Research Collection
Doctoral Thesis
Enzymatic Synthesis and Hydrolysis of Linear Alkyl and Steryl Hydroxycinnamic Acid Esters
Author(s): Schär, Aline Lea
Publication Date: 2016
Permanent Link: https://doi.org/10.3929/ethz-a-010670414
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ETH Library DISS. ETH NO. 23265
Enzymatic Synthesis and Hydrolysis of Linear Alkyl and Steryl Hydroxycinnamic Acid Esters
A thesis submitted to attain the degree of
DOCTOR OF SCIENCES of ETH ZURICH
(Dr. sc. ETH Zurich)
presented by
Aline Lea Schär
MSc ETH in Food Science, ETH Zurich
born on 24.01.1987
citizen of Madiswil (BE)
accepted on the recommendation of
Prof. Dr. Laura Nyström, examiner
Dr. Pierre Villeneuve, co-examiner
Prof. Dr. Evangelos Topakas, co-examiner
2016
Wir treten auf. Wir spielen. Wir treten ab.
Moritz Leuenberger
Abstract
Phenolic acids are natural antioxidants found widely in the plant kingdom in various forms. In the focus of this thesis were hydroxycinnamic acids, namely ferulic acid, caffeic acid, sinapic acid and p-coumaric acid. In multiphase food systems, the polarity of the phenolic antioxidant is a crucial property, which can be adjusted through esterification. Nowadays, an enzymatic procedure is often preferred for this purpose over a chemically catalyzed reaction. However, a phenolic hydroxyl group in para-position in combination with an unsaturated side chain makes enzymatic esterification of hydroxycinnamic acids by lipases challenging. Since this is the case for the hydroxycinnamic acids mentioned above, it is of interest to find efficient ways to enzymatically esterify them.
Using the immobilized lipase from R. miehei, the esterifications of ferulic acid with ethanol and decanol in n-hexane were optimized applying surface response methodology. With an incubation time of 72 hours, the yields for ethyl ferulate and decyl ferulate were 76% and 88%, respectively. Furthermore, esters of primary alcohols and ferulic acid with varying chain lengths from C2 to C18 were synthesized, yielding 76% to 92% ferulate esters. The ethylations of other hydroxycinnamic acids were also optimized; leading to the conclusion that, for R. miehei lipase, two phenolic hydroxyl groups strongly decrease the yield and a saturated side chain strongly increases the esterification yield of hydroxycinnamic acid derivatives. Overall, the lipase from R. miehei proved to be an efficient catalyst for the esterification of hydroxycinnamic acids with ethanol.
Other than linear alkyl esters, steryl phenolates are also prominent examples of lipophilic hydroxycinnamic acid esters. The plant sterol part of the molecule esterified to phenolic acid brings cholesterol lowering properties as an additional health benefit. Rice bran is often used as source for steryl phenolates extraction, which leads to a limited sterol and phenolic acid pattern available. Therefore, we investigated a simple enzymatic esterification method to produce steryl ferulates. We optimized the direct esterification of ferulic acid and the transesterification of ethyl ferulate, yielding steryl ferulates. The lipase from C. rugosa was used as catalyst for these reactions. Yields of 35% and 55% for the direct esterification and transesterification, respectively, were measured after five days of incubation, both following a similar time course. For other hydroxycinnamic acids, the transesterification yields were significantly lower, especially in the case of a hydroxyl group in para-position without a neighboring methoxy group. The evaluation of the antioxidant activity of steryl hydroxycinnamates in comparison to their linear C18 esters leads to the conclusion that esterification to the sterol does not necessarily improve their antioxidant activity. Overall, an
i enzymatic synthesis of steryl ferulates was investigated and other hydroxycinnamic acids were evaluated as substrates for C. rugosa lipase.
One important group of enzymes in the metabolism of hydroxycinnamic acids are feruloyl esterases. They are well known for their ability to release ferulic acid from polar plant cell wall components but little is known about their capability of hydrolyzing nonpolar ferulates. The previously synthesized alkyl ferulates were therefore evaluated as substrates for four feruloyl esterases and a control lipase. A decrease in the kinetic constants Km and kcat was observed for an increasing lipophilicity of the ferulic acid esters. Moreover, only one feruloyl esterase from C. thermocellum and the lipase showed hydrolytic activity against the linear C18 alkyl ferulate. It is therefore suggested that feruloyl esterases are not able to hydrolyze nonpolar ferulate esters.
This study provides simple and efficient methods for the enzymatic esterification of ferulic acid with sterols and linear alcohols including ethanol. Moreover, hydroxycinnamic acids were esterified and transesterified using the lipases from R. miehei and C. rugosa, revealing very different activity profiles towards hydroxycinnamic acids. For further improvements enzyme engineering may offer an approach to achieve more efficient and better applicable processes. Overall, the enzymatic synthesis is a promising solution to generate steryl phenolates, which can be used as standards, substrates for research, and finally as food additives.
ii Zusammenfassung
Phenolcarbonsäuren sind natürlich Antioxidantien und im Pflanzenreich in verschiedenen Formen weit verbreitet. Im Fokus dieser Arbeit standen Hydroxyzimtsäuren, nämlich Ferulasäure, Kaffeesäure, Sinapinsäure und p-Coumarsäure. In mehrphasigen Lebensmittelsystemen spielt die Polarität der phenolischen Antioxidantien eine zentrale Rolle, welche durch Veresterung entsprechend angepasst werden kann. Heutzutage wird ein enzymatisches Verfahren oft bevorzugt gegenüber einer chemisch katalysierten Reaktion. Jedoch erschwert eine phenolische Hydroxygruppe in der para-Position in Kombination mit einer ungesättigten Seitenkette die enzymatische Veresterung von Hydroxyzimtsäuren durch Lipasen. Dies ist der Fall für die bereits genannten Hydroxyzimtsäuren. Es ist darum von grossem Interesse effiziente enzymatische Veresterungen für Hydroxyzimtsäuren zu entwickeln.
Die Veresterungen von Ferulasäure mit Ethanol und Decanol durch die immobilisierte R. miehei Lipase wurden optimiert mit einer Response Surface Methode. Innerhalb von 72 Stunden waren die Ausbeuten für Ethylferulat 76% und für Decylferulat 88%. Des Weiteren wurden primäre Alkohole mit verschiedenen Kettenlängen von C2 bis C18 mit Ferulasäure verestert. Die Ausbeuten in diesen Experimenten betrugen von 76% bis 92%. Die Ethylierung von anderen Hydroxyzimtsäuren wurden ebenso optimiert. Dies führte zu der Schlussfolgerung, dass für die R. miehei Lipase bei der Veresterung von Hydroxyzimtsäuren zwei phenolische Hydroxygruppen die Ausbeute stark reduzieren und eine gesättigte Seitenkette die Ausbeute deutlich erhöht. Insgesamt ist die R. miehei Lipase ein effizienter Katalysator für die Veresterung von Hydroxyzimtsäuren mit Ethanol.
Nebst linearen Alkylestern sind Sterylphenolate bedeutende Beispiele von lipophilen Hydroxyzimtsäureestern. Der mit der Phenolcarbonsäure veresterte Pflanzensterolteil bringt eine cholesterinsenkende Wirkung als zusätzlichen Gesundheitsnutzen. Reiskleie dient oft als Ausgangsmaterial für die Extraktion von Sterylphenolaten, was zu einem limitierten Sterol- und Phenolcarbonsäureprofil führt. Deshalb wurde eine einfache enzymatische Methode zur Herstellung von Sterylferulaten entwickelt. Dazu wurden die direkte Veresterung der Ferulasäure und die Umesterung von Ethylferulat zu Sterylferulaten optimiert. Die C. rugosa Lipase katalysierte diese Reaktionen. Nach einer Inkubationszeit von fünf Tagen wurde für die Veresterung eine Ausbeute von 35% und für die Umesterung eine Ausbeute von 55% erreicht. Beide Reaktionen verliefen ähnlich über die Reaktionszeit. Für andere Hydroxyzimtsäuren waren die Umesterungsraten deutlich tiefer, speziell wenn sich die phenolische Hydroxygruppe in para-Position befand ohne eine benachbarte
iii Methoxygruppe. Die Ermittlung der antioxidativen Wirkung der Hydroxyzimtsäuresterole im Gegensatz zu ihren linearen C18 Estern zeigte, dass die Veresterung mit Sterolen nicht zwingend zu einer erhöhten antioxidativen Wirkung führt. Zusammenfassend, es wurde eine enzymatische Synthese für Sterylferulate entwickelt und andere Hydroxyzimtsäuren konnten als Substrate für die C. rugosa Lipase evaluiert werden.
Eine bedeutende Gruppe von Enzymen im Metabolismus von Hydroxyzimtsäuren sind Ferulasäure-Esterasen. Sie sind dafür bekannt, dass sie die Fähigkeit besitzen Ferulasäure von polaren Pflanzenzellwandbestandteilen freizusetzen. Jedoch ist wenig bekannt über ihre Fähigkeit auch apolare Ferulate zu hydrolysieren. Die bereits synthetisierten Alkylferulate wurden als Substrate für vier Ferulasäure-Esterasen und eine Kontrolllipase analysiert. Eine
Verminderung von den kinetische Konstanten Km und kcat konnte beobachtet werden mit einer steigenden Lipophilie. Für das lineare C18 Alkylferulat konnte nur mit der C. thermocellum Ferulasäure-Esterase und mit der Kontrolllipase hydrolytische Aktivität verzeichnet werden. Diese Beobachtungen führen zur Schlussfolgerung, dass Ferulasäure- Esterasen nicht in der Lage sind apolare Ferulasäureester zu hydrolysieren.
Diese Studie stellt einfache und effiziente Methoden zur enzymatischen Veresterung von Ferulasäure mit Sterolen und linearen Alkoholen inklusive Ethanol vor. Zudem wurden Hydroxyzimtsäuren verestert und umgeestert mit R. miehei und C. rugosa Lipasen, was ein sehr unterschiedliches Aktivitätsprofil gegenüber Hydroxyzimtsäuren aufzeigte. Für weitere Verbesserungen könnte Enzym-Engineering einen Ansatz bieten um effizientere und besser anwendbare Prozesse zu erreichen. Abschliessen ist die enzymatische Synthese ein vielversprechender Ansatz um den Bedarf an Sterylphenolaten zu decken, welche benötigt werden als Standards, Ausgangsmaterial für weitere Forschung und schliesslich als Lebensmittelzusatzstoffe.
iv Contents
Abstract ...... i
Zusammenfassung ...... iii
Contents ...... v
Introduction ...... 1
PART A - Review of Literature ...... 3
1 Hydroxycinnamic acids ...... 3
1.1 Structure ...... 3
1.2 Occurrence in plants ...... 4
1.2.1 Alkyl hydroxycinnamates...... 5
1.2.2 Steryl hydroxycinnamates ...... 6
1.3 Antioxidant activity of hydroxycinnamic acids ...... 8
1.4 Bioavailability and health benefits ...... 9
2 General enzymatic reactions ...... 12
2.1 Kinetics of enzymatic reactions ...... 12
2.2 Lipase catalysis in organic solvent ...... 13
2.3 Properties of lipases used for lipophilization reactions ...... 15
3 Enzymatic lipophilization of hydroxycinnamic acids ...... 17
3.1 Using lipases ...... 17
3.2 Using other enzymes ...... 22
4 Esterification of phytosterols ...... 25
4.1 Enzymatic phytosterol fatty acid esters synthesis ...... 25
v
4.2 Steryl phenolates ...... 29
4.2.1 Chemical synthesis ...... 29
4.2.2 Chemoenzymatic synthesis ...... 31
4.2.3 Enzymatic synthesis ...... 33
5 Feruloyl esterases ...... 35
5.1 Occurrence in nature ...... 35
5.2 Classification ...... 36
5.3 Hydrolysis of nonpolar substrates ...... 37
References ...... 39
Part B - Research Papers ...... 53
High yielding and direct enzymatic lipophilization of ferulic acid using lipase from Rhizomucor miehei ...... 55
Enzymatic synthesis of steryl ferulates ...... 75
Enzymatic synthesis of steryl hydroxycinnamates and their antioxidant activity .... 97
Hydrolysis of nonpolar n-alkyl ferulates by feruloyl esterases ...... 117
Conclusion ...... 131
Outlook ...... 132
Acknowledgements ...... 134
vi Introduction
Introduction
Hydroxycinnamic acids can be found widely amongst plants and are known for their antioxidant activity and are therefore attributed to the prevention of chronic diseases including cancer and cardiovascular disease (Zhao & Moghadasian, 2010). Amongst cereal grains ferulic acid, a hydroxycinnamic acid, is the most common one and can be found in free form, solubly conjugated, or insolubly bound form (Manach et al., 2004; Shahidi & Chandrasekara, 2009). As part of the solubly conjugated form, various alkyl ferulates occur naturally. Ethyl ferulate was detected in wine and in sake and a homologous series of C16-C30 ferulates can be found in suberin waxes, an extractable part of suberized cells in plants (Graça, 2010; Hashizume et al., 2013b; Hixson et al., 2012). One special alky phenolate is the steryl phenolate. The phenolic acid is in this case esterified to a plant sterol. Steryl ferulates can be mainly found in cereal grains such as rice, wheat and corn (Mandak & Nyström, 2012). Due to the phytosterol part, cholesterol lowering properties are associated to these compounds (Wilson et al., 2007). As the sterol pattern is limited in rice, the most common source, an enzymatic synthesis for further research and later on food application would be of great interest.
For food and pharmaceutical applications the use as antioxidants is of high importance. The polarity displays a major property of the antioxidant, especially when applying in multiphase systems. This property of phenolic acids can be adjusted through esterification with a polar or nonpolar compound and this esterification can be achieved through chemical or enzymatic catalysis. The enzymatically catalyzed reactions are known for being more environmental friendly, as they are more specific and fewer solvents are required for purification and overall the use of non-toxic catalysts is a plus. However, the esterification of hydroxycinnamic acids has been shown to be a rather challenging esterification for lipases (Figueroa-Espinoza & Villeneuve, 2005). Due to the conjugation of the hydroxyl group with the acid group, the electrophilic center of the carboxylic acid is deactivated (Buisman et al., 1998; Guyot et al., 1997). One approach is to perform a transesterification starting from methyl or ethyl phenolate, where the side product can be simply evaporated (Villeneuve, 2007), thus leading to a two-step enzymatic synthesis.
Moreover, other enzymes than lipases have been already applied for the esterification of phenolic acids, namely feruloyl esterases. This group of enzymes gained of interest as they can improve the saccharification of cereal based products for bioalcohol and animal feed production. Feruloyl esterases can liberate ferulic acid from plant cell wall polysaccharides and make it thus available for other degradative enzymes. Main sources are of microbial
1 Introduction
origin, but also in plants and in the human gut feruloyl esterase activity has been reported (Faulds, 2010). However, if nonpolar alkyl ferulates also display a substrate for feruloyl esterases, has not been researched systematically yet.
It was therefore the aim of this thesis to find an efficient esterification system for the most common hydroxycinnamic acids, mainly ferulic acid. Not only the ethylation, which can be used to produce an intermediate product, but also the esterification with other primary alcohols was aimed for. Further, the fully enzymatic synthesis of steryl hydroxycinnamates should be investigated. Thirdly, the evaluation of feruloyl esterases on their ability to hydrolyze nonpolar alkyl ferulates was of interest.
2 PART A - Review of Literature
PART A - Review of Literature
1 Hydroxycinnamic acids
1.1 Structure
Phenolic acids are composed of an aromatic ring bearing at least one phenolic hydroxyl group and a carboxylic acid attached to the aromatic ring (Figueroa-Espinoza & Villeneuve, 2005). There are two main classes, namely hydroxybenzoic acid derivatives and hydroxycinnamic acid derivatives, which can be differentiated based on the length of the side chain (Figure 1) (Figueroa-Espinoza & Villeneuve, 2005; Manach et al., 2004). While the hydroxybenzoic acid derivatives are composed of a C6-C1 skeleton, the hydroxycinnamic acid derivatives have a C6 – C3 structure.
Figure 1: Examples of the hydroxybenzoic acid derivatives (p-hydroxybenzoic acid, left) and the hydroxycinnamic acid derivatives (p-coumaric acid, right).
The hydroxycinnamic acid derivatives are more common than the hydroxybenzoic acid derivatives (Manach et al., 2004). Moreover, the focus will be on hydroxycinnamic acids as they are in the core of interest in this study. The main representatives of the hydroxycinnamic acid group are ferulic acid, p-coumaric acid, caffeic acid and sinapic acid (Figure 2) (Manach et al., 2004). Their differences are the number and positions of hydroxyl and methoxy groups.
Figure 2: Major hydroxycinnamic acids: ferulic acid, p-coumaric acid, caffeic acid, and sinapic acid (from left to right).
3 Hydroxycinnamic acids
1.2 Occurrence in plants
Hydroxycinnamic acids occur in the plant kingdom widely distributed (El-Seedi et al., 2012). Therefore, they can be found ubiquitously in plant based foods such as fruits, vegetables, cereals, nuts, legumes, oilseeds, and beverages (Shahidi & Chandrasekara, 2009). Ferulic acid is dominant in cereals and caffeic acid in most fruits (Manach et al., 2004). In Brassica vegetables sinapic acid and sinapic acid derivatives are particularly frequent (Nićiforović & Abramovič, 2014). Amongst the coumaric acid derivatives, p-coumaric is most abundant in foods (Shahidi & Chandrasekara, 2009). However, also o-coumaric acid has been reported in foods such as oat and peanut and m-coumaric acid in small berries (Shahidi & Chandrasekara, 2009; Zadernowski et al., 2005). Hydroxycinnamic acids are therefore important phenolics in our diet.
Overall, hydroxycinnamic acids can be found free, conjugated but soluble, and in insoluble- bound form (Shahidi & Chandrasekara, 2009). As summarized by Zhao and Moghadasian the free form is less abundant, simple esters are found in fruits and vegetables, in contrast to cereals where they occur mostly as insoluble esters (Zhao & Moghadasian, 2010). The insoluble-bound hydroxycinnamic acids are covalently linked to structural parts of the plant cell wall i.e. cellulose, lignin, and proteins (Shahidi & Chandrasekara, 2009). In the cytoplasm more commonly the soluble forms are located (El-Seedi et al., 2012). Apart from the alkyl and steryl esters discussed below, many other simple esters appear naturally. Examples are hydroxycinnamic acid amides such as 4-coumaroyltyramine and feruloyltryptamine (Facchini et al., 2002). Further hydroxycinnamic acid amides are avenanthramides in oats, which are esters 5-hydroxyanthranilic acid and one hydroxycinnamic acid (p-coumaric, ferulic acid or caffeic acid) (Shahidi & Chandrasekara, 2009). Sinapine and sinapoyl malate are common esters of sinapic acid, as well as sinapoyl glucose (Nićiforović & Abramovič, 2014). Caffeic acid can appear esterified with a quinic acid, also called chlorogenic acid (El-Seedi et al., 2012). Rosmarinic acid, an ester of caffeic acid and 3,4-dihydroxyphenyllactic acid is also a known form (Shahidi & Chandrasekara, 2009). Finally, a third example of a caffeate is the caffeic acid phenethyl ester (Shahidi & Chandrasekara, 2009). This list could still be extended further, but it already shows the large variability of hydroxycinnamic acid esters in plants.
The biosynthesis of hydroxycinnamic acids in plants has been reviewed recently (El-Seedi et al., 2012). Briefly, phenylalanine and tyrosine are synthesized via the shikimate pathway, starting from phosphoenolpyruvate and erythrose-4-phosphate. These amino acids can be deaminated leading to cinnamic acid and p-coumaric acid. The cinnamic acid can also be
4 Hydroxycinnamic acids
converted into p-coumaric acid. From the p-coumaric acid caffeic acid is synthesized, which is again the precursor of ferulic acid. Finally, sinapic acid is formed from ferulic acid (El-Seedi et al., 2012).
1.2.1 Alkyl hydroxycinnamates
Alkyl hydroxycinnamates are widely present in plants. The name alkyl hydroxycinnamate is rather vague but often associated with esters of hydroxycinnamic acids and primary, acyclic, often saturated and long-chain alcohols. An overview on the occurrence of alkyl hydroxycinnamates in plants has recently been published (He et al., 2015). Mainly studies are listed, which identified ferulic, p-coumaric, or caffeic acid alkyl esters (C14-C32) in the bark, root, or leave fibers (He et al., 2015). Overall, alkyl hydroxycinnamates can be found in suberin and in its associated waxes as summarized by Graça. Suberin is a biopolymer of suberized cells, which are a barrier against water loss. Hydroxycinnamates can be found in the polymeric suberin and in non-polymeric extractable suberin waxes. Apart from linear alkyl ferulates also hydroxycinnamic esters of ω-hydroxyacids and glycerol can be found in depolymerized suberin (Graça, 2010). Recently aliphatic waxes associated to suberized cells of various plants were analyzed on alkyl hydroxycinnamates. Except in carrot roots, in all analyzed samples alkyl hydroxycinnamates were found. The distribution of alkyl ferulates, coumarates and caffeates is very different between the plants. Alkyl chain length was only even numbered and ranging mostly from C18 to C22 with some longer and shorter exception. In rice for example, where only ferulates were found the chain length varied from C20 to C28. On the other hand in sweet potato, for example, all three alkyl hydroxycinnamates were detected (Kosma et al., 2015). Therefore, alkyl hydroxycinnamates are widespread in plants in certain tissues such as the periderm.
The biosynthesis of long-chain alkyl hydroxycinnamates has been assessed and enzymes involved in it have been identified. First, an enzyme from the outermost cell layers of wound- healing potatoes was extracted, which transesterified ferulic acid from feruloyl-CoA in vitro to ω-hydroxyfattyacids and 1-alkanols (C10-C18). Sinapoyl-CoA and p-coumaroyl-CoA were also accepted as substrates, whereas caffeoyl-CoA was not accepted (Lofty et al., 1994). Later on genes encoding for a feruloyl-coenzyme A transferase in Arabidopsis were identified. Almost complete elimination of ester-linked ferulates in association with suberin was found in knockout mutants. Recombinant enzymes catalyzed the transesterification of feruloyl-CoA to ω-hydroxyfatty acids and fatty alcohols (Molina et al., 2009). Later on the identification of an fatty alcohol:caffeoyl-CoA caffeoyl transferase was achieved, showing higher activity towards caffeoyl-CoA than coumaroyl-CoA and feruloyl-CoA (Kosma et al., 2012). This indicates that separate acyl transferases are involved in the biosynthesis of alkyl
5 Hydroxycinnamic acids
hydroxycinnamates, depending on the acid and that an overlap exists between suberin and root wax biosynthesis (Kosma et al., 2012). However, the physiological function of alkyl hydroxycinnamate esters is hardly understood so far (Kosma et al., 2012). Overall, the role of alkyl hydroxycinnamates and their detailed formation and localization in the plant still needs to be elucidated.
Apart from long chain alkyl ferulates also methyl and ethyl esters have been reported in plants or foods. For example methyl sinapate and methyl ferulate were identified in rapeseeds, methyl sinapate being one of the two major phenols (Fang et al., 2012). Further, methyl caffeate and vinyl caffeate were isolated from perilla frutescens leaves and stems (Tada et al., 1996). Although it remains questionable if these compounds could also arise from extraction or working solvents. Finally, ethyl ferulate and ethyl p-coumarate have been quantified in wine and ethyl ferulate in sake (Hashizume et al., 2013b; Hixson et al., 2012). In sake the formation of ethyl ferulate has been associated with a rice koji enzyme (Hashizume et al., 2013a). Apparently also methyl and ethyl hydroxycinnamates can be compounds of our diet.
1.2.2 Steryl hydroxycinnamates
Hydroxycinnamic acid derivatives may also occur as plant sterol esters. The biosynthesis of phytosterols, their biological function and their importance to human nutrition has been reviewed by Piironen and co-workers. Plant sterols are a diverse group containing over 250 different sterols, mostly β-sitosterol, stigmasterol and campesterol. They can be separated into groups by different properties such as the saturation of the ring structure (stanols), the position of the unsaturation (sterols) or the presence of methyl groups (4α-monomethyl sterols and 4,4-dimethyl sterols). Sterols in the plant cell membrane help controlling the fluidity of the membrane with changing temperature. Their biosynthesis occurs via the isoprenoid pathway and as consumed by humans they lower plasma cholesterol and LDL cholesterol (Piironen et al., 2000). Combining hydroxycinnamic acids and phytosterols, sterol phenolic acid esters are therefore promising compounds. The research about their potential health benefits is discussed in another chapter (1.4).
An overview of identified steryl hydroxycinnamates (including steryl cinnamate) is given in Table 1. The main focus was to provide an overview of the most important studies showing a range of plant materials with the main attention on the steryl phenolates varieties. As discussed above, plant sterols are also a diverse group and in combination with hydroxycinnamic acids lead to an even more complex group. However, differences in the sterol profile are not discussed here. Many studies describing contents and composition of
6 Hydroxycinnamic acids
steryl ferulates mainly in rice, but also in wheat and corn are not shown neither and only representative contents are shown.
Steryl ferulates (also known as γ-oryzanol) are the predominant group of steryl hydroxycinnamates (Table 1). The main focus is on cereal grains, as highest steryl ferulates contents are found therein. The content of steryl phenolates in rice and wild rice are very high (262-627 and 850 -1352 μg/g, respectively) compared to for example corn (31-70 μg/g) (Miller & Engel, 2006; Norton, 1995; Seitz, 1989). The content in corn bran is much higher with 70-540 μg/g, which is one example that steryl phenolates are concentrated in the bran of cereal grains. Often no contents are reported in studies where the focus laid on the identification of new compounds.
