Journal of the American Oil Chemists' Society

, Volume 89, Issue 2, pp 189–198

Optimization of an Analytical Procedure for Extraction of Lipids from Microalgae

  • Eline Ryckebosch
  • Koenraad Muylaert
  • Imogen Foubert
Original Paper

DOI: 10.1007/s11746-011-1903-z

Cite this article as:
Ryckebosch, E., Muylaert, K. & Foubert, I. J Am Oil Chem Soc (2012) 89: 189. doi:10.1007/s11746-011-1903-z

Abstract

An optimized procedure for extraction of total and non-polar lipids from microalgae is proposed. The effects of solvent, pretreatment (lyophilization, inactivation of lipases, and addition of antioxidants) and cell-disruption (liquid nitrogen, sonication, and bead beating) on total lipid content, lipid class, and fatty acid composition were examined. Chloroform–methanol 1:1 was shown to be the best solvent mixture for extraction of total lipids from microalgae. When performing this extraction, lyophilized algae can be used, no pretreatment with isopropanol to inactivate the lipases is needed and addition of antioxidants is not necessary. Furthermore, cell-disruption is not essential, although in that case two extractions must be performed in series to ensure that, irrespective of the microalgal species, all lipids are extracted. Determination of non-polar lipid content should be performed by separation of the total lipid extract on an SPE column. Extraction using petroleum ether is only appropriate when a bead beater is used for pretreatment.

Keywords

AlgaeLipid extractionSolvent mixturePretreatmentCell-disruptionFat content

Introduction

Microalgae are sunlight-driven cell factories that convert carbon dioxide into potential biofuels, foods, feed, and high-value bioactives. Compared with traditional crops, they have a high areal productivity, a relatively high oil and protein content, and do not depend on arable land and freshwater. Today, there is a strong interest in lipid production using microalgae. Microalgae are theoretically capable of producing much more lipids than any conventional crop and are, therefore, attractive as a potential source of biodiesel. Because lipids from algae are often rich in the long-chain omega-3 fatty acids EPA and DHA, algae may also be a more sustainable source of these fatty acids for use in food or feed compared with fish oil.

Lipids have been recovered from microalgae via a multitude of extraction methods described in the literature. Because of the nature of microalgae, regular extraction methods (used for example for food) may not be applicable. First of all, microalgae are single cells, each surrounded by an individual cell wall. Furthermore, they often contain “unusual” lipid classes and fatty acids differing from those in higher animal and plant organisms [1]. For these reasons, it is necessary to have a profound look at methods for extraction of lipids from microalgae.

The method of Folch et al. [2], which is mostly used for the extraction of total lipids from microalgae, was originally optimized for isolation and purification of total lipids from animal tissues. It uses chloroform–methanol 2:1 for extraction of the lipids and water to remove non-lipid substances from the extract. The method of Bligh and Dyer [3], also often used for the extraction of total lipids from microalgae, was originally optimized for extraction of phospholipids from fish muscle. It uses chloroform–methanol 1:2 followed by an extraction with chloroform. Iverson et al. [4] compared the Bligh and Dyer and Folch methods for determination of total lipids in marine tissue, although marine algae were not investigated. Nevertheless, their figures showed that the Bligh and Dyer method can give rise to major underestimation for samples containing more than 2–10% lipids. The unmodified Bligh and Dyer extraction is thus not suitable for microalgae, because their lipid content often exceeds 10%. Unfortunately, the method is still used by several authors. Guckert et al. [5] compared three solvent systems for extraction of lipids from the green alga Chlorella. Soxhlet extraction (dichloromethane–methanol, 3 h reflux) recovered more non-polar lipids than the other methods, but was equivalent to the room temperature modified Bligh and Dyer extraction in polar lipid recovery. The Soxhlet method, however, led to significantly lower recovery of many polyunsaturated fatty acids. The hexane–isopropanol method gave poor recovery of all lipid classes. Lee et al. [6] also tested different solvent systems (chloroform–methanol 2:1, hexane–isopropanol 3:2, dichloroethane–ethanol 1:1, and acetone–dichloromethane 1:1) for extraction of lipids from the green alga Botryococcus braunii. The highest lipid content was obtained with chloroform–methanol 2:1. Grima et al. [7] compared the fatty acid profile of the biomass of Isochrysis galbana, obtained by direct saponification, with that of the lipids extracted by use of different solvent systems followed by KOH-catalyzed saponification. The solvent mixtures compared were chloroform–methanol–water 1:2:0.8, hexane–ethanol 1:2.5, hexane–ethanol 1:0.9, butanol, ethanol, ethanol–water 1:1, and hexane–isopropanol 1:1.5. The highest yield was obtained with chloroform–methanol–water 1:2:0.8, by use of which 93.8% of the fatty acids were extracted. Matyash et al. [8] proposed the use of methyl-tert-butyl ether (MTBE) as an alternative to the carcinogenic chloroform. Because of the low density of MTBE, it forms the upper phase in the two-phase partitioning system, which makes it easier to collect. For almost all major lipid classes this method results in recoveries similar to or better than those of the “golden standards” (Folch or Bligh and Dyer recipes). They tested their method on E. coli, mouse brain, Caenorhabditis elegans, and human blood plasma. The non-polar lipid class was not considered.

