mainsubstrates, is described in work of Amon et al. (2004).Laboratory digesters under mesophilic conditions were pro-cessing a basic mixture, which included 31% of maize silage,15% of corn maize and 54% of pig manure, together withaddition of various levels of g-phase (3, 6, 8 and 15%).The methane yield from the basic mixture without glycerineaddition reached 0.335 m3/kg VS [86]. Addition of 3% ofglycerine increased the methane yield by 20% and achieved0.411 m3/kg VS. The addition of 6% of glycerine resultedin the highest methane yield of 0.440 m3/kg VS. Additionof more than 6% glycerine to the basic mixture had onlya low positive influence on the methane yield. Additionof 15% glycerine even decreased the methane yield to0.400 m3/kg VS and the duration of fermentation increased.Methane formation at the start of the experiments wasdelayed. Analysis of the VFA concentrations in the mixtureduring the experiments resulted in the hypothesis that theinhibition of methane formation was caused by increasedconcentration of propionic and butyric acids. The largeamounts of these acids were built during decomposition ofmethanol. VFA accumulation reflects a kinetic uncouplingbetween acid producers and consumers and is typical forstress situations [92]. The main cause of the toxic effects ofhigh VFA concentrations on the anaerobic digestion processis generally considered to be the resulting drop in pH.

Long-term operation of anaerobic digester for cofermen-tation of maize silage and crude glycerol was studied in workSpalkova et al. (2009). Two laboratory models of a volumeof 6 liters were fed by maize silage and a mixture of maizesilage with crude glycerol and operated under mesophilicconditions [87]. During the operation period, no negativeinfluence of supplementation of the feed with crude glycerolwas observed. Biogas production as well as the sludge waterquality (pH, concentrations of COD, VFA, ammonia andphosphate) was similar in both reactors. Maximum portionof g-phase added formed 41.5% of total daily COD dose(together with maize silage). Specific biogas productionachieved was approximately 0.40 m3/kg (COD) in the caseof both sole maize silage and a mixture of maize silage withg-phase, meaning that both the maize silage and g-phase hadsimilar specific biogas productions per unit quantity of COD.

A positive effect of glycerol as a cofermentation mediumis supported by Amon et al. (2006). Biogas productions frompig manure, crude glycerol and a mixture of 94% of manurewith 6% of glycerol were compared in the study [88]. A 6%supplementation of glycerol to pig manure and maize silageresulted in a significant increase in methane production from0.569 to 0.679 m3/kg (VS). The methane yield of the mixturesupplemented with glycerine was higher than the combinedmethane yields of both substrates if digested separately.Increase in the specific methane production could not be justcorresponding to supplemented glycerol, but was also resultof the improved anaerobic degradation caused by the effectof codigestion. Co-digestion of various substrates provides


in many cases suitable option for anaerobic processingfor various technical reasons. One of the main reasons isthe stability of pH and sufficient buffer capacity. Lack ofnutrients or high concentration of inhibitory agents can alsobe improved by sensible choice of cosubstrates. A particularlystrong reason for codigestion of feedstock is the adjustmentof the carbon-to-nitrogen (C/N) ratio. Digestion of hardlydegradable substances was found to be faster by the additionof easily degradable substrates. Moreover some previouslyproblematic wastes were found digestible if digested in amixture of other waste.

The work by Hutnan et al. (2009) showed that crudeglycerol is also a suitable cosubstrate in the full scalebiogas plant for anaerobic treatment of maize silage [80].Reactor was operated under mesophilic conditions and theeffective volume of a full scale anaerobic reactor was 2450 m3.Evaluated specific production of biogas from crude glycerolwas about 0.890 m3/kg of crude glycerol added. Dose ofcrude glycerol, which represented only 5.2% of overall doseto biogas plant, produced almost 15% of overall biogasproduction. A significant influence on positive economicalbalance of biogas plant using this cosubstrate has beendemonstrated in this study. At the electrical power outputof cogeneration unit 300 kW is the daily share of electricityproduced from crude glycerol 1067 kWh. At the current priceof 0.15 C per 1 kWh, this represents a daily profit of 156.55 Cand a saving of almost 15% silage (1865 kg at a price around60 C for 1 ton).

The possible inhibition effects, resulting from the sub-strate composition, have to be considered by anaerobictreatment of crude glycerol. Metabolism of the anaerobicmicroorganisms may be negatively affected mainly by thehigh salinity of g-phase [79, 80]. The relatively high contentof sodium or potassium salts (ca 20–100 g/L) originatesfrom the catalyst, used for the biodiesel production. Higherconcentrations of sodium in the anaerobic reactor canseriously inhibit the microbial activity [93].

Biological processing of organic materials in the presenceof salts have been studied mainly as an alternative possibilityof treating wastewater from industrial processes [94–97](meat canning, pickled vegetables, dairy products, olive andfish processing industries, petroleum, textile and leatherindustries). The anaerobic digestion of industrial salineeffluents, predominantly from seafood processing, at saltconcentrations ranging from 10 to 71 g/L has been studiedrecently,using different processes, such as an anaerobic filter,UASB reactor and an anaerobic contact system [94, 95, 98].The COD removal efficiencies obtained, generally remainedbetween 70% and 90%, with OLR ranging from 1 to15 kg/m3·d (COD).

The concentration of sodium exceeding 10 g/L was fora long time generally considered to strongly inhibit methano-genesis [94]. However, the anaerobic digestion in the highsalinity level was proven to be possible for treatment of fish-processing effluent [99, 100], if a suitable strategy for adapt-ing the methanogenic biomass was applied. Furthermore, itwas shown, that the toxicity of sodium in sludge depends onseveral factors, such as the type of methanogenic substrateused, the antagonistic or synergistic effects of other ions,

the nature and the progressive adaptation of sludge to highsalinity and reactor configuration [93, 101].

These factors may be the cause of different results achi-eved in different studies. Some of the researchers reportedthe concentrations from 0.9 g/L to 8 g/L to be slightlyinhibitory (reduction in methane production by 10%), usingdifferent types of substrates [102]. In other experiments, theconcentrations in the range of 5.6–53 g/L, depending on theconditions, have been documented to cause the decrease ofmethane production to a half [93, 101–103].

Adequate adaptation of the sludge appears to be ofextreme importance, hence the continuous exposure ofmethanogenic sludge seams to lead to the tolerance of ahigher salinity compared to the sludge exposed to salt shocks.The adaptation includes gradual increase of salt concen-trations in the sludge, by low organic loading, providingadequate conditions for internal structural changes in thepredominant species of methanogens, to adapt to higherosmolarity [104]. Hence the startup period may take severalmonths [94]. According to Gebauer [97] is the adaptation tohigh sodium concentrations more likely to happen as a resultof selection of tolerant species than by adaptation of everysingle microorganism.

Methanogenic microorganisms seem to be more affectedby sodium toxicity than other populations, such as pro-pionate utilizes [101]. The most sensitive appeared to bethe nitrogen removing microorganisms [98]. Provided thebiomass is acclimated, high salinity is reportedly not anobstacle to its growth and it has no negative influence on sed-imentation properties of the sludge or the granulated sludgeviscosity. The lack of macronutrients (nitrogen, phosphorus,sulfur) in the medium was found to have a more pronouncednegative effect on biomass under the saline conditions. Onthe other hand, the absence of micronutrients did not furtherreduce biomass activity under salinity [101, 105].

Another important concern about the anaerobic diges-tion of crude glycerol should be the concentration ofnitrogen-rich substances. Ammonium concentration up to200 mg/L in the anaerobic reactor is considered to be benefi-cial [93], since nitrogen is an essential nutrient for microor-ganisms. Considering the low concentration of nitrogen inthe crude glycerol, it may be necessary to supply the nitrogen-rich substances into the reactor. Urea or NH4Cl are mostfrequently used as external source of ammonium nitrogen.

4. Oil Cakes

Oil cakes or oil meals are solid residues obtained afteroil extraction from the seeds. Their composition widelyvaries depending on the quality of seeds or nuts, growingconditions and extraction methods. Oil cakes can be eitheredible or nonedible. Edible cakes have a high protein contentranging from 15 to 50%. The chemical compositions of oilcakes originating from different types of plants are listed inTable 3 [106].

In our geographic area (EU), rapeseed and sunflower arethe most frequently used substrates, hence we are focusingon the by-products from their processing.


Table 3: Composition of oil cakes.

Oil cake Dry matter % Crude protein % Crude fibre % Ash % Calcium % Phosphorus %

Canola oil cake 90 33.9 9.7 6.2 0.79 1.06

Coconut oil cake 88.8 25.2 10.8 6.0 0.08 0.67

Cottonseed cake 94.3 40.3 15.7 6.8 0.31 0.11

Groundnut oil cake 92.6 49.5 5.3 4.5 0.11 0.74

Mustard oil cake 89.8 38.5 3.5 9.9 0.05 1.11

Olive oil cake 85.2 6.3 40.0 4.2 — —

Palm kernel cake 90.8 18.6 37 4.5 0.31 0.85

Sesame oil cake 83.2 35.6 7.6 11.8 2.45 1.11

Soy bean cake 84.8 47.5 5.1 6.4 0.13 0.69

Sunflower oil cake 91 34.1 13.2 6.6 0.30 1.30

Table 4: Composition of rapeseed, rapeseed cake after extraction of 60, 70, and 75% of oil and rapeseed meal, in percents of total solids[111].

Feed stock Rapeseed Rapeseed cake Rapeseed meal

Portion of extracted oil 60% 70% 75%

Crude oil 45 24.7 19.7 17 4.5

Crude protein 23 31.5 33.6 34.7 40

Crude fibre 7 9.6 10.2 10.6 12.3

Ash 5 6.8 7.3 7.5 7.7

Depending on the method of oil extraction from theseeds, two basic types of solid by-products are generated.Oil cakes are produced when simple oil pressing system isused. In case that pressing is followed by advanced extractiontechniques, residues are usually referred to as oil meals.As can be seen in Table 4, main difference between the oilcakes and oil meals is based on the content of fats. Themore effective is the extraction process, the fewer lipidsremain in the cakes. About 12% of fats (or even 20% incase of small processing facilities) may remain in the oilcake when simple pressing method is employed. Secondpressing, sometimes accompanied by water vapor extraction,can lower the content of fats to approximately 8%. If theextraction using hexane is engaged, oil meal with fats contentabout 1–3% can be generated.

