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Potential of using organic fertilizer to cultivate Chlorella vulgaris for biodiesel production
Applied Energy 94 (2012) 303–308
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Potential of using organic fertilizer to cultivate Chlorella vulgaris for biodiesel production Man Kee Lam, Keat Teong Lee ⇑ School of Chemical Engineering, Universiti Sains Malaysia, Engineering Campus, Seri Ampangan, 14300 Nibong Tebal, Pulau Pinang, Malaysia
a r t i c l e
i n f o
Article history: Received 20 September 2011 Received in revised form 27 January 2012 Accepted 31 January 2012 Available online 25 February 2012 Keywords: Biodiesel Microalgae Chlorella vulgaris Organic fertilizer
a b s t r a c t Cultivating microalgae at industrial scale for biodiesel production required substantial amount of mineral fertilizer, typically nitrogen and phosphorus. In fact, the production of mineral fertilizer implies the usage of energy and fossil fuels resulting to unsustainable practise in a long term. On the other hand, organic fertilizer which is derived from food waste, biomass or manure also contains high value of nutrients that can support microalgae growth. Hence, in the present study, the potential of using organic fertilizer as an alternative nutrient source to cultivate Chlorella vulgaris was investigated. Under the supplement of organic nutrients, it was found that C. vulgaris grown favourably under the following conditions: initial nitrate content of 26.67 mg/L, 24 h of continuous illumination and pH of 5. Nevertheless, slow growth rate was observed when cultivating C. vulgaris under open environment; a reduction of 27% was recorded in comparison with controlled environment. On the other hand, it was possible to reutilize the water to re-cultivate C. vulgaris. This observation reflects the high adaptability of C. vulgaris towards the surrounding environment and suitability to be grown under outdoor conditions. Total lipid of 18.1% from C. vulgaris biomass was successfully extracted and the fatty acids methyl ester profile was proven to be suitable for making biodiesel. Ó 2012 Elsevier Ltd. All rights reserved.
1. Introduction Sustainable energy development has become one of the key challenges in this century. Depletion of fossil fuel in the near future and the effect of green house gases towards human and environment have escalated the search of renewable energies, such as solar, wind, hydro, wave and geothermal power [1]. However, these renewable energies are heavily dependent on regional or local condition that can be very unpredictable and inconsistent. On the other hand, biodiesel being recognized as a green and alternative renewable fuel has attracted great interest from researchers, governments, local and international traders. Some of the advantages of using biodiesel instead of fossil diesel are it is a non-toxic fuel, biodegradable and lower emission of green house gases when burned in diesel engine [2,3]. Normally, biodiesel is produced through transesterification reaction, in which vegetable oil reacts with short chain alcohol (e.g. methanol) in the presence of catalyst (e.g. sodium hydroxide, NaOH) [4,5]. Edible oils such as soybean, sunflower, rapeseed and palm oil are the common feedstock for biodiesel production. However, continuous growth of the biodiesel industries in the last ten years has raised a tragic social issue: the food versus fuel dispute [6]. It is strongly believed that further ⇑ Corresponding author. Tel.: +60 4 5996467; fax: +60 4 5941013. E-mail address: [email protected] (K.T. Lee). 0306-2619/$ - see front matter Ó 2012 Elsevier Ltd. All rights reserved. doi:10.1016/j.apenergy.2012.01.075
expansion of biodiesel industries in the global market will result to more undernourished people suffering from hunger and malnutrition [7,8]. Currently, microalgae have been identified as a superior feedstock for biodiesel production, mainly due to their fast growth rate (100 times faster than terrestrial plant) and able to double their biomass in less than one day under favourable culture conditions [9]. Apart from that, certain microalgae strains are able to accumulate sufficient amount of lipid that can be expected to overcome the terrestrial crops [10]. For example, microalgae with 30% of lipid content have the potential to produce 54 tonnes of oil/ha/year, whereas palm oil and jatropha are only able to produce 3.62 and 4.13 tonne of oil/ha/year, respectively [11,12]. Thus, cultivating microalgae for biodiesel production requires only a minimum land area and up hold an important key for a more sustainable land utilization [13]. Apart from that, an added advantage of cultivating microalgae is the ability of the microorganism to act as a carbon sink assimilating CO2 from atmosphere and flue gases through photosynthesis; a golden opportunity for carbon credit program [14–16]. However, one of the limitations to cultivate microalgae in industrial scale is the availability of nutrients sources. Chemical or inorganic fertilizers are commonly used to achieve promising growth rate of microalgae. Nevertheless, a life cycle assessment on microalgae cultivation has underlined that 50% of energy use
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and green house gases emissions are associated with chemical fertilizer production [17]. Meanwhile, utilizing secondary or tertiary wastewater as nutrients source to cultivate microalgae appears as a promising choice to reduce the overall energy input [18–21]. This is because wastewater contains high concentration of nitrate and phosphate that promote the growth of microalgae and at the same time, microalgae can act as a purifier to further remove the nutrients from wastewater before discharging to water sources. Nevertheless, the key challenges of using wastewater are susceptible contamination and inconsistency of nutrients compositions; these factors will directly retard the growth of microalgae. On the other hand, using organic fertilizer as a nutrient source offers an alternative and convenient way to minimize the contamination level in water. Organic fertilizer can be obtained through a series of composting and refining processes from waste materials such as manure, sewage sludge, biomass and food [22]. Although CO2 is released during composting process, however, the amount is lower compared to the production of inorganic fertilizer. It was estimated that 4–82 kg of CO2 and 4–67 kg of CO2 could be saved by composting one tonne of food waste and garden waste, respectively [23]. In addition, the emitted CO2 from composting process can be recycled as an alternative carbon source to cultivate microalgae. Normally, organic fertilizers available in the market are made in dry pellet form for easy transportation and storage. Hence, recycle and reuse the waste materials to cultivate microalgae for biodiesel production appears as a greener solution to drive the industry towards sustainable growth. Thus, the objective of the present study is to evaluate the potential of using organic fertilizer to cultivate Chlorella vulgaris. Various culture parameters such as amount of nitrates, illumination duration, pH, outdoor cultures and the effect of reuse the recycling water were systematically investigated. All experiments were performed using 5 L photobioreactor (except seed culture) instead of using lab scale conical flask and tap water was used rather than using distilled or sterilized water. This approach was implemented to promote an easier up-scaling and to minimize the overall energy consumption in cultivating microalgae in commercial scale.
paper (Double Rings 101). The resulting organic fertilizer medium was dark-brown in colour with typical characteristics as shown in Table 1. From the Table, the concentrated organic medium contains two types of nutrients which are equally important to support microalgae growth: (1) macronutrients, such as nitrogen, phosphorus, potassium, calcium and magnesium and (2) micronutrients, such as manganese, boron and iron. In addition, the prepared medium also contains exceptional high level of Chemical Oxygen Demand (COD) and Biochemical Oxygen Demand (BOD), in which those values are similar to untreated wastewater [24]. On the other hand, the filtered particulate solids (organic compost) can be further used as soil amendment to increase the water and nutrients holding capacity and to improve soil physical properties by providing better aeration [25]. Subsequently, a pre-determined volume of the organic fertilizer medium was introduced into a photobioreactor with 5 L of tap water (without sterilization) and the pH of the medium was adjusted according to pre-determined values. Then, C. vulgaris with initial cell concentration of 0.3 106 cells (around 10 mL from the seed culture) was introduced into the photobioreactor. The photobioreactor was aerated with compressed air continuously and illuminated with cool-white fluorescent light (Philip TL-D 36W/865, light intensity of 60–70 lmol m2 s1).