For lipid extraction from microalgae, different pretreatment methods have also been used, the most important ones being lyophilization, inactivation of lipases, and addition of antioxidants. Lyophilization increases the surface area of the sample, leading to a better lipid extraction [9]. Furthermore, lyophilized preparations do not require special storage conditions and are easily rehydrated [10]. Finally, lyophilized microalgae are easily weighed and handled. Lipases potentially present in samples of plant or animal origin, which could cause lipolysis, can be denatured by plunging the sample into boiling water, isopropanol, or dilute acetic acid for short periods [11]. Heat treatment can also deactivate these enzymes [12]. When the microalgae are lyophilized, removal of almost all water may, however, enable omission of this step, because lipases require a minimum amount of water for full expression of their enzymatic activity [13]. Precaution should also be taken to eliminate oxidation of lipids during extraction, especially when the sample contains polyunsaturated fatty acids. Use of an antioxidant or antioxidant system may therefore prove beneficial [12]. Natural tissue antioxidants, for example tocopherols, afford some protection to lipid extracts, but it is usually advisable to add further synthetic antioxidants, for example butylhydroxytoluene (BHT), to storage solvents [11]. Algae are however, known to contain a large amount of natural antioxidants [14], leading to a large antioxidant capacity of microalgal extracts (our own preliminary results). This may enable omission of addition of a synthetic antioxidant. Because microalgae consist of individual cells surrounded by thick cell walls, appropriate cell-disruption before extraction may be necessary. Until now, few investigations have been performed on the effect of cell-disruption on total lipid extraction. The lipid content of Botryococcus braunii was determined by use of the solvent system chloroform–methanol 2:1 and six different cell-disruption methods (none, sonication, homogenization, French press, bead-beating, lyophilization) [6]. The highest lipid content was obtained by use of bead beating. Comparable research was performed on different microalgae using the solvent system chloroform–methanol 1:1 [15]. Large differences in extraction efficiency were obtained, depending on species (Botryococcus sp., Chlorella vulgaris, and Scenedesmus sp.) and cell-disruption method (none, autoclaving, bead-beating, microwaves, sonication, osmotic shock).

The objective of this research was to optimize an analytical procedure for total lipid extraction from microalgae. The focus was on solvent mixture, pretreatment, and cell-disruption method. Not only the amount of extracted lipids was considered, but also the potential effect of the extraction procedure on degradation of the lipids, the lipid class, and fatty acid composition. Because the extraction procedure should be valid for all microalgae, the effect of cell-disruption was studied using different microalgae belonging to different taxonomic classes and with a different cell wall structure.

Because non-polar lipids are the most important in, e.g., the biodiesel industry, the possibility of using a direct extraction technique to determine non-polar lipid content was also investigated.

Materials

HPLC-grade chloroform, methanol, dichloromethane, and acetone were purchased from Labscan (Boom, Meppel, The Netherlands). GC-grade hexane, petroleum ether p.a. (bp 30–50 °C), diethyl ether p.a., acetic acid p.a., ethyl acetate p.a., isopropanol p.a., ethanol p.a., sulfuric acid p.a., TBHQ, anhydrous sodium sulfate, potassium chloride, rhodamine 6G, sodium chloride, diethylamine, N,N-diisopropylethylamine, ethanol p.a., and methyl-tert-butyl ether p.a. were purchased from Sigma–Aldrich (Bornem, Belgium). Arachidic acid (C20:0) was purchased from Nu-check Prep, (Minnesota, USA). Bis(2-methoxyethyl)aminosulfur trifluoride was purchased from TCI Europe (Zwijndrecht, Belgium).

Experimental Procedures

Microalgae Cultivation and Harvest

Phaeodactylum tricornutum Pt1 8.6 was obtained from CCMP2561 in the Provasoli–Guillard National Center for Culture of Marine Phytoplankton. The cells were grown in batch culture in 30-L Plexiglas columns, containing 25 L WC medium [16] to which 30 g/L artificial sea salt (Homarsel, Zoutman Industries, Roeselare, Belgium) was added. The culture was continuously illuminated with two fluorescent lamps (Sylvania Gro-Lux, 36 W) and aerated with 0.3 μm-filtered air. Nannochloropsis salina SAG 40.85, Chlorella vulgaris SAG 211-11b, and Arthrospira platensis SAG 85.79 were obtained from Sammlung von Algenkulturen Goettingen (SAG), Germany. Scenedesmus obliquus CCAP 276/3A was obtained from Culture Collection of Algae and Protozoa, UK. N. salina and C. vulgaris were grown in batch culture in 2-L glass bottles at 21.9 ± 0.5 °C in WC medium [16]. For N. salina 30 g/L artificial sea salt was added. A. platensis was grown in 2 L batch cultures at 21.9 ± 0.5 °C in Spirulina medium (SAG, Germany). The cultures were illuminated 12 h/day with fluorescent lamps (Philips, Master TL5 HE, 35 W/840) and aerated continuously. Growth was monitored by measuring the optical density at 550 nm. The microalgae were harvested by centrifugation in the early stationary phase (P. tricornutum––day 14, N. salina, S. obliquus––day 15, C. vulgaris––day 9, A. platensis––day 13). The pellet obtained was either used directly (fresh algae) or lyophilized for further analysis.