Oil cakes have been currently in use predominantlyfor feed applications to poultry, ruminant, fish and swineindustry [107–110]. Some of them are considered to be suit-able organic nitrogenous fertilizers. Several cakes have beenutilized for production of proteins, enzymes, antibiotics,mushrooms, ethanol [107–112]. Biotechnological applica-tions of oil cakes also include production of vitamins andantioxidants [111, 113].

Current prices of oil cakes are relatively high (rapeseedcake and meal are in Europe worth approximately 166 and161 Euros per ton, resp.), compared to other agroindustrialby-products and wastes, which could be also used as sub-strates for biogas production. However experts are warningagainst their expected drop due to possible overproduction[114–116]. Moreover, with increasing emphasis on costreduction of industrial processes and value addition to

agroindustrial residues, alternative utilization for oil cakeshas been required.

Utilization of oil cakes as an energy source is underexamination for now. Some of the oil cakes have been studiedas possible feedstocks for biogas production, combustionor pyrolysis [117]. Considering the high content of fats, oilcakes have a high energetic value. They could be suitablesubstrates for combustion, however because of the largequantity of ash and high emissions of nitrogen oxides,advanced purification technology is required.

Oil cakes and meals contain a high portion of digestiblesubstances, which makes them suitable substrates for theproduction of biogas. Nutritional content should not besignificantly affected by the anaerobic degradation (nutrientssuch as nitrogen and phosphorus stay in the digestate afterdegradation) and the digestate should be a convenient agri-cultural fertilizer. In addition, the plant nutrients containedin cakes are more easily available after the biodigestion.

5. Anaerobic Digestion of Rapeseed Oil Cake

Rapeseed cake and rapeseed meal are degradable organicsubstances. They are suitable for anaerobic digestion, how-ever supplementation of other organic substrates might berequired to achieve better process performance and particu-lar problems of digestion should be more closely studied.

Rapeseed cake is a protein-rich substrate, hence thedecomposition and conversion to biogas takes longer timethan decomposition of substrates rich in carbohydrates. Incase of protein degradation, hydrolysis is the limiting step,specifically the cracking of proteins into amino acids and


Table 5: Methane production potential from rapeseed, rapeseed cake and meal in m3/kg VS [111].

Feed stock Rapeseed Rapeseed cake Rapeseed meal

Portion of extracted oil 60% 70% 75%

Saccharides 0.11 0.16 0.17 0.17 0.20

Lipids 0.43 0.24 0.19 0.16 0.04

Proteins 0.12 0.16 0.17 0.18 0.20

Together 0.66 0.55 0.53 0.51 0.45

Together (kg) 0.47 0.39 0.37 0.36 0.32

Calorific value (MJ) 23.4 19.5 18.6 18.1 15.8

polypeptides by extracellular enzymes. The hydrolysis ofcarbohydrates takes place within a few hours, while thehydrolysis of proteins within few days. Rate and readinessof degradation of different types of carbohydrates can quitevary. Fats are often decomposed completely. Hemicelluloseand lignin, forming the shells of rapeseed, could be quitedifficult to decompose in the process of biogas production.

Accumulation of free fatty acids can cause a problemby digestion of materials with higher content of oil, con-sidering that the fats decomposition step is faster thanthe methanogenesis. Generally, hydrolytic and acidogenicmicroorganisms are growing about ten times faster thanmethanogens. Co-digestion of rapeseed cake or meal withother feedstocks, such as manure, provides an alternativesolution. Improvement of the biogas production is expected,based on the high oil content.

Rapeseed cake and rapeseed meal are nitrogen-richmedia, they content about 35–40% of nitrogen substances[106]. These substances are predominantly proteins, con-taining amino acids. Expressed in the terms of carbon-to-nitrogen ratio, this makes about 5–8 in case of rapeseed cake.Compared to other materials, the C/N ratio of lignocellulosicmaterials is in the range of 60–400, grass and silages haveC/N about 20–40, swine manure about 12–15 and sunflowercake about 12-13 [118]. The C/N ratio of substrate isan important parameter to be considered by anaerobicdegradation, since high content of nitrogen may cause toohigh content of ammonium nitrate in the biogas reactor.Ammonium nitrogen levels of about 4 g/L of wet sludge bringthe risk of process inhibition. If the ammonium content istoo high, it is necessary to dilute the substrate with water ornitrogen-poor material.

Phytotoxic effects of rapeseed cake, caused by the con-tent of glucosinolates, must also be considered. They play animportant role in the process of digestion, since in hig-her concentrations they may have harmful effect on metha-nogenic. The risk of inhibition is getting less serious withdecreasing of the glucosinolates level in the rapeseed mealand cake.

There is not much information about experimental anae-robic processing of rapeseed cake or meal in the availableliterature.

Bernesson et al. estimated the potential biogas pro-duction from rapeseed, rapeseed meal and rapeseed cakeafter 60–75% extraction of oil [111]. Table 5 indicates,that with the increased amount of oil extracted in the

process, the possible biogas production from rapeseed cake isdecreasing.

Antonopoulou et al. carried out batch mesophilicbiochemical methane potential tests using rapeseed andsunflower residues as a substrate [119]. The experimentsindicated that the biological methane potential of rapeseedand sunflower meal were 0.450 m3/kg and 0.481 m3/kg,respectively. Compared to commonly used substrate maizesilage, the potential of these oil meals are about 40% higher,so it suggests interesting substrates for the production ofbiogas. Various pretreatment methods, such as thermal,chemical (through alkali or acid addition) or combination ofthe above methods were also tested in the effort to enhancethe methane productivity and yield. Thermal pretreatmentmethod was conducted at 121◦C for 60 minutes in a pressurecooker. Acid or alkali pretreatment of the feedstocks wasconducted by the addition of 2% w/v H2SO4 or NaOH,respectively, for 60 minutes at a temperature of 25◦C orat 121◦C for 60 min in a pressure cooker (thermal acidor thermal alkali pretreatment). The experiments showedthat the pretreatment methods tested did not enhance themethane potential of the rapeseed and sunflower residues.This could be attributed to the inhibitory compounds whichwere possibly released during the pretreatment.

6. Anaerobic Digestion of Sunflower Oil Cake

Sunflower oil cakes and meals are also feasible feedstocks foranaerobic digestion. Raposo et al. examined their anaerobicdegradability, biochemical methanogenic potential and theinfluence of substrate to inoculum ratio in batch laboratory-scale digesters [120, 121]. High stability of the anaerobicdigestion process of sunflower oil cake under mesophilicconditions was demonstrated.

The experimental study, with the duration of 7 days, wascarried out in a multibatch reactor system [120, 121], whichconsisted of continuously stirred flasks with an effectivevolume of 250 mL. The six different inoculum to substrateratios were tested: 3.0, 2.0, 1.5, 1.0, 0.8 and 0.5. The ultimatemethane yield decreased considerably with the inoculum tosubstrate ratio. The yield of methane was in the range from0.227 m3/kg for the ratio of 3.0 to 0.107 m3/kg (VS) for theratio of 0.5. Biodegradability copied this trend, from 86%to 41% was achieved. Higher contribution of substrate maycause lower methane yield due to higher energy consumption


in the hydrolytic-acidogenic stage. However, the net VSremoved only varied from 42% to 36%, when the ratiodecreased from 3.0 to 0.5, which demonstrated the adequateoperation of the hydrolytic-acidogenic stage.

The increase in CODs concentrations presented 780–6100 mg/L for the inoculum to substrate ratios of 3.0–0.5,respectively. In case of inoculum-substrate ratio of 3.0 or2.0, the final values of total VFA were proportional to theamount of substrate added, and no accumulation occurred.However, when the ratio was lower than 2.0, an imbalance ofthe process was observed, when the VFAs increased to 2050or 5500 mg/L (for ratios 0.8 and 0.5, resp.). The dissolvedCODs also increased, reaching the levels of 3380–12100 mg/Lafter seven days for the inoculum-substrate ratios of 3.0–0.5,respectively. The trend in the increase of COD with digestiontime observed was due mainly to the accumulation of VFA,which reflects a kinetic uncoupling between acid formersand consumers and is typical for a stress situation. Thismeans that the hydrolytic-acidogenic stage was carried outsatisfactorily and the imbalance of the process was due to thestress of methanogenic microorganisms.

The net production of total ammonia nitrogen increasedwith the load added, as a consequence of degradation ofproteinsfrom the sunflower oil cake, achieving a maximumvalue of 1085 mg/L at the ratio of 0.5 (198 mg/L at the ratio3.0). However, the specific total ammonia nitrogen reachedin all ratios similar production of about 40 mg/g VS added.

Identification of the individual VFA may also providevaluable information on the metabolic pathways involved inthe process. The high influence of inoculum-substrate ratioon the composition and concentration of the different VFAwas shown. By the inoculum to substrate ratio of 3 and 2,the predominant VFA were valeric and butyric acids, butthe residual compound was the latter. The absence of aceticand propionic acids indicated, that the methanogenic stagewas not disturbed and the formation of methane from theseintermediates was quick.

When the ratios of 1.5, 1.0 and 0.8 were applied, thepredominant VFA during the first few days were aceticand propionic acids, followed by valeric and butyric acids.Although in the end valeric acid dominated. This perfor-mance demonstrated that the lower inoculum-substrate ratiocauses the greater accumulation of the longer chain VFA.

By the ratio of 0.5 the predominant VFA were aceticand propionic acids during the first few days, followed by adecrease in acetic acid with time, with a significant residualconcentration of propionic, valeric and butyric acids. TheVFA profile obtained is a consequence of the imbalance inthe methanogenic stage.

De La Rubia et al. investigated also influence of thehydraulic retention time (HRT, it is a measure of the lengthof time that sludge remains in reactor.) and OLR on theperformance of the hydrolytic-acidogenic step of a two-stageanaerobic digestion process of sunflower oil cake [122]. Theexperiments were performed in laboratory-scale completelystirred tank reactors, with a working volume of 2 L, atmesophilic (35◦C) temperature. Digesters were operatedover a total period of approximately 350 days. Six OLRs(ranging from 4 to 9 kg/m3·d(VS)) for four HRTs (8, 10,

12 and 15 days) were tested to check the effect of eachoperational variable. Hydrolysis yields obtained for all HRTsand OLRs assayed were in the range of 20.5–30.1%.

