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Comparison of harvesting methods for microalgae Chlorella sp. and its potential use as a biodiesel feedstock a

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A.L. Ahmad , N.H. Mat Yasin , C.J.C. Derek & J.K. Lim

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School of Chemical Engineering, Engineering Campus, Universiti Sains Malaysia, Seri Ampangan, 14300 Nibong Tebal, Seberang Perai Selatan, Pulau Pinang, Malaysia Published online: 29 Mar 2014.

To cite this article: A.L. Ahmad, N.H. Mat Yasin, C.J.C. Derek & J.K. Lim (2014) Comparison of harvesting methods for microalgae Chlorella sp. and its potential use as a biodiesel feedstock, Environmental Technology, 35:17, 2244-2253, DOI: 10.1080/09593330.2014.900117 To link to this article: http://dx.doi.org/10.1080/09593330.2014.900117

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Environmental Technology, 2014 Vol. 35, No. 17, 2244–2253, http://dx.doi.org/10.1080/09593330.2014.900117

Comparison of harvesting methods for microalgae Chlorella sp. and its potential use as a biodiesel feedstock A.L. Ahmad∗ , N.H. Mat Yasin, C.J.C. Derek and J.K. Lim School of Chemical Engineering, Engineering Campus, Universiti Sains Malaysia, Seri Ampangan, 14300 Nibong Tebal, Seberang Perai Selatan, Pulau Pinang, Malaysia

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(Received 3 December 2013; accepted 27 February 2014 ) Three methods for harvesting Chlorella sp. biomass were analysed in this paper – centrifugation, membrane microfiltration and coagulation: there was no significant difference between the total amount of biomass obtained by centrifugation and membrane microfiltration, i.e. 0.1174 ± 0.0308 and 0.1145 ± 0.0268 g, respectively. Almost the same total lipid content was obtained using both methods, i.e. 27.96 ± 0.77 and 26.43 ± 0.67% for centrifugation and microfiltration, respectively. However, harvesting by coagulation resulted in the lowest biomass and lipid content. Similar fatty acid profiles were obtained for all of the harvesting methods, indicating that the main components were palmitic acid (C16:0), oleic acid (C18:1) and linoleic acid (C18:2). However, the amounts of the individual fatty acids were higher for microfiltration than for centrifugation and coagulation; coagulation performed the most poorly in this regard by producing the smallest amount of fatty acids (41.61 ± 6.49 mg/g dw). The harvesting method should also be selected based on the cost benefit and energy requirements. The membrane filtration method offers the advantages of currently decreasing capital costs, a high efficiency and low maintenance and energy requirements and is thus the most efficient method for microalgae harvesting. Keywords: centrifugation; microfiltration; coagulation; microalgal biomass; biodiesel

1. Introduction Biodiesel production has received considerable attention in recent years as a strong potential alternative to diesel. Biodiesel consists of monoalkyl esters of long-chain fatty acids derived from the chemical reaction (transesterification, pyrolysis, gasification, etc.) of renewable feedstocks such as vegetable oil or animal fats. Although biodiesel has become an economically viable alternative energy source, production costs are still a major obstacle for its large-scale commercial application. Raw material costs account for approximately 75% of the total cost of biodiesel production [1]; therefore, it is essential to selecting the best feedstock to ensure low biodiesel production costs. The feedstock should be available at the lowest price possible and in plenty.[2] Biodiesel production should also use feedstocks that do not compete with food crops or lead to land-clearing and provide reductions in greenhouse gases.[3] Microalgae are currently considered to be one of the most promising alternative primary feedstocks for biodiesel. Converting a microalgal lipid to biodiesel by transesterification has the potential to dramatically improve our environment, while being economically practical and feasible for large-scale cultivation and harvesting. However, there are still many challenges, including process recovery costs, which have to be tackled wisely: for ∗ Corresponding

author. Email: [email protected]