Experimental Setup: Effect of Solvent Mixture and Ratio

We investigated which solvent mixture and which ratio was best for extraction of lipids from microalgae. Therefore, the following solvents and mixtures, often used in literature, were tested on C. vulgaris:
  • Chloroform–methanol 1:1

  • Chloroform–methanol 2:1

  • Dichloromethane–ethanol 1:1

  • Hexane–isopropanol 3:2

  • Acetone

  • Diethyl ether

  • Methyl-tert-butyl ether–methanol 10:3

Total lipid content extracted with the different solvent systems was compared.

Experimental Setup: Effect of Pretreatments

First, it was investigated whether lyophilization causes any changes to the lipid composition of P. tricornutum. Therefore, the total lipid content, the lipid class composition, and the total fatty acid composition of fresh and lyophilized algae were compared. Lyophilization was performed on the microalgae pellet, stored at −80 °C. Second, we tested whether endogenic lipases cause lipolysis and their deactivation is thus necessary before extraction. The effect of addition of isopropanol (to deactivate the lipases [11]) to C. vulgaris on the total lipid content and the lipid class composition was investigated. Therefore, 1 mL isopropanol was added to 80–100 mg microalgae (dry weight, DW) before the total lipid extraction was performed, and the result was compared with that from a sample with no isopropanol added. The effect of addition of isopropanol was also determined for P. tricornutum. Therefore, fresh and lyophilized biomass were stored at 4 °C with or without addition of isopropanol. The free fatty acid (FFA) content of the microalgal biomass was determined on days 0, 7, and 17. Finally, to examine whether significant oxidation occurred during extraction, we examined the effect of addition of the synthetic antioxidant TBHQ on the total fatty acid composition of the alga C. vulgaris to which eicosapentaenoic acid (EPA, C20:5n-3) had been added. TBHQ was added at a concentration of 500 ppm to the methanol used during extraction.

Experimental Setup: Cell-Disruption Methods

We investigated whether algal cells need to be disrupted to achieve a complete extraction and which method is best. Therefore, three methods, often found in literature, were tested on P. tricornutum:
  • Lyophilized microalgae were frozen in liquid nitrogen and then thawed, three times.

  • Lyophilized microalgae were suspended in 0.5 mL methanol and bead beaten twice (Qiagen Tissuelyser, Venlo, The Netherlands) for 30 s at 60 Hz.

  • Lyophilized microalgae were sonicated (sonication bath EIA USCP 100SB) for 15 min after addition of 4 mL methanol, 2 mL chloroform, and 0.4 mL water.

Because an optimum procedure should be valid for all microalgae, the effect of cell-disruption was studied using different microalgae. The following microalgae were used, because they belong to different taxonomic classes and have a cell wall structure different from that of P. tricornutum (taxonomic class: Bacillariophyceae––cell wall structure: organic cell wall with up to 10 silica bands embedded in its surface in the girdle region [17]):

  • N. salina (taxonomic class: Eustigmatophyceae––cell wall structure: algaenans containing trilaminar outer cell wall (TLS) [18]);

  • S. obliquus (taxonomic class: Chlorophyceae––cell wall structure: glucose (major), galactose, mannose rigid cell wall, glucose wall matrix [19]);

  • C. vulgaris (taxonomic class: Trebouxiophyceae––cell wall structure: no TLS [20], glucosamine rigid cell wall, no fucose matrix [21]); and

  • A. platensis (taxonomic class: Cyanophyceae––cell wall structure: four-layered cell wall with a fibrillar β-1,2-glucan inner layer, a murein one, another fibrillar layer and a structured outer layer [22, 23]).

For time-saving reasons and because the different cell-disruption methods resulted in no significant differences for the lipid content of P. tricornutum (Section "Effect of Cell-Distruption Method on Total Lipid Extraction") for the remaining algae all cell-disruption methods were performed in series and compared with lyophilized algae without further cell-disruption.

Analysis of Total Lipid Content––Testing Solvents and Mixtures

The different solvents and mixtures described above were tested. Briefly, the tested solvent or mixture (6 mL) was added to 100 mg lyophilized microalgae and the tube was vortex mixed for 30 s. The solvent or mixture (2 mL) and water (2 mL) were then added and the tube was vortex mixed again and subsequently centrifuged at 2,000 rpm for 10 min. The aqueous layer was removed and the solvent layer was transferred into a clear tube. The remaining solids were re-extracted with 4 mL solvent or mixture. The combined solvent layers were passed through a layer of anhydrous sodium sulfate using Whatman No. 1 filter paper in a funnel. The solvent was removed by rotary evaporation at 40 °C after which the lipid content was determined gravimetrically. The extraction was performed in quadruplicate. The resulting percentage of extracted lipids is the sum of three extractions performed in series.