Variations inHRT did not affect the COD solubiliza-tion of this substrate withinthe HRT range (15–8 days)researched. Variations in OLR affect the organic matter lique-faction slightly, the highest value (30.1%) being achieved forHRT of 10 days andOLR of 6 kg/m3·d(VS). The acidificationyield increased with OLR up to 6 kg/m3·d(VS), the highestvalue (83.8%) being achieved for HRT of 10 days and anOLR of 6 kg/m3·d(VS). However, higher loading provokes adecrease in the acidification yield, probably due to the factthat the acidogenic bacteria could have been affected andinhibited at the highest OLR studied.

7. Washing Water

Another possible by-product from biodiesel productionoffers the water, generated by washing of raw biodiesel.Under the conventional process (alkali-catalyzed transester-ification) for every 100 L biodiesel produced about 20 L ofwashing water is discharged(or more in case of prior acidpretreatment) [123].

Washing water (usually referred to as biodiesel wastewa-ter) is a viscous liquid with an opaque white color similar toaqueous soap. It contains significant amounts of methanol,glycerol and soaps. Methyl esters bound with soap, NaOH orKOH from the catalyst, sodium or potassium salts and tracemono, di- and triglycerides bound up with the soap are alsocontained in the water.

A great variety of systems for biodiesel purification isavailable commercially and new alternative technologies arealso being investigated. The possible options include drywashing. In this case, the impurities from biodiesel (freeglycerol, soap, free fatty acids, catalyst,glycerides, etc.) areabsorbed to form a solid waste product instead of a liquid.Dry washing replaces water with an ion exchange resin ora magnesium silicate powder. Both these methods are beingused in industrial plants [124]. No regeneration is normallyapplied and the spent material has to be disposed of tolandfill or other applications (compost, potential animal feedadditive and potential fuel).

Relatively expensive ultrafiltration or reverse osmosiscould also be applied for purification of biodiesel. Yet, thewashing with water remains the most convenient alternative[125–127].

Besides crude glycerol, oil cakes and biodiesel wastewater,other potential by-products can be generated in the biodieselindustry. These products are specific, depending on theprocessing technologies in biodiesel plants. For example,some facilities utilize citric acid solutions in order to washreactors and other equipment, which produces possibleadditional waste.

Like the raw glycerol, washing water has also highlevels of COD, values in the range of 18–800 g/L have beenreported [123, 128–130]. High content of degradable organicsubstances makes it a suitable source of carbon for microbio-logical processes, however some issues have to be considered.


The wastewater is basic (alkaline), due to the significant levelsof residual KOH, and contains a high level of oil and greaseand has a high solid content. Nutrients for microbial growth(such as nitrogen and phosphorus) are not abundant inwashing water, except for the carbon source. Together thesecomponents inhibit the growth of most microorganismsmaking this wastewater difficult to degrade naturally [123].Focusing on anaerobic degradation, long-chain fatty acids,which are present on a high level in the washing water, havebeen reported to be inhibitors of the digestion process [93].To reduce this effect, electrocoagulation has been proposedas a successful pretreatment for oily wastewater with asubsequent anaerobic treatment [129, 130].

With the likely expansion of biodiesel production byplants using the conventional method, comes the inherentneed to treat the wastewater. The main component ofthe wastewater is the residual remaining oil, thus, suchwastewater should not be discharged into public drainagebecause the oil causes plugging of the drainage and decreasesbiological activity in sewage treatment. Some of the typicalcommercially available treatments of oily wastewater employa dissolved air floatation technique or oil and grease trap unit[130]. Currently, several processes have been developed totreat the biodiesel wastewater, such as the use of chemicalrecovery approach and electrochemical treatment [128, 130],but also the employment of microbiological processes [123,131–134] and anaerobic digestion [129].

In the work of Jaruwat et al. (2010), the management ofraw biodiesel wastewater was carried out at a laboratory scaleat ambient temperature by a combined protonation basedchemical recovery of biodiesel followed by electrochemicaltreatment of the residual wastewater [128]. The combinedtreatment completely removed COD and oil and grease, andreduced BOD (biologic oxygen demand) levels by more than95%.

In the study, carried out by Chavalparit and Ong-wandee (2009), electrocoagulation was adopted to treat thebiodiesel wastewater [130]. This study demonstrates that theelectrocoagulation process using an aluminum anode andgraphite cathode is effective in reducing oil and grease andsuspended solids by more than 95% in the washing water.However, the COD removal is achieved by 55% due to lesssignificant removal of glycerol and methanol. Therefore, theelectrocoagulation process is possibly suitable for a primarytreatment for biodiesel wastewater and it still requires afurther biological treatment process. Authors believe thatpretreatment with electrocoagulation followed by a biolog-ical treatment process is feasible and competitive comparedwith evaporation or pure physicochemical treatments. Itrequires less energy consumption, short process time, nochemical addition and less sludge production.

The biological treatment of washing water was inves-tigated by Suehara et al. (2005). For the microbiologicaldegradation using a 10-L fermentor, oil degradable yeast,Rhodotorula mucilaginosa, was used and the optimumconditions were determined [123]. The pH was adjustedto 6.8 and several nutrients such as a nitrogen source(ammonium sulfate, ammonium chloride or urea), KH2PO4

and MgSO4·7H2O were added to the wastewater. To avoid

the inhibition of the microbial growth, the raw biodieselwastewater was diluted with the same volume of water. Theoptimal initial concentration of yeast extract was 1 g/L andthe optimal C/N ratio was between 17 and 68 when usingurea as a nitrogen source. Authors suggest this biologicaltreatment system to be useful for small-scale biodieselproduction plants, because it is simple and no controllers,except for a temperature, are necessary.

Kato et al. (2005) proposed a continuous-type consor-tium bioreactor for treatment of washing water [131]. Themain component of this reactor was bacteria-fixed ceramicmaterial with high-oil degrading capability. A series of oildecomposition tests was carried out using the consortiumsystem, in which the most important bacteria types wereAcinetobacter, Bacillus and Pseudomonas. The optimal con-ditions for operation were confirmed by batch tests: airagitation, pH of around 6 and water temperature of 30◦C.This reactor operated almost maintenance-free for one year.The field test results for washing water showed that oil andgrease concentrations decreased from an initial 120 g/L to atreated range of 10–30 mg/L.

Papanikolaou et al. investigated valorization of soapsfrom washing water for the production of microbial lipidsof specific structure [132–134]. Several oleaginous yeasts andmolds are able to accumulate in abundance storage lipid, andat the same time modify the composition of the fat utilizedas the carbon source. In the case of various crude fats orfatty wastewaters of low value, this may be an industriallyand financially interesting approach. Potential productionof cocoa-butter substitute by Yarrowia lipolytica was studiedand the cell growth and lipid accumulation of Y. lipolyticawas investigated [132].

The anaerobic codigestion of glycerol and wastewaterderived from biodiesel manufacturing was studied in batchlaboratory-scale reactors of 1 L volume, inoculated by gran-ular biomass, at mesophilic (35◦C) temperature [129]. Themain purpose of this study was to evaluate the performance,stability, biodegradability, methane yield coefficient, kineticsof methane production, inoculum-substrate ratio and OLRof the anaerobic codigestion of these by-products derivedfrom biodiesel manufacturing.

Prior to the biological treatment, glycerol was acid-ified with H3PO4 in order to recover the alkaline cat-alyst employed in the transesterification reaction (KOH)as agricultural fertilizer. Wastewater was subjected to anelectrocoagulation process in order to reduce its oil content.The pretreated washing water was mixed with glycerol at aproportion of 85–15 (COD), until obtaining a final solubleCOD of 300 g/L. After mixing, the anaerobic revalorizationof the wastewater was studied employing inoculum-substrateratios ranging from 5.02 to 1.48 kilogram of VSS (volatilesuspended solids) per kilogram of COD and OLR of0.27–0.36 kg/kg·d (COD/VSS). Biodegradability was foundto be around 100%, while the methane yield coefficientwas 0.310 m3/kg COD removed. The results showed thatanaerobic codigestion reduces the clean water and nutrientrequirement, with the consequent economical and environ-mental benefit.


8. Conclusions

The process of biodiesel production is predominantly car-ried out by catalyzed transesterification. Besides desiredmethylesters, this reaction provides also a few other products,including crude glycerol, oil-pressed cakes and washingwater. Although their composition widely varies dependingon the parameters and substrates used for biodiesel pro-duction, all these by-products provide valuable feedstocksfor biogas generation. The possibility and performance ofanaerobic digestion of these materials have been studiedto various extents. The results can be summarized in fewpoints:

(i) Crude glycerol from biodiesel production was provento be a suitable substrate for anaerobic degradation.A couple of studies have demonstrated the possibilityof biogas production, using g-phase as a singlesubstrate.

(ii) G-phase has also shown a great potential as acosubstrate by anaerobic treatment of different typesof organic waste: organic fraction of municipal solidwaste, mixture of olive mill wastewater and slaughter-house wastewater and potato processing wastewater.Positive effect of crude glycerol on the enhancementof anaerobic processes was observed by treatment ofcorn maize, maize silage, and swine manure.

(iii) Oil cakes and oil meals can also be used as feasibleand economically interesting substrates for biogasproduction. The possibility of methane productionfrom rapeseed and sunflower oil cakes, which deservethe most interest in our area, has been suggestedlately.

(iv) Tests of anaerobic degradability, biochemicalmethanogenic potential and influence of substrateto inoculum ratio demonstrated high stability ofthe anaerobic digestion of sunflower oil cake undermesophilic conditions.

(v) The potential biogas production from rapeseed mealand rapeseed cake was estimated. It was shown,that with the increased amount of oil gained in theextraction process, the possible biogas productionfrom rapeseed cake decreases.

(vi) No significant effect of the pretreatment (thermaland chemical), of neither sunflower nor rapeseedresidues, on the enhancement of methane yield hasbeen observed so far.