2. Material and methods
l ðday1 Þ ¼
2.1. Pure microalgae strain and culture conditions A wild-typeC. vulgaris was isolated from local freshwater located at Penang, Malaysia. The microalgae was preserved and grown in Bold’s Basal Medium (BBM), consisting of: (1) 10 mL per liter of culture medium with the following chemicals: NaNO3 (25 g/L), CaCl22H2O (2.5 g/L), MgSO47H2O (7.5 g/L), K2HPO4 (7.5 g/L), KH2PO4 (17.5 g/L), NaCl (2.5 g/L) and (2) 1 mL per liter of culture medium with the following chemicals: EDTA anhydrous (50 g/L), KOH (31 g/L), FeSO47H2O (4.98 g/L), H2SO4 (1 mL), H3BO3 (11.4 g/ L), ZnSO47H2O (8.82 g/L), MnCl24H2O (1.44), MoO3 (0.71 g/L), CuSO45H2O (1.57 g/L), Co(NO3)26H2O (0.49 g/L). The initial pH of the medium was adjusted to 6.8. The seed culture was grown in a 100 mL Erlenmeyer flask containing 50 mL of medium, aerated with compressed air, surrounding temperature of 25–28 °C and illuminated with cool-white fluorescent light (Philip TL-D 36W/ 865, light intensity of 60–70 lmol m2 s1) continuously. 2.2. Cultivating microalgae with organic fertilizer Baja Serbajadi Humus 27 (organic fertilizer or compost) with granular shape was purchased from a local market. A 10 g of the fertilizer was immersed in 600 mL tap water and stirred for 24 h using magnetic stirrer. Non-soluble particulate solids were observed after the stirring process and were filtered using filter
2.3. Measurement of microalgae growth A correlation between the optical density of C. vulgaris and biomass was pre-determined. Optical density was measured daily at 540 nm using spectrophotometer (Shimadzu UV mini-1240) [26]. Then, 10 mL of sample were centrifuged at 10 1000g for 5 min. The supernatant was slowly decanted back into the culture medium whereas the microalgae biomass were dried in an oven at 100 °C for 24 h. All the samplings were performed in triplicate to ensure the accuracy of the data. The correlation is shown below:
Dry weight ðg=LÞ ¼ 0:4541 OD540 ; R2 ¼ 0:9890
ð1Þ
The specific growth rate (l) was measured by using below:
lnðN2 =N1 Þ t2 t1
ð2Þ
where N1 and N2 are defined as biomass (g/L) at time t1 and t2, respectively. Table 1 Characteristics of organic fertilizer medium (obtained by mixing 10 g of organic fertilizer with 600 mL tap water). Parameter
Unit
Concentration
CODa BOD – 5 days test at 20 °Cb Nitrogenc Phosphorusd Potassiume Calciumf Magnesiumg Manganeseh Boroni Ironj
ppm ppm ppm ppm ppm ppm ppm ppm ppm ppm
1729.9 576.0 1323.2 213.6 634.4 269.9 54.5 1.0 4.1 1.3
Testing method: a APHA 5220 B (2005). b APHA 5210 B (2005). c APHA 4500-NH3F. d APHA 4500-P E (2005). e APHA 3111 B (2005). f APHA 3111 B (2005). g APHA 3111 B (2005). h APHA 3111 B (2005). i APHA 4500-B C (2005). j APHA 3111 B (2005).
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2.4. Measurement of nitrate content in medium Nitrate concentration was determined according to the Ultraviolet Spectrophotometric Screening Method [27]. A sample (1 mL) collected from photobioreactor was centrifuged at 10 1000 g for 5 min. The supernatant was collected and optical density was measured at 275 nm and 220 nm by using spectrophotometer (Shimadzu UV mini-1240). Then, the absorbance reading at 275 nm was subtracted two times from the reading at 220 nm to obtain the actual absorbance caused by the presence of nitrate. Dry potassium nitrate (KNO3) at different concentrations was used for calibration purposes. 2.5. Microalgae harvesting and biomass collection When C. vulgaris had grown to stationary phase, air aeration to the culture medium was stopped. The microalgae were let to settle naturally to the bottom of photobioreactor for two days. Two distinguished layers were observed, in which the upper layer consist of water with suspended microalgae cells and the bottom layer was microalgae biomass. The upper layer water was slowly decanted, leaving behind the microalgae biomass which was further dried in an oven at 80 °C for 24 h. The dried microalgae biomass were collected and sealed in an empty container for lipid extraction.
þ sources such as nitrate ðNO 3 Þ, ammonia ðNH4 Þ or urea (NH2–CO– NH2) appear to be the most significant contributor to sustain microalgae growth [30]. In the present study, synthesized nutrients from organic fertilizer were diluted to different concentration with the purpose to study the ability of C. vulgaris in up taking the nutrients. Fig. 1 depicts the growth of C. vulgaris with different nitrate concentration. From the figure, C. vulgaris was found to grow even under limited nutrients condition (5 and 10 mL of nutrients), however, the quantity of biomass produced were unsatisfactory: 0.1– 0.14 g/L after 12 days of cultivation. In comparison, C. vulgaris grew much better with sufficient supplement of nutrients (80 and 100 mL). The biomass produced were 0.29–0.31 g/L, which was nearly 150% of increment. Apart from that, specific growth rate of C. vulgaris was also improved due to supplement of ample nutrients. For example, the specific growth rate was 0.132 day1 and 0.229 day1 for 5 and 100 mL of nutrients added respectively; an increment of 73% was observed. This result indicates that large quantities of nutrient sources are the mandatory step to promote microalgae growth and consequently to attain higher biomass yield. On the other hand, the growth curve for C. vulgaris with excess amount of nutrients (150 mL) exhibited the same trend as 100 mL, in which no further improvement of biomass yield was observed. This is because other parameters such as photoperiod and pH control will become the growth controlling factors if adequate nutrients are supplied.
2.6. Microalgae lipid extraction
3.2. Effect of photoperiod
Ten grams of dried C. vulgaris biomass were placed in a cellulose thimble and extraction process was performed using Soxhlet extractor. Four types of solvent were used to compare the extraction efficiency, such as n-hexane, methanol, ethanol and mixed methanol–chloroform with volume ratio of 2:1 [28]. A total of 250 mL for each solvent was placed in Soxhlet extractor and heated at 60–65 °C for 24 h. After that, the solvent was evaporated in a rotary evaporator and the leftover lipid was collected. The residues (solid material) after evaporation were subjected to repeated extraction twice using the same solvent. Thereafter, crude microalgae lipids were measured gravimetrically. All samplings were performed in triplicate.
The effect of photoperiod on microalgae cultivation exhibited a significant influence towards microalgae photosynthetic activity and growth rates in a typical cultivation system [31,32]. However, it should be noted that microalgae with excessive of light exposure would result to unwanted electricity consumption (energy waste) and also inhibit the growth of microalgae [33]. Fig. 2 shows the effect of photoperiod towards the growth of C. vulgaris. From the figure, the growth of C. vulgaris was strongly affected by photoperiod, with the highest biomass yield attained at 0.31 g/L through continuous 24 h light illumination for 12 days (specific growth rate of 0.228 day1). In addition, for photoperiod of 3 h, 6 h and 9 h, the biomass attained were only 0.03–0.1 g/L which was exceptional unsatisfactory. This result indicates that longer photoperiod will result to continuous uptake of nutrients by C. vulgaris through photosynthesis and subsequently, increase their biomass at the same cultivation time. In other words, this result also indicates
Transesterification was performed using 1 g of crude C. vulgaris, methanol to lipid molar ratio of 15:1 and 3 wt.% of concentrated sulfuric acid (H2SO4) as catalyst. The reaction was carried out in a water bath shaker at 60 °C for 3 h. Upon completion of the reaction, 1 lL of the reaction product was subjected to gas chromatography-mass spectrometry (GC–MS; PerkinElmer Clarus 600) analysis. The GC was equipped with flame ionization detector (FID) and Elite 5-MS column (30 m 0.25 mm 0.25 mm). The initial oven temperature was 65 °C, held for 2 min and raised to 280 °C at ramping rate of 8 °C/min and held at 280 °C for 10 min, while the injector temperature was set to 250 °C. The compounds detected were identified and quantified using NIST Mass Spectral Search Program.
50 mL 80 mL
5 mL 10 mL
0.35
20 mL 30 mL
100 mL 150 mL
0.3
Biomass (g /L)
2.7. Transesterification reaction and fatty acid methyl esters (FAME) analysis
0.25 0.2 0.15 0.1 0.05
3. Result and discussion
0 0
3.1. Effect of nutrients concentration Sufficient supplement of nutrients for microalgae to grow is the first key step to produce bulk quantity of microalgae biomass [29]. Among all the nutrients required, adequate supply of nitrogen
2
4
6
8
10
12
Cultivation time (days) Fig. 1. Effect of initial nitrate concentration towards the growth of C. vulgaris. Other culture conditions: pH = 7 and illuminated for 24 h continuously. Initial nitrate content (mg/L): 5 mL = 1.33, 10 mL = 2.69, 20 mL = 5.38, 30 mL = 7.98, 50 mL = 13.09, 80 mL = 20.94, 100 mL = 26.67, 150 mL = 40.71.