Analysis of Total Lipid Content––Testing Pretreatment and Cell-Disruption

The different pretreatments and cell-disruption methods were incorporated into the general chloroform–methanol 1:1 extraction method, which was selected as optimum (section “Effect of Solvent”). Briefly, 400 mg fresh microalgae paste or 100 mg lyophilized microalgae was mixed with 4 mL methanol. Chloroform (2 mL) and water (0.4 mL) were added and the mixture was vortex mixed for 30 s. Chloroform (2 mL) and water (2 mL) were added and the mixture was vortex mixed again. The tubes were centrifuged at 2,000 rpm for 10 min. The upper layer was removed and the lower layer was transferred into a clear tube. The remaining solids were re-extracted with 4 mL chloroform–methanol 1:1. The combined solvent layers were passed through a layer of anhydrous sodium sulfate using Whatman No. 1 filter paper in a funnel. The solvent was removed by rotary evaporation at 40 °C after which the lipid content was determined gravimetrically. The extraction was performed in quadruplicate. Taking the high lipid content of algae into account, the extraction was repeated four times in series on the same algae, so possible saturation of the solvent mixture with lipids could be detected. Lipid content was determined separately for each extraction.

Analysis of Non-polar Lipid Content

Non-polar lipids were extracted with petroleum ether [11, 12]. The extraction method was similar to the total lipid extraction method, with replacement of the methanol and chloroform with petroleum ether. No water was added, unless otherwise stated. Considering the high non-polar lipid content of algae, the extraction was repeated five times in series on the same algae, so possible saturation of the solvent with lipids could be detected. Lipid content was determined separately for each extraction. The extractions were performed in quadruplicate.

Analysis of Lipid Class Composition

To determine the effects of the pretreatments on lipid class composition, thin-layer chromatography (TLC) as described by Christie [11] was performed. Briefly, lipids were separated by spotting the total extract on a silica gel 60 F254 plate (Merck, Darmstadt, Germany). Non-polar lipids were separated by use of hexane–diethyl ether–acetic acid 80:20:2 as mobile phase and polar lipids were separated by use of ethyl acetate–isopropanol–chloroform–methanol–0.25% KCl in water 25:25:25:10:9 as mobile phase. Spots were visualized by use of rhodamine 6G solution in ethanol (0.5 g/L). Tentative identification of the lipid classes was performed using standards that were spotted next to the samples.

To determine the effects of cell-disruption methods on lipid class composition, silica solid phase extraction (SPE) (Refs. [11, 24], with slight adjustments) was performed. Briefly, the SPE column was conditioned with 10 mL chloroform. Then, approximately 10 mg lipids in 100 μL chloroform was applied to the column. Elution with 10 mL chloroform yielded the non-polar lipids (NL), 10 mL acetone gave the glycolipid fraction (GL), and 10 mL methanol yielded the phospholipids (PL). Each class was determined gravimetrically. Quantitative determination of lipid class composition was performed in triplicate.

Analysis of Fatty Acid Composition

To determine the effects of the pretreatments and the cell-disruption methods on fatty acid composition, methylation was performed according to Ref. [11], with slight adjustments. Briefly, 5 mg lipid sample was dissolved in 1 mL toluene and 2 mL 1% sulfuric acid in methanol was added. The mixture was left overnight in a stoppered tube at 50 °C. Aqueous sodium chloride solution (5%, 5 mL) was then added and the required methyl esters were extracted with 3 mL hexane. Necessary dilutions were made before injection for GC analysis.

The fatty acid methyl esters (FAMEs) obtained were separated by gas chromatography with cold on-column injection and flame ionization detection (FID) (Carlo Erba Instruments, Interscience, Louvain-la-Neuve, Belgium; GC8000 series instrument). An EC Wax column of length 30 m, ID 0.32 mm, film 0.25 μm (Grace) was used with the following time–temperature program: 100–180 °C at 10 °/min, 180–215 °C at 2 °/min, 215 °C (44 min). Peak areas were quantified with Chromcard for Windows software (Interscience, Louvain-la-Neuve, Belgium). Standards (Nu-check, Elysian, USA) containing a total of 35 different FAMEs were analyzed for provisional peak identification. Peak identification was confirmed by use of GC–MS (Trace GC, Thermo Scientific) using an Rxi-5 Sil MS column of length 30 m, ID 0.25 mm, film 0.25 μm (Restek, Belgium).

