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Kinetic studies and thermodynamics of oil extraction and transesterification of Chlorella sp. for biodiesel production a
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A.L. Ahmad , N.H. Mat Yasin , C.J.C. Derek & J.K. Lim
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School of Chemical Engineering, Engineering Campus, Universiti Sains Malaysia, Seri Ampangan, 14300 Nibong Tebal, Seberang Perai Selatan, Pulau Pinang, Malaysia Published online: 19 Nov 2013.
To cite this article: A.L. Ahmad, N.H. Mat Yasin, C.J.C. Derek & J.K. Lim (2014) Kinetic studies and thermodynamics of oil extraction and transesterification of Chlorella sp. for biodiesel production, Environmental Technology, 35:7, 891-897, DOI: 10.1080/09593330.2013.855263 To link to this article: http://dx.doi.org/10.1080/09593330.2013.855263
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Environmental Technology, 2014 Vol. 35, No. 7, 891–897, http://dx.doi.org/10.1080/09593330.2013.855263
Kinetic studies and thermodynamics of oil extraction and transesterification of Chlorella sp. for biodiesel production A.L. Ahmad∗ , N.H. Mat Yasin, C.J.C. Derek and J.K. Lim School of Chemical Engineering, Engineering Campus, Universiti Sains Malaysia, Seri Ampangan, 14300 Nibong Tebal, Seberang Perai Selatan, Pulau Pinang, Malaysia
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(Received 26 June 2013; final version received 8 October 2013 ) In this work, a mixture of chloroform and methanol (1:1, v/v) was applied to oil extraction from Chlorella sp. at 30, 40, 50 and 60◦ C for 150 min extraction times. Kinetic studies revealed that the values of n and the rate constants were found to depend strongly on temperature. The activation energy was Ea = 38.893 kJ/mol, and the activation thermodynamic parameters at 60◦ C were S = = −180.190 J/mol K, H = = 36.124 kJ/mol and G = = 96.128 kJ/mol. Both H and S yielded positive values, whereas G was negative at 60◦ C, indicating that this process is endothermic, irreversible and spontaneous. The acidic transesterification process was also investigated by gas chromatographic analysis of the microalgae fatty acid methyl esters (biodiesel) at different temperatures and reaction times. The fatty acid profile indicated that the main components were palmitic, linoleic and linolenic acids. The concentration of linolenic acid increased and oleic acid decreased as the temperature increased. Two-hour transesterification is the best reaction time for biodiesel production because it produces the highest percentage of unsaturated fatty acids (74%). These results indicate the potential of Chlorella sp. to produce biodiesel of good quality. Keywords: microalgal biomass; kinetics; thermodynamics; oil extraction; biodiesel
1. Introduction Biodiesel has been receiving considerable attention in recent years due to its potential as a sustainable, biodegradable, nontoxic fuel and environmental-friendly alternative to petrodiesel.[1] Biodiesel derived from oil crops is potentially renewable and a carbon-neutral alternative to petroleum fuels. Unfortunately, biodiesel from oil crops, waste cooking oil and animal fat cannot realistically satisfy even a small fraction of the existing demand for transportation fuels. The relatively high cost of this biodiesel renders the resulting fuels unable to compete with petroleum-derived fuel. Raw materials and ways to reduce the production costs of biodiesel should, therefore, be explored. Microalgae (microscopic unicellular organisms with oxygenic photosynthetic capabilities) appear to be the only source of renewable biodiesel that has the potential to replace the petroleum-derived transportation fuels completely. Microalgae are a specialized group of microorganisms that live in diverse ecological habitats such as freshwater, brackish and marine ecological systems. Similar to plants, microalgae use sunlight to produce oils, but they do so more efficiently than crop plants. Oil productivity from many microalgae greatly exceeds the oil productivity of the best-producing oil crops. Researchers are gaining ∗ Corresponding
author. Email: [email protected]
© 2013 Taylor & Francis
more interest in the cultivation of microalgae because of the valuable metabolites and lipids that can be found in them. The advantages of culturing microalgae as a resource for biomass are listed elsewhere.[2–6] Various methods to extract the oil from algae, such as expeller/press, solvent extraction and supercritical fluid extraction, are available.[7] Extraction using supercritical fluid provides very high purity and good product concentration. However, the operating and investment cost is high.[8] Solvent extraction, a common and efficient technique for producing oil for biodiesel production, involves the transfer of a soluble fraction from a solid material to a liquid solvent. Solvent extraction has a relatively low operating cost compared with supercritical fluid extraction. There are three methods that have been identified for solvent extraction: (i) hot water extraction, (ii) Soxhlet extraction and (iii) ultrasonication technique.[9] However, the solvent extraction has some disadvantages such as poor extraction of polar lipids, long time required for extraction and hazards of boiling solvents.[10] In fact, the lipid extraction step from microalgae still represents a bottleneck of the whole process of producing biodiesel from microalgae. In particular, the main limitation related to the use of organic solvents for extracting lipids from microalgae is their high cost and the need of their continuous supply
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since not all of them can be suitably recycled. For this reason, a rigorous quantification of the extraction kinetics is critical to optimize the extraction process and consequently increase the economical feasibility of the process. Moreover, the organic solvents used to perform extraction are typically toxic and thus the optimization of the amounts used during the extraction step might result in a higher environmental sustainability of the process. The most popular chemical for solvent extraction is hexane, which is relatively inexpensive. Suganya and Renganathan [11] studied the efficiency of algal oil extraction using six different extraction methods with 12 different solvent systems and showed that the extraction of oil using an ultrasound pretreatment method with 1% diethyl ether and 10% methylene chloride in hexane achieved a high yield of oil. Numerous methods are currently available and have been adopted for the production of biodiesel fuel. The four primary ways and processes to produce biodiesel include direct use and blending, microemulsion, thermal cracking (pyrolysis) and transesterification.[12,13] The most common method for converting the oils to biodiesel is transesterification due to its low cost and simplicity. Methanol is usually the preferred alcohol for producing biodiesel because of its low cost and its physical and chemical advantages. In the present investigation, oil extraction from Chlorella sp. with hexane with acidic transesterification for converting the oils to biodiesel was studied. The objectives of this study were to (i) determine the rate constant and activation energy; (ii) determine the enthalpy value and other thermodynamic parameters for oil extraction from Chlorella sp. biomass and (iii) study the effect of temperature and reaction time for acidic transesterification on the fatty acid (biodiesel) yield.
2. Materials and methods 2.1. Preparation of microalgae biomass Chlorella sp. belongs to the family of green algae and is considered part of phytoplankton.[14] Chlorella sp. is the strain most favoured by researchers for biodiesel production and has been described as a good option for biodiesel production.[15] Chlorella sp. was grown in Bold’s Basal Medium and incubated in 2 l Erlenmeyer flasks filled with 2000 ml distilled water for 14 days of cultivation. The medium and flasks were sterilized in an autoclave for 15 min at 121◦ C to prevent any contamination during the early stages of culture growth. The details of the specific measurements and characterization of the Chlorella sp. cultures were reported in our previous research.[15–17] After growth, Chlorella sp. was harvested by centrifugation at 4000 rpm for 10 min using a model Megafuge 40 centrifuge (Thermo Scientific, Germany) and then dried overnight with a freeze dryer (model 7754030, Labconco, US).