© 2014 Taylor & Francis

example, the downstream recovery of microalgal biomass can be substantially more expensive than culturing algae. Algal cells are typically very small, with diameters ranging from 3 to 30 μm, which has further complicated the dewatering mechanism and made biomass recovery difficult. In addition, the culture broths are generally relatively dilute (< 0.5 kg/m3 dry biomass), which requires that large volumes be used to harvest a small amount of biomass. In certain cases, the total investment dedicated to microalgae biomass recovery can sometimes reach as high as 60% of the total production cost, whereas biomass cultivation only contributes the other 40%.[4] Thus, universal harvesting techniques for recovering microalgae biomass from the culture broth are highly desirable. An optimal harvesting technique should be independent of the cultured species, consume little energy and few chemicals and not damage the valuable products in the extraction process.[5] Because of all these constraints, various harvesting techniques, such as membrane filtration,[6] magnetic separation,[7] centrifugation,[8] coagulation [9], etc. have been developed. A major challenge in the downstream processing of microalgae lies in harvesting microalgae from their growth medium, which consists of small individual cells in a large volume of the culture medium.[10] Centrifugation, filtration

Environmental Technology Table 1.

Summary of advantages and disadvantages of techniques used for harvesting microalgal biomass.

Technique Centrifugation

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Advantages

Disadvantages

References

Rapid, easy, efficient

High cost, energy consumption, cell [11–14] damage Chemical flocculation Low cost, low cell damage, ease of use Biomass toxicity, produces a large volume [4,15,16] of sludge that is difficult to dehydrate Bio-flocculation High efficiency, no cell damage Requires a higher energy input than other [17,18] flocculants Filtration Low cost, water reuse Slow process, which fails to recover [19] organisms approaching bacterial dimensions Crossflow membrane filtration Water reuse, less energy, no cell damage Membrane fouling and clogging [12–14,20,21] Submerged membrane filtration Low cost, reduced shear stress, reduced Potential problems with scale-up [22] membrane fouling Coagulation-membrane filtration Improved filtration yields, no cell Membrane surface charge can be [23,24] damage neutralized or increased by coagulants

and coagulation/flocculation are the most common harvesting methods. Here, we summarize the advantages and disadvantages of conventional techniques that are currently used for harvesting microalgal biomass (Table 1). Centrifugation is the most rapid and efficient method available. Large volumes can be rapidly processed and the biomass can remain fully contained during recovery. However, the centrifugal effect damages cells.[10] The harvesting technique should not change the microalgal population significantly or affect microalgal viability [12]; thus, centrifugal recovery is only feasible if the targeted metabolite is a high-value product because the process is highly energy-intensive. In addition, using this technique at a large scale is problematic because of high power consumption, which increases production costs.[13] Therefore, centrifugation has not been implemented at large scales.[14] Coagulation/flocculation is more effective than centrifugation because large culture volumes can be treated without high energy consumption.[20] This technique is not expensive, the process control is easy and only a settling tank is required for a large-scale implementation.[13] However, the technique can induce toxic effects, such as chemical contamination of the biomass, which can kill and prevent the growth of microalgae.[11,13] For these reasons, membrane filtration may represent an alternative solution for the concentration of fragile cells.[12,21] Operating under a low transmembrane pressure (TMP) makes the process less energy-intensive than centrifugation, and the long membrane lifespan makes recovery relatively more cost-effective in the long term. Membrane-based separation processes also pose several challenges such as membrane fouling and clogging and electrostatic repulsion from the negative surface charges of the membrane surface and algal cells.[4,13,21] The most efficient harvesting method for microalgae biomass has not yet been determined. Thus, a current centrifugation method was compared with membrane microfiltration and coagulation to determine the most efficient method for separating microalgae biomass from the culture

medium. This study was also used to determine whether the coagulant in the coagulation method interferes with the lipid extraction process and the fatty acid profiles of freshwater microalgae, which are very significant for biodiesel production. 2. Materials and methods 2.1. Microalgae The first crucial step in developing a microalgal process is to select an appropriate species. Pulz and Gross [25] stated that successful microalgal biotechnology primarily depends on choosing a microalga with specific culture conditions and products. Several factors must be considered when choosing a strain of microalgae that can be used for biodiesel production, such as the environmental conditions, available resources and the choice of a culture system.[26] The Chlorella sp. is a eukaryotic, unicellular, non-motile freshwater green microalga [27] and is the strain most favoured by researchers. Chlorella sp. cells are spherical in shape and loosely aggregated [28] with cell sizes ranging from 2 to 4 μm and are considered to be a form of phytoplankton.[29] We chose to investigate this species in this study because it appears to be a good option for biodiesel production [19] and is easily cultured in the laboratory.[12] Additionally, as a green microalga, Chlorella sp. has a relatively high growth rate and a high lipid content and is robust to environmental conditions.[30,31] 2.2.