Analysis of FFA Content

The FFA content was determined by selective formation of diethyl amide derivatives according to Kangani et al. [25]. To do this, 0.45 mg arachidic acid (C20:0) in chloroform (150 μL) was added as internal standard before extraction. The extracted lipids were then dissolved in 750 μL dichloromethane and transferred into a screw-capped tube. After addition of 10 μL diisopropylethylamine and 30 μL diethylamine, the solution was cooled to 0 °C. Bis(2-methoxyethyl)aminosulfur trifluoride (10 μL) was added dropwise and the solution was vortex mixed for 5 s. The solution was kept at 0 °C for 5 min, subsequently warmed to room temperature, and kept there for 15 min. Water (2 mL) and hexane (4 mL) were added and the tubes were vortex mixed for 1 min. After centrifugation for 10 min at 2,000 rpm, the organic layer was collected and transferred into a vial for GC analysis. A blank analysis was performed by use of the same method, but without addition of bis(2-methoxyethyl)aminosulfur trifluoride.

The diethyl amide derivatives were analyzed by gas chromatography with cold on-column injection and FID (GC8000; Carlo Erba Instruments). An EC Wax column of length 30 m, ID 0.32 mm, film 0.25 μm (Grace) was used with the following temperature program: 100–160 °C at 10 °/min, 160–240 °C at 2 °/min, 240 °C (7 min). Peak areas were quantified with Chromcard for Windows software. The areas of the peaks that were not present in the blank were summed and compared with the area of the internal standard (C20:0).

Statistics

Results were statistically evaluated by use of one-way analysis of variance (ANOVA) and a post-hoc Tukey’s test with α = 0.05 (Sigmaplot 11, Systat Software).

Results and Discussion

Effect of Solvent

The extracted total lipid percentage was highly dependent on the solvent or mixture used (Fig. 1). Solvent mixtures containing a polar and a non-polar solvent extracted a greater amount of lipids. In these cases, the polar solvent releases the lipids from their protein–lipid complexes, and the lipids subsequently dissolve in the non-polar solvent [26, 27]. Depending on the extent of release by the polar solvent and the nature of the non-polar solvent, different extracted percentages are obtained [28]. Chloroform–methanol 1:1 gave the highest lipid yield. Chloroform–methanol 2:1 extracted only 76.5% of the lipids extracted with the 1:1 mixture. This is in contrast with the results of Lee et al. [29], who stated that solvent ratio has no effect on lipid recovery from fish when a high solvent-to-sample ratio is used. Only the non-polar lipids are dissolved in the relative non-polar diethyl ether, explaining the low percentage extracted when using this solvent [28]. Given its highest lipid yield, chloroform–methanol 1:1 was thus used to test the effects of pretreatments and cell-disruption method.
https://static-content.springer.com/image/art%3A10.1007%2Fs11746-011-1903-z/MediaObjects/11746_2011_1903_Fig1_HTML.gif
Fig. 1

Effect of solvent mixture and/or ratio on the total amount of lipids extracted (mean ± SD; n = 3; total of three extractions) from C. vulgaris. Solvent systems: 1 chloroform–methanol 1:1, 2 dichloromethane–ethanol 1:1, 3 hexane–isopropanol 3:2, 4 chloroform–methanol 2:1, 5 acetone, 6 diethyl ether, 7 methyl-tert-butyl ether–methanol 10:3. a, b show significance of difference

Effect of Pretreatment

First, the effect of lyophilization was tested. Total lipids extracted from fresh (24 ± 2% of DW) and lyophilized algae (26 ± 1% of DW) were not significantly different. Thus, in contrast with what Nielsen et al. [9] stated, lyophilization does not lead to better lipid extraction. The same lipid classes were detected with similar intensities by use of TLC. The fatty acid composition, also, was not significantly different (Table 1). This indicates that lyophilization is a pretreatment that can be used without altering the lipid composition. This is convenient, because it makes the microalgae easier to handle.
Table 1

Fatty acid composition as a percentage of total fatty acids extracted (mean ± SD; n = 2) from fresh (No Lyo) and lyophilized (Lyo) P. tricornutum

 

No Lyo

Lyo

C14:0

7.3 ± 0.4

7.7 ± 0.2

C16:0

26.9 ± 0.1

27.9 ± 0.4

C16:1

48.2 ± 0.3

48.7 ± 0.1

C16:3

3.2 ± 0.1

3.0 ± 0.1

C18:1

3.4 ± 0.1

3.4 ± 0.1

C20:5

11.0 ± 0.2

9.3 ± 0.5

Next, we tested whether endogenic lipases present cause lipolysis and whether their deactivation is therefore necessary before extraction to eliminate any effect on lipid content and composition. To do this, the effect of addition of isopropanol (known to deactivate the lipases) on the total lipid content and lipid class composition of C. vulgaris was investigated. Total extracted lipids of lyophilized algae (28 ± 1% of DW) and lyophilized algae treated with isopropanol (29.6 ± 0.9% of DW) were not significantly different. The same lipid classes were detected with similar intensities by use of TLC. No spot corresponding to the free fatty acid standard was detected. Testing of P. tricornutum showed that lipases were active in the fresh microalgal biomass that was not treated with isopropanol: in this case the amount of FFA increased over time. Lyophilization or treatment with isopropanol led to a much smaller FFA content, suggesting deactivation of the lipases by both techniques. The tests on both microalgal species thus indicated that addition of isopropanol to deactivate endogenic lipases is not necessary when using lyophilized biomass. Most probably, the lipases have already (partly) been deactivated by elimination of the water during lyophilization, because lipases require a minimum amount of water for full expression of their enzymatic activity [13]. Furthermore, the microalgae were extracted rapidly after harvest and lyophilization, leaving the lipases limited time to work.

