Combining phospholipases and a liquid lipase for one-step biodiesel production using crude oils
© Cesarini et al.; licensee BioMed Central Ltd. 2014
Received: 23 October 2013
Accepted: 10 February 2014
Published: 26 February 2014
Enzymatic biodiesel is becoming an increasingly popular topic in bioenergy literature because of its potential to overcome the problems posed by chemical processes. However, the high cost of the enzymatic process still remains the main drawback for its industrial application, mostly because of the high price of refined oils. Unfortunately, low cost substrates, such as crude soybean oil, often release a product that hardly accomplishes the final required biodiesel specifications and need an additional pretreatment for gums removal. In order to reduce costs and to make the enzymatic process more efficient, we developed an innovative system for enzymatic biodiesel production involving a combination of a lipase and two phospholipases. This allows performing the enzymatic degumming and transesterification in a single step, using crude soybean oil as feedstock, and converting part of the phospholipids into biodiesel. Since the two processes have never been studied together, an accurate analysis of the different reaction components and conditions was carried out.
Crude soybean oil, used as low cost feedstock, is characterized by a high content of phospholipids (900 ppm of phosphorus). However, after the combined activity of different phospholipases and liquid lipase Callera Trans L, a complete transformation into fatty acid methyl esters (FAMEs >95%) and a good reduction of phosphorus (P <5 ppm) was achieved. The combination of enzymes allowed avoidance of the acid treatment required for gums removal, the consequent caustic neutralization, and the high temperature commonly used in degumming systems, making the overall process more eco-friendly and with higher yield. Once the conditions were established, the process was also tested with different vegetable oils with variable phosphorus contents.
Use of liquid lipase Callera Trans L in biodiesel production can provide numerous and sustainable benefits. Besides reducing the costs derived from enzyme immobilization, the lipase can be used in combination with other enzymes such as phospholipases for gums removal, thus allowing the use of much cheaper, non-refined oils. The possibility to perform degumming and transesterification in a single tank involves a great efficiency increase in the new era of enzymatic biodiesel production at industrial scale.
KeywordsEnzymatic biodiesel Liquid lipase Crude oils Phospholipases
Lipase-catalyzed biodiesel production is an intensive area of research because of its potential to generate eco-friendly fuel. Indeed, employment of lipases as biocatalysts in the transesterification of triacylglycerides allows mild reaction conditions and easy recovery of glycerol, without need for further purification or chemical waste production. In addition, the enzymatic process tolerates the common water content of oil and increases the biodiesel yield, avoiding the typical soap formation due to alkaline transesterification . Various lipases from different sources, generally in their immobilized form, have been investigated for transesterification using several substrates . To date, high lipase activity and stability, with very good conversion rates in short reaction times, have been reported [3–6]. However, the high costs of enzyme immobilization remain a major drawback for lipase-catalyzed biodiesel production. The immobilization process and the supports used make the enzymatic transesterification economically not competitive against the chemical process [3, 7]. According to Ghaly et al. (2010), the cost of the enzyme makes up to 90% of the total cost of the enzymatic process . Moreover, lipase inhibition due to binding of the released glycerol to the supports has been reported . In order to make the process more competitive and sustainable, the possibility of using soluble, non-immobilized lipases in biodiesel production is being investigated nowadays [9–11]. Soluble lipases can also be used under mild reaction conditions, have a faster reaction time, display higher conversion rates than immobilized enzymes , and allow, without any fatty acid methyl esters (FAMEs) yield loss, the presence of water in the process, which is required for oil pretreatment or during downstream steps . Furthermore, liquid lipases can be produced and sold at a much lower price [12, 13], and can also be re-used after recovery from the glycerin phase. For instance, Callera Trans L maintained up to 95% activity after recycling (AR Madsen, personal communication). But in contrast with immobilized lipases, the necessary number of re-uses of liquid lipases could be much lower due to the large difference in price of both types of enzymes. The use of liquid instead of immobilized enzyme has resulted in a significant simplification of oil transesterification and is the background for the new, cost-effective process developed by Novozymes (Bagsværd, Denmark) using liquid lipase Callera Trans L, which is now being scaled-up at several production plants for biodiesel synthesis from different feedstocks .
