Polar Lipid Profile of Nannochloropsis oculata Determined Using a

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The polar lipid profile of Nannochloropsis oculata determined using a variety of lipid extraction procedures Kelly Servaes, Miranda Maesen, Barbara Prandi, Stefano Sforza, and Kathy Elst J. Agric. Food Chem., Just Accepted Manuscript • DOI: 10.1021/acs.jafc.5b00241 • Publication Date (Web): 24 Mar 2015 Downloaded from http://pubs.acs.org on March 30, 2015

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Journal of Agricultural and Food Chemistry

JF-2015-00241s

Revision

Original manuscript: JF-2014-031295

The polar lipid profile of Nannochloropsis oculata determined using a variety of lipid extraction procedures K. Servaesa, M. Maesena, B. Prandib, S. Sforzab, K. Elsta*

a

Flemish Institute for Technological Research (VITO), Industrial Innovation, Unit Separation

and Conversion Technology, Boeretang 200, 2400 Mol, Belgium b

University of Parma, Department of Food Science, Viale delle Scienze 59/A University

Campus, 43124-Parma, Italy

* Corresponding author: Tel.: +32 14 33 56 17 Fax: +32 14 32 11 86 E-mail address: [email protected]

1 ACS Paragon Plus Environment


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Abstract

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Lipid compositions obtained from microalgae species are affected by both the cultivation

3

conditions and the extraction method used. In this study the extraction of lipids from

4

Nannochloropsis oculata using traditional and modern extraction technologies with several

5

solvents has been compared. Because important polyunsaturated fatty acids (PUFA) are bound to

6

polar lipids, these polar lipids were the main focus of this study.

7

The dominant compounds in the glycolipid fractions were monogalactosyldiglycerides (MGDG)

8

and digalactosyldiglycerides (DGDG), bearing fatty acid chains containing at least one site of

9

unsaturation. Phosphatidylcholine (PC) and trimethylhomoserines (DGTS) were detected in the

10

phospholipid fractions. The fatty acid profile comprised large fractions of C16:0, C16:1, C20:5

11

and C18:3.

12

Extraction of specific compounds was determined by extraction efficiency as well as differences

13

in selectivity of the method used. It was observed that the composition derived from a glycolipid

14

fraction was affected to a greater extent by the method used than the phospholipid fraction.

15 16 17 18 19

Keywords: microalgae; extraction; lipids; galactolipids; phospholipids; phosphatidylcholine

20 21 22 23


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Introduction Microalgae are a heterogeneous group of cellular organisms, that convert carbon dioxide into

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potential biofuels, food, feed and high-value bioactives by means of sunlight. Compared to

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traditional crops, the cultivation of microalgae is characterized by a high growth rate, short

28

growth time, high biomass production and minimal land use.1,2

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Microalgae are a rich source of organic micromolecules, such as lipids, proteins, carbohydrates

30

and pigments.3,4 Thus, the applied cultivation conditions (medium composition, temperature,

31

illumination intensity, etc.) have a major impact on the productivity, quality and exact

32

composition of the final products, e.g. fatty acid pattern of lipids. Therefore, by tuning the

33

cultivation conditions it is possible to control the content of specific types of compounds.5,6

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Because of this microalgae can be an important and sustainable source of feedstock chemicals

35

with numerous applications within the food and feed, cosmetic, pharmaceutical and fuel

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industries.7,8

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Polyunsaturated fatty acids (PUFAs) and more specifically omega-3 long-chain fatty acids

38

(omega-3 LC-PUFA), like eicosapentaenoic acid (EPA; C20:5) and docosahexanoic acid (DHA;

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C22:6), are known to be beneficial to human health due to their unique pharmaceutical properties.

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Despite the rising awareness of their importance, their daily intake is in most countries still below

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the recommended dose.4,8,9-11 Fish oil is currently the major source of these omega-3 LC-PUFAs.

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However, with depletion of fish stocks, it will become increasingly difficult to meet the human

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demand for these omega-3 LC-PUFAs by fish oil alone.10-13 Microalgae are an abundant source

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of these omega-3 LC-PUFAs and hence can form a valuable alternative for fish oil.10,11,13 Recent

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studies have shown that a significant portion of these omega-3 LC-PUFAs are associated with

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polar lipids, i.e. glyco- and phospholipids.4,10,11 Their presence in the polar lipid fraction is

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potentially interesting since it may lead to both an increased absorption of omega-3 LC-PUFAs

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and better oxidative stability of the oil compared to triacylglycerol (TAG) oil.10,11



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Whereas in the literature significant attention is paid to the presence of free fatty acids and TAG

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in algae extracts, primarily related to the production of biodiesel,1,4,8,14-22 information on the

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occurrence of glyco- and phospholipids is rather scarce.11,12,15,21,22 Phospholipids and glycolipids

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are important components of cellular membranes.21 Certain bioactive phospholipids are applied in

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the nutritional and pharmaceutical fields to improve human health or to prevent certain diseases.

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Moreover, being amphiphilic molecules and natural surfactants, commercial phospholipids are

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used in food applications as among others baking improver, wetting enhancer and additives to

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chocolate to reduce viscosity and prevent crystallization as well as stabilizer of margarine.23,24

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Since the quantity of lipids in microalgae is relatively small (on average in the range 15-30%

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depending on the species), the extraction procedure used needs to be as efficient as possible in

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order to maximize the extent the lipids are extracted.25 Different extraction technologies and

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solvent systems have already been described in literature2,5,14, although the majority have been

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performed in the framework of biodiesel production, and thus focused only on the recovery of the

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neutral lipids (TAG). The methods range from the classical organic solvent extraction1,10,11,17,26,

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Soxhlet extraction13,20,25,27, pressurized fluid extraction4 (PFE, sold as “Accelerated Solvent

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Extraction”, ASE), through to microwave-assisted28 and ultrasound-assisted26,27 organic solvent

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extraction to supercritical fluid extraction (SFE).13,16,17 The majority use organic solvents, such as

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chloroform, pure alcohols (methanol, ethanol, isopropanol), hexane, dichloromethane, ethyl

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acetate, etc. or mixtures thereof. A combination of chloroform and methanol has been reported in

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the literature as the best option to recover total lipids from microalgae. These mixtures extract

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both neutral lipids by chloroform and polar lipids by methanol.10,11,26 A chloroform/methanol

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mixture is for example used in the method of Folch et al.29 for the extraction of total lipids from

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microalgae. This method, originally optimized for the isolation and purification of total lipids

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from animal tissues, uses chloroform/methanol (2:1) for lipid extraction and water to remove

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non-lipid substances from the extract.29,30 Recently, hexane/isopropanol (3:2) has been suggested


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as an alternative to the chloroform/methanol system.10,14,18 The mixture works in a similar fashion

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as the chloroform/methanol system, with a higher selectivity towards neutral lipids compared to

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chloroform/methanol.14,18

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However these traditional extraction methods are coupled with some disadvantages: the use of

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hazardous and flammable liquid organic solvents, high-purity solvents that are costly and the

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potential of toxic emissions during extraction. The extraction processes, e.g. Soxhlet, are often

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slow and time-consuming. Furthermore, an energy-intensive evaporation step is required for

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solvent removal. Additionally, these organic solvents can cause adverse health and environmental

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effects. In order to minimize these drawbacks, alternative, greener extraction techniques have

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been investigated.5,16 These disadvantages have led to the development of other methods of

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extraction considered more sustainable, such as PFE and SFE.