To examine whether significant oxidation occurs during extraction, the effect of addition of the synthetic antioxidant TBHQ on the total fatty acid composition of the alga C. vulgaris to which eicosapentaenoic acid (EPA, C20:5n-3) was added, was examined. Although oxidation of exogenous EPA may not be the same as oxidation of endogenous complex lipids, more rapid oxidation of exogenous added fatty acids is expected. Performing the extraction procedure with or without addition of TBHQ yielded no difference in the fatty acid profile of the alga nor in the amount of EPA (Table 2). Therefore, because no oxidation of exogenous EPA was observed, it is supposed that, when using the same procedure, no oxidation of endogenous fatty acids will take place either. Addition of an antioxidant during lipid analysis is thus not necessary. This can be explained by the large antioxidant capacity of microalgal extracts (our own preliminary results), confirming that the large amount of natural antioxidants, for example tocopherols, carotenoids, polyphenols, … , known to be present in microalgae [14], protect the lipid extracts. However, these results must be regarded with caution, because for this test the algae were extracted rapidly (<1 week) after harvest and lyophilization. When storage time is significant, other results can be expected.
Table 2

Fatty acid composition as a percentage of total fatty acids extracted (mean ± SD; n = 2) from lyophilized C. vulgaris to which EPA was added, with (With TBHQ) or without (Without TBHQ) addition of TBHQ

 

Without TBHQ

With TBHQ

C16:0

17.7 ± 0.1

17.7 ± 0.4

C16:1

9.8 ± 0.5

9.4 ± 0.3

C16:2

2.5 ± 0.1

2.6 ± 0.02

C16:3

13.5 ± 0.8

13.2 ± 0.3

C18:1

11.5 ± 1.1

11.6 ± 0.4

C18:2

5.6 ± 0.2

5.4 ± 0.2

C18:3n-3

30.2 ± 1.5

28.9 ± 0.8

C20:5n-3

9.2 ± 1.9

11.2 ± 1.8

Effect of Cell-Disruption Method on Total Lipid Extraction

The amount of total lipids extracted from P. tricornutum (Fig. 2) was used as an indication of the efficiency of the cell-disruption method used. No significant differences could be detected between the different cell-disruption methods. This indicates that the cell wall is penetrated or dissolved by the solvents used, so it does not need to be destroyed for optimum extraction. However, when looking at the first extraction only, bead beating led to more efficient extraction of lipid, although the difference was rather small (<7.5%). This result is consistent with the result of Lee et al. [6], who extracted once only. During the second extraction, a smaller amount of lipids is extracted when bead beating is used as a cell-disruption method. It is most likely that the same result would have been obtained using microwaves or autoclaving, when a second extraction would have been performed by Lee et al. [15]. Irrespective of the cell-disruption method, more than 92% of the lipids is extracted during the first extraction. Two extractions in series extract more than 96% of the total lipids and is thus sufficient, because more extractions make the procedure too time-consuming. Ideally, one extraction only is sufficient to determine the complete lipid profile of the microalgae. This is, however, only possible if there is no selective extraction of specific lipid classes or fatty acids during the first or second extraction. To check this, the lipid class composition of the first and second extracts was determined (Fig. 3). No significant difference was observed, meaning that no lipid class was favoured during extraction with chloroform–methanol 1:1. Subsequently, the fatty acid composition of the lipid classes obtained was determined (Table 3). The fatty acid composition of the non-polar lipids obtained during the first and second extraction was not significantly different. The glycolipids and phospholipids extracted during the first and second extractions did not have the same fatty acid composition, however. Some fatty acids were not present in the same amounts in both extracts. Apparently, there is some selectivity toward fatty acids during total lipid extraction. However, there seems to be no trend toward (un)saturation or chain length of the fatty acids extracted first. The differences are thus probably because of selectivity in lipid class or molecular species extracted. Possibly these differences are not visible from Fig. 3, because of the high variability or because the differences are at the molecular species level (e.g., type of phospholipid or glycolipid); these were not identified by the technique used. Taking into account the biological variability and the time effort necessary for a second extraction, the differences are probably too small to conclude that a second extraction must be performed when analyzing the fatty acid composition.
https://static-content.springer.com/image/art%3A10.1007%2Fs11746-011-1903-z/MediaObjects/11746_2011_1903_Fig2_HTML.gif
Fig. 2

Effect of cell-disruption method on total lipids extracted from, and separate extractions of, P. tricornutum (mean ± SD; n = 4). Cell-disruption methods: 1 none (fresh algae), 2 lyophilization, 3 lyophilization and sonication, 4 lyophilization and liquid nitrogen, 5 lyophilization and bead beating. a, b show significant differences between amounts of total lipids extracted. x, y show significant differences between amounts of lipids extracted during first extraction

https://static-content.springer.com/image/art%3A10.1007%2Fs11746-011-1903-z/MediaObjects/11746_2011_1903_Fig3_HTML.gif
Fig. 3

