Open Access
Issue
OCL
Volume 24, Number 4, July-August 2017
Article Number D405
Number of page(s) 6
Section Lipids of the future / Lipides du futur
DOI https://doi.org/10.1051/ocl/2017011
Published online 26 April 2017

© T. Mnasri et al., published by EDP Sciences, 2017

Licence Creative Commons
This is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/4.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

1 Introduction

Lysoglycerophospholipids (or lysophospholipids, LPLs) are glycerol-based lipids containing one fatty acyl moiety at either the sn1 or sn2 position and a phosphate group at the sn3 position. Except for lysophosphatidic acid (LPA), the phosphate group is esterified to an alcohol or amino-alcohol (mainly choline, ethanolamine, inositol and serine) leading to various LPL species, such as lysophosphatidylcholine (LPC), the most abundant one in nature, or lysophosphatidylethanolamine (LPE). Depending on the acylated position, two isomers are distinguished, namely 1-acyl-2-lyso-phospholipid (or 2-LPL) and 1-lyso-2-acyl-phospholipid (or 1-LPL). For example, the structures of 1-LPC and 2-LPC isomers are shown in Figure 1.

LPA has first attracted attention as an ubiquitous mediator in cellular signal transduction (Valentine et al., 2008; Lin et al., 2010). However, other LPL species have been studied since then and are also now recognized as essential bioactive lipids involved in a large variety of both normal and pathological processes such as carcinogenesis, neurogenesis, immunity, vascular development or regulation of metabolic diseases (Grzelczyk and Gendaszewska-Darmach, 2013). Moreover, LPLs circulate in the plasma mainly bound to albumin and their concentration can significantly vary in relation with some diseases. They are thus considered as potential biomarkers for the early detection of ovarian cancer (Fan et al., 2016) or colorectal cancer (Zhao et al., 2007).

Interestingly, most often LPL effects are remarkably acyl chain-dependent, i.e. related to acyl-chain length and degree of unsaturation (Brkić et al., 2012; Rao et al., 2013). For instance, among the various species of LPCs in plasma, 16:0-, 18:1- and 20:4-LPC induce an increase of endothelial prostacyclin production in vitro (1.4-, 3- and 8.3-fold, respectively) although C18:2-LPC is inactive (Riederer et al., 2010). Moreover, several studies have contributed to show that LPLs are convenient carriers for optimal transport of docosahexaenoic acid (DHA, C22:6 ω3) to the brain where it plays a number of important functions (Bernoud et al., 1999; Lagarde et al., 2015). DHA-LPLs have also been proved to be anti-inflammatory (Huang et al., 2010; Hung et al., 2011) and anti-angiogenic (Tsushima et al., 2012) lipids.

LPLs have also industrial and pharmaceutical uses such as emulsifiers and wetting agents for the food-processing (Kasinos et al., 2014) and cosmetic industries (Yahagi et al., 2011), components of liposomes for use in drug delivery (Koklic and Trancar, 2012) and adjuvant in vaccines (Cmielewski et al., 2010).

Due to their diverse biological roles and industrial applications, efficient methods for production of LPLs are necessary. In literature, various chemical, enzymatic or chemo-enzymatic ways for LPL synthesis have been described and reviewed (D'Arrigo and Servi, 2010; Pencreac'h et al., 2013). Enzymatic pathways, using lipases or phospholipases, are of particular interest due to their stereoselective activities although building the lyso-PL chiral structure is a crucial issue when using chemical routes. Moreover, enzymatic reactions usually occur under milder conditions, mainly lower temperatures, as compared to chemical reactions.

Lipases (triacylglycerol acyl hydrolases, E.C.3.1.1.3) are ubiquitous enzymes which natural function is to catalyze the hydrolysis of ester bonds in long chain triacylglycerols. They are widely distributed among animal, plant and microbial kingdoms. Lipases are classified into 2 groups regarding their regioselectivity: 1,3-regioselective lipases preferentially catalyze hydrolysis of ester bonds at the 1 and 3 positions in triacylglycerols, and non-specific lipases catalyze the hydrolysis of the three ester bonds similarly. Interestingly, in low aqueous conditions, such as in organic solvent, ionic liquid or supercritical CO2, microbial lipases also efficiently catalyze the reverse reaction, i.e. ester bond synthesis. Moreover, lipases display broad substrate specificity and catalyze a large number of various reactions (Kapoor and Gupta, 2012). All these reasons make lipases one of the most widespread biocatalysts used in biotechnological applications. Many lipases from bacterial and fungal sources are commercially available from several suppliers under either free or immobilized form.