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Despite the fact that PFE requires temperatures between 50 and 200 °C and pressures of 10 to 15

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MPa, the associated increased lipid solubility, improved cell wall penetration by solvent, the

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relatively low solvent use compared to other methods and reduced emission of volatile organic

88

compounds (VOC) due to the required pressure all add up to make this technique somewhat

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greener than the more traditional methods. Further advantages include its ability to be automated

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and the lack of light and oxygen within the process which is beneficial when working with

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bioactive molecules.4,5,31

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The vast majority of SFE applications use supercritical CO2 (scCO2) as the primary solvent. The

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low critical temperature (31.1 °C) and moderate critical pressure (7.4 MPa) make it particularly

94

useful for extractions of thermolabile compounds.5,14,16 Numerous literature examples of scCO2

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extractions on different natural sources have been reported.7,8,17,32-34 A common practice in SFE,

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which has to be mentioned in relation to the physicochemical properties of supercritical fluids, is

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the use of modifiers (cosolvents). These cosolvents, e.g. methanol, ethanol, are added to the

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primary fluid to enhance the extraction efficiency. For example, the addition of 1-10% methanol



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or ethanol to scCO2 expands its extraction range to include more polar lipids. The addition of

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these modifiers is often required to assist scCO2 in the extraction of highly polar compounds.16 A

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disadvantage of scCO2 extraction, however, is its high capital cost (CAPEX) due to its pressure-

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driven character.

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It has also been reported in the literature that differences in lipid extraction arise due to the

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method and solvent used.15,35 Furthermore, the extraction efficiency of a particular solvent

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(system) cannot, in general, be extrapolated to microalgae species, as indicated by the study of

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Ryckebosch et al.10 The main factor that determines recovery from different microalgae species

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seems to be the permeability of the cell wall.

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The purpose of the present study was therefore to compare traditional (Soxhlet, Folch) and

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modern (PFE, scCO2) extraction techniques for the extraction of lipids from Nannochloropsis

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oculata. Thus, the main focus was set on the differences in fatty acid profile of the glyco- and

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phospholipids. The Soxhlet extraction method was included due to its simplicity in operation,

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relative safety and potential for scale up to industrial plant level. Chloroform/methanol (2:1) was

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evaluated as it is used as extraction solvent in the Folch method, while hexane was selected as it

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is currently the most commonly applied for both commercial extraction of food lipids and

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extraction of omega-3 LC-PUFA containing triglycerides from heterotrophic microalgae.10

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Ethanol was evaluated due to its low cost, it is volatile and has a strong affinity to membrane-

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associated lipid complexes because of its ability to form hydrogen bonds.14 Based on UPLC-MS

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data the nature of glyco- and phospholipids present in the different extracts has been identified.

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Therefore, this study aims at providing new insights to the existing literature by giving a

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comprehensive view on the influence of the extraction method used on the lipid profile of the

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polar lipids at molecular level.


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Materials and methods The microalgae species Nannochloropsis oculata was used in this study to perform the different

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extractions. Lyophilized algae were kindly provided by Proviron Industries NV (Hemiksem,

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Belgium) (lot: CCAP 849/1). The algae were first concentrated in the microfiltration unit and

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subsequently centrifuged, resulting in approximately 9.44 L with a dry weight of approximately

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12.3%. They were then subjected to a freeze drying process without delay by pouring

128

concentrated microalgae mixtures on to 600 mL trays (approximately 1 cm thichness). The freeze

129

drying process was repeated once in order to completely dry the algae. Finally the freeze-dried

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algae were vacuum packed. The analysis of Proviron Industries NV performed using the

131

procedure described by Ryckebosch et al.1, specified a total lipid fraction of the Nannochloropsis

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oculata amounting to approximately 31%, with the following subdivision into the different lipid

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classes: 34% of neutral lipids, 24% of glycolipids and 42% of phospholipids. This data, obtained

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by the procedure of Ryckebosch et al.1, i.e. extraction with chloroform/methanol (1:1), will be

135

considered as the reference.

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GC-grade chloroform, methanol, ethanol and hexane were purchased from VWR International

137

(Leuven, Belgium). All weight determinations were carried out on a calibrated analytical balance.

138 139

Extraction conditions All extractions were performed in triplicate using 7.5 g of freeze-dried Nannochloropsis oculata

140

after homogenizing them with a mortar and pestle. For the Folch extraction, however, only 1 g of

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freeze-dried Nannochloropsis oculata was used. For the PFE extraction and the Soxhlet

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extraction 3 solvents were selected: hexane, ethanol and chloroform/methanol (2:1). The

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conventional Folch extraction was performed with chloroform/methanol (2:1) and the SFE

144

extractions were done with scCO2 and scCO2 with 30% ethanol as cosolvent.

145

More detailed description of the different extraction procedures is given below. After each

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replicate extraction experiment, with the exception of the Folch extraction, the extract was split



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into two fractions and these fractions were weighed accurately. Both fractions were dried under

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nitrogen atmosphere until visibly dry and stored in a vacuum oven at 23 °C overnight.

149 150

Soxhlet extraction For the Soxhlet extraction the freeze-dried algae were placed in a 22 mm x 80 mm extraction

151

thimble (Whatman, VWR, Leuven) and extracted with 80 mL of solvent (hexane, ethanol,

152

chloroform/methanol (2:1)) for 6 h in a 30 mL Soxhlet apparatus.

153 154

Pressurized fluid extraction Lipids were extracted from the lyophilized and homogenized algae using an automated Dionex

155

ASE200 Accelerated Solvent Extraction (ASE) system. The algae (60%) were mixed with celite

156

545 (40%) (0.02-1 mm, Merck, VWR, Leuven) prior to being loaded into Dionex standard 33 mL

157

stainless steel extraction thimbles. After a small amount of celite was added to the extraction cell,

158

the sample mix (12.5 g) was loaded. Sea sand (Merck pa, VWR, Leuven) was finally used to fill

159

any extra space in the cell. Extractions were performed with hexane, ethanol and

160

chloroform/methanol (2:1) as extraction solvent. For all solvents, the same conditions were used:

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1500 psi (103 bar) and 125 °C, with a 5 min static extraction (after 5 min preheating) and 4 static

162

cycles. On completion of the extraction, the thimble was flushed with solvent (50%) and purged

163

with nitrogen for 120 s. The solvent was collected in 60 mL vials with Teflon septa.

164 165

Conventional Folch extraction 1 g of freeze-dried algae was weighed and put into glass vessels. 1 volume of chloroform, 1

166

volume of methanol and 1 volume of chloroform were added in sequence to the algae sample.

167

The mixture was shaken manually after each solvent addition. After addition of the 3 solvents,

168

the mixture was shaken for 15-20 min at room temperature with high speed. Subsequently

169

phosphate buffered saline (PBS) (pH 7-7.5) (0.8 volume) was added to obtain a final

170

concentration of 1:20 (sample weight/solution volume; g/mL). This PBS was added to increase

171

the ionic strength of the upper phase in order to improve phase separation.29 The mixture was


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then centrifuged at 1000xg for 1 min to allow the organic and aqueous layers to separate. After

173

removing and collecting the organic layer, the aqueous layer was extracted twice more by adding

174

only 2 volumes of chloroform, followed by manual shaking and centrifugation to induce phase

175

separation. The chloroform phases were collected in a tared glass vial. The extract was dried

176

under nitrogen atmosphere until visibly dry and put in a vacuum oven at 23 °C overnight.