Lipid class composition of the first and second total lipid extract from P. tricurnutum (mean ± SD; n = 4). a, b show significant differences. NL non-polar lipids, GL glycolipids, PL phospholipids

Table 3

Fatty acid composition as a percentage of total fatty acids (mean ± SD; n = 2) for the lipid classes from the first (Ex1) and second (Ex2) total lipid extractions of P. tricornutum

 

Non-polar lipids

Glycolipids

Phospholipids

Ex1

Ex2

Ex1

Ex2

Ex1

Ex2

C14:0

5.3 ± 0.4

5.3 ± 0.00

1.2 ± 0.2

1.2 ± 1.2

14.4 ± 3.6

9.6 ± 1.1

C15:0

tr

tr

2.1 ± 2.1

1.4 ± 0.7

C16:0

29.9 ± 1.2

30.7 ± 0.1

9.0 ± 1.7

18.1 ± 2.1a

25.2 ± 5.3

20.4 ± 0.4

C16:1b

46.5 ± 0.3

45.9 ± 0.2

23.3 ± 3.4

16.1 ± 2.5

23.3 ± 2.1

25.2 ± 6.4

C16:2b

tr

tr

3.4 ± 0.8

3.3 ± 0.3

C16:3b

tr

tr

22.5 ± 2.8

19.0 ± 2.1

tr

2.2 ± 1.8

C16:4b

1.7 ± 2.0

2.2 ± 0.1

C18:0

tr

tr

1.6 ± 1.1

6.5 ± 0.4a

tr

1.8 ± 0.9

C18:1b

7.6 ± 0.4

7.8 ± 0.1

1.6 ± 0.2

3.6 ± 0.2a

10.3 ± 0.3

8.1 ± 0.00a

C18:2b

tr

tr

2.5 ± 0.5

1.8 ± 0.3

C18:3n-6

tr

tr

tr

tr

C18:3n-3

tr

tr

1.3 ± 0.1

1.3 ± 0.1

tr

tr

C20:4n-6

tr

tr

tr

tr

tr

tr

C20:3n-3

tr

tr

C20:5n-3

6.1 ± 0.9

6.1 ± 0.1

34.3 ± 0.5

28.4 ± 1.1a

12.3 ± 7.8

18.0 ± 6.9

C22:2b

tr

tr

C24:0

4.9 ± 0.1

6.5 ± 0.6

C22:6n-3

tr

tr

1.8 ± 0.3

3.2 ± 2.0

a Indicates significantly different data

b Indicates that the position of the double bond(s) could not be determined or a mixture of different isomers was detected

tr indicates that the component is <1%. – indicates that the component was not detected

To test whether this procedure is applicable to different microalgae from different taxonomic classes and with different cell wall structure and composition, the total lipid content of different microalgae was determined with or without use of cell-disruption. The results are summarized in Table 4. No significant difference was observed between total lipid content with or without cell-disruption from N. salina, C. vulgaris, and A. platensis when two extractions in series were taken into account. As for P. tricornutum, this indicates that the cell wall is penetrated or dissolved by the solvents used. For N. salina, the first extraction is slightly more efficient after extra cell-disruption whereas for C. vulgaris cell-disruption results in a much greater amount of lipids extracted during the first extraction. This indicates that a different cell wall structure gives rise to different infiltration by the solvents. C. vulgaris also forms many extracellular polysaccharides [30], which possibly have a large effect on infiltration of the solvents. For A. platensis, the first extraction was slightly less efficient when cell-disruption was performed. This can be explained by the formation of a thick, sticky algae–chloroform layer after cell-disruption. Even after addition of extra chloroform, it was not possible to completely separate the chloroform–lipid layer from the algae, leading to incomplete recovery of the lipids. During the second extraction, this phenomenon was not observed, leading to complete recovery of the lipids. Thus, the same total (after two extractions) amount of lipids was extracted as when no cell-disruption was performed. For S. obliquus a small difference was observed after two total lipid extractions. In contrast with what might be expected, use of cell-disruption yielded a lower total lipid content. Most probably, losses occurred during multiple manipulation of the algae during the cell-disruption steps. The first extraction was slightly more efficient after performing cell-disruption.
Table 4

Lipids (as a percentage of dry microalgal weight) extracted during the first (Ex1) and second (Ex2) total lipid extraction (mean ± SD; n = 3)

 