Particularly, lipases are widely used in the field of enzymatic modifications of phospholipids. Indeed, they are efficient biocatalysts for hydrolysis of ester bonds (Haas et al., 1994; Hara et al., 1997; Gutiérrez-Ayesta et al., 2007) or acyl chain exchange in phospholipids (Haraldsson and Thorarensen, 1999; Vikbjerg et al., 2005).

This paper describes the various ways using lipases for the production of LPLs with special emphasis on our works dealing with the production of DHA-LPLs via hydrolysis of DHA-rich phospholipids extracted from the microalga Isochrysis galbana.

thumbnail Fig. 1

Structures of lysophosphatidylcholine isomers (R = acyl chain).

2 Lipase-catalyzed deacylation of phospholipids

LPLs can be produced via enzyme-catalyzed partial deacylation of phospholipids, i.e. hydrolysis of one of the two ester bonds of phospholipids. Most often, the enzyme used for this purpose is phospholipase A2 and the phospholipidic substrates are natural phospholipids, the so-called lecithins, from soybean (Aura et al., 1994), sunflower (Cabezas et al., 2012) or egg yolk (Kim et al., 2001). In these cases, the acyl chain released is the one linked to the sn2 position due to the sn2 regioselectivity of phospholipase A2.

However, in some cases, it is advantageous to deacylate the sn1 position rather than the sn2 position of phospholipids using 1,3-regioselective lipases. This occurs when phospholipids contain fatty acids with health or industrial benefit linked to the sn2 position. The sn1 specific deacylation therefore allows the production of 1-LPL rich in the desired fatty acids. Several studies have implemented this approach for the production of DHA-rich LPLs. Indeed, DHA moiety in DHA-containing phospholipids from natural sources is almost exclusively located at the sn2 position (Farkas et al., 2000; Chen and Li, 2007). Thus, the 1,3-regioselective lipase from Rhizomucor miehei (Lipozyme RM-IM®, Novozymes, Denmark) has been used to catalyze the hydrolysis of DHA-rich phospholipids extracted from squid skin (Ono et al., 1997; Tsushima et al., 2012) or from egg yolk enriched in DHA obtained from fish oil-fed hens (Ono et al., 1997; Hosokawa et al., 1998). In our laboratory, we have shown that I. galbana (strain 927/1, CCAP, Ambleside, UK), a well-known DHA-rich marine microalga, is a suitable source of phospholipids for the lipase-catalyzed preparation of DHA-LPLs. Indeed, in this microalga, 75% of the DHA contained in total lipids are contained in the phospholipidic fraction and DHA represents 50% of the total fatty acid profile of phospholipids. Moreover, DHA is actually located at the sn2 position of the glycerol backbone (Devos et al., 2006).

In order to achieve lipase-catalyzed partial hydrolysis of DHA-rich phospholipids, the main challenge is to select a lipase that releases all acyl chains but DHA from the glycerol backbone i.e. which discriminates against DHA. Among twelve lipases tested, the immobilized lipases from Rhizopus oryzae (Lipase F AP-15®) and Mucor javanicus (Lipase M®), provided by Amano Enzymes (Japan), efficiently catalyze ester bond hydrolysis in phospholipids with, however, low DHA release. Both lipases are 1,3-regioselective towards triacylglycerols although Lipase M® may also slightly release some fatty acids from the sn2 position (technical data sheets from the supplier). The time course of the reaction with Lipase F AP-15® is shown in Figure 2 (from Devos et al., 2006). It can be seen that DHA ratio among total phospholipid fatty acids increased rapidly, from 50% to around 70% within the first 15 min of reaction, and afterwards more slowly to reach 77% after 3 h of reaction. Regarding DHA recovery, i.e. recovery of DHA initially present in phospholipids, a decrease was observed down to 85% during the first hour of reaction with no further modification with time.

Further experiments were performed in view to determine whether the DHA chain remains located at the sn2 position or migrates spontaneously to the sn1 position as presumed. Indeed, 2-LPLs are thermodynamically more stable than 1-LPLs and intramolecular acyl migration from the sn2 to the sn1 position was shown to readily occur (Plückthun and Dennis, 1982). In our case, a multi-step enzymatic method was developed to address this issue (Poisson et al., 2009). The LPLs produced by the selective hydrolysis of I. galbana phospholipids with Lipase F AP-15® were further hydrolyzed using a phospholipase A2. No DHA release was observed proving that no DHA was attached to the sn2 position in LPLs. The LPLs were then hydrolyzed using the Pseudomonas aeruginosa lipase (Lipase PS, Amano Enzyme Europe, UK), which is known to be non-specific (Devos et al., 2006). In this case, DHA was efficiently released proving that DHA had actually migrated from the sn2 to the sn1 position. Figure 3 represents the general scheme of 1,3-regioselective lipase-catalyzed hydrolysis of phospholipids, illustrated with phosphatidylcholine, taking into account the acyl migration occurring in the LPCs produced.