177 178

Supercritical extraction An ISCO SFX 220 Supercritical Fluid Extractor equipped with an ISCO Model 260 D Syringe

179

pump, ISCO Restrictor Temperature Controller and an ISCO SFX 200 Controller (BRS) was

180

used. The supercritical extraction with either CO2 or CO2/30% ethanol (co-solvent) was

181

conducted at a pressure of 400 bar and a temperature of 40 °C with a 240 min dynamic

182

extraction. The flow rate of scCO2 and scCO2/30% ethanol was set to 3 mL/min and 2.1 mL/min,

183

respectively. The microalgal powder was brought into a high-pressure extraction cell. Carbon

184

dioxide was liquefied by a cooler and pressurized to operation pressure using a high-pressure

185

pump. The pressurized CO2 was then pumped into the heated extraction cell during the SFE

186

process. The extract was collected in a glass tube by venting scCO2. The temperature of the

187

restrictor (2 mL/min TCR 0.75) was set at 90 °C.

188 189

Lipid part of the extract – Analysis of total lipid content With the exception of the Folch extraction, two fractions were obtained from each replicate

190

extraction. One of these fractions was stored at -20 °C under nitrogen. Since it is known that

191

some solvent (mixtures) may not only extract lipids but also non-lipid polar compounds like

192

carbohydrates and proteins, the other half was treated with 10 mL chloroform/methanol (2:1) in

193

order to separate the non-lipid and lipid fractions in the extract.11,15 The redissolved extract was

194

shaken for 2 min and then centrifuged for 5 min at 3000xg. The supernatant was removed and the

195

residue was washed twice more with the chloroform/methanol mixture. The lipid fractions of the

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3 washing steps were collected, dried under nitrogen atmosphere until visibly dry and finally



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stored in a vacuum oven overnight. The residue was accurately weighed in order to determine the

198

total lipid content. After weighing, the extract was resuspended in 10 mL chloroform/methanol

199

(2:1) and divided into different fractions: one fraction was used for the analysis of the

200

phosphorous content, and the other one for the fractionation of lipid classes (see further). After

201

drying and weighing, the fraction for the fractionation and the additional sample were stored at -

202

20 °C under nitrogen to prevent lipid oxidation.

203 204

Separation of lipid classes In order to determine the influence of the different extraction methods and conditions on the lipid

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class composition, the lipid class content of the extracts was determined using silica solid-phase

206

extraction (SPE), performed according to the procedure described by Ryckebosch et al.1

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Briefly, a 500 mg/6mL Grace PureTM SPE Silica (Grace, Lokeren, Belgium) was first conditioned

208

by elution with 10 mL of chloroform. About 10 to 20 mg of lipids, dissolved in 500 µL

209

chloroform, was brought onto the silica column. Elution with 10 mL chloroform yielded the

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neutral lipids (NL), while 10 mL of acetone was used for elution of the glycolipids (GL). Finally,

211

the phospholipids (PL) were eluted with 10 mL of methanol. The yield of each class was

212

determined gravimetrically in triplicate.

213 214

Characterization of the lipid extracts and fractions The phosphorous concentration was determined in the total lipid extract (after washing),

215

dissolved in chloroform/methanol (2:1) by means of ICP following DIN EN 14107.

216

A known quantity of each fraction (NL, GL or PL) was dissolved in isopropanol/acetonitrile

217

(90:10) with 0.1% formic acid. The analysis was based on Ultra Performance Liquid

218

Chromatography (UPLC)-Mass Spectrometry (MS). A Waters Acquity UPLC system (Waters,

219

Milford, MA, USA) was used, equipped with a Acquity UPLC BEH300 C18 column (150 mm x

220

2.1 mm; 1.7 µm). The column temperature was maintained at 35 °C. Optimum separation was

221

obtained with a binary mobile phase constituted of acetonitrile/water (50:50) (eluent A) and


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isopropanol/acetonitrile (90:10) (eluent B), both solvent mixtures buffered with 0.1% formic acid.

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The gradient elution program was: 0-1 min: from 100% A to 45% A; 1-25 min: from 45% A to

224

30% A; 25-26 min: from 30% A to 0% A; 26-30 min: 100% B; 30-31 min: from 0% A to 100%

225

A and 31-40 min: 100% A (return to initial conditions and equilibration of the column). The flow

226

rate of the mobile phase was 0.2 mL/min. Different injection volumes were used in order to be in

227

the linear range for quantification.

228

The UPLC system was coupled to a Waters Single Quadrupole mass spectrometer (Waters,

229

Milford, MA, USA), which was operated in the positive electrospray ionisation mode (ESI+) in

230

full scan acquisition. The parameters of the mass spectrometer were as follows: electrospray

231

source block and desolvation temperature 150 °C and 300 °C, respectively; capillary voltage 4.5

232

kV; cone voltage 85 V.

233

Positive identification of the compounds was based on chromatographic retention times,

234

molecular weight and their characteristic m/z ions generated by in-source fragmentation. An

235

overview of the retention times and m/z ions of the different compounds is given in the

236

Supporting Information (Table S1). The ion chromatogram of each m/z ratio, corresponding to a

237

particular compound, was derived from the total ion chromatogram using the MassLynx Mass

238

Spectrometry Software (Waters). The area of the resulting single peak was then integrated.

239

Finally, a quantitative determination of the phospholipids was made using an analytical standard.

240

Thus, a phospholipid mixture for HPLC from Glycine max (soybean) in chloroform, containing

241

phosphatidylcholine (PC), lyso-phosphatidylcholine, phospatidylethanolamine (PE) and

242

phosphatidylinositol (PI), was purchased from Sigma-Aldrich (Bornem, Belgium). The

243

phospholipid standard was dried under nitrogen to remove chloroform and reconstituted with

244

isopropanol/acetonitrile 90:10 with 0.1% formic acid (eluent B). Five dilutions were made in

245

duplicate to obtain the following PC concentrations: 25, 50, 100, 150 and 200 µg/mL. These

246

calibration solutions were injected into the UPLC-MS system using the instrumental conditions



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described above. The calibration curve was made by plotting the total peak area of PC

248

components in function of the total PC concentration reported on the certificate of analysis of the

249

analytical standard. Linear calibration curves were constructed with a squared regression

250

coefficient R² of 0.99. The limit of detection (LOD) and limit of quantification (LOQ) of the

251

applied method were calculated considering the peak-to-peak signal-to-noise ratio in the

252

chromatogram of the standard solution with the lowest concentration of PC (25 µg/mL). Since

253

various PCs bearing different fatty acids were present in the analytical standard, the signal-to-

254

noise ratio of the lowest chromatographic peak was determined (worst case scenario). The LOD

255

and LOQ were defined as the concentration that would give a signal-to-noise ratio of 3 and 10,

256

respectively. Extrapolating from the signal-to-noise ratio observed for the lowest peak, the LOD

257

and LOQ were calculated to be equal to 4 µg/mL and 13 µg/mL, respectively.

258 259

Calculations and statistical analysis The presence of PC was assessed using the phosphorous content of the different lipid fractions.

260

The phosphorous content was converted into a PC concentration, expressed as g PC/100 g algae,

261

using the molecular weight of phosphorous (30.97 g/mol) and of oleoyl-palmitoyl-

262

phosphatidylcholine (PC 16:0/18:1) (760 g/mol). The PC 16:0/18:1 was used for this purpose

263

since Wang et al. observed that C16:0 and C18:1 are the major fatty acids in PC in

264

Nannochloropsis lipids.15

265

The total lipid fraction as well as the NL, GL and PL fractions were calculated on a weight basis

266

as g/100 g algae and the results are expressed as %.