Nowadays, biodiesel is mainly produced by transesterification of edible oils such as those from soybean, rapeseed, sunflower, or palm . The cost of feedstocks is another important economical factor in biodiesel production, which indicates that selection of the appropriate raw material is of major importance for ensuring the feasibility of the process at industrial scale . Thus, exploring the use of low value or non-edible feedstocks is a goal for biodiesel producing partners. In this context, non-refined (crude) soybean oil is an alternative solution already investigated for enzymatic transesterification [6, 11, 17]. However, crude, non-degummed oils contain impurities as phosphorous compounds (phospholipids), which have to be removed for efficient transesterification. Crude soybean oil generally has a phosphorus (gums) content of 800 to 1,200 ppm, equivalent to 2 to 3% of phospholipids , which can cause problems during storage due to precipitate formation and water accumulation. Moreover, according to the legal specifications, phosphorus concentration in final biodiesel must be reduced to less than 10 ppm (EN14214:2003; ASTM D6751:2012). For biodiesel production, a high content of phospholipids in the raw feedstocks means a concomitant loss of yield in FAMEs production, as the fatty acids enclosed into the phospholipid molecules are not accessible to the lipase for transesterification. Phospholipids are commonly removed by previously developed refining/degumming processes of physical or chemical nature. In the 1990s the first industrial enzymatic oil-degumming process was launched with EnzyMax® (Lurgi AG, Frankfurt, Germany), based on phospholipase conversion of non-hydratable phospholipids (NHPs) to hydratable lyso-phospholipids . NHPs, which consist of phospholipids combined with metal cations, such as Ca2+ or Mg2+, can be chemo-enzymatically converted into hydratable compounds and simply removed by water washing and centrifugation . The easily hydratable phospholipids are phosphocholine (PC), phosphatidylinositol (PI), and lysophosphatidylcholine (LPC), whereas phosphatidylethanolamine (PE) and phosphatidic acid (PA) are generally considered as NHPs . A guideline for industrial enzymatic degumming was reported by Cowan and Nielsen in 2008, suggesting a first step of citric acid treatment, where the acid is distributed with a high-shear mixer in the oil at high temperatures, to chelate the metals and to open the phospholipid micelles; following the citric acid treatment, the oil is cooled down to the optimal temperature for a final phospholipase treatment. Water (up to 1.5 to 2.5%) is added together with the enzyme and sodium hydroxide, and the enzymatic reaction is carried out for 2 to 6 h .
Different types of microbial phospholipases have been described depending on the acyl ester bond they hydrolyze in the phospholipid molecule , which could be applied to enzymatic degumming. Phospholipase A1 (PLA1), which catalyzes the hydrolysis of the fatty acid at position 1 in the phospholipid molecule, releasing a lyso-phospholipid and a free fatty acid (FFA), has already been applied to the degumming process [18, 22, 23] also in combination with phospholipase C (PLC), which cleaves the phosphorus–oxygen bond between glycerol and phosphate, releasing a diacylglycerol (DAG) and the phosphate ester group .
Properties and working conditions in transesterification and degumming of all enzymes used
Callera Trans® L (Novozymes A/S)
Lecitase® Ultra (Novozymes A/S)
LLPL-2 (Novozymes A/S)
4.5 to 5.5
4.5 to 4.8
50 to 55°C
40 to 45°C
4 to 6 h
250 to 500 ppm
2 to 3.5%
1 to 4%
0.065% Citric acid/0.025% Phosphoric acid
2 eqs NaOH
Results and discussion
Citric acid effect on lipase activity
Experimental plan and results (FAMEs release) from testing acid, NaOH addition, and NaOH excess additions
Extra NaOH (ppm)
Effect of methanol on degumming
The second main parameter to be investigated for the combined degumming and transesterification reaction was the possible phospholipase inhibition due to the presence of methanol in the reaction mixture. In order to preserve the stability of the lipase, methanol was pumped into the reaction mixture following a slow continuous gradient of 0.4 ml/h during 10 h, for a total amount of 1.5 molar equivalents of methanol to the total fatty acids (in glycerides and FFAs) in the oil. Taking into consideration that during an efficient transesterification reaction methanol is consumed by the lipase to form methyl esters, this compound is never present in the reaction mixture at the amount of 1.5 eqs. Knowing that 1 eq of methanol is required for a theoretical complete transesterification, the maximum amount of methanol left in the system that might inhibit phospholipases, would be 0.5 eqs.