Table 1: Overview of identified steryl hydroxycinnamates (including steryl cinnamate) and their contents in plants. Steryl hydroxy- Individual Total content Plant materials Reference cinnamates content (μg/g) (μg/g) (Miller & Brown rice Steryl ferulates 262-627b Engel, 2006) Steryl ferulates (Fang et al., Rice bran n.r. n.r. Steryl caffeates 2003) Steryl ferulates (Zhu & Cargo rice, wild rice Steryl p-coumarates n.r. n.r. Nyström, Steryl sinapate 2015) Steryl ferulates 670-1029 (Aladedunye Wild rice kernels Steryl caffeates 79-182 850 -1352 et al., 2013) Steryl cinnamates 73-141 Rye kernels, 0.8a,c 92a wheat kernels, Steryl ferulates 2a,c 142a (Esche et spelt kernels, Steryl p-coumarates 7 Hydroxycinnamic acids Apart from steryl ferulates also steryl p-coumarates, steryl caffeates, steryl cinnamates and one steryl sinapate have been reported (Table 1). In rice steryl caffeates and a steryl sinapate have been reported but not quantified (Fang et al., 2003; Zhu & Nyström, 2015). In wild rice steryl caffeates and steryl cinnamate were quantified and were found in a similar content (79-182 and 73-141 μg/g, respectively), making each up to 10% of steryl phenolate content (Aladedunye et al., 2013). Correctly, steryl cinnamates do not belong to the group of phenolates but are still included in the table and calculation. The steryl p-coumarates on the other hand only make very few percent of total steryl phenolates in rye, wheat, spelt and corn (Esche et al., 2012). In contrast also higher proportion of steryl p-coumarates were reported in corn bran and related fractions reaching up to 11.5% of total steryl phenolates (Norton, 1995) and in a similar range in whole grain corn (Seitz, 1989). To conclude steryl esters of cinnamic acid and all main hydroxycinnamic acids have been reported in plants, although in very different quantities. About the function and biosynthesis of steryl hydroxycinnamates little to no information is available. The biosynthesis of both the hydroxycinnamic acids and sterols are described above. It is proposed that the esterification takes place afterwards; although no enzyme responsible for this reaction has been described so far. It is also possible that the CoA- phenolate is transesterified to the sterol in a similar way as described for long chain alkyl ferulates, in suberin formation (Bernards, 2002). The function of steryl phenolates in the plant has not been revealed so far. Possibly they are not involved in the regulation of fungal activity in the grains (Seitz, 1989) but might be attributed to drought tolerance (Kumar et al., 2014). Still, there is a lack of research in terms of biosynthesis and function of steryl hydroxycinnamates. 1.3 Antioxidant activity of hydroxycinnamic acids Their function as antioxidants is one of the key interests of hydroxycinnamic acids. Due to the phenolic hydroxyl group they have the ability to form stable phenolic free radicals after hydrogen donation. This property makes them to a free radical scavenger and chain breaking antioxidants (Decker, 1998). The antioxidant activity of hydroxycinnamates has been reviewed recently (Shahidi & Chandrasekara, 2009). Generally, amongst major hydroxycinnamates caffeic acid shows the highest and p-coumaric acid the lowest antioxidant activity (Shahidi & Chandrasekara, 2009). In addition to the type of hydroxycinnamic acid the polarity also influences the antioxidant activity strongly. Connected to this property two theories are of interest, the polar paradox and the cutoff effect. The polar paradox was proposed by Porter and co-workers. It states that in nonpolar systems such as 8 Hydroxycinnamic acids bulk oil polar antioxidants show higher antioxidant activity. Whereas in polar systems such as emulsions more nonpolar antioxidants are of higher efficiency (Porter et al., 1989). This phenomenon was later on explained by interfacial oxidation and the presence of colloids also in bulk oil (Chaiyasit et al., 2007). Only a few years ago the so called cutoff effect has been proposed for antioxidants (Laguerre et al., 2009). In emulsified systems there is a nonlinear relationship between the chain length of the antioxidant and the antioxidant activity. First, the activity increases and after reaching a maximum a decrease is observed. This behavior has been attributed to the location of the antioxidant in the system. However, the broad literature evaluating antioxidant activities in various systems with diverse methods will not be discussed here further. The practical applications of hydroxycinnamates as antioxidants are rather few and have been summarized recently (Figueroa-Espinoza et al., 2013). Although the amount of research conducted on hydroxycinnamates is large, only two benzoic acid derivatives and their esters are approved for food application. Namely gallic acid and esters (E310-E312) as antioxidants and p-hydroxybenzoic acid (E214-E219) and its esters and sodium salts as antimicrobial preservatives. Other than that, some ferulates (including ethyl ferulate) are applied as UV filter, skin conditioner or antioxidants. Further, also ethyl caffeate found application as skin conditioner. Due to their cost and availability, the application in foods might be challenging (Figueroa-Espinoza et al., 2013). 1.4 Bioavailability and health benefits The bioavailability of hydroxycinnamates has been reviewed by Zhao and Moghadasian, 2010. Briefly, it is suggested from In situ or ex vivo absorption models that hydroxycinnamic acids are absorbed in the stomach, jejunum, ileum and colon of rats. Generally the absorption efficiencies of ferulic acid and p-coumaric acid are better than the one of caffeic acid. The mechanism of the absorption is not fully elucidated yet. Two have been suggested, passive diffusion but also an H+-driven transport system. However, a large proportion of dietary hydroxycinnamic acids are not provided in free form. It has been shown in rats, that diferulates and rosmarinic acid can be absorbed as intact molecules. But also mucosal esterases in rats were detected, which can liberate ferulic acid from oligosaccharides and microflora enzymes can hydrolyze feruloyl polymers in the colon (Zhao & Moghadasian, 2010). Overall the absorption rate strongly depends on the form in which the hydroxycinnamic acid is provided. The measured bioavailability ranges from 0.4-98% for ferulic acid. Such as from tomatoes a bioavailability of 11-25% was measured. In contrast 9 Hydroxycinnamic acids from cereal products it seems to be below 3% (El-Seedi et al., 2012). The composition of hydroxycinnamates therefore has to be kept in mind when thinking about bioavailability. The potential health benefits of hydroxycinnamates, which are possibly mostly due to their antioxidant activity, have also been reviewed by El Seedi and co-workers. In vitro and animal studies investigate effects such as prevention of cardiovascular diseases, prevention and treatment of cancer, side effect reduction in chemotherapy, antimicrobial activity, and antiosteoclast activity. However, the recorded effects were mostly at rather high concentrations of hydroxycinnamic acid. Epidemiological studies suggest a negative correlation between the consumption of food high in hydroxycinnamic acids (fruits, tea, coffee, and wine) and the occurrence of Alzheimer’s disease and cancer. Further, clinical trials suggest anti-inflammatory and analgesic activities of hydroxycinnamic acids. Apart from their antioxidant activity in food preservation, they may therefore also help to prevent some human disorders (El-Seedi et al., 2012). The main potential health benefit attributed to steryl ferulates is the cholesterol lowering property. Three important studies are discussed here concerning the bioactivity of steryl ferulates. Berger and co-workers conducted a human study with mildly hypercholesterolemic men. Rice bran oil containing γ-oryzanol reduced total plasma cholesterol. However the two evaluated γ-oryzanol concentrations did not show significant difference (Berger et al., 2005). Another study conducted in hamsters showed that γ-oryzanol lowered plasma lipid and lipoprotein cholesterol concentrations and aortic cholesterol ester accumulation. This effect was higher for γ-oryzanol compared to ferulic acid (Wilson et al., 2007). Finally, Lubinus and colleagues evaluated the recovery of steryl ferulates in the feces after human consumption. Almost 80% could be detected intact in the feces. Hydrolyzed sterols and fecal metabolites could only be detected from desmethyl steryl ferulates (Lubinus et al., 2013). Overall, there are indications that γ-oryzanol possesses cholesterol lowering properties similar to free phytosterols or phytosterol esters, although they seem to be absorbed poorly or not at all; however full prove in human studies has not been provided yet. The potential health benefits of alkyl hydroxycinnamates, apart from their antioxidant activity, have been studied in vitro in a few reports. It has been shown that alkyl caffeates and alkyl ferulates inhibited tumor cell proliferation and COX enzyme, with differences between the different alkyl esters (Jayaprakasam et al., 2006). In another study it was shown, that the anticancer activity was higher for linear side chains compared to branched side chains of ferulic and caffeic acid (Li et al., 2012). Hexyl ferulate and caffeate and feruloyl- and caffeolyhexylamide showed cytotoxicity towards human breast cancer cell lines, whereas the 10 Hydroxycinnamic acids parent free acid did not show activity (Serafim et al., 2011). The anti-inflammatory activity was analyzed of alkyl caffeates and revealed that length and size of the alkyl part influenced nitric oxide production in macrophages (Uwai et al., 2008). Finally, the antiamyloidal activities of caffeic, chlorogenic, ferulic and sinapic acid esters were analyzed in vitro, which also showed an effect of the lipophilicity of the hydroxycinnamic acid derivatives (Kondo et al., 2014). However, although these in vitro studies show some evidence, many further studies on the health benefits and potential toxicity of alkyl hydroxycinnamates need to be performed to make a clear and full picture. . 11 General enzymatic reactions 2 General enzymatic reactions 2.1 Kinetics of enzymatic reactions The basics in enzyme kinetics were extracted from two text books (Belitz et al., 2009; Bugg, 2012). To compare and to evaluate enzymes often the kinetic constants Km and kcat are determined. They derive from the Michaelis-Menten model, which is based on the following scheme: Scheme 1: Michaelis-Menten model. This model indicates that only one substrate is binding to the enzyme, which leads to the reversible formation of an enzyme-substrate complex (ES). Further, there is only one kinetically significant step, which leads to the product formation and is irreversible. Also not many enzymes fit these criterions exactly; it is a suitable model for a broad range of enzymes. A steady state approximation, which means that the concentration of the intermediate species ES remains constant, leads to the Michaelis-Menten equation (1). It describes the dependency of the initial reaction rate (v0) of an enzyme to the substrate concentration [S] as illustrated in Figure 3. 푣푚푎푥∙[푆] 푣0 = (1) 퐾푚+[푆] Figure 3: Initial reaction rate as a function of the substrate concentration based on the Michaelis-Menten equation. The Michaelis constant Km is defined as the substrate concentration at which half of the maximum velocity can be observed. Finally, vmax divided by the total enzyme concentration leads to kcat, the turnover number, describing the number of substrates converted per enzyme per time. If the reaction proceeds via an enzyme-acyl complex the Michaelis-Menten model can be adapted accordingly (Zerner & Bender, 1964). The catalytic step is divided into two steps, the 12 General enzymatic reactions formation (k2) of the acyl-enzyme intermediate (EI) and the deacylation (k3). In case of an ester, P1 represents the alcohol and P2 the acid. This leads to the following scheme: Scheme 2: Adapted Michaelis-Menten model including an enzyme-acyl complex, adapted from (Zerner & Bender, 1964). In this case also the definition of the kinetic constants changes, where Ks represents the substrate binding constant: 푘2푘3 푘3 푘푐푎푡 = (2) 퐾푚 = 퐾푆 (3) 푘2+푘3 푘2+푘3 From there two cases can be distinguished, depending on the values of k2 and k3 or the rate- determining step. If k2 >> k3, the deacylation is limiting it follows: 푘3 푘푐푎푡 = 푘3 (4) 퐾푚 = 퐾푆 (5) 푘2 In the case of k3 >> k2 where the formation of the acyl-enzyme intermediate is limiting it leads to: 푘푐푎푡 = 푘2 (6) 퐾푚 = 퐾푆 (7) In case of chymotrypsin for certain amide substrates kcat is dominated by the formation of the intermediate and for certain ester substrates by the deacylation (Zerner & Bender, 1964). If esters of the same acid show similar rate constants, this can be explained by the deacylation of a common intermediate, which is rate-determining (Zerner et al., 1964). Comparisons of kinetic constants of enzymatic reactions including acyl-enzyme intermediates can therefore give information about the mechanism of the enzymatic catalysis. 2.2 Lipase catalysis in organic solvent Early reviews on enzyme catalysis in monophasic organic solvents were published in the late 1980ies, discussing the “new” technique of enzymatic catalysis in almost anhydrous solvents (Dordick, 1989). The main possible advantages of this technique were listed and discussed including substrate solubility, shifting of thermodynamic equilibria, easier product recovery and increased enzyme stability at higher temperatures. The main issues of optimizing the efficiency of the system included the role of water, the biocatalyst preparation, and the effect 13 General enzymatic reactions and choice of the solvent (Dordick, 1989). These parameters are still key factors for the optimization of such systems today. For the application of lipases, these factors are discussed below in more detail. Enzymes as catalysts in organic solvents can be used in two forms, either in free form or immobilized. The immobilization of lipases has been reviewed recently (Adlercreutz, 2013). Normally, lipases are insoluble in organic solvents and are thus in the solid state, as it is the case for a lyophilized lipase powder. Non-immobilized lipases usually show rather low activity and tend to aggregate, which may lower mass transfer. Through immobilization of lipases the activity can be increased also probably due to conformational changes during immobilization. Common techniques to immobilize lipases include adsorption, entrapment, covalent coupling, and cross-linking of the enzyme. The immobilized enzyme should be evaluated based on its catalytic activity, the yield at the end of the reaction and its stability during the process. However, there is not the one optimal immobilization technique for lipases, each is unique in its properties and therefore also immobilization has to be adapted (Adlercreutz, 2013). The choice of solvent can influence the system drastically. As a rule of thumb solvents with a log P value larger than three are preferred, as the enzyme is deactivated less and stays active longer (Villeneuve, 2007). However, the solubility of the substrate has to be kept in mind and should be selected in a way that the substrates are at least partially soluble. Apart from the very nonpolar solvents such as n-hexane or isooctane also more polar solvents are used as co-solvents or pure. Often applied candidates are tertiary alcohols, as they do not participate in the reaction (Villeneuve, 2007). Further, it can also help to select a solvent, which solubilizes the product best. This can make the reverse reaction unfavorable and therefore increase the yield (Zeuner et al., 2012). Overall, the selection of solvent is also very much dependent on the system of interest. The water activity (aw) is another key factor in enzymatic catalysis in organic solvents. The positive effects (i.e. activation due to increased flexibility of the enzyme) and the negative effects (i.e. favoring hydrolysis, building up a diffusion barrier or inhibition) have to be in balance leading to an optimum for esterification reactions (Adlercreutz, 2013). However, to control the water activity during a course of reaction is not a fully solved problem yet, although several solutions have been proposed (Villeneuve, 2007). For initial water activity equilibration with saturated salt solutions can be applied. For the control during the reaction systems such as the use of membranes between the reaction media and the salt solution or a controlled air stream have been evaluated. However, the efficiency can be a problem as 14 General enzymatic reactions mass transfer between the phases can be limited (Villeneuve, 2007). Finally, these systems require special equipment, which is especially a problem during screenings. Moreover, the application of drying agents (i.e. molecular sieve) is used to remove water, which is formed during the reaction. However, this is far from fine-tuning the water activity (Villeneuve, 2007). 2.3 Properties of lipases used for lipophilization reactions Lipases [E.C.3.1.1.3] are also known as triacylglycerol ester hydrolases and as their name predicts, they naturally hydrolyze ester bonds of triacylglycerols (Adlercreutz, 2013; Villeneuve, 2007). Lipases usually act on organic–aqueous interface at which they are even activated. Lipases have a broad substrate specificity and high activity and stability in organic media can be achieved easier than with many other enzymes. This leads to many applications of lipases in organic media. The most commonly used lipases are from Burkholderia cepacia (Lipase PS), Candida antarctica (Novozym 435 in immobilized form), Candida rugosa, Rhizomucor miehei (formerly Mucor miehei, Lipozyme RM IM in immobilized form), Rhizopus oryzae, and Thermomyces lanuginosus (Lipozyme TL IM in immobilized form) (Adlercreutz, 2013). As the lipases applied in this thesis are the ones from C. rugosa and R. miehei, they will be discussed in more detail below. The properties and applications of the lipase from R. miehei in fats and oils modifications and in chemical processes have been reviewed recently (Rodrigues & Fernandez-Lafuente, 2010a, 2010b). The lipase from R. miehei is an extracellular enzyme and naturally appears in two forms, which differentiate by partial deglycosylation. R. miehei lipase is commercially available for example from Novozymes in free and immobilized form. A weak anion- exchange resin serves as carrier for the immobilized lipase. The enzyme is composed of one polypeptide chain of 269 amino acids, which makes a molecular weight of 31’600 Da. In the active center a catalytic triad is located (Ser144, Asp203, His257). Lipase from R. miehei shows high esterification activity, even in anhydrous systems. Further, the lipase from R. miehei is sn-1,3-specific. It found many applications in the modification of fats and oils but also as catalyst for various ester formations, the resolution of racemic mixtures and also in the use of its regioselectivity. Overall, the lipase from R. miehei lipase seems to be a suitable catalyst for esterification reactions (Rodrigues & Fernandez-Lafuente, 2010a, 2010b). The characteristics of C. rugosa lipase have also been summarized (Dominguez de Maria et al., 2006). One of the key characteristics of C. rugosa lipase is the presence of several isoenzymes. At least seven genes are involved of the lipase production and enzymes expressed from five genes have been biochemically characterized. Also commercially available C. rugosa lipase preparations contain isoenzymes, although Lip1 in highest amount 15 General enzymatic reactions amongst the analyzed preparations. The fermentation parameters of C. rugosa during lipase production can strongly influence the lipase quantity and quality including isoenzyme profile. Additionally, C. rugosa applies a non-universal codon for serine. This makes production of recombinant lipases challenging. Site-directed mutagenesis or even complete synthesis of the required gene have been applied to overcome this challenge. However, amongst the isoenzymes the homology of the 534 residues long peptide chain is high (ca. >70%), differences in the biocatalytic behavior could be observed. Although, the characterized C. rugosa lipases show a catalytic triad (Ser209-Glu341-His449). This lipase has a tunnel for the substrate, which is rather L-shaped suitable for oleic acid. Thus, it has a broad specificity for fatty acids but low activity for long, polyunsaturated fatty acids. Finally, it is proposed that in organic medium, Lip1 rather prefers linear alcohols and Lip2 and Lip3 catalyze the esterification of sterically hindered alcohols. Although there are several drawbacks using C. rugosa lipase, it bears a great potential for biotechnological applications (Dominguez de Maria et al., 2006). 16 Enzymatic lipophilization of hydroxycinnamic acids 3 Enzymatic lipophilization of hydroxycinnamic acids Through esterification new hydroxycinnamic acid esters can be created. In case the second substrate to the phenolic acid is lipophilic, this process is also called lipophilization (Figueroa- Espinoza & Villeneuve, 2005). The interest of this modification is mainly an adjusted polarity. This leads to an increased solubility in lipophilic systems. Secondly, the hydrophobicity is a key property if phenolic acids are applied as antioxidants in a multi-phase system including emulsions (Laguerre et al., 2013). Thirdly, there are indications that the bioactivity differs between hydroxycinnamic acid esters (Jayaprakasam et al., 2006). Efficient and direct esterification systems are therefore required. Two main approaches of esterification can be differentiated, chemically and enzymatically catalyzed reactions. Enzymatic catalyzed reactions are typically more specific and therefore less side products are formed. This reduces the cost for waste treatment and simplifies purification. Further, the reactions usually occur under milder reactions (Villeneuve et al., 2000). However, enzymes are often more expensive than traditional chemical catalysts (Figueroa-Espinoza & Villeneuve, 2005). Conclusively, enzyme catalyzed reactions are more environmental friendly and should be optimized to reduce enzyme costs. 3.1 Using lipases One class of enzymes applied in the enzymatic lipophilization of hydroxycinnamic acids are lipases. In Table 2 a selection of studies including the enzymatic lipophilization of hydroxycinnamic acids by lipases are listed. In the chapter below the applications of other enzymes are discussed. The focus of this chapter lays on lipophilization. The enzymatic esterification of hydroxycinnamic acid with saccharides has been reviewed recently (Zeuner et al., 2012) and will not be discussed here. Neither will be discussed the enzymatic synthesis of glycerol hydroxycinnamates that leads to the formation of more hydrophilic compounds. As one possibility of the esterification to glycerol, the incorporation of hydroxycinnamic acids into triglycerides is presented. The collected studies were grouped into four categories, namely the esterification of short and medium chain alcohols, the esterification of long chain alcohols, caffeic acid esterification to phenyl alcohols, and esterification to acylglycerols. 17 Enzymatic lipophilization of hydroxycinnamic acids Table 2: Overview of selected studies investigating enzymatic lipophilization of hydroxycinnamic acid derivatives catalyzed by lipases. Substrates Enzymes Solvents / conditions References Esterification of short and medium chain alcohols Hydroxycinnamic acid derivatives, butanol, octanol, dodecanol, oleyl Novozym 435 Solvent-free (Guyot et al., 1997) alcohol Ferulic acid, ethanol, octanol/ t-Butanol, toluene, (Compton et al., Novozym 435 ethyl ferulate, octanol, monoolein triolein solvent-free (triolein) 2000) Novozym 435, Hydroxycinnamic acid and benzoic acid (Stamatis et al., Lipozyme RM IM Solvent-free derivatives, octanol 1999, 2001) C. rugosa lipase Ferulic acid, ethanol and Novozym 435 Isooctane (Lee et al., 2006) p-methoxycinnamic acid, 2-ethyl hexanol Immobilized lipase Ferulic acid, pentanol, hexanol and Solvent-free, (Yoshida et al., from C. antarctica heptanol continuous system 2006) Chirazyme L-2 C2 Hydroxycinnamic acid derivatives, Novozym 435, Ionic liquid or (Katsoura et al., methanol, ethanol, propanol, butanol, Lipozyme RM IM hexane and acetone 2009) hexanol, octanol, geraniol Lipases from R. miehei and Hydroxycinnamic acid and benzoic acid (Zoumpanioti et al., C. antarctica in Solvent-free derivatives, octanol 2010) modified cellulose organogels Steapsin (Kumar & Kanwar, Ferulic acid, ethanol DMSO immobilized on celite 2011) Dihydrocaffeic acid, ferulic acid, caffeic Hexane/butanone (Yang et al., 2012b; acid, butanol, hexanol, octanol, decanol, Novozym 435 mixtures or ionic Yang et al., 2012c) dodecanol, octadecanol liquids Deep eutectic Methyl p-coumarate, methyl ferulate, (Durand et al., Novozym 435 solvent–water binary octanol 2013) mixtures R. oryzae lipase on Ferulic acid, ethanol Isooctane or hexane (Kumar et al., 2013) Fe3O4-chitosan Methyl caffeate, propanol Novozym 435 Ionic liquid (Pang et al., 2013) B. licheniformis p-Coumaric aid, methanol, ethanol, (Sharma et al., SCD11501 lipase on Solvent-free propanol, butanol 2014) celite Caffeic acid, 2-pentanol, 2-heptanol, Novozym 435 Isooctane (Xiao et al., 2014) 2-octanol (Sandoval et al., Ferulic acid, ethanol, dodecanol Novozym 435 Diisopropyl ether 2015) Ionic liquid, (Wang et al., Caffeic acid, methanol Novozym 435 ultrasound 2015a) irradiation Esterification of long chain alcohols Dihydrocaffeic acid, ferulic acid, linolenyl Hexane/2-butanone (Sabally et al., Novozym 435 alcohol 75:25 or 65:35 (v/v) 2005) Hydroxycinnamic acid derivatives, (Vosmann et al., Lipozyme RM IM, methyl or ethyl esters thereof, oleyl Solvent-free, 80 kPa 2006) (Weitkamp et Novozym 435 alcohol al., 2006) 18 Enzymatic lipophilization of hydroxycinnamic acids Table 2 continued: Solvents / Substrates Enzymes References conditions Ionic liquid/ Ferulic acid, oleyl alcohol Novozym 435 isooctane binary (Chen et al., 2011a) system Caffeic acid esterification with phenyl alcohols Cinnamic acid and hydroxy and methoxy (Stevenson et al., derivatives, phenylethanol, 4-methoxy Novozym 435 t-Butanol 2007) phenylethanol, tyrosol (Widjaja et al., Caffeic acid, 2-phenylethanol Novozym 435 Isooctane 2008) Methyl caffeate, 2-cyclohexylethanol, 3-cyclohexyl-1-propanol, 4-phenylbutanol, Novozym 435 Ionic liquid, 845 hPa (Kurata et al., 2010) 5-phenylpentanol (Chen et al., 2010a; Caffeic acid, 2-phenylethanol, octanol Novozym 435 Isooctane Chen et al., 2010b) Continuous ultrasound-assisted Caffeic acid, 2-phenylethanol Novozym 435 packed-bed reactor, (Chen et al., 2011b) in isooctane/ t-butanol 9:1 Caffeic acid, phenethyl alcohol Novozym 435 Ionic liquid (Ha et al., 2013) Ionic liquid, (Wang et al., 2014; Methyl caffeate, propanol, 2-phenylethanol Novozym 435 continuous flow Wang et al., 2013) microreactor 2% DMSO in ionic Caffeic acid, 2-phenylethanol Novozym 435 (Gu et al., 2014) liquid Esterification with acylglycerols (Laszlo & Compton, Ethyl ferulate, soybean oil Novozym 435 Solvent free 2006; Laszlo et al., 2003) p-Hydroxyphenyl acetic acid, p-coumaric Hexane/2-butanone acid, sinapic acid, ferulic acid and Novozym 435 (Safari et al., 2006) 85:15 (v/v) 3,4-dihydroxybenzoic acid, triolein Ethyl ferulate, tributyrine Novozym 435 Toluene (Zheng et al., 2008) Hexane/2-butanone (Karboune et al., Hydroxycinnamic acid derivatives, ethyl Novozym 435 85:15 or solvent- 2008; Sorour et al., ferulate, flaxseed oil free, surfactants 2012) Ferulic acid, cinnamic acid, flaxseed oil Novozym 435 Hexane (Choo et al., 2009) Ethyl ferulate, triolein Novozym 435 Solvent-free (Theng et al., 2009) (Ciftci & Saldana, Ferulic acid, flaxseed oil Novozym 435 Supercritical CO 2 2012) Ethyl ferulate, glycerol, fish oil Novozym 435 Solvent-free (Yang et al., 2012a) Solvent-free, (Sun et al., 2012; Ethyl ferulate, distearin, monostearin Novozym 435 10 mm Hg Sun & Zhou, 2014) Toluene/chloroform Ethyl ferulate, phosphatidylcholine Novozym 435 (Yang et al., 2013) 9:1 (v/v) p-Coumaric acid, triolein, seal blubber oil, Hexane/2-butanone (Wang & Shahidi, Novozym 435 menhaden oil 3:1 (v/v) 2014a, 2014b) If not indicated otherwise, primary alcohols were used as substrates. Novozym 435 corresponds to immobilized lipase B from C. antarctica, Lipozyme RM IM corresponds to immobilized lipase from R. miehei. 19 Enzymatic lipophilization of hydroxycinnamic acids The enzymatic esterification has been achieved in various solvents. Apart from organic solvents, also non-conventional media such as ionic liquids, deep eutectic solvents and supercritical CO2 were applied. Finally, solvent-free systems were chosen, where the solvent represents the second substrate. Concerning organic solvents a wide variety found application in lipophilization of hydroxycinnamic acids. From quite polar solvents such as DMSO, acetone or t-butanol also nonpolar solvents such as hexane, toluene or isooctane were applied. Many studies evaluated different solvents (Chen et al., 2011a; Katsoura et al., 2009; Lee et al., 2006; Stamatis et al., 1999; Yang et al., 2012b). However, as can also be seen in Table 2, the conclusions were quite different but often in favour on nonpolar solvents, even if the solubility of free hydroxycinnamic acids is low. Low substrate solubility increases the net binding energy to the enzyme. Further, the more hydrophobic product is stabilized in the nonpolar solvent and the reverse reaction is less favoured (Zeuner et al., 2012). Several times ionic liquids were found to be the better solvent, leading to improved yields (Chen et al., 2011a; Katsoura et al., 2009; Yang et al., 2012c). The disadvantage of ionic liquids is that the products have to be extracted after the reaction, as the reaction solvent cannot easily be evaporated, as well a their high costs (Zeuner et al., 2012). Overall, many different solvent systems have already been evaluated in the enzymatic lipophilization of hydroxycinnamic acids. Novozym 435 has been applied most often as catalyst (Table 2). Secondly, Lipozyme RM IM was used to esterify hydroxycinnamic acids. Finally, also other immobilized lipases have been applied such as steapsin or lipases from R. oryzae or B. licheniformis. Several studies evaluated different lipase preparations. Often Novozym 435 was found most active (Sun et al., 2012; Vosmann et al., 2006; Weitkamp et al., 2006; Yang et al., 2013) or the only enzyme able to catalyze the reaction (Compton et al., 2000). However, it has also been shown for the esterification of ferulic acid that Lipozyme RM IM leads to higher yields in solvent-free system (Stamatis et al., 1999) or in hexane (Katsoura et al., 2009). Further interesting results were reported on the enzyme activity in the solvent-free esterification of 4-methoxycinnamic acid with oleyl alcohol in vacuo. For the direct esterification the activity measured after 2h for Novozym 435 was double compared to Lipozyme RM IM. In contrast to the transesterification of methyl 4-methoxycinnamate under similar conditions, where the activity of Novozym 435 was almost six times higher (Vosmann et al., 2006; Weitkamp et al., 2006). The comparison of lipase activity may therefore be also strongly dependent if the substrate is directly esterified or transesterified. Overall, to name the best enzyme catalyst for hydroxycinnamic acid lipophilization is not possible as such; it depends on the detailed substrates and system. 20 Enzymatic lipophilization of hydroxycinnamic acids Lipophilization was achieved in two main ways, by direct esterification of the free acid or by transesterification of a short chain alcohol hydroxycinnamate (often methyl or ethyl esters). However, there are not so many studies directly comparing the efficiency of these two approaches. In the study of Compton and colleagues the yield of octyl ferulate catalyzed by Novozym 435 was increased from 14% to 50% when going from direct esterification to transesterification of ethyl ferulate. The yield was even increased further by applying vacuum every 24 h to remove the formed ethanol (Compton et al., 2000). Later on the transesterifcation activity of Novozym 435 towards methyl ferulate and 1-hexadecanol was 56 times more compared to the esterification activity under the same conditions (Weitkamp et al., 2006). Finally, in ionic liquids the yield of propyl caffeate was increased from 41% to 52% and 99% when ethyl caffeate or methyl caffeate were used, respectively (Pang et al., 2013). In contrast also a lower yield was measured for ethyl ferulate compared to free ferulic acid when transesterified to flaxseed oil in organic solvent (Karboune et al., 2008). The use of activated esters as substrates for enzymatic transesterification such as vinyl ferulate has been evaluated (Yu et al., 2010) and is discussed in chapter 4.2.2. Overall, mostly increased activities and/or yields can be observed if hydroxycinnamic acids are transesterified. The esterification yield may strongly depend on the structure of the hydroxycinnamic acid derivative. This phenomenon was first described by Guyot and co-workers. They observed lipase inhibition in case of a simultaneous presence of a double bond in the side chain and a para-hydroxylation, which they attributed to electronic effects (Guyot et al., 1997). However, they conducted the esterifications solvent-free with 1-butanol. Dihydrocaffeic acid was esterified to 78%, ferulic acid to traces and for caffeic acid no reaction was measured (Guyot et al., 1997). Later on Yang and colleagues observed a very similar behavior in hexane- butanone mixtures. Dihydrocaffeic acid was almost fully converted in 3 days but the yield for caffeic acid under similar conditions was around 12% in 6 days (Yang et al., 2012b). However, in ionic liquid the ratio was not as drastic. The yield for caffeic acid was 8% and for diyhdrocaffeic acid 36% (Katsoura et al., 2009). This difference was even less pronounced in the solvent-free esterification to oleyl alcohol. The yield of the caffeic acid was similar, but reaction time was double (6 days) (Vosmann et al., 2006). Apparently, the strength of the electronic effect of the conjugated acid group with the phenolic hydroxyl group is dependent on the polarity of the reaction mixture. Further, the ratio of yields between different cinnamic acid derivatives depends on the lipase. Stamatis and colleagues measured the esterification activity of Novozym 435 and Lipozyme RM IM for several hydroxycinnamic acid derivatives. Cinnamic acid was esterified by both lipases most efficiently, as well as the m-coumaric acid amongst the coumaric acids. 