267

The chromatographic peak area of each compound belonging to a certain class was determined

268

and related to the amount injected. This enabled the various extraction methods could be directly

269

compared on the base of their effects on each single component. An assessment of the total

270

amount of the different lipid classes present was made by the sum of all chromatographic peak

271

areas in the UPLC chromatogram belonging to the lipid class under consideration. The total peak


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area per mass algae extracted was then calculated by taking into account the relative contribution

273

of that particular lipid class to the total lipid fraction. Finally, the peak area ratio of one specific

274

compound relative to the total peak area of the lipid class under consideration yields the relative

275

lipid composition (area in %).

276

The PC concentration, expressed in µg/mL, was converted into w/w % (mass PC/mass algae)

277

using the sample volume and the weight of the PL fraction.

278

All results are the average of three replicate measurements. Each replicate measurement was

279

obtained from an independent extraction and fractionation. In addition, the standard deviation

280

was calculated, which is indicated by the error bars.

281

Results and discussion

282 283

Total lipid fraction and lipid class composition The total lipid fractions were obtained after the washing step with chloroform/methanol (2:1).

284

Data on the influence of this washing step on the total lipid fraction are shown in the Supporting

285

Information (Figure S1). In general, the total lipid fractions originating from extractions with

286

more polar solvents (e.g. ethanol) were the most affected by the washing step with

287

chloroform/methanol (2:1), resulting in a larger mass loss in comparison to non-polar extractions.

288

The polar extractions are indeed more prone to the co-extraction of non-lipid, more polar matrix

289

materials (e.g. proteins, carbohydrates).

290 291

The final total lipid fractions, i.e. after washing with chloroform/methanol (2:1), extracted by the

292

different methods from Nannochloropsis oculata are shown in Figure 1. Chloroform/methanol

293

(2:1) as extraction solvent, either with PFE, Soxhlet or Folch extraction, resulted in comparable

294

total lipid fractions in the range of 26-32%. These total lipid fractions correspond well with the

295

reference value of 31%, obtained by Ryckebosch et al.1 via extraction with chloroform/methanol

296

(1:1). The mixture chloroform/methanol (2:1) is the most frequently used solvent mixture for



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lipid extraction from any living tissue and is applied in methods such as Folch and Bligh and

298

Dyer1,14,29,30. This mixture is able to extract both non-polar lipids by chloroform and polar lipids

299

by methanol. For this reason, mixtures of chloroform/methanol tend to be more efficient in the

300

extraction of total lipids.26

301

The highest total lipid fraction is extracted from Nannochloropsis oculata using PFE extraction

302

with ethanol (49%). It can be assumed that, although a washing step has been performed, co-

303

extracted (polar) matrix components, such as phytochemicals, are still present in the extract, as

304

also observed by Mulbry et al.36

305 306

Based on the results of the total lipid fractions, it can be concluded that the polarity of the

307

extraction solvent clearly influences the crude lipid yield using both the classical extraction

308

methods (Soxhlet, Folch) and the modern techniques (PFE, SFE). Non-polar solvents can only

309

extract very non-polar lipids, whereas more polar solvents (mixtures) extract lipids of a broader

310

polarity range, resulting in a higher lipid yield.

311

With either PFE or Soxhlet extractions, the following order of total lipid fraction was observed:

312

ethanol > chloroform/methanol (2:1) > hexane. The addition of 30% ethanol as cosolvent to pure

313

scCO2 increased the total lipid yield relative to the extraction with pure scCO2. Extractions that

314

use only scCO2 usually give good recoveries of non-polar lipids. Polar lipids, however, have low

315

solubility in scCO2 and hence are not extracted. Adding ethanol expands the extraction range of

316

scCO2 to include the more polar lipids, resulting in a higher extraction yield.

317

These observations are in accordance with literature data on the influence of solvent polarity on

318

the extraction of lipids from microalgae10,11,20,34,36,37 For example, McNichol et al. observed that

319

hexane extracted approximately 3% of lipids from Nannochloropsis granulate, while ethanol and

320

chloroform/methanol (2:1) extracted > 25% of lipids on a dry weight basis.20 In the study of

321

Balasubramanian et al. the use of hexane alone gave only 16% lipid yield, while the addition of a


Page 15 of 37


322

polar solvent improved yield.37 Furthermore, no significant difference in the lipid extraction

323

efficiency was observed between different extraction methodologies (sonication, PFE,

324

homogenisation, Soxhlet), all using chloroform/methanol (2:1) as the extraction solvent.

325 326

To gain insight into the lipid class composition, the different extracts have been fractionated into

327

three portions using SPE in order to determine the NL, GL and PL (Figure 1).1 The lipid class

328

composition of the chloroform/methanol (2:1) extracts, obtained either by Soxhlet, PFE or Folch,

329

is in accordance with the specifications on Nannochloropsis oculata applying the reference

330

method (chloroform/methanol 1:1). The distribution of the lipids among the different classes is as

331

follows: phospholipids > neutral lipids > glycolipids. The phospholipids are predominantly

332

present in these extracts, in the range 40-45% of total lipids.

333

Low polarity solvents are the best for the extraction of neutral lipids, including waxes and

334

pigments, which are characterized by a very low polarity. In contrast, lipids from the chloroplast

335

and membrane, which respectively contain GL en PL, are more effectively extracted by more

336

polar organic solvents, such as ethanol or methanol.10,37,38. The affinity of polar lipids (GL and

337

PL) for more polar solvents (mixtures) is also observed in the present study. The PL fraction in

338

the ethanol extracts is the most abundant, while the yield of PL using hexane is almost negligible

339

due to the apolar character of the solvent. Also for SFE the same trend is observed. Using pure

340

scCO2 the lipid fraction consists mainly of NL, which demonstrates the selectivity of scCO2

341

towards non-polar molecules.16,39 Its non-polar nature makes pure scCO2 unable to interact with

342

either polar lipids or neutral lipids that form complexes with polar lipids.14 By the addition of

343

ethanol, the affinity towards GL and PL is enhanced, although the NL fraction is still

344

predominant.

345

The findings in this study are consistent with literature data. In the study of Ryckebosch et al.

346

comparing the extraction efficiency of hexane/isopropanol (3:2) and hexane for the extraction of



Page 16 of 37

347

lipids from different microalgae species, the NL content was higher in the hexane extract and

348

lower in the chloroform/methanol (1:1) and hexane/isopropanol (3:2) extracts for each

349

microalga.10 Hence, the amount of polar lipids (GL and PL) was lowest in the hexane extract.

350

Ryckebosch et al. also compared dichloromethane/ethanol with nonhalogenated solvent systems

351

for the extraction of lipids from Nannochloropsis gaditana.11 Dichloromethane/ethanol extracted

352

the three lipid classes with more or less the same efficiency, while all nonhalogenated solvent

353

(mixtures) more easily extracted NL than the polar lipids (GL+PL). Of the nonhalogenated

354

solvents, the highest recovery of NL was obtained with hexane/isopropanol (3:2) and ethyl

355

acetate/hexane. Also for the GL, hexane/isopropanol (3:2) proved to be the most efficient solvent,

356

while for the PL ethanol was best.11 Similar results have also been observed by Balasubramanian

357

et al.37 and Mendes34 and coworkers for which Mendes et al. evokes solvent polarities to explain

358

these observations.

359 360

The phosphorous content of the lipid extracts gives insight in the presence of phospholipids.

361

According to the literature PC was reported to be the major component of phospholipids in many

362

microalgae, though its content might vary due to species-specificity or culture conditions.8,11

363

The phosphorous content of the different crude lipid extracts (Figure S2) is in accordance with

364

the lipid class composition. The extraction of PL is favoured by a more polar extraction solvent.