Phosphorus content of samples treated or untreated with methanol
AD + PLA1c
AD + PLCc
AD + PLA1 + PLCc
AD + PLA1 + LLPL-2c
Given the widespread use of soybean oil in industrial biodiesel production, the evidence of solubilization of phospholipids by methanol acquires a great importance in the process. It means that gums can be dissolved by methanol, with no requirement for a conventional acid treatment, thus making the released phospholipids more available for enzymatic transesterification, that is, methanol seems to break apart the micelles. Therefore, presence of methanol, used here as a substrate, may allow to abolish the acid degumming step without any loss of performance. This assumption was demonstrated with the combination of enzymatic degumming and transesterification, performed without citric acid treatment, as described in the following sections.
Combining enzymatic degumming and transesterification
Oil degumming is a requirement to obtain refined, edible oils, but it is also essential for biodiesel production. For immobilized Candida antarctica lipase, Watanabe and co-workers reported that crude (non-degummed) oil does not undergo enzymatic-catalyzed methanolysis . Depending on the raw materials used, degumming becomes an indispensable step for biodiesel production to achieve phospholipids removal and to reduce the final phosphorus content below the specified limits. In addition to an extra tank, the degumming process involves the use of acids and high temperatures, all factors boosting the process costs. Moreover, during the degumming process there is an unavoidable loss of oil that migrates to the gums during removal. For instance, for crude soybean oil containing an average 900 ppm P, gums represent a 2.5% loss of total oil; being the current market price US$1,100 per ton, this corresponds to a loss of US$27.5 per ton of oil. These drawbacks could be overcome with the unification of degumming and transesterification in the same tank . For this purpose, a single-step enzymatic degumming and transesterification process using phospholipases and liquid lipase Callera Trans L, with no need for a conventional acid degumming treatment, could provide a solution to such problems.
Biodiesel resulting from the combined degumming/transesterification process
85.2 ± 1.4
823 ± 56
PLA1 + TE
98.2 ± 2.1
8.0 ± 3.0
PLC + TE
90.8 ± 4.0
12.8 ± 1.8
PLA1 + LLPL-2 + TE
97.8 ± 0.3
6.0 ± 3.0
PLA1 + PLC + TE
96.6 ± 0.8
4.6 ± 1.7
Study of phospholipases activity
Analysis of phosphatides by ultra-performance liquid chromatography tandem mass spectrometry (UPLC/MS/MS) in the oil and glycerin phases obtained from the combined reactions, confirmed the enzymatic activity of phospholipases, excluding the possibility that the phospholipids present in the original raw material were separated to the glycerin phase or to the interphase as entire molecules, without any gain of oil for further lipase-catalyzed transesterifications.
The final glycerin composition of all samples was analyzed for each type of phosphorous compound, and the mode of action of the phospholipases used could be confirmed (see Additional files 1, 2, 3, 4, and 5). As expected, PC (RT = 5 min; m/z 184) could be found only in the glycerin phase derived from the reactions where PLC was present (PLC + TE and PLA1 + PLC + TE), proving the specific activity of this phospholipase for the phosphodiester bond (see Additional file 3 and Additional file 5). Instead, as a result of PLA1 activity, lyso-phosphocholine should appear at RT = 1.88 min, extracting ions 496.3, 518.3, 520.3, and 522.3 m/z (corresponding to LPC bearing the 16:0, 18:1, 18:2, or 18:3 fatty acids, respectively), but it could also be detected in very small amounts in all glycerin samples, suggesting a kind of instability of the LPC molecule. In fact, looking at glycerol-phosphocholine (RT = 11 min; m/z 258), this product should be present only in the glycerin phase of the reaction PLA1 + LLPL-2 + TE, resulting from the combined activity of these enzymes. Nevertheless, it was also found in all other samples, supporting the hypothesis of an extra activity of PLA1 in processing the fatty acid at position sn-2, or being the consequence of a kind of instability of the LPC molecule (see Additional files 1, 2, 3, 4, and 5). However, LLPL-2 activity could be confirmed by the peak of glycerol-phosphocholine found in sample PLA1 + LLPL-2 + TE, which was in fact higher than that found for the other samples (see Additional file 4). From these results we can conclude that parallel analysis by UPLC/MS/MS of the phosphatides found in the oil and glycerin phases, resulting from the combined reactions performed, turned out as a powerful tool for the study of such a complex system, and allowed to isolate the activity of each enzyme applied, confirming the importance of phospholipases in the combined degumming and transesterification process.
