Effective extraction of microalgae lipids from wet

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20 Mar 2014 - Producing biodiesel from lipid extracted from microalgae is a promising approach for sustainable ... In the second stage, the lipid content was enhanced by six-fold after three weeks ... demand and inevitable depletion of fossil fuel reserves [1,2]. ... drying is the most commonly used method, as it directly uti-.
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Effective extraction of microalgae lipids from wet biomass for biodiesel production Hanifa Taher a, Sulaiman Al-Zuhair a,*, Ali H. Al-Marzouqi a, Yousef Haik b, Mohammed Farid c a

Chemical and Petroleum Engineering Department, UAE University, Al-Ain, United Arab Emirates Mechanical Engineering Department, UAE University, Al-Ain, United Arab Emirates c Chemical and Materials Engineering Department, University of Auckland, New Zealand b

article info

abstract

Article history:

Producing biodiesel from lipid extracted from microalgae is a promising approach for

Received 23 September 2013

sustainable fuel production. However, this approach is not yet commercialized due to the

Received in revised form

high costs of upstream processes that are associated with the time consuming and/or

28 January 2014

energy intensive drying, and lipid extraction processes. In this study, the possibility of

Accepted 28 February 2014

avoiding the drying process, and extracting the lipid directly from the wet concentrated

Available online 20 March 2014

cells, using enzymatic disruption to enhance the extraction, has been tested. Results showed that lysozyme and cellulase were both efficient in disrupting cell walls and

Keywords:

enhancing lipid extraction from wet samples, with highest lipid extraction yield of 16.6%

Enzymes

achieved using lysozyme. The applicability of using supercritical CO2 (SC-CO2) in extracting

Cell disruption

lipid from wet biomass was also tested and the highest yield of 12.5% was achieved using

Wet microalgae

lysozyme. In addition, a two-step culturing process was applied, using Scenedesmus sp., to

Separation

combine both high biomass growth and lipid content. The strain was able to increase its

Nitrogen starvation

biomass productivity in the first stage, reaching 174 mg l1 d1, with almost constant lipid

Biodiesel

content. In the second stage, the lipid content was enhanced by six-fold after three weeks of nitrogen starvation, but with lower biomass productivity. ª 2014 Elsevier Ltd. All rights reserved.

1.

Introduction

Biodiesel, a series of mono-alkyl fatty acid esters, has received an increasing attention, especially with the increasing energy demand and inevitable depletion of fossil fuel reserves [1,2]. Conventionally, biodiesel is produced from oils extracted from oil crops. This is highly controversial and competes with their use as a food stock. In addition, the oil crops require large arable land development, fertilization, and fresh water

* Corresponding author. Tel.: þ971 35319. E-mail address: [email protected] (S. Al-Zuhair). http://dx.doi.org/10.1016/j.biombioe.2014.02.034 0961-9534/ª 2014 Elsevier Ltd. All rights reserved.

irrigation. These significant drawbacks limit further industrialization and urge to find new feedstocks. Waste cooking oils and animal fats have been suggested as alternatives; however the supply of such feedstocks is not consistent and cannot satisfy the large demand for biodiesel production. In addition, poor cold flow properties (cloud and pour points) of oils and saturated fatty acid contents of animal fats reduce the quality of produced biodiesel [3]. On the other hand, oil extracted from microalgae has recently emerged as a potential alternative source due to the microalgae high growth rate, high