Interestingly, the observation that the Lipase F AP-15® actually discriminates towards DHA even though DHA chains migrate to the sn1 position indicates that this lipase displays a clear typoselectivity against DHA.

thumbnail Fig. 2

Time course of DHA enrichment of LPC via selective hydrolysis of DHA-rich phospholipids from the microalga Isochrysis galbana.

thumbnail Fig. 3

Reaction scheme of 1,3-regioselective lipase-catalyzed hydrolysis of phosphatidylcholine.

3 Lipase-catalyzed acylation of glycerophosphoryl moiety

Besides partial hydrolysis of phospholipids, lipases have also been used to synthesize LPA, LPC and LPE via acylation of one of the two hydroxyl groups of glycerophosphatidic acid (GPA, disodium salt form), glycerophosphocholine (GPC) and glycerophosphoethanolamine (GPE), respectively. The reactions were implemented in low-aqueous environment in order to favour the synthesis reaction vs. the hydrolysis reaction. The acyl donors were either free fatty acids for direct esterification reactions (Han and Rhee, 1995; Virto et al., 1999; Kim and Kim, 2000; Hong et al., 2011) or fatty acid vinyl esters for transesterification reactions (Virto et al., 1999; Virto and Adlercreutz, 2000). Figure 4 represents, as an example, the scheme of GPC esterification with free fatty acids.

From these studies, it is clear that lipases, and more particularly immobilized lipases, are suitable catalysts for such reactions. For the esterification of GPC with free palmitic acid, two immobilized lipases, namely R. miehei lipase and Candida antarctica lipase B (Lipozyme RM-IM® and Novozym 435®, respectively, Novozymes, Denmark,) were shown to be efficient catalysts with 68% and 42% of GPC conversion to LPC, respectively (Kim and Kim, 1998). Among three free lipases tested for the same reaction, Aspergillus lipase (Lipase AP-6®, Amano Enzymes, Japan) offered similar activity (50%) as the immobilized lipases although the other two, namely R. oryzae lipase and R. niveus lipase (Lipase F-AP15® and Lipase N®, respectively, Anamo Enzymes, Japan) displayed lower activities (34–36%). Hong et al. (2011) also compared several enzymes for the esterification of GPC with conjugated linoleic acids: three immobilized lipases (Novozym 435®, Lipozyme TL-IM®, Lipozyme RM-IM®), a free phospholipase A1 (Lecitase Ultra®, Novozymes, Denmark) and a free phospholipase A2 (Lecitase 10L®, Novozymes, Denmark). Interestingly, the two phospholipases are less effective (<10% GPC conversion) than the immobilized lipases in the reaction conditions used (approximately 40%, 18% and 15%, respectively).

Regarding the implementation of these reactions, trials were first performed by mixing the substrates and the enzyme in various organic solvents commonly used as non-aqueous media for lipase-catalyzed reactions (Han and Rhee, 1995; Kim and Kim, 2000). It was observed that the esterification reaction proceeded very slowly, or even did not proceed at all depending on the solvent used, due to the insolubility of GPA (Han and Rhee, 1995) and GPC (Kim and Kim, 2000) in the solvent. Indeed, GPA and GPC are highly polar molecules nearly insoluble in organic solvents such as hexane, 2-methyl-2-butanol or acetonitrile. Experiments have therefore been further performed in solvent-free systems in which the free fatty acids or fatty acid esters form the liquid phase. However, GPC solubilisation in these conditions was still not complete. Some solid GPC particles were still visible in the reaction mixture and they slowly solubilized as the reaction proceeded (Kim and Kim, 2000). Similarly, GPA was reported to form a separated gel phase or to be dispersed in the media depending on the fatty acid used (Virto et al., 1999).

In acylation reactions in low-aqueous environment, another crucial parameter is the water present in the reaction medium (Stergiou et al., 2013). Water is indeed essential to maintain the adequate hydration state of the catalyst for maximal activity. Moreover, in direct esterification reaction, water is a product of the reaction which can shift the equilibrium towards hydrolysis if not removed from the reaction medium. This is why, in studies dealing with the esterification of GPA or GPC, water amount is controlled by either performing the reaction in open reactors (Han and Rhee, 1995), under low pressure (Hong et al., 2011), adding a convenient co-solvent such as dimethylformamide (Kim and Kim, 2000) or buffering the water amount during the reaction by the use of salt hydrate pairs (Han and Rhee, 1998).