Combined degumming/transesterification process using crude, difficult vegetable oils
Combined degumming/transesterification biodiesel process applied to difficult oils
PLA1 + TE
PLC + TE
PLA1 + LLPL-2 + TE
PLA1 + PLC + TE
When crude canola oil was used, a high transesterification rate was achieved but the final phosphorus content was not reduced below 130 ppm (Table 5). This oil is known to have a very high content of Ca2+ and Mg2+, and is especially difficult to degum. Crude canola oil is considered one of the most difficult oils for degumming because it is mostly composed by NHPs. These NHPs are barely attacked by phospholipases without the aid of an acid treatment. Moreover, they need a strong chelating agent such as citric acid at higher concentrations (0.1%) to achieve a suitable phosphorus reduction. These conditions can hardly be reached with milder conditions of citric acid or even with phosphoric acid (D Cowan, personal communication), thus making it difficult to reach the required phosphorus content in the final biodiesel.
A successful, completely enzymatic process has been investigated resulting in a more economic and eco-friendly biodiesel production. Combination of crude oil degumming and transesterification in a unique step is possible by using phospholipases and liquid lipase Callera Trans L. In the combined process, an important cost reduction can be achieved. In addition to the US$27.5 per ton savings in the case of soybean oil, costs can be substantially lowered by avoiding the extra tank commonly required for oil degumming pretreatments, and by using mild temperatures (35°C). Moreover, citric acid treatment has been eliminated and only low sodium hydroxide concentrations are used, thus increasing the savings of the whole process. Therefore, the developed method meets the conditions for being easily scaled-up and is suitable for most crude vegetable oils.
Crude soybean oil (FFA = 1%; P = 900 ppm; pH = 6.8) was obtained from Cargill (Iowa Falls, IA, USA). Corn oil from bioethanol production (FFA = 6%; P = 62 ppm; pH = 4.5) was provided by Blue Sun (St Joseph, MO, USA). Crude canola oil (FFA = 1%; P = 250 ppm; pH = 5.7) was kindly donated by Richardson (Winnipeg, MB, Canada). Crude soybean oil was chosen as a cheaper raw material and because it is still the most commonly used oil in the industrial biodiesel production. Corn oil was considered interesting for this study because it derives, as a residue, from the bioethanol industrial production, thus to close the hypothesis to unify both bioethanol/biodiesel production processes. Finally, crude canola oil was tested to verify the potentialities of the suggested process towards oils rich in NHPs; the canola oil used here contained approximately 130 ppm of P in NHPs over the total 250 ppm of initial P content.
Enzymes and chemicals
Soluble lipase Callera Trans L, phospholipase A1, PLA1 (Lecitase® Ultra), and lyso-phospholipase, LLPL-2 (patent WO 2001027251 A1) used in this work were from Novozymes A/S. Phospholipase C, PLC (Purifine®) was purchased from Verenium (San Diego, CA, USA). All chemicals used were from Sigma Aldrich (St Louis, MO, USA).
When only enzymatic degumming was performed, an adaptation of the Cowan and Nielsen protocol was used]. Acid treatment was performed by adding citric acid (0.065%) to 20 g of pre-heated oil (55°C) and mixed with an Ultra Turrax® T25 (IKA, Staufen, Germany) for 10 s at 12,000 rpm. The emulsion was incubated for 30 min at 55°C and 250 rpm in a horizontal shaker. Caustic neutralization was performed with addition of NaOH (2 eqs to citric acid) and 3.5% water. Enzymatic degumming was completed by applying phospholipases, with the recommended dosage indicated in Table 1. Lyso-phospholipase was used only in combination with PLA1 at a concentration of 400 ppm. Combined degumming/transesterification reactions were run at 35°C for 24 h with 250 rpm agitation, according to the optimum transesterification conditions, instead of the recommended incubation for degumming of 55°C for 2 h.