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lipid content, and ability to grow in seawater. Furthermore, they do not require development of agriculture lands. The lipid productivity of microalgae is reported to be ten times higher than that of the best oil crop, which makes it a promising alternative [1,3]. In spite of their obvious advantages over oil crops, microalgae-biodiesel production processes are not yet commercialized. Generally, microalgae can produce both neutral lipids, composed mainly of triglycerides, and polar lipids such as phospholipids, which commonly produced in cellular membrane, whereas the former usually accumulated as droplets in the cytoplasm [4]. The biodiesel production process from microalgae consists of biomass cultivation, harvesting, drying, and lipid extraction, followed by the conversion of extracted lipids to biodiesel, and finally purification of the produced biodiesel. Among the main challenges in the biodiesel production from microalgae are separating the cultivated biomass from the growth media and extracting lipids from the harvested biomass. The biomass concentration of Scenedesmus is reported to be low in outdoor cultures, reaching 0.02e0.12 g l1 [5] and increases to about 0.3e3.4 g l1 [6] in photobioreactors. The currently used harvesting methods are centrifugation, sedimentation and flocculation. These methods have been proven to be effective but are costly, representing 20e35% of total biomass production cost [7]. On the other hand, a drying step is usually required prior to lipid extraction from the harvested microalgae cells. Sun drying is the most commonly used method, as it directly utilizes the solar energy. However, it is a time consuming process and the drying rate remains the main challenge of such process. Other faster processes are energy intensive and/or can alter the lipid structure and the protein rich leftover biomass, which affects their quality. It has been reported that the drying step accounts for 89% of the required energy input [8], and 70e75% of total processing cost [9]. Thus, it is considered as the major bottleneck of algae based biodiesel production [10]. Therefore, positive net energy from microalgae biodiesel could be obtained if wet extraction is used. It would be economically favorable to avoid the drying step while maintaining an effective lipid extraction from the wet biomass. Indeed, it has been estimated that the energy required to produce 1 kg of biodiesel from dried biomass is 4000 times more than that from wet biomass [11]. This requires expensive and high energy consumption in both the up- and downstream processes in order to get high quality biodiesel. It is vital therefore to develop a cost and energy effective processes, which is green and environmental friendly, which can overcome the main technical and economical barriers of the conventional techniques. In addition, for microalgae utilization process to be feasible, the extraction process should not negatively affect the de-lipidated leftover biomass to allow it to be used in useful applications, or its conversion to a valuable product. Several extraction methods can be used to extract the lipid from dried microalgae biomass. Among the common used techniques are Folch [12], Bligh and Dyer [13], and Soxhlet [14]. However, these techniques are complicated and time consuming, requiring about 1e3 days for completion. In addition, an extra solvent evaporation unit is required to

separate the extracted lipid from the solvent. Furthermore, contamination of the de-lipidated with traces of solvent is inevitable, which limits their further uses. Recently, supercritical CO2 (SC-CO2) received considerable attention due to its tunable solvation power, mild operating conditions, and environmental benign features. In addition, the process is fast, and extraction that requires 24 h using conventional techniques is completed with 30e60 min. This technique also does not require solvent separation, as CO2 is separated from the extracted lipid by simple depressurization. Furthermore, there is no contamination of the leftover biomass with the extraction solvent. On the other hand, the rigid and tough cell walls of microalgae cells hinder the extraction of the cells lipids. These cell walls are mainly composed of 24e74% neutral sugars, 1e24% uronic acids, 2e16% protein, and 0e15% glucosamine [15]. In addition, the presence of water in wet harvested biomass forms a film preventing the solvent from reaching the lipid, which further prevents efficient lipid extraction. Thus, for an efficient extraction of the lipid, the microalgae cells have to be disrupted to liberate the lipid and allow them to come into contact with the solvent. This is conventionally done using wet milling [16], ultrasonication [17,18], beadbeating [14], microwaves [18], autoclaving at a high temperature and pressure, and osmotic disruption by treatment with sodium chloride [19]. All these processes however, are energy intensive and may affect the properties of the triglycerides causing downstream difficulties in their transesterification to biodiesel. In addition, mechanical disruption usually results in excessive heat generation, which requires cooling. Another technique, which is commonly used is acid treatment [20]. Usually, this is performed by soaking the biomass in diluted acid, commonly sulfuric acid, and then heating it at high temperature for a certain time. In many cases, this is followed by alkaline addition, commonly sodium hydroxide, thus leading to pores swelling and decreasing in cellulose crystallinity. Although this process is efficient in lignocellulose degradation, sulfuric acid is toxic and corrosive, which makes this process not recommended, neither for fuel production nor for the left biomass applications. Freeze drying has also been suggested [18], wherein harvested cells are lyophilized, resulting in dried powder, prior to lipid extraction. Lyophilization, however, is an energy intensive process that is not justified in energy production processes. For an-economical and effective lipid production, efficient cell disruption, mild extraction conditions, and leftover biomass utilization are essential. Some enzymes have the potential to facilitate the cell disruption. They can operate at low temperatures with high selectivity and fewer side products. Utilizing enzymes for cell disruption is expected to enhance the efficiency of lipid extraction at mild conditions. Operation at mild conditions is less energy intensive and also has a minimum effect on the triglycerides structure or the leftover residual biomass that can be utilized in pharmaceutical, food and fuel applications. Several lysis enzymes can be used, such as cellulase that can effectively hydrolyze the cellulosic structure of the cell walls, and lysozyme that can hydrolyze the linkage between peptidoglycan residues in the cell walls. Specifically, it degrades polymers containing N-acetylglucosamine, which is a