21 Enzymatic lipophilization of hydroxycinnamic acids However, the ferulic acid was esterified better compared to p-coumaric acid by Lipozyme RM IM, while it was the other way around for Novozym 435 (Stamatis et al., 1999). Also comparing these two lipases in ionic liquid Katsoura and colleagues found similar results. The yield for the cinnamic acid was similar for both lipases, but Novozym 435 was almost not esterifying sinapic acid, while the yield of octyl sinapate for Lipozyme RM IM was more than half of the yield of cinnamic acid (Katsoura et al., 2009). This confirmed the good activity of Lipozyme RM IM for methoxylated hydroxycinnamic acids. In the study of Stevenson and co- workers also a mixture of hydroxycinnamic acid derivatives was esterified by Novozym 435 to various alcohols. Even in a mixture similar behavior as described above could be observed although ferulic acid was slightly better esterified than p-coumaric acid (Stevenson et al., 2007). Finally, it was detected that using secondary alcohols as substrates for the esterification of caffeic acid by Novozym 435 optically pure caffeic acid esters were produced (Xiao et al., 2014). Overall, the enzymatic esterification of hydroxycinnamic acid is not only dependent on the structure but also on the enzyme-substrate combination. The enzymatic esterification of hydroxycinnamic acids is overall regarded as challenging and yields are often low or very high amounts of enzyme are added. The mentioned difficulties including electronic effects and steric hindrance reduce the activity of the lipases. Especially the combination of an unsaturated side chain with a para-hydroxylation leads to a deactivation of the carboxylic acid. However, there are differences between lipases, which also suggest that steric hindrance could contribute the reduced activity. It is therefore of interest to also evaluate other enzymes than lipases on their esterification activity against hydroxycinnamic acids, as it is discussed in the next chapter. 3.2 Using other enzymes Apart from lipases other enzymes have been applied to esterify hydroxycinnamic acids. In Table 3 the studies conducting enzymatic lipophilization by other enzymes than lipases are listed. Studies including only the esterification with for example sugars or glycerol were again not included. Mostly feruloyl esterases were evaluated on their ability to esterify or transesterify hydroxycinnamic acid. But also commercial mixtures containing feruloyl esterase activity, a cutinase and a rice koji enzyme were used (Table 3). Feruloyl esterases (for more details see chapter 5) can release hydroxycinnamic acids from plant fibers (Faulds, 2010). Depending on their substrate specificity for the most common hydroxycinnamic acids (ferulic acid, sinapic acid, p-coumaric acid, and caffeic acid) and further properties they can be separated into groups (Crepin et al., 2004). For the esterification of phenolic acids they were mainly applied in microemulsion system and/or 22 Enzymatic lipophilization of hydroxycinnamic acids immobilized. So called surfactantless microemulsions or ternary systems are usually composed of hexane and water and a short chain alcohol. This short chain alcohol can be a tertiary one, and another substrate like a sugar can be added (Topakas et al., 2005) or e.g. primary or secondary butanol can be used to from the microemulsion and as substrate in one (Vafiadi et al., 2008a). In an emulsion containing surfactant, namely cetyltrimethylammoniumbromide (CTAB), the synthesis of pentyl ferulate was achieved catalyzed by A. niger feruloyl esterase (Giuliani et al., 2001). The drawback of these emulsified systems is often a limited choice of alcohols as substrates. Table 3: Overview of studies conducting lipophilization through esterification or transesterification of hydroxycinnamic acids with other enzymes than lipases. Substrates and solvents Enzymes Reference Ferulic acid, pentanol in cetyltrimethyl- Feruloyl esterase (Giuliani et al., 2001) ammoniumbromide microemulsion from A. niger Cinnamic, p-coumaric, ferulic, F. oxysporum p-hydroxyphenyl propionic acid, 1-octanol, esterase, F. solani (Stamatis et al., 2001) solvent free cutinase p-Hydroxyphenylacetic acid, p-hydroxyphenylpropionic, cinnamic acid, F. oxysporum (Topakas et al., 2003) p-coumaric acid, ferulic acid, 1-propanol in feruloyl esterase n-hexane/1-propanol/water Methyl ferulate, methyl p-coumarate, methyl caffeate, methyl sinapate, S. thermophile (Topakas et al., 2005) 1-butanol, L-arabinose in feruloyl esterase n-hexane/butanol/water Methyl ferulate, methyl p-coumarate, CLEAs of A. niger methyl caffeate, methyl sinapate, type A feruloyl (Vafiadi et al., 2008a) 1-butanol, 2-butanol in esterase n-hexane/butanol/water CLEAs of Ultraflo L, Methyl ferulate, 1-butanol in Depol 670L, Depol (Vafiadi et al., 2008b) n-hexane/butanol/water 740L Depol 740L on Methyl ferulate, 1-butanol and 7.5% buffer (Thorn et al., 2011) mesoporous silica Ferulic acid, sinapic acid, caffeic acid, (Hashizume et al., p-coumaric acid, ethanol, methanol, Rice koji enzyme 2013a) 1-propanol in buffer CLEA: cross-linked enzyme aggregate 23 Enzymatic lipophilization of hydroxycinnamic acids Immobilization techniques were also of relevance amongst the esterification studies with feruloyl esterases. One approach was the immobilization of a feruloyl esterase on mesoporous silica, which could then be used in an almost solvent-free system for glyceryl ferulate synthesis with only small addition of buffer (Thorn et al., 2011). Also feruloyl esterase CLEAs found application in surfactantless microemulsions for the synthesis of butyl ferulate showing higher and more stable synthetic activity (Vafiadi et al., 2008b). Both, transesterifications of methyl hydroxycinnamates and direct esterifications, were applied. The yields for the direct esterifications were rather low after long incubation times. Except in the study of Giuliani, where higher yields (50-60%) in 8.3 h were reached (Giuliani et al., 2001). Generally, immobilization of feruloyl esterases improves esterification activity and enzyme stability and transesterification may improve the yield. The synthesis of long chain alkyl ferulates by non-lipase enzymes has not been researched on yet. The longest alkyl is 1-octanol which was esterified by an esterase and a cutinase in a solvent free system (Stamatis et al., 2001). However, the yields were only 10% or below for p-coumaric acid and ferulic acid. The application of longer alcohols in the hydroxycinnamic acid ester synthesis by feruloyl esterase has not been reported yet and could be further explored. Further, feruloyl esterases were applied to synthesize glyceryl ferulate (Tsuchiyama et al., 2006; Zeng et al., 2014). The enzymatic synthesis of glyceryl ferulate is a promising approach, as these compounds are occurring naturally (Graça & Pereira, 2000). In this way the water solubility of the ferulate is improved. However, as this is not the core of this work it will not be discussed further. Neither discussed is the synthesis of sugar esters by feruloyl esterase, which has also been studied several times (Couto et al., 2010; Couto et al., 2011; Vafiadi et al., 2005). Further, the formation of ethyl ferulate from ferulic acid and ethanol by a rice koji enzyme has been showed in a buffer system (Hashizume et al., 2013a). Enzymes naturally catalyzing the esterification of ferulic acid have also been purified and evaluated including hydroxycinnamoyl-CoA transferases discussed in chapter 1.2.1. Also another enzyme has been purified from rice and in vitro catalyzed the formation of feruloyl arabinoxylan- trisaccharide from feruloyl CoA (Yoshida-Shimokawa et al., 2001). Later on a hydroxycinnamoyltransferase from rice has been expressed in E. coli, which catalyzed the acid transfer from p-coumaroyl-CoA, caffeoyl-CoA, and feruloyl-CoA to glycerol or shikimic acid (Kim et al., 2012). However, the relevance of such enzymes for possible large-scale applications is difficult to judge, mainly due to the requirement of the CoA hydroxycinnamate. 24 Esterification of phytosterols 4 Esterification of phytosterols 4.1 Enzymatic phytosterol fatty acid esters synthesis The enzymatic esterification of phytosterols is challenging due to the structure of the sterol. Sterols are secondary alcohols and bulky substrates. The enzyme supposed to catalyze the reaction has to be able to accommodate such substrates. However, many different approaches have been suggested for the enzymatic esterification of phytosterols with fatty acids (Table 4). If a screening of various lipases was conducted only the one with the highest yield or the one chosen for most experiments is listed. The most commonly applied lipase is from C. rugosa in free form but also immobilized on various carriers. Generally, enzymes from Candida genus were able to catalyze the esterification of plant sterols. Further, lipases from the genera Pseudomonas, Rhizomucor and Thermomyces were applied. In a few studies lipases from the genera Alcaligene and Burkholderia and also from papaya were used (Table 4). Conclusively, various lipases seem to have a good potential as catalyst for the enzymatic synthesis of steryl esters. Cholesterol esterases on the other hand have been used rarely for the esterification of phytosterols. One explanation for this could be the low tolerance of cholesterol esterases towards organic solvent that helps to solubilize substrates such as sterols. The first organic solvent tolerant cholesterol esterase has only been reported several years ago (Takeda et al., 2006). Another issue with cholesterol esterases can be the sterol specificity thus allowing less flexibility for sterol substrates. Morinaga and colleagues reported that a cholesterol esterase from Trichoderma sp. AS59 showed 50% esterification activity towards stigmasterol compared to cholesterol (Morinaga et al., 2011). Similar observations were recorded earlier for a porcine pancreas homogenate, where the esterification yield with oleic acid compared to the cholesterol after 2 h was 41% for β-sitosterol and only 15% and 12% for stigmasterol and ergosterol, respectively (Swell et al., 1954). The application of sterol esterases therefore bears several challenges. 25 Esterification of phytosterols Table 4: Overview of studies conducting enzymatic esterification of phytosterols with fatty acids or fatty acid esters. Substrates, conditions Enzymes References Immobilized lipase in solvent system Phytostanols, fatty acids C12:0, Immobilized C. antarctica (He et al., 2010) C14:0, C16:0, C18:0 in hexane lipase B, Novozym 435 Immobilized T. lanuginosus (Sengupta & Ghosh, β-Sitosterol, fish oil in hexane lipase, Lipozyme TL IM 2011) Candida sp. 99–125 Phytosterols, oleic acid in isooctane (Pan et al., 2012) immobilized on textile Phytosterols, lauric acid in hexane C. rugosa lipase on (Jiang et al., 2013) with trehalose addition macroporous resin β-Sitosterol, fatty acids C2:0-C18:0 Immobilized C. antarctica (Panpipat et al., 2013) in hexane lipase A β-Sitosterol, conjugated linoleic acid Chirazyme L-2 c.-f. C2 (Li et al., 2010) in hexane (from C. antartica) Phytosterols, triglycerides and free C. rugosa lipase (Zheng et al., 2012a; fatty acids from sunflower, rapeseed, immobilized on various Zheng et al., 2014; corn, tea seed, linseed and rice carriers (functionalized Zheng et al., 2012b; bran; free fatty acids (C16:0, C18:1, silica particles or polymer Zheng et al., 2013; C18:2, C18:3, conjugate linoleic particles) Zheng et al., 2012c; acid) in isooctane or hexane Zheng et al., 2015) Non-immobilized lipase in solvent system Canola phytosterols, oleic acid, C. rugosa lipase (Villeneuve et al., 2005) methyl oleate in hexane Physotsterols, oleic acid in hexane C. rugosa lipase (Kim & Akoh, 2007) Phytosterols, caprylic acids in C. rugosa lipase (Tan et al., 2012) hexane Phytosterols, docosahexaenoic acid Lipoprotein lipase 311 (Tan & Shahidi, 2012b) in hexane Non-immobilized lipase in solvent-free system Lipase QLM (Alcaligenes Phytosterols, sunflower oil (Negishi et al., 2003) sp.) Soybean oil deodorizer distillate, (Teixeira et al., 2011, olive oil deodorizer distillates, refined C. rugosa lipase 2012, 2014; Torres et olive oil, oleic acid al., 2007) 26 Esterification of phytosterols Table 4 continued : Substrates, conditions Enzymes References Solvent-free system under reduced pressure Cholesterol, sitostanol, Immobilized lipases from: T. stigmasterol, 5α-cholestan-3β-ol, lanuginosus, R. miehei, C. (Weber et al., 2001a, methyl oleate, oleic acid, triolein, antarctica; non-immobilized 2001b, 2002, 2003) methyl fatty acid esters, rapeseed lipases from: C. rugosa lipase, oil, soybean oil, 20-40 mbar Carica papaya lipase Wood sterols, sunflower fatty acid (Martinez et al., Lipase from P. stutzeri PL-836 methyl esters, 2 mbar 2004) Phytosterols, fatty acids from butter C. rugosa lipase on octylsilica (Torrelo et al., 2009) oil, 100 mbar Phytosterols, tributyrine, ethyl C. rugosa lipase, P. stutzeri butyrate, fatty acid ethyl esters (Torrelo et al., 2012) lipase from butter fat, 100-350 mbar Phytosterols, fatty acid from pine C. rugosa lipase on Lewatit VP (No et al., 2013) nut, 80 kPa OC 1600 Non-conventional reaction medias Cholesterol, cholestanol, and (Shimada et al., sitosterol, fatty acids C22:6, C20:5, Pseudomonas sp. lipase 1999) C18:3, C18:2, and 30% water Sitostanol, C8:0, C10:0, C12:0, Lipase from Burkholderia (King et al., 2001) C16:0, C18:0 in supercritical CO2 cepacia, Chirazyme L-1 β-Sitosterol, C6:0, C8:0, C10:0, C12:0, conjugated linoleic acid, and C. rugosa lipase (Vu et al., 2004) 0.3 mL/gsterol water or hexane Phytosterols, fatty acids C12:0, C14:0, C16:0, C18:0, C18:1 in C. rugosa lipase (Zeng et al., 2015) water-in-ionic liquid microemulsion Phytosterols, oleic acid in C. rugosa lipase immobilized on isooctane, under microwave ZnO nanowires/macroporous (Shang et al., 2015) irradiation SiO2 Phytosterols, soybean oil in Immobilized C. antarctica (Hu et al., 2015) supercritical CO2 lipase, Novozym 435 Sterol esterases Dihydrocholesterol, cholesterol, β-sitosterol, sitosterol, stigmasterol, Homogenate from hog (Swell et al., 1954) ergosterol, butyric acid, oleic acid, pancreas in buffer containing bile salts Phytosterols, caprylic acid Sterol esterase from A. oryzae (Hellner et al., 2010) sunflower oil, solvent-free Cholesterol, stigmasterol, stearic Cholesterol esterase from (Morinaga et al., acid, in buffer or biphasic hexane- Trichoderma sp. AS59 2011) water system 27 Esterification of phytosterols The substrates esterified are very diverse, both sterol and fatty acid. Concerning the sterol substrate most studies use a mixture of phytosterols, the major compound being β-sitosterol (Table 4). This is probably also due to the lack of commercially available single plant sterols. Further, sterols from plant oil deodorizer distillates have been used as source for sterols, in solvent-free systems. They are cheap sources of phytosterols but also bring some challenges for the enzymatic esterification, due to variable compositions (Teixeira et al., 2014). Similar trends can be observed for the fatty acid substrate. While there are some studies using pure and saturated or monounsaturated fatty acids, newer studies focus on the use of plant oil as fatty acid source or aim at the esterification of polyunsaturated fatty acids. The combination of the sterol with the polyunsaturated fatty acids leads to a combination of the health benefits of two molecules in one. The application of possible substrates is therefore numerous and could even be further explored. Concerning the sterol specificity of lipases the series from Weber and co-workers can be highlighted. They applied various lipases in solvent-free system in vacuo. Apart from sitostanol and cholesterol also other sterols were evaluated such as 5α-cholestan-3β-ol, thiocholesterol, stigmasterol, ergosterol, 7-dehydrocholesterol and lanosterol. Lanosterol with its 4,4-dimethyl substituents was esterified only to a small extend by R. miehei lipase and thiocholesterol was not esterified by C. rugosa lipase (Weber et al., 2001a, 2001b). One special acid donor was ethyl dihydrocinnamate, which was transesterified with cholesterol by R. miehei lipase to 56% in 96 h (Weber et al., 2001b). Using sterol ester as substrates for transesterification only yielded low amounts of products (Weber et al., 2001a). Overall, it would still be of interest to deeper study the sterol specificity of lipases. As broad as the enzymes and substrates, as broad were also the conditions of the reaction system. Quite a number of studies are using a monophasic solvent system with the enzyme either immobilized or in free form (Table 4). The water content was controlled in some studies by the addition of small amounts of water or molecular sieve as drying agent (e.g. He et al., 2010; Liu et al., 2014; Zheng et al., 2012a). Or the water activity was adjusted before the reaction (Shang et al., 2015) or during the reaction (Teixeira et al., 2011, 2012) with saturated salt solutions. But also solvent-free systems have a potential. The reaction can occur under atmospheric pressure and under reduced pressure. The application of a reduced pressure helps to reduce the melting point, without further increasing the temperature. Another possibility to reduce the melting point of the system is the addition of a fatty acid with a low melting point such as oleic acid. This has been conducted with soybean oil deodorizer distillates (Torres et al., 2007). Further, also non-conventional medias were applied such as supercritical CO2 (Hu et al., 2015; King et al., 2001). Almost solvent-free systems were 28 Esterification of phytosterols applied as well, where only little solvent was added. In the study of Vu and co-workers small amounts of water or hexane was added to the sterol-fatty acid mixture. After an incubation of 6 h no significant difference between the two systems could be measured (Vu et al., 2004). Finally, also systems such as a water-in-ionic liquid microemulsion (Zeng et al., 2015) or in solvent under microwave irradiation (Shang et al., 2015) have been applied. To conclude, the enzymatic esterification of phytosterol with fatty acids has been studied widely. Often yields above 90% in relatively short incubation times were recorded. There is still potential concerning the sterol specificity of lipases and the use of impure substrates such as plant oils or oil deodorizer distillates. However, many studies also use non- commercial enzymes or non-commercial enzyme carriers and are therefore challenging to reproduce by other laboratories. Finally, the commercialization of such processes has to be promoted. 4.2 Steryl phenolates 4.2.1 Chemical synthesis There are several published procedures for the chemical synthesis of steryl phenolates, usually including the protection of the phenolic hydroxyl group, followed by a coupling reaction with the sterol and finally a deprotection. The first process was published by Kondo and co-workers in 1988 (Kondo et al., 1988). In 2001 Condo and colleagues presented a revised procedure, which was even further optimized by Winkler-Moser in 2015 (Condo et al., 2001; Winkler-Moser et al., 2015). Furthermore, a procedure without a protection and deprotection step was presented recently (Fu et al., 2014). Finally, also one process employing coupling of unprotected phenolic aldehydes has been published long time ago (Elenkov et al., 1995). In the work of Kondo and colleagues, trans-4-O-acetylferulic acid was transformed into trans- 4-O-acetylferuoyl chloride by SOCl2 in chloroform. After evaporation, the residue was redissolved in pyridine with stigmastanol and was allowed to stand over night. The crude product was subjected to silica gel chromatography. Finally, deprotection occurred with NaBH4 in chloroform:methanol 1:1 and final silica gel chromatography and recrystallization yielded stigmastanyl trans-ferulate. The coupling reaction gave a yield of 61.6% and the deprotection 82% (Kondo et al., 1988). The main limitation of this method is the synthesis of the highly reactive trans-4-O-acetylferuoyl chloride, which is difficult to purify and has to be handled with special care (Condo et al., 2001). Furthermore, the uncommon deprotection step with NaBH4 could also be improved further (Condo et al., 2001). However, a similar 29 Esterification of phytosterols procedure was applied only recently. The protected caffeic acid or p-coumaric acid was reacted with oxalyl chloride instead of SOCl2 and later with γ-oryzanol sterols (which have been produced by hydrolyzing γ-oryzanol). The deprotection also occurred with NaBH4 (D'Ambrosio, 2013). In 2001 Condo and co-workers had set up e new procedure for the synthesis of steryl ferulates. Protection of the phenolic hydroxyl group in the ferulic acid was achieved with acetic anhydride in pyridine. The trans-4-O-acetylferulic acid was condensed with the phytosterol mixture in the presence of N,N-dicyclohexylcarbodiimide and 4-(dimethylamino)- pyridine in dichloromethane. The separation of the trans-4-O-acetylferulate products from the byproduct N,N-dicyclohexylurea was achieved through preparative liquid chromatography. However, an additional chromatographic step still had to be included to remove further byproducts. A selective deprotection was achieved with K2CO3 in a methanol-chloroform mixture. The yield of the condensation reaction was 43-61% and 71% of the deprotection and purification (Condo et al., 2001). This procedure was further improved by Winkler-Moser and colleagues. First, the synthesis of trans-4-O-acetylferulic acid was optimized. The addition of 4-(dimethylamino)pyridine reduced the reaction time and the product was washed with water and methanol to increase the purity. The condensation step was improved mainly by the purer starting material and an increased addition of 4-(dimethylamino)pyridine. This reduced the reaction time to 1.5 h. The purification was also slightly improved by precipitating the byproduct 1,3-dicyclohexylurea with hexanes, followed by a column chromatography. The yield for the protecting step was 92%, for the coupling reaction 77-90%, and the deprotection yielded 81-97% steryl ferulates (Winkler-Moser et al., 2015). Overall yields and reaction times were improved, however the procedure still includes three synthetic and two chromatographic steps. Another procedure without a protection step, but still including three steps, has been published long time ago. The coupling of the sterol occured from (carbocholesteryloxymethyl)-triphenyl phosphonium bromide with the unprotected phenolic aldehyde by the Wittig reaction (Elenkov et al., 1995). However, the Witting substrate ((carbocholesteryloxymethyl)-triphenyl phosphonium bromide) has to be produced by two synthethic steps including one chromatographic step. This leads to a procedure similar in complexity and workload as the one discussed above. A different approach was presented by Fu and colleagues. Avoiding the protection steps, they coupled gallic acid directly with the phytosterols in tetrahydrofuran in the presence of N,N-dicyclohexylcarbodiimide. The residue was redissolved in ethyl acetate, washed with 30 Esterification of phytosterols brine and subjected to column chromatography. This very simplified procedure gave an overall yield of around 20% (Fu et al., 2014). Although the yield is quite low, this procedure may find its application for laboratory purposes as it is much less labour intensive. To conclude, several procedures for the chemical synthesis of steryl phenolates have been presented. To reach a high yield, labor intensive procedures are required. The shortened procedure of Fu and co-workers on the other hand provides a simple solution, if starting materials are cheap and available in large quantities. However, the overall problematic aspects of a chemical synthesis including formation of byproducts and therefore extended purification requirement cannot be neglected. 4.2.2 Chemoenzymatic synthesis One way applied to improve the yield of an enzymatic esterification is the use of vinyl esters. The liberated vinyl alcohol tautomerizes into acetaldehyde, which makes the process irreversible (Scheme 3) (Villeneuve, 2007). However, it has been shown that acetaldehyde can inhibit certain microbial lipases (Weber et al., 1995). For ferulic acid the difference between vinyl ferulate and ethyl ferulate in lipase catalyzed reactions has been studied in detail (Yu et al., 2010). In this study the two ferulate esters were compared in transesterification reactions with triolein in toluene catalyzed by immobilized C. antarctica lipase B. They concluded that regardless the conditions, greater effectiveness and efficiency were observed for vinyl ferulate over ethyl ferulate in enzymatic feruloylated lipid synthesis. For example the maximum conversion obtained with ethyl ferulate was 70% in 96 h and for vinyl ferulate 91% in 62h. However, not only in this study but also in all studies cited in Table 5 the vinyl ferulate synthesis was catalyzed by mercury acetate. This toxic heavy metal catalyst requires thorough purification if the products should be applied in food. The feasibility of these vinyl esters as substrates for food additive synthesis is therefore questionable. Overall, the use of vinyl esters allows for improved enzymatic reaction yields but their feasibility for large scale applications is doubtful for the reasons discussed. Scheme 3: General lipase catalyzed transesterification of a vinyl ester. The chemoenzymatic synthesis of steryl phenolates has been part of several studies (Table 5). They all followed the procedure discussed above including the synthesis of a vinyl phenolate. This vinyl phenolate synthesis was followed by a purification step on a silica gel column. The yielding vinyl phenolate was then further transesterified enzymatically to the free 31 Esterification of phytosterols sterol. Different sterol substrates were used. In the study from Chigorimbo-Murefu and colleagues dihydrocholesterol and 5α-androstane-3β,17β-diol were used, while the other used different mixtures of phytosterols, containing mainly β-sitosterol. The range of sterol concentration was similar for all studies and was from 7.6 to 20 mg/mL. In contrast to the molar substrate ratio, this ranged from 7.6 times excess of vinyl phenolate to twice the amount of sterol molecules. However, only the study from Wang and co-workers contains data of other substrate ratios leading to the conclusion that an equimolar ratio of both substrates is most suitable (Wang et al., 2015b). This is also the case for other reaction parameters such as the solvent, temperature and time. All studies applied a lipase from C. rugosa as catalyst for the transesterification reaction. They all tested different lipases finding that C. rugosa was the only one catalyzing the reaction. Except Wang and colleagues who also found low activity for other lipases and medium activity for Amano lipase PS IM for the synthesis of steryl cinnamate (Wang et al., 2015b). However, all studies came to the conclusion to apply a non-immobilized lipase from C. rugosa, although at very different concentrations (0.085-100 mg/mL). Of course it is possible that different C. rugosa lipases have been used, the activity is not reported in all studies, but they were all purchased from Sigma-Aldrich. Steryl ferulate were synthesized in all conditions leading to yields from 45 to 90%. Unfortunately in the study from Chigorimbo- Murefu and colleagues no incubation time was reported (Chigorimbo-Murefu et al., 2009). It is therefore difficult to compare the efficiency of the system to the two others. Between the method from Tan and Shahidi and Wang and co-workers the main differences are the enzyme amount and the incubation time (Tan & Shahidi, 2011; Wang et al., 2015b). The enzyme amount was 24 times higher and the incubation time was 10 times longer in the studies from Tan and Shahidi; although the sterol concentration was higher but less than a factor two. With these facts in mind the yields of the two comparable phenolates, vanillate and ferulate, are very high from Wang and co-workers. To summarize all studies applied C. rugosa lipase measuring very different transesterification efficiencies. Finally, the transesterification efficiency of C. rugosa lipase in dependency of the vinyl phenolate structure was mainly studied by Wang and co-workers. However, also the studies of Tan and Shahidi give some information. The yield of steryl caffeate was only about half compared to the steryl ferulate. The second hydroxyl group instead of the methoxy group therefore decreased the yield. In contrast to the sinapate, with an additional methoxy group, where the yield measured was similar to the steryl ferulate. Finally, also the vanillate with the shorter side chain was transesterified to a similar extend (Tan & Shahidi, 2011, 2012a, 2013). These findings were not all confirmed by Wang and colleagues. The yield of the steryl 32 Esterification of phytosterols vanillate was only half to the steryl ferulate, which could be due to the shorter incubation time. Additionally the yield of the vanillate was higher than the p-hydroxybenzoate without the methoxy group in meta-position. Methoxy groups in comparison to hydroxyl groups seem to rather increase the yield. This was the case for the steryl ferulate in contrast to the phytosteryl 3,4-dimethoxycinnamate. Further, the authors concluded that a longer saturated side chain rather decreases the yield and that a double bond in the side chain increases the yield. This was for example the case for cinnamic acid and hydrocinnamic acid. However, one has to keep in mind that the system was optimized for cinnamic acid and it is therefore not surprising that the yield was higher thereof. Overall, the main structure elements influencing the transesterification yield are the length and structure of the side chain, the position and number of hydroxyl groups in combination with methoxy groups. 4.2.3 Enzymatic synthesis The direct enzymatic synthesis of steryl ferulates has been described in 1987 very briefly in an meeting abstract (Seino, 1987). They describe a reaction of cholesterol, β-sitosterol or stigmasterol at 40°C. In conclusion the reaction was more efficient in cyclohexane than in buffer solution and the lipase from Candida showed the highest activity amongst the examined lipases. However, detailed information is missing to perform the reaction accordingly. Another reaction described, which comes close is the transesterification of ethyl dihydrocinnamate with cholesterol catalyzed by immobilized R. miehei lipase (Weber et al., 2001b). However, as the dihydrocinnamic acid is lacking a phenolic hydroxyl group, these reactions cannot be fully compared to the enzymatic synthesis of a steryl ferulate. Mainly due to the structural reasons discussed in the previous chapter. To the best of our knowledge a fully enzymatic synthesis of steryl phenolates, including steryl ferulates, has not been described in detail yet. 33 Esterification of phytosterols - References (Chigorimbo Murefu et al., 2009) (Tan Shahidi, & 2012a, 2011, 2013) (Wang al., et 2015b) 44% 88% 56% 90% 50% 80% yield 1.79% 69.49% 17.31% 23.64% 38.04% 31.95% 21.56% 0 72.11% 27.47% 45.41% Isolated - diol - androstane - Sitosterol(76% Sitosterol(90% - - Sterols Dihydrocholesterol, 5α 3β,17β β pure with other sterols) β 10% with other sterols) hoxycinnamate dimet - coumarate hydroxybenzoate chlorophenylacetate phenylbutyrate phenylvalerate - - - - - ferulate m ferulate Vinyl Vinyl caffeate Vinyl sinapate Vinyl vanillate Vinyl 4 Vinyl vanillate Vinyl 4 Vinyl hydrocinnamate Vinyl 4 Vinyl 5 Vinyl cinnamate Vinyl Vinyl Vinyl 3,4 Vinyl Vinyl phenolates Vinyl ferulate Vinyl methyl methyl - 1:1, hexane 1:1, sitosterol, - butyl substrate ratio substrate - , Conditions butanone (9:1, v/v), v/v), (9:1, butanone v/v), (8:2, butanone - - mg/mLand mg/mLβ mg/mL steroid mM) mg/mL phytosterols, .1 10 7.6 (26 tert 8.7:1, ether,45°C 20 substratehexane ratio 1:2, 2 and 45°C,10 days 13.8 substrateratio 2 and 55°C,24h lipase, type VII, lipase, mg/mL U/mL, mg/mL Table 5: Overview of studiesconducting chemoenzymatic steryl phenolates synthesis. Enzymetype and amount rugosa C. 100mg/mL rugosa C. 8% of thetotal substratesweight, 6 rugosa C. 100 0.085 Substrateratio to refers molarratio substrate vinyl of phenolate to sterol. 