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0.4
3h
12 h
9h
6h
0.6
24 h
8
7
6
9
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0.3
Biomas (g /L)
Biomass (g /L)
0.35
5
4
3
0.25 0.2 0.15 0.1
0.4 0.3 0.2 0.1
0.05 0 0
0 0
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Cultivation time (days)
that C. vulgaris is weak in storing substantial amount of light energy for later use in the dark period (exergonic reactions) [32]. Hence, surface area of illumination should be taken into consideration when designing a photobioreactor to cultivate C. vulgaris (typically outdoor cultivation), so that optimum light energy can be efficiently utilized by the microalgae. From Fig. 2, it was clear to observe that the quantity of biomass attained at 12 h of photoperiod time (equivalent to outdoor sunlight cycle) were nearly double the quantity of biomass attained at photoperiod of 3 h, 6 h and 9 h. This result proved that it is feasible to cultivate C. vulgaris under outdoor conditions; however, it is still dependent on the local weather condition and temperature. 3.3. Effect of initial pH One of the controlling factors to cultivate microalgae is the pH of the culture medium [34]. Generally, pH of 7–8 is plausible for microalgae to grow [34]. Nevertheless, it is worth to mention that suitable microalgae strains for biodiesel production should not only contain high lipid yield, but should be able to withstand pH changes during cultivation cycle. For example, high pH value (9.0–9.5) was observed in photobioreactor due to shortage of inorganic carbon after rapid consumption by microalgae [35]. Sometimes, pH can also drop drastically if high concentration of CO2 is continuously supplied to the culture medium and caused adverse effect towards the growth of microalgae [19]. More importantly, in the near future, when flue gases are used to cultivate microalgae in industrial scale, the selected microalgae strains should be able to tolerate inconsistent concentration of CO2 in the flue gases that indirectly resulted to pH variation in the culture medium [36]. Fig. 3 shows the effect of initial pH in the culture medium for C. vulgaris. From the figure, C. vulgaris exhibited almost a linear growth at pH of 4, 5 and 8 with maximum biomass of 0.47–0.51 g/L obtained after 12 days of cultivation. Furthermore, higher specific growth rate was attained at pH 4 (0.265 day1), 5 (0.270 day1) and 8 (0.263 day1) compared to culture growth at neutral condition, pH 7 (0.229 day1). This result indicates that C. vulgaris can adapt very well even at low or high pH. One of the advantages of cultivating microalgae at low or high pH is the simultaneous elimination of contaminants such as fungus and therefore, able to sustain their growth naturally. Nevertheless, at pH 3, the growth of C. vulgaris was reverted and no significant increment of biomass was detected after 12 days of cultivation. On the other hand, although the growth of C. vulgaris was satisfactory at pH 6, 7 and 9, a lag phase was observed from day 1 to day 4 and the growth was almost stagnant from day 9 to day 12 with a maximum 0.33 g/L of biomass obtained.
8
10
12
Fig. 3. Effect of initial pH towards the growth of C. vulgaris. Other culture conditions: Initial nutrients volume = 100 mL (nitrate content of 26.67 mg/L), illuminated for 24 h continuously.
3.4. Comparison of the growth of C. vulgaris with organic and inorganic fertilizer under indoor and outdoor conditions Up to date, research on microalgae is focussed on indoor culture where a control environment is relatively easy to achieve. Nevertheless, for the purpose of cultivating microalgae in a commercial scale, outdoor cultivation system is the most feasible option due to easy access to sunlight [37]. This strategy will help to reduce the overall energy input (mainly for illumination and temperature control) and consequently, enhance the cost effectiveness to produce biodiesel from microalgae. However, one of the limitations to cultivate microalgae under outdoor conditions is the inconsistent changes of local weather, temperature and light intensity. Therefore, microalgae strains that are able to adapt and grow well under these extreme cultivation conditions will be an added advantage. Fig. 4 shows a comparison of the growth of C. vulgaris (supplied with organic or inorganic nutrients) under indoor and outdoor conditions. From the figure, C. vulgaris was found to grow much faster with inorganic nutrients compared to organic nutrients, especially when cultivated under indoor conditions with specific growth rate of 0.30 day1 and 0.27 day1, respectively. The
0.8
Indoor (organic)
Indoor (inorganic)
Outdoor (organic)
Outdoor (inorganic)
0.7 0.6
Biomass (g /L)
Fig. 2. Effect of photoperiod (hours of light) towards the growth of C. vulgaris. Other culture conditions: initial nutrients volume = 100 mL (nitrate content of 26.67 mg/ L) and pH = 7.
6
Cultivation time (day)
0.5 0.4 0.3 0.2 0.1 0 0
2
4
6
8
10
12
14
16
Cultivation time (day) Fig. 4. Comparison of indoor and outdoor culture towards the growth of C. vulgaris. Culture conditions: (a) organic nutrients; initial nutrients volume = 100 mL (nitrate content of 26.67 mg/L) and pH = 5 and (b) inorganic nutrients; BBM medium and pH = 5. Photoperiods: (a) indoor: 24 h illumination and (b) outdoor: 12 h light and 12 h dark.
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result is rather expected due to controlled conditions and ample macro- and micro-nutrients present in the inorganic nutrients. Apart from that, by referring to Fig. 4, C. vulgaris was observed to grow slower under outdoor environment, either supplied with organic or inorganic nutrients. Under uncontrolled environment, the temperature at outdoor varied from minimum 23 °C to maximum 35 °C. In addition, a lag phase was detected from day 1 to day 3 of culture, showing that the microalgae required some time to adapt themselves to the new environment. As a consequence, the specific growth rate of microalgae cultivated under outdoor conditions was dropped to 0.20 day1 for both organic and inorganic nutrients. This observation indicates that outdoor conditions will become the key controlling factor towards microalgae growth when sufficient nutrients supplement is available. In this regards, using organic fertilizers to cultivate microalgae under uncontrolled environment will be an added advantage since the final biomass attained was nearly the same with the inorganic nutrients. Apart from that, it was found that the C. vulgaris cultivated with organic nutrients under outdoor conditions attained higher biomass yield (0.37 g/L) on day 12 compared to indoor culture with 12 h illumination (0.21 g/L). This may be due to the intensity of sunlight that is much more suitable to cultivate C. vulgaris compared to illumination by fluorescent lamp.
3.5. Effect of using recycling water Effective uses of water resources play a significant role in microalgae cultivation since microalgae are aquatic microorganisms that need water to survive [38]. In large scale microalgae cultivation system, huge amount of water is required to reach optimum productivity of microalgae biomass. Therefore, to recycle and reuse the water not only can save the overall operation cost, but also helps to preserve the water for other purposes [39]. However, not all microalgae strains are able to re-grow effectively in recycled water due to susceptible contamination by fungus and bacteria (e.g. open pond system). Hence, the selected microalgae strains for biodiesel production should be able to grow rapidly and withstand other contaminants. Fig. 5 shows the effect of using recycled water from the previous batch culture (indoor and outdoor, respectively) without any pre-treatment or purification process. The recycled water still contained minimum amount of nutrients and microalgae cells which were not completely harvested. From the figure, the growth of C. vulgaris under indoor conditions was not affected by the use of recycled water and produced satisfactory
Indoor
Outdoor
0.6
3.6. Lipid extraction and biodiesel production from C. vulgaris Up to now, lipid extraction using chemical solvent is the most reliable method to determine the overall lipid content presence in microalgae. Extraction efficiency by chemical solvent is higher than physical method (mechanical pressing machine), typically due to the high polarity of fatty acids toward the chemical solvent. Nevertheless, the effectiveness of different type of chemical solvent used is strongly dependent on microalgae strains, especially when the existence of cell wall in microalgae will indirectly impede chemical solvent extraction efficiency [40]. In the present study, four different types of solvent were used for lipid extraction study: n-hexane, methanol, ethanol and methanol mixed with chloroform with volume ratio of 2:1 (Bligh and Dyer method). Bligh and Dyer method gave the highest extraction efficiency, in which 18.1% of lipid was successfully extracted. This value is comparable to the total lipid of C. vulgaris reported in the literature, which lies between 5% and 40% under phototrophic culture condition [24]. The solvent extraction efficiency in descending order was methanol (15.5%), ethanol (10.7%) and n-hexane (3.2%). Although n-hexane has been widely used in oil extraction study, nevertheless, it is not a suitable choice to extract lipid from C. vulgaris. This is because n-hexane is a non-polar solvent which has poor permeability in cell wall and hence only extracellular lipids are extracted [41]. The fatty acids methyl ester compositions of C. vulgaris mainly consisted of C16:0, C16:1, C16:2, C18:1, C18:2 and C18:3 as shown in Fig. 6. From the figure, C16:0 (palmitic acid methyl ester), C18:1 (oleic acid methyl ester) and C18:2 (linoleic acid methyl ester) represent the major portion of fatty acid methyl esters compositions, accounted for total of 85.6%. These fatty acids are naturally found in oil bearing crops, such as soybean, sunflower, cottonseed and palm oil, in which the fatty acids are suitable for biodiesel production. Apart from that, unsaturated fatty acids methyl esters (C16:1, C16:2, C18:1, C18:2, C18:3) were predominant in the fatty acids profile, accounted for 74%. It is important to note that higher compositions of unsaturated fatty acids methyl ester can reduce the pour point of biodiesel and making it feasible to be used in cold climate countries [42]. Other saturated fatty acids methyl ester
0.4 0.3 0.2 0.1 0 0
2
4
6
8
10
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14
16
Cultivation time (day) Fig. 5. Effect of reuse the recycling water towards the growth of C. vulgaris. Culture conditions: Initial nutrients volume = 100 mL (nitrate content of 26.67 mg/L) and pH = 5. Photoperiods: (a) indoor: 24 h illumination and (b) outdoor: 12 h light and 12 h dark.