(vii) Washing water from biodiesel purification is alsoa promising material for anaerobic degradation,considering the high content of readily degradableorganic substances. However, the possibility of biogasgeneration has not been sufficiently studied.

(viii) The specific inhibition effects, resulting from thesubstrates composition, have to be considered byanaerobic treatment of biodiesel by-products. Incase of anaerobic digestion of crude glycerol, highsalinity of the substrates may negatively affect the

methanogenic microorganisms. The concentration ofammonium should also be monitored. Since nitrogenis an essential nutrient for microorganisms, the lowconcentration in the crude glycerol and washingwater has to be compensated by ammonium sup-plement. On the other hand, rapeseed cake containsa high portion of nitrogen-rich substances, whichmay cause inhibition of digestion due to ammoniumaccumulation in the reactor.

Utilization of the by-products as a potential source ofenergy, rather then treat them as a waste, seems to be aconvenient way of lowering the costs of biodiesel and makingit more competitive.

Acknowledgment

This work was supported by the Slovak Grant Agency forScience VEGA (Grant 1/0145/08).

References

[1] J. Janaun and N. Ellis, “Perspectives on biodiesel as asustainable fuel,” Renewable and Sustainable Energy Reviews,vol. 14, no. 4, pp. 1312–1320, 2010.

[2] European Biodiesel Board: 2009-2010, EU Biodiesel IndustryRestrained Growth in Challenging Times, 2010.

[3] A. Demirbas, “Comparison of transesterification methodsfor production of biodiesel from vegetable oils and fats,”Energy Conversion and Management, vol. 49, no. 1, pp. 125–130, 2008.

[4] J. Van Gerpen, “Biodiesel processing and production,” FuelProcessing Technology, vol. 86, no. 10, pp. 1097–1107, 2005.

[5] D. Y. C. Leung, X. Wu, and M. K. H. Leung, “A reviewon biodiesel production using catalyzed transesterification,”Applied Energy, vol. 87, no. 4, pp. 1083–1095, 2010.

[6] M. Fangrui and A. Milford, “Biodiesel production: a review,”Bioresource Technology, vol. 70, no. 1, pp. 1–15, 1999.

[7] M. Balat and H. Balat, “Progress in biodiesel processing,”Applied Energy, vol. 87, no. 6, pp. 1815–1835, 2010.

[8] F. I. Gomez-Castro, V. Rico-Ramirez, J. G. Segovia-Hernandez, and S. Hernandez, “Feasibility study of athermally coupled reactive distillation process for biodieselproduction,” Chemical Engineering and Processing, vol. 49,no. 3, pp. 262–269, 2010.

[9] J. S. Lee and S. Saka, “Biodiesel production by heterogeneouscatalysts and supercritical technologies,” Bioresource Technol-ogy, vol. 101, no. 19, pp. 7191–7200, 2010.

[10] J. Ye, S. Tu, and Y. Sha, “Investigation to biodiesel productionby the two-step homogeneous base-catalyzed transesterifica-tion,” Bioresource Technology, vol. 101, no. 19, pp. 7368–7374,2010.

[11] F. E. Kiss, M. Jovanovic, and G. C. Boskovic, “Economic andecological aspects of biodiesel production over homogeneousand heterogeneous catalysts,” Fuel Processing Technology, vol.91, no. 10, pp. 1316–1320, 2010.

[12] Z. Helwani, M. R. Othman, N. Aziz, W. J. N. Fernando, andJ. Kim, “Technologies for production of biodiesel focusingon green catalytic techniques: a review,” Fuel ProcessingTechnology, vol. 90, no. 12, pp. 1502–1514, 2009.

[13] E. Li and V. Rudolph, “Transesterification of vegetable oilto biodiesel over MgO-functionalized mesoporous catalysts,”Energy and Fuels, vol. 22, no. 1, pp. 145–149, 2008.


[14] G. Arzamendi, I. Campo, E. Arguinarena, M. Sanchez, M.Montes, and L. M. Gandıa, “Synthesis of biodiesel withheterogeneous NaOH/alumina catalysts: comparison withhomogeneous NaOH,” Chemical Engineering Journal, vol.134, no. 1–3, pp. 123–130, 2007.

[15] W. M. Antunes, C. D. O. Veloso, and C. A. Henriques,“Transesterification of soybean oil with methanol catalyzedby basic solids,” Catalysis Today, vol. 133–135, no. 1–4, pp.548–554, 2008.

[16] X. Liu, H. He, Y. Wang, S. Zhu, and X. Piao, “Transesterifi-cation of soybean oil to biodiesel using CaO as a solid basecatalyst,” Fuel, vol. 87, no. 2, pp. 216–221, 2008.

[17] G. Ondrey, “Biodiesel production using a heterogeneouscatalyst,” Chemical Engineering, vol. 111, no. 11, p. 13, 2004.

[18] A. Bajaj, P. Lohan, P. N. Jha, and R. Mehrotra, “Biodieselproduction through lipase catalyzed transesterification: anoverview,” Journal of Molecular Catalysis B, vol. 62, no. 1, pp.9–14, 2010.

[19] T. Tan, J. Lu, K. Nie, L. Deng, and F. Wang, “Biodieselproduction with immobilized lipase: a review,” BiotechnologyAdvances, vol. 28, no. 5, pp. 628–634, 2010.

[20] S. Al-Zuhair, A. Almenhali, I. Hamad, M. Alshehhi, N.Alsuwaidi, and S. Mohamed, “Enzymatic production ofbiodiesel fromused/waste vegetable oils: design of a pilotplant,” Renewable Energy. In press.

[21] P. D. Patil, V. G. Gude, A. Mannarswamy et al., “Optimizationof direct conversion of wet algae to biodiesel under supercrit-ical methanol conditions,” Bioresource Technology, vol. 102,no. 1, pp. 118–122, 2011.

[22] S. Saka, Y. Isayama, Z. Ilham, and X. Jiayu, “New process forcatalyst-free biodiesel production using subcritical acetic acidand supercritical methanol,” Fuel, vol. 89, no. 7, pp. 1442–1446, 2010.

[23] R. Sawangkeaw, K. Bunyakiat, and S. Ngamprasertsith, “Areview of laboratory-scale research on lipid conversion tobiodiesel with supercritical methanol (2001–2009),” Journalof Supercritical Fluids, vol. 55, no. 1, pp. 1–13, 2010.

[24] A. Demirbas, “Biodiesel from waste cooking oil via base-catalytic and supercritical methanol transesterification,”Energy Conversion and Management, vol. 50, no. 4, pp. 923–927, 2009.

[25] J. C. Thompson and B. B. He, “Characterization of crudeglycerol from biodiesel production from multiple feed-stocks,” Applied Engineering in Agriculture, vol. 22, no. 2, pp.261–265, 2006.

[26] T. Kocsisova and J. Cvengos, “G-phase from methyl esterproduction-splitting and refining,” Petroleum and Coal, vol.48, pp. 1–5, 2006.

[27] L. Bournay, D. Casanave, B. Delfort, G. Hillion, and J.A. Chodorge, “New heterogeneous process for biodieselproduction: a way to improve the quality and the value of thecrude glycerin produced by biodiesel plants,” Catalysis Today,vol. 106, no. 1–4, pp. 190–192, 2005.

[28] N. Pachauri and B. He, “Value added utilization of crudeglycerol from biodiesel production: a survey of currentresearch activities,” in Proceedings of the ASABE AnnualInternational Meeting, 2006.

[29] A. Singhabhandhu and T. Tezuka, “A perspective on incor-poration of glycerin purification process in biodiesel plantsusing waste cooking oil as feedstock,” Energy, vol. 35, no. 6,pp. 2493–2504, 2010.

[30] J. A. Weber, The economic feasibility of community-bas-ed biodiesel plants, M.S. thesis, University of Missouri,Columbia, 1993.

[31] N. Luo, X. Fu, F. Cao, T. Xiao, and P. P. Edwards, “Glycerolaqueous phase reforming for hydrogen generation overPt catalyst—effect of catalyst composition and reactionconditions,” Fuel, vol. 87, no. 17-18, pp. 3483–3489, 2008.

[32] D. Pyle, Use of biodiesel-derived crude glycerol for the produc-tion of omega-3 polyunsaturated fatty acids by the microalgaSchizochytriumlimacinum, Ph.D. thesis, Biological SystemsEngineering, 2008.

[33] Z. Chi, D. Pyle, Z. Wen, C. Frear, and S. Chen, “Alaboratory study of producing docosahexaenoic acid frombiodiesel-waste glycerol by microalgal fermentation,” ProcessBiochemistry, vol. 42, no. 11, pp. 1537–1545, 2007.

[34] G. P. da Silva, M. Mack, and J. Contiero, “Glycerol: apromising and abundant carbon source for industrial micro-biology,” Biotechnology Advances, vol. 27, no. 1, pp. 30–39,2009.

[35] Y. Feng, Q. Yang, X. Wang, Y. Liu, H. Lee, and N. Ren,“Treatment of biodiesel production wastes with simultane-ous electricity generation using a single-chamber microbialfuel cell,” Bioresource Technology, vol. 102, no. 1, pp. 411–415,2010.

[36] T. Valliyappan, N. N. Bakhshi, and A. K. Dalai, “Pyrolysisof glycerol for the production of hydrogen or syn gas,”Bioresource Technology, vol. 99, no. 10, pp. 4476–4483, 2008.

[37] F. Freitas, V. D. Alves, J. Pais et al., “Production of a newexopolysaccharide (EPS) by Pseudomonas oleovorans NRRLB-14682 grown on glycerol,” Process Biochemistry, vol. 45, no.3, pp. 297–305, 2010.

[38] S. J. Yoon, Y. C. Choi, Y. I. Son, S. H. Lee, and J. G. Lee,“Gasification of biodiesel by-product with air or oxygen tomake syngas,” Bioresource Technology, vol. 101, no. 4, pp.1227–1232, 2010.

[39] M. Hajek and F. Skopal, “Treatment of glycerol phase formedby biodiesel production,” Bioresource Technology, vol. 101,no. 9, pp. 3242–3245, 2010.