2.2. Microalgal cells wall disruption Freeze-dried samples of 20 mg of Chlorella sp. biomass along with 5.5 ml distilled water were placed into capped test tubes and then sonicated (model 8510, Branson, USA) for 5 min. The optical microscope (model BX53F, Olympus, Japan) was used to characterize the Chlorella sp. biomass before and after sonication to identify the changes in the shape of the Chlorella sp. cells. 2.3. Lipid extraction The total lipids were extracted by a mixture of chloroform/methanol (1:1, v/v) using a slightly modified version of Bligh and Dyer’s method.[18] Approximately 12 ml of the mixed solvents was used in each extraction step. The extraction was executed on a water bath preheated to the set temperatures (30, 40, 50 and 60◦ C) on a heating plate using a magnetic stirrer. The total extraction time was 150 min with 30 min time intervals. All experiments were conducted in triplicate. 2.4. Determination of oil extraction yield The amount of oil extracted relative to the time at different temperatures was determined gravimetrically by measuring the weight of the residue after drying the solvent under a flow of nitrogen gas. The oil extraction yield (% w/w) was calculated using the following equation: Oil extraction yield (%) =
Weight of oil extracted (g) × 100 Weight of biomass (g) (1)
2.5. Transesterification In this experiment, temperature and reaction time were the important factors in the transesterification process. The molar ratio of methanol to oil (56:1) and the mixing rate were fixed. An acid catalyst (hydrochloric acid) was used in this research because the alkali catalyst was not suitable for the transesterification of microalgal oil, most likely because of the high acid value of microalgal oil.[19] Zhang et al.[20] also reported that the alkali catalytic process is sensitive to water. The presence of water under alkaline conditions may cause ester saponification. A standard reaction mixture consisting of 4.25 ml methanol, 5 ml hexane and 215 μl HCl (37% vol.) was added to the oil and mixed at 500 rpm. After heating for the specified period, the reaction mixture was allowed to cool and then centrifuged at 3000 rpm for 2 min, resulting in the separation of the two layers. The upper layer consisted of methyl esters (biodiesel), and the lower layer contained the glycerol, the excess methanol and the remaining catalyst. Each experiment was conducted in triplicate. The biodiesel from each experiment was analysed individually in triplicate to calculate the mean values of the experimental data.
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into the inner surfaces of the Chlorella sp. cells. In addition, the Chlorella sp. cells were also dispersed into smaller structures after sonication method as shown in Figure 1.
2.6. Gas chromatographic analysis A fatty acid composition analysis was performed using a gas chromatograph (model 7890A, Agilent, China) equipped with a flame ionization detector, splitless automatic injector and fused silica capillary column (30 m × 0.25 mm × 0.25 μm) (Omegawax 250, Supelco, USA). Injector and detector temperatures were kept constant at 250 and 280◦ C, respectively. The oven temperature program started at 100◦ C for 1 min and then ramped to 190◦ C at 25◦ C/min and remained constant at this temperature for 20 min. Each fatty acid component was identified by comparing the retention time and peak area with the retention time and peak area in the standard solutions, and the composition was calculated as the percentage of the total fatty acids present in the sample. Six fatty acids (C16:0, C16:1, C18:0, C18:1, C18:2 and C18:3) were used as the standard materials. Three replicates of each fatty acid analysis were performed.
3.
3.2. Extraction kinetics A reaction rate equation for oil extraction from Chlorella sp. biomass can be written as [21] dY (2) = kY n dt where Y is the oil extraction yield (%), t is the time of extraction (min), k is the extraction rate constant (min−1 ) and n is the order of the reaction. As the percentage of oil extraction increases over time, the term dY /dt has a positive sign.[21] Using the value in Table 1 and applying the differential method, plots of ln (dY /dt) versus ln Y at different temperatures were found to be linear according to Equation (2). The order of the reaction was determined from the slopes of the straight lines in Figure 2, and the reaction rate constants were calculated from the intercept of the linear plot. A fractional kinetic (with order of about 1.5) was found from the values of n obtained with an average regression coefficient R2 = 0.969. The obtained values of n vary from a minimum of 1.363 to a maximum of 1.638 as shown in Figure 2 and Table 1. From the analysis of the data, the oil extraction yield was found to increase with an increase in extraction time and temperature, showing the influence of these two factors on lipid extraction. The reaction rate constant, k, the time required to obtain maximum extraction of oil from the Chlorella sp. biomass, increases with increasing temperature, and this trend is obvious in Table 1. This observation appears to suggest that the reaction rate constants are often found to depend strongly on temperature. This trend was also found by Suganya and Renganathan with marine macroalgae strain.[11] These authors reported that an increase in the reactivity of the solvent enhances the rate of extraction. However, in their works, the increase in k values is significant at the temperature higher than 45◦ C. As can be noticed, in comparison with other strains such as sunflower seeds [21] and Jatropha Curcas,[22] an increase in k values is consistent even at high temperature. This phenomenon is similar with our findings; there were very slight
Results and discussion
3.1. Sonication pretreatment To increase the cell wall damage in the Chlorella sp. cell, a sonication pretreatment method was implemented on the Chlorella sp. biomass before the oil extraction. The advantages of this sonication method are reduced extraction time, reduced solvent consumption, greater penetration of solvent into cellular materials and improved release of cell contents into the bulk medium.[9] The method also disrupts the cell wall, leading to an increase in the oil extraction yield from the Chlorella sp. biomass due to the penetration of solvent
(a)
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(b)
Figure 1. Optical observation of Chlorella sp. cells (a) before and (b) after sonication (Mag = 20×).
Table 1. The oil extraction yield (%) from Chlorella sp. biomass and the reaction rate constants at different temperatures with respect to extraction time. Oil extraction yield (%) Temp (◦ C) 30 40 50 60
Reaction rate constant, k (min−1 )
n
30 min
60 min
90 min
120 min
150 min
4.7 × 10−4 1.12 × 10−3 1.23 × 10−3 2.1 × 10−3
1.638 1.488 1.504 1.363
8.43 ± 0.14 9.65 ± 0.11 10.48 ± 0.05 10.89 ± 0.07
8.93 ± 0.10 10.73 ± 0.30 12.14 ± 0.20 12.81 ± 0.04
9.49 ± 0.05 12.24 ± 0.14 13.98 ± 0.19 15.64 ± 0.22
10.11 ± 0.012 13.96 ± 0.15 16.37 ± 0.10 19.54 ± 0.18
10.79 ± 0.14 15.96 ± 0.15 19.78 ± 0.10 24.04 ± 0.37
Note: Values are means ± standard deviations of three replicates.