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derivative of glucosamine, major component of cell wall. The enzymes have been tested on Chlorella species and outer cell wall disruption was observed [21e24]. In this work, the two enzymes were tested to disrupt cell wall, and liberate the lipid, which would enhance the extraction process. To the best of our knowledge, enzymatic disruption combined with SC-CO2 extraction has not been reported in literature. The use of such process is less time consuming than conventional solvent extraction processes, avoids the use of toxic chemicals, and does not require solvent separation unit. In addition, the process allows the utilization of the protein rich leftover biomass. In addition to the extraction challenges, the lipid content of the microalgae cells is important. The overall effectiveness of biodiesel production from microalgae depends on the lipid productivity, which is a combined effect of biomass productivity and lipid content. The reported lipid content of microalgae is in the range of 20e30% (dry basis) when the microalgae are grown under controlled conditions with sufficient nutrients. However, under stress conditions, microalgae may accumulate larger lipid content [25e28]. The primary stress applied to green microalgae is nitrogen deficiency, where accumulations of more than 50% (dry basis) have been reported [1,29]. This is mainly due to the lack of nitrogen required for protein synthesis, and the excess carbon from photosynthesis is then diverted into lipid production pathway [30]. The objective of this study is to assess the effectiveness of using enzymes for cell disruption and compare it conventional methods. In addition, investigate the possibility of extracting lipids from wet biomass using SC-CO2, avoiding the high organic solvent consumption, the time consuming drying step and the energy intensive mechanical cell disruption. In addition, lipid enhancement by nitrogen starvation, combined with efficient biomass production is also studied to confirm Scenedesmus sp. applicability for biodiesel production. The success of this work adds great enhancement to the extraction process, which in turn has a positive effect on the overall algae biodiesel production process.

2.

Materials and methods

2.1.

Chemicals and enzymes

n-Hexane, methanol, acetone, sulfuric acid, sodium hydroxide, HEPES (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid), 14% boron tetraflouridemethanol mixture, Nile Red (9diethylamino-5-benzo[a] phenoxazinone), dimethyl sulfoxide (DMSO), were purchased from SigmaeAldrich Inc., USA. Liquefied CO2 with a purity of 99.95% was supplied by Abu-Dhabi Oxygen Company, UAE. Ultra high purity helium and zero-air were supplied by Air Product Company, UAE. Lysozyme from chicken egg white (activity > 40,000 U mg1, according to the supplier’s definition) and cellulase from Trichoderma longibrachiatum (activity  1.0 U mg1, according to the supplier’s definition) were purchased from SigmaeAldrich Inc., USA, and stored below 8  C and above 0  C according to the supplier’s instructions, respectively. Standard of high purity fatty acid methyl esters (FAMEs), used for gas chromatography (GC)

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calibration and fatty acid compositional analysis, comprised of; 3.9% myristic acid methyl ester (C14:0), 9.9% palmitic acid methyl ester (C16:0), 6.0% stearic acid methyl ester (C18:0), 10% elaidic acid methyl ester (C18:1), 24.8% cis-9-oleic methyl ester (C18:1), 36.1% linoleic acid methyl ester (C18:2n6c), 1.9% linolelaidic acid methyl ester (C18:2), 2.1% arachidic acid methyl ester (C20:0), and 2.1% behenic acid methyl ester (C22:0) were obtained from Sigma Aldrich, USA.

2.2.

Strains and culture medium

Scenedesmus sp. culture was obtained from Algal Oil Company, Philippines, and cultivated in a modified bassel medium (þ3NBBM) for two weeks, followed by three weeks of stressing in nitrogen-deprivation medium (N-BBM). The modified medium, þ3N-BBM, comprised of (in mM); 8.82 NaNO3, 0.17 CaCl2$2H2O, 0.3 MgSO4$7H2O, 1.29 KH2PO4, 0.43 K2HPO4, 0.43 NaCl, 1 ml l1 of Vitamine B12, and 6 ml l1 of P-IV solution that consisted of 2 Na2EDTA$2H2O, 0.36 FeCl3$6H2O, 0.21 MnCl2$4H2O, 0.37 ZnCl2, 0.0084 CoCl2$6H2O, and 0.017 Na2MoO4$2H2O. In N-BBM, the nitrate source (NaNO3) was removed.

2.3.