Fatty acids methyl ester concentration
Biomass (g /mL)
0.5
quantity of biomass within 16 days of cultivation time. Apart from that, for C. vulgaris cultivated at outdoor using recycled water, the lag phase from day 1 to day 3 was greatly minimized if compared with the use of fresh water (Fig. 4). This is because C. vulgaris cells that were not harvested and remain in the recycled water have already adapted to the outdoor environment. Therefore, once the new organic nutrients were supplied to the culture medium, the microalgae can utilize the nutrients immediately. Consequently, higher biomass quantity (0.55 g/L) was obtained compared to the use of fresh water under outdoor conditions (0.37 g/L).
C16:0
C16:1
C16:2
C18:1
C18:2
C18:3
Fatty acids methyl ester profile Fig. 6. Fatty acids methyl ester profile of C. vulgaris.
Others
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except C16:0 (palmitic acid methyl ester) were also identified, such as C14:0 (myristic acid methyl ester), C15:0 (pentadecanoic acid methyl ester) and C18:0 (stearic acid methyl ester). However, the compositions were minimum and they accounted for only 1.1% of the overall lipid compositions. 3.7. Estimation of nutrients cost in cultivating C. vulgaris Based on the optimum cultivating conditions of C. vulgaris using organic and inorganic fertilizer, a simple cost analysis was conducted. The cost of organic fertilizer purchased at local market was 1.2 USD per pack (400 g) whereas the overall cost to prepare the inorganic fertilizer (BBM) was 15–25 USD per liter. In addition, for the preparation of organic nutrients, electricity cost for stirring and cost for solid residue filtration were included. Consequently, the total cost of nutrients required to produce 1 kg of C. vulgaris biomass was estimated to be 2.5–3 USD and 60–85 USD for organic and inorganic fertilizer, respectively. Hence, utilization of cheap nutrient sources such as organic fertilizer will be more beneficial in large scale microalgae cultivation system, typically in terms of cost saving and better environmental protection. 4. Conclusions The result in this study showed that it is feasible to use organic fertilizer to cultivate C. vulgaris for biodiesel production. This approach offers several advantages in terms of environmental perspective and cost effectiveness. In addition, this method can be also easily adopted since organic fertilizers are widely available in market. More importantly, C. vulgaris was found to be able to reproduce when grown under outdoor conditions (uncontrolled environment). Besides, reutilize the water to re-cultivate C. vulgaris is possible and this will further drive the microalgae cultivation system towards a more sustainable process. Acknowledgements The authors would like to acknowledge the funding given by Universiti Sains Malaysia (Research University Grant No. 814146, Postgraduate Research Grant Scheme No. 8044031 and USM Vice-Chancellor’s Award) for this project. References [1] Leung DYC, Wu X, Leung MKH. A review on biodiesel production using catalyzed transesterification. Appl Energy 2010;87:1083–95. [2] Demirbas A. Progress and recent trends in biodiesel fuels. Energy Convers Manage 2009;50:14–34. [3] Qiu F, Li Y, Yang D, Li X, Sun P. Biodiesel production from mixed soybean oil and rapeseed oil. Appl Energy 2011;88:2050–5. [4] Vasudevan PT, Briggs M. Biodiesel production – current state of the art and challenges. J Ind Microbiol Biotechnol 2008;35:421–30. [5] Lam MK, Lee KT, Mohamed AR. Homogeneous, heterogeneous and enzymatic catalysis for transesterification of high free fatty acid oil (waste cooking oil) to biodiesel: a review. Biotechnol Adv 2010;28:500–18. [6] Wu X, Leung DYC. Optimization of biodiesel production from camelina oil using orthogonal experiment. Appl Energy 2011;88:3615–24. [7] Srinivasan S. The food versus fuel debate: a nuanced view of incentive structures. Renew Energy 2009;34:950–4. [8] Amaro HM, Guedes AC, Malcata FX. Advances and perspectives in using microalgae to produce biodiesel. Appl Energy 2011;88:3402–10. [9] Tredici MR. Photobiology of microalgae mass cultures: understanding the tools for the next green revolution. Biofuels 2010;1:143–62. [10] Rasoul-Amini S, Montazeri-Najafabady N, Mobasher MA, Hoseini-Alhashemi S, Ghasemi Y. Chlorella sp.: a new strain with highly saturated fatty acids for biodiesel production in bubble-column photobioreactor. Appl Energy 2011;88:3354–6. [11] Chisti Y. Biodiesel from microalgae. Biotechnol Adv 2007;25:294–306. [12] Lam MK, Lee KT. Renewable and sustainable bioenergies production from palm oil mill effluent (POME): Win–win strategies toward better environmental protection. Biotechnol Adv 2010;29:124–41.