365

Indeed, the highest phosphorous content is observed for ethanol, followed by

366

chloroform/methanol (2:1) and hexane. Furthermore, as pure scCO2 is not polar enough to extract

367

the phospholipids none were detected above the LOQ in the scCO2 extract. The increase in

368

polarity by the addition of 30% ethanol, however, resulted in an enhancement of the phosphorous

369

concentration, comparable to the content in the lipid extract of PFE with hexane.


Page 17 of 37


370

Compound identification in the polar lipid fractions GL and PL

371 372

GL fractions An overview of the total chromatographic peak area integrated for the GLs in the different

373

extracts is given in Figure 2. This data also indicates that by increasing the polarity of the solvent,

374

more glycolipids are being extracted, as indicated by the higher total chromatographic peak area.

375

For all extraction methods used, the majority of the GLs, i.e. highest total peak area/mass algae

376

extracted, is detected in the corresponding fractions. The analysis of the different PL fractions

377

however revealed that quite a persistent proportion of GL is also present in these fractions (Figure

378

2). Thus, all extraction methodologies lead to higher yields of GL than the analysis of the GL

379

fractions indicate. Similarities in chemical structure, and consequently polarity, between GL and

380

PL lead to the observed incomplete separation, especially when using SPE.

381 382

Glycolipids and more specifically monoglycodiglycerides and diglycodiglycerides were detected

383

as sodium adducts in the GL and PL fractions (Figure 2). Since galactolipids are the predominant

384

lipids in photosynthetic membranes, they could be assumed to be monogalactosyldiglycerides

385

(MGDG) and digalactosyldiglycerides (DGDG). The structures of MGDG and DGDG are given

386

in Figure 2. In these natural compounds the galactopyranose moiety is linked to the glycerol

387

backbone, bearing two long fatty acid chains.

388

As can be seen from Figure 3, the methodology used has a significant effect upon the extraction

389

yield of the galactolipids, shifting it more or less towards MGDG or DGDG. Both the SFE with

390

pure scCO2 and the Soxhlet extraction with hexane predominantly yield MGDGs with 98% and

391

92% of the total peak area, respectively, belonging to MGDG. The additional galactose moiety

392

makes DGDG sufficiently polar to prevent its extraction with a non-polar solvent. In contrast, the

393

PFE extraction with hexane does extract DGDGs (53% MGDG, 47% DGDG), which can be

394

explained by the higher extraction temperature and pressure. The extraction yield of DGDG is



Page 18 of 37

395

enhanced by increasing the solvent polarity, with DGDG contributing > 60% for the PFE or

396

Soxhlet extraction with chloroform/methanol (2:1).

397 398

Linked to MGDG and DGDG are different types of fatty acid (Figure 3). All galactolipids

399

identified contain at least one site of unsaturation, with a strong presence of C20:5. Among the

400

saturated fatty acids, palmitic acid (C16:0) is the most abundant. PUFAs such as C20:5 are prone

401

to oxidation30 and the incidence of C20:5 in the GL fractions indicates that the extraction

402

methods used are able to preserve this intact. This occurrence of C20:5 is in accordance with

403

literature data, where it is stated that PUFAs of microalgae are predominantly linked to polar

404

lipids, especially glycolipids (MGDG, DGDG, sulphoquinovosyldiacylglycerol (SQDG)).4,10,11

405

Also within every class, the abundance of one (or more) MGDG or DGDG compound(s) with a

406

particular fatty acid is favoured depending on the extraction method and solvent used (Figure 3).

407

For example, DGDG 16:0/16:1 is the principle DGDG extracted with chloroform/methanol (2:1)

408

as solvent, using either Soxhlet or PFE. The SFE-scCO2 extraction on the other hand

409

preferentially extracts MGDG 20:5/20:5. These observations demonstrate again that a more polar

410

solvent (mixture) extracts more polar lipids, i.e. more sugars and shorter fatty acid chains,

411

whereas the use of a less polar solvent yields less polar lipids, i.e. single sugar, longer fatty acid

412

chains.

413

Wang et al. characterized the lipid profile of Nannochloropsis oculata extracted with ethanol (80

414

°C, 30 min), followed by a separation of the lipid and non-lipid fractions using the Folch’s

415

procedure.15 DGDG was the major GL class, accounting for 28.4% of all quantified polar lipids.

416

The contribution of MGDG amounted to 2.1%. Hence, Wang et al. obtained an excess of DGDG

417

in the ethanol extract, whereas MGDG and DGDG were more equally distributed in the ethanol

418

extracts (PFE/Soxhlet) in the present study. According to Wang et al. the major fatty acids bound

419

were C16:0 (52.3%), C14:0 (14.0%) and C18:1 (10.8%) in MGDG and C16:0 (51%) and C16:1


Page 19 of 37


420

(19.1%) in DGDG.15 In this study, however, different types of fatty acids are linked to MGDG

421

and DGDG. Whereas C16:0 and C16:1 are common in both studies, the present study reveals the

422

predominance of particular PUFAs, like C18:2, C18:3 and C20:5. This difference in fatty acid

423

profile of the galactolipids can be explained by the cultivation, harvesting (sedimentation,

424

chemical flocculation, etc.) and storage (freeze, dry, freeze-dry, wet, etc.) conditions of the

425

microalgae used. The extent of production of biomass, lipids, carotenoids and carbohydrates as

426

well as the exact composition of the final products (e.g. fatty acid pattern of lipids) can vary

427

considerably depending on the prevailing conditions in the cultivation medium.5,6 In addition,

428

differences in fatty acid profile15,35 of the extracts also arise due to extraction methodology used,

429

i.e. traditional vs. PFE must also be taken into account, which can result in a different fatty acid

430

profile.

431 432

PL fractions A typical chromatogram of a PL fraction is shown in Figure 4. In the PL fractions two different

433

classes of polar lipids were detected: trimethylhomoserines (DGTS; 1,2-diacylglyceryl-O-

434

(N,N,N-trimethyl)-homoserine) and phoshatidylcholines (PC). The chemical structure of both

435

classes is given in Figure 4. DGTS belongs to the betaine lipids, a recently discovered group of

436

complex lipids, primarily found in the plant kingdom.40 Ether-linked glycerolipids containing a

437

betaine moiety occur naturally in algae, bryophytes, fungi and in some primitive protozoa and

438

photosynthetic bacteria. These lipids contain a polar group linked by an ether bond in the sn-3

439

position of the glycerol moiety, with the fatty acids esterified in the sn-1 and sn-2 positions

440

(Figure 4). A positively charged trimethylammonium group and a negatively charged carboxyl

441

group give them zwitterionic character at neutral pH. At least three types of betaine lipids have

442

been described with differing permethylated hydroxyamino acids linked to diacylglycerols

443

through an ether bond: DGTS, 1,2-diacylglyceryl-3-O-2’-(hydroxymethyl)-(N,N,N-trimethyl)-β-

444

alanine and 1,2-diacylglyceryl-3-O-carboxy-(hydroxymethyl)choline. Of these, the first is by far



Page 20 of 37

445

the most common in nature.40,41 There is evidence in the literature that supports betaine lipids (as

446

well as glycolipids) being used to replace part of the membrane phospholipids in the case of

447

phosphorous starvation, as observed in the photosynthetic bacterium Rhodobacter sphaeroides42

448

or in the plant Arabidopsis thaliana.43

449

DGTS can also be present in their lysated form due to the loss of a fatty acid residue, referred to

450

as lyso-DGTS.