Medium and culture conditions

Chlorella sp. cells were cultured in Bold’s basal medium (BBM) by adding 10 mL of BBM (I) and 1 mL each of BBM (II), BBM (III), BBM (IV) and BBM (V) to 1 L of sterilized distilled water (DW). BBM (I) contained the following compounds per 200 mL of DW: 5.0 g of NaNO3 , 1.5 g of MgSO4 .7H2 O, 0.5 g of NaCl, 1.5 g of K2 HPO4 , 3.5 g of KH2 PO4 and 2.28 g of H3 BO3 . BBM (II) contained the following compounds per litre of DW: 8.82 g of ZnSO4 .7H2 O,

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A.L. Ahmad et al. Chlorella sp. cultivation

Harvesting

Centrifugation

Membrane filtration

Coagulation

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Dry microalgae biomass

Add 5.5 ml water to each 30 mg of microalgae

Sonication for 10 min

Add solvent consisting of chloroform:methanol (1:1, v/v) [35]

Extraction for 24 hrs

Centrifuge at 3000 rpm for 2 min

Methanol layer + water

Chloroform layer + lipid

Residues

Evaporate solvent to obtain microalgal oil (lipid)

Figure 1.

The flowchart representing the harvesting method and lipid extraction procedure from Chlorella sp. cultivation.

1.18 g of MnCl2 .2H2 O, 1.193 g of Na2 MoO4 .2H2 O, 1.0 g of CuSO4 and 0.401 g of CoCl2 .6H2 O. BBM (III) contained the following compounds per 100 mL of DW: 5.0 g of EDTA-Na2 and 3.1 g of KOH. BBM (IV) contained the following compounds per litre of DW: 4.48 g of FeSO4 .7H2 O and 1.0 g of concentrated H2 SO4 . BBM (V) contained 0.01 g/L of vitamin B12 (cyanocabalamin) and 0.2 g/L of vitamin B1 (thiamine–HCl). Inoculums of 19 × 106 cells/mL (approximately 5 mL in volume) were resuspended in 2 L of culture medium and maintained at 25◦ C under 2000 lux of illumination in the laboratory. Charge neutralization is the basis of coagulation and membrane filtration; thus, it is important to determine the optimum harvesting time for microalgae. Thus, the Chlorella sp. cells were harvested on the ninth day, when

the cell density reached 4.86 × 109 cells/mL and the cells had reached their maximal electronegative strength.[10,21] 2.3. Harvesting Three harvesting methods, centrifugation (as a control), membrane microfiltration and coagulation, for the Chlorella sp. were compared by dividing 2 L of microalgae cultures so that all of the experiments were conducted on the same batch cultures (Figure 1). Centrifugation was carried out at 4000 rpm for 10 min using a Model Megafuge 40 centrifuge (Thermo Scientific, Germany) and then dried overnight with a freeze dryer (model 7754030, Labconco, USA). Membrane filtration was carried out in a specially fabricated crossflow configuration module using a circular,