[13] Huang G, Chen F, Wei D, Zhang X, Chen G. Biodiesel production by microalgal biotechnology. Appl Energy 2010;87:38–46. [14] Li Y, Horsman M, Wu N, Lan CQ, Dubois-Calero N. Biofuels from microalgae. Biotechnol Prog 2008;24:815–20. [15] Wang B, Li Y, Wu N, Lan CQ. CO2 bio-mitigation using microalgae. Appl Microbiol Biotechnol 2008;79:707–18. [16] Demirbas A. Biodiesel from oilgae, biofixation of carbon dioxide by microalgae: a solution to pollution problems. Appl Energy 2011;88: 3541–7. [17] Clarens AF, Resurreccion EP, White MA, Colosi LM. Environmental life cycle comparison of algae to other bioenergy feedstocks. Environ Sci Technol 2010;44:1813–9. [18] Levine RB, Costanza-Robinson MS, Spatafora GA. Neochloris oleoabundans grown on anaerobically digested dairy manure for concomitant nutrient removal and biodiesel feedstock production. Biomass Bioenergy 2010;35:40–9. [19] Kong QX, Li L, Martinez B, Chen P, Ruan R. Culture of microalgae chlamydomonas reinhardtii in wastewater for biomass feedstock production. Appl Biochem Biotechnol 2010;160:9–18. [20] Mata TM, Martins AA, Caetano NS. Microalgae for biodiesel production and other applications: a review. Renew Sust Energy Rev 2010;14:217–32. [21] Rawat I, Ranjith Kumar R, Mutanda T, Bux F. Dual role of microalgae: phycoremediation of domestic wastewater and biomass production for sustainable biofuels production. Appl Energy 2011;88:3411–24. [22] Marzouk HA, Kassem HA. Improving fruit quality, nutritional value and yield of Zaghloul dates by the application of organic and/or mineral fertilizers. Sci Hortic 2010;127:249–54. [23] Boldrin A, Andersen JK, Møller J, Christensen TH, Favoino E. Composting and compost utilization: accounting of greenhouse gases and global warming contributions. Waste Manage Res 2009;27:800–12. [24] Chinnasamy S, Bhatnagar A, Hunt RW, Das KC. Microalgae cultivation in a wastewater dominated by carpet mill effluents for biofuel applications. Bioresource Technol 2010;101:3097–105. [25] Tisserat B, O’kuru RH, Cermak SC, Evangelista RL, Doll KM. Potential uses for cuphea oil processing byproducts and processed oils. Ind Crop Prod 2012;35:111–20. [26] Gouveia L, Oliveira AC. Microalgae as a raw material for biofuels production. J Ind Microbiol Biotechnol 2009;36:269–74. [27] Greenberg A, Clesceri L, Eaton A. Standard methods for the examination of water and wastewater. NO3-ultraviolet spectrophotometric screening method. American Public Health Association, America Water Works Association and Water Environment Federation, Washington, 1992; 18th ed., 4–87. [28] Bligh EG, Dyer WJ. A rapid method of total lipid extraction and purification. Can J Biochem Physiol 1959;37:911–7. [29] Xin L, Hong-ying H, Ke G, Ying-xue S. Effects of different nitrogen and phosphorus concentrations on the growth, nutrient uptake, and lipid accumulation of a freshwater microalga Scenedesmus sp. Bioresour Technol 2010;101:5494–500. [30] Hsieh C-H, Wu W-T. Cultivation of microalgae for oil production with a cultivation strategy of urea limitation. Bioresour Technol 2009;100:3921–6. [31] Janssen M, De Winter M, Tramper J, Mur LR, Snel J, Wijffels RH. Efficiency of light utilization of Chlamydomonas reinhardtii under medium-duration light/ dark cycles. J Biotechnol 2000;78:123–37. [32] Jacob-Lopes E, Scoparo CHG, Lacerda LMCF, Franco TT. Effect of light cycles (night/day) on CO2 fixation and biomass production by microalgae in photobioreactors. Chem Eng Process 2009;48:306–10. [33] Chen CY, Yeh KL, Su HM, Lo YC, Chen WM, Chang JS. Strategies to enhance cell growth and achieve high-level oil production of a C. vulgaris isolate. Biotechnol Prog 2010;26:679–86. [34] Zeng X, Danquah MK, Chen XD, Lu Y. 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Applied Energy journal homepage: www.elsevier.com/locate/apenergy
Potential of using organic fertilizer to cultivate Chlorella vulgaris for biodiesel production Man Kee Lam, Keat Teong Lee ⇑ School of Chemical Engineering, Universiti Sains Malaysia, Engineering Campus, Seri Ampangan, 14300 Nibong Tebal, Pulau Pinang, Malaysia
a r t i c l e
i n f o
Article history: Received 20 September 2011 Received in revised form 27 January 2012 Accepted 31 January 2012 Available online 25 February 2012 Keywords: Biodiesel Microalgae Chlorella vulgaris Organic fertilizer
a b s t r a c t Cultivating microalgae at industrial scale for biodiesel production required substantial amount of mineral fertilizer, typically nitrogen and phosphorus. In fact, the production of mineral fertilizer implies the usage of energy and fossil fuels resulting to unsustainable practise in a long term. On the other hand, organic fertilizer which is derived from food waste, biomass or manure also contains high value of nutrients that can support microalgae growth. Hence, in the present study, the potential of using organic fertilizer as an alternative nutrient source to cultivate Chlorella vulgaris was investigated. Under the supplement of organic nutrients, it was found that C. vulgaris grown favourably under the following conditions: initial nitrate content of 26.67 mg/L, 24 h of continuous illumination and pH of 5. Nevertheless, slow growth rate was observed when cultivating C. vulgaris under open environment; a reduction of 27% was recorded in comparison with controlled environment. On the other hand, it was possible to reutilize the water to re-cultivate C. vulgaris. This observation reflects the high adaptability of C. vulgaris towards the surrounding environment and suitability to be grown under outdoor conditions. Total lipid of 18.1% from C. vulgaris biomass was successfully extracted and the fatty acids methyl ester profile was proven to be suitable for making biodiesel. Ó 2012 Elsevier Ltd. All rights reserved.
1. Introduction Sustainable energy development has become one of the key challenges in this century. Depletion of fossil fuel in the near future and the effect of green house gases towards human and environment have escalated the search of renewable energies, such as solar, wind, hydro, wave and geothermal power [1]. However, these renewable energies are heavily dependent on regional or local condition that can be very unpredictable and inconsistent. On the other hand, biodiesel being recognized as a green and alternative renewable fuel has attracted great interest from researchers, governments, local and international traders. Some of the advantages of using biodiesel instead of fossil diesel are it is a non-toxic fuel, biodegradable and lower emission of green house gases when burned in diesel engine [2,3]. Normally, biodiesel is produced through transesterification reaction, in which vegetable oil reacts with short chain alcohol (e.g. methanol) in the presence of catalyst (e.g. sodium hydroxide, NaOH) [4,5]. Edible oils such as soybean, sunflower, rapeseed and palm oil are the common feedstock for biodiesel production. However, continuous growth of the biodiesel industries in the last ten years has raised a tragic social issue: the food versus fuel dispute [6]. It is strongly believed that further ⇑ Corresponding author. Tel.: +60 4 5996467; fax: +60 4 5941013. E-mail address: [email protected] (K.T. Lee). 0306-2619/$ - see front matter Ó 2012 Elsevier Ltd. All rights reserved. doi:10.1016/j.apenergy.2012.01.075
expansion of biodiesel industries in the global market will result to more undernourished people suffering from hunger and malnutrition [7,8]. Currently, microalgae have been identified as a superior feedstock for biodiesel production, mainly due to their fast growth rate (100 times faster than terrestrial plant) and able to double their biomass in less than one day under favourable culture conditions [9]. Apart from that, certain microalgae strains are able to accumulate sufficient amount of lipid that can be expected to overcome the terrestrial crops [10]. For example, microalgae with 30% of lipid content have the potential to produce 54 tonnes of oil/ha/year, whereas palm oil and jatropha are only able to produce 3.62 and 4.13 tonne of oil/ha/year, respectively [11,12]. Thus, cultivating microalgae for biodiesel production requires only a minimum land area and up hold an important key for a more sustainable land utilization [13]. Apart from that, an added advantage of cultivating microalgae is the ability of the microorganism to act as a carbon sink assimilating CO2 from atmosphere and flue gases through photosynthesis; a golden opportunity for carbon credit program [14–16]. However, one of the limitations to cultivate microalgae in industrial scale is the availability of nutrients sources. Chemical or inorganic fertilizers are commonly used to achieve promising growth rate of microalgae. Nevertheless, a life cycle assessment on microalgae cultivation has underlined that 50% of energy use
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and green house gases emissions are associated with chemical fertilizer production [17]. Meanwhile, utilizing secondary or tertiary wastewater as nutrients source to cultivate microalgae appears as a promising choice to reduce the overall energy input [18–21]. This is because wastewater contains high concentration of nitrate and phosphate that promote the growth of microalgae and at the same time, microalgae can act as a purifier to further remove the nutrients from wastewater before discharging to water sources. Nevertheless, the key challenges of using wastewater are susceptible contamination and inconsistency of nutrients compositions; these factors will directly retard the growth of microalgae. On the other hand, using organic fertilizer as a nutrient source offers an alternative and convenient way to minimize the contamination level in water. Organic fertilizer can be obtained through a series of composting and refining processes from waste materials such as manure, sewage sludge, biomass and food [22]. Although CO2 is released during composting process, however, the amount is lower compared to the production of inorganic fertilizer. It was estimated that 4–82 kg of CO2 and 4–67 kg of CO2 could be saved by composting one tonne of food waste and garden waste, respectively [23]. In addition, the emitted CO2 from composting process can be recycled as an alternative carbon source to cultivate microalgae. Normally, organic fertilizers available in the market are made in dry pellet form for easy transportation and storage. Hence, recycle and reuse the waste materials to cultivate microalgae for biodiesel production appears as a greener solution to drive the industry towards sustainable growth. Thus, the objective of the present study is to evaluate the potential of using organic fertilizer to cultivate Chlorella vulgaris. Various culture parameters such as amount of nitrates, illumination duration, pH, outdoor cultures and the effect of reuse the recycling water were systematically investigated. All experiments were performed using 5 L photobioreactor (except seed culture) instead of using lab scale conical flask and tap water was used rather than using distilled or sterilized water. This approach was implemented to promote an easier up-scaling and to minimize the overall energy consumption in cultivating microalgae in commercial scale.