451 452

The total peak area of polar lipids observed in the PL fractions of the different algae extracts is

453

given in Figure 4 with a distinction between PC, DGTS and lyso-DGTS. DGTS and PC are the

454

major contribution to the total peak area of polar lipids. The highest total peak area was found for

455

the PFE extraction with ethanol, followed by the conventional Folch extraction, with the highest

456

affinity for DGTS. As for the GL fractions, a trend of increasing total peak area with increasing

457

solvent polarity is observed, both for the traditional solvent systems and scCO2.

458 459

Figure 5 shows the fatty acid profile of the DGTS, comprising large fractions of C20:5, C16:0

460

and C16:1. Based on these results, we can state that the various extraction methods under

461

consideration are able to produce DGTS mixtures with slightly different composition in fatty

462

acids. However, in general the methodology of extraction seems to affect the composition less

463

than in the case of the galactolipids. This observation can possibly be explained by on one hand

464

the amphiphilic structure of PC and DGTS, both bearing a positively charged group (quaternary

465

ammonium), thus increasing the affinity for polar solvents such as ethanol. On the other hand,

466

sufficient solubility is maintained in more non-polar solvents such as chloroform/methanol

467

mixtures due to the presence of the two acyclic moieties. Hence, a very similar lipid distribution

468

is generated among the different extraction methods. One exception is the extraction of DGTS

469

16:0/16:1 which is particularly favoured by Soxhlet extraction with hexane.


Page 21 of 37


470

In addition, two lysated forms of DGTS were identified in the different extracts, i.e. lyso-DGTS

471

16:0 and lyso-DGTS 20:5, with C16:0 and C20:5, respectively, as the remaining fatty acids

472

(Figure 5). The majority of extraction methods favour the extraction of lyso-DGTS 20:5. The

473

trend is however inverted for Soxhlet extraction with hexane and scCO2 extraction. The non-polar

474

character of the solvent used in the latter favours the extraction of lyso-DGTS bearing short chain

475

fatty acids.

476

Due to the availability of a PC analytical standard, a quantitative assessment of the PCs in the

477

different extracts was carried out (Table 1). The highest PC concentration was observed in the PL

478

fraction of the PFE extract with ethanol. No PCs were measured above LOQ (LOQ 14 µg/mL) in

479

the pure scCO2 extract. The addition of 30% ethanol to scCO2 however results in an increase of

480

the PC extracted, with a PC content of 0.33% (w/w).

481

A total concentration of PC in the range of only 1.5-10% (w/w) in the PL fractions, which

482

corresponds to 0.1-2.5% in the algae, raised the question about the nature of other compounds

483

present and their contribution to the total concentration of PLs. The presence of homoserines

484

(DGTS, lyso-DGTS) can be stipulated with certainty, though due to the lack of an analytical

485

standard their quantification was not possible. The presence of other PL classes, i.e.

486

phosphatidylinositols and phosphatidylethanolamines, was investigated. However, these

487

compounds were not detected in a quantifiable amount.

488

The chromatogram of the different PL fractions indicating the considerable presence of polar

489

compounds, combined with the low solubility of the PL samples, even in a large volume of

490

isopropanol/acetonitrile (90:10) with 0.1% formic acid, clearly suggest the presence of very polar

491

compounds, possibly salts.

492 493

C16:0, C16:1, C20:5 and C18:3 are the dominant fatty acids linked to PCs in the different

494

extracts (Table 1). The PL fractions of all PFE extractions and the scCO2/30% ethanol extraction



Page 22 of 37

495

show a rather similar PC pattern, dominated by PCs 16:0/16:1, 16:0/18:1 and 16:0/18:3. In

496

contrast, the PL fraction originating from the conventional Folch extraction is rich in PC

497

16:0/20:5, PC16:1/16:1 and PC 16:1/20:5. From the Soxhlet extract PC16:0/16:1, PC 16:1/16:1,

498

PC 16:1/20:5 and PC 18:1/16:1 are the most abundant, independent of the solvent used. Hence, in

499

analogy to DGTS, it can be concluded that the different methodologies seem to be able to extract

500

PCs with slightly different fatty acid profile. Compared to the GL fractions, however, the fatty

501

acid pattern seems to be less affected by the extraction methodology applied. Therefore, it can be

502

stated that the different methods are able to extract PCs with similar fatty acid distribution.

503 504

Purity control of NL fractions The UPLC-MS method here applied, was specifically developed to assess the presence of GLs

505

and PLs, which were the main focus of this study. Hence, TAGs could not be detected and

506

identified. The NL fractions, however, were also analysed using this method in order to establish

507

the purity of these fractions. As expected, no polar lipids (GL + PL) were detected in the different

508

NL fractions. The chromatogram of the NL fraction originating from the PFE extraction with

509

chloroform/methanol (2:1) is given as an example in Figure 6, which clearly shows the absence

510

of peaks belonging to GLs and PLs. Hence, the SPE fractionation of lipid classes results in a

511

complete separation of the neutral lipids from the more polar ones.

512

The peaks present in the chromatograms of the NL fractions can be attributed to pigments, i.e.

513

oxygenated carotenoids, commonly known as xanthophylls. Since the presence of carotenoids in

514

the different algae extracts are not the focus of the present study, no further attention has been

515

paid to the detailed analysis of the carotenoids present.

516 517

Cultivation, harvesting and storage conditions significantly influence the lipid and fatty acid

518

profile of microalgae species. This study clearly indicates that the extraction methodology also

519

affects the composition of the lipid profile of Nannochloropsis oculata, affecting both the


Page 23 of 37


520

efficiency and selectivity of extraction. Non-polar solvents show the highest affinity for neutral

521

lipids and the extraction of polar lipids is favoured by increasing solvent polarity.

522

Special emphasis has been given to the identification of the polar lipids present in the GL and PL

523

fractions. Within these lipid classes, the extraction methodology used affects the molecular

524

composition of the extract, both in structure and fatty acid composition. This effect was more

525

pronounced in the GL fractions. Thus, the extraction of lipids from microalgae is not only a

526

matter of “how many”, but also of “which ones”, according to the different methodologies and

527

solvents used.

528

Acknowledgment

529

The authors would like to acknowledge and thank Proviron NV for the samples of the

530

Nannochloropsis oculata.

531 532

Supporting Information Available: phosphorous content of different extracts, influence of the

533

chloroform/methanol (2:1) washing step and details on the instrumental analysis (retention time,

534

m/z ions) are included in the Supporting Information. This material is available free of charge via

535

the Internet at http://pubs.acs.org.

536



Page 24 of 37

537

References

538 539 540 541 542 543 544 545 546 547 548 549 550 551 552 553 554 555 556 557 558 559 560 561 562 563 564 565 566 567 568 569 570 571 572 573 574 575 576 577 578 579 580 581 582 583 584 585

1

Ryckebosch, E.; Muylaert, K.; Foubert, I. Optimization of an analytical procedure for extraction of lipids from microalgae. J. Am. Oil Chem. Soc. 2012, 89, 189-198.

2

Mercer, P.; Armenta, R.E. Developments in oil extraction from microalgae (review article). Eur. J. Lipid Sci. Technol. 2011, 113, 539-547.

3

Bong, S.C.; Loh, S.P. A study of fatty acid composition and tocopherol content of lipid extracted from marine microalgae, Nannochloropsis oculata and Tetraselmis suecica, using solvent extraction and supercritical fluid extraction. Int. Food Res. J. 2013, 20, 721-729.