34 Feruloyl esterases 5 Feruloyl esterases Feruloyl esterases [E.C. 3.1.1.73] are also known as ferulic acid esterases, cinnamoyl esterases, cinnamic acid hydrolases, or chlorogenate esterases (Faulds, 2010). As their name suggests, they are able to liberate cinnamic acid derivatives, including ferulic acid, from plant cell wall polysaccharides (Benoit et al., 2008). To quantify the feruloyl esterase activity various substrates found application such as feruloylated oligosaccharides, de-starched wheat bran, or methyl or ethyl esters of hydroxycinnamic acids, mainly ferulic acid but also sinapic, p-coumaric, and caffeic acid (Topakas et al., 2007). Mostly the liberated hydroxycinnamic acid is then quantified. Structurally some feruloyl esterases have been shown to have a catalytic triad in the active site and to resemble lipases (Faulds et al., 2005; Hermoso et al., 2004; Schubot et al., 2001; Topakas et al., 2012). They gained interest as feruloyl esterases can help improving saccharification of cereal-derived products, which is important for bioalcohol and animal feed production (Faulds, 2010). Further, they can improve bioavailability of phytonutrients from foods and be a tool to recover and purify ferulic acid from plant materials (Faulds, 2010; Gopalan et al., 2015). As biomass refining, ferulic acid production and plant metabolism are not key points of this thesis only their described naturally occurrence, their classification and their reported ability to accept nonpolar hydroxycinnamates as substrates will be discussed here. 5.1 Occurrence in nature Most isolated feruloyl esterases so far are from fungal origin, less were identified from bacteria or plants (Udatha et al., 2011). Feruloyl esterases produced from microorganisms were reviewed by Topakas and colleagues in 2007. To induce feruloyl esterase production of the microorganism a suitable substrate is crucial. Substrates with high amounts of esterified ferulic acid such as wheat bran, maize bran, or sugar beet pulp and many more have been applied so far. Feruloyl esterases from various genera have been produced such as Aspergillus, Bacillus, Lactobacillus, and Streptomyces. Overall, numerous feruloyl esterases from microorganisms have been produced, purified, and characterized with very diverse substrate specificities (Topakas et al., 2007). Feruloyl esterase activity has also been reported in plants. Earliest it has been quantified in crude barley extract, from barley grains and from malted barley (Sancho et al., 1999). Later on a crude extract from barley malt was partially purified for a feruloyl esterase hydrolyzing glyceryl ferulate (Humberstone & Briggs, 2002). Further, from malted finger millet also a feruloyl esterase has been purified and characterized (Madhavi Latha et al., 2007). If this is a coincidence or not all the reported feruloyl esterase activities in plants are in Poaceae. 35 Feruloyl esterases Further research on feruloyl esterase activity in more plants would be of interest, including characterization of the enzymes involved in this measured activity. Feruloyl esterases involved in the human digestion have been described from two main origins, namely from mucosa and from gut microbiota. For further detail, the dietary implications including feruloyl esterases from gut microbiota have been reviewed recently (Faulds, 2010). Briefly, Andreasen and co-workers showed that esterases all along the intestinal tract of mammals are present, which are able to hydrolyze hydroxycinnamate esters. Mucosa cell-free extracts, with feruloyl esterase activity, gave first indication of human cinnamoyl esterases. Additionally, the feruloyl esterase activity was also measured in the lumen. Further, chlorogenic acid was only cleaved by colonic microbial esterases but not by mucosal esterases (Andreasen et al., 2001a). Moreover, activity towards diferulates also from rats and human colonic microflora and cell-free extracts from intestine mucosa was shown (Andreasen et al., 2001b). Esterases able to hydrolyze hydroxycinnamic esters and diferulates were reported extracellular and intracellular of Caco-2 cells (Kern et al., 2003). There is therefore evidence that human epithelial cells exhibit feruloyl esterase activity. Additionally feruloyl esterases have been extracted from human gut microflora. In a human model colon including the fermentation of wheat bran microbial ferulic acid esterase activity was present (Kroon et al., 1997). In another human colon model extracellular feruloyl esterase activity was measured induced by water-unextractable arabinoxylan (Vardakou et al., 2007). Moreover, isolates from human fecal bacteria hydrolyzed ethyl ferulate and were identified as strains from E. coli, Bifidobacterium lactis and Lactobacillus gasseri (Couteau et al., 2001). Also further intestinal bacterial strains were identified to produce feruloyl esterases such as Lactobacillus acidophilus (Wang et al., 2004). With a growing interest in health promoting foods the role of these enzymes involved in the digestion of substrates such as hydroxycinnamates and derivatives need to be investigated further (Faulds, 2010). 5.2 Classification An early classification of feruloyl esterases into two groups, type A and type B, was based on substrate specificity and the ability to release diferulates. Type A feruloyl esterases are induced by growth on xylan, are able to release diferulates, and prefer methyl hydroxycinnamates with methoxy substitutions. Whereas type B feruloyl esterases are rather induced by growth on sugar beet pulp, do not release diferulates, and prefer methyl hydroxycinnamates with hydroxyl substitution (Crepin et al., 2003; Faulds, 2010; Faulds & Williamson, 1994; Kroon et al., 1997; Kroon et al., 1999). This classification was further improved with the identity of the primary sequences by Crepin and co-workers (Table 6). Not 36 Feruloyl esterases only based on substrate specificity towards methyl hydroxycinnamates and the ability to release diferulates, also primary sequence similarities were taken into account to classify feruloyl esterases into 4 groups named A-D. Based on a phylogenetic tree the earlier classification was mostly supported (Crepin et al., 2004). In 2008 a classification into seven subfamilies has been proposed based on sequences of known and putative genes encoding for feruloyl esterases in fungal genomes. Though, only three of them contain biochemically characterized feruloyl esterases (Benoit et al., 2008). Even further analysis led to a classification into twelve families based on amino acid sequence information (Udatha et al., 2011). Nevertheless, the biochemical classification proposed by Crepin and co-workers still finds wide application in scientific papers (Gopalan et al., 2015). Table 6: Classification of microbial feruloyl esterases as proposed by Crepin et al., 2004. Type A Type B Type C Type D M. thermophila T. stipitatus P. fluorescens Example A. niger FaeA FaeB FaeC XYLD Ferulic acid, Ferulic acid, Ferulic acid, Ferulic acid, Hydrolyze caffeic acid, caffeic acid, sinapic acid, caffeic acid, methyl ester of p-coumaric acid, p-coumaric acid, p-coumaric acid p-coumaric acid sinapic acid sinapic acid Release of Yes (5-5’) No No Yes (5-5’) diferulic acid Chlorogenate Sequence Acetyl xylan Lipase esterase, Xylanase similarity to esterase tannase Content adapted from (Crepin et al., 2004). 5.3 Hydrolysis of nonpolar substrates As discussed above, enzyme activity of feruloyl esterases is determined by quantification of released ferulic acid is from methyl or ethyl ferulate, sugar esters or even biological samples such as wheat straw. The data on more nonpolar samples is rather scarce. In an early study Aliwan and colleagues analyzed a feruloyl esterase FAE-III (later on renamed to AnFaeA (Faulds, 2010)) from A. niger. As the primary sequence of these enzymes shows similarities to fungal lipases, they analyzed its lipase activity in comparison to two lipases and two ferulic acid substrates. Against methyl ferulate low activity of the lipases was measured, while the feruloyl esterase showed very high activity. For the natural diglycerides, a lipase substrate, it was exactly the opposite; the hydrolytic activity of FAE-III was very low. And for olive oil 37 Feruloyl esterases triglycerides no activity of the feruloyl esterase could be measured at all. They concluded that FAE-III does not exhibit significant lipase activity (Aliwan et al., 1999). In a study of Koseki and co-workers a feruloyl esterase from A. amawori was engineered and the substrate specificity was evaluated. As nonpolar substrates α-naphthyl esters were used. The wild type enzyme did not show any hydrolytic activity against decanoic acid ester and longer acid esters, while some mutants and R. miehei lipase still hydrolyzed these substrates. Finally, also the kinetic parameters of the enzymes towards α-naphthyl butyrate and α-naphthyl caprylate were determined. For the all enzymes Km and kcat were lower for α-naphthyl caprylate (Koseki et al., 2005). However, also in this study, the wild-type feruloyl esterase did not show activity towards long-chain α-naphthyl esters. After the two studies discussed above using type A feruloyl esterases and non-ferulated, nonpolar substrates, several studies were published using ferulate esters up to C4 linear and branched esters for a type B (Topakas et al., 2012) and three type C feruloyl esterases (Moukouli et al., 2008; Vafiadi et al., 2006; Vafiadi et al., 2005). The affinity towards branched and sterically more demanding esters was higher and they were hydrolyzed more efficiently by StFaeC (Vafiadi et al., 2005). Further, FoFaeC showed least affinity towards n-butyl ferulate, and methyl ferulate was hydrolyzed the fastest and with highest efficiency compared with other ferulates (Moukouli et al., 2008). Similarly, TsFaeC showed also lowest affinity towards n-butyl ferulate and ethyl ferulate was hydrolyzed the fastest and most efficient (Vafiadi et al., 2006). Finally, the type B feruloyl esterase from M. thermophila (earlier S. thermophile) showed highest affinity towards methyl ferulate and secondly towards the butyl ferulates. Highest kcat was observed for n-propyl ferulate and highest catalytic efficiency for n-butyl ferulate (Topakas et al., 2012). 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C., Ma, F. L., & Feng, Y. Q. (2015). Enzymatic deacidification of the rice bran oil and simultaneous preparation of phytosterol esters-enriched functional oil catalyzed by immobilized lipase arrays. Rsc Advances, 5(86), 70073-70079. 51 References Zheng, Y., Quan, J., Ning, X., & Zhu, L. M. (2008). Enhancement in the Synthesis of Novel Feruloyl Lipids (Feruloyl Butyryl Glycerides) by Enzymatic Biotransformation Using Response Surface Methodology. Journal of Agricultural and Food Chemistry, 56(23), 11493-11498. Zhu, D., & Nyström, L. (2015). Differentiation of rice varieties using small bioactive lipids as markers. European Journal of Lipid Science and Technology, 117(10), 1578-1588. Zoumpanioti, M., Merianou, E., Karandreas, T., Stamatis, H., & Xenakis, A. (2010). Esterification of phenolic acids catalyzed by lipases immobilized in organogels. Biotechnology Letters, 32(10), 1457-1462. 52 Part B - Research Papers Part B - Research Papers Aline Schär and Laura Nyström (2015). High yielding and direct enzymatic lipophilization of ferulic acid using lipase from Rhizomucor miehei. Journal of Molecular Catalysis B: Enzymatic, 118, 29-35. Aline Schär and Laura Nyström (2016). Enzymatic synthesis of steryl ferulates. European Journal of Lipid Sciences and Technology. doi: 10.1002/ejlt.201500586. Aline Schär, Silvia Liphardt and Laura Nyström. Enzymatic synthesis of steryl hydroxycinnamates and their antioxidant activity. Submitted manuscript (June 2016). Aline Schär, Isabel Sprecher, Evangelos Topakas, Craig B. Faulds and Laura Nyström. Hydrolysis of nonpolar alkyl ferulates by feruloyl esterases. Submitted manuscript (June 2016). 53 Part B: Enzymatic esterification of ferulic acid High yielding and direct enzymatic lipophilization of ferulic acid using lipase from Rhizomucor miehei Reprinted with permission from Aline Schär and Laura Nyström (2015). Journal of Molecular Catalysis B: Enzymatic, 118, 29-35. Copyright (2015) Elsevier. Abstract Ferulic acid is an abundant phenolic acid and a good antioxidant that occurs naturally in free form or esterified. The structure of this hydroxycinnamic acid, with a hydroxyl group in para-position, makes enzymatic esterification with lipases challenging. Adjusted lipophilicity of the ferulic acid as an antioxidant is crucial for complex food matrices, calling for a simple esterification method. Esterification of ferulic acid with ethanol and decanol in n-hexane using immobilized lipase from R. miehei was optimized using surface response methodology. After 72 h, the yields were 76% and 88% for ethyl ferulate and decyl ferulate, respectively. Furthermore, ferulate esters of primary alcohols with varying chain lengths from C-2 to C-18 were also synthesized, with yields ranging from 76% to 92%. Finally, ferulic acid was preferably esterified to short chain alcohols in a mixture of primary alcohols. This study provides simple and efficient methods for the enzymatic esterification of ferulic acid. Keywords: Phenolic acid lipophilization / Ferulic acid / R. miehei lipase / Alkyl ferulates / Enzymatic esterification Highlights Ferulic acid was efficiently esterified with primary alcohols by R. miehei lipase Yields were 76% and 88% for ethyl ferulate and decyl ferulate at optimized conditions The syntheses of C3, C4, C6, C8, C14 and C18 ferulate esters were also successful Short alcohols were preferentially esterified with ferulic acid by R. miehei lipase 55 Part B: Enzymatic esterification of ferulic acid 1. Introduction Phenolic acids are secondary plant metabolites and are powerful, hydrophilic antioxidants present in particular in vegetables, fruits, spices, and grains (Figueroa-Espinoza & Villeneuve, 2005; Yu et al., 2010; Zoumpanioti et al., 2010). There is evidence that ferulic acid, which is abundant in plant cell walls (Yu et al., 2010), has potential to treat Alzheimer’s disease, cancer, cardiovascular disease, diabetes mellitus, and skin disease (Mancuso & Santangelo, 2014). For applications in oil-based or multiphase systems, the lipophilicity of the antioxidant, which can be adjusted through lipophilization, is crucial (Laguerre et al., 2013). Especially in multiphase food systems a critical chain length of the esterified alcohol must be found to reach highest antioxidant activity (Laguerre et al., 2013). Alkyl ferulates appear in nature in suberin, a specific plant cell wall component, in which ferulic acid esters of the 1-alkanols of C-16 to C-30 can be found (Bernards, 2002), for example, in potato tubers (Yunoki et al., 2004). The ethanol ester of ferulic acid, ethyl ferulate, has been quantified in wine and in sake (Hashizume et al., 2013; Hixson et al., 2012). Furthermore, differences in bioactivity within alkyl ferulates and between free and esterified hydroxycinnamates have been shown by several studies (Cione et al., 2008; Garrido et al., 2012; Jayaprakasam et al., 2006; Kondo et al., 2013). To fully capitalize on the antioxidant activity and bioactivity of ferulic acid, an enzymatic esterification process is needed to produce various alkyl ferulates. Ferulic acid belongs to the family of hydroxycinnamic acids with an unsaturated side chain and one hydroxyl group in para-position. It has been observed several times that this combination either partially or fully inhibits esterification (Guyot et al., 1997; Stamatis et al., 2001). Earlier studies have shown low activity and yields from trace amounts to 30% for ferulic acid esterification reactions in solvent-free systems using commercial lipase Novozym® 435 (immobilized Candida antarctica lipase B) (Guyot et al., 1997; Stamatis et al., 1999, 2001), in anhydrous solvents such as n-hexane, butanone, or mixtures thereof (Compton et al., 2000; Katsoura et al., 2009; Sabally et al., 2005; Safari et al., 2006; Yang et al., 2012b), and similar results with immobilized R. miehei lipase (Katsoura et al., 2009; Stamatis et al., 1999, 2001). Candida antarctica lipase B and Rhizomucor miehei lipase immobilized in organogels did not show any esterification activity towards ferulic acid in a solvent-free system (Zoumpanioti et al., 2010). A yield of 87% ethyl ferulate from ferulic acid and ethanol after 48 h was reached in a study by Lee et al., 2006, where also lipase B from C. antarctica was used, but in isooctane (Lee et al., 2006). Finally, Yoshida et al. developed a continuous solvent-free system to esterify ferulic acid and 1-pentanol, 1-hexanol, or 1-heptanol by Novozym® 435 (Yoshida et 56 Part B: Enzymatic esterification of ferulic acid al., 2006). However, efficient enzymatic esterification reaction systems have only rarely been described therefore, additional possibilities are needed. As an alternative to lipases, other enzymes, such as ferulic acid esterases (FAEs), have also been applied to directly esterify ferulic acid in microemulsions (Giuliani et al., 2001), or to transesterify methyl ferulate in detergentless microemulsions (Vafiadi et al., 2008). Unfortunately, microemulsion systems and also solvent-free systems are often restricted to a narrow range of alcohol chain length that can be utilized. A reaction system in an organic solvent, on the other hand, offers higher flexibility. In addition to direct esterification, lipases (Compton et al., 2000; Sun et al., 2012; Yang et al., 2012a; Yu et al., 2010; Zheng et al., 2009) and feruloyl esterases (Vafiadi et al., 2008) have been applied in transesterification reactions using ethyl or methyl ferulate as substrates. These studies generally demonstrated higher yields by transesterification compared to direct esterification, however these studies have not fully solved the problem of an enzymatic synthesis of ferulate esters, which would be more environmentally friendly, specific, and require less purification steps (Figueroa- Espinoza & Villeneuve, 2005). There are indications that the immobilized lipase from R. miehei is higher in efficiency than the C. antarctica lipase B (Katsoura et al., 2009; Stamatis et al., 1999, 2001). Notably in the study performed by Katsoura et al. the esterification yield of ferulic acid with ethanol in n-hexane were 11.4% and 24.3% for the immobilized C. antarctica lipase B and R. miehei lipase, respectively, by applying the same mass of immobilized lipase (Katsoura et al., 2009). Additionally R. miehei lipase has been investigated much less frequently on its ability to directly esterify ferulic acid. However, most of the studies thus far have shown rather unsatisfactory yields for enzymatic synthesis of alkyl ferulates. The aim of this study was to determine a direct and efficient process for esterification of ferulic acids with various primary alcohols using the immobilized lipase from R. miehei (Lipozyme® RM IM). The main factors affecting esterification yield, such as ferulic acid and alcohol concentrations, temperature, reaction time, and enzyme-to-substrate ratio have been investigated on the synthetic reaction of ethyl and decyl ferulate. Reaction conditions were explored using surface response methodology. Finally the esterification activity for various alcohols was evaluated. 57 Part B: Enzymatic esterification of ferulic acid 2. Materials and Methods 2.1 Chemicals and enzymes Ferulic acid, ≥99% was purchased from Sigma-Aldrich, Switzerland. Methyl ferulate, 99% and ethyl ferulate, 98% were purchased from Alfa Aesar, Switzerland. Wako Pure Chemical Industries, Japan provided the γ-oryzanol (min. 98%). All used solvents were of HPLC grade. Lipase from R. miehei (formerly known as Mucor miehei) immobilized on macroporous ion- exchange resin , >30 U/g (lot result: 63 U/g, product number: 62350) was provided by Sigma- Aldrich, Switzerland. 1 U refers to the amount of enzyme, which liberates 1 μmol stearic acid per minute at pH 8.0 and 70 °C from tristearin. Figure 1: General enzymatically catalyzed reaction examined in this study. 2.2 Enzymatic synthesis For the enzymatic esterification of ferulic acid the total reaction volume was 3 mL. The ferulic acid and the enzyme were weighed into a 10 mL glass tube with a Teflon-lined screw cap before the n-hexane, dehydrated over 4 Å molecular sieve before use, and finally the alcohol were added. Before incubation the samples were shaken thoroughly and then incubated without shaking (Figure 1). For a typical ethyl ferulate synthesis experiment, 5 mg of ferulic acid, 12.5 mg enzyme, 2.95 mL of n-hexane and 50 µL of ethanol were incubated together. Whereas for a typical decyl ferulate synthesis experiment 7.5 mg of ferulic acid, 18.8 mg enzyme, 2.85 mL of n-hexane and 150 µL of decanol were combined. Blank reactions were carried out under similar conditions, where no product could be detected. Aliquot samples of 50 µL were collected during incubation, and the samples were evaporated under a gentle nitrogen stream at 50°C. Ferulates were redissolved in 500 µL solvent B (see HPLC- conditions below) and filtered through a 0.45 µm PTFE filter into a HPLC vial. The synthesis of ethyl ferulate and decyl ferulate was optimized using a 3-level-5-factor design. Different solvents were tested for the synthesis of ethyl ferulate. Based on the optimal conditions determined for ethyl ferulate and decyl ferulate, optimal conditions for the other alcohols were derived based on the lengths of the alcohol chain. The primary factors 58 Part B: Enzymatic esterification of ferulic acid affecting optimal conditions were found to be the concentrations of alcohol and ferulic acid, which was then related, linearly, to the alcohol chain length to find optimal conditions for all alcohol chain lengths used. 2.3 HPLC analysis and quantification Ferulic acid and ferulate esters were analyzed using high performance liquid chromatography (HPLC, Agilent 1100, Switzerland). A reverse phase xBridgeTMPhenyl column from Waters, with a particle size of 3.5 µm and gradient elution at room temperature was used for separation of the ferulates. Solvent A was 1% acetic acid in water and solvent B composed of acetonitrile, water, butanol, and acetic acid with a ratio of 88:6:4:2, respectively. The flow was set to 0.6 mL/min, and the elution program was a linear gradient from 75:25 (A:B) to 100% B for 3 min, isocratic flow of 100% B for 5 min, 4 min of a linear gradient to 75:25 (A:B) and 2 min isocratic 75:25 (A:B). For the analysis of dodecyl ferulate and γ-oryzanol, the isocratic flow of 100% B was extended to 11 min and the subsequent linear gradient from to 75:25 (A:B) was shortened to 3 min. Detection of the alkyl ferulates was achieved with a diod array detector (DAD) at 325 nm. For quantification, an external calibration (0.1-13 nmol/injection) was conducted using ferulic acid and commercially available ferulate esters (methyl ferulate, ethyl ferulate, and steryl ferulates), which were used to create one calibration curve of the response versus the molar concentration. This led to a linear regression with a correlation factor of R2=0.996. The UV response originated from the ferulic acid and was not influenced by the alcohol esterified to it. Therefore, quantification of all ferulate esters was calculated based on this calibration. The yield was calculated based on the amount of synthesized ester detected and is presented as averages with standard deviation in brackets. Identification was supported by the specific UV spectra of ferulic acid. Additionally, the reaction product identities were confirmed by mass spectrometry using a SynaptTM G2 time-of-flight mass spectrometer from Waters. The sample was introduced by direct infusion using ESI negative ion mode with the following settings: the voltages of capillary, sampling cone and extraction cone were 2.5 kV, 60 V and 4 V, respectively. The temperature was 120°C for source and 250°C for desolvation. The nitrogen flow rates for cone and desolvation were 20 and 800 L/h, respectively 2.4 Experimental design and statistical analysis A 3-level-5-factor Box-Behnken design was employed. This design requires 40 experiments and a center sample (all coded variables equal to 0), which was repeated six times, making overall 46 measurements. The experiments, which are shown in Table 1, were performed in 59 Part B: Enzymatic esterification of ferulic acid random order. The variables used were time (24-72 h), temperature (55-65°C), enzyme-to-substrate ratio (1-4 g/g, i.e. 1-4 g of the immobilized enzyme preparation per gram of ferulic acid substrate or 0.012-0.049 U/µmol or 100-400% (wt% of ferulic acid)), and the concentrations of ferulic acid and alcohol. For the optimization of ethyl ferulate synthesis, the ferulic acid concentration varied from 0.833 to 2.5 mg/mL and ethanol concentration varied from 8.33 to 25 µL/mL. For the decyl ferulate synthesis, the ferulic acid concentration was 1.67 to 5 mg/mL and the decanol concentration ranged from 25 to 75 µL/mL. These parameters were chosen based on preliminary experiments (data not shown). Optimal conditions were confirmed in triplicate analysis.The experimental data collected was analyzed using the software The Unscrambler X (CAMO Software, Oslo, Norway) to fit the second-order polynomial equation 1: 5 5 4 5 2 푌 = 훽푘0 + ∑ 훽푘푖푥푖 + ∑ 훽푘푖푖푥푖 + ∑ ∑ 훽푘푖푗푥푖푥푗 (1) 푖=1 푖=1 푖=1 푗=푖+1 where 푌 corresponds to the response (molar yield %), 훽푘0 , 훽푘푖 , 훽푘푖푖, and 훽푘푖푗 are constant coefficients, and the uncoded independent variables are represented by 푥푖. Table 1: Coded experiments conducted following the 3-level-5-factor Box-Behnken design with five variables, excluding the center sample. Coded variables are: time (24, 48, 72 h), temperature (55, 60, 65°C), enzyme-to-substrate ratio (1, 2.5, 4 g/g), ferulic acid (2.5, 5, 7.5 mg/3 mL for ethyl ferulate, 5, 10, 15 mg/ 3 mL for decyl ferulate), and alcohol (25, 50, 75 µL/3 mL for ethanol and 75, 150, 225 µL/ 3 mL for decanol). ID 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 x1: temperature -1 0 0 0 0 0 0 1 -1 -1 -1 -1 -1 -1 0 0 0 0 0 0 x2: time -1 -1 -1 -1 -1 -1 -1 -1 0 0 0 0 0 0 0 0 0 0 0 0 x3: ferulic acid 0 -1 0 0 0 0 1 0 -1 0 0 0 0 1 -1 -1 -1 -1 0 0 x4: alcohol 0 0 0 -1 1 0 0 0 0 0 -1 1 0 0 0 -1 1 0 -1 1 x : enzyme: 5 0 0 -1 0 0 1 0 0 0 -1 0 0 1 0 -1 0 0 1 -1 -1 substrate ratio ID 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 x1: temperature 0 0 0 0 0 0 1 1 1 1 1 1 -1 0 0 0 0 0 0 1 x2: time 0 0 0 0 0 0 0 0 0 0 0 0 1 1 1 1 1 1 1 1 x3: ferulic acid 0 0 1 1 1 1 -1 0 0 0 0 1 0 -1 0 0 0 0 1 0 x4: alcohol -1 1 0 -1 1 0 0 0 -1 1 0 0 0 0 0 -1 1 0 0 0 x : enzyme: 5 1 1 -1 0 0 1 0 -1 0 0 1 0 0 0 -1 0 0 1 0 0 substrate ratio 60 Part B: Enzymatic esterification of ferulic acid 3. Results and Discussion 3.1 Optimization of ethyl ferulate synthesis Direct esterification of ferulic acid with ethanol using immobilized lipase from R. miehei in n-hexane was studied, and the experimental conditions were optimized. Pre-experiments showed a suitable range for the reaction parameters. One of these factors that differs strongly from previous studies is ferulic acid concentration, which depends on the solvent and could be set higher for solvent-free systems. The solubility of ferulic acid in hexane is limited, but enzymatic esterification with high yields can still be reached. Also an addition of a 3Å powdered molecular sieve was tested. The addition of 10 mg/mL and 20 mg/mL resulted in around one third and one sixth of the yield without an addition of molecular sieve. The factor molecular sieve addition was therefore excluded from the optimization procedure. For optimization of ethyl ferulate synthesis, the second-order polynomial model (eq. 1) was fitted to the experimental data. The model represented an adequate explanation of variance in the data, as displayed in the statistically insignificant lack of fit (p=0.22, Table 2). Furthermore, most linear (except temperature) and all quadratic predictors had a significant influence on the model (Table 3), and the model exhibited a high coefficient of determination (0.96), which indicates a good fit of the experimental data with the calculated model. The alcohol concentration had the highest β-coefficient of the linear and quadratic factors, indicating high influence. Of the interaction factors studied, only three were found to have significant influence on the yield, namely interactions of ferulic acid and ethanol concentration, ferulic acid and enzyme-to-substrate ratio, and ethanol concentration and enzyme-to-substrate ratio. Based on these results, the model chosen appears to be suitable to predict the yield from enzymatic ethyl ferulate synthesis. Table 2: Analysis of variance for the Box-Behnken designs for ethyl ferulate synthesis and decyl ferulate synthesis. ss: sum of squares, df: degree of freedom, p-value = level of significance. Ethyl ferulate Decyl ferulate ss df p-value ss df p-value Model 10241.80 20 0.00 5801.10 20 0.00 Linear 6949.98 5 0.00 3223.64 5 0.00 Interaction 683.45 10 0.00 1296.45 10 0.00 Quadratic 3703.46 5 0.00 1847.65 5 0.00 Lack of fit 378.73 20 0.22 630.39 20 0.05 R-square 0.96 0.90 61 Part B: Enzymatic esterification of ferulic acid Figure 2: Contour plots of molar yield of ethyl ferulate synthesis at constant temperature (61°C), which has the lowest β-coefficient for this model. The gray scale indicates the predicted molar conversion at given conditions for the ethyl ferulate synthesis. □: < 30%, ■: 35-45%; ■: 45-55%; ■: 55-65%; ■: 65-70%; ■: 70-75%. 62 Part B: Enzymatic esterification of ferulic acid In addition to the numerical results, the influences of the single factors on the yield are presented in the contour plots of the model (Figure 2). The contour plots represent a fixed temperature of 61°C, the temperature with the highest predicted yield, and the factor with the smallest linear β-coefficient. A yield above 70% was reached at a concentration of 50 µL/3 mL ethanol for an enzyme-to-substrate ratio equal to or greater than 2.5 g/g, and with rather low ferulic acid concentrations. There was no clear increase in yield when the enzyme-substrate-ratio of 2.5 was changed to 4 g/g, but a clear trend towards a better yield was seen at low ferulic acid concentrations. A maximum conversion was predicted at medium concentration of ethanol, which is logical because at a mid-level concentration the inhibitory effects are balanced with the positive effects of a high substrate ratio and increased solubility of ferulic acid. Furthermore, a trend towards maximum yield at medium incubation time was also observed. It is possible that after a certain time the reaction would tend towards hydrolysis, but for the model applied, this explanation does not seem plausible because the solubility of ferulic acid is drastically lower compared to ethyl ferulate in hexane and only very little water is present. The overall optimal conditions for ethyl ferulate synthesis based on the fitted model were identified as: 61°C, 52 h, 3.75 mg/3 mL ferulic acid, 57.5 µL/3 mL ethanol, and an enzyme-to- substrate ratio of 2.5 g/g, which predicted a yield of 74.7%. In a second step, the influence of the solvent was tested by applying the optimized conditions described above using various solvents: hexane, cyclohexane, octane, toluene, butanone, and acetone, as well as a solvent-free treatment in ethanol, each conducted in duplicate. The observed yields after 52 h were 64.35, 48.0, 49.3, and 31.7% for hexane, cyclohexane, octane, and toluene, respectively. In butanone, acetone, or ethanol very little to no ethyl ferulate was detected. This corresponds well with former studies (Lee et al., 2006; Zheng et al., 2009), in which higher yields for esterification or transesterification of ferulic acid was found in nonpolar solvents using the lipase B from C. antarctica. Furthermore, in solvent-free systems using the lipase from R. miehei low yields or no reactions were observed for the esterification of ferulic acid (Stamatis et al., 2001; Zoumpanioti et al., 2010). The results of this study clearly showed the highest yields in hexane, for which solvent the synthesis was optimized. 63 Part B: Enzymatic esterification of ferulic acid Table 3: β-coefficients and corresponding p-values of the ethyl ferulate and decyl ferulate synthesis optimization. The variables refer to x : temperature; 1 x : time; x : ferulic acid; x : alcohol; x : enzyme-to- 2 3 4 5 substrate ratio. p-value = level of significance. Ethyl ferulate Decyl ferulate β-coefficient p-value β -coefficient p-value 69.06 88.46 β0 x1 1.89 0.0788 4.09 0.0040* x 4.22 0.0004* 12.25 0.0000* 2 x3 -10.14 0.0000* -0.76 0.5611 x4 14.66 0.0000* 5.03 0.0006* x 9.76 0.0000* 3.00 0.0286* 5 x1*x2 -1.11 0.5968 -4.39 0.1012 x1*x3 -1.16 0.5796 -1.44 0.5822 x *x 1.96 0.3504 -0.11 0.9663 2 3 x1*x4 -1.36 0.5163 1.41 0.5887 x2*x4 -3.31 0.1213 0.12 0.9625 x3*x4 5.61 0.0117* -0.53 0.8373 x1*x5 -3.85 0.0735 -10.43 0.0004* x2*x5 1.64 0.4339 -12.70 0.0000* x3*x5 4.62 0.0343* -5.53 0.0421* x4*x5 -9.02 0.0002* -0.06 0.9816 2 x1 -3.51 0.0188* -5.71 0.0031* 2 x2 -4.94 0.0016* -6.35 0.0013* 2 x3 -7.78 0.0000* -2.93 0.1056 2 x4 -16.72 0.0000* -2.17 0.2246 2 x5 -6.89 0.0000* -11.20 0.0000* *significant at p = 0.05 64 Part B: Enzymatic esterification of ferulic acid Due to the significant influence of the ethanol concentration, which is demonstrated by the high linear and quadratic β-coefficient of 14.7 and -16.7 observed in the first model, this factor was reexamined by testing the following ethanol concentrations in triplicates: 42.5, 50, 57.5, 65 µL/3 mL. Molar conversions of 62.6 (1.7)%, 69.3 (1.1)%, 64.5 (2.5)%, and 56.1 (2.6)%, respectively were found after 52 h. The predicted values for these conditions were 68.1, 72.9, 74.7, and 73.5%, respectively. Generally, the values measured were somewhat lower than the predicted values, and the optimum slightly shifted. Repeating experiments showed maximum conversion rather at 50 µL/3mL than as by the model predicted at 57.5 µL/3mL. Additionally, the factor time needed further examination. The model predicts a slight decrease of yield when moving from 52h to 72h. To confirm this, the sample with 57.5 µL/3mL ethanol was incubated for 72h. The molar conversion after 52 h was 64.5 (2.5)% and after 72h a conversion of 76.2 (2.0)% was detected (predicted yield 70.6%). This experiment shows that the model-predicted decrease in yield over longer incubation times does not correspond well with reality. Therefore a longer incubation for 72h is more suitable to reach a higher yield. After method validation, the optimal ethanol content was observed to be 50 µL/3 mL and the optimal time 72h, while other factors remained the same as predicted (61°C, 3.75 mg/3 mL ferulic acid, and an enzyme-to-substrate ratio of 2.5 g/g). The predicted value for the conversion of ethanol and ferulic acid to ethyl ferulate with these conditions was 70.6%, and the actual measured value was 76.2 (2.0)%. Compared to other studies using lipase from R. miehei, this is the first time a reasonable yield of enzymatic ferulic acid esterification in a relatively short time has been reported. The main difference between this study and those reported in current literature is the use of a hexane system instead of a solvent-free system (Stamatis et al., 1999) or a polar solvent (Compton et al., 2000). 