[40] K. Zamzow, T. K. Tsukamoto, and G. C. Miller, “Biodieselwaste fluid as an inexpensive carbon source for bioreactorstreating acid mine drainage (California-Nevada, UnitedStates),” in Proceedings of the IMWA Symposium: Water inMining Environments, Cagliari, Italy, 2007.

[41] I. Bodık, A. Blstakova, S. Sedlacek, and M. Hutnan, “Biodieselwaste as source of organic carbon for municipal WWTPdenitrification,” Bioresource Technology, vol. 100, no. 8, pp.2452–2456, 2009.

[42] N. A. Krueger, R. C. Anderson, L. O. Tedeschi, T. R. Callaway,T. S. Edrington, and D. J. Nisbet, “Evaluation of feedingglycerol on free-fatty acid production and fermentationkinetics of mixed ruminal microbes in vitro,” BioresourceTechnology, vol. 101, no. 21, pp. 8469–8472, 2010.

[43] N. Rahmat, A. Z. Abdullah, and A. R. Mohamed, “Recentprogress on innovative and potential technologies for glyc-erol transformation into fuel additives: a critical review,”Renewable and Sustainable Energy Reviews, vol. 14, no. 3, pp.987–1000, 2010.

[44] M. Slinn, K. Kendall, C. Mallon, and J. Andrews, “Steamreforming of biodiesel by-product to make renewable hydro-gen,” Bioresource Technology, vol. 99, no. 13, pp. 5851–5858,2008.

[45] P. J. Lammers, B. J. Kerr, T. E. Weber et al., “Digestible andmetabolizable energy of crude glycerol for growing pigs,”Journal of Animal Science, vol. 86, no. 3, pp. 602–608, 2008.

[46] A. K. Forrest, R. Sierra, and M. T. Holtzapple, “Effectof biodiesel glycerol type and fermentor configuration on


mixed-acid fermentations,” Bioresource Technology, vol. 101,no. 23, pp. 9185–9189, 2010.

[47] S. Papanikolaou, S. Fakas, M. Fick et al., “Biotechnologicalvalorisation of raw glycerol discharged after bio-diesel (fattyacid methyl esters) manufacturing process: production of1,3-propanediol, citric acid and single cell oil,” Biomass andBioenergy, vol. 32, no. 1, pp. 60–71, 2008.

[48] C. E. Nakamura and G. M. Whited, “Metabolic engineeringfor the microbial production of 1,3-propanediol,” CurrentOpinion in Biotechnology, vol. 14, no. 5, pp. 454–459, 2003.

[49] K. K. Cheng, J. A. Zhang, D. H. Liu et al., “Pilot-scale pro-duction of 1,3-propanediol using Klebsiella pneumoniae,”Process Biochemistry, vol. 42, no. 4, pp. 740–744, 2007.

[50] W. D. Deckwer, “Microbial conversion of glycerol to 1,3-propanediol,” FEMS Microbiology Reviews, vol. 16, no. 2-3,pp. 143–149, 1995.

[51] C. J. Sullivan, D. C. Dehm, E. E. Reich, and M. E. Dillon,“Polyester resins based upon 2-methyl-1,3-propanediol,” TheJournal of Coatings Technology, vol. 62, pp. 37–45, 1990.

[52] C. Forschner, D. E. Gwyn, C. Frisch, J. Wang, P. Jackson,and A. Sendijarevic, Utilization of 1,3-Propanediol in Thermo-plastic Polyurethane Elastomers (TPUs), Shell Chemicals Ltd.,1999.

[53] U. Witt, R. J. Muller, J. Augusta, H. Widdecke, and W.D. Deckwer, “Synthesis, properties and biodegradabilityof polyesters based on 1,3-propanediol,” MacromolecularChemistry and Physics, vol. 195, pp. 793–802, 1994.

[54] A. P. Zeng and H. Biebl, “Bulk chemicals from biotechnology:the case of 1,3-propanediol production and the new trends,”Advances in biochemical engineering/biotechnology, vol. 74,pp. 239–259, 2002.

[55] S. Igari, S. Mori, and Y. Takikawa, “Effects of molecular struc-ture of aliphatic diols and polyalkylene glycol as lubricants onthe wear of aluminum,” Wear, vol. 244, no. 1-2, pp. 180–184,2000.

[56] Carlo Erba Reagents: 100th edition of Catalogue Chemicals,Section 2, Green Solvents—Sustainable chemistry, pp. 45–50,2004.

[57] F. Barbirato, E. H. Himmi, T. Conte, and A. Bories, “1,3-propanediol production by fermentation: an interesting wayto valorize glycerin from the ester and ethanol industries,”Industrial Crops and Products, vol. 7, no. 2-3, pp. 281–289,1998.

[58] B. Solomos, A.-P. Zeng, H. Biebl, H. Schlieker, C. Posten, andW.-D. Deckwer, “Comparison of the energetic efficiencies ofhydrogen and oxochemicals formation in Klebsiellapneumo-niae and Clostridium butyricum during anaerobic growth onglycerol,” Journal of Biotechnology, vol. 39, pp. 107–117, 1995.

[59] A. Hiremath, M. Kannabiran, and V. Rangaswamy, “1,3-Propanediol production from crude glycerol from jatrophabiodiesel process,” New Biotechnology, vol. 28, no. 1, pp. 19–23, 2011.

[60] T. Ito, Y. Nakashimada, K. Senba, T. Matsui, and N. Nishio,“Hydrogen and ethanol production from glycerol-containingwastes discharged after biodiesel manufacturing process,”Journal of Bioscience and Bioengineering, vol. 100, no. 3, pp.260–265, 2005.

[61] S. Adhikari, S. D. Fernando, and A. Haryanto, “Hydrogenproduction from glycerin by steam reforming over nickelcatalysts,” Renewable Energy, vol. 33, no. 5, pp. 1097–1100,2008.

[62] M. D. Blankschien, J. M. Clomburg, and R. Gonzalez,“Metabolic engineering of Escherichia coli for the production

of succinate from glycerol,” Metabolic Engineering, vol. 12, no.5, pp. 409–419, 2010.

[63] S. Ethier, K. Woisard, D. Vaughan, and Z. Wen, “Continu-ous culture of the microalgae Schizochytriumlimacinum onbiodiesel-derived crude glycerol for producing docosahexa-neoic acid,” Bioresource Technology, vol. 102, pp. 88–93, 2011.

[64] B. S. Fernandes, G. Peixoto, F. R. Albrecht, N. K. Saavedradel Aguila, and M. Zaiat, “Potential to produce biohydrogenfrom various wastewaters,” Energy for Sustainable Develop-ment, vol. 14, no. 2, pp. 143–148, 2010.

[65] M. Navratil, J. Tkac, J. Svitel, B. Danielsson, and E. Sturdık,“Monitoring of the bioconversion of glycerol to dihydroxy-acetone with immobilized gluconobacter oxydans cell usingthermometric flow injection analysis,” Process Biochemistry,vol. 36, no. 11, pp. 1045–1052, 2001.

[66] H. Hu and T. K. Wood, “An evolved Escherichia coli strain forproducing hydrogen and ethanol from glycerol,” Biochemicaland Biophysical Research Communications, vol. 391, no. 1, pp.1033–1038, 2010.

[67] A. Kivisto, V. Santala, and M. Karp, “Hydrogen productionfrom glycerol using halophilic fermentative bacteria,” Biore-source Technology, vol. 101, no. 22, pp. 8671–8677, 2010.

[68] K. O. Yu, S. W. Kim, and S. O. Han, “Engineering of glycerolutilization pathway for ethanol production by Saccharomycescerevisiae,” Bioresource Technology, vol. 101, no. 11, pp. 4157–4161, 2010.

[69] Z. Yuan, J. Wang, L. Wang et al., “Biodiesel derived glycerolhydrogenolysis to 1,2-propanediol on Cu/MgO catalysts,”Bioresource Technology, vol. 101, no. 18, pp. 7088–7092, 2010.

[70] Y. Zhu, J. Li, M. Tan et al., “Optimization and scale-upof propionic acid production by propionic acid-tolerantPropionibacterium acidipropionici with glycerol as the carbonsource,” Bioresource Technology, vol. 101, no. 22, pp. 8902–8906, 2010.

[71] A. Andre, P. Diamantopoulou, A. Philippoussis, D. Sarris,M. Komaitis, and S. Papanikolaou, “Biotechnological conver-sions of bio-diesel derived waste glycerol into added-valuecompounds by higher fungi: production of biomass, singlecell oil and oxalic acid,” Industrial Crops and Products, vol.31, no. 2, pp. 407–416, 2010.

[72] S. Papanikolaou and G. Aggelis, “Lipid production byYarrowia lipolytica growing on industrial glycerol in a single-stage continuous culture,” Bioresource Technology, vol. 82, no.1, pp. 43–49, 2002.

[73] S. Fakas, S. Papanikolaou, A. Batsos, M. Galiotou-Panayotou,A. Mallouchos, and G. Aggelis, “Evaluating renewable carbonsources as substrates for single cell oil production by Cun-ninghamella echinulata and Mortierella isabellina,” Biomassand Bioenergy, vol. 33, no. 4, pp. 573–580, 2009.

[74] A. Makri, S. Fakas, and G. Aggelis, “Metabolic activitiesof biotechnological interest in Yarrowia lipolytica grown onglycerol in repeated batch cultures,” Bioresource Technology,vol. 101, no. 7, pp. 2351–2358, 2010.

[75] S. Papanikolaou and G. Aggelis, “Biotechnological valoriza-tion of biodiesel derived glycerol waste through productionof single cell oil and citric acid by Yarrowia lipolytica,” LipidTechnology, vol. 21, no. 4, pp. 83–87, 2009.

[76] S. Papanikolaou and G. Aggelis, “Lipid production byYarrowia lipolytica growing on industrial glycerol in a single-stage continuous culture,” Bioresource Technology, vol. 82, no.1, pp. 43–49, 2002.

[77] S. Papanikolaou, L. Muniglia, I. Chevalot, G. Aggelis, and I.Marc, “Yarrowia lipolytica as a potential producer of citric


acid from raw glycerol,” Journal of Applied Microbiology, vol.92, no. 4, pp. 737–744, 2002.