4

Pieber, S.; Schober, S.; Mittelbach, M. Pressurized fluid extraction of polyunsaturated fatty acids from the microalga Nannochloropsis oculata. Biomass Bioenergy 2012, 47, 474-482.

5

Mubarak, M.; Shaija, A.; Suchithra, T.V. A review on the extraction of lipid from microalgae for biodiesel production (Review article). Algal Res. 2015, 7, 117-123.

6

Koller, M.; Salerno, A.; Tuffner, P.; Koinigg, M.; Böchzelt, H.; Schober, S.; Pieber, S.; Schnitzer, H.; Mittelbach, M.; Braunegg, G. Characteristics and potential of micro algal cultivation strategies: a review. J. Cleaner Prod. 2012, 37, 377-388.

7

Mouahid, A.; Crampon, C.; Toudji, S.-A. A.; Badens, E. Supercritical CO2 extraction of neutral lipids from microalgae: Experiments and modelling. J. Supercrit. Fluids 2013, 77, 7-16.

8

Andrich, G.; Nesti, U.; Venturi, F.; Zinnai, A.; Fiorentini, R. Supercritical fluid extraction of bioactive lipids from the microalga Nannochloropsis sp. Eur. J. Lipid Sci. Technol. 2005, 107, 381-386.

9

Yen, H.-W.; Hu, I.-C.; Chen, C.-Y.; Ho, S.-H.; Lee, D.-J.; Chang, J.-S. Microalgae-based biorefinery – From biofuels to natural products. Bioresour. Technol. 2013, 135, 166-174.

10

Ryckebosch, E.; Bruneel, C.; Termote-Verhalle, R.; Muylaert, K.; Foubert, I. Influence of extraction solvent system on extractability of lipid components from different microalgae species. Algal Res. 2014, 3, 36-43.

11

Ryckebosch, E.; Cuéllar Bermúdez, S.P.; Termote-Verhalle, R.; Bruneel, C.; Muylaert, K.; Parra-Saldivar, R.; Foubert, I. Influence of extraction solvent system on the extractability of lipid components from the biomass of Nannochloropsis gaditana. J. Appl. Phycol. 2014, 26, 1501-1510.

12

Chen, G.-Q.; Jiang, Y.; Chen, F. Fatty acid and lipid class composition of the eicopentaenoic acid-producing microalga, Nitzchia laevis. Food Chem. 2007, 104, 1580-1585.

13

Cheung, P.C.K.; Leung, A.Y.H.; Ang, Jr., P.O. Comparison of supercritical carbon dioxide and Soxhlet extraction of lipids from a brown seaweed, Sargassum hemiphyllum (Turn.) C. Ag. J. Agric. Food Chem. 1998, 46, 4228-4232.

14

Halim, R.; Danquah, M.K.; Webley, P.A. Extraction of oil from microalgae for biodiesel production: A review. Biotechnol. Adv. 2012, 30, 709-732. 24 ACS Paragon Plus Environment

Page 25 of 37

586 587 588 589 590 591 592 593 594 595 596 597 598 599 600 601 602 603 604 605 606 607 608 609 610 611 612 613 614 615 616 617 618 619 620 621 622 623 624 625 626 627 628 629 630 631 632 633


15

Wang, G.; Wang, T. Characterization of lipid components in two microalgae for biofuel application. J. Am. Oil Chem. Soc. 2012, 89, 135-143.

16

Sahena, F.; Zaidul, I.S.M.; Jinap, S.; Karim, A.A.; Abbas, K.A.; Norulaini, N.A.N.; Omar, A.K.M. Application of supercritical CO2 in lipid extraction – A review. J. Food Eng. 2009, 95, 240-253.

17

Cheng, C.-H.; Du, Tz.-B.; Pi, H.-C.; Jang, S.-M.; Lin, Y.-H.; Lee, H.-T. Comparative study of lipid extraction from microalgae by organic solvent and supercritical CO2. Bioresour. Technol. 2011, 102, 10151-10153.

18

Halim, R.; Gladman, B.; Danquah, M.K.; Webley, P.A. Oil extraction from microalgae for biodiesel production. Bioresour. Technol. 2011, 102, 178-185.

19

Samburova, V.; Lemos, M.S.; Hiibel, S.; Hoekman, S.K.; Cushman, J.C.; Zielinska, B. Analysis of triacylglycerols and free fatty acids in algae using ultra-performance liquid chromatography mass spectrometry. J. Am. Oil Chem. Soc. 2013, 90, 53-64.

20

McNichol, J.; MacDougall, K.M.; Melanson, J.E.; McGinn, P.J. Suitability of Soxhlet extraction to quantify microalgal fatty acids as determined by comparison with in situ transesterification. Lipids 2012, 47, 195-207.

21

Olmstead, I.L.D.; Hill, D.R.A.; Dias, D.A.; Jayasinghe, N.S.; Callahan, D.L.; Kentish, S.E.; Scales, P.J.; Martin, G.J.O. A quantitative analysis of microalgal lipids for optimization of biodiesel and omega-3 production. Biotechnol. Bioeng. 2013, 110, 2096-2104.

22

Olmstead, I.L.D.; Kentish, S.E.; Scales, P.J.; Martin, G.J.O. Low solvent, low temperature method for extracting biodiesel lipids from concentrated microalgal biomass. Bioresour. Technol. 2013, 148, 615-619.

23

Guo, Z.; Vikbjerg, A.F.; Xu, X. Enzymatic modification of phospholipids for functional applications and human nutrition. Biotechnol. Adv. 2005, 23, 203-259.

24

van Hoogevest, P.; Wendel, A. The use of natural and synthetic phospholipids as pharmaceutical excipients. Eur. J. Lipid Sci. Technol. 2014, 116, 1088-1107.

25

Ramluckan, K.; Moodley, K.G.; Bux, F. An evaluation of the efficacy of using selected solvents for the extraction of lipids from algal biomass by the soxhlet extraction method. Fuel 2014, 116, 103-108.

26

dos Santos, R.R.; Moreira, D.M.; Kunigami, C.N.; Aranda, D.A.G.; Teixeira, C.M.L.L. Comparison between several methods for total lipid extraction from Chlorella vulgaris biomass. Ultrason. Sonochem. 2015, 22, 95-99.

27

Montes D’Oca, M.G.; Viêgas, C.V.; Lemões, J.S.; Miyasaki, E.K.; Morón-Villarreyes, J.A.; Primel, E.G.; Abreu, P.C. Production of FAMEs from several microalgal lipidic extracts and direct esterification of the Chlorella pyrenoidosa. Biomass Bioenergy 2011, 35, 1533-1538.



634 635 636 637 638 639 640 641 642 643 644 645 646 647 648 649 650 651 652 653 654 655 656 657 658 659 660 661 662 663 664 665 666 667 668 669 670 671 672 673 674 675 676 677 678 679 680 681 682 683

Page 26 of 37

28

Balasubramanian, R.K.; Allen, J.D.; Kanitkar, A.; Boldor, D. Oil extraction from Scenedesmus obliquus using a continuous microwave system – design, optimization, and quality characterization. Bioresour. Technol. 2011, 102, 3396-3403.

29

Folch, J.; Lees, M.; Stanley, G.H.S. A simple method for the isolation and purification of total lipids from animal tissues. J. Biol. Chem. 1957, 226, 497-509.

30

Ryckebosch, E.; Muylaert, K.; Eeckhout, M.; Ruyssen, T.; Foubert, I. Influence of drying and storage on lipid and carotenoid stability of the microalga Phaeodactylum tricornutum. J. Agric. Food Chem. 2011, 59, 11063-11069.