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A.L. Ahmad et al. Table 2. The thermodynamic activation parameters for Chlorella sp. biomass oil extraction at various temperatures. Temp (K)
S = (J/mol K)
H = (kJ/mol)
G = (kJ/mol)
−179.405 −179.675 −179.936 −180.190
36.374 36.291 36.208 36.124
90.733 92.529 94.327 96.128
303 313 323 333
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theory [21]: Figure 2. A plot of ln (dY /dt) versus ln Y at different temperatures for oil extraction from Chlorella sp. biomass. The error bars indicate the standard deviations among the replicates.
RT S = /R e Nh = Ea − RT
(5)
G = = H = − T S =
(6)
A= H =
(4)
where N is Avogadro’s constant, h is Planck’s constant, S = is the activation entropy (J/mol K), H = is the activation enthalpy (kJ/mol) and G = is the activation free energy or Gibb’s energy (kJ/mol). These activation thermodynamic parameters are shown in Table 2 for each temperature. 3.5. Figure 3. Activation energy calculation from the plot of ln k versus 1/T (K−1 ).
difference in the results of k values at the temperature of 40, 50 and 60◦ C as shown in Table 1. 3.3. Calculation of activation energy The change in the rate constant can be described by the Arrhenius equation [21]: k = Ae−Ea/RT
(3)
where k is the reaction rate constant (min−1 ), A is the Arrhenius constant (s−1 ), Ea is the activation energy (kJ/mol), R is the universal gas constant (J/mol K) and T is the absolute temperature (K). A plot of ln k versus 1/T yields a straight line with the slope representing the activation energy of extraction (-Ea/R) and an intercept as the Arrhenius constant, ln A (Figure 3). The activation energy, Ea, and the Arrhenius constant, A, were calculated as 38.893 kJ/mol and 2684 s−1 , respectively. 3.4.
Calculation of thermodynamic activation parameters The thermodynamic activation parameters (S = , H = and G = ) for microalgal oil extraction were calculated using the following equations according to transition state
Calculation of thermodynamic parameters
The thermodynamic parameters (S, H and G) for the extraction of microalgal oil can be estimated using the expressions below [21]: YT Yu G 1 H 1 S ln K = − =− + R T R T R K=
(7) (8)
where K is the equilibrium constant, YT is the percent oil yield (%) at t = 150 min at temperature T , Yu is the percent unextracted oil (%), H is the enthalpy change (kJ/mol), S is the entropy change (1/mol K) and G is the free energy or Gibb’s energy (kJ/mol). At equilibrium, none of the oil yields changes with time. The plot of ln YT versus 1/T at 150 min yields a straight line of ln YT = −HR(1/T ) + C with the slope representing the enthalpy change of extraction, – HR. The enthalpy change was calculated to be H = 0.308 kJ/mol for Chlorella sp. biomass oil extraction (Figure 4). The H value obtained indicated the physicochemical nature of the oil extraction process.[21] The positive value for enthalpy indicates that the oil extraction process is endothermic and requires energy input during the process. Other thermodynamic parameters (S and G) and the equilibrium constant values for Chlorella sp. biomass oil extraction are given in Table 3 for each temperature. An increase in the entropy change and a decrease in the free energy indicate that the process is spontaneously forward. The positive value of entropy change (S > 0) at
Environmental Technology
summarized in Table 4. Based on the previous report,[24] the most common fatty acids contained in biodiesel were palmitic, stearic, oleic and linolenic acid. However, in our study on the Chlorella sp. biomass, palmitic acid (C16:0), linoleic acid (C18:2) and linolenic acid (C18:3) were the major components, followed by stearic acid (C18:0), oleic acid (C18:1) and small amounts of palmitoleic acid (C16:1) was also identified. To study the effect of temperatures, experiments were carried out at temperatures of 70, 80 and 90◦ C because the transesterification is not affected at low temperature. This is in agreement with the results presented in the previous research by Rashid and Anwar [25]; an increase in the reaction temperature leads to a higher yield of fatty acid. In spite of this, good quality of biodiesel could be obtained at a higher temperature. However, at very high temperature (above 90◦ C) it could burn some of the oil before completion of the alcoholysis. Thus, only three temperatures were chosen in this experiment. Unlike the two major fatty acids, special attention should be taken to the content of linolenic acid (C18:3) because the European regulation EN 14103 has set 12% as the maximum allowed content of C18:3 for quality biodiesel. Furthermore, fatty acid with a high oleic acid (C18:1) has been reported to have a reasonable balance of fuel properties due to increases of oxidative stability for longer storage.[26,27] As shown in Table 4 linolenic acid content increased as reaction temperature increased, while opposite trend was observed in oleic acid content. These findings differ from the reported lipid content of Nannochloropsis oculata and Chlorella vulgaris, where the increase in temperature led to an increased content of oleic acid (C18:1).[14] Therefore, these two fatty acids (C18:1 and C18:3) might be useful as a good indicator for various temperatures. The influence of reaction time was also analysed by varying the reaction time up to 5 hours while keeping the optimized temperature at 90◦ C. The trends observed when varying the reaction time from 1 to 5 hours were generally inconsistent for most fatty acids. As noted by Knothe [27] and Griffiths and Harrison,[28] the carbon chain length and the degree of unsaturation are important characteristics
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Figure 4. Enthalpy change calculation from the plot of ln YT versus 1/T (K−1 ). Table 3. The equilibrium constant (K) and the thermodynamic parameters (S and G) for Chlorella sp. biomass oil extraction at different temperatures. Temp (K) 303 313 323 333
K 0.372 0.669 0.987 1.522
G (kJ/mol) 2.494 1.048 0.036 −1.164
S (1/mol K) −7.214 −2.362 0.841 4.420
T ≥ 50◦ C indicates that the process is irreversible, whereas the value of S is negative for the T < 40◦ C. Thus, the extraction process at the lower temperature is not suitable for oil extraction of Chlorella sp. because the equilibrium constant is very low at the temperature lower than 40◦ C as shown in Table 3. The different types of results were previously reported by Amin et al.[22] and Liauw et al.[23] for the extraction of oil from Jatropha Curcas and Azadirachta indica A. Juss. In their works, S is positive and G is negative for all conditions. The equilibrium constant is also high (K > 1) even at low temperature. 3.6. Fatty acid compositions The fatty acid composition of the Chlorella sp. was tested at various temperatures and reaction times, and the results are Table 4.