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Environmental Technology flat membrane with an effective area of 7.07 × 10−4 m2 . The membrane used in this study was supplied by Sterlitech and was made of cellulose acetate with a 1.2 μm nominal pore size and a 47 mm diameter. The TMP was constant at 1.5 bar and the crossflow velocity (CFV) was maintained at 0.4 ms−1 . We have previously described crossflow microfiltration in detail.[21,28,32] At the end of the concentration step, the microalgae biomass that stuck to the membrane was recovered by remove the cells from the membrane surface with a suitable brush before drying step. The coagulant selected in the coagulation method was chitosan because it is a non-toxic substance and presents no risk for consumer use in subsequent applications.[33] It is bionature, biodegradable and biocompatible and has attractive adsorption properties.[34] The chitosan was obtained from Hunza Pharmaceutical Sdn. Bhd., Malaysia as an offwhite, fine powder with a mesh size of less than 120. The chitosan was dissolved in 1% dilute acetic acid solution and mixed with a magnetic stirrer at room temperature to form a 1% standard solution. The coagulation procedure followed our previous work [34] with the following parameters: a chitosan concentration of 10 ppm, a mixing time of 20 min, a mixing rate of 150 ppm and a sedimentation time of 20 min. After the coagulation process, the flocculated cells at the bottom of a beaker were recovered by remove the clear solution and then the flocs were dried overnight with a freeze dryer. Five replicate measurements were conducted for each harvesting method. 2.4. Total lipids extraction The lipid extraction was performed on the dry biomass. After harvesting, the biomass was dried overnight with a Model 7754030 freeze dryer (Labconco, USA) to measure the cell dry weight and the total lipids content. Figure 1 shows that approximately 30 mg of the freeze-dried biomass were placed into capped test tubes. A 5.5 mL volume of DW was added to the test tubes, which were then sonicated using a Model 8510 sonicator (Branson, USA) for 5 min to lyse the cells. The lipids were then extracted by mixing 12 mL of chloroform: methanol (1:1, v/v) using a slightly modified version of Bligh and Dyer’s method [35] at 60–65◦ C for 24 h. The mixture was then centrifuged at 3000 rpm for 2 min, which resulted in separation into three layers. The upper and middle layers (the methanol layer and residues) were removed and the lower layer (the chloroform layer including the lipids) was collected. The solvent was then evaporated under a nitrogen gas flow to obtain the microalgal oil (the lipids). The weight of the lipid extract was determined gravimetrically. All of the chemicals used were of analytical grade. 2.5. Acidic transesterification of microalgal oil An acid catalyst (hydrochloric acid) was used in this study because the high acidic value of microalgal oil made the use

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of an alkali catalyst for the transesterification of microalgal oil unsuitable.[36] Zhang et al. [37] have also reported that the alkali catalyst process is sensitive to water. The presence of water under alkaline conditions may cause ester saponification. A standard reaction mixture consisting of 4.25 mL of methanol, 5 mL of hexane and 215 μL of HCl (37% vol) were added to the oil, mixed at 750 rpm and heated to the reaction temperature at 80–85◦ C. The reaction mixture was heated for 2 h, allowed to cool down and then centrifuged at 3000 rpm for 2 min, resulting in the separation into two layers. The upper layer consisted of methyl ester (biodiesel) and the lower layer contained glycerol, excess methanol and the remaining catalyst. The biodiesel from each harvesting method was individually analysed in five replicate measurements to calculate the mean values for the experimental data. 2.6. Gas chromatography analysis Fatty acid methyl esters (FAMEs) were analysed using a Model 7890A gas chromatograph (Agilent, China) equipped with a flame ionization detector, a splitless automatic injector and a fused silica capillary column (30 m × 0.25 mm × 0.25 μm) (Omegawax 250, Supelco, USA). The injector and detector temperatures were maintained constant at 250◦ C and 280◦ C, respectively. The oven temperature programme was started at 100◦ C for 1 min and then ramped at 25◦ C/min to 190◦ C, at which it was kept constant for 20 min. Each of the fatty acid components were identified by comparing the retention times and peak areas with those for standard solutions. Six fatty acids (C16:0, C16:1, C18:0, C18:1, C18:2 and C18:3) were used as the standard materials. Five replicates of each fatty acid analysis were conducted. 2.7. Analytical methods In this study, a Model BX53F optical microscope (OLYMPUS, Japan) was used to observe the microalgae biomass. Samples were taken from the bottom of the beakers at the end of the coagulation process, for which microscopic images of the formed flocs of the Chlorella sp. cells were obtained. 3. Results and discussion 3.1. Comparison of biomass and lipid content for different harvesting methods The method used to harvest microalgae cultures is a major factor affecting the cost and quality of the final products.[38] However, the harvesting method of choice depends both on the fatty acids profile and on the entire recovery process; thus, factors such as the microalgae dry biomass and the lipid content should be considered. A high lipid content is one of the key criteria in selecting a harvesting method. Therefore, in the first part of this study, the amount of total