FAMEs synthesis reactions were carried out in 100 ml squared bottles for 24 h at 35°C with 250 rpm agitation in a horizontal shaker. The reaction mixtures consisted of 20 g oil, 1% w/w Callera Trans L lipase solution, 3.5% H2O, and 10 ppm of NaOH. Total methanol (MeOH) per reaction was 1.5 eqs of oil, added continuously by a syringe pump system (SP220 IZ, WPI) with a flow rate of 0.4 ml/h during 10 h.
One-step enzymatic degumming and transesterification
All enzymes (lipase and phospholipases) were added at a time to 20 g of oil. Reaction mixtures included water (3.5%), NaOH (10 ppm), and 1.5 eqs of MeOH, pumped in a linear gradient for 10 h. Combination of degumming and transesterification was incubated following the transesterification conditions.
Statistical design of experiments
The effect of citric acid was studied by a RSM, where two variables were analyzed at three different levels: 1) choice of acid (citric/phosphoric); 2) the equivalents of NaOH, necessary to balance the pH (1 to 1.5 to 2 eqs); and 3) extra NaOH, generally helpful for transesterification of difficult oils (0 to 10 to 20 ppm). These conditions were combined with the citric/phosphoric acid possibility. Distribution of the experimental patterns analyzed is shown in Table 2. Each pattern corresponds to a single batch reaction, where the acid treatment, followed by caustic neutralization, was combined directly with the transesterification without any phospholipase addition. Citric acid, at a fixed concentration of 0.065% w/w of oil, or phosphoric acid at 0.025% w/w, were added to 20 g of oil, mixed by high shearing and incubated for 30 min at 55°C, with 250 rpm agitation. After incubation, the oil was cooled down and Callera Trans L (1% w/w) was added together with 3.5% water and NaOH, as indicated in Table 2. Incubation was prolonged for 24 h at 35°C with a linear gradient pumping of methanol (0.4 ml/h for 10 h). Each pattern was analyzed for FAMEs production and final phosphorus content. Data were analyzed with JMP software (SAS Institute Inc.).
Analysis of FAMEs production was performed by gas chromatography (GC). After 24 h incubation, 1 ml reaction mixture was taken and evaporated in a Heto Vacuum concentrator at 60°C for 1½ h to remove excess methanol.
For phosphorus analysis, the whole reaction volume was transferred to a 50 ml tube and centrifuged at low speed (2,000 rpm) for 5 min to simulate the sedimentation step used to separate the final products in an industrial production plant. After centrifugation, 4 ml were taken from the upper oil phase and analyzed through ICP-OES. The bottom phase (glycerin) was analyzed by UPLC/MS/MS to study the phosphatides composition resulting from the reactions.
Determination of FAMEs (%) was performed according to the EN14103 standard method on a Varian Chrompack CP-3900 GC with flame ionization detectors (FIDs), equipped with a Varian ‘Select Biodiesel for FAMEs’ (30 m, 0.32 id) column. Methyl heptadecanoate was used as internal standard, as indicated by EN14103. The solution was prepared at a concentration of 10 mg/ml in acetone. After methanol evaporation, 50 mg of the oil phase were used for each analysis.
Phosphorus content quantification
Phosphorus content was determined by the ICP-OES method at the department of Analytical Development, Novozymes (Kalundborg, Denmark). Accordingly, 0.2 g of each sample were initially destructed in 4.5 ml concentrated HNO3 (69%) and heated for 4 to 5 h at 105°C for further dissolution in a total volume of 10 ml Milli-Q water. Treated samples were analyzed in a Varian Vista MPX system for ICP, using yttrium as internal standard. Resulting data were processed with ICPExpert version 4.1 software, and phosphorus concentration expressed in ppm = mg/kg.
Analysis of phosphatides
Phosphatides content in the oil and glycerin phase was analyzed by UPLC/MS/MS in a Q-Tof Premier (Waters, Milford, MA, USA). Two chromatographic systems were set up: one with a hydrophilic interaction liquid chromatography (HILIC) column (Acquity BEH Amide, 1.7 μm, 2.1 mm × 150 mm), and the second with a reverse phase (RP) column (Acquity CSH C18, 1.7 μm, 2.1 mm × 100 mm). Samples were analyzed in positive and negative mode on the UPLC-UV-Tof and data processed using MassLynx version 4.1 software (Waters). The RP-chromatography was set up with a 0.25 ml/min flow of eluent A containing acetonitrile/ isopropyl alcohol/HCOOH (50:50:0.15) and eluent B containing isopropyl alcohol/HCOOH (100:0.15). The gradient was running isocratic for 1 min at 99% A, followed by a 49 min gradient to 1% A. The 1% A was running isocratic for 5 min, followed by a 5 min gradient back to initial settings (that is, 99% A). The HILIC was set up with a 0.35 ml/min flow and an A-eluent with acetonitrile/HCOOH (100:0.15%) and B-eluent with acetonitrile/MQ water/HCOOH (50:50:0.15). The method was running isocratic for 20 min with 95% A. For both procedures, the MS was set to scan from 95 to 1,500 m/z ions in both positive and negative mode.