65 Part B: Enzymatic esterification of ferulic acid 3.2 Optimization of decyl ferulate synthesis The optimization, which was performed for ethyl ferulate synthesis was repeated in similar manner for decyl ferulate. However, due to essential differences in the solubility of the product and the molar mass of the alcohol, higher ferulic acid and higher volumetric alcohol concentrations were applied for the decyl ferulate synthesis. Also, in the case of decyl ferulate a second-order polynomial model (equation 1) was fitted to the experimental data. This time the coefficient of determination was calculated to 0.90, which is somewhat lower than that of the ethyl ferulate synthesis. In addition, the p-value of the lack of fit is, in this case, lower, specifically 0.05, which is just the required level of insignificance to demonstrate an adequate explanation of variance by the model. Indicating that technically the requirements are met but that there is a lot of variance in the data which cannot be explained by the fitted model. For the optimization of decyl ferulate synthesis all linear factors except the ferulic acid concentration had a significant impact on the yield (Table 3), with time exhibiting the greatest influence. Concerning the quadratic factors, the ferulic acid concentration and the alcohol concentration squared did not have significant influence. The enzyme-to-substrate ratio squared had a very high β-coefficient, indicating a strong influence. Two interaction terms showed very high β-coefficients: temperature*enzyme-to-substrate ratio, and time*enzyme- to-substrate ratio, and therefore had a significant influence on the molar conversion. Generally, the β-coefficients for the decyl ferulate were somewhat lower than those in the ethyl ferulate model, which indicates a lower sensitivity of the yield with respect to changing factors. However, in order to predict the yield of decyl ferulate, this model appears to be adequate. 66 Part B: Enzymatic esterification of ferulic acid Figure 3: Contour plots of molar of molar yield of decyl ferulate synthesis at 8.5 mg ferulic acid / 3 mL hexane, which has the lowest β-coefficient for this model. The gray scale indicates the predicted molar conversion at given conditions. □: < 50%, ■: 50-60%; ■: 60-70%; ■: 70-80%; ■: 80-90%; ■: 90-100%. 67 Part B: Enzymatic esterification of ferulic acid The influence of the independent variables on the yield can be further examined in the contour plots in Figure 3. The plots are displayed with a fixed ferulic acid concentration, which was the only linear factor with insignificant influence on the yield. In calculating the expected yields, several maxima above 95% yield over the entire space were observed. One optimum was at a similar temperature and enzyme-substrate ratio as was found for the ethyl ferulate synthesis. For the decyl ferulate synthesis, a reasonably clear pattern of higher yields at longer incubation times was observed. Concerning the enzyme-to-substrate ratio, a slight increase from 1 to 2.5 g/g was exhibited, mainly on the time scale. However, a higher enzyme-to-substrate ratio of 4 g/g did not result in a higher, but rather a lower yield. This phenomenon has been observed several times before and was attributed to catalyst aggregation at excess enzyme and therefore mass transfer limitations (Šabeder et al., 2006; Sun et al., 2012). Further, the water content, which is increased with an increasing lipase load, may play a role (He et al., 2012; Šabeder et al., 2006). The increased amount of water in the reaction system may cause the reverse reaction and lead to a decreased yield. This seems most likely for this reaction system, especially since this phenomenon was only observed in the case of decyl ferulate, where higher substrate and therefore higher absolute concentrations of enzyme were applied. However, a higher enzyme-to-substrate ratio did not appear necessary, because good yields were already reached at smaller enzyme amounts. The contour plots at an enzyme-substrate ratio of 1 were generally steeper, indicating that the reaction system is more sensitive to small changes in the reaction conditions. Therefore, an enzyme-substrate ratio of 2.5 was defined as optimal. Unlike for the ethanol, no clear optimum for the decanol concentration was observed in the contour plots. However, it seems that the higher the decanol concentration, the higher the yield. Overall, the results of the optimization are not as clear as for the ethyl ferulate synthesis. Therefore, the optimal conditions were set similar to those from the ethyl ferulate synthesis: 61°C, 72 h, 8.5 mg/3 mL ferulic acid, 75 µL/mL decanol, and an enzyme-to-substrate ratio of 2.5 g/g. The calculated yield for these conditions obtained from the model is 97.6% and the measured yield, in triplicate, was significantly lower at 88 (2.0)%, which is not sufficiently similar to confirm the model. However, the low p-value for the lack of fit and the reasonably low R-square value also indicated that the model was not a very good fit. Furthermore, the optimal conditions found lay on the edge of the design space, where the model is not as strong. Nevertheless, it can be said that the synthesis of decyl ferulate is less sensitive to changing factors as the synthesis of ethyl ferulate, efficient reaction conditions could still be found. For the synthesis of decyl ferulate a good yield (88%) could be reached, which is a little higher than the one for ethyl ferulate (76%), and a higher concentration of ferulic acid 68 Part B: Enzymatic esterification of ferulic acid can be used. However, further optimization leading to even higher yields may still be possible. 3.3 Esterification of various alcohols After the optimization of the ethyl ferulate and decyl ferulate synthesis, esterification of ferulic acid with other alcohols was tested. The concentrations applied were adjusted linearly up to C10, based on the optimal conditions found for C2 and C10 as described above, and the conditions applied for the esterification of tetradecanol and octadecanol were equal to the ones for decyl ferulate. For all reactions, the temperature was held constant at 61°C, reaction time was 72 h, and the enzyme-to-substrate ratio was 2.5 g/g. In Figure 4, the molar yields for the esterification of ferulic acid with ethanol, propanol, butanol, hexanol, octanol, decanol, tetradecanol, octadecanol, isopropanol, and 2-octanol are presented, which ranged from 76(2)% for ethyl ferulate to 92(5.2)% for hexyl ferulate. The yields of the esters with longer alcohols did not significantly differ and varied from 84-90%. The higher yield for the longer ferulate ester may be explained by the higher concentration of alcohol which can be applied without negative effects on enzyme activity. This leads not only to a higher substrate concentration but also to an increased solubility of ferulic acid. Figure 4: Molar yield of ferulic acid ester synthesis based on carbon chain length of the alcohol (n=3, error bars referring to standard deviation). Reaction conditions were: 72h, 61°C, enzyme to substrate ratio 2.5, ferulic acid and alcohol concentration linearly increasing from C2 to C10 from 6.4mM and 0.29M to 14.6mM and 0.39M, respectively. For C14 and C18 the conditions of C10 were applied. 69 Part B: Enzymatic esterification of ferulic acid The immobilized lipase from R. miehei was also tested for its ability to esterify ferulic acid with secondary alcohols, such as isopropanol and 2-octanol, but for these secondary alcohols the observed yields were drastically lower at 29(1.1)% and 11(1.2)%, respectively (Figure 4). Lower yields of the secondary esters could be expected due to the 1,3-specificity of R. miehei lipase. This 1,3-specificity can be translated to a lower activity towards secondary alcohols for other ester bond hydrolysis than triglycerides (Hari Krishna & Karanth, 2002). Additionally, secondary alcohols are sterically more hindered, which also influences their reactivity. Reflecting to that the yield for isopropyl ferulate at 29% is rather high, although the yield seems to decrease with a decreasing polarity of the alcohol. Generally, it can be said that using this process, all primary alcohols (from C2 on) can be directly esterified to ferulic acid. Compared to solvent-free systems, as variously applied in previous studies, the primary advantage of the hexane system is flexibility of the alcohol, as has been demonstrated in this study. This allows users to directly esterify the requested alcohol, which would be necessary for the application in question, and a subsequent transesterification can, therefore, be avoided. 3.4 Esterification with alcohol mixture The esterification yield with longer alcohols was shown to be higher than for short alcohols, and the preference of R. miehei lipase for various alcohols was studied with a mixture of alcohols as substrates. When the experimental conditions optimized for ethyl ferulate or decyl ferulate were applied to a mixture of alcohols, esterification was observed to favor shorter alcohols such as propanol, ethanol, and butanol (Figure 5). The molar concentration of all primary alcohols was the same and when summed, equaled the optimal alcohol concentration. For the lower alcohol concentrations, which corresponded to the optimal conditions for the ethyl ferulate synthesis, the difference was even higher. Although higher yields were reached for the esterification with longer alcohols, the short alcohols were esterified preferably. One explanation for this phenomenon may be the slower diffusion rate of the longer alcohols through the immobilization material as previously reported (Ghamgui et al., 2004). If a mixture of alcohols was added to the lipase from R. miehei, the shorter alcohols were esterified to ferulic acid more quickly, but all primary alcohols provided were esterified. 70 Part B: Enzymatic esterification of ferulic acid Figure 5: Molar yields of ferulate esters (C-2 to C-18) at 61°C over time with an alcohol mixture (n=3, error bars referring to standard devation). The concentration of each of the alcohols was equal. Left: the ferulic acid and total alcohol concentration were 6.4 mM and 0.29 M, respectively, consistent with the optimal conditions for ethyl ferulate synthesis. Right: ferulic acid and total alcohol concentration were 14.6 mM and 0.39 M, respectively, consistent with optimal conditions for decyl ferulate. 4. Conclusion The synthesis in n-hexane using the immobilized lipase from R. miehei (Lipozyme RM IM) was optimized, which lead to maximal molar conversions of 76(2.0)% and 88(2.0)% after 72 h were reached for ethyl ferulate and decyl ferulate, respectively. The main differences in optimal reaction conditions were in the concentrations of the ferulic acid and the alcohol representing the substrates. Based on these optimizations, the esterification of ferulic acid with other alcohols, such as primary propanol, butanol, hexanol, octanol, tetradecanol and octadecanol and the branched alcohols isopropanol and 2-octanol, were tested. All primary alcohols were esterified to an expected extent. Specifically, increasing esterification from ethyl ferulate to hexyl ferulate was observed, and then it remained constant up to the 18 C long ester of ferulic acid. The branched alcohols did not esterify to ferulic acid as efficiently using R. miehei lipase. In a mixture of primary alcohols, the shorter ones from ethanol to butanol were esterified significantly quicker than the longer ones. The method developed in this study can be applied to enzymatically synthesize various alkyl ferulates, which opens new possibilities for further analysis of these compounds and future application as antioxidants in various systems. 71 Part B: Enzymatic esterification of ferulic acid References Bernards, M. A. (2002). Demystifying suberin. Canadian Journal of Botany, 80(3), 227-240. Cione, E., Tucci, P., Senatore, V., Perri, M., Trombino, S., Iemma, F., Picci, N., & Genchi, G. (2008). 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Biotechnology Letters, 32(10), 1457-1462. 73 Part B: Enzymatic synthesis of steryl ferulates Enzymatic synthesis of steryl ferulates Reprinted with permission from Aline Schär and Laura Nyström (2016). European Journal of Lipid Science and Technology. doi:. 10.1002/ejlt.201500586 Copyright (2016) Wiley. Abstract Steryl ferulates are plant sterols esterified to ferulic acid, a common phenolic acid. This esterification leads to sterol esters with improved biological properties, such as antioxidant activity. Commercially available and extracted steryl ferulates from rice bran are often limited in their sterol profiles. For further research and later food applications a simple enzymatic esterification could address the lack of availability of single steryl ferulates. Whereas several enzymatic procedures for the esterification of steryl fatty acid esters have been published, no fully enzymatic procedure for steryl ferulates has been reported so far. We optimized both direct esterification of β-sitosterol with ferulic acid as well as transesterification with ethyl ferulate yielding steryl ferulates. The reaction was catalyzed by a lipase from Candida rugosa, which lead to yields of 35% and 55% for the direct esterification and transesterification, respectively. Moreover, both reactions followed a similar time course over incubation. The enzyme activity was rather low, which is probably due to the specificity of the different isoenzymes of C. rugosa lipase. However, successful conditions for a fully enzymatic synthesis of steryl ferulates are reported for the first time. Practical applications: This enzymatic procedure leads to steryl ferulates, which do not need thorough purification, as no toxic catalysts were applied. This is especially an advantage when animal or human studies are conducted, which are needed for further evaluation of the potential health benefits of steryl ferulates. Further, it is less labor intensive than earlier published procedures using vinyl esters as substrates, which have to be synthesized and chromatographically purified. Keywords: Steryl ferulates / Candida rugosa lipase / Enzymatic esterification / Enzymatic transesterification / Phenolic acid lipophilization 75 Part B: Enzymatic synthesis of steryl ferulates 1. Introduction Steryl ferulates are esters of various plant sterols and ferulic acid, which are suggested to posses many health benefits, and which appear mainly in cereal brans(Mandak & Nyström, 2012). Steryl ferulates have been shown to lower total plasma cholesterol and LDL cholesterol in hypercholesterolemic hamsters (Wilson et al., 2007), and they are also known for their antioxidant activity (Nyström et al., 2007). The esterification of the antioxidative ferulic acid leads to an increased solubility in oil based systems and also allows high temperature applications (Nyström et al., 2007). Steryl ferulates extracted from rice are commonly known as γ-oryzanol (Mandak & Nyström, 2012). γ-oryzanol is predominantely composed of the two 4,4’-dimethyl sterol esters 24-methylenecycloartanyl ferulate and cycloartenyl ferulate, whereas the sterol pattern of steryl ferulates in wheat and corn is dominated by the desmethyl sterols, namely sitosterol, campesterol, and their saturated forms (Mandak & Nyström, 2012). Most commercially available steryl ferulates are extracts from rice and are therefore limited in their sterol pattern. Several studies have shown differences in antioxidant activity for different steryl ferulates (Nyström et al., 2005; Winkler- Moser et al., 2012; Xu et al., 2001). Further, in vitro hydrolysis studies indicate a difference in their potential metabolism between dimethyl and desmethyl steryl ferulates (Miller et al., 2004; Moreau & Hicks, 2004; Nyström et al., 2008). Therefore, procedures for the production of single steryl ferulates are required for research and later on maybe also for food and pharmaceutical applications. Chemical synthesis of steryl ferulates generally includes protection of the phenolic hydroxyl group, followed by esterification, and finally a deprotection step. The main disadvantage of the first method published included the synthesis of the highly reactive trans-4-O-acetylferuloyl chloride, which is rather difficult to handle (Condo et al., 2001; Kondo et al., 1988). This procedure was improved by Condo and co-workers (Condo et al., 2001), introducing a condensation of trans-4-O-acetylferulic acid with the sterol in the presence of N,N-dicyclohexylcarbodiimide and 4-(dimethylamino)-pyridine and a selective transesterification of the acetyl protecting group. However, the method still included long incubation times and did not result in satisfactory yields (39-61% for the coupling reaction). Recently, Winkler-Moser and colleagues (Winkler-Moser et al., 2015), proposed further improvements to the method, which included reduced reaction times, faster removal of the byproduct 1,3-dicyclohexylurea from the coupling reaction and finally higher yields of 77- 90%. Nevertheless, the improved chemical synthesis of steryl ferulates included three synthetic and two chromatographic steps, which overall lead to a high reagent and solvent consumption and a labor intensive procedure. 76 Part B: Enzymatic synthesis of steryl ferulates A combination of enzymatic and chemical synthesis of steryl ferulates has been applied earlier (Chigorimbo-Murefu et al., 2009; Tan & Shahidi, 2011; Wang et al., 2015). In all cases the intermediate product vinyl ferulate was chemically synthesized. This step required mercury acetate as catalyst and the synthesis was followed by purification using column chromatography. The resulting product vinyl ferulate was then transesterified enzymatically using the lipase of Candida rugosa. Vinyl esters are used in transesterification due to the fact that the liberated vinyl alcohol tautomerizes to acetaldehyde, thus, making the reaction irreversible. In the first study, the reached yield of the enzymatic transesterification was 56% (Chigorimbo-Murefu et al., 2009), in the second study 90% in 10 days (Tan & Shahidi, 2011) and in a recent study 45% in 24h (Wang et al., 2015). However, this method is still a multistep procedure and requires, apart from the enzyme, a heavy metal catalyst. Apart from the steryl phenolate synthesis discussed above, C. rugosa lipase has not been mentioned often so far as catalyst for phenolic acid lipophilization. In an early study, C. rugosa lipase was compared to other lipases on different cinnamic acid derivatives, showing 8% conversion to 1-octanol in 12 days for ferulic acid in solvent-free system (Stamatis et al., 2001). Later on, applying C. rugosa lipase in solvent-free condition under reduced pressure (80 kPa) for the esterification of 4-methoxycinnamic acid with oleyl alcohol lead to no or very low esterification activity (Vosmann et al., 2006). However, also a reasonable conversion of 26% has been reported for the transesterification of ethyl ferulate with tributyrin in toluene in 4 days (Zheng et al., 2009). Overall, the data available on the esterification activity of C. rugosa lipase on hydroxycinnamic acids is rather scarce and could be explored furthee, including the corresponding specificities of the isoenzymes. Plant sterols appear naturally in free form or covalently bound to a fatty acid, a sugar or a phenolic acid (Piironen et al., 2000). Their ability to lower plasma cholesterol and LDL cholesterol after human ingestion is the key nutritional interests of phytosterols (Piironen et al., 2000). The esterification with fatty acids and sterols has been investigated before. In two studies Weber and co-workers explored the esterification activity of lipases for sitosterol and oleic acid under reduced pressure in solvent free systems. The lipases from Rhizomucor miehei, C. rugosa and lipase B from C. antarctica were evaluated, showing that the activity of Candida rugosa lipase was highest (Weber et al., 2001a, 2001b). Later also in an organic solvent system it was found that C. rugosa lipase was most efficient and a yield of almost 85% steryl esters in 72 h was reported (Villeneuve et al., 2005). In another study a yield of 79.3% was reached in the esterification of plant sterols with lauric acid in 96h using Novozym 435 in n-hexane (He et al., 2010). Recently, Panpipat and co-workers demonstrated that C. antarctica lipase A shows superior catalytic activity to other lipases (C. rugosa lipase was not 77 Part B: Enzymatic synthesis of steryl ferulates evaluated in this study) such as C. antarctica lipase B. Yields of 93-98% were demonstrated for the esterification of β-sitosterol with fatty acids (C8-C18) within 24h in n-hexane (Panpipat et al., 2013). The enzymatic esterification of the plant sterols with fatty acids is therefore widely studied and well established. On the other hand the enzymatic esterification or transesterification of other acids than fatty acids with plant sterols has only been demonstrated rarely. In the study of Weber and co-workers the transesterification of ethyl dihydrocinnamate with cholesterol using Lipozyme (immobilized lipases from Rhizomucor miehei) showed a yield of 56% (Weber et al., 2001b). Also its structure is related to phenolic acids it does not belong to the group of phenolic acids, which possess at least one hydroxyl group on the aromatic ring (Figueroa-Espinoza & Villeneuve, 2005). The aim of this study was to elucidate the possibilities for enzymatic production of steryl ferulates. We focused on the use of C. rugosa lipase, which was successfully applied for the production of steryl ferulates via direct esterification of β-sitosterol with ferulic acid, as well as transesterification of ethyl ferulate. Both of the reactions were further optimized for reaction parameters using, surface response methodology. 2. Materials and Methods 2.1 Chemicals and enzymes Ferulic acid, ≥99% and β-sitosterol, ≥70% (impurities being mainly campesterol and β-sitostanol) were obtained from Sigma-Aldrich, Switzerland. γ-oryzanol, ≥ 99%, was purchased from Wako Pure Chemical Industries, Japan. Ethyl ferulate was purchased from Alfa Aesar, Germany (98% purity), and also synthesized as reported earlier (Schär & Nyström, 2015). All used solvents were of HPLC grade and all enzymes were purchased from Sigma-Aldrich, Switzerland. Five lipases were used in this study, namely lipase from Rhizomucor miehei (formerly known as mucor miehei) immobilized on macroporous ion-exchange resin, >30 U/g (1 U sets free 1 μmol stearic acid at pH 8.0 and 70°C per minute), lipase from C. rugosa type VII, ≥11.7 U/mg (at pH 7.2 and 37°C 1 U will hydrolyze 1.0 microequivalent of fatty acid from a triglyceride per minute), lipase A from C. antarctica immobilized on Immobead 150, recombinant from Aspergillus oryzae ≥500 U/g (1 U corresponds to the amount of enzyme which liberates 1 μmol butyric acid per minute at pH 10.0 and 40°C), lipase type B from C. antarctica, recombinant from A. niger, immobilized on acrylic resin ≥5,000 U/g (propyl laurate units), and lipase from C. rugosa, immobilized on 78 Part B: Enzymatic synthesis of steryl ferulates Immobead 150, ≥100 U/g (1 U corresponds to the amount of enzyme which liberates 1 μmol butyric acid from tributyrin per minute at pH 10.0 and 40°C). 2.2 Enzyme screening The four immobilized lipases (R. miehei lipase, C. antarctica lipase A, C. antarctica lipase B and C. rugosa lipase) were evaluated for their ability to transesterify ferulic acid from ethyl ferulate to sitosteryl ferulate. A solution of β-sitosterol in n-hexane was prepared (2.5 mg/mL), aliquots containing 12.5 mg of β-sitosterol were transferred into glass tubes and the hexane evaporated under a stream of nitrogen. Subsequently 10 mg of ethyl ferulate and 100 mg of immobilized enzyme were weighed into the tubes, followed by 3 mL of dehydrated (over 4 Å molecular sieve) hexane. The duplicate samples in addition to blanks without enzyme were shaken thoroughly before incubation without shaking at 50°C for 5 days. After incubation the solvent was evaporated and the whole sample was redissolved in 10 mL acetone. Aliquots of 350 µL were taken in duplicates, evaporated and redissolved in 1 mL of solvent B for HPLC analysis. 2.3 HPLC analysis, quantification and identification Steryl ferulates, ethyl ferulate and ferulic acid were analyzed using high performance liquid chromatography (HPLC, Agilent 1100, Switzerland), as described earlier ((Schär & Nyström, 2015)). Briefly, separation of analytes was achieved with reverse phase xBridgeTM Phenyl column from Waters (particle size of 3.5 µm) at room temperature, and detection was done at 325 nm with a diode array detector (DAD). Gradient elution with two solvents was used, where solvent A was 1% acetic acid in water, and solvent B a mixture of acetonitrile:water:1-butanol:acetic acid (88:6:4:2 v/v/v/v). The elution sequence was composed of a 3 min linear gradient from 75:25 (A:B) to 100% B, isocratic flow of 100% B for 7 min, 3 min linear gradient to 75:25 (A:B) and 2 min isocratic flow 75:25 (A:B) at a flow of 0.6 mL/min. For the quantification external calibration (0.05-6 nmol/injection) of ferulic acid, ethyl ferulate and γ-oryzanol were used. Identification was achieved by standard compounds, as well as the specific UV spectrum of ferulic acid. Molar yield was calculated based on the amount steryl ferulates quantified in the sample in comparison to the amount of sterols added (β-sitosterol and sterol impurities, campesterol and β-sitostanol). Additionally, as control, recoveries of the ferulic acid and ethyl ferulate were calculated for the samples analyzed with the full sample method (for more details see 2.4). Finally, mass spectrometry was used to confirm products identities of selected samples by showing presence of the expected mass. A SynaptTM G2 high resolution time-of-flight (TOF) mass spectrometer (Waters Corporation, Milford, MA, USA) was used applying direct and 79 Part B: Enzymatic synthesis of steryl ferulates electron spray ionization (ESI) in the negative ion mode. The voltages of capillary, sampling cone and extraction cone were 2.5 kV, 60 V and 4 V, respectively. The applied temperature was 120°C at the source and 550°C for desolvation, with a nitrogen flow of 20 and 800 L/h for the cone and desolvation, respectively. Figure 1: Esterification (R=H) and transesterification (R=Et) reaction of ferulic acid with sitosterol to sitosteryl ferulate. 2.4 Optimization of steryl ferulate synthesis For a typical reaction (Figure 1) a solution of β-sitosterol in n-hexane was prepared and the required amount transferred into the glass tubes. After evaporation of the hexane under a stream of nitrogen, ferulic acid or ethyl ferulate and C. rugosa lipase type VII were weighed into the glass tubes. After the addition of n-hexane, dehydrated over 4 Å molecular sieve, the tubes were shaken thoroughly using a vortex. The samples were incubated standing at the requested temperature without shaking. General reaction volume was 3 mL, except only for the optimization design for the transesterification reaction, 1.5 mL was used. In this study two different methods were used for sample analysis. For the aliquot sampling method, the tubes were cooled to room temperature and shaken thoroughly. Aliquot samples of 50 µL were taken and evaporated under a stream of nitrogen at 50°C. The residue was redissolved in 500 µL solvent B and filtrated before HPLC analysis. For the second method, the full sample method, the cooled samples were evaporated to dryness under a stream of nitrogen. To re-dissolve product and educts, 10 mL of acetone were added and the tubes shaken thoroughly. For this method the sample analysis was performed in duplicates and average values were calculated. For that purpose aliquots of 350 µL were transferred into another glass tube. The acetone was evaporated under a nitrogen stream at 50°C. Finally, the residue was redissolved in 1 mL solvent B and filtrated. For the control of optimal conditions Data is presented as mean with standard deviation in parentheses. 2.5 Experimental design and surface response methodology The Unscramble X from CAMO Software, Oslo, Norway was used to design the experiments and to evaluate the data. A 3-level-4-factor Box-Behnken design was used in this study. The experimental data was fitted to the second-order polynomial equation 1: 80 Part B: Enzymatic synthesis of steryl ferulates 4 4 2 3 4 푌 = 훽푘0 + ∑푖=1 훽푘푖푥푖 + ∑푖=1 훽푘푖푖푥푖 + ∑푖=1 ∑푗=푖+1 훽푘푖푗푥푖푥푗 (1) Y refers to the response (molar yield %), 훽푘0 , 훽푘푖 , 훽푘푖푖, and 훽푘푖푗 are constant coefficient and 푥푖represents the coded independent variables. The used variables were: temperature (x1), enzyme-to-sterol ratio (x2), sterol amount (x3), molar substrate ratio (x4). A center sample (all coded variables equal zero) was included and analyzed in triplicates. The parameters and the ranges thereof were chosen based on preliminary experiments (data not shown). The ranges of the variables are listed in Table 1 for the optimization of both, the direct esterification and the transesterification. The conducted experiments for the optimization are listed in Table 2 and the corresponding experimental data after 120 h of incubation. Table 1: Range of variables for the conducted optimizations of the direct esterification of ferulic acid (FA) with β-sitosterol and the transesterificaion of ethyl ferulate (EF) with β-sitosterol in hexane using C. rugosa lipase. Variable Direct esterification Transesterification x1: temperature 50-65°C 45-65°C x2: enzyme-to-sterol ratio 1-3 g/g 1-3 g/g x3: sterol amount 10-30 mg/3 mL 5-15 mg/3 mL x4: substrate ratio 1-5 mol FA / mol β-sitosterol 1-3 mol EF / mol β-sitosterol 2.6 Two-step synthesis of steryl ferulates To confirm the fully enzymatic synthesis of steryl ferulates involving the formation of ethyl ferulate followed by transesterification with sterol, a two-step reaction was carried out. Enzymatic synthesis of ethyl ferulate was carried out as described earlier (Schär & Nyström, 2015). Ferulic acid and ethanol were incubated for 72 h the in hexane with the immobilized lipase from R. miehei. After incubation samples were cooled to room temperature and filtered to remove the enzyme. This ferulic acid esterification was performed in triplicates. The concentration of ethyl ferulate in the filtrate was determined, after which hexane and ethanol were then evaporated under nitrogen at 50°C. To ensure total ethanol evaporation, dry hexane was added and evaporated again. The crude product containing the produced ethyl ferulate was subjected to transesterification as described above. 