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Table 2. Lipid content (percentage of dry weight biomass = % dw) in Chlorella sp. from different harvesting methods. The values shown are the mean ± standard deviation of five replicates. Harvesting method

Water removal (%)

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Centrifugation Membrane microfiltration Coagulation

Dry weight biomass (g)

Total lipids (% dw)

100 90

0.1174 ± 0.0308 27.96 ± 0.77 0.1145 ± 0.0268 26.43 ± 0.67

95

0.0943 ± 0.0227 15.38 ± 2.61

Figure 2. Lipid content in 30 mg of biomass after extraction for (a) centrifugation, (b) microfiltration and (c) coagulation methods.

lipids was determined after lipid extraction from the cell. The results from different harvesting methods were compared to determine the highest dry biomass weight and the total lipid percentage. Table 2 shows that there was no significant difference between the total amounts of biomass of 0.1174 ± 0.0308 and 0.1145 ± 0.0268 g obtained from centrifugation and membrane microfiltration, respectively. The lipid yields were 27.96 ± 0.77% and 26.43 ± 0.67% for centrifugation and microfiltration, respectively; thus, the total lipids were almost the same for both methods and notably close to the total lipids from previous studies.[30] The authors of these studies reported that the lipid content obtained after 20 days of incubation of Chlorella vulgaris was significantly higher (29.53%) than that obtained after 15 days of incubation (26.71%). However, in our study, the surface charge of the Chlorella sp. cells is an important criterion in determining the optimum harvesting time, as we have previously discussed.[21,34]

Figure 3.

Centrifugation and microfiltration exhibit the strongest potential among current harvesting methods because they strike a balance between a high biomass and a high lipid content. It could be seen from Figure 2 that the lipid contents in 30 mg of microalgae biomass for centrifugation and microfiltration almost show the similar quantity of lipid compared to the lipid content after the coagulation method. The lowest total lipids (15.38 ± 2.61%) were obtained by coagulation, and this value seems to deviate from the amount of dry biomass which is 0.0943 ± 0.0227 g. This result can be attributed to the weight of dry biomass being made up of the weights of both the Chlorella sp. cells and the chitosan. To confirm this hypothesis, Chlorella sp. cells were observed under an optical microscope. Figure 3 shows the microscopic images of the Chlorella sp. cells before and after coagulation. The microalgae are present as single cells and no floc formation is observed in Figure 3(a). The microscopic images also show that the majority of the Chlorella sp. cells were trapped in flocs formed by the bridging of chitosan and Chorella sp. cells, and only a few cells remained in the suspension after the addition of chitosan (Figure 3(b)). This phenomenon has already been observed by Salim et al. [39] for harvesting microalgae by bio-flocculation. These authors reported that the positively charged polymers bind partially or completely to negatively charged microalgal cells, thereby bridging the cells to produce a network of polymers and microalgal cells. These observations, combined with a comparison of the pictures in Figure 3(a) and 3(b), confirm that adding chitosan as a coagulant affected the weight of the dry biomass and thus interfered with the lipid yield from the cells. One way of resolving this issue is to use an additional method to break the attachment of the flocculated chitosan to the Chlorella cells. Thus, further research should be carried out to gain insight into this phenomenon. Figure 4 shows the microalgae culture and the microscope images of the Chlorella sp. cells before and after microfiltration for 2 L of microalgae culture. After 4 h of membrane filtration, the numbers of Chlorella sp. cells per millilitre of culture under a × 20 magnification increased significantly, and the volume of the microalgae culture in the beakers was reduced to only 10% of the original

Microscopic picture of (a) individual Chlorella sp. cells before coagulation and (b) flocculated Chlorella cells after coagulation.

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Environmental Technology

Figure 4.