Summary of fatty acid compositions in Chlorella sp. at various temperatures and reaction times (unit % of total fatty acid). Temperature (◦ C)
Fatty acids C16:0 C16:1 C18:0 C18:1 C18:2 C18:3 SFA (%) UFA (%)
895
Reaction time (h)
70
80
90
1
2
3
4
5
29.07 ± 0.51 0.83 ± 0.04 9.48 ± 0.57 2.66 ± 0.22 38.75 ± 0.96 19.22 ± 0.64 38.55 ± 0.29 61.45 ± 0.59
30.08 ± 1.89 0.89 ± 0.09 12.77 ± 0.17 2.55 ± 0.14 33.49 ± 0.74 20.22 ± 3.78 42.85 ± 1.12 57.15 ± 1.43
26.01 ± 0.12 2.62 ± 0.08 3.01 ± 0.28 2.44 ± 0.05 39.45 ± 1.46 26.49 ± 1.19 29.01 ± 0.11 70.99 ± 1.31
28.41 ± 4.11 0.55 ± 0.33 1.82 ± 0.19 3.04 ± 0.02 42.53 ± 0.58 23.65 ± 2.61 30.22 ± 2.53 69.78 ± 0.26
24.25 ± 1.56 0.75 ± 0.28 1.50 ± 0.00 4.73 ± 1.17 45.59 ± 3.58 23.17 ± 0.28 25.75 ± 1.35 74.25 ± 3.1
37.56 ± 2.90 1.09 ± 0.17 1.58 ± 0.44 2.87 ± 0.36 36.83 ± 4.30 20.07 ± 0.07 39.14 ± 1.28 60.86 ± 1.51
25.15 ± 1.37 1.41 ± 0.34 1.78 ± 0.45 4.12 ± 1.10 42.26 ± 2.41 25.28 ± 1.76 26.93 ± 1.31 73.07 ± 1.28
28.41 ± 3.58 1.73 ± 0.57 2.21 ± 0.23 4.14 ± 0.85 41.01 ± 4.57 22.51 ± 1.76 30.62 ± 1.58 69.38 ± 2.35
SFA, saturated fatty acids; UFA, unsaturated fatty acids. Note: Data are expressed as mean ± standard deviations of three replicates.
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of fatty acids for biodiesel production and may affect the quality of the biodiesel produced. Table 4 shows that the Chlorella sp. lipids for all reaction times are composed mainly of unsaturated fatty acids (60–75%), with a significant percentage of palmitic acid also present (25–38%). Among the five reaction times applied in our study, the shortest reaction time (2 h) shows the highest percentage of UFAs, and special attention should be paid to the linoleic acid because the C18:2 unsaturated chains were the predominant fatty acids (45.6%) followed by C18:3 (25.2%) and C16:0 (24.3%). It is also important to highlight that higher compositions of UFAs can reduce the pour point of biodiesel and making it feasible to be used in cold climate countries.[29] However, when the UFA with double bond ≥ 3, their high content is undesirable since it reduces the chemical and oxidative stability of the resulting biodiesel.[27] Therefore, the higher content of unsaturated chains (such as C18:1 and C18:2) and the lower content of C18:3 in Chlorella sp. at 2 h of reaction time may be expected to produce a good quality of biodiesel. Further study will be necessary to uncover the feasibility and effectiveness of various harvesting methods for the production of high-quality biodiesel from Chlorella sp., then the physical and fuel properties may be comparable to diesel fuel and the ASTM biodiesel standard.
Conclusion The kinetics for oil extraction from Chlorella sp. biomass by a mixture of chloroform/methanol (1:1, v/v) were found to be a fractional kinetic with the order of about 1.5 by the differential method and strongly dependent on temperature. The activation energy was Ea = 38.893 kJ/mol and the thermodynamic activation parameters at 60◦ C were S = = −180.190 J/mol K, H = = 36.124 kJ/mol and G = = 96.128 kJ /mol. The enthalpy value was found to be positive (H = 0.308 kJ/mol), indicating that the process is endothermic and requires the input of energy during the process. The other thermodynamic parameters at 60◦ C were calculated to be S = 4.420 1/mol K and G = −1.164 kJ/mol. Our study also demonstrated that palmitic acid (C16:0), linoleic acid (C18:2) and linolenic acid (C18:3) are major fatty acids found in Chlorella sp., indicating a good potential for use of the Chlorella sp. lipids in the production of biodiesel. The variation of temperature influenced the oleic acid (C18:1) and linolenic acid (C18:3) contents strongly, as an increase from 70 to 90◦ C increased the content of linolenic acid from 19.22 to 26.49% and decreased the oleic acid from 2.66 to 2.44%. Finally, 2 h of reaction time yields the most appropriate fatty acid composition in terms of unsaturated fatty acids such as C18:1 and C18:2. The next research effort will address the harvesting of Chlorella sp. cells using appropriate and efficient methods, making Chlorella sp. biomass a good candidate for biodiesel production.
Acknowledgements All authors are affiliated with the Membrane Science and Technology Cluster USM.
Funding This material is based on the work supported by a Research University (RU) [Grant No. 1001/PJKIMIA/814060) and a Postgraduate Research (PRGS) [Grant No. 1001/JKIMIA/8044014] from Universiti Sains Malaysia (USM). N.H. Mat Yasin gratefully acknowledges Universiti Malaysia Pahang [SLAB 2011] for a scholarship.
References [1] Vicente G, Martinez M, Aracil J. Integrated biodiesel production: a comparison of different homogeneous catalysts systems. Bioresour Technol. 2008;99:9009–9012. [2] Ahmad AL, Mat Yasin NH, Derek CJC, Lim JK. Microalgae as a sustainable energy source for biodiesel production: a review. Renew Sust Energ Rev. 2011;15:584–593. [3] Mata TM, Martins AA, Caetano NS. Microalgae for biodiesel production and other applications: a review. Renew Sust Energ Rev. 2010;14:217–232. [4] Khan AK, Rashmi HMZ, Prasad S, Banerjee UC. Prospects of biodiesel production from microalgae in India. Renew Sust Energ Rev. 2009;13:2361–2372. [5] Janaun J, Ellis N. Perspectives on biodiesel as a sustainable fuel. Renew Sust Energ Rev. 2009;14:1312–1320. [6] Brennan L, Owende P. Biofuels from microalgae – a review of technologies for production, processing and extractions of biofuels and co-products. Renew Sust Energ Rev. 2010;14:557–577. [7] Demirbas A. Production of biodiesel from algae oils. Energ Sources A. 2009;31:163–168. [8] Mongkholkhajornsilp D, Donglas PL, Elkamel A, Tepparitoon W, Pongamphair S. Supercritical CO2 extraction of nimbim from neem seeds – a modeling study. J Food Eng. 2004;71:331–340. [9] Achten WMJ, Verchit L, Mathijs Franken YJ, Singh E, Aerts VP, Muys RB. Jatropha bio-diesel production and use. Biomass Bioenerg. 2008;32(12):1063–1084. [10] Kirrolia A, Bishnoi NR, Singh R. Microalgae as a boon for sustainable energy production and its future research & development aspects. Renew Sust Energ Rev. 2013;20: 642–656. [11] Suganya T, Renganathan S. Optimization and kinetic studies on algal oil extraction from marine macroalgae Ulva lactuca. Bioresour Technol. 2012;107:319–326. [12] Ma F, Hanna MA. Biodiesel production: a review. Bioresour Technol. 1999;70:1–15. [13] Singh SP, Singh D. Biodiesel production through the use of different sources and characterization of oils and their esters as the substitute of diesel: a review. Renew Sust Energ Rev. 2010;14:200–216. [14] Converti A, Casazza AA, Ortiz EY, Perego P, Borghi MD. Effect of temperature and nitrogen concentration on the growth and lipid content of Nannochloropsis oculata and Chlorella vulgaris for biodiesel production. Chem Eng Process Process Intensif. 2009;48:1146–1151. [15] Ahmad AL, Mat Yasin NH, Derek CJC, Lim JK. Crossflow microfiltration of microalgae biomass for biofuel production. Desalination. 2012;302:65–70. [16] Ahmad AL, Mat Yasin NH, Derek CJC, Lim JK. Optimization of microalgae coagulation process using chitosan. Chem Eng J. 2011;173(3):879–882.