81 Part B: Enzymatic synthesis of steryl ferulates 3. Results and Discussion 3.1 Optimization of transesterification In a first step different immobilized lipases were tested on their activity to transesterify ferulic acid from ethyl ferulate to β-sitosteryl ferulate. For the duplicate sample with the immobilized C. rugosa lipase an average molar yield of 9.2% was measured. For the lipase A from C. antarctica and the lipase from R. miehei a very small amount of steryl ferulates could be detected, however, smaller than the quantification limit. In the samples with C. antarctica lipase B no clear difference to the blank could be measured after 5 days of incubation. This findings correspond well with other studies, were C. rugosa lipase has been the only lipase able to catalyze the synthesis of steryl ferulates starting from vinyl ferulate (Chigorimbo- Murefu et al., 2009; Tan & Shahidi, 2011). Recently the C. antarctica lipase A was shown to esterify sterols with fatty acids most efficiently (Panpipat et al., 2013). However, based on this data it seems that more complex acid substrates such as hydroxycinnamic acids are not amongst good substrates of C. antarctica lipase A. Conclusively, a lipase from C. rugosa was selected for later use. However, since the yield of steryl ferulates with the immobilized lipase was rather low and the enzyme amount very high, a non-immobilized enzyme preparation with a higher activity was chosen with the lipase type VII from C. rugosa. The transesterification of ferulic aid from ethyl ferulate to sitosteryl ferulate using C. rugosa lipase was optimized regarding four parameters: temperature, enzyme-to-sterol ratio, sterol amount, and substrate ratio (Table 2). The three center points gave a yield of 48.7(2.9)% and the experimental data was fitted to the second-order polynomial equation (equation 1). The analysis of variance (Table 3) shows a strong correlation between the model and the experimental data, as indicated by the low p-values for all model variables and a very high R2 and lack of fit. 82 Part B: Enzymatic synthesis of steryl ferulates Table 2: Conducted coded experiment following the Box-Behnken design with four variables and the experimental data, center samples (x1-4=0) are not included. Uncoded variables and further conditions are listed in Table 1 and experimental data. 1 2 3 4 5 6 7 8 9 10 11 12 x1: temperature 0 0 0 0 -1 1 -1 1 -1 1 -1 1 x2: enzyme-to-sterol ratio 0 0 0 0 -1 -1 1 1 0 0 0 0 x3: sterol amount -1 1 -1 1 0 0 0 0 0 0 0 0 x4: substrate ratio -1 -1 1 1 0 0 0 0 -1 -1 1 1 Yield direct esterification [%] 13.6 23.9 16.9 20.6 7.0 9.5 22.6 25.3 13.5 26.0 15.4 20.5 Yield transesterification [%] 34.5 39.1 40.8 40.8 23.4 32.3 43.4 53.4 32.6 43.3 38.6 46.0 13 14 15 16 17 18 19 20 21 22 23 24 x1: temperature 0 0 0 0 0 0 0 0 -1 1 -1 1 x2: enzyme-to-sterol ratio -1 1 -1 1 -1 1 -1 1 0 0 0 0 x3: sterol amount -1 -1 1 1 0 0 0 0 -1 -1 1 1 x4: substrate ratio 0 0 0 0 -1 -1 1 1 0 0 0 0 Yield direct esterification [%] 7.6 28.8 12.7 22.0 12.0 29.5 13.3 24.9 12.5 13.3 16.6 24.9 Yield transesterification [%] 25.0 51.0 29.6 51.3 31.1 42.9 24.1 53.7 37.6 44.0 33.6 52.0 Table 3: Analysis of variance of models calculated for direct esterification system and transesterification system. ss: sum of squares, df: degree of freedom. Direct esterification Transesterification ss df p-value ss df p-value Model 1037.3 14 0.0001 1627.1 14 0.0000 Linear 844.2 4 0.0000 1449.0 4 0.0000 Interaction 82.8 6 0.1105 120.2 6 0.0095 Quadratic 173.3 4 0.0037 96.8 4 0.0073 Lack of fit 73.3 10 0.0404 34.4 10 0.8332 R2 0.9335 0.9707 83 Part B: Enzymatic synthesis of steryl ferulates The β-coefficients and the corresponding p-values can be found in Table 4. All linear factors except the sterol amount have a significant influence on the yield, with the temperature and the enzyme-to-sterol ratio having the highest β-coefficients and therefore, a strong influence on the yield. Similarly for the quadratic factors all factors are significant and have a medium influence on the yield. But only two interactions, namely temperature x sterol amount and enzyme-to-sterol ratio x substrate ratio, were found to have a significant influence on the yield. Table 4: β-Coefficients and corresponding p-values of the fitted models for the direct esterification and transesterification reaction yielding steryl ferulates catalyzed by C. rugosa lipase in hexane. Variables referring to: x1: temperature, x2: enzyme-to-sterol ratio, x3: sterol amount and x4: substrate ratio. Direct esterification Transesterification β-coefficient p-value β-coefficient p-value β0 23.33 48.73 x1 2.66 0.00* 5.15 0.00* x2 7.58 0.00* 10.85 0.00* x3 2.33 0.01* 1.13 0.06 x4 -0.58 0.44 1.71 0.01* x1* x2 0.05 0.97 0.28 0.78 x1* x3 1.88 0.16 3.00 0.01* x2* x3 -2.98 0.03* -1.08 0.28 x1* x4 -1.85 0.16 -0.82 0.40 x2* x4 -1.48 0.26 4.45 0.00* x3* x4 -1.65 0.21 -1.15 0.25 2 x1 -3.82 0.00* -3.68 0.00* 2 x2 -2.80 0.02* -6.05 0.00* 2 x3 -3.03 0.02* -3.79 0.00* 2 x4 -0.94 0.40 -5.27 0.00* *significant at p < 0.05 In the contour plots (Figure 1 in supporting information) the calculated yield for the corresponding conditions after 120 h of incubation are illustrated. A clear trend towards high enzyme-to-sterol ratio and a rather high temperature can be found. For the sterol amount and the substrate ratio a trend towards the middle can be found. Based on the determined β- coefficients optimal conditions were calculated (Table 5). For the transesterification reaction they are at 63°C, with an enzyme-to-sterol ratio of 3 g/g, a sterol amount of 11.2 mg/3mL, and a substrate ratio of 2.5 mol/mol. The predicted yield for these conditions is 57.2%, which 84 Part B: Enzymatic synthesis of steryl ferulates was confirmed experimentally leading to a yield of 54.9(2.5)% (Table 5). Conclusively, the optimization was successful and the built model is displaying the experimental data well. Other studies have shown an increased activity of C. rugosa lipase when some percentages of water (w/w% of substrate) were added to the organic solvent ((Shieh et al., 1996)), and so an addition of water was also tested in this study. With an addition of 10% and 20% (w/w of enzyme) only a decrease in transesterification activity could be detected, when performed in a previous optimization design (data not shown), and thus water was not added to later experiments. Furthermore, a possible addition of 4 Å molecular sieve was evaluated, but excluded as a factor in the optimization design, as a small amount (1-2 pellets, approximately 5-20 mg) was found to have no significant influence in the screening design. After optimization it was tested again for both reactions by an addition of 50 mg /3 mL 4 Å molecular sieve, which lead to reactions with almost no yield. Therefore, neither an addition of water, nor an addition of molecular sieve was included as factor in the optimization designs. Finally also the addition of the co-solvent butanone was evaluated, as a previous study used 10% butanone in n-hexane (Tan & Shahidi, 2011). But already a butanone addition of 5% (v/v) lead to a decrease of the molar yield of around 50% for both, the transesterification and direct esterification, reactions. This indicates that the inhibition of the enzyme through the butanone is stronger compared to the possibly improved reactivity due to the increased solubility of the ferulic acid or ethyl ferulate. Table 5: Optimal conditions for direct esterification and transesterification (from ethyl ferulate) reactions with C. rugosa lipase in hexane to produce steryl ferulates, their predicted yields, and confirmed results; standard deviations in parentheses. Direct esterification Transesterification x1: temperature [°C] 63 63 x2: enzyme-to-sterol ratio [g/g] 3 3 x3: sterol amount [mg/3 mL] 23.8 11.2 x4: substrate ratio [mol/mol] 1 2.5 Predicted Yield [%] 31.0 57.2 Measured Yield [%] 34.8 (1.5); n=10 54.9 (2.5); n=9 n= number of conducted replicates, yield reflects the molar percentage of sterols (β-sitosterol and sterol impurities campesterol and β-sitostanol) converted to steryl ferulates. 85 Part B: Enzymatic synthesis of steryl ferulates 3.2 Two-step synthesis of steryl ferulates To confirm the fully enzymatic, two-step synthesis of steryl ferulates, a reaction was carried out, where ferulic acid, ethanol and sterol were used as substrates. The optimal conditions for the synthesis of ethyl ferulate as reported earlier (61°C, 72h, 3.75 mg/3mL ferulic acid, 50 µL/3mL ethanol, and an enzyme-to-sterol ratio of 2.5 g/g) were applied with an expected yield of 76.2% (Schär & Nyström, 2015). Therefore, to synthesize 15 mg of ethyl ferulate, approximately 18.5 mg ferulic acid is needed. After the incubation the synthesized ethyl ferulate was quantified and used for transesterification as described above. The conversion of ferulic acid to ethyl ferulate observed was 82.4(2.7)% after incubation. After evaporation the requested amount of β-sitosterol, enzyme, and hexane were added to reach condition similar as the optimal conditions mentioned above. For the transesterification the samples were incubated for 120 h at 63°C and finally the concentration of steryl ferulates was determined with the aliquot sampling method. The measured yield was 56.9(3.4)%, which corresponds well with the predicted yield and the yields reached with commercial ethyl ferulate. However, this value is slightly higher than the others measured with commercial ethyl ferulate. This is probably due to the different sampling method, which was here the aliquot sampling method, thus leading to a slight overestimation (see discussion below). Conclusively, the fully enzymatic, two-step synthesis of steryl ferulates was successfully investigated. 3.3 Optimization of direct esterification The direct esterification of β-sitosterol with ferulic acid using C. rugosa lipase was optimized for four parameters: temperature, enzyme-to-sterol ratio, sterol amount, and substrate ratio (Table 2). The yields of steryl ferulates in the replicates in the center of the design were 23.3(0.6)%. The model (equation 1) was fitted to the experimental data, and the analysis of variance (Table 3) indicates that the model is significant and represents the relationship between the variables and the yield adequately. However, the lack of fit is just below the level of significance, which indicates that the variance in the data cannot be fully explained by the model. This may also be caused by the very small variation among the replicates of the center point compared to the possibly higher variation of the other data points. Looking at the β-coefficients and the corresponding p-values (Table 4), all linear and quadratic factors have a significant influence, except the linear and quadratic factor of ferulic acid to sterol ratio. This seems logical, as the solubility of the ferulic acid in the hexane system is very low and thus a higher amount of ferulic acid in the overall system does not lead to a higher concentration available for the enzyme. Of the interaction factors only the enzyme-to-sterol ratio x sterol amount has a significant influence on the yield. In the contour 86 Part B: Enzymatic synthesis of steryl ferulates plots (Figure 2 in supporting information) the full picture of the built model over the design space can be seen. Clearly there is a trend for higher yields towards a high enzyme-to-sterol ratio. As already indicated by the insignificant β-coefficient of the substrate ratio, only a small increase in the yield towards a small substrate ratio could be found. The calculated optimal conditions for the direct esterification system were at 63°C, with an enzyme-to-sterol ratio of 3 g/g, a sterol amount of 23.8 mg/3 mL, and a substrate ratio of 1 mol/mol for which a calculated yield of 31% can be expected. This yield was confirmed several times with different batches of enzyme and was found to be 34.8(1.5)% after 120h. This yield is generally a bit higher than calculated by the model, but still fitting the expected range. Therefore, the enzymatic esterification of ferulic acid with β-sitosterol was successfully optimized. 3.4 Comparison of direct esterification and transesterification The time courses of both reactions follow a similar trend (Figure 2). All time points were analyzed in triplicates using the full sample method, requiring preparation of three individual samples for each time point. The main difference between the two reactions is the reached yield, but for both reactions 5 days seems to be a time where the maximum is reached. It is therefore not the case that the direct esterification is just slower, but actually really seems to lead to a lower yield. The esterification of phenolic acids has been reviewed by Figueroa-Espinoza and Villeneuve in 2005 (Figueroa-Espinoza & Villeneuve, 2005). They highlight the challenging factors of enzymatic phenolic acid esterification with lipases, including the fact that an unsaturation in the side chain conjugated with a hydroxyl group in para-position can lead to lipase inhibition. Therefore, the direct esterification of free phenolic acids is rather challenging and slow, which can be addressed by performing transesterification of methyl, ethyl or vinyl phenolates. As in the study of Compton and colleagues where the yield could be increased from 14% to 50% for the synthesis of octyl ferulate from free ferulic acid and ethyl ferulate, respectively (Compton et al., 2000). Also in another study Weitkamp and co-workers transesterified phenolic acids with fatty alcohols in a solvent free system. They found that the transesterification was up to 56 times faster than direct esterification in the case of ferulic acid (Weitkamp et al., 2006). The results of this study are rather in the range of the study of Compton and co-workers. The yield increased from around 35% to 55% by going from direct esterification to transesterification of ferulic acid. Apart from the comparison between direct esterification and transesterification also the transesterification of ethyl ferulate and vinyl ferulate has been compared before (Yu et al., 87 Part B: Enzymatic synthesis of steryl ferulates 2010). That study showed that the vinyl ferulic acid ester was more efficiently transesterified (91%) with triolein, unlike the ethyl ferulate, where the transesterification yield was only 70%. In previous studies the transesterification of vinyl ferulate with sterols using C. rugosa lipase (Chigorimbo-Murefu et al., 2009; Tan & Shahidi, 2011; Wang et al., 2015) lead to a yield between 45% and 90%. In the study of Tan and Shahidi the samples were incubated for 10 days (Tan & Shahidi, 2011). The yield of around 55% is therefore well in the range, which could be expected based on the comparison of the transesterification ability of ethyl ferulate compared to vinyl ferulate (Yu et al., 2010). Figure 2: Time course of transesterification (●) and direct esterification (♦) reaction yielding steryl ferulates at optimal conditions (see Table 5) catalyzed by C. rugosa lipase in hexane, means of n=3, error bars representing standard deviation, sample analysis was conducted with the full sample method (see section 2.4). For both systems the enzyme amount applied was enormous. As the applied enzyme preparation is not immobilized, an enzyme-to-sterol ratio of 3 g/g is really high. Although this is of course a cost factor, the applied lipase preparation is rather cheap and impure. We estimated the protein content of the enzyme preparation using Bradford assay with bovine serum albumin as standard, and found it to be only about 2%. This lies in the range of protein contents for C. rugosa lipases from the same supplier determined earlier (0.8-6%) (Domínguez de María et al., 2006; Lopez et al., 2004). It is a known problem that these C. rugosa lipase preparations are usually low in their purity and protein content (Dominguez de Maria et al., 2006). The measured lipase activity was 0.06 and 0.04 U/g (1 U equals 1 μmol of steryl ferulate formed per minute at 63°C). This is indeed a low activity but not too 88 Part B: Enzymatic synthesis of steryl ferulates far from the activities determined earlier for the esterification of sterols with saturated fatty acids (0.1-32.3 U/g) (Weber et al., 2001a). One explanation for this low activity could be found in the fact that C. rugosa lipase contains several isoenzymes and type 3 is known to exhibit cholesterol esterase activity (Lopez et al., 2004; Tenkanen et al., 2002). Cholesterol esterases have been purified from various microbial sources, C. rugosa being one of them (Maeda et al., 2008). This type 3 lipase was found to make up to 11% of the commercially available C. rugosa lipase type VII from Sigma (Lopez et al., 2004). In addition to that, the lipase 3 from C. rugosa was found to be still active after immobilization in isooctane system (Lopez et al., 2004). This leads to the possible conclusion that only the isoenzyme type 3 lipase is responsible for the esterification of ferulic acid and sterols. In this study two different sampling methods were applied, the aliquot sampling method and the full sample method. Both methods have their advantages and disadvantages. The aliquot sampling method has the advantage, that the reaction progress of the same samples can be observed over time. But the risk of errors is rather high. First, especially at long incubation times and incubation temperatures close to the boiling point of the solvent, there is a risk of evaporating solvent and therefore an overestimation of the yield. Additionally, the sampling volume has to be rather small to not change the reaction system, which makes the pipetting error relatively high. The full sample method on the other hand has the disadvantage, that only one time point per sample can be analyzed an therefore, especially when it comes to time courses, is more labor intensive. But the risk of overestimation is minimized and the recovery of the substrates can also be calculated as control or even to calculate the yield. Recoveries from 92-109% were found for this study. Here both methods were applied and overestimations of the aliquot sampling method of 0-12% were observed, and the overestimation increased with time. Overall, both sampling methods can be suitable, if one is aware of the limitations. The purification after incubation also differs for the direct esterification and transesterification systems. In the case of the direct esterification system the remaining ferulic acid can be removed from the hexane system simply by washing the hexane phase with water and an additional drying step. The free sterols can be separated from the steryl ferulates with a base-acid wash (Evershed et al., 1988; Hakala et al., 2002). In the case of the transesterification system the separation of the remaining ethyl ferulate and the steryl ferulates is more challenging and requires a chromatographic step (i.e. reverse phase C18 solid phase extraction). Although the yield of the transesterification is higher, for laboratory purpose the direct esterification may be the choice as the purification is less labor intensive. Conclusively, the transesterification of ferulic acid to steryl ferulates leads to a higher yield 89 Part B: Enzymatic synthesis of steryl ferulates over the direct esterification, but the choice which system is most suitable relies also on other factors such as necessity of purification, whether the phenolic acid ester is commercially available, and the price of the sterol substrate (more needed for the direct esterification). 4. Conclusions In this study we presented the first fully enzymatic synthesis of steryl ferulates. The direct esterification of ferulic acid and the transesterification from ethyl ferulate to steryl ferulates was optimized leading to yields of 35% and 55%, respectively. In combination with the enzymatic esterification of ferulic acid with ethanol using an immobilized lipase from R. miehei, this leads to a fully enzymatic two-step synthesis of steryl ferulates. Although the yield for the transesterification system is higher, both systems should be considered for future applications and the selection can be made based on several arguments discussed above. The main differences found for the optimal reaction conditions are the sterol amount, which can be set higher for the direct esterification system, and the substrate ratio, which is of less importance for the direct esterification system. The process developed in this study allows for a simple enzymatic synthesis of steryl ferulates on a laboratory scale and also provides basics for further improvement to later on implement larger scale applications. 5. Acknowledgements We gratefully acknowledge the financial support of the Swiss National Science Foundation, SNSF (Project 200021_141268) and ETH Zurich. The authors declare no conflict of interest. 90 Part B: Enzymatic synthesis of steryl ferulates References Chigorimbo-Murefu, N. T. L., Riva, S., & Burton, S. G. (2009). Lipase-catalysed synthesis of esters of ferulic acid with natural compounds and evaluation of their antioxidant properties. Journal of Molecular Catalysis B: Enzymatic, 56(4), 277-282. Compton, D. L., Laszlo, J. A., & Berhow, M. A. (2000). Lipase-catalyzed synthesis of ferulate esters. Journal of the American Oil Chemists Society, 77(5), 513-519. Condo, A. M., Baker, D. C., Moreau, R. A., & Hicks, K. B. (2001). 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Maeda, A., Mizuno, T., Bunya, M., Sugihara, S., Nakayama, D., Tsunasawa, S., Hirota, Y., & Sugihara, A. (2008). Characterization of novel cholesterol esterase from Trichoderma sp. AS59 with high ability to synthesize steryl esters. Journal of Bioscience and Bioengineering, 105(4), 341-349. Mandak, E., & Nyström, L. (2012). The Effect of In Vitro Digestion on Steryl Ferulates from Rice (Oryza sativa L.) and Other Grains. Journal of Agricultural and Food Chemistry, 60(24), 6123- 6130. Miller, A., Majauskaite, L., & Engel, K. H. (2004). Enzyme-catalyzed hydrolysis of gamma-oryzanol. European Food Research and Technology, 218(4), 349-354. Moreau, R. A., & Hicks, K. B. (2004). The in vitro hydrolysis of phytosterol conjugates in food matrices by mammalian digestive enzymes. Lipids, 39(8), 769-776. 91 Part B: Enzymatic synthesis of steryl ferulates Nyström, L., Achrenius, T., Lampi, A. M., Moreau, R. A., & Piironen, V. (2007). 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Villeneuve, P., Turon, F., Caro, Y., Escoffier, R., Barea, B., Barouh, B., Lago, R., Piombo, G., & Pina, M. (2005). Lipase-catalyzed synthesis of canola phytosterols oleate esters as cholesterol lowering agents. Enzyme and Microbial Technology, 37(1), 150-155. Vosmann, K., Weitkamp, P., & Weber, N. (2006). Solvent-free lipase-catalyzed preparation of long- chain alkyl phenylpropanoates and phenylpropyl alkanoates. Journal of Agricultural and Food Chemistry, 54(8), 2969-2976. Wang, Z., Hwang, S. H., & Lim, S. S. (2015). Lipophilization of phenolic acids with phytosterols by a chemoenzymatic method to improve their antioxidant activities. European Journal of Lipid Science and Technology, 117(7), 1037-1048. Weber, N., Weitkamp, P., & Mukherjee, K. D. (2001a). Fatty acid steryl, stanyl, and steroid-esters by esterification and transesterification in vacuo using Candida rugosa lipase as catalyst. Journal of Agricultural and Food Chemistry, 49(1), 67-71. Weber, N., Weitkamp, P., & Mukherjee, K. D. (2001b). Steryl and stanyl esters of fatty acids by solvent-free esterification and transesterification in vacuo using lipases from Rhizomucor miehei, Candida antarctica, and Carica papaya. Journal of Agricultural and Food Chemistry, 49(11), 5210-5216. 92 Part B: Enzymatic synthesis of steryl ferulates Weitkamp, P., Vosmann, K., & Weber, N. (2006). Highly efficient preparation of lipophilic hydroxycinnamates by solvent-free lipase-catalyzed transesterification. Journal of Agricultural and Food Chemistry, 54(19), 7062-7068. Wilson, T. A., Nicolosi, R. J., Woolfrey, B., & Kritchevsky, D. (2007). Rice bran oil and oryzanol reduce plasma lipid and lipoprotein cholesterol concentrations and aortic cholesterol ester accumulation to a greater extent than ferulic acid in hypercholesterolemic hamsters. Journal of Nutritional Biochemistry, 18(2), 105-112. Winkler-Moser, J. K., Hwang, H. S., Bakota, E. L., & Palmquist, D. A. (2015). Synthesis of steryl ferulates with various sterol structures and comparison of their antioxidant activity. Food Chemistry, 169, 92-101. Winkler-Moser, J. K., Rennick, K. A., Palmquist, D. A., Berhow, M. A., & Vaughn, S. F. (2012). Comparison of the Impact of gamma-Oryzanol and Corn Steryl Ferulates on the Polymerization of Soybean Oil During Frying. Journal of the American Oil Chemists Society, 89(2), 243-252. Xu, Z. M., Hua, N., & Godber, J. S. (2001). Antioxidant activity of tocopherols, tocotrienols, and gamma-oryzanol components from rice bran against cholesterol oxidation accelerated by 2,2 '-azobis(2-methylpropionamidine) dihydrochloride. Journal of Agricultural and Food Chemistry, 49(4), 2077-2081. Yu, Y., Zheng, Y., Quan, J., Wu, C. Y., Wang, Y. J., Branford-White, C., & Zhu, L. M. (2010). Enzymatic Synthesis of Feruloylated Lipids: Comparison of the Efficiency of Vinyl Ferulate and Ethyl Ferulate as Substrates. Journal of the American Oil Chemists Society, 87(12), 1443- 1449. Zheng, Y., Wu, X. M., Branford-White, C., Ning, X., Quan, J., & Zhu, L. M. (2009). Enzymatic synthesis and characterization of novel feruloylated lipids in selected organic media. Journal of Molecular Catalysis B: Enzymatic, 58(1-4), 65-71. 93 Part B: Enzymatic synthesis of steryl ferulates Supporting information Figure 1: Contour plots of molar yield of transesterification reaction after 120h, generated by fitting the experimental data (Table 2) to equation 1. The gray scale indicates the predicted molar yield of steryl ferulates from ethyl ferulate and β-sitosterol catalyzed by C. rugosa lipase at given conditions. Substrate ratio refers to mol ethyl ferulate / mol β-sitosterol. □: < 10%, ■: 10-20%; ■: 20-30%; ■: 30-40%; ■: 40-50%, ■: 50-60% 94 Part B: Enzymatic synthesis of steryl ferulates Figure 2: Contour plots of molar yield of direct esterification reaction after 120h, generated by fitting the experimental data (Table 2) to equation 1. The gray scale indicates the predicted molar yield of steryl ferulates from ferulic acid and β-sitosterol catalyzed by C. rugosa lipase at given conditions. Substrate ratio refers to mol ferulic acid / mol β-sitosterol. □: < 10%, ■: 10-20%; ■: 20-30%; ■: 30-40%; ■: 40-50%, ■: 50-60% 95 Part B: Enzymatic synthesis of steryl ferulates A B Figure 3: ESI-MS/MS spectra of sitosteryl ferulate synthesized through transesterification from ethyl ferulate (A) and through direct esterification from ferulic acid (B). The most abundant species refers to [M-H]- and [M-H-Me]-. Further ions are related to the ferulic acid part. This is in accordance to previously published data (Zhu & Nyström, 2015). MS- conditions can be found in section 2.3. Reference: [1] Zhu, D., & Nyström, L. (2015). Differentiation of rice varieties using small bioactive lipids as markers. European Journal of Lipid Science and Technology, 117(10), 1578-1588. 96 Part B: Esterification of hydroxycinnamic acids and their antioxidant activity Enzymatic synthesis of steryl hydroxycinnamates and their antioxidant activity Aline Schär, Silvia Liphardt and Laura Nyström Submitted manuscript (June 2016). Abstract Steryl hydroxycinnamates are of increasing interest as they are antioxidant esters of phytosterols with potential cholesterol lowering properties. Apart from ferulates, also other plant steryl hydroxycinnamates have been identified in natural products. In this study hydroxycinnamic acid derivatives were ethylated enzymatically using R. miehei lipase, and transesterified by lipase from C. rugosa to yield steryl hydroxycinnamates. The influence of the structural differences between the hydroxycinnamic acid derivatives on the esterification yields was very different for the two lipases applied. Furthermore, the antioxidant activity of steryl and stearyl hydroxycinnamates was evaluated by DPPH radical scavenging activity and in two methyl linoleate systems. In bulk methyl linoleate free sinapic acid showed the highest antioxidant activity over other sinapates, whereas in emulsified methyl linoleate, stearyl sinapate was highest. In conclusion, the enzymatic synthesis of steryl hydroxycinnamates is highly structure dependent and their antioxidant activity is not necessarily improved through esterification with sterols. Keywords: Steryl ferulates / C. rugosa lipase / R. miehei lipase / steryl phenolates / phenolic acid lipophilisation / lipophilic antioxidants 97 Part B: Esterification of hydroxycinnamic acids and their antioxidant activity 1. Introduction Phenolic acids are effective antioxidants due to their phenolic hydroxyl group and therefore possess the ability to form stable phenoxy radicals after donation of hydrogen (Decker, 1998). Phenolic acids (as reviewed by Manach and colleagues in 2004) can be separated into two groups, the benzoic acid derivatives and cinnamic acid derivatives. Hydroxycinnamic acids, mainly p-coumaric, caffeic, ferulic, and sinapic acid are more common than hydroxybenzoic acids and are mostly found in bound form, like ferulic acid esterified to cell wall polysaccharides such as arabinoxylan. The most abundant phenolic acids in fruits is caffeic acid and in cereal grains ferulic acid (Manach et al., 2004). Hydroxycinnamic acids occur naturally also as esters of fatty alcohols, and plant sterols. For instance long chain alkyl ferulates (C16 to C30) occur in suberin, a cell wall component of plants (Bernards, 2002), and hexadecyl, octadecyl, and eicosyl p-coumarates in vine and root latex of sweet potato (Snook et al., 1994), and many others sources in the plant kingdom, as recently reviewed (He et al., 2015). Sterol esters of hydroxycinnamic acids, on the other hand, are most abundantly found in cereals such as rice, wheat, rye, and corn, where they commonly occur as ferulic acid esters (Mandak & Nyström, 2012; Norton, 1995). In addition to steryl ferulates also other hydroxycinnamic acid sterol esters have been identified, such as caffeic sterol esters in canary seeds (Takagi & Iida, 1980), and p-coumaric acid sterol esters in corn (Norton, 1995; Seitz, 1989). Plant sterols in general have gained significant interest due to their ability to lower plasma cholesterol and LDL cholesterol (Piironen et al., 2000), and this effect has also been demonstrated for ferulic acid esters of sterols in hamsters (Wilson et al., 2007). To summarize, phytosteryl hydroxycinnamates are natural and lipophilic antioxidants with potential health benefits. Antioxidants have been studied for many years, including phenolic acids and their derivatives, in a range of oxidation systems to evaluate the link between polarity and antioxidant activity. An early theory raised in this context is the polar paradox, which states that in nonpolar media, such as bulk oil, the highest antioxidant activity for homologous series of antioxidants with varying polarities can be observed for the polar compounds (Porter et al., 1989). Similarly, in more polar systems such as oil-in-water emulsions, nonpolar antioxidants show higher antioxidant activity. This theory was later on explained by the presence of colloids in bulk oils, at which oxidation is likely to occur and where polar antioxidants are preferentially located (Chaiyasit et al., 2007). However, not all studies on structure-activity relationships of similar antioxidants found results following this polar paradox, thus further research is still needed. Recent advances in the field have 98 Part B: Esterification of hydroxycinnamic acids and their antioxidant activity demonstrated the so-called cutoff effect, which was first illustrated for chlorogenic acid (Laguerre et al., 2009). In an emulsified system there is a critical chain length of the esterified alcohol at which the antioxidant activity is highest. This chain length determines where the antioxidant is located in the system. However, not only the chain length but also the type of emulsifier may change its distribution in the system and correspondingly its antioxidant activity (Stockmann et al., 2000). Therefore, the antioxidant activity of phenolic acids strongly depends on their lipophilicity and the system of application. To alter the lipophilicity of phenolic acids, to improve the antioxidant activity, the acid group may be esterified with a nonpolar alcohol. The enzymatic esterification of phenolic acids has been reviewed a few years ago (Figueroa-Espinoza & Villeneuve, 2005). An enzymatic procedure brings several advantages over a chemical esterification such as less intermediate steps and side products that overall lead to a reduced solvent usage and waste production. The comparison of several phenolic acids esterified by several enzymes was conducted by Stamatis and co-workers (Stamatis et al., 1999). In solvent-free system the esterification yield of R. miehei lipase of 1-octanol with hydroxycinnamic acids decreased in the following order: cinnamic acid > m-coumaric acid > ferulic acid > p-coumaric acid > o-coumaric acid > caffeic acid, whereas the order was changed for C. antarctica lipase. Apart from possible steric effects, this reactivity was attributed to the presence and position of the hydroxyl group and the unsaturation of the side chain. A conjugated phenolic hydroxyl group with the carboxylic group (as it is the case for a para-hydroxyl group in combination with an unsaturated side chain) leads to a deactivation of the electrophilic carbon center for a nucleophilic attack of the alcohol (Buisman et al., 1998). However, apart from linear alcohols, also the esterification of ferulic acid with sterols is of interest. Three approaches have been studied so far: a chemical synthesis, which was optimized only recently (Winkler-Moser et al., 2015), a chemoenzymatic approach including the transesterification of vinyl phenolic acid esters (Chigorimbo-Murefu et al., 2009; Tan & Shahidi, 2011, 2012, 2013; Wang et al., 2015), or a fully enzymatic approach (Schär & Nyström, 2016). In this latest study, two enzymatic methods were compared, the direct esterification of ferulic acid and the transesterification of ferulic acid from ethyl ferulate to steryl ferulates. Therefore, further information about the potential of the fully enzymatic synthesis of steryl hydroxycinnamates and about the structure dependency of the esterification yield of hydroxycinnamates with different lipases are needed. The aim of this study was to assess the influence of the structure of hydroxycinnamic acid derivatives on the enzymatic esterification with ethanol by R. miehei lipase, and to evaluate their transesterification efficacy to sterols catalyzed by C. rugosa lipase. The synthesized 99 Part B: Esterification of hydroxycinnamic acids and their antioxidant activity products were analyzed for their antioxidant capacity as hydroxycinnamic acids, as well as their stearyl and steryl esters. 2 Materials and Methods 2.1. Chemicals All solvents used were of HPLC grade or of higher purity. All hydroxycinnamic acid derivatives (ferulic acid ≥ 99%, caffeic acid ≥ 95%, sinapic acid ≥ 98%, p-coumaric acid ≥ 98%, m-coumaric acid 99%, o-coumaric acid 97%, phloretic acid 98%, cinnamic acid ≥99%, hydrocinnamic acid 99%), the α-tocopherol ≥ 96%, pyrogallol (puriss.), the DPPH (2,2-diphenyl-1-picrylhydrazyl), and Tween® 20 were purchased from Sigma-Aldrich, Buchs, Switzerland. Methyl caffeate, methyl ferulate 99% and ethyl ferulate 99% were obtained from Alfa Aesar, Karlsruhe, Germany. β-Sitosterol ≥ 70% (main impurities: campesterol and β- sitostanol) was purchased from Sigma-Aldrich, Switzerland. γ-Oryzanol was from Wako Pure Chemical Industries, Osaka, Japan. Methyl linoleate > 99% was purchased from Nu-Chek Prep, Elysian, MN. 2.2 Enzymes The lipases were purchased from Sigma-Aldrich, Buchs, Switzerland, namely lipase from Rhizomucor miehei (formerly known as Mucor miehei) immobilized on macroporous ion- exchange resin, >30 U/g (1 U sets free 1 μmol stearic acid at pH 8.0 and 70 °C per minute), lipase from Candida rugosa type VII, ≥11.7 U/mg (at pH 7.2 and 37 °C 1 U will hydrolyze 1.0 microequivalent of fatty acid from a triglyceride per minute), and Lipase A from Candida antarctica immobilized on Immobead 150, recombinant from Aspergillus oryzae ≥500 U/g (1 U corresponds to the amount of enzyme, which liberates 1 μmol butyric acid per minute at pH 10.0 and 40°C). 