Growth experiments

For an economical biodiesel production from microalgae, high biomass productivity and lipid content are important. These two factors are difficult to achieve simultaneously, as conditions favoring high biomass productivity usually result in low lipid accumulation, and vice versa. To overcome this, a two stage cultivation approach has been used, wherein the first stage, the cells were allowed to grow in a nutrient rich medium (þ3N-BBM) for two weeks to enhance the biomass productivity, and in the second stage cells in their exponential growth phase were transferred to a nitrogen-deficient medium (N-BBM) for three weeks to enhance the lipid accumulation. The cultivations in both stages were done in a 5 l bubble column photobioreactor with an internal illumination. All cultivations in this work were autotrophic, with CO2 naturally present in air bubbled through the system being the sole carbon source. Prepared media were sterilized in autoclave (Hirayama HV50, Japan) at 121  C for 15 min and cooled to room temperature prior to use. The microalgae was first grown in 200 ml of þ3NBBM placed in 500 ml Erlenmeyer flasks with filtered air bubbling at a constant temperature of 25  1  C under light intensity of 75 mmol m2 s1 determined using a light meter (model 472990, Extech Instruments, Massachusetts) with an initial cell concentration of 3.2  104 cells ml1. Details about cell concentration determination are mentioned later. After 11 days of growth, the culture was transferred to the photobioreactor, where the growth was monitored with time. The photobioreactor was illuminated with one 50 cm, 60 W, white fluorescent light at a light intensity of 120 mmol m2 s1, measured using the light meter under 12 h light/dark photoperiod automatically controlled by 24 h timer (S2402, China). The photobioreactor had an outer diameter of 10 cm, an inner diameter of 5 cm and 40 cm height. The cell growth was monitored daily by measuring the optical density at 680 nm using a spectrophotometer

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(Shimadzu UVe1800 UV, Kyoto, Japan). The samples were measured twice and the average values were considered. The cell concentration (cells ml1) at any given cultivation time was calculated from a pre-prepared calibration curve of the optical density at 680 nm versus cell concentration determined using Neubauer Hemocytometer, placed on a microscope (Eclipse LV100 Pol, Nikon, Japan). The dry weight of algal biomass was also determined by filtering the algal suspension using a, prewashed and dried Whatman filter paper, dried overnight at 105  C in an oven (Memmert, Germany) until constant weight. On day 14, while cells were still in their exponential growth phase, they were allowed to settle at the bottom of the photobioreactor, and were then concentrated by centrifugation at 3000 r min1 for 15 min using multispeed centrifuge (IEC CL31, Thermo Scientific, USA), and washed twice with distilled water. Concentrated microalgae cells, after centrifugation, were cultivated back in a similar photobioreactor in N-BBM medium to enhance the lipid accumulation. Samples were collected at regular intervals, centrifuged and re-diluted in distilled water to obtain a 4 ml of cell suspension. For lipid testing, the accumulations were monitored by staining the constant concentration samples (1.5  106 cells ml1) with Nile Red that emits a yellow fluorescent signal in the presence of the lipid, and the fluorescents were visualized using fluorescence microscope (Olympus). The Nile Red stock solution was prepared as described by Siaut et al. [31], by dissolving 0.1 mg of Nile Red in 1 ml acetone, and the solution was stored in the dark at 4  C. Culture samples (500 ml) were placed in an eppendorf tube, span in a centrifuge (Sigma 113, Germany) for 30 s at 4000 r min1 and 410 ml of the supernatant were taken. DMSO (10 ml) was then added to promote the accessibility of Nile Red into the cells. The culture was then vortexed and 1 ml of Nile Red solution was added followed by 20 min incubation in the dark. The lipid accumulations were then quantified using Multi-label Plate Reader (PerkineElmer, Boston) with black 96-well plates. Fluorescences were measured before and after Nile Red staining and for Nile Red stained N-BBM medium. The intensity was considered after subtracting the stained medium and sample before staining intensities from the stained sample intensity at excitation and emission wavelengths of 485 and 590 nm, respectively.

2.4.

Cells disruptions

Wet cell biomass samples (1 g containing 6.8% solids) were subjected to three different disruption methods, namely: (1) lyophilization, (ii) enzymatic disruption and (iii) acid treatment. The lyophilization was carried out in a freeze drier (Telstar, Terrassa, Spain) operated at 54  C and 0.02 mbar for 6 h. The enzymatic pre-treatments were carried as follow; 3.25 ml of 10 mg ml1 of enzymatic (lysozyme or cellulase) solution was added to the wet biomass in the presence of 7.5 ml HEPES buffer solution (pH ¼ 7.48). This corresponds to lysozyme and cellulase loadings of 1.92  104 and 0.48 U mg1 dry biomass, respectively, with 6.78 g l1 biomass loading. The mixtures were then incubated in SI-300 Shaker at 37  C and 100 r min1 for 30 min. The acid treatment was done by adding 1 ml of sulfuric acid (1 M) to the wet biomass and heated to 90  C for 30 min in shaking water bath (LabTech, Daihan

LabTech Co., Ltd., Korea), followed by addition of 1 ml sodium hydroxide (5 M) solution and further incubation at 90  C for 30 min [20].

2.5.