[78] J. Lopez, M. Santos, A. Perez, and A. Martın, “Anaerobicdigestion of glycerol derived from biodiesel manufacturing,”Bioresource Technology, vol. 100, no. 23, pp. 5609–5615, 2009.

[79] I. Bodık, M. Hutnan, T. Petheoova, and A. Kalina, “Anaerobictreatment of biodiesel production wastes,” in Proceedingsof the Book of Lectures on 5th International Symposiumon Anaerobic Digestion of Solid Wastes and Energy Crops,Hammamet, Tunisia, 2008.

[80] M. Hutnan, N. Kolesarova, I. Bodık, V. Spalkova, and M.Lazor, “Possibilities of anaerobic treatment of crude glycerolfrom biodiesel production,” in Proceedings of 36th Interna-tional Conference of Slovak Society of Chemical Engineering,Tatranske Matliare, 2009.

[81] Y. Yang, K. Tsukahara, and S. Sawayama, “Biodegradationand methane production from glycerol-containing syntheticwastes with fixed-bed bioreactor under mesophilic andthermophilic anaerobic conditions,” Process Biochemistry,vol. 43, no. 4, pp. 362–367, 2008.

[82] M. S. Fountoulakis and T. Manios, “Enhanced methaneand hydrogen production from municipal solid waste andagro-industrial by-products co-digested with crude glycerol,”Bioresource Technology, vol. 100, no. 12, pp. 3043–3047, 2009.

[83] M. S. Fountoulakis, I. Petousi, and T. Manios, “Co-digestionof sewage sludge with glycerol to boost biogas production,”Waste Management, vol. 30, no. 10, pp. 1849–1853, 2010.

[84] J. Ma, M. Van Wambeke, M. Carballa, and W. Verstraete,“Improvement of the anaerobic treatment of potato pro-cessing wastewater in a UASB reactor by co-digestion withglycerol,” Biotechnology Letters, vol. 30, no. 5, pp. 861–867,2008.

[85] J. A. Alvarez, L. Otero, and J. M. Lema, “A methodology foroptimising feed composition for anaerobic co-digestion ofagro-industrial wastes,” Bioresource Technology, vol. 101, no.4, pp. 1153–1158, 2010.

[86] T. Amon, V. Kryvoruchko, B. Amon, and M. Schreiner,Untersuchungen zur Wirkung von Rohglycerin aus derBiodieselerzeugung als leistungssteigerndes Zusatzmittelzur Biogaserzeugung aus Silomais, Kornermais, Raps-presskuchen und Schweinegulle, Ergebnisbericht Depart-ment fur Nachhaltige Agrarsysteme Institut fur Landtechnik,Universitat fur Bodenkultur, Wien, Austria, 2004.

[87] V. Spalkova, M. Hutnan, and N. Kolesarova, “Selected prob-lems of anaerobic treatment of maize silage,” in Proceedingsof 36th International Conference of Slovak Society of ChemicalEngineering, Tatranske Matliare, 2009.

[88] T. Amon, B. Amon, V. Kryvoruchko, V. Bodiroza, E. Potsch,and W. Zollitsch, “Optimising methane yield from anaerobicdigestion of manure: effects of dairy systems and of glycerinesupplementation,” International Congress Series, vol. 1293,pp. 217–220, 2006.

[89] A. I. Qatibi, A. Bories, and J. L. Garcia, “Sulfate reduction andanaerobic glycerol degradation by a mixed microbial culture,”Current Microbiology, vol. 22, no. 1, pp. 47–52, 1991.

[90] A. I. Qatibi, R. Bennisse, M. Jana, and J.-L. Garcia, “Anaerobicdegradation of glycerol by Desulfovibrio fructosovorans and D.carbinolicus and evidence for glycerol-dependent utilizationof 1,2- propanediol,” Current Microbiology, vol. 36, no. 5, pp.283–290, 1998.

[91] C. Gallert and J. Winter, “Bacterial metabolism in wastewatertreatment systems,” in Environmental Biotechnology. Conceptsand Applications, H. J. Jordening and J. Winter, Eds., Wiley-VCH, Weinheim, Germany, 2005.

[92] B. K. Ahring, M. Sandberg, and I. Angelidaki, “Volatilefatty acids as indicators of process imbalance in anaerobicdigestors,” Applied Microbiology and Biotechnology, vol. 43,no. 3, pp. 559–565, 1995.

[93] Y. Chen, J. J. Cheng, and K. S. Creamer, “Inhibition of anaer-obic digestion process: a review,” Bioresource Technology, vol.99, no. 10, pp. 4044–4064, 2008.

[94] O. Lefebvre and R. Moletta, “Treatment of organic pollutionin industrial saline wastewater: a literature review,” WaterResearch, vol. 40, no. 20, pp. 3671–3682, 2006.

[95] G. D. Boardman, J. L. Tisinger, and D. L. Gallagher,“Treatment of clam processing wastewaters by means ofupflow anaerobic sludge blanket technology,” Water Research,vol. 29, no. 6, pp. 1483–1490, 1995.

[96] O. Lefebvre, N. Vasudevan, M. Torrijos, K. Thanasekaran,and R. Moletta, “Anaerobic digestion of tannery soak liquorwith an aerobic post-treatment,” Water Research, vol. 40, no.7, pp. 1492–1500, 2006.

[97] R. Gebauer, “Mesophilic anaerobic treatment of sludge fromsaline fish farm effluents with biogas production,” BioresourceTechnology, vol. 93, no. 2, pp. 155–167, 2004.

[98] O. Lefebvre, N. Vasudevan, M. Torrijos, K. Thanasekaran,and R. Moletta, “Halophilic biological treatment of tannerysoak liquor in a sequencing batch reactor,” Water Research,vol. 39, no. 8, pp. 1471–1480, 2005.

[99] M. Soto, R. Mendez, and J. M. Lema, “Biodegradabilityand toxicity in the anaerobic treatment of fish canningwastewaters,” Environmental Technology, vol. 12, no. 8, pp.669–677, 1991.

[100] F. Omil, R. Mendez, and J. M. Lema, “Anaerobic treatmentof seafood processing waste waters in an industrial anaerobicpilot plant,” Water SA, vol. 22, no. 1, pp. 173–181, 1996.

[101] G. Feijoo, M. Soto, R. Mendez, and J. M. Lema, “Sodiuminhibition in the anaerobic digestion process: antagonismand adaptation phenomena,” Enzyme and Microbial Technol-ogy, vol. 17, no. 2, pp. 180–188, 1995.

[102] Y. Liu and D. R. Boone, “Effects of salinity on methanogenicdecomposition,” Bioresource Technology, vol. 35, no. 3, pp.271–273, 1991.

[103] M. Soto, R. Mendez, and J. M. Lema, “Sodium inhibitionand sulphate reduction in the anaerobic treatment of musselprocessing wastewaters,” Journal of Chemical Technology andBiotechnology, vol. 58, no. 1, pp. 1–7, 1993.

[104] R. D. Sleator and C. Hill, “Bacterial osmoadaptation: therole of osmolytes in bacterial stress and virulence,” FEMSMicrobiology Reviews, vol. 26, no. 1, pp. 49–71, 2002.

[105] I. Vyrides and D. C. Stuckey, “Adaptation of anaerobicbiomass to saline conditions: role of compatible solutesand extracellular polysaccharides,” Enzyme and MicrobialTechnology, vol. 44, no. 1, pp. 46–51, 2009.

[106] S. Ramachandran, S. K. Singh, C. Larroche, C. R. Soc-col, and A. Pandey, “Oil cakes and their biotechnologicalapplications—a review,” Bioresource Technology, vol. 98, no.10, pp. 2000–2009, 2007.

[107] E. Molina-Alcaide and D. R. Yanez-Ruiz, “Potential use ofolive by-products in ruminant feeding: a review,” AnimalFeed Science and Technology, vol. 147, no. 1–3, pp. 247–264,2008.

[108] K. Franke, U. Meyer, H. Wagner, H. O. Hoppen, and G.Flachowsky, “Effect of various iodine supplementations,rapeseed meal application and two different iodine specieson the iodine status and iodine excretion of dairy cows,”Livestock Science, vol. 125, no. 2-3, pp. 223–231, 2009.


[109] T. Mushtaq, M. Sarwar, G. Ahmad et al., “Influence ofsunflower meal based diets supplemented with exogenousenzyme and digestible lysine on performance, digestibilityand carcass response of broiler chickens,” Animal Feed Scienceand Technology, vol. 149, no. 3-4, pp. 275–286, 2009.

[110] N. M. Soren and V. R. B. Sastry, “Replacement of soybeanmeal with processed karanj (Pongamia glabra) cake on thebalances of karanjin and nutrients, as well as microbialprotein synthesis in growing lamb,” Animal Feed Science andTechnology, vol. 149, no. 1-2, pp. 16–29, 2009.

[111] S. Bernesson, “Fields of application for the by-productsof extraction and tranesterification of rapeseed oil,” Tech.Rep. 1652-3237, Department of Biometry andEngineering,Uppsala, Sweden, 2007.

[112] J. M. Cervero, P. A. Skovgaard, C. Felby, H. R. Sørensen, andH. Jørgensen, “Enzymatic hydrolysis and fermentation ofpalm kernel press cake for production of bioethanol,” Enzymeand Microbial Technology, vol. 46, no. 3-4, pp. 177–184, 2010.

[113] T. Vastag, L. Popovic, S. Popovic, V. Krimer, and D. Pericin,“Production of enzymatic hydrolysates with antioxidant andangiotensin-I converting enzyme inhibitory activity frompumpkin oil cake protein isolate,” Food Chemistry, vol. 124,no. 4, pp. 1316–1321, 2011.

[114] J. N. Ferrer and E. A. Kaditi, “The EU added value of agri-cultural expenditure—from market to multifunctionality—gathering criticism and success stories of the CAP,” EuropeanParliament, Policy Department on Budgetary Affairs, Brus-sels, Belgium, 2007.

[115] Solar Oil Systems: Rape cake: more than just a by-product,2007, http://www.solaroilsystems.nl/publicaties/img/1184614337 2007-6-Rapeseed cake - more than just byproduct.pdf.