paper (Double Rings 101). The resulting organic fertilizer medium was dark-brown in colour with typical characteristics as shown in Table 1. From the Table, the concentrated organic medium contains two types of nutrients which are equally important to support microalgae growth: (1) macronutrients, such as nitrogen, phosphorus, potassium, calcium and magnesium and (2) micronutrients, such as manganese, boron and iron. In addition, the prepared medium also contains exceptional high level of Chemical Oxygen Demand (COD) and Biochemical Oxygen Demand (BOD), in which those values are similar to untreated wastewater [24]. On the other hand, the filtered particulate solids (organic compost) can be further used as soil amendment to increase the water and nutrients holding capacity and to improve soil physical properties by providing better aeration [25]. Subsequently, a pre-determined volume of the organic fertilizer medium was introduced into a photobioreactor with 5 L of tap water (without sterilization) and the pH of the medium was adjusted according to pre-determined values. Then, C. vulgaris with initial cell concentration of 0.3 106 cells (around 10 mL from the seed culture) was introduced into the photobioreactor. The photobioreactor was aerated with compressed air continuously and illuminated with cool-white fluorescent light (Philip TL-D 36W/865, light intensity of 60–70 lmol m2 s1).
2. Material and methods
l ðday1 Þ ¼
2.1. Pure microalgae strain and culture conditions A wild-typeC. vulgaris was isolated from local freshwater located at Penang, Malaysia. The microalgae was preserved and grown in Bold’s Basal Medium (BBM), consisting of: (1) 10 mL per liter of culture medium with the following chemicals: NaNO3 (25 g/L), CaCl22H2O (2.5 g/L), MgSO47H2O (7.5 g/L), K2HPO4 (7.5 g/L), KH2PO4 (17.5 g/L), NaCl (2.5 g/L) and (2) 1 mL per liter of culture medium with the following chemicals: EDTA anhydrous (50 g/L), KOH (31 g/L), FeSO47H2O (4.98 g/L), H2SO4 (1 mL), H3BO3 (11.4 g/ L), ZnSO47H2O (8.82 g/L), MnCl24H2O (1.44), MoO3 (0.71 g/L), CuSO45H2O (1.57 g/L), Co(NO3)26H2O (0.49 g/L). The initial pH of the medium was adjusted to 6.8. The seed culture was grown in a 100 mL Erlenmeyer flask containing 50 mL of medium, aerated with compressed air, surrounding temperature of 25–28 °C and illuminated with cool-white fluorescent light (Philip TL-D 36W/ 865, light intensity of 60–70 lmol m2 s1) continuously. 2.2. Cultivating microalgae with organic fertilizer Baja Serbajadi Humus 27 (organic fertilizer or compost) with granular shape was purchased from a local market. A 10 g of the fertilizer was immersed in 600 mL tap water and stirred for 24 h using magnetic stirrer. Non-soluble particulate solids were observed after the stirring process and were filtered using filter
2.3. Measurement of microalgae growth A correlation between the optical density of C. vulgaris and biomass was pre-determined. Optical density was measured daily at 540 nm using spectrophotometer (Shimadzu UV mini-1240) [26]. Then, 10 mL of sample were centrifuged at 10 1000g for 5 min. The supernatant was slowly decanted back into the culture medium whereas the microalgae biomass were dried in an oven at 100 °C for 24 h. All the samplings were performed in triplicate to ensure the accuracy of the data. The correlation is shown below:
Dry weight ðg=LÞ ¼ 0:4541 OD540 ; R2 ¼ 0:9890
ð1Þ
The specific growth rate (l) was measured by using below:
lnðN2 =N1 Þ t2 t1
ð2Þ
where N1 and N2 are defined as biomass (g/L) at time t1 and t2, respectively. Table 1 Characteristics of organic fertilizer medium (obtained by mixing 10 g of organic fertilizer with 600 mL tap water). Parameter
Unit
Concentration
CODa BOD – 5 days test at 20 °Cb Nitrogenc Phosphorusd Potassiume Calciumf Magnesiumg Manganeseh Boroni Ironj
ppm ppm ppm ppm ppm ppm ppm ppm ppm ppm
1729.9 576.0 1323.2 213.6 634.4 269.9 54.5 1.0 4.1 1.3
Testing method: a APHA 5220 B (2005). b APHA 5210 B (2005). c APHA 4500-NH3F. d APHA 4500-P E (2005). e APHA 3111 B (2005). f APHA 3111 B (2005). g APHA 3111 B (2005). h APHA 3111 B (2005). i APHA 4500-B C (2005). j APHA 3111 B (2005).
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2.4. Measurement of nitrate content in medium Nitrate concentration was determined according to the Ultraviolet Spectrophotometric Screening Method [27]. A sample (1 mL) collected from photobioreactor was centrifuged at 10 1000 g for 5 min. The supernatant was collected and optical density was measured at 275 nm and 220 nm by using spectrophotometer (Shimadzu UV mini-1240). Then, the absorbance reading at 275 nm was subtracted two times from the reading at 220 nm to obtain the actual absorbance caused by the presence of nitrate. Dry potassium nitrate (KNO3) at different concentrations was used for calibration purposes. 2.5. Microalgae harvesting and biomass collection When C. vulgaris had grown to stationary phase, air aeration to the culture medium was stopped. The microalgae were let to settle naturally to the bottom of photobioreactor for two days. Two distinguished layers were observed, in which the upper layer consist of water with suspended microalgae cells and the bottom layer was microalgae biomass. The upper layer water was slowly decanted, leaving behind the microalgae biomass which was further dried in an oven at 80 °C for 24 h. The dried microalgae biomass were collected and sealed in an empty container for lipid extraction.
þ sources such as nitrate ðNO 3 Þ, ammonia ðNH4 Þ or urea (NH2–CO– NH2) appear to be the most significant contributor to sustain microalgae growth [30]. In the present study, synthesized nutrients from organic fertilizer were diluted to different concentration with the purpose to study the ability of C. vulgaris in up taking the nutrients. Fig. 1 depicts the growth of C. vulgaris with different nitrate concentration. From the figure, C. vulgaris was found to grow even under limited nutrients condition (5 and 10 mL of nutrients), however, the quantity of biomass produced were unsatisfactory: 0.1– 0.14 g/L after 12 days of cultivation. In comparison, C. vulgaris grew much better with sufficient supplement of nutrients (80 and 100 mL). The biomass produced were 0.29–0.31 g/L, which was nearly 150% of increment. Apart from that, specific growth rate of C. vulgaris was also improved due to supplement of ample nutrients. For example, the specific growth rate was 0.132 day1 and 0.229 day1 for 5 and 100 mL of nutrients added respectively; an increment of 73% was observed. This result indicates that large quantities of nutrient sources are the mandatory step to promote microalgae growth and consequently to attain higher biomass yield. On the other hand, the growth curve for C. vulgaris with excess amount of nutrients (150 mL) exhibited the same trend as 100 mL, in which no further improvement of biomass yield was observed. This is because other parameters such as photoperiod and pH control will become the growth controlling factors if adequate nutrients are supplied.
2.6. Microalgae lipid extraction
3.2. Effect of photoperiod
Ten grams of dried C. vulgaris biomass were placed in a cellulose thimble and extraction process was performed using Soxhlet extractor. Four types of solvent were used to compare the extraction efficiency, such as n-hexane, methanol, ethanol and mixed methanol–chloroform with volume ratio of 2:1 [28]. A total of 250 mL for each solvent was placed in Soxhlet extractor and heated at 60–65 °C for 24 h. After that, the solvent was evaporated in a rotary evaporator and the leftover lipid was collected. The residues (solid material) after evaporation were subjected to repeated extraction twice using the same solvent. Thereafter, crude microalgae lipids were measured gravimetrically. All samplings were performed in triplicate.