Ex1

Ex2

Total lipids

% 1st Ex

Lyo

Lyo + CD

Lyo

Lyo + CD

Lyo

Lyo + CD

Lyo

Lyo + CD

N. salina

29.3 ± 0.6

32.2 ± 1.0 *

5.1 ± 0.1

2.9 ± 0.3 *

34.4 ± 0.6

35.1 ± 0.8

85.2 ± 0.2

91.8 ± 1.0 *

S. obliquus

23.6 ± 0.4

26.0 ± 0.4 *

6.1 ± 0.4

2.7 ± 0.01 *

29.7 ± 0.1

28.7 ± 0.4 *

79.6 ± 1.4

90.5 ± 0,2 *

C. vulgaris

10.2 ± 0.4

18.1 ± 1.7 *

9.6 ± 2.2

3.6 ± 0.4 *

19.9 ± 1.6

21.7 ± 2.0

51.8 ± 7.1

83.3 ± 0,5 *

A. platensis

11.3 ± 1.1

8.9 ± 0.6 *

1.6 ± 0.7

2.7 ± 0.8

13.2 ± 2.1

11.6 ± 0.8

87.8 ± 3.2

76.6 ± 5.9

Total lipids are the sum of Ex1 and Ex2. % 1st Ex is the percentage of the total lipids extracted during the first extraction. Lyo shows the results for the lyophilized algae, Lyo + CD shows the results for the lyophilized algae treated with liquid nitrogen, bead beater, and sonication

*Indicates significantly different result

In conclusion it can be said that for the different microalgae, the first extraction yields a different percentage of total lipids. Especially for C. vulgaris a second extraction step is essential for accurate determination of lipid content and comparison with other microalgae. To be able to expand the method to microalgae in general, two extraction steps must therefore be incorporated in the final procedure.

Effect of Cell-Disruption Method on Non-polar Lipid Extraction

The amount of non-polar lipids extracted from P. tricornutum was used as an indication of the efficiency of the cell-disruption method used during non-polar lipid extraction (Fig. 4). Very large differences were detected between the different cell-disruption methods. Bead beating was most efficient. This indicates that petroleum ether cannot sufficiently penetrate the cell wall or dissolve components in the cell wall, making it impossible to extract all the non-polar lipids. Bead beating is the only technique that damages the cell wall sufficiently to recover all the non-polar lipids. When using bead beating, the first extraction released 92% of the non-polar lipid content. The lipid class composition of the total non-polar lipids extracted from lyophilized bead-beaten algae was determined by use of SPE (Fig. 5). It was observed that co-extraction of polar lipids occurs: 8.2% of the extracted lipids were not non-polar lipids. Because non-polar lipid extraction only can be performed by use of a bead beater, it is very time consuming. Furthermore, it does not extract the non-polar lipids only. Therefore, it is easier, less time consuming, and more accurate to determine non-polar lipids by separation of the total extract into lipid classes by use of an SPE column.
https://static-content.springer.com/image/art%3A10.1007%2Fs11746-011-1903-z/MediaObjects/11746_2011_1903_Fig4_HTML.gif
Fig. 4

Effect of cell-disruption method on non-polar lipids extracted from, and the separate extractions of, P. tricornutum (mean ± SD; n = 4). Cell-disruption methods: 1 water (fresh algae), 2 none (fresh algae), 3 lyophilization, 4 lyophilization and sonication, 5 lyophilization and liquid nitrogen, 6 lyophilization and bead beating. a, b show significant differences between the total amounts of extracted non-polar lipids. v, w show significant differences between amounts of non-polar lipids obtained with one extraction

https://static-content.springer.com/image/art%3A10.1007%2Fs11746-011-1903-z/MediaObjects/11746_2011_1903_Fig5_HTML.gif
Fig. 5

Lipid class composition of the lipids extracted with petroleum ether from lyophilized bead-beaten P. tricornutum (mean ± SD; n = 4)

Conclusion

An optimized analytical procedure was developed for total lipid extraction from microalgae. The amount of lipids extracted from microalgal biomass is highly dependent on the solvent or mixture used. Chloroform–methanol 1:1 gave the highest lipid content and is thus the preferred solvent mixture for determination of total lipids. When performing this analysis, lyophilized algae can be used, which makes weighing easier. There is no need for pretreatment with isopropanol to inactivate the lipases or for addition of antioxidants. No cell-disruption method is necessary. In general, two extractions must be performed in series. For some microalgae however, one extraction may be sufficient. Non-polar lipids should, preferably, be determined by separation of the total lipid extract on an SPE column. If extracting with petroleum ether, bead beating is necessary as a cell-disruption step.

Acknowledgments

The research presented in this paper was financially supported by the Institute for the Promotion of Innovation by Science and Technology—Strategic Basic Research (IWT-SBO) project Sunlight and K.U.Leuven Kulak. We acknowledge IS-X (Interscience, Louvain-la-Neuve, Belgium) for the use of the GC–MS at their demolab.

Copyright information

© AOCS 2011

Authors and Affiliations

  • Eline Ryckebosch
    • 1
  • Koenraad Muylaert
    • 2
  • Imogen Foubert
    • 1
  1. 1.K. U. Leuven Kulak, Research Unit Food and Lipids, Department of Molecular and Microbial SystemsLeuven Food and Nutrition Research Centre (LFoRCe)KortrijkBelgium
  2. 2.K. U. Leuven Kulak, Lab Aquatic BiologyBiology DepartmentKortrijkBelgium