Lipid extraction

Microalgae lipids were extracted using two extraction solvents, namely n-hexane and SC-CO2. In the static n-hexane extraction, where no continuous flow or circulation of the extraction solvent was employed compared to Soxhlet, 30 ml of n-hexane was added to 1 g (containing 6.8% solids) of the disrupted biomass (wet basis), incubated in an SI-300 Benchtop Shaker at 50  C and 100 r min1 overnight. The total lipid content of the biomass was determined from lyophilized biomass, as a base line, by Soxhlet extraction using n-hexane for 8 h. The wet samples had a water content of 93.2% determined by drying a pre-weighed sample on a pre-dried Whatman filter paper and drying overnight at 60  C until a constant weight was reached. For the enzymatic and acidic treated samples, n-hexane solvent was added directly to the treated sample, without removing the treating solutions. The mixture was then centrifuged at 3000 r min1 for 5 min. The upper nhexane layer containing the lipid was collected and the lipid content was determined gravimetrically after solvent evaporation. Same approach was carried out with lyophilized samples, where lyophilized cells were extracted directly without any additional cell disruption step, as lyophilization can simultaneously dry and disrupt the cell. The extraction was performed in a duplicate and average values were considered. The SC-CO2 extractions were conducted at a pressure of 500 bar, temperature of 50  C and 3 ml min1 flow of solvent in ISCO supercritical extraction unit (SFX-220, USA), which are the optimal operating conditions determined in our previous study [32]. The high pressure used was to ensure highest lipid extraction. Unlike in the static n-hexane extraction, in the SCCO2 extraction, separating the wet biomass from the enzyme solution, prior to extraction, was required. After incubation in the enzymatic solution for the desired time, the sample was centrifuged at 3000 r min1 for 5 min. The supernatant was removed and residual treated biomass was collected. The disrupted wet biomass was placed in 10 ml extraction cell with glass wool placed in both extraction vessel sides. CO2 was pressurized to the operation pressure using a high-pressure syringe pump (Model 260D, ISCO, USA), and the pressurized CO2 was then heated to the operation temperature and pumped into the extraction vessel. The extract was collected in a vial after depressurization via micro-metering valve (HIP 15-12AF1-V), and lipid content was determined gravimetrically after solvent evaporation. Further details about SC-CO2 extraction procedure were reported elsewhere [32,33].

2.6.

Fatty acid methyl ester (FAME) profile

To quantify the fatty acids present in the extracted lipid, which affects the properties of the produced FAMEs and their applicability to replace petroleum fuel, lipids were esterified to FAME using 14% boron tetraflourideemethanol mixture as described by Rule [34]. FAME were identified and quantified by GC (Varian, CP-3800, USA), fitted with CP-Sil 88 FAME capillary

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column (100 m  0.25 mm  0.2 mm, Varian, USA), flame ionization detector (FID) and equipped with auto-injector (CP 8410. Varian, USA). The oven initial temperature was held at 150  C for 1 min and then increased to 250  C at 4  C min1. Helium and zero air were used as the carrier gases with a split ratio of 40:1. Both the injector and the detector temperatures were set at 260  C.

2.7.

Statistical analysis

Experiments were conducted with at least duplicate treatments. The data were analyzed using one-way (unstacked) analysis of variance (ANOVA) followed by Fisher’s Least Significant Differences (LSD). p-Value 5% was considered as significant, and data were presented as mean  standard deviation.

3.

Results and discussion

3.1.

Strain growth and productivity

Scenedesmus sp. cultivation was carried in two stages; in the first the cells were allowed to grow in a nutrient rich media (þ3N-BBM) for 14 d to enhance the biomass productivity, and in the second the cells were transferred to nitrogen deficient medium (N-BBM) to enhance the lipid accumulation. Fig. 1 shows the growth and the correspondence time course of lipid accumulation in the two cultivation stages. The figure shows that the lag phase was short, and the growth entered

the exponential growth phase almost immediately. As expected, the lag phase was short as the inoculum into the photobioreactor was from the stock broth of cells grown in the same medium. In stage 1, the specific growth rate was 0.195 d1, determined from the slope of the logarithmic growth line (not shown here). The lipid content was determined at the beginning and end of this stage and found to remain almost constant at 12.6  0.8% (dry basis). The lipid content was determined after cells lyophilization at 54  C and 0.02 mbar for 6 h in the freeze drier followed by SC-CO2 extraction at 50  C, 500 bars and 3 ml min1 with 100 ml total CO2 passed. Multiplying the biomass productivity, determined from the slope of the growth curve, by lipid content resulted in overall lipid productivity of 19.5 mg l1 d1. Similar lipid content was also reported in the work of Rodolfi et al. [35] done on a strain of a genus similar to the one used in this work. The exact species of the strain used in this work was not identified. However, slightly higher biomass productivity was obtained by Rodolfi et al. [35], which resulted in higher overall lipid productivity. This is mainly because the cultivation was done with 5% CO2 enrichment, whereas in this work, the cultivation was autotrophic with CO2 being the sole carbon source. In addition, this slightly higher productivity is due to the preferable illumination intensity and duration, which were in this work 120 mmol m2 s1 under 12 h light/dark photoperiod, respectively, compared to 100 mmol m2 s1 under continuous illumination applied in Ref. [35]. These results are also comparable to those found by Ren et al. [36] for the same strain but grown heterotrophically.