The effect of photoperiod on microalgae cultivation exhibited a significant influence towards microalgae photosynthetic activity and growth rates in a typical cultivation system [31,32]. However, it should be noted that microalgae with excessive of light exposure would result to unwanted electricity consumption (energy waste) and also inhibit the growth of microalgae [33]. Fig. 2 shows the effect of photoperiod towards the growth of C. vulgaris. From the figure, the growth of C. vulgaris was strongly affected by photoperiod, with the highest biomass yield attained at 0.31 g/L through continuous 24 h light illumination for 12 days (specific growth rate of 0.228 day1). In addition, for photoperiod of 3 h, 6 h and 9 h, the biomass attained were only 0.03–0.1 g/L which was exceptional unsatisfactory. This result indicates that longer photoperiod will result to continuous uptake of nutrients by C. vulgaris through photosynthesis and subsequently, increase their biomass at the same cultivation time. In other words, this result also indicates
Transesterification was performed using 1 g of crude C. vulgaris, methanol to lipid molar ratio of 15:1 and 3 wt.% of concentrated sulfuric acid (H2SO4) as catalyst. The reaction was carried out in a water bath shaker at 60 °C for 3 h. Upon completion of the reaction, 1 lL of the reaction product was subjected to gas chromatography-mass spectrometry (GC–MS; PerkinElmer Clarus 600) analysis. The GC was equipped with flame ionization detector (FID) and Elite 5-MS column (30 m 0.25 mm 0.25 mm). The initial oven temperature was 65 °C, held for 2 min and raised to 280 °C at ramping rate of 8 °C/min and held at 280 °C for 10 min, while the injector temperature was set to 250 °C. The compounds detected were identified and quantified using NIST Mass Spectral Search Program.
50 mL 80 mL
5 mL 10 mL
0.35
20 mL 30 mL
100 mL 150 mL
0.3
Biomass (g /L)
2.7. Transesterification reaction and fatty acid methyl esters (FAME) analysis
0.25 0.2 0.15 0.1 0.05
3. Result and discussion
0 0
3.1. Effect of nutrients concentration Sufficient supplement of nutrients for microalgae to grow is the first key step to produce bulk quantity of microalgae biomass [29]. Among all the nutrients required, adequate supply of nitrogen
2
4
6
8
10
12
Cultivation time (days) Fig. 1. Effect of initial nitrate concentration towards the growth of C. vulgaris. Other culture conditions: pH = 7 and illuminated for 24 h continuously. Initial nitrate content (mg/L): 5 mL = 1.33, 10 mL = 2.69, 20 mL = 5.38, 30 mL = 7.98, 50 mL = 13.09, 80 mL = 20.94, 100 mL = 26.67, 150 mL = 40.71.
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0.4
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9h
6h
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Biomas (g /L)
Biomass (g /L)
0.35
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0.05 0 0
0 0
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Cultivation time (days)
that C. vulgaris is weak in storing substantial amount of light energy for later use in the dark period (exergonic reactions) [32]. Hence, surface area of illumination should be taken into consideration when designing a photobioreactor to cultivate C. vulgaris (typically outdoor cultivation), so that optimum light energy can be efficiently utilized by the microalgae. From Fig. 2, it was clear to observe that the quantity of biomass attained at 12 h of photoperiod time (equivalent to outdoor sunlight cycle) were nearly double the quantity of biomass attained at photoperiod of 3 h, 6 h and 9 h. This result proved that it is feasible to cultivate C. vulgaris under outdoor conditions; however, it is still dependent on the local weather condition and temperature. 3.3. Effect of initial pH One of the controlling factors to cultivate microalgae is the pH of the culture medium [34]. Generally, pH of 7–8 is plausible for microalgae to grow [34]. Nevertheless, it is worth to mention that suitable microalgae strains for biodiesel production should not only contain high lipid yield, but should be able to withstand pH changes during cultivation cycle. For example, high pH value (9.0–9.5) was observed in photobioreactor due to shortage of inorganic carbon after rapid consumption by microalgae [35]. Sometimes, pH can also drop drastically if high concentration of CO2 is continuously supplied to the culture medium and caused adverse effect towards the growth of microalgae [19]. More importantly, in the near future, when flue gases are used to cultivate microalgae in industrial scale, the selected microalgae strains should be able to tolerate inconsistent concentration of CO2 in the flue gases that indirectly resulted to pH variation in the culture medium [36]. Fig. 3 shows the effect of initial pH in the culture medium for C. vulgaris. From the figure, C. vulgaris exhibited almost a linear growth at pH of 4, 5 and 8 with maximum biomass of 0.47–0.51 g/L obtained after 12 days of cultivation. Furthermore, higher specific growth rate was attained at pH 4 (0.265 day1), 5 (0.270 day1) and 8 (0.263 day1) compared to culture growth at neutral condition, pH 7 (0.229 day1). This result indicates that C. vulgaris can adapt very well even at low or high pH. One of the advantages of cultivating microalgae at low or high pH is the simultaneous elimination of contaminants such as fungus and therefore, able to sustain their growth naturally. Nevertheless, at pH 3, the growth of C. vulgaris was reverted and no significant increment of biomass was detected after 12 days of cultivation. On the other hand, although the growth of C. vulgaris was satisfactory at pH 6, 7 and 9, a lag phase was observed from day 1 to day 4 and the growth was almost stagnant from day 9 to day 12 with a maximum 0.33 g/L of biomass obtained.
8
10
12
Fig. 3. Effect of initial pH towards the growth of C. vulgaris. Other culture conditions: Initial nutrients volume = 100 mL (nitrate content of 26.67 mg/L), illuminated for 24 h continuously.
3.4. Comparison of the growth of C. vulgaris with organic and inorganic fertilizer under indoor and outdoor conditions Up to date, research on microalgae is focussed on indoor culture where a control environment is relatively easy to achieve. Nevertheless, for the purpose of cultivating microalgae in a commercial scale, outdoor cultivation system is the most feasible option due to easy access to sunlight [37]. This strategy will help to reduce the overall energy input (mainly for illumination and temperature control) and consequently, enhance the cost effectiveness to produce biodiesel from microalgae. However, one of the limitations to cultivate microalgae under outdoor conditions is the inconsistent changes of local weather, temperature and light intensity. Therefore, microalgae strains that are able to adapt and grow well under these extreme cultivation conditions will be an added advantage. Fig. 4 shows a comparison of the growth of C. vulgaris (supplied with organic or inorganic nutrients) under indoor and outdoor conditions. From the figure, C. vulgaris was found to grow much faster with inorganic nutrients compared to organic nutrients, especially when cultivated under indoor conditions with specific growth rate of 0.30 day1 and 0.27 day1, respectively. The
0.8
Indoor (organic)
Indoor (inorganic)
Outdoor (organic)
Outdoor (inorganic)
0.7 0.6
Biomass (g /L)
Fig. 2. Effect of photoperiod (hours of light) towards the growth of C. vulgaris. Other culture conditions: initial nutrients volume = 100 mL (nitrate content of 26.67 mg/ L) and pH = 7.
6
Cultivation time (day)
0.5 0.4 0.3 0.2 0.1 0 0
2
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14
16
Cultivation time (day) Fig. 4. Comparison of indoor and outdoor culture towards the growth of C. vulgaris. Culture conditions: (a) organic nutrients; initial nutrients volume = 100 mL (nitrate content of 26.67 mg/L) and pH = 5 and (b) inorganic nutrients; BBM medium and pH = 5. Photoperiods: (a) indoor: 24 h illumination and (b) outdoor: 12 h light and 12 h dark.
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result is rather expected due to controlled conditions and ample macro- and micro-nutrients present in the inorganic nutrients. Apart from that, by referring to Fig. 4, C. vulgaris was observed to grow slower under outdoor environment, either supplied with organic or inorganic nutrients. Under uncontrolled environment, the temperature at outdoor varied from minimum 23 °C to maximum 35 °C. In addition, a lag phase was detected from day 1 to day 3 of culture, showing that the microalgae required some time to adapt themselves to the new environment. As a consequence, the specific growth rate of microalgae cultivated under outdoor conditions was dropped to 0.20 day1 for both organic and inorganic nutrients. This observation indicates that outdoor conditions will become the key controlling factor towards microalgae growth when sufficient nutrients supplement is available. In this regards, using organic fertilizers to cultivate microalgae under uncontrolled environment will be an added advantage since the final biomass attained was nearly the same with the inorganic nutrients. Apart from that, it was found that the C. vulgaris cultivated with organic nutrients under outdoor conditions attained higher biomass yield (0.37 g/L) on day 12 compared to indoor culture with 12 h illumination (0.21 g/L). This may be due to the intensity of sunlight that is much more suitable to cultivate C. vulgaris compared to illumination by fluorescent lamp.