Regarding the acyl moiety of the synthetized LPLs, most are saturated chains. Capric (C10:0), lauric (C12:0), myristic (C14:0) and palmitic (C16:0) fatty acids (Han and Rhee, 1998; Kim and Kim, 2000) and their vinyl ester counterparts (Virto and Adlercreutz, 2000) have thus been studied as acyl donors. In comparison, to our knowledge, only one study has specifically focused on the synthesis of LPLs with an unsaturated chain, namely linoleoyl chain (Hong et al., 2011). Oleic acid has also been envisaged as an acyl donor for the esterification of GPA in comparison with lauric acid vinyl ester (Virto et al., 1999). In our laboratory, experiments have shown that oleoyl-LPC can be efficiently produced by direct esterification of GPC and free oleic acid.

Finally, the reaction parameters such as molar ratio of substrates, enzyme amount and reaction temperature markedly influence the catalytic activity of the lipase used. In optimized reaction conditions, high yields were obtained as claimed in above mentioned studies.

thumbnail Fig. 4

Reaction scheme of 1,3-regioselective lipase-catalyzed esterification of glycerophosphocholine with free fatty acid.

4 Lipase-catalyzed alcoholysis of phospholipids

Lipase-catalyzed alcoholysis of phospholipids has also been implemented in some studies for the preparation of LPLs. As an example, the reaction scheme for LPC synthesis is presented in Figure 5.

This approach was first developed in 1994 using Lipozyme IM-60® (immobilized lipase from M. miehei, Novozymes, Denmark) (Sarney et al., 1994). Other lipases tested were inactive in the conditions used. The initial phospholipid, namely synthetic dipalmitoyl phosphatidylcholine, was dissolved in 95/5 alcohol/water. The alcohol therefore serves both as substrate of the reaction and as the liquid phase of the reaction medium. When ethanol, 2-propanol or 1-butanol were used, high conversion yields were achieved (>95%) although no significant activity was detected with methanol. Interestingly, in these conditions, only 1-LPC was produced suggesting that no acyl migration occurred. Moreover the authors highlighted some advantages of alcoholysis vs. hydrolysis of phospholipids such as the homogeneous reaction mixture thus simplifying the overall process control and allowing continuous operation.

Lipase-catalyzed alcoholysis of natural phospholipids from soybean was also investigated (Ghosh and Bhattacharyya, 1997) using Lipozyme IM-20® (immobilized lipase from M. miehei, Novozymes, Denmark) and various short- and long-chain alcohols (C4 to C18). The aim of the authors was to simultaneously produce LPLs and fatty acid esters of individual alcohols, the latter having also important industrial applications. The reaction yields for both desired products were higher than 70% whatever the alcohol used.

More recently, Lipozyme TL-IM® and Novozym 435® were shown to catalyze ethanolysis of purified phosphatidylcholine as efficiently as Lipozyme RM-IM® (Yang et al., 2015). Reaction was carried out either directly in ethanol as in studies mentioned above or in hexane where phosphatidylcholine and ethanol were dissolved. In both systems, high LPC yields (>90%) were reached with the three lipases tested, Novozym 435® allowing the highest yields (about 98%) with shorter reaction times (6 h and 4 h in ethanol and hexane, respectively).

thumbnail Fig. 5

Reaction scheme of 1,3-regioselective lipase-catalyzed alcoholysis of phosphatidylcholine.

5 Conclusion

Lipases have been shown to be efficient biocatalysts for LPL synthesis mainly via hydrolytic reactions or direct ester bond synthesis reactions. Interestingly, the substrates for lipase-catalyzed synthesis of LPLs such as vegetable lecithins for hydrolysis and GPC and free fatty acids for esterification, are easily available materials. Commercial lecithins are even by-products of the edible oil processing industry. Therefore, lipase-catalyzed methodologies for LPL synthesis are consistent with the concept of biomass biorefinery for the production of active molecules and ingredients widely developed nowadays.

Acknowledgements

This work was financially supported by “Laval Agglomération” and the “Conseil Général de la Mayenne”.

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Cite this article as: Mnasri T, Hérault J, Gauvry L, Loiseau C, Poisson L, Ergan F, Pencréac'h G. 2017. Lipase-catalyzed production of lysophospholipids. OCL 24(4): D405.

All Figures

thumbnail Fig. 1

Structures of lysophosphatidylcholine isomers (R = acyl chain).

In the text
thumbnail Fig. 2

Time course of DHA enrichment of LPC via selective hydrolysis of DHA-rich phospholipids from the microalga Isochrysis galbana.

In the text
thumbnail Fig. 3

Reaction scheme of 1,3-regioselective lipase-catalyzed hydrolysis of phosphatidylcholine.

In the text
thumbnail Fig. 4

Reaction scheme of 1,3-regioselective lipase-catalyzed esterification of glycerophosphocholine with free fatty acid.

In the text
thumbnail Fig. 5

Reaction scheme of 1,3-regioselective lipase-catalyzed alcoholysis of phosphatidylcholine.

In the text

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