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Environmental Technology [17] Ahmad AL, Mat Yasin NH, Derek CJC, Lim JK. Microfiltration of Chlorella sp.: influence of material and membrane pore size. Membrane Water Treat. 2013;4(2):143–155. [18] Bligh EG, Dyer WJ. A rapid method of lipid extraction and purification. Can J Biochem Physiol. 1959;37:911–917. [19] Miao X, Wu Q. Biodiesel production from heterotrophic microalgal oil. Bioresour Technol. 2006;97:841–846. [20] Zhang Y, Dube MA, Mclean DD, Kates M. Biodiesel production from waste cooking oil: 2. Economic assessment and sensitivity analysis. Bioresour Technol. 2003;90:229–40. [21] Topallar H, Gecgel U. Kinetics and thermodynamics of oil extraction from sunflower seeds in the presence of aqueous acidic hexane solutions. Turk J Chem. 2000;24:247–253. [22] Amin AK, Hawash S, El Diwani G, El Rafei S. Kinetics and thermodynamics of oil extraction from Jatropha Curcas in aqueous acidic hexane solutions. J Am Sci. 2010;6(11): 293–300. [23] Liauw MA, Natan FA, Widiyanti P, Ikasari D, Indraswati N, Soetaredjo FE. Extraction of neem oil (Azadirachta indica A. Juss) using n-hexane and ethanol: studies of oil
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quality, kinetic and thermodynamic. ARPN J Eng Appl Sci. 2008;3(3):49–54. Knothe G. “Designer” biodiesel: optimizing fatty ester composition to improve fuel properties. Energ Fuel. 2008;22(2):1358–1364. Rashid U, Anwar F. Production of biodiesel through optimized alkaline-catalyzed transesterification of rapeseed oil. Fuel. 2008;87:265–273. Rashid U, Anwar F, Moser BR, Knothe G. Moringa oleifera oil: a possible source of biodiesel. Bioresour Technol. 2008;99:8175–8179. Knothe G. Dependence of biodiesel fuel properties on the structure of fatty acid alkyl esters. Fuel Process Technol. 2005;86:1059–1070. Griffiths MJ, Harrison STL. Lipid productivity as a key characteristic for choosing algal species for biodiesel production. J Appl Phycol. 2009;21:493–507. Abdelmalik AA, Abbott AP, Fothergill JC, Dodd S, Harris RC. Synthesis of a base-stock for electrical insulating fluid based on palm kernel oil. Ind Crop Prod. 2011;33:532–536.
Environmental Technology Publication details, including instructions for authors and subscription information: http://www.tandfonline.com/loi/tent20
Kinetic studies and thermodynamics of oil extraction and transesterification of Chlorella sp. for biodiesel production a
a
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A.L. Ahmad , N.H. Mat Yasin , C.J.C. Derek & J.K. Lim
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School of Chemical Engineering, Engineering Campus, Universiti Sains Malaysia, Seri Ampangan, 14300 Nibong Tebal, Seberang Perai Selatan, Pulau Pinang, Malaysia Published online: 19 Nov 2013.
To cite this article: A.L. Ahmad, N.H. Mat Yasin, C.J.C. Derek & J.K. Lim (2014) Kinetic studies and thermodynamics of oil extraction and transesterification of Chlorella sp. for biodiesel production, Environmental Technology, 35:7, 891-897, DOI: 10.1080/09593330.2013.855263 To link to this article: http://dx.doi.org/10.1080/09593330.2013.855263
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Environmental Technology, 2014 Vol. 35, No. 7, 891–897, http://dx.doi.org/10.1080/09593330.2013.855263
Kinetic studies and thermodynamics of oil extraction and transesterification of Chlorella sp. for biodiesel production A.L. Ahmad∗ , N.H. Mat Yasin, C.J.C. Derek and J.K. Lim School of Chemical Engineering, Engineering Campus, Universiti Sains Malaysia, Seri Ampangan, 14300 Nibong Tebal, Seberang Perai Selatan, Pulau Pinang, Malaysia
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(Received 26 June 2013; final version received 8 October 2013 ) In this work, a mixture of chloroform and methanol (1:1, v/v) was applied to oil extraction from Chlorella sp. at 30, 40, 50 and 60◦ C for 150 min extraction times. Kinetic studies revealed that the values of n and the rate constants were found to depend strongly on temperature. The activation energy was Ea = 38.893 kJ/mol, and the activation thermodynamic parameters at 60◦ C were S = = −180.190 J/mol K, H = = 36.124 kJ/mol and G = = 96.128 kJ/mol. Both H and S yielded positive values, whereas G was negative at 60◦ C, indicating that this process is endothermic, irreversible and spontaneous. The acidic transesterification process was also investigated by gas chromatographic analysis of the microalgae fatty acid methyl esters (biodiesel) at different temperatures and reaction times. The fatty acid profile indicated that the main components were palmitic, linoleic and linolenic acids. The concentration of linolenic acid increased and oleic acid decreased as the temperature increased. Two-hour transesterification is the best reaction time for biodiesel production because it produces the highest percentage of unsaturated fatty acids (74%). These results indicate the potential of Chlorella sp. to produce biodiesel of good quality. Keywords: microalgal biomass; kinetics; thermodynamics; oil extraction; biodiesel
1. Introduction Biodiesel has been receiving considerable attention in recent years due to its potential as a sustainable, biodegradable, nontoxic fuel and environmental-friendly alternative to petrodiesel.[1] Biodiesel derived from oil crops is potentially renewable and a carbon-neutral alternative to petroleum fuels. Unfortunately, biodiesel from oil crops, waste cooking oil and animal fat cannot realistically satisfy even a small fraction of the existing demand for transportation fuels. The relatively high cost of this biodiesel renders the resulting fuels unable to compete with petroleum-derived fuel. Raw materials and ways to reduce the production costs of biodiesel should, therefore, be explored. Microalgae (microscopic unicellular organisms with oxygenic photosynthetic capabilities) appear to be the only source of renewable biodiesel that has the potential to replace the petroleum-derived transportation fuels completely. Microalgae are a specialized group of microorganisms that live in diverse ecological habitats such as freshwater, brackish and marine ecological systems. Similar to plants, microalgae use sunlight to produce oils, but they do so more efficiently than crop plants. Oil productivity from many microalgae greatly exceeds the oil productivity of the best-producing oil crops. Researchers are gaining ∗ Corresponding
author. Email: [email protected]
© 2013 Taylor & Francis
more interest in the cultivation of microalgae because of the valuable metabolites and lipids that can be found in them. The advantages of culturing microalgae as a resource for biomass are listed elsewhere.[2–6] Various methods to extract the oil from algae, such as expeller/press, solvent extraction and supercritical fluid extraction, are available.[7] Extraction using supercritical fluid provides very high purity and good product concentration. However, the operating and investment cost is high.[8] Solvent extraction, a common and efficient technique for producing oil for biodiesel production, involves the transfer of a soluble fraction from a solid material to a liquid solvent. Solvent extraction has a relatively low operating cost compared with supercritical fluid extraction. There are three methods that have been identified for solvent extraction: (i) hot water extraction, (ii) Soxhlet extraction and (iii) ultrasonication technique.[9] However, the solvent extraction has some disadvantages such as poor extraction of polar lipids, long time required for extraction and hazards of boiling solvents.[10] In fact, the lipid extraction step from microalgae still represents a bottleneck of the whole process of producing biodiesel from microalgae. In particular, the main limitation related to the use of organic solvents for extracting lipids from microalgae is their high cost and the need of their continuous supply