[116] International Fertilizer Industry Association (IFA) Task Forceon Bioenergy, “A survey of the anticipated impact of biofueldevelopment on short-, medium- and long-term fertilizerdemand,” in Proceedings of the 75th IFA Annual Conference,Istanbul, Turkey, 2007.

[117] S. Yorgun, S. Sensoz, and O. M. Kockar, “Flash pyrolysis ofsunflower oil cake for production of liquid fuels,” Journal ofAnalytical and Applied Pyrolysis, vol. 60, no. 1, pp. 1–12, 2001.

[118] F. Straka, P. Jenıcek, J. Zabranska, and M. Dohanyos,“Anaerobic fermentation of biomass and wastes with respectto sulfur and nitrogen contents in treate materials,” inProceedings of the11th International Waste Management andLandfill Symposium, Cagliari, Italy, 2007.

[119] G. Antonopoulou, K. Stamatelatou, and G. Lyberatos,“Exploitation of rapeseed and sunflower residues formethane generation through anaerobic digestion: the effectof pretreatment,” in Proceedings of the 2nd InternationalConference of Industrial Biotechnology, Padua, Italy, 2010.

[120] F. Raposo, R. Borja, M. A. Martın, A. Martın, M. A. de laRubia, and B. Rincon, “Influence of inoculum-substrate ratioon the anaerobic digestion of sunflower oil cake in batchmode: process stability and kinetic evaluation,” ChemicalEngineering Journal, vol. 149, no. 1–3, pp. 70–77, 2009.

[121] F. Raposo, R. Borja, B. Rincon, and A. M. Jimenez, “Assess-ment of process control parameters in the biochemicalmethane potential of sunflower oil cake,” Biomass andBioenergy, vol. 32, no. 12, pp. 1235–1244, 2008.

[122] M. A. De La Rubia, F. Raposo, B. Rincon, and R. Borja,“Evaluation of the hydrolytic-acidogenic step of a two-stage mesophilic anaerobic digestion process of sunflower oilcake,” Bioresource Technology, vol. 100, no. 18, pp. 4133–4138,2009.

[123] K. I. Suehara, Y. Kawamoto, E. Fujii, J. Kohda, Y. Nakano,and T. Yano, “Biological treatment of wastewater dischargedfrom biodiesel fuel production plant with alkali-catalyzedtransesterification,” Journal of Bioscience and Bioengineering,vol. 100, no. 4, pp. 437–442, 2005.

[124] M. Berrios and R. L. Skelton, “Comparison of purificationmethods for biodiesel,” Chemical Engineering Journal, vol.144, no. 3, pp. 459–465, 2008.

[125] M. Hayyan, F. S. Mjalli, M. A. Hashim, and I. M. AlNashef, “Anovel technique for separating glycerine from palm oil-basedbiodiesel using ionic liquids,” Fuel Processing Technology, vol.91, no. 1, pp. 116–120, 2010.

[126] J. Saleh, A. Y. Tremblay, and M. A. Dube, “Glycerol removalfrom biodiesel using membrane separation technology,” Fuel,vol. 89, no. 9, pp. 2260–2266, 2010.

[127] N. Sdrula, “A study using classical or membrane separation inthe biodiesel process,” Desalination, vol. 250, no. 3, pp. 1070–1072, 2010.

[128] P. Jaruwat, S. Kongjao, and M. Hunsom, “Management ofbiodiesel wastewater by the combined processes of chemicalrecovery and electrochemical treatment,” Energy Conversionand Management, vol. 51, no. 3, pp. 531–537, 2010.

[129] J. A. Siles, M. A. Martın, A. F. Chica, and A. Martın,“Anaerobic co-digestion of glycerol and wastewater derivedfrom biodiesel manufacturing,” Bioresource Technology, vol.101, no. 16, pp. 6315–6321, 2010.

[130] O. Chavalparit and M. Ongwandee, “Optimizing electroco-agulation process for the treatment of biodiesel wastewaterusing response surface methodology,” Journal of Environmen-tal Sciences, vol. 21, no. 11, pp. 1491–1496, 2009.

[131] S. Kato, H. Yoshimura, K. Hirose, M. Amornkitbamrung, M.Sakka, and I. Sugahara, “Application of microbial consortiumsystem to wastewater from biodiesel fuel generator,” IEA-Waterqual, Singapore, CD-ROM ,2005.

[132] S. Papanikolaou, I. Chevalot, M. Komaitis, G. Aggelis, andI. Marc, “Kinetic profile of the cellular lipid compositionin an oleaginous Yarrowia lipolytica capable of producinga cocoa-butter substitute from industrial fats,” Antonie vanLeeuwenhoek, vol. 80, no. 3-4, pp. 215–224, 2001.

[133] S. Papanikolaou and G. Aggelis, “Selective uptake of fattyacids by the yeast Yarrowia lipolytica,” European Journal ofLipid Science and Technology, vol. 105, no. 11, pp. 651–655,2003.

[134] S. Papanikolaou and G. Aggelis, “Modeling lipid accumu-lation and degradation in Yarrowia lipolytica cultivated onindustrial fats,” Current Microbiology, vol. 46, no. 6, pp. 398–402, 2003.


Research Article

A Simple and Fast Method for the Production andCharacterization of Methylic and Ethylic Biodieselsfrom Tucum Oil via an Alkaline Route

Marcelo Firmino de Oliveira,1 Andressa Tironi Vieira,2 Antonio Carlos Ferreira Batista,3

Hugo de Souza Rodrigues,3 and Nelson Ramos Stradiotto2

1 Departamento de Quımica, Faculdade de Filosofia, Ciencias e Letras de Ribeirao Preto, Universidade de Sao Paulo,14040-901 Ribeirao Preto, SP, Brazil

2 Departamento de Quıımica Analıtica, Instituto de Quımica, UNESP, 14800-900 Araraquara, SP, Brazil3 Laboratorio de energias Renovaveis e Meio ambiente do Pontal (LERMAP), UFU, 38302-000 Ituiutaba, MG, Brazil

Correspondence should be addressed to Marcelo Firmino de Oliveira, [email protected]

Received 31 August 2010; Revised 6 February 2011; Accepted 9 March 2011

Academic Editor: Stanley Brul

Copyright © 2011 Marcelo Firmino de Oliveira et al. This is an open access article distributed under the Creative CommonsAttribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work isproperly cited.

A simple, fast, and complete route for the production of methylic and ethylic biodiesel from tucum oil is described. Aliquots ofthe oil obtained directly from pressed tucum (pulp and almonds) were treated with potassium methoxide or ethoxide at 40◦C for40 min. The biodiesel form was removed from the reactor and washed with 0.1 M HCl aqueous solution. A simple distillation at100◦C was carried out in order to remove water and alcohol species from the biodiesel. The oxidative stability index was obtainedfor the tucum oil as well as the methylic and ethylic biodiesel at 6.13, 2.90, and 2.80 h, for storage times higher than 8 days.Quality control of the original oil and of the methylic and ethylic biodiesels, such as the amount of glycerin produced duringthe transesterification process, was accomplished by the TLC, GC-MS, and FT-IR techniques. The results obtained in this studyindicate a potential biofuel production by simple treatment of tucum, an important Amazonian fruit.

1. Introduction

The search for alternative fuels to replace oil-derived energysources has received increased attention worldwide over thelast decades, more specifically since the oil crisis in the20th century. Fossil fuels constitute a finite energy source,and they emit a large amount of pollutants [1]. Interestingalternatives have been presented as possible substitutes forfossil fuels, such as fuel ethanol [1] and biofuels fromanimal and plant oils. Nowadays, these oils appear as apromising replacement for diesel oil, and special attentionhas been devoted to biodiesel [2]. This biofuel consists ofmonoalkylesthers of long chain fatty acids, derived fromplant or animal oils [2]. A schematic diagram for theproduction of biodiesel from triacylglycerols is shown inFigure 1.

Biodiesel production offers several advantages, since thisfuel is renewable and environmentally friendly, can be mixedwith diesel, not to mention its numerous animal [3–7] andplant sources [8–14].

Of all the reported sources of plant oils for biodieselproduction, tucum (Astrocaryum vulgate Mart) must behighlighted [15]. The fruit of tucum trees is extensively usedby Amazonian communities for the production of food andcosmetics [15–17] among other applications.

The oil is commonly extracted from tucum by pressingthe entire fruit (pulp and almonds together). Some scientificstudies about its use as a biofuel have been reported [16, 17].In terms of chemical composition, the tucum oil is rich inbetacarotene [16] and contains oleic and palmitic acids asthe major fatty acids, apart from a small level of linoleic acid[15]. This oil is considered an important raw material for


O C

O

O

OC

C

O

O

R3R2

R1

HOR

C

O

R3

C

OR2

C

OR1 OH

HO

HORO

RO

RO

Solvent

Catalyst++

Figure 1: Schematic diagram of triacylglycerol transesterificationby alcoholysis, in the presence of catalysts.

the production of oleochemicals, and it is stable to oxidativedeterioration.

Despite its interesting advantages from an oleochemicalview point, application of tucum oil as a raw material forbiodiesel production has not been extensively reported. Inthis context, Lima et al. [18] studied the production ofbiodiesel from tucum by transesterification with anhydrousethanol and methanol, using NaOH as catalyst. The authorsconcluded that the properties of the two types of biodiesel(ethanolic and methanolic) were very similar to those of oil-derived diesel.

In this context, research on biodiesel production fromtucum fruit is justified, once this constitutes an importantmotivational factor for Amazonian rural communities. Inthis way, the Amazonian rural population could obtain theirown biofuel in a sustainable fashion. Therefore, the aimof this work was to develop a simple, fast and efficientalkaline (KOH) route for the production of methylic andethylic biodiesels from tucum oil, extracted from Amazoniantrees by local communities in a sustainable way, in order toprovide a local biofuel production.


2.1. Biodiesel Production. The tucum oil was obtained bypressing the entire fruit (pulp and almonds) in a conven-tional mechanic press. The obtained oil was filtered througha glass tube of 1 cm diameter filed with cotton. Potassiummethoxide and potassium ethoxide were prepared usingpotassium hydroxide P.A., and methanol P.A., and ethanolP.A. purchased from Sinth, respectively, following a similarexperimental process described in the literature [18]. Themethoxide form was obtained after adding 9.5 g of KOHat 120 mL of methanol under stirring until the completedissolution (exothermic reaction). The ethoxide form alsowas obtained after adding 9.5 g of KOH at 150 mL of ethanolunder stirring until the complete dissolution.