31

Plaza, M.; Santoyo, S.; Jaime, L.; García-Blairsy Reina, G.; Herrero, M.; Señoráns, F.J.; Ibáñez, E. Screening for bioactive compounds from algae. J. Pharm. Biomed. Anal. 2010, 51, 450-455.

32

Herrero, M.; Cifuentes, A.; Ibañez, E. Sub- and supercritical fluid extraction of functional ingredients from different natural sources: Plants, food-by-products, algae and microalgae. Food Chem. 2006, 98, 136-148.

33

Mendes, R.L.; Nobre, B.P.; Cardoso, M.T.; Pereira, A.P.; Palavra, A.F. Supercritical carbon dioxide extraction of compounds with pharmaceutical importance from microalgae. Inorg. Chim. Acta 2003, 356, 328-334.

34

Mendes, R.L.; Reis, A.D.; Palavra, A.F. Supercritical CO2 extraction of γ-linolenic acid and other lipids from Arthrospira (Spirulina) maxima: Comparison with organic solvent extraction. Food Chem. 2006, 99, 57-63.

35

Guckert, J.B.; Cooksey, K.E.; Jackson, L.L. Lipid solvent systems are not equivalent for analysis of lipid classes in the microeukaryotic green alga, Chlorella. J. Microbiol. Methods 1988, 8, 139-149.

36

Mulbry, W.; Kondrad, S.; Buyer, J.; Luthria, D.L. Optimization of an oil extraction process for algae from the treatment of manure effluent. J. Am. Oil Chem. Soc. 2009, 86, 909-915.

37

Balasubramanian, R.K.; Doan, T.T.Y.; Obbard, J.P. Factors affecting cellular lipid extraction from marine microalgae. Chem. Eng. J. 2013, 215-216, 929-936.

38

Ferraz, T.P.L.; Fiúza, M.C.; dos Santos, M.L.A.; Pontes de Carvalho, L.; Soares, N.M. Comparison of six methods for the extraction of lipids from serum in terms of effectiveness and protein preservation. J. Biochem. Biophys. Methods 2004, 58, 187-193.

39

Crampon, C.; Mouahid, A.; Toudji, S.-A. A.; Lépine, O.; Badens, E. Influence of pretreatment on supercritical CO2 extraction from Nannochloropsis oculata. J. Supercrit. Fluids 2013, 79, 337-344.

40

Dembitsky, V.M. Betaine ether-linked glycerolipids: Chemistry and biology. Prog. Lipid Res. 1996, 35, 1-51.

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Christie, W.W. Betaine lipids: occurrence, biochemistry and analysis. URL (http://lipidlibrary.aocs.org/Lipids/betaine/index.htm (July 2014) 26 ACS Paragon Plus Environment

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Benning, C.; Huang, Z.H.; Gage, D.A. Accumulation of a novel glycolipid and betaine lipid in cells of Rhodobacter sphaeroides grown under phosphate limitation. Arch. Biochem. Biophys. 1995, 317, 103-111.

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Li, M.; Welti, R.; Wang, X. Quantitative profiling of Arabidopsis polar glycerolipids in response to phosphorus starvation. Roles of phospholipases Dζ1 and Dζ2 in phosphatidylcholine hydrolysis and digalactosyldiacylglycerol accumulation in phosphorusstarved plants. Plant Physiol. 2006, 142, 750-761.



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Figure captions

703 704

Figure 1. Total amount of lipids and lipid class composition, obtained by the different extraction

705

methods (mean ± SD; n = 3) from Nannochloropsis oculata (top). In the separate graphs the total

706

chromatographic peak area of NL, GL and PL fractions are shown (mean ± SD; n = 3). (CHL:

707

chloroform; MeOH: methanol; EtOH: ethanol)

708 709

Figure 2. Total Ion Current (TIC) chromatogram of the GL fraction for the PFE extraction with

710

chloroform/methanol (2:1). In the separate graph the total chromatographic peak area is shown

711

(mean ± SD; n = 3). The chemical structure of the compounds identified in the GL fractions is

712

shown. The ESI(+) mass spectrum of a compound is given as an example. (CHL: chloroform;

713

MeOH: methanol; EtOH: ethanol)

714 715

Figure 3. Relative distribution, expressed as % of total peak area, of the different MGDG and

716

DGDG in the GL fraction. The results are the average of 3 replicate measurements (mean ± SD).

717

(CHL: chloroform; MeOH: methanol; EtOH: ethanol)

718 719

Figure 4. Total Ion Current (TIC) chromatogram of the PL fraction for the PFE extraction with

720

chloroform/methanol (2:1). In the separate graph the total chromatographic peak area is shown

721

(mean ± SD; n = 3). The chemical structure of the compounds identified in the PL fractions is

722

shown. The ESI(+) mass spectrum of a compound is given as an example. (CHL: chloroform;

723

MeOH: methanol; EtOH: ethanol)

724


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725

Figure 5. Relative distribution over different DGTS and lyso-DGTS, expressed as % of total

726

peak area of DGTS and lyso-DGTS, respectively. Results are the average of 3 replicate

727

measurements (mean ± SD). (CHL: chloroform; MeOH: methanol; EtOH: ethanol)

728 729

Figure 6. The NL fractions: (a) Total Ion current chromatogram of the PFE extraction with

730

chloroform/methanol (2:1); (b) MS spectrum of astaxanthin glucoside as an example; (c)

731

chemical structure of the compounds identified.



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Table 1. Total Concentration, Expressed as % w/w, of Phosphatidylcholine (PC) and Relative Distribution, as %, among Different Combinations of Fatty Acid Chains in the Different Polar Lipid Fractions. Results Are the Average of 3 Replicate Measurements (mean ± SD).

Extraction method

Total PC (% w/w)

Hexane

PFE Ethanol

0.14

2.51

CHL/MeOH (2:1) 0.42

Hexane 0.01

Relative distributionb (%) among PCs with different fatty acid chains PC 16:0/16:1 24.6 25.7 26.7 34.0 PC 16:0/18:1 19.0 15.3 19.0 4.2 PC 16:0/18:2 ---1.5 PC 16:0/18:3 16.9 20.9 15.7 3.5 PC 16:0/20:5 6.6 8.3 7.2 9.1 PC 16:1/16:1 7.8 11.9 8.9 14.1 PC 16:1/18:0 ---1.2 PC 16:1/20:5 5.6 5.7 4.5 8.2 PC 16:2/18:2 ---2.7 PC 18:1/16:1 15.3 7.5 14.4 13.3 PC 20:3/16:0 4.1 4.8 3.7 8.3

Soxhlet Folch Ethanol CHL/MeOH CHL/MeOH (2:1) (2:1) 0.48 0.18 0.60

17.7 9.0 0.5 8.4 7.8 20.3 1.6 14.8 2.3 8.7 8.9

a

LOQ of PC analysis: 13 µg/mL

b

calculated using the relative peak area of each PC compound and the total PC concentration

12.9 5.5 1.4 11.3 6.8 26.2 4.7 10.8 6.5 6.4 7.6

18.9 10.6 -6.4 11.1 15.9 -15.9 -9.8 11.5

SFE scCO2/30% EtOH < LOQa 0.33 scCO2

------------

27.3 17.6 -23.7 8.7 4.2 -7.3 -6.7 4.6

30

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Figure 1.

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Figure 2.

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Figure 3.

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Figure 4.

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Figure 5.

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Figure 6.

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Table of contents graphics


Polar Lipid Profile of Nannochloropsis oculata Determined Using a

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