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Microalgae culture and microscopic picture of Chlorella sp. (a) before and (b) after microfiltration. Table 3.

Fatty acid composition of Chlorella sp. for various harvesting methods. Amount of fatty acid (mg/g dw)

Fatty acid C16:0 C16:1 C18:0 C18:1 C18:2 C18:3 Others Total SFA Total UFA Total FAME

Centrifugation

Microfiltration

Coagulation

17.17 ± 2.11 (21.72) 0.51 ± 0.06 (0.95) 1.97 ± 0.35 (3.00) 8.13 ± 1.86 (15.68) 23.55 ± 1.54 (47.28) 0.74 ± 0.03 (10.87) 0.21 ± 0.04 (0.50) 19.15 ± 2.46 (25.22) 32.93 ± 3.49 (74.78) 52.07 ± 5.95 (100)

20.49 ± 2.43 (22.20) 0.49 ± 0.03 (0.98) 2.48 ± 1.74 (3.87) 11.43 ± 1.75 (15.30) 24.32 ± 0.93 (45.35) 0.83 ± 0.02 (11.69) 0.23 ± 0.02 (0.50) 22.97 ± 4.17 (26.67) 37.07 ± 2.73 (73.33) 60.05 ± 6.90 (100)

14.66 ± 2.35 (22.83) 0.40 ± 0.01 (1.11) 0.89 ± 0.05 (1.58) 7.81 ± 2.90 (12.81) 17.09 ± 1.07 (46.53) 0.74 ± 0.12 (14.62) 0.09 ± 0.01 (0.52) 15.56 ± 2.4 (24.93) 26.05 ± 4.09 (75.07) 41.61 ± 6.49 (100)

Note: (): Fatty acid composition (wt %). SFA, saturated fatty acid; UFA, unsaturated fatty acid. The data expressed as the mean ± standard deviation for five replicates.

volume of the thick cell suspension. These observations show that membrane filtration is a convenient and efficient method because it can be used to harvest a small amount of biomass from the large volumes of culture. The microfiltration method also resulted in 90% water removal (Table 2), which can be considered to be a good percentage removal. However, increasing the membrane filtration time to 6 h could increase the water removal up to 98%.

3.2.

Fatty acid profiles from various harvesting methods The fatty acid profiles for the Chlorella sp. studied for the various harvesting methods were determined by gas chromatography analysis (Table 3). A previous study [40] showed that oleic acid and linoleic acid were the most common fatty acids in the C. vulgaris strain. However, in our biomass study, the major components of Chlorella sp. were palmitic acid (C16:0), oleic acid (C18:1) and linoleic acid (C18:2), followed by stearic acid (C18:0) and linolenic acid (C18:3) and small amounts of palmitoleic acid (C16:0). Table 3 shows that slightly lower amounts

of total FAMEs were obtained from centrifugation compared to microfiltration, corresponding to 52.07 ± 5.95 and 60.05 ± 6.90 mg/g dw, respectively. The amounts of each of the FAME components were also higher using microfiltration than centrifugation and coagulation. Coagulation produced the worst FAME because it resulted in the small amount of fatty acids (41.61 ± 6.49 mg/g dw). This result may have been because of the chitosan contents interferes with the lipid extraction process, which may have affected the fatty acids profile measurements. Thus, chitosan can interfere with final biomass use (e.g. biodiesel) or further biomass processing (e.g. lipid extraction). Griffiths and Harrison [26] and Knothe [41] noted that the length of the carbon chain and the degree of unsaturation are important fatty acid characteristics for biodiesel production and may affect biodiesel properties such as the cetane number, the oxidative stability and the cold-flow properties. Table 3 shows that unsaturated FAMEs (C16:1, C18:1, C18:2 and C18:3) were predominant in the fatty acids profile (> 70%) and that a significant percentage of palmitic acid was also present (> 20%). Other components were also identified such as myristic acid (C14:0) and pentadecanoic

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Table 4.

Comparison of microalgae harvesting methods.