3.2.

Fig. 1 e Cell growth (C D3N-BBM and B LN-BBM) and lipid content (-) curves of Scenedesmus sp. grown in 5 l photobioreactor

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Effect of nitrogen stressing on lipid productivity

In the second stage, cells were grown in a nitrogen deficient medium (N-BBM), which resulted in a sharp drop in the specific growth rate to 0.0165 d1, as shown in Fig. 1. The slight drop in biomass concentration observed at the beginning of the second stage was due to loss of some cells during the harvesting, after collecting the cells and re-culturing them in the new medium. However, having grown in stage 1 to a sufficient concentration, the accumulation of lipid was the main objective of this stage. This reduces the net energy and increases biodiesel productivity [8]. Real outdoor cultivations may have lower growth rates due to self-shading and other effects. However, even if lower growth rates are, the proposed technique will still be applicable. The total lipid content in the first day of starvation was 12.6% based on dried cell weight. The lipid accumulation was monitored by staining with Nile Red. Fig. 2 shows the fluorescence microscopic images in different days after starvation, where the brightness in the photos indicates the amount of lipids. It is clearly shown that by day 11, cells started to accumulate the lipid, which enhanced further until day 23. This has been also proven by quantitative measurements using micro-plate reader as shown in Fig. 3. The lipid content was also determined gravimetrically by Soxhlet extraction from lyophilized samples and a calibration curve between the fluorescence intensity and lipid content was prepared. The results shown in Fig. 3 further confirmed that lipid accumulation started after day 11 of starvation and increased exponentially thereafter. After 3 weeks of starvation, a six-fold increase in lipid content

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Fig. 2 e Fluorescence microscopy images of Scenedesmus sp. cells stained with Nile Red and showing lipid accumulation after 1 (a), 11 (b), 14 (c), 20 (d) and 23 (e) days of nitrogen starvation in cultures. The brightness in the photos indicates the amount of lipids.

was observed, reaching 73% on a dry weight basis. The proposed two step process results in lowering the harvesting and lipid extraction costs. Table 1 shows the FAME compositions of extracted lipid, after SC-CO2 extraction, rapid transesterification to FAME, and analysis using GC-FID, from the growth in nitrogen sufficient (þ3N-BBM) and nitrogen deficient (N-BBM) media. The results show that the compositions of the extracted lipid were almost similar, but the triglycerides content (fatty acid contents) was higher in the lipid extracted from cells grown in N-BBM confirming that the natural lipid are accumulated by starvation.

3.3.

Pretreatment effect on lipid extraction yield

The use of chemical solvents, such as n-hexane, has several drawbacks, which include the leftover biomass contamination with the solvent, long extraction time and the need of additional separation units. These drawbacks can be overcome by using SC-CO2 extraction. It was also assumed that operating at the supercritical conditions may allow the solvent (CO2) to penetrate through the water film and reach the lipid. Therefore, SC-CO2 was tested with both lyophilized and wet samples. The results, shown in Fig. 4, indicated that the extraction yield from wet and lyophilized samples was close to those obtained using n-hexane. The reproducibility of the results was confirmed by doing the experiments in at least in a

Fig. 3 e Relative fluorescence intensity (C) determined using micro-plate reader and lipid content in dry weight basis (B) in the nitrogen starvation stage.

duplicate, and the results presented in Fig. 4 are the average values with the error bars representing the respective standard deviations. As mentioned earlier, drying is a time consuming step and in many cases an energy intensive process. Avoiding this step would significantly enhance the microalgae-biodiesel production process. Thus, in this study, at first, the lipid extracted from lyophilized cells was compared to that extracted from wet biomass. Soxhlet extraction from lyophilized biomass using n-hexane was used to determine the total lipid content in the biomass. It was found that the total lipid content was 21.1  1.5%. The lipid extraction was also performed using static n-hexane, as a solvent and left overnight in an incubator at 50  C. Fig. 4 shows the lipid extraction yield expressed as percentage of lipid extracted per dry biomass weight. As shown in the figure, lipid extraction yield from wet sample was only 4% compared to 12.6% achieved from lyophilized sample. The lower lipid extraction yield from the wet sample is due to the water film formed over the lipid, which prevents the solvent from contacting it. In addition to the drying effect that enhanced the extraction yield, lyophilization also disrupts algal cell and makes cell walls more porous [37].