3.5. Effect of using recycling water Effective uses of water resources play a significant role in microalgae cultivation since microalgae are aquatic microorganisms that need water to survive [38]. In large scale microalgae cultivation system, huge amount of water is required to reach optimum productivity of microalgae biomass. Therefore, to recycle and reuse the water not only can save the overall operation cost, but also helps to preserve the water for other purposes [39]. However, not all microalgae strains are able to re-grow effectively in recycled water due to susceptible contamination by fungus and bacteria (e.g. open pond system). Hence, the selected microalgae strains for biodiesel production should be able to grow rapidly and withstand other contaminants. Fig. 5 shows the effect of using recycled water from the previous batch culture (indoor and outdoor, respectively) without any pre-treatment or purification process. The recycled water still contained minimum amount of nutrients and microalgae cells which were not completely harvested. From the figure, the growth of C. vulgaris under indoor conditions was not affected by the use of recycled water and produced satisfactory
Indoor
Outdoor
0.6
3.6. Lipid extraction and biodiesel production from C. vulgaris Up to now, lipid extraction using chemical solvent is the most reliable method to determine the overall lipid content presence in microalgae. Extraction efficiency by chemical solvent is higher than physical method (mechanical pressing machine), typically due to the high polarity of fatty acids toward the chemical solvent. Nevertheless, the effectiveness of different type of chemical solvent used is strongly dependent on microalgae strains, especially when the existence of cell wall in microalgae will indirectly impede chemical solvent extraction efficiency [40]. In the present study, four different types of solvent were used for lipid extraction study: n-hexane, methanol, ethanol and methanol mixed with chloroform with volume ratio of 2:1 (Bligh and Dyer method). Bligh and Dyer method gave the highest extraction efficiency, in which 18.1% of lipid was successfully extracted. This value is comparable to the total lipid of C. vulgaris reported in the literature, which lies between 5% and 40% under phototrophic culture condition [24]. The solvent extraction efficiency in descending order was methanol (15.5%), ethanol (10.7%) and n-hexane (3.2%). Although n-hexane has been widely used in oil extraction study, nevertheless, it is not a suitable choice to extract lipid from C. vulgaris. This is because n-hexane is a non-polar solvent which has poor permeability in cell wall and hence only extracellular lipids are extracted [41]. The fatty acids methyl ester compositions of C. vulgaris mainly consisted of C16:0, C16:1, C16:2, C18:1, C18:2 and C18:3 as shown in Fig. 6. From the figure, C16:0 (palmitic acid methyl ester), C18:1 (oleic acid methyl ester) and C18:2 (linoleic acid methyl ester) represent the major portion of fatty acid methyl esters compositions, accounted for total of 85.6%. These fatty acids are naturally found in oil bearing crops, such as soybean, sunflower, cottonseed and palm oil, in which the fatty acids are suitable for biodiesel production. Apart from that, unsaturated fatty acids methyl esters (C16:1, C16:2, C18:1, C18:2, C18:3) were predominant in the fatty acids profile, accounted for 74%. It is important to note that higher compositions of unsaturated fatty acids methyl ester can reduce the pour point of biodiesel and making it feasible to be used in cold climate countries [42]. Other saturated fatty acids methyl ester
0.4 0.3 0.2 0.1 0 0
2
4
6
8
10
12
14
16
Cultivation time (day) Fig. 5. Effect of reuse the recycling water towards the growth of C. vulgaris. Culture conditions: Initial nutrients volume = 100 mL (nitrate content of 26.67 mg/L) and pH = 5. Photoperiods: (a) indoor: 24 h illumination and (b) outdoor: 12 h light and 12 h dark.
Fatty acids methyl ester concentration
Biomass (g /mL)
0.5
quantity of biomass within 16 days of cultivation time. Apart from that, for C. vulgaris cultivated at outdoor using recycled water, the lag phase from day 1 to day 3 was greatly minimized if compared with the use of fresh water (Fig. 4). This is because C. vulgaris cells that were not harvested and remain in the recycled water have already adapted to the outdoor environment. Therefore, once the new organic nutrients were supplied to the culture medium, the microalgae can utilize the nutrients immediately. Consequently, higher biomass quantity (0.55 g/L) was obtained compared to the use of fresh water under outdoor conditions (0.37 g/L).
C16:0
C16:1
C16:2
C18:1
C18:2
C18:3
Fatty acids methyl ester profile Fig. 6. Fatty acids methyl ester profile of C. vulgaris.
Others
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except C16:0 (palmitic acid methyl ester) were also identified, such as C14:0 (myristic acid methyl ester), C15:0 (pentadecanoic acid methyl ester) and C18:0 (stearic acid methyl ester). However, the compositions were minimum and they accounted for only 1.1% of the overall lipid compositions. 3.7. Estimation of nutrients cost in cultivating C. vulgaris Based on the optimum cultivating conditions of C. vulgaris using organic and inorganic fertilizer, a simple cost analysis was conducted. The cost of organic fertilizer purchased at local market was 1.2 USD per pack (400 g) whereas the overall cost to prepare the inorganic fertilizer (BBM) was 15–25 USD per liter. In addition, for the preparation of organic nutrients, electricity cost for stirring and cost for solid residue filtration were included. Consequently, the total cost of nutrients required to produce 1 kg of C. vulgaris biomass was estimated to be 2.5–3 USD and 60–85 USD for organic and inorganic fertilizer, respectively. Hence, utilization of cheap nutrient sources such as organic fertilizer will be more beneficial in large scale microalgae cultivation system, typically in terms of cost saving and better environmental protection. 4. Conclusions The result in this study showed that it is feasible to use organic fertilizer to cultivate C. vulgaris for biodiesel production. This approach offers several advantages in terms of environmental perspective and cost effectiveness. In addition, this method can be also easily adopted since organic fertilizers are widely available in market. More importantly, C. vulgaris was found to be able to reproduce when grown under outdoor conditions (uncontrolled environment). Besides, reutilize the water to re-cultivate C. vulgaris is possible and this will further drive the microalgae cultivation system towards a more sustainable process. Acknowledgements The authors would like to acknowledge the funding given by Universiti Sains Malaysia (Research University Grant No. 814146, Postgraduate Research Grant Scheme No. 8044031 and USM Vice-Chancellor’s Award) for this project. References [1] Leung DYC, Wu X, Leung MKH. A review on biodiesel production using catalyzed transesterification. Appl Energy 2010;87:1083–95. [2] Demirbas A. Progress and recent trends in biodiesel fuels. Energy Convers Manage 2009;50:14–34. [3] Qiu F, Li Y, Yang D, Li X, Sun P. Biodiesel production from mixed soybean oil and rapeseed oil. Appl Energy 2011;88:2050–5. [4] Vasudevan PT, Briggs M. Biodiesel production – current state of the art and challenges. J Ind Microbiol Biotechnol 2008;35:421–30. [5] Lam MK, Lee KT, Mohamed AR. Homogeneous, heterogeneous and enzymatic catalysis for transesterification of high free fatty acid oil (waste cooking oil) to biodiesel: a review. Biotechnol Adv 2010;28:500–18. [6] Wu X, Leung DYC. Optimization of biodiesel production from camelina oil using orthogonal experiment. Appl Energy 2011;88:3615–24. [7] Srinivasan S. The food versus fuel debate: a nuanced view of incentive structures. Renew Energy 2009;34:950–4. [8] Amaro HM, Guedes AC, Malcata FX. Advances and perspectives in using microalgae to produce biodiesel. Appl Energy 2011;88:3402–10. [9] Tredici MR. Photobiology of microalgae mass cultures: understanding the tools for the next green revolution. Biofuels 2010;1:143–62. [10] Rasoul-Amini S, Montazeri-Najafabady N, Mobasher MA, Hoseini-Alhashemi S, Ghasemi Y. Chlorella sp.: a new strain with highly saturated fatty acids for biodiesel production in bubble-column photobioreactor. Appl Energy 2011;88:3354–6. [11] Chisti Y. Biodiesel from microalgae. Biotechnol Adv 2007;25:294–306. [12] Lam MK, Lee KT. Renewable and sustainable bioenergies production from palm oil mill effluent (POME): Win–win strategies toward better environmental protection. Biotechnol Adv 2010;29:124–41.
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