UPLC/MS/MS analysis of phosphatide compounds used as standards
Fatty acid methyl ester
Free fatty acid
Flame ionization detector
Hydrophilic interaction liquid chromatography
Inductively coupled plasma optical emission spectrometry
Tandem mass spectrometry
Response surface methodology
Ultra-performance liquid chromatography.
The authors thank Novozymes A/S (Denmark) for kindly providing the enzymes and research facilities. This work was financed by the Scientific and Technological Research Council (MINECO, Spain), grant CTQ2010-21183-C02-02/PPQ, by the IV Pla de Recerca de Catalunya, grant 2009SGR-819, by PCI-AECID project A203563511, and by the Generalitat de Catalunya to the ‘Xarxa de Referència en Biotecnologia’ (XRB). SC acknowledges a doctoral fellowship from the Spanish Ministry of Science and Education (AP2008-04579).
- Dizge N, Aydiner C, Imer DY, Bayramoglu M, Tanriseven A, Keskinlera B: Biodiesel production from sunflower, soybean, and waste cooking oils by transesterification using lipase immobilized onto a novel microporous polymer. Bioresour Technol. 2009, 100 (6): 1983-1991. 10.1016/j.biortech.2008.10.008.View ArticleGoogle Scholar
- Fjerbaek L, Christensen KV, Norddahl B: A review of the current state of biodiesel production using enzymatic transesterification. Biotechnol Bioeng. 2009, 102 (5): 1298-1315. 10.1002/bit.22256.View ArticleGoogle Scholar
- Ghaly AE, Dave D, Brooks MS, Budge S: Production of biodiesel by enzymatic transesterification: review. Am J Biochem Biotechnol. 2010, 6 (2): 54-76. 10.3844/ajbbsp.2010.54.76.View ArticleGoogle Scholar
- Kaieda M, Samukawa T, Matsumoto T, Ban K, Kondo A, Shimada Y, Noda H, Nomoto F, Ohtsuka K, Izumoto E, Fukuda H: Biodiesel fuel production from plant oil catalyzed by Rhizopus oryzae lipase in a water-containing system without an organic solvent. J Biosci Bioeng. 1999, 88 (6): 627-631.View ArticleGoogle Scholar
- Kawakami K, Oda Y, Takahashi R: Application of a Burkholderia cepacia lipase-immobilized silica monolith to batch and continuous biodiesel production with a stoichiometric mixture of methanol and crude Jatropha oil. Biotechnol Biofuels. 2011, 4 (1): 42-10.1186/1754-6834-4-42.View ArticleGoogle Scholar
- Watanabe Y, Shimada Y, Sugihara A, Tominaga Y: Conversion of degummed soybean oil to biodiesel fuel with immobilized Candida antarctica lipase. J Mol Catal B Enzym. 2002, 17 (3–5): 151-155.View ArticleGoogle Scholar
- Robles-Medina A, González-Moreno PA, Esteban-Cerdán L, Molina-Grima E: Biocatalysis: towards ever greener biodiesel production. Biotechnol Adv. 2009, 27 (4): 398-10.1016/j.biotechadv.2008.10.008.View ArticleGoogle Scholar
- Xu Y, Nordblad M, Nielsen PM, Brask J, Woodley JM: In situ visualization and effect of glycerol in lipase-catalyzed ethanolysis of rapeseed oil. J Molec Catal B. 2011, 72 (3–4): 213-219.View ArticleGoogle Scholar
- Chen X, Du W, Liu D, Ding F: Lipase-mediated methanolysis of soybean oils for biodiesel production. J Chem Technol Biotechnol. 2008, 83 (1): 71-76. 10.1002/jctb.1786.View ArticleGoogle Scholar
- Lv D, Du W, Zhang G, Liu D: Mechanism study on NS81006-mediated methanolysis of triglyceride in oil/water biphasic system for biodiesel production. Process Biochem. 2010, 45 (4): 446-450. 10.1016/j.procbio.2009.10.017.View ArticleGoogle Scholar