3.4.

Lipid extraction from pre-treated wet biomass

Microalgal cell wall rigidity and toughness is the main barrier to microalgal lipid extraction, and cell walls need to be disrupted in order to enhance the extraction. The main parameters that determine disruption method suitability are process cost, scalability and product contamination with other cell components. Lyophilization is the most commonly used technique, and as shown in Fig. 4, the lipid yield increased from only 4% using wet biomass to 12.6% from lyophilized sample. However this technique requires high energy to freeze the samples and its operation and maintenance costs are relatively high, which is usually not justified in energy production processes. Acid treatment has also been suggested to disrupt the cells and liberate the lipid [20]. Pretreatment using diluted sulfuric acid was tested and resulted in a lipid extraction yield of 10%. However, using acids is neither recommended in fuel production nor for the leftover biomass applications. In addition, using acids require special materials of construction, which is not economical for large-scale applications.

0.05 0.7 0.4 0.9 0.5 0.7 0.6 0.8 0.6 0.02           3.20 17.4 6.30 12.1 8.20 0.60 40.7 8.40 1.20 1.90 78.2 2.3 0.4 0.2 0.7 0.3 01

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7.00 18.6 9.30 4.70 e 7.00 41.9 4.70 4.70 2.30 76.1 1.4 0.6 0.2

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3.50  16.5  4.70  11.8  9.40  e 44.7  7.10  e 2.40  69.6 2.7 0.4 0.3 0.7 0.9 0.4 0.8 0.9 0.1 0.4 6.00  11.9  13.1  4.80  7.10  6.00  45.2  2.40  3.60  e 79.0 1.9  0.8  0.1

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7.5  0.7 16.4  0.08 13.4  0.1 6.0  0.7 7.5  0.96 4.5  0.8 41.8  0.1 e e 3.0  0.6 82.2 1.5  0.5  0.2  0.4

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 0.1

Extraction from lyophilized cell grown in þ3N-BBM. Extraction from lyophilized cell grown in N-BBM. Unsaturation ratio. The standard deviation.

7.50 15.1 11.3 e 9.40 1.90 50.9 e 1.90 1.90 79.7 1.7 0.8 0.5 0.8 0.9 0.7 0.03 0.7 0.4 6.10  12.2  8.20  2.00  12.2  2.00  53.1  4.10  e e 76.9 2.8 0.4 0.09 0.5 0.1 0.4 0.6 0.9       

3.20 21.0 8.10 12.9 8.10 1.60 43.5 e e 1.60 84.3 2.0 C14:0 C16:0 C18:0 C18:1 C18:1 (trans-9) C18:2 C18:2 (n6) C18:3 (n3) C20:0 C22:0 Total fatty acids U/Sc

Lysozyme Wet Lyophilizationa

Wet

Acid treatment

Lysozyme

Cellulase

Lyophilizationa

SC-CO2 extraction  εd n-Hexane extraction  εd Fatty acid

Table 1 e Fatty acid composition (% w/w) of the lipid extracted using SC-CO2 and n-hexane with different disruption methods.