Harvesting method Centrifugation Membrane filtration Coagulation a Data

Percentage of Duration a ,b Relative energy Maintenance (min) require Reliability Efficiency required recovery a (%) 99 90–98 70–80

30 45 40

High Low Very low

Good Good Poor

High High Low

High Low Low

Remarks

References

One-step harvesting Continuous method Coagulant required

[48] [49,50] [48]

from experimental results. for 600 mL volume of microalgae culture.

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b Duration

acid (C15:0) which accounted for only 0.5% of the overall compositions. Note that high unsaturated FAME contents can reduce the pour point of biodiesel, enabling it to be used in countries with cold climates.[42] Polyunsaturated FAMEs (C18:2 and C18:3) have very good cold-flow properties but are more susceptible to oxidation.[43] However, the high content of unsaturated fatty acids (UFAs) with double bond ≥ 3 is undesirable since it reduces the chemical and oxidative stability of the resulting biodiesel.[41] Unlike the two major UFAs (C18:1 and C18:2), special attention should be taken to the content of linolenic acid (C18:3) due to the European regulation (EN 14213 and EN 14214) has set at 12% the maximum allowed content of C18:3 for quality biodiesel.[44] Furthermore, fatty acids with high oleic acid (C18:1) have been reported to have a reasonable balance of fuel properties due to increases in oxidative stability for a longer storage.[41,45] As shown in Table 3, significant changes in the amount of C18:3 were observed by varying the harvesting method of Chlorella sp. The percentage of C18:3 in Chlorella sp. by centrifugation and microfiltration as harvesting method makes the biodiesel derived from Chlorella sp. able to meet the level of European regulation for transportation use [44]; thus, Chlorella sp. biomass is a good candidate for biodiesel production. However, the percentage of C18:3 unsaturated chains exceed the maximum allowed when applying coagulation as the harvesting method.

3.3.

Economic analysis of harvesting methods

Basic considerations in selecting a suitable harvesting method are cost estimation and evaluating the energy requirements. Researchers often forget that the successful commercialization of a harvesting technology depends on economic factors. The harvesting technique must, at a minimum, be comparable in overall cost and preferably be lower in cost to existing technologies.[46] In the centrifugation process, the microalgae culture is subjected to centrifugal forces that drive cell motion through the liquid. Several centrifugal devices (i.e. rotating wall devices) have been studied in the literature for potential use in algae separation or concentration processes.[47] These devices were found to be very efficient in a one-step harvesting process. Although centrifugation is a highly reliable and efficient

method (Table 4), one should keep in mind that a high operational cost can be incurred because the devices need to be stopped and most often, cleaned manually.[51] Up to 99% of microalgae cells can be recovered via centrifugation with only 30 min of duration using 50 mL of centrifuge tubes; however, the process involves high energy costs because of increased power consumption for a large-scale production and potentially higher maintenance requirements because of freely moving parts, which make this method uneconomical. Microalgae separation by coagulation is considered to be an inexpensive process that requires only a coagulant and has low maintenance requirements (Table 4). In largescale biomass harvesting, minimal costs are incurred for the energy consumed by pumping the suspension from the growth chamber to the settling tank [13]; however, coagulation has a very low reliability and efficiency.[51] To remove large microalgae cells such as Spirulina sp., which exhibit a reasonable settling velocity in a suspension, gravity sedimentation under free settling is satisfactory. However, to remove fine cells with a diameter of a few microns, such as Scenedesmus and Chlorella sp., practical coagulation operation may be induced by adding multivalent metal salts to form larger cells with a reasonable settling velocity, which increases the percentage of recovery. Costs will increase if a higher quality coagulant is used. The selection of the appropriate coagulant is important for the process of biodiesel production. Therefore, the coagulant utilized must be cheap, nontoxic and effective at low application dosages. The only coagulant that meets these criteria is chitosan, which being bionatural and environmental friendly can ensure the survival of the microalgae during the separation process.[34] Thus, the percentage of recovery is only in the 70–80% range, which cannot compete with the percentage of recovery from centrifugation. In addition, after coagulation, some of the microalgae cells remain on the top surface and have a tendency to float instead of settling to the bottom, preventing complete harvesting. Rashid et al. [49] have made similar observations. These authors proposed additional mixing to destabilize the floating cells, such that they can settle to the bottom and be successfully harvested from the solution. However, additional mixing represents an additional energy cost, which is unfavourable from an economic perspective. As presented in Table 2, the percentage of water removal for the coagulation method reached 95%. Coagulation is a