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since not all of them can be suitably recycled. For this reason, a rigorous quantification of the extraction kinetics is critical to optimize the extraction process and consequently increase the economical feasibility of the process. Moreover, the organic solvents used to perform extraction are typically toxic and thus the optimization of the amounts used during the extraction step might result in a higher environmental sustainability of the process. The most popular chemical for solvent extraction is hexane, which is relatively inexpensive. Suganya and Renganathan [11] studied the efficiency of algal oil extraction using six different extraction methods with 12 different solvent systems and showed that the extraction of oil using an ultrasound pretreatment method with 1% diethyl ether and 10% methylene chloride in hexane achieved a high yield of oil. Numerous methods are currently available and have been adopted for the production of biodiesel fuel. The four primary ways and processes to produce biodiesel include direct use and blending, microemulsion, thermal cracking (pyrolysis) and transesterification.[12,13] The most common method for converting the oils to biodiesel is transesterification due to its low cost and simplicity. Methanol is usually the preferred alcohol for producing biodiesel because of its low cost and its physical and chemical advantages. In the present investigation, oil extraction from Chlorella sp. with hexane with acidic transesterification for converting the oils to biodiesel was studied. The objectives of this study were to (i) determine the rate constant and activation energy; (ii) determine the enthalpy value and other thermodynamic parameters for oil extraction from Chlorella sp. biomass and (iii) study the effect of temperature and reaction time for acidic transesterification on the fatty acid (biodiesel) yield.
2. Materials and methods 2.1. Preparation of microalgae biomass Chlorella sp. belongs to the family of green algae and is considered part of phytoplankton.[14] Chlorella sp. is the strain most favoured by researchers for biodiesel production and has been described as a good option for biodiesel production.[15] Chlorella sp. was grown in Bold’s Basal Medium and incubated in 2 l Erlenmeyer flasks filled with 2000 ml distilled water for 14 days of cultivation. The medium and flasks were sterilized in an autoclave for 15 min at 121◦ C to prevent any contamination during the early stages of culture growth. The details of the specific measurements and characterization of the Chlorella sp. cultures were reported in our previous research.[15–17] After growth, Chlorella sp. was harvested by centrifugation at 4000 rpm for 10 min using a model Megafuge 40 centrifuge (Thermo Scientific, Germany) and then dried overnight with a freeze dryer (model 7754030, Labconco, US).
2.2. Microalgal cells wall disruption Freeze-dried samples of 20 mg of Chlorella sp. biomass along with 5.5 ml distilled water were placed into capped test tubes and then sonicated (model 8510, Branson, USA) for 5 min. The optical microscope (model BX53F, Olympus, Japan) was used to characterize the Chlorella sp. biomass before and after sonication to identify the changes in the shape of the Chlorella sp. cells. 2.3. Lipid extraction The total lipids were extracted by a mixture of chloroform/methanol (1:1, v/v) using a slightly modified version of Bligh and Dyer’s method.[18] Approximately 12 ml of the mixed solvents was used in each extraction step. The extraction was executed on a water bath preheated to the set temperatures (30, 40, 50 and 60◦ C) on a heating plate using a magnetic stirrer. The total extraction time was 150 min with 30 min time intervals. All experiments were conducted in triplicate. 2.4. Determination of oil extraction yield The amount of oil extracted relative to the time at different temperatures was determined gravimetrically by measuring the weight of the residue after drying the solvent under a flow of nitrogen gas. The oil extraction yield (% w/w) was calculated using the following equation: Oil extraction yield (%) =
Weight of oil extracted (g) × 100 Weight of biomass (g) (1)
2.5. Transesterification In this experiment, temperature and reaction time were the important factors in the transesterification process. The molar ratio of methanol to oil (56:1) and the mixing rate were fixed. An acid catalyst (hydrochloric acid) was used in this research because the alkali catalyst was not suitable for the transesterification of microalgal oil, most likely because of the high acid value of microalgal oil.[19] Zhang et al.[20] also reported that the alkali catalytic process is sensitive to water. The presence of water under alkaline conditions may cause ester saponification. A standard reaction mixture consisting of 4.25 ml methanol, 5 ml hexane and 215 μl HCl (37% vol.) was added to the oil and mixed at 500 rpm. After heating for the specified period, the reaction mixture was allowed to cool and then centrifuged at 3000 rpm for 2 min, resulting in the separation of the two layers. The upper layer consisted of methyl esters (biodiesel), and the lower layer contained the glycerol, the excess methanol and the remaining catalyst. Each experiment was conducted in triplicate. The biodiesel from each experiment was analysed individually in triplicate to calculate the mean values of the experimental data.
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Environmental Technology
into the inner surfaces of the Chlorella sp. cells. In addition, the Chlorella sp. cells were also dispersed into smaller structures after sonication method as shown in Figure 1.
2.6. Gas chromatographic analysis A fatty acid composition analysis was performed using a gas chromatograph (model 7890A, Agilent, China) equipped with a flame ionization detector, splitless automatic injector and fused silica capillary column (30 m × 0.25 mm × 0.25 μm) (Omegawax 250, Supelco, USA). Injector and detector temperatures were kept constant at 250 and 280◦ C, respectively. The oven temperature program started at 100◦ C for 1 min and then ramped to 190◦ C at 25◦ C/min and remained constant at this temperature for 20 min. Each fatty acid component was identified by comparing the retention time and peak area with the retention time and peak area in the standard solutions, and the composition was calculated as the percentage of the total fatty acids present in the sample. Six fatty acids (C16:0, C16:1, C18:0, C18:1, C18:2 and C18:3) were used as the standard materials. Three replicates of each fatty acid analysis were performed.
3.