Lyophilizationb

b i o m a s s a n d b i o e n e r g y 6 6 ( 2 0 1 4 ) 1 5 9 e1 6 7

165

To test enzymatic treatment, two enzymes have been tested, namely lysozyme, naturally used to disrupt bacterial cell walls, and cellulase, which catalyzes the hydrolysis of cellulose. The latter was tested to disrupt the cellulosic structure of the cell walls. Fig. 4 shows that lipid extraction yields from wet samples using both enzymes, separately, resulted in a better yield than even the lyophilized samples, with values of 16.6 and 15.4% using lysozyme and cellulase, respectively. This is a significant finding as it shows that using either enzyme, with lysozyme showing slightly better results, lipid can be extracted from wet sample without the need for the time consuming and energy intensive drying step. The result of this work clearly shows the superiority of using lysozyme, or cellulase, in the extraction of lipid from wet microalgae, while avoiding the drying step. The synergic effect of both enzymes was then tested using 50% mixture of lysozyme and cellulase; however lower yield of 12% was achieved. This could be due to the formation of a product from one enzymatic reaction that inhibits the other enzymatic reaction. Cellulase, from the same source as the one used in this study, was also tested by Liang et al. [38] to assist lipid extraction from pre-ultrasonicated wet cells of Chlorella vulgaris and compared to extraction from lyophilized cells using Bligh and Dyer method. It was found that the lipid extraction yield increased with the addition of the enzyme. However the highest lipid recovery (lipid extracted to actual lipid content) was 49%, which was significantly lower than the one obtained in this study, which reached 72%. In addition, the solid concentration was significantly higher, where the solids were concentrated to 18% compared to only 6.8% in this work. Furthermore, Liang et al. ultra-sonicated the lyophilized biomass, which was shown in this work to be not required. The one-way ANOVA proved that there is a significant difference in the yield with p-value equal to 0.038 between treatments and the Fisher’s (95% confidence intervals) multiple comparison demonstrated that the yield of individual enzyme use is close, while the enzyme mixture yield is not. Although the use of the enzymes has shown to be effective, their industrial implementation for low-value products is often limited due to their high cost. It is worth mentioning though that the price of lysozyme is relatively not expensive, compared to other enzymes [39]. In addition, it can be repeatedly used for many cycles without any significant lose of activity when used in an immobilized forms. Lysozyme immobilized on extrudate-shaped NaY zeolite has been successfully used for 12 cycles [40]. Immobilization of the enzyme also has the advantage of enhancing the recovery and purification steps. Due to SC-CO2 advantages over conventional solvent extraction, the application of enzymatic treatment with SCCO2 extraction from wet cells using lysozyme has been tested. The lipid extraction yield was 12.5%, which is almost 75% of that extracted using n-hexane. The significant difference in the yield was also confirmed by one way ANOVA, where a pvalue of 0.033 was obtained. This relatively lower yield can be attributed to some free lipid that may have been lost in the enzymatic aqueous solution during the disruption step after centrifugation. This did not occur with n-hexane extraction, as the solvent was added before the centrifugation, which was not possible in the SC-CO2 extraction. Nevertheless, due to its

166

b i o m a s s a n d b i o e n e r g y 6 6 ( 2 0 1 4 ) 1 5 9 e1 6 7

Fig. 4 e Yield of extracted lipid by n-hexane and SC-CO2 with different treatments.

favorable features, the successful use of SC-CO2 with wet biomass is still a major finding. Both operation time and cost are critical parameters in process commercialization. SC-CO2 approach has a shorter extraction time with less solvent consumption compared to organic solvents extraction. Although the operation cost might be higher, associated to the pumping cost, this will be counterbalanced by lower down-streaming cost, as no solvent recovery unit is required and the solvent can be separated by simple depressurizing and recycled back. Using this environmental friendly and recycled approSCCO2 was also tested with lyophilized biomass and compared to n-hexane. It was found that the difference in yield was insignificant compared to n-hexane (p-value ¼ 0.724).

3.5.

Acknowledgment The authors would like to express their sincere appreciation to Prof. Kourosh Salehi-Ashtiani, Associate Professor of Biology at New York University Abu Dhabi, for his valuable assistance in visualizing the lipid accumulation of the microalgae strains.

Analysis of extracted lipid

Complete analysis of the fatty acid composition of the extracted lipid by n-hexane and SC-CO2 was carried out using GC-FID analysis, as shown in Table 1. Mass fractions are normalized according to the total fatty acids found in the GC analysis. It is clearly shown that the extracts were mainly composed of linoleic acid (42e55%), palmitic acid (12e19%) and oleic acid (7e12%). The obtained unsaturation ratio was in the range of 1.5e3. No significant change in composition was observed by using different treatments and extraction techniques. However, the total fatty acid of the lipid extracted by SC-CO2 was slightly lower in all treatments, which is mainly due to the solubility of other pigments in SC-CO2. When considering the extract yield, quality, costs and environmental impacts of using chemical solvents, the enzymatic disruption followed by SC-CO2 extraction could probably be the most efficient for biodiesel production from wet biomass. This is especially true if the leftover, protein rich, biomass after lipid extraction is used in pharmaceutical or food industries.

4.

water. The highest lipid extraction yield of 16.6% was obtained from enzymatically disrupted cells using lysozyme. The SCCO2 extraction was also successful in extracting lipid from enzymatic disrupted biomass, but with a slightly less yield compared to n-hexane. In addition, the study also shows that lipid content was enhanced in nitrogen starvation medium.

Conclusion

The work looked at the possibility of extracting lipid from microalgae, while avoiding the time consuming and energy intensive drying step. It was shown that by using enzymatic cell disruption, using lysozyme or cellulase, the lipid extraction was achieved from the wet sample that contained 93%

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