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Environmental Technology two-step harvesting process in which coagulation is used to preconcentrate the biomass before a physical method is used for the final dewatering.[50] Most of the water can be removed during the first step; however, the second dewatering step is quite complicated because some of the cells stay on the top surface of the solution, while other cells rest at the bottom. Therefore, microalgal flocs should have a high sedimentation rate resulting in rapid settling. This technology requires an extra harvesting unit and thus can increase investment costs. Tran et al. [52] reported a higher water content of harvested microalgae from coagulation than from centrifugation. This result was obtained because centrifugation can remove as much water as possible, i.e. up to 100%, from the solution. Centrifugation, however, is too expensive and has high energy requirements because of the large volumes of culture medium that need to be processed. Continuous harvesting by membrane filtration is recommended to produce high-quality microalgae. Microfiltration and ultrafiltration have been shown to be more efficient and suitable for harvesting fragile microalgal cells.[4,19] The longer time to harvest 600 mL volume of Chlorella sp. suspensions for membrane filtration was needed as compared to centrifugation and coagulation methods (Table 4). However, this disadvantage balanced against their benefits. Their considerable operational costs, which include the capital and maintenance costs of membrane installation as well as energy costs, should be compared with the benefits offered by the end-product. A detailed description of the cost estimation for membrane filtration has been presented elsewhere.[48,53] Schafer et al. [48] estimated that the membrane life is on the order of five years; thus, the membrane cost is both a maintenance and an investment cost. Their study also showed that membranes offer the advantage of currently decreasing capital costs and have low maintenance requirements. This decrement results from allocating capital costs only to the initial cost of membrane installation and the cost of the skids, housing, pumps, valves, automation and other associated hardware.[53] Capital costs differ from operating costs, which increase every year. The operating costs are calculated by summing the costs of energy, chemicals and membrane replacement, where the energy costs are calculated considering the applied TMP, the energy required to maintain a specified CFV, the type of membrane process used, etc.[53] Operating under a low TMP appears to be a cost-effective option for small facilities for which membrane filtration becomes less energy intensive than centrifugation.[53] 4. Conclusion The freshwater microalgae Chlorella sp. was used to study harvesting methods because it is easy to grow and has a significant lipid content. High dry weights of 0.1174 ± 0.0308 and 0.1145 ± 0.0268 g and high lipid yields of 27.96 ± 0.77% and 26.43 ± 0.67% were obtained by centrifugation and microfiltration, respectively. Using chitosan

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in the coagulation process affected the weight of the dry biomass and interfered with the lipid content results. This new finding is very important for biodiesel production. A qualitative analysis of FAMEs showed that there were slight differences between the results obtained by centrifugation and microfiltration, i.e. 52.07 ± 5.95 and 60.05 ± 6.90 mg/g dw, respectively. The highest amount of linoleic acid (at approximately 24 mg/g dw) was found in Chlorella sp., and the amount of linolenic acid that meet the level of European regulation indicating that this microalgae species could be a potential biodiesel feedstock. The final selection of a suitable microalgae harvesting method for production should consider all of the cost estimates and the energy requirements. Membrane filtration has the advantage of currently decreasing capital costs, a higher efficiency, and comparably low maintenance and energy requirements relative to other methods. Thus, membrane filtration was concluded to be the most efficient method for microalgae harvesting.

Funding The authors would like to acknowledge financial support for this work from the USM Research University (RU) Grant [Grant No. 1001/PJKIMIA/814060], the USM Postgraduate Research (PRGS) Grant [Grant No. 1001/PJKIMIA/8044014] and the USM Membrane Science and Technology Cluster. In addition, N.H. Mat Yasin gratefully acknowledges a scholarship from the Universiti Malaysia Pahang (SLAB 2011).

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