Each aliquot the of oil-alcohol-KOH mixture (methylicor ethylic form) was kept under stirring at 40◦C for40 min. The transesterification process was monitored bythin layer chromatography: the development of separation inhexane/ethyl acetate 95%/5% solution can exhibit a decreaseof oil band and increase of biodiesel bands through the time.

After the decantation step, each biodiesel form wasseparated from glycerin and washed with HCl 0.1 M solution.A simple distillation at 100◦C was performed, in order toremove both water and alcohol from the biodiesel samples.

123456789

1011121314

Abu

nda

nce

4 6 8 10 12 14 16 18 20 22

Time

Lau

ric Mir

isti

c Pal

mit

ic Ole

ic

×104

Figure 2: Chromatogram with mass spectrum identificationobtained for the free fatty acids of tucum oil-identification of itsmajor oleochemical components.

2.2. Oxidative Stability Studies. The importance of this studyconsists of finding out up to which time point there isno formation of secondary compounds of oxidation, andto establish when the onset of increasing oxidation rate,peroxide index, oxygen absorption, and formation of volatilesubstances take place.

The oxidative stability measurements were carried outusing a 873 Rancimat equipment from Metrohm. The tucumoil and the methylic and ethylic biodiesel samples weresubmitted to an air gas flow of 10 L/h, under a continuousheating at 110◦C ± 0.3◦C.

2.3. GC-MS Analysis. The chromatographic analysis of thebiodiesel samples such as the original oil and of the glycerinobtained during the transesterification process was carriedout using an HP gas chromatograph, model CG 5890, seriesII, equipped with an HP1 column (100% dimethyl polysilox-ane) with 30 m length and 0.2 mm internal diameter. Themobile phase consisted of H2 and N2 (30 L/min) and air(300 L/min). An injection volume of 0.5 µL was used inall the measurements. An injector temperature of 200◦Cwas employed, and analysis was accomplished by using atemperature ramp from 80 to 200◦C. The mass spectraof the main chromatographic peaks monitored in a massspectrometer HP model 5988A, which was coupled to thechromatograph.

2.4. FT-IR Analysis. The spectroscopic measurements for allthe studied species were recorded on an infrared spectropho-tometer from Perkin Elmer, model 1430, in the spectral workrange from 4500 to 450 cm−1.


The methylic route was more efficient for biodiesel produc-tion (75.1%) than the ethylic route (66.7%). Both resultsallowed us to propose this methodology as an alternativeroute for biodiesel production. The presence of Lauric,Miristic, Palmitic, and Oleic fatty acids as the major com-ponents of the tucum oil, detected retention times range


00

102030405060708090

1 2 3 4 5 6 7 8

(h)

(µS/

cm)

6.13

(a)

0 1 2 3 4 5 6 7 80

255075

100125150175 2.9

(µS/

cm)

(h)

(b)

0 1 2 3 4 5 6 70

255075

100125150175200

(µS/

cm)

2.8

(h)

(c)

Figure 3: oxidative stability index measurements for: (a) tucum oil,(b) methylic biodiesel, and (c) ethylic biodiesel.

from 12 to 23 min, was confirmed by GC-MS, as indicatedin Figure 2, in good agreement with previous works reportedin the literature [15–18].

Biodiesel production was also confirmed by the FT-IRtechnique. In this case, it is possible to check the locationof the bands of the carbonyls of the oils in relation to theones of the biodiesel. The main difference observed betweenthe infrared spectrum of tucum oil and those of its biodieselswas a small displacement of the C=O stretching band of thebiodiesel samples to lower energy (1747–1740 cm−1), whichcan be explained in terms of the substitution of glycerol bythe methoxilic radical [18].

The studies of oxidative stability index indicated loss ofchemical stability for the biodiesels in relation to the originaloil, as indicated in Figure 3. The results about stabilityindex and storage time are reported in Table 1. While itwas possible to observe a stability time of 6.13 h for theoil samples, only 2.90 h and 2.80 h were observed for themethylic and ethylic biodiesels. According to Brazilian laws,for example [19], the indexes obtained for biodiesels in thiswork are under the recommended values (6 h at 110◦C). Theoil form is more stable because it is rich in carotenoids.

The storage time differs from the oxidative stabilityindex values once no induced oxidation was provided or

Table 1: Oxidative stability index and storage time studies obtainedfor the tucum oil and the methylic and ethylic biodiesels.

Sample Oxidative stability index(h)

Storage time at 20◦C(days)

Tucum oil 6.13 35.2

Methylic biodiesel 2.90 9.52

Ethylic biodiesel 2.80 8.73

accelerated. It consists in the storage time at 20◦C. In thiscase, it was possible to observe a storage time of 35 days forthe tucum oil, which is possible due to the presence of waterand the high acidity of the sample.

4. Conclusions

The proposed methodology successfully produced methylicand ethylic biodiesel from the tucum oil using KOH asreagent for alkoxide production. In spite of being used inseveral ways by local Amazonian communities, the tucumoil can also be converted into its biodiesel form, consistingof an interesting self-production route for biofuel and thuscontributing to sustainable exploitation of this species. Theoxidative stability indexes and storage time make this oil agood precursor for local biodiesel production, even thoughfurther studies are needed to establish its application in dieselengines, for example.

Acknowledgment

The authors thank the financial support of CAPES, CNPq,FINEP, and FAPEMIG.

References

[1] M. F. de Oliveira, A. A. Saczk, L. L. Okumura, and N. R.Stradiotto, “Analytical methods employed at quality control offuel ethanol,” Energy and Fuels, vol. 23, no. 10, pp. 4852–4859,2009.

[2] F. C. C. Oliveira, P. A. Z. Suarez, and W. L. P. Santos, “Biodiesel:possibilidades e desafios,” Quimica Nova na Escola, vol. 28, pp.3–8, 2008.

[3] O. Andersen and J. E. Weinbach, “Residual animal fat and fishfor biodiesel production. Potentials in Norway,” Biomass andBioenergy, 2010.

[4] Y. Liang, N. Sarkany, Y. Cui, and J. W. Blackburn, “Batch stagestudy of lipid production from crude glycerol derived fromyellow grease or animal fats through microalgal fermentation,”Bioresource Technology, vol. 101, no. 17, pp. 6745–6750, 2010.

[5] C. L. Bianchi, D. C. Boffito, C. Pirola, and V. Ragaini, “Lowtemperature de-acidification process of animal fat as a pre-stepto biodiesel production,” Catalysis Letters, vol. 134, no. 1-2, pp.179–183, 2010.

[6] I. Rivera, G. Villanueva, and G. Sandoval, “Produccionde biodiesel a partir de residuos grasos animales por vıaenzimatica,” Grasas y Aceites, vol. 60, no. 5, pp. 468–474, 2009.

[7] C. Oner and S. Altun, “Biodiesel production from inedibleanimal tallow and an experimental investigation of its use asalternative fuel in a direct injection diesel engine,” AppliedEnergy, vol. 86, no. 10, pp. 2114–2120, 2009.


[8] N. C. Om Tapanes, D. A. Gomes Aranda, J. W. de MesquitaCarneiro, and O. A. Ceva Antunes, “Transesterification ofJatropha curcas oil glycerides: theoretical and experimentalstudies of biodiesel reaction,” Fuel, vol. 87, no. 10-11, pp.2286–2295, 2008.

[9] C. da Silva, F. de Castillos, J. V. Oliveira, and L. C. Filho, “Con-tinuous production of soybean biodiesel with compressedethanol in a microtube reactor,” Fuel Processing Technology,vol. 91, pp. 1274–1281, 2010.

[10] H. Joshi, B. R. Moser, S. N. Shah, A. Mandalika, and T. Walker,“Improvement of fuel properties of cottonseed oil methylesters with commercial additives,” European Journal of LipidScience and Technology, vol. 112, no. 7, pp. 802–809, 2010.

[11] J. A. Melero, L. F. Bautista, G. Morales, J. Iglesias, and R.Sanchez-Vazquez, “Biodiesel production from crude palmoil using sulfonic acid-modified mesostructured catalysts,”Chemical Engineering Journal, vol. 161, no. 3, pp. 323–331,2010.

[12] J. Zhang, S. Chen, R. Yang, and Y. Yan, “Biodiesel productionfrom vegetable oil using heterogenous acid and alkali catalyst,”Fuel, vol. 89, pp. 2939–2944, 2010.

[13] M. Peri and L. Baldi, “Vegetable oil market and biofuel policy:an asymmetric cointegration approach,” Energy Economics,vol. 32, no. 3, pp. 687–693, 2010.

[14] A. Demirbas, “New biorenewable fuels from vegetable oils,”Energy Sources Part A, vol. 32, no. 7, pp. 628–636, 2010.

[15] F. O. J. Oboh, “The oleochemical potential of tucum (Astro-caryum vulgare Mart) pulp oil and stearin,” Rivista Italianadelle Sostanze Grasse, vol. 86, no. 1, pp. 39–47, 2009.

[16] D. B. Rodriguez-Amaya, J. Amaya-Farfan, and M. Kimura,“Carotenoid composition of Brazilian fruits and vegetables,”Acta Horticulturae, vol. 744, pp. 409–416, 2007.

[17] F. O. J. Oboh and R. A. Oderinde, “Analysis of the pulp andpulp oil of the tucum (Astrocaryum vulgare Mart) fruit,” FoodChemistry, vol. 30, no. 4, pp. 277–287, 1988.

[18] J. R. D. O. Lima, R. B. da Silva, E. Miranda de Moura, andC. V. Rodarte de Moura, “Biodiesel of tucum oil, synthesizedby methanolic and ethanolic routes,” Fuel, vol. 87, no. 8-9, pp.1718–1723, 2008.

[19] EN 14112:2003, “Fat and Oil Derivatives—Fatty Acid MethylEsters (FAME),” Determination of oxidation stability (acceler-ated oxidation test), 2003.

journal of biomedicine and biotechnologydownloads.hindawi.com/journals/focusissues/748976.pdfthe...

Documents