- Cesarini S, Diaz P, Nielsen PM: Exploring a new, soluble lipase for FAMEs production in water-containing systems using crude soybean oil as a feedstock. Process Biochem. 2013, 48 (3): 484-487. 10.1016/j.procbio.2013.02.001.View ArticleGoogle Scholar
- Tufvesson P, Lima-Ramos J, Nordblad M, Woodley JM: Guidelines and cost analysis for catalyst production in biocatalytic processes. Org Process Res Dev. 2010, 15 (1): 266-274.View ArticleGoogle Scholar
- Nielsen PM, Brask J, Fjerbaek L: Enzymatic biodiesel production: technical and economical considerations. Eur J Lipid Sci Technol. 2008, 110 (8): 692-700. 10.1002/ejlt.200800064.View ArticleGoogle Scholar
- Nielsen PM: Proceedings of the 104th AOCS Annual Meeting and Expo: April 28-May 1. 2013, Montréal, QCGoogle Scholar
- Hama S, Kondo A: Enzymatic biodiesel production: an overview of potential feedstocks and process development. Bioresour Technol. 2013, 135: 386-395.View ArticleGoogle Scholar
- Haas M, McAloon A, Yee W, Foglia T: A process model to estimate biodiesel production costs. Bioresour Technol. 2006, 97 (4): 671-678. 10.1016/j.biortech.2005.03.039.View ArticleGoogle Scholar
- Encinar JM, Sanchez N, Martinez G, Garcia L: Study of biodiesel production from animal fats with high free fatty acid content. Bioresour Technol. 2011, 102 (23): 10907-10914. 10.1016/j.biortech.2011.09.068.View ArticleGoogle Scholar
- Cowan D, Nielsen PM: Enzymatic Degumming Of Edible Oils And Fats. Bleaching and Purifying Fats and Oils: Theory and Practice. Edited by: Patterson HBW. 2009, Urbana, IL: AOCS Press, 216-235.Google Scholar
- Aalrust E, Beyer W, Ottofrickenstein H, Penk G, Plainer H, Reiner R: Enzymatic Treatment of Edible Oils. 1993, US Patent: 5,264,367Google Scholar
- Dijstra A: Proceedings of the World Conference on Oilseed Technology and Utilization. 1993, Champaign, IL: American Oil Chemists’ SocietyGoogle Scholar
- van Nieuwenhuyzen W, Tomás MC: Update on vegetable lecithin and phospholipid technologies. Eur J Lipid Sci Technol. 2008, 110 (5): 472-486. 10.1002/ejlt.200800041.View ArticleGoogle Scholar
- Clausen K: Enzymatic oil-degumming by a novel microbial phospholipase. Eur J Lipid Sci Technol. 2001, 103 (6): 333-340. 10.1002/1438-9312(200106)103:6<333::AID-EJLT333>3.0.CO;2-F.View ArticleGoogle Scholar
- Holm HC, Nielsen PM, Christensen MW: Production of Fatty Acid Alkyl Esters. 2008, US Patent: US 2008/0199924 A1Google Scholar
- Dayton CLG, Galhardo F: Enzymatic Degumming Utilizing a Mixture of PLA and PLC Phospholipases. 2008, US Patent: US 2008/0182322 A1Google Scholar
- Pinisetty D, Moldovan D, Devireddy R: The effect of methanol on lipid bilayers: an atomistic investigation. Ann Biomed Eng. 2006, 34 (9): 1442-1451. 10.1007/s10439-006-9148-y.View ArticleGoogle Scholar
- Daicheng L, Fucui M: Soybean Phospholipids. Recent Trends for Enhancing the Diversity and Quality of Soybean Products, Volume 22. Edited by: Krezhova D. 2011, Rijeka: InTechGoogle Scholar
- Hitchman T: Purifine® PLC: industrial application in oil degumming and refining. Oil Mill Gazetteer. 2009, 115: 2-4.Google Scholar
This article is published under license to BioMed Central Ltd. This is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/2.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly credited. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.