2.3 Esterification of hydroxycinnamic acids with ethanol The esterification of the hydroxycinnamic acid derivatives (Figure 1) was performed in a similar manner as published earlier for ferulic acid (Schär & Nyström, 2015). The hydroxycinnamic acid and the immobilized R. miehei lipase were mixed with hexane and ethanol in a glass tube with a Teflon-lined screw cap, and the samples were incubated in an oil bath for selected times. The esterification reaction conditions (for details see Table 1) for all other hydroxycinnamic acid derivatives were optimized using surface response methodology similarly as previously reported for ferulic acid (Schär & Nyström, 2015). The temperature was kept from the previous study at 61°C and a fixed time (72 h) was chosen. 100 Part B: Esterification of hydroxycinnamic acids and their antioxidant activity Therefore, a three-factors-three-levels Box-Behnken design was carried out for each of the hydroxycinnamic acids (data not shown). After the ethylation, the hexane and ethanol were evaporated under a stream of nitrogen at 50°C. Hexane was added and evaporated a second time to ensure full ethanol evaporation. Figure 1: Structural formulas of hydroxycinnamic acid derivatives used in this study: ferulic acid (a), caffeic acid (b), sinapic acid (c), p-coumaric acid (d), m-coumaric acid (e), o-coumaric acid (f), phloretic acid, (g) hydrocinnamic acid (h), and cinnamic acid (j). R may correspond to either R=H (free acid), R=CH3 (methyl ester), R=CH2CH3 (ethyl ester), R=(CH2)17CH3 (stearyl ester) or R=Rsteryl (steryl ester). 2.4 Esterification of hydroxycinnamic acids with stearyl alcohol The hydroxycinnamic acids were directly esterified with stearyl alcohol to C18 esters using the immobilized lipase from R. miehei as described earlier (Schär & Nyström, 2015). Incubation took place as described above. For all hydroxycinnamic acids similar conditions were applied, namely 14.6 mM hydroxycinnamic acid, 0.38 M stearyl alcohol, 21.5 mg/3 mL of enzyme in hexane for 72 h at 61°C. Caffeic acid was not directly esterified but transesterified from methyl caffeate to the stearyl alcohol. A base-acid work-up was used for purification (Hakala et al., 2002). for which the hexane was evaporated under a stream of nitrogen and 400 µL of the sample were redissolved in 16 mL methanol. After the addition of 1.33 mL 1.2% aqueous KOH, remaining free alcohol was extracted six times with each 12.8 mL hexane. Afterwards, the methanol phase was acidified by the addition of 1.6 mL 6 M HCl and the hydroxycinnamic acid esters were extracted three times with 12.8 mL hexane. Absence of free hydroxycinnamic acids was confirmed by RP-HPLC as described below and a reduced concentration of free alcohol was observed by NP-HPLC and RI detection (Luna 101 Part B: Esterification of hydroxycinnamic acids and their antioxidant activity HILIC column (Phenomex, Torrance, CA), hexane and isopropanol (99:1) at isocratic conditions, 0.5 mL/min). 2.5 Transesterification of hydroxycinnamic acids with sterols Transesterification of hydroxycinnamic acids was achieved using the product from the ethylation reaction. The residue after evaporation was used directly for the transesterification reaction as published earlier for the ferulic acid (Schär & Nyström, 2016). β-Sitosterol, C. rugosa lipase and solvents were added to the ethyl hydroxycinnamates followed by incubation in an oil bath. The reaction conditions varied for the different hydroxycinnamic acids (for details see Table 2). For low yielding transesterification reactions, small optimizations were performed such as the addition of butanone (5-20%). Phloretic acid was transesterified in a similar manner as published earlier (Panpipat et al., 2013). The steryl esters applied in the antioxidant assay, namely steryl ferulate and steryl sinapate, were directly esterified as previously reported (Schär & Nyström, 2016). The purification was achieved with a base-acid work-up as described for the C18 esters. Again purity was confirmed by RP-HPLC to ensure absence of free phenolic acids. 2.6 HPLC Analyses and quantification of hydroxycinnamates Samples of esterification reactions, after purification and for antioxidant assays were analyzed by RP-HPLC as described earlier (Schär & Nyström, 2015). The Solvent was evaporated and the sample redissolved in solvent B composed of acetonitrile, water, n- butanol, acetic acid in a ratio of 88:6:4:2. The HPLC was equipped with a xBridgeTM Phenyl column (Waters) with a particle size of 3.5 µm at room temperature. The detection was achieved at 325 nm or 280 nm with a diode array detector (DAD). A gradient of solvent A (1% acetic acid in water) and solvent B was applied: 3 min linear gradient from 75:25 (A:B) to 100% B, isocratic flow of 100% B for 11 min, 4 min linear gradient to 75:25 (A:B) and 2 min isocratic flow 75:25 (A:B) at a flow rate of 0.6 mL/min. For the detection of only free and ethylated hydroxycinnamic acids the isocratic flow of solvent B was shortened to 2 min. For the quantification external calibration (0.05-13 nmol/injection) of the free hydroxycinnamic acid was used and also applied for esterified hydroxycinnamates (Schär & Nyström, 2015). Previously, similar response for ferulic acid and ferulate esters was shown, which allows for the use of a single calibration curve for the hydroxycinnamic acid and its esters. Similar behavior was also confirmed for caffeic acid and methyl caffeate as well as cinnamic acid and ethyl cinnamate and thus later only a single calibration curve was applied for each hydroxycinnamic acid and its conjugates. Identification was supported by the specific UV spectra of the hydroxycinnamic acids. The identity of the products applied for the antioxidant assays were verified by detection of the expected mass in a UPLC-MS system applying the 102 Part B: Esterification of hydroxycinnamic acids and their antioxidant activity conditions as published earlier (Zhu & Nyström, 2015). Further, also the masses of steryl esters of o-coumaric acid, m-coumaric acid, and phloretic acid were confirmed the same way. 2.7 DPPH radical scavenging activity assay Solutions of all antioxidants of 1 mM and 3 mM concentration were prepared in either methanol or ethyl acetate. The antioxidant solution was added (25 µL) to 1.475 mL of a DPPH solution (0.045 mg/L) in a cuvette, making final antioxidant concentrations of 16.7 µM and 50 µM. The absorbance at 517 nm was recorded for 10 min for the methanol and 60 min for the ethyl acetate solutions using a Cary 100 UV-Vis spectrophotometer (Agilent Technologies, Basel, Switzerland). A 4 mM solution of pyrogallol was used as positive control and its scavenging activity was set to 100%. The radical scavenging activity (RSA%) in percent was calculated as following: RSA% = (A0-At)/(A0-Ap)*100, where A0 corresponds to the absorbance before the addition of antioxidant, At to the absorbance after 10 min or 60 min for methanol and ethyl acetate, respectively, Ap represents the absorbance at the end of the pyrogallol measurement. Samples were analyzed in triplicate and results are presented as mean with standard deviation in parentheses. For all antioxidant assays γ- oryzanol was used as control for commercially available steryl ferulates and α-tocopherol as positive control. 2.8 Antioxidant activity in bulk methyl linoleate Antioxidant activity measurements in bulk and emulsified methyl linoleate (including HPLC analyses of hydroperoxides) were adapted from a previous study (Nyström et al., 2005). The water content of methyl linoleate substrate was analyzed in quadruplicate by Karl Fischer titration (784 KFP Titrino, Metrohm, Herisau, Schweiz). An aliquot of 100 µL of an antioxidant solution (10 mM in acetone) was added to 1 g of methyl linoleate in a 4 mL glass vial (15 mm diameter). For control samples pure acetone was applied. After the solvent was evaporated at 40°C under a stream of nitrogen, a final antioxidant concentration of 1 µmol/g was reached. The open vials were oxidized in a dark oven at 40°C. Oxidation was monitored by measuring the formation of hydroperoxides with NP-HPLC. For this purpose 50 mg aliquots were diluted with hexane in a 5 mL volumetric flask at suitable intervals. For the percentage of inhibition the sample was compared to the control without antioxidant addition at the same time point. Results are presented as means of triplicates. 2.9 Antioxidant activity in emulsified methyl linoleate Methyl linoleate (0.5 g) was weighed into a falcon tube and 50 µL of 10 mM antioxidant solution was added. The solvent was evaporated under a stream of nitrogen at 50°C before 103 Part B: Esterification of hydroxycinnamic acids and their antioxidant activity adding 50 mg of Tween 20 in 4.45 mL water. The mixture was emulsified by sonication (UP200s, Hielscher, Teltow, Germany) (3x 30s) in an ice bath. Droplet size was analyzed using laser diffraction (Beckman Coulter, California, USA) and volumetric median was found to be X50,3=0.56 µm before and X50,3=0.87 µm after incubation of 11 days. After homogenization, the emulsions were transferred into 25 mL glass vials with screw caps and oxidized in a dark oven at 40°C with moderate stirring by magnetic bars. Again oxidation was monitored by analyzing hydroperoxides by HPLC. Aliquots of 500 mg were weighed into a test tube, and 2 mL of methanol and a few drops of aqueous saturated sodium chloride solution were added. Lipids were extracted by three times with 2 mL of hexane. All extracts were combined and diluted to 10 mL in a volumetric flask. Dry sodium sulfate was added before filtration for HPLC analysis. Antioxidant activity in emulsion was measured in triplicate. 2.10 HPLC determination of hydroperoxides The methyl linoleate hydroperoxides (methyl-13-hydroperoxy-cis-9-trans-11- octadecadienoate, methyl-13-hydroperoxy-trans-9-trans-11-octadecadienoate, methyl-9- hydroperoxy-cis-10-trans-12-octadecadienoate, and methyl-9-hydroperoxy-trans-10-trans- 12-octadecadienoate) were analyzed by HPLC (Agilent technologies 1200 series equipped with a SupelcosilTM LC-SI column from Supelco, 5 µm particle sice, and dimensions of 250 mm x 2.1 mm). Detection was achieved using a DAD with a wavelength of 234 nm. The mobile phase consisted of 12% diethyl ether in hexane with a flow rate of 0.4 mL/min. With every batch an in house reference sample (mixture of hydroperoxides from methyl linoleate) was analyzed to ensure consistency of the chromatographic system. Results are presented as sum of the areas of the four hydroperoxides peaks. 2.11 Statistical analysis Statistical analysis of the DPPH radical scavenging activity and the bulk methyl linoleate oxidation inhibition was performed using SPSS version 22. One-way analysis of variance was used and a significance level of p<0.05 between groups was accepted as statistically different. As homogeneity of variance between groups was not given, comparisons of the means were performed using the Games-Howell post hoc Test. 104 Part B: Esterification of hydroxycinnamic acids and their antioxidant activity 3. Results and Discussion 3.1 Esterification of cinnamic acid derivatives by R. miehei lipase The esterification of the hydroxycinnamic acid was optimized for each acid individually. The yield of ethyl sinapate in comparison with ethyl ferulate was slightly lower, also the amount of immobilized enzyme needed was little higher (enzyme-to-substrate ratio (m/m) of 3.1 instead of 2.5) (Table 1). Thus, the second methoxy group does not influence the enzymatic esterification strongly. On the other hand, esterification yield was drastically lower for caffeic acid (yield 16.1%), which has a second hydroxyl group in meta-position, instead of a methoxy group. This is in accordance to previous published work, where immobilized R. miehei lipase was employed in ionic liquid to esterify hydroxycinnamic acids with octanol (Katsoura et al., 2009). In this earlier study, the yields of octyl ferulate and octyl sinapate were similar, whereas the yield of octyl caffeate was significantly lower. Also in solvent-free reaction system with 1-octanol, ferulic acid was esterified more efficiently than caffeic acid by immobilized R. miehei lipase (Stamatis et al., 1999). Table 1: Molar yields of the enzymatic esterifications of various hydroxycinnamic acid derivatives with ethanol using R. miehei lipase at 61°C. The hydroxycinnamic acid ethylations were optimized and optimal conditions are listed. Results are presented as a mean of triplicate analysis with standard deviation in parentheses. Hydroxy- Enzyme/ Hexane Ethan Butanon Time Yield cinnamic substrate [µL] ol [µL] e [µL] [h] [%] acid [mg] [mg/mg] Ferulic acid 3.7 2.5 2950 50 0 72 76.2 (2.0)a Caffeic acid 2.5 7 2575 75 350 72 16.1 (1.2) Sinapic acid 2.5 3.12 2935 65 0 72 66.7 ( 1.4 ) m-Coumaric acid 2.5 2.52 2968 32 0 72 69.0 ( 1.7 ) o-Coumaric acid 2.5 4 2963 37 0 72 60.6 ( 0.6 ) p-Coumaric acid 2.5 3.72 2950 50 0 72 61.4 ( 1.4 ) Phloretic acid 3.3 2.61 2975 25 0 8 97.3 ( 2.3 ) a: (Schär & Nyström, 2015) When comparing the esterification yields of the coumaric acids (Table 1), m-coumaric acid was esterified most efficiently: not only was the yield higher, but also the amount of enzyme needed to reach this yield was lower compared to p-coumaric acid and o-coumaric acid. This is again in accordance to the previously published solvent-free esterification with 1-octanol by immobilized R. miehei lipase, where the yield for m-coumaric acid was also the highest amongst the coumarates (Stamatis et al., 1999). 105 Part B: Esterification of hydroxycinnamic acids and their antioxidant activity Phloretic acid with a saturated side chain and a hydroxyl group in para-position was esterified much faster compared to the hydroxycinnamic acid derivatives with an unsaturated side chain (Table 1). After optimization the reaction time was reduced to 8 h where almost full conversion was measured. Esterification of cinnamic acid and hydrocinnamic acid was also evaluated applying the same conditions as for ferulic acid, without further optimization. However, the variation in the yield measured was very high, thus no values are published here. This variation might be a consequence of a factor, such as possibly the water content, which influenced these reactions strongly and could not be controlled fully. It has been described earlier that cinnamic acid is esterified faster by immobilized R. miehei lipase than p-coumaric acid or ferulic acid in ionic liquid (Katsoura et al., 2009), or in solvent- free system (Stamatis et al., 1999). It is generally considered that a combination of a para- hydroxyl group and an unsaturated side chain in hydroxycinnamic acids leads to a decreased yield of enzymatic esterification by lipases (Guyot et al., 1997; Stamatis et al., 1999). This was confirmed again in the present study; however, the ortho- and para-hydroxyl group had a similar impact on the yield. In fact significant increase in reaction speed was measured when the side chain of the substrate was saturated. Overall, all hydroxycinnamic acid derivatives could be enzymatically ethylated using the immobilized lipase from R. miehei, although with significant differences observed in yield and reaction time. 3.2. Transesterification of hydroxycinnamic acid derivatives with sitosterol Transesterification of ferulic acid was optimized systematically in an earlier study (Schär & Nyström, 2016), and the process was slightly adjusted by the addition of some butanone or slight changes of the substrate concentrations for other hydroxycinnamic acids to improve the yield. Overall, the yield for the steryl ferulate was the highest with almost 55% (Table 2). Ethyl sinapate was also transesterified quite efficiently by C. rugosa lipase to steryl sinapate (31.1%). Interestingly, of the three coumaric acids m-coumaric acid and o-coumaric acid were transesterified to the according steryl ester in similar efficiency (18.8% and 18.7%, respectively), but p-coumaric acid was transesterified not to a quantifiable extent. For the m- and o-coumaric acid, addition of some butanone increased the yield from below 10% to almost 19%, compared to the ferulic acid where it only decreased the yield (Schär & Nyström, 2016). Commercial methyl caffeate had to be used as starting material for the transesterification of caffeic acid. However, the yield of the steryl caffeate was very low. Earlier only the transesterification of vinyl caffeate using C. rugosa lipase with sterols has been applied, also leading to a better purified yield for steryl ferulate (90%) and steryl sinapate (80%) than steryl caffeate (50%) (Tan & Shahidi, 2011, 2012, 2013). 106 Part B: Esterification of hydroxycinnamic acids and their antioxidant activity Table 2: Molar yields of transesterification reactions of ethyl hydroxycinnamates with sitosterol using C. rugosa lipase with the following conditions: Sitosterol (11 mg/3 mL) was incubated with ethyl hydroxycinnamate (molar ratio of substrates was ethyl hydroxycinnamate/sitosterol = 2.5 (mol/mol)) at 63°C for 120 h in hexane with an enzyme loading of 3 mg/mg (enzyme/sitosterol), or as described differently below. Results are presented as average of triplicate analysis with standard deviation in parentheses. Hydroxycinnamic acid Yield [%] derivative Ferulic acid 54.9 (2.5)d Sinapic acid 31.1 (2.5) m-Coumaric acida 18.8 (2.0) o-Coumaric acida 18.7 (0.6) p-Coumaric acid >LOQ Caffeic acidb >LOQ Phloretic acidc 21.3 (0.7) a: 10% butanone b: 5 mg methyl caffeate, 5 mg sitosterol and 10 mg C. rugosa lipase were incubated for 120 h at 63°C in 1.5 mL hexane including 10 % butanone. c: The synthesis of steryl phloretate was achieved by incubation of 15 mg ethyl phloretate, 18.4 mg sitosterol, 36.8 mg C. antarctica lipase A in 5 mL hexane for 96 h at 50°C. d:(Schär & Nyström, 2016) >LOQ: Below limit of quantification Phloretic acid, which is considered as a rather simple substrate for the esterification, was not transesterified by C. rugosa lipase to a measurable extent. But applying the C. antarctica lipase A in similar conditions as published earlier (Panpipat et al., 2013), lead to a yield of 21.3% of steryl phloretate (Table 2). It has been stated before that the double bond in the side chain improves the yield, for transesterification of vinyl phenolates with sterols by C. rugosa lipase (Wang et al., 2015). In another study it has been shown that in solvent-free system p-coumaric acid was esterified more efficiently to 1-octanol by C. rugosa lipase, compared to ferulic acid (Stamatis et al., 2001). However, this is not in agreement with the observations in this study, where it appears that the 3-methoxy group is of high importance for the C. rugosa lipase to accept the hydroxycinnamic acid as substrate. The yield decreases drastically from ferulic acid (55%) to p-coumaric acid (below quantification limit). Interestingly the o-coumaric acid was transesterified better than the p-coumaric acid. This indicates that the low reactivity of the phenolic acids with the hydroxyl group in para-position is rather due to steric hindrance than electron donating effects. 107 Part B: Esterification of hydroxycinnamic acids and their antioxidant activity 3.3 Radical scavenging activity DPPH radical scavenging activity was tested in two different solvents at two concentrations for caffeic acid, sinapic acid, ferulic acid and p-coumaric acid and their C18 and steryl esters, excluding steryl caffeate and steryl p-coumarate, which were not obtained in sufficient amounts due to very low yields. α-Tocopherol was used as positive control and γ-oryzanol served as control for commercially available steryl ferulates. p-Coumaric acid and its C18 ester showed hardly any DPPH-radical scavenging activity, but for other compounds significant activities were measured (Table 3). The control α-tocopherol showed equal activity in methanol and in ethyl acetate, but for hydroxycinnamic acids and their derivatives the values are lower in ethyl acetate than in methanol. From the higher concentration employed for caffeic acid and its C18 ester no clear tendency can be seen as the values are all close to 100%. However, for the lower caffeates concentration in methanol a higher DPPH radical scavenging activity for the C18 ester was observed compared to its free acid, whereas no difference in ethyl acetate was measured. For the sinapic acid the results showed a different trend. In methanol for the free acid a higher activity was measured at both concentrations. On the other hand in ethyl acetate the values were similar for the sinapates at the lower concentration, but at the higher concentration the free acid was less active. The ferulates in methanol showed similar behavior, the free acid was also more active. However, in ethyl acetate the radical scavenging activity of steryl ferulate was higher than that of γ-oryzanol, which served as control for steryl ferulates. This is the only point where a difference between steryl ferulate and γ-oryzanol has been measured, which is still a topic under discussion. Earlier studies reported both, there are indications for differences in the antioxidant activity between individual steryl ferulates (Nyström et al., 2005; Winkler-Moser et al., 2015), as well as studies reporting no differences (Xu & Godber, 2001). It has been shown earlier that the solvent can influence the DPPH radical scavenging activity for protocatechuic acid (3,4- dihydroxybenzoic acid) and its esters (Saito et al., 2004). For example in acetone, DPPH radical scavenging activity was similar for the free acid and its short chain esters, compared to the activity measured in methanol, where the opposite was observed (Saito et al., 2004). This was also the case for the lower concentration tested here. The antioxidant activity of the free hydroxycinnamic acid was different in methanol (higher for sinapic acid and ferulic acid and lower for caffeic acid) and the same in ethyl acetate compared to their esters. Kikuzaki and colleagues measured the DPPH radical scavenging activity of ferulic acid and its esters in ethanol (Kikuzaki et al., 2002). The activity for free ferulic acid was also found to be higher than the radical scavenging activity of the alkyl ferulates. In an earlier study comparing the DPPH radical scavenging activity of the free acids and their sterol ester in ethanol, a higher activity was found for steryl caffeate, but a lower activity for steryl sinapate compared to the 108 Part B: Esterification of hydroxycinnamic acids and their antioxidant activity corresponding free acid (Tan & Shahidi, 2014). To conclude, the type of solvent influences the DPPH radical scavenging activity for esterified and free hydroxycinnamic acids. Based on these experiments p-coumaric acid and its C18 ester were excluded from further experiments in methyl linoleate systems, as they essentially showed no radical scavenging activity at tested concentrations. Table 3: DPPH-radical scavenging activity of hydroxycinnamic acids and their esters at two concentration levels in methanol and in ethyl acetate. Pyrogallol (66.67 µM final concentration) was used as a reference for 100% activity. RSA % = (A0 – At)/(A0 – AP), At = Absorbance after 10 min for methanol, absorbance after 60 min for ethyl acetate, A0 = DPPH blank, mean of triplicate analysis, standard deviation in parenthesis. RSA [%] in methanol RSA [%] in ethyl acetate Antioxidant 16.67 µM 50 µM 16.67 µM 50 µM Caffeic acid 41.1 (2.0) f 97.6 (1.1) g 38.3 (0.9) d 91.9 (0.2) f C18-Caffeate 59.5 (0.6) g 100.0 (0.6) g 38.0 (1.0) d 97.2 (0.0) g Sinapic acid 31.9 (0.2) e 74.0 (2.1) f 18.1 (0.2) c 36.5 (0.1) cd C18-Sinapate 19.9 (0.7) cb 50.7 (0.3) b 15.6 (0.6) c 47.4 (0.4) e Steryl sinapate 19.1 (0.5) cb 64.9 (5.4) bcdef 17.6 (0.4) c 48.2 (2.7) de Ferulic acid 26.4 (0.5) d 58.1 (0.6) e 10.5 (0.8) b 28.9 (1.3) bc C18-Ferulate 20.4 (0.2) c 46.5 (0.6) c 9.9 (0.7) b 23.9 (0.5) b Steryl ferulate 17.8 (0.2) b 41.8 (0.5) d 10.4 (0.1) b 38.4 (0.7) d γ-Oryzanol 21.5 (0.6) c 42.2 (1.9) bcd 11.5 (0.3) b 23.8 (0.3) b p-Coumaric acid 3.5 (0.4) a 5.7 (0.3) a 2.3 (0.6) a 3.0 (0.4) a C18-p-Coumarate 2.3 (0.5) a 1.7 (0.7) a 2.3 (0.3) a 2.5 (0.7) a α-Tocopherol 38.8 (1.8) f 100.2 (0.5) g 34.3 (1.4) d 92.1 (0.0) f Values within a column followed by the same letter are not significantly different (p< 0.05). 3.4. Antioxidant activity in bulk methyl linoleate The increase in methyl linoleate hydroperoxides was followed over 60 days (Figure 2) and inhibition thereof calculated after 10 days (Table 4). γ-Oryzanol was used as control for commercially available steryl ferulates, α-tocopherol as positive control and a blank without any antioxidant as negative control. A water content of 0.03% was measured in the methyl linoleate, indicating presence of interfaces also in the bulk oil. The control without any antioxidant oxidized from the very beginning. The group of samples, which could retard oxidation only slightly, is composed of all ferulates being free ferulic acid, C18-ferulate, steryl ferulate and γ-oryzanol. The differences between free ferulic acid and its esters are small. On the other hand, in bulk methyl linoleate the C18 sinapate and steryl sinapate retarded oxidation significantly less compared to the free sinapic acid. The caffeic acid and the C18 109 Part B: Esterification of hydroxycinnamic acids and their antioxidant activity caffeate were able to inhibit oxidation very strongly and no increase in peroxides could be determined over the full experimental period. Table 4: Percentages of oxidation inhibition determined by hydroperoxides formation in bulk methyl linoleate after 10 days of incubation at 40°C. Concentrations of antioxidants were 1 µmole per gram methyl linoleate and results are presented as mean of triplicate analysis with standard deviation in parenthesis. Antioxidant Inhibition (10 days) [%] Caffeic acid 98.7 (0.1) g C18-Caffeate 98.2 (0.1) f Sinapic acid 98.0 (0.1) f C18-Sinapate 92.1 (0.2) d Steryl sinapate 91.2 (0.1) c Ferulic acid 73.1 (1.6) b C18-Ferulate 70.2 (1.3) b Steryl ferulate 64.5 (1.1) a γ-oryzanol 69.6 (0.2) ab α-Tocopherol 95.9 (0.0) e Values followed by the same letter are not significantly different (p< 0.05). Following the polar paradox, the more polar free phenolic acids would have a higher antioxidant activity in this bulk methyl linoleate. This was the case for the sinapates. For the caffeates no conclusion can be drawn, as no formation of hydroperoxides was detected in both caffeate samples. For the ferulates the only significant difference was that the steryl ferulate was significantly lower (64.5%) than the ferulic acid and the C18 ferulate (73.1% and 70.2% inhibition after 10 days, respectively). Similar antioxidant activities for free ferulic acid and steryl ferulates has been observed earlier for lower antioxidant concentrations in bulk methyl linoleate (Nyström et al., 2005). Only at the higher concentration the free ferulic acid showed stronger antioxidant activity. The concentration of antioxidants applied in this study (1 µmol/g) is between the two concentrations applied earlier (0.52 mM - 2.58 mM) (Nyström et al., 2005). However, formation of hydroperoxides was retarded only little, which may not be enough to show the effect of the antioxidant paradox. Overall the antioxidant activity measurement in bulk methyl linoleate reflects the data from the DPPH radical scavenging activity regarding the order of caffeates being the strongest antioxidants, followed by the sinapates and the ferulates. 110 Part B: Esterification of hydroxycinnamic acids and their antioxidant activity Figure 2: Formation of hydroperoxides during antioxidant activity assay in bulk methyl linoleate at 40°C. The concentration of all antioxidants is 1 µmol/g. Means of triplicate analysis are presented. Figure 3: Formation of hydroperoxides during antioxidant activity assay in emulsified methyl linoleate at 40°C. The concentration of all antioxidants refers to 1 µmol per gram methyl linoleate. Means of triplicate analyses are presented, except the time points above 200 h where only duplicate analysis was performed. 111 Part B: Esterification of hydroxycinnamic acids and their antioxidant activity 3.5 Antioxidant activity in emulsified methyl linoleate For the antioxidant activity in emulsified methyl linoleate the same controls and antioxidants as for the bulk methyl linoleate were applied. Formation of hydroperoxides was again followed over time (Figure 3). In general, the free phenolic acids could not retard the oxidation in comparison to the control sample without any antioxidant added. The nonpolar ferulates could inhibit oxidation only very little. Surprisingly, the C18 ester of caffeic acid and the steryl sinapate follow a similar trend. The C18 ester of sinapic acid was most efficient in retarding oxidation of all the hydroxycinnamates applied. The noteworthy fact is the large difference between the steryl sinapate and the C18 sinapate. In an emulsified system it could be expected that the polar free hydroxycinnamic acids only have little to no antioxidant effect, as they are probably mainly located in the water phase as measured earlier for chlorogenic acid (Laguerre et al., 2009). In the same study Laguerre and co-workers found a decreasing antioxidant activity if the chain length was too high. For C18 and C20 esters of chlorogenic acid a decreased antioxidant capacity and an increase of chlorogenic acid esters in the water phase could be measured, probably due to formation of aggregates with the emulsifier (Laguerre et al., 2009). The different type of emulsifier and hydroxycinnamic acid could lead to the fact that the C18 ester of sinapic acid is better located in the system than the sterol ester and therefore exhibits better antioxidant activity. Overall the nonpolar antioxidants were more efficient in the emulsified system with the C18 sinapate showing the highest activity. 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Journal of the American Oil Chemists Society, 78(6), 645-649. Zhu, D., & Nyström, L. (2015). Differentiation of rice varieties using small bioactive lipids as markers. European Journal of Lipid Science and Technology, 117(10), 1578-1588. 115 Part B: Hydrolysis of alkyl ferulates by feruloyl esterases Hydrolysis of nonpolar n-alkyl ferulates by feruloyl esterases Aline Schär, Isabel Sprecher, Evangelos Topakas, Craig B. Faulds and Laura Nyström Submitted manuscript (June 2016). Abstract Ferulic acid is one of the major phenolic acids in plants and can be found esterified to plant cell wall components, but also as long-chain n-alkyl and steryl esters. Microbial feruloyl esterases may play a role in the bioavailability of phenolic acids during human and animal digestion. It is therefore of interest if feruloyl esterases are capable of hydrolyzing nonpolar ferulic acid esters. A series of n-alkyl ferulates with increasing lipophilicity were enzymatically synthesized and the kinetic constants of their hydrolysis by four feruloyl esterases and a lipase as control were determined. A decrease in Km and kcat could be observed with decreased substrate polarity for all the feruloyl esterases. Only one feruloyl esterase and the control lipase showed hydrolytic activity towards octadecyl ferulate. These results led to the conclusion that lipophilic ferulates are poor substrates for known feruloyl esterases and more specific esterases/lipases need to be identified. Keywords: Feruloyl esterase / Alkyl ferulates / A. niger feruloyl esterase / C. thermocellum feruloyl esterase / R. miehei lipase / Ferulic acid Highlights: Kinetics of four feruloyl esterases with five alkyl ferulates were determined. Km decreases with increasing lipophilicity of the substrate. Octadecyl ferulate was hydrolyzed by only one feruloyl esterase. R. miehei lipase can hydrolyze alkyl ferulates and is thus a suitable control. 