3.2. Extraction kinetics A reaction rate equation for oil extraction from Chlorella sp. biomass can be written as [21] dY (2) = kY n dt where Y is the oil extraction yield (%), t is the time of extraction (min), k is the extraction rate constant (min−1 ) and n is the order of the reaction. As the percentage of oil extraction increases over time, the term dY /dt has a positive sign.[21] Using the value in Table 1 and applying the differential method, plots of ln (dY /dt) versus ln Y at different temperatures were found to be linear according to Equation (2). The order of the reaction was determined from the slopes of the straight lines in Figure 2, and the reaction rate constants were calculated from the intercept of the linear plot. A fractional kinetic (with order of about 1.5) was found from the values of n obtained with an average regression coefficient R2 = 0.969. The obtained values of n vary from a minimum of 1.363 to a maximum of 1.638 as shown in Figure 2 and Table 1. From the analysis of the data, the oil extraction yield was found to increase with an increase in extraction time and temperature, showing the influence of these two factors on lipid extraction. The reaction rate constant, k, the time required to obtain maximum extraction of oil from the Chlorella sp. biomass, increases with increasing temperature, and this trend is obvious in Table 1. This observation appears to suggest that the reaction rate constants are often found to depend strongly on temperature. This trend was also found by Suganya and Renganathan with marine macroalgae strain.[11] These authors reported that an increase in the reactivity of the solvent enhances the rate of extraction. However, in their works, the increase in k values is significant at the temperature higher than 45◦ C. As can be noticed, in comparison with other strains such as sunflower seeds [21] and Jatropha Curcas,[22] an increase in k values is consistent even at high temperature. This phenomenon is similar with our findings; there were very slight
Results and discussion
3.1. Sonication pretreatment To increase the cell wall damage in the Chlorella sp. cell, a sonication pretreatment method was implemented on the Chlorella sp. biomass before the oil extraction. The advantages of this sonication method are reduced extraction time, reduced solvent consumption, greater penetration of solvent into cellular materials and improved release of cell contents into the bulk medium.[9] The method also disrupts the cell wall, leading to an increase in the oil extraction yield from the Chlorella sp. biomass due to the penetration of solvent
(a)
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(b)
Figure 1. Optical observation of Chlorella sp. cells (a) before and (b) after sonication (Mag = 20×).
Table 1. The oil extraction yield (%) from Chlorella sp. biomass and the reaction rate constants at different temperatures with respect to extraction time. Oil extraction yield (%) Temp (◦ C) 30 40 50 60
Reaction rate constant, k (min−1 )
n
30 min
60 min
90 min
120 min
150 min
4.7 × 10−4 1.12 × 10−3 1.23 × 10−3 2.1 × 10−3
1.638 1.488 1.504 1.363
8.43 ± 0.14 9.65 ± 0.11 10.48 ± 0.05 10.89 ± 0.07
8.93 ± 0.10 10.73 ± 0.30 12.14 ± 0.20 12.81 ± 0.04
9.49 ± 0.05 12.24 ± 0.14 13.98 ± 0.19 15.64 ± 0.22
10.11 ± 0.012 13.96 ± 0.15 16.37 ± 0.10 19.54 ± 0.18
10.79 ± 0.14 15.96 ± 0.15 19.78 ± 0.10 24.04 ± 0.37
Note: Values are means ± standard deviations of three replicates.
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S = (J/mol K)
H = (kJ/mol)
G = (kJ/mol)
−179.405 −179.675 −179.936 −180.190
36.374 36.291 36.208 36.124
90.733 92.529 94.327 96.128
303 313 323 333
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theory [21]: Figure 2. A plot of ln (dY /dt) versus ln Y at different temperatures for oil extraction from Chlorella sp. biomass. The error bars indicate the standard deviations among the replicates.
RT S = /R e Nh = Ea − RT
(5)
G = = H = − T S =
(6)
A= H =
(4)
where N is Avogadro’s constant, h is Planck’s constant, S = is the activation entropy (J/mol K), H = is the activation enthalpy (kJ/mol) and G = is the activation free energy or Gibb’s energy (kJ/mol). These activation thermodynamic parameters are shown in Table 2 for each temperature. 3.5. Figure 3. Activation energy calculation from the plot of ln k versus 1/T (K−1 ).
difference in the results of k values at the temperature of 40, 50 and 60◦ C as shown in Table 1. 3.3. Calculation of activation energy The change in the rate constant can be described by the Arrhenius equation [21]: k = Ae−Ea/RT
(3)
where k is the reaction rate constant (min−1 ), A is the Arrhenius constant (s−1 ), Ea is the activation energy (kJ/mol), R is the universal gas constant (J/mol K) and T is the absolute temperature (K). A plot of ln k versus 1/T yields a straight line with the slope representing the activation energy of extraction (-Ea/R) and an intercept as the Arrhenius constant, ln A (Figure 3). The activation energy, Ea, and the Arrhenius constant, A, were calculated as 38.893 kJ/mol and 2684 s−1 , respectively. 3.4.
Calculation of thermodynamic activation parameters The thermodynamic activation parameters (S = , H = and G = ) for microalgal oil extraction were calculated using the following equations according to transition state
Calculation of thermodynamic parameters
The thermodynamic parameters (S, H and G) for the extraction of microalgal oil can be estimated using the expressions below [21]: YT Yu G 1 H 1 S ln K = − =− + R T R T R K=
(7) (8)
where K is the equilibrium constant, YT is the percent oil yield (%) at t = 150 min at temperature T , Yu is the percent unextracted oil (%), H is the enthalpy change (kJ/mol), S is the entropy change (1/mol K) and G is the free energy or Gibb’s energy (kJ/mol). At equilibrium, none of the oil yields changes with time. The plot of ln YT versus 1/T at 150 min yields a straight line of ln YT = −HR(1/T ) + C with the slope representing the enthalpy change of extraction, – HR. The enthalpy change was calculated to be H = 0.308 kJ/mol for Chlorella sp. biomass oil extraction (Figure 4). The H value obtained indicated the physicochemical nature of the oil extraction process.[21] The positive value for enthalpy indicates that the oil extraction process is endothermic and requires energy input during the process. Other thermodynamic parameters (S and G) and the equilibrium constant values for Chlorella sp. biomass oil extraction are given in Table 3 for each temperature. An increase in the entropy change and a decrease in the free energy indicate that the process is spontaneously forward. The positive value of entropy change (S > 0) at
Environmental Technology
summarized in Table 4. Based on the previous report,[24] the most common fatty acids contained in biodiesel were palmitic, stearic, oleic and linolenic acid. However, in our study on the Chlorella sp. biomass, palmitic acid (C16:0), linoleic acid (C18:2) and linolenic acid (C18:3) were the major components, followed by stearic acid (C18:0), oleic acid (C18:1) and small amounts of palmitoleic acid (C16:1) was also identified. To study the effect of temperatures, experiments were carried out at temperatures of 70, 80 and 90◦ C because the transesterification is not affected at low temperature. This is in agreement with the results presented in the previous research by Rashid and Anwar [25]; an increase in the reaction temperature leads to a higher yield of fatty acid. In spite of this, good quality of biodiesel could be obtained at a higher temperature. However, at very high temperature (above 90◦ C) it could burn some of the oil before completion of the alcoholysis. Thus, only three temperatures were chosen in this experiment. Unlike the two major fatty acids, special attention should be taken to the content of linolenic acid (C18:3) because the European regulation EN 14103 has set 12% as the maximum allowed content of C18:3 for quality biodiesel. Furthermore, fatty acid with a high oleic acid (C18:1) has been reported to have a reasonable balance of fuel properties due to increases of oxidative stability for longer storage.[26,27] As shown in Table 4 linolenic acid content increased as reaction temperature increased, while opposite trend was observed in oleic acid content. These findings differ from the reported lipid content of Nannochloropsis oculata and Chlorella vulgaris, where the increase in temperature led to an increased content of oleic acid (C18:1).[14] Therefore, these two fatty acids (C18:1 and C18:3) might be useful as a good indicator for various temperatures. The influence of reaction time was also analysed by varying the reaction time up to 5 hours while keeping the optimized temperature at 90◦ C. The trends observed when varying the reaction time from 1 to 5 hours were generally inconsistent for most fatty acids. As noted by Knothe [27] and Griffiths and Harrison,[28] the carbon chain length and the degree of unsaturation are important characteristics
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Figure 4. Enthalpy change calculation from the plot of ln YT versus 1/T (K−1 ). Table 3. The equilibrium constant (K) and the thermodynamic parameters (S and G) for Chlorella sp. biomass oil extraction at different temperatures. Temp (K) 303 313 323 333
K 0.372 0.669 0.987 1.522
G (kJ/mol) 2.494 1.048 0.036 −1.164
S (1/mol K) −7.214 −2.362 0.841 4.420
T ≥ 50◦ C indicates that the process is irreversible, whereas the value of S is negative for the T < 40◦ C. Thus, the extraction process at the lower temperature is not suitable for oil extraction of Chlorella sp. because the equilibrium constant is very low at the temperature lower than 40◦ C as shown in Table 3. The different types of results were previously reported by Amin et al.[22] and Liauw et al.[23] for the extraction of oil from Jatropha Curcas and Azadirachta indica A. Juss. In their works, S is positive and G is negative for all conditions. The equilibrium constant is also high (K > 1) even at low temperature. 3.6. Fatty acid compositions The fatty acid composition of the Chlorella sp. was tested at various temperatures and reaction times, and the results are Table 4.
Summary of fatty acid compositions in Chlorella sp. at various temperatures and reaction times (unit % of total fatty acid). Temperature (◦ C)
Fatty acids C16:0 C16:1 C18:0 C18:1 C18:2 C18:3 SFA (%) UFA (%)
895
Reaction time (h)
70
80
90
1
2
3
4
5
29.07 ± 0.51 0.83 ± 0.04 9.48 ± 0.57 2.66 ± 0.22 38.75 ± 0.96 19.22 ± 0.64 38.55 ± 0.29 61.45 ± 0.59
30.08 ± 1.89 0.89 ± 0.09 12.77 ± 0.17 2.55 ± 0.14 33.49 ± 0.74 20.22 ± 3.78 42.85 ± 1.12 57.15 ± 1.43
26.01 ± 0.12 2.62 ± 0.08 3.01 ± 0.28 2.44 ± 0.05 39.45 ± 1.46 26.49 ± 1.19 29.01 ± 0.11 70.99 ± 1.31
28.41 ± 4.11 0.55 ± 0.33 1.82 ± 0.19 3.04 ± 0.02 42.53 ± 0.58 23.65 ± 2.61 30.22 ± 2.53 69.78 ± 0.26
24.25 ± 1.56 0.75 ± 0.28 1.50 ± 0.00 4.73 ± 1.17 45.59 ± 3.58 23.17 ± 0.28 25.75 ± 1.35 74.25 ± 3.1
37.56 ± 2.90 1.09 ± 0.17 1.58 ± 0.44 2.87 ± 0.36 36.83 ± 4.30 20.07 ± 0.07 39.14 ± 1.28 60.86 ± 1.51
25.15 ± 1.37 1.41 ± 0.34 1.78 ± 0.45 4.12 ± 1.10 42.26 ± 2.41 25.28 ± 1.76 26.93 ± 1.31 73.07 ± 1.28
28.41 ± 3.58 1.73 ± 0.57 2.21 ± 0.23 4.14 ± 0.85 41.01 ± 4.57 22.51 ± 1.76 30.62 ± 1.58 69.38 ± 2.35
SFA, saturated fatty acids; UFA, unsaturated fatty acids. Note: Data are expressed as mean ± standard deviations of three replicates.
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of fatty acids for biodiesel production and may affect the quality of the biodiesel produced. Table 4 shows that the Chlorella sp. lipids for all reaction times are composed mainly of unsaturated fatty acids (60–75%), with a significant percentage of palmitic acid also present (25–38%). Among the five reaction times applied in our study, the shortest reaction time (2 h) shows the highest percentage of UFAs, and special attention should be paid to the linoleic acid because the C18:2 unsaturated chains were the predominant fatty acids (45.6%) followed by C18:3 (25.2%) and C16:0 (24.3%). It is also important to highlight that higher compositions of UFAs can reduce the pour point of biodiesel and making it feasible to be used in cold climate countries.[29] However, when the UFA with double bond ≥ 3, their high content is undesirable since it reduces the chemical and oxidative stability of the resulting biodiesel.[27] Therefore, the higher content of unsaturated chains (such as C18:1 and C18:2) and the lower content of C18:3 in Chlorella sp. at 2 h of reaction time may be expected to produce a good quality of biodiesel. Further study will be necessary to uncover the feasibility and effectiveness of various harvesting methods for the production of high-quality biodiesel from Chlorella sp., then the physical and fuel properties may be comparable to diesel fuel and the ASTM biodiesel standard.
Conclusion The kinetics for oil extraction from Chlorella sp. biomass by a mixture of chloroform/methanol (1:1, v/v) were found to be a fractional kinetic with the order of about 1.5 by the differential method and strongly dependent on temperature. The activation energy was Ea = 38.893 kJ/mol and the thermodynamic activation parameters at 60◦ C were S = = −180.190 J/mol K, H = = 36.124 kJ/mol and G = = 96.128 kJ /mol. The enthalpy value was found to be positive (H = 0.308 kJ/mol), indicating that the process is endothermic and requires the input of energy during the process. The other thermodynamic parameters at 60◦ C were calculated to be S = 4.420 1/mol K and G = −1.164 kJ/mol. Our study also demonstrated that palmitic acid (C16:0), linoleic acid (C18:2) and linolenic acid (C18:3) are major fatty acids found in Chlorella sp., indicating a good potential for use of the Chlorella sp. lipids in the production of biodiesel. The variation of temperature influenced the oleic acid (C18:1) and linolenic acid (C18:3) contents strongly, as an increase from 70 to 90◦ C increased the content of linolenic acid from 19.22 to 26.49% and decreased the oleic acid from 2.66 to 2.44%. Finally, 2 h of reaction time yields the most appropriate fatty acid composition in terms of unsaturated fatty acids such as C18:1 and C18:2. The next research effort will address the harvesting of Chlorella sp. cells using appropriate and efficient methods, making Chlorella sp. biomass a good candidate for biodiesel production.
Acknowledgements All authors are affiliated with the Membrane Science and Technology Cluster USM.
Funding This material is based on the work supported by a Research University (RU) [Grant No. 1001/PJKIMIA/814060) and a Postgraduate Research (PRGS) [Grant No. 1001/JKIMIA/8044014] from Universiti Sains Malaysia (USM). N.H. Mat Yasin gratefully acknowledges Universiti Malaysia Pahang [SLAB 2011] for a scholarship.
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