117 Part B: Hydrolysis of alkyl ferulates by feruloyl esterases 1. Introduction In plant tissues, ferulic acid is one of the most abundant hydroxycinnamic acids (Faulds & Williamson, 1999). The phenolic acids in plants occur as soluble free acids, soluble conjugated phenolic acids, and as insoluble bound phenolic acids (Li et al., 2008). In wheat for instance the major group is the insoluble bound form, which is composed of phenolic acids bound to insoluble cell wall components (Adom et al., 2005), such as arabinoxylan or pectin (Benoit et al., 2008). The soluble conjugated phenolates, like the nonpolar alkyl ferulates, are covalently bound to low-molecular weight components, and can be analyzed through extraction and hydrolysis afterwards (Li et al., 2008). Prominent examples are steryl ferulates, where the phenolic acid is esterified to a plant sterol, which can be found for example in cereal grains, such as rice, wheat, and corn (Mandak & Nyström, 2012). In addition to steryl ferulates, also other nonpolar alkyl ferulates can be found in suberin waxes, a non-polymeric extract of low polarity from suberized tissues (Graça, 2010). Ferulic acid esters of 1-alkanols in suberin waxes are long-chain (C16-C30) and mostly possess even- number of carbons in the alkyl chain (Bernards, 2002; Graça, 2010). A summary of the occurrence of alkyl hydroxycinnamate in plants has been published recently (He et al., 2015). Furthermore, these compounds are known for their antioxidant activity, which is dependent on the chain length and the type of hydroxycinnamic acid (Sorensen et al., 2014). Overall, phenolic acids can be found esterified to various compounds with very different properties. Feruloyl esterases have a significant impact on plant processing by not only improving the bioavailability of phytonutrients, but also by optimizing the saccharification of cereal derived raw materials for feed and bioalcohol production (Faulds, 2010). It has been shown that esterases extracted from human intestinal mucosa are capable of hydrolyzing esters of dietary hydroxycinnamic acids (Andreasen et al., 2001). Further, a feruloyl esterase has been extracted and characterized also from a typical human intestinal bacterium Lactobacillus acidophilus (Wang et al., 2004), and esterases with hydroxycinnamates- hydrolyzing activity characterized from intestinal Eschericia coli, Bifidobacterium lactis and Lactobacillus gasseri (Couteau et al., 2001). The substrate specificity of feruloyl esterases is therefore of interest for a broad range of areas including the human digestion of plant materials containing phenolic acid esters. Feruloyl esterases can be classified into at least four groups, as suggested by Crepin and co-workers (Crepin et al., 2004). Their activity on different hydroxycinnamic acid methyl esters, the capability to release 5,5′-diferulic acid from various substrates, and amino acid sequence similarities are key criteria for this grouping. The feruloyl esterase from Aspergillus 118 Part B: Hydrolysis of alkyl ferulates by feruloyl esterases niger (AnFaeA) is a typical representative of a Type-A feruloyl esterase, showing preference for methyl hydroxycinnamates with methoxy groups on the aromatic ring, such as ferulic and sinapic acid (Faulds & Williamson, 1994; Kroon et al., 1997). Further, AnFaeA shows structural similarities to lipases (Hermoso et al., 2004). However, AnFaeA did not show lipase activity on olive oil triglycerides and very little hydrolytic activity on diglycerides (Aliwan et al., 1999). Type-B feruloyl esterases, such as the one from Myceliophthora thermophila (Topakas et al., 2012), on the other hand prefer methyl hydroxycinnamates with one or two hydroxyl groups such as p-coumaric acid or caffeic acid and show only very low to no activity against methyl sinapate (Crepin et al., 2004). In addition, the type of sugar, the length of oligosaccharide chain and the location of the ester link between the acid and the sugar has a strong impact on the specificity of feruloyl esterases (Faulds et al., 1995). Thus, feruloyl esterases of different classes may show strongly varying activities towards a range of substrates. Apart from methyl hydroxycinnamates, methyl esters of various phenylalkanoic and cinnamic acids have also been evaluated as substrates for feruloyl esterases (Kroon et al., 1997; Topakas et al., 2005; Vafiadi et al., 2006). While the influence of the acid moiety of the substrate on the feruloyl esterase activity has been studied several times, there are less studies available related to the effect of alcohol moiety on the enzyme activity. For two type-C and one type-B feruloyl esterases short-chain alkyl ester substrates up to butyl ferulate were evaluated (Moukouli et al., 2008; Topakas et al., 2012; Vafiadi et al., 2006; Vafiadi et al., 2005), but for more lipophilic substrates the data is scarce. For example, the activity of type-A feruloyl esterase from A. awamori against α-naphthyl esters was evaluated and no activity was detected for acids longer than eight carbon atoms such as caprylic acid (Koseki et al., 2005). However, the chain length of the fatty acid was varied and the alcohol α-naphthol remained the same. Enzymatic activity of feruloyl esterases on lipophilic substrates is further influenced by co-solvents (Faulds et al., 2011). For AnFaeA the activity towards methyl ferulate decreased to around 60% if the buffer solution contained 5% DMSO (v/v). On the other hand for the substrate p-nitrophenyl acetate the activity increased to almost 180% by the addition of 5% DMSO. Therefore, for water insoluble substrates a treatment with 10-30% DMSO was proposed beneficial to the activity of feruloyl esterases (Faulds et al., 2011). Consequently it is of interest if feruloyl esterases can also hydrolyze nonpolar n-alkyl ferulates, but this question has until now not been systematically evaluated for chain lengths longer than four. To approach this problem a series of n-alkyl ferulates with increasing 119 Part B: Hydrolysis of alkyl ferulates by feruloyl esterases lipophilicity were synthesized and evaluated as substrates for four types of feruloyl esterases and one lipase as control. 2. Materials and Methods 2.1 Chemicals Ferulic acid (≥99%), MOPS (3-(N-morpholino)propanesulfonic acid, ≥99.5%) and MES (2-(N- morpholino)ethanesulfonic acid, ≥99%) were obtained from Sigma-Aldrich, Buchs, Switzerland. Methyl ferulate (99%) and ethyl ferulate (98%) were purchased from Alfa Aesar, Germany. γ-Oryzanol was obtained from Wako Pure Chemical Industries, Osaka, Japan. All solvents used were of HPLC grade or of higher purity. 2.2 Enzymes Lipozyme® RM IM was provided by Novozymes A/S, Bagsvaerd, Denmark. Feruloyl esterases from rumen microorganism, ROFae (600 U/mL where 1 U corresponds to 1 µmol ferulic acid released from ethyl ferulate per minute at pH 6.5 and 40°C) and from XynZ domain of Clostridium thermocellum, CtFae (10 U/mL where 1 U corresponds to 1 µmol ferulic acid released from ethyl ferulate per minute at pH 6 and 50°C) were obtained from Megazyme, Bray, Ireland. Recombinant feruloyl esterase type-A from A. niger, AnFaeA, was produced according to Juge and co-workers (Juge et al., 2001). The lyophilized enzyme was redissolved in buffer (MOPS, pH 6). The type-B feruloyl esterase from Myceliophthora thermophila, MtFaeB, was prepared according to Topakas et al. without the chromatographic purification (Topakas et al., 2012). Lipase from Rhizomucor miehei (≥20000 U/g) was purchased from Sigma-Aldrich, Buchs, Switzerland. Protein contents of enzyme preparations were analyzed by Bradford assay using Bradford reagent from Sigma-Aldrich, Buchs, Switzerland and bovine serum albumin as standard. 2.3 Preparation of n-alkyl ferulates Propyl, hexyl, decyl and octadecyl ferulates (Figure 1) were enzymatically esterified using Lipozyme® RM IM as published earlier (Schär & Nyström, 2015). To remove the ferulic acid from the propyl ferulate, the reaction mixture in n-hexane was washed with water. After evaporation of the unreacted propanol and the solvent n-hexane at 50°C, the propyl ferulate product was redissolved in acetone and ready for hydrolytic reactions. The other ferulates were purified by a base-acid wash adapted from Hakala and co-workers (Hakala et al., 120 Part B: Hydrolysis of alkyl ferulates by feruloyl esterases 2002). In this procedure, n-hexane was evaporated and 100 µL of the remaining alcohol including the ferulic acid and the n-alkyl ferulate were redissolved in 4 mL of methanol. After the addition of 666 µL of 0.6% KOH (0.6% (v/v) aqueous saturated KOH diluted in water) the methanol was washed ten times with 3.2 mL n-hexane to remove the unreacted alcohol. Finally, the methanol phase was acidified with 400 µL 6 M aqueous hydrochloric acid and the n-alkyl ferulates were extracted five times with 3.2 mL n-hexane. For the octadecyl ferulate the following minor changes in the base-acid wash were conducted: 333 µL of 1.2% KOH, only five times washing of the basic methanol and the whole procedure was performed twice. Products were analyzed by NP-HPLC (Luna HILIC column from Phenomex, USA, isocratic flow of hexane and isopropanol (99:1) at 0.5 mL/min) equipped with a refractive index detector (RID) to control the removal of the free alcohol. Figure 1: Structural formula of ferulic acid esters. For the enzymatic esterification n corresponds to 2, 5, 9 or 17 and for the hydrolysis by feruloyl esterases n equals 0, 1, 2, 5, 9 or 17. 2.4 Hydrolysis of n-alkyl ferulates by feruloyl esterases An aliquot of a solution of n-alkyl ferulates in acetone was transferred into a glass tube and the solvent was removed under a stream of nitrogen at 50°C. The volume of substrate solution in acetone was calculated based on the amount needed for the hydrolysis experiments in accordance to the concentration determined, as described below. First the DMSO was added followed by the buffer to reach the total reaction volume, final concentrations were 5% DMSO, 1 mM MOPS or 5 mM MES buffer and varying n-alkyl ferulate concentrations. The reactions with AnFaeA and MtFaeB were conducted at pH 6 with MES buffer and the others (lipase, CtFae, ROFae) with MOPS buffer at pH 7. Concentrations of n-alkyl ferulates ranged from 3.5 µM to 6 mM, depending on the enzyme, and final protein concentrations were 1.5 nM, 0.6 nM, 35.2 nM, 0.9 nM, and 3.7 µM for AnFaeA, MtFaeB, CtFae, ROFae, and lipase, respectively. For each enzyme and substrate six or more different substrate concentrations were analyzed in triplicates. The sample was preheated in a water bath at 40°C before the enzyme was added to start the hydrolytic 121 Part B: Hydrolysis of alkyl ferulates by feruloyl esterases reaction. After 15 minutes the reaction was terminated again by the addition of acetonitrile in a ratio of 1:1 to the reaction volume and filtration for HPLC analysis. 2.5 Quantification of substrates and ferulic acid by RP-HPLC and data analysis A standard substrate concentration was measured in the same way without incubation and enzyme addition to determine the substrate concentration in the acetone. The activity of the enzyme solution was periodically monitored with a standard assay based on methyl ferulate. If the activity decreased significantly a new solution was prepared. Ferulic acid and n-alkyl ferulates were quantified by RP-HPLC as published earlier (Schär & Nyström, 2015). Briefly, an xBridgeTM Phenyl column from Waters was used with a gradient elution of 1% acetic acid in water and acetonitrile, water, butanol, acetic acid in a ratio of 88:6:4:2. Calibration was achieved for all ferulates by creating one calibration curve for ferulic acid, methyl ferulate, ethyl ferulate and γ-oryzanol (0.006-2.6 nmol/injection). Kinetic constants were estimated by fitting them to Michaelis-Menten kinetics using SigmaPlot (Version 12.5 Systat Software, Inc., San Jose, CA, USA), which includes an estimation of the standard error for the calculated parameters. The used molecular masses for the calculation of kcat were the following: 30 kDa for AnFaeA (Juge et al., 2001), 39 kDa for MtFaeB (Topakas et al., 2012), 31.6 kDa for the lipase (Wu et al., 1996), and 29 kDa for CtFae and 29 kDa for ROFae, according to the provided data sheets. 3. Results and Discussion The kinetic constants using the Michaelis-Menten equation were determined for four feruloyl esterases and one control lipase using methyl, ethyl, propyl, hexyl, and decyl ferulate as substrates (Table 1). For the substrate with the longest alkyl chain, the octadecyl ferulate, no hydrolysis could be measured for AnFaeA, MtFaeB and ROFae, even if the incubation time was increased to 24h. In contrast, CtFae and the control lipase liberated ferulic acid, however the activity was too low to determine kinetic constants. Generally, Km and kcat values decreased with increasing chain length for the feruloyl esterases. Although with increasing lipophilicity of the substrate Km is decreasing stronger compared to the kcat values, the catalytic efficiency kcat/Km is increasing mainly in the case of AnFaeA and MtFaeB. For CtFae and the control lipase the pattern was not as clear. Also the coefficient of determination (R2) of the experimental data fitted to the Michaelis-Menten kinetics showed a decreasing trend with increasing chain length of the n-alkyl ferulate. 122 Part B: Hydrolysis of alkyl ferulates by feruloyl esterases The kinetic constants of the different feruloyl esterases for methyl ferulate differed quite strongly. MtFaeB and ROFae show very high affinity to methyl ferulate with Km values of 51 µM and 134 µM, respectively. On the other hand, AnFaeA and CtFae showed only low affinity towards methyl ferulate, even lower than R. miehei lipase. The kinetic constants for AnFaeA against methyl ferulate have been determined before and were found to be 780 µM, -1 -1 -1 70.74 s and 91 mM ∙s for Km, kcat and kcat/Km, respectively (Faulds et al., 2005). This Km is slightly lower than the value determined in this study, which could be a result of the 5% DMSO in the reaction system, as shown for another feruloyl esterase (Faulds et al., 2011). The turnover number measured here was quite low, which may result again from the DMSO addition, as it was shown in an earlier study for AnFaeA, where addition of 8% DMSO lead to a decrease of 50% of the original activity (Faulds et al., 2011). Moreover, the different molecular masses, which were determined earlier for AnFaeA can lead to differences in kcat values depending on the method. The molar mass determined by mass spectroscopy was 29.7 kDa, while following SDS-PAGE a molecular mass of 36 kDa was found (deVries et al., 1997). Furthermore, the kinetic constants of MtFaeB for methyl ferulate were determined -1 -1 -1 earlier and were found to be 270 µM, 6.4 s and 23.7 mM ∙s for Km, kcat and kcat/Km, respectively (Topakas et al., 2012). Comparing to that study, the turnover number obtained -1 matches quite well (8.8 s ), however Km found in this study is lower (51 µM). This difference may again result from the DMSO addition, as not all feruloyl esterases show the same effect of activity on the addition of this aprotic solvent (Faulds et al., 2011). Overall, the determined kinetic constants for methyl ferulate as substrate are in the range that could be expected based on previous results. 123 Part B: Hydrolysis of alkyl ferulates by feruloyl esterases Table 1: Kinetic constants of feruloyl esterases (type-A from A. niger (AnFaeA), type B from M. thermophila (MtFaeB), from C. thermocellum (CtFae), and from rumen microorganism (RoFae)) and the control lipase from R. miehei for different n-alkyl ferulates Methyl Ethyl Propyl Hexyl Decyl Octadecyl ferulate ferulate ferulate ferulate ferulate ferulate Km [µM] 1123 (71) 611 (60) 245 (18) 40 (3.9) 8 (2.0) n.d. k [s-1] 32.9 (1.2) 29.4 (1.2) 44.6 (1.2) 9.8 (0.3) 4.6 (0.3) cat AnFaeA k /K [mM-1∙s-1] 29 (2) 48 (5) 182 (14) 243 (25) 547 (136) cat m R2 0.996 0.985 0.989 0.948 0.709 n 9 9 11 13 11a Km [µM] 51 (3.4) 48 (2.7) 27 (1.7) 10 (0.9) >0 n.d. k [s-1] 8.8 (0.3) 11.2 (0.4) 12.1 (0.4) 8.9 (0.3) cat MtFaeB k /K [mM-1∙s-1] 173 (13) 236 (15) 452 (32) 906 (89) cat m R2 0.988 0.991 0.985 0.918 n 9 9 9 13 Km [µM] 2472 (170) 2578 (152) 1237 (358) 29 (5) 125 (27) >0 k [s-1] 8.0 (0.3) 5.7 (0.2) 3.2 (0.5) 0.2 (0.006) 0.4 (0.02) cat CtFae k /K [mM-1∙s-1] 3.2 (0.3) 2.2 (0.1) 2.6 (0.8) 5.6 (0.9) 3.2 (0.7) cat m R2 0.994 0.996 0.93 0.909 0.907 n 6 6 10 11 10 c Km [µM] 134 (17) 149 (16) 81 (8) 27 (2.5) 3.3 (0.8) n.d. k [s-1] 33.5 (4.9) 30.7 (4.5) 31.7 (4.6) 6.1 (0.9) 2.6 (0.4) cat ROFae k /K [mM-1∙s-1] 250 (48) 206 (37) 391 (68) 225 (39) 780 (213) cat m R2 0.962 0.973 0.976 0.937 0.636 n 8 8 9 13 11 1848b K [µM] 413 (79) 636 (168) 88 (25) 146 (33) >0 m (401) -1 0.002 0.004 0.022 0.006 0.010 k [s ] cat (0.0002) (0.0004) (0.0030) (0.0004) (0.0009) Lipase -1 -1 0.005 0.006 0.012 k /K [mM ∙s ] 0.07 (0.02) 0.07 (0.02) cat m (0.001) (0.002) (0.003) R2 0.941 0.939 0.979 0.811 0.894 n 7a 7a 10a 9a 8 Numbers in parentheses represent the estimated standard errors. R2 reflects the coefficient of determination between the experimental data and the calculated Michaelis-Menten kinetics. n: number of different substrate concentrations analyzed in triplicates n.d.: amount of ferulic acid released was below limit of detection >0: amount of ferulic acid released was below limit of quantification a: at one substrate concentration only duplicates were available b : Km above tested substrate concentrations c : Km below tested substrate concentrations 124 Part B: Hydrolysis of alkyl ferulates by feruloyl esterases Several trends in the kinetic constants for the different feruloyl esterases could be observed for a varied lipophilicity of the ferulate substrate. There is a trend of a decreasing Michaelis constant (Km) with increasing lipophilicity of the substrate for all tested feruloyl esterases. Furthermore, the turnover number was also shown to decrease with increasing chain length of the alcohol. For CtFae the turnover number behaves in a similar way as the Michaelis constant, which results in a rather stable catalytic efficiency with varying lipophilicity of the substrate. If kcat decreases less than Km, the catalytic efficiency increases. This was the case for ROFae, where the catalytic efficiency is around 3 times higher for decyl ferulate than for methyl ferulate. For AnFaeA, the stronger decrease in Km than in kcat is most pronounced, leading to a much higher catalytic efficiency for decyl ferulate. The kinetic constants of MtFaeB for decyl ferulate could not be determined as hydrolysis was observed, but no clear change of initial reaction rate over the measured substrate concentrations could be observed. For MtFaeB, the kinetic constants have been determined earlier for also ethyl, propyl and butyl ferulates (Topakas et al., 2012). However, due to DMSO addition comparisons are difficult between similar reaction systems, as discussed above for methyl ferulate. The lipase from R. miehei has been applied as positive control. For this lipase no clear trend within the kinetic constants concerning the lipophilicity of the substrate could be observed. The Michaelis constant and the turnover number of the lipase were at a maximum with propyl ferulate. Michaelis-Menten kinetics seemed appropriate, as low substrate concentrations and therefore monophasic conditions were applied. However, the R. miehei lipase seems to be a suitable control enzyme for the hydrolysis of n-alkyl ferulates, although its hydrolytic activity is low. For decyl ferulate, Km was higher for CtFae and for the lipase compared to the other enzymes tested. Although this would indicate lower affinity, these were the two enzymes where still some activity against octadecyl ferulate could be measured. Interestingly, the type A feruloyl esterase AnFaeA, which structurally resembles the R. miehei lipase (Faulds et al., 2005; Hermoso et al., 2004), was not able to hydrolyze octadecyl ferulate. This might be explained by the structure of AnFaeA. Although the catalytic serine is exposed to the solvent in a large cavity, the region around shows, similarly to carbohydrate-binding proteins, a highly negative electrostatic potential (Hermoso et al., 2004). Earlier it has also been shown that the catalytic efficiency of the same enzyme (AnFaeA earlier FAE-III) is generally higher for sugar esters than for methyl ferulate (Faulds et al., 1995; Ralet et al., 1994). Therefore, the findings of this study correspond well with the general idea of feruloyl esterases preferring polar ferulates. Furthermore, the coefficient of determination was very low for 125 Part B: Hydrolysis of alkyl ferulates by feruloyl esterases AnFaeA and ROFae with decyl ferulate, which is probably due to the fact, that only few samples below Km were measured. This also increases the relative error and therefore the uncertainty of the determined constants. A lower Km value for feruloyl esterases with decyl ferulate could therefore not directly be connected to a higher affinity for non-polar substrates. The Michaelis constant decreased with an increasing lipophilicity of the substrate for all tested feruloyl esterases, which could have several reasons. Firstly, as the solubility of the long-chain n-alkyl ferulates in the reaction system was very low, aggregation of substrate can be one source of error. The apparent Km in this case would rather represent the solubility of the substrate than the affinity of the enzyme to the substrate, because above the limit of solubility the substrate in solution would stay constant, even if the substrate amount would be increased. However, since the Michaelis constants determined in this study for decyl ferulate were quite different between the enzymes ranging from 3.3 to 146 μM, this factor can be excluded. Secondly, a more pronounced decrease in Km with increasing lipophilicity compared to kcat indicates a reduced k-1 (rate constant for dissociation of enzyme-substrate complex) or an increased k1 (rate constant for formation of enzyme-substrate complex) for more hydrophobic substrates. This could lead to the hypothesis that a decreasing Km with increasing lipophilicity of the substrate is not only an indication for the specificity to the enzyme, but also reflects the solubility of the substrate in the aqueous system. The substrate undergoes desolvation when binding to the enzyme, which is energetically more favored for less soluble substrates (Klibanov, 1997; Zeuner et al., 2012). Accordingly, the reverse process (k-1 ) is less favored. In this case, the declining Km may therefore be misleading, concerning the specificity of feruloyl esterases. On a mechanistic basis feruloyl esterases show similarities. All feruloyl esterases evaluated in this study, except ROFae, have been shown to have a catalytic triad in the active site (Hermoso et al., 2004; Schubot et al., 2001; Topakas et al., 2012), as well as the lipase (Derewenda et al., 1992). Therefore, a covalent enzyme-acyl intermediate is formed during the hydrolysis. Identical catalytic rate constants can result from a common acyl-enzyme intermediate and a rate limiting deacylation (Zerner et al., 1964). As the acyl group was always ferulic acid, the catalytic rate should always be similar if the deacylation is rate limiting. However, this was often only the case for short-chain ferulic acid esters. Examples are ROFae and AnFaeA where similar kcat for methyl, ethyl and propyl ferulates were measured, while a decrease in rate constant was observed for longer chains. In this case, the rate limiting step probably shifted partially or fully to the formation of the acyl-enzyme complex, which could be explained by a less suitable position of the long-chain ester for the nucleophilic attack of the catalytic serine. However, as the feruloyl esterases are structurally 126 Part B: Hydrolysis of alkyl ferulates by feruloyl esterases very different one would have to study the interaction of the nonpolar substrate in more detail individually. Overall this supports the hypothesis that long-chain n-alkyl ferulates are poor substrates for feruloyl esterases. A systematic evaluation of the activity of feruloyl esterases from different classes on nonpolar n-alkyl ferulates was carried out to evaluate if microbial feruloyl esterases are capable of hydrolyzing naturally occurring n-alkyl ferulates. This led to the conclusion that for feruloyl esterases, nonpolar ferulic acid esters such as long-chain n-alkyl ferulates are very poor substrates. Only very little or no activity was determined for octadecyl ferulate. This conclusion is supported by earlier studies, which showed no activity of a feruloyl esterase against olive oil triglycerides or in a second study against long-chain (>C10) α-naphthyl esters. Further evaluations of more feruloyl esterases would support this conclusion. Finally, studies using biological samples containing long-chain n-alkyl ferulates would be of interest to evaluate the in vivo activity in a more complex environment. 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Kinetic Evidence for Formation of Acyl-Enzyme Intermediates in -Chymotrypsin-Catalyzed Hydrolyses of Specfic Substrates. Journal of the American Chemical Society, 86(18), 3674-3679. Zeuner, B., Kontogeorgis, G. M., Riisager, A., & Meyer, A. S. (2012). Thermodynamically based solvent design for enzymatic saccharide acylation with hydroxycinnamic acids in non- conventional media. New biotechnology, 29(3), 255-270. 130 Conclusion and Outlook Conclusion This study shows that the esterification of hydroxycinnamic acids, mainly ferulic acid, can be achieved in an n-hexane system using the immobilized lipase from R. miehei as catalyst. The reaction system was optimized yielding, after 72 h of incubation, 76% and 88% of ethyl ferulate and decyl ferulate, respectively. The optimal conditions estimated by surface response methodology mainly differ in the amount of ferulic acid and alcohol, which could be set higher for the decyl ferulate synthesis. Based on the optimal conditions for the model compounds ethyl and decyl ferulate, other linear alcohols from C3 to C18 were esterified with ferulic acid. The yield increased from C2-C6 up to 92% and did not significantly change for the longer alcohols. The secondary alcohols isopropanol and 2-octanol reacted only to a little extent catalyzed by R. miehei lipase, which probably reflects the 1,3-specificity of the lipase. Moreover, in a mixture of primary alcohols, the ones shorter than C6 reacted significantly faster compared to the longer ones. Overall, this developed esterification method for ferulic acid provides the possibility to efficiently apply ferulic acid in multiphase systems as antioxidant. Also, standards for the analysis of biological samples can be produced with this method. As a second achievement the fully enzymatic synthesis of steryl ferulates was investigated. The two optimized systems were the direct esterification and the transesterification from ethyl ferulate yielding 35% and 55% steryl ferulates, respectively. In combination with the method discussed above, this leads to a fully enzymatic two-step synthesis of steryl ferulates. Both systems seem promising, although the yield of the transesterification is higher. However, the sterol concentration of the direct esterification system can be set higher and the purification is more straightforward. Therefore, both systems can be applied and give a basis for further development of this enzymatic synthesis. Overall, the main achievement is that vinyl ferulate, which often requires a heavy metal catalyst in the synthesis, can be avoided. In a third study different hydroxycinnamic acid derivatives were evaluated as substrates for the R. miehei and C. rugosa lipases. The activity profile towards hydroxycinnamic acid derivatives for the two lipases was very different. For the R. miehei lipase the yield increased when the side chain was saturated and decreased if two phenolic hydroxyl groups were present. On the other hand, for the C. rugosa lipase the yield decreased if there was a hydroxyl group in para-position without a neighboring methoxy group. If the side chain is saturated the yield rather decreases as well. The ethylations catalyzed by R. miehei lipase were optimized individually. Yields above 60% for all tested hydroxycinnamic acids were reached, except for ethyl caffeate, which had a lower yield. For the steryl hydroxycinnamates 131 Conclusion and Outlook synthesis catalyzed by C. rugosa lipase, the steryl ferulates conditions were applied with slight modifications. In this case p-coumaric acid, caffeic acid and phloretic acid were hardly accepted as substrates and yields were therefore not measurable. In general, the yields of the steryl hydroxycinnamates syntheses were rather small and the steryl ferulates conditions could not be easily transferred to other hydroxycinnamic acids. The antioxidant activities of some synthesized alkyl and steryl hydroxycinnamates were evaluated in three systems, namely in DPPH radical scavenging activity, bulk methyl linoleate and emulsified methyl linoleate. The radical scavenging activities of hydroxycinnamic acids and their esters depend on the solvent. It is therefore important to actively decide, which solvent suits best for the application of interest. In bulk methyl linoleate the free acids showed highest antioxidant activity, according to the polar paradox. In the emulsified methyl linoleate the C18 sinapate showed superior activity to the steryl sinapate. This could be due to the cutoff effect, which would need further investigation with other sinapate esters in the same system. Overall, the antioxidant activity of hydroxycinnamates depends on the system of application. In the last study the synthesized alkyl ferulates were evaluated as substrates for feruloyl esterases. Especially for the long chain, nonpolar ferulates very little or no activity was measured. Only the feruloyl esterase from C. thermocellum and the control lipase showed hydrolytic activity towards octadecyl ferulate. It can be assumed that naturally occurring alkyl ferulates are not hydrolyzed by feruloyl esterases and rather lipase are responsible for this reaction. On the whole, the conducted studies provide methods for simple enzymatic synthesis of analytical standards and of substrates for further studies, including antioxidant assays for the alkyl ferulates or animal and cell studies for the steryl hydroxycinnamates. However, further improvements are required, especially for the steryl hydroxycinnamates synthesis to increase the yield and therefore the capacity. Outlook The products of the enzymatic alkyl hydroxycinnamates synthesis can be used as standards for further analysis of biological samples on their alkyl hydroxycinnamate content and profile. Of special interest are food products, which have been already shown to contain steryl ferulates or other steryl hydroxycinnamates. Furthermore, it would be interesting to focus on the distribution within the plant, and in particular during growth, to investigate possible links 132 Conclusion and Outlook between steryl hydroxycinnamates and alkyl hydroxycinnamates. As a totally different application, a more thorough understanding of the so-called cutoff effect could be achieved with the alkyl hydroxycinnamates. Factors such as the surfactant type and concentration, antioxidant concentration, or oil phase properties could be investigated. The enzymatic synthesis of steryl hydroxycinnamates may also be applied for the synthesis of standards. Uncommon sterols or phenolic acids can be used as substrates to produce internal standards. However, for further optimization of the enzymatic process, the C. rugosa lipase should be optimized first. The initial step would be to test the single isoenzymes of C. rugosa lipase. The most efficient isoenzyme should then be expressed as recombinant, to be able to produce the pure isoenzyme more easily. In case of unsatisfying yields or efficiencies, immobilization or even enzyme engineering could be tried. By modelling the substrate-enzyme interaction, an optimized amino acid sequence could be determined and adjusted recombinant enzymes could be produced. By doing so, the non-universal codon for serine of C. rugosa should be taken into account. The synthesized steryl hydroxycinnamates could be used to improve research on these interesting compounds, reaching an official health claim would further increase the interest on steryl hydroxycinnamates. Concerning the use of nonpolar substrates for feruloyl esterases, the evaluation of more feruloyl esterases would be of interest, with particular attention on the still missing groups. Furthermore, their activity on biological samples could be analyzed to gain data in a more complex environment. Samples containing long-chain alkyl ferulates could be treated with feruloyl esterases and the concentration thereof monitored over time. Also, fungi degrading such samples could be applied to evaluate if the long chain ferulates are hydrolyzed. Moreover, the synthetic activity of feruloyl esterases would be of interest, in particular if they are able to esterify ferulic acid with nonpolar alcohols. For this purpose, microemulsion systems or enzyme immobilization would have to be applied. 133 Acknowledgements This thesis was only achieved with the help and support of some people, which I would like to acknowledge here. Further, financial support was provided by the Swiss National Science Foundation, SNSF (project 200021_141268) and ETH Zurich. Without Prof. Dr. Laura Nyström this thesis would not exist. She introduced me to scientific research and woke my fascination to work on a topic in a depth like this. The good teamwork convinced me to start and also finalize my thesis with her. Thank you for always being available for my questions and my concerns; and for letting me enough freedom to fulfill my own ideas and to develop myself. I further thank Dr. Pierre Villeneuve for accepting to be a co-examiner of this thesis. A special thank goes to Prof. Dr. Evangelos Topakas for also being a co-examiner and for hosting me during a visit in his laboratory in 2013. You introduced me to a more biotechnological perspective of enzyme catalysis. A very big thank you goes to Dan from the “steryl ferulates team”. We had many fruitful conversations on and off topic. Also the mass spectroscopic measurement could only be conducted with the help of her. Then I would like to thank Samy for many discussions about the chemical synthesis of steryl ferulates. Linda is acknowledged for implementing several systematic ways of working and Attila for bringing a different view on many things into the group. Thank you Marie for open my mind to sterol oxidation. I further want to thank Nadja, Elena, Melanie, Nese and all current and former members of the group for the nice working atmosphere. Acknowledged for their support in running the group and lab smoothly are Daniela, Aida and Teresa. I further thank Pascal Guillet for the Karl Fischer measurements and Nathalie Scheuble for the particle size determinations. I would also like to acknowledge my students for turning my ideas into practice and for questioning and broadening my knowledge: Francesca Molinaro, Lisa Schwarz, Lorena Taddei, Lisa Menet, Fabiola Alig, Nico Kummer, and Fabienne Michel. Especially acknowledged are Silvia Liphardt and Isabel Sprecher who also became co-authors in two of my papers. Further, I thank Diana Gongora and Savitha Gayathri for the practical help in my projects. Above all, I want to thank my parents Doris and René for their support during all my life. You showed me a life in which one should never stop learning. Last but not least I thank Leo for going with me through all the ups and downs. Thank you